The role of arginine methylation of hnRNPUL1 in the DNA
damage response pathway
Gayathri Gurunathan
Faculty of Medicine
Division of Experimental Medicine
McGill University, Montreal, Quebec, Canada
August 2014
A Thesis Submitted to McGill University in Partial Fulfillment of the Requirements for the Degree of
Master of Science
© Gayathri Gurunathan 2014
Abstract
Post-translational modifications play a key role in mediating the DNA damage response
(DDR). It is well-known that serine/threonine phosphorylation is a major post-translational
modification required for the amplification of the DDR; however, less is known about the role of
other modifications, such as arginine methylation. It is known that arginine methylation of the
DDR protein, MRE11, by protein arginine methyltransferase 1 (PRMT1) is essential for the
response, as the absence of methylation of MRE11 in mice leads to hypersensitivity to DNA
damage agents. Herein, we identify hnRNPUL1 as a substrate of PRMT1 and the methylation of
hnRNPUL1 is required for DNA damage signalling. I show that several RGG/RG sequences of
hnRNPUL1 are methylated in vitro by PRMT1. Recombinant glutathione S-transferase (GST)
proteins harboring hnRNPUL1 RGRGRG, RGGRGG and a single RGG were efficient in vitro
substrates of PRMT1. Moreover, I performed mass spectrometry analysis of Flag-hnRNPUL1
and identified the same sites methylated in vivo. PRMT1-depletion using RNA interference led
to the hypomethylation of hnRNPUL1, consistent with PRMT1 being the only enzyme in vivo to
methylate these sequences. We replaced the arginines with lysine in hnRNPUL1 (FlaghnRNPUL1RK) such that this mutant protein cannot be methylated by PRMT1. Indeed FlaghnRNPUL1RK was undetected using specific dimethylarginine antibodies. Flag-hnRNPUL1RK did
not co-immunoprecipitate with PRMT1, as expected, since PRMT1 is known to associate with
its substrates. Flag-hnRNPUL1RK had reduced affinity to NBS1, a subunit of the MRE11RAD50-NBS1, a DDR complex. Finally, Flag-hnRNPUL1RK had an aberrant localization at
DNA damage breaks using laser microirradiation. Collectively, my findings provide insight into
how the arginine methylation of hnRNPUL1 plays a significant role in the DNA damage
response.
2
Sommaire
Les modifications post-traductionnelles jouent un rôle fondamental, notamment dans la
régulation des mécanismes de réponse aux dommages causés à l’ADN. Il a été démontré que la
méthylation
des
arginines
de
la
protéine
MRE11
par
l’enzyme
protéine-arginine
méthyltransférase 1 (PRMT1) est essentiel dans la voie de signalisation des dommages causés à
l’ADN. Dans cette étude, nous décrivons l’identification d’un nouveau substrat de l’enzyme
PRMT1 : la protéine hnRNPUL1.
Dans un premier temps, nous avons identifié que la déplétion de l’enzyme PRMT1 induit
une hypométhylation de la protéine hnRNPUL1. De plus la protéine hnRNPUL1 contient
plusieurs motifs RGG/RG, une séquence consensus méthylée par les PRMTs. Un essai de
méthylation in vitro a montré que plusieurs de ces séquences consensus sont méthylées par
l’enzyme PRMT1. Une analyse in vivo de la protéine Flag-hnRNPUL1 par spectrométrie de
masse a confirmée les sites de méthylation des motif RGG/RG identifiés précédemment in vitro.
Ces résultats furent supportés par la génération d’un mutant Flag-hnRNPUL1RK dépourvue
d’arginine dans les séquences RGG/RG méthylées identifiées. En utilisant un anticorps
spécifique reconnaissant les arginines diméthylées, nous avons découvert que le mutant FlaghnRNPUL1RK était hypométhylé. Nous avons également observé que le mutant ne pouvait plus
interagir avec PRMT1 par co-immunoprécipitation, contrairement à la protéine non-muté FlaghnRNPUL1. Ensemble ces résultats démontrent une association physique de la protéine
hnRNPUL1 avec PRMT1 requise pour sa méthylation directe par PRMT1 sur les résidus
arginines de ces motifs RGG/RG.
3
Nous avons ensuite découvert que le mutant hypométhylé Flag-hnRNPUL1RK présente
une affinité de liaison réduite avec la sous-unité NBS1 du complexe de recrutement MRE11RAD50-NBS1(MRN), impliqué dans la réponse aux dommages à l’ADN. En effet, nous avons
été capable de co-immunoprécipiter la sous-unité NBS1 avec la protéine Flag-hnRNPUL1, mais
pas avec son mutant Flag-hnRNPUL1RK. Finalement, la localisation de Flag-hnRNPUL1RK est
aberrante en présence aux coupures des brins d’ADN. Nous pensons que ce mécanisme est
régulé par la méthylation des motifs RGG/RG de hnRNPUL1 par PRMT1.
De façon globale, ces résultats illustrent la façon dont la méthylation des arginines de la
protéine hnRNPUL1 par l’enzyme PRMT1 joue un rôle de premier plan au niveau de la réponse
aux dommages de l’ADN produite par les coupures doubles brins.
4
Preface
This Master of Science thesis was written in accordance with the Guidelines for Thesis
Preparation from the Faculty of Graduate Studies and Research of McGill University.
The experiments and studies were conducted under the supervision of Dr.Stéphane
Richard. As well, each contributor who has collaborated in this research is acknowledged in the
following section and throughout the remainder of the text.
The mass spectrometry analysis discussed in this thesis was performed by Dr. Éric
Bonneil (Figure 6B). The laser scissor damage experiment discussed in this thesis was
performed by Yan Coulombe in Dr. Jean-Yves Masson laboratory (Université de Laval) (Figure
9A).
5
Acknowledgements
I would first like to extend my sincere gratitude and appreciation to my supervisor, Dr.
Stéphane Richard, for giving me the remarkable opportunity to work on such an interesting
project and guiding me every step of the way. His continuous support and encouragement has
truly been the propelling force that has motivated and inspired me to learn various technical
skills and explore the field without limiting myself, thereby allowing me to grow as a young
scientist and pursue my studies at McGill University in an active and dynamic environment.
I would also like to thank the members of my thesis advisory committee, who have
provided me with valuable guidance and support while monitoring my progress throughout my
studies. Their input and feedback throughout my annual committee meetings have been truly
noteworthy and have enabled me to push myself a step higher in pursuing my scientific goals
while constantly driving myself forward.
My fellow colleagues (both past and present) in the Richard lab have provided me with
immense support, guidance and care, which I am truly grateful for. Whether it is moral support
during times of need, experimental expertise when troubleshooting experiments, or simply
providing a friendly and exciting atmosphere to work in on a daily basis, I truly appreciate their
support. I would like to extend my special thanks and sincere gratitude to Dr. Zhenbao Yu, who
has provided me with continuous support in helping me develop the necessary skills required to
perform various experiments and for his valuable input in the troubleshooting aspect of my
experimental techniques. His guidance and experimental expertise has made a significant
contribution to my project and allowed for the completion of my thesis. As well, my utmost
gratitude goes out to Palaniraja Thandapani who has been extremely supportive in the
6
stimulation of scientific discussions and in providing moral support. All of my dear lab members
have been the constant driving force in improving my research strategies in all aspects and their
immense care, generosity and unconditional support has truly allowed me to acquire various
skills and values both inside and outside the professional environment which are tools that shall
be useful throughout my life. I also like to extend my thanks to the all the students, technicians
and post-doctoral fellows who I have met at the Lady Davis Institute throughout my studies, for
their insight into my project, moral support and encouragement.
I would like to extend my gratitude to Dr. Éric Bonneil for performing the mass
spectrometry analysis which has significantly contributed to my project. As well, the laser
scissor damage research provided by Yan Coulombe under the guidance of Dr. Jean-Yves
Masson has been crucial for the completion of my thesis and I would like to sincerely thank them
for their valuable contribution to my project.
I would like to acknowledge the Canadian Institutes of Health Canada (CIHR) for
providing financial support for the research presented in this thesis.
Last but not least, I would like to thank my parents and brother for being there for me and
supporting me throughout my graduate studies and schooling. I am forever grateful for the love
and care that they have provided me with. Their continuous support and motivation have always
encouraged me to chase my dreams in life.
7
Table of Contents
ABSTRACT ............................................................................................................2
SOMMAIRE ...........................................................................................................3
PREFACE ...............................................................................................................5
ACKNOWLEDGEMENTS ..................................................................................6
TABLE OF CONTENTS .......................................................................................8
LIST OF FIGURES AND TABLES ...................................................................10
LIST OF ABBREVIATIONS .............................................................................. 11
INTRODUCTION................................................................................................15
Part I: Arginine Methylation ..............................................................................15
PRMT1 ...........................................................................................................19
PRMT2 ...........................................................................................................20
PRMT3 ...........................................................................................................20
PRMT4 ...........................................................................................................21
PRMT5 ...........................................................................................................21
PRMT6 ...........................................................................................................22
PRMT7 ...........................................................................................................22
PRMT8 ...........................................................................................................22
PRMT9 ...........................................................................................................23
Part II: RNA binding proteins ............................................................................24
Part III: hnRNPs .................................................................................................26
hnRNP purification .........................................................................................26
Structural architecture of hnRNP family members.........................................27
8
Involvement of hnRNPs in a multitude of pathways .........................................30
Part IV: hnRNP U and hnRNPUL1 ....................................................................33
Part V: The DNA damage response ....................................................................37
Genotoxic stress and DNA damage ................................................................38
Types of damage and cellular response ..........................................................39
Signal transduction in the DNA damage response .........................................43
ATM pathway specifics ..................................................................................45
ATR pathway specifics ...................................................................................45
Apoptosis ........................................................................................................46
Human disorders and cancer...........................................................................47
Therapeutic applications and personalized clinical intervention ....................49
EXPERIMENTAL PROCEDURES ...................................................................52
RESULTS..............................................................................................................58
RGG/RG motifs of hnRNPUL1 are methylated by PRMT1 ..........................58
PRMT1 methylates and associates with hnRNPUL1 in vivo .........................60
Arginine methylation regulates the hnRNPUL1/NBS1 interaction................61
The methylation of the hnRNPUL1 RGG/RG motifs regulates its
recruitment at sites of DNA damage...................................…………………62
DISCUSSION .......................................................................................................70
REFERENCES .....................................................................................................77
9
List of Figures and Tables
Figure 1: The three flavours of protein arginine methyltransferases. ................................... 16
Figure 2: The nine members of the protein arginine methyltransferase family .................. 18
Figure 3: Conserved structural domains in the hnRNP family.. ............................................ 29
Figure 4: Types of damage and repair mechanisms. ............................................................... 41
Figure 5: The RGG/RG motif of hnRNPUL1 is methylated by PRMT1 in vitro.................. 65
Figure 6: hnRNPUL1 harbors methylated arginine residues identified in vivo via mass
spectrometry analysis ................................................................................................................. 66
Figure 7: The RGG/RG motif of hnRNPUL1 is methylated by PRMT1 in vivo via a
physical association ..................................................................................................................... 67
Figure 8: Methylation of hnRNPUL1 is required for its interaction with NBS1 .................. 68
Figure 9: The methylation of the hnRNPUL1 RGG/RG motifs regulates its recruitment at
sites of DNA damage ................................................................................................................... 69
