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TRANSCRIPT
Control of Cardiomyocyte Proliferation by p53/MDM2-Regulated microRNAs
by
Shanna Stanley-Hasnain
A thesis submitted in conformity with the requirements for the degree of Masters of Science – Medical Science
Institute of Medical Science, Cardiovascular Collaborative Program University of Toronto
© Copyright by Shanna Stanley-Hasnain 2016
ii
Control of Cardiomyocyte Proliferation by p53/Mdm2-Regulated
microRNAs
Shanna Stanley-Hasnain
Masters of Science – Medical Science
Institute of Medical Science, Cardiovascular Collaborative Program University of Toronto
2016
Abstract
Defining the roadblocks responsible for adult cardiomyocyte cell cycle arrest lies at the core of
developing cardiac regenerative therapies. Inactivation of the p53/MDM2 tumor suppressor
circuitry in the heart caused a significant increase in cardiomyocyte proliferation, through an
upregulation of factors involved in cell cycle re-entry. These factors may be regulated by
microRNAs (miRNAs), in quiescent cardiomyocytes. Therefore, we hypothesized that
inactivation of p53/MDM2-regulated miRNAs could promote the expression of cell cycle
activators and induce proliferation of cardiomyocytes. Comparison of miRNA expression
profiles from cardiac specific p53/MDM2 double knockout (DKO) mouse hearts and wild type
controls revealed 11 miRNAs that were downregulated in the "proliferative" DKO hearts and
enriched for mRNA targets involved in cell cycle regulation. Knockdown of these 11 miRNAs in
neonatal rat cardiomyocytes significantly increased the occurrence of cytokinesis, revealing a
novel subset of p53/MDM2-regulated miRNAs responsible for maintaining cell cycle arrest in
the heart.
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Acknowledgments
Thank you to my beautiful friends and family for their unending support throughout my academic endeavors. Your confidence and belief in me has been my driving force throughout the last 6 years of my life when I began my undergraduate degree at the University of Toronto. I
have a few specific acknowledgments to individuals who have really helped shape the person I am over the course of my life and years studying.
Firstly. I’d like to thank Effie Argyropoulos, my roommate of 5 years, and best friend. She has been there for me every step of the way, through every all-nighter, and every stressful moment, since I began studying in Toronto. I am thankful to her for listening to me practice countless
presentations, and guide me through many important life decisions. She has been my rock, and everything I do in life, I do in hopes of making her proud.
Next, I want to acknowledge Jonah Chevrier, because without him, I would not be the person I am today, or have the love for science that I currently hold. We have been side by side throughout almost our entire post-secondary careers, and have managed to teach each other many
things, support each other and importantly, make some of the best memories I’ll ever have throughout this time. Thank you for the ~1,000s of hours we spent together at Gerstein, you
made them fun. I would not be the person I am today if I had not met you.
Next, I have to recognize Keith Dadson, the post-doc in the Billia Lab, for making every day an absolute treat to come into the lab. Keith was always there to talk and kept me motivated, happy
and in a great mood throughout my days in the lab. I am thankful that I was able to learn a great deal from him through his many years of wisdom and understanding of the field and of research
as a whole. On top of this, I am grateful for his kindness, generosity, and patience. He was always there to lend a helping hand, and to listen. His attitude towards research and life is something I hope to carry with me through the rest of my time. I’m also thankful that he was
around to make sure that I took the time to eat lunch, and for all our trips to Fast Fresh.
Thank you to all my girlfriends back in Ottawa, for always being there for me. I am blessed to know that I will always have them on my side, despite any distance between us. Their successes
push me to be a better person every day, and I am so lucky that we all ended up as friends at De La Salle, because everyone of us made each other a better person. When I look at us now, I am
proud that we have all made it to where we are, together.
I also need to thank Taras Lesiuk, for being the most wonderful distractio n imaginable. Next, are the summer students Amelia Fung and Shelly Chauhan, for always brightening up the lab!
Most importantly, thank you to Joanne Stanley and Sadiq Hasnain, my parents. They are the
reason that I was able to follow my dreams, and the number one reason for where I am today. Nothing would be possible without them. Other important individuals are my step father Klaas Van Weringh and my father’s partner Sasha Sadilova. Thank you for taking care of my family
while I have been away, and for raising me along side of my parents. Your support and love means the world to me.
Lastly, thank you for taking the time to read my thesis thus far.
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Contributions
Dr. Phyllis Billia, my mentor, and supervisor contributed holistically to this project through her undying support, insight and wisdom in my research endeavors and in my life as a whole. She helped to carefully craft a unique and exciting research project that I was passionate and excited
about, and was there to guide me every step of the way throughout my thesis. She supported me throughout all the experiment planning, trouble shooting, talk giving, poster presenting and
scholarship applications, as well as in going to conferences and participating in extracurricular activities. Furthermore, she guided me through the thesis writing process and providing me extremely valuable insight and thorough edits. It is thanks to her that I was able to experience a
well-rounded, inspiring and successful two year Master’s degree. I am eternally grateful that she believed in me, and gave me a shot in her lab, which turned out to be two of the best years of my
life. Phyllis is a role model to me, and much of my efforts in the future will be to make her proud of the scientist that blossomed under her supervision, guidance and mentorship.
Daniela Grothe, our incredible lab manager, taught me everything I needed to know about
working in the lab. She trained me exhaustively in all the techniques that I would require for the experiments in my thesis in an incomparably in-depth manner. Working along her side was a
once in a life time opportunity to learn the practice of meticulous lab bench work at a level that I have never seen before. She taught me the importance of trouble shooting and always using logic to make sure that what you are doing actually makes sense, as opposed to blindly following
protocols. She also taught me that it is always of benefit to try new things, and never to remain stuck in old methods and techniques, because the field of science is always advancing. Most
importantly, she was there for me every step of the way in all the experiments I conducted throughout my thesis, to either help me directly in the lab, answer my questions (there were a lot), or talk out and rationalize the upcoming experiments until I was 100%n confident that I
knew what I was doing. From RNA isolation, to spending hours at the microscope, she was always there to lend a helping hand. Together, we established and worked on many protocols that the lab now uses on a regular basis, including microRNA RT-qPCR, immunoprecipitations
and cell transfections. I am grateful for the weekends she sacrificed to come in and help me with countless cardiomyocyte isolations, especially on the ones where I was away and completely
relying on her for cells to keep my project moving. It is with her support and wisdom that a large majority of my experiments were successful, and that any complications I encountered, we were able to easily fix and surmount. I am forever grateful to everything she has taught me and done
for me over the past two years, and know that there is no way that my thesis could have turned out the way it did without her by my side. Finally, and maybe most importantly, she has been
there for me and a friend and confident through two very important years of my life. I will never forget all the hours we spent together in the lab, singing, laughing, dancing and talking. Her positive attitude and energy made it an absolute joy to work along her side, and made my two
years in the Billia lab fly by (almost too quickly).
Ludger Hauck, the Research Associate in the Billia lab has also been a major support throughout
my Master’s degree. He helped me countless times to plan out experiments, trouble shoot, and understand how to critically analyze my data. His brilliant mind helped guide and expand my project, and I could always count on him when I was stuck and didn’t know in what direction to
turn. His passion for high quality research and thinking outside of the status quo of a given field has shaped the kind of researcher that I now am. I am extremely grateful to him for his wisdom,
and the many hours he sacrificed to help me work through my project and understand my results,
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as well as the weekends he gave up to help with my cardiomyocyte isolations. His patience and ability to trouble shoot with the help of Dani were the factors that allowed us to finally perfect
our cardiomyocyte isolation technique. On top of this, I am extremely thankful for the time he took to read my thesis from front to back, and provide me with extensive suggestions and edits
that largely improved my work. Finally, I am thankful that he believed in me, and in my abilit ies to execute this project. I am overjoyed to see how proud he is in the scientist that I have become over the past two years, thanks to him and the other members of the Billia lab.
My program advisory committee members Dr. Anthony Gramolini and Dr. Vivek Rao, for the support, wisdom and knowledge they have provided with me throughout my Master’s. Together,
they pushed me to be a better critic of my own work and guided my project, ensuring that my goals were feasible. I am forever grateful for the time they have sacrificed to push me as a young scientist and for all their advice and insight that they provided following my PAC meetings in
terms of future experiments, as well as for my thesis.
Finally, thank you to Cedric Manlhiot, for your help with statistics. I greatly appreciate you
taking the time out of your day to set me in the right direction regarding in my statistic calculations for RT-qPCR.
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Table of Contents
Acknowledgments........................................................................................................................... iii
Contributions................................................................................................................................... iv
Table of Contents ............................................................................................................................ vi
List of Tables ................................................................................................................................... x
List of Figures ................................................................................................................................. xi
List of Abbreviations .....................................................................................................................xiii
Chapter 1 Introduction .....................................................................................................................1
1 Literature Review ........................................................................................................................1
1.1 Anatomy...............................................................................................................................1
1.2 Diastolic and Systolic Performance .....................................................................................1
1.3 Cardiomyocyte Structure and Function ...............................................................................2
1.4 Cardiovascular Disease ........................................................................................................4
1.4.1 General Overview ....................................................................................................4
1.4.2 Ischemic Heart Disease ............................................................................................5
1.4.3 Myocardial Infarction ..............................................................................................5
1.4.4 Heart Failure ............................................................................................................7
1.4.5 Ventricular Remodeling ...........................................................................................8
1.4.5.1 Cardiomyocyte Hypertrophy .....................................................................9
1.4.5.2 Scar Formation ..........................................................................................9
1.4.6 Current Treatments for Ischemic Heart Disease and Heart Failure .........................9
1.5 Cell Cycle Regulation ........................................................................................................10
1.5.1 The Cell Cycle .......................................................................................................10
1.5.2 G1 Phase ................................................................................................................12
1.5.3 S Phase ...................................................................................................................13
1.5.4 G2 Phase ................................................................................................................15
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1.5.5 M Phase..................................................................................................................16
1.6 Cardiogenesis and Cardiomyocyte Cell Cycle Regulation ................................................20
1.6.1 Cardiogenesis .........................................................................................................20
1.6.2 Cardiomyocyte Regeneration.................................................................................21
1.7 MicroRNAs ........................................................................................................................22
1.7.1 miRNA Nomenclature ...........................................................................................23
1.7.2 miRNA Biogenesis ................................................................................................23
1.7.3 miRNA Mechanism and Function .........................................................................25
1.7.4 MicroRNAs and the Heart .....................................................................................27
1.7.4.1 MicroRNAs, Cardiac Development and Homeostasis ............................27
1.7.4.2 MicroRNAs and cardiovascular disease..................................................29
1.7.5 MicroRNA Therapeutics........................................................................................31
1.7.5.1 Delivery Methods for miRNA Based Therapies .....................................31
1.7.5.2 miRNA Mimics .......................................................................................32
1.7.5.3 miRNA Antagomirs.................................................................................33
1.7.6 MicroRNAs and the cell cycle ...............................................................................34
1.8 Tumor Suppressors – A Focus on p53 ...............................................................................36
1.8.1 The p53/Mdm2 Tumor Suppressor Circuitry ........................................................38
1.8.2 Canonical p53 Pathways ........................................................................................39
1.8.3 Non-Canonical p53 Pathways ................................................................................40
1.8.4 Post-Translational Regulation of p53 by MDM2 ..................................................41
1.8.5 p53/MDM2 and the Heart ......................................................................................42
1.8.6 p53 and miRNAs....................................................................................................45
Chapter 2 Rational Hypothesis ......................................................................................................50
2 Hypothesis and Thesis Aims .....................................................................................................50
2.1 Hypothesis..........................................................................................................................51
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2.2 Thesis Aims........................................................................................................................51
Chapter 3 Data Chapter..................................................................................................................52
3 Control of Cardiomyocyte Proliferation by p53/MDM2-regulated microRNAs......................52
3.1 Introduction ........................................................................................................................53
3.2 Methods..............................................................................................................................57
3.2.1 MDM2 and p53 conditional mutant mice ..............................................................57
3.2.2 RNA Isolation ........................................................................................................58
3.2.3 Nanostring nCounter miRNA assay.......................................................................59
3.2.4 miRNA RT-qPCR ..................................................................................................59
3.2.5 Western Blot ..........................................................................................................60
3.2.6 Neonatal rat cardiomyocyte and fibroblast isolation and culture ..........................60
3.2.7 miRNA antagomir development ............................................................................60
3.2.8 Neonatal rat cardiomyocyte transfection ...............................................................61
3.2.9 Cell Cycle RT-qPCR Array ...................................................................................61
3.2.10 Immunofluorescence ..............................................................................................61
3.2.11 Statistical Analyses ................................................................................................62
3.3 Results ................................................................................................................................63
3.3.1 The tumor suppressor circuitry p53/MDM2 regulate expression of a unique subset of miRNAs within the heart. .......................................................................63
3.3.2 p53/MDM2-regulated miRNAs are enriched for mRNA gene targets involved in cell cycle progression.........................................................................................67
3.3.3 p53/MDM2-regulated miRNAs are novel cardiomyocyte cell cycle inhibitors ....71
3.3.4 p53/MDM2-regulated miRNAs inhibit cardiomyocyte proliferation by downregulating target genes involved in the progression of all 4 phases of the
cell cycle. ...............................................................................................................78
3.4 Discussion ..........................................................................................................................81
3.4.1 p53 and MDM2 regulate the expression of muscle specific miRNAs in the
heart........................................................................................................................81
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3.4.2 p53 and MDM2 maintain cardiomyocyte quiescence through the regulation of a unique group “anti-proliferative” miRNAs. ........................................................82
3.4.3 p53/MDM2-regulated miRNAs maintain cardiomyocyte cell cycle arrest through the inhibition of multiple cell cycle regulators. ........................................84
3.5 Supplementary Figures and Tables ....................................................................................87
Chapter 4 Conclusion.....................................................................................................................96
4 Conclusion and Future Directions.............................................................................................96
4.1 General Discussion ............................................................................................................96
4.1.1 Thesis summary .....................................................................................................96
4.1.2 The 11 p53/MDM2-regulated miRNAs are well characterized tumor suppressors .............................................................................................................98
4.1.2.1 The miR-30 family ..................................................................................98
4.1.2.2 The let-7 family .......................................................................................98
4.1.2.3 miR-181a .................................................................................................99
4.1.2.4 miR-26b ...................................................................................................99
4.1.2.5 miR-204 ...................................................................................................99
4.1.2.6 miR-149 .................................................................................................100
4.1.2.7 miR-194 .................................................................................................100
4.1.3 Tumor suppressive p53/MDM2-regulated miRNAs may function
synergistically to promote cardiac cell cycle arrest .............................................101
4.1.4 Mechanisms of cardiac regeneration with a therapeutic goal in mind .................102
4.1.4.1 Cell-based cardiac regenerative therapies .............................................102
4.2 Limitations and Future Directions ...................................................................................109
4.3 Overall Conclusion ..........................................................................................................114
5 References ...............................................................................................................................116
Copyright Acknowledgements.....................................................................................................136
Permissions granted for use of external figures: ..............................................................136
x
List of Tables
Table 3.1 - miRNAs significantly downregulated in DKO vs. vehicle control…………………66
Table 3.2 - Final list of p53/MDM2-regulated miRNAs……….……..………………………...70
Supplementary Table 3.1 - Normalized RT-qPCR Ct values for miRNA expression levels
following 11 antagomir transfection……………………………………………………………..95
xi
List of Figures
Figure 1.1 - Anatomy of the cardiac sarcomere…………………………………………………..3
Figure 1.2 - The main molecular proponents of myocardial ischemia reperfusion injury………..7
Figure 1.3 - The major events of the cell cycle are regulated by transitions between CDK/cyclin
complexes…………………………………………………………………………………..……11
Figure 1.4 - Assembly of the pre-replicative and pre-loading complex………………...………14
Figure 1.5 - The main stages of mitosis………………………………………………………....16
Figure 1.6 – Aurora B Localization During Mitosis and Cytokinesis…………………………..19
Figure 1.7 - The Canonical Pathway of miRNA Biogenesis……………………………………25
Figure 1.8 - An overview to cell cycle control by microRNAs………………………………....35
Figure 1.9 - Regulation of the cell cycle by p53-induced miRNAs……………………………..47
Figure 3.1 - Significant miRNA transcriptional changes in the absence of p53 and/or MDM2
create unique miRNA profiles…………………………………………………………………...64
Figure 3.2 - miRNAs downregulated in the DKO are enriched for target genes within the cell
cycle pathway…………………………………………………………………………………....67
Figure 3.3 - Genes within the cell cycle pathway are redundantly targeted by the miRNAs
downregulated in the DKO……………………………………………………………………....69
Figure 3.4 - Antagomirs are specific between miRNA families but redundant within miRNA
families…………………………………………………………………………………………...72
Figure 3.5 - Inhibition of 11 target miRNAs by antagomir cocktail upregulates cardiomyocyte
cytokinesis………………………………………………………………………………………..74
Figure 3.6 - Treatment of cardiomyocytes with cocktail of 11 antagomirs promotes
cardiomyocyte cytokinesis……………………………………………………………………….77
Figure 3.7 - Inhibition of 11 target miRNAs causes upregulation of many miRNA target genes
within the cell cycle pathway…………………………………………………………………....80
Supplemental Figure 3.1 - Validation of cardiac specific MDM2 and p53 knock-down
following Tamoxifen injection …………………………………………………………………..87
Supplemental Figure 3.2. Biological pathway analysis for the 11 p53/MDM2-regulated
miRNAs………………………………………………………………………………………….88
Supplemental Figure 3.3 - 11 miRNAs hits are downregulated in cardiac fibroblasts………...89
Supplemental Figure 3.4 - Efficient transfection of neonatal rat cardiomyocytes with red
xii
fluorescent positive control……………………………………………………………………...90
Supplemental Figure 3.5 - Raw count data for cytokinesis events following antagomir
treatment………………………………………………………………………………………...91
Supplemental Figure 3.6 - Average Cytokinetic events and average RFU show similar %
change trends in transfected cardiomyocytes…………………………………………………...92
Supplemental Figure 3.7 - RT-qPCR for cardiomyocyte housekeeping genes………………..93
Supplemental Figure 3.8 - MirSystem miRNA target gene biological pathway ranking score.94
xiii
List of Abbreviations
ACE Angiotensin-converting enzyme
AGO Argonaute
AIP1 Actin-interacting protein 1
AKT AKT serine/threonine kinase
AMPK 5' AMP-activated protein kinase
ANOVA Analysis of variance
APAF1 Apoptotic peptidase activating factor 1
APC Anaphase promoting complex
APC Adenomatous polyposis coli
ARF Alternate reading frame protein
ATM Ataxia telangiectasia mutated
ATP Adenosine triphosphate
APAF1 Apoptotic peptidase activating factor 1
BCL2 B-cell lymphoma 2
BIN1 Bridging integrator 1
BRCA1 Breast cancer 1
BRMS Breast cancer metastasis suppressor 1
BAX BCL2 Associated X Protein
CDK Cyclin-dependent kinase
CKI Cyclin-dependent kinase inhibitor
C-kit KIT Proto-Oncogene Receptor Tyrosine Kinase
CMG Cdc45-MCM-GINS
Ca2+ Calcium
CDC Cell division cycle
xiv
CDH1 Cadherin 1
CRISPR Clustered regularly interspaced short palindromic repeats
DAPI 4',6-diamidino-2-phenylindole
DDC2 DNA damage checkpoint protein 2
DDK Dbf4-dependent Cdc7 kinase
DDX5 DEAD-Box helicase 5
DGCR8 DiGeorge Syndrome critical region gene 8
DKO Double knockout
DR5 Death receptor 5
DP-1 E2f dimerization partner 1 / Transcription factor Dp-1
Drosha Drosha ribonuclease III
MED23 Mediator complex subunit 23
MEF2A Myocyte enhancer factor 2A
MEF2C Myocyte enhancer factor 2C
DBF4 Dbf4 Zinc finger
DPB11 DNA replication regulator Dpb11
eIf4E Eukaryotic translation initiation factor 4E
eIF6 Eukaryotic translation initiation factor 6
E2F E2 Factor
EF Ejection fraction
ERBB2/4 Erb-b2 receptor tyrosine kinase 2/4
ERK Extracellular signal-regulated kinase
EDE1 EH domain-containing and endocytosis protein 1
EI24 Etoposide induced 2.4/Autophagy associated transmembrane protein
FA Franconi anemia related tumor suppressor
FGF Fibroblast growth factor
xv
G1 Gap phase 1
G2 Gap phase 2
GINS Go Ichi Nii and San complex
GADD45A Growth arrest and DNA damage inducible alpha
GATA4 GATA Binding Protein 4
GlUT1/4 Glucose transporter 1/4
GTP Guanosine-5'-triphosphate
HIF-1 Hypoxia-inducible factor 1-alpha
INN1 Ingression protein 1
iPSC Induced pluripotent stem cell
ISL1 ISL LIM Homeobox 1
IRS2 Insulin receptor substrate 2
KCND2 Potassium voltage-gated channel subfamily D member 2
Ki-67 Marker of proliferation Ki-67
KRAS KRAS proto-oncogene GTPase
LNA Locked nucleic acid
LV Left ventricle
MAD2L1 MAD2 mitotic arrest deficient- like 1
MAPK Mitogen-activated protein kinase
MLH1 MutL Homolog 1
MPTP Mitochondrial permeability transition pore
MSH2 MutS Homolog 2
Mcm Tamoxifen inducible Cre recombinase (MerCreMer)
MCM2-7 Minichromosome maintenance proteins 2-7
MDM2 Murine double minute-2
Mdm2KO Murine double minute-2 knockout
xvi
MDM4 Mouse double minute 4 / Mdm2-like p53-binding protein
miRNA microRNA
mTOR Mammalian/mechanistic target of rapamycin
MULE MCL-1 ubiquitin ligase E3
MYC V-Myc avian myelocytomatosis viral oncogene homolog
Na+ Sodium
NBS1 Nisbrin
NFAT Nuclear factor of activated T-cells
NPT Nuclear protein, ataxia-telangiectasia locus
NET1 Neuroepithelial cell transforming 1
NKX2.5 NK2 homeobox 5
NOXA Phorbol-12-Myristate-13-Acetate-Induced protein 1
NRG1 Neuregulin 1
OACIS Osaka Acute Coronary Insufficiency Study
PCNA Proliferating cell nuclear antigen
PGC1β PPARG coactivator 1 beta
PIK3CB Phosphatidylinositol-4,5-bisphosphate 3-kinase catalytic subunit beta
PI3K Phosphatidylinositide 3-kinase
PTEN Phosphatase and tensin homolog
PERP TP53 apoptosis effector
PIDD1 P53-Induced death domain protein 1
PiH3 Phosphorylated histone H3
PTPRV Protein tyrosine phosphatase, receptor type V, Pseudogene
PUMA P53 up-regulated modulator of apoptosis
p107 Retinoblastoma-like protein 1
p130 Retinoblastoma-like protein 2
xvii
p21Cip1 Cyclin-dependent kinase inhibitor 1A
p27Kip1 Cyclin-dependent kinase inhibitor 1B
p38 MAPK p38 mitogen-activated protein kinase
p53AIP1 Tumor protein P53 regulated apoptosis inducing protein 1
p53KO p53 knockout
PCR Polymerase chain reaction
RAB22A RAB22A, member RAS oncogene family
RAD21 RAD21 cohesin complex component
RalA RALA Ras like proto-oncogene A
RB Retinoblastoma
RELA RELA proto-oncogene, NF-kB subunit
RISC RNA-induced silencing complex
RIZ1 Retinoblastoma protein-interacting zinc finger protein
RFU Relative fluorescence units
RhoA Ras homolog family member A
ROCK Rho associated coiled-coil containing protein kinase 1
SA1/STAG1 Stromal antigen 1
SA2/STAG2 Stromal antigen 2
SCA1 Ataxin-1
SERCA2 ATPase sarcoplasmic/endoplasmic reticulum Ca2+ transporting 2
SIRT1 Sirtuin 1
SSSK2 S-phase kinase-associated protein 2, E3 ubiquitin protein ligase
SNAI1 Snail family transcriptional repressor 1
SOX4 SRY-box 4
SR Sarcoplasmic reticulum
SLD2/3/7 DNA replication regulator Sld2/3/7
xviii
SMC1/SMC3 Structural maintenance of chromosomes 1/3
TAC Trans-aortic constriction
TBX5 T-box protein 5
TIGAR TP53 Induced glycolysis regulatory phosphatase
TIMP1 Tissue inhibitor of metalloproteinases 1
TRAF6 Tumor necrosis factor receptor associated factor 6
Tam 4-Hydroxytamoxifen
UTR Un-translated region
WNT Wingless-type MMTV integration site family
XPC XPC complex subunit, DNA damage recognition and repair factor
YAP1 Yes associated protein 1
YWHAS tyrosine 3-monooxygenase/tryptophan 5-monooxygenase activation protein sigma
/ Stratifin
1
Chapter 1
Introduction
1 Literature Review
1.1 Anatomy
The structure and function of the heart, like in any organ, are inseparable, both in health and
disease. Accordingly, Kresh and Armour advocated that the “heart should be considered as self-
regulating functional system, which is greater than the sum of its constitutive parts”1. This infers
that the loss of structural integrity in one area of the heart can have devastating effects on heart
function as a whole.
The heart pumps on average 5 L of blood a minute2, requiring tight synergy of its electrical and
mechanical function. It is composed of four chambers; the smaller left and right atria, which
receive the incoming blood, and the larger left and right ventricles, which send the blood to be
oxygenated by the lungs, and to the rest of the body, respective ly. The ventricular myocardial
fibers are organized in a helicoid structure that form two spiral turns, known as the ventricular
myocardial band3. It is this configuration that allows the heart to perform the dual function of
both ejection and suctioning blood3. In other words, the intricate organization of the heart is
fundamental for its proper role as a muscular pump that functions to supply oxygen and
metabolites to the peripheral tissues of the body. This tightly regulated process of contraction
and relaxation that drive blood flow throughout the body is called the cardiac cycle. Diastole is
the portion of the cardiac cycle where the heart refills with blood following systole, the portion
of the cycle primarily involved in contraction.
1.2 Diastolic and Systolic Performance
Systolic performance is the ability of the ventricles to empty. This is influenced by myocardial
contractile function, ventricular load, and ventricular configuration4. As the systolic performance
of the left ventricle (LV) is required for supplementing the entire body with blood, it is generally
of main focus. The ability of the LV to empty can be calculated as the LV emptying fraction or
ejection fraction (stroke volume divided by end-diastolic volume).4 Consequently, impaired LV
emptying, indicative of systolic dysfunction, can be measured by a decrease in ejection fraction
2
(<50%)5. When studying murine cardiac contractility however, fractional shortening, which
determines the change in ventricular diameter between the contracted and relaxed state (%
Fractional Shortening = LV end-diastolic diameter – LV end-systolic diameter / LV end-diastolic
diameter x 100) is measured instead of ejection fraction6. A fractional shortening less than 30%
is considered impaired diastolic performance6.
As an effective pump, the ventricles must not only eject but also fill with blood. Effectively,
normal LV diastolic function is defined as its sufficient filling to produce a cardiac output
proportionate to the body’s requirements4. Thus, to maintain an adequate cardiac output when
LV ejection fraction is decreased, a larger end-diastolic volume is required, indicating that the
amount of filling required is dependent on LV systolic performance5. The LV fills in diastole in
response to the pressure gradient from the left atrium to the LV. This occurs two times during the
cardiac cycle: early in diastole after mitral valve opening and late in diastole during atrial systole.
Most of LV filling occurs early in diastole, and less than 25% of the LV stroke volume enters the
LV during atrial systole. The rate of early LV filling pressure is determined by two factors: the
rate of LV relaxation and left atrial pressure at the time of mitral valve opening4.
1.3 Cardiomyocyte Structure and Function
Cardiomyocytes, the contracting functional cellular unit of the heart, occupy an average of 70%
of the ventricular wall under normal physiological conditions, making them the major structural
component of the myocardium7. In the adult, 80% of these long ellipsoidal cells are binucleated,
and are interconnected by gap junctions to allow for coordinated contractile activity8,9,10.
Cardiomyocytes also form close interactions with the extracellular matrix, primarily composed
of fibroblasts, endothelial cells and vascular smooth muscle cells to form a complex 3D network
of cells9.
Intracellularly, the sarcomere forms the basic contractile unit of the cardiomyocyte, whereby
repeating molecules of actin (thin filaments) and myosin (thick filaments) are the building blocks
of this structure (Figure 1.1)9. The sarcomere itself consists of ~20 proteins, but forms
connections with many others linking the myocytes and their extracellular matrix to regulate
contractility9. One sarcomere unit is defined as a segment between two Z-discs (anchoring plane
that cross-links thin filaments from opposing sarcomere halves through a lattice formed by
3
α-actinin and many other proteins)9. The thick filaments sit in between two thin filaments, in the
middle of the sarcomere, and are linked to the Z-discs by the protein titin11. The Z-disc is thought
to be crucial in facilitating cardiomyocyte response to mechanical stress by acting as the liaison
between the sarcomeres and the extracellular matrix.
At the level of the individual cardiomyocyte, contraction is coordinated by the flux of calcium
(Ca2+) into and out of the cell and the sarcoplasmic reticulum through spatially defined ion
channels and exchangers12. Systole will occur when an action potential is fired from pacemaker
cells within sinoatrial node of the heart12. These specialized non-contractile cardiomyocytes
generate repetitive action potentials and dictate the heart rate12. The action potential will first
spread throughout the atria and then to the ventricles leading to depolarization of the
cardiomyocytes12. Depolarization causes voltage gated Ca2+ channels within the sarcolemma to
open and allow influx of Ca2+ into the cytoplasm12. The build-up of Ca2+ causes ryanodine
receptors within the sarcoplasmic reticulum (SR) to release more Ca2+ from the SR into the
cytoplasm (Ca2+ induced Ca2+ release)11. Cytoplasmic Ca2+ will interact with troponin C of the
troponin complex (major thin filament proteins), resulting in a conformational change in the
Figure 1.1 Anatomy of the cardiac sarcomere9. (A) Diagram of the basic organization of the sarcomere.
(B) Representation of the major proteins of the sarcomere. Originally published by Harvey, P. A. &
Leinwand, L. A. J. Cell Biol. 194, 355–65 (2011)9. Used with permission of the author and publisher.
4
position of tropomyosin on the thin filament12. This conformational change reveals sites where
actin and myosin filaments can interact to form cross-bridges, and results in the shortening of the
myofilaments and force generation12. Diastole occurs when Ca2+ is removed from the cytosol
through SR re-uptake by the Ca+ transporter (SERCA2) and extrusion through the sarcolemma
by the Na+/Ca2+ exchanger12. Removal of Ca2+ causes tropomyosin to resume its original
configuration resulting in cross bridge detachment11. Coordination of this response between
individual cardiomyocytes to allow for synchronized contraction is dependent on the cardiac
conduction system and direct electrical contact between the myocytes11. Highlighting the
importance of the precise regulation of the cardiac cycle, dysfunction or mutation of almost any
protein involved in this process has been shown to induce or increase the risk of heart failure.
1.4 Cardiovascular Disease
1.4.1 General Overview
Cardiovascular diseases are the leading cause of death worldwide despite many advances in
pharmaceutical treatments and surgical procedures13. Cardiovascular disease is an overarching
term used to describe groups of ailments of the heart or blood vessels. The most common
cardiovascular diseases are: hypertensive heart disease caused by increased blood pressure13,
ischemic heart disease elicited by the narrowing of the coronary arteries14 and cerebrovascular
disease - pertaining to blockages of the blood vessels in the brain15. Cardiovascular disease poses
a significant burden on health care systems globally as they are the primary cause of death.
Importantly, it is known that adopting healthy life style changes such as smoking cessation,
maintaining a balanced diet and increasing physical activity, all of which help to lower blood
pressure (the primary risk factor for cardiovascular disease) on top of their own individual
benefits, can postpone the onset of heart disease by up to 14 years16. It was estimated that in
2011 one person died every 7 minutes from heart disease or stroke (66,000 Canadians) and that
1.3 million were living with heart disease16. This costs the Canadian economy more than $20.9
billion a year, accounting for a significant burden on health care services, lost wages and
decreased productivity16.
5
1.4.2 Ischemic Heart Disease
Ischemic heart disease (otherwise known as coronary artery disease) is a major public health
problem, as it is the most common cause of death in most western countries17. Ischemic heart
disease manifests itself as ischemia as a result of restricted blood supply to the heart tissue
followed by angina (substernal pains/discomfort caused by reduced blood flow to the heart). This
can lead to myocardial infarction, arrhythmias, heart failure, and sudden death if left untreated17.
The most common cause of ischemic heart disease is atherosclerosis, a condition in which
plaques of fatty material are deposited on the inner walls of arteries. Build-up of atherosclerotic
plaques in the coronary artery system can lead to reduced blood flow and, consequently, reduced
oxygen transportation to areas of the heart. The mismatch between supply and demand for
oxygen results in myocardial ischemia18. Chronic oxygen supply-demand imbalance will have
important myocardial consequences over time. At the cellular level, cardiac remodeling
(hypertrophy), and cardiomyocyte autophagy (catabolic degradation of organelles and
macromolecules within the cell) are known to occur during chronic ischemic heart disease19,20.
