University of Iowa
Iowa Research Online
Theses and Dissertations
Summer 2014
A role for the CSN/COP9 signalosome in
synaptonemal complex assembly and meiotic
progression
Heather Marie Brockway
University of Iowa
Copyright 2014 Heather Marie Brockway
This dissertation is available at Iowa Research Online: http://ir.uiowa.edu/etd/1296
Recommended Citation
Brockway, Heather Marie. "A role for the CSN/COP9 signalosome in synaptonemal complex assembly and meiotic progression." PhD
(Doctor of Philosophy) thesis, University of Iowa, 2014.
http://ir.uiowa.edu/etd/1296.
Follow this and additional works at: http://ir.uiowa.edu/etd
Part of the Genetics Commons
A ROLE FOR THE CSN/COP9 SIGNALOSOME IN SYNAPTONEMAL COMPLEX
ASSEMBLY AND MEIOTIC PROGRESSION
By
Heather Marie Brockway
A thesis submitted in partial fulfillment
of the requirements for the Doctor of
Philosophy degree in Genetics
in the Graduate College of
The University of Iowa
August 2014
Thesis Supervisor: Assistant Professor Sarit Smolikove
Copyright by
HEATHER MARIE BROCKWAY
2014
All Rights Reserved
Graduate College
The University of Iowa
Iowa City, Iowa
CERTIFICATE OF APPROVAL
_______________________
PH.D. THESIS
_______________
This is to certify that the Ph.D. thesis of
Heather Marie Brockway
has been approved by the Examining Committee
for the thesis requirement for the Doctor of Philosophy
degree in Genetics at the August 2014 graduation.
Thesis Committee: __________________________________
Sarit Smolikove, Thesis Supervisor
__________________________________
Andrew Forbes
__________________________________
Robert Malone
___________________________________
Amy Sparks
__________________________________
Tina Tootle
__________________________________
Lori Wallrath, Chair
To Alan M Brockway, you made this possible. You told me that despite my reservations,
I needed to make the most of the opportunity to finally get my PhD. Your encouragement
to pursue my dream will never be forgotten.
To Walter E Schubert, my dearest grandfather, you fed my passion for science since I
was a child. My fondest memories of my childhood are stargazing with you, especially
watching Halley’s Comet one cold clear winter night.
I miss both of you dearly and wish you could have been here to share in my success.
ii There’s as many atoms in a single molecule of your DNA as there are stars in the typical
galaxy. We are, each of us, a little universe.
Neil deGrasse Tyson
Cosmos
iii ACKNOWLEDGMENTS
I have traveled a very long road to this point and while the journey has been
difficult, many have made it bearable. I would like to thank Sarit Smolikove for giving
me the opportunity to follow my passion for reproductive genetics. I would also like to
thank past and present Smolikove lab members: especially Kristi Walker-Dexter and
Marcus Tatum for their assistance with my early work on the RNAi screen and Martha
Dean and Nathan Balukoff for their work on the CSN/COP9 project. Yizhi Yin for being
a gracious lab mate, reminding me there is always a bright side to everything, and most of
all being a good friend. Thank you to the rest of the Smolikove lab members who are not
mentioned, but will never be forgotten. I would also like to thank the members of my
committee for their input and insight into the various projects I have worked on. It is
much appreciated and has helped me grow as a scientist. To all of our colleagues who
kindly provided reagents, worm strains and fly stocks, thank you and your assistance is
much appreciated. Lastly, to my friends and colleagues, past and present, no mere words
can express the thanks I owe you all. You never let me quit and your encouraging words
carried me through even the darkest of days in the lab. For this, I will be forever grateful.
iv ABSTRACT
Defects in meiotic prophase I events, resulting in aneuploidy, are a leading cause
of birth defects in humans; however, these are difficult to study in mammalian systems
due to their occurrence very early in development. The nematode, Caenorhabditis
elegans, is an excellent model for prophase I studies as its gonad is temporally and
spatially organized around these meiotic events. Homolog pairing, synapsis, meiotic
recombination and crossover formation are essential to the proper segregation of
chromosomes into the respective gametes, either the egg or sperm. Disturbances in these
events leads to missegregation of chromosomes in the gametes in the meiotic divisions.