Table 1: Methylated arginine residues of hnRNPUL1 identifiedin vivo via mass
spectrometry ................................................................................................................................ 64
10
List of Abbreviations
53BP1: p53 binding protein 1
AA: amino acid
AdoMet: S-Adenosyl methionine
aDMA: asymmetric dimethyl-arginine
ATCC: american type culture collection
ATM: ataxia telangiectasia mutated
ATP: adenosine-5'-triphosphate
ATR: ataxia telangiectasia and Rad3 related
BBS: BRD-7 binding site
BER: Base excision repair
BLAST: basic local alignment search tool
BLM: Bloom’s syndrome helicase
BRCA1: breast cancer 1
BRCT: BRCA1 C-terminus
BRD7: bromodomain-Containing Protein 7
CARM1: coactivator-associated arginine methyltransferase 1
CBP: nuclear cap-binding protein
CDC25A: cell division cycle 25 homolog A
CDK: cyclin-dependent kinases
cDNA: complementary DNA
CHK1: checkpoint kinase 1
CHK2: checkpoint kinase 2
Co-IP: co-immunoprecipitation
CRB2: crumbs homolog 2
CSK: cytoskeleton striping buffer
11
CtIP: CtBP-interacting protein
DAPI: 4',6-diamidino-2-phenylindole
DMEM: Dulbecco's Modified Eagle's Medium
DNA: deoxyribonucleic acid
DNA-PK: DNA-dependent protein kinase
DSB: double-strand break
E1B-AP5: adenoviral early region 1B-associated protein 5
G0 phase: quiescent state, in cell cycle context
G1 phase: cell cycle state between mitosis and DNA replication
G1/S: cell cycle state boundary between G1 phase and S phase
G2 phase: cell cycle state between DNA replication and mitosis
G2/M: cell cycle state boundary between G2 phase and M phase
GAR: glycine and arginine rich region
GFP: green fluorescent protein
GST: glutathione-S-transferase
H2A: histone H2A
H2AX: H2A histone family, member X
H2B: histone H2B
H3: histone H3
H3K4: H3 lysine 4
H4: histone H4
hnRNP: heterogeneous nuclear ribonucleoproteins
hnRNPUL1: heterogeneous nuclear ribonucleoprotein U-like 1
HR: homologous recombination
IgG: immunoglobulin G
IFN: interferon receptor
IP: immunoprecipitation
12
IPTG: isopropyl β-D-1-thiogalactopyranoside
IR: ionizing radiation
IRES: internal ribosomal entry site
IRIF: ionizing radiation induced foci
KO: knockout
LncRNA: long non coding RNA
M phase: mitosis state in cell cycle
MEF: mouse embryonic fibroblasts
MMA: monomethyl arginine
MMR: mismatch repair
MDC1: mediator of DNA damage checkpoint 1
MRE11: meiotic recombination 11
MRN: Mre11-Rad50-Nbs1
NBS1: nibrin
NCBI: national center for biotechnology information
NER: nucleotide excision repair
NHEJ: non-homologous end joining
p53: tumor protein 53
PBS: phosphate buffered saline
PCR: polymerase chain reaction
PCNA: proliferating cell nuclear antigen
PDB: protein data bank
PML: promyelocytic leukemia
PP: proline rich region
PRMT: protein arginine methyltransferase
PST: proline-serine-threonine rich
PTM: post-translational modification
13
RAD50:DNA repair protein RAD50
RAD51: DNA repair protein RAD51 homolog 1
RAD54:DNA repair and recombination protein RAD54-like
RBP: RNA binding protein
RCSB: research collaboratory for structural bioinformatics
RGG/RG: arginine glycine rich region
RNA: ribonucleic acid
ROS: reactive oxygen species
S phase: DNA synthesis state in cell cycle
SAF-A: scaffold attachment factor A
S/TQ: serine or threonine followed by glutamine site
SAM: S-adenosyl methionine
SAM68: Src-associated in mitosis 68 kDa protein
SAP: SAF-A/B Acinus and PIAS motif
sDMA: symmetric dimethyl-arginine
SDS: sodium dodecyl sulfate
SDS-PAGE: SDS- polyacrylamide gel electrophoresis
SH3: SRC homology 3
siRNA: short interfering RNA
SPRY: SPIa/Ryanodine receptor domain
ssDNA: single-stranded DNA
ORF: open reading frame
UTR: untranslated region
XRCC4: X-ray repair complementing defective repair in chinese hamster cells 4
γH2AX: phosphorylated H2AX
14
Introduction
Part I: Arginine Methylation
The first evidence of arginine methylation dates back to 1964 when the Allfrey et al.,
group demonstrated the structural modifications of histones via the addition of either acetyl or
methyl groups (Allfrey, Faulkner et al. 1964). Moreover, these two modifications (i.e. acetylation
and methylation) have been shown to take place post-translationally upon completion of the
polypeptide chain of proteins (Allfrey, Faulkner et al. 1964). Arginine is a positively charged
amino acid that is unique in its nature as it has hydrogen bonding potential: its guanidino group
has five potential hydrogen bond donors that are favorably positioned to mediate interactions
with biological hydrogen bond acceptors (Bedford and Clarke 2009). It has also been
demonstrated to engage in amino aromatic interactions with nucleic acids and proteins. Arginine
methylation is a common post-translational modification (PTM) that takes place in eukaryotic
cells, such that the arginine residues may be modified to contain methyl groups (Gary and Clarke
1998). This variation may occur in three flavours, specifically ω-NG-monomethylarginine
(MMA),
ω-NG,
NG-asymmetric
dimethylarginine
(ADMA)
andω-NG’,
NG-symmetric
dimethylarginine (SDMA) where S-adenosylmethionine (SAM) is used as the methyl group
donor (Bedford and Clarke 2009) (Atkinson and Murray 1967) (Figure 1). Furthermore, due to
the addition of the methyl groups onto the arginine residues, there are numerous implications
with respect to protein-protein interactions, both in a positive and negative manner, among
others. The bulkiness of the arginine residue increases causing greater steric hindrance leading to
decreased hydrogen bonding potential which places an increasing hydrophobic property to a
protein. Nevertheless, the positive charge of the arginine residue does not change.
15
Figure 1.The three flavours of protein arginine methyltransferases -Adapted from (Yang
and Bedford 2013)
16
This covalent post-translational modification is catalyzed by a family of intracellular enzymes
known as the protein arginine methyltransferases (PRMTs). The PRMTs are a family consisting
of nine members which have a highly conserved methyltransferase catalytic domain which is
known to oligomerize into a ring-like structure (Weiss, McBride et al. 2000) (Zhang and Cheng
2003) (Figure 2). Furthermore, the PRMTs may be classified into three categories; namely, type
I, type II or type III. Type I enzymes are known to catalyze the addition of two methyl groups to
the same terminal nitrogen of the arginine residue in order to generate ADMA and MMA
(Bedford 2007). PRMT1, PRMT3, PRMT4 (also known as CARM1), PRMT6 and PRMT8 fall
into this category. PRMT2 has been recently shown to have Type I arginine methyltransferase
activity, however, little is known about its respective substrate repertoire (Lakowski and Frankel
2009). Meanwhile, type II enzymes catalyze the formation of SDMA and MMA via the addition
of the second methyl group to the other terminal nitrogen of the arginine residue. PRMT5 and
PRMT9 classify in this listing (Bedford 2007). Finally, type III PRMTs generate exclusively
MMAs and to date only PRMT7 has been established as a member of this category (ZuritaLopez, Sandberg et al. 2012).
17
Figure 2. The nine members of the protein arginine methyltransferases family -Adapted
from (Nicholson, Chen et al. 2009, Yang and Bedford 2013)
18
PRMT1
Protein arginine methyltransferase 1 is the major arginine methyltransferase that is responsible
for catalyzing the formation of 85% of total arginine residues methylated in the mammalian cell
(Tang, Kao et al. 2000). Moreover, it is principally a nuclear protein that exhibits a preference
for the methylation of RGG/RG motifs of RNA binding proteins in addition to other proteins
(Tang, Kao et al. 2000) (Boisvert, Dery et al. 2005). It falls under the type I category of arginine
methylation flavours such that it catalyzes the formation of ω-monomethylarginine as an
intermediate and asymmetric ω-NG, NG-dimethylarginine as the product. In various tissues,
PRMT1 has been shown to have varying catalytic activities as well as distinct substrate
specificity since the PRMT1 gene has 12 coding exons which allows for the generation of 7
different isoforms via splicing (Goulet, Gauvin et al. 2007).
It has been demonstrated that PRMT1 is essential for early development via a conditional null
allele of PRMT1 in mouse models; a PRMT1 null embryo has been shown to be lethal and die at
embryonic day 6.5 (Yu, Chen et al. 2009) (Pawlak, Scherer et al. 2000). In addition, it was
observed that a conditional knockout of PRMT1 in mouse embryonic fibroblasts (MEFs) causes
spontaneous DNA damage, checkpoint activation defects, cell cycle progression defects, among
others. This further suggests a significant role for PRMT1 in the DNA damage response pathway
(Yu, Chen et al. 2009). Cell cycle defects include a reduced intra- S phase, numerous
chromosomal translocations, and an elevated hypersensitivity to chemically induced DNA
damage (Yu, Chen et al. 2009). The deregulation of PRMT1, whether it be overexpressed or
aberrantly spliced, leads to numerous types of pathologies of several diseases, mainly cancers
including breast, prostate, lung, colon, bladder cancer as well as leukemia (Goulet, Gauvin et al.
19
2007) (Seligson, Horvath et al. 2005, Le Romancer, Treilleux et al. 2008) (Cheung, Wan et al.
2007, Shia, Okumura et al. 2012) (Yoshimatsu, Toyokawa et al. 2011) (Guendel, Carpio et al.
2010, Butler, Gore et al. 2011) (Mathioudaki, Scorilas et al. 2011, Tzifi, Economopoulou et al.
2012) (Zou and Elledge 2003).
PRMT2
PRMT2 was initially identified via sequence homology to the rat protein arginine
methyltransferase PRMT1 gene (Katsanis, Yaspo et al. 1997). It has a Src homology 3 (SH3)
binding domain, which is associated with many processes (Baldwin, Morettin et al. 2014). In
particular, it has been shown to interact with hnRNPUL1, also known as the adenovirus early
protein E1B-55 kDa (Kzhyshkowska, Schutt et al. 2001). The substrate repertoire of PRMT2 has
not been fully elucidated as any methyltransferase activity has yet to be detected (Baldwin,
Morettin et al. 2014). Numerous alternatively spliced isoforms of PRMT2 have been identified
(Baldwin, Morettin et al. 2014). Also, PRMT2 has been shown to interact with a variety of
proteins including RARα, PPAR-γ, estrogen receptor α, among others (Qi, Chang et al. 2002).
PRMT3
Yeast two hybrid analysis was performed to identify PRMT3 as a rat PRMT1 interacting protein.
Although they vary in their subcellular localizations, PRMT3 shares sequence similarity with
PRMT1 and is predominantly cytoplasmic. It is known to have type I arginine methyltransferase
activity (Tang, Gary et al. 1998). The crystal structure of PRMT3 has been reported and is
established as having an Ado-Met domain as well as a barrel-like domain (Zhang, Zhou et al.
20
2000). The cone-shaped active site is localized between the two domains (Zhang, Zhou et al.
2000).
PRMT4/CARM1
PRMT4 is also known as a Co-activator-associated arginine methyltransferase 1 (CARM1) and it
was first identified as an interacting protein to GRIP1, which is the p160 steroid receptor
coactivator (Chen, Huang et al. 2000). It has type I arginine methyltransferase activity and is
involved in a multitude of cellular processes including RNA metabolism such as transcription
and pre-mRNA processing, as well as a role in the DNA damage response pathway (Baldwin,
Morettin et al. 2014). Unlike other methyltransferases in the type I category, PRMT4 has a
preference for PGM methylation motifs in substrates (Baldwin, Morettin et al. 2014). PRMT4
functions as a transcriptional coactivator by binding to p160 including estrogen responsive genes
(Chen, Huang et al. 2000) (Teyssier, Chen et al. 2002).
PRMT5
PRMT5 has type II arginine methyltransferase activity and is the predominant type II
methyltransferase in mammals (Pawlak, Scherer et al. 2000). PRMT5 is mainly
compartmentalized in the cytoplasm and is in a complex with MEP50 and pICln involved in
snRNP biogenesis (Friesen, Massenet et al. 2001) (Krapivinsky, Pu et al. 1998). PRMT5 has
been shown to interact with a wide repertoire of proteins and complexes including Rad9a, FEN1,
and p53 (Jansson, Durant et al. 2008) (Guo, Zheng et al. 2010) (He, Shi et al. 2011). Moreover,
PRMT5 KO mice demonstrate lethality between 3.5 and 6.5 embryonic days (Tee, Pardo et al.
2010).
21
PRMT6
PRMT6 was identified on chromosome 1 of the human genome as a novel methyltransferase
displaying unique functionality compared to its other type I enzyme family members, in that it
exhibited automethylation activity (Frankel, Yadav et al. 2002). It was the first PRMT to
demonstrate such activity and it was shown to be uniquely compartmentalized in the cell nucleus
as identified during its association with the green fluorescent protein (GFP) marker (Frankel,
Yadav et al. 2002).
PRMT7
PRMT7 was initially identified from a genetic suppressor elements screen in Chinese hamster
cells, where PRMT7 was presented as a gene encoding two proteins, namely, p77 and p82,
which was later renamed PRMT7α and β due to significant sequence homology with the PRMT
enzymes family (Gros, Delaporte et al. 2003). A role for PRMT7 in drug resistance has been
reported(Zheng, Schmidt-Ott et al. 2005). Although PRMT7 was initially classified as a type II
protein arginine methyltransferase, recent findings show PRMT7 is a type III enzyme due to the
generation of a monomethyl arginine, which is deemed to be an intermediate during the
methylation reactions (Zurita-Lopez, Sandberg et al. 2012).
PRMT8
PRMT8 has a particular tissue specificity in that it is anchored to the plasma membrane in the
brain and has two automethylation sites as well as a myristoylation site that are localized in the
N-terminus (Lee, Sayegh et al. 2005) (Dillon, Rust et al. 2013). Although the complete
mechanism of PRMT8, as well as its substrate repertoire are not fully established, a role for the
22
N-terminus of PRMT8 in its automethylation activity by reducing its binding affinity for
AdoMet has been reported (Dillon, Rust et al. 2013).
PRMT9
This protein arginine methyltransferase has not fully been characterized as of yet (Yang and
Bedford 2013). Nevertheless, being identified at the same time as PRMT8, its structural
architecture is similar to that of PRMT7, since it has Ado-Met binding motifs. In addition, it
contains two tetratricopeptide repeats (Lee, Sayegh et al. 2005) (Yang and Bedford 2013).
