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Destabilization of the replication fork protection complex is associated with meiotic defects in fission yeast

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Destabilization of the replication fork protection complex is associated with meiotic defects in fission yeast

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DESTABILIZATION OF THE REPLICATION FORK PROTECTION
COMPLEX IS ASSOCIATED WITH MEIOTIC DEFECTS IN
FISSION YEAST

By: Wilber Escorcia Torres

December 2016

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)

i

Acknowledgements
First and foremost, I would like to thank my mother, Cecilia Torres, for always
supporting me unconditionally. She has taught me to work hard and to persevere through
hardship. She is an excellent role model and no academic degree will ever match the broad
education in life she has provided to me. I would also like to thank my wife, Wendy Alvarez, for
being incredibly understanding of my line of work and for loving me even when this thesis took
over my body and soul. Her letters of encouragement have always given me an emotional boost
more potent than any caffeine-infused product I could procure. I am vastly thankful for having
Susan L. Forsburg as my academic advisor. Her patience and committed mentorship transformed
me into the budding scientist I am today. I would also like to extend my gratitude to my step
father, Marcos A. Turcios, for his steadfast belief in my abilities. His excellent driving saved me
several times from being late to final exams. I would also like to thank my in-laws, Francisco
Alvarez and Maria Alvarez, for being amazing individuals who have provided a nourishing
atmosphere at home in which I could thrive personally. I am also grateful for everything Frankie
Alvarez, my brother-in-law, has done for Wendy and me through the process of completing this
thesis.
I am equally indebted to my thesis committee members for all the guidance they provided
to me through the years. Sean Curran’s sideline talks have inspired me to learn from mistakes
proficiently and quickly move on to more promising ventures. Matt Michael’s questions and
suggestions have greatly helped me to address inquiries I had not anticipated. His input has been
key in allowing me to see many a blind spot in my work and thinking. Andrea Armani has been
extremely generous with her time and common sense. Her recommendation on how to
effectively manage vacation time so it does not affect one’s work output upon returning to the
ii

bench is one life hack I will always remember. Moreover, I would like to thank other USC
faculty for their contribution to my academic development. George Sanchez has been
instrumental in facilitating funding for my Ph.D. for five years. His goal of enhancing diversity
at USC has positively impacted my and the lives of many other minority students. Albert Herrera
has given me the opportunity during two consecutive summers to hone my teaching and science
communication skills. I feel less intimidated while giving talks before my peers due in large part
to his excellent career advice. Bob Baker is by far the best counselor one could ever ask for.
While 1960s scientists had Kennedy to thank for getting them fascinated with science, I have
Bob Baker. Our many conversations have allowed me to fully appreciate the intrinsic value of all
the work I do at bench.
I am indebted to everyone I met during grad school, but some individuals deserve
specific mention, for their influence in my work is invaluable. Amanda M. Jensen has been a
terrific friend since she recruited me to USC and to Susan’s lab. Our talks over the years have
enabled me to gain extraordinary personal and professional insights. I could not have figured out
many science problems nor could I have verbally communicated my work in understandable
ways without the help of her sharp, quick mind. Nimna Ranatunga is not only the lab’s unofficial
record keeper, but also a fantastic friend on whose lab experience one could always rely to
troubleshoot impossible experiments. Yael Freiberg has been a phenomenal source of
constructive criticism. Over the years, we have held numerous interesting conversations ranging
from the scientific to the mundane. Her presence during the lab’s night shift hours has been an
essential factor in breaking the monotony of many grueling whole-day experiments.
Kuo-Fang Shen’s mastery of microscopy analysis has greatly enhanced the quality of the
work presented here. Though he is an Ohio State football fan, I have enjoyed our various talks
iii

about college sports and authentic Taiwanese food. Ji-Ping Yuan is an indispensable component
of the Forsburg lab. Without his help, my work would have been excruciatingly difficult. Tara
Mastro’s mentorship has taught me most of what I know about fission yeast meiosis. Her
impetus as a senior graduate student gave me the motivational boost to focus on my thesis work
more efficiently. Sarah Sabatinos provided me with an excellent primer on fission yeast genetics
and physiology. Her superb instruction has given me the tools to become a more meticulous and
observant researcher. Marc Green was the lab’s microscope master and with his training I have
been able to ask and address yeast genetics questions that have fueled the progress of most of my
research. Ruben Petreaca is a remarkable human and a true older brother to me, albeit from
Romanian parents. His wise words have resonated well in my head and thanks to them I feel
more mature about my professional and personal life. I often miss our cigar-fueled conversations
on Plato’s theory of forms and spooky physic theories. Ashley Yi, my undergraduate mentee, is
an academically and technically gifted young researcher whose hard work in lab has made a
tremendous difference in my push to finish this thesis. Lastly, the MCB class of 2010 has made
my Ph.D. experience quite memorable. Thank you Melina Butuci, Jordan Eboreime, Ana
Carolina Dantas Machado, Jackie Lo, and Hank Cheng for being amazing class mates, studying
partners, and tail gate drinking buddies. Grad school would have been truly awful without your
shenanigans. Now, let’s drink to finishing up!
And to those whom I forgot to mention here, please know that I am thankful for your help
and support. I shall, in due time, say so personally!

iv

Table of Contents

Acknowledgements i
List of Abbreviations vi
List of Tables vii
List of Figures viii
List of Movies ix
Abstract xi

Chapter 1: Introduction 1
1.1 Meiosis 1
1.2 The fork protection complex (FPC) 13
1.3 Chapter 1 Bibliography 20
Chapter 2: Destabilization of the replication fork protection complex
is associated with meiotic defects. 31
2.1 Overview 31
2.2 Introduction 31
2.3 Results 36
2.4 Discussion 54
2.5 Materials and methods 63
2.6 Supplemental data 68
2.7 Legends for movies 72
2.8 Chapter 2 Bibliography 73
Appendix 1: Tetrad dissection in fission yeast 81
1.1 Overview 81
1.2 Introduction 81
1.3 Materials 82
1.4 Methods 84
v

1.5 Notes 89
1.6 Appendix 1 Bibliography 92
Appendix 2: Random spore analysis in fission yeast 93
2.1 Overview 93
2.2 Introduction 93
2.3 Materials 95
2.4 Methods 97
2.5 Notes 100
2.6 Appendix 2 Bibliography 101
Appendix 3: Genetic screen for suppressors of dfp1-rad35,
a truncation allele of dfp1+ with meiosis-specific defects 103
3.1 Overview 103
3.2 Introduction 103
3.3 Results 106
3.4 Discussion 106
3.5 Materials and methods 106
3.6 Appendix 3 Bibliography 108

vi

List of Abbreviations
Ana I Anaphase I
Ana II Anaphase II
ChIP Chromatin immunoprecipitation
CO Crossover
HJ Holiday Junction
DAPI 4',6-diamidino-2-phenylindole
DSB Double strand break
EMM Edinburgh minimal media
FACS Fluorescent assisted cell sorting
HR Homologous recombination
HT Horse tailing
HU Hydroxyurea
IR Ionizing radiation
MeiS Pre-meiotic S-phase
MI Meiosis I
MII Meiosis II
MMS Methyl methanesulfonate
MT Metaphase
NCO Non-crossover
PCR Polymerase chain reaction
PFGE Pulse field gel electrophoresis
PMG Pombe glutamate medium
prDSB Programmed double strand breaks
S-phase DNA synthesis phase
SC Synaptonemal complex
SCE Sister chromatid exchange
UV Ultraviolet radiation
YES Yeast extract plus supplements

vii

List of Tables
Table 2.1 Recombination dynamics in FPC mutants. 43
Supplemental Table 2.1 List of strains used in this work 70

viii

List of Figures
Figure 1.1 Meiosis is a cell division program that reduces ploidy. 1
Figure 1.2 Cytological stages of fission yeast during meiosis. 7
Figure 1.3 Dynamics of chromosome segregation in meiosis. 10
Figure 1.4. The FPC is required for replication at various genomic regions. 14
Figure 1.5. Destabilization of the fork leads to DNA damage in FPC mutants. 19
Figure 2.1 Spore viability and replication dynamics in FPC mutants. 38
Figure 2.2 Persistence of DNA damage in FPC mutants. 41
Figure 2.3 Meiotic segregation in FPC mutants. 45
Figure 2.4 MI equational division in FPC mutants. 47
Figure 2.5 Non-disjunction in MI and MII in FPC mutants. 50
Figure 2.6 Elimination of the linear elements in FPC mutants. 51
Figure 2.7 Duration of Sgo1 in metaphase in FPC mutants. 53
Figure 2.8 Persistence of centromeric cohesion in FPC mutants. 55
Figure 2.9 Recruitment of heterochromatin to the centromere in FPC mutants. 57
Figure 2.10 Model of FPC contribution to meiotic segregation. 60
Supplemental Figure 2.1 Replication dynamics in FPC mutants lacking Rec12. 68
Supplemental Figure 2.2 Chromosome fragmentation in FPC mutants. 69
Figure 3.1 Tetrad dissection diagram. 87
Figure 3.2 Tetrad types resulting from recombination between two linked loci. 88
Figure 3.3 Fate of dissected tetrad products. 91
Figure 4.1 Counting and plating spores for RSA. 99
Figure 5.1. Schematic of genetic screen for suppressors of dfp1-rad35. 104
Figure 5.2. Suppressors of dfp1-rad35. 107

ix

List of movies
Movies can be viewed and/or downloaded at pombe.net
Movie 2.1: Representative live cell imaging of meiosis in a WT heterothallic cross of cells
homozygous for RPA-YFP, Rad52-CFP, and H3-mRFP.
Movie 2.2: Representative live cell imaging of meiosis in a ∆swi1 heterothallic cross of cells
homozygous for RPA-YFP, Rad52-CFP, and H3-mRFP.
Movie 2.3: Representative live cell imaging of meiosis in a ∆swi3 heterothallic cross of cells
homozygous for RPA-YFP, Rad52-CFP, and H3-mRFP.
Movie 2.4: Representative live cell imaging of meiosis in a WT heterothallic cross of cells
homozygous for histone H3-mRFP.
Movie 2.5: Representative live cell imaging of meiosis in a ∆swi1 heterothallic cross of cells
homozygous for histone H3-mRFP.
Movie 2.6: Representative live cell imaging of meiosis in a ∆swi3 heterothallic cross of cells
homozygous for histone H3-mRFP.
Movie 2.7: Representative live cell imaging of meiosis in a WT heterothallic cross of cells
heterozygous for LacI-GFP and lacO near centromere I and homozygous for H3-mRFP.
Movie 2.8: Representative live cell imaging of meiosis in a ∆swi1 heterothallic cross of cells
heterozygous for LacI-GFP and lacO near centromere I and homozygous for H3-mRFP.
Movie 2.9: Representative live cell imaging of meiosis in a ∆swi3 heterothallic cross of cells
heterozygous for LacI-GFP and lacO near centromere I and homozygous for H3-mRFP.
Movie 2.10: Representative live cell imaging of meiosis in a WT heterothallic cross of cells
homozygous for LacI-GFP and lacO near centromere I as well as for H3-mRFP.
Movie 2.11: Representative live cell imaging of meiosis in a ∆swi1 heterothallic cross of cells
homozygous for LacI-GFP and lacO near centromere I as well as H3-mRFP.
Movie 2.12: Representative live cell imaging of meiosis in a ∆swi3 heterothallic cross of cells
homozygous for LacI-GFP and lacO near centromere I as well as H3-mRFP.
Movie 2.13: Representative live cell imaging of meiosis in a WT heterothallic cross of cells
homozygous for Rec27-GFP and H3-mRFP.
Movie 2.14: Representative live cell imaging of meiosis in a ∆swi1 heterothallic cross of cells
homozygous for Rec27-GFP and H3-mRFP.
Movie 2.15: Representative live cell imaging of meiosis in a ∆swi3 heterothallic cross of cells
homozygous for Rec27-GFP and H3-mRFP.
Movie 2.16: Representative live cell imaging of meiosis in a WT heterothallic cross of cells
homozygous for Sgo1-GFP and H3-mRFP.
x

Movie 2.17: Representative live cell imaging of meiosis in a ∆swi1 heterothallic cross of cells
homozygous for Sgo1-GFP and H3-mRFP.
Movie 2.18: Representative live cell imaging of meiosis in a ∆swi3 heterothallic cross of cells
homozygous for Sgo1-GFP and H3-mRFP.
Movie 2.19: Representative live cell imaging of meiosis in a WT heterothallic cross of cells
homozygous for Rec8-GFP and H3-mRFP.
Movie 2.20: Representative live cell imaging of meiosis in a ∆swi1 heterothallic cross of cells
homozygous for Rec8-GFP and H3-mRFP.
Movie 2.21: Representative live cell imaging of meiosis in a ∆swi3 heterothallic cross of cells
homozygous for Rec8-GFP and H3-mRFP.
Movie 2.22: Representative live cell imaging of meiosis in a WT heterothallic cross of cells
homozygous for Swi6-GFP and Cnp1-mCherry.
Movie 2.23: Representative live cell imaging of meiosis in a ∆swi1 heterothallic cross of cells
homozygous for Swi6-GFP and Cnp1-mCherry.
Movie 2.24: Representative live cell imaging of meiosis in a ∆swi3 heterothallic cross of cells
homozygous for Swi6-GFP and Cnp1-mCherry.

xi

Abstract
In addition to its essential function in replication initiation, the DDK protein kinase plays
a role in the response to replication stress. A truncation allele in the Dfp1 subunit of DDK is
proficient for replication, but is MMS sensitive with defects in induced mutagenesis. It is also
defective at multiple steps in meiosis including the initiation of double strand breaks,
transcription, and cleavage of Rec8 (the alpha-kleisin subunit of cohesin), accompanied by a
severe loss of viability. We have taken a candidate approach to examine meiosis in mutants
affecting DDK-associated proteins that also affect MMS sensitivity and fork stability. In this
thesis, we examine components of the replication Fork Protection Complex (FPC): Swi1 (Tof1)
and Swi3 (Csm3). We show that destabilization of the FPC decreases spore viability, increases
sister chromatid exchange, reduces homologous recombination, and induces abnormal
segregation in MI and MII divisions. We find that the elimination of Rec27 (a linear elements
component), duration of Sgo1 before anaphase I and persistence of centromeric Rec8 are
disrupted in FPC mutants. Moreover, as they progress though meiosis, FPC mutants show Rad11
(RPA) and Rad22 (Rad52) signals that indicate persistent DNA damage. Thus, the FPC stability
is required for proper meiotic segregation. Our results suggest that the FPC suppresses excess
ssDNA and helps to stabilize cohesion at the centromere, indicating that events in meiosis are
sensitive to disruptions in replication fork stability.

1

Chapter 1: Introduction
1.1 Meiosis
Meiosis is a cell division program that generates gametes (spores in the yeasts) carrying
half the genetic content of progenitor cells. By contrast, mitosis produces daughter cells with
identical amounts of DNA as parent cells. The difference between these processes stems from
the need to reduce ploidy in cells that participate in sexual reproduction. Without meiosis, there
would be an untenable expansion of chromosome number from one generation to the next.
Circumventing ploidy amplification requires that two consecutive nuclear divisions follow one
round of DNA replication. While reciprocal exchange of genetic material between homologous
chromosomes makes meiosis indispensable for the creation of genetic diversity, it is also
essential for the proper distribution of chromosomes to gametes (Figure 1.1) (Zickler and
Kleckner 1999; Cavalier-Smith 2002; Hochwagen 2008; Ohkura 2015).

Figure 1.1 Meiosis is a cell division program that reduces ploidy. Whereas mitosis generates two daughter cells
with similar amounts of genetic material as the parent cell, meiosis produces four daughter cells with half the
parental genetic content. This results from two nuclear divisions that follow one round of DNA replication.
Generation of DSBs and subsequent recombination between homologues ensures proper reductional division (MI).
Persistence of centromeric cohesion is essential to guarantee correct equational division (MII), where sister
chromatids segregate similarly as they do in mitosis. Diagram was adapted from Hochwagen 2008.
2

Homologous recombination sets meiosis apart from mitosis because it calls for the
programmed execution of DNA double strand breaks (DSBs), the repair of which influences the
correct segregation of chromosomes during the first meiotic division (MI). The physical
connection between homologous chromosomes ensures their correct bi-orientation at the
metaphase plate as well as their subsequent segregation to opposite poles of the cell. Whereas in
mitosis they separate after replication, in meiosis sister chromatids are held together by cohesin
until the second nuclear division (MII). The gradual loss of cohesion that first occurs on
chromosome arms before MI and then at the centromeres preceding MII distinguishes meiotic
from mitotic segregation, for it makes use of components specific to meiosis that safeguard sister
chromatid segregation until the right conditions for cell division are met (Zickler and Kleckner
1999; Davis and Smith 2001; Marston and Amon 2004; Miller et al. 2013).
In the yeasts, nutrient deprivation is crucial to induce cells to enter meiosis (Yamamoto et
al. 1996; Honigberg et al. 2003; Wu and Nurse 2014). In fission yeast, nutrient depletion
activates Atf1/Pcr1 (a pair of regulatory transcription factors) and downregulates the kinase
Pka1. These two events increase Ste11 levels, which in conjunction with the mating pheromone
signaling pathway, promote the production of Mei3. The meiotic program is then initiated when
Mei3 disables the phosphorylation-dependent repression of Mei2 (an RNA-binding protein) by
the kinase Pat1 (Watanabe et al. 1994; Li et al. 1996; Davis and Smith 2001; Kitamura et al.
2001; Watanabe et al. 2002; Harigaya et al. 2006).
Once cells have committed to enter meiosis from the G1-phase, they engage in DNA
replication. The meiotic budding yeast CDKs along with Clb5 and Clb6 prompt the start of meiS
phase in a manner resembling pre-mitotic S-phase. Not only are similar kinases used, but the
same origins of replication are also employed. This can perhaps explain why meiotic and mitotic
3

replication rates in this yeast are, to a large degree, comparable (Collins et al. 1994; Dirick et al.
1998; Stuart et al. 1998; Benjamin et al. 2003; Offir et al. 2004; Kohl et al. 2013). The same is
not true for other organisms, however. The duration of meiS phase in most higher eukaryotes as
well as in fission yeast can be significantly longer. Differing lengths of replication can be
attributed to factors that are unique to meiotic events as well as to how organisms align and
segregate their chromosomes; processes that are closely linked to DNA synthesis (Cha et al.
2000; Forsburg and Hodson 2000; Murakami and Nurse 2001; Wu and Nurse 2014).
In many aspects, pre-meiotic and pre-mitotic DNA replication function similarly. In both
yeasts, the two cell division programs make use of mini-chromosome maintenance (MCM)
proteins to set up the pre-replication complex (pre-RC) and employ polymerases α, δ, ε and other
replication factors (including Swi1/Tof1 and Swi3/Csm3) to carry out DNA synthesis (Hopwood
and Dalton 1996; Zou and Stillman 1998; Matsumoto et al. 2002). While in the yeasts cell
division cycle 45 (Cdc45) is necessary for mitotic replication initiation (Simchen 1974; Aparicio
et al. 1999; Lindner et al. 2002; Matsumoto et al. 2011), its orthologues in Xenopus laevis and
Arabidopsis thaliana (CDC45 in both) are essential for efficient loading of Polα onto DNA and
proper flower bud formation, respectively (Ortega et al. 2003; Stevens et al. 2004; Hashimoto et
al. 2012). Without CDC45, A. thaliana plants exhibit embryonic lethality and its depletion leads
to fragmented chromosomes, which reduce fertility significantly. Since these phenotypes likely
stem from incomplete DNA replication, they suggest that the meiS phase is equally crucial for
correct chromosome segregation as is pre-mitotic replication (Ortega et al. 2003; Stevens et al.
2004; Huang et al. 2016).
DNA replication is also a crucial step for homologous chromosomes to physically
interact via meiotic recombination, chromosome alignment, and synaptonemal complex (SC)
4

formation, a structural change that further stabilizes pairing of homologues (Borde et al. 2000;
Smith et al. 2001). Following bulk DNA synthesis, the transesterase sporulation protein Spo11
creates DSBs (Keeney et al. 1997; Borde et al. 2000; Cervantes et al. 2000). In the absence of
budding yeast replisome components Tof1 and Csm3, DSB formation is compromised. The same
is true for cells in which DNA synthesis is reduced by the elimination of Clb5 and Clb6,
suggesting that reduction in DSB formation is proportional to the extent DNA synthesis is
decreased (Stuart et al. 1998; Borde et al. 2000; Smith et al. 2001; Murakami and Keeney 2014).
Fission yeast differs in this respect, as compromised replication does not entirely abrogate
generation of DSBs by the Spo11 orthologue Rec12. In fact, DSB levels in cells where DNA
synthesis is disrupted remain significant. Unlike budding yeast, fission yeast does not have a true
SC. This may explain the different dependencies on replication that each yeast has for DSB
formation (Cha et al. 2000; Cervantes et al. 2000; Murakami and Nurse 2001). A shared
characteristic among various species including both yeasts is the requirement for meiotic
replication to establish sister chromatid cohesion. The importance of this step is evident in the
absence of Rec8 (a meiotic cohesin subunit) where cells show problems with chromosome
pairing, recombination, and DSB formation. Consequently, replication stability in meiS phase is
crucial for the proper function of downstream processes that ensure correct nuclear division
(Molnar et al. 1995; Uhlmann and Nasmyth 1998; Klein et al. 1999; Watanabe et al. 2001;
Murakami and Keeney 2014; Wu and Nurse 2014).
DSBs in meiosis differ from those in mitosis in that the former are programmed
(pmDSBs) and are necessary for meiotic recombination, whereas the latter result from insults to
DNA such as ionizing radiation or, more commonly, from replication failure (Keeney et al.
1997). Although there are differences among species in the way DSBs are processed, they
5

proceed via a recombination program that is mostly conserved in eukaryotic organisms.
Following DSB formation by Spo11, the MRN complex, CtIP, Exo1, and Dna2 carry out
resection of the 3’ end (Symington and Gautier 2011; Youds and Boulton 2011; Huang et al.
2016). The resulting ssDNA is coated by RPA. Rad52 replaces RPA and promotes the binding of
Rad51 (a RecA orthologue), while simultaneously preventing the association of Dmc1 (another
RecA orthologue). Swi5-Sfr1 facilitates the loading of Dmc1, thereby enabling the participation
of Rad51-Dmc1 in the invading nucleoprotein filament (Murayama et al. 2013). Rad54
stimulates strand invasion in areas of high sequence homology (Youds and Boulton 2011).
Though Rdh54 (a Rad54 paralogue) has overlapping functions with Rad54, it also helps to
stabilize Dmc1 assembly (Catlett and Forsburg 2003; Nimonkar et al. 2012). The divergent roles
in these recombination proteins is important because it determines many of the attributes that
define partner choice. For instance, Rad51-Rad54 appear to promote intersister exchange at
recombination hotspots, while Dmc1-Rdh54 facilitate interhomologue recombination at
recombination cold spots. The interplay between these two choices defines how DSBs are
repaired (Catlett and Forsburg 2003; Hyppa et al. 2008; Hyppa and Smith 2010; Youds and
Boulton 2011; Nimonkar et al. 2012).
The meiotic phase that encompasses recombination is characterized by low meiotic CDK
activity and the formation of chiasma between homologous chromosome pairs. The
chromosomal linkage required for efficient segregation in MI begins with DNA breaks induced
by Spo11 (Keeney and Kleckner 1997). These breaks are processed after DNA synthesis by
recombination pathways leading to crossovers (COs, which involve reciprocal exchange of
DNA) and non-crossovers (NCOs). Only the first pathway, however, results in chiasma.
Disruptions in this pathway not only reduce links between homologues, but also interfere with
6

correct chromosome migration during anaphase I (Zickler and Kleckner 1999; Pâques and
Harber 1999; Bishop and Zickler 2004). In budding yeast, there is a mechanism in place that
monitors the extent to which DSBs are repaired. This recombination or pachytene checkpoint
arrests cells at steps preceding MI by blocking CDK activation and silencing meiotic gene
expression (Bailis and Roeder 2000; Murakami and Nurse 2000). The checkpoint is implemented
by Swe1’s (Wee1) inhibition of Cdc28 activity, inactivation of Ndt80 (a meiotic transcription
factor) and the downregulation of CLB genes by the meiotic repressor Sum1 (Leu and Roeder
1999; Tung et al. 2000; Pak and Segall 2002). In fruit fly oocytes, arrest in response to
unrepaired DSBs is induced through the phosphorylation of BAF by NHK-1, which effectively
blocks chromosome clustering during the post-recombination period (Lancaster et al. 2010).
Though not solely responding to recombination status, germ cells of metazoan females are also
equipped with a program that halts meiotic progression at the diplotene stage. Oocyte maturation
resumes only after the organism is ready for reproduction (Kishimoto 2003).
Once cells have completed DNA replication, DSB formation, and recombination, they are
poised to undergo nuclear division. The first meiotic division (MI) is reductional because it
decreases ploidy number by half (Ohkura 2015). Contrary to MII and mitotic equational
divisions where sister chromatids separate, in MI homologous chromosomes segregate to
opposite sides. To achieve this, three crucial requirements must be fulfilled. First, chiasmata
must link homologues to ensure their correct alignment on the MI spindle (Zickler and Kleckner
1999). Second, centromeric cohesion must be preserved between sister chromatids until the onset
of MII to keep their alignment intact and prevent their premature separation (Ishiguro and
Watanabe 2007). Third, sister kinetochores must attach to microtubules radiating from one
7

spindle pole body (SPB) in MI and from two in MII (Yamagishi et al. 2014) (See Figure 1.2 for
fission yeast meiotic cytological progression).

Figure 1.2 Cytological stages of fission yeast during meiosis. Following nutrient deprivation, fission yeast haploid
cells of opposite mating type conjugate and combine their genetic material through nuclear fusion. DNA replication
occurs during a period that overlaps late nuclear fusion and early horse tailing, a series of telomere-led oscillations
necessary for chromosome pairing. Alignment of homologous chromosomes and DSB formation encompass the
majority of horsetailing. Metaphase is characterized by the end of oscillatory movement and the beginning of
nuclear compaction. At this stage, crossover recombination is completed. Division of the main nuclear mass happens
in meiosis I, where homologous chromosomes segregate to opposite sides. Likewise, an additional round of nuclear
division occurs during sister chromatid separation in meiosis II. Finally, sporulation marks the end of fission yeast
meiosis.
Anaphase I is preceded by the step-wise removal of cohesion along chromosome arms.
This transition is regulated by the meiosis-specific subunits of cohesin, the protein complex
composed of Smc1, Smc3, Scc1, and Scc3 that binds sister chromatids together following DNA
replication (Klein et al. 1999; Ishiguro and Watanabe 2007). Rad21 (fission yeast Scc1
orthologue) is replaced by Rec8 and Pcs3 (Scc3 orthologue) by Rec11, except at the centromere
(Watanabe and Nurse 1999; Parisi et al. 1999; Kitajima et al. 2003). The dual substitution of
these cohesin subunits is pivotal in promoting recombination activity, as casein kinase 1 (CK1)
phosphorylation of Rec11 enables its interaction with Rec10, which recruits Rec25 and Rec27
(the other linear elements, LinEs) to the axes of homologous chromosomes. Together, LinEs
8

stimulate the assembly of the DSB apparatus, which generates DNA cuts through the action of
Rec12 (Sakuno and Watanabe 2015; Phadnis et al. 2015). This step is crucial for later
segregation events, since cells lacking Rec8 and Rec12 show important reductions in
homologous recombination and chiasma formation (DeVeaux and Smith 1994; Krawchuk et al.
1999; Davis and Smith 2001; Molnar et al. 2001; Ellermeier and Smith 2005; Hirose et al. 2011;
Phadnis et al. 2015). Contrary to some mammals including humans, which depend on the
prophase pathway for cohesion elimination from chromosome arms, both yeasts, round worm,
and mouse rely on the cleavage activity of separase (fission yeast Cut1) (Buonomo et al. 2000;
Sumara et al. 2002; Losada et al. 2002; Davis et al. 2002; Herbert et al. 2003; Kitajima et al.
2003). During metaphase I of fission yeast, when chromosomes align on the MI spindle, the
APC/C targets securin (Cut2) for destruction, which frees and activates separase. Rec8,
phosphorylated by CK1, polo kinase 1 (Plk1) and DDK at various sites, is then bound by
separase, which cleaves it and prompts its chromatin dissociation from chromosome arms. This,
along with the dissolution of chiasmata, allows homologues to move to opposite poles as cells
progress to anaphase I (Figure 1.3) (Kitajima et al. 2003; Watanabe 2003; Losada and Hirano
2005; Nasmyth and Haering 2005; Le et al. 2013; Sakuno and Watanabe 2015; Phadnis et al.
2015).
Centromeric cohesion must be retained until MII to prevent premature sister separation.
This is accomplished by preventing local destruction of cohesin at the centromere during the
period it is actively removed from chromosome arms (Buonomo et al. 2000; Kitajima et al.
2003). In the fruit fly, MEI-332 makes centromeric cohesin resistant to separase, while in the
yeasts shugoshin 1 (Sgo1) takes on that protective role. Although these proteins have similar
functions, they act at different meiotic stages. While fission yeast Sgo1 is active between
9

metaphase I and anaphase I, MEI-332 and budding yeast Sgo1 function from prophase to
metaphase II. This difference may reflect the various degrees of dependence each organism has
on their centromeric cohesin protector (Kerrebrock et al. 1992; Katis et al. 2004; Kitajima et al.
2004; Kitajima et al. 2004; Marston et al. 2004; Rabitsch et al. 2004). Interestingly, fission yeast
Sgo2 is active in both mitosis and MI, but not in MII. Its dual roles in both cellular division
programs may be explained by its involvement in regulating chromosome attachment at the
spindle, a role independent of centromeric cohesion retention (Vanoosthuyse et al. 2007). Sgo1’s
protective activity in fission yeast is evident when cells containing non-cleavable Rec8 fail to
separate sister chromatids in MII. In addition, in the absence of Sgo1, Rec8 is not retained at the
centromere and thus sister chromatids segregate randomly after anaphase II (Kitajima et al.
2004; Rabitsch et al. 2004). Centromere and kinetochore proteins also assist in regulating
cohesion protection. In budding yeast, increased mini-chromosome loss 3 (Iml3) and
chromosome loss 4 (Chl4) facilitate the establishment of cohesin at the centromere (Marston et
al. 2004). Fission yeast Bub1 (whose spindle checkpoint function assures appropriate
chromosome segregation in MI) and Swi6 localize Sgo1 to the pericentromere, where in
conjunction with Clr4, help to retain the Rec8-Pcs3 cohesin duo (Bernard et al. 2001; Nonaka et
al. 2003; Sakuno and Watanabe 2003; Kitajima et al. 2004; Kim et al. 2015). As previously
alluded, the presence of Pcs3 and not Rec11 at the centromere suggests a different regulatory
mechanism for maintaining not only centromeric cohesion, but also for keeping low levels of
centromeric recombination (Figure 1.3) (Sakuno and Watanabe 2015; Nambiar and Smith
2016). For successful chromosome segregation in each meiotic division, sister kinetochores must
face the same pole (mono-orientation or co-orientation) in MI and opposite poles (bi-orientation)
in MII. Controlling this change in kinetochore orientation is intricate and involves many key
10

players (Nasmyth 2001; Watanabe 2012). In both yeasts, cells with significant reductions in
chiasmata resulting from Spo11 (Rec12) elimination show normal kinetochore co-orientation.
However, in fission yeast kinetochore orientation is disrupted when Rec12 is absent in conditions
that destabilize the centromere (Klein et al. 1999; Kitajima et al. 2003; Davis and Smith 2003;
Yamamoto and Hiraoka 2003). Moreover, sister chromatid cohesion is crucial for co-orientation.
The absence of Rec8 causes kinetochores not to mono-orient in MI, which leads to equational
division in MI and random segregation of sister chromatids in MII. Swapping Rec8 with the
mitotic cohesin does not yield proper kinetochore orientation in fission yeast as it does in
budding yeast, albeit with an important reduction in centromeric cohesion (Klein et al. 1999;
Watanabe and Nurse 1999; Yokobayashi 2003).

Figure 1.3 Dynamics of chromosome segregation in meiosis. During chromosome pairing, the linear elements
(LinEs) localize along the axes of homologous chromosomes. Cohesin rings bind sister chromatids together, while
chiasmata resulting from recombination connect homologous chromosomes. At the centromere, the cohesin subunit
Rec8 interacts with Moa1 and together help sister kinetochores co-orient toward the same pole. The shugoshins
(Sgo1 and Sgo2, collectively labeled Sgo) stabilize the phosphatase PP2A at the pericentromere, which prevents
cleavage of centromeric cohesin by separase. During MI onset, cohesion on chromosome arms but not the
centromere is eliminated and homologous chromosomes separate to opposite sides. As cells enter MII, centromeric
cohesion is removed and sister chromatids segregate to different poles. Diagram was adapted from Ishiguro and
Watanabe 2007.
11

More directly involved in defining kinetochore orientation are the three components of
budding yeast monopolin: monopolar microtubule attachment during meiosis I (Mam1),
chromosome segregation in meiosis I (Csm1), and loss of rDNA silencing 4 (Lrs4). Mam1
localizes at the kinetochores during prophase and metaphase I only. Csm1 and Lrs4 are not
specific to meiosis, as they are found in the nucleolus in mitosis, but join Mam1 at the
kinetochores in prophase and metaphase I (Tóth et al. 2000; Rabitsch et al. 2003). Apart from its
roles in cohesin removal and processing of recombination intermediates, Cdc5 (polo kinase 1)
ensures correct kinetochore orientation in MI by releasing Lrs4 from the nucleolus and localizing
monopolin to the kinetochores through phosphorylation, especially of Mam1 (Clyne et al. 2003;
Lee and Amon 2003). Although Pcs1 is the fission yeast orthologue of Csm1 and is found at both
the kinetochores and nucleolus, its role in aiding correct chromosome segregation is confined to
mitosis and MII, not MI (thus excluding it from regulating mono-orientation) (Rabitsch et al.
2003). Instead, fission yeast co-orientation of sister kinetochores is achieved in a cohesion-
mediated manner. Moa1 and Rec8 physically interact and co-localize at the central core of the
centromere. Elimination of Rec8 disrupts mono-orientation in much the same way as does the
lack of Moa1. Therefore, Moa1’s role in ensuring co-orientation of sister kinetochores in MI
requires the presence of Rec8 to facilitate central core cohesion (Yokobayashi and Watanabe
2005; Kim et al. 2015). Properly engaging monopolin and Rec8-Moa1 at the kinetochores and
centromere, respectively, before MI, therefore, prevents activation of the spindle checkpoint
through Mad2, which then induces Aurora B (budding yeast Ipl1, fission yeast Ark1) to
destabilize any faulty kinetochore-microtubule connections until suitable tension has been re-
established, at which point cells can proceed to anaphase I (Figure 1.3) (Biggins 2001; Shonn
2000; Shonn 2003; Dewar 2004; Hauf et al. 2007).
12

As cells transition from MI to MII, a balance of CDK activity must be established. While
low CDK levels induce the disassembly of the MI spindle, high CDK activity is required to
prevent the formation of pre-replicative complexes (Pre-RCs) (Marston and Amon 2004). Work
in frog oocytes indicates that MII occurs when the APC/C is inhibited and CDK is partially
inactivated. However, a non-degradable form of Cyclin B halts the completion of MI. This
implies that successful MI-MII transition does not require total elimination of CDK, but may
depend, on the temporal equilibrium of Cyclin B production and degradation (Huchon et al.
1993; Furuno et al. 1994; Iwabuchi et al. 2000; Taieb et al. 2001). In budding yeast, failure to
downregulate CDK activity may result in MII division, but chromosome segregation is highly
abnormal. Moreover, Cdc14 is required for the degradation of B-type cyclins and for the
completion of MI. Absence of Cdc14 or the presence of non-degradable B-type cyclins cause a
failure in the breakdown and formation of MI and MII spindles, respectively. Therefore, CDK
downregulation during the MI-MII transition is vital for properly placing chromosome
segregation within the context of meiotic spindle dynamics (Buonomo et al. 2003; Marston et al.
2003). In fission yeast, cells without Mes1 manage to complete MI, but fail to start MII because
they are unable to inhibit co-activators of the APC/C. This phenotype is similar to those incurred
by cells lacking B-type cyclins in budding yeast. Consequently, Mes1 is necessary to retain CDK
activity during the MI-MII transition (Shimoda et al. 1985; Grandin et al. 1993; Dahmann et al.
1995; Kimata et al. 2011).
During MII, the second phase of chromosome segregation takes place. Division at this
stage is dependent on the resurgence of CDK activity, which in itself relies on B-type cyclin
turnover (Hochegger et al. 2001). Bi-orientation of sister chromatid kinetochores is facilitated by
the disappearance of monopolin in budding yeast and a shift in Moa1/Rec8 localization
13

dynamics in fission yeast (Tóth et al. 2000; Yokobayashi and Watanabe 2005). Centromeric
cohesion dissipates during the onset of anaphase II as a result of two critical events: the
reactivation of separase and the removal of cohesin protection. Centromeric Rec8 is targeted for
degradation, allowing the spindles to pull sister chromatids in opposite directions (Rabitsch et al.
2003; Ishiguro and Watanabe 2007; Yamagishi et al. 2014). Though most mammalian oocytes
arrest meiotic progression at metaphase II in anticipation of fertilization, both yeasts proceed to
sporulation (Davis and Smith 2001; Kishimoto 2003).
1.2 The fork protection complex
During S-phase, cells must duplicate their DNA faithfully despite many obstacles the
replisome encounters. The fork protection complex (FPC) facilitates smooth transit through
difficult-to-replicate DNA regions and also mediates many processes vital to genomic integrity.
Its localization at the replication fork enables the FPC to travel with the replisome and coordinate
multiple transactions with chromatin and with chromatin-associated proteins (Figure 1.4).
The FPC consists of two primary proteins: Swi1 and Swi3 (Timeless and Tipin in
metazoans or Tof1 and Csm3 in budding yeast). In fission yeast as well as in other organisms,
the FPC components behave as a heterodimeric complex with interdependent regulation (Ünsal-
Kaçmaz et al. 2007; Gotter et al. 2007; Noguchi et al. 2004; Chou et al. 2006). At the beginning
of S-phase, the FPC associates with origins of replication, subunits of the mini-chromosome
maintenance (MCM) helicase, DNA polymerases δ and ε, replication protein A (RPA), Dfp1-
Hsk1 (DDK), structural maintenance of chromosome 1/3 (Smc1/3) and other proteins (Ünsal-
Kaçmaz et al. 2007; Chou et al. 2006; Gotter et al. 2007; Yoshizawa-Sugata et al. 2007;
Noguchi et al. 2004; Calzada et al. 2005; Errico et al. 2007; Gambus et al. 2006; Katou et al.
2006; Matsumoto et al. 2005; Tanaka et al. 2009; Leman et al. 2010). These interactions
14

highlight the FPC’s involvement in different functions including normal DNA replication,
replication at fragile regions, pausing and stalling of the fork, the replication checkpoint and
sister chromatid cohesion (Leman and Noguchi 2012).