Synapsis is especially critical in meiosis as it precedes and is required for meiotic
recombination in C. elegans. The formation of the synaptonemal complex (SC) is
fundamental to chromosomal synapsis, yet the molecular mechanisms of synaptonemal
complex morphogenesis are largely unknown. The investigations described in this thesis
were undertaken to better understand the molecular contributions to synaptonemal
complex morphogenesis. Chapter One reviews knowledge of morphogenesis and its
relationship to the events of meiotic prophase I. Recent studies in our laboratory have
implicated AKIRIN, a nuclear protein with multiple biological functions, as having a role
in synaptonemal complex disassembly, specifically preventing the aggregation of
synaptonemal proteins (Clemons et al., 2013). As a result of our efforts to discern the
mechanism by which AKIRIN regulates disassembly, we found that the highly conserved
CSN/COP9 signalosome has a role in SC assembly, leading to defects in prophase I
events and in MAPK signaling , leading to the arrest of nuclei in the later stages of
meiosis. While the CSN/COP9 signalosome has been implicated in general fertility in C.
elegans (Pintard et al., 2003), no role had been defined in earlier meiotic stages until this
study. Chapter Two describes an RNAi enhancer/suppressor screen undertaken in the
v akir-1 mutant background. Several RNAi clones were selected for future study based on a
reduction in brood size; one of which, csn-5, is the focus of the analysis presented in
Chapter 3. Chapter Three describes the phenotypic characterization of two CSN/COP9
signalosome subunits, csn-2 and csn-5. Alleles of both genes display synaptonemal
complex protein aggregation and defects in mitotic cell proliferation, homologous
chromosome pairing, meiotic recombination and crossover formation, leading to an
increase in apoptosis. Oocyte maturation is also disrupted by a lack of MAPK signaling,
resulting in a lack of viable oocytes, which renders the csn mutant homozygotes sterile.
These findings support a model suggesting the CSN/COP9 signalosome has an essential
role in regulating meiotic prophase I events and oocyte maturation. Chapter 4 describes
the methodology used in this study. Chapter 5 provides a summary of the thesis findings
and examines the future directions to extend this work.
vi TABLE OF CONTENTS
LIST OF TABLES ............................................................................................................. ix
LIST OF FIGURES ............................................................................................................ xi
LIST OF ABBREVIATIONS ..........................................................................................xiii
CHAPTER 1 INTRODUCTION ......................................................................................... 1
Meiosis matters ................................................................................................ 1
C. elegan: A model for meiotic study .............................................................. 4
Early meiotic prophase I events in C. elegans ................................................. 8
Replication and the establishment of cohesion ......................................... 8
Pairing ....................................................................................................... 9
Synapsis................................................................................................... 13
Meiotic recombination and crossover formation .................................... 18
Late meiotic prophase I events ....................................................................... 23
The meiotic divisions .................................................................................... 25
The meiotic divisions .................................................................................... 27
CHAPTER 2 RNAI SCREEN FOR ENHANCERS AND SUPPRESSORES OF
MEIOTIC DEFECTS OF AKIR-1 MUTANTS ........................................... 36
Introduction .................................................................................................... 36
Results ............................................................................................................ 39
An RNAi screen identified potential genes
that interact with akir-1 ........................................................................... 39
Cytological analyses of chromosome II clones ....................................... 43
F10G7.4 ........................................................................................... 44
ZC239.6 ........................................................................................... 45
Cytological analyses of chromosome IV clones ..................................... 45
F38H4.9 ........................................................................................... 46
Y66H1B.2 ....................................................................................... 47
C09G4.3 .......................................................................................... 47
B0547.1 ........................................................................................... 48
Discussion ...................................................................................................... 49
F10G7.4 .................................................................................................. 49
F38H4.9 .................................................................................................. 51
Y66H1B.2 .............................................................................................. 52
C09G4.3 ................................................................................................. 53
Evaluation of screen design.................................................................... 55
CHAPTER 3 A NOVEL ROLE FOR THE CSN/COP9 SIGNALOSOME IN SC