PRMTs are known to have a preference for methylation of proteins that harbor a glycine arginine
rich (RGG/RG) motif (Najbauer, Johnson et al. 1993). Nevertheless, certain protein arginine
methyltransferases such as PRMT4 (CARM1) have substrate specificity in that they do not
methylate at glycine-arginine rich regions (Lee and Bedford 2002) Instead, PRMT4 prefers
methylation at proline-glycine and methionine rich sites (Chen, Huang et al. 2000).
There are numerous substrates of PRMT1 in the cell, of which, histones consist of a major
portion. Histone 4 at arginine 3 has been shown to be dimethylated by PRMT1 and this has
numerous implications in chromatin structure and function (Strahl, Briggs et al. 2001) (Wang,
Huang et al. 2001). Moreover, other substrates of PRMT1 include RNA binding proteins,
transcription factors, and DNA damage proteins (Boisvert, Dery et al. 2005).
Many players that are involved in the DNA damage response pathway have been shown to be
methylated by PRMT1 which include the following: 53BP1, MRE11 and BRCA1 (Boisvert,
Hendzel et al. 2005, Boisvert, Rhie et al. 2005) (Guendel, Carpio et al. 2010). Furthermore, one
such example of the importance of arginine methylation is regulation of MRE11 in the DNA
23
damage response pathway. The RGG/RG motif of MRE11 is methylated by PRMT1 both in
vitro as well as in vivo (Boisvert, Hendzel et al. 2005). Arginine methylation of MRE11 is
essential for both its location to DNA damage foci as well as its exonuclease activity (Boisvert,
Dery et al. 2005).
Part II: RNA binding proteins
RNA binding proteins (RBPs) are involved in numerous aspects of RNA metabolism
whether it be to ensure the proper and effective processing, folding, stabilization and localization
of RNA, as well as correct mRNA translation (Lukong, Chang et al. 2008). Numerous RBPs
contain RGG/RG motifs making them good candidate substrates for arginine methylation.
More recently, a prominent role for RBPs in the DNA damage response in both RNA and DNA
metabolism has been reported (Dutertre, Lambert et al. 2014). RBPs regulate post-transcriptional
gene expression in a variety of ways such as binding to RNA sequences, regulatory long
noncoding RNAs (ncRNAs), and secondary structures in pre-mRNAs (Glisovic, Bachorik et al.
2008) (Lunde, Moore et al. 2007). Furthermore, upon DNA damage, there is a global decrease in
gene expression due to transcriptional inhibition via introduction of premature stop codons
resulting in nonsense mediated mRNA decay (Dutertre, Lambert et al. 2014).
RBPs have been shown to be vital mediators in the prevention of genomic instability by
inhibiting the formation of RNA and DNA hybrids, also known as R-loops. These loops, which
are prompted by G-rich DNA repeats, are a threat to genomic stability when in contact with
replication forks (Aguilera and Garcia-Muse 2012). Moreover, the mechanism by which the Rloop formation is bypassed involves packaging of pre-mRNA in the earlier stages in order to
24
prevent unprocessed transcripts from binding to the DNA template (Dutertre, Lambert et al.
2014).
In addition, a role for RBPs in localizing to the sites of DNA damage, as well as associating with
other DNA repair factors and proteins has been observed (Dutertre, Lambert et al. 2014). One
such RBP which binds to mismatched DNA and engages in the DNA mismatch repair machinery
is YB-1. (Kim, Paek et al. 2003, Gaudreault, Guay et al. 2004, Bergmann, Royer-Pokora et al.
2005). YB-1 has been reported to participate in endonuclease activity and strand separation in
vitro (Gaudreault, Guay et al. 2004) (Kim, Paek et al. 2003, Bergmann, Royer-Pokora et al.
2005).
A variety of enzymatic activities have been attributed to RBPs involved in the DNA damage
response pathway. Many of the RBPs undergo extensive post-translational modifications
including phosphorylation, ADP-ribosylation, acetylation, ubiquitination, and sumoylation
(Dutertre, Lambert et al. 2014). Such modifications are vital for effective signalling and cascade
action to take place, such that the appropriate DNA damage repair response is propagated. Major
DNA damage molecular sensors such as ATR kinase and ATM kinase are examples of
complexes which post-translationally modify the RBPs in order to elicit a choreographed DNA
repair response.
Stress responses often trigger the relocalization of numerous RBPs which also includes the
process of nuclear-cytoplasmic shuttling (Cammas, Lewis et al. 2008). This subnuclear
localization enables RBPs to take part in numerous aspects of cellular processes, thereby,
increasing efficiency and likelihood of proper cell functioning. An example of such a scenario is
the relocalization of the splicing factor Sam68 (Src-associated in mitosis) to nuclear stress
25
granules upon DNA damage (Busa, Geremia et al. 2010). In addition, a highly coordinated and
intricate network is enabled upon effective RBPs redistribution in the cell post DNA damage;
thereby, facilitating mRNA stability. Numerous changes in alternative splicing patterns take
place upon RBP relocalization and redistribution in the cell (Cammas, Lewis et al. 2008).
Part III: hnRNPs
A growing importance for hnRNPs in numerous biological events has emerged such that
gene regulation is a key and prominent cellular event among them (Piccolo, Corona et al. 2014).
hnRNP proteins are a group of multifunctional nuclear-cytoplasmic shuttling and highly
conserved family of RBPs. They play a variety of roles in a multitude of regulatory pathways
including transcription and translation regulation, telomere elongation, chromatin remodeling,
and DNA repair, among others (Carpenter, MacKay et al. 2006). hnRNP proteins have conserved
structural domains and the individual members of this family have the potential to elicit tumor
progression through their involvement in mechanisms that cause cell attack and apoptosis
prevention (Carpenter, MacKay et al. 2006).
Furthermore, hnRNPs and their respective cellular activities take place both in the nuclear and
cytoplasmic compartments; that is, processes such as pre-mRNA processing and transcription
which are initiated in the cell nucleus and shift location upon completion to the cytoplasm, in
order to take part in mRNA translation and turnover (Krecic and Swanson 1999).
hnRNP purification
Some of the earliest studies had established an effective purification method for the hnRNPs
(Pinol-Roma, Choi et al. 1988). These hnRNP complexes/particles are a combination of
26
heterogeneous nuclear RNAs (hnRNAs) which are associated with other proteins in the cell in
order to form hnRNP complexes (Dreyfuss 1986). An efficient immunoprecipitation method
allowed for the purification of hnRNP particles using monoclonal antibodies to hnRNPs (PinolRoma, Choi et al. 1988). In order to categorize and name the various components of the hnRNP,
affinity chromatography and two dimensional gel electrophoresis permitted for the effective
visualization of the various polypeptides and their corresponding molecular weights i.e. 34,000
to 120,000 daltons (Pinol-Roma, Choi et al. 1988). The approximate 30 spots identified on the
two dimensional gel allowed for the naming of the many members of the hnRNP family from
hnRNP A1 to U. Finally, the interaction between proteins and the associated RNA was
visualized using a covalent cross-linking method in the presence of UV light (Dreyfuss 1986).
Structural architecture of hnRNP family members
Extensive studies have established a significant role for hnRNPs in a multitude of regulatory
pathways in the cell, whether it be DNA regulation, RNA regulation, or many others (Carpenter,
MacKay et al. 2006). Alternative splicing patterns enable the generation of numerous isoforms of
hnRNPs. There is a conservation of numerous structural domains amongst all the members of the
hnRNP family as depicted below (Carpenter, MacKay et al. 2006) (Figure 3). Namely, the RNA
binding domain (also coined the RNA recognition motif), which is located at the N-terminus,
allows for the recognition and binding of DNA or RNA (Dreyfuss 1986). Other common
structural elements that are found in many hnRNP family members include the RGG/RG motif
as well as KH domains. Furthermore, the RGG/RG motifs span 25 amino acid sequences at a
time, with conserved Arg-Gly-Gly tripeptide repeats and play a role in protein-protein
interactions, RNA binding, and transcriptional regulation (Dreyfuss 1986). In terms of the KH
27
domain, it is ~50 amino acids in length with stretches of octapeptide repeats of Ile-Gly-X2-GlyX2-Ile where X can be any amino acid (Iwanaga, Sueoka et al. 2005). Moreover, this KH domain
was first reported in hnRNP K, where it was shown to play a role in RNA binding.
28
Figure 3- Conserved structural domains in the hnRNP family -Adapted from (Carpenter et
al., 2005)
29
Involvement of hnRNPs in a multitude of pathways
As mentioned previously, many roles for hnRNPs in numerous pathways have been reported. For
example, hnRNPs have been shown to play a role in DNA regulation such that many hnRNPs
participate in the DNA repair pathways in response to cellular damage and stress. Certain colocalization experiments demonstrated the interaction between hnRNPs and DNA repair factors;
one such example being hnRNP B1 and DNA-PK (Iwanaga, Sueoka et al. 2005). As well, a role
for hnRNPs in telomere regulation and biogenesis has been reported. The mechanism involves
hnRNPs in the recruitment of telomerases to telomeric ends of chromosomes (Ford, Wright et al.
2002)
Gene regulation needs to be tightly monitored and catered to appropriate cellular conditions in
order to propagate the most efficient signalling cascade whether it is during normal cellular
conditions or during stressful periods such as oncogenesis. Thus, there is a need for chromatin
remodeling and transcription. This being said, a role for hnRNPs during this process has been
reported. In this context, hnRNP U was reported to be involved in a direct binding interaction
with chromosomal DNA in vivo (Goehring and Morabia 1997).
hnRNPs are involved in numerous aspects of RNA metabolism as well. Firstly, in mRNA
turnover, nascent pre-mRNA requires 3`end processing and polyadenylation (Proudfoot and
O'Sullivan 2002). Effective cleavage of pre-mRNA is enhanced via hnRNPs, namely hnRNP H`,
such that cleavage stimulation factors (CstF) are recruited to the sites on the pre-mRNA in order
to mediate this process efficiently and ensure generation of mature mRNA (Bagga, Arhin et al.
1998). In such a way, many other members of the hnRNP family contribute to mRNA
biogenesis, whether it is positive or negative regulation.
30
Numerous hnRNPs have been reported to bind the internal ribosomal entry site (IRES) which
allows for the effective recruitment of the ribosome to the mRNAs; thereby, regulating the
translation of the mRNA into proteins (Kim, Paek et al. 2003). An example of this includes the
binding of hnRNP C to the IRES of the c-myc mRNA, such that the translation of the c-myc
mRNA is upregulated (Kim, Paek et al. 2003). The importance of this binding interaction
between hnRNP C within the IRES may be highlighted as a c-myc mutant mRNA which did not
contain the hnRNP C binding site demonstrated a decrease in the translation of the c-myc mRNA
to protein compared to the wildtype mRNA (Kim, Paek et al. 2003).
hnRNPs can regulate apoptosis either positively or negatively depending on their interaction
partners. Bcl-x is a protein that belongs to the apoptotic pathway and two isoforms may be
obtained via pre-mRNA alternative splicing. The two forms, Bcl-xS and Bcl-xL are a promoter
and inhibitor of apoptosis, respectively. This being said, hnRNP F and hnRNP H mediate the
selection of the isoforms via binding to the exonic regions (Garneau, Revil et al. 2005).
Furthermore, one feature of cancer cells is metastasis which requires the regulation of the
spreading initiation centre (de Hoog, Foster et al. 2004). The destruction of focal adhesion
complexes enables the cancer cells to detach from the basal membrane and relocalize itself to
other parts of the body. It has been reported that hnRNPs play a role in this process in that certain
hnRNPs, (hnRNP P2, hnRNP K, and hnRNP E1) were found in the spreading initiation centres
(de Hoog, Foster et al. 2004). Interestingly, the alteration with any of these hnRNPs triggered an
increase in metastasis. Another aspect of cancer progression includes the process of angiogenesis
whereby tumor growth is dependent on their own blood source. Factors involved in this process
include fibroblast growth factor (FGF), vascular endothelial growth factor (VEGF), among
31
others and these have been shown to be regulated by members of the hnRNP family; primarily
hnRNP L (Nyberg, Xie et al. 2005) (Shih and Claffey 1999).
Various types of cancers express different members of the hnRNP family. The expression of
hnRNP I and hnRNP C1/C2 is increased in brain and lung cancers, respectively (Carpenter,
MacKay et al. 2006). A role for hnRNPs in tumor development and progression has also been
reported. Upon literature analysis via databases, it was found that the hnRNP promoter regions;
namely, those of hnRNP A1, A2, D, F, H, H’ and K, all contained an upstream binding element
for oncogenes (Carpenter, MacKay et al. 2006). Such binding elements included E2F AP1, cmyc, and acute myeloid leukemia (AML). Thus, this phenomenon further suggests that hnRNPs
are regulated by oncogenes (Carpenter, MacKay et al. 2006). In the instance of cellular
proliferation which is a prominent feature in cancer cells, members of the hnRNP family
including hnRNP A1 and hnRNP A2/B1 have their expression regulated throughout the various
stages during cell-cycle transitioning in Colo 16 squamous carcinoma cells and HaCaT
immortalized keratinocytes (He, Brown et al. 2005). The proliferation rate in these cells was
drastically diminished upon hnRNP A1 and A2 reduction via siRNA targeting, suggesting these
genes have oncogenic potential (He, Brown et al. 2005). Another member of the hnRNP family
has been classified as a tumor suppressor, namely, hnRNP E4. Studies have demonstrated that
expression levels of hnRNP E4 are upregulated upon DNA damage in a p53-dependent manner
(Zhu, Zhang et al. 2000).
