epigenetic reprogramming and dna demethylation · however reprogramming of imprinting is only...
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Epigenetic Reprogramming and DNA Demethylation
HAKAN BAGCI
Imperial College London
Faculty of Medicine
MRC Clinical Sciences Centre
Doctor of Philosophy
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I, Hakan Bagci, hereby declare that this thesis is my own work and that work performed by others has been appropriately acknowledged and referenced.
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The copyright of this thesis rests with the author and is made available under a Creative Commons Attribution Non-Commercial No Derivatives licence. Researchers are free to copy, distribute or transmit the thesis on the condition that they attribute it, that they do not use it for commercial purposes and that they do not alter, transform or build upon it. For any reuse or redistribution, researchers must make clear to others the licence terms
of this work.
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Abstract Embryonic development begins with fertilization of the egg, a progressive process
that gives rise to the zygote and subsequently to the formation of somatic tissues.
Normally once cells acquire a fate, it is stably maintained. Conversion back to a
multipotent state occurs rarely in-vivo, but can be achieved experimentally by inducing
‘reprogramming’. In this study I am looking at the epigenetic mechanisms that underlie
reprogramming and, in particular, DNA methylation and demethylation. To address this I
am taking advantage of the cellular fusion system. Fusion of pluripotent cells with
differentiated cells results in the formation of transient heterokaryon and hybrid cells,
where the somatic partner is efficiently reprogrammed. This gives me the opportunity to
monitor early and late events in pluripotent conversion, in which global remodelling of
chromatin and changes in DNA methylation occur.
Here, I examine changes in DNA methylation that are induced at imprinted loci and
pluripotency-associated genes when somatic cells are fused with either mouse embryonic
stem (ES) or embryonic germ (EG) cells. I focus on defining the factors and order of events
that accompany reprogramming. I show that acquisition of pluripotency is an early process
occurring at the heterokaryon stage, and is followed by imprint erasure later in hybrids.
However reprogramming of imprinting is only induced by EG, but not ES cells, and it
requires sequential steps of 5-methylcytosine oxidation mediated by Tet proteins and
nucleotide exchange upon several rounds of DNA synthesis. I provide evidence that Tet
proteins are dispensable for pluripotent reprogramming using CRISPR-Cas9 genome
editing to abrogate the expression of both Tet1 and Tet2. This result suggests that either
DNA demethylation can occur without TET activity (implying a redundancy with other
demethylating agents and routes), or that DNA demethylation is not required for inducing
pluripotency. Finally, I describe how CRISPR/Cas9 approaches were used to demonstrate
that non-canonical Wnt signalling components are downstream targets of JARID2 in ES
cells.
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Acknowledgments
I would first like to express my gratitude to Mandy and Matthias for giving me the opportunity to be a part of the scientifically stimulating, socially interacting, internationally competent Lymphocyte Development Team at the CSC. I am especially grateful to Mandy, for teaching me how to be open-minded, patient and collaborative in science, how to ask the right questions and how to approach them. Thank you very much for encouraging curiosity and innovation, always being tolerant, thoughtful and supportive with me. I would also like to thank the Medical Research Council, for financially supporting me and my PhD work.
I would like to thank all the past and present members of the LympDeve, for collectively creating a joyful and friendly scientific environment, for help, advice, support and long discussions, and for making my four years unforgettable. I would like to say “air hair lair” to Allifia, thank you for not only being a valuable friend, but also for trusting me and for providing me shelter in the last months of my thesis. I cannot put into words how great it feels to know you will be there whenever I am in trouble. Thank you Amélie for our little conversations, for your help with experiments, and for bringing order to our laboratory; without you, we would all be lost. Andy, my desk buddy, who by now should have become a YoYo master, thank you for your friendship, for marvellous memories, your jokes and puns that will always make me giggle. Thank you for bringing colour and laughter to the PhD office (also thank you to your iPod!). Feng, thank you for your humour, I wish we made a list, but I believe it would be a bit inappropriate to be written in here. Thank you Irene for long discussions, and for never being tired of sharing your extensive knowledge with me (and for delicious tiramisus). Thank you Jorge for your scientific help, for sharing your ideas, for always being kind, supportive and positive. Lee the east Londoner, thank you for great house parties, for teaching me the contemporary usage of the English language and exposing me to the alternative London lifestyle. Lesly, thank you for setting limits to our bad jokes. Preksha, I have always enjoyed discussing with you, on any subject, and thank you for always being there since the first day of our PhD. Thais, thank you for your never-ending kindness, and for our dialogues while cycling to the west. Ziwei, it has always been a pleasure talking to you, and thank you for your help with experiments. I am thankful to the present members Anne-Céline, Grainne, Isabel, Kotryna, Liz, Ludovica, Matt, Sergi, Tom, Vlad, and to the past members Antoine, Bryony, Cynthia, Hegias, Luke, Rory, for making my time more enjoyable in the laboratory, you will never be forgotten. I would specially like to thank Francesco, David and Karen, for instructive discussions and for your help and contribution to this thesis. I would finally like to thank James in the FACS facility, Zoe in the Transgenics facility, Microscopy facility and Sequencing facility for their help in conducting my PhD work.
Special thanks to my great friends Matteo, Joana, Silvia and Joao. Thank you for delicious dinners, for pub nights, for party nights, for laughters, for concerts, for travels, for festivals starting from the first days of our PhD… Our bond will never be broken.
Thank you Mélanie for your love, for your patience with me, for everything we shared and for all of our cheerful memories. Je t’envoie pleins de bisous!
And I would finally like to thank my parents. Annem ve babama: her ne kosulda olursa olsun her zaman arkamda oldugunuz icin, beni her kararimda desteklediginiz ve buralara kadar gelmemde en buyuk rolu oynadiginiz icin cok tesekkur ederim.
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Contents
Abstract........................................................................................................................... . 4
Acknowledgments.......................................................................................................... 5
Contents.......................................................................................................................... . 6
List of Figures & Tables ................................................................................................ 9
Abbreviations................................................................................................................. 11
Chapter 1. Introduction ............................................................................................... 13
1.1. Pluripotent embryonic stem cells .............................................................................. 13 1.1.1. Chromatin dynamics in embryonic stem cells ................................................................................... 15 1.1.2. Polycomb regulation and bivalency ................................................................................................... 16
1.2. DNA methylation dynamics in mammals ................................................................... 17 1.2.1. Maintenance and de novo establishment of DNA methylation ......................................................... 17 1.2.2. Roles of DNA methylation in gene regulation .................................................................................... 19 1.2.3. Genomic Imprinting ........................................................................................................................... 20 1.2.4. DNA demethylation ........................................................................................................................... 23
1.2.4.1. Passive demethylation ................................................................................................................ 23 1.2.4.2. Active demethylation ................................................................................................................. 25 1.2.4.3. TET protein mediated 5-mC oxidation in passive and active demethylation ............................. 27 1.2.4.4. TET-associated DNA demethylation dynamics in embryonic development and pluripotency .. 33
1.3. Reprogramming cell fate ........................................................................................... 38 1.3.1. Transdifferentiation ........................................................................................................................... 39 1.3.2. Pluripotent conversion of somatic cells ............................................................................................. 40
1.3.2.1. Nuclear transfer .......................................................................................................................... 40 1.3.2.2. Induced pluripotent stem cells ................................................................................................... 42 1.3.2.3. Cell fusion ................................................................................................................................... 43
1.4. Aims of this study ..................................................................................................... 47
Chapter 2. Materials and Methods .............................................................................. 48
2.1. Materials ................................................................................................................... 48 2.1.1. Cell lines ............................................................................................................................................. 48 2.1.2. Antibodies .......................................................................................................................................... 49
2.2. Methods.................................................................................................................... 49 2.2.1. Cell culture ......................................................................................................................................... 49 2.2.2. Cell fusion experiments ..................................................................................................................... 50 2.2.3. Fluorescence activated cell sorting (FACS) ........................................................................................ 51 2.2.4. Quantitative Reverse Transcription Polymerase Chain Reaction (qRT-PCR) Analysis ....................... 51
2.2.4.1. RNA extraction and reverse transcription .................................................................................. 51 2.2.4.2. Quantitative PCR ........................................................................................................................ 52
2.2.5. DNA methylation and hydroxymethylation analyses......................................................................... 52 2.2.5.1. Bisulfite sequencing analysis ...................................................................................................... 53 2.2.5.2. 5-hmC quantification by enzyme protection assay .................................................................... 53
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2.2.6. Imaging analysis ................................................................................................................................. 54 2.2.6.1. Immunofluorescence and confocal microscopy imaging ........................................................... 54 2.2.6.2. X-gal staining .............................................................................................................................. 54
2.2.7. Western Blot analysis ......................................................................................................................... 54 2.2.8. Chromatin Immunoprecipitation (ChIP) analysis ............................................................................... 55 2.2.9. Plasmid construction and delivery into ES cells ................................................................................. 56 2.2.10. CRISPR/Cas9 genome editing system .............................................................................................. 57
2.2.10.1. CRISPR/Cas9 plasmid construction ........................................................................................... 57 2.2.10.2. Surveyor and RFLP Assays ........................................................................................................ 58
Chapter 3. Pluripotency Gene Demethylation during Reprogramming ................... 59
3.1. Introduction ............................................................................................................... 59
3.2. Reprogramming of human B lymphocytes upon fusion with mouse embryonic stem cells ................................................................................................................................. 59
3.3. DNA methylation profiles of pluripotency associated genes in human B lymphocytes and human ES cells ......................................................................................................... 61
3.4. Changes in DNA methylation of OCT4 accompanies reprogramming but the extent is variable ............................................................................................................................ 62
3.5. Reprogramming human fibroblasts and OCT4 induction without detectable changes in DNA methylation ............................................................................................................. 64
3.6. No evidence of DNA methylation changes at site upstream of the OCT4 transcription start site. .......................................................................................................................... 66
3.7. DNA demethylation kinetics of somatic Oct4 transgene in reprogrammed mouse hybrids. ............................................................................................................................ 67
3.8. Summary and Discussion ......................................................................................... 68
Chapter 4. Mechanisms of Imprint Erasure in Somatic Cells mediated by Embryonic Germ Cell Fusion ...................................................................................... 72
4.1. Introduction ............................................................................................................... 72
4.2. Imprint erasure in somatic cells induced by embryonic germ cell fusion. ................... 72
4.3. Using dual reporter (2rB) somatic cells to assess the kinetics of imprint erasure during EG-reprogramming. ......................................................................................................... 73
4.4. EG cell capacity to induce demethylation is not restricted to imprinted genes. .......... 75
4.5. Imprint erasure is not seen in fusions with mouse ES cells or female ES cells that are globally hypomethylated. ................................................................................................. 76
4.6. Hydroxymethylation at imprinted loci upon fusion with mouse EG cells. ................... 78
4.7. Tet regulated 5-mC oxidation at imprinted loci upon fusion with mouse EG cells. ..... 79
4.8. Summary and Discussion ......................................................................................... 81
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Chapter 5. Analysis of TET Protein Requirement in Mouse Embryonic Stem Cell Induced Reprogramming of Human B Lymphocytes................................................ 84
5.1. Introduction ............................................................................................................... 84
5.2. Tet Knockdown in mouse ES cells and cell fusion .................................................... 84 5.2.1. Tet1 knockdown and fusion ............................................................................................................... 84 5.2.2. Tet2 knockdown and fusion ............................................................................................................... 87 5.2.3. Tet1 and Tet2 double knockdown and fusion .................................................................................... 89
5.3. CRISPR/Cas9 mediated Tet gene editing and cellular fusion ................................... 91 5.3.1. CRISPR/Cas9 system construction against Tet1 and Tet2 genes and delivery into mES cells ............ 91 5.3.2. Surveyor Assay for analysis of CRISPR/Cas efficiency ........................................................................ 94 5.3.3. Restriction Fragment Length Polymorphism screen on CRISPR/Cas9 targeted mES cells for Tet1 and Tet2 .............................................................................................................................................................. 95 5.3.4. Sequencing of Tet1&Tet2 CRISPR targeted ES clones ........................................................................ 97 5.3.5. Reprogramming capacity of CRISPR/Cas9 mediated Tet1 and Tet2 mutant ES cell clones upon cell fusion ........................................................................................................................................................... 99
5.4. Summary and Discussion ....................................................................................... 100
Chapter 6. CRISPR/Cas Editing of Jarid2 and Non-Canonical WNT Pathway Components…………………………………………………………………………………. 106
6.1. Introduction ............................................................................................................. 106
6.2. CRISPR/Cas9 editing of Jarid2 and Prickle1/Fzd2/Wnt9a in mouse embryonic stem cells ............................................................................................................................... 106
6.2.1. Guide RNA design and delivery into mouse ES cells ........................................................................ 107 6.2.2. Surveyor Assay for analysis of CRISPR/Cas9 efficiency .................................................................... 108 6.2.3. Clonal screens and sequencing for targeted Jarid2 locus in mouse ES cells .................................... 108 6.2.4. Clonal screens and sequencing for targeted Prickle1, Fzd2 and Wnt9a loci in mES cells. ............... 110
6.3. JARID2 deficiency in mouse ES cells can be phenocopied by Prickle1/Fzd2/Wnt targeting ........................................................................................................................ 113
6.4. Summary and Discussion ....................................................................................... 115
Chapter 7. General Discussion ................................................................................. 118
7.1. DNA methylation dynamics in reprogramming ........................................................ 118
7.2. Genome editing and the use of CRISPR/Cas9-based approaches ......................... 120
7.3. Future Studies ........................................................................................................ 123 7.3.1. Droplet-based microfluidics and cell fusion ..................................................................................... 124 7.3.2. Single-cell heterokaryon analysis ..................................................................................................... 126
Bibliography................................................................................................................ 128
Appendix...................................................................................................................... 155
Publications................................................................................................................. 158
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List of Figures & Tables
FIGURES
Figure 1.1. Maintenance/replication-coupled loss of DNA methylation
Figure 1.2. Mechanisms of dynamic modifications of cytosine
Figure 1.3. TET-induced demethylation in mouse embryonic development and ES cells
Figure 1.4. In-vitro strategies for nuclear reprogramming to pluripotency
Figure 2.1. Vectors used for delivery
Figure 2.2. px330 vector and the guide RNA sequence
Figure 3.1. Interspecies cell fusion and reprogramming of human B lymphocyte by mouse ES cells
Figure 3.2. Bisulfite sequencing of OCT4, NANOG and CRIPTO promoters in human B lymphocytes and human ES cells
Figure 3.3. Transcript analysis and bisulfite sequencing of human OCT4 in heterokaryons after 72 hours of fusion
Figure 3.4. Transcript analysis and bisulfite sequencing of human OCT4 in heterokaryons after 72 hours of fusion in four different experiments
Figure 3.5. Fusion of human fibroblasts with mouse ES cells and transcript and bisulfite sequencing analyses
Figure 3.6. Bisulfite sequencing of human OCT4 upstream region in human fibroblasts before and after fusion and in human ES cells
Figure 3.7. DNA demethylation kinetics upon reprogramming in mouse hybrids
Figure 4.1. CpG methylation analysis of imprinted H19 locus upon reprogramming mediated by mouse EG cells
Figure 4.2. CpG methylation analysis of imprinted Peg1 locus upon pluripotent reprogramming mediated by mouse EG cells
Figure 4.3. Functional resetting of somatic imprints mediated by mouse EG cells
Figure 4.4. CpG methylation analysis of LINE1 repeats upon pluripotent reprogramming mediated by mouse EG cells
Figure 4.5. CpG methylation analysis of imprinted H19, Peg3 and Gtl2/Dlk1 loci upon pluripotent reprogramming induced by mouse ES cells
Figure 4.6. CpG methylation analysis of imprinted H19, Peg3 and Gtl2/Dlk1 loci upon pluripotent reprogramming induced by Pgk12.1 female mouse ES cells
Figure 4.7. Acquisition of 5-hmC at human B lymphocyte ICRs upon fusion with mouse EG cells
Figure 4.8. Roles of TET proteins in the acquisition of 5-hmC at human B lymphocyte ICRs upon fusion with mouse EG cells.
Figure 5.1. Effect of Tet1 knockdown on reprogramming
Figure 5.2. Effect of Tet2 knockdown on reprogramming
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Figure 5.3. Effect of Tet1/Tet2 double knockdown on reprogramming
Figure 5.4. Tet1 and Tet2 targeting by CRISPR/Cas
Figure 5.5. Workflow for CRISPR/Cas mediated gene targeting in mouse ES cells
Figure 5.6. Schematic Representation of Surveyor Assay.
Figure 5.7. Surveyor Assay on Tet1 and Tet2 in wild type and CRISPR/Cas targeted mES cells, treated with Puroymcin or mCherry sorted upon co-transfection
Figure 5.8. Schematic Representation of RFLP
Figure 5.9. RFLP Assay on WT and Tet1&Tet2 CRISPR co-targeted 32 clones
Figure 5.10. DNA sequencing results on Tet1&Tet2 CRISPR co-targeted loci
Figure 5.11. Effect of Tet1/Tet2 knockout on reprogramming
Figure 5.12. HP1α redistribution in mESxhF heterokaryons
Figure 6.1. Jarid2, Prickle1, Fzd2 and Wnt9a targeting by CRISPR/Cas
Figure 6.2. Surveyor Assay on Puromycin treated populations of Jarid2 single and Prickle1, Wnt9a, Fzd2 triple CRISPR/Cas targeted mES cells
Figure 6.3. Surveyor Assay on CRISPR/Cas targeted single mES cell clones for Jarid2
Figure 6.4. DNA sequencing results on Jarid2 CRISPR/Cas targeted locus
Figure 6.5. Western Blot detection of Jarid2 in wild type and CRISPR/Cas targeted clones
Figure 6.6. Surveyor Assay on CRISPR/Cas triple targeted single mES cell clones for Fzd2
Figure 6.7. Surveyor Assay on CRISPR/Cas triple targeted single mES cell clones for Prickle1 and Wnt9a
Figure 6.8. DNA sequencing results on Prickle1, Fzd2 and Wnt9a CRISPR/Cas co-targeted loci
Figure 6.9. mRNA levels of CRISPR/Cas targeted Prickle1, Fzd2 and Wnt9a in selected clones and wild type cells
Figure 6.10. mRNA levels of Prickle1, Fzd2 and Wnt9a in Jarid2 mutant lines and wild type cells
Figure 6.11. Flow Cytometry analysis of Nanog expression in mES cells Clones 12C and 2D
Figure 6.12. Off-target identification of CRISPR/Cas targets
Figure 7.1. Schematic representation of RNA-guided Cas9 targeting on DNA.
TABLES
Table 1 Primers for transcript analysis by quantitative RT-PCR
Table 2 Primers for bisulfite sequencing analysis
Table 3 Primers for enzyme protection assay
Table 4 Primers for ChIP assay
Table 5 Primers for genomic DNA amplification for Surveyor and RFLP Assays
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Abbreviations
2i Small molecule inhibitors of MEK and GSK3β 3C Chromosome Conformation Capture 5-caC 5-carboxycytosine 5-fC 5-formylcytosine 5-hmC 5-hydroxymethylcytosine 5-hmU 5-hydroxymethyluracil 5-mC 5-methylcytosine β-gal β-galactosidase µ Micro Ac acetyl group BER Base excision repair bp base pair CAGE Cap analysis of gene expression Cas CRISPR-associated cDNA Complementary DNA CGI Cytosine Guanine dinucleotide islands CHD Chromodomain helicase DNA-binding ChIP Chromatin immunoprecipitation CpG Cytosine Guanine dinucleotide Ct Threshold Cycle CRISPR clustered regularly interspaced short palindromic repeats Ctrl Control DAPI 4,6-diaminido-2-phenylindole DMEM Dulbecco’s Modified Eagle DMR Differentially methylated region DNA Deoxyribonucleic acid DNMT DNA methyltransferase DSB Double strand break E Embryonic day EC Embryonic carcinoma EDTA Ethylene diamine tetraacetic acid EdU 5-ethynyl-2-deoxyuridine EG Embryonic germ ES Embryonic stem FACS Fluorescence activated cell sorting FBS Fetal Bovine Serum FGF Fibroblast Growth Factor FRAP Fluorescence recovery after photobleaching g Gram GFP Green Fluorescent Protein H Histone hB human B HCP High CpG density promoter HDAC Histone deacetylase HDR Homology-directed break HP1α Heterochromatin Protein 1α HR Homologous recombination ICM Inner cell mass ICP Intermediate CpG density promoter
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ICR Imprinting control region Indel Insertion / Deletion iPS Induced pluripotent stem kb Kilo base K Lysine KRAB Krüppel associated box KO Knock-out l Litre LCP Low CpG density promoter LINE Long interspersed element LIF Leukemia Inhibitory Factor m Milli M Molar MBD Methyl binding domain Me methyl group MET mesenchymal-to-epithelial NEAA Non-essential aminoacids n Nano Neo Neomycin NHEJ Non-homologous end joining ntES Nuclear-transfer-embryonic stem NURD Nucleosome Remodelling Deacetylase PAM Protospacer Adjacent Motif PBS Phosphate Buffered Saline PCR Polymerase Chain Reaction PEG Polyethylene Glycol PGC Primordial germ cell PN Pronuclear PRC Polycomb repressor complex RFLP Restriction Fragment Length Polymorphism RNA Ribonucleic acid RNAi RNA interference RT Reverse Transcription S Serine SAM S-adenosyl-l-methionine SCNT Somatic-cell nuclear transfer sgRNA single guide RNA shRNA short hairpin RNA SNP Single nuclear Polymorphism SWI/SNF SWItch/Sucrose NonFermentable TF Transcription Factor TKO Triple Knock-out tracrRNA trans-activating crRNA U Unit WT Wild type X X-chromosome
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Chapter 1
Chapter 1. Introduction
The faithful transmittance of the genetic material from mother to daughter cells
ensures that every cell of a higher eukaryotic organism shares the same genomic
sequence, originated from a single fertilised egg. Yet, each cell type performs a discrete
function that depends on information decoded from the DNA. Cell fate is acquired in the
course of development, characterised by a distinct gene expression profile, and is
maintained by epigenetic mechanisms.
1.1. Pluripotent embryonic stem cells
Cells within the inner cell mass (ICM) of the embryo are pluripotent and possess
the ability to differentiate into all three germ layers. Although the ICM exists transiently in
the blastocysts during pre-implantation development, it is possible to derive cells from this
stage (Evans and Kaufman, 1981; Martin, 1981). These in-vitro counterparts are known
as embryonic stem (ES) cells, they are pluripotent and mouse ES cells can be indefinitely
propagated in culture on feeder embryonic fibroblasts or in the presence of leukaemia
inhibitory factor (LIF). Remarkably, once injected into the blastocysts, ES cells can give
rise to both adult somatic and germ cells following mouse chimera production. Although
human ES cells share main properties with mouse ES cells including self-renewal and
pluripotency, they are morphologically different, proliferate with slower kinetics, require
fibroblast growth factor 2 (FGF2) and Activin/Nodal pathway activity and proposed to
exhibit similar characteristics to mouse epiblast stem cells (Schnerch et al., 2010).
ES cell state is maintained by tight control over gene regulation with OCT4, SOX2
and NANOG providing the core transcription factors (TFs) that underwrite the pluripotency
network (Boyer et al., 2005). The equilibrium between pluripotency and differentiation is
established by the relative levels of TF expression. For example, a modest increase of
Oct4 expression triggers mesodermal and endodermal differentiation, while its decrease
directs differentiation towards a trophectoderm lineage (Niwa et al., 2000). On the other
hand, mono-allelic disruption of Oct4 results in increased genomic OCT4 protein
occupancy, maintenance of a stable pluripotent state and a delay in differentiation
(Karwacki-Neisius et al., 2013). Similarly, changes in SOX2 levels cause deregulation of
pluripotency (Kopp et al., 2008). Interestingly, ES cells regularly oscillate between high
and low Nanog expression levels, where the former state associates with efficient self-
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Chapter 1
renewal, while the latter is linked to a propensity to differentiate (Chambers et al., 2007).
Similar fluctuations have been observed for additional pluripotency network components
including Essrb, Klf4, Tbx3, Rex1 and Stella (Cahan and Daley, 2013). Homogeneous
high expression of pluripotency genes can be achieved by culturing mouse ES cells in the
presence of two small-molecule inhibitors (PD0325901 and CHIR99021, denoted as 2i)
that selectively target mitogen-activated protein kinase kinase (MEK) and glycogen
synthase kinase-3 (GSK3) (Wray et al., 2010; Ying et al., 2008). Moreover, mouse ES
cells cultured under 2i condition exhibit reduced levels of lineage-specific genes, reduced
bivalent (H3K27me3/H3K4me3) marking (see below) and genome-wide DNA
hypomethylation (see section 1.2) (Leitch et al., 2013; Marks et al., 2012). On this basis,
ES cells maintained in 2i conditions are proposed to exist in a ground state of pluripotency
(Wray et al., 2010).
The OCT4 interactome is composed of over 160 proteins, including pluripotency-
associated transcription factors SALL4, ESRRB, DAX1 and TCFCP2I1 and chromatin
modifiers, such as NURD and SWI/SNF complex components (van den Berg et al., 2010).
Acute depletion of OCT4 results in reduced recruitment of transcription factors on their
genomic targets, suggesting that OCT4 provides an anchor point for the assembly and
co-localisation of associated factors. Core transcription factors share a significant number
of common targets and can regulate their own expression, which may be a key concept
in pluripotent homeostasis (Boyer et al., 2005). Interestingly, genes that show a high level
of transcription factor occupancy tend to be active, while those bound by only a few
transcription-factors tend to be silent, implying that gene regulation is linked to
combinatorial localisation of pluripotency network components (Kim et al., 2008).
Moreover, core transcription factors and their interactors show preferential binding at
enhancers and NANOG-OCT4-SOX2 binding can recruit p300, a histone
acetyltransferase to the enhancer regions (Chen et al., 2008). These ES-cell specific
enhanceosomes can interact with the target gene promoters to induce gene expression
(Chen et al., 2008), and this interaction may be facilitated by mediator and cohesin
complexes forming a DNA loop structure, as demonstrated by chromosome conformation
capture (3C) technique (Kagey et al., 2010). Inspection of several ES-specific gene
enhancers has revealed that they comprise clusters of constituent enhancers spanning
large distances (as much as 50 kb). These so-called “super-enhancers” are enriched for
OCT4, SOX2, NANOG, KLF4 and ESRRB binding, and contain high levels of the Mediator
coactivator complex that is shown to be associated with general transcription factors and
RNA polymerase II (Whyte et al., 2013).
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Chapter 1
1.1.1. Chromatin dynamics in embryonic stem cells
The basic repeating unit of eukaryotic chromatin is the nucleosome, which consists
of a central histone octamer core (normally composed of two copies of each of the
histones H2A, H2B, H3 and H4) wrapped by a segment of ~147 base pairs of DNA and a
linker DNA associated with histone H1 (Luger et al., 1997). Gene expression activity
correlates with post-translational modifications of the histone residues that can impact
expression by two main mechanisms. The first one is via influencing the physical structure
of the chromatin, and the second by creating docking sites for the binding of effector
proteins (Bannister and Kouzarides, 2011). These modifications occur at the N-terminal
histone tails or within the globular domains on specific residues [such as lysine (K) or
serine (S)] and include acetylation, mono- di- and tri-methylation, phosphorylation and
ubiquitylation, that can crosstalk or fine-tune the transcriptional readout (Lee et al., 2010).
Originally termed by Emil Heitz in 1928 (Heitz, 1928) according to the staining profiles,
chromatin exists in two forms; densely packed, transcriptionally inactive heterochromatin
and de-condensed, transcription permissive euchromatin. Both structures can be
recognised by their signature histone marks; while H3K27me3 and H3K9me3 are
generally associated with heterochromatin, H3K4me3 and histone acetylation often
associate with euchromatin. Collectively, these histone modifications, and others, have
been postulated to establish a ‘histone code’ that forms the basis of epigenetic regulation
of gene expression (Jenuwein and Allis, 2001).
Embryonic stem cells have been reported to have a generalised ‘open chromatin
structure’, a feature that is also observed in the ICM, which gradually condenses in the
course of lineage commitment (Ahmed et al., 2010; Azuara et al., 2006; Meshorer et al.,
2006). Remarkably, core histones and structural proteins of the chromatin, including HP1,
are hyperdynamic in pluripotent cells, as demonstrated by sensitivity to salt extraction and
by fluorescence recovery after photobleaching (FRAP) (Bhattacharya et al., 2009;
Meshorer et al., 2006). Consistently, electron microscopy indicates that heterochromatic
regions are infrequent in ES cells, where H3K9me3 modification is underrepresented
(Efroni et al., 2008; Meshorer et al., 2006). In addition, histone H3K27me3, which is not
abundant in ES cells, is significantly increased upon ES cell differentiation (Hawkins et al.,
2010; Zhu et al., 2013), accompanied by reduced H3K9 acetylation levels (Krejci et al.,
2009). From a higher-order genome organization perspective, OCT4, SOX2 and NANOG
binding sites co-localise at three dimensional space establishing transcriptionally active
regions in the nucleus (Denholtz et al., 2013), while inactive regions form less long-range
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Chapter 1
interactions in ES cells than differentiated cells (de Wit et al., 2013). In pluripotency, an
open chromatin landscape may actively be maintained by chromatin remodelling factors
including CHD (chromodomain helicase DNA-binding) family members that can slide or
eject nucleosomes in an ATP-dependent manner to promote transcription (Clapier and
Cairns, 2009). In this regard, Chd1 knockdown in ES cells leads to accumulation of
heterochromatin, and disrupted differentiation capacity (Gaspar-maia et al., 2009).
1.1.2. Polycomb regulation and bivalency
Although ES cell chromatin is open and dynamic, expression of lineage-specific
genes must be restricted for the maintenance of pluripotency. Silencing of developmental
regulators in ES cells is primarily mediated by polycomb repressor complex (PRC)
proteins which exist in two sub-complexes. PRC1 catalyses the mono-ubiquitinylation of
histone H2A on position 119 (H2AK119ub) and PRC2 is responsible for H3K27me3
deposition (Boyer et al., 2006). It has recently been demonstrated that CXXC domain
containing proteins are involved in targeting PRC components for gene repression.
Notably, KDM2B has been shown to recruit non-canonical PRC1 components on CpG
island promoters by associating with unmethylated DNA via its CXXC domain, leading to
targeted mono-ubiquitinylation of H2AK119 (Farcas et al., 2012; Wu et al., 2013a). This
results in recruitment of PRC2 that in turn generate H3K27me3 for repression of gene
expression (Blackledge et al., 2014). Consequently, a positive feedback loop is
established, in which H3K27me3 further promotes binding of the canonical PRC1 on the
chromatin to expand the PRC silenced domain (Kalb et al., 2014). Recently, a Jumonji
histone demethylase family member JARID2 has been identified to associate with PRC2
and is critical for mouse development and ES cell differentiation (Landeira and Fisher,
2011). Differentiation defects observed in Jarid2-/- ES cells could reflect the lack of serine
5 phosphorylated RNA polymerase II enrichment at the bivalent domains (Landeira et al.,
2010). More recent analyses have revealed that, although catalytically inactive (Klose et
al., 2006) JARID2 exhibits nucleosome-binding activity and promotes PRC2 recruitment
to nucleosomes (Son et al., 2013), which is in part mediated by long noncoding RNAs
(Kaneko et al., 2014).
In ES cells, while developmentally regulated genes are silent, they possess the
capacity to be rapidly activated upon external differentiation signals. These silenced
lineage-specific regulators possess both repressive H3K27me3 and active H3K4me3
histone marks at their promoters, a property known as bivalency, which is thought to be
16
Chapter 1
important for fast gene reactivation (Azuara et al., 2006; Bernstein et al., 2006). Of
importance, bivalent gene promoters are occupied by serine 5 phosphorylated RNA
polymerase II, which marks transcription initiation, but not elongation, indicating that the
transcription is ‘poised’ but ready to be initiated at these genes (Brookes et al., 2012; Stock
et al., 2007).