ASSEMBLY ................................................................................................ 65
Introduction .................................................................................................... 65
CSN/COP9 signalosome ......................................................................... 65
Results ............................................................................................................ 68
csn mutants exhibit defects in SC morphogenesis and meiotic
progression .............................................................................................. 68
csn-2 and csn-5 mutants similarly affect CR assembly .......................... 71
vii csn-2 and csn-5 are required for gonad proliferation and fertility .......... 72
csn-2 and csn-5 are required for pairing stabilization ............................. 73
Meiotic recombination and crossover formation are preturbed in
csn mutants ............................................................................................. 74
Apoptosis is increased in csn mutatns and dpMPK-1 levels are
reduced ................................................................................................... 76
Discussion ...................................................................................................... 79
CSN/COP9 is required for meiotic prophase I events ............................ 79
CSN/COP9 is required for the normal levels of germline
proliferation .............................................................................................. 80
CSN/COP9 is required for meiotic progression and pachytene exit ........ 84
How the CSN/COP9 signalosome regulates chromosome synapsis ........ 86
The CSN subunits and their roles in meiotic events ................................ 89
Summary .................................................................................................. 93
CHAPTER 4 METHODS AND MATERIALS ............................................................. 122
C. elegans strains and culture conditions ..................................................... 122
PCR validation of strains.............................................................................. 122
RNAi screen for identification of genes enhancing or suppressing the
akir-1 phenotype........................................................................................... 123
Quantification of F1 progeny for RNAi screen ............................................ 124
RNAi feeding protocols................................................................................ 125
Microscopy ................................................................................................... 125
Quantification of immunostained foci.......................................................... 126
Fluorescence in situ hybridization (FISH) ................................................... 127
Western Analyses ......................................................................................... 127
Apoptotic analyses ....................................................................................... 129
CHAPTER 5 CONCLUSIONS AND FUTURE DIRECTIONS ................................... 132
A role for the CSN/COP9 signalosome in meiotic prophase I ..................... 133
The role for the CSN/COP9 signalosome in MAPK signaling .................... 136
Determining the substrates of the CSN/COP9 signalosome regulating
SC formation ............................................................................................... 137
APPENDIX A RNAI SCREEN METADATA ............................................................... 140
APPENDIX B THE ROLE OF AKIRIN IN DROSOPHILA OOGENESIS .................. 163
APPENDIX C EXPERIMENTS PERFORMED FOR CLEMONS ET AL., 2013 ....... 194
REFERENCES ............................................................................................................... 196
viii LIST OF TABLES
Table1-1: Conservation of meiotic factors ............................................................... 35
Table2-1: Potential Candidate genes from RNAi screen ......................................... 64
Table 3-1: Average lengths from PMT to diplotene and p-values for pairwise
comparisons between single mutant control and double mutants .......................... 104
Table 3-2: The average number of oocytes in diakinesis for csn mutants and
csn mutant apoptosis checkpoint double mutants .................................................. 106
Table 3-3: The average number of eggs laid for csn mutants and apoptosis
checkpoint double mutants ..................................................................................... 107
Table3-4: Number of nuclei counted for FISH analyses ........................................ 110
Table 3-5: p-values calculated by Fisher’s Exact Test for all pairwise
comparisons of FISH data ...................................................................................... 111
Table 3-6: p-values and total number of nuclei counted per zone for RAD-51
analyses .................................................................................................................. 113
Table 3-7: p-values and total number of nuclei counted in late pachytene zone
for apoptotic analyses ............................................................................................. 118
Table 3-8: Predicted post-translational modifications sites for SYP proteins........ 120
Table 4-1: Strains used in this study ...................................................................... 129
Table 4-2: Primer sequences and PCR conditions for validation of deletion
strains...................................................................................................................... 131
Table A-1: Chromosome II RNAi clones that were quantified.............................. 140
Table A-2: Chromosome IV RNAi clones that were quantified ............................ 147