Acutely, if an atherosclerotic plaque ruptures, exposure of its fatty core provides a potent
substrate for activation of the coagulation cascade21. Coagulation cascade activation leads to the
formation of a platelet-rich thrombus (blood clot) which can grow to the point of vessel
occlusion21. Alternatively, a thromboembolism may occur, whereby the blood clot breaks loose
from the vessel and travels through the circulatory system, risking occlusion of a distal blood
vessel21. The result of complete coronary artery occlusion is acute myocardial infarction21.
1.4.3 Myocardial Infarction
Myocardial infarction afflicts 7.6 million Americans each year, increasing an individual’s
susceptibility to a subsequent myocardial infarction, and often precedes adverse cardiac
remodeling, hypertrophy and heart failure22. A myocardial infarction occurs when blood flow to
an area of the heart is reduced to a critical level whereby normal cellular function can no longer
occur due to lack of oxygen23. A series of biochemical changes in the myocardium ensue with
lack of oxygen; halted oxidative phosphorylation leading to mitochondrial membrane
depolarization, ATP depletion, and subsequently, inhibition of cardiomyocyte contractile
function23. Cellular metabolism switches to anaerobic glycolysis, resulting in a buildup of lactate
6
and a reduction in intracellular pH23. The acidic intracellular environment eventually leads to
Na+ and Ca2+ overload within the cell23. If blood flow is not restored (reperfusion), and
ischemia persists for a prolonged period of time, irreparable myocyte damage and/or cell death
can occur23. Coronary artery occlusion due to a thrombus superimposed on a ruptured or unstable
atherosclerotic plaque, is the most common cause for myocardial infarction23. Six primary risk
factors for myocardial infarction and coronary artery disease have been identified:
hyperlipidemia, diabetes mellitus, hypertension, tobacco use, male gender, and family history of
atherosclerosis.
Cell death primarily occurs in the area of the heart most distal to the blocked arterial blood
supply, the endocardium24. Over the duration of the occlusion, the area of cell death will enlarge
into the myocardium and finally the epicardium24. The magnitude of a myocardial infarction is
defined by the extent of myocardial cell death24. Removing the occlusion and restoring blood
flow to the ischemic area of the heart as quickly as possible is essential to salvage the viable
myocardium24. However, reperfusion itself can also induce cardiomyocyte death, known as
myocardial-reperfusion injury23. This significantly reduces the efficiency of reperfusion as a
treatment for myocardial infarction, making myocardial reperfusion injury an important
therapeutic target. The major mediators of reperfusion injury are oxidative stress, intracellular
Ca2+ overload, rapid pH restoration causing hypercontraction (reperfusion with an acidic buffer
in an animal model of ischemia reperfusion injury can reduce myocardial infarction size25),
mitochondrial permeability transition pore (MPTP) opening (blocking MPTP opening during
reperfusion is associated with greater post ischemic recovery26) and inflammation23 (Figure 1.2).
All of these stimulators of myocardial reperfusion injury seem to occur within the first few
minutes following reperfusion23. However, processes such as apoptosis and inflammation
initiated at this time, as well as during ischemia, will continue to occur over several hours or
even days following reperfusion23. Halting these processes to minimize the extent of
cardiomyocyte cell death is another important therapeutic option to be considered, as the extent
of cardiomyocyte cell death during ischemia and reperfusion is an important risk factor for the
development of heart failure following myocardial infarction. The severity of cardiomyocyte
loss, and lack of regenerative capacity of the heart is highlighted by the fact that 33% of
myocardial infarction survivors will develop heart failure over time24.
7
.
1.4.4 Heart Failure
Heart failure, a disease affecting over 5 million North Americans, occurs when the heart can no
longer generate adequate contractile force to sustain the required cardiac output of an individual
at normal filling pressures8. It is the culminating pathway for numerous diseases that affect the
heart. Irreversible damage to the heart muscle caused by myocardial infarction is the most
common risk factor for heart failure, followed by hypertension27. Furthermore, the Framingham
Heart Study suggests that coronary artery disease is the most common cause of heart failure,
albeit coronary artery disease comes often hand in hand with hypertension and elevated lipids27.
pC
Figure 1.2 The main molecular proponents of myocardial ischemia reperfusion injury23. During acute
myocardial infarction, the cell metabolism switches to anaerobic respiration, leading to the buildup of lactate and
a drop in intracellular pH. The Na+-H+ exchanger will consequently extrude H+, resulting in intracellular
Na+ overload. This activates the Na+-Ca2+ exchanger to function in reverse to extrude Na+ and leads to intracellular
Ca2+ overload. During reperfusion, the electron transport chain is reactivated, generating ROS, which mediates
myocardial reperfusion injury by inducing the opening of the MPTP, acting as a neutrophil chemoattractant, and
mediating dysfunction of the sarcoplasmic reticulum. Reperfusion and reactivation of the Na+-H+ exchanger result
in washout of lactic acid, resulting in the rapid restoration of physiological pH, which releases the inhibitory effect
on MPTP opening and cardiomyocyte contraction. The restoration of the mitochondrial membrane potential drives
calcium into the mitochondria, which can also induce MPTP opening. Originally published by Hausenloy, D. J. &
Yellon, D. M. J. Clin. Invest. 123, 92–100 (2013)23. Used with permission of the author and publisher.
8
Heart failure is a clinical syndrome that can arise from diverse causes, and is a results of a
complex interplay of structural and functional abnormalities. As such, heart failure has been
described as a cardio-renal disorder causing excessive salt and water retention, a hemodynamic
disorder with peripheral vasoconstriction and reduced cardiac output, a hormonal disorder with
the activation of the renin-angiotensin-aldosterone system and adrenergic nervous system, an
inflammatory syndrome characterized by increased circulating and local inflammatory cytokines,
or a direct myocardial disorder instigated by injury to the heart followed by pathological
ventricular remodeling24. Yet, these factors are not mutually exclusive and often many of them
combined cause the onset and progression of heart failure28. The causative injury can either be
acute and obvious (e.g. acute myocardial infarction), or stealthy and asymptomatic (e.g. chronic
hypertension)27. Following this, a series of compensatory mechanisms are initiated, but
eventually these mechanisms will be detrimental to the heart, resulting in heart failiure28.
1.4.5 Ventricular Remodeling
Left ventricular remodeling is a process by which neurohormonal, mechanical and genetic
factors alter ventricular function, shape and size29. This process occurs following numerous types
of injury and can continue with the onset and progression of heart failure. It is an adaptive
process during development, but becomes pathological following injuries such as myocardial
infarction, cardiomyopathy, hypertension or valvular disease30. As myocardial infarction is the
injury of focus in this thesis, ventricular remodeling will be overviewed in this context.
The acute loss of myocardium following myocardial infarction causes a sudden increase in
cardiac loading conditions which produce a unique pattern of cardiac remodeling in the infarct
border zone as well as in the distant non-infarcted myocardium30. This increase in cardiac
loading triggers a cascade of intracellular signaling pathways that initiate reparative changes
such as hypertrophy, dilatation, and collagen deposition to form a scar30. Injury caused by
myocardial infarction and reperfusion also result in the migration of immune cells such as
macrophages and neutrophils into the infarct zone30. These cells will instigate intracellular
inflammatory signaling and neurohormonal activation, both of which play an important role in
remodeling30. The remodeling process has been divided into an early (within 72 hours) phase
9
characterized by infarct expansion, and late (beyond 72 hours) phase associated with more global
ventricular dilatation and hypertrophy29,30.
1.4.5.1 Cardiomyocyte Hypertrophy
Cardiomyocyte hypertrophy is initiated during early remodeling and continues throughout the
late remodeling process. It can be visualized by microscopy, whereby up to a 70% increase in
myocyte size has been noted30. It is instigated by the neurohormonal activation and
cardiomyocyte stretching that occurs during the remodeling process. Primarily, it functions as an
adaptive process to offset increased load, and compensate for the lost cardiomyocytes, but
ultimately cardiomyocytes hypertrophy can become detrimental if not halted30. The mitogen-
activated protein kinase (MAPK) cascade activation via diverse growth factors (insulin- like
growth factor, fibroblast growth factor, epidermal growth factor, insulin, platelet derived growth
factor, etc.) is the major molecular executor of transcriptional and morphological changes that
occur during cardiomyocytes hypertrophy30.
1.4.5.2 Scar Formation
Myofibroblasts are responsible for reconstruction of the collagen network and fibrotic scar post-
myocardial infarction30. This reparative process is triggered by many stimuli including cytokines
released from necrotic/apoptotic or injured cardiomyocytes (such as transforming growth factor-
β1), hormonal activity (Angiotensin II), fibroblast growth factor (FGF), nitric oxide release etc30.
Angiotensin II released by myofibroblasts stimulates aldosterone production which subsequently
stimulates the transcription of collagen type I and III. Several weeks following myocardial
infarction, a stable fibrotic structure, composed primarily of collagen type I and type III, will
have formed in the infarcted zone31. Accordingly, this process can be blocked by angiotensin II
receptor blockade32.
1.4.6 Current Treatments for Ischemic Heart Disease and Heart Failure
Due to the complex mechanisms and myriad of causes underlying heart failure, and
cardiovascular disease as a whole, the development of effective treatments has been challenging.
Currently, pharmaceutical and surgical options are available to manage symptoms of ischemic
heart disease and heart failure, in addition to lifestyle modifications. However, no cures are
10
available. Management of ischemic heart disease is largely through symptom relief (angina), and
lifestyle modifications such as increased physical activity, improved diet and quitting smoking29.
Surgical interventions such as angioplasty/stents or coronary artery bypass grafts are common if
plaque buildup begins to significantly restrict coronary blood flow, heightening risk for acute
myocardial infarction29. As for heart failure, angiotensin converting enzyme (ACE)-inhibitor
treatment to inhibit the renin-angiotensin-aldosterone pathway and reduce the work load of the
heart is one of the first steps in patients with heart failure29. The major goal is to minimize the
process of maladaptive remodeling. As heart failure progresses, β-blocker treatment, aldosterone
antagonists, and ionotropes can be added to the treatment regimen29. Patients with poor
prognosis despite optimal medical therapy will likely be considered for candidacy for heart
transplantation33. However, it is important to note that even heart transplantation is not a cure.
One year survival rates following heart transplantation are 90%, and the median length of
survival following transplantation is 11 years34. For selected patients who are too ill to wait for a
heart donor, ventricular assist devices are often utilized as a life-saving intermediate. These
devices are a mechanical option that serve to unload the failing heart and help maintain
perfusion to critical organs35. Heart transplantation is also inaccessible to many patients due to
donor organ unavailability, long wait lists and strict patient candidacy requirements.
Consequently, with the improvement in ventricular assist device technology over the past
decade, this therapy has shifted from merely a bridge to transplant, to providing lifetime support
for certain patients with end-stage heart failure35, as destination therapy. Despite major advances
in the field of transplantation and implanted mechanical devices, regenerative options to mitigate
cardiomyocyte loss or repair damaged cardiac tissue remains a sought after goal in cardiology.
Such options would strive to prevent the development of heart failure, and reduce the current
excessive demand for heart transplant.
1.5 Cell Cycle Regulation
1.5.1 The Cell Cycle
The cell cycle is the universal process by which cells reproduce, and lies at the core of growth,
development and tissue regeneration of all living organisms36. It consists of two main stages,
DNA synthesis (S phase), and chromosomal segregation/mitosis (M phase), interspersed by
quality control gap phases (G1 and G2 phase)37. These events culminate at the physical division
11
of the cell into two daughter cells (cytokinesis) in the order G1, S, G2, M, cytokinesis37. Mitosis
can further be subdivided into five discrete phases: prophase, prometaphase, metaphase,
anaphase and telophase37.
DNA replication and mitosis are tightly regulated by, and dependent on, an interplay of many
cell cycle proteins, some of the most important being cyclin dependent kinases (CDKs) and
cyclins38. CDKs are serine-threonine-specific kinases and their activity and substrate specificity
is controlled by diverse cyclins, which serve as the regulatory subunit for CDKs38. Binding of
cyclins to CDKs promotes a conformational change in the kinase, allowing it to bind to ATP and
to their phosphorylation substrate. This interaction occurs in a specific orientation that facilitates
the transfer of the terminal phosphate group of ATP to the target serine or threonine residue of
the bound substrate37. The involvement of these proteins in the cell cycle was first illustrated in
yeast, in which a single CDK could promote the transition between different phases of the cell
cycle, based on which cyclin it was associated with38,39. The analogy was accordingly made that
CDKs act as the engine which drives cell cycle progression, and cyclins serve as the gears
changed between each phase to assist in the transition. As in any vehicle, brakes are essential to
maintain control. In this case, cyclin dependent inhibitors (CKIs) are the brakes of the cell cycle,
which regulate the kinase activity of CDK/cyclin complexes, and can halt cell cycle progression
under unfavorable conditions38. Tightly regulated oscillations between levels and activities of
CDK/cyclin complexes within the cell are the foundation upon which cell cycle progression is
built (Figure 1.3).
Figure 1.3. The major events of the cell cycle are regulated by transitions between CDK/cyclin complexes 37.
G1-cyclins (D type cyclins) complexed with CDKs are required to initiate the cell cycle and activate B-type-
cyclin-CDK activity. Low levels of B-type-cyclin-Cdk2 activity are sufficient to trigger S phase. CDK1 activation
and high levels of B-type-cyclin-CDK1 triggers mitosis and activates the anaphase promoting complex, triggering
anaphase and forming a negative feedback loop that in turn inhibits CDK1 activity. CDK1 inactivation allows
exit from mitosis and re-entry into G1 phase. Originally published by Rhind, N. & Russell, P. Cold Spring Harb.
Perspect. Biol. 4, a005942 (2012)37. Used with permission of the author and publisher.
12
1.5.2 G1 Phase
In the first gap phase (G1 phase), the longest phase of the cell cycle, cells grow and prepare for
the onset of DNA synthesis40. Importantly, it is during this phase that the cell commits to
entering into a new cell cycle41. Specifically, it is a checkpoint late in the G1 phase called the
restriction point that the cell must pass in the presence of the appropriate growth factors, to
proceed into the next stages of the cycle42,43. Conversely, if the appropriate growth factors are
not present, the cell cycle will arrest at the restriction point, and entre the quiescent G0 phase42.
The tumor suppressor protein retinoblastoma (RB) is a potent inhibitor of the transition from G1
to S phase, preventing the initiation of DNA replication43. RB is found within a family of tumor
suppressor proteins including retinoblastoma-like protein 1 (p107) and retinoblastoma-like
protein 2 (p130), all of which are fundamental in the regulation of cell cycle and proliferat ion41.
RB is a nuclear phosphoprotein that can be phosphorylated by diverse CDK/cyclin complexes,
and is active when found in a hypophosphorylated state. In this state, early in the G1 phase, RB
can bind and inhibit the function of E2F transcription activators (E2F1, E2F2, E2F3A).
Furthermore, it recruits histone deacetylases to E2F regulated promoters which epigenetically
alter their conformation, rendering them inaccessible to the transcriptional machinery41,43.
Conversely, E2F transcriptional repressors (E2F4 and E2F5) are bound by p130 and p107, which
allows their nuclear localization where they can inhibit transcription of S phase related genes43.
RB confines the transcriptional activity E2F1-3 until the later stages of the G1 phase because this
family of proteins activates the transcription of genes that promote the transition from the G1
phase to the S phase. In the mid G1 phase, RB becomes hyperphosphorylated by CDK/cyclin
complexes (CDK4/6 and cyclin D1, D2 and D3), rendering it inactive until M phase41. This
allows activator E2Fs to induce transcription of many genes required for DNA replication such
as the E2f dimerization partner 1 (DP-1), DNA polymerase-α, proliferating cell nuclear antigen
(PCNA, a DNA clamp that functions with DNA polymerase), the DNA cutting enzyme
topoisomerase type 1, as well as genes essential for cell cycle progression like the proto-
oncogene avian myelocytomatosis viral oncogene homolog (c-MYC), cyclin D1, cyclin A, cyclin
E, CDK2, CDK1 (also known as CDC2), and CDC2541. D-type cyclins will reach their peak
expression level by late G1 phase. The degradation of cyclin D1 before this time point disables
the cell from progressing into S phase41, which highlights the requirement of E2F transcriptional
13
activity for cell cycle progression, enabled by CDK-cyclin D1 dependent inhibition of RB. While
D-type cyclins are important for the progression of the G1 phase (marked by the switch from RB
activation to inactivation) cyclin Es and cyclin As are thought to be required at the G1 to S
transition40,41. Cyclin E’s expression peaks at this transition, and is subsequently degraded during
S phase40,41. Cyclin A at this time, will bind and activate CDK2, triggering S phase initiation40,41.
Its expression begins at late G1 phase following as previously mentioned and accumulates
throughout S and G2 phase, then disappears during M phase40,41.
1.5.3 S Phase
In eukaryotic cells, replication propagates bidirectionally from thousands of sites along the
genome (origins of replication)44. These origins of replication are first recognized by the origin
of replication complex (ORC1-6) during G1 phase44. Recognition of these sites by the ORCs will
localize the loading of the replicative DNA helicase minichromosome maintenance proteins 2-7
Complex (MCM2-7) to the origins44. Loading of the MCM2-7 also requires the proteins cell
division cycle 6 (CDC6) and chromatin licensing and DNA replication factor 1 (CDT1), which
are associated to the ORC complex and MCM2-7 respectively44. This event is often referred to as
“Origin Licensing”, and together the assembled proteins form the pre-replicative complex44.
During S phase, the MCM2-7 DNA helicase must become activated, which is followed by the
assembly of the replisome. The replisome consists of this helicase, which unwinds DNA, DNA
polymerases, which synthesize the leading and lagging strands of DNA, and a primase, which
facilitates the initiation of DNA synthesis44. Loading proteins, DNA replication regulator
SLD2/3/7 (SLD2, SLD3, and SLD7) are then recruited to the re-replicative complex44.
Phosphorylation of the pre-replicative complex (inactive DNA helicase) by Dbf4-dependent
Cdc7 kinase (DDK), and the loading proteins SLD3 and SLD7, will stimulate the recruitment of
cell division cycle 45 (CDC45), an essential helicase activator44. S-phase-CDKs such as CDK2
will subsequently phosphorylate SLD3 and SLD245. Phosphorylated Sld2 results in its interaction
with the DNA replication regulator DPB11 (DPB11)45. It is thought that this interaction
stimulates the recruitment of DNA polymerase-ε (polymerase that extends the leading strand of
DNA during replication) and GINS (a multi-protein complex), leading to the formation of what
is known as the pre-loading/initiation complex4546. The pre-loading complex will dock onto
phosphorylated SLD3 via DPB11 and deliver GINS and DNA polymerase-ε to the nascent
14
replication complexes45. GINS, the second necessary helicase activator, will compete with SLD3
for binding to CDC45 and MCM2-747, resulting in the unloading of SLD3 from this complex and
the formation of the functional, replicative DNA helicase, CDC45/MCM2-7/GINS, otherwise
known as the CMG complex44. DNA polymerase-ε will bind directly to the CMG48.
Furthermore, although its time of assembly to the replisome is debated45, MCM10 is required for
recruiting the remaining DNA-polymerase-α/primase and DNA-polylmerase-δ44. As DNA
polymerases require a primer on which to begin DNA synthesis, the recruitment of DNA-
polymerase-α/primase, following the unwinding of the DNA strands by the CMG complex,
accomplishes this task49. The RNA primase will synthesize a ~10-nucleotide RNA primer
followed by 10 to 20 DNA bases. DNA polymerase-ε and δ will subsequently begin their tasks
of replicating the leading and lagging strands, respectively49,50 (Figure 1.4).
Figure 1.4 Assembly of the pre-replicative and pre-loading complex50. The origin of recognition complex
(ORC1-6) serves as a platform for the assembly of the pre-replication complex. CDC6 and CDT1 bind to the
ORC (origin licensing) which facilitates the loading of the inactive helicase complex proteins MCM 2-7. This is
known as the pre-replicative complex (PreRC). Helicase activator CDC45 is then recruited by SLD3, SDLD7 and
DDK. Phosphorylation of SLD2 and SLD3 by CDK2 promotes their interaction with DPB11 and the recruitment
of GINS to form the re-loading/initiation complex (PreIC). Unloading of SLD3 from the complex results in the
formation of the mature, functional DNA helicase, CDC45/Mcm2-7/GINS. Recruitment of Mcm10 and DNA
polymerases subsequently leads to the commencement of DNA replication. Modified from the original
publication by Aladjem, M. I. Nat. Rev. Genet. 8, 588–600 (2007)50. Used with permission of the author and
publisher.
SLD7
CDK2
P P
DDK
15
G1 and S phase cyclins and CDKs are crucial for the sophisticated choreography of DNA
replication. Together they help to assemble and activate the previously mentioned prereplication
complex at the origins of replication of the genome51. Cyclin E-CDK2 and cyclin A-CDK2
complexes are of the most crucial for S phase initiation and progression51. Cyclin E-CDK2
phosphorylates many components of the prereplication complex which regulates initiation of
DNA replication, as well as proteins like nuclear protein ataxia-telangiectasia locus (NPAT)
which are required to activate the transcription of histones51. In turn, cyclin A-CDK2 is found
with replicating foci of DNA, and phosphorylates many components of the replication machinery
such as DNA polymerase α,δ, and PCNA51. Further evidence of cyclin A’s importance in DNA
replication is demonstrated by the fact that its ectopic overexpression is sufficient to promote S
phase progression52 and concordantly, injection of anti-sense DNA constructs for cyclin A
inhibits S phase entry53.
1.5.4 G2 Phase
The G2-M phase checkpoint is essential to ensure replication of the genome has occurred
faithfully and to its entirety, thereby preventing the division of mutated cells, cells lacking
genetic material, or cells with DNA damage. During this phase, cell growth continues, and
proteins are synthesized for the execution of mitosis. Again, this crucial checkpoint is tightly
regulated by CDKs42, where the transition from G2-M phase and entry into prophase is initiated
by the activation of CDK1 complexed to B-type cyclins (cyclin B1 and B2)54. Knockdown of
either of these components can cause G2 arrest54,55,56. Therefore during G2, CDK1 activity is
dampened by various mechanisms until the cell is completely prepared for M phase. Primarily,
this occurs through retaining CDK1-cyclin B in the cytoplasm, where its activity can be
negatively regulated by the WEE1 G2 checkpoint kinase (WEE1)37. WEE1 phosphorylates
CDK1 on the tyrosine and threonine residues, Y15 and T14 respectively, which inhibits its
catalytic activity by disrupting its substrate binding ability37. During the G2/M transition,
following CDK-activating kinase (CAK) phosphorylation, the CDK1-cyclin B complex will
translocate into the nucleus where inhibitory phosphate residues can be removed by the nuclear
bound phosphatase cell division cycle 25 (CDC25) such that it can proceed to phosphorylate its
many nuclear targets37. Dephosphorylation of Y15 and T14 by Cdc25 is the rate limiting step for
progression into mitosis37.
16
1.5.5 M Phase
CDK1 has hundreds of downstream phosphorylation targets including many different effector
kinases necessary for the progression of mitosis37. For example, the initial chromosomal
condensation in prophase is dependent on the phosphorylation of the condensin II protein by
CDK157. This process can be visualized microscopically as string-like structures within the
nuclei, and is a hallmark of M phase initiation57.
M phase can be broken down into two large events; nuclear division (mitosis), followed by
cellular division (cytokinesis). During mitosis, the newly replicated sister chromatids are
separated equally for the formation of two daughter cells58,59. It is the most dramatic phase of the
cell cycle, as nearly all the inner contents of the cell become reorganized. Mitosis is
conventionally separated into 5 stages: prophase, prometaphase, metaphase, anaphase, and
telophase59 (Figure 1.5).
Figure 1.5 The main stages of mitosis 37. Prophase: CDK1 activation followed by CDK1-cylin B trafficking
into the nucleus leads to chromosomal condensation. Prometaphase: Breakdown of the nuclear envelope allows
formation of the mature spindle with centrosomes on either side of the cell. Kinetochore microtubules interact
with kinetochore protein complexes formed at the centromere, moving chromosomes to the spindle midzone.
Metaphase: Formation of the metaphase plate at the middle of the spindle, chromosomes become “bi-oriented”
through attachment to kinetochore microtubules. Unattached chromosomes produce a checkpoint signal that
prevents metaphase/anaphase transition. Anaphase: Once all chromosomes are attached and oriented properly,
the anaphase promoting complex ubiquitin ligase is activated, leading to ubiquitin -dependent proteolysis of
inhibitory protein, securin, allowing activation of a protease called separase which cleaves the cohesion proteins
that maintain cohesion between sister chromatids, allowing the chromosomes to separate to opposite poles of the
cell. Telophase: Following chromosome segregation, CDK1 becomes inhibited through anaphase promoting
complex mediated degradation cyclin B, and activation of the CDK1-antagonizing phosphatase CDC14. Dramat ic
decrease in CDK1 activity leads to reconstruction of the nuclear envelop, chromosomal decondensation and entry
into G1 following cytokinesis. Originally published by Rhind, N. & Russell, P. Cold Spring Harb. Perspect. Biol.
4, a005942 (2012)37. Used with permission of the author and publisher.
17
During prophase, chromosomal condensation begins and the centrosomes (microtubule
organizing organelles) move to opposite sides of the nucleus. There, the centrosomes will serve
as the two poles of the mitotic spindle (microtubule structure responsible for separating the
chromosomes to the two opposite ends of the cell)37, which begins its construction during late
prophase. The condensed sister chromatids are held together at the centromere, the DNA region
where proteins will later attach to form the kinetochore, a structure that mediates attachment of
the chromosomes to the mitotic spindle37. The striking breakdown of the nuclear envelope marks
the end of prophase and the commencement of prometaphase, freeing the sister chromatids from
the nucleus37. In prometaphase, the mature mitotic spindle is formed with poles residing at the
two opposite sides of the cell37. Kinetochore microtubules will emanate from the spindle poles
and interact with the assembled kinetochores of the sister chromosomes37. This process triggers
the brief onset of metaphase, where the chromatids are aligned at the mid zone of the mitotic
spindle, known as the metaphase plate37,59. Transition into anaphase is triggered by activation of
the anaphase promoting complex (APC)37. The APC is a multi-subunit E3 ubiquitin ligase
complex that targets many mitotic proteins for proteolytic degradation. Its activation at the
metaphase-anaphase transition is mediated by the binding of CDC20, and APC-CDC20 will
proceed to degrade key proteins that function to prevent the separation of the sister chromatids,
thus permitting anaphase to occur37. A key substrate of APC-CDC20 at this time is the protein
securin37. Securin is a protein that inhibits separase, which is the protease responsible for
cleaving the proteins within the cohesin complex37 (a group of proteins including structural
maintenance of chromosomes 1 (SMC1), structural maintenance of chromosomes 3 (SMC3),
RAD21 cohesin complex component (RAD21), stromal antigen 1 (SA1/STAG1) and stromal
antigen 2 (SA2/STAG2), which maintain cohesion between sister chromatids following
replication)60. Loss of chromosomal cohesion by separase-dependent cleavage of this complex is
necessary to permit chromatid segregation, allowing the mitotic spindle to pull the sister
chromatids to opposite poles37. The end of mitosis is marked by telophase, the stage in which the
chromosomes decondense and the nuclei reform around the sister chromatids37. Nuclear envelop
reconstruction is permitted by the inactivation of CDK, as a result of cyclin B ubiquitination and
degradation through APC bound to cadherin 1 (CDH1)37.
18
Cytokinesis, the mechanical process that cleaves the newly replicated mother cell into two
separate daughter cells, is the final step of mitosis59. During cytokinesis, the cell goes through
significant shape changes that must be tightly controlled, spatially and temporally, to ensure
proper segregation of genetic and cellular material61. The events of cytokinesis initiate during
anaphase, and are mediated by the formation of a contractile ring of actin and myosin filaments
beneath the plasma membrane, known as the cleavage furrow59. The ring is formed
perpendicularly to the mitotic spindle. Its contraction results in cleavage of the cell in the plane
that passes through the metaphase plate, pulling the plasma membrane inward and eventually
pinching the cell in half59.
Much like the rest of the cell cycle, cytokinesis is regulated by waves of cyclin and CDK
expression62. In budding yeast, degradation of mitotic cyclins by the APC, inactivation of CDK1
and upregulation of the major CDK antagonizing phosphatase CDC14, promotes each sequential
step of cytokinesis, beginning with furrow ingression, followed by plasma membrane resolution
and ultimately cell separation62. Experiments executed in HeLa cells expressing non degradable
cyclin B263, or temperature sensitive CDC14 mutants in yeast64,65, resulted in cells arresting in
late anaphase, without the completion of cytokinesis. CDC14 is sequestered into the nucleolus by
the protein neuroepithelial cell transforming 1 (NET1), throughout the majority of the cell cycle,
keeping it inactive66. However, in anaphase an abrupt increase in phosphorylation of NET1 by
the mitotic cyclin B1-CDK1 complex releases CDC14 it into the nucleus and cytoplasm where it
can now access its substrates67,68.
Recently, Kuilman et al. (2015), conducted phosphoproteome analyses to determine which
mitotic CDK substrates required dephosphorylation by CDC14 for the proper execution of
cytokinesis62. Dephosphorylation of actin interacting protein 1 (AIP1), EH domain-containing
and endocytosis protein 1 (EDE1), and Ingression protein 1 (INN1) were deemed necessary62.
They proposed that phosphorylated AIP1 acts as a negative regulator of acto-myosin ring
contraction, (dephosphorylation of AIP1 inhibits its function). Conversely, dephosphorylated
EDE1 acts as a positive regulator of cytokinesis (deletion of EDE1 but not AIP1 results in
increased cytokinetic defects), and dephosphorylation of INN1 promotes INN1’s timely
recruitment to the bud neck region of the dividing cell where it is required for ingression of the
plasma membrane62.
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Following furrow ingression, dividing cells are partitioned into two distinct daughters, but
remain connected by a thin intracellular bridge-like structure called the midbody69. This
structure arises from the central mitotic spindle (the microtubule array assembled between the
separating sister chromatids during anaphase), known as the midzone69. Conversion of the
midzone into the midbody corresponds with furrow ingression, as it is thought that this
contraction process compacts the microtubule bundles of the midzone into a single large
structure, creating the core of the midbody69. The serine-threonine kinase Aurora B, localizes to
the midzone and midbody structures, and also plays a role in chromosomal condensation through
histone H3 phosphorylation (PiH3)70. Due to its colocalization with the highly distinctive
midzone and midbody structures, immunofluorescence with antibodies against this protein is a
robust method to visualize cells undergoing mitosis and cytokinesis71. RNAi ablation of this
kinase results in a wide range of defects in prometaphase chromosomal condensation72,73,
anaphase chromosomal segregation73 and cytokinesis74 (Figure 1.6).
Figure 1.6 Aurora B Localization During Mitosis and Cytokinesis 71. HeLa cells were fixed and stained to
reveal Aurora B (red), tubulin (green), and DNA (blue). Aurora B translocates from the mitotic chromosomes in
metaphase to the central spindle early in anaphase. In early telophase when the cleavage furrow begins to form,
the cell can be seen pinching inwards in the same plane as the metaphase plate. At this stage Aurora B highlights
the spindle midzone. As telophase progresses, cleavage furrow ingression proceeds and the midzone is converted
into the midbody structure which is clearly marked by Aurora B that accumulates at the midbody arms in mid and
late telophase. Scale bars: 10 μm. Originally published by D’Avino, P. P. & Capalbo, L. Front. Oncol. 5, 221
(2015)71. Used with permission of the author and publisher.
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It is clear by the strict regulation of this complex process occurring at the all levels within the
cell, that the precise execution of the cell cycle is crucial for the proper function of every organ.
It is through the multiple signaling pathways and checkpoints described that critical events of
cell division occur in the proper order, and that quality control prevents cells from dividing with
damaged DNA or misaligned chromosomes. Certain cell types in the human body can re-enter
the cell cycle when stimulated by specific cues, to regenerate lost tissue, such as the liver, or are
in continuous turnover, such as endothelial cells. Other cells, such as adult cardiomyocytes, do
not have this luxury, and remain under tight cell cycle arrest from early in development. The
differences in regenerative potential between cells of different organs have important
implications for the types of diseases that can afflict them.
1.6 Cardiogenesis and Cardiomyocyte Cell Cycle Regulation
Adult cardiomyocytes are terminally differentiated cells that have exited the cell cycle, and
exhibit extremely poor regenerative potential. As previously discussed, different insults to the
heart such as ischemia (myocardial infarction), hypertension, atherosclerosis, and many others
can lead to heart disease, dysfunction or loss of cardiac muscle cells, and eventually death. The
problem lies in part, in the fact that the adult mammalian heart cannot efficiently generate new
cardiomyocytes in response to injury. Understanding early cardiogenesis, and the mechanisms of
cardiomyocyte cell cycle exit are crucial for the development of future therapies that involve
cardiac repair, as opposed to the current treatments that only transiently improve the function of
pre-existing cells.