1.2. DNA methylation dynamics in mammals
DNA methylation mainly involves cytosine residues. Cytosine methylation on the
fifth carbon (abbreviated as 5-mC) is an essential epigenetic mark for mammalian
development and homeostasis, and is often found in the context of symmetrical CpG
dinucleotides (Bird, 2002). While non-clustered CpGs are often methylated, clustered
CpGs (so called CpG islands, CGIs, which are on average 1000 base pairs long with high
C+G density) exhibit hypomethylation (Deaton and Bird, 2011). It is important to note that
CpG dinucleotide sequences outside CGIs are evolutionarily underrepresented in the
animal genome due to spontaneous deamination of methylcytosine to thymidine and
accumulation of TpG dinucleotides because of inaccurate mismatch repair (Bird, 1980;
Lander et al., 2001). DNA methylation is involved in key developmental processes that
involve regulation of gene expression, repetitive element silencing, X-inactivation and
imprinting (Smith and Meissner, 2013).
1.2.1. Maintenance and de novo establishment of DNA methylation
Methylation of the cytosine residues is catalysed by a group of DNA
methyltransferase (DNMT) enzymes; DNMT1 is responsible for maintenance, and
DNMT3A and DNMT3B mediate establishment of de novo methylation patterns (Goll and
Bestor, 2005). DNMT2, despite possessing sequence and structural characteristics of
DNA methyltransferases, methylates transfer RNA, but not DNA (Goll et al., 2006). A last
member, DNMT3L, is catalytically inactive and lacks DNA affinity, but stimulates DNMT3A
and 3B activities by direct interaction (Suetake et al., 2004).
Maintenance of DNA methylation is based on the recognition of hemi-methylated
CpGs, sites that are generated by incorporation of the unmodified cytosine into the newly
synthesised DNA during semiconservative DNA replication. DNMT1 can then methylate
these sites to propagate the information at every cell cycle. DNMT1 was the first DNA
17
Chapter 1
methyltransferase to be cloned and sequenced (Bestor et al., 1988), and its depletion
results in embryonic lethality accompanied with significantly reduced 5-mC levels in the
embryo (Li et al., 1992). Similarly, DNMT1 deficient ES cells undergo rapid genome-wide
demethylation, yet are viable and retain low level but stable DNA methylation patterns (Lei
et al., 1996). DNMT1 is highly abundant in mitotic cells (Kishikawa et al., 2003) and is
recruited to the replication foci during DNA replication (Leonhardt et al., 1992).
Recruitment is mediated by its tethering factor UHRF1 (also known as NP95), which
specifically recognises and binds hemi-methylated CpG sites (Bostick et al., 2007; Sharif
et al., 2007). This results in the flipping of the 5-mC out of the DNA helix that is positioned
in the SRA domain of UHRF1, which then correctly orients unmethylated cytosine to
DNMT1 for methylation (Arita et al., 2008; Avvakumov et al., 2008; Hashimoto et al.,
2008). In addition, UHRF1 can recognise H3K9me2/me3 marks, which is proposed to
enhance DNMT1 recruitment and DNA methylation (Liu et al., 2013). Of importance, Uhrf1
deletion results in substantially decreased DNA methylation levels due to lack of
maintenance and Uhrf1 deficiency causes embryonic lethalily (Sharif et al., 2007).
DNMT3A and DNMT3B, the de novo methyltransferases, are required for
establishment of DNA methylation patterns in early mammalian development (Okano et
al., 1998, 1999). DNMT3B was first shown to specifically methylate pericentromeric
regions (Okano et al., 1999) and is expressed in the ICM, epiblast and the embryonic
ectoderm, while DNMT3A is not detected at these stages (Watanabe et al., 2002). Instead
Dnmt3a is expressed between E10.5-E14.5 embryos (Watanabe et al., 2002), partly
related to its role in the establishment of Imprint Control Regions (ICRs) (Kaneda et al.,
2004). Although both proteins share homology, deletion of Dnmt3b results in embryonic
death, while Dnmt3a-null mice die after birth (Okano et al., 1999). However, combined
deletion of both genes causes earlier embryonic lethality, revealing some level of partial
redundancy between the two enzymes and overlapping functions during embryogenesis
(Okano et al., 1999). In ES cells that lack both DNMT3A and 3B, loss of methylation is
gradual and requires long term passaging (Chen et al., 2003). In addition,
Dnmt1/Dnmt3a/Dnmt3b triple KO (TKO) ES cells exhibit extensive loss of CpG
methylation, but retain their self-renewal capacity and undifferentiated state, suggesting
that in ES cells other epigenetic mechanisms are sufficient to maintain stable
heterochromatin and chromosome structures (Tsumura et al., 2006). However, although
TKO cells can contribute to the blastocyst stage and differentiate into extraembryonic
tissues in vivo, they do not contribute to the embryonic lineages (Sakaue et al., 2010).
18
Chapter 1
1.2.2. Roles of DNA methylation in gene regulation
CpG islands constitute essential platforms for DNA methylation to exert its role as
transcriptional repressor, in combination with chromatin modifications (Deaton and Bird,
2011). Around 70% of human gene promoters are associated with CpG islands (Saxonov
et al., 2006). These high CpG density promoters (HCPs) are involved in regulation of
housekeeping as well as key developmental genes, and are generally free of DNA
methylation (Saxonov et al., 2006). This is evidenced by cap analysis of gene expression
(CAGE) technique that revealed the correlation between initiation of transcription and
presence of CGIs (Carninci et al., 2006). It is important to mention that CGI promoters can
be methylated and methylation provides long-term stabilization of silencing, including
imprinted and inactive X-chromosome genes. Two additional classes are intermediate and
low CpG density promoters (ICPs and LCPs, respectively), which exhibit more frequent
DNA methylation (Weber et al., 2007). Although LCPs are generally methylated,
methylation does not have an influence on gene expression, which remains active (Borgel
et al., 2010; Weber et al., 2007). On the other hand, ICP methylation results in gene
silencing, examples of which include repression of pluripotency-associated and germline
specific genes during differentiation (Borgel et al., 2010; Farthing et al., 2008; Meissner et
al., 2008).
CGI promoters are enriched for H3K4me3, a chromatin mark of actively transcribed
genes. However it is important to note that not all of the analysed promoters carrying this
mark are active (Guenther et al., 2007; Mikkelsen et al., 2007). Association of H3K4me3
with the CGIs is in part mediated by CFP1, which binds to the unmethylated CpGs upon
its interaction with SETD1, a H3K4 methyltransferase (Clouaire et al., 2012; Thomson et
al., 2010). H3K4me3 and histone variant H2A.Z, which also mark nucleosome depleted
transcription start sites, protect promoters from acquisition of de novo methylation (Ooi et
al., 2007; Zilberman et al., 2008). Similarly, KDM2A binding to unmodified CpGs in CGIs
results in depletion of di-methylated H3K36, necessary for a permissive environment for
transcriptional machinery (Blackledge et al., 2010). On the other hand, another H3K36
demethylase ,KDM2B, is required for recruiting PRC1 to the unmethylated CGIs and is
associated with silencing of genes involved in embryonic development and cellular
differentiation in ES cells (as described earlier, (Farcas et al., 2012)).
Gene repression mediated by CpG methylation can be attributed to the
combinatorial effects of inhibition of transcription factor binding and recruitment of
chromatin re-modellers by methyl-binding domain (MBD) proteins (Klose and Bird, 2006).
19
Chapter 1
However, it is speculated that gene silencing precedes DNA methylation, which then acts
as a lock to stabilise the repressed status (Jones, 2012). For example, silencing of Oct4
and Nanog genes during carcinoma cell differentiation is accompanied with chromatin
remodelling of regulatory sequences that are subsequently methylated by DNMT3A (You
et al., 2011). This is in agreement with the fact that G9A, a H3K9 di-methyltransferase,
promotes de novo methylation during differentiation, suggesting that DNA methylation
follows changes in chromatin structure and histone marks (Dong et al., 2008; Epsztejn-
Litman et al., 2008; Tachibana et al., 2008). For example, ectopic targeting of HP1α at the
Oct4 locus in ES cells results in H3K9 tri-methylation and gene silencing that are later
followed by DNA methylation (Hathaway et al., 2012). Furthermore, H3K9 tri-methylation
is an early event in X-inactivation leading to gene repression, which precedes CGI
methylation (Mermoud et al., 2002). Histone marks around methylated CpGs are also
important for mitotic inheritance of DNA methylation. This is exemplified by the
requirement of UHRF1 binding to the methylated H3K9 to enable the recruitment of DNA
methylation maintenance machinery (Rothbart et al., 2012). Therefore, DNA methylation
can be considered as a provider of high-fidelity epigenetic memory by stabilising gene
repression.
1.2.3. Genomic Imprinting
Genomic imprinting ensures mono-allelic gene expression that is dependent on the
parental origin. DNA methylation is the major epigenetic component of genomic imprinting
and marks differentially methylated regions (DMRs) in the genome. These DMRs are
established during gametogenesis (denoted as germline DMR) or after fertilization
(denoted as somatic DMRs) and can functionally control expression of imprinted genes in
cis within clusters, acting as imprinting control regions (ICRs) (Ferguson-Smith, 2011).
While the majority of methylated germline DMRs are maternally inherited, thus far only
four germline DMRs, H19-Igf2, Dlk1, Rasgrf1, and Zdbf2, are characterised as paternally
silenced (John and Lefebvre, 2011). Since the initial discovery of imprinted Igf2r, Igf2 and
H19 loci (Barlow et al., 1991; Bartolomei et al., 1991; DeChiara et al., 1991; Ferguson-
Smith and Cattanach, 1991) more than a hundred genes in mouse and human have been
identified as imprinted (Henckel and Arnaud, 2010). Genomic imprinting is involved in
many processes such as embryonic growth (Smith et al., 2006a), placental development
(Frost and Moore, 2010), metabolism (Radford et al., 2011) and behaviour (Wilkinson et
20
Chapter 1
al., 2007), and perturbations are linked to several human disorders including Prader-Willi
and Angelman syndromes (Butler, 2009).
For new gamete-specific methylation to be established in the germline, parental-
specific DNA methylation marks at the DMRs must first be erased. This occurs during
proliferation and migration of the primordial germ cells (PGCs) to the gonads and is
completed by embryonic day 13.5 in both male and female mouse embryos (Guibert et
al., 2012; Hajkova et al., 2002; Popp et al., 2010; Seisenberger et al., 2012). At that point
parent-of-origin-specific de novo germline DMR methylation starts and is completed during
the early neonatal period in females and late foetal development in males (Hajkova et al.,
2008; Lucifero et al., 2004). During this period, DNA methyltransferase DNMT3A is
recruited at DMRs via interaction with DNA methyltransferase-like protein DNMT3L, and
establishes de novo methylation of both maternal and paternal imprinted loci (Hata et al.,
2002; Kaneda et al., 2004). DMR methylation is then maintained through DNA replication
by DNMT1.
One important difference between maternal and paternal DMRs is their genomic
location; while maternal DMRs are intragenic and are mostly found at transcription start
sites of protein-coding or non-coding RNA genes, paternal DMRs are intergenic (Edwards
and Ferguson-Smith, 2007). In male foetal germ cells, promoter associated maternal
DMRs are transcriptionally active and are enriched for H3K4me3 (Henckel et al., 2012).
Mechanistically, interaction of DNMT3L with the DNA is strongly inhibited by this histone
modification thus preventing the recruitment of DNMT3A for de novo methylation (Ooi et
al., 2007). This correlates with the requirement of KDM1B, a H3K4 demethylase that is
highly expressed in oocytes, for proper establishment of DNA methylation at several
maternal DMRs (Ciccone et al., 2009). Many other histone marks and trans-acting
elements are collectively involved in the establishment of imprinting, and consequent
designation of actual parental-specific imprinting occurs during the global DNA
demethylation wave that occurs soon after fertilization (Kelsey and Feil, 2013).
Global erasure of DNA methylation happens shortly after fertilization in both
parental pronuclei. This global DNA demethylation is thought to be necessary for the
acquisition of totipotency and the establishment of the developmental programme. This
occurs asymmetrically; CpG methylation levels are rapidly reduced in the paternal
pronucleus at the one-cell stage, while the maternal pronucleus undergoes a gradual loss
of DNA methylation throughout pre-implantation development (Santos et al., 2002). At this
stage, imprinted genes must be selectively protected from the global wave of DNA
demethylation. It has recently demonstrated that paternal pronucleus undergoes
21
Chapter 1
extensive TET3-mediated CpG hydroxylation (Gu et al., 2011; Iqbal et al., 2011; Wossidlo
et al., 2011), while the maternal pronucleus remains protected from TET3 activity by a
maternal factor, STELLA (also known as PGC7 or DPPA3) (Nakamura et al., 2012). In
addition, STELLA also protects maternal germline DMRs at Peg1, Peg3, Peg10 loci as
well as paternal germline DMRs at H19 and Rasgrf1 loci from demethylation (Nakamura
et al., 2007). However, STELLA’s specificity for imprinted genes is questionable as its
protective function is genome-wide (Nakamura et al., 2012). ZFP57, a zinc finger protein
with a KRAB (Krüppel associated box) domain, stands as a strong candidate for
imprinting-associated maintenance and its deficiency in mouse embryos results in
hypomethylation of both parental germline DMRs (Li et al., 2008). ZFP57 binds methylated
alleles of germline DMRs in mouse ES cells, via sequence-specific recognition of a
methylated hexanucleotide motif (TGCCGC) found in all murine ICRs (Quenneville et al.,
2011). ZFP57 belongs to a family of proteins that interact with TRIM28 (also known as
KRAB-associated protein 1, KAP1) a component of a multifunctional repressive complex,
which in turn brings repressive histone marks and DNA methyltransferases
(DNMT1/3A/3B) on to the zinc finger bound DNA (Iyengar and Farnham, 2011). Indeed,
ChIP experiments in mouse ES cells revealed co-localisation of ZFP57, TRIM28 and
H3K9me3 at methylated alleles of ICRs (Quenneville et al., 2011). Moreover, maternal
loss of Trim28 leads to ICR hypomethylation and embryonic lethality (Messerschmidt et
al., 2012). Further investigation of DNA methylation on each individual cell of the 8-cell
blastomeres revealed that maternal TRIM28 deficiency causes mosaic demethylation at
the DMRs thus confirming its importance in pre-implantation development
(Lorthongpanich et al., 2013). Interaction of TRIM28 with ZFP57 leads to sequence-
specific recruitment of the DNA methylation maintenance machinery to the imprinted
genes (Zuo et al., 2012) and therefore ensures protection of methylation at imprinted loci
during pre-implantation embryo development (Messerschmidt, 2012).
Unmethylated alleles of DMRs must also be protected from de novo methylation.
One example is the association of zinc finger protein CTCF with H19 DMR. By specifically
binding onto the unmethylated CTCF binding sites in the maternal allele, CTCF brings the
H19 enhancer and promoter together (Murrell et al., 2004) to ensure transcriptional activity
that in turn prevents acquisition of DNA methylation (Engel et al., 2006). In addition, loop
formation at this region, stabilized by Cohesin (Nativio et al., 2009), prevents the
enhancers interacting with the promoter of Igf2 gene (which is therefore not activated on
the maternal allele). As CTCF does not bind to the methylated H19 DMR in the paternal
allele, Igf2 gene remains active via interaction between enhancers (Nativio et al., 2009).
22
Chapter 1
Early nuclear transplantation studies have demonstrated that mouse embryos that
contain biparental gynogenones (two female pronuclei) or biparental androgenones (two
male pronuclei) failed to undergo successful embryogenesis (McGrath and Solter, 1984;
Surani et al., 1984). This shows the importance and requirement of one paternal and one
maternal genome for normal development. Although the origins and evolution of genomic
imprinting are still being debated (Patten et al., 2014), it constitutes an ideal model system
to study epigenetic mechanisms and their interplay.
1.2.4. DNA demethylation
DNA methylation is a vital epigenetic modification in mammalian development.
Nonetheless, despite its chemical and hereditary stability, rapid loss of methylation is
observed in the zygote and during PGC development. Understanding the cause and
consequences of this vital switch is essential for elucidating the molecular details of life
cycle.
The sixth carbon of a cytosine nucleotide plays an important role in the chemistry
of methylation as it has electrophilic characteristics and it is therefore prone to be attacked
by nucleophilic cysteine thiolate, which is found on the catalytic site of DNA
methyltransferases. Enzyme binding on the sixth carbon of the cytosine leads to the
activation and increased nucleophilicity of the fifth carbon. Upon induction, the fifth carbon
receives an electrophilic attack mediated by the cofactor S-adenosyl-l-methionine (SAM),
which transfers a methyl group. This process is known as cytosine methylation, and its
stability arises from the established carbon-carbon bond (Smith et al., 1992). For this
reason, direct removal of the methyl group has been speculated to be energetically
unfavourable (Ooi and Bestor, 2008). Yet, especially in the past decade, several
mechanisms that mediate loss of DNA methylation have been reported, and are grouped
into two major pathways: (1) passive demethylation, via DNA replication-mediated dilution
of 5-methylcytosine; (2) active demethylation, via replication-independent enzymatic
activities. Interestingly, recent investigations have also revealed that both pathways can
co-exist and lead to loss of DNA methylation.
1.2.4.1. Passive demethylation
There are several ways for the passive DNA demethylation to occur: inhibition,
downregulation or exclusion of maintenance machinery, which is composed of DNMT1
23
Chapter 1
and UHRF1, or further modification of 5-mC that impairs its recognition (Figure 1.1). As a
consequence, upon DNA replication, the nascent cytosine that is incorporated into the
newly synthesized strand remains unmodified. One pharmacological implication of
passive demethylation is the usage of 5-azacytidine for treatment of diseases that stem
from hyper-methylation. 5-azacytidine is a cytosine derivative, where the fifth carbon atom
is substituted by a nitrogen atom, and can be incorporated into the newly synthesized
DNA. Although DNMT1 can recognise 5-azacytidine, it gets trapped on the DNA due to
chemically unresolved methylation reaction. This triggers DNA damage signalling
resulting in degradation of DNMT1. Therefore lack of nascent cytosine methylation leads
to gradual loss of DNA methylation via cell division (Stresemann and Lyko, 2008). Passive
demethylation has been described both in plants and animals. In plants, downregulation
of Dnmt1 homologue MET1 is essential for activation of imprinted genes during female
gametogenesis (Jullien et al., 2008). MET1 downregulation also in part explains activation
of transposable elements in the endosperm of Arabidopsis Thaliana (Slotkin et al., 2009).
Figure 1.1. Maintenance/replication-coupled loss of DNA methylation. (A) DNA replication at the methylated CpG sites results in hemimethylated DNA, which is restored to full methylation by maintenance machinery composed of UHRF1 and DNMT1. Downregulation, inhibition, or nuclear exclusion of UHRF1 and/or DNMT1 leads to replication-coupled dilution of 5-mC mark as incorporated cytosine to the newly synthesized DNA remains unmodified. (B) Iterative oxidation of the 5-mC mark prevents its recognition by the maintenance machinery, which similarly leads to progressive loss of methylation.
Similarly, nuclear exclusion and/or transcriptional repression of UHRF1 (which is the
tethering factor of DNMT1) during mouse germ cell development has been suggested to
24
Chapter 1
result in genome-wide DNA demethylation (Kagiwada et al., 2013; Seisenberger et al.,
2012). Moreover, nuclear exclusion of maintenance machinery causes progressive loss
of methylation levels in the mouse maternal DNA soon after fertilization (Cardoso and
Leonhardt, 1999). These examples indicate that passive demethylation is a common
mechanism to achieve relatively rapid and efficient resetting of global methylation
programme.
1.2.4.2. Active demethylation
Methylation-dependent binding protein MBD2 was initially proposed to be able to
directly remove the methyl group of the 5-methylcytosine in-vitro (Bhattacharya et al.,
1999; Ramchandani et al., 1999) and in-vivo (Cervoni and Szyf, 2001). Nevertheless, this
suggestion proved to be unreliable as the results were not reproduced in subsequent
studies, and mice lacking MBD2 showed normal methylation patterns (Hendrich et al.,
2001; Santos et al., 2002). Since then, although no protein with such enzymatic activity
has been identified, multi-layered enzymatic pathways that could lead to DNA
demethylation have been proposed, as outlined below (Ooi and Bestor, 2008).
Active demethylation is well characterized in plants. DNA glycosylases DME,
ROS1, DML2 and DML3 are involved in DNA demethylation of imprinting, silenced
transgenes as well as specific genomic loci. These enzymes can recognise and remove
5-mC from double stranded DNA irrespective of sequence context. They additionally
possess apyrimidinic lyase activity to cut off the abasic site that is left after 5-mC excision.
A nascent cytosine is then introduced to the excision site via DNA polymerase and DNA
ligase enzymes that are components of base excision repair (BER) pathway (Ikeda and
Kinoshita, 2009).
Mammalian orthologs of such DNA glycosylases have not yet been identified and
a universal pathway that is responsible for DNA demethylation has not been found. In
recent years, studies in many model systems have led to several different hypotheses with
contrasting mechanisms. BER pathway constitutes one of the proposed mechanisms for
active DNA demethylation in mammals (Wu and Zhang, 2010). For example, BER
components were shown to be recruited to the paternal pronucleus of the zygote where
DNA breaks occur after fertilization (Hajkova et al., 2010; Wossidlo et al., 2010). BER-
mediated DNA repair results in incorporation of unmodified cytosine, and the inhibition of
BER components such as PARP1 and APE1 was shown to interfere with loss of DNA
methylation (Hajkova et al., 2010). PARP1 was also reported to be part of active DNA
25
Chapter 1
demethylation in PGCs in DNA damage-dependent and –independent ways (Ciccarone
et al., 2012; Hajkova et al., 2010; Kawasaki et al., 2014). A screening in Xenopus
suggested that Gadd45a is a key regulator of active DNA demethylation in non-dividing
cells, by promoting DNA repair (Barreto et al., 2007). The link between BER and GADD45
was further elucidated in Danio rerio (Zebrafish) (Rai et al., 2008). GADD45 lacks
enzymatic activity, but in this study it was demonstrated to interact with MBD4, a thymine
glycosylase, and DNA deaminases AID (activation-induced deaminase) and APOBEC
(apolipoprotein B mRNA editing catalytic polypeptide) family members such as
APOBEC2A and APOBEC2B. According to the suggested model, 5-mC deamination
gives rise to a T:G mismatch, which is then repaired by MBD4-mediated removal of
thymine and incorporation of an unmodified cytosine (Rai et al., 2008). GADD45A was
also shown to interact with AID and thymine DNA glycosylase (TDG) that collectively
exhibit active DNA demethylation activity in a similar two-step model (Cortellino et al.,
2011). Interestingly, DNMT3A and 3B were reported to exhibit 5-mC deaminase activity
for the establishment of a dynamic TDG-coupled DNA demethylation/methylation process
upon external stimuli (Kangaspeska et al., 2008; Métivier et al., 2008). It should be noted
that although Mbd4 knockout mice are viable and fertile (Millar et al., 2002), Tdg deficiency
is associated with embryonic lethality (Cortellino et al., 2011). In the meantime, conflicting
results were reported regarding the involvement of GADD45A in active DNA
demethylation (Jin et al., 2008). Furthermore, knock-out mice for Gadd45a and its
homologous Gadd45b are viable and show no significant change in methylation levels
(Engel et al., 2009; Ma et al., 2009). However this does not rule out the possibility of
redundant roles of Gadd45 members (Niehrs and Schäfer, 2012). Although GADD45 is
not involved in global control of DNA demethylation, site-specific Gadd45-mediated DNA
demethylation may be induced upon various external stimuli, including neurogenesis in
adult mouse brain (Ma et al., 2009). Gene-specific loss of methylation mediated by
GADD45A were further demonstrated upon targeted recruitment by inhibitor of growth
protein 1 (ING) (Schäfer et al., 2013) and by long non-coding RNA TARID (Arab et al.,
2014).
The mechanism of AID-mediated deamination of 5-mC to thymine as an
intermediate for DNA demethylation was suggested to occur in PGCs, where AID
deficiency resulted in less efficient loss of DNA methylation compared to wild type mice
(Popp et al., 2010). The fact that AID knock-out mice are viable (Muramatsu et al., 2000)
suggests the presence of redundant mechanisms of DNA demethylation exerted by
APOBEC family members (Popp et al., 2010) or by other pathways (see below).
26
Chapter 1
Interestingly, AID was shown to be essential for experimental cell fusion-based
reprogramming, especially for demethylation and transcriptional reactivation of
pluripotency-associated genes OCT4 and NANOG (Bhutani et al., 2010). However, this
finding was later revoked by Foshay et. al., who demonstrated that Aid was not expressed
in the cell lines used in the former study. Furthermore, the authors reported that Aid over-
expression did not alter the reprogramming kinetics nor changed the DNA demethylation
kinetics after cell fusion (Foshay et al., 2012). Similar results have recently been shown
regarding the role of AID in induced reprogramming. While one report suggested that
acute loss of AID affects reprogramming efficiency (Bhutani et al., 2013), subsequent
reports could not reproduce this finding, and suggested that AID is dispensable for early
reprogramming of iPS cells (Habib et al., 2014; Shimamoto et al., 2014), but might be
required later for the stabilisation of the pluripotency programme (Kumar et al., 2013). It is
worth mentioning that biochemical analysis of AID/APOBEC activity revealed a
substantially lower propensity (~10-fold) for 5-mC deamination relative to their canonical
substrate, cytosine (Nabel et al., 2012). Moreover, AID/APOBEC requires single-stranded
DNA for efficient deamination that has not been found in the investigated model systems
(Bransteitter et al., 2003). Therefore, further investigation is necessary to clarify the
intermediary effect of AID/APOBEC mediated 5-mC deamination for active DNA
demethylation.
1.2.4.3. TET protein mediated 5-mC oxidation in passive and active demethylation
As described above, several mechanisms for DNA demethylation have been
postulated in the past, although a consensus mechanism has not been evident. This may
be because certain mechanisms are restricted to particular biological settings, or that
conflicting results are genuinely unreliable (Wu and Zhang, 2014, 2010). However, the
recent discovery of 5-hydroxymethylcytosine (5-hmC) as a proposed intermediate for DNA
demethylation, has captivated attention (Pastor et al., 2013).
In 2009, two laboratories independently demonstrated the presence of 5-hmC in
the mammalian genome (Kriaucionis and Heintz, 2009; Tahiliani et al., 2009). The
presence of an “unusual” nucleotide in mammals was first observed in mouse Purkinje
and granule cell nuclei by thin layer chromatography (found in ~0.6% and ~0.2% of all CG
dinucleotides, respectively), and was confirmed to be 5-hmC, the oxidative derivative of
5-mC, by mass spectrometry (Kriaucionis and Heintz, 2009). Nucleotide oxidation was
previously characterised in Trypanasome brucei, where J-binding proteins (JBPs)
27
Chapter 1
catalyse thymine oxidation into 5-hydroxymethyluracil (Borst and Sabatini, 2008). In a
computational screen for mammalian homologs of JBPs, Tahiliani et al. identified three
paralogous human proteins TET1, TET2 and TET3 (Tahiliani et al., 2009). The authors
further demonstrated that TET1 overexpression resulted in decreased genomic 5-mC
levels, and that this was due to TET-mediated 5-mC conversion into 5-hmC (Tahiliani et
al., 2009). Similarly, mouse TET proteins (TET1-3) were shown to catalyse 5-hmC
production (Ito et al., 2010). Following studies reported that TET proteins can further
oxidize 5-hmC to yield 5-formylcytosine (5-fC) and 5-carboxycytosine (5-caC) (~15 5-fC
and ~3 5-caC for every 103 5-hmC) (He et al., 2011; Ito et al., 2011; Pfaffeneder et al.,
2011). Structurally, TET proteins contain CXXC-type zinc finger domain that has affinity
for clustered CpG sites, and a carboxy-terminal catalytic domain that provides Fe(II) and
2-oxoglutarate-dependent dioxygenase activity (Iyer et al., 2009). In jawed vertebrates
TET2 underwent chromosomal gene inversion, which resulted in detachment of the exon
containing the CXXC domain that became an independent gene encoding IDAX (inhibition
of the Dvl and axin complex) protein (Iyer et al., 2009; Ko et al., 2013). Interestingly, TET
proteins can target methylated and hemimethylated DNA in a CpG or non-CpG context
for catalytic oxidation (Ficz et al., 2011; Tahiliani et al., 2009)
A broad analysis in mouse tissues revealed that 0.03% to 0.69% of cytosines are
hydroxymethylated, with highest levels in the brain tissues (Globisch et al., 2010).
Similarly, pluripotent embryonic stem cells contain significant levels of 5-hmC, attributed
to the presence of TET1 and TET2 (Ito et al., 2010; Koh et al., 2011; Tahiliani et al., 2009).
Upon retinoic acid-mediated ES cell differentiation, Tet1 and Tet2 genes are
downregulated, accompanied by the upregulation of Tet3, suggesting differential and
developmental regulation of Tet genes (Koh et al., 2011). Indeed, during pre-implantation
development while Tet3 expression is limited to the zygote, Tet1 and Tet2 expression
programme is initiated in two cell embryos (Wossidlo et al., 2011) and maintained in the
inner cell mass of the blastocysts (Ito et al., 2010). Moreover, Tet1 and Tet2 are expressed
in the PGCs, whereas Tet3 is found particularly in the somatic cells during PGC
development (Yamaguchi et al., 2012). This dynamic control can in part be explained by
the presence of large number of pluripotency-associated transcription factor binding sites
at the Tet promoter regions (Ficz et al., 2011) and can be exemplified by OCT4-SOX2
complex-mediated regulation of Tet1 and Tet2 expression in mouse ES cells (Koh et al.,
2011).
To elucidate the relationship between pluripotency and 5-mC oxidation, several
studies have focused on genome-wide mapping of 5-hmC and TET binding in mouse ES
28
Chapter 1
cells. Due to the presence of CXXC domain, TET1 preferentially associates with CpG
islands found at the promoter regions of actively transcribed genes, co-localising with
H3K4me3, yet no enrichment of 5-hmC is observed at these loci (Williams et al., 2011;
Wu et al., 2011; Xu et al., 2011). Confirming this, TET1 depletion results in 5-mC
accumulation at many TET1-associated regions (Wu et al., 2011; Xu et al., 2011).
Interestingly, both TET1 and 5-hmC co-localise at Polycomb repressed CpG-rich bivalent
gene promoters, and this is proposed to protect these regions from the acquisition of de
novo DNA methylation, providing an additional layer of epigenetic control over lineage
commitment (Pastor et al., 2011; Williams et al., 2011; Wu et al., 2011). It should be noted
that, TET1 depletion results in similar levels of upregulated and downregulated genes (Wu
et al., 2011). This suggests that in addition to its catalytic role, TET1-mediated gene
regulation may take place in a catalytic activity-independent manner. In this regard TET1
was demonstrated to recruit SIN3A (Deplus et al., 2013; Vella et al., 2013; Williams et al.,
2011) and MBD3-NURD repressor complexes (Yildirim et al., 2011). In addition, recent
studies have documented TET protein interaction with O-linked N-acetylglucosamine (O-
GlcNAc) transferase (OGT) enzyme, which catalyses GlcNAc addition to serine/threonine
residues of proteins including histones. It was also revealed that TET mediated OGT
recruitment is associated with positive regulation of gene expression (Balasubramani and
Rao, 2013; Chen et al., 2013b; Deplus et al., 2013; Vella et al., 2013). Furthermore, most
of the active promoters that contain high CpG density are devoid of 5-hmC, 5-fC and 5-
caC modifications (Shen et al., 2013; Wu et al., 2011; Yu et al., 2012). Enrichment of TET1
in these regions as well as in active distal enhancers suggests that TET1 might be acting
as a safe-guard machinery to remove any randomly occurring de novo methylation (Shen
et al., 2013). On the other hand, 5-hmC is mainly found at the promoters that contain lower
CpG densities, bearing lower transcriptional activity (Yu et al., 2012). Similarly, 5-hmC is
enriched at poised enhancers, which may require rapid DNA demethylation during lineage
specification (Shen et al., 2013). On the contrary, a strong link exists between gene body
enrichment of 5-hmC and active transcription. TET2 protein, which lacks the CXXC
domain, has been shown to be responsible for 5-hmC deposition at these regions of the
active genes (Huang et al., 2014), as confirmed by TET2 depletion (Chen et al., 2013b).
