Alkyladenine DNA Glycosylase (Aag)-Dependent
Cell-Specific Responses to Alkylating Agents
by
Carrie Marie Margulies
B.A. Chemistry
Dartmouth College, 2008
Submitted to the Department of Biological Engineering
in Partial Fulfillment of the Requirements for the Degree of
DOCTOR OF PHILOSOPHY
at the
MASSACHUSETTS INSTITUTE OF TECHNOLOGY
February 2016
@ 2016 Massachusetts Institute of Technology. All rights reserved.
Signature of
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Department of Bioko'cal Engineering
Jnuary 8 th 2016U
~zignatu re redactedi--j--_
.... . . . . 
Leona D. Samson
Professor of Biological Engineering and Biology
Thesis Supervisor
Accepted by: Signature redacted
Chair, Biologic
.........................
Forest White
Professor of Biological Engineering
al Engineering Graduate Committee
MASSACHUSETTS INSTITUTEOF TECHNOLOGY-
MAY 2 6 2015
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Alkyladenine DNA Glycosylase (Aag)-Dependent
Cell-Specific Responses to Alkylating Agents
By
Carrie Marie Margulies
Submitted to the Department of Biological Engineering on January 81h, 2016 in
Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy in
Biological Engineering
Abstract
Methylating agents are ubiquitous in our internal and external environments and
can cause damage to all cellular components, including our DNA. If left
unrepaired, methylated DNA can cause mutations, cell death, and disease, such
as cancer and neurodegeneration. The majority of DNA lesions caused by
methylating agents are repaired by the base excision repair (BER) pathway,
which is initiated by the lesion-specific alkyladenine glycosylase (Aag). Loss of
Aag in embryonic stem (ES) cells renders them sensitive to the methylating
agent MMS (methyl methanesulfonate) compared to wild-type (WT). Surprisingly,
this phenotype is reversed in hematopoietic myeloid progenitors and cerebellar
granule neurons (CGNs) where Aag' cells are resistant to MMS induced killing
compared to WT. In this study, we investigated how Aag can cause cell-specific
responses to alkylating agents.
We generated new WT, Aag', and Aag overexpressing (mAagTg) 129 and
C57B1/6 ES cells and showed that inbred genetic background did not affect
sensitivity to MMS, indicating this to be a cell-intrinsic response. Moreover, we
found that cells overexpressing Aag were even more sensitive to MMS than
Aag' cells, suggesting that ES cells endure methylation treatment best when
they express Aag within an optimal range.
To study Aag-dependent neural sensitivity to methylating agents, we optimized
protocols for the isolation and culture of primary cerebellar granule neurons and
determination of cell death after drug treatment by high-throughput imaging.
CGNs isolated from WT, Aag, and mAagTg mice exhibited cell sensitivity to
MMS treatment that was dependent on Aag and Parp activity, thus recapitulating
in vivo results and proving that CGN death is cell-intrinsic. Cell death was
independent of caspases, mitochondrial depolarization, and AIF translocation.
We did observe the formation of enlarged mitochondria and are investigating
whether mitochondrial dynamics are causative of cell death in an Aag-dependent
manner.
Finally, we used in vitro hematopoietic and neuronal differentiation to monitor cell
responses to MMS as a function of cellular development. Three different
methods all successfully generated mature neurons based on morphology,
2
immunochemical staining, and Aag expression. Though we successfully
differentiated ES cells into cell types of interest, we are continuing to optimize
methods for the assessment of alkylation sensitivity in the resulting
heterogeneous populations.
Thesis Supervisor: Leona D. Samson
Title: Professor of Biological Engineering and Biology
3
Acknowledgements
I would like to begin by thanking my advisor, Leona Samson, for her endless
support and guidance throughout my graduate career. Through all the ups and
down of research, she was a constant figure I could rely and depend on. I would
also like to thank my committee members Bevin Engelward and Doug
Lauffenburger for their assistance and contributions to my project.
I would not have been able to complete this without the assistance of all the
members of the Samson lab, past and present. Whether during group meeting,
over lunch/coffee, or outside of the lab, they have been wonderfully helpful on all
aspects of my scientific and personal life. They made the lab an enjoyable place
to work everyday and I look forward to staying friends with them for a long time.
Next, I want to acknowledge all my close friends who have been there for me
throughout the past 6 years. To the BE Class of 2009, there is no way I would
have survived at MIT without your assistance in the dungeon during the first year!
In the years since then, each one of you is the reason MIT is exciting,
invigorating, and full of laughter. To all my close friends in the Boston area,
particularly all my Dartmouth hockey friends, thank you for reminding me to take
breaks and relax to make cookies, dance, reminisce, and be 'me', no matter how
crazy that is. To Sue, thank you for your 'wellness checks', Dan and I would not
be the same without them.
Finally, I would like to thank my family. To my wonderful husband Dan, thank you
for pushing me to become all that you know I can be and supporting me every
step along the way. To my doggies Ted and Ruby, thank you for always being
happy to see me, making me smile, and reminding me of the importance to relax
and enjoy life. To my parents, Matt and Julie, thank you for your infinite support,
financially and emotionally, and dedication toward all the endeavors I have
undertaken, from hockey to academics. I would not have achieved everything I
4
have without your love and confidence in my abilities. To my brothers, Bud and
Jake, thank you for your competitive spirits, without which I would have never
learned how to push myself to the limits. To my aunt Trudy, though I have not
always appreciated you as much as you deserve, you have been nothing but
generous, dependable, and encouraging. To the rest of my enormous family, all
the grandparents, aunts, uncles, cousins, etc., you have given me the love,
support, and motivation I have needed to accomplish my goals and I hope I can
offer you all the same. Finally, I'd like to thank my new family. Thank you to
David, Else, Maja, and Nik for taking me in and providing me a family in Boston.
In particular, thank you David for all of the scientific insight and support you have
provided me in the past few years.
5
Table of Contents
Acknowledgements ........................................................................................... 4
Table of Contents .............................................................................................. 6
L ist o f F ig u re s .................................................................................................... . 9
L ist o f T a b le s .................................................................................................. . . 12
Chapter I: Introduction ..................................................................................... 16
Intro d u ctio n ................................................................................................ . . 16
Alkylating Agents in our Endogenous and Exogenous Environments Cause
Cytotoxic DNA Damage ............................................................................ 16
Methylating Agent Mechanism of Action .................................................. 17
DNA Repair Pathways for Alkylation Damage ......................................... 18
Alkyladenine Glycosylase (Aag) Initiates Repair of Methylated DNA Bases2l
Cellular Consequences of Imbalanced BER ........................................... 22
Alkylation Sensitivity of Cells with Altered Aag Expression.......................23
Poly (ADP-Ribose) Polymerase 1 Can Mediate Cell Death Caused by BER
Im b a la n c e s ................................................................................................... 2 5
Model System to Study Aag-Dependent Alkylating Sensitivity.................27
Overview of the Presented Study..............................................................27
F ig u re s ....................................................................................................... . . 2 9
T a b le s ......................................................................................................... . . 3 6
R e fe re n ce s .................................................................................................. . 3 7
Chapter II: Generation of WT, Aag' and mAagTg C57B1/6 and 129 Embryonic
Stem Cells and Sensitivity to Alkylating Agents ............................................. 51
In tro d u ctio n ................................................................................................ . . 5 1
Materials and Methods..................................................................................54
R e s u lts ....................................................................................................... . . 6 2
Generation of C57B1/6 and 129 Mouse Embryonic Stem Cells and Validation
of Pluripotency.......................................................................................... 62
6
BER Gene Expression and Aag Activity in WT, Aag-'- and mAagTg ES Cells
..................................................................................................................... 6 3
Sensitivity of ES Cell Lines to Alkylating Agents....................................... 64
Discussion..................................................................................................... 65
Figures ......................................................................................................... 69
References................................................................................................... 78
Chapter III: In vitro Hematopoietic and Neuronal Differentiation of Mouse
Em bryonic Stem Cells ..................................................................................... 86
Introduction .................................................................................................. 86
M aterials and Methods................................................................................. 89
Results ......................................................................................................... 95
In Vitro Hematopoietic Differentiation of 129 ES Cells.............................. 95
In vitro Differentiation of Cerebellar Granule Neurons (CGNs) ................ 96
In vitro Adherent Differentiation of Cortical Neurons................................. 98
In vitro Neural Differentiation Following 'Bibel' et al. (2007).......................100
D is c u s s io n ...................................................................................................... 1 0 1
F ig u re s ........................................................................................................... 1 0 5
T a b le s ............................................................................................................ 1 1 5
References.....................................................................................................116
Chapter IV: Primary Mouse Cerebellar Granule Neuron Sensitivity to MMS is
Dependent on Aag and Parp1 ........................................................................... 123
In tro d u c tio n .................................................................................................... 1 2 3
M aterials and Methods...................................................................................127
R e s u lts ........................................................................................................... 1 3 3
Differential Expression of Aag in Primary Cerebellar Granule Neurons
causes changes M MS Sensitivity ex vivo...................................................133
Aag-Dependent CGN Sensitivity to MMS is mediated through Poly (ADP-
Ribose) Polymerase Activity.......................................................................134
Downstream Modulators of Neuron Sensitivity to M MS ............................. 136
Base Excision Repair Protein Expression in Primary CGNs......................138
D is c u s s io n ...................................................................................................... 1 3 9
7
F ig u re s ........................................................................................................... 1 4 4
A p p e n d ix ........................................................................................................ 1 6 4
CGN Sensitivity to Glutamate Excitotoxicity and Oxidative Stress is
Independent of Aag Activity........................................................................164
Genetic Deletion of Alkbh7 Reduces Female CGN Sensitivity to MMS.....165
R e fe re n c e s ..................................................................................................... 16 8
Chapter V: Discussion ....................................................................................... 180
D is c u s s io n ...................................................................................................... 1 8 0
Imbalances in DNA Repair alter Embryonic Stem Cell Sensitivity to
A lky la tin g A g e nts ........................................................................................ 18 0
Aag and Parp1 Mediate Cerebellar Granule Neuron Sensitivity to MMS... 184
Aag-Dependent Cell-Specific Responses .................................................. 187
R e fe re n c e s ..................................................................................................... 18 9
Appendix: Development of a Method to Assay DNA Repair Capacity for
Mammalian Cells using High-Throughput Sequencing ..................................... 196
In tro d u c tio n .................................................................................................... 1 9 6
Manuscript: Multiplexed DNA repair assays for multiple lesions and multiple
doses via transcription inhibition and transcriptional mutagenesis. ............... 197
8
List of Figures
Figure 1.1: Structures of Aag DNA Substrates................................................29
Figure 1.2: Base Excision Repair Pathway. ..................................................... 30
Figure 1.3: Alternate DNA Repair Pathways for Methylation Damage. ........... 32
Figure 1.4: Parp1 utilizes NAD' to generate PAR polymers.............................33
Figure 1.5: Alternative mechanisms of Cell Death Caused during Base Excision
R e p a ir (B E R ). ............................................................................................... . . 34
Figure 2.1: Generation of new Embryonic Stem (ES) Cells. ........................... 69
Figure 2.2: Karyotyping of C57B1/6 ES Cells.................................................. 70
Figure 2.3: Fluorescent Immunocytochemical Staining for Pluripotency Markers
in C 57B1/6 and 129 ES C ells. ......................................................................... 71
Figure 2.4: Aag Expression and Activity in C57BI/6 ES Cells. ......................... 72
Figure 2.5: Aag Expression and Activity in 129 ES Cells. ............................... 73
Figure 2.6: Expression of Base Excision Repair genes in ES cells.................74
Figure 2.7: Sensitivity of C57B1/6 ES Cells to Alkylating Agents. ..................... 75
Figure 2.8: Sensitivity of 129 ES Cells to Alkylating Agents............................76
Figure 2.9: MMS Sensitivity of 129 Hematopoietic Progenitors, Cerebellar
Granule Neurons, and Retinal Photoreceptors is Aag-dependent. ................. 77
Figure 3.1: Mouse embryonic stem cells will be differentiated in vitro into
hematopoietic progenitors and cerebellar granule neurons. ............................. 105
Figure 3.2: In vitro Hematopoietic Differentiation of 129 ES cells. .................... 106
Figure 3.3: In vitro hematopoietic differentiation of VVT and Aag-'- 129 ES Cells
a nd M M S se nsitiv ity ........................................................................................... 10 7
Figure 3.4: In vitro Differentiation of Cerebellar Granule Neurons. ................... 108
Figure 3.5: In vitro Differentiation of Cortical Neurons.......................................109
9
Figure 3.6: Representative images of cells as they differentiate from ES cells into
n e u ro n s .............................................................................................................. 1 1 0
Figure 3.8: In vitro Neuronal Differentiation.......................................................112
Figure 3.9: Aag Expression Changes During In vitro Neuronal Differentiation.. 113
Figure 4.1: MMS Induced Cerebellar Degeneration in vivo is dependent on Aag
a nd P a rp 1 exp re ssio n ........................................................................................ 14 4
Figure 4.2: Isolation of Primary Cerebellar Granule Neurons (CGNs). ............. 145
Figure 4.3: Development of a High-Throughput Method to Determine Primary
Neuron Sensitivity to Drug Treatment. .............................................................. 146
Figure 4.4: Sensitivity to MMS treatment ex vivo is dependent on Aag. ........... 147
Figure 4.5: Aag is differentially expressed in primary CGNs from Aag', WT, and
m A a g Tg m ice . ................................................................................................... 14 8
Figure 4.6: Aag Glycosylase Activity is Significantly Different between Aag<, WT,
and m A ag Tg prim ary C G N s. ............................................................................. 149
Figure 4.7: Primary CGN Sensitivity to MMS is dependent on Parp1 activity. .. 150
Figure 4.8: PAR formation post-MMS treatment (1 mM) is dependent on the
expressio n leve l of A ag . .................................................................................... 15 1
Figure 4.9: Parp inhibitor Veliparib inhibits PAR formation after MMS treatment.
........................................................................................................................... 1 5 2
Figure 4.10: Neither supplementation of NAD' nor pyruvate rescues WT or
mAagTg primary CGN sensitivity to MMS.........................................................153
Figure 4.11: Downstream molecular mechanisms of CGN Aag-dependent MMS
s e n s itiv ity . .......................................................................................................... 1 5 4
Figure 4.12: There is no loss of mitochondrial permeability in Aagt, WT, or
mAagTg neurons 1-7 hours after MMS treatment. ............................................ 155
Figure 4.13: No evidence of AIF nuclear translocation after MMS treatment.... 156
Figure 4.14: Expression of BER genes in primary CGNs..................................157
10
Figure 4.15: Aag expression is not induced after MMS treatment in WT neurons.
........................................................................................................................... 1 5 8
Figure 4.16: Glycolysis and Tricarboxylic Acid (TCA) Cycle Schematic............159
Figure 4.17: Aag activity does not contribute to ex vivo sensitivity to glutamate
e x c ito to x ic ity . ..................................................................................................... 16 0
Figure 4.18: Aag does not contribute to ex vivo CGN sensitivity to treatment with
hyd rogen pe roxide (H 20 2). ................................................................................ 16 1
Figure 4.19: Female AlkbhT'CGNs are relatively resistant to MMS treatment ex
vivo compared to male AlkbhT', female WT and male WT neurons. ............... 162
Figure 4.20: WT and Alkbh T' CGNs are rescued from MMS sensitivity by Parp
inhibition but not by Caspase inhibition. ............................................................ 163
11
List of Tables
Table 1.1: Methylation patterns in single- and double-stranded DNA after
treatment with MMS represented as percentages of total methylation............36
Table 3.1: Attem pts at In vitro Cortical Neuronal...............................................115
12
CHAPTERS
13
Chapter I: Introduction
14
Table of Contents
C hapter 1: Introd uction ................................................................................... . . 16
In tro d u ctio n ................................................................................................ . . 16
Alkylating Agents in our Endogenous and Exogenous Environments Cause
Cytotoxic DNA Damage ............................................................................ 16
Methylating Agent Mechanism of Action .................................................. 17
DNA Repair Pathways for Alkylation Damage ......................................... 18
Alkyladenine Glycosylase (Aag) Initiates Repair of Methylated DNA Bases2l
Cellular Consequences of Imbalanced BER ........................................... 22
Alkylation Sensitivity of Cells with Altered Aag Expression...................... 23
Poly (ADP-Ribose) Polymerase 1 Can Mediate Cell Death Caused by BER
Im ba la nce s ............................................................................................ . . 2 5
Model System to Study Aag-Dependent Alkylating Sensitivity..................27
Overview of the Presented Study............................................................. 27
F ig u re s ....................................................................................................... . . 2 9
T a b le s ......................................................................................................... . . 3 6
R e fe re n ce s ................................................................................................. . . 3 7
15
Chapter I: Introduction
Introduction
Alkylating Agents in our Endogenous and Exogenous Environments Cause
Cytotoxic DNA Damage
The cells in our body are under constant attack from both endogenous and
exogenous sources. At the genomic level, cells can accumulate around 100,000
DNA lesions per day (Ciccia, 2010). Fortunately, our cells have multiple repair
mechanisms to restore DNA to its native sequence. Without repair, most of these
events have the capacity to result in a heritable mutation that can contribute to
cancer, degenerative conditions, and other maladies. Alkylating agents represent
a broad category of DNA damaging agents that contribute to both the
endogenous and exogenous sources of cellular damage.
Methylating agents are ubiquitous in our external and internal environments.
Endogenous sources of alkylation damage include lipid peroxidation by-products
and cellular cofactors. S-adenosyl-methionine is one potential source of aberrant
DNA methylation, inducing 3-methyladenine (3MeA) and 7-methylguanine
(7MeG) at rates of 600 and 4,000 lesions per cell per day, respectively (Fig. 1.1)
(Fu et al., 2012b; Lindahl, 2000). The nitrosation of amines on amino acids,
peptides, or polyamines is another potential source of methylating agents
(Sedgwick, 1997). Moreover, the observation that alkylation DNA repair enzymes
are conserved throughout evolution indicates that DNA alkylation must be
naturally and continuously occurring. Exogenous alkylating agents include methyl
halides derived from burning biomass and decaying vegetation, cigarette smoke,
food, and other occupational exposures (Hamilton et al., 2003). Finally, patients
are intentionally exposed to extremely high concentrations of alkylating agents
16
during chemotherapy for various cancers, including glioblastoma, lymphomas,
and leukemias (Fu et al., 2012b).
Methylating Agent Mechanism of Action
Methylating agents act by covalently adding methyl (-CH 3) groups to all
components of our cells, including proteins, lipids, and most importantly, our
DNA. Methylation on DNA bases occurs on both the ring nitrogens and exocyclic
oxygens. Methylating agents are categorized as either SN1 or SN2 agents based
on their mode of chemical reaction and this classification is indicative of the
proportion and type of DNA adducts they form. The predominant DNA adducts
formed by all alkylating agents are on ring nitrogens. Methylation at the highly
nucleophilic N7-position of guanine produces the most common methyl lesion,
7MeG (Table 1.1). 7MeG is neither mutagenic nor cytotoxic in itself; however,
methylation of purines at the N7 position causes base destabilization,
spontaneous depurination, and the formation of toxic and mutagenic abasic sites.
The other primary N-methylation product in double-stranded (ds) DNA is 3MeA.
Though 3MeA adducts are less common than 7MeG, they are toxic owing to their
ability to block DNA replication by high fidelity replicative polymerases
(Engelward et al., 1998; Groth et al., 2010; Johnson et al., 2007), although they
can be bypassed by certain translesion DNA polymerases (Glassner et al., 1998;
Johnson et al., 2007; Lange et al., 2011). When bypassed, 3MeA lesions are
potentially mutagenic and can lead to A:T to T:A transversions (Fronza and Gold,
2004). The rarer 1-methyladenine (1MeA) lesion can be formed preferentially in
single-stranded (ss)DNA and can block replication (Fu et al., 2012b). Methylation
can also occur on exocyclic oxygen atoms. In addition to N-alkylation, SN
agents methylate the 06-position of guanine to generate 06-methylguanine (06-
meG). Even though these lesions are relatively rare, they are potentially more
hazardous since O6meG can mispair with thymine during DNA replication leading
to downstream mutagenicity and cytotoxicity.
17
DNA Repair Pathways for Alkylation Damage
Fortunately, our cells have evolved multiple conserved DNA repair pathways
through evolution to combat the mutagenic and cytotoxic effects of methylated
bases. Below I discuss a few of the relevant pathways.
Base Excision Repair. The two most common methylation adducts, 7MeG and
3MeA, are repaired by the base excision repair (BER) pathway, a complex and
multi-enzyme process (Fig. 1.2). This pathway is initiated by lesion-specific DNA
glycosylases, which search the genome and recognize and excise damaged
bases. DNA glycosylases are classified as either bi- or mono-function based
upon their catalytic functions. Monofunctional glycosylases can remove damaged
bases while bifunctional glycosylases can both remove a damaged base and
cleave the DNA backbone. In mammalian cells, 11 different glycosylases have
been identified, often with overlapping substrates for repair redundancy (Jacobs
and Schar, 2012). However, there is only one glycosylase that acts on alkylated
bases, the monofunctional alkyladenine glycosylase (Aag; also known as Mpg).
Aag hydrolyzes the destabilized glycosyl bond between the DNA base and ribose
in the sugar-phosphate backbone to generate an abasic site (AP site). The Aag
glycosylase will be described in more detail below. In the next step of BER, the
abasic site is processed by the mammalian endonuclease Apel to form a single
strand break (SSB) with a 3'-OH and 5'-deoxyribosephosphate (5'-dRP). The
bifunctional DNA polymerase P (Pol P) then carries out two important functions.
First, Pol P removes the remaining sugar moiety left at the 5' end by APE1 with
its 5'-dRP lyase activity (Sobol et al., 2000). Secondly, Pol P fills in the missing
complementary nucleotides. BER is finalized by ligation of the single strand
break by a DNA ligase. X-ray cross-complementing protein 1 (Xrccl) acts as a
molecular scaffold during BER to recruit downstream enzymes and facilitate
processing of repair intermediates. It should be noted that the pathway described
above is the simplest form of BER and is termed short-patch (SP-) BER since
only one nucleotide is replaced. In the alternative pathway, long-patch (LP-)
18
BER, 2 to 12 nucleotides are replaced spanning the damaged DNA base (Kim
and Wilson, 2012). Replication is thought to be mediated by replicative DNA
polymerases Pol 6 and Pol E in combination with the accessory protein
proliferating cell nuclear antigen (PCNA). The displaced DNA is then cleaved by
flap endonuclease 1 (Feni) before the nick is sealed to complete repair (Fig.
1.2).
Nucleotide Excision Repair. Nucleotide excision repair (NER) is another multi-
enzyme and multi-step process generally reserved for the repair of bulky helix-
distorting DNA lesions. NER is initiated via two sub-pathways: global genome
repair (GGR) or transcription-coupled repair (TCR). The pathways vary in the
way they detect DNA damage but converge to the same repair mechanism. After
detection of a DNA lesion, NER proteins are recruited and the heterodimer
Erccl-Xpf makes an incision 5' to the lesion. DNA replication is then initiated at
the single-strand break thus displacing the lesion-containing DNA. Next, a
second incision is created 3' to the lesion by Xpg, releasing the damaged DNA.
Repair is finalized after all nucleotides are replaced and the DNA nick has been
ligated (Fig. 1.3CB) (Scharer, 2013). Interestingly, NER has been shown to
compensate in the repair of alkylated DNA in the absence of BER, primarily
through the global genome repair subpathway (Huang et al., 1994; Memisoglu
and Samson, 2000; Plosky et al., 2002; Samson et al., 1988).
Direct reversal proteins Mgmt and Alkbh2. The simplest repair strategy is to
directly remove the methyl group from the DNA, thus repairing the lesion without
affecting the overall DNA structure or sugar-phosphate backbone. Methylguanine
methyl transferase (Mgmt) is one such 'suicide' protein that repairs toxic 06-meG
lesions by transferring the methyl group to an internal cysteine residue, thus
targeting the protein for ubiquitin-dependent degradation (Fig. 1.3A) (Gerson,
2004; Liu et al., 2002; Pegg et al., 1991; Xu-Welliver and Pegg, 2002). Given the
highly cytotoxic and mutagenic nature of 06-meG lesions, Mgmt expression and
promoter methylation is highly correlated to cell sensitivity after treatment with
19
SN1 methylating agents (Brandes et al., 2008; Gerson, 2004; Hegi et al., 2004;
Maze et al., 1996). As mentioned above, 06-meG lesions readily mispair with
thymine during DNA replication, thus creating a DNA mismatch that is recognized
by the MutSa heterodimer, which initiates DNA mismatch repair (MMR). MMR
removes the newly synthesized DNA base (T); however, polymerases repeatedly
insert T opposite the 06-meG lesions, creating perpetual rounds of MMR and
generating single strand DNA breaks as MMR intermediates (futile cycling)
(Mojas et al., 2007); moreover, these single-strand DNA breaks can activate
downstream DNA damage signaling responses (Hickman and Samson, 1999b).
A second mechanism for direct DNA damage reversal is through the AlkB
homologue (Alkbh) family of proteins, some of which can repair 1MeA and 3-
methylcytosine (3MeC) in ss- and ds-DNA through an oxidative demethylation
mechanism that results in the release of formaldehyde as a byproduct (Fig. 1.3A)
(Begley and Samson, 2003; Falnes et al., 2002; Trewick et al., 2002).
Double Strand Break Repair. Double strand breaks (DSBs) can occur by a
variety of mechanisms. Though treatment with alkylating agents does not cause
DSBs directly, they can be generated by replication forks progressing past
single-strand breaks that can be created during BER (Ma et al., 2011; Pascucci
et al., 2005). Therefore, the ability of a cell to repair such DSBs is a determinate
of sensitivity to alkylation DNA damage (Kondo et al., 2009; Roos et al., 2009).
Two separate pathways can repair DSBs: error-prone non-homologous end
joining (NHEJ) and the more accurate homologous recombination (HR). NHEJ is
a relatively simple mechanism that ligates together double-stranded DNA ends
with little to no homology (Fig. 1.3D). HR, on the other hand, functions primarily
by using homologous DNA on sister chromatids as a template to resynthesize
any missing DNA across the break, and is therefore only functional during S or
G 2/M phases of the cell cycle (Fig. 1.3C). This cellular action results in the
appearance of sister chromatid exchanges (SCEs) which can be measured after
alkylation treatment and serve as a measurement for HR activity. Additionally,
some of the proteins involved in homologous recombination, most notably
20
Rad5l, have been shown to dually function in the regression of replication forks
as a means to bypass replication-blocking lesions (Adelman et al., 2013;
Hashimoto et al., 2010; Petermann et al., 2010; Schlacher et al., 2011; Shukla et
al., 2005; Zellweger et al., 2015). Upon encountering replication stress, the
parental DNA strand can re-anneal while the newly synthesized DNA strands
unwind and eventually anneal to each other, forming a 4-way Holliday junction.
The regression of the replication fork allows for DNA repair, replication past DNA-
lesions by using an undamaged template, or filling in of single-strand DNA gaps
(Neelsen and Lopes, 2015).
Alkyladenine Glycosylase (Aag) Initiates Repair of Methylated DNA Bases
Though Aag was first identified by its ability to remove alkylated DNA bases, it
has since been demonstrated to initiate repair of a structurally diverse set of DNA
lesions, including 3MeA, 7MeG, deaminated adenine (hypoxanthine), 1-
methyladenine (1 MeA), 1-methylguanine (1MeG), 3-methylcytosine (3MeC), 8-
oxoguanine (8-oxoG) and cyclic etheno adducts (EA; 1,2-EG) (Fig. 1.1) (Bessho
et al., 1993; Lee et al., 2009a; Shrivastav et al., 2010b; Wyatt et al., 1999). The
ability to effectively excise such a wide variety of DNA lesions while excluding
normal bases makes Aag unique. Aag has been shown to employ both hopping
and sliding mechanisms to move along double stranded DNA to search for rare
DNA lesions among other normal undamaged bases. (Hedglin and O'Brien,
2008; Hedglin and O'Brien, 2010). During this search, Aag adopts a low-affinity
conformation that is characterized by a disordered active site and non-specific
hydrogen bonding to DNA (Setser et al., 2012). Upon binding an appropriate
DNA lesion substrate, Aag assumes a higher-affinity conformation in which
residue Tyr-162 is intercalated into the double helix causing the DNA lesion to be
flipped into the glycosylase active site (Lau et al., 1998; Setser et al., 2012). A
water molecule is appropriately placed to be activated by Asp-238 and act as a
nucleophile to catalyze the hydrolysis of the N-glycosyl bond, causing the release
of the damaged base (Lau et al., 1998). The active site is lined with hydrophobic,
21
electron-rich aromatic amino acid residues that effectively bind and stabilize
positively charged alkylated bases (e.g. 3MeA, 7MeG), which additionally act as
good leaving groups during nucleophilic elimination (Lau et al., 2000). However,
not all of Aag's substrates are positively charged, such as the neutral lesions
hypoxanthine and EA (Fig. 1.1). It seems a second requirement for base
cleavage is dependent on the DNA lesion containing a hydrogen bond acceptor
for residue His-136 (Lau et al., 2000). Moreover, release of neutral bases
requires both a base to activate the water molecular and an acid to protonate the
leaving group, while positively charged lesions only need a base for cleavage
initiation (O'Brien and Ellenberger, 2003).
Cellular Consequences of Imbalanced BER
BER requires the tight coordination of multiple enzymes since the pathway
intermediates have been shown to by cytotoxic if allowed to accumulate. AP sites
and SSBs both inhibit transcription and replication machinery and can lead to the
generation of DSBs (Fig. 1.5B) (Boiteux and Guillet, 2004). Though translesion
polymerases exist to replicate past AP sites and avoid cytotoxic DSBs, this
process often produces point mutations (Avkin et al., 2002; Pages et al., 2008;
Schaaper et al., 1983; Weerasooriya et al., 2014). SSBs are rendered even more
toxic during BER if the 5'-dRP termini is not cleared by the lyase activity of Pol B.
Polfl' cells are hypersensitive to methylating agents, however reintroduction of
the Pol P lyase domain, without an active polymerase domain, rescues sensitivity
to near WT levels (Sobol et al., 2000). Moreover, Polf3' cells are only methylation
sensitive if BER is initiated by Aag; Polf5'Aag-' cells exhibit the same sensitivity
to MMS as WT cells (Sobol et al., 2003).
To help shuttle intermediates through repair to completion, Xrccl acts as a
molecular scaffold to recruit and trigger the activity of BER proteins to avoid
cellular accumulation of abasic sites or single-strand breaks. Xrccl is an
essential protein has been shown to interact with glycosylases (including Aag)
22
(Campalans et al., 2005; Mutamba et al., 2011), Apel (Vidal et al., 2001), Pol P
(Caldecott et al., 1996; Kubota et al., 1996), poly (ADP-ribose) polymerase
(Parp1) (Caldecott et al., 1996; Masson et al., 1998), and DNA ligase III (Lig Ill)
(Caldecott et al., 1994; Tebbs et al., 1999). The presence of XRCC1 in human
cells was shown to facilitate AAG-dependent excision of hypoxanthine and 1,N -
ethenoadenine (sA) (Fig. 1.1) (Mutamba et al., 2011). Moreover, Xrccl competes
with Apel and Parp1 to bind DNA repair intermediates themselves (Nazarkina et
al., 2007). Knockout of Xrccl is embryonic lethal and knockdown of the protein
leads to enhanced sensitivity to oxidative and alkylation treatment (Caldecott,
2003).
Even in the presence of Xrccl, BER pathway imbalances can occur due to
increased pathway initiation via glycosylase activity or decreased activity of any
downstream step. Decreases in Apel activity cause increases in AP sites and
sensitivity to alkylation treatment (Ensminger et al., 2014; Silber et al., 2002). As
mentioned above, downregulation of Pol P activity causes increases in SSBs,
double-strand breaks, and sensitivity to methylating agents (Senejani et al.,
2012; Sobol et al., 2000). Below, we will discuss in more detail the effects that
altered Aag expression has on cell sensitivity to alkylating agents.
Alkylation Sensitivity of Cells with Altered Aag Expression
Given all that is known about BER and cellular responses, it is conceivable that
Aag-' cells could react in disparate ways to alkylating agents. Since BER is
initiated by the adduct-specific Aag DNA glycosylase, cells lacking Aag
accumulate cytotoxic and mutagenic DNA methylation lesions 3MeA and 7MeG.
Though 3MeA has been shown to block DNA polymerases, potentially leading to
double strand breaks at collapsed replication forks that can be repaired by
homologous recombination (Hendricks et al., 2002), translesion polymerases can
bypass these lesions during DNA synthesis (Johnson et al., 2007; Lange et al.,
2011). On the other hand, the loss of Aag could mask BER imbalances resulting
23
in accumulation of toxic intermediates that typically signal for cell death in wild-
type (WT) cells.
In fact, there is a range of responses to alkylation damage in cells lacking the
Aag DNA glycosylase. Aag' mouse embryonic stem ES cells are more sensitive
to the SN2 alkylating agent methyl methanesulfonate and exhibit more alkylation
induced sister chromatid exchanges, p53 stabilization, and apoptosis compared
to wild-type ES cells (Engelward et al., 1998; Engelward et al., 1996). Primary
mouse embryonic fibroblasts (MEFs) from Aag' mice are more sensitive than
wild-type MEFs to Me-Lex, an agent that specifically induces 3MeA lesions, but
exhibit no difference in sensitivity to MMS (Engelward et al., 1997; Sobol et al.,
2003). Similarly, human cervical carcinoma (HeLa) cells and glioma cell lines
with significantly reduced Aag protein expression are sensitized to
chemotherapeutic methylating agents (Agnihotri et al., 2011; Paik, 2005).
Conversely, the increased sensitivity of Aag' ES cells does not extend to the
adult Aag' mouse. Aag' mice are not more sensitive to alkylation-induced
lethality compared to wild type nor do they exhibit significantly more spontaneous
or alkylation induced cancers (Roth and Samson, 2002) (unpublished data).
Surprisingly, the loss of Aag actually renders particular adult mouse tissues
resistant to MMS induced cell death. Aag' myeloid progenitor cells of the
hematopoietic lineage are resistant to MMS compared to WT progenitors in ex
vivo colony-forming assays (Meira et al., 2009a; Roth and Samson, 2002). Aag'
mice also exhibit hematopoietic resistance to MMS in vivo as measured by total
bone marrow cell numbers and the micronucleus assay, a measure for large-
scale erythropoietic chromosomal damage. A similar phenotype was observed in
Aag' retinal photoreceptors, which demonstrated remarkable resistance to
MMS-induced degeneration and blindness (Meira et al., 2009b). Finally,
treatment of WT mice with methylating agents induces neurodegeneration of
cerebellar granule neurons, yet this cell death is completely abolished in Aag-
mice (Calvo et al., 2013b; Kisby et al., 2009).
24
These results suggest that initiation of BER in certain WT tissues causes cell
death that is suppressed upon loss of Aag. Indeed, overexpression of Aag
renders these tissues even more sensitive to MMS induced degeneration by
creating a more severe imbalance in the BER pathway due to increased pathway
initiation without concomitant increases in downstream enzymes (Calvo et al.,
2013b). The Aag dependent sensitivity of hematopoietic progenitors, retinal
photoreceptors, and cerebellar granule neurons is similarly completely
suppressed in the absence of Parp1, independent of the level of Aag expressed,
indicating that Parpi mediates cell death caused by BER imbalances in these
tissues (Calvo et al., 2013b). All in all, these results support the notion that cell
fate in response to methylating agents is both Aag-dependent and cell-specific.
Poly (ADP-Ribose) Polymerase I Can Mediate Cell Death Caused by BER
Imbalances
Parp1 was first identified as a DNA repair protein with the ability to catalyze the
formation of long poly (ADP-ribose) (PAR) polymers on itself and other proteins
(Satoh and Lindahl, 1992). Parpl strongly binds single-strand breaks within
minutes of their generation though two zinc-fingers and one zinc-binding domain
that mediate DNA-dependent enzymatic activation (Langelier et al., 2008). PAR
polymers are created through the hydrolysis of NAD' and transfer of the ADP-
ribose moiety to protein acceptors (Fig. 1.4). The polymers are highly negatively
charged and function both as posttranslation modifications as well as signaling
molecules on their own right.
Upon binding to SSBs, Parp1 undergoes auto-PARylation that subsequently
stimulates its release from DNA due to negatively charged interactions between
the DNA and the PAR polymers. PARylation also occurs on multiple DNA
damage repair and signaling proteins including Aag, Xrccl and the ataxia
telangiectasia and Rad3 related kinase (ATR) (Jungmichel et al., 2013; Kedar et
25
al., 2008; Masson et al., 1998); however, the functional role of PARylation of Aag
has not yet been explored. Though Parp1 is not required for accurate completion
of BER, activation of Parp1 helps recruit Xrccl and can stimulate DNA repair
(Dantzer et al., 1999; El-Khamisy et al., 2003; Prasad et al., 2015; Prasad et al.,
2001). It should be noted that Parp1's cellular function extends beyond DNA
repair as it has been found to play a role in transcription, metabolism,
inflammation, cell differentiation, and regulation of chromatin structure (Bai and
Canto, 2012; Ditsworth et al., 2007; Ji and Tulin, 2010; Krishnakumar and Kraus,
2010; Schreiber et al., 2006).
While at moderate levels of DNA damage Parp1 activation can assist in DNA
repair and cell survival, upon excessive levels of DNA damage and formation of
SSBs Parp1 hyperactivation can cause cell death through caspase-independent
programmed necrosis (Berghe et al., 2014). Historically, Parp1 mediated cell
death has been hypothesized to be caused by cellular depletion of NAD+ through
PARylation and subsequent ATP loss leading to bioenergetic failure (Fig. 1.5B).
Indeed, in certain cases repletion of bioenergetic substrates such as NAD+ and
pyruvate does rescue cell sensitivity to alkylating agents (Alano et al., 2010;
Tang et al., 2010; Tang et a!., 2009). However, recent publications have
challenged this hypothesis by demonstrating that ATP depletion precedes NAD+
loss. In fact, PAR polymers were shown to translocate from the nucleus to the
mitochondria and bind directly to hexokinase (HK), the initiating enzyme of
glycolysis, thus inhibiting its activity (Andrabi et al., 2014; Fouquerel et al., 2014).