Table A-3: Clones selected for cytological analysis from Chromosome II
RNAi screen ........................................................................................................... 153
Table A-4: Chromosome II cytological defect analyses-wild-type........................ 155
Table A-5: Chromosome II cytological defect analyses-akir-1(rj1). ..................... 157
Table A-6: Clones elected for cytological analyses from Chromosome IV
RNAi screen ........................................................................................................... 159
Table A-7: GO Terms for all 2007 Chromosome IV clones selected for
cytological analyses ................................................................................................ 160
Table B-1: Strains used in the Drosophila Akirin project...................................... 177
ix Table B-2: Primers used for RT-PCR analyses ...................................................... 178
Table B-3: Conditions for RT-PCR and amplicon length ...................................... 178
Table B-4: Number of trans-heterozygote F1 adult flies expected and the pvalues Fishers Exact test for the comparisons to the trans-heterozygous
controls ................................................................................................................... 180
Table B-5: Results of the fecundity study for selected trans-heterozygote
models and p-values Fishers Exact test .................................................................. 182
Table B-6: Akirin quantification, n values and p-values Fishers Exact test .......... 186
Table B-7: Quantification of C(3)G, n values and p-values for Fishers Exact
test .......................................................................................................................... 188
Table B-8: Quantification of C(3)G staining for akirin EY08097-e82 models. ............ 190
Table B-9: Comparison between wild-type and Akirin expressing
backgrounds for C(3)G staining ............................................................................ 191
Table B-10: Quantification of RNAi experiment .................................................. 193
x LIST OF FIGURES
Figure 1-1: Events in meiosis ................................................................................... 28
Figure 1-2: Organization of the C. elegans gonad ................................................... 29
Figure 1-3: A representation of the SUN-KASH proteins in C. elegans ................. 30
Figure 1-4: A representation of the general structure of the synaptonemal
complex .................................................................................................................... 31
Figure 1-5: A simplified diagram of meiotic recombination c ................................. 32
Figure 1-6: C. elegans specific progression through the events of meiotic
prophase.................................................................................................................... 33
Figure 1-7: Synaptonemal complex disassembly and bivalent defects in akir-1
mutants ..................................................................................................................... 34
Figure 2-1: A schematic representation of the RNAi enhancer/suppressor
screen ........................................................................................................................ 57
Figure 2-2: Bivalent defects in unc-85(RNAi) and quantification of defects ........... 58
Figure 2-3: Bivalent defects in unc-85(RNAi) and unc-85(ok2125) ........................ 59
Figure 2-4: Bivalent defects in ZC239.6 mutants .................................................... 60
Figure 2-5: The lack of SYP-1 and chromosome defects in let-29(RNAi) ............... 61
Figure 2-6: Y66H1B.2(RNAi) and Y66H1B.2(ok2611) mutants. ............................. 62
Figure 2-7: SC and bivalent defects in C09G4.3(RNAi) mutants............................. 63
Figure 3-1: CSN/COP9 signalosome structure and function ................................... 94
Figure 3-2: SC central element assembly defects in csn mutants.. .......................... 95
Figure 3-3: csn mutants have smaller gonads and lack of oocytes progressing
through diakinesis..................................................................................................... 96
Figure 3-4: HTP-3 not aggregate in csn mutants. .................................................... 97
Figure 3-5: HIM-3 does not aggregate in csn mutants. ........................................... 98
Figure 3-6: SYP-4 aggregates in csn mutants .......................................................... 99
Figure 3-7: P-granule components kgb-1 and glh-1(gk100) do not have SYP-1
aggregation phenotype. .......................................................................................... 100
Figure 3-8: Quantification of the SYP-1 aggregates .............................................. 101
xi Figure 3-9: Representative images of nuclei and aggregation phenotypes. ........... 102
Figure 3-10: Quantification of SYP-1 protein in csn mutants................................ 103
Figure 3-11. Quantification of the lack of oocytes. ................................................ 105
Figure 3-12. Pairing stabilization is affected in csn mutants. ................................ 108
Figure 3-13: High magnification micrographs of individual nuclei....................... 109
Figure 3-14: Accumulation of recombination intermediates in csn mutants. ........ 112
Figure 3-15: Reduced crossover formation in csn mutants. ................................... 114
Figure 3-16: High magnification micrographs from RAD-51 and COSA-1
analyses. ................................................................................................................. 115
Figure 3-17: Apoptosis and MPK-1 expression are altered in csn mutants. .......... 116
