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Using epigenetic toggle switches to repress tumor-promoting gene expression
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Using epigenetic toggle switches to repress tumor-promoting gene expression
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Content
Using Epigenetic Toggle Switches to
Repress Tumor-promoting Gene Expression
By
Karly Nisson
Mentor: Dr. Peggy J. Farnham
A Thesis Presented to the
FACULTY OF THE USC GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the Requirements for the Degree
MASTER OF SCIENCE
(Biochemistry and Molecular Medicine)
August 2019
II
Acknowledgements
I would like to first thank my advisor, Dr. Peggy Farnham, for the invaluable support,
enthusiasm, and guidance she gave me throughout my MS career. She presented me with many
opportunities to develop and present my work, all while serving as a role model whose expertise
and methods of leadership I one day hope to emulate.
I would also like to give my heartfelt thanks to the rest of the Farnham lab, who each contributed
to an incredibly supportive and stimulating environment. I especially thank Dr. Charles Nicolet
and Andrew Perez, who welcomed me onto the toggle switch project and provided me with
continuous guidance and support in approaching experimental challenges. The latter served as
my close mentor, and I thank him for his constant encouragement and willingness to provide me
with helpful feedback. I thank Jenevieve Polin and Stephanie Weiya Ni for their collaboration in
the realm of ZFX, Dr. Suhn Rhie for her computational expertise, and Daniel Bsteh of Dr. Bell’s
lab for his help in navigating the FACS machines.
I would also like to express my gratitude to our collaborators, Dr. Henriette O’Geen and Dr.
David Segal of UC Davis, for allowing me to contribute to such an exciting, rigorous research
project.
Finally, I would like to thank the members of my committee, Dr. Oliver Bell and Dr. Michael
Stallcup, for offering their continuing support and expertise as I developed my project and career
aspirations.
III
Table of Contents
Acknowledgements......................…………………………………………………....………...…II
Table of Contents………………………………………...……...……………………………….III
List of Figures…………………………...……………......………………………………………V
List of Tables…………………………........………….………………………...……………….VI
Abstract..........................………..........…………………………………………………………VII
List of Abbreviations…………………………….……………………….……………………VIII
Chapter 1. Introduction……………………....……………………………………………………1
1.1 Epigenetic memory is initiated and maintained via a dynamic interplay between
histone modifications and DNA methylation…………......................……………………1
1.2 Aberrant epigenetic landscapes contribute to gene expression signatures associated
with cancer…………………………………………......................……………………….4
1.3 Epigenetic therapies for prostate cancer……………...........………………………….5
1.4 dCas9-based epigenome editing can be used to repress tumor-promoting
genes…………………………..………………………..…………………………………7
1.5 Depositing different epigenetic modifications may elicit variable transcriptional
outputs depending on the local chromatin environment……….…...........………………10
Chapter 2. Materials and Methods.................................................................................................16
2.1 dCas9 expression plasmid constructs……….….........………..……………………...16
2.2 Cell culture and transfection……………......……...…...…………..….…………….18
2.3 RNA extraction and reverse-transcription quantitative PCR (RT-qPCR)...................18
Chapter 3. Transient Repression of Promoters..............................................................................20
3.1 Experimental workflow to study transient repression.................................................20
3.2 Transient repression of the TRPM4 promoter is achieved with all targeted effector
domains..............................................................................................................................21
3.3 Very modest transient repression can be achieved when the dCas9-system is targeted
to GRP78............................................................................................................................24
3.4 Transient repression of ZFX can be achieved with dCas9-KRAB +/-
DNMTs..............................................................................................................................27
3.5 Summary and Future Studies.......................................................................................29
Chapter 4. Persistent Repression of Promoters..............................................................................31
4.1 Persistent repression may require the deposition of both histone modifications and
DNA methylation...............................................................................................................31
4.2 Persistent repression of TRPM4 is only achieved through the combined targeting of
histone modifiers and DNA methyltransferases................................................................32
4.3 Achievable persistent repression of GRP78 is comparable to levels of transient
repression achieved with the same effector domains.........................................................35
4.4 Transient repression of ZFX is not maintained regardless of the effector domains
targeted...............................................................................................................................37
4.5. dCas9-mediated repression of TRPM4 can be maintained for at least thirty days.....38
4.6 Summary and Future Studies.......................................................................................40
Chapter 5. Discussion and Future Studies.....................................................................................42
5.1 Achieving significant transient or persistent repression of a target gene relies on the
optimization of the dCas9 toggle system to individual chromatin
environments......................................................................................................................42
IV
5.2 More robust persistent repression may be achieved through increased transfection
efficiency............................................................................................................................45
5.3 Further analysis of persistent repression achieved via the dCas9 toggle system in
TRPM4...............................................................................................................................46
5.4 Introduction of toggle switches into patient tumors....................................................48
References......................................................................................................................................50
V
List of Figures
Figure 1.1 One model of epigenetic regulation by endogenous cellular machinery.......................4
Figure 1.2 Proposed mechanism of CRISPR-mediated epigenetic repression via modulating
H3K9me3 and DNA methylation levels............................................................................11
Figure 1.3 dCas9-ED fusions.........................................................................................................12
Figure 1.4. Proposed mechanism of EZH2-mediated transcriptional repression..........................13
Figure 1.5. Proposed mechanism of FOG1-mediated transcriptional repression..........................13
Figure 1.6. DNA methylation of promoter CpG islands in TRPM4, ZFX and GRP78.................14
Figure 2.1. Expression vectors used to clone either effector domain or gRNA sequences...........16
Figure 3.1. Transient repression workflow....................................................................................21
Figure 3.2. TRPM4 gRNA schematic............................................................................................22
Figure 3.3 Transient repression of TRPM4 can be achieved with all dCas9-ED constructs in
C42B and HEK293T cell lines..........................................................................................24
Figure 3.4. GRP78 gRNA schematic.............................................................................................25
Figure 3.5 Transient repression of GRP78 is ED- and cell type-specific......................................27
Figure 3.6. ZFX gRNA schematic.................................................................................................28
Figure 3.7 Transient repression of the ZFX promoter is influenced by cell type..........................29
Figure 4.1 Persistent repression workflow.....................................................................................32
Figure 4.2 Persistent repression of the TRPM4 promoter can be achieved with dCas9-KRAB +
DNMTs and dCas9-EZH2 + DNMTs................................................................................35
Figure 4.3. Persistent repression is not achieved with any dCas9-ED in either cell type..............37
Figure 4.4 Significant 14d persistent repression of ZFX cannot be maintained with any of the
dCas9-ED fusions in either cell line..................................................................................38
Figure 4.5. 30-day persistent repression of TRPM4 can be achieved with dCas9-KRAB +
DNMTs in C42B cells.......................................................................................................39
Figure 4.6. The dCas9-KRAB expression vector is no longer detectable by 14d.........................40
VI
List of Tables
Table 2.1. Sequences and locations of gRNAs targeting the TRPM4 promoter or first
enhancer.........................................................................................................................................17
Table 2.2. Sequences and locations of gRNAs targeting the GRP78 promoter............................17
Table 2.3 Sequences and locations of gRNAs targeting the ZFX promoter..................................18
Table 2.4 Sequences of qPCR primers for TRPM4, ZFX, GRP78 and GAPDH..........................19
VII
Abstract
Gene expression patterns are modulated by reversible changes to the epigenome. Such epigenetic
changes take the form of histone modifications and DNA methylation, which may work in
concert to stably activate or repress a gene. In cancer, the epigenetic landscapes of many tumor-
repressor genes and oncogenes are distinctive from those in normal cells. Abnormal epigenetic
signatures may thus drive carcinogenesis by altering the expression of key genes involved in
driving or suppressing tumorigenesis. Attempts to reverse these disease-associated expression
changes through artificial deposition of specific marks have largely focused on generating
transient results. Targeting powerful histone modifiers such as H3K9me3 or H3K27me3
depositors to a site of interest can achieve potent, specific gene repression. Maintaining this
induced repression, however, may require that histone modifiers be targeted in conjunction with
DNA methyltransferases. Here, I use a modified CRISPR-dCas9 system to explore whether
lasting, or persistent, repression of three tumor-promoting genes can be achieved with a variety
of epigenetic modifiers. I report that efficient DNA methylation in conjunction with the
repressive histone modification H3K9me3 can persistently silence a promoter for at least 30 days
post-transfection and the degree of achievable persistent repression is dependent on both the
target gene and cell type.
VIII
List of Abbreviations
AAV Adeno-associated virus
cDNA Complementary DNA
CRISPR/Cas9 Clustered, regularly interspaced, short palindromic repeat/ CRISPR
associated protein 9
CRISPRi CRISPR interference
DNMT3a DNA methyltransferase 3 alpha
DNMT3L DNA methyltransferase 3 like
DNMTs DNA Methyltransferases
ED Effector domain
EZH2 Enhancer of zeste homolog 2
FOG1 Friend of GATA-1
GRP78 Glucose-regulated protein, 78kD
gRNA Guide RNA
H3K4me3 Tri-methylation of histone 3 on lysine 4
H3K9ac Acetylation of histone 3 on lysine 9
H3K9me3 Tri-methylation of histone 3 on lysine 9
H3K27me3 Tri-methylation of histone 3 on lysine 27
HAT Histone acetyltransferase
HDAC Histone deacetylase
KRAB Kruppel-associated box
KRAB-ZNF Kruppel-type zinc finger
NuRD Nucleosome remodeling and deacetylase complex
PCa Prostate Cancer
PRC2 Polycomb repressive complex 2
TALEN Transcription activator-like effector nucleases
TRPM4 Transient receptor potential cation channel subfamily M member 4
TSS Transcription start site
ZFX Zinc finger protein X-linked
ZNF Zinc finger protein
1
Chapter 1. Introduction
1.1 Epigenetic memory is initiated and maintained via a dynamic interplay between histone
modifications and DNA methylation.
Genetic material is both compacted and dynamically regulated within chromatin, a complex of
DNA, RNA and associated proteins. The basic structural unit of chromatin is the nucleosome,
which contains approximately 147 bp of DNA wrapped around four core histone proteins.
Protruding from each histone are N- and C- terminal tails that can be post-translationally
modified to initiate remodeling of the local chromatin environment. These histone modifications
work in conjunction with DNA methylation to shift genomic regions between “open” and
“closed” conformations. In actively transcribed genes, chromatin exists in an “open” state that is
conducive to interactions between genetic material and cellular machinery. Silenced regions,
such as transposable elements that threaten genomic stability (Belancio, Deininger and Roy-
Engel 2009), are sequestered from this machinery in a “closed” conformation. Both histone
modifications and DNA methylation are essential components of the epigenome and serve as
mediators of epigenetic regulation (King et al. 2016).
Depositors of histone modifications and DNA methylation are represented as “writers”
correlated with either active or repressed transcriptional states. Epigenetic “readers” interpret
these chemical modifications and may then recruit remodeling complexes capable of altering the
local chromatin architecture. Finally, epigenetic “erasers” remove epigenetic modifications.
Specific patterns of modifications have been associated with active or repressed transcriptional
states. For instance, histone acetylation is characteristic of active gene promoters, where it may
function as a docking site for regulatory proteins that promote transcription (Verdone, Caserta
and Di Mauro 2005).
Histone acetylation is catalyzed by histone acetyltransferases (HATs), such as CBP or p300, and
can be removed by histone deacetylases (HDACs) (Bannister and Kouzarides 1996, Ogryzko et
al. 1996, Taunton, Hassig and Schreiber 1996, Rundlett et al. 1996). Additionally, histone tails
may either be mono- or di-methylated at arginine residues (Byvoet 1972) by members of the
2
PRMT family of histone methyltransferases or mono-, di- or tri-methylated at lysine residues
(MURRAY 1964) by members of the SET domain containing proteins (Rea et al. 2000) or Dot1
like proteins (Feng et al. 2002). Gene expression changes enacted by histone methylation can be
reversed by histone demethylases such as the amine oxidase KDM1, which can remove mono- or
di-methylation (Metzger et al. 2005, Shi et al. 2004). Demethylation of tri-methylated histones
can be accomplished by demethylases containing the Jumonji catalytic domain (Cloos et al.
2006, Tsukada et al. 2006, Whetstine et al. 2006, Yamane et al. 2006). The effects of histone
methylation on transcriptional regulation are context dependent (Chen et al. 2017, Greer and Shi
2012, Zhang and Reinberg 2001, O'Geen et al. 2019) and the degree and location of methylation
determines whether the mark is activating versus repressive. For instance, tri-methylation of
histone 3 on lysine 4 (H3K4me3) is generally associated with active promoters or bivalent
promoters poised for activation, while trimethylation of histone 3 on lysine 27 and 9 (H3K27me3
and H3K9me3, respectively) is associated with repressed chromatin.
DNA methylation is also an important regulator of transcriptional output and plays roles in many
biological processes including X-chromosome inactivation (Sharp et al. 2011, Nesterova et al.
2003, Hellman and Chess 2007) and loss of pluripotency (Nazor et al. 2012, Bibikova et al.
2006, Li 2002, Boland, Nazor and Loring 2014). In mammalian cells, methylation at the 5
position of cytosines within a CpG dinucleotide located in a promoter or enhancer is generally
correlated with gene repression (Mohn et al. 2008, Aran and Hellman 2013). Conversely, DNA
methylation of gene bodies is closely associated with active transcription states (Kungulovski et
al. 2015, Jurkowska, Jurkowski and Jeltsch 2011, Jones 2012).
Transcriptional regulation results from the combined effects of histone modifications and DNA
methylation, though the precise mechanism of this coordination has not been well established.
Recent evidence suggests that the relationship between DNA methyltransferases and some
histone modifications is bi-directional and mutually reinforcing (Soshnev, Josefowicz and Allis
2016). For instance, DNMTs have been shown to recruit H3K9 methyltransferases, including
those associated with heterochromatin formation, in a positive regulatory loop (Fuks et al.
2003a).
3
The relationship between regulatory marks is of particular interest when examining how
established chromatin states are maintained through cell divisions. The “piggy-back” model
proposes that histone modifying machinery is recruited to pre-existing, specific patterns of DNA
methylation. In support of this, MeCP2, a protein capable of binding to methylated CpGs, directs
SUV39H1 to target genes where it can deposit H3K9me3 marks (Fuks et al. 2003b). Further
studies have linked methyl-CpG binding H3K9 methyltransferases SETDB1 (Sarraf and
Stancheva 2004) and G9a (Estève et al. 2006) to DNMT1-dependent interactions with replication
forks, suggesting a model for the cooperation of histone and DNA modifications in chromatin
assembly and epigenetic inheritance.
Alternatively, studies demonstrating that EZH2 interacts with and directs DNMTs to target sites
(Viré et al. 2006) suggest that DNA methylation acts downstream of histone modifications. In
line with this hypothesis, the creation of a foundation suitable for an adaptor molecule such as
HP1 by H3K9 (Lehnertz et al. 2003, Fuks et al. 2003a) may recruit DNMTs to catalyze
methylation in nearby regions. This methylation may then attract HDAC complexes to remove
H3K9Ac and thus prepare additional, nearby nucleosomes for receiving the H3K9me3 mark
(Rea et al. 2000, Fuks 2005). This would allow spreading of the repressive chromatin
environment (Ayyanathan et al. 2003). A detailed illustration of these interactions can be found
in Figure 1.1.
4
Figure 1.1. One model of epigenetic regulation by endogenous cellular machinery. Step 1: A
KRAB domain-containing zinc finger transcription factor (KRAB-ZFP) may recruit KAP-1 and
associated HMTs and HDACs. Step 2: HDACs can remove the active H3K9Ac to allow the
HMTs to put on the repressive H3K9me3 mark, making the chromatin more accessible to
DNMTs. Step 3: DNMTs may then methylate nearby CpGs, locking the repressive effect of the
histone modifications in place and allowing the new chromatin state to be maintained through
future cell divisions.