This glycolytic inhibition was independent of loss of NAD+ and glycolysis could be
restored through the addition of tricarboxylic acid (TCA) cycle substrates
pyruvate and glutamine (Andrabi et al., 2014). Furthermore, PAR translocation to
the mitochondria has been shown to cause the mitochondrial release of
apoptosis-inducing factor (AIF) (Hong and Dawson, 2004; Yu et al., 2006). AIF
translocates to the nucleus where it interacts with histone H2AX, leading to
chromatinolysis through interaction with cyclophilin A (Artus et al., 2010). It
26
should be noted, however, that AIF translocation is not a necessary component
of Parp1-mediated programmed necrosis (Tang et al., 2010).
Model System to Study Aag-Dependent Alkylating Sensitivity
Despite the known and obvious cell-specific differences in Aag-dependent
sensitivity to methylating agents, no attempts to understand the underlying
biology have been made. One option is to study pure populations of cells
exhibiting varied Aag-dependent alkylation responses, for example, mouse
embryonic stem (ES) cell lines and primary neurons. Another possibility,
however, is to utilize the pluripotency of one of the cell types of interest - mouse
ES cells. ES cells have the capacity to differentiate in vitro into any cell type of
interest. Moreover, if in vitro differentiation recapitulates in vivo development then
Aag* MMS-sensitive ES cells will transform to become Aag4 MMS-resistant
hematopoietic progenitors and cerebellar neurons. Benefits of this method
include a shorter timeline of differentiation in vitro compared to in vivo, abrogating
the need for maintaining mouse colonies, and offering the option of assessing
Aag' sensitivity as a function of development. Limitations of this method include
heterogenous populations of cells after differentiation and the inability to know
forthright whether in vitro differentiation will reflect in vivo work.
Overview of the Presented Study
In the presented work, we explore the Aag dependent DNA-damage responses
to methylating agents in embryonic stem cells compared to primary neurons. To
this end, new ES cells have been generated and validated for pluripotency. We
show that Aag' sensitivity to MMS compared to WT is independent of inbred
genetic background. Surprisingly, we find that overexpression of Aag in ES cells
renders them even more sensitive to MMS than loss of Aag, indicating that
methylation sensitivity in ES cells is not Aag gene dose dependent as seen in
27
primary neurons or retinal photoreceptors. Additionally, we studied in detail the
Aag- and Parp1-dependent cell sensitivity of mouse cerebellar granule neurons
in ex vivo conditions, demonstrating that methylation responses are cell-intrinsic
phenomena that can be studied outside of brain tissue. Neurons undergo
caspase-independent cell death after MMS treatment and do not exhibit
mitochondrial permeabilization or AIF nuclear translocation. Finally, we employed
in vitro differentiation of hematopoietic progenitors and cerebellar neurons in an
attempt to recapitulate the transition in Aag-' MMS-responses from sensitivity in
ES cells to resistance in differentiated cells.
28
0(D N
</ NH
NN N NH 2
7meG
N
NH
N
Hypoxanth ine
0
H
N
NNH
N NH 2
8-oxoG
NH 2
N
NN
3meA
0
N
NN NH-2
ImeG
N
N N
1,N6-E A
NH 2
N
NN NN
lmeA
NH
N
3meC
0
N
/ 
N
N N
1,N2-EG
Figure 1.1: Structures of Aag DNA Substrates.
7MeG, 7-methylguanine; 3MeA, 3-methyladenine; 1MeA, 1-methyladenine;
1MeG, 1-methylguanine; 3MeC, 3-methylcytosine; 8-oxoG, 8-oxoguanine; 1,N -
EA, 1,N6-ethenoadenine; 1,N2-G, 1,N2-ethenoguanine.
I'I
29
Figures
DNA Base Damage
4
Mono-functional
UNG) YI ISMLJ61
MBD4) AAGIMPGj
3'
A G T T C A
T A C A A G T
3 I 1 , 1 5
T--- 3
A G T
5 - -T 3'
A'7 G T T C A
T A C A A G T
yj __
T A AG G
3 5
A G T T C A
T A C A A G TL G la/XRCC1
5 -111 3
SP-BER
Bi-functional (P)
(OGG1 NTH1
Bi-functional (0,6)
riNE1L I 2
1 1 1 1 35'--T~~
A G T T C A A G T T C A
T A C A A G T T A 
C A A GT
3 5 3' 5
APE1' PNKP
5' - -T 1 3
5 A C T T C A
T A C A A G T
3~~~~ 1~..L A . I.. 5'
PCNA
Po6
PoIE
C
5 -- T - 3
33A ' ~ ' C A
T A C A A G T
3' 1 1 1 5'
FENI
A G C A
TA C A 
AG T
LIGI
---- ---, -r---'r 3'
A ; ' C A
T A C A A GI
LP-BER
Figure 1.2: Base Excision Repair Pathway.
Base excision repair (BER) pathway is initiated by the mono-functional Aag
glycosylase that remove a methylated base, leaving an abasic site. The abasic
site is cleaved by AP endonuclease 1 (Apel) resulting in a 5'-
deoxyribosephosphase (5'-dRP) that is cleared by the lyase activity of Pol 1,
which also synthesizes any missing DNA nucleotides. Finally the single-strand
break is sealed together by a ligase (Lig). The whole process is coordinated by
Xrccl. LP-BER is initiated by bi-functional DNA glycosylases that remove a
damaged base and also cleave the DNA backbone. Inhibitory DNA termini are
removed by either Apel or Pnkp and DNA replication is completed by either Pol
P, Pol 6, or Pol E. The displaced DNA is removed by the flap endonuclease 1
(Fen1) and the nick is sealed. Figure adapted from (Kim and Wilson, 2012).
30
A Direct repair (MGMT)
CH1
MGMT .
CysCH,
MGMT
- Cys-CH,
IIIIIIIIII
C
Direct repair (ALKBH) B NER
CH XPC-RAD23B
mifhrm17l
ALKBH OH
H ,
LK -HCOH
0
HR
XPA
RPA
5 fnc sion 3' incsion
XPA TFIH
ERCCXG
XPE
Repair synthesis
Pol 6.
or L
D=I|
PCNA
Ligation
niwiQItTIi
NHEJ
DNA-PK binding
and synapsis
Processing by Artemis,
PNK and other proteins
Recruitment of LIG4-
XRCC4 and rejoining
31
Figure 1.3: Alternate DNA Repair Pathways for Methylation Damage.
(A) Direct Reversal repair mechanisms include MGMT and the AlkBH
proteins. MGMT directly removes methyl groups from the 06-position of
guanine to an internal cysteine residue. AlkBH proteins remove methyl
lesions through an oxidative demethylation mechanism that results in the
release of formaldehyde.
(B) Nucleotide excision repair (NER) pathway identifies DNA damage through
global genome repair subpathway. Single-strand nicks are created 5' and
3' of the damage so that a fragment of lesion-containing DNA can be
removed. Polymerases (Pol 6 or Pol E) fill in the empty segment before
ligation. TFIIH, Transcription factor IIH; XP, xerodera pigmentosum
proteins.
(C)A replication fork passing a single-strand DNA break will generated a one
ended double-strand break. The 3' overhang of the double-strand break
will invade the complementary sequence on the sister chromatid to
generate a D-loop and DNA is synthesized past the single-strand break.
The resulting Holliday junction will be cleaved, thus restoring the
replication forks.
( n- g IIU JUIIIIId jIn  I I J) Is Iniliated by the binding of Ku
proteins (green circles) to double-stranded DNA ends, recruitment of
DNA-PKcs (taupe cylinder), processing of the ends by Artemis (red
triangle), and eventual ligation by Lig4-XRCC4 complex (orange ovals).
DNA-PKcs, DNA-dependent protein kinase catalytic subunit; Lig4, DNA
ligase 4; XRCC4, X-ray repair cross-complementing protein 4.
Figures adapted from (Fu et al., 2012b; O'Driscoll and Jeggo, 2006, Helleday, et
al., 2007)
32
0 0
NHH
N H N A D 
NH
H NAD Nicotinamide
OHN O
Parp1l
Acceptor Acceptor
Protein Protein
SE. D, or K
NHC=O
<N N IUHydrohise
W 
NH
O OH
NN
0 OH
NHARG AR
NHO N OHOHI
OOH
OH OH
Figure 1.4: Parp1 utilizes NAD* to generate PAR polymers.
Parp1 catalyzes the addition of the ADP-ribose portion of NAD* to an acceptor
protein, releasing nicotinamine in the process. Long polymers of ADP-ribose are
called poly (ADP-ribose) or PAR polymers. Polymers can be either branched or
linear and are primarily hydrolyzed by poly (ADP-ribose) glycohydrolase (PARG).
Figure adapted from (Krishnakumar and Kraus, 2010)
33
Alkylating Agent
AAG
APE
- ~ OH
jG
Alkylating Agent
.3'OH
.3'OH
( LID
DNA Survival
replication Mutationblock
DNA replication DSB
and fork collapse
Survival
Mutation
NHEJ HR
Survival
SCE
TLS Surviva
Mutation
Parpi Parp1
Activation Hyperactivation
DNA NAD'/ATP
Repair depletion
4.riv C 4lDe
Figure 1.5: Alternative mechanisms of Cell Death Caused during Base Excision
Repair (BER).
Alkylating agents cause the formation of DNA damage that is repair by the BER
pathway (described in the text). DNA lesions and abasic sites (AP sites) inhibit
34
__________ _____________I
A
%A
0
1A
E
4,
12
Apoptosis
Necrosis
Cell
death
DNA replication but can be bypassed by translesion synthesis (TLS), resulting in
survival albeit with mutations. Single-strand breaks, however, can cause death
through two different mechanisms. (A) SSBs strong inhibit replication fork
progress leading to cytotoxic double-strand breaks which can be repaired by the
error-prone NHEJ or more accurate HR. DSBs can also cause cell death, often
through apoptotic mechanisms. (B) Alternatively, SSBs can be bound by Parp1,
leading to activation. At low levels of DNA damage, Parp activation stimulates
DNA repair; however, Parp hyperactivation can cause cell death through
energetic failure and programmed necrosis. Figures adapted from (Calvo et al.,
2013b; Fu et al., 2012b).
35
Tables
Table 1.1: Methylation patterns in single- and double-stranded DNA after
treatment with MMS represented as percentages of total methylation.
nd = not detected. Data adapted from (Beranek, 1990; Drablos et al., 2004).
Site of Methylation ssDNA/RNA
Adenine
N1- 18 3.8
N3- 1.4 10.4
N7- 3.8 1.8
Guanine
N3- -1 0.6
o6- 
- 0.3
N7- 68 83
Cytosine
2
-
N3-
nd
10
Diester 2
nd
<1
0.8
36
dsDNA
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48
Chapter II: Generation of WT, Aag' and
mAagTg C57B1/6 and 129 Embryonic Stem
Cells and Sensitivity to Alkylating Agents
Carrie M. Margulies, Jennifer J. Jordan, Isaac Alexander Chaim,
Jennifer A. Calvo, Leona D. Samson
Contributions: C.M.M. and L.D.S. designed experiments. C.M.M. conducted all
experiments unless otherwise noted. J.J.J. measured BCNU and MNU sensitivity
of C57B11/6 ES cells and helped with chromosome spreads and quantification.
J.J.J. also did the 129 ex vivo hematopoietic colony forming assays. I.A.C.
measured in vivo Aag activity by host cell reactivation assays. J.A.C. assayed in
vivo sensitivity of 129 cerebellar granule neurons and retinal photoreceptors.
49
Chapter 11 Table of Contents
Chapter II: Generation of WT, Aag' and mAagTg C57B11/6 and 129 Embryonic
Stem Cells and Sensitivity to Alkylating Agents .................................................. 51
Introduction .................................................................................................. 51
M aterials and M ethods................................................................................. 54
Results......................................................................................................... 62
Generation of C57B1/6 and 129 Mouse Embryonic Stem Cells and Validation
of Pluripotency.......................................................................................... 62
BER Gene Expression and Aag Activity in WVT, Aag- and mAagTg ES Cells
..................................................................................................................... 6 3
Sensitivity of ES Cell Lines to Alkylating Agents....................................... 64
Discussion..................................................................................................... 65
Figures......................................................................................................... 69
References................................................................................................... 78
50
Chapter II: Generation of WT, Aag~' and mAagTg C57B11/6 and 129
Embryonic Stem Cells and Sensitivity to Alkylating Agents
Introduction
Embryonic stem (ES) cells are derived from the inner cell mass of a blastocyst,
making them pluripotent cells that have the capacity to generate all three germ
layers in vivo or in vitro (endoderm, ectoderm, and mesoderm). Because ES cells
grow rapidly and differentiate to produce all cells of a complete organism, it is
postulated that they must possess robust and accurate mechanisms to repair
DNA damage since any deleterious mutation incurred early in development that
does not cause cell death will be passed on to all daughter cells. In practice, ES
cells exhibit -100-fold lower spontaneous mutation rate compared to somatic
cells (Cervantes et al., 2002; Sugiura et al., 2014). Moreover, they have higher
antioxidant levels and enhanced stress defenses compared to differentiated
cells, thus reducing endogenous sources of DNA damage, such as reactive
oxygen species (ROS) (Armstrong et al., 2010; Saretzki et al., 2004; Saretzki et
al., 2008).
Alkylating agents represent another constant threat to our DNA from both
endogenous and exogenous sources (Fu et al., 2012b; Hamilton et al., 2003;
Lindahl, 2000; Sedgwick, 1997). If left unrepaired, alkylated DNA can cause
mutations, cancer, degenerative diseases, or teratogenesis in the case of a
growing embryo. The two most commonly formed methylation adducts, N7-
methylguanine (7MeG) and N3-methyladenine (3MeA), are repaired by the base
excision repair (BER) pathway, a coordinated multi-enzyme process. The
pathway is initiated by the monofunctional lesion-specific alkyladenine
glycosylase, Aag, which hydrolyzes the destabilized glycosyl bond between the
damaged DNA base and the ribose in the sugar-phosphate backbone,
generating an abasic site. The abasic site is cleaved by the apurinic/apyrimidinic
endonuclease 1 (Apel) to form a single strand break with a 3'-OH and 5'-
51
deoxyribosephosphate (5'-dRP). Bifunctional DNA polymerase P (Pol P) then
carries out two important functions. First, Pol P removes the remaining sugar
moiety left by Apel with its 5'-dRP lyase activity in a rate-limiting step and
secondly, it fills in any missing DNA nucleotides. BER is finalized by ligation of
the single strand break by DNA ligase (Lig). Since the repair process depends
upon the activity of multiple enzymes functioning in a specific sequence, Xrccl
acts as a scaffold to recruit, interact, and facilitate processing of repair
intermediates.
ES cells express high levels of many BER proteins compared to differentiated
mouse embryonic fibroblasts (MEFs), including Apel, DNA ligase 111, Xrccl and
the supporting protein poly-(ADP-ribose) polymerase (Parp1). These increases at
the protein level yield enhanced in vitro BER repair capacity, agreeing with the
theory that ES cells have heightened DNA repair and damage monitoring
mechanisms to avoid the induction of mutations (Tichy et al., 2011). Additionally,
mouse ES cells express high levels of p53 mRNA and protein, which is
subsequently lost during cellular differentiation and may play a role in modulating
cell-specific variation in alkylation sensitivity (Solozobova and Solozobova,
2010). Furthermre, p53 has been shown to induce ES cell ddirentiation after
DNA damage through suppression of the pluripotency regulating transcription
factor Nanog (Abdelalim and Tooyama, 2014; Lin et al., 2004).
ES cells lacking the Aag glycosylase on a 129 inbred genetic background are
sensitive to the monofunctional Sn2 methylating agent methyl methanesulfonate
(MMS), and exhibit increased sister chromatid exchanges, p53 induction, and
apoptosis after treatment compared to wild-type ES cells (Engelward et al., 1998;
Engelward et al., 1996). Interestingly, Aag' ES cells are also sensitive to the
bifunctional interstrand crosslinking (ICL) agents psoralen, 1,3-bis(2-chloroethyl)-
1-nitrosourea (BCNU), and mitomycin C (MMC); in the case of psoralen, Aag-
cells are not sensitive to their corresponding monofunctional equivalent,
implicating a still undefined role for Aag in the repair of DNA interstrand
52
crosslinks (ICLs) (Allan et al., 1998; Engelward et al., 1996; Maor-Shoshani et
al., 2008).
In this study, we generated new WT and Aag' C57B11/6 ES cells and
characterized their response to alkylating agents to determine if, and how,
genetic background affects cell sensitivity. We found that Aag dependent cell
sensitivity to alkylation treatment is independent of genetic background (129 vs.
C57B11/6), with the exception of BCNU. Moreover, we also generated new WT,
Aag- and transgenic Aag overexpressing (mAagTg) 129 ES cells. Surprisingly,
we find that both loss and overexpression of Aag render ES cells sensitive to
alkylating agents compared to WT. These differences in sensitivity can be
attributed solely to variations in Aag expression and activity since all downstream
BER proteins were expressed at similar levels between the genotypes.
53
Materials and Methods
Materials and Antibodies
We used the following materials in this study: leukemia inhibitory factor (LIF;
Millipore), MMS (Sigma, 129925), BCNU (Sigma, C0400), Mitomycin C (Sigma,
M4287), and N-methyl-N'-nitro-nitrosoguanidine (MNNG; Sigma).
The following primary antibodies were used for immunocytochemical staining and
western blots: Sox2 (AbCam, ab97959, 1:500), Nanog (AbCam, ab80892,
1:200), Sseal (Cell Signaling Technologies, 4744, 1:100), Apel (AbCam, ab82,
1:1000), Pol B (AbCam, ab26343, 1:700), Xrccl (AbCam, ab9147, 1:1000),
Parp1 (AbCam, ab6079, 1:400), and Tubulin (AbCam, ab6160, 1:5000). H2AX,
p43
Generation of ES Cells
Mice were mated and 3.5 days post-coitus (dpc) female mice were brought to the
ES Cell and Transgenics Core Facility at the Koch Institute for Integrative Cancer
Research where blastocysts were harvested according to previously published
methods (Markoulaki et al., 2008). ES cells were derived in media containing
knock-out serum replacement (KSOR) and a Mek1 inhibitor (15% KOSR
(Invitrogen), LIF, 50 pM Mek1 inhibitor (PD98059, Cell Signaling Technologies),
1% NEAA, 1% L-glut, 1% penicillin/streptomycin in DMEM) and later converted to
growth on complete ES media (see below).
Cell Culture
MEFs were grown in DMEM plus 10% heat-inactivated fetal bovine serum (HI-
FBS) (Sigma), 1% L-glutamine and 1% penicillin/streptomycin. To generate
feeder layers, primary MEFs were irradiated with 45 Gy with a Cobalt-60 source
and frozen at 1 x 10 7 cells/vial. ES cells were grown in DMEM plus 15% HI-FBS
(Sigma), 1% L-glutamine, 1% non-essential amino acids, 1% sodium pyruvate,
100 pM 3-mercaptoethanol (Sigma, M3148), 1% penicillin/streptomycin and
54
recombinant mouse leukemia inhibitory factor (LIF). All components were
purchased from Invitrogen unless noted. ES cells were grown either on an
irradiated MEF feeder layer or on tissue culture plates coated with 0.1% gelatin.
Karyotyping
ES cells were grown on irradiated MEFs in the presence of colcemid (0.01
ug/mL; Sigma, D1925) for 3 hours prior to trypsinization to arrest cells in mitosis.
Cells were harvested by trypsinization and resuspended dropwise in hypotonic
lysis buffer (5% FBS, 0.2 mg/mL sodium citrate, 0.2 mg/mL potassium chloride)
while simultaneously flicking the tube to ensure single cells. After a 15 minute
incubation at 370 C and centrifugation for 5 minutes at 100g, cells were fixed
three times in Carnoy's fixative (3:1 ratio of methanol to acetic acid, following the
same protocol as above for the hypotonic solution. Finally, cells were dropped
from 12-18 inches onto glass microscope slides that were pre-labeled and dipped
in water. Slides were dried overnight in the dark and stained with 5% Giemsa
(Sigma, GS500) in Sorensen's buffer (pH 6.8) for 8-10 minutes, washed gently
with water and allowed to dry completely. Slides were then mounted with
coverslips and imaged on a Nikon Eclipse E800 microscope with a Qlmaging
Retiga EXi Fast1394 camera at 10OX magnification. Professional G-band
karyotypic analyses were conducted by Cell Line Genetics (Madison, WI).
Colony Forming Assays
ES cells were seeded at increasing densities in 12 well plates coated with
irradiated MEFs in duplicate (200 - 200,000 cells/well). 14 hours later, cells were
treated for 1 hour at 370 C with varying concentrations of drug in serum-free ES
media. For MNNG, drug was added in complete ES media and left for the
duration of the experiment. After 6-8 days of growth, plates were washed twice
with PBS and dried overnight in a 370 C incubator and stained with 1/20 dilution
of Giemsa in PBS for 2-3 hours. Colonies with >50 cells were counted using
ImageJ (NIH) and percent survival was calculated based on the number of
colonies formed in untreated wells and the number of single cells plated.
55
Host Cell Reactivation Assay
The host cell reactivation is based upon the production of fluorescence from an
exogenous plasmid with a site-specific lesion. In the case of Aag, plasmids were
engineered to contain a hypoxanthine (Hx) lesion in the fluorophore codon. In the
absence of Aag, Hx exhibits transcriptional mutagenesis and miscodes for a C,
leading to a fluorescent protein. If Aag repairs Hx, the appropriate U is
incorporated and no fluorescent in exhibited. For the Apel assay, a
tetrahydrofuran (THF) was incorporated in the plasmid. THF blocks transcription
and mimics an abasic site; thus fluorescence would only be exhibited if cells
contained function Ape 1.
For engineering substrates reporting via transcriptional mutagenesis, non-
fluorescent variants of the different reporter plasmids containing a single
mutation in a site coding for their respective chromophores were identified. In
order to produce ssDNA, we followed previously described methods with minor
modifications (Nagel et al., 2014b). Reporter plasmids were nicked with either
Nb.Bstl or Nt.Bstl (New England Biolabs, depending on the lesion containing
strand, see table X). The nicked strand then was digested with exonuclease Ill,
and the remaining single-stranded circular DNA (ssDNA) was purified by using a
1% agarose gel. 15 picomoles of the respective phosphorylated lesion-
containing-oligonucleotide were combined with 3.2 pg of the corresponding
single-stranded plasmid in 1X Pfu polymerase AD buffer (Agilent
Biotechnologies) in a final volume of 46 pL. The mixture was heated to 85 'C in a
thermal cycler for 6 min, and then allowed to anneal by cooling to 40 *C at 1 0C
per minute. To extend the primer, 5 units of Pfu polymerase AD (Agilent) and 0.4
pM dNTP were added. The reaction is then cleaned up with a Qiagen PCR
Purification kit column and eluted in EB buffer and subsequently combined with
1X Ligase Buffer (New England Biolabs - NEB), 0.4 pM dNTP, 1 pM ATP, 1X
BSA, 1.5 units T4 DNA Polymerase and 80 units T4 DNA Ligase (NEB) and
56
incubated for an additional hour at 16 0C to yield closed circular plasmid. Finally,
the product was purified from a 1 % agarose gel using a Qiagen gel extraction kit.
To test for the presence of Hypoxanthine in GFP-C289T-Hx and its negative
controls (GFP-WT and GFP-C289T), 150 ng of the plasmids were incubated with
10 units ApaLl (NEB) in 1X Cutsmart buffer for 1 hour at 37 *C, followed by 20
min at 65 0C for heat-inactivation. Products were run in a 1% agarose gel for
visualization.
To test for the presence of THF in GFP-617-THF and mOrange-A215C-THF and
their negative controls (GFP-WT and mOrange-WT, respectively), 150 ng of the
plasmids were incubated with 10 units APE1 (NEB) in 1X #4 buffer for 1 hour at
37 0C, followed by 20 min at 65 0C for heat-inactivation. Products were run on a
1 % agarose gel for visualization.
ES cells grown on gelatin in a 12-well plate gelatin and were transfected with
Lipofectamine LTX (Invitrogen) according to manufacturer's instructions. Briefly
2.5 pg of total plasmid DNA were mixed with 2.5 pL Plus reagent and Opti-MEM,
further mixed with 6.2 pL lipofectamine LTX in Opti-MEM and incubated at room
temperature for 5 min before adding 200pL of the transfection reaction on top of
the cells. Transfected cells were incubated for 18 hours at 37 0C and 5% CO2.
Following incubation cells were trypsinized and resuspended in a total of 500 pL
of complete media containing TO-PRO-3 and transferred to a FACS tube for
analysis.
Cells suspended in culture media were analyzed for fluorescence on a BD LSR 11
cytometer running FACSDiva software. Cell debris, doublets, and aggregates
were excluded based on their side- and forward-scatter properties.
Every experimental setup consisted of two sets of transfections: A control
transfection (CT) and a sample transfection (ST) containing the one or more
reporters with DNA lesions. Both transfections included the same color
combination with the same undamaged reporter to normalize each set for
57
transfection efficiency.
Fluorescence Index (FI) for a given reporter within one transfection was
calculated using Eq. X:
FI = CF x MFI
CL
where CF iS the number of positive fluorescent cells for that given fluorophore,
MFI is the mean fluorescence intensity of the CF, and CL is the total number of
live cells.
The normalized fluorescence index for a given reporter Flo was calculated using
Eq. X:
FIE
where Fl" corresponds to the F1 of a reporter normalized to the F! of the
transfection efficiency normalization plasmid, FIE.
Normalized reporter expression from a sample transfection, FOST, and that from
the same reporter plasmid in control transfection, F/0CT, were used to compute
the percent reporter expression (%R.E.) using Eq. X:
FI0 sT% R. E. = FIOS X 100
FIOCT
Immunofluorescence
ES cells were seeded on MEFs in 24-well glass bottom plates (Invitrosci) and
fixed in 4% paraformaldehyde (Electron Microscopy Sciences #15700) for 15
58
minutes. Cells were permeabilized with 0.1% Triton-X in CSK buffer (100 mM
NaCl, 300 mM sucrose, 10 mM PIPES, 3 mM MgC 2 , 1 mM EGTA) for 20
minutes, blocked with 4% bovine serum albumin (BSA), and stained overnight at
4' C with appropriate dilutions of antibodies. Images were taken with a
Hamamatsu ORCA-R 2 CCD camera on a Zeiss Axio Observer.Z1 Microscope.
Gene Expression Analysis
ES cells grown on gelatin were collected by trypsinization and resuspended in
TRIzol and RNA was isolated with the RNeasy Mini Kit (Qiagen). 1 pg of RNA
was converted to cDNA with Omniscript reverse transcriptase following
manufacturer's instructions (Qiagen).
cDNA was measured using a ABI Fast Cycler and the following Taqman probes
(Invitrogen): Pol P (Mm00448234_ml), Apel (Mm01319526_g1), XRCC1
(Mm00494222_ml), Parp1 (Mm01321084), Aag (Mm00447872_ml), Lig3
(Mm00521933_ml) and GAPDH (Mm99999915_g1).
Aag mRNA Fluorescence In Situ Hybridization (FISH)
ES cells were plated on 28-mm glass bottom dishes (Invitrosci) (200,000/plate)
that were coated with a 1:30 dilution of Matrigel in cold media for 1 hour at room
temperature (Sigma, E1270). The following day cells were fixed with 4%
paraformaldehyde for 20 minutes at room temperature. Cells were permeabilized
with 70% ethanol at 4* C overnight and washed the following morning for 5
minutes with wash buffer (25 % formamide, 2X SSC). Cells were hybridized with
40 20-nucleotide-long fluorescently labeled (Alexa-647) oligonucleotides
designed against the Aag transcript overnight at 370 C in a heavily humidified
chamber in hybridization buffer (100 mg/mL dextran sulfate, Sigma, D8906: 0.5
mg/mL E.coli tRNA, Sigma, R4251: 0.5 mg/mL ssDNA, Sigma D9156: 1 mg/mL
Ultapure BSA, Ambion, AM2616: 10 mM VRC, New England Biolabs, S1402S:
25 % formamide, Ambion, AM9342: 2X SSC, Ambion, AM9763). Cells were
stained with DAPI (2 pg/mL) in wash buffer for 30 minutes at 37* C before
59
imaging. All the previous steps were done in nuclease-free water (Ambion,
AM9932) to avoid RNA degradation.
Images were taken with a Hamamatsu ORCA-R 2 CCD Camera on a Zeiss Axio
Observer.Z1 Microscope. 21 Z-stack images were taken, each 0.3 pM away from
the previous. Images were compiled and foci per cell were counted with a
MATLAB algorithm adapted from previously published methods (Raj et al., 2008).
Western Blots
Exponential growth phase ES cells were harvested by trypsinization, flash frozen
with liquid nitrogen and kept at -80' C until use. Cells were resuspended in 2-3x
the pellet volume with cell lysis buffer (20 mM Tris-HCI pH 8.0, 137 mM NaCl,
10% glycerol, 1% Nonidet P-40, 10 mM EDTA, protease inhibitors (Roche
04693159001), 10 mM NaF, 1 mM DTT, 1 mM sodium orthovanadate) and left
on ice for 30 minutes. Cells were then sonicated on ice (Branson Model 450
Digital Sonifier: Amplitude 20%, 3 cycles each 2 seconds long) and centrifuged
for 10 minutes at max speed. The supernatant was removed and quantified with
Micro BCA assay kit (Thermo Scientific Pierce). 30 ug of denatured protein was
run per lane on precast NuPAGE@ Novex@ Bis-Tris gels (invitrogen) in MOPS
buffer for 1 hour at 200 V. Proteins were transferred to Immuno-Blot PVDF
Membranes (Bio-Rad) for 1 hour at 100 V in transfer buffer (49.1 mM Tris base,
38.6 mM glycine, 20% methanol). Membranes were blocked with Odyssey
blocking buffer (LI-COR Biosciences) and then stained with primary antibodies in
a 1:1 mixture of blocking buffer and PBS-tween (PBS + 10% Tween-20) and
imaged on an Odyssey Intrared Imaging System (Licor Biosciences).
Ex vivo Hematopoietic Colony Forming Assays
Bone marrow was extracted from the femurs of adult mice 8-12 weeks of age
using a 25 G needle into RPMI media (Invitrogen). Cells were pipetted to a single
cell solution, counted and diluted to 2 x 106 cells/mL in RPMI. Two millions cells
60
were aliquoted out per drug treatment and treated with MMS in RPMI media
without serum for 1 hour at 370 C. After treatment, samples were spun down at
1200 rpm for 5 minutes and washed once with RPMI to remove any extra MMS.
Cells were then resuspended and added to complete hematopoietic
methylcellulose (R&D Systems; Cat. # HSCO07), vortexed, and 1.1 mL was
added to 35 mm culture dishes in duplicate with a 16G blunt end needed (Stem
Cell Tech.). Two 35 mm cell containing dishes and a third filled with water
(without lid) were placed in a 10 cm dish and kept at 370 C for 8 days. Colonies
were counted by eye through a microscope at 1OX magnification.
In vivo Sensitivity to MMS
Mice (8-12 weeks old) were treated with varying doses of MMS by i.p injection.
Tissues were processed at the David H. Koch Institute for Integrative Cancer
Research, Histology Core Facility where they were paraffin-embedded, sectioned
at 5 pm and stained with hematoxylin and eosin (H&E) prior to imaging.
61
Results
Generation of C57BI/6 and 129 Mouse Embryonic Stem Cells and Validation
of Pluripotency
We generated new WT, Aagt, and mAagTg embryonic stem (ES) cells on either
a 129 or C57B1/6 genetic background with the assistance of the ES Cell and
Transgenics Facility at the Koch Institute for Integrative Cancer Research.
Previously published ES cell results demonstrating Aag sensitivity to MMS were
conducted on cells with a 129 genetic background (Allan et al., 1998; Engelward
et al., 1998; Maor-Shoshani et al., 2008), yet all in vivo mouse experiments
showing Aag' resistance to MMS have been done on a C57B11/6 genetic
background (Calvo et al., 2013b; Meira et al., 2009b). Thus, we wanted to
determine whether Aag dependent responses were influenced by genetic
background. Blastocysts were isolated from C57B1/6 (WT & Aag') or 129
females (WT/mAagTg & Aag-) 3.5 days post-coitus and coaxed into new ES cell
lines. Recent advances in our understanding of ES cell pluripotency have
increased the efficiency of ES cell generation and only 1 female was needed per
genotype in order to produce multiple cell lines (Fig. 2.1B) (Hassani et al., 2012;
Ying et al., 2008).
Mouse ES cells are often genetically unstable and altered clonal populations can
rapidly overtake otherwise healthy cell lines. We first examined 5 C57B11/6 cell
lines within our lab and found none to contain obvious numerical chromosomal
abnormalities (Fig. 2.2A and 2.2B). However, up to 38% of mES cell lines contain
structural genetic changes that are not detected by simple chromosome spreads,
such as chromosomal translocations (Kim et al., 2013). We sent out four C57B1/6
ES cell lines for professional karyotyping (Cell Line Genetics; Madison, WI).
Neither of the WT cell lines examined had any chromosomal aberrations;
however, both Aag' cell lines had at least one metaphase spread with non-clonal
aberrations (Fig. 2.2C). Aag' C57B11/6 #1 cell line exhibited trisomy 15, which
does not pose a significant risk to cell line health. Aag' #3, on the other hand,
62
had three non-clonal aberrations, the most worrisome being trisomy 8, a
chromosomal aberration that confers an in vitro growth advantage, interferes with
differentiation potential, and can account for up to 89% of all ES cell genetic
abnormalities (Kim et al., 2013; Liu et al., 1997). We therefore chose to use cell
lines WT #1 and Aag'#1 for all future experiments. All cell lines were found to
be male (XY).
We verified that all cell lines being used were pluripotent by immunocytochemical
staining against the pluripotency associated proteins markers SSEA-1, Nanog
and Sox2. All ES cells (129 & C57B1/6) were stained positive for these markers
while differentiated primary MEFs did not exhibit any staining (Fig. 2.3).
BER Gene Expression and Aag Activity in WT, Aag' and mAagTg ES Cells
Next, we sought to verify and quantify the differences in Aag expression and Aag
glycosylase activity between cell lines. AagV C571B1/6 cells showed little to no
expression of Aag as determined by qPCR, which translated into significant
differences in Aag glycosylase activity (Fig. 2.4). Similarly, Aag expression was
significantly different between Aag', WT, and mAagTg 129 ES cell lines as were
intracellular Aag activity levels (Fig. 2.5).
We measured the levels of downstream BER proteins to determine whether there
were inherent imbalances in BER that would affect pathway throughput and
therefore sensitivity to alkylating agents. There were no significant differences in
expression of Apel, Xrccl, Parpi, Pol P, and Lig 3 between cell lines (Fig. 2.6).
Moreover, we measured the protein activity levels of Apel in 129 ES cells and
found no significant differences in activity (Fig. 2.5C). Taken together, these
results indicate that neither Aag expression nor protein activity influence the
expression or activity of other downstream BER proteins in ES cells.
63
Sensitivity of ES Cell Lines to Alkylating Agents
It has been previously shown that the loss of Aag in 129 ES cells renders them
sensitive to the alkylating agents MMS, BCNU, Mitomycin C (MMC), and
psoralen. We now show that genetic background does not affect Aag' sensitivity
to MMS or MMC, as Aag' C57B1/6 ES cells were sensitive compared to WT (Fig.
2.7A and 2.7C). We also measured sensitivity to methylnitrosourea (MNU),
another monofunctional methylating agent. Similar to MMS treatment, we found
that loss of Aag increases cellular sensitivity to MNU (Fig. 2.7D). Interestingly,
Aag' C57B1/6 ES cells showed no enhanced sensitivity to the alkylating agent
BCNU, a contrast to 129 ES cells (Engelward et al., 1996), indicating that
sensitivity to at least some DNA damaging agents appears to be dependent on
genetic background (Fig. 2.7B).
Overexpression of Aag has been shown to sensitize many mouse and human
cell lines to treatment with alkylating agents (Calvo et al., 2013b; Tang et al.,
2011; Trivedi et al., 2008). Though the lab has assessed the effect of
overexpression of Aag in various differentiated tissues post MMS treatment in
vivo and seen severe sensitivity compared to WT, we have not determined how
Aag overexpression alters ES cell sensitivity, if at all. We were surprised to find
that both loss of Aag and overexpression of Aag renders ES cells sensitive to
MMS. Furthermore, cells overexpressing Aag were even more sensitive than
Aag' ES cells (Fig. 2.8A). These results agree with hypotheses that ES cells,
because of their stem-cell pluripotent nature, are exquisitely sensitive to
imbalanced BER and the formation of toxic AP sites and SSBs that could cause
hazardous mutations, which would then be passed along to all subsequent
daughter cells. Likewise, we found that treatment with high doses of the SN1-type
methylating agent MNNG causes the same Aag dependent cell sensitivity profile
as seen for the SN2-type alkylating agent MMS (Fig. 2.8B). Overexpression of
Aag similarly caused increased cell sensitivity to the interstrand crosslinking (ICL)
agent mitomycin C, albeit to the same extent as loss of Aag (Fig. 2.8C).
64
Discussion
Embryonic stem cells are pluripotent cells that have the capacity for unlimited
self-renewal. To maintain the genomic integrity of all potential daughter cells, it is
postulated that these cells must possess enhanced mechanisms to limit DNA
damage accumulation and mutations compared to differentiated cells. Indeed,
the DNA repair pathways BER and mismatch repair (MMR) are more active in ES
cells compared to MEFs, and ES cells preferentially utilize the more accurate
homologous recombination to repair DSBs than error-prone non-homologous end
joining (NHEJ) (Tichy et al., 2011; Tichy and Stambrook, 2008). Moreover, ES
cells are more sensitive than MEFs after treatment with gamma irradiation and
the methylating agent N-methyl-N'-nitro-nitrosoguanidine (MNNG) (de Waard et
al., 2008; Roos et al., 2007). In this study, we explored the effects of genetic loss
or overexpression of the Aag glycosylase in ES cells from two different inbred
genetic mouse models.
Previous work has demonstrated that loss of Aag in 129 ES cells causes
sensitivity to MMS. Conversely, loss of Aag in C571B1/6 cerebellar granule
neurons, retinal photoreceptors, and hematopoietic progenitors renders these
cells totally resistant to MMS induced death (Calvo et al., 2013b; Meira et al.,
2009b; Roth and Samson, 2002). We sought to determine whether these
differences in sensitivity were dependent upon cell type or genetic background.