Oxidation of 5-mC has also been linked to regulation of gene expression,
independent of DNA demethylation. DNA methylation is recognised by methyl binding
domain proteins, which recruit chromatin remodellers to prevent gene expression (Klose
and Bird, 2006). Oxidation of 5-methylcytosine strongly inhibits association of MBDs
(including MBD1, MBD2 and MBD4) with the DNA that in turn may positively affect
29
Chapter 1
transcription (Jin et al., 2010; Valinluck et al., 2004). Interestingly, although methyl-CpG-
binding protein 2 (MeCP2) associates with 5-mC to exhibit repressive functions, it can
bind 5-hmC with similar high affinity and this association facilitates transcription in neurons
(Mellén et al., 2012). On the other hand, in-vitro binding assays revealed that MBD3 is
able to associate with 5-hmC but not 5-mC, which may lead to recruitment of repressive
NURD complex for negative regulation of transcription (Yildirim et al., 2011). However this
finding was not confirmed in a later report, where readers of 5-mC and its oxidized
derivatives were screened by unbiased quantitative mass spectroscopy (Spruijt et al.,
2013). It was further demonstrated that there is a limited overlap between readers (which
are expressed in a cell type specific manner) of each of the 5-mC, 5-hmC, 5-fC and 5-
caC, suggesting individual and cell specific roles of such epigenetic modifications (Spruijt
et al., 2013). Therefore, although a general mechanism has not yet been elucidated,
growing evidence indicates that relative stability of 5-hmC in the genome can influence
gene expression.
5-hmC and passive demethylation
The major excitement of 5-mC oxidation stems from the fact that 5-hmC can act as
an intermediate of DNA demethylation, involved in both DNA replication-dependent and
independent pathways (Figures 1.1 and 1.2). DNA methylation is stably inherited via
collaborative action of DNMT1 with its partner UHRF1 (as described earlier). However
modifications on the methylated cytosine may interfere with the maintenance machinery.
A previous study demonstrated that oxidation of 5-mC prevents methylation of the newly
synthesised DNA, resulting in heritable changes in DNA methylation patterns (Valinluck
and Sowers, 2007). However, whether this is due to the inability of UHRF1 in recognition
of the hemi-hyrdoxymethylated CpG sites has been controversial. While Frauer et al.
reported that UHRF1 can bind to hemi-methylated and hemi-hydroxymethylated DNA with
similar affinities, Hashimoto et al. demonstrated that the efficiency is tenfold less in the
latter case (Frauer et al., 2011; Hashimoto et al., 2012). Nevertheless, DNMT1 is
significantly inefficient in catalytic methylation of the unmodified cytosine in hemi-
hydroxymethlated DNA (60-fold less compared to hemimethylated DNA) (Hashimoto et
al., 2012; Otani et al., 2013). This shows that 5-mC oxidation coupled passive dilution can
occur even in the presence of maintenance machinery. It should be noted that in cells
where TET activity is high (such as neurons and ES cells), 5-mC and 5-hmC levels may
30
Chapter 1
be necessary to be stably maintained. In this regard, it has been shown that de novo
methyltransferases DNMT3A and 3B can efficiently methylate hemi-methylated and hemi-
hydroxymethylated CpG sites (Hashimoto et al., 2012; Otani et al., 2013).
5-hmC and active demethylation
Oxidative derivatives of 5-mC have recently been suggested to be the
intermediates of replication-independent DNA demethylation. Several enzymatic
pathways have been proposed for this process, with a particular emphasis on BER.
The first DNA-repair based pathway that has been proposed is the one involving
AID/APOBEC family members (Figure 1.2). Guo et al. demonstrated that 5-hmC levels
induced by TET1 in HEK293 cells were significantly reduced upon AID over-expression.
In addition, AID and some of the APOBEC family members were able to demethylate
reporter DNA in the presence of TET1. According to the authors, demethylation pathway
first involves TET mediated hydroxylation of 5-mC that is followed by deamination to yield
5-hydroxymethyluracil (5-hmU). The resulting 5-hmU:G mismatch is then repaired by BER
components and 5-hmU is replaced by unmodified cytosine (Guo et al., 2011). In line with
this report, Cortellino et al. demonstrated that SMUG1 (single-strand-selective
monofunctional uracil DNA glycosylase 1) and TDG glycosylases interact with AID and
perform in-vitro 5-hmU removal on double stranded DNA (Cortellino et al., 2011). However
subsequent studies showed conflicting evidence on AID/APOBEC-mediated 5-hmU
generation. Firstly, no enrichment of 5-hmU was detected in mammalian cells using
sensitive mass spectroscopy, suggesting that either this is a very short-lived nucleotide
base or deamination of 5-hmC does not occur (Globisch et al., 2010; Pfaffeneder et al.,
2011). Secondly, although AID/APOBEC members have significantly reduced activity
towards 5-mC, no activity for 5-hmC was detected neither in vitro nor upon overexpression
in cells (Nabel et al., 2012). Mechanistically, AID does not show affinity on 5-hmC due to
steric hindrance and increased electron cloud size (Rangam et al., 2012). Therefore,
involvement of AID/APOBEC deaminase activity in 5-hmC associated DNA demethylation
seems unlikely to occur.
The second DNA repair-mediated pathway involved in active DNA demethylation
is based on TET ability to perform iterative 5-mC oxidation that results in the formation of
5-fC and 5-caC (Figure 1.2) (He et al., 2011; Ito et al., 2011; Pfaffeneder et al., 2011). 5-
fC and 5-caC can be removed by TDG glycosylase activity and this will result in DNA
31
Chapter 1
demethylation upon incorporation of unmodified cytosine by the BER components (He et
al., 2011; Maiti and Drohat, 2011; Zhang et al., 2012). Compared to 5-hmC, TDG exhibits
great affinity for 5-fC and 5-caC (paired with guanine in double stranded DNA) (Ito et al.,
2011; Maiti and Drohat, 2011; Zhang et al., 2012). Supporting this, TDG deficiency causes
significantly increased levels of 5-caC in mouse ES cells (He et al., 2011; Shen et al.,
2013; Song et al., 2013). This collaborative action has been further demonstrated in germ
cell development (Nettersheim et al., 2013), in embryonic stem cells (Okashita et al., 2014)
and in somatic cell reprogramming (Hu et al., 2014). Therefore, TDG stands out as an
essential component in active DNA demethylation and epigenetics not only because it
interacts with numerous transcription factors and chromatin remodellers (Dalton and
Bellacosa, 2012) but also because its deletion in mice is embryonically lethal (Cortázar et
al., 2011; Cortellino et al., 2011). An alternative hypothesis is the direct removal of
carboxyl group from 5-caC. Although chemically feasible, such decarboxylase has not
been found yet (Schiesser et al., 2012).
Figure 1.2. Mechanisms of dynamic modifications of cytosine. 5-methylcytosine (5-mC) generated from cytosine by DNA methyltransferases (DNMTs), can further be oxidized by TET proteins to yield 5-hydroxymethylcytosine (5-hmC), 5-formylcytosine (5-fC) and 5-carboxylcytosine (5-caC). 5-fC and 5-caC can be excised by thymine DNA glycosylase (TDG) and replaced by an unmodified cytosine via base excision repair (BER) pathway. Alternatively, 5-mC and 5-hmC deamination by activation-induced deaminase (AID) and/or apolipoprotein B mRNA editing enzyme catalytic popylpeptide (APOBEC) proteins have been proposed yield thymine and 5-hydroxymethyluracil for DNA demethylation after excision by TDG and SMUG1 (single-strand-selective monofunctional uracil DNA glycosylase) followed by BER. However this model has been controversial (see the main text).
32
Chapter 1
1.2.4.4. TET-associated DNA demethylation dynamics in embryonic development and pluripotency
Following the discovery of TET-mediated 5-mC oxidation, the mechanisms of
genome-wide DNA demethylation occurring in pre-implantation embryos and during germ
cell development have been re-visited.
In the zygote
The first wave of demethylation occurs soon after fertilization allowing the
activation of the transcriptional programme in the totipotent zygote. Initial reports
demonstrated that the paternal pronucleus undergoes rapid loss of 5-mC before the onset
of DNA replication, whereas DNA methylation at the maternal pronucleus is gradually lost
during cleavage stages (Mayer et al., 2000; Oswald et al., 2000; Santos et al., 2002).
Notably, however, later analyses revealed that the observed paternal loss of methylation
signal was actually due to formation of 5-hmC as well as 5-fC and 5-caC, mediated by
TET3 protein (Gu et al., 2011; Inoue and Zhang, 2011; Iqbal et al., 2011; Wossidlo et al.,
2011). 5-hmC begins to be significantly accumulated in the paternal genome at around
pronuclear stage PN3 of the zygote, which coincides with the loss of cytosine methylation
signal, whereas the maternal genome is protected from 5-mC oxidation (Wossidlo et al.,
2011). At this stage, while Tet1 and Tet2 genes are silenced, Tet3, which is transcribed in
the oocyte, is present at high levels (Iqbal et al., 2011; Wossidlo et al., 2011). Importantly,
zygotic RNA interference-mediated silencing of Tet3 prevents 5-hmC generation
accompanied by increased 5-mC signal in the paternal pronuclei (Wossidlo et al., 2011).
Confirming this finding, generation of 5-hmC was not detected in the zygote upon maternal
deletion of Tet3 and blocked demethylation of paternal pluripotency-associated genes and
prevented subsequent gene activation (Gu et al., 2011). Furthermore, maternal Tet3
mutant embryos are severely degenerated and possess morphological abnormalities, and
homozygous Tet3 deletion results in prenatal lethality (Gu et al., 2011). It is important to
note that Tet3 is specifically associated with paternal pronuclei, indicating that it is actively
excluded from the maternal genome (Gu et al., 2011). Later it was shown that protection
of the maternal genome from oxidation is mediated by STELLA, which prevents TET3
binding by possibly altering chromatin configuration (Nakamura et al., 2012). STELLA
preferentially binds maternal chromatin due to its association with H3K9me3 that is
predominantly in the maternal pronucleus. From this point, replication-dependent
33
Chapter 1
demethylation of the paternal DNA occurs as evidenced by gradually decreased levels of
5-hmC during cleavage stages (Inoue and Zhang, 2011). This is further supported by the
limited affinity shown by DNMT1 for hemi-hydroxymethylated DNA and its exclusion of
maintenance machinery (Hashimoto et al., 2012; Hirasawa et al., 2008). Furthermore,
iterative oxidation products (5-fC and 5-caC) of 5-mC that are also present in the paternal
pronucleus can contribute to passive demethylation (Inoue et al., 2011). It is worth
mentioning that biological significance of genome-wide paternal 5-mC oxidation is not well
elucidated and TET3 deficiency does not affect global transcription levels in the embryo
(Inoue et al., 2012).
In the Primordial Germ Cells
The second wave of global DNA demethylation takes place during primordial germ
cell development and migration to the genital ridges, which is essential for the
establishment of totipotency in the next generation. Loss of DNA methylation in PGCs
follows a strict bi-phasic temporal order; demethylation first starts at the pluripotency
associated and germ cell specific genes, and only after involves ICRs, LINE repeats and
X-chromosome linked loci once PGCs settle in the gonads (Guibert et al., 2012;
Seisenberger et al., 2012). The bulk of the DNA demethylation occurs in the first phase
between E6.5 to E9.5, where overall CpG methylation level decreases from ~70% to 30%
(Seisenberger et al., 2012). It affects promoters of the genes that are expressed early in
PGC development, including transcription factors of pluripotency network genes such as
Nanog, while DMRs of the imprinted genes and X-linked CGIs maintain methylation at this
stage (Seisenberger et al., 2012). De novo methyltransferases are silenced in PGCs, while
DNMT1 is present and is localised in the nucleus (Hajkova et al., 2002; Kurimoto et al.,
2008). However DNMT1 tethering factor UHRF1 is significantly downregulated and the
remaining protein is cytoplasmic (Kurimoto et al., 2008; Seisenberger et al., 2012). This
suggests that genome-wide DNA demethylation may occur passively due to largely
impaired maintenance machinery, which may specifically protect some loci (including
DMRs of imprinted genes) from loss of methylation during this phase (Seisenberger et al.,
2012).
The second phase of DNA de-methylation further decreases overall CpG
methylation to 14% in male and 7% in female PGCs at E13.5 (Seisenberger et al., 2012)
and it seems to involve TET1 and TET2 mediated 5-mC oxidation (Hackett et al., 2013;
Yamaguchi et al., 2013a). Expression of Tet1 and Tet2 genes reaches maximum levels
34
Chapter 1
between E9.5 and E10.5, and is accompanied by substantial amount of 5-hmC generation
in the PGC nuclei and increases progressively until 5-mC is no more detectable at E11.5
(Hackett et al., 2013; Yamaguchi et al., 2012, 2013a). Genome-wide mapping
demonstrated that 5-hmC is accumulated at several late demethylated loci, including
DMRs of the imprinted genes (Hackett et al., 2013). Interestingly unlike ES and somatic
cells, 5-hmC is also enriched at heterochromatic satellite repeats in PGCs (Hackett et al.,
2013). From E11.5 onward, 5-hmC signal is progressively lost in the PGC nuclei as
evidenced by time-course immunostaining analysis. As PGCs undergo cell division at
every 12.6 hours at this stage (Kagiwada et al., 2013), this gradual loss is suggested to
occur by a replication-coupled mechanism where unmodified cytosine is incorporated
during DNA synthesis (Hackett et al., 2013). It is worth mentioning that, BER pathway
components including Tdg are expressed in PGCs, (but not Aid or Apobec1, calling into
question their contribution in loss of methylation as previously proposed (Popp et al.,
2010)), suggesting a potential role for these factors in DNA demethylation (Hajkova et al.,
2002; Kagiwada et al., 2013). The bi-phasic character of DNA demethylation and
contribution of 5-mC oxidation during germ cell development was further confirmed in-vitro
by ES cell into PGC differentiation (Vincent et al., 2013). This study revealed that TET
proteins are dispensable for the first phase, as evidenced by the presence of global
demethylation that occurred in TET1 and TET2 deficiency. On the other hand, the second
phase of locus-specific loss of methylation is TET-dependent (Vincent et al., 2013).
Tet1 depletion interferes with demethylation of meiosis related gene promoter and
transcriptional activation in female PGCs. This results in loss of oocytes, decreased
fertilization and smaller litters (Yamaguchi et al., 2012). Although male Tet1 deficiency
does not interfere with testes development, E13.5 PGCs exhibit locus specific
hypermethylation, mainly observed in late-demethylated group of imprinted genes
(Yamaguchi et al., 2013a). In the progeny derived from crossing Tet1 knockout males with
wild type females, aberrant hypermethylation is observed at the DMRs of imprinted genes
including Peg3 and Peg10. Dysregulation of imprinting is in part responsible for various
phenotypes observed in the new heterozygous generation, such as placental, foetal and
post-natal defects as well as embryonic lethality (Yamaguchi et al., 2013a). Tet1/Tet2
double knockout PGCs are completely devoid of 5-hmC and the progenies display
compromised imprinting, supporting the role of Tet activity in erasure of paternal imprints
(Dawlaty et al., 2013). The majority of Tet1/Tet2-null mice die within the first two days of
birth; however those who survive to adulthood are fertile, suggesting a compensatory role
35
Chapter 1
of TET3. Indeed, double knockout male sperm contains almost normal levels of 5-hmC
(Dawlaty et al., 2013).
In embryonic stem cells
Since the discovery of relatively high levels of 5-hmC in ES cells, studies have
focused on the roles of TET proteins in ES cell-specific properties that are self-renewal
and pluripotency. Initially, shRNA-mediated Tet1 knockdown in mouse ES cells was
shown to result in Nanog downregulation and defects in stem cell maintenance (Ito et al.,
2010). Furthermore, acute depletion of Tet1 resulted in impaired LIF/Stat3 signalling with
adverse effects on ES cell identity (Freudenberg et al., 2012). In contrast, several other
studies reported that Tet1 knockdown does not affect the expression of key pluripotency-
associated factors including Nanog. The discrepancy between different studies may stem
from culture conditions, use of different cell lines and RNA interference based off-target
effects (Ficz et al., 2011; Koh et al., 2011; Williams et al., 2011). More evidence was
provided upon genetic deletion of Tet1, where knockout ES cells exhibit normal
undifferentiated ESC morphology and express pluripotency markers (Dawlaty et al.,
2011). To rule out possible redundant functions and compensation by TET2 in chronic
Tet1 depletion, Dawlaty et al. further deleted Tet2 gene and generated double Tet1/Tet2
knockout ES cells (Dawlaty et al., 2013). This resulted in complete depletion of 5-hmC
signal, while the cells remained pluripotent, as evidenced by their capacity to generate
tissues from all embryonic layers by teratoma assay (Dawlaty et al., 2013). Interestingly,
teratomas formed from Tet1 single and Tet1/Tet2 double knockout ES cells are
haemorrhagic and enriched for throphoblast cells, suggesting a skewed differentiation
potential toward extraembryonic tissues. It should be noted that these cells can efficiently
contribute to the chimaeras upon blastocyst injection, indicating that differentiation
problems can be overcome during embryonic development (Dawlaty et al., 2013).
Similarly, ES cells deficient of all three members of TET proteins maintain normal
morphology and express pluripotency markers, however their differentiation potential is
severely restricted, as reported by lack of endodermal and mesodermal structures in
teratomas (Dawlaty et al., 2014). Moreover, triple knockout ES cells contribute poorly to
developing embryos upon blastocyst injection, probably due to hypermethylation of
promoter regions of several developmental genes. Confirming the importance of TET
proteins in this process, ectopic Tet1 expression in TET-null ES cells rescues their
differentiation defects and reinstates chimeric contribution. (Dawlaty et al., 2014). Overall
36
Chapter 1
this data demonstrate partial redundancy between TET proteins and their critical roles in
proper differentiation.
TET-assisted DNA demethylation has recently been shown to contribute to the
acquisition of an ICM-like naïve state in mouse ES cells. Traditionally mouse ES cells are
isolated and grown in serum supplemented with leukaemia inhibitory factor (LIF), however
they exhibit heterogeneity with two distinguishable populations. These are primed
epiblast-like and naïve ICM-like states, as exemplified by biphasic Nanog expression
(Chambers et al., 2007). Interestingly ICM-like ground state homogeneity can be
established by growing mouse ES cells in media supplemented with two kinase inhibitors
(2i) (Ying et al., 2008) and the epigenetic basis of this naïve status has recently begun to
be elucidated. For example, ES cells isolated in the presence of 2i display genome-wide
hypomethylation that span CGIs, LINE1 elements, as well as minor and major satellite
sequences, conversely to what observed in serum supplemented media (Leitch et al.,
2013). Similarly, 2i addition to serum grown ES cells results in a rapid shift of methylation
status, accompanied by downregulation of Dnmt3a, Dnmt3b and Dnmt3l gene expression
that is partly mediated by PRDM14 (Leitch et al., 2013). Consequently, genome-wide 5-
hmC accumulation catalysed by TET1/TET2, coupled with impaired methylation
maintenance and disabled de novo methyltransferase activity, results in global replication-
coupled passive loss of methylation in 2i condition (Ficz et al., 2013; Habibi et al., 2013).
In addition, global methylation data sets confirmed that ES cells grown in 2i show
similarities to ICM cells at E3.5 or migratory PGCs at E9.5 (Ficz et al., 2013; Habibi et al.,
2013), reminiscent of TET activity during embryonic development.
Interestingly, in 2i condition further demethylation can occur in the presence of a
rather unexpected supplement, vitamin C. Actually, vitamin C is a necessary ingredient of
human embryonic stem cell media and activates genes implicated in growth, proliferation
and pluripotency by regulating methylation dynamics at their promoters (Chung et al.,
2010). Interestingly, mouse embryonic fibroblasts supplemented with vitamin C undergo
significant 5-mC oxidation, while TET levels remain stable. This is due to enhanced
enzymatic TET activity upon vitamin C association with TET catalytic domains that results
in structural change and enhanced product recycling (Minor et al., 2013). In mouse ES
cells where Tet1 and Tet2 genes are active, vitamin C supplementation leads to 40%
decrease in genome-wide 5-mC levels due to 5-hmC generation, affecting previously
methylated gene promoters and results in upregulation of germline genes (Blaschke et al.,
2013; Yin et al., 2013). Yet, imprinted genes and intracisternal A particle retroelements
37
Chapter 1
resist demethylation induced by vitamin C, reminiscent of pre-implantation embryo
development (Blaschke et al., 2013).
Figure 1.3. TET-induced demethylation in mouse embryonic development and ES cells. TET proteins are involved in both genome-wide demethylation waves that occur in the pre-implantation embryo and in primordial germ cells (PGCs). TET-assisted demethylation is also observed in mouse ES cells cultured in 2i media and in the presence of Vitamin C. TET involvement in the transitions is highlighted as surf board and DNA methylation dynamics are indicated. The methylated cytosine is shown in red, unmodified cytosine in grey. Adapted and modified from (Bagci and Fisher, 2013).
1.3. Reprogramming cell fate
Early mammalian development involves sequential cell fate decisions
accompanied by specialization and progressive restriction of potential, giving rise to the
germ lineage and over two hundred different types of somatic cells. It has long been
thought that cell differentiation is an irreversible process, with a few exceptional
pathological cases such as metaplasia and malignancy. However, decades of research
have now challenged this view, unravelling the hidden potential of differentiated cells.
38
Chapter 1
1.3.1. Transdifferentiation Transdifferentiation is a process of direct switch of cell fate, which transforms one
cell type into another without passing through a pluripotent state. A very important
example of this switch was demonstrated by treating immortalized fibroblasts with 5-
azacytidine (inhibitor of DNA methylation), which resulted in their conversion into
adipocytes and chondrocytes (Taylor and Jones, 1979). This suggested that lineage-
specific transcription factors are tightly controlled by DNA methylation. It was later
demonstrated that, a single factor, MyoD, can convert fibroblasts into myocytes via ectopic
expression (Davis et al., 1987). The blood system has been an informative platform in
terms of cell conversion analyses. For example, high level expression of the erythroid-
megakaryocyte-affiliated transcription factor GATA1 in monocytes (macrophage
precursors) does not only downregulate monocyte-associated factors, but also activates
erythroid-megakaryocyte gene expression (Kulessa et al., 1995; Visvader et al., 1992).
Interestingly, conversion towards the opposite direction can also occur; expression of
PU.1 in erythroid-megakaryocytic cells promotes their switch to monocytes by repressing
GATA1 (Nerlov and Graf, 1998). Furthermore, the granulocyte/macrophage-specific
transcription factor C/EBPα can very efficiently convert primary B-cell progenitors into
macrophages, or to a lesser extent immunoglobulin-producing B lymphocytes (Xie et al.,
2004). Identification of factors that can modulate cellular epigenome for fate switching has
implications in regenerative therapy. For example, pancreatic β-cells are key targets for
treatment of diabetes, and a transcription factor screen in pancreatic tissue revealed three
important genes: Pdx1, Ngn3 and MafA for their normal function. In vivo adenoviral
delivery of these factors for transient expression in fully differentiated pancreatic exocrine
cells resulted in conversion into β-cells with an efficiency of 20% (Zhou et al., 2008). A
similar candidate-based approach was used to identify Gata4, Mef2c and Tbx5 genes that
can directly convert mouse cardiac and dermal fibroblasts into cardiomyocyte-like cells
(Ieda et al., 2010). Additional examples include Ascl1-mediated fibroblast-to-neuron
conversion (Vierbuchen et al., 2010) and fibroblast-to hepatocyte conversion by defined
factors (Huang et al., 2011; Sekiya and Suzuki, 2011).
These examples have demonstrated powerful trans-acting functions of
transcription factors. This is especially interesting considering the presence of higher order
chromatin organization that packs the DNA to repress gene expression and to prevent
accessibility of activating factors. Although cell division has been proposed to allow
transcription factor binding due to unwinding of the chromatin during DNA replication, in
39
Chapter 1
some cases cell fate can rapidly be converted in the absence of cell proliferation. This is
proposed to be accomplished by ‘pioneer transcription factors’ which can target repressive
lineage-specific regulatory sites, to create a permissive environment for accessibility and
binding of additional activators (Zaret and Carroll, 2011). It is important to note that
transdifferentiation efficiency depends on developmental proximity between two cell types
and degree of their lineage specificity. Cell fate conversion between different germ layers
has been more challenging due to genome-wide differences in epigenetic status.
Therefore, a distant switch may require additional transcription factors and epigenetic
modulators (Graf and Enver, 2009).
1.3.2. Pluripotent conversion of somatic cells
Cellular differentiation is accompanied by the acquisition of lineage-specific
transcription programme. Although mechanisms of epigenetic memory ensures
conservation of fate during proliferation, cells remain surprisingly plastic. This does not
only render the process of transdifferentiation possible as described earlier, but also
allows pluripotent conversion of differentiated cells. Once a pluripotent state is acquired,
it is possible to generate cells from different germ lines, which has significant therapeutic
implications. Cellular plasticity was demonstrated by early studies of nuclear transfer and
cell fusion, and recently by factor mediated induction of pluripotency. It is crucial to
elucidate molecular details of pluripotent conversion, to better understand epigenetic
mechanisms that govern cellular differentiation and establishment of pluripotency.
1.3.2.1. Nuclear transfer
Once transplanted into an enucleated oocyte, the nucleus of a differentiated cell
undergoes extensive nuclear reprogramming that can lead to the generation of an entire
organism, a process also known as cloning. Pioneering work was conducted in
amphibians, where blastocyst cell implantation into enucleated oocyte resulted in clones
of swimming tadpoles (Briggs and King, 1952). Although the efficiency was lower when
more differentiated intestinal epithelium cell nuclei were transferred (~1% compared with
~30% in the case of blastocyst cells), adult frog clones could still be obtained (Gurdon,
1962a, 1962b). These initial results proved the plasticity and reversibility of differentiated
nuclear state, and revealed that all the genetic information to generate an entire individual
40
Chapter 1
Figure1.4. In-vitro strategies for nuclear reprogramming to pluripotency. Nuclear transfer. This approach involves the injection of somatic cell nucleus into an enucleated oocyte. Factors in the oocyte cytoplasm act on the somatic nucleus and reprogram it to a pluripotent state. This oocyte can then be further cultured to obtain genetically matched nuclear transfer embryonic stem (ntES) cells or to obtain a clone animal upon implantation into a surrogate mother. Induction of pluripotency factors. Ectopic expression of pluripotency-associated genes in a somatic cell can initiate reprogramming, and resulting “induced pluripotent stem cells” (IPSCs), which have embryonic stem cell-like properties can be stably propagated in culture. Cell fusion. In this approach, a somatic cell can be reprogrammed to pluripotency after fusion with a pluripotent cell type such as embryonic stem or embryonic germ cell. The fusion first yields a heterokaryon where parental nuclei are separated but share the same cytoplasm. The merging of the nuclei results in the formation of the pluripotent hybrid, which, in the case of an intraspecies fusion, can stably be cultured.
is stably kept in the specialized cell nucleus. The same approach in mammalians gave
rise to the first cloned sheep (named after Dolly Parton) (Wilmut et al., 1997), and a year
later led to cloning of mice (Wakayama et al., 1998). Mice clones could further be
generated by somatic-cell nuclear transfer (SCNT), using mature B and T cells as well as
olfactory sensory neurons as donors (Eggan et al., 2004; Hochedlinger and Jaenisch,
2002). It was notably, however, that reprogramming of terminally differentiated cell was
41
Chapter 1
very inefficient and a two-step model was used. In this system, nuclear transplantation of
the somatic cell gives rise to the blastocysts, from which nuclear-transfer-embryonic stem
(ntES) cells are derived and retransplanted into enucleated oocytes. It should be noted
that cloned animals display common abnormalities that involve premature death (Thuan
et al., 2010). That is most probably due to a defect in proper erasure of epigenetic memory
that has been imposed during differentiation by DNA methylation and histone
modifications (Simonsson and Gurdon, 2004). Supporting this, inhibition of histone
deacetylases improved the efficiency of cloning, possibly by promoting histone acetylation
that leads to structural changes in the chromatin and enhanced DNA demethylation
(Kishigami et al., 2006). In the past decade, a wide range of species have been cloned
(Thuan et al., 2010), and ntES cells have been successfully derived from non-human
primates (Byrne et al., 2007). Although several attempts to isolate human ntES cells failed
in the past (Egli et al., 2011; Noggle et al., 2011), a recent study demonstrated their
isolation by using high quality enuclated human oocytes as recipients and human
fibroblast nuclei as donors (Tachibana et al., 2013). This has important implications in
derivation of patient-specific pluripotent cells for developing specific therapies.
1.3.2.2. Induced pluripotent stem cells
In 2006, Takahashi and Yamanaka reported the derivation of induced pluripotent
stem (iPS) cells from mouse embryonic fibroblasts (MEFs) and skin cells (Takahashi and
Yamanaka, 2006). They conducted a systematic candidate-based interrogation of ES cell-
associated genes for their potential to induce pluripotency in adult cells and identified four
factors: OCT4, SOX2, KLF4 and c-MYC (OSKM). This breakthrough in reprogramming
was quickly embraced and iPS cells were derived upon ectopic expression of OSKM from
human fibroblasts and from a plethora of differentiated cells in subsequent studies
(Yamanaka, 2009). iPS cells form compact ES cell-like colonies with distinct borders, can
propagate in culture indefinitely and possess the capacity to form each of the three
embryonic germ layers (Hanna et al., 2010). It is important to note that the differentiation
potential of iPS cells, which would be a prerequisite for therapy, is influenced by the
parental-origin. For example, monocyte-derived iPS cells can differentiate more efficiently
along the haematopoietic lineages compared to fibroblast-derived iPS cells. The fact that
their potential can be enhanced by HDAC inhibitors and 5-azacytidine indicates a defect
in erasure of epigenetic memory during reprogramming (Kim et al., 2010). Somatic cell
reprogramming is an inefficient and lengthy process (Hochedlinger and Plath, 2009), and
42
Chapter 1
has been suggested to be composed of three phases: initiation (marked by mesenchymal-
to-epithelial (MET) transition), maturation and stabilization (marked by expression of
genes associated with embryonic development and maintenance) (Samavarchi-Tehrani
et al., 2010). OSKM activation first induces stochastic regulation of genes involved in
pluripotency, in MET, in proliferation and in metabolism, and is possibly the rate-limiting
step of reprogramming. Those cells with the appropriate combination of gene expression
can then enter a hierarchical phase, in which pluripotency network is activated leading to
a decrease in intercellular variation (Buganim et al., 2012). Transcriptional changes that
are observed during pluripotent conversion are driven by epigenetic alterations in the
nucleus. These involve histone modifications, chromatin reorganization and DNA
demethylation. Remarkably, these events are ordered: although histones are immediately
modified after induction (Koche et al., 2011). DNA demethylation occurs late in the process
and coincides with the acquisition of a stably reprogrammed state (Polo et al., 2012).
OSKM can be in part, or fully replaced by other transcription factors or chemicals for the
induction of pluripotency, yet a wide discrepancy exists between studies, which stems
from the stochasticity and the inefficiency of the reprogramming process (Theunissen and
Jaenisch, 2014).
1.3.2.3. Cell fusion
Cell fusion experiments were initiated in 1960s. Since then this methodology has
provided valuable information on molecular details of nuclear-cytoplasmic interaction,
dominant effects of trans-acting factors, nuclear plasticity and regulatory mechanisms of
pluripotency. Fusion of two or more cell types creates a single cytoplasmic entity called a
heterokaryon, which harbours both parental nuclei. Heterokaryons are generally transient,
and upon further culture, the nuclei merge and give rise to hybrid cells which undergo cell
division. It should be noted that stability of hybrids depends on the genetic background
compatibility of the fusion partners. While fusion between same species can generate
stably proliferating hybrids, interspecies hybrids are susceptible to chromosomal
abnormalities and loss.
Based on the fact that certain animal viruses can induce the formation of
multinucleated cells, Harris and Watkins used Sendai virus to induce interspecies fusion
between human HeLa and mouse Ehrlich ascites tumour cells (Harris and Watkins, 1965).
Tritiated uridine and leucine labelling experiments demonstrated that both sets of nuclei
could produce RNA in the resulting heterokaryons that also contain normal protein levels.
43
Chapter 1
This suggested that fundamental cellular functions are not altered upon fusion.
Furthermore, although heterokaryons did not undergo cell division, tritiated thymidine
staining revealed the presence of partially synchronised DNA synthesis in unfused nuclei
(Harris and Watkins, 1965). This was further confirmed in a systematic approach, where
a parental cell line undergoing DNA replication was shown to induce premature DNA
synthesis on its fusion partner nucleus (Rao and Johnson, 1970). In the meantime, cell
fusion system provided evidence on regulation of gene expression by trans-acting
elements. This was demonstrated by ceased melanin and tyrosine amino-transferase
synthesis in hamster melanocytes or rat hepatocytes, respectively, after fusion with mouse
fibroblasts, revealing very important initial clues on dominant action of a cell type on its
fusion partner (Davidson et al., 1966; Weiss and Chaplain, 1971). Dominance was further
evaluated in fusion experiments conducted between malignant with non-malignant cells,
and tumorigenicity was found to be suppressed in hybrid cells by the possible trans-acting
functions of anti-oncogenes (Harris and Miller, 1969). Previously silenced genes can also
be activated upon cell fusion, as initially analysed in hybrid cells (Davidson, 1972;
Peterson and Weiss, 1972). One caveat of these early reports arises from the instability
of interspecies hybrids, where chromosomal loss or rearrangement may result in the
observed phenotypes. However, enhanced sensitivity of detection techniques facilitated
the analysis of early events, and led subsequent studies to focus on heterokaryons. For
example, heterokaryons formed between human amniocytes and mouse muscle cells
started to produce human specific muscle proteins that are normally repressed in
amniocytes, as early as 3 days of fusion (Blau et al., 1983). As parental nuclei remain
distinct and retain their chromosomes in heterokaryons, observed gene activation could
be directly attributed to the mouse muscle specific trans-acting factors in the cytoplasm.
Furthermore, muscle cells were able to alter the differentiated state of cells originated from
different embryonic lineages (mesoderm, ectoderm and endoderm), revealing the nuclear
plasticity of diverse cell types (Blau et al., 1985). It is important to note that frequency and
kinetics of reprogramming is a cell-type specific phenomenon, for example mesoderm-
derived human cells (which are more closely related to muscle cells) initiated muscle
specific gene expression sooner compared to ectoderm or endoderm derived cells (Blau
et al., 1985). Reprogramming of distant lineages by muscle cells (such as hepatocytes
derived from the endoderm) is dependent on the gene dosage, as lowest reprogramming
efficiency was observed in heterokaryons when hepatocyte nuclei proportion was higher
(Blau et al., 1985). In addition, HeLa cells that could not be reprogrammed by muscle cells,
were only reprogrammed following 5-azacytidine treatment, increasing their
44
Chapter 1
responsiveness for muscle specific trans-acting regulatory factors (Chiu and Blau, 1985).
These reports demonstrated that somatic cells can influence the transcription
programme of their fusion partners, by the action of cell specific master regulators. Early
studies also addressed whether pluripotency can be induced in differentiated cells. For
example, when mouse embryonic carcinoma (EC) cells were fused with mouse
thymocytes, not only the features of the somatic partner were extinguished, but also
pluripotent properties of the parental EC cells (Miller and Ruddle, 1976), including
reactivation of the silent X-chromosome (Takagi, 1993), were acquired in the generated
hybrids. Dominant pluripotent conversion capacity was later extended by Tada and
colleagues, by fusing mouse thymocytes with mouse EG (Tada et al., 1997) or mouse ES
cells (Tada et al., 2001) that resulted in pluripotent hybrids. It is worth mentioning that
these studies revealed an important finding; somatic originated imprinted genes remained
methylated when the fusion partner was ES cells, but were demethylated upon fusion with
EG cells. This resembles to the regulation of imprinted gene methylation in early embryo
and PGC development, from which ES and EG cells are derived (Tada et al., 1997, 2001).
Subsequent studies demonstrated that ES cells from human origin can successfully
reprogram fibroblasts, and resulting stable tetraploid hybrids possess the characteristics
of parental ES cells. They actively express OCT4 gene, whose promoter is
hypomethylated and can differentiate into each embryonic germ layer (Cowan et al.,
2005). Cell fusion-based reprogramming is a fast process as evidenced by rapid activation
of Oct4-GFP transgene embedded in the somatic genome (Do and Schöler, 2004; Han et
al., 2008; Silva et al., 2006; Wong et al., 2008). Remarkably, pluripotent-specific
transcription program is initiated in interspecies heterokaryons as early as 24 hours of
fusion, indicated by induction of OCT4, NANOG, CRIPTO, ESRRB, TLE1 and REX1
genes from the somatic nucleus (Bhutani et al., 2010; Pereira et al., 2008; Soza-Ried and
Fisher, 2012). Regulation of gene expression is accompanied by substantial changes in
the chromatin structure. For instance, somatic DNA in pluripotent heterokaryons and
hybrids acquire histone H3 and H4 acetylation as well as di- and tri-methylated H3K4me3,
all of which are marks of a pluripotent state (Kimura et al., 2004; Piccolo et al., 2011).
Although cell fusion based reprogramming kinetics are similar to nuclear transfer,
the latter method is technically challenging (Yamanaka and Blau, 2010). The speed and
extent of reprogramming by fusion are not surprising considering the effectiveness of the
pluripotency-associated network in maintaining pluripotent identity by regulating gene
expression (Ng and Surani, 2011). This already established powerful network can quickly
act on the somatic nucleus. It is noteworthy that Oct4, Klf4 and Sox2 have recently been
45
Chapter 1
identified as pioneering transcription factors, suggesting that their modulatory functions
are more powerful than previously anticipated (Soufi et al., 2012). This is exemplified by
rapid localisation of the OCT4 protein on the somatic DNA after fusion with ES cells and
reprogramming cannot be achieved in its absence (Pereira et al., 2008). In this regard,
cell fusion model establishes a platform to screen for important factors that drive
pluripotent conversion. For example, fusion of ES cells that overexpress Nanog with
neural stem cells yields at least 200-fold increased number of pluripotent hybrids (Silva et
al., 2006). Similarly Polycomb-group proteins are essential for successful reprogramming
of lymphocytes by ES cells (Pereira et al., 2010). AID has also been demonstrated to be
crucial in pluripotent conversion by actively demethylating OCT4 and NANOG gene
promoters in somatic DNA (Bhutani et al., 2010); however this observation has been
revoked by subsequent studies (Foshay et al., 2012).
46
Chapter 1
1.4. Aims of this study
Pluripotent conversion of differentiated cells involves the installation of a
pluripotency-associated transcription program which is mediated by nuclear
reorganisation, chromatin remodelling and resetting of DNA methylation patterns. The
overall aim of this thesis is to elucidate molecular mechanisms of DNA demethylation in
the course of initiation and propagation of successful reprogramming. In this regard, cell-
fusion based reprogramming provides a tractable experimental platform that helps
dissection of early events leading to pluripotent conversion. For this reason, I will generate
experimental heterokaryons and hybrids between pluripotent and somatic cells, and
analyse the epigenetic stages of reprogramming.
Initially, I will focus on the dynamics of DNA demethylation in heterokaryons
(generated between mouse embryonic stem cells and B lymphocytes or fibroblasts) to
understand its role in initiation of pluripotency-associated gene expression. Then I will
investigate the molecular details of mouse embryonic germ cell mediated imprint erasure
in somatic genome upon fusion. I will finally analyse the involvement of TET proteins at
the early stages of successful reprogramming, by using gene knockdown and knockout
approaches. For this, I will explore CRISPR/Cas9-induced gene editing system, which I
will also use as part of a technical chapter, to generate mouse embryonic stem cell lines
deficient of non-canonical Wnt pathway components.
47
Chapter 2
Chapter 2. Materials and Methods
2.1. Materials
2.1.1. Cell lines
Mouse embryonic stem cells
E14tg2A : Feeder-free, hypoxanthine guanine phophoribosyltransferase (HPRT) –deficient mouse
embryonic stem cell line (Hooper et al., 1987). The main mouse embryonic stem cell line used throughout this
thesis unless otherwise specified. Pgk12.1 : Feeder-free female embryonic stem cell line, kindly provided by Professor Neil Brockdorff
(Penny et al., 1996).
Mouse embryonic germ cells
58G : Mouse embryonic germ cell line derived from primordial germ cells of female E12.5 embryos,
kindly provided by Professor Takashi Tada (Tada et al., 1997).
Mouse B lymphocytes
mB : Mouse pre-B cell line derived from transgenic (GOF-18/ΔPE/GFP) mice (Yoshimizu et al.,
1999). These cells were Aberson transformed and contain a Puromycin resistance cassette. The main mouse B
lymphocyte line used throughout this thesis unless otherwise specified. 2rB : Mouse pre-B cell line derived from transgenic mouse generated by mating male GOF-
18/ΔPE/GFP (Yoshimizu et al., 1999) with female Peg1M-β-gal (Lefebvre et al., 1998) mice. These cells were
Aberson transformed and contain a Puromycin resistance cassette.
Human embryonic stem cells
H7 : Human embryonic stem cell line, derived from human blastocysts.
Human B lymphocytes
hB : Epstein-Barr Virus transformed human adult B cell line, kindly provided by Emily R. Eden (Eden
et al., 2002).
Human fibroblasts
IMR90 : Female human fibroblasts (Coriell), immortalised by telomerase reverse transcriptase (pBABE-
hTERT-blastocydin plasmid).
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Chapter 2
2.1.2. Antibodies
mTet1 : Rabbit Polyclonal anti-Tet1 (Millipore, 09-872). WB at 1:1000
mTet2 : Rabbit Polyclonal anti-Tet2 (Santa Cruz Biotechnology, sc136926). WB at 1:250 mOct4 : Goat Polyclonal anti-Oct-3/4 (Santa Cruz Biotechnology, sc-8628). WB at 1:2000.
mNanog : Rabbit Polyclonal anti-Nanog (Cosmo Bio, REC-RCAB0002P-F). WB at 1:5000
mTubulin : Rabbit Monoclonal anti-α Tubulin (Sigma, T9026). WB at 1:10000
mJarid2 : Rabbit Polyclonal anti-Jarid2 (Abcam, ab48137). WB at 1:750
mLamin B : Goat Polyclonal anti-Lamin B (Santa Cruz Biotechnology, sc-6216). WB at 1:20000
H3 : Rabbit Polyclonal anti-histone 3 (Abcam, ab1791). ChIP, 2µg per IP
H3K4me3 : Rabbit Polyclonal anti-histone 3 trimethyl K4 (Millipore, 07-473). ChIP, 2µg per IP
2.2. Methods
2.2.1. Cell culture
All of the reagents used for tissue culture were supplied from Gibco (Life
Technologies), unless otherwise specified. All of the cells were maintained at 37oC and
5% (v/v) CO2, unless otherwise specified.
Mouse embryonic stem cells were cultured in Knockout Dulbecco’s Modified Eagle
Medium (KO-DMEM), supplemented with 10% (v/v) Fetal Bovine Serum (FBS), 1mM Non-
Essential Amino Acids (NEAA), 2mM L-Glutamine, antibiotics (100U/ml penicillin &
100µg/ml streptomycin), 50µM β-mercaptoethanol and 1000U/ml Leukemia Inhibitory
Factor (LIF) (ESGRO-LIF, Millipore). The cells were maintained on 0.1% gelatin (Sigma)
coated dishes. The cells were routinely passaged at a 1:8 dilution by treatment with 0.05%
Trypsin-EDTA.
Mouse embryonic germ cells were cultured in DMEM/F12, supplemented with 20%
(v/v) FBS, 1mM NEAA, 2mM L-Glutamine, antibiotics (100U/ml penicillin & 100µg/ml
streptomycin), 50µM β-mercaptoethanol, 1mM Sodium Pyruvate, 0.12% Sodium
Bicarbonate, 1% (v/v) Nucleosides (stock solution is prepared in distilled water: 0.08%
(m/v) Adenosine, 0.073% (m/v) Cytidine, 0.024% (m/v) Thymidine, 0.085% (m/v)
Guanosine, 0.073% (m/v) Uridine (Sigma)) and 1000U/ml Leukemia Inhibitory Factor (LIF)
(ESGRO-LIF, Millipore). The cells were maintained on γ-irradiated primary mouse
49
Chapter 2
embryonic fibroblast feeder layers and were routinely passaged at a 1:8 dilution by
treatment with 0.05% Trypsin-EDTA. Embryonic germ cells were treated with Plasmocin
(25µg/ml, Invivogen) for 1 week, and all the experiments were performed between
passages 24 and 28.
Mouse B lymphocytes were cultured in Roswell Park Memorial Institute-1640
(RPMI-1640), supplemented with heat inactivated 20% (v/v) FBS, 1mM NEAA, 2mM L-
Glutamine, antibiotics (100U/ml penicillin & 100µg/ml streptomycin) and 50µM β-
mercaptoethanol.
Human embryonic stem cells were cultured in mouse embryonic fibroblast
conditioned KO-DMEM, supplemented with 20% (v/v) Knockout Serum Replacement
(KSR), 1mM NEAA, 2mM L-Glutamine, antibiotics (100U/ml penicillin & 100µg/ml
streptomycin), 50µM β-mercaptoethanol, 1mM Sodium Pyruvate, 40ng/ml bFGF
(Peprotech) and 5ng/ml Activin A (Peprotech). The cells were maintained on dishes
overnight coated with 0.5mg/mL growth-factor-reduced matrigel (BD Biosciences) and
were routinely passaged by a 1:3 dilution by treatment with 0.02% EDTA.
Human B lymphocytes were cultured in RPMI-1640, supplemented with heat
inactivated 10% FBS, 1mM NEAA, 2mM L-Glutamine and antibiotics (100U/ml penicillin
& 100µg/ml streptomycin).
Human fibroblasts were cultured in DMEM, supplemented with supplemented with
10% FBS, 1mM NEAA, 2mM L-Glutamine and antibiotics (100U/ml penicillin & 100µg/ml
streptomycin). The cells were cultured in 3% (v/v) oxygen and were routinely passaged at
a 1:3 dilution by treatment with 0.05% Trypsin-EDTA.
Cells were frozen for later usage using 10% dimethyl sulfoxide (DMSO) in FBS.
2.2.2. Cell fusion experiments
Inter-species or intra-species cell fusion experiments were performed between
mouse ES or mouse EG cells and either human B, mouse B, 2rB lymphocytes or IMR90
fibroblasts as previously described (Pereira and Fisher, 2009), with minor modifications.
Briefly, cells were mixed in a conical 20-ml universal tube with a 1:1 ratio and washed
twice with KO-DMEM. After the last wash, the supernatant was completely removed, the
cell pellet was gently broken and 1mL of polyethylene glycol (PEG 1500, Sigma) was
added onto the cell pellet over a 60 second period. The PEG-cell mixture was incubated
at 37oC for 90 seconds with constant stirring. Then PEG was diluted by addition of 4mL of
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Chapter 2
KO-DMEM over a 4 minute period, followed by additional 10 mL KO-DMEM, and
incubation at 37oC for 3 minutes. The cells were then centrifuged at 1350 rpm for 5 minutes
followed by the removal of the supernatant and the cells were let to swell in complete
mouse ES or mouse EG cell media for 3 minutes before resuspension. Cell mixtures were
plated under conditions that promote maintenance of pluripotency at 0.5x106cells/cm2
density. Non-fused mouse ES or mouse EG cells were eliminated by the addition of
puromycin (1µg/ml, Sigma), after 6-12 hours fusion and onwards. In the case of mouse
ES (H2BmCherry) with IMR90 (HP1αGFP) fusion, double positive heterokaryons were
isolated by Fluorescence Activated Cell Sorting (FACS, see section 2.1.3) after 48 hours
of fusion and plated for further culture. GFP positive reprogrammed inter-species hybrid
colonies generated between mouse ES or mouse EG cells and mouse B or 2rB
lymphocytes were picked (between 8 to 10 days of fusion) under fluorescence microscope
(Leica DM IRE2), dissociated with 0.05% Trypsin-EDTA, plated back and further cultured.
At earlier time points, GFP positive reprogrammed cells were isolated by FACS and further
processed for DNA isolation.
2.2.3. Fluorescence activated cell sorting (FACS)
Cells were resuspended in Phosphate Buffered Saline -/- (PBS, Gibco)
supplemented with 2% FBS and 2mM EDTA (FACS buffer) and run in the FACS
instrument (BD FACSAria III). Sorted cells were collected in FACS buffer, and plated in
appropriate culture conditions.
2.2.4. Quantitative Reverse Transcription Polymerase Chain Reaction (qRT-PCR) Analysis
2.2.4.1. RNA extraction and reverse transcription Cell lysates were homogenised with the QIAshredder columns (Qiagen), total RNA
was extracted using the RNeasy Mini kit (Qiagen) and residual DNA was eliminated with
the DNA-free kit (Ambion), following manufacturer’s instructions. RNA concentration was
measured using NanoDrop (Thermo). Reverse transcription of the total RNA was
performed using SuperScript III First-Strand Synthesis system (Invitrogen). Briefly, up to
51
Chapter 2
3µg of RNA was diluted in RNase free water containing 1µl of 10mM dNTP mix and 1µl of
oligo(dT)12-18 (Invitrogen) to a final volume of 11µl. The mixture was incubated at 65oC for
5 minutes and kept at 4oC for 2 minutes, while 4µl of 5X first strand buffer, 1µl of 0.1M
dithiothreitol (DTT), 1µl of RNase OUT and 1µl of 200U/µl of SuperScript III (Invitrogen)
were added. An additional mixture was also prepared without the reverse transcriptase as
a negative control. The mixture was then incubated at 25oC for 5 minutes, 50oC for 50
minutes and 75oC for 15 minutes to generate cDNA.
2.2.4.2. Quantitative PCR
The PCR mixture was prepared as follows: 2µl of cDNA, 300nM of forward and
reverse primers, 1X QuantiTect Sybr Green PCR mix (Qiagen) completed to 20µl with
DNase free water. An additional mixture was also prepared without cDNA as a negative
control for primer dimer formation. The cDNA in the mixture was amplified (in duplicates)
in Chromo 4 PCR instrument (Bio-Rad), using Opticon Monitor 3 software (MJ Research)
under the following cycling conditions: initial denaturation at 95oC for 15 minutes, 40 cycles
of denaturation at 94oC for 15 seconds, annealing at 60oC for 30 seconds and elongation
at 72oC for 30 seconds. Fluorescence quantification was performed at 72oC, 75oC, 78oC
and 83oC, and melting curve was determined from 70oC to 90oC, at 0.2oC intervals. The
qPCR data analysis was performed using the Opticon Monitor 3 software and the relative
cDNA abundance of the samples (to GAPDH and Ubc housekeeping genes) was
calculated with the obtained Ct values (threshold cycle number in which the fluorescence
resulting from the amplification becomes detectable above background) using the formula
2ΔΔCt. Primer sequences for qPCR analysis can be found in the Appendix Table 1.
For quantitative PCR analysis of ChIP samples similar settings were used except
for instead of the cDNA, 5µl of DNA was added in the reaction mixture. Primer sequenced
for ChIP analysis can be found in the Appendix Table 4.
2.2.5. DNA methylation and hydroxymethylation analyses
All the reagents described below were supplied from Sigma, unless otherwise
specified. For genomic DNA isolation, cells were first lysed by overnight incubation at 55oC
in 500µl of lysis buffer that contains 10mM NaCl, 10mM Tris-HCl pH 7.5, 10mM EDTA,
0.5% Sarcosyl and 200µg/ml Proteinase K. The genomic DNA containing aqueous
52
Chapter 2
solution was collected upon subsequent treatments with Phenol,
Phenol/Chloroform/Isoamyl Alcohol and Chloroform and the DNA was precipitated using
1/10 volume 3M Sodium Acetate pH 5.2 and 2.5X volume Ethanol, followed by
resuspension in Tris-EDTA Buffer (10mM Tris. 1mM EDTA). DNA concentration was
measured by NanoDrop (Thermo).
2.2.5.1. Bisulfite sequencing analysis
Up to 1.5µg of genomic DNA was used for bisulfite conversion reaction that was
performed using the EZ DNA Methylation Kit (Zymo Research), following manufacturer’s
instructions. Bisulfite converted DNA was recovered by columns provided in the kit in 10µl
elution buffer (provided in the kit). 2µl of bisulfite converted DNA was used for PCR
amplification, using the Taq PCR Kit (New England BioLabs) following manufacturer’s
instructions, with 10µM primers listed in Appendix Table 2. PCR primers were designed
to amplify bisulfite converted species-specific DNA and tested in-silico using the Bisearch
Web Server (http://bisearch.enzim.hu). Amplified PCR products were cloned into pGEM-
T easy vector (Promega), following manufacturer’s instructions, and competent DH5α
bacteria were transformed by heat shock. At least ten bacterial colonies were randomly
picked, incubated overnight in lysogeny broth at 37oC with agitation and plasmid DNA was
isolated using Wizard SV 96 plasmid purification system (Promega). DNA sequencing was
performed by the MRC Clinical Sciences Centre Sequencing Facility and obtained
sequences were analysed by CLC Main Workbench software (Qiagen).
2.2.5.2. 5-hmC quantification by enzyme protection assay
10µg of genomic DNA was either treated with T4 Phage β-glucosyltransferase (T4-
BGT, New England BioLabs) (T4+) or not (T4-) following manufacturer’s instructions.
Glucosylated and non-glucosylated genomic DNA were either treated with methylation
insensitive 100U of MspI restriction enzyme (New England BioLAbs), or not treated with
enzyme (mock digestion) at 37oC for 4 hours, followed by Proteinase K treatment for 30
minutes at 40oC. Primers that span the MspI restriction site were used to quantify by
quantitative PCR the T4+ and T4- samples, normalised individually to the amplification of
a control region that does not contain MspI site, and subtraction between the two
normalised levels translates into percentage 5-hmC levels. Primer sequences for enzyme
protection assay can be found in the Appendix Table 3.
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Chapter 2
2.2.6. Imaging analysis
2.2.6.1. Immunofluorescence and confocal microscopy imaging
IMR90 (HP1αGFP) fibroblasts, mouse ES (H2BmCherry) cells and heterokaryons
were grown on glass coverslips coated with 0.1% gelatin. The coverslips were washed
with PBS and the cells were fixed in 2% paraformaldehyde (Sigma) for 15 minutes. Fixed
samples were washed three times in PBS for 5 minutes and the cells were permeabilized
with 0.4% Triton X-100 (Sigma) for 5 minutes. The samples were incubated in blocking
solution [2.5% bovine serum albumin (BSA, Sigma), 0.05% Tween 20 (Sigma), normal
goat serum (Sigma) in PBS], supplemented with Alexa Fluor 647 Phalloidin (1:50 dilution,
Invitrogen), for 1 hour in humid chamber. After three washes in washing buffer (0.2% BSA,
0.05% Tween 20, in PBS) for five minutes, coverslips were mounted using Vectashield
medium supplemented with DAPI (Vector Laboratories). Samples were visualised with a
SP5 Leica laser-scanning confocal microscope and processed using Leica Application
Suite Software and Adobe Photoshop CS5.
2.2.6.2. X-gal staining
For X-gal staining all reagents were supplied from Sigma. mEG/2rB hybrids were
fixed for 15 minutes in 0.1M PBS supplemented with 5mM ethylene glycol tetraacetic acid
(EGTA), 2mM MgCl2, 0.2% gluteraldehyde, followed by washing with 0.1M PBS
supplemented with 2mM MgCl2, and incubated in staining buffer (0.1M PBS supplemented
with 2mM MgCl2, 5mM potassium ferrocyanide, 5mM potassium ferricyanide and 1mg/ml
X-gal) for 16 hours at 37oC. After three washes, stained plates were analysed by inverted
microscope with 10X objective (Leica).
2.2.7. Western Blot analysis
All reagents were supplied from Sigma unless otherwise specified. Whole cell
protein extract was prepared by cell lysis in sample buffer (50mM Tris-HCl pH6.8, 1%SDS,
10% Glycerol) by incubation at 95oC and vortexing. After quantification (Pierce BCA
assay), Bromophenol Blue and 5% β-mercaptoethanol were added (0.001% and 5%,
respectively) into the samples. Bio-Rad minigel system was used to perform Sodium
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Chapter 2
dodecyl sulphate – polyacrylamide gel electrophoresis (SDS-PAGE). 10 to 50µg of protein
was loaded on acrylamide (Bio-Rad) stacking gel (4% acrylamide, 0.125M Tris-HCl pH
6.8, 0.1% SDS, 0.1% ammonium persulphate and 0.1% tetramethylethylenediamine
(TEMED), and separated in a 10% acrylamide resolving gel (10% acrylamide, 0.4M Tris
pH8.8, 0.1%SDS, 0.1% ammonium persulphate and 0.1% TEMED), using Tris-glycine
buffer (25mM Tris, 192mM glycine, 0.1% SDS). Resolved proteins in acrylamide gels were
blotted to Protran nitrocellulose transfer membrane (Schleicher&Schuell Bioscience), in
transfer buffer (48mM Tris, 39mM glycine, 0.037% SDS and 20% methanol), using the
trans-blot semi-dry electrophoretic transfer instrument (Bio-Rad). The membranes were
incubated for 1 hour with blocking buffer (5% fat free milk powder, 20mM Tris pH 7.5,
150mM NaCl), followed by incubation with primary antibody diluted in blocking buffer for
1 hour at room temperature. After three washes in washing buffer (20mM Tris pH 7.5,
150mM NaCl, 0.1% Tween 20), membranes were incubated with horseradish peroxidase-
coupled secondary antibodies (anti-rabbit at 1:5000, anti-mouse at 1:2000 dilutions, both
from Amersham, anti-goat at 1:2000 dilution, from Santa Cruz), diluted in blocking buffer
for 1 hour. ECL Prime western blotting detection kit (Amersham) was used for signal
detection using Kodak Carestream photographic films and Kodak X-Omat Developer.
2.2.8. Chromatin Immunoprecipitation (ChIP) analysis
ChIP analysis was performed using the LowCell# ChIP Kit from Diagenode (all
reagents described below were supplied from Diagenode). Briefly, Anti-H3, Anti-H3K4me3
and IGG (as negative control) antibodies were bound to magnetic beads following
manufacturer’s instructions. Up to 100,000 cells were resuspended in PBS, fixed with
formaldehyde for 8 minutes before the solution was quenched with glycine, using the
concentrations advised in the kit. Fixed cells were washed twice in cold PBS and lysed
and sonicated (Bioruptor Plus, Diagenode) for 12 cycles of 30 seconds ON and 30
seconds pause. Sheared chromatin was overnight incubated with antibody coated
magnetic beads at 4oC. After the washes, magnetic bead-bound chromatin was eluted,
and the DNA was isolated using IPure kit (Diagenode). Isolated DNA was later processed
by quantitative PCR as described in section 2.2.4.2.
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2.2.9. Plasmid construction and delivery into ES cells
Short hairpin RNA (shRNA) plasmids were constructed using pSuper.GFP.Neo
backbone vector (Oligoengine, Figure 2.1.A). Synthetic primer pairs (sense and
antisense) that contain the shRNA targeting sequence were annealed in a reaction
mixture that contain 1X T4 ligase buffer (Invitrogen), by incubation at 90oC for 4 minutes,
and slowly cooling down to room temperature. The annealed oligo inserts were ligated
into BglII and HindIII digested pSuper.GFP.Neo vector by T4 ligation reaction (Invitrogen).
H2BmCherry sequence was kindly provided by Dr. Nobuaki Kudo. The sequence
was amplified with primers that contain XhoI and NotI sites and ligated into XhoI and NotI
digested pCAGIresPuro vector (Figure 2.1.B) (Niwa et al., 2002).
HP1αGFP sequence was kindly provided by Dr. Jesus Gil. The sequence was
amplified with primers that contain BglII and EcoRI and ligated into BglII and EcoRI
digested pMIP retroviral vector (kindly provided by Bradley Cobb, Figure 2.1.C).
All of the above mentioned ligation products were transformed into competent
DH5α bacteria by heat shock. Bacteria colonies were picked, incubated overnight in
lysogeny broth at 37oC with agitation and plasmid DNA was isolated by QIAprep Spin
Miniprep Kit (Qiagen). DNA sequencing was performed by the MRC Clinical Sciences
Centre Sequencing Facility and obtained sequences were analysed for correct insertion
by CLC Main Workbench software (Qiagen).
Figure 2.1. Vectors used for delivery. (A) short hairpin RNA sequence was cloned downstream of H1 RNA Polymerase III promoter. NeoR: Neomycin resistance Cassette, GFP: Green Fluorescence Protein. (B) H2BmCherry coding sequence was cloned downstream of Chicken Beta Actin (CBA) Promoter and upstream of IRES (Internal Ribosome Entry Site). PuroR: Puromycin resistance cassette. (C) Hp1αGFP coding sequence was cloned downstream of Murine Stem Cell Virus (MSCV) promoter.
shRNA constructs were delivered into mouse ES and mouse EG cells by
electroporation using Amaxa Nucleofector 2b system (VPH-1001, programme A-024).
PCAGIresPuro-H2BmCherry plasmid was similarly electroporated into mouse ES cells
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and Puromycin resistant mCherry positive single clones were picked and expanded. For
retroviral infection, pMSCVIresPuro-HP1αGFP plasmid was transfected into 293T cells
with helper p10A1 plasmid using calcium phosphate. Culture supernatants containing the
retrovirus were harvested post-transfection and added on IMR90 fibroblast culture
together with 2 µg/ml polybrene. Cells were treated with Puromycin (1.5µg/ml) and
expanded.
2.2.10. CRISPR/Cas9 genome editing system
2.2.10.1. CRISPR/Cas9 plasmid construction
All of the reagents used for plasmid construction were supplied from New England
BioLabs, unless otherwise specified. The plasmid used for cloning was supplied from
Addgene (px330, plasmid 42230, Figure 2.2) (Cong et al., 2013). The targeting guide RNA
sequence was designed using a bioinformatics software available online at
http://crispr.mit.edu/ as sense and antisense complementary oligonucleotides as
illustrated in Figure 2.2.
Figure 2.2. px330 vector and the guide RNA sequence. Annealed sense and antisense guide RNA sequences contain 5’ and 3’ overhangs for ligation into BbsI digested px330 plasmid. The ligation results in chimeric single guide RNA sequence, transcribed by the RNA U6 Polymerase III promoter. Chicken beta actin promoter drives the transcription of human codon optimised Streptococcus Pyogenes Cas9, which contains nuclear localisation signal (NLS) sequences on both ends.
The px330 plasmid was digested with BbsI restriction enzyme (Thermo, #ER1011)
at 37oC for 1 hour, heat inactivated, dephosphorylated by Antarctic Phosphatase, heat
inactivated and gel purified (QIAquick Gel Extraction Kit, Qiagen). In the meantime
complementary guide sequences were annealed in 1X T4 Ligase buffer (Invitrogen), by
incubating at 95oC for 3 minutes, and slowly cooling down to room temperature. After
phosphorylation by T4 PNK, annealed oligos were ligated to the linearized vector using
T4 DNA ligase (Invitrogen) and competent DH5α bacteria were transformed by heat
shock. Bacteria colonies were picked, incubated overnight in lysogeny broth at 37oC with
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agitation and plasmid DNA was isolated by QIAprep Spin Miniprep Kit (Qiagen). DNA
sequencing was performed by the MRC Clinical Sciences Centre Sequencing Facility and
obtained sequences were analysed for correct insertion by CLC Main Workbench software
(Qiagen).
2.2.10.2. Surveyor and RFLP Assays
The Surveyor assay, as described in Chapter 5.3.2, was used to detect the
presence of indel mutations at the CRISPR target locus. The target locus was PCR
amplified (primers are listed in Appendix Table 5) using Phusion High-Fidelity DNA
Polymerase (New England BioLabs) followed by PCR purification (QIAquick PCR
Purification Kit, Qiagen). 360ng of purified amplicons were mixed with 1X Taq Buffer (New
England BioLabs), denatured and reannealed [as described in (Ran et al., 2013a)] and
treated with Surveyor DNA endonuclease (Transgenomic), following manufacturer’s
instructions. Samples were run on 2% Agarose gel for detection.
The Restriction Fragment Length Polymorphism (RFLP) was used where
applicable, as described in Chapter 5.3.3. CRISPR target locus was amplified followed by
PCR purification as described above. PCR amplicons were digested by suitable restriction
enzymes and ran on 2% Agarose gel for visualisation.
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Chapter 3
Chapter 3. Pluripotency Gene Demethylation during
Reprogramming
3.1. Introduction
The fate of pluripotent cells gradually becomes restricted as they acquire specific
properties and differentiate to become part of a particular tissue or organ. Under normal
conditions, once a cell is differentiated, it appears to be epigenetically protected in the
sense that its identity remains stable in-vivo and propagated to daughters upon cell
division. We know that although cells don’t generally change their identity, cells remain
flexible, and their epigenetic status can be reset (Reik, 2007). This can be achieved by
specific experimental approaches including transcription factor induced-reprogramming
and nuclear transfer. A third method is cell fusion, where a pluripotent cell can dominantly
alter the identity of the differentiated fusion partner towards pluripotency (Yamanaka and
Blau, 2010). Our laboratory and others have shown that pluripotent mouse embryonic
stem (mES) cells can efficiently induce reprogramming of somatic cells upon fusion, over
a short period of time (Pereira et al., 2008, 2010; Tada et al., 2001). Although it has
previously been reported that DNA demethylation is critical for the acquisition of
pluripotency (Mikkelsen et al., 2008; Simonsson and Gurdon, 2004), how DNA
demethylation is achieved early in reprogramming is not yet clear. In this Chapter, I use
cell fusion-based reprogramming to examine the changing status of DNA methylation at
pluripotency associated genes during pluripotent conversion of the somatic nucleus.
3.2. Reprogramming of human B lymphocytes upon fusion with mouse embryonic stem cells
Our laboratory has previously demonstrated that interspecies fusion between
mouse ES and human B lymphocytes results in efficient reprogramming of the somatic
nuclei, and that this happens at the early heterokaryon stage, in which discrete nuclei
originating from both fusion partners are apparent (Pereira et al., 2008, 2010) (Figure
3.1.A). This transient period lasts up to 72 hours after fusion and terminates when the two
nuclei merge to form a tetraploid hybrid. Because of genetic incompatibility, although
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interspecies hybrids can divide, they are not karyotypically stable and are not viable for
long-term. In addition, although cell division does not occur in heterokaryons, we have
recently shown that DNA synthesis is widespread at this stage, and that it has a positive
impact on reprogramming efficiency (Tsubouchi et al., 2013).
To begin to examine DNA methylation changes during cell fusion-based
reprogramming, I fused mouse ES cells (E14tg2A) and human B lymphocytes (Epstein -
Barr virus transformed adult B cells), using polyethylene glycol (PEG). The advantage of
performing interspecies fusion is that it allows us to analyse species-specific gene
expression. By using primers specific for human genes including pluripotency factors
(OCT4, NANOG, CRIPTO), I analysed the induction of human pluripotency gene
expression by performing qRT-PCR in human B lymphocytes (0h, before fusion) and in
heterokaryons (72h after fusion) (Figure 3.1.B).
Figure 3.1. Interspecies cell fusion and reprogramming of human B lymphocyte by mES cells. (A) Schematic representation of interspecies fusion between mES cells (light grey) and human B lymphocytes (dark grey). Nuclei originating from both cell types are separate at the heterokaryon stage (persisting for ~72 hours after fusion) and tetraploid hybrids are formed upon nuclei merging. (B) qRT-PCR analysis of pluripotency factor gene expression (OCT4, NANOG and CRIPTO) in hB lymphocytes before (0h) and at 72 hours after fusion with mES cells. Data were normalised to human GAPDH and represent mean and SEM of 3 different experiments.
Transcript analysis revealed that pluripotency network is silent in B lymphocytes,
as evidenced by the lack of expression of OCT4, NANOG, and CRIPTO before fusion (0h)
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(Figure 3.1.B). This profile was altered as early as 72 hours after fusion with mES cells,
where I observed significant upregulation of pluripotency-associated gene expression
originating from human nucleus. These results together with previous work (Pereira et al.,
2008, 2010) show that cell fusion mediated reprogramming provides an opportunity to
study the molecular details of early stages of pluripotent induction.
3.3. DNA methylation profiles of pluripotency associated genes in human B lymphocytes and human ES cells
Among many of the epigenetic regulators, DNA methylation is a stable modification
and is associated with inhibition of gene expression (Klose and Bird, 2006). To understand
the status of DNA methylation at the promoters of pluripotency-associated genes in
human B lymphocytes and human ES cells (H7), I conducted bisulfite sequencing analysis
(Figure 3.2). This method enables the assessment of individual 5-methylcytosine residues
in DNA strands. It is based on bisulfite-induced modification of genomic DNA, where
cytosine molecules are converted to uracil while 5-methylcytosine remains nonreactive
Figure 3.2. Bisulfite sequencing of OCT4, NANOG and CRIPTO promoters in human B lymphocytes and human ES cells. Genomic DNA from human B lymphocytes (top) and human ES cells (bottom) were bisulfite converted, and PCR amplicons for OCT4, NANOG and CRIPTO promoters were sequenced. Transcription start sites for each gene were taken as reference and relative positions of CpG sites were indicated. Unfilled (white) circles and filled (black) circles represent un-methylated and methylated CpGs respectively.
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that can be distinguished by sequencing of the PCR products (Frommer et al., 1992). H7
human ES cells express high levels of pluripotency-associated factors, including OCT4,
and have the capacity to differentiate into cell types from all the three germ layers
(Thomson, 1998; Xu et al., 2001). In differentiated cells (human B lymphocytes), the
promoters of pluripotency-associated OCT4, NANOG and CRIPTO genes were highly
methylated (Figure 3.2, top). Consistently, these genes lacked CpG methylation in
pluripotent human ES cells (Figure 3.2, bottom).
Overall, these data suggest that there is a correlation between pluripotency-
associated gene expression and lack of cytosine methylation.
3.4. Changes in DNA methylation of OCT4 accompanies reprogramming but the extent is variable
Upon fusion with mES cells, human B cell nucleus is rapidly reprogrammed and
starts expressing components of pluripotency network genes. To assess how DNA
methylation signatures were altered in the course of pluripotent reactivation, I conducted
genomic bisulfite sequencing using human specific primers for amplification and
sequencing of the OCT4 promoter. Figure 3.3 demonstrates a single cell fusion
experiment between mES cells and human B cells, where I analysed gene expression
levels and methylation status of OCT4 in heterokaryons after 72 hours of fusion.
Successful reprogramming was evident as shown by transcript analysis, and this was
accompanied by partial demethylation of human OCT4 promoter (Figure 3.3).
Figure 3.3. Transcript analysis and bisulfite sequencing of human OCT4 in heterokaryons after 72 hours of fusion. (Left) qRT-PCR analysis of pluripotency factor gene expression (OCT4, NANOG and CRIPTO) in hB lymphocytes before (0h) and at 72 hours after fusion with mES cells, data were normalised to human GAPDH. (Right) Bisulfite sequencing analysis on human OCT4 promoter in heterokaryons after 72 hours of fusion, where relative positions of CpG sites to transcription start site were indicated. Unfilled (white) circles and filled (black) circles represent un-methylated and methylated CpGs respectively.
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Even though this data demonstrated a relation between demethylation and
induction of gene expression in heterokaryons, a second experiment I conducted showed
no correspondence between OCT4 expression and cytosine demethylation at the OCT4
promoter (Figure 3.4.A). I performed this analysis in three more experiments, but again
saw no strong correlation between gene reactivation and DNA demethylation, at least at
the population level in heterokaryons (Figure 3.4.B).
Figure 3.4. Transcript analysis and bisulfite sequencing of human OCT4 in heterokaryons after 72 hours of fusion in four different experiments. (A) (Left) qRT-PCR analysis of pluripotency factor gene expression (OCT4, NANOG and CRIPTO) in hB lymphocytes before (0h) and at 72 hours after fusion with mES cells, data were normalised to human GAPDH. (Right) Bisulfite sequencing analysis on human OCT4 promoter in heterokaryons after 72 hours of fusion, where relative positions of CpG sites to transcription start site were indicated. Unfilled (white) circles and filled (black) circles represent un-methylated and methylated CpGs respectively. (B) Same analysis as in (A) in 3 more experiments.
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3.5. Reprogramming human fibroblasts and OCT4 induction without detectable changes in DNA methylation
In order to repeat previous investigations in a different cell type, I used human
fibroblasts as fusion partners of mouse ES cells. I generated human fibroblasts (IMR90)
and mouse ES cells constitutively expressing HP1α-GFP and H2B-mCherry respectively.
After two days of fusion, I isolated double positive heterokaryons by Fluorescence
Activated Cell Sorting (FACS) and plated them back for two more days of culture. Then I
re-sorted double positive heterokaryons, and used these to do transcript and bisulfite
sequencing analyses (Figure 3.5.A).
Gene expression analysis after 96 hours of fusion demonstrated upregulation of
pluripotency-associated genes OCT4, NANOG and CRIPTO originating from human
fibroblast (Figure 3.5.B). Despite this, bisulfite analysis showed that the human OCT4
promoter remained methylated at this stage as assessed in two different experiments
(Figure 3.5.C). Collectively these data suggest that demethylation of the human OCT4
promoter may not be essential for expression.
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Figure 3.5. Fusion of human fibroblasts with mouse ES cells and transcript and bisulfite sequencing analyses. (A) Schematic demonstration of the experimental approach where human fibroblasts (IMR90) expressing HP1α-GFP were fused with mouse ES cells expressing H2B-mCherry. Double positive cells (GFP+&mCherry+) were sorted after 2 days of fusion (Q2 population) and plated. 2 days later, same sorting settings were applied and the re-sorted population was analysed for gene expression and methylation profiles. (B) qRT-PCR analysis of pluripotency factor gene expression (OCT4, NANOG and CRIPTO) in hFibroblasts before (0h) and at 96 hours after fusion with mES cells. Data were normalised to human GAPDH and represent mean and SEM of 2 different experiments. (C) Bisulfite sequencing analysis on human OCT4 promoter in human fibroblasts before (0h) and at 96 hours after fusion (in 2 independent experiments), where relative positions of CpG sites to transcription start site were indicated. Unfilled (white) circles and filled (black) circles represent un-methylated and methylated CpGs respectively.
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3.6. No evidence of DNA methylation changes at site upstream of the OCT4 transcription start site.
Using similar system, settings and cell lines, Bhutani et al. have previously reported
DNA demethylation at the OCT4 upstream region upon fusion (Bhutani et al., 2010). The
analysed region in this report is localised between the OCT4 promoter and the proximal
enhancer sites (Nordhoff et al., 2001) (Figure 3.6.A). Therefore, I investigated the
methylation status of the same locus in human fibroblast DNA before and after fusion.
Human fibroblast DNA was fully methylated as expected, however I did not detect a loss
of DNA methylation after fusion with mES cells as demonstrated in two independent
experiments (Figure 3.6.B). Interestingly, this region (unlike the promoter), was mostly
methylated in H7 human ES cells (Figure 3.6.C). In addition, the same DNA methylation
pattern at the OCT4 upstream was previously reported in pluripotent human embryonal
carcinoma cells (Deb-Rinker et al., 2005), calling into question the published role of this
site in regulating OCT4 expression.
Figure 3.6. Bisulfite sequencing of human OCT4 upstream region in human fibroblasts before and after fusion and in human ES cells. (A) Schematic representation of human OCT4 locus. The arrow indicates transcription start site (TSS), and red lines indicate locations of the CpG sites relative to the TSS. Location of the five CpG sites analysed by Bhutani et al. and here is marked. Bisulfite sequencing in (B) human fibroblasts (IMR90) before (0h) and at 96 hours after fusion (in 2 independent experiments) and in (C) human embryonic stem cells (H7) where relative positions of CpG sites to transcription start site were indicated. Unfilled (white) circles and filled (black) circles represent un-methylated and methylated CpGs respectively.
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3.7. DNA demethylation kinetics of somatic Oct4 transgene in reprogrammed mouse hybrids.
To better understand the relationship between Oct4 re-expression and locus
demethylation during reprogramming, I made use of a mouse indicator B cell line in which
re-expression of Oct4 was evident by GFP expression. In order to track reprogrammed
cells, I used B cell lines isolated from transgenic mice bearing GFP coding sequence
under the control of 18 kb genomic mouse Oct4 fragment (GOF-18/ΔPE/GFP) (Yoshimizu
et al., 1999) (Figure 3.5.7, left). This fragment was previously shown to be sufficient to
recapitulate endogenous expression patterns of Oct4 in embryonic development, thus
GFP expression under its regulation can be considered as a direct sign for pluripotency
(Yeom et al., 1996). Upon fusion with mouse ES cells, the silent transgene carried by the
mouse B cells was reactivated, leading to a prominent GFP expression in hybrid colonies
(Figure 3.7.A, right). To assess the methylation status of the transgene in mouse B cells
and to track how it is altered during pluripotency induction, I conducted bisulfite
sequencing on the region which spans exogenous Oct4 promoter site and GFP coding
sequence. As expected, the transgene was fully methylated in mouse B lymphocytes
(Figure 3.7.B), in line with the lack of GFP transgene expression (data not shown). To
examine the kinetics of DNA demethylation of the transgene in the course of
reprogramming, I isolated GFP expressing hybrid cells by FACS at different time points
(3, 6 and 13 days after fusion). Then I conducted bisulfite conversion on each sample and
sequenced the exogenous Oct4-GFP genomic fragments (by using PCR primers that
specifically span the promoter/GFP region). Analysis of DNA methylation revealed that
among GFP positive cells isolated 3 days after fusion, partial demethylation was seen. By
6 days, DNA methylation was completely lost, and the region remained un-methylated at
least until 13 days of fusion. (Figure 3.7.C).
These data demonstrated that reprogramming was rapidly induced following intra-
species cell fusion and resulted in GFP expression under the control of the exogenous
Oct4 promoter. Interestingly, this was accompanied by partial demethylation (as observed
3 days after fusion), followed later by complete loss of DNA methylation.
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Figure 3.7. DNA demethylation kinetics upon reprogramming in mouse hybrids. (A) (Left) Schematic representation of interspecies fusion between mES cells and mouse B lymphocytes carrying an exogenous Oct4-GFP fragment. Nuclei originating from both cell types are separate at the heterokaryon stage (persisting for ~72 hours after fusion) and tetraploid hybrids are formed upon nuclei merging. (Right) Bright field and fluorescent microscopy images of a hybrid colony, cultured for 10 days after fusion. (B) Bisulfite sequencing analysis on Oct4-GFP transgene in mouse B lymphocytes and (C) in GFP expressing hybrids at indicated time points. Unfilled (white) circles and filled (black) circles represent un-methylated and methylated CpGs respectively.
3.8. Summary and Discussion
Cell fusion allows an assessment of the impact of one cell type on another. This is
achieved by trans-acting factor binding to DNA sites that can lead to reversing of cell fate.
Our laboratory and others have previously demonstrated that differentiated cells can
successfully acquire a pluripotent-like identity upon fusion with mouse embryonic stem
cells (Pereira et al., 2008, 2010; Tada et al., 2001). In this Chapter I have re-examined
these findings with a focus on DNA methylation dynamics that occur during
reprogramming.
The activation of silenced genes in interspecies heterokaryons was first
demonstrated by fusing mouse muscle cells with human amniotic cells, where initiation of
previously silent human muscle-specific gene expression was detected (Blau et al., 1983).
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Here I confirm that a silent pluripotent program can be reactivated in somatic human B
lymphocytes after fusion with mouse ES cells (Pereira et al., 2008). Pluripotency-related
gene activation is detected as early as 24 hours of fusion in heterokaryons.
How trans-acting factors remodel DNA elements and relieve repressive chromatin
structure is still largely unknown. DNA methylation is a key epigenetic mechanism involved
in stable transcriptional silencing of gene expression. Here I demonstrated that DNA
methylation was inversely correlated with pluripotency-associated gene expression in
human B lymphocytes. During reprogramming, it might be assumed that DNA
demethylation would occur in parallel to gene re-activation. Indeed, nuclear
transplantation experiments have revealed that DNA demethylation is necessary and
precedes Oct4 transcription during pluripotent conversion (Simonsson and Gurdon, 2004).
However, in my interspecies heterokaryon experiments I did not see a direct correlation
between promoter DNA demethylation and transcriptional activation of OCT4 gene.
Instead, I observed either lack of, or partial demethylation of OCT4, despite OCT4 gene
re-expression. Similar to these results, a study has demonstrated that when 293T cells
were treated by pluripotent embryonic carcinoma cell extract, OCT4 gene reactivation was
accompanied by a mosaic demethylation pattern after 4 weeks of culture (Freberg et al.,
2007). In a related study, Foshay et. al. analysed changes in gene expression in rat
fibroblasts after fusion with mouse ES cells, and concluded that reprogramming of cis-
silenced genes occurs with rather slow kinetics and requires DNA synthesis (Foshay et
al., 2012). These authors suggested that this was due to repressive histone marks, and
the presence of DNA methylation around genes such as Oct4 and Nanog. Moreover, the
rate of Oct4 promoter demethylation was slow when analysed at different times after
fusion (at population level), and was only complete in selected (and cultured) stable
hybrids (Foshay et al., 2012). One explanation for my observations might be that the
initiation of OCT4 gene expression may take place even in the presence of methylated
CpGs and be regulated by epigenetic features and mechanisms such as active histone
modifications (See Chapter 5.4 for discussion). A second rationale for the lack of
demethylation might be due to the fact that the analysis was conducted at a population
level; in a group of heterokaryons, it is possible that some fused cells may not undergo
reprogramming. This would obscure the bisulfite analysis, even though mRNA transcripts
for OCT4 can be detected in the population. To overcome this issue, it would be crucial to
conduct a single heterokaryon gene expression analysis coupled with single cell DNA
methylation profiling [which are now possible with the advent of microfluidic platforms
(Lorthongpanich et al., 2013)]. I am currently looking at the feasibility of this analysis.
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Previously, Bhutani et. al. showed demethylation of OCT4 upstream region in
human fibroblasts soon after fusion with mouse ES cells (Bhutani et al., 2010). The
authors claimed that this was mediated by AID, as Aid knockdown impaired
reprogramming and OCT4 demethylation. Using similar approach and cell lines, I did not
observe demethylation at the OCT4 transcription initiation site, nor in the upstream region
highlighted in the research article. Instead, I detected CpG methylation at this upstream
locus in human ES cells, which raises a major concern about the validity of this region in
regulating gene expression. Furthermore, the role of AID in DNA demethylation and
reprogramming has recently been re-evaluated. For example, in vitro examination
revealed a substantially reduced activity of AID on 5mC relative to cytosine, caused by
the adverse effect of steric hindrance on its function (Nabel et al., 2012). In addition, iPS
cells can successfully be generated from Aid deficient fibroblasts without any significant
changes in the DNA methylome (Habib et al., 2014; Shimamoto et al., 2014). Consistent
with these reports, I did not see any effect of Aid knockdown in reprogramming upon cell-
fusion and Aid mRNA levels in ES cells were already very low (data not shown), as also
observed in a similar study (Foshay et al., 2012).
I have demonstrated the kinetics of DNA demethylation of an exogenous Oct4-
GFP fragment integrated into mouse B cell genome, upon fusion with mouse ES cells.
With this system, successful reprogramming can be visualised using GFP marker as
previously demonstrated (Do and Schöler, 2004; Han et al., 2008; Silva et al., 2006; Wong
et al., 2008). Although the transgene was silent and fully methylated in mouse B cells, loss
of methylation was evident by 3 days of fusion, and was completed by 6 days. In these
experiments I sorted GFP-positive cells and therefore the bisulfite sequencing is biased
to successfully reprogrammed cells. Interestingly, at Day 3, there was a substantial
number of methylated clones, despite the expression of GFP. This can be attributed to a
couple of different possibilities. First, the transgene might have multiple copies in the
mouse B genome, and according to where the transgene was integrated, different kinetics
of demethylation would be encountered according to genomic context. Secondly, although
DNA methylation is symmetrical, in the course of loss of methylation one strand might be
demethylated while the complementary strand remains unmodified. This profile would be
expected to be seen during passive demethylation, and could be visualised using a hairpin
bisulfite sequencing strategy (Laird et al., 2004). Thirdly, the partial demethylation might
reflect ongoing conversion of 5-mC to 5-hmC and to subsequent derivatives either
actively, or through passive mechanisms.
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In conclusion my results demonstrated that reprogramming of somatic nuclei by
cellular fusion is a rapid process, evidenced by initiation of pluripotency associated gene
expression. Although this system allows us to study the early events that contribute to the
pluripotent conversion, exactly how gene reactivation is synchronised with DNA
demethylation is unclear due to heterogeneity in the heterokaryon population. One
contribution to this heterogeneity comes from the various cell cycle stages of the fusion
partners. Indeed we have shown that mouse ES cells in S/G2 phase are more efficient in
reprogramming human B lymphocytes upon fusion (Tsubouchi et al., 2013). This is due to
their capacity to induce DNA synthesis in the somatic nuclei. To better understand
sequential events taking place during early stages of reprogramming, it is important to
collect data from single heterokaryons in a high-throughput manner to collect transcript,
DNA methylation and reprogramming (imaging) analysis. The combination of those would
provide us a global understanding of progressive conversion towards pluripotency.
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Chapter 4
Chapter 4. Mechanisms of Imprint Erasure in Somatic
Cells mediated by Embryonic Germ Cell Fusion
4.1. Introduction
It has previously been shown that despite their dominant pluripotent
reprogramming potential, mouse ES cells were not able to induce DNA demethylation of
imprinted regions in the somatic genome upon fusion (Tada et al., 2001). This is especially
interesting considering that some mouse embryonic germ (mEG) cells derived from
primordial germ cells, can on the other hand, re-set imprints (Tada et al., 1997). In this
Chapter, I analyse the kinetics of imprint erasure in somatic cells following fusion with
mouse EG cells, and show some of the molecular mechanisms likely to be involved in this
process.
4.2. Imprint erasure in somatic cells induced by embryonic germ cell fusion.
Embryonic germ (EG) cells are pluripotent in-vitro counterparts of primordial germ
cells (PGCs). They express many pluripotency-associated factors similar to ES cells (Mise
et al., 2008), but interestingly, EG cells exhibit genome-wide DNA hypomethylation that
includes imprinted domains (Labosky et al., 1994; Tada et al., 1998). An additional
difference is that EG cells, unlike ES cells, were reported to induce imprint erasure in
thymocyte DNA upon fusion in reprogrammed hybrids (Tada et al., 1997). With Francesco
Piccolo, I have examined the DNA methylation status of imprinted H19 loci in
reprogrammed hybrids generated between mouse B lymphocytes (bearing Oct4-GFP
transgene) and mouse EG cells. To assess whether imprint resetting would occur and if
so what its kinetics would be, I isolated GFP positive hybrids at 7, 12 and 21 days after
fusion, and conducted bisulfite sequencing analysis on those samples (Figure 4.1). The
hypomethylation observed in 21 day hybrids confirmed the erasure of imprints, induced
by mouse EG cells (1.6%, compared to initial 31.2%). Interestingly, methylation level was
maintained until at least 7 days of fusion (43%), and exhibited a gradual decrease after
this point (17.4% at 12 Days, Figure 4.1).
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Figure 4.1. CpG methylation analysis of imprinted H19 locus upon pluripotent reprogramming mediated by mouse EG cells. Genomic bisulfite sequencing of H19 ICR in mEG and mB lymphocytes (1:1 mixture) before fusion (0h) and in GFP sorted hybrids at 7, 12 and 21 Days after fusion. Corresponding methylation levels are depicted as percentage. Unfilled (white) circles and filled (black) circles represent un-methylated and methylated CpGs respectively.
4.3. Using dual reporter (2rB) somatic cells to assess the kinetics of imprint erasure during EG-reprogramming.
To elucidate whether EG cells were able to functionally reset imprinting in somatic
cells, I used an alternative mouse B lymphocyte cell line (2rB) as a fusion partner. The
2rB cells, in addition to Oct4-GFP transgene, possess a maternal LacZ knock-in allele of
imprinted Peg1 gene (Peg1M-β-gal) (Lefebvre et al., 1998). Peg1 is paternally expressed,
while the maternal allele is silent. In this setting β-galactosidase activity would be detected
only if the maternal imprinting is reset and the gene is reactivated. To first assess whether
Peg1 imprint can be reset by EG cell fusion (as in the case of H19), I analysed methylation
status of the Peg1 DMR in hybrids generated between mouse EG and 2rB cells. I isolated
GFP positive hybrids at consecutive time points (7, 12 and 21 days of fusion) and
conducted bisulfite sequencing. Although DNA methylation was maintained until at least
7 day hybrids (30%), it gradually decreased at further time points (13.3% at Day 12 and
7% at Day 21, Figure 4.2). This result demonstrated that, Peg1 imprint can efficiently be
re-set in EG/2rB hybrids.
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Figure 4.2. CpG methylation analysis of imprinted Peg1 locus upon pluripotent reprogramming mediated by moue EG cells. Genomic bisulfite sequencing of H19 ICR in mEG and 2rB lymphocytes and in GFP sorted hybrids at 7, 12 and 21 Days after fusion. Corresponding methylation levels are depicted as percentage. Unfilled (white) circles and filled (black) circles represent un-methylated and methylated CpGs respectively.
To assess the functional outcome of EG-mediated imprint erasure at the 2rB
somatic nucleus, Karen Brown and David Landeira investigated Peg1 driven β-
galactosidase (β-gal) activity in hybrids. In GFP positive hybrids at 12 days of fusion, β-
gal activity was not detected (as assessed by X-gal staining, Figure 4.3.A-B), indicating
that Peg1M-β-gal remained silent. This result was expected as Peg1 is only expressed in
differentiated cells (Lefebvre et al., 1998). To this end, EG/2rB hybrids were differentiated
by removing LIF from the culture media. Differentiated 22-day hybrids were positive for β-
gal activity as analysed by X-gal staining (Figure 4.3.A-B). This indicated that Peg1 gene
could be expressed from previously methylated 2rB maternal allele that was re-set in
hybrids by EG-cell fusion, however this was not observed in hybrids formed by ES-cell
fusion (Figure 4.3.C, see Chapter 4.5)
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Figure 4.3. Functional resetting of somatic imprints mediated by mouse EG cells. (A) Schematic illustration of the experimental strategy to assess imprint erasure in 2rB nucleus in EG-cell hybrids. In addition to Oct4-GFP transgene, 2rB cells carry a maternal LacZ knock-in allele of Peg1. (B) A mEG/2rB hybrid colony expresses GFP which is driven by an Oct4 promoter, however lacks β-gal activity as assessed by X-gal staining). 10 days after differentiation by LIF removal, EG/2rB hybrids are positive for β-gal activity (blue). (C) mES/2rB hybrids, differentiated by LIF removal for 10 days after 12 days of fusion, negative for β-gal activity as assessed by X-gal staining.
4.4. EG cell capacity to induce demethylation is not restricted to imprinted genes.
To understand whether EG cell-induced erasure of DNA methylation is limited to
imprinted genes or occurs on additional sites in the somatic nucleus, we analysed the
methylation status of long interspersed element (LINE) repeats in hybrids. Mouse EG cells
exhibited less DNA methylation at LINE1 repeats compared to 2rB lymphocytes. In GFP
positive hybrids 21 days of fusion of 2rB and EG cells, LINE1 methylation was significantly
reduced (15.8%) (Figure 4.4).
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Figure 4.4. CpG methylation analysis of LINE1 repeats upon pluripotent reprogramming mediated by mouse EG cells. Genomic bisulfite sequencing of LINE1 in mouse EG and 2rB lymphocytes and in GFP sorted hybrids at 21 days after fusion. Corresponding methylation levels are depicted in as percentage. Unfilled (white) circles and filled (black) circles represent un-methylated and methylated CpGs respectively.
4.5. Imprint erasure is not seen in fusions with mouse ES cells or female ES cells that are globally hypomethylated.
In 2001, Tada and colleagues showed that in hybrids generated between
thymocytes and mES cells, although reprogrammed, imprinted domains remained
methylated. By conducting methylation-sensitive restriction enzyme analysis, they showed
that the imprint control region (ICR) of the H19 locus maintained CpG methylation in hybrid
clones (Tada et al., 2001).
With Francesco Piccolo, we confirmed that, in addition to H19, Peg3 and Gtl2/Dlk1
ICRs were also produced in mouse ES x mouse B cell fusions. Bisulfite analysis on H19,
Peg3 and Gtl2/Dlk1 imprinted loci in 1 : 1 mixture of mES and mouse B lymphocytes
bearing an Oct4-GFP transgene (denoted as 0h, before fusion) demonstrated initial level
of methylation (56%, 32% and 35%, respectively, Figure 4.5). It is important to note that
although expected 50% methylation of imprinting is observed in mouse B cells, this level
is around 25-30% in mES cells (Piccolo et al., 2013). Then we conducted bisulfite analysis
at the same three loci in GFP positive reprogrammed hybrids generated between mES
and mB lymphocytes after 21 days of fusion. The survey revealed that ICR methylations
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of H19, Peg3 and Gtl2/Dlk1 genes were not altered in pluripotent hybrids. (45%, 27% and
46%, respectively, Figure 4.5).
Figure 4.5. CpG methylation analysis of imprinted H19, Peg3, Gtl2/Dlk1 loci upon pluripotent reprogramming induced by mouse ES cells. Genomic bisulfite sequencing of mES and mB lymphocytes (1:1 mixture) before fusion (0h) and corresponding methylation levels (depicted as percentage) (Top). Genomic bisulfite sequencing of 21 days hybrids of mES fused with mB lymphocytes and corresponding methylation levels (depicted as percentage) (Bottom). Unfilled (white) circles and filled (black) circles represent un-methylated and methylated CpGs respectively.
To understand whether the genome-wide methylation status of the mES cells
would have an effect on imprint erasure, I repeated this analysis in hybrids formed
between mB lymphocytes and a female mouse embryonic stem cell line, Pgk12.1
(Zvetkova et al., 2005). Female mES cell lines were previously shown to exhibit global
DNA hypomethylation, including imprint control regions (Shovlin et al., 2008; Zvetkova et
al., 2005). Bisulfite sequencing revealed that H19 and Peg3 lost CpG methylation and
Gtl2/Dlk1 contained moderate levels of DNA methylation in Pgk12.1 cells (9%, 0% and
36%, respectively, Figure 4.6.A). Then I analysed whether these cells can induce imprint
erasure upon fusion with mouse B lymphocytes. Initial methylation levels of 1 : 1 mixture
of Pgk12.1 and mB cells (before fusion) were 26.1%, 22.1% and 37.1% for H19, Peg3 and
Gtl2/Dlk1, respectively (Figure 4.6.B, black dots). Bisulfite analysis of three different hybrid
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clones at 21 days after fusion showed that no significant change in methylation levels
occurred in these three loci (Figure 4.6.B, red dots). Overall, these data demonstrated that
although mES cells (male or female origin) can induce pluripotent conversion in somatic
cells upon fusion, they fail to reset imprinting.
Figure 4.6. CpG methylation analysis of imprinted H19, Peg3, Gtl2/Dlk1 loci upon pluripotent reprogramming induced by Pgk12.1 female mouse ES cells. (A) Genomic bisulfite analysis of H19, Peg3 and Gtl2/Dlk1 in Pgk12.1 female ES cells. Unfilled (white) circles and filled (black) circles represent un-methylated and methylated CpGs respectively. (B) Genomic bisulfite analysis of H19, Peg3 and Gtl2/Dlk1 before (0h, black dots) and after fusion where three individual hybrid clones were examined (21 Days, red dots). Each dot represents the percentage methylation acquired from at least 16 respective bisulfite converted DNA sequences. Percentage methylation values at 21 Days hybrids for H19 are 18.4%, 17.7% and 25%; for Peg3 are 23.1%, 29.3% and 14.7%; for Gtl2/Dlk1 are 17.9%, 31.1% and 29.2%.
4.6. Hydroxymethylation at imprinted loci upon fusion with mouse EG cells.
In mouse EG cell hybrids generated with mouse B lymphocytes, we noticed that
imprint erasure started long after the induction of gene re-expression. The gradual loss of
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ICR methylation led us hypothesise that this process might involve active oxidation of 5-
methylcytosine (to 5-hydroxymethylcytosine “5-hmC”) coupled to replication-dependent
passive DNA demethylation (Bagci and Fisher, 2013). To explore this, and to understand
early events that might contribute to imprint erasure, we conducted interspecies fusion
between mouse EG cells and human B lymphocytes. This provided us the opportunity to
specifically analyse modifications that may occur to human DNA. Francesco Piccolo in the
lab conducted restriction enzyme protection assays to quantify 5-hmC levels at several
imprinted loci. These results demonstrated that 5-hmC accumulated at human ICRs of
H19, Peg3 and SNRPN/SNURF imprinted genes, as early as 48 hours after fusion (Figure
4.7), providing a possible route for active conversion of 5-mC to 5-hmC at these ICRs.
Figure 4.7. Acquisition of 5-hmC at human B lymphocyte ICRs upon fusion with mouse EG cells. qPCR analysis of restriction enzyme protection assay (see Materials&Methods) to quantify 5hmC levels at human ICRs before (0h) and at 48 and 72 hours after fusion with mouse EG cells.
4.7. Tet regulated 5-mC oxidation at imprinted loci upon fusion with mouse EG cells.
Oxidation of 5-methylcytosine is catalysed by the mammalian TET family
members, composed of paralogous Tet1, Tet2 and Tet3 proteins that share significant
homology (Tahiliani et al., 2009). Pluripotent cells mainly express Tet1 and Tet2 proteins
and have been shown to be responsible for the genomic abundance of 5-hmC (Ito et al.,
2010), while Tet3 has been reported to play the major role in conversion of 5-mC into 5-
hmC in the pre-implantation embryo (Gu et al., 2011; Iqbal et al., 2011; Wossidlo et al.,
2011). In mouse EG cells, I found that both proteins were as abundant as in mES cells as
detected by western blotting. As expected TET proteins were not detected in differentiated
mB lymphocytes (Figure 4.8.A). To investigate whether Tet1 and Tet2 proteins were
involved in the acquisition of 5-hmC at somatic ICRs in heterokaryons, I used RNA
interference to separately downregulate Tet1 and Tet2 expression in mouse EG cells
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(shTet1 and shTet2). This resulted in efficient depletion of Tet1 and Tet2 proteins,
detected both by Western Blotting and qRT-PCR (Figure 4.8.A-B). Upon downregulation
of either of the genes, pluripotency-associated Oct4 and Nanog gene expression were not
significantly altered, implying that these cells retained pluripotency (Figure 4.8.B).
Following this, together with Francesco Piccolo, we fused shRNA transfected mEG cells
with human B lymphocytes, and quantified the change in the levels of 5-hmC at the ICRs
of H19, Peg3 and SNRPN/SNURF imprinted genes (48h and 72h, Figure 4.8.C). The
analysis demonstrated that Tet1 depletion resulted in lack of 5-hmC acquisition at the
corresponding ICRs compared to the control fusion, whereas Tet2 downregulation did not
result in a significant change (Figure 4.8.C).
These data suggested that imprint erasure was dependent, at least to some extent
on TET1 activity. Interestingly, although TET1 was present in mouse ES cells, we did not
observe 5-hmC accumulation at the ICRs when the B cells were fused with mES cells,
rather than EG cells (Piccolo et al., 2013). It is important to note that, Tet1 depletion in
mEG cells did not however interfere with their pluripotent reprogramming activity (Piccolo
et al., 2013).
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Figure 4.8. Roles of TET proteins in the acquisition of 5-hmC at human B lymphocyte ICRs upon fusion with mouse EG cells. (A) Western Blot detection of Tet1 (left), Tet2 (right), Oct4, Nanog and Tubulin in whole cell extracts of mES, mB and mEG cells upon transfection with shControl, shTet1 (left), or shTet2 (right) RNA interference plasmids. Tubulin antibody was used as a loading control (See Materials&Methods for details). (B) qRT-PCR analysis of Tet1 (left), Tet2 (right), Oct4 and Nanog in mEG cells, 72 hours after transfection with either empty (shCtrl, grey bars) or shTet1 (green bars, left) or shTet2 (blue bars, right) plasmids. Data were normalised to mouse Ubc and represent mean and SEM of four to five independent experiments. (C) qPCR analysis of restriction enzyme protection assay (see Materials&Methods) to quantify 5hmC levels at human ICRs before (0h) and at 48 and 72 hours after fusion with mEG cells transfected either by shTet1 (green dots, upper) or shTet2 (blue dots, lower) plasmids compared with shControl (grey dots) transfected mEG cells.
4.8. Summary and Discussion
In this Chapter, by using cell fusion system, I showed that although mES and
mouse EG cells can both efficiently reprogram somatic cells towards pluripotency, only
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mouse EG cells possess the ability to induce imprint erasure. This contrast is especially
interesting considering that both cell types are similar at the level of their transcriptomes
(Leitch et al., 2013). It is important to note that, mouse EG cells we used in our analyses
were derived late in the PGC development (embryonic day 12.5). We also discovered that
these cells can induce imprint reprogramming only at early passage number (< passage
30) (Piccolo et al., 2013). Although we do not yet understand the reasons behind this loss
of ability, culture conditions have been shown to lead to major transcriptome and
methylome changes in these cells that may alter their intrinsic properties (Leitch et al.,
2013). In addition, other EG cells isolated earlier during PGC development do not appear
to possess this imprint erasing capability although they can reprogram (Piccolo et al.,
2013).
Pluripotent conversion of somatic nucleus upon fusion with mouse EG cells occurs
early at the heterokaryon stage. Using mB lymphocytes that contain Oct4-GFP transgene
as fusion partners, I was able to obtain GFP positive hybrids as early as 3 days of fusion.
Here I showed that the imprinted genes are “tagged” by 5-mC oxidation at the early stages
of mouse EG cell induced reprogramming and that this is mediated by Tet1 protein.
However, I also determined that Tet1 is present at similar levels in mES cells. This shows
that imprint resetting induced by mouse EG cell fusion cannot be attributed solely to the
presence of Tet proteins. Over the last years, several studies reported that Tet family
members can interact with various proteins or be part of protein complexes. Examples
include interactions with SIN3A (Williams et al., 2011), Mbd3/NURD complex (Yildirim et
al., 2011), NANOG (Costa et al., 2013), O-linked N-acetylglucosamine transferase (Chen
et al., 2013b; Vella et al., 2013), and long non-coding RNA TARID together with Gadd45a
(Arab et al., 2014). It would be important to compare Tet interaction partners in mES and
mouse EG cells, also to do profiling by mass spectroscopy of EG cells at early and late
passages, to shed light on the discrepancy of their imprint resetting potentials.
Investigation of imprint erasure kinetics in the hybrids formed between mouse EG
cells and mB lymphocytes revealed an interesting result. It was not until at least 7 days
after fusion that demethylation at the somatic ICRs was detected that gradually continued
until 21 days after fusion. This delay of imprint erasure compared to pluripotent
reprogramming can be considered as a reminiscent of PGC development in the embryo.
PGC precursors can first be detected in the epiblast at E6.25 by the expression of specific
marker Blimp1 (Ohinata et al., 2005). By E7.5, Oct4 expression is restricted to the newly
emerged PGCs in the embryo and PGCs continue to express Oct4 during their migration
to form the genital ridges. (Scholer et al., 1990; Yeom et al., 1996). In the meantime,
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waves of genome-wide DNA demethylation accompanies major changes in the chromatin
context that is reminiscent of acquisition of pluripotency (Hajkova et al., 2008). It is
important to note that in PGC development DNA demethylation is a temporally ordered
process; first promoters of pluripotency markers and germ cell specific genes undergo
demethylation, which is later followed by loss of methylation at the ICRs (Guibert et al.,
2012; Seisenberger et al., 2012). These events strikingly resemble our observations that
mouse EG cell mediated pluripotent conversion occurs ahead of imprint reprogramming
in somatic cells after fusion. In addition, it has been reported that PGCs express significant
levels of Tet1 and Tet2, which may contribute to the replication-coupled removal of 5-mC
upon conversion into 5-hmC. 5-hmC is no longer be recognised by the UHRF1 (Hackett
et al., 2013; Hashimoto et al., 2012), therefore 5-hmC is replaced by unmodified cytosine
during DNA synthesis. In-vivo analysis Tet1 knock-out is also shown to result in
abnormalities in genomic imprinting, due to the presence of hypermethylated ICRs
(Yamaguchi et al., 2013b).
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Chapter 5. Analysis of TET Protein Requirement in
Mouse Embryonic Stem Cell Induced Reprogramming of
Human B Lymphocytes
5.1. Introduction
In this Chapter, I investigate the importance of TET proteins for reprogramming
efficiencies. I have used two strategies to disturb Tet activity. The first approach (Chapter
5.2), is based on RNA interference. Upon knockdown Tet mRNA levels are decreased in
mES cells, and these cells are then fused to hB lymphocytes. The reprogramming
efficiencies of Tet -sh and -control mES cells are compared. The advantage of this system
is that it can circumvent any potential long-term effects of knockdown of Tets on mES cell
identity. For complete withdrawal of TET activity in mES cells, my second approach
(Chapter 5.3) is based on CRISPR/Cas9 genome editing system. Here I report a detailed
work-flow of activity to generate mutant ES cells and analyse the capacity of these knock-
out lines to reprogram hB lymphocytes to pluripotency.
5.2. Tet Knockdown in mouse ES cells and cell fusion
In order to knockdown Tet1 and Tet2 genes separately or together in mES cells, I
took advantage of RNA interference system using shRNA vectors (Tet1 alone, Tet2 alone
and Tet1/Tet2 double knockdown), as previously demonstrated (Chapter 4) (Ito et al.,
2010; Williams et al., 2011). To determine how the Tet knockdown affected the ability of
mES cells to convert hB cells to pluripotency, I compared gene expressions in
heterokaryons.
5.2.1. Tet1 knockdown and fusion
In order to reduce Tet1 expression levels, I electroporated mES cells with
pSUPER.neo.GFP + Tet1 shRNA vector (shTet1), and used FACS to sort successfully
transfected cells after 24 hours based on GFP expression (Figure 5.1.A). I used the same
FACS settings to sort mES cells transfected with empty vector (shCtrl) for comparison. 72
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hours after transfection (at the time of fusion), qRT-PCR showed (grey compared to green)
that Tet1 mRNA levels had decreased to 35% of the original value (Figure 5.1.B).
Interestingly, this was accompanied by a slight increase of Tet2 expression (Figure 5.1.B).
Although Oct4 expression was not altered in these cells, Nanog mRNA levels were
augmented (Figure 5.1.B), a result which has been observed in a previous research study
(Williams et al., 2011). Previously Ito et al. demonstrated that upon Tet1 knockdown,
Nanog expression level was substantially decreased (Ito et al., 2010), but Williams et al.
reported that this was due to off-target effects of the shRNA used in their study (Williams
et al., 2011).
ES cells transfected with shCtrl or shTet1 (72 hours) were fused with hB
lymphocytes, and the changes in gene expression in the human B nuclei were analysed
(Figures 5.1.C-D). As expected, silent human pluripotency genes OCT4, NANOG and
CRIPTO were induced upon fusion with shCtrl transfected mES cells (Figure 5.1.C, grey
bars). Likewise, shTet1 transfected mES cells were able to reprogram hB cells upon
fusion, more efficiently (i.e. better than controls, Figure 5.1.C, green bars compared to
grey wild type). In line with expectations, hB specific gene CD19 expression decreased
upon fusion with either shCtrl or shTet1 transfected mES cells (Figure 5.1.D). The
decrease was slightly more prominent with Tet1 knockdown ES cells (also CD45 gene
downregulation) suggesting that Tet1 withdrawal may enhance reprogramming.
Collectively, these results demonstrate that Tet1 downregulation does not worsen
the reprogramming efficiency in heterokaryons, suggesting that Tet1 does not play a
crucial role in the early stages of cell fusion mediated pluripotent conversion. If anything,
reprogramming was slightly enhanced in the absence of TET1, which might be due to
unknown secondary effects of the knockdown, or because Tet2 and Nanog expressions
were elevated in these targeted mES cell lines. This contrasts with what has been shown
in a previous report (Costa et al., 2013).
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Figure 5.1. Effect of Tet1 knockdown on reprogramming. (A) FACS profile of pSUPER.neo.GFP + Tet1 (shTet1) transfected mES cells. The gate was chosen to sort medium/high GFP expressing cells (plots in blue). (B) qRT-PCR analysis of Tet1, Tet2, Oct4 and Nanog in mES cells, 72 hours after transfection with either empty (shCtrl, grey bars) or shTet1 (green bars) plasmids. Data were normalised to mouse Ubc and represent mean and SEM of 4 different experiments. (C) qRT-PCR analysis of pluripotency factor gene expression (OCT4, NANOG and CRIPTO) in hB lymphocytes before (0h) and at 48 and 72 hours after fusion with mES cells transfected with either shCtrl (grey bars) or shTet1 plasmids (green bars). Data were normalised to human GAPDH and represent mean and SEM of 2 different experiments. (D) qRT-PCR analysis of lymphocyte specific factor gene expression (CD19 and CD45) in hB lymphocytes before (0h) and at 48 and 72 hours after fusion with mES cells transfected with either shCtrl (grey bars) or shTet1 plasmids (green bars). Data were normalised to human GAPDH and represent mean and SEM of 2 biological replicates.
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5.2.2. Tet2 knockdown and fusion
To knockdown the Tet2 gene in mES cells I used a similar approach, where I
transfected the cells with pSUPER.neo.GFP + Tet2 shRNA vector (shTet2) and sorted
them after 24 hours based on GFP expression (Figure 5.2.A). I used the same FACS gate
settings to sort empty vector (shCtrl) transfected mES cells as a control. qRT-PCR
analysis of the transfected cells demonstrated a decrease in Tet2 gene expression level
compared to shCtrl transfection, without a major change in Tet1, Oct4 and Nanog levels
(Figure 5.2.B).
To determine whether the decrease in Tet2 mRNA level compromises the
reprogramming ability of mES cells, I conducted cellular fusions and analysed the gene
expression in resulting heterokaryons. Using shCtrl transfected mES cells as controls,
(Figure 5.2.C, grey bars), shTet2 transfected mES cells induced a similar level of
pluripotency gene expression in hB cells (Figure 5.2.C, blue bars). In accord with this,
silencing of CD19 gene expression was comparable in both samples, and there was no
major difference in the extinction of human CD45 expression. These data suggest that the
reduced Tet2 mRNA does not impair the reprogramming potential of mES cells, but is
implicated in mEG mediated reprogramming (Chapter 4) and may also have roles in iPS
conversion (Doege et al., 2012).
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Figure 5.2. Effect of Tet2 knockdown on reprogramming. (A) FACS profile of pSUPER.neo.GFP + Tet2 (shTet2) transfected mES cells. The gate was chosen to sort medium/high GFP expressing cells (plots in blue). (B) qRT-PCR analysis of Tet1, Tet2, Oct4 and Nanog in mES cells, 72 hours after transfection with either empty (shCtrl, grey bars) or shTet2 (blue bars) plasmids. Data were normalised to mouse Ubc and represent mean and SEM of 4 different experiments. (C) qRT-PCR analysis of pluripotency factor gene expression (OCT4, NANOG and CRIPTO) in hB lymphocytes before (0h) and at 48 and 72 hours after fusion with mES cells transfected with either shCtrl (grey bars) or shTet2 plasmids (blue bars). Data were normalised to human GAPDH and represent mean and SEM of 2 different experiments. (D) qRT-PCR analysis of lymphocyte specific factor gene expression (CD19 and CD45) in hB lymphocytes before (0h) and at 48 and 72 hours after fusion with mES cells transfected with either shCtrl (grey bars) or shTet2 plasmids (blue bars). Data were normalised to human GAPDH and represent mean and SEM of 2 biological replicates.
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5.2.3. Tet1 and Tet2 double knockdown and fusion
In order to rule out the possible redundant functions of Tet1 and Tet2 in
reprogramming, it was important to simultaneously decrease their expression in mES
cells. Therefore, I modified the Tet1 RNAi plasmid, by replacing the neo.GFP with mCherry
coding sequence and I co-transfected mES cells with pSuper.mCherry + Tet1 (shTet1)
and pSUPER.neo.GFP + Tet2 (shTet2) vectors. After 24 hours of transfection, I sorted the
cells expressing both fluorescent markers. (Figure 5.3.A). I applied the same settings for
mES cells co-transfected with empty vectors expressing GFP and mCherry (shCtrls). After
72 hours of shTet1&shTet2 co-transfection, qRT-PCR analysis revealed that the
expression levels of Tet1 and Tet2 were both decreased (by 75% and 55% respectively),
Oct4 level remained constant, and Nanog level was slightly increased, all compared to
empty vector co-transfected mES cells (Figure 5.3.B, purple bars –shTet1&shTet2-
compared to grey bars –shCtrls-).
To answer whether knocking down both of Tet1 and Tet2 genes had an effect on
reprogramming, I fused these mES cells with hB lymphocytes, and evaluated the
efficiency of pluripotent conversion by qRT-PCR in heterokaryons. Interestingly, shRNA
mediated decrease of Tet1 and Tet2 expression did not have a negative effect on
reprogramming, as judged by OCT4, NANOG and CRIPTO expression (Figure 5.3.C), or
extinction of human B cell genes (CD19 and CD45). Thus, inhibition of Tet1 and Tet2 does
not interfere with the capacity of mES cells to reprogram hB cells upon fusion. These data
raise the question whether 5-methylcytosine oxidation is necessary for experimental
reprogramming by cell fusion. However, it is noteworthy that RNAi-based systems do not
necessarily reduce protein or enzymatic activities of targeted genes rapidly. Although it is
conceivable that even a slight decrease in expression of a crucial factor would result in a
detectable deficiency of pluripotent induction, complete removal of TET proteins may not
be achieved with this approach. For this reason, in the following section I describe how I
generated Tet Knockout mES cell lines using CRISPR/Cas9 mediated genome editing
system, and compare reprogramming potentials of these lines.
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Figure 5.3. Effect of Tet1/Tet2 double knockdown on reprogramming. (A) FACS profile of pSUPER.mCherry+ Tet1 (shTet1) and pSUPER.neo.GFP + Tet2 (shTet2) co-transfected mES cells. The gate was chosen to sort medium/high GFP & mCherry expressing cells (plots in blue). (B) qRT-PCR analysis of Tet1, Tet2, Oct4 and Nanog in mES cells, 72 hours after co-transfection with either empty (shCtrls, grey bars) or shTet1&shTet2 (purple bars) plasmids. Data were normalised to mouse Ubc and represent mean and SEM of 3 different experiments. (C) qRT-PCR analysis of pluripotency factor gene expression (OCT4, NANOG and CRIPTO) in hB lymphocytes before (0h) and at 48 and 72 hours after fusion with mES cells co-transfected with either shCtrls (grey bars) or shTet1&shTet2 plasmids (blue bars). Data were normalised to human GAPDH and represent mean and SEM of 2 different experiments. (D) qRT-PCR analysis of lymphocyte specific factor gene expression (CD19 and CD45) in hB lymphocytes before (0h) and at 48 and 72 hours after fusion with mES cells co-transfected with either shCtrls (grey bars) or shTet1&shTet2 plasmids (purple bars). Data were normalised to human GAPDH and represent mean and SEM of 2 different experiments.
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5.3. CRISPR/Cas9 mediated Tet gene editing and cellular fusion
First discovered as a constituent of bacterial adaptive immunity, Type II
CRISPR/Cas system has recently been used to modify genomes of higher species (Sorek
et al., 2013; Terns and Terns, 2014). Simplicity and cost-effectiveness are among the
advantages of CRISPR-based technologies, providing the opportunity to target multiple
genes simultaneously both in-vivo and in-vitro. This feature has substantially shortened
the time it would take with conventional methods (Wang et al., 2013a). The main role of
CRISPR/Cas system is to generate double-strand break at a particular location in the
genome. The resulting DNA damage is then recognized by the cell’s own repair
mechanism which in return acts on the break. There are two main repair pathways, and
the system can be repurposed according to the aim. First pathway is the non-homologous
end joining, an error-prone mechanism, leading to nucleotide insertions or deletions at the
break site. This can be particularly useful to rapidly inactivate genes by causing mutations
that lead to frameshifts if coding sequences are targeted, or can be used to disrupt
particular protein binding sites. On the other hand, the second system is a high-fidelity
repair mechanism, mediated by homologous recombination which can be exploited to
insert, remove, alter or replace specific sequences with a provided template.
In this section I describe generation of mES cell lines by CRISPR/Cas9, where I
concomitantly target Tet1 and Tet2 genes for non-homologous end joining and report data
on their ability to reprogram somatic cells in experimental fusion system.
5.3.1. CRISPR/Cas9 system construction against Tet1 and Tet2 genes and delivery into mES cells
The Type II CRISPR system is a ribonucleoprotein complex and has two
complementary parts. The first part is composed of CRISPR-RNA (crRNA), a 20
nucleotide guide sequence complementary to the DNA, and Trans-activating crRNA
(tracrRNA) which is complementary to the crRNA and is involved in RNA processing. The
second part is the Cas protein, which is an RNA-guided endonuclease (Deltcheva et al.,
2011). Recently Jinek et al. reported the merging of the two RNA components into a single
RNA chimera (single guide RNA –sgRNA-), which targets the Cas endonuclease with
similar efficiency to the DNA (Jinek et al., 2012).
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The px330 plasmid contains all the components of Type II CRISPR system (Cong
et al., 2013); a human codon-optimized Streeptococcus pyogenes Cas9 under strong
hybrid chicken beta-actin promoter, and chimeric guide RNA under the control of U6
promoter. I separately cloned Tet1 and Tet2 sgRNAs into px330 plasmids (see Materials
& Methods) (Figure 5.4) that have previously been shown to target Tet genes efficiently
(Wang et al., 2013a; Yang et al., 2013).
Figure 5.4. Tet1 and Tet2 targeting by CRISPR/Cas. Schematic representation of mouse Tet1 and Tet2 genes, and the DNA target sites. Nucleotides in green represent the PAM sequence, necessary for the target recognition by Cas9 endonuclease of Streptococcus pyogenes (-NGG). Nucleotides in blue depict sgRNA targeted DNA sites. Red arrows demonstrate the locations (3 base pairs downstream of PAM) of double strand breaks upon Cas9 endonuclease activity, which are positioned within SacI and EcoRV restriction enzyme recognition sites in Tet1 and Tet2 respectively.
I co-transfected mES cells by electroporation with (1) px330+sgTet1 vector, (2)
px330+sgTet2 vector and finally (3) pH2BmCherry-Ires-Puro vector which enables the
selection of successfully transfected cells by either of two different methods; FACS and
Puromycin drug treatment (see strategy outlined in Figure 5.5.A).
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Figure 5.5. Workflow for CRISPR/Cas mediated gene targeting in mouse ES cells. (A) mES cells are co-transfected by electroporation. (H2BmCherry-IRES-Puro expression cassette is under the control of highly active chicken beta-actin promoter. Transfected cells are splitted and either sorted by FACS according to mCherry expression or treated with Puromycin for 2 days after 48 hours of transfection. A total of 32 colonies were picked for further analysis. (B) FACS profile for transfected mES cells. The gate has been chosen to sort cells expressing high levels of mCherry.
Two days after Puromycin treatment, I used one plate (low cell density) to pick up
colonies, and the other plate (high cell density) for Surveyor Assay to determine the
mutation efficiency. I used the third plate to apply FACS on the mES cells that express
high level of mCherry (Figure 5.5.B), and sorted one cell per well of a 96-well plate, where
after one week, 20-25% of the wells contained viable clones. I sorted the rest of the cells
and plated back for 4 days and collected them to conduct Surveyor Assay. In total I
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obtained 32 clones (24 from Puromycin treated, 8 from mCherry sorted) to be further
analysed for the mutation identities.
5.3.2. Surveyor Assay for analysis of CRISPR/Cas efficiency
Surveyor Assay helps to analyse the presence of mutations at a specific locus on
the DNA and is based on the Surveyor DNA endonuclease (Transgenomic), which is a
member of CEL nuclease family isolated from celery (Qiu et al., 2004). Surveyor nuclease
has the capacity to recognise and cleave the 3’ end side of any mismatch site in a given
DNA duplex. Its sensitivity allows the examination of small mismatches such as single
nuclear polymorphisms (SNPs). As non-homologous end joining upon CRISPR/Cas
endonuclease activity creates random indels at the target site, various mutations are
expected to be encountered in a transfected population of cells. Following the
amplification of the target site by PCR, denaturation and reannealing create a vast amount
of mismatches, which then can be recognised and cleaved by the Surveyor nuclease, as
illustrated in Figure 5.6.
Figure 5.6. Schematic Representation of Surveyor Assay. As a result of CRISPR/Cas endonuclease activity on a specific target, random mutations are created by non-homologous end repair (Left). Upon PCR, denaturation and annealing of the DNA from a population of targeted cells (Centre), DNA duplexes contain mismatches, which will then be digested by CEL-1 (Right). The cleaved and non-cleaved bands can be distinguished by gel electrophoresis, where the upper band will be the WT bands (un-cleaved) and the lower bands will be the mismatch bands (cleaved). To conduct the Surveyor Assay, I PCR amplified Tet1 and Tet2 targeted loci of
from the DNA of ‘Puromycin Treated’ or ‘mCherry Sorted’ samples (Day 6, Figure 5.5). I
denatured the PCR amplicons at high temperature, and let the emerging single stranded
DNA randomly reanneal by gradually decreasing the temperature (See Materials &
Methods). This results in DNA duplexes with mismatches, and to visualise this I migrated
the samples on agarose gel after treating with Surveyor endonuclease. In non-transfected
wild type cells, as Tet1 and Tet2 amplicons do not contain mismatches (unless there are
naturally occurring SNPs on these particular loci), no cleavage took place (Figure 5.7;
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WTs for Tet1 and Tet2, 460 bp and 466 bp respectively). On the other hand, in
CRISPR/Cas transfected cell population, efficient cleavage by Surveyor Endonuclease
was observed, unveiling the presence of mismatches that were resulted in non-
homologous end joining at the target sites (Figure 5.7; ~235 bp and ~225 bp for Tet1,
~252 bp and ~214 bp for Tet2). These results indicated successful targeting of Tet1 and
Tet2 genes as previously demonstrated (Wang et al., 2013a).
Figure 5.7. Surveyor Assay on Tet1 and Tet2 in wild type and CRISPR/Cas targeted mES cells, treated with Puroymcin or mCherry sorted upon co-transfection. Tet1 PCR produces 460 bp amplicon, Tet2 PCR produces 466 bp amplicon. PCR primers were designed to centre the sgRNA target sites in both Tet1 and Tet2 (Wang et al., 2013a). In all conditions, the upper bands are the undigested amplifications (either wild type or no mismatch mutant amplicons –see Figure 5.6-). The lower bands demonstrate the cleaved products of the amplicons containing mismatches upon Surveyor Assay.
It is important to note that cleavage efficiencies in ‘Puromycin Treated’ and
‘mCherry Sorted’ populations were very similar for both targets, so clones isolated from
either of the conditions could be used for further analysis.
5.3.3. Restriction Fragment Length Polymorphism screen on CRISPR/Cas9 targeted mES cells for Tet1 and Tet2
In order to analyse whether single clones that I selected from CRISPR/Cas
targeting (a total of 32; 24 from puromycin selection, 8 from mCherry sorting) had acquired
indels, I conducted a Restriction Fragment Length Polymorphism (RFLP) screen. This
allows an investigation to quickly assess mutated alleles, as the targeted Tet1 and Tet2
loci contain restriction enzyme recognition sites that can be used to identify mutations in
the analysis (Figure 5.4). Disruption of these sites by indels demonstrates successful
error-prone non-homologous end joining upon cleavage by Cas9 (Figure 5.8).
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Figure 5.8. Schematic Representation of RFLP. A wild type allele is expected to be digested by restriction enzyme. On the contrary, the disruption of the recognition site by indels, prevents the cleavage of the DNA amplicon. Agarose gel electrophoresis reveals the identity of allelic targeting by CRISPR/Cas9.
The screen of the single clones is based on PCR amplification of the targeted Tet1
and Tet2 loci (same primer couples as in the Surveyor Assay, the uncleaved products are
460 bp and 466 bp for Tet1 and Tet2 respectively) and restriction digestion. The Tet1
sgRNA targets SacI recognition site in exon 4, and the Tet2 sgRNA targets EcoRV
recognition site in exon 3. Double strand breaks are expected to take place in these
recognition sites, which are then corrected by non-homologous end joining that results in
indel acquisition. There are three outcomes of the clonal screen by RFLP, (1) total
cleavage of the PCR product proving that both alleles are wild type, (2) partial cleavage
of the PCR product showing that one allele is wild type and one allele is mutated, (3) no
cleavage of the PCR product, demonstrating that the both alleles are likely mutated.
As shown in Figure 5.9, wild type ES cell samples show PCR amplicons for both
Tet1 and Tet2 targeted loci that have been efficiently cleaved by the corresponding
restriction enzymes (Figure 5.9; left WT column). Almost all of the co-targeted clones
contained bi-allelic mutations at the Tet1 and Tet2 target sites as evidenced by the lack
of restriction digestion of the amplicons (Figure 5.9). This observation is in line with the
efficiency determined by the Surveyor Assay at the population level.
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Figure 5.9. RFLP Assay on WT and Tet1&Tet2 CRISPR co-targeted 32 clones. Tet1 PCR produces 460 bp amplicon, Tet2 PCR produces 466 bp amplicon. For RFLP on the Tet1 target locus, the PCR amplicons were digested with Sac1 restriction enzyme, which in WT condition results in bands at sizes of ~235 bp and ~225 bp. For RFLP on the Tet2 target locus, the PCR amplicons were digested with EcoRV restriction enzyme, which in WT condition results in bands at sizes of ~252 bp and ~214 bp. PCR amplicon sizes may vary due to the nature of acquired indels.
5.3.4. Sequencing of Tet1&Tet2 CRISPR targeted ES clones
Acquired indels upon error prone non-homologous end-joining may have various
outcomes. As the exons are targeted, base insertion or base deletion may cause a frame-
shift in the coding sequence. Alternatively, these modifications may lead to in-frame
mutations, in which case the protein activity may or may not change, depending on the
length of alteration in the DNA sequence. To characterise the indel identities of the
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CRISPR targeting, I sequenced the Tet1 and Tet2 target loci in 6 clones (5A, 5B, 5C, 8A,
8B, 8E). To do so, I cloned the PCR amplicons into pGEM®-T Easy Vector (Promega),
which I transformed into competent bacteria. Per transformation, I used at least 8 bacterial
colonies for sequencing to cover both alleles (Figure 5.10.A-B).
Figure 5.10. DNA sequencing results on Tet1&Tet2 CRISPR co-targeted loci. (A) Table demonstrating the size of indels in each allele in corresponding clones. Purple colour indicates frame-shift acquisition in one allele, green colour indicates frame-shift acquisition in both alleles, and red colour indicates in-frame mutations in both alleles. (B) DNA sequences of the WT (blue) and targeted clones. Green nucleotides in WT indicate the restriction sites SacI and EcoRV in Tet1 and Tet2 respectively. Deletions are indicated as hyphens, insertions are indicated in red, and the indel sizes are indicated on the right for each sequence.
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In line with the RFLP screen, sequences of Tet1 and Tet2 CRISPR target loci in all
of the 6 clones underwent error prone repair upon double strand break by Cas9 which
distorted the restriction enzyme recognition sites (Figure 5.10.B, nucleotides in green in
WTs). DNA sequencing revealed various genotypes among 6 clones which may lead to
different phenotypes in the ES cells. For example clone 8B contains indels that result in
bi-allelic frame-shifts both in Tet1 and Tet2 genes, causing early termination of translation
by the stop codons in the new frames. On the other hand, in clones 5A and 5B this is the
case only in one of the alleles of Tet1 and Tet2 genes, while second alleles for these
genes acquired an in-frame mutation.
The majority of the observed indels are deletions, and it is interesting to note that
some mutations reoccur in different clones, which may indicate the presence of micro-
homology directed repair in these loci (McVey and Lee, 2008). Overall, the presence of
mutations in all of the sequenced and screened clones suggests a high efficiency of
CRISPR/Cas co-targeting using the protocol described here.
5.3.5. Reprogramming capacity of CRISPR/Cas9 mediated Tet1 and Tet2 mutant ES cell clones upon cell fusion
As a result of CRISPR/Cas9 mediated gene editing, I obtained ESC clones with
various genotypes for Tet1 and Tet2 as previously described. Clone 5A contains one
frame-shift and one in-frame allele for both Tet1 and Tet2 genes, while in the clone 8B
both alleles for both genes acquired frame-shift mutations which are expected to abrogate
TET1 and TET2 activity. To understand how these changes affect reprogramming
potential of ES cells, I conducted cell fusions in which I fused human B cells with clones
5A or 8B or wild type cells. I analysed human gene expression in heterokaryons formed
after 48 and 72 hours of fusion. As expected, pluripotency genes OCT4, NANOG and
CRIPTO that were not expressed in human B cells at the time of fusion but were induced
in heterokaryons upon fusion with WT mES cells (Figure 5.11.A). Interestingly at 48 hours,
both 5A and 8B clones were able to induce the expression of these factors either as
efficiently as or more efficiently than compared to WT cells. However even though WT
heterokaryons were further reprogrammed at 72 hours of fusion, pluripotency gene
expression levels of clone 5A and clone 8B heterokaryons remained unchanged (Figure
5.11.A). In addition, human B specific CD19 and CD45 genes were downregulated to
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comparable levels upon fusion with WT, clone 5A and clone 8B ES cells at 48 and 72
hours (Figure 5.11.B).
Figure 5.11. Effect of Tet1/Tet2 knockout on reprogramming. (A) qRT-PCR analysis of pluripotency factor gene expression (OCT4, NANOG and CRIPTO) in hB lymphocytes before (0h) and at 48 and 72 hours after fusion with WT mES cells, Clone 5A and Clone 8B. Data were normalised to human GAPDH and represent mean and SEM of 2 different experiments. (D) qRT-PCR analysis of lymphocyte specific factor gene expression (CD19 and CD45) in hB lymphocytes before (0h) and at 48 and 72 hours after fusion with WT mES cells, Clone 5A and Clone 8B. Data were normalised to human GAPDH and represent mean and SEM of 2 different experiments.
These results demonstrate that partial or complete ablation of Tet activity do not
hinder ES cells’ capacity of reprogramming B cells upon fusion in heterokaryons.
However, the fact that pluripotent gene expression does not appear to be increasing
between 48 and 72 hours of fusion raises questions. This might be due to defects in
stabilisation of pluripotent induction, or due to the stress the clones have been through in
the course of CRISPR/Cas editing.
5.4. Summary and Discussion
Developing mammalian embryos undergo two waves of genome-wide DNA
demethylation, occurring soon after fertilization and during PGC development. Since the
discovery of TET proteins as catalytic modulators of 5-mC to 5-hmC conversion (Tahiliani
et al., 2009; Kriaucionis and Heintz, 2009), many studies have focused on their roles in
embryonic development, as 5-hmC has been suggested to be an intermediate of
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replication-dependent and –independent loss of methylation (Pastor et al., 2013). These
waves of DNA demethylation lead to in-vivo acquisition of pluripotency, and it is crucial to
have a better understanding of how lineage identity is reset.
In this chapter, I have not only shown that TET is dispensable in experimental cell
fusion based reprogramming assay, but also described the CRISPR/Cas system as an
effective gene knock-out tool. Firstly, RNA interference-mediated Tet1 and Tet2
knockdown (separately or together), did not interfere with the capacity of mouse ES cells
to reprogram hB lymphocytes. Since RNA interference does not completely abolish mRNA
level of the target gene, I also created Tet1 and Tet2 knock-out mES cell lines using the
CRISPR/Cas system. Interestingly, these ES cells were as successful as wild type cells
in initiating B cell reprogramming, but the induction of pluripotent gene expression did not
develop fully. Reprogramming was initiated in these cells, apparently even in the presence
of cytosine methylation at OCT4, NANOG and CRIPTO genes. This observation deserves
more attention, and I will return to it later in this document.
Recent in-vitro reprogramming strategies have strived to elucidate the contribution
of Tet proteins to pluripotent conversion. These have however, resulted in different and
often opposing findings. The involvement of Tet activity in iPS cells was first reported by
Doege et al., where they reported an increased global 5-hmC distribution upon factor
induced (Oct4, Sox2, Klf4 and c-Myc –OSKM-) reprogramming of mouse embryonic
fibroblasts (Doege et al., 2012). Further analysis revealed 5-hmC enrichment in
endogenous pluripotency genes (such as Nanog an Esrrb), accompanied by gene
expression. This is in line with the early activation of endogenous Tet2 expression (by day
3 of induction), while interestingly Tet1 remained silent. In addition, Tet2 downregulation
upon shRNA knockdown completely abrogated iPS cell colony formation, suggesting that
TET2 protein is responsible for the global and locus specific DNA hydroxylation, necessary
for successful reprogramming (Doege et al., 2012). In a similar approach, Costa et al.
postulated that Tet1 knockdown inhibited factor-induced iPS cell generation from mouse
embryonic fibroblasts (Costa et al., 2013). Moreover, upon fusion with hB cells, Tet1
Knock-out ES cells were less efficient in their ability to reprogram compared to WT ES
cells. Finally, the authors demonstrated that overexpression of TET1 or TET2 proteins
enhanced iPS cell formation, mainly via downstream effects of physical interaction with
NANOG (Costa et al., 2013). A third study revealed that Tet1 expression was gradually
increased during iPS cell reprogramming (while Tet2 levels remained relatively stable)
and Tet1 downregulation by RNA interference abolished colony formation (Gao et al.,
2013). These investigators showed that in the course of reprogramming, TET1
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hydroxylated the methylated Oct4, leading to its demethylation and subsequent activation,
and that TET1 could substitute OCT4, in the OSKM factor cocktail (Gao et al., 2013). A
fourth approach demonstrated that, during human iPS cell reprogramming 5-hmC levels
were significantly increased, mediated mainly by the induced expression of endogenous
TET1 gene; its knockdown resulted in a decreased number of reprogrammed colonies
(Wang et al., 2013b). Another study analysed TET function modulation by Vitamin C
during pluripotent conversion of MEFs, and concluded that Tet1 deficiency resulted in
enhanced reprogramming in the presence of, and reduced reprogramming in the absence
of Vitamin C (Chen et al., 2013a). This was interesting as Vitamin C has previously been
shown to enhance Tet activity (Blaschke et al., 2013; Yin et al., 2013). The authors
concluded that cooperative action of Tet1 and Vitamin C stands as a barrier of
mesenchymal-to-epithelial transition in the course of reprogramming (Chen et al., 2013a).
A recent study has provided data that conflicts with all previous reports. It
demonstrated that Tet1&Tet2 and Tet1&Tet3 double mutant MEFs were successfully
reprogrammed into pluripotent colonies. However, simultaneous depletion of all Tet
constituents completely abolished this conversion, which the authors ascribed to a lack of
hydroxylation of methylated miR-200 family gene promoters, followed by demethylation
(Hu et al., 2014). In wild type cells, these activities are exerted by redundant Tet function
in cooperation with thymine deglycosylation by TDG (Hu et al., 2014). By narrowing down
the Tet requirement in factor-induced reprogramming into a miRNA family, this study has
raised doubt as to the necessity of Tet activity in reactivating pluripotency genes in the
course of reprogramming. In view of this, I have examined whether DNA demethylation of
OCT4 is a prerequisite for pluripotent reprogramming.
DNA demethylation was noted in iPS cell creation (Wernig et al., 2007) and was
also shown to be a perquisite for Oct4 reactivation in nuclear transfer-mediated
reprogramming studies (Simonsson and Gurdon, 2004). It is noteworthy that other
epigenetic markers undergo global changes in pluripotent conversion. For example,
genome-wide remodelling is evident in very early stages of factor induced conversion in
somatic cells, where active histone marks such as H3K4 methylation are rapidly
accumulated in pluripotency and early development genes (Koche et al., 2011). In
addition, our lab has previously shown that upon fusion, various chromatin features
change markedly, such as histone acetylation levels (Piccolo et al., 2011) and
heterochromatin marker HP1α is rapidly redistributed in the somatic hB nucleus (Pereira
et al., 2010). I re-evaluated these findings by using mouse ES cells expressing H2B
tagged with mCherry fused with human fibroblasts expressing HP1α tagged with GFP
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(Figure 5.12). Hp1α is a nuclear protein and its distribution in the somatic fibroblast is
punctuate (Figure 5.12, top left), marking the heterochromatic loci, and the H2B
distribution is nuclear in mouse ES cells as expected from a histone protein (middle). In
heterokaryons, the mouse-derived nucleus (here mouse ES) can be distinguished from
human derived nucleus (here human fibroblast) on the basis of DAPI staining profile
(Pereira et al., 2008) (Figure 5.12, bottom right). Imaging reveals that upon fusion with
mES cell, HP1α punctuate profile is lost by fibroblast nuclei (white arrow), but mouse ES
cell nuclei (red arrows) in the same heterokaryon show a punctate H2B distribution (Figure
5.12, bottom left white and red arrows, respectively). Similarly, H2BmCherry originating
from mouse nucleus accumulates in the human fibroblast partner.
Figure 5.12. HP1α redistribution in mESxhF heterokaryons. Top row, human IMR90 fibroblast expressing Hp1αGFP fusion protein. Middle row, mES cells expressing H2BmCherry, Bottom row, a heterokaryon with two mES cell nuclei (demonstrated by red arrows) and one human fibroblast nucleus (demonstrated by white arrow), distinguished by the DAPI staining profile. Right column, merge image of GFP and mCherry as well as DAPI (blue) which stains nuclear DNA and Phalloidin (yellow) which stains filamentous actin. Scale bar = 10µm.
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This result recapitulates previous findings that HP1α is re-distributed in somatic nuclei
upon fusion, and demonstrates the global alteration in the epigenetic landscape at the
early stages of reprogramming. Using tagged proteins of this sort allows live cell imaging
of chromatin remodelling events in heterokaryons. Considering the global change in
chromatin signatures, is it possible that pluripotency genes can be re-expressed even in
the presence of DNA methylation? Interestingly a precedent for this has previously been
reported during C/EBPα-induced B cell to macrophage transdifferentiation; where silent
macrophage-specific genes were activated, even though their promoters remained highly
methylated (Rodríguez-Ubreva et al., 2012). In addition, these genes were shown to
acquire H3K4 trimethylation (H3K4me3) as well as H3K9/H3K14 acetylation (features of
active transcription) all of which are active histone marks. With this in mind, the authors
have suggested that repressive effect of DNA methylation may be overcome by additional
epigenetic factors such as active histone modifications (Rodríguez-Ubreva et al., 2012).
I have previously shown that by day 3, heterokaryon populations still show high
levels of DNA methylation at the human OCT4 promoter although the OCT4 gene
expression is detected as early as 24 hours after fusion. To explore whether a locus-
specific histone remodelling is occurring in these cells, I used chromatin precipitation
(ChIP) analysis. This is technically difficult as the number of heterokaryons generated after
fusion is low and can be masked by the presence of unfused cells in these cultures. To
avoid this, I used H2BmCherry expressing mouse ES cells and dye labelled human B cells
(CellTrace Violet, Invitrogen) as fusion partners, and I enriched double-labelled
heterokaryons by FACS. I also adapted a low cell ChIP protocol (see Materials and
Methods) to look for H3K4me3 enrichment at the human OCT4 promoter locus, in both
human B cells and heterokaryons using human specific primers. Preliminary ChIP data
demonstrated that H3K4me3 was not present at the OCT4 promoter in human B cells, in
line with the gene’s repressed status, but was highly enriched at the same region in
heterokaryons (Figure 5.13). Although these are preliminary data, it is important that
acquisition of H3K4me3 is detected at the OCT4 promoter of the somatic nucleus upon
reprogramming. It will be essential to expand this analysis in future to see whether other
active histone marks have also been incorporated, and to assess whether other
pluripotency gene loci are also remodelled early after cell fusion. Eventually we hope to
conduct a genome-wide ChIP sequencing analysis of these changes (for discussion see
Chapter 7).
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Figure 5.13. H3K4me3 ChIP on human B lymphocytes and Day 3 heterokaryons. H3K4me3 is analysed in human B cells and heterokaryons at β-ACTIN as positive control, at OCT4 and at TSH2B (testis-specific histone 2B) as negative control. H3Kme3 data are normalized to total H3, and as negative control, IgG data are normalized to input.
Overall my results, and examples from the literature, support a rather unexpected
model of gene re-activation upon reprogramming. DNA methylation has been regarded as
a relatively stable modification (due to the presence of strong Carbon-Carbon bond
between the cytosine and the methyl group). In a compact chromatin structure where DNA
methylation and repressive histone marks coexist, histone tails could be viewed as being
more physically and chemically accessible, which may render them prone to modification
by external cues. Accumulation of active marks on histone tails may lead to recruitment of
transcription factors that could kick-start low levels of gene transcription. This may be
followed by DNA demethylation, required to stabilise gene expression. As reprogramming
in heterokaryons is considered fast and efficient, as compared to other systems, it might
be difficult to accurately order the sequence of these events. With developing technology
and sensitive detection systems, single cells have increasingly been used to avoid the
loss of information due to averaging. In the future, such analysis could be done on single
heterokaryons that would provide valuable information on the nature of reprogramming.
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Chapter 6. CRISPR/Cas Editing of Jarid2 and Non-
Canonical WNT Pathway Components
6.1. Introduction
Jarid2 (Jumonji, AT rich interactive domain 2) is a component of the Polycomb
Repressor Complex 2 (PRC2) in ES cells and has been implicated in regulating
pluripotency networks in embryonic stem cell differentiation (Landeira and Fisher, 2011).
In addition, recent work by David Landeira and colleagues in our laboratory has
demonstrated that Jarid2 deficient mES cells (Shen et al., 2009) possess elevated levels
of Nanog expression (Landeira et. al, submitted) and are extremely good at
reprogramming through cell fusion (Pereira et al., 2010). Analysis of publically available
mES cell ChIP datasets revealed a lack of JARID2 binding at the Nanog promoter,
suggesting that Nanog was indirectly regulated by JARID2. To identify factors that were
differentially expressed in JARID2 deficient mES cells, gene expression profiling was
performed, and several Wnt signalling components were determined to be significantly
de-regulated. Among these were Prickle1 and Fzd2 (downregulated in the absence of
Jarid2) involved in non-canonical Wnt pathway and Wnt9a, all of which were confirmed to
be directly bound by Jarid2 (Landeira et. al, submitted). To understand whether loss of
these non-canonical pathway components could phenocopy the Jarid2 deficiency in
mouse ES cells, I took advantage of loss-of-function approach by using CRISPR/Cas9
gene editing system. In this Chapter, I describe CRISPR design process for Jarid2,
Prickle1, Fzd2 and Wnt9a, and caution their subsequent modification in mouse ES cells.
6.2. CRISPR/Cas9 editing of Jarid2 and Prickle1/Fzd2/Wnt9a in mouse embryonic stem cells
In Chapter 5, I demonstrated how CRISPR/Cas system can be used to modify Tet1
and Tet2 genes using published sgRNA sequences (Wang et al., 2013a). In this Chapter
I describe de-novo design of targeting sgRNAs to establish knockout mES cell lines for
Jarid2 (single) and for Prickle1, Fzd2 and Wnt9a (triple). The process of design is
especially important in view of the undesired off-target potential of CRISPR/Cas9 system.
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6.2.1. Guide RNA design and delivery into mouse ES cells
CRISPR/Cas9 DNA targeting generally leads to acquisition of random mutations
as a result of double strand break followed by error-prone non-homologous end joining
pathway. When an exon of a gene is targeted, acquired mutations can cause frameshifts
in the downstream coding sequence. This results in impaired protein function, an early
stop codon, or degradation. For this reason it is wise to target the start of the coding
sequence, paying attention to the presence of splice variants and alternative start codons
(which can be visualised at the UCSC Genome Browser available at
http://genome.ucsc.edu/).
In order to determine guide RNA sequences for targeting Jarid2, Prickle1, Fzd2
and Wnt9a, I used a bioinformatics tool which is available online at http://crispr.mit.edu/
(Hsu et al., 2013). This tool ranks candidate sgRNAs based on the number of potential
off-targets. Furthermore, I performed analysis of Genome-Wide Tag Scan (Iseli et al.,
2007) to evaluate the identities of the possible off-targets (See Chapter 6.4 for discussion).
I therefore eliminated all the sgRNA sequences with a high number of potential off-targets,
as well as those with off-targets falling into intragenic regions. Using the best candidates,
I constructed the px330 plasmid for the expression of sgRNA along with Cas9
endonuclease (Figure 6.1).
Figure 6.1. Jarid2, Prickle1, Fzd2 and Wnt9a targeting by CRISPR/Cas. Schematic representation of mouse Jarid2, Prickle1, Fzd2 and Wnt9a genes, and the DNA target sites. Nucleotides in red represent the PAM sequence, necessary for the target recognition by Cas9 endonuclease of Streptococcus pyogenes (-NGG). Nucleotides in blue depict sgRNA targeted DNA sites. Red arrows demonstrate the locations (3 base pairs downstream of PAM) of double strand breaks upon Cas9 endonuclease activity. Green arrows represent the transcription start sites. Sequences are not to scale.
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Here, the aim is to generate mES cells deficient for JARID2 as a control, and mES
cells simultaneously deficient for PRICKLE1, FZD2 and WNT9a. For this reason, I either
transfected mES cells with px330+sgJarid2 or co-transfected with px330+sgPrickle1,
px330+sgFzd2 and px330+sgWnt9a, along with pH2BmCherry-Ires-Puro vector for
selection. I used the same workflow depicted in Figure 5.5, to obtain samples for Surveyor
Assay and single colonies (32 colonies in each case).
6.2.2. Surveyor Assay for analysis of CRISPR/Cas9 efficiency
To determine whether the designed guide RNAs can efficiently target Cas9 to the
loci of interest and whether this results in random acquisition of mutations, I conducted
Surveyor Assay at population level. Strong appearance of cleavage products of the
targeted Jarid2 locus in mES cells indicated successful indel acquisition (Figure 6.2, left).
Similarly, sgRNA sequences tested for Prickle1, Wnt9a and Fzd2 efficiently targeted
corresponding loci in co-transfected mES cells (Figure 6.2, right).
Figure 6.2. Surveyor Assay on Puromycin treated populations of Jarid2 single and Prickle1, Wnt9a, Fzd2 triple CRISPR/Cas targeted mES cells. Jarid2 PCR produces 503 bp amplicon, Prickle1 PCR produces 499 bp amplicon, Wnt9a PCR produces 500 bp amplicon and Fzd2 PCR produces 488 bp amplicon. Respective cleavage product lengths are: ~203 and ~300, ~224 and ~275, ~302 and ~198, ~284 and ~204. In all conditions, the upper bands are the undigested amplifications (either wild type or no mismatch mutant amplicons).
6.2.3. Clonal screens and sequencing for targeted Jarid2 locus in mouse ES cells
Jarid2 CRISPR/Cas target location does not contain a restriction enzyme cut site.
For this reason, instead of using RFLP, I conducted Surveyor Assay at the clonal level.
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To do so, I mixed wild type DNA together with DNA derived from each clone in 1 to 1 ratio.
A major drawback of this screen (contrary to RLFP) is that the presence of cleavage
products does not necessarily demonstrate whether the mutations are mono- or bi-allelic.
The Surveyor Assay showed that the majority of the clones acquired mutations at the
targeted Jarid2 locus, with at least one allele modified (Figure 6.3).
Figure 6.3. Surveyor Assay on CRISPR/Cas targeted single mES cell clones for Jarid2. The assay was conducted on both wild type, and 32 clones selected upon Jarid2 targeting. Jarid2 PCR produces 503 bp amplicon with cleavage products at ~203 and ~300 bp.
To characterise the identities of the random mutations and determine those which
cause frame-shift at the coding sequence of Jarid2, I sequenced the target locus in eight
of the clones (Figure 6.4).
Figure 6.4. DNA sequencing results on Jarid2 CRISPR/Cas targeted locus. DNA sequences of the WT (blue) and targeted clones. Deletions are indicated as hyphens, insertions are indicated in red, and the indel sizes are indicated on the right for each sequence.
Sequencing of the Jarid2 target locus revealed that none of the selected clones
contained a wild type allele. Furthermore most of the acquired mutations caused shift in
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the frame of the coding sequence. Interestingly, one of the clones (Clone 9G) contained
three different alleles, which was most probably not due to a karyotypic instability, but due
to the concept of mosaicism. This occurs when the transfected single cell undergoes cell
division before its DNA could be targeted, which results in two genetically different
populations in the same colony.
To demonstrate the effect of frame-shift mutations, together with Amélie Feytout
we analysed the presence of JARID2 protein in three selected clones (9D, 11C and 12C)
by Western Blotting. The lack of full size JARID2 protein in the mutant clones confirmed
efficient CRISPR/Cas targeting in these ES cell lines (Figure 6.5).
Figure 6.5. Western Blot detection of Jarid2 in wild type and CRISPR/Cas targeted clones. Western blotting is conducted on whole-cell extracts of wild type mouse ES cells, and Clones 9D, 11C and 12C, using antibodies against Jarid2, and Lamin B as control.
6.2.4. Clonal screens and sequencing for targeted Prickle1, Fzd2 and Wnt9a loci in mES cells.
I selected 32 colonies from triple targeted mES cells, and screened those clones
for presence of mutations. The Fzd2 target locus contains a Tsp45i restriction enzyme
recognition site, allowing me to conduct RFLP screen on the clones (Figure 6.6).
Figure 6.6. Surveyor Assay on CRISPR/Cas triple targeted single mES cell clones for Fzd2. The assay was conducted on both wild type, and 32 clones selected upon triple targeting. Fzd2 PCR produces 488 bp amplicon with cleavage products at ~284 and ~204 bp, indicated by white arrows (wild type, left).
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Even though PCR amplification was not observed in some of the, many others
contained mono- or bi-allelic mutations at the Fzd2 target locus (demonstrated by the lack
of cleavage) (Figure 6.6). To further screen the clones for mutations on Prickle1, and
Wnt9a, I selected 13 clones where Fzd2 gene was modified in both alleles, and I
conducted Surveyor Assay (Figure 6.7.A-B).
Figure 6.7. Surveyor Assay on CRISPR/Cas triple targeted single mES cell clones for Prickle1 and Wnt9a. (A) Surveyor Assay on Prickle1 target locus on wild type (right) and 13 clones. Prickle1 PCR produces 499 bp amplicon with cleavage products at ~224 and ~275 bp. (B) Surveyor Assay on Wnt9a target locus on wild type (right) and 13 clones. Wnt9a PCR produces 500 bp amplicon with cleavage products at ~302 and ~198 bp.
These results revealed that majority of the selected 13 clones contained mutations
at both Prickle1 and Wnt9a target loci (Figure 6.7.A-B). However, it is not known whether
these mutations are mono- or bi-allelic, as this cannot be distinguished by the Surveyor
Assay. To determine the mutations, I went on to sequence co-targeted three loci in five
selected clones (Figure 6.8). Sequencing results revealed that almost all co-targeted sites
acquired indels. Interestingly, 3 of the 5 clones appeared to be mosaic for the targeted
Wnt9a locus (depicted by the presence of more than 2 alleles). Collectively, around two
thirds of the acquired indels resulted in frame-shifts, however it is difficult to predict
whether some of the in-frame mutations would cause a conformational change in the
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protein and decrease its functionality (such as 18 base deletions at the Allele 3 of targeted
Wnt9a in Clone 3A; Figure 6.8).
Figure 6.8. DNA sequencing results on Prickle1, Fzd2 and Wnt9a CRISPR/Cas co-targeted loci. DNA sequences of the WTs (blue) and targeted clones. Deletions are indicated as hyphens, insertions are indicated in red, and the indel sizes are indicated on the right for each sequence.
Because antibodies to PRICKLE1, WNT9A and FZD2 are not available, I was not
able to determine the effect of frame-shift mutations at the protein level. Instead, together
with Amélie Feytout we used an alternative approach to visualise gene knock-out. The
shift in the coding frame may lead to appearance of premature stop codons causing early
termination of translation. In turn, this affects the stability of mRNA at the translational
level, resulting in degradation. This concept is known as nonsense mediated decay
(Losson and Lacroute, 1979). We conducted transcript analysis for the targeted genes,
and showed that mutations in Prickle1 cause efficient decrease of mRNA levels. We also
observed this effect in Fzd2 and Wnt9a transcripts, yet not as efficiently as in the case of
Prickle1. This can be explained by the fact that Fzd2 and Wnt9a coding sequence sizes
are much smaller than Prickle1, possibly providing stability and prevention from decay
(Figure 6.9).
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Figure 6.9. mRNA levels of CRISPR/Cas targeted Prickle1, Fzd2 and Wnt9a in selected clones and wild type cells. qRT-PCR analysis of Prickle1, Fzd2, and Wnt9a expression in clones 2A, 2D, 3A, 3C, 3H and wild type parental cells. Data were normalised to Hmbs and represent mean and SEM of 3 biological replicates. Asterisks indicate statistical significance (p<0.05; Student’s t-test).
6.3. JARID2 deficiency in mouse ES cells can be phenocopied by Prickle1/Fzd2/Wnt targeting
Recent study by David Landeira (Landeira et. al, submitted) showed that non-
canonical Wnt pathway components were downregulated in previously established Jarid2
deficient mES cells (Shen et al., 2009). To investigate whether I can reproduce these data
in mES cells targeted by CRISPR/Cas9 for Jarid2, together with Amelie Feytout we have
conducted transcript analysis. We analysed the gene expression levels of Prickle1, Fzd2
and Wnt9a in three of the Jarid2 mutant mES cell clones (9D, 11C and 12C).
Figure 6.10. mRNA levels of Prickle1, Fzd2 and Wnt9a in Jarid2 mutant lines and wild type cells. qRT-PCR analysis of Prickle1, Fzd2, and Wnt9a expression in Jarid2 deficient clones 9D, 11C and 12C. Data were normalised to Hmbs and represent mean and SEM of 3 biological replicates. Asterisks indicate statistical significance (p<0.05; Student’s t-test).
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Compared to the parental wild type mES cells, Jarid2 mutants showed decreased
expression levels of Prickle1, Fzd and Wnt9a, recapitulating previous analysis (Figure
6.10).
Mouse ES cells that lack Jarid2 contain increased level of Nanog expression
(Landeira et. al, submitted). Based on this observation, together with Jorge-Soza Ried,
we analysed Nanog levels in mES cell clones 12C (Jarid2 mutant) and 2D (non-canonical
Wnt pathway mutant), both generated upon CRISPR/Cas9 editing. Flow cytometry
analysis demonstrated that CRISPR/Cas9 mediated Jarid2 deficiency resulted in
increased number of Nanog-high cells, in agreement with previously observed data
(Figure 6.11, left). Interestingly, knocking-out non-canonical Wnt pathway components
also lead to elevated number of Nanog-high cells in the population (Figure 6.12), as also
observed by immunofluorescence (data not shown). This suggests that Jarid2 might be
controlling steady state level of Nanog expression in wild type mES cells via non-canonical
Wnt pathway.
Figure 6.11. Flow Cytometry analysis of Nanog expression in mES cells Clones 12C and 2D. Flow cytometry analysis demonstrating Nanog expression at the population level in mES cell clones targeted for Jarid2 (12C) or Prickle1/Fzd2/Wnt9a (2D) (green traces) compared to wild-type parental ES cells (filled grey). The black line depicts negative control.
Furthermore, by collaborating with Transgenics Facility and Karen Brown, we
analysed individual contribution of parental wild-type cells and clones 12C and 2D to the
developing embryo. We injected wild-type blastocysts with either parental wild-type,
mutant Clone 12C, or mutant Clone 2D ES cells and cultured for 16 hours before
inspection. Remarkably, we observed that blastocysts injected with 12C or 2D clones
initiated the formation of more than one ICM (in around 40% of the blastocysts for either
of the clones), which was not detected in any of the control blastocysts injected with
parental wild-type cells (Landeira et. al, submitted).
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Collectively these data suggest that Jarid2 might be regulating Wnt signalling
components that in turn control steady state levels of Nanog expression and proper
development of the pre-implantation embryo.
6.4. Summary and Discussion
As a tool of genome engineering, CRISPR/Cas9 system has gained extensive
popularity due to its easiness, accuracy and efficiency. These features have enabled
scientists to simultaneously target multiple genes in a very short time, both for in-vivo and
in-vitro studies. In this chapter, I have not only demonstrated CRISPR/Cas9 design
process for DNA targeting, but also shown efficient generation of knock-out mES cell lines
for single (Jarid2) and multiple (Prickle1/Fzd2/Wnt9a) genes.
A major concern on CRISPR/Cas9 system has been the extent of potential off-
target mutagenesis it may cause. Cas endonuclease is brought on to the DNA by
complementation of 20-nucleotide long guide RNA and a study on bacterial genome has
revealed that mismatches outside the 12-base ‘seed region’ can be tolerated for targeting
(Jiang et al., 2013). This rises the chances of encountering off-target mutagenesis, even
though DNA cleavage will occur only if a PAM sequence is situated on the downstream of
the target sequence (-NGG for Cas9 from Streptococcus pyogenes). For this reason it is
important to use bioinformatics tools that help to rank guide RNA sequences according to
the possession of least off-targets, which preferentially fall into inter-genic non-conserved
areas in the genome (Hsu et al., 2013). I designed my targeting sequences according to
this ranking, and conducted a genome-wide homology search by using the ‘seed region’
to identify off-target locations (Iseli et al., 2007) Figure (6.12). This analysis demonstrated
that designed guide RNAs possessed limited number of off-targets, none of which
targeted a coding or conserved region.
As off-target potential is a primary drawback of the CRISPR/Cas9 system, many
studies have strived to minimize this effect. Feng Zhang lab has developed a ‘double
nicking’ system, where Cas9 nickase mutant was used to create single strand break upon
targeting. DNA editing is possible only a second Cas9 nickase mutant is targeted to the
complementary sequence at the same location. With this strategy, single strand off-target
breaks can be repaired with high fidelity by base-excision repair (Ran et al., 2013b). In
addition, Shen and colleagues have demonstrated that double nicking has significantly
reduced the number of double strand off-target cleavage compared to wild type Cas9, as
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shown by decreased H2AX recruitment to the cut sites (Shen et al., 2014). Development
of such strategies are crucial, especially for translational usage of CRISPR/Cas9 system.
Figure 6.12. Off-target identification of CRISPR/Cas9 targets. Table showing guide RNA sequences targeting Jarid2, Prickle1, Fzd2 and Wnt9a loci together with in-silico identified off-targets and their corresponding genomic coordinates.
Analysis of published ChIP-seq data for Jarid2 binding in mES cells has revealed
that Jarid2 has a strong preference for binding on Wnt signalling and Wnt-related pathway
components (Pasini et al., 2010). Among these are Prickle1, Fzd2 and Wnt9a, also bound
by Ezh2, which were downregulated in Jarid2 knock-out mES cells (Landeira et. al,
submitted) and upon JARID2 depletion by CRISPR/Cas9. This suggests that Polycomb
group proteins might be positively regulating active transcription of these particular genes,
as part of a recently reported feature that involves PRC2-dependent H3K27 mono/di-
methylation (Ferrari et al., 2014). It is important to note that Nanog gene is not bound by
JARID2, yet Nanog levels were significantly increased in mES cells that lack JARID2. My
results upon Prickle1/Fzd2/Wnt9a depletion suggest that this indirect regulation might
pass through non-canonical Wnt signalling pathway.