Figure 3-18: Analysis of SYP-1 aggregate phenotype in csn mutants and
apoptosis checkpoint double mutants ..................................................................... 117
Figure 3-19: Quantification of dpMPK-1 expression in csn mutants .................... 119
Figure 3-20: Model for CSN/COP9 signalosome function in meiotic prophase
I ............................................................................................................................... 121
Figure B-1: Identifying potential trans-heterozygote mutant combinations to
model akirin deficiency in Drosophila. ................................................................. 176
Figure B-2: Akirin is required for fertility. ............................................................ 179
Figure B-3: The number of hatched eggs is reduced in akirin transheterozygote mutants .............................................................................................. 181
Figure B-4:.A schematic of Drosophila oogenesis in prophase I .......................... 183
Figure B-5: Immunostaining of Drosophila follicles with Akirin specific
antibodies................................................................................................................ 184
Figure B-6: Quantification of Akirin staining ........................................................ 185
Figure B-7: Quantification of C(3)G staining.. ...................................................... 187
Figure B-8: Quantification of C(3)G staining for akirin EY08097-e82 mutants.. ......... 189
Figure B-9: Comparison of pooled wild-type samples, akirinDG/KG, and
akirinKG/EX82.. .......................................................................................................... 192
xii LIST OF ABBREVIATIONS
BPA
CO
CR
CRIPSR
DAPI
DNA
DSB
DTC
EMB
EMO
FET
GFP
HIM
LE
MAPK
MI
MII
MW
NCO
NHEJ
PC
PMT
RNA
RNAi
SC
SMC
TZ
Bisphenol A
Crossover
Central region
Clustered Regularly interspaced short plaindromic
repeats
4',6-diamidino-2-phenylindole
Deoxyribose nucleic acid
Double strand break
Distal tip cell
Embryonic lethal
Endomitotic
Fishers Exact Test
Green Fluorescent Protein
High Incidence of Males
Lateral element
Mitogen activated protein kinase
Meiotic Division I
Meiotic Division II
Mann Whitney test
Non-crossover
Non-homologous end joining
Pairing center
Pre-meiotic tip
Ribose nucleic acid
RNA interference
Synaptonemal complex
Structural Maintenance of Chromosomes
Transition zone
xiii 1 CHAPTER 1
INTRODUCTION
Meiosis matters
The formation of haploid gametes is critical for reproduction in most eukaryotic
organisms. Meiosis is the specialized cellular division leading to the formation of
gametes, which in metazoans are eggs and sperm. Unlike mitosis, meiosis has one round
of chromosome replication followed by two divisions: the first division is referred to as
MI, in which homologous chromosomes segregate from each other, and the second
division is referred to as MII, where sister chromatids segregate (Figure 1-1). It is
essential that chromosome segregation during the divisions occurs correctly or an
aberrant number of chromosomes will be present in the gametes, resulting in aneuploid
eggs or sperm and consequently aneuploid offspring (Hassold and Hunt 2001; Sears,
Hegemann, and Hieter 1992; Koehler et al., 1996; Hassold, Abruzzo, and Adkins 1996;
Sheppard et al., 2012; Sheltzer et al., 2011). While plants can tolerate changes in ploidy
which can lead to new genes via duplication events and even new species, animals are
much less tolerant of changes in ploidy which typically leads to lethality (Ramsey and
Schemske 1998; D. E. Soltis and Soltis 1999; Powers et al., 1991). While getting the
correct number of chromosomes into the gametes is critical, meiosis also important for
genetic recombination that provides new allele combinations in species. These new allele
combinations can provide raw material for evolutionary section to act upon(Ramsey and
Schemske 1998; D. E. Soltis and Soltis 1999; Powers et al.,, 1991; Van de Peer, Maere,
and Meyer 2009; Magadum et al., 2013). Therefore, meiosis is not only about
gametogenesis, but it also works as driver of evolution, providing the genetic variation
through recombination and subsequent random segregation of chromosomes into
gametes.
In meiotic prophase I, preceding the first division three key events take place,
homologous chromosomes pair (associate), synapse (stably associate throughout their
2 length via the synaptonemal complex), and form crossovers (recombine the genetic
material) although the order of the later two events can vary between organisms (Figure
1-1). Crossovers and sister chromatid cohesion result in chiasma, the visually detectable
connections between homologous chromosomes observed in late prophase I. Chiasma
allow homologs to align properly at the metaphase plate during meiosis I and
subsequently segregate to opposite poles (Hirose et al., 2011). All prophase I steps are
highly regulated, ensuring that meiotic prophase proceeds correctly. Disruption of
meiosis prophase I events can result in apoptosis or cell cycle arrest as a result of
activating checkpoints or more commonly, result in aneuploid gametes.
Missegregation of homologous chromosomes during meiosis I is the most
common reason for aneuploidy in humans and other organisms. Aneuploidy in gametes
usually leads to developmentally deficient or non-viable progeny. The incidences of
aneuploidy in gametes are 1-2% in human males and up to 20% in human females
(Hassold and Hunt 2001; Hunt and Hassold 2008; Nagaoka, Hassold, and Hunt 2012;
Hassold, Hall, and Hunt 2007). Furthermore, up to 35% of human spontaneous abortions
are the results of aneuploidy (Hassold and Hunt 2001). The 0.3% of human live
aneuploid births result in Down Syndrome (trisomy 21), Edward Syndrome (trisomy 18),
Patau syndrome (trisomy 13), Turner syndrome (X0), and Klinefelter syndrome (XXY).