Collectively, each model demonstrates a necessity for crosstalk between DNA methylation and
histone modifications in establishing epigenetic memory. However, it is apparent that further
details of this dialogue must be determined to better define a relationship between epigenetics
and transcriptional regulation.
1.2 Aberrant epigenetic landscapes contribute to gene expression signatures associated with
cancer.
Appropriate deposition and maintenance of epigenetic modifications is essential to ensure
normal gene expression patterns (Arechederra et al. 2018). However, epigenetic modifiers are
highly mutated in cancer cells (Morgan and Shilatifard 2015), which have an epigenome distinct
from normal cells (Berdasco and Esteller 2010). Both aberrant DNA methylation and histone
modifications have been implicated in many cancers (Esteller 2005, Goering, Kloth and Schulz
2012, Massie, Mills and Lynch 2017, Rauch et al. 2012, Ashktorab and Brim 2014, Nakazawa et
al. 2012, Silverman and Shi 2016) and may occur in a wide range of cellular pathways and stages
of cancer development (Esteller 2008).
5
The genomes of cancer cells are globally hypomethylated when compared with normal DNA
methylation patterns. Such aberrant patterns have been associated with inappropriate activation
of genomic regions that promote chromatin instability, increased chromosomal rearrangements,
aneuploidy, and activation of transposable elements (Berdasco and Esteller 2010, Eden et al.
2003, Karpf and Matsui 2005). In support of this, inactivating mutations in the DNA
methyltransferase 3 alpha (DNMT3a) have been linked to acute monocytic leukemias (Yan et al.
2011). In contrast, gene promoters can be either hypermethylated or hypomethylated in cancers.
For example, the CpG island promoters of tumor-suppressor genes are largely hypermethylated
in cancer cells and contribute to silencing in a tissue-specific manner (Esteller 2008, Futscher et
al. 2002, Berdasco and Esteller 2010). In contrast, CpG island promoters of genes that promote
cell division are frequently hypomethylated (Kanwal and Gupta 2012), contributing to an altered
transcriptional landscape that favors tumor-promoting gene expression. One of the most well-
known genes that is hypomethylated in cancer is the oncogenic transcription factor c-Myc
(Tsujiuchi et al. 1999, Cheah, Wallace and Hoffman 1984).
Mutations in histone modifiers are also associated with cancer diagnosis and outcomes. For
instance, mutated or overexpressed HDACs contribute to aberrant histone acetylation in multiple
tumor types (Juan et al. 2000, Fraga et al. 2005, Yasui et al. 2003, Choi et al. 2001, Halkidou et
al. 2004). Decreased frequency of other activating marks, such as H3K4me2 or H3K18ac, are
indicative of poor prognoses (Seligson et al. 2009). Additionally, aberrantly deposited repressive
modifications may recruit DNMTs, contributing to abnormal patterns of hypermethylation and
inappropriate gene silencing (Kanwal and Gupta 2012).
1.3 Epigenetic Therapies for Prostate Cancer.
The altered epigenetic landscape of prostate cancer (PCa) cells is distinctly relevant because of
its potential to reveal disease susceptibility and druggable gene targets. Despite a 100% survival
rate for localized PCa, prognoses for distant metastasis remain at a 29% 5-year survival rate.
These statistics illustrate the necessity for early diagnoses and intervention to prevent life-
threatening disease progression in an otherwise treatable disease.
6
Recent investigations into the altered gene expression patterns of PCa cells have identified
promising cancer-related biomarkers. Among these are upregulated tumor-promoting genes
whose contributions to PCa progression may be minimized by altering regulatory epigenetic
modifications. This study will focus on three such genes, all of which have been shown to
accelerate rates of PCa cell proliferation and migration in vitro (Sagredo et al. 2018, Holzmann
et al. 2015, Lu et al. 2015, Song et al. 2018). The first of these genes is transient receptor
potential cation channel, subfamily M, member 4 (TRPM4), a nonselective Ca
2+
-activated and
voltage dependent ion channel that regulates Ca
2+
influx by modulating the membrane potential
(Nilius et al., 2006). TRPM4 is overexpressed in prostate cancer, including the transitionary
phase of prostatic intraepithelial neoplasia’s (PIN) evolution to prostate cancer (Ashida et al.,
2004; Singh et al., 2006), and has been associated with androgen-independent prostate cancer
progression (Schinke et al., 2014). Though a detailed mechanism of TRPM4’s role in PCa is still
to be elicited, it has so far been implicated in the regulation of β-catenin signaling and epithelial
to mesenchymal transition (Sagredo A et al., 2018).
Another potential target for future PCa therapies is a 78-kDa glucose-regulated protein (GRP78),
also known as BiP and HSPA5. GRP78 has been associated with a wide range of cancers,
including glioma (Kang BR et al., 2016), pancreatic cancer (Niu Z et al., 2015), colon cancer
(Mhaidat NM et al., 2016) and PCa (de Ridder G et al., 2011), as well as neurological disorders
(Wang M et al., 2009). Multiple reports have identified it as a promising drug target (Staquicini
et al. 2018, Sato et al. 2010) because of its elevated expression on the cell surfaces of cancer
versus normal cells (Ni, Zhang and Lee 2011). In normal cells, GRP78 functions as an
endoplasmic reticulum chaperone by translocating newly synthesized polypeptides across the ER
membrane, assisting in their folding and targeting any misfolded proteins for degradation. In
addition to its roles as an ER chaperone, GRP78 acts as a master regulator for ER stress by
controlling the activation of the unfolded protein response. This response prevents apoptosis
under acute stress and triggers cell death when stress is severe or prolonged (Wang et al. 2009).
Notably, the microenvironment of tumor cells has been shown to simulate this physiologic stress,
thus resulting in inappropriate activation of the pro-survival pathways of the UPR (Koumenis
and Wouters 2006). However, the tumor-promoting, anti-apoptotic properties of GRP78 are
significantly reduced when its expression levels are decreased to 50%. Specifically, GRP78
7
haploinsufficiency has been shown to suppress acinar-to-ductal metaplasia and pancreatic
tumorigenesis in mutant Kras mice (Shen et al. 2017). In PCa, knockdown of GRP78 via
asymmetric small interfering RNAs increases rates of apoptosis and reduces proliferation in PC-
3 cells (Lu et al. 2015).
Of additional mounting therapeutic interest is the highly conserved transcriptional activator ZFX,
a zinc finger protein that has been associated with many human cancers, including PCa, breast
cancer, glioma, non-small cell lung carcinoma and colorectal cancer (Cai et al. 2018, Yang et al.
2014, Zhou et al. 2011, Jiang et al. 2012). ZFX may function as an oncogene (Rhie et al. 2018),
and recent studies have explored its potential mechanistic contributions to tumorigenesis. ZNFs
are generally involved in protein-protein interactions and sequence-specific binding of DNA and
RNA (Stubbs, Sun and Caetano-Anolles 2011, Najafabadi et al. 2015), though the biological
implications of these roles are yet to be defined. ZFX contains both a DNA-binding domain and
a large transcriptional activation domain at its N-terminus and has been shown to bind
downstream of transcription start sites (TSS) at the majority of CpG island promoters (Rhie et al.
2018). In PCa, ZFX expression is significantly increased when compared with expression levels
in benign prostate hyperplasia. siRNA mediated knockdown of ZFX in PCa cells results in
decreased proliferation and number of colonies in colony forming assays (Song et al. 2018).
1.4 dCas9-based epigenome editing can be used to repress tumor-promoting genes.
Gene-editing tools such as CRISPR/Cas9 (clustered, regularly interspaced, short palindromic
repeat/ CRISPR associated protein 9), engineered ZNFs and TALENs have significantly
improved efforts to better understand and treat a wide range of diseases (Panfil et al. 2018, Flynn
et al. 2015, Wang et al. 2014, Xu et al. 2015, Schwank et al. 2013, Carroll 2011) by altering
specific genes or pathways. For instance, in the field of cancer immunotherapy, CRISPR/Cas9
has been implemented to engineer therapeutic immune cells including chimeric antigen receptor
T cells and programmed cell death 1 protein knockout (Xia et al. 2019). Additionally, ZNFs have
been used to target functional copies of CCR5 in clinical HIV treatment trials (Tebas et al. 2014).
8
Though promising, the potentially damaging off-target effects of such technologies will remain a
substantial risk until the local and genome-wide implications of nuclease-based treatments are
better understood. For instance, recent studies report that CRISPR-Cas9-induced double stranded
breaks activate repair pathways that result in large deletions and complex rearrangements
(Kosicki, Tomberg and Bradley 2018), and CRISPR-Cas9 mediated knock-ins have resulted in
unexpected genomic rearrangements at target loci (Rezza et al. 2019). Both findings emphasize
that CRISPR-Cas9 will require rigorous screening for off-target effects and additional
modifications before it can be applied in routine clinical settings. Technologies that instead focus
on the alteration of the epigenome may allow for an effective alternative to gene editing.
Considering the widespread epigenetic alterations that occur in cancer cells, introduced
modifications to the epigenome may allow for tumor-specific reversals of oncogene activation
and tumor-suppressor gene silencing. One of many recent, innovative manipulations of the
CRISPR-Cas9 technology has allowed for such site-specific deposition of epigenetic
modifications. Cas9 can be made catalytically inactive through the introduction of two single
amino acid mutations (D10A, H849A). The resulting protein is nuclease-deficient (dCas9),
targetable via RNA guides, and can be fused to a wide range of epigenetic editors (Jinek et al.
2012).
Previous reports have fused both transcriptional activators and repressors to dCas9. Explorations
into the applications of these dCas9-effector domain fusions (dCas9-EDs) are ongoing, but
recent studies have used them to identify endogenous gene regulatory elements and combat
aberrant transcriptional outputs associated with disease. For instance, fusing the human
acetyltransferase p300 to dCas9 has allowed researchers to identify genetic regulatory elements
through gain-of-function screens in HEK293T cells (Klann et al. 2017). Additionally, dCas9
fusions incorporating the powerful transcriptional repressor KRAB can be used to robustly
silence oncogenes (Wang et al. 2018, O'Geen et al. 2017) and disrupt the activity of gene
regulatory elements (Thakore et al. 2015, Klein and O'Neill 2018, Gao et al. 2014, Kearns et al.
2015).
The potency of dCas9-ED induced transcriptional repression is especially promising in the
context of cancer. In cases where aberrant genetic activation promotes tumorigenesis, dCas9
9
technologies may be used to improve current therapies by repressing cancer-driver genes. This
study will investigate whether dCas9 fused to repressive effector domains can effectively repress
TRPM4, GRP78, and ZFX. By focusing on genes upregulated in PCa, I hope to demonstrate the
therapeutic potential of the dCas9 technology, here referred to as a toggle switch in reference to
its ability to induce reversible modulations in gene expression.
Past studies detailing the ability of dCas9-EDs to reduce gene repression have largely focused on
short term experiments (O'Geen et al. 2017, Gilbert et al. 2013, Jinek et al. 2012, Larson et al.
2013) or on long term experiments using lentiviral constructs to stably introduce dCas9-EDs
(Amabile et al. 2016, Parsi et al. 2017, Gilbert et al. 2013). While both techniques may reveal the
repressive capacity of the dCas9 system, neither explores whether the cell is able to persistently
maintain the introduced epigenetic modifications in the absence of continued expression of the
toggle switch. To evolve into a clinically relevant treatment, the toggle switch must be able to
deposit marks that are both effective in repressing gene expression and persistent through
multiple rounds of replication. Thus, for modifications to be placed and to persist, the chromatin
environment of the cell must be both conducive to dCas9-binding at the target site and capable of
preserving the introduced modifications. The latter may entail the availability of cellular
enzymes required for sustaining and copying the marks over multiple cell divisions.
Transient repression achieved with various dCas9-ED fusions in previous reports has resulted
from the combined effects of CRISPR interference (CRISPRi) and epigenetic remodeling (Zheng
et al. 2018, Larson et al. 2013, Gilbert et al. 2013). CRISPRi can be defined as a targeted
silencing method that relies on steric hindrance of the RNA polymerase during transcription
(Larson et al. 2013). While capable of achieving rapid gene repression similar to that of RNA
interference, CRISPRi can only transiently impact transcription; as soon as the dCas9 protein is
diluted via replication, the gene expression will return to control levels. Achieving lasting
repression will therefore require an understanding of how CRISPRi versus epigenetic editing
individually contribute to dCas9-ED generated transient repression. This knowledge may allow
the development of more efficient dCas9-ED fusions while exploring how different epigenetic
modifications work in conjunction to regulate gene expression.
10
1.5 Depositing different epigenetic modifications may elicit variable transcriptional outputs
depending on the local chromatin environment.
I hypothesize that achieving persistent repression will require imitating the dynamic interplay
between histone modifications and DNA methylation that takes place in a normally functioning
cell. Consequently, I will assay whether histone modifiers or DNA methyltransferases alone are
sufficient to induce persistent repression, or whether they must be co-targeted to deposit self-
sustaining modifications. It is possible that the effectiveness of each mark in generating
repression may be influenced by the chromatin environments of the regions that are targeted. For
this reason, I will compare repression achieved with a variety of dCas9-ED combinations,
including four histone modifiers and two DNA methyltransferases, at 3 different promoters.
dCas9-KRAB has been demonstrated to initiate potent repression of target genes (O'Geen et al.
2017) through recruitment of endogenous chromatin modifying complexes that both deposit
repressive H3K9me3 marks and deacetylate histones and also deposit DNA methylation (Figure
1.2). Recently, the fusion of KRAB to MeCP2, a protein containing a methyl-CpG binding
domain, has produced dCas9-KRAB-MeCP2 (Yeo et al. 2018). This fusion has been cited as an
enhanced version of dCas9-KRAB because of its ability to bind methylated CpGs, therefore
augmenting KRAB-mediated chromatin remodeling. Additionally, MeCP2 has been known to
interact with SIN3A to recruit histone deacetylases that facilitate transcriptional repression
(Jones PL et al., 1998; Nan X et al., 1998). Therefore, I will compare the transient and persistent
repression achieved when KRAB plus or minus MeCP2 is targeted to the TRPM4, GRP78 and
ZFX promoters via C-terminal dCas9 fusions (Figure 1.3).
11
Figure 1.2. Proposed mechanism of CRISPR-mediated epigenetic repression via modulating
H3K9me3 and DNA methylation levels. The dCas9-based toggle switch may enact repression by
taking advantage of the chromatin remodeling complexes present in the cell. A dCas9 fused to a
KRAB domain from a KRAB-ZNF TF can be directed to a target site via a gRNA. Once bound
to DNA, the KRAB domain can recruit endogenous chromatin remodeling complexes that
maintain and further spread the deposited modification.
I will also explore the ability of H3K27me3 writers to induce transient and persistent
transcriptional repression. Recently, dCas9 fusions to Enhancer of Zeste Homolog 2 (EZH2) and
Friend of GATA-1 (FOG1) have been shown to induce potent transient repression of oncogenic
promoters (O’Geen H et al., 2017). EZH2 contains a SET-domain that is responsible for the
methyltransferase activity of the Polycomb repressive complex 2 (PCR2). Only fusions
containing the full-length EZH2 (EZH2-FL) were capable of depositing H3K27me3 marks.
While dCas9 fusions to the isolated SET domain were not able to modify the chromatin, they
resulted in levels of transient repression similar to those attained with EZH2-FL. In this case,
assaying for persistent repression may answer the question of whether H3K27me3 marks can
contribute to transcriptional repression independent of steric hindrance. Therefore, I will
compare the transient and persistent repression achieved when EZH2-FL is targeted to the
TRPM4, GRP78 and ZFX promoters via an N-terminal dCas9 fusion (Figure 1.3).
Figure adapted from Fig. 2 of Ecco et al., 2017
12
Figure 1.3. dCas9-ED fusions. The indicated effector domains were fused to dCas9 at the N-, C-
or N- and C- termini. Each fusion contains nuclear localization sequences (NLS) at both termini,
a 3Xflag epitope tag and two 15-aa linker sequences (GGS)5.
O’Geen et al., 2017 have demonstrated that fusions to the N- and C- terminal residues of FOG1,
a transcription factor associated with deacetylation and H3K27me3 activity, produced robust
transient repression that was dependent on the deposition of H3K27me3. Despite laying down
the same marks, the repressive capabilities of the dCas9-EZH2 and FOG1 fusions differ in how
they induce chromatin remodeling. EZH2 is tethered directly to dCas9, allowing it to directly
modify the DNA via its intrinsic catalytic activity (Figure 1.4). FOG1 instead recruits
endogenous modifying complexes in a manner that better resembles the strategies used by
natural transcription factors (Figure 1.5).
13
Figure 1.4. Proposed mechanism of EZH2-mediated transcriptional repression. Step 1: EZH2
may recruit the PRC2 remodeling complex to the target site, where associated HDACs may
remove activating marks. Step 2: dCas9-bound EZH2 may then directly tri-methylate H3K27.
Step 3: DNMTs recruited by the PRC2 complex may sustain this repression by methylating
nearby CpGs.
Figure 1.5. Proposed mechanism of FOG1-mediated transcriptional repression. Step 1: dCas9-
bound FOG1 may recruit the PRC2 and NuRD complexes which (Step 2) may remove local
activating marks and deposit H3K27me3 marks. Step 3: DNA methylation deposited by DNMTs
recruited by PRC2 and NuRD may further this chromatin silencing.
In addition to assaying the repression achieved with the individual histone modifiers, I will test
whether the addition of dCas9-DNMT3a plus DNMT3L influences persistent chromatin
silencing. DNMT3a is one of two de novo methyltransferases that methylate CpG sites to
regulate gene expression. Despite lacking catalytic activity, DNMT3L has been shown to
increase the rate and incidence of DNMT3a mediated methylation (Chedin F et al., 2002;
Amabile et al., 2016; Stepper et al., 2017). Targeted DNA methylation has been widely used to
14
epigenetically reprogram cancer cells (Rivenbark AG et al., 2012) and I suspect it will be
essential in persistently repressing the CpG island promoters of TRPM4, GRP78 and ZFX in a
prostate cancer cell line (Figure 1.6). I hypothesize that directing methylation to active,
hypomethylated regions will be essential in distinguishing the contribution of methylation to the
repression achieved with the histone modifiers alone. Although some of the dCas9-EDs may, in
theory, also recruit DNMTs, I expect that directly targeting DNMTs to the promoter may be a
more effective strategy than relying on secondary recruitment.
Figure 1.6. DNA methylation of promoter CpG islands in TRPM4, ZFX and GRP78. The
promoter CpG islands of each gene are hypomethylated in C42B cells, suggesting dCas9-
DMNT3a+3L mediated methylation may help to silence the nearby chromatin. LGALS3 has
been included as a hypermethylated promoter comparison.
In summary, I will use a variety of different toggle switches that will place several different
types of repressive epigenetic marks on three different promoters that drive genes implicated in
15
prostate tumorigenesis. The overall goal will be to determine which dCas9-ED combinations that
achieve the greatest amount of repression among the promoters tested. I will assay gene
expression at four, fourteen and thirty days to determine whether any transient repression I
achieve can be maintained by the cell.
16
Chapter 2. Materials and Methods
2.1 dCas9 expression plasmid constructs
Vectors containing dCas9 fused to KRAB, EZH2[FL], FOG1 or DNMT3a in the orientations
outlined in Figure 1.3 were obtained from Dr. Henriette O’Geen (Department of Pharmacology
and the Genome Center, UC Davis). dCas9-fusions were constructed by introducing effector
domain sequences upstream of a C-terminus 3X Flag tag in the dCas9 plasmid (Addgene
#41815, Figure 2.1A) using Gibson cloning (O'Geen et al. 2015, O'Geen et al. 2017). Dr.
O’Geen also provided a pcDNA-DNMTL expression plasmid, originally a gift from Dr. Fred
Chedin (Department of Molecular and Cellular Biology and the Genome Center, UC Davis), for
use in conjunction with dCas9-DNMT3a.
Figure 2.1. Expression vectors used to clone either effector domain (A) or gRNA (B) sequences.
The effectiveness of each modification may largely depend on the ability of the dCas9-ED
fusions to bind to the DNA. For this reason, I have considered the chromatin states of our
selected target regions in designing our assays. Specifically, I have used previous data marking
DNase hypersensitive sites (Consortium 2012, Thurman et al. 2012), active promoter H3K27ac
17
modifications and RNA expression levels to ensure our gRNAs are targeting accessible open
chromatin.
gRNAs were preferentially designed to target regions that did not already appear to be
hypermethylated. gRNA-encoding plasmids were constructed by cloning 20 bp target sequences
into AflII-digested gRNA cloning vector (Addgene #41824, Figure 2.1B) using Gibson
Assembly. Either three or four gRNA sequences were designed to promoter or enhancer regions
of open chromatin, as indicated by DHS tracks and H3K27me3 enrichment, in TRPM4 (Table
2.1, Figure 3.2A), GRP78 (Table 2.2, Figure 3.3A) and ZFX (Table 2.3, Figure 3.4A) using the
online tool CRISPOR.
Table 2.1. Sequences and locations of gRNAs targeting the TRPM4 promoter (TP).
gRNA Sequence Location relative to TSS
TP 1 GTCTCTGTCCCCCTCTCCGT -570 bp
TP 2 TAAATTGTCCCCTCCCTGTC -233 bp
TP 3 CGGGTCCCAGGCCGCGATAA -65 bp
TP 4 AGCAGGTGAGCGCCGGACCA +130 bp
Table 2.2. Sequences and locations of gRNAs targeting the GRP78 promoter (GP)
gRNA Sequence Location relative to TSS
GP 1 CTTGCCGTTCAAGGTTCGAC +693 bp
GP 2 CGGTGGTCGGCATCGACCTG +350 bp
GP 3 TAGCAGCCAATGAATCAGCT -136 bp
GP 4 CAGGGCCGTTCGTTGCTCAC -558 bp
18
Table 2.3 Sequences and locations of gRNAs targeting the ZFX promoter (ZP)
gRNA Sequence Location relative to TSS
ZP 1 GACTCACCGGACGGACGTGC +139 bp
ZP 2 TAGCGCGGGAGGGCGCCTTG +232 bp
ZP 3 GTTGTGCCGGAGGCGGTCGA +387 bp
ZP 4 ATAGCGAAAATCGGCCCGGC +514 bp
2.2 Cell culture and transfection
The human prostate cancer cell line C42B (ATCC #CRL-3315) was grown in RPM11640
medium supplemented with 10% fetal bovine serum and 1% penicillin/streptomycin. The human
embryonic kidney cell line HEK293T (ATCC #CRL-3216) was grow in Dulbecco’s Modified
Eagle Medium supplemented with 10% fetal bovine serum and 1% penicillin/streptomycin. Both
cell lines were maintained at 37°C and 5% CO2 and verified by the Bioreagent and Cell Culture
Core at the USC Norris Cancer Center.
Cells were transfected at 60-70% confluency using Lipofectamine 3000 (Thermo Fisher)
following the manufacturer’s instructions. Transfections were performed in 12-well plates using
100 ng pBABE-puro, 600 ng dCas9 expression vector and 300 ng of equimolar pooled gRNA
expression vectors. Cells were treated with 2 μg/ml puromycin at 24h post transfection and
harvested after four days. In experiments assaying persistent repression, 4d cells were washed
with 1X Dulbecco’s phosphate-buffered saline, trypsinized and passaged into a new 12-well
plate. Cells were continuously passaged at 70-80% confluency until 14 or 30d post-transfection,
at which they were harvested in the same manner as the 4d cells.
2.3 RNA extraction and reverse-transcription quantitative PCR (RT-qPCR)
At either 4, 14 or 30d, transfected cells were washed with 1XDPBS, trypsinized and pelleted at
430 RCF for five minutes. Total RNA was isolated using TRIzol according to the manufacturer’s
instructions. 500-800 ng RNA were reversed-transcribed using the iScript Reverse Transcription
Supermix for RT-qPCR (BioRad) according to the manufacturer’s instructions. RT-qPCR was
19
performed in triplicate with PerfeCTa SYBR Green Super Mix (VWR) using the BioRad CFX96
Real-Time system. TRPM4, ZFX and GRP78 expression levels were measured in reference to
GAPDH using two intron spanning primers each. qPCR primer sequences for all genes can be
found in Table 2.4.
Table 2.4 Sequences of qPCR primers for TRPM4, ZFX, GRP78 and GAPDH.
Gene Forward Reverse
TRPM4 GATGCACACACCACGGAGAA AGAGCCGGAGGAAATTGCTG
ZFX TGAGCTGTGCTTTACGCT CCCATCTTCATCCATGGC
GRP78 GAACGTCTGATTGGCGATGC TCAACCACCTTGAACGGCAA
GAPDH AATCCCATCACCATCTTCCA CTCCATGGTGGTGAAGACG
20
Chapter 3. Transient Repression of Promoters
3.1 Experimental workflow to study transient repression.
A dCas9-based epigenetic editor has the potential to reverse oncogenic gene expression changes.
In addition to the therapeutic possibilities of further developing this tool, however, we can use it
to better distinguish the relationship between epigenetic modifications and changes in gene
expression. In this chapter I will target different epigenetic toggle switch combinations to the
promoters of TRPM4, ZFX and GRP78 and compare the efficiencies of each combination in
transiently repressing each gene. Targeted regions of the promoter will exist in an open
chromatin conformation – as indicated by the DNA methylation and active histone modification
data available for these cell lines – and lie either upstream or downstream of the transcription
start site. Achievable transient repression may indicate both the potential effectiveness of
developing certain toggle switches as future cancer therapies, and the receptivity of different
chromatin environments to the introduction of different repressive modifications.
In line with the results of previous studies outlined in Chapter 1 (Gilbert et al. 2014, Kungulovski
et al. 2015), I hypothesize that transient repression may result from both induced chromatin
remodeling and CRISPRi. To better understand the individual contributions of each mechanism,
3 different promoters were targeted with an RNA-guided dCas9 without an effector domain. Any
disruption of the transcriptional machinery by dCas9 alone may indicate that steric hindrance and
the consequential blockage of RNA polymerase II contributes to transient repression.
Additionally, experimental samples were normalized to cells transfected with untargeted dCas9
in order to account for any gene expression changes introduced by the system alone. Figure 3.1
provides a full overview of the workflow I used to achieve transient repression.
Transient experiments were performed in human prostate cancer cell line C42B to assay the
ability of the epigenetic toggle system to repress genes specifically upregulated in prostate
cancer. C42B cells are advanced androgen independent prostate cancer cells and thus serve as a
promising system in which to target genes associated with advanced cancers. Additionally, these
experiments were performed in human embryonic kidney HEK293T cells to examine the extent
21
to which gRNA precision, transfection efficiency, and different chromatin environments
contribute to achievable repression.
Figure 3.1. Transient repression workflow. Cells are transfected with plasmids containing
puromycin resistance, gRNA or an effector domain. In this diagram, the effector domain may be
a histone modifier, DNA methyltransferase, or both, though in the latter case each ED would be
transfected on a separate plasmid. Selection for cells that have taken up antibiotic resistance
plasmids is carried out via 72 hours of puromycin treatment. Cells are harvested and gene
expression is analyzed via RT-qPCR at four days.
3.2 Transient repression of the TRPM4 promoter is achieved with all targeted effector
domains.
To determine the effectiveness of different histone modifications in transiently repressing
TRPM4, and whether this repression was influenced by exogenous DNMTs, dCas9-KRAB,
dCas9-KRAB-MeCP2, dCas9-EZH2 or dCas9-FOG1 was targeted to the promoter in the
presence or absence of dCas9-DNMT3a+DNMT3L. Each dCas9-ED was directed by a pool of
four gRNAs. gRNAs were designed to regions of open chromatin, as indicated by enrichment of
ENCODE-derived active transcription mark H2K27ac and LNCaP DHS tracks (Consortium
2012, Thurman et al. 2012). Because C42B DHS tracks were not available via ENCODE, I used
the respective data sets from LNCaP cells to predict optimal gRNA targets. C42B cells derive
from LNCaP cells, which are androgen-sensitive human prostate carcinoma cells that have
metastasized to a lymph node. Additionally, gRNAs were designed to hypomethylated regions of
promoter CpG islands following the hypothesis that hypermethylated areas would be less
22
receptive to further methylation. CpG islands were also predicted from LNCaP data. gRNA
positions relative to known epigenetic modifications are illustrated in Figure 3.2.
Figure 3.2. TRPM4 gRNA schematic. Four gRNAs were targeted to hypomethylated regions of
open chromatin in the TRPM4 promoter, as indicated by the respective tracks taken from the
UCSC genome browser. Tracks indicating methylation and H3K27ac marks were taken from
C42B data while CpG islands and DNase HS tracks were predicted from LNCaP datasets.
When targeted with dCas9-KRAB, TRPM4 mRNA levels were reduced to <10% of those
obtained with the untargeted dCas9 control in C42B cells; the addition of the DNMTs did not
improve the repression obtained with the histone modifier (Figure 3.3A). Similarly, targeting
dCas9-KRAB-MeCP2 to the promoter reduced TRPM4 expression to 20% of the starting
amount, and this was not heightened by the addition of DNMTs (Figure 3.3B). dCas9-EZH2
and dCas9-FOG1 also reduced mRNA levels to 20% regardless of the addition of DNMTs
(Figures 3.3C and 3.3D).
It is important to note that TRPM4 mRNA levels were reduced to 40-60% with targeted dCas9
alone (i.e. a dCas9 having no added effector domain). This suggests that some of the repression
observed with the dCas9 fusion constructs may be due to steric hindrance rather than deposition
of the expected modifications. Clearly the effect of the CRISPRi captured in the dCas9 controls
is less dramatic than the repression achieved with the epigenetic modifiers, but whether this
repression is definitively the result of the introduced modifications will require further
verification. Experiments needed to validate this hypothesis are discussed below.
23
To determine whether the repression of TRPM4 achieved in C42B cells may be cell-type
specific, I performed toggle switch experiments in HEK293T cells. Specifically, I hypothesized
that differences in the chromatin environments of cancerous versus non-cancerous cells may
influence the degree to which certain chromatin modifications are able to induce chromatin
remodeling. For instance, the deposition of repressive modifications may be hindered by the
presence of activating marks or a diminished availability of enzymes necessary to cause or
maintain chromatin restructuring. Additionally, I considered that the rapid growth rates and
relatively high transfection efficiencies of HEK293T cells may increase my ability to repress
TRPM4 if such factors were limiting in C42B cells.
Comparison of TRPM4 repression achieved with dCas9-KRAB +/- DNMTs (Figure 3.3A and
3.3E) in C42B vs HEK293T cells demonstrates that neither technical concern had a substantial
impact on toggle switch efficiency. dCas9-KRAB is able to repress TRPM4 to ~20% in both
C42B and HEK293T cells, with or without the addition of dCas9-DNMT3a+3L. As in C42B
cells (Figure 3.3C), dCas9-EZH2+DNMTs can repress TRPM4 to ~30% in HEK293T cells
(Figure 3.3F), but only upon the addition of DNMTs. Targeting the TRPM4 promoter with
dCas9-EZH2 alone achieves repression similar to that achieved with the dCas9 control,
suggesting that steric hindrance rather than deposition of epigenetic modifications may be
responsible.
24
Figure 3.3 Transient repression of TRPM4 can be achieved with all dCas9-ED constructs in
C42B (A-D) and HEK293T (E-F) cell lines. Shown is the TRPM4 mRNA expression (relative to
GAPDH) in cells co-transfected with a pool of four gRNAs targeted to the TRPM4 promoter and
the indicated dCas9-ED fusions. The fusions have been abbreviated as follows: dCas9-K
indicates dCas9-KRAB, dCas9-K-M indicates dCas9-KRAB-MeCP2, dCas9-E indicates dCas9-
EZH2, dCas9-F indicates dCas9-FOG,1 and the addition of 3a,3L to any of these indicates co-
transfection with plasmids encoding dCas9-DNMT3a and DNMT3L.
3.3 Very modest transient repression can be achieved when the dCas9-system is targeted to
GRP78.
Encouraged by my ability to achieve transient repression of the TRPM4 promoter with all
dCas9-ED fusions, I next investigated whether the system could be applied to a gene that has
been lauded as a promising therapeutic target in many cancer types (Van Hoesen et al. 2017,
Araujo, Hebbar and Rangnekar 2018, Casas 2017) including PCa. Recent reports have shown
that GRP78 haploinsufficiency in Pdx1-Cre; KrasG
12D/+
; p53
f/+
mice significantly reduces
*P<0.01, **P<0.001
25
pancreatic tumor growth and suppresses AKT, S6, ERK and STAT3 activation (Shen et al.
2017). In line with these results, we explored whether our toggle system could be used to achieve
50% repression of GRP78 in PCa cells. To test this, the GRP78 promoter was targeted following
the same strategy as described above for TRPM4; a pool of four gRNAs designed to regions of
open chromatin that were not already hypermethylated (Figure 3.4), along with dCas9-KRAB,
dCas9-KRAB-MeCP2 and dCas9-EZH2, all +/- dCas9-DNMT3a+3L, were directed to the
GRP78 promoter.
Figure 3.4. GRP78 gRNA schematic. Four gRNAs were targeted to the GRP78 promoter, as
indicated by the UCSC genome browser tracks.
In C42B cells, targeting the dCas9 alone could not repress the GRP78 promoter and only very
modest repression could be achieved using the H3K9me3 depositors dCas9-KRAB and dCas9-
KRAB-MeCP2 (~80% remaining activity) that was not improved by the addition of the DNMTs
(Figure 3.5 A and B). Repression could also not be achieved with dCas9-EZH2 +/- DNMTs
(Figure 3.5C). To determine whether the lack of achievable repression with either induced
chromatin remodeling or CRISPRi was caused by inefficient gRNA binding (e.g. due to
mutations or SNPs in the GRP78 promoter in C42B cells), I tested seven different gRNA
sequences spanning 1.3kb of the promoter. I found no differences between the repression
achieved with different gRNA pools (data not shown). Because it is highly unlikely that there
were SNPs or single nucleotide mutations at the genomic locations of all 7 guide RNAs, I
concluded that the lack of repression was not due to the genomic sequences.
I next investigated whether the lack of repression of the GRP78 promoter was cell type specific
by repeating these experiments in HEK293T cells. Targeting dCas9-KRAB +/- DNMTs to the
26
GRP78 promoter in HEK293T cells (Figure 3.5D) achieved dramatically different results than
those observed in C42B cells (Figure 3.5A). I found that GRP78 could be transiently repressed
to nearly 30%, independent of the addition of dCas9-DNMT3a+3L, and that this repression was
significantly greater than that achieved with the targeted dCas9 control (Figure 3.5D). ~50%
repression was observed when dCas9-KRAB-MeCP2 + DNMTs were targeted to the promoter,
though this was not significantly different than the repression observed with targeted dCas9
alone (Figure 3.5E). The H3K27me3 depositor EZH2, however, was not able to repress GRP78
(Figure 3.5F). I have not yet tested the ability of the dCas9-FOG1 +/- DNMTs to repress
GRP78.
The apparent difference in the ability of the dCas9-ED toggle system to repress GRP78 in the
two cell lines suggests that achievable repression of some promoters may be cell-type specific. It
is not yet clear if the differences in repression of GRP78 in the two cell lines is influenced by
differences in the ability of the epigenetic modifiers to put down their marks. Further analysis, as
discussed below, will be required to validate whether the expected marks have been deposited in
either cell type.
27
Figure 3.5 Transient repression of GRP78 is ED- and cell type-specific. Shown is the GRP78
mRNA expression (relative to GAPDH) in C42B cells co-transfected with a pool of four gRNAs
targeting the GRP78 promoter and the indicated dCas9-ED fusions for C42B (A-C) and
HEK293T cells (D-F).
3.4 Transient repression of ZFX can be achieved with dCas9-KRAB +/- DNMTs.
Finally, I applied our dCas9-ED system to the ZFX promoter with the goal of better
understanding how different promoter regions and chromatin environments may influence
achievable repression. Like TRPM4 and GRP78, ZFX is upregulated in multiple cancers,
including PCa, in which ZFX knockdown decreases both cell proliferation and migration (Liu et
al. 2015). I investigated whether the toggle system could be used to effectively repress ZFX by
*P<0.01, **P<0.001
28
targeting dCas9-KRAB, dCas9-KRAB-MeCP2, dCas9-EZH2 and dCas9-FOG1, all +/- dCas9-
DNMT3a+3L to the ZFX promoter via four gRNAs (Figure 3.6) in C42B and HEK293T cells.
Figure 3.6. ZFX gRNA schematic. Four gRNAs were targeted to the ZFX promoter, as indicated
by the UCSC genome browser tracks.
I found that targeting the ZFX promoter with dCas9-KRAB +/- DNMTs resulted in ~40%
remaining promoter activity in C42B cells (Figure 3.7A) and in HEK293T (Figure 3.7E) cells.
Unexpectedly, the added CpG binding capabilities of MeCP2 did not enhance the transient
repression achieved with dCas9-KRAB in C42B cells (Figure 3.7B) but did slightly increase
repression when co-transfected with dCas9-DNMT3a+3L in HEK293T cells (Figure 3.7F).
Targeting dCas9-EZH2 +/- DNMTs could not repress ZFX in C42B cells (Figure 3.7C) but
modest repression was observed using dCas9-EZH2 +/- DNMTs in HEK293T cells (Figure
3.7G). The effects of targeting dCas9-FOG1 to the promoter also appear to be cell type specific:
very modest repression of ZFX is achieved with dCas9-FOG1 +/- DNMTs in C42B cells (Figure
3.7D), whereas the combination of dCas9-FOG and DNMTs is needed to substantially repress
the ZFX promoter in HEK293T cells (Figure 3.7H).
29
Figure 3.7 Transient repression of the ZFX promoter is influenced by cell type. Shown is the
ZFX mRNA expression (relative to GAPDH) in C42B cells (A-D) or HEK293T cells (E-H) co-
transfected with a pool of four gRNAs targeting the ZFX promoter and the indicated dCas9-ED
fusions.
3.5 Summary and Future Studies
Transiently transfecting both C42B and HEK293T cells with histone modifiers resulted in
repression that was gene-specific and mostly independent of the addition of DNMTs. Among the
genes tested, dCas9-KRAB was universally the best repressor, though similar levels of
repression could be achieved with dCas9-KRAB-MeCP2 and dCas9-EZH2 when GRP78 and
ZFX were targeted, respectively. I observed cell-type specific effects when targeting GRP78:
only a very modest reduction in GRP78 mRNA levels could be achieved in C42B cells while a
~50% reduction could be achieved in HEK293T cells.
*P<0.01, **P<0.001
30
To understand the factors responsible for the transient repression achieved at each promoter in
each cell line, the next step should be to determine if the expected modifications have been
deposited. Chromatin immunoprecipitation qPCR (ChIP-qPCR) assays can be used to determine
if the dCas9-ED proteins have successfully deposited their respective marks. Specifically, cells
transfected with dCas9-EZH2 and dCas9-FOG1 should be analyzed for H3K27me3 at the target
promoters using ChIP-qPCR. Similarly, cells transfected with dCas9-KRAB and dCas9-KRAB-
MeCP2 should be analyzed for H3K9me3 at the target promoters. To assay whether the addition
of the DNA methyltransferases DNMT3a and 3L have resulted in increased methylation at the
target sites, bisulfite conversion followed by sequencing could be performed.
Verification of the deposited modifications will aid in distinguishing the contributions of
CRISPRi versus chromatin remodeling to the levels of transient repression achieved. Further,
differences in efficient deposition of marks among promoters and cell lines may contribute to our
understanding of how the toggle system behaves in different chromatin environments. The
implications of variability in mark deposition among these environments will be explored further
in Chapter 5.1.
Distinguishing between the roles of CRISPRi and chromatin remodeling in transient repression
may not always be necessary, however. For instance, when short term repression is all that is
desired, the toggle switch method may serve as a valuable alternative to routine in vitro
techniques such as siRNA. Experiments relying on efficient gene knockdown may benefit from
demonstrating that knockdown can be achieved through the alteration of more than one pathway.
Still, for the toggle system to be considered as a potential PCa therapy, it must be able to induce
repression that can be maintained beyond four days.
31
Chapter 4. Persistent Repression of Promoters
4.1 Persistent repression may require the deposition of both histone modifications and
DNA methylation.
In line with assessing the therapeutic potential of the toggle technology, I next investigated
whether the transient repression I observed could persist through multiple cell passages. Such
persistence would require that the deposited marks be self-sustaining, or conducive to
maintenance by the cellular machinery. Comparison of maintainable repression induced by
different dCas9-ED combinations among the different promoters and cell lines may also reveal
capabilities and limitations of the toggle technology. For instance, persistent repression may be
influenced by the degree of transient repression achieved under specified conditions. If
observable transient repression is the culmination of CRISPRi and restructuring of the local
chromatin environment, gene expression may increase as the exogenous dCas9-EDs are removed
from the nuclear environment during cell passages. The loss of bound dCas9 may decrease
instances of CRISPRi, thus allowing us to answer the question of whether our epigenetic toggle
switch can induce substantial repression apart from steric hindrance.
Additionally, the ability to achieve persistent repression may contribute to our understanding of
how repressive modifications are maintained. Based on the proposed mechanisms discussed in
Chapter 1.1 and outlined in Figure 1.1, I hypothesize that both histone modifications and DNA
methylation will be necessary to induce maintainable gene repression. Previous studies have
reported that neither histone modifications nor DNA methylation alone can induce persistent
repression. (Kungulovski et al. 2015) demonstrated that targeted deposition of either H3K9me2/3
or DNA methylation could induce robust transient repression, but neither could be maintained
after the exogenous methyltransferases were removed. O’Geen et al. (2017) was able to
demonstrate persistent repression following transient lipofectamine transfection of dCas9-ED
fusions, but only when cells were co-transfected with dCas9-EZH2 and dCas9-DNMT3a+3L. In
these experiments, targeted histone modifiers such as dCas9-KRAB and dCas9-FOG1 achieved
substantial transient but not persistent repression.
32
I have analyzed persistent repression using histone modifiers +/- DNMTs of the TRPM4, GRP78
and ZFX promoters after fourteen days. This length of time should be sufficient for reduction of
the transfected dCas9-ED through cell passaging, and the observable repression may
consequently result from successfully deposited and maintained epigenetic modifications. As in
the transient assays, I co-transfect cells with a pool of three to four gRNAs and selected dCas9-
ED combinations. Cells are treated with puromycin for 72 hours to select for those that have
taken up the plasmids. 80% of cells are harvested after three days of puromycin selection for
transient assays. At this point, 20% of cells continue to be passaged until fourteen days, at which
all are harvested and assayed for persistent repression. A complete diagram of this workflow can
be found in Figure 4.1.
Figure 4.1 Persistent repression workflow. Cells are transfected with plasmids containing
puromycin resistance, sgRNAs, and/or a dCas9-ED at day zero. At four days, a portion of cells
are harvested, and transient gene repression analyzed via RT-qPCR. Persistent repression is
analyzed by passaging a portion of cells at four days and performing RT-qPCR at fourteen or
thirty days.
4.2 Persistent repression of TRPM4 is only achieved through the combined targeting of
histone modifiers and DNA methyltransferases.
Because I was able to achieve the highest degrees of transient repression when targeting the
TRPM4 promoter, I first used this promoter for persistence assays. dCas9-KRAB, dCas9-KRAB-
MeCP2, dCas9-EZH2 and dCas9-FOG1 were targeted to the TRPM4 promoter using the same
33
gRNA pool as in the transient assays. The ability of each histone modifier to induce persistent
repression was assayed individually and in the presence of dCas9-DNMT3a+3L. Cells were
harvested at four or fourteen days, allowing for the direct comparison of transient and persistent
repression (Figure 4.2).
In C42B cells, targeting dCas9-KRAB in conjunction with dCas9-DNMT3a+3L to the TRPM4
promoter resulted in nearly 60% repression at 14 days (Figure 4.2A). Additionally, I tested
whether dCas9-DNMT3a+3L alone was sufficient to induce persistent repression but found no
significant change in expression after fourteen days (Data not shown).
Repression at 14 days achieved with dCas9 alone was similar to that using dCas9-KRAB,
indicating that CRISPRi may still be contributing to the observable repression. However, the
degree of persistent repression was drastically reduced from that achieved transiently with
dCas9-KRAB +/- DNMTs. This discrepancy may result from the cell’s inability to maintain the
induced chromatin remodeling. To further explore the mechanisms responsible for the persistent
repression achieved in C42B cells, we repeated the dCas9-KRAB +/- DNMTs experiments in
HEK293T cells. Interestingly, nearly the same amount of persistent repression could be achieved
with dCas9-KRAB + DNMTs in HEK293T cells as in C42B cells (~60%), but none of the
transient repression obtained with dCas9 alone or dCas9-KRAB could be maintained (Figure
4.2E). These results emphasize the necessity for both DNA methylation and histone
modifications in creating lasting repression.
I next explored whether the persistent repression achieved with KRAB could be augmented by
the addition of MeCP2 in C42B cells. Following the hypothesis that spreading of silent
chromatin may be essential to maintaining a closed chromatin state, I targeted cells with dCas9-
KRAB-MeCP2 +/- DNMTs. When I did this, I discovered that the addition of MeCP2 did not
enhance achievable persistent repression (Figure 4.2B). Despite ~80% repression achieved with
dCas9-KRAB-MeCP2 +/- DNMTs after four days, only 20% repression could be persistently
maintained. No persistent repression was observed with either the targeted dCas9 control or the
dCas9-KRAB-MeCP2 alone, again suggesting a requirement for the combined deposition of
histone modifications and DNA methylation in attaining persistent repression.
34
Co-targeting dCas9-EZH2 and dCas9-DNMT3a+3L to the TRPM4 promoter resulted in ~60%
persistent repression, only a 20% increase in expression from the 80% repression attained in the
transient assays (Figure 4.2C). This effect was not seen when dCas9+EZH2 alone was targeted
to the TRPM4 promoter. Instead, expression levels returned to those achieved with the
untargeted dCas9 control. Whether or not dCas9-EZH2 may have activating properties in the
absence of DNMTs as seen in transient assays targeting the ZFX promoter will require further
exploration.
This activating effect of EZH2 was amplified in HEK293T cells, wherein cells treated with
dCas9-EZH2 alone expressed TRPM4 near two-fold times expression levels recorded in the
dCas9+EGV control (Figure 4.2F). When combined with DNMTs, however, the levels of
transient repression could be completely maintained. No persistent repression was observed in
the targeted dCas9 control.
The success of EZH2 in achieving persistent repression in both cell lines prompted us to explore
whether the other H3K27me3 depositor, dCas9-FOG1, could induce lasting repression. Despite
significant repression achieved in transient assays (~80% with dCas9-FOG1 +/- DNMTs),
expression levels returned to normal in all conditions after fourteen days (Figure 4.2D). To
determine whether the success of the transient assays derived solely from dCas9-induced steric
hindrance, further assays will be necessary to verify the deposition of the H3K27me3 marks.
35
Figure 4.2 Persistent repression of the TRPM4 promoter can be achieved with dCas9-KRAB +
DNMTs and dCas9-EZH2 + DNMTs. Shown is the TRPM4 mRNA expression level (relative to
GAPDH) in C42B (A-D) or HEK293T (E-F) cells co-transfected with a pool of four gRNAs
targeting the TRPM4 promoter and the indicated dCas9-ED combinations at 4d and 14d.
4.3 Achievable persistent repression of GRP78 is comparable to levels of transient
repression achieved with the same effector domains.
I next explored whether the transient repression achieved by targeting GRP78 in C42B cells
could persist beyond four days. I found that when dCas9-KRAB +/- DNMTs (Figure 4.3A),
dCas9-KRAB-MECP2 +/- DNMTS (Figure 4.3B) or dCas9-EZH2 +/- DNMTs (Figure 4.3C)
were targeted to the GRP78 promoter, only very modest levels of repression were observed at 14
days (~20%), similar to what I observed in the transient assays. Comparably, no difference in
four- and fourteen-day repression could be distinguished in samples treated with dCas9-EZH2
+/- DNMTs. The very modest transient repression achieved with dCas9-KRAB-MeCP2 +/-
DNMTs disappeared after four days.
*P<0.01, **P<0.001
36
Considering I was able to achieve nearly 70% transient repression of GRP78 in HEK293T cells
at 4 days, I next explored whether this repression could be maintained. However, very little
persistent repression was observed in any combination of toggle switches at 14 days in
HEK293T cells. When targeted with dCas9-KRAB, the robust transient repression faded to 30%
in dCas9-KRAB and 10% in dCas9-KRAB + DNMTs (Figure 4.3D). Similarly, the transient
repression achieved with dCas9-KRAB-MeCP2 +/- DNMTs is not efficiently maintained after
fourteen days (Figure 4.3E). When targeted with dCas9-EZH2 +/- DNMTs, the modest levels of
transient repression achieved were not maintained (Figure 4.3F). As is the case in TRPM4,
targeting the GRP78 promoter with dCas9-DNMT3a+3L alone did not result in persistent
repression.
37
Figure 4.3. Persistent repression is not achieved with any dCas9-ED in either cell type. Shown is
the GRP78 mRNA expression level (relative to GAPDH) in C42B (A-C) or HEK293T (D-F)
cells co-transfected with a pool of four gRNAs targeting the GRP78 promoter and the indicated
dCas9-ED combinations at 4d and 14d.
4.4 Transient repression of ZFX is not maintained regardless of the effector domains
targeted.
Despite achieving ~50% transient repression by targeting dCas9-KRAB +/- DNMTs to the ZFX
promoter, only a very modest percentage of this repression could persist to fourteen days in
C42B cells (Figure 4.4A) or in HEK293T cells (Figure 4.4D). Only very modest persistent
repression could be achieved in either cell line with any of the tested dCas9-EDs. Targeting
*P<0.01, **P<0.001
38
dCas9-EZH2 +/- DNMTs achieved the strongest of this repression in both C42B and HEK293T
cells, though ~80% activity still remained.
Figure 4.4 Significant 14d persistent repression of ZFX cannot be maintained with any of the
dCas9-ED fusions in either cell line. (A-F) Relative ZFX mRNA expression in either C42B or
HEK293T cells co-transfected with a pool of three gRNAs targeting the ZFX promoter and the
indicated dCas9-ED combinations at 4d and 14d.
4.5. dCas9-mediated repression of TRPM4 can be maintained for at least thirty days.
Among the different conditions tested, the dCas9-ED system appears to induce both gene and
cell-type specific effects. These effects are especially apparent in the persistence assays, where
*P<0.01, **P<0.001
39
repression may be less influenced by steric hindrance than the ability of a unique chromatin
environment to spread and maintain the deposited marks. To further characterize the factors
responsible for robust persistent repression, I next investigated whether the marks could be
maintained for even longer. To explore this, I assayed whether the repression achieved with the
dCas9-KRAB plus DNMTs combination targeting TRPM4 in C42B cells could be maintained
for thirty days. Importantly, the persistent repression recorded at fourteen days with dCas9-
KRAB + DNMTs was almost completely maintained while that recorded in cells targeted with
dCas9 or dCas9-KRAB returned to normal (Figure 4.5). These results demonstrate that
persistent repression can be achieved through transient deposition of epigenetic modifications,
but only when both histone modifications and DNA methylation are introduced.
Figure 4.5. 30-day persistent repression of TRPM4 can be achieved with dCas9-KRAB +
DNMTs in C42B cells. Shown is TRPM4 expression at 4, 14 and 30d (relative to GAPDH) after
cells were co-transfected with a pool of four gRNAs targeting the TRPM4 promoter and the
indicated effector domains.
It remained possible, however, that the persistent repression achieved at 14 and 30d was the
result of expression vectors that had integrated into the genome and continued to deposit marks.
To ensure that this was not the case, I assayed levels of the dCas9-KRAB plasmid in four,
*P<0.01, **P<0.001
40
fourteen and thirty-day C42B cells in which TRPM4 was targeted with dCas9-KRAB and
dCas9-DNMT3a+3L. I performed qPCR on cDNA collected from the thirty-day experiment
depicted in Figure 4.5.
Figure 4.6 demonstrates that the expression of the dCas9-KRAB vector significantly decreased
from four to fourteen days and no expression of the plasmid could be detected in the thirty-day
cDNA samples. I further validated these results by running the qPCR products on a 1.5% agarose
gel (Figure 4.6C). The intensity of the four, fourteen- and thirty-day bands reflect the
information yielded by the qPCR data: dCas9-KRAB expression is significantly reduced by
fourteen days and nearly undetectable in the thirty-day samples.
Figure 4.6. The dCas9-KRAB expression vector is no longer detectable by 14d. C42B cells
targeted with dCas9+EGV or dCas9-KRAB+DNMTs were harvested at 4d, 14d and 30d and RT-
qPCR was performed to assay dCas9-KRAB vector expression (relative to GAPDH). (A)
Expression of dCas9-KRAB at 4d and 14d. (B) Expression of dCas9-KRAB at 14d. (C) qPCR
products from the 4d, 14d and 30d samples were further analyzed on a 1.5% agarose gel.
4.6 Summary and Future Studies.
I have found that repression can be achieved after 30 days at the TRPM4 promoter in C42B cells.
The next important step is to determine if the expected epigenetic marks have been deposited by
the respective dCas9-ED combinations. As previously described, ChIP-qPCR can be used to
assay whether the appropriate histone modifications have been deposited in the thirty-day
samples. Detecting methylation deposited by dCas9-DNMT3a+3L will require bisulfite
41
sequencing. If the expected epigenetic marks are present, it will be of interest to determine how
far they have spread from the region targeted by the guide RNAs.
I note that it is possible that instead of the repression observed at fourteen and thirty days being
due to the maintenance of epigenetic marks, it could result from marks being continuously
deposited by dCas9-ED complexes that have integrated into the genome. Notably, for integrated
plasmids to have a substantial effect on observable repression, both the gRNA and effector
domain plasmids would have to integrate and maintain expression for 30 days of culture in the
absence of a selection agent. Another, perhaps remote, possibility is that there is some residual
dCas9-ED and guide RNA plasmids in the media that manages to get into the cells over time
(although this possibility seems very remote because the media is changed with each passage).
While these results strongly suggest that the expression of exogenous dCas9-KRAB is not
responsible for the majority of persistent repression achieved, a Western Blot could be
performed. However, the sensitivity of the Western blot is much lower than that of RT-PCR and
thus I do not expect that any protein could be detected.
Further, it should be noted that the 50% repression I see at fourteen and thirty days represents an
average of the repression achieved among individual populations. Dissecting the contribution of
multiple populations to the overall repression observed may indicate the effectiveness of the
toggle switch in generating stably maintained epigenetic loci. For instance, is robust repression
achieved in a small percentage of cells lowering the average expression levels, or is TRPM4
reduced by ~50% in a large percentage of cells? To determine this, it may be of use to perform
flow cytometry on transfected cells at each recorded timepoint. FACS analysis may be
performed using a PE-conjugated TRPM4 antibody to detect whether a shift in TRPM4
expression occurs between four and thirty days. Additionally, because I will be measuring
protein levels, this sensitive technique will allow me to verify whether the changes in gene
expression translate to changes in the amount of protein produced.
42
Chapter 5. Discussion and Future Studies
5.1 Achieving significant transient or persistent repression of a target gene relies on the
optimization of the dCas9 toggle system to individual chromatin environments.
Targeting epigenetic toggle switches to tumor-promoting genes may provide a new inroad to
developing a PCa therapy. In line with this hypothesis, I targeted multiple combinations of
epigenetic editors to the promoters of three genes that are upregulated in PCa and assayed the
degree to which each combination could enact transient or persistent repression. Among the
different conditions tested, I found that both short- and long-term repression were highly
dependent on the target promoter, the dCas9-ED combination, and the cell type. These results
strongly suggest that the epigenetic toggle system must be optimized to the unique chromatin
environment of the target gene in order to achieve robust persistent repression. Among
everything I tested, the greatest persistent repression I achieved was by co-targeting dCas9-
KRAB and dCas9-DNMT3a+3L to the TRPM4 promoter in C42B cells. Alternatively, only very
modest fourteen-day repression could be attained when ZFX was targeted with the same dCas9-
ED fusions, and none of the transient repression achieved in GRP78 could be maintained.
It is possible that the differences I observed could be due to inherent cell type-specific
differences in depositing or maintaining the deposited marks at the specific promoter regions.
This may in turn reflect differences in the chromatin architecture of each promoter in each cell
type. It should be noted that the marks will be deposited at positions near where the guide RNAs
bind. Previous studies employing dCas9-KRAB mediated repression have noted the importance
of well-designed gRNAs in attaining robust repression (Zheng et al. 2018, Lo and Qi 2017).
Importantly, I did ensure that the gRNAs I used were strategically designed to target regions of
hypomethylated open chromatin in the core promoter region. However, I did not assay the
gRNAs for their individual abilities to achieve repression but instead used them in pools of three
to four. It is possible that this approach of targeting dCas9-ED fusions via multiple gRNAs may
have resulted in low affinity guides having a negative impact on high affinity guide RNAs in the
same pool. I also note that the gRNAs were designed based on sequences available on the UCSC
genome browser and not on the genomic sequences of the specific cell line I was using. Though
43
this design strategy did not account for somatic mutations or SNPs that may have occurred in the
gene promoters of our cell lines, using a pool of gRNAs should have increased the likelihood of
successful binding.
It may be important to achieve a higher persistent repression if the toggle switch method is to be
used as a therapy. One way to approach this would be to combine epigenetic silencing of
promoters with the additional targeting of other regions in the transcribed gene. As discussed
above, some of the repression I observed could have been due to steric hindrance of the dCas9-
ED (i.e. due to CRISPRi). Perhaps one could try to exploit CRISPRi by combining the targeted
epigenetic modification of the promoter with additional guide RNAs within the transcribed
region. Additional guide RNAs could be made to regions of open chromatin throughout the gene
to try to block the RNA polymerase II that manages to transcribe from the epigenetically
modified promoter.
In line with this hypothesis, I targeted gRNAs to a total of four intronic regions within TRPM4
and compared the amount of repression I achieved to that obtained with the promoter alone.
Targeted regions were located 4kb, 8kb, 20kb and 43kb downstream of the promoter. Of these,
only the 8kb and 43kb sites were in regions of open chromatin as marked by H3K27Ac.
Targeting the 8kb region with dCas9-KRAB resulted in an 80% reduction in TRPM4 expression
after four days, and 20% of this persisted to fourteen days. Only modest transient repression
could be achieved by targeting the other three regions. Co-targeting the 8kb region with the
promoter, however, was not able to repress TRPM4 as efficiently as targeting the promoter
alone. To fully test the potency of this application, in the future it may be of use to target more
intronic regions in different genes, or individually assay the affinity of each gRNA to ensure high
affinity promoter gRNAs are not being diluted out by low affinity, less potent intronic gRNAs.
Additionally, it may be necessary to consider the normal expression levels of the target gene in
the cell line being tested. The differences in persistent repression achieved in TRPM4, GRP78
and ZFX strongly suggest that the nature of the target gene influences the impact the toggle
system has on transcription. Even the degree of CRISPRi is highly specific to the target gene
(Gilbert et al. 2013). It may be that highly expressed genes are more difficult to repress. In line
44
with this, I compared the expression levels of the three genes. Normal expression levels in C42B
and HEK293T cells correlated with the persistent repression achieved in each cell line. GRP78
was expressed at nearly 9-fold higher levels than TRPM4 in C42B cells, which may explain the
substantial differences in repression achieved among the two genes. The importance of the gene
to the cell may also influence achievable repression. GRP78 is a housekeeping gene and
reducing its expression may affect the viability of the cell. Consequently, it may be that I have
selected for cells in which the dCas9-mediated knockdown was unsuccessful, or there may be
additional pathways that revive normal expression levels despite successful dCas9-ED
repression. While this may present one explanation for modest GRP78 repression, it is important
to note that selection may not occur by the four-day timepoint. To fully test this hypothesis, I
would thus need to perform a shorter experiment assaying for repression. Collecting a two-day
timepoint, for instance, may indicate whether repression was occurring early enough for me to
observe selection effects at four days.
Similarly, the activating capacity of ZFX may be essential to cell viability. Previous reports have
identified ZNF711 and ZFY as transcription factors that are functionally redundant to ZFX.
These findings, combined with the knowledge that ZFX is highly conserved throughout
evolution, suggests a crucial role for the transcriptional activators in gene regulation. In C42B
cells, however, ZFY is not expressed and ZNF711 is only expressed at very low levels. Thus,
repressing ZFX may lead to non-viable cells, and the repression I observe would reflect only the
mRNA levels of cells expressing ZFX at or near normal levels. Though I was able to reduce ZFX
expression >50% with dCas9-KRAB + DNMTs in C42B cells, this repression cannot be
maintained. Perhaps cells with reduced ZFX levels are non-viable or grow at slower rates than
those with normal ZFX expression, resulting in transient repression that is selected against by
day fourteen. To explore whether the absence of redundant transcription factors affected my
ability to repress ZFX, I performed this experiment in HEK293T cells, which express
functionally redundant ZNF711 at higher levels than C42B. The results I obtained with both cell
lines, however, were nearly identical. Alternatively, it may be that cells can salvage ZFX
expression through additional pathways over the course of fourteen days. As with GRP78,
however, it may be necessary to perform a two-day experiment to determine whether the effects
of selection could be observed in a four-day timepoint.
45
In summary, the ability of the dCas9 toggle system to enact persistent repression appears to be
highly dependent on the unique chromatin architecture of the target site. This, in turn is
influenced by the gene, region and cell type targeted.
5.2 More robust persistent repression may be achieved through increased transfection
efficiency.
The amount of persistent repression I can achieve appears to be influenced by the degree of
transient repression I initially observe. Perhaps this transient repression reflects the frequency of
successful dCas9 binding, which may be influenced both by the chromatin structure of the target
site and the ability of the plasmids to successfully enter the cell and replicate. As discussed
above, optimizing the dCas9 toggle system to the chromatin architecture of the target region may
increase the amount of persistent repression I can achieve. The latter factor, however, may be
improved by increasing transfection efficiency. dCas9-ED-mediated repression appears to be
cell-type specific, as best exemplified by the increased transient repression observed in
HEK293T versus C42B cells when GRP78 or ZFX is targeted with dCas9-KRAB +/- DNMTs.
HEK293T cells are widely lauded for being highly transfectable (Thomas P and Smart TG, 2005;
Ooi et al 2016), and differences in observable transient repression may thus reflect a difference
in the number of plasmids that can enter the cells.
The dCas9-ED, gRNA and puromycin resistance sequences were introduced into cells on
separate plasmids via cationic-lipid mediated transfections. Importantly, for the dCas9-ED
system to have its intended effects on gene expression, each of these plasmids must enter the cell
during transfection. Because the puromycin resistance gene is introduced via its own plasmid,
there may be instances where I select for cells that are expressing the antibiotic resistance
plasmid, but none of those required for the toggle machinery. However, the significant
differences in transient repression of the target genes between the dCas9+EGV control and the
dCas9-ED combinations suggest that the introduction of multiple plasmids is not a major
impediment to achieving repression. In future studies, however, it may be worth considering
whether introduction of the necessary sequences on fewer plasmids, each with its own selection
46
marker, may select against cells that haven’t taken up a fully intact, functional system.
Alternatively, each plasmid may be fused to a fluorophore that would allow for isolation of cells
that have taken up all plasmids via FACS sorting. Cells may either be sorted before the analyses
of each timepoint, or at four days, after which only cells with low expression of the target gene
are passaged and analyzed.
In addition to a cationic-lipid mediated transfection, electroporation may be applied to increase
the number of plasmids that enter the cell. This may be particularly beneficial in C42B cells,
where transfection efficiency may have been a limiting factor in the degree of transient
repression I could achieve. Similarly, performing multiple sequential transfections may increase
the amount of dCas9 machinery that enters the cell. For instance, following the initial
transfection, cells may be transfected a second time once enough time has passed to allow them
to recover and take up the initial dose of plasmids. If effective in repressing the target gene
beyond that achieved with a single transfection, this may be performed multiple times.
5.3 Further analysis of persistent repression achieved via the dCas9 toggle system in
TRPM4.
Evaluation of TRPM4 expression at thirty days demonstrated that the degree of repression
achieved at fourteen days could be maintained for longer. It remains unclear, however, what this
repression represents: is TRPM4 robustly repressed in a few populations while the majority of
cells show only modest reduction in TRPM4 levels, or is TRPM4 universally repressed by
~50%? Answering this question may provide insight into how the toggle switch initiates
repression and the degree to which cells can maintain it. Additionally, it may allow me to
determine the extent to which selection for cells with modest TRPM4 repression contributes to
the persistent repression I observe. For instance, if selection has a substantial influence on the
persistent repression I achieve, I may see reductions in the number of populations with robust
TRPM4 expression by fourteen or thirty days.
As discussed above, analysis of TRPM4 levels in individual cell populations may be
accomplished via flow cytometry. Previous experiments by collaborators Dr. Henriette O’Geen
47
and Dr. David Segal (Department of Biochemistry and Molecular Medicine, UC Davis) used this
method to analyze the expression of HER2 following dCas9-EZH2 +/- DNMTs mediated
knockdown at fifty-four days in HCT116 cells. They found that cells were separated into two
distinct populations when compared to the dCas9 and unstained controls: 31% of cells showed
robust repression of HER2 and the remaining cells showed modest to no repression of HER2.
Rather than observe a universal shift in HER2 expression, which might indicate a uniform loss of
repression over time in all cells, they observed that certain cell populations could completely
maintain robust levels of repression. Thus, I may expect to find that TRPM4 levels are robustly
reduced in only a fraction of cells, but these cells can completely maintain their silenced
chromatin states through multiple passages.
Further characterization of the cells’ ability to maintain this chromatin state in different
conditions, however, may improve the practicality of developing the toggle switch into an
eventual therapy. For instance, whether cells that show persistent repression are able to maintain
this repression after being frozen is yet to be explored. In addition to improving the convenience
of the toggle switch assay in routine biological assays and therapy development, freezing and
thawing the cells may indicate whether the new chromatin state can persist through periods of
intense cell stress. Again, consideration of gene target and cell model would be necessary in
evaluating cells’ ability to maintain repression throughout this process. GRP78’s role as the
master regulator of ER stress, for instance, may be upregulated as part of pathways that become
activated in response to acute cell stress.
Additionally, a toggle switch-based therapy may hold more promise if cells can be re-repressed
following an initial transfection. Among the conditions I tested, the general trend appears to be a
partial loss of the repression I observe at 4d by 14d. If this transient repression can be restored
through additional transfections, a future gene therapy may be able to incorporate fewer, more
potent doses. This feature may increase the cost efficiency of the treatment and make routine
delivery more feasible. To test this, future investigations may incorporate multiple, sequential
transfections. Following an initial transfection, additional transfections may be performed at 2d
and 4d, after which gene expression may be assayed and compared to that achieved with a single
48
transfection. If increased persistent repression can be achieved via this method, the toggle switch
may translate to an effective, practical gene therapy.
5.4 Introduction of toggle switches into patient tumors.
Ultimately, the potential of the epigenetic toggle switch as a gene therapy will depend on an
effective and safe delivery method. I have shown that the dCas9-ED system can persistently
silence an oncogene, TRPM4, in a PCa cell line by 50%. Whether this result can be applied in
different systems will rely on future optimization of the toggle switch to the unique chromatin
environment of the target. Additionally, future analysis of repression achieved with sequential
transfections may be important to determine whether the toggle switch can persistently reduce
mRNA levels beyond 50%.
To be developed as a gene therapy, however, these in vitro optimizations of the toggle system
must be translatable to patient tumors. Consequently, the method of delivery will highly impact
the functionality and specificity of the toggle switch in vivo. Though practical for developing and
characterizing a toggle system in laboratory assays, plasmid-mediated delivery may be
suboptimal for use in a clinical setting. Previous reports have shown that plasmid-based delivery
is accompanied by a myriad of complications, such as potentially dangerous immune responses
and unwanted integration of the plasmid into the host genome (Hemmi et al. 2000, Wagner 2001,
Ramakrishna et al. 2014). A plasmid-based approach would also require the introduction of a
bulky array of transfection tools that may complicate the delivery process. Fortunately, several
other techniques present promising alternatives to this method and may result in efficient
delivery of the toggle system into patient tumors.
Cell penetrating peptide (CPP)-mediated delivery of Cas9 and gRNA may pose an efficient and
safe alternative to other current delivery systems. Ramakrishna et al. 2014 developed CPP-
conjugated Cas9 and positively charged CPP-conjugated sgRNA complexes that can efficiently
edit human cells. They show that treatment of HEK293T cells with both complexes leads to
specific mutations in the target sequences without the need for additional delivery tools or drastic
off-target effects relative to plasmid-mediated delivery. Unlike ZNFs and TALENs, this system
49
does not rely on laborious de novo protein synthesis for each therapeutic application. CPP-
mediated delivery of the toggle switch may thus present an encouraging means of in vivo
epigenome editing.
An additional approach to delivering the toggle system may involve exosome-liposome hybrid
nanoparticles. Exosomes are nanoscale membrane vesicles that are naturally secreted by a wide
variety of cells and can pass through stringent biological barriers (Ibrahim and Marbán 2016,
Alvarez-Erviti et al. 2011). They have been shown to stably carry drugs through circulation and
cross vascular epithelium in order to deliver them to target cells (van den Boorn et al. 2011).
Though exosomes are too small to encapsulate the CRISPR/Cas9 system on their own, Lin et al.
2018 has developed hybrid-exosomes capable of carrying and delivering the bulky system to
target sites. These hybrids consist of exosomes that have fused with liposomes, enabling them to
contain large plasmid DNA such as that encoding the dCas9-based toggle system.
Both methods present promising delivery systems that may circumvent the potentially dangerous
immune reactions associated with adeno-associated viral (AAV) delivery. Though multiple
studies (Nelson et al. 2019, Colella, Ronzitti and Mingozzi 2018) have demonstrated AAV-based
delivery as a highly efficient method of introducing CRISPR/Cas9 into cells, the immune
responses associated with such vectors remain a substantial concern (Lin et al. 2018).
Following optimization of the toggle switch to each target gene, either CPP-based or liposome-
exosome hybrid delivery may be used to deliver the toggle system into patient cells. Through
early treatment of patient tumors with dCas9-ED fusions targeted to genes upregulated in PCa,
the toggle system may be a promising candidate for future epigenome-based therapies.
50
References
Alvarez-Erviti, L., Y. Seow, H. Yin, C. Betts, S. Lakhal & M. J. Wood (2011) Delivery of
siRNA to the mouse brain by systemic injection of targeted exosomes. Nat Biotechnol,
29, 341-5.
Amabile, A., A. Migliara, P. Capasso, M. Biffi, D. Cittaro, L. Naldini & A. Lombardo (2016)
Inheritable Silencing of Endogenous Genes by Hit-and-Run Targeted Epigenetic Editing.
Cell, 167, 219-232.e14.
Aran, D. & A. Hellman (2013) DNA methylation of transcriptional enhancers and cancer
predisposition. Cell, 154, 11-3.
Araujo, N., N. Hebbar & V. M. Rangnekar (2018) GRP78 Is a Targetable Receptor on Cancer
and Stromal Cells. EBioMedicine, 33, 2-3.
Arechederra, M., F. Daian, A. Yim, S. K. Bazai, S. Richelme, R. Dono, A. J. Saurin, B. H.
Habermann & F. Maina (2018) Hypermethylation of gene body CpG islands predicts
high dosage of functional oncogenes in liver cancer. Nat Commun, 9, 3164.
Ashktorab, H. & H. Brim (2014) DNA Methylation and Colorectal Cancer. Curr Colorectal
Cancer Rep, 10, 425-430.
Ayyanathan, K., M. S. Lechner, P. Bell, G. G. Maul, D. C. Schultz, Y. Yamada, K. Tanaka, K.
Torigoe & F. J. Rauscher (2003) Regulated recruitment of HP1 to a euchromatic gene
induces mitotically heritable, epigenetic gene silencing: a mammalian cell culture model
of gene variegation. Genes Dev, 17, 1855-69.
Bannister, A. J. & T. Kouzarides (1996) The CBP co-activator is a histone acetyltransferase.
Nature, 384, 641-3.
Belancio, V. P., P. L. Deininger & A. M. Roy-Engel (2009) LINE dancing in the human genome:
transposable elements and disease. Genome Med, 1, 97.
Berdasco, M. & M. Esteller (2010) Aberrant epigenetic landscape in cancer: how cellular
identity goes awry. Dev Cell, 19, 698-711.
Bibikova, M., E. Chudin, B. Wu, L. Zhou, E. W. Garcia, Y. Liu, S. Shin, T. W. Plaia, J. M.
Auerbach, D. E. Arking, R. Gonzalez, J. Crook, B. Davidson, T. C. Schulz, A. Robins, A.
Khanna, P. Sartipy, J. Hyllner, P. Vanguri, S. Savant-Bhonsale, A. K. Smith, A.
Chakravarti, A. Maitra, M. Rao, D. L. Barker, J. F. Loring & J. B. Fan (2006) Human
embryonic stem cells have a unique epigenetic signature. Genome Res, 16, 1075-83.
Boland, M. J., K. L. Nazor & J. F. Loring (2014) Epigenetic regulation of pluripotency and
differentiation. Circ Res, 115, 311-24.
Byvoet, P. (1972) In vivo turnover and distribution of radio-N-methyl in arginine-rich histones
from rat tissues. Arch Biochem Biophys, 152, 887-8.
Cai, L., Y. H. Tsai, P. Wang, J. Wang, D. Li, H. Fan, Y. Zhao, R. Bareja, R. Lu, E. M. Wilson,
A. Sboner, Y. E. Whang, D. Zheng, J. S. Parker, H. S. Earp & G. G. Wang (2018) ZFX
Mediates Non-canonical Oncogenic Functions of the Androgen Receptor Splice Variant 7
in Castrate-Resistant Prostate Cancer. Mol Cell, 72, 341-354.e6.
Carroll, D. (2011) Genome engineering with zinc-finger nucleases. Genetics, 188, 773-82.
Casas, C. (2017) GRP78 at the Centre of the Stage in Cancer and Neuroprotection. Front
Neurosci, 11, 177.
Cheah, M. S., C. D. Wallace & R. M. Hoffman (1984) Hypomethylation of DNA in human
cancer cells: a site-specific change in the c-myc oncogene. J Natl Cancer Inst, 73, 1057-
65.
51
Chen, N. M., A. Neesse, M. L. Dyck, B. Steuber, A. O. Koenig, C. Lubeseder-Martellato, T.
Winter, T. Forster, H. Bohnenberger, J. Kitz, K. Reuter-Jessen, H. Griesmann, J.
Gaedcke, M. Grade, J. S. Zhang, W. C. Tsai, J. Siveke, H. U. Schildhaus, P. Ströbel, S.
A. Johnsen, V. Ellenrieder & E. Hessmann (2017) Context-Dependent Epigenetic
Regulation of Nuclear Factor of Activated T Cells 1 in Pancreatic Plasticity.
Gastroenterology, 152, 1507-1520.e15.
Choi, J. H., H. J. Kwon, B. I. Yoon, J. H. Kim, S. U. Han, H. J. Joo & D. Y. Kim (2001)
Expression profile of histone deacetylase 1 in gastric cancer tissues. Jpn J Cancer Res,
92, 1300-4.
Cloos, P. A., J. Christensen, K. Agger, A. Maiolica, J. Rappsilber, T. Antal, K. H. Hansen & K.
Helin (2006) The putative oncogene GASC1 demethylates tri- and dimethylated lysine 9
on histone H3. Nature, 442, 307-11.
Colella, P., G. Ronzitti & F. Mingozzi (2018) Emerging Issues in AAV-Mediated. Mol Ther
Methods Clin Dev, 8, 87-104.
Consortium, E. P. (2012) An integrated encyclopedia of DNA elements in the human genome.
Nature, 489, 57-74.
Eden, A., F. Gaudet, A. Waghmare & R. Jaenisch (2003) Chromosomal instability and tumors
promoted by DNA hypomethylation. Science, 300, 455.
Esteller, M. (2005) Aberrant DNA methylation as a cancer-inducing mechanism. Annu Rev
Pharmacol Toxicol, 45, 629-56.
--- (2008) Epigenetics in cancer. N Engl J Med, 358, 1148-59.
Estève, P. O., H. G. Chin, A. Smallwood, G. R. Feehery, O. Gangisetty, A. R. Karpf, M. F.
Carey & S. Pradhan (2006) Direct interaction between DNMT1 and G9a coordinates
DNA and histone methylation during replication. Genes Dev, 20, 3089-103.
Feng, Q., H. Wang, H. H. Ng, H. Erdjument-Bromage, P. Tempst, K. Struhl & Y. Zhang (2002)
Methylation of H3-lysine 79 is mediated by a new family of HMTases without a SET
domain. Curr Biol, 12, 1052-8.
Flynn, R., A. Grundmann, P. Renz, W. Hänseler, W. S. James, S. A. Cowley & M. D. Moore
(2015) CRISPR-mediated genotypic and phenotypic correction of a chronic
granulomatous disease mutation in human iPS cells. Exp Hematol, 43, 838-848.e3.
Fraga, M. F., E. Ballestar, A. Villar-Garea, M. Boix-Chornet, J. Espada, G. Schotta, T. Bonaldi,
C. Haydon, S. Ropero, K. Petrie, N. G. Iyer, A. Pérez-Rosado, E. Calvo, J. A. Lopez, A.
Cano, M. J. Calasanz, D. Colomer, M. A. Piris, N. Ahn, A. Imhof, C. Caldas, T.
Jenuwein & M. Esteller (2005) Loss of acetylation at Lys16 and trimethylation at Lys20
of histone H4 is a common hallmark of human cancer. Nat Genet, 37, 391-400.
Fuks, F. (2005) DNA methylation and histone modifications: teaming up to silence genes. Curr
Opin Genet Dev, 15, 490-5.
Fuks, F., P. J. Hurd, R. Deplus & T. Kouzarides (2003a) The DNA methyltransferases associate
with HP1 and the SUV39H1 histone methyltransferase. Nucleic Acids Res, 31, 2305-12.
Fuks, F., P. J. Hurd, D. Wolf, X. Nan, A. P. Bird & T. Kouzarides (2003b) The methyl-CpG-
binding protein MeCP2 links DNA methylation to histone methylation. J Biol Chem, 278,
4035-40.
Futscher, B. W., M. M. Oshiro, R. J. Wozniak, N. Holtan, C. L. Hanigan, H. Duan & F. E.
Domann (2002) Role for DNA methylation in the control of cell type specific maspin
expression. Nat Genet, 31, 175-9.
52
Gao, X., J. C. Tsang, F. Gaba, D. Wu, L. Lu & P. Liu (2014) Comparison of TALE designer
transcription factors and the CRISPR/dCas9 in regulation of gene expression by targeting
enhancers. Nucleic Acids Res, 42, e155.
Gilbert, L. A., M. A. Horlbeck, B. Adamson, J. E. Villalta, Y. Chen, E. H. Whitehead, C.
Guimaraes, B. Panning, H. L. Ploegh, M. C. Bassik, L. S. Qi, M. Kampmann & J. S.
Weissman (2014) Genome-Scale CRISPR-Mediated Control of Gene Repression and
Activation. Cell, 159, 647-61.
Gilbert, L. A., M. H. Larson, L. Morsut, Z. Liu, G. A. Brar, S. E. Torres, N. Stern-Ginossar, O.
Brandman, E. H. Whitehead, J. A. Doudna, W. A. Lim, J. S. Weissman & L. S. Qi (2013)
CRISPR-mediated modular RNA-guided regulation of transcription in eukaryotes. Cell,
154, 442-51.
Goering, W., M. Kloth & W. A. Schulz (2012) DNA methylation changes in prostate cancer.
Methods Mol Biol, 863, 47-66.
Greer, E. L. & Y. Shi (2012) Histone methylation: a dynamic mark in health, disease and
inheritance. Nat Rev Genet, 13, 343-57.
Halkidou, K., L. Gaughan, S. Cook, H. Y. Leung, D. E. Neal & C. N. Robson (2004)
Upregulation and nuclear recruitment of HDAC1 in hormone refractory prostate cancer.
Prostate, 59, 177-89.
Hellman, A. & A. Chess (2007) Gene body-specific methylation on the active X chromosome.
Science, 315, 1141-3.
Hemmi, H., O. Takeuchi, T. Kawai, T. Kaisho, S. Sato, H. Sanjo, M. Matsumoto, K. Hoshino, H.
Wagner, K. Takeda & S. Akira (2000) A Toll-like receptor recognizes bacterial DNA.
Nature, 408, 740-5.
Holzmann, C., S. Kappel, T. Kilch, M. M. Jochum, S. K. Urban, V. Jung, M. Stöckle, K. Rother,
M. Greiner & C. Peinelt (2015) Transient receptor potential melastatin 4 channel
contributes to migration of androgen-insensitive prostate cancer cells. Oncotarget, 6,
41783-93.
Ibrahim, A. & E. Marbán (2016) Exosomes: Fundamental Biology and Roles in Cardiovascular
Physiology. Annu Rev Physiol, 78, 67-83.
Jiang, M., S. Xu, W. Yue, X. Zhao, L. Zhang, C. Zhang & Y. Wang (2012) The role of ZFX in
non-small cell lung cancer development. Oncol Res, 20, 171-8.
Jinek, M., K. Chylinski, I. Fonfara, M. Hauer, J. A. Doudna & E. Charpentier (2012) A
programmable dual-RNA-guided DNA endonuclease in adaptive bacterial immunity.
Science, 337, 816-21.
Jones, P. A. (2012) Functions of DNA methylation: islands, start sites, gene bodies and beyond.
Nat Rev Genet, 13, 484-92.
Juan, L. J., W. J. Shia, M. H. Chen, W. M. Yang, E. Seto, Y. S. Lin & C. W. Wu (2000) Histone
deacetylases specifically down-regulate p53-dependent gene activation. J Biol Chem,
275, 20436-43.
Jurkowska, R. Z., T. P. Jurkowski & A. Jeltsch (2011) Structure and function of mammalian
DNA methyltransferases. Chembiochem, 12, 206-22.
Kanwal, R. & S. Gupta (2012) Epigenetic modifications in cancer. Clin Genet, 81, 303-11.
Karpf, A. R. & S. Matsui (2005) Genetic disruption of cytosine DNA methyltransferase enzymes
induces chromosomal instability in human cancer cells. Cancer Res, 65, 8635-9.
53
Kearns, N. A., H. Pham, B. Tabak, R. M. Genga, N. J. Silverstein, M. Garber & R. Maehr (2015)
Functional annotation of native enhancers with a Cas9-histone demethylase fusion. Nat
Methods, 12, 401-403.
King, A. D., K. Huang, L. Rubbi, S. Liu, C. Y. Wang, Y. Wang, M. Pellegrini & G. Fan (2016)
Reversible Regulation of Promoter and Enhancer Histone Landscape by DNA
Methylation in Mouse Embryonic Stem Cells. Cell Rep, 17, 289-302.
Klann, T. S., J. B. Black, M. Chellappan, A. Safi, L. Song, I. B. Hilton, G. E. Crawford, T. E.
Reddy & C. A. Gersbach (2017) CRISPR-Cas9 epigenome editing enables high-
throughput screening for functional regulatory elements in the human genome. Nat
Biotechnol, 35, 561-568.
Klein, S. J. & R. J. O'Neill (2018) Transposable elements: genome innovation, chromosome
diversity, and centromere conflict. Chromosome Res, 26, 5-23.
Kosicki, M., K. Tomberg & A. Bradley (2018) Repair of double-strand breaks induced by
CRISPR-Cas9 leads to large deletions and complex rearrangements. Nat Biotechnol, 36,
765-771.
Koumenis, C. & B. G. Wouters (2006) "Translating" tumor hypoxia: unfolded protein response
(UPR)-dependent and UPR-independent pathways. Mol Cancer Res, 4, 423-36.
Kungulovski, G., S. Nunna, M. Thomas, U. M. Zanger, R. Reinhardt & A. Jeltsch (2015)
Targeted epigenome editing of an endogenous locus with chromatin modifiers is not
stably maintained. Epigenetics Chromatin, 8, 12.
Larson, M. H., L. A. Gilbert, X. Wang, W. A. Lim, J. S. Weissman & L. S. Qi (2013) CRISPR
interference (CRISPRi) for sequence-specific control of gene expression. Nat Protoc, 8,
2180-96.
Lehnertz, B., Y. Ueda, A. A. Derijck, U. Braunschweig, L. Perez-Burgos, S. Kubicek, T. Chen,
E. Li, T. Jenuwein & A. H. Peters (2003) Suv39h-mediated histone H3 lysine 9
methylation directs DNA methylation to major satellite repeats at pericentric
heterochromatin. Curr Biol, 13, 1192-200.
Li, E. (2002) Chromatin modification and epigenetic reprogramming in mammalian
development. Nat Rev Genet, 3, 662-73.
Lin, Y., J. Wu, W. Gu, Y. Huang, Z. Tong, L. Huang & J. Tan (2018) Exosome-Liposome
Hybrid Nanoparticles Deliver CRISPR/Cas9 System in MSCs. Adv Sci (Weinh), 5,
1700611.
Liu, T. Y., W. Gong, Z. J. Tan, W. Lu, X. S. Wu, H. Weng, Q. Ding, Y. J. Shu, R. F. Bao, Y.
Cao, X. A. Wang, F. Zhang, H. F. Li, S. S. Xiang, L. Jiang, Y. P. Hu, J. S. Mu, M. L. Li,
W. G. Wu, B. Y. Shen, L. X. Jiang & Y. B. Liu (2015) Baicalein inhibits progression of
gallbladder cancer cells by downregulating ZFX. PLoS One, 10, e0114851.
Lo, A. & L. Qi (2017). F1000Res, 6.
Lu, T., W. Yang, Z. Wang, Z. Hu, X. Zeng, C. Yang, Y. Wang, Y. Zhang, F. Li, Z. Liu, D. Wang
& Z. Ye (2015) Knockdown of glucose-regulated protein 78/binding immunoglobulin
heavy chain protein expression by asymmetric small interfering RNA induces apoptosis
in prostate cancer cells and attenuates migratory capability. Mol Med Rep, 11, 249-56.
Massie, C. E., I. G. Mills & A. G. Lynch (2017) The importance of DNA methylation in prostate
cancer development. J Steroid Biochem Mol Biol, 166, 1-15.
Metzger, E., M. Wissmann, N. Yin, J. M. Müller, R. Schneider, A. H. Peters, T. Günther, R.
Buettner & R. Schüle (2005) LSD1 demethylates repressive histone marks to promote
androgen-receptor-dependent transcription. Nature, 437, 436-9.
54
Mohn, F., M. Weber, M. Rebhan, T. C. Roloff, J. Richter, M. B. Stadler, M. Bibel & D.
Schübeler (2008) Lineage-specific polycomb targets and de novo DNA methylation
define restriction and potential of neuronal progenitors. Mol Cell, 30, 755-66.
Morgan, M. A. & A. Shilatifard (2015) Chromatin signatures of cancer. Genes Dev, 29, 238-49.
MURRAY, K. (1964) THE OCCURRENCE OF EPSILON-N-METHYL LYSINE IN
HISTONES. Biochemistry, 3, 10-5.
Najafabadi, H. S., S. Mnaimneh, F. W. Schmitges, M. Garton, K. N. Lam, A. Yang, M. Albu, M.
T. Weirauch, E. Radovani, P. M. Kim, J. Greenblatt, B. J. Frey & T. R. Hughes (2015)
C2H2 zinc finger proteins greatly expand the human regulatory lexicon. Nat Biotechnol,
33, 555-62.
Nakazawa, T., T. Kondo, D. Ma, D. Niu, K. Mochizuki, T. Kawasaki, T. Yamane, H. Iino, H.
Fujii & R. Katoh (2012) Global histone modification of histone H3 in colorectal cancer
and its precursor lesions. Hum Pathol, 43, 834-42.
Nazor, K. L., G. Altun, C. Lynch, H. Tran, J. V. Harness, I. Slavin, I. Garitaonandia, F. J. Müller,
Y. C. Wang, F. S. Boscolo, E. Fakunle, B. Dumevska, S. Lee, H. S. Park, T. Olee, D. D.
D'Lima, R. Semechkin, M. M. Parast, V. Galat, A. L. Laslett, U. Schmidt, H. S.
Keirstead, J. F. Loring & L. C. Laurent (2012) Recurrent variations in DNA methylation
in human pluripotent stem cells and their differentiated derivatives. Cell Stem Cell, 10,
620-34.
Nelson, C. E., Y. Wu, M. P. Gemberling, M. L. Oliver, M. A. Waller, J. D. Bohning, J. N.
Robinson-Hamm, K. Bulaklak, R. M. Castellanos Rivera, J. H. Collier, A. Asokan & C.
A. Gersbach (2019) Long-term evaluation of AAV-CRISPR genome editing for
Duchenne muscular dystrophy. Nat Med, 25, 427-432.
Nesterova, T. B., C. M. Johnston, R. Appanah, A. E. Newall, J. Godwin, M. Alexiou & N.
Brockdorff (2003) Skewing X chromosome choice by modulating sense transcription
across the Xist locus. Genes Dev, 17, 2177-90.
Ni, M., Y. Zhang & A. S. Lee (2011) Beyond the endoplasmic reticulum: atypical GRP78 in cell
viability, signalling and therapeutic targeting. Biochem J, 434, 181-8.
O'Geen, H., S. L. Bates, S. S. Carter, K. A. Nisson, J. Halmai, K. D. Fink, S. K. Rhie, P. J.
Farnham & D. J. Segal (2019) Ezh2-dCas9 and KRAB-dCas9 enable engineering of
epigenetic memory in a context-dependent manner. Epigenetics Chromatin, 12, 26.
O'Geen, H., I. M. Henry, M. S. Bhakta, J. F. Meckler & D. J. Segal (2015) A genome-wide
analysis of Cas9 binding specificity using ChIP-seq and targeted sequence capture.
Nucleic Acids Res, 43, 3389-404.
O'Geen, H., C. Ren, C. M. Nicolet, A. A. Perez, J. Halmai, V. M. Le, J. P. Mackay, P. J.
Farnham & D. J. Segal (2017) dCas9-based epigenome editing suggests acquisition of
histone methylation is not sufficient for target gene repression. Nucleic Acids Res, 45,
9901-9916.
Ogryzko, V. V., R. L. Schiltz, V. Russanova, B. H. Howard & Y. Nakatani (1996) The
transcriptional coactivators p300 and CBP are histone acetyltransferases. Cell, 87, 953-9.
Panfil, A. R., J. A. London, P. L. Green & K. E. Yoder (2018) CRISPR/Cas9 Genome Editing to
Disable the Latent HIV-1 Provirus. Front Microbiol, 9, 3107.
Parsi, K. M., E. Hennessy, N. Kearns & R. Maehr (2017) Using an Inducible CRISPR-dCas9-
KRAB Effector System to Dissect Transcriptional Regulation in Human Embryonic Stem
Cells. Methods Mol Biol, 1507, 221-233.
55
Ramakrishna, S., A. B. Kwaku Dad, J. Beloor, R. Gopalappa, S. K. Lee & H. Kim (2014) Gene
disruption by cell-penetrating peptide-mediated delivery of Cas9 protein and guide RNA.
Genome Res, 24, 1020-7.
Rauch, T. A., Z. Wang, X. Wu, K. H. Kernstine, A. D. Riggs & G. P. Pfeifer (2012) DNA
methylation biomarkers for lung cancer. Tumour Biol, 33, 287-96.
Rea, S., F. Eisenhaber, D. O'Carroll, B. D. Strahl, Z. W. Sun, M. Schmid, S. Opravil, K.
Mechtler, C. P. Ponting, C. D. Allis & T. Jenuwein (2000) Regulation of chromatin
structure by site-specific histone H3 methyltransferases. Nature, 406, 593-9.
Rezza, A., C. Jacquet, A. Le Pillouer, F. Lafarguette, C. Ruptier, M. Billandon, P. Isnard Petit, S.
Trouttet, K. Thiam, A. Fraichard & Y. Chérifi (2019) Unexpected genomic
rearrangements at targeted loci associated with CRISPR/Cas9-mediated knock-in. Sci
Rep, 9, 3486.
Rhie, S. K., L. Yao, Z. Luo, H. Witt, S. Schreiner, Y. Guo, A. A. Perez & P. J. Farnham (2018)
ZFX acts as a transcriptional activator in multiple types of human tumors by binding
downstream of transcription start sites at the majority of CpG island promoters. Genome
Res.
Rundlett, S. E., A. A. Carmen, R. Kobayashi, S. Bavykin, B. M. Turner & M. Grunstein (1996)
HDA1 and RPD3 are members of distinct yeast histone deacetylase complexes that
regulate silencing and transcription. Proc Natl Acad Sci U S A, 93, 14503-8.
Sagredo, A. I., E. A. Sagredo, C. Cappelli, P. Báez, R. E. Andaur, C. Blanco, J. C. Tapia, C.
Echeverría, O. Cerda, A. Stutzin, F. Simon, K. Marcelain & R. Armisén (2018) TRPM4
regulates Akt/GSK3-β activity and enhances β-catenin signaling and cell proliferation in
prostate cancer cells. Mol Oncol, 12, 151-165.
Sarraf, S. A. & I. Stancheva (2004) Methyl-CpG binding protein MBD1 couples histone H3
methylation at lysine 9 by SETDB1 to DNA replication and chromatin assembly. Mol
Cell, 15, 595-605.
Sato, M., V. J. Yao, W. Arap & R. Pasqualini (2010) GRP78 signaling hub a receptor for
targeted tumor therapy. Adv Genet, 69, 97-114.
Schwank, G., B. K. Koo, V. Sasselli, J. F. Dekkers, I. Heo, T. Demircan, N. Sasaki, S. Boymans,
E. Cuppen, C. K. van der Ent, E. E. Nieuwenhuis, J. M. Beekman & H. Clevers (2013)
Functional repair of CFTR by CRISPR/Cas9 in intestinal stem cell organoids of cystic
fibrosis patients. Cell Stem Cell, 13, 653-8.
Seligson, D. B., S. Horvath, M. A. McBrian, V. Mah, H. Yu, S. Tze, Q. Wang, D. Chia, L.
Goodglick & S. K. Kurdistani (2009) Global levels of histone modifications predict
prognosis in different cancers. Am J Pathol, 174, 1619-28.
Sharp, A. J., E. Stathaki, E. Migliavacca, M. Brahmachary, S. B. Montgomery, Y. Dupre & S. E.
Antonarakis (2011) DNA methylation profiles of human active and inactive X
chromosomes. Genome Res, 21, 1592-600.
Shen, J., D. P. Ha, G. Zhu, D. F. Rangel, A. Kobielak, P. S. Gill, S. Groshen, L. Dubeau & A. S.
Lee (2017) GRP78 haploinsufficiency suppresses acinar-to-ductal metaplasia, signaling,
and mutant. Proc Natl Acad Sci U S A, 114, E4020-E4029.
Shi, Y., F. Lan, C. Matson, P. Mulligan, J. R. Whetstine, P. A. Cole & R. A. Casero (2004)
Histone demethylation mediated by the nuclear amine oxidase homolog LSD1. Cell, 119,
941-53.
Silverman, B. R. & J. Shi (2016) Alterations of Epigenetic Regulators in Pancreatic Cancer and
Their Clinical Implications. Int J Mol Sci, 17.
56
Song, X., M. Zhu, F. Zhang, Y. Zhang, Y. Hu, L. Jiang, Y. Hao, S. Chen, Q. Zhu, W. Huang, J.
Lu, J. Gu, W. Gong, M. Li & Y. Liu (2018) ZFX Promotes Proliferation and Metastasis
of Pancreatic Cancer Cells via the MAPK Pathway. Cell Physiol Biochem, 48, 274-284.
Soshnev, A. A., S. Z. Josefowicz & C. D. Allis (2016) Greater Than the Sum of Parts:
Complexity of the Dynamic Epigenome. Mol Cell, 62, 681-94.
Staquicini, D. I., S. D'Angelo, F. Ferrara, K. Karjalainen, G. Sharma, T. L. Smith, C. A.
Tarleton, D. E. Jaalouk, A. Kuniyasu, W. B. Baze, B. K. Chaffee, P. W. Hanley, K. F.
Barnhart, E. Koivunen, S. Marchiò, R. L. Sidman, J. E. Cortes, H. M. Kantarjian, W.
Arap & R. Pasqualini (2018) Therapeutic targeting of membrane-associated GRP78 in
leukemia and lymphoma: preclinical efficacy in vitro and formal toxicity study of BMTP-
78 in rodents and primates. Pharmacogenomics J, 18, 436-443.
Stubbs, L., Y. Sun & D. Caetano-Anolles (2011) Function and Evolution of C2H2 Zinc Finger
Arrays. Subcell Biochem, 52, 75-94.
Taunton, J., C. A. Hassig & S. L. Schreiber (1996) A mammalian histone deacetylase related to
the yeast transcriptional regulator Rpd3p. Science, 272, 408-11.
Tebas, P., D. Stein, W. W. Tang, I. Frank, S. Q. Wang, G. Lee, S. K. Spratt, R. T. Surosky, M.
A. Giedlin, G. Nichol, M. C. Holmes, P. D. Gregory, D. G. Ando, M. Kalos, R. G.
Collman, G. Binder-Scholl, G. Plesa, W. T. Hwang, B. L. Levine & C. H. June (2014)
Gene editing of CCR5 in autologous CD4 T cells of persons infected with HIV. N Engl J
Med, 370, 901-10.
Thakore, P. I., A. M. D'Ippolito, L. Song, A. Safi, N. K. Shivakumar, A. M. Kabadi, T. E.
Reddy, G. E. Crawford & C. A. Gersbach (2015) Highly specific epigenome editing by
CRISPR-Cas9 repressors for silencing of distal regulatory elements. Nat Methods, 12,
1143-9.
Thurman, R. E., E. Rynes, R. Humbert, J. Vierstra, M. T. Maurano, E. Haugen, N. C. Sheffield,
A. B. Stergachis, H. Wang, B. Vernot, K. Garg, S. John, R. Sandstrom, D. Bates, L.
Boatman, T. K. Canfield, M. Diegel, D. Dunn, A. K. Ebersol, T. Frum, E. Giste, A. K.
Johnson, E. M. Johnson, T. Kutyavin, B. Lajoie, B. K. Lee, K. Lee, D. London, D.
Lotakis, S. Neph, F. Neri, E. D. Nguyen, H. Qu, A. P. Reynolds, V. Roach, A. Safi, M. E.
Sanchez, A. Sanyal, A. Shafer, J. M. Simon, L. Song, S. Vong, M. Weaver, Y. Yan, Z.
Zhang, B. Lenhard, M. Tewari, M. O. Dorschner, R. S. Hansen, P. A. Navas, G.
Stamatoyannopoulos, V. R. Iyer, J. D. Lieb, S. R. Sunyaev, J. M. Akey, P. J. Sabo, R.
Kaul, T. S. Furey, J. Dekker, G. E. Crawford & J. A. Stamatoyannopoulos (2012) The
accessible chromatin landscape of the human genome. Nature, 489, 75-82.
Tsujiuchi, T., M. Tsutsumi, Y. Sasaki, M. Takahama & Y. Konishi (1999) Hypomethylation of
CpG sites and c-myc gene overexpression in hepatocellular carcinomas, but not
hyperplastic nodules, induced by a choline-deficient L-amino acid-defined diet in rats.
Jpn J Cancer Res, 90, 909-13.
Tsukada, Y., J. Fang, H. Erdjument-Bromage, M. E. Warren, C. H. Borchers, P. Tempst & Y.
Zhang (2006) Histone demethylation by a family of JmjC domain-containing proteins.
Nature, 439, 811-6.
van den Boorn, J. G., M. Schlee, C. Coch & G. Hartmann (2011) SiRNA delivery with exosome
nanoparticles. Nat Biotechnol, 29, 325-6.
Van Hoesen, K., S. Meynier, P. Ribaux, P. Petignat, F. Delie & M. Cohen (2017) Circulating
GRP78 antibodies from ovarian cancer patients: a promising tool for cancer cell targeting
drug delivery system? Oncotarget, 8, 107176-107187.
57
Verdone, L., M. Caserta & E. Di Mauro (2005) Role of histone acetylation in the control of gene
expression. Biochem Cell Biol, 83, 344-53.
Viré, E., C. Brenner, R. Deplus, L. Blanchon, M. Fraga, C. Didelot, L. Morey, A. Van Eynde, D.
Bernard, J. M. Vanderwinden, M. Bollen, M. Esteller, L. Di Croce, Y. de Launoit & F.
Fuks (2006) The Polycomb group protein EZH2 directly controls DNA methylation.
Nature, 439, 871-4.
Wagner, H. (2001) Toll meets bacterial CpG-DNA. Immunity, 14, 499-502.
Wang, G., M. L. McCain, L. Yang, A. He, F. S. Pasqualini, A. Agarwal, H. Yuan, D. Jiang, D.
Zhang, L. Zangi, J. Geva, A. E. Roberts, Q. Ma, J. Ding, J. Chen, D. Z. Wang, K. Li, J.
Wang, R. J. Wanders, W. Kulik, F. M. Vaz, M. A. Laflamme, C. E. Murry, K. R. Chien,
R. I. Kelley, G. M. Church, K. K. Parker & W. T. Pu (2014) Modeling the mitochondrial
cardiomyopathy of Barth syndrome with induced pluripotent stem cell and heart-on-chip
technologies. Nat Med, 20, 616-23.
Wang, H., R. Guo, Z. Du, L. Bai, L. Li, J. Cui, W. Li, A. R. Hoffman & J. F. Hu (2018)
Epigenetic Targeting of Granulin in Hepatoma Cells by Synthetic CRISPR dCas9 Epi-
suppressors. Mol Ther Nucleic Acids, 11, 23-33.
Wang, M., S. Wey, Y. Zhang, R. Ye & A. S. Lee (2009) Role of the unfolded protein response
regulator GRP78/BiP in development, cancer, and neurological disorders. Antioxid Redox
Signal, 11, 2307-16.
Whetstine, J. R., A. Nottke, F. Lan, M. Huarte, S. Smolikov, Z. Chen, E. Spooner, E. Li, G.
Zhang, M. Colaiacovo & Y. Shi (2006) Reversal of histone lysine trimethylation by the
JMJD2 family of histone demethylases. Cell, 125, 467-81.
Xia, A. L., Q. F. He, J. C. Wang, J. Zhu, Y. Q. Sha, B. Sun & X. J. Lu (2019) Applications and
advances of CRISPR-Cas9 in cancer immunotherapy. J Med Genet, 56, 4-9.
Xu, P., Y. Tong, X. Z. Liu, T. T. Wang, L. Cheng, B. Y. Wang, X. Lv, Y. Huang & D. P. Liu
(2015) Both TALENs and CRISPR/Cas9 directly target the HBB IVS2-654 (C > T)
mutation in β-thalassemia-derived iPSCs. Sci Rep, 5, 12065.
Yamane, K., C. Toumazou, Y. Tsukada, H. Erdjument-Bromage, P. Tempst, J. Wong & Y.
Zhang (2006) JHDM2A, a JmjC-containing H3K9 demethylase, facilitates transcription
activation by androgen receptor. Cell, 125, 483-95.
Yan, X. J., J. Xu, Z. H. Gu, C. M. Pan, G. Lu, Y. Shen, J. Y. Shi, Y. M. Zhu, L. Tang, X. W.
Zhang, W. X. Liang, J. Q. Mi, H. D. Song, K. Q. Li, Z. Chen & S. J. Chen (2011) Exome
sequencing identifies somatic mutations of DNA methyltransferase gene DNMT3A in
acute monocytic leukemia. Nat Genet, 43, 309-15.
Yang, H., Y. Lu, Y. Zheng, X. Yu, X. Xia, X. He, W. Feng, L. Xing & Z. Ling (2014) shRNA-
mediated silencing of ZFX attenuated the proliferation of breast cancer cells. Cancer
Chemother Pharmacol, 73, 569-76.
Yasui, W., N. Oue, S. Ono, Y. Mitani, R. Ito & H. Nakayama (2003) Histone acetylation and
gastrointestinal carcinogenesis. Ann N Y Acad Sci, 983, 220-31.
Yeo, N. C., A. Chavez, A. Lance-Byrne, Y. Chan, D. Menn, D. Milanova, C. C. Kuo, X. Guo, S.
Sharma, A. Tung, R. J. Cecchi, M. Tuttle, S. Pradhan, E. T. Lim, N. Davidsohn, M. R.
Ebrahimkhani, J. J. Collins, N. E. Lewis, S. Kiani & G. M. Church (2018) An enhanced
CRISPR repressor for targeted mammalian gene regulation. Nat Methods, 15, 611-616.
Zhang, Y. & D. Reinberg (2001) Transcription regulation by histone methylation: interplay
between different covalent modifications of the core histone tails. Genes Dev, 15, 2343-
60.
58
Zheng, Y., W. Shen, J. Zhang, B. Yang, Y. N. Liu, H. Qi, X. Yu, S. Y. Lu, Y. Chen, Y. Z. Xu, Y.
Li, F. H. Gage, S. Mi & J. Yao (2018) Author Correction: CRISPR interference-based
specific and efficient gene inactivation in the brain. Nat Neurosci, 21, 894.
Zhou, Y., Z. Su, Y. Huang, T. Sun, S. Chen, T. Wu, G. Chen, X. Xie, B. Li & Z. Du (2011) The
Zfx gene is expressed in human gliomas and is important in the proliferation and
apoptosis of the human malignant glioma cell line U251. J Exp Clin Cancer Res, 30, 114.
Abstract (if available)
Abstract
Gene expression patterns are modulated by reversible changes to the epigenome. Such epigenetic changes take the form of histone modifications and DNA methylation, which may work in concert to stably activate or repress a gene. In cancer, the epigenetic landscapes of many tumor-repressor genes and oncogenes are distinctive from those in normal cells. Abnormal epigenetic signatures may thus drive carcinogenesis by altering the expression of key genes involved in driving or suppressing tumorigenesis. Attempts to reverse these disease-associated expression changes through artificial deposition of specific marks have largely focused on generating transient results. Targeting powerful histone modifiers such as H3K9me3 or H3K27me3 depositors to a site of interest can achieve potent, specific gene repression. Maintaining this induced repression, however, may require that histone modifiers be targeted in conjunction with DNA methyltransferases. Here, I use a modified CRISPR-dCas9 system to explore whether lasting, or persistent, repression of three tumor-promoting genes can be achieved with a variety of epigenetic modifiers. I report that efficient DNA methylation in conjunction with the repressive histone modification H3K9me3 can persistently silence a promoter for at least 30 days post-transfection and the degree of achievable persistent repression is dependent on both the target gene and cell type.
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Asset Metadata
Creator
Nisson, Karly Ann
(author)
Core Title
Using epigenetic toggle switches to repress tumor-promoting gene expression
School
Keck School of Medicine
Degree
Master of Science
Degree Program
Biochemistry and Molecular Medicine
Publication Date
07/23/2019
Defense Date
06/07/2019
Publisher
University of Southern California
(original),
University of Southern California. Libraries
(digital)
Tag
cancer,CRISPR,dCas9,DNA methylation,epigenetics,gene expression,gene repression,histone modifications,OAI-PMH Harvest,persistent repression,promoter,prostate cancer,toggle switch,Tumor
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application/pdf
(imt)
Language
English
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Electronically uploaded by the author
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Advisor
Farnham, Peggy (
committee chair
), Bell, Oliver (
committee member
), Stallcup, Michael (
committee member
)
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karlyann@att.net,knisson@usc.edu
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https://doi.org/10.25549/usctheses-c89-187041
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UC11660819
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etd-NissonKarl-7583.pdf (filename),usctheses-c89-187041 (legacy record id)
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Nisson, Karly Ann
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Tags
CRISPR
dCas9
DNA methylation
epigenetics
gene expression
gene repression
histone modifications
persistent repression
promoter
prostate cancer
toggle switch