1.6.1 Cardiogenesis
During embryogenesis in mice, two populations of cardiac progenitors derived from a common
precursor in the mesoderm contribute to the formation of the heart. Cardiac fate of these cells is
induced by bone morphogenic protein produced by the endodermal layer, whereas signals from
the wingless-type MMTV integration site family (WNT proteins, large family of secreted
glycoproteins which regulate crucial aspects of cell fate determination) from the neural tube and
notochord suppress cardiomyocyte specification75,76. At embryonic day 7.5, the primary heart
field, or cardiac crescent, is derived from the anterior mesoderm75,76. The secondary heart field is
derived from the pharyngeal mesoderm, located anterior and medial to the primary field75. Cells
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from the primary heart field migrate to form a linear heart tube at embryonic day 8, which acts as
the initial scaffold for further cardiac growth, including posterior and anterior expansion from
cells migrating from the secondary heart field75. The heart tube subsequently undergoes
rightward looping, which is the basis for the formation of the ventricles and atria and eventually
leads to the proper development of the four heart chambers75. Heart maturation includes septum
and valve formation. GATA Binding Protein 4 (GATA4) and NK2 homeobox 5 (NKX2.5) are
fundamental transcription factors in both heart fields, and two of the earliest cardiac lineage
markers, whereas T-box protein 5 (TBX5) and ISL LIM Homeobox 1 (ISL1) are uniquely
expressed in the primary and secondary heart fields, respectively75. Insight into these lineage
markers has been essential for the production of cardiomyocytes derived from induced
pluripotent stem cells (iPSCs) as well as for promoting cardiac repair in injured adult hearts.
1.6.2 Cardiomyocyte Regeneration
Throughout embryogenesis, mammalian cardiomyocytes undergo cell division such that the
heart grows through cardiomyocyte proliferation77. After birth however, this drastically changes.
Cardiomyocytes undergo a final round of DNA synthesis (karyokinesis), but do not complete
cytokinesis, rendering the vast majority of cells binucleated, which marks their terminal
differentiation77. Morphologically, terminal differentiation of cardiomyocytes is characterized by
binucleated cells with increased size, myofibril density, and mature intercalated discs77. At this
time, there is a transition from hyperplastic (cell proliferation) to hypertrophic (cell enlargement)
cardiomyocyte growth. This was highlighted in a study which showed that cardiomyocytes
isolated from the hearts of 1 day old rats increased in number by 68% over the first three days in
vitro and subsequently remained constant. In contrast, from day 3 to day 12, cardiomyocytes
increased in volume by 2.5 fold, as opposed to in number78. In addition to an increased
proliferative capacity, neonatal hearts also retain a heightened regenerative capacity in response
to injury, similar to amphibians and zebrafish79. To demonstrate this, murine ventricular apexes
were resected 1 day after birth and at 21 days following resection, the entire apical defect
incurred by these mice had been replaced by new cardiomyocytes, accompanied by increased
markers of proliferation, restored systolic function, with the absence of hypertrophy or fibrosis79.
Interestingly, resection at 7 days after birth, a time point which aligns with cardiomyocyte cell
cycle withdrawal, does not promote cardiac regeneration79. Following surgery, these hearts did
22
not show any indication of increased cardiomyocyte proliferation, and instead experienced large
amounts of fibrosis79. These results indicate that there is a drastic switch in the proliferative and
regenerative capacity of cardiomyocytes within the first week after birth. This loss of the
proliferative capacity is associated with an exit from the cell cycle. The cell cycle exit observed
has been shown to correspond with a stark upregulation of CDK inhibitors leading to a dramatic
downregulation of many cyclins and their CDKs to almost undetectable levels (their expression
is known to be high in embryonic, proliferative hearts)77. Tane et al. (2015) demonstrated that
the upregulation of two CDK inhibitors, cyclin-dependent kinase inhibitor 1A and 1B
(CDKN1A/p21Cip1 and CDKN1B/p27Kip1) mediate cell cycle exit in postnatal cardiomyocytes,
and the inhibition of two cell cycle activators, cyclin D1 and CDK1, mainta ins cell cycle arrest
throughout life80,81. Firstly, they demonstrated that p21Cip1 and p27kip1 levels peak at post-natal
day 581. Upregulation of these inhibitors coincides with when cyclin D1, cyclin E and cyclin A
activities begin to decrease, which are essential for G1 entry, progression and transition into S
phase81. Next they showed that over expression of cyclin D1 in mice using a Cre-loxP system
allowed adult cardiomyocytes to re-enter into the cell cycle, but these cells became arrested in M
phase due to lack of CDK1 activation80. They therefore proposed that two blockades of cell cycle
progression exist in adult cardiomyocytes, a G1 blockade, due to cyclin D1 downregulation, and
an M phase blockade, due to inhibition of CDK1 activation80. It is clear from this work that a
multi-pronged approach that could induce the upregulation of a battery of cell cycle activating
genes, and/or reduce key cell cycle inhibitors is necessary to promote cardiomyocyte
proliferation, but targeting solely one cell cycle regulator is not sufficient, as road-blocks at
multiple levels are in place.
1.7 MicroRNAs
MicroRNAs (miRNAs) are a group of small non-coding RNAs with the capability of expansive
genetic regulation. This class of endogenous 21-25 nucleotide long single-stranded RNA
molecules function as guide molecules in RNA silencing through base pairing to their
complementary mRNA targets82,83,84. By targeting specific mRNAs for degradation or
translational repression, miRNAs acts as inhibitors of gene expression, and are involved in
almost all developmental and pathological processes in animals. Since the discovery of the first
miRNA in Caenorhabditis elegans, lin-4, in 199385, cloning and size-fractionated RNA
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techniques have allowed for the identification of thousands of miRNAs, many of which have
been shown to have regulatory roles in cell cycle regulation, proliferation, cell death, cardiac
development, metabolism, neuronal patterning, hematopoietic differentiation and the immune
system82. Recently, high throughput sequencing, computational techniques and bioinformatics
algorithms have allowed for more in depth studies of the diverse target genes of miRNAs, and it
is known that one given miRNA can bind hundreds of target mRNAs. Based on the potential for
vast impact on gene expression, miRNA biogenesis is tightly regulated at multiple levels within
the cell.
1.7.1 miRNA Nomenclature
Over time, nomenclature of miRNAs has been inconsistent. Early discovered miRNAs were
named after their phenotypes (e.g. lin-4 or let-7), whereas miRNAs discovered through cloning
or sequencing were given numerical names (e.g. the lin-4 homologues in other species are called
miR-125)84. miRNAs generated from different loci that have identica l “seed” sequences from
nucleotides 2-7 are considered a family of miRNAs, and generally arose through gene
duplication84. For example, in humans, there are 14 paralogous loci (encoding “miRNA sisters”
that belong to the let-7 family, and are indicated by lettered suffixes (e.g. let-7a, let-7f)84. Each
locus produces two mature miRNAs; one from the 5’ strand and one from the 3’ strand of the
pre-miRNA (e.g. miR-181a-5p and miR-181a-3p), but one arm is then chosen as the guide
strand, and is usually 96-99% more prevalent and more biologically active than the other,
passenger strand76.
1.7.2 miRNA Biogenesis
miRNA genes are transcribed by RNA polymerase II or III as a long (> 1 kilobase pairs) primary
transcript (pri-miRNA) containing a hairpin structure in which the miRNA sequence is
embedded83. Still within the nucleus, the pri-miRNA is recognized by the double stranded RNA
binding domain regions of the DiGeorge Syndrome critical region gene 8 protein (DGCR8),
which will guide the class 2 RNAse III enzyme Drosha ribonuclease III (Drosha) to cleave the
RNA molecule83. Cleavage by Drosha liberates a 60-70 nucleotide long stem structure known as
the pre-miRNA. Proper cleavage by Drosha is crucial as it defines the terminus of an miRNA,
and thus, its specificity83. Drosha will cleave the pri-miRNA ~11 base pairs from the basal
24
junction, which serves as the major reference point for cleavage site determination, and then
again ~22 base pairs from the apical junction83,86. The cleaved pre-miRNA has a staggered end
with a ~2 nucleotide single stranded overhang at its 3’ end. This overhang allows it to be
recognized by exportin-5 which will mediate its transport into the cytoplasm for further
maturation83. Here, Dicer, a second RNAse III endonuclease, will process the pre-miRNA into
its mature form through specific cleavage ~22 nucleotides from its preexisting 5’ terminus (in
mammals)83. The approximately 22 nucleotide miRNA duplex is subsequently loaded onto the
Argonaute (AGO) protein complex, and one strand of the duplex (passenger strand) is degraded
by the endo-nucleolytic enzyme activity of AGO87, whereas the remaining single stranded RNA
molecule becomes the mature guide strand/miRNA. The mature miRNA bound to the AGO
complex is known as the RNA-induced silencing complex (RISC). Guide versus passenger
strand selection is mediated partially by the thermodynamic stability of the two ends of the
duplex, whereby the strand with lower internal stability at the 5’ terminus is preferentially bound
by AGO and retained in the RISC complex88. Additionally, AGO proteins are biased to selecting
guide strands that contain a uridine at their 5’ terminus. However, it is known that these rules of
strand selection are not completely strict, as passenger strand selection (arm switching) has been
observed with varying frequencies between species89,90. The mature miRNA serves to guide the
RISC complex to its target mRNA, and complementary binding of the miRNA to its target
gene’s mRNA transcript will lead to silencing of the mRNA through its direct degradation, by
translational repression, or by promoting mRNA decay84,91 (Figure 1.7).
Mature miRNAs are thought to be quite stable within the cell, as they can still be detected 48
hours after depletion of miRNA processing factors92. Binding of mature miRNAs to AGO
proteins enhances their stability even further93. Different miRNAs are seen to be degraded at
different rates within the cell, and degradation occurs at the precursor level as well as of mature
miRNAs, indicating that this may be yet another layer of regulation in the processing pathway92.
However, nucleases that degrade human miRNAs remain to be uncovered92.
25
1.7.3 miRNA Mechanism and Function
The RISC complex, guided by a mature miRNA to recognize specific mRNAs, post-
transcriptionally downregulates gene expression by one of three mechanisms: 1) inhibition of
mRNA translation, 2) mRNA cleavage, or 3) acceleration of miRNA decay84. miRNAs bind to
their targets through Watson-Crick base pairing, most frequently in the 3’ un-translated region
Figure 1.7 The Canonical Pathway of miRNA Biogenesis.91 (A) RNA polymerase II or III mediated
transcription of the primary miRNA transcript (pri-miRNA) and cleavage of the pri-miRNA by the
microprocessor complex Drosha-DGCR8 within the nucleus begins miRNA biogenesis. This results in the
formation of the pre-miRNA which is exported into the cytoplasm by exportin-5. In the cytoplasm, the RNase
Dicer complexed with TRBP will cleave the pre-miRNA into its mature 21-25 nucleotide length. The guide
miRNA strand is loaded onto the Argonaute proteins (AGO) to form the RNA -induced silencing complex
(RISC). The RISC complex, guided by the mature miRNA will silence target mRNAs through mRNA cleavage,
translational repression or mRNA deadenylation83,91. (B) The microprocessor complex measures ~11 bp from
the basal junction and ~ 22 bp from the apical junction and will cleave the RNA strand at this position to generate
the pre-miRNA83,100. Originally published by Winter, J et al. Nat. Cell Biol. 11, 228–234 (2009)91. Used with
permission of the author and publisher.
A B
26
(UTR) of target mRNAs84. Binding is not always 100% complementary, and mismatches and
bulges can be observed84. The degree of complementarity between miRNA and mRNA is the
major determinant of whether the mRNA becomes cleaved or if translation is simply inhibited84.
A high degree of complementarity is necessary to facilitate AGO-catalyzed mRNA cleavage,
whereas increased mismatching leads to translational repression or mRNA decay84.
The exact mechanism of translational inhibition by RISC has not been completely elucidated.
However, density gradient centrifugation has been used to show that translation can be inhibited
by miRNAs at either the translation initiation step (mRNA found complexed with RISC and
translational initiation factors) or at the elongation step (larger, incomplete polysomes
detected)84. Peterson et al. 2006 proposed that RISC binding can promote early ribosome
dissociation from mRNAs, thus interfering with the elongation process94.
Two different models have been proposed for miRNA inhibition of translational initiation. First,
RISCs have been shown to compete with eukaryotic translation initiation factor 4E (eIF4E) for
binding to the 5’ cap of the mRNA, resulting in failure to initiate translation90,94. The second
model proposes that binding of the RISC complex to the mRNA and 40S ribosomal subunit
preinitiation complex recruits the eukaryotic translation initiation factor 6 (eIF6), a ribosome
inhibitory protein. eIF6 binds to the 60S ribosomal subunit and prevents productive assembly of
80S ribosomes, and it was shown that depletion of Eif6 abrogates the Caenorhabditis elegans lin-
4 mediated translational repression of endogenous LIN-4 mRNA.
mRNA decay is promoted by RISC destabilization of the mRNA transcript. This occurs when
RISC binding to certain targets leads to rapid deadenylation of the 3’ Poly(A) tail of the
mRNA95,96. 3’ Deadenylation destabilizes the mRNA and facilitates its decay in the cytoplasm.
The let-7 family has been shown to function through this mechanism95,96.
Overall, the exact mechanism of silencing, be it translational repression, mRNA decay through
destabilization of its structure, or AGO mediated mRNA cleavage, remains to be elucidated for
the large majority of mammalian miRNAs. It is clear, however, that their influence on gene
expression is potent and widespread.
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1.7.4 MicroRNAs and the Heart
1.7.4.1 MicroRNAs, Cardiac Development and Homeostasis
Zhao et al. (2007)97 and Chen et al. (2008)98 were the first to reveal the crucial role of miRNAs
in heart development and function through the conditional, tissue-specific knockout of Dicer, the
RNase III enzyme essential for the biogenesis of mature miRNAs. Dicer knockout led to early
embryonic97 or post-natal98 lethality (depending on time of knockout), due to severe
developmental defects in the cardiovascular system. Using Cre recombinase technology under
the control of the endogenous NKX2.5 promoter, knockout of Dicer, specifically in cardiac
progenitors by embryonic day 8.5 (E8.5), caused pericardial edema, poorly developed ventricles
myocardium and death by E12.597. Although early cardiac markers such as TBX5 remained
unchanged, microarray analysis of hearts at E11.5 revealed upregulation and downregulation of
numerous genes97. For example, the endoderm marker α-fetoprotein and the skeletal muscle
troponin, were upregulated, whereas myoglobin and the potassium voltage-gated channel
subfamily D member 2 (KCND2) were strongly downregulated97. Interestingly, ablation of Dicer
using Cre recombinase under the control of the α-myosin heavy chain promoter (α-MHC), which
directs cardiac specific expression, did not lead to embryonic lethality. Instead, all mice died
shortly after birth, by day 498. The discrepancy between these two studies likely lies in the timing
of Dicer knockout, as use of the α-MHC Cre recombinase system leads to knockout by E14.5 as
opposed to E8.5 when using the NKX2.5 promoter. This indicates that miRNAs play a role in
early and in late cardiac development. Dicer knockout by E14.5 led to the development of
substantially larger hearts compared to non-mutated littermates, due to dramatic left ventricle
dilation with intracardiac thrombus98. Decreased myocardial organization, decreased
cardiomyocyte integrity, and disarrayed myofibrils and sarcomeres, due to a significant decrease
in expression of contractile proteins were also observed, indicating that miRNAs are essential for
the development of proper heart structure and contractile function98. Disarrayed sarcomeres were
also dramatically shorter and decreased in abundance in mutant mice98. Specifically, there was a
strong decrease in myosin heavy chain proteins (except for β-myosin heavy chain) and cardiac
troponin T, which likely accounts for the structural phenotypes observed98. Furthermore, there
was a drop in heart rate also observed, suggestive of conductive defects98. Concordant with these
molecular changes, Dicer mutant mice also suffered from severe left ventricular dilatation and
28
decreased fractional shortening98. In all, it was concluded that loss of miRNA production in
cardiomyocytes by E14.5 results in a phenotype suggestive of dilated cardiomyopathy with the
concomitant development of heart failure, shortly following birth. This demonstrates that proper
expression of miRNAs is absolutely essential for the intricate development of the cardiomyocyte
contractile apparatus, and thus heart function98.
To determine the role miRNAs play in the maturing and adult heart, da Costa Martins et al.
(2008) subsequently knocked out Dicer in mice at 3 and 8 weeks of age99. Cardiac-specific
deletion of Dicer 3 weeks following birth results in sudden death, with only mild cardiac
remodeling, whereas deletion 8 weeks after birth caused extensive hypertrophy and the
accumulation of fibrotic lesions99. In addition, deletion of Dicer at 3 weeks of age led to mild
inflammatory cell infiltration, but no myofibril disarray, hypertrophy or fibrosis99. A
downregulation of the KCND2 calcium channel was also observed (similarly observed in Dicer
knockout at E8.597), along with enhanced arrhythmias, which likely accounts for the sudden
death experienced by these mice99. This revealed a necessity for miRNA function not only in the
embryonic heart, but also during cardiac maturation99.
Knockout of Dicer in 8 week old adult mouse hearts however led to a severely different
phenotype, with mutant mice developing heart weights twice the mass of their non-mutant
littermates, severely hypertrophic and disarrayed cardiomyocytes, strong inflammatory
infiltration and interstitial fibrosis99. Four weeks following Dicer depletion, mice experienced a
50% decrease in fractional shortening and significant dilation of the left ventricle, indicating that
Dicer depletion in the adult myocardium led to rapid pathological remodeling, and a phenotype
resembling heart failure99. In both mutants, a strong upregulation of pro-hypertrophic and fetal
genes occurred99. Together, these three studies indicate that subsets of miRNAs play different
but crucial roles in cardiac function at all stages of development as well as in cardiac
maintenance/homeostasis. Based on the requirement of miRNAs to maintain mature cardiac
homeostasis, Rao et al. (2009) sought to profile the population of miRNAs expressed in the adult
murine heart using Illumina/Solexa deep sequencing of a small RNA library100. Unexpectedly,
analysis of over 7 million reads from each sample of 6-8 week old hearts revealed that 18
miRNAs make up more than 90% of all expressed small RNAs, and the miR-1/206 family
accounted for almost 40% of all known miRNA reads in the heart100. The miR-30, let-7, miR-29
29
and miR-26 families were the next most abundant miRNA families expressed in the adult murine
heart100. The full list of top 20 most enriched miRNAs, and miRNA families can be found in
Figure 1. of Rao et al. (2009)100. miRNAs expressed within the introns of important cardiac
genes (e.g. miR-378 encoded within an intron of the PPARG coactivator 1 beta (PGC1-β) gene, a
crucial regulator of mitochondrial biogenesis) were also found to have a relatively high
expression in the heart, likely reflecting the high levels of transcription of these genes100. These
results indicate that either a limited number of highly expressed miRNAs regulate cardiac
maintenance/homeostasis, or a large number of low-abundance miRNAs with overlapping
functions, as well as the few highly abundant ones, are involved in these processes101. In line
with this study, Porrello et al. (2011) showed that from day 1 after birth to day 7, there is a
drastic switch in the miRNA profile in mice102. By day 7, the miR-15, let-7 and miR-30 families
were all significantly upregulated, correlating with the time of cardiomyocyte maturation and
cell cycle exit102. Overall, it is now well documented that the timely expression of specific
miRNAs is crucial for cardiac development and homeostasis.
1.7.4.2 MicroRNAs and cardiovascular disease
Given the importance of miRNAs in development, it is evident that deregulation of miRNA
expression could be implicated in human disease. Recently, miRNAs have been found
circulating in blood, and have been emerging as important biomarkers in numerous
cardiomyopathies103–105. Understanding the role of these miRNAs in the pathogenesis of
cardiovascular disease may has opened a new avenue for the development of novel therapies and
diagnostic tools.
The ischemic process during and following myocardial infarction is the major cause of global
transcriptomic changes within the heart, and triggers an inflammatory response22. Subsequently,
maladaptive cardiac remodeling plays an important pathologic role in the development of heart
failure post-myocardial infarction. The role of miRNAs in cardiac remodeling post-myocardial
infarction came to light when da Costa Martins et al. (2008) demonstrated that the miRNA
biogenesis pathway was essential for overall cardiac function, and that its loss appeared to
contribute to maladaptive cardiac remodeling and heart failure99. Several studies subsequently
demonstrated a crucial role for miRNAs in regulating angiogenesis, fibrosis and hypertrophy,
30
following myocardial infarction, linking altered miRNA expression to cardiomyocyte
ischemia106,107,108. The deregulation of miRNAs following ischemic insult highlights their
involvement in the regulation of normal heart function, suggesting that detecting alterations in
miRNA expression could be a sensitive prognostic marker in early the stages of heart failure.
The use of high-sensitive real-time PCR (RT-qPCR) or small RNA array techniques facilitate
miRNA detection in blood samples, even at very low concentrations in the circulation22. For
example, miR-208b was found to be increased by ~1600 fold in the plasma of patients with acute
myocardial infarction, compared to patients presenting with atypical chest-pain without
myocardial infarction104. Furthermore, miR-499-5p levels were found to be elevated in acute
myocardial infarction patients and in animal models of myocardial infarction109. Increased levels
of miR-499-5p were also positively correlated with 12-month mortality in a group of elderly
myocardial infarction subjects, highlighting its potential as a prognostic biomarker109. Moreover,
patients in the Osaka Acute Coronary Insufficiency Study (OACIS), who died due to a cardiac
cause within a year of discharge, had a ~3-4 fold upregulation of miR-155 and miR-380*,
suggesting measuring their levels in patients following acute myocardial infarction may have
prognostic value105. It was also shown that OACIS patients surviving acute myocardial
infarction, who developed de novo heart failure within one year, had significantly upregulated
levels of the p53-responsive miRNAs-192, -194 and -34a (see section 1.8.6 p53 and miRNAs) in
their sera103.
Not only have miRNAs been implicated in the onset of heart failure, post-ischemic injury, but a
plethora of miRNAs are also dysregulated during the progression of heart failure22. Multiple
studies have demonstrated that subsets of miRNAs that become upregulated in heart failure
promote the formation of interstitial fibrosis110,111,112 , hypertrophic extracellular signal-regulated
kinase (ERK)/MAPK110, and nuclear factor of activated T-cells (NFAT)/calcineurin signaling113,
the fetal gene program114, glucose metabolism115,116, apoptosis116, cytoskeletal remodeling117 and
arrythmogenesis116 (often by miRNA-induced inhibition of negative regulators of these
pathways). Conversely, miRNAs which maintain normal low physiological expression levels of
target genes that promote cytoskeletal and myofibrillar rearrangements (hallmarks of
hypertrophy), such as the RAS homolog gene family member A (RhoA)118 and CDC42119,
31
become downregulated in hypertrophy and heart failure (such that these processes become
upregulated)120.
Given these observations, miRNAs are involved in a myriad signaling and biological processes
critical for the regulation of cardiac tissue homeostasis and function. Thus, exploiting these
functions through miRNA-based therapies is poised to lead to novel approaches to treat patients
following myocardial infarction or with heart failure.
1.7.5 MicroRNA Therapeutics
Over the years, as it has become more apparent that miRNAs function as master regulators of the
genome, and are differentially regulated in multiple disease states, many groups have sought to
take advantage of their function for the development of miRNA based therapeutics against
different types of diseases. Through over-expression by miRNA mimics or by knock down with
the use of antagomirs, using either viral vectors or lipid based techniques as delivery methods,
the therapeutic potential of miRNAs is extensive.
1.7.5.1 Delivery Methods for miRNA Based Therapies
Viral-based delivery systems usually exploit retroviruses, lentiviruses, adenoviruses or adeno-
associated viruses as delivery vectors to overexpress a given gene of interest121. These systems
can also be employed for the overexpression of miRNAs, often in a tissue-specific manner. Viral
vectors have three major advantages: 1) high infection efficiency, 2) high and constant
expression of the miRNA mimic or antagomir of interest and 3) certain viruses can be
engineered to have a particular tissue specificity depending on the receptors they use to enter
into cells121. Furthermore, certain genomic modifications which inhibit their ability to replicate
have improved the safety of this delivery system121.
Transfection of miRNA inhibitors or mimics using lipid based systems is one of the most
common ways to study miRNA overexpression or downregulation121. In this approach, a cell-
membrane-like lipid surface will encapsulate the nucleic acids which need to be delivered into
the cell, and the membrane- like qualities of the capsule allow the liposome to pass through the
cell’s membrane, into the cytoplasm121. Using a lipid-based approach circumvents many of the
problems of viral vectors, but lipid based techniques still cause a certain amount of toxicity to the
32
cell, the transfection efficiency of this method is often less potent, and the liposome particles
have a short half-life121. Yet, liposomes remain as the more clinically relevant method to alter
miRNA expression. Commonly, miRNA mimics, which act to overexpress a given miRNA, or
miRNA antagomirs (anti-miRs), which act to downregulate the expression of a given miRNA,
are transfected into cells using this approach121.
1.7.5.2 miRNA Mimics
miRNA mimic (miR-mimic) technology is an innovative approach to gene silencing, through
exploitation of miRNA function122,123. This is achieved by designing a ~22 nucleotide RNA
fragment with its 5’ end partially complementary to a selected unique sequence in the 3’ UTR of
the target gene of choice123. Once transfected into the cell, this RNA molecule, mimicking an
endogenous miRNA, will be picked up by the RISC complex, bind specifically to its target gene,
and induce post-transcriptional repression of the gene123. Alternatively, a miRNA mimic can be
designed to have the same sequence as a known miRNA, which allows you to study the effect of
having increased amounts of a given miRNA within a cell122. miRNA mimics have been used
extensively to study the consequences of miRNA overexpression in the heart, and have been
utilized in a therapeutic framework to substitute protective miRNAs that become depleted in a
diseased state. For example, Huang et al. (2016) transfected neonatal rat cardiomyocytes with a
cocktail of miR-21-mimic and miR-146a-mimic, two miRNAs previously known to have anti-
apoptotic and anti-inflammatory properties, respectively124, following hypoxia in vitro, and
myocardial infarction in vivo. Co-transfection of these two miRNA mimics reduced the degree of
cardiomyocyte apoptosis124. Furthermore, this treatment reduced infarct size and improved
cardiac function post-myocardial infarction124. Looking at gene expression in the heart following
treatment, they found that upregulation of these miRNAs significantly reduced the expression of
the phosphatase and tensin homolog (PTEN) and tumor necrosis factor receptor associated factor
6 (TRAF6), important apoptotic regulators, leading to the increased phosphorylation of p38
MAPK and a consequential downregulation of caspase 3 activity (involved in the signal
transduction cascade responsible for apoptosis execution)124. This study highlights the
therapeutic value and feasibility of upregulating cardioprotective miRNAs using miRNA
mimics.
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1.7.5.3 miRNA Antagomirs
miRNA antagomirs employ the opposite approach to mimics, and function by intercepting
overexpressed, detrimental miRNAs122. As their name suggests, antagomirs antagonize the
function of miRNAs, and have served as a promising strategy to target oncogenic miRNAs for
the treatment of cancer, or upregulated pathogenic miRNAs in other diseases122. This technology
was pioneered by Krutzfeldt et al. 2005, who designed chemically modified, cholesterol-
conjugated single-stranded RNA analogues complementary to miRNAs, and gave them the name
antagomirs125,126. These molecules can be transfected or transduced into cells in vitro or in vivo,
and function by intercepting and degrading mature miRNAs, preventing the functional miRNA
effect, that is silencing of their respective target genes125,126. Antagomir nucleotide modification
was also optimized by this group, including 2’O-methyl nucleotides for enhanced miRNA-
mRNA duplex stability, a phosphorothioate backbone to prevent RNase-mediated antagomir
degradation and conjugation to a cholesterol molecule for improved delivery across cell
membranes in vivo without the requirement of liposomes125. Further improvements to antagomir
technology have been made over the years, such as utilizing fully locked nucleic acid (LNA)-
modified oligonucleotides122. LNAs are RNA nucleotides that are modified to contain an extra
bridge on the ribose moiety, locking the ribose in a conformation which significantly improved
hybridization properties, therefore improving the potency of antagomirs122.
The therapeutic potential of miRNA antagomirs in the heart has been highlighted by many
studies, showing that knockdown of specific miRNAs can improve heart function in a battery of
cardiovascular disease models. For example, Wahlquist et al. (2014) demonstrated that miR-25
becomes significantly upregulated in human myocardial samples from patients with severe heart
failure. This miRNA strongly targets and downregulates expression of the Ca2+ pump SERCA2
(Ca2+ uptake is impaired due to a downregulation in SERCA2 expression in heart failure)127.
They next showed that transduction of anti-miR-25 in vivo into trans-aortic constriction (TAC)-
induced failing mouse hearts led to a significant increase in SERCA2 levels, restored fractional
shortening to normal levels (~20% increase), stabilized the heart weight to body weight ratio and
drastically improved survival (7/8 of anti-miR-25+TAC mice surviving vs. 7/22 TAC mice
surviving)127. Due to their ability to enter cells without the requirement of liposomes or viral
vectors, antagomirs may be a safe and viable avenue for drug development in cardiovascular
34
disease as in other diseases. Current research is now beginning to focus on non-oligonucleotide
based methods to modulate miRNA expression levels, such as through small molecules.
1.7.6 MicroRNAs and the cell cycle
Similar to embryogenesis and development, the carefully timed expression of cell cycle
activators and inhibitors is essential for the proper regulation of the cell cycle. Furthermore, as
cells of different organs proliferate at different rates, or are completely removed from the cell
cycle, it is clear that the expression level of these cell cycle regulators is highly tissue specific.
Thus, it is not surprising that miRNAs, with the capability to fine tune the expression level of
hundreds of genes, are key players in cell cycle regulation. The differential expression of
miRNAs in the different tissues of the body likely plays an important role on regulating that
tissue’s proliferative capacity. Consequently, over the years, miRNAs have been emerging as
novel regulators of tumor suppression and oncogenesis.
Each stage of the cell cycle is regulated by a battery of miRNAs, targeting mRNA transcripts of
both cell cycle activating and inhibiting genes128. One of the first links to implicate miRNAs with
the cell cycle was made by Calin et al. (2002) who uncovering the anti-proliferative effects of
the miR-15a-16-1 cluster (which expresses mature miRNAs miR-15a and miR-16), and their
implication in cancer129. This cluster was identified as the target of specific chromosome
aberrations in chronic lymphocytic leukemia patients, and is deleted or downregulated in ~70%
of these patients129. Further studies then found that it was also deleted or downregulated in
pituitary adenomas130, gastric cancer131,132 and prostate cancer133, providing supporting evidence
for an important role in tumor suppression. It is thought that these two miRNAs maintain cell
cycle arrest at the G1 phase by targeting CDK1, CDK2, CDK as well as cyclins D1, D3 and
E1132,134,135. Following this, a plethora of other miRNAs were found to target these major cell
cycle kinase complexes. It is now clear that at each phase of the cell cycle, many important cell
cycle regulators are targeted by multiple miRNAs in a largely redundant manner. This highlights
the importance of keeping a tight regulation over these key genes (Figure 1.8).
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Based on the knowledge that miRNAs have a strong influence on the cell cycle, Eulalio et al.
(2012) sought to determine which miRNAs play a role in maintaining cell cycle arrest in
cardiomyocytes136. To do this, they performed a fluorescence-microscopy-based screen of 875
miRNA mimics in neonatal rat cardiomyocytes to systematically identify miRNAs that trigger
cardiomyocyte cell cycle re-entry136. Staining for the proliferation antigen Ki-67 and for 5-
Figure 1.8 An overview to cell cycle control by microRNAs 128. let-7 (let-7a-f) and miR-15 (miR-15, miR-16
and miR-195) indicate several members of the corresponding miRNA family. miRNAs with proliferative potential
are shown in red whereas antiproliferative miRNAs are in blue. S = S-phase, M = Mitosis, G1 and G2 indicate the
gap phases of the cell cycle and G0 indicates quiescent cells . Originally published by Bueno, M. J. & Malumbres,
M. Biochim. Biophys. Acta - Mol. Basis Dis. 1812, 592–601 (2011)128. Used with permission of the author and
publisher .
36
ethynyl-29-deoxyuridine (EdU, a uridine analogue that is incorporated into newly synthesized
DNA), they found that 204 miRNA mimics significantly increased neonatal cardiomyocyte
proliferation by more than two-fold, and 331 mimics decreased Edu incorporation and Ki-67
positivity from the baseline of 12.5% to 0%136. Interestingly, the top miRNAs promoting
cardiomyocyte proliferation were not among those affecting cardiomyocyte size or
hypertrophy136. In this study, the top 4 pro-proliferative miRNAs which significantly increased
both EdU incorporation and Ki-67 positivity were miR-590-3p, miR-199a-3p, miR-1825 and
hsa-miR-33b*136. Conversely, the top 4 miRNAs with tumor suppressor properties, that
significantly decreased cardiomyocyte proliferation were miR-1287, miR-34c, miR-885 and
miR-449136. The authors subsequently showed that treatment of mice with mimics to these
potentially pro-regenerative miRNAs (miR-590-3p and miR-199a) significantly boosted the
repair of adult cardiomyocytes post-infarct, and that the increase in cardiomyocyte proliferation
was correlated with improved functional outcomes136. The overall implication of these results are
twofold. Firstly, that a vast number of miRNAs must be differentially regulated in adult
cardiomyocytes to maintain cell cycle arrest, whereby the majority of pro-proliferative miRNAs
are downregulated and miRNAs promoting cell cycle arrest are more prominently expressed.