In case study of 1000 human trisomic fetuses/live born individuals conducted in the mid1990s, it became clear that the majority of cases originated from maternal MI errors
(Hassold and Hunt 2001; Hassold and Hunt 2009). There is a correlation between the
human maternal age and the incidence of aneuploidy, with women under the age of 25
years only having approximately 2% of trisomic pregnancies while women over the age
of 40 years have approximately 30-35% of trisomic pregnancies (Hassold and Hunt 2001;
Hunt and Hassold 2008). When coupled with the current trend of women delaying
childbirth until later in life, this increase in aneuploidy with maternal age becomes a
significant reproductive complication (Hunt and Hassold 2008). Furthermore, recent
3 work examining the detrimental affects of xenoestrogens, such as Bisphenol A (BPA), on
meiotic prophase I events demonstrates that age alone is not the only contributing factor
to aneuploidy, but also environmental factors play a role (Hunt et al.,, 2003; Hunt et al.,,
2009; Allard and Colaiácovo 2010).
In humans, we can readily observe the outcomes of meiotic prophase I errors.
However, determining the source of those errors is not easy, especially since female
meiotic prophase I occurs in the embryo between weeks 11-12, where meiotic
chromosomes undergo pairing, synapsis and then arrest in late prophase I phase until
puberty (Hassold and Hunt 2001; Hunt and Hassold 2008). Therefore, a woman is born
with a specific number of arrested oocytes that have already passed through meiotic
prophase I. In males, meiotic entry does not occur until puberty, therefore, many of the
human studies in meiosis are centered on spermatogenesis as it is easier to study. While
many of the spermatogenesis studies have yielded insights into human meiotic prophase I
[the link between synapsis and recombination defects leading to male infertility
(Schramm et al., 2011; M. A. Handel and Schimenti 2010)], we lack fundamental
knowledge of the origins of human female meiotic prophase I errors that lead to
aneuploidy.
To gain insights into the general mechanisms of meiotic prophase I events, model
organisms such as S. cerevisiae, C. elegans, and D. melanogaster have been examined.
Though studies of meiosis in these model organisms, insights have been gained into the
mechanisms of prophase I events such as the specifics of homologous chromosome
pairing, the details of synapsis and the morphology of the synaptonemal complex, the
mechanisms of genetic exchange via homologous recombination and its role in the
segregation of chromosomes during the meiotic divisions. Just as important, it has been
determined that while all these events do indeed take place in meiosis, the timing and
order vary between organisms for example; in yeast and mammals, recombination and
synapsis are mutually dependent, while in flies and worms, synapsis is independent of
4 recombination (Bhalla and Dernburg 2008). Despite the differences in the various
organisms, overall the processes are highly conserved: chromosomes must pair, synapse,
recombine, and form crossovers to ensure proper segregation (Page and Hawley 2004).
Disruption in any of these events will lead to aneuploidy in any sexually reproducing
organism; therefore, studying meiosis in multiple organisms will provide not only insight
into the general mechanisms, but also species specific mechanisms as well.
C. elegans: a model for meiotic study
Since its introduction as a model organism in 1974 by Sydney Brenner (Brenner
1974), C. elegans has become a powerful model system for the study of basic biological
processes as well as a model for human disease. With 60-80% of its genes showing
conservation with humans, this 1.5 mm soil nematode has been utilized to study
neurobiology, development, and is a model for various diseases such as diabetes, cancer
and is being utilized in toxicological studies (Kaletta and Hengartner 2006; Boulin and
Hobert 2011). The advantages of using C. elegans are numerous. The transparency of its
cuticle allows visualization of internal organs and processes without dissection. A
detailed analysis of somatic cell linages revealed that an adult only contains 959 somatic
cells. In addition, worms reproduce rapidly and prolifically; development from egg to
adult is only three days. C. elegans are self-fertilizing hermaphrodites, which contain
reproductive organs of both sexes, generating eggs and sperm, They can produce 300-350
progeny through self-fertilization which is useful for the study of recessive genes where
one quarter of the progeny will be recessive (Kaletta and Hengartner 2006; Boulin and
Hobert 2011). Furthermore, the existence of males allow for generation of cross progeny
(Kaletta and Hengartner 2006). C. elegans is ideal for genetic screens (Boulin and Hobert
2011): many worms can be grown in low cost (due to its small size, no need of crosses,
short generation time and simple media and conditions for cultivation) and variety of
mutangenesis techniques are available. C. elegans is highly amenable to genome editing
technologies (Wood et al., 2011) (Waaijers et al., 2013); cytological analyses(Boulin and
5 Hobert 2011); transgenesis by integration (Frøkjær-Jensen, Davis, and Ailion 2012) and
expression of genes somatically from extra chromosomal arrays (Boulin and Hobert
2011).
