ROLE OF SAM68 IN DNA DAMAGE RESPONSES AND TUMORIGENESIS
by Xin Sun
A dissertation submitted to Johns Hopkins University in conformity with the requirements for the degree of Doctor of Philosophy
Baltimore, Maryland
Dec 2016
© 2016 Xin Sun All Rights Reserved
Abstract
The ability to recognize and repair DNA damage through rapid and appropriate DNA damage responses is pivotal to safeguard genomic information, which is persistently challenged by internal and environmental offenses. DNA lesion initiated poly(ADP- ribosyl)ation (PARylation), catalyzed primarily by poly(ADP-ribose) polymerase 1
(PARP1), is one of the earliest post-translational modifications to orchestrate downstream DNA damage response (DDR) signaling. However, the precise mechanisms through which PARP1 is activated and poly(ADP-ribose) (PAR) is robustly synthesized are not fully understood. Converging evidence support the emerging role of RNA- binding proteins (RBPs) in promoting DNA damage repair at different stages of DDR.
We discovered Src-associated substrate during mitosis of 68 kDa (Sam68) as a novel- signaling molecule in DDR. In the absence of Sam68, DNA damage-triggered
PARylation and PAR-dependent DNA repair signaling were dramatically diminished.
With serial cellular and biochemical assays, we revealed that Sam68 is recruited to and significantly overlaps with PARP1 at DNA lesions and that the interaction between
Sam68 and PARP1 is crucial for DNA damage-initiated and PARP1-conferred
PARylation. Utilizing cell lines and knockout mice, we demonstrated that Sam68- deleted cells and animals are hypersensitive to genotoxicity caused by DNA-damaging agents. In addition, loss of Sam68 delays basal cell carcinoma (BCC) onset and progression in a mouse model for basal cell carcinoma (BCC) of the skin, suggesting
Sam68 is required for BCC tumorigenesis, likely through promoting tumor cell survival.
Together, our findings demonstrate that Sam68 plays a crucial role in DDR via
ii regulating DNA damage-initiated PARylation, and that Sam68 is crucial in protecting normal cells from environmental genotoxic stress under physiological conditions and facilitating tumor cells to undergo malignant transformation in BCC.
Thesis Readers
Dr. Fengyi Wan
Dr. Pierre Coulombe
Dr. Alan Scott
Dr. Luis Garza
Dr. Michael Matunis
Dr. Jay Bream
Dr. Xuhang Li
Preface
My passion has always been applying new discoveries from the bench to the diagnosis and treatment of human tumors at bedside. There are six biological competencies acquired during the initiation and progression of human tumors, including ‘‘sustaining proliferative signaling, evading growth suppressors, resisting cell death, enabling replicative immortality, inducing angiogenesis, and activating invasion
iii and metastasis’‘, as summarized by Drs. Douglas Hanahan and Robert A. Weinberg in the prominent document ‘‘Hallmarks of Cancer: The Next Generation’‘ (Hanahan and
Weinberg 2011). Remarkably, alteration of genomic information underlies all six hallmarks of cancer to generate mutations that promote every aspect of tumorigenesis and progression.
With my thesis, I wanted to explore the underlying mechanisms of this multifaceted disease in relation to DNA damage responses (DDRs). I first examined the hypothesis that Src-associated substrate during mitosis of 68 kDa (Sam68), a previously known RNA binding protein (RBP), is a novel DNA damage-signaling molecule, governing poly(ADP-ribose) polymerase 1 (PARP1)-mediated poly(ADP-ribosyl)ation.
Having made the observation that genetic ablation of Sam68 decreases tumor burden in the mouse colon (Fu, Sun et al. 2016), I then assessed whether Sam68 is also involved in skin basal cell carcinoma (BCC) tumorigenesis.
In this dissertation, the main focus has been to investigate the role of Sam68 as a novel DNA damage response (DDR) protein. The resulting thesis is divided into six parts. Chapter 1 introduces the background information and our scientific questions. It first describes the concept of DDR, and then discusses the essential role of PARP1 and
PARylation during DDR and the importance in its tight regulation. Lastly it introduces
Sam68, our protein of interest, and our hypothesis that Sam68 may be a previously unappreciated DDR protein regulating PARylation. Chapter 2 reports on the approaches and observations in addressing our scientific questions. Chapter 3 discusses conclusions
iv we drew from the study, the significance of identifying Sam68 as a regulator of PARP1 catalytic activity, the emerging of RBPs in different phases of DDR and the uniqueness of Sam68 that distinguishes it from other RBPs in the context of DDR. In addition, I reviewed what are the burning questions that are left unanswered, and proposed how we can further uncover the mechanism through which Sam68 is activated and the mechanism through which Sam68 regulate PARP1. Chapter 4 summarizes the unpublished data about the potential role of Sam68 in BCC as an appendix to further discuss the involvement of Sam68 in tumorigenesis. This dissertation represent an expanded and updated version of the recent by Sun, X., Fu, K., Hodgson, A., Wier, E. M.,
Wen, M. G., Kamenyeva, O., Xia, X., Koo, L.Y., and Wan, F., published in PLoS Biology in 2016. Sam68 Is Required for DNA Damage Responses via Regulating Poly(ADP- ribosyl)ation. PLoS Biol, 14(9), e1002543, with the addition of unpublished data. Figures
1.1, 1.3 and 1.5 were summaried from current literature and made by myself; Figures 1.2,
1.4 and 1.6 were adopted from the cited literature and remade by myself; Figures 2.1, 2.2,
2.3, 2.4, 2.5, 2.6B, 2.6D-E, 2.8, 2.11E-G, 2.13, 2.14, 2.15, 2.16, 2.18, 3.1, 4.1D-E and 4.3 were generated entirely by my own data; Figure 2.6A, 2.6C-D, 2.7, 2.11B, 2.11H-I, 2.12, 4.1A-B were generated entirely by Dr. Kai Fu; Figures 2.11A and 2.11C were generated in collaboration with Dr. Kai Fu; Figures 2.6F-G, 2.9, and 2.10 were generated by myself with help from Drs. Lily Y. Koo and Olena Kamenyeva; Figues 4.1C and 4.2 were generated by myself with help from Dr. Kai Fu, Mr. Matthew Wen and Ms. Xin Guo;
Figure 3.3 was summarized from two of our published manuscripts and made by myself
(Fu, Sun et al. 2016, Sun, Fu et al. 2016).
v Acknowledgement
Graduate school has been a long voyage with many obstacles along the way. But it is not a lonely journey. With the warm company and generous support from my amazing mentors, colleagues, friends and family, and with a determined mind, my thesis has become a reality, and I have been rewarded. It is all the laughers and tears that allow me to uncover my potential and keep moving forward in science and in life.
Foremost, I would like to express my gratefulness to the great set of mentors that
I have. I cannot thank my advisor Dr. Fengyi Wan enough for taking me into his lab and granting a first year graduate student plentiful trust to open a new field in the lab. He provided me not only the techniques, but also the intellect to tackle the scientific questions. He always gave me an opportunity to describe my plan, before offering suggestions and explaining how he would improve it. It is all the open discussions we had over the years that fostered my independent scientific thinking. Fengyi’s passion about science has driven everyone in the lab to always try our best. I feel lucky to have such an incredible advisor, and I appreciate his tremendous guidance and support during my graduate career. I also would like to thank members of my thesis committee,
Drs. Pierre Coulombe, Michael Matunis, Alan Scott, and Luis Garza, who have encouraged, challenged and guided me to be faithful to science and myself. I want to thank Dr. Pierre Coulombe for mentoring me through our fruitful collaboration and our invaluable joint lab meetings. He showed me that a true scientist is one with broad knowledge, a talent who continuously ask insightful questions and a mentor who is
vi always approachable for advise and help. My special thanks also goes to Dr. Michael
Matunis. His great suggestions and substantial support through the whole duration of graduate school helped shape my thesis work and career. Conversations in his office or small talks in the hallway prepared me for my meetings and career development.
I want to thank our collaborators Drs. Stephane Richard from McGill University,
Johnny He from University of North Texas Health Science Center, Jeremy Stark from
Beckman Research Institute of City of Hope, John Pascal from University of Montreal,
Anthony Leung from Johns Hopkins University and Zhaoqi Wang from Fritz Lipmann
Institut for sharing experimental material; Dr. Cory Brayton and Ms. Xin Guo from
Johns Hopkins University for help with histological analyses; Drs. Owen Schwartz, Lily
Y. Koo and Olena Kamenyeva from National Institutes of Health for help with laser micro-irradiation microscopy. Your help significantly improved my thesis project.
Thank you to all my friends and colleagues inside and outside of school. I want to thank all the previous and current members of the Wan Lab, Dr. Kai Fu, Dr. Andrea
Hodgson, Dr. Eric Wier, Xue (Summer) Xia, Yue (Harry) Liu, Matthew Wen, Dr. Wenxin
Zheng and Jackline Lasola. You build such an incredible positive and collaborative working environment that I learned a lot from each of you. My graduate school would not be so rewarding without your friendship and support. I especially want to thank Kai for our fruitful and enjoyable collaboration on so many projects over the years. Kai is so knowledgable with hands on bench techniques that he is the MASTER of western blotting and immunoprecipitation in our lab. He is always willing to lend a helping
vii hand. His contribution to my thesis project significantly enhanced our understanding of the molecular mechasim. And thank you Andrea, for your generous personality, providing me rides from the airport to home after my experiments at NIH, and keeping me company at 12am in the emergency room. I also want to thank Eric for great advices in improving my scientific writing. In addition, I want to thank Dr. Tao Zhang, Summer,
Harry, my classmates Elizabeth Alexander, Fengrong Wan, Chynna Broxton and Nicole
Parker and my friends outside of school for their comforting and reassuring friendship and support. Thank you for accompanying me in the library late at night and during weekend, studying for exams and writing thesis, for creating so many game nights, potluck nights, baking nights and outings to keep me positive and recharge me.
I would also like to extend my sincere thanks to my family. To my father, who is so passionate about life, who continues to learn and grow, and who has always been a source of inspiration to me throughout my life. You taught me to focus on the big picture and not be afraid of temporal setback. To my mother, who is so caring for me, who respects and supports every decision I make, and who I know is always there waiting for me with her warm arms no mater what happens. To my parents-in-law, who lent me great help when I struggled in getting balance between work and life. To my grand mother and my entire family back in China, who support me along the way.
Last but not least, I want to thank my dear husband Dr. Guofan Hu and precious baby Jeremy Jialin Hu, who give me a loving home in a foreign land. Thank you Guofan.
I am truly grateful for your unconditional love. Knowing you since the age of 15 in high
viii school, together we added the roles of husband and wife and then father and mother.
Your unwavering support carried me through rough patches in life. I would not be who
I am today without you looking at me with a smile in your eyes. Thank you Jialin. You have been a constant source of bliss and joy through the battle of publishing my thesis work. And you are the reason I want to stay strong and become a better self. I cannot wait to enjoy a life seeing you two everyday and build our future together!
ix Table of Contents
1 Introduction ...... 1
1.1 The Prompt and Appropriate DNA Damage Responses (DDRs) are Essential
for the Cellular DNA Damage Repair and Survival...... 1
1.2 DNA lesion-initiated, PARP1-conferred PARylation is one of the earliest post-
translational modifications to orchestrate downstream DNA damage response
signaling...... 6
1.3 Sam68 is a KH Domain-containing RNA Binding Protein with Versatile
Cellular Functions...... 10
1.4 Figures and Figure Legends ...... 15
2 Sam68 is Required for DNA Damage Responses via Regulating Poly(ADP- ribosyl)ation...... 22
2.1 Sam68 Plays a Critical Role in Repairing DNA Strand Breaks...... 22
2.2 Sam68 Deficiency Impairs DNA Damage-initiated PARylation and Signaling
Cascade...... 24
2.3 DNA Damage Enhances Interaction Between Sam68 and PARP1...... 25
2.4 Sam68 is Recruited to and Regulate PARylation at DNA Damage Sites...... 26
2.5 Sam68 is Required for PARP1 Catalytic Activation and Its Release from DNA
Damage Sites...... 28
2.6 Sam68 stimulates PARP1 Catalytic Activity via the Interation Between their N-
termini...... 30
x 2.7 PARP1 Deficiency/Inhibition and Sam68 Deficiency Share Similar Effects on
DNA Repair...... 33
2.8 Sam68 Deletion Attentuates PARylation and PAR-dependent Repair Signaling
in Radiodamaged Thymi...... 34
2.9 Sam68 Deficient Mice are Hypersensitive to Genotoxic Stresses...... 35
2.10 Figures and Figure Legends ...... 38
2.11 Materials and Methods ...... 71
2.12 Summary ...... 81
3 Conclusion and Future Perspective ...... 83
3.1 Sam68 is a Novel DNA Damage Repair Signaling Protein that Governs PARP1-
conferred PARylation...... 83
3.2 Sam68 is the Master Regulator of DDR via Stimulating PARP1 Catalytic
Activation...... 86
3.3 Sam68 Bridges Different Stages in the Process of Upregulating DNA Damage-
initiated Gene Expression...... 87
3.4 RBPs are Emerging as a New Group of Important Effectors in DDR...... 88
3.4.1 RBPs Ensure Precise DNA Replication to Safeguard Genomic Information...... 90
3.4.2 RBPs Coordinate Gene Expression as a Long-term Remedy to DNA Damage. .... 94
3.4.3 RBPs Play an Important and Direct Role in DNA Damage Repair...... 97
3.5 Sam68 May Serve as a Potential Therapeutic Target by PARP1 Inhibition. .. 101
3.6 Future Perspective ...... 102
3.6.1 Virtual Regulation Between Sam68 and ATM ...... 102
3.6.2 Structural Basis for the Binding of Sam68 to Damaged DNA and Its Interaction
with PARP1 ...... 103
3.7 Conclusion ...... 104
xi 3.8 Figures and Figure Legends ...... 106
4 Appendix: Sam68 is a Potential Therapeutic Target for Basal Cell
Carcinoma...... 108
4.1 Introduction ...... 108
4.2 Results ...... 109
4.2.1 Sam68 is Significantly Upregulated in Basal Cell Carcinoma...... 109
4.2.2 Loss of Sam68 Delays BCC Tumorigenesis...... 110
4.2.3 Sam68 is crucial for skin tumor cell survival and malignant
transformation...... 111
4.3 Figures and Figure Legends ...... 112
4.4 Discussion ...... 118
4.5 Material and Methods...... 118
5 References ...... 122
6 Curriculum Vitae ...... 137
xii Table of Figures
Figure 1.1 DNA lesions and repair pathways...... 15
Figure 1.2 Schematic diagram representing the DNA damage responses in cells...... 16
Figure 1.3 The coordination of the homologous recombination (HR) pathway and cell-cycle transition signaling cascade in response to DNA double-stranded breaks (DSBs)...... 17
Figure 1.4 The structural and functional domains of human PARP1...... 19
Figure 1.5 PARP1 interacts with multiple HR and NHEJ proteins and regulates their recruitment and activity through PARylation...... 20
Figure 1.6 The structural and functional domains of Sam68...... 21
Figure 2.1 Sam68 is required for repairing DNA strand breaks...... 38
Figure 2.2 Sam68 is required for clonogenic survival of mouse embryonic fibroblasts following exposure to H2O2...... 41
Figure 2.3 Sam68 is critical for repairing DNA damage in U2OS reporter cell lines...... 42
Figure 2.4 Sam68 deficiency attentuates DNA damage-initiated repair signaling in cell culture...... 43
Figure 2.5 Sam68 deficiency attenuates DNA repair signaling in cell culture...... 45
Figure 2.6 Sam68 is recruited to and regulates PARylation at DNA damage sites...... 46
Figure 2.7 Sam68 interacts with early DNA damage sigaling molecules PARP1 and ATM...... 49
Figure 2.8 Chromatin fractionation assays showing dynamics of Sam68 on damaged chromatin...... 50
Figure 2.9 Sam68 localizes to sites of DNA damage in DDR...... 51
Figure 2.10 Sam68 regulates PARP1 retention at but not PARP1 recruitment to DNA damage sites...... 53
Figure 2.11 Sam68 enhances the damaged DNA-dependent PARP1 activation and PARylation in vitro. 55
Figure 2.12 The DNA-dependent PARP1-mediated PARylation in vitro...... 58
Figure 2.13 PARP1 inhibition attenuates DNA damage-induced repair signaling in thymocytes...... 60
Figure 2.14 PARP1 is critical for repairing DNA strand breaks...... 61
Figure 2.15 PARP1 knockdown does not impact the sensitivity of Sam68 deleted cells in response to DNA damage...... 64
xiii Figure 2.16 Sam68 deletion dampens DNA repair signaling in the radiodamaged thymus...... 65
Figure 2.17 Sam68 KO mice are hypersensitive to genotoxic stresses...... 67
Figure 2.18 Sam68 deficiency does not affect thymus and small intestine development in mice, but causes more severe damage post γ-irradiation...... 69
Figure 3.1 Schematic model representation of Sam68 functioning as a signaling molecule in DNA damage repair...... 106
Figure 3.2 Schematic model representation of Sam68 functioning as the master regulator of DDR...... 107
Figure 4.1 Sam68 is upregulated in basal cell carcinoma...... 112
Figure 4.2 Loss of Sam68 delays basal cell carcinoma tumorigenesis...... 114
Figure 4.3 Sam68 is required for skin cancer cell survival and malignant transformation...... 116
xiv 1 Introduction
1.1 The Prompt and Appropriate DNA Damage Responses (DDRs) are Essential for
the Cellular DNA Damage Repair and Survival.
Genome integrity is constantly threatened by both internal and external insults.
Intrinsic deamination or depurination on a DNA single strand happens at a rate of
~10,000 events per cell/per day, respectively generating significant inter-substitutions and loss of nucleotide bases on DNA (Bauer, Corbett et al. 2015). Extrinsic reactive oxygen species (ROS) and carcinogens originating from cellular metabolism, chemical agents used in cancer chemotherapy, and ultraviolet (UV) light B (UVB) can confer DNA damage to a single strand (Demple and Harrison 1994, Ciccia and Elledge 2010, Budden and Bowden 2013). Other extrinsic damage, such as γ-irradiation, UVA, UVC, and bifunctional alkylating agents that generate interstrand cross-links (ICLs) will lead to damage to both strands of chromatin (Sinha and Hader 2002, Ciccia and Elledge 2010,
Budden and Bowden 2013, Bauer, Corbett et al. 2015) (Fig 1.1).
Cells are endowed with an elaborate surveillance network, termed DNA damage responses (DDRs) to cope with damages. During DDR, cells sense the presence of DNA damage and transduce damage signaling to the recruit effector proteins. Immediate repair of the damage through the appropriate pathway (Fig 1.1), accurate cell cycle control, precise transcriptional/post-transcriptional regulation of gene expression and
1 correct cell fate choice are essential to safeguard their genetic information (Zhou and
Elledge 2000) (Fig 1.2).
Intrinsic DNA damage including deamination or depurination of the DNA bases and alkylating agent-induced small lesions on DNA single strands are repaired by base excision repair (BER) (Liu, Prasad et al. 2007). The chemical nature of the lesions determines whether short-patch BER or long-patch BER will be utilized. These two pathways share similar initial steps. DNA glycosylases recognize specific damage and cleave the N-glycosidic bond, generating abasic sites (AP sites). Apurinic endonuclease 1
(APE1) cleaves the phosphodiester bond of these AP sites, generating a 3’-OH and a 5’- phosphorylated sugar. During short-patch BER, polymerase β (pol β) cleaves the sugar residue by β-elimination and puts in the correct nucleotide using the undamaged strand as a template. Finally, the nick remaining in the damaged DNA strand is rejoined by
DNA ligase III. In long-patch BER, pol β, together with replication factor C (RFC) and proliferating cell nuclear antigen (PCNA), produce a long patch of nucleotides by replicating the undamaged strand as the template. Flap structure-specific endonuclease
1 (FEN1) then removes the old strand. Finally, DNA ligase I rejoins the damaged DNA strand (Fortini and Dogliotti 2007).
“Bulky” lesions such as pyrimidine dimers and large chemical adducts on a
DNA single strand are repaired by nucleotide excision repair (NER) (Le May, Egly et al.
2010). There are two NER subpathways, global genome NER (GGNER) and transcription-coupled NER (TC-NER). During GGNER, the XPC complex subunit, DNA
2 damage recognition and repair factor (XPC)-hHR23B complex detects DNA lesions, which are then opened by TFIIH-associated helicases ERCC excision repair 3, TFIIH core complex helicase subunit (ERCC3/XPB) and ERCC excision repair 2, TFIIH core complex helicase subunit (ERCC2/XPD). After RPA binds the undamaged strand, endonuclease
ERCC excision repair 4, endonuclease catalytic subunit (ERCC4/XPF) and ERCC excision repair 5, endonuclease (ERCC5/XPG) make incisions on both sides of the lesion on the
DNA strand. The remaining gap is filled by pol δ/ε, PCNA and RFC, using the undamaged strand as the template. Finally, DNA ligase I rejoins the nick. During TC-
NER, RNA Polymerase II initiates repair upon getting stalled at DNA lesions (Le May,
Egly et al. 2010, Budden and Bowden 2013).
A DNA double strand break (DSB) is one of the most deleterious forms of DNA damage, caused by a variety of intrinsic and extrinsic factors (Ciccia and Elledge 2010).
Non-homologous end joining (NHEJ) and homologous recombination (HR) are the two major pathways to repair DSBs (Jackson and Bartek 2009, Symington and Gautier 2011).
NHEJ is an error-prone repair pathway, which does not require the use of a sister chromatid as a template. Instead, incompatible DNA DSB ends, with little or no terminal microhomology, are recognized by poly(ADP-ribose) polymerase 1 (PARP1) and the
Ku70/Ku80 heterodimers. The resulting recruitment and activation of DNA-dependent protein kinase, catalytic subunit (DNA-PKcs) promotes the Artemis/DNA-PKcs complex to cleave a single-strand tail, or facilitate Pol μ to add extra nucleotides to the DNA ends.
Then the DNA ligase IV/ X-ray repairs cross-complementing protein (XRCC)/XRCC4-
3 like factor (XLF) complex promotes direct ligation of the DNA ends (Jackson and Bartek
2009). Hence, NHEJ can drive DSB repair during any phase of the cell cycle. On the contrary, HR only occurs during late S phase or G2 phase of the cell cycle, when the sister chromatids are present. This allows faithful restoration of the broken DNA sequence from the replication of the template within its sister chromatid. During HR, the presence of DSBs activates PARP1 (Tartier, Spenlehauer et al. 2003), which promotes Mre11-
Rad50-Nbs1 (MRN) complex recruitment (Haince, McDonald et al. 2008), leading to activation of kinase ataxia telangiectasia mutated (ATM) and the phosphorylation of its substrates histone 2AX (H2AX), checkpoint kinase 1 (CHK1) and checkpoint kinase 2
(CHK2). Phosphorylated CHK1 and CHK2 depart the proximity of DNA DSB sites to orchestrate the cell cycle through regulating CDKs and cyclins (Kastan and Bartek 2004).
Phosphorylated H2AX, termed γH2AX, marks DSB sites and serves as the scaffold to recruit mediator of DNA damage checkpoint 1 (MDC1). MDC1 phosphorylation by
ATM switches phosphorylation signals to ubiquitylation/SUMOylation signals by recruitment of RING-finger ubiquitin ligase ring finger protein 8 (RNF8)/ ubiquitin conjugating enzyme 13 (Ubc13) and ring finger protein 168 (RNF168)/Ubc13, as well as
PIAS1/4 E3 ligase, to ubiquitylate histone 2A (H2A) and H2AX surrounding the break.
Receptor-associated protein 80 (RAP80) binds to the ubiquitylated scaffold, which promotes recruitment of breast cancer 1 (BRCA1), downstream effector protein breast cancer 2 (BRCA2) and RAD51 to conduct synthesis-dependent strand annealing of DSBs
(Bergink and Jentsch 2009, Ciccia and Elledge 2010) (Fig 1.3).
4 Interstrand DNA crosslinks (ICLs) are lesions that covalently link the Waston-
Crick strands of DNA (Hashimoto, Anai et al. 2016), making them more difficult to repair than other lesions. ICLs are mostly detected by DNA polymerase during DNA replication, or by RNA polymerase during transcription. It has been shown that multiple
DNA repair pathways are involved in repairing ICLs, including factors of HR, NER and translesion DNA synthesis (TLS), all of which are integrated and categorized as the
Fanconi anemia (FA) pathway (Moldovan and D'Andrea 2009, Hashimoto, Anai et al.
2016). A family of proteins, including FANCA, FANCB, FANCC, FANCE, FANCF,
FANCG, FANCL and FANC, are the building blocks of the FA core complex. The repair of ICLs involves the initial incision of the lesions by NER factors and then repair of the incision by HR and/or NER factors (Moldovan and D'Andrea 2009).
The complex DNA repair network is not only capable of repairing DNA damage, but is also essential in maintaining telomere homeostasis to prevent ageing, generating receptor diversity through V(D)J recombination, causing class-switching recombination and somatic hyper-mutation in immune responses, and creating genetic diversity through meiotic sister chromatin exchange during gametogenesis (Jackson and Bartek
2009). Failure to engage appropriate DDR can lead to loss of genomic integrity in cells, and further cause premature aging, heritable diseases, neurodegenerative diseases, immune deficiency, infertility, metabolic syndrome, and cancer (Ciccia and Elledge
2010). Therefore, it is important to understand the regulatory mechanisms of DDR in more detail.
5 1.2 DNA lesion-initiated, PARP1-conferred PARylation is one of the earliest post-
translational modifications to orchestrate downstream DNA damage response
signaling.
PARP1 has been recognized as an indispensable sensor of different types of DNA damage (Schreiber, Dantzer et al. 2006). As the founding member of PARP superfamily,
PARP1 is composed of six domains (Fig 1.4). Among them, the two zinc-binding domains, Zn1 and Zn2, enable PARP1 to recognize and bind to damaged DNA
(Eustermann, Videler et al. 2011, Langelier, Planck et al. 2011). The Zn3 domain is not essential for DNA binding. Instead, together with the tryptophan-, glycine-, arginine- rich (WGR) domain, it bridges the conformational change in the DNA binding domain
(DBD) upon binding of damaged DNA to the catalytic domain (CAT), which triggers the decrease in the thermal stability of the CAT helical subdomain (HD) (Langelier, Planck et al. 2012). The auto-modification domain (AD) possesses a BRCA1 carboxyl-terminal
(BRCT) motif responsible for protein-protein interactions during DDR, and the majority of auto-modification residues (glutamate and lysine) act as acceptors of ADP-ribose moieties (Rouleau, Patel et al. 2010). Finally, the C-terminal CAT domain sequentially adds one or more (up to a few hundred) negatively charged ADP-ribose moieties from nicotinamide adenine dinucleotide (NAD+) to target proteins, including PARP1 itself, histones, chromatin regulators, DNA repair proteins and transcription factors
(Jungmichel, Rosenthal et al. 2013, Zhang, Wang et al. 2013).
6 Remarkably, PARP1 has been shown to be recruited to DNA damage sites within milliseconds following laser micro-irradiation in the nucleus, and is one of the earliest events in DDR (Tartier, Spenlehauer et al. 2003). Binding to DNA strand breaks results in the activation of the C-terminal CAT of PARP1, which dramatically elevates its catalytic activity (D'Amours, Desnoyers et al. 1999, Langelier, Planck et al. 2012). The immediate activation of PARP1 by DNA breaks and the resulting poly(ADP- ribosyl)ation (abbreviated as PARylation) serve as the first defense during DNA damage recognition and repair (Schreiber, Dantzer et al. 2006). First, PARylation labels the damaged sites to signify the occurrence as well as the magnitude of DNA damage.
Second, PARylation of the core histones amino-terminal tails favors relaxation of local chromatin structure around DNA damage sites (Poirier, de Murcia et al. 1982, Messner,
Altmeyer et al. 2010). Third, the branched and elongated poly(ADP-ribose) (PAR) chains function as a docking scaffold for the rapid recruitment of DNA repair effector proteins.
Equally important, localized accumulation of highly negatively charged PAR chains on
PARP1 itself also initiates its departure from the damaged sites promptly after fulfilling its role, acting as a feedback regulation (Mortusewicz, Ame et al. 2007).
Mounting evidence suggests that PARP1 and its function in PARylation are indispensable for multiple DNA repair pathways, including BER, single strand break
(SSB) repair, mismatch repair, HR, and NHEJ (Masson, Niedergang et al. 1998, Leppard,
Dong et al. 2003, Okano, Lan et al. 2003, Audebert, Salles et al. 2006, Aguilar-Quesada,
Munoz-Gamez et al. 2007, Haince, Kozlov et al. 2007, Haince, McDonald et al. 2008,
7 Couto, Wang et al. 2011, Spagnolo, Barbeau et al. 2012, Ying, Chen et al. 2016). PARP1 has been shown to either directly interact with multiple essential players from the aforementioned pathways or regulate their recruitment to and activation at sites of DNA damage through PARylation (Fig 1.5). The critical roles of PARP1 in DNA damage repair have been illustrated by a great deal of studies from genetic (especially three independent PARP1 deficient mouse models), pharmacological, and epidemiological studies (Durkacz, Omidiji et al. 1980, Jameson, Frey et al. 1980, de Murcia, Niedergang et al. 1997, Wang, Stingl et al. 1997, Masutani, Nozaki et al. 1999). Loss of PARP1 activity, due to genetic ablation or chemical inhibition, consequently delays DNA repair, which gives rise to hypersensitivity to both alkylating agents and γ-irradiation at both cellular and animal levels. Moreover, PARP1 inhibition has arisen as a promising novel therapeutic approach to treat human cancers and inflammatory diseases that are associated with abnormal DNA repair activities (Anders et al., 2010; Papeo et al., 2009).
One of the PARP1 inhibitors, Olaparib, has recently been approved by FDA for treating certain types of ovarian cancer (Oza, Cibula et al. 2015).
Despite the extensive knowledge about the importance of PARP1 in DNA damage repair pathways, the precise mechanisms of stimulation and regulation of
PARP1 catalytic activity remain elusive. The biggest challenge in studying the mechanism of PARP1 activation is that the rapid and highly dynamic nature of PARP1 recruitment to DNA damage sites and catalytic activation makes it difficult to zoom in to the sites of DNA damage within an extremely narrow time window. Nevertheless,
8 PARP1 activation needs to be fine-tuned upon DNA damage for cells to respond properly. The amount of PAR signals the presence and severity of DNA damage.
Hyperactivated PARP1 is on the other hand hazardous to cells, when the intracellular pools of NAD+ are drained and ATP production is disrupted (Eguchi, Shimizu et al.
1997, Leist, Single et al. 1997, Ha and Snyder 1999, Herceg and Wang 2001).
Recent studies demonstrate that post-translational modifications and interactions with other proteins at DNA damage foci are essential for fine-tuning PARP1 function during DDR. Some key findings include that PARP1 is required to be phosphorylated by mitogen-activated protein kinase (MAPK) and extracellular-signal-regulated kinase
(ERK) for its maximal activation during DNA damage repair (Kauppinen, Chan et al.
2006); that high mobility group N1 (HMGN1) interacts with PARP1 in untreated mouse fibroblast cells and it can boost PARP1 catalytic activity in vitro (Masaoka, Gassman et al.
2012); that protein inhibitor of activated STAT protein y (PIASy)-mediated SUMOylation of PARP1 facilitates DNA damage-induced NF-κB activation (Stilmann, Hinz et al.
2009); and that the deacetylase Sirtuin 6 (SIRT6), through mono-ADP-ribosylation, stimulates PARP1 activity in DNA damage repair under oxidative stress (Mao, Hine et al. 2011). These studies provide novel insight into, and also suggest a very complicated mechanism(s) regarding the functional regulation of PARP1 in the cellular response to
DNA damage. Hence, it is imperative to identify the regulators of PAPR1 prior to, during, and after its binding to DNA strand breaks, allowing one to comprehend the complexity in regulatory network of PAPR1 function.
9 1.3 Sam68 is a KH Domain-containing RNA Binding Protein with Versatile Cellular
Functions.
The Src-associated substrate during mitosis of 68 kDa (Sam68) belongs to the heteronulear ribonuleoprotein particle K (hnRNP K) homology (KH) domain family of
RNA-binding proteins (RBPs). In 1990, it was originally discovered as p62, a tyrosine phosphoprotein (Moran, Koch et al. 1990). It was not until 1994 that p62 was identified as a substrate of Src during mitosis and noticed to actually migrate at 68 kDa (Fumagalli,
Totty et al. 1994, Taylor and Shalloway 1994). Therefore, p62 was designated as Sam68 from that point forward. In 2002, a new name of Sam68 was released by the Human
Genome Organization gene nomenclature committee as KH domain containing, RNA binding, signal transduction associated 1 (KHDRBS1). However, because most of the past and current literatures still use the name Sam68, I will follow the name Sam68 to avoid confusion.
The single heteronulear ribonuleoprotein particle K (hnRNP K) homology (KH) domain, together with its flanking and highly conserved N-terminal of KH (NK) and C- terminal of KH (CK), mediates Sam68’s binding to RNA (Wong, Muller et al. 1992). In addition, Sam68 harbors six proline-rich motifs that are potential binding sites for SH3 and WW domain-containing proteins (Kay, Williamson et al. 2000). Moreover, the C- terminus of Sam68 possesses several tyrosine residues that have been shown to be phosphorylated by tyrosine kinases (Fumagalli, Totty et al. 1994, Taylor and Shalloway
1994, Lang, Mege et al. 1997, Derry, Richard et al. 2000), and a nuclear localization signal
10 (NLS) that determines the predominant nuclear localization of Sam68 in most cell types.
(Fig 1.6)
Mounting evidence indicates that Sam68 has versatile functions in a variety of cellular processes (Lukong and Richard 2003). Sam68 was revealed to play an important role in cell cycle regulation, promoting S phase entry through its KH domain (Richard
2010). Transient expression of a spliced isoform of Sam68 that is naturally expressed in quiescent cells inhibits entry of NIH3T3 cells to the S phase due to decreased cyclin D1 expression (Barlat, Maurier et al. 1997). In addition, Sam68-deficient chicken B cell line
DT40 exhibit a slow growth phenotype because of an elongated G2-M phase (Li, Haga et al. 2002). Sam68 was also discovered as a regulator of alternative splicing in response to extracellular signals. It binds the exonic splice-regulatory elements of the CD44 exon V5 and promotes the inclusion of that exon (Matter, Herrlich et al. 2002). Sam68 also binds to intron 4 in the cyclin D1 gene to mediate the recruitment of the spliceosomal component U1-70K during splicing (Paronetto, Cappellari et al. 2010). Additionally,
Sam68 binds to intronic splice elements within intron 5 of the mTOR gene to regulate its alternative splicing during adipogenesis (Huot, Vogel et al. 2012). Furthermore, Sam68 mediates the inclusion of the human papillomavirus (HPV) 16 E6 exon during splicing in response to epidermal growth factor (EGF) signaling pathway activation
(Rosenberger, De-Castro Arce et al. 2010). A comprehensive exploration of Sam68 pre- mRNA targets in mouse neuroblastoma identified twenty-four novel exons regulated by
Sam68, most of which are known mediators in neurogenesis (Paronetto, Achsel et al.
11 2007). Sam68 also has other functions, such as participating in the exportation of retroviral RNAs that possess a constitutive transport element (CTE) from the nucleus to the cytoplasm (Coyle, Guzik et al. 2003). Most of these functions have been attributed to its RNA binding property (Lukong and Richard 2003, Richard 2010).
Sam68 has been long acknowledged as an almost strictly nuclear protein with an identified NLS (Ishidate, Yoshihara et al. 1997). However, the potential function of
Sam68 in the nuclear-initiated signaling pathways has not been intensively investigated.
Recently, an increasing amount of evidence supports the role of Sam68 in signaling transduction and gene transcription, independent of its RNA binding ability (Richard
2010, Adamson, Smogorzewska et al. 2012). Sam68 is capable of serving as an adaptor to transmit signal from activated membrane-associated receptors. Sam68 has been shown to participate in the T cell receptor (TCR)-coupled signaling pathway through binding to
Src family kinases FYN proto-oncogene, Src family tyrosine kinase (Fyn) and LCK proto- oncogene, Src family tyrosine kinase (Lck) via their SH2 and SH3 domains (Fusaki,
Iwamatsu et al. 1997). Sam68 also bridges signaling from the leptin receptor insulin receptor substrate 1 (IRS-1) to phosphatidylinositol 3-kinase (PI3K) activation (Martin-
Romero and Sanchez-Margalet 2001). Moreover, Sam68 connects signaling pathways to gene transcription, which is in line with the findings that Sam68 is predominantly a nuclear protein and binds to both single-stranded and double-stranded DNA (Wong,
Muller et al. 1992, Bielli, Busa et al. 2011, Frisone, Pradella et al. 2015). In particular, we previously showed that Sam68 is a non-Rel component of the nuclear factor-kappa B
12 (NF-κB) complex and is required for NF-κB-dependent CD25 transcription in T cells (Fu,
Sun et al. 2013). Sam68 also binds to the CH3 domain of transcriptional cofactor CREB- binding protein (CBP) and represses CBP-dependent transcription (Hong, Resnick et al.
2002). Sam68 interacts with hnRNP K and inhibits hnRNP K-mediated transcriptional activation of the CT-promoter (Yang, Reddy et al. 2002). Sam68 is also able to form transcriptional complex with mixed lineage leukemia (MLL) and protein arginine methyltransferase 1 (PRMT1) to promote hematopoietic stem cells self-renewal (Cheung,
Chan et al. 2007). These findings indicate that Sam68 may possess more previously unappreciated functions in the nucleus.
Converging evidence suggests that Sam68 could play an important role in DNA damage-initiated signaling in the nucleus. First, Sam68 was shown to bind single- stranded and double-stranded DNA (Wong, Muller et al. 1992), which provides the basis for Sam68 to recognize and bind to damaged DNA. Secondly, Sam68 bears a consensus motif recognized by ATM, ATM and Rad3-related (ATR), and DNA-dependent protein kinase (DNA-PK), thus being proposed as a putative substrate of ATM/ATR/DNA-PK.
Indeed, up-regulated phosphorylation sites and acetylation sites have been identified in
Sam68 in response to DNA damage in a recent mass spectrometry study (Beli,
Lukashchuk et al. 2012), and Sam68 phosphorylation post γ-irradiation treatment has been shown to be ATM dependent from another proteomic study (Bensimon, Schmidt et al. 2010). This evidence indicates that Sam68 may possess certain functions that need to be regulated in an ATM-dependent and/or acetylation-controlled manner during DDR.
13 Moreover, Sam68 was observed among the PAR-associated proteins from two independent proteomic studies (Gagne, Isabelle et al. 2008, Jungmichel, Rosenthal et al.
2013), which further supports the notion that Sam68 may participate in the nuclear- initiated signaling to repair DNA damage.
We therefore examined the hypothesis that Sam68 is a novel signaling molecule in DDR. More specifically, we assessed the impact of Sam68 on DNA damage repair by evaluating the effect of Sam68 ablation on the DNA damage-caused downstream signal amplifying, repair completion, and cell viability and proliferation. We also explored the mechanism through which Sam68 plays a role in DNA damage repair through investigating the interplays between Sam68 and other signaling molecules in the DNA damage response, especially PARP-1, to characterize the function of Sam68 in DNA damage repair. Furthermore, we determined the pathophysiological significance of
Sam68 in the DNA damage response and tumorigenesis in mice.
14 1.4 Figures and Figure Legends
Figure 1.1 DNA lesions and repair pathways.
DNA damage type (Demple and Harrison 1994, Sinha and Hader 2002, Budden and Bowden 2013, DNA damage repair pathway Bauer, Corbett et al. 2015, Hashimoto, Anai et al. 2016) Depurination of N-glycosidic bond, generating an abasic site Base Excision Repair (BER) (Bauer, Corbett et -9 Intrinsic (spontaneous rate = 3X10 /min) al. 2015) damage Deamination, generating a new base different than the original one Base Excision Repair (BER) (Bauer, Corbett et (spontaneous rate = 3.6X10-9/min) al. 2015) Direct Reversal Repair or Base Excision Repair Alkylating agents (BER) (Bauer, Corbett et al. 2015) Damage to a Reactive oxygen species (ROS), oxidizing purines, generating 8-Oxo- Base Excision Repair (BER) (Demple and single strand Deoxyguanosine or 8-Oxo-Deoxyadenosine Harrison 1994, Bauer, Corbett et al. 2015) Extrinsic Nucleotide Excision Repair (NER) (Le May, Egly Metabolically activated carcinogens, generating "bulky" lesions damage et al. 2010) Nucleotide Excision Repair (NER) (Le May, Egly Platinum, generating intrastrand cross-links et al. 2010) Ultraviolet (UV) light B (UVB), catalyzing intrastrand photodimer Nucleotide Excision Repair (NER) (Sinha and formation from pyrimidine Hader 2002) Ultraviolet Light A (UVA) and ultraviolet Light C (UVC) inducing double Homologous Recombination (HR) or strand breaks (DSBs) through clustered local oxidative single-stand Nonhomologous End Joining (NHEJ) breaks (Hartlerode and Scully 2009) Fanconi Anemia (FA) pathway (Moldovan and Damage to Extrinsic Bifunctional alkylating agents, generating interstrand cross-links (ICLs) D'Andrea 2009, Ciccia and Elledge 2010, both strands damage Hashimoto, Anai et al. 2016) Homologous Recombination (HR) or γ-irradiation, generating double strand breaks (DSBs) Nonhomologous End Joining (NHEJ) (Hartlerode and Scully 2009)
15 Figure 1.2 Schematic diagram representing the DNA damage responses in cells.
Endogenous and environmental genotoxic insults (represented by lightning bolt) lead to damage on chromatin (represented by DNA helix). Damage will first be recognized by sensor proteins, and the interconnected transducers in the complex damage signaling pathway will signal the presence of DNA damage to recruit effectors and orchestrate
DNA damage responses, which includes repair of the damage, upregulation of anti- apoptotic genes’ transcription. Failure to do so may lead to senescence of cell cycle
(represented by stop sign) or apoptosis (represented by tombstone). This figure is adopted from the cited literature and remade by myself (Zhou and Elledge 2000).
16 Figure 1.3 The coordination of the homologous recombination (HR) pathway and cell- cycle transition signaling cascade in response to DNA double-stranded breaks
(DSBs).
The DSBs-induced poly(ADP-ribose)polymerase 1(PARP1) activation promotes ataxia telangiectasia mutated (ATM) and Mre11-Rad50- Nbs1 (MRN) complex recruitment and activation, which further trigger and orchestrate repair of DSBs through HR and cell- cycle transition through checkpoint mediators. This figure is summaried from cited literature and made by myself (Kastan and Bartek 2004, Bergink and Jentsch 2009, Ciccia and Elledge 2010). ATM, ataxia telangiectasia mutated; BRCA1, breast cancer 1; CDC25a, cell division cycle 25a; CDKs, cyclin-dependent kinases; CHK1, checkpoint kinase 1;
CHK2, checkpoint kinase 2; H2A, histone 2A; H2AX, histone 2AX; MDC, mediator Of
DNA damage checkpoint; MRN, Mre11-Rad50-Nbs1 complex; PARP1, poly(ADP- ribose)polymerase 1; RAP80, receptor-associated protein 80; RNF168, ring finger protein
168; RNF8, ring finger protein 8; Ubc13, ubiquitin conjugating enzyme.
17
18 Figure 1.4 The structural and functional domains of human PARP1.
Poly(ADP-ribose)polymerase 1(PARP1) is composed of three essential domains: the the
DNA binding domain (DBD), the automodification domain (AD) and the catalytic domain (CAT). Within the BD, there are three zinc fingers (Zn). The AD is mainly comprised with the BRCA1 carboxy-terminal repeat motif (BRCT). The CAT contains one tryptophan-, glycine-, arginine-rich (WGR) domain, one helical subdomain (HD) and the signature ADP-ribosyltransferase domain (ART). This figure is adopted from the cited literature and remade by myself (Rouleau, Patel et al. 2010, Langelier, Planck et al.
2012).
19 Figure 1.5 PARP1 interacts with multiple HR and NHEJ proteins and regulates their recruitment and activity through
PARylation.
DDR pathway effector proteins function PAR synthesis is required for DNA damage-induced ATM activation (Aguilar-Quesada, Munoz- ATM Homologous Gamez et al. 2007, Haince, Kozlov et al. 2007) Recombination PARP1 is crucial for Mre11 and Nbs1 recruitment to DNA damage sites (Haince, McDonald et al. MRN complex 2008) Binding to PARP1 facilitates conformational changes in the DNA-PK synaptic dimer assembly DNA-PK (Spagnolo, Barbeau et al. 2012) Non-homologous PARylation promotes recruitment and/or retention of Ku at DNA double strand breaks (Couto, Ku70/Ku80 End Joining Wang et al. 2011) Polynucleotide PNK and PARP1 are co-recruited to DNA ends together (Audebert, Salles et al. 2006) kinase (PNK)
20 Figure 1.6 The structural and functional domains of Sam68.
The Src-associated substrate during mitosis of 68 kDa (Sam68) protein is composed of six consensus proline-rich motifs (P), a tripartite of a single RNA binding KH domain flanked by the N-terminal of KH region (NK) and the C terminal of KH region (CK), and a C-terminal nuclear localization signal (NLS). This figure is adopted from the cited literature and remade by myself (Richard 2010).
21 2 Sam68 is Required for DNA Damage Responses via Regulating
Poly(ADP-ribosyl)ation.
2.1 Sam68 Plays a Critical Role in Repairing DNA Strand Breaks.
Hypersensitivity to DNA-damaging agents is one of the hallmarks of defective
DDR. To address the role of Sam68 in repair of DNA strand breaks, we first examined the effect of Sam68 deletion in clonogenic survival of mouse embryonic fibroblasts
(MEFs) following exposure to genotoxic stresses. Sam68 deletion in MEFs led to an increased sensitivity to etoposide (a DNA-damaging agent that inhibits DNA topoisomerase II), γ-irradiation, and H2O2 compared with wild-type cells (Fig 2.1A–2.1B and Fig 2.2). To ascertain whether Sam68 is essential for the completion of DNA repair, we performed single-cell gel electrophoresis-based alkaline comet assays, a sensitive method for detecting DNA strand breaks (Olive and Banath 2006). No comet tails were observed in mock-irradiated Sam68-/- and Sam68+/- primary thymocytes, suggesting
Sam68 deletion does not spontaneously cause DNA damage (Fig 2.1C and 2.1D). The vast majority of γ-irradiated thymocytes, in the presence or absence of Sam68, showed prominent comet tail moments, an indicator of DNA damage severity, at 15 min post γ- irradiation (Fig 2.1C and 2.1D). The comet tails lessened in a time-dependent manner in
Sam68+/- thymocytes, and almost no comet tails were detected at 3 h post γ-irradiation.
Strikingly, the comet tails remained prominent in Sam68-/- thymocytes during the same
22 time period (Fig 2.1C and 2.1D), which strongly supports an indispensable role of Sam68 in repairing DNA breaks.
DNA double-strand breaks (DSBs) are the most severe form of damage to DNA, and HDR and NHEJ have been proposed as the major mechanisms used to repair DSBs
(Gunn and Stark 2012). We sought to directly test whether Sam68 facilitates DNA repair through one or more such specific signaling pathways. To this end, we utilized U2OS cell lines that contain chromosomally integrated green fluorescent protein (GFP) reporters with recognition sites for the rare-cutting endonuclease I-SceI to assess the rates of HDR and NHEJ, as GFP positivity by flow cytometry analysis suggests that repair has occurred in these cells (Gunn and Stark 2012). Upon down-regulation of
Sam68 by small interference RNA (siRNA), we observed the repair efficiency of the
NHEJ and HDR pathways was reduced to 42.3% and 12.2%, respectively, in comparison to the nonspecific control siRNA (Fig 2.1E and 2.1F). The efficiency of Sam68 silencing in the U2OS reporter cell lines was verified by IB (Fig 2.1F), and Sam68 down-regulation exhibited no substantial effect on transfection efficiency of these cells (Fig 2.1E).
Moreover, ectopic expression of a Sam68 construct that is resistant to siRNA markedly rescued the repair efficiency of the NHEJ and HDR pathways in the Sam68 knockdown cells (Fig 2.3). Together, these results suggest that Sam68 deletion causes defective DNA repair, thus resulting in persistent DSBs and hypersensitivity to DNA-damaging agents.
23 2.2 Sam68 Deficiency Impairs DNA Damage-initiated PARylation and Signaling
Cascade.
As Sam68 knockdown has a more profound impact on the efficiency of HDR (Fig
2.1E and 2.1F), which is the most important error-free pathway for repairing DSBs, we sought to examine how Sam68 deficiency affects the signaling cascade in response to
DSBs caused by γ-irradiation. Phosphorylation of the histone variant H2AX (γH2AX), a
DNA damage marker that promotes the recruitment of chromatin-modifying complexes and downstream repair factors (Rogakou, Boon et al. 1999, Fernandez-Capetillo, Celeste et al. 2003, Bonner, Redon et al. 2008), was robustly increased in Sam68-sufficient MEFs, primary thymocytes, and U2OS cells treated with γ-irradiation (Fig 2.4A–2.4C and Fig
2.5). In contrast, such DNA damage-triggered γH2AX accumulation was severely attenuated in Sam68-deficient cells (Fig 2.4A–2.4C and Fig 2.5). Moreover, the γ- irradiation-induced phosphorylation of ATM (which is the kinase of H2AX) and its substrates Chk1 and Chk2, all of which are essential DNA damage signaling transducers
(Polo and Jackson 2011), was consistently more robust in Sam68-sufficient MEFs, thymocytes, and U2OS cells in comparison to Sam68-deficient cells (Fig 2.4A and 2.4D and Fig 2.5). These observations further suggest that Sam68 deficiency leads to decreased DNA repair signaling. As DSB-induced ATM phosphorylation is known to rely on the activation of the DNA damage sensor PARP1 and subsequent PARylation
(Aguilar-Quesada, Munoz-Gamez et al. 2007, Haince, Kozlov et al. 2007), we examined
PAR chain formation in γ-irradiated cells in the presence and absence of Sam68. As
24 expected, PAR chains were rapidly and vigorously built up in Sam68-sufficient thymocytes and MEFs following γ-irradiation (Fig 2.4C and 2.4E–2.4F). However, while
Sam68 deficiency did not affect PARP1 levels (Fig 2.4E and 2.4F), γ-irradiation triggered
PARylation was diminished in Sam68-deficient cells (Fig 2.4C and 2.4E–2.4F), which demonstrates that Sam68 is crucial for DNA damage-initiated PAR chain formation.
Moreover, supplementing with exogenous Sam68 markedly restored the DNA damage- initiated PAR synthesis and the PAR-dependent phosphorylation of ATM, Chk1, Chk2, and H2AX in Sam68 knockout (KO) MEFs following γ-irradiation (Fig 2.4A and 2.4E), highlighting the crucial role of Sam68 in the DSB-triggered PARylation and PAR- dependent signaling to DNA repair.
2.3 DNA Damage Enhances Interaction Between Sam68 and PARP1.
Given that Sam68 deletion almost abolished PARylation following γ-irradiation
(Fig 2.4E and 2.4F), we first performed immunoprecipitation assays to examine whether
Sam68 interacts with PARP family proteins in DDR thus affecting the DNA damage- initiated PAR formation. Indeed, at 5 min post γ-irradiation, Sam68 substantially associated with PARP1 and ATM (Fig 2.6A and Fig 2.7); in contrast, no detectable interaction between Sam68 and PARP2, PARP3, and PARP5a/b was observed under the same conditions (Fig 2.7B). Given the observed interaction and that PARP1 is the primary nuclear enzyme that transfers PAR chains to various target proteins in response to DNA damage (Gibson and Kraus 2012), we further characterized the Sam68-PARP1
25 interaction following DNA damage. The γ-irradiation-triggered Sam68-PARP1 association was almost identical in cells pretreated with the PARP inhibitor PJ-34 compared to control (Fig 2.6B), which suggests that PAR chain formation is dispensable for DNA damage-induced Sam68-PARP1 interaction. In contrast, the Sam68-PARP1 interaction in response to γ-irradiation was greatly reduced in the presence of the DNA- binding agent, ethidium bromide (Fig 2.6C), demonstrating that damaged DNA is critical for the DNA damage-induced Sam68-PARP1 interaction.
2.4 Sam68 is Recruited to and Regulate PARylation at DNA Damage Sites.
The fact that PARP1 is rapidly recruited to DNA damage sites (Mortusewicz,
Ame et al. 2007) and damaged DNA enhances the Sam68-PARP1 interaction in DDR (Fig
2.6A and 2.6C) led us to assess whether Sam68 can be recruited to DNA lesions where it interacts with PARP1 and regulates PAR formation. As illustrated by chromatin fractionation assays, Sam68 was remarkably enriched in chromatin fractions in MEFs and U2OS cells following γ-irradiation (Fig 2.6D and Fig 2.8). To ascertain whether the damaged chromatin-enriched Sam68 is recruited to DNA lesions rather than generally bound to chromatin, we performed chromatin immunoprecipitation (ChIP) assays to monitor the DNA damage-triggered Sam68 recruitment to the proximity of a DSB that is actively generated at a unique chromosomally integrated I-SceI site in HDR-GFP reporter U2OS cells (Fig 2.6E). While no association of Sam68 or of PARP1 was detected with chromatin near the I-SceI site in U2OS cells without I-SceI transfection, Sam68 and
26 PARP1 were indeed enriched at the site near the generated DSB in the I-SceI-expressing
U2OS reporter cells (Fig 2.6E). In contrast, we did not detect an enrichment of Sam68 or
PARP1 at the site far away from the cut I-SceI site (Fig 2.6E), which suggests that Sam68 and PARP1 are specifically recruited to the DNA damage site. Moreover, we carried out laser micro-irradiation microscopy assays to visualize the recruitment of Sam68 to local
DNA strand breaks, using MEFs expressing red fluorescent protein (RFP)-tagged Sam68 as well as GFP-tagged PARP1. Consistent with the previous studies (Gibson and Kraus
2012), accumulation of ectopically expressed GFP-PARP1 at DNA lesions was manifested as cytological discernable foci in laser-irradiated Sam68 KO MEFs (Fig 2.6F and Fig 2.9A). Interestingly, RFP-Sam68 formed discrete, cytologically detectable foci, which significantly overlap with the damage foci formed by GFP-PARP1 and endogenous γH2AX, after laser micro-irradiation in parallel samples all collected at the same time post micro-irradiation (Fig 2.6F and Fig 2.9A). In contrast, the RFP control failed to form discrete foci, although GFP-PARP1 formed damage foci as expected, under the same condition, in cells expressing RFP and GFP-PARP1 (Fig 2.6F). Of note, endogenous Sam68 and PARP1 also formed discrete DNA damage foci that significantly overlapped with those manifested by γH2AX, after laser micro-irradiation (Fig 2.9B), further supporting our assertion that Sam68 localizes at sites of damaged DNA significantly overlapping with PARP1 as well as γH2AX during the cellular responses to
DNA damage. To assess the impact of Sam68 on the PAR formation at local DNA damage sites, we carried out immunofluorescence staining assays for PARylation at local DNA strand breaks generated by laser micro-irradiation in the nucleus in the
27 presence and absence of Sam68. As expected, shortly (1 min) after laser micro- irradiation, vigorous PARylation was detected locally in the nuclear regions where laser beams were introduced in WT MEFs (Fig 2.6G). In striking contrast, PAR formation was barely observed in the laser micro-irradiated regions in Sam68 KO MEFs (Fig 2.6G), which suggests that Sam68 is crucial for local PAR synthesis at DNA lesions.
Collectively, these results demonstrate that Sam68 is recruited to and significantly overlaps with PARP1 at DNA damage sites, where it regulates local PARylation in the cellular responses to DNA damage.
2.5 Sam68 is Required for PARP1 Catalytic Activation and Its Release from DNA
Damage Sites.
The findings that Sam68 deficiency attenuates DNA damage-induced global
PARylation (Fig 2.4E and 2.4F) and focal PAR formation (Fig 2.6G) and that Sam68 and
PARP1 interact and overlap at DNA strand breaks after DNA damage (Fig 2.6A and
2.6F) led us to hypothesize that Sam68 reguates damaged-DNA-initiated PARP1 activation. The binding of PARP1 to DSBs is a prerequisite for triggering PARP1 activation and subsequent PAR synthesis (D'Amours, Desnoyers et al. 1999). We therefore sought to examine the impact of Sam68 on the dynamics of PARP1 at sites of
DNA breaks by laser micro-irradiation microscopy. As previously reported (D'Amours,
Desnoyers et al. 1999), GFP-tagged PARP1 rapidly accumulated at DNA damage sites introduced locally by laser-irradiation in Sam68 wild-type and knockout MEFs, reaching
28 maximal accumulation within 1 min (Fig 2.10A and 2.9C). The GFP-PARP1 fluorescence intensities after laser micro-irradiation were comparable in wild-type and Sam68-deleted cells (Fig 2.10A and 2.10C), indicating that Sam68 is not required for PARP1 localization to sites of damaged DNA. Similarly, PARP1 inhibition with Olaparib, compared to the vehicle control, did not affect the recruitment of GFP-PARP1 at sites of damaged DNA
(Fig 2.9C). In addition, Olaparib treatment augmented, rather than attenuated, the localization of RFP-Sam68 at DNA damage foci in the MEFs expressing RFP-Sam68 and
GFP-PARP1 (Fig 2.9D). These results suggest that Sam68 is not required for the recognition and binding of PARP1 to DSBs and vice versa.
It has been acknowledged that the binding of PARP1 to DSBs triggers conformational changes in PARP1 and dramatically elevates its activity to vigorously transfer ADP-ribosyl polymers to numerous substrates including PARP1 itself
(Langelier, Planck et al. 2012). While PARylated PARP1 promotes the recruitment of additional PARP1 as well as other DNA damage-responding molecules to DSBs, auto-
PARylation of PARP1 stimulates its dissociation from DNA damage sites, which probably arises from electrostatic repulsion between the DNA and the highly negatively charged PAR chains (Polo and Jackson 2011). As expected, the GFP-PARP1 foci that formed promptly following laser micro-irradiation faded away rapidly in wild-type
MEFs (Fig 2.10A and 2.10B, Fig 2.10E and 2.10F). Strikingly, the retention time of GFP-
PARP1 at DNA damage sites was substantially prolonged in Sam68 knockout MEFs (Fig
29 2.10A and 2.10B, Fig 2.10E and 2.10F). These results suggest that Sam68 deficiency significantly retards the departure of inactivated PARP1 from local DSBs.
2.6 Sam68 stimulates PARP1 Catalytic Activity via the Interation Between their N-
termini.
The results that Sam68 is not required to target PARP1 to DNA damage sites (Fig
2.10A and 2.10C), that Sam68 and PARP1 interact and overlap at DNA strand breaks after DNA damage (Fig 2.6A and 2.6F) and that Sam68 deficiency traps inactivated
PARP1 at DNA damage sites (Fig 2.10A and 2.10B, Fig 2.10D and 2.10E) suggest that
Sam68 governs DNA damage-initiated PARylation via directly stimulating the catalytic activity of PARP1. To test this hypothesis, we performed in vitro PARylation assays using recombinant PARP1 and Sam68 proteins. As expected, damaged DNA-activated
PARP1 automodified itself with the addition of PAR moieties from the supplemented nicotinamide adenine dinucleotide (NAD+), as indicated by the robust PAR chain formation (Fig 2.11A and Fig 2.12A). Strikingly, incubation of recombinant Sam68 protein, compared to the GST control, with PARP1 dramatically boosted PARP1 activation and PARylation in a dose-dependent manner (Fig 2.11A, compare lane 3 with lane 7, and Fig 2.12B) in the presence of damaged DNA and NAD+. Moreover, in the absence of PARP1, Sam68 recombinant protein was not able to produce a PAR chain with supplemented damaged DNA and NAD+ (Fig 2.12C), indicating that Sam68 per se does not possess the enzymatic activity to transfer ADP-ribosyl polymers. Hence, our
30 results demonstrate that Sam68 simulates PARP1 activation and subsequent PARylation in the presence of damaged DNA and NAD+.
We sought to understand the key domain(s) in Sam68 critical for the interaction with PARP1 and the PARP1-stimulatory function using various Sam68 truncates. We detected the association of the full-length, ΔC, and ΔKH truncated Sam68 to endogenous
PARP1, but not GFP vehicle (Fig 2.11B). In contrast, deletion of the N-terminal amino acids 1–102 (ΔN) of Sam68 almost abolished the association of Sam68 to PARP1 (Fig
2.11B), suggesting a key role of the N-terminus for the interaction between Sam68 and
PARP1. Moreover, we carried out pulldown assays using GST-fused full-length and ΔN truncated Sam68 together with PARP1 recombinant proteins. While full-length Sam68 pulled down PARP1, the ΔN truncated Sam68 failed to do so (Fig 2.11C), which further confirms the critical role of the N-terminus of Sam68 for the Sam68-PARP1 interaction.
To examine whether the Sam68 N-terminus-mediated Sam68-PARP1 interaction is important for the PARP1-stimulatory function, we conducted the in vitro PARylation assays, supplementing PARP1 with full-length or ΔN mutant Sam68 recombinant protein. Indeed, the stimulatory effect of Sam68 for PARP1 activation and PARylation was substantially reduced when Sam68 (ΔN) recombinant protein was utilized, in comparison to full-length Sam68 (Fig 2.11D, compare lane 11 with lane 15). This result underscores an important role of the Sam68 N-terminus in controlling PARP1 activity in vitro. Moreover, to further examine whether the Sam68-PARP1 interaction is functionally important for DNA damage-initiated PARylation in cells, we compared the
31 γ-irradiation triggered PARylation in Sam68 KO MEFs transiently expressing full-length or ΔN truncated Sam68, both of which share strict nuclear localization (Fig 2.11E). The expression of full-length Sam68, but not GFP control, significantly restored the DNA damage-initiated PAR synthesis in Sam68 KO MEFs (Fig 2.11F), consistent with our previous observation (Fig 2.4E). However, ectopic expression of Sam68 (ΔN) mutant failed to restore the γ-irradiation triggered PARylation in Sam68 KO MEFs (Fig 2.11F).
Thus, our results demonstrate that the N-terminus of Sam68, which is critical for the
Sam68-PARP1 interaction, is functionally important for PARP1-catalyzed PARylation.
We then performed structural-functional analyses using full-length and truncated PARP1 proteins to explore the domain(s) on PARP1 essential for Sam68 interaction and Sam68-stimulated PARP1 activation. In contrast to the strong interaction between full-length PARP1 and Sam68, PARP1 (663–1,014) truncated protein, containing the catalytic domain, barely bound Sam68 (Fig 2.11G), which indicates that the stimulatory function of Sam68 on PARP1 activity may not be conferred via a direct interaction between Sam68 and PARP1 catalytic domain. Conversely, the PARP1 (1–662) truncated protein, which contains the DNA-binding domain and automodification domain, associated with Sam68 to a similar extent as full-length PARP1 (Fig 2.11G), indicative of a potentially important role of the PARP1 N-terminus for the stimulatory function of Sam68. Indeed, in striking contrast to the robust PAR chain formation by
Sam68 and PARP1 in the presence of damaged DNA and NAD+, incubation of Sam68 with either PARP1 (1–662) or PARP1 (663–1,014) failed to form detectable PAR chains
32 (Fig 2.11H, compare lanes 18, 20, and 22), suggesting that the Sam68-PARP1 interaction and PARP1 catalytic domain are both required for Sam68 to stimulate PARP1 activation.
Moreover, in the absence of damaged DNA, incubation of Sam68 with PARP1 exhibited no detectable PARP1 activation (Fig 2.11I and Fig 2.12D), which demonstrates that
Sam68 primarily stimulates the DNA-dependent PARP1 activation. Altogether, our results suggest that Sam68 stimulates the DNA-dependent PARP1 activation and subsequent PARylation through the interaction between their N-termini.
2.7 PARP1 Deficiency/Inhibition and Sam68 Deficiency Share Similar Effects on
DNA Repair.
To recapitulate our observed phenotypes due to diminished PAR synthesis in the absence of Sam68, we performed parallel experiments in PARP1-deficient or -inhibited cells. Indeed, pretreatment of thymocytes with PARP1 inhibitors, Olaparib and PJ-34, significantly impeded γ-irradiation-triggered PARylation and the downstream phosphorylation of ATM and Chk1 (Fig 2.13), mirroring the impaired PAR synthesis and DNA repair signaling in Sam68-deficient cells (Fig 2.4). As a result, PARP1 deletion in MEFs, in comparison to wild-type cells, resulted in increased sensitivity to genotoxicity of etoposide and γ-irradiation (Fig 2.14A and 2.14B), which is comparable to Sam68 KO in MEFs (Fig 2.1A and 2.1B). Moreover, alkaline comet assays revealed that
DNA damage repair was substantially delayed in the PARP1-inhibited thymocytes compared to the control cells, as illustrated by the persistent comet tail moments within
33 3 h post γ-irradiation (Fig 2.14C and 2.14D), further supporting a similar effect of Sam68 deletion and PARP inhibition on DNA repair of γ-irradiated lesions in thymocytes.
Furthermore, PARP1 down-regulation by siRNA significantly attenuated the repair efficiency of NHEJ and HDR in the U2OS reporter cell lines, in comparison to the nonspecific control siRNA (Fig 2.14E), which mirrors the impact of Sam68 silencing on the specific signaling pathways that repair DSBs (Fig 2.1F). These results suggest that deficiency in either PARP1 or Sam68 similarly causes defective DNA repair, thus resulting in persistent DSBs and hypersensitivity to DNA-damaging agents. To examine whether PARP1 loss further impacts the hypersensitivity of Sam68 KO MEFs to DNA- damaging agents, we down-regulated PARP1 expression by siRNA in Sam68 KO MEFs.
Indeed, following exposure to DNA-damaging agents, etoposide and γ-irradiation, the clonogenic survival of Sam68 KO MEFs expressing PARP1-specific siRNA was almost identical to those expressing control siRNA (Fig 2.15). These results further highlight the crucial role of Sam68 in controlling the PARP1-catalyzed PARylation in DDR.
2.8 Sam68 Deletion Attentuates PARylation and PAR-dependent Repair Signaling
in Radiodamaged Thymi.
In light of the crucial role of Sam68 in mediating DNA repair in cultured cells, we additionally examined the thymi, which is known to be hypersensitive to radiotoxicity
(Barlow, Hirotsune et al. 1996, de Murcia, Niedergang et al. 1997, Gannon, Woda et al.
2012), derived from mice at various periods post whole body γ-irradiation (WBIR) to
34 assess the impact of Sam68 on DNA repair signaling in damaged organs in vivo. To assess the immediate effect of Sam68, we harvested the thymi from mock- and γ- irradiated mice at 20 min post WBIR. As visualized by immunohistological staining, the
DNA repair signaling in mock-irradiated Sam68+/- and Sam68-/- mice was comparable (Fig
2.16A and 2.16C). Shortly after WBIR, vigorous PAR synthesis and phosphorylation of
ATM, Chk1, Chk2, and H2AX were detected in the thymi derived from Sam68+/- mice
(Fig 2.16A and 2.16C). In striking contrast, such a response was greatly diminished in the γ-irradiated Sam68-/- thymi (Fig 2.16A and 2.16C). Moreover, these results were further supported by IB of thymocyte lysates derived from γ-irradiated Sam68+/- and
Sam68-/- mice (Fig 2.16B and 2.16D). Hence Sam68 is essential for DNA damage-initiated
PARylation and repair signaling in radiodamaged thymus.
2.9 Sam68 Deficient Mice are Hypersensitive to Genotoxic Stresses.
To determine the importance of Sam68 in DDR in vivo, we assessed the impact of
Sam68 deletion on γ-irradiation-caused genotoxicity in mice. Sam68+/- male mice subjected to a sublethal dose of WBIR initially showed a modest loss in body weight and consistently regained their weight by 18 d post WBIR, correlating with 100% survival from radiotoxicity (Fig 2.17A). In striking contrast, more severe weight loss and mortality were observed in Sam68-/- mice, with a survival rate of 17% over a period of 45 d post WBIR (Fig 2.17A). Likewise, Sam68-/- female mice exhibited hypersensitivity to radiotoxicity compared with their Sam68 sufficient littermates, reflected in a sharp
35 reduction in both body weight and survival rate (Fig 2.17B). Moreover, we monitored mouse mortality following intraperitoneal administration of N-methyl-N-nitrosourea
(MNU), a DNA alkylating agent that causes genotoxicity (de Murcia, Niedergang et al.
1997, Masutani, Nozaki et al. 2000). Over a period of 14 d, Sam68-/- mice had higher and accelerated mortality compared to Sam68+/- controls (Fig 2.17C). Together, these results demonstrate a crucial role of Sam68 in protecting mice from genotoxic challenges by γ- irradiation and alkylating chemicals.
We further compared the morphology of the thymus and small intestine in mock- and γ-irradiated animals by gross dissection and histological staining. Of note, the thymic size, structure, and lymphocyte subpopulations of mock-irradiated Sam68+/- and Sam68-/- mice were comparable, showing that Sam68 deficiency does not impair thymus development (Fig 2.17D and 2.17E and S2.11A Fig). Fourteen d post WBIR, the thymi in Sam68+/- mice were indistinguishable to those in mock-irradiated mice in size and morphology (Fig 2.17D), and all exhibited a well-delineated cortex and medulla zones by histological staining (Fig 2.17E), indicative of successful DNA repair in the thymi. In contrast, the thymi in γ-irradiated Sam68-/- mice were severely damaged, with reduced size and abolished cortex–medulla conjunctions, compared with those from γ- irradiated Sam68+/- or mock-irradiated mice (Fig 2.17D and 2.17E). Similarly, the small intestine develops normally regardless of Sam68 sufficiency in mice (Fig 2.17F and
S2.11B Fig). However, Sam68-/- mice subjected to WBIR had more widespread damage to the shortened small intestine, particularly to the structure of the villi and crypts in the
36 duodenum (Fig 2.17F and S2.11C Fig). The severe damage in the thymus and intestine in
Sam68-/- mice strongly supports that Sam68 is essential for radioprotection in mice.
37 2.10 Figures and Figure Legends
Figure 2.1 Sam68 is required for repairing DNA strand breaks.
(A, B) Survival fraction of wild-type (WT) and Sam68 knockout (KO) mouse embryonic fibroblasts (MEFs) 96 h post treatment with indicated concentrations of etoposide for 20 h (A) or indicated doses of γ-irradiation (IR)(B).
(C) Representative microphotographs of alkali comet assay of Sam68+/- and Sam68-/- thymocytes at indicated time points following 4 Gy of IR or mock-treated.
(D) Quantification of tail moments in C, with summarized data from 40–60 cells within
15 random fields for each time point.
(E) Flow cytometric detection of effect of Sam68 knockdown on DNA damage repair efficiency. U2OS reporter cell lines specifically designed to repair DNA damage through nonhomologous end joining (NHEJ) and homology-directed repair (HDR), were transfected with nonspecific control (si-NC) or Sam68-specific (si-Sam68) small interference RNA, together with (+) or without (−) I-SceI plasmid, or green fluorescent protein (GFP) control. Shown are representative flow cytometry analyses of the frequency of GFP+ cells in indicated reporter cell lines 72 h following transfection.
(F) Quantification of relative repair efficiency (normalized to si-NC and I-SceI cotransfected cells) in indicated reporter cell lines, summarized from three independent experiments. The Sam68 knockdown efficiency was examined by immunoblot (IB), with
38 β-actin as a loading control, in indicated reporter cell lines (bottom). Results in (A), (B),
(D), and (F) are expressed as mean and standard error of the mean (SEM). ns, nonsignificant difference; *, p < 0.05; **, p < 0.01; ***, p < 0.001 by Student’s t tests. Data are representative of at least three independent experiments.
39
40 Figure 2.2 Sam68 is required for clonogenic survival of mouse embryonic fibroblasts following exposure to H2O2.
Survival fraction of Wild-type (WT) and Sam68 knockout (KO) mouse embryonic fibroblasts (MEFs) 96 h post treatment with indicated concentrations of H2O2 for 15 min.
Results are expressed as mean and s.e.m. ***, p < 0.001; ****, p < 0.0001 by Student’s t tests.
41 Figure 2.3 Sam68 is critical for repairing DNA damage in U2OS reporter cell lines.
U2OS reporter cell lines, specifically designed to repair DNA damage through nonhomologous end joining (NHEJ) and homologous directed recombination (HDR), were transfected with non-specific control (si-NC) or Sam68-specific (si-Sam68) small interference RNA. 48 h later, cells were transfected with siRNAs and RFP or siRNA- resistant RFP-Sam68, as indicated, together with (+) or without (-) I-SceI plasmid.
Another 72 h later, cells were harvested for flow cytometric analyses of the DNA damage repair efficiency in the indicated reporter cell lines. The relative repair efficiency
(normalized to si-NC, RFP, and I-SceI cotransfected cells) was quantified from three independent experiments. Results are expressed as mean and s.e.m. ns, non-significant difference; *, p < 0.05; **, p < 0.01; by Student’s t tests.
42 Figure 2.4 Sam68 deficiency attentuates DNA damage-initiated repair signaling in cell culture.
(A, E) WT and Sam68 KO MEFs expressing red fluorescent protein (RFP) or RFP-Sam68 were γ-irradiated (IR) at 10 Gy, and whole cell lysates were derived at indicated time points and immunoblotted (immunoblot, IB) for indicated proteins, with β-actin as a loading control.
(B, D, F) Primary thymocytes isolated from Sam68+/- and Sam68-/- mice were γ-irradiated at 4 Gy, and whole cell lysates were derived at indicated time points and immunoblotted for indicated proteins, with β-actin as a loading control.
(C) Immunofluorescence micrographs of PAR and γH2AX in thymocytes collected at 15 min post mock- or IR (4 Gy), with nuclei counterstained by 4′, 6-diamidino-2- phenylindole (DAPI). Scale bar, 10 μm.
43
44 Figure 2.5 Sam68 deficiency attenuates DNA repair signaling in cell culture.
(A, B) U2OS cells transiently transfected with non-specific control (si-NC) or Sam68- specific (si-Sam68) small interference RNA (A) or wild-type (WT) and Sam68 knockout
(KO) mouse embryonic fibroblasts (MEFs) (B) were treated with 4 Gy of γ-irradiation
(IR). Whole cell lysates were derived at indicated time points following IR and immunoblotted (IB) for indicated proteins, with β-actin as a loading control.
45 Figure 2.6 Sam68 is recruited to and regulates PARylation at DNA damage sites.
(A) Coimmunoprecipitation showing the inducible Sam68-PARP1 interaction. WT and
Sam68 KO MEFs were γ-irradiated (IR) at 10 Gy, and whole cell lysates (Input) derived at indicated time points post IR were immunoblotted directly or after immunoprecipitated (immunoprecipitation, IP) with Sam68 antibody for indicated proteins.
(B) WT MEFs were pretreated with DMSO or PJ-34 (20 μM) for 1h, followed by at 10
Gy. Whole cell lysates (Input) derived at the indicated periods post IR were immunoblotted directly or after immunoprecipitated with Sam68 antibody for the indicated proteins.
(C) WT MEFs were γ-irradiated at 10 Gy. Cells were harvested at the indicated periods post IR and lysed in the lysis buffer supplemented with phosphate-buffered saline (PBS) or ethidium bromide (EtBr, 50 μg/ml). The derived whole cell lysates (Input) were immunoblotted directly or after immunoprecipitated with Sam68 antibody for the indicated proteins.
(D) Chromatin fractionation assays showing dynamics of Sam68 on damaged chromatin.
WT and Sam68 KO MEFs were γ-irradiated at 10 Gy, and the chromatin, soluble (Sol. fr.), and insoluble (Ins. fr.) subcellular fractions were derived at indicated time points post IR and immunoblotted for indicated proteins. Casp-3, Caspase-3; H3, Histone H3.
(E) Chromatin immunoprecipitation (ChIP) analyses showing Sam68 recruitment to the
46 DNA double-strand break (DSB) at I-SceI site in HDR-GFP U2OS reporter cells. Upper, diagram shows the locations of I-SceI site and PCR products amplified by primer set 1
(P1), P2, and P3 on HDR-GFP chromosome. HDR-GFP U2OS reporter cells were transfected with (+) or without (−) I-SceI plasmid for 20 h, and the nuclear extracts
(Input) were derived and subjected to ChIP assays with isotype (Iso), Sam68 (68), or
PARP1 (P) antibody. PCR evaluation of the enrichment of Sam68 or PARP1 (as a positive control) adjacent to the DSB using indicated primer sets. Bottom, inputs, and chromatin immunoprecipitants were immunoblotted for Sam68 and PARP1, respectively.
(F) Fluorescence micrographs of Sam68 KO MEFs transiently expressing RFP-Sam68 or
RFP together with GFP-PARP1, before (Pre-IR) and after laser micro-irradiation (Micro-
IR).
(G) Immunofluorescence micrographs of endogenous PAR in WT and Sam68 KO MEFs at 1 min post laser micro-irradiation, with nuclei counterstained by DAPI. As positive controls, γH2AX form damage foci (indicated by arrows).
47
48 Figure 2.7 Sam68 interacts with early DNA damage sigaling molecules PARP1 and
ATM.
(A) Co-immunoprecipitation showing the inducible Sam68-PARP1 interaction. Wild- type (WT) mouse embryonic fibroblasts (MEFs) were γ-irradiated (IR) at 10 Gy and whole cell lysates (Input) derived at indicated time points post IR were immunoblotted
(IB) directly or after immunoprecipitated (IP) with PARP1 or isotype control antibody for indicated proteins.
(B) WT and Sam68 knockout (KO) MEFs were γ-irradiated (IR) as in (A). Whole cell lysates (Input) derived at the indicated periods post IR were IB directly or after IP with
Sam68 antibody for the indicated proteins.
49 Figure 2.8 Chromatin fractionation assays showing dynamics of Sam68 on damaged chromatin.
U2OS cells were γ-irradiated (IR) at 10 Gy and the chromatin, soluble (Sol. fr.), and insoluble (Ins. fr.) subcellular fractions were derived at indicated time points following
γ-irradiation, and immunoblotted (IB) for indicated proteins. Casp-3, Caspase-3; H3,
Histone H3.
50 Figure 2.9 Sam68 localizes to sites of DNA damage in DDR.
(A) Sam68 knockout (KO) mouse embryonic fibroblasts (MEFs) transiently expressing
RFP-Sam68 together with GFP-PARP1 were subjected to laser micro-irradiation (Micro-
IR). Cells were fixed at 1 min post Micro-IR and stained for endogenous γH2AX. Shown are fluorescence micrographs of RFP-Sam68, GFP-PARP1, and endogenous γH2AX at
DNA damage foci.
(B) Immunofluorescence micrographs of endogenous Sam68, PARP1, and γH2AX in wild-type MEFs at 1 min post Micro-IR.
(C) WT MEFs expressing GFP-PARP1 were pretreated with DMSO or 10 μM of Olaparib for 90 min, followed by Micro-IR. The increase in relative fluorescence intensity (RFI) of
GFP-PARP1 at damage foci ~10 sec post Micro-IR versus pre-Micro-IR in DMSO- or
Olaparib-treated cells were graphed, normalized to DMSO controls.
(D) Sam68 KO MEFs expressing RFP-Sam68 were pretreated with DMSO or 10 μM of
Olaparib for 90 min, followed by Micro-IR. The increase in relative fluorescence intensity
(RFI) of RFP-Sam68 at damage foci ~10 sec post Micro-IR versus pre-Micro-IR in DMSO- or Olaparib-treated cells were graphed, normalized to DMSO controls. Results in (C-D) are expressed as mean and s.e.m. ns, non-significant difference; *, p < 0.05 by Student’s t tests.
51
52 Figure 2.10 Sam68 regulates PARP1 retention at but not PARP1 recruitment to DNA damage sites.
(A) Live cell imaging of wild-type (WT) and Sam68 knockout (KO) mouse embryonic fibroblasts (MEFs) expressing GFP-PARP1 (A), or Sam68 KO MEFs expressing GFP-
PARP1 together with red fluorescent protein (RFP), or RFP-Sam68, following laser micro-irradiation (Micro-IR) (indicated by white boxes). Scale bars, 10 μm.
(B) The relative fluorescence intensity (RFI) of GFP-PARP1 at damage foci, normalized to peak fluorescence intensity, in WT and Sam68 KO MEFs were graphed.
(C) Quantification of individual initial (10 sec post Micro-IR) GFP-PARP1 fluorescence intensity at damage foci in WT (n = 20) and Sam68 KO (n = 22) MEFs.
(D) Quantification of individual GFP-PARP1 fluorescence intensity at damage foci at 5 min post Micro-IR, relative to the peak fluorescence intensity, in WT (n = 20) and Sam68
KO (n = 22) MEFs.
(E) Histogram of GFP-PARP1 residence time measured as decay in GFP fluorescence intensity at damage foci to 50% from the peak fluorescence intensity in WT (n = 20) and
Sam68 KO (n = 22) MEFs.
53 54 Figure 2.11 Sam68 enhances the damaged DNA-dependent PARP1 activation and
PARylation in vitro.
(A, D) Recombinant Sam68, Sam68 (ΔN), or GST control protein was incubated in reaction buffer containing damaged DNA alone or with recombinant PARP1 protein in the presence and absence of NAD+, with indicated final concentrations. Where indicated,
PARP inhibitor PJ-34 was added to the reaction mixture. The reaction mixture was separated by SDS-PAGE and subjected to immunoblotting with the indicated antibodies.
(B) Schematic diagram of the GFP-tagged full-length (residues 1–443) and truncated mutants (ΔN lacks residues 1–102, ΔC lacks 347–443, and ΔKH lacks 165–224) of Sam68.
KH, hnRNP K homology; NLS, nuclear localization signal. Whole cell lysates (Input) from HEK293T cells expressing GFP or indicated Sam68 fusion proteins were immunoblotted directly or after immunoprecipitated with GFP antibody for the indicated proteins.
(C) Whole cell lysates (Input) containing His-PARP1, GST, GST-Sam68, or GST-Sam68
(ΔN) recombinant proteins were immunoblotted directly, or following GST pulldown, for indicated proteins.
(E) Immunofluorescence micrographs of Sam68 KO MEFs expressing GFP, GFP-Sam68, or GFP-Sam68 (ΔN) proteins, with nuclei counterstained by DAPI. Scale bar, 10 μm. (F)
Sam68 KO MEFs expressing GFP, GFP-Sam68, or GFP-Sam68 (ΔN) proteins were mock-
55 or γ-irradiated (IR) at 10 Gy, and whole cell lysates were derived at 5 min post γ- irradiation and immunoblotted for indicated proteins, with β-actin as a loading control.
(G) Schematic diagram of the PARP1 full-length (residues 1–1,014), with indicated DNA- binding, automodification, and catalytic domains, and truncated mutants (residues 1–
662 and 663–1,014) of PARP1. Whole cell lysates (Input) from HEK293T cells expressing myc-tagged PARP1, PARP1 (1-662), or PARP1 (663-1,014) protein were immunoblotted directly, or after immunoprecipitation with myc antibody, for indicated proteins. The nonspecific antibody heavy chains are labeled with an asterisk.
(H, I) Recombinant proteins were incubated in reaction buffer in the presence and absence of damaged DNA and NAD+, with indicated final concentrations. The PARP1,
PARP1 (1-662), PARP1 (663-1014) proteins and spontaneously degraded PARP1 (1-662) species are labeled with asterisks and cycles, respectively (H). The reaction mixture was separated by SDS-PAGE and subjected to IB for the indicated proteins.
56
57 Figure 2.12 The DNA-dependent PARP1-mediated PARylation in vitro.
(A) Recombinant PARP1 protein was incubated in reaction buffer in the presence and absence of damaged DNA, NAD+, and PARP inhibitor PJ-34, as indicated. The reaction mixture was separated by SDS-PAGE and subjected to immunoblotting (IB) with the
PAR antibody.
(B) GST control or increasing amount of GST-Sam68 recombinant proteins were incubated with recombinant PARP1 protein in reaction buffer containing damaged DNA and NAD+, with indicated final concentrations. The reaction mixture was separated by
SDS-PAGE and subjected to IB with the PAR and GST antibodies. Right, quantification of relative PARP1 activity based on PAR band density (normalized to the GST control), summarized from three independent experiments. Results are expressed as mean and s.e.m. *, p < 0.05 by Student’s t tests.
(C) The indicated recombinant proteins were incubated in reaction buffer containing damaged DNA and NAD+. The reaction mixture was separated by SDS-PAGE and subjected to IB with the PAR antibody.
(D) The indicated recombinant proteins were incubated in reaction buffer containing
NAD+ in the presence and absence of damaged DNA. The reaction mixture was separated by SDS-PAGE and subjected to IB with the PAR and GST antibodies.
58 59 Figure 2.13 PARP1 inhibition attenuates DNA damage-induced repair signaling in thymocytes.
(A, B) Sam68+/- thymocytes pretreated with Olaparib (20 μM) (A), PJ-34 (20 μM) (B), or
DMSO for 2 h were γ-irradiated (IR) at 4 Gy and whole cell lysates were derived at indicated time points and immunoblotted (IB) for the indicated proteins, with β-actin as a loading control.
60 Figure 2.14 PARP1 is critical for repairing DNA strand breaks.
(A, B) Survival fraction of Wild-type (WT) and PARP1 knockout (KO) mouse embryonic fibroblasts (MEFs) 96 h post treatment with indicated concentrations of Etoposide for 20 h (A) or indicated doses of γ-irradiation (IR) (B).
(C) WT thymocytes were pretreated with DMSO or 20 uM of Olaparib, followed by 4 Gy of IR or mock-irradiation. Cells were harvested at the indicated time points post IR and subjected to alkali comet assay. Shown are representative microphotographs.
(D) Quantification of tail moments in (C), with summarized data from 40-60 cells within
15 random fields for each time point.
(E) U2OS reporter cell lines, specifically designed to repair DNA damage through nonhomologous end joining (NHEJ) and homologous directed recombination (HDR), were transfected with non-specific control (si-NC) or PARP1-specific (si-PARP1) small interference RNA, together with (+) or without (-) I-SceI plasmid, or GFP control. 72 h later, cells were harvested for flow cytometric analyses of the effect of PARP1 knockdown on DNA damage repair efficiency in the indicated reporter cell lines. The relative repair efficiency (normalized to si-NC and I-SceI cotransfected cells) was quantified from three independent experiments. The PARP1 knockdown efficiency was examined by immunoblotting (IB), with β-actin as a loading control, in the indicated reporter cell lines (bottom).
61 Results in A-B and D-E are expressed as mean and s.e.m. ns, non-significant difference;
*, p < 0.05; **, p < 0.01; ***, p < 0.001; ****, p < 0.0001 by Student’s t tests. Data are representative of two independent experiments.
62
63 Figure 2.15 PARP1 knockdown does not impact the sensitivity of Sam68 deleted cells in response to DNA damage.
(A, B) Survival fraction of Sam68 knockout (KO) mouse embryonic fibroblasts (MEFs) silenced with non-specific control (si-NC) or PARP1-specific (si-PARP1) small interference RNA (siRNA) 96 h post treatment with indicated concentrations of
Etoposide for 20 h (A) or indicated doses of γ-irradiation (IR) (B).
(C) The PARP1 knockdown efficiency was examined by immunoblot (IB), with β-actin as a loading control, in Sam68 KO MEFs at 96 h after siRNA transfection.
64 Figure 2.16 Sam68 deletion dampens DNA repair signaling in the radiodamaged thymus.
(A, C) Immunofluorescence micrographs of PAR (A) and phosphorylation of indicated
DNA repair signaling molecules (C) in thymi collected from Sam68+/- and Sam68-/- mice at
20 min following mock-irradiation (Mock) or whole body γ-irradiation (WBIR), with nuclei counterstained by DAPI. Scale bars, 10 μm.
(B, D) Whole cell lysates derived from thymocytes collected as in (A, C) were immunoblotted for indicated proteins, with β-actin as a loading control.
65
66 Figure 2.17 Sam68 KO mice are hypersensitive to genotoxic stresses.
(A, B) Kaplan-Meier analysis of the survival rate (left) and normalized body weight changes (right) in male Sam68-/- mice and their littermate Sam68+/- controls following
6.5 Gy of WBIR (A) and female mice following 7 Gy of WBIR (B). p = 0.0016 (A) and p <
0.0001 (B), respectively, Gehan-Breslow-Wilcoxon test.
(C) Kaplan-Meier analysis of the survival rate in male Sam68+/- and Sam68-/- mice following N-methyl-N-nitrosourea (MNU) administration at 165 mg/kg of body weight. p = 0.0081, Gehan-Breslow-Wilcoxon test.
(D) Photographs of thymi derived from Sam68+/- and Sam68-/- mice at 14 d post mock- irradiation (Mock) or WBIR. Thymi were outlined with dashed green lines and indicated by arrows. H, heart. Scale bar, 5 mm.
(E) Hematoxylin and eosin (H&E) staining of thymi as in (D). C, cortex; M, medulla.
Scale bar, 100 μm.
(F) H&E staining of duodenum from Sam68+/- and Sam68-/- mice irradiated as in (D). V, villi; C, crypts; M, muscularis externa. Scale bar, 100 μm.
67 68 Figure 2.18 Sam68 deficiency does not affect thymus and small intestine development in mice, but causes more severe damage post γ-irradiation.
(A) Representative flow cytometry analysis of indicated immune cell subpopulations in thymocytes derived from naïve Sam68+/- and Sam68-/- mice.
(B, C) Photographs of small intestines collected from Sam68+/- and Sam68-/- mice at 14 days post mock-irradiation (Mock) (B) or whole-body γ-irradiation (WBIR) (C). Scale bars, 2 cm.
69 70 2.11 Materials and Methods
Mice All animal experiments were performed according to protocol number MO13-
H349, approved by the Johns Hopkins University’s Animal Care and Use Committee and in direct accordance with the NIH guidelines for housing and care of laboratory animals. Sam68-/− (Sam68 KO) mice and their gender-matched littermate Sam68+/− heterozygous mice (occasionally substituted with gender-matched littermate Sam68+/+ mice when Sam68+/− ones were lacking but referred to as Sam68+/− alone for simplicity) were produced using heterozygous breeding pairs and were genotyped for disrupted or wild-type Sam68 gene, as previously described (Huot, Vogel et al. 2012). Mice were maintained in a specific pathogen-free facility and fed autoclaved food and water ad libitum.
Cell Culture, Reagents, and Plasmids Wild-type, Sam68 KO, and PARP1 KO MEFs were kindly shared by Drs. Stephan Richard (McGill University, Canada) and Zhao-Qi
Wang (Fritz Lipmann Institute, Germany). Wild-type U2OS cells and the DR-GFP and
EJ5-GFP reporter U2OS cells (Gunn and Stark 2012) were kindly provided by Drs.
Michael Matunis (Johns Hopkins University) and Jeremy Stark (Beckman Research
Institute of City of Hope). MEF and U2OS cells were cultured in DMEM medium containing 10% fetal calf serum, 2 µM glutamine, and 100 U/ml each of penicillin and streptomycin, except for the addition of 10 mM HEPES (pH 7.2–7.5) for U2OS culture.
U2OS reporter cells were cultured using similar medium as for U2OS cells, except without sodium pyruvate. Antibodies used were as follows: Sam68, GST, PARP2,
71 PARP3, and PARP5a/b from Santa Cruz Biotechnology (Dallas, Texas); β-actin and myc from Sigma-Aldrich (St. Louis, Missouri); ATM, PARP1, Caspase-3, Chk1, Chk2, p-Chk1
(Ser345), p-Histone H3 (Ser10), and β-Catenin from Cell Signaling Technology (Danvers,
Massachusetts); PAR from Trevigen (Gaithersburg, Maryland); γH2AX from Millipore
(Billerica, Massachusetts); H2AX from Bethyl Laboratories (Montgomery, Texas); p-
ATM from Rockland (Gilbertsville, Pennsylvania); p-Chk2 from Novus Biologicals
(Littleton, Colorado); RFP from GenScript (Piscataway, New Jersey); Sam68 from
GeneTex (Irvine, California); Histone3 from Abcam (Cambridge, Massachusetts); CD4 and CD8α from BioLegend (San Diego, California). MNU, etoposide (VP16), 4′, 6- diamidino-2-phenylindole (DAPI), and ethidium bromide (EtBr) were obtained from
Sigma-Aldrich. 4-[(3-[(4-cyclopropylcarbonyl)piperazin-4-yl]carbonyl)-4- fluorophenyl]methyl(2H)phthalazin-1-one (Olaparib) and N-(6-oxo-5,6- dihydrophenanthridin-2-yl)-N, N-dimethylacetamide-HCl (PJ-34) were purchased from
Fisher Scientific (Pittsburgh, Pennsylvania) and Enzo Life Sciences (Farmingdale, New
York), respectively. The GFP-PARP1, RFP-Sam68, and I-SceI plasmids were kindly shared by Drs. Anthony Leung (Johns Hopkins University), Johnny He (University of
North Texas Health Science Center), and Jeremy Stark (Beckman Research Institute of
City of Hope), respectively. The GFP, GFP-Sam68, GFP-Sam68 (ΔC), GFP-Sam68 (ΔN),
GFP-Sam68 (ΔKH), GST, GST-Sam68, and GST-Sam68 (ΔN) constructs were described previously (Fu, Sun et al. 2013).
γ-Irradiation The γ-irradiation on MEFs, U2OS cells, and primary thymocytes was
72 performed using a 137Caesium source (dose rate 4 Gy/min). WBIR in mice was performed as previously described (de Murcia, Niedergang et al. 1997, Masutani, Nozaki et al.
2000). Briefly, 6–8 wk-old Sam68+/- and Sam68-/- mice were subjected to a single dose of sublethal γ-irradiation from an MSD Nordion Gammacell 40 Exactor, with a dual
137Caesium source (dose rate 1 Gy/min). The body weight, mortality, and survival of mice were monitored post irradiation, and in some circumstances, the γ-irradiated mice were sacrificed at indicated time points post WBIR for histological and immunohistological analyses.
Proliferation Assays The in vitro proliferation assays were performed as previously described (Wu, Chen et al. 2011). Briefly, 1 103 MEF cells were γ-irradiated at the indicated dose and were seeded in the wells of a 6-well plate immediately after γ- irradiation. In certain cases, 1 103 MEF cells seeded in DMEM medium were treated with or without indicated concentrations of etoposide for 20 h, followed by extensive washes with phosphate-buffered saline (PBS). After incubation for an additional 96 h, the surviving cells were accounted using a Z1 Coulter Particle Counter (Beckman
Coulter, Indianapolis, Indiana), and the survival fractions were calculated by comparing the live cell numbers in treated cultures to those in untreated controls.
Alkaline Comet Assays Comet assays were conducted by using the Comet Assay Kit
(Trevigen) following the manufacturer’s instructions. Briefly, isolated primary mouse thymocytes were mock-treated or γ-irradiated with indicated doses and then allowed to recover in normal DMEM culture medium containing 10 mM HEPES (pH 7.2–7.5) for
73 indicated periods at 37 °C. Cells were collected and washed once with PBS, and 3 105 cells were combined with 1% molten LMAgarose at 37 °C at a ratio of 1:10 (v/v) and immediately pipetted onto slides. Slides were then immersed in prechilled lysis solution for 1 h on ice to lyse cells, followed by alkaline unwinding of chromatin. Alkaline electrophoresis of gelled slides was performed using an Econo-Sub Horizontal System
(C.B.S. Scientific, Del Mar, California) at 24V (0.7 V/cm) at 4 °C for 30 min. The DNA was visualized by SYBR Green staining, and images were taken under an Axio Observer fluorescence microscope (Zeiss, Oberkochen, Germany) and analyzed by CometScore software (TriTek, Sumerduck, Virginia).
RNA Interference and Transfection The siRNAs targeting human Sam68 were described previously (Fu, Sun et al. 2013). Human and mouse PARP1 siRNAs were purchased from Santa Cruz Biotechnology. Transient transfection of siRNA or plasmids in cells was performed using Lipofectamine RNAiMax and Lipofectamine 2000 (Life
Technologies, Frederick, Maryland), respectively, according to the manufacturer's instructions.
DNA Double-Strand Break Repair Reporter Assays DNA double-strand break (DSB) repair assays were conducted as previously described (Gunn and Stark 2012). In brief, 6
× 105 U2OS cells (DR-GFP/homologous-directed repair and EJ5-GFP/NHEJ) were transfected with nonspecific control or Sam68-specific siRNAs. Forty-four hours later, cells were transfected again with siRNAs together with I-SceI (to generate a DSB at the unique I-SceI site) or GFP (to indicate transfection efficiency) plasmids. Seventy-two
74 hours later, cells were collected and subjected to flow cytometry analysis for GFP positive cells.
Flow Cytometry For flow cytometry, 0.5–2 × 106 cells were washed twice with PBS, resuspended in staining buffer (1% fetal bovine serum in PBS), and stained with appropriate antibodies for cell surface markers on ice for 30 min. Following staining and extensive washes with staining buffer, cells were analyzed on a FACSCalibur (BD
Biosciences, San Jose, California). Events were collected and analyzed with the FlowJo software (Tree Star, Ashland, Oregon).
Isolation of Primary Thymocytes Freshly excised thymi from mice were gently teased with a syringe and forceps in DMEM containing 10 mM HEPES (pH 7.2–7.5). The mechanically disrupted cell clumps were poured through 70 μm nylon mesh cell strainers (BD Falcon, Bedford, Massachusetts) to remove connective tissue and prepare single cell suspensions. The cell suspensions were washed once with PBS, and the red blood cells were lysed with Ammonium-Chloride-Potassium (ACK) buffer. The remaining thymocytes were counted, washed twice in PBS, and recovered in DMEM containing 10% fetal calf serum, 2 µM glutamine, 100 U/ml each of penicillin and streptomycin, and 10 mM HEPES (pH 7.2–7.5) for 1 h, followed by further treatments as indicated.
Immunoprecipitation and Immunoblot Immunoprecipitation and immunoblot assays were conducted as previously described (Fu, Sun et al. 2013). In brief, cells were harvested and lysed on ice by 0.4 ml of lysis buffer (50 mM Tris-HCl [pH 8.0], 150 mM
75 NaCl, 1% NP-40 and 0.5% sodium deoxycholate, 1 complete protease inhibitor cocktail
[Roche Applied Science, Indianapolis, Indiana]) for 30 min. The lysates were centrifuged at 10,000 g at 4 °C for 10 min. The protein-normalized lysates were subjected to immunoprecipitation by adding 10 mg/ml of the appropriate antibody, 30 μl of protein
G-agarose (Roche Applied Science), and rotating for more than 2 h at 4 °C. The precipitates were washed at least four times with cold lysis buffer followed by a separation by SDS-PAGE under reduced and denaturing conditions. The resolved protein bands were transferred onto nitrocellulose membranes, probed as described previously (Wan, Weaver et al. 2011, Wier, Neighoff et al. 2012), developed by the Super
Signaling system (Thermo Scientific, Waltham, Massachusetts) according to the manufacturer's instructions, and imaged using a FluorChem E System (Protein Simple,
Santa Clara, California).
Immunofluorescence Microscopy Immunofluorescence microscopy was performed as previously described (Wan, Anderson et al. 2007). Briefly, cells were fixed with 4% paraformaldehyde in PBS and then Cellspin mounted onto slides. After a 5-min permeabilization with 0.05% Triton X-100 in PBS and a 30-min blocking with 5% goat serum, the fixed cells were stained with the appropriate primary antibodies for 1 h, and with fluorescence dye-conjugated second antibodies (Life Technologies) for 1 h together with 1 µg/ml of DAPI (Sigma-Aldrich) for 5 min at 25 °C. The slides were then rinsed with PBS three times and cover mounted for fluorescence microscopy.
Chromatin Fractionation Cells were harvested at indicated time points after γ-
76 irradiation, and cell pellets were resuspended in the NETN buffer (20mM Tris–HCl [pH
8.0], 100 mM NaCl, 1mM EDTA, and 0.5% NP-40) and incubated on ice for 20 min.
Supernatant after 3,000 × g for 10 min was collected as soluble fraction. Pellets were recovered and resuspended in 0.2 M HCl on ice for 30 min and sonicated for 10 s to release chromatin-bound proteins; then, the soluble fractions were neutralized with 1 M
Tris–HCl (pH 8.5) and collected as chromatin fraction, and the pellets were collected as insoluble fraction for further analysis, as described previously (Wu, Chen et al. 2011,
Liu, Wu et al. 2013).
ChIP ChIP assays were performed as previously described (Wan, Anderson et al. 2007).
Briefly, U2OS DR-GFP cells were transfected with a plasmid encoding I-Scel endonuclease to create a DSB. At 20 h post transfection, 4.5 × 106 cells were fixed by 1% formaldehyde at room temperature for 10 min, and the cross-linking reaction was stopped by 125 mM of glycine. Cells were homogenized in cell lysis buffer (5 mM PIPES,
85 mM KCl, 0.5% NP-40, 1 × protease inhibitor cocktail [Roche]), and the nuclei were resuspended in nuclei lysis buffer (50 mM Tris-HCl [pH 8.1], 10 mM EDTA, 1% SDS, 1 × protease inhibitor cocktail) on ice for 10 min and sonicated by Bioruptor UCD-200 (Life
Technologies). The extract was then clarified by centrifugation and diluted 10-fold with dilution buffer (16.7 mM Tris [pH 8.1], 167 mM NaCl, 1.2 mM EDTA, 1.1% Triton X-100,
0.01% SDS) to yield the solubilized chromatin. For immunoprecipitation, anti-Sam68, anti-PARP1, or IgG control antibody, and Protein AG magnetic beads (Thermo
Scientific) were added to the soluble chromatin, and the mixture was incubated at 4 °C
77 overnight. The beads were then washed sequentially with TSE-150 mM NaCl (20 mM
Tris [pH 8.1], 2 mM EDTA, 1% Triton X-100, 0.1% SDS, and 150 mM NaCl), TSE-500 mM
NaCl, buffer III (10 mM Tris [pH 8.1], 0.25 M LiCl, 1% NP-40, 1% deoxycholate, 1 mM
EDTA) and three times with TE (10 mM Tris 8, 1 mM EDTA). The immunoprecipitants were eluted by elution buffer (1% SDS, 50 mM NaHCO3, 20 µg/ml glycogen), and DNA was extracted with phenol-chloroform and resuspended in TE. PCR (30–35 cycles) was performed using MyTaq Red Mix (Bioline, Taunton, Massachusetts) with following primers at or adjacent to the unique I-SceI site: P1F, 5′-GAGCAAGGGCGAGGAGCTGT-
3′; P1R, 5′-CCGTAGGTCAGGGTGGTCAC-3′; P2F, 5′-
TCTTCTTCAAGGACGACGGCAACT-3′; P2R, 5′-TGCCGTTCTTCTGCTTGTC-3′; P3F,
5′-CCGCGACGTCTGTCGAGAAG-3′; and P3R, 5′-GCCGATGCAAAGTGCCGATA-3′.
GST Pulldown Assays The GST pulldown assays were performed as previously described (Wan, Anderson et al. 2007). Briefly, 1 μg of GST/Ni2+ affinity-purified recombinant protein, as indicated, was applied to glutathione Sepharose 4B resins
(Amersham Pharmacia) and incubated for 1 h at 4 °C. After washing, bound proteins were eluted and subjected to SDS-PAGE, followed by immunoblotting.
Laser micro-irradiation Microscopy Laser micro-irradiation microscopy assays were performed as previously described (Mortusewicz, Ame et al. 2007, Kong, Mohanty et al.
2009), with some modifications. Briefly, near infrared (NIR) excitation was provided by a Mai Tai HP Ti:Sapphire laser (Spectra Physics, Santa Clara, California) tuned to 800 nm, 125 fs pulses, and an 80 MHz repetition rate. The laser beam was introduced
78 through a Leica DMi8 confocal microscope (Leica Microsystems, Mannheim,
Germany). The pulsed NIR beam was focused to a diffraction-limited spot with a 63 × oil-immersion objective (1.4 NA). An electro-optic modulator (EOM) was used to create regions of interest (ROIs) targeted for microirradiation by position-dependent power adjustment of the NIR beam. Prior to every experiment, the NIR beam power at the objective was measured to ensure repeatable laser settings that generated a detectable
DDR restricted to ROIs without noticeable cytotoxicity. A coverglass-bottomed dish
(MatTek Corporation, Ashland, Massachusetts) plated with cells was placed into a temperature-controlled chamber (37 °C, 5% CO2) (Life Imaging Services, Basel,
Switzerland) enclosing the entire microscope stand. Immediately after microirradiation, the DDR was captured through time-lapse acquisition of the GFP- or RFP-fused protein recruitment using the FRAP application unit of LAS X acquisition software. All images were processed using the LAS AF Lite software (Leica Microsystems), and we quantified the pre- and post-microirradiation fluorescence intensity of the ROI and an adjacent nuclear area with a similar pre-microirradiation fluorescence intensity, to correct fluorescence intensity for transfection efficiency and photobleaching.
MNU Administration MNU injection-induced genotoxic stress in mice was carried out as previously described (de Murcia, Niedergang et al. 1997, Masutani, Nozaki et al.
2000). Briefly, 6–8-wk-old Sam68+/- and Sam68-/- mice were administrated a single dose of
165 mg/kg body weight of MNU in 200 μl of PBS by intraperitoneal injection. The body weight, mortality, and survival of mice were monitored until 14 d post injection.
79 Histology After euthanizing mice, the thymus was removed, washed once with ice-cold
PBS, immersed in optimal cutting temperature media (Tissue-Tek, Elkhart, Indiana), and frozen in dry ice to preserve the tissue. The duodenum and colon were removed, washed once with ice-cold PBS, fixed in 4% PFA at room temperature for 24 h, and embedded in paraffin. Five-micron sections were cut for all tissues and processed for hematoxylin and eosin (H&E) staining. Stained sections were microphotographed to perform histomorphometric analyses, as previously described (Hodgson, Wier et al.
2015).
Immunohistology Immunohistology was carried out as previously described (Hodgson,
Wier et al. 2015). In Brief, after euthanizing mice, the thymus was excised under aseptic conditions and frozen in optimal cutting temperature media (Tissue-Tek). Five-micron frozen sections were cut using a Microm HM 550 Cryostat (Thermo Scientific), collected on coated slides, fixed in 4% paraformaldehyde, washed with PBS, and blocked with appropriate sera in PBS. After incubating with appropriate antibodies, sections were washed and incubated with fluorescence dye-conjugated second antibodies and 1 µg/ml of DAPI (Sigma). Stained sections were washed and mounted under a coverslip using
Fluoro-gel with Tris Buffer (Electron Microscopy Sciences, Hatfield, Pennsylvania) and examined using an Axio Observer fluorescence microscope (Zeiss).
In vitro PARylation Assays For in vitro PARylation assays, PARP1, PARP1 (1–662),
PARP1 (663–1,014), GST, GST-Sam68, and GST-Sam68 (ΔN) recombinant proteins were purified as previously described (Langelier, Planck et al. 2012) using three
80 chromatographic steps: GST/Ni2+ affinity, heparin-sepharose, and gel filtration. In vitro
PARylation assays were performed as previously described (Zaniolo, Desnoyers et al.
2007). Briefly, the indicated recombinant proteins were incubated for 20 min at 30 °C in a standard assay buffer (100 mM Tris-HCl [pH 8.0], 10 mM MgCl2, 10% (v/v) glycerol, and
1.5 mM DTT) in the presence and absence of damaged DNA (sonicated) and NAD+. The reaction was terminated by the addition of SDS sample buffer (Life Technologies), and the boiled samples were subjected to SDS-PAGE. When indicated, the PARP inhibitor
PJ-34 was added to the reaction mixture at a final concentration of 1 μM for 15 min prior to the reaction.
Statistical Analysis All statistical analysis was performed using GraphPad Prism version 6.0 (GraphPad Software, San Diego, California). The differences between treated and control groups were examined by unpaired Student’s t tests, except Gehan-Breslow-
Wilcoxon tests were used for Kaplan-Meier survival curves. Standard errors of means
(SEMS) were plotted in graphs. ns means nonsignificant difference, and significant differences were considered * at p < 0.05; ** at p < 0.01; *** at p < 0.001; and **** at p <
0.0001.
2.12 Summary
Herein, we report that Sam68, as a novel signaling molecule in DDR, plays a crucial function in governing DNA strand break-triggered PARP1 activation and
81 PARylation. Upon DNA damage, Sam68 is recruited to and significantly overlaps with
PARP1 at DNA damage sites. Interaction between Sam68 and PARP1 via their N-termini is critical for DNA dependent-PARP1 activation and PARylation in vivo and in vitro. In line with the attenuated PAR-dependent repair signaling, DNA damage is poorly repaired in Sam68 deficient cells and animals in comparison to the Sam68 sufficient controls. As a consequence, Sam68 knockout mice are hypersensitive to genotoxicity caused by γ-irradiation and DNA alkylating agents. Hence, our data reveal an unexpected function for Sam68 in DNA damage-initiated early signaling and provide a novel mechanism on the activation and regulation of PARP1 in DDR.
82 3 Conclusion and Future Perspective
3.1 Sam68 is a Novel DNA Damage Repair Signaling Protein that Governs PARP1-
conferred PARylation.
The evidence that Sam68 interacts with PARP1 and is recruited to and substantially overlaps with PARP1 at sites of damaged DNA and that Sam68 deletion diminishes the DSB-initiated PARylation suggests that Sam68 regulates PARP1 activity in
DDR. The similarity in the phenotypes of Sam68- and PARP1-deficient/inhibited cells and animals in response to DNA damage further supports this notion. First, the dramatically attenuated PARylation in response to DNA damage in Sam68-deficient cells (Fig 2.4 C and 2.4E-F) is similar to the abolished DNA damage-triggered PAR synthesis caused by
PARP inhibitors (Fig 2.13). The role of Sam68 in controlling PARP1 activity is further supported by the evidence that Sam68 deficiency retards inactivated PARP1 release from the DNA damage sites (Fig 2.10) and that recombinant Sam68 is sufficient to boost
PARP1 activity in the presence of damaged DNA (Fig 2.11A). Secondly, the PAR- dependent DNA repair signaling cascade is dampened in PARP1 KO cells (Aguilar-
Quesada, Munoz-Gamez et al. 2007, Haince, Kozlov et al. 2007), PARP1-inhibited cells
(Fig 2.13), and Sam68-deficient cells (Fig 2.4A–2.4D and Fig 2.5). Moreover, the delayed repair of DNA strand breaks, as illustrated by comet assays, is similarly observed in
PARP1-deficient (Trucco, Oliver et al. 1998) or -inhibited cells (Fig 2.14C and 2.14D) and
Sam68-deficient cells (Fig 2.1C and 2.1D), in line with the PARP1 malfunction-impeded
83 DNA repair signaling and recruitment of the repair machinery. Hence, Sam68-deleted cells are hypersensitive to genotoxic stress (Fig 2.1A and 1B), similar to what has previously been reported in cells with repressed PARP1 activity through the use of chemical inhibitors or transdominant mutations (Kupper, Muller et al. 1995) and in
PARP1-deleted cells (Fig 2.14A and 2.14B). Furthermore, owing to its key function in
DDR, PARP1 is known to be required for protecting mice from genotoxicity. PARP1 KO mice were reported previously to be hypersensitive to the DNA alkylating agent MNU and γ-irradiation (de Murcia, Niedergang et al. 1997, Masutani, Nozaki et al. 2000), and
γ-irradiation has been shown to cause severe acute damage to the small intestine in
PARP1 KO mice (de Murcia, Niedergang et al. 1997). Interestingly, Sam68 KO mice are also hypersensitive to MNU and γ-irradiation challenges and exhibit acute radiodamage in the small intestine and the thymus (Fig 2.16 and Fig 2.17), thus exactly recapitulating the phenotypes in PARP1 KO mice.
One hallmark of DDR factors is that they are rapidly recruited proximal to sites of damaged DNA, and laser micro-irradiation microscopy provides an ideal assay to visualize this highly dynamic process (Jasin 1996, Essers, Houtsmuller et al. 2006,
Mortusewicz, Leonhardt et al. 2008, Polo and Jackson 2011). It has been demonstrated that activated PARP1 PARylates itself, which further amplify its recruitment to DNA damage sites (Fig 2.6F) (Gibson and Kraus 2012). Notably, in our laser micro-irradiation microscopy assays, the accumulation of ectopically expressed and endogenous Sam68 also leads to the formation of discrete, cytologically detectable foci at sites of damaged
84 DNA that significantly overlap with the damage foci formed by GFP-PARP1 or endogenous γH2AX (Fig 2.6F and Fig 2.9A–2.9B). In addition, we demonstrate that
Sam68 and PARP1 are specifically enriched on the chromatin close to the unique DSB at the I-SceI site (Fig 2.6E). Moreover, Sam68 and PARP1 are capable of physically interacting with each other (Fig 2.11C), and DNA damage enhances interactions between endogenous Sam68 and key initial components of DDR, i.e., PARP1 and ATM (Fig 2.6A and Fig 2.7). Furthermore, recombinant Sam68 is sufficient to facilitate the DNA- dependent PARP1 activity in vitro (Fig 2.11A and Fig 2.12D), and the N-terminus of
Sam68 is both critical for the Sam68-PARP1 interaction and functionally important for
PARP1-catalyzed PARylation (Fig 2.11B–2.11D). Together, these results suggest that
Sam68, functioning as a key initial signaling molecule, is crucial in regulating PARP1 activity and the PAR-dependent signaling cascade in DDR.
In summary, our data demonstrate that the deletion of Sam68 attenuates DNA damage-triggered PARP1 activation, PARylation, and PAR-dependent DNA repair signaling cascades, in line with our observations of an indispensable role for Sam68 in the
HDR and NHEJ repair signaling pathways. Moreover, we show that Sam68 KO mice are hypersensitive to genotoxic challenges by DNA damaging agents. Mechanistically, upon
DNA damage, Sam68 is recruited to DNA damage sites, where Sam68 stimulates the catalytic activity of PARP1 at DNA lesions. Based on our data, we propose a working model (Fig 3.1). In the presence of itself, Sam68 co-localizes with PARP1 at DNA damages sites, where it directly stimulates PARP1 catalytic activity. Activated PARP1
85 produces sufficient PAR chains to signal the occurrence of damage and recruit downstream effectors to repair damage. As a consequence, tissue can maintain genome integrity and tissue homeostasis. However, in the absence of Sam68, PARP1 is incapable of building up PAR chains, although it can still be recruited to DNA damage sites. This leads to an attenuated signaling cascade and inefficient damage repair, which further causes genomic instability and tissue injury.
3.2 Sam68 is the Master Regulator of DDR via Stimulating PARP1 Catalytic
Activation.
In addition to its indispensible role in facilitating DNA damage repair completion by stimulating PARP1 activation (Sun, Fu et al. 2016), we have also illustrated that loss of
Sam68 dramatically decreases phosphorylation of Chk1 and Chk2, which are essential kinases for controlling cell-cycle checkpoints, suggesting Sam68 is critical for the rapid yet transient cell cycle arrest in order to repair damaged DNA and prevent passaging damaged genetic information (Sun, Fu et al. 2016). Moreover, we have discovered that
Sam68 is required for genotoxic stress-induced NF-κB activation to enhance anti- apoptotic gene transcription (Fu, Sun et al. 2016). We reported that loss of Sam68 impairs
PARP1-catalyzed PARylation, which further leads to diminished NF-κB signalosome formation and attenuated NF-κB activation. Therefore, dampened NF-κB-dependent transcription of anti-apoptotic gene is inadequate to protect cells and tissues from
86 genotoxicity. These findings indicate that Sam68 is also critical for transcription activation and preventing cells that are repairing damaged DNA from undergoing apoptosis.
Together, our data illustrates that the recruitment of Sam68 to DNA damage sites and its role in stimulating PARylation are essential for repair completion, cell-cycle control, transcription activation and cell-fate decision, all of which constitute the four fundamental branches of DDR (Fig 1.2). Hence, our model proposes Sam68 as a master regulator of all DDR aspects in cells, essential for survival and proliferation (Fig 3.2).
3.3 Sam68 Bridges Different Stages in the Process of Upregulating DNA Damage-
initiated Gene Expression.
Sam68 harbors versatile functions in cells, which include initiating DNA damage repair (Sun, Fu et al. 2016), activating NF-κB-dependent transcription in response to DNA damage (Fu, Sun et al. 2016, Sun, Fu et al. 2016), modulating the promoter specificity of
NF-κB-dependent transcription (Fu, Sun et al. 2013), and participating in pre-mRNA processing (Lukong and Richard 2003). It is tempting to speculate that these seemingly random functions are actually into a comprehensive program, in space and time, during
DDR. Initially, Sam68 is recruited to DNA damage sites, where it stimulates PARylation upon induction of DNA damage. However, the association of Sam68 with the damaged chromatin is transient in that it departs the damaged sites prior to the completion of both
DNA damage repair and genotoxic-induced NF-κB activation, indicating that Sam68 may be cued to debut its other functions after being post-transnationally modified at the sites of DNA damage. Based on the findings that Sam68 is a non-Rel subunit of the NF-κB
87 complex and that Sam68 mediates the promoter specificity of NF-κB, it is logical to hypothesize that Sam68 binds to the activated NF-κB complex to promote anti-apoptotic and/or DNA damage repair/cell cycle-related gene transcription in the second phase.
Finally, as an RNA-binding Protein (RBP), Sam68 facilitates pre-mRNA processing in the nucleus and chaperons mRNA translocation from the nucleus to the cytoplasm and participates in further regulation of mRNA and translation in the third phase. This hypothesis draws a broad picture of how an RBP can participates in different phases of
DDR with its distinct yet coupled functions.
3.4 RBPs are Emerging as a New Group of Important Effectors in DDR.
RBPs are a class of proteins that harbor at least one RNA-binding domain and bind single- and/or double-stranded RNA to form ribonucleoprotein complexes in cells
(Lunde, Moore et al. 2007). Eukaryotic cells encode approximately 500 RBPs with diverse
RNA binding specificity and protein-protein interaction affinity (Lunde, Moore et al.
2007). RBPs are known to play important roles in regulating the biogenesis, transportation, cellular localization, turnover, and function of RNA in cells (Glisovic,
Bachorik et al. 2008).
On top of these versatile functions in regulating RNA, increasing amount of RBPs, in addition to Sam68, have been suggested to play important roles in DDR. Over the last decade, innumerable research reports have provided a long list of post-translational
88 modifications (PTMs) on different RBPs in response to DNA damage (Adamson,
Smogorzewska et al. 2012). In particular, several large-scale mass spectrometry-based proteomics screenings conducted recently have identified RBPs as a major group of target proteins that are prone to various PTMs in the cellular response to DNA damage
(Matsuoka, Ballif et al. 2007, Gagne, Isabelle et al. 2008, Bennetzen, Larsen et al. 2010,
Bensimon, Schmidt et al. 2010, Beli, Lukashchuk et al. 2012). PTMs, such as phosphorylation and PARylation, occur rapidly upon DNA damage to expand the amino acids’ chemical repertoire by supplementing new functional groups, resulting in efficient
DNA damage repair (Dantuma and van Attikum 2016). To date, over one hundred RBPs have been shown to be phosphorylated upon induction of DNA damage. Combining antibodies to phosphor-SQ or phosphor-TQ sites, Matsuoka and colleagues identified over 124 human and mouse mRNA-transcription/processing related proteins that are phosphorylated in response to DNA damage (Matsuoka, Ballif et al. 2007). Bensimon and colleagues discovered 753 phosphorylation sites mapping to 394 proteins that are modulated in response to treatment with the radiomimetic drug neocarzinostatin. Most of the identified phosphorylated protein are related to nucleic acid metabolism, especially
RNA Processing (Bensimon, Schmidt et al. 2010). In another mass spectrometry study combined with affinity enrichment of phosphopeptides, Bennetzen and colleagues reported on the temporal profiles of 594 regulated phosphorylation sites on 209 proteins in cells exposed to γ-irradiation, further highlighting the relevance of RNA processing proteins in DDR (Bennetzen, Larsen et al. 2010). Finally, Beli and colleagues discovered that 1470 high confidence phosphorylation sites were upregulated following treatment of
89 etoposide, a topoisomerase II inhibitor that primarily causes double-strand DNA breaks, or γ-irradiation, many of which were RBPs (Beli, Lukashchuk et al. 2012). RBPs have also been revealed as DDR effector proteins through mass spectrometry-based, PARylation- focused screens. Jungmichel and colleagues discovered that about 30% of all significantly
PARylated proteins in response toD oxidative DNA damage are involved in RNA metabolic processes and process RNA-binding activity (Jungmichel, Rosenthal et al.
2013). RBPs involved in RNA metabolism and RNA splicing were shown to be PAR- associated by Gagné and colleagues (Gagne, Hunter et al. 2003). Notably, few RBPs identified in these studies have been experimentally confirmed to be essential in DDR.
Indeed, a genome-wide siRNA screen by Paulsen and colleagues revealed that about 50
RBPs, including mRNA processing factors, ribonucleoproteins, and others, are required for maintaining genome stability (Paulsen, Soni et al. 2009).
Collectively, the evidence available indicate that RBPs contribute to almost all aspects of DDR: not only through fine-tuning gene expression as a long-term remedy after damage, but also in preventing accumulation of DNA breaks as a safeguard, and even directly participating in DNA damage repair.
3.4.1 RBPs Ensure Precise DNA Replication to Safeguard Genomic Information.
DNA replication is a fundamental process that ensures precise duplication of genetic information in all cells (McCulloch and Kunkel 2008). As part of the replication process, however, there are many perturbations that cause damage to DNA (Ganai and
Johansson 2016). Several RBPs have been reported to help the DNA replication
90 machinery overcome the numerous barriers encountered while sliding along the template strand, allowing accurate reproduction of the genetic information and thereby safeguarding the DNA code (Dutertre, Lambert et al. 2014).
Okazaki fragments are newly synthesized short DNA fragments on the lagging template strand during DNA replication. Maturation of Okazaki fragments is pivotal to high-fidelity DNA replication (Sakabe and Okazaki 1966). Previous studies show that heterogeneous nuclear ribonucleoprotein A1 (hnRNP A1) interacts with flap endonuclease-1 (FEN-1) and stimulates its nuclease activities to produce a suitable substrate during Okazaki fragment ligation (Chai, Zheng et al. 2003).
Collision between replication forks and transcription complexes can occur on very long genes that need more than one cell cycle to be transcribed, or genes that are transcribed during S phase (Helmrich, Ballarino et al. 2013). The consequences of transcription-replication collision favor the initiation or stabilization of R loops, which occur when newly synthesized RNA hybridizes with the template DNA strand and displace the non-template DNA strand to form a three-strand nuclei acid structure
(Westover, Bushnell et al. 2004). The presence of R loops may lead to DNA damage and further give rise to cell death or tumor formation (Prado and Aguilera 2005, Tuduri,
Crabbe et al. 2009, Aguilera and Garcia-Muse 2012). Several RBPs have been reported to prevent the formation of stable R loops by assisting mRNA ribonucleoprotein (mRNP) particles in packaging the nascent mRNAs and prevent its invasion into the DNA duplex
(Aguilera and Garcia-Muse 2012). Alternative splicing factor/splicing factor 2 (ASF/SF2),
91 also known as serine/arginine-rich splicing factor 1 (SRSF1), associates with transcription complexes and organizes pre-mRNP particles formation. Depletion of ASF/SF2 leads to the destabilization of the pre-mRNP particles, exposing certain regions of the RNA to template DNA, and allowing the formation of R loops (Li and Manley 2005). Mutations in the components of the conserved eukaryotic mRNA transcription/processing complex, including Tho2 (Piruat and Aguilera 1998), Hpr1 (Huertas and Aguilera 2003), mitochondria fusion protein targeting factor 1 (Mft1) and Thp2 (Chavez, Beilharz et al.
2000), as well as mutations in components of the mRNA export machinery such as
Sub2/UAP56 (Fan, Merker et al. 2001) and Yra1/always early (ALY) (Jimeno, Rondon et al. 2002), show strong transcription-dependent hyperrecombination phenotypes, which could be a sign of genome instability caused by R loops. Moreover, RBPs are capable of preventing R loop formation by coordinating with DNA topoisomerase I (TOP-1). TOP-1 is a key mediator at the interface between DNA replication and mRNA transcription by directly relaxing DNA topology (Rossi, Labourier et al. 1996, Tuduri, Crabbe et al. 2009).
A recent proteomic analysis identified 36 nuclear proteins that were associated with TOP-
1, most of which were RBPs (Czubaty, Girstun et al. 2005). Among these, Nucleolin/Nsr1 was shown to interact with TOP-1 through its N terminus, where it regulates TOP-1 cellular localization (Edwards, Saleem et al. 2000). ASF/SF2 is required to mediate TOP-1
DNA binding or cleavage (Andersen, Tange et al. 2002). Pyrimidine tract binding protein- associated splicing factor (PSF) stimulates the topoisomerase activity of TOP-1 (Straub,
Grue et al. 1998). This evidence strongly supports the notion that RBPs are crucial for preventing DNA damage occurring during collision between replication and
92 transcription, by preventing R loop formation via packaging the nascent mRNAs, and by regulating TOP-1 function by various mechanisms.
Telomeres are essential in protecting chromosome ends from shortening or fusion to other chromosomes during mitosis and meiosis (Blackburn and Gall 1978). The telomerase holoenzyme, containing reverse transcriptase, the telomerase RNA, and other accessory factors, maintains telomere length by adding telomeric repeats to the 3’ end of the chromosome (Blackburn, Epel et al. 2015, Schmidt and Cech 2015). RBPs inside and outside of the holoenzyme play important roles in protecting telomere and chromosome integrity (Han, Tang et al. 2010). Suppressors with morphogenetic defects in genitalia
(SMG) proteins have been shown to enrich at telomeres in vivo and negatively regulate telomeric repeat-containing RNA (TERRA) association with chromatin to protect chromosome ends from telomere loss (Azzalin, Reichenbach et al. 2007). In yeast, the Est1 protein directly binds to both the telomerase RNA and the single-stranded telomeric
DNA binding protein Cdc13p. Loss of Est1 attenuate telomere elongation and further leads to senescence (Lundblad and Szostak 1989, Pennock, Buckley et al. 2001). When overexpressed, the human homologs of yeast Est1, hEST1A and hEST1B, have been shown to be specifically associate with almost all active telomerase in cell extracts and cause chromosome-end fusion due to uncap chromosome ends (Reichenbach, Hoss et al.
2003, Snow, Erdmann et al. 2003). Up-frameshift (UPF1) binds to telomeres during the S and G2/M phases, therefore UPF1 depletion causes telomere instability and cell arrest in S phase (Azzalin and Lingner 2006, Chawla, Redon et al. 2011). In addition, several
93 observations imply that members of the heterogeneous nuclear ribonucleoprotein
(hnRNP) family play important roles in telomere maintenance, in particular, hnRNPs A1,
A2/B1, A3 and D act as molecular adapters that regulate the interactions between telomerase and telomeres by binding to both telomerase RNA and single-stranded telomere DNA (Ford, Wright et al. 2002, Han, Tang et al. 2010). Altogether, these observations indicate that RBPs are important in the synthesis and maintenance of the telomeres, which further prevent damage to the DNA.
3.4.2 RBPs Coordinate Gene Expression as a Long-term Remedy to DNA Damage.
Upon DNA damage, eukaryotic cells modify the global gene expression pattern to orchestrate an effective DDR. On one hand, DNA damage triggers a transient global transcriptional repression to prevent intensive energy use during transcription and translation and to avoid the generation of proteins with potentially dangerous mutations
(Heine, Horwitz et al. 2008). On the other hand, DNA damage prompts the expression of a variety of genes involved in DNA repair, cell cycle control and anti-/pro-apoptosis
(Roos, Thomas et al. 2016).
To achieve precise regulation in gene expression, cells execute both transcriptional and post-transcriptional control mechanisms. An increasing number of links between
RBP-mediated transcriptional and DNA damage have been established. In response to
DNA damage, heterogeneous nuclear ribonucleoprotein K (hnRNP K) is recruited to the
94 promoters of p53-responsive genes as a transcriptional cofactor, where it coordinates the transcriptional response to DNA damage in an ATM/ATR-dependent manner (Moumen,
Masterson et al. 2005). UV treatment increases phosphorylation of the C-terminal domain of RNA polymerase II (RNAPII), leading to a transient repression of gene transcription in cells by blocking RNAP II recruitment to the transcription initiation complex (Rockx,
Mason et al. 2000). γ-irradiation induced DNA breaks cause RNA polymerase I (RNAPI) transcription repression through an ATM/Nbs1/MDC1-dependent pathway (Kruhlak,
Crouch et al. 2007). DNA damage also induces CstF to form the CstF/BRCA1 associated
RING domain 1 (BARD1)/BRCA1 complex, and cause a transient inhibition of mRNA 3’ end processing, providing a link between transcription-coupled RNA processing and
DNA repair (Kleiman and Manley 2001, Mirkin, Fonseca et al. 2008).
Moreover, increased attention has been devoted to RNA splicing and its role in
DDR (Shkreta and Chabot 2015). During pre-mRNA splicing, introns are removed from a pre-mRNA transcript and exons are joined to produce one isoform of a mature mRNA.
Alternative splicing allows a single gene-originated pre-mRNA to be spliced in different ways to generate distinct mRNAs (Black 2003). Changes in post-translational modification, localization and activity of RBPs, which are component of the spliceosome or splicing regulators, promote alternative splicing during DDR (Shkreta and Chabot
2015). Phosphorylation and deacetylation of SRSF2, which belongs to an important class of splicing regulators of the serine/arginine-rich (SR) protein family, leads to alternative splicing of caspase-8 pre mRNA, generating a competitive inhibitor of caspase-8, in
95 response to cisplatin treatment (Edmond, Moysan et al. 2011). CPT, a DNA topoisomerase I inhibitor that causes DNA strand breaks, induces alternative splicing of
TATA-box binding protein associated factor 1 (TAF1) through ATR-dependent degradation of a subset of splicing-regulatory proteins, such as transformer 2 (Tra2) (Katzenberger,
Marengo et al. 2009). Upon UV irradiation, the Ewing sarcoma (EWS) protein dissociates from ABL proto-oncogene 1, non-receptor tyrosine kinase (ABL1) mRNA and relocalizes to the nucleoli to regulate of ABL1 expression by alternative splicing (Paronetto, Minana et al.
2011). UV irradiation also results in hnRNP A1 cytoplasmic accumulation and phosphorylation by the MAP kinase (MKK)3/6-p38 pathway, paralleled by alternative splicing pattern change of an adenovirus E1A pre-mRNA reporter (van der Houven van
Oordt, Diaz-Meco et al. 2000).
In addition, RBPs also participate in regulation of gene expression through mediating mRNA turnover and translation. DNA damage caused by UVC can increase the stability of a subset of mRNA and destabilize another subset of mRNA to regulate gene expression (Wang, Furneaux et al. 2000, Fan, Yang et al. 2002). On the translational level, Lu and colleague showed that γ-irradiation induces gene expression changes, and this could involve RBPs mediated shuttling of existing mRNAs to and away from polysomes (Lu, de la Pena et al. 2006). Indeed, evidence from four different individual studies on the translation of p53 mRNA support this hypothesis. One of these studies has shown that HuR binds to the 3’ untranslated region (UTR) of p53 mRNA and enhances p53 translation (Mazan-Mamczarz, Galban et al. 2003). A similar effect has been observed
96 in ribosomal protein L26 (RPL26). RPL26 binds to the 5’ UTR of p53 mRNA and increases p53 translation after DNA damage to induce G1 cell-cycle arrest and augment γ- irradiation-induced apoptosis (Takagi, Absalon et al. 2005). Mdm2 is also known to be phosphorylated by ATM at Ser395 to bind p53 mRNA in the nucleus, which leads to an increase in p53 translation in response to genotoxic stress (Gajjar, Candeias et al. 2012).
Additionally, during the treatment of cancer with the chemotherapy medicine doxorubicin, polypyrimidine tract binding protein (PTB) translocates from the nucleus to the cytoplasm, where it binds to p53 mRNA internal ribosome entry sites (IRES) to facilitate p53 translation (Grover, Ray et al. 2008). In summary, RBPs are essential in orchestrating gene expression post DNA damage by participating both post- transcriptional and translational regulation.
3.4.3 RBPs Play an Important and Direct Role in DNA Damage Repair.
DNA lesions are so deleterious that cells have evolved an efficient regulation system that recruits many proteins to the sites of damage to repair the lesions (Zhou and
Elledge 2000). In addition to influencing DNA homeostasis via ensuring proper DNA replication (1.3.1) and coordinating gene expression (1.3.2), an increasing number of RBPs have been reported at the sites of damage, where they directly play important roles to repair damage. Fused in sarcoma/translocated in liposarcoma (FUS) has been long believed to promote D-loop formation, an essential step in HR (Baechtold, Kuroda et al.
1999). Moreover, Fus-/- mice exhibit phenotypes that are typically cause by DSB repair
97 defects, such as defective B cell development and activation, chromosomal instability, male sterility, and radiosensitivity (Hicks, Singh et al. 2000, Kuroda, Sok et al. 2000).
Recently, the observation of the PAR-dependent recruitment of FUS to sites of double- strand breaks (DSBs) locally introduced by laser microirradiation (Mastrocola, Kim et al.
2013) further suggests that FUS might directly participate in DNA damage repair. In addition, several heterogeneous nuclear ribonucleoproteins (hnRNPs) have been reported to be recruited at sites of DNA lesions. For instance, hnRNP C partially relocates to DNA damage sites, where it interacts with BRCA2 and BRCA1, to facilitate HR
(Anantha, Alcivar et al. 2013). The hnPNP U-like (hnRNP UL) protein is recruited to
DSBs in MRN- and PAR-dependent manners, where it promotes the recruitment of the
BLM helicase, a prerequisite for DSB resection (Polo, Blackford et al. 2012, Hong, Jiang et al. 2013). There is also evidence that the non-POU domain containing, octamer-binding protein (NONO) is also directly involved in DNA repair. It forms a heterodimer with splicing factor, proline- and glutamine-rich protein (SFPQ/PSF), and is rapidly recruited to sites of DNA damage induced by laser micro-irradiation (Salton, Lerenthal et al. 2010), in a PAR-dependent manner (Krietsch, Caron et al. 2012). The findings that NONO promotes NHEJ while repressing HR (Krietsch, Caron et al. 2012), and that both binding partners were identified as a candidate DNA double-strand break rejoining factor
(Bladen, Udayakumar et al. 2005) support the notion that NONO plays a direct role in
DNA DSB repair pathway decision at sites of DNA damage.
98 Nevertheless, it is noteworthy that not all RBPs recruited to sites of damage participate in repairing DNA lesions directly. It is most likely that those RBPs, especially whose roles are known to be involved in splicing, are relocated to the proximity of DNA lesions, where they are post-translational modified by damage-activated PTM enzymes and then released to coordinate splicing during damage repair. For example, RBMX, a hnRNP protein that regulates alternative splicing, is recruited to sites of damage shortly after micro-irradiation in a PAR-dependent manner, and is removed from damaged DNA to form an ‘‘anti-stripe’‘. RNA binding motif protein, X-linked (RBMX) promotes homologous recombination repair of DNA damage through facilitating BRCA2 expression (Adamson, Smogorzewska et al. 2012). DEAD-box helicase 17 (DDX17), a
DEAD-box RNA helicase, is phosphorylated in response to γ-irradiation (Matsuoka,
Ballif et al. 2007), and is also recruited to sites of micro-irradiation damage (Adamson,
Smogorzewska et al. 2012). Given that DDX17 is known to regulate alternative splicing of several DNA- and chromatin-binding factors (Dardenne, Pierredon et al. 2012), it is logical to speculate that DDX17 could be recruited to damage sites, and then phosphorylated DDX17 is released to mediate gene expression.
There is another possible venue by which RBPs could be recruited to the proximity of DNA damage sites and promote damage repair. DSBs induce the production of ∼21-nucleotide small RNAs from sequences in the vicinity of DSB sites.
These small noncoding small RNAs (ncRNAs) are required for DNA damage foci formation and ATM signaling pathway activation (Francia, Michelini et al. 2012, Wei, Ba
99 et al. 2012). RNA polymerase IV (Pol IV), dicer 1, ribonuclease III (DICER) and drosha ribonuclease III (DROSHA) have been implicated to play important roles in generation of these small ncRNAs. The biogenesis, maintenance and function of these small ncRNAs at sites of DNA damage most likely involve other not-yet-identified RBPs.
Taken together, these observations highlight the emerging functions of RBPs in preventing DNA damage generation and facilitating DNA damage repair, beyond their known functions in RNA-related bioprocesses, which suggests a more comprehensive mechanisms through which RBPs function in DDR. Of note, the RBPs that participate in
DNA damage repair, all function after PARP1 catalytic activation and PAR formation.
These findings may indicate a potential common characteristic of RBPs in DDR, in particular they could use the same motif for binding to both RNA and PAR (Teloni and
Altmeyer 2016). Based on the similarity in the composition and structure of RNA and
PAR, PAR could compete with RNA for the RNA-bind motifs within RBPs to halt mRNA splicing and switch to directly participate in DNA damage repair post PARP1 activation.
In striking contrast, Sam68 is a previously unappreciated early signaling molecule that functions prior to, or at least in parallel to, PARP1 activation in the cellular response to
DNA damage (Sun, Fu et al. 2016). More importantly, Sam68 is capable to stimulate the catalytic activity of PARP1 in vitro (Fu, Sun et al. 2016).
100 3.5 Sam68 May Serve as a Potential Therapeutic Target by PARP1 Inhibition.
In light of their crucial roles in various DNA damage repair-signaling pathways, the inhibition of PARP1 as well as other PARP family proteins has emerged as a promising therapeutic approach for treating multiple human diseases associated with impaired DNA repair activities (Papeo, Forte et al. 2009, Anders, Winer et al. 2010).
However, the current classes of PARP inhibitors undergoing clinical trials and the FDA- approved Olaparib are all based on a competitive binding strategy first observed with nicotinamide (Wahlberg, Karlberg et al. 2012), and almost all PARP inhibitors are derivatives of this natural PARP inhibitor (Luo and Kraus 2012). There is an urgent need for improving their specificity for each individual PARP family member and lowering their off-target effect and toxicity (Gibson and Kraus 2012). That said, the nature of
PARP1’s rapid recruitment (within milliseconds) to DNA damage sites upon sensing lesions combined with the fact that activated PARP1 vigorously catalyzes the PARylation reaction, makes it difficult to elucidate the mechanism of PARP1 activation in DDR.
Although post-translational modifications and interactions with other proteins have been proposed to fine-tune PARP1 activity in DDR (Kauppinen, Chan et al. 2006, Mao, Hine et al. 2011), the stimulatory mechanisms of PARP1 activation at DNA damage sites remain largely unknown. Our proposed model that Sam68, as a previously unrecognized stimulatory factor beyond DNA strand breaks, stimulates PARP1 activation and
PARylation at DNA damage sites could provide a novel strategy to develop a new category of PARP1 inhibitors and therapeutics for human diseases involving DNA repair.
101 3.6 Future Perspective
3.6.1 Virtual Regulation Between Sam68 and ATM
It is noteworthy that our work elucidates a novel role for the poorly-characterized
N-terminus of Sam68 in DDR signaling, that it is required for interaction with PARP1 and crucial for the activation of PARP1. Albeit not appearing to harbor any well-defined functional domains or motifs, the N-terminus of Sam68 does contain a cluster of several serine and threonine residues. This cluster could make Sam68 a potential target of serine/threonine phosphorylation. The evidence that Sam68 interacts with ATM shortly post IR (Fig 2.7B) hints that serine/threonine phosphorylation of Sam68 by ATM may serve as an important functional switch that allows Sam68 to be recruited to DNA damage sites, where it interact with PARP1 and regulates PARP1-catalyzed PARylation in DDR. This hypothesis is worthy of experimental validation in future studies. We could first test whether ATM is the kinase that catalyzes Sam68 serine/threonine phosphorylation during DDR. Using in vitro kinase assay with recombinant proteins, we can easily assess the direct enzymatic reaction. We can further use cell culture under conditions of ATM knockdown or ATM inhibition to exam the changes in Sam68 phosphorylation level. Next, we can assess the functional consequences of Sam68 phosphorylation during DDR. Under conditions of ATM knockdown or ATM inhibition, we can test whether the recruitment of Sam68 to the damage sites are dependent on
ATM-mediated phosphorylation, also whether the inducible interaction between PARP1 and Sam68 relies on ATM-mediated phosphorylation of the latter.
102 Reversely, based on the evidence that Sam68 deficiency attenuates ATM activation during DDR, which is crucial for the initiation of cell-cycle checkpoints and
DNA repair factor recruitment (Lee and Paull 2007), it is also worthwhile to examine whether the attenuation is solely through decreased PARP1 catalytic activity or it is partially caused by the direct regulatory role of Sam68 on ATM. To this end, in vitro kinase assay again serves as a definitive method to test whether supplementing recombinant Sam68 can boost ATM activation.
3.6.2 Structural Basis for the Binding of Sam68 to Damaged DNA and Its Interaction with
PARP1
We have visualized the recruitment of Sam68 to DNA damage sites through live cell imaging of microlaser-irradiated cells. To further understand the details of this binding, we could test Sam68’s binding affinity to single-stranded DNA, double-stranded
DNA and the single-stranded/double-stranded DNA junction through electrophoretic mobility shift assays (EMSA); we could also examine the exact localization of Sam68 on damaged DNA by exonuclease assay to test which region is protected by Sam68 from enzymatic digestion. The ultimate method to uncover the details of this binding is to solve the structure of Sam68 when binding to damaged DNA, which can provide insights into the details of the binding and help us identify the crucial residues in Sam68 that are important for the binding.
103 In addition, we have shown that Sam68 binds to the N-terminus of PARP1 (Fig
2.11G), which contains the ZnF1, ZnF2 and ZnF3 of the DNA binding domain and the automodification domain. We can first carefully map the inducible interaction between
Sam68 and PARP1 in DDR by co-IP and pull-down assays. Then we can move forward to solve the structure of the complex containing interacting domains.
With the evidence that Sam68 deficiency, like PARP1 inhibitors, renders hypersensitive in colorectal cancer cell lines and the evidence that Sam68 is upregulated in colorectal tumor and required for colorectal tumorigenesis (Fu, Sun et al. 2016), we could design Sam68 inhibitors that prevent Sam68’s binding to DNA or that hinder the interaction between Sam68 and PARP1, based on the structure, for potentially targeted therapy.
3.7 Conclusion
The identification of more RBPs in DDR through both large-scale proteomic analysis and functional studies on individual protein uncovers a previously unappreciated systemic link between RNA biology and DNA damage repair. Adding these RBPs into different aspects of DNA damage repair unfolds a much broader panorama of DDR in cells. A great deal has been learned about RBPs in DDR in the past few decades, with much more to be discovered in the future. There is a growing need for more comprehensive dissection of the functions and regulations of the RBPs in DDR to
104 better understand DDR and to identify potential therapeutic targets for treating diseases involving dysfunctional DDR, such as various cancers and neurodegenerative diseases.
105 3.8 Figures and Figure Legends
Figure 3.1 Schematic model representation of Sam68 functioning as a signaling molecule in DNA damage repair.
Shown are cellular repair signaling cascades in response to DNA damage in the presence (A) and absence (B) of Sam68.
106 Figure 3.2 Schematic model representation of Sam68 functioning as the master regulator of DDR.
Shown are efficient cellular DDR in the presence of Sam68, resulting in maintenance of genome stability, survival and proliferation.
107 4 Appendix: Sam68 is a Potential Therapeutic Target for Basal Cell
Carcinoma.
4.1 Introduction
Skin cancer is the most prevalent form of all human cancers (Ananthaswamy,
Loughlin et al. 1997). There are three major types of skin cancers, including basal cell carcinoma (BCC), which accounts for almost 80% of all skin cancers, squamous cell carcinoma (SCC), and melanoma (Linares, Zakaria et al. 2015). The pathogenesis of BCC tumors are associated with constitutive activation of the Sonic hedgehog (Shh) signaling pathway both in mouse and in human (Booth 1999). Gli2tg mice, the mouse model for
BCC-like skin tumors, spontaneously develop BCC-like tumors with 100% penetrance, due to overexpression of the Shh signaling downstream transcription factor Gli2 in keratinocytes of epidermal basal layer and hair follicle under the control of a K5 promoter (Grachtchouk, Mo et al. 2000). BCC-like lesions on the ears of Gli2tg mice features consecutively hyperkeratosis (flaking), thickening and hyperpigmentation
(Depianto, Kerns et al. 2010).
We have previously shown that Sam68, a KH domain containing RBP is crucial for colorectal tumorigenesis (Fu, Sun et al. 2016). Our findings demonstrate that Sam68 facilitates tumor cells survival through stimulating PARP1-conferred PARylation during
DDR (Fu, Sun et al. 2016, Sun, Fu et al. 2016). Emerging evidence suggests that elevated
Sam68 expression is associated with aggressive progression and poor prognosis in many
108 types of cancer (Richard, Vogel et al. 2008, Li, Yu et al. 2012, Liao, Liu et al. 2013, Zhang,
Xu et al. 2014, Li, Ngo et al. 2016). Here, we report that Sam68 is upregulated in both human BCC and SCC tumor tissues. The increasing Sam68 protein levels are correlated with BCC-like lesion development in mice. Genetic ablation of Sam68 protein delays
BCC-like skin tumorigenesis in mice, with dampened mitotic activity and reduced blood vessel expansion. Preliminary data from cell culture suggests that Sam68 is important for skin cancer cell survival and tumor malignant transformation.
4.2 Results
4.2.1 Sam68 is Significantly Upregulated in Basal Cell Carcinoma.
First, we performed indirect immunofluorescence staining of human tissue to assess Sam68 expression. Quantification of the fluorescence intensity revealed that Sam68 was significantly upregulated in both basal cell carcinoma samples and squamous cell carcinoma samples, comparing to normal skin tissues (Fig 4.1A-4.1B). To verify whether this finding also applied to the BCC mouse model, we examined Sam68 expression in the
Gli2tg mice. Sam68 expression level was dramatically increased in the epidermis of BCC, comparing to dermis and adjacent non-tumor tissue (Fig 4.1C), and this was further confirmed by IB of protein samples from skin tissue (Fig 4.1D). It has been reported that, on average, onset of lesions occurs at 65 days in male Gli2tg mice (Depianto, Kerns et al.
2010). To investigate the timeline for Sam68 upregulation, we sampled both WT and
109 Gli2tg mice ears at different postnatal time. Sam68 protein level increased gradually from
P60 in Gli2tg mice, whereas no significant change was detected in WT mice, suggesting increasing Sam68 protein levels are correlated with BCC development in mice (Fig 4.1E).
4.2.2 Loss of Sam68 Delays BCC Tumorigenesis.
To assess whether Sam68 is required for BCC tumorigenesis, we monitord BCC- like lesion onset and progression in gender-matched littermates Gli2tg/+Sam68+/-,
Gli2tg/+Sam68-/-, Gli2+/+Sam68+/-, Gli2+/+Sam68-/- mice. In WT mice, ears appeared normal regardless of Sam68 expression, indicating presence of Sam68 itself in skin tissue does not drive tumorigenesis. In the background of Gli2 transgene, tumors arose with 100% penetrance, on average at P78, in the presence of Sam68. In the absence of Sam68, the average age of ear tumor onset was P106, indicating a significant delay (Fig 4.2A-4.2C).
Strikingly, in comparison to Gli2tg/+Sam68+/- mice, which exhibited profound pigmentation
(Fig 4.2A) and hyperplasia (Fig 4.2D-4.2E) with 100% penetrance, 27% of the Gli2tg/+Sam68-
/- mice never developed BCC-like lesions, up to P180 of age (Fig 4.2C).
Elevated proliferation of the tumor cells and exaggerated angiogenesis are essential for the progression of tumor. Indirect immunofluorescence staining of phosphorylated Histone 3 (p-H3), a marker for cells that are in M phase, revealed that loss of Sam68 is associated with a decrease in mitotic activity in the ear tissues of
Gli2tg/+Sam68-/- mice, comparing to Gli2tg/+Sam68+/- mice (Fig 4.2F). Staining of PECAM, a
110 marker for blood vessel, showed that loss of Sam68 reduces blood vessel expansion in the ear tissues of Gli2tg/+Sam68-/- mice, comparing to Gli2tg/+Sam68+/- mice, as well (Fig 4.2F).
Together, these data demonstrated that Sam68 is required for BCC tumorigenesis through promoting tumor cell proliferation and angiogenesis.
4.2.3 Sam68 is crucial for skin tumor cell survival and malignant transformation.
To further probe unto the mechanism through which Sam68 contributes to the growth of BCC-like tumors in mouse skin, we first investigated the role of Sam68 on skin tumor cell survival by comparing total cell number and percentage of early-stage apoptotic cells in A431 cells. In comparison to nonspecific control, Sam68 knockdown in
A431 cells resulted in reduced total cell number and increased apoptosis (Fig 4.3A-4.3C), suggesting Sam68 is required for tumor cell survival. Sam68 knockdown efficiency was confirmed by IB (Fig 4.3D). Next, we assessed the role of Sam68 in tumor cell anchorage independent growth by performing soft agar colony formation assay. Sam68 knockdown in A431 cells significantly decreased both colony number and colony size (Fig 4.3E-4.3G), comparing to nonspecific control, suggesting Sam68 is required for tumor cell malignant transformation.
111 4.3 Figures and Figure Legends
Figure 4.1 Sam68 is upregulated in basal cell carcinoma.
(A) Immunostaining of Sam68 in human BCC, SCC and normal tissue. Basal cell carcinoma (BCC), squamous cell carcinoma (SCC).
(B) Quantification of fluorescence intensity in (A), with BCC and SCC combined as non- melanoma skin cancer (NMSC). Results are represented as mean and s.e.m.
(C) Immunostaining of Sam68 in mouse BCC and adjacent non-tumor tissue. Histone 3 serves as a loading control. epi, epidermis; derm, dermis.
(D) Immunoblot of Sam68 in mouse BCC and adjacent non-tumor tissue.
(E) Immunoblot of Sam68 in WT and Gli2tg/+ mouse ear tissues. β-actin serves as a loading control.
112
113 Figure 4.2 Loss of Sam68 delays basal cell carcinoma tumorigenesis.
(A) Macroscopic images of mouse ears on gender matched littermate Gli2tg/+Sam68+/-,
Gli2tg/+Sam68-/-, Gli2+/+Sam68+/-, Gli2+/+Sam68-/- mice.
(B) Quantification of individual BCC lesion onset in Gli2tg/+Sam68+/- mice (n = 13) and their littermate control Gli2tg/+Sam68-/- mice (n = 11). Results are represented as mean and s.e.m.
(C) Kaplan-Meier analysis of the BCC lesion free survival rate Gli2tg/+Sam68+/- mice (n = 13) and their littermate control Gli2tg/+Sam68-/- mice (n = 11). p = 0.0162, Gehan-Breslow-
Wilcoxon test.
(D) Hematoxylin and eosin (H&E) staining of ears tissue sections from gender-matched littermates Gli2tg/+Sam68+/-, Gli2tg/+Sam68-/-, Gli2+/+Sam68+/-, Gli2+/+Sam68-/- mice.
(E) Quantification of maximum ear tissue thickness as in (D). Results are represented as mean and s.e.m.
(F) Immunostaining of phosphorylated histone H3 (p-H3; green) in mouse ear tissue sections. DNA was stained with DAPI (blue).
(G) Immunostaining of PECAM (green) in mouse ear tissue sections. DNA was stained with DAPI (blue).
114 115 Figure 4.3 Sam68 is required for skin cancer cell survival and malignant transformation.
(A) Representative flow cytometry analysis of Annexin V/PI staining in A431 cells 72 hours post transfection with nonspecific control (si-NC) or Sam68-specific (si-Sam68) small interference RNA.
(B) Quantification of total cell number as in (A).
(C) Quantification of Annexin V+/PI- cell percentage as in (A).
(D) The Sam68 knockdown efficiency was examined by immunoblot (IB), with β-actin as a loading control, in A431 cells at 72 h after siRNA transfection.
(E) Micrographs of A431 cells at two weeks after siRNA transfection forming colonies in soft agar.
(F) Quantification of total colony number per field as in (A). Ten random fields were chosen for each treatment.
(G) Quantification of average colony size per field as in (A). Ten random fields were chosen for each treatment.
116
117 4.4 Discussion
We discovered elevated and aberrant Sam68 expression in both human and Gli2tg mice BCC samples. Our mouse model demonstrated that Sam68 is crucial for BCC-like skin tumorigenesis. Preliminary data from tissue culture suggested that Sam68 plays important roles in tumor cell survival and malignant transformation. Together, the findings that loss of Sam68 delays BCC-like lesion onset and progression, reduces mitotic activity and angiogenesis suggest that Sam68 could serve as a potential novel therapeutic target for BCC. Future work is needed to provide detailed mechanism through which
Sam68 promotes BCC tumorigenesis.
4.5 Material and Methods
Mice. Gender-matched littermates Gli2tg/+Sam68+/-, Gli2tg/+Sam68-/-, Gli2+/+Sam68+/-,
Gli2+/+Sam68-/- mice were produced by breeding of C57BL/6 Gli2tg/+Sam68+/- male mice with
Gli2+/+Sam68+/- female mice and genotyped. Mice were housed in a pathogen-free facility and fed autoclaved food and water ad libitum. All animal experiments were performed in accordance with protocols approved by the Johns Hopkins Institutional Animal Care and
Use Committee.
Cell culture. A431 cells and SIMEK cells were kindly provided by Dr. Pierre Coulombe
(Johns Hopkins University). A431 Cells were cultured in DMEM medium containing 10%
118 SIMEK cells were cultured in flavin adenine dinucleotide (FAD) medium as previously described (Feng and Coulombe 2015).
Antibodies and reagents. Antibodies used were: Sam68 from Santa Cruz Biotechnology;
β-actin from Sigma-Aldrich (St. Louis, MO); p-Histone H3 from Cell Signaling
Technology (Danvers, MA); Histone3 from Abcam (Cambridge, MA); PECAM from
Chemicon. 4', 6-diamidino-2-phenylindole (DAPI) were obtained from Sigma-Aldrich.
Histology and immunohistology. Ears were surgically removed from mice littermates and immersed in optimal cutting temperature media (Tissue-Tek, Elkhart, IN) without fixation, and then frozen in dry ice to preserve the tissue. 5-micron sections were cut using a Microm HM 550 Cryostat (Thermo Scientific), collected on coated slides, fixed in
4% paraformaldehyde, washed with PBS, and processed for Hematoxylin and Eosin
(H&E) staining or indirect immunofluorescence. H&E stained sections were microphotographed to perform histomorphometric analyses and Maximum epithelial thickness measurement, as previously described (Depianto, Kerns et al. 2010). Tissue sections for indirect immunofluorescence were blocked with normal goat serum in PBS.
After incubating with appropriate primary antibodies, sections were stained with Alexa
Fluor-conjugated second antibodies (Invitrogen) and 1 µg/ml of DAPI (Sigma). Stained sections were mounted under a coverslip using Fluoro-gel with Tris Buffer (Electron
Microscopy Sciences, Hatfield, PA) and examined using an Axio Observer fluorescence microscope (Zeiss, Oberkochen, Germany).
Immunoblot. After euthanizing mice, ears were excised under aseptic conditions, and
119 lysed on ice with 0.4 ml of lysis buffer (50 mM Tris-HCl [pH 8.0], 150 mM NaCl, 1% NP-
40 and 0.5% sodium deoxycholate, 1 complete protease inhibitor cocktail [Roche
Applied Science, Indianapolis, IN]) for 30 min, boiled at 95°C for 20min. Then tissues were sonicated and boiled again for 5min. All samples were separated by SDS-PAGE under reduced and denaturing conditions. The resolved protein bands were transferred onto nitrocellulose membranes, probed as described previously, developed by the Super
Signaling system (Thermo Scientific) according to the manufacturer's instructions, and imaged using a FluorChem E System (Protein Simple, Santa Clara, CA).
RNA Interference and transfection. The siRNAs targeting human Sam68 were described previously (Sun, Fu et al. 2016) and transient transfection of siRNA into A431 cells was performed using Lipofectamine 2000 (Life Technologies) according to the manufacturer's instructions.
Flow Cytometry. 0.5–2 × 106 cells were washed twice with cold PBS, resuspended in
1XBinding Buffer (BD Biosciences FITC Annexin V Apoptosis Detection Kit) and stained with FITC Annexin V and PI at room temperature for 15min. Cells were then analyzed on a FACSCalibur (BD Biosciences, San Jose, CA). Events were collected and analyzed with the FlowJo software (Tree Star, Ashland, OR).
Soft agar colony formation assay. 24 hours post RNAi, 10,000 siNC cells or siSam68 cells were mixed with 1.5ml 0.3% agar in normal medium, plated on top of a bottom layer containing 2ml 0.5% agar in normal medium in 6-well plate, and covered with 500ul normal medium. Cells were allowed to grow for 2 weeks during which the top medium
120 was changed every 3 days. Then the whole field was scanned under microscope and colony number was counted and plotted for analysis.
Statistical analysis. All statistical analysis was performed using GraphPad Prism version
6.0 (GraphPad Software, San Diego, CA). The difference between treated and control groups were examined by unpaired Student’s t tests, except Gehan-Breslow-Wilcoxon tests were used for Kaplan-Meier survival curves. Standard errors of means (s.e.m.) were plotted in graphs. n.s. means non-significant difference and significant differences were considered * at p < 0.05; ** at p < 0.01; and *** at p < 0.001.
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