Secondly, that upregulating cardiomyocyte proliferation following injury to stimulate heart
regeneration is a viable therapeutic option that merits further investigation.
1.8 Tumor Suppressors – A Focus on p53
Neoplasia and tumorigenesis represent a loss of control in the processes of cell proliferation and
differentiation137. Tumor suppressor genes were first identified because they were inactivated in
mammalian tumors, and subsequent experimentation demonstrated their ability to protect against
neoplastic transformation137,138. These crucial regulatory genes are now known to influence
fundamental cellular processes such as development, growth and cell cycle, genome
maintenance, differentiation, cell death and tissue regeneration137,138. There are four main
mechanisms by which tumor suppressors may act to protect cells from neoplastic transformation:
suppression of cell division, DNA damage repair, induction of apoptosis or senescence, and
inhibition of metastasis139
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Tumor suppressors that suppress cell cycle and division include RB, adenomatosis polyposis coli
(APC), alternate reading frame (ARF), retinoblastoma protein-interacting zinc finger protein
(RIZ1), cyclin-dependent kinase inhibitor 2B (CDKN2B/p15ink4b), cyclin-dependent kinase
inhibitor 2A (CDKN2A/p16ink4a), cyclin-dependent kinase inhibitor 2C (CDKN2C/p18ink4c),
p21Cip1, p27Kip1, and Trp53 (p53)140–150. Inhibition of aberrant cell division is the main
mechanism for most tumor suppressors, and many of these genes are frequently mutated in all
types of cancer. RB was the first discovered tumor suppressor. It inhibits the transcription of
factors required for cell cycle progression through repressing the activity transcription factors
like the E2Fs140,141. p15INK4B, p16INK4A, p18INK4C, p21Cip1 and p27Kip1 function by
inhibiting CDKs, which in turn inhibit RB and promote cell cycle progression144–148. Thus,
inhibition of CDKs permits RB activity, allowing it to maintain G1 phase arrest. The tumor
suppressor p53, is a transcription factor that stimulates the expression of many cell cycle
inhibitors and pro-apoptotic genes, among many other roles which will be discussed further150.
ARF in turn functions as tumor suppressor by stimulating the p53 pathway through neutralizing
the effects of p53’s two negative regulators murine double minute 2 (MDM2) and MCL-1
ubiquitin ligase E3 (MULE)149. Furthermore, independently of p53, ARF functions to antagonize
the function of major cell cycle activators such as c-MYC and E2F1149.
The proper regulation of apoptosis maintains normal homeostasis and suppresses cancer.
Consequently, cancerous cells are often resistant to apoptosis through a downregulation or
deletion of key pro-apoptotic tumor suppressors151. Tumor suppressors that promote apoptosis of
aberrant cells are p53, APC, cluster of differentiation 95 (CD95), bridging integrator 1 (BIN1),
and PTEN152–156.
The ability to fix DNA damages is known as the DNA damage response. Although the
replication of mammalian cells is a high-fidelity process with multiple checkpoints in place, the
genome is at constant risk of mutation by many sources such as ultraviolet light, ionizing
radiation and reactive oxygen species151. The ability of genes within the DNA damage response
pathway to monitor the integrity of their genome and repair damaged DNA is crucial to prevent
the transmission of mutations to daughter cells during replication, which can contribute to
malignant transformation and tumorigenesis if the given mutation is found within a tumor
suppressor or oncogene. Tumor suppressors that maintain genome integrity are mutS homolog 2
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(MSH2), mutL homolog 1 (MLH1), Ataxia-telangiectasia-mutated gene product (ATM), breast
cancer protein (BRCA), Nijmegen breakage syndrome 1 (NBS1), Fanconi-Anemia-related tumor
suppressor (FA) and p53157–161.
Although tumorigenesis is promoted by loss of regulation of the first three mechanisms of tumor
suppression discussed above, the majority of cancer deaths are caused by metastasis (the spread
of a cancer to a different organ)151. During metastasis, cancerous cells interact with endothelial
cells, signaling to initiate angiogenesis and break down vascular walls which facilitates their
spread throughout the body151. Genes like p53, metastin, breast cancer metastasis suppressor 1
(BRMS1), tissue inhibitor of metalloproteinases (TIMP1) mediator complex subunit 23
(MED23) and CD82 promote endothelial integrity, inhibit angiogenesis and the epithelial to
mesenchymal transition, and exert many other functions to suppress metastasis162–167.
It is hypothesized that in a context-specific manner, some tumor suppressors may also have an
effect on suppression of regeneration in cells that lack strong regenerative capacity137. Studying
tumor suppressors in this context could allow us to promote regeneration in organs such as the
heart that largely lack this ability137. The tumor suppressor p53 will be of focus for the rest of
this literature review as it functions in all 4 main tumor suppressor mechanisms, is mutated and
or inactivated in nearly half of all sporadic human cancers, and has been found to be implicated
in tissue regeneration and the injury response in numerous organisms.
1.8.1 The p53/Mdm2 Tumor Suppressor Circuitry
Trp53 (p53) is a tumor suppressor gene encoding for a transcription factor that is the most
frequent target for mutations in tumors168. Due to this fact, the increased susceptibility for cancer
in patients with Li-Fraumeni syndrome (individuals who inherit a mutant p53 allele, and the
spontaneous development of tumors in p53 null mice169, the importance of p53’s role in tumor
suppression is unequivocal168. It is also known that in addition to its tumor suppressive function,
p53 plays an important role in homeostasis as a cellular stress sensor. It can trigger transient or
permanent cell cycle arrest, apoptosis or DNA repair in response to many stimuli including
hypoxia, DNA damage, ribonucleotide depletion and nutrient starvation, and is involved in the
regulation of metabolism and autophagy168. Depending on the stress experienced by the cell, the
phenotypic end result following p53 activation, due to displacement of its negative regulators
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MDM2/MDM4 or MULE, is either cell recovery and survival, or death137. Additionally, p53
belongs to a family of transcription factors including p63 and p73 which have overlapping and
distinct roles168.
Much of our initial understanding of p53’s tumor suppressive roles came from the generation of
p53-knockout mice. These mice, although developmentally normal, experience early onset,
spontaneous tumors (primarily CD4+CD8+ T cell lymphomas) at 100% penetrance170–172.
Moreover, reminiscent of patients with Li-Fraumeni syndrome, p53-null heterozygous mice have
a heightened predisposition to cancer compared to wild type mice, and primarily develop
sarcomas, as well as some lymphomas and carcinomas170–172. Subsequently, more refined studies
of tissue-specific p53 knockouts have helped to highlight p53’s role in individual tissues of the
body.
1.8.2 Canonical p53 Pathways
p53’s most thoroughly characterized biochemical role within the cell is as a transcription
activator, however it also is known to repress transcription of certain genes (estimated ~15% of
target genes), as well as to help promote mitochondrial membrane permeabilization to trigger
apoptosis168. More than 80% of p53 mutations in human tumors occur within the DNA binding
domains of this protein, compromising its ability for sequence-specific DNA binding and its
function as a transcription factor168. p53 forms tetramers through a carboxy-terminus
tetramerization domain which will subsequently bind to p53 response elements within the
genome to activate transcription of p53-dependent genes168. The consensus DNA binding
sequence for p53 consists of two repeats of the 10bp motif 5’-PuPuPuC(A/T)(A/T)GPyPyPy-3’
(Pu=purine, Py=pyrimidine) typically separated by a spacer of 0-13 nucleotides168,173. These
p53-dependent genes have largely been elucidated through genetic studies and chromatin
immunoprecipitation experiments, to help determine p53’s precise role within the cell168.
p53’s contribution to tumor suppression through regulation of cell cycle arrest lies in its
transcriptional activation of important cell cycle inhibitors168. Activation of p21Cip1 by p53
occurs in response to DNA damage, which leads to G1 phase cell cycle arrest, through p21Cip1
mediated inhibition of CDK1, CDK2 and PCNA174,175. p53 also regulates other cell cycle genes
such as protein tyrosine phosphatase receptor type V (PTPRV) which is also involved in G1
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arrest176, growth arrest and DNA-damage inducible 45 (GADD45A)177 and stratifin (YWHAS)178
which control the G2/M transition175.
To promote apoptosis, p53 activates the transcription of pro-apoptotic genes such as BCL2
associated X protein (BAX)179, CD95180, death receptor 5 (DR5)173, etoposide induced 2.4
(EI24)181, p53-Induced death domain protein 1 (PIDD1)173, tumor protein P53 regulated
apoptosis inducing protein 1(p53AIP1)182, p53 up-regulated modulator of apoptosis (PUMA)183,
phorbol-12-Myristate-13-Acetate-Induced protein 1 (NOXA)184 and p53 apoptosis effector
(PERP)185,173. These genes are the downstream effectors of p53 mediated apoptosis, which is
dependent on the apoptotic peptidase activating factor 1 (APAF1)/Capsase-9 pathway and
cytochrome c release from the mitochondria173.
On top of p53’s ability to trigger cell cycle arrest or apoptosis in response to DNA damage, it
further earns its name as “guardian of the genome” through its critical role in nucleotide excision
repair186. p53 mediates nucleotide excision repair in response to DNA damage through
transcriptional activation of two genes in this pathway, DNA damage checkpoint protein 2
(DDC2)187 and XPC complex subunit, DNA damage recognition and repair factor (XPC)188,
which recognize and process UV photoproducts in the genome.
1.8.3 Non-Canonical p53 Pathways
Non-canonical roles of p53 such as its involvement in metastasis, metabolism and autophagy
have been emerging as essential for tumor suppression and homeostasis168. It is now clear that
loss of p53 not only leads to tumor development, but the ability of tumors to spread throughout
the body168,162. Loss of p53 causes an increase in expression of GTP-bound (active) RhoA, a
small GTPase that regulates cell migration, and Rho associated coiled-coil containing protein
kinase 1 (ROCK1), its main effector protein162. These proteins control actin dynamics which
promote tumor cell invasiveness. Furthermore, loss of p53 also stimulates the epithelial-to-
mesenchymal transition of tumor cells, the activation of genes that are part of the adhesive
machinery required for motility and invasion, and degradation of the extracellular matrix162.
p53 also has a widespread role over metabolism within the cell189. Several studies have shown
that p53 can regulate both glycolysis and oxidative phosphorylation189. Reportedly, p53 inhibits
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glycolysis (tumor cells have a high metabolic demand and require heightened sugar uptake,
which is facilitated loss of p53) by inhibiting the expression of the glucose transporters GLUT1
and GLUT4190, while activating the expression of the TP53 induced glycolysis regulatory
phosphatase (TIGAR, a negative regulator of glycolysis)191. p53’s restraint on glycolysis comes
with an opposing stimulation of oxidative phosphorylation, by activating transcription of many
genes involved in mitochondrial maintenance189.
Owing to reports of autophagy playing a role in tumorigenesis and in the control of metabolic
stress in healthy cells, it is not surprising that p53 has been found to directly activate this
process191. p53 can induce transcription of the 5' AMP-activated protein kinase (AMPK) and
inhibit the expression of the mechanistic target of rapamycin (mTOR), thereby promoting the
process of autophagy192. Furthermore, canonical p53 target genes such as BAX and PUMA can
also act as positive regulators of autophagy, in addition to their role as mediators of p53-
dependent apoptosis193. As metabolic dysregulation and autophagy have been implicated in
cardiovascular disease and heart failure194,195, these relatively novel functions of p53, along with
p53’s regulation of canonical signaling pathways in the heart, are interesting therapeutic avenues
to be pursued.
1.8.4 Post-Translational Regulation of p53 by MDM2
Due to the vast effect that p53 can exert on the cell, it is imperative that its activity is tightly
regulated. p53 is subject to multiple modifications including phosphorylation, ubiquitination,
methylation, acetylation, sumoylation, and neddylation, all of which intricately regulate p53
function at an epigenetic level196,197. Among these, ubiquitination, phosphorylation and
acetylation are the most extensively studied, and are involved in the regulation of all 3 steps
required for p53 activation: stabilization of the p53 protein, p53 binding to regulatory DNA
recognition elements and transcriptional activation of target gene expression196,197.
Ubiquitination is the main contributor to p53 stability, and this job is largely assumed by the
oncoprotein, MDM2.
p53’s function is predominantly regulated by its protein stability, and has a short half-life of 5-30
minutes in normal unstressed cells196. This knowledge led to the search for a protein that could
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degrade p53, thereby explaining this very short intracellular half-life196. Immunoprecipitation
experiments identified this protein as MDM2, which forms a 90kDa complex with p53198.
Mdm2 is an E3 ubiquitin ligase that targets p53 for proteolytic degradation196,197,199. MDM2 was
first discovered as a gene that was overexpressed by more than 50-fold in a spontaneously
transformed mouse cell line (3T3-DM)200. The reason for the transformation potential of
overexpressed MDM2 was discovered, when it was shown to bind p53 and inhibit its
transcriptional transactivation capacity198. Similarly, it was found that MDM2 gene amplification
occurs in over 1/3 of human sarcomas that maintain wild type p53201.
Ubiquitination of p53 by MDM2 occurs in a series of steps that involves 3 enzymes196,197. The
E1 enzyme binds and activates the ubiquitin protein, E2 then accepts the activated ubiquitin and
transfers it to the E3 enzyme (MDM2 in this case), which is a substrate-specific ligase that
covalently bonds ubiquitin to its substrate (p53)196,197,199. MDM2 monoubiquitinates p53 at
multiple lysine residues, which is the critical step in mediating its degradation by nuclear and
cytoplasmic proteasomes196,197,199.
Mdm2 itself is the product of a p53 inducible gene197. Therefore, the two proteins form an
autoregulatory negative feedback loop that tightly maintains p53 levels within the cell197.
Moreover, this relationship limits the duration and severity of p53-dependent biological
responses following a non-lethal stressor to the cell199. In turn, low levels of p53 activity result in
constitutively low MDM2 levels197. To escape the negative regulation of MDM2 upon cellular
stress, p53 and MDM2 both become phosphorylated by diverse kinases, preventing MDM2’s
ability to bind its substrate202,203. Unlike the p53 knockouts, MDM2-null mice experience a very
early embryonic lethality, due to an overactive p53, impairing organ development204,205.
Consequently, concurrent knockout of p53 and MDM2 rescues the embryonic lethality of
MDM2-null mice204. MDM2 therefore plays an essential role in tumorigenesis as well as
homeostasis, by antagonizing the function of p53.
1.8.5 p53/MDM2 and the Heart
The development of primary cardiac tumors is extremely rare, with a prevalence of 0.001–0.03%
in autopsy series206. Based on this low prevalence of tumorigenesis, and the hearts low
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regenerative capacity, tumor suppressors may be exerting an important growth-limiting role
within this organ. The exact function of the p53/MDM2 tumor suppressor circuitry in the heart
has remained elusive. Some involvement of p53 and cardiovascular disease however, has been
inferred. It has been shown that cardiomyocyte-specific deletion of p53 in the heart is capable of
protecting against the transition of hypertrophy to heart failure207. p53 ablation resulted in an
upregulation of the pro-angiogenic hypoxia-inducible-factor-1 (HIF-1) which increased the
number of microvessels in the heart, and improved systolic function compared to wild type mice
subjected to the same insult207. Furthermore, the authors showed that following pressure
overload, p53 and phosphorylated p53 levels accumulate in the heart, and concluded that this
upregulation may be partially responsible for the detrimental effects on heart function
experienced by these mice207. Four studies have also demonstrated that p53 becomes
upregulated in response to cardiac ischemia, and that inhibition of p53 conveyed a protective
effect through reduction of apoptosis, via a decrease in BAX expression, and ameliorated
ventricular remodeling208–211. More recently, Yoshida et al. (2015) demonstrated that the
accumulation of p53 during cardiac pressure overload occurs also in cardiac endothelial cells212.
Upregulation of p53 in endothelial cells resulted in an upregulation of the intercellular adhesion
molecule-1, which is essential for the infiltration of inflammatory cells into the heart upon
pressure overload212. Therefore, upregulation of p53 following pressure overload contributes to
the inflammatory response induced by this insult, and exacerbates cardiac dysfunction212.
Although some research has been conducted on the effect of p53-dependent apoptosis in the
heart following pressure overload or hypoxia induced stress, gaps of knowledge exist in our
understanding of other functions that this tumor suppressor circuitry exerts on the heart, such as
cell cycle, regeneration and metabolism. Furthermore, few studies have examined the
combinatorial effect of p53 and MDM2 in the heart. The Billia lab hypothesized that based on
p53 and MDM2’s expansive control over cell cycle in many other organs, an explanation for the
heart’s low regenerative potential must lie, at least in part, within this pathway213. Research in
our laboratory sought to determine if the p53/MDM2 complex acts as an intrinsic inhibitor of
adult cardiomyocyte proliferation. To do this, p53 and MDM2 were inactivated specifically in
the adult mouse heart employing the Cre-loxP recombination system213. Using an inducible
cardiomyocyte-specific transgenic mouse, mcm, in which the cardiac muscle α-myosin heavy
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chain 6 promoter drives the expression of the Cre recombinase fused to two mutant estrogen-
receptor ligand binding domains, when exposed to 4-Hydroxytamoxofen (Tam) injections,
MDM2 (Mdm2KO), p53 (p53KO), and both proteins (double knockout, DKO), were knocked
out, specifically in the adult mouse heart213.
These three different mice had strikingly different phenotypes214. As expected, the p53KO did
not immediately show any adverse effects, but exhibited mortality by 6 months post-Tam. By
this time point, these mice exhibited hypertrophy and increases in fetal gene expression. Loss of
p53 also led to a decrease in two cardiac transcription factors, GATA4 and NKX2.5, identifying
these genes as previously unknown targets of p53 in the heart. As previously mentioned, the role
of p53 in controlling energy production and mitochondrial function has recently been brought to
light. Since the heart is a biomechanical pump with an enormous demand for ATP, this requires a
high mitochondrial content within the cardiomyocyte. It was therefore hypothesized that p53’s
role in energy regulation would be crucial within the heart. Indeed, by the 6 month time period,
ATP content significantly dropped. Thus, p53 is clearly essential within the heart to protect
cardiomyocytes against aberrant growth and metabolic transformation. Next, p53’s canonical
role of promoting cell cycle arrest was investigated in the heart. Interestingly, knockout of p53
alone was not sufficient to induce cardiomyocyte proliferation.
Conversely, the Mdm2KO mice had high early onset mortality by 14 days214. As expected, due
to high levels of p53 in the absence of MDM2, these mutant hearts exhibited significantly
increased cardiomyocyte apoptosis, accompanied by a reduced fractional shortening.
Furthermore, unlike the p53KO mice, energy balance defects were less pronounced.
Additionally, loss of MDM2 did not induce cardiomyocyte proliferation, as expected, due to
elevated levels of p53214.
Tam-induced homozygous ablation of both p53 and MDM2 caused dramatic early onset death of
all mice by day 10214. These results indisputably reveal that p53 and MDM2 are integral players
in a pathway that maintains normal cardiac performance. DKO mice developed dilated
cardiomyopathy with wall thinning, increased chamber sizes and substantial fibrosis, but had no
indications of hypertrophy. Furthermore, the cardiomyocytes exhibited sarcomeric structure
abnormalities, corresponding to a markedly impaired fractional shortening. Many genes within
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the fetal gene program were reactivated, with a concurrent decline in the expression of essential
cardiac genes. Similar to the p53KO, the DKO mice experienced a drop in energy production,
but at a much more dramatic rate. By 8 days post-Tam, ATP levels dropped significantly, and
this corresponded with a drastic reduction in mitochondrial copy number, mitochondrial gene
transcription, and a significant downregulation in mitochondrial Complex I-V genes.
Furthermore, as previously mentioned, p53 inhibits glycolysis and promotes fatty acid oxidation
(the main mechanism of energy production in the heart), and loss of p53 led to a significant
decrease in canonical p53 target genes that maintain fatty acid oxidation including AMPKα, and
TIGAR, concordant with an increase in glycolytic markers such as the RELA proto-oncogene,
NF-kB subunit (RELA, a crucial activator of the glycolytic program). These results revealed that
p53 and MDM2 are crucial for the maintenance of mitochondrial bioenergetics and energy
metabolism. Investigation of cardiomyocyte proliferation in the DKO mice revealed another
striking and unexpected phenotype. These mice had a significant upregulation of
cardiomyocytes in S phase, M phase and undergoing cytokinesis, in all four chambers of the
heart. Thus, it appears reasonable to assume that on top of p53, MDM2 regulates important cell
cycle activators within the heart, and that removal of both the negative regulator (p53) and the
inhibitor of positive regulators (MDM2) is required to induce cardiomyocyte proliferation. It
seems as though in the heart, multiple roadblocks are in place that must be removed
simultaneously to stimulate cell cycle re-entry214.
In all, there is an abundant amount of literature that demonstrates the role of p53 as an essential
contributor to growth arrest and apoptosis in response to various types of stress. Through
analysis of three heart specific mutant mice (p53KO, Mdm2KO and DKO) Billia et al. (2016)
established p53 and MDM2 to also be necessary under physiological conditions of low stress in
the heart, where they act to control heart function through the maintenance of cardiomyocyte
differentiation, hypertrophy, energy production/balance, and cell cycle arrest214.
1.8.6 p53 and miRNAs
It has been hypothesized that miRNAs evolved, to allow organisms to deal with stress, as
miRNAs have been shown to play integral roles in cell cycle, apoptosis, tumor
suppression/oncogenesis, metabolism, and much more215,216. In keeping with this, miRNAs have
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emerged as important executors of p53 dependent tumor suppression and stress responses217,218.
miRNAs are heavily interconnected in the p53 stress response pathway, not only because they
can be induced by p53, but because p53 is a downstream target of many miRNAs217.
The mechanism by which p53 mediates repression of target genes has been unclear, but it is now
thought that miRNAs contribute to a large majority of the mRNA and protein expression
downregulation observed following p53 activation217. Besides regulating the expression of
specific miRNA-coding genes, p53 also affects the processing of miRNAs directly through
interaction with DDX5 to enhance its interaction with the Drosha complex, promoting miRNA
biogenesis219. The discovery of many p53-dependent miRNAs has therefore expanded p53’s role
within the cell allowing it to indirectly regulate the expression of a multitude of genes. In
response to stress, p53-regulated miRNAs become expressed, engage in many ‘feedforward’ and
‘feedback’ loops that control amplification, robustness, fine-tuning and buffering of signals, all
of which contribute to the appropriate cellular response that occurs.
Several groups initially identified miR-34a and miR-34b/c as the first direct miRNA
transcriptional targets of p53, and have demonstrated that they help mediate many of functions of
p53 including induction of apoptosis, senescence and cell cycle arrest220–226. Since then, many
other p53-dependent miRNAs have been discovered including miR-15a/16-1, miR-145, miR-
200, miR-192, miR-104, miR-215, the let-7 family, miR-107, miR-149227, miR-29, miR-605217.
Recently, Hunten et al. (2015) undertook a high throughput approach, utilizing pulsed stable
isotope labeling, next generation sequencing, and chromatin immunoprecipitation, to determine
the genome wide effect of p53 upregulation in a colorectal cancer cell line228. They found that
p53 induction (using a vector based technique to overexpress p53) led to widespread differential
regulation of proteins (542 up, 569 down), mRNAs (1258 up, 415 down), miRNAs (111 up, 95
down), and long non-coding RNAs (270 up, 123 down)228. Transcriptionally induced genes
displayed more occupied p53 binding sites than repressed genes, suggesting indirect mechanisms
of repression, such as through miRNAs, are at play. Moreover, 50% of the transcriptionally
repressed genes contained seed-matching sequences to p53-induced miRNAs in their 3’-
UTRs228. This study further supports the notion that p53-dependent gene repression is mediated
47
indirectly through p53-induced miRNAs. A full table of p53-regulated miRNAs can be found in
Supplemental Figure 11 of Hunten et al. (2015)228.
Many p53-regulated miRNAs have been shown to have robust effects on cell cycle, and
themselves act as tumor suppressors217. Numerous studies have shown distinct p53-regulated
miRNAs to be downregulated by genetic or epigenetic mechanisms in diverse human tumors,
and that their re-introduction through miRNA mimic technology can mitigate tumor cell
proliferation and improve cancer outcomes217. Cell cycle-related factors have bioinformatically
been shown to be over-represented in the profiles of mRNAs targeted by p53-regulated miRNAs
such as miR-34a220–222. For example, CDK4, CDK6, cyclin E2, E2F3, c-MYC and n-MYC are
all downregulated by miR-34a220–222. p53 also promotes the expression of miR-145 such that it
can downregulate MYC, CDK4 and CDK5229. As c-MYC is also repressed by p53 independently
of miRNAs230, miR-145 and miR-34a engage in a feedforward loop that ensures the robust
suppression of c-MYC expression. It therefore follows that their downregulation in cancer would
lead to the relief of inhibition over a large number of cell cycle regulators that can promote
tumorigenesis (Figure 1.9).
In addition to proliferation, the miR-34 family has been shown to induce apoptosis, when
upregulated, in multiple cell lines, likely by repressing the expression of many mRNA targets
Figure 1.9 Regulation of the cell cycle by p53-induced miRNAs217. Cell cycle regulators and effectors
repressed by p53-induced miRNAs are grouped according to the cell cycle phase in which they are mainly
expressed or most functionally relevant. p53-induced miRNA mediated inhibition of these factors may cause or
contribute to cell cycle arrest in the respective phase of the cell cycle in which they are required. Downregulation
of these miRNAs may therefore promote cell cycle progression. Originally published by Hermeking, H. Nat. Rev.
Cancer 12, 613–626 (2012)217. Used with permission of the author and publisher.
48
including B-cell lymphoma 2 (BCL2)222, survivin231, sirtuin 1 (SIRT1)232 and snail family
transcriptional repressor 1 (SNAI1)233. As apoptosis is a fundamental feature of ageing
cardiomyocytes, Boon et al. (2013) investigated if the p53-dependent miR-34 family played a
role in cardiac ageing234. They discovered that the miR-34 family is significantly upregulated in
aged mouse hearts, and that miR-34 expression, but not muscle specific miR-1 or miR-133
expression, was correlated with age in human cardiac biopsies235. With this knowledge, they
inhibited the miR-34 family using antagomirs in 18 month old mice, and saw that this treatment
significantly reduced age-associated cell death in the heart235. Young mice treated with anti-miR-
34 were phenotypically similar to their littermates, whereas old anti-miR-34 treated mice
exhibited less hypertrophic cardiomyocytes, reduced cardiomyocyte apoptosis and preserved
cardiac function, compared to non-treated littermates235. They concluded that the upregulation of
miR-34 by p53 in aged hearts contributes to the progressive decline in heart function throughout
the aging process235. Since this study, it was shown that p53-dependent miRNAs are not only
involved in cardiac ageing, but that circulating p53 responsive miRNAs become upregulated in
the serum of patients post-AMI who subsequently develop heart failure103. Matsumoto et al.
(2013) found that miR-192, miR-194 and miR-34a all become upregulated in the serum of
patients from the OACIS study who developed heart failure following myocardial infarction236.
As the development of heart failure following myocardial infarction is positively correlated with
infarct size, it follows that a larger stressor may more strongly activate the p53 stress response,
leading to an upregulation of p53-responsive miRNAs in these patients. They concluded that
these miRNAs may be of value for predicting the risk of heart failure development following
myocardial infarction103.
Based on these studies, understanding the role of p53-regulated miRNAs can provide insight into
the mechanisms of aging and cardiovascular disease in the heart. Furthermore, it may be possible
to exploit the function of p53-regulated miRNAs in the heart for the development of cardiac
related therapies. Knowledge gaps reside in our understanding of whether MDM2 plays a role in
miRNA regulation, and which miRNAs are regulated by the p53/MDM2 tumor suppressor
complex specifically in the heart under physiological conditions of low stress. Identifying the
specific miRNAs which are regulated by p53/MDM2 in the heart may provide more insight into
49
how these proteins exert their crucial homeostatic roles of preserving cardiac function,
bioenergetics and cardiomyocyte cell cycle exit.
50
Chapter 2
Rational Hypothesis
2 Hypothesis and Thesis Aims
Adult cardiomyocytes are terminally differentiated cells that exist in a non-proliferative, post-
mitotic state237,238. Cell cycle exit occurs shortly after birth, and coincides with a concomitant
decrease in the expression of cell cycle activators and increase in the expression of cell cycle
inhibitors, resulting in adult cardiomyocytes that are refractory to proliferative stimuli81. Billia et
al. (2016) have demonstrated that inactivating both the tumor suppressor p53 and its negative
regulator MDM2, specifically in the heart (double knockout mouse, DKO), can allow
differentiated cardiomyocytes to regain their proliferative capacity, through an upregulation of
factors involved in cell cycle re-entry (cyclins and cyclin-dependent kinases) and a
downregulation of cell cycle inhibitors. Specifically, DKO mice exhibited a significant
upregulation in CDK2 kinase activity, as well as an upregulation in the expression of cyclin E,
both of which are crucial for cell cycle reactivation. Concurrently, the expression of p21Cip1 and
p27Kip1, two major inhibitors of cell cycle progression, was downregulated by up to 90%. These
changes within the cell were accompanied by a significant increase in cardiomyocytes in all
phases of the cell cycle, including cytokinesis. Importantly, knockout of either gene individually
(p53KO and Mdm2KO) failed to promote cardiomyocyte cell cycle re-entry. Further
transcriptome analysis revealed that loss of p53 and MDM2 caused a widespread upregulation in
cell cycle regulators, indicating that p53/MDM2 repress their expression in wild type
cardiomyocytes, promoting their quiescence. The ability of p53 to directly repress gene
expression, however, is debated. What is known is that p53 can mediate transcriptional
repression indirectly, through the activation of microRNAs (miRNAs). As many miRNAs have
been shown to function as potent tumor suppressors, we hypothesized that in the heart, there are
a subset of miRNAs regulated by the dual presence of both p53 and MDM2, which function to
downregulate important cell cycle activators and maintain adult cardiomyocyte quiescence.
Upon knockout of p53 and MDM2 in the heart, the expression of this subset of p53/MDM2-
regulated miRNAs is lost, which alleviates the inhibition on multiple cell cycle regulators and
permits cardiomyocytes to regain proliferative capacity.
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2.1 Hypothesis
In the heart, p53 and MDM2 regulate a set of “anti-proliferative” microRNAs that maintain
cardiomyocyte cell cycle arrest. Inactivation of these p53/MDM2-regulated miRNAs will
alleviate their inhibition of crucial cell cycle regulators and thus, promote cardiomyocyte cell
cycle re-entry and proliferation.
2.2 Thesis Aims
Aim 1: Identification of p53/MDM2-regulated miRNAs involved in the regulation of cell cycle.
a) Perform a miRNA microarray analysis comparing hearts from p53KO,
Mdm2KO and DKO mice to identify miRNAs regulated by both p53 and
MDM2.
b) Perform target gene analysis of identified p53/MDM2 regulated miRNAs to
determine their involvement in cell cycle regulation.
c) Develop antagomirs that can inhibit identified “anti-proliferative” p53/MDM2-
regulated miRNAs.
Aim 2: Inhibition of identified miRNAs in vitro in rat neonatal cardiomyocytes using
antagomirs.
a) Transfect cardiomyocytes with a cocktail of antagomirs targeting the identified
p53/MDM2-regulated miRNAs and determine if this treatment promotes
cardiomyocyte proliferation.
b) Determine if putative miRNA target genes involved in cell cycle regulation
become upregulated following antagomir treatment to elucidate which factors
support cardiomyocyte cell cycle re-entry.
52
Chapter 3
Data Chapter
3 Control of Cardiomyocyte Proliferation by p53/MDM2-regulated microRNAs
Authors: Shanna Stanley-Hasnain1,2, Ludger Hauck2, Daniela Grothe2, and Filio Billia1,2,3,4*
Affiliations: 1Institute of Medical Science, University of Toronto, 1 King's College Circle, Toronto, Ontario Canada M5G 1A8 2Toronto General Research Institute, Toronto, Ontario Canada, 100 College St., M5G 1L7 3 Division of Cardiology, University Health Network (UHN), Toronto, Ontario, Canada, 200
Elizabeth St., Toronto, Ontario Canada, M5G 2C4 4Heart and Stroke Richard Lewar Centre of Excellence, University of Toronto
Formatted for submission to the journal Molecular Therapy – Nucleic Acids
53
3.1 Introduction
Trp53 (p53) is a tumor suppressor gene encoding a transcription factor that is the most frequent
target for mutations in tumors168. In addition to its tumor suppressive abilities, p53 acts as a
cellular stress sensor239–241, and plays an important role in homeostasis213 and senescence242–244.
It can trigger transient or permanent cell cycle arrest, apoptosis or DNA repair in response to
many stimuli including hypoxia, DNA damage, ribonucleotide depletion and nutrient starvation,
and is involved in the regulation of metabolism and autophagy168,245. Based on the observations
that p53-knockout mice develop early onset spontaneous tumors with 100% penetrance171, that
individuals with Li-Fraumeni syndrome who inherit a mutant p53 allele have heightened
susceptibility for tumorigenesis169, and that cells with dysfunctional p53 exhibit aberrantly high
levels of proliferation246, the importance of p53’s role in growth control is unequivocal.
Due to the vast effect that p53 can exert within the cell, it is imperative that its activity is tightly
regulated. Post-translational ubiquitination is one of the means by which to control p53 stability.
This is largely mediated by the oncoprotein murine double minute 2 (MDM2)247. MDM2 is an
E3 ubiquitin ligase that monoubiquitinates p53 at multiple lysine residues, a critical step in
promoting its degradation by nuclear and cytoplasmic proteasomes196,197,199. Interestingly,
MDM2 itself is the product of a p53 inducible gene, creating a negative feedback loop that
tightly maintains p53 levels within the cell197. In turn, low levels of p53 activity result in
constitutively low MDM2 levels197. This feedback loop allows the cell to limit the duration and
severity of p53-dependent biological responses199. Unlike p53 knockouts, MDM2-null mice
experience a very early embryonic lethality, due to high levels of p53, impairing organ
development through an excessive induction of apoptosis and diminished cell proliferation204,205.
The most thoroughly characterized biochemical role of p53 within the cell is as a transcriptional
activator, however it also is known to repress transcription of certain genes (estimated ~15% of
target genes). As a transcription factor, p53 mediates cell cycle arrest through activation of
important cell cycle inhibitors168. Cell cycle target genes transcriptionally regulated by p53
include cyclin-dependent kinase inhibitor 1A (p21Cip1), and upregulation of p21Cip1 expression by
p53 leads to G1 phase cell cycle arrest, through p21Cip1-mediated inhibition of cyclin-dependent
kinase 1 (CDK1), cyclin-dependent kinase 2 (CDK2) and proliferating cell nuclear antigen
54
(PCNA)174,175. Further genes activated by p53 which have cell cycle regulatory functions include
the protein tyrosine phosphatase receptor type V (PTPRV) which is also involved in G1 arrest176,
growth arrest and DNA-damage inducible 45 (GADD45a)177 and Stratifin (YWHAS)178 which
control the G2/M transition175.
In contrast, p53-mediated transcriptional repression has been a contentious area of research, and
direct mechanisms of p53 repression have recently been refuted in a computational meta-analysis
of 6 genome-wide analyses of p53 dependent gene expression, published by Fischer et al.
(2014)248. Regardless, it has been noted that activation of p53 leads to the downregulation of cell
cycle activators, beyond those targeted by p21Cip1 activation228.
Recently, microRNAs (miRNAs) have been suggested as the main mediators of p53-regulated
gene repression228. miRNAs are a group of small non-coding RNAs with the capability of
expansive genetic regulation. This class of endogenous 21-25 nucleotide long, single-stranded
RNA molecules function as guide molecules in RNA silencing through base pairing to their
complementary mRNA targets82–84. By targeting specific mRNAs for degradation or translational
repression, miRNAs act as inhibitors of gene expression, and are involved in almost all
developmental and pathological processes in animals, including multiple pathways regulated by
the tumor suppressor p5382. Hunten et al. (2015) demonstrated that upon p53 activation, 111
miRNAs become upregulated, indicating that there is subset of miRNAs transcriptionally
activated by p53 (p53-regulated miRNAs)228. Furthermore, in this study, of the 415 genes
downregulated following p53 activation, 50% contained complementary seed-matching
sequences to p53-regulated miRNAs in their 3’-UTRs228. This supports the notion that p53-
dependent gene repression can be mediated indirectly through p53-regulated miRNAs.
Many p53-regulated miRNAs have been shown to have robust effects on cell cycle, and
themselves act as tumor suppressors217. Numerous studies have demonstrated the downregulation
of distinct p53-regulated miRNAs by genetic or epigenetic mechanisms in diverse human
tumors, and that their re-introduction through miRNA mimic technology can mitigate tumor cell
proliferation and improve cancer outcomes249–252. Cell cycle-related factors have
bioinformatically been shown to be over-represented in the profiles of mRNAs targeted by p53-
regulated miRNAs such as miR-34a220–222. For example, cyclin-dependent kinase 4 (CDK4),
55
cyclin-dependent kinase 6 (CDK6), cyclin E2, E2 factor 3 (E2F3), V-Myc avian
myelocytomatosis viral oncogene homologs (c-MYC and n-MYC) are all downregulated by
miR-34a220–222. p53 also promotes the expression of miR-145 such that it can downregulate c-
MYC, CDK4 and cyclin-dependent kinase 5 (CDK5)229. Furthermore, certain p53-regulated
miRNAs have been shown not only to play a role in cancer, but in other diseases such as
cardiovascular disease103,253. As miRNAs are involved in a myriad of signaling and biological
processes which regulate growth and regeneration, cardiac function, homeostasis, and the
progression of cardiac remodeling, hypertrophy and heart failure, exploiting their function
through miRNA based therapies have been previously studied in cardiovascular disease and heart
failure254.
Heart failure is a leading cause of morbidity and mortality in Canada. Adult cardiomyocytes are
terminally differentiated cells that exist in a non-proliferative, post-mitotic state237,238. Thus, the
regenerative capacity of the heart is poor, and cardiac tissue is particularly vulnerable to injury.
This is emphasized by the fact that cardiomyocyte loss due to insults such as myocardial
infarction is a major contributing factor to the development of heart failure255,256. As the quality
of life and prognosis for these patients is poor, therapies to prevent and treat heart failure are
urgently needed28. Therefore, determining the roadblocks responsible for maintaining
cardiomyocyte cell cycle exit lies at an important crossroad in the development of potential
regenerative therapies to promote cardiac repair following injury. Given p53’s crucial role in
promoting growth arrest, Billia et al (2016) sought to determine the function of the p53/MDM2
tumor suppressor circuitry in the heart213,214. They demonstrated that inactivation of both p53
and MDM2 specifically in the heart (double knockout mouse, DKO) can allow differentiated
cardiomyocytes to regain proliferative capacity, through an upregulation of factors involved in
cell cycle re-entry (cyclins and cyclin-dependent kinases) and a downregulation of cell cycle
inhibitors. Importantly, knockout of either two genes individually (p53KO and Mdm2KO) was
not sufficient to promote cardiomyocyte cell cycle re-entry. Based on p53’s known ability to
indirectly repress gene expression through the activation of miRNAs, and the ability of miRNAs
to function as potent tumor suppressors, we hypothesized that in the heart, there are a subset of
miRNAs regulated by the dual presence of both p53 and MDM2, that function to downregulate
important cell cycle activators and maintain adult cardiomyocyte quiescence. Upon knockout of
56
p53 and MDM2 in the heart, the expression of this subset of p53/MDM2-regulated miRNAs is
lost, which alleviates the ihibition on multiple cell cycle regulators and permits cardiomyocytes
to regain proliferative capacity.
The aim of this study was to characterize the effect of p53 and MDM2 in the heart through their
ability to regulate the expression of miRNAs. Herein, we identified a unique subset of miRNAs
that become downregulated upon the loss of both p53 and MDM2, in the heart. This subset of
miRNAs were enriched for target genes involved in cell cycle regulation, and their inhibition
using antagomirs significantly promoted wild type neonatal rat cardiomyocyte proliferation.
These data support the existence of a novel role for p53 and MDM2 in the heart, in maintaining
cardiomyocyte cell cycle arrest through regulating a subset of anti-proliferative miRNAs.
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3.2 Methods
3.2.1 MDM2 and p53 conditional mutant mice
All animal usage in this study was in accordance with approved institutional animal care
guidelines of the UHN (AUP 1815/1379, Canadian Council in Animal Care). The mcm
transgenic mice on a C57BL/6J background were obtained from Jackson (Bar Harbor,
ME04609 USA; strain name: B6.FVB(129)-Tg(Myh6-cre/Esr1*)1Jmk/J; stock number:
005657). The Mdm2f/f mice (strain number 01XH9) and p53f/f mice (strain number 01XC2)
were obtained from the mouse repository of the National Cancer Institute/National
Institutes of Health at Frederick (Rockville, MD 20852 USA). The Mdm2f/f and p53f/f strains
were backcrossed onto C57BL/6J inbred mice (Jackson; stock number 000664) for at least eight
generations. Age-matched syngeneic adult male mice (12-13-week-old; 22-27 g body weight)
were used in this study. All experiments utilized controls of matched age and sex. Genotyping
was carried out by PCR employing alkaline hydrolysis of genomic DNA isolated from tail tips
using the Terra PCR Direct Kit (mcm, Mdm2f/f, p53f/f; Clontech) or AccuStart II GelTrack PCR
Supermix and the following oligonucleotides:
mcm forward 5’-AGGTGGACCTGATCATGGAG-3’
mcm reverse 5’-ATACCGGAGATCATGCAAGC-3’
Mdm2f/f forward 5’-CTGTGTGAGCTGAGGGAGATGTG-3’
Mdm2f/f reverse 5’-CCTGGATTTAATCTGCAGCACTC-3’
p53f/f forward 5’-CACAAAAACAGGTTAAACCCAG-3’
p53f/f reverse 5’-AGCACATAGGAGGCAGAGAC-3’
To achieve inactivation of MDM2 and p53 in the adult mouse heart, we used the Cre-loxP
recombination system of bacteriophage P1. We employed an inducible cardiomyocyte-specific
transgenic mouse in which the cardiac muscle D-myosin heavy chain 6 (Myh6) promoter
drives the expression of tamoxifen (Tam)-inducible Cre recombinase protein fused to two
mutant estrogen- receptor ligand-binding domains (mcm). In this strain, the mcm fusion
protein is expressed only in cardiomyocytes, not in non-cardiomyocytes, and is retained in the
58
cytoplasm. Administration of Tam induces mcm nuclear translocation thereby permitting Cre-
mediated recombination. This system allowed us to explore conditional mutations of
p53/Mdm2 that might differ from the germ line absence of both of these factors in the adult
heart. Next, we crossed mcm transgenic mice with mice carrying conditional Mdm2f/f and
p53f/f alleles to obtain Mdm2f/f; p53f/f;mcm (DKO), Mdm2f/f;mcm (Mdm2KO) and p53f/f;mcm
(p53KO) mice on a C57BL/6J background. In Mdm2KO mice, Mdm2 exons 7 to 9 are flanked
with loxP sites to facilitate inactivation of the RING finger domain which exerts ubiquitin
ligase activity towards p53. p53KO mice carry loxP sites in p53 introns 1 to 10 to ensure Cre-
mediated elimination of the majority of the coding sequence. In the absence of Tam, animals of
all these strains develop normally and lack an obvious phenotype.
We injected an ethanol/peanut oil (P2144, Sigma-Aldrich) emulsion of 4-Hydroxytamoxifen
(H6278, Sigma-Aldrich) intraperitoneally. Tam-treated adult Mdm2f/f, p53f/f, Mdm2f/f; p53f/f and
mcm mice were analyzed by immunohistology and echocardiography rendering no evidence of
fibrosis or cardiac dysfunction. To achieve conditional genetic ablation of MDM2 and p53
in adult cardiomyocytes in vivo, Tam was intraperitoneally injected for four consecutive days.
The day of the fourth Tam injection was arbitrarily set as day zero.
3.2.2 RNA Isolation
Total RNA, including the small RNA fraction, was isolated from mouse hearts (n= 3 hearts per
strain) or neonatal rat cardiomyocytes using the standard Trizol method (Invitrogen). To isolate
RNA from cultured neonatal rat cardiomyocytes, Trizol was also used. 300 µl of trypsin was
added per well of a 6 well plate and incubated for 5 minutes at 37 degrees Celsius. The reaction
was halted by 700 µl of media containing 10% horse and fetal bovine serum. Cells were
subsequently scraped and pelleted by centrifugation. 500ml of Trizol was added to each cell
pellet, and a 28 gauge syringe was utilized to rupture the cell membranes and release the nucleic
acids. Phase Lock Gels (5PRIME) were utilized to ensure a pure and distinct organic and
aqueous phase separation (gel creates a physical barrier between the two phases), leading to the
isolation high quality non-contaminated RNA. Cardiomyocyte and fibroblast RNA concentration
and purity was measured both by the NanoDrop 2000 spectrophotometer (Thermo Scientific)
and the Qubit 2.0 fluorometer (Thermo Fisher Scientific) prior to utilization for RT-qPCR. The
59
RNA isolated from mouse hearts which was used for the Nanostring miRNA assay was passed
through a Bioanalyzer (Agilent) by the Princess Margaret Genomics Centre core facility to more
accurately assess the quality of the RNA. Only RNA with an RNA Integrity Number (RIN) of
≥9.0 (out of 10), was utilized for the miRNA assay.
3.2.3 Nanostring nCounter miRNA assay
33 ng of total RNA from each heart was used as input for the nCounter mouse miRNA assay and
was and processed according to the manufacturer’s protocols. For each sample, a high-density
scan (1155 fields of view) was performed. Raw array data was processed using the Nanostring
nSolver Analysis Software 2.5. The mean of the negative controls plus one standard deviation
was subtracted for background correction. Normalization of the data for sample/RNA content
was performed using the geometric mean of the top 100 most highly expressed genes.
Normalized miRNA expression levels were transformed from ratios (treatment vs. vehicle
control) to fold change values and analyzed using a t-test to identify significantly differentially
expressed miRNA between samples. Heatmaps were generated with the fold change values using
Euclidean distance as the distance metric and the average as the linkage method. Pathway
analysis was executed with the online platform miRSystem ver. 20150312.
3.2.4 miRNA RT-qPCR
Total RNA isolated from total hearts, cardiomyocytes or fibroblasts was reverse transcribed
using the qScript microRNA cDNA Synthesis Kit (Quanta BioSciences, Inc.) using an initial
input of 500ng of RA. The RNA first undergoes a polyadenylation step, followed by reverse
transcription with an oligo-dT adapter primer which has a unique sequence at its 5’ end allowing
amplification of the cDNAs in quantitative PCR reactions (qPCR). qPCR is carried out using the
Roche Light Cycler 480, a PerfeCTa miRNA assay unique for each individual miRNA along
with the PerfeCTa Universal PCR Primer (specific to the unique sequence of the oligo-dT
adapter primer) and PerfeCTA SYBR Green SuperMix (5ng cDNA/25µl reaction). Relative
miRNA expression and fold change was calculated by the standard ΔΔCt method, and was
normalized to the endogenous control RNU6 (small RNA component of spliceosome) using the
LC480 SW 1.5.1 analysis software.
60
3.2.5 Western Blot
Protein was isolated from mouse hearts and neonatal rat cardiomyocytes using the NE-PER
Nuclear and Cytoplasmic Extraction Kit (ThermoFisher) according to the manufacturer’s
protocol. For Western Blot, we used SDS/PAGE gels (4–12% Bolt Bis-Tris Plus; Invitrogen),
PVDF membranes (iBlot; Invitrogen), primary antibodies against p53 (Enzo) and Mdm2
(Sigma), horseradishperoxidase (HRP) conjugated secondary antibodies (Cell Signaling), and the
Luminata Crescendo HRP substrate for chemiluminescence detection (EMD Millipore). Blots
were imaged with the BioRad ChemiDoc XRS+ and analyzed with Image Lab 5.0 software.
3.2.6 Neonatal rat cardiomyocyte and fibroblast isolation and culture
Left ventricular cardiomyocytes from 2 to 3 day old wild type Wistar rats (Charles River) were
isolated using collagenase II (0.5 mg/mL, Invitrogen) and pancreatin (1 mg/mL, Sigma). To
obtain highly pure cardiomyocyte cultures, one round or “pre-plating” was utilized. The cardiac
digestion mixture was plated onto 10cm dishes and incubated for 50 minutes. This is sufficient
time to allow cardiac fibroblasts to adhere to the plates, but not cardiomyocytes, as described
previously257. The supernatant containing a purified population of cardiomyocytes was removed
from the 10 cm plates and plated onto Collagen I coated cell culture dishes (1x106 cells/well in 6
well plates and150,000 cells/well in 48 well plates) (Corning). Cardiomyocytes were cultured in
DMEM/F12 medium containing 3 mM Na-pyruvate, 2 mM glutamine, antibiotics (Gibco), 0.2%
bovine serum albumin (Sigma), 0.1 mM ascorbic acid, 10% fetal bovine serum (Sigma), 10%
horse serum (Sigma) and 25 µM arabinosylcytosine (AraC; Sigma) to inhibit remaining non-
cardiomyocyte proliferation for 48 hours prior to treatment or analysis. 500,000 fibroblasts were
seeded/well in 6 well plates, and cultured in DMEM/F12 medium containing 10% fetal bovine
serum (Sigma), 10% horse serum (Sigma) and antibiotics. Cells were cultured to confluency
prior to isolation of RNA for RT-qPCR.
3.2.7 miRNA antagomir development
miRIDIAN microRNA Hairpin Inhibitors were ordered from Dhamarcon, Thermo Scientific
(5nmol). The Anti-cel-67 targeting a Caenhabdoartisi elegans miRNA not expressed in
mammals was employed as a negative control.
61
3.2.8 Neonatal rat cardiomyocyte transfection
Cardiomyocytes were transfected using the TransIT TKO Transfection Reagent (Mirus Bio) and
50 nM of BLOCK-iT Alexa Fluor 555 Red Fluorescent Oligo (Positive Control) to determine
transfection efficiency. Transfection efficiency was assessed fluorescent microscopy, calculating
the number of MEF2A positive, Alexa Fluor 555 positive cells/well. Transfection of antagomirs
(25nM each, Dhamarcon) was carried out using the standard transfection protocol provided by
Mirus Bio., 48 hours following plating of cells, in cardiomyocyte media (DMEM/F12)
containing 1% fetal bovine serum, 1% horse serum and antibiotics. 24 hours after transfection,
media was replaced, and 48 hours following transfection, cells were fixed for
immunofluorescence or RNA was isolated.
3.2.9 Cell Cycle RT-qPCR Array
RNA was purified using the method described above from cardiomyocytes transfected with
the cocktail of all 11 miRNA antagomirs (25nM each), the optimized final 9 antagomir
cocktail (25nM each), or a negative control (cel-67 100 nM) RNA concentrations were
determined using a NanoDrop 2000 spectrophotometer (Thermo Scientific). Equal amounts of
RNA (400ng/sample) were converted into cDNA using the First Strand cDNA Synthesis
(Qiagen) kit. The PCR Array profiled the expression of 87 cell cycle genes, 5 housekeeping
genes, controls for genomic DNA contamination, and efficiency of both the (RT-PCR: PCR)
reactions
3.2.10 Immunofluorescence
Cardiomyocytes were fixed with 4% PBS-buffered formalin for 7 minutes, permeabilized with
0.5% Triton-X-100 in 1X-Tris-buffered saline. Cells were stained for 2 hours with primary
antibodies diluted in 0.5% Triton-X-100 in 1X-Tris-buffered saline: Aurora B (Sigma), Gata4
(BD), Phalloidin (Thermo Fisher Scientific). Cells were then washed and incubated for 45
minutes in their respective secondary antibodies conjugated to Alexa Fluor- and Alexa Fluor-555
(Life Technologies) and Genomic DNA was stained with 4',6-diamidino-2-phenylindole
dihydrochloride (DAPI; Life Sciences). Finally, cells were fixed again with 4% PBS-buffered
formalin for 7 minutes. Cells were imaged using the Zeiss Axio Observer inverted fluorescent
microscope. Images were acquired with the MetaMorph Microscopy Automation & Image
62
Analysis Software and analyzed using ImageJ. The cells were also run through the Hidex Sense
microplate reader to detect Alexa Fluor-488 and 555 fluorescence levels (measured in Relative
Fluorescent Units; RFU).
3.2.11 Statistical Analyses
Numerical data are expressed as either the ratio change or % change of the means ± the standard
error of the mean (SEM). The Student’s t-test was utilized to determine significance between
experiments with only 2 variables. One-way analysis of variance (ANOVA) with Tukey’s
Multiple Comparison Post-Test was employed to analyze data between multiple independent
(unrelated) groups using the GraphPad Prism 5.01 software. Level of significance was first
determined by ANOVA analyses (p < 0.05), and subsequently significance between every
pairwise group in the experiment was determined by Tukey’s Multiple Comparison Test.
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3.3 Results
3.3.1 The tumor suppressor circuitry p53/MDM2 regulate expression of a unique subset of miRNAs within the heart.
To evaluate the role of p53 and MDM2 as regulators of miRNA expression within the heart, total
miRNA expression was profiled from cardiac-specific conditional p53/MDM2 DKO mouse
hearts, and was compared to single knockouts (p53KO and Mdm2KO), and vehicle injected
controls, using the Nanostring nCounter miRNA platform (n=3 hearts per group). p53 and
MDM2 knockout following tamoxifen injections was confirmed for all three strains by Western
blot (Supplemental Figure 3.1).
The nCounter miRNA array profiled over 600 mouse miRNAs. Heat map construction by
agglomerative clustering (the "bottom up" hierarchical clustering approach where each miRNA
starts in its own cluster, and pairs of clusters are merged when moving up the hierarchy), using
the nSolver analysis software, revealed that the Mdm2KO and DKO miRNA profiles were
closely related, whereas the p53KO’s miRNA profile was distinct from the other two knockouts
(Figure 3.1 A). Furthermore, the p53KO miRNA profile was more closely related to the miRNA
prolife of vehicle- injected control hearts, whereas the profiles in the Mdm2KO and DKO hearts
were more distantly related (Figure 3.1 G). Setting the miRNA expression fold change cut off as
greater than 1.2 or less than -1.2, a total of 89 significantly changed miRNA transcripts out of
600 (~15%) were identified in the three mutant hearts compared to the vehicle- injected controls,
with a P-value of ≤ 0.01 (Figure 3.1 B) (p53KO: 3 up, 12 down, Mdm2KO: 29 up, 19 down,
DKO: 36 up, 29 down).
The top 20 differentially regulated miRNAs were determined for each strain. Members of the
miR-15 family, miR-15a and miR-16, well-characterized p53-regulated miRNAs258, were in the
top 12 downregulated miRNAs of the p53KO. These miRNAs were also significantly
downregulated in the DKO, but were upregulated in the Mdm2KO, likely due to the known
higher levels of p53. This highlights the importance of MDM2 in maintaining normal levels of
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p53-regulated genes within the cell by dampening p53’s activity. Interestingly, miR-1 and miR-
133b, classic muscle specific miRNAs which play a critical role in myogenic differentiation259,
were in the top 10 downregulated miRNAs of DKO mouse hearts. miR-1 was also in the top 10
downregulated miRNAs in Mdm2KO hearts. Furthermore, loss of p53 seemed to have little
effect on miRNA upregulation, as the expression of only a total of 3 miRNAs was significantly
increased in the p53KO. Conversely, in the Mdm2KO and DKO, a total of 29 and 36 miRNAs
respectively, were significantly upregulated compared to control hearts (Figure 3.1 D-F).
Although the change in expression of many miRNAs overlapped between the three different
strains, each knockout also had a subset of uniquely up and downregulated miRNAs (Figure 3.1
C). For this study, the subset of interest were the miRNAs uniquely downregulated in the DKO
strain, because we sought to determine if inhibition of these miRNAs in wild type
cardiomyocytes could promote proliferation. In DKO mouse hearts, 15 uniquely downregulated
miRNAs were identified with a significant fold change of less than -1.2, that were not
significantly changed either of the single knockout mice. These became the target miRNAs of
interest, and were defined as p53/MDM2-regulated miRNAs (Table 3.1).
Figure 3.1. Significant miRNA transcriptional changes in the absence of p53 and/or MDM2
create unique miRNA profiles. (A) Agglomerative clustering was preformed to generate a heat
map demonstrating miRNA fold change in the heart between the three different strains post Tamoxifen injection vs. the vehicle- injected control. 600 miRNAs were profiled using the nCounter mouse miRNA panel (Nanostring). Fold change values are depicted by color intens ity,
where blue = repressed and red = induced. Mice were analyzed at 6 months post-Tam (p53KO), 14d post Tam (Mdm2KO) and 8d post-Tam (DKO), n=3 mice per strain. (B) Heat map
representing significant (P≤0.01) miRNA fold changes between the three different strains post Tamoxifen injection vs. control with a fold change ≥ 1.2 (±) . 81 miRNA transcripts (rows) out of 600 (13.5%) were significantly changed in at least one of the three strains (columns). (C) Venn
diagram analysis illustrating the overlap of significantly up and downregulated miRNAs between the 3 KOs vs. control. The 15 miRNAs uniquely downregulated in the DKO are the target
miRNAs of interest. (D-F) Top 20 differentially regulated genes in (D) p53KO vs. control, (E)
Mdm2KO vs. control and (F) DKO vs. control. (P≤0.01) (G) Scatter plot matrix representing normalized Log2 counts for each miRNA profiled of p53KO (green), Mdm2KO (pink) and DKO
(blue) (Y axis) compared to Log2 counts of vehicle injected control (X axis). Each point represents one miRNA. R2 values indicate that the p53KO miRNA profile is more closely related
to that of the control than of the Mdm2KO or DKO profiles.
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Table 3.1. miRNAs significantly downregulated in DKO vs. vehicle control. Using the nCounter Analysis Software, miRNAs with a fold change ≥ -1.2 and p value of ≤ 0.01 that were
uniquely downregulated in the DKO vs. vehicle control were determined. Mmu-miR-1931 had no predicted target genes (determined using the miRSystem online software) and was thus eliminated from further analysis. mmu-miR-133a and mmu-miR-133b did not target genes within the cell
cycle pathway (determined using the miRSystem software) and were also eliminated from further analysis. mmu-miR-212 expression change determined by Nanostring array was not validated by
RT-qPCR (See Figure 2) and was therefore also removed from further analysis. The 11 remaining miRNAs are the final “hits” from the miRNA screen.
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3.3.2 p53/MDM2-regulated miRNAs are enriched for mRNA gene targets involved in cell cycle progression.
To characterize enriched pathways and functions regulated by the target genes of the 15
p53/MDM2-regulated miRNAs identified by the Nanostring array, the online miRNA pathway
analysis platform, miRSystem, was employed (Figure 3.2 B-C). miRSystem integrates 7
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miRNA target gene prediction algorithms (DIANA, miRanda, miRBridge, PicTar, PITA, rna22
and TargetScan) as well as 2 experimental validation databases to predict miRNA target genes.
A valid miRNA target gene is required to be identified by 4 out of the 7 prediction programs. A
P-value is then assigned to each biological pathway (KEGG and Biocarta pathways) represented
by the target genes of the input group of miRNAs. This P-value is based on how many miRNA
target genes are found within a given pathway and the overall number of genes within that
pathway. Enriched biological pathways are subsequently ranked on a score calculated by
multiplying the P-value of a given pathway by the weighted expression values of the miRNAs
that target genes within this pathway. (Supplemental Figure 3.8)260.
In this analysis, miR-1931 did not yield any predicted target genes and was eliminated from
analysis. Only 2 out of the 14 remaining miRNAs (muscle specific miR-133a and miR-133b) did
not have target genes within the cell cycle pathway and were also removed from further analysis,
whereas 12/14 miRNAs targeted at least one gene within this pathway. To validate that the 12
miRNA hits were truly downregulated in the DKO compared to control hearts as seen by the
Nanostring array, RT-qPCR was performed using the same RNA sampled for the array. RT-
qPCR revealed that 11/12 miRNAs were significantly downregulated in the DKO compared to
the vehicle control. Only 1 miRNA, miR-212, showed an opposite expression pattern than what
Figure 3.2. miRNAs downregulated in the DKO are enriched for target genes within the
cell cycle pathway. (A) Reverse transcription and quantitative PCR (RT-qPCR) was performed
to validate miRNA expression values measured by the Nanostring nCounter mouse miRNA
panel. All 12 miRNA hits of interest were measured. Data are represented as the miRNA
expression ratio change in DKO mouse hearts compared to controls for each primer using the
ΔΔCt method ± s.e.m, n=3. A significant difference between the mean normalized delta Ct
values for the DKO vs. control is indicated by * p ≤ 0.05, ** p ≤ 0.01, *** p ≤ 0.001 determined
by the Student’s t-test. The 11 validated miRNAs correspond to the final miRNA hits extracted
from the Nanostring microarray, whereas miR-212 was removed from further analysis. (B-C)
miRSystem Software was utilized to determine the predicted target genes of the 11 miRNA hits
within the KEGG cell cycle pathway (125 total genes) (B) Percent of each miRNA’s total target
genes found within the cell cycle pathway (*P≤0.05, **P≤ 0.01). (C) The number of cell cycle
activating and inhibiting genes targeted by the 11 miRNA hits.
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was determined by the array, and was upregulated compared to vehicle control (Figure 3.2 A).
As the expression level of this miRNA could not be confirmed, it was eliminated from further
study. Cell cycle target genes of the remaining 11 miRNAs were subsequently investigated in
further depth. The most enriched biological pathways that the target genes of these miRNAs
Figure 3.3 Genes within the cell cycle pathway are redundantly targeted by the miRNAs
downregulated in the DKO. (A) Schematic of the 26 cell cycle activating genes targeted by the
12 miRNA hits as determined by the miRSystem software. 81% of cell cycle genes targeted play a role in cell cycle activation. (B) Schematic of the 6 cell cycle inhibiting genes targeted by the
12 miRNA hits as determined by the miRSystem software. 19% of cell cycle genes targeted play a role in cell cycle inhibition.
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belong to are illustrated in Supplemental Figure 3.2. The cell cycle pathway was within the top
30th percentile of biological pathways for this group of miRNAs. Target genes of these miRNAs
also belonged to many other biological pathways that regulate the cell cycle, including mitogenic
signaling pathways, the wingless-type MMTV integration site family (WNT) signaling pathway,
and pathways involved in regulating the actin cytoskeleton. These 11 miRNAs were enriched
for target genes that function directly within the cell cycle. For example, approximately 10% of
miR-30e’s target genes were found within this pathway (Figure 3.2 B). Furthermore, there was a
high amount of redundancy between the 11 miRNAs and their targets within the cell cycle,
where some genes were targeted by up to 6 of the identified miRNAs (Figure 3.2 C). This
redundancy was apparent for target genes involved in cell cycle activation, whereas genes
involved cell cycle inhibition were only targeted by a maximum of 4 miRNAs. Furthermore, 26
out of the 32 miRNA target genes found within the cell cycle pathway were involved in cell
cycle activation (81%), whereas only 6 out of 32 were involved in cell cycle inhibition (Figure
3.2 C, Figure 3.3 A, B), suggesting that upon downregulation of these 11 miRNAs, the
inhibition over a battery of key cell cycle activating genes may be alleviated, promoting their re-
expression within the cell.
Therefore, the 11 validated miRNAs, uniquely downregulated in DKO mouse hearts vs. controls,
who target at least one gene within the cell cycle pathway were defined as the final “hits” from
this miRNA screen, and are listed in Table 3.2. These miRNAs were hypothesized to function as
cell cardiac cell cycle inhibitors.
Table 3.2. Final list of p53/MDM2-regulated miRNAs. List of the 11 miRNAs downregulated uniquely in DKO mouse hearts vs. controls with a fold change of ≥ -1.2 and a P value of ≤ 0.01
determined by the Nanostring miRNA array, that were validated by RT-qPCR, and target at least one gene within the cell cycle. These are the final 11 miRNAs selected to be inhibited by antagomirs, and
are defined as p53/Mdm2-regulated miRNAs.
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3.3.3 p53/MDM2-regulated miRNAs are novel cardiomyocyte cell cycle inhibitors
Primary rat neonatal cardiomyocytes were utilized as an in vitro model to explore the role of the
11 p53/MDM2-regulated miRNAs in modulating cardiac cell cycle arrest. To support the
hypothesized “anti-proliferative” function of these miRNAs, their expression levels were
examined in both wild type cardiomyocytes, which do not proliferate, and cardiac fibroblasts,
which are known to proliferate extensively in response to cardiac injury261. It was thus
hypothesized that the expression of these 11 cell “cycle-inhibitory” miRNAs would be
downregulated in cardiac fibroblasts compared to non-proliferative cardiomyocytes. RT-qPCR
revealed that the expression of all 11 miRNAs was significantly decreased in fibroblasts
compared to cardiomyocytes by up to 80% (Supplemental Figure 3.3). This strengthens the
notion that inhibition of these miRNAs in wild type cardiomyocytes could promote cell cycle re-
entry and proliferation, as they are found to be downregulated in cardiomyocytes that have re-
entered the cell cycle (in DKO mouse hearts), as well as in wild type fibroblasts which
proliferate naturally.
Antagomirs (chemically modified, cholesterol-conjugated, single-stranded RNA analogues
complementary to miRNAs that antagonize the function of miRNAs125,126), targeting the 11
p53/MDM2-regulated miRNAs were subsequently utilized to investigate if the identified
miRNAs regulate cell cycle in wild type cardiomyocytes..
To study the specificity and inhibitory effect of antagomirs in cardiomyocytes, we transfected
cells with 2 antagomirs (anti-miR-30b and anti-let-7f) separately (cardiomyocyte transfection
efficiency of 93% ± 6%, see Supplemental Figure 3.4). This was to determine if antagomirs are
cross-reactive between and within miRNA families, as 4 of the miRNA hits are part of the miR-
30 family, and 2 belong to the let-7 family. Antagomir inhibition of their target miRNA was
highly potent: anti-miR-30b and anti-let-7f downregulated the expression of miR-30b by 97%
(P≤0.0001, n=4) and let-7f by 99% (P≤0.0001, n=4) respectively, as measured by RT-qPCR.
Conversely, transfection of anti-miR-30b had no significant effect on the expression of let-7f.
Similarly, anti-let-7f did not significantly alter the expression of miR-30b in rat neonatal
cardiomyocytes (Figure 3.4 C). This suggests that these two antagomirs do not inhibit off target,
unrelated miRNAs. However, assessing the expression levels of other miR-30 and let-7 family
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members in cardiomyocytes transfected with anti-mir-30b and anti-let-7f, respectively, revealed
that that antagomirs downregulate off target miRNAs from within the same miRNA family as
their target miRNA to varying degrees of potency. For example, cardiomyocytes transfected with
anti-miR-30b also exhibited a significant downregulation of miR-30c (P≤0.0001, n=4), but not
of miR-30e, and anti-let-7f induced a potent downregulation in the expression of let-7f’s sister
miRNA, let-7a (P≤0.0001, n=4) (Figure 3.4 A, B).
Figure 3.4. Antagomirs are specific between miRNA families but redundant within
miRNA families (A) RT-qPCR for cardiomyocytes transfected with 50 nM of anti-miR30b or anti-cel-67 (negative control that targets Caenorhabditis elegans specific miRNA). Anti-miR-
30b decreases expression of its target miRNAs (miR-30b) significantly, and the expression of miRNAs from within the miR-30 family to varying degrees of potency. Data are represented as
miRNA expression ratio change of the antagomir treated group compared to the anti-cel-67 negative control group for each primer ± s.e.m, **P≤0.01, ***P≤0.001, n=4, and analyzed by Student’s t-test. (B) RT-qPCR for cardiomyocytes transfected with 50nM of anti-let-7f or anti-
cel-67. Anti-let-7f decreases expression of its target, let-7f, as well as let-7a (off target within the same family) significantly (ratio change values are ± s.e.m, **P≤0.01, ***P≤0.001, n=4,
analyzed by Student’s t-test). (C) RT-qPCR for cardiomyocytes transfected with either 50 nM of anti-let-7f (checkered bars), anti-miR-30b (solid grey bars) or anti-cel-67 Antagomirs significantly decrease expression of their target miRNA, but do not significantly decrease
expression of off target miRNAs of different miRNA families. Data was analyzed by one-way ANOVA (P≤0.0001) and Tukey’s Multiple Comparison post hoc test to identify differences
between groups and are represented as miRNA expression ratio change of the antagomir treated
group compared to the cel-67 negative control group for each primer± s.e.m, ***P≤0.001, ****P≤0.0001, n=4.
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To determine if antagomirs can effectively inhibit their target miRNA’s expression levels when
delivered as a cocktail, cardiomyocytes were next transfected with a combination of all 11 anti-
miRs. Inhibition of the 11 target miRNAs remained highly effective (Figure 3.5 A). Different
antagomirs exhibit varying degrees of inhibitory potency. Antagomirs targeting the miR-30
family were the most potent, and this could possibly be due to combined cross reactivity between
the different antagomirs within this family, as noted in Figure 3.4 A. These results confirmed
that transfecting cardiomyocytes with a cocktail of antagomirs was a feasible approach to knock
down multiple miRNAs at one time.
Knowing that the selected antagomirs effectively inhibit their target miRNAs, we proceeded to
determine whether knockdown of all 11 p53/MDM2-regulated miRNAs in cardiomyocytes could
promote cell cycle re-entry and proliferation. Cardiomyocyte proliferation was assessed by
immunofluorescence, staining for Aurora B (cytokinesis marker), co-stained with α-sarcomeric-
actin (cardiomyocyte-specific sarcomere marker), and DAPI (nuclear DNA marker) (Figure 3.6
A-H). Indeed, transfecting cardiomyocytes with a cocktail of antagomirs targeting all 11
p53/Mdm2-regulated miRNAs led to a striking increase in the occurrence of cytokinetic events
(Aurora B positive mid-body structures) by over 90%, compared to the negative control
transfected wells (~50,000 cells per well) (P≤0.01, n=3) (Figure 3.5 B, Supplemental Figure
3.5 A, B). This result underscores the role of these 11 miRNAs as cell cycle inhibitors in
cardiomyocytes.
Cardiomyocytes were also transfected with varying combinations of antagomirs to determine
which of the 11 were necessary to induce proliferation. To determine the minimal number of
antagomirs required to produce this proliferative phenotype, a “process of elimination” similar to
the concept pioneered by Takahashi & Yamanaka (2006) for determining the factors necessary in
pluripotent stem cell induction was utilized. Each antagomir was removed individually from the
cocktail and proliferation was assessed by immunofluorescence (Figure 3.5 B, Supplemental
Figure 3.5 A, B). If removing a specific antagomir reduced proliferation significantly compared
to the cocktail containing all 11 antagomirs (P≤ 0.05), it was deemed an essential member of the
antagomir cocktail. In contrast, if removing a specific antagomir had no effect on proliferation
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(i.e. a statistically significant number of cytokinetic events was maintained compared to negative
control transfected wells), this antagomir was noted as non-essential for the induction of
cardiomyocyte proliferation, and removed from the cocktail. Following this screen, it was
determined that antagomirs targeting miR-30c and let-7a were non-essential to promote
cardiomyocyte proliferation, whereas anti-miR-30b, anti-let-7f, anti-miR-181a, anti-miR-149
and anti-miR-194 were necessary components of the cocktail. To increase the throughput of the
screening process, we tested if the use of a microplate reader (picking up overall red 555
fluorescent signal in each well, corresponding to the Aurora B antibody) would parallel the
microscopy findings. As indicated by relative fluorescent units (RFU), the microplate reader also
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determined that cells transfected with all 11 antagomirs had increased fluorescence compared to
the negative control transfected wells, however the machine was not sensitive enough to detect
significant changes (Supplemental Figure 3.6 A, B). One-way ANOVA and Tukey’s Multiple
Comparison Post-Test were the statistical manipulations utilized to determine significance for
these experiments.
To confirm that anti-miR-30c and anti-let-7a were non-essential to promote cardiomyocyte
Figure 3.5. Inhibition of 11 target miRNAs by antagomir cocktail upregulates
cardiomyocyte cytokinesis. (A) Transfection with 11 antagomir cocktail efficiently knocks
down all target miRNAs. RT-qPCR performed for cardiomyocytes transfected with either a cocktail of all 11 antagomirs (25nM each) or anti cel-67 (negative control, 100 nM). Data are represented as represented as miRNA expression ratio change of the antagomir treated group
compared to the anti-cel-67 negative control group for each primer ± s.e.m, ** p ≤ 0.01, *** p ≤ 0.001, **** p ≤ 0.0001, n=3 and analyzed by Student’s t-test. (B) Cardiomyocytes were
transfected with either anti-cel-67, a cocktail of all 11 antagomirs (25nM each, red bar), a cocktail containing 10/11 antagomirs (label under bar indicates antagomir excluded from cocktail) or not transfected (black bar). Cardiomyocytes were subsequently stained with
antibodies against Aurora B, α-actinin and DAPI. The total number of cardiomyocyte cytokinetic events (indicated by Aurora B positive mid-body structures between two cells with
α-actinin positive sarcomeric structures) were counted in each well (~50,000 cells/well). Blue bars indicate the cocktails that retained a statistically significant number of cytokinetic events compared to the negative control, despite removal of one antagomir. These antagomirs are
hypothesized to be unnecessary for induction of cardiomyocyte proliferation. Light grey bars indicate cocktails where removal of the given antagomir significantly reduced cardiomyocyte
proliferation compared to the cocktail containing all 11 antagomirs. These antagomirs are hypothesized to be essential within the cocktail for induction of cardiomyocyte proliferat ion. Data represented as % change of Aurora B positive mid-body structures in treated cells
compared to negative control. Data analysis by one-way ANOVA (p=0.0035) and Tukey’s Multiple Comparison Test to identify differences between groups. ** indicates p ≤ 0.01, *** p
≤0.001 for antagomir treatment compared to negative control treatment. # indicates P≤0.05, ##P≤0.01, ### P≤0.001 for antagomir treatment compared to all 11 antagomir cocktail treatment, ± s.e.m, n=3. (C) Cardiomyocytes were treated according to the + and – symbols
under the x axis. Transfection with only anti-miR-30c+anti- let-7a significantly reduced cardiomyocytes proliferation compared to the cocktail containing all 11 antagomirs.
Cardiomyocytes transfected with all antagomirs except anti-miR-30c+anti- let-7a retained significant upregulation in proliferation compared to negative control. Data analysis by one-way ANOVA (P≤0.0001) and Tukey’s Multiple Comparison Test. *** indicates p ≤0.001 for
antagomir treatment compared to negative control treatment, ### indicates P≤0.001 for antagomir treatment compared to all 11 antagomir cocktail treatment, ± s.e.m, n=3. (D) List of
9 antagomirs in the finalized antagomir cocktail. anti-miR-30c and anti-let-7a were elimina ted.
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proliferation, a follow-up experiment was conducted where both antagomirs were removed from
the cocktail simultaneously. In this experiment, cardiomyocytes transfected with this cocktail of
only 9 antagomirs (containing all antagomirs except for anti-miR-30c and anti-let-7a) and
cardiomyocytes transfected with the cocktail containing all 11 antagomirs experienced over a
100% increase in cytokinetic events, compared to negative control treated cells. Accordingly,
transfection with a cocktail containing only anti-let-7a and anti-miR-30c significantly decreased
cardiomyocyte proliferation compared to cells transfected with all 11 antagomirs, and led to an
8% decrease in proliferation compared to the negative control (not significant). Similarly,
treatment of cardiomyocytes with either of those antagomirs alone caused a significant decrease
in proliferation compared to cardiomyocytes transfected with all 11 antagomirs. In fact,
transfection of cardiomyocytes with anti-miR-30c alone caused a significant ~23% decrease in
cytokinetic events compared to negative control treated cells, indicating that downregulating
miR-30c may be hindering cardiomyocyte proliferation in the cocktail containing all 11
antagomirs (Figure 3.5 C, Supplemental Figure 3.4 C, D). Thus, anti-miR-30c and anti-let-7a
were considered unnecessary members of the optimal antagomir cocktail required to induce
cardiomyocyte proliferation.
Next, we tested whether transfecting cardiomyocytes with only the five antagomirs deemed
“necessary” (anti-miR-30b, anti-let-7f, anti-miR-181a, anti-miR-149 and anti-miR-194) was
sufficient to promote cell cycle re-entry. Treatment of cardiomyocytes with this cocktail induced
a significant upregulation in proliferation of ~60%, however this was also significantly less
potent than either the cocktail containing 11 or 9 antagomirs. This indicated that antagomirs
other than those deemed necessary (dark grey bars, Figure 3.5 A: anti-miR-30a, anti-miR-30e,
anti-miR-204 or anti-miR-26b) were also adding to the ability of cardiomyocytes to proliferate.
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Figure 3.6. Treatment of cardiomyocytes with cocktail of 11 antagomirs promotes cardiomyocyte cytokinesis.
(A-H) Two 40x confocal immunofluorescence images of cardiomyocytes transfected with a cocktail of all 11
antagomirs (25nM each) undergoing cytokinesis. Cells were fixed and stained for (A,E) α-sarcomeric actin (red, cardiomyocyte specific sarcomere marker), (B,F) DAPI (blue), and (C,G) Aurora B (green, cytokinesis marker). (D,H) Merged image of all 3 channels. White arrows indicate Aurora B positive mid-body structures between two
cells undergoing cytokinesis
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We therefore sequentially removed each of these antagomirs from the cocktail of 9 antagomirs
(lacking the two unnecessary anti-miRs; anti-let-7a and anti-miR-30c) to determine if any were
revealed to be essential within this cocktail. Interestingly, removal of any of the 4 antagomirs
from the cocktail of 9 led to a significant decrease in cardiomyocyte proliferation compared to
cells treated with all 11 or 9 inhibitors. From this we concluded that treatment of cardiomyocytes
with 9/11 antagomirs (without anti-let-7a and anti-miR-30c) was the lowest number of
antagomirs required in the cocktail to promote proliferation (Figure 3.5 C, Supplemental
Figure 3.4 D). Thus, inhibition of p53/MDM2 regulated miR-30a, miR-30b, miR-30e, let-7f,
miR-204, miR-26b, miR-149, and miR-194 simultaneously was necessary for cardiomyocytes to
regain proliferative ability. A cocktail of 9 antagomirs targeting these specific miRNAs was
identified as the optimized final cocktail sufficient to promote cell cycle reactivation in
cardiomyocytes (Figure 3.5 D).
3.3.4 p53/MDM2-regulated miRNAs inhibit cardiomyocyte proliferation by downregulating target genes involved in the progression of all 4 phases of the cell cycle.
To elucidate how treatment of cardiomyocytes with either the cocktail of all 11 antagomirs, or
the final optimized cocktail of 9 antagomirs mechanistically promotes cell cycle reactivation, a
cell cycle pathway specific RT-qPCR array (Qiagen) which profiles the expression of 86 cell
cycle related genes was utilized. We were interested to determine if inhibition of these miRNAs
could promote re-expression of their targets genes within the cell cycle (listed in Figure 3.2 C
and 3.3 A, B). This would assist to confirm their role as cell cycle inhibitory and/or tumor
suppressive miRNAs.
Of the 32 genes within the cell cycle pathway targeted by the 11 p53/MDM2-regulated miRNAs,
20 were contained in the RT-qPCR cell cycle array. This array would therefore allow us to
confirm certain predicted miRNA target genes, but also help us to identify novel genes regulated
by these miRNAs that have not been previously characterized.
Four samples from each treatment group (11 antagomir cocktail, final 9 antagomir cocktail and
negative anti-cel-67 control) were profiled by the cell cycle array. This array revealed that
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inhibition of p53/MDM2-regulated miRNAs in both treatment cocktails causes a wide spread
upregulation in factors involved in cell cycle re-entry and progression. Specifically, inhibition of
all 11 miRNAs caused a significant (P≥0.05) upregulation of 32 genes involved in cell cycle
regulation (~36% of genes on the array). Genes necessary at each of the four phases of the cell
cycle became significantly upregulated, including cyclin D2 which promotes G1 phase
progression, minichromosome maintenance proteins 2/4 (MCM2/4) and origin of replication
complex subunit 2 (ORC2) which are necessary for DNA replication in S phase, cell division
cycle 25 (CDC25) which controls the rate limiting step of CDK1 activation in late G2 phase, and
importantly, CDK1, the absolutely essential kinase that induces M phase initiation and
progression37. Only one gene from this array, cyclin D1, was found to be significantly
downregulated by both treatments. This highlights the canonical role of miRNAs as inhibitors of
gene expression, such that their inhibition by antagomirs allows for a widespread upregulation of
their target genes. Notably, in cardiomyocytes transfected with the cocktail of all 11 antagomirs,
40% (8 out of 20) of the predicted miRNAs’ cell cycle target genes profiled by the array were
upregulated (Figure 3.7A).
Cells treated with the final antagomir cocktail also exhibited a significant upregulation in 25 out
of the 86 cell cycle genes profiled by this array, 7 of which were predicted target genes of the
miRNAs inhibited by the treatment (Figure 3.7 B). This treatment also induced the upregulation
of genes required for cell cycle progression at G1, S, G2 and M phase that are turned off in non-
treated cardiomyocytes, likely accounting for their quiescence. Importantly, treatment with both
cocktails produced a significant overlap in the change in gene expression elicited. 15 genes out
of the total number of genes significantly changed by both treatments were the same, indicating
that these two cocktails likely upregulate cell cycle re-entry and proliferation through a similar
mechanism. The extensive effects that inhibition of p53/MDM2-regulated miRNAs have on the
expression of a plethora of crucial cell cycle genes supports their role as strategic mediators of
cell cycle arrest in cardiomyocytes.
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Figure 3.7. Inhibition of 11 target miRNAs causes upregulation of many miRNA target genes within the cell
cycle pathway. Significantly (≤0.05) changed cell cycle related genes (ratio change of >1.2 or < 0.5) following treatment of cardiomyocytes with a cocktail of all 11 antagomirs or the final cocktail of 9 antagomirs (without unnecessary anti-miR-30c and anti-let-7a). The Qiagen Cell Cycle RT2 Profiler PCR Array was used to profile 86 cell
cycle relates genes following antagomir treatment compared to negative control treatment (anti-cel-67) by RT-qPCR. (A) Significantly changed cell cycle related genes following treatment of cardiomyocytes with all 11 antagomirs vs.
control treatment. (B) Significantly changed cell cycle related genes following treatment of cardiomyocytes with the final cocktail of 9 antagomirs vs. control treatment. White bars in (a) and (b) reflect genes that were significantly changed by both treatments. Values are represented as the ratio change of treatment vs. control ± s.e.m, n=4 and
analyzed by the Student’s t-test.
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3.4 Discussion
Our data show that a subset of 9 to 11 miRNAs regulated by p53/MDM2 function within
cardiomyocytes to maintain cell cycle arrest. The implications of these findings are two-fold.
Firstly, they reveal a novel role for p53 and MDM2 within the heart, of maintaining
cardiomyocyte cell cycle control through the regulation of miRNAs. Secondly they uncover
group of anti-proliferative miRNAs actively regulating cardiomyocyte quiescence.
3.4.1 p53 and MDM2 regulate the expression of muscle specific miRNAs in the heart
Following the conditional, cardiac-specific knockout of the tumor suppressor circuitry p53 and
MDM2, we were able to identify distinct populations of miRNAs that are regulated by each gene
individually, as well as in tandem. Loss of p53 led to the downregulation of 12 miRNAs whereas
19 and 29 miRNAs were downregulated in the Mdm2KO and DKO, respectively. Furthermore,
knockout of p53 produced an upregulation of only 3 miRNAs, whereas the Mdm2KO had 29
upregulated genes, and DKO had 36, many of which overlapped between the two strains,
indicating that the over-activation of p53 in the Mdm2KO is not the sole cause of the gene
expression changes observed. This highlights a potentially new role for MDM2, in regulating the
stability of other miRNA transcriptional activators yet to be identified. Among the
downregulated miRNAs in DKO hearts were the muscle specific miR-1 (also downregulated in
Mdm2KO) and miR-133 families. These miRNAs are expressed in adult skeletal and cardiac
muscle, but not in other tissues, and have been shown to play key roles in repressing the fetal
gene program in the adult heart120,262. Knockout of these two miRNAs in the adult heart was
shown to induce a re-expression of many fetal genes including Glycogenin 1, Glucan Branching
Enzyme 1, smooth muscle actin-α2, myosin heavy chain-β and myosin light chain-β120,262.
Interestingly, our lab had demonstrated that the double knockout of p53 and MDM2 specifically
in the heart also leads to the re-expression of the fetal gene program, and entails a metabolic shift
from fatty acid synthesis to glycolysis214 and our current study brings to light a possible
mechanism through which this may be occurring. Thus, the elucidation of p53 and MDM2
regulated miRNAs has helped to construct a specific role for these two genes within the heart of
maintaining cardiac differentiation and metabolism, apart from their canonical roles in tumor
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suppression. Future studies revealing the distinct interplay between p53, MDM2 and these
muscle specific miRNAs may reveal novel insights into the dedifferentiation and metabolic shift
exhibited by the heart during the progression of heart failure.
3.4.2 p53 and MDM2 maintain cardiomyocyte quiescence through the regulation of a unique group “anti-proliferative” miRNAs.
It has been widely accepted that a large number of miRNAs function as tumor suppressors within
the cell128. The cell cycle is a molecular cascade that entails numerous feedback loops, involving
an expansive number of regulators with overlapping functions. The robustness of this network
highlights the sheer evolutionary importance of this pathway, as the cell does not rely solely on a
small number of proteins for this process to occur, but instead utilizes the tight interplay of a
large number of proteins with degrees of functional redundancy, such that the loss of a single
factor does not result in a mitotic catastrophe. For example, knockout of CDK2, CDK4 or CDK6
alone does not produce a significant effect on cell proliferation263–265. CDK1 has been shown to
compensate for the loss of CDK2 and is capable to bind cyclin E in S phase266, and CDK2 was
shown to assume the functions of CDK4/6 upon their loss. It therefore follows that miRNAs play
an important role in maintaining quiescence of differentiated cells, as their capacity to fine tune
or dampen the expression of an expansive number of genes gives them the ability to
simultaneously downregulate the expression of a multitude of cell cycle regulators, overcoming
the internal redundancy of this pathway. Unpublished research from our lab suggests that by 7
days after birth, when cardiomyocytes are known to exit the cell cycle, there is a blanket
downregulation in almost all factors regulating the cell cycle pathway, including activators and
inhibitors, making the cells refractory to growth stimuli. This comprehensive shutdown of the
cell cycle pathway coincides with a switch in miRNA expression profiles between day 1 and day
10 post-natal mouse hearts, as demonstrated by Porrello et al (2011)102. It is possible that during
this switch in miRNA profiles, there is a bias towards the upregulating of anti-proliferative
miRNAs which contributes to the widespread downregulation of cell cycle regulators noted.
Based on previous research in our laboratory demonstrating a reactivation of the cardiomyocyte
cell cycle upon specific deletion of both p53 and MDM2 in the heart, and p53’s known ability to
regulate the expression of tumor suppressive miRNAs, we sought to determine if in the heart,
p53 and MDM2 may be promoting cell cycle arrest through the activation of “anti-proliferative
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miRNAs”. We found that the loss of both p53 and MDM2 led to the downregulation of a unique
subset of 15 miRNAs which were not significantly changed in the hearts of either single KOs.
We therefore hypothesized that these miRNAs may be crucial inhibitors of cell cycle in wild
type mouse hearts, and their downregulation following the loss of p53/MDM2 facilitates the cell
cycle re-entry we observe in these mutant mice (not observed in either of the single KOs).
Indeed, through bioinformatic target gene analysis, we found that 11 of these miRNAs (miR-30a,
miR-30b, miR-30c, let-7a, let-7f, miR-181a, miR-26b, miR-204, miR-149 and miR-194) possess
target genes within the cell cycle, and that a large majority of these genes (81%), are involved in
cell cycle activation. The implication of this finding is that downregulation of these miRNAs,
such as what is seen in DKO hearts, could remove the inhibition over a large group of key cell
cycle factors necessary for reactivation of this pathway and proliferation of cardiomyocytes. We
therefore inhibited these 11 miRNAs all together in wild type cardiomyocytes using a cocktail of
11 antagomirs, which elicited over a 100% increase in cytokinesis (determined by the presence
of Aurora B positive mid-body structures as opposed to indirect measurements of proliferation
such as Ki-67 or PiH3). This indicates that these miRNAs have a broad effect over cell cycle
control, and that their inhibition not only induces re-entry, but the completion of this pathway to
its entirety, resulting in the formation of new cardiomyocytes. This is important because it is
known that cardiomyocytes have cell cycle roadblocks at multiple phases in the pathway, and
although the upregulation of a single factor can promote cell cycle re-entry, it is extremely rare
that they are shown to complete the process and physically divide into two separate cells81. This
data also fits in well with the research presented by Porrello et al., as they demonstrated that the
miR-30 and let-7 families were prominently upregulated in P10 hearts coinciding with cell cycle
arrest, and antagomirs targeting 5 miRNAs from these families make up a large portion of our
antagomir cocktail102.
The approach we have taken to promote cardiomyocyte proliferation demonstrates that targeting
miRNAs, who have an overarching, all-inclusive control over biological pathways with the
capability to inhibit the expression of hundreds of genes, may be an efficient method to promote
a desired phenotype. Furthermore, the redundancy in target genes between different miRNAs
(e.g. the stromal antigen 1 (STAG1) is a target of 6/11 miRNAs) supports the cocktail method
that we utilized. Targeting simply one miRNA may not be sufficient to upregulate all the cell
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cycle factors required to promote cardiomyocyte proliferation, because other anti-proliferative
miRNAs remain, who have overlapping target genes, and can maintain their inhibition despite
the loss of one miRNA. miRNAs likely do not function in isolation124. Rather, the expression of
multiple miRNAs that target genes with a certain degree of overlap promotes a synergistic
function, enhancing their potency within the cell to shut off biological pathways, while
enhancing others124. With this notion in mind, we were interested in defining the potential for a
smaller cocktail of antagomirs. Through sequential removal of each antagomir from the cocktail
individually, we concluded that the presence of 9 antagomirs was sufficient to upregulate
cardiomyocyte proliferation to a level comparable with the cocktail of all 11 antagomirs. It is
interesting that the two non-essential antagomirs (anti-miR-30c and anti-let-7a) inhibit miRNAs
who have other family members also being targeted in the cocktail, whereas inhibition of each
unique miRNA was necessary. Two plausible explanations can be deduced: (1) within the
cocktail there are antagomirs targeting three other miR-30 family members and one other let-7
family member, and inhibition of these other family members causes a sufficiently potent
upregulation of their target genes, such that miR-30c and let-7a need not be inhibited (miR-30c
target genes 100% overlap with those of miR-30a, let-7a target genes 100% overlap with those of
let-7f), (2) antagomirs targeting miR-30a, miR-30b or miR-30e non-specifically inhibit miR-30c,
and the antagomir targeting let-7f non-specifically inhibits let-7a such that anti-miR-30c and
anti-let-7a are not necessary.
3.4.3 p53/MDM2-regulated miRNAs maintain cardiomyocyte cell cycle arrest through the inhibition of multiple cell cycle regulators.
To determine a mechanism by which our antagomir cocktails upregulated cardiomyocyte
proliferation, we investigated the expression of 86 key cell cycle regulators following treatments,
which included 20 out of the 32 of the predicted cell cycle target genes of the 11 miRNAs. Both
treatments (all 11 antagomirs or optimized final cocktail with 9 antagomirs) significantly
upregulated the expression of a multitude of cell cycle regulators required at each phase of the
cycle, including a significant percentage of miRNA cell cycle target genes predicted by the target
gene prediction software that we utilized. Treatment of cardiomyocytes with all 11 antagomirs
upregulated the expression of 32 cell cycle regulators, and the optimized 9 antagomir cocktail
promoted re-expression of 24 genes, whereas only cyclin D1 was significantly downregulated by
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both treatments. This demonstrates that the knockdown of these miRNAs alleviates their
inhibition over a large number of genes essential for cell cycle entry (early G1 phase),
progression (G1/S transition, S phase, G2/M transition, M phase) and completion (cytokinesis).
The majority of the genes upregulated were cell cycle activators, but key cell cycle inhibitors
such as retinoblastoma (RB), retinoblastoma-like protein 1 (RBL1/p107) and retinoblastoma-like
protein 2 (RBL2/p130) were also induced. This finding indicates that the treatments promote
cardiomyocytes to begin cycling through the cell cycle, whereby a large majority of the cells are
likely in G1 phase (the longest phase of the cell cycle) in which the expression of these cell cycle
inhibitors are important for the timely coordination of E2F mediated gene expression267. In line
with this, the inhibitory E2F5 gene is also upregulated by both treatments, and therefore can be
bound by p130 allowing it to traffic to the nucleus and inhibit important S phase genes while the
cell is preparing itself for cell cycle commitment in early G1 phase267. Factors that promote G1/S
phase transition such as CDK6 and the S-phase kinase-associated protein 2 E3 ubiquitin protein
ligase (SKP2) were also upregulated. CDK6 phosphorylates RB, which inhibits its function and
allows the cell to progress into S phase. The E3 ubiquitin ligase SKP2 plays a similar role, and
degrades p27Kip1, a protein which promotes G1 phase arrest by inhibiting the catalytic activity of
Cyclin-CDK complexes268. Crucial S phase executors such as MCM2 and MCM4 are induced.
These proteins are components of the DNA helicase and form the foundation for the construction
of active replisomes, allowing DNA replication to proceed44. The essential G2/M transition
mediator, cell division cycle 25B (CDC25B) is also upregulated. CDC25B removes the
inhibitory phosphate groups from CDK1, which is the rate limiting step for the initiation of
mitosis269. Importantly, this step would be redundant if CDK1 was not also expressed (it is
downregulated in adult hearts), as M phase cannot progress without this key kinase57. Indeed,
both treatments significantly upregulated the expression of CDK1 in cardiomyocytes, indicating
that not only can the cells enter into the cell cycle, but can also proceed through mitosis. Other
regulators of M phase such as the MAD2 mitotic arrest deficient-like 1 protein (MAD2L1), were
increased as well. MAD2L1 is a component of the mitotic checkpoint complex, which monitors
chromosome alignment and is necessary to halt transition into anaphase if chromosomes become
misaligned37. Moreover, as we observed an increased number of cells undergoing cytokinesis,
we presumed key regulators of this process would be upregulated. Indeed stathmin (STMN1), an
important mediator of microtubule dynamics, whose activity is necessitated during cytokinesis to
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modulate the drastic cytoskeletal changes that occur during this process, was significantly
upregulated following both antagomir treatments270. In all, cell cycle progression requires an
intricate orchestration of cell cycle activators, inhibitors and checkpoint regulators, many of
which we have shown to be inhibited in cardiomyocytes by a group of “anti-proliferative”
miRNAs regulated by p53/MDM2. The inhibition of these identified miRNAs permits the
upregulation of a widespread group of cell cycle regulators that as a whole, allow the re-entry
and controlled cycling of cardiomyocytes through each phase of the cell cycle.
The ability of miRNAs to inhibit many genes offers a potential therapeutic advantage over other
agents that act only on a single gene, especially when striving to regulate a process that requires
the reactivation of multiple genes. Furthermore, targeting a group of miRNAs who function in a
similar pathway may be important in effectively modulating complex physiological processes.
We have identified a group of miRNAs who function in the heart to promote cardiomyocyte
quiescence, under the regulation of the tumor suppressor circuitry p53 and MDM2. These data
provide a new level of understanding to how cell cycle arrest is regulated in the heart, and how
p53 and MDM2 participate in the mediation of this process. In addition, this approach has the
possibility to be exploited for the development of novel regenerative therapies to replenish lost
heart cells following injurious events like myocardial infarction.
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3.5 Supplementary Figures and Tables
Supplemental Figure 3.1. Validation of cardiac specific MDM2 and p53 knock-down
following Tamoxifen injection. To specifically knock-down p53 and Mdm2 in the adult mouse heart, a tamoxifen (Tam) inducible Cre-loxP system where MerCreMer recombinase
(mcm) expression is regulated by the cardiomyocyte specific myosin heavy chain 6 promoter was used to generate the following knockout (KO) strains: p53KO p53f/f;mcm; Mdm2KO,
Mdm2f/f;mcm; DKO, Mdm2f/f;p53f/f;mcm. Tam was injected intraperitoneally once daily for 4 consecutive days into 8 week old mice. Day 0 = day of last Tam injection. Total protein isolated from the left ventricle of the 3 KO strains was probed with primary antibodies against
p53 and Mdm2 as indicated (left). An antibody against nucleophosmin (Npm1) was reprobed for on all membranes to verify equal protein loading into all lanes.
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Supplemental Figure 3.2. Biological pathway analysis for the 11 p53/MDM2-regulated miRNAs. miRNA
target genes were grouped into GSEA and KEGG biological pathways. Represented are the top 100 enriched pathways that the 11 miRNAs regulate, ranked based on their target genes and expression levels (fold change) (miRSystem Pathway ranking Score – see Supplemental Figure 6 for score equation). The KEGG “Cell Cycle”
pathway is in the top 30 biological pathways that the p53/MDM2-regulated miRNAs regulate
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Supplemental Figure 3.3. 11 miRNAs hits are downregulated in
cardiac fibroblasts. RT-qPCR of cardiomyocytes and fibroblas ts probing for the 11 miRNA hits. Data are represented as the miRNA
expression ratio change in cardiac fibroblasts compared to wild type cardiomyocytes for each primer ± s.e.m, n=3. A significant difference
between the mean normalized delta Ct values for fibroblasts vs. cardiomyocytes is indicated by * p ≤ 0.05, ** p ≤ 0.01, *** p ≤ 0.001, **** p ≤ 0.0001, n=3. Data analyzed by Student’s t-test.
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Supplemental Figure 3.4. Efficient transfection of neonatal rat cardiomyocytes with red fluorescent positive
control. (A-D) 10x immunofluorescence images of cardiomyocytes transfected with a positive control (Alexa Fluor®555 Red Fluorescent Control siRNA, 100 nM). Transfection efficiency = 93% ± 6%. (A,E) Nuclear DAPI
stain, (B,D) cardiomyocyte specific nuclear stain, MEF2A, (C.G) Alexa Fluor®555 Red Fluorescent Control siRNA within the cytoplasm of the cardiomyocytes, and (D,H) a merged image of all 3 channels. (E-H) Zoomed
in view of one section of micrographs A-D. The fluorescent positive control can be seen within the cytoplasm of
each cell.
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Supplemental Figure 3.5. Raw count data for cytokinesis events following antagomir
treatment. (A) Total number of cytokinetic events for graphs shown in Figure 4 B, (B) Total number of cytokinetic events for graphs shown in Figure 4B, (C) Total number of cytokinet ic events for graphs shown in Figure 4C, and (d) Total number of cytokinetic events for graphs shown
in Figure 4C. Each graph indicates a separate experiment. Data analysis by one-way ANOVA (A)
p=0.0035, (B) p<0.0001, (C) p<0.0001 (D) p<0.0001 and Tukey’s Multiple Comparison Posttest to
identify differences between groups. ** indicates p ≤ 0.01, *** p ≤0.001 for antagomir treatment compared to negative control treatment. # indicates P≤0.05, ## P≤0.01, ### P≤0.001 for antagomir
treatment compared to all 11 antagomir cocktail treatment, ± s.e.m, n=3
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Supplemental Figure 3.6. Average Cytokinetic events and average RFU show similar % change trends in
transfected cardiomyocytes. For each transfected cocktail, the % change of mean cytokinetic events and RFU was
calculated compared to the negative control. (A) The same cardiomyocytes transfected and stained in Figure 4 b were passed through a fluorescent microplate reader. The Relative Fluorescent Units (RFU) were measured to determine the fluorescence intensity of Aurora B in each well. Aurora B fluorescence was highest in the
cardiomyocytes transfected with all 11 antagomirs. One-way ANOVA analysis (p=0.1) revealed that this method was not sufficiently sensitive to pick up significant changes in RFU between treatments. (B) The % change
calculated by each method shows similar trends between cocktails despite lack of significance in microplate reader
data.
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Supplemental Figure 3.7. RT-qPCR for cardiomyocyte housekeeping genes. (A)
Cardiomyocytes were transfected with a cocktail of all 11 antagomirs (25nM, n=2), anti-cel-67 negative control (100nM, n=2) or not transfected (n=2), and probed for RNU6 (small nuclear RNA component of the spliceosome) by RT-qPCR. RNU6 expression was not strongly affected
by antagomir treatments and was thus chosen as the housekeeping gene to be used for normalization for all subsequent miRNA RT-qPCR experiments. (B) Cardiomyocytes were
transfected with a cocktail of all 11 antagomirs (25nM, n=4), the final cocktail of 9 antagomirs (25nM, n=4) or anti-cel-67 negative control (100nM, n=4) or not transfected (n=3). Out of 5 housekeeping genes contained in the Qiagen RT2 profiler array (B2M, ACTB, LDHA, HPRT1
and RPLP1), the expression of LDHA and HPRT1 remained the most consistent between all three treatments and were thus chosen as the housekeeping genes to be used for normalization of
all subsequent genes within the array.
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Supplemental Figure 3.8. MirSystem miRNA target gene biological
pathway ranking score. For a given set of miRNA, expression levels are
used as the weight (ѿi). The weight of one miRNA is calculated by dividing its absolute expression value by the absolute sum of the expression values of all input miRNAs. Next, after identifying the target genes by the prediction
algorithms, for each functional category (biological pathway), the ranking score is obtained by summation of miRNAs times its enrichment –log (P-
value) from the predicted target genes
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Supplemental Table 3.1. Normalized RT-qPCR Ct values for miRNA expression levels
following 11 antagomir transfection. RT-qPCR for cardiomyocytes transfected with either
a cocktail of all 11 antagomirs (25nM each) or anti cel-67 as a negative control (100 nM). Data are represented as mean Ct values normalized to small RNA housekeeping gene RNU6.
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Chapter 4
Conclusion
4 Conclusion and Future Directions
4.1 General Discussion
4.1.1 Thesis summary
The heart is an organ with limited regenerative capacity. Shortly after birth, proliferation in the
mammalian heart almost completely comes to a halt271. The implication of this is that following
injury, any lost cardiac muscle cells will not be replaced. Rather, the heart will attempt to heal by
cardiac hypertrophy and scar formation. The precise evolutionary mechanisms that set us apart
from amphibians such as newts, that can regenerate their cardiac tissue, remains to be defined271.
Various principal investigators have postulated that it is either due to an evolutionary trade off
preventing the development of cancer272, which is an extremely rare occurrence within the heart.
Others suggest that an evolutionary prioritization of hemostasis, hypertrophy and fibrosis over
regeneration is the cause of the adult heart’s inherent inability to activate this process271.
Remarkably, we have been able to demonstrate that the cardiac-specific knockout of p53 and
MDM2 can cause cardiomyocytes to re-enter into the cell cycle, in stark contrast to wild type
hearts214. However, loss of both p53 and MDM2 in the heart also caused early onset lethality,
due to many perturbations in cardiac metabolism and structural integrity214. We therefore sought
to determine the mechanism responsible for this proliferative response. We reasoned that if we
could modulate a single part of the downstream system, as opposed to the entire circuitry
regulated by p53/MDM2, this could provide a more directed approach to promote cardiomyocyte
proliferation while avoiding the other negative outcomes. p53 is known to activate the expression
of miRNAs that act as tumor suppressors in other organs217. Intriguingly, a canonical p53-
regulated miRNA family, miR-15, was shown to be capable of regulating the post-natal mitotic
arrest of cardiomyocytes, among other important miRNAs102. Thus, the specific aim of this thesis
was to identify the miRNAs regulated by p53 and MDM2 that maintain cardiomyocyte cell cycle
arrest in the adult mammalian heart.
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Through comparison of miRNA expression profiles, we found that a distinct set of deregulated
miRNAs could be identified in each of the three cardiac-specific mutant strains of p53 and
MDM2. As expected, the cardiac miRNA profile of DKO mice had some overlap with the two
single knockouts, but also a subset of uniquely deregulated miRNAs were present. More
specifically, 11 miRNAs from this group become downregulated in the DKO hearts, and are
enriched for target genes within the cell cycle pathway. As miRNAs exert their function post-
transcriptionally through inhibition of their target genes’ mRNA transcripts, we hypothesized
that these miRNAs may function as “anti-proliferative” miRNAs, by inhibiting multiple target
genes within the cell cycle pathway. This would help to explain the cell cycle re-entry exhibited
by DKO mice, where these miRNAs are downregulated, and therefore no longer exerting their
anti-proliferative effects within the cell.
To further characterize these miRNAs as regulators of cell cycle arrest, we determined their
expression levels in a proliferative cell type in the heart, that is, cardiac fibroblasts. Cardiac
fibroblasts have a strong proliferative capacity in vitro, and we expected that this group of
miRNAs might be downregulated in these cells as compared to the non-proliferative
cardiomyocytes. Indeed, each miRNA was significantly downregulated in fibroblasts, supporting
the idea that they form a roadblock for cell cycle re-entry in cardiomyocytes. We therefore
hypothesized that inhibiting these 11 miRNAs in isolated cardiomyocytes might promote a
phenotype similar to what was observed in the DKO, and allow for cell cycle re-entry. To test
this hypothesis, we knocked down the expression of the 11 miRNAs using a cocktail of
antagomirs in cultured neonatal rat cardiomyocytes. We found that antagomir treatment did lead
to significantly increased numbers of cardiomyocytes undergoing cytokinesis, through the
upregulation of a widespread number of cell cycle regulators. Therefore, we were able to clearly
demonstrate that the identified 11 p53/MDM2-regulated miRNAs negatively regulated the
expression of crucial cell cycle mediators required by cardiomyocytes to undergo cell division.
We also demonstrated that two antagomirs, anti-let-7a and anti-miR-30c, were not necessary
within the initial cocktail to promote cardiomyocyte proliferation, although the reason behind
this remains to be elucidated, as discussed in the Discussion section 3.4.2 of Chapter 3.
These p53/MDM2-regulated miRNAs seem to act in concert to maintain cardiomyocyte cell
cycle arrest, as at least 9 out of the 11 must be inhibited to promote proliferation. Notably, each
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of these miRNAs have been shown to function as tumor suppressors in cell types more amenable
to ectopic cell cycle re-entry, whereby their upregulation can halt the progression of diverse
types of cancers.
4.1.2 The 11 p53/MDM2-regulated miRNAs are well characterized tumor suppressors
4.1.2.1 The miR-30 family
miR-30a, miR-30b, miR-30c and miR-30e all act as tumor suppressors in breast cancer,
colorectal cancer, chondrosarcoma, and multiple myeloma, through their inhibition over
oncogenic factors such as metadherin, the insulin receptor substrate 2 (Irs2), SRY-box 4 (SOX4),
and the WNT/β-catenin pathway. In each of these distinct cancer types, the expression of one or
more miR-30 family members was decreased, and the ectopic administration of mimics for these
miRNAs halted cancer progression through induction of cell cycle arrest and inhibition of
metastasis273–279.
4.1.2.2 The let-7 family
The let-7 family is also widely characterized as a group of individual tumor suppressor miRNAs.
Primarily, members of this family including let-7a and let-7f, were shown to be downregulated in
lung cancer which was correlated with an upregulation of the RAS oncogene280. Subsequently, it
was demonstrated that these miRNAs were also downregulated in neuroblastomas, breast cancer,
renal cell carcinoma and many others, and that they inhibit numerous oncogenes, the most
notable being c-MYC281–286. Interestingly, Ohshima et al (2010) showed exosomic secretion of
the let-7 family in a metastatic gastric cancer cell line, presumably for protection of the
cancerous cells against their tumor suppressive effects287. This highlights that the let-7 family
likely exerts a potent anti-tumor effect, that cancer cells must evolve mechanisms to circumvent.
Re-expression of the let-7 family in cancers has thus been a highly researched topic in cancer
therapeutics, and synthetic let-7 administration in multiple cancer cell lines has been shown to
promote cancer cell quiescence and an accumulation of cells in the G0 phase288. Furthermore, in
agreement with the research presented in this master’s thesis, upregulation of the let-7 family in
liver and in lung cancer cell lines promoted either directly or indirectly a downregulation of
many cell cycle regulating factors including cyclin A2, E2F5, CDC25 and SKP2288. Following
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treatment of rat neonatal cardiomyocytes with our cocktail of antagomirs, including two that
inhibit members of the let-7 family, we also saw a significant re-expression of all the genes listed
above.
4.1.2.3 miR-181a
miR-181a also has notable tumor suppressor properties. This miRNA is downregulated in
gliomas, lung cancer, and chronic myelogenous leukemia289–291. In lung cancer cells (A549
cells), and leukemic cells (K562 cells), re-expression of miR-181a was shown to directly
downregulate two oncogenes, KRAS proto-oncogene GTPase (KRAS)290 and RALA Ras like
proto-oncogene A (RalA)291 respectively, thereby inhibiting proliferation of these cells. miR-
181a overexpression of RalA in leukemic cells caused G2 phase arrest and induction of
apoptosis291. Thus, inhibiting this miRNA in cardiomyocytes may have therapeutic value, not
only in promoting cell cycle reactivation, but also by mitigating apoptosis, which is key in the
pathogenesis of many cardiac diseases.
4.1.2.4 miR-26b
miR-26b similarly exerts a strong tumor suppressive effect over multiple types of cancer. This
miRNA has been found significantly downregulated in colon cancer, breast cancer, melanoma,
and bladder cancer, and has been documented as a potent inhibitor of cell proliferation in many
of these diseases292–295. Other studies have shown that miR-26b potently inhibits the cell cycle at
the G1/S phase transition by repression of cyclin D2, cyclin E1 and CDK6 mRNA transcripts in
fibroblasts and spermatocyte derived cells296,297. In support of these findings, the data presented
in this thesis show a significant upregulation of both cyclin D2 and CDK6 following antagomir
cocktail treatment, which includes an inhibitor against miR-26b.
4.1.2.5 miR-204
Reduced expression of miR-204 in gastric cancer, breast cancer and acute myeloid leukemia
predicts poor patient prognosis, based on three different studies which indicate that this miRNA
clearly plays an important role in preventing cancer progression298–300. It has also been identified
to be strongly downregulated in bladder and colorectal cancers, where its re-expression promotes
the inhibition of the anti-apoptotic BCL2 gene301,302, and a member of the RAS oncogene family
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RAB22A301, sensitizing these cancer cells to chemotherapy by upregulating apoptosis and
inducing cell cycle arrest.
4.1.2.6 miR-149
Overwhelming amounts of research support miR-149 as a tumor suppressor that inhibits
proliferation and mediates cell cycle arrest when re-expressed in multiple cancers. It therefore
follows that it is downregulated in numerous cancers including gliomas, gastric cancers, breast
cancers, colorectal cancers, ovarian cancers among many others301,303–307. Furthermore, miR-149
was shown to be downregulated in cancer-associated fibroblasts, presumably to promote the
creation of a tumor microenvironment that is more amenable for tumor growth and metastasis. In
line with this view, miR-149 re-expression in cancer-associated fibroblasts inhibited the pro-
inflammatory cytokine, IL-6, that functions to promote the cross talk between tumor cells and
fibroblasts308. Thus, downregulation of this miRNA in cardiomyocytes following cardiac injury
may promote not only the beneficial effect of upregulating proliferation and repair, but also by
promoting an anti-inflammatory environment.
4.1.2.7 miR-194
Finally, miR-194 has also been well characterized as a tumor suppressor in various cancers
including oral squamous cell carcinoma, colorectal cancer and hepatocellular carcinoma309–311.
On top of this, low levels of miR-194 are now known as an independent risk factor for the
reoccurrence of colorectal adenomas, with a high sensitivity and specificity of >70% as a
predictor312. It is interesting to note that this miRNA has also been found to work in a feedback
loop with p53. Wang et al. (2015) demonstrated that this miRNA can directly target the mitogen-
activated protein kinase kinase kinase kinase 4 (MAP4K4)309. Intriguingly, MDM2 is a
downstream target of MAP4K4, and inhibition of this kinase by miR-194 downregulated the
expression of MDM2, in turn, leading to p53 stabilization. This finding serves to explain the
broad downregulation of miR-194 in many cancers.
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4.1.3 Tumor suppressive p53/MDM2-regulated miRNAs may function synergistically to promote cardiac cell cycle arrest
With the discussion above, there is a large amount of evidence from the cancer field that the 11
p53/MDM2-regulated miRNAs identified in this thesis are crucial inhibitory molecules
regulating the pathways of growth and proliferation. This is supported by the miRNA target gene
analysis we conducted that listed “pathways in cancer” to be the second most enriched biological
pathway regulated by this group of miRNAs, preceded only by the MAPK pathway. If the
postulated evolutionary reasoning behind the lack of regenerative potential in the heart is due to
a trade off in the prevention of cancer, it follows well that these potent tumor suppressor
miRNAs would be upregulated in the heart. The downregulation of just one of these miRNAs
has the ability to heighten an individual’s risk for the development of cancer in tissues that are
more amenable to cell cycle re-entry. It is therefore plausible that the downregulation of all these
inhibitors together is required in cardiomyocytes to remove the rigorous block in the cell cycle
that they are subject to. Furthermore, the expression levels of these miRNAs within a cell may
play a role in defining their regenerative capacity, e.g. upregulated in cells which are strongly
refractory to proliferative stimuli such as cardiomyocytes, lower levels of expression in cells
which retain the natural ability to proliferate such as fibroblasts and lowest levels of expression
in cells which undergo aberrant growth/proliferation such as tumor cells. These findings also
highlight that the potency of p53 as a tumor suppressor may be strengthened through the
activation of multiple downstream tumor suppressor miRNAs. The upregulated expression of
these miRNAs in cardiomyocytes likely allows them to work synergistically to enhance p53’s
function as a tumor suppressor, all together creating a potent inhibitory network surveying
aberrant cell cycle re-entry through multiple different avenues.
The synergistic function of miRNAs has been previously described by Zu et al (2013), who show
that the overlapping target genes of miRNAs potentiates their ability to repress expression of
these genes313. Studies regarding the synergistic function of miRNAs have since focused on the
delivery of two miRNA mimics to produce a more potent phenotype over the delivery of either
one individually. For example, to promote tumor suppression, combinations of miRNA mimics
of miR-34+let-7314, miR-143+miR-145315 and miR-10b+miR-21316 synergistically inhibited the
proliferation of tumor cells. In contrast, the combination of miR-133+miR-499 potentiates
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cardiomyocyte differentiation317, and the synergistic combination of miR-21+miR-146a
attenuates cardiac apoptosis following ischemia reperfusion124. However, in the present study,
we exploit this mechanism in reverse, whereby the inhibition of a large group of miRNAs
removes their synergistic repression over numerous cell cycle regulators, thereby promoting
cardiomyocytes proliferation, whereas inhibition of these miRNAs in isolation is not sufficiently
potent to promote this effect.
4.1.4 Mechanisms of cardiac regeneration with a therapeutic goal in mind
The findings described in this thesis shed light on a group of tumor suppressive miRNAs
previously unknown to function within the heart, that have the potential to be utilized in the
development of novel regenerative cardiovascular therapies. Following cardiac injury, such as
myocardial infarction, millions of cardiomyocytes can be lost during the ischemic process as
well as during reperfusion318. Such dramatic losses significantly disrupt the structural and
functional integrity of the heart. Thus, future cardiovascular research should be aimed at the
replacement of lost cardiomyocytes as opposed to preventing further damage from occurring by
mitigating the work-load of the heart. This would prevent the negative repercussions that ensue
when the heart attempts to compensate for these losses by fibrosis and hypertrophy. Ultimately,
the goal would be to prevent or reverse the development of heart failure by intrinsic regeneration.
The focus of regenerative cardiovascular therapies has historically revolved around two main
themes: promotion of endogenous repair through reactivation of the cardiomyocyte cell cycle or,
promotion of exogenous repair through delivery of cell therapies.
4.1.4.1 Cell-based cardiac regenerative therapies
The transplantation of multiple different cell types into the heart has been proposed as a means to
promote myocardial repair and regeneration. Although research has yielded promising results, it
has also revealed many caveats that come along with introducing exogenous cells into the heart.
Much research is still focused on determination of the optimal cell type to be used in
transplantation, promotion of proper cell integration, and understanding the mechanisms by
which benefit is being provided.
Some of the earliest work in this field focused on transplanting skeletal myoblasts (progenitor
cells found in skeletal muscle) into the heart, with the hope that they would differentiate into
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cardiomyocytes when provided the change in microenvironment319. Rather, these cells
differentiated into myotubes (early developmental stage of muscle fiber formed by the fusion of
multiple myoblasts to create a syncytium), and were unsuccessful at integrating electrically with
the host myocardium319. Nevertheless, multiple clinical trials were undertaken due to the
observations of functional improvements in pre-clinical trials. However, results suggested that
these cells had limited integration efficiency into human hearts and thus, put patients at risk for
arrhythmias319. These studies highlighted that due to the tightly organized electrical conductive
system in the heart, not present in other organs, a major issue for cardiac cell-based therapies
would be to optimize graft integration to avoid the onset of arrhythmogenesis,
Following this, most work has been focused on transplanting cardiac derived cell types, obtained
from either embryonic stem cells, induced pluripotent stem cells, or arising from resident or non-
resident cardiac progenitor cells320.
Due to their unlimited capacity for self-renewal, and their ability to be efficiently differentiated
into cardiomyocytes, embryonic stem cells were an appealing candidate for cardiac stem cell
based therapies320. Undifferentiated embryonic stem cells cannot be injected into tissues, due to
the development of teratomas (tumors derived from cells not normally present in host organ).
However, injection of embryonic stem cell derived cardiomyocytes into immunocompatible
hearts were able to form intracardiac grafts that improved heart function in mice, rats and other
small animal models320. These cells were also able to achieve electrical integration, and co-
delivery with paracrine factors or bio-engineered scaffolds improved engraftment and survival of
these cells321,322. Numerous drawbacks were also identified for the use of embryonic stem cell-
derived cardiomyocytes, including the ethical issues of cell procurement, determining
immunocompatibility and the likely requirement for immunosuppression, and the immaturity of
the cardiomyocytes derived from embryonic stem cells.323 This last drawback likely prevents
optimal cardiac integration and function. Furthermore, the safety of this therapeutic option is in
question, due to the unknown potential for the development of teratomas, the longevity of the
grafts and the possibility of de-integration leading to arryhthmogenesis323.
Induced pluripotent stem cells have more recently come to the forefront as a more advantageous
source of cells from which to derive cardiomyocytes for transplanting purposes323. The protocol
for cardiomyocyte derivation from induced pluripotent stem cells has been largely fine-tuned
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over the last few years, such that the cardiomyocytes derived from these cells more closely
resemble adult cardiomyocytes in terms of their calcium handling and contractile ability323.
Furthermore, deriving cardiomyocytes by this method side steps the ethical issues surrounding
embryonic stem cells and the requirement for immunosuppression (as induced pluripotent stem
cells can be generated autologously). Despite these improvements, logistical issues arise for
developing therapies based on the transplantation of cardiomyocytes derived from induced
pluripotent stem cells. Firstly, generating cardiomyocytes from autologous induced pluripotent
stem cells would require numerous months of preparation. For this reason, their use to treat the
immediate damage that occurs in acute illnesses such as myocardial infarction would not be
possible. Secondly, a uniform manufacturing protocol to generate induced pluripotent stem cell
derived cardiomyocytes is a large regulatory hurdle for their use in the clinic.
The search for a population of resident cardiac progenitor cells has been contentious. Although
these cells have been shown to exist, their function in the heart is poorly understood, as they are
clearly not efficiently activated following cardiac injury to regenerate the heart, unlike skeletal
muscle cell progenitors. Many different populations of cardiac progenitor cells have been
reported, expressing diverse stem cell markers such as KIT Proto-Oncogene Receptor Tyrosine
Kinase (C-kit) + cardiac progenitor cells324, ataxin-1 (SCA1) + cardiac progenitor cells325, ISL1+
cardiac progenitor cells326, and cardiosphere-derived cells327. The cell type of main focus has
been C-kit+ cells found within the heart. C-kit is a receptor tyrosine kinase that is highly
expressed on hematopoietic stem cells, mature circulating cells, mast cells, some endothelial
cells, and immature cardiomyocytes323. Therefore, this marker has the issue of lack of specificity
to cardiac lineage cells. However, differentiation of these cells into the three major cell types of
the heart, cardiomyocytes, smooth muscle cells and endothelial cells, has been reported328. As
such, when injected into the heart, they were found to form not only new cardiomyocytes, but
also new blood vessels, enhancing vascularization of the damaged areas328. However, the
cardiomyogenic activity of these C-kit+ cells, which arise from the bone marrow, has been
contested by many groups323. Despite this, a prominent clinical trial was carried out, whereby
autologous C-kit+ cells expanded from myocardial tissue harvested from patients who underwent
a coronary artery bypass graft, were transplanted back into these patients to determine if this
could improve the outcomes of ischemic heart failure237. Minor improvements of left ventricular
ejection fraction were observed (7% increase), but it was hypothesized that this improvement
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was due to the secretion of paracrine factors from the transplanted cells, as opposed to a direct
cardiomyogenic contribution328.
Non-resident cardiac progenitor cells have also been studied for their potential to promote
myocardial repair323. C-kit+, hematopoietic lineage negative, bone marrow cells were isolated
and transplanted into the border zone of myocardial infarcts in mice, and were shown to generate
new myocardium in the infarct zone329. Unfortunately, the differentiation of c-kit+ bone marrow
cells into cardiomyocytes could not be reproduced in other studies330,331. It has been
hypothesized that the transplanted cells may fuse to resident cardiomyocytes, and this could
account for the “differentiation” that was observed323. Due to the ease of access of these cells,
despite the discrepant findings in small animal models, clinical trials were carried out that
injected different types of bone marrow derived cells into the heart, including mesenchymal
stromal cells, hematopoietic cells, and endothelial progenitor cells332–334. The outcomes of these
trials were modest at best in terms of cardiac functional improvement332–334.
Another interesting regenerative concept has been the reprograming of cardiac fibroblasts into
cardiomyocytes. It raises the appealing possibility that the same cells which produce deleterious
scarring following myocardial infarction, could be utilized to replenish the lost cardiac cells323.
Retroviral delivery of three factors crucial in cardiomyocyte differentiation, TBX5, myocyte
enhancer factor 2C (MEF2C), and GATA4, has been shown to successfully reprogram murine
fibroblasts into “cardiomyocyte-like cells”, suggesting that this could also occur in vivo335.
Hurdles to overcome still remain for this novel regenerative option, such as delivery of these
factors through other mechanisms than integrating viruses which pose issues for clinical
translation.
The mechanisms behind how different cell-based therapies have led to modest improvements in
heart function remain to be elucidated. The initial goal of these methods were to replenish lost
cardiomyocytes through the differentiation of the injected cells into cardiomyocytes, or directly
injecting differentiated cardiomyocytes into the tissue. However, all of this work has
demonstrated that only a low, permanent cellular engraftment level is achievable. For example,
25% of injected C-kit + cardiac progenitor cells present at 5 minutes following transplantation
remained after 24 hours, and only 2.8% remained after 35 days336. Furthermore, engraftment
efficiency is even lower when cells are delivered via an intracoronary injection as opposed to
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directly into the myocardium, and only a limited number of cells can be injected directly into the
myocardium at one time323. Based on the low level of cellular engraftment, it is hypothesized
that the number of cardiomyocytes formed directly from these cells is too low to be conveying
the functional benefits reported. These observations led to the concept that the injected cells are
secreting paracrine factors that act on the endogenous myocardium by either affecting the non-
cardiomyocytes to promote angiogenesis, or function on the cardiomyocytes themselves to
promote survival or cell division323.
If one of the major hypothesized mechanisms by which cell-based therapies improve cardiac
function is through stimulating endogenous cardiomyocyte proliferation by the secretion of
paracrine factors, then upregulating this process by methods that circumvent the logistical issues
surrounding cell transplantation may prove to be a more efficient method for promoting
cardiomyocyte regeneration.
Numerous studies have attempted to drive cardiomyocytes into cell division, through the
manipulation of cell cycle activators or inhibitors or pro-mitogenic signaling pathways.
Overexpression of cell cycle activators such as cyclin D1, D2 or D3 in cardiac specific
transgenic mice successfully resulted in the upregulation of cardiomyocyte DNA synthesis and
potentially, cell cycle re-entry337. Transgenic delivery of cyclin D2 mice was the only model (out
of the three) where persistent DNA synthesis following myocardial infarction was observed,
albeit, at a low efficiency337. However, it is important to note that this study did not provide any
evidence that cyclin D2 overexpression upregulated the actual incidence of cardiomyocyte cell
division, and only DNA synthesis assays were performed. A more recent study demonstrated that
cyclin D1 overexpression in cardiomyocyte resulted in increased DNA synthesis but that these
cells were blocked from entering M phase due to a lack in activation of CDK181. Thus, it is
possible that hearts of cyclin D2 transgenic mice had cardiomyocytes increasing in cell size
through re-entry into G1 and S phase, but had no increase in the number of newly formed cells,
due to the block at M phase. In fact, the demonstration of cardiomyocytes actually undergoing
cytokinesis has been rare in this field of research. More commonly surrogate markers of
proliferation such as phosphorylated histone H3, as a marker of M phase, and the proliferation
antigen Ki-67 are utilized. This highlights one of the strengths of our study, in that we utilized
Aurora B-positive midbody structures as our primary marker for cardiomyocyte proliferation,
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which is to date some of the strongest evidence to suggest that cardiomyocytes are undergoing
the process of cytokinesis leading to the formation of new cells, as the mid-body is the last
structure linking the two daughter cells together70.
The manipulation of mitogenic signaling pathways that regulate proliferation is proposed to be a
more efficient method to promote cardiomyocyte cell division, as opposed to targeting one
specific cell cycle regulating factor323. This is a similar concept to what we have proposed:
targeting multiple miRNAs that have a broad effect on regulating cell cycle at different levels,
leading to the upregulation of multiple cell cycle factors, as opposed to one in particular. Studies
have shown that targeting the Hippo pathway, a highly evolutionarily conserved signaling
pathway that regulates proliferation, organ size, and cell survival, can have a strong effect on
cardiomyocyte proliferation338,339. Specifically, it has been shown that knockdown of the
upstream effector proteins in the Hippo pathway, or over expression of the downstream target
protein, yes associated protein 1 (YAP1), can promote cardiomyocyte proliferation and
ventricular wall thickening339. Conversely, inhibition of YAP1 during the course of cardiac
development led to cardiomyocyte hypoplasia and lethality340. Another study also showed that
the downstream target of YAP1, phosphatidylinositol-4,5-bisphosphate 3-kinase catalytic subunit
beta (PIK3CB) can upregulate cardiomyocyte proliferation when overexpressed338. They utilized
Aurora B as a marker for cardiomyocyte cytokinesis in vivo to demonstrate that cardiac specific
PIK3CB gain of function could promote the occurrence of cytokinesis in 0.01% of
cardiomyocytes, indicating how rare this process is, whereas the presence of PiH3 positive and
BrdU positive nuclei were increased 0.3% and 20% respectively338. The rate of cytokinesis
observed in this study is comparable with our observations. It would be interesting to determine
if combining both these approaches could have an additive effect on cardiomyocyte proliferation.
Administration of signaling molecules to the myocardium has also promoted cardiomyocyte cell
cycle re-entry. For example, treatment of cardiomyocytes with the cytokine fibroblast growth
factor-1, along with an inhibitor to the proapoptotic p38 MAPK pathway, promotes
cardiomyocyte proliferation, through a mechanism that promoted initial sarcomere
dedifferentiation, followed by proliferation341. It is interesting to note that a certain number of the
cells undergoing cytokinesis in our experiments had disorganized sarcomeric structures, whereas
cells that appeared to be in the very late stages of this process tended to have more maturely
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constructed sarcomeres. This observation is likely due to the massive cytoskeletal
rearrangements that arise throughout cytokinesis. In another approach, infusion of the molecule
neuregulin (NRG1) which signals through the ERBB2 receptor tyrosine kinase 2/4
(ERBBB2/ERBB4) pathway, was also shown to promote fetal and adult cardiomyocyte
proliferation, and these studies warranted a clinical trial342. NRG1 was subsequently infused into
patients with heart failure and promoted an acute improvement of cardiac function that lasted for
up to three months343. However, the mechanism of this functional improvement remains to be
elucidated.
As previously discussed, miRNAs also have emerged as important regulators of cardiac
development and the mediation of cell cycle arrest. Overexpression of miR-590 and miR-199a
was capable of promoting cardiomyocyte proliferation136. Others have demonstrated that
downregulating the expression of certain miRNAs such as the miR-15 family can promote
cardiomyocyte cell cycle re-entry and upregulate cytokinesis102. miRNAs are attractive
therapeutic targets because the miRNA mimic and antagomir technologies that have been
developed can be delivered as synthetic small molecules that have the capability of entering into
cells and exerting their function in vivo127. Another benefit to utilizing antagomirs or mimics is
that their effect is only transient, as opposed to utilizing viral vectors or other genome editing
techniques such as CRISPR. This is important because maintaining controlled levels of
proliferation is necessary to retain the structural integrity of the heart, and sustained proliferation
over extensive periods of time could have detrimental effects on heart function.
In all, the technique of stimulating endogenous cardiomyocyte cell cycle re-entry and
proliferation is very attractive in the field of cardiac regeneration, because it is through this
mechanism that regeneration occurs in lower vertebrates, as well as in neonatal mammals271.
This method promotes the generation of new, autologous cardiomyocytes, which implies that
they will be mechanically, electrically and vascularly integrated into the myocardium, because
they arise from cells that already have these properties, as opposed to exogenous cells that have
to surpass the hurdles posed by engraftment. Future work remains to determine whether the level
of cardiomyocyte proliferation that has been successfully induced in our lab and in others, is
sufficient to expand the cardiomyocyte population in a clinically meaningful manner.
Importantly, the optimal method for cardiomyocyte cell cycle reactivation is still up for debate. It
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is possible that taking a multi-pronged approach by combining multiple methods outlined in this
thesis e.g. targeting miRNAs, in tandem with the administration of certain cytokines such as
fibroblast growth factor-1, might be more effective than any one method on their own. The work
that is presented herein provides insight into the potential mechanisms by which cardiomyocytes
maintain cell cycle arrest, and offers novel targets that could be manipulated in a clinical setting
to promote cardiac regeneration. Ultimately, our final goal would be to initiate a phase III
clinical trial with the use of an optimized cocktail of antagomirs, delivered via an intra-coronary
injection at the time of reperfusion following presentation of patients with acute myocardial
infarction. By this approach, the antagomirs could reach the area of damage and transiently
reactivate the cell cycle in cardiomyocytes to help replenish the lost cardiomyocyte population,
while possibly also promoting other beneficial effects such as inhibiting apoptosis and
controlling inflammation.