The study of meiotic prophase I events in humans is challenging, especially the
study of oogenesis. However, C. elegans has advantages that make is a good model for
the study of oogenesis despite these differences between the two organisms because the
global processes are the same. In C. elegans assaying non-disjunction events is easy
because missegregation of the X chromosome leads to an increase in male progeny, a
high incidence of males (HIM) phenotype. In C. elegans, sex is determined by the
number of X chromosomes present in a worm: males are XO and hermaphrodites are XX.
In the wild-type background, males are produced at a very low frequency (<0.2%) (Lui
and Colaiácovo 2013). However, if segregation is disrupted, then the frequency of males
increases (a HIM phenotype). If autosomes are also affected an increase in the embryonic
lethality (EMB) is observed as a result of non-viable progeny due to aneuploidy.
The germline accounts for more than half the nuclei in the worm and contains
about 2000 nuclei (Colaiácovo 2006). The availability of this large number of nuclei for
analysis allows the study of meiotic prophase I defects in detail (Figure 1-2A and B).
Meiotic prophase I consists of five sequential stages: leptotene, zygotene, pachytene,
diplotene and diakinesis (Page and Hawley 2004). These stages can be differentiated by
chromosome morphology. The C. elegans germline is temporally and spatially organized
around the events of meiotic prophase I. There are two gonadal arms, which are U-shaped
and tubular and consist of the germline and gonadal somatic tissues with a
distal/proximal polarity (Figure 1-2A and B). At the distal end of the germline,
mitotically dividing stem cells provide a source of germ line cells (about 20 rows of cells,
see Figure 1-2B), which constitute the pre-meiotic tip (PMT). Throughout this distal part
of the germline, the germ cells are syncytial and connected to each other through a central
canal, the rachis, and share a common cytoplasm. (Figure 1-2C) (McCarter et al., 1997;
6 D. H. Hall et al., 1999). The transition zone (TZ) marks the entry of nuclei into meiotic
prophase I, beginning with leptotene and zygotene (these to stages cannot be
distinguished in C. elegans). The TZ is about 8-12 rows of cells long and is defined by
the clustered or polarized configuration of the chromosomes to one side of the nucleus
(Figure 1-2A and B). It is at this stage that replication, cohesion establishment and
pairing occur, and synapsis is initiated. As the nuclei progress through the cell cycle and
the germ line, they enter an extended pachytene stage, which accounts for well over half
the cells in the germline (Figure 1-B). Synapsis is completed [the synaptonemal complex
(SC) is fully formed] during pachytene and meiotic recombination is completed
(McCarter et al., 1997; D. H. Hall et al., 1999). As the nuclei progress towards the
proximal end of the gonad, the rachis narrows and the nuclei enter the “loop” region of
the gonad where MAPK signaling initiates the exit from pachytene (Church, Guan, and
Lambie 1995; McCarter et al., 1997). Here, apoptosis reduces the number of nuclei and
the surviving nuclei start to transition to diplotene, chromosomes start to condense and
form bivalents, and desynapsis ensues. In C. elegans wild-type nuclei, what is referred to
as physiological apoptosis occurs, reducing the over all number of germ cells that
transition into diplotene and involves the activation of core apoptotic machinery
(Gumienny et al., 1999; Gartner et al., 2000). The trigger for physiological apoptosis of
remains elusive and its not yet clear what targets specific nuclei to cell death via this
pathway. The current model put forth for the reduction in nuclei involves the notion that
the excess germ cells are “nurse cells” that provide cytoplasmic components (through the
syncytial connection with the rachis) to the surviving germ cells and allowing for the
excess cytoplasm to be utilized by the maturing oocyte (Gumienny et al., 1999). In
aberrant conditions, as found in mutants and/or after exposure to genotoxic agents,
apoptosis is not random. In this cases defects in synapsis or recombination can initiate
two distinct pathways leading to apoptosis. This is considered to serve as a means to clear
defective nuclei that fail to synapse properly or to properly repair DNA damage
7 (Gumienny et al., 1999; Gartner et al., 2000). In the last stage of prophase, diakinesis,
there is an increase in cytoplasmic contents such as mitochondria, ribosomes and other
cellular organelles (D. H. Hall et al., 1999). As the oocytes near the distal end of the
gonad, they gain yolk and the most proximal oocyte is the most mature, ready for
fertilization. Cellularization occurs in these last three oocytes at the proximal end of the
gonad (D. H. Hall et al., 1999). These oocytes are commonly referred to as the late
diakinesis oocytes. The most proximal oocyte just about to be fertilized is called
diakinesis -1 (D-1); the next oocyte distal to D-1 is diakinesis -2 (D-2) and the third
oocyte is called diakinesis -3 (D-3) (Figure 1-2).