Figure 1.4. The FPC is required for replication at various genomic regions. The replisome encounters many
obstacles as it travels along the genome. These include difficult-to-replicate regions like the centromere and
telomeres, which feature repetitive sequences and protein-DNA complexes. Cohesin rings, fork barriers with and
without Myb-domain proteins and DNA structures distorted by exogenous damage are also in the way. The FPC
mediates interactions with all these obstacles, allowing for smooth pausing and passage of the replisome through
otherwise treacherous domains that may interfere with replication stability. Diagram was adapted from Errico and
Costanzo 2012.
Elimination of Swi1 and Swi3 abolishes a DSB break at the mate-type locus. This leads
to mate-type switching defects, which was used to identify the two FPC components in fission
yeast (Egel et al. 1984). Examination of the Swi1 orthologue in budding yeast (Tof1) reveals an
interaction with topoisomerase 1 (hence the name), which suggests a genome maintenance
function for the FPC (Park et al. 1999). In both yeasts, the absence of Tof1 and Swi1-Swi3
during exposure to hydroxyurea (HU, a ribonucleotide reductase inhibitor) is associated with a
failure to activate the replication checkpoint via the effector kinases Rad53 and Cds1,
respectively (Foss et al. 2001; Murakami et al. 1995; Noguchi et al. 2003; Noguchi et. al 2004).
15

Although the human replication checkpoint functions differently to that of the yeasts,
disruption of Timeless-Tipin sensitizes cells to genotoxic agents like HU, UV, gamma rays and
camptothecin because Chk1 phosphorylation decreases, failing to activate the checkpoint (Ünsal-
Kaçmaz et al. 2007; Chou et al. 2006; Gotter et al. 2007; Yoshizawa-Sugata et al. 2007; Leman
et al. 2010; Ünsal-Kaçmaz et al. 2005). However, in a destabilized FPC without exogenous
replication stress, Chk1 activity increases due to the buildup of unstable DNA structures (Ünsal-
Kaçmaz et al. 2007; Yoshizawa-Sugata et al. 2007; Smith et al. 2009). Evidence suggests,
therefore, that the FPC is important for maintaining genome integrity for two reasons: 1). When
present, it helps to activate Chk1 upon replication stress; 2). When absent, it leads to DNA
configurations that induce Chk1 activation.
In metazoans, the FPC’s role as a mediator of the replication checkpoint is corroborated
by its interactions with other proteins involved in stabilizing the fork. During replication stress,
long stretches of ssDNA are generated. These are bound by RPA, a subunit of which (RPA34)
associates with Tipin (Ünsal-Kaçmaz et al. 2007; Gotter et al. 2007; Nakaya et al. 2010). Loss of
Tipin, in turn, is linked to decreased binding of Claspin (Mrc1 in both yeasts) to the fork and
biochemical examination has confirmed the association of Tipin with Claspin (Kemp et al.
2010). Moreover, Claspin is required for Chk1 phosphorylation in conditions that augment
ssDNA (Chini et al. 2003; Kumagai et al. 2000). Meanwhile, Timeless appears to stabilize Tipin,
since cells without Timeless downregulate Tipin (Chou et al. 2006; Yoshizawa-Sugata et al.
2007). These results suggest that Tipin functions like a protein platform that facilitates the
activation of Chk1 in the presence of RPA-ssDNA complexes. In this scenario, Claspin relays
the stress signal and Timeless ensures that the checkpoint apparatus remains integral via its
regulation of Tipin at the fork.
16

In both yeasts, the FPC is implicated, through its interaction with DDK, in the regulation
of origins of replication. In budding yeast, DNA damage leads to the phosphorylation of Dbf4
(binding partner of Cdc7 in the DDK complex) by Rad53. This prevents the association of Cdc45
and other replication factors to the fork, thereby blocking late-origin firing (Lopez-Mosqueda et
al. 2010; Zegerman and Diffley 2010; Masai et al. 2010). In fission yeast, the DDK components,
Hsk1 and Dfp1, physically interact with Swi1 and Swi3. Cells lacking Hsk1 or a C-terminal
segment of Dfp1 are sensitive to sources of replication stress like HU and methyl
methanesulfonate (MMS, an alkylation agent) (Matsumoto et al. 2005; Dolan et al. 2010).
Similar to FPC mutants, but epistatic to them, cells devoid of Hsk1 fail to arrest S-phase
progression in response to MMS (Matsumoto et al. 2005; Zou et al. 2000; Masai et al. 1995;
Brown et al. 1998; Shimmoto et al. 2009; Sommariva et al. 2005). It appears, thus, that
interaction between the FPC and DDK serves to inhibit replication origins when cells incur
replication stress and DNA damage.
Destabilization of the FPC has been observed to impede correct fork progression. DNA
combing and DNA fiber analysis in budding yeast and human cells, respectively, show that the
corresponding absence of Tof1 and Timeless results in shorter replication tracks and slower
replication rates (Ünsal-Kaçmaz et al. 2007; Tourrière et al. 2005). To achieve steady fork
progression, the FPC is thought to stabilize fork structures and replisome components during
DNA replication, especially in difficult-to-replicate areas of the genome where fork
destabilization can induce recurring DNA breakage (Leman et al. 2012). Tof1 and Csm3, for
instance, are crucial in allowing the fork to pause firmly at rDNA fork block sites (Calzada et al.
2005). Without the FPC, HU-treated cells show an increase in fork velocity. In addition, Cdc45
localization with respect to replicated areas in Tof1 deficient cells reveal the uncoupling of
17

helicase from polymerase activity. Consequently, extensive exposure of ssDNA ensues, which
elicits the recruitment of a checkpoint mediator, Rad9 (Errico et al. 2007). The FPC, therefore,
helps to prevent unregulated DNA unwinding, which contributes to the formation of DNA
structures that are prone to damage.
Elimination of the FPC in fission yeast is associated with the emergence of deleterious
genomic rearrangements. Lack of Swi1 in cells growing without genotoxic stress causes the fork
to collapse at rDNA pausing sites with a concomitant increase of Holliday junctions. These
recombination structures also suggest the occurrence of genomic rearrangements, which is
corroborated by elevated Rad22 (Rad52) foci and augmented sister chromatid exchange in the
absence of both Swi1 and Swi3 (Noguchi et al. 2004; Noguchi et al. 2003; Dolan et al. 2010). In
mouse cells depleted of Timeless, a similar increase in Rad51 and Rad52 foci is seen along with
higher sister chromatid recombination levels (Urtishak et al. 2009). These observations support
the conclusion that the FPC prevents fork collapse and damaging forms of DNA recombination
during normal and perturbed replication (Noguchi et al. 2004; Noguchi et al. 2003; Dolan et al.
2010).
Apart from its function in stabilizing the fork during replication, the FPC is also involved
in preserving sister chromatid cohesion (SCC). In budding yeast, cells lacking Csm3 (the Tipin
orthologue) undergo improper chromosome segregation in meiosis (hence the name), which
contributes to their low spore viability (Rabitsch et al. 2001). Moreover, in Caenorhabditis
elegans meiosis absence of TIM-1 (the Timeless orthologue) is linked to faulty chromosome
cohesion and segregation. These phenotypes are substantiated by the mass spectrometry finding
that SMC-1 interacts with TIM-1. Additionally, TIM-1 is necessary for loading non-SMC
18

subunits to the cohesin complex (Chan et al. 2003). Consequently, TIM-1’s role in meiosis
appears to ensure correct SCC and normal segregation.
The FPC mediates transactions at the fork that influence SCC. Budding yeast Tof1 and
Csm3 are purified as a complex and cells without one of these components are inviable in the
absence of cohesion factors Ctf4 or Ctf8. Interestingly, simultaneous elimination of Ctf8 and the
helicase Chl1 (the orthologue of human ChlR1) is also synthetically lethal (Mayer et al. 2001;
Warren et al. 2004; Mayer et al. 2004). In fission yeast, lack of Ctf18, a replication maintenance
factor, in cells devoid of Swi1 is deadly. Moreover, chl1 overexpression rescues the HU and
MMS sensitivity of Swi1 deficient cells, which further supports genetic studies in both yeasts
suggesting that Swi1/Tof1 and Chl1 share a common functional pathway (Ansbach et al. 2008;
Xu et al. 2007). Results in human cells linking ChlR1 insufficiency to Warsaw breakage
syndrome (a cohesinopathy-related disease) along with the observation that fission yeast cells
lacking Swi1 or Swi3 show premature centromere separation in metaphase indicate that the
FPC’s role in stabilizing the fork is coupled to coordinating active replication with SCC
establishment (Petronczki et al. 2004; Noguchi et al. 2004; Parish et al. 2006; Ansbach et al.
2008; Leman et al. 2010; van der Lelij et al. 2010).
Vertebrates also show disruption of SCC upon FPC destabilization. Xenopus egg extracts
containing reduced levels of Smc3 show similar defects in pairing sister chromatids as samples
depleted of Timeless or Tipin (Tanaka et al. 2009; Errico et al. 2009). In human epithelial cells,
reduction of Timeless or Tipin increases SCC defects (Leman et al. 2010; Dheekollu et al.
2011); while in cell extracts, these FPC components co-immunoprecipitate with Smc1 and Smc3,
both of which show a stronger interaction with Timeless. In addition, ChlR1 physically interacts
with Fen1 (a flap endonuclease required for processing Okazaki fragments), components of the
19

cohesin complex and Timeless, which explains why reduction of the latter is linked to
dissociation of ChlR1 and cohesin subunits from chromatin. Coincidentally, a decrease in Fen1
levels is associated with cohesion defects and its budding yeast orthologue associates with Chl1
and Eco1 (an acetyltransferase involved in cohesion establishment) (Farina et al. 2008; Leman et
al. 2010; Rudra et al. 2012). These observations support the hypothesis that Timeless mediates
recruitment of ChlR1 to the fork to facilitate lagging strand synthesis and processing. In support
of this, evidence shows that Timeless suppresses generation of excessive ssDNA, inhibiting
possible formation of loop structures that may block replisome passage through the cohesin ring.
Therefore, it appears that the FPC role in ensuring genomic stability stems from differing roles of
its primary constituents. Tipin helps to activate the replication checkpoint through its association
with RPA and also anchors Timeless to the lagging strand. Timeless, on the other hand,
synchronizes lagging strand processing and SCC via protein-protein interactions with elements
that regulate these functions (Figure 1.5) (Lengronne et al. 2006; Smith et al. 2009; Uhlmann et
al. 2009; Sherwood et al. 2010; Noguchi et al. 2011; Leman et al. 2012; Rudra et al. 2012).

Figure 1.5. Destabilization of the fork leads to DNA damage. Upon encountering a replication block, the FPC
mediates the activation of the replication checkpoint effector kinase Cds1. However, in the absence of Swi1 the
20

helicase uncouples from the polymerases, which generates excess ssDNA. This in turn leads to unstable DNA
structures that collapse the fork and generate DSBs. Diagram was adapted from Leman and Noguchi 2012.
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31

Chapter 2: Destabilization of the replication fork protection complex is associated with
meiotic defects
2.1 Overview
Cells must duplicate their genomes under conditions that are frequently suboptimal.
Failure to perform replication with high fidelity leads to unstable forms of DNA associated with
cancer in humans. The replication fork protection complex (FPC) is important to replication
stability because it coordinates multiple processes at the fork that are especially crucial for
unimpeded passage of the replisome through different DNA barriers and difficult-to-replicate
areas of the genome. In this work, we examine the function of Swi1 and Swi3, fission yeast’s
primary FPC components, to elucidate how replication stability contributes to DNA integrity in
meiosis. We report that destabilization of the FPC results in reduced spore viability, delayed
replication and DSB formation, shift in partner choice, and chromosome mis-segregation in MI
and MII. These phenotypes are linked to accumulation and persistence of DNA damage markers
in meiosis and to problems with cohesion stability at the centromere. These findings reveal a
vital connection between meiotic replication and chromosome segregation, two processes with
major implications to human reproductive health.
2.2 Introduction
Meiosis is a cell division program important for reducing ploidy and for producing
genetic diversity. Whereas in mitosis progenitor cells generate daughter cells with identical
amounts of DNA, in meiosis daughter cells carry half the genetic content of parent cells. To
accomplish this, each phase of meiotic DNA replication is followed by two consecutive rounds
of chromosome segregation. This reduction in ploidy allows organisms that reproduce sexually
32

to combine their chromosomes to propagate diploid offspring (reviewed in Zickler and Kleckner
1999; Cavalier-Smith 2002; Hochwagen 2008; Ohkura 2015).
Unlike mitosis, meiosis executes programmed double strand breaks (prDSBs) that
facilitate recombination between homologous chromosomes (Keeney et al. 1997). The physical
connections (i.e. chiasmata) established in this step are vital for proper chromosome segregation
(Sharif et al. 2002; Davis and Smith 2003). Also important to meiotic nuclear divisions are the
orientation of sister kinetochores and the gradual elimination of cohesion (Kitajima et al. 2003;
Brar et al. 2006; Sakuno et al. 2009). Separation of homologs to opposite poles of the cell after
anaphase I requires that sister kinetochores mono-orient, such that they migrate in the same
direction on the meiotic spindle. At the onset of anaphase II, bi-oriented kinetochores enable
sister chromatids to segregate as they do in mitosis (Yokobayashi and Watanabe 2005; Hauf et
al. 2007; Sakuno et al. 2009; Kim et al. 2015).
Cohesion changes also define meiotic divisions, for a gradual loss of cohesin, first on
chromosome arms and then at the centromere, facilitates the onset of MI and MII, respectively
(Kitajima et al. 2003; Brar et al. 2006). Before MI, Rec8, the meiosis-specific cohesin subunit, is
degraded everywhere along the chromosome except the centromere, where it is protected by
shugoshin (Marston et al. 2004; Kitajima et al. 2004; Ishiguro et al. 2010). This step-wise
process promotes chiasma resolution, necessary for segregation of homologs, while keeping
sister chromatids tethered at the centromere until MII. When cells near MII, Rec8 is entirely
removed from the centromeres, allowing the spindle to pull sister chromatids apart (Kitajima et
al. 2003; Brar et al. 2006; Katis et al. 2010).
The replication fork protection complex (FPC) is important for diverse functions at the
replisome. It helps to maintain genome integrity by participating in crucial roles such as normal
33

replication, replication at centromeres and telomeres, programmed fork pausing, fork stalling and
restart after replication stress, activation of the replication checkpoint, and sister chromatid
cohesion (Reviewed in Leman and Noguchi 2012). Its primary components are Swi1/Tof1
(human Timeless) and Swi3/Csm3 (human Tipin), which form a heterodimeric complex with
interdependent regulation (Ünsal-Kaçmaz et al. 2007; Gotter et al. 2007; Noguchi et al. 2004;
Chou et al. 2006). The FPC is conserved among eukaryotes and has been found to interact with
the mini-chromosome maintenance (MCM) helicase (Chou et al. 2006; Roseaulin et al. 2013),
DNA polymerase α, ε and δ (Errico et al. 2009; Roseaulin et al. 2013), replication protein A
(RPA) (Kemp et al. 2010; Witosh et al. 2014), Dfp1-Hsk1 (DDK) (Matsumoto et al. 2005;
Shimmoto 2009), structural maintenance of chromosome 1/3 (Smc1/3) (Chan et al. 2003; Leman
et al. 2010) and other proteins that also contribute to genome integrity (Tanaka et al. 2010;
Leman et al. 2010; Wang et al. 2013).
In fission yeast, Swi1 and Swi3 were identified through their role in generating a DSB
important for mate-type switching (Egel et al. 1984). A vital insight related to the FPC’s function
in safeguarding genome integrity came from work on Tof1, the budding yeast orthologue of
Swi1, which was found to interact with topoisomerase 1 (Park and Sternglanz 1999). Further
examination of Tof1, Swi1, and Swi3 during hydroxyurea-induced replication stress revealed
their involvement in activating the replication checkpoint (Foss et al. 2001; Murakami et al.
1995; Noguchi et al. 2003; Noguchi et. al 2004). Likewise, reports on the FPC in humans
showed that it facilitates Chk1 phosphorylation upon exposure to genotoxic agents like
hydroxyurea, UV, gamma rays, and camptothecin. Additionally, depletion of the FPC
components was observed to increase Chk1 activity due to accumulated unstable DNA structures
(Ünsal-Kaçmaz et al. 2007; Chou et al. 2006; Gotter et al. 2007; Yoshizawa-Sugata et al. 2007;
34

Leman et al. 2010; Ünsal-Kaçmaz et al. 2005; Smith et al. 2009). Close inspection of proteins
that interact with Timeless-Tipin such as RPA (Gotter et al. 2007; Nakaya et al. 2010) and
Claspin (Kemp et al. 2010) support a model where Timeless stabilizes Tipin at the fork (Chou et
al. 2006; Yoshizawa-Sugata et al. 2007). Tipin senses RPA bound to ssDNA (generated by
replication stress) (Witosch et al. 2014) and communicates with Claspin, which relays the
message to checkpoint effector kinases (Chini et al. 2003; Kumagai et al. 2000). Thus, the role
of the FPC in various organisms appears to focus on sensing replication stress and repressing
DNA structures that are prone to DNA damage.
Another instrumental function of the FPC is to inhibit replication origins after cells suffer
replication stress and DNA damage. The FPC in both yeasts interacts with components of DDK
(Dbf4-Cdc7 in budding yeast and Dfp1-Hsk1 in fission yeast) (Matsumoto et al. 2005;
Shimmoto et al. 2009; Murakami and Keeney 2014). When cells are exposed to sources of DNA
damage like hydroxyurea and methyl-methanesulfonate (MMS, an alkylating agent), DDK
regulates late origins of replication by blocking association of various replication factors to the
fork (Lopez-Mosqueda et al. 2010; Zegerman and Diffley 2010; Wu and Nurse 2014; Murakami
and Keeney 2014). This measure along with other FPC functions are necessary to prevent
unregulated helicase activity. Analysis of FPC-deficient budding yeast cells treated with
hydroxyurea revealed that fork velocity increases with a concomitant uncoupling of Cdc45 and
Tof1 (Ünsal-Kaçmaz et al. 2007; Tourrière et al. 2005). Thus, the FPC is crucial for linking
replisome and DNA helicase activities, which prevent DNA conformations that promote
unregulated genomic rearrangements. Indeed, fission yeast cells lacking Swi1 that undergo
genotoxic stress show fork collapse at rDNA pausing sites as well as a related increase of
Holliday Junctions. Furthermore, destabilization of the FPC was observed to induce elevated
35

levels of Rad52 foci and sister chromatid exchange (Noguchi et al. 2004; Noguchi et al. 2003;
Dolan et al. 2010), which confirms that the FPC is important to prevent fork collapse and toxic
forms of DNA recombination.
The FPC has also been found to function in the regulation of sister chromatid cohesion, a
process intimately linked to DNA replication. In budding yeast, the cohesion factors Ctf4 and
Ctf8 are essential in cells devoid of the FPC components. In addition, absence of the helicase
Chl1 (the yeast homologue of human ChlR1) and Ctf8 was seen to induce synthetic lethality
(Mayer et al. 2001; Warren et al. 2004; Mayer et al. 2004). This result corroborates work in
fission yeast that suggested a common pathway between Swi1 and Chl1 (Ansbach et al. 2004;
Xu et al. 2007). Moreover, it lends support to observations that associate absence of the FPC in
fission yeast with premature centromere separation, and ChlR1 insufficiency in humans to
Warsaw breakage syndrome, a cohesinopathy-related disease (Ansbach et al. 2008; Parish et al.
2006; Leman et al. 2010; van der Lelij et al. 2010). Furthermore, Xenopus egg extracts depleted
of Smc3 or Timeless or Tipin exhibit similar defects in sister chromatid pairing (Tanaka et al.
2009; Errico et al. 2009). These results confirm studies in human were the FPC components
were found to physically interact with Smc1/3 and to cause cohesion defects when depleted
(Leman et al. 2010). These observations support a model proposing that the FPC helps to recruit
ChlR1 to the fork via Timeless to help regulate lagging strand synthesis and processing, since
ChlR1 was found to interact with Fen1 (required for Okazaki fragment processing), cohesin, and
Timeless (Farina et al. 2008; Leman et al. 2010; Rudra and Skibbens 2012). This FPC function
would in turn facilitate smooth replisome passage through cohesion rings by reducing excess
ssDNA generated during replication stress (discussed in Leman and Noguchi 2012).
36

Replication instability is deleterious to genomic integrity and is associated with cancer in
humans (Durkin et al. 2008; Allera-Moreau et al. 2012; Coschi et. al. 2014; van der Crabben et
al. 2016). When it occurs in meiosis, it can reduce fitness of gametes and negatively affect
reproductive health (Baudat et al. 2000; Nudell et al. 2000; Baarends et al. 2001; Gu et al. 2007).
Although the effects of abnormal replication have been widely studied in mitosis, less is known
about the consequences it has in meiosis. Recent work on replication stability genes in fission
and budding yeast, however, has shown a link between alterations to normal replication and
meiotic defects (Dolan et al. 2010; Le et al. 2013; Wu and Nurse 2014; Murakami and Keeney
2014; Mastro and Forsburg 2014). In this report, we examine how an intact replication fork
facilitates the proper execution of the meiotic program. We used cells carrying full deletions of
Swi1 and Swi3 and examined the outcome of their meioses by various means. We observed that
though fork destabilization did not halt meiotic progression, it was associated with problems in
DSB formation and recombination. Moreover, lack the FPC components resulted in abnormal
chromosome segregation in both meiotic divisions, which may explain the substantial reduction
in spore viability in these strains. Lastly, we found evidence that support a model in which the
FPC contributes to genome stability in meiosis by suppressing DNA damage and promoting
proper centromeric cohesion.
2.3 Results
In fission yeast, meiosis is coupled to sporulation. Thus, defective spores may suggest
underlying issues with meiosis. We crossed heterothallic ∆swi1 and ∆swi3 strains (FPC mutants)
to examine their relative spore viability by random spore analysis. Compared to wild type cells
(100% ± 0.47), the FPC mutants showed a substantial drop in spore viability (∆swi1 33.28 ±
0.47%; ∆swi1 37.6 ± 0.49%) (Figure 2.1A). These reductions were similar to those of the ∆swi1
37

∆swi3 (31.78 ± 0.87%) and ∆rec12 mutants (35.64 ± 1.58%). Intriguingly, absence of Rec12
moderately alleviated the spore viability defects of both ∆swi1 (45.45 ± 1.71%) and ∆swi3
(54.33 ± 1.78%) cells. This result may reflect use of DNA damage accumulated in the FPC
mutants as substrates for recombination to compensate for the lack of programmed double strand
breaks (prDSBs) in ∆rec12 cells, which lack the meiosis-specific endonuclease Rec12/Spo11
(discussed in Farah et al. 2005; Pankratz and Forsburg 2005). Moreover, plating efficiency
revealed no substantial decrease in viability in the FPC mutants as compared to wild type cells
(wild type, 100.00 ± 2.46%; ∆swi1, 102.01 ± 2.45%; ∆swi3, 96.72 ± 2.46%; ∆rec12, 122.96 ±
3.45%; ∆rec12 ∆swi1, 125.29 ± 2.81%; ∆rec12 ∆swi3, 111.12 ± 3.10%). Fork destabilization,
thus, is associated with a defective meiotic program.
Since the FPC is necessary for stable replication (Noguchi et al. 2003), we asked if the
observed meiotic defects in the ∆swi1 and ∆swi3 strains were related to problems in meiotic S
phase (meiS phase). For this, we induced synchronous meiosis by trapping cells in G1 through
nitrogen starvation and by heat-inactivating Pat1, a negative meiotic regulator. Increasing the
temperature to 34
o
C causes either haploid or diploid cells carrying the pat1-114 allele to enter
meiosis. To avoid any confounding effects associated with meiotic induction of haploid cells
(described in Yamamoto and Hiraoka 2003; Pankratz and Forsburg 2005), we generated stable h-
/mat2-102 pat1-114/pat1-114 diploids in ∆swi1 and ∆swi3 backgrounds that could only enter
meiosis by temperature shift and which did not sporulate (detailed in Pankratz and Forsburg
2005; Le et al. 2013). We harvested cells over the course of eight hours and used them to
examine replication dynamics, DSB formation and repair, and meiotic progression. We
monitored completion of meiS phase by measuring DNA content changes with fluorescence-
activated cell sorting (FACS) (described in Forburg and Sabatinos 2009).
38

Figure 2.1 Spore viability and replication dynamics in FPC mutants. (A) Bulk spore germination of
heterothallic cells homozygous for the presence or absence of the FPC and Rec12. At least 6 trials were conducted
per genotype. Significance was determined using chi-squared analysis. P-values are reported as follows: p<0.05 *,
p<0.01 **, p<0.001 ***. Error bars represent 95% confidence intervals. Synchronous meiosis in mat2-102/h- pat1-
114/pat1-114 diploids was performed for 8 hours. Cells were harvested every hour and examined as indicated in (B-
D). (B) FACS profiles showing progression of replication through meiotic induction. DNA doubling is denoted as
the change from 2C to 4C DNA content. Images are representative of three independent trials. (C) Pulsed field gel
electrophoresis used to separate whole chromosomes by size and to assess formation and repair of meiotic DSBs.
Smears migrating faster than chromosome III represent DSBs. Images of ethidium bromide-stained agarose gels
39

represent three different trials. (D) DAPI-stained nuclei were counted for each time point to ascertain progression
through meiotic divisions, which are reported as follows: black stands for 1 nucleus, dark-gray for 2 nuclei, and
light-gray for 3 or more nuclei (3+). 2 and 3+ nuclei indicate onset of MI and MII, respectively.
Wild type cells finished bulk DNA replication 3 hours after meiotic induction, while the
FPC mutants exhibited a slanted peak at that time point, which indicates a modest delay in DNA
synthesis (Figure 2.1B). After 4 hours, both FPC mutants finished replication similarly to the
wild type diploid. Although ∆rec12 cells also show a slight delay in replication by hour 3, the
∆rec12 ∆swi3 mutant did not completely finish meiS by hour 5 (Supplemental Figure 2.1). It
should be noted that though results for the ∆rec12 ∆swi1 mutant are still pending, it is likely they
will resemble those of the ∆rec12 ∆swi3 strain given the similarities of the single mutant
profiles. These observations indicate that the FPC is mostly dispensable for replication
progression in unperturbed meiS phase, except when Rec12 is removed, in which case DNA
synthesis is substantially retarded.
Even though fork destabilization did not impede meiS phase progression, it is possible
that programmed DSB formation and repair, which follow replication, could be affected. To
address this, we carried out pulsed field gel electrophoresis (PFGE) to separate chromosomes
and the signature smears of DSB generation by size (described in Young et al. 2004). In wild
type cells, DSBs were initiated 3 hours after induction. By hour 4, the majority of DSBs were
created, as evidenced by a strong signal beneath chromosome III. DBS resolution occurred in
hour 5 with a concomitant return of chromosome signals. By hour 6 all DSBs were repaired,
which is consistent with full meiotic entry (Figure 2.1C). The FPC mutants initiated and
sustained the bulk of DSB formation in hours 3 and 4, respectively. However, DSB repair was
not entirely finished by hours 5 and 6. In the absence of Rec12, cells predictably failed to create
any DSBs (described in Cervantes et al. 2000; Ogino et al. 2006). By contrast, ∆rec12 ∆swi3
cells exhibited a fast-migrating smear with unchanging signal intensity at all time points. This
40

agrees with a constitutive level of DNA damage and DNA breaks, which is consistent with
previous observations involving ∆rec12 and other replication stability mutants (Dolan et al.
2010; Le et al. 2014) (Supplemental Figure 2.1). These results suggest that the FPC is
important for proper induction and repair of prDSBs during meiosis.
Since fork destabilization delayed DSB repair into anaphase I (hour 6) (Figure 2.1C), we
asked if meiotic progression was affected. We stained nuclei with DAPI and scored for changes
in the number of DAPI dots over the course of meiosis (described in Prankratz and Forsburg
2003; Le et al. 2014). Wild type cells had a single DAPI dot until hour 4 (97.70 ± 0.50%) and
began to show 2 dots by hour 5, when 31.4 ± 1.55% of cells entered MI. From hour 6 onward,
cells continued to divide their nuclei until 92.6 ± 0.87% of cells showed 3 or more DAPI dots,
indicating completion of MII (Figure 2.1D). The FPC mutants showed similar kinetics to wild
type cells, except for ∆swi3 cells which were moderately delayed entering MI by hour 5 (17.3 ±
2.47%). Moreover, the ∆rec12 mutant progressed unimpeded through meiosis like wild type
cells and the single FPC mutants. By contrast, ∆rec12 ∆swi3 cells had reduced efficiency in MII
dynamics as seen previously in other replication mutants (Davis and Smith 2003) (Supplemental
Figure 2.1). These results suggest that fission yeast cells complete meiosis even when fork
integrity is compromised. Taken together, these observations indicate that destabilizing the fork
by eliminating Swi1 or Swi3 is associated with defects in meiotic prophase that though
seemingly moderate may disproportionately affect downstream events.
In mitosis, absence of the FPC components results in DNA damage accumulation
following replication (Noguchi et al. 2004). We asked if the same was true in meiS phase and
whether such damage persisted through meiosis. We used cells whose histones and markers of
DNA damage (Hht1-mRFP/Histone3, Rad11-CFP/RPA and Rad22-YFP/Rad52) were
41

fluorescently tagged and followed their meioses by live cell microscopy (described in Lisby et
al. 2004; Sabatinos et al. 2012; Mastro and Forsburg 2014) (Figure 2.2A-B).

Figure 2.2 Persistence of DNA damage in FPC mutants. (A) Live-cell images of meiotic cells carrying
fluorescently tagged histone3 (Hht1-mRFP), RPA (Rad11-CFP), and Rad52 (Rad22-YFP). Panels are shown for
individual fluorescent channels. A false-color image is rendered to show merged signals. Dotted cell outlines are
overlaid on panels for easier visualization of meiotic cells. Representative images for each reported phenotype were
42

used. Selected time frames were chosen for optimal representation of nuclear dynamics. The analyzed meiotic time
window encompasses horse-tailing (HT), metaphase (MT), meiosis I (MI) and meiosis II (MII). Upright scale bars
denote 5 µm. (B) Cartoon depicting DNA damage persistence through meiosis in FPC mutants. Gray and Black dots
represent RPA and Rad52, respectively. (C) & (D) Quantification of RPA and Rad52 signals in cells from
microscopy pictures shown in (A). More than 150 cells were scored from at least two independent movies for each
genotype. Chi-squared analysis was used to determine significance. Compared to the wild type strain, ∆swi1 and
∆swi3 cells show a significant increase in overall RPA and Rad52 levels during meiosis (p<0.001 for both). Error
bars represent 95% confidence intervals. RPA and Rad52 signals are binned as follows: light-gray for 1 focus, white
for 2 foci, and black for more than 3 foci.
Wild type cells exhibited RPA signals during most of horse tailing (HT) (RPA foci,
100.00%), while a third of cells showed Rad52 signals in late HT (Rad52 foci, 31.41 ± 4.88%)
(Figure 2.2D-C). This agrees with the timing of chromosome pairing and recombination that
ensues after DNA synthesis. When cells reached metaphase, RPA signals began to decrease, but
Rad52 signals showed a transitional increase (RPA foci, 45.82 ± 5.24%; Rad52 foci, 45.82 ±
5.24%) (Figure 2.2D-C). This is an expected outcome during the period when most
recombination is finalized. As cells entered MI, RPA and Rad52 signals disappeared in most
cells (RPA foci, 25.36 ± 4.58%; Rad52 foci, 25.36 ± 4.58%) and were mostly not seen again by
the time MII was completed (RPA foci, 16.43 ± 3.90%; Rad52 foci, 16.43 ± 3.90%) (Figure
2.2D-C).
By contrast, and compared to the wild type strain, higher levels of DNA damage that
persisted through meiosis were observed in ∆swi1 and ∆swi3 cells (Figure 2.2D-C). In the ∆swi1
mutant, RPA levels did not significantly decrease from HT to the end of MII (RPA foci in HT,
100%; MT, 95.51 ± 3.04%; MI, 85.96 ± 5.10%; MII, 75.28 ± 6.34%) (Figure 2.2D-C). Rad52
levels increased from the end of HT to metaphase and gradually returned to similar HT levels by
the end of meiosis, remaining relatively high (Rad52 foci in HT, 81.46 ± 5.71%; MT, 95.51 ±
3.04%; MI, 86.52 ± 5.02%; MII, 75.84 ± 6.29%) (Figure 2.2D-C). In ∆swi3 cells, the same
pattern of moderate RPA level reduction from HT to MII was observed as in the ∆swi1 strain,
albeit in lower overall amounts (RPA foci in HT, 100%; MT, 78.02 ± 6.02%; MI, 69.23 ±
43

6.71%; MII, 54.95 ± 7.23%)(Figure 2.2D-C). Likewise, ∆swi3 cells recapitulated the ∆swi1
mutant’s Rad52 signal level trend through meiosis, but the cell proportion showing the marker
from HT to MII was lower (Rad52 foci in HT, 60.44 ± 7.10%; MT, 76.37 ± 6.17%; MI, 68.70 ±
6.74%; MII, 54.95 ± 7.23%) (Figure 2.2D-C). Three important differences could thus be
appreciated between the FPC mutants and wild type cells: DNA damage was greater, more
persistent, and at more loci (as evidenced by the larger fractions of cells showing multiple RPA
and Rad52 foci during and after MT). These results suggest that, upon fork destabilization, DNA
damage accumulates and remains unrepaired through meiosis, which may negatively affect
events important for meiotic segregation (described in Mastro and Forsburg 2014).

Table 2.1 Recombination dynamics in FPC mutants. Intergenic and intragenic recombination are shown in top
half of table and sister chromatid recombination figures are shown in bottom half. Significance for genetic distance
was determined using a 2-tailed student t-test. Chi-squared analysis was employed for frequencies. At least 5
independent trials were carried out per genotype. Error margins represent 95% confidence intervals. P-values are
shown in the right-hand columns. Fold change values are relative to wild type outcomes. At least 5 independent
trials were performed per genotype.
Fission yeast Rad22/Rad52 is a RecA-like protein important for facilitating
recombination (Muris et al. 1997; van den Bosch et al. 2001). Since we observed abnormal
Rad52 signals before and during meiosis upon fork destabilization, we asked if recombination
dynamics were affected. To address this question, we examined a pair of linked loci in crosses
between parents carrying different alleles and assayed homologous exchange by scoring for
recombinant colonies (detailed in Dolan et al. 2010; Le et al. 2013; Mastro and Forsburg 2014).
44

Relative to wild type cells, the FPC mutants showed an overall decrease in recombination at four
distinct intergenic intervals (Table 2.1). We also assessed intragenic recombination at an ade6
locus containing a recombination hotspot (described in Catlett and Forsburg 2003; Pankratz and
Forsburg 2005). A similar reduction in recombination was observed, especially in the absence of
Swi3. This observation prompted us to ask if fork destabilization shifted recombination
dynamics to favor sister chromatids over homologous chromosomes. For this, we set up a cross
that produced ade+ offspring only if one the parent cells performed intra- or inter-sister
recombination during meiosis (described in Osman et al. 2000; Catlett and Forsburg 2003;
Mastro and Forsburg 2014) (Table 2.1). Interestingly, in the absence of Swi1 and Swi3 there was
an increase in sister chromatid exchange (Table 2.1). These results indicate that fork
destabilization promotes a shift in partner choice during recombination, presumably through
DNA damage generated in replication.
Chromosome segregation is sensitive to changes in recombination (Murakami and Nurse
2001; Watanabe and Nurse 2001; Hirose et al. 2012; Mastro and Forsburg 2014). Thus, we asked
if fork destabilization resulted in abnormal nuclear division in MI and MII. We used cells with
fluorescently tagged histones (Htt1-mRFP) and followed the outcome of their meioses by live
cell microscopy (Figure 2.3A-B) (described in Cooper et al. 1998; Chikashige et al. 2001;
Klutstein et al. 2015). Whereas few wild type cells showed abnormal nuclear masses after MI
and MII (MI 5.21 ± 2.28%, MII 10.68 ± 3.17%) (Figure 2.3C-D), the FPC mutants revealed
increases in cells containing asymmetric nuclei and nuclei with unincorporated DNA masses
( ∆swi1 MI 19.40 ± 5.14%, MII 40.10 ± 6.38%; ∆swi3 MI 16.61 ± 4.25%, MII 33.22 ± 5.37%)
(Figure 2.3C-D). Moreover, in the absence of Swi1 and Swi3, cells exhibited more chromosome
fragmentation following MI and MII (wild type MI 9.04 ± 2.94%, MII 9.59 ± 3.03%; ∆swi1 MI
45

29.96 ± 5.96%, MII 40.10 ± 6.38%; ∆swi3 MI 18.64 ± 4.44%, MII 21.02 ± 4.65%)
(Supplemental Figure 2.2A-B). These observations reveal that the FPC is important for proper
chromosome segregation in meiosis.

Figure 2.3 Meiotic segregation in FPC mutants. (A) Live-cell images of meiotic cells carrying fluorescently
tagged histone3 (Hht1-mRFP). Panels are shown for a fluorescent channel and DIC. A false-color image is rendered
46

to show merged signals. Dotted cell outlines are overlaid on panels for easier visualization of meiotic cells. Images
are representative of each phenotype. Displayed time frames were chosen for optimal representation of nuclear
dynamics. The meiotic phases shown are: horse-tailing (HT), metaphase (MT), meiosis I (MI) and meiosis II (MII).
Upright scale bars stand for 5 µm. (B) Cartoon depicting the products of normal and abnormal divisions in MI and
MII. (C) & (D) Quantification of abnormally segregated asci from microscopy images shown in (A). More than 150
cells were scored from at least two independent movies for each genotype. Chi-squared analysis was used to
determine significance. P-values are reported as follows: p<0.05 *, p<0.01 **, p<0.001 ***. Error bars represent
95% confidence intervals.
Segregation problems could emerge from premature sister chromatid separation or
chromosome non-disjunction (Yokobayashi and Watanabe 2005; Hauf et al. 2007; Hirose et al.
2011). To address the first possibility, we crossed cells heterozygous for a GPF marking near the
centromere of chromosome I and carrying histones tagged as indicated above. If MI division
occurred normally (i.e. reductional division), one of the two resulting nuclear spots would
contain a GFP dot. However, if equational division ensued (i.e. abnormal MI division), both
nuclear spots would show GFP dots (described in Yokobayashi and Watanabe 2005; Hauf et al.
2007; Hirose et al. 2011) (Figure 2.4A-B). Compared to the wild type strain (5.8 ± 2.53%),
∆swi1 (20.0 ± 5.93%) but not ∆swi3 (9.50 ± 3.34%) cells showed an important increase in
equational division (Figure 2.4C). This result indicates that Swi1 helps to prevent untimely
separation of sister chromatids, possibly by contributing to correct monopolar attachment, as was
reported previously for Mrc1 (a secondary FPC factor) in conjunction with Rec12 (Hirose et al.
2011). To examine if fork destabilization is associated with chromosome non-disjunction, we
used a similar strategy as indicated above, but in a cross homozygous for the semi-centromeric
GFP marking. If division occurred normally in both MI and MII, GFP dots would be distributed
symmetrically. However, if non-disjunction took place, GFP distribution would be asymmetric in
both divisions (described in Yokobayashi and Watanabe 2005; Hauf et al. 2007; Hirose et al.
2011) (Figure 2.5A-B). While only ∆swi3 cells show elevated non-disjunction issues in MI
(wild type 2.40 ± 1.90%; ∆swi1 5.98 ± 3.43%; ∆swi3 9.86 ± 4.00%) (Figure 2.5C), both FPC
mutants feature increases in cells that fail to disjoin sister chromatids in MII (wild type 8.00 ±
47

3.36%; ∆swi1 16.30 ± %; ∆swi3 15.02 ± 4.80%) (Figure 2.5D). These results suggest that Swi3
is necessary for proper homolog segregation in MI, while both FPC components contribute to
normal sister chromatid separation in MII.

Figure 2.4 MI equational division in FPC mutants. (A) Live-cell images of meiotic cells carrying fluorescently
tagged histone3 (Hht1-mRFP) and LacI-GFP on a LacO repeat at lys1 near the centromere of chromosome I. Panels
48

are divided into individual fluorescent channels. A false-color image was generated to show merged signals. Dotted
cell outlines are overlaid on panels for easier visualization of meiotic cells. Images are representative of each
phenotype. Displayed time frames were chosen for optimal representation of nuclear dynamics. The meiotic phases
shown are: horse-tailing (HT), metaphase (MT), meiosis I (MI) and meiosis II (MII). Upright scale bars are 5 µm.
(B) Cartoon depicting a pair of homologous chromosomes where one is marked with GFP near the centromeres.
Symmetrical distribution of GFP indicates equational division. (C) Quantification of cells showing equational
division in MI. More than 150 cells were scored from at least two independent movies for each genotype. Chi-
squared analysis was used to determine significance. P-values are reported as follows: p<0.05 *, p<0.01 **, p<0.001
***. Error bars represent 95% confidence intervals.
Reduction in homologous recombination explains some of the mis-segregation
phenotypes observed upon fork destabilization, as is the case when meiS phase is perturbed
(described in Watanabe and Nurse 2001). However, cohesion problems are also consistent with
abnormal meiotic divisions (Watanabe and Nurse 1999). To determine if fork destabilization
affects processes important to cohesion stability, we used live cell microscopy to examine cells
with fluorescently tagged histones as well as Rec27-GFP (a component of the linear elements),
Sgo1-GFP (shugoshin, a centromeric cohesion protector) and Rec8-GFP (the -kleisin subunit of
meiotic cohesin). These proteins are associated with different aspects of cohesin function and
regulation (described in Watanabe and Nurse 1999; Rozalén et al. 2008; Kitajima et al. 2004;
Kitajima et al. 2006). Moreover, meiotic Rec8 stability is compromised between MI and MII
when Dfp1 is dysfunctional (Le et al. 2013).
Thus, we reasoned that any changes to the temporal dynamics of Rec27, Sgo1, and Rec8
due to fork destabilization would indicate cohesion-related sources of chromosome mis-
segregation. Rec27 functions downstream of Rec8 and upstream of Rec12 (Sakuno and
Watanabe 2015; Phadnis et al. 2015). It is important for alignment of homologous chromosomes
and for promoting DSB formation (Ellermeier and Smith 2005; Davis et al. 2008; Phadnis et al.
2015), thus serving as a convenient readout for these processes (Figure 2.6A-B). Compared to
wild type cells, Rec27 elimination in the FPC mutants was less gradual and considerably faster
(wild type, 28.26 ± 1.31 min.; ∆swi1, 22.68 ± 1.41 min.; ∆swi3, 21.22 ± 1.63 min.) (Figure
49

2.6C). These Rec27 defects are suggestive of changes to the structural integrity of chromosome
axes affecting DSB formation and repair. Because fork destabilization has been associated with
cohesion defects before the onset of mitosis (Ansbach et al. 2008), these observations may also
indicate changes to cohesin stability in meiosis.

50

Figure 2.5 Non-disjunction in MI and MII in FPC mutants. (A) Live-cell images of meiotic cells carrying
fluorescently tagged histone3 (Hht1-mRFP) and LacI-GFP on a LacO repeat at lys1 near the centromere of
chromosome I. Panels are divided into individual fluorescent channels. A false-color image is provided to show
merged signals. Dotted cell outlines are overlaid on panels for easier visualization of meiotic cells. Images are
representative of each phenotype. Displayed time frames were chosen for optimal representation of nuclear
dynamics. The meiotic phases shown are: horse-tailing (HT), metaphase (MT), meiosis I (MI) and meiosis II (MII).
Upright scale bars are 5 µm. (B) Cartoon depicting a pair of homologous chromosomes marked with GFP near the
centromeres. Asymmetric distribution of GFP indicates non-disjunction events. (C-D) Quantification of cells
showing chromosome non-disjunction in MI and MII. More than 150 cells were scored from at least two
independent movies for each genotype. Chi-squared analysis was used to determine significance. P-values are
reported as follows: p<0.05 *, p<0.01 **, p<0.001 ***. Error bars represent 95% confidence intervals.
In fission yeast, Sgo1 protects centromeric cohesion from separase before anaphase I
(Kitajima et al. 2004; Ishiguro et al. 2010). Given that that ∆swi1 cells show equational division
in MI, which is linked to issues with centromeric cohesion stability (Yokobayashi et al. 2005;
Hauf et al. 2007), we asked if fork destabilization would affect the temporal dynamics of Sgo1
(Figure 2.7A-B). Intriguingly, we observed that the ∆swi3 mutant (14.42 ± 1.46 min.), and not
∆swi1 (18.04 ± 1.36 min.), exhibited a shorter duration of Sgo1 signal as compared to wild type
cells (17.50 ± 1.88 min.) (Figure 2.7C). These results suggest Swi3 is necessary for stability of
Sgo1 at the centromere and further highlight the separate functions of the FPC components.
Rec8 is a subunit of cohesin that helps to keep sister chromatids together following meiS
phase (Watanabe and Nurse 1999; Watanabe and Nurse 2001). During anaphase I, Rec8 is
removed from chromosome arms, which facilitates chiasma resolution and segregation of
homologous chromosomes (Buonomo et al. 2000; Kitajima et al. 2003; Brar et al. 2006).
However, Rec8 is retained at the centromere until right before anaphase II to ensure correct
segregation of sister chromatids (Kitajima et al. 2003; Kitajima et al. 2004; Riedel et al. 2006).
Moreover, Rec8 along with Moa1 facilitate mono-orientation of kinetochores in MI, thus
promoting segregation of homologous chromosomes to opposite poles of the cell (Yokobayashi
and Watanabe 2005; Hauf et al. 2007). Since lack of Swi1 and Swi3 are associated with
abnormal MI segregation and decreased timing of Sgo1 before MI, respectively, we asked if fork
51

destabilization would disrupt the temporal dynamics of centromeric cohesion. For this, we
followed Rec8-GFP from late metaphase I until right before MII (Figure 2.8A-B).

Figure 2.6 Elimination of the linear elements in FPC mutants. (A) Live-cell images of meiotic cells carrying
fluorescently tagged histone3 (Hht1-mRFP) and Rec27-GFP. Panels are divided into individual fluorescent
channels. A false-color image was rendered to show merged signals. Dotted cell outlines are overlaid on panels for
easier visualization of meiotic cells. Images are representative of each phenotype. Displayed time frames were
chosen for optimal representation of nuclear dynamics. The meiotic phases shown are: metaphase (MT) and meiosis
I (MI). Upright scale bars are 5 µm. (B) Cartoon depicting elimination of Rec27-GFP during metaphase and before
MI. (C) Quantification of cells showing the timing of Rec27-GFP elimination. More than 150 cells were scored
52

from at least two independent movies for each genotype. Chi-squared analysis was used to determine significance.
P-values are reported as follows: p<0.05 *, p<0.01 **, p<0.001 ***. Error bars represent 95% confidence intervals.
In wild type cells, GFP signal goes from pan-nuclear (observed in late metaphase I to
early anaphase I) to a pair of foci that localize at the centromeres and mostly persist for 32.22 ±
1.24 min. (Figure 2.8C). This signal pattern following MI corresponds to Rec8 elimination at
chromosome arms, persistence at the centromere, and degradation before MII (detailed in
Watanabe and Nurse 1999; Kitajima et al. 2003; Le et al. 2013). In the FPC mutants, however,
persistence of centromeric cohesion is substantially shorter ( ∆swi1, 20.91 ± 1.49 min.; ∆swi3,
24.02 ± 1.69 min) and Rec8 loss is more precipitous than that of wild type cells (Figure 2.8C).
To confirm the centromeric cohesion defect observed upon fork destabilization, we
synchronized cells by Pat1 inactivation and performed chromatin immunoprecipitation followed
by PCR to determine if Rec8 enrichment at the centromere would be affected over the period that
encompasses the metaphase I-anaphase I transition. Relative to wild type cells (1.51 fold), the
∆swi1 (0.17 fold) and ∆swi3 (0.26 fold) mutants respectively showed an important and modest
reduction in centromeric Rec8 enrichment (Figure 2.8D). These results along with the
observation that sister chromatids in the FPC mutants are left untethered at the centromeres for
longer ( ∆swi1, 31.55 ± 1.64 min.; ∆swi3, 28.30 ± 1.76 min.) than in wild type cells (16.84 ± 1.07
min.) suggest that fork destabilization accelerates elimination of centromeric cohesion, which not
only precipitates early separation of sister chromatids in MI (via Swi1 elimination), but also
contributes to their abnormal segregation in MII.
Swi6 (the fission yeast homologue of human HP1) is essential for heterochromatin
formation and for the recruitment of Rec8 and Sgo1 to the centromere (Ekwall et al. 1995;
Kitajima et al. 2003; Kawashima et al. 2007; Yamagishi et al. 2008). To further corroborate the
centromeric cohesion defects thus far observed, we asked if fork destabilization would impact
53

centromeric localization of Swi6. We used live cell microscopy to monitor co-localization of
Cnp1-mCherry (the fission yeast homologue of mammalian CENPA, a centromere-specific
histone variant) and Swi6-GFP following MI and MII (Figure 2.9A-B).

Figure 2.7 Duration of Sgo1 in metaphase in FPC mutants. (A) Live-cell images of meiotic cells carrying
fluorescently tagged histone3 (Hht1-mRFP) and Sgo1-GFP. Panels are divided into individual fluorescent channels.
A false-color image was generated to show merged signals. Dotted cell outlines are overlaid on panels for easier
visualization of meiotic cells. Images are representative of each phenotype. Displayed time frames were chosen for
optimal representation of nuclear dynamics. The meiotic phases shown are: horse-tailing (HT), metaphase (MT) and
54

meiosis I (MI). Upright scale bars are 5 µm. (B) Cartoon depicting duration of Sgo1-GFP during metaphase and
before MI. (C) Quantification of cells showing the timing of Sgo1-GFP signal in metaphase. More than 150 cells
were scored from at least two independent movies for each genotype. Chi-squared analysis was used to determine
significance. P-values are reported as follows: p<0.05 *, p<0.01 **, p<0.001 ***. Error bars represent 95%
confidence intervals.
We used this approach because it is difficult to distinguish Cnp1-Swi6 co-localization
before MI by this method. Additionally, we reasoned that co-localization dynamics of these
proteins after MI and MII should reflect the overall ability of cells to recruit heterochromatin to
the centromere. We observed that more ∆swi1 (MI, 6.77 ± 4.27%; MII, 12.03 ± 5.53%), but not
∆swi3 (MI, 4.76 ± 2.53%; MII, 5.13 ± 2.62%) or wild type (MI, 0.69± 1.35%; MII, 4.17 ±
3.26%) cells had defective Cnp1-Swi6 co-localization after each meiotic division (Figure 2.9C-
D). Although the effect is modest, it nonetheless suggests Swi1 is necessary for correct
recruitment of heterochromatin to the centromere.
2.4 Discussion
The fork protection complex (FPC) travels with the replisome and is important for
coupling the MCM helicase with polymerases (Leman et al. 2011). This function helps to
prevent excessive ssDNA during replication stress. Areas with extended ssDNA segments are
prone to fork reversal and unregulated genomic recombination (Noguchi et al. 2003; Noguchi et
al. 2004). In fission yeast, the FPC’s primary components are Swi1 and Swi3 (Noguchi et al.
2004). Swi1 associates with chromatin through its DDT domain, which is also important for
recruiting Swi3 (Rapp et al. 2012). Stable association of Swi1 and Swi3 facilitates binding to the
fork of Mrc1, a secondary FPC component necessary for replication processivity and activation
of the replication checkpoint (Shimmoto et al. 2010). Together, these proteins act as sensors of
replication stress that protect genomic integrity during DNA duplication. Furthermore, Swi1 and
Swi3 are also involved in restarting the fork after exposure to S-phase stressors and play a role in
55

establishing and maintaining proper sister chromatid cohesion in mitosis (Noguchi et al. 2004;
Matsumoto et al. 2005; Ansbach et al. 2008; Rapp et al. 2010).

Figure 2.8 Persistence of centromeric cohesion in FPC mutants. (A) Live-cell images of meiotic cells carrying
fluorescently tagged histone3 (Hht1-mRFP) and Rec8-GFP. Panels are divided into individual fluorescent channels.
A false-color image was generated to show merged signals. Dotted cell outlines are overlaid on panels for easier
visualization of meiotic cells. Images are representative of each phenotype. Displayed time frames were chosen for
56

optimal representation of nuclear dynamics. The meiotic phases shown are: metaphase (MT), meiosis I (MI) and
meiosis II (MII). Upright scale bars are 5 µm. (B) Cartoon depicting gradual elimination of Rec8, first at
chromosome arms and then at the centromere. Rec8 dynamics are shown in the context of MI and MII division. (C)
Quantification of cells showing persistence of Rec8-GFP foci after MI. More than 150 cells were scored from at
least two independent movies for each genotype. Chi-squared analysis was used to determine significance. P-values
are reported as follows: p<0.05 *, p<0.01 **, p<0.001 ***. Error bars represent 95% confidence intervals. (D)
Quantification of chromatin immunoprecipitation followed by PCR. Cells were synchronized by Pat1 inactivation.
ChIP values were calculated as follows: (IP/input at dg) / (IP/input at mes1); to generate fold values, figures for hour
5 (ana I) were divided by those for hour 6 (MI).
In fission yeast, meiS phase takes longer to complete than S-phase in vegetative
conditions (Wu and Nurse 2013). DNA damage induced at this stage does not elicit checkpoint
arrest and can be repaired through meiotic recombination. In checkpoint mutants, however,
excess damage may persist until at least the first meiotic division (Pankratz and Forsburg 2005).
Moreover, perturbations to meiS phase are associated with downstream recombination and
segregation problems (Watanabe and Nurse 2001; Wu and Nurse 2014). Since the FPC
contributes to replication processivity and response to DNA damage, we asked if absence of its
components would disrupt any aspect of the meiotic program.
We observed that the FPC mutants showed decreased spore viability, which is consistent
with previous reports of similar outcomes in other replication stability mutants (Dolan et al.
2010; Le et al. 2013; Mastro and Forsburg 2014; Murakami and Keeney 2014; Wu and Nurse
2014). The reduction seen in the ∆swi1 ∆swi3 strain is similar to that of ∆swi1 cells and agrees
with Swi1’s function of stabilizing Swi3 at the fork (Leman et al. 2011; Rapp et al. 2012).
Interestingly, combining ∆rec12 with ∆swi1 and ∆swi3 suppressed some spore unviability
observed in the single mutants. This observation suggests that the damage-induced
recombination intermediates previously seen during vegetative growth in the FPC mutants
(Noguchi et al. 2003; Noguchi et al. 2004) may also be present during meiotic prophase and may
serve as recombination sources when prDSBs are abolished.

57

Figure 2.9 Recruitment of heterochromatin to the centromere in FPC mutants. (A) Live-cell images of meiotic
cells carrying fluorescently tagged Cnp1-mCherry and Swi6-GFP. Panels are divided into individual fluorescent
channels. A false-color image was generated to show merged signals. Dotted cell outlines are overlaid on panels for
58

easier visualization of meiotic cells. Images are representative of each phenotype. Displayed time frames were
chosen for optimal representation of nuclear dynamics. The meiotic phases shown are: metaphase (MT), meiosis I
(MI) and meiosis II (MII). Upright scale bars are 5 µm. (B) Cartoon depicting localization of Cnp1-mCherry and
Swi6-GFP in metaphase, MI, and MII. (C) Quantification of cells showing mis-localized Cnp1-mCherry and Swi6-
GFP dots in both meiotic divisions. More than 130 cells were scored from at least two independent movies for each
genotype. Chi-squared analysis was used to determine significance. P-values are reported as follows: p<0.05 *,
p<0.01 **, p<0.001 ***. Error bars represent 95% confidence intervals.
Absence of the FPC components did not halt meiotic progression and was associated with
a modest delay in bulk DNA synthesis. Given that DSB formation follows replication and that
the FPC mutants showed delays in DSB repair, it is possible both events were affected by
unstable DNA structures resulting from large ssDNA gaps or rearranged forks (as discussed in
Noguchi et al. 2003; Noguchi et al. 2004). This would confirm the constitutive DNA damage
bands seen in PFGE images of ∆swi3 cells that lack Rec12. Deletion of ∆swi1 and ∆swi3 was
found to produce increased RPA and Rad52 levels in vegetative cells (Noguchi et al. 2004).
Thus, we decided to visualize these DNA damage markers in FPC mutants undergoing meiosis.
We observed elevated signals for both markers that intensified in late HT and persisted until MII.
This result is consistent with the FPC’s role in surveilling excess ssDNA levels and in
maintaining stability at the fork.
A secondary consequence of accumulating large pools of Rad52 is the impact this may
have on recombination, since its dissociation is crucial for Rad51 binding (Shinohara and Ogawa
1998; New et al. 1998). We analyzed four distinct intergenic loci and one intragenic locus to
assess recombination activity in the FPC mutants. We observed a general decrease in
homologous recombination, which prompted us to ask if the same was true for sister chromatid
exchange. Deletion of the FPC components in meiosis led to increased recombination between
sister chromatids. This change in partner choice, while ensuring resolution of recombination
substrates, may compromise downstream meiotic activity which depends on chiasma formation
for proper chromosome segregation (Hirose et al. 2011).
59

We used live cell imaging to follow FPC mutant cells through meiosis. We observed that
without Swi1 and Swi3, fission yeast cells have increased chromosome mis-segregation in both
meiotic divisions. We asked if the observed division anomalies resulted from premature sister
separation or chromosome non-disjuction. Interestingly, we found separate functions of Swi1
and Swi3 during MI. Swi1 appears to facilitate accurate monopolar attachment, since its absence
is associated with an increase in equational division (Yokobayashi et al. 2005). By contrast,
Swi3 seems to mediate correct disjunction of homologous chromosomes. In MII, both FPC
mutants exhibit increased non-disjunction of sister chromatids. Issues with recombination have
been previously reported to affect meiotic segregation (Davis and Smith 2003). Thus, the mis-
segregation phenotypes of the FPC mutants may be related to their reduced homologous
recombination.
Another possibility is that these phenotypes originated due to cohesion problems
(Watanabe and Nurse 1999; Yokobayashi et al. 2003; Kitajima et al. 2003). We addressed this
option by examining aspects of cohesin regulation that may affect meiotic divisions. We looked
at the timing of Rec27 elimination because it coincides with DSB repair and chiasma resolution.
A component of the linear elements that facilitates correct pairing of homologs, Rec27 functions
downstream of Rec8, but upstream of Rec12 (Davis et al. 2008; Sakuno and Watanabe 2015;
Phadnis et al. 2015). Therefore, the finding that without the FPC Rec27 elimination is
accelerated confirms the DSB repair defects seen by PFGE and suggest possible cohesion issues
(Sakuno and Watanabe 2015; Phadnis et al. 2015). Since Sgo1 protects centromeric cohesion
before the onset of anaphase I (Kitajima et al. 2004; Ishiguro et al. 2010), we decided to examine
its temporal dynamics in metaphase. We observed that deletion of Swi3 but not Swi1 decreases
the timing of Sgo1’s signal. This result indicates that Swi3 is important for normal protection of
60

centromeric cohesion and that this function may be independent of its association with Swi1 at
the fork.

Figure 2.10 Model of FPC contribution to meiotic segregation. Proposed model explaining the mis-segregation
phenotypes of FPC mutants. Swi1 stabilizes Swi3 at the fork. Both Swi1 and Swi3 help to suppress generation of
large ssDNA gaps bound by excess RPA. Normal RPA levels facilitate proper binding of Rad52 to nucleoprotein
filament, which in turn promotes Rad51 binding. Both Rad51 and Dmc1 promote normal homologous
recombination, which guarantees proper disjunction of homologous chromosomes in Mi. Swi1 is necessary for
normal recruitment of Swi6 to the centromere. Centromeric Swi6 helps recruit Sgo1, which stabilizes Rec8 at the
centromere. Stable centromeric cohesion ensures monopolar attachment of sister kinetochores, which is necessary
for correct reductional segregation in MI. Suppression of DNA damage and maintenance of cohesion stability at the
centromere ensures accurate separation of sister chromatids.
61

Rec8 is the cohesin subunit whose separase-dependent degradation ensures proper
segregation of sister chromatids in MII (Kitajima et al. 2003; Ishiguro et al. 2010). Because both
FPC mutants showed abnormal MII division, we decided to follow, by live cell imaging, the
duration of centromeric Rec8 from MI to MII (as described in Le et al. 2013). In the FPC
mutants, we observed a precipitous disappearance of most Rec8 foci within the first 20 minutes
of meiosis. This meant that for the remaining 30-50 minutes before MII, cells lacking either FPC
component were untethered at the centromere. This may explain the high proportion of cells that
exhibited non-disjunction issues in MII. To corroborate this finding, we examined the
enrichment of Rec8 at the centromere during the metaphase I-anaphase I transition period.
Relative to wild type cells, we observed a reduction of Rec8 that agrees with microscopy
observations. Furthermore, it has been established that the heterochromatin protein Swi6 helps to
recruit both Sgo1 to the centromere (Yamagishi et al. 2008). Therefore, we asked if Swi6
recruitment to centromeres would be affected in the FPC mutants during MI and MII. We found
that without Swi1, cells mis-colocalize Swi6 and Cnp1 (a centromeric marker). This observation
confirms that the FPC is necessary to maintain cohesion stability at the centromere, a result that
has been previously reported for pre-mitotic cells (Ansbach et al. 2008).
In light of these results, we propose a model where the FPC contributes to proper meiotic
segregation by suppressing unregulated generation of ssDNA and by ensuring cohesion stability
at the centromere. These two possibilities seem to agree with the herein observed separate
functions of the FPC components. Because Swi1 regulates Swi3’s binding to the fork, we
reasoned that phenotypes shared by both mutants can be attributed to the absence of Swi3.
However, Swi1 may retain independent functions that may, if disrupted, compound those of a
Swi3 molecule unbound to the fork. Consistent with observations previously made of Tipin
62

(reviewed in Leman and Noguchi 2012), Swi3 likely monitors ssDNA content by sensing RPA
binding. In the absence of Swi3, RPA levels increase and so does the amount of Rad52, which
antagonizes Dmc1 (a RecA-like protein important for homologous recombination) (Murayama et
al. 2013). To test this possibility, we examined whether ∆dmc1 cells would recapitulate the spore
viability and recombination results of both FPC mutants. While reduction in spore viability was
modest relative to wild type cells (wild type 100% ± 0.47; ∆dmc1 85.81 ± 1.57), the ∆dmc1
mutant showed a similar decrease in recombination at the his4-lys4 intergenic interval as the
∆swi1 mutant (wild type 6.81 ± 0.92 cM; ∆swi1 4.81 ± 0.78 cM; ∆swi3 2.43 ± 0.83 cM; ∆dmc1
4.70 ± 0.82 cM). This suggests that decreased recombination may lay behind some of the
homolog non-disjunction issues observed in both FPC mutants, but particularly in ∆swi3 cells.
In the absence of Swi1, we observed problems with Swi6 recruitment to the centromere.
As shown previously, this effect may disrupt centromeric stability of Rec8 by preventing Sgo1
association (Yamagishi et al. 2008). Moreover, ∆swi3 cells showed decreased Sgo1 timing in
metaphase and both FPC mutants had problems with persistence of centromeric Rec8. Thus, it
seems that Swi1 is important for monopolar attachment and reductional division in MI by
maintaining Rec8 stability at the centromere. This connection with Rec8 dynamics may also
confirm the observed prDSB and recombination problems in the FPC mutants via disruption of
Re27 elimination. Given that the FPC physically interacts with DDK and DDK has a role in
regulating replication, DSB formation and cohesion (Matsumoto et al. 2005; Matos et al. 2008;
Katis et al. 2010; Shimmoto et al. 2010; Le et al. 2013), our model predicts DDK or a yet
unknown DDK substrate as the link that mechanistically connects replication with meiotic
segregation, which would confirm similar observation made in budding yeast (Murakami and
Keeney 2015). Further research is necessary to elucidate how this is executed in fission yeast.
63

2.5 Materials and Methods
Cell growth and culture
Supplemental Table 2.1 shows the strains used in this work. Detailed descriptions of
general fission yeast culture conditions, media and standard techniques are found in Sabatinos
and Forsburg (2010). Liquid and solid yeast extract plus supplements (YES) was used for the
construction and maintenance of all strains, except for when pombe minimal glutamate (PMG)
with the appropriate supplements was required. For plating efficiency, cells were picked from
single colonies and grown in 10-ml cultures at 32
o
C to mi-log phase. Cells were counted with a
hemocytometer and 500 cells were spread with glass beads on each plate. Plates were incubated
at 32
o
C for 3-5 days, after which colonies were counted.
Spore viability and recombination
Spore viability and recombination assays were carried out by picking single colonies
from mate-type heterothallic strains, which were subsequently mixed in 5 µl sterile water on malt
extract (ME) plates. Mated strains were incubated at 25
o
C for 2-3 days. Colony matter from
individual mating patches was swiped and resuspended in a 1-ml 0.5% glusulase solution
(PerkinElmer; Boston, MA). The glusulase suspension was incubated with constant rotation at
25
o
C for 12-16 hours. Spores were counted and 500-1000 spores were dispersed per YES plate.
Spores were allowed to germinate and proliferate at 32
o
C for 3-5 days. Colonies were counted
and, where suitable, replica plated onto solid PMG media with appropriate supplements. For
assays involving tetrad dissection, cell matter was taken from individual mating patches and
spread on YES plates. Asci were dissected using a microscope with a micromanipulator, after
which spores were treated as mentioned above. To distinguish diploid cells, phloxin B was added
64

to YES plates in which diploids look red, while haploids look pale pink. Moreover, in
recombination assays diploid spore formation was monitored by combining parent strains with or
without his3-D1 and leu1-32 on the same chromosome. Thus, only colonies that were His+ Leu-
or His- Leu+ arose from actual recombination events and not from diploids that survived
glusulase treatment. Other recombination experiments examined two linked loci in parent strains
where one of the loci is dysfunctional (e.g. his4-239 lys4+ or his4+ lys4-95). Recovered colonies
that are His+ Lys+ or His- Lys- are products of recombination. Genetic distance was calculated
using Haldane’s formula as described in Smith (2009): cM = -50 ln (1-2R), where genetic
distance is in cM and R is the recombinant fraction. Restoration of the Ade+ phenotype was used
as a measure of intragenic recombination between parent strains respectively carrying the ade6-
M26 and ade6-52 alleles. Experiments were repeated at least six times, plating at least 1000
spores per trial. Sister chromatid recombination was calculated as previously described by Catlett
and Forsburg (2003). To account for post meiotic recombination between sister chromatids,
spores were plated on YE plates. Any white colonies that emerged resulted from true meiotic
events, while sectored colonies arose only after the first cell division. A two-tailed t-test was
employed to determine significance for genetic distances. A chi-squared test was used in all
other instances yielding frequency and proportion data, except for when sample size merited use
of Fisher’s exact test.
Pulse field gel electrophoresis (PFGE)
Stable diploids were created using ade6-M210/M216 complementation in parent strains
carrying the mat2-102 and pat1-114 alleles to perform synchronous meiosis as described by
Catlett and Forsburg (2003) with a few adjustments. Briefly, cultures were grown in Edinburgh
Minimal Medium (EMM) plus supplements, except adenine, to an OD595=0.7-1.0, after which
65

cells were starved for 16.5-17 hours in EMM minus nitrogen. Starved cultures were refed EMM
containing a 1:2 dilution of supplements (minus adenine) plus NH4CL and shifted from 25
o
C to
34
o
C for meiotic induction. Subsequently, cells were harvested every hour (FACS samples were
also taken), treated with sodium azide and washed in phosphate-buffered saline (PBS) and
citrate-phosphate sorbitol EDTA (CSE). A lysing enzymes solution containing 0.2 mg/ml 100T
Zymolyase (Nacalai Tesque; Kyoto, Japan) and 0.45 mg/ml Trichoderma harzianum lysing
enzymes (Sigma, St. Louis, MO) was used to digest the cell wall. Following the first time point,
the lysing solution was titrated to 50% and 25% in the second and all subsequent time points,
respectively, as performed by Cervantes et al. (2000). Lysate from cells was mixed with 2%
CleanCut Agarose (Bio-Rad; Hercules, CA) to generate pulse field gel plugs, which were then
incubated at 55
o
C in a Proteinase K (Bioline Reagents Ltd.; Taunton, MA)/sarkosyl-EDTA
solution for 48 hours, with an intermediate change of protease solution after 24 hours. The DNA
plugs were then washed in Tris-EDTA (TE) and equilibrated in Tris-acetate-EDTA (TAE). A
Bio-Rad Chef II Pulse Field Machine was employed to separate DNA by size using a 1% Mega
Base Agarose (Bio-Rad; Hercules, CA)/TAE gel. The pulse field apparatus was run for 48 hours
at 2 V/cm, with an 1800-sec switch time, and a 106
o
angle. The gel was stained in ethidium
bromide, visualized in a ChemiDoc Imager (Bio-Rad; Hercules, CA), and analyzed with ImageJ
(National Institutes of Health; Bethesda, MD).
Live-cell imaging
For live cell imaging, mate-type heterothallic strains were grown in PMG with
appropriate supplements at 25
o
C or 32
o
C to late log phase (OD595=~0.8). Cells were concentrated
and washed in liquid ME and incubated at 25
o
C for 12-20 hours in an air shaker. Starved cells
were pelleted in a microfuge and spread on pads made with 2% agarose in liquid sporulation
66

media (SPAS) atop glass slides and sealed with 1:1:1 Vaseline/lanolin/paraffin (VaLaP, at
indicated ratios by weight). Imaging was carried out as described in Mastro and Forsburg (2014)
and Sabatinos et al. (2015) with a few modifications. Cells were staged at 25
o
C using a
DeltaVision microscope with softWorRx version 4.1 (GE Healthcare; Issaquah, WA) equipped
with a 60x (NA 1.4 PlanApo) lens, solid-state illuminator, and 12-bit CCD camera. Fluorescent
proteins were excited and detected with the following filter sets and exposure times: Green
Fluorescent Protein (GFP): (ex)475/28, (em)525/50, 0.15 seconds; DsRed or mCherry or Red
Fluorescent Protein (RFP): (ex)575/25, (em)632/60, 0.08 seconds; Cerulean or Cyan Fluorescent
Protein (CFP): (ex)438/24, (em)470/24, 0.15 seconds; Yellow Fluorescent Protein (YFP):
(ex)513/17, (em)559/38, 0.15 seconds. The following polychroic mirrors were used:
GFP/mCherry Chroma ET C125705 approximately: 520/50-630/80; Semrock CFP/YFP/DsRed
61008 bs approximately: 415/20-462/32-535/5—635/74. Long-term time-lapse videos used
thirteen 0.5-µm z-sections. Images were deconvolved and maximum-intensity projected
(softWoRx). Projected fluorescence images were combined with transmitted light images.
Images were contrast-adjusted using an equivalent histogram stretch on all samples. Focus
discrimination was performed by applying a signal threshold twice as large as average nuclear
background. For images featuring fluorescent dots (i.e. smaller than foci), only the average
nuclear background was eliminated. When examining the spatial and temporal dynamics of
fluorescently tagged proteins, an event was determined to have changed if it failed to occur again
after three consecutive frames (each frame is 10 minutes long). At least two independent
experiments and five fields per experiment were assessed for each sample. Cell counts were
pooled and presented as proportions ± 95% confidence interval, which was calculated using sum
67

squared error rules. Significance was evaluated using a chi-squared test or Fisher’s exact test,
depending on sample size.
Flow cytometry and microscopy
Whole-cell fluorescence-activated cell sorting (FACS) was carried out as described in
detail in Sabatinos et al. (2009, 2015) with minor changes. To perform cell cycle analysis or
microscopy, cells were fixed in 70% ethanol. Following rehydration in 50 mM sodium citrate,
cells were treated with 1 µM SytoxGreen (Invitrogen; Carlsbad, CA) plus 10 µg/ml RNase A and
incubated at 36
o
C for 1-2 hours. Samples were then sonicated before being analyzed on a
FACScan machine (BD Biosciences; San Jose, CA). 4',6-diamidino-2-phenylindole (DAPI)
staining was done by first rehydrating fixed cells in phosphate-buffered saline (PBS) and placing
them to dry on positively charged glass slides. Mount solution (50% glycerol and 1 µg/ml DAPI)
was then applied before photographing cells on a Leica DMR wide-field epifluorescence
microscope equipped with a 63x objective lens (numerical aperture [NA] 1.62 Plan Apo), 100-W
Hg arc lamp for excitation, and a 12-bit Hamamatsu ORCA-100 charge-coupled device (CCD)
camera. Images were acquired using OpenLab version 3.1.7 (ImproVision; Lexington, MA)
software and analyzed with ImageJ (National Institutes of Health; Bethesda, MD).
Chromatin Immunoprecipitation
ChIP was performed as described in Rougemaille et al. (2008) with minor changes.
Briefly, synchronous meiosis was achieved as mentioned above. Cells were harvested every
hour, treated with 37% formaldehyde followed by quenching with 0.25 M glycine, and washed
in Tris-Buffered Saline (TBS) prior to storage at -80
o
C. Pelleted cells were lysed by bead-beating
in a FastPrep machine (MP Biomedicals, LLC; Santa Ana, CA) and the resulting lysate was
68

sonicated four times for 15 seconds (15% duty cycle) with a Branson Sonifier 450 microtip
sonicator. Protein concentration was determined using the Bradford method. Some of the lysate
was stored as input, while the rest was treated with 1:200 mouse anti-GFP antibody (abcam 290)
(abcam; Cambridge, MA) and later combined with protein A Dynabeads (Invitrogen; Carlsbad,
CA). After several washes (in lysis buffer, wash buffer, and Tris-EDTA), the protein-DNA
portion was separated from the Dynabeads using TE and 1% SDS at 65
o
C for 30 minutes,
followed by centrifugation. To reverse the formaldehyde crosslink, the resulting elution was then
incubated at 65
o
C for at least 12 hours (input included). Afterward, 2 mg/ml Proteinase K
(Bioline Reagents Ltd.; Taunton, MA) was used for 2 hours at 37
o
C to eliminate any remnant
protein from the samples. DNA was isolated with a Qiagen PCR Purification Kit and analyzed
by PCR/agarose gel electrophoresis. ImageJ (National Institutes of Health; Bethesda, MD) was
used for sample quantification.
2.6 Supplemental Data

WT ∆swi3
69

Supplemental Figure 2.1 Replication dynamics in FPC mutants lacking Rec12. Synchronous meiosis in mat2-
102/h- pat1-114/pat1-114 ∆rec12 diploids. (A) FACS profiles showing replication progression following meiotic
induction. (B) Agarose gel images showing results from PFGE of indicated strains. (C) Quantification of meiotic
progression by DAPI-nuclei counts.

Supplemental Figure 2.2 Chromosome fragmentation in FPC mutants. (A) Cartoon depicting the products of
divisions showing temporary or permanent chromosome fragmentation in MI and MII. (B) Quantification of live-
cell images of meiotic cells carrying fluorescently tagged histone3 (Hht1-mRFP) and displaying nuclear
fragmentation during chromosome segregation. More than 150 cells were scored from at least two independent
movies for each genotype. Chi-squared analysis was used to determine significance. P-values are reported as
follows: p<0.05 *, p<0.01 **, p<0.001 ***. Error bars represent 95% confidence intervals.
Supplemental Table 2.1 List of strains used in this work. FY number is the digit identification given to fission
yeast strains stored in the Forsburg Lab cryogenic collection. Genotype denotes the gene deletions and specific
alleles of each collection strain. Source indicates whether a strain was taken out of the lab stock or if it was newly
constructed for the experiments in this study. Strains are listed from oldest to most recent, as indicated by their
collection identification number. Although most strains have been thoroughly genotyped, those without a specific
allele show a phenotype notation (e.g. leu-). (See next page for table)

70

71

72

2.7 Movie legends
Movie 2.1: Representative live cell imaging of meiosis in a WT heterothallic cross of cells homozygous for RPA-
YFP, Rad52-CFP, and H3-mRFP. Green is used as false color for RPA-YFP, blue for Rad52-CFP, and magenta for
histone H3-mRFP. Scare bar is 5 µm. Time frames are every 10 minutes.
Movie 2.2: Representative live cell imaging of meiosis in a ∆swi1 heterothallic cross of cells homozygous for RPA-
YFP, Rad52-CFP, and H3-mRFP. Green is used as false color for RPA-YFP, blue for Rad52-CFP, and magenta for
histone H3-mRFP. Scale bar is 5 µm. Time frames are every 10 minutes.
Movie 2.3: Representative live cell imaging of meiosis in a ∆swi3 heterothallic cross of cells homozygous for RPA-
YFP, Rad52-CFP, and H3-mRFP. Green is used as false color for RPA-YFP, blue for Rad52-CFP, and magenta for
histone H3-mRFP. Scale bar is 5 µm. Time frames are every 10 minutes.
Movie 2.4: Representative live cell imaging of meiosis in a WT heterothallic cross of cells homozygous for histone
H3-mRFP. Magenta is used as false color for histone H3-mRFP. Scale bar is 5 µm. Time frames are every 10
minutes.
Movie 2.5: Representative live cell imaging of meiosis in a ∆swi1 heterothallic cross of cells homozygous for
histone H3-mRFP. Magenta is used as false color for histone H3-mRFP. Scale bar is 5 µm. Time frames are every
10 minutes.
Movie 2.6: Representative live cell imaging of meiosis in a ∆swi3 heterothallic cross of cells homozygous for
histone H3-mRFP. Magenta is used as false color for histone H3-mRFP. Scale bar is 5 µm. Time frames are every
10 minutes.
Movie 2.7: Representative live cell imaging of meiosis in a WT heterothallic cross of cells heterozygous for LacI-
GFP and lacO near centromere I and homozygous for H3-mRFP. Green is used as false color for LacI-GFP and
magenta for histone H3-mRFP. Scale bar is 5 µm. Time frames are every 10 minutes.
Movie 2.8: Representative live cell imaging of meiosis in a ∆swi1 heterothallic cross of cells heterozygous for LacI-
GFP and lacO near centromere I and homozygous for H3-mRFP. Green is used as false color for LacI-GFP and
magenta for histone H3-mRFP. Scale bar is 5 µm. Time frames are every 10 minutes.
Movie 2.9: Representative live cell imaging of meiosis in a ∆swi3 heterothallic cross of cells heterozygous for LacI-
GFP and lacO near centromere I and homozygous for H3-mRFP. Green is used as false color for LacI-GFP and
magenta for histone H3-mRFP. Scale bar is 5 µm. Time frames are every 10 minutes.
Movie 2.10: Representative live cell imaging of meiosis in a WT heterothallic cross of cells homozygous for LacI-
GFP and lacO near centromere I as well as for H3-mRFP. Green is used as false color for LacI-GFP and magenta
for histone H3-mRFP. Scale bar is 5 µm. Time frames are every 10 minutes.
Movie 2.11: Representative live cell imaging of meiosis in a ∆swi1 heterothallic cross of cells homozygous for
LacI-GFP and lacO near centromere I as well as H3-mRFP. Green is used as false color for LacI-GFP and magenta
for histone H3-mRFP. Scale bar is 5 µm. Time frames are every 10 minutes.
Movie 2.12: Representative live cell imaging of meiosis in a ∆swi3 heterothallic cross of cells homozygous for
LacI-GFP and lacO near centromere I as well as H3-mRFP. Green is used as false color for LacI-GFP and magenta
for histone H3-mRFP. Scale bar is 5 µm. Time frames are every 10 minutes.
Movie 2.13: Representative live cell imaging of meiosis in a WT heterothallic cross of cells homozygous for Rec27-
GFP and H3-mRFP. Green is used as false color for Rec27-GFP and magenta for histone H3-mRFP. Scale bar is 5
µm. Time frames are every 10 minutes.
73

Movie 2.14: Representative live cell imaging of meiosis in a ∆swi1 heterothallic cross of cells homozygous for
Rec27-GFP and H3-mRFP. Green is used as false color for Rec27-GFP and magenta for histone H3-mRFP. Scale
bar is 5 µm. Time frames are every 10 minutes.
Movie 2.15: Representative live cell imaging of meiosis in a ∆swi3 heterothallic cross of cells homozygous for
Rec27-GFP and H3-mRFP. Green is used as false color for Rec27-GFP and magenta for histone H3-mRFP. Scale
bar is 5 µm. Time frames are every 10 minutes.
Movie 2.16: Representative live cell imaging of meiosis in a WT heterothallic cross of cells homozygous for Sgo1-
GFP and H3-mRFP. Green is used as false color for Sgo1-GFP and magenta for histone H3-mRFP. Scale bar is 5
µm. Time frames are every 10 minutes.
Movie 2.17: Representative live cell imaging of meiosis in a ∆swi1 heterothallic cross of cells homozygous for
Sgo1-GFP and H3-mRFP. Green is used as false color for Sgo1-GFP and magenta for histone H3-mRFP. Scale bar
is 5 µm. Time frames are every 10 minutes.
Movie 2.18: Representative live cell imaging of meiosis in a ∆swi3 heterothallic cross of cells homozygous for
Sgo1-GFP and H3-mRFP. Green is used as false color for Sgo1-GFP and magenta for histone H3-mRFP. Scale bar
is 5 µm. Time frames are every 10 minutes.
Movie 2.19: Representative live cell imaging of meiosis in a WT heterothallic cross of cells homozygous for Rec8-
GFP and H3-mRFP. Green is used as false color for Rec8-GFP and magenta for histone H3-mRFP. Scale bar is 5
µm. Time frames are every 10 minutes.
Movie 2.20: Representative live cell imaging of meiosis in a ∆swi1 heterothallic cross of cells homozygous for
Rec8-GFP and H3-mRFP. Green is used as false color for Rec8-GFP and magenta for histone H3-mRFP. Scale bar
is 5 µm. Time frames are every 10 minutes.
Movie 2.21: Representative live cell imaging of meiosis in a ∆swi3 heterothallic cross of cells homozygous for
Rec8-GFP and H3-mRFP. Green is used as false color for Rec8-GFP and magenta for histone H3-mRFP. Scale bar
is 5 µm. Time frames are every 10 minutes.
Movie 2.22: Representative live cell imaging of meiosis in a WT heterothallic cross of cells homozygous for Swi6-
GFP and Cnp1-mCherry. Green is used as false color for Swi6-GFP and magenta for Cnp1-mCherry. Scale bar is 5
µm. Time frames are every 10 minutes.
Movie 2.23: Representative live cell imaging of meiosis in a ∆swi1 heterothallic cross of cells homozygous for
Swi6-GFP and Cnp1-mCherry. Green is used as false color for Swi6-GFP and magenta for Cnp1-mCherry. Scale
bar is 5 µm. Time frames are every 10 minutes.
Movie 2.24: Representative live cell imaging of meiosis in a ∆swi3 heterothallic cross of cells homozygous for
Swi6-GFP and Cnp1-mCherry. Green is used as false color for Swi6-GFP and magenta for Cnp1-mCherry. Scale
bar is 5 µm. Time frames are every 10 minutes.
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81

Appendix 1: Tetrad Dissection in Fission Yeast
The work in this chapter was submitted for review to Methods in Molecular Biology (Springer)
in October 2016.
1.1 Overview
Tetrad dissection is a powerful tool in yeast genetics that allows the analysis of products
of a single meiosis. With just a few tetrads, it is possible to determine linkage, identify unique
phenotypes associated with double mutants, or assess specific meiotic defects. Strains are
crossed on nitrogen-limiting medium for 3 days. With the help of a micromanipulator, ripe asci
are isolated to spots 5 mm apart on a YES plate. Incubation at 36
o
C for about 3-5 hours is
necessary for the ascus walls to break down. Once the spores are released, they are individually
placed in a row containing four tetrad products, separated by 5 mm. The spores are then put in
the appropriate temperature for the cross until colonies form, and phenotypes are assessed by
replica plating or microscopic analysis.
Key Words: fission yeast, Schizosaccharomyces pombe, meiosis, tetrad dissection, tetrad
analysis, linkage analysis, spore fate analysis
1.2 Introduction
Tetrad dissection is one of the great tools of yeast genetics because it allows analysis of the
products of a single meiosis. Combined with the principles of classical Mendelian genetics, this
provides substantial information. As few as ten tetrads dissected on a plate can determine linkage
or isolate unique interactions. The frequency of single versus double crossovers in a known
genetic interval can be assessed, and the effect of different mutations on the progression of
meiosis can be determined (Forsburg 2003; Forsburg and Rhind 2006; Moreno et al. 1991).
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Tetrad analysis is ideal at any time when it is necessary to identify a double mutant
unambiguously. For example, consider a situation where the phenotype of double mutants is
unknown or possibly lethal in a cross between ∆yfg1::ura4
+
and ∆yfg2::ura4
+
. Using tetrad
dissection, a non-parental ditype (NPD) tetrad that contains 2 Ura+ and 2 Ura- spores can be
isolated, which will occur on average in 1/6 tetrads for unlinked genes (Kohli et al. 1977). Even
if the double mutant is inviable, Mendelian genetics indicates that the presence of 2 Ura- spores
means the remaining two spores in that tetrad must have been the double mutants yfg1::ura4
+

yfg2::ura4
+
. An alternative method, random spore analysis (see chapter on RSA in fission yeast),
does not allow this certainty. A detailed description of the analysis of meiotic products can be
found in (Smith 2009).
Tetrad dissection also provides an opportunity to observe phenotypes associated with meiotic or
post-meiotic processes. For example, a tetrad showing a non-Mendelian, 1:3 segregation pattern
for a given marker is indicative of gene conversion. Mixed phenotypes observed in single
colonies suggest post-meiotic segregation (Forsburg 2003; Smith 2009). By spatially separating
tetrad products, it becomes easier to follow the fate of each spore and thus decipher the effect of
specific mutations on meiotic and post-meiotic outcomes.
1.3 Materials
For efficient tetrad dissection and analysis, the parent strains must grow in conditions that ensure
optimal cell viability. This requirement is often met by using rich medium, although minimal
medium can be employed if selection is required. Contrary to S. cerevisiae, mating and meiosis
are coupled in S. pombe. Consequently, it is not necessary to isolate diploids first. All that is
needed is to cross two strains on a nitrogen-limiting medium at a temperature range of 25
o
C-
83

29
o
C and to let them proceed to sporulation (Forsburg 2003; Forsburg and Rhind 2006; Moreno
et al. 1991; Sabatinos and Forsburg 2010).
Culture Media
1. Yeast extract plus supplements (YES): YE base plus 225 mg/l each of adenine, L-
histidine, L-leucine, uracil, and L-lysine. The corresponding solid medium contains 2%
(w/v) agar.
2. Edinburgh minimal medium (EMM): 3 g/l potassium hydrogen pthallate, 2.2 g/l
Na2HPO4, 5 g/l NH4CL, 20 g/l glucose, 20 ml/l salts (50x stock), 1 ml/l vitamins (1000x
stock), 0.1 ml/l minerals (10,000x). The corresponding solid medium contains 2% (w/v)
agar.
3. Malt extract (ME): 30 g/l bacto-malt extract plus 225 mg/l each of adenine, histidine,
leucine, and uracil. Adjust to pH 5.5. The corresponding solid medium contains 2% (w/v)
agar.
4. Sporulation agar with supplements (SPAS): 10 g/l glucose, 1 g/l KH2PO4, 1 ml/l 1000x
vitamin stock, 45 mg/l of each: adenine, histidine, leucine, uracil, and lysine
hydrochloride. The corresponding solid medium contains 2% (w/v) agar.
5. 50x salt sock: 522.2 g/l MgCl2●6H2O, 0.735 g/l CaCl2●2H2O, 50 g/l KCl, 2 g/l Na2SO4.
6. 1000x vitamin stock: 1 g/l pantothenic acid, 10 g/l nicotinic acid, 10 g/l inositol, and 10
mg/l biotin.
7. 10,000x mineral stock: 5 g/l boric acid, 4 g/l MnSO4, 4 g/l ZnSO4●7H2O, 2 g/l
FeCl2●6H2O, 1 g/l KI, 0.4 g/l molybdic acid, 0.4 g/l CuSO4●5H2O, 10 g/l citric acid.
8. 2000x phloxin B: 5 mg/l phloxin B (Sigma, P4030). The corresponding volume in solid
medium contains 0.5 ml/l.
84

Preparation of tetrads
1. Sterile H2O
2. Sterile flat toothpicks
3. Microscope with micromanipulator
4. Stereo microscope with plate-holding stage
5. 32
o
C or 36
o
C Incubator
6. 25
o
C-29
o
C incubator
1.4 Methods
It is important to note that in S. pombe it is not necessary to select diploids to do a simple cross,
because mating is normally followed by sporulation without further manipulation. Two haploids
can be mixed on malt extract and after a few days the mating cells will form a banana-shaped
zygotic ascus and then proceed directly into meiosis. If a diploid must be created first as is the
case when crossing an h
90
cell (which prefers to cross with itself) with either an h
+
or h
-
cell, it is
recommended to use complementing markers in the parent strains (e.g. h
90
ade6-M210 X h
+

ade6-M216) to ensure efficient mating. The azygotic ascus resulting from this cross will look
linear and upon subsequent sporulation, the h
90
allele will segregate in a 2:2 Mendellian ratio
with the h
+
allele. Sometimes a zygotic ascus can arise from a diploid strain, but this usually
represent two diploids mating (Forsburg 2003; Moreno et al. 1991; Sabatinos and Forsburg
2010).
Although tetrad dissection is identical to that in S. cerevisiae, including the use of the same
apparatus, S. pombe asci do not need prior digestion with glusulase because spores fall apart on
their own. From a mating plate, ripe asci are transferred to rich solid medium. With the aid of a
85

micromanipulator, individual tetrads are isolated and then incubated at 32
o
C to 36
o
C for 3-5
hours to allow for the ascus walls to break down. Released spores are then picked and separated
in a single tetrad line. Once all tetrads are dissected, they are incubated at the appropriate
temperature for optimal growth (Forsburg 2003; Forsburg and Rhind 2006).
The use of a micromanipulator to separate tetrad products takes time and practice to master, but
the vast amounts of information investigators can obtain make it an invaluable genetic tool. In
addition to manual micromanipulators associated with table top microscopes (Zeiss, Micro Video
Instruments, Tritech), there are also automated systems (Singer Instruments). The critical
component is the glass needle, which can be hand-pulled from thin glass pipettes, or pre-made
from fiber optics (e.g., Singer, CoraStyles.com). Given that S. pombe does not have sticky
spores, it is necessary to use a high quality needle with a completely flat plane to ensure surface
tension makes spores adhere to the needle
Strain Cross Mating
1. Use a sterile toothpick to swipe a sample of cells from strain 1 and make a small patch on
a mating (ME) or sporulation (SPAS) plate. Use a different sterile toothpick to take a
similar amount of cells from strain 2 and add it to the previous patch.
2. Add 5 µl sterile water and mix cell matter gently on the surface of the agar with a sterile
toothpick.
3. Incubate patch plate at 25
o
C to 29
o
C for as long as it is appropriate for the cross and type
of plate.
Tetrad Dissection
86

1. Using a microscope, evaluate an ME or SPAS plate which has been incubated for 2 to 3
(zygotic) or 1 to 2 (azygotic) days at 25
o
C to 29
o
C to identify ripe asci. These should be
intact but must have clearly visible spores.
2. With a sterile toothpick, transfer and spread cells across the edge of the YES plate
(Figure 3.1A-C). On occasion, it may be necessary to use selective media, although
germination tends to be less efficient so YES is preferred.
3. Once intact ripe asci are identified, use a dissecting needle to pick up and move each
ascus to a fixed location on the plate (Figure 3.1A-C).
4. Incubate the plate a few hours to stimulate ascus breakdown. The process can be
accelerated by incubation at 36
o
C for two hours, if the strain is not temperature sensitive.
5. After appropriate incubation, return the plate to the micromanipulator and identify tetrads
that have released four large, round spores and which have left behind the faint skin of an
ascus. For each tetrad, place spores down the plate side-by-side in a line with an
intervening distance of 5 mm between them (Figure 3.1 A-C). Repeat step until all
isolated tetrads have popped.
6. When the plate is completed, incubate at the appropriate temperature (32
o
C for wild type,
25
o
C for temperature-sensitive mutants). Germination takes place within a few hours,
while it takes 2 to 5 days to observe colonies, depending on incubation temperature.
7. Spore fate can be monitored under a bench-top microscope once colonies are visible (2-3
days post dissection). Cells can be followed from the time of initial observation until no
more colonies emerge.
87

8. To screen the genotype resulting from each tetrad product, the colonies on the tetrad plate
must be stamped onto the appropriate media by replica plating. Score each colony for its
specific genotype and count each tetrad outcome type (see sections 3.3 and 3.4).
Linkage Analysis
1. Individual tetrads are classified by determining the segregation of pairs of markers into
parental ditype (PD), which is the configuration associated with the parents; non-parental
ditype (NPD), which contains completely recombinant spores, and tetratype (TT) which
contains 1 of each class (Figure 3.2). For unlinked markers, the number of PD is equal
to the number of NPD. As long as at least one marker is not centromere linked, the ratio
PD: NPD: TT will be 1:1:4. For linked markers, PD>NPD (Perkins 1949; Sherman
2002).

88

Figure 3.1 Tetrad dissection diagram. (A). Colonies from spores dissected on a YES plate. (B). Diagram showing
the main features of a tetrad dissection plate. The borderline (dashed line) of the “ripe asci spread zone” is 25 mm
from the right edge of the plate. In this zone, cell matter containing ripe asci is spread by streaking. The “ascus
breakdown zone” is where selected ripe asci (at least two) are placed with the micromanipulator for the release of
spores from their respective ascus. The “tetrad dissection zone” is where spores are deposited after ascus
breakdown. These are spaced apart every 5 mm. The grid produced in this zone shows each tetrad in individual rows
and their products are separated by each column. (C). Micrograph of two ripe asci found in the “ripe asci spread
zone.”
2. Given that fission yeast does not exhibit crossover interference, as in budding yeast,
mapping functions using the Perkins formula (see below) gives a reasonable
approximation of genetic distance (centimorgans, cM), but caution must be placed in
outcomes yielding distances higher than 60 cM. Conversely, outcome types resulting in
distances approximating or equaling 0 cM (most, if not PDs only) indicate a high degree
of linkage (Perkins 1949).
3. Perkin’s formula:
cM = [TT + 6NPD] / [PD + NPD + T], where genetic distance is in cM, tetratype is TT,
non-parental ditype is NPD, and parental ditype is PD (Perkins 1949).

89

Figure 3.2 Tetrad types resulting from recombination between two linked loci. In a cross between AB x ab
three tetrad classes can be obtained: parental ditype (PD), Non-parental ditype (NPD), and tetratype (T). The
presumed molecular events that produce these classes are classified as no crossover, single crossover, and double
crossovers involving multiple strands. The ratios shown below suggest the occurrence of linkage. If PD>NPD, the
observed genetic loci are linked. If loci are linked to their respective centromeres, the proportion of T decreases. A
1:1:4 ratio for PD:NPD:T indicates independent assortment.
Analysis of spore fate
1. After colonies begin to appear, check under a bench-top microscope the phenotypes
associated with no growth. These can take the form of a spore, a spore with a germinating
tube, or a small cluster of cells that form a micro colony (Figure 3.3 A-D).
2. Count all cell types (including those giving rise to visible colonies) from each dissected
tetrad. If missing colonies failed to grow due to spore-related cells types, spore failure
can be attributed for the phenotype. However, if a micro-colony forms, the problem may
stem from a post-meiotic event.
3. Compare spore fate ratios with those of strains with known phenotypes. Tetrads of wild
type strains produce spores that germinate and generate colonies more than 95% the time.
On the other hand, nearly ¾ of all tetrads in ∆rec12 mutants, which fail to create double
strand breaks in meiotic prophase, produce failed spores. Consequently, tetrad outcomes
showing reduced colony formation and numerous failed spores may suggest problems
specific to meiosis.
1.5 Notes
1. For multiple matings using the same strains, swipe a generous amount of each and dip
into 100 µl sterile water. Then, use a pipet to dispense 5 µl of each onto a mating patch
and mix each cross gently with a toothpick (Sabatinos and Forsburg 2010).
2. Mating of strains growing in liquid medium is possible but requires at 2- 3 washes in
nitrogen-free medium to avoid mating suppression.
90

3. In cases where cross mating is inefficient, dense growth in liquid medium is
recommended. After appropriate washing of 500 µl samples, resuspend cell precipitates
in 10 µl nitrogen-free medium. Combine resuspensions of the two strains and place 5 µl
of the mix onto an ME or SPAS plate.
4. To construct a diploid, mate cells carrying complementing alleles (e.g. ade6-M210 and
ade6-M216) on either an ME or SPAS plate. After 12 and 24 hours of mating, streak
some cells onto EMM plates lacking adenine to select for diploid cells. Since diploids are
prone to sporulate in minimal media, it is useful to restreak them onto YES plates with
phloxin B, which retards sporulation and where diploids stain a darker pink. Once
identified, streak diploids onto YES and store at 4
o
C for up to two weeks, after which
fresh diploids must be created anew to avoid selecting for sporulation-deficient mutants
(Forsburg 2003; Forsburg and Rhind 2010).
5. Mating and sporulation should be performed at 25
o
C (maximum of 29
o
C) for optimal
outcomes, given the temperature sensitivity of these processes.
6. When examining mating patches to identify ripe asci, it may be useful to spread cells
along the agar with a toothpick to make them more visible.
7. If tetrad dissection cannot be done when asci are ripe, the ME or SPAS plate can be
stored in the refrigerator for a day or two to retard ascus breakdown.
8. For efficient tetrad dissection with the micromanipulator, it is recommended that YES
plates are poured thin (15 ml to 20 ml), have a very flat surface, and are sufficiently dry
(at least a week old) to prevent asci sticking to the agar surface.
9. Once tetrads have been identified and isolated, ascus breakdown will vary according to
the strain’s temperature requirements. If not temperature sensitive, it can be incubated at
91

warmer temperatures (32
o
C to 36
o
C), which should considerably accelerate the process.
Otherwise, the YES plate can be incubated at room temperature or 17
o
C overnight.

Figure 3.3 Fate of dissected tetrad products. Spores were incubated at 32
o
C post dissection for 2-5 days. They
were monitored under the microscope for 2-3 days after colonies began to form. The following four spore fates were
observed: (A). Failed pore; (B). Spore with germination tube; (C). Micro-colony; (D). Colony.
10. Since it can be difficult to find the asci set aside for dissection after ascus breakdown, it is
advised that the agar surface be carefully perforated with the needle to mark the location
where each tetrad will be dissected.
11. To ensure spores adhere efficiently to the needle, tap lightly on the base of the
micromanipulator, which will cause vibration to facilitate the task. Likewise, careful
tapping will enable the release of spores from the needle to their designated locations on
the plate.
12. Replica plating onto solid media containing specific drugs or lacking individual nutrients
will enable genotype determination and will elucidate certain gene interaction types.
13. Despite its overall usefulness, the Perkin’s formula can at times underestimate genetic
distance because it does not account for multiple crossovers. To address this limitation, it
may be pertinent to use the Papazian’s equation (see below), which uses the Poisson
distribution to estimate the presence of multiple crossovers (Papazian 1952).
14. Papazian’s equation:
92

cM = 100 x -0.5 ln [PD - NPD] / [PD + NPD + TT] where genetic distance is in cM,
tetratype is TT, non-parental ditype is NPD, and parental ditype is PD (Papazian 1952).
1.6 Appendix 1 Bibliography
Forsburg, Susan L. "Growth and manipulation of S. pombe." Current Protocols in
Molecular Biology (2003): 13-16.
Forsburg, Susan L., and Nicholas Rhind. "Basic methods for fission yeast." Yeast 23, no.
3 (2006): 173-183.
Kohli, J., Hottinger, H., Munz, P., Strauss, A., & Thuriaux, P. “Genetic mapping in
Schizosaccharomyces pombe by mitotic and meiotic analysis and induced haploidization.”
Genetics 87, no 3 (1977) 471-489.
Moreno, Sergio, Amar Klar, and Paul Nurse. "Molecular genetic analysis of fission yeast
Schizosaccharomyces pombe." Methods in enzymology 194 (1991): 795-823.
Papazian, H. P. "The analysis of tetrad data." Genetics 37, no. 2 (1952): 175.
Perkins, David D. "Biochemical mutants in the smut fungus Ustilago maydis." Genetics
34, no. 5 (1949): 607.
Sabatinos, Sarah A., and Susan L. Forsburg. "Molecular genetics of
Schizosaccharomyces pombe." Methods in enzymology 470 (2010): 759-795.
Sherman, Fred. “Getting started with yeast.” Methods in enzymology 350 (2002): 3-41.
Smith, Gerald R. "Genetic analysis of meiotic recombination in Schizosaccharomyces
pombe." Meiosis: Volume 1, Molecular and Genetic Methods (2009): 65-76.

93

Appendix 2: Random Spore Analysis in Fission Yeast
The work in this chapter was submitted for review to Methods in Molecular Biology (Springer)
in October 2016.
2.1 Overview
Random spore analysis (RSA) is a tool that allows for the screening of a large number of
meiotic products. It requires only limited effort, and is often the method of choice for
constructing strains with unambiguous genotypes. It is also useful to identify the frequency of
rare events. Strains are crossed on nitrogen-limiting medium for three days. Mated cells are
observed under the microscope to check for the presence of ripe asci. To release spores from
their ascus, a sample of the cross is taken from the mating plate and resuspended in enzyme
solution overnight at 25
o
C-29
o
C. Spores are then counted using a hemocytometer before plating
an appropriate number. Incubation at the appropriate temperature follows until colonies form.
Key Words: fission yeast, Schizosaccharomyces pombe, meiosis, random spore analysis, strain
construction, linkage analysis, plasmid recovery, lethal allele analysis, plasmid complementation
2.1 Introduction
Random Spore Analysis (RSA) is an efficient method to screen large numbers of meiotic
products in S. pombe. Treatment of ripe asci with glusulase (snail gut enzyme) not only breaks
down the ascus wall to release spores, but also kills any vegetative cells that did not mate.
Moreover, unlike S. cerevisiae, S. pombe spores do not stick to one another following digestion
and are readily dispersed. This facilitates the analysis of a large number of spores in a manner
that is both time and labor efficient compared to tetrad dissection. RSA is particularly useful for
screening rare outcomes. However, it is not always the appropriate choice: for example, RSA
94

should not be employed if a desired double mutant cannot be unambiguously identified in the
pool, or if the genotypes of offspring are not equally viable, or if the products of a single meiosis
need to be compared. Although most common strain construction requirements are satisfied by
RSA, when in doubt, it is recommended to pull tetrads (Forsburg 2003; Forsburg and Rhind
2006; Gould 2004; Sabatinos and Forsburg 2010).
During strain construction, RSA can be particularly helpful if the alleles of interest in the desired
strain are marked differently. For example, the double mutant offspring of a cross between
yfg1::ura4
+
and yfg2::leu1
+
can be easily identified through isolation of strains that are Ura+ and
Leu+. However in a cross between from mating yfg1::ura4
+
with yfg2::ura4
+
, the double
mutant would show an Ura4+ phenotype that is undistinguishable from the single mutant. In the
latter example, tetrad dissection is required to discriminate the double mutant by isolating the
non-parental ditype (NPD) tetrad carrying 2 ura4+ and 2 ura4- spores (see Chapter on Tetrad
dissection).
RSA is helpful to examine linkage. For instance, by calculating the recombination
frequencies between leu1 and his7 from a pool of many individual crosses, the degree of linkage
between the two genes can be determined. Since there is little recombination between linked
genes, RSA affords sufficient numbers for robust statistics to elucidate the appropriate
segregation ratios. Indeed, processing large number of crosses with relative ease makes RSA
ideal for determining recombination frequencies in large mapping studies (Forsburg 2003;
Forsburg and Rhind 2006; Kohli et al.1977). Using the same logic, the frequency of any rare
meiotic event can be readily determined by this method.
RSA can also be employed in recovering spores containing plasmids. S. pombe, unlike S.
cerevisiae, has centromeres that are too large for shuttle vectors to carry and are typically
95

unstable in meiosis. As a result, roughly 10% of spores contain the plasmid (Forsburg 2003;
Forsburg and Rhind 2006). Since RSA allows screening of large numbers, desired strains are
readily isolated. Finally, RSA can be employed in spore germination experiments to analyze the
phenotype of lethal mutations (Hayles and Nurse 1992; Liang et al. 1999). In this strategy
consider a diploid cell heterozygous for yfg3
+
and yfg3::ura4
+
following sporulation. RSA
and/or tetrad analysis shows no viable Ura+ offspring on plates. By scaling up the experiment
and inoculating the RSA in liquid culture selecting for Ura+, only the cells containing the
disruption will be able to germinate. Samples are taken during a time course and monitored for
cell and nuclear morphology as well as DNA content (Liang et al. 1999).
2.3 Materials
It is recommended that the strains to be crossed grow robustly first on YES plates (or
selection plates where required) to ensure optimal mating results. Unlike S. cerevisiae, S. pombe
cells can grow as haploids and readily mate when starved of nitrogen, which is required for entry
into meiosis.
Culture Media
9. Yeast extract plus supplements (YES): YE base plus 225 mg/l each of adenine, L-
histidine, L-leucine, uracil, and L-lysine. The corresponding solid medium contains 2%
(w/v) agar.
10. Edinburgh minimal medium (EMM): 3 g/l potassium hydrogen pthallate, 2.2 g/l
Na2HPO4, 5 g/l NH4CL, 20 g/l glucose, 20 ml/l salts (50x stock), 1 ml/l vitamins (1000x
stock), 0.1 ml/l minerals (10,000x). The corresponding solid medium contains 2% (w/v)
agar.
96

11. Malt extract (ME): 30 g/l bacto-malt extract plus 225 mg/l each of adenine, histidine,
leucine, and uracil. Adjust to pH 5.5. The corresponding solid medium contains 2% (w/v)
agar.
12. Sporulation agar with supplements (SPAS): 10 g/l glucose, 1 g/l KH2PO4, 1 ml/l 1000x
vitamin stock, 45 mg/l of each: adenine, histidine, leucine, uracil, and lysine
hydrochloride. The corresponding solid medium contains 2% (w/v) agar.
13. 50x salt sock: 522.2 g/l MgCl2●6H2O, 0.735 g/l CaCl2●2H2O, 50 g/l KCl, 2 g/l Na2SO4.
14. 1000x vitamin stock: 1 g/l pantothenic acid, 10 g/l nicotinic acid, 10 g/l inositol, and 10
mg/l biotin.
15. 10,000x mineral stock: 5 g/l boric acid, 4 g/l MnSO4, 4 g/l ZnSO4●7H2O, 2 g/l
FeCl2●6H2O, 1 g/l KI, 0.4 g/l molybdic acid, 0.4 g/l CuSO4●5H2O, 10 g/l citric acid.
16. 2000x phloxin B: 5 mg/l phloxin B (Sigma, P4030). The corresponding volume in solid
medium contains 0.5 ml/l.
17. Glusulase solution (PerkinElmer #NEE154001)
Preparation of spores
1. Sterile H2O
2. Sterile flat toothpicks
3. Light microscope
4. 25
o
C-29
o
C incubator
5. Hemocytometer

97

2.4 Methods
Crossing cells for strain construction or analysis of meiotic products involves mixing two
strains of opposite mating type on starvation medium and letting them undergo meiosis at 25
o
C-
29
o
C (Forsburg 2003; Forsburg and Rhind 2006; Hayles and Nurse 1992). If very efficient
sporulation is required or the cross involves one h
90
parent (which prefers to cross with itself), a
diploid can be generated using complementing nutritional markers. S. pombe has a convenient
pair of ade6 (ade6-M210 and ade6-M216) alleles which when put together offer diploid cells an
Ade+ phenotype via intragenic complementation. After identifying a diploid colony, place on
starvation medium to induce sporulation. In the absence of ade6 markers, any pair of auxotrophic
markers can be employed to construct a diploid, but caution must be exercised to verify that the
recovered strain is in fact a diploid and not simply a recombinant haploid spore (Forsburg 2003;
Forsburg and Rhind 2006; Gould 2004; Moreno et al. 1991).
Strain Cross Mating
1. Using a sterile toothpick, pick some cell matter from each of two strains and create a
mating patch by mixing them with 5 µl H2O on an ME (or SPAS) plate.
2. Incubate mating plate at 25
o
C to 29
o
C until colonies form.
3. Matings can also be performed in liquid media.
Random Spore Analysis
1. Observe under a microscope (phase contrast, 20-40x objective) for the presence of ripe
asci in the mating patch. Zygotic asci, resulting from the cross of two haploid cells with
opposite mating types, are banana-shaped, while azygotic asci, arising from diploid cells,
are linear. For ripe tetrads, spores inside the asci should be clearly visible.
98

2. When the cross has sufficiently sporulated, pick a substantial amount of cell matter from
a mating patch with a toothpick and put it into a 0.5% glusulase solution (1:10 dilution of
5% glusulase in sterile H2O). Vortex the glusulase suspension and incubate for 12-16
hours at 25
o
C.
3. Dilute the glusulase suspension 1:5-1:20 in sterile H2O and count spore number using a
hemocytometer.
4. It is important to pay close attention to the cells under observation. Spores are small,
round, and highly refractive. Vegetative cells are rod-shaped and look dark from the
glusulase treatment (Figure 4.1A).
5. Plate 200-1000 spores onto YES plates and incubate at the appropriate temperature until
colonies form (Figure 4.1B).
6. Replicate plate onto appropriate media to determine candidates for analysis.
7. The remaining spores can be washed in sterile H2O and stored at 4
o
C for up to a month,
should further plating be required.
Spore Viability Analysis
1. Count the number of colonies that grow on each YES plate.
2. Divide colony counts by microscope counts (number of spores plated) to derive the
proportion of spores that are viable (Smith 2009).
Viable Spore Yield Analysis
1. Before making a cross, use a hemocytometer to count the number of cells in the parent
with the least cell density.
99

2. Divide the number of spores in the glusulase suspension by the number of cells in the less
dense parent of the cross. This gives the proportion of viable spores per viable cell
(theoretically 2 per cell).
3. Compare figures to that of the wild type control to obtain a relative value that better
indicates mating efficiency of the cross and the extent of mitotic growth on starvation
medium (Smith 2009).

Figure 4.1 Counting and plating spores for RSA. (A). One of 25 hemocytometer squares showing three spores
(white arrows). Notice the presence of cell debris (black open arrow) and abnormally shaped spores (black arrow).
91 spores where counted in this sample (within the linear range: 50-100), which came from a 1/10 dilution. Thus,
the number of spores per µl in the original glusulase suspension is: 91 (spores counted) x 10 (dilution factor) x 10
(hemocytometer factor) = 9,100. This number is then multiplied by 1/200, which is the dilution from which spores
are plated (9,100 x 1/200 = 45.5 spores/µl). To obtain the volume required from the 1/200 dilution to dispense 500
spores calculate as follows: 500 spores/45.5 spores/µl = 10.9 µl. Place 89.1 µl sterile H 2O onto a YES plate
containing glass beads and add 10.9 µl spore suspension. (B). Colony growth resulting from spores counted in A.
There are 259 viable colonies, which gives an estimated 51.8% spore viability for this strain (i.e. [259 colonies/500
counted spores] x 100). Take note of the colony crowding in just 259 viable spores. Figures above 500 make colony
counts unreliable.
Recombination Frequency Analysis
1. Calculate the number of recombinant colonies and divide this value by the total number
of spores screened to obtain the recombinant frequency.
100

2. Convert recombinant frequency into genetic distance (centimorgans, cM) using
Haldane’s formula (see below), which is appropriate in fission yeast due to lack of
crossover interference (Smith 2009).
3. Haldane’s formula:
cM = -50 ln (1-2R), where genetic distance is in cM and R is the recombinant fraction (a
figure between 0-0.5) (Smith 2009).
2.5 Notes
1. To improve mating efficiency, grow each strain in 2 ml liquid YES. Take 500 µl sterile
H2O and wash cells in nitrogen-free medium. Resuspend cell pellets in 10 µl of the same
washing medium. Combine 5 µl of resuspension from each strain and mix the patch with
a pipette.
2. To generate a diploid, cross strains with complementing alleles (e.g. ade6-M210 and
ade6-M216) on either an ME or SPAS plate. After 12 and 24 hours of mating, streak
some cells onto EMM plates lacking adenine to select for diploid cells. Diploids are
prone to sporulate in minimal media. Thus it is useful to restreak them onto YES plates
with phloxin B, which will delay sporulation and will stain diploids a darker pink. Once
confirmed, streak diploids onto YES and store at 4
o
C for up to two weeks, after which
fresh diploids must be created again to avoid selecting for sporulation-deficient mutants
(Forsburg 2003; Forsburg and Rhind 2006).
3. If faster glusulase treatment is required, incubate glusulase cell suspension at 32
o
C-36
o
C
for 4-6 hours. Dilute the suspension 1:5-1:20 in sterile H2O and count spores with a
hemocytometer before dispersing them on YES plates. This is only recommended for
strain construction and not for spore viability analysis.
101

4. Avoid counting spores that do not look round-shaped or which no longer refract light
well, as these have most likely been compromised by glusulase (Figure 4.1A).
5. Make sure to use appropriate dilution practices when counting spores to avoid being
outside the hemocytometer’s linear range. For example, VWR Scientific Counting
Chamber’s range is between 50-100 cells. Divide the upper and lower end of the range by
25 (the number of squares within the hemocytometer grid) to obtain the optimal number
of spores per square; in this case 2-4 (Figure 4.1A). Consult the manufacturer’s manual
to find your hemocytometer’s specific range.
6. Spore dispersal with glass beads ensures greater control of spore suspension volume loss.
Use the same number of beads (e.g. 7) and make sure to shake plates evenly on a
horizontal plane.
7. The number of spores plated depends upon the purpose of the cross. If all spores are
expected to be viable, and screened by replica plating, between 100- 500 should be plated
on a standard sized Petri dish to ensure sufficient separation of colonies (Figure 4.1B).
On the other hand, if there is a direct screen to be applied, with only a few viable colonies
expected on selective media, several thousand can be plated. In this case, a non-selective
control plate for total viability of the population should also be determined.
2.6 Appendix 2 Bibliography
Forsburg, Susan L. "Growth and manipulation of S. pombe." Current Protocols in
Molecular Biology (2003): 13-16.
Forsburg, Susan L., & Nicholas Rhind. "Basic methods for fission yeast." Yeast 23, no. 3
(2006): 173-183.
Gould, K. L. “Protocols for experimentation with Schizosaccharomyces pombe.”
Methods 33, no. 3 (2004): 187-188.
Hayles, J., & Nurse, P. “Genetics of the fission yeast Schizosaccharomyces pombe.”
Annual review of genetics 26, no. 1 (1992): 373-402.
102

Kohli, J., Hottinger, H., Munz, P., Strauss, A., & Thuriaux, P. “Genetic mapping in
Schizosaccharomyces pombe by mitotic and meiotic analysis and induced haploidization.”
Genetics 87, no.3 (1977): 471-489.
Liang, D. T., Hodson, J. A., & Forsburg, S. L. “Reduced dosage of a single fission yeast
MCM protein causes genetic instability and S phase delay.” Journal of Cell Science 112, no.
4 (1999): 559-567.
Moreno, Sergio, Amar Klar, & Paul Nurse. "Molecular genetic analysis of fission yeast
Schizosaccharomyces pombe." Methods in enzymology 194 (1991): 795-823.
Sabatinos, Sarah A., and Susan L. Forsburg. "Molecular genetics of
Schizosaccharomyces pombe." Methods in enzymology 470 (2010): 759-795.
Smith, Gerald R. "Genetic analysis of meiotic recombination in Schizosaccharomyces
pombe." Meiosis: Volume 1, Molecular and Genetic Methods (2009): 65-76.

103

Appendix 3: Genetic screen for suppressors of dfp1-rad35, a truncation allele of dfp1+ with
meiosis-specific defects
3.1 Overview
In this screen, we sought to find targets of DDK that functioned either downstream of
dfp1 or in a parallel pathway. The purpose was to find candidates acting in nucleotide
metabolism or proximal pathways that could explain rad35’s sensitivity to alkylation damage
and its multiple meiotic defects. We took a random approach by transforming plasmids carrying
cDNA pieces from transcripts harvested in meiosis. We exposed rad35 cells to methyl-
methanesulfonate (MMS, an alkylating agent) on YES plus phloxin B plates. This allowed us to
distinguish cells that were MMS-sensitive from those that were not. For candidates that
suppressed rad35’s MMS sensitivity, we confirmed that such phenotype was due to
overexpression of the cDNA piece and not due to other circumstances by re-streaking the clone
with and without plasmid induction. After corroborating potential candidates by re-streaking, we
isolated each plasmid and sequenced the cDNA within in. Subsequently, we performed a
homology search to find out the identity each rad35 suppressor. Lastly, we re-introduced a
different plasmid carrying each positive candidate and checked if it recapitulated suppression of
rad35’s MMS sensitivity (Figure 1).
3.2 Introduction
DDK is an essential kinase with multiple functions that ensure genome stability in
organisms from yeast to human. In fission yeast, it has been found to play an important role in
DNA replication, DNA repair, and checkpoint functions (Duncker and Brown, 2003; Kim et al.
2003; Labib, 2010; Sclafani, 2000). DDK is composed of Dfp1 and Hsk1. The budding yeast and
human orthologues are Dbf4 and Cdc7, respectively. Since Dbf4 regulates the activity of Cdc7,
the catalytic subunit, by targeting it to various substrates, the complex is better known as Dbf4-
104

Dependent Kinase or DDK for short. Dfp1’s multiple domains have been associated with distinct
functions. For instance, the N-terminal domain is involved in the replication checkpoint (Takeda
et al., 1999; Duncker et al. 2002). The middle domain (M) and the MIR motif are
correspondingly linked to general replication and replication at the centromere via association
with Swi6. In fact, Dfp1’s requirement for proper sister chromatid cohesion also stems from its
interaction with Swi6 (Bailis and Forsburg, 2004; Varrin et al., 2005). The C-terminal domain,
on the other hand, is dispensable for replication activities, but is necessary for responding to
alkylation damage and for normal meiosis (Fung et al., 2002; Gabrielse et al. 2006; Harkins et
al., 2009; Dolan et al., 2010).

Figure 5.1. Schematic of genetic screen for suppressors of dfp1-rad35. Fission yeast rad35 and wild type cells
were transformed with inducible plasmids (hence the ON & OFF labels throughout) carrying a library of cDNA
105

pieces harvested from meiotic cells. Transformed cells were spread on plates with MMS and phloxin B, where
MMS-resistance could be distinguished. After identification of possible rad35 suppressors, the phenotype was
confirmed by shutting the plasmid on and off. Once suppressors were confirmed by plating and streaking, plasmids
were isolated and cDNA pieces sequenced. Re-transformation of rad35 and wild type cells was performed to ensure
MMS-resistance phenotype was indeed correct for each positive candidate.
Work in budding yeast shows that DDK is required for meiotic replication initiation and
recombination (Buck et al., 1991; Sclafani et al., 1998). In both yeasts, DDK has been
implicated in the recruitment of Rec12/Spo11, which is responsible for generating programmed
double strand breaks (prDSBs) following replication (Ogino et al. 2006; Wan et al., 2006;
Sasanuma et al. 2008; Wan et al., 2008). A recent report in budding yeast demonstrates that this
recruitment is dependent on the fork protection complex (FPC) (Murakami and Keeney, 2014),
which is consistent with biochemical data in fission yeast that show physical association between
the FPC and DDK (Matsumoto et al., 2005; Shimmoto et al. 2009). Additionally, budding yeast
cells lacking Cdc7 exhibit faulty meiotic transcription and an overproduction of diploid dyads,
resulting from inappropriate monopolar attachment of sister kinetochores in MI (Tóth et al.,
2000; Rabitsch et al., 2003; Lo et al., 2008; Matos et al., 2008; Lo et al., 2012).
Mis-segregation phenotypes appear to be linked to DDK’s regulation of sister chromatid
cohesion. Following pre-meiotic S-phase (meiS), cells without DDK undergo anaphase I arrest.
However, when the meiotic cohesin subunit Rec8 is also removed, chromosome segregation is
restored. This may be directly connected to the observations that both Cdc7 and casein kinase 1
(CK1) separately function in the degradation of Rec8 (Valentin et al. 2006; Ishiguro et al. 2010;
Katis et al., 2010). In fact, a few pieces of evidence in fission yeast lend support to this idea.
Cells harboring a C-terminal truncation in Dfp1 (i.e. dfp1-rad35 cells) have low spore viability,
reduced prDSBs, delayed expression of meiotic genes, and abnormal chromosome segregation.
Some of these meiotic defects may originate from Dfp1’s inability to modulate phosphorylation
and degradation of Rec8, for dfp1-rad35 cells exhibit a high frequency of dyad formation and an
106

overall failure to progress past the MII division (Le et al. 2013). Put together, these data
highlight DDK’s important contribution to the correct execution of different meiotic events,
especially those dependent on the role of Dfp1/Dbf4.
3.3 Results
We were unable to identify suppressors that were immediately connected to DDK’s
known roles in replication, the replication and DNA damage checkpoints or in meiosis (Figure
5.2A-C). However, the identified genes come from various cellular processes including 5 that
function in and outside the nucleus, 3 in the mitochondria, and 2 in the Golgi. Interestingly, 1
candidate has a function in cytoskeleton dynamics, which could play a distant role in the mis-
segregation phenotypes of rad35 cells.
3.3 Discussion
Candidates that function in cellular energy and metabolic activities dominated the overall
breakdown of rad35 suppressors. It is possible that sick rad35 cells physiologically benefited
from a boost in functions important for cellular housekeeping. Other than speculating on how
DDK pathways may overlap with other cellular processes, we are at present incapable of making
direct connections that may offer mechanistic insights into how a dysfunctional dfp1 allele was
suppressed in conditions that render cells vulnerable to replication stress and DNA damage.
3.4 Materials and Methods
Detailed descriptions of general fission yeast culture conditions, media and standard
techniques such as electroporation are found in Sabatinos and Forsburg (2010). Liquid and solid
yeast extract plus supplements (YES) was used for the growth and maintenance of all strains,
except for when pombe minimal glutamate (PMG) with the appropriate supplements was
107

required. Cells were spread with glass beads on each plate. Plates were incubated at 25
o
C for 3-5
days, after which colonies were examined.
6
Figure 5.2. Suppressors of dfp1-rad35. (A) List of all rad35 suppressors including the gene name, function and
gene ontology description. (B) Percentage breakdown of cellular processes affected by the identified rad35
suppressors. (C) Diagram depicting the information presented in (A) & (B) in the context of cellular and
chromosome locations.

108

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Abstract (if available)

Abstract In addition to its essential function in replication initiation, the DDK protein kinase plays a role in the response to replication stress. A truncation allele in the Dfp1 subunit of DDK is proficient for replication, but is MMS sensitive with defects in induced mutagenesis. It is also defective at multiple steps in meiosis including the initiation of double strand breaks, transcription, and cleavage of Rec8 (the alpha-kleisin subunit of cohesin), accompanied by a severe loss of viability. We have taken a candidate approach to examine meiosis in mutants affecting DDK-associated proteins that also affect MMS sensitivity and fork stability. In this thesis, we examine components of the replication Fork Protection Complex (FPC): Swi1 (Tof1) and Swi3 (Csm3). We show that destabilization of the FPC decreases spore viability, increases sister chromatid exchange, reduces homologous recombination, and induces abnormal segregation in MI and MII divisions. We find that the elimination of Rec27 (a linear elements component), duration of Sgo1 before anaphase I and persistence of centromeric Rec8 are disrupted in FPC mutants. Moreover, as they progress though meiosis, FPC mutants show Rad11 (RPA) and Rad22 (Rad52) signals that indicate persistent DNA damage. Thus, the FPC stability is required for proper meiotic segregation. Our results suggest that the FPC suppresses excess ssDNA and helps to stabilize cohesion at the centromere, indicating that events in meiosis are sensitive to disruptions in replication fork stability.

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Creator Escorcia Torres, Wilber (author)

Core Title Destabilization of the replication fork protection complex is associated with meiotic defects in fission yeast

School College of Letters, Arts and Sciences

Degree Doctor of Philosophy

Degree Program Molecular Biology

Publication Date 11/15/2016

Defense Date 10/12/2016

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

Tag aneuploidy,chromosome fragmentation,cohesion,DNA damage,equational division,genome integrity,genome stability,meiosis,nondisjunction,OAI-PMH Harvest,reductional division,replication,replication fork protection complex,replication stability,segregation,Swi1,Swi3

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Advisor Forsburg, Susan L. (committee chair), Armani, Andrea (committee member), Curran, Sean (committee member), Michael, Matthew (committee member)

Creator Email escorcia@usc.edu,wilber.escorcia@gmail.com

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