4.2 Limitations and Future Directions
Despite the exciting findings revealed throughout this thesis, there are a number of limitations
that warrant further discussion, to pave the way for future experimentation and directions.
Firstly, to determine the initial miRNA expression profiles between the three knockout mice
(p53KO, Mdm2KO and DKO) in our study, we utilized RNA isolated from whole hearts, as
opposed to cardiomyocyte specific RNA. Although knockout of the three genes was carried out
using a cardiac-specific promoter, there is a high degree of cross talk between fibroblasts and
cardiomyocytes in the myocardium. For this reason, it is possible that miRNA gene expression
changes were also occurring in the fibroblasts, and by our method, there was no way to
determine if downregulation of the 11 identified “anti-proliferative” miRNAs was occurring
solely in cardiomyocytes. In an attempt to partially address this issue, we determined the
expression profile of these miRNAs in wild type cardiac-derived fibroblasts, and observed that
the miRNAs were already significantly downregulated, as compared to neonatal cardiomyocytes.
Therefore we hypothesized that the downregulation of these miRNAs that we observed between
DKO and wild type hearts was occurring in the cardiomyocytes, which allowed them to regain
their proliferative capacity. Another method to circumvent this limitation would be to
specifically isolate DKO cardiomyocytes by either magnetic assisted sorting or negative
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selection columns (which bind fibroblasts and endothelial cells but allow cardiomyocytes to pass
through). Performing the Nanostring miRNA array using RNA from these purified adult DKO
cardiomyocytes and wild type cardiomyocytes would provide us with a more precise depiction of
the miRNA gene expression changes occurring in cardiomyocytes specifically, in response to
loss of p53 and MDM2.
Secondly, to identify miRNA target genes, we utilized a target gene prediction software called
miRSystem. Computational prediction tools to determine miRNA target genes scan for Watson-
Crick base pairing nucleotides between the seed regions of miRNAs (nucleotides 2-7) and
3′UTR of genomic sequences, to identify putative miRNA binding sites344,345. Most prediction
tools now also make use of thermodynamics to predict the likelihood of these interactions to
occurs, as well as evolutionary conservation at the binding sites identified, binding site structural
accessibility and nucleotide composition of the regions flanking the binding sites to minimize
false positives, thereby increasing the specificity of target gene prediction344. Although use of
computational techniques is the most rapid and cost efficient method to determine putative
miRNA target genes, to date, miRNA target prediction tools continue to produce a high rate of
false positive hits, and an unknown number of false negatives344. Additionally, the predicted
target genes of a given miRNA derived from different prediction algorithms often do not yield
overlapping results. Due to these limitations, algorithm based miRNA target prediction tools
likely fall substantially short of capturing in complete detail the physical, temporal and spatial
requirements of miRNAs and their target mRNAs344. In effort to mitigate this limitation, we
utilized a software that combines data from 7 miRNA target gene prediction algorithms as well
as 2 experimental validation databases, whereby the target gene was required to be predicted by
4 out of the 7 algorithms to be considered a putative target. This likely produces a more accurate
representation of miRNA target genes compared to using only one of the algorithms. However,
the putative list of cell cycle target genes that we obtained from this program likely does not
reflect the exact mRNA transcripts targeted by the 11 identified miRNAs in vivo. In order to
improve the accuracy of the target genes predicted for the miRNAs of focus in this thesis, certain
experimental procedures are available. Traditionally, the identification of precise miRNA target
genes has been executed with the use of luciferase assays345. In this method, the target gene of
interest is linked to a luciferase reporter gene, and following overexpression of the miRNA of
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interest in vitro, luciferase activity is monitored (reduction in luciferase activity indicates
miRNA binding to target gene)345. While this method is highly specific, it is labour intensive, has
low throughput and requires pre-existing knowledge of the miRNA target gene. Alternatively,
many groups have utilized microarray platforms or next generation sequencing following the
immunoprecipitation of AGO2 or other proteins of the RISC complex, to capture all the mRNA
transcripts bound by RISC complexes within a cell (RIP-Chip, or RIP-Seq)345. Although this
technique is high-throughput, can illuminate cell type specific transcripts being silenced, and can
expose changes in gene silencing following diverse treatments, it still relies on computational
techniques to determine which miRNAs are targeting the mRNAs that were sequenced345.
Recently, a moderate throughput method has been described which allows the detection of
specific miRNA target genes by the use of synthetic miRNA-duplexes346. In this method, cells
are transfected with biotinylated miRNA duplexes, then, streptavidin coated beads are utilized to
pull down these duplexes, and the mRNAs bound to the duplexes are sequenced to determine the
direct mRNA targets for a given miRNA346. In our study, we carried out a cell cycle RT-qPCR
array following our antagomir treatments, and were capable of detecting an acceptable amount of
overlap between the target genes predicted by the miRSystem software, and the gene expression
changes detected by RT-qPCR. In the future, to determine the precise target genes of the 11
p53/MDM2-regulated miRNAs in cardiomyocytes, the technique described above could be
utilized. This would allow us to construct a more detailed model which accurately illustrates the
gene silencing effects that these miRNAs exert within the heart.
Thirdly, the detection method that we utilized to determine cardiomyocyte proliferation could be
optimized. Our method was based on the presence of Aurora B kinase positive mid-body
structures located between two cardiomyocytes. By fluorescence microscopy, the number of
these structures were counted in each well containing treated or non-treated cells (48 well plates
were utilized). Unfortunately, this method is inherently biased due to the fact that an individual
must determine what corresponds to an “Aurora B positive mid-body structure”. This can
sometimes be a difficult task due to the fact that the structures change in appearance depending
on the stage of cytokinesis that is occurring. Furthermore, cell counting was not executed in a
blinded manner, and the treatments in each well were known, which further increased bias. The
screening was also only carried out in one plane, and thus any proliferating cardiomyocytes
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found in a different plane could be missed, possibly resulting in an underestimation in the total
number of proliferating cells per well. Although the method utilized to detect proliferation is
more accurate than utilizing surrogate markers such as phosphorylated histone H3 or Ki-67,
alternative methods could be employed to minimize bias within the results obtained. For
example, the use of an automated microscope that takes a Z-stack of each well in its entirety, and
a computer program that mathematically computes the number of Aurora B positive signals
found in each well (based on a threshold defined manually), would be the ideal way to calculate
the number of proliferating cells in each well. For future experimentation, we will utilize the
Opera Phenix High-Content Screening System (PerkinElmer) that is now available. This system
contains a fluorescent confocal microscope that allows the screening of cells in many different
formats of plates (including 48 well), as opposed to only glass slides, and can be automated to
take in focus Z-stack images in each well. Using this system would largely increase the
reliability of our results.
Finally, there are limitations in the in vitro model system that we employed. Based on ease of
isolation and culture, we utilized neonatal rat cardiomyocytes to assess the ability of the diverse
antagomir cocktails to promote proliferation. Although this in vitro system has been extensively
utilized to characterize cardiomyocyte proliferation and cell cycle arrest, it is not optimal based
on the fact that neonatal cardiomyocytes have a higher proliferative capacity compared to adult
cardiomyocytes. Thus, upregulation of proliferation would be inherently easier to induce in these
cells, compared to cells isolated from adult hearts that have been arrested from the cell cycle for
many weeks or months. We transfect our cells at 4-5 days following birth, and it has been
suggested that by this time point cardiomyocytes have already exited the cell cycle, however it is
likely that over time, they become increasingly refractory to proliferative stimuli. Thus, to
provide strong evidence that the 11 identified “anti-proliferative” miRNAs are in fact regulating
cell cycle arrest in adult hearts, it is of utmost importance that the antagomir treatments we have
formulated be tested in either adult cardiomyocyte cultures, or in vivo, in adult mouse or rat
hearts.
Based on the limitations defined, the most important future experiment to be carried out would
be to determine the effect of the antagomir treatments in vivo. If positive results were obtained
following in vivo studies, whereby the antagomir treatments significantly upregulated adult
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cardiomyocyte proliferation, this would validate the results obtained from our in vitro screen.
LNA-modified antagomirs have been developed by the company Exiqon347. These molecules
potently downregulate miRNA targets when injected in vivo without the requirement of
transfection reagents, which can be highly toxic to cells. We would inject both the final
optimized cocktail of 9 antagomirs, and the 11 antagomir cocktail into mice to determine the
effect of downregulating p53/MDM2-regulated miRNAs in vivo. These injections could be
executed intramyocardially, to determine the precise effect of the treatment on the heart, as well
as intravenously, to determine if other tissues are also affected. For example, it will be
interesting to determine if cardiac fibroblast proliferation is upregulated by the antagomir
cocktails. We showed that the expression of the 11 identified miRNAs is already downregulated
in fibroblasts, and therefore their further downregulation may not have a potent effect.
Alternatively, this could significantly upregulate fibroblast proliferation to a deleterious amount
in the heart. It would also be of interest to determine if the antagomirs can cross the blood brain
barrier and promote proliferation in the nervous system, where certain cell types such as neurons
are also notorious for their lack of regenerative capability348. In vivo experiments would also help
elucidate the effect that inhibition of p53/MDM2-regulated miRNAs has on overall heart
function and survival, and would allow us to investigate other phenotypes such as hypertrophy,
and apoptosis that may be up or downregulated by the treatments. During these experiments, we
could investigate other pathways that these miRNAs have been predicted to regulate, such as the
MAPK pathway, the ERBB pathway, FGF signaling, and the phosphatidylinositide 3-kinase/
AKT serine/threonine kinase (PI3K/AKT) pathway, all of which have been shown to regulate
cardiomyocyte proliferation, as described above. The miRNAs identified in this thesis also have
multiple putative targets within each of these pathways, including the PIK3CB protein which
was shown to be an important regulator of YAP1 induced cardiomyocyte proliferation338
Certainly, the p53/MDM2-regulated miRNAs identified in the work presented herein have a
much more broad regulation over the cell cycle than just the precisely defined cell cycle pathway
itself. Delving into other signaling pathways which ultimately influence the cell cycle pathway
would provide us with a more holistic understanding of how these miRNAs regulate cell cycle
arrest within cardiomyocytes.
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Ultimately, if the antagomir treatments we have devised successfully promote adult
cardiomyocyte proliferation, we could begin to determine their therapeutic value. We could
investigate the use of these treatments in models of ischemia/reperfusion or myocardial
infarction, through intracoronary injection (in larger animal models), or by peri-infarct injections.
Subsequently, proliferation of cardiomyocytes would be assessed to determine if the cocktails
can upregulate cell cycle re-entry following injury, and if this has therapeutic value based on
improved cardiac functional outcomes.
Identifying miRNAs regulated by other tumor suppressors or oncogenes could provide us with
further insight into cardiac cell cycle regulation. Our lab has multiple other cardiac specific
knockout strains for crucial tumor suppressors such as ARF and RB. Eventually, we would like
to utilize the same approach taken throughout this study to elucidate novel miRNA subsets
regulated by these proteins. It will be intriguing to determine if the miRNA profiling in these
mutant mice will have overlaps with the miRNA profiles from p53/MDM2 knockout mice, as the
ARF/RB and p53/MDM2 signaling pathways are highly interconnected349. Alternatively,
inhibiting miRNAs regulated by both pathways uniquely could produce an additive effect on
cardiomyocyte proliferation. These future avenues of research highlight that the intricately
interconnected pathways governing the cell cycle in cardiomyocytes remain largely unknown,
and warrant substantial additional research.
4.3 Overall Conclusion
To conclude, the research presented in this thesis demonstrated that the cardiac cell cycle is
tightly regulated by complex pathways including tumor suppressors and their downstream small
RNA effectors. We have identified 11 miRNAs regulated by the presence of the crucial tumor
suppressor circuitry of p53 and MDM2 in cardiomyocytes, which function to promote cell cycle
arrest within these cells. Importantly, we highlight the concept of synergism within the cell,
whereby multiple regulators of a pathway function together to promote a specific phenotype.
This research breaks away from the mold of “one gene, one phenotype” or the classical miRNA
studies that conclude one given miRNA target gene is the effector of the intracellular changes
elicited following deregulation of the miRNA. Instead, it shows that regulation of the cell cycle
within cardiomyocytes involves a complex network of genes mediating this process with varying
115
degrees of redundancy, explaining the limited success in cell cycle reactivation through
modulating a single activator or inhibitor. Importantly, this thesis has provided an increased
understanding on how cardiomyocytes maintain cell cycle arrest, and paves the way for the
development of novel cardiovascular therapies by inhibiting groups of miRNAs who
synergistically dampen the regenerative potential in the heart.
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Biochim. Biophys. Acta - Mol. Basis Dis. 1812, 364–380 (2011).
349. Sherr, C. J. & McCormick, F. The RB and p53 pathways in cancer. Cancer Cell 2, 103–112 (2002).
136
Copyright Acknowledgements
Permissions granted for use of external figures:
Figure 1.1 - Anatomy of the cardiac sarcomere. Harvey, P. A. & Leinwand, L. A. The cell biology of disease: cellular mechanisms of
cardiomyopathy. J. Cell Biol. 194, 355–65 (2011).
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Shanna Stanley-Hasnain
Today 2:13 PM
Sent Items
August 4, 2016 Dear Dr. Leinwand,
I am a University of Toronto graduate student at the Institute of Medical Science completing my Master’s thesis entitled “Control of Cardiomyocyte Proliferation by p53/Mdm2-Regulated microRNAs”.
My thesis will be available in full text on the internet for reference, study and / or copy. Except in situations where a thesis is under embargo or restriction, the electronic version will be accessible through the U of T Libraries web pages, the Library’s web catalogue, and also through web search engines. I will also be granting Library and Archives Canada and ProQuest/UMI a non-exclusive license to reproduce, loan, distribute, or sell single copies of my thesis by any means and in any form or format.
I am requesting your permission to allow inclusion of the following material in my thesis: Figure 1. Anatomy of the cardiac sarcomere, from the published article Harvey, P. A. & Leinwand, L. A. The cell biology of disease: cellular mechanisms of cardiomyopathy. J. Cell Biol. 194, 355–65 (2011).
The material will be attributed through a citation. Please confirm in writing or by email that these arrangements meet with your approval.
Thank you very much for your time.
Sincerely, -- Shanna Stanley-Hasnain
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I believe we assigned copyright to the Journal of Cell Biology. I always ask journal when I want to reproduce a figure from a published article, so I think you have to go through JCB as we no long own the rights.
Journal of Cell Biology / Rockefeller University Press Copyright Policy:
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Figure 1.2 - The main molecular proponents of myocardial ischemia reperfusion injury. Hausenloy, D. J. & Yellon, D. M. Myocardial ischemia-reperfusion injury: a neglected
therapeutic target. J. Clin. Invest. 123, 92–100 (2013).
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Dear Dr. Yellon, I am a University of Toronto graduate student at the Institute of Medical Science completing my Master’s thesis entitled “Control of Cardiomyocyte Proliferation by p53/Mdm2-Regulated microRNAs”.
My thesis will be available in full text on the internet for reference, study and / or copy. Except in situations where a thesis is under embargo or restriction, the electronic version will be accessible through the U of T Libraries web pages, the Library’s web catalogue, and also through web search engines. I will also be granting Library and Archives Canada and ProQuest/UMI a non-exclusive license to reproduce, loan, distribute, or sell single copies of my thesis by any means and in any form or format.
I am requesting your permission to allow inclusion of the following material in my thesis: Figure 1. Schematic illustrating the main proponents of acute myocardial IRI, from the published article Hausenloy, D. J. & Yellon, D. M. Myocardial ischemia-reperfusion injury: a neglected therapeutic target. J. Clin. Invest. 123, 92–100 (2013).
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Thank you very much for your time. Sincerely, -- Shanna Stanley-Hasnain
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University of Toronto
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Yellon, Derek <[email protected]>
Dear Shana Thank you for your email. Let me confirm that you have my full permission to include Figure 1 from our J.
Clin. Invest. 123, 92–100 (2013) paper for your thesis. I hope it all goes well. Best wishes Derek Professor Derek Yellon Professor of Molecular & Cellular Cardiology Director of the Hatter Cardiovascular Institute UCL, Programme Director (Cardiometabolic), NIHR UCLH Biomedical Research Centre. University College London & UCLH 67 Chenies Mews, London WC1E 6HX Tel: +44 203 447 9591 Fax: +44 203 447 9818 PA:[email protected]
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Figure 1.3 - The major events of the cell cycle are regulated by transitions between CDK/cyclin complexes.
Rhind, N. & Russell, P. Signaling pathways that regulate cell division. Cold Spring Harb. Perspect. Biol. 4, a005942 (2012).
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Dear Dr. Rhind,
I am a University of Toronto graduate student at the Institute of Medical Science completing my Master’s thesis entitled “Control of Cardiomyocyte Proliferation by p53/Mdm2-Regulated microRNAs”.
My thesis will be available in full text on the internet for reference, study and / or copy. Except in situations where a thesis is under embargo or restriction, the electronic version will be accessible through the U of T Libraries web pages, the Library’s web catalogue, and also through web search engines. I will also be granting Library and Archives Canada and ProQuest/UMI a non-exclusive license to reproduce, loan, distribute, or sell single copies of my thesis by any means and in any form or format.
I am requesting your permission to allow inclusion of the following material in my thesis: Box 1. The main stages of mitosis, and Figure 1. The major events of the cell cycle, from the published article Rhind, N. & Russell, P. Signaling pathways that regulate cell division. Cold Spring Harb. Perspect. Biol. 4, a005942 (2012).
The material will be attributed through a citation. Please confirm in writing or by email that these arrangements meet with your approval.
Thank you very much for your time. Sincerely, -- Shanna Stanley-Hasnain
Master's Candidate | Institute of Medical Science, Cardiovascular Sciences Collaborative Program University of Toronto
The University Health Network
Toronto General Research Institute
Billia Lab, MaRS Centre, Max Bell Research Center
(647)-985-0770
Address:
232 Gerrard Street East, Apartment D Toronto, ON, Canada M5A 2E8
142
Reply all| Today 3:02 PM
Shanna Stanley-Hasnain;
Dear Shanna,
You have my permission to use the requested figure.
Good luck with your thesis,
Nick
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Figure 1.4 - Assembly of the pre-replicative and pre-loading complex. Aladjem, M. I. Replication in context: dynamic regulation of DNA replication patterns in
metazoans. Nat. Rev. Genet. 8, 588–600 (2007).
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Dear Dr. Aladjem
I am a University of Toronto graduate student at the Institute of Medical Science completing my Master’s thesis entitled “Control of Cardiomyocyte Proliferation by p53/Mdm2-Regulated microRNAs”.
My thesis will be available in full text on the internet for reference, study and / or copy. Except in situations where a thesis is under embargo or restriction, the electronic version will be accessible through the U of T Libraries web pages, the Library’s web catalogue, and also through web search engines. I will also be granting Library and Archives Canada and ProQuest/UMI a non-exclusive license to reproduce, loan, distribute, or sell single copies of my thesis by any means and in any form or format.
I am requesting your permission to allow inclusion of the following material in my thesis: Figure 1. Assembly of pre-replication and pre-initiation protein complexes, from the published article Aladjem, M. I. Replication in context: dynamic regulation of DNA replication patterns in metazoans. Nat. Rev. Genet. 8, 588–600 (2007).
The material will be attributed through a citation. Please confirm in writing or by email that these arrangements meet with your approval.
Thank you very much for your time. Sincerely, -- Shanna Stanley-Hasnain
Master's Candidate | Institute of Medical Science, Cardiovascular Sciences Collaborative Program
University of Toronto
The University Health Network
Toronto General Research Institute Billia Lab, MaRS Centre, Max Bell Research Center
(647)-985-0770
Address: 232 Gerrard Street East, Apartment D
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Aladjem, Mirit (NIH/NCI) [E] <[email protected]>
Dear Ms. Stanley-Hasnain Thank you for contacting me. I am happy that you are using this figure, and I will be happy to grant you permission, however have you checked that you do not need permission from Nature Reviews Genetics? If the publisher states that you need permission from me, you have my permission. Best of luck wit your thesis. Sincerely, Mirit I. Aladjem, Ph.D. Senior Investigator Head, DNA replication Group, Developmental Therapeutics Branch Center for Cancer Research, NCI, NIH Bldg. 37, Rm 5056 37 Convent Dr. Bethesda, MD 20892-4255 Tel. 301-435-2848 Fax 301-402-0752
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Figure 1.5 - The main stages of mitosis. Rhind, N. & Russell, P. Signaling pathways that regulate cell division. Cold Spring Harb.
Perspect. Biol. 4, a005942 (2012).
Permission to Use Copyrighted Material in a Master’s Thesis Shanna Stanley-Hansain
Today 2:31 PM
Sent Items
August 4, 2016
Dear Dr. Rhind,
I am a University of Toronto graduate student at the Institute of Medical Science completing my Master’s thesis entitled “Control of Cardiomyocyte Proliferation by p53/Mdm2-Regulated microRNAs”.
My thesis will be available in full text on the internet for reference, study and / or copy. Except in situations where a thesis is under embargo or restriction, the electronic version will be accessible through the U of T Libraries web pages, the Library’s web catalogue, and also through web search engines. I will also be granting Library and Archives Canada and ProQuest/UMI a non-exclusive license to reproduce, loan, distribute, or sell single copies of my thesis by any means and in any form or format.
I am requesting your permission to allow inclusion of the following material in my thesis: Box 1. The main stages of mitosis, and Figure 1. The major events of the cell cycle, from the published article Rhind, N. & Russell, P. Signaling pathways that regulate cell division. Cold Spring Harb. Perspect. Biol. 4, a005942 (2012).
The material will be attributed through a citation. Please confirm in writing or by email that these arrangements meet with your approval.
Thank you very much for your time. Sincerely, -- Shanna Stanley-Hasnain
Master's Candidate | Institute of Medical Science, Cardiovascular Sciences Collaborative Program
University of Toronto The University Health Network
Toronto General Research Institute
Billia Lab, MaRS Centre, Max Bell Research Center
(647)-985-0770
Address:
232 Gerrard Street East, Apartment D Toronto, ON, Canada
M5A 2E8
149
Reply all| Today 3:02 PM
Shanna Stanley-Hasnain;
Dear Shanna,
You have my permission to use the requested figure.
Good luck with your thesis,
Nick
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Figure 1.6 – Aurora B Localization During Mitosis and Cytokinesis. D’Avino, P. P. & Capalbo, L. New Auroras on the Roles of the Chromosomal Passenger
Complex in Cytokinesis: Implications for Cancer Therapies. Front. Oncol. 5, 221 (2015).
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[email protected]; [email protected]
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Dear Dr. D'Avino and Dr. Capalbo
I am a University of Toronto graduate student at the Institute of Medical Science completing my Master’s thesis entitled “Control of Cardiomyocyte Proliferation by p53/Mdm2-Regulated microRNAs”.
My thesis will be available in full text on the internet for reference, study and / or copy. Except in situations where a thesis is under embargo or restriction, the electronic version will be accessible through the U of T Libraries web pages, the Library’s web catalogue, and also through web search engines. I will also be granting Library and Archives Canada and ProQuest/UMI a non-exclusive license to reproduce, loan, distribute, or sell single copies of my thesis by any means and in any form or format.
I am requesting your permission to allow inclusion of the following material in my thesis: Figure 1. The CPC shows dynamic localization during mitosis and cytokinesis, from the published article D’Avino, P. P. & Capalbo, L. New Auroras on the Roles of the Chromosomal Passenger Complex in Cytokinesis: Implications for Cancer Therapies. Front. Oncol. 5, 221 (2015). The material will be attributed through a citation. Please confirm in writing or by email that these arrangements meet with your approval.
Thank you very much for your time. Sincerely, -- Shanna Stanley-Hasnain
Master's Candidate | Institute of Medical Science, Cardiovascular Sciences Collaborative Program
University of Toronto
The University Health Network
Toronto General Research Institute Billia Lab, MaRS Centre, Max Bell Research Center
(647)-985-0770
Address: 232 Gerrard Street East, Apartment D
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Dear Frontiers of Oncology Journal, I am a University of Toronto graduate student at the Institute of Medical Science completing my Master’s thesis entitled “Control of Cardiomyocyte Proliferation by p53/Mdm2-Regulated microRNAs”. My thesis will be available in full text on the internet for reference, study and / or copy. Except in situations where a thesis is under embargo or restriction, the electronic version will be accessible through the U of T Libraries web pages, the Library’s web catalogue, and also through web search engines. I will also be granting Library and Archives Canada and ProQuest/UMI a non-exclusive license to reproduce, loan, distribute, or sell single copies of my thesis by any means and in any form or format. I am requesting your permission to allow inclusion of the following material in my thesis: Figure 1. The CPC shows dynamic localization during mitosis and cytokinesis, from the published article D’Avino, P. P. & Capalbo, L. New Auroras on the Roles of the Chromosomal Passenger Complex in Cytokinesis: Implications for Cancer Therapies. Front. Oncol. 5, 221 (2015). The material will be attributed through a citation. I was not able to request permission through the Copyright Clearance Center because the option for requestion to use a single figure in an academic thesis was not available. On the site it said "Copyright Clearance Center has upgraded its republication service. Unfortunately, the rightsholder (copyright owner) is not currently enrolled. Consider contacting the rightsholder directly for assistance." Please confirm in writing or by email that these arrangements meet with your approval. Alternatively, it would be greatly appreciated if you could direct me to the appropriate individual to approach for obtaining permission to use this figure in my thesis. Thank you very much for your time. Sincerely, -- Shanna Stanley-Hasnain Master's Candidate | Institute of Medical Science, Cardiovascular Sciences Collaborative Program
University of Toronto
The University Health Network
Toronto General Research Institute
Billia Lab, MaRS Centre, Max Bell Research Center
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Pier Paolo D'Avino <[email protected]>
Dear Shanna,
Sorry for the delay.
We are happy for you to use our Figure in your thesis, providing that our paper is cited in
the Figure itself or in its legend.
Regards,
Paolo D'Avino and Luisa Capalbo
Reply all| Today 1:48 AM
Shanna Stanley-Hasnain
Frontiers Editorial Office <[email protected]>
Dear Mrs Stanley-Hasnain,
Thank you for your inquiry. All frontiers articles are published under the CC-BY 4.0 license,
which allows re-use provided that the original creators are correctly attributed.
As a courtesy, I would recommend that you inform the originators of the work, Pier Paolo
D’Avino, ([email protected];) and Luisa Capalbo, ([email protected]).
Best regards,
Faraz Alam, PhD
Ethics & Integrity Specialist
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Figure 1.7 - The Canonical Pathway of miRNA Biogenesis. Winter, J., Jung, S., Keller, S., Gregory, R. I. & Diederichs, S. Many roads to maturity:
microRNA biogenesis pathways and their regulation. Nat. Cell Biol. 11, 228–234 (2009).
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August 4, 2016
Dear Dr. Diederichs
I am a University of Toronto graduate student at the Institute of Medical Science completing my Master’s thesis entitled “Control of Cardiomyocyte Proliferation by p53/Mdm2-Regulated microRNAs”.
My thesis will be available in full text on the internet for reference, study and / or copy. Except in situations where a thesis is under embargo or restriction, the electronic version will be accessible through the U of T Libraries web pages, the Library’s web catalogue, and also through web search engines. I will also be granting Library and Archives Canada and ProQuest/UMI a non-exclusive license to reproduce, loan, distribute, or sell single copies of my thesis by any means and in any form or format.
I am requesting your permission to allow inclusion of the following material in my thesis: Figure 1. The ‘linear’ canonical pathway of microRNA processing, from the published article Winter, J., Jung, S., Keller, S., Gregory, R. I. & Diederichs, S. Many roads to maturity: microRNA biogenesis pathways and their regulation. Nat. Cell Biol. 11, 228–234 (2009). The material will be attributed through a citation. Please confirm in writing or by email that these arrangements meet with your approval.
Thank you very much for your time. Sincerely, -- Shanna Stanley-Hasnain
Master’s Candidate | Institute of Medical Science, Cardiovascular Sciences Collaborative Program
University of Toronto
The University Health Network
Toronto General Research Institute Billia Lab, MaRS Centre, Max Bell Research Center
(647)-985-0770
Address: 232 Gerrard Street East, Apartment D
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Diederichs, Sven <[email protected]>
Dear Ms. Stanley-Hasnain, thank you for your email and your interest in our research. If you cite the source accordingly as well as for non-commercial purposes, I grant you permission to use our figure. Best regards, Sven Diederichs
________________________________________ Prof. Dr. Sven Diederichs University Professor - Cancer Research (University Hospital Freiburg) Division Head - RNA Biology & Cancer (German Cancer Research Center Heidelberg) Albert-Ludwigs-University Freiburg Medical Center - Department of Thoracic Surgery Division of Cancer Research Breisacher Str. 115, 79106 Freiburg, Germany German Cancer Research Center (DKFZ) Foundation under Public Law Division of RNA Biology & Cancer Im Neuenheimer Feld 280 (B150), 69120 Heidelberg, Germany Management Board: Prof. Dr. Michael Boutros (interim), Prof. Dr. Josef Puchta, VAT-ID No.: DE143293537 [email protected] www.diederichslab.org
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Figure 1.8 - An overview to cell cycle control by microRNAs. Bueno, M. J. & Malumbres, M. MicroRNAs and the cell cycle. Biochim. Biophys. Acta - Mol.
Basis Dis. 1812, 592–601 (2011).
Permission to Use Copyrighted Material in a Master’s Thesis Shanna Stanley-Hasnain
Today 3:16 PM
Sent Items
August 4, 2016 Dear Dr. Malumbres,
I am a University of Toronto graduate student at the Institute of Medical Science completing my Master’s thesis entitled “Control of Cardiomyocyte Proliferation by p53/Mdm2-Regulated microRNAs”.
My thesis will be available in full text on the internet for reference, study and / or copy. Except in situations where a thesis is under embargo or restriction, the electronic version will be accessible through the U of T Libraries web pages, the Library’s web catalogue, and also through web search engines. I will also be granting Library and Archives Canada and ProQuest/UMI a non-exclusive license to reproduce, loan, distribute, or sell single copies of my thesis by any means and in any form or format.
I am requesting your permission to allow inclusion of the following material in my thesis: Figure 1. An overview to cell cycle control by microRNAs, from the published article Bueno, M. J. & Malumbres, M. MicroRNAs and the cell cycle.Biochim. Biophys. Acta – Mol. Basis Dis. 1812, 592–601 (2011). The material will be attributed through a citation. Please confirm in writing or by email that these arrangements meet with your approval. Thank you very much for your time. Sincerely, -- Shanna Stanley-Hasnain
Master’s Candidate | Institute of Medical Science, Cardiovascular Sciences Collaborative Program
University of Toronto The University Health Network
Toronto General Research Institute
Billia Lab, MaRS Centre, Max Bell Research Center
(647)-985-0770
Address: 232 Gerrard Street East, Apartment D Toronto, ON, Canada
M5A 2E8
159
Reply all| Today 3:17 PM
Shanna Stanley-Hasnain;
Dear Shanna, Sure, I am glad you are interested in this figure. Please use it at will! Best Marcos
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Figure 1.9 - Regulation of the cell cycle by p53-induced miRNAs. Hermeking, H. MicroRNAs in the p53 network: micromanagement of tumour suppression. Nat.
Rev. Cancer 12, 613–626 (2012).
Permission to Use Copyrighted Material in a Master’s Thesis Shanna Stanley-Hasnain
Today 3:33 PM
Sent Items
August 4, 2016 Dear Dr. Hermeking,
I am a University of Toronto graduate student at the Institute of Medical Science completing my Master’s thesis entitled “Control of Cardiomyocyte Proliferation by p53/Mdm2-Regulated microRNAs”.
My thesis will be available in full text on the internet for reference, study and / or copy. Except in situations where a thesis is under embargo or restriction, the electronic version will be accessible through the U of T Libraries web pages, the Library’s web catalogue, and also through web search engines. I will also be granting Library and Archives Canada and ProQuest/UMI a non-exclusive license to reproduce, loan, distribute, or sell single copies of my thesis by any means and in any form or format.
I am requesting your permission to allow inclusion of the following material in my thesis: Figure 2. Regulation of the cell cycle, metastasis and stemness by p53-induced microRNAs, from the published article Hermeking, H. MicroRNAs in the p53 network: micromanagement of tumour suppression. Nat. Rev. Cancer 12, 613–626 (2012). The material will be attributed through a citation. Please confirm in writing or by email that these arrangements meet with your approval. Thank you very much for your time. Sincerely, -- Shanna Stanley-Hasnain
Master’s Candidate | Institute of Medical Science, Cardiovascular Sciences Collaborative Program University of Toronto
The University Health Network
Toronto General Research Institute
Billia Lab, MaRS Centre, Max Bell Research Center
(647)-985-0770
Address: 232 Gerrard Street East, Apartment D Toronto, ON, Canada M5A 2E8
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Today 4:56 AM
Shanna Stanley-Hasnain
Hermeking, Heiko Prof.Dr. <[email protected]>
Ok with me.
Prof. Heiko Hermeking
Experimental and Molecular Pathology
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