The somatic tissues of the germline also exist in the distal to proximal orientation.
At the distal end of each arm in the gonad, resides the single distal tip cell (DTC). The
somatic DTC regulates the differentiation and proliferation of the nucleus population in
the PMT (Pepper et al., 2003; Albert Hubbard 2007). This regulation occurs through the
GLP-1/Notch signaling pathway: GLP-1/Notch receptors in the germline are activated by
the LAG-2 ligand, which is expressed by the DTC (Pepper et al., 2003; Albert Hubbard
2007). Laser ablation of the DTC and loss-of-function mutations in the signaling pathway
lead to early entry into meiosis by the nuclei (Pepper et al., 2003; Crittenden and Kimble
2008; Kimble and Crittenden 2005). A more detailed explanation of proliferation is
provided in Chapter 3.
The majority of the germline is covered with a layer of cells termed sheath
cells(five cells total) (Figure 1-2A); however, the PMT is considered a “bare” region and
lacks a sheath covering. These sheath cells are closely associated with the germline and
are essential to germline proliferation, development and oocyte maturation (McCarter et
al., 1997; D. H. Hall et al., 1999). Laser ablated sheath cells result in a disorganized
gonad morphology (Crittenden et al., 1994) and a lack of a central rachis (McCarter et
al., 1997). In the proximal gonad, sheath cells are responsible for oocyte maturation
through MAPK signaling (discussed in Chapter 3) as well as engulfment of nuclei that
8 under go apoptosis, which removes them from the germline (Gartner et al., 2000). As
oocytes mature, it is hypothesized that the sheath cells assist with signal transduction
from the spermatheca to the rest of the gonad to facilitate oocyte progression and
maturation (Church, Guan, and Lambie 1995; McCarter et al., 1997). Not only is the C.
elegans gonad a model for meiotic studies, but it can allow us to understand the
interactions between the germline and the somatic gonad as well.
Early meiotic prophase I events in C. elegans
Replication and the establishment of cohesion
Establishing the cohesion landscape along the homologous chromosomes is
absolutely critical for the execution of prophase I events. The nuclei leave the PMT and
enter meiosis in the TZ, with fully replicated chromosomes (each composed now of two
sister chromatids). At this point, cohesion must be already established to hold the
chromatids together (Peters, Tedeschi, and Schmitz 2008; Wood, Severson, and Meyer
2010). Sister chromatid cohesion is essential to the proper segregation of chromosomes
during the meiotic divisions. Without cohesion, sister chromatids would prematurely
separate at the MI division (Lui and Colaiácovo 2013).
Cohesion occurs in both meiotic and mitotic divisions and is immediately
established after replication. The highly conserved cohesion complexes in most
organisms are comprised of SMC (structural maintenance of chromosomes) subunits and
an α-kleisin subunit (Peters, Tedeschi, and Schmitz 2008; Wood, Severson, and Meyer
2010). The SMC subunits form a structure to encircle the chromatids, binding them
together. The α-kleisin acts as a latch to the SMC complex (Wood, Severson, and Meyer
2010). While the SMC subunits are the same for both mitotic and meiotic cohesion
complexes, the α-kleisins are specific to the type of division: Scc1 for mitosis and Rec8
for meiosis (Wood, Severson, and Meyer 2010).
For the most part, the meiotic cohesion complex in C. elegans is no different than
those in other organisms, consisting of SMC subunits: SMC-1/HIM-1 and SMC-3; SCC 9 3, but interestingly it has three meiotic α-kleisins REC-8, COH-3 and COH-4 (Severson
et al., 2009; Csankovszki et al., 2009; J. S. Bickel et al., 2010). The kleisins, REC-8,
COH-3 and COH-4 appear to have redundant function and only when all three are
depleted, is sister chromatid cohesion disrupted (Severson et al., 2009). Depletion of the
SMