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Response to alkylation damage linked to meiotic progression

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Response to alkylation damage linked to meiotic progression

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RESPONSE TO ALKYLATION DAMAGE LINKED TO MEIOTIC
PROGRESSION

By: Tara Mastro

May 2015

________________________________________________________________________
A Dissertation Presented to the
FACULTY OF THE USC GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
(MOLECULAR BIOLOGY)

ii

Acknowledgements

First, I thank Susan L. Forsburg for being a fantastic mentor. Not only has she
taught me how to think, do, and talk science, but also, she has supported so many other
aspects of my professional development. Susan has supported me in exploring my own
ideas and paths through my projects while somehow still keeping me on track. She has
taught me to take things a step further and to be bold in with experiments. Outside of
her mentoring at the bench Susan has been exceptionally supportive of my personal
goals outside of lab and without this I surely could not have accomplished as much
professionally or personally in my time in her lab.
I thank my committee for the guidance and support through not only my thesis
work but also my oral examinations. Thank you to Oscar Aparicio for helpful feedback
and the use of his lab equipment such as the Facscan. I am grateful to Michelle
Arbeitmen and Xuelin Wu for their helpful advise. I thank Steven Finkel for reminding
me to take breaks from lab work to put it all in perspective. I also thank Steven Finkel
for being a huge supporter of the Molecular and Computational Biology Graduate
Student Association. His mentoring of this organization made it possible to have many
great development and social events for students.
All the members of the Forsburg lab have been great to work with and learn from.
I am grateful to Ji-Ping Yuan for his excellent lab management. Ji-Ping has also very
patiently taught me many of the techniques I have used including pulse field gel, tetrad
dissection, frogging, and much more. I appreciate, Marc Green and all his help with
imaging. Sarah Sabatinos has been an invaluable resource of knowledge about all

iii

molecular techniques. Sarah was also a great mentor and collaborator in the work we
did together on the toxicity of base analogs in chapter 5. I thank Poa-Chen Li and Anh-
Huy Le for leaving me with detailed notes on what they have done in the past allowing
me to apply it to my thesis work. I also thank Anh-Huy for his collaboration in the work
with Dfp1 meiosis (Chapter 2). Nimna Ranatunga has be a great friend to me in and out
of the lab. Nimna has always been willing to provide me with an extra set of hands
when I need them. I thank Wilber Escorcia for helping make many of the meiosis
movies in chapter 4. Wilber has also read and provided feedback on many versions of
the manuscripts I have worked on. Yoona Jung is an undergraduate in the lab that has
been working with me for the last several years. She has been a joy to work with and
has been an invaluable set of skilled hands in the lab. I thank for Bijan Givechian, who
is also an undergraduate, been working with me on the project in chapter 4 as well.
Thank you to past and present lab members that have helped in various ways and were
great to work with; Ruben Petreaca, Rebecca Nugent, Lin Ding, Amanda Jensen, and
Kuo-Fang Shen.
Thank you to WiSE for all the support for my growing family. WiSE has provided
me with funding for parental leave and childcare that has made managing a career in
science and having a family possible.
Thank you to all my MCB classmates. Specifically Anna Skylar, Asen Erdem,
and Nimna Ranatunga for being fantastic friends and great resources for scientific
discussion.
I thank my husband, Eamon, for his support. I know it was not always easy to be
supportive of midnight journeys to the lab or 24-hour time courses, and yet, he has. I

iv

thank him for trying to participate in conversations about experimental approach during
meals even though I was mostly talking to myself. Most of all I thank him for his comic
relief during setbacks and continuous positive attitude.

v

Table of Contents
Acknowledgements ii
List of Abbreviations vii
List of Tables ix
List of Figures xi
List of Movies xiii
Abstract xv

Chapter 1: Introduction

1
1.1 Introduction 1
1.2 Events during meiosis 3
1.2.1 Meiotic cohesion 4
1.2.2 Programmed meiotic double strand break repair 6
1.3 DNA Damage Response 10
1.3.1 Nucleotide excision repair 12
1.3.2 Post replication repair 14
1.3.3 DNA damage checkpoints 15
1.3.4 Structure specific endonucleases and genome stability 17
1.4 Conclusion 18
1.5 Chapter 1 Bibliography 20

Chapter 2: The C terminus of S. pombe DDK subunit Dfp1 is required for
meiosis-specific transcription and cohesin cleavage
34
2.1 Overview 35
2.2 Introduction 35
2.3 Results 38
2.4 Discussion 57
2.5 Materials and Methods 63
2.6 Legends for Movies 73
2.7 Chapter 2 Bibliography 74

Chapter 3: Increased Meiotic Crossovers and Reduced Genome Stability in
Absence of Schizosaccharomyces pombe Rad16 (XPF)
85
3.1 Overview 86
3.2 Introduction 86
3.3 Results 90
3.4 Discussion 122
3.5 Materials and Methods 133
3.6 Legends for Movies 140
3.7 Chapter 3 Bibliography 142

vi

Chapter 4: Translesion synthesis polymerases contribute to meiotic
chromosome segregation in S. pombe

160
4.1 Overview 161
4.2 Introduction 161
4.3 Results 164
4.4 Discussion 177
4.5 Materials and Methods 181
4.6 Legends for Movies 188
4.7 Chapter 4 Bibliography 190

Chapter 5: A mammalian-like DNA damage response of fission yeast to
nucleoside analogs
198
5.1 Overview 199
5.2 Introduction 199
5.3 Results 202
5.4 Discussion 221
5.5 Materials and Methods 236
5.6 Legends for Movies 246
5.7 Chapter 2 Bibliography 247

Comprehensive Bibliography 257

vii

List of Abbreviations

BER Base excision repair
BrdU 5-bromo-2-deoxyuridine
ChIP Chromatin immunoprecipitation
CO crossover
CPT camptothecin
DDR DNA damage response
dHJ Double Holliday junction
DMSO Dimethyl sulfoxide
DSB Double strand break
EdU 5-Ethynyl-2´-deoxyuridine
EMM Edinburgh Minimal Media
FA Fanconi anemia
FACS Fluorescent assisted cell sorting
HR Homologous recombination
HU Hydroxyurea
I Incorporator
ICL Interstrand crosslink
ICL Inter-strand crosslink
inc Incorporator
IR Ionizing radiation
MI Meiosis I
MII Meiosis II
MMR Mismatch repair
MMS methyl methanesulfonate
NCO Non-crossover
NER Nucleotide excision repair
NHEJ Non homologous end joining
NI Non- incorporator
PFGE Pulse field gel electrophoresis

viii

pmDSBs Programmed meiotic double strand breaks
PMG Pombe Glutamate medium
prDSB Programmed double strand breaks
PRR Post replication repair
qRT-PCR Quantitative reverse transcription polymerase chain reaction
S-phase Synthesis phase
SC Synaptonemal complex
SCE Sister chromatid exchange
sHJ Single Holliday junction
SSA Single strand annealing
TBZ Thiabendazole
Thy thymidine
TLSP Translesion synthesis polymerase
UV Ultraviolet radiation
XP Xeroderma pigmentosum
YES Yeast extract with supplements

ix

List of Tables

Table 1.1 Comparison of DNA damage repair pathways 11
Table 2.1 Spore Viability 40
Table 2.2 Summary of classes from Figure 2.5 51
Table 2.3 Chromosome Segregation monitored with LacI-GFP. 52
Table 2.4 Strains used in this study. 69
Table 2.5 Oligos used for RT-PCR. 72
Table 3.1 Tetrad analysis of recombination between His4 and Lys4. 95
Table 3.2 Recombination and spore viability between his4-239 and lys4-95,
and ade6.
106
Table 3.3 Recombination and spore viability of ade6 heteroallele. 108
Table 3.4 Genetic interactions: phenotype of double mutants with rad16-249 112
Table 3.5 Distribution of cell length measurements binned. 118
Table 3.6 Analysis of H3-mRFP Taz1-GFP mitotic live cell movies 119
Table 3.7 Analysis of Rad52 and RPA. 122
Table 3.8 Mitotic recombination events in heteroallele spore germination. 128
Table 3.9 Strains used in this study. 133
Table 4.1 Taz1-GFP signal for quad∆ H3-mRFP mis-segregating bodies. 172
Table 4.2 Strains used in this study. 182
Table 5.1 Mutation rates for spontaneous and BrdU-induced canavanine
forward mutation analysis
228

x

Table 5.2 Frequency of hsv-tk+ loss or sectoring in incorporating wild-type
and spd1Δ cultures, with or without BrdU treatment
236
Table 5.3 Fission yeast strains used in this study. 243

xi

List of Figures

Figure 1.1 Meiotic vs. mitotic chromosome segregation 2
Figure 1.2 Rec8 cleavage during meiosis 5
Figure 1.3 Meiotic Double Strand Break Repair and Recombination
Pathways
10
Figure 1.4 Pathways for nucleotide excision repair 14
Figure 1.5 Check points in S. pombe compared to other organisms. 16
Figure 2.1 Terminal Meiotic Phenotype of dfp1-r35 Mutants. 39
Figure 2.2 Synchronous Meiosis in mat2-102/h- pat1-114/pat1-114 diploids
and pat1-114 haploids.
42
Figure 2.3 Transcriptional Expression of Meiotic Genes. 44
Figure 2.4 prDSBs in dfp1-r35 Mutants 47
Figure 2.5 Chromosome Segregation in dfp1-r35 Mutants compared to
rec8∆ and rec12∆.
50
Figure 2.6 Rec8-GFP Stability. 55
Figure 3.1 Long Term Viability in Presence of Drug. 91
Figure 3.2 Spore Viability and Chromosome Segregation. 92
Figure 3.3 Meiotic DSBs in Diploids. 100
Figure 3.4 Timing of Synchronous Meiotic Events. 103
Figure 3.5 Recombination of Intergenic and Intragenic Intervals and spore
viability.
105
Figure 3.6 Check Point Activation and DNA Damaging Drug Sensitivity. 114

xii

Figure 3.7 Growth Rates and Drug Sensitivity for rad16-249 Double Mutants. 116
Figure 3.8 Western blot of Chk1-HA using 16B12 anti-HA antibody. 120
Figure 3.9 Visualization of DNA Damage via Rad52 and RPA foci. 121
Figure 4.1 Spore and Mitotic Viability. 165
Figure 4.2 Recombination and Synchronous Meiosis. 168
Figure 4.3 Meiotic chromosome segregation. 170
Figure 4.4 Rec8 dynamics. 174
Figure 4.5 Rad21-GFP in meiosis. 177
Figure 5.1 BrdU and EdU doses affect signal, viability, and cell division. 205
Figure 5.2 Media formulation alters BrdU and EdU sensitivity. 207
Figure 5.3 BrdU and EdU dose affects cell viability 209
Figure 5.4 BrdU and EdU cause prolonged DNA synthesis, cell-cycle
slowing, and DNA damage.
212
Figure 5.5 BrdU and EdU cause prolonged DNA synthesis, cell cycle slowing
and DNA damage.
214
Figure 5.6 BrdU exposure triggers the DNA damage response. 217
Figure 5.7 BrdU and EdU induce a DNA damage response. 221
Figure 5.8 BrdU pretreatment changes sensitivity to DNA damaging drugs. 224
Figure 5.9 BrdU pre-treatment changes sensitivity to mutagens. 226
Figure 5.10 Spd1 protects cells from division and mutation during dNTP
imbalance.
230
Figure 5.11 Spd1 protects cells from division and mutation during dNTP
imbalance
233

xiii

List of Movies

Movie 2.1 Live cell imaging of WT meiosis LacI-GFP
Movie 2.2 Live cell imaging of dfp1-r35 meiosis LacI-GFP
Movie 2.3 Live cell imaging of rec12∆ meiosis LacI-GFP
Movie 2.4 Live cell imaging of rec8∆ meiosis LacI-GFP
Movie 2.5 Live cell imaging of dfp1-r35 rec12∆ meiosis LacI-GFP
Movie 2.6 Live cell imaging of dfp1-r35 rec8∆ meiosis LacI-GFP
Movie 2.7 Live cell imaging of WT Rec8-gfp meiosis
Movie 2.8 Live cell imaging of dfp1-r35 Rec8-gfp meiosis
Movie 2.9 Live cell imaging of rec12∆ Rec8-gfp meiosis
Movie 3.1 Live cell imaging of WT meiosis.
Movie 3.2 Live cell imaging of rad16-249 meiosis.
Movie 3.3 Live cell imaging of rhp14∆ meiosis.
Movie 3.4 Live cell imaging of rad13∆ meiosis.
Movie 3.5 Live cell imaging of mus81∆ meiosis.
Movie 3.6 Live cell imaging of mus81∆ meiosis.
Movie 3.7 Live cell imaging of rec12Δ meiosis.
Movie 3.8 Live cell imaging of rad16-249 mitosis.
Movie 4.1 Live cell imaging of WT LacI-GFP, H3-mRFP meiosis
Movie 4.2 Live cell imaging of quad∆ LacI-GFP, H3-mRFP meiosis
Movie 4.3 Live cell imaging of WT Rec8-GFP meiosis
Movie 4.4 Live cell imaging of quad∆ Rec8-GFP meiosis

xiv

Movie 4.5 Live cell imaging of WT Rad21-GFP, H3-mRFP meiosis
Movie 4.6 Live cell imaging of quad∆ Rad21-GFP, H3-mRFP meiosis
Movie 5.1 WT hsv-tk+ hENT+ Rad52 foci BrdU with treatment
Movie 5.2 cds1∆ hsv-tk+ hENT+ Rad52 foci BrdU with treatment
Movie 5.3 WT hsv-tk+ hENT+ Rad52 foci BrdU with treatment
Movie 5.4 cds1∆ hsv-tk+ hENT+ Rad52 foci BrdU with treatment

xv

Abstract:
Mechanisms that maintain genome stability are essential for human health. Loss
of genome stability is associated with cancer and birth defects. This dissertation uses a
model fission yeast system to investigate how cells preserve chromosome integrity
during the specialized differentiation process of meiosis. In this work mutants sensitive
to alkylation damage were examined for their ability to proceed through meiosis. It was
found that these genes contribute to meiotic progression. However, their contribution is
not isolated to a single process or mechanism. Rather it was seen that these mutants
that are sensitive to meiotic alkylation damage have a diverse role in meiotic
progression.

1

Chapter 1: Introduction
1.1 Introduction
This dissertation project is concerned with mechanisms that maintain
genome stability, particularly during meiosis. The work I shall describe touches
not only on the mechanisms of meiosis, but also mechanisms of DNA repair that
contribute to the integrity of the genome during this differentiation process.
Meiosis is common to sexually reproducing eukaryotes. During meiosis,
diploid progenitor cells give rise to four genetically distinct haploid gametes.
Meiosis differs from mitosis in that it has one replication event followed by two
cell divisions. The first meiotic division is reductional (homologous chromosomes
separate). The second division is equational (sister chromatids separate) and
produces haploid gametes (Figure 1). In order ensure faithful transmission of
genetic material from one generation to another though sexual reproduction,
successful meiotic chromosome segregation must occur. Sister chromatids and
homologous chromosomes must align and then separate in a well-orchestrated
manner. Both cohesion and programmed meiotic double strand breaks
(pmDSBs) are essential to this process. When either of these processes is
compromised, meiotic errors ensue which often results in aneuploidy. In humans
most aneuploidies are lethal. However, in some cases aneuploidy may be
tolerated, but with the consequence of developmental defects such as in the
case of trisomy 21 (Down syndrome).

2

Figure 1.1: Mitotic (a) vs meiotic (b) chromosome segregation. Nature
Reviews Molecular Cell Biology 5, 984 (2004).
Humans are shockingly bad at meiosis compared to other metazoans.
Saccharomyces cerevisiae meiotic chromosome mis-segregtion is 1 in 10,000
(SEARS et al. 1992). In Drosophilia melanogaster mis-segregations are estimated
at a rate of 1 in 1,700 (KOEHLER et al. 1996). In mammals meiotic errors are
inferred from observing aneuploidies in fertilized eggs. In mice, aneuploidy in
fertilized eggs from young females is between 3-4% (MERRIMAN et al. 2012).
However in humans, nearly 10-30% of all fertilized eggs are thought to have
aneuploidies (HASSOLD AND HUNT 2001). Interestingly, among young woman (age
<35) there is a higher rate of aneuploidy if a previous aneuploidy has occurred

3

that cannot be explained due to germline mosaicism (MUNNE et al. 2004). This
suggests a potential genetic predisposition toward aneuploidy.
My thesis work has been focused on determining genetic risk factors for
meiotic failures. To achieve this I have employed a model system, the fission
yeast, Schizosacchromyces pombe. Meiosis is a well-conserved process from
yeast to humans. Many of the genes and activities required for meiosis in
humans are conserved in fission yeast. S. pombe provides excellent genetic and
molecular tools. Cells can be induced to enter meiosis synchronously, and in
large numbers. This allows live cell analysis throughout the process. This makes
fission yeast a perfect system to study factors important for proper meiosis and
their mechanism of action.
1.2. Events during meiosis
In order to promote proper segregation of the chromosomes during
meiosis, homologous chromosomes need to be linked, so that they can
segregate from one another properly during the MI division. In contrast, sister
chromatids remain associated during MI. This requires modification of several of
the mechanisms associated with chromosome segregation during mitosis. First
the cohesion system that links sister chromatids must be adapted so the sisters
remain associated during the MI division. There must be a monopolar spindle
attachment to each homolog. Finally, physical linkage between the homologs to
promote proper pairing and segregation is required, initiated through pmDSBs
and homologous recombination.

4

1.2.1 Meiotic cohesion
Sister chromatid cohesion in meiosis is required to promote segregation of
homologous chromosomes in the MI reductional division, and is degraded to
allow the segregation of sister chromatids in the MII equational division. In
mitosis the cohesin complex is loaded before replication but is not converted to
the active complex which links sister chromatids until S-phase (ISHIGURO AND
WATANABE 2007). The cohesion complex forms a ring structure consisting of four
subunits, Psm1, Psm3, Rad21 and Psc3 (LOSADA AND HIRANO 2005; NASMYTH
AND HAERING 2005). In meiosis the Kleisin subunit Rad21 is mostly replaced by
the meiosis specific Rec8. The Psc3-like Rec11 protein forms a complex with
Psm1, Psm3, and Rec8 at arm regions while Psc3 is maintained in centromeric
regions (WATANABE 2004; NASMYTH AND HAERING 2005).
As in mitosis, cohesin cleavage occurs allowing for segregation. In
meiosis, the separase-mediated cleavage of Rec8 occurs only on the arm
regions in MI while centromeric Rec8 remains protected from cleavage by the
conserved Shugoshin (Sgo1) (LEE et al. 2005; NASMYTH AND HAERING 2005;
WATANABE 2005; WATANABE AND KITAJIMA 2005). Sgo1 recruits PP2A
phosphatase to centromeres. The interaction of Sgo1 and PP2A in meiosis is
required for Rec8 centromeric protection (KITAJIMA et al. 2006; RIEDEL et al.
2006). Rec8 phosphorylation is necessary and sufficient for separase cleavage
(KATIS et al. 2010). Several kinases are required for phosphorylation of Rec8

5

driving cleavage, including CK1 (ISHIGURO et al. 2010; RUMPF et al. 2010) and
Dbf4-dependent Cdc7 kinase (DDK) (KATIS et al. 2010; LE et al. 2013). Cleavage
of the centromeric Rec8 allows MII to proceed with sister chromatid segregation.
Thus, Rec8 stepwise cleavage in different regions ensures proper segregation
during MI and MII (Figure 2).

Figure 1.2: Rec8 cleavage duing meiosis. Graphical abstract (KATIS et al.
2010) of mechanism of Rec8 cleavage during meiosis.

6

1.2.2 Programmed meiotic double strand breaks
In meiosis unlike in proliferative vegetative growth, DNA damage in the
form of pmDSBs is deliberately induced. The meiosis specific endonuclease
Rec12 (Spo11) induces pmDSBs (LIN AND SMITH 1994; BERGERAT et al. 1997;
KEENEY et al. 1997). These pmDSBs act as substrates for recombination
between homologous chromosomes and thus the production of genetic variation
among the haploid progeny. Further, in most eukaryotes, the formation and
recombinational repair of the induced DSBs is required for proper homolog
pairing and thus chromosome segregation.
pmDSBs directly follow and are linked to pre-meiotic DNA replication. It is
unclear whether replication itself if required for pmDSB formation or merely the
presence of certain replication proteins. In budding yeast, pmDSBs do not occur
when replication is inhibited through the use of Hydroxyurea (HU; causes
nucleotide starvation) or inhibition of cyclins (proteins that drive cell cycle and
meiotic progression). However, in both budding and fission yeast, when origins
are suppressed using pre-replication complex mutants, pmDSBs do form (BORDE
et al. 2000; MURAKAMI AND NURSE 2001; SMITH et al. 2001). In both yeasts, DDK
(S.p. Hsk1) is required for Rec12/Spo11 recruitment and thus pmDSB formation
(OGINO et al. 2006; WAN et al. 2006; SASANUMA et al. 2008; WAN et al. 2008).
One interesting observation, which is conserved across several model
systems, is that there are several more breaks made across the genome than
there are observable crossovers (CO) (MOENS AND PRINCE 2002; PHADNIS et al.

7

2011; SERRENTINO AND BORDE 2012). When the number of pmDSB is reduced;
however, the number of CO remain constant at the expense of non-crossover
(NCO) resolution (MARTINI et al. 2006). This phenomenon is termed crossover
homeostasis (MARTINI et al. 2006; CHEN et al. 2008; KAN et al. 2011). Thus far the
molecular mechanism of crossover homeostasis is unknown. Another example of
CO control which may contribute to CO homeostasis, CO invariance, is
prominent in S. pombe (HYPPA AND SMITH 2010). Here recombination is not
through the homologous chromosome, but through the sister chromatid, which is
undetectable due to sequences being identical (HYPPA AND SMITH 2010; PHADNIS
et al. 2011). Specifically CO invariance allows for pmDSB hotspots to exist
across the chromosome but maintain a regular CO frequency overall.
Paradoxically, the DSB hotspots have a higher proportion of repair being done
through the sister (YOUNG et al. 2002; CROMIE et al. 2007; HYPPA AND SMITH
2010), while the colder regions of the genome have an increased rate through
the homolog. Thus, though the number of pmDSBs in a region of a chromosome
may be high, the actual interval between CO is actually invariant. This balance
of recombination between the sister and the homolog may contribute to overall
CO homeostasis as any sister event would be invisible and thus termed as a
NCO (MARTINI et al. 2006). CO homeostasis ensures that the number and
location of CO are distributed ensure proper homolog segregation (MARTINI et al.
2006).
Mechanisms to inhibit excessive CO formation also exist. For example, in
some organisms CO homeostasis is regulated by the synaptonemal complex

8

(SC) and CO interference (SHINOHARA et al. 2008). The synaptonemal complex
forms a connection between paired homologs during meiosis, which may
constrain the number of CO in that region (PAGE AND HAWLEY 2004). Not all CO
are subject to CO interference. In budding yeast there are genetically distinct
classes of COs. ZMM proteins define a group of diverse genes required for
synapsis. The absence of any one of these proteins will inhibit ZMM dependent
COs, while NCOs remain normal (LYNN et al. 2007; SHINOHARA et al. 2008).
ZMM dependent COs are subject to CO interference. In contrast, COs that are
MUS81 dependent escape CO interference. Mus81 is an endonuclease that acts
as a Holiday Junction (HJ) resolvase and is genetically required for a subset of
CO formation in budding yeast and worms (HOLLINGSWORTH AND BRILL 2004;
YOUDS et al. 2010). Fission yeast lacks a full SC but maintains rudimentary
structure called linear elements, which are related to the axial elements of the SC
(LORENZ et al. 2004; LOIDL 2006). Interestingly, there is no CO interference and
the majority of CO are Mus81 dependent (BODDY et al. 2001; MARTINEZ-PEREZ
AND COLAIACOVO 2009).
The pmDSBs are repaired via recombination (Figure 1.3). Once a DSB is
made by Rec12/Spo11, the 3’ end undergoes resection by the MRN complex,
CtIP, Exo1, and Dna2 (SYMINGTON AND GAUTIER 2011; YOUDS AND BOULTON
2011). Rad52 displaces RPA from the 3’ end promoting the loading of the RecA
homolog, Rad51 and inhibiting the loading of the meiosis specific RecA homolog,
Dmc1 (MURAYAMA et al. 2013). Dmc1 loading is instead promoted by Swi5-Sfr1
(MURAYAMA et al. 2013). Rad51 forms a complex with Rad54 to promote

9

invasion to the homologous sequences (YOUDS AND BOULTON 2011).
Rdh54/Tid1, a Rad54 homolog, has overlapping functions with Rad54, but
primarily promotes Dmc1 assembly (CATLETT AND FORSBURG 2003; NIMONKAR et
al. 2012). Rad54 and Rad51 appear to promote primarily intersister
recombination while Dmc1 and Rdh54 are thought to promote interhomolog
recombination (CATLETT AND FORSBURG 2003; HYPPA AND SMITH 2010; YOUDS AND
BOULTON 2011; NIMONKAR et al. 2012). In S. pombe the Rad51 mediator
complexes Rad55 and Rad57 appear to work in both pathways (HYPPA AND
SMITH 2010). A pmDSB may be repaired by either a double (dHJ) or single
holiday junction (sHJ). A sHJ occurs when resolution occurs before second end
recapture has occurred. Studies have shown predominantly dHJs in budding
yeast (BELL AND BYERS 1983; SCHWACHA AND KLECKNER 1995), while fission yeast
primarily has sHJ (CROMIE et al. 2006). The current model suggests that a NCO
can arise via specific cleavage of either a dHJ or sHJ, or by SDSA, while a CO
can form only through specific cleavage of either a dHJ or sHJ (PHADNIS et al.
2011; YOUDS AND BOULTON 2011).

10

Figure 1.3: Meiotic double strand break repair and recombination
pathways. Adapted from (PHADNIS et al. 2011)

1.3 DNA damage repair
DNA damage may occur in many ways, some of which include normal cell
metabolism, DNA replication, exogenous insults, such as, drugs or ultraviolet
radiation (UV). The cell must respond to the damage by restraining the cell cycle,
and activating repair pathways. This initial response works via the monitoring
system known as checkpoints, which activates distinct signal transduction
pathways to prevent cell division and promote repair.

11

There are classically six major pathways of DNA damage repair (Table
1.1): base excision repair (BER), nucleotide excision repair (NER), mismatch
repair (MMR), post replication repair (PRR) and double strand break repair
through either nonhomologous end joining (NHEJ) or homologous recombination
(HR) (FRIEDBERG 2005; CICCIA AND ELLEDGE 2010).
Table 1.1: Comparison of DNA damage repair pathways. Adapted from
(FRIEDBERG 2005; BRANZEI AND FOIANI 2008).
BER NER MMR PRR HR NHEJ
Damaging
agent
X-rays,
alkylation,
oxygen
radicals
UV, oxygen
radicals
Replication
errors
X-rays,
alkylation,
oxygen
radicals,
UV
X-rays, UV,
replication
fork collapse
X-rays, UV
Lesions Abasic site,
8-
oxoguanine
, single
strand
breaks,
uracil
Bulky adducts,
6-4
photoproduct,
cyclobutane
pyrimidine
dimer
A-G and C-T
mismatch,
insertions,
deletions
Various
leasions
found in
BER and
NER
DSB, inter-
strand
corsslink
DSB, inter-
strand
corsslink
Cell-cycle G1 G1 S S G2-M, S G1

MMR and BER are employed under conditions of minimal DNA damage
distortions, while NER and HR are required to rectify larger DNA lesions such as
those caused by alkylation damage (MEMISOGLU AND SAMSON 2000; ALSETH et al.
2005; KANAMITSU AND IKEDA 2011). PRR is a damage tolerance pathway that
allows bypass of the lesion and subsequent repair by fork regression,
recombination, or other repair pathways (FRIEDBERG 2005; WICKLIFFE et al. 2006;

12

DOLAN et al. 2010. In S. pombe aside from the major contribution of NER and
HR, BER acts synergistically with these two repair systems to repair alkylation
damage (MEMISOGLU AND SAMSON 2000; OSMAN et al. 2003; RIBAR et al. 2004;
ALSETH et al. 2005; SUGIMOTO et al. 2005; KANAMITSU AND IKEDA 2011). Alkylation
damage can be induced by specific drugs such as methyl methanesulfonate
(MMS) or can commonly be due to exposure to environmental hazards such as
tobacco smoke (CICCIA AND ELLEDGE 2010).

1.3.1 Nucleotide Excision Repair
There are two major branches for NER. These branches use many of the
same downstream core factors for repair however, the mechanism by which
lesions are sensed differ (Figure 1.4). In transcription coupled NER the stalling of
RNA polymerase II serves as a damage-sensing signal whereas in global
genome NER damage is recognized due to DNA being thermodynamically
unstable (DE LAAT et al. 1999; FAGBEMI et al. 2011). Regardless of how the
damage is recognized, once it is, helicases are recruited to allow access by other
NER repair factors (DE LAAT et al. 1999; FAGBEMI et al. 2011). Once DNA is
open, XPA, RPA, and XPG are recruited (Figure 1.4). XPA serves to recruit the
5’ endonuclease XPF, while the 3’ endonuclease XPG is recruited separately
(VOLKER et al. 2001; TSODIKOV et al. 2007; ORELLI et al. 2010). XPG has both
catalytic and structural roles in NER though the structural roles are not entirely
defined (FAGBEMI et al. 2011). XPF is the last factor to be recruited to the site of

13

damage and dual incision of the bulky lesion at the 5’ and 3’ end is mediated by
the presence of XPF (FAGBEMI et al. 2011). It is unclear whether the dual incision
is step-wise or simultaneous, but current data support a model in which XPF cuts
prior to XPG (WAKASUGI et al. 1997; CONSTANTINOU et al. 1999; TAPIAS et al.
2004; STARESINCIC et al. 2009).
Gap filling synthesis proceeds by an unknown mechanism. It is however
known that DNA polymerases are required: the replicative polymerases pol δ and
pol ε, and the error prone pol κ are required for synthesis during NER (OGI AND
LEHMANN 2006). The replicative clamp, PCNA, and its ubiquitination by Rad18
are also required (OGI et al. 2010; FAGBEMI et al. 2011). These genetic
requirements are reminiscent of post replication repair and polymerase switching
leaving the formal possibility that this may be the mechanism by which NER gap
filling synthesis proceeds.

14

Figure 1.4: Pathways for nucleotide excision repair. Adapted from (DE LAAT et
al. 1999)
1.3.2 Post replication repair
Completion of DNA replication is a prerequisite for successful cell division.
In the case that the replication fork is blocked, there must exist a mechanism to

15

allow replication to continue. The mechanisms by which a replication fork may
proceed past fork stalling damage is term post replication repair or DNA damage
tolerance. There are largely two pathways for replication blocks to be resolved.
The first is the error-prone mechanism, which recruits lower fidelity translesion
synthesis DNA polymerases and the second is through an error free fork
regression or recombinational repair (FRIEDBERG 2005; WICKLIFFE et al. 2006;
DOLAN et al. 2010). At the crux of this pathway is PCNA. The ubiquitination state
of PCNA mediates pathway choice (FRIEDBERG 2005; CICCIA AND ELLEDGE 2010;
DOLAN et al. 2010). Mono-ubiquitination at K164 of PCNA preformed by Rhp18
will shunt repair toward the error prone (FRIEDBERG 2005; CICCIA AND ELLEDGE
2010; DOLAN et al. 2010). However, if the poly-ubiquitination occurs via Ubc13,
error free repair will occur through a fork regression mechanism (FRIEDBERG
2005; CICCIA AND ELLEDGE 2010; DOLAN et al. 2010). It is also possible to restart
the fork through homologous recombination (Figure 1.5) (FRIEDBERG 2005; CICCIA
AND ELLEDGE 2010; DOLAN et al. 2010).
1.3.3 DNA damage checkpoints
DNA damage checkpoints are part of a signal transduction that monitors
the condition of the DNA to ensure that division only occurs when the genome is
stable. In S. pombe, the damage checkpoint kinase Chk1 controls the G2/M
transition and is activated via phosphorylation by the upstream sensor Rad3
(ATR homolog) in response to DSBs and single stranded DNA (WALWORTH et al.
1993; AL-KHODAIRY et al. 1994; WALWORTH AND BERNARDS 1996). Chk1
phosphorylation can occur when cells are challenged with genome insults such

16

as UV, ionizing radiation (IR), MMS, Campothecin (CPT), and Cisplatin (WAN AND
WALWORTH 2001; LATIF et al. 2004; PAPARATTO et al. 2009). Cds1, the
Rad53/Chk2 homolog, is the core component of the replication checkpoint that is
activated by Rad3 in times of replication stress such as when nucleotide pools
are depleted by HU or single stranded DNA present at replication forks (LINDSAY
et al. 1998; SABATINOS AND FORSBURG 2013).

Figure 1.5: Check points in S. pombe compared to other organisms.
Adapted from (SABATINOS AND FORSBURG 2013)

17

1.3.4 Structure specific endonucleases and genomic stability
Structure-specific endonucleases cleave specific DNA structures that are
generated during the response to DNA damage or during repair. These
endonucleases show specificity for different and overlapping DNA templates.
There are many structure specific endonucleases in eukaryotes implicated in
diverse processes including different repair pathways and processing of
recombination intermediates (SCHWARTZ AND HEYER 2011).
Mus81-Eme1 has been shown to cleave D-loops, nicked Holliday
junctions, 3′ flaps, and Y- DNA in vitro (BODDY et al. 2001; CHEN et al. 2001;
CICCIA et al. 2008). Gen1/Yen1 is a monomeric XPG family endonuclease having
5’ flap activity with the unique feature in this family to also cleave Holliday
junctions (HARRINGTON AND LIEBER 1994; LIEBER 1997; HOSFIELD et al. 1998;
JOHNSON et al. 1998). Gen1
Hs
and its budding yeast orthologue Yen1
Sc
overlap
with Mus81 (BLANCO et al. 2010; HO et al. 2010; TAY AND WU 2010). Though S.
pombe lacks an obvious Gen1/Yen1 homolog, expression of Gen1 will rescue
chromosome segregation defects in the absence of Mus81 (LORENZ et al. 2010).
XPF–ERCC1 also cleaves 3′ flaps and is the 5’ endonuclaese in NER (CICCIA et
al. 2008; FEKAIRI et al. 2009; CICCIA AND ELLEDGE 2010). SLX4–SLX1 can cleave
HJs in vitro and localizes to regions of replication stress such as telomeres in
vivo (FEKAIRI et al. 2009; SVENDSEN et al. 2009; KASHIYAMA et al. 2013; WAN et al.
2013; WILSON et al. 2013). Further, SLX4 has a scaffolding function that binds to
the three endonucleases, MUS81–EME1, XPF–ERCC1, and SLX1 (MUNOZ et al.
2009). These endonucleases are termed the swiss army knife of the cell in that

18

all three contribute to genomic stability and form a macro complex for some
repairs such as interstrand crosslinks (ICLs) in mammalian systems (HANADA et
al. 2006; FEKAIRI et al. 2009; MUNOZ et al. 2009; SVENDSEN et al. 2009)

1.4 Conclusion
In the following chapters, I will describe my work on the meiotic effects of
a number of mutants first characterized for their role in the response to alkylation
damage. This project initiated in previous work in the lab that identified an MMS-
sensitive mutant allele of the DDK kinase subunit Dfp1 (dfp1-r35) as part of a
larger screen for MMS-sensitive mutations (DOLAN et al. 2010). In a collaboration
with a former student in the lab, Anthony Le, we showed that this allele of dfp1
has profound defects at multiple stages in meiosis (Chapter 2, (LE et al. 2013)).
In Chapter 3, I examined another strain from the same screen that identified
dfp1-r35; this mutant lacks the function of the structure specific endonuclease
Rad16/XPF. I found a novel meiotic phenotype that I term “fragmentation” and
showed that Rad16 plays a critical role in genome stability both in mitosis and
meiosis, with evidence for overlap with Mus81 (Chapter 3, (MASTRO AND
FORSBURG 2014)). In Chapter 4, I describe recent work investigating meiotic
contributions of another group of MMS-sensitive mutations affecting the error-
prone trans-lesion synthesis polymerases. Finally, in chapter 5, I describe a
collaboration with postdoc Sarah Sabatinos examining the effects associated
with treatment with nucleoside analogues BrdU and EdU. Together, this body of

19

work shows that genes, which are important for alkylation damage in mitosis do
have substantial contributions to meiotic progression. One of the studies find a
contribution of Rad16/XPF to DSB repair, while two of these studies show a role
for alkylation damage genes in meiotic cohesion (Chapter 2 and Chapter 4). This
implies that despite a diversity of mechanisms that may link alkylation damage
tolerance and meiotic progression, some may be more prominent than others.

.

20

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33

Wickliffe, J. K., L. A. Galbert, M. M. Ammenheuser, S. M. Herring, J. Xie et al.,
2006 3,4-Epoxy-1-butene, a reactive metabolite of 1,3-butadiene, induces
somatic mutations in Xpc-null mice. Environ Mol Mutagen 47: 67-70.
Wilson, J. S., A. M. Tejera, D. Castor, R. Toth, M. A. Blasco et al., 2013
Localization-dependent and -independent roles of SLX4 in regulating
telomeres. Cell Rep 4: 853-860.
Youds, J. L., and S. J. Boulton, 2011 The choice in meiosis - defining the factors
that influence crossover or non-crossover formation. J Cell Sci 124: 501-
513.
Youds, J. L., D. G. Mets, M. J. McIlwraith, J. S. Martin, J. D. Ward et al., 2010
RTEL-1 enforces meiotic crossover interference and homeostasis.
Science 327: 1254-1258.
Young, J. A., R. W. Schreckhise, W. W. Steiner and G. R. Smith, 2002 Meiotic
recombination remote from prominent DNA break sites in S. pombe. Mol
Cell 9: 253-263.

34

Chapter 2

The C terminus of S. pombe DDK subunit Dfp1 is required for meiosis-
specific transcription and cohesin cleavage

The work in this chapter was originally published as
Le, Anh-Huy, Tara L. Mastro, and Susan L. Forsburg. “The C-Terminus of
S. Pombe DDK Subunit Dfp1 Is Required for Meiosis-Specific
Transcription and Cohesin Cleavage.” Biology Open 2.7 (2013): 728–738.
PMC. Web. 18 Dec. 2014.

T. L. Mastro performed the experiments in Figure 2.2 a and b, 2.3 b, 2.4 b, 2.5,
2.6 a, b, c, e and f, and Movies 2.1-9; and contributed to the writing and editing of
the manuscript

35

2.1 Overview
The DDK complex is a conserved kinase complex, consisting of a catalytic
subunit, Hsk1 (Cdc7), and its regulatory subunit Dfp1 (Dbf4). This kinase is
essential for DNA replication. In this work, we show that dfp1-r35, which
truncates the Dfp1 C-terminus zinc finger, causes severe meiotic defects,
including reduced spore viability, reduced formation of programmed double
strand breaks, altered expression of meiotic genes, and disrupted chromosome
segregation. There is a high frequency of dyad formation. Mutants are also
defective in the phosphorylation and degradation of the meiotic cohesin Rec8,
resulting in a failure to proceed through the MII division. These defects are more
pronounced in a haploid meiosis model than in a normal diploid meiosis. Thus,
several critical meiotic functions are linked specifically to the C-terminus of Dfp1,
which may target specific substrates for phosphorylation by Hsk1.
2.2 Introduction
The S. pombe DDK (Dbf4-Dependent Kinase) complex is a conserved,
essential kinase consisting of a catalytic subunit Hsk1 (Cdc7 in humans or
budding yeast) and its regulatory subunit Dfp1 (Dbf4 in humans or budding
yeast) (reviewed in (Duncker and Brown, 2003; Kim et al., 2003; Labib, 2010;
Sclafani, 2000)). In vegetative cells, DDK has an essential function in the
initiation of DNA replication as well as DNA repair and checkpoint activities
(reviewed in (Duncker and Brown, 2003; Kim et al., 2003; Labib, 2010; Sclafani,
2000)). S. pombe dfp1
+
is regulated transcriptionally and post-transcriptionally to

36

restrict kinase activity to S phase (Brown and Kelly, 1999; Takeda et al., 1999;
Tatebe et al., 2001). Dbf4 and its orthologues contain several well conserved
domains to which different functions have been linked (Bailis and Forsburg,
2004; Dolan et al., 2010; Duncker et al., 2002; Fung et al., 2002; Gabrielse et al.,
2006; Harkins et al., 2009; Takeda et al., 1999; Varrin et al., 2005). These
include an N-terminal domain which is required for checkpoint response, a
middle domain M which is associated with replication activities, the short MIR
motif which is required for association with the Swi6 heterochromatin protein
contributing to sister chromatid cohesion and timely replication in the centromere,
and the extreme C-terminal domain which is required for normal response to
alkylation damage and required for meiosis. A simple model suggests that these
different domains of Dfp1 target the DDK to different substrates, although the C-
terminal mutations are also associated with reduced kinase activity (Fung et al.,
2002; Harkins et al., 2009).
DDK has been linked to multiple roles throughout meiosis. In S.
cerevisiae, early studies on a temperature sensitive allele of CDC7 showed the
kinase is required for meiotic recombination (Buck et al., 1991; Sclafani et al.,
1988). More recent experiments have used selectively timed inactivation to
demonstrate that DDK complex is required for meiotic replication initiation
(Valentin et al., 2006; Wan et al., 2006), but also functions in recombination (Lo
et al., 2008; Matos et al., 2008; Ogino and Masai, 2006; Wan et al., 2008; Wan et
al., 2006). In S. cerevisae and in S. pombe, DDK is required to recruit the
nuclease SpRec12/ScSpo11 to the chromosome to generate programmed

37

double strand breaks (prDSBs) (Ogino et al., 2006; Sasanuma et al., 2008; Wan
et al., 2008; Wan et al., 2006). The meiotic transcriptional program is also
regulated by Cdc7 (Lo et al., 2012). S. cerevisiae cdc7 mutants that complete
meiosis often produce diploid dyads, which is linked to the failure of monopolar
attachment at the kinetochore (Lo et al., 2008; Matos et al., 2008; Rabitsch et al.,
2003; Toth et al., 2000). Loss of Rec8 has been shown to relieve the anaphase I
arrest in DDK-depleted cells after meiS phase (Valentin et al., 2006). Recently,
S. cerevisae Cdc7 and CK1 kinases were found to independently collaborate in
the degradation of Rec8 (Ishiguro et al., 2010; Katis et al., 2010).
In this work, we make use of a previously isolated S. pombe dfp1
+
allele
lacking the C-terminus (dfp1-(1-519), also known as rad35) to study the role of
DDK in meiosis. For simplicity, we will refer to this allele as dfp1-r35. Earlier, we
showed that this is a separation-of-function allele that shows a subset of DDK-
associated phenotypes; dfp1-r35 cells are proficient for replication, but defective
in response to alkylation damage (Dolan et al., 2010). Here, we show that dfp1-
r35 cells are proficient for meiotic S phase, but show subsequent defects in the
meiotic program including reduced spore viability, reduced prDSBs, delayed
expression of meiotic genes, and disrupted chromosome segregation. There is a
high frequency of dyad formation. Mutants are also defective in the
phosphorylation and degradation of the meiotic cohesin Rec8, resulting in a
failure to proceed through the MII division. Thus, several critical meiotic
functions are linked specifically to the C-terminal zinc-finger of Dfp1, suggesting it
targets the Hsk1 kinase to several different meiosis-specific substrates. Finally,

38

we demonstrate that the meiotic phenotype of dfp1-r35 varies depending on the
presence of homologous chromosomes.

2.3 Results
2.3.1 Defective meiosis in dfp1-r35 cells
The Dfp1 protein is an essential regulatory subunit for the Hsk1 (Cdc7)
kinase, and targets the kinase to different substrates (reviewed in (Duncker and
Brown, 2003; Labib, 2010)). The C-terminus of S. pombe Dfp1 contains a
putative zinc finger similar in sequence to the C-terminus of S. cerevisiae Dbf4
(Fig 1A). Several truncations of this C-terminus have been constructed; the
mutants are viable and therefore competent for S phase and the replication
function of DDK, but they are defective in response to alkylating damage (Dolan
et al., 2010; Fung et al., 2002; Ogino et al., 2001). Thus, they function similar to
separation of function alleles. The dfp1-r35 mutant, which truncates just 21
amino acids from the C-terminus of the Dfp1 protein (Figure 2.1A), was
previously reported to have defects in the response to alkylation damage (Dolan
et al., 2010).

39

Figure 2.1. Terminal Meiotic Phenotype of dfp1-r35 Mutants. (A) Structure of
Dfp1. dfp1-r35 mutation is a truncation within a zinc-finger domain that is
conserved between S. pombe and S. cerevisiae (B) Nuclear morphology of
terminal meiotic products of dfp1-r35 (FY1154) compared to WT (FY155)
visualized with DAPI staining. The size bar is 10 mm. (C) Quantification of
terminal meiotic phenotypes as horsetailing (HT), 2 DAPI stained bodies, 4 DAPI

40

stained bodies, or abnormal. Defects of dfp1-r35 are suppressed by ectopically
expressed dfp1
+
. 300 asci for each strain were analyzed and error bars represent
standard deviation.
Wild type homothallic (h90) cells undergoing meiosis produce asci with
four regularly shaped spores and evenly segregated nuclei (Figure 2.1B).
However, h90 dfp1-r35 cells form asci with aberrant morphologies (Figure 2.1B).
Only about 24% of observed asci produce four spores. The majority (58%)
produce two spored-asci. In about 14% of asci, at least one spore contains
fragmented or multiple nuclei (Figure 2.1C). Generally, spores are of different
sizes, and spore viability is significantly reduced (1.81±1.46 %) even when
compared to rec12∆ mutants (20.99±5.17 %) that fail to induce prDSBs (Table
2.1). The aberrant ascus morphologies of dfp1-r35 are rescued by a plasmid
containing dfp1
+
, but not by a plasmid containing hsk1
+
, indicating that the
observed effects are specific to the absence of the Dfp1 C-terminus (Figure
2.1C).
Table 2.1 Spore Viability. Fixed numbers of spores were plated after counting.
The standard deviation was calculated from four trials of the experiment
% Viable Spores

Total
Plated
Total
CFU Absolute Normalized
Std.
Dev.
Fold
Reduction
WT 19000 11081 58.32 100.00 15.72
rec12∆ 100000 12240 12.24 20.99 5.17 4.76
dfp1-r35 400000 4225 1.06 1.81 1.46 55.22

41

In order to test how dfp1-r35 affects the progression of meiotic S phase (meiS
phase), we used the temperature sensitive pat1-114 mutation to induce a
synchronous meiosis. Pat1 kinase is an inhibitor of meiosis, and shifting the
temperature to 34°C will cause the mutant to enter meiosis from either a haploid
or a diploid state. While the haploid is frequently used as a meiotic model, the
absence of homologous chromosomes and mating type heterozygosity leads to
some differences in meiotic dynamics (Pankratz and Forsburg, 2005; Yamamoto
and Hiraoka, 2003). Therefore, we constructed a stable h-/ mat2-102 pat1-114/
pat1-114 diploid that maintain heterozygosity and diploidy, and can only enter
meiosis when Pat1 is inactivated. This allows synchronous meiotic progression
as in (Pankratz and Forsburg, 2005). We compared the effects of dfp1-r35 to
wild type and rec12∆ in homozygous h-/mat2-102 diploids.
As expected, wild type and rec12∆ mutants enter and complete meiotic S
phase between two and three hours. There was no evident delay in meiotic DNA
replication in dfp1-r35 mutants following induction of meiosis (Figure 2.2B),
consistent with our prior observations in vegetative cells (Dolan et al. 2010). We
monitored progression through meiotic divisions by counting the number of
nuclei, where the MI division yields 2 DAPI staining spots and the MII division
yields ≥ 3. Over the time course of this experiment, wild type strains showed MI
occurring at 5 hours and MII at 7 hours with nearly all cells completing both
divisions. In contrast, approximately 25% of dfp1-r35 pat1 mutants did not
complete the MII division. There was also a 1hour delay in the MI division
(Figure 2.2A). We observed normal MI dynamics in rec12∆ mutants, but a

42

reduced efficiency of MII, similar to previous reports (Figure 2.2A and (Davis and
Smith, 2003)). However, this defect was not as severe as dfp1-r35.

Figure 2.2. Synchronous Meiosis in mat2-102/h- pat1-114/pat1-114 diploids
and pat1-114 haploids. (A) Comparison of DAPI stained profiles categorizing 1,
2, or ≥3 DAPI stained bodies for WT, dfp1-r35, and rec12∆ following meiotic
induction using pat1-114. (B) FACS profiles demonstrating the progression of
meiotic replication through meiotic induction. DNA content moves from 2C to 4C.
(C) Comparison of haploid meiotic induction of dfp1-r35 compared to wild type.
Both FACs and DAPI staining was done as in A and B to monitor meiotic
progression. Strains: wild type (FY6332/FY6336), dfp1-r35 (FY6347/FY6378),
rec12∆ (FY6530/FY6531), wild type (FY4129), dfp1-r35 (FY4396).
We compared these results to the same experiment in dfp1-r35 pat1-114
haploids, in which cells are induced to undergo meiosis in the absence of
homologous chromosomes. These also proceeded normally through meiotic S
phase, but showed a more striking delay of MI. There was no evidence for MII

43

even after 30 hours (Figure 2.2C). Thus, the haploid pat1-induced meiosis is
more sensitive to dfp1-r35 mutation than the diploid, and in both conditions, the
defect appears to occur after meiS phase.
2.3.2 dfp1-r35 disrupts meiotic transcription
The delay in meiosis in dfp-r35 haploid mutants could reflect a delay in the
overall meiotic program. We examined the timing of meiotic transcript
accumulation, which occurs in characteristic waves (Mata et al., 2007). Early
meiotic gene expression depends in part on the meiosis-specific DNA synthesis
control-like transcription factor complex (DSC1) (Cunliffe et al., 2004; Mata et al.,
2007), which regulates rec12
+
, dfp1
+
, and numerous other genes. There is also
DSC1-independent early transcription of several genes including rec25
+
and
rec27
+
(Mata et al., 2007). The middle wave depends on expression of mei4
+
, a
transcription factor that itself regulates later gene expression including cdc25
+

and mde10
+
(Horie et al., 1998; Murakami-Tonami et al., 2007; Nakamura et al.,
2004). Transcript accumulation is also affected by the presence of regulatory
sequences called DSR elements, which target the mRNA for turnover. These
are found on a number of genes including rec8
+
and mei4
+
and prevent
premature accumulation of their transcripts (Yamamoto, 2010). We investigated
progression through the meiotic program by monitoring accumulation of
messages from several meiosis-specific genes: the DSC targets psm3
+
, rec12
+

and dfp1
+
, a non-DSC early gene rec25
+
, middle meiotic genes mei4
+
, and
mei4
+
-dependent transcripts cdc25
+
and mde10
+
(Figure 2.3).

44

Figure 2.3. Transcriptional Expression of Meiotic Genes. RT-PCR of pat1-
114 (FY4129), pat1-114 rec12∆ (FY2008), pat1-114 rec8∆ (FY1955) and pat1-
114 dfp1-r35 (FY4396) cells undergoing synchronous haploid meiosis shows
expression with altered timing of meiotic markers: mid/late transcripts (cdc25
+
,
mde10
+
, mei4
+
), early rep1-independent (rec25
+
) and -dependent (psm3
+
,
rec12
+
, dfp1
+
).

45

In pat1-114 control cells, psm3
+
and rec12
+
message levels peaked at 2
hours, quickly decreased again by 6 hours, and showed a slight increase again
by 8 hours. Transcription of both mei4
+
and mde10
+
peaked at 6 hours.
Expression of cdc25
+
, another mei4
+
-target, peaked between 4 and 6 hours.
This is consistent with previous descriptions of meiotic gene expression (Mata et
al., 2002; Mata et al., 2007).
These patterns were significantly changed in dfp1-r35 mutants. Both
psm3
+
and rec12
+
messages were induced with normal timing but decreased at a
slower rate and did not experience a rebound at 8 hours. Transcription of mei4
+
,
mde10
+
, cdc25
+
and failed to cycle as in wild type; however, levels of mei4
+
and
mde10
+
increased throughout the time course. Additionally, dfp1
+
transcription
itself was altered, as it did not decrease within the time course. Thus, while
meiosis-specific gene expression occurs in dfp1-r35 cells, it is significantly
disrupted relative to the normal program.
We compared these results to transcription patterns in rec8∆ , which lacks
the meiotic cohesin and rec12∆ which lacks the ability to make DSBs (Cervantes
et al., 2000; Parisi et al., 1999; Watanabe and Nurse, 1999). Expression of the
meiosis-specific transcripts was not affected in rec12∆, despite the lack of
prDSBs, indicating that the transcription program is largely independent of Rec12
or DSB formation (Figure 2.3). In rec8∆ pat1-114 mutants, we observed that the
peak of all but rec25
+
occurred similar to wild type; however, the levels were

46

noticeably higher. Interestingly for both rec8∆ and rec12∆, dfp1
+
levels were
elevated compared to wild type and were already increased upon meiotic
induction. Thus, both rec8∆ and dfp1-r35 cause dysregulation of the meiotic
transcriptional program, although the patterns are different. Importantly, the
expression of meiosis-specific transcripts also confirms that the dfp1-r35 cells
have entered meiosis.
2.3.3 Recombination and Induction of prDSBs in Dfp1 C-terminal
Truncation mutants.
Previous studies have shown that DDK is required for recombination and
induction of prDSBs in both budding and fission yeast (Ogino et al., 2006;
Sasanuma et al., 2008; Wan et al., 2008; Wan et al., 2006). We examined
recombination in known genetic intervals in dfp1-r35 relative to wild type diploids.
Intergenic recombination was determined in the interval between lys4-95 and
his4-239, located on chromosome II, and intragenic recombination in the ade6
hotspot was measured using the ade6-52 and ade6-26 point mutations located
on chromosome III (Figure 2.4A and (Ponticelli et al., 1988)). In wild type cells,
3.98% of germinating spores were both His
+
and Lys
+
recombinants. As
expected, except for one colony, all the His
+
Lys
+
spores recovered from rec12∆
were diploids. In dfp1-r35 mutants, a high fraction of diploids was also
recovered. In the few haploids, a sharp reduction in intergenic recombination
was observed; however unlike rec12∆, it was still detectable at 0.108% His
+
Lys
+
.
We observed a similar reduction in intragenic recombination; in wild type cells,
about 0.285 % of germinating spores were Ade
+
. For rec12∆, we recovered no

47

Ade
+
cells, and for dfp1-r35 mutants we recovered one colony, for a rate of 0.007
%.

Figure 2.4. prDSBs in dfp1-r35 Mutants (A) Pulse Field Gel Electrophoresis
(PFGE) of WT (FY4129), rec12∆ (FY2008), rec8∆ (FY1955) or dfp1-r35
(FY4396) pat1-114 cells undergoing synchronous meiosis. dfp1-r35 mutants ,

48

similar to rec12∆ mutants, cannot induce prDSBs. (B) Quantification of PFGE in
A using the ratio of breaks/ chromosome signal as previously described in (Borde
et al., 2000a)

hsk1 mutants are defective in the induction of prDSBs because they
disrupt the localization of the endonuclease Rec12 (Green et al., 2009;
Sasanuma et al., 2008; Wan et al., 2008). To test whether the recombination
defect in dfp1-r35 was due to the inability to induce prDSBs, we performed
pulsed field gel analysis on agarose plugs prepared from cell cultures undergoing
synchronous pat1 meiosis (Figure 2.4B). Wild type cells typically exhibit a smear
below chromosome III that is observed between 2-4 hours, which has been
shown to represent prDSBs. We observed that prDSBs are significantly reduced
or absent in the negative control strains pat1-114 rec8∆, and pat1 rec12∆
mutants, consistent with previous observations (Cervantes et al., 2000; Ogino et
al., 2006). We found that pat1-114 dfp1-r35 mutants exhibit a smear in all time
points; this is consistent with a constitutive level of DNA damage and DNA
breaks, which was also observed in vegetative cells (Dolan et al., 2010).
However, the smear does not increase in intensity as in wild type, suggesting
that the C-terminus of Dfp1 is important for inducing prDSBs during meiosis.
Chromosome Segregation Defects in Dfp1 C-terminal Truncation mutants.
Importantly, the meiotic phenotype of dfp1-r35 suggests that disruption in
prDSB formation is not its only defect, because rec12∆ mutants, which are

49

completely defective for prDSB formation, nevertheless complete both meiotic
divisions in with a normal timing and transcriptional program. Though rec12∆
mutants also form dyads (Figure 2.2A and (Davis and Smith, 2003)), the level of
dyad formation of the dfp1-35 mutant is dramatically higher than that in the
rec12∆ cells. Consistent with previous findings (Davis and Smith, 2003), rec12∆
spores are about 20 % viable. In contrast, dfp1-r35 mutants retain only about 2%
spore viability. The loss of viability as well as the disruption in segregation and
spore formation suggests that Dfp1 contributes to additional activities,
independent of Rec12-driven prDSBs.
Because dfp1 mutants have a high fraction of dyad asci, we asked
whether this division resembles MI (reductional) or MII (equational) divisions, by
analyzing sister chromatid segregation. We constructed diploid strains
heterozygous for lacO binding sites at the centromere-linked lys1
+
marker
(Tatebe et al., 2001). Upon expression of a lacI-GFP reporter, this array can be
visualized via a single GFP focus associated with one of the chromosome I
homologues.
Using live cell imaging, we monitored meiosis and spore formation. In wild
type diploids, we observed that 97.44% of the meiotic events produced a four
spore ascus (Figure 2.5 and Table 2.2; Movies 2.1-6). 92.31% of wild type
showed the expected reductional segregation at MI, in which the lacI-GFP signal
remains in one nucleus (Figure 2.5 and Table 2.2). During the reductional MII
division, the lacI-GFP signal separates, and because fission yeast tetrads are

50

ordered, this results in two adjacent spores containing the signal (Yokobayashi et
al., 2003).

Figure 2.5. Chromosome Segregation in dfp1-r35 Mutants compared to
rec8∆ and rec12∆. (A) Representative images from live cell analysis of LacI-
GFP chromosome segregation assay in diploid cells (see movies). The indicated
strains heterozygous for lys1
+
::lacO lacI-GFP adjacent to centromere I were
grown at (32˚C) and plated on agaraose pads for image acquisition (see
Materials and methods.). Strains: wild type (FY6221xFY6331), rec8∆
(FYFY5916xFY6131), rec12∆ (FY5197xFY6134), dfp1-r35 (FY6236xFY6441),
dfp1-r35 rec8∆ (FY6204xFY6416), dfp1-r35 rec12∆ (FY6143xFY6175). Scale
bar is 10µm. Selected panels display predominate segregation phenotype for

51

each mutant observed. (B) Graphical representation of quantification of live cell
imaging classes. See Table 2.3 for class descriptions and quantities.
In contrast, while rec8∆ mutants also mostly formed four spores, in
91.67% of cells, the MI division was equational and the lacI-GFP signal split
(Figure 2.5 and Table 2.2). This is similar to the data reported from terminal
phenotype analysis for rec8∆ mutants (Molnar et al., 2001; Yokobayashi et al.,
2003).
Table 2.2 Summary of classes from Figure 2.5
WT
dfp1-
r35 rec12∆ rec8∆
dfp1-
r35
rec12∆
dfp1-r35
rec8∆
1 spore (J) 0.00 0 0 0 0 10.34
2 spores
(A,C,D,E,I) 2.56 75.00 42.55 0.00 44.44 44.83
3 spores (F) 0.00 2.78 10.64 0.00 0.00 0.00
4 spores (B,G,H) 97.44 22.22 46.81 100.00 55.56 44.83

1st is Reductional
(A,B,C,F,G,I) 92.31 80.56 95.74 8.33 55.56 10.34
1st is Equational
(D,E,H,J) 7.69 19.44 4.26 91.67 44.44 89.66

Both dfp1-r35 and rec12∆ diploids exhibited a variety of aberrant
chromosome segregation phenotypes in live analysis. Both dfp1-r35 (75%) and
rec12∆ (43%) formed dyads at a high rate. Interestingly, there was a distinctive

52

difference in segregation in these dyads. The predominant class in dfp1-r35
mutants came from asci that underwent a single reductional division (55.57%
Table 2.3 class A). However, in rec12∆ mutants, the predominant dyads two
meiotic divisions, but formed a single spore around two nuclear bodies, resulting
in an apparent bi-nuclear dyad (44.68% Table 2.3 class B).
Table 2.3 Chromosome Segregation monitored with LacI-GFP.
Chromosome segregation assay using live cell imaging and parental strains
heterozygous for LacO array. Cells were sorted into different segregation
Class Definitions Percentage of Classes
Class
#
Divisions
1st
Division
2nd
Division
#
Spores WT
dfp1-
r35 rec12∆ rec8∆
dfp1-
r35
rec12∆
dfp1-
r35
rec8∆
A 1 R no 2 0.00 55.56 6.38 0.00 22.22 0.00
B 2 R E 4 89.74 16.67 44.68 8.33 33.33 6.90
C 2 R E 2 2.56 2.78 29.79 0.00 0.00 0.00
D 2 E yes 2 0.00 2.78 0.00 0.00 0.00 0.00
E 1 E no 2 0.00 13.89 4.26 0.00 22.22 41.38
F 2 R E 3 0.00 2.78 10.64 0.00 0.00 0.00
G 2 R R 4 0.00 2.78 2.13 0.00 0.00 0.00
H 2 E yes 4 7.69 2.78 0.00 91.67 22.22 37.93
I 2 R R 2 0.00 0.00 2.13 0.00 0.00 3.45
J 1 E no 1 0.00 0.00 0.00 0.00 0.00 10.34
Total cells analyzed 39 36 47 36 27 29

53

classes dependent on whether a reductional or equational division occurred at MI
and how many spores were formed in the terminal product. Data is pooled from
at least 3 biological replicates.

2.3.4 Rec8 Dynamics in Dfp1 C-terminal Truncation mutants.
The differences in chromosome segregation defects in dfp1-r35 and
rec12∆ are consistent with data from budding yeast suggesting that DDK is
required to phosphorylate and inactivate the Rec8 cohesin that maintains
association between sister chromatids (Green et al., 2009; Lo et al., 2008; Matos
et al., 2008; Molnar et al., 1995; Parisi et al., 1999; Rabitsch et al., 2003; Toth et
al., 2000; Watanabe and Nurse, 1999)). Deletion of S. cerevisiae rec8 partly
rescues the phenotype associated with a cdc7 mutant (Valentin et al., 2006). We
constructed homozygous double mutant diploids dfp1-r35 rec12∆ or dfp1-r35
rec8∆. Both of these partly rescued the dfp1-r35 dyad phenotype from 75% to
44.44% and 44.83% respectively (Figure 2.5 and Table 2.2). The dfp1-r35 rec8∆
double mutant showed the same fraction of equational MI divisions as the rec8∆
single mutant, indicating that rec8∆ is epistatic to dfp1-r35 (Figure 2.5 and Table
2.2).
The dfp1-r35 phenotype resembles the phenotype observed in mutants
containing a non-cleavable form of Rec8 (Kitajima et al., 2003), and the partial
rescue of dfp1-r35 dyad formation by rec8∆ suggests that some of its defect
reflects the inability to inactivate Rec8. We therefore examined whether Rec8

54

stability is affected by deletion of the C-terminus of Dfp1, using live cell imaging
and western blot analysis in wild type and mutant cells.
As previously described (Watanabe and Nurse, 1999), we observed that wild
type diploids accumulate a pan-nuclear signal of rec8-GFP during meiosis. This
signal is reduced to two puncta at the MI division, which completely disappear
following MII (Figure 2.6A; Movies 2.7-9). We noted a slight delay in the
disappearance of Rec8-GFP in dfp1-r35 cells relative to the MI division. In wild
type the pan-nuclear signal decreased to a single focus at 10min post MI;
however, dfp1-r35 showed this with an average timing of 15.5min. Also, the
dfp1-r35 cells had a large variation in the timing, while the wild type cells
behaved more consistently (Figure 2.6F). Surprisingly, we saw an even more
pronounced delay in rec12∆ mutants, in which the pan-nuclear signal persisted
for an extended time of an average of 50.5 mins. Again we observed a larger
variation in the mutant compared to wild type; however, the difference between
wild type and rec12∆ is significant using a two-tailed t-test with a p-value of 8 x
10
-7
(Figure 2.6F). In wild type, Rec8-GFP foci disappeared 48mins after the
conversion from pan-nuclear signal to single focus; while this disappearance
occurred at 56.5min in dfp1-r35, and 55.5 min in rec12∆ (Figure 2.6F).

55

Figure 2.6. Rec8-GFP Stability. (A) Representative images from live cell
imaging observing diploid meiosis in cells containing Rec8-GFP (see movies).
The indicated strains were grown at (32˚C temperature) and plated on agaraose
pads for image acquisition (see Materials and methods.). Strains: wild type
(FY6173xFY6174), dfp1-r35 (FY6241xFY6242), rec12∆ (FY6217x6218). Scale
bar is 10µm. (B) Western blot of Rec8-GFP during synchronous meiosis in h-
/mat2-102 pat1-114/pat1-114 stable diploid. Strains: wild type (FY6332/FY6336),
dfp1-r35 (FY6347/FY6378) rec12∆ (FY6530/FY6531). (C) Quantification of B
using GFP signal/ PCNA. (D) Western blot of Rec8-GFP in haploid pat1-114
meiosis. Strains: wild type (FY4129) and dfp1-r35 (FY4396). (E) Quantification

56

of D as done in C. (F) Graphical representation of quantification of Rec8-GFP
timing in live cell imaging in A. Pan-nuclear signal disappearance occurred at an
average of 10mins, 15.5mins, and 50.5mins post MI for wild type, dfp1-r35, and
rec12∆ respectively with p-values using a two-tailed t-test of 0.0000008 and
0.102 for rec12∆ and dfp1-r35 respectively. For the disappearance of the
nuclear focus relative to the disappearance of the pan-nuclear signal the timing
was an average of 48mins, 56.6mins, and 55.5mins for wild type, dfp1-r35, and
rec12∆ with p-values using a two-tailed t-test of 0.258 and 0.599 for rec12∆ and
dfp1-r35 respectively.
In order to get a higher resolution analysis of Rec8-GFP dynamics in both
dfp1-r35 and rec12∆ mutants, we induced synchronous meiosis in a mat2-102
stable diploid using pat1-114 temperature inactivation, and used western blots to
visualize protein levels (Figure 2.6B and C). In wild type cells, Rec8-GFP protein
levels peak at four hours and are undetectable by 6 hours (Figure 2.6B and C).
We observed an electrophoretic mobility shift in Rec8-GFP at four and five hours
in wild type cells, prior to its disappearance. This corresponds to a previously
reported phospho-shift found to be important for Rec8 degradation (Ishiguro et
al., 2010; Parisi et al., 1999; Rumpf et al., 2010). We observed that the amount of
Rec8-GFP in dfp1 mutants levels did not increase to the level of wild type. The
peak of Rec8-GFP in dfp1-r35 was observed at three hours rather than four as in
wild type, and a low level of Rec8-GFP was still detectable in the mutant at later
time points. The Rec8 mobility shift was undetectable in dfp1 mutant. rec12∆
diploids were similar to wild type in terms of Rec8-GFP dynamics and mobility

57

shift. Thus, the prolonged pan-nuclear staining we observe during MI is not due
to changes in protein level, but to defects in localization.
We observed that the dfp1-r35 Rec8-GFP phenotype is more pronounced
in haploids compared to diploids (Figure 2.6D and E). There was a reduced level
of total protein, and Rec8-GFP persisted even after 24 hours post meiotic
induction, suggesting that the inability of haploid pat1 dfp1-r35 mutants to
undergo a second meiotic division is due to the persistence of Rec8 in this strain.
Interestingly, the mobility shift associated with phosphorylation was much less
apparent in the haploid than the diploid.
2.4 Discussion
DDK was first identified as an S phase specific kinase essential for DNA
replication (Duncker and Brown, 2003; Kim et al., 2003; Labib, 2010; Sclafani,
2000)). Subsequent studies have linked it to a wide range of activities temporally
linked to S phase. Separation of function alleles in the fission yeast DDK
regulatory subunit dfp1
+
suggest that different functions may map to different
Dfp1 domains (Bailis et al., 2003; Dolan et al., 2010; Fung et al., 2002; Hayashi
et al., 2009). Previously, the Dfp1 C-terminus was shown to be important to the
response to alkylation damage, possibly through mediating trans-lesion synthesis
repair (Dolan et al., 2010; Fung et al., 2002). Our current work shows that the C-
terminus of Dfp1, which contains in a zinc finger domain truncated by the dfp1-
r35 mutation, influences DDK activity during meiosis, leading to multiple defects

58

including loss of programmed double strand breaks, disruption in the meiotic
transcription program, and defects in turnover of the meiotic cohesin Rec8.
Early studies on DDK in meiosis in S. cerevisiae linked DDK to meiotic
recombination, rather than replication (Buck et al., 1991; Sclafani et al., 1988).
Subsequently, it was determined that DDK is important for activation of the
Rec12 /Spo11 endonuclease in both budding and fission yeasts (Ogino and
Masai, 2006; Sasanuma et al., 2008; Wan et al., 2008; Wan et al., 2006). In
budding yeast, DDK phosphorylates Mer2 (Sasanuma et al., 2008; Wan et al.,
2008), although a homologue of this substrate has not been identified in fission
yeast. We show that S. pombe dfp1-r35 mutants are proficient for meiotic S
phase, but defective in prDSB formation, with dramatically reduced
recombination. The residual recombination in dfp1-r35 relative to rec12∆ may
reflect the presence of a low level of DNA damage and double strand breaks in
dfp1-r35 (Dolan et al., 2010); there is evidence that mutations that cause low
levels of DNA damage may provide Rec12-independent substrates for
recombination (Farah et al., 2005a; Farah et al., 2005b; Pankratz and Forsburg,
2005).
The defects of dfp1-r35 in meiosis are far more severe than those
observed in rec12∆ cells, including substantially reduced spore viability and
disruptions of segregation, indicating it plays roles beyond induction of
programmed double strand breaks. Consistent with this, we also observed
dysregulation of meiosis-specific gene expression in dfp1 mutants. Early
transcripts (including rec8
+
, rec12
+
, and dfp1
+
itself) accumulate with normal

59

timing. However, accumulation of mid-meiotic transcripts, including mei4
+
, is
noticeably delayed in dfp1-r35 mutants. Mei4 is a fork-head transcription factor
that is responsible for expression of numerous downstream genes required for
meiotic divisions and sporulation (Horie et al., 1998; Mata et al., 2002).
Expression of mei4
+
occurs early in meiosis, but there is no accumulation of its
transcript due to regulated turnover by the Mmi1 protein, operating through the
DSR sequence element (Harigaya et al., 2006). The same Mmi1 system
contributes to the turnover of early meiotic transcripts such as rec8
+
(Yamanaka
et al., 2010). Because the rec8
+
transcript accumulates normally in dfp1-r35
cells, the failure to accumulate mei4
+
cannot be due to a failure to inactivate
Mmi1.
There is evidence in budding yeast that its mid-meiosis transcription
factor, Ndt80, is regulated by DDK through an indirect mechanism (Lo et al.,
2012). Although ScNdt80 performs a similar function to Sp Mei4, the proteins are
not structurally related. It is possible that the DDK regulation of the mid-meiotic
transcription factors occurs through other, more conserved proteins.
Alternatively, there may be broad functional similarities but molecularly distinct
mechanisms involved.
We also find that dfp1 cells largely bypass the MII division, forming dyads.
Loss of viability and dyad formation can be partly rescued by deletion of Rec8
cohesin, suggesting that persistence of Rec8 in dfp1 hinders meiotic progression.
Previously, we showed that Dfp1 interacts with Psc3, a component of the mitotic
cohesin complex (Bailis et al., 2003), which could suggest a direct interaction

60

with the cohesins. We find that the characteristic Rec8 phosphorylation mobility
shift is reduced in dfp1 cells, and the Rec8 protein persists until late in meiosis.
This is particularly apparent in haploid cells, and suggests that Rec8 turnover
may be DDK-dependent. Turnover of Rec8 is driven by phosphorylation that
depends in part upon casein kinase 1 (Ishiguro et al., 2010; Parisi et al., 1999;
Rumpf et al., 2010)(Katis et al., 2010). Our study indicates that Rec8
phosphorylation also depends upon DDK. This is consistent with work from
budding yeast suggesting that DDK collaborates with CK1 in Rec8
phosphorylation (Katis et al., 2010). A priori , we cannot conclude whether DDK
acts directly upon Rec8, or whether it regulates the activity of CK1, although
based on analogy to budding yeast, the former seems likely.
Together, these observations link DDK to multiple functions in fission
yeast meiosis as in budding yeast: induction of programmed DSBs, regulation of
the mid-meiotic transcription program, and turnover of the Rec8 cohesin required
for proper MII division. This is particular striking as the overall logic of meiotic
regulation, and many drivers of meiotic progression, are not conserved in the two
yeasts.
How does one domain of Dfp1 target the DDK to such diverse activities?
One possibility is that this mutation simply reduces kinase activity, as proposed in
(Fung et al., 2002; Harkins et al., 2009). However, dfp1-r35 cells are competent
for replication in both vegetative and meiotic cells, which shows that that
sufficient kinase activity remains for this essential function, even though meiosis
is severely disrupted. If a loss of kinase activity is responsible for all the defects

61

we observe, it would suggest a substantial difference in the threshold kinase
activity required for different activities. Another possibility is that the C-terminal
zinc finger domain truncated in dfp1-r35 contributes to substrate recognition.
We showed previously that Dfp1 chromatin association in response to MMS
treatment is disrupted by loss of this domain (Dolan et al., 2010). Since meiotic
cohesion and prDSB formation are both linked to progression of S phase (Doll et
al., 2008; Kugou et al., 2009; Ogino and Masai, 2006), it is plausible that both
Rec8 and Rec12 meet the replication fork, and possibly, a fork-linked DDK.
An additional finding of our study is a significant difference between the
phenotype of dfp1-r35 mutants in haploid and diploid driven meiosis, with the
haploid being more severely affected. These differences are particularly relevant
because the pat1 temperature-induction system in haploids is frequently used to
model meiosis in fission yeast. Previously, it was shown that monopolar spindle
attachment prior to the MI division, which is essential for reductional segregation,
has different requirements depending on the ploidy of the cell (Yamamoto and
Hiraoka, 2003). In haploids, monopolar attachment depends on Pat1
inactivation, pheromone signaling, and Rec8. However, it is independent of
Rec12 and recombination. In contrast, in diploids, monopolar attachment is
independent of pheromone signaling but requires Rec12 and recombination, in
addition to Rec8 and Pat1 inactivation.
In our study, we observe that haploid pat1-114 dfp1-r35 cells fail to
proceed through MII of meiosis, accompanied by failure to degrade the Rec8
cohesin. Interestingly, we see is a more modest mobility shift of Rec8 in wild type

62

haploids compared to diploids. We speculate that this might indicate that the role
of CK1 in Rec8 phosphorylation is reduced in haploids, although this remains to
be tested.
In contrast, in diploid cells homozygous for pat1-114 dfp1-r35, even
though the majority of cells do not complete MII, we see only a modest reduction
of Rec8 turnover; a similar phenotype is observed for a pat1
+
dfp1-r35 diploid.
The MII arrest is partly (but not completely) rescued by rec8∆, indicating that
diploids homozygous for dfp1-r35 have additional defects beyond difficulties in
regulating Rec8. These likely include disruptions in recombination and the
transcriptional program, as we observe, or changes in chromosome or rDNA
segregation as in (Bailis and Forsburg, 2004; Sullivan et al., 2008).
By analogy to the effect on monopolar spindle attachment reported by
(Matos et al., 2011; Yamamoto and Hiraoka, 2003), our data may also suggest
that a Rec12-dependent pathway in the diploid ameliorates the effects due to
dfp1-r35 just as it contributes to monopolar attachment. Interestingly, Rec12
protein persists and has been suggested to play a role in MII (DeWall et al.,
2005), so there may be some additional effects of proper pairing of homologous
chromosomes in the context of recombination that is independent of DDK. These
results provide a caveat in the use of haploid pat1 strains to model meiotic
progression.

63

2.5 Materials and Methods
2.5.1 Strains and Media
S. pombe strains are indicated in Table S1. Standard genetic techniques
and media were used to construct and maintain strains (Sabatinos and Forsburg,
2010).
2.5.2 Microscopy
For terminal meiotic phenotypes, microscopy was performed on live or
ethanol fixed cells stained with DAPI (Sabatinos and Forsburg, 2010) using a
63X oil-immersion lens (PLApo, NA=1.32) on a Leica DMR fluorescence
microscope. Images were collected with Openlab 3.6.1 contrast adjusted and
assembled in Canvas 12.
For LacI-GFP chomosome segregation live cell imaging, cells were plated
on agarose pads as in (Green et al., 2009). Images were acquired with a
DeltaVision Core wide field deconvolution microscope (Applied Precision,
Issaquah, WA) using an Olympus 60X/1.40, PlanApo, NA=1.40 objective lens
and a 12-bit Photometrics CoolSnap HQII CCD, deep-cooled, Sony ICX-285
chip. The system x-y pixel size is 0.1092µm x-y. softWoRx v4.1 (Applied
Precision, Issaquah, WA) software was used at acquisition electronic gain=1.0
and pixel binning 1x1. Excitation illumination was from a Solid-state illuminator
(7 color version), GFP was excited and detected with a (ex)475/28,(em)525/50
filter set and a 0.2 second exposure. A polychroic mirror was used GFP/mCherry
Chroma ET C125705 roughly: 520/50–630/80. Nine z sections at 0.5µm were

64

acquired. 3-D stacks were deconvolved with manufacturer provided OTFs using
a constrained iterative algorithm and images were maximum intensity projected.
Images were contrast adjusted using a histogram stretch with an equivalent scale
and gamma for comparability. Brightfield images were acquired with DIC.
Images were assembled using softWoRx Explorer v. 1.3.0 and Photoshop CS3 v.
10.0.1.
2.5.3 Spore Viability Assay and Recombination Assays
The recombination assay was performed as described (Catlett and
Forsburg, 2003). Indicated strains were mated on ME agar plates for three days
at 25°C. Asci were treated with 0.5% glusulase (Perkin Elmer) solution and
incubated on a rotating platform overnight at room temperature. For each trial
spores for all strains were distributed over ten plates. For each trial,
approximately 5,000, 35,000, or 100,000 spores were plated for wild type,
rec12∆, or dfp1-r35 respectively. Spore viability is represented as the fraction of
resulting colonies over the total number of plated spores. The standard deviation
was calculated from four trials. Spore viability of mutants was normalized to wild
type, which was defined as 100% viable. YES plates were replica plated onto
EMM plates lacking histidine, lysine, adenine, or histidine and lysine. The fraction
of autotrophs on the respective media was tabulated to determine recombination
frequencies. Diploids were distinguished from haploids via cell morphology, color
on phloxinB, and FACs. Data were pooled from four trials.

65

2.5.4 Induction of meiosis
For homothallic h90 strains, cultures were grown overnight at 25°C to an
OD
595
nm of 0.8 in 10 ml of EMM media with appropriate supplements. Cells were
washed twice in 10 ml EMM-N media, and then starved in ME media for 30 hrs.
1ml of cells suspension was fixed in 70% ethanol, and stained with
DAPI(Sabatinos and Forsburg, 2010).
Induction of meiosis for live cell imaging was accomplished by co-culturing
h- and h
+
strains of appropriate genotype independently at 32˚C to OD of ~1.
Cells were then washed 2x in EMM-N. Cell pellets of each were combined into
12ml ME media at 25˚C for 12-16hrs. Cells were then centrifuged at low speed
(1000 x g) and placed on a SPAS 2% agarose pad for imaging at 25˚C.
Induction of synchronous meiosis was carried out using a pat1
temperature sensitive allele as described (Forsburg and Hodson, 2000). Briefly,
200 ml of cell cultures were grown in EMM media supplemented with appropriate
nutritional supplements overnight to reach an OD595nm of ~0.8. Cultures were
washed twice in EMM-N media and were starved at 25°C in 200 ml of EMM-N
Media for ~17 hrs in order to arrest cells in G1. Haploid strains were
supplemented with 7 mg/ml adenine. To induce meiosis in cell cultures, the
temperature was shifted to 34°C by adding an equal volume of pre-heated (36°C)
EMM media containing 1 g/l of NH4Cl and 70 mg/ml uracil, adenine and leucine
to starved cultures, and by incubating cultures in a water bath at 34°C. Samples
for nuclear counts, FACS, RT-PCR, and Westerns were taken at indicated times.

66

2.5.5 Flow Cytometry (FACS)
Flow cytometry was performed as described (Sabatinos and Forsburg,
2009). Briefly, 200 ml of cell suspension fixed in 70% ethanol were rehydrated by
washing twice in 1 ml of 0.05 M sodium citrate solution and removing the
supernatant after each wash. Rehydrated cells were resuspended in 500 ml of
0.05 M sodium citrate solution with 0.1 mg/ml RNaseA, and incubated for 90
minutes at 36°C. Cells were stained with Sytox Green (Invitrogen) by adding 500
ml 0.05 M sodium citrate solution with 0.1 mM Sytox Green. Cell suspensions
were sonicated with three pulses at 20% amplitude, and vortexed once more
shortly before submission of samples to FACS analysis.
2.5.6 Reverse Transcriptase PCR
RNA was extracted using QIAGEN’s RNAse Easy RNA extraction kit.
Extracted RNA was quantified on a spectrophotometer (Nanodrop, ND1000).
RNA was converted to cDNA using Roche kit. Relative transcription levels were
determined by performing multiplex PCR using target primer and actin primers as
described above (Table S2). Actin primer sequences from (Kloc et al., 2008)
were used. Quantification was done using QuantOne software version 4.6.9
taking a ratio of target to actin signal.
2.5.7 Detection of Meiotic prDSBs by Pulsed Field Gel Electrophoresis
(PFGE)
Agarose plugs were prepared as described (Cervantes et al., 2000).
Briefly, 50 ml of cultures were stopped by adding 500 ml of 20% sodium azide

67

solution to the culture and incubating the culture on ice for 5 minutes. Harvested
cultures were spun down, washed once in 10 ml of PBS buffer, and once in 10 ml
of CSE buffer (20 mM citric acid, 20 mM Na2HPO4, 40 mM EDTA, 1.2 M sorbitol
at pH 5.6). Each culture was digested in CSE media containing 0.45 mg/ml of
Lysing Enzyme, and 0.02 mg/ml of zymo 100T for 20-40 minutes. Digestion was
monitored by microscopy. Plugs were prepared from digested cell pellets that
were resuspended in TSE (10 mM Tris pH 7.5, 45 mM EDTA, pH 8.0, 0.9 M
sorbitol) to make a cell suspension of 3.0x10 9 cells/ml. Plugs were treated with
Protinase K in Sarkosyl-EDTA (1% sarkosyl, 0.5 M EDTA, 1mg/ml Protease K,
calibrated to a pH of 9.5) solution for 48 hours. The Sarkosyl-EDTA-Protease K
solution was changed once after 24 hours. Plugs were washed three times in 10
ml TE Buffer, and three times in 10 ml TAE buffer (40 mM Tris acetate, 1 mM
EDTA at pH 8.0).
PFGEs were run on a Biorad Chef II Pulse Field machine with the
following specifications: 48 hours at 2 Volts/cm at an angle of 106° and pulse
times ramping from 1200 to 1800 seconds. The gel was stained in 200 ml of
TAE Buffer containing SYBR Green for 20 minutes, and bands were visualized in
a ChemiDoc Machine (XRS, BioRad). Quantification of breaks using the ratio of
breaks/ chromosome signal as previously described in (Borde et al., 2000a;
Borde et al., 2000b) with BioRad QuantOne software version 4.6.9

68

2.5.8 Determination of Rec8 Stability
We performed video microscopy on diploid cells entering meiosis
expressing Rec8-GFP and quantified these by scoring the time at which the
Rec8-GFP signal decreased from pan-nuclear to a focus in relation to the MI
division. The time at which the Rec8-GFP focus disappeared was measured
from the point at which the pan-nuclear signal became reduced. A two-tailed t-
test was applied to determine significance in the different strains.
For analysis of protein, TCA precipitation was performed as described
(Foiani et al., 1994). Cell cultures were stopped by adding 10x STOP buffer
containing sodium azide solution to harvested culture and incubating the cultures
on ice for 10 minutes. Cells were washed in PBS buffer (137 mM NaCl, 2.7 mM
KCl, 4.3 mM Na2HPO4,1.47 mM KH2PO4) and then MQ water. Protein was
extracted using TCA precipitation. Protein extracts were quantified using BCA
and an equal amount of protein for each sample were run on a 8% SDS PAGE
gel containing 1.25% crosslinker. Membrane was blocked in 5% PBST (1xPBS
solution with 2% Tween-20). To detect Rec8-GFP, membrane was incubated in
5% PBST milk containing a 1:2000 dilution of JL8 monoclonal antibodies
(Clontech) overnight at 4˚C, and washed three times for 10 minutes in 10 ml
PBST each time. Blots were incubated in 5% PBST milk solution with 1:3000
secondary goat anti-mouse HRP antibodies (Millipore). Blots were developed
using ECL (Pierce). Quantification was done using QuantOne software version
4.6.9 taking a ratio of GFP to PCNA signal.

69

Table 2.4. Strains used in this study.
Name Genotype Reference
FY155 h90 968 Our stock
FY1154 h90 dfp1-r35 ura4-D18 leu1-32
Henning
Schmidt
FY1319 h
-
∆rec8::ura4
+
leu1-32 ura4-D18 ade6-M210 Our stock
FY2008 h
-
pat1-114 ∆rec12::LEU2 leu1-32 ade6-M216 Our stock
FY3973 h
-
dfp1-r35 leu1-32 ura4-D18 ade6-M210 This work
FY4129 h
-
pat1-114 ura4-D18 ade6-M216 This work
FY4396 h
-
pat1-114 dfp1-r35 ade6-M216 This work
FY4397 h
-
dfp1-r35 ura4-D18 ade6-M52 lys4-95 This work
FY4398 h
-
ura4-D18 ade6-M52 lys4-95 This work
FY4405 h
+
leu1-32 ade6-M26 his4-239 This work
FY4518 h
+
dfp1-r35 leu1-32 ade6-M26 his4-239 This work
FY4561
h
+
∆rec12::LEU2 ura4-D18 leu1-32 ade6-M26 his4-
239 This work
FY4843
h
+
dfp1-r35 sad1
+
::DsRed-LEU2 his7
+
::lacI-GFP
This work

70

lys1
+
::lacO leu1-32 ura4-D18 ade6-M216
FY4915
h
+
∆rec8::ura4
+
sad1
+
::DsRed-LEU2 his7
+
::lacI-GFP
lys1
+
::lacO leu1-32 ura4-D18 ade6-M216 This work
FY4991
h
-
pat1-114 rec8GFP::Kan leu1-32 ade6-M210
can1-1 This work
FY5164
h
-
pat1-114 dfp1-r35 rec8GFP::Kan leu1-32 ade6-
M210 This work
FY5268 h
-
∆rec12::ura4
+
ura4-D18 leu1-32 ade6-M52 lys4-95 This work
FY5916
h
+
∆rec8::ura4
+
his7
+
::lacI-GFP lys1
+
::lacO ura4-D18
leu1-32 This work
FY5917
h
+
∆rec12::ura4
+
his7
+
::lacI-GFP lys1
+
::lacO leu1-32
ura4-D18 This work
FY6131
h
+
leu1-32 ura4-D18 rec8::ura4
+
arg3
+
::D0817-
mCherry::his5
+
his5D? This work
FY6134
h
-
ura4-D18 rec12::ura4
+
arg3
+
::D0817-
mCherry::his5
+
leu1-32 hsi5D? This work
FY6143
h
-
∆rad35-271 rec12::ura4
+
his7
+
::lacI-GFP lys1
+
::lacO leu1-32 ura4-D18 This work
FY6173 h
+
rec8-GFP-kan leu1-32 ade6-M210 This work

71

FY6174 h
-
rec8-GFP-kan leu1-32 ade6-M210 This work
FY6175
h
-
ura4-D18 rad35-271 rec12::ura4
+
arg3
+
::D0817-
mCherry::his5
+
leu1-32 hsi5D? This work
FY6204
h
-
rad35-271 ∆rec8::ura4
+
arg3
+
::D0817-
mCherry::his5
+
his5D? ura4-D18 leu-32 This work
FY6217
h
+
rec8-GFP-kan ∆ rec12::ura4
+
ade6-M210 leu1-32
ura4-D18 This work
FY6218
h
-
rec8-GFP-kan ∆rec12::ura4
+
ade6-M210 leu1-32
ura4-D18 This work
FY6221 h
+
his7
+
::lacI-GFP lys1
+
::lacO leu1-32 This work
FY6236
h
+
rad35-271 arg3
+
::D0817-mCherry::his5
+
his5D?
leu1-32 This work
FY6241 h
-
rad35-271 rec8-GFP-kan leu1-32 This work
FY6242 h
+
rad35-271 rec8-GFP-kan leu1-32 This work
FY6331 h
-
arg3
+
::D0817-mCherry::his5
+
leu1-32 his5D This work
FY6332 h90 mat2-102 pat1-114 rec8-GFP-kan ade6-M210 This work
FY6336 h
-
pat1-114 rec8-GFP-kan ade6-M216 This work
FY6347 h
-
pat1-114 rad35-271 rec8-GFP-kan ade6-M216 This work

72

FY6416
h
+
rad35-271 ∆rec8::ura4
+
his7
+
::lacI-GFP lys1
+
::lacO ura4-D18 leu1-32 This work
FY6441 h
+
rad35-271 his7
+
::lacI-GFP lys1
+
::lacO leu1-32 This work
FY6478
h90 mat2-102 pat1-114 rad35-271 rec8-GFP-kan
leu1-32 ade6-M210 This work
FY6530
h90 mat2-102 pat1-114 rec12∆::ura4
+
ura4-D18 rec8-
GFP-kan ade6-M210 This work
FY6531
h
-
pat1-114 rec12∆::ura4
+
ura4-D18 rec8-GFP-kan
ade6-M216 This work

Table 2.5 Oligos used for RT-PCR.
Target Oligo Sequence (5'
-
3') Source
mei4 TTGGAGATGAAATGGCGGGCTTTG This work
mei4 GTAGCGAAACGTGTTGCGAATCCA This work
cdc25 GCACAGAGCACCTCATTTGGCATT This work
cdc25 AACATGCGAAGCATCGTTCATCGG This work
mde10 CAGTTCGCTTTGCGATGCTCAAGA This work
mde10 AACGATGCCGTTTCCACATGTTCC This work
rec25 GAGAGAGACACCGGACATGAT This work
rec25 CCCTTGTATAAGCTGAGACCCTTG This work

73

psm3 TTCAAGGTCGCTGTTGAGGCTACT This work
psm3 TGGCATCAGGATAAGTAACCGCCT This work
rec12 CCGAAGAGGCTTAGCGGATACAA This work
rec12 ACTGTGCGGTTCATGACTGTAGG This work
act1 TGCACCTGCCTTTTATGTTG (Kloc et al. 2008)
act1 TGGGAACAGTGTGGGTAACA (Kloc et al. 2008)

2.6 Legends for Movies
Movies 2.1-6: Representative movie of live cell imaging. These movies are the
primary data for the time-lapse images in Figure 5. Yellow signal is LacI-GFP.
Image is fusion of transmitted light and GFP signal. Scale bar is 10 microns.
Movies 1-6 show the following genotypes in order: wild type, dfp1-r35, rec12∆,
rec8∆, dfp1-r35 rec12∆, dfp1-r35 rec8∆.
Movies 2.7-9: Representative movie of live cell imaging of Rec8-GFP in
asynchronous meiosis. These movies are the primary data for the time-lapse
images in Figure 6. Green signal is Rec8-GFP. Image is fusion of transmitted
light and GFP signal. Scale bar is 10 microns. Movies 7-9 show the following
genotypes in order: wild type, dfp1-r35, rec12 ∆.

74

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Chapter 3

Increased Meiotic Crossovers and Reduced Genome Stability in Absence
of Schizosaccharomyces pombe Rad16 (XPF).

The work in this chapter was originally published as
Increased Meiotic Crossovers and Reduced Genome Stability in Absence of
Schizosaccharomyces pombe Rad16 (XPF). Mastro TL, Forsburg SL. Genetics.
2014 Dec;198(4):1457-72. doi: 10.1534/genetics.114.171355. Epub 2014 Oct 6.

T. L. Mastro performed all the experiments, and contributed to the writing and
editing of the manuscript.

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3.1 Overview
Schizosaccharomyces pombe Rad16 is the orthologue of the XPF
structure-specific endonuclease, which is required for nucleotide excision repair
and implicated in the single strand annealing mechanism of recombination. We
show that Rad16 is important for proper completion of meiosis. In its absence,
cells suffer reduced spore viability and abnormal chromosome segregation with
evidence for fragmentation. Recombination between homologous chromosomes
is increased, while recombination within sister chromatids is reduced, suggesting
that Rad16 is not required for typical homologue crossovers but influences the
balance of recombination between the homologue and the sister. In vegetative
cells, rad16 mutants show evidence for genome instability. Similar phenotypes
are associated with mutants affecting Rhp14
XPA
but are independent of other
nucleotide excision repair proteins such as Rad13
XPG
. Thus, the XPF/XPA
module of the nucleotide excision repair pathway is incorporated into multiple
aspects of genome maintenance even in the absence of external DNA damage.
3.2 Introduction
The XPF/ERCC1 protein complex is one of several structure-specific
endonucleases that function broadly as resolvases in the repair of damaged DNA
(SCHWARTZ AND HEYER 2011). Rad16/Swi9 is the fission yeast orthologue of
endonuclease XPF, which forms a complex with Swi10/Rad23 (ERCC1) (CARR et
al. 1994). XPF has a conserved role in nucleotide excision repair (NER) to

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remove UV induced lesions in the DNA (CAMENISCH et al. 2006; GREGG et al.
2011). In humans, mutation of XPF is associated with Xeroderma Pigmentosum
(XP), which causes sun sensitivity and high rates of skin cancer, as well as
premature aging and neurological disorders (CAMENISCH et al. 2006; GREGG et al.
2011). Recent studies suggest XPF mutations are also associated with Fanconi
Anemia (FA), which requires repair of interstrand crosslinks (ICL) (BOGLIOLO et
al. 2013; KASHIYAMA et al. 2013). The canonical model for NER suggests that
upon recognition of helix-distorting lesions, the DNA is unwound to form a bubble
around the damage. XPA (SpRhp14) loads XPF (SpRad16) and ERCC1
(SpSwi10); XPF cleaves at the 5’ end of the bubble while XPG (SpRad13),
another endonuclease, cleaves at the 3’ end to remove the offending segment
(FAGBEMI et al. 2011; SCHWARTZ AND HEYER 2011). Thus, S. pombe rad16
mutants show a decrease in (6-4) photoproduct excision (MCCREADY et al. 1993;
CARR et al. 1994).
Consistent with these clinical effects, XPF is implicated in multiple
mechanisms of genome maintenance (PAQUES AND HABER 1997; GREGG et al.
2011; SCHWARTZ AND HEYER 2011). XPF is required for Single Strand Annealing
(SSA), a form of double strand break (DSB) repair distinct from typical
Homologous Recombination (HR) (MA et al. 2003; KASS AND JASIN 2010). This
occurs when short regions of homology exposed by resection are able to pair,
leaving non-homologous 3’ overhangs as substrates for XPF cleavage. In
budding yeast, recruitment of ScRad1
XPF
and ScRad10
ERCC1
in this pathway
depends on interactions with other proteins including the recombination mediator

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Rad52, and a scaffold provided by Saw1 and Slx4 (rev in (LYNDAKER AND ALANI
2009)). Recent studies have also implicated ScRad1
XPF
in recombination
between dispersed repeats (SYMINGTON et al. 2000; MAZON et al. 2012) and in
sister chromatid recombination to repair replication-induced double strand breaks
(MUNOZ-GALVAN et al. 2012; PARDO AND AGUILERA 2012). There is evidence that
XPF is recruited to the replisome even in undamaged DNA, suggesting an
intimate role in genome maintenance during DNA replication(GILLJAM et al.
2012). Despite this, mammalian XPF is not essential for viability (BROOKMAN et
al. 1996; TIAN et al. 2004), possibly because it overlaps with another structure-
specific endonuclease, Mus81. In chicken DT40 cells that lack Mus81, XPF is
essential for viability and inactivation leads to chromosome breakage and failure
at a late stage of HR (KIKUCHI et al. 2013). Mammalian XPF is also implicated in
telomere protection (ZHU et al. 2003; MUNOZ et al. 2005).
These observations are consistent with the phenotypes of S. pombe rad16
mutants, which are defective for gene conversion associated with mating type
switching and for repair of radiation-induced DNA damage (EGEL et al. 1984;
SCHMIDT et al. 1989; PRUDDEN et al. 2003). There is increased instability between
direct repeats, leading to increased gene conversion as well as deletions (OSMAN
et al. 2000). S. pombe Rad16 promotes recombination repair of broken
replication forks using ectopic donor sequences, in contrast to Mus81, which
promotes repair via sister chromatids (ROSEAULIN et al. 2008).
Evidence is mixed regarding XPF function in meiosis. In Drosophila, the
DmMei9
XPF
homologue functions as a Holliday junction resolvase during meiosis;

89

flies deficient in mei9 show reduced meiotic recombination and loss of viabile
progeny (YILDIZ et al. 2002). In C. elegans, XPF functions redundantly with other
structure-specific endonucleases MUS-81 and SLX-1 in resolution of crossovers,
and suppresses formation of abnormal structures (AGOSTINHO et al. 2013; O'NEIL
et al. 2013; SAITO et al. 2013). In humans and in budding yeast, the structure-
specific endonucleases SLX1/Slx1, MUS81/Mus81, and GEN1/Yen1, but not
XPF, are linked to crossover resolution of Holliday junctions (KALIRAMAN et al.
2001; FRICKE AND BRILL 2003; WYATT et al. 2013). Thus, Sc rad1∆ mutants show
no decrease in spore viability, leading to the conclusion that there is no general
meiotic function for ScRad1
XPF
(HIGGINS et al. 1983), although it is implicated in
resolving insertions within heteroduplex DNA (KIRKPATRICK et al. 2000; KEARNEY
et al. 2001)
In fission yeast, Mus81 functions as the primary Holliday Junction
resolvase during meiosis (BODDY et al. 2000; BODDY et al. 2001; SMITH et al.
2003), and there is no obvious GEN1/Yen1 orthologue (IP et al. 2008). Rad16 is
required for short-patch repair of C/C mismatches in meiotic recombination
intermediates, consistent with its role in NER (FLECK et al. 1999). Loss of rad16
reduces gene conversion at an ade6 hotspot that additionally contains unpaired
heteroduplex DNA (FARAH et al. 2005; FARAH et al. 2009). However, a detailed
analysis of its function in typical meiotic recombination has not been carried out.
In this study, we examine the effect of rad16 mutation on fission yeast
meiosis. We show that rad16 mutants have reduced spore viability accompanied
by chromosome mis-segregation and apparent chromosome fragmentation at

90

both meiotic divisions. While the gross dynamics of DNA double strand break
repair appear intact, there is a modest increase in the rate of meiotic inter-
homologue crossover (CO) exchange, accompanied by a reduction in events that
use the sister chromatid. These phenotypes are shared with rhp14∆ mutants that
disrupt the XPA loading factor, but not in rad13∆ mutants that disrupt the XPG
endonuclease. Importantly, rad16 mutants also cause genome instability during
vegetative growth, accompanied by chronic activation of the DNA damage
checkpoint in the absence of external stress. Synthetic interactions with DNA
repair and replication mutants suggest that Rad16 contributes to genome stability
even in an unperturbed cell cycle.
3.3 Results
3.3.1 rad16 mutation reduces spore viability and perturbs meiotic
chromosome segregation.
The rad16-249 allele was originally identified in a screen for mutants
sensitive to alkylation damage (DOLAN et al. 2010). This allele carries a
truncation of the 877 amino acid protein at residue 118, and eliminates all
conserved domains. The rad16-249 allele is recessive, and behaves identically
to a disruption allele that removes aa313-798 (Figure 3.1).

91

Figure 3.1: Long Term Viability in Presence of Drug. Representative image.
Equal concentrations of cells plated in 5x serial dilutions from left to right. Drug
plates supplemented with indicated concentration of drug + PhloxinB.

In a cross between rad16-249 strains, we observed relative spore viability
dropped to 59% compared to wild type (assayed by random spore analysis;
Figure 3.2A). This is approximately four-fold higher viability than observed for
rec12∆ cells, which lack the meiosis-specific endonuclease Rec12
Spo11
and
cannot generate meiosis-specific DSBs (FARAH et al. 2005). Relative viability of
the double mutant rec12∆ rad16-249 is not significantly different from that of
rec12∆ (11.35% +/- 4.27 and 14.19% +/- 9.06, respectively). This suggests that
Rad16 operates in a Rec12-dependent pathway.

92

Figure 3.2: Spore Viability and Chromosome Segregation (continued on
next page).

93

Figure 3.2: Spore Viability and Chromosome Segregation (continued). (A)
Bulk spore germination of homozygous h+/h- meiosis from homologue

94

recombination data. Error bars represent standard error of the mean. At least 9
trials for each genotype for a total of 24600 and 38600 spores plated for wild type
and rad16-249 respectively, and 156000 for rec12∆ and rad16-249 rec12∆. (B
and C). Quantification of MI and MII segregations defects respectively. –Taz
indicates no Taz1-GFP signal on histone body +Taz1 indicates at least one
Taz1-GFP signal associated with histone body. 2x MII category refers to a single
spore encapsulating the both daughter nuclei of an MII division. (D)
Representative images selected from live cell analysis of meiosis in homozygous
h+/h- meiosis for wild type, rad16-249, rhp14∆, rad13∆, mus81∆, and rec12∆
homozygous h+/h- meiosis. Still image frames are taken from live cell movies at
indicated times relative to first image panel in series labeled 0min. White box
indicates portion of image at higher magnification in bottom row showing
fragmentation in rad16-249. White arrows indicate fragments. White dots outline
cell wall and spore walls. Magenta is signal from H3-mRFP and yellow from
Taz1-GFP. Scale bar represents 15µm. Live cell analysis was performed on at
least 30 cells from 3 different biological replicates (WT: MI=46 MII=47, rad16-
249: MI=69 MII=71, rad13∆: MI=68 MII=69, rhp14∆: MI=31 MII=36, rec12∆:
MI=81 MII=77, mus81∆: MI=50 MII=44).

Spore viability determined using tetrad dissetion was greater than that of a
random spore analysis, likely because normal-appearing four spored asci are
preferentially selected in tetrad analysis (Table 3.1). However, even with this
bias, only 39% of rad16-249 asci from a homozygous cross had four viable

95

germinating spores, and some had no viable spores at all, indicating an important
role for normal meiotic progression.
Table 3.1. Tetrad analysis of recombination between His4 and Lys4.

Viable
Spores/Tetrad WT rad16-249
0.00 0.40% 5.66%
1.00 4.37% 4.31%
2.00 15.48% 15.90%
3.00 7.94% 35.04%
4.00 71.83% 39.08%
cM 7.18 11.73
Relative Viability 100.00% 85.90%
Ratios of Colony Types 4 Spore Tetrads

his+lys- 224.00 207.00
his+lys- 231.00 205.00
his-lys- 28.00 38.00
his+lys+ 28.00 36.00

We examined the dynamics of meiosis in live rad16-249 cells in a cross
between h
+
and h
-
parents compared to wild type (Figure 3.2B-D, Movie 3.1 and
3.2). We visualized two fluorescent markers: mRFP-labeled histone H3 to label
the chromatin, and the telomere associated Taz1-GFP to identify any defects in
telomere clustering and bouquet formation, which can lead to disruptions in
meiosis and recombination (COOPER et al. 1997; COOPER et al. 1998; CHIKASHIGE
AND HIRAOKA 2001; TOMITA AND COOPER 2007). We saw no obvious

96

abnormalities in telomere clustering or in the characteristic horse-tailing
movement in the rad16-249 mutant, suggesting that telomere organization is
normal.
However, there are striking defects apparent during both MI and MII
divisions, in which fragments of H3-mRFP separate from the bulk of nuclear
signal (29% in MI and 37% in MII; Figure 3.2B-D). In some cases, these
fragments were enclosed as extra-nuclear spots within the spores (included), but
at other times, they remained outside of the spore wall (excluded, 50% in MI and
61% in MII) (Figure 3.2B,C). Approximately 15% MI and 46% MII fragments
contained one or two Taz1-GFP signals, as would be expected if they contain full
length chromosomes (Figure 3.2B,C). The absence of Taz1 in the remainder
suggests they result from some form of chromosome breakage.
3.3.2 Defects in Meiotic Chromosome segregation observed in other DNA
repair mutants
The fragmentation phenotype is visually similar to the abnormal
segregation and >4 nuclear spots that we reported previously for rad54∆ rdh54∆
double mutants, which are completely deficient in DSB repair. However, in that
case all the spores were inviable, consistent with a catastrophic failure of DNA
repair (CATLETT AND FORSBURG 2003). We investigated the formation of
fragments in other mutants.
During NER, the XPF endonuclease is recruited to the DNA by the XPA
protein, encoded by rhp14
+
in fission yeast (HOHL et al. 2001; CROTEAU et al.

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2008). Similar to rad16-249, we observed fragments in MI and MII divisions in
rhp14∆ (Figure 3.2B-D and Movie 3.3), suggesting that XPA and XPF also act
together during meiosis. In contrast, we observed no disruptions in meiosis in
rad13∆ mutants, which lack the downstream NER endonuclease XPG (Figure
3.2B-D and Movie 3.4).
We compared this to the meiotic phenotype of mus81∆ cells, which lack
the Holliday junction resolvase and cannot resolve chiasmata (Figure 3.2B-D,
Movie 3.5 and 3.6) (BODDY et al. 2000; BODDY et al. 2001; SMITH et al. 2003).
Previous immunofluorescence analysis of fixed mus81∆ cells showed evidence
for entangled chromosomes and dramatic disruption of divisions (BODDY et al.
2001; GAILLARD et al. 2003; OSMAN et al. 2003b). Consistent with this, we
observe extensively disordered MI divisions in live mus81∆ cells, with a failure of
nuclear division in 52% of cells. By following live cells through the time course,
we were able to observe them as they entered MII based on timing, regardless of
the MI outcome. In 27% of the cells in MII, we observed no nuclear division. In
those that did divide, segregation was highly unequal with multiple defects. We
observed fragments, which we defined as extra spots of histone-mRFP apart
from the main body of the nuclei. These were observed in 18% of MI and 32% of
MII divisions. About half of MI fragments and nearly all of the MII fragments
contained a Taz1 signal, indicating the presence of telomeres.
Finally, we examined rec12∆ mutants, which fail to create meiosis-specific
DSBs. In most cells, both meiotic divisions occurred with irregularities,
generating additional histone signals in 38% of cells (Figure 3.2B-D and Movie

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3.7). As expected, these extra spots appeared to be intact chromosomes , as
they always contained at least one Taz1 signal, with the majority containing two
or four Taz1 signals (Figure 3.2B,C). About half of these fragments were not
encapsulated into spores. We also observed a background level of dyad asci
(39%), likely diploids as reported previously (DAVIS AND SMITH 2003). In about
73% of these dyads we observed encapsulation of two distinct histone signals
inside a single spore. Only 23% of dyads resulted from an apparent failure to
undergo an MII division, where as the remainder underwent an MII division but
both products were incorporated into a single spore.
3.3.3 Meiotic repair dynamics in rad16-249
We reasoned that the defect in meiotic progression in rad16-249 reflects
defects in repair of programmed meiotic DSBs defining a function that is
important, but not essential. We therefore examined the formation of
programmed DSBs using pulsed field gel electrophoresis. In these experiments,
programmed breaks are typically observed as a smear below the three
chromosome pairs (YOUNG et al. 2002; CATLETT AND FORSBURG 2003). We used
temperature shift to induce a synchronous meiosis in h-/mat2-102 pat1/pat1
diploids, which maintain ploidy and mating type signaling, but allow temperature-
dependent synchronous meiotic progression (KOHLI et al. 1977; YAMAMOTO AND
HIRAOKA 2003; PANKRATZ AND FORSBURG 2005).
Consistent with previous studies, we observed the induction of meiotic
DSBs in wild-type cells beginning 3 hours after temperature shift with the majority

99

being repaired by 4 hours (Figure 3.3A). To quantify the amount of DNA in the
breaks, we determined the signal intensity in the DSB smear relative the total
signal observed in the whole chromosomes that enter the gel, indicated by the
boxes in lane 0 (BORDE et al. 2000). In the characteristic wild-type pattern, there
is a transient reduction of the signal corresponding to whole chromosomes,
which is caused by DSBs and also by chromosomes with unresolved
recombination intermediates that are retained in the well. Repair of the DSBs and
resolution of recombination leads to restored migration of the intact
chromosomes by 5 hours.

100

Figure 3.3: Meiotic DSBs in Diploids. Data from representative experiment
selected from 3 independent trials. (A) Representative image of three pulse field
gel experiments of synchronous mat2-102 pat1-114 diploid meiosis indicating
chromosome I, II, III, and DSBs for 0,1,2,3,4,5,6 hours. (B) Quantification of gel
in (A) showing the ratio of DSBs/total Chromosome signal for each time point
once the local background has been subtracted.

101

Figure 3.3: Meiotic DSBs in Diploids (continued). (C) Representative panels
of live cell imaging for RPA-CFP and Rad52-YFP. Scale bar is 15 microns. (D)
Quantification of RPA-CFP and Rad52-YFP focus persistence in meiotic cells.
Each bar is a single cell. Dark bars indicate cells in which there was an
abnormal segregation event and light bars represent apparently normal meiotic
progression.

102

The overall pattern in rad16-249 cells is roughly similar to wild type. The
bulk of the population shows restored chromosome entry, although the
DSB:whole chromosome ratio remains slightly elevated even at 6 hours (Figure
3.3A,B). The DSBs we see in rad16-249 depend upon Rec12 (Figure 3.3A), and
the timing of meiS phase, MI and MII divisions in rad16-249 are similar to wild
type (Figure 3.4), suggesting this is a post-Rec12 effect. This result contrasts
with PFGE performed on with repair-defective mutants such as mus81∆, rad54∆
or rad54∆ rdh54∆ , in which the DSB smear persists throughout the entire time
course, consistent with their catastrophic failure to repair or resolve the broken
DNA (CATLETT AND FORSBURG 2003; YOUNG et al. 2004).

103

Figure 3.4: Timing of Synchronous Meiotic Events. (A) Nuclear counts
visualized with DAPI to determine times of MI (2 signals) and MII (3+ signals)
divisions. (B) FACS analysis for samples used in A, B, and C showing the timing
and completion of meiotic replication as DNA content moves from 2C to 4C.
As an independent assay to see whether damage persists in rad16-249
cells, we examined the formation and resolution of DNA damage markers during
meiosis in wild type and rad16-249 diploids. Cells with DNA damage show

104

increased numbers of foci of fluorescently tagged RPA and Rad52 , so that these
markers provide a metric for unresolved DSBs (e.g., (LISBY et al. 2004;
SABATINOS et al. 2012). In wild type cells the RPA and Rad52 signals were
resolved by the MI division in the majority of cells (71% and 67% respectively).
In rad16-249, we observed resolution in only 22% of cells. Instead, the majority
of the rad16-249 cells (77%) had RPA and Rad52 signals that persisted through
nuclear divisions and as far as spore formation (Figure 3.3D). The presence of
fragments during meiotic divisions largely correlated with the persistence of both
RPA and Rad52 signals, suggesting failure to resolve recombination structures
leads to the abnormal segregation.
3.3.4 rad16-249 alters recombination frequencies
We investigated recombination outcomes in rad16-249 by examining three
different intergenic regions; his4
+
- lys4
+
and his7
+
– leu1
+
on chromosome II,
and ura1
+
- leu2
+
on chromosome I. Surprisingly, we observed a modest but
statistically significant increase in crossovers among the surviving spores in
rad16-249 when compared to wild type in all intervals tested (Figure 3.5A and
Table 3.2). The wild-type genetic distance was 9.05 cM, 2.02 cM, and 4.94 cM
for the his4-lys4, leu2-ura2, and his7-leu1 intervals respectively, while rad16-249
was 11.03 cM (p-value = 0.0024), 6.79 cM (p-value = 0.027), and 9.36 cM (p-
value = 0.018). The increased recombination in the his4 –lys4 interval was also
verified by tetrad dissections. Again, there was a statistically significant
difference (Χ
2
= 5.898 , alpha between 0.01 to 0.02) between WT ( 7.18 cM) and
rad16-249 (11.73 cM ; Table 3.1). Importantly, we saw no evidence for dyad

105

formation or diploid offspring that might affect these ratios. We observed no
striking difference in gene conversion events (3:1 or 1:3 segregation ratios) in
rad16-249 (3%) compared to wild type (1%, similar to other reports (RUDOLPH et
al. 1999)).
Figure 3.5: Recombination of Intergenic and Intragenic Intervals and spore

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viability. Significance established by two-tailed t-test. (A) Recombination
frequencies for homologue intergenic recombination. At least 4 trials were done
for each interval for a total of at least 5000 spores analyzed. p-values are
*0.0023, **0.027, and ***0.018 respectively for rad16-249 compared to WT for
each interval. (B) Diagram indicating types of repair between the sister and the
homologue using the ade-his-ade allele. (C) p-values for rad16-249 and rdh54∆
compared to WT are *0.028 and **0.016 respectively. (D) Spore viability
determined via random spore analysis. (E) Absolute viability assayed via tetrads.
Error bars represented as standard error. * X2=6.01, alpha=0.02 for swi5∆ rad16-
249 double mutant compared to swi5∆ single mutant.
Although a modest increase in crossovers was apparent in several genetic
intervals, we did not see the same effects when we measured gene conversion
at a single locus induced by the ade6-M26 hotspot allele (GUTZ 1971; GOLDMAN
AND SMALLETS 1979; PONTICELLI et al. 1988). We observed a modest decrease
in the recovery of Ade+ recombinants in rad16-249 relative to wild type (Table
3.2).
Table 3.2: Recombination and spore viability between his4-239 and lys4-
95, and ade6.
Strains Genotype
Total
germinated
Spores
Plated
Mean
Viability
Relative
to WT
Average
cM
His4
Lys4
Average
cM
Leu2
Ura2
Average
cM
His7
Leu1
Average
%ade+
1251 x
5107 WT 8940 24600
100.00
% 9.07 __ __ 0.40%
5192 x
5205
rad16-
249 7158 38600 58.87% 11.03 __ __ 0.19%
5268 x
rec12∆ 1041 15600 14.19% 0 __ __ 0

107

4561
5176 x
5180
rec12∆
rad16-
249 782 15600 11.35% 0 0
6917 x
6923 WT 19314 __ __ __ 1.84 __ __
6915 x
6924
rad16-
249 11409 __ __ __ 5.24 __ __
168 x
6919 WT 8011 __ __ __ __ 4.75 __
4707 x
6921
rad16-
249 5307 __ __ __ __ 7.73 __
Typically, meiotic DSBs can be repaired using either the homologous
chromosome, or the sister chromatid, a distinction referred to as “partner choice”.
Only recombination with the homologous chromosome has the potential to
generate a genetic crossover (CO) event, although these chiasmata can also be
resolved as non-crossovers (NCO). With the increased rate of homologous CO
events in rad16-249, we reasoned that the balance of repair between the
homologous chromosome and the sister chromatid might be disrupted. To test
this, we employed a diploid in which one copy of chromosome 3 contains a
double point mutant ade6-M375-M210, while the other copy contains tandem
heteroallelic ade6-L469/pUC8/his3+/ade6-M375 (as described in (CATLETT AND
FORSBURG 2003; PANKRATZ AND FORSBURG 2005). In this configuration, an ade6+
allele can typically only be recovered via inter-sister or intra-sister exchange
(because M210 and M469 are only 2bp apart; (SZANKASI et al. 1988) and R.
MacFarlane and W. Wahls, pers. comm.), so this can be used as a rough metric
for intra-chromatid or intra-sister events (Figure 3.5B). We found that rad16-249
has a 1.9 fold decrease in sister exchanges, measured by the recovery of Ade+

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spore clones (Figure 3.5C). We examined the types of SCE events recovered
by scoring the presence of the his3
+
marker. In mitotic cells, conversion events
that keep his3
+
represent short tracts of recombination between the repeats,
while deletion events are thought to result from SSA, non-conservative one-sided
invasion, replication slippage, intra-chromatid crossing-over, or unequal sister
chromatid crossing-over (OSMAN et al. 2000). There was a modest increase in
the proportion of conversion types (Ade
+
His
+
) in rad16-249 cells compared to
wild type (ELLERMEIER et al. 2004), although this was not statistically significant
(Table 3.3, Figure 3.5C).
Table 3.3: Recombination and spore viability of ade6 heteroallele.
WT rad16-249 rdh54∆
radh54 rad16-
249
Total spores counted 1399 1863 2270 1754
Total ade+ colonies recovered 104 41 478 256
STDEV 12.43 11.54 5.92 5.18
STError 6.21 5.77 2.96 2.59
Total plated 7000 22000 20000 20000
Relative Viability to WT 100.00 42.37 56.79 43.88
Average %ade6+ .48 .23 1.58 1.0
Fold ∆ 1.9 3.5 2.2
p-value Ade+ 0.028 0.016 0.068
Average % his+ ade+ 13.07 64.32 18.06 18.24
Fold ∆ 4.9 1.4 1.4
p-value His+Ade+ 0.067 0.163 0.210

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Previous studies have examined the effect of rad16 in haploids containing
a similar ade6-his+-ade6 allele and have observed a modest increase in
recombination in rad16 cells compared to wild type during vegetative growth
(OSMAN et al. 2003a). Since we see the opposite effect in meiosis than was
observed in mitosis, we infer that this is unlikely to represent rearrangements
during mitotic expansion of the spore clones. To test this, we repeated the
experiment, plating the spores on low adenine media where Ade+ colonies are
white and Ade- colonies are pink (Table 3.8). We reasoned that rearrangements
that occur during meiosis should generate a clonally pure Ade+ colony, which
should appear completely white, while rearrangements that occur during mitotic
expansion should generate a colony with white and pink sectors. We observed
sectors in approximately 10% of wild type Ade+ colonies, and close to 50% of
rad16 Ade+ colonies. The frequency of non-sectored Ade+ colonies (generated
by rearrangement during meiosis) is just under 0.5% in wild type, and
approximately 0.17% in rad16, reflecting the trend we observed in the larger
experiment. Further, consistent with the report of (OSMAN et al. 2003a), the
frequency of sectored Ade+ colonies reflecting rearrangements during mitosis
was higher in rad16 (0.11%) than wild type (0.06%).
In a previous study, we showed that the meiosis-specific homologous
recombination mutant rdh54∆ causes an increase in sister exchanges using this
assay, with no mitotic phenotype observed (CATLETT AND FORSBURG 2003). Since
this is the opposite of rad16-249, we constructed a double mutant and found that
rad16-249 reduces the frequency of intra-homologue exchanges in rdh54∆,

110

thought it still remains elevated over wild type. Spore viability is not changed,
with both single mutants and the double mutant each showing 50% spore viability
compared to wild type (Figure 3.5D).
Interhomologue events, both CO and NCO, depend primarily on the Swi5
/Sfr1 complex to mediate Rad51 filament formation, particularly at hotspots
(AKAMATSU et al. 2003; HYPPA AND SMITH 2010). Rad55 and Rad57 play a minor
role in both interhomologue and intrasister exchanges at DSB-poor regions,
while Rad52/Rti1 are implicated primarily in intra-sister events (AKAMATSU et al.
2003; OCTOBRE et al. 2008; HYPPA AND SMITH 2010). We constructed double
mutants to investigate whether rad16-249 could be placed genetically in either of
these pathways. We performed tetrad analysis and found that spore viability in
swi5∆ rad16-249 mutants was significantly reduced compared to either single
mutant, while rad57∆ rad16-249 did not show a significant change (Figure 3.5E).
This synthetic phenotype suggests that Rad16 functions in a pathway separate
from the Swi5-mediated interhomologue events, consistent with a role in
intersister exchanges.
The choice between NCO and CO resolution for resolution of chiasmata in
the homologous chromsomes depends upon the Fml1 (FANCM) translocase,
which limits CO in favor of NCO (Lorenz et al. 2012). Thus, fml1∆ mutants also
show evidence of increased homologous exchange. We constructed a double
mutant between fml1∆ and rad16-249 and performed tetrad analysis. We see a
modest decrease in spore viability in fml1∆ that is unchanged in the fml1∆ rad16-
249 double mutant (Figure 3.5E).

111

3.3.5 rad16-249 has genetic interactions with other DNA damage repair
mutants
In Drosophila, the XPF orthologue Mei9 functions as a Holliday junction
resolvase in meiosis (YILDIZ et al. 2002). In fission yeast, the primary meiotic
resolvase is Mus81 (BODDY et al. 2001; DOE et al. 2002; GAILLARD et al. 2003;
SMITH et al. 2003; GASKELL et al. 2007), and in its absence, cells are unable to
complete meiosis (BODDY et al. 2001; OSMAN et al. 2003b; SMITH et al. 2003) and
Figure 3.3). We were unable to construct a double mutant between rad16-249
and mus81∆ (Table 3.4); this synthetic lethality suggests that lesions produced in
mus81∆ mutants in vegetative growth absolutely depend upon Rad16 for
resolution and vice versa. This is consistent with data suggesting that Rad16 and
Mus81 overlap to maintain genome stability in metazoans (MAZON et al. 2012;
MUNOZ-GALVAN et al. 2012; KIKUCHI et al. 2013; SAITO et al. 2013). In vegetative
fission yeast cells, Rad16 functions as a template specific resolvase during repair
of replication forks, along with Mus81 (ROSEAULIN et al. 2008).

112

Table 3.4: Genetic interactions: phenotype of double mutants with rad16-
249
Viability UV sensitivity MMS sensitivity Comments
rhp51∆ Lethal - -
mus81∆ Lethal - -
rad3∆ Lethal - -
rqh1∆ Lethal - -
pcn1-K164R Lethal - -
rhp18∆ Lethal - -
rad2∆ Viable Enhanced Enhanced c
mms2∆ Viable Enhanced Enhanced b c
ubc13∆ Viable Enhanced Enhanced b c
eso1 Viable Enhanced Like rad16-249 c
kpa1∆ Viable Enhanced Enhanced
apn2∆ Viable Enhanced Enhanced b c
nth1∆ Viable Enhanced Enhanced c
saw1∆ Viable Like rad16-249
a
Like rad16-249
a

slx4∆ Viable Like rad16-249
a
Like rad16-249
a

a One parent is not noticeably sensitive to drug.
b Colony size smaller than either parent indicating poor growth.
c Colonies darker pink than either single mutant on PhloxinB indicating poor health.

The Mus81 endonuclease is essential for viability in rqh1∆ mutants, which
lack the RecQ helicase that restrains recombination in mitotic cells (DOE et al.

113

2002). If Rad16 and Mus81 overlap, we reasoned that rad16-249 rqh1∆ should
also be lethal, and this was observed (Table 3.4). Similarly, rad16-249 and
rad51∆ double mutants are synthetically lethal. These data indicate that Rad16
plays an important role to preserve genome stability even in unperturbed cells.
Next, we investigated the spectrum of damage sensitivity associated with
mutation of Rad16
XPF
, Rhp14
XPA
, or Rad13
XPG
. Consistent with their role in NER,
we observed similar sensitivity to UV and MMS in rad16-249, rhp14∆, and
rad13∆ (Figure 3.6A). We also observed sensitivity to CPT in rad16-249 and
rhp14∆ mutants, but not in the rad13∆ mutant. This is consistent with previous
work in S. cerevisiae that identified a role for Rad1
XPF
in resolution of
topoisomerase-bound intermediates caused by CPT treatment (VANCE AND
WILSON 2002). Finally, we observed sensitivity to hydroxyurea (HU) in rad16-249
and rhp14∆ mutants, but again not in rad13∆. HU causes fork stalling due to
nucleotide depletion, and restart occurs via recombination-based mechanisms
(MEISTER et al. 2007; LAMBERT et al. 2010; SABATINOS et al. 2012).

114

Figure 3.6: Check Point Activation and DNA Damaging Drug Sensitivity. (A)
Representative image of long-term viability and sensitivity. Equal concentrations
of exponentially growing cells in YES plated in 5x serial dilutions from left to right.
* Indicates rad16-249 mutation in a background comparable to the background of
rhp14∆ mutant while no * is rad16-249 mutation in background comparable to
rad13∆. (B) Representative image of long-term viability and sensitivity. Equal

115

concentrations of exponentially growing cells in YES plated in 5x serial dilutions
from left to right. (C) Cell length distribution of exponentially gorwing cells in
YES. (D) Western blot of Chk1-HA using 12CA5 anti-HA antibody with and
without treatment of 0.01% MMS. * indicates non specific bands, > is Chk1-HA
specific band, ¬ indicates modified Chk1-HA band.
Slx4 and Saw1 are proposed to function as scaffolds that assemble XPF
and other structure specific endonucleases (LYNDAKER AND ALANI 2009;
KASHIYAMA et al. 2013; LI et al. 2013; WAN et al. 2013). In contrast to budding
yeast (FLOTT et al. 2007; LI et al. 2008), neither slx4∆ nor saw1∆ are sensitive to
UV or MMS in fission yeast (Figure 3.7, and (COULON et al. 2006). We observed
no synthetic growth defects in double mutants between rad16-249 and either
slx4∆ or saw1∆, and little if any effect on MMS or UV sensitivity compared to
rad16-249 alone (Table 3.4 and Figure 3.7).

116

Figure 3.7: Growth Rates and Drug Sensitivity for rad16-249 Double

117

Mutants. Representative images of cells were plated in 5x serial dilutions from
equal starting concentrations on YES plates containing PhloxinB and noted drug
concentrations.

Finally, we examined interactions with components of post-replication
repair pathways. We found that rad16-249 is synthetically lethal with rhp18∆ or
pcn1-K164R (Table 3.4), which affect both error-prone (translesion synthesis)
and error-free post-replication repair pathways (FRAMPTON et al. 2006). We
found synthetically lethal interactions similar to that of rad16-249 in double
mutants with rhp14∆, but not rad13∆.
When rad16-249 is combined with error-free PRR pathway mutants
mms2∆ and ubc13∆ (BROWN et al. 2002), we do not observe synthetic lethality,
but rather increased sensitivity to UV and MMS (Table 3.4 and Figure 3.7). When
rad16-249 is combined with kpa1∆ (error prone DNA polymerase kappa; (KAI AND
WANG 2003)) we also observe increased sensitivity to damage caused by UV
and MMS. There was an enhancement of UV sensitivity when rad16-249 was
combined with and eso1 allele that deletes the polymerase eta domain, but no
change in MMS sensitivity. There were modest synthetic growth defects and
increased damage sensitivity in double mutants between rad16-249 and other
repair mutants including the flap endonuclease rad2∆ (YONEMASU et al. 1997),
the base excision repair mutant nth1∆ (OSMAN et al. 2003a), the base excision
repair mutant apn2∆ (FRASER et al. 2003), all consistent with linked repair
functions.

118

3.3.6 The DNA damage checkpoint is constitutively active in rad16-249
The rad16-249 mutants have an elongated cell morphology, which is
typically evidence of checkpoint activation from intrinsic DNA damage (Figure
3.6C and Table 3.5; rev. in (GOMEZ AND FORSBURG 2004). We examined nuclear
morphology during the vegetative cell cycle using RFP-histone, and observed
fragmented or lagging histone signals in both rad16-249 and rhp14∆, although
less frequently than in meiosis (9.62% and 8.25% of cells for rad16-249 and
rhp14∆ respectively; Table 3.6 and Movie 3.8). This, along with the double
mutant analysis, suggests Rad16 is required for chromosome stability even in the
absence of external perturbations. However, despite this, rad16-249 cells
maintain a high level of viability, with plating efficiency of 93.5% (SD +/- 7%)
relative to wild type.
Table 3.5: Distribution of cell length measurements binned.
WT rad16-249 chk1∆ rad16-
249
5 - 9.99 µm 60 16 69
10 – 14 µm 38 39 27
> 14 µm 2 45 4
Average 9.52 14.08 9.22
N 102 127 100

119

Table 3.6: Analysis of H3-mRFP Taz1-GFP mitotic live cell movies
WT rad16-249 rhp14∆
counted % counted % Counted %
Total scored 132 156 97
Normal 131 99.24 141 90.36 88 90.72
Included fragment w/ 1 Taz1
signal
0 0.00 5 3.21 1 1.03
Excluded fragment w/out Taz1
signal
0 0.00 1 0.64 3 3.09
Included fragment w/ 2 Taz1
signals
0 0.00 6 3.85 0 0.00
Anaphase bridging 1 0.76 3 1.92 4 4.12
Total abnormal 1 0.76 15 9.62 8 8.25

We constructed double mutants between rad16-249 and cds1∆, which
disrupts the intra-S phase checkpoint (LINDSAY et al. 1998; RHIND AND RUSSELL
2000); chk1∆, required for the DNA damage checkpoint pathway (WALWORTH AND
BERNARDS 1996; RHIND AND RUSSELL 2000); and rad3∆, the ATR homologue at
the apex of both pathways that is required for other damage responses as well
(BENTLEY et al. 1996; EDWARDS et al. 1999; RHIND AND RUSSELL 2000; DU et al.
2003; ROZENZHAK et al. 2010). We observed no additional growth defect in the
rad16-249 cds1∆ double mutants compared to either single mutant (Figure 3.6B).
In contrast, the rad3∆ rad16-249 double mutant is synthetically lethal (Table 3.4),
again consistent with chronic DNA damage caused by rad16-249.
The rad16-249 chk1∆ double mutants were viable, but slow-growing, with
reduced cell size. These cells also showed heightened sensitivity to UV and
MMS sensitivity (Figure 3.6B). Chk1 is activated by phosphorylation which
causes a characteristic mobility shift on SDS-PAGE(WALWORTH AND BERNARDS
1996). We observed a slower-migrating Chk1-HA in rad16-249 asynchronously
growing cultures in both the presence and absence of MMS , consistent with

120

intrinsic damage (Figure 3.6D). Similar results were observed for rhp14∆
(Figure 3.8). We did not see Chk1 activation in rad13∆, consistent with
previously reported results (HERRERO et al. 2006).

Figure 3.8: Western blot of Chk1-HA using 16B12 anti-HA antibody.
Lanes (+) 2,4,6,8,10 from cultures exposed to 0.01%MMS for 4 hours; lanes (-)
1,3,5,7,9 from cultures not exposed to drug. * indicates non specific bands, > is
Chk1-HA specific band, ¬ indicates modified Chk1-HA band.

To determine whether Chk1 activation in rad16-249 reflects increased
DNA damage, we examined live vegetative cells containing fluorescently-tagged
damage markers. We observed single stranded DNA by RPA-CFP and repair
foci marked by Rad52-YFP (e.g. (LISBY et al. 2004; SABATINOS et al. 2012)). In
wild-type cells, we found 28% contained a single RPA focus and 25% contained
a single Rad52 focus (Figure 3.9, e.g,(SABATINOS et al. 2012)). However, we
saw an increased number of cells with a single RPA (42%) and Rad52 (41%)
focus in rad16-249. Similar results were observed for rhp14∆, but not rad13∆.

121

Figure 3.9: Visualization of DNA Damage via Rad52 and RPA foci. (A)
Representative images of Rad52-YFP and RPA-CFP foci in exponentially
growing cultures in YES media for designated genotypes. Number of nuclei
analyzed for wild type, rad13∆, rad16-249, rhp14∆, rhp14∆ rad16-249 is 1816,
689, 1131, 854, and 843 respectively. See Table 3.7 for standard error and
confidence interval. (B) Distribution of nuclei containing 1,2, or 3+ Rad52-YFP
and/or RPA-CFP foci.

122

Table 3.7: Analysis of Rad52 and RPA.
Percent Nuclei with Rad11
foci
Standard Error 95% Confidence
Interval
1 2 3+ Any 1 2 3+ Any 1 2 3+ Any
WT 28 3 0 31 1.04 0.39 0.15 1.08 2.04 0.76 0.30 2.11
rad13∆ 26 4 0 31 1.02 0.48 0.09 1.07 2.00 0.94 0.17 2.10
rad16-249 42 14 4 60 1.14 0.81 0.46 1.13 2.24 1.58 0.91 2.22
rhp14∆ 44 15 2 61 1.15 0.84 0.29 1.13 2.26 1.64 0.58 2.21
rhp14∆
rad16-249
48 10 3 60 1.16 0.69 0.39 1.13 2.27 1.35 0.76 2.22
Percent Nuclei with Rad52
foci
Standard Error 95% Confidence
Interval
1 2 3+ Any 1 2 3+ Any 1 2 3+ Any
WT 25 2 0 28 1.01 0.34 0.11 1.04 1.98 0.67 0.21 2.04
rad13∆ 30 5 0 36 1.07 0.52 0.12 1.83 2.09 1.02 0.24 3.58
rad16-249 41 11 2 54 1.14 0.74 0.30 1.48 2.24 1.44 0.58 2.09
rhp14∆ 42 19 9 71 1.14 0.92 0.67 1.55 2.24 1.80 1.32 3.04
rhp14∆
rad16-249
43 12 5 61 1.15 0.76 0.52 1.68 2.25 1.49 1.02 3.29

3.4 Discussion
XPF is a conserved, structure-specific endonuclease with multiple
functions in genome maintenance (SCHWARTZ AND HEYER 2011). Originally linked
to nucleotide excision repair, XPF, its binding partner ERCC1, and its loading
factor XPA are also implicated in repair of ICLs, in SSA, homologous
recombination, and telomere maintenance (BOGLIOLO et al. 2013; KASHIYAMA et
al. 2013). XPF appears to be particularly important to trim unpaired ssDNA at the
boundaries of limited homology domains, and cleaves the 3’ end of non-
homologous flaps (PAQUES AND HABER 1997; HOLLINGSWORTH AND BRILL 2004;
FAGBEMI et al. 2011; SCHWARTZ AND HEYER 2011; MAZON et al. 2012). The role of
XPF in meiosis varies considerably in different species. In Drosophila, Mei9
XPF
is
essential for resolution of meiotic chiasmata (YILDIZ et al. 2002), while in C.

123

elegans, XPF overlaps with two other nucleases, Mus81 and SLX-1 in meiosis
(AGOSTINHO et al. 2013; O'NEIL et al. 2013; SAITO et al. 2013). In budding yeast,
Rad1
XPF
appears to have no function in meiosis (HIGGINS et al. 1983). Another
structure-specific endonuclease, Yen1, functionally overlaps with ScMus81 and
ScRad1
XPF
in budding yeast mitosis (BLANCO et al. 2010; MUNOZ-GALVAN et al.
2012), and functions in meiosis as to resolve late COs (MATOS et al. 2011), but
there is no obvious Yen1 orthologue in fission yeast (IP et al. 2008).
3.4.1 S. pombe Rad16
XPF
is required for normal meiosis
We investigated the role of the S. pombe XPF nuclease Rad16 in meiosis,
using a newly-characterized truncation allele that eliminates all the conserved
domains of the protein. We find that rad16-249 mutants undergo meiosis with
normal timing, including formation and repair of Rec12-dependent DSBs, but
nevertheless show a reduction in spore viability to about half of wild type levels.
Loss of viability was also reported in a previous study, using a different allele
(LORENZ et al. 2012). In tetrad dissection, only about one third of four-spored
tetrads are capable of germinating all four spores (4:0 viable), while the
remainder range from 3:1 to 0:4 viability. This indicates a role in meiosis that is
important, but not absolutely essential.
Using live-cell imaging, we observed that rad16-249 mutants suffer
chromosome segregation abnormalities during both MI and MII (Figure 3.2).
These are apparent as extra spots of histone-RFP or DAPI, which are smaller
and less bright than a full nucleus. Many, but not all of these apparent fragments

124

lack the telomere marker Taz1, suggesting they are chromosome fragments,
rather than full chromosomes. They often are left outside of the spore wall,
consistent with being disconnected chromosome fragments that are not attached
to a kinetochore. We see similar fragmentation phenotypes for rhp14∆
XPA
, but not
for rad13∆
XPG
. This agrees with data showing that XPA and XPF have functions
independent of XPG and other NER proteins (PAQUES AND HABER 1999;
LYNDAKER AND ALANI 2009), and implicates XPA and XPF in a distinct meiotic
function.
The cells that produce fragments are more likely to display persistent RPA
and Rad52 foci during meiotic divisions whereas these signals are generally
resolved prior to divisions in wild type cells, or cells without fragments (Figure
3.3). This suggests that the fragmentation phenotype is associated with a failure
to properly resolve DNA damage, either due to intrinsic stress or defects in
resolution of a subset of recombination structures. The progression through
meiotic divisions despite the presence of damage signals is consistent with
previous observations suggesting that the damage checkpoint is not triggered in
meiosis (PANKRATZ AND FORSBURG 2005).
We compared the rad16 phenotype to rec12∆ mutants (Figure 3.2), which
do not create programmed double strand breaks (SHARIF et al. 2002). In contrast
to rad16, the rec12∆ fragments always contain at least one Taz1 signal,
consistent with aberrant segregation of intact chromosomes. We infer therefore
that Taz1-minus chromosome fragments in rad16 result from unrepaired breaks,
or from damage that occurs due to aberrant segregation of unresolved

125

recombination intermediates. This is consistent with our previous report of
chromosome fragments during meiosis in rad54∆ rdh54∆ double mutants, which
are completely deficient in meiotic DSB repair and produce no viable spores
(CATLETT AND FORSBURG 2003). We examined the phenotype of mus81∆ cells,
which lack the main Holliday junction resolvase in fission yeast and fail to
segregate their chromosomes due to unresolved entanglements (BODDY et al.
2000; BODDY et al. 2001; OSMAN et al. 2003b; SMITH et al. 2003). Using live cell
analysis, we confirmed that the majority of Mus81 cells fail to undergo
chromosome segregation particularly in MI, with only a small percentage showing
evidence for Taz1-minus chromosome fragments. Thus, the rad16-249
phenotype is clearly distinct from that of mutants that fail to form DSBs (rec12∆,
which segregates whole chromosomes) or fail to resolve crossovers (mus81∆,
which remains largely entangled), and more closely resembles the phenotype of
rad54∆ rdh54∆, which is deficient in repair, although the phenotype in rad16 is
much less penetrant.
3.4.2 The majority of breaks are repaired in rad16 mutants
We used a PFGE assay to examine DSB formation and resolution more
closely (Figure 3.3). During DSB formation and resolution, the three
chromosomes show reduced migration; replaced by a smear of low molecular
weight DNA represents breakage, and joint molecules that are retained in the
well. We observe that the rad16-249 mutants have roughly normal timing for the
formation and disappearance of the majority of programmed double strand
breaks and repair, as measured by PFGE. However, at later time points, there is

126

a modest but persistent background of low molecular weight DSB signal (Figure
3.3). By comparison, in mus81∆, failure to resolve joint molecules reduces the
whole chromosome signal and strikingly increases in the smear of DSBs
throughout the entire timecourse (YOUNG et al. 2004). Similar observations of
reduced whole chromsomes and persistant DSBs were made in rad54∆ rdh54∆
which is also completely deficient in DSB repair (e.g. (CATLETT AND FORSBURG
2003)). We conclude that most DSBs are repaired in rad16∆, but a subset
remains.
One possibility is that rad16-249 simply has more DSBs due to intrinsic
genome instability in this strain (see below). If this were the case, there would be
a fraction of DSBs occurring that are independent of Rec12. Mutations that
cause breaks due to genome instability can partially rescue the viability defect
associated with rec12∆ mutants (FARAH et al. 2005; PANKRATZ AND FORSBURG
2005), but we see no evidence of that in rec12∆ rad16 double mutants.
Additionally, we observed no DSB smear in rad16-249 rec12∆ indicating that the
breaks observed are Rec12-dependent.
3.4.3 rad16 increases crossovers between homologous chromosomes
We observe a statistically significant increase in the rate of COs between
homologous chromosomes in several genetic intervals in rad16 compared to wild
type (Figure 3.5). This indicates that rad16 cells are competent for some form of
DSB repair, consistent with the PFGE result. An increased rate could represent
a shift in the balance between crossover and non-crossover resolution of

127

chiasmata between homologous chromosomes. Alternatively, or in addition, it
could indicate a shift in partner choice from the sister chromatid to the
homologous chromosomes.
In contrast to the interhomologue recombination data, we observe different
results for gene conversion involving the ade6-M26 hotspot, in which rad16-249
reduces the frequency of Ade
+
spores recovered. A similar modest reduction at
the ade6 hotspot was also observed by (Lorenz et al. 2012). Biased conversion
and marker effects at the ade6 hotspot have linked to defects in mismatch repair
(SCHUCHERT AND KOHLI 1988; SZANKASI AND SMITH 1995; FLECK et al. 1999),
which remains a possible explanation for the results with the ade6 hotspot.
Previously, rad16 was reported to reduce gene conversion at an ade6 hotspot
that also contained unpaired heteroduplex DNA (FARAH et al. 2005; FARAH et al.
2009); this would be consistent XPF function at unpaired flaps (SCHWARTZ AND
HEYER 2011). Why rad16 has different effects at normal intervals than at the
hotspot is unclear.
To examine whether the increase in homologous CO reflects a change in
partner choice, we investigated of sister-chromatid events using a substrate in
which only intra- or inter-chromatid events can give an Ade+ colony (CATLETT AND
FORSBURG 2003; PANKRATZ AND FORSBURG 2005). We observed reduced
frequency of Ade+ colonies in rad16-249 mutants compared to wild type (Figure
3.5). Although this reduction in recombination might reflect the role of Rad16 in
processing heterologous flap structures, and thus be an artifact of the tandem
allele construct we used, we consider this unlikely. In vegetative haploids,

128

mutation of rad16 actually increases recovery of Ade
+
at the ade6-
L469/pUC8/his3+/ade6-M375 locus (OSMAN et al. 2003a); therefore there is no
intrinsic impediment to resolution in the absence of rad16. In agreement with
this, we observe an increase in Ade+ sectoring during mitotic growth in rad16
haploids compared to wild type (Table S8). We suggest that the rad16 mutant is
impaired in sister chromatid recombination during meiosis.
Table 3.8: Mitotic recombination events in heteroallele spore germination.
Total Spores Ade+ Sectored
% of Total
Sectored
% of Ade+
Sectored
WT 4175 20 2 0.048 10
rad16-249 1625 4 2 0.12 50

Previously, we observed several situations in which recombination using
this sister construct was increased rather than decreased. In rdh54∆ mutants, a
modest increase in use of the sister is accompanied by a modest decrease in
recombination with the homologue, consistent with a role in partner choice
(CATLETT AND FORSBURG 2003). We observe that the rad16-249 mutant partly
reverses the rdh54∆ effect without rescuing its spore viability (Figure 3.5). This
also suggests that rad16-249 is deficient in the resolution of sister recombination
events. Increased recovery of Ade+ offspring is also seen in DNA checkpoint
mutants, which we inferred is due to sister-mediated repair of genome instability,
similar to that occurring in vegetative cells (PANKRATZ AND FORSBURG 2005).
However, in that case, this leads to Rec12-independent DNA damage and

129

rescue of rec12∆ viability. Despite the genome instability associated with rad16,
we see no evidence for rad16- induced DNA damage during meiosis.
A substantial fraction of the joint molecules in fission yeast are formed
between sister chromatids, not between homologues (CROMIE AND SMITH 2008),
and this is particularly true for areas of efficient DSB formation (HYPPA AND SMITH
2010). The Swi5/Sfr1 recombination mediator appears to be particularly
important for interhomologue exchanges in regions of DSB hotspots, while
Rad22/Rti1 are proposed to function at intersister exchanges (AKAMATSU et al.
2007; HYPPA AND SMITH 2010). The Rad55/Rad57 mediator, which is distinct
from Swi5/Sfr1, may be more important for exchanges in cold regions, affecting
both interhomologue and intersister events (KHASANOV et al. 1999; AKAMATSU et
al. 2003; KHASANOV et al. 2008; HYPPA AND SMITH 2010). We find that rad16-249
swi5∆ double mutants have reduced spore viability compared to the single
mutants, which suggests that Rad16 operates in a pathway that is separate from
Swi5. We see little additive effect in rad16-249 rad57∆ double mutants. Thus we
suggest that Rad16 may play a role in resolution of events mediated by
Rad55/57, particularly those involving the sister.
Consistent with this, mutations that reduce sister chromatid
exchanges in C. elegans without affecting crossovers between homologues
generate DNA fragments (BICKEL et al. 2010), leading to the suggestion that
homologue –independent recombination is important to preserve genome
stability in meiosis. We conclude that events involving the sister chromatid,

130

rather than the homologous chromosome, are similarly important for meiotic
genome stability in fission yeast.
3.4.5 Genome instability in vegetative rad16 cells
Rad16 is clearly important for genome stability even in an otherwise
unperturbed vegetative fission yeast cells. The rad16-249 mutants suffer
disordered segregation, with increased damage foci from markers Rad52-YFP
and RPA-CFP, and constitutive activation of the DNA damage checkpoint kinase
Chk1. Rad16 contributes to replication fork recovery in response to different
replication stress conditions (ROSEAULIN et al. 2008; MUNOZ-GALVAN et al. 2012).
This implies that replication stresses are intrinsic to normal cell cycle
progression, and require Rad16 for effective management. The synthetic
lethality of rad16-249 rad3∆ indicates that in addition to Chk1 activation, other
repair activities initiated by Rad3 are also important for rad16-249 viability. These
could include histone H2A(X) phosphorylation, upstream checkpoint proteins, or
chromatin effectors (EDWARDS et al. 1999; DU et al. 2003; ROZENZHAK et al.
2010).
We observe CPT and HU sensitivity in both rhp14∆ and rad16-249
mutants. This contrasts with budding yeast in which Rad14
XPA
, the XPF loading
factor, is not required for CPT resistance (VANCE AND WILSON 2002). Instead, Sc
Slx4 and Saw1 are proposed to provide an alternative XPF loading complex, and
Slx4 with Slx1 forms another structure specific endonuclease (LYNDAKER AND
ALANI 2009; SCHWARTZ AND HEYER 2011). In fission yeast, Slx4 -Slx1

131

endonuclease is linked to rDNA maintenance via a Rad51-independent
recombination mechanism (COULON et al. 2006), but so far there is no evidence
that Slx4 and Saw1 affect Rad16
XPF
in S.pombe. Significantly we and others
observe no damage sensitivity in slx4∆ or saw1∆ mutants (Figure 3.7, and
(COULON et al. 2006),) and no change in nucleolar morphology in rad16-249
(data not shown) suggesting they operate independently.
The instability of rad16-249 mutants even in an unperturbed mitotic cell
cycle suggests that its role in recombination may be an important component to
normal genome maintenance. A potential collaborator may be PCNA, the
replication clamp that ensures processive replication. Ubiquitylation of PCNA on
K164 by SpRhp18 (Sc Rad18) is required for post replication repair. Poly-
ubiquitylation is required for error-free PRR (FRAMPTON et al. 2006). We
observed synthetic lethality between rad16-249 and rhp18∆ or pcn1-K164R, but
not with genes that affect its polyubiquitylation, which suggests that PCNA mono-
ubiquitylation is essential for viability in the absence of Rad16. Interestingly,
several studies implicate PCNA modification in maintenance of repeat stability
(MOTEGI et al. 2006; DAEE et al. 2007; PUTNAM et al. 2010) and in Exo-1 mediated
resection (CHEN et al. 2013). The XPF loading factor XPA binds PCNA and XPA
and XPF are found as constituents of the replication fork in unperturbed cells
(GILLJAM et al. 2012). Indeed, the decreased stability of the ade6 heteroallele in
mitosis may reflect a rad16-related instability of the replication fork in repetitive
sequences.

132

DNA replication is intrinsically a source of DNA damage (rev. in (LEHMANN
AND FUCHS 2006). Structure-specific endonucleases such as Mus81 and XPF
actively contribute to recombination events that rescue damaged replication forks
or other structures, thus promoting genome stability (e.g., (ROSEAULIN et al. 2008;
WILLIS AND RHIND 2009; MUNOZ-GALVAN et al. 2012). The synthetic lethality we
observe between rad16-249 and mus81∆ is consistent with data in metazoans
that argues for redundancy between these two enzymes, with deficiency leading
to increased double strand DNA breaks (KIKUCHI et al. 2013). Yet these same
proteins contribute actively to gross chromosome rearrangements including
translocations, which are typical of cancer (MAZON et al. 2012; PARDO AND
AGUILERA 2012).
The choice of helpful or harmful pathway may reflect access to repetitive
sequences that facilitate SSA forms of repair. Typically, regions closest to the
DSB are used preferentially for intrachromatid repair by the SSA pathway (RAY et
al. 1988; NICKOLOFF et al. 1989; SUGAWARA AND HABER 1992; FRANKENBERG-
SCHWAGER et al. 2009), and evidence suggests that the extent of resection in
meiosis influences choice of repair pathways (NEALE et al. 2002). There is likely
to be a closely-regulated interplay between resection, helicase-driven resolution
of recombination structures, and the activity of structure specific endonucleases
that determines the outcome of these events.

133

3.5 Materials and Methods
3.5.1 Cell growth and culture: General culture conditions and media are
described in (SABATINOS AND FORSBURG 2010). Strains used listed in Table 3.9.
Cells were grown from single colonies in 5ml cultures at 32˚C overnight to mid-
log phase for RPA and Rad52 focus imaging and serial dilution plating assays to
determine drug sensitivities. Drug plates were incubated at 32˚C for 2-4days
before being imaged using flatbed scanner. For imaging, cells were concentrated
at 6000rpm in microfuge and spread on PMG agar on glass slide for imaging
(SABATINOS et al. 2012). Heterothallic strains were grown independently for
meiotic movies in PMG with appropriate supplements as 32˚C until culture was in
late log-phase (OD595~0.8). Cells were pelleted and washed in EMM-N and
resuspended in ME and incubated 12-20hrs in a 25˚C airshaker. Cells were
concentrated using microfuge and spread on SPAS agar pads on glass slides.
Imaging was performed at 25˚C.
Table 3.9: Strains used in this study.
Strains
6 h- leu1-32 ade6-704 ura4-294 Our Stock
118 h90 ura4-D18 leu1-32 ade6-M216 Our Stock
168 h+ ade6-704 leu1-32 Our Stock
416 h- ade6-704 leu1-32 ura4-D18 rad13::ura4 Our Stock
421 h- ade6-704 leu1-32 ura4-D18 ∆chk1::ura4 Tony Carr
527 h- his3-D1 ade6-M216 ura4-D18 leu1-32 Our Stock
528 h+ his3-D1 ade6-M210 ura4-D18 leu1-32 Our Stock
865 h- ∆cds1::ura4 ura4-D18 leu1-32 Tony Carr
915 h- leu1-32 ade6-M210 Our Stock

134

941 h- ∆rad2::ura4+ leu1-32 ade6-704 ura4-D18 Our Stock
1107 h- ∆rad3::ura4+ ura4-D18 leu1-32 ade6-M216 Our Stock
1251 h+ ade6-M26 his4-239 Gerry Smith
1256 h- mad2∆::ura4+ ade6-M210 leu1-32 ura4-D18 Our Stock
1893 h- ade6-M375-M210 leu1-32 ura4-D18 his3-D1
(Catlett and
Forsburg 2003)
1898
h- rdh54∆::ura4+ ade6-L469/pUC8/his3+/ade6-M375 ura4-D18 leu1-
32
(Catlett and
Forsburg 2003)
1902 h+ ade6-L469/pUC8/his3+/ade6-M375 ura4-D18 leu1-32 his3-D1
(Catlett and
Forsburg 2003)
1942 h+ rdh54∆::ura4+ ade6-M375-M210 leu1-32 ura4-D18 his3-D1
(Catlett and
Forsburg 2003)
2057 h- pat1-114 ade6-M216 can1-1 Our Stock
2111 h- pat1-114 rec12∆::ura4+ ura4-D18 ade6-M216 Our Stock
2170 h90 mat2-102 pat1-114 rec12∆::ura4+ ura4-D18 ade6-M210 Our Stock
3490 h- ∆swi10::kanMX ura4-D18 leu1-32 ade6-704 Tony Carr
3500 h90 mat2-102 pat1-114 ade6-M210 Our Stock
3766 h- ∆swi5::his3+ ade6-M210 ura4-D18 leu1-32 his3-D1 Our Stock
3767 h+ ∆swi5::his3+ ade6-M210 ura4-D18 leu1-32 his3-D1 Our Stock
3769 h+ ∆rhp57::ura4+ ade6-M210 ura4-D18 leu1-32 his3-D1 Our Stock
3770 h- ∆rhp57::ura4+ smt0 ade6-M210 ura4-D18 leu1-32 his3-D1 Our Stock
3876 h- apn2::kanMX6 ura4-D18 leu1-32 his3-D1
Mathew
O’Connell
3877 h+ nth1::ura4 ura4-D18 leu1-32 his3-D1 arg3-D1
Mathew
O’Connell
3884 h- exo1::ura4 ura4-D18
Mathew
O’Connell
3887 h- rhp14::kanMX6 ade6-704 leu1-32 ura4-D18
Mathew
O’Connell
3958 h- rad35-271 ura4-D18 leu1-32 Our Stock
4415 h+ ∆reb1::kanMX ade6-M216 ura4-D18 leu1-32 Our Stock
4504 h+ rad16-249 ura4-D18 leu1-32 This Study
4505 h+rad16-249 his3-D1 ura4-D18 leu1-32 ade6-M210 =rad16 This Study

135

4561 h+ Delta-rec12::ura4+ ura4-D18 his4-239 ade6-M26 This Study
4562 h- rad16-249 ura4-D18 ade6-M210 This Study
4661 h- rad16-249 his3-D1 ura4-D18 leu1-32 ade6-M216 This Study
4707 h- rad16-249 leu1-32 ade6-M210 This Study
4707 h- rad16-249 leu1-32 ade6-M210 This Study
4708 h+ rad16-249 leu1-32 ade6-M210 This Study
4839 h90 Rad16-249 Hht2-GFP-ura4+ ura4-D18 leu1-32 ade6-M216 This Study
4941 h90 ura4-D18 rad16::ura4+ Henning Schmidt
4983 h+ ∆mms2::LEU2+ rad16-249 leu1-32? ura4-D18 ade6-52 This Study
4984 h+ ∆srs2::KanMX6 rad16-249 ura4-D18 ade6-M210 This Study
4985 h- ∆kpa1::bleMX6 ura4-D18 This Study
4986 h- rad16-249 rad35-271 ura4-D18 leu1-32 This Study
4987 h- rad16-249 ∆ubc13::ura4+ ura4-D18 ade6-52 This Study
5136 h- ∆swi10::kanMX rad16-249 ura4-D18 ade6-704 This Study
5146 h- eso1::kanMX6 rad16-249 ura4-D18 ade6- This Study
5147 h- ∆kpa1::bleoMX6 rad16-249 ura4-D18 ade6- This Study
5165 h- apn2::kanMX6 rad16-249 ura4-D18 leu1-32 his3-D1 This Study
5166 h- nth1::ura4+ ura4-D18 rad16-249 ade6-52 This Study
5172 h- rad16-249 exo1::ura4 ura4-D18 This Study
5176 h- ∆rec12::ura4+ ura4-D18 rad16-249 lys4-95 ade6-52 This Study
5180 h+ ∆rec12::ura4+ siw9-249 ura4-D18 his4-239 ade6-M26 This Study
5181 h- ∆rad2::ura4+ rad16-249 ura4-D18 leu1-32 ade6- This Study
5182 h+ ∆srs2::kanMX6 ura4-D18 ade6-M210 This Study
5186 h- ∆ubc13::ura4+ ura4-D18 ade6-M210 This Study
5191 h+ ∆mms2::leu2 ura4-D18 ade6-M210 This Study
5192 h+ rad16-249 his4-239 ade6-M26 This Study
5193 h- ∆slx4::kanMX4 his3-D1 leu1-32 ura4-D18 ade6-M216 This Study
5194 h+ rad16-249 ura4-D18 ade6-M210 This Study
5204 h- ∆swi10::kanMX ura4-D18 ade6-704 This Study

136

5205 h- rad16-249 lys4-95 ade6-52 This Study
5206 h- eso1::kanMX6 ura4-D18 ade6-704 This Study
5207 h- lys4-95 ade6-52 This Study
5208 h- rad16-249 ade6-M210 This Study
5221 h90 mat2-102 pat1-114 rad16-249 ade6-M216 This Study
5241 h- rad16-249 ∆chk1::ura4 ade6-704 leu1-32 ura4-D18 This Study
5243 h- rad16-249 ∆cds1::ura4 ura4-D18 leu1-32 This Study
5245 h- rad16-249 ∆slx4::kanMX4 his3-D1 leu1-32 ura4-D18 ade6-M210 This Study
5247 h- rad16-249 mad2∆::ura4+ ura4-D18 leu1-32 This Study
5257 h- ∆saw1::kanMX4 his3-D1 ura4-D18 leu1-32 ade6-M216
This Study-
Bioneer derived
5268 h- ∆rec12::ura4+ ura4-D18 ade6-52 lys4-95 This Study
5287
h- ∆saw1::kanMX4 rad16-249 his3-D1 ura4-D18 leu1-32 ade6-
M216/210?
This Study-
Bioneer derived
5497 h- pat1-114 rad16-249 Drec12::ura4+ ura4-D18 ade6-M216 This Study
5530 h+ rad16-249 ∆reb1::kanMX ade6-M210 leu1-32 ura4-D18 This Study
5580 h- rhp14::kanMX6 rad16-249 ura4-D18 leu1-32 This Study
5600
h90 mat2-102 pat1-114 rad16-249 Drec12::ura4+ ura4-D18 ade6-
M210
This Study
5800 h- rad16-249 pat1-114 ade6-M210 This Study
5809
h+ rad16-249 ∆rdh54::ura4+ ade6-M375-M210 leu1-32 ura4-D18 his3-
D1
This Study
5811 h+ rad16-249 ade6-M375-M210 leu1-32 ura4-D18 his3-D1 This Study
5814
h- rad16-249 ade6-L469/pUC8/his3+/ade6-M375 ura4-D18 leu1-32
his3-D1
This Study
5816
h- rad16-249 ∆rdh54::ura4+ ade6-L469/pUC8/his3+/ade6-M375 ura4-
D18 leu1-32 his3-D1
This Study
5825 h- rad16-249 ade6-704 leu1-32 ura4-D18 rad13::ura4 This Study
6257 h- ∆fml1::natMX4 ura4-D18 his3-D1 leu1-32 ade6-M216 Our Stock
6258 h+ ∆fml1::natMX4 ura4-D18 his3-D1 leu1-32 ade6-M216 Our Stock
6915 h- rad16-249 leu2-120 This Study
6917 h+ leu2-120 ade6-M210 This Study

137

6919 h- his7-36 ade6- This Study
6921 h- rad16-249 his7-36 ade6- This Study
6923 h- ura2-10 ade6- This Study
6924 h+ rad16-249 ura2-10 ade6- This Study
7376
h- ∆rhp57::ura4+ rad16-249 ade6-M210 ura4-D18 leu1-32 his3-
D1
This Study
7377
h+ ∆rhp57::ura4+ rad16-249 ade6-M210 ura4-D18 leu1-32 his3-
D1
This Study
7378
h- ∆swi5::his3+ rad16-249 ade6-M210 ura4-D18 leu1-32 his3-
D1
This Study
7379
h+ ∆swi5::his3+ rad16-249 ade6-M210 ura4-D18 leu1-32 his3-
D1
This Study
7467 h- rad16-249 ∆fml1::natMX4 ura4-D18 his3-D1 leu1-32 This Study
7468 h+ rad16-249 ∆fml1::natMX4 ura4-D18 his3-D1 leu1-32 This Study
7475 h- lys4∆::kanMX ura4-D18 leu1-32 ade6-
This Study-
Bioneer derived
7515 h- lys4∆::kanMX rad16-249 ura4-D18 leu1-32 ade6-
This Study-
Bioneer derived

3.5.2 Spore viability and Recombination:
Spore viability and recombination were performed by mating strains on
SPAS agar for 2-3days at which point the mating patch was scraped from plate
and diluted in 1ml 0.5% glusolase. This was digested for 16-20hours rotating at
room temperature. Spores were plated on YES media and grown at 32˚C for 3-
5days before counting and replica plating colonies onto PMG media with
appropriate supplements. Phloxin B was included to identify any diploids; for
rad16-249, no diploids or dyad asci were observed. We also assayed diploid
spore formation in rad16-249 and wildtype in recombination assays using a lys4
deletion marked with kanMX in combination with the his4-239 point mutation. In

138

this way when His+Lys+ colonies were recovered, any colonies which were also
diploid would be resistant to G418. We found no colonies that were His+Lys+
and G418 resistant. His+Lys+, Leu+Ura+, or His+Leu+ progeny were identified
and genetic distance was calculated by (2(His+Lys+)/total colonies)*100. Within
the same strains intragenic recombination was assessed using the ade6-M26
and ade6-52 alleles and scoring for the restoration of the Ade
+
phenotypes (GUTZ
1971; PONTICELLI et al. 1988). The experiment was repeated 9 times plating 2000
spores for each genotype each trial. For tetrad analysis genetic distance was
calculated as using Perkin’s formula as described (SMITH 2009). Sister
recombination was determined as in assay described previously (CATLETT AND
FORSBURG 2003). Significance was calculated for genetic distances using two
tailed t-test. To test mitotic recombination in the sister recombination assay,
sectored colonies were counted after germination of spores on YE and EMM low
ade media. For tetrad dissection spore viability, asci were dissected after mating
on SPAS media and germinated on YES for 3-5 days. Significance was
determined using the Chi Squared test.
3.5.3 Imaging:
Images were acquired with a DeltaVision Core widefield deconvolution
microscope (Applied Precision, Issaquah, WA) using an Olympus 60X/1.40,
PlanApo, NA=1.40 objective lens and a 12-bit Photometrics CoolSnap HQII CCD,
deep-cooled, Sony ICX-285 chip. The system x-y pixel size is 0.1092µm x-y.
softWoRx v4.1 (Applied Precision, Issaquah, WA) software was used at
acquisition electronic gain=1.0 and pixel binning 1x1. Excitation illumination was

139

from a Solid-state illuminator (7 color version), GFP was excited and detected
with a (ex)475/28,(em)525/50 filter set and a 0.2 second exposure, RFP was
excited and detected with a (ex)575/25, (em)632/60 filter set and a 0.2 second
exposure exposure, CFP was excited and detected with a (ex)438/24,(em)470/24
filter set and a 0.15 second exposure, and YFP was excited and detected with a
(ex)513/17,(em)559/38 filter set and a 0.15 second exposure for meiotic time
courses; for mitotic still imaging CFP was excited and detected with a
(ex)438/24,(em)470/24 filter set and a 0.5 second exposure, and YFP was
excited and detected with a (ex)513/17,(em)559/38 filter set and a 0.5 second
exposure. Suitable polychroic mirrors was used, GFP/mCherry Chroma ET
C125705 roughly: 520/50–630/80 and Semrock CFP/YFP/DsRed 61008 bs
roughly: 415/20–462/32–535/50–635/74. Thirteen z sections at 0.5µm were
acquired. 3-D stacks were deconvolved with manufacturer provided OTFs using
a constrained iterative algorithm and images were maximum intensity projected.
For live cell imaging time points were taking 10mins apart for the length of
experiment. Images were contrast adjusted using a histogram stretch with an
equivalent scale and gamma for comparability. Brightfield images were acquired
with DIC. Whole cell SytoxGreen flow cytometry (FACS) was performed as
described in (SABATINOS AND FORSBURG 2009).

3.5.4 Western Blot:
Western blot analysis of cell extracts was taken from cultures grown to
early mid-log phase (OD595~0.3) in YES media at 32˚C. Cultures were split in
equal volumes and treated with 0.01%MMS and untreated and grown for 4 hours

140

at 32˚C at which point 10XSTOP buffer was added. Extracts were prepared
using TCA (FOIANI et al. 1994). Protein extracts were quantified using pierce
BCA and 100ug protein was run on a 8% acrylamide w/ 1.25% crosslinker SDS
PAGE gel and probed with 16B12 α-HA (Covance) or 12CA5 α-HA (Abcam) at
1:1500 or 1:1000 respectively dilution in PBSt antibody.
3.5.5 Pulse Field Gel Electrophoresis:
Synchronous diploid meiosis was achieved using the mat2-102 and pat1-
114 alleles as in (CATLETT AND FORSBURG 2003) to create stable diploids using
ade6-M210/M216 complementation. Pulse field gel plugs were created by
digesting the cell wall with 0.2mg/ml 100T Zymolase and 0.45mg/ml Sigma
Lysing Enzymes titrated to 50% and 25% of original strength for time points 1-2
and 3-6 respectively as in (CERVANTES et al. 2000). Pulse field gel using Biorad
Chef II Pulse Field machine was run for 48hours using 2 V/cm, 1800 seconds
switch time, and a 106˚ angle. DNA was visualized via ethidium bromide. DSB
quantification was done by quantification using BioRad Quant One software
representing the DSB breaks as a ratio to total chromosome signal with the local
background subtracted as in (BORDE et al. 2000).
3.6 Legends for Movies:
Movie 3.1: Representative of live cell imaging of WT using H3-mRFP (Magenta)
and Taz1-GFP (cyan) to view meiotic progression.
Movie 3.2: Representative of live cell imaging of rad16-249 using H3-mRFP
(magenta) and Taz1-GFP (cyan) to view meiotic progression.

141

Movie 3.3: Representative of live cell imaging of rhp14Δ using H3-mRFP
(magenta) and Taz1-GFP (cyan) to view meiotic progression.
Movie 3.4: Representative of live cell imaging of rad13Δ using H3-mRFP
(magenta) and Taz1-GFP (cyan) to view meiotic progression.
Movie 3.5 and 3.6: Representative of live cell imaging of mus81Δ using H3-
mRFP (magenta) and Taz1-GFP (cyan) to view meiotic progression. Movie 5
shows mus81Δ that complete an MI division and Movie 6 shows MI division
failure.
Movie 3.7: Representative of live cell imaging of rec12Δ using H3-mRFP
(magenta) and Taz1-GFP (cyan) to view meiotic progression.
Movies 3.8: Representative of live cell imaging of rad16-249 using H3-mRFP
(red) and Taz1-GFP (green) to view mitosis.

142

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Chapter 4

Translesion synthesis polymerases contribute to meiotic chromosome
segregation in S. pombe

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4.1 Overview
In this study the role of translesion synthesis polymerases in
Schizosaccromyces pombe meiotic progression was analyzed. Previous studies in
budding yeast have implicated translesion synthesis polymerases in meiotic
recombination at the level of double strand break repair yet there is no striking loss
of meiotic viability. Through cell physiology, live cell imaging and molecular
techniques I showed that the absence of all four translesion synthesis polymearsaes
in the fission yeast lead to a drastic loss of meiotic viability and defects in
chromosome segregation, while meiotic double strand breaks and repair appear
unaffected. However, there is an apparent mis-regulation of the meiosis specific
cohesin Rec8 implicating translesion synthesis polymerases in the process of
meiotic cohesin establishment, activation, and/ or maintenance.

4.2 Introduction
Transmission of genetic information across generations requires faithful
replication and maintenance of the genome. This is accomplished through the use
of high fidelity polymerases. These polymerases catalyze DNA synthesis by an
error free mechanism involving 3’-5’ proof reading activities (PRAKASH et al. 2005;
RATTRAY AND STRATHERN 2005). However, efficient these replicative polymerases
are at synthesizing DNA under unchallenged conditions they are unable to replicate
through helix distorting lesions such as abasic sites, base dimmers, and bulky

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adducts (PRAKASH et al. 2005; RATTRAY AND STRATHERN 2005). In instances of these
lesions, a replicative polymerase will stall potentially leading to deleterious DSBs.
Translesion synthesis polymerases (TLSP) are error prone. Many have error
rates exceeding 10
-3
(GOODMAN 2002). Despite these large error rates in replication,
these polymerases are important for preserving genome stability. These TLSPs
have the unique property to replicate across DNA lesions with various error rates.
Thus DNA replication is allowed to continue avoiding a potential genomic
catastrophe due to blocked replication (WATERS et al. 2009). Alternatively, TLSPs
contribute to genomic stability by gap filling (HELLER AND MARIANS 2006). There are
now well over a dozen described TLSPs in human cells, several of which are
conserved in the budding and fission yeasts (PRAKASH et al. 2005; RATTRAY AND
STRATHERN 2005).
Fission yeast have four known and described TLSPs: Rev1, Pol ζ, Eso1, and
Pol κ. Eso1 is unique in S. pombe in that it is the fusion of what are two gene
products in other organisms, the essential cohesion acetyltransferase Eco1, and
Pol η, which may be post-translationally split (TANAKA et al. 2000; CHEN et al. 2014).
The advantage, if any, to this fusion is unclear.
Though TLSPs have the ability to bypass lesions, sometimes even quite
accurately, overall they are mutagenic. Increasing the expression of TLPs is causes
hypermutation (KIM et al. 1997; OGI et al. 1999; BERGOGLIO et al. 2002; BAVOUX et al.
2005). Thus regulation of TLSPs at the transcriptional level is important for genome
stability. In S. pombe gene expression of several TLSPs is increased in

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synchronous meiotic cells (MATA et al. 2002). This increase is even more profound
that that seen in many environmental stress situations such as H
2
O
2
(MATA et al.
2002; CHEN et al. 2003; KAWAMOTO et al. 2005). Transcriptional up-regulation in
meiotic cells is not limited to fission yeast. In humans, pol η has enriched
expression in the mouse testis, specifically in the spermatotids (MCDONALD et al.
1999). This transcriptional up-regulation may be indicative of a meiotic role for these
polymerases.
One attractive model for what TLSPs may be doing in meiosis implicates
them in meiotic double strand break repair. Two studies have demonstrated in vitro
that Pol η can perform D-loop extension during homologous recombination (HR) in
double strand break repair (DSB) (MCILWRAITH et al. 2005; LI et al. 2009). It has also
been shown in budding yeast that TLSPs contribute to mutagenesis during HR
during meiosis, potentially aiding in genetic diversification (ARBEL-EDEN et al. 2013).
Previously a connection between TLSPs and meiosis in S. pombe was shown
through the regulatory subunit of the DDK kinase, Dfp1 (LE et al. 2013), Dfp1 is
required for the error prone pathway of post replication repair (PRR) where the
TLSPs also function (LE et al. 2013). A truncation allele, dfp1-r35, is defective in
induced mutagenesis, indicating that it has a role in TLSP activity. This allele also
has striking defects in meiosis, including disruptions of replication, induction of
programmed double strand breaks, and chromosome segregation. In dfp1-r35

truncation mutants, phosphorylation and cleavage of meiotic cohesin is
compromised, leading to defects in the MII division (LE et al. 2013).

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In this study I investigated the role of TLSPs in S. pombe meiotic progression.
Here I describe a phenotype of chromosome mis-segregation due to the loss of all
four TLSPs in S. pombe. Further, I provide evidence that these TSLPs do not
substantially affect meiotic DSB repair and recombination, but suggest a role in
cohesion dynamics.

4.3 Results
4.3.1 Spore viability is reduced in TLS mutants
In order to determine to magnitude of a contribution to meiotic progression, if
any, of the TSLPs in S. pombe I first analyzed spore viability using random spore
analysis. The largest reduction in spore viability was in the quadruple TLS mutant
lacking all four proteins (Figure 4.1A). Each single TLS mutants showed a modest,
but statistically significant reduction in spore viability. eso1∆ (deletion of the pol η
homology region only) and rev1∆ showed similar reductions in viability compared to
wild-type of 64% and 61% respectively. kpa1∆ showed a very modest reduction of
81% while rev3∆ (encodes a subunit of Pol ζ) had a 58% viability. The quad∆
mutant had a reduction 16% viability. This is comparable to the 19% observed in
rec12∆, which fails to make meiosis specific DSBs and thus lacks all recombination
(Figure 4.1A and (SHARIF et al. 2002)).

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Figure 4.1: Spore and Mitotic Viability. (A) Sore viability as analyzed via random
spore analysis plating at least 10,000 spores with at least 6 biological replicates per
genotype. Error bars are standard error calculated for categorical data. (B) Mitotic
plating efficiency relative to wild type. At least 6 different biological replicates were
used and 9,000 cells plated per genotype. Error bars are standard error calculated
for categorical data. (C) Long term drug sensitivity assays. Cells plated in 5x serial
dilutions on minimal media containing the concentrations of the indicated drugs.

166

In contrast there was little to no reduction in mitotic viability assessed via
plating efficiency in either single or the quadruple mutant, which indicates that the
failure to observe spore colonies is not due to subsequent mitotic defects but
represents a failure in meiosis (Figure 4.1B). Each mutant has a characteristic
sensitivity to DNA damaging agents during vegetative growth (Figure 4.1C). It was
observed, as was previously reported, that eso1∆ shows enhanced sensitivity to
ultraviolet radiation (TANAKA et al. 2000). kpa1∆ shows enhanced sensitivity to the
alkylation due to methyl methanesulfonate (MMS), which is consistent with a
predicted role in nucleotide excision repair (OGI AND LEHMANN 2006). In contrast,
rev3∆ has a significant sensitivity to the topoisomerase inhibitor, Camptothecin,
which causes S phase specific breaks, and rev1∆ has sensitivity to the spindle
poison, Thiabendazole, Interestingly, the quad∆ shows a less severe phenotype
when challenged with these drugs. It appears that in some situations the lack of all
four of the polymerases is less deleterious than the absence of one.
4.3.2 Quadruple TLS mutant has normal meiotic recombination, DSB repair,
and progression
There is increased expression of TLSPs in meiosis but with differing
dynamics (MATA et al. 2002). Kpa1 and Eso1 are expressed early in the meiotic
program while Rev1 and Rev3 reach a maximum around MI/MII. I examined
whether the spore viability defect in the quad∆ reflects defects in recombinational
repair, which occurs early in the meiotic program. Further, in a mutant, rec12∆, that

167

completely lacks meiotic DSBs, there is a loss of spore viability to 19% that of wild
type similar to that of the quadruple deletion mutant (Figure 4.1A, and (SHARIF et al.
2002; PANKRATZ AND FORSBURG 2005)). In contrast, if meiotic DSBs are made, but
there is a catastrophic failure of DSB repair, spore viability drops to near 0 (BODDY et
al. 2001; CATLETT AND FORSBURG 2003; OSMAN et al. 2003; CROMIE et al. 2006;
LORENZ et al. 2012). In order to determine whether the viability loss in the quad∆
mutant represents a defect in meiotic recombination, the formation of DSBs was
analyzed via pulse field gel looking at whole chromosomes in a synchronous diploid
meiosis culture (PANKRATZ AND FORSBURG 2005; LE et al. 2013). In wild type cells,
meiotic DSBs, visualized as a smear below the intact chromosomes, occur between
3 and 4 hours and are resolved by 5 hours. This trend is apparent in the quad∆
(Figure 4.2A). Thus there appears to be no gross disruption in meiotic DSB
occurrence or repair dynamics in absence of all the TLSPs. Consistent with this, the
overall dynamics of premeiotic S phase and meiotic divisions were similar in both
wild type and quad∆ (Figure 4.2B, 4.2C).
Next, the resolution of these breaks was examined looking at interhomolog
recombination between two markers on chromosome II that are approximately 76kb
apart. In wild type cells there was a genetic distance of 8.79 cM. All the single
mutant TLSPs and quadruple mutant maintain similar levels of recombination as the
wild type with no statistically significant differences with the exception of
kpa1∆, which has a slight reduction in homolog recombination in this interval (Figure
4.2D). Thus, the defects causing failure of meiosis are not due to gross defects in
the program of meiotic recombination.

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Figure 4.2: Recombination and Synchronous Meiosis. (A) Representative
image of three biological replicate pulse field gel electrophoresis experiments
showing chromosome I, II, and III with lower molecular weight programmed meiotic
double strand breaks. (B) FACs data showing replication progression during meiotic

169

time course. Representative of at least three biological replicates. (C) DAPI staining
of meiotic progression of 1, 2, 3+ Dapi stained nuclear masses, representative of
three biological replicates. (D) Graph of meiotic recombination rates between his4-
239 and lys4-95 on chromosome II. Error bars are standard deviation. * is
significance of 2.79E-4 and ** 4.94E-12 using a two-tailed t-test.

4.3.3 Quadruple TLS mutant has disrupted chromosome segregation in
meiosis
Dynamics of meiotic progression were examined more closely using live cell
imaging. Visualizing histone tagged with mRFP, it was observed that the quad∆
showed the appearance of extranuclear histone bodies or fragments during MI and
MII divisions (10.47% and 13.25% of divisions respectively) (Figure 4.3A,B, and
Movie 4.2). This fragmentation or mis-segregation is rarely observed in wild type MI
and MII (4.85% and 3.88% of divisions) (Figure 4.3A,B, and Movie 4.1). There was
an increased incidence of uneven nuclear divisions, based on qualitative
segregation of H3-mRFP during an MI or MII division in the quad∆ (MI=10.47%,
MII=3.61%) compared to the wild type (MI=0.75%, MII=0.00%) (Figure 4.3A,B). This
suggests there are defects in overall chromosome segregation, which could
represent chromosome fragmentation, nondisjunction, or premature separation.

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Figure 4.3: Meiotic chromosome segregation. (A) Representative images
showing H3-mRFP fragments in quad∆ (top), uneven segregation in quad∆ (middle),

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and wild-type (bottom). Scale bars are 10 microns. (B) Quantification of meiotic
errors observed in live cell imaging. (C) Quantification of LacI-GFP segregation in
MI and MII. * Indicates significance using two-tailed t-test of 0.009.

In order to determine whether the mis-segregating histone bodies observed in
the LacI-GFP H3-mRFP experiments are whole chromosomes or fragments, I
examined segregation in a strain with histone H3-mRFP and the telomere protein
Taz1-GFP. I reasoned that whole chromosomes would always contain a telomere,
but a chromosome fragment might not. It was previously shown that mutants that
suffer from failures in DSB repair have mis-segregating histone bodies that lack
Taz1-GFP signals, while mutants with failures in whole chromosome separation,
such as rec12∆, have a Taz1-GFP signal on the mis-segregating histone body
(MASTRO AND FORSBURG 2014). I observed a difference in Taz1-GFP signal between
MI and MII mis-segregations in the quad∆ (Table 4.1). Only 55% of mis-segregated
histone bodies had a Taz1-GFP signal associated, suggesting that a fraction of
these are some sort of chromosome fragment, while all MII mis-segregating histone
bodies had at least one Taz1-GFP signal, suggesting that these segregations
involve a whole chromosome.

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Table 4.1: Taz1-GFP signal for quad∆ H3-mRFP mis-segregating bodies.
Type of H3-mRFP Body MI % MI MII %MII
Taz1-GFP-plus 11 55 20 100
Taz1-GFP-minus 9 45 0 0
Total 20 20

In order to look at the segregation of a specific chromosome, a strain
heterozygous for a derivative of chromosome I with a lacO array integrated proximal
to the centromere was examined. When present in a genetic background
expressing LacI-GFP, this allows examination of reductional (MI) vs equational (MII)
division. In MI division in wild type, the sister chromatids remain associated, so that
there is no segregation of the lacI-GFP focus. In the MII division, the sister
chromatids come apart, leading to separation of two lacI-GFP foci into adjacent
spores. The reductional and equational divisions are dependent on meiotic cohesin
Rec8, and its step-wise cleavage (YOKOBAYASHI et al. 2003). In a rec8∆ mutant, the
absence of proper cohesion at the centromere leads to an equational MI division and
premature sister chromatid separation (YOKOBAYASHI et al. 2003). Conversely, if
there is a failure of Rec8 cleavage so that the protein persists, the sisters will not
separate during MII, and this nondisjunction resembles a reductional division
(KITAJIMA et al. 2003a). The quad∆ strain showed a statistically significant increase
in nondisjunction or non-separation in MII, indicating a failure to segregate sister
chromatids properly during the equational division (Figure 4.3C).

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4.3.4. Rec8-GFP dynamics and recruitment are altered in quad∆
One explanation for the chromosome mis-segregation seen in the quad∆
mutant is that the meiosis specific cohesin, Rec8, could be mis-regulated. It was
previously shown that that DDK regulatory subunit Dfp1 contributes to the error
prone TLS pathway of PRR and also has a mis-regulation of Rec8 (DOLAN et al.
2010; LE et al. 2013). A disruption of Rec8 may account for the changes seen in
meiotic segregations. Therefore, live cell imaging of asynchronous diploid meiotic
cultures was used to assay the dynamics of Rec8-GFP.
Rec8 has very characteristic visual patterns as cells go through meiosis. As
was seen in previous studies, wild-type shows a pan-nuclear signal which is reduced
to two single puncta shortly following the MI division, indicating Rec8-GFP had been
released from the chromosome arms but maintained at the centromeres (Figure
4.4A, and Movie 4.3) (WATANABE AND NURSE 1999; LE et al. 2013). These puncta
disappear just prior to MII, indicating that Rec8-GFP had been fully removed from
the chromatin. These dynamics are dramatically altered in the quad∆. There was a
modest but significant delay in the disappearance of the pan nuclear signal in the
quad∆ compared to wild type (17 and 10mins respectively). Conversely, it was
observed that the quad∆ had a significantly shorter period for the Rec8-GFP focus to
go away (32 mins and 55 mins respectively) (Figure 4.4B and Movie 4.4). This
suggests there may be a delay in the removal of Rec8 from chromosome arms
during MI, but that it may be prematurely removed from the centromere during MII.

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Figure 4.4: Rec8 dynamics. (A) Live cell imaging of Rec8-GFP schematic of pan-
nuclear vs focus formation during meiosis. (B) Quantification of the Rec8-GFP live
cell imaging. * Indicates significance using two-tailed t-test of 2.95E-09 and **
7.91E-17. (C) Representative western blot of Rec8-GFP of synchronous diploid

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meiosis using pat1-114/ mat2-102. (D) Representative quantification of western
blot using GFP/Tubulin. (E) RT-PCR quantified using end point PCR GFP/Actin.
(F) ChIP of Rec8-GFP at dh or act1. Error bars are standard error.

Rec8 dynamics are regulated through phosphorylation. Phosphorylation by
the kinases DDK and CK1 is required for Rec8 cleavage at the centromere
(ISHIGURO et al. 2010; LE et al. 2013). In contrast, de-phosphorylation by Sgo1-
PP2A prevents Rec8 cleavage (ISHIGURO et al. 2010). Rec8-GFP phosphorylation
and overall protein levels through meiosis were assessed by using a pat1-114 –
driven synchronous meiosis in diploids (Figure 4.4D). Rec8-GFP was visible at
hours 2 through 5 with the slower migrating phosphorylated species present in wild
type at hours 3 through 5 (PARISI et al. 1999; ISHIGURO et al. 2010; RUMPF et al.
2010; LE et al. 2013). The phosphorylation as well as protein levels in the quad∆
were modestly reduced compared to wild type (Figure 4.4C and D). Thus, there was
an observable change in Rec8-GFP dynamics in meiosis as well as a reduction in
phosphorylation and protein levels of Rec8 in the quad∆.
One explanation for the reduction in protein levels is that the transcriptional
regulation of rec8 may be altered. Using qRT-PCR the rec8 transcript levels in wild-
type and quad∆ were shown not to be significantly different indicating the differences
in the levels of Rec8 protein are not at the transcript level (Figure 4.4E). An
alternative explanation for the reduced protein levels and phosphorylation of Rec8 is
that Rec8 is not effectively deposited on the chromatin in the absence of the TLSPs.

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It is possible that Rec8 phosphorylation and subsequent cleavage requires that jt is
bound to the chromatin. In order to address the levels of Rec8-GFP bound to the
chromatin I performed chromatin immunoprecipitation (ChIP) looking at the
centromere dh region and a distal euchromatic locus of act1. There was a
significant reduction in Rec8-GFP at the pericentromeric region, dh in the quad∆
compared to wild-type (Figure 4.4F). The effect is modest at early time points, but
dramatic at later time points. There was not a significant difference in the non-
centromeric locus, act1 (Figure 4.4F). Thus, the dynamics of Rec8-GFP at the
centromere measured molecularly resemble the dynamics observed visually:
premature removal of the Rec8-GFP cohesion from the centromere in the quad∆
mutant.
In order to determine whether the disruption of meiotic cohesin in the quad∆
is specific to meiotic specific cohesins, the dynamics of the mitotic cohesin, Rad21
tagged with GFP, was analyzed. In wild type, Rad21-GFP is visible in the
horsetailing phase of meiosis and disappears right before MI ((DING et al. 2006),
Figure 4.5 and Movie 4.5). The Rad21-GFP signal is localized to the leading edge
of the horsetail nucleus in the rDNA region ((DING et al. 2006), Figure 4.5 and Movie
4.5). There was no disruption in localization or timing detected in the quad∆
compared to wild type (Figure 4.5, Movie 4.5 and Movie 4.6).

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Figure 4.5: Rad21-GFP in meiosis. Representative images of live cell imaging of
Rad21-GFP and H3-mRFP showing horsetailing and MI division. Green is Rad21-
GFP and magenta is H3-mRFP. Scale bars are 10 microns.

4.4 Discussion
In this chapter, it has been shown that the TSLPs contribute to genome
stability in meiosis, with defects in chromosome segregation in MI and MII leading to
reduced spore viability. Spore viability of the quad∆ is dramatically reduced
compared to the single mutants (Figure 4.1A), suggesting there are overlapping
functions of the TLSPs in meiosis. In mitosis, this same trend is not observed, as

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the drug sensitivities to CPT and TBZ as well as the plating efficiency of the quad∆
are less severe than the single mutants (Figure 4.1B,C).
These data did not support a role of TLSPs in meiotic recombination or DSB
repair. Normal levels of DSBs and their repair were observed and no changes in
interhomolog recombination rates were apparent (Figure 4.2A,D). This does not
discount the possibility that the TSLPs could be involved in a minor way that was
undetectable by our methods. In budding yeast TSLPs have been implicated in
meiotic recombination and DSB repair by contributing to increased mutation rates
around DSB sites (ARBEL-EDEN et al. 2013). Mutation rate at DSB sites were not
specifically tested in this study so the possibility that TSLPs are involved in this in S.
pombe cannot be ruled out.
Defects in chromosome segregation during meiosis, as well as a disruption in
the dynamics associated with the meiotic cohesion Rec8 were apparent in this
study. Normally, Rec8 is removed from the chromosome arms but remains
protected at the centromere in MI, finally being phosphorylated and cleaved to allow
MII progression (LEE et al. 2005; NASMYTH AND HAERING 2005; WATANABE 2005;
WATANABE AND KITAJIMA 2005; KATIS et al. 2010). Specifically there was a reduction
of Rec8 meiosis specific cohesin loading and/or maintenance at the pericentromeric
region but no change at distal locations. Further the Rec8 that is loaded at the
centromere appears unstable; I do not observe significant phosphorylation yet it
turns over more rapidly especially following MI.

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One possibility is that the TLSPs act as a recruitment/stabilization factor for
the Rec8. It has been shown that Rec8 is loaded to some extent prior to meiotic
replication (WATANABE et al. 2001). Activation of cohesin occurs separately from
loading and requires the acetyltransferase Eco1 (KENNA AND SKIBBENS 2003;
SKIBBENS 2005). Eco1 has been shown to interact directly with the replicative clamp
(PCNA) and the clamp loader (RFC), implying a link between replication and cohesin
activation (KENNA AND SKIBBENS 2003; SKIBBENS 2005). Eco1 is required for the
acetylation of the cohesin subunit Psm3, which localizes at the centromere in
meiosis with Rec8 (KAGAMI et al. 2011). Without this acetylation, monopolar
kinetochore attachment is inhibited (KAGAMI et al. 2011). In S. pombe, eso1 is a
fusion of polη and the acetyltransferase homologue eco1, although the protein
product may be split post-translationaly (CHEN et al. 2014). Still, at the
transcriptional and translational levels these genes are co-regulated (CHEN et al.
2014), A further link between cohesion establishment and replication is in S.
cerevisiae where two essential genes, trf4 and trf5 that are required for mitotic sister
cohesin have been shown to interact with Pol ε (WANG Z 2000; EDWARDS et al.
2003). Thus the replication fork may act to enhance cohesin loading, activate
cohesin, or redistribute cohesin for enrichment in centromeric regions.
Replication fork mediated cohesin activation may be regulated through the
TLSPs in some way. It has been shown in S. cerevisiae that Pol η is required for
damage induced genome wide cohesion (ENERVALD et al. 2013). In that study there
was no requirement for PCNA nor any of the other TLSPs in S. cerevisiae (Rev1 or
Polζ) (ENERVALD et al. 2013).

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The premature loss of Rec8 in the TLSP mutant is accompanied by an
increase in nondisjunction in MII. Paradoxically, this is similar to the phenotype
observed in a non-cleavable form of Rec8, rather than a loss of the protein (KITAJIMA
et al. 2003b). Thus, the presenting defect does not appear to be premature loss of
centromere cohesion. An alternative interpretation of the MII nondisjunction could
be that the lack of Rec8 at the centromere disrupts proper bipolar spindle
attachment, as a residual amount of Rec8 at the centromere in MII is required for
bipolar attachment of the kinetochore (KITAJIMA et al. 2004; RABITSCH et al. 2004;
KITAJIMA et al. 2006; RIEDEL et al. 2006). If this is the explanation, then the apparent
nondisjunction is due to an error in spindle attachment rather than an error in
resolving cohesion. For example, destabilization of the centromere through mutation
of swi6∆ (required for pericentromeric heterochromatin) attenuates Rec8 loading,
resulting in a mono-oriented kinetochore attachment in MII, leading to chromosome
mis-segregation in MII (KITAJIMA et al. 2003b; YOKOBAYASHI et al. 2003; KITAJIMA et
al. 2006; KAWASHIMA et al. 2007). Interestingly, Swi6 is also required for the
recruitment of shugoshin and PP2A to the centromere through a direct interaction
(YAMAGISHI et al. 2008). Thus a disruption in centromeric structure could also cause
the lack of Rec8 phosphorylation observed in this study.
The disruption observed in Rec8 meiosis specific cohesin is not
observed for Rad21. Rather the Rad21 dynamics observed through live cell imaging
are normal. In rec8∆ there is an invasion of Rad21-GFP from the rDNA region into
the chromosome arms (DING et al. 2006). Even in the rec8∆ this effect is minor
(DING et al. 2006). It is then not surprising that there was not a detectible disruption

181

of Rad21-GFP in the quad∆ as Rec8 is largely retained in the quad∆ while its
presence at the centromere is compromised.
A third possibility that remains is that the TSLPs may be required for efficient
replication through the centromere and thus centromere stability in meiosis. In this
case, a failure to segregate sister chromatids in MII could represent entanglements
that link the chromosomes together due to unresolved replication intermediates or
repair structures. There were no gross meiotic replication problems in the quad∆
mutant identified in this study; however, a more detailed analysis of replication at the
centromere may be warranted.

4.5 Materials and Methods:
4.5.1 Cell growth and culture:
General culture conditions and media are described in (SABATINOS AND FORSBURG
2010). Drug plates were incubated at 32˚C for 2-4days before being imaged using
flatbed scanner. For imaging, cells were concentrated at 6000rpm in microfuge and
spread on PMG agar on glass slide for imaging (SABATINOS et al. 2012).
Heterothallic strains were grown independently for meiotic movies in PMG with
appropriate supplements as 32˚C until culture was in late log-phase (OD595~0.8).
Cells were pelleted and washed in EMM-N and resuspended in ME and incubated
12-20hrs in a 25˚C airshaker. Cells were concentrated using microfuge and spread
on SPAS agar pads on glass slides. Imaging was performed at 25˚C. Strains used
are listed in Table 4.1.

182

Table 4.2: Strains used in this study.
Strain Genotype Source
1251 h+ ade6-M26 his4-239 Gerry Smith
2057 h- pat1-114 ade6-M216 can1-1
(MASTRO AND
FORSBURG
2014)
3500 h90 mat2-102 pat1-114 ade6-M210
(MASTRO AND
FORSBURG
2014)
5207 h- lys4-95 ade6-52
(MASTRO AND
FORSBURG
2014)
5259 h- rev3::hphMX6 lys4-95 ade6-52 This Study
5262 h- eso1::kanMX6 lys4-95 ade6-52 This Study
5263 h+ rev3::hghMX6 his4-239 ade6-M26 This Study
5269 h+ eso1::kanMX6 his4-239 ade6-M26 This Study
5401 h+ ∆rev1::ura4+ ura4-D18 his4-239 ade6-M26 This Study
5466 h- ∆rev1::ura4+ ura4-D18 lys4-95 ade6-52 This Study
5608 h- hht1-mRFP:KanMX6 leu1-32 ura4-D18
(MASTRO AND
FORSBURG
2014)
5787
h+ hht1-mRFP:kanMX his7+::lacI-GFP lys1+::lacO
leu1-32 ura4-D18
(MASTRO AND
FORSBURG
2014)
6137 h+ rec8-GFP-kan(YW) ade6-M210 This Study
6138 h- rec8-GFP-kan(YW) ade6-M210 This Study
6332
h90 mat2-102 pat1-114 rec8-GFP-kan(YW) ade6-
M210
(MASTRO AND
FORSBURG
2014)

183

6336 h- pat1-114 rec8-GFP-kan(YW) ade6-M216
(MASTRO AND
FORSBURG
2014)
6664
h- his4-239 eso1::kanMX6 dinB::bleMX6
rev3::hphMX6 ∆rev1::ura4+ ura-D18 ade6- This Study
6671
h90 mat2-101 pat1-114 eso1::kanMX6
kpa1::bleMX6 rev3::hphMX6 ∆rev1::ura4+ ura-D18
leu1-32 ade6-M210 This Study
6703
h+ eso1::kanMX6 kpa1:bleMX6 rev3::hphMX6
∆rev1::ura4+ ura-D18 his4-239 ade6-m26? This Study
6716
h+ eso1::kanMX6 kpa1:bleMX6 rev3::hphMX6
∆rev1::ura4+ ura-D18 lys4-95 ade6-52 This Study
6717
h- eso1::kanMX6 kpa1:bleMX6 rev3::hphMX6
∆rev1::ura4+ ura-D18 lys4-95 ade6-52 This Study
7117
h- eso1::kanMX6 dinB::bleMX6 rev3::hphMX6
∆rev1::ura4+ his7+::lacI-GFP lys1+::lacO hht1-
mRFP:natMX6 ura4-D18 leu1-32 ade6- This Study
7167
h- pat1-114 eso1::kanMX6 kpa1::bleMX6
rev3::hphMX6 ∆rev1::ura4+ ura-D18 leu1-32 ade6-
M216 This Study
7168
h+ eso1::kanMX6 dinB::bleMX6 rev3::hphMX6
∆rev1::ura4+ hht1-mRFP:natMX6 ura4-D18 leu1-
32 ade6- This Study
7402
h- pat1-114 rec8-GFP::kanMX6 (WW)
eso1::kanMX6 kpa1::bleMX6 rev3::hphMX6
∆rev1::ura4+ ura-D18 ade6-M216 This Study
7501
h90 mat2-102 pat1-114 eso1::kanMX6
kpa1∆::bleMX6 rec8-gfp:kanMX rev3::hphMX6
∆rev1::ura4+ ura-D18 ade6-M210 This Study
7616 h+ kpa1:bleMX6 his4-239 ade6- This Study
7685 h- kpa1:bleMX6 lys4-95 ade6- This Study
7691
h- eso1::kanMX6 dinB::bleMX6 rev3::hphMX6
∆rev1::ura4+ taz1-GFP:KanMX6 hht1-
mRFP:KanMX6 ura4-D18 leu1-32 ade6- This Study
7692
h- eso1::kanMX6 dinB::bleMX6 rev3::hphMX6
This Study

184

∆rev1::ura4+ taz1-GFP:KanMX6 hht1-
mRFP:KanMX6 ura4-D18 leu1-32 ade6-

4.5.2 Viability and Recombination:
Spore viability and recombination were performed by mating strains on SPAS agar
for 2-3days at which point the mating patch was scraped from plate and diluted in
1ml 0.5% glusolase. This was digested for 16-hours rotating at room temperature.
Spores were plated on YES media and grown at 32˚C for 3-5days before counting
and replica plating colonies onto PMG media with appropriate supplements. Phloxin
B was included to identify any diploids; no diploids or dyad asci were observed in
TLSP mutants. His+Lys+ progeny were identified and genetic distance was
calculated by (2(His+Lys+)/total colonies)*100. Experiment was repeated at least 6
times plating a least 1000 spores for each genotype each trial. Significance was
calculated for genetic distances using two tailed t-test. Mitotic viability was assayed
via determining plating efficiency after cells were grown to an OD of ~0.6 in YES
media at 32˚C. Experiment was repeated at least 6 times for every genotype plating
at least 1000 cells per trial.
4.5.3 Imaging:
Images were acquired with a DeltaVision Core widefield deconvolution microscope
(Applied Precision, Issaquah, WA) using an Olympus 60X/1.40, PlanApo, NA=1.40
objective lens and a 12-bit Photometrics CoolSnap HQII CCD, deep-cooled, Sony
ICX-285 chip. The system x-y pixel size is 0.1092µm x-y. softWoRx v4.1 (Applied
Precision, Issaquah, WA) software was used at acquisition electronic gain=1.0 and

185

pixel binning 1x1. Excitation illumination was from a Solid-state illuminator (7 color
version), GFP was excited and detected with a (ex)475/28,(em)525/50 filter set and
a 0.2 second exposure; RFP was excited and detected with a
(ex)575/25,(em)632/60 filter set and a 0.2 second exposure exposure;. A suitable
polychroic mirror was used GFP/mCherry Chroma ET C125705 roughly: 520/50–
630/80. Thirteen z sections at 0.5µm were acquired. 3-D stacks were deconvolved
with manufacturer provided OTFs using a constrained iterative algorithm and images
were maximum intensity projected. Images were contrast adjusted using a
histogram stretch with an equivalent scale and gamma for comparability. Brightfield
images were acquired with DIC. Whole cell SytoxGreen flow cytometry (FACS) was
performed as described in (SABATINOS AND FORSBURG 2009).
4.5.4 Western Blot:
Cells were grown and synchronous meiosis was induced as in (CATLETT AND
FORSBURG 2003) Cell cultures were stopped by adding 10x STOP buffer containing
sodium azide solution to harvested culture and incubating the cultures on ice for 10
minutes. Cells were washed in PBS buffer (137 mM NaCl, 2.7 mM KCl, 4.3 mM
Na2HPO4,1.47 mM KH2PO4) and then MQ water. Extracts were prepared using
TCA (trichloroacetic acid; (FOIANI et al. 1994). Protein extracts were quantified using
pierce BCA and 100ug protein was run on a 8% acrylamide w/ 1.25% crosslinker
SDS PAGE gel and probed with 5% PBST milk containing a 1:2000 dilution of JL8
monoclonal antibodies (Clontech) overnight at 4˚C, and washed three times for 10
minutes in 10 ml PBST each time. For secondary, blots were probed with 5% PBST
milk solution with 1:3000 secondary goat anti-mouse HRP antibodies (Millipore).

186

Blots were developed using ECL (Pierce). Quantification was done using GelEval
Version 1.37 (1.37).
4.5.6 Pulse Field Gel Electrophoresis:
Synchronous diploid meiosis was achieved using the mat2-102 and pat1-114 alleles
as in (CATLETT AND FORSBURG 2003) to create stable diploids using ade6-
M210/M216 complementation. Pulse field gel plugs were created by digesting the
cell wall with 0.2mg/ml 100T Zymolase and 0.45mg/ml Sigma Lysing Enzymes
titrated to 50% and 25% of original strength for time points 1-2 and 3-6 respectively
as in (CERVANTES et al. 2000). Pulse field gel using Biorad Chef II Pulse Field
machine was run for 48hours using 2 V/cm, 1800 seconds switch time, and a 106˚
angle. DNA was visualized via ethidium bromide.
4.5.7 RT-PCR
RNA was extracted using QIAGEN’s RNAse Easy RNA extraction kit. Extracted
RNA was quantified on a spectrophotometer (Nanodrop, ND1000). RNA was
converted to cDNA using Roche kit. Relative transcription levels were determined by
performing multiplex PCR using target primer for GFP and actin primers. Actin
primer sequences from (KLOC et al. 2008) were used. Quantification was done using
GelEval Version 1.37 (1.37) taking a ratio of target to actin signal.
4.5.8 ChIP
Protocol modified from (LI et al. 2013). Strains were harvested and cross-linked for
15 min with 1% formaldehyde at room temperature with rotating. Quenching was

187

done with 0.25 m glycine for 5 min at room temperature. Cells were harvested by
centrifugation at 1500 rpm for 3 min at 4 °C and washed once with 1× ice-cold Tris-
buffered saline. Cells were resuspended in 1× Tris-buffered saline and transferred to
a screw-cap tube. After centrifugation, the supernatant was discarded, and the pellet
was frozen and stored at –80 °C. Pellets were resuspended in 500 µl of lysis buffer
(50 mm Hepes/KOH (pH 7.5), 150 mm NaCl, 1 mm EDTA, 1% Triton X-100, 0.1%
sodium deoxycholate, 1x Calbio phosphatase inhibitor, and 1× Sigma fungal
protease inhibitor mixture), and cells were lysed by bead beating ten times for 1 min
with 5-min rests on ice. Tubes were punctured and spun into tubes for 1 min at 1000
rpm. The flow-through was transferred to a microcentrifuge tube and sonicated four
times for 15 s 12-15% amplitude duty cycle resting on ice for 5mins in between
sonications to achieve shearing between 500-750bps. Samples were then spun at
14,000 rpm for 5 min at 4 °C, the supernatant was transferred to a new tube and
spun again for 10mins at 4˚C. Supernatant was transferred to a new tube and
quantified using Bradford reagent. 1-2mg of crude lysate diluted to 400ul in ChIP
lysis buffer was precleared using 30ul Invitrogen Protein A beads rotating at 4˚C for
2hours. 20ul was set aside as the input control. 100 µl of the lysis buffer plus a 1:20
dilution of anti-GFP antibody (Abcam 290) or with no antibody (Mock) was added
and rotated at 4˚C overnight. 15ul of NEB Protein A magnetic beads were added to
lysate and rotated for 1 hour at 4˚C. Beads were washed twice for 5 min with lysis
buffer, twice for 5 min with high salt lysis buffer (lysis buffer with 500 mm NaCl),
once for 5 min with wash buffer (10 mm Tris-HCl (pH 8.0), 0.25 m LiCl, 0.5%
Nonidet P-40, 0.5% sodium deoxycholate, and 1 mm EDTA), and once for 5 min

188

with 10 mm Tris-HCl (pH 8.0) and 1 mm EDTA. Samples were eluted by addition of
100 µl of elution buffer (50 mm Tris (pH 8.0), 10 mm EDTA, and 1% SDS) and
incubation for 30 min at 65°C with agitation every 5mins. Samples crosslink was
reverse by incubation at 55˚C overnight. 200ug proteinase K was added and
incubated at 37˚C for 2hours. Samples were purified using a Qiagen PCR
purification kit. DNAs were quantified by end point quantitative PCR with primers
specific for the dh region and act1. PCR products were separated using a 4% gel
and visualized via syber green staining and scanning on biorad FX scanner.
Quanitfication was through GelEval Version 1.37 (1.37) using IP/Input – Mock/Input.

4.7 Movie Legends
Movie 4.1: Representative live cell imaging of WT heterozygote LacI-GFP and lacO
at centromere I with H3-mRFP in meiosis. Yellow is LacI-GFP and magenta is H3-
mRFP.
Movie 4.2: Representative live cell imaging of quad∆ heterozygote LacI-GFP and
lacO at centromere I with H3-mRFP in meiosis. Yellow is LacI-GFP and magenta is
H3-mRFP.
Movie 4.3: Representative live cell imaging of WT with Rec8-GFP in meiosis.
Green is Rec8-GFP.
Movie 4.4: Representative live cell imaging of quad∆ with Rec8-GFP in meiosis.
Green is Rec8-GFP.

189

Movie 4.5: Representative live cell imaging of WT with Rad21-GFP and H3-mRPF
in meiosis. Yellow is Rad21-GFP and magenta is H3-mRFP.
Movie 4.6: Representative live cell imaging of quad∆ with Rad21-GFP and H3-
mRPF in meiosis. Yellow is Rad21-GFP and magenta is H3-mRFP.

190

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Chapter 5

A Mammalian-Like DNA Damage Response of Fission Yeast to Nucleoside
Analogs

The work in this chapter was originally published as
Sabatinos, Sarah A. et al. “A Mammalian-Like DNA Damage Response of
Fission Yeast to Nucleoside Analogs.” Genetics 193.1 (2013): 143–157. PMC.
Web. 18 Dec. 2014.

T. L. Mastro performed the experiments in Figure 5.1 b, d, and e, 5.2, 5.3, 5.4 a, b,
d, 5.5 c, d, e, 5.7 a, 5.8 e, Table 5.1; and contributed to the writing and editing of the
manuscript

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5.1 Overview
Nucleoside analogs are frequently used to label newly synthesized DNA.
These analogs are toxic in many cells, with the exception of the budding yeast. We
show that Schizosaccharomyces pombe behaves similarly to metazoans in
response to analogs 5-bromo-29-deoxyuridine (BrdU) and 5-ethynyl-29-deoxyuridine
(EdU). Incorporation causes DNA damage that activates the damage checkpoint
kinase Chk1 and sensitizes cells to UV light and other DNA-damaging drugs.
Replication checkpoint mutant cds1Δ shows increased DNA damage response after
exposure. Finally, we demonstrate that the response to BrdU is influenced by the
ribonucleotide reductase inhibitor, Spd1, suggesting that BrdU causes dNTP pool
imbalance in fission yeast, as in metazoans. Consistent with this, we show that
excess thymidine induces G1 arrest in wild-type fission yeast expressing thymidine
kinase. Thus, fission yeast responds to nucleoside analogs similarly to mammalian
cells, which has implications for their use in replication and damage research, as
well as for dNTP metabolism.
5.2 Introduction
Understanding mechanisms that maintain DNA replication fidelity is key to
understanding cancer (Barlogie et al. 1976; Johnson et al. 1981; Christov and
Vassilev 1988). Replication must be controlled to avoid mutations, promote DNA
repair, and restrain rereplication. Predictably, abnormally replicating and proliferating
cells are a hallmark of tumorigenesis (Barlogie et al. 1976; Johnson et al. 1981;
Christov and Vassilev 1988). Thus, accurate analysis of replication states is
essential to understanding the response of a cell population under study.

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Studies of replication dynamics rely on manipulations of DNA nucleotide
metabolism. For example, the drug hydroxyurea (HU) is commonly used to inhibit
nucleotide synthesis. This causes replication fork stalling and checkpoint activation
until dNTP levels are reestablished (Santocanale and Diffley 1998; Kim and
Huberman 2001; Lopes et al. 2001; Alvino et al. 2007; Poli et al. 2012).
Dysregulation of nucleotide levels is associated with disruptions in cell-cycle and
checkpoint control (Kumar et al. 2010, 2011; Davidson et al. 2012; Poli et al. 2012).
The regulatory response to nucleotide levels is different between the yeasts
Saccharomyces cerevisiae and Schizosaccharomyces pombe. Budding yeast S.
cerevisiae is highly resistant to HU, and its checkpoint proteins directly regulate
nucleotide biosynthesis during the normal cell cycle (Chabes et al. 2003; Poli et al.
2012). In contrast, fission yeast S. pombe is sensitive to much lower levels of HU
and uses checkpoint-independent mechanisms to control nucleotide levels during
normal cell growth (Hakansson et al. 2006; Nestoras et al. 2010).
Exposure to exogenous nucleosides alters metazoan dNTP metabolism and
cell-cycle dynamics (Meuth and Green 1974a,b), which is a potential problem in
assays that rely on incorporation of modified or antigenic nucleoside analogs. By
measuring incorporation of analog, the replicative capacity of a culture is inferred
(Bohmer and Ellwart 1981; Crissman and Steinkamp 1987; Frum and Deb 2003),
providing a direct metric of DNA synthesis. Thymidine analogs 5-bromo-29-
deoxyuridine (BrdU) and 5-ethynyl-29- deoxyuridine (EdU) (Diermeier-Daucher et al.
2009) are commonly used nucleosides. BrdU is detected using antibodies (Hodson
et al. 2003), while EdU is covalently labeled by bio-orthogonal, copper-based

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chemistry (Buck et al. 2008). In yeasts, which lack a thymidine salvage pathway,
analog incorporation requires an exogenous thymidine kinase (e.g., herpes simplex
virus, hsv-tk
+
) (Sclafani and Fangman 1986; Hodson et al. 2003; Sivakumar et al.
2004; Viggiani and Aparicio 2006).
A disadvantage to this method is the inherent toxicity of nucleoside analogs
such as BrdU, FdU, and IdU (Sivakumar et al. 2004). Observations in bacteria and
mammals indicate that BrdU is both a mutagen and a teratogen (Davidson and
Kaufman 1978; Kaufman and Davidson 1978; Lasken and Goodman 1984), able to
cause T-C transition mutations (Goodman et al. 1985). However, up to 400 mg/ml
BrdU may be added to a culture of S. cerevisiae before toxic effects are observed
(Lengronne et al. 2001). In fission yeast, both EdU and BrdU analogs show toxicity
at much lower doses and may activate a Rad3 (ATR/Mec1)-dependent damage
response pathway (Hodson et al. 2003; Sivakumar et al. 2004; Hua and Kearsey
2011). Differences in dosage sensitivity may reflect the differences in nucleotide
metabolism between these different yeast species.
We examine the range of toxicity and the induction of DNA damage in S.
pombe cells that incorporate either BrdU or EdU. Checkpoint mutants rad3Δ, chk1Δ,
and cds1Δ show hypersensitivity to BrdU and EdU, which is exacerbated in minimal
media. Cells exposed to low doses of BrdU are sensitized to additional DNA-
damaging agents and show increased mutation rates. Even in low-dose analog, we
observe induction of damage markers including phosphorylated histone H2A and
Rad52 foci. Chk1 and Cdc2 phosphorylation indicates the DNA damage checkpoint
is activated, showing that analog incorporation generates DNA damage.

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Consistent with this, we observe that completion of S phase and entry to the
next cell cycle is delayed with increased analog dose. This effect is not limited to
analogs, however, because fission yeast cells exposed to thymidine undergo G1
arrest, similar to thymidine synchronization in human cells. Finally, the ribonucleotide
reductase (RNR) inhibitor Spd1 modulates cell response to BrdU and EdU. While
spd1Δ cells are less sensitive to chronic exposure, acute viability is decreased and
surviving spd1Δ cells frequently carry mutations. Thus, the power of using analog
incorporation to detect new DNA synthesis must be tempered by appropriate
cautions to maximize relevant conclusions and minimize disruptions to normal
nucleotide metabolism.
5.3 Results
5.3.1 Cell signal and viability vary with analog dose
Acute nucleoside analog toxicity is proportional to dose in human cells
(Popescu 1999). To compare this with fission yeast we used BrdU and EdU
concentrations typically used in replication studies. Our strains contained hsv-tk
+
and the human equilibrative nucleoside transporter (hENT), which allowed us to use
lower analog doses and achieve effective transport into cells (Hodson et al. 2003;
Sivakumar et al. 2004). Since BrdU dose is frequently presented in micrograms per
milliliter, we began with a starting BrdU dose of 100 mg/ml (326 mM), half the dose
used in fission yeast genome labeling work without hENT (e.g., Hayano et al. 2011)
and less than commonly used in budding yeast (e.g., Lengronne et al. 2001).
However, to compare between BrdU (typically milligrams per milliliter) and EdU

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(typically micromolar), we report all doses in molarity. The starting dose for each
(326 mM BrdU and 10 mM EdU) was determined by previous protocols.

We compared BrdU signals, using flow cytometry on HU-blocked cells
released into BrdU (32.6 or 326 mM) or EdU (1 or 10 mM), to determine whether
there was a corresponding decrease in the resulting signal (Figure 5.1). Low-dose
BrdU (32.5 mM) produced a similar signal to the full dose of 326 mM, but with better
viability (Figure 5.1A). EdU toxicity appeared at a dose of 10 mM (Figure 5.1D),
consistent with results in Hua and Kearsey (2011), yet a 10-fold decrease in EdU
dose to 1 mM significantly decreased EdU signal intensity (Figure 5.1C). Thus,
minimum doses of 32.6 mM BrdU and 10 mM EdU are required for optimal short-
term labeling in hsv-tk
+
hENT
+
strains. Higher doses of nucleoside analog may be
required to detect small incorporation differences during short incubations, but
enhanced signal comes at a cost to cell viability.
Disruptions in DNA replication and repair activate checkpoint pathways
(reviewed in Sabatinos and Forsburg 2012). The master regulator in fission yeast is
the Rad3
ATR
kinase, which activates Chk1 kinase at DNA double-strand breaks
(DSBs). During replication fork stalling, Cds1 kinase is activated by Rad3 via the fork
processivity factor Mrc1. Mutations in these checkpoint proteins cause sensitivity to
drugs that activate the replication checkpoint. Previous work indicated that Rad3 is
required for viability in EdU, but did not investigate the downstream pathways
required (Hua and Kearsey 2011).
We examined cell viability in replication checkpoint mutant cds1Δ and DNA
damage checkpoint mutant chk1Δ, compared to wild-type and rad3Δ incorporating

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strains (Figure 5.1, B and D). We found chk1Δ and rad3Δ are strikingly sensitive to
BrdU and EdU treatment. While cds1Δ behaved similarly to wild-type cells during the
first 4 hr of incubation, prolonged BrdU exposure (8 hr) caused decreased cds1Δ
viability (similar results for mrc1Δ, data not shown). Lower BrdU and EdU doses
improved viability in all genotypes (Supporting Information, Figure 5.3), yet rad3Δ
and chk1Δ cells were still the most EdU sensitive.
Wild-type cells without the incorporation cassette continued to divide normally
during exposure (Figure 5.1E). Wild-type hsv-tk
+
hENT
+
cells had reduced division
during analog exposure, as did cds1Δ, rad3Δ, and mrc1Δ (data not shown). In
contrast, the chk1Δ incorporating strain continued to divide in BrdU and EdU,
suggesting that Chk1 activation is required to inhibit cell division during exposure.

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Figure 5.1: BrdU and EdU doses affect signal, viability, and cell division. (A)
Time course of incorporation at 32.6 or 326 mM BrdU in HU-synchronized cells after
release. Asynchronous (AS) cultures were blocked for 4 hr in HU (HU time point)
before release at 32˚C in medium with BrdU at the indicated concentration. BrdU
signal was detected in isolated nuclei. (B) Relative viability during 32.6 mM BrdU
incubation, comparing non-incorporating (wt) or hsv-tk+ hENT+ cells, both wild type
(wt-inc) and checkpoint mutants. Means +/- SEM of three experiments are shown.

206

(C) As in A, time course of EdU incorporation at 1 and 10 mM doses in HU-
synchronized cells post-release. Whole cells were treated with ClickIt reaction before
flow cytometry. (D) Relative viability in 10 mM EdU treatment over time for
nonincorporating (wt) or incorporating wild-type (wt-inc) or checkpoint-mutant
incorporating cells. Means +/- SEM of three experiments are shown. (E) Cells were
counted during BrdU or EdU treatment to determine proliferation in nonincorporating
(wt) or hsv-tk+ hENT+ wild-type (wt-inc) and checkpoint-mutant cells. Cell
concentrations were normalized to the 0-hr sample for each cell line/condition and
are shown as means +/- SEM (n = 3).

Checkpoint and cell-cycle phenotypes can be sensitive to media composition.
We examined sensitivity to BrdU and EdU in spot tests on three media types: rich
YES, defined EMM, and lower-nitrogen PMG (Figure 5.2 and Figure 5.3). On YES,
chk1Δ incorporating cells show modest sensitivity at 16.3 mM, while growth
inhibition in rad3Δ and cds1Δ began to show at 32.6 mM (Figure 5.2A). EdU
sensitivity was highest in chk1Δ on YES (5 mM) followed by cds1Δ (10 mM).
Unexpectedly, rad3Δ sensitivity was similar to wild type at 10 mM EdU in YES
(Figure 5.2A).

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Figure 5.2: Media formulation alters BrdU and EdU sensitivity. (A) Serial
dilution assay in YES medium of nonincorporating wildtype (WT) and hsv-tk+ hENT+
WT and checkpoint-mutant strains cds1Δ, chk1Δ, rad3Δ, and mrc1Δ. Plates
containing BrdU, thymidine (Thy) control, or EdU were compared using 1/5 dilutions
of cells, grown at 32˚C for 3 days. (B) As in A, spot tests on defined nitrogen rich
EMM medium. Refer to Figure 5.3 for PMG media effects.

208

On EMM, which contains high-level nitrogen, we found that wild-type, chk1Δ,
and rad3Δ hsv-tk
+
hENT
+
cells were all BrdU sensitive (Figure 5.2B). Yet, mrc1Δ
and cds1Δ were less sensitive. Wild-type, chk1Δ, and rad3Δ hsv-tk
+
hENT
+
cells
were most sensitive at 10 mM EdU in EMM, followed by cds1Δ and mrc1Δ. Non-
incorporating cells were not sensitive to EdU or BrdU on any media.
PMG medium contains low levels of nitrogen. All incorporator strains were
sensitive to 16.3 mM BrdU in PMG (Figure 5.3), above which there was little growth.
Similarly, EdU sensitivity was higher in PMG and chk1Δ cells were most sensitive to
EdU in PMG, followed by cds1Δ. Thus, cells are less sensitive to analogs on rich
medium than on minimal medium, while chk1Δ restricts growth in all cases.

209

Figure 5.3 BrdU and EdU dose affects cell viability (refer to Figures 5.1 and
5.2). (A) Relative viability of hsv-‐tk+ hENT+ wild-‐type, cds1Δ, chk1Δ, and rad3Δ
cells at 16.3µM BrdU (compare with Figure 5.1B) with non-incorporating (N.I.)
control. Shown are the means of two independent experiments ±SEM. (B) As in (A)
viability in lower dose EdU (5µM) (compare with Figure 5.1D). Shown are the
means of two independent experiments ±SEM. (C) Spot tests on poor-nitrogen
source medium PMG for non-incorporating wild-type (wt) and hsv-tk+ hENT+cells of
indicated genotype (compare with Figure 5.2).

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5.3.2 S-phase progression is slowed by analog incorporation
We next tested whether analog incorporation has an effect on DNA synthesis.
We blocked cells in early S phase with HU and released them into medium with low-
or high-dose BrdU or EdU to monitor replication. Non-incorporating cells completed
S phase by 1 hr postrelease, with the appearance of a 2C DNA peak. A second S
phase was detected 1.5–2 hr postrelease by 4C DNA content and a septation index
(Figure 5.3, A and B, and Figure 5.5A). We detected a shift in forward scatter (FSC)
to small cells (Figure 5.5B), correlating with cell division at 2 hr post-release.
BrdU exposure at both doses delayed S phase completion in incorporating
strains. The DNA peak moved slowly to a 2C position, completing by 2 hr
postrelease. Septation was highest at 3 hr, coincident with a small 4C peak
observed by FACS (Figure 5.3A and Figure 5.5A). FSC confirmed the BrdU-
incorporated cells were elongated (Figure 5.5B), consistent with incomplete
septation.
Cells treated with 1 mM EdU completed the first S phase by 1 hr and entered
the second S phase at 2 hr postrelease with increased septa and 4C DNA (Figure
5.3B and Figure 5.5A). Cells in 10 mM EdU completed a first S phase within 2 hr but
had much slower transit through the second S phase, which started at 2 hr and was
not resolved by 3 hr (Figure 5.5A). Thus, EdU causes a dose-dependent inhibition of
cell-cycle progression in incorporating cells.

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Thymidine is commonly used in human cell cultures to arrest cells in G1

(Harper 2005), but does not affect yeast since S. pombe does not have a thymidine
salvage pathway (non-incorporating cells). We asked whether thymidine changes
cell cycle in hENT
+
hsv-tk
+
strains (Figure 5.4C). We treated cells with 2 mM
thymidine (Harper 2005) and saw no effect in non-incorporating cells or in cells with
hENT
+
but no hsv-tk. However, an incorporating strain treated with 2 mM thymidine
arrested with a 1C DNA content. After thymidine removal, cells shifted toward 2C
DNA content. Thus, thymidine reversibly arrests hENT
+
hsv-tk
+
S. pombe cells with
a 1C DNA content, as in mammals.

212

213

Figure 5.4 BrdU and EdU cause prolonged DNA synthesis, cell-cycle slowing,
and DNA damage. (A) DNA synthesis profiles of wild-type non-incorporating (non-
inc) and incorporating (Inc) cells, out of hydroxyurea arrest (HU), released into
medium with 32.6 or 326 mM BrdU to detect DNA replication. Left, whole-cell DNA
content (Sytox- Green) FACS profiles. Right, septation index for non-inc (NI) and Inc
(I) cells stained with aniline blue and DAPI, at different BrdU doses (mM). (B) As in
A, cells released from HU were released into medium with 1 or 10 mM EdU. Left,
FACS profiles of whole-cell DNA content. Right, septation index at different EdU
doses (mM). (C) Asynchronous (AS) cells were treated with 2 mM thymidine (+Thy)
or DMSO (vehicle control) for 3 hr (32˚C) and then released for 0.75 hr. DNA content
(SytoxGreen FACS) for each time point, analyzed by FACS, is shown at each point.
DMSO control was to test response to DMSO only and was not released. Similar
results were seen with aqueous thymidine solution (not shown). (D) Cells were
exposed to 32.6 mM BrdU for 2 hr (32_) and then processed for BrdU and phospho-
histone H2A (p-H2A) immunofluorescence. DNA was counterstained with DAPI.
Merged image is BrdU and p-H2A signals. Bar, 10 mm.

214

Figure 5.5 BrdU and EdU cause prolonged DNA synthesis, cell cycle slowing
and DNA damage (related to Figure 5.4). (A) Sytox Green stained cells (from
Figures 5.4A, 5.4B) were analyzed by flow cytometry to highlight progression to 4C
DNA (second S phase; using modified cytometer settings). Non-incorporating (non-
inc) or hsv-tk+ hENT+ cells (Inc) at indicated doses of BrdU or EdU. Note that 4C
peak accumulation is consistent with septation index peaks (Figure 5.4A, 5.4B),
indicating the second S phase after release. (B) Forward scatter (FSC) dynamics of
cells in (A) indicating cell size during experiment. Left-‐shift toward smaller cell size
(M phase) occurs slightly later than septation (S phase; Figure 5.4A, 5.3B). (C)

215

Cells were stained with DAPI and aniline blue to detect nuclei and septa,
respectively, before or after 6h of BrdU or EdU treatment. wild-type (wt)
incorporating cells elongate during prolonged exposure. Both chk1Δ and rad3Δ
hsv-‐tk+ hENT+ cells continue to septate and divide, and many cells mis-segregate
DNA (indicated by arrows). mrc1Δ and cds1Δ hsv-tk+ hENT+ cells show an
intermediate phenotype in EdU. Scale bar 10 µm. (D) Abnormal DNA segregation
events were scored as the percentage of cut or anucleate cells in the total
population during BrdU treatment. Shown are combined data from 2 independent
experiments, displayed as proportion of abnormal segregants ±95% CI. (E)
Abnormally segregated nuclei during EdU exposure. Shown are combined data
from 2 independent experiments, displayed as proportion of abnormal segregants
±95% CI.

5.3.3 Nucleoside analogs induce DNA damage response
Reduced viability of checkpoint mutants and delayed S-phase progression
during analog incorporation suggested that DNA damage was generated. We
examined molecular markers of DNA damage after BrdU treatment. First, we
detected histone H2A phosphorylated at S129 (p-H2A) and BrdU. Rad3 and Tel1
kinases phosphorylate p-H2A in response to DNA DSBs and replication stress
(Nakamura et al. 2004; Bailis et al. 2008). Nonincorporating cells had few p-H2A foci
(Figure 5.4D), while BrdU-labeled cells frequently costained with p-H2A. These
results suggest that BrdU incorporation activates a DNA damage response, which is
enhanced in cds1Δ cells that cannot properly respond to S-phase stress.

216

Next, we looked for activation of checkpoint components. We examined HA-
tagged Chk1 protein (Figure 5.6A), the G2-M. kinase of the S. pombe DNA damage
response (DDR). We observed that Chk1HA shows a characteristic phospho-shift
(Capasso et al. 2002) after 1 hr exposure to 326 mM BrdU in hENT
+
hsv-tk
+
cells
(Figure 5.6A) or after 3 hr in 32.6 mM BrdU. Thus, BrdU dose influences. However,
Cds1-myc kinase, required for response to replication stress, was not significantly
modified relative to overall protein level in BrdU (Figure 5.6A).
We observed a phospho-shift in the upstream G2-DDR checkpoint mediator
Crb2 in BrdU (Figure 5.6A), again consistent with G2 checkpoint activation.
Activated Chk1 leads to Cdc2 phosphorylation at tyrosine 15 to maintain G2
checkpoint arrest (O’Connell et al. 1997), and we saw phospho-Cdc2 accumulated
in BrdU-treated incorporating cells (Figure 5.46). A dose of 326 mM BrdU was lethal
to most cells during 3 hr exposure (Figure 5.6B), suggesting the damage is
catastrophic.
Similar results were seen in EdU, causing Chk1HA and Crb2 phospho-shifts
(Figure 5.6C) and increased phospho-Cdc2 (Figure 5.6D), which was proportional to
EdU dose (Figure 5.6E). These data indicate that the DNA damage checkpoint is
fully activated during analog exposure.

217

Figure 5.6: BrdU exposure triggers the DNA damage response. (A) Cells were
treated with the indicated dose of BrdU and harvested for protein extraction hourly.
Chk1-HA was detected with anti-HA (16B12); the asterisk indicates nonspecific
background signal and the open arrow indicates phospho-Chk1HA. Crb2
modification is also indicated by an open arrow. Cds1-myc was detected with
antimyc (solid arrow). Phospho-Cdc2 (p-Cdc2, open arrow) and Cdc2 were also
detected. PCNA and b-tubulin are loading controls. The solid line indicates a split
between two independent gels, with identical lysates. (B) Incorporating strain viability
(FY5031) proportional to BrdU dose is shown as means of three independent
experiments +/- SEM. (C) Chk-HA and Crb2 phosphorylation after 3 hr EdU (mM), in

218

wild-type hsv-tk+ or nonincorporating cells. BrdU (326 mM) is included as a control.
Chk1-HA and Crb2 band shifts are indicated with open arrows. The asterisk
indicates nonspecific background band (above Chk1HA) detected using a different
antibody from A (aHA, 12CA5). (D) Phospho-Cdc2 (p-Cdc2) after 3 hr BrdU or EdU
exposure (doses mM). Below bands are the quantified band intensities of p-Cdc2,
normalized to total Cdc2 (below). (E) Quantification of p-Cdc2, relative to total Cdc2
levels, from three independent experiments. Mean values are +/-SEM.

5.3.4 Loss of the replication checkpoint influences damage response
Our data suggest that the Chk1-DNA damage checkpoint pathway is a
primary response to analogs. However, the modest sensitivity caused by cds1Δ
indicated a role for the replication checkpoint during treatment. Cds1 is particularly
important for replication fork stalling and restart (e.g., Lindsay et al. 1998; Kim and
Huberman 2001; Miyabe et al. 2009). The homologous recombination protein Rad52
forms nu- clear foci in response to a variety of lesions, including double-strand
breaks and collapsed or restarting replication forks (Meister et al. 2005; Bailis et al.
2008). We monitored Rad52-YFP focus formation in wild-type and cds1Δ
incorporating strains after 3 hr in 32.6 or 326 mM BrdU (Figure 5.7A). Incorporating
cells formed more Rad52 foci than without BrdU treatment, consistent with increased
damage (Figure 5.7B). This effect was dose dependent, and more foci were
detected at 326 mM BrdU (wild type) and at all BrdU doses (cds1Δ).
Live cell analysis in a microfluidics chamber allowed us to track individual
cells during and after analog exposure (Movie 5.1-4). We monitored Rad52 foci

219

during 3 hr exposure to 32.6 mM BrdU (Movie 5.1 and 5.2) or 10 mM EdU (Movie
5.3 and 5.4) in wild-type and cds1Δ hsv-tk
+
hENT
+
cells. We used a dose of 32.6
mM BrdU be- cause this was the lowest dose that produced a difference in Rad52
foci (Figure 5.7B) and was close to the 10 mM EdU dose required to effectively label
cells for replication studies. BrdU movies recapitulated our static time-point data
(Figure 5.7, A and B), confirming that cds1Δ cells generated more Rad52 foci during
analog exposure. We also monitored recovery during 3 hr after analog was
removed, observing that multiple Rad52 foci (two or more) frequently resolved into
one focus. Further, cells that septated and presumably entered the next cell cycle
during BrdU (Movie 5.1 and 5.2) or EdU exposure (Movie 5.3 and 5.4) promptly
formed Rad52 foci, suggesting an immediate response is mounted during S phase.
However, cds1Δ cells that entered S phase during analog exposure frequently lysed
during recovery (Figure 5.7, C and D).
We next asked whether differences in Rad52 foci levels in different BrdU
doses could be attributed to different levels of BrdU incorporation in DNA. We
extracted total DNA from cells after 3 hr exposure to 32.6 or 326 mM BrdU and
blotted several amounts of heat-denatured DNA to probe with BrdU antibody (Figure
5.7E). Surprisingly, we found that the amount of BrdU incorporated at either 32.6 or
326 mM did not change. Thus, BrdU incorporation is saturated at 32.6 mM,
consistent with our earlier FACS results (Figure 5.1A). However, wild-type cell
viability was higher in 32.6 mM BrdU than in 326 mM BrdU (Figure 5.6B). This
suggests that additional stress occurs at the higher dose of BrdU that does not
involve BrdU-base substitution. Interestingly, we detected less BrdU incorporation in

220

cds1Δ compared to wild type in the samples with the higher levels of DNA. Thus,
enhanced analog sensitivity in cds1Δ is accompanied by less efficient BrdU
incorporation. This could reflect replisome disruption in cds1Δ or Cds1-mediated
interactions with dNTP metabolism.

221

Figure 5.7: BrdU and EdU induce a DNA damage response. (A) Rad52-YFP foci
were monitored in untreated cells (untrt) or after 3 hr BrdU at 32˚C. Rad52 foci (left)

222

or DAPI-stained nuclei (right) are shown on a transmitted light background. Bar, 10
mm. (B) Quantification of three independent experiments in A. Shown are
proportions of nuclei with two or more Rad52-YFP foci after 3 hr BrdU 6 95%
confidence interval (C.I.). (C and D) Time points selected from movies of wild-type
(C; and Movie 5.3) or cds1Δ (D; and File S4) cells treated with 10 mM EdU. Arrow
(in D) indicates cell that forms foci and lyses. Bar, 10 mm. (E) BrdU incorporated is
similar at 32.6 and 326 mM doses. Shown is the mean BrdU signal per dot (+/- SEM,
three independent experiments) at 2.5, 0.25, or 0.025 mg of heat-denatured total
DNA, blotted and detected with BrdU antibody. Below, example of BrdU detection on
DNA spots. (F) Wild-type nonincorporating or hsv-tk+ hENT+ cells were treated
BrdU doses for 3 hr, plated on YES, and then irradiated with 100 J/m2 UV light.
Comparison plates were not treated with BrdU, to calculate percentage of viability
after BrdU+UV treatment. Shown is the mean viability after BrdU+UV relative to
BrdU only, for three independent experiments +/- SEM.

5.3.5 Enhanced sensitivity to “second-hit” damage after BrdU exposure
Since hsv-tk
+
hENT
+
cells acquired DNA damage and G2

arrest signals, we
investigated whether BrdU incorporation changes cell sensitivity to other DNA
damage, as reported for human cells (e.g., Ackland et al. 1988; Cecchini et al.
2005). UV treatment on BrdU-substituted human cells induces DSBs, demonstrating
that BrdU-DNA is sensitized to additional lesions. To test this in S. pombe, we
treated cells with 3 hr of low-dose (32.6 mM) BrdU for minimal toxicity with saturated

223

incorporation and examined viability with UV treatment (Figure 5.7F).
Nonincorporating cells experienced a slight viability decrease, to 88 6 3%, after UV
dose of 100 J/m
2
. However, BrdU incorporation significantly decreased UV survival
and further, was dose dependent: cells treated with 3.2 mM BrdU were slightly more
UV sensitive (58% viability), while saturating doses of 32.6 or 326 mM had the same
effect on UV survival to 20% viability.
Next, we incubated cells with 32.6 mM BrdU for 2 hr and then examined their
sensitivity to a panel of DNA-damaging drugs by a spot test (Figure 5.8A). First, we
examined sensitivity to phleomycin, a radiomimetic (Figure 5.8B). All cells showed
increased sensitivity to phleomycin after BrdU pretreatment. DDR pathway mutants
rad3Δ and chk1Δ were most severely affected at lower doses, followed by mrc1Δ
and cds1Δ and then wild-type hsv-tk
+
hENT
+
cells.
BrdU substitution also enhanced sensitivity to camptothecin (CPT) in chk1Δ
(5 mM) and cds1Δ (10 mM) (Figure 5.8C). CPT causes single-strand DNA breaks by
covalently linking topoisomerase I to DNA, damage that is later converted to a DSB
(Hsiang et al. 1989). Although rad3Δ and mrc1Δ are highly sensitive to CPT, we
detected no increased CPT sensitivity following BrdU exposure in rad3Δ and mrc1Δ
cells.

224

Figure 5.8 BrdU pretreatment changes sensitivity to DNA damaging drugs. (A–
D) Cells were either untreated (untrt) or pretreated (+BrdU) with 32.6 mM BrdU for 2
hr at 32˚C and then spotted onto drug plates in a 1/5 serial dilution. All plates are
YES medium. Arrows, far right, indicate strains that were more sensitive to drug

225

following BrdU pretreatment. Strains FY3454, -2317, -3179, -5148, -5149, and -5150
are shown. Also refer to Figure S3. (A) YES control for plating efficiency. (B)
Sensitivity to phleomycin. (C) Camptothecin (CPT) sensitivity. (D) Hydroxyurea (HU)
sensitivity. (E) Forward mutation analysis for loss of Can1 wild-type status. Cells
were incubated with 32.6 mM BrdU for 2hr (32˚C) and then plated to assess colony
number on titer dishes. Remaining culture was plated on PMG + canavanine and
incubated 7 days at 32˚C. Mutation rate, per 107 generations, was calculated
comparing can1- mutants that grew on canavanine plates to the total number plated.
Refer to Table 5.1 for significance results.

The alkylating agent methanemethyl sulfonate (MMS) methylates DNA bases
and stalls replication forks. The rad3Δ cells were already highly sensitive to MMS
regardless of BrdU pretreatment (Figure 5.9). However, the remaining hsv-tk
+
hENT
+
strains were all more MMS sensitive following BrdU incorporation, with the
highest sensitivity in chk1Δ (Figure 5.9). HU stalls replication forks, but acts through
dNTP de- pletion as opposed to DNA base damage (Figure 5.8D). BrdU
pretreatment enhanced HU sensitivity only in the chk1Δ hsv-tk
+
hENT
+
strain. UV
lesions stall replication and transcrip- tion, so we looked at different doses of UV
post-BrdU. All strains except rad3Δ showed increased UV sensitivity after BrdU, with
chk1Δ being the most affected (Figure 5.9).

226

Figure 5.9 BrdU pre-treatment changes sensitivity to mutagens (refer to Figure
6). (A, B) Cells were untreated (untrt) or pre-treated (+BrdU) with 32.6 µM BrdU (2h
at 32°C), and then spotted onto drug plates (YES), 1/5 dilutions. Arrows indicate
greater sensitivity to drug +BrdU. Refer to Figure 6A for control. (A) Sensitivity to UV,
irradiated after plating yeast. (B) Sensitivity to MMS after BrdU treatment. (C)

227

Analysis of can1- isolates from forward mutation study. Strains were pooled to
assess can1 amplification and RFLP ± BrdU; no differences were seen between
genotypes. The can1 locus was amplified by PCR, and produces a 3.2 kb band by
agarose gel electrophoresis. PCR product was digested with EcoRI, producing 4
restriction fragments, which were screened on 8% TBE-PAGE gels. Lane 1 (top) is a
negative (water) control for PCR. Lanes 2 and 3 are non-incorporating strains that
were known can1+ or can1-1 genotypes. Restriction fragment length differences
were not detected in any of the can1- isolates. Instead, a minority of BrdU treated
isolates failed to amplify a detectable can1 band (8.6% of all BrdU treated isolates).

We then determined spontaneous mutation frequencies after 32.6 mM BrdU
treatment. We used the can1
+
gene, which encodes an arginine transporter that
imports the toxic precursor canavanine; can1
-
mutants are resistant to canavanine
(Fantes and Creanor 1984). Increased can1 mutation occurred after BrdU exposure
in wild-type and cds1Δ hsv-tk
+
hENT
+
cells (Figure 5.8E and Table 5.1), but not in
chk1Δ and nonincorporating cells. Intriguingly, rad3Δ and mrc1Δ hsv-tk
+
hENT
+
cells had a significantly lower mutation rate after BrdU treatment.

We amplified the can1 locus from can1
+
colonies in all genotypes (with or
without BrdU) to determine whether can1 mutation occurred by gross chromosomal
rearrange- ment. Instead, we saw a 3.8-kb can1 band in 91% of BrdU-treated hsv-
tk
+
hENT
+
can1
-
isolates (Figure 5.9). We checked for smaller deletions by
restriction fragment length polymorphism (RFLP) analysis and did not see

228

differences between fragments. Thus, we infer that BrdU-induced mutation in wild-
type or cds1Δ cells at can1 is largely due to point mutations, as in metazoans
(Goodman et al. 1985).
Table 5.1: Mutation rates for spontaneous and BrdU-induced canavanine
forward mutation analysis
Strain Strain
Treatment
a

No.
mutation
(m)
b

can1- rate/
10
7

generations
b

95% C.I.
b
t-test
c

Wild type,
non-inc
(FY3454)
Untreated (n =
8)
4.669 5.59 3.02/8.70

*
+BrdU (n = 8) 4.77 6.82 3.70/10.60
Wild type
(FY2317)
Untreated (n =
8)
5.601 9.56 5.37/14.59 P
<0.001
+BrdU (n = 7) 9.079 14.19 8.41/21.00
cds1∆
(FY5148)
Untreated (n =
8)
5.617 9.84 5.53/15.01 P <
0.025
+BrdU (n = 8) 7.350 14.21 8.43/21.05
chk1∆
(FY5149)
Untreated (n =
8)
8.079 15.82 9.55/23.19 *
+BrdU (n = 8) 8.714 16.80 10.28/24.45
rad3∆
(FY5150)
Untreated (n =
8)
4.272 9.68 5.12/15.24 P <
0.025
+BrdU (n = 8) 5.868 9.56 5.43/14.52
mrc1∆
(FY3179)
Untreated (n =
8)
6.525 11.36 6.59/17.04 P <
0.025
+BrdU (n = 8) 8.393 9.23 5.61/13.48
a
Experiments were plated in duplicate and results summed for analysis. Total
number of biological replicate experiments is indicated (“n”) for each sample.
b
Number of mutations (m), mutation rate, and 95% confidence interval (C.I.)
calculated using the Ma–Sandri–Sarkar maximum-likelihood estimator (MSS-MLE)
method FALCOR calculator (http://www.keshavsingh.org/protocols/FALCOR.html).
Data are from eight independent assays (Hall et al. 2009).

229

c
Pairwise two-tailed t-test calculated within genotypes with or without BrdU from
MSS-MLE results, using mutation number (m) and calculated variance, with
(n
withBrdU+
n
withoutBrdU
- 2) d.f. *Not significant, P > 0.05.

5.4.6 dNTP pools influence BrdU toxicity and mutagenesis
We next asked whether BrdU affects dNTP pools, which would cause toxicity
or mutation. Fission yeast Spd1 inhibits RNR to regulate RNR activity throughout the
cell cycle and in response to DNA damage and repair. Previous work has shown that
spd1Δ cells have higher endogenous dNTP pools (Holmberg et al. 2005). We
reasoned that increased pools of normal dNTPs might dampen the sensitivity to
exogenous nucleotides. Consistent with this prediction, we found that spd1Δ hENT
hsv-tk
+
cells were less sensitive to thymidine, BrdU, or EdU than wild type (Figure
5.10A and Figure 5.11). spd1Δ cells were minimally sensitive to BrdU or EdU in YES
(Figure 5.11A). In EMM with BrdU or EdU, we observed two colony sizes in spd1Δ
strains: large colonies, similar to other incorporating cells, amid a high background
population of small colonies (Figure 5.11B). We found that spd1Δ cells incorporate
BrdU similarly to wild type and produce p-H2A foci in BrdU-labeled nuclei (Figure
5.11C).

230

Figure 5.10: Spd1 protects cells from division and mutation during dNTP
imbalance. (A) Comparison between wild-type and spd1Δ hsv-tk+ hENT+ strains by
spot test on EMM plates containing BrdU, EdU, or thymidine. DMSO is a vehicle
control for EdU. Shown is the minimal dose where wildtype cells began to show
sensitivity to analogs. Strains FY2317, -3454, and -6247 are shown. (B) Cultures

231

were treated with 32.6 mM BrdU and plated to calculate viability relative to 0 hr.
Wild-type (wt) and spd1Δ strains express hsv-tk+ hENT+ (FY2317 and -6247), while
the nonincorporating control (non-inc, FY3454) does not. Shown are mean viability
values from three independent experiments +/- SEM. (C) Proliferation was monitored
by counting cell concentration during BrdU treatment for cultures as in A, in addition
to rad3Δ hsv-tk+ hENT+ (FY5150). Shown are mean viability values (n = 3
experiments) +/- SEM. (D) Canavanine mutation was scored for incorporating wild-
type and spd1Δ strains (FY2317 and -6247), with or without 32.6 mM BrdU
treatment (2 hr, 32˚C). Lea and Coulson fluctuation analysis was used to calculate
the rate of can1- forward mutation (per 107 generations) in independent cultures
over three experiments (wt, n = 12; spd1Δ, n = 15). Shown are median mutation
rates with quartile bounding boxes and 95% C.I. error whiskers. Significance was
assessed by two-tailed pairwise Mann–Whitney U-tests, *P = 0.0001, **P<0.0001.
(E) Colonies of wild-type or spd1Δ cells, untreated (no drug) or following 2 hr in 32.6
mM BrdU, were grown on YES and then replicated onto medium with FUdR to score
for hsv-tk+ loss. Significance was assessed by two-tailed Z-test (**P < 0.0002). (F)
Enhanced sectoring of colonies on FUdR was noted for spd1Δ cells either untreated
or following BrdU exposure as in E (two-tailed Z-test, ** P < 0.0002), compared to
BrdU-treated wild-type cells. Inset, example of spd1Δ colony with hsv-tk+ loss (*) or
sectored area (arrow). Frequencies were calculated from independent experiments,
presented with 95% C.I. (G) Model for the effect of exogenous thymidine (Thy) and
nucleoside analogs in fission yeast cells expressing a reconstituted thymidine
salvage pathway (hsv-tk+). Details are in Discussion.

232

Because spd1Δ hsv-tk
+
hENT
+
cells were less sensitive to analogs than wild-
type incorporating cells in chronic exposure, we were surprised to find that spd1Δ
viability was lower than wild type during acute BrdU treatment in liquid culture
(Figure 5.10B). However, spd1Δ hsv-tk
+
hENT
+
cells continued to divide in 32.6 mM
BrdU, while wild-type and rad3Δ cells delayed division (Figure 5.10C). We examined
can1 reversion, in a direct comparison between wild type and spd1Δ after 32.6 mM
BrdU for 2 hr at 32˚C (Figure 5.10D). Consistent with Figure 6E, wild-type hsv-tk
+
hENT
+
cells experienced an increase in can1 mutation with 32.6 mM BrdU
treatment. Notably, spd1Δ cells already had a higher rate of mutation before BrdU,
which increased a further 10-fold after BrdU incorporation.
We reasoned that enhanced spd1Δ survival on plates (despite poor viability
and enhanced mutagenesis in liquid) might be attributed to genetic selection
combined with the increased rate of mutation, leading to loss of the hsv-tk
+
cassette.
If this were to happen, a subpopulation of cells would accumulate that is insensitive
to further analog incorporation, which could explain the background small cells on
streaked plates (Figure 5.11B). Cells expressing hsv-tk
+
are dNTP pools influence
BrdU toxicity and mutagenesis

233

Figure 5.11: Spd1 protects cells from division and mutation during dNTP
imbalance (refer to Figure 7). (A) On YES medium, spd1Δ cells withstand high
doses of EdU and BrdU. DMSO is a vehicle control for EdU. Note that wild-type (wt)
hsv-‐tk+ hENT+ cells were also resistant to 50 µM thymidine on rich media. (B)
Strains (FY 2317, 3454, 6427, 5030, 5031, 5150, 5149, 5148) were streaked onto
supplemented EMM with thymidine, BrdU or EdU to assess growth on plates. spd1Δ
hsv-tk+ hENT+ cells formed some large colonies, but also a background of small
colonies, also seen for cds1Δ hsv-‐tk+ hENT+. (C) Immunofluorescence of spd1Δ
hsv-tk+ hENT+ cells, after 2h BrdU treatment for nuclei (DAPI), BrdU incorporation,

234

phospho-‐histone H2A (p-H2A), and merged BrdU/p-H2A. Scale 10 µm. (D) Addition
of 2 mM thymidine over prolonged periods in non-incorporating wild-type (wt), wt and
spd1Δ hsv-tk+ hENT+ cells (Inc). The 1C and 2C DNA content peaks are indicated;
G1 arrest causes a shift toward the 1C peak.

We next asked whether BrdU affects dNTP pools, which would cause toxicity
or mutation. Fission yeast Spd1 inhibits RNR to regulate RNR activity throughout the
cell cycle and in response to DNA damage and repair. Previous work has shown that
spd1Δ cells have higher endogenous dNTP pools (Holmberg et al. 2005). We
reasoned that increased pools of normal dNTPs might dampen the sensitivity to
exogenous nucleotides. Consistent with this prediction, we found that spd1Δ hENT
hsv-tk
+
cells were less sensitive to thymidine, BrdU, or EdU than wild type (Figure
5.10A and Figure 5.11). spd1Δ cells were minimally sensitive to BrdU or EdU in YES
(Figure 5.11). In EMM with BrdU or EdU, we observed two colony sizes in spd1Δ
strains: large colonies, similar to other incorporating cells, amid a high background
population of small colonies (Figure 5.11B). We found that spd1Δ cells incorporate
BrdU similarly to wild type and produce p-H2A foci in BrdU-labeled nuclei (Figure
5.11C).
Because spd1Δ hsv-tk
+
hENT
+
cells were less sensitive to analogs than wild-
type incorporating cells in chronic exposure, we were surprised to find that spd1Δ
viability was lower than wild type during acute BrdU treatment in liquid culture
(Figure 5.10B). However, spd1Δ hsv-tk
+
hENT
+
cells continued to divide in 32.6 mM

235

BrdU, while wild-type and rad3Δ cells delayed division (Figure 5.10C). We examined
can1 reversion, in a direct comparison between wild type and spd1Δ after 32.6 mM
BrdU for 2 hr at 32˚C (Figure 5.10D). Consistent with Figure 6E, wild-type hsv-tk
+
hENT
+
cells experienced an increase in can1 mutation with 32.6 mM BrdU
treatment. Notably, spd1Δ cells already had a higher rate of mutation before BrdU,
which increased a further 10-fold after BrdU incorporation.
We reasoned that enhanced spd1Δ survival on plates (despite poor viability
and enhanced mutagenesis in liquid) might be attributed to genetic selection
combined with the increased rate of mutation, leading to loss of the hsv-tk
+
cassette.
If this were to happen, a subpopulation of cells would accumulate that is insensitive
to further analog incorporation, which could explain the background small cells on
streaked plates (Figure 5.11B). Cells expressing hsv-tk
+
are sensitive to FudR, so
loss of the cassette can be scored by assessing the frequency of FudR-resistant
colonies or sectors (Kiely et al. 2000). We observed that untreated spd1Δ cells give
rise to FudR-resistant colonies infrequently, at a rate similar to wild type (Figure
5.10E and Table 5.2). However, BrdU treatment caused a significant increase in
FudR resistance in spd1Δ compared to wild-type cells. Significantly, higher colony
sectoring (Figure 5.10F) is also consistent with an increase in mutation frequency
and suggests that spd1Δ has higher genomic plasticity even within single colonies.

236

Table 5.2: Frequency of hsv-tk+ loss or sectoring in incorporating wild-type
and spd1Δ cultures, with or without BrdU treatment
Strain Treatment FUdR resistance
(%)
a

Sectored colonies
a

Wild type
(FY2317)
Untreated (n = 8422) 0.226 (+/- 0.101) 1.35 (+/- 0.25)
+BrdU (n = 5474) 0.274 (+/- 0.138) 1.41 (+/- 0.31)
spd1∆
(FY6247)
Untreated (n = 6968) 0.172 (+/- 0.097) 2.65 (+/- 0.38)
+BrdU (n = 612) 2.78 (+/- 1.30) 33.50 (+/- 3.74)
a
FUdR resistance and sectoring frequencies are presented with 95% confidence
intervals (C.I.). Data from two (wild type) or three (spd1Δ) independent experiments
are shown.

5.4 Discussion
Previous studies in budding yeast indicated that BrdU incorporation does not
perturb yeast cell growth (Lengronne et al. 2001), although specific mutants may be
more sensitive to BrdU during replication (Hodgson et al. 2007). We propose that
BrdU, EdU, and thymidine all skew dNTP pools in hsv-tk+ hENT+ fission yeast. Our
data show that S. cerevisiae and S. pombe are different in their response to
nucleotide analogs and to dNTP imbalance. This is consistent with a 10-fold
difference in HU sensitivities between the yeasts; HU also acts by depleting dNTPs.
Studies in metazoans found that BrdU treatment inhibits RNR and causes a “dCTP-
less” state (Meuth and Green 1974b) (Figure 5.10G, part 1). BrdU-induced
mutagenesis is attributed to low levels of dCTP (Hopkins and Goodman 1980; Meuth
1989), which is suppressed by co-culture with exogenous cytidine (Meuth and Green

237

1974b; Davidson and Kaufman 1978). Thymidine arrest in mammalian cells (Harper
2005) is also caused by decreased dCTP levels, which are restored once excess
thymidine is removed from the medium (Meuth 1989; Kunz and Kohalmi 1991).
Unlike BrdU, thymidine is minimally toxic over short time periods (Lockshin et al.
1985), although high doses (0.2 mM) induce mutation (Phear and Meuth 1989),
polyploidy, and chromosomal aberrations (Potter 1971). Interestingly, lower BrdU
concentrations (e.g., 32.6 mM) skew dNTP pools similarly to much higher thymidine
doses (2 mM) (Meuth and Green 1974a; Meuth et al. 1976).
We hypothesize that exposure to exogenous nucleosides creates dNTP pool
imbalance in S. pombe. During thymidine incubation, hsv-tk+ hENT+ cells arrest with
1C DNA content and release into S phase within 45 min. Previous work by Mitchison
and Creanor (1971) determined that thymidine had no effect on S. pombe,
consistent with the absence of a thymidine salvage pathway in wild-type cells.
However, our incorporating strains have a reconstituted thymidine salvage pathway
and are able to convert exogenous thymidine, BrdU, or EdU into nucleotides.
Mitchison and Creanor (1971) also demonstrated that deoxyadenosine
treatment induces a short-term G1 arrest in wild-type cells (Mitchison and Creanor
1971). This is consistent with our data and suggests that dNTP pools are skewed by
exogenous nucleosides in S. pombe if they are converted into useable nucleotides,
to create a transient G1 arrest. We propose that Br- and ethynyl-dUTP (BrdU and
EdU, respectively) or thymidine, in a hsv-tk+ strain, cause an apparent increase in
dTTP. Excess thymidine or analog may also inhibit RNR as in human cells (Meuth
and Green 1974a,b), depleting the dCTP pool (Figure 5.10G, parts 1 and 2). If so,

238

BrdU and cytidine co-treatment will suppress toxicity and mutagenesis as in
metazoans (Davidson and Kaufman 1978; Popescu 1999).
Future work will determine how dNTP pools are altered with these
nucleosides, but we predict that thymidine, BrdU, and EdU all cause a dCTP-less
state as in human cells. Interestingly, the fission yeast dCMP deaminase mutant
(dcd1Δ) has an opposite effect: levels of CTP pools become high while that of dTTP
is low. The effects on cell fitness are similar to what we observed. Consistent with
our model, dcd1Δ cells expressing hsv-tk+ increase dTTP pools, relieving growth
defects and dcd1Δ sensitivities to UV, HU, and bleomycin (Sanchez et al. 2012).
A second observation supports dNTP pool imbalance as a cause of observed
phenotypes in S. pombe hsv-tk+ hENT+ cells: while BrdU .32.6 mM does not
increase BrdU substitution in DNA, cell viability strikingly decreases above this dose.
We observe no evidence for additional BrdU substitution, which excludes further T^C
point mutations as the source of high-dose sensitivity. Instead, we propose that this
dosage sensitivity reflects increased changes in cellular dNTP pools, perhaps via
negative allosteric inhibition of RNR by analogs/thymidine (Meuth and Green
1974b). To confirm this hypothesis, measurements of cellular dNTPs and RNR
activity during BrdU and EdU are required.
Changes in dNTP pools cause point mutations in metazoans (Meuth 1989;
Phear and Meuth 1989), by substituting the nucleotide in excess during replication
(Phear and Meuth 1989). We find that BrdU treatment increases can1 forward
mutations in wild-type, spd1Δ, and cds1Δ hsv-tk+ hENT+ cells. However, the rate of
mutagenesis is not significantly changed in chk1Δ, while rad3Δ and mrc1Δ

239

experience decreased mutation. We suggest that the replication checkpoint does not
actively prevent BrdU-dependent mutation, while an active G2/M checkpoint (chk1+)
promotes mutagenesis. This may reflect the increased survival of checkpoint-intact
cells or checkpoint effects on Spd1 and nucleotide metabolism (see below). We
assume that can1- mutations are A•T ^G•C transitions caused by Br-dUTP
substitution for the lowered dCTP analog or G•C^A•T transitions via enhanced Br-
dUTP pairing with guanine in template DNA (Hopkins and Goodman 1980; Lasken
and Goodman 1984; Goodman et al. 1985) (Figure 7G, part 3). This is confirmed by
our RFLP analysis of can1 mutants, which did not show any gross structural
changes.
Fission yeast dNTP metabolism is controlled by RNR, which is inhibited by
Spd1 (Holmberg et al. 2005; Hakansson et al. 2006; Nestoras et al. 2010). Spd1 is
ubiquitinated during stress so that the dNTP pool is expanded, which ties nucleotide
metabolism to the cell cycle and DDR (Moss et al. 2010; Nestoras et al. 2010). We
found that spd1Δ cells are less sensitive to chronic nucleoside exposure, but their
relative viability in acute BrdU is worse. Intriguingly, spd1Δ cells showed a 10-fold
increase in mutations of can1+ or hsv-tk+ and frequent sectoring, indicating that
spd1Δ cells are unstable and prone to mutation. Previous work showed spd1Δ
suppresses mutation in ddb1Δ cells that are incapable of activating RNR (Holmberg
et al. 2005). However, we show that spd1Δ cells are intrinsic mutators, an effect
worsened after BrdU treatment. Increased basal dNTP pools in spd1Δ (Holmberg et
al.
2005) could promote this mutagenesis by facilitating replication during exogenous

240

thymidine/dUTP treatment and allowing additional Br-dUTP incorporation opposite G
(Lasken and Goodman 1984; 1985). Alternatively, transient changes in one dNTP
during spd1Δ replication may not prompt arrest. Our data are consistent with models
linking dNTP regulation to genome stability in S. pombe (Holmberg et al. 2005; Moss
et al. 2010; Nestoras et al. 2010). While chk1Δ hsv-tk+ hENT+ cells do not show
increased mutation post-BrdU, we hypothesize that a spd1Δ chk1Δ double mutant
may have a very high mutation rate due to extra damage and larger dNTP pools.
Rad3 is required to survive BrdU as reported in Hua and Kearsey (2011). We
show this occurs through the Chk1 G2-DDR path downstream of Rad3, resulting in
Cdc2 phosphorylation and Rad52 focus formation. BrdU also increases phospho-
histone H2A, a signal of DSBs and/or replication fork stress (e.g., Bailis et al. 2008;
Rozenzhak et al. 2010). Our model suggests that halogenated dUTP causes DNA
damage (Figure 5.10G, part 4), perhaps via removal of substituted bases by base
excision repair (BER) (Krych et al. 1979; Szyszko et al. 1983; Morgan et al. 2007).
BER under dNTP depletion could additionally cause single-strand DNA breaks
(SSBs) that stall replication forks and/or convert to DSBs during S phase. This would
promote cell accumulation in S phase and a modest requirement for the Cds1
pathway.
Alternatively, topoisomerase I has RNaseH activity on a double-strand DNA
template with only one dUTP substitution (Sekiguchi and Shuman 1997) and could
contribute to SSB and DSB accumulation. While BrdU-induced DNA damage in
human cells is documented, the cause is not known (Dillehay et al. 1984; Ackland et
al. 1988). The role of postreplication repair in surviving BrdU may prove essential.

241

As in mammals, we observe that a second DNA-damaging agent is more
dangerous after BrdU substitution. Sensitivity of chk1Δ cells to other drugs following
BrdU incorporation suggests that BrdU substitution increases the DNA damage
“load” in treated cells. In mammalian cells, UV exposure following BrdU substitution
causes DSBs and interstrand cross-links (Murray and Martin 1989; Cecchini et al.
2005) and enhanced sensitivity to bleomycin (Ackland et al. 1988) and cisplatin
(Russo et al. 1986). Budding yeast is also UV sensitive after BrdU exposure
(Sclafani and Fangman 1986). Some part of this sensitivity may result from S-phase
slowing in BrdU, a more vulnerable time for DNA damage. We show increased
sensitivity in cds1Δ strains, implying that replication fork stability is diminished,
perhaps from imperfect BrdU base pairing (Lasken and Goodman 1985). We find
that rad3Δ response to CPT, UV, MMS, or HU is unchanged with BrdU pretreatment,
probably because of catastrophic failure of rad3Δ in these drugs.
Our results point to challenges in the use of nucleoside analogs to analyze
DNA replication. While we do not directly compare between the two analogs, our
results indicate that fission yeast is extremely sensitive to nucleoside analogs BrdU
and EdU, when treated at doses similar to those used in human cells. Consistent
with replication stability problems, the increased toxicity of EdU, and cellcycle effects
at lower doses, may reflect a larger ethynyl side group and thus greater steric
interference during replication. The long-term effects associated with BrdU and EdU
exposure mean that these analogs are most useful for analysis in a single cell cycle.
Further, thymidine may be a potential reversible blocking agent in S. pombe, yet its
effects must be more clearly described. In all cases, appropriate care must be taken

242

to mitigate analog effects, lest disruption of nucleotide levels interfere with the very
process under study.

5.5 Materials and Methods
5.5.1 Yeast strains, analog addition, growth, and mutagenesis
Fission yeast strains are described in Table 5.3. Cells were grown as
described in Sabatinos and Forsburg (2010), and BrdU (5 mg/ml in water) or EdU
(10 mM stock; Invitrogen, Carlsbad, CA) was added as required to liquid cultures or
plates. Media were yeast extract with supplements (YE5S, hereafter “YES”),
Edinburgh minimal medium (EMM– ammonium chloride nitrogen source), or S.
pombe minimal glutamate medium (PMG– sodium glutamate nitrogen source)
(Sabatinos and Forsburg 2010). Physiology experiments in- cluding Chk1 protein,
Rad52 foci, mutagenesis, and flow cytometry were all performed in liquid EMM
cultures. Protein extracts were prepared from 0.3 M NaOH-treated cells and lysed in
2· SDS–PAGE sample buffer by boiling for 5 min, as described in Sabatinos and
Forsburg (2010). The choice of HA monoclonal produced a different nonspecific
background band location on Chk1-HA blots (Roche 12CA5 or Covance 16B12).
Mutation rate was determined by splitting cultures (6BrdU), treating them with
32.6 mM BrdU for 2 hr, and then plating them on YES and PMG + canavanine.
Canavanine plates (70 mg/ml in PMG+ supplements) were scored after 8 days
growth (32) and numbers of Can1
-
colonies were compared to the concentration
derived from titer plates (YES). Grouped experiments for Table 5.1 and Figure 7

243

were performed independently and rates calculated using FALCOR
(http://www.keshavsingh.org/protocols/FALCOR.html) and the Ma–Sandri–Sarkar
Maximum-Likelihood Estimator (MSS-MLE) or the Lea–Coulson method of the
median. MSS-MLE results were analyzed by a two-tailed t-test, and Mann–Whitney
U-tests were used for Lea–Coulson significance testing
(http://vassarstats.net/utest.html). Frequency of hsv-tk
+
loss was scored as the
number of FUdR-resistant or sectored colonies per total, and the proportions were
assessed with Z-tests (http://vassarstats.net/propdiff_ind. html).
Table 5.3: Fission yeast strains used in this study.
Strain Genotype Source
FY2317
h+ leu1-32::hENT1-leu1+(pJAH29) his7-366::hsv-tk-
his7+(pJAH31) ura4-D18 ade6-M210
Hodson et
al. (2003)
FY3179
h+ mrc1Δ::ura4+ leu1-32::hENT1-leu1+(pJAH29)
his7-366::hsv-tk-his7+(pJAH31) ura4-D18 ade6-M210 This study
FY3454 h+ ura4-D18 ade6-M210 This study
FY5148
h+ cds1Δ::ura4 leu1-32::hENT1-leu1+(pJAH29) his7-
366::hsv-tk-his7+(pJAH31) ura4-D18 ade6-M210 This study
FY5149
h+ chk1Δ::ura4+ leu1-32::hENT1-leu1+(pJAH29) his7-
366::hsv-tk-his7+(pJAH31) ura4-D18 ade-704 This study
FY5150
h+ rad3Δ::ura4+ leu1-32::hENT1-leu1+(pJAH29)
his7-366::hsv-tk-his7+(pJAH31) ura4-D18 ade6-M210 This study
FY5155
h2 cds1Δ::ura4+ pola-FLAG::ura4+ rad11-
myc::KanMX6 rad22-YFP::natMX leu1-32::[hENT-
leu1+] his7-366::[hsv-tk his7+] ura4-D18 ade6-M210 This study
FY5159
h- pola-FLAG::ura4+ rad11-myc::KanMX6 rad22-
YFP::natMX leu1-32::[hENT-leu1+] his7-366::[hsv-tk
his7+] ura4-D18 ade6-M210 This study
FY5030
h- cds1-13myc::KanMX leu1-32::[hENT leu1+] his7-
366::[his7+] ade6-M210 ura4-D18 This study
FY5031
h+ cds1-13myc::KanMX chk1HA leu1-32::[hENT
leu1+] his7-366::[hsv-tk his7+] ade6-M216 ura4-D18 This study
FY6247
h+ Δspd1::ura4+ ura4-D18 leu1-32::hENT1-
leu1+(pJAH29) his7-366::hsv-tk-his7+(pJAH31) ade6- This study

244

5.5.2 Microscopy
Cells were fixed in cold 70% ethanol for cell-cycle analysis or microscopy.
Cells were rehydrated in water and incubated for 10 min in 1 mg/ml of aniline blue
[Sigma (St. Louis) M6900]. Cells in mount (50% glycerol, 1 mg/ml DAPI, and 1
mg/ml p-phenylenediamine) were photographed on a Leica DMR wide-field
epifluorescent microscope, using a 63· ob- jective lens (NA 1.32 Plan Apo), a 100-W
Hg arc lamp for excitation, and a 12-bit Hamamatsu ORCA-100 CCD cam- era.
OpenLab v3.1.7 (Improvision, Lexington, MA) software was used at acquisition and
ImageJ (Schneider et al. 2012) for analysis. DAPI counterstaining did not
significantly affect BrdU or EdU signal intensity.
Live-cell Rad52-YFP foci were imaged on a DeltaVision Core microscope
(with softWoRx v4.1; Applied Precision, Issaquah, WA), using a 60· (NA1.4 Plan
Apo) lens, a solid- state illuminator, and a 12-bit Roper CoolSnap HQII CCD camera.
The system’s x-y pixel size was 0.1092 mm. YFP fluorescence for single time points
was acquired as eighteen 0.3-mm z-sections. Long-term time-lapse movies used
nine z-steps of 0.5 mm. Rad52-YFP images were deconvolved and maximum-
intensity projected (softWoRx). Movies were per- formed in CellAsics (Hayward, CA)
microfluidics plates (Y04C series), with supplemented EMM medium. Transmit- ted
light images were fused with DAPI or Rad52-projected images. Images were
contrast adjusted using a histogram stretch and equivalent scale in all samples. A
threshold of 2· over the average nuclear YFP signal was used for focus
discrimination. Rad52 foci are presented as the proportion of nuclei with Rad52 foci
695% confidence interval (C.I.). Significance was assessed with chi-square tests.

245

5.5.3 Flow cytometry
Whole-cell SytoxGreen flow cytometry (FACS) was per- formed as described
in Sabatinos and Forsburg (2009). Whole-cell FACS for EdU was performed using
Click-iT (Invitrogen) with AlexaFluor 488 on rehydrated cells. FACS for BrdU was
performed on “ghosts” (Carlson et al. 1997), prepared by spheroplasting cells in 0.1
M KCl with 5 mg/ml Zymolyase 20T and then 1% Triton X-100 before sonicating to
release nuclei. Nuclei were blocked (10% FCS, 1% BSA) for 30 min and then
incubated with mouse anti-BrdU (Becton Dickinson; B44). Secondary antibody was
Alexa- 488–conjugated anti-mouse (Invitrogen), and nuclei were counterstained with
propidium iodide for total DNA content.

246

5.6 Legends for Movies
Movie 5.1 Wild-type hsv-tk+ hENT+ cells in a microfluidics chamber were treated
with 32.6µM BrdU for 3h (pink border) and switched to BrdU-free medium for 3h
afterward, to monitor Rad52-‐YFP foci (yellow).
Movie 5.2 cds1Δ hsv-tk+ hENT+ cells in a microfluidics chamber were treated with
32.6µM BrdU for 3h (pink border) and switched to BrdU-free medium for 3h
afterward, to monitor Rad52-‐YFP foci (yellow).
Movie 5.3 Wild-type hsv-tk+ hENT+ cells in a microfluidics chamber were treated
with 10µM EdU for 3h (pink border) and switched to EdU-free medium for 3h
afterward, to monitor Rad52-‐YFP foci (yellow).
Movie 5.4 cds1Δ hsv-tk+ hENT+ cells in a microfluidics chamber were treated with
10µM EdU for 3h (pink border) and switched to EdU-free medium for 3h afterward,
to monitor Rad52-‐YFP foci (yellow).

247

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Abstract (if available)

Abstract Mechanisms that maintain genome stability are essential for human health. Loss of genome stability is associated with cancer and birth defects. This dissertation uses a model fission yeast system to investigate how cells preserve chromosome integrity during the specialized differentiation process of meiosis. In this work mutants sensitive to alkylation damage were examined for their ability to proceed through meiosis. It was found that these genes contribute to meiotic progression. However, their contribution is not isolated to a single process or mechanism. Rather it was seen that these mutants that are sensitive to meiotic alkylation damage have a diverse role in meiotic progression.

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Creator Mastro, Tara (author)

Core Title Response to alkylation damage linked to meiotic progression

School College of Letters, Arts and Sciences

Degree Doctor of Philosophy

Degree Program Molecular Biology

Publication Date 02/18/2015

Defense Date 01/07/2015

Publisher University of Southern California (original), University of Southern California. Libraries (digital)

Tag checkpoint,Chromosomes,DNA damage,genetics,meiosis,OAI-PMH Harvest,S. pombe

Format application/pdf (imt)

Language English

Contributor Electronically uploaded by the author (provenance)

Advisor Forsburg, Susan L. (committee chair), Craft, Cheryl M. (committee member), Goodman, Myron F. (committee member), Michael, Matthew (committee member)

Creator Email mastro@usc.edu,tara.mastro@gmail.com

Permanent Link (DOI) https://doi.org/10.25549/usctheses-c3-535465

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