Previous studies have demonstrated that the majority of NG,NG-dimethylarginine found in
mammalian cells, approximately 65%, is enclosed by hnRNPs (Kzhyshkowska, Schutt et al.
2001).
32
Part IV: hnRNP U and hnRNPUL1
hnRNPUL1 is known as heterogeneous nuclear ribonucleoprotein U-like protein and
belongs to the hnRNP family. It contains significant sequence homology with hnRNP U, which
is also identified as scaffold attachment factor A (SAF-A) (Bode, Goetze et al. 2003). hnRNP U
has also been previously demonstrated to be an inhibitor of RNA polymerase II elongation (Kim,
Paek et al. 2003).
hnRNPUL1 was first identified as adenoviral early region 1B-associated protein 5 (E1B-AP5)
since it was known to associate with the adenovirus early protein E1B-55kDa (Ad5EE1B55K)
during lytic infection (Gabler, Schutt et al. 1998). Viral growth and propagation depends on the
accumulation of viral mRNA in the cytoplasm. This recruitment of mRNA relies on the binding
interaction between E1B-55 kDa protein
and
hnRNPUL1
(Gabler, Schutt
et
al.
1998).hnRNPUL1 has been reported to act on both viral and cellular promoters, with
overexpression causing adenoviral mRNA export while host cell mRNA shuffling is shut down
in lytically infected cells (Gabler, Schutt et al. 1998). Alongside this previously mentioned role,
hnRNPUL1 has been suggested to act as a cofactor when it functions as a regulator of gene
expression.
hnRNP U (SAF-A) is known to be multi-functionally involved in both DNA and RNA binding
with a specific affinity for scaffold associated DNA regions which are known to be sequestered
with A-T amino acids (Xiao, Tang et al. 2012) (Ahn and Whitby 2003). In addition, hnRNP U is
a key structural component involved in chromatin modeling and organization and has been
shown to be methylated by PRMT1 in its RGG/RG motif; a particular site of interest spanning
amino acids 778 to 793, where there are four RG rich sites (Herrmann, Bossert et al. 2004).
33
Moreover, hnRNP U is ubiquitously methylated and this may be seen as constitutive because the
unmethylated form of hnRNP U is rapidly degraded post translation. Methylated proteins in cells
may be recycled and reused in order to maintain a constant pool of resources for the newly
synthesized gene products during various developmental and cellular processes (Herrmann,
Bossert et al. 2004).
hnRNP U has been shown to be a basic transcriptional inhibitor (Kzhyshkowska, Rusch et al.
2003). Studies demonstrated roles in regulating transcription of hormone receptors such as the
glucocorticoid receptor (GR), which is stimulated via ligand binding (Eggert, Michel et al.
1997). The inhibition of transcriptional activation may be seen when hnRNP U physically
interacts with the glucocorticoid receptor. Overexpression experiments of hnRNP U hinders the
transcription and may function to sequester and store intranuclear glucocorticoid receptors
(Eggert, Michel et al. 1997).
As well, another protein that was shown to co-localize with hnRNPUL1 and physically associate
is the bromodomain (BD)-containing protein, BRD7 (Kzhyshkowska, Rusch et al. 2003). It has
been demonstrated that the interaction between BRD7 and hnRNPUL1 is essential in order for
hnRNPUL1 to maintain its characteristic as a transcriptional activator. Deletion of the region
where BRD7 binds to hnRNPUL1 changes hnRNPUL1 from an activator to a strong
transcriptional repressor of GR-responsive promoters. Removing the RGG region of hnRNPUL1
results in modification of its repression activity, but it was shown that the N-terminal of
hnRNPUL1 was responsible for this repression. In addition, hnRNPUL1 has been shown to
interact with the TAP protein on its N-terminus, which allows for the regulation of RNA nuclear
export substrates and its nuclear pore complexes (Kzhyshkowska, Rusch et al. 2003).
34
An adenovirus’ replication cycle may be divided into two stages based on viral DNA replication;
namely, early and late phase. During the early stage of adenovirus replication, viral DNA
replication takes place while, the late stage of replication is marked by cellular protein synthesis
shutdown since the host cell mRNAs have been inhibited (Schenk, Lieberburg et al. 1996). The
late phase of the adenovirus infection may be characterized by the favored production of viral
proteins resulting in changes to gene expression profiles in terms of mRNA transport as well as
protein biogenesis (Dobner and Kzhyshkowska 2001).
Previous studies demonstrated that the methylation of hnRNPUL1 changes during the later phase
of the adenovirus lytic infection. In particular, a late adenoviral product, L4-100 kDa gets
methylated by PRMT1 (Kzhyshkowska, Kremmer et al. 2004). An in vitro methylation assay
confirmed the methylation of this late viral product, which is 100 kDa in molecular weight.
Many functions have been attributed to this product such as RNA binding as well as assistance in
increasing the translation efficiency of the late viral transcripts (Hayes, Telling et al. 1990,
Andrade, Bull et al. 2001). Various kinds of cells generate unique methylation profiles for L4100 kDa during lytic infection. Nevertheless, the enzymatic activity of numerous enzymes of the
protein arginine methyltransferase family play a role as all methylation profiles generated
throughout adenoviral lytic infections are not generated by just a singular enzyme
(Kzhyshkowska, Kremmer et al. 2004).
PARP1 is a protein that has a key role in the maintenance of genomic stability (Hong, Jiang et al.
2013). Once DSBs arise, PARP1 in addition many protein kinases such as ATM and ATR, bind
to the double stranded breaks and mediate the signal cascade to effectively repair the damaged
DNA. PARP1 has also been shown to associate with the MRE11 subunit of the MRN molecular
35
sensor complex in order to get recruited to the DNA damage sites (Haince, McDonald et al.
2008). PARP1 poly(ADP-ribosyl)ates itself and other proteins to contribute to the DNA damage
response (Schreiber, Dantzer et al. 2006) (Jagtap and Szabo 2005). hnRNPUL1 is one among the
many proteins to which PARP1 binds and recruits to the DNA damage sites via poly(ADPribosyl)ation. It was shown that upon hnRNPUL1 depletion, PARP1 expression levels were
lessened suggesting a role for hnRNPUL1 in PARP1 expression (Hong, Jiang et al. 2013). A
newer role for hnRNPUL1 was identified as a regulator of the ATR pathways during lytic
adenoviral infection, in that, hnRNPUL1 plays a role at the viral replication centres (Blackford,
Bruton et al. 2008). hnRNPUL1 has also shown to co-localize with ATRIP, which is an ATRinteracting protein as well as RPA 32, which is an ATR substrate replication protein A 32.
hnRNPUL1 has been shown to play a role in cancer progression such that it was reported to
interact with the tumor suppressor p53 and have an inhibitory effect on its transcription (Barral,
Rusch et al. 2005).
hnRNPUL1 contains a SAP domain which is known as the DNA/RNA binding domain and is
involved in higher order chromosomal organization. It also has a SPRY domain which is
implicated in various protein-protein interactions. Its other domains include BBS, which are the
BRD7 binding site, RGG/RG domain as well as some lower complexity regions.
Initial findings have shown that hnRNPUL1 is methylated in its RGG rich box, located in the Cterminal part of the protein, via PRMT1 in vitro, which is responsible for regulating proteinRNA interactions (Kzhyshkowska, Schutt et al. 2001). These methylation sites have yet to be
mapped, however, PRMT1 was not responsible for the methylation of hnRNPUL1 in vivo.
Further experiments demonstrated that hnRNPUL1 and PRMT2 occurs interacted in vivo in
36
order to methylate hnRNPUL1 in its RGG box domain (Kzhyshkowska, Schutt et al. 2001). This
was seen via a yeast-two hybrid system where the interaction between hnRNPUL1 and PRMT2
was occurring through hnRNPUL1’s SH3 domain. The SH3 domain has been shown to be a
significant player in modulating the stability of PRMT2. hnRNPUL1 also contains a proline-rich
sequence right after the RGG motif. This stretch of proline amino acids have been known to be
involved in protein-protein interactions (Kzhyshkowska, Schutt et al. 2001).
A role for hnRNPUL1 in the DNA damage pathway has been identified (Polo, Blackford et al.
2012). It was shown to be required for double strand break resection and ATR signalling. It was
proposed that there are two pools of hnRNPUL1 in the cell, such that one pool is involved in
RNA metabolism while the other participates in DSB pair (Polo, Blackford et al. 2012). The
MRN complex was seen as a key player in this process as the localization of hnRNPUL1 to sites
of DSBs required the presence of MRN complex. Furthermore, depletion of hnRNPUL1
displayed defective ATR signalling and recruitment of ATRIP to the damage sites, while ATM
signalling remained unchanged. There was a deficiency in RPA accumulation as well as ssDNA
formation in the hnRNPUL1-depleted cells. Further experiments demonstrated that hnRNPUL1
proteins function in a pathway that is downstream of MRN and CtIP, however, upstream of BLM
such that hnRNPUL1 proteins promote the DSB resection by regulating BLM recruitment. The
RGG motif of hnRNPUL1 is essential for its mobilization to and from the sites of DNA damage,
in a MRN dependent and RNA independent manner (Polo, Blackford et al. 2012).
Part V: The DNA damage response
The ultimate objective of propelling life forward relies on the successful transmission of
accurate and intact genetic material to progeny cells from one generation to the next
37
(Kerzendorfer and O’Driscoll 2009). Thus, in order to ensure organisms’ survival and fitness in
the next generation, a faithful surveillance of the genetic information must be maintained. Many
endogenous and exogenous agents place genotoxic stress on DNA such that genomic integrity is
constantly challenged (Jackson and Bartek 2009). Therefore, precision and accuracy in DNA
duplication, chromosome distribution as well as responding to DNA damage issues are of main
concern. Genetic control of cell-cycle transitions along with the evolution of cellular systems that
monitor DNA damage have allowed for normal human development in hope of maximizing
genomic fidelity while minimizing the propagation of heritable mutations to daughter cells
(Jackson and Bartek 2009). Pioneering work has established that the DNA damage response
(DDR) pathway is a multifaceted and intermeshed network which consists of DNA damage
detection, signal initiation and activation of DNA repair mechanisms (Harper and Elledge 2007).
Failure to repair defective DNA and bypass of cellular senescence may result in a transmission
of damaged DNA and lethal mutations to progeny. Genomic instability increases the likelihood
for the development of numerous human illnesses (Huen and Chen 2010).
Genotoxic stress and DNA damage
The highly reactive nature of DNA exposes itself as a prime target for a plethora of genotoxic
agents. The number of modifications that occur to human genomic DNA in a single day from
genotoxic stress surpasses 100,000 (Rich, Allen et al. 2000). A persistent source of exogenous
DNA damage is ultraviolet light (UV). UV-A and UV-B are parts of the solar UV spectrum that
are present in sunlight and they have a harming potential of approximately 100,000 lesions per
exposed cell per hour (Jackson and Bartek 2009). Apart from ultraviolet radiation, other
environmental agents that cause DNA damage include tobacco products, industrial substances,
38
warfare chemicals (ex: mustard gas) among other drugs that are used in cancer treatment and
chemotherapy. The latter play a large role in causing DNA-damage as the ailments used in
cancer therapies initiate covalent crosslinks among DNA bases (ex: carboplatin and cisplatin),
drugs that add alkyl groups to DNA bases (ex: temozolomide), and those that sequester
topoisomerase I or II enzymes onto the DNA to create single (SSB) or double strand breaks
(DSB) (Lord and Ashworth 2012). In addition, endogenous sources of DNA damage include
cellular metabolic processes that produce reactive oxygen species (ROS) that oxidize DNA bases
and generate SSBs. To avoid such damage, DNA is embedded in chromatin proteins which
provide some DNA damage protection. However, during essential cellular processes such as
DNA replication and transcription, chromatin rearrangement occurs in such a way that DNA
becomes exposed to damaging agents; hence, becoming more vulnerable (Rich, Allen et al.
2000). During replication, DSBs may be generated due to stalled replication forks and collisions
further increasing DNA’s proneness to damage. This being said, ionization radiation (IR) is also
another means of DSBs, which is the most toxic and lethal form of DNA damage seeking
attention from the DDR pathway (Finn, Lowndes et al. 2012).
