UNIVERSITY OF CINCINNATI
Date:___March 7, 2005_____
I, _____Jacob Robison______, hereby submit this work as part of the requirements for the degree of: __Doctor of Philosophy (Ph.D.)______in: __Department of Environmental Health______It is entitled: _Interaction of the Mre11/Rad50/Nbs1 (MRN) Complex and______Replication Protein A (RPA) in Response to DNA Damage______
This work and its defense approved by:
Chair: __Kathleen Dixon, Ph.D.______Co-chair: __John Bissler, M.D.______Michael Borchers, Ph.D.______Mario Medvedovic, Ph.D______Gregory Oakley, Ph.D.______James Stringer, Ph.D.______
Interaction of the Mre11/Rad50/Nbs1 (MRN) Complex and Replication Protein A (RPA) in Response to DNA Damage
A dissertation submitted to the
Division of Research and Advanced Studies of the University of Cincinnati
in partial fulfillment of the requirements for the degree of
DOCTOR OF PHILOSOPHY (Ph.D.)
in the Department of Environmental Health of the College of Medicine
Monday, March 7, 2005
At 10:00 a.m. in 121 Kettering
by
Jacob Robison
B.S. Brigham Young University
Committee Chair: Kathleen Dixon, Ph.D. Committee Co-Chair: John Bissler, M.D. Committee: Michael Borchers, Ph.D. Mario Medvedovic, Ph.D. Gregory Oakley, Ph.D. James Stringer, Ph.D.
ABSTRACT
Both replicative stress and DNA damage initiate cellular processes collectively termed
the DNA damage response. These processes include activation of appropriate DNA repair
mechanisms, cell cycle checkpoints, and in some cases, apoptosis. Accurate and efficient
operation of the DNA damage response is essential for preventing mutations that may
lead to oncogenic transformation or some types of inherited diseases. The DNA damage
response involves sensing the damage, activation of specific kinases that transduce the
activation signal via protein phosphorylation, and activation of effector proteins that carry
out the functional aspects of the response. Two hallmarks of the DNA damage response
are phosphorylation of key regulatory proteins and aggregation of multiprotein
complexes into foci at or near the site of damage. The proteins that are phosphorylated
and the composition of the foci depend upon the nature of the DNA lesion, and changes
as the damage is recognized, processed and then repaired. Although different types of
DNA damage activate specific repair proteins and pathways, some proteins respond to
multiple types of lesions. Two protein complexes essential for the response to many
lesions types are the Mre11/Rad50/Nbs1 (MRN) complex and replication protein A
(RPA). Evidence supports the hypothesis that both of these complexes have multiple
roles in the DNA damage response, including initial DNA damage recognition, activation
of the signal transducing kinases and functional roles in DNA repair pathways. Although the MRN complex and RPA both become phosphorylated and form foci in response to multiple types of DNA lesions, we found that they co-localize to nuclear foci only in response to a subset of lesions. However, depletion of RPA via siRNA abrogates the ability of the MRN complex to form foci. These data suggest that the MRN complex and
RPA have functional activities that can be both dependent and independent of each other.
Understanding the determinant of whether or not the MRN complex and RPA interact, as well as the functional consequence of this interaction, will help elucidate the cellular
responses to different types of DNA lesions and provide crucial information that may
allow us to intervene to prevent the negative effects of DNA damage.
ACKNOWLEDGMENTS
First, I would like to acknowledge and thank my committee members Dr. Kathleen
Dixon, Dr. John Bissler, Dr. Michael Borchers, Dr. Mario Medvedovic, Dr. Gregory
Oakley and Dr. James Stringer for their support and time as they willingly met with me on a frequent basis to discuss the progress of my research and the scientific quality of my work. I especially would like to thank my advisor Dr. Dixon and my co-advisor Dr.
Bissler. Dr. Dixon allowed me the opportunity to begin my dissertation work in her laboratory, and gave me the freedom to choose and design a project of my own. Through her mentorship, I was able to get a great start on the dissertation before moving over to
Dr. Bissler’s lab. Dr. Bissler has provided a great environment for me to finish my dissertation, and more importantly, to better prepare for my future. Dr. Dixon and Dr.
Bissler through the entire course of my graduate work have been great mentors, advisors and friends; and I know those relations will continue into the future. I owe them a sincere thank you for their time and efforts to help make my graduate work a very educational and positive experience.
Additionally, I need to express gratitude for my wife Aliess and her support as I have
taken this long road of training to obtain the career that I dream of. Her love and
understanding have made this experience much easier and more enjoyable. Our children
Reid, Alexis, Elizabeth and Kurt also deserve recognition. There is nothing like the enthusiasm and love of a young child to help you remember what is really important in life.
i
TABLE OF CONTENTS
ABBREVIATIONS ...... iii
LIST OF FIGURES AND TABLES...... v
CHAPTER 1. Background and Introduction ...... 1
CHAPTER 2. Replication Protein A (RPA) and the Mre11/Rad50/Nbs1 (MRN)
Complex Co-localize and Interact at Sites of Stalled Replication Forks...... 12
I. INTRODUCTION...... 12
II. MATERIALS AND METHODS...... 15
III. RESULTS ...... 19
IV. DISCUSSION...... 41
CHAPTER 3. DNA Lesion-Specific Co-localization of the Mre11/Rad50/Nbs1 (MRN)
Complex and Replication Protein A (RPA) to Repair Foci...... 45
I. INTRODUCTION...... 45
II. MATERIALS AND METHODS...... 48
III. RESULTS ...... 50
IV. DISCUSSION...... 65
CHAPTER 4: Loss of Replication Protein A (RPA) Prevents Phospho-Nbs1 Foci
Formation Following Etoposide-Induced DNA Damage ...... 70
I. INTRODUCTION...... 70
II. MATERIALS AND METHODS...... 72
III. RESULTS ...... 75
IV. DISCUSSION...... 86
CHAPTER 5. Recombinant Proteins and Future Directions...... 92 ii
I. INTRODUCTION...... 92
II. ATM and ATR ...... 92
A. RATIONALE...... 92
B. MATERIALS AND METHODS...... 93
C. RESULTS...... 96
D. DISCUSSION ...... 99
III. MRN Complex...... 100
A. RATIONALE...... 100
B. MATERIALS AND METHODS...... 101
C. RESULTS...... 103
D. DISCUSSION ...... 108
IV. RPA...... 109
A. RATIONALE...... 109
B. MATERIALS AND METHODS...... 109
C. RESULTS...... 112
D. DISCUSSION ...... 116
V. DISCUSSION ...... 116
CHAPTER 6. Conclusion ...... 120
REFERENCES ...... 128
iii
ABBREVIATIONS
A-T …………………. Ataxia telangiectasia
ATLD …………………. Ataxia telangiectasia-like disorder
ATM …………………. Ataxia telangiectasia mutated
ATP …………………. Adenosine triphosphate
ATR …………………. ATM- and Rad3-related
BSA …………………. Bovine serum albumin
CAMPT ……………….. Camptothecin
CIP …………………. Calf intestinal phosphatase
CP …………………. Creatine phosphate
CPK …………………. Creatine phospho-kinase
DAPI …………………. 4′,6′-deanidino-2-phenylindole
DMEM ………………… Dulbecco’s modified eagles medium
DNA-PK ………………. DNA-dependent protein kinase
DSB …………………. Double-strand break dsDNA ……………….... Double-stranded DNA
DTT …………………. Dithiolthreitol
EDTA …………………. Ethylenediaminetetraacetic acid
EtBr ……………….…. Ethidium bromide
ETOP ………………….. Etoposide
FBS …………….……. Fetal bovine serum
γH2AX …………………. Phosphorylated histone H2AX iv
HU ………………….. Hydroxyurea
IR ………………….. Ionizing radiation
λPPase …………………. Lambda phosphatase
MEM ………………….. Minimum essential medium
MMC ………………….. Mitomycin C
MRN Complex …….…... Mre11/Rad50/Nbs1 complex
NBS …………….……. Nijmegen breakage syndrome
PAGE ………………….. Polyacrlyamide gel electrophoresis
PBS ………………….. Phosphate buffered saline
PIKKs ………………….. Phosphatidylinositol 3-kinase-like kinases
PMSF ………………….. Phenylmethylsulfonyl fluoride
PS ………………….. Penicillin-streptomycin
RPA ……….…………. Replication protein A
SDS ………………….. Sodium dodecyl sulfate
SSB …………………. Single-strand breaks ssDNA ……….………… Single-stranded DNA
UV ……….………… Ultraviolet light
WRN ……….………… Werner’s Helicase
v
LIST OF FIGURES AND TABLES
Figure 1 DNA damage response ………………………………………………...... 1
Figure 2 Role of the MRN complex and RPA in the DNA damage response .….... 8
Figure 3 Phosphorylation of RPA and Mre11 after HU and UV treatment ….. 28-29
Figure 4 Phosphorylated RPA and Mre11 are chromatin-bound ………….…. 30-31
Figure 5 RPA and Mre11 co-localize to discrete nuclear foci following HU and
UV treatment ……………………………………………………...…32-33
Figure 6 Co-immunoprecipitation of RPA and Mre11 ……………………..…34-36
Figure 7 Phosphatase treatment disrupts the RPA/MRN complex interaction ..37-38
Figure 8 Model of the interaction of RPA and the MRN complex in response to
stalled replication forks …………………………………………...…39-40
Figure 9 HU-, CAMPT-, ETOP- and MMC-induced foci formation …………55-56
Figure 10 HU-, CAMPT-, ETOP- and MMC-induced hyperphosphorylation of RPA-
p34 …………………………………………………………………...57-58
Figure 11 ETOP- and MMC-induced generation of DNA DSBs ………………59-60
Figure 12 Prolonged exposure to HU induced DNA DSBs and loss of phospho-Nbs1
and RPA foci co-localization ……………………………...…………61-62
Figure 13 Model of the interaction of the MRN complex and RPA in response to
different DNA lesions …………………………………………..……63-64
Figure 14 Time- and dose-response of ETOP-induced Nbs1 phosphorylation and
RPA hyper-phosphorylation………………………………………….78-79
Figure 15 Depletion of RPA and Mre11 proteins using targeted siRNAs………80-81 vi
Figure 16 Treatment with siRPA or siMre11 does not affect ETOP-induced hyper-
phosphorylation of RPA-p34…………………………………………82-83
Figure 17 Depletion of RPA abrogates the ability of cells to form phospho-Nbs1 and
RPA-p34 foci following ETOP treatment……………………………84-85
Figure 18 Generation of recombinant ATM and ATR …………………………97-98
Figure 19 Generation of recombinant Mre11, Rad50 and Nbs1 …………….104-105
Figure 20 Generation of phosphorylated recombinant Mre11 ………………106-107
Figure 21 One- and two-dimensional SDS-PAGE of HeLa and recombinant RPA
demonstrate the presence of multiple RPA-p34 phosphorylations .114-115
Figure 22 Model depicting the interaction of the MRN complex and RPA in
response to DNA damage ……………………………………………...124
Table 1 Comparison of ataxia-telangiectasia (A-T), ataxia-telangiectasia-like
disorder (ATLD) and Nijmegen breakage syndrome (NBS) ……………. 4 1
CHAPTER 1. Background and Introduction
Cells within our body are continuously exposed to agents, both endogenous and exogenous, that damage the cellular DNA. This damage, if left unrepaired can cause mutations that may then lead to oncogenic transformation of the cell and progress to the development of cancer. In order to prevent this from occurring, cells have evolutionarily evolved complex and intricate pathways that respond to, and repair the damage before mutations occur.
DNA Damage Response: In response to DNA damage, cells activate an intricate web of signaling pathways known as the “DNA damage response” [1]. This damage response proceeds through a stepwise activation of specific proteins. First, sensor proteins recognize the DNA lesion or damaged DNA/ altered chromatin chromatin alterations and then activate structure transducers. Transducers convey the sensors damage signal to downstream transducers effectors, via protein phosphorylation, culminating in activation of cell-cycle effectors checkpoints, appropriate DNA repair cell cycle arrest DNA repair apoptosis pathways, or, in certain instances, Figure 1. DNA damage response. apoptosis [1, 2]. These repair activities are extremely important, emphasized by the fact that unrepaired DNA damage can be mutagenic, cytotoxic and/or carcinogenic [3]. The major protein transducers in the DNA 2
damage response are the phosphatidylinositol 3-kinase-like kinases (PIKKs) ataxia-
telangiectasia mutated (ATM), ATM- and Rad3-related (ATR) and DNA-dependent
protein kinase (DNA-PK). These kinases transduce the signal by phosphorylating key
effector proteins such as BRCA1, p53, MDC1, 53BP1, γH2AX, Nbs1, Mre11, RPA or
other kinases such as Chk1 and Chk2 [4, 5]. The sensors for the DNA damage response
are not definitively known and may vary depending on the type of damage. The list of
effector proteins continues to grow with time as new pathways and new components of
these pathways are identified.
Replication Stress: Stalled replication forks threaten DNA replication fidelity and
genomic integrity. Stalled forks can occur following deficiencies in replication substrates,
inhibition of replication machinery, or blocking of the replication machinery by DNA
damage or DNA-protein complexes [6, 7]. Replication fork stalling is also thought to
occur during normal DNA replication, particularly at DNA sequences that are prone to
form secondary structures (e.g. tRNA genes) or in regions where transcription complexes
may collide [7]. Stalled replication forks, if unresolved, progress to collapsed forks and
the generation of DNA strand breaks and genomic instability [8]. In order to prevent such
instability, replication stress activates the DNA damage response.
Foci: Many DNA damage response proteins localize to sites of damage where they form large protein aggregations termed nuclear foci. The role and function of these foci are not
entirely known, but have been hypothesized to be important for signal amplification and
control of damage-induced checkpoints, facilitation of repair of persistent damage, 3
compartmentalization of activated proteins, or simply a high local concentration of repair
proteins which may be required for certain biochemical steps of the repair process [1, 9-
11]. The composition of the foci depends on the nature of the lesion, and changes over
time as the DNA damage response progresses from damage recognition and signaling to
repair and resolution [12].
Mre11/Rad50/Nbs1 (MRN) Complex: The MRN complex (composed of Mre11, Rad50 and Nbs1) is highly conserved throughout evolution and orthologues of Mre11 and
Rad50 are found in all taxonomic kingdoms [13]. Interestingly, Nbs1 is only seen in higher eukaryotic systems and its orthologue, Xrs2, is only found in yeast [13]. The MRN complex plays a major role in the repair of DNA double-strand breaks (DSBs) [14, 15], dealing with misfolded DNA secondary structures that arise during DNA replication [16,
17], maintaining telomere length [18, 19] and activating cell cycle checkpoints [20-22].
Petrini and Stracker have postulated that the MRN complex acts as a sensor of DNA damage needed to activate the DNA damage response [23]. Recent data supports this hypothesis and suggests that the MRN complex functions as an activator of the ATM and
ATR kinases, as well as acting as an effector downstream of ATM/ATR in double-strand break repair and cell cycle checkpoints [20, 24-28]. In response to DNA damage induced by a variety of agents, the MRN complex re-localizes to nuclear foci [9, 29-32] where it is believed to play a role both in the recruitment of other protein factors to the foci as well as possible enzymatic activity within the foci [12, 14].
4
Genome Instability, Clinical Disease and the MRN Complex: Loss of ATM Clinical feature A-T ATLD NBS Ataxia + + - kinase activity leads to the genome Telangiectasia + - - Dysarthria + + - Abnormal eye movements + - - instability and neurodegenerative Immunodeficiency + - + Neurodegeneration + + - disease ataxia-telangiectasia (A-T) Pulmonary infections + NK + Cancer predisposition + NK + Skin abnormalities + NK + [33]. This disease is characterized by Microcephaly - - + Craniofacial abnormalities - - + ataxia, ocular telangiectasias, loss of Congenital malformations - - +
Cellular feature cerebellar function, immune defects, Chromosome translocations + + + Radiosensitivity + + + sterility, radiosensitivity and cancer Radioresistant DNA synthesis + + + Slow p53 accumulation + - - predisposition [34]. In 1999 it was +:presenceoffeature;-:absence;NK:notknown realized that there were a subset of Table 1. Comparison of ataxia-telangiectasia (A- T), ataxia-telangiectasia-like disorder (ATLD) and individuals diagnosed with A-T that Nijmegen breakage syndrome (NBS). had normal levels of ATM. Further study found that these patients actually had hypomorphic mutations for Mre11 [35]. These individuals were reclassified as having an ataxia-telangiectasia-like disorder (ATLD). The clinical features of patients with ATLD are very similar to those of A-T, but ATLD is characterized by a later onset of the neurological features and a slower progression of the disorder, giving an overall appearance of a milder condition than A-T in the early years [36] (Table 1). The clinical and molecular similarities between A-T and ATLD indicate that ATM and Mre11 probably function in similar pathways within the cell. It is interesting to note that hypomorphic mutations in the Nbs1 gene also cause a genomic instability disease, termed
Nijmegen breakage syndrome (NBS). NBS was first described in 1981 [37], but it wasn’t until 1998 that mutations in the Nbs1 gene were identified as the underlying cause of the 5
disease [38]. While NBS is clinically distinct from A-T and ATLD, there are a number of
clinical and cellular characteristics that the three diseases have in common (Table 1).
Mutations in Rad50 have recently been identified, with the phenotype resulting in
classification of the disease as a variant of NBS [1].
Mre11: Mre11 is the core of the MRN complex and interacts independently with both
Rad50 and Nbs1 [39-41]. Mre11 is best known for its nuclease activity (both exonuclease and endonuclease), but also has DNA binding, strand-dissociation and strand-annealing activities [42-47]. The functions of Mre11 are modulated in part by the presence/absence
of Rad50, Nbs1 and ATP [42, 43, 48-50]. Mre11 is phosphorylated in a DNA damage-
and DNA replication-dependent manner [16, 51-53]. The functional significance of this
phosphorylation is not known, but it has been suggested that phosphorylation may
increase the nuclease activity of Mre11 [16].
Rad50: The overall structure of Rad50 is similar to a class of proteins known as the
‘structural maintenance of chromosome’ (SMC) proteins, which are involved in
chromosome condensation and sister-chromatid cohesion [54]. Located on the amino-
and carboxy-terminal ends of Rad50 are two ATP-binding domains termed Walker A and
B respectively. Rad50 forms an anti-parallel homodimer that brings together the Walker
A and B binding motifs of separate Rad50 molecules to create a functional ATP-binding
domain [47]. The proposed main function of Rad50 is to bind DNA ends and hold them
in close proximity [44].
6
Nbs1: As mentioned previously, Nbs1 is found in higher eukaryotes and its orthologue
Xrs2 is found in yeast, while other taxonomical kingdoms to date do not have a known
orthologue or functional equivalent. The amino-terminal region of Nbs1 contains a
forkhead-associated domain (FHA domain) and a BRCA1 carboxy-terminal domain
(BRCT domain) [38], both of which are involved in protein-protein interactions in a phosphorylation-dependent manner [55-58]. These domains are essential for interaction of the MRN complex with γH2AX, WRN and possibly other proteins. Loss of the FHA domain blocks the ability of the MRN complex to form foci at sites of DNA damage [59].
The carboxy-terminal region of Nbs1 binds to the Mre11/Rad50 complex [59]. Nbs1 also contains the nuclear localization signal for the entire MRN complex, and without Nbs1,
Mre11/Rad50 remains cytoplasmic [59]. Nbs1 is phosphorylated in a DNA damage-
dependent manner on serine residues 278 and 343 both in vitro and in vivo [60].
Phosphorylation on these residues following IR-induced damage is predominantly via
ATM, but ATR may also play a role, particularly at later time points [41, 61-63]. Site directed mutagenesis ablating the ability of Nbs1 phosphorylation at serine 343 prevents the S-phase checkpoint that is normally activated following IR treatment and also the phosphorylation of SMC1 and FANCD2, both downstream targets of ATM [64-67]. It is not known if other functional significance is associated with Nbs1 phosphorylation.
Replication Protein A (RPA): Replication Protein A (RPA), the major single-stranded
DNA (ssDNA) binding protein in eukaryotic cells, is a heterotrimer composed of 70
(p70), 34 (p34) and 14 kDa (p14) subunits [68]. RPA is essential for DNA replication,
recombination and repair [68]. RPA is also able to bind to double-stranded DNA 7
(dsDNA) and RNA, though with an affinity at least three orders of magnitude lower than that for ssDNA [69-71]. RPA has been shown to unwind dsDNA, but the data is consistent with this activity being due to a helix-destabalizing activity instead of a helicase-like activity [72-74]. This unwinding is stimulated by phosphorylation of RPA
[72]. RPA undergoes a conformational change when it binds to ssDNA which involves both the p70 and p34 subunits, and results in RPA becoming a better substrate for phosphorylation [75, 76]. The p34 subunit is phosphorylated in a cell cycle- and DNA damage-dependent manner. A phosphorylated form, often termed the mitotic form [77], is present beginning at the G1/S transition and extending until late mitosis [78, 79].
Following DNA damage by a wide variety of agents, such as ionizing radiation (IR) and ultraviolet radiation (UV), RPA-p34 is hyper-phosphorylated [77, 80-84]. These phosphorylation events occur predominantly within the N-terminal 33 residues.
Phosphorylation does not appear to alter the ability of RPA to bind to ssDNA [85, 86], but does induce a conformational change due to altered intersubunit interactions that may regulate the interaction of RPA with DNA and other proteins [85]. RPA accumulates along stretches of ssDNA generated by stalled replication forks and/or DNA damage [31,
68], suggesting that the RPA/DNA complex created by the accumulation of RPA at these sites may signal the presence of damage and activate the DNA damage response. In support of this sensor activity, Dart et al. have shown that recruitment of ATR to nuclear foci following replication fork stalling is dependent on RPA [87]. Additionally RPA promotes DNA binding and activation of ATR/ATRIP in vitro [88]. RPA is also required to recruit and activate Rad17 complexes for checkpoint signaling in vivo [89]. Further evidence that RPA acts as a common intermediate for signaling stalled replication 8
forks/DNA damage is demonstrated by the RPA-dependent binding of Cut5 to chromatin following DNA damage and its subsequent recruitment of ATR and DNA polymerase α to chromatin [90]. In that report, Parrilla-Castellar and Karnitz put forth a model suggesting that ssDNA generated at a stalled replication fork is coated with RPA, leading to Cut5 chromatin association. Cut5 then facilitates the chromatin association of ATR and DNA Polα, which, in turn, leads to the loading of the Rad9-Rad1-Hus1 complex
[90]. These data have given credence to the hypothesis that RPA-coated ssDNA acts as a common intermediate for signaling stalled replication forks and/or DNA damage, and subsequent recruitment and activation of DNA damage response proteins. It is likely that
RPA plays a dual role in the damage response network: that of a sensor of damage as well as an effector. Replication protein A is an essential component of most DNA repair processes.
MRN Complex and RPA in the DNA
damaged DNA/ Damage Response: Both the MRN altered chromatin structure complex and RPA are thought to M R N play critical roles in the DNA RPA sensors damage response by initially sensing ATM transducers ATR the damage and facilitating the effectors activation of ATM and ATR. These M R N RPA protein complexes may also be cell cycle arrest DNA repair apoptosis classified as effectors. Both Figure 2. Roles of the MRN complex and RPA in the DNA damage response. complexes can be phosphorylated by 9
ATM and ATR following DNA damage, and phosphorylation modifies their function.
Both the MRN complex and RPA are also found in nuclear foci during DNA replication
and following DNA damage.
Specific Goals and Questions: Evidence supports the hypothesis that the MRN complex
and RPA have some similar functions in the DNA damage response, and have led us to
ask the question: Do the MRN complex and RPA interact in the DNA damage response?
Demonstration that the MRN complex and RPA interact has lead to additional questions
addressing this interaction. (1) Is this interaction similar for all forms of DNA damage?
(2) Does protein phosphorylation play a role in this interaction? (3) What is the functional
consequence or significance of this interaction?
Environmental Health Relevance: A variety of different DNA lesions arise by three main
mechanisms [91]. (1) Environmental electromagnetic energy such as ionizing radiation
(IR), ultraviolet light (UV) and numerous environment genotoxicants can damage DNA.
Left unrepaired, this damage can lead to mutations that may progress to neoplastic
transformation. It has been estimated that as many as 85% of cancer deaths can be attributed to environmental carcinogens [92]. (2) Byproducts of normal cellular metabolism, such as reactive oxygen species, can alter the DNA. (3) Some chemical bonds in DNA tend to spontaneously disintegrate under physiological conditions. These include deamination reactions that result in miscoding base changes as well as hydrolysis of nucleotides that lead to generation of abasic sites.