To this end, we generated new WT and Aag-* ES cells from pure C57B1/6 mice
and showed that, similar to 129 ES cells, loss of Aag causes increased cell death
post-alkylation treatment, indicating this to be an ES cell specific response and
not dependent on genetic background. Likewise, Aag~ C57B11/6 ES cells were
sensitive to the methylating agent MNU. Moreover, we have shown that MMS-
sensitivity of hematopoietic progenitors, cerebellar granule neurons, and retinal
photoreceptors from 129 mice is Aag-dependent, further corroborating that Aag-
dependent responses to MMS are cell-specific and not contingent on genetic
background (Fig. 2.9).
65
Genetic background did, however, affect ES cell sensitive to BCNU. 129 ES cells
lacking Aag are sensitive to the chemotherapeutic drug, while there was no
difference in sensitivity between WT and Aag' C57B11/6 ES cells. Under certain
circumstances, a single genetic manipulation is sufficient to cause identical
phenotypes in various inbred mouse strains, yet it is becoming more common to
observe variations in phenotype due to modifier genes that act in combination
with the gene of interest (Montagutelli, 2000). BCNU is a complex bifunctional
alkylating agent that produces both mono-adducts and interstrand crosslinks
(ICLs) and Aag-initiated BER represents only one of many relevant DNA repair
pathways potentially involved in ICL repair. In particular, the direct reversal repair
protein 06-methylguanine methyltransferase (MGMT) has been shown to be a
strong predictor of cell sensitivity to BCNU by repairing mono-adducts before
they are converted to ICLs (Bodell et al., 1986; Phillips et al., 1997; Samson et
al., 1986). It is possible that there are differences in expression or activity of
MGMT between 129 and C57B1/6 ES cells, as up to 2% of all genes have been
shown to be differentially express between genetic background (Kraus et al.,
2012; Turk et al., 2004). Moreover, 129 cells and mice do not express functional
polymerase Iota, an error-prone translesion polymerase that has been Implicated
in the bypass of ICLs (Ho and Scharer, 2010; McDonald et al., 2003; Smith et al.,
2012). Interestingly, the influence of genetic background is specific to BCNU
damage since there was no difference in sensitivity to mitomycin C, another
bifunctional ICL generating drug.
Though there are numerous examples of how loss of DNA repair genes affects
ES cell sensitivity to DNA damaging agents, there are minimal studies assessing
the effects of increased DNA repair in pluripotent stem cells compared to WT. It
is reasonable to hypothesize that enhanced DNA repair would reduce sensitivity
to DNA damaging agents assuming ES cells strive to maintain genetic integrity.
On the other hand, increasing the activity of only one component of a multi-
enzyme process may cause pathway imbalance and increased sensitivity to DNA
66
damage. We found that overexpression of Aag was more harmful to ES cells
than WT Aag levels or not initiating DNA repair at all after treatment with the
alkylating agents MMS and MNNG. In fact, overexpression of Aag has been
shown to sensitize numerous different cell types to alkylation treatment, including
murine neurons, hematopoietic progenitors, retinal photoreceptors, and human
breast cancer cells (Calvo et al., 2013b; Harrison et al., 2007; Meira et al., 2009b;
Rinne et al., 2005; Trivedi et al., 2008).
The molecular mechanisms leading to cell death downstream of Aag
hyperactivity vary depending on the cell type. Initiation of BER in mouse
cerebellar neurons and retinal photoreceptors activates poly (APD-ribose)
polymerase 1 (Parpi), which is the sole mediator of Aag-dependent cell death
after alkylation treatment (Calvo et al., 2013b; Meira et al., 2009b). In ES cells,
however, cell death is most likely due to replication stress and downstream
apoptosis. Aag' and mAagTg ES cells will both experience increased replication
blocks compared to WT cells, the former due to 3MeA lesions remaining in the
genome unrepaired and the latter from abasic sites and single strand breaks
(SSBs) created during imbalanced BER. Each of these replication blocks can
cause highly cytotoxic double strand breaks (DSBs) when encountered by a
replication fork (Ensminger et al., 2014). In fact, Aag' ES cells were shown to
have increased sister chromatid exchanges and chromosomal breaks post-MMS
treatment compared to WT, both of which are indicative of DSBs (Engelward et
al., 1998). It would therefore appear that WT ES cells are optimally devised to
complete BER after initiation without accumulation of toxic intermediates. This
suggests that ES cells must strive to maintain an appropriate level of Aag activity
within some optimal range such that BER is effectively completed after initiation.
Nonetheless, it is interesting that we observed increased cell sensitivity upon Aag
overexpression compared to loss of Aag (though both were more sensitive than
WT), indicating either differences in perceived cellular DNA damage or cell death
mechanism. Upon treatment with the same dose of MMS, mAagTg cells may
67
sense higher levels of DNA damage compared to WT or Aag' cells due to
unbalanced initiation of BER causing persistent abasic sites and SSBs.
Additionally, overexpression of Aag has been shown to remove the otherwise
non-toxic DNA lesion 7MeG after alkylation treatment, thus initiating DNA repair
and exposing the cell to toxic repair intermediates that will not have been
significant in WT or Aag' cells (Rinne et al., 2005; Sobol et al., 2003).
Alternatively, downstream cell death signaling pathways might vary between cell
lines. Aag and Apel activity are responsible for the generation of SSBs during
BER which was shown to trigger the DNA damage response through the kinases
ataxia telangiectasia-mutated (ATM) and checkpoint kinase 2 (Chk2) in human
293T and HeLa cells (Chou et al., 2015). Aag has also been shown to directly
interact with and inhibit p53's ability to act as a transcription factor for pro- and
anti-apoptotic genes (Song et al., 2012). Ultimately, however, it may be a
combination of effects that causes increased cell death upon enhanced DNA
damage repair.
68
3.5 dpc
Pregnant Mouse
2-3 Weeks
Isolate Blastocysts Embryonic StemCell Lines
B
Genotype Blastocysts Blastocysts thatIsolated Generated ES Cell Lines
WT C57B1/6 8 6
AagG C57B/6 5 5
mAagTg/WT 129 7 3
Aag'- 129 5 2
Figure 2.1: Generation of new Embryonic Stem (ES) Cells.
(A) Schematic of the isolation and generation of ES cells. Blastocysts were
isolated from pregnant females 3.5 days post-coitus and plated on a bed
of irradiated mouse embryonic fibroblasts (MEFs). ES cells were derived
from the inner cell mass (ICM) of the blastocyst.
(B) Blastocysts from a variety of genotypes and genetic backgrounds were
harvested and ES cells were generated.
69
Figures
A
C57B1/6 ES Cell Chromosome Counts
44-
42-
40-
38-
36-
Eamon
No
-"
Uo A
I **
K.
N)I' ~
B
N=40
'5
2.2: Karyotyping of C57B1/6 ES Cells.
Counting of metaphase spreads revealed no major numerical
chromosomal abnormalities between cell lines.
Representative chromosome spread with the expected 40 chromosomes.
Results of professional G-band karyotypic analysis. Based on these
results, cell lines WT #1 and Aag- #1 were chosen for future experiments.
A
0
0
0
0
C
Metaphase # Apparently Normal Non-Clonal Aberrations KaryotypeSpreads (0X) NnCoa brain aytp
Analyzed '
WT #1 20 20 40, XY
WT #2 20 20 40, XY
Aag-1- #1 20 19 41, XY, +15 40, XY
41, XY, +8
Aag4 - #3 20 17 40, XY, del(19)(C2) --
41, XY, add(8)(E1), +mar
Figure
(A)
(B)
(C)
70
AA
A A A A A
. I
WT Aag- IR MEFs WT Aag- IR MEFs
HoechstHecs
B
WT Aag' mAagTg IR MEFs
Figure 2.3: Fluorescent Immunocytochemical Staining for Pluripotency Markers
in C571B1/6 and 129 ES Cells.
(A) WT and Aagl C57BI/6 ES cells, and not irradiated MEFs, exhibit staining
for pluripotency transcription factor Sox2 (left) or the cell surface
pluripotency marker SSEA-1 (right)
(B)WT, mAagTg, and Aag' 129 ES cells, and not irradiated MEFs, exhibit
staining for pluripotency transcription factor Sox2 or the cell surface
pluripotency marker SSEA-1
71
B~zz~
~1~~
C:
0
aI)
LU
0
a)
01
0
C,)
a) 1.0-
0.5-
U)[If
40-
30-
20-
10-
Aag Activity
_T1__
0-I-
Figure 2.4: Aag Expression and Activity in C57B1/6 ES Cells.
(A)Aag-'- C57B1/6 have significantly reduced expression of Aag as measured
by qRT-PCR. (Errors bars denote standard deviation from the mean, n=3,
*** p<0.001 using Student's standard two-tailed T-test)
(B) Host cell-reactivation assay demonstrates significant difference in Aag
glycosylase activity. (Errors bars denote standard deviation from the
mean, n=3, *** p<0.001 using Student's standard two-tailed T-test)
72
___________- 
--
_ _ 
I
A
0.0045
O.OL
B
Aag Activity
0-
LU
A If
A "A 0
0 A~
0 A]A
0 'AA
S.A
20-
15-
10-
5-
0-
T1.
C
Apel Activity in 129 ES Cells
1.5-1
I
'S.
.5
t5
U)
0.
U)
Cu
1 0-
0.5-
0.0.
Figure 2.5: Aag Expression and Activity in 129 ES Cells.
(A)Aag' and mAagTg ES cells have significantly different expression of Aag
compared to WT. Each data point on the graph represents the number of
individual Aag transcripts in a single cell as measure by mRNA FISH.
(Errors bars denote standard deviation from the mean, n>35, *** p<0.001
using Student's standard two-tailed T-test)
(B) Host cell-reactivation assay demonstrates significant difference in Aag
glycosylase activity and no significant difference in Apel activity (C).
(Errors bars denote standard deviation from the mean, n=3, *** p<0.001
using Student's standard two-tailed T-test)
73
A
100-
hr
<C)
50-
oJ
0
I
ell-
B
129 ES Cells
6- M 129 WVT
M 129 Aag
129 mAagTg
4-
2-
C
0
U)
(D
uJ
a)
M
6-
4-
2-
C57B/6 ES Cells
M C57BI/6 WT
D C57BI/6 Aag-
U-.
e;~
01
Figure 2.6: Expression of Base Excision Repair genes in ES cells.
Aag is the only gene differentially expressed between Aag", WT, and mAagTg
129 (A) and C57BI/6 (B) ES cells. (Errors bars denote standard deviation from
the mean, n=3)
74
A
C
0
LU
<Z)
0
B
100.0
1 0-
10 0
0.01 i . I ,
0 1 2 3
MMS (mM)
100'
10
0.1
0.01
0.001 .
0.0 0.5 1.0 1.5
Mitomycin C (ug/ml)
D
0o
Figure 2.7: Sensitivity of C57B1/6 ES Cells to Alkylating Agents.
Sensitivity of WT (black circles) and Aag-'- (red triangles) C57B1/6 ES cells to
alkylating agents. (A) MMS (n=6), (B) BCNU (n=3), (C) Mitomycin C (n=3), and
(D) MNU (n=3). (Errors bars denote standard deviation from the mean, n=3, **
p< 0 .0 1 using Student's standard two-tailed T-test comparing a single dose to
WT)
75
A
-0
C
0U
100-
10
1
0.1
0.01
0 100 200 300
BCNU (uM)
100
10-
1~
0.0 0.5 1.0 1.5
MNU (mM)
B
100
10
1
-W WT
Aag-
-0- mAagTg
0.01*
0.001
0.0 0.5 1 0 1.5 2,0 2.5
MMS (mM)
C
100k%
10
1- Aag
-0- mAagTg
0 5 10 15 20
MNNG (uM)
0U
00
10
-0- WT
1 Aag
-9- mAagTg
0.0 0.5 1 0 1 5
Mitomycin C (ug/ml)
Figure 2.8: Sensitivity of 129 ES Cells to Alkylating Agents.
Sensitivity of WT (black circles), Aag' (red triangles), and mAag Tg ES cells (blue
squares) to alkylating agents using colony forming assaya. (A) MMS (n=3), (B)
MNNG (n=2), and (C) Mitomycin C (n=4). (Errors bars denote standard deviation
from the mean, * p<0.05 using Student's standard two-tailed T-test comparing a
single dose to WT)
I
76
A
C,
1
A 1001B MMS 150 mg/kg, 24h
Untreated Aag mAagTg
10_
Aag-'
-0- WT
-W- mAagTg
0.1 I I
0.0 0.5 1.0 1.5 2.0
MMS (mM)
C MMS 75mg/kg 7d
Untreated WT Aag'- mAagTg
Figure 2.9: MMS Sensitivity of 129 Hematopoietic Progenitors, Cerebellar
Granule Neurons, and Retinal Photoreceptors is Aag-dependent.
(A) Ex vivo hematopoietic colonies forming assays from WT, Aag-, and
mAagTg 129 mice. Errors bars denote standard deviation from the mean,
n=3.
(B) H&E stained cerebellar sections of Aag- and mAagTg 129 mice 24 hours
post MMS treatment (150 mg/kg). White areas represents areas of edema
surrounding dying pyknotic cells. Scale bar is 100 pm. All untreated mice
appeared identical.
(C) H&E stained retinal sections of WT, Aag-l, and mAagTg 129 mice 7 days
following MMS treatment (75 mg/kg). Yellow arrowheads indicate outer
nuclear layer and yellow stars indicate inner nuclear layer. Scale bar is
100 pm. All untreated mice appeared identical; the representative
untreated sample shown here is WT.
77
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83
Chapter III: In vitro Hematopoietic and
Neuronal Differentiation of Mouse
Embryonic Stem Cells
Carrie M. Margulies, June Criscione, Danielle Wall, Leona D. Samson
Contributions: C.M.M. and L.D.S. designed experiments. C.M.M. conducted all
experiments unless otherwise noted. J.C. and D.W. assisted with in vitro
neuronal differentiation experiments.
84
Chapter III Table of Contents
Chapter 11I: In vitro Hematopoietic and Neuronal Differentiation of Mouse
Em bryonic Stem Cells ..................................................................................... 86
Introduction .................................................................................................. 86
Materials and Methods................................................................................. 89
Results ......................................................................................................... 95
In Vitro Hem atopoietic Differentiation of 129 ES Cells.............................. 95
In vitro Differentiation of Cerebellar Granule Neurons (CGNs) ................ 96
In vitro Adherent Differentiation of Cortical Neurons................................. 98
In vitro Neural Differentiation Following 'Bibel' et al. (2007).......................100
D is c u s s io n ...................................................................................................... 1 0 1
F ig u re s ........................................................................................................... 1 0 5
T a b le s ............................................................................................................ 1 1 5
References.....................................................................................................116
85
Chapter III: In vitro Hematopoietic and Neuronal Differentiation of Mouse
Embryonic Stem Cells
Introduction
DNA damaging sources are ubiquitous in our external and internal environments,
presenting constant threats to the integrity of our DNA. One type of damage,
methylation damage, is caused by the covalent addition of methyl groups (-CH 3)
to reactive sites on all components of our cells; however, it is the methylation of
DNA that is particularly toxic. Thankfully, our cells have devised numerous repair
mechanisms throughout evolution to avoid cytotoxicity and DNA mutations that
can manifest in disease. One such DNA repair pathway is base excision repair
(BER) (Drablos et al., 2004).
BER is initiated by a lesion-specific glycosylase that searches the genome for
damage and upon recognition, excises the DNA base lesion by hydrolyzing the
glycosyl bond. In the case of methylation damage, BER is initiated by the
alkyladenine DNA glycosylase (Aag). Excision of the damaged base leaves an
abasic (AP) site that is cleaved by the apurinic/apyrimidinic endonuclease 1
(Apel) resulting in a single-strand break (SSB) with 3'-OH and 5'-
deoxyribosephosphate (5'-dRP) ends. Polymerase P (Pol P) removes the toxic 5'-
dRP termini and catalyzes the insertion of any missing nucleotides before the
SSB is ligated to complete repair.
Interestingly, results from our lab have demonstrated that loss of Aag can cause
opposite effects depending on the cell type. Aag was first knockout in pluripotent
mouse embryonic stem (ES) cells and loss of the DNA repair protein rendered
these cells sensitive to the methylating agent methyl methanesulfonate (MMS),
among other alkylating agents, compared to wild-type (WT) (Allan et al., 1998;
Engelward et al., 1998; Engelward et al., 1996). Yet surprisingly, Aag' mature
mice display striking resistance to MMS-induced tissue degeneration in particular
86
tissues compared to WT; even though there is no difference in overall organismal
sensitivity (Calvo et al., 2013b; Roth et al., 2002). Among these Aag- MMS-
resistant tissues are hematopoietic myeloid progenitors, cerebellar granule
neurons, retinal photoreceptors, pancreatic @-cells, and splenocytes (Calvo et al.,
2013b; Meira et al., 2009a; Roth et al., 2002). Thus it would seem that there are
cell-specific inherent qualities that determine how loss of a DNA repair protein
affects sensitivity to DNA damaging agents.
Though pluripotent mouse embryonic stem cells were first generated in 1981, the
recent advent of induced pluripotent stem (iPS) cells has stimulated increased
investigation of their in vitro differentiation capacity in hopes of generating
patient-specific cells and tissues to cure human disease and assist in drug
discovery and development (Evangelos Kiskinis, 2010; Evans and Kaufman,
1981; Inoue and Yamanaka, 2011; Takahashi and Yamanaka, 2006; Wu and
Hochedlinger, 2011). In addition, generation of these differentiation protocols has
enabled investigators to study a broad array of isogenic somatic cells types in
vitro, therefore eliminating the need to maintain mouse colonies to study different
tissues or cell types.
Currently, there are protocols available for the differentiation of -200 different cell
types (Inoue and Yamanaka, 2011). Within the neuronal field, differentiation of
retinal cells, striatal neurons, dopaminergic neurons, motor neurons,
oligodendrocytes, astrocytes, peripheral neurons and cerebellar granule neurons
has been documented (Mattis and Svendsen, 2011; Salero and Hatten, 2007).
This battery of specialized neuronal differentiation techniques has been applied
to study neurological conditions such as Parkinson's disease, spinal muscular
atrophy, RETT syndrome, Huntington's disease and Friedreich ataxia, in patient-
specific iPSCs where resulting 'disease' neurons display distinct phenotypes
correlating to the disease (Wu and Hochedlinger, 2011).
87
In vitro hematopoietic differentiation is one of the oldest differentiation techniques
described, spurred by an early desire to treat blood disorders with healthy stem
cells derived in culture instead of from blood donors. In vitro hematopoietic
differentiation was first documented in 1991 and has been characterized in detail
over the past two decades (Keller, 2005; Schmitt et al., 1991; Wiles and Keller,
1991). It is one of the more detailed differentiation methods studied, with
commercially available protocols, materials, and kits.
For our purposes, in vitro differentiation of genetically altered ES cell provides an
avenue to investigate cell specific responses exhibited in adult mice. In particular,
we are interested in studying how Aag' MMS-sensitive ES cells differentiate to
become Aag' MMS-resistant hematopoietic progenitors and cerebellar granule
neuron (CGNs) compared to their respective WT cells (Fig. 3.1). Though it is
possible to study these cell-specific responses to MMS in homogenous
populations of primary cells, in vitro differentiation offers the opportunity to track
cell response as a function of cellular development. Here we describe our efforts
to differentiate WT and Aag' ES cells into hematopoietic progenitors and
cerebellar granule neurons and study cell responses to MMS treatment. Though
it has proved challenging, we have SuCCessfully differentiated both cell types of
interest and are pursuing new techniques to allow for the study of their sensitivity
to MMS-induced genotoxicity.
88
Materials and Methods
Materials and Antibodies
Methyl methanesulfonate (MMS; Cat. #129925) was purchased from Sigma.
Unless otherwise noted, cell culture components were purchased from
Invitrogen. All cytokines were purchased from R&D Systems unless otherwise
noted.
The following antibodies and their dilutions were used: GFAP (AbCam, ab7779-
500, 1:500), Map2 (AbCam, ab5392, 1:20,000), Tuj1 (AbCam, ab7751, 1:500),
yH2AX (AbCam, ab2893, 1:1,000), Nestin (AbCam, ab6142, 1:500), and cleaved
caspase 3 (Cell Signaling Tech, 9661S, 1:500).
Cell Culture
MEFs were grown in DMEM plus 10% heat-inactivated fetal bovine serum (HI-
FBS) (Sigma), 1% L-glutamine and 1% penicillin/streptomycin. To generate
feeder layers, primary MEFs were irradiated with 45 Gy with a Cobalt-60 source
and frozen at 1 x 10 7 cells/vial. ES cells were grown in DMEM plus 15% HI-FBS
(Sigma), 1% L-glutamine, 1% non-essential amino acids, 1% sodium pyruvate,
100 uM P-mercaptoethanol (P-ME) (Sigma, M3148), 1% penicillin/streptomycin
and recombinant mouse leukemia inhibitory factor (LIF, Millipore). All
components were purchased from Invitrogen unless noted. ES cells were grown
either on an irradiated MEF feeder layer or on tissue culture plates coated with
0.1% gelatin.
In Vitro Hematopoietic Differentiation
The following method was adapted from Stem Cell Technologies 'In Vitro
Hematopoietic Differentiation of mESCs & miPSCs' technical manual, version
3.1.0' (method available at this address:
http://www.stemcell.com/-/media/Technical%20Resources/0/28415MAN 3 1 0.
89
pdfla=en). ES cells were grown for one passage in IMDM based ES complete
media prior to initiation of hematopoietic differentiation (Same media formulation
as complete ES media but substituting IMDM for DMEM). Upon -75%
confluence, ES cells were trypsinized and resuspended at 50,000 cell/mL in
IMDM hanging drop media (15% HI-FBS, 1% L-glutamine, and 100 pM P-
mercaptoethanol in IMDM media). Using a multichannel pipet, 15 pL drops were
initiated on the bottom of a 15 cm Petri dish, being careful to keep each drop
separate. The tops were replaced and the plates were carefully, but swiftly,
inverted so that the media and cells hung as individual drops. The plates were
incubated at 370 C and after two days, one embryoid body (EB) could be
observed in each drop. EBs were collected in 10% HI-FBS/IMDM by gentle
pipetting into a 50 mL conical tube and allowed to settle to the bottom before very
gently aspirating the excess media. EBs were resuspended in minimal amount of
10% HI-FBS/IMDM and transferred to methylcellulose based media (1%
methylcellulose (Stem Cell Technologies), 15% HI-FBS, 2 mM L-Glutamine, 150
pM mono-thioglycerol (MTG: Sigma), 40 ng/mL murine stem cell factor (mSCF;
Cell Signaling Tech.) in IMDM). EBs in methylcellulose media were added to 35
mm non-tissue culture treated dishes using blunt-end needles (Stem Cell Tech.,
cat #28110) and asyrin ge. EBs are allowed to grow at 370 C for 5 days at which
point they were fed with 500 pL/plate of the following media (0.5%
methylcellulose, 15% HI-FBS, 150 pM MTG, 160 ng/mL mSCF, 30 ng/mL murine
interleukin-3 (mlL-3), 20 ng/mL human interleukin-6 (hlL-6), 3 U/mL human
erythropoietin (rhEPO) in IMDM).
By 5 more days growth, large EBs had developed and many had begun beating
due to the formation of myocardial muscle cell differentiation. EBs were
harvested with IMDM media to dilute the methylcellulose and digested with
collagenase (Stem Cell Tech. cat #07902) for 1 hour at 370 C. To ensure a
single-cell suspension, cells were pipetted through an 18G needle and run
through a 70 pm filter before plating. For drug treatment, 3 million cells were
counted and treated with MMS in serum-free IMDM media for 1 hour at 370 C.
90
Following treatment, cells were plated in complete murine hematopoietic
methylcellulose media using a blunt-end syringe and grown for 7-10 days at 370
C, after which time colonies were counted under the microscope (1.4%
methylcellulose, 15% HI-FBS, 2% bovine serum albumin (BSA), 2 mM L-
glutamine, 50 pM P-ME, 200 pg/mL human transferrin, 5 U/mL rhEPO, 10 pg/mL
recombinant human insulin, 10 ng/mL mlL-3, 10 ng/mL mlL-6, 50 ng/mL mSCF,
in IMDM media). Complete methylcellulose media was purchased from R&D
Systems (Cat. # HSCO07).
In vitro Neuronal Differentiation based on Salero and Hatten, 2007
The first neuronal differentiation method we employed was based on Salero and
Hatten, 2007. Differentiation was initiated by plating -2 x 106 ES cells in a 10 cm
Petri dish in DFK5 media (50%: DMEM/F12 + 1% P/S, 1% L-Glut, 0.1 mM P-ME,
1X N2 supplement (Invitrogen), 20 nM progesterone (Sigma), and 60 pM
putrescine (Sigma) and 50%: complete ES media without LIF and HI-FBS but
containing 10% knock-out serum replacement (Gibco)). After 1 day in vitro (DIV),
2pM all-trans retinoic acid (RA; Sigma) and 100 ng/mL fibroblast growth factor 8b
(FGF8b) were added directly to the media. After 4 DIV, EBs were carefully
collected and resuspended in DMEM with N2 Supplement, 100 ng/mL FGF8b,
100 ng/mL FGF4, and 20 ng/mL bFGF. EBs were plated on surfaces coated with
laminin (1 pg/mL; Sigma) and allowed to adhere and grow for 2 days at which
point the EBs were carefully fed with DMEM with 100 ng/mL FGF8b, 50 ng/mL
Wntl, and 50 ng/mL Wnt 3a. After 5 more DIV, EBs were fed with DMEM plus 1X
N2 supplement B, 1X B27 supplement (Invitrogen), 100 ng/mL BMP7, 100 ng/mL
GDF7, and 20 ng/mL BMP6.
In Vitro Neuronal Differentiation based on Gaspard, et al. (2009)
ES cells growing on gelatin were plated at low density (125,000 - 500,000 cells
for 129 and 750,000 cells for C57B1/6) on gelatin coated 6 cm plates in complete
ES media plus LIF (Gaspard et al., 2009; Ying et al., 2003a). The following day,
91
each plate was washed twice with PBS to remove any residual FBS and fed with
5 mL of N2B27 media (0.5X N2 Supplement (Invitrogen), 1X B27 supplement
without vitamin A (Invitrogen), 1% non-essential amino acids, 1% sodium
pyruvate, 1% L-glutamine, 1% penicillin/streptomycin, 2.5% tissue culture grade
BSA (Invitrogen), and 100 pM P-ME in 50% DMEM/F12 and 50% Neurobasal
media). Every 2 days cells were washed 2-3 times with PSB and fed with N2B27
media until day 8/12 when cells were split. On the day of splitting, cells were fed
with N2B27 media 1 hour before trypsinization. Cell were trypsinized, collected,
and centrifuged for 5 minutes at 290x g. Cells were resuspended in N2B27
media and pipetted sequentially with a P1000, P200 and P20 to break up cell
clusters before being run through a 100 pm filter. Finally, cells were plated on
laminin (1 pg/mL) and poly-D-lysine (PDL; 50 pg/mL) coated surfaces.
Cells were treated at day 14 with MMS in serum-free media for 1 hr at 370 C (1:1
mix of DMEM/F12 and Neurobasal media with 1% each of the following: NEAA,
sodium pyruvate, L-glutamine, pen/strep).
In Vitro Neuronal Differentiation based on Bibel, et al. (2007)
Embryoid bodies were initiated by plating 4 x 1 cells in a 10 cm bacteriological
Petri dish in 15 mL CA media (DMEM, 10% HI-FBS, 1% L-Glutamine, 1% NEAA,
100 uM s-ME). After 2 DIV at 37* C, hundreds of EBs had readily formed and
were washed by gently collecting them in 50 mL conical tubes and allowing the
EBs to fall to the bottom (15-20 minutes). Media was very carefully aspirated and
EBs were resuspended in 15 mL CA media and returned to a new Petri dish for 2
more days of growth. At 4 and 6 DIV, EBs were washed and fed similar to Day 2
but retinoic acid (RA) was added to the media (5 pM). EBs were split on Day 8
with 0.05% (w/v) trypsin (Sigma, Cat. #T8802) in 0.05% (w/v) EDTA/PBS for 5-8
minutes in a 370 C water bath. Aliquots of trypsin were kept at -800 C and thawed
immediately before use. Cells were then resuspended in N2B27 media, pipetted
thoroughly to achieve a single-cell suspension, and centrifuged for 5 minutes at
200 g. Cells were resuspended in a minimal amount of media, run through a 70
92
pm filter, counted and plated at 2 x 105 cells/cm 2 on poly-ornithine (100 pg/mL)
and laminin (1 pg/mL) coated surfaces in N2B27 media.
Immunofluorescence
Differentiated cells were plated in 24- or 96-well glass bottom plates (Invitrosci)
treated with laminin/PDL/poly-ornithine and fixed with 4% paraformaldehyde for
15-30 minutes. Cells were then permeabilized with 0.1% Triton-X in CSK buffer
(100 mM NaCl, 300 mM sucrose, 10 mM PIPES, 3 mM MgC 2, 1 mM EGTA) for
20 minutes, blocked with 4% bovine serum albumin (BSA) for 1 hour at 370 C,
and stained overnight at 40 C with appropriate dilutions of antibodies in 4% BSA
in PBS. Secondary antibodies were added for 2 hours at room temperature and
nuclei were stained with Hoechst for 15 minutes. Images were taken with a
Hamamatsu ORCA-R 2 CCD Camera on a Zeiss Axio Observer.Z1 Microscope.
Cell staining was quantified by eye using ImageJ (NIH).
Aag mRNA Fluorescence In Situ Hybridization (FISH)
Differentiated neurons (Bibel method) were plated on PDL-coated 28-mm glass
bottom dishes (Invitrosci) and fixed the following morning with 4%
paraformaldehyde for 20 minutes at room temperature. Cells were permeabilized
with cold 70% ethanol overnight at 40 C and washed the following morning for 5
minutes with wash buffer (25 % formamide, 2X SSC). Neurons were stained with
40 20-nucleotide-long fluorescently labeled (Alexa-647) oligonucleotides
designed against the Aag transcript overnight at 37* C in a heavily humidified
chamber in hybridization buffer (100 mg/mL dextran sulfate, Sigma, D8906: 0.5
mg/mL E.coli tRNA, Sigma, R4251: 0.5 mg/mL ssDNA, Sigma D9156: 1 mg/mL
Ultapure BSA, Ambion, AM2616: 10 mM VRC, New England Biolabs, S1402S:
25 % formamide, Ambion, AM9342: 2X SSC, Ambion, AM9763). Finally, cells
were stained with DAPI (2 pg/mL) in wash buffer for 30 minutes at 370 C before
imaging. All the previous steps were done in nuclease-free solutions to avoid
RNA degradation.
93
Images were taken with a Hamamatsu ORCA-R 2 CCD Camera on a Zeiss Axio
Observer.Z1 Microscope. 21 Z-stack images were taken per focal area, each 0.3
uM away from the previous. Images were compiled and foci per cell were
counted with an automated MATLAB algorithm adapted from previously
published methods (Raj et al., 2008).
94
Results
In Vitro Hematopoietic Differentiation of 129 ES Cells
Given the availability of detailed protocols and commercial products available for
in vitro hematopoietic differentiation, we attempted this differentiation technique
first. In the first step, ES cells are induced to form embryoid bodies (EB) (Fig.
3.2A). EBs roughly resemble the developing embryo and contain features of the
primitive streak, a region of the embryo that will initiate gastrulation, or the
generation of the three germ layers: endoderm, ectoderm, and mesoderm. The
original method we followed induced the formation of embryoid bodies by plating
trypsinized ES cell directly into methylcellulose-based semi-solid media (In Vitro
Hematopoietic Differentiation of mESCs & miPSCs technical manual, version
3.1.0). Upon attempting this method multiple times, however, we were not
successful in generating EBs.
EBs can be formed via numerous different methods, all of which are based upon
the following characteristics: ES cells need to aggregate and grow without
adherence in the absence of the pluripotency maintaining molecular leukemia
inhibitory factor (LIF) (Rungarunlert, 2009). Based on our options, we decided to
try forming EBs through the hanging-drop method. In this procedure, ES cells are
dissociated into single-cells and small drops (15 pL) are dispensed onto the
bottom of a Petri dish, fitting as many drops in a dish as possible without the
drops coalescing into large volumes. The dish is then carefully inverted so that
the media is hanging from the top and the ES cells will fall to the bottom by
gravity and aggregate (Fig 3.2B). Using this method, we found that -90% of
hanging drops had formed EBs after two days of inverted culture (Fig. 3.2C). At
this point, the EBs were very carefully transferred to methylcellulose-based
media to encourage hematopoietic growth.
Within the EB, the cells will naturally differentiate to Flk-1 mesoderm containing
various hematopoietic progenitors and it has been estimated that up to 50% of
95
cells in EBs are Flk-1 (Kabrun et al., 1997). After 12 days of growth, the EBs
were dissociated with collagenase into a single-cell suspension and plated in
methylcellulose-based hematopoietic differentiation media. To test for MMS
sensitivity, cells were treated prior to plating in serum free media. The
methylcellulose media is supplemented with cytokines and growth factors that
specifically allow growth and differentiation of hematopoietic progenitors,
including IL-3, IL-6, murine stem cell factor, and erythropoietin (Fig. 3.2A). After
another 12-14 days of growth, colonies were quantified under a microscope to
determine hematopoietic differentiation efficiency and sensitivity to MMS.
Interestingly, the Aag' 129 ES cells differentiated into hematopoietic progenitors
with relative ease compared to the WT cells, where colonies were much more
rare. We were able to generate a killing curve for Aag' hematopoietic
progenitors differentiated in vitro that mimicked ex vivo cell sensitivity of Aag'
hematopoietic progenitors after in vivo differentiation (Fig. 3.3)(Calvo et al.,
2013b; Roth et al., 2002). Unfortunately, due to low colony numbers from
inefficient differentiation, we were not able to generate a complete WT killing
curve for comparison; however, preliminary results at low MMS doses suggests
that the WT hematopoietic progenitors were more sensitive than Aag' as seen in
x vivo killing assays (Fig. 3.3).
Ultimately, we did not pursue differentiation of hematopoietic progenitors any
further and I focused my efforts on in vitro neuronal differentiation given the work
I was simultaneously conducting on primary CGNs. Hematopoietic differentiation
was handed over to a post-doc.
In vitro Differentiation of Cerebellar Granule Neurons (CGNs)
Despite our preliminary successes at in vitro hematopoietic differentiation, we
turned our attention to neuronal differentiation of cerebellar granule neurons
because of our simultaneous study of ex vivo primary neuronal cultures (Chapter
IV). There has been only one published method for the differentiation of
96
cerebellar granule neurons (CGN) (Salero and Hatten, 2007), possibly because
CGNs are among the last neurons to fully develop in vivo and thus represent a
late and challenging differentiation target ex vivo (Wang et al., 2001). The
protocol to generate CGNs is roughly 3 weeks in length and applies the use of
numerous cytokines known to impact the development of CGNs in vivo (Fig.
3.4A).
As with hematopoietic differentiation, embryonic stem cells are first grown in the
absence of LIF to induce formation of embryoid bodies (EBs). The EBs are then
cultured in the presence of fibroblast growth factor 8b (Fgf8b) and retinoic acid
(RA) to induce expression of neural and general cerebellar markers. Cells at this
step have been induced down the neural, and more specifically, cerebellar
lineage. To stimulate the cells to develop into granule neuron precursors, they
are cultured with additional fibroblast growth factors, Wnt signaling proteins, and
bone morphogenic proteins (BMPs). To recapitulate the rapid expansion of these
cells seen in vivo, sonic hedgehog (Shh) and JAG1 are added to the media.
Finally, the cells are induced to differentiate into mature post-mitotic cerebellar
granule neurons by the addition of brain-derived neurotrophic factor (BDNF). In
total, in vitro differentiation takes 24 days to complete.
After plating the EBs to allow for attachment and growth, cells with long axons
resembling neurons emanated out of the EBs (Fig. 3.4B). Immunocytochemical
staining demonstrated the presence of mature neurons and astrocytes at day 16
of differentiation, among other non-neuronal cell types (Fig. 3.4C). Despite the
presence of mature differentiated neurons, we decided to move to a new method
that avoided differentiation through EBs. EB are inherently heterogeneous and
represent a significant origin of non-neuronal replicative cells that took over
cultures during the extended differentiation protocol. Moreover, neurons
differentiated from adherent EBs have significant 3-dimensional morphology and
the neurons often end up in tightly wound clusters. These structures make
staining, imaging and determinations of cell death much more challenging.
97
In vitro Adherent Differentiation of Cortical Neurons
Neuronal differentiation has long been considered the 'default' differentiation
pathway, since it was shown to be stimulated upon growth without serum (Stern,
2006; Suter and Krause, 2008). Ying, et al. (2003) developed a simple and
elegant method for the differentiation of neuroectodermal precursors based on
monolayer culture and the exclusion of any factors that would induce non-
desirable differentiation. Neuroectodermal cells are the earliest neural tissue
generated during development. Building upon this protocol, Gaspard, et al.
(2008, 2009) demonstrated the differentiation of mature cortical neurons using
this protocol with up to 90% neural cells. Cortical neurons are found in the
mammalian cerebral cortex not the cerebellum; however, our goal was to use this
protocol schematic for the highly efficient generation of mature neurons
combined with the cytokines already published for the specific generation of
CGNs.
ES cells are plated at very-low density in media without serum and allowed to
grow for 12 days. The protocol recommends use of Neurobasal with N2
supplement (Invitrogen) but we found this media was not able to support ES cell
growth and adherence, thus we modified the method by using media containing
the two neural supplements N2 and B27 (Invitrogen). The protocol avoids the
formation of EBs, thus reducing heterogeneity, but also contains a step for
dissociation of differentiating cultures at day 12 (Fig. 3.5A). Dissociation and re-
plating is beneficial when aiming to compare drug treatments to ensure that all
cells are at the same stage of differentiation and went through the process in the
same manner.
In initial experiments, we tested the efficiency of the differentiation method
through immunocytochemical staining at differentiation days 14, 21, and 28.
98
Cultures at each timepoint were fixed and stained with antibodies to mark neural
progenitors (Nestin) and mature neurons (Tuj1 and Map2). We found that ES
cells on a 129 background differentiated much more efficiently than C57B1/6 ES
cells, independent of the genotype (WT vs. Aag-) (Fig. 3.5B). At Day 14 of
differentiation, we had -60% neural progenitors and -25% mature neurons in
129 ES cells, numbers that closely resemble differentiation efficiencies previously
published (Gaspard et al., 2009). Representative fluorescent images
demonstrate that the cells at Day 14 are evenly dispersed over the surface and
are not present in the small cell clusters that remained after dissociation (Fig.
3.6). However, as we let the cells progress to day 21 and 28 of differentiation, we
noticed that the small cell clumps present at day 14 de-differentiated and grew to
become embryoid bodies themselves (Fig. 3.6). A similar phenotype was
observed during hematopoietic differentiation if there was incomplete dissociation
resulting in cell clusters (data not shown). We made numerous attempts to
account for the incomplete dissociations, however, we had a lot of difficulty
recreating these initial results in subsequent experiments for a variety of reasons
(See differentiation attempts in Appendix).
Nevertheless, a few experiments successfully progressed to Day 14 where a
majority of the cells are Nestin+ neural progenitors and we assessed their
responses to MMS. Cells were treated with MMS and queried for the formation of
yH2AX, a marker of double-stranded breaks (DSBs), in Nestin+ cells 2 hours
after MMS treatment. We saw a slight difference in the number of cells positively
stained for yH2AX at 1.5 mM dose; however, this difference was not maintained
through other doses (Fig. 3.7A). There was no difference in the percentage of
Nestin+ cells between genotypes or doses (data not presented, 60-70% of the
cells were Nestin+). We also looked at cell sensitivity to MMS by counting
pyknotic nuclei 8 hours after treatment with MMS. Similarly, we found no
difference in sensitivity between genotypes (Fig. 3.7B). It should be noted that we
do not know whether these results recapitulate in vivo differences since we have
never looked at the sensitivity of neural progenitors, only mature cerebellar
99
neurons. Nevertheless, there was no difference in preliminary experiments and
as mentioned above, we had difficulties optimizing differentiation to retest these
experiments.
In vitro Neural Differentiation Following 'Bibel' et al. (2007)
Based on our difficulties optimizing the adherent differentiation method described
above, we simultaneously started using a third published method (Bibel et al.,
2007; Bibel et al., 2004). This method utilizes retinoic acid, a commonly applied
molecule known to stimulate differentiation of ES cells toward the neural lineage.
ES cells are allowed to aggregate into EBs and are grown in Petri dishes to avoid
adherence (Fig. 3.8). Though EBs are used in the induction of neural lineage,
they are dissociated after 8 days of growth and plated onto surfaces coated with
laminin and poly-ornithine such that only neurons should adhere and survive in
neuronal media. We only recently began using this method but cellular
morphology after differentiation mimics the structure of ex vivo cultured primary
cerebellar granule neurons (Fig. 3.8B).
In preliminary differentiation experiments, we assessed single-cell Aag
expression before and after differentiation using mRNA fluorescence in situ
hybridization (FISH). In addition to quantifying individual Aag transcripts, we also
stained cultures for Map2 to accurately determine which cells were neuronal and
which were non-neuronal (Fig. 3.9B; red arrows indicate non-neuronal cells). WT
ES cells expressed on average 11 transcripts per cell. We found that Aag was
downregulated during differentiation to an average of 2.17 and 6.03 transcripts in
neurons and non-neuronal cells, respectively (Fig. 3.9A and C). This reduction in
Aag expression in in vitro differentiated neurons is similar to Aag expression
patterns in primary cerebellar granule neurons ex vivo (Fig. 3.9C). Therefore, in
all aspects considered so far in vitro differentiated neurons resemble primary
neurons ex vivo.
100
Discussion
In vitro differentiation of pluripotent stem cells is widely regarded as one of the
next big advances in personalized medicine. The use of patient-specific induced
pluripotent stem cells (iPSCs), for example, allows experimentalists the
opportunity to generate new and healthy tissues ex vivo to treat disease without
fear of rejection upon transplantation back into the patient of origin. Moreover,
isogenic stem cells can be differentiated into an array of cells to study how a
gene of interest affects cell physiology. In our case, we used in vitro
differentiation of mouse embryonic stem (ES) cells as a means to study how and
when responses to DNA damage change over the course of cell development as
a function of the presence or absence of the DNA repair protein Aag. Here we
present the results of our experiments to differentiate WT and Aag' mouse ES
cells into hematopoietic progenitors and cerebellar neurons.
Hematopoiesis in the developing mouse embryo occurs in two regions with
differing hematopoietic potential. The yolk sac generates a limited subset of
primitive blood cells to support the embryo during early development. It is the
aorta-gonad-mesonephros (AGM), however, that is the original location of
definitive hematopoiesis that will eventually be relocated to the bone marrow for
the remainder of the organism's life (Dzierzak and Dzierzak, 2008; Orkin, 2008).
Experiments have demonstrated that in vitro hematopoietic differentiation more
closely resembles the primitive hematopoiesis of the yolk sac in both gene
expression patterns and developmental timeline (Keller et al., 1993).
Furthermore, hematopoietic stem cells capable of reconstituting lethally irradiated
adult mice have only been located in the AGM and not in the yolk sac (Medvinsky
and Dzierzak, 1996; Muller et al., 1994). Consequently, even though we were
not able to generate killing curves for both genotypes, the hematopoietic
progenitors generated in vitro from this method may not have fully recapitulated
killing curves and DNA damage responses of hematopoietic progenitors isolated
from adult mice.
101
We successfully generated mature neurons from mouse embryonic stem cells
using three separate methods (Bibel et al., 2007; Gaspard et al., 2009; Salero
and Hatten, 2007). The in vitro differentiated neurons resembled primary murine
neurons in all aspects regarded, including morphology, immunocytochemical
staining, and Aag expression patterns. Ultimately, however, we faced challenges
getting the differentiated cultures to a position where neurotoxicity assays could
be completed. Based on extensive work developing toxicity assays for primary
cerebellar granule neurons (Thesis Chapter IV), we aimed for certain criteria in
our differentiated cultures. These criteria included: relatively homogenous and
predictable cultures, little to no variation in cell heterogeneity between treatment
conditions, a method to identify neurons before and after treatment without the
need for fixation, and even monolayer distribution of neurons on the growth
surface. The sole method published for the differentiation of murine CGN's did
not fulfill these criteria (Salero and Hatten, 2007). Though we generated mature
neurons, they were often located in tightly wound 3-dimensial bunches or
emanating from highly heterogenous embryoid bodies, rendering many
microscopy techniques for assessing cell identity and viability difficult.
Next, we instead focused our attempts on fulfilling the criteria mentioned above,
namely the generation of more homogenous populations of neurons that would
be spread evenly on the cell culture surface. Despite using neuronal media
without FBS, we often encountered significant populations of replicative non-
neuronal cells overtaking mature post-replicative neurons. Nonetheless, the
second and third differentiation techniques attempted resulted in healthy mature
monolayer neurons at relatively high percentages. However, the ability of these
methods to yield predictable cultures of neurons was low (Appendix Table 1).
Assessing neurotoxicity, whether in primary or in vitro differentiated neuronal
cultures, is not an easy task. Neurons are post-mitotic and thus not subject to
many of the same types of viability assays used for replicative cell lines, such as
102
colony forming assays and growth inhibition. Moreover, large numbers of cells
are needed to get robust differences in sensitivity to drug treatment. One of the
major challenges in assessing neuron viability in heterogeneous cultures is
identifying dead cells as neurons. Based on our work studying primary cerebellar
granule neuron sensitivity to MMS, neurons lose their classifying morphology and
specific immunocytochemical staining patterns upon cell death. However,
primary CGN cultures are nearly -95% homogenous, meaning that most, if not
all, dead cells were originally mature neurons (Chapter IV). In vitro differentiated
cultures rarely provide such homogenous and predictable neural cultures. This is
one possible reason explaining the abundant presence of many disease-specific
in vitro differentiation models aimed at preventing neurodegeneration compared
to differentiation models for the downstream assessment of neurotoxicity
(Kindberg et al., 2014; Visan et al., 2012; Wu and Hochedlinger, 2011; Zeng et
al., 2006). One possible technique to avoid such issues is live-cell imaging.
Neurons could be specifically stained prior to drug treatment and monitored over
time, allowing cell type and sensitivity to be resolved over other cells. The
challenge here is to locate neuron-specific extracellular markers. To date, there
are very few stains that fulfill these requirements (Er et al., 2015; Menon et al.,
2014; Turac et al., 2013). Nonetheless, this is an exciting future direction we are
pursuing.
It should be noted that other in vitro cell models exist for the production of mature
neurons. Often, these replicative cell-lines can be cultured under defined
conditions and mature to differentiated neurons upon the addition or removal of
particular chemicals. Examples of such commonly used cell lines include human
neuroblastoma SH-SY5Y cells or rat PC-12 cells (Barbosa et al., 2015; Biedler et
al., 1973; Greene and Tischler, 1976). More recently, neural stem cell (NSC)
lines, derived from ES cells or replicative areas of the brain, have been adapted
for indefinite monolayer culture while maintaining their multipotent status,
circumventing the need to propagate NSCs as partially differentiated
neurospheres (Conti and Cattaneo, 2010; Conti et al., 2005). Upon the removal
103
of fibroblast growth factor 2 (FGF-2) and epidermal growth factor (EGF), NSCs
differentiate into neurons, astrocytes and oligodendrocytes (Conti et al., 2005).
Thus, another future direction may be to create stable NSC lines from our
genetically altered ES cells for continual derivation of mature neurons.
In the presented work, we differentiated WT and Aag ES cells into
hematopoietic progenitors and neurons in hopes of recapitulating Aag-dependent
transitions in sensitivity to MMS compared to WT. Despite encouraging
preliminary results with hematopoietic differentiation, we focused most of our
efforts of neuronal differentiation. After numerous attempts and multiple methods,
we were still unable to reliably and efficiently differentiate relatively homogenous
populations of neurons for assessment of alkylation sensitivity. In experiments
moving forward, we plan to utilize live-cell imaging methods to identify neurons
prior to treatment and then specifically follow their responses after MMS.
Similarly, we hope to establish NSC lines from our genetically altered ES cell
lines, which will provide a shorter and faster developmental pathway to the
production of mature neurons.
104
Embryonic Stern
Cells
Embryoid Bodies
Hernatopoietic Progenitors
Cerebellar Neurons
Aag-' MMS-Sensitive Aag' MMS-Resistant
Figure 3.1: Mouse embryonic stem cells will be differentiated in vitro into
hematopoietic progenitors and cerebellar granule neurons.
If in vitro differentiation recapitulates in vivo development, then Aag-' MMS-
sensitive ES cells will become Aag-'- MMS-resistant progenitors and neurons.
105
Figures
Embryonic Stem
Cells
Days
Soluble
Factors
Ernbryoid Bodies
12- 14
Hematopoietic
Colonies
24-28
mSCL
IL-3
11-6
P)
C
15 pL Drops
150-375 ES Cells per Drop
I _
- -~ let
0)
0)C U)CU CO
0
100-
80-
60-
40-
20-
Number of Cells in Each Drop
Figure 3.2: In vitro Hematopoietic Differentiation of 129 ES cells.
(A) Protocol schematic for hematopoietic differentiation. ES cells are allowed
to aggregate into embryoid bodies (EBs) and grown for 2 weeks in culture.
At this point, EBs are dissociated and plated in semi-solid media with
cytokines to specifically support the growth of hematopoietic progenitors.
(B) EB formation is induced with hanging drops of ES cells in media. Each
drop is 15 pL and contains between 150-375 cells.
(C) Nearly 90% of hanging drops have formed an EB after two days in culture
independent of genotype.
106
A
B
-T
-Aag-'
1001
Aag-'-
3 10-
Cl)
0.0 0.5 1.0 1.5 2.0
MMS (mM)
Figure 3.3: In vitro hematopoietic differentiation of WT and Aag-' 129 ES Cells
and MMS sensitivity.
VVT and Aag' ES cells were differentiated in vitro to hematopoietic progenitors
and assayed for their sensitivity to MMS. Not enough WT colonies were formed
after 1.5 and 2 mM MMS to accurately determine sensitivity. Errors bars denote
standard deviation from the mean. WT n=2, Aag'l n=3. * p<0.05 using Student's
standard two-tailed T-test.
107
AEmbryonic Stem Emhbrvoid Bodies Neural Lineage Cerehellar lineage lGranule Neurons
ells Precursors
Days
7 12 16 24
Soluble LFF
Factors LF Fgf 8 Fgf Fgf 9 Fgf 9
RA Wntl/3 Wnt]/' Wil/3
Biup6 Bmp6
Bmp7 Bmp7
Gdf7 Gdf7
Slhh
Jagl
BDNF
B C NT,
Figure 3.4:/n vitro Differentiation of Cerebellar Granule Neurons.
(A) Differentiation schematic as adapted from published method (Salero and
Hatten, 2007). ES cells are allowed to form EBs which are exposed
sequentially to cytokines known to impact cerebellar differentiation in vivo.
(B) Phase contrast image of EBs after adherence to laminin coated surfaces
at day 10 of differentiation. EBs are identified by red arrows and neural
cellular outgrowths can be observed.
(C) Representative image of cells differentiated following this protocol at Day
16. Remnants of an embryoid body can be seen in the blurry growth on
the right with neural outgrowth emanating to the left. Blue: Hoechst, Red:
GFAP+ Astrocytes, Green: Tujl+ mature neurons.
108
A
Neura Induction Neurogenesis
Embryonic Stem Neural Lineage Neurons
Cells
Days
1 0 2 10 12 14 21 28
S 9e d E'S C es Iph se
Media____ - -- N2/B27 Neuronal Media - --- -
Media
B
80 W Nestin Positive
Tuj1 Positive
a_ 60- MAP2 Positive
CU
5 40
W 20-
0
Figure 3.5:/n vitro Differentiation of Cortical Neurons.
(A) Differentiation schematic adapted from previously published methods
(Gaspard et al., 2009; Ying et al., 2003a). ES cells are plated at low
density in neural media (N2B27) without serum and allowed to grow for 12
days before dissociation and plating on laminin coated surfaces.
(B) Efficiency of differentiation at Day 14 based on immunocytochemical
staining for neural progenitors (Nestin) and mature neurons (Tujl/Map2).
Error bars denote standard deviation from the mean.
109
Nestin Hoechst Tu Hc
Day 14
Day 21
Day 28
Figure 3.6: Representative images of cells as they differentiate from ES cells into
neurons.
Images in the left column show the presence of neural progenitors as evidenced
by staining for the marker Nestin. As time progresses after Day 14, the few
clusters of cells remaining after dissociation de-differentiate to become
heterogeneous EBs. Images in the column on the right show the presence of
mature neurons as evidenced by staining with the neural markers Tuj1 (green)
and Map2 (magenta). At Day 14, neurons are spread evenly about the surface
but are integrated into large EBs at Day 21 and Day 28. Nuclei in both images
are stained with Hoechst (blue).
110
oechst
301
mr Aag-/-
[~ Aag-/-
20-
) 0 50-
+ 10-
C
a)
z
0
0 0
Dose MMS (mM) Dose MMS (mM)
Figure 3.7: Sensitivity of in vitro differentiated neural progenitors to MMS.
Day 14 in vitro differentiated neural progenitors (Nestin+) were treated with MMS
and assessed for the formation of yH2AX (2 hours) and cellular survival (8
hours). Preliminary results show no differences between WT and Aag- neural
progenitor responses to MMS. 5 images were taken per treatment and at least
1000 cells per treatment were counted.
111
A B
A
® (6)split
Embryonic Stemn Embryoid Bodies Neurons
Cells
8 Days Growth +
Retinoic Acid
B
ES Cells In vitro Differentiated Neurons Primary Neurons
Figure 3.8: In vitro Neuronal Differentiation.
(A) Differentiation schematic adapted from published method (Bibel et al.
2007). ES cells are aggregated into EBs that are dissociated after 8 days
of growth in media containing retinoic acid. Dissociated cells are plated on
laminin/poly-lysine coated surfaces.
(B) Representative images of cells before and after differentiation. ES cells
grown on gelatin form tight colonies with distinct edges. After
differentiation, individual cells containing neural projections can be seen.
These cells resemble primary CGNs isolated from mouse cerebellum.
112
A
(D
0
.
I-
z
E
40-
30-
20-
10-
0
00
000
00 0
00000
*
* 00 AA. ____
'K
'K
B
C
WT ES Cell 11 10 1 33
In vitro Differentiated VVT Neuron 2.17 2 0 7
In vitro Differentiated WT Non-Neuron 6.03 5.5 0 23
Primary WT Neuron 1.21 1 0 5
Figure 3.9: Aag Expression Changes During In vitro Neuronal Differentiation.
(A) Aag expression is downregulated during in vitro differentiation of WT 129
ES cells to neurons and non-neuronal cells. Each point on the figure
113
represents an individual cell. Errors bars represent mean +/- 95%
confidence interval. *** p<0.001 using Student's standard two-tailed T-
test. N 35 cells per category.
(B) Representative image of cells post-differentiation. Large, diffuse nuclei are
from non-neuronal cells (red arrows) while neurons have smaller circular
nuclei and are stained by the neuron marker Map2 (Magenta). Individual
transcripts can be seen as red foci.
(C) Quantification of Aag transcripts in WT ES cells, in vitro differentiated WT
neurons and non-neuronal cells compared to primary VVT cerebellar
neurons.
114
Tables
Table 3.1: Attempts at /n vitro Cortical Neuronal.
115
Attempt # Date Initiated Results
1 12.8.2013 (1/6) Initial experiments done to determine optimal numbers of cells to plate at Day 1 and optimal media
2 12.10.2013 (BI/6) Initial experiments done to determine optimal numbers of cells to plate at Day 1 and optimal media
3 12.31.2013 (1/6) Initial experiments done to determine optimal numbers of cells to plate at Day 1 and optimal media
129/16 cells; differentiation went well Cultures shown to be 60% neural progenitors, 15-20%
4 1.15.2014 neurons at Day 14.
5 5.02.2014 (B11/6) Nothing adhered after splitting
6 5.05.2014 (B11/6) All cells had detached and died by Day 12
7 5.19.2014 (B1/6 & 129) All cells had detached and died by Day 12
Differentiation went well. Neural progenitors were treated with MMS and assessed for y-H2AX
8 7.18.2014 formation
Differentiation went well. Neural progenitors were treated with MMS and assessed for y-H2AX and
9 7.20.2014 cleaved caspase-3
10 9.15.2014 Cells became contaminated
11 9.16.2014 Cells became contaminated
12 9.17.2014 Cells became contaminated
WT cells stopped growing and became large, flat, filled with vacuoles. Aag were looking good so I
13 10.9.2014 continued differentiation for protocol optimization. Split cells at Day 8/12 - nothing adhered
14 10.10.2014 WT cells stopped growing - became filled with vacuoles.
15 10.12.2014 WT cells stopped growing - became filled with vacuoles.
Cells were overgrown but remained on plate, neural rosettes formed, cells adhered after splitting.
16 11.19.2014 Cells on 24-well plate were grown in different cytokine combinations
Cells on 12-well plate were treated with MMS.
17 11.21.2014 Cells were overgrown before Day 12 split and peeled off the plate during routine washings
Differentiation was looking great morphologically (neural rosettes, networks formed), split a few
18 11.25.2014 wells at Day 8: Nothing Adhered... Split the remaining cells on Day 12: Nothing Adhered
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Chapter IV: Ex vivo Mouse Cerebellar
Granule Neuron Sensitivity to MMS is
Dependent on Aag and Parpi
Carrie M. Margulies, Isaac Alexander Chaim, Jennifer J. Jordan,
Aprotim Mazumder, June Criscione, Danielle Wall, Leona D. Samson
Contributions: C.M.M. and L.D.S. designed experiments. C.M.M. conducted all
experiments unless otherwise noted. J.J.J. bred mice and assisted with AlkbhT'~
experiments. I.A.C. developed the HCR assay to measure Aag activity and
helped establish the method in primary neurons. A.M. developed the Aag mRNA
FISH method and applied it to primary neurons. J.C. took ICC images of PAR
staining after MMS treatment. D.W. helped imaged and quantified GFAP+ cells.
121
Chapter IV Table of Contents
Chapter IV: Primary Mouse Cerebellar Granule Neuron Sensitivity to MMS is
Dependent on Aag and Parpi ........................................................................... 123
In tro d u c tio n .................................................................................................... 12 3
Materials and Methods...................................................................................127
R e s u lts ........................................................................................................... 1 3 3
Differential Expression of Aag in Primary Cerebellar Granule Neurons
causes changes MMS Sensitivity ex vivo...................................................133
Aag-Dependent CGN Sensitivity to MMS is mediated through Poly (ADP-
Ribose) Polymerase Activity.......................................................................134
Downstream Modulators of Neuron Sensitivity to MMS.............................136
Base Excision Repair Protein Expression in Primary CGNs......................138
D is c u s s io n ...................................................................................................... 1 3 9
F ig u re s ........................................................................................................... 14 4
A p p e n d ix ........................................................................................................ 16 3
CGN Sensitivity to Glutamate Excitotoxicity and Oxidative Stress is
Independent of Aag Activity........................................................................164
Genetic Deletion of Alkbh7 Reduces Female CGN Sensitivity to MMS.....165
R e fe re n c e s ..................................................................................................... 16 8
122
Chapter IV: Primary Mouse Cerebellar Granule Neuron Sensitivity to MMS
is Dependent on Aag and Parpi
Introduction
Our cells are constantly exposed to DNA damaging agents from our endogenous
and exogenous environments. Moreover, some cancer patients are intentionally
treated with high levels of DNA damaging agents in the form of cancer
chemotherapy. For example, the methylating agent temozolomide (TMZ) is the
current frontline therapy for patients with glioblastoma multiforme, a highly
infiltrative and aggressive brain cancer (Fu et al., 2012b; Stupp et al., 2005).
Though the drug is effective in killing the mitotically active cancerous cells
thereby leading to tumor shrinkage, healthy neighboring cells are simultaneously
exposed to cytotoxic DNA damage. Thus, it is important to understand not only
the responses of cancer cells to treatment with methylating agents, but also the
responses of healthy neurons to minimize chemotherapy side effects such as
'chemo brain' (Simo et al., 2013).
Methylating agents function by covalently adding methyl (-CH 3) groups to cellular
components, including proteins, lipids and importantly, DNA. These DNA lesions
can be mutagenic and cytotoxic if left unrepaired. Fortunately, our cells have
evolved several different DNA repair mechanisms to combat these hazardous
effects. Some repair pathways are relatively simple, involving the direct removal
of the methyl group, such as the methylguanine methyltransferase (Mgmt)
suicide enzyme that repairs cytotoxic 06-methylguanine lesions (Gerson, 2004).
However, other more complex DNA repair pathways involve the coordination of
many proteins in a concerted effort, such as double-strand break (DSB) repair
and base excision repair (BER). The two most common DNA lesions, cytotoxic
and mutagenic 3-methyladenine (3MeA) and the otherwise harmless 7-
methylguanine (7MeG), are repaired by the BER pathway.
123
BER repairs a wide variety of DNA lesions, including alkylated, oxidized and
deaminated bases. It is able to cover such a wide variety of DNA damage due to
a repertoire of cellular glycosylases that find and remove specific damaged DNA
bases, thus initiating the BER pathway (Stivers and Jiang, 2003). Alkyladenine
glycosylase (Aag) is one such protein that recognizes and excises structurally
diverse DNA adducts, including the methyl lesions 3MeA and 7MeG (Wyatt et al.,
1999). Removal of the DNA base through hydrolysis of the N-glycosyl bond
leaves an abasic (AP) site. The apurinic/apyrimidinic endonuclease 1 (Apel)
cleaves the phosphodiester backbone, generating a single-strand break (SSB)
with 3'-OH and 5'-deoxyribosephosphate (5'-dRP) ends. Polymerase B removes
the 5'-dRP end and catalyzes the addition of any removed DNA nucleotides.
Finally, the SSB is sealed together by Ligase I or the XRCC1/ Ligase Ilila
complex, thus completing the repair pathway.
Coordination of BER is tightly regulated since many of the intermediate DNA
structures are toxic if left unrepaired. AP sites and SSBs block transcription and
replication and can lead to the formation of double-stranded breaks (DSBs) in the
context of a passing replication fork (Ensminger et al., 2014). Moreover, SSBs
are bound tightly by poly (ADP-Ribose) polymerase I (Parp1), which catalyzes
the addition of poly-ADP ribose (PAR) polymers to itself and other target
proteins, which include necessary downstream BER components (Sousa et al.,
2012). This catalysis utilizes cellular NAD' as a substrate and Parp1
hyperactivation was hypothesized to cause bioenergetic failure through inhibition
of ATP production and cell death. Parp1 can also cause cell death through an
independent mechanism in which PAR polymers induce mitochondrial release
and nuclear translocation of apoptosis-inducing factor (AIF), thus initiating cell
death through programmed necrosis (Cregan et al., 2002; Hong and Dawson,
2004; Yu et al., 2006).
It is reasonable to hypothesize that a cell lacking the Aag glycosylase would be
sensitive to methylation DNA damage, such as that caused by methyl
124
methanesulfonate (MMS), since cytotoxic DNA lesions would persist unrepaired
throughout the genome. Surprisingly, however, certain Aag' mouse cells and
tissues display remarkable resistance to cell death after treatment with MMS,
where the equivalent dose causes cell death and tissue degeneration in wild-type
(WT) mice (Calvo et al., 2013a; Meira et al., 2009a; Roth and Samson, 2002).
Furthermore, overexpression of Aag in these cell types confers an even more
severe degenerative phenotype after MMS treatment. These results suggest that
initiation of BER through Aag causes cell death and that the original unrepaired
DNA damage is relatively non-toxic in these cell types.
Among these Aag' resistant tissues is the cerebellum, and more specifically the
cerebellar granule neuron (CGN) (Calvo et al., 2013a). The cerebellum is a
region of the brain primarily responsible for coordinating muscle control and the
cerebellar granule neuron is the sole excitatory neuron in this structure. CGNs
are among the smallest neurons in the murine brain and represent nearly 50% of
all neurons, making them the most abundant (Wang et al., 2001). After MMS
treatment, mice display cerebellar neurodegeneration and consequently motor
function impairment in an Aag gene dosage dependent manner, where Aag'
mice are strikingly resistant to neurodegeneration, WT are sensitive and mAagTg
are hypersensitive. Similarly, genetic disruption of poly (ADP-Ribose) polymerase
1 (Parp1) completely rescues mice from cerebellar neurodegeneration after
alkylation treatment even when Aag is expressed at high levels in the mAagTg
mice (Fig. 4.1). Hence CGN sensitivity to MMS is dependent upon the initiation of
BER by Aag and is mediated through the downstream activity of Parp1.
To study the downstream molecular mechanisms of Aag and Parp1-mediated
CGN sensitivity we optimized a method for the isolation and culture of primary
cerebellar granule neurons (CGNs) from post-natal day 5-8 mouse pups. We
show that alkylation induced cell death ex vivo is dependent on Aag and Parp1
activity. These results also establish that cultured CGNs recapitulate in vivo
results and therefore represent a viable system for the study of cell death
125
pathways. We show that this CGN sensitivity to MMS is independent of NAD+
consumption, glycolytic inhibition, and AIF translocation, thus suggesting an as
yet undefined role for Parp1 in DNA damage induced cell death in mouse CGNs.
126
Materials and Methods
Materials and Antibodies
The following materials were purchased from Sigma-Aldrich: Methyl
methanesulfonate (129925), Zvad-fmk (V1 16), Cyclosporin A (30024), BAPTA-
AM (A1076), P-Nicotinamide adenine dinucleotide hydrate (NAD+, N7004), B-
Estradiol (E2758), necrostatin-1 (N9037). Sodium pyrvutate was purchased from
Invitrogen. Parp inhibitors Veliparib and Olaparib were purchased from Selleck
Chem. All cell culture components were purchased from Invitrogen unless
otherwise noted.
The following antibodies were used in this study: GFAP (AbCam, ab7779-500,
1:500), Map2 (AbCam, ab5392, 1:20,000), Tuj1 (AbCam, ab7751, 1:500), PAR
(Trevigen, 4335-MC-100, 1:2,000), AIF (Cell Signaling Tech., 5318, 1:500), and
CoxlV (Cell Signaling Tech., 11967, 1:200).
Isolation of Cerebellar Granule Neurons
Cerebellar granule neurons were isolated as previously described (Lee et al.,
2009b). Neurons were isolation from WT, Aag-', mAagTg and AlkbhT' mouse
pups at post-natal day 5-8. Each cerebellum was isolated and plated separately
and blind to genotype, representing distinct biological replicates. Tails were
collected to genotype at a later time. Cerebella were carefully isolated under a
dissecting microscope in HBSS-glucose (6 g/L) and the meninges and choroid
plexus were removed. The cerebella were dissociated with papain & DNAse
(Worthington Biochemical) for 30-45 minutes at 370 C, after which the tissue was
further dissociated by trituration with a P1000 pipet. Cells were spun down,
resuspended in Earle's Balanced Salt Solution (EBSS, Worthington),carefully
pipetted atop albumin-ovomucoid inhibitor solution (Worthington Biochemical)
and centrifuged for 6 minutes at 70x g. Cells were then resuspended in neuronal
media containing FBA and run through a 70 pm filter to remove large cells and
127
clumps. Remaining cells were pre-plated on poly-d-lysine (PDL) coated 12-well
plates (100 pg/mL) for 2 consecutive 30-minute incubations at 370 C to remove
non-neuronal cells, which adhere readily in the presence of FBS. Cells were
collected, counted, and diluted to 1 x 106 cell/mL in complete neuronal media
(B27/Neurobasal plus 1x L-Glutamine, 1x penicillin/streptomycin, and 250 pM
KCI). Cells were plated on PDL coated dishes (500 pg/mL) at 3,000 cells/mm2 .
For sensitivity assays, cells were plated on optical bottom 96-well plates (Costar
3603) and for immunocytochemistry neurons were plated on glass bottom dishes
(Invitrosci).
All PDL coated plates were incubated for 2 hours at room temperature. They
were then washed twice with water and allowed to dry completely before use.
Determination of Sex and Genotype
Tails were removed from pups during the neuronal isolation procedure and DNA
was isolated to determine genotype. Primers used for determination of Aag
status (null vs. WT) were: (5'- ACC CCG CTT TAC AGA GAA -3', 5'- GCC AAG
ACA GAG ATG AGA CC -3', and 5'- TGG GGG TGG GAT TAG ATA -3').
Primers used for am plifcation of the Aag transgene were: (5'- AGA GAG ACT
CCT GTC GAC CC -3' and 5'- GGT CCT TGA GGG AAC GGC CG -3').
Sex was determined by PCR using primers to amplify TSH-Beta as a control (5'-
AAC GGA GAG TGG GTC ATC AC-3' and 5'-CAT TGG GTT AAG CAC ACA
GG- 3') and SRY to determine sex (5'- AGA GAT CAG CAA GCA GCT CC -3'
and 5'-TCT TGC CTG TAT GTG ATG GC-3').
Treatment of Neurons
Neurons were treated 6-8 days post-plating in neuronal media without B27 for 1
hour at 370 C. Cells were pre-treated with Parp inhibitors (Veliparib/Olaparib) for
1 hour before MMS treatment and Parp inhibitors were included in the media
during and after MMS treatment. Neurons were similarly pre-treated with zVad-
128
fmk (100 pM, 3 hours, left in media during treatment), cyclosporine A (50 pM, 1
hour pretreatment and not left in media post-treatment), necrostatin-1 (50 uM,
left in media post-treatment), BAPTA (20 pM, 1 hour pretreatment only), NAD+
(various doses, 1 hour pretreatment and present during and after MMS
treatment), or P-estradiol (various doses, 3 or 24 hours pretreatment) in complete
media at 370 C.
Determination of Cell Death by Calcein/PI
Cell survival was assessed 24 hours post-drug treatment by staining with
Calcein-AM (Invitrogen, C3099, 1 pM) and propidium iodide (Invitrogen, 20 pM)
for 30 minutes at 370 C. 96-well plates (Costar 3603) were imaged with the
ArrayScan XTI High Content Platform (Life Technologies) and the included
software algorithm was optimization to identify and count live and dead cells from
the green and red fluorescent channels, respectively. Data presented is from
biological replicates, where each data point is the average of 2-4 wells, with each
well imaged 7 times.
JC-1 Mitochondrial Permeabilization
Cells were treated with MMS for 1 hour in serum-free media and JC-1 (3 pg/mL)
was added at different times post-treatment. Mitochondria were stained for 40
minutes at 370 C, stain was removed and fluorescence was immediately
measured with a fluorescent plate reader (VWR SpectraMax ® M3 Multi-Mode
Microplate Reader). Mitochondrial decoupler FCCP (carbonyl cyanide 4-
(trifluoromethoxy) phenylhydrazone; Sigma C2920; 10 pM) was added during JC-
1 incubation and used as a positive control for mitochondrial depolarization.
Immunofluorescence
Neurons were plated in 24- or 96-well glass bottom plates (Invitrosci) and fixed
with 4% paraformaldehyde for 15-30 minutes. Cells were then permeabilized with
0.1% Triton-X in CSK buffer (100 mM NaCl, 300 mM sucrose, 10 mM PIPES, 3
mM MgC 2 , 1 mM EGTA) for 20 minutes, blocked with 4% bovine serum albumin
129
(BSA) for 1 hour at 370 C, and stained overnight at 40 C with appropriate dilutions
of antibodies in 4% BSA in PBS. Secondary antibodies were added for 2 hours at
and neurons were stained with Hoechst for 15 minutes. Images were taken with
a Hamamatsu ORCA-R 2 CCD Camera on a Zeiss Axio Observer.Z1 Microscope.
Neurons stained with anti-PAR were imaged and quantified with the ArrayScan
XTI High Content Platform (Life Technologies). 7 images were taken per well at
1oX magnification and the included software algorithm was used to quantify the
total nuclear fluorescence using Hoechst staining as a mask.
Aag mRNA Fluorescence In Situ Hybridization (FISH)
Neurons were plated on PDL-coated 28-mm glass bottom dishes (Invitrosci) and
fixed 7 days later with 4% paraformaldehyde for 20 minutes at room temperature.
Cells were permeabilized with cold 70% ethanol overnight at 4* C and washed
the following morning for 5 minutes with wash buffer (25 % formamide, 2X SSC).
Neurons were stained with 40 20-nucleotide-long fluorescently labeled (Alexa-
647) oligonucleotides designed against the Aag transcript overnight at 370 C in a
heavily humidified chamber in hybridization buffer (100 mg/mL dextran sulfate,
Sigma, D8906: 0.5 mg/mL E.C01i tRNA, Sigma, R425 .. I mg/mL ssDNA, Sigma
D9156: 1 mg/mL Ultapure BSA, Ambion, AM2616: 10 mM VRC, New England
Biolabs, S1402S: 25% formamide, Ambion, AM9342: 2X SSC, Ambion,
AM9763). Finally, cells were stained with DAPI (2 pg/mL) in wash buffer for 30
minutes at 370 C before imaging. All the previous steps were done in nuclease-
free solutions to avoid RNA degradation.
Images were taken with a Hamamatsu ORCA-R 2 CCD Camera on a Zeiss Axio
Observer.Z1 Microscope. 21 Z-stack images were taken, each 0.3 pm away from
the previous. Images were compiled and foci per cell were counted with a
MATLAB algorithm adapted from previously published methods (Raj et al., 2008).
130
Host Cell Reactivation
The host cell reactivation is based upon the production of fluorescence from an
exogenous plasmid with a site-specific lesion. In the case of Aag, plasmids were
engineered to contain a hypoxanthine (Hx) lesion in the fluorophore codon. In the
absence of Aag, Hx exhibits transcriptional mutagenesis and miscodes for a C,
leading to a fluorescent protein. If Aag repairs Hx, the appropriate U is
incorporated and no fluorescent in exhibited.
Neurons were plated on 96 well plates and transfected with Lipofectamine 3000
(Invitrogen) 6-7 days post isolation according to manufacturer's instructions.
Briefly 2.5 pg of total plasmid DNA were mixed with 2.5 pL Plus reagent and
Opti-MEM, further mixed with 6.2 pL lipofectamine 3000 in Opti-MEM and
incubated at room temperature for 5 min before adding 100 pL of the transfection
reaction to the cells. 5 wells per pup were transfected with damaged or
undamaged plasmids to get as many cell counts as possible. The plate was
immediately placed into the IncuCyte Zoom (Essen Bioscience) live cell
automated fluorescent imager. 4 images per well were taken every hour (phase
contrast and red/green fluorescent filters). Included software was used to identify
cells and quantify integrated cellular fluorescence.
For engineering substrates reporting via transcriptional mutagenesis, non-
fluorescent variants of the different reporter plasmids containing a single
mutation in a site coding for their respective chromophores were identified. In
order to produce ssDNA, we followed previously described methods with minor
modifications (Nagel et al., 2014b). Reporter plasmids were nicked with either
Nb.Bstl or Nt.BstI (New England Biolabs, depending on the lesion containing
strand, see table X). The nicked strand then was digested with exonuclease Ill,
and the remaining single-stranded circular DNA (ssDNA) was purified by using a
1 % agarose gel. 15 picomoles of the respective phosphorylated lesion-
containing-oligonucleotide were combined with 3.2 pg of the corresponding
single-stranded plasmid in 1X Pfu polymerase AD buffer (Agilent
131
Biotechnologies) in a final volume of 46 pL. The mixture was heated to 85 'C in a
thermal cycler for 6 min, and then allowed to anneal by cooling to 40 0C at 1 0C
per minute. To extend the primer, 5 units of Pfu polymerase AD (Agilent) and 0.4
pM dNTP were added. The reaction is then cleaned up with a Qiagen PCR
Purification kit column and eluted in EB buffer and subsequently combined with
1X Ligase Buffer (New England Biolabs - NEB), 0.4 pM dNTP, 1 pM ATP, 1X
BSA, 1.5 units T4 DNA Polymerase and 80 units T4 DNA Ligase (NEB) and
incubated for an additional hour at 16 *C to yield closed circular plasmid. Finally,
the product was purified from a 1 % agarose gel using a Qiagen gel extraction kit.
To test for the presence of Hypoxanthine in GFP-C289T-Hx and its negative
controls (GFP-WT and GFP-C289T), 150 ng of the plasmids were incubated with
10 units ApaLl (NEB) in 1X Cutsmart buffer for 1 hour at 37 *C, followed by 20
min at 65 0C for heat-inactivation. Products were run in a 1% agarose gel for
visualization.
Gene Expression Analysis
Neurons were harvested for RNA isolation and gene expression analysis 1 week
after plating in a 6 cm plate. Neurons were washed twice with HBSS and
harvested by gentle cell scraping. Cells were resuspended in TRIzol and RNA
was isolated with the RNeasy Mini Kit (Qiagen). 1 ug of RNA converted to cDNA
with Omniscript reverse transcriptase following manufacturer's instructions
(Qiagen).
cDNA was measured using a ABI Fast Cycler and the following Taqman probes
(Invitrogen): Pol B (Mm00448234_ml), Apel (Mm01319526_g1), XRCC1
(Mm00494222_ml), Parp1 (Mm01321084), Aag (Mm00447872_ml), Lig3
(Mm00521933_ml) and MAP2 (Mm99999915_g1).
132
Results
Differential Expression of Aag in Primary Cerebellar Granule Neurons
causes changes MMS Sensitivity ex vivo
In order to study the molecular mechanisms of MMS-induced cell death ex vivo,
we needed to verify that neuronal responses to alkylation treatment were cell-
intrinsic properties and not dependent on the environmental context of the
cerebellum, which has very distinct layers and connectivity. To this end, we
optimized a method for the isolation of cerebellar granule neurons (CGNs) from
mouse pups to study the effects of alkylation treatment in individual cells ex vivo
(Fig. 4.2A). Immunocytochemical analyses demonstrated that the resulting
primary cultures were homogenous with minimal contaminating GFAP+
astrocytes (Fig. 4.2B and 4.2C).
Since mature neurons do not replicate, they are not amenable to many traditional
cell sensitivity analyses post-drug treatment. Therefore, we developed a method
to assess neuron sensitivity to drug treatment based on staining with the
fluorescent dyes Calcein-AM (Invitrogen) and propidium iodide (PI), which
independently stain live and dead neurons, respectively. Fluorescent imaging
and live/dead cell counting was conducted with the ArrayScan XTI High Content
platform (Thermo Scientific) (Fig. 4.3). Using this methodology, we confirmed that
CGN sensitivity to MMS is a cell-dependent phenomenon that can be studied ex
vivo. Aag- neurons are resistant to MMS-induced death compared to WT,
whereas overexpression of Aag renders neurons extremely sensitive to MMS
(Fig. 4.4).
We next sought to determine the difference in Aag expression and activity
between the genotypes. We performed mRNA fluorescence in situ hybridization
(FISH) to determine absolute expression levels of Aag mRNA in individual
neurons. Aag expression was significantly different between the three genotypes,
133
with average numbers of transcripts per cell of 0.49, 1.21, and 37.04 in Aag-,
WT, and mAagTg neurons, respectively (Fig. 4.5). Given the near complete MMS
resistance in Aag' neurons, we were surprised to find that the difference in Aag
expression between WT and Aag' was relatively small, but nonetheless
statistically significant. Most striking, however, was the massive increase in Aag
expression in the mAagTg neurons, with up to 100 Aag transcripts in a single
neuron.
We correlated expression of Aag to protein glycosylase activity by using a host-
cell reactivation (HCR) based method. Plasmids with site-specific hypoxanthine
lesions were transfected into primary neurons and fluorescent expression was
monitored over time with the IncuCyte Zoom Live Cell imager (Essen
BioScience). In this experiment, the presence of Aag will lead to reduced
reported expression, while the absence of Aag will lead to increase reporter
expression. We calculated Aag activity overtime and found that after an initial
acclimation period of roughly 20 hours, the percent reporter expression stabilized
within genotypes change figure numbers (Fig. 4.6A). Looking at 24 hours post
transfection, we found significant differences in Aag activity (Fig. 4.6B). The fold
differences n Aag expression between genotypes were remarkably similar tL the
fold changes in Aag activity as measured by HCR. Fold differences between WT
and Aag- Aag expression and activity were 2.5 and 2.05, respectively, while fold
changes between WT and mAagTg Aag expression and activity were 30.6 and
30.42, respectively.
Aag-Dependent CGN Sensitivity to MMS is mediated through Poly (ADP-
Ribose) Polymerase Activity
Parp1 activity is known to modulate the severity of many neurodegenerative
diseases, including stroke and Alzheimer's disease (Martire et al., 2015;
Strosznajder et al., 2012; van Wijk and Hageman, 2005). Moreover, our lab has
previously demonstrated that genetic loss of PARP1 completely rescues
134
cerebellar and retinal degeneration post-MMS treatment, independent of the
activity level of Aag (Calvo et al., 2013a). To test whether PARP modulates
MMS-induced CGN death ex vivo, we used two clinically relevant PARP
inhibitors, Veliparib (AbbVie) and Olaparib (AstraZeneca). Addition of increasing
amounts of either PARP inhibitor rescued neuronal sensitivity to MMS to control
levels (Fig. 4.7A and 4.7B). Similarly, Parp inhibitors conferred significant rescue
of WT and mAagTg neurons as the dose of MMS was increased, indicating that
Parpi activity mediates alkylation induced cell death at low and high levels of
DNA damage (Fig. 4.7C). Surprisingly, Parp inhibitors provided slight rescue
even in Aag neurons, though not to a statistically significant amount. Damaged
DNA bases such as 7-methylguanine or 3-methyladenine spontaneously
depurinate (Drablos et al., 2004; Lindahl, 1993) which can initiate BER
independent of Aag activity leading to Parpi activation; thus Parp inhibitors can
still facilitate MMS rescue even in the absence of Aag. We conclude that Aag-
initiated CGN death post-MMS treatment is facilitated through Parpi activity ex
vivo, further corroborating in vivo results and indicating that CGN response to
MMS is cell-intrinsic.
Parp1 mediates cell death after DNA damage by binding tightly to single-strand
DNA breaks; such tight binding stimulates Parp catalyze the formation of PAR
polymers on itself and other protein targets. Since single-strand breaks are
transiently created during base-excision repair, we anticipate that Aag activity
should correlate with the magnitude of PAR formation after MMS treatment. We
assessed PAR formation during MMS treatment by immunocytochemical (ICC)
staining and saw a large and statistically significant increase in nuclear PAR 15
minutes post-MMS addition in mAagTg neurons which was reduced to basal
levels by 60 minutes post-treatment (Fig. 4.8). There was a slight but non-
significant increase in PAR formation in WT neurons 30 minutes post treatment
and, interestingly, we saw a similar consistent increase in PAR formation 60
minutes post-treatment in Aag~1 neurons. As mentioned above, this PAR
formation is most likely due to spontaneous depurination of damaged bases and
135
reinforces the observation that Parp inhibitors were able to provide slight viability
rescue in Aag-' neurons. The addition of PARP inhibitor Veliparib completely
prevented PAR formation after treatment with MMS in all genotypes and at all
timepoints (Fig. 4.9).
At moderate levels of DNA damage, PAR formation stimulates DNA repair by
recruiting necessary downstream repair proteins; however, PARP hyperactivation
in response to excessive DNA damage is hypothesized to cause cell death due
to excessive consumption of cellular NAD*, leading to bioenergetic failure (Sousa
et al., 2012). Some have reported that addition of exogenous NAD+ or pyruvate
can rescue PARP-mediated neuronal death by bypassing the reduction in
glycolysis and directly supplementing the TCA cycle (Fig. 4.16) (Alano et al.,
2010; Andrabi et al., 2014). However, we found that neither the addition of NAD+
nor pyruvate was able to rescue WT or mAagTg CGN sensitivity to MMS (Fig.
4.10), suggesting that PARP is causing cell death through another pathway.
Downstream Modulators of Neuron Sensitivity to MMS
We next sought to investigate the downstream mechanisms of Parp1 mediated
alkylation sensitivity. Cell death caused by Parpi hyperactivation is considered a
form of programmed necrosis and is often, though not always, executed
independently of caspase activity (Vandenabeele et al., 2010; Zhang et al.,
2012). Indeed, we found that addition of the pan-caspase inhibitor zVad-fmk did
not affect mAagTg CGN sensitivity to MMS (Fig. 4.11 A).
Programmed necrosis can be initiated by numerous different stimuli, resulting in
a variety of cellular outcomes that ultimately conclude in what appears as
necrotic cell death (Berghe et al., 2014; Vandenabeele et al., 2010). The
canonical extra-cellular ligand initiated programmed necrosis (necroptosis) by
tumor necrosis factor (TNF) is mediated through the receptor-interacting protein
kinase 1 (Ripkl) and Ripk3. Necrostatin-1 (Nec-1) is a potent small molecule
136
kinase inhibitor of Ripk1 that was identified by its ability to inhibit necrotic cell
death (Vandenabeele et al., 2013). Pre-incubation of neurons with Nec-1 was
similarly not able to rescue mAagTg sensitivity to MMS (Fig. 4.11 B). This is not
entirely surprisingly given that Ripk1 acts upstream of Parp1 and they represent
two distinct cell death pathways (Sosna et al., 2013; Xu et al., 2010).
Finally, we looked at other known modulators of programmed necrosis. Calcium
fluxes can cause neuronal death through the activation of the calcium-dependent
cysteine proteases calpains, which can cleave mitochondrial AIF leading to its
translocation to the nucleus (Blenn et al., 2010; Cao et al., 2007; Kouzoukas et
al., 2012). However, in our system, there was no effect on cell death when
calcium was chelated to prevent fluxes (Fig. 4.11C). We also queried whether
mitochondrial permeability transition (MPT) was occurring through opening of the
MPT pore (MPTP) since MPT is known to play an important role in neuronal
necrosis in response to excitotoxicity and ischemia-reperfusion injury (Abramov
and Duchen, 2008; Baines et al., 2005; Vaseva et al., 2012). The MPTP sits
within the inner mitochondrial membrane and upon opening, creates a pore
connecting the cytosol and inner mitochondria that is permeable to all molecules
under 1.5 kDa. Inhibition of MPT with cyclosporine A (CsA) did not rescue the
sensitivity of CGNs to MMS (Fig. 4.11 D).
Even though a MPT did not occur, tthis does not preclude the possibility that
mitochondria were becoming depolarized through other mechanisms. We
monitored mitochondrial permeability 1-7 hours post-MMS treatment with the
fluorescent mitochondrial polarization marker JC-1. We did not find any evidence
of mitochondrial depolarization within this timeframe. The positive control,
mitochondrial decoupler FCCP (carbonyl cyanide 4-(trifluoromethoxy)
phenylhydrazone), demonstrated significant mitochondrial depolarization,
showing that depolarization is possible in this cell type (Fig. 4.12).
137
Translocation of apoptosis inducing factor (AIF) from the mitochondria to the
nucleus is often seen in Parp1-mediated alkylation sensitivity and believed to
stimulate cytotoxic nuclear DNA fragmentation (Hong and Dawson, 2004; Yu et
al., 2006). However, we found no evidence of AIF translocation in WT neurons
either 3 or 6 hours post-MMS treatment (Fig. 4.13). Surprisingly, we did observe
the development of an interesting mitochondrial structure at these timepoints,
which have been termed 'donut' or 'blob' mitochondria. These abnormal
mitochondrial shapes have been strongly associated with mitochondrial stress in
many cell types, including neurons after hypoxia-reoxygenation damage (Ahmad
et al., 2013; Liu and Hajnoczky, 2011).
Base Excision Repair Protein Expression in Primary CGNs
We measured the expression of various BER proteins to exclude the possibility
that repair imbalances exist downstream of Aag; such imbalances may contribute
to cell death and Parp activation. As expected, there were large differences in the
expression of Aag but there were no other significant changes between
genotypes in the expression of Apel, Xrccl, Pol 0, Parp1, and Lig3 (Fig. 4.14).
Moreover, Aag expression does not change after MMS treatment, indicating that
the basal Aag expression levels and protein activity are sufficient to cause
significant variability in cell sensitivity to MMS (Fig. 4.15).
138
Discussion
We report the development of a high-throughput fluorescent imaging method for
the assessment of primary mouse cerebellar granule neuron (CGN) sensitivity to
treatment with DNA damaging agents ex vivo. Initiation of the BER pathway by
the Aag glycosylase after methylation DNA damage caused primary CGN cell
death mediated through Parp1 hyperactivation. Pharmacological inhibition of
Parp activity was sufficient to rescue neuronal sensitivity and PAR polymer
formation (Fig. 4.7 and 4.9). Cell death was independent of NAD' and pyruvate
consumption, caspase and Rip1K activity, calcium fluxes, and mitochondrial
depolarization.
We found that inhibition of Parp rescued MMS-induced neuron sensitivity, similar
to that seen in vivo upon genetic deletion of Parp1 (Calvo et al., 2013a). It is
important to note that Parp'- and Parp inhibition do not always cause identical
phenotypes such as in this case. Parp inhibitors, including Veliparib and
Olaparib, function by competing with NAD' for the NAD' binding site within Parp
proteins. Inhibition of Parp1 after binding to single-strand breaks prevents auto-
PARylation and the subsequent release of Parp1 from DNA, thus creating a
Parpl-DNA complex, which in some cases can be converted to a covalent link
(Kedar et al., 2012; Prasad et al., 2014). Such protein-DNA crosslinks can
themselves be cytotoxic by creating an impediment to the transcription and
replication machinery, resulting in double-strand breaks (Heacock et al., 2010;
Horton et al., 2005; Jelinic and Levine, 2014). Olaparib has been shown to be a
more potent creator of Parpl-DNA complexes than Veliparib, yet we found no
difference in MMS rescue between the two inhibitors (Murai et al., 2012)(Fig.
4.8A and 4.8B). Furthermore, double-strand break formation after Parp inhibition
and DNA damage induction is dependent on cells being in S-phase of the cell
cycle (Horton et al., 2005). Given the post-replicative nature of mature neurons, it
is not surprising that Parp inhibition did not exacerbate sensitivity to alkylating
agents.
139
On the other hand, Parp1 hyperactivation is a well-known mediator of
programmed necrosis (Baines, 2010). After excessive DNA damage, Parp1
catalyzes the formation of PAR polymers onto itself and other proteins,
consuming NAD* in the process. It has been postulated that this reduction in
NAD , and subsequent loss of ATP, is sufficient to cause cell death (Alano et al.,
2010). Yet, we found that the addition of NAD+ did not rescue Aag and Parp1
mediated neuronal sensitivity (4.1 1A). Furthermore, recent publications have
demonstrated that this timeline of events is not always the case since ATP
depletion can precede NAD+ loss. In fact, PAR polymers were shown to
translocate to the mitochondria and bind directly to hexokinase (HK), the initiating
enzyme of glycolysis, thus inhibiting its activity (Andrabi et al., 2014; Fouquerel et
al., 2014). This glycolytic inhibition was independent of loss of NAD+ and
glycolysis could be restored through the addition of tricarboxylic acid (TCA) cycle
substrates pyruvate and glutamine (Fig. 4.16) (Andrabi et al., 2014). Again, we
saw no rescue in neuronal viability after MMS treatment upon the addition of
pyruvate (Fig. 4.11B).
PAR translocation to the mitochondria has similarly been shown to cause the
mitochondrial release of AIF (Hong and Dawson, 2004; Yu et al., 2006). AIF
translocates to the nucleus where it interacts with histone H2AX, leading to
chromatinolysis through interaction with cyclophilin A (Artus et al., 2010).
Similarly, there was no evidence of AIF nuclear translocation after MMS
treatment in our cultures (Fig. 4.13). However, it is possible that we did not
choose the relevant timepoint and this failed to capture AIF translocation.
It is widely accepted that the PAR polymers causing these mitochondrial effects
are formed in the nucleus; however it cannot be ignored that Parpi has been
shown to partially localize to the mitochondrial under physiological conditions (Du
et al., 2003; Rossi et al., 2009). Parpl maintains mitochondrial genetic stability
and was identified in a DNA-binding protein complex containing DNA Ligase Ill
140
(Rossi et al., 2009). Mitochondrial specific PARylation was also found on a
variety of other proteins involved in cellular bioenergetics and homeostasis after
oxidative damage (Pankotai et al., 2009). Finally, inhibition of mitochondrial PAR
relieved cell death caused by oxidative damage (Du et al., 2003), suggesting that
the PAR polymers found in the mitochondria may not be derived from the
nucleus and that modulation of mitochondrial protein activity through PARylation
might be more common than expected.
Inhibition of calcium fluxes via calcium chelation did not alter cell sensitivity to
MMS. Increases in calcium concentration occurring during programmed necrosis
are responsible for the activation of the calpains, calcium dependent intracellular
cysteine proteases, that can cleave AIF and the pro-apoptotic protein Bid to tAIF
and tBid, respectively. There are debates in the literature, but under certain
circumstances, calpain activation, not PAR translocation, is essential for the
release of tAIF from the mitochondria after alkylation treatment (Cabon et al.,
2011; Moubarak et al., 2007; Wang et al., 2009). Calcium chelation has also
been shown previously to rescue oxidation but not alkylation induced death
(Bentle et al., 2006). Given the variety of results seen in the literature, it is likely
that the downstream mediators of programmed necrosis are dependent on cell
type and treatment conditions.
Though we found no large-scale evidence of mitochondrial depolarization after
MMS treatment, we did observe the formation of novel 'donut' or 'blob'
mitochondrial structures. This mitochondrial arrangement is associated with
mitochondrial swelling, alterations in mitochondrial fusion dynamics, increased
ROS and overall poor mitochondrial health (Ahmad et al., 2013; Liu and
Hajnoczky, 2011). Additionally, donut shapes only form in individually
depolarized mitochondria (Liu and Hajnoczky, 2011); therefore it is possible that
we missed these events by taking aggregate permeabilization measurements.
Donut mitochondria formation has been documented after ischemia/reperfusion,
oxidative stress, and glutamate excitotoxicity (Gray et al., 2013; Kageyama et al.,
141
2012; Liu and Hajnoczky, 2011; Rintoul et al., 2003; Sharma et al., 2014). Finally,
'donut' mitochondria withstand stress better than tubular mitochondria, possibly
due to their ability to share and mix matrix components more easily, so their
formation may simply be an attempt at survival post-methylation treatment (Liu
and Hajnoczky, 2011). It remains to be seen whether spherical mitochondria
formation is dependent on Aag and/or Parp1 activity.
It is not completely clear, however, whether these mitochondrial structures are
causative of cell death or merely correlated. Mitochondrial fission and fusion
dynamics are important physiological events in healthy cells, particularly in
neurons due to their increased respiratory needs. Diseases characterized by
altered mitochondria fusion/fission often exhibit a neurodegenerative component
and more recently, several neurological diseases have demonstrated altered
mitochondrial dynamics (Alexander et al., 2000; Chen and Chan, 2009; Delettre
et al., 2000; Zuchner et al., 2004). Loss of the mitochondrial fission protein Drpl
in cerebellar Purkinje cells results in large spherical mitochondria with increased
production of ROS due to impaired respiratory function which was blocked by the
addition of the antioxidant N-acetylcysteine (NAC) (Kageyama et al., 2012).
Division dynamics are directly altered after oxidative stress or UxcitotLxAcity
through the cleavage and inactivation of the fusion protein Opal or
transcriptional downregulation of fusion protein Mfn2 (Gray et al., 2013; Martorell-
Riera et al., 2014). Recently, the mitochondrial phosphatase PGAM5 has been
identified as an important component of programmed necrosis signaling
pathways after both extrinsic and intrinsic stimuli (Wang et al., 2012). After
oxidative stress, phosphorylation of PGAM5 leads to the activation of the
mitochondrial fission protein Drpl and subsequent mitochondrial fragmentation
and ultimately cell death. Importantly, inhibition of Drpl activity attenuated cell
death (Wang et al., 2012). Finally, Huntington's disease was similarly attenuated
by inhibition of mitochondrial fission (Guo et al., 2013). These results suggest
that mitochondrial fission/fusion dynamics may be a causative mechanism during
programmed necrosis.
142
Our results demonstrate that ex vivo cultures of primary murine CGNs
recapitulate in vivo Aag- and Parp1- dependent sensitivity to MMS treatment,
thus representing an accurate model within which to study molecular
mechanisms of cell death. We have shown that the cell death is independent of
caspase or Rip1 K activity and there was no evidence of AIF nuclear translocation
or mitochondrial depolarization. Moreover, the addition of exogenous NAD' and
pyruvate was not able to rescue Aag-dependent MMS sensitivity. In future work,
we plan to more closely dissect how mitochondrial fission/fusion dynamics may
be contributing to cell death and whether glycolysis is inhibited in a Parp-
dependent manner.
143
Figures
A WT Aag Parp -
Cerebellum
Untreated
Cerebellum #
150 mg/kg MMS 4
24h
AagTg AagTglParp
~W
0" "F
~J
.4
Figure 4.1: MMS Induced Cerebellar Degeneration in vivo is dependent on Aag
and Parp1 expression.
H&E stained cerebellar sections from WT, Aag<, Parp', AagTg, and
AagTg/Parp' from untreated mice or 24 hours post-MMS treatment. Scale bar is
15 pm. Figure from (Calvo et al., 2013a)
144
A
Dissect out cerebella
Remove meninges
Dissociate cerebella with papain
Plate neurons on poly-D-lysine
coated surfaces
P5-8 pups
B C
U)
0
a)
0
0~
0~
U-
0
10-
8-
4-
2-
0-
n = 10
Figure 4.2: Isolation of Primary Cerebellar Granule Neurons (CGNs).
(A) Neurons were isolated from mouse cerebella at post-natal day 5-8. Each
pup was isolated and processed separately to represent a distinct
biological replicate.
(B) Representative image shows the relative amount of neurons (Map2,
green) and astrocytes (GFAP, red). Nuclei are depicted in blue from
Hoechst staining.
(C) GFAP+ astrocytes were quantified from images from 10 pups isolated on
different days. Error bars represent standard deviation from the mean.
145
A
3-5 Pups Per Genotype 2-4 Wells/Condition 7 Images/Wells
Grayscale Image Object Identification
Figure 4.3: Development of a High-Throughput Method to Determine Primary
Neuron Sensitivity to Drug Treatment.
(A) 3-5 biological replicates per genotype were isolated and plated on 96-well
optical bottom plates. Each treatment condition was applied to 2-4
wells/pup and each well was imaged 7 times at 1OX magnification through
red and green fluorescent filters.
(B) Images were processed by the ArrayScan software which was optimized
to identify and count individual live and dead cells in the green and red
fluorescent channels, respectively. Blue circled objects were accepted by
the algorithm while red circled objects were rejected based on algorithm
criteria.
146
B
Composite
Calcein
(FITC)
Pi
(TRITC)
*100
**
CO)
* *
Aag-1-
-- WT *
-U- mAagTg *
10 .
0.00 0.25 0.50 0.75 1.00
MMS (mM)
Figure 4.4: Sensitivity to MMS treatment ex vivo is dependent on Aag.
mAagTg neurons (blue) displayed enhanced sensitivity to MMS while Aag* (red)
neurons are relatively resistant compared to WT. Errors bars denote standard
error from the mean, Aag-/-: n=6 ,WT: n=5, mAagTg: n=3. * p<0.05, ** p<0.01,
p<0.0001 using Student's standard two-tailed T-test comparing a single dose to
WT.
147
A B
mRNA
WVT
C
Aag Expression in Primary CGNs Aag'
1000
100-
< 10- ~gT< 0 AAA mAagTg
E E MOEN
Figure 4.5: Aag is differentially expressed in primary CGNs from Aag<, WT, and
mAagTg mice.
(A) mRNA fluorescence in situ hybridization (FISH) was used to quantify the
absolute number of Aag transcripts in individual cells.
(B) Representative images of primary neurons from Aag--, WT, and mAagTg
primary neurons. Nuclei are in blue, Aag transcripts in red, and cell
boundaries depicted in green (WGA).
(C) Transcripts were counted per cell and are plotted in, where each data
point represents one cell. Errors bars denote standard deviation from the
mean, n > 50 cells, *** p<0.001 using Student's standard two-tailed T-test.
148
100-,
10-
- V
-n7AagTg
Aag
0 20 40
Hours Post-Transfection
Figure 4.6: Aag Glycosylase Activity is Significantly Different between Aag-'-, WT,
and mAagTg primary CGNs.
(A) Glycosylase activity was monitored by over time by cellular fluorescent
output after transfection with a site-specific hypoxanthine containing
reporter plasmid (See methods for detailed explanation). Decreased
reporter expression indicates an increase in Aag activity. Solid lines
denote mean while dashed lines indicate standard deviation from the
mean.
(B) WNT, Aag' and mAagTg neurons have significantly different Aag activity at
24 hours post-transfection. Errors bars denote standard deviation from the
mean. WT n=3, mAagTg n=2, Aag' n=2. ** p<0.01 using Student's
standard two-tailed T-test)
149
A B
0
C,)
U)
X
0
80
r'Th
60-
40-
20-
C/)
(D)
uLJ
-C
0
0
60
0- ~1~~~~l' . . -
Aag-
TW
Control 10- 10 1 1-
Olapan b M)
100- 1 1001
Aaq
Aag +- Velipa 
b
0.00 0 M 00 5 1 b0
MIMS (mM)
-- WVT
101 -1- WVT + Velparb
000 025 0 50 075 1 00
MMS (mM)
1001
-+r mAagTg
--- mAagTg Veliparib
10
0.00 0 25 050 0,75 1.00
MMS (mM)
Figure 4.7: Primary CGN Sensitivity to MMS is dependent on Parp1 activity.
Aag-, WT, and mAagTg neurons rescued from MMS toxicity after pretreatment
with Parp inhibitor Veliparib (A) or Olaparib (B) (1 mM MMS). (C) Parp inhibitor
Veliparib rescues cell sensitivity at all doses of MMS in WT and mAagTg
neurons. Errors bars denote standard error from the mean, Aag<: n=6 ,WT: n=5,
mAagTg: n=3. * p<0.05, ** p<0.01, *** p<0.0001 using Student's standard two-
tailed T-test comparing sensitivity with or without Veliparib at a particular dose of
MMS.
150
A
100-
U)
B
/K
Aag-g
-,r mAagTg
TZ
1.O
Control 10- 10 10-6
Veliparib (M)
10 10
C
U'2
A MMS Treated B
15 Mins 30 Mins 60 Mins Fold Change in Nuclear PAR Formation
Post MMS Treatment
Aag^ CGNs
20- Aagl
-M- V\/T
-A- mAagTg
1 5-
mAagTg CGNs C
0 100
0 20 40 60
Time Post MMS Treatment (Minutes)
Figure 4.8: PAR formation post-MMS treatment (1 mM) is dependent on the
expression level of Aag.
(A) PAR formation (green) was visualized in Aag , WT, and mAagTg neurons
0, 15, 30, or 60 minutes after the addition of MMS by immunocytochemical
staining.
(B) Elevation in nuclear PAR was quantified and reflects qualitative trends in
(A). Errors bars denote standard error from the mean. Average nuclear
fluorescence from each biological replicate represents aggregation of
>1000 nuclei. Aag-: n=4 ,WT: n=3, mAagTg: n=2. ** p<0.01 using
Student's standard two-tailed T-test comparing nuclear fluorescence at a
particular timepoint compared to untreated cells.
151
A Untreated MMS Treated (1 mM)
15 Mins
mAag CGNs mAag CGNs mAag CGNs
Veliparib + Veliparib
B Green = PAR Staining
2,0- 20- 2 0-
Aag" -0- WV -o- mAagTg
0 Aag- + Veliparb W VVT + Veliparib mAagTg + Velipanb
UI) I) Cl)
a_ a. 0.
11 5-
n.s.
z z z
CeCO Ce
0) 0)0
104 10 10
0 20 40 60 0 20 40 60 0 20 40 60
Time Post MMS Treatment (Minutes) Tme Post MMS Treatment (Minutes) Time Post MMS Treatment (Minutes)
Figure 4.9: Parp inhibitor Veliparib inhibits PAR formation after MMS treatment.
(A) Qualitative images of PAR formation in mAagTg neurons either left
untreated, or 15 minutes after MMS treatment with or without the addition
of Veliparib (5 pM).
(B) Quantification of changes in nuclear PAR formation in Aag, WT, or
mAagTg neurons with or without Veliparib. mAagTg neurons
demonstrated an immediate and significant increase in PAR formation 15
minutes after MMS addition that was significantly inhibited upon the
addition of Parp inhibitor. Errors bars denote standard error from the
mean. Average nuclear fluorescence from each biological replicate
represents aggregation of >1000 nuclei. Aag->: n=4 ,WT: n=3, mAagTg:
n=2. * p<0.05 using Student's standard two-tailed T-test comparing
nuclear fluorescence with or without Veliparib at a particular time post
MMS treatment.
152
BgT
mmAagTg
CO
100-
80-
60-
40-
20-
100-
80-
60-
40-
20-
c,, x x x x x
C\,' ~
mr
mmAagTg
x)) x x
q \ q
<0~
Figure 4.10: Neither supplementation of NAD' nor pyruvate rescues WT or
mAagTg primary CGN sensitivity to MMS.
(A) Neurons were preincubated with NAD' for 3 hours in complete neuron
media before addition of 1 mM MMS. NAD' was included in the MMS
treatment and during recovery. NAD' treatment alone did not reduce
neuron viability (results not shown). There was no evidence of sensitivity
rescue in either genotype. Errors bars denote standard deviation from the
mean. WT n=4, mAagTg n=3.
(B) Neurons were pretreated with pyruvate for 3 hours in complete neuron
media before addition of MMS (1 mM). Veliparib significantly rescued
neuron sensitivity while the addition of pyruvate played no effect. Errors
bars denote standard deviation from the mean. WT n=5, mAagTg n=1. ***
p<0.001 using Student's standard two-tailed T-test comparing MMS and
MMS + Veliparib.
153
A
I. in inin.I
(I)
fl -IM-
A B C D
Caspase Inhibitor Rip k nhibitor Caic um Cheator MPTP nhibitor
100-
50-
0.1 0
0\; K
100-
50-
100-
50-
Figure 4.11: Downstream molecular mechanisms of CGN Aag-dependent MMS
sensitivity.
Primary mAagTg CGN sensitivity to MMS is not dependent on caspase activation
(A), Rip1 kinase activity (B), calcium fluxes (C), or mitochondrial permeability
transition (MPT) (D). In each experiment, Veliparib was included to demonstrate
cell death dependence on Parp activation. Inhibitors used include (A) zVad-fmk,
n=6, (B) necrostatin-1, n=3, (C) BAPTA-AM, n=3, and (D) cyclosporin A, n=3.
Errors bars denote standard error from the mean. *** p<0.001 using Student's
standard two-tailed T-test comparing MMS to MMS+Veliparib.
154
CO
100-
50
0-
X)
Loss of JC-1 Red Fluorescence After MMS Treatment
100
0
0)-
Aag-
-e- VVT
-*- mAagTg
Control 1 Hr 2 Hr 3 Hr 4 Hr 5 Hr 6 Hr FCCP
Time Post 1 mM MMS Treatment
Figure 4.12: There is no loss of mitochondrial permeability in Aag', WT, or
mAagTg neurons 1-7 hours after MMS treatment.
Cells were treated with MMS for 1 hour and then assayed for mitochondrial
depolarization using the fluorescent dye JC-1, measuring by fluorescent plate
reader. Co-incubation of cells with JC-1 and the mitochondrial decoupler FCCP
as a positive control demonstrated significant loss of fluorescence. Errors bars
denote standard deviation from the mean. Aag- n=2, WNT n=3, mAagTg n=2. **
p<0.01 using Student's standard two-tailed T-test comparing FCCP to control. *
p<0.05 using Student's standard one-tailed T-test comparing FCCP to control.
155
MMS Treated (1 mM)
3 Hours 6 Hours
Figure 4.13: No evidence of AIF nuclear translocation after MMS treatment.
WT CGNs treated with MMS do not show evidence of AIF (red) translocation
from the mitochondria (green; CoxIV) to the nucleus (blue; Hoechst). Instead, the
mitochondria adopt an unusual blob structure after treatment associated with
stress and dysfunction. 63X Magnification.
156
Untreated
15-
10
5-
0
I
D Aag-'- CGN
VCGN
mAagTg CGN
.1 ~ - - - -
, N
Figure 4.14: Expression of BER genes in primary CGNs.
Expression of Base Excision Repair genes demonstrates that Aag is the only
gene differentially expressed between Aag<, WT, and mAagTg primary neurons.
Errors bars denote standard deviation from the mean, Aag- n=4, WT n=5,
mAagTg n=1.
157
C-0
U)
U)
0)
LUJ
a)
1 0\
I
QPN 4 C)
MMS
(0.75 mM
1 hr
t = -1
Complete
Media
1 hr
t = 0
B
z
tX
E
t = 1
1.2-
1.0
0.8-
0.6
0.4
0.2-
0.0
-J Mean
Control MMSOhr MMS1hr
I
Figure 4.15: Aag expression is not induced after MMS treatment in WT neurons.
(A) Aag expression was assessed either before, immediately after, or an hour
after MMS treatment.
(B) There was no difference in Aag expression at any timepoint. Errors bars
denote standard deviation from the mean.
158
A
glucose
Glycolysis
pyruvatePDH
PC Acetyl-CoA
oxaloacetate citrate
TCA Cycle
a KG
GDH glutamate glutamine
glutaminase
Figure 4.16: Glycolysis and Tricarboxylic Acid (TCA) Cycle Schematic.
Cells can generate energy through the enzymatic breakdown of glucose during
glycolysis, which results in the formation of pyruvate that is shuttled into the TCA
cycle after conversion to Acetyl-CoA. Glutamine can be broken down during
glutaminolysis to glutamate which can also supplement the TCA cycle after
conversion to a-ketoglutarate (aKG). PC, pyruvate carboxylase; PDH, pyruvate
dehydrogenase. Figure adapted from (Newsholme et al., 2013).
159
100- 12 Aagf
M WT
mAagTg
C) 50-
0
4% C
Figure 4.17: Aag activity does not contribute to ex vivo sensitivity to glutamate
excitotoxicity.
Errors bars denote standard deviation from the mean. N=3 for all genotypes.
160
100- U U A
inWT
mAagTg
CO)
50
01
&g ov r r rn
C,
N'
Figure 4.18: Aag does not contribute to ex vivo CGN sensitivity to treatment with
hydrogen peroxide (H 2 02).
Errors bars denote standard deviation from the mean. WT n=5, mAagTg n=1.
161
WT Male
WT Female
Alkbh7' Male
Alkbh7-- Female
100-
Dose MMS (mM)
Figure 4.19: Female A/kbh7-CGNs are relatively resistant to MMS treatment ex
vivo compared to male Alkbh T', female WT and male WT neurons.
Error bars denote standard error from the mean. WT male n=6, WT female n=10,
AlkbhT' male n=12, Alkbh7 female n=6.
162
MJ
CD
MI
75-
5 50-
C')
25-
0-
I
.4.
0 0.75
I
10.5
I am
I
A B
10
8
6
m VT Male
IliVVT Female
| Alkbh7-Male
|J AIkbh7-l- Female
(U
0
0-
0-
0-
0
C-
10
8
6
4
2
**- *
intill
x.
Figure 4.20: WT and AlkbhT CGNs are rescued from MMS sensitivity by Parp
inhibition but not by Caspase inhibition.
(A) AlkbhT' and WT primary CGNs are not rescued from MMS sensitivity by
treatment with the pan-caspase inhibitor zVad-fmk. Error bars denote
standard error from the mean. WT male n=1, WT female n=3, AlkbhT'
male n=4, AlkbhT A female n=2.
(B) Parp inhibitor Veliparib rescues AlkbhT' and WT primary CGNs from
MMS toxicity. Error bars denote standard error from the mean. WNT male
n=6, WT female n=10, AlkbhT' male n=12, AlkbhT ' female n=6.***
p<0.001 using Student's standard two-tailed T-test comparing MMS to
MMS+Veliparib for each genotype and sex.
163
0-
0-
0-
0-
0
CO)
imn
A
Appendix
CGN Sensitivity to Glutamate Excitotoxicity and Oxidative Stress is
Independent of Aag Activity
Ischemia reperfusion (l/R) injury is characterized by the acute loss of blood flow
and consequently reperfusion of blood, oxygen, and other components into the
affected area. /R injury often leads to necrosis of the injured tissue and is
thought to be mediated by the formation of DNA damage caused by two main
sources: excitotoxicity induced by overactivation of glutamate receptors
(explained below) and the generation of reactive oxygen species (ROS) during
reoxygenation. Recent work from our lab has shown that Aag' mice are resistant
to cerebral ischemia-reperfusion injury (IRI) compared to WT mice (Ebrahimkhani
et al., 2014). Thus, we sought to determine whether the expression of Aag in ex
vivo CGN cultures dictated sensitivity to glutamate excitotoxicity or ROS.
During the hypoxia stage of I/R injury, excitotoxicity occurs as a consequence of
neurons becoming depolarized, leading to increased cellular uptake of Ca2+ and
the simultaneous release of glutamate and impaired glutamate re-uptake (Choi
and Rothman, 1990). It is the calcium overload that is particularly hazardous to
the cells; Ca2+ leads to the activation of
nitric oxide synthase (nNOS) (Dawson
Samdani et al., 1997b). nNOS generates
superoxide to form reactive oxygen and
DNA (Beckman and Koppenol, 1996).
excitotoxicity from /R injury or treatment
activity, supporting the notion that DNA
death (Andrabi et al., 2011; Mandir et
Zhang et al., 1994). Yet, we found no
numerous proteins, including neuronal
et al., 1996; Samdani et al., 1997a;
nitric oxide (NO) radicals that react with
nitrogen species (RONS) that damage
Moreover, neuronal death caused by
with nitric oxide is mediated by Parp1
damage is the ultimate cause of cell
al., 2000; Szabo and Dawson, 1998;
differences in sensitivity to glutamate
excitotoxicity between Aag, WT, and mAagTg neurons (Fig. 4.17).
164
Tissues undergoing I/R are exposed to high levels of reactive oxygen species
(ROS) during reoxygenation due to the rapid influx of blood and concurrent
inflammatory response (Eltzschig and Eckle, 2011). Though Aag was identified
as a glycosylase for alkylated bases, the mouse protein has been shown to bind
and excise 8-oxoguanine (8-oxoG), which is formed under conditions of
inflammation and oxidative stress (Bessho et al., 1993; Wyatt et al., 1999). Yet,
we found that the level of Aag activity did not affect cellular sensitivity upon
treatment with the ROS agent hydrogen peroxide, H2 0 2 (Fig. 4.18).
Even though Aag activity did not influence primary CGN sensitivity to glutamate
excitotoxicity or ROS ex vivo, this does not preclude the already identified role for
Aag in vivo after ischemia-reperfusion. /R is a highly complex injury involving
specific phases of hypoxia and reperfusion that may not be accurately
recapitulated ex vivo after treatment with glutamate of hydrogen peroxide.
Genetic Deletion of Alkbh7 Reduces Female CGN Sensitivity to MMS
Recent work from the lab has identified the human AlkB homolog 7 (ALKBH7)
protein as necessary for alkylation induced programmed necrosis (Fu et al.,
2013). Knockdown of ALKBH7 rendered cells resistant to necrosis induced by
the methylating agents MMS, temozolomide, and MNNG. NAD* and ATP levels
dropped in both WT or ALKBH7-depleted due to PARP hyperactivation post-
treatment, yet only ALKBH7-depleted cells were able to recovery NAD* and ATP
levels to basal levels, thus maintaining mitochondrial polarization and overall
viability.
Given that in vivo cerebellar degeneration post-MMS treatment proceeds via a
Parp1-dependent programmed necrotic pathway, experiments were conducted to
determine whether Alkbh7 modulates neurodegeneration in mice. Surprisingly,
we found that Alkbh7 exerts rescue in a sex-specific manner. Specifically,
AIkbh7' male mice were preferentially rescued from MMS-induced cerebellar
165
degeneration after a high dose of MMS compared to WT male, WT female and
AlkbhT' females.
To study the alkylation induced cerebellar degeneration at a cellular level, we
isolated primary neurons from male and female wild type and AlkbhT' pups.
Initial experiments were aimed at determining whether ex vivo cultures of CGNs
accurately recapitulated adult in vivo responses to MMS. Surprisingly, our ex
vivo results contradicted in vivo trends in MMS induced neuronal sensitivity. We
found that in ex vivo CGN cultures, female AlkbhT' neurons were resistant to
MMS induced cell death compared to WT male, WT female or AIkbhT' male
neurons (Figure 4.19). These results are intriguing given that ex vivo CGN MMS
induced sensitivity does mimic in vivo results demonstrating a dependence on
Aag and Parp1 activity, suggesting that Alkbh7 may be differentially expressed or
function in distinct roles in adult compared to immature neurons.
Both Alkbh7 and Parp1 have been shown to play a role in programmed necrosis
downstream of alkylation damage where genetic loss of Parp1 completely
rescues cerebellar degeneration in vivo after MMS treatment (Berghe et al.,
2014; Calvo et al., 2013a; Fu et al., 2013). We next sought to determine whether
Parp1 acts upstream or downstream of Alkbh7 by modulating Parp activity with
the clinical inhibitor Veliparib. Veliparib rescued cell death in all genotypes and
sexes, albeit to different extents (Figure 4.20B). Inhibition of Parp preferentially
rescued WAT male neurons from MMS-induced death compared to VVT female
neurons. These results agree with previous reports demonstrating that male
mice, unlike female mice, are rescued from ischemia-reperfusion injury by Parp
inhibition or genetic deletion (Du et al., 2004; Hagberg et al., 2004).
Finally, we investigated the role of caspases in alkylation induced neuronal
death. Addition of the pan-caspase inhibitor zVad-FMK prior to treatment with
MMS did not rescue neuron sensitivity in any of the samples, indicating that
alkylation induced neuronal death is caspase-independent (Fig. 4.20A). These
166
results were surprising given the suggestion that male neurons prefer to undergo
AIF mediated programmed necrosis while female neurons preferentially undergo
caspase-dependent apoptosis (Du et at., 2004).
167
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Chapter V:
178
Discussion
Table of Contents
Chapter V: Discussion ....................................................................................... 180
D is c u s s io n ...................................................................................................... 1 8 0
Imbalances in DNA Repair alter Embryonic Stem Cell Sensitivity to
A lky la tin g A g e nts ........................................................................................ 18 0
Aag and Parp1 Mediate Cerebellar Granule Neuron Sensitivity to MMS ... 184
Aag-Dependent Cell-Specific Responses .................................................. 187
R e fe re n c e s .................................................................................................... 1 8 9
179
Chapter V: Discussion
Discussion
Imbalances in DNA Repair alter Embryonic Stem Cell Sensitivity to
Alkylating Agents
In the presented work, we investigated how alterations in base excision repair
initiation by the Aag glycosylase affects cell sensitivity upon treatment with
alkylating agents. We began by generating new embryonic stem cell (ES) lines
from WT, Aag' and transgenic mAagTg female mice and demonstrating their
pluripotency. We found that genetic background (129 vs C57B1/6) played a
minimal effect on Aag-dependent phenotypes and our results agreed with
previous publications showing that loss of Aag in ES cells causes an increase in
cell sensitivity to alkylating agents (Allan et al., 1998; Engelward et al., 1998;
Engelward et al., 1996; Maor-Shoshani et al., 2008). Moreover, results generated
from cells overexpressing Aag have extended our understanding of DNA repair
coordination in stem cells. Interestingly, we found that methylation sensitivity had
a more complex relationship with Aag gene-dosage in that overexpression of Aag
caused higher sensitivity than loss of Aag compared to WT (mAagTg > Aag* >
WT). To our knowledge, this is the first documented case where both loss and
enhancement of DNA repair initiation has caused increased cell sensitivity to
DNA damage compared to WT in mammalian cells and provides a unique model
to study DNA repair capacity and cellular toxicity. Interestingly, a similar Aag-
dependent phenotype was previously published in E. coli upon the loss or
increased expression of the AIkA DNA glycosylase, which excises similar
substrates as Aag (Kaasen et al., 1986).
Though ES cells have more robust and efficient DNA damage repair compared to
differentiated cells (Tichy et al., 2011; Tichy and Stambrook, 2008), they also die
at a higher rate after treatment with DNA damaging agents, including gamma
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irradiation and MNNG (de Waard et al., 2008; Roos et al., 2007). These
particular cell responses are hypothesized to be due to the pluripotent nature of
stem cells, which will give rise to all daughter cells and thus strive to avoid
mutations and maintain genetic stability. Similar to replicative somatic cells, stem
cells can halt cell cycle progression to allow for DNA repair prior to mitosis. It is
widely accepted, however, that mouse ES cells lack a G1 cell cycle checkpoint
and rely primarily on S or G2 phase checkpoints (Aladjem et al., 1998). This fact
may contribute to enhanced apoptotic response in ES cells given that at any
time, -75% of ES cells are in S-phase (Savatier et al., 2001). A more recent
publication suggests that ES cells do maintain an active G1/S checkpoint that is
regulated by ES cell-specific miRNAs of the miR-290 family (Wang et al., 2008).
Nevertheless, the highly replicative nature of ES cells most likely contributes to
their sensitivity to DNA damage.
How does enhanced DNA repair initiation cause more cell death than a lack of
DNA repair? We hypothesize that mAagTg and Aag' ES cells sense a higher
level of DNA damage than WT cells after identical treatment doses due to
increases in replication stress. Aag" cells lack the ability to remove the
replication blocking DNA lesion 3MeA and subsequent repair through BER.
Previous work from the lab has demonstrated that these lesions are converted
into cytotoxic double-strand breaks (DSBs), some of which are repaired by
homologous recombination (HR) during S phase as evidenced by an increase in
sister chromatid exchanges in Aag' cells compared to WT (Engelward et al.,
1998; Hendricks et al., 2002). In the situation where Aag is overexpressed, both
cytotoxic 3MeA lesions and otherwise innocuous 7MeG lesions are shuttled
through BER (Rinne et al., 2005). However, given the overabundance of Aag
compared to downstream proteins, AP sites and single strand breaks will
accumulate and cause DSBs during replication. mAagTg cells will perceive
higher levels of DNA damage than Aag' due to BER processing of both 3MeA
and 7MeG, whereas in Aag' cells only 3MeA lesions will cause replication
stress. WT cells, on the other hand, likely express appropriate amounts of each
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of the BER proteins to maintain genomic stability. Experiments are ongoing to
test this hypothesis by examining SCE formation and the generation of SSBs and
AP sites after MMS treatment. It is also surprising that such a large increase in
MMS sensitivity is seen in mAagTg cells given that they only express roughly 4
times as much Aag as WT cells (Fig. 2.5A). It remains to be seen whether there
is an Aag expression threshold above, or below, which MMS sensitivity is
changed compared to WT, or whether the relationship between Aag activity and
sensitivity is linear and proportional.
It will also be important to assess the timecourse of cell death post-MMS
treatment as it is entirely possible that Aag", WT, and mAagTg ES cells are
dying via different mechanisms. Interestingly, both WT and Aag' ES cells treated
with MMS exhibited a maximal percentage of apoptotic cells 75 hours post-
treatment, which correlates to roughly 6 cell cycles assuming a 12 hour cycling
time (Engelward et al., 1998; Engelward et al., 1996). On the other hand,
treatment of cells with Me-Lex, a 3MeA specific alkylating agent, caused cell
death that reached a plateau as early as 20 hours post-treatment (Engelward et
al., 1998). These results indicate that 3MeA is most likely causing cell death
during the first S-phase due to replication blockage. However, the greater variety
of cellular lesions generated by MMS appears to affect cell sensitivity through
other mechanisms that may or may not depend on the absolute amount of Aag
present. It may be the case that mAagTg cells are not dying due to replication
stress caused by BER intermediates, but through more complex downstream
mechanisms in later cell cycles. It was recently shown in human cells that AAG
and APE1 activity are essential for the' phosphorylation and activation of the
important DNA damage response kinases ataxia-telangiestasia mutated (ATM) at
S1981 and checkpoint kinase 2 (CHK2) at T68 (Chou et al., 2015). If this
signaling component remains valid in mouse ES cells then we would expect to
see robust Atm/Chk2 activation only in WT and mAagTg cells, but not Aag.
Additionally, ATM was also shown to regulate and increase Aag activity through
phosphorylation at S172, suggesting that ATM may play an important role in
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regulating BER function along with subsequent cellular consequences (Agnihotri
et al., 2011).
In addition to replication stresses, another possibility is that downstream DNA
damage signaling responses may differ between genotypes after MMS
treatment. Aag-' ES cells have been shown to induce a more robust induction of
p53 after MMS treatment than WT cells (Engelward et al., 1998). It has also been
shown, however, that p53 does not translocate to the nucleus efficiently in ES
cells post-DNA damage despite robust stabilization (Aladjem et al., 1998).
Recently, p53 translocation was shown to be dependent on the deacetylase
Sirt1. Upon DNA damage, Sirt1 deacetylates p53, causing translocation to the
mitochondria, not the nucleus, to enact apoptosis (Han et al., 2008).
Nevertheless, some p53 does enter the nucleus of ES cells to alter
transcriptional response after DNA damage. p53 has been shown to directly
suppress transcription of the pluripotency regulating transcription factor Nanog
(Lin et al., 2004). Furthermore, not only does p53 reduce expression of
numerous genes associated with pluripotency, it simultaneously promotes
expression of genes linked to differentiation (Li et al., 2012). Moreover, the N-
terminal region of human Aag has been shown to directly inhibit p53's ability to
bind to DNA and serve as a transcription factor in unstressed cells. Interestingly,
binding of Aag to p53 selectively inhibited transcription of the pro-cell cycle arrest
genes p21, 14-3-3y, and Gadd45 but had no effect on pro-apoptotic gene
expression (Puma, Noxa) (Song et al., 2012). Overexpression of Aag reduced
expression of target genes while depletion of Aag increased expression. These
results suggest the possibility that Aag itself may play a role in modulating p53
during the DNA damage response.
Finally, though all the ES cells lines are derived from inbred 129 mice and should
be isogenic except for variations in Aag expression, we cannot rule out the
possibility that there are gene expression changes between cell lines that alter
sensitivity to DNA damage. Each ES cell line was derived from an individual
183
blastocyst and therefore experienced a slightly different derivation process that
could affect genomic expression or epigenetic patterns. Though we ruled out the
possibility of inherent expression differences in BER transcripts, there are
numerous other genes, or combinations of genes, that upon differential
expression might affect responses to MMS.
Aag and Parp1 Mediate Cerebellar Granule Neuron Sensitivity to MMS
Here we've shown that, unlike ES cells, Aag causes cerebellar granule neuron
(CGN) sensitivity to MMS in a gene-dose dependent manner where loss of Aag
leads to complete rescue from sensitivity to MMS. Furthermore, Parp1
hyperactivation mediates this cell death through binding to single-strand breaks
generated during imbalanced BER initiated by Aag. Pharmacological inhibition of
Parp1 completely rescued sensitivity of WT and Aag overexpressing neurons.
These results mimic in vivo work and thus represent a viable model within which
to study downstream molecular mechanisms of programmed necrosis. This cell
death was not rescued by NAD' or pyruvate supplementation and was found to
be independent of caspase activity, mitochondrial depolarization, and AIF
translocation.
We showed that two different clinically relevant Parp inhibitors, Olaparib and
Veliparib, were able to rescue neuron sensitivity to MMS ex vivo to the same
extent. Moreover, recent unpublished work from our lab demonstrated that both
these inhibitors were able to rescue cerebellar degeneration post-MMS treatment
in vivo. Interestingly, however, investigation into published literature revealed that
Olaparib has been shown to be a substrate for the multidrug resistance protein 1
(Mdrl) efflux pump located on the blood brain barrier and is therefore excluded
from the central nervous system of mice and rats after a single bolus dose
(Chalmers et al., 2014); Veliparib, on the other hand, was shown to efficiently
cross the blood-brain barrier in mice (Donawho et al., 2007). Nevertheless,
Olaparib provided rescue from alkylation-induced cerebellar degeneration
184
indicating that the drug was crossing the blood brain barrier. These results
suggest that Parp inhibitors may minimize DNA damaging cancer chemotherapy
side effects found in the cerebellum. Indeed, magnetic resonance imaging (MRI)
has revealed that during cancer chemotherapy the cerebellum has significant
loss of gray matter, which can contribute to symptoms of 'chemobrain'
(McDonald et al., 2010; Simo et al., 2013).
The ability of NAD* supplementation to rescue Parp1 mediated programmed
necrosis is debated. Though Parp1 hyperactivation and subsequent cell death
has been attributed to NAD* consumption and bioenergetic death for decades,
not all publications support this hypothesis. Nearly 50% of cellular NAD+ is stored
in the mitochondria in mouse neurons and Parp1 activation post-methylation
DNA damage specifically consumes the cytosolic portion, leaving the
mitochondrial portion unaltered unless depolarization occurs (Alano et al., 2007).
In some situations, NAD* depletion without Parp1 activation has been sufficient
to cause programmed necrosis in neurons, complete with AIF nuclear
translocation and mitochondrial depolarization (Alano et al., 2010). Parpl-
mediated cell death has also been shown to cause an inhibition of glycolysis,
thus NAD+ depletion was thought to be the culprit (Ying et al., 2005; Ying et al.,
2003b). More recently, however, two independent publications have
demonstrated that NAD+ loss is not sufficient to cause glycolytic inhibition and
cell death (Andrabi et al., 2014; Fouquerel et al., 2014). Furthermore, they
identified the initiating enzyme of glycolysis, hexokinase-1 (HK1), as a PAR-
binding partner. Nuclear PAR polymers formed in response to DNA damage
were shown to translocate to the mitochondria where they directly bind HK1,
thereby inhibiting its activity independent of, and prior to, NAD+ loss (Andrabi et
al., 2014; Fouquerel et al., 2014). However, these results conflict with the
observation that NAD+ repletion can rescue glycolytic inhibition and Parp1-
mediated cell death since NAD' depletion should be a consequence and not a
cause of suppressed glycolysis. Nevertheless, cell death may be a result of
combined loss of NAD* and inhibition of glycolysis, as supplementation of cells
185
with metabolic substrates that function downstream of glycolysis has similarly
been shown to rescue Parp1 mediated cell death (Alano et al., 2010; Alano et al.,
2007; Andrabi et al., 2014; Zong et al., 2004). Interestingly, we didn't find any
evidence that either NAD' or pyruvate addition rescued CGN sensitivity to MMS
despite multiple attempts, suggesting that either a higher concentration of these
compounds is needed to see rescue or that Parp1 is mediating cell death through
a separate mechanism. Zong, et al. (2004) has suggested that DNA damage
dependent Parp1-mediated cell death only occurs in cells that primarily utilize
glucose for their ATP needs via glycolysis, whereas cells that can employ
oxidative phosphorylation are resistant to the bioenergetic effects of Parp1
hyperactivation (Zong et al., 2004). This theory has not been built upon
unfortunately and it remains to be seen whether metabolic capabilities are the
key to Aag-dependent cell-specific responses.
Despite numerous attempts at experiments to dissect cell death mechanisms, we
ultimately came up with many negative results. Cell death was not rescued by
NAD' or pyruvate and was independent of caspase activity, calcium fluxes, AIF
translocation and mitochondrial permeabilization, many of the key features of
Parp1 mediated cell death. This begs the question: how are Aag and Parp
causing cell death in neurons after alkylation damage? And what are the next
steps to investigate downstream cell death components? We have assumed that
NAD+ and ATP are being depleted, however, we have not verified this fact.
Though it is very likely that we will see an Aag dependent reduction in NAD+
post-MMS, given the kinetics of PAR formation we have seen post-treatment, it is
possible that the cells recover functional levels prior to cell death, indicating that
reductions in NAD* are not causing sensitivity.
Our main focus moving forward is to investigate more closely how mitochondria
and mitochondrial function are involved in MMS-induced neural toxicity. Though
we did not see major alterations in mitochondrial polarization post-treatment, it
may be more informative and relevant to look at the level of individual cells or
186
mitochondria. We observed the formation of 'donut' and 'blob' shaped
mitochondria after drug treatment, which is correlated with mitochondrial stress
and dysfunction (Ahmad et al., 2013; Liu and Hajnoczky, 2011). Moreover,
reports describing this mitochondrial structure demonstrated that not all
mitochondria exhibited loss of polarization after treatment; rather, it was only
those mitochondria that became 'donut' shaped that had been depolarized and
often these mitochondria were able to regain polarization (Liu and Hajnoczky,
2011). Future experiments will be aimed at determining whether these
mitochondrial shapes are Aag and Parp1 dependent and whether hexokinase 1
activity is altered after MMS treatment, which would suggest PAR translocation
and direct inhibition.
Aag-Dependent Cell-Specific Responses
Ultimately, we have yet to determine a simple, definitive answer for why some
cells become sensitive to alkylation treatment upon the loss of Aag and why
some become resistant. Though neurons and retinal photoreceptors are post-
mitotic, myeloid progenitors are replicative, eliminating the hypothesis that
sensitivity to alkylating agents in the absence of Aag is solely dependent on an
active cell cycle. It seems to be the more common outcome for Aag-'- cells is to
become resistant to alkylation treatment or show no difference in sensitivity
compared to WT. The only documented mammalian cell types where loss of Aag
renders cells sensitive to MMS are murine ES cells and human glioblastoma and
carcinoma cells, all highly replicative cell types (Agnihotri et al., 2011; Engelward
et al., 1998; Paik, 2005). Interestingly, there are even conflicting results from
human glioma cells about whether loss of Aag causes sensitivity or resistance to
alkylating agents, indicating that predicting response will take more insight than
simply knowing the cell type (Agnihotri et al., 2011; Tang et al., 2011). On the
other hand, all reports of Aag overexpression have caused increased sensitivity
to alkylating agents, independent of cell type or replicative status (Calvo et al.,
2013b; Chou et al., 2015; Fishel et al, 2003; Fishel et al., 2007; Tang et al.,
187
2011; Trivedi et al., 2005; Trivedi et al., 2008). It is apparent, nonetheless, that
the mechanisms of cell death vary between cell types as evidenced by how Parp
inhibition affects cell sensitivity. In mouse neurons and photoreceptors, Parp
inhibition rescues mAagTg sensitivity by suppressing exaggerated programmed
necrosis (Calvo et al., 2013b). In human glioma cells overexpressing Aag, Parp
inhibition was instead shown to exacerbate alkylation sensitivity, likely due to
increased replication stress caused by the accumulation of AP sites/SSBs and
Parp inhibitor trapping onto the DNA (Murai et al., 2012). Therefore, the larger
question may be why do these cells die by different mechanisms? If we could
predict the mode of cell death, it is likely that we could determine how the cells
would respond upon loss of Aag.
Given the widespread use of alkylating drugs in chemotherapy, it would be
enormously beneficial to determine how alterations in Aag activity would change
cell-specific responses. Not only would we be able to decide whether the
treatment would be successful for a particular type of cancer, but we could
simultaneously determine how non-malignant cells would respond to
chemotherapy and minimize some of the debilitating side effects. Moreover,
inhibition of Aag in particular cell types would be an approach to enhance
desirable responses to alkylation treatment. Though no Aag inhibitors have been
developed to date, modulation of other downstream proteins in the BER pathway
is already being pursued in combination with alkylation chemotherapy (Agnihotri
et al., 2011; Jaiswal et al., 2011; Rouleau et al., 2010; Simeonov et al., 2009;
Tang et al., 2011). Thus, Aag represents an important druggable target for future
studies.
188
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Development of a Method to
Assay DNA Repair Capacity in Mammalian
Cells using High-Throughput Sequencing
195
Appendix:
Appendix: Development of a Method to Assay DNA Repair Capacity for
Mammalian Cells using High-Throughput Sequencing
Introduction
Prior to initiating my thesis research, I was involved in the development of a
method to assay DNA repair capacity in human cells in a high-throughput
manner. Though I contributed to the development of a plasmid based fluorescent
protein method, the majority of my lab-based work was focused on development
of a high-throughput sequencing based method to determine cellular repair
capacity and the ability of particular DNA lesions to cause transcriptional
mutagenesis. The manuscript below details the rationale, optimization, and
results of methods to assay DNA repair capacity based on the production of
fluorescent proteins or complete transcripts from transfected plasmids and
measured by flow cytometry or sequencing, respectively.
The work was published in 2014 and can be found in the included manuscript
below (Nagel et al., 2014a).
196
Manuscript: Multiplexed DNA repair assays for multiple lesions and
multiple doses via transcription inhibition and transcriptional mutagenesis.
Z. D. Nagel1'2 , C. M. Marulies', 1. A. Chaim', S. K. McRee2, P. Mazzucatol'2Aha, 2, R. 2,a3li',4 Chi 2,3S K Mce 2 ,P1 2,34A. Ahmad , R. P. Abo' , V. L. Butty' 3 4, A. L. Forget', L. D. Samson'
'Department of Biological Engineering, 2Center for Environmental Health
Sciences, 3Department of Biology, 4The David H. Koch Institute for Integrative
Cancer Research, Massachusetts Institute of Technology, Cambridge, MA
02139, USA
*To whom correspondence should be addressed: Address: 77 Massachusetts
Avenue, MIT Building 56-235, Cambridge, MA 02139, USA; Phone: 617-253-
6220; email: Isamson@mit.edu
High-Throughput DNA Repair Assays
Key Words: Host cell reactivation; DNA repair capacity, flow cytometry, high
throughput sequencing, transcriptional mutagenesis, multiplex assays
ABSTRACT: The capacity to repair different types of DNA damage varies among
individuals making them more or less susceptible to the detrimental health
consequences of such exposures.. Current methods for measuring DNA repair
capacity (DRC) are relatively labor intensive, often indirect and usually limited to
a single repair pathway. Here we describe a fluorescence-based multiplex flow-
cytometric host cell reactivation assay (FM-HCR) that measures the ability of
human cells to repair plasmid reporters each bearing a different type of DNA
damage or different doses of the same type of DNA damage. FM-HCR
simultaneously measures repair capacity in any four of the following pathways,
NER, MMR, BER, NHEJ, HR and MGMT. We show that FM-HCR can measure
interindividual DRC differences a panel of 24 cell lines derived from genetically
diverse apparently healthy individuals, and we show that FM-HCR can be used
to identify inhibitors or enhancers of DRC. We further develop a next generation
sequencing-based HCR assay (HCR-Seq) that detects rare transcriptional
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mutagenesis events due to lesion bypass by RNA polymerase, providing an
added dimension to DRC measurements. FM-HCR and HCR-Seq provide
powerful tools for exploring relationships among global DRC, disease
susceptibility, and optimal treatment.
SIGNIFICANCE STATEMENT: DNA, the blueprint of the cell, is constantly damaged
by chemicals and radiation. Because DNA damage can cause cell death or
mutations that can lead to diseases like cancer, cells are armed with an arsenal
of several distinct mechanisms for repairing the many types of DNA damage that
occur. DNA repair capacity (DRC) varies among individuals, and reduced DRC
is associated with disease risk, however the available DRC assays are labor
intensive and only measure one pathway at a time. Herein, we present powerful
new assays that measure DRC in multiple pathways in a single assay. We use
the assays to measure inter-individual DRC differences, inhibition of DNA repair,
and to uncover unexpected error-prone transcriptional bypass of a thymine
dimer.
INTRODUCTION:
DNA is under constant assault from damaging agents that produce a vast
array of lesions. Left unrepaired, these lesions have the potential to alter cellular
function and compromise the health of the organism, leading to degenerative
diseases, cancer, and premature aging (Ciccia and Elledge, 2010; Fu et al.,
2012a; Hoeijmakers, 2001; Jackson and Bartek, 2009; Lindahl and Wood, 1999).
Consequently, inter-individual variations in DNA repair capacity (DRC) are
thought to contribute to the fact that people have different susceptibilities to these
diseases (Athas et al., 1991; Decordier et al., 2010; Jalal et al., 2011; Ralhan et
al., 2007; Wilson et al., 2011). Furthermore, the efficacy of cancer chemotherapy
with DNA damaging agents is dependent on the DRC of the targeted cells
(Sarkaria et al., 2008). Thus, DRC measurements could potentially be used to
personalize both treatment and prevention of disease.
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We define DRC as the basal ability of cells to eliminate DNA damage from
the genome, however it should be noted that some DRC assays, such as
mutagen sensitivity assays, may also reflect changes in gene expression and the
activation of non-DNA repair pathways upon treatment of cells with DNA
damaging agents. A wide variety of methods are used to estimate DRC. Many
studies focus on indirect assessments of DRC through transcriptional profiling,
proteomics, and single nucleotide polymorphism (SNP) screens (Chin and Gray,
2008; Ellis, 2003; Li et al., 2013; van 't Veer and Bernards, 2008; Warmoes et al.,
2012). However, SNPs in DNA repair genes are not informative if the relevant
gene is not expressed. Likewise, gene expression data are not informative for
cases in which the gene product is inactive or cannot be assembled into a
functional complex. More direct measurements of DRC in vitro using cell lysates
overcome some of this complexity by integrating these factors into a single
readout (Geng et al., 2011; Georgiadis et al., 2012; Leitner-Dagan et al., 2012;
Redaelli et al., 1998); however these methods require separate assays for
measurements in more than one repair pathway, and may not be representative
of DRC in intact cells. Measuring the consequences of DNA repair in intact cells
by monitoring sister chromatid exchanges, chromosome aberrations, or DNA
strand breaks by comet assays also integrate complexity into a single readout,
but require labor-intensive analyses that make them refractory to implementation
in a clinical setting (Evans and Norman, 1968; Parshad et al., 1983; Perry and
Evans, 1975). Although recently developed high throughput comet assays
provide an excellent alternative, they are limited to DNA damage that leads to, or
can be converted to DNA strand breaks (Wood et al., 2010).
Host cell reactivation (HCR) assays report the ability of cells to repair DNA
damage that blocks transcription of a transiently transfected reporter gene (Athas
et al., 1991; Decordier et al., 2010; Li et al., 2009; Mendez et al., 2011; Qiao et
al., 2002; Ramos et al., 2004). Repair of the transcription-blocking lesion
reactivates reporter gene expression, thus providing a quantitative readout for
DRC. However, HCR assays cannot generally report repair of DNA lesions that
do not block the progression of the RNA polymerase, and current methods for
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measuring DRC are further limited by the need for separate experiments to
measure repair capacity in more than one pathway, or at more than one dose of
DNA damage. Herein we introduce a new high-throughput, fluorescence-based
multiplex HCR assay (FM-HCR) for measuring DRC in living cells that
overcomes these limitations. We first present a multicolor fluorescence assay
that simultaneously measures DNA repair at multiple doses of DNA damage. We
then demonstrate simultaneous DRC measurements for up to four repair
pathways in human and rodent cells, using reporters for the repair of both
transcription-blocking lesions and lesions that are bypassed by RNA polymerase.
To demonstrate potential applications of FM-HCR to population studies, we
measure interindividual DRC differences in five pathways in a panel of 24
lymphoblastoid cell lines derived from apparently healthy individuals. We also
show that FM-HCR can be used to identify agents that inhibit or enhance DRC.
Finally, to further increase throughput and to establish a single generalized
detection method for the repair of any lesion that alters transcription of a reporter
gene, we added to the HCR reporter protein paradigm a reporter transcript assay
that leverages the extraordinary power of next generation sequencing (HCR-
Seq). We use HCR-Seq to measure 20 independent reporter signals in a single
assay, and detect error-prone transcriptional bypass at a bulky DNA lesion in
human cells. FM-HCR and HCR-Seq provide rapid, high throughput methods of
assessing DRC in multiple pathways and represent a major improvement over
standard methods currently used in basic, clinical and epidemiological research
addressing the relationship among DNA damage, DNA repair and disease
susceptibility.
RESULTS:
Validation of FM-HCR
FM-HCR was used to assay DRC in 55 cell lines (Table 1). Expression
levels of five fluorescent reporter proteins were quantitated simultaneously using
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flow cytometry (Fig. S1). Use of 96-well electroporation plates reduced the time
required for transfection to less than an hour per plate, and a BD High
Throughput Sampler permitted data acquisition in less than 10 minutes active
time.
In vitro treatment of plasmids with UVC light resulted in a dose-dependent
reduction in reporter expression. When each of the five fluorescent reporter
plasmids was treated with a unique dose of UVC (Plasmid combination #1 in
Table 2), and subsequently co-transfected into cells, a dose-response curve was
generated from a single experiment that required only two transfections (Fig. Ia).
Dose-response curves spanning up to 3 decades of percent reporter expression
(%R.E.) were obtained for 7 lymphoblastoid cell lines (Fig. 1b,c), chosen
because they were previously characterized over 20 years ago for their capacity
to repair UV-irradiated plasmid DNA by another much more laborious method
(Athas et al., 1991). Differences in DRC were most pronounced at the highest
dose to plasmid (800 J/m2 ), with % reporter expression (%R.E.) values varying
over a range of about 100-fold among the cell lines. As expected, the highest
DRC was observed for lymphoblastoid cell lines derived from apparently healthy
individuals, referred to as wild type (WT) (Table 1). Moderately reduced DRC
was observed for two XPC cell lines, and severe defects were evident for XPA
and XPD cell lines. Between 18 and 40 hours, %R.E. increased for most cell
lines (Fig. Ib,c), consistent with time-dependent repair of transcription blocking
lesions.
The FM-HCR data presented in Fig. Ic reproduce those from the previous
study that also monitored transcription inhibition on UV-damaged plasmids 40
hours after transfection (Athas et al., 1991). In that study, chloramphenicol acetyl
transferase (CAT) levels in cell-free extracts were used as the reporter. Two
complementary methods were used to compare our data to the historical data.
First, the percent CAT expression (%CAT) reported at a single dose of UV
irradiation (300 J/m2 in ref. (Athas et al., 1991) ) was highly correlated (R2 = 0.92,
p = 0.0006) with %R.E. at a single dose (400 J/m 2) in the present study (Fig. 1d).
The relative repair capacity of multiple cell lines can also be compared by
201
calculating the parameter Do, corresponding to the dose at which HCR falls
below 37% R.E. (Jagger, 1976). The Do values calculated from our experimental
data were also highly correlated with the historical Do values (R2 = 0.92, p <
0.0001) (Fig. le).
To confirm that the dose response curves in Figs. lb and 1c could be
obtained independently of the choice of fluorescent reporters, the experiment
was repeated with the plasmids shuffled with regard to which plasmid received a
particular UV dose (Plasmid combination #2 in Table 2). The resulting dose
response curves obtained at 18 and 40 hours are presented in Figs. If and Ig,
respectively. Once again, the FM-HCR data collected at 40 hours reproduce the
historical data (Athas et al., 1991) (Fig. 1h). Likewise, the FM-HCR data
collected with plasmid combination #2 reproduce those obtained using the
plasmid combination #1 (Fig. Ii).
FM-HCR assays were also carried out on 7 primary untransformed skin
fibroblast cell lines and compared to Epstein-Barr virus transformed
lymphoblastoid cell lines derived from the same individuals (represented as
individuals i-vii in Table 1); cells were from 4 apparently healthy individuals and 3
XP patients. A similar pattern of dose response curves was obtained for both
fibroblasts and lymphoblastoid cells (Figs. 1j and I k, respectively). Overall,
absolute NER capacity measured in fibroblasts appeared to be somewhat higher
than that in the lymphoblastoid cell lines; however, a comparison of DRC
measured at 800 J/m 2 indicated that NER phenotype is strongly correlated (R2
0.94, p = 0.0003) between the two cell types (Fig. 11).
Development of FM-HCR assays for DNA mismatch repair and direct
reversal of 06-MeG
Fluorescent plasmid reporters for direct reversal of O6-MeG and mismatch
repair capacity were developed (Fig. S2). For mismatch repair assays, a G:G
mismatch containing plasmid was constructed; it has previously been shown that
G:G mismatches are repaired inefficiently in extracts from MMR deficient cells
202
(Parsons et al., 1993), and we confirmed this observation in MMR-deficient
HCT116 cells and MMR-proficient HCT116+3 cells that have been
complemented with human chromosome 3 (Fig. S2d), as well as MMR-deficient
MT1 cells that lack MSH6, and MMR-proficient TK6 cells (Fig. 2). This reporter
expresses a non-fluorescent protein unless a repair event restores a cytosine in
the transcribed strand, leading to wild type orange fluorescent protein. Because
the plasmid lacks a strand discrimination signal, the theoretical upper limit of
reporter expression (relative to a similarly constructed wild type homoduplex
control) is 50%. For direct reversal of 06-MeG, a plasmid that encodes a non-
fluorescent protein in the absence of the lesion was prepared. Introduction of a
site-specific 06-MeG lesion into the transcribed strand causes transcription errors
(Burns et al., 2010), producing transcripts encoding the wild type mPlum
fluorescent protein. Thus, cells deficient for methylguanine methyltransferase
(MGMT) mediated 06-MeG repair express relatively high levels of the mPlum
reporter, whereas cells expressing MGMT remove the source of transcription
errors, thus reducing reporter expression.
Simultaneous measurement of DRC in 3 pathways with FM-HCR
DRC for three pathways, namely NER, mismatch repair (MMR), and the
direct reversal of 06-MeG (MGMT) was measured in five lymphoblastoid cell
lines (Fig. 2). DRC for each pathway was first measured in separate
experiments. The severe NER defect for the XPA-deficient GM02344 cell line
was reproduced, whereas relatively high NER capacity was confirmed for the
four other cell lines with no known NER defect (GM01953, MT1, TK6, and
TK6+MGMT). The lymphoblastoid cell line MT1, known to be deficient for
mismatch repair (Kat et al., 1993), expressed the MMR reporter at a level up to
10-fold lower than that of the other four cell lines that have no known MMR
defects. As expected, the 06-MeG direct reversal reporter was highly expressed
in MT1 and TK6, owing to the absence of MGMT. Expression of the same
reporter was reduced nearly 1000-fold in TK6+MGMT cells expressing a high
203
level of MGMT, and was reduced ~ 8-fold and 250-fold in GM01953 and
GM02344, respectively, both of which express active MGMT, but at different
levels (Zhukovskaya et al., 1992).
One of our goals is to increase the throughput of DRC assays by
measuring the activity of multiple DNA repair pathways in a single assay. To test
whether our DRC reporters could be combined in a single experiment without
affecting the accuracy of the assay, we co-transfected three reporter plasmids,
each targeting a different pathway, along with an internal transfection control
(plasmid combination #3 in Table 2). This yielded nearly identical DRC profiles
for NER, MMR and MGMT as those obtained when the reporters were
transfected in separate experiments (Fig. 2b).
Simultaneous measurement of four DNA repair pathways including BER
and NHEJ.
We next sought to measure DRC in four pathways including BER or DSB
repair (Fig. 3). To add a fourth pathway to the FM-HCR in Fig. 2, a BFP reporter
for repair of a double strand break by the NHEJ pathway was developed (Fig. 3a
and Fig. S3). This assay was validated using the M059J and M059K cell lines,
which were derived from a single glioblastoma (Allalunis-Turner et al., 1993).
The M059J cell line is deficient for DNA PKcs that is required for NHEJ
(Leesmiller et al., 1995). As expected, M059J cells expressed -40-fold lower
levels of the NHEJ reporter relative to the wild type M059K cells when the
reporter was transfected separately from other reporters (Fig. 3b). To test
whether DRC could be measured in four pathways simultaneously, we co-
transfected the NHEJ reporter with the reporters described above for NER, MMR,
and MGMT (plasmid combination #4 in Table 2). As with the three-pathway
measurements described above, the co-transfected reporters yielded nearly
identical DNA repair profiles as the separately transfected reporters (Figs. 3b
and 3c). MMR and MGMT capacity was similar in the two cell lines, however
NER capacity was reduced in M059J by approximately 7-fold relative to M059K.
204
It has been observed previously that the XPC and ERCC2 genes are
overexpressed in M059J vs M059K cells, and that the M059J cells are slightly
more sensitive than M059K cells to UV irradiation (Galloway and Allalunis-
Turner, 2000). Inefficient NER in the presence of excess XPC protein has also
been noted in vitro (Mu et al., 1996).
To further demQnstrate the versatility of FM-HCR, we performed an assay
that includes BER, NER, MMR and MGMT (Fig. 3d). An mOrange fluorescent
reporter for base excision repair of 8-oxoguanine (8-oxoG) was developed that
produces wild type mOrange transcripts when RNA polymerase incorporates
adenine opposite a site-specific 8-oxoG lesion. Deficient 8-oxoG repair is
expected to result in a higher reporter expression. In keeping with this
expectation, mouse embryonic fibroblasts (MEFs) deficient for 8-oxoG DNA
glycosylase (Ogg1) expressed an approximately 20-fold higher level of mOrange
than wild type MEFs when the reporter was transfected separately from the other
reporters (Fig. 3e). When the 8-oxoG reporter was co-transfected with three
other reporters for NER, MMR and MGMT (plasmid combination #5 in Table 2),
we once again observed DRC profiles that were nearly identical to those
obtained when the reporters were transfected separately (Figs. 3e and 3f).
MMR and NER capacity were similar in the two cell lines, but MGMT reporter
expression was approximately 5-fold higher in the wild type MEFs. Since the
wild type MEFS were derived from C57BL/6J mice, the differences in MGMT
repair capacity could be due to the mixed (C57BL/6J and 129SV) background of
the Ogg1-/ mouse from which the MEFs were derived (Klungland et al., 1999).
Analysis of DRC for five pathways in a panel of 24 cell lines derived from
apparently healthy individuals
A previously described assay for HR was incorporated into FM-HCR
experiments and was validated using cell lines with known defects in double
strand break repair (Fig. S4) (Kiziltepe et al., 2005). Assays for NER, MMR,
MGMT, HR and NHEJ capacity (see plasmid combinations #3 and #6) were then
205
carried out on a panel of 3 control cell lines (TK6, MT1, and TK6+MGMT) and 24
human lymphoblastoid cell lines derived from apparently healthy individuals of
diverse ancestry (Collins et al., 1998). Each of the 24 cell lines exhibited a unique
DNA repair profile (Figure 4a), and a range of DRC was observed across the 24
cell lines for each pathway (MGMT, 285-fold; MMR, 4.4-fold; HR, 3.7-fold; NER,
3.2-fold, and NHEJ, 2.1-fold). To further validate the FM-HCR assay for MGMT,
transcript levels measured by TaqMan qPCR for the cell lines in Fig. 4a were
compared with fluorescent reporter expression. A non-linear relationship was
observed between MGMT FM-HCR % Reporter Expression and transcript levels
(not shown), however log-transformed FM-HCR data correlated extremely well
with MGMT transcript levels (Fig. 4b).
Application of FM-HCR to assays for DRC inhibition
To further demonstrate the potential applications of FM-HCR, the assay
was used to detect inhibition of DRC by metals and a small molecule. Cadmium
and arsenic have been shown previously to inhibit NER (Hartwig, 2010; Hartwig
et al., 1997). FM-HCR confirmed NER inhibition in the presence of low
micromolar concentrations of the two metals (Fig. 4c), whereas no effect on NER
was detected in the presence of Compound 401, known to inhibit a critical NHEJ
factor, namely the DNA dependent protein kinase catalytic subunit (DNA PKCs)
(Griffin et al., 2005). Moreover, Compound 401 was shown to inhibit NHEJ in a
dose dependent manner (Fig. 4d), while MMR, NER, and HR were unaffected.
Taken together, the FM-HCR data demonstrate a versatile method for
measuring in a single assay the repair of multiple DNA damage substrates, with
either different doses or with different types of damage. While measuring four
DNA repair pathways simultaneously represents a significant advance, the
degree of multiplexing possible for the FM-HCR assay is dependent upon the
number of fluorescent reporters that can be measured simultaneously. We
therefore developed an additional assay that does not require the detection of
fluorescent proteins.
206
Deep sequencing analysis of cells transfected with reporter plasmids
To increase the potential number of reporters that can be detected in a
single assay we developed HCR-seq, a method of distinguishing and quantifying
multiple full-length reporter transcripts using next generation sequencing. Two
cell lines exhibiting a large difference in their NER capacity (GM02344 and
GM01953) were selected for a direct comparison of DRC measured by HCR-seq
versus FM-HCR (Fig. 5). Each cell line was transfected with plasmid
combination #7 (Table 2), and at 18 hours cells were analyzed by both HCR-seq
and FM-HCR. Plasmid combination #7 included a modified GFP reporter
containing a single site-specific cyclobutane pyrimidine dimer (CPD). This
reporter was included to allow a focused analysis of possible transcriptional
errors induced by a bulky DNA lesion.
For HCR-seq analysis, total RNA was isolated and subjected to standard
Illumina mRNAseq sample preparation and analysis (Mortazavi et al., 2008). A
total of 358,281,302 reads were generated for two replicates of 4 multiplexed
samples, with each replicate analyzed in a separate HiSeq lane (Table SI).
Between 30 and 50 million reads were assigned to each original sample.
315,574,792 reads (88%) mapped properly to genes annotated for the human
genome plus the five reporter sequences. In each sequencing lane, all 5 reporter
transcripts were detected for each of the 4 samples; each sequencing lane
simultaneously measured expression levels for 20 reporters. Alignment
statistics, the criteria used to define proper alignment and reasons for excluding
the remaining 12% of reads from subsequent analysis are detailed in Tables S2-
S5.
DNA repair and transcriptional mutagenesis detected by RNA sequencing
Relative transcript levels in WT (GM01953) versus XPA (GM02344) cell
lines were determined for both host genes and plasmid reporter genes. Plasmid
reporters were found to be among the most highly expressed genes (Fig. S5a).
207
As expected, reporter expression from UV-treated plasmids was reduced in a
dose-dependent manner (Fig. 6a), and the reduced expression for the XPA cell
line (GM02344) was far greater than that for W-T cells (GM01953). More
importantly, reporter transcript expression mirrored closely the dose response
curves obtained from the same transfected cells using FM-HCR (Fig. 6b).
Reporter expression from plasmids containing a single site-specific CPD in the
transcribed strand was likewise reduced relative to that from undamaged
plasmids (Fig. S5b).
With respect to global gene expression in the transfected cells, fewer than
10 host transcripts showed a > 2-fold change in expression in the presence of UV
treated plasmids versus undamaged plasmids (Table S5). Among these, only
three (SMNI, RPL21, and RN5-8S1) were observed in both cell lines, but in no
case was a change in the same direction observed for both replicates. Thus, no
significant transcriptional response to the presence of DNA damage in plasmids
was evident under our experimental conditions. However, consistent with the
FM-HCR data (Fig. 2) that suggested higher MGMT activity in GM02344 cells
compared with GM01953 cells, the mRNAseq data indicated an approximately 3-
fold higher expression of the MGMT transcript in GM02344 versus GM01953.
Furthermore, XPA transcripts were expressed at lower levels in GM02344 versus
GM01953, and they were only rarely spliced correctly in GM02344 (Fig. S5c).
These data reproduce previously reported splicing errors in GM02344 due to a
homozygous 555G>C mutation in the XPA gene (Satokata et al., 1992). Finally,
to assess the potential for DNA contamination in RNAseq samples, the density of
reads aligning to intergenic regions (which are not expected to be represented in
transcripts) was compared to the density of reads aligning to exons, and the ratio
of exonic/intergenic reads was found to be greater than 1000, indicating an RNA
purity >99.9%.
Sequence-level analysis of mRNAseq data revealed base substitutions in
reporter transcripts at the position corresponding to the site-specific CPD; this
was true for both cell lines (Fig. 6c). The most frequently observed base
change, an A->G mutation at the 5' Adenine in the ApA sequence opposite the
208
CPD (hereafter AA->GA), was detected at a frequency of 1.3% in cells with no
known repair defect (GM01953), and 5.8% in NER-deficient GM02344 cells. In
transcripts expressed from the undamaged plasmid, the frequency of the
AA+GA mutations at this position was less than 0.2%. A potential experimental
concern is that trace contaminating plasmid DNA might be amplified during
Illumina sample preparation, thus giving rise to DNA fragments with base
substitutions due to error-prone CPD bypass by DNA polymerase. However,
nearly identical frequencies for AA+GA mutations were found using a second
sample preparation method that excludes the possibility of contaminating lesion-
containing plasmid (Fig. S5d). It therefore appears that human RNA polymerase
can bypass a thymine dimer in vivo, albeit in an error prone manner.
AA-+GA mutations were also induced in a dose-dependent manner in
transcripts expressed from UV-irradiated reporter plasmids containing thymine
dimers that were not site-specific (Fig. 6d). As expected for randomly induced
DNA damage, the absolute frequency of the base substitution was much lower
than that observed for transcripts expressed from the reporter with the site-
specific thymine dimer. Once again, base changes occurred at a higher
frequency in NER-deficient versus wild type cells. These data provide additional
evidence for error prone transcriptional bypass of thymine dimers.
DISCUSSION
We have established new methods that enable rapid, high throughput
measurements of DRC for DNA lesions that either block RNA polymerase II-
mediated transcription, or alter the sequence of reporter transcripts. These
methods do not require the generation of cell lysates or the use of in vitro assays,
and can simultaneously measure in vivo DRC at multiple doses or in multiple
repair pathways. As a result, these assays outperform current methods of
measuring DRC (Leitner-Dagan et al., 2012; Li et al., 2009), and have the
potential to be used to personalize the prevention and treatment of cancer and
other diseases caused by inefficient repair of DNA damage.
209
We have demonstrated an application of the FM-HCR to the question of
whether NER capacity in human lymphoblastoid cells is representative of repair
capacity in other tissues. Lymphoblastoid cells provide a convenient source of
cells for use in human variability studies, however the extent to which they
represent a faithful surrogate for other cells in primary tissues has been called
into question (Choy et al., 2008; Davis and Kohane, 2009; Stark et al., 2010).
The present data indicate a strong correlation between NER capacity in primary
human skin fibroblasts and transformed B-lymphoblastoid cells from the same
individuals (Fig. 11). The strong correlation further illustrates that the assay can
be carried out reproducibly in primary or transformed cells from multiple tissues.
To our knowledge, our use of HCR to simultaneously measure
combinations of NER, MMR, BER, NHEJ, HR or MGMT capacity is the first
example of a quantitative assay capable of measuring repair of DNA damage by
multiple distinct pathways in parallel. One of the strengths of FM-HCR is that it
yields a single readout (fluorescence) in place of multiple unique outputs from
very different experimental procedures that have been used previously to
characterize the same repair pathways in the cell lines for which data are
presented (Fig. 2 and Fig. 3) (Athas et al., 1991; Hickman and Samson, 1999a;
Kat et al., 1993; Klungland et al., 1999; Leesmiller et al., 1995; Zhukovskaya et
al., 1992). The fluorescent reporters for direct reversal of 06-MeG and BER of 8-
oxoG illustrate the use of transcriptional mutagenesis to measure the repair of
DNA lesions that are bypassed in an error-prone manner by RNA polymerase.
This paradigm, where the presence of a DNA lesion changes the expressed
reporter sequence to one that encodes a functional protein, holds promise as a
general method of measuring DRC, because a wide variety of toxic and
mutagenic DNA lesions are known to induce transcriptional errors (Bregeon and
Doetsch, 2006; Bregeon et al., 2009).
We have presented several applications of FM-HCR to demonstrate the
broad utility of the assay. The flow cytometric fluorescence-based FM-HCR
method accurately reproduces data collected previously for a set of cell lines
known to differ in NER capacity (Athas et al., 1991). A screen of a much larger
210
set of cell lines derived from apparently healthy individuals illustrates the
potential to efficiently measure DRC in multiple pathways in large sets of
samples (Fig. 4a), and to identify agents that inhibit or enhance DRC (Figure 4c
and 4d). Screens for DRC inhibitors or enhancers are expected to identify some
agents for which the mechanism of action is unknown; indeed, uncertainty
remains as to the precise mechanisms by which arsenic and cadmium exposure
lead to reduced DRC (Bhattacharjee et al., 2013; Hartwig, 2010); the strength of
FM-HCR lies in the ability to measure changes in DRC as an important functional
endpoint.
By using multiple fluorescent reporters, a 96-well format, and automated
flow cytometric sample processing, the method is rapid and less labor intensive
than the standard CAT-based HCR assay. For example, the total active
laboratory time required to perform the analysis to generate the triplicate data in
Fig. 1b and Ic is approximately 12 hours, or 1-2 hours per cell line, using flow
cytometers equipped with a high throughput sampler to enable automated data
acquisition. In addition, experimental error is reduced by co-transfection of
reporters, allowing normalization of expression from a damaged plasmid to that
of an undamaged control plasmid included in every transfection. Through these
technical improvements, FM-HCR removes a major barrier to epidemiological
studies of DRC that include large populations and multiple DNA repair pathways.
Furthermore, because standard oncology labs are equipped with flow
cytometers, the assay also has the potential to be of use in a clinical setting.
The use of next generation sequencing to essentially count reporter
transcripts (HCR-seq), rather than measuring their fluorescent translation
products, presents an opportunity to vastly increase throughput, and overcomes
important limitations on assay throughput and versatility that are otherwise
imposed by the need to detect fluorescent reporter proteins. We have validated
the HCR-seq approach by showing that HCR of UV-irradiated plasmids analyzed
by mRNAseq yields a pattern of dose-response curves similar to those obtained
previously using a CAT-based HCR assay (Athas et al., 1991), as well as those
obtained in the current study by FM-HCR analysis (Fig. 1). Because next
211
generation sequencing can be used to quantitate the expression levels of
thousands of transcripts simultaneously, our assay has the potential to measure
expression of dozens of reporters for multiple individuals in a single experiment;
this would make characterization of global DRC in large populations both efficient
and affordable (See supplementary note).
HCR-seq constitutes a paradigm shift in the quantitation of DRC because
of the ability to measure the repair of any lesion that either inhibits transcription
or induces transcriptional mutagenesis. Base misincorporation opposite DNA
lesions that are bypassed by DNA polymerase during replication has been
extensively studied for many lesions (Shrivastav et al., 2010a). Misincorporation
during transcription by RNA polymerase has been documented for a growing
number lesions, and often mirrors that of DNA polymerase during replication
(Bregeon and Doetsch, 2011). As a result, most mutagenic lesions can be
expected to have a transcriptional mutagenic signature. The HCR-seq strategy
should therefore be useful in DRC measurements for nearly any pathway. The
data in Fig. 6 also illustrate the power of this unbiased approach to detect rare
events that are specific to transcription of damaged DNA.
The two major applications to human health that we foresee for these
assays relate to personalized prevention and treatment of cancer. The available
published data indicate that DRC is an important factor both in cancer
susceptibility and in the efficacy of cancer treatment with DNA damaging agents,
and that plasmid-based HCR assays can readily be applied to primary human
tissue samples, including stimulated peripheral blood mononuclear cells (Athas
et al., 1991; Decordier et al., 2010; Jalal et al., 2011; Leitner-Dagan et al., 2012;
Li et al., 2009; Sarkaria et al., 2008). FM-HCR and HCR-Seq now open the door
to a comprehensive analysis of DRC as a biomarker for disease susceptibility.
For personalized disease prevention, FM-HCR could be applied to human blood
cells to identify individuals who may have a higher risk of disease. In terms of
personalized treatment, the assays could be used to measure DRC in blood cells
to predict patient tolerance for a particular cancer therapy (Alapetite et al., 1999),
or to measure DRC in cancer cells to predict the efficacy of treatment with DNA
212
damaging chemotherapeutic agents in a manner analogous to using MGMT
promoter methylation to predict the response of cancers to alkylating
chemotherapy agents such as temozolomide (Hegi et al., 2005). Indeed, the
data in Fig. 4 show that FM-HCR data reproduce the results of a standard
TaqMan qPCR assay for MGMT gene expression in lymphoblastoid cell lines.
The functional FM-HCR and HCR-Seq assays might be expected to outperform
promoter methylation assays because (i) they provide a direct, quantitative
readout of repair activity rather than an indirect estimate of DNA repair gene
expression, and (ii) they provide data for repair capacity in additional pathways
such as MMR, which also contributes importantly to alkylation sensitivity (Kat et
al., 1993). Finally, the ability of FM-HCR to identify agents that either inhibit or
enhance DRC in human cells (Fig. 4c and 4d) opens the door to screens for
novel compounds that could be used either to potentiate the effects of DNA
damage-based anticancer agents or to mitigate the deleterious effects of
environmental exposure to DNA damaging agents (Kim et al., 2011; Srinivasan
and Gold, 2012; Zellefrow et al., 2012).
In addition to the possible clinical applications described above, HCRseq
has the potential to reveal new biological phenomena in the basic research
setting. The mRNAseq data presented here provide evidence that transcriptional
errors result when human RNA polymerase I bypasses a CPD. Because the
plasmids are not replicated in the cell, and transcript sequence changes were
observed at an elevated rate in repair deficient cells, these changes are likely to
reflect transcriptional mutagenesis events due to unrepaired DNA lesions in the
transcribed DNA strand. While it has been reported previously that in vivo
bypass of a CPD by RNA polymerase may result in rare deletions, and bypass of
a bulky 8,5'-cyclo-2'-deoxyadenosine lesion may result in both deletions and
base substitutions (Marietta and Brooks, 2007), our observation of frequent base
misincorporation opposite a CPD by RNA polymerase II appears to be without
precedent. A recent in vitro analysis indicated that transcriptional CPD bypass
followed a so-called A-rule, resulting in error-free bypass (Walmacq et al., 2012);
athough base misincorporation was observed, subsequent extension of
213
transcripts beyond the misincorporated base was strongly inhibited. The present
data provide in vivo evidence of error-prone transcriptional bypass of bulky DNA
lesions in human cells followed by completion and polyadenylation of the
transcript. A lower limit (about 6%) for the frequency of bypass events resulting
in an AA->GA mutation can be estimated from the data in Fig. 6c. Since it is
expected that reporter plasmids that have already been repaired will be
transcribed at a higher rate, and because error-free bypass (according to an A-
rule) cannot be distinguished from transcripts arising from repaired plasmid, the
rate of bypass is likely higher than 6%.
CONCLUSIONS:
FM-HCR and HCRseq represent powerful new tools for high throughput
measurements of human DRC and provide a rapid functional characterization
that complements existing, indirect measures of DRC. FM-HCR permits the
simultaneous measurement of repair for up to four different doses of DNA
damage, or types of DNA damage, in a single assay. HCR-seq has the potential
to measure thousands of reporter sequences in a single assay, with barcodes
providing unique identifiers for the type or dose of DNA damage as well as for the
individual whose cells are being analyzed. Both methods expand the scope of
lesions whose repair can be measured to include those that do not block
transcription, and as additional substrates are developed, we anticipate that our
assays will permit measurements of DRC in all of the major DNA repair pathways
in a single assay. Our assays hold an advantage over in vitro assays because
the transcription-based reporters limit the readout to DNA that has been repaired
in vivo in chromatinized DNA, thus increasing the likelihood of recapitulating
physiological DNA repair phenotypes. The assays have the potential to reduce
the cost and labor required for DRC measurements to a level compatible with
large-scale epidemiological studies and clinical diagnostic/prognostic
applications. The data presented herein also illustrate the utility of the assays as
a research tool that can reveal mechanisms of DNA repair and damage tolerance
214
and that can provide a new means of screening chemical libraries for inhibitors or
enhancers of DRC.
MATERIALS AND METHODS:
DNA Repair Reporter Plasmids
Detailed methodology for the construction of reporter plasmids and the methods
used to transfect plasmids into cells can be found in the supplemental
information (Figs S2-S4). Briefly, Plasmids for expression of the fluorescent
proteins AmCyan, EGFP, mOrange, and mPlum were purchased from Clontech,
and that for tagBFP was purchased from Axxora. Reporter genes were
subcloned into the pmax cloning vector (Lonza). NER reporters were prepared
by irradiating plasmids with UVC light. The resulting DNA damage prevents
fluorescent reporter expression by blocking transcription; repair by NER restores
reporter expression. The NHEJ reporter comprised a linearized fluorescent
reporter; because double strand breaks constitute an absolute block to
transcription, NHEJ-dependent recircularization of the plasmid is required to
restore reporter expression. MMR reporters consisted of heteroduplex DNA
engineered such that the transcribed strand encoded a non-fluorescent protein.
Repair of a single, site-specific mismatch restores the wild type sequence to the
transcribed strand, and results in fluorescent reporter expression. Reporters for
repair of 8-oxoG or 06-MeG were engineered such that transcriptional
mutagenesis in the presence of the DNA lesion lead to expression of wild type
fluorescent reporter protein. Because repair removes the source of
transcriptional mutagenesis, repair of these plasmids is inversely proportional to
the measured fluorescence.
Flow Cytometry
215
Cells suspended in culture media were analyzed for fluorescence on a BD LSRIl
cytometer, running FACSDIVA software. Cell debris, doublets and aggregates
were excluded based on their side scatter and forward scatter properties.
TOPRO-3 was added to cells 5-10 minutes prior to analysis, and used to exclude
dead cells from the analysis. The following fluorophores and their corresponding
detectors (in parentheses) were used: tagBFP (Pacific Blue), AmCyan
(AmCyan), EGFP (FITC), mOrange (PE), mPlum (PE-Cy5-5), and TOPRO-3
(APC). The linear range for the corresponding photomultiplier tubes was
determined using BD Rainbow fluorescent beads and unlabeled polystyrene
beads based on the signal-to-noise ratio, %CV, and M1/M2 parameters as
previously described (Perfetto et al., 2006). Compensation was set using single
color controls. Regions corresponding to cells positive for each of the 5
fluorescent proteins were established using single color dropout controls. For
reporters that required compensation in more than one detector channel,
fluorescence in the reporter channel was plotted separately against each of the
channels requiring compensation. Using these plots, both single controls and the
dropout control (in which the reporter of interest was excluded from the
transfection) were used to establish regions corresponding to positive cells (Fig.
Sla). Equations used to calculate fluorescent reporter expression are detailed in
the supplemental information.
mRNAseq
Total RNA was isolated using standard procedures detailed in the
supplemental information. Total RNA samples were submitted to the MIT
BioMicroCenter for preparation and sequencing. Briefly, total RNA was poly-A
purified, fragmented, and converted to cDNA using the Illumina Tru-SeqT M
protocol. Library construction from cDNA was performed using the Beckman
Coulter SPRI-works system. During library amplification, a unique bar-code was
introduced for each of 8 samples corresponding to the four transfections
performed in duplicate (Table S1), and from which total RNA was generated.
Four samples from each replicate were clustered on a separate sequencing lane
216
and run on an Illumina HiSeq 2000 instrument. Image analysis, base calling and
sequence alignment to a synthetic genome consisting of the human genome and
the five fluorescent reporter genes were performed using the Illumina Pipeline.
Aberrant expression of the XPA gene in GM02344 cells provided an internal
confirmation of the identity of the cell lines; reduced expression and an expected
lack of regular splicing junction reads spanning intron 4 of the XPA gene from
GM02344 was observed (Fig. S5c), confirming a previously reported missplice in
XPA transcripts due to the homozygous 555G>C mutation (Satokata et al.,
1992).
To ensure that trace DNA contamination of the RNAseq samples did not
contribute significantly to the observed frequency of base substitutions in
transcripts expressed from reporter plasmids (Fig. 6), a second complementary
sample preparation was performed and analyzed by Illumina sequencing.
Details of the experimental procedures are available in the supplemental
information; briefly, mRNA isolated from cells transfected with reporter plasmids
was treated with DNAse and reverse transcribed to generate a cDNA library.
PCR amplification of reporter cDNA was not detected in when mRNA that was
not reverse transcribed was used as a template (Fig. S5d), confirming cDNA as
the template for PCR amplification, and hence ruling out significant plasmid
contamination. Amplicons were fragmented and submitted for standard Illumina
sample preparation.
Next Generation Sequencing Data Analysis
Illumina sequencing data were analyzed using the Tuxedo software
suite. Mapped reads were aligned to the hg19 human genome assembly and the
five reporter gene sequences using Tophat v 2.0.6, and junction reads
determined. Additional details of all analyses including input parameters are
available in tables S2-S6. Cufflinks v 2.0.2 was run to quantify reads in terms of
reads per kilobase of transcript per million mapped reads (RPKM) (Trapnell et al.,
2012). Samtools mpileup (v 0.1.16 r963:234) was used to aggregate reads at all
217
positions in the alignment file. Using the pileup file as input, single nucleotide
variants, as well as insertions and deletions (indels) present in the mRNAseq
data were identified using the software package VarScan v2.3.4 (Koboldt et al.,
2012). All positions meeting a minimum read depth of 8 were considered further,
however no minimum variant frequency threshold was set in order to detect rare
variants and to establish the sequencing error rate. Custom Python scripts were
used to generate a list of all deletions spanning an ApA sequence. The
frequencies for base substitutions at each ApA sequence in the reporter
transcripts were also determined.
Statistics
Statistics were performed with the GraphPad Prism 5.0 software package.
The correlations between data sets in Figs. I and 4 were assessed using a
linear regression model that reports R2 for the goodness of fit and a p value for
the slope of the line being significantly different from zero. The p values in Fig. 6
were calculated from a two-tailed unpaired t-test. Error bars in figs. 1,2,3,4 and
5 report the standard deviation of at least three biological replicates
ACKNOWLEDGEMENTS: We acknowledge funding support from NIH grant DP1-
ES022576. We thank Dr. Jennifer Calvo for establishing the immortalized MEF
cell lines, Prof. Penny Jeggo for the V79 and xrs6 cell lines, and Prof. Ryan
Jensen for the VC8 cell line. Authors declare no competing interests.
218
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225
FIGURE LEGENDS:
Fig. 1. Measurements of DRC by FM-HCR. (a) DNA lesions are introduced into
fluorescent reporter plasmids in vitro. Numbers labeling the plasmids represent
the dose (in J/m 2) of UV radiation. Following treatment, plasmids were combined
and co-transfected into cells. After 18 or 40 hours incubation, cells were assayed
for fluorescence by flow cytometry. Comparison of fluorescence signals to those
from cells transfected with undamaged plasmids yields a dose-response curve
(experimental data for GM02344 with plasmid combination #1 in Table 2) (b)
Dose-response curves for seven cell lines 18 hours after transfection with
plasmid combination #1 (Table 2) (c) Dose response curves for the cells in (b) at
40 hours. (d) Comparison of % reporter expression as measured by FM-HCR at
400 J/m 2 is plotted against %CAT as measured by conventional HCR for the
same cell lines at 300 J/m 2. (e) Do values calculated from FM-HCR data plotted
against those reported in the literature. Error bars represent the standard
deviation calculated from biological triplicates. (f) Dose-response curves for
seven cell lines 18 hours after transfection with plasmid combination #2 (Table
2). (g) Dose response curves at 40 hours. (h) Comparison of % reporter
expression as measured by FM-HCR at 400 J/m 2 with plasmid combination #2 is
plotted against %CAT as measured by conventional HCR for the same cell lines
at 300 J/m 2 . (i) Comparison of FM-HCR data for plasmids treated at 400 J/m 2 in
experiments #1 and #2. (j) Dose-response curves generated by FM-HCR for
lymphoblastoid cell lines 40 hours after transfection with plasmid combination #2
(Table 2). (k) Corresponding dose response curves for primary skin fibroblasts
from the same seven individuals. (1) Correlation between % reporter expression
from plasmids irradiated at 800 J/m 2 in the lymphoblastoid and fibroblast cells
isolated from the same individuals. Each color in panels j, k and I corresponds to
one of the individuals (i-vii) in Table 1. Error bars represent the standard
deviation calculated from biological triplicates. See also Fig S1.
226
Fig. 2. Simultaneous measurements of DRC in three pathways. (a) Plasmids
used in the multi-pathway FM-HCR (also see plasmid combination #3 in Table
2). (b) DRC for several cell lines obtained by assaying each pathway in a
separate transfection experiment (left) simultaneously following co-transfection of
the reporter plasmids (right). Error bars represent the standard deviation
calculated from biological triplicates. See also Fig S2.
Fig. 3. Simultaneous measurements of DRC in four pathways. (a) Plasmids
used in the FM-HCR for NHEJ, NER, MMR and MGMT (also see plasmid
combinations #4 Table 2). Note that the undamaged plasmid (AmCyan) included
to control for transfection efficiency is not shown. (b) DRC for several cell lines
obtained by assaying each pathway in a separate transfection experiment. (c)
DRC measured simultaneously following co-transfection of the reporter plasmids.
(d) Plasmids used in the FM-HCR for NER, MMR, BER, and MGMT (also see
plasmid combinations #5 Table 2). (e) DRC for several cell lines obtained by
assaying each pathway in a separate transfection experiment. (f) DRC measured
simultaneously following co-transfection of the reporter plasmids. Error bars
represent the standard deviation calculated from biological triplicates. See also
Fig S3.
Fig. 4. Applications of FM-HCR to interindividual DRC differences and
identifying DNA repair inhibitors and enhancers. (a) FM-HCR analysis of repair
capacity in 5 pathways for 27 cell lines. Cells were transfected with plasmid
combination #3 or plasmid combination #6. (b) Correlation between MGMT
transcript levels measured by taqMan qPCR and % Reporter Expression (log
transformed) from the MGMT HCR. (c) FM-HCR measurements of NER
inhibition. (d) FM-HCR measurements of Compound 401 inhibition of DNA repair
capacity in four pathways. For all experiments, cells were assayed by flow
cytometry 18 hours after transfection. See also Fig. S4.
227
Fig. 5. Workflow for experiments comparing two methods of analyzing reporter
expression. Following transfection, an aliquot of cells is analyzed by flow
cytometry. From the remaining cells, RNA is isolated and an aliquot is subjected
to Illumina sample preparation and sequencing. FM-HCR analysis of fluorescent
reporter expression, is compared to HCR-Seq analysis of reporter transcript
expression, measured as RPKM. See also Tables S1-5.
Fig. 6. mRNAseq analysis of reporter expression. (a) Dose response curves for
reporter expression from randomly damaged plasmids generated from mRNAseq
analysis. (b) Dose response curves for the same cells generated from flow
cytometric (FM-HCR) analysis. (c) Sequence variants detected in transcripts at
the position corresponding to the site-specific thymine dimer in the absence (top)
or presence of the lesion (bottom). Frequencies are reported for the expected
sequence (AA) as well as all variants that were observed in at least one sample.
(d) Frequencies of AA->GA mutations in transcripts expressed from randomly
damaged plasmids as a function of dose (combination #7 in Table 2). The
undamaged case (0 J/m 2) refers to the frequency of mutations as measured in
transcripts expressed from the BFP transfection control. "U" refers to HCR-Seq
data from cells in which all of the reporter plasmids were undamaged, and "D"
refers data for cells transfected with reporters irradiated as indicated in Table 2.
Symbols (*) represent differences that were deemed to be statistically significant
(p<0.05) by a t test. See also Fig. S5.
228
FIGURES
Fig. 1
Transfection -, Time
-M*"
Cytometry
-MM R
100
Do 
- WT
XPc
XPA
XPD
XPA
200 400 600 800
WT
10
XPc
XPA
XPD
0 200 400 600 600XPA
1001WT
XPc
10
XPB
11 XPA
0 200 400 600 800
C
g
k
10a
D,- WT
10
XPc
XPA
XPD
XPA0 200 400 600 80
-WT100
XPA
XPD
XPA
0 200 400 W O 000
1()oWT
10 XPC
XPB
11 XPA
0 400 600 800
d
U
Ir
h
U-
L~i
CL
E
Dose to Plasmid, J/m 2
60
40-
20
0 2o 40 60 80
% R.E. CAT-HCR
80
60-
4 0
20-
00 20 40 60 8O
% R.E. CAT-HCR
60-
40 T
20-
0',T
0 20 40 60 80
% R.E. Fibro.
e
900
W
r600.
300
0 300 600 900
D., CAT HCR
c.
3t
80-
60
40
20
0 20 40 60 80
% R.E. #1
Panels b-i Panels j-1
10 01953(0W) "i (VNT)
G?0J36w/ (W01) 1i(0)
C 2240 (XPC) vi (WT)
G002246 (XPC) i (vw)
GM02344 (XPA) v(XPCJ
GM2253(XPD) v (XPB)
GM02345 (XPA) t (XPA)
229
b
0
u.J
L0-
0
f i
I
Fig. 2
a Plasmids
NER Repaired
800 J/m2 GFP
G
C
R1 p 
r0
morng
0
4-J
L..
0
a-
a)0
DR Repaired;
06MeG 
_ _non
fluorescent
NER Separate
20-
MMR Separate
OhMeG Separate
U1 III0
10~ ~
(1C9
NER Together
20
15]
MMR Together
ObMeG Together
10
MMR
mGt
-nsac
230
b FM-HCR
a Plasmid combo #4
N HEJ
NER
Qe
G
MMR
MG MT
d Plasmid combo #5
Oo NER
6 ER
I BER0
Oe! MGMT
b Separate
100-
0
LU
t
0~Q)
01
c Together
M059K M059
(WT) (DNA PKcs)
e Separate
1001
.0
01
0.a- 0
0 01
1 00-
10-
0.1
M059K M059J
(WT) (DNA PKcs)
f Together
100
10
1
0.1
WT Ogg1- - 0 WT Ogg 1'
Fig. 3
0s1
0 sau6ose
w NHEJ
3 NER
m' MMR
- MGMT
m NER
c MMR
r- BER
- MGMT
231
NHEJ
S MMR
& A MGMT
Cell lines 2 HR
00Q
c
(,020
0010
0010
U
=0.81 c
U
4 -2 0 2 4
In(MGMT HCR % R.E.)
Lu
z
()
cr-
1 0d
0.5t
0.0 
C
cpI 1K
100
U
0- 10(3
HR
MMR
NER
NHEJ
0 10 20 30
Compound 401], uM
Fig. 4
a
0
U,
0.
1c
60
50
40
30
20
0
01
b
0
-J
CL
.7-
cvM- (M0
P0
232
-~ -2
Fig. 5
Transfection
c
0
L
A)
0-
0
aO
.5:
Lf)
4-J
I1M r RNA Isolation
Illumina mRNA library
L -------------- I 
Cytometry RNA sequencing
Ir
F FM-HCR analysis of HCR-Seq Analysis of reporter
reporter expression expression levels and
levels transcriptional mutagenesis
233
------
i
I
I I
00
10
1
0 400 80
b
0WT U
x
w
XPA
0
-00,
1 00
0
0 400 800
C
WT
PA VI,
_0
0)
0.20 0 i/r 2  0.20 200 i/r 2
0.15 0.15
0.10 0.10
0.05 0.05
0.00 0.00
UD UL) UD U D
400 J/m2  800 J/M 2
0.20 0.20
0.15 0.15
0.10 0.10
0.05 0.05
0.00 U D U b 0.00 U D U D
No Lesion
10
95
2.
~~e 0
100 Site-Specific T
95
90
6
4,
0i
Observed Sequence
Fig. 6
-WT
tXPA
1
C0
d
0
=3
E
0
<>T
234
a
1
TABLES:
Table 1. 55 cell lines used for this study. To facilitate comparison of data, the
seven individuals from whom both lymphoblastoid and fibroblast cultures were
derived have been assigned indexes i through vii.
235
Cell Line
GM01630 (i)
GM01 953
GM02246
GM02249
GM02253
GM02344 (i)
GM02345
GM03657 (ii)
GM03658 (ii)
GM07752 (iii)
GM07753 (iii)
GM14878 (iv)
GM14879 (iv)
GM21071 (v)
GM21148 (v)
GM21677 (vi)
GM21833 (vii)
GM23249 (vi)
GM23251 (vii)
TK6
MT1
TK6+MGMT
HCT116
HCT1 16+3
M059J
M059K
WT MEFS
Ogg1- MEFS
V79 (Hamster)
VC8 (Hamster)
xrs6 (Hamster)
GM15029 (#1)
GM 15036 (#2)
Cell Type
Fibroblast
Lymphoblastoid
Lymphoblastoid
Lymphoblastoid
Lymphoblastoid
Lymphoblastoid
Lymphoblastoid
Lymphoblastoid
Fibroblast
Lymphoblastoid
Fibroblast
Lymphoblastoid
Fibroblast
Fibroblast
Lymphoblastoid
Lymphoblastoid
Lymphoblastoid
Fibroblast
Fibroblast
Lymphoblastoid
Lymphoblastoid
Lymphoblastoid
Colorectal
Carcinoma
Colorectal
Carcinoma
Glioblastoma
Glioblastoma
MEFs
MEFs
Fibroblasts
Fibroblasts
CHO
Lymphoblastoid
Lymphoblastoid
Genotype
XPA
WT
XPC
XPC
XPD
XPA
XPA
WT
vr
WT
WT
XPC
XPC
XPB
XPB
Wr
WT
WT
WT
MGMT
MGMT,
MSH6
WT
MLH1
WT
DNA PKcs
WT
WT
Ogg1
WT
BRCA2
Ku80
WT
WT
Repair Defect
NER, severe
None
NER, moderate
NER, mild
NER, severe
NER, severe
NER, severe
None
None
None
None
NER, very mild
NER, very mild
NER, severe
NER, severe
None
None
None
None
DR of O6MeG
MMR and DR of
O6MeG
None
MMR
None
NHEJ
None
None
BER of 8-oxoG
None
HR
NHEJ
None
None
GM15215 (#3) Lymphoblastoid WT None
GM15223 (#4) Lymphoblastoid WT None
GM15245 (#5) Lymphoblastoid WT None
GM15224 (#6) Lymphoblastoid WT None
GM15236 (#7) Lymphoblastoid WT None
GM15510 (#8) Lymphoblastoid WT None
GM15213 (#9) Lymphoblastoid WT None
GM15221 (#10) Lymphoblastoid WT None
GM 15227 (#11) Lymphoblastoid WT None
GM15385 (#12) Lymphoblastoid WT None
GM15590 (#13) Lymphoblastoid WT None
GM15038 (#14) Lymphoblastoid WT None
GM15056 (#15) Lymphoblastoid WT None
GM15072 (#16) Lymphoblastoid WT None
GM15144 (#17) Lymphoblastoid WT None
GM15216 (#18) Lymphoblastoid WT None
GM15226 (#19) Lymphoblastoid WT None
GM15242 (#20) Lymphoblastoid WT None
GM15268 (#21) Lymphoblastoid WT None
GM15324 (#22) Lymphoblastoid WT None
GM15386 (#23) Lymphoblastoid WT None
GM15061 (#24) Lymphoblastoid WT None
Table 2. Combinations of reporter plasmids and types of DNA damage used in
each experiment.
Combination
#1
#2
#3
#4
#5
#6
#7
tagBFP
600 J/m2
No lesion
No lesion
DSB
800 J/m 2
DSB
No lesion
AmCyan
No lesion
200 J/m 2
No lesion
No lesion
200 J/m 2
EGFP
800 J/m2
400 J/m 2
800 J/m 2
800 J/m 2
A:CA
DSB
T<>TB
mPlum
40 J/m2
800 J/m 2
06-MeGD
O6-MeGD
06-MeG"
No Lesion
800 J/m
2
AA:C mismatchBSite specific thymine dimer
cG:G mismatch
D 0 6
-MeG
236
mOrange
200 J/m
600 J/m 2
G:Gc
G:GC
8-oxoG
400 J/m2
SUPPLEMENTAL FIGURES
a _GF.P b AmnCyan c mOrange d Minus Green
-8 C- P8/ / P8 p
'IT 0' 1 3V10 3311 1 F) 10,
FITC-A FIT-A FT
13. 1 1 11 , 0 10 103 10 1' 01 13 30 1031 0 1
FllA FIT3>0 A TC- FIT'4
e Plasmid ChLP
Lymphoblasts (GM21833)
transfected with pMax-mCherry
(D
0
E
c
0
U_
1000-
100-
10
I * -I-=1
Fibroblasts (GM23251)
transfected with pMax-mCherry
1000]
(~
0
E
U)
0c
LL
antibody PCR primer combination
100-
10.
It1
antibody PCR primer combination
Fig. S1; related to Fig. 1. Transfection controls and method of establishing the
positive region for a fluorophore requiring compensation in two or more channels.
237
(.2~8
E
1-
6 e e
Both the mOrange and AmCyan reporters overlap significantly with the GFP
reporter; they result in significant signal in the GFP (FITC) channel even when
compensation is applied, creating the potential for false positives. In this
example, we illustrate why no single gate is sufficient to identify true positive GFP
cells in the presence of mOrange and AmCyan. The problem is overcome by
establishing a region that represents the union of two or more gates; cells must
appear in both gates to be counted as positive for GFP. Each panel in the figure
indicates the fluorescent reporter or mixture of reporters being expressed in the
cells analyzed. The detector channels are as follows: AmCyan = Cyan, FITC =
GFP, PE YG = mOrange. a) Flow cytometry plots for cells transfected with the
GFP reporter. Bright fluorescence is detected in the FITC channel b) Flow
cytometry plots for cells tranfected with the AmCyan reporter plasmid.
Application of compensation leads to a typical "funnel" shape (the brightest cells
in the AmCyan channel fan out into the FITC channel). Gate P8 is drawn in such
a way as to exclude them as false positives. However, on a plot of fluorescence
in the FITC channel against that in the PE YG channel, these cells are
indistinguishable from GFP positive cells, and would be counted as false
positives if gate P9 were used alone as the region corresponding to GFP positive
cells. c) Plots for cells transfected with the mOrange reporter. Some cells
appearing in gate P8 are indistinguishable from the GFP positive cells in panel
(a). When fluorescence in the same cells is plotted as FITC vs. PE YG, it is seen
that the false positives arise from the same "funnel" phenomenon occurring for
AmCyan in in the upper plot in panel (b). As a result, using P8 alone would also
be insufficient for the identification of true GFP positive cells in the presence of
the other two fluorophores (mOrange and AmCyan). However, these false
positives can be excluded using gate P9. Taken together, the observations in (b)
and (c) indicate that a GFP positive region defined as the union of regions P8
and P9 represents true positives. d) Plots for cells transfected with all reporters
except for the GFP reporter. If needed, gates are further adjusted to minimize
the number of cells appearing in the region P8+P9. e) Chromatin
immunoprecipitation of plasmid DNA. The lymphoblastoid cell line GM21833 and
238
the primary human fibroblast culture GM23251 were electroporated with the
pmax:mCherry reporter plasmid and analyzed for plasmid chromatinization.
Antibodies raised against the human histone H3 and H4 were used in all
experiments. H3 precipitation yielded an enrichment of at least 40-fold for the
reporter plasmid DNA in both cell types compared with chromatin precipitated
with the nonspecific antibody IgG. Error bars represent the standard deviation of
triplicate measurements. Similar enrichment was observed for the host gene
glyceraldehyde 3-phosphate dehydrogenase (GAPDH). When DNA was
immunoprecipitated with the antibody to histone H4, an enrichment of at least 2-
fold was observed for the plasmid DNA and similar enrichment was found for
GAPDH. Overall, the results are consistent with incorporation of plasmid DNA
into nucleosomes.
239
GMMR C
mismatch
A AAAA 3
-AAAA
O5MeG MGMT 01
Extendj
LgateQ
b
C
1 kb
G IGGCCC-. Gs M
PspOMl C
-GGGCCC- -GGGCCC-
-CCCGGG- - -CCCGGG- -
Nb Btsl
Deture, +O Anneal, Up~te
0
d,:5 20
4 67 915
fX
1
X 3
10 kb- 1 2 3 4 5 6 7 8
4kb
2 kb
Fig. S2; related to Fig. 2. Construction and validation of plasmid reporters for
MMR and MGMT. a) Fluorescent reporters for mismatch repair and MGMT. Top,
repair of a G:G mismatch restores the wild type sequence to the transcribed
strand, and results in expression of orange fluorescent protein. Bottom, use of
transcriptional mutagenesis to measure repair of a site-specific 06-
methylguanine lesion. The lesion induces misincorporation of uracil into
transcripts. The uracil containing transcripts are translated into wild type protein.
Following repair of the 06-methylguanine lesion, transcripts contain cytosine at
the relevant position, and encode a non-fluorescent protein. b) Synthetic method
for generation of heteroduplex plasmids. The transcribed strand of the reporter
plasmid is nicked with a strand specific nicking endonuclease. The nicked strand
is then digested with exonuclease Ill. The resulting closed circular single
stranded DNA (ssDNA) is then combined with linearized double stranded DNA
prepared from a second plasmid that differs by a single nucleotide; this sequence
change prevents expression of fluorescent protein. The two molecules are
240
0
0
6
denatured with sodium hydroxide, and then brought to neutral pH to facilitate
annealing between the circular ssDNA and the complementary strand from the
linearized DNA. Unwanted side products (ssDNA and linear DNA) are selectively
digested by plasmid safe ATP dependent nuclease (Epicentre). The desired
heteroduplex DNA is finally ligated to produce a closed circular double stranded
heteroduplex. c) Gel analysis of starting materials, intermediates and products of
MMR substrate preparation. The material in lane 9 was used for HCR assays.
Lane 1, Uncut; lane 2, Nb.BtsI (Nicked DNA); lane 3, Nb.BtsI + Exolli (ssDNA);
lane 4, Nhel (linear DNA); lane 5, Annealing product of #3 and #4; lane 6, #5 +
PSAD; lane 7, #6 following gel extraction; lane 8, #7 following ligation; lane 9, #8
following gel extraction. d) G:G mismatch repair in MMR-proficient HCT1 16+3
cells and MMR-deficient HCT1 16 cells. e) Synthetic method for 06-
methylguanine reporter, and molecular basis for resistance to reporter cleavage
by PspOMI. An oligo containing a site-specific 06-methylguanine lesion is
annealed to single stranded DNA (non-transcribed strand), followed by primer
extension and ligation to yield closed circular DNA. In the absence of 06-
methylguanine, PspOMI cleaves the mPlum C207G:T208C reporter plasmid at
the recognition site GGGCCC. The presence of 06-methylguanine prevents
recognition by the enzyme, thus the reporter plasmid containing the site specific
lesion in the transcribed strand is resistant to cleavage. f) Assay for site specific
incorporation of 06-methylguanine into the C207G:T208C mutant of the mPlum
reporter plasmid. This assay was performed as an independent confirmation of
the presence of the lesion, and to minimize the possibility that fluorescent signal
might arise due to the presence of unmodified bases or other lesions. Primer
extension reactions were performed with single stranded DNA from the
C207G:T208C mutant of the mPlum reporter plasmid, which contains the
GGGCCC recognition site for the restriction enzyme PspOMI. It has been shown
previously that 06 -methylguanine can abolish recognition of restriction sites (Wu
et al., 1987). In the analytical digest shown below, we find that 06-
methylguanine blocks cleavage of the plasmid to at least 90%, and that removal
of the lesion by MGMT restores PspOMI cleavage at the restriction site to near
241
100%. The data are consistent with greater than 90% incorporation of the
desired lesion at the intended position. Lane 1, 06-MeG plasmid, no treatment;
lane 2, 06-MeG plasmid + PspOMI; lane 3, 06-MeG plasmid + MGMT; lane 4,
06-MeG plasmid + MGMT + PspOM; lane 5, Homoduplex; lane 6, Homoduplex
+ PspOMI; lane 7, Homoduplex + MGMT; lane 8, Homoduplex + MGMT +
PspOMI.
242
aONt.BspQl ExoIll
2.5-
2.0-
0. 
5O ~Extend, a
Ligate Anneal CL 1.5-
Sca)
00 1.0
00
cci 
-GAT AGT ACT )
T6 MT1
(WT) (MSH6)
Fig. S3; related to Fig. 3. Construction and validation of reporters for MMR and
NHEJ. a) Synthetic method for A:C mismatch in a GFP reporter. The non-
transcribed strand of the C289T mutant GFP reporter plasmid is nicked with
Nt.BspQ, and digested from the plasmid with Exoll. An oligo with the wild type
sequence (G at the position opposite C289) and complementary to the region of
the plasmid that has been mutated is annealed and extended to form a
heteroduplex with an A:C mismatch. b) Validation of GFP reporter for MMR
repair of an A:C mismatch. As expected, reduced reporter expression was
observed in the MSH6-deficient MT1 cell line relative to the WT TK6 cell line. C)
Structure of the BFP reporter for NHEJ. An Scal recognition site (AGTACT) was
inserted upstream of the reporter gene (BFP, represented by a blue arrow).
Plasmids were linearized with the Scal restriction enzyme, which leaves a blunt-
end double strand break in the 5' untranslated region of the reporter plasmid.
243
ab C 100
NHEJ Deficient NHEJ
M IMMR
3 0 10 10.
s HR Deficient
19. 0.1
V79 VC xrs6
00
EGP(WT) (BRCA2) (Ku8O) 0.01 -Together Separate
Fig. S4; related to Fig. 4. Homologous recombination plasmid reporters and
assay validation. (a) Recombination between two plasmids (one circular and one
linearized) that express truncated non-fluorescent proteins results in a plasmid
that expresses full-length green fluorescent protein. NHEJ-mediated repair of the
linearized reporter plasmid does not give rise to fluorescent protein expression.
(b) BRCA2-deficient VC8 hamster cells exhibit deficient homologous
recombination relative to wild type V79 hamster cells. Ku8O-deficient xrs6
hamster cells are NHEJ deficient, and elevated HR is observed, consistent with
the competition between NHEJ and HR for repair of DSBs. (c) Cells were
transfected with a combination of reporter plasmids for NHEJ, HR and MMR
either in separate transfections or together in a single transfection. % Reporter
expression under the two conditions was indistinguishable within the
experimental error of the measurements (error bars represent the standard
deviation of 3 experiments).
244
GM01953 GM0236A b 8 0  1 %3 H,,Mhy
601GM02344 C
-40
20 GM01953
GM01953 GM02344 0
M Intron 4 Exon 4
0- PA Exon 5 Intron 4 Exon 4
W X~~PAI . . .
/o /GM02344 SO M 2%0 "I kined invon 98%
Log2 gene expression in cells transfected GM019S3
with undamaged plasmid Splwd 48%
d 100 1 2 3 4 e f800
600 
"
6 -GM01953
10 = GM02344 +
62 4
2
0,
Fig. S5; related to Fig. 6. RNAseq analysis of transfected cells and construction
of a plasmid with a site-specific thymine dimer. a) Gene expression profile of
cells transfected with damaged or undamaged reporter plasmids. Panels a and b
represent the two replicate measurements, for the cell lines indicated above each
plot. Levels of expression in cells that were transfected with damaged plasmids
are plotted on the vertical axis, and expression levels in cells transfected with the
undamaged (control) plasmids are plotted on the horizontal. Genes expressed at
the same level under both conditions appear on the diagonal, and this is
overwhelmingly the case for endogenous transcripts (black and gray circles),
indicating no major changes in transcription in cells in response to the presence
of damaged plasmid DNA. Reporter transcripts are colored in blue, cyan,
orange, green, and magenta. These reporters are seen to be among the most
highly expressed in all samples. Reduced expression in the presence of DNA
damage (due to transcription blocking lesions) is reflected in these points falling
245
below the diagonal. b) Expression of GFP mRNAfrom reporter plasmid
containing a site-specific thymine dimer in the transcribed strand. Expression
was assayed by flow cytometry or RNAseq. Error bars represent the standard
deviation of two biological replicates. c) XPA read coverage and junction reads
for GM02344 and GM01953. Exons 4 and 5 are shown. A 555G>C mutation in
exon 4 of the XPA gene has previously been reported to induce transcript
splicing errors in GM02344 (Satokata et al., 1992). Reads (indicated as small red
and blue bars) are aligned to the region of the genome that encodes the XPA
gene. The majority of reads in transcripts from the wild type cell line GM01953
align, as expected, to the exons. Read coverage for GM01953 is overall higher
than that for GM02344; XPA expression was ~ 2.5-fold higher in in GM01953.
The expected intron-spanning reads (indicated as light gray lines that run
between exons) are abundant for GM01953, and ~ 50% of the transcripts were
correctly spliced. By contrast, read coverage is lower, intron-spanning reads are
nearly absent, and many reads fall within the introns for GM02344. Only -2% of
the XPA transcripts in the mutant cell line were correctly spliced. d) Agarose gel
(top) and sequencing (bottom) analysis of PCR amplicons generated from
reporter cDNAs. In the gel, PCR products were analyzed from reactions where
the template was excluded (lane 1), plasmid DNA (lane 2), purified mRNA that
was not reverse transcribed (lane 3), or cDNA generated by reverse transcription
of mRNA (lane 4). No product was observed in the absence of reverse
transcription, and amplicons generated from cDNA templates migrated 100-200
bp below amplicons generated from plasmid DNA templates; this size difference
corresponds to a 136 bp intron expected to be absent from cDNA, but retained in
plasmid DNA. The frequency of AA->GA base substitutions in transcripts
expressed in GM01953 and GM02344 from plasmids containing a site-specific
thymine dimer are within experimental error of the frequencies measured for
AA-GA substitutions using RNAseq (Fig. 6c). e) Preparation of a reporter with
a site-specific thymine dimer. The GFP reporter plasmid contains two
recognition sites, 18 bp apart, for the strand specific nicking endonuclease
Nb.Bpul0. The plasmid is nicked with this enzyme, heated to melt the
246
oligonucleotide away from the plasmid, and then rapidly cooled to prevent the
oligo from re-annealing. The oligo is then removed from the mixture using a
Qiagen PCR cleanup kit. Plasmid is then combined with an excess of a 5'-
phosphorylated oligonucleotide containing a site specific thymine dimer, heated
to 80 C, and cooled slowly to facilitate annealing of the oligonucleotide. Finally,
the resulting nicked plasmid is ligated and the desired closed circular DNA is
purified from the mixture by gel electrophoresis. f) Assay for site-specific
incorporation of a cyclobutane pyrimidine dimer into the pmax:GFP plasmid. 500
ng of plasmid was incubated in a 50 pL volume for 16 hours at 37*C with 40 units
of the bifunctional thymine dimer specific glycosylase / AP lyase (T4 PDG, New
England Biolabs), which nicks DNA that contains thymine dimers. Untreated
plasmid (lane 2) is resistant to cleavage, whereas plasmid irradiated at 800 J/m2
is completely digest (lane 6). A second band in lane 6 migrates as linearized
DNA (-3.7 kb) and likely reflects nicking at closely opposed DNA lesions. Nearly
complete digest of the plasmid in Lane 4 to nicked DNA is consistent with at least
95% incorporation of the lesion.
247
SUPPLEMENTAL TABLES
Table SI; related to Fig. 5. Samples submitted for next generation sequencing.
8 total samples were submitted for complete RNA-seq using 40bp paired end
libraries. These included damaged and undamaged samples for the two cell
lines and two replicates for each cell line.
Sample ID Sample name Cell line Replicate
D12-4969 WT undam GM01953
D12-4970 WT dam GM01953
1
D1 2-4971 XPA mut undam GM02344
D1 2-4972 XPA mut dam GM02344
D12-4973 WT undam GM01953
D12-4974 WT dam GM01953
2
D1 2-4975 XPA mut undam GM02344
D12-4976 XPA mut dam GM02344
248
Table S2; related to Fig. 5. Cufflinks and Tophat parameters.
Tophat Value Description
parameter
--min-anchor-length 6 TopHat will report junctions spanned by reads with at least this
many bases on each side of the junction. Note that individual
spliced alignments may span a junction with fewer than this many
bases on one side. However, every junction involved in spliced
alignments is supported by at least one read with this many bases
on each side. This must be at least 3 and the default is 8.
--splice-mismatches 0 The maximum number of mismatches that may appear in the
"anchor' region of a spliced alignment. The default is 0.
--min-intron-length 10 minimum intron size allowed in genome
--max-intron-length 1000000 maximum intron size allowed in genome
--min-isoform-fraction 0.0 The minimum frequency of any isoform to consider. The default is
0.15
--max-multihits 20 Instructs TopHat to allow up to this many alignments to the
reference for a given read, and suppresses all alignments for
reads with more than this many alignments. The default is 20 for
read mapping.
--no-novel-juncs True Only look for reads across junctions indicated in the supplied GFF
or junctions file.
--segment-length 20 Each read is cut up into segments, each at least this long. These
segments are mapped independently. The default is 25.
--library-type fr- library prep used for input reads
unstranded
--solexal.3-quals True As of the Illumina GA pipeline version 1.3, quality scores are
encoded in Phred-scaled base-64. Use this option for FASTQ files
from pipeline 1.3 or later.
--mate-inner-dist 200 This is the expected (mean) inner distance between mate pairs.
For, example, for paired end runs with fragments selected at
300bp, where each end is 50bp, you should set -r to be 200. The
default is 50bp.
--mate-std-dev 100 The standard deviation for the distribution on inner distances
between mate pairs. The default is 20bp.
Cufflinks Value Description
parameters
249
--min-intron-length 10 minimum intron size allowed in genome
--max-intron-length 1000000 maximum intron size allowed in genome
--min-isoform-fraction 0.0 The minimum frequency of any isoform to consider. The default is
0.15
--library-type fr- library prep used for input reads
unstranded
--compatible-hits-norm True count hits compatible with reference RNAs only
--multi-read-correct True use 'rescue method' for multi-reads (more accurate)
--frag-bias-correct True use bias correction - reference fasta required
Table S3; related to Fig. 5. RPKM values for the five reporter genes across
samples.
RPKM values
Replicate Reporter gene XPA mut undam XPA mut dam Norm undam Norm dam
BFP 1737.01 1213.61 1607.05 2076.39
AmCyan 14975.3 5993.3 13792 16456.5
1 GFP_615 1434.24 463.966 1251.22 984.1
mOrange 2192.32 239.698 1853.39 2164.26
mPlum 4257.75 162.343 3723.82 2928.89
BFP 1652.01 1096.74 1467.33 1608.85
AmCyan 18999.2 4596.23 15349 16521.4
2 GFP_615 1283.83 447.682 1314.83 652.47
mOrange 1998.8 193.741 2119.65 1532.83
mPlum 3706.3 108.021 4299.3 2027.84
250
Table S4; related to Fig. 5. Read counts for RNA-seq samples and numbers of
aligned reads using TopHat.
Norm Norm XPA mut XPA mut Norm Norm XPA mut XPA mutSample names
undam dam undam dam undam dam undam dam
Replicate 2
Sample ID D12-4969 D12-4970 D12-4971 D12-4972 D12-4973 D12-4974 D12-4975 D12-4976
4119316
Total sequences 46727196 42643848 42485978 41062392 49343810 45472418 49352500
0
4897171
Total mapped reads 55847402 50879779 52191219 47970477 57600436 54420536 58422772
0
3585149
Total 41264548 37791752 38687218 35630500 42814870 40359416 43174990
8
BFP 23230 26733 22671 11979 21405 19408 23312 13147
AmCyan 245632 265119 232280 78184 277105 275550 357715 71185
Mapped GFP_616 16261 12068 17476 3728 17887 7859 16816 5011
properly mOrange 21258 22148 24496 1596 26262 13090 23339 1978
mPlum 35921 29101 41173 1194 44851 17103 38084 1531
Plasmids 342302 355169 338096 96681 387510 333010 459266 92852
3575864Other genes 40922246 37436583 38349122 35533819 42427360 40026406 42715724
6
Unmapped reads 245774 243730 228043 192470 337249 241637 215238 164415
1312021Total 14582854 13088027 13504001 12339977 14785566 14061120 15247782
2
Other
Read mapped, 1449291 1385025 1359536 1159168 1606305 1427621 1463587 1257407
genes
mate
Plasmid
unmapped 98603 75626 79587 57255 94179 71455 100645 20241
5
Pair not
Read and Other
mapped 7914228 6810016 7155202 7004002 7688172 7239820 8356254 7280056
mate mapped, genes
properly insert size too Plasmid
718 736 818 676 720 564 802 276large s
Read and Other
5118728 4815446 4907660 4117100 5395452 5321062 5325798 4561628
mate mapped genes
to different Plasmid
1286 1178 1198 1776 738 598 696 604
chrom s
251
Table S5; related to Fig. 5. Genes with log2 fold change >= 1 when comparing
cells transfected with undamaged plasmid to those transfected with damaged
plasmids.
Sample Replicat Gene Log2 Fold
e Name Chr Bp Change
GFP_615 GFP_615 0-951 -1.62
AmCyan AmCyan 0-921 -1.32
MALAT1 chr1 1 65265232-65273939 -1.29
SCARNA9 chrl 1 93454679-93455032 -1.06
RPL21 chrl3 27825691-27830702 -1.38
NDUFA3 chr1 9 54606159-54610281 -1.13
IN080B-WBP1 chr2 74682149-74688018 -1.00
GM0234 HIST1H4H chr6 26285353-26285727 -1.76
chrUn_gIO002
RN5-8S1 20 112024-112180 -4.07
chrUn-gIOO02
RN5-8S1 20 155996-156152 -4.07
mOrange mOrange 0-942 -3.19
mPium mPlum 0-912 -4.70
GFP_615 GFP_615 0-951 -1.52
2 AmCyan AmCyan 0-921 -2.04
mOrange mOrange 0-942 -3.37
mPlum
C1orf86
NUBP2
ATP5D
FAM108A1
TMEM160
SCAND1
C4orf48
TMUB1
C9orf16
FBXWS
BCYRN1
RPL21
SMN1
SMN1
RN5-8S1
mPlum
chri
chrl6
chrl9
chr19
chr19
chr20
chr4
chr7
chr9
chr9
chrX
chr13
chr5
chr5
chrUn-g0002
0-912
2115898-2139172
1832932-1839192
1241748-1244824
1876974-1885518
47549166-47551882
34541538-34543281
2043719-2045697
150778171-
150780620
130922538-
130926207
139834884-
139839206
70430034-70948962
27825691-27830702
69345349-69373418
70220767-70248838
112024-112180
-5.10
1.02
1.05
1.00
1.05
1.05
1.34
1.42
1.00
1.02
1.01
1.41
2.53
1.50
1.43
1.92
GM0195
3
2
252
1
20
chrUn_gIO002
RN5-8S1 20 155996-156152 1.92
GFP_615 GFP_615 0-951 -1.01
C16orfl3 chrl6 684428-686347 -1.01
ATP5D chrl9 1241748-1244824 -1.07
LOC100129250 chr9 32551141-32553015 -1.00
mPlum mPlum 0-912 -1.08
SUPPLEMENTAL EXPERIMENTAL PROCEDURES
Plasmids
The AmCyan, EGFP, mOrange, and mPlum and tagBFP reporter genes were
subcloned into the pmax cloning vector (Lonza) between the KpnI and Sacl
restriction sites in the multiple cloning site. The Kozak translation initiation
consensus sequence and an additional Nhel restriction site were introduced at
the 5' end of each reporter, and a Hindll restriction site was added to the 3' end.
The pmax cloning vector places reporter genes under the CMV Intermediate-
Early promoter. Plasmids were amplified using E. coli DH5a (Invitrogen), and
purified using Qiagen endotoxin-free maxi and giga kits. Constructs were
confirmed by DNA sequencing and restriction enzyme digestion.
UV-Irradiated Substrates
Plasmids were irradiated in TE buffer (10 mM Tris-HCI, 1 mM EDTA, pH 7.0) at a
DNA concentration of 50 ng/pL in a volume of 1.5 mL in 10 cm polystyrene petri
dishes (without lids) with UVC light generated by a Stratalinker 2000 box.
Following treatment, reporter plasmids were combined in the following ratio: 1
part tagBFP, 10 parts AmCyan, 1 part GFP, 2 parts mOrange, and 4 parts
mPlum; these proportions were used to compensate for weaker fluorescence
intensities observed for some of the reporters. The same plasmid mixture
without UV irradiation was prepared as described above, except without UV-
253
treatment. Data obtained following transfections with the plasmid mixture
containing irradiated plasmids have been labeled "damaged" and those from
untreated plasmids have been labeled "undamaged"; however every transfection
included an undamaged reporter. The undamaged transfection reporter was
used to normalize transfection efficiency. Further details regarding the UVC
dose delivered to each plasmid are available in Table 2. Plasmid mixtures were
ethanol precipitated, washed with 70% ethanol, and redissolved in TE buffer at ~
1.5 pg/pL; damaged and undamaged plasmid mixtures were adjusted to the
same final concentration, confirmed using a Nanodrop spectrophotometer.
Substrates containing a G:G mismatch
Substrates were prepared using a method based on a previously published
protocol (Baerenfaller et al., 2006). The pmax:mOrange plasmid was nicked with
Nb.Bst (New England Biolabs) to generate a single strand break in the
transcribed strand (Fig. S2b). The nicked strand was then digested with
exonuclease Ill, and the remaining single stranded circular DNA (ssDNA) purified
using a 1 % agarose gel. 20 pg of the ssDNA was combined with 40 pg of G299C
mutant pmax:mOrange plasmid linearized with Nhel (New England Biolabs); the
mixture was denatured by addition of 0.3N sodium hydroxide, and then returned
to neutral pH to facilitate annealing between wild type ssDNA and the
complementary strand of the linearized mutant sequence to yield a heteroduplex
containing a G:G mismatch at position 299 of the mOrange gene. Subsequently,
reactions were cleaned up using a Qiagen PCR cleanup kit, and unwanted linear
and single stranded DNA side products were digested with Plasmid Safe ATP
dependent DNAse (Epicentre). Nicked plasmid was purified using a 1 % agarose
gel, and ligated using 800 units T4 DNA ligase (New England Biolabs). Finally, a
second gel purification was performed to isolate closed circular products (Fig.
S2c). Homoduplex DNA was prepared using the same procedure, except that
linearized DNA was prepared from the wild type pmax:mOrange.
254
Substrates containing a site-specific 06-MeG
A nonfluorescent variant of the pmax:mPlum reporter (T208C) was identified.
This construct was further modified with a mutation that does not change the
encoded protein sequence (C207G) to generate a unique recognition site
(GGGCCC) for the restriction enzyme PspOMI (New England Biolabs).
Substrates were prepared based on a previously described method (Baerenfaller
et al., 2006), with minor modifications (Fig. S2e). Single stranded DNA was
prepared as described above. 5 picomoles of a phosphorylated 06-MeG
containing oligonucleotide (5'-CACGTAGGCCTTGGXCCCGTACATGATCTG-3',
where X = 06-MeG) was combined with 2.5 pg of single stranded
pmax:mPlum:C207G:T208C plasmid DNA in 1X Pfu polymerase buffer (Agilent
Biotechnologies) in a total volume of 50 pL. The mixture was heated to 850C in a
thermal cycler for 6', and then allowed to anneal by cooling to 400C at 10C per
minute. To extend the primer, 2.5 units Pfu polymerase (Agilent) and 0.2 pM
dNTPs were added and the reaction, and then incubated for 1 hour at 680C. The
reaction was then cooled to 370C, and supplemented with an additional 0.5 pM
dNTPs, 1 mM ATP, 1.5 units T4 DNA polymerase (New England Biolabs), and
40 units T4 DNA ligase, and incubated for an additional hour at 370C to yield
closed circular plasmid. Finally, the product was purified from a 1% agarose gel
using a Qiagen gel extraction kit. A homoduplex control plasmid that expresses
the wild type mPlum fluorescent reporter protein was prepared using identical
conditions with single stranded pmax:mPlum:C207G and the following
oligonucleotide: 5'-CACGTAGGCCTTGGACCCGTACATGATCTG-3'. The
plasmid containing 06-MeG was resistant to cleaveage by the restriction enzyme
PspOMI, whereas the lesion free homoduplex generated with the same ssDNA
was readily digested under the same conditions. Treatment of the 06-MeG
containing plasmid with human methylguanine methyltransferase (hMGMT)
(Alexis Biochemicals) resulted in >95% PspOMI cleavable material (Fig. S2f).
Substrates containing a site-specific 8-oxoG
255
A non-fluorescent variant of mOrange (A215C) that lacks a critical tyrosine that
forms the chromophore was identified. The plasmid encoding this non-
fluorescent protein was used to prepare ssDNA using the same protocol
described above for preparation of the 06-MeG containing plasmid. The following
8-oxoG containing oligonucleotide was annealed and extended using the same
conditions that were used to prepare the 0-MeG containing plasmid: 5'-
GTAGGCCTTGGAGCCGXAGGTGAACTGAGG-3', where X represents the 8-
oxoG lesion. An mOrange homoduplex was prepared using the following oligo
with ssDNA prepared from the wild type mOrange plasmid: 5'-
GTAGGCCTTGGAGCCGTAGGTGAACTGAGG-3'.
Substrates containing an A:C mismatch
Substrates were prepared using a method similar that described above for
preparation of the 06-MeG and 8-oxoG containing plasmids (Fig. S3a). A non-
fluorescent GFP variant (C289T) was identified. The protein expressed from this
construct lacks a conserved arginine that is required for chromophore maturation.
Single stranded DNA was prepared from this plasmid as described above, except
the nicking enzyme was Nt.BspQ, which nicks the non-transcribed strand. Thus,
after Exoill digest, the remaining ssDNA comes from the transcribed strand. The
following 5'-phosphrylated oligonucleotide was annealed to the ssDNA: 5'-P-
GGCTACGTCCAGGAGCGCACCATCTTCTTC-3'. Primer extension was carried
out as described above, except the extension temperature was lowered to 610C
to increase yield. The resulting substrate is a heteroduplex in which the
transcribed strand has the mutant sequence (A) and the non-transcribed strand
has the wild type sequence (C) at position 289. Mismatch repair activity restores
the wild type sequence to the transcribed strand and leads to GFP expression. A
wild type homoduplex was prepared identically using wild type plasmid DNA as
the starting material, and the substrates were validated using the MT1 and TK6
cell lines (Fig. S3b).
256
Substrates containing a blunt-end double strand break
A unique Scal restriction site was inserted into the 5' untranslated region of the
pmax:BFP reporter plasmid, immediately 5' of the reporter gene (Fig. S3c).
Plasmids were linearized with Scal restriction enzyme, purified by
phenol/chloroform extraction and ethanol precipitation, and digest completeness
was confirmed by gel electrophoresis. The uncut pmax:BFP reporter with the
Scal restriction site was used as the undamaged control in experiments
measuring repair of the linearized reporter.
Substrates and methods for measuring homologous recombination
GFP-based HR reporter plasmids have been described previously (Kiziltepe et
al., 2005), and were a generous gift from Prof. Bevin Engelward. The D5G
plasmid expresses a non-fluorescent GFP reporter that is truncated at the 5' end
of the gene (Fig S4a). This plasmid was modified to include a Stul restriction site
so that a blunt-end double strand break could be introduced into the plasmid with
the goal of reducing the likelihood of unwanted re-ligation of the plasmid,
because this process does not yield fluorescent signal. Cells were transfected
with 0.5 micrograms of Stul-linearized D5G, 5 micrograms of D3G, plus 0.5
micrograms of an undamaged plasmid that was used as a control for transfection
efficiency. Homology directed repair of the DSB in the D5G reporter plasmid that
uses the D3G plasmid as a donor sequence leads to a full length GFP-encoding
gene, and results GFP expression.
Substrates containing a site-specific thymine dimer
A site-specific thymine dimer spanning positions 614-615 of the GFP sequence
was successfully introduced into the transcribed strand of the pmax GFP reporter
plasmid using previously described methods (Kitsera et al., 2011) (Fig. S5e).
257
Briefly, two nicking sites for the enzyme Nb.Bpul0 (Thermo Scientific) near the
3' end of the GFP reporter gene were used to excise a single stranded
oligonucleotide of 18 bp in length: 5'-TCAGGGCGGATTGGGTGC-3'. The
nicking sites flank a silent mutation that was introduced to generate a TpT
sequence in the transcribed strand of the plasmid. A synthetic oligonucleotide 5'-
TCAGGGCGGAT<>TGGGTGC-3' containing a thymine-thymine cis-syn
cyclobutane dimer indicated by T<>T (synthesized by TriLink BioTechnologies
using a cis-syn thymine dimer phosphoramidite (Glenn Research)) was annealed
and ligated into the gapped plasmid. Incorporation of the site-specific thymine
dimer in the plasmid was confirmed by endonucleolytic digestion with thymine
dimer specific glycosylase / AP lyase (T4 PDG, New England Biolabs). Greater
than 95% of the resulting product migrated as nicked plasmid DNA (Fig. S5f),
indicating at least 95% of the plasmids contained the lesion.
Isolation of total RNA for mRNAseq
At 18 hours, transfected cells were harvested by centrifugation, washed three
times with PBS, and resuspended in 1 mL Trizol reagent. The suspension was
extracted with 200 pL chloroform. The aqueous phase was removed, combined
with one volume of absolute ethanol, and applied to a Qiagen RNeasy mini-prep
spin column. The column was then washed two times with 500 pL buffer RPE
(Qiagen), and finally eluted in 40 pL diethylpyrocarbonate (DEPC) treated water.
From this point forward, RNA was handled in Eppendorf DNA LoBind tubes to
minimize loss of material. The quality of the RNA preparation was determined
using a bioanalyzer to confirm a RIN of at least 9.0. 1 pg of total RNA was stored
in TE Buffer at -80*C until submission for mRNAseq.
Isolation of mRNA and synthesis of cDNA
mRNA was isolated using a Qiagen Oligotex kit, using the manufacturer's
protocol, but substituting Eppendorf DNA LoBind tubes for those provided with
the kit. In the final step, mRNA was eluted in 20 uL buffer OEB preheated to 70
258
0C. 5 pL of the eluate was transferred to a LoBind tube, combined with 1 pL of
DNAse buffer and 1 unit of DNAsel (Invitrogen). The mixture was brought up to a
10 pL volume with DEPC treated water, and incubated for 15 minutes at room
temperature. DNAse was inactivated by addition of 1 pL of 25 mM EDTA,
followed by incubation at 65 0C for 10 minutes. A cocktail comprised of 2X RT
buffer (Qiagen), oligo-dT(12-18) (125 ng/uL; invitrogen), 4 units of RNAse
inhibitor (Qiagen), 5 mM dNTPs, and 4 units of reverse transcriptase (Omniscript;
Qiagen) was prepared, and 8 pL added to the DNAse digest. The reaction was
incubated for 1 hour at 37 *C. No-RT controls were performed identically, except
for the exclusion of the reverse transcriptase.
Specific amplification of reporter cDNA by PCR
cDNA samples were amplified with primers specific to the 3' and 5' UTR
regions of the pMax vector. The following primers were synthesized for specific
amplification of reporter cDNA:
5UTR: 5'- TTG CTA ACG CAG TCA GTG CT -3'
3UTR: 5'- GCA TTC TAG TTG TGG TTT GTC C -3'
1.5 pL of cDNA was PCR amplified in a 25 pL reaction volume with 1X PCR
buffer (Denville), 0.5 pM primers, 0.2 mM dNTPs, and 1 unit Taq polymerase
(Denville). Specific amplification was confirmed by gel electrophoresis and
analysis on a bioanalyzer chip. Water and EGFP encoded plasmids were used
as negative and positive controls, respectively. Finally, reactions were cleaned
up using a Qiagen PCR cleanup kit according to the manufacturer's protocol, and
eluted in 50 uL of TE.
Fragmentation of DNA
250 ng of PCR product was diluted to a total volume of 130 uL in TE
buffer. The DNA was fragmented in a Covaris microTUBE using a Covaris S2
sonicator (Duty Cycle 10%, Intensity 5, 200 cycles per burst, 180 seconds.
259
Fragmentation to a target base pair peak of 150 bp was checked using a Agilent
BioAnalyzer.
Validation of MGMT FM-HCR
Total RNA was isolated as described immediately above for mRNAseq
from the panel of cell lines for which data are presented in Fig. 4a. Poly-dT
oligonucleotides were used to generate cDNA, and MGMT was quantified by
TaqMan qPCR as described previously (Kitange et al., 2009). MGMT transcript
levels were quantitated using the DDCT method, and GAPDH was used as the
internal control. Transcript levels are reported in Fig. 4b relative to transcript
levels in the cell line TK6 +MGMT, which overexpresses MGMT and was used a
positive control.
Supplementary Note
In this manuscript we have presented a proof of concept for high
throughput DNA repair capacity assays. Because our experiments required only
a small fraction of the theoretical maximum throughput achievable by HCR-seq,
they were relatively expensive to perform. In what follows, we describe how the
cost of performing HCR-seq on a per sample basis decreases as more samples
are added to a given experiment.
Whereas FM-HCR allowed for the simultaneous detection of 5
independent repair reporters, the HCR-seq permitted the measurement of 20
reporters (5 reporter genes x 4 bar-codes) in a single experiment. The 20
RNAseq reporters were detected at sufficient coverage to obtain highly
reproducible dose response curves (Fig. 5). Because these transcripts
represented less than 1 % of the total mapped reads, it can be estimated that at
least 2000 reporters (or 200 dose-response curves) could be independently
assayed on a single lane if host transcripts were excluded from the assay.
The four dose response curves derived from sequencing data and
presented in Figure 5a were acquired at a cost of approximately $800 per curve.
260
However, several considerations would reduce the cost of sequencing-based
assays if deployed in large-scale population studies. As cost of sequencing
continues to fall, and particularly if a large number of samples is multiplexed on
single lane, sample preparation can be expected to dominate the cost of the
assay, with sequencing accounting for a small fraction of the overall cost. In the
present work bar-codes were introduced as part of the Illumina sample
preparation pipeline, however an equivalent means of distinguishing among
samples would be to introduce bar-codes into the library of reporter plasmids.
This configuration would permit sample pooling prior to sequencing library
preparation, leaving the cost of cell culture and transfection reagents as the
major contribution to the remaining cost of the assay. Such a workflow would
also essentially limit the active time in the research laboratory to that required to
handle and transfect the cells of interest. Furthermore, selective amplification of
reporter transcripts could be used to exclude host transcripts, which represented
the overwhelming majority of mapped sequences in the present work.
261
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