Types of damage and cellular response
Firstly, the DNA damage response pathway is a complex network of signal transduction that
allows for interaction among the various pathway molecules which include sensors, transducers
and effectors in order to maximize cell survival. Once the damage is recognized, it is assessed
via checkpoints and either repair and/or apoptosis take place accordingly. The cell-cycle
checkpoint arrest provides the cell with some time to repair the DNA damage, however, if the
appropriate response is implemented yet the repair is unsuccessful, then either cellular
39
senescence or apoptosis takes place to prevent the transmission of damaged genetic information
to daughter cells during cell division (Huen and Chen 2010). Checkpoints are vital players in the
DDR as they work to preserve genomic integrity by serving as barriers that halt the
dissemination of defective genomes. This process is essential as a lack of quality control may
result in embryonic lethality (due to heightened sensitivity to DNA-damaging mediators) and
genomic instability which are hallmark features of cancer (Rich, Allen et al. 2000).
40
Figure 4: Types of damage and repair mechanisms - Adapted from (Zhou and Elledge
2000)
41
The presence of multiple DNA- repair mechanisms may be attributed to the fact that DNA repair
is tailored to the type of DNA lesion that has occurred (Hakem 2008). Figure 4 depicts an
overview of the various repair mechanisms based on the type of lesion, the corresponding stage
in the cell cycle, and their mode of action. The different types of damage include SSBs, DSBs,
bulky adducts, base mismatches and base alkylation. As well, the various repair mechanisms
include base excision repair, homologous recombination and non-homologous end joining (both
for DSBs), nucleotide excision repair, mismatch repair and direct reversal. The major proteins
that play a role in the DDR pathway associated with each kind of break is shown, resulting in
particular tumors if these respective DDR proteins are defective. Thus, drugs listed below have
been developed to resolve these clinical conditions (Lord and Ashworth 2012).
Double Stranded Breaks
DSBs are the most toxic and lethal form of DNA damage seeking attention from the DDR
pathway (Finn, Lowndes et al. 2012). Two main mechanisms which regulate the repair of DSBs
include non-homologous end joining (NHEJ) and homologous recombination (HR) (Ferguson
and Alt 2001). NHEJ’s distinct feature is that it ligates DNA ends together with minimal
processing. This mechanism occurs throughout the cell cycle, since it does not require template
DNA for the repair process (Caldecott 2008). Thus, it remains a preferred method of repair in
mammalian cells, particularly quiescent cells. However, despite its simplicity, the likelihood of
mutagenesis is higher compared to HR: if ligation does not occur properly, then chromosomal
translocations or fusions may be the outcome (Huen and Chen 2010). Ku protein heterodimers
bind to the DSBs to stabilize the lesions. Then, it recruits the protein kinase DNA-PKcs allowing
for activation and recruitment of the end-processing enzymes (DNA-PK holoenzyme). This
nucleoprotein complex triggers the alignment and rejoining of DNA via DNA ligase IV-XRCC4
42
(Rich, Allen et al. 2000). There is also a Ku-independent NHEJ pathway known as
microhomology-mediated end-joining (MMEJ), which generates deletions in sequences,
however, this pathway remains a subject of study (Ciccia and Elledge 2010).
HR is favored mainly at the S and G2 phases of the cell cycle (Liu and Huang 2014). The DNA
surrounding the DSB is resected to expose the ssDNA. This is promoted by the MRE11-RAD50NBS1 (MRN) complex performing nucleolytic processing. The ssDNA binding protein,
Replication Protein A (RPA) quickly sequesters onto the ssDNA which then dislodge and form a
RAD51-nucleoprotein filament (Liu and Huang 2014). With the combined effort of some other
proteins such as breast-cancer susceptibility proteins (BRCA1 and BRCA2), strand invasion
occurs and a homologous sister chromatid serves as the template for DNA synthesis to ensure
effective repair with the help of polymerases, nucleases and helicases (Daley, Kwon et al. 2013).
Overall, the coordinated assembly of repair factors leads to the tight orchestration of events to
maximize repair efficiency and ensure genomic integrity.
Signal transduction in the DNA damage response
Key components of the signal transduction pathway are phosphoinositol-3-kinase-like protein
kinases ATM (Ataxia telangiectasia) and ATR (Ataxia telangiectasia and Rad3- related)
(Kerzendorfer and O’Driscoll 2009). They are the major regulators in the DNA damage response
network and orchestrate a plethora of phosphorylation events of over 700 proteins. These protein
kinases are activated upon DNA lesion detection caused by genotoxic stress (Huen and Chen
2010). As seen earlier, ionizing radiation generates the most harmful type of DNA damage which
is DSBs. Many of the proteins that take part in the signal cascade and repair mechanisms form
foci structures when triggered by ionizing radiation. RAD51 forms foci structures upon DNA
43
damage, allowing phosphorylation of the histone variant H2AX and recruitment along with an
assembly of other DNA repair proteins at the sites of DNA lesions where the ionizing radiation
induced foci (IRIF) are present (Tarsounas, Davies et al. 2003). Thus, this phosphorylation event
of H2AX (a sensitive marker used to detect DNA damage) is an essential trigger required for the
enhancement of DNA repair (Sharma, Singh et al. 2012). Studies have shown that H2AX
deficient embryos are immunologically compromised and have delays in development, leading to
tumor susceptibility (Huen and Chen 2010).
In mammalian cells, the BRCT domain-containing protein mediator of DNA damage checkpoint
1 (MDC1) established its importance as a key checkpoint regulator and ‘molecular adaptor’ since
it is one of the primary proteins that makes its way to the DNA lesion sites in the γH2AX –
dependent pathway (Stucki, Clapperton et al. 2005). It is a key protein that aids in the
recruitment of other repair proteins such as BRCA1 and 53BP1 to lesion sites where foci have
formed.
The ATM and ATR pathway work in parallel to one another (Marechal and Zou 2013). Their
phosphorylation substrates often overlap (ex: p53, BRCA1, FANC-D2). ATM is primarily
responsible dealing with DNA damage caused by IR, whereas the ATR mediated pathway is
activated with other kinds of stress. ATR knockout mice die early during embryogenesis which
establishes ATR as a vital protein kinase involved in development (Zhou and Elledge 2000).
ATR is known to take on the role of a haploinsufficient tumor suppressor, such that genomic
integrity is maintained and tumor incidence is lessened. Furthermore, loss of ATR generates
genomic instability called DNA fragile Site expression, which are regions in the genome which
usually span over 100 kb in size and demonstrate instability and high chances of breakage during
44
replicative stress (Kerzendorfer and O’Driscoll 2009). Meanwhile, ATM null mice demonstrate
viability despite some growth defects such as infertility (Zhou and Elledge 2000).
ATM pathway specifics
At DSB sites, the Mre11-Rad50-Nbs1 (MRN) complex is used to recruit ATM, alongside other
signal mediators such as MDC1 and 53BP1. Chk2 is an effector kinase protein that is
phosphorylated downstream of ATM, where Chk2 then phosphorylates Cdc25 which triggers it
for ubiquitin-mediated degradation (Falck, Mailand et al. 2001). Due to the fact that Cdc25 is a
key regulator in the activation of Cdk-cyclins, the degradation of Cdc25 does not allow for the
removal of inhibitory phosphates on Cdk1-Cyclin B and Cdk-cyclin E present on Cdk-cyclin
complexes (Falck, Mailand et al. 2001). Hence, the checkpoint arrest occurs at the G2/M and
G1/S phases. Moreover, the p53 tumor suppressor gene is phosphorylated by ATM, which
greatly enhances its stability. This induces its target p21, a Cdk-inhibitor, such that DNA
replication is prohibited (Kerzendorfer and O’Driscoll 2009).
ATR pathway specifics
ATR interacts with ATRIP and is a heterodimer that gets recruited by the Replication Protein A
heterotrimer (RPA1-3) to the SSBs (Zou, Liu et al. 2006). ATR phosphorylates the MRN
complex alongside the Rad17/Rfc2-5 and Rad9/Rad1/Hus1 complexes and all are recruited to
DNA damage sites (Zou and Elledge 2003). ATR kinase activity is increased by TopBP1, since
the complexes mentioned earlier are recruited to the damage sites independent of ATR/ATRIP
complex.
45
ATR is able to phosphorylate other molecules such as p53 and BRCA1. Chk1 is an effector
kinase that works in a similar manner as Chk2. This overlap in pathways highlights the fact that
defects in both Chk1 and Chk2 are required for the cell-cycle checkpoint failure as each on its
own is insufficient to initiate (Kerzendorfer and O’Driscoll 2009). Overall, the forced arrest of
the cell cycle enables DNA repair of damaged cells to take place before moving forward.
Apoptosis
The major cause of programmed cell death is DNA damage (Rich, Allen et al. 2000). Studies
have shown that DNA damage may elicit apoptosis in order to deal with tissue organization
issues. That is, cell-cell communication is vital, especially when cellular processes such as DNA
replication and transcription take place, and thus, complications with these processes has
detrimental effects on the cell. Changes in cellular polarity and sensing are the basis of tumors
such as adenomas (Royer and Lu 2011). Instances like this require apoptosis to take place
compared to other modes of death.
The quality of the genome that is passed down from one generation to the next is closely
monitored via checkpoints and repair mechanisms. In the case that genome integrity is
challenged and successful repair has not taken place, then apoptosis takes place to ensure
transmission of damaged genetic material does not occur. Sometimes, it also happens that mixed
signals are carried through such that even cells where damage has been rectified face the death
signalling pathway.
Apoptosis allows for the quick deletion of cells from tissues while targeting them for
phagocytosis. Parts of cells may be recycled for regeneration via low energy usage (uncoupling
of catalytic and DNA-binding domains of PARP) (Elmore 2007).
46
Telomeres are guanine-rich repetitive DNA sequences that are present at the ends of
chromosomes. DNA repair proteins tend to cluster at telomeric ends perhaps because the cells
consider them as DSBs. The telomeres interact with other proteins to form a loop-like structure
called the shelterin complex (process known as ‘capping’) (Donate and Blasco 2011).With age,
telomere lengths shorten (telomere attrition) and this causes chromosome ends to become
exposed which may result in ligation, however, the shelterin complex prevents this from
happening. After a certain point, the telomere ends surpass minimal length and thus, the shelterin
complex is not able to protect the chromosome anymore. This is lethal for the cell and thus
triggers checkpoints to begin apoptosis to prevent unwanted cell proliferation. Nevertheless,
there are instances when the p53 apoptosis pathway is bypassed, resulting in end-end ligations.
This chromosome fusion and translocation events are harmful for the cell as they promote
tumorigenesis (Donate and Blasco 2011). Such altered chromosomes and shortened telomeres
are key characteristics of persistent cancers. Thus, the integrity of the telomeric DNA challenges
genomic instability (Lord and Ashworth 2012).
Human disorders and cancer
Unfaithful repair of DNA damage and transmission of flawed genomic DNA to progenitor cells
enhance rates of tumor development. A key feature of a tumor cell is its genomic instability
(Zhou and Elledge 2000). Previous studies have demonstrated that alongside genomic instability
in most cancers, there is chromosome instability as well, which refers to any alterations in
chromosome number and structure. When such chromosomal changes occur, it may result in
‘driver’ genes present in the tumor, which leads to overall alterations in cell behaviour increasing
47
the likelihood of disease. Cancer hallmarks include various other types of genomic changes that
occur as well (Zhou and Elledge 2000).
The following diseases may be grouped under the classification of defective DNA signalling
response mechanism:
Ataxia Telangiectasia (A-T)
A-T is a progressive neurological disorder which has a clinical phenotype that consists of the
following: progressive loss of muscle coordination (ataxia), dysfunctional eye coordination
(oculomotor apraxia), impairment of speech and articulation, different kinds of specific
immunoglobulin deficiencies, as well as an increased susceptibility to developing lymphomas
(Kerzendorfer and O’Driscoll 2009). Patients with this condition show a typical defect in the cell
cycle checkpoint G1/S, intra-S and G2/M phases when the DDR pathway is triggered by DSBs.
As well, one of the other hallmark characteristics of this human disorder is that p53 and Chk2 are
unsuccessful in being phosphorylated. Despite the fact that ATM is a vital player in the DDR of
‘heterochromatin associated double stranded breaks’, many components of the DDR are
dysfunctional in this disorder (Kerzendorfer and O’Driscoll 2009). This highlights the
importance of genomic integrity and stability. Functional chromatin is required in order to
prevent atypical ATM signalling. Cell-cycle arrest is vital to allow the cell to repair any DNA
lesions and breaks present. However, this A-T disorder has combined dysfunctional DNA repair
and defective cell-cycle checkpoints, which together create the onset of chromosomal damage
which greatly promotes tumor formation. Thus, the intricate network of DDR, required
checkpoint activation and effective repair are all vital for faithful genome maintenance in order
to prevent such human disorders and promote normal human development (Kerzendorfer and
O’Driscoll 2009).
48
Nijmegen breakage syndrome (NBS)
The MRN (Mre11-Nbs1-Rad50) complex and its substrates are vital players in the signalling
pathway of ATM. Patients with this syndrome exhibit microcephaly, growth impedance, multiple
infections, amenorrhea, ovarian dysgenesis along with a predisposition for lymphoma
(Chrzanowska, Gregorek et al. 2012). Those with NBS syndrome demonstrate extreme
sensitivity to DNA damaging agents along with defects in the following cell-cycle checkpoints:
G1/S, intra-S and G2/M. The characteristics of this disorder are mostly similar to those patients
with A-T syndrome as ATM is a vital player in the DDR pathway to DSBs (Chrzanowska,
Gregorek et al. 2012).
Therapeutic applications and personalized clinical intervention
Thus far, the sites of cancer and its particular clinical features have been used to classify cancer
and analyze patient’s cases (Lord and Ashworth 2012). However, the more effective approach
would be to use techniques such as whole-genome sequencing in order to assess the ‘pathogenic
mutation’ patterns for every cancer. As well, further elucidating the intricate DDR network will
allow for the discovery of novel biomarkers for the early detection of cancer and may provide
personalized treatment options for each patient, on a case-by-case basis. Detection of ‘driver’
genes affected by DNA damage will allow for the effective classification of cancer. Furthermore,
by obtaining the DNA sequence for the cancer (which is often referred to as a ‘sequence scar’), it
is possible to identify which mutagens have driven mutagenesis in the tumor and, which repair
mechanisms have been triggered in response to this damage (Lord and Ashworth 2012). By
following such an approach, patients will be treated based on the assessment of their specific
49
DNA repair mechanism defect and biomarkers will be used to recognize such defects allowing
target based therapy to be used.
Hallmark features of cancer include changes in cell and tissue architecture, and genomic
instability. The nucleus of the cell is where numerous supercomplexes are present which engage
in various aspects of the DNA repair mechanisms as well as apoptotic pathways (Rich, Allen et
al. 2000). The onset of cancer brings about many changes with DNA metabolism as well as
organisation of the nucleus. With such rapid mutations, with numbers ranging from 103 to 105
mutations per tumor, it may be difficult to fully characterize the source of lesions (Rich, Allen et
al. 2000). Nevertheless, certain techniques and advances in cancer research have led to the
emergence of techniques such as karyotyping to visualize and perform chromosomal analysis.
Other immunoprecipitation techniques as well as RNAi screening coupled to functional readouts
may help explore novel components in the DDR pathway (Huen and Chen 2010).
Despite the fact that many components of the DDR pathway have already been analyzed, there is
still need for identifying and characterizing components and interactions taking part in this
intricate and complex network comprising of signalling, repair, checkpoints, and response. A
closer look at novel interactions and DNA damage in vivo studies may help discern between
different medical conditions. This will allow for a more effective and personalized therapeutic
approach in order to maximize the chances of a successful clinical intervention. In order to
optimize the DNA damage response and cater the repair to the type of damage, a network of
sensors is required. Moreover, it is vital to ensure that once repair is initiated, the sensory
component is able to determine when the repair process is complete. The latter is also an
important aspect to consider as the repair mechanism eventually needs to be stopped once the
DNA damage has been rectified (Zhou and Elledge 2000). Constitutive repair is not beneficial
50
for the cell: cells may falsely classify certain components of DNA replication as damage and as a
result, disruptions in DNA synthesis and other deleterious effects are plausible (Zhou and
Elledge 2000). Although unraveling new substrates and their interactions in the DDR pathway
remain enigmatic, it is a key step forward in this field of study, creating scope for advances in
medical research.
51
Experimental Procedures
Antibodies, immunoprecipitations and immunoblotting
Rabbit anti-hnRNPUL1 was purchased from Proteintech (Chicago, IL). Mouse anti-Flag (M2)
antibody, anti--tubulin antibody, and protein-A-Sepharose beads were purchased from Sigma
(St. Louis, MO). Anti-GFP antibody was purchased from Novus Biologicals (Littleton, CO).
Rabbit anti-PRMT1 antibody and ASYM25b were described previously (Cote, Boisvert et al.
2003, Yu, Chen et al. 2009). Species-specific immunoglobulin G (IgG) control antibodies were
from Invitrogen (Carlsbad, CA). Protease and phosphatase inhibitor cocktails were obtained
from Roche (Mississauga, ON). Streptavidin agarose beads were purchased from Life
Technologies (Burlington, ON). Immunoprecipitations and immunoblotting were performed as
previously described (Boisvert, Cote et al. 2002). Briefly, cells were lysed in 50 mM HEPES pH
7.4, 150 mM NaCl, and 1% Triton X-100 on ice for 15 min. After removal of the Triton
insoluble matter by centrifugation, the supernatant was incubated with the indicated antibodies
on ice for 2 h. The bound proteins were immunopurified using protein A Sepharose beads
tumbled at 4˚C for 1 h and separated by SDS-PAGE, transferred to nitrocellulose membranes
and immunoblotted with the indicated antibodies as previously described (Boisvert, Cote et al.
2002).
Plasmids, siRNAs and transfection
Flag-hnRNPUL1 was subcloned into the pcDNA 3.1 vector. The hnRNPUL1 human cDNA
clone was purchased from ORIGENE (Rockville, MD). The primers (Sigma Aldrich) used for
the subcloning were 5'- GGG GGA TCC GAT GTG CGC CGT CTG AAG GTG- 3' and 5'GGG GTC GAC CTA CTG TGT ACT TGT GCC ACC- 3'. Flag-hnRNPUL1RK (R612K,
52
R618K, R620K, R639K, R645K, R656K and R661K in pcDNA 3.1 was generated from FlaghnRNPUL1 by Mutagenex Inc. (Hillsborough, NJ). GFP-hnRNPUL1 and GFP-hnRNPUL1RK
were also generated by Mutagenex Inc. (Hillsborough, NJ). All DNA constructs were entirely
sequenced.
GST-fusion proteins of hnRNPUL1 peptides were constructed by inserting synthesized
oligonucleotide duplexes into the PGEX-3X vector at the Bam HI and Eco RI cut sites.
Oligonucleotide sequences (Sigma Aldrich) of GST fusion proteins of hnRNPUL1 are as
follows:
Di-RG:
5'- GATCTATGAAGAAAACCGGGGACGGGGGTACTTTGAGCACTGA-3' and
5'-TCG ATCAGTGCTCAAAGTACCCCCGTCCCCGGTTTTCTTCATA-3'
RRGR:
5'-GATCCACCGAGAGGATAGGAGGGGCCGCTCTCCTCAGCCTTGA-3' and
5'-TCGATCAAGGCTGAGGAGAGCGGCCCCTCCTATCCTCTCGGTG-3'
RIRG:
5'-GATCCCCCTTAGTGAGCGTATCCGGGGCACCGTTGGACCATGA-3' and
5'-TCGATCATGGTCCAACGGTGCCCCGGATACGCTCACTAAGGGG-3'
Tri-RG:
5'GATCTTTGACAACCGAGGTGGTGGTGGCTTCCGGGGCCGCGGGGGTGGTGGTGGCTT
CCAGTGA-3' and
5'- TCGATCACTGGAAGCCACCACCACCCCCGCGGCCCCGGAAGCCACCACCACCTC
GGTTGTCAAA-3'
53
Di-RGG:
5'-GATCCCTGGAGGCAACCGTGGCGGCTTCCAGAACCGAGGGGGAGGCAGCGGTG
GAGGATGA-3' and
5'-TCGATCATCCTCCACCGCTGCCTCCCCCTCGGTTCTGGAAGCCGCCACGGTTGCC
TCCAGG-3'
Mono-RGG:
5'-GATCGGAGGAGGCAACTACCGAGGAGGTTTCAACCGCAGCGGAGGTGGTGGCT
GA-3' and
5'-TCGATCAGCCACCACCTCCGCTGCGGTTGAAACCTCCTCGGTAGTTGCCTCC
TCC-3'
siRNAs purchased from Dharmacon Inc. (Lafayette, CO) along with their respective target
sequences were as follows: siLuciferase (siCTL, 5'-CGU ACG CGG AAU ACU UCG A dTdT3'), siGFP (siCTL, 5'-AAC ACU UGU CAC UAC UUU CUC UU dTdT -3') sihnRNPUL1 (5'GCA GUG GAA CCA GUA CUA U dTdT -3'), and siPRMT1 (5'-CGT CAA AGC CAA CAA
GTT A dTdT - 3').
Cell culture, transfection and drug treatments
HEK293T and U2OS were purchased from the American Type Culture Collection (Manassas,
VA). Cells were cultured in Dulbecco’s modified Eagle medium (DMEM; Invitrogen)
supplemented with 10% fetal bovine serum (FBS; BioSera), 1 mM sodium pyruvate and
antibiotics under typical culture conditions. Plasmids were transfected with Lipofectamine 2000
(Invitrogen), and siRNAs were transfected with Lipofectamine RNAiMAX (Invitrogen) as per
54
the manufacturer’s instructions. Cells were analyzed 48 h or 72 h after transfection depending on
the experiment to be performed. Stable cell lines were selected with 200 g/ml of Geneticin
(G418; Invitrogen) and a few single clones were selected via western blotting analysis and
immunofluorescence confirmation of stably transfected cells. Campthothecin drug treatments,
using 1 μM as the final concentration in media were performed as described previously (Polo,
Blackford et al. 2012). One hour post drug treatment, cells were washed with PBS and
supplemented with new DMEM media; cells were harvested at the various time points post drug
treatment accordingly.
Expression and purification of GST fusion proteins
GST fusion proteins were purified from 500 ml of E.coli DH5α cells grown in a rich 2XYT
medium with the appropriate antibiotic. Subsequently, following five hours of vigorous shaking
post induction via IPTG (Invitrogen), bacteria were centrifuged at 5000 rpm 7 min at 4˚C. Cells
were lysed with lysis buffer (50 mM Tris pH 7.5, 150 mM NaCl, 1mM PMSF, 5mM EDTA,
0.1% Triton X-100) and samples were sonicated 3 times for 30 sec each, followed by a
centrifugation at 12000 rpm for 15 min. The lysates were incubated with gluthathione-Sepharose
beads on a tumbler at 4˚C overnight. The following day, beads were washed with 20 mM Tris
pH 7.5, 0.1 % Triton X-100, with a final wash with just PBS. GST-fusion proteins were eluted
with elution buffer (10 mM reduced gluthathione in 50 mM Tris pH 8.0). The eluted proteins
were dialysed overnight in PBS.
55
In vitro methylation assays
The methylation of the hnRNPUL1 RGG/RG motifs by PRMT1 was performed using
recombinant proteins. GST-RGG/RG proteins were incubated with GST-PRMT1 and 0.55 µCi
of [methyl-3H] S-adenosyl-L-methionine in the presence of 25 mM Tris- HCl at pH 7.4 for 1 h in
a final total volume of 30 μl (Cote, Boisvert et al. 2003). The reactions were stopped by adding
30 μl of 2X Laemmli buffer, followed by boiling the samples at 100˚C for 10 min. The
methylation of the GST-RGG/RG motifs was assessed by polyacrylamide gel electrophoresis
and stained with Coomassie Blue. The destained gel was soaked in EN3HANCE for 1 h,
followed by a gentle agitation in cold water 30 min. The washed gel was dried at 60˚C for 1.5 h
and visualized by fluorography, as described (Cote, Boisvert et al. 2003).
Affinity pull-down assays
For the affinity pull down assay, 300 μl of cellular lysates were incubated with 2 μl of 1 mM
unmethylated and methylated forms of a biotinylated DiRGG and DiRGGme2a peptides,
respectively, on ice for 1 h. Then, 40 µl of 50% Streptavidin Agarose slurry was added and
incubated at 4 ºC for 1 h with constant end-over-end mixing. Beads were washed with lysis
buffer containing increasing salt concentrations. The samples were then boiled with 40 µl of 2X
SDS PAGE sample buffer, resolved in SDS polyacrylamide gels, transferred to nitrocellulose
membranes and subjected to immunoblotting with the indicated antibodies. As previously
mentioned, peptides used were as follows:
DiRGG Biotin-RRGRGRGRGFRGARGGRGGGGAPRG-NH2
DiRGGme2a Biotin-RRGRGRGRGFRGAR(Me2a)GGR(Me2a)GGGGAPRG-NH2
where R (Me2a) signifies a methylation arginine residue.
56
Laser Scissor
HeLa cells were transfected with 400 ng of GFP-hnRNPUL1 and GFP-hnRNPUL1RK plasmids
with Effectene transfection reagent from Qiagen (Toronto, ON). 16 hours post transfection,
FRAP analysis was performed. Fluorescence was observed on a Leica TCS SP5 II confocal
microscope. A 405-nm ultraviolet laser was used to create laser-induced DNA damage. GFP
fluorescence was visualized and observed within the micro-irradiated nuclear region using a 488
nm excitation filter with a 63X objective. Volocity software was used in order to monitor
background and perform photo-bleaching corrections to every dataset, respectively.
57
Results:
RGG/RG motifs of hnRNPUL1 are methylated by PRMT1
Arginine glycine repeat sequences have been found in more than 1400 human proteins
(Thandapani, O'Connor et al. 2013). They can be classified into 4 types of RGG/RG motifs
namely, Di-RG, Di-RGG, Tri-RG and Tri-RGG (Thandapani, O'Connor et al. 2013). RGG/RG
motifs regulate protein/protein, protein/RNA and protein/DNA interactions and are preferred
sites of methylation by protein arginine methyltransferases (PRMTs) (Thandapani, O'Connor et
al. 2013). The arginine methylation of the MRE11 RGG/RG motif is physiologically required for
optimal MRE11 nuclease activity to activate the ATR signalling cascade (Yu, Vogel et al. 2012).
To further define the role of RGG/RG motif and its methylation in DNA damage signalling, we
chose to study heteronuclear ribonucleoprotein (hnRNP) U like protein 1. hnRNPUL1 was
recently shown to participate in ATR activation following DNA damage signalling (Polo,
Blackford et al. 2012). hnRNPUL1 harbors a DiRGG and a TriRG motif as well as single RG
sequences (Fig. 5A). We examined whether these sequences were methylated by PRMT1. In
vitro methylation assays were performed where each of the RGG/RG motifs of hnRNPUL1 was
fused to GST. These GST-RGG/RG fusion proteins were incubated with GST-PRMT1 and
[methyl-3H]-S-adenosyl-L-methionine, as the methyl donor. The methylated proteins were
visualized by fluorography after separation by SDS-PAGE and equal loading was observed by
Coomassie Blue staining (Fig. 5B, left panel). The TriRG, DiRGG as well as monoRGG of
hnRNPUL1 were methylated by PRMT1. GST-GAR of MRE11 was used as a positive control
and GST alone as a negative control (Fig. 5B, right panel). All of the three methylated RGG/RG
motifs are located nearby to each other from amino acid 612 to 666.
58
To investigate whether hnRNPUL1 harbors any methylarginines in vivo, we performed mass
spectrometry analysis. U2OS cells transfected with an expression vector encoding Flag-epitope
tagged hnRNPUL1 were lysed under denaturing conditions with a lysis buffer containing 1%
SDS in order to disrupt protein-protein interactions and thus limit proteins that coimmunoprecipitate with hnRNPUL1. Such denaturing conditions allow for a reduction in
background, since we were interested uniquely in studying the post-translational modifications of
hnRNPUL1 and not in identifying other unspecific protein-protein interactions. The SDS was
diluted to 0.1% with lysis buffer devoid of SDS, such that Flag-hnRNPUL1 could be
immunoprecipitated with anti-Flag antibodies coupled to agarose beads. The immunopurified
hnRNPUL1 was subjected to mass spectrometry with focus on modified hnRNPUL1 peptides
(Fig. 6B). Mass spectrometry analysis detected 49 peptides with 62% coverage of hnRNPUL1
and 42 of the total 66 arginines in the protein were detected (Fig. 6A). Amongst the 42 arginines
detected, 5 were dimethylated (R584, R618, R620, R645, R656), while some were
monomethylated (R661, R685, R690) (Table 1). Importantly, R645 resides in a Di-RGG motif
and R618 and R620 reside within the Tri-RG motif of hnRNPUL1. The presence of
monomethylated R661, R685 and R690, suggests that monomethylarginines may represent
intermediates for the final asymmetrical dimethylation step by PRMT1.
Moreover, the methylation sites observed in vitro correlate and are included in the list of arginine
residues that were shown to be methylated in vivo via mass spectrometry analysis i.e. R584, 618,
620, 645, 656, 661, 685 and 690, which further reinforces our findings.
59
PRMT1 methylates and associates with hnRNPUL1 in vivo.
We next examined whether PRMT1 was the enzyme responsible for the methylation of
endogenous hnRNPUL1 in vivo. HEK293 cells were transfected with control (siLuciferase) or
PRMT1 siRNAs and cellular extracts were prepared for immunoprecipitation with control (IgG)
or anti-hnRNPUL1 antibodies. The bound proteins were separated by SDS-PAGE and
immunoblotted with ASYM25b, a specific asymmetric dimethylarginine antibody. Equivalent
hnRNPUL1 was immunoprecipitated from siLuc and siPRMT1 cells (Fig. 7A). ASYM25b only
recognized hnRNPUL1 in siLuc, but not siPRMT1 transfected cells, demonstrating that
hnRNPUL1 was hypomethylated in PRMT1-depleted cells. These findings suggest that PRMT1
is the physiological enzyme responsible for the arginine methylation of hnRNPUL1 in vivo.
We identified hnRNPUL1 arginines 612, 618, 620, 639, 645, 656, and 661 to be arginine
methylated and we next proceeded to substitute all these 7 arginines with lysines to generate a
mutant hnRNPUL1 (RK) that is devoid of any methylarginines (Fig. 6 and Table 1). We chose
lysine to maintain the charge and lysine is not a substrate of PRMTs. We initially examined
whether Flag-hnRNPUL1RK is indeed hypomethylated in vivo. HEK293 cells transfected with
Flag-hnRNPUL1 or Flag-hnRNPUL1RK were lysed and anti-Flag immunoprecipitations were
performed. Immunoblotting of the Flag immunoprecipitations with ASYM25b revealed that
hnRNPUL1 was abundantly arginine methylated, but not Flag-hnRNPUL1RK (Fig. 7B).
Immunoblotting lysates with anti-tubulin antibodies confirmed equivalent loading and
expression of the Flag epitope tagged proteins. These findings show that the arginines that reside
within RGG/RG motifs are the only sequences that contain methylarginines in hnRNPUL1.
We tested whether endogenous PRMT1 associated with Flag-hnRNPUL1 and FlaghnRNPUL1RK. HEK293 transfected with Flag-hnRNPUL1 or Flag-hnRNPUL1RK were lysed and
60
co-immunoprecipitation analyses were performed. Untransfected cells were used as the negative
control. We observed that PRMT1 co-immunoprecipitated with Flag-hnRNPUL1, but not FlaghnRNPUL1RK (Fig. 7B). Immunoblotting lysates with anti-tubulin and anti-Flag antibodies
confirmed equivalent loading and expression of the Flag epitope tagged proteins (Fig. 7B). The
fact that PRMT1 did not associate with Flag-hnRNPUL1RK may explain why the protein is
indeed hypomethylated. These findings suggest that the 7 arginines located within the RGG/RG
motifs of hnRNPUL1 are the sites of methylation by PRMT1.
Arginine methylation regulates the hnRNPUL1/NBS1 interaction
Previous studies identified an interaction between the RGG/RG region of hnRNPUL1
and NBS1, but it was not known that hnRNPUL1 was arginine methylated or was required for
this interaction (Polo, Blackford et al. 2012). To determine whether arginine methylation of
hnRNPUL1 affects the NBS1 interaction, methylated and unmethylated DiRGG biotinylated
peptides were used in affinity pull-down assays (Fig. 8A). Cellular lysates were prepared from
HEK293 cells transfected with YFP-NBS1 and incubated with DiRGG and DiRGGme2a
biotinylated peptides bound to Streptavidin beads. The bound proteins were separated by SDSPAGE and immunoblotted with anti-GFP antibodies to detect YFP-NBS1. We observed that the
methylated DiRGG peptide had a higher relative affinity towards NBS1 in comparison to the
unmethylated peptide and this was confirmed by washing with increasing concentrations of NaCl
(Fig. 8A and 8B).
We tested whether hypomethylated hnRNPUL1 interacted with NBS1. U2OS cells were cotransfected with control pcDNA3.1, Flag-hnRNPUL1 or Flag-hnRNPUL1RK and YFP-NBS1
expression vectors. Once the cells were lysed, co-immunoprecipitation experiments were
61
performed. NBS1 associated with the Flag-hnRNPUL1, as expected, however, NBS1 had a
reduced affinity for Flag-hnRNPUL1RK (Fig. 8C).
To further determine whether arginine methylation of hnRNPUL1 regulates its interaction with
NBS1, we examined the interaction in PRMT1-deficient cells. HEK293 cells were transfected
with control or PRMT1 siRNAs along with YFP-NBS1 and Flag-hnRNPUL1. The lysed cells
were immunoprecipitated with anti-Flag antibodies and the bound proteins were separated by
SDS-PAGE and immunoblotted with anti-GFP antibodies. We observed a reduced
NBS1/hnRNPUL1 interaction in PRMT1 siRNA treated cells (Fig. 8D). Immunoblots of whole
cell lysates (WCL) from control and PRMT1 knockdown cells with anti-GFP, anti-FLAG and
anti-PRMT1 antibodies confirmed equal expression, as well as the PRMT1 knockdown. These
findings suggest a role for arginine methylation in the interaction between NBS1 and
hnRNPUL1.
The methylation of the hnRNPUL1 RGG/RG motifs regulates its recruitment at sites of
DNA damage
hnRNPUL1 exhibits two types of dynamics at DNA damage sites depending on the
presence or absence of RNA (Polo, Blackford et al., 2012). hnRNPUL1 is rapidly excluded from
laser microirradiation sites, but in the presence of transcription inhibitors, it is effectively
recruited at DNA damage tracks. An RGG deletion mutant of hnRNPUL1 was shown to be
defective in these dynamic properties (Polo, Blackford et al., 2012). To assess the role of the
arginine and their methylation within the RGG motifs in the exclusion and recruitment at sites of
DNA damage, we transfected U2OS cells with GFP-hnRNPUL1 or GFP- hnRNPUL1RK. The
cells were examined for the recruitment to laser microirradiated nuclear regions in the absence
62
and presence of the transcriptional inhibitor,
5,6 dichloro--D-ribofuranosylbenzimidazole
(DRB). Interestingly, both GFP-hnRNPUL1 and hnRNPUL1RK were excluded from laser
microirradiated nuclear regions in U2OS cells (Figure 9A). However, GFP-hnRNPUL1 was
effectively recruited at sites of DNA damage, while GFP-hnRNPUL1RK was not recruited at
these DNA damage sites (Figure 9A, 9B). Furthermore, these observations demonstrate a role for
the arginine methylation of RGG/RG motif of hnRNPUL1 in the recruitment of hnRNPUL1 at
DNA damage sites.
63
Table 1. Methylated arginine residues of hnRNPUL1 identified in vivo via mass
spectrometry
64
Figure 5. The RGG/RG motif of hnRNPUL1 is methylated by PRMT1 in vitro
A) SAP designates the SAF- A/B, Acinus and PIAS motif, while BBS denotes BRD7-binding
site. The sequence of 549-696 is shown: underlined and bold arginines were identified to be
methylated. B) GST with the indicated peptide sequences were methylated by recombinant
3
PRMT1 in vitro using H –SAM. Proteins were resolved by SDS-PAGE, stained with Coomassie
blue (left panel), dried and analyzed by fluorography (right panel). GST-GAR (glycine arginine
rich) and GST were used as the positive and negative control, respectively. The indicated
sequences (RGG or RG) were fused to GST and thus migrate at a similar molecular mass as
GST. GST-GAR harbors extra ~60aa and migrates close to 33kDa.
65
A
B
Figure 6. hnRNPUL1 harbors methylated arginine residues identified in vivo via mass
spectrometry analysis
A) Protein sequence of hnRNPUL1 with the arginine residues that were detected via mass
spectrometry highlighted in purple. B) Schematic of a peptide analyzed by mass spectrometry is
depicted. Flag-hnRNPUL1 in U2OS cells was immunoprecipitated and subjected to mass
spectrometry analysis. The corresponding mass spectrum demonstrates the addition of the methyl
groups to the respective arginine residues. The addition of 28 atomic mass units denotes the
addition of a methyl group to the arginine residue as indicated by the x-axis mass over charge
ratio. Moreover, the peak size as indicated on the y-axis of the spectrum correlates with intensity
and abundance of the peptide.
66
Figure 7: The RGG/RG motif of hnRNPUL1 is methylated by PRMT1 in vivo via a
physical association
A) Whole cell lysates from HEK293 cells transfected with siPRMT1 and siControl were
subjected to immunoprecipitation 48 h post transfection with the anti-hnRNPUL1 antibody.
Whole cell lysates (WCL) and immunoprecipitants were immunoblotted with the indicated
antibodies. B) HEK293 cells were transfected with Flag-hnRNPUL1 wildtype and FlaghnRNPUL1RK in which the arginine residues R612, R618, R620, R639, R645, R656, and R661
located in the RGG/RG motif and identified to be methylated were mutated to lysines. Whole
cell lysates (WCL) and Flag immunoprecipitants were subjected to western blot analysis using
the antibodies indicated. Tubulin was used as loading control.
67
Figure 8. Methylation of hnRNPUL1 is required for its interaction with NBS1
A) HEK293 cells were transfected with YFP-NBS1. Cellular lysates were incubated with 2 μl of
1 mM DiRGG and DiRGGme2a peptides and subjected to streptavidin pull-down. Whole cell
lysates (WCL) and Streptavidin-immunoprecipitants were immunoblotted with the anti-GFP
antibody. B) HEK293 cells were transfected with YFP-NBS1. Cellular lysates were incubated
with 2 μl of 1 mM DiRGG and DiRGGme2a peptides and subjected to streptavidin pull-down.
Beads were washed with lysis buffer containing increasing concentrations of NaCl. Whole cell
lysates (WCL) and Streptavidin-immunoprecipitates were immunoblotted with the anti-GFP
antibody. C) U2OS cells were transfected with empty vector (pcDNA 3.1), Flag-hnRNPUL1
wildtype, Flag-hnRNPUL1 R-K mutant and YFP-NBS1. Whole cell lysates (WCL) and Flagimmunoprecipitants were immunoblotted with the indicated antibodies. D) HEK293 cells were
transfected with siControl and siPRMT1 along with YFP-NBS1 and Flag-hnRNPUL1. The
whole cell lysates (WCL) and Flag immunoprecipitates were subjected to western blot analysis
using the antibodies indicated.
68
Figure 9. The methylation of the hnRNPUL1 RGG/RG motifs regulates its recruitment at
sites of DNA damage
A) U2OS cells were transfected with GFP-hnRNPUL1 wildtype and GFP-hnRNPUL1RK and
subjected to laser micro irradiation treatment. The cells were examined for the recruitment to
laser microirradiated nuclear regions in the absence and presence of the transcriptional inhibitor,
5,6 dichloro--D-ribofuranosylbenzimidazole (DRB). Live imaging was used to observe the
exclusion and recruitment patterns post laser scissor damage. B) Quantification of percentage of
cells with hnRNPUL1 recruitment. Quantification of recruitment patterns of GFP-hnRNPUL1
(WT) and GFP-hnRNPUL1RK (R/K mutant) respectively.
69
Discussion:
RNA binding proteins (RBPs) have been increasingly associated with a multitude of
pathways, including, but not limited to RNA metabolism, DNA metabolism, signal transduction
and more recently DNA repair (Dutertre, Lambert et al. 2014). Their roles in such pathways
include the regulation and controlled monitoring of post-transcriptional gene regulation through
binding to RNA sequences and secondary structures in untranslated and coding regions of
mRNAs and association with non-coding RNAs (Lunde, Moore et al. 2007, Glisovic, Bachorik
et al. 2008). Post-translational modifications of proteins play an integral part of the DNA damage
response pathway. RBPs are sensitive to post-translational modifications by regulating their
cellular localization, stability and recruitment at sites of DNA damage. Moreover, these
modifications can elicit tight control (both spatial and temporal) of the key players involved in
the DNA damage response such that protein-protein interactions may be facilitated allowing
effective signal amplification and a final cellular response which include DNA repair, cell cycle
arrest, and apoptosis.
My findings have elucidated a role for arginine methylation of hnRNPUL1, a member of the
hnRNP family, in the DNA damage response pathway. The family of hnRNPs have been
implicated in numerous cellular processes including RNA metabolism, telomere elongation,
DNA repair, and chromatin reorganization, in addition to among many other regulatory pathways
(Carpenter, MacKay et al. 2006). hnRNPUL1 has sequence homology to hnRNP U, a protein
which has been shown to be a participant in the DNA damage response pathway (Polo,
Blackford et al. 2012).
70
hnRNP U, also known as scaffold attachment factor A (SAF-A), is a nucleic acid binding
ribonucleoprotein which has a characteristic binding preference for scaffold associated DNA
regions including areas rich in A-T amino acids (Fackelmayer, Dahm et al. 1994).
hnRNP U contains a unique site from 778 to 793 amino acids bearing four RG rich regions
which are preferred sites for arginine methylation (Herrmann, Bossert et al. 2004). PRMT1 has
been reported to be the enzyme responsible for the methylation of hnRNP U in its RGG/RG rich
motif (Herrmann, Bossert et al. 2004). In addition, the unmethylated form of hnRNP U is
unstable, further suggesting the methylation of this protein is constitutive. As methylated
proteins may be recycled and reused, this process allows for the constant replenishment of the
resource pool, making it available for the newly synthesized gene products throughout a
multitude of cellular processes that occur in the cell (Herrmann, Bossert et al. 2004). In the
present study, we have defined a role for the methylation of the RGG/RG motif of hnRNPUL1 in
DNA damage signalling. We show that the TriRG, DiRGG and monoRGG of hnRNPUL1 are
methylated by PRMT1 in vitro using [methyl-3H] S-adenosyl-L-methionine as the methyl donor.
Defects in PRMT1-deficient cells have been previously characterized (Yu, Chen et al. 2009). It
has been shown that PRMT1 is essential for early development using a conditional null allele of
PRMT1 in mice. PRMT1 null embryos die at embryonic day 6.5 (Yu, Chen et al. 2009). PRMT1deficient MEFs display genomic instability and exhibit spontaneous DNA damage, checkpoint
defects, and delays in cell cycle progression. This further supports a crucial role for PRMT1 in
the DNA damage response pathway (Yu, Chen et al. 2009).
We further showed via mass spectrometry analysis of Flag-tagged hnRNPUL1 in U2OS cells the
presence of methylarginines in vivo. The identification of mono-methylarginine suggests these
marks serve as potential priming sites for the subsequent dimethylation by PRMT1. These
71
findings further reinforced our previously obtained analysis in vitro, where we observed that
PRMT1 was responsible for the in vitro methylation of the RGG/RG rich sequences of
hnRNPUL1 (Fig. 5B).
The role of arginine methylation and PRMTs in the DDR pathway has not fully been
characterized. Nevertheless, we observed the methylation status of hnRNPUL1 in PRMT1
knockdown HEK293 cells using siRNA. hnRNPUL1 was shown to be hypomethylated in the
PRMT1-deficient cells, demonstrating that PRMT1 is the physiological enzyme responsible for
the methylation of hnRNPUL1 in its RGG/RG motif. Previous studies have implicated
hnRNPUL1 to be required for resection at double strand breaks and to be required for optimal
activation of ATR. It was proposed that there are two pools of hnRNPUL1 in the cell, such that
one pool is involved in RNA metabolism, while the other participates in DSB repair (Polo,
Blackford et al. 2012). Perhaps arginine methylation divides these two pools.
Earlier studies have demonstrated a role for arginine methylation of the MRE11 subunit of the
MRN molecular sensor complex, an important complex in the DDR, to be methylated in its
RGG/RG motif by PRMT1 both in vitro as well as in vivo (Boisvert, Dery et al. 2005). The
exonuclease activity of MRE11 is regulated via its methylation by PRMT1 (Boisvert, Dery et al.
2005). A previously generated mouse knock-in allele of Mre11, where the methylated arginines
within its RGG/RG motif are substituted to lysines, displayed hypersensitivity to γ-irradiation
(IR) (Yu, Vogel et al. 2012). In addition, genomic instability and cell cycle checkpoint defects
were also observed. An analogous experimental design, using charge conservation, an
hnRNPUL1 (RK) mutant protein was generated such that it was devoid of methylated arginines
at the previously identified sites. We were able to show that the Flag-hnRNPUL1RK was
72
hypomethylated in vivo, whereas the Flag-hnRNPUL1 was copiously methylated, which further
signifies the presence of methylarginines within the context of the RGG/RG motif.
PRMT1 is known to associate with its substrates (Zhang and Cheng 2003). Moreover, it
preferentially methylates RGG/RG motifs of RBPs in addition to other proteins (Tang, Kao et al.
2000). Our findings demonstrate, using endogenous PRMT1 in co-immunoprecipitation studies
that PRMT1 physically interacts with Flag-hnRNPUL1, but not with Flag-hnRNPUL1RK.
It has been shown that the localization and recruitment of hnRNPUL1 to DSB damage sites is
dependent on the presence of the MRN complex (Polo, Blackford et al. 2012). It has also been
reported that hnRNPUL1 functions in a pathway that is downstream of MRN and CtIP, however,
upstream of BLM such that hnRNPUL1 promotes DSB resection by regulating BLM recruitment
(Polo, Blackford et al. 2012). Previous findings have also demonstrated an interaction between
the C-terminal of NBS1 and the RGG/RG motif of hnRNPUL1. Nonetheless, the role that
methylation plays in this interaction was now known. We used methylated and unmethylated
forms of DiRGG biotinylated peptides in affinity pull-down assays. We observed an increase
binding between the methylated peptide and NBS1, as opposed to the unmethylated form of the
peptide. These findings suggest the methylation of hnRNPUL1 increases the interaction with
NBS1. To explore this further in vivo, we used Flag-hnRNPUL1 and evaluated its association
with endogenous NBS1. Interestingly, the affinity for this interaction was significantly lessened
for Flag-hnRNPUL1RK (Fig. 8C). In addition, using siRNA PRMT1-depleted cells in which both
YFP-NBS1 and Flag-hnRNPUL1 were overexpressed, we observed a reduced binding
interaction between NBS1 and hnRNPUL1.
Previous studies have shown that the RGG/RG motif of hnRNPUL1 is essential for its
mobilization to and from sites of DNA damage, in an MRN-dependent and RNA-independent
73
manner (Polo, Blackford et al. 2012). Moreover, it has been observed that upon laser scissor
damage, hnRNPUL1 forms unique exclusion and recruitment patterns, such that they either
become excluded from the breakage sites or are recruited to them (Polo, Blackford et al. 2012).
Nevertheless, as numerous post-translational modifications have been shown to play a role in the
DDR pathway, this collectively indicates the relevance of further exploring the roles of arginine
methylation in the context of DNA repair pathways. Using post-laser scissor damage, we
observed a prominent role for the arginine methylation of hnRNPUL1, by observing U2OS cells
transfected with GFP-hnRNPUL1 or GFP-hnRNPUL1RK and their distribution patterns
accordingly, whether it is exclusion or recruitment. The cells were examined for the recruitment
to laser microirradiated nuclear regions in the absence and presence of the transcriptional
inhibitor, 5,6 dichloro--D-ribofuranosylbenzimidazole (DRB). An interesting pattern that was
observed was the following: both GFP-hnRNPUL1 and hnRNPUL1RK were excluded from laser
microirradiated nuclear regions in U2OS cells (Figure 9A). Nevertheless, GFP-hnRNPUL1 was
effectively recruited at sites of DNA damage, while GFP-hnRNPUL1RK was not recruited at
these DNA damage sites (Figure 9A, 9B). These observations had confirmed that initial thought
that perhaps the methylation of the arginine residues located in the RGG/RG motif of
hnRNPUL1 could be required for the recruitment of hnRNPUL1 to the sites of DNA damage.
Therefore, these findings suggest that the arginines within the RGG/RG motifs are required for
the recruitment at sites of DNA damage in the presence of transcription inhibitors, but are not
required for exclusion of hnRNPUL1 from sites of DNA damage. Further studies in this matter
should be performed to further explore whether arginine methylation and PRMT1 is required for
the localization of hnRNPUL1 to DNA damage sites. This may be done by using U2OS cells
74
transfected with PRMT1 siRNA and the recruitment of GFP-hnRNPUL1 at laser microirradiated
nuclear regions may be assessed accordingly.
Collectively, our findings demonstrate a role for arginine methylation of hnRNPUL1 in the DNA
damage response pathway. Nevertheless, there still remains much more to be determined to fully
delineate the role that arginine methylation plays in the DDR. For example, it would be
important to perform functional studies comparing Flag-hnRNPUL1 and Flag-hnRNPUL1RK
while removing the endogenous hnRNPUL1 via siRNA or using CRISPR/Cas9 technology. This
will enable us to further monitor and assess the defects that hypomethylation of hnRNPUL1 may
cause to the DDR. In particular, in order to understand the role of arginine methylation in the
regulation of hnRNPUL1 functions, mutants harboring R-K mutations of the identified methyl
arginine sites may be used to study defects in checkpoint activation, genomic instability and
DNA damage response. For example, in the case of checkpoint activation studies, a role for the
RGG/RG motif of hnRNPUL1 may be elucidated, if we are to predict that the hnRNPUL1RK will
have defects in carrying out proper cell-cycle transitioning. Survival assays such as colony
formation assays post campthothecin drug treatment (causing DNA damage) may be used in
order to assess any differences between the wild type and mutants forms of hnRNPUL1. In
perspective, measuring the half-life of hnRNPUL1 wild type and mutants will enable us to
comment on how the methylation of hnRNPUL1 will affect overall protein stability.
Another interesting question that we may expand on is whether the methylation of hnRNPUL1 is
modulated upon DNA damage. Further experiments need to be performed in order to determine
if the arginine methylation of hnRNPUL1 is regulated upon DNA damage. SILAC experiments
allow for the effective comparison between the relative levels of arginine methylation between
DNA-damage treated and non-treated samples. Thus, large quantities of protein are required for
75
the accurate detection of peptides covering the region of the methylated residues. Our SILAC
experiment was not sensitive enough to compare the arginine methylation between the IR-treated
cells and control cells. Thus, further experimentation will allow us to shed some light in this area
of study.
Although many components of the hnRNPUL1 mechanism are being gradually uncovered, the
overall contribution of this post-translational modification necessitates further studies to gain
perspective on the significance of hnRNPUL1.
76
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