10
It is now becoming well documented that mutagenesis is the “fundamental cornerstone of
the molecular basis of all forms of cancer” [93]. Under normal conditions, cells activate the DNA damage response to ensure the damaged DNA is repaired and genome integrity maintained, or that the cell is eliminated via apoptosis. When the DNA damage response is compromised in any way, there is an increased rate of accumulation of mutations, and therefore an increased risk of development of cancer due to exposure to environmental or endogenous agents [92]. Mutations in a number of different genes whose protein products are involved in the DNA damage response lead to clinical disease syndromes characterized by chromosomal instability, increased sensitivity to environmental agents and an increased risk of cancer. In addition to the three syndromes discussed previously
(A-T, ATLD and NBS), the most notable include xeroderma pigmentosum (XP), hereditary non-polyposis colon cancer (HNPCC), BRCA1 gene-associated forms of breast cancer, Bloom’s syndrome, Werner syndrome and Fanconi anemia [92]. Loss of other proteins involved in the DNA damage response have not been observed in the human population, and the genetic knock-out of these genes in laboratory animals results in embryonic lethality. These proteins include members of the MRN complex, ATR,
Rad51 and others [94-97], verifying the importance of these proteins in maintaining genomic stability.
Clinical Relevance: A comprehensive understanding of the DNA damage response will provide critical insights that may translate into treatments for patients with cancer predisposition diseases as well as treatments or preventive measures for the general population. Errol C. Friedberg, M.D., a clinical pathologist world-renowned for his 11
research in DNA repair responses, noted the potential for research in this field with the
following statement:
“The study of biological responsiveness to DNA damage embraces DNA repair, mutagenesis, damage tolerance, cell-cycle checkpoint control, programmed cell death, and other cellular responses to genomic insult. This integrated field is now deciphering the complex regulatory pathways transduced by signaling mechanisms that detect DNA damage and/or arrested DNA replication. As these pathways become better understood, parallel technological gains in gene therapy and therapeutic intervention by rational drug design will offer new strategies for blocking the unwanted consequences of DNA damage, especially cancer.” [93]
Applying a detailed understanding of the molecular mechanisms underlying cancer for targets of therapeutic intervention is becoming a reality. The United States Food and
Drug Administration recently approved a small molecule kinase inhibitor for treatment of patients with chronic myeloid leukemia and gastrointestinal stromal tumors (for Review, see [98]). The conceptual basis of this specific therapy stemmed from the understanding of the molecular events associated with these cancers, and how these events deviate from normal cellular processes. In a review discussing these two specific cases and other potential molecules for “targeted therapy”, Charles Sawyers, MD stated, “The era of molecular targeted cancer therapy has clearly arrived… The successes of the past few years illustrate the power of the approach and should reinforce the need to continue basic studies of the molecular underpinnings of human cancers.” [99] 12
CHAPTER 2. Replication Protein A (RPA) and the Mre11/Rad50/Nbs1 (MRN)
Complex Co-localize and Interact at Sites of Stalled Replication Forks
I. INTRODUCTION
Stalled replication forks threaten DNA replication fidelity and genomic integrity. Stalled forks occur following deficiencies in replication substrates, inhibition of replication machinery, or blocking of the replication machinery by DNA damage or DNA-protein complexes [6, 7]. Replication fork stalling, with subsequent replication stress, is also thought to occur during normal DNA replication, particularly at DNA sequences that are prone to form secondary structures (e.g. tRNA genes) or in regions where transcription complexes may collide [7]. Failure to resolve this replication stress may result in the collapse of stalled forks and genomic instability [8]. In order to prevent such instability, replication stress triggers the activation of a DNA damage response. This response involves the recruitment and activation of proteins involved in DNA repair and cell cycle regulation. This DNA damage response uses proteins to detect damage, signal the site of damage, and transduce and amplify the signal in order to activate needed effector proteins. Many proteins involved in these three aspects accumulate and form large nuclear foci following DNA damage. Examples of such proteins include γH2AX, 53BP1,
ATM, ATR, BRCA1, Werner’s protein, the MRN complex, and RPA [100, 101]. The functions of these foci are not fully understood, but they may represent sites of fork reactivation, protein activation, DNA repair and/or non-repairable damage [9, 17, 29, 30].
13
Replication Protein A (RPA), the major single-stranded DNA (ssDNA) binding protein in eukaryotic cells, accumulates along stretches of ssDNA generated by stalled replication forks and/or DNA damage [31, 68]. It has been suggested that the RPA/DNA complex created by the accumulation of RPA at these sites may signal the presence of damage and activate the DNA damage response. In support of this, Dart et al. have shown that recruitment of ATR to nuclear foci following replication fork stalling is dependent on
RPA [87]. Additionally, it has been shown that RPA promotes DNA binding and activation of ATR/ATRIP in vitro [88]. RPA is also required to recruit and activate
Rad17 complexes for checkpoint signaling in vivo [89]. Further evidence that RPA acts as a common intermediate for signaling stalled replication forks/DNA damage is demonstrated by the RPA-dependent binding of Cut5 to chromatin following DNA damage and its subsequent recruitment of ATR and DNA polymerase α to chromatin
[90]. In that report, Parrilla-Castellar and Karnitz put forth a model suggesting that ssDNA generated at a stalled replication fork is coated with RPA, leading to Cut5 chromatin association. Cut5 then facilitates the chromatin association of ATR and DNA
Polα, which, in turn, leads to the loading of the 9-1-1 complex [90]. These data have given credence to the hypothesis that RPA-coated ssDNA acts as a common intermediate for signaling stalled replication forks and/or DNA damage, and subsequent recruitment and activation of DNA damage response proteins. It is likely that RPA plays a dual role in the damage response network: that of a sensor of damage and also as an effector.
Replication protein A is an essential component of most, if not all, DNA repair processes.
14
Petrini and Stracker have postulated that, similar to RPA, the MRN complex acts as a
sensor of DNA damage needed to activate the DNA damage response [23]. Recent data
suggests that the MRN complex functions as a damage sensor upstream of ATM/ATR
activation, in addition to acting as an effector downstream of ATM/ATR in double strand
break repair and cell cycle checkpoints [20, 24, 25, 102]. Although it is known that the
MRN complex binds to DNA ends, there is no clearly defined mechanism by which the
MRN complex recognizes other types of DNA damage.
Since both RPA and the MRN complexes are believed to be involved in the sensing and
signaling of DNA damage, we postulated that these two complexes might interact at sites
of stalled replication forks and DNA damage. To test this hypothesis, we investigated the interaction of these proteins in response to hydroxyurea- (HU) and ultraviolet light- (UV) induced replication stress and DNA damage. Following treatment with these agents, RPA and Mre11 became phosphorylated and co-localized to discrete chromatin-bound nuclear foci. RPA and the MRN complex also co-immunoprecipitated together, suggesting that these proteins interact, either directly or indirectly. The interaction between RPA and the
MRN complex was abrogated by phosphatase treatment. Taken together, our data suggest that following replication stress induced by HU or UV, RPA and the MRN complex co- localized and interacted at sites of stalled replication forks, and that the protein
interaction may be mediated, at least in part, by protein phosphorylation.
15
II. MATERIALS AND METHODS
Cell lines and treatments: HeLa cells were obtained from American Type Culture
Collection (ATCC; Manassas, VA) and maintained at 37°C and 5% CO2 in Dulbecco’s
Modified Eagles Medium (DMEM; Gibco, Gaithersburg, MD) supplemented with 10%
fetal bovine serum (FBS; Hyclone, Logan, UT) and 1% penicillin-streptomycin (Gibco).
For UV exposure, the growth medium was removed (and held at 37°C) and cells were
washed with phosphate buffered saline (PBS). The PBS was replaced with minimum
essential medium (MEM; Gibco) without phenol red and cells were treated with 30 J/m2
UVC (for asynchronous cells) or 20 J/m2 (for cells synchronized in S-phase) using a low-
pressure mercury lamp (Mineralight lamp; model UVG-11; UVP, Inc., San Gabriel, CA)
with a maximal output at 254 nm. Following UV exposure, the MEM was removed and
replaced with the original growth medium, and cells were allowed to recover for 8 h at
37°C before harvesting. For hydroxyurea (HU; Sigma-Aldrich, St. Louis, MO)
treatment, cells were incubated in growth medium containing HU (2 mM) for 3 h before harvesting.
Western Immunoblots: Cell lysates and immunoprecipitates were separated on 12% SDS- polyacrlyamide gels and transferred to PVDF membranes (Millipore Corp., Bedford,
MA). Membranes were probed using the following primary antibodies: anti-Mre11
(Novus Biological, Littleton, CO; 1:20,000), anti-Mre11 (GeneTex, San Antonio, Texas;
1:10,000), anti-RPA-p34 (Neomarkers, Freemont, CA; 1:5000) and anti-RPA-p34-SP4-
SP8 (Bethyl Laboratories, Inc., Montgomery, TX; 1:10,000). Secondary antibodies were
horseradish peroxidase-linked anti-rabbit and anti-mouse (Amersham Biosciences, 16
Buckinghamshire, England; 1:3000), and bound antibodies were visualized using
chemiluminescent detection.
Cell Synchronization: Cells were synchronized in S-phase and G1-phase of the cell cycle
as previously described [77]. To synchronize in S-phase, cells were incubated in growth
medium containing aphidocolin (final concentration of 1 µM; Sigma-Aldrich) for 15 h.
Following incubation, the aphidocolin-containing medium was removed, cells were washed with PBS, and then incubated in fresh medium without aphidocolin for an
additional 2 h at 37°C. For nocodazole synchronization in G1- or S-phase of the cell
cycle, cells were incubated in medium containing nocodazole (0.3 µM final concentration; Sigma-Aldrich) for 17 h. The mitotic cells, which become detached from the culture dish as they enter M-phase, were collected and pelleted at 500 x g. Mitotic cells were released from nocodazole treatment by incubating in fresh medium. We have shown previously that HeLa cells enter G1-phase about 5 h after release and enter S- phase about 12 h after release [77]. For the experiments reported here, cells were treated
with HU from 4.5 h to 7.5 h after release (for G1-phase) or from 12 h to 15 h after release
(for S-phase) and then harvested.
Subcellular Fractionation: The cellular protein fractionation protocol was performed as previously described with slight modifications [103]. Briefly, S-phase synchronized
HeLa cells were treated with either HU, UV, or mock-treated. The free cytoplasmic/nucleoplasmic fraction was prepared by allowing cells to lyse for 10 min on ice in 0.5% Triton X-100 in cell lysis buffer (50 mM Tris-HCl, pH 7.5; 150 mM NaCl; 17
0.1% NP-40; 5 µl/mL pepstatin; 5 µg/mL leupeptin; 5 µg/mL aprotonin; 10 mM NaF; 10 mM β-glycerophosphate; 1 mM Na3VO4; 1 mM PMSF). The insoluble fraction was pelleted by centrifugation at 13,000 x g for 10 min at 4○C and the supernatant (free cytoplasmic/nucleoplasmic; fraction FCN) was collected. The pellet was washed with
PBS, and treated with 100 µg/mL DNase I in cell lysis buffer at 37○C for 15 min, followed by addition of ammonium sulfate (250 mM final concentration) and further incubation at room temperature for 10 min. The insoluble fraction was pelleted by centrifugation at 13,000 x g for 10 min at 4○C and the supernatant was collected and designated as the chromatin-bound fraction (fraction CB). The remaining insoluble material was washed with PBS, suspended in SDS buffer (2% SDS; 20 mM Tris-HCl, pH
7.5; 1 M β-mercaptoethanol; 10% glycerol), and designated the nuclear matrix fraction
(fraction NM). The presence of glucose-3-phosphate dehydrogenase, Histone H1 and
Lamin A/C within the free nucleoplasmic/cytoplasmic fraction, chromatin-bound fraction and nuclear matrix fraction respectively verified the validity of this fractionation method.
Immunofluorescence: Cells were grown on 18 mm or 12 mm coverslips (Becton
Dickinson Labware, Bedford, MA) for 20-24 h prior to treatment. Cells were treated with
30 J/m2 UVC (asynchronous cells) and allowed to recover for 8 h or with 2 mM HU for 3 h. After treatment, cells were washed with PBS, then washed with PBS containing 0.5%
Triton X-100, and fixed for 5 min with PBS containing 3% paraformaldehyde (Fisher
Scientific, Hampton, NH). Cells were then blocked for 1 h in PBS containing 15% FBS.
Primary antibody dilutions used are as follows: anti-RPA-p34-SP4-SP8 1:2000 (Bethyl
Laboratories, Inc.), anti-RPA 1:1000 (Neomarkers), anti-Mre11 1:500 (Novus 18
Biological), anti-Mre11 1:500 (Genetex), anti-γH2AX 1:300 (Upstate Cell Signaling
Solutions, Waltham, MA) and anti-Wrn 1:300 (Novus Biological). Secondary antibody dilutions are as follows: anti-rabbit Alexa Fluor 488 1:250 and anti-mouse Alexa Fluor
568 1:250 (Molecular Probes, Eugene, OR). Images were captured with a Nikon inverted fluorescent microscope with attached CCD camera at 100 X magnification and processed using Photoshop 7.0 (Adobe) software.
Co-immunoprecipitation assays: For co-immunoprecipitation reactions, 50 µL of protein
G-agarose beads (Invitrogen, Carlsbad, CA) were incubated with 3.0-5.0 µg of anti-
Mre11 (Novus Biological), anti-RPA-p70 (Bethyl Laboratories, Inc.), or normal rabbit
IgG (Oncogene, San Diego, CA) antibodies in PBS for 1 h at room temperature with end- over-end mixing. Following the addition of approximately 1000 µg of cell lysate, the immunoprecipitation reactions were incubated for 20-24 h at 4oC with end-over-end mixing. The immunoprecipitates were separated from the supernatant by centrifugation and washed with PBS containing 0.05% NP-40. Proteins were extracted from the agarose beads by resuspending in 1 X Laemmli gel-loading buffer and separated on 12%
SDS-polyacrlyamide gels.
When antibodies cross-linked to the agarose beads were used, 50 µL of protein G-agarose beads (Invitrogen) were incubated with 3.0 – 5.0 µg of anti-Mre11 (Novus Biological) for
1 h at room temperature with end-over-end mixing. The beads were washed twice with
200 mM triethanolamine (Sigma-Aldrich) and then incubated in 20 mM dimethylpimelimidate (DMP; Sigma-Aldrich) in 200 mM triethanolamine for 30 min at 19
room temperature. The DMP solution was replaced with 50 mM Tris-HCl, pH 7.5 for an
additional 15 min. The Tris-HCl was removed and the beads were suspended in PBS and
stored at 4°C until addition of cell lysate.
Phosphatase Treatment: For phosphatase treatments of cellular lysates, 10 X NEBuffer 3
(New England BioLabs, Beverly, MA; 100 mM NaCl, 50 mM Tris-HCl, 10 mM MgCl2,
1 mM dithiothreitol) was added to approximately 1000 µg total protein to give a final concentration of 1 X NEBuffer. 200 U of calf intestinal alkaline phosphatase (CIP) was added to the mixture and then incubated for 2 h at 37°C. For phosphatase treatment of immunoprecipitates, samples were prepared by washing immunoprecipitates with PBS to remove non-specifically bound proteins and remaining medium. For CIP treatment, the pellets were resuspended in 1X NEBuffer 3 with the addition of 50 U CIP. The reaction mixtures were incubated at 37°C for 2 h before collection of the Mre11-associated pellet and supernatant. For lambda phosphatase (λPPase) treatment, the pellets were resuspended in 1X λPPase buffer (New England Biolabs) with 2 mM MnCl2 or 1X
λPPase buffer with 1 mM Na2VO4 and 10 mM NaF (known inhibitors of λPPase). 400 U
of λPPase was added to each reaction tube and then incubated at 30°C for 30 min.
III. RESULTS
Replication stress induces hyperphosphorylation of RPA and phosphorylation of Mre11.
Both RPA-p34 and Mre11 are phosphorylated in a DNA damage-dependent and a DNA replication-dependent manner [16, 104]. To verify that phosphorylation of RPA-p34 and 20
Mre11 occurred after exposure of HeLa cells to 2 mM HU for 3 h and 20 J/m2 or 30 J/m2
UV, whole cell lysates from treated cells were visualized for RPA-p34 and Mre11 via western blotting. Similar to previous reports, these treatments resulted in the
hyperphosphorylation of the RPA-p34 subunit and phosphorylation of Mre11 (Figure 3A,
lanes 2-3, 5-6; Figure 3B, lanes 2-3, 8-9). When cells were synchronized in S-phase prior to treatment with HU or UV, the proportion of hyperphosphorylated-RPA and phosphorylated-Mre11 increased (Figure 3A, lanes 5-6; Figure 3B, lanes 8-9). This increase in RPA and Mre11 phosphorylation in treated S-phase cells is most likely due to
the replication stress induced by these agents. An antibody that recognizes
phosphorylated serine 4 and serine 8 of RPA-p34 specifically recognized only the DNA
damage-dependent hyperphosphorylated form of RPA-p34 (Figure 3B, lanes 4-6, 10-12).
The phosphorylation of RPA and Mre11 suggests that these proteins are involved in the cellular response to replication stress.
Phosphorylated RPA and Mre11 are more tightly bound to the chromatin. It has been previously reported that following DNA damage, RPA becomes re-distributed within the nucleus and is found predominantly bound to the chromatin [103]. Mre11, which has increased chromatin binding in S-phase, does not show a re-distribution within the nucleus following DNA damage in any stage of the cell cycle [29, 105]. In order to investigate the nuclear re-distribution of the phosphorylated isoforms of RPA and Mre11 following replication fork stalling and DNA damage, cellular proteins of S-phase synchronized cells were separated into three fractions: (1) free cytoplasmic/nucleoplasmic, (2) chromatin-bound, and (3) nuclear matrix fractions. To 21
confirm the separation of the three fractions, we verified the presence of glucose-3-
phosphate dehydrogenase (a cytosolic protein), histone H1 and lamin A/C (a nuclear
matrix protein) within the free nucleoplasmic/cytoplasmic fractions, chromatin-bound
fractions and nuclear matrix fractions, respectively (data not shown). In mock-treated
cells, approximately 60% of the total RPA was in the free cytoplasmic/nucleoplasmic
fraction; whereas Mre11 was present in all three fractions, with the chromatin-bound fraction containing the largest percent (Figure 4A, lanes 2-3; Figure 4B). Following HU
or UV treatment, there was a redistribution of RPA to the chromatin-bound fraction
(Figure 4A, lanes 6-8, 10-12; Figure 4B), with the percent of total RPA changing from
40% chromatin-bound in mock-treated cells to approximately 85% in HU or UV treated
cells. As previously reported [101], the damage-induced hyperphosphorylated isoform of
RPA was found predominantly in the chromatin-bound fraction. While the distribution of
Mre11 between the different fractions did not change after damage, the majority of the
damage-induced phosphorylated form of Mre11 was found in the chromatin-bound
fraction (Figure 4A, lanes 7, 11; Figure 4B).
RPA and Mre11 co-localize to discrete nuclear foci following HU or UV treatment.
Following the induction of DNA damage, many proteins associated with DNA damage
signaling, DNA repair and cell cycle checkpoints become localized to sites of damage
and form nuclear “foci” [100]. In order to visualize these foci and to determine if Mre11
and RPA were present in these “repair foci,” we used immunofluorescent staining. Pre-
extraction of cells with a detergent-containing buffer removes the nucleoplasmic and
cytoplasmic proteins, leaving behind the chromatin-bound and matrix-associated proteins 22
[29]. In mock-treated cells stained with antibodies against RPA and Mre11, there was a
diffuse nuclear staining. Following HU or UV treatment, there was a redistribution of
RPA and Mre11 to discrete foci that co-localized (Figure 5A, Panels D, H and L).
Although studies have shown that both RPA and the MRN complex are able to form
nuclear foci following DNA damage by various genotoxic agents [9, 29, 31, 32, 106-
108], no previous reports have indicated that RPA and Mre11 foci co-localize. We also
looked for co-localization of Mre11 with hyperphosphorylated-RPA using the RPA-p34-
SP4-SP8 phospho-specific antibody. Under mock-treatment conditions, there is no
staining with the phospho-specific RPA antibody as expected (Figure 5B, Panel B).
Following treatment with HU or UV, similar to staining with the antibody that recognizes
all isoforms of RPA, phosphorylated-RPA aggregated into nuclear foci that co-localized
with Mre11 (Figure 5B, Panels D, H and L).
In addition to co-localization of RPA and hyperphosphorylated-RPA with Mre11, and
similar to previous reports, these foci also showed limited co-localization with
phosphorylated histone H2AX (γH2AX) [29, 32, 109, 110] and with Werner’s protein
(Wrn) (data not shown) [101, 106]. γH2AX foci formation is a marker of double-strand
breaks (DSBs) and has also been reported to form following replication stress and at
replication forks [17, 29, 111]. Wrn protein is a RecQ-class DNA helicase that localizes
to sites of stalled replication where it is involved in the resolution and prevention of
aberrant recombination events [101], and is able to directly interact with RPA [101, 112]
and the Mre11 complex [113]. Co-localization of hyperphosphorylated-RPA and the
MRN complex with γH2AX and Wrn, along with previous reports [29, 101] indicate that 23
these foci are at sites of stalled replication forks, which at the time points investigated may have progressed to the point of collapse and generation of DSBs.
RPA and the MRN complex interact. The observation of co-localization of RPA and the
MRN complex raised the possibility that these proteins may directly interact. We
performed co-immunoprecipitation experiments using whole cell lysates of S-phase-
synchronized HeLa cells from HU, UV or mock-treated cells to probe for such
RPA/MRN complex interactions. Rabbit anti-Mre11, anti-Rad50 or anti-Nbs1 antibodies were able to immunoprecipitate RPA, and rabbit anti-RPA-p70 antibodies were able to
immunoprecipitate Mre11 (Figure 6A, lanes 7-12; Figure 6B, lanes 3-5; Figure 6C).
Normal rabbit IgG did not immunoprecipitate RPA or Mre11 (Figure 6A, lanes 4-6;
Figure 6B, lane 6), demonstrating the co-immunoprecipitation of RPA and Mre11 was
not due to non-specific antibody binding.
RPA is a very abundant protein in the cell. Therefore we considered the possibility that the apparent co-immunoprecipitation of RPA with Mre11 using anti-Mre11 antibodies could be non-specific and simply due to RPA’s abundance. In addition, the co- immunoprecipitation of Mre11 with RPA using anti-RPA-p34 antibodies was barely detectable and co-immunoprecipitation using anti-RPA-p70 antibodies was present, but at very low levels. To demonstrate that the co-immunoprecipitation of RPA and MRN was specific, we carried out the following experiment. Cell lysates were incubated with anti-
Mre11 antibodies cross-linked to protein-G agarose beads, bound protein was eluted with sodium citrate (pH 3.0), neutralized and then immunoprecipitated again with anti-RPA- 24
p70, anti-Mre11 or anti-IgG antibody coated agarose beads. The sequential
immunoprecipitations removed the excess RPA from the lysate, allowing for a more
stoichometrically equal amount of RPA and MRN complex to interact in the second
round. The ability of anti-RPA-p70 antibodies to immunoprecipitate Mre11 was
increased under these conditions (Figure 6B, lanes 3 and 4), confirming the specificity of
the interaction.
In addition to co-immunoprecipitation of RPA and MRN in lysates from damaged cells,
co-immunoprecipitation was also observed in lysates from mock-treated cells (Figure 6A,
lane 7; Figure 6B, lane 3). This suggested that the interaction might not be dependent on
DNA damage. It has been shown that the MRN complex is required for the resolution of
breaks that occur spontaneously during DNA replication [16], so we considered the
possibility that the co-immunoprecipitation in the mock-treated samples might be an S- phase phenomenon due to normal replication. To investigate this possibility,
immunoprecipitation reactions were done using whole cell lysates from mock-treated and
HU-treated cells synchronized in G1-phase or S-phase of the cell cycle. The amount of
RPA that co-immunoprecipitated with Mre11 in mock-treated cells in S-phase of the cell
cycle was six-times more than mock-treated G1-phase cells as measured by densitometry
(Figure 6D, lanes 1 and 3). Following HU treatment, the amount of immunoprecipitated
RPA increased eight-fold when comparing S-phase to G1-phase. (Figure 6D, lanes 2 and
4). This suggests that the interaction between RPA and the MRN complex, while present
in both G1- and S-phase of the cell cycle, is increased when cells are in S-phase, and
increased to a greater extent following damage in S-phase. The increase in interaction in 25
mock-treated cells in S-phase may represent interactions that occur at spontaneously
stalled replication forks during normal DNA replication.
We also considered the possibility of an indirect interaction between RPA and the MRN
complex mediated by independent binding of both protein complexes to DNA. To test
this possibility, we pre-treated lysates with ethidium bromide (EtBr) or DNase I, or
treated immunoprecipitates with DNase I. EtBr is a DNA intercalator known to disrupt
DNA-protein interactions [114, 115]. Pretreatment of lysates with 50 µg/mL EtBr did not
alter the ability of anti-Mre11 antibodies to immunoprecipitate RPA (Figure 6E, lane 9).
DNase I treatment of lysates or immunoprecipitates, using the same conditions as the sub-cellular fractionation protocol, did not affect the RPA/MRN complex interaction as well (Figure 6E, lanes 3 and 6 respectively), suggesting this interaction is not indirect via
DNA-protein interactions.
Phosphatase treatment abrogates the RPA/ MRN complex interaction. The Nbs1 protein of the MRN complex contains two domains associated with protein-protein interactions, the forkhead-associated domain (FHA) and the BRCA1 carboxy-terminal domain
(BRCT) [38, 116]. FHA and BRCT domains are both known to mediate protein-protein interactions in a phosphorylation-dependent manner [55, 57], which is demonstrated in the interaction of Nbs1 with γH2AX and BRCA1 with BACH1 [57, 109]. Since increased
RPA/MRN complex interactions occur under conditions of increased RPA-p34 phosphorylation, specifically RPA hyperphosphorylation, we wanted to investigate if protein phosphorylation played a part in the RPA/MRN complex interaction. To address 26
this question, whole cell lysates from HU-treated cells were incubated for 2 h at 37°C with or without calf intestinal phosphatase (CIP) before incubation with anti-Mre11 antibody coated agarose beads. Pre-treatment with CIP abrogated the ability of anti-
Mre11 antibodies to immunoprecipitate RPA (Figure 7A, lane 3), while the 2 h incubation at 37°C had no effect (Figure 7B, lane 2). To verify that the CIP treatment
resulted in protein de-phosphorylation, the supernatant from the CIP treated sample was
analyzed for RPA-p34. Loss of the hyper-phosphorylated form of RPA-p34 and retention
of non-phosphorylated RPA (Figure 7B, lane 4) indicated that protein de-phosphorylation
had occurred.
To continue investigating the effect of phosphatase activity on the RPA/MRN complex interaction, we immunoprecipitated proteins from whole cell lysates of HU-treated cells with anti-Mre11 antibody coated beads. The resulting immunoprecipitate treated with
CIP demonstrated a loss of RPA in the Mre11-associated pellet and appearance of non- phosphorylated RPA in the supernatant (Figure 7B, lanes 2 and 3).
To verify that the abrogation of the RPA/MRN interaction was dependent upon phosphatase activity and not just the physical presence of the phosphatase enzymes, we used another phosphatase, lambda phosphatase (λPPase), and specific phosphatase inhibitors. Immunoprecipitates from HU-treated lysates immunoprecipitated with anti-
Mre11 antibodies were treated with lambda phosphatase (λPPase) with (Figure 7C, lanes
5 and 6) and without the presence of specific λPPase inhibitors (sodium orthovanadate and sodium fluoride; Figure 7C, lanes 3 and 4). The addition of the λPPase inhibitors 27
preserved the RPA/MRN complex interaction (and the presence of phosphorylated-RPA) with the anti-Mre11 antibody coated beads (Figure 7C, lanes 5 and 6), suggesting that the abrogation of the RPA/MRN complex interaction was indeed dependent upon phosphatase activity. Together, these data suggest that the RPA/MRN complex interaction may be mediated by protein phosphorylation. These data, however do not address whether the abrogation of the RPA/MRN complex interaction is due to de- phosphorylation of RPA, the MRN complex or some other protein. 28
Figure 3. Phosphorylation of RPA and Mre11 following HU and UV treatment.
Asynchronous: Asynchronous, sub-confluent HeLa cells were treated with 2 mM HU for
3 h; or 30 J/m2 UV with 8 h recovery. S-Phase: HeLa cells synchronized with aphidicolin
(1 µM, 15 h) and allowed to enter S-phase were treated with 2 mM HU for 3 h; or 20
J/m2 UV with 8 h recovery. Whole cell lysates prepared from these cells were separated
on a 12% SDS-PAGE gel. A, Mre11 was visualized with a polyclonal antibody against
Mre11. Lanes 1 and 4, no treatment; lanes 2 and 6, UV treated; lanes 3 and 5, HU treated.
B, RPA-p34 was visualized using a monoclonal antibody against RPA-p34 (lanes 1-3, 7-
9) or a polyclonal antibody specific for RPA-p34 phosphorylated on serine 4 and serine 8
(anti-RPA-SP4-SP8, lanes 4-6 and 10-12). At least five different forms of the p34 subunit
of RPA can be visualized; form 1 is the fastest migrating band and is unphosphorylated,
while form 5 is the slowest migrating band and is the DNA damage-induced
hyperphosphorylated form. 29
FIGURE 3
A. Asynchronous S-Phase HU – – + – + – UV ––+ – – +
Mre11 Phosphorylated Mre11
1 23 4 5 6
B. Asynchronous S-Phase Anti-RPA Anti-RPA-SPP4-S 8 Anti-RPA Anti-RPA-SPP4-S 8 HU – – + – – + – + – – + – UV – + – ––+ ––+ – – +
RPA- Hyperphosphorylated p34 RPA-p34
1423 567 89 1011 12
30
Figure 4. Phosphorylated RPA and Mre11 are chromatin-bound. HeLa cells synchronized with aphidicolin (1 µM, 15 h) and allowed to enter S-phase were treated with 2 mM HU for 3 h, 20 J/m2 UV with 8 h recovery, or mock-treated. A, Whole cell lysates (WC; lanes 1, 5 and 9) and fractionated extracts were prepared as described in
“Materials and Methods.” Three different fractions were obtained: free cytoplasmic/nucleoplasmic (Fraction FCN, lanes 2, 6 and 10), chromatin-bound (Fraction
CB, lanes 3, 7 and 11) and nuclear matrix fractions (Fraction NM, lanes 4, 8 and 12).
Whole cell extracts and cell fractions were separated on a 12% SDS-PAGE gel and visualized with anti-Mre11 and anti-RPA antibodies. B, Densitometry measurements of the western blot depicted in panel A. The numbers reported are: percent of total Mre11 or RPA-p34 protein for each treatment group in the indicated lane, the ratio of the phosphorylated to non-phosphorylated form of Mre11 (form 2/form 1) for each lane, or the ratio of the hyper-phosphorylated to non-phosphorylated form of RPA-p34 (form
5/form 1) for each lane. 31
FIGURE 4
A. Mock-Treated HU UV Fraction WC FCN CB NM WC FCN CB NM WC FCN CB NM 2 Mre11 1
5 RPA 1 12 3 4 5 67 8 9101112
B. Mock-Treated HU UV
Fraction WC FCN CB NM WC FCN CB NM WC FCN CB NM
Anti-G3PDH (cytoplasmic)
Anti-H1 (chromatin)
Anti-Lamin A/C (nuclear matrix)
C. Mock-Treated HU UV Lane 2 3 4 6 7 8 10 11 12 Mre11 % of total Mre11 27 59 14 26 59 15 26 67 7 ratio of phosphorylated: non- phosphorylated (2/1) 0 0 0 0.013 0.1 0 0.051 0.24 0 RPA %oftotalRPA 61 39 0 19 81 0 13 87 0 ratio of hyper- phosphorylated: non- phosphorylated (5/1) 0 0 0 1.2 6.9 0 1.4 9.2 0 32
Figure 5. RPA and Mre11 co-localize to discrete nuclear foci following HU and UV treatment. Asynchronous HeLa cells were treated with 2 mM HU for 3 h, 30 J/m2 UV
and allowed 8 h to recover or mock-treated. Following extraction of cytoplasmic and
nucleoplasmic proteins with PBS containing 0.5% Triton X-100, cells were fixed in
paraformaldahyde, incubated in primary and secondary antibodies, and visualized by
fluorescent microscopy. A, RPA-p34 and Mre11 co-localize to HU- and UV- induced
nuclear foci. Cells were stained with anti-Mre11 antibodies (Green, Panels B, F and J)
and anti-RPA-p34 antibodies (Red, Panels C, G and K). Panels A, E and I are the DAPI-
stained nuclei, and Panels D, H and L are the merged images of the anti-Mre11 and anti-
RPA stained cells. B, Hyperphosphorylated-RPA-p34 and Mre11 co-localize to HU- and
UV- induced nuclear foci. Cells were stained with damage-induced hyperphosphorylation
specific anti-RPA-p34-SP4-SP8 antibodies (Green, Panels B, F and I) and anti-Mre11
antibodies (Red, Panels C, G and K). Panels A, E and I are the DAPI-stained nuclei, and
Panels D, H and L are the merged images of the anti-RPA-p34-SP4-SP8 and anti-Mre11 stained cells. 33
FIGURE 5
A. DAPI RPA-p34 ABA C D
E E F G H
+HU
I I J KL +UV
B. RPA-p34- DAPI S4PP-S8 ABC D
EFG H +HU
I J KL +UV 34
Figure 6. Co-immunoprecipitation of RPA and Mre11. A, HeLa cells synchronized with aphidicolin (1 µM, 15 h) and allowed to enter S-phase were treated with 2 mM HU for 3 h, 20 J/m2 UV and allowed 8 h to recover, or mock-treated. Whole cell lysates were incubated with agarose beads coated with non-specific anti-rabbit IgG (lanes 4-6), anti-
Mre11 (lanes 7-9) or anti-RPA-p70 (lanes 10-12) antibodies for 20-24 h. 10% of whole cell lysate volumes used for IP reactions were included as loading controls (Input, lanes
1-3). Proteins from the immunoprecipitates were detected by western blotting using anti-
RPA-p34 and anti-Mre11antibodies. B, S-phase synchronized HeLa cells were either treated with 2 mM HU for 3 h (lanes 2, 4-6) or mock-treated (lanes 1 and 3). Whole cell lysates were incubated with agarose beads cross-linked with anti-Mre11 antibodies as the primary IP. The proteins were eluted from the beads using sodium citrate and then incubated with anti-RPA-p70 (lanes 3 and 4), anti-Mre11 (lane 5) or anti-IgG (lane 6) antibody coated agarose beads for the secondary IP. The proteins were eluted from the second set of antibody-coated beads with Laemmli loading buffer and separated on a
12% SDS-PAGE gel. Proteins were visualized by western blotting using anti-RPA-p34 and anti-Mre11 antibodies. C, S-phase synchronized HeLa cells were treated with 2 mM
HU for 3 h. Whole cell lysates were incubated with agarose beads coated with anti-
Mre11, anti-Rad50 or anti-Nbs1 antibodies. The immunoprecipitated proteins were eluted with Laemmli loading buffer and subjected to western blotting using anti-RPA-p34 antibodies. D, HeLa cells were synchronized in M-phase of the cell cycle with nocodazole and allowed time to enter G1-phase (lanes 1 and 2) or S-phase of the cell cycle with aphidicolin (lanes 3 and 4) as described in “Materials and Methods.” Cells were either mock-treated or treated with 2 mM HU for 3 h before harvesting and whole 35
cell lysate formation. Lysates were incubated with anti-Mre11 antibody coated agarose beads and eluted off using Laemmli gel loading buffer. The immunoprecipitated proteins were subjected to western blotting using anti-RPA-p34 and anti-Mre11 antibodies. E, S- phase synchronized HeLa cells were treated with 2 mM HU for 3 h. Whole cell lysates were either treated with 100 µg/ml DNase I for 20 min at 37°C (lane 3) or 50 µg/ml ethidium bromide (EtBr) on ice for 30 min (lane 9). Pre-treated lysates, lysates without
DNase or EtBr incubated under similar conditions (lanes 2 and 8), as well as lysates with no previous incubation were immunoprecipitated with anti-Mre11 antibody coated agarose beads. The two IP reactions with non-treated lysates were washed with PBS and incubated for 20 min at 37°C with or without DNase I in cell lysis buffer (100 µg/ml, lanes 5 and 6 respectively). Proteins from all the immunoprecipitation reactions were eluted from the beads with Laemmli gel loading buffer and separated on 12% SDS gels.
The immunoprecipitated proteins were subjected to western blotting using anti-RPA-p34 antibodies. 36
FIGURE 6
A. B. st Input (10%) IP-IgG IP-Mre11 IP-RPA-p70 Input (10%) 1 IP-Mre11 HU ––+ –+ ––+ ––+ – HU ––+++ + UV –+– + – – – – + – – + 2nd IP ––RPA RPA Mre11 IgG
Mre11 Mre11
RPA RPA
1 23456789 1011 12 1 23456
C. D. G1-Phase S-Phase IP-Mre11 IP-Rad50 IP-Nbs1 HU ––+ + HU + + + Mre11
RPA
RPA 1 2 3 12 3 4 E.
Input IP-Mre11 Input IP-Mre11 Input IP-Mre11 HU + + + + + + HU + ++ DNase – – + – – + EtBr – – +
RPA
1 2 3 4 5 6 7 8 9 37
Figure 7. Phosphatase treatment disrupts the RPA/MRN complex interaction. A,
Lysates from S-phase synchronized HeLa cells subjected to 2 mM HU were either treated
with calf intestinal phosphatase (CIP) prior to incubation with anti-Mre11 antibody coated agarose beads (lane 3) or mock treated (lane 2). 10% of the whole cell lysate was used as a loading control (lane 1) and 10% of the supernatant (S) from the IP of the CIP treated lysate was loaded to verify protein de-phosphorylation (lane 4). B, Whole cell lysates from S-phase synchronized HeLa cells treated with 2 mM HU for 3 h were immunoprecipitated with anti-Mre11 antibody coated agarose beads. The immunoprecipitates were incubated with (lane 2) or without CIP (lane 1) and the proteins associated with the anti-Mre11 pellet (P) and the supernatant (S) (lane 3) were analyzed by western blotting with anti-RPA-p34 antibodies. C, S-phase synchronized HeLa cells treated with 2 mM HU were incubated with anti-Mre11 antibody coated agarose beads.
Immunoprecipitate pellets were washed with PBS and resuspended in phosphatase buffer with (lanes 3-6) or without λ phosphatase (lanes 1 and 2) or specific phosphatase inhibitors (lanes 5 and 6) as indicated. Samples were incubated at 30°C for 30 min and the immunoprecipitate pellets (P) and supernatants (S) separated by centrifugation and subjected to western blotting using anti-RPA-p34 antibodies. Whole cell lysate (10% of total protein used in IP reaction) was included as a loading control (Input, lane 1). 38
FIGURE 7
A. B. CIP – – ++ IP-Mre11 Input PSP CIP – + + PPS Mre11
RPA RPA 1 2 3 1 2 3 4
C. IP-Mre11 λ Phosphatase – – ++++ PPase Inhibitors ––––++ Input PP S P S
RPA
1 2 3 456 39
Figure 8. Model of the interaction of RPA and the MRN complex in response to stalled replication forks (see “Discussion” for details). 40
FIGURE 8
MRN complex associated with replication fork DNA lesion
MRN Pol δ Normal Replication Fork
RPA coated ssDNA
MRN Pol δ Stalled Replication Fork; Generation of a long stretch of RPA coated ssDNA
Recruitment and activation of ATR/ATRIP
P MRNP Pol δ P
P P P ATR Phosphorylation of RPA and P ATRIP P the MRN complex P ATR P ATRIP P P Recruitment of additional P MRN complex and other factors (e.g. Rad17, Rad9- Rad1-Hus1 complex, Wrn…)
P MRN Pol δ P P Phosphorylation-dependent P ATR P P ATRIP interaction of RPA and the P ATR P ATRIP P MRN complex P P P MRN P P
Checkpoint Resolution of NHEJ-mediated Recombination- activation Holliday junctions repair of like repair of and continued subsequent DSBs subsequent DSBs replication (Rad51- independent) 41
IV. DISCUSSION
The results presented here offer insight into the role of RPA and the MRN complex in the
replicative stress-induced DNA damage response. Upon treatment with HU or UV, RPA
becomes hyperphosphorylated and Mre11 is phosphorylated. The relative proportion of
the phosphorylated isoforms of RPA and Mre11 were increased when cells were synchronized in S-phase of the cell cycle prior to treatment. The phosphorylated isoforms of RPA and Mre11 were chromatin-bound; most likely at sites of stalled and/or collapsed replication forks [29]. While redistribution of phosphorylated RPA to chromatin-bound fractions has been previously reported [103], this may seem to be in contradiction to reports that show phosphorylated RPA has a decreased affinity for double-stranded DNA
(dsDNA) [85, 86]. Two possible explanations may account for this apparent discrepancy.
First, stalled replication forks lead to the generation of large regions of ssDNA. While
phosphorylated RPA has decreased affinity for dsDNA, it shows no difference in ssDNA
binding activity [86]. Alternatively, following replication fork stalling and/or DNA
damage, RPA may have increased interactions with a protein that is itself associated with the chromatin (possibly the MRN complex or some other, as yet, unidentified protein).
Either scenario could explain the increased chromatin-association of phosphorylated RPA
we observed following HU or UV treatment.
Consistent with a previous report showing constitutive chromatin association of the MRN
complex [105], we saw similar amounts of Mre11 in the chromatin-bound fraction in
mock-treated, HU and UV treated samples. This is also consistent with a report that
shows increased chromatin-association of the MRN complex in S-phase as compared to 42
G1 or G2/M, but no change in S-phase chromatin-association with or without damage
[29]. Interestingly, the damage-dependent phosphorylated Mre11 is predominantly
contained within the chromatin-bound Mre11 pool. These data suggest that phosphorylated Mre11 may have increased DNA binding affinity or that Mre11 may have to be chromatin-bound to be phosphorylated.
While studies have shown that both RPA and MRN form nuclear foci following
treatment with a variety of genotoxic agents [9, 29, 31, 32, 106-108], no one has
previously reported that these foci co-localize. Mre11 foci formation following UV-
induced damage has been reported to occur only in xeroderma pigmetosum variant
(XPV) cells, and not in ‘normal’ cells [9, 32, 117]. We observed UV-induced Mre11 foci
in both HeLa and a normal-telomerase transformed cell line (data not shown) 8 h
following UV irradiation. It has recently been reported that the large T-antigen in SV40
transformed cell lines disturbs the formation of Mre11 nuclear foci [56]. While previous
studies that reported Mre11 foci only in XPV cell lines investigated similar time points and doses of UV, their use of SV40 transformed cells may explain the discrepancies with our results.
Both RPA and the MRN complex are known to bind DNA and interact with numerous
other proteins. However, our data demonstrate that DNA binding is not required for the
RPA/MRN complex interaction. In addition, we observed that protein phosphorylation
enhanced RPA/MRN complex interaction, similar to the reported phosphorylation-
enhanced BRAC1/BACH1 interaction [57]. 43
Our data, as well as previous reports [87, 88, 118], have led to our proposed model of
events associated with stalled replication forks (Figure 8). RPA and the MRN complex
are normally associated with replication forks during S-phase [29, 68, 119]. Stalled
replication forks generate stretches of ssDNA that become coated with RPA, leading to the recruitment and activation of ATR/ATRIP [87, 88], and possibly ATM [87]. The close proximity of these proteins at stalled forks would allow for ATR/ATM phosphorylation of RPA and the MRN complex. This enhances recruitment of additional
MRN complex and phospho-dependent RPA/MRN interaction. Phosphorylation most likely has functional consequences in addition to altering protein interactions. For example, phosphorylation of Mre11 is thought to increase its nuclease activity [16, 87], which could increase DNA processing necessary for resolution of the stalled fork.
Phosphorylation of RPA is known to decrease its unwinding or destabilization activity of dsDNA but does not effect ssDNA binding ability [85, 86]. This may help prevent further migration of the replication fork and provide the opportunity for further protein recruitment. The recruited proteins, RPA/ATR/ATRIP/Mre11/Rad50/Nbs1 (and possibly
ATM), may then initiate DNA repair and act as a scaffold to activate other proteins such as Rad17, Rad9-Rad1-Hus1 or Wrn [89, 101]. Possible outcomes may include cell cycle arrest, resolution of Holliday junctions and continuation of replication, non-homologous end joining (NHEJ)-mediated repair of subsequent DSBs, or recombination-like repair of subsequent DSBs. This model suggests that RPA and the MRN complex work together at sites of stalled or collapsed replication forks, and that this cooperative interaction occurs via MRN’s ability to interact with RPA in a phosphorylation-dependent manner. 44
In this study, a single time point for each DNA-damaging agent was selected based on peak incidence of RPA-p34 hyperphosphorylation (data not shown). Future work needs to investigate additional time points to determine the exact role of RPA and MRN in the replication stress-induced damage response, and if additional proteins mediate the
RPA/MRN complex interaction. 45
CHAPTER 3. DNA Lesion-Specific Co-localization of the Mre11/Rad50/Nbs1
(MRN) Complex and Replication Protein A (RPA) to Repair Foci
I. INTRODUCTION
Both replicative stress and DNA damage initiate cellular processes collectively termed
the DNA damage response. These processes include activation of appropriate DNA repair
mechanisms, delay of the cell cycle in order to allow sufficient time for repair, and in
some cases, apoptosis [1, 2]. One hallmark of the DNA damage response is the
aggregation of multiprotein complexes into foci or repair centers. The composition of the
foci depends upon the nature of the DNA lesion and is temporally dynamic; changing as
the damage is first recognized, processed and then repaired [11, 12]. The assembly of foci
appears to be largely governed by a network of protein-protein interactions rather than
DNA-protein interactions [11, 12], providing an explanation for the dynamic composition
of the foci. Although different types of DNA damage activate specific repair pathways
and therefore specific proteins, there are some protein factors that respond to multiple
types of lesions [11, 12, 91, 93]. These global response proteins are organized in a
distinct spatiotemporal fashion depending upon the nature of the DNA lesion [11, 12].
Two protein complexes that are intimately involved in the DNA damage response to
multiple types of lesions are the MRN complex and RPA.
The Mre11/Rad50/Nbs1 (MRN) complex is best known for its role in DNA double-strand break (DSB) repair [120], but more recent evidence has expanded this role to include 46
participation in DNA replication, cell cycle checkpoints, telomere maintenance,
signaling/sensing of DNA damage and response to stalled replication forks [11, 14-19,
23, 121]. The MRN complex forms foci at sites of DNA replication, DNA damage and
DNA repair [9, 11, 29-32, 121]. Both Mre11 and Nbs1 become phosphorylated following treatment of cells with genotoxic agents [47, 60], and Mre11 is phosphorylated in a DNA replication-dependent manner [16]. The functional significance of these phosphorylation events is not entirely known, but Nbs1 phosphorylation affects cell cycle checkpoint activation [61, 67, 122], and Mre11 phosphorylation may increase its nuclease activity
[16].
Replication protein A (RPA), a three subunit protein (p70, p34 and p14; named for their
apparent molecular weights), is the major single-stranded DNA (ssDNA) binding protein
in mammalian cells (Replication Factor A or RFA in yeast). RPA is an essential protein
that plays a crucial role in DNA replication, repair and recombination [68]. RPA coats the
ssDNA found at sites of DNA replication [123-127], DNA damage and DNA repair [11,
31, 106-108, 128]. It has been postulated that this RPA-DNA intermediate is a common
signal necessary to activate the DNA damage response [24, 88]. The p34 subunit of RPA
is hyperphosphorylated in response to replicative stress and DNA damage [80, 104, 121,
129]. This hyperphosphorylation is postulated to function as a “switch” to direct RPA
activity from DNA replication to DNA repair [85, 86, 130].
We have previously shown that the MRN complex and RPA co-localize to stalled
replication forks where they interact following replicative stress induced by hydroxyurea 47
(HU) or ultraviolet light (UV) [121]. We wanted to determine if the interaction of the
MRN complex with RPA was specific to replicative stress responses, or if other types of
DNA damage also stimulate the interaction. To achieve this we used topoisomerase inhibitors and cross-linking agents. While HU leads to replication fork stalling due to depletion of deoxyribonucleotide substrates [131, 132], camptothecin (CAMPT), a topoisomerase I inhibitor [133, 134], leads to ssDNA breaks. Both HU and CAMPT have the potential to ultimately induce DNA DSBs: HU through the collapse of stalled replication forks [8, 135, 136] and CAMPT through the replication of nicked template
DNA [134, 137, 138]. Double-strand breaks also can be caused by etoposide (ETOP), a potent inhibitor of topoisomerase II [133, 139-141], and mitomycin C (MMC), a DNA cross-linking agent [142, 143], that generates DNA DSBs through the repair process of the cross-links [144-146].
Here we demonstrate that co-localization of the MRN complex and RPA in nuclear foci differs depending on the damaging agent used. HU and CAMPT treatment led to phospho-Nbs1 foci that co-localized with RPA foci. In contrast, ETOP treatment induced both phospho-Nbs1 and RPA foci, but not within the same cell. Treatment with MMC induced phospho-Nbs1 foci, but did not induce any RPA foci formation until the latest time point investigated. These localization differences suggest that although the DNA damage response is activated (evidenced by Nbs1 phosphorylation) in response to HU,
CAMPT, ETOP and MMC, the interaction of the MRN complex and RPA at the level of co-localization into repair foci is limited to responses to HU and CAMPT.
48
II. MATERIALS AND METHODS
Cell lines and treatments: HeLa cells were obtained from American Type Culture
Collection (ATCC; Manassas, VA) and maintained at 37°C and 5% CO2 in Dulbecco’s
Modified Eagles Medium (DMEM; Gibco, Gaithersburg, MD) supplemented with 10%
fetal bovine serum (FBS; Hyclone, Logan, UT) and 1% penicillin-streptomycin (Gibco).
For hydroxyurea (HU; Sigma-Aldrich, St. Louis, MO) treatment, asynchronous cells
were incubated in growth medium containing 4 mM HU for 1-24 hours before harvesting.
For camptothecin (CAMPT; Sigma-Aldrich) treatment, the drug was added directly to the
cell medium at a final concentration of 1 µM for 1-6 hours before cell harvest. For etoposide (ETOP; Sigma-Aldrich) treatment, cells were incubated in growth medium containing 25 µM ETOP for 1-6 hours before harvesting, while for mitomycin C (MMC;
Sigma-Aldrich) treatment, the drug was added to the media at a final concentration of 1
µg/ml for 1-6 hours before cell harvest.
Immunofluorescent detection of foci: Cells were grown on 12 mm coverslips (Becton
Dickinson Labware, Bedford, MA) for 48 hours prior to treatment. Cells were treated
with 4 mM HU for 1-24 hours, 1 µM CAMPT for 1-6 hours, 25 µM etoposide for 1-6 hours, or 1 µg/ml MMC for 1-6 hours. After treatment, cells were fixed with PBS containing 4% paraformaldehyde (Electron Microscopy Sciences, Hatfield, PA), permeablized with PBS containing 0.5% Triton X-100 (Sigma-Aldrich), and then blocked in PBS containing 15% FBS. After incubation in primary antibody solutions overnight at
4°C, cells were washed with PBS, incubated in secondary antibody solutions at room 49
temperature, washed with PBS and then placed on slides using gel mounting medium
(Biomedia, Foster City, CA) containing DAPI as a counter stain. Primary antibody dilutions used are as follows: anti-RPA 1:1,500 (Neomarkers) and anit-Nbs1-SP343
1:1,500 (Novus Biological). Secondary antibody dilutions are as follows: anti-rabbit
Alexa Fluor 488 1:250 and anti-mouse Alexa Fluor 596 1:250 (Molecular Probes,
Eugene, OR). The percentages of cells containing foci were determined using a Nikon inverted fluorescent microscope, and images of the foci were captured using a Zeiss
Axiovert 100M equipped with a Zeiss LSM 510 scanning confocal microscope. Images were processed using Adobe Photoshop 7.0 (Adobe, San Jose, CA). For each experiment condition, at least 100 cells were analyzed, and experiments were repeated three times.
Western immunoblots: Cell lysates were separated on 12% SDS-polyacrlyamide gels and transferred to PVDF membranes (Millipore Corp., Bedford, MA). Membranes were probed using anti-RPA-p34 (Neomarkers, Freemont, CA; 1:5,000) and horseradish peroxidase-linked anti-mouse antibodies (Amersham Biosciences, Buckinghamshire,
England; 1:3,000). Bound antibodies were visualized using chemiluminescent detection.
Comet assays: Neutral comet assays were performed as previously described [147, 148] with slight modifications. Briefly, 12,000-15,000 cells from treated cultures were suspended in low-melting point agarose in PBS (Invitrogen; final concentration of
0.75%) and layered on microscope slides. The slides were then placed in neutral lysis buffer (1 mM tris-HCl, pH 7.4; 150 mM NaCl; 4 mM EDTA; 18 mM N-laurylsarcosine) for 3 minutes and then into deionized H2O for 10 minutes. Electrophoresis was done at 20 50
V, 200 mA for 10 minutes in electrophoresis buffer (230 mM tris, 180 mM boric acid, 0.2 mM EDTA). Following electrophoresis, cells were immersed in a propidium iodide solution (PI; 2.5 µg/ml; Sigma-Aldrich) for 20 minutes, rinsed in deionized H2O, and coverslips were placed on the gels. Slides were analyzed using a Nikon inverted fluorescent microscope with attached CCD camera. Images were saved as bitmap (BTM) files and tail moments were determined using CometScore Freeware from TriTek Corp.
(Sumerduck, VA).
III. RESULTS
HU and CAMPT induce co-localization of phospho-Nbs1 and RPA foci. The MRN complex and RPA both form foci at stalled replication forks and sites of DNA damage [9,
11, 29-32, 106-108, 121]. We have previously reported that Mre11 and RPA foci co- localize following replicative stress [121], and consistent with that report we observed co-localization of phospho-Nbs1 and RPA foci following hydroxyurea (HU) treatment
(Figure 9A, panels D-F). In a similar fashion, cells treated with camptothecin (CAMPT), a topoisomerase I inhibitor [133, 134], also formed phospho-Nbs1 and RPA foci that co- localized (Figure 9A, panels G-I). To determine the extent of foci formation, we calculated the percentage of cells containing phospho-Nbs1 foci, RPA foci, or both.
While not addressing co-localization, the percent of cells expressing foci provided information on the ability of the MRN complex and RPA to form foci independently of each other. The percentage of cells that contained phospho-Nbs1, RPA, or both phospho-
Nbs1 and RPA foci showed a marked similarity between HU and CAMPT treatment. The percentage of cells containing phospho-Nbs1 foci increased from approximately 40% at 1 51
hour to 60% at 6 hours. Notably the cells that contained phospho-Nbs1 foci also
contained RPA foci, and the reverse was also true (Figure 9B). With these agents, the
phospho-Nbs1 and RPA foci co-localized, implying that the MRN complex and RPA
responded jointly at both stalled replication forks and DNA single-strand breaks (SSBs).
ETOP and MMC induce phospho-Nbs1 and RPA foci that do not co-localize. To
determine the extent of phospho-Nbs1 and RPA foci formation and to determine if these
proteins co-localize at other DNA lesions, we investigated two additional agents that
damage DNA by different mechanisms: etoposide (ETOP), a topoisomerase II inhibitor
[133, 139]; and mitomycin C (MMC), a DNA cross-linking agent [142, 143]. In contrast
to the HU- and CAMPT-induced foci, ETOP and MMC treatment did not lead to
phospho-Nbs1 and RPA foci co-localization (Figure 9A, panels L, O and R). Following
treatment with ETOP the majority of cells contained either phospho-Nbs1 or RPA foci,
but not both (Figure 9A, panels J-O; Figure 9B). After 1 hour of ETOP treatment,
approximately 60% of cells had phospho-Nbs1 foci, and about 20% had both phospho-
Nbs1 and RPA foci. At later time points an equivalent percentage of cells (about 50-55%)
demonstrated either phospho-Nbs1 or RPA predominant foci. However, the percentage
of cells that contained both phospho-Nbs1 and RPA foci dropped to 5% at 3 hours, and
by 6 hours had risen to about 20% (Figure 9B).
Treatment with MMC led to an increase in the percentage of cells containing phospho-
Nbs1 foci from about 25% after 1 hour of treatment to 95% after 6 hours of treatment
(Figure 9B). Notably RPA foci were not seen until the 6 hour time point, where only 52
about 20% of cells which contained phospho-Nbs1 foci also contained RPA foci (Figure
9B). Coupled with the ETOP data, this difference in co-localization suggested that the
response of the MRN complex and RPA was not equivalent for all types of lesions. While
the MRN complex responded to every type of lesion and at every time point investigated,
the response of RPA to MMC was very weak and only occurred at the latest time point
investigated despite the robust MRN response. The difference in response of the MRN
complex and RPA is also supported by the observation that while both the MRN complex
and RPA responded to ETOP, their responses were independent of each other, as
demonstrated by the ability of each to form foci in the absence of the other.
HU, CAMPT, ETOP and MMC induce RPA-p34 hyperphosphorylation. We have previously shown that HU treatment leads to damage-dependent phosphorylation of
Mre11 and hyperphosphorylation of RPA-p34 [121]. Formation of phospho-Nbs1 foci in the immunofluorescence experiments verified that HU, CAMPT, ETOP and MMC all induced the phosphorylation of Nbs1. In order to determine if these agents induced hyperphosphorylation of RPA-p34, we performed western immunoblotting of whole-cell lysates and looked for the characteristic hyperphosphorylated band. HU, CAMPT, ETOP and MMC all led to hyperphosphorylation of RPA-p34, with a time-dependent increase in the amount of the hyperphosphorylated form (Figure 10, lanes 2-4, 6-8, 10-12 and 16;
Form 5 represents the hyperphosphorylated band). Consistent with the foci data that showed RPA did not respond to MMC treatment until 6 hours, RPA hyperphosphorylation was only seen at the 6 hour time point and was much less robust than with the other agents (Figure 10, lane 16). 53
ETOP and MMC induce DNA double-strand breaks (DSBs). The foci data suggested that
different types of lesions generated specific responses that differed in the extent of RPA
involvement. One difference, comparing HU and CAMPT with ETOP and MMC
treatment, is the generation of DNA double-strand breaks (DSBs). ETOP is known to be
a potent inducer of DNA DSBs through the inhibition of topoisomerase II [140, 141], and
repair of MMC-induced damage progresses through a DNA DSBs intermediate [144-
146]. In contrast, HU leads to replication fork stalling due to depletion of
deoxyribonucleotide substrates [131, 132], and camptothecin induces DNA single-strand
breaks (SSBs) through inhibition of topoisomerase I [134, 138]. It must be noted
however, that both HU and CAMPT have the potential to induce DNA DSBs: HU
through the collapse of stalled replication forks [8, 135, 136], and CAMPT through the
replication of nicked template DNA [134, 137, 138]. To investigate the extent of DNA
DSBs induced by these four agents, we utilized the neutral comet assay. The neutral comet assay allows for visualization and quantification of DNA DSBs from individual cells. The fold increase in the amount of DNA DSBs above non-damaged controls was compared for the different experimental groups using tail moment measurements obtained by computer analysis of comet images using CometScore. A two-tailed student t-test was used to compare the experimental to non-damaged control samples, and a p value of less than 0.02 was considered significant. As expected, there was no difference in the amount of DNA DSBs in control cells and HU-treated cells (Figure
11A). Even though CAMPT treatment led to an apparent three-fold increase in the amount of DNA DSBs (Figure 11B), only the 1 hour time point was significantly 54
different from the non-damaged controls (1 hour, p=0.014; 3 hours, p=0.126; 6 hours, p=0.041). There was a time-dependent increase in ETOP-induced DSBs culminating in a seven-fold increase over controls (1 hour, p=0.002; 3 hours, p=0.001; 6 hours, p=0.005).
Likewise, MMC-induced DSBs culminated in a six-fold increase in DSBs over controls by 6 hours (1 hour, p=0.172; 3 hours, p=0.051; 6 hours, p=0.014) (Figure 11C and D).
These data suggested that the presence of DNA DSBs may play a role in the loss of co- localization of the MRN complex and RPA.
Prolonged HU exposure induces DNA DSBs and loss of co-localization of phospho-Nbs1
and RPA foci. Short-term HU exposure causes replication fork arrest and co-localization of phospho-Nbs1 and RPA foci. Prolonged HU exposure induces DNA DSBs through the collapse of stalled forks. We therefore utilized such prolonged HU exposure to determine if the co-localization results were dependent on the presence or extent of DSBs [8, 135,
136]. After 24 hours of continuous HU treatment, cells contained similar amounts of
DSBs as ETOP and MMC treated cells (Figure 12A). At the immunofluorescence level, all of the HU-treated cells contained both phospho-Nbs1 and RPA foci; however, the foci no longer co-localized as seen at earlier time points (Figure 12B). Loss of co-localization in the prolonged HU-treated cells, together with DNA DSBs induced by ETOP and MMC leading to a very small percentage of foci positive cells expressing both phospho-Nbs1 and RPA foci, suggests that the MRN complex and RPA respond to DNA DSBs independently of one another. 55
Figure 9. HU-, CAMPT-, ETOP- and MMC-induced foci formation. HeLa cells were
treated for 3 hours with 4 mM HU, 1µM CAMPT, 25 µM ETOP, 1 µg/ml MMC or mock-treated. Following fixation in 4% paraformaldehyde, the cytoplasmic and nucleoplasmic proteins were extracted with PBS containing 0.5% Triton X-100, the cells
incubated in primary and secondary antibodies, and visualized by confocal microscopy.
A, HU- and CAMPT-induced phospho-Nbs1 and RPA foci co-localization (panels F and
I). In contrast, ETOP-induced phospho-Nbs1 and RPA foci did not form in the same cells
(panels J-O); cells were either positive for phospho-Nbs1 foci or RPA foci, but not both.
MMC treatment induced formation of phospho-Nbs1 foci, but not RPA foci (panels P-R).
B, Quantification of the percentage of cells containing phospho-Nbs1, RPA, both
phospho-Nbs1 and RPA foci, or neither following HU, CAMPT, ETOP or MMC
treatment. Cells treated with HU and CAMPT showed an increase from about 40% of
cells with phospho-Nbs1 and RPA foci to 55-60% as exposure time increased from 1
hour to 6 hours. Treatment with ETOP led to expression of either phospho-Nbs1 or RPA
foci, but the percentage of cells with both types of foci remained much lower. Treatment
with MMC led to increasing percentage of cells with phospho-Nbs1 foci with time, and a
small percentage of cells with both phospho-Nbs1 and RPA foci at 6 hours. 56
FIGURE 9
57
Figure 10. HU-, CAMPT-, ETOP- and MMC-induced hyperphosphorylation of
RPA-p34. HeLa cells were treatment with 4 mM HU, 1 µM CAMPT, 25 µM ETOP or 1
µg/ml MMC for 1-3 hours. HU, CAMPT, ETOP and MMC led to the hyperphosphorylation of RPA-p34 (lanes 2-4, 6-8, 10-12, and 16). RPA-p34 hyperphosphorylation was determined by the appearance of a slower migrating band
(form 5) on the western blot. 58
FIGURE 10
59
Figure 11. ETOP- and MMC-induced generation of DNA DSBs. HeLa cells treated
with HU, CAMPT, ETOP, MMC or mock-treated were analyzed for the presence of
DNA DSBs using the neutral comet assay. Results were presented as fold increase of
DNA DSBs above non-damaged controls. A, 4 mM HU treatment did not induce DNA
DSBs above control levels at the time points presented. B, 1 µM CAMPT induced an
apparent three-fold increase in DSBs, but overall was not statistically different from non-
damaged controls. C, 25 µM ETOP treatment induced a time dependent increase in DSBs
with an approximately seven-fold increase in DNA DSBs at 6 hours. D, 1 µg/ml MMC treatment led to a time-dependent increase in the amount of DNA DSBs culminating at a six-fold increase in DSBs.
60
FIGURE 11
61
Figure 12. Prolonged exposure to HU induced DNA DSBs and loss of phospho-Nbs1 and RPA foci co-localization. A, A time course of prolonged HU treatment showed that at 24 hours continuous exposure to 4 mM HU, cells had DNA DSBs equivalent in amount to cells treated with ETOP or MMC. B, At 24 hours of continuous 4 mM HU treatment, phospho-Nbs1 and RPA foci were visualized by confocal microscopy. Cells contained both phospho-Nbs1 and RPA foci, but these foci no longer co-localized. 62
FIGURE 12
63
Figure 13. Model of the interaction of the MRN complex and RPA in response to different DNA lesions (see “Discussion” for details). 64
FIGURE 13
65
IV. DISCUSSION
Stalled replication forks induced by HU treatment and DNA single-strand breaks (SSBs)
induced by CAMPT both activate the DNA damage response in a similar manner with the
phosphorylation of Nbs1, hyperphosphorylation of RPA-p34, and the formation of foci
containing both the MRN complex and RPA. DNA double-strand breaks (DSBs) induced
by ETOP or MMC treatment also activated the DNA damage response; however, the
involvement of the MRN complex and RPA was distinctly different from that seen with
HU and CAMPT. In contrast to the HU- and CAMPT-induced response, ETOP-induced
phosphorylation of Nbs1, hyperphosphorylation of RPA and foci that contained either
phospho-Nbs1 or RPA, but not both. In further contrast, MMC treatment induced
phosphorylation of Nbs1 and formation of phospho-Nbs1 foci, but not
hyperphosphorylation and foci formation of RPA. This suggested that the activation and localization of the MRN complex and RPA was dependent on the type of lesion. Lesions
that contained stalled replication forks or SSBs induced activation and co-localization of
the MRN complex and RPA, but lesions that contained crosslinked DNA or DSBs
induced a different RPA response.
The response to DNA DSB leads to activation of one of two different pathways to repair the damaged DNA; non-homologous end joining or homologous recombination. The
Rad52 epistasis group of proteins plays important and critical roles in homologous recombination. It has been shown previously that both Rad51 (member of the Rad52 epistasis group) and Mre11 form foci following ionizing radiation (IR) treatment, but that
Rad51 and Mre11 foci were not found in the same cell [9]. Maser et al. suggested that 66
these data support the idea that the MRN complex and Rad51 have independent roles in
DSB repair [9]. More recently, Lisby et al. showed that the protein composition of foci changed dynamically at the site of a single endonuclease-induced DSB. MRX, the yeast
MRN complex, formed foci very early at the site of the break, but MRX had disappeared from the foci by the time Rad51 and Rad52 were recruited to the same site [11]. In that study, they also demonstrated that RPA was recruited to the site of the break before the departure of MRX, and that formation of Rad51 and Rad52 foci at the site of damage was dependent on the presence of RPA. With these facts in mind, we investigated the role of
DNA DSBs in the observed differences in phospho-Nbs1 and RPA foci co-localization.
Treatment with either ETOP or MMC led to DNA DSBs and to phospho-Nbs1 foci in the absence of RPA foci. To determine if previously co-localized phospho-Nbs1 and RPA foci at stalled forks continued to co-localize following collapse of the stalled fork and generation of a DSB, prolonged HU treatment of cells was employed. After 24 hours of continuous exposure to HU, cells contained amounts of DNA DSBs comparable to those induced by ETOP and MMC treatment. At this later time point, phospho-Nbs1 and RPA foci were both still present in the nucleus, but they no longer co-localized. This suggested that the transition from stalled replication fork to DNA DSB resulted in the loss of co- localization of the MRN complex and RPA. This loss of co-localization may represent independent functions of the MRN complex and RPA in the repair of DSBs, or may represent a “snapshot” of the temporally dynamic composition of foci as reported by
Lisby et al [11].
67
The type of DNA damage may play a critical role in determining whether RPA is bound to the damaged DNA, which in turn affects the ability of RPA to become hyperphosphorylated [75, 149]. The binding of RPA to ssDNA has been well characterized. A 30 nucleotide long stretch of ssDNA is required for usual RPA binding
[70, 150], but under certain circumstances RPA can bind with only 8-10 nucleotides [75,
151]. The 8-nucleotide binding mode is likely a precursor to the more stable 30- nucleotide binding [75], and is consistent with the observed lower affinity of RPA for shorter oligonucleotides in vitro [152]. In our assays, the lack of RPA foci only indicates that the RPA content in the foci is below the level of resolution of this assay, and cannot be taken to mean that RPA is completely absent. Therefore the differences in RPA binding are likely due to relative amounts. With HU treatment, large sections of ssDNA are generated at the sites of stalled forks [131], providing a substrate for RPA binding.
With CAMPT treatment, the inhibition of topoisomerase I allowed for continued replication, and the generation of additional ssDNA. Inhibition of topoisomerase II by
ETOP leads to the production of DNA DSBs, which may not have significant amounts of ssDNA at the broken ends initially. The MRN complex is probably recruited to these sites to tether the broken ends together [44, 153], and then help recruit other necessary protein factors, including a nuclease that processes the ends to generate ssDNA that is then bound by RPA [11]. Interstrand crosslinking may prevent the accumulation of significant ssDNA because the interstrand tethering would inhibit DNA unwinding, and thus prevent the formation of a ssDNA platform to which RPA could bind. The mitomycin C experiments presented here, which demonstrated a lack of RPA foci formation and RPA hyperphosphorylation, are consistent with such a model. This lack of 68
a response by RPA also occurs with cisplatinum, another agent that causes interstrand
crosslinks (John Turchi, personal communication), indicating that there is something
about the interstrand crosslink that either inhibits RPA binding and activation, or is not
recognized by a signaling and repair pathway that includes RPA.
These results, combined with observations made by others, are compatible with a model
of DNA repair that considers the availability of ssDNA for RPA binding and subsequent
hyperphosphorylation (Figure 13) to explain the foci data presented here. This model
depicts the MRN complex as a sensor for damage and bestows a protector, sensor, and
effecter function on RPA. RPA protects ssDNA and is hyperphosphorylated to identify
replication stress, SSBs and some types of DSBs. The replication stress foci, if
unrepaired, lead to replication fork collapse and subsequent DNA DSBs and formation of
early DNA repair foci (similar to Mirzoeva and Petrini’s Type II foci [30]). These type II
foci contain MRN and can also contain RPA depending on the type of DNA damage and
amount of ssDNA. Subsequently, the foci evolve into type III DNA repair foci that
represent repair of DSBs either through a recombinatorial pathway using RAD51/RAD52 or non-homologous end joining. These processes of DSB repair may occur within the same cell, or different cells, thus leading to differences in protein composition of the foci.
We have demonstrated that the MRN complex responds to multiple types of lesions and
that RPA is involved in the response to a sub-set of these lesions, probably those that
contain significant stretches of ssDNA. Additionally, these data demonstrate that the
MRN complex and RPA can act independently of each other, and probably have different 69
roles in the repair of DSBs that are separated physically and/or temporally. These data
however only include a small proportion of time points and agents that could be
investigated to further elucidate the nature of the MRN complex and RPA interaction and the importance of that interaction on repair of specific types of lesions. Ongoing work to quantitate the amount of ssDNA present in the different lesions will help determine if this is the explanation for the presence or absence of RPA foci and the unexpected absence of
RPA in the response to interstrand crosslinking agents. 70
CHAPTER 4: Loss of Replication Protein A (RPA) Prevents Phospho-Nbs1 Foci
Formation Following Etoposide-Induced DNA Damage
I. INTRODUCTION
In response to DNA damage and replicative stress, cells initiate processes, collectively
termed the DNA damage response, that culminate in the activation of repair proteins and
cell cycle checkpoints. Incorrect initiation and/or function of the DNA damage response
increases the frequency of mutations and leads to genomic instability, a known precursor
of cancer [91, 154-156]. Many of the proteins activated in the DNA damage response
localize to the sites of damage to form large protein aggregations called nuclear foci. The
function of these foci is not entirely known, but may be important in controlling damage-
induced checkpoints, facilitating repair of persistent damage, compartmentalizing
activated proteins, or concentrating repair proteins that may require a threshold level for
certain biochemical steps in the repair process [1, 9, 10, 12]. The foci composition depends on the nature of the lesion, and changes over time as the DNA damage response progresses from damage recognition and signaling to repair and resolution [12]. In
addition, the assembly of foci is largely governed by protein-protein interactions rather
than DNA-protein interactions [11, 12]. Although different types of DNA damage
activate specific repair pathways, there are some protein factors that respond to multiple
types of lesions [11, 12, 91, 93]. Two such broad response protein complexes are the
Mre11/Rad50/Nbs1 (MRN) complex and replication protein A (RPA).
71
The MRN complex (composed of Mre11, Rad50 and Nbs1) is best known for its role in
DNA double-strand break (DSB) repair, but also responds to DNA replicative stress,
DNA single-strand breaks (SSBs) and interstrand cross-links [157] [47]. As part of the
response to these different types of damage, the MRN complex functions to activate the
cellular kinases ATM and ATR, initiate cell cycle checkpoints, and help maintain
telomeres [18-22, 24-28]. The MRN complex is proposed to be one of the damage
sensors that, upon sensing DNA damage, activate the appropriate processes [23]. We
have shown that the MRN complex is able to form foci at sites of damage induced by a wide variety of agents, and that the formation of these foci can be visualized in the absence of RPA foci [157].
Replication protein A (RPA) is the major single-stranded DNA (ssDNA) binding protein in eukaryotic cells, and is essential for DNA replication, repair and recombination [68,
130]. RPA is composed of three subunits with different molecular weights: 70 kDa (p70),
34 kDa (p34), and 14 kDa (p14). The p34 subunit of RPA can be phosphorylated in a cell cycle-dependent manner and in response to DNA damage [130]. While the functional consequences of this phosphorylation are not clear, it is known that phosphorylation causes a conformational change in RPA via intersubunit interactions, which may then regulate RPA-protein interactions and DNA binding [85]. Additionally, RPA-coated ssDNA is a common structure generated at sites of DNA damage that plays an important role in the recruitment of other DNA repair proteins such as ATR-ATRIP [88], Rad17
[89] and Cut5 [90].
72
Both the MRN complex and RPA are hypothesized to act as sensors of DNA damage as
well as to have additional functional roles in the DNA damage response. Because the
MRN complex is reported to be required for activation of ATM and ATR, the two kinases believed to phosphorylate RPA following DNA damage [130], and because the
MRN complex is recruited to sites of DNA DSBs prior to RPA recruitment [11], we wanted to investigate the ability of RPA to form foci and to become phosphorylated when cells were depleted of Mre11. We also wanted to determine if the ability of the MRN complex to form foci and become phosphorylated in response to DNA damage was altered under conditions of RPA depletion. Using a siRNA approach for targeted protein depletion, we demonstrate that loss of Mre11 did not alter the ability of RPA-p34 to become hyper-phosphorylated or form foci following etoposide (ETOP) treatment.
Depletion of RPA-p70 did not affect the ability of the remaining RPA-p34 to become
hyper-phosphorylated, but abrogated the ability of RPA-p34 and the MRN complex to
form repair foci following ETOP treatment, suggesting that MRN foci formation may be
dependent on the presence of RPA.
II. MATERIALS AND METHODS
Cell lines and treatments: HeLa cells were obtained from American Type Culture
Collection (ATCC; Manassas, VA) and maintained at 37°C and 5% CO2 in Dulbecco’s
Modified Eagles Medium (DMEM; Gibco, Gaithersburg, MD) supplemented with 10%
fetal bovine serum (FBS; Hyclone, Logan, UT) and 1% penicillin-streptomycin (Gibco).
For etoposide (ETOP; Sigma-Aldrich) treatment, cells were incubated in growth medium
containing 10-50 µM ETOP for 1-3 hours before harvesting. 73
Immunofluorescent detection of foci: Cells were grown on 12 mm coverslips (Becton
Dickinson Labware, Bedford, MA) for 48 hours prior to treatment with siRNAs. After 45
hours of siRNA treatment, cells were exposed to 25 µM etoposide for 3 hours and then
subjected to immunofluorescent staining for foci as previously described [157]. Briefly,
cells were fixed with phosphate-buffered saline (PBS) containing 4% paraformaldehyde
(Electron Microscopy Sciences, Hatfield, PA), permeablized with PBS containing 0.5%
Triton X-100 (Sigma-Aldrich), and then blocked in PBS containing 15% FBS. After
incubation in primary antibody solutions overnight at 4°C, cells were washed with PBS,
incubated in secondary antibody solutions at room temperature, washed with PBS and
then placed on slides using gel mounting medium (Biomedia, Foster City, CA). Primary
antibody dilutions used are as follows: anti-RPA 1:1,500 (Neomarkers, Freemont, CA)
and anit-Nbs1-SP343 1:1,500 (Novus Biological, Littleton, CO). Secondary antibody
dilutions are as follows: Alexa Fluor 488 1:250 and Alexa Fluor 594 1:250 (Molecular
Probes, Eugene, OR). Foci images were captured using a Zeiss Axiovert 100M equipped
with a Zeiss LSM 510 scanning confocal microscope. Images were processed using
Adobe Photoshop 7.0 (Adobe, San Jose, CA).
Western immunoblots: Cell lysates were separated on 12% SDS-polyacrlyamide gels and
transferred to PVDF membranes (Millipore Corp., Bedford, MA). Membranes were
probed using anti-RPA-p34 (Neomarkers; 1:5,000), anti-RPA-p34-SP4-SP8 (Bethyl
Laboratories; 1:20,000), anti-Mre11 (Novus; 1:20,000), anti-Nbs1-SP343 (Novus;
1:5000), anti-glyceraldehyde-3-phosphate dehydrogenase (G3PDH; Trevigen, 74
Gaithersburg, MD; 1:10,000) and horseradish peroxidase-linked anti-mouse or anti-rabbit
antibodies (Amersham Biosciences, Buckinghamshire, England; 1:3,000). Bound
antibodies were visualized using chemiluminescent detection. Quantitation of protein
amounts was done using ImageQuant software (version 5.1; Molecular Dynamics,
Piscataway, NJ) and G3PDH as a loading control.
siRNA treatment: HeLa cells were seeded in 24 well plates at a density of 1 x 104 cells
per well or 6 well plates at a density of 2.5 x 104 cells per well in DMEM medium
containing 10% FBS. Cells were allowed to attach for 24 hours prior to transfection with
siRNA. Transfection-ready siRNA duplexes were purchased from Dharmacon Research
using previously published sequences. Two different sequences directed at RPA-p70
were used, termed RPA#1 [87], and RPA#2 [158]. An equal mixture of three different
sequences was used for Mre11 [159]. Control (non-silencing) siRNA was purchased from
Qiagen (Valencia, CA). Working concentrations of 20 µM were used for each siRNA, with cells exposed to a final concentration of 333 nM for RPA#1 and RPA #2, and 500 nM for Mre11. The siRNAs were transfected into cells using Lipofectamine 2000 reagent
(Invitrogen) following the manufacture’s protocol. Briefly, siRNA was suspended in
DMEM without serum or antibiotics, gently mixed with lipofectamine (13 µg/mL final concentration exposed to cells) and incubated at room temperature for 20 minutes. Just prior to the addition of the siRNA/lipofectamine to the cells, the medium in the wells was replaced with DMEM containing 12% FBS and no antibiotics (10% FBS final concentration after addition of siRNA solution). The cells were then incubated in a humidified 37°C incubator with 5% CO2. At 5 hours and 24 hours after the addition of 75
the siRNA, the medium in each well was replaced with fresh DMEM with 10% FBS and
1% penicillin-streptomycin. After 45 hours total incubation, the ETOP-treated cells were exposed to ETOP for 3 hours, and then harvested for foci staining and/or western
immunoblotting.
III. RESULTS
ETOP treatment induces phosphorylation of Nbs1 and hyper-phosphorylation of RPA-
p34. We have previously reported that ETOP treatment leads to the formation of
phosphorylated Nbs1 foci and hyper-phosphorylation of RPA-p34 [157]. Here we present
a time and dose response with 10-50 µM ETOP for 1-3 hours. The level of phosphorylated Nbs1 was essentially the same for all times and doses investigated
(Figure 14, lanes 2-10). For RPA-p34 hyper-phosphorylation, there was a time- and dose- dependent increase in the relative amount of hyper-phosphorylated RPA (form 5)
compared to the non-phosphorylated isoform (form 1; Figure 14, lanes 2-10). The robust
phosphorylation of Nbs1 and RPA following ETOP treatment, along with the previously
reported observation that ETOP-induced phosphorylation of RPA is independent of DNA
replication [129], led us to use 25 µM ETOP treatment for 3 hours (Figure 14, lane 6) to
activate the DNA damage response in siRNA-treated cells.
siRNA directed at RPA-p70 decreases RPA protein levels. Treatment of cells with one of
two different siRNA sequences directed at RPA-p70 (siRPA#1 and siRPA#2) decreased
RPA-p70 protein levels to about 10% of cells treated with a non-silencing control siRNA
(siControl) (Figure 15A and 15C). Levels of RPA-p34 decreased as well, but not to the 76
same extent, decreasing 12-32% of siControl (Figure 15A and 15C). This is similar to a
previous report using the same sequence as siRPA#2 [158]. Since the siRPA#2 led to a greater decrease in protein levels, siRPA#2 was used for all future experiments (hereafter referred to as siRPA).
siRNA directed at Mre11 decreases Mre11 protein levels but does not affect RPA levels.
Treatment of cells with an equal mixture of three different siRNA sequences all directed
at Mre11 (siMre11) led to a decrease in Mre11 protein levels to about 15% of siControl-
treated cells (Figure 15B and 15C). The RPA protein levels were not affected by siMre11
treatment as demonstrated by normal levels of RPA-p34 (Figure 15B and 15C). Previous
reports using the same mixture of siMre11 saw no effect on Nbs1 and Rad50 protein
levels [159, 160].
siRPA and siMre11 do not affect ETOP-induced hyper-phosphorylation of RPA-p34.
Cells pre-treated with either siRPA or siMre11 were treated with 25 µM ETOP for 3
hours, and the phosphorylation status of RPA-p34 was monitored. ETOP induced RPA-
p34 hyper-phosphorylation to a similar extent in cells treated with siControl, siRPA or
siMre11 (Figure 16). While this suggested that RPA-p34 hyper-phosphorylation was
independent of Mre11, the possibility remains that within individual cells the level of
Mre11 protein may vary greatly, with some cells having complete depletion of Mre11
protein, and others having normal levels. RPA-p34 shown in Figure 16 is pooled from a
whole cell population, which may contain cells with different Mre11 status. To try and 77
determine if RPA-p34 response to ETOP is truly independent of Mre11, we wanted to look within individual cells using immunofluorescence.
Depletion of Mre11 does not affect the ability of cells to form RPA-p34 foci. Cells treated with siMre11 were assessed for their ability to form RPA-p34 and phospho-Nbs1 foci following ETOP treatment using immunofluorescence. Depletion of Mre11 led to an abrogation of phospho-Nbs1 foci formation, which was expected due to a previous report that Nbs1 phosphorylation is dependent on the presence of Mre11 [159]. However, siMre11 treatment did not decrease the formation of RPA foci, but rather increased the percentage of cells with RPA foci from about 35% in siControl cells to approximately
55% (Figure 17). These data were consistent with the RPA-p34 hyper-phosphorylation data in Figure 16 showing that RPA response to ETOP treatment was independent of
Mre11. Although these data suggested that the MRN complex and RPA functioned independently following ETOP treatment, we needed to address the question if the MRN response was dependent on RPA.
Depletion of RPA abrogates phospho-Nbs1 foci formation following ETOP treatment. As expected, the depletion of RPA led to a decrease in the percentage of cells that contained
RPA foci. Interestingly, siRPA treatment also decreased the ETOP-induced phospho-
Nbs1 foci formation to levels equivalent to negative controls (Figure 17). While this suggests that the MRN complex foci formation may be dependent upon RPA, we cannot rule out the possibility that the loss of phospho-Nbs1 foci is due to loss of phosphorylation of Nbs1 and not an inhibition of foci formation. 78
Figure 14. Time- and dose-response of ETOP-induced Nbs1 phosphorylation and
RPA hyper-phosphorylation. Cell lysates from HeLa cells treated with 10-50 µM
ETOP for 1-3 hours were subjected to western immunoblotting using anti-phospho-Nbs1
and anti-RPA-p34 antibodies. Nbs1 was phosphorylated at every time and dose investigated. RPA-p34 hyper-phosphorylation (form 5) increased in a time- and dose- dependent manner. 79
FIGURE 14
80
Figure 15. Depletion of RPA and Mre11 proteins using targeted siRNAs. A, Western immunoblot demonstrating two different siRNA sequences directed at RPA-p70 led to dramatically decreased protein levels of RPA-p70 and a modest reduction in RPA-p34. B,
Western immunoblot demonstrating a decrease in Mre11 protein levels following treatment with siRNA directed at Mre11. RPA protein levels were not affected by this treatment. C, Quantitation of protein levels in the western immunoblots depicted in A and
B. The number for each protein is the percent of that protein relative to the protein level in the siControl treated cells (all relative to the loading control G3PDH). Treatment with both siRPA sequences led to a reduction of RPA-p70 to about 10% of siControl, and treatment with siMre11 decreased Mre11 protein levels to about 15% of siControl levels. 81
FIGURE 15
82
Figure 16. Treatment with siRPA or siMre11 does not affect ETOP-induced hyper-
phosphorylation of RPA-p34. The presence of RPA-p34 hyper-phosphorylation was
assessed following ETOP treatment in HeLa cells pre-treated with siControl, siRPA or
siMre11. Hyper-phosphorylation was monitored by using the damage-dependent
phospho-specific antibody anti-RPA-p34-SP4-SP8, and G3PDH was used as loading control. The proportion of RPA that became hyper-phosphorylated following ETOP
treatment was not altered by pre-treatment with siRPA or siMre11.
83
FIGURE 16
84
Figure 17. Depletion of RPA abrogates the ability of cells to form phospho-Nbs1 and
RPA-p34 foci following ETOP treatment. HeLa cells treated with siControl, siRPA, or siMre11 were either mock-treated (control) or treated with 25 µM ETOP for 3 hours. A.
Confocal microscopy images showing ETOP-induced phospho-Nbs1 and RPA foci in siControl treated cells, an absence of phospho-Nbs1 and RPA foci in siRPA treated cells, and only RPA foci in siMre11 treated cells. B. Quantification of the percentage of cells expressing phospho-Nbs1, RPA, both or neither types of foci following siRNA and
ETOP treatment.
85
FIGURE 17
86
IV. DISCUSSION
DNA damage induced by agents such as etoposide (ETOP) activates the DNA damage response, resulting in phosphorylation of key effector proteins and formation of repair foci. While it is not known what proteins are responsible for initially sensing the presence of the damage and activating the response, it has been hypothesized that the MRN complex and RPA may play such a role [23, 24]. Here we have shown that following induction of DNA damage by ETOP, the MRN complex and RPA both become phosphorylated and form repair foci. However, cells contain either phospho-Nbs1 foci or
RPA foci, but not both. Additionally, in response to mitomycin C-induced damage, phospho-Nbs1 foci formed but RPA foci did not [157]. These observations suggested that the MRN complex and RPA may be able to respond to damage independently of each other. In order to further investigate the dependency or independency of response of these two protein complexes to DNA damage, we have used siRNA-mediated depletion of
RPA-p70 and Mre11. Following depletion of these proteins and treatment with ETOP, we then assessed the ability of cells to phosphorylate RPA-p34 and to form repair foci.
Depletion of RPA-p70 and Mre11 did not alter the ability of ETOP to induce hyper- phosphorylation of RPA-p34 as determined by western immunoblotting. These results may be due to the nature of the assay. The cellular lysates used in the western immunoblots represent total cellular protein from a population of cells, and may mask what is occurring within an individual cell. In order to try and assess the affect of siRNA treatment at an individual cell level, we also utilized immunofluorescence.
87
Using immunofluorescence, we saw that depletion of RPA-p70 led to loss of both RPA
and phospho-Nbs1 foci. In contrast, depletion of Mre11 led to loss of phospho-Nbs1 foci, but the percentage of cells containing RPA foci increased. These data suggest that RPA foci formation is independent of MRN complex, but that phospho-Nbs1 foci formation is dependent on the presence of RPA. There are multiple possible explanations for the lack of phospho-Nbs1 foci following siRPA treatment. The first is that the MRN complex recognizes ssDNA coated by RPA and binds to RPA instead of binding to the damaged
DNA. Loss of RPA would then prevent the formation of MRN complex foci. Another alternative is that RPA is necessary for the activation of the kinase(s) that phosphorylates
Nbs1. Lack of phospho-Nbs1 foci does not rule out the possibility that non-
phosphorylated Nbs1 and/or other members of the MRN complex are able to form foci.
The data presented here does not fully answer the question if the response to ETOP-
induced DNA damage by the MRN complex and RPA are dependent upon each other,
though they do suggest that the MRN response is dependent on RPA and that RPA is
independent of the MRN complex. In order to more fully understand this relationship,
and to be able to generate a model to describe these events, further experiments are
necessary. Potentially beneficial experiments include: (1) Cell cycle differences in the
percent of cells that contain RPA or phospho-Nbs1 foci following treatment with ETOP.
(2) The ability of Nbs1 and phospho-RPA-p34 to form foci following siRNA and ETOP
treatment. (3) Comet assay to determine if siRNA treatment causes DNA damage or if
siRNA treatment alters the ability of ETOP to induced DNA double-strand breaks
(DSBs). (4) Immunofluorescence to visualize γH2AX foci, a known marker of DNA 88
DSBs. (5) Auto-phosphorylation of ATM on serine residue 1981 as a marker of
activation of the DNA damage response. These proposed assays, the rationale for doing
them and the contribution they may make to this study are discussed below.
(1) Cell cycle differences in the percent of cells that contain RPA or phospho-Nbs1 foci
following treatment with ETOP. We have previously shown that ETOP treatment induces
about 50-55% of cells to express phospho-Nbs1 foci, 40-45% to express RPA foci and
5% to express both phospho-Nbs1 and RPA foci. We have hypothesized that this
exclusive formation of phospho-Nbs1 or RPA foci may be due to different modes of
DNA DSB repair, either non-homologous end joining (NHEJ; which may represent the
phospho-Nbs1 foci) or homologous recombination (HR; which may represent the RPA
foci). If this is the case, it is known that HR occurs in S-phase and G2/M phase of the cell
cycle, due to the availability of a homologous chromosome, while NHEJ occurs
predominantly in G1 and early S-phase of the cell cycle. Treatment with siRPA and
siMre11 slows down the progression of cells through the cell cycle, and probably results
in cells accumulating in G1 phase. If the ability of phospho-Nbs1 or RPA to form foci
following ETOP treatment is dependent on the stage of the cell cycle, the foci data may
simply reflect the cell cycle-dependent formation of foci rather than a dependency of
MRN complex and RPA on each other.
(2) The ability of Nbs1 and phospho-RPA-p34 to form foci following siRNA and ETOP
treatment. Loss of Mre11 and/or RPA may decrease or prevent the activation of kinases involved in the DNA damage response such as ATM and ATR. If ATM and/or ATR are 89
not activated, the integrity of the DNA damage response is compromised and the damage
is not repaired appropriately. siRPA treatment blocked formation of phospho-Nbs1 foci,
but it is not known if siRPA blocked phosphorylation of Nbs1, or if it blocked MRN foci
formation. Additionally, siMre11 did not block RPA foci formation, but it may prevent
the formation of phospho-RPA foci.
(3) Comet assay to determine if siRNA treatment causes DNA damage or if siRNA treatment alters the ability of ETOP to induced DNA double-strand breaks (DSBs).
Depletion of Mre11 or RPA may lead to a situation where the cell is more sensitive to damaging agents and not able to repair the damage that does occur. Conversely, siRNA- treated cells may not progress through the cell cycle at the same rate and therefore any replication-dependent processes may be substantially decreased. With these ideas in mind, it is important to know the extent of DNA damage induced by siRNA treatment alone and siRNA treatment in conjunction with ETOP treatment.
(4) Immunofluorescence to visualize γH2AX foci, a known marker of DNA DSBs. A
commonly used marker for DSBs is γH2AX foci. The formation of γH2AX foci is
dependent upon protein phosphorylation, and is therefore similar to phospho-Nbs1 foci
formation. To help determine if the lack of phospho-Nbs1 foci following siRNA and
ETOP treatment is due to a lack of foci formation in general, inhibition of kinase activity,
or a specific inhibition of MRN complex foci formation, the ability of cells to form
γH2AX foci can be used as a positive control.
90
(5) Auto-phosphorylation of ATM on serine residue 1981 as a marker of activation of the
DNA damage response. Prior to activation of the DNA damage response, ATM is in an inactive homo-dimer complex. The dissociation of ATM to active monomers following activation is concurrent with auto-phosphorylation of ATM on serine 1981 and can be monitored using an antibody specific for phosphorylated serine 1981 [161]. It is possible that γH2AX will not be seen following siRPA and siMre11 treatment because ATM cannot become activated in the absence of RPA and/or Mre11. Determining the activation of ATM following siRNA and ETOP treatment will help determine if the lack of phospho-Nbs1 foci, and possible lack of γH2AX foci, are due to inability of cells to activate the necessary kinases and not necessarily due to the inability to form foci at sites of damage. Additionally, monitoring the auto-phosphorylation of ATM will help determine if RPA and/or the MRN complex are necessary for ATM activation following
ETOP treatment.
The combined results of the data presented and from the proposed experiments would provide a more complete picture of what is occurring within the cells following depletion of Mre11 and/or RPA. At this point, the data suggest that there is a dependency of the
MRN complex on RPA for foci formation at sites of DNA damage. This may seem to be contradictory to our earlier work that showed independent phospho-Nbs1 and RPA foci formation. This may be a limitation of the immunofluorescent foci assay rather than an independent response of these proteins. While RPA foci are not visible following mitomycin C treatment, this does not exclude the possibility that RPA is present at those foci at a level below the threshold for immunofluorescent detection. 91
A possible model incorporating the data presented here proposes that RPA coats the ssDNA ends at sites of DSBs, and that the MRN complex recognizes and binds this RPA- ssDNA complex. Subsequently, MRN activates ATM and ATR, resulting in the phosphorylation of downstream targets such as the MRN complex and RPA. This phosphorylation may then switch the MRN complex and RPA functions from detection to effector mode. At that point, the nature of the lesion and the stage of the cell cycle may determine what particular pathway(s) of repair is activated and what proteins are recruited to the foci in high enough concentration to be viewed by immunofluorescence.
If the MRN complex and RPA have effector functions in different repair processes, they may then form visually detectable foci independently of each other.
92
CHAPTER 5. Recombinant Proteins and Future Directions
I. INTRODUCTION
Proteins involved in the DNA damage response are often classified into three broad categories: sensors/initiators, signal transducers, and effectors. It is hypothesized that the
MRN complex and RPA have roles as both sensors/initiators and effectors. Their roles as sensors is supported by data that indicates the MRN complex and RPA are necessary for correct activation of the ATM and ATR kinases [20, 24-26, 28, 88, 89, 162], the two major signal transducers in the DNA damage response. Both the MRN complex and RPA are also phosphorylated in the damage response, suggesting they have effector functions downstream of the ATM/ATR kinases. We were interested in using purified recombinant proteins for in vitro experiment as a method to help delineate MRN complex/RPA protein-protein interactions as well as the functional consequences of this interaction. In order to proceed in this direction, we have begun to generate purified recombinant ATM,
ATR, MRN complex, and RPA. Acquisition of these purified proteins will allow for future exploration into the interaction of the MRN complex and RPA, the role of these proteins in activation of ATM/ATR, and the functional consequence of MRN complex and/or RPA phosphorylation by ATM/ATR.
II. ATM and ATR
A. RATIONALE
The ATM and ATR protein kinases are considered to be the major signal transducers
within the DNA damage response [4]. ATM and ATR are both members of the 93
phosphatidylinositol 3-kinase-like kinase (PIKK) family, along with DNA-dependent protein kinase (DNA-PK) and ATX/hSMG-1 [4]. ATM responds primarily to DNA
DSBs, and ATR responds to DNA DSBs, UV-induced damage and stalled replication forks [4], though there is considerable overlap and redundancy between the activation and targets of ATM and ATR. Recent reports have indicated that the MRN complex is necessary for full activation of ATM and ATR [20, 24, 26, 27], and that RPA is necessary for the activation of ATR [88]. In addition to activating these kinases, the MRN complex
(specifically Mre11 and Nbs1) and RPA are downstream targets of ATM and ATR [41,
51, 61, 163]. Generation of active recombinant ATM and ATR will facilitate the study of the in vitro activation of these kinases by the MRN complex and RPA, the ability of
ATM and/or ATR to phosphorylate the MRN complex and RPA, as well as possible functional consequences of MRN and RPA phosphorylation.
B. MATERIALS AND METHODS
Cell line: 293T cells (human embryonic kidney cells) were obtained from Michael
Kastan (St. Jude Children’s Research Hospital, Memphis, TN) and maintained at 37°C and 5% CO2 in Dulbecco’s Modified Eagles Medium (DMEM; Gibco, Gaithersburg,
MD) supplemented with 10% fetal bovine serum (FBS; Hyclone, Logan, UT) and 1%
penicillin-streptomycin (Gibco).
Preparation of ATM expressing extracts: Full-length, FLAG-tagged ATM cDNA cloned
into a CMV eukaryotic expression vector was obtained from Michael Kastan. The cDNA
was amplified by electroporating into E. coli SURE cells (Stratagene, La Jolla, CA) and 94
incubating for 24-48 hours in LB-AMP medium. The bacterial cells were collected and the cDNA extracted using QIAGEN plasmid purification kits according to manufacture’s
guidelines (QIAGEN Plasmid Mega and Giga kits; QIAGEN, Valencia, CA). The ATM
cDNA was transfected into 293T cells using calcium phosphate. 48 hours after transfection, the 293T cells were collected and cell lysates prepared using buffer A (50 mM Tris-HCl, pH 7.5; 150 mM NaCl; 10 mM β-glycerophosphate; 10% glycerol; 1%
Tween-20; 0.1% NP-40; 1 mM NaVO4; 1 mM NaF). Lysates were sonicated and cleared
by centrifugation at 25,000 x g for 30 minutes, and stored at -70°C until purification on
an anti-FLAG affinity gel column (Sigma-Aldrich, St. Louis, MO).
Preparation of ATR expressing extracts: A full-length, FLAG-tagged ATR cDNA plasmid construct was obtained from Karlene Cimprich (Stanford University School of
Medicine, Stanford, CA). ATR cDNA was amplified, collected, transfected into 293T cells, and cellular extracts prepared in an identical manner as described for ATM.
Anti-FLAG affinity column: FLAG-tagged ATM and FLAG-tagged ATR were purified from recombinant protein-expressing cell extracts following the protocol of Rhodes et al. with slight modifications [164]. Briefly, cell extracts were loaded onto an anti-FLAG M2 affinity gel column (Sigma-Aldrich) and washed with 15-20 column volumes of buffer A.
The proteins were eluted from the column using 15 ml of FLAG peptide solution (100
µg/ml in buffer A; Sigma-Aldrich). The fractions collected from the column were tested for the presence of ATM/ATR by western immunoblots and for purity by coomassie stained gels. Fractions containing the protein of interest were pooled together and 95
dialyzed into buffer B (10 mM Hepes/NaOH, pH 7.5; 50 mM NaCl; 10 mM β-
glycerophosphate; 10% glycerol; 1 mM NaVO4; 1 mM NaF; 1 mM DTT) and frozen at -
70°C.
Kinase reaction: Kinase reactions using recombinant ATM, recombinant ATR or commercially available DNA-PK were preformed following the protocol of Sarkaria et
al. [165] with slight modifications. Recombinant ATR was obtained as purified fractions
from the anti-FLAG column, and recombinant ATM was obtained by
immunoprecipitating from FLAG-ATM-expressing 293T cell lysates with anti-FLAG
antibodies (Simga-Aldrich) or anti-ATM antibodies (Oncogene). Kinase activity was
determined by the ability of the kinases to phosphorylate PHAS-1, a known substrate of
all three kinases [165]. Briefly, ATM, ATR or DNA-PK were mixed in kinase reaction
buffer (20 mM Hepes, pH 7.4; 100 µM NaCl; 20 mM MgCl2; 20 mM MnCl2; 2 mM
DTT) containing 10 µM ATP, 10 µCi γ-ATP (Perkin Elmer Life Sciences, Boston, MA),
0.5 µg DNA, 1 µg PHAS-1 or 1 µg recombinant RPA (generously provided by John
Turchi, Wright State University, Dayton, OH) and incubated for 30 minutes at 37°C.
Kinase activity was inhibited by the addition of 12 µM wortmanin. To terminate the
reaction, LDS sample buffer was added to each reaction and the samples boiled for 5
minutes. The samples were loaded onto 12% Bis-Tris gels and subjected to
electrophoresis and phospho-imaging.
96
C. RESULTS
Recombinant FLAG-tagged ATM and ATR were generated by transfecting the appropriate cDNA into 293T cells. The recombinant proteins were purified by loading
ATM- or ATR-expressing 293T lysates onto an anti-FLAG affinity column. Eluted fractions containing the recombinant proteins were collected and assessed for purity by coomassie staining of SDS-PAGE gels (Figure 18C and D). To test for kinase activity of the recombinant proteins, in vitro kinase activities were run with FLAG-ATM immunoprecipitated from 293T cells, or purified recombinant FLAG-ATR. In both cases, the recombinant kinases phosphorylated PHAS-1, and ATR was able to phosphorylate
RPA as well (Figure 18B and D).
97
Figure 18. Generation of recombinant ATM and ATR. Recombinant ATM and ATR
were generated using FLAG-tagged cDNA constructs transfected into human 293T cells.
A, Western immunoblot showing increased amounts of ATM in 293T cells transfected
with FLAG-ATM cDNA. B, In vitro kinase reaction using DNA-PK, ATM
immunoprecipitated from 293T cells with anti-ATM antibodies, or FLAG-ATM
immunoprecipitated from transfected 293T cells with anti-FLAG antibodies. PHAS-1
was used as a substrate and phosphorylation was visualized by auto-radiography of
incorporated γ-ATP. C, Coomassie stained gel of fractions eluted from an anti-FLAG
column loaded with FLAG-ATM-expressing 293T lysate. D, Coomassie stained gel of
FLAG-ATR transfected 293T lysate, anti-FLAG column flow-through, and pooled fractions eluted from an anti-FLAG column loaded with FLAG-ATR-expressing 293T
lysate. E, In vitro kinase reaction using DNA-PK and purified recombinant FLAG-ATR.
PHAS-1 and recombinant hRPA were used as substrates and phosphorylation was visualized by auto-radiography of incorporated γ-ATP. Wortmanin was used as an
inhibitor of DNA-PK and ATR kinase activity. 98
FIGURE 18
99
D. DISCUSSION Accomplishments: We have generated ATM- and ATR-expressing 293T lysates. With
ATM, we were able to immunoprecipitate the recombinant protein using anti-FLAG
antibodies and show that the protein had kinase activity towards PHAS-1. We were also
able to run some of the ATM-expressing lysates on an anti-FLAG column, and purify the recombinant ATM. With regards to ATR, we generated highly purified and active ATR
that was able to phosphorylate PHAS-1 and recombinant RPA in vitro.
Technical limitations: The major limitation for generation of ATM was the relative
instability of the cDNA vector. Frequent recombination events and/or mutations,
visualized by changes in fraction lengths following restriction enzyme digests, occurred
during bacterial amplification that disrupted the gene and led to production of an inactive
final product. For ATR, the major limitation was time.
Future directions/conclusions: Once purified and active kinases are available,
experiments can be performed to look at the ability of the kinases to phosphorylate
specific proteins, such as Mre11 and RPA. The ability of the kinases to phosphorylate
Mre11 and RPA in vitro is only suggestive of what may actually occur in vivo, but is an
important first step in determining the complex interactions that occur in the DNA
damage response. The activity of these kinases to known substrates (such as PHAS-1, p53, Chk2, and others) can be monitored in the presence/absence of the MRN complex
and/or RPA. This may provide additional information on the ATM/ATR activating properties/potential of the MRN complex and RPA. Another use of the purified kinases is 100
to phosphorylate recombinant Mre11 and/or RPA, and then use these phosphorylated
proteins for further studies (elaborated on in later sections).
III. MRN Complex
A. RATIONALE
The MRN complex is important in the DNA damage response, and hypothesized to be
involved in multiple steps or stages of the response. In order to conduct in vitro
experiments to investigate the function of this complex, we needed to generate the
purified, recombinant complex. A baculvirus expression system for the production and
purification of the MRN complex has been previously generated and was generously
provided by Tanya Paull (University of Texas at Austin, Austin, TX) [42]. This system
produces 6 x His-tagged proteins that can be purified by column chromatography. As
reported previously [42], Mre11 can be produced by itself, but Rad50 and Nbs1 can only be produced when cells are co-infected with Mre11-encoding baculoviruses. This leads to the possibility of producing the following combinations: Mre11, Mre11/Rad50,
Mre11/Nbs1, or Mre11/Rad50/Nbs1. The ability to produce different combinations allows for the possibility of investigating the specific activity/function of the member(s) of the complex, as well as the member(s) of the complex that interacts with other proteins of interest, such as RPA.
101
B. MATERIALS AND METHODS
Cell line and treatment: Sf9 insect cells (S. frugiperdoa ovarian tissue) were obtained
from Yolanda Sanchez (University of Cincinnati, Cincinnati, OH) and maintained at
27°C in Sf-900 II serum free medium (Gibco). For viral amplification, virus was added to
the cells at a cell density of 5.0x105 cells per ml. For protein expression, virus was added to the cells at a cell density of 1.0x106 cells per ml.
Viral amplification: High-titer stocks of Mre11, Rad50 and/or Nbs1 baculoviruses were
prepared as previously described [42]. Briefly, viruses were added to cultures of Sf9 cells
at dilutions of 1:100-1:200 and incubated at 27°C for 4 days. Since baculoviruses are
lytic viruses, mature viruses are released into the medium after cell lysis. Following the
incubation period, cellular debris was centrifuged out of solution and the virus-enriched
supernatant was collected. A total of 4 serial amplifications were done in order to
generate high-titer stock solutions.
Protein expression: Protein expression was also done in Sf9 cells using viral dilutions of
1:20 (Mre11), 1:5 (Rad50) and 1:10 (Nbs1). Following an incubation of 48 hours, the
cells were pelleted, washed with PBS, and resuspended in insect cell lysis buffer (20 mM
potassium phosphate, pH 7.4; 10% glycerol; 500 mM KCl; 0.2 mM EDTA; 1 mM PMSF;
and 5 mM β-mercaptoethanol). Following sonication, cellular lysates were cleared by
centrifugation at 25,000 x g for 2 hours and then dialyzed into purification buffer (20 mM
potassium phosphate, pH 7.4; 10% glycerol; 50 mM KCl, and 5 mM β-mercaptoethanol). 102
Protein purification: Recombinant proteins were purified from dialyzed supernatants by
utilization of a nickel-NTA superflow resin column (Qiagen). After loading the dialyzed
supernatants onto the column and washing with 10-15 column volumes of purification
buffer containing 5 mM imidazole, the proteins of interest were eluted from the column
using purification buffer supplemented with 100 mM imidazole. Fractions with purified
Mre11, Rad50 and/or Nbs1 were determined by western immunoblotting and coomassie
stained gels. Those fractions containing the desired protein(s) were pooled together and
dialyzed into buffer B (20 mM Tris-HCl, pH 8.0; 50 mM NaCl; 0.05% Tween-20; 10%
glycerol; and 1 mM DTT).
In vitro Mre11 phosphorylation reaction: A fractionated HeLa extract was prepared as
previously described [166]. The phosphorylation reaction entailed incubating purified
recombinant Mre11 (dialyzed into HeLa extract dialysis buffer (45 mM Hepes, pH 7.8;
70 mM KCl; 7.4 mM MgCl2; 0.9 mM DTT; 0.4 mM EDTA, 3.4% glycerol)) in the HeLa
extract supplemented with 50 µg/ml creatine phosphate kinase (CPK; Sigma-Aldrich), 22
mM creatine phosphate (CP; ICN Biomedicals, Aurora, OH), 2 mM ATP, 500 µg/ml lambda DNA, 4 µM okadeic acid (Calbiochem), 2.5 mM (-)-p-Bromotetramisole oxalate
(Sigma-Aldrich), 500 µM cantharidin (Sigma-Aldrich), and 500 nM microcystin LR
(Sigma-Aldrich) for 3 hours at 30°C. Following the 3 hour incubation, the recombinant phosphorylated Mre11 was purified from the extract by re-running the reaction mixture through the Ni-NTA column as described previously.
103
Phosphatase treatment: Phosphorylated Mre11 eluted from the Ni-NTA column was
treated with 50 units of alkaline calf intestinal phosphatase (CIP; New England Biolabs,
Beverly, MA) in 1 X NEBuffer 3 (10 mM NaCl, 5 mM Tris-HCl, 1 mM MgCl2, 0.1 mM
DTT) at 37°C for 35 minutes. The non-CIP treated samples were incubated at 37°C for
30 minutes in 1 X NEBuffer 3 in the absence of CIP.
C. RESULTS
Ni-NTA column purification: Recombinant Mre11, Mre11/Rad50 and Mre11/Nbs1 were generated using the baculovirus expressing system. Mre11-expressing and Mre11/Nbs1- expressing lysates were purified using a Ni-NTA column (Figure 19A, B, C and D).
Mre11/Rad50-expressing lysates were generated (Figure 19E), but never purified.
In vitro Mre11 phosphorylation: Use of the modified HeLa extract as a means of phosphorylating recombinant proteins in vitro has previously been used (Nuss et al., submitted manuscript). Using the same HeLa extract, we were able to produce a slower migrating form of Mre11 that upon phosphatase treatment returned to normal migration
(Figure 20A and B). These data suggested that we were able to phosphorylate recombinant Mre11 in vitro.
104
Figure 19. Generation of recombinant Mre11, Rad50 and Nbs1. Recombinant His-
tagged Mre11, Rad50 and Nbs1 were generated using recombinant protein-expressing
baculoviruses in Sf9 insect cells. A, Western immunoblot showing the presence of Mre11 in eluted fractions from a Ni-NTA column loaded with Mre11-expressing Sf9 lysates. B,
Coomassie stained gel of eluted fractions from a Ni-NTA column loaded with Mre11- expressing Sf9 lysates. C, Western immunoblot for Mre11 and Nbs1 from pooled fractions eluted from a Ni-NTA column loaded with Mre11/Nbs1-expressing Sf9 lysates.
D, Coomassie stained gel of fractions eluted from a Ni-NTA column loaded with
Mre11/Nbs1-expressing Sf9 lysates. E, Western immunoblot of Sf9 lysate and 293T lysate (as a positive control) demonstrating the expression of recombinant Rad50. 105
FIGURE 19
106
Figure 20. Generation of phosphorylated recombinant Mre11. Recombinant His-
tagged Mre11 was phosphorylated in vitro by incubating for 3 hours in a HeLa nuclear extract described in “Materials and Methods”. The phosphorylated Mre11 was purified from the reaction mixture by running the reaction volume over the Ni-NTA column as described previously. A, Western immunoblot showing 3 hour incubation in HeLa extract leads to generation of a slower migrating form of Mre11. B, The slower migrating form of Mre11 returned to a faster migrating form following phosphatase (CIP) treatment. C,
Western immunoblot of phosphorylated Mre11 eluted from the Ni-NTA column. D,
Coomassie stained gel of phosphorylated Mre11 eluted for the Ni-NTA column. 107
FIGURE 20
108
D. DISCUSSION
Accomplishments: We produced Mre11, Mre11/Nbs1 and Mre11/Rad50 expressing Sf9
lysates. We were also able to purify Mre11 and Mre11/Nbs1 using column
chromatography, and to phosphorylate recombinant Mre11 in vitro using a modified
HeLa nuclear extract.
Technical limitations: The generation of a Rad50-expressing lysate was difficult. After
obtaining a second aliquot of Rad50 baculovirus from Dr. Paull, we were able to observe
Rad50 expression by western blot. The purity of the proteins eluted from the Ni-NTA
column was suboptimal. Phosphorylated recombinant Mre11 was very difficult to purify.
Following elution from the Ni-NTA column, we had problems recovering phospho-
Mre11 after dialysis. We used different dialysis membranes and columns, both of which
worked well with the non-phosphorylated Mre11, but had a persistent inability to recover
the phosphorylated protein. The cause of this loss is still not known, but may have to do with protein solubility or protein adherence to the membranes.
Future directions/conclusions: Sf9 lysates expressing Mre11, Mre11/Rad50 or
Mre11/Nbs1 have been generated, whereas lysates expressing Mre11/Rad50/Nbs1 still need to be generated. These recombinant proteins need to be purified from the lysates by column chromatography for use in in vitro experiments. In particular, work needs to be done to generate the entire complex in order to study a more physiologically relevant complex, and to generate in vitro phosphorylated MRN complex in order to study the role of protein phosphorylation on the function of the complex. 109
IV. RPA
A. RATIONALE
Recombinant RPA was generated as previously described [167] and generously provided
by John Turchi (Wright State University, Dayton, OH). An in vitro phosphorylated form of RPA has also been generated in the Turchi laboratory, and a collaborative effort has been made to compare the in vitro phosphorylated-RPA and in vivo phosphorylated-RPA.
Our collaborative efforts have resulted in a submitted manuscript demonstrating that the in vitro phosphorylated RPA is a reasonable representation of what occurs in vivo, and more physiologically relevant compared to serine-to-aspartate mutants often used to study the effects of protein phosphorylation (Nuss et al., submitted manuscript).
B. MATERIALS AND METHODS
Cell lines and treatments: HeLa cells were obtained from American Type Culture
Collection (ATCC; Manassas, VA) and maintained at 37°C and 5% CO2 in DMEM
(Gibco) supplemented with 10% FBS (Hyclone) and 1% penicillin-streptomycin (Gibco).
Cells were synchronized in S-phase of the cell cycle as previously described [77].
Briefly, cells were incubated in growth medium containing aphidocolin (final concentration of 1 µM; Sigma-Aldrich) for 15 hours. Following incubation, the aphidocolin-containing medium was removed, cells were washed with PBS, and then incubated in fresh medium without aphidocolin for an additional 2 hours at 37°C. For UV exposure, growth medium from S-phase cells was removed (and held at 37°C) and the 110
cells washed with PBS. The PBS was replaced with minimum essential medium (MEM;
Gibco) without phenol red and cells were treated with 20 J/m2 UVC using a low-pressure
mercury lamp (Mineralight lamp; model UVG-11; UVP, Inc., San Gabriel, CA) with a
maximal output at 254 nm. Following UV exposure, the MEM was removed and
replaced with the original growth medium, and cells were allowed to recover for 8 hours
at 37°C before harvesting. For hydroxyurea (HU; Sigma-Aldrich) treatment, S-phase
cells were incubated in growth medium containing 2 mM HU for 3 hours before harvesting.
Cell lysate formation and immunoprecipitation: Following treatment, cells were harvested and lysed using SDS-PAGE lysis buffer (50 mM Tris-HCl, pH 7.5; 150 mM
NaCl; 0.1% NP-40; 5 µg/ml pepstatin; 5 µg/ml leupeptin; 5 µg/ml aprotonin; 10 mM
NaF; 10 mM β-glycerophosphate; 1 mM Na3VO4; 1 mM PMSF). Approximately 1000
µg total protein from each sample was immunoprecipitated using 6 µg of anti-RPA-p34
antibodies (Bethyl Laboratories, Inc., Montgomery, TX) cross-linked to 60 µL of protein
G-agarose beads (Invitrogen, Carlsbad, CA). The antibodies were cross-linked to the
beads by incubating together for 1 hour at room temperature with end-over-end mixing.
The beads were then washed twice with 200 mM triethanolamine (Sigma-Aldrich) and
then incubated in 20 mM dimethylpimelimidate (in 200 mM triethanolamine; DMP;
Sigma-Aldrich) for 30 minutes at room temperature. The DMP solution was replaced
with 50 mM Tris-HCl, pH 7.5 for an additional 15 minutes. The Tris-HCl was removed
and the cell lysates were then added to the beads. Following 14-16 hours incubation at
4°C with end-over-end mixing, the immunoprecipitates were washed once with PBS, and 111
the proteins were eluted from the beads using rehydration buffer (7 M urea, 2 M thiourea,
4% CHAPS, 40 mM DTT, 2.0% IPG buffer).
First dimension/isoelectric focusing: For recombinant RPA, approximately 1.5 µg of recombinant human RPA (rhRPA), hyper-phosphorylated RPA (hypRPA), or a mixture of 1.5 µg each were mixed with 2D rehydration buffer. For HeLa lysates, the proteins were eluted from the immunoprecipitation reaction using the 2D rehydration buffer. The protein-containing rehydration buffer was placed in 13 cm strip holders (Amersham
Biosciences, Buckinghamshire, England) and Immobiline DryStrip polyacrlyamide strips with an immobilized pH gradient of pH 3-10 (Amersham Biosciences) were immersed in the buffer; mineral oil was layered on top of each strip, the cover added, and the holders then placed in a Pharmacia Biotech IPGphor isoelectric focusing system
(Amersham Biosciences) under the following conditions: 20°C, 50 µA/strip. The electrophoretic program was run in four steps: Step 1 – 50 V, 12 hours; Step 2 – 500 V, 1 hour; Step 3 – 1000 V, 1 hour; Step 4 – 8000 V, 8 hours.
Second Dimension: Following isoelectric focusing, the strips were removed from the strip holders, rinsed in deionized H2O, and then incubated in equilibration buffer 1 for 15
minutes at room temperature (50 mM Tris-HCl, pH 8.8; 6 M urea; 30% glycerol; 2%
SDS; 1% DTT). The strips were then transferred to equilibration buffer 2 (50 mM Tris-
HCl, pH 8.8; 6 M urea; 30% glycerol; 2% SDS; 2.5% iodoactamide) and incubated for an additional 15 minutes at room temperature. The strips were placed on top of 12% polyacrlyamide SDS gels with 0.5% agarose layered on top of the strips. The 2D gels 112
were electrophoresed at 16°C for 15 minutes at 10 mA/gel and 300 V; then 25 mA/gel
and 300 V for 5 hours.
Western Immunoblots: The SDS-polyacrlyamide gels were transferred to PVDF
membranes (Millipore Corp., Bedford, MA), and the membranes probed using anti-RPA-
p34 antibody (Neomarkers, Freemont, CA; 1:5000). The secondary antibody was
horseradish peroxidase-linked anti-mouse (Amersham Biosciences; 1:3000), and bound
antibodies were visualized using chemiluminescent detection.
C. RESULTS
Two-dimensional (2D) gel electrophoresis that separated proteins in the first dimension
based on isoelectric focusing point (PI point), and in the second dimension by molecular
weight was very useful in visualizing the different phosphorylated forms of RPA-p34.
With the addition of phosphate, RPA shifted to a more acidic PI (to the left) and increased molecular weight (slower migrating) producing a series of spots that shift to the
left and up following addition of phosphate groups (Figure 21B). In one-dimensional
SDS-PAGE, RPA-p34 hyper-phosphorylation shows 5 (possibly 6) distinct bands of
RPA-p34 (Figure 21A). 2D SDS-PAGE shows 6 (possibly 7-8) levels of RPA-p34 phosphorylation, and multiple variants (individual dots) at each level (Figure 21C). While the HU- and UV- induced 2D patterns were very similar, the array of dots indicate the complexity of RPA-p34 phosphorylation (Figure 21C). The in vitro phosphorylated RPA produced a pattern of dots that was very similar to the top two levels of dots produced by
UV-induced in vivo phosphorylation of RPA-p34 (Figure 21D), suggesting that the in 113
vitro phosphorylated RPA recapitulates the in vivo RPA phosphorylation following treatment with genotoxic agents. 114
Figure 21. One- and two-dimensional SDS-PAGE of HeLa and recombinant RPA
demonstrate the presence of multiple RPA-p34 phosphorylations. HeLa cells were
treated as described in “Materials and Methods” and lysates prepared and probed for
RPA-p34. A, One-dimensional SDS-PAGE and western blot analysis of extracts prepared
from control (lane 1), UV- (lane 2) and HU- (lane 3) treated cells. Increasing
concentrations of purified in vitro hyper-phosphorylated RPA (hypRPA; lanes 4-6) or
rhRPA (lanes 7-9). B, Two-dimensional SDS-PAGE of HeLa lysates (same lysates as depicted in A). C, Two-dimensional SDS-PAGE of rhRPA, hypRPA and a mixture of
rhRPA and hypRPA. D, Enlarged picture of 2-D SDS-PAGE of hypRPA and HeLa UV
treated lysates for comparison. Circles indicate spots that appear to migrate in a similar
pattern. 115
FIGURE 21
116
D. DISCUSSION
Accomplishments: The 2D gels of the in vivo, UV-induced phosphorylated RPA-p34 and in vitro hyper-phosphorylated RPA-p34 showed remarkable concordance. The pattern of dots produced suggested that many of the same residues are phosphorylated in vitro and in vivo.
Technical limitations: It has been reported that RPA-p70 is also phosphorylated [168,
169]. The 2D analysis of RPA-p70 did not work well enough to distinguish the phosphorylated forms of RPA-p70. This is probably due to solubility issues and would require additional optimization of the procedure for visualization.
Future directions/conclusion: Being able to generate hyper-phosphorylated recombinant
RPA in vitro that resembles the genotoxin-induced in vivo hyper-phosphorylated RPA allows us the opportunity to do in vitro assays trying to delineate the significance of
RPA-p34 hyper-phosphorylation. The next step is to do protein interaction assays to compare the ability of recombinant non-phosphorylated RPA and recombinant hyper- phosphorylated RPA to bind to the MRN complex in order to determine if RPA phosphorylation plays a role in this interaction.
V. DISCUSSION
The goal of this dissertation is to investigate the interplay of the MRN complex and RPA in the DNA damage response. The specific questions being addressed are: 117
(1) Do the MRN complex and RPA interact in the DNA damage response?
(2) Is this interaction similar for all forms of DNA damage?
(3) Does protein phosphorylation play a role in this interaction?
(4) What is the functional consequence or significance of this interaction?
While cell culture work is the main model in which we address these questions, the
potential contribution of recombinant proteins for exploring each question is discussed
below.
Question 1: Do the MRN complex and RPA interact in the DNA damage response?
In cell culture, co-immunoprecipitation reactions were used to demonstrate that
antibodies against the MRN complex were able to precipitate RPA, and antibodies against RPA precipitated Mre11. However, those experiments do not rule out the possibility that other proteins are mediating the interaction between the MRN complex
and RPA. The use of purified recombinant proteins in co-immunoprecipitation reactions
and/or ELISAs will help to address the question of a direct interaction between these
proteins. In addition, since the MRN complex can be purified as subcomponents (Mre11,
Mre11/Rad50, Mre11/Nbs1 and Mre11/Rad50/Nbs1), the subunit of the MRN complex
responsible for this interaction can be determined.
Question 2: Is this interaction similar for all forms of DNA damage?
This question will not be significantly enhanced by the use of recombinant proteins
unless there is a difference in the phosphorylation status of the MRN complex and RPA
following different types of damage, and if those phosphorylated forms can be 118
recapitulated in vitro. The data thus far suggest that there is not a difference in genotoxin-
induced phosphorylation in vivo that can be recapitulated in vitro.
Question 3: Does protein phosphorylation play a role in this interaction?
Examination of the similarities between the hyper-phosphorylated RPA generated in vitro with the in vivo damage-induced hyper-phosphorylated RPA will help verify if the in vitro phosphorylation protocol is physiologically similar to what occurs in vivo. If this protocol does produce a physiologically relevant phosphorylated protein, the protocol may then be used to phosphorylate the MRN complex in vitro, although validation will need to be done for the phosphorylated MRN complex in a manner similar to that done for RPA. Once the phosphorylated recombinant proteins have been generated, pull-down reactions similar to those described to address question 1 can be done to determine if phosphorylation of the MRN complex and/or RPA modulate the ability of these proteins to interact.
Question 4: What is the functional consequence or significance of this interaction?
The MRN complex and RPA both have functional activity that can be measured in vitro: the MRN complex has DNA binding, nuclease, strand-dissociation, strand-annealing and
DNA tethering activities; and RPA has DNA binding and unwinding capabilities. The interaction of the MRN complex and RPA may modulate any one of these activities.
With recombinant proteins, in vitro assays can be done to compare the activities of the
MRN complex and RPA separately with activities when the proteins are combined.
119
In addition to what has been described for the MRN complex and RPA, recombinant
ATM and ATR have been generated. These two kinases can be useful in different ways.
First, ATM and ATR can be used to phosphorylate the MRN complex and RPA in vitro.
This will help determine whether these kinases are able to phosphorylate these proteins, as well as generate sufficient quantities of the phosphorylated proteins to identify phosphorylated residues by proteomics methods (e.g. mass spectrometery, two- dimensional gel electrophoresis). Identification of the phosphorylated sites may then be used to generate, by site-directed mutagenesis, expression vectors that block protein phosphorylation. These mutated proteins may then be used in functional assays to help determine the significance of MRN and RPA phosphorylation. Second, ATM and ATR kinase activity towards other known substrates can be investigated in the presence/absence of the MRN complex and RPA. Recently Lee and Paull reported that addition of the MRN complex to an in vitro reaction mixture containing ATM led to increased ATM kinase activity towards p53 and Chk2 [26].
The in vitro plans proposed here, using recombinant ATM, ATR, MRN complex and
RPA, will provide confirming data to that which has been obtained through cell culture methods and provide additional data, particularly on functional implications of the MRN complex/RPA interaction. The combination of cell culture work and in vitro experimentation with recombinant proteins will give a more precise understanding of how the MRN complex and RPA interact, the role of phosphorylation on this interaction, and the functional consequences of this interaction. 120
CHAPTER 6. Conclusion
The pivotal question addressed by this dissertation is whether or not the MRN complex
and RPA interact in the DNA damage response; our data indicate that they do. We have
demonstrated this interaction by the co-localization of the MRN complex and RPA at
sites of stalled replication forks and some forms of DNA damage as well as by the co-
immunoprecipitation of these proteins, with an increased interaction observed in both S-
phase of the cell cycle and following replicative stress. The co-immunoprecipitation
experiments, including consecutive co-immunoprecipitations, suggest a direct interaction
between the MRN complex and RPA and not an interaction bridged by structural
components such as DNA or a large multi-protein complex.
Evidence that the MRN complex and RPA interact fostered the following questions which will be addressed below:
(1) Is the interaction between the MRN complex and RPA similar for all forms of
DNA damage?
(2) Does protein phosphorylation play a role in the MRN complex and RPA
interaction?
(3) What is the functional consequence or significance of the MRN complex and
RPA interaction?
In addition to addressing these questions, the data discussed here suggests a new paradigm about the DNA damage response that emphasizes the importance of the MRN complex and RPA as sensors and effectors for multiple types of DNA damage. 121
(1) Is the interaction between the MRN complex and RPA similar for all forms of DNA damage?
To determine if the interaction of the MRN complex and RPA is similar for all forms of
DNA damage, we treated cells with different types of DNA-damaging agents, including ultraviolet light (UV), hydroxyurea (HU), camptothecin (CAMPT), etoposide (ETOP) and mitomycin C (MMC) and then assayed for the ability of the MRN complex and RPA to co-localize to nuclear foci. Following treatment with UV, HU and CAMPT the MRN complex and RPA co-localized to a very high degree. In contrast, treatment with ETOP or MMC led to MRN complex and RPA foci formation without co-localization.
Specifically, after ETOP treatment cells contained either phospho-Nbs1 or RPA foci, but not both; and after MMC treatment cells contained only phospho-Nbs1 foci. Prolonged exposure to HU, which led to replication fork collapse and generated DNA double-strand breaks (DSBs), resulted in loss of phospho-Nbs1 and RPA foci co-localization even though they both formed independently within the same cell. These results suggested that the interaction of the MRN complex and RPA was dependent upon the type of damage induced within the cell, and may not occur in the response to DSBs. These data also suggested that the MRN complex and RPA may be able to function independently of each other.
(2) Does protein phosphorylation play a role in the MRN complex and RPA interaction?
We demonstrated that phosphatase treatment decreased the ability of the MRN complex and RPA to interact. Co-immunoprecipitation of the MRN complex and RPA after 122
replicative stress was abrogated by phosphatase treatment of lysates prior to
immunoprecipitation and by phosphatase treatment of the immunoprecipitate pellet. Co-
treatment with phosphatase and phosphatase inhibitors did not affect the interaction of the
MRN complex and RPA. These data suggested that the interaction was dependent on
protein phosphorylation, but did not reveal whether the critical factor was MRN
phosphorylation, RPA phosphorylation or both.
(3) What is the functional consequence or significance of the MRN complex and RPA interaction?
The functional consequence(s) of the interaction between the MRN complex and RPA is still not known. Our data demonstrate that the MRN complex can form foci independent of RPA foci, but that siRNA-directed depletion of RPA abrogates phospho-Nbs1 foci formation. These data suggest that the MRN complex formation is dependent upon the presence of RPA, though the level of RPA may be below the threshold for detection using immunofluorescence. While this suggests one possible function of the interaction between the MRN complex and RPA, further work needs to be done to investigate other possible functional implications of this interaction. Our efforts to produce purified recombinant proteins were the initial step to investigate these types of questions using in vitro functional assays, but we have currently not progressed to the point of being able to complete these assays.
Historically the MRN complex has been best known for its role in DNA DSB repair, but
our data, and other recent reports, clearly show that the MRN complex has many 123
additional roles. It has been demonstrated that the MRN complex plays a role in DNA
replication, cell cycle checkpoints, telomere maintenance, signaling/sensing of DNA
damage, and response to stalled replication forks [11, 14-19, 23, 121]. Our data suggest
that the MRN complex may act as a sensor of multiple types of DNA lesions by
recognizing and binding to RPA-coated damaged DNA. This binding may be necessary
for the correct initiation and full activation of the DNA damage response. Investigating
the role of the MRN complex in the DNA damage response, and the effect its interaction
with RPA has on this role, has many implications for DNA repair. More specifically, this
understanding will provide better information on the cellular components necessary for
maintaining genomic stability. In addition to sensor functions, the MRN complex and
RPA also have roles as effectors in the damage response. How the MRN complex/RPA
interaction affects the sensor and effector functions of MRN complex and/or RPA
remains to be elucidated.
The data that we have collected has led to the generation of a model of events that takes place following induction of replicative stress and/or DNA damage (Figure 22). This model considers the availability of single-stranded DNA (ssDNA) for RPA binding and subsequent hyper-phosphorylation to explain the immunofluorescent foci and co- immunoprecipitation data presented earlier. The MRN complex and RPA are both sensors of DNA damage. Additionally, RPA protects ssDNA from nuclease-mediated degradation and becomes hyper-phosphorylated to signal replication stress, single-strand breaks (SSBs) and some types of double-strand breaks (DSBs). Stalled replication forks, if unrepaired, lead to replication fork collapse and subsequent DNA DSBs and formation 124
of early DNA repair foci (similar to Mirzoeva and Petrini’s Type II foci [30]). These type II foci contain MRN and may also contain visually detectable RPA depending on the type of DNA damage and amount of ssDNA. Subsequently, the foci evolve into type III
DNA repair foci that represent repair of DSBs either through a recombinatorial pathway using Rad51/Rad52 or non-homologous end joining. These processes of DSB repair may occur within the same cell, or different cells, thus leading to differences in protein composition of the foci. The role of the MRN complex and RPA as sensors and/or effectors in the DNA damage response probably occurs concurrently within a cell population and within a single cell. Up to this point, we have not tried to distinguish or
Figure 22. Model depicting the interaction of the MRN complex and RPA in response to DNA damage 125
differentiate the possible sensor/effector functions of these proteins and if the observed
MRN/RPA interaction is limited to a subset of these functions.
These observations have given rise to further questions. One area of work that needs to be
expanded is the work with siRNA-mediated depletion of Mre11 or RPA. Data collected
from such experiments will help address the role of this interaction on the activation of
the DNA damage response. Another question focuses on the difference in RPA foci
formation following treatment with different genotoxic agents. Our model proposes that
the difference is due to the amount of ssDNA present at the site of damage. Further
efforts are required to address this question. Useful approaches may include chromatin-
immunoprecipitation experiments in which DNA bound by the MRN complex and/or
RPA is precipitated and the nature of the DNA (specifically the single-stranded vs.
double-stranded nature) is examined. Additionally, the functional significance of the
MRN complex/RPA interaction needs to be addressed at a biochemical level, probably
through in vitro functional assays. Potential experiments to investigate this may include
measuring the DNA binding ability of Mre11 and RPA together, the ability of MRN to
tether or bind separate fragments of DNA in the presence/absence of RPA, and the change in Mre11 nuclease activity in the presence of RPA.
This work has been conducted in cell culture models using transformed cell lines. Further
work needs to be done to determine if similar findings occur in primary cell lines and
animal models. In animal models, it would be of interest and value to understand the
organ-specific responses to the agents we have investigated. HU, CAMPT, ETOP and 126
MMC are all currently employed as chemotherapeutic agents. Further understanding of
the cellular response to these agents will provide information to help physicians utilize
these agents in a manner that increases effectiveness and decreases the detrimental side
effects. Additionally, tumor drug resistance can occur, and understanding the cellular
response to these agents will help identify the processes behind this resistance and may
suggest approaches to overcome this resistance and increase the effectiveness of these
agents.
Direct application of the importance of the MRN complex /RPA interaction and the DNA
damage response in a more clinically oriented manner can be seen in the kidney. In the
kidney medulla, the concentration of NaCl is increased about two-fold in order to
concentrate the urine. This high concentration of NaCl leads to the trans-localization of
Mre11 from the nucleus into the cytoplasm and induces DNA DSBs which do not
activate the DNA damage response and are not repaired [170-172]. These observations
have clinical relevance with regards to renal carcinomas, polycystic kidney diseases and
tuberous sclerosis complex. These diseases are all associated with somatic mutations in renal tubule epithelial cells. The altered response to DNA damage in renal cells may explain why autosomal dominant polycystic kidney disease and tuberous sclerosis complex have phenotypic expression within the kidney. While the cause/effect relationship of the loss of Mre11 from the nucleus and loss of activation of the DNA damage response are not known, our data that suggests the MRN complex and RPA function together as sensors of damage may explain this loss of activation of the DNA damage response. The studies that looked at the trans-localization of Mre11 did not look 127
at RPA or a host of other protein factors in the DNA damage response. Further studies to determine if the loss of Mre11 and/or disruption of the MRN complex/RPA interaction
plays a role in these responses to DNA damage in the kidney will be beneficial for trying
to develop therapeutic approaches to delay renal involvement and subsequent renal
failure. 128
REFERENCES
1. Barzilai, A. and K. Yamamoto, DNA damage responses to oxidative stress. DNA
Repair (Amst), 2004. 3(8-9): p. 1109-1115.
2. Zhou, B.B. and S.J. Elledge, The DNA damage response: putting checkpoints in
perspective. Nature, 2000. 408(6811): p. 433-439.
3. Begley, T.J. and L.D. Samson, Network responses to DNA damaging agents.
DNA Repair (Amst), 2004. 3(8-9): p. 1123-1132.
4. Shiloh, Y., ATM and related protein kinases: safeguarding genome integrity.
Nat.Rev.Cancer, 2003. 3(3): p. 155-168.
5. Kurz, E.U. and S.P. Lees-Miller, DNA damage-induced activation of ATM and
ATM-dependent signaling pathways. DNA Repair (Amst), 2004. 3(8-9): p. 889-
900.
6. Rothstein, R., B. Michel, and S. Gangloff, Replication fork pausing and
recombination or "gimme a break". Genes Dev., 2000. 14(1): p. 1-10.
7. Bierne, H. and B. Michel, When replication forks stop. Mol.Microbiol., 1994.
13(1): p. 17-23.
8. Michel, B., S.D. Ehrlich, and M. Uzest, DNA double-strand breaks caused by
replication arrest. EMBO J., 1997. 16(2): p. 430-438.
9. Maser, R.S., et al., hMre11 and hRad50 nuclear foci are induced during the
normal cellular response to DNA double-strand breaks. Mol.Cell Biol, 1997.
17(10): p. 6087-6096.
10. Nelms, B.E., et al., In situ visualization of DNA double-strand break repair in
human fibroblasts. Science, 1998. 280(5363): p. 590-592. 129
11. Lisby, M., et al., Choreography of the DNA Damage Response; Spatiotemporal
Relationships among Checkpoint and Repair Proteins. Cell, 2004. 118(6): p. 699-
713.
12. Lisby, M. and R. Rothstein, DNA damage checkpoint and repair centers.
Curr.Opin.Cell Biol, 2004. 16(3): p. 328-334.
13. Connelly, J.C. and D.R. Leach, Tethering on the brink: the evolutionarily
conserved Mre11-Rad50 complex. Trends Biochem Sci., 2002. 27(8): p. 410-418.
14. van den, B.M., R.T. Bree, and N.F. Lowndes, The MRN complex: coordinating
and mediating the response to broken chromosomes. EMBO Rep., 2003. 4(9): p.
844-849.
15. Yamaguchi-Iwai, Y., et al., Mre11 is essential for the maintenance of
chromosomal DNA in vertebrate cells. EMBO J., 1999. 18(23): p. 6619-6629.
16. Costanzo, V., et al., Mre11 protein complex prevents double-strand break
accumulation during chromosomal DNA replication. Mol.Cell, 2001. 8(1): p.
137-147.
17. Wang, Y., et al., BASC, a super complex of BRCA1-associated proteins involved
in the recognition and repair of aberrant DNA structures. Genes Dev., 2000.
14(8): p. 927-939.
18. Zhu, X.D., et al., Cell-cycle-regulated association of RAD50/MRE11/NBS1 with
TRF2 and human telomeres. Nat.Genet., 2000. 25(3): p. 347-352.
19. Ritchie, K.B. and T.D. Petes, The Mre11p/Rad50p/Xrs2p complex and the Tel1p
function in a single pathway for telomere maintenance in yeast. Genetics, 2000.
155(1): p. 475-479. 130
20. Carson, C.T., et al., The Mre11 complex is required for ATM activation and the
G2/M checkpoint. EMBO J., 2003. 22(24): p. 6610-6620.
21. Petrini, J.H., The Mre11 complex and ATM: collaborating to navigate S phase.
Curr.Opin.Cell Biol, 2000. 12(3): p. 293-296.
22. Petrini, J.H., The mammalian Mre11-Rad50-nbs1 protein complex: integration of
functions in the cellular DNA-damage response. Am.J.Hum.Genet, 1999. 64(5):
p. 1264-1269.
23. Petrini, J.H. and T.H. Stracker, The cellular response to DNA double-strand
breaks: defining the sensors and mediators. Trends Cell Biol, 2003. 13(9): p. 458-
462.
24. Uziel, T., et al., Requirement of the MRN complex for ATM activation by DNA
damage. EMBO J., 2003. 22(20): p. 5612-5621.
25. Mochan, T.A., et al., 53BP1 and NFBD1/MDC1-Nbs1 function in parallel
interacting pathways activating ataxia-telangiectasia mutated (ATM) in response
to DNA damage. Cancer Res., 2003. 63(24): p. 8586-8591.
26. Lee, J.H. and T.T. Paull, Direct activation of the ATM protein kinase by the
Mre11/Rad50/Nbs1 complex. Science, 2004. 304(5667): p. 93-96.
27. Horejsi, Z., et al., Distinct functional domains of Nbs1 modulate the timing and
magnitude of ATM activation after low doses of ionizing radiation. Oncogene,
2004. 23(17): p. 3122-3127.
28. Cerosaletti, K. and P. Concannon, Independent roles for nibrin and Mre11/Rad50
in the activation and function of Atm. J.Biol Chem., 2004. . 131
29. Mirzoeva, O.K. and J.H. Petrini, DNA replication-dependent nuclear dynamics of
the Mre11 complex. Mol.Cancer Res., 2003. 1(3): p. 207-218.
30. Mirzoeva, O.K. and J.H. Petrini, DNA damage-dependent nuclear dynamics of the
Mre11 complex. Mol.Cell Biol, 2001. 21(1): p. 281-288.
31. Raderschall, E., E.I. Golub, and T. Haaf, Nuclear foci of mammalian
recombination proteins are located at single-stranded DNA regions formed after
DNA damage. Proc.Natl.Acad.Sci.U.S.A, 1999. 96(5): p. 1921-1926.
32. Limoli, C.L., et al., UV-induced replication arrest in the xeroderma pigmentosum
variant leads to DNA double-strand breaks, gamma -H2AX formation, and Mre11
relocalization. Proc.Natl.Acad.Sci.U.S.A, 2002. 99(1): p. 233-238.
33. Savitsky, K., et al., A single ataxia telangiectasia gene with a product similar to
PI- 3 kinase. Science, 1995. 268(5218): p. 1749-1753.
34. McKinnon, P.J., Ataxia-telangiectasia: an inherited disorder of ionizing-
radiation sensitivity in man. Progress in the elucidation of the underlying
biochemical defect. Hum.Genet, 1987. 75(3): p. 197-208.
35. Stewart, G.S., et al., The DNA double-strand break repair gene hMRE11 is
mutated in individuals with an ataxia-telangiectasia-like disorder. Cell, 1999.
99(6): p. 577-587.
36. Taylor, A.M., A. Groom, and P.J. Byrd, Ataxia-telangiectasia-like disorder
(ATLD)-its clinical presentation and molecular basis. DNA Repair (Amst), 2004.
3(8-9): p. 1219-1225. 132
37. Digweed, M. and K. Sperling, Nijmegen breakage syndrome: clinical
manifestation of defective response to DNA double-strand breaks. DNA Repair
(Amst), 2004. 3(8-9): p. 1207-1217.
38. Varon, R., et al., Nibrin, a novel DNA double-strand break repair protein, is
mutated in Nijmegen breakage syndrome. Cell, 1998. 93(3): p. 467-476.
39. Carney, J.P., et al., The hMre11/hRad50 protein complex and Nijmegen breakage
syndrome: linkage of double-strand break repair to the cellular DNA damage
response. Cell, 1998. 93(3): p. 477-486.
40. Usui, T., et al., Complex formation and functional versatility of Mre11 of budding
yeast in recombination. Cell, 1998. 95(5): p. 705-716.
41. Gatei, M., et al., ATM-dependent phosphorylation of nibrin in response to
radiation exposure. Nat.Genet., 2000. 25(1): p. 115-119.
42. Paull, T.T. and M. Gellert, The 3' to 5' exonuclease activity of Mre 11 facilitates
repair of DNA double-strand breaks. Mol.Cell, 1998. 1(7): p. 969-979.
43. Trujillo, K.M., et al., Nuclease activities in a complex of human recombination
and DNA repair factors Rad50, Mre11, and p95. J.Biol Chem., 1998. 273(34): p.
21447-21450.
44. de Jager, M., et al., Human Rad50/Mre11 is a flexible complex that can tether
DNA ends. Mol.Cell, 2001. 8(5): p. 1129-1135.
45. Furuse, M., et al., Distinct roles of two separable in vitro activities of yeast Mre11
in mitotic and meiotic recombination. EMBO J., 1998. 17(21): p. 6412-6425.
46. Paull, T.T. and M. Gellert, A mechanistic basis for Mre11-directed DNA joining
at microhomologies. Proc.Natl.Acad.Sci.U.S.A, 2000. 97(12): p. 6409-6414. 133
47. D'Amours, D. and S.P. Jackson, The Mre11 complex: at the crossroads of dna
repair and checkpoint signalling. Nat.Rev.Mol.Cell Biol., 2002. 3(5): p. 317-327.
48. Paull, T.T. and M. Gellert, Nbs1 potentiates ATP-driven DNA unwinding and
endonuclease cleavage by the Mre11/Rad50 complex. Genes Dev., 1999. 13(10):
p. 1276-1288.
49. de Jager, M., et al., DNA-binding and strand-annealing activities of human
Mre11: implications for its roles in DNA double-strand break repair pathways.
Nucleic Acids Res., 2001. 29(6): p. 1317-1325.
50. Lee, J.H., et al., Regulation of Mre11/Rad50 by Nbs1: effects on nucleotide-
dependent DNA binding and association with ataxia-telangiectasia-like disorder
mutant complexes. J.Biol Chem., 2003. 278(46): p. 45171-45181.
51. Dong, Z., Q. Zhong, and P.L. Chen, The Nijmegen breakage syndrome protein is
essential for Mre11 phosphorylation upon DNA damage. Journal of Biological
Chemistry, 1999. 274(28): p. 19513-19516.
52. Stewart, G.S., et al., Residual ataxia telangiectasia mutated protein function in
cells from ataxia telangiectasia patients, with 5762ins137 and 7271T-->G
mutations, showing a less severe phenotype. J.Biol Chem., 2001. 276(32): p.
30133-30141.
53. Yuan, S.S., et al., Arsenic-induced Mre11 phosphorylation is cell cycle-dependent
and defective in NBS cells. DNA Repair (Amst), 2002. 1(2): p. 137-142.
54. Hirano, T., The ABCs of SMC proteins: two-armed ATPases for chromosome
condensation, cohesion, and repair. Genes Dev., 2002. 16(4): p. 399-414. 134
55. Durocher, D., et al., The FHA domain is a modular phosphopeptide recognition
motif. Mol.Cell, 1999. 4(3): p. 387-394.
56. Digweed, M., et al., SV40 large T-antigen disturbs the formation of nuclear DNA-
repair foci containing MRE11. Oncogene, 2002. 21(32): p. 4873-4878.
57. Yu, X., et al., The BRCT domain is a phospho-protein binding domain. Science,
2003. 302(5645): p. 639-642.
58. Hofmann, K. and P. Bucher, The FHA domain: a putative nuclear signalling
domain found in protein kinases and transcription factors. Trends Biochem Sci.,
1995. 20(9): p. 347-349.
59. Tauchi, H., et al., The forkhead-associated domain of NBS1 is essential for
nuclear foci formation after irradiation but not essential for hRAD50[middle
dot]hMRE11[middle dot]NBS1 complex DNA repair activity. J.Biol Chem., 2001.
276(1): p. 12-15.
60. Kobayashi, J., et al., NBS1 and its functional role in the DNA damage response.
DNA Repair (Amst), 2004. 3(8-9): p. 855-861.
61. Lim, D.S., et al., ATM phosphorylates p95/nbs1 in an S-phase checkpoint
pathway. Nature, 2000. 404(6778): p. 613-617.
62. Zhao, S., et al., Functional link between ataxia-telangiectasia and Nijmegen
breakage syndrome gene products. Nature, 2000. 405(6785): p. 473-477.
63. Wu, X., et al., ATM phosphorylation of Nijmegen breakage syndrome protein is
required in a DNA damage response [see comments]. Nature, 2000. 405(6785): p.
477-482. 135
64. Kim, S.T., et al., Substrate specificities and identification of putative substrates of
ATM kinase family members. Journal of Biological Chemistry, 1999. 274(53): p.
37538-37543.
65. Yazdi, P.T., et al., SMC1 is a downstream effector in the ATM/NBS1 branch of the
human S-phase checkpoint. Genes Dev., 2002. 16(5): p. 571-582.
66. Nakanishi, K., et al., Interaction of FANCD2 and NBS1 in the DNA damage
response. Nat.Cell Biol, 2002. 4(12): p. 913-920.
67. Lee, J.H., et al., Distinct functions of Nijmegen breakage syndrome in ataxia
telangiectasia mutated-dependent responses to DNA damage. Mol.Cancer Res.,
2003. 1(9): p. 674-681.
68. Wold, M.S., Replication protein A: a heterotrimeric, single-stranded DNA-
binding protein required for eukaryotic DNA metabolism. Annu.Rev.Biochem.,
1997. 66: p. 61-92.
69. Brill, S.J. and B. Stillman, Yeast replication factor-A functions in the unwinding
of the SV40 origin of DNA replication. Nature, 1989. 342(6245): p. 92-95.
70. Kim, C., R.O. Snyder, and M.S. Wold, Binding properties of replication protein A
from human and yeast cells. Mol.Cell Biol, 1992. 12(7): p. 3050-3059.
71. Wold, M.S., et al., Identification of cellular proteins required for simian virus 40
DNA replication. Journal of Biological Chemistry, 1989. 264(5): p. 2801-2809.
72. Georgaki, A. and U. Hubscher, DNA unwinding by replication protein A is a
property of the 70 kDa subunit and is facilitated by phosphorylation of the 32 kDa
subunit. Nucleic Acids Res., 1993. 21(16): p. 3659-3665. 136
73. Treuner, K., U. Ramsperger, and R. Knippers, Replication protein A induces the
unwinding of long double- stranded DNA regions. J.Mol.Biol, 1996. 259(1): p.
104-112.
74. Georgaki, A., et al., DNA unwinding activity of replication protein A. FEBS Lett.,
1992. 308(3): p. 240-244.
75. Blackwell, L.J., J.A. Borowiec, and I.A. Masrangelo, Single-stranded-DNA
binding alters human replication protein A structure and facilitates interaction
with DNA-dependent protein kinase. Mol.Cell Biol, 1996. 16(9): p. 4798-4807.
76. Gomes, X.V., L.A. Henricksen, and M.S. Wold, Proteolytic mapping of human
replication protein A: evidence for multiple structural domains and a
conformational change upon interaction with single-stranded DNA.
Biochemistry, 1996. 35(17): p. 5586-5595.
77. Oakley, G.G., et al., UV-induced hyperphosphorylation of replication protein a
depends on DNA replication and expression of ATM protein. Mol.Biol.Cell, 2001.
12(5): p. 1199-1213.
78. Dutta, A. and B. Stillman, cdc2 family kinases phosphorylate a human cell DNA
replication factor, RPA, and activate DNA replication. EMBO J., 1992. 11(6): p.
2189-2199.
79. Din, S., et al., Cell-cycle-regulated phosphorylation of DNA replication factor A
from human and yeast cells. Genes Dev., 1990. 4(6): p. 968-977.
80. Liu, V.F. and D.T. Weaver, The ionizing radiation-induced replication protein A
phosphorylation response differs between ataxia telangiectasia and normal
human cells. Mol.Cell Biol, 1993. 13(12): p. 7222-7231. 137
81. Shao, R.G., et al., Replication-mediated DNA damage by camptothecin induces
phosphorylation of RPA by DNA-dependent protein kinase and dissociates
RPA:DNA-PK complexes. EMBO J., 1999. 18(5): p. 1397-1406.
82. Wang, Y., et al., Roles of replication protein A and DNA-dependent protein
kinase in the regulation of DNA replication following DNA damage. J.Biol
Chem., 1999. 274(31): p. 22060-22064.
83. Liu, J.S., et al., Adozelesin triggers DNA damage response pathways and arrests
SV40 DNA replication through replication protein A inactivation. Journal of
Biological Chemistry, 2000. 275(2): p. 1391-1397.
84. Wang, H., et al., Replication protein A2 phosphorylation after DNA damage by
the coordinated action of ataxia telangiectasia-mutated and DNA-dependent
protein kinase. Cancer Res., 2001. 61(23): p. 8554-8563.
85. Binz, S.K., et al., The phosphorylation domain of the 32-kDa subunit of
replication protein A (RPA) modulates RPA-DNA interactions. Evidence for an
intersubunit interaction. J.Biol Chem., 2003. 278(37): p. 35584-35591.
86. Oakley, G.G., et al., RPA phosphorylation in mitosis alters DNA binding and
protein-protein interactions. Biochemistry, 2003. 42(11): p. 3255-3264.
87. Dart, D.A., et al., Recruitment of the Cell Cycle Checkpoint Kinase ATR to
Chromatin during S-phase. J.Biol Chem., 2004. 279(16): p. 16433-16440.
88. Zou, L. and S.J. Elledge, Sensing DNA damage through ATRIP recognition of
RPA-ssDNA complexes. Science, 2003. 300(5625): p. 1542-1548. 138
89. Zou, L., D. Liu, and S.J. Elledge, Replication protein A-mediated recruitment and
activation of Rad17 complexes. Proc.Natl.Acad.Sci.U.S.A, 2003. 100(24): p.
13827-13832.
90. Parrilla-Castellar, E.R. and L.M. Karnitz, Cut5 is required for the binding of Atr
and DNA polymerase alpha to genotoxin-damaged chromatin. J.Biol Chem.,
2003. 278(46): p. 45507-45511.
91. Hoeijmakers, J.H., Genome maintenance mechanisms for preventing cancer.
Nature, 2001. 411(6835): p. 366-374.
92. Digweed, M., Response to environmental carcinogens in DNA-repair-deficient
disorders. Toxicology, 2003. 193(1-2): p. 111-124.
93. Friedberg, E.C., DNA damage and repair. Nature, 2003. 421(6921): p. 436-440.
94. Xiao, Y. and D.T. Weaver, Conditional gene targeted deletion by Cre
recombinase demonstrates the requirement for the double-strand break repair
Mre11 protein in murine embryonic stem cells. Nucleic Acids Res., 1997. 25(15):
p. 2985-2991.
95. Luo, G., et al., Disruption of mRad50 causes embryonic stem cell lethality,
abnormal embryonic development, and sensitivity to ionizing radiation.
Proc.Natl.Acad.Sci.U.S.A, 1999. 96(13): p. 7376-7381.
96. Zhu, J., et al., Targeted disruption of the Nijmegen breakage syndrome gene
NBS1 leads to early embryonic lethality in mice. Curr.Biol, 2001. 11(2): p. 105-
109.
97. de Klein, A., et al., Targeted disruption of the cell-cycle checkpoint gene ATR
leads to early embryonic lethality in mice. Curr.Biol, 2000. 10(8): p. 479-482. 139
98. Ross, D.M. and T.P. Hughes, Cancer treatment with kinase inhibitors: what have
we learnt from imatinib? Br J Cancer, 2004. 90(1): p. 12-9.
99. Sawyers, C., Targeted cancer therapy. Nature, 2004. 432(7015): p. 294-7.
100. Rouse, J. and S.P. Jackson, Interfaces between the detection, signaling, and repair
of DNA damage. Science, 2002. 297(5581): p. 547-551.
101. Constantinou, A., et al., Werner's syndrome protein (WRN) migrates Holliday
junctions and co-localizes with RPA upon replication arrest. EMBO Rep., 2000.
1(1): p. 80-84.
102. Petrini, J.H., K.G. Huwiler, and D.T. Weaver, A wild-type DNA ligase I gene is
expressed in Bloom's syndrome cells. Proc.Natl.Acad.Sci.U.S.A, 1991. 88(17): p.
7615-7619.
103. Liu, J.S., et al., DNA damage by the enediyne C-1027 results in the inhibition of
DNA replication by loss of replication protein A function and activation of DNA-
dependent protein kinase. Biochemistry, 2001. 40(48): p. 14661-14668.
104. Carty, M.P., et al., UV light-induced DNA synthesis arrest in HeLa cells is
associated with changes in phosphorylation of human single- stranded DNA-
binding protein. EMBO J., 1994. 13(9): p. 2114-2123.
105. Zhao, S., W. Renthal, and E.Y. Lee, Functional analysis of FHA and BRCT
domains of NBS1 in chromatin association and DNA damage responses. Nucleic
Acids Res., 2002. 30(22): p. 4815-4822.
106. Sakamoto, S., et al., Werner helicase relocates into nuclear foci in response to
DNA damaging agents and co-localizes with RPA and Rad51. Genes Cells, 2001.
6(5): p. 421-430. 140
107. Golub, E.I., et al., Interaction of human rad51 recombination protein with single-
stranded DNA binding protein, RPA. Nucleic Acids Res., 1998. 26(23): p. 5388-
5393.
108. Vassin, V.M., M.S. Wold, and J.A. Borowiec, Replication protein A (RPA)
phosphorylation prevents RPA association with replication centers. Mol.Cell
Biol, 2004. 24(5): p. 1930-1943.
109. Kobayashi, J., et al., NBS1 localizes to gamma-H2AX foci through interaction
with the FHA/BRCT domain. Curr.Biol, 2002. 12(21): p. 1846-1851.
110. Paull, T.T., et al., A critical role for histone H2AX in recruitment of repair factors
to nuclear foci after DNA damage. Curr.Biol., 2000. 10(15): p. 886-895.
111. Ward, I.M. and J. Chen, Histone H2AX is phosphorylated in an ATR-dependent
manner in response to replicational stress. J.Biol Chem., 2001. 276(51): p.
47759-47762.
112. Shen, J. and L.A. Loeb, Unwinding the molecular basis of the Werner syndrome.
Mech.Ageing Dev., 2001. 122(9): p. 921-944.
113. Cheng, W.H., et al., Linkage between Werner syndrome protein and the Mre11
complex via Nbs1. J.Biol Chem., 2004. 279(20): p. 21169-21176.
114. Schroter, H., et al., DNA intercalators induce specific release of HMG 14, HMG
17 and other DNA-binding proteins from chicken erythrocyte chromatin. EMBO
J., 1985. 4(13B): p. 3867-3872.
115. Lai, J.S. and W. Herr, Ethidium bromide provides a simple tool for identifying
genuine DNA-independent protein associations. Proc.Natl.Acad.Sci.U.S.A, 1992.
89(15): p. 6958-6962. 141
116. Bork, P., et al., A superfamily of conserved domains in DNA damage-responsive
cell cycle checkpoint proteins. FASEB J., 1997. 11(1): p. 68-76.
117. Limoli, C.L., et al., Inaugural article: polymerase eta deficiency in the xeroderma
pigmentosum variant uncovers an overlap between the S phase checkpoint and
double-strand break repair. Proc.Natl.Acad.Sci.U.S.A, 2000. 97(14): p. 7939-
7946.
118. Canman, C.E. and D.S. Lim, The role of ATM in DNA damage responses and
cancer. Oncogene, 1998. 17(25): p. 3301-3308.
119. Maser, R.S., et al., Mre11 complex and DNA replication: linkage to E2F and sites
of DNA synthesis. Mol.Cell Biol, 2001. 21(17): p. 6006-6016.
120. Haber, J.E., The many interfaces of Mre11. Cell, 1998. 95(5): p. 583-586.
121. Robison, J.G., et al., Replication Protein A and the Mre11{middle
dot}Rad50{middle dot}Nbs1 Complex Co-localize and Interact at Sites of Stalled
Replication Forks. J.Biol Chem., 2004. 279(33): p. 34802-34810.
122. Buscemi, G., et al., Chk2 activation dependence on Nbs1 after DNA damage.
Mol.Cell Biol, 2001. 21(15): p. 5214-5222.
123. Kenny, M.K., et al., The role of human single-stranded DNA binding protein and
its individual subunits in simian virus 40 DNA replication. J.Biol Chem., 1990.
265(13): p. 7693-7700.
124. Adachi, Y. and U.K. Laemmli, Identification of nuclear pre-replication centers
poised for DNA synthesis in Xenopus egg extracts: immunolocalization study of
replication protein A. J.Cell Biol, 1992. 119(1): p. 1-15. 142
125. Cardoso, M.C., H. Leonhardt, and B. Nadal-Ginard, Reversal of terminal
differentiation and control of DNA replication: cyclin A and Cdk2 specifically
localize at subnuclear sites of DNA replication. Cell, 1993. 74(6): p. 979-992.
126. Brenot-Bosc, F., et al., Changes in the subcellular localization of replication
initiation proteins and cell cycle proteins during G1- to S-phase transition in
mammalian cells. Chromosoma, 1995. 103(8): p. 517-527.
127. Yan, H. and J. Newport, An analysis of the regulation of DNA synthesis by cdk2,
Cip1, and licensing factor. J.Cell Biol, 1995. 129(1): p. 1-15.
128. Alani, E., et al., Characterization of DNA-binding and strand-exchange
stimulation properties of y-RPA, a yeast single-strand-DNA-binding protein.
J.Mol.Biol, 1992. 227(1): p. 54-71.
129. Block, W.D., Y. Yu, and S.P. Lees-Miller, Phosphatidyl inositol 3-kinase-like
serine/threonine protein kinases (PIKKs) are required for DNA damage-induced
phosphorylation of the 32 kDa subunit of replication protein A at threonine 21.
Nucleic Acids Res., 2004. 32(3): p. 997-1005.
130. Binz, S.K., A.M. Sheehan, and M.S. Wold, Replication protein A phosphorylation
and the cellular response to DNA damage. DNA Repair (Amst), 2004. 3(8-9): p.
1015-24.
131. Sogo, J.M., M. Lopes, and M. Foiani, Fork reversal and ssDNA accumulation at
stalled replication forks owing to checkpoint defects. Science, 2002. 297(5581): p.
599-602. 143
132. Bianchi, V., E. Pontis, and P. Reichard, Changes of deoxyribonucleoside
triphosphate pools induced by hydroxyurea and their relation to DNA synthesis.
J.Biol Chem., 1986. 261(34): p. 16037-16042.
133. D'Arpa, P. and L.F. Liu, Topoisomerase-targeting antitumor drugs.
Biochim.Biophys.Acta, 1989. 989(2): p. 163-177.
134. Liu, L.F., et al., Mechanism of action of camptothecin. Ann N Y Acad Sci, 2000.
922: p. 1-10.
135. Cox, M.M., et al., The importance of repairing stalled replication forks. Nature,
2000. 404(6773): p. 37-41.
136. Lundin, C., et al., Different roles for nonhomologous end joining and homologous
recombination following replication arrest in mammalian cells. Mol.Cell Biol,
2002. 22(16): p. 5869-5878.
137. Furuta, T., et al., Phosphorylation of histone H2AX and activation of Mre11,
Rad50, and Nbs1 in response to replication-dependent DNA double-strand breaks
induced by mammalian DNA topoisomerase I cleavage complexes. J.Biol Chem.,
2003. 278(22): p. 20303-20312.
138. Falk, S.J. and P.J. Smith, DNA damaging and cell cycle effects of the
topoisomerase I poison camptothecin in irradiated human cells. Int.J.Radiat.Biol,
1992. 61(6): p. 749-757.
139. Hande, K.R., Topoisomerase II inhibitors. Cancer Chemother Biol Response
Modif, 2003. 21: p. 103-25. 144
140. Chen, G.L., et al., Nonintercalative antitumor drugs interfere with the breakage-
reunion reaction of mammalian DNA topoisomerase II. J.Biol Chem., 1984.
259(21): p. 13560-13566.
141. Sestili, P., F. Cattabeni, and O. Cantoni, Simultaneous determination of DNA
double strand breaks and DNA fragment size in cultured mammalian cells
exposed to hydrogen peroxide/histidine or etoposide with CHEF electrophoresis.
Carcinogenesis, 1995. 16(4): p. 703-706.
142. Tomasz, M. and Y. Palom, The mitomycin bioreductive antitumor agents: cross-
linking and alkylation of DNA as the molecular basis of their activity. Pharmacol
Ther, 1997. 76(1-3): p. 73-87.
143. Tomasz, M., Mitomycin C: small, fast and deadly (but very selective). Chem Biol,
1995. 2(9): p. 575-9.
144. McHugh, P.J., W.R. Sones, and J.A. Hartley, Repair of intermediate structures
produced at DNA interstrand cross-links in Saccharomyces cerevisiae. Mol Cell
Biol, 2000. 20(10): p. 3425-33.
145. De Silva, I.U., et al., Defects in interstrand cross-link uncoupling do not account
for the extreme sensitivity of ERCC1 and XPF cells to cisplatin
Repair of DNA interstrand crosslinks: molecular mechanisms and clinical relevance
Defining the roles of nucleotide excision repair and recombination in the repair of DNA
interstrand cross-links in mammalian cells
Repair of intermediate structures produced at DNA interstrand cross-links in
Saccharomyces cerevisiae. Nucleic Acids Res, 2002. 30(17): p. 3848-56. 145
146. McHugh, P.J., V.J. Spanswick, and J.A. Hartley, Repair of DNA interstrand
crosslinks: molecular mechanisms and clinical relevance. Lancet Oncol, 2001.
2(8): p. 483-90.
147. O'Neill, K.L., et al., Critical parameters influencing hyperthermia-induced
apoptosis in human lymphoid cell lines. Apoptosis., 1998. 3(5): p. 369-375.
148. Robison, J.G., et al., Decreased DNA repair rates and protection from heat
induced apoptosis mediated by electromagnetic field exposure.
Bioelectromagnetics, 2002. 23(2): p. 106-112.
149. Gomes, X.V. and M.S. Wold, Functional domains of the 70-kilodalton subunit of
human replication protein A. Biochemistry, 1996. 35(32): p. 10558-10568.
150. Kim, C. and M.S. Wold, Recombinant human replication protein A binds to
polynucleotides with low cooperativity. Biochemistry, 1995. 34(6): p. 2058-2064.
151. Blackwell, L.J. and J.A. Borowiec, Human replication protein A binds single-
stranded DNA in two distinct complexes. Mol.Cell Biol, 1994. 14(6): p. 3993-
4001.
152. Kim, C., B.F. Paulus, and M.S. Wold, Interactions of human replication protein A
with oligonucleotides. Biochemistry, 1994. 33(47): p. 14197-14206.
153. Lobachev, K., et al., Chromosome Fragmentation after Induction of a Double-
Strand Break Is an Active Process Prevented by the RMX Repair Complex. Curr
Biol, 2004. 14(23): p. 2107-12.
154. Sarasin, A. and A. Stary, Human cancer and DNA repair-deficient diseases.
Cancer Detect.Prev., 1997. 21(5): p. 406-411. 146
155. Miyagawa, K., Genetic instability and cancer. Int.J.Hematol., 1998. 67(1): p. 3-
14.
156. Taylor, A.M., Chromosome instability syndromes. Best.Pract.Res.Clin.Haematol.,
2001. 14(3): p. 631-644.
157. Robison, J.G., et al., DNA lesion-specific co-localization of the
MRE11/RAD50/NBS1 (MRN) complex and replication protein A (RPA) to repair
foci. J Biol Chem, 2005. 14: p. 14.
158. Dodson, G.E., Y. Shi, and R.S. Tibbetts, DNA Replication Defects, Spontaneous
DNA Damage, and ATM-dependent Checkpoint Activation in Replication Protein
A-deficient Cells. J.Biol Chem., 2004. 279(32): p. 34010-34014.
159. Pichierri, P., A. Franchitto, and F. Rosselli, BLM and the FANC proteins
collaborate in a common pathway in response to stalled replication forks. EMBO
J., 2004. 23(15): p. 3154-3163.
160. Pichierri, P. and F. Rosselli, The DNA crosslink-induced S-phase checkpoint
depends on ATR-CHK1 and ATR-NBS1-FANCD2 pathways. EMBO J., 2004.
23(5): p. 1178-1187.
161. Bakkenist, C.J. and M.B. Kastan, DNA damage activates ATM through
intermolecular autophosphorylation and dimer dissociation. Nature, 2003.
421(6922): p. 499-506.
162. Delia, D., et al., MRE11 mutations and impaired ATM-dependent responses in an
Italian family with Ataxia-Telangiectasia Like Disorder (ATLD). Hum.Mol.Genet,
2004. . 147
163. Yuan, S.S., et al., Neocarzinostatin induces Mre11 phosphorylation and focus
formation through an ATM- and NBS1-dependent mechanism. Toxicology, 2002.
177(2-3): p. 123-130.
164. Rhodes, N., T.M. Gilmer, and T.J. Lansing, Expression and purification of active
recombinant ATM protein from transiently transfected mammalian cells. Protein
Expr.Purif., 2001. 22(3): p. 462-466.
165. Sarkaria, J.N., et al., Inhibition of phosphoinositide 3-kinase related kinases by
the radiosensitizing agent wortmannin. Cancer Research, 1998. 58(19): p. 4375-
4382.
166. Ariza, R.R., et al., Reversible protein phosphorylation modulates nucleotide
excision repair of damaged DNA by human cell extracts. Nucleic Acids Res.,
1996. 24(3): p. 433-440.
167. Patrick, S.M. and J.J. Turchi, Replication protein A (RPA) binding to duplex
cisplatin-damaged DNA is mediated through the generation of single-stranded
DNA. Journal of Biological Chemistry, 1999. 274(21): p. 14972-14978.
168. Treuner, K., et al., Phosphorylation of replication protein A middle subunit
(RPA32) leads to a disassembly of the RPA heterotrimer. Journal of Biological
Chemistry, 1999. 274(22): p. 15556-15561.
169. Brush, G.S. and T.J. Kelly, Phosphorylation of the replication protein A large
subunit in the Saccharomyces cerevisiae checkpoint response. Nucleic Acids Res,
2000. 28(19): p. 3725-32. 148
170. Dmitrieva, N.I., D.V. Bulavin, and M.B. Burg, High NaCl causes Mre11 to leave
the nucleus, disrupting DNA damage signaling and repair. Am J Physiol Renal
Physiol, 2003. 285(2): p. F266-74. Epub 2003 Apr 08.
171. Dmitrieva, N.I., Q. Cai, and M.B. Burg, Cells adapted to high NaCl have many
DNA breaks and impaired DNA repair both in cell culture and in vivo. Proc Natl
Acad Sci U S A, 2004. 101(8): p. 2317-22.
172. Dmitrieva, N.I. and M.B. Burg, Living with DNA breaks is an everyday reality for
cells adapted to high NaCl. Cell Cycle, 2004. 3(5): p. 561-3. Epub 2004 May 19.