How does Wnt signalling control Nanog expression? NANOG is a core element of
pluripotency network and its expression fluctuates in mES cells leading to a bimodal
distribution (Chambers et al., 2007). mES cells expressing constitutively high levels of
Nanog are associated with naïve pluripotency, and this can be achieved by the usage of
Mitogen-activated protein kinase kinase (MEK) and Glycogen Synthase Kinase 3 (Gsk3)
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inhibitors (known as 2 inhibitors; 2i) (Ying et al., 2008). When not inhibited, Gsk3 leads to
the proteolysis of β-catenin (Aberle et al., 1997), a key component of Wnt signalling that
has been suggested to play major roles in pluripotent self-renewal. Upon Wnt activation,
β-catenin is accumulated in the nucleus where it interacts with Tcf/Lef (T-cell
factor/lymphoid enhancer factor) transcription factors and it has been reported that Gsk3
inhibition diminishes Tcf3 repression exerted on pluripotency network (Wray et al., 2011).
In addition, Tcf3 ablation has been shown to replace the requirement of Gsk3 inhibition in
terms of self-renewal (Yi et al., 2011). In line with these observations, activation of
canonical Wnt signalling reduces the levels of TCF3 which results in increased Nanog
levels and defects in differentiation (Atlasi et al., 2013). This is mainly because TCF3 acts
as a direct repressor of Nanog gene expression, and it is believed that this action provides
steady-state levels of NANOG for the maintenance of differentiation potential of
pluripotent cells (Pereira et al., 2006). It has been suggested that non-canonical Wnt
pathway may be negatively regulating the canonical pathway through the Tcf/Lef
transcription factors (Kühl et al., 2000). Additionally, Osei-Sarfo et al. demonstrated that
retinoic acid mediated mES cell differentiation lead to the activation of non-canonical Wnt
pathway, while canonical pathway is inhibited. This resulted in significant accumulation of
TCF3 on the promoters of pluripotency-associated genes and their subsequent repression
(Osei-Sarfo and Gudas, 2014).
Collectively, my data support a view that JARID2 might be regulating the interplay
between the canonical and non-canonical Wnt pathways that controls the self-renewal
and pluripotency of mES cells. Chromatin immunoprecipitation experiments would unravel
whether Nanog upregulation in Jarid2 deficiency is due to Tcf3 dissociation, regulated by
Prickle1/Fzd2/Wnt9a downregulation. Tcf3 displacement can occur upon β-catenin
mediated phosphorylation by homeodomain interacting protein kinase 2 (HIPK2) (Hikasa
et al., 2010), or by direct interaction with β-catenin (Solberg et al., 2012). Alternatively,
Tcf3 expression might be regulated at the expression level by changes in Wnt Signalling
(Atlasi et al., 2013).
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Chapter 7. General Discussion
In mammals, CpG methylation has been associated with gene repression.
Although the mechanisms of DNA methylation (both de-novo and maintenance) are well
characterised, how this is reversed in-vivo remained elusive. Recent studies have started
to unravel how DNA demethylation occurs initially by focusing on biological processes
such as pre-implantation embryo development and germ cell specification. In addition,
investigators have studied DNA demethylation that occurs during in-vitro pluripotent
reprogramming. It is now widely accepted that loss of DNA methylation takes place by two
distinct, but interconnected pathways: (1) passive, DNA replication-dependent
demethylation, and (2) active, DNA replication-independent demethylation. Passive
demethylation is based on a gradual dilution of 5-mC that occurs with successive rounds
of DNA replication and cell division when the methylation maintenance machinery is
disabled. Active demethylation involves enzymatic activity that modifies 5-mC, and
ultimately results its replacement by an unmodified cytosine residue. Here I have focused
on DNA methylation/demethylation dynamics during pluripotent reprogramming, where
DNA methylation has been considered to pose a major roadblock. Cell type specific DNA
methylation needs to be re-set for the acquisition of pluripotency (Pasque et al., 2011). In
this regard, cell fusion-based reprogramming provides a tractable experimental platform
to investigate the first signs of pluripotent conversion (within newly formed heterokaryons)
as the pluripotency-associated transcriptional programme is initiated.
7.1. DNA methylation dynamics in reprogramming
Our laboratory and others have demonstrated that pluripotent stem cells can
dominantly reprogram somatic cells upon fusion [(Soza-Ried and Fisher, 2012) and
recapitulated in this thesis]. These studies have suggested that the direction of conversion
(dominance) can be predicted and probably reflects the action of trans-acting factors that
also maintain the ‘stemness’ of pluripotent cells. Pluripotent cells possess so-called “open
chromatin” structure, which may allow cells to rapidly react to external differentiation
signals (Meshorer and Misteli, 2006). This property is maintained by global chromatin
remodellers that mediate histone tail modifications, nucleosome positioning and
reorganization, and interact pluripotency-associated network components (Gaspar-Maia
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et al., 2011). Interestingly, mouse ES cells treated with histone deacetylase (HDAC)
inhibitors were shown to be more efficient in reprogramming upon fusion with somatic cells
(Hezroni et al., 2011). In addition, ES cells that lack PRC activity have reprogramming
defects, consistent with a role for chromatin modifiers in pluripotent conversion (Pereira et
al., 2010).
Following differentiation, lineage restriction normally ensures the stable repression
of genes, particularly those involved in pluripotency. During this process, nucleosome-
depleted regions in mouse ES cells may be subject to nucleosome assembly followed by
acquisition of DNA methylation (You et al., 2011). In Chapter 3 of this study, I confirmed
the hypermethylated status of pluripotency-related OCT4, NANOG and CRIPTO genes in
differentiated human B lymphocytes and in human fibroblasts. Intriguing, although these
genes are rapidly induced upon cell fusion and reprogramming, bisulfite sequencing of the
promoter region of the OCT4 gene in heterokaryons did not reveal any significant loss of
DNA methylation. This could be attributed to the technical limitations (for example, 5-hmC
cannot be differentiated from 5-mC by bisulfite sequencing), or to the presence of
unreprogrammed heterokaryons within the population. Alternatively, loss of DNA
methylation may not be required for the initial activation of OCT4 gene expression. In
mouse, Oct4 gene expression and silencing is tightly controlled during embryogenesis
and silencing occurs via a series of events; repressor binding results in G9A-mediated H3
lysine 9 methylation that recruits HP1 that is eventually followed by de-novo DNA
methylation (Feldman et al., 2006). Gene reactivation may occur through a sequential
reversal of these events; histone remodelling followed by DNA demethylation. Upon
reprogramming by cell fusion, it is possible that the chromatin structure is relaxed and
modifications may help initiate OCT4 gene expression. A rapid global increase in H3K9
acetylation and H4 pan-acetylation have been reported as a distinguishing feature of
somatic cell-reprogramming by cell fusion (Piccolo et al., 2011). In addition, my preliminary
results (Chapter 5) suggest that active H3 lysine 4 tri-methylation is rapidly acquired on
the OCT4 gene promoter of somatic nuclei upon fusion with mouse ES cells. It has also
been shown that similarly, C/EBPα mediated pre-B cell conversion into macrophages
initiates macrophage specific gene expression programme with histone modification while
DNA methylation is maintained (Rodríguez-Ubreva et al., 2011). This could perhaps also
explain why the reprogramming potential of mouse ES cells depleted of TET proteins is
unchanged (at least at early stages), as gene induction may not be dependent on the
removal of DNA demethylation (as demonstrated in Chapter 5). It is noteworthy that a
recent study reported that TET activity was necessary in iPS cells only for the induction of
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miR-200 family members (which then facilitates the transition from mesenchymal to
epithelial state), but not for reactivation of pluripotency genes (Hu et al., 2014).
Tet proteins, on the other hand, may be required for mouse EG cell mediated
imprint erasure in mouse B lymphocyte genome upon fusion, (as shown in Chapter 4).
Here I showed that in mEG-hB heterokaryons TET activity resulted in 5-hmC accumulation
at imprinted control region DMRs, yet it is not until at least 7 days of fusion that I detected
DNA demethylation. This suggests an orchestrated action of both active and passive
demethylation processes; where actively hydroxylated 5-mC at DMRs may be passively
diluted with continuous cell replication. This mechanism may also be operating in-vivo
during PGC development, where TET enzymatic action and replication-coupled loss of
methylation has been evoked (Hackett et al., 2013).
7.2. Genome editing and the use of CRISPR/Cas9-based approaches
Genetic engineering is the process of targeted modification of genetic material.
Genome editing is usually achieved by homologous recombination (HR) in which an
externally provided DNA molecule serves as a template. The low efficiency of this event
(Capecchi, 1989) was shown to substantially increase by providing a DNA double-strand
break (DSB) at the target region. This occurs because the cell’s intrinsic DNA repair
mechanism can recognise the externally provided DNA as a template for correction via
homology-directed repair (HDR) pathway (Rouet et al., 1994). In the absence of a repair
template, non-homologous end joining pathway (an error-prone repair mechanism),
ligates cleaved DNA ends by inducing insertion and deletion mutations (indels) at the DSB
site (Moore and Haber, 1996). Both repair pathways have been extensively repurposed
for genome editing, however the major challenge has been to deliberately induce site-
specific DSBs among billions of DNA bases of the eukaryotic genome. For this purpose,
in the last decade four main strategies have been developed which exploit DNA-binding
proteins; meganucleases (Smith et al., 2006b), zinc finger nucleases (Urnov et al., 2005),
transcription activator-like effector nucleases (Miller et al., 2011) and recently, RNA-
guided Cas9 endonuclease, originating from microbial acquired immune system CRISPR
(clustered regularly interspaced short palindromic repeats) (Cong et al., 2013; Mali et al.,
2013). Thanks largely to basic research on bacteria, the CRISPR system has now been
re-purposed for gene editing, and in the last couple of years a plethora of studies have
proven its efficacy, despite initial off-target effects (see below). In chapters 5 and 6 of this
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document, I used CRISPR/Cas9 to generate a panel of mouse ES cell lines in which
Jarid2, Tet1/Tet2 and Prickle1/Fzd2/Wnt9a were targeted.
CRISPR originates in studies performed in the 1980s, when Atsuo Nakata and
colleagues encountered mysterious “unusual” repeats in Escherichia Coli while
sequencing the iap gene (Ishino et al., 1987). In the following years these repeats were
discovered in various prokaryotes, however it was not until 2002 that they were united
under the acronym CRISPR (Jansen et al., 2002). Briefly, CRISPR sequences are
composed of direct repeats of 21 to 37 base pairs with a loose dyad symmetry, and are
interspaced by similarly sized variable sequences (spacers) (Jansen et al., 2002). In
addition, adjacent to CRISPR loci, Jansen et. al. identified the presence of CRISPR-
associated (cas) genes which possess nuclease motifs, suggesting a possible functional
relation with the repeats (Jansen et al., 2002). Although well conserved among species,
at that time biological significance of CRISPR and Cas genes had remained elusive. Later,
several in-silico approaches identified spacers as originating from foreign DNA elements,
which led scientists to hypothesise that they may be involved in acquired immunity against
bacteriophages. Finally in 2007, direct evidence came from Horvath laboratory, where the
authors demonstrated that CRISPR is a prokaryotic resistance mechanism against
invading DNA molecules mainly originating from viruses, which involves recognition,
destruction and adaptation (Barrangou et al., 2007; Sorek et al., 2013). Studies on
Streptococcus thermophiles revealed three basic components of Type II CRISPR system
(Type I and Type III have slightly different mechanisms). These are Cas9 (or Cas5)
endonuclease that mediates DNA cleavage (Garneau et al., 2010), CRISPR-RNA (crRNA)
transcribed and processed from CRISPR array and trans-activating crRNA (tracrRNA) that
forms a hybrid with crRNA to guide Cas9 for targeting to the homologous DNA sequence
(Deltcheva et al., 2011). In-vitro studies demonstrated that Type II components can
efficiently be used to target and cleave plasmid DNA and to further simplify the system,
single guide RNA (sgRNA) was generated upon fusion of crRNA with tracrRNA (Jinek et
al., 2012) (Figure 7.1). Soon after, two studies simultaneously reported engineering of
Type II CRISPR system from Streptococcus pyogenes for mammalian genome
manipulation (Cong et al., 2013; Mali et al., 2013).
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Figure 7.1. Schematic representation of RNA-guided Cas9 targeting on DNA. The Cas9 endonuclease from Streptococcus Pyogenes (in beige) is recruited to the genomic DNA (in black) by sgRNA which is generated by fusion of 20-nt guiding crRNA (in purple) and scaffolding tracrRNA (in green). The guide RNA forms a heteroduplex with the DNA directly upstream of the 5’-NGG motif (Protospacer Adjacent Motif, PAM), which is a strict requirement for the Cas9 nuclease activity. Red triangles show the DSB locations mediated by Cas9, located at the 3-bp upstream of the PAM sequence.
High-resolution structural investigations showed that RNA binding (crRNA-
tracrRNA complex, or sgRNA) is necessary for structural rearrangement and consequent
activation of Cas9 to exhibit DSB upon targeting (Jinek et al., 2014; Nishimasu et al.,
2014). Single molecule imaging using DNA curtains demonstrated that RNA-guided Cas9
first interacts with Protospacer Adjacent Motif (PAM, Figure 7.1) upon random collisions
along the DNA (Sternberg et al., 2014). PAM recognition is followed by DNA strand
separation and sequential extension of the guide RNA-DNA heteroduplex starting from
the PAM (Sternberg et al., 2014). This mechanism also explains the importance of 8-12
nucleotide-long seed sequences on the downstream of the guide RNA where any
nucleotide mismatch would terminate the heteroduplex formation, while upstream
mismatches can be tolerated (Jiang et al., 2013; Sternberg et al., 2014).
In the last couple of years, numerous laboratories have conducted CRISPR/Cas9
mediated genome manipulation in both cell lines and animal models and the number of
such studies have been expanding at a dazzling pace (Sander and Joung, 2014). One of
the biggest advantage of this technique is the possibility to simultaneously edit multiple
genes -a process that would take a long time using conventional methods. For example
Rudolph Jaenisch and colleagues simultaneously disrupted five genes (Tet1, Tet2, Tet3,
Sry and Uty) in mouse ES cells based on indel acquisition upon NHEJ (Wang et al.,
2013a), and rapidly generated reporter cell lines by HDR (Yang et al., 2013). In addition,
upon zygotic injection of Cas9 mRNA and sgRNAs, the authors have been able to
generate knock-in or knock-out mice without any need for further breeding (Wang et al.,
2013a; Yang et al., 2013). Moreover, it has been possible to correct genetic diseases in
human cells (Schwank et al., 2013) and in mouse zygotes (Long et al., 2014; Wu et al.,
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2013b) and new reports demonstrated the proof-of-principle that CRISPR/Cas9 system
can be used in adult mice by hydrodynamic tail injection of the components (Xue et al.,
2014; Yin et al., 2014). Alternatively, catalytically inactive Cas9 can be used to alter
epigenetic signature of a targeted site upon tethering to transcriptional regulators, to
activate (Maeder et al., 2013; Perez-pinera et al., 2013), or to repress (Gilbert et al., 2013)
gene expression. In addition, lentiviral sgRNA libraries enabled genome-wide knock-out
screens (Shalem et al., 2014; Wang et al., 2014).
One major concern in CRISPR based gene editing has been the potential of
unwanted off-target mutagenesis as mismatches in target recognition by guide RNA can
be tolerated to some extent. Several studies have reported that undesired modifications
can occur at different sites in the genome with higher levels than expected (Fu et al., 2013;
Hsu et al., 2013; Pattanayak et al., 2013; Shen et al., 2014). However recent studies
demonstrated that careful design of guide RNA sequence (Cho et al., 2014) can
significantly reduce off-target potential as analysed by whole-genome sequencing in
human pluripotent ES and IPS cells (Smith et al., 2014; Suzuki et al., 2014; Veres et al.,
2014). These studies caution that careful analysis and design are required to minimise the
adverse off-target effects of CRISPR targeting.
Here, I used CRISPR/Cas9 to generate single (Jarid2), double (Tet1/Tet2) and
triple (Prickle1/Fzd2/Wnt9a) knock-out mouse ES cell lines. In-vitro and in-vivo analyses
showed that Prickle1/Fzd2/Wnt9a knock-out mouse ES cells phenocopied the effects of
Jarid2 depletion in terms of altered blastocyst development and aberrant cell sorting.
Using these novel cell reagents, I intend to extend these studies and examine the roles of
non-canonical Wnt pathway components in embryogenesis in future studies.
7.3. Future Studies
To dissect the early molecular events that are required for reprogramming I will
explore two main approaches in the future. The first approach is based on microfluidic
systems and aims to establish a high-throughput platform for heterokaryon generation.
This will be important to allow us to conduct epigenomic (ChIP for histone modifications
and transcription factor binding) studies that require a relatively high number of fused cells.
The second approach is based on non-destructive imaging of single heterokaryons to
elucidate the kinetics and order of events underpinning chromatin reorganisation, and
DNA demethylation during reprogramming.
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7.3.1. Droplet-based microfluidics and cell fusion
Although cell fusion-based reprogramming is an efficient process, the major
limitation of conventional fusion protocols has been the low number of 1:1 heterokaryons
obtained. In fusion with PEG, electric pulse or virus-mediated, cell pairing is random, and
the resulting population contain a heterogeneous mixture of both cell types. This
eventually leads to unwanted fusion products as a result of pairing of same kinds of cells.
Fused cells ofthen have to be selected by using chemical selection, or by cell sorting using
flow cytometry. Therefore, the protocol is time consuming and yields only small numbers
of viable heterokaryons of the desired type. To find a solution we decided to look into
microfluidic systems to increase fusion efficiency by improving the cell pairing and
membrane fusion in a high-throughput manner.
Microfluidics is the science of fluids confined to a miniature scale and has been
used to analyse small sample volumes in drug discovery, medical diagnostics, genomics,
molecular biology and high-throughput screening [with the idea of miniaturising a
laboratory into a chip (Whitesides, 2006)]. Microfluidic platforms have recently been
repurposed to precisely control and manipulate cells in microenvironments leading to next-
generation living-cell microarrays (Yarmush and King, 2009). In addition to many
applications, innovative microfluidic designs have been tested in the past for efficient cell
fusion. One study reported a microfluidic electrofusion system, where cell conjugation was
achieved by biotin-streptavidin coating and paired cells were flowed in a specific
microfluidic channel under continuous direct current, which consisted of narrow and wide
sections. The field intensity at the centre of the narrow channel enabled paired cells to
fuse during their passage (Wang and Lu, 2006). Electrofusion has been used in various
microfluidic formats, including micro-electrode arrays (Cao et al., 2008; Hu et al., 2011;
Qu et al., 2011), micro-orifices (Kimura et al., 2011) and in several other platforms (Hu et
al., 2013). An alternative approach has been the fabrication of thousands of microscaled
polydimethyl siloxane traps into a flow-through channel to increase the pairing efficiency.
The traps were designed to capture both cell type that are sequentially loaded into the
chip. Once the cells were paired and immobilized, PEG or electric field were applied for
fusion (Dura et al., 2014; Kemna et al., 2011; Skelley et al., 2009). However as yet high-
throughput reprogramming has not been established. Droplet-based microfluidic platforms
offer the ability to process millions of individual assays in very short times and high
reproducibility (Huebner et al., 2008). These systems are briefly based on combining two
immiscible phases (water and oil) by segmented-flow, where shear force and interfacial
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tension spontaneously generate microdroplets at rates of up to several kHz (Huebner et
al., 2008).
Together with Dr. Andrew deMello’s research team we have begun to generate
prototype droplet-based high-throughput cell fusion systems. This involves encapsulation
of either of the fusion partners in droplets (Köster et al., 2008) followed by droplet merging
(Niu et al., 2008), sorting droplets that contain both cell types (Baret et al., 2009) and
passing them through electric field to induce cell fusion. Although cell encapsulation
follows Poisson statistics (Köster et al., 2008), it is possible to defeat stochastic loading
by engineering alternative micro-channel designs to ensure proper cell ordering (Edd et
al., 2008; Hur et al., 2010; Kemna et al., 2012). Together with ultra-high-speed droplet
generation and cell encapsulation, it would be possible to process millions of events only
in a short period of time, significantly increasing the number of cell fusion events. This
could provide us with a bespoke of “pure” heterokaryon population for molecular analyses.
One of these is chromatin immunoprecipitation experiments to determine the
initiation and the kinetics of reprogramming at the chromatin level. Although I have been
able to demonstrate a trend of H3K4me3 acquisition at the human OCT4 promoter in B
cells upon fusion with mouse ES cells, it would be of interest to analyse further histone
modifications in various loci, and eventually to conduct ChIP-sequencing at a global level.
Furthermore, analysis of how mouse specific pluripotency-associated factors originating
from mouse ES cells bind to and act on the human genome of the somatic cell upon fusion
would shed light on how pluripotent conversion is induced. Specifically, our laboratory has
previously shown that mouse Oct4 protein is very rapidly accumulated on the somatic
DNA as early as 6 hours after fusion. ChIP experiments on mouse Oct4 would provide a
valuable spatio-temporal information on the establishment of pluripotency network at the
somatic DNA. It is noteworthy that recent advances in ChIP sensitivity suggest that it may
be possible to conduct similar experiments using relatively smaller number of (fused) cells
(Lara-Astiaso et al., 2014).
A second approach for which high number of heterokaryons would be needed is
the analysis of the chromatin environment of the newly replicated DNA in heterokaryons.
In our laboratory we have shown that cell cycle stages of the fusion partners affect the
efficiency of reprogramming, and require DNA replication by the somatic nucleus
(Tsubouchi et al., 2013). By using 5-ethynyl-2’-deoxyuridine (EdU) labelling of newly
replicated DNA, together with click-chemistry, EdU containing DNA can be pulled-down
(with the surrounding chromatin intact) so that we can characterise the first epigenetic
events in cell-type conversion (Kliszczak et al., 2011).
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Chapter 7
7.3.2. Single-cell heterokaryon analysis
Although maximizing number of “purified” heterokaryons is a prerequisite for such
techniques, recent developments have enabled valuable information to be gained from
studying single cells. Therefore, during the process of establishing a fully functional
microfluidics based high-throughput cell fusion system, I will also be conducting single cell
transcription and DNA methylation analyses on heterokaryons.
Microfluidics based transcription analysis arrays (Fluidigm) have recently been
used to measure expression levels of tens of genes in hundreds of single cells in parallel.
One example is by Guo et. al. where the authors investigated the differentiation process
in embryogenesis (Guo et al., 2010). They analysed mRNA levels of 48 genes on single
cells starting from 1-cell zygote and have been able to characterize three distinct cell types
in 64-cell blastocysts according to their expression profiles (Guo et al., 2010). Similar
setting was later used in iPS cell reprogramming by Rudolph Jaenisch and colleagues,
where the analysis of 48 genes in single cells helped to identify the order of events during
pluripotent conversion (Buganim et al., 2012). According to that study, reprogramming first
starts as a stochastic event and single cells exhibit great heterogeneity which is then
followed by a hierarchical phase, where activation of key pluripotency factors lead to a
predictable series of events for the acquisition of pluripotency (Buganim et al., 2012).
Single cell mRNA investigation of heterokaryons will provide informative data on the
kinetics of gain of pluripotency-associated and loss of somatic-cell-specific gene
expression profiles. This will help me determine the timing of key events, and identify the
fundamentals of reprogramming. In addition, as the pluripotent conversion is much quicker
compared to iPS cell reprogramming, by using RNAi or CRISPR based approaches we
can easily scan for necessary factors and their consecutive roles upon cell fusion. By
combining single cell microarrays with next-generation sequencing it is now possible to
conduct RNA-sequencing with decreasing complexity and cost (Kalisky et al., 2011;
Shapiro et al., 2013). This will deliver a global view on how single heterokaryons behave
at the transcriptional level, and lead to the identification of the concept of cellular
dominance.
With recent advances in miniaturisation, it is now possible to analyse the status of
DNA methylation at a single cell level. The first example of this technique came from Axel
Schumacher laboratory, where a micro-reaction slide was used to conduct PCR based
methylation-sensitive restriction assay to detect DNA methylation levels at particular loci
on single cells (Kantlehner et al., 2011). A similar approach was used to simultaneously
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Chapter 7
investigate methylation levels of 6 imprinted genes in each single cell of 8-cell pre-
implantation embryos, in order to determine how the loss of Trim28, involved in DNMT1
recruitment, can result in defects in imprint maintenance (Lorthongpanich et al., 2013).
Based on this idea, I will conduct single-heterokaryon DNA methylation analysis on
important pluripotency loci, to analyse the kinetics of DNA demethylation. Comparison of
single cell gene expression and spatiotemporal methylation levels will give information of
epigenetic dynamics of reprogramming, which is not easy to obtain by averaging the
population. In addition, I will be able to screen for candidate factors thought to be directly
or indirectly involved in DNA demethylation in a robust and controllable system.
Furthermore, in the long term, single cell genome-wide bisulfite sequencing can be
conducted which can presently detect DNA methylation at almost half of the CpG sites
(Smallwood et al., 2014). Finally, imaging of heterokaryons can give clues on the real-time
kinetics of reprogramming. As discussed (in Chapter 5), I have conducted live cell imaging
on heterokaryons generated between mouse ES cells expressing H2BmCherry and
human fibroblasts expressing HP1αGFP and demonstrated fusion of distinct nuclei
(Cantone et al, submitted). I hope that using these cell lines and microfluidic platforms, I
will be able to apply single-cell live imaging to study hundreds of cells in a single chip
(Kellogg et al., 2014). These approaches will aim to identify the initial early events that
dictate the conversion of one cell type into another, and provide a platform that allows
these events to be interrogated and tested.
127
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Appendix
Appendix Oligonucleotides
Table 1. Primers for transcript analysis by quantitative RT-PCR
Human-specific primers Gene Sequence 5’ – 3’
GAPDH Forward TCT GCT CCT CCT GTT CGA CA Reverse AAA AGC AGC CCT GGT GAC C
OCT4 Forward TCG AGA ACC GAG TGA GAG GC Reverse CAC ACT CGG ACC ACA TCC TTC
NANOG Forward CCA ACA TCC TGA ACC TCA GCT AC Reverse GCC TTC TGC GTC ACA CCA TT
CRIPTO Forward AGA AGT GTT CCC TGT GTA AAT GCT G Reverse CAC GAG GTG CTC ATC CAT CA
CD19 Forward GCT CAA GAC GCT GGA AAG TAT TAT T Reverse GAT AAG CCA AAG TCA CAG CTG AGA
CD45 Forward CCC CAT GAA CGT TAC CAT TTG Reverse GAT AGT CTC CAT TGT GAA AAT AGG CC
Mouse-specific primers Gene Sequence 5’ – 3’
Ubc Forward GTC TGC TGT GTG AGG ACT GC Reverse GTC TTG CCT GTC AGG GTC TT
Oct4 Forward CGT GGA GAC TTT GCA GCC TG Reverse GCT TGG CAA ACT GTT CTA GCT CCT
Nanog Forward GAA CTA TTC TTG CTT ACA AGG GTC TGC Reverse GCA TCT TCT GCT TCC TGG CAA
Tet1 Forward GAG CCT GTT CCT CGA TGT GG Reverse CAA ACC CAC CTG AGG CTG TT
Tet2 Forward TGT TGT TGT CAG GGT GAG AAT C Reverse TCT TGC TTC TGG CAA ACT TAC A
Prickle1 Forward ATG GAT TCT TTG GCG TTG TC Reverse TGA CGG TCT TGG CTT GCT
Fzd2 Forward GAC ACC AGG GCT GAA GAG TG Reverse AAG GGC ACT TAG AAA AGT CGA G
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Appendix
Wnt9a Forward CGA GTG GAC TTC CAC AAC AA Reverse GGC ATT TGC AAG TGG TTT C
Table 2. Primers for bisulfite sequencing analysis
Human-specific primers Locus Sequence 5’ – 3’
OCT4 Forward AAA GTT TTT GTG GGG GAT TTG TAT T Reverse AAA ACC TAA CCC AAC CCC CAA
NANOG Forward TTA ATT TAT TGG GAT TAT AGG GGT G Reverse AAA CCT AAA AAC AAA CCC AAC AAC
CRIPTO Forward GGA GGA TTG AAA TGT TAG GTG AG Reverse AAA TTT ATC TCA ACC TCC CAA CTC
OCT4 (Bhutani et al., 2010) Forward GGA GAG GGG GTT AAG TAT TTG GGT TTT Reverse TCC ACT TTA TTA CCC AAA CTA A
Mouse-specific primers
Locus Sequence 5’ – 3’
Oct4-Gfp Forward GGG GTT AGA GGT TAA GGT TAG AGG Reverse ACC AAA ATA AAC ACC ACC CC
H19 Forward AAG GAG ATT ATG TTT TAT TTT TGG A Reverse AAA AAA ACT CAA TCA ATT ACA ATC C
Peg1 Forward GAT TAG AGA TTT ATA AGG AAA GAG Reverse CAA CAA AAA CAA CAA ACA ACA AC
LINE1 Forward GTT AGA GAA TTT GAT AGT TTT TGG AAT AGG Reverse TCA AAC ACT ATA TTA CTT TAA CAA TTC CCA
Peg3 Forward TTG ATA ATA GTA GTT TGA TTG GTA GGG TGT Reverse ATC TAC AAC CTT ATC AAT TAC CCT TAA AAA
Gtl2/Dlk1 Forward GGA AGG AAA AGA TAA AAT GTA GAA A Reverse CAT AAA TAA ATA AAC CCA TAA TCC C
Table 3. Primers for enzyme protection assay
Human-specific primers Locus Sequence 5’ – 3’
H19 Digestion Forward ACT GAA GCC CTC GGA GTG T
Reverse AGA TCT TCA GGT CGG GCA TT
Normalisation Forward GAT AAT GCC CGA CCT GAA GA Reverse GGG GTC ATC TGG GAA TAG GA
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Appendix
PEG3 Digestion Forward AAA ACC CCT ACA GGC AGG AC
Reverse GCG AAA ATG CCC CTT CCT
Normalisation Forward GAA AAC CCC TAC AGG CAG GA Reverse TTG TTT GCC GCA GTG GTG
SNRPN/SNURF Digestion Forward ACT GCG GCA AAC AAG CAC
Reverse CTC CTC AGA CAG ATG CGT CA
Normalisation Forward ACT GCG GCA AAC AAG CAC Reverse CAG GCT TCG CAC ACA TCC
Table 4. Primers for ChIP assay
Human-specific primers Locus Sequence 5’ – 3’
β-ACTIN Forward GAT CAG CAA GCA GGA GTA TGA CG Reverse AAG GGT GTA ACG CAA CTA AGT CAT AG
OCT4 Forward TTG CCA GCC ATT ATC ATT CA Reverse TAT AGA GCT GCT GCG GGA TT
TSH2B Forward Diagenode, pp-1041–500 Reverse
Table 5. Primers for genomic DNA amplification for Surveyor and RFLP Assays
Mouse-specific primers Gene Sequence 5’ – 3’
Tet1 Forward TTG TTC TCT CCT CTG ACT GC Reverse TGA TTG ATC AAA TAG GCC TGC
Tet2 Forward CAG ATG CTT AGG CCA ATC AAG Reverse AGA AGC AAC ACA CAT GAA GAT G
Jarid2 Forward GGC ACA GGG TAG AAG GAA AA Reverse ATT CCA GGG GTC CTT GAG TT
Prickle1 Forward TGG CCA TTG GCT TAT TTT TC Reverse AAC ACA ACC CAC AGG AAA GC
Fzd2 Forward ACA TCG CCT ACA ACC AGA CC Reverse GAG ATA GGA CGG CAC CTT GA
Wnt9a Forward GTG CTC TGG CTC CTC TGT TC Reverse TGT GCC CAG TAG AAG GGT TT
157
Publications Part of the work presented here has been published as follows:
Piccolo, F.M., Bagci, H., Brown, K.E., Landeira, D., Soza-Ried, J., Feytout, A., Mooijman, D., Hajkova, P., Leitch, H.G., Tada, T., Kriaucionis, S., Dawlaty, M.M., Jaenisch, R., Merkenschlager, M., and Fisher, A.G. (2013). Different roles for Tet1 and Tet2 proteins in reprogramming-mediated erasure of imprints induced by EGC fusion. Mol. Cell 49, 1023–1033.
Tsubouchi, T., Soza-Ried, J., Brown, K., Piccolo, F.M., Cantone, I., Landeira, D., Bagci, H., Hochegger, H., Merkenschlager, M., and Fisher, A.G. (2013). DNA synthesis is required for reprogramming mediated by stem cell fusion. Cell 152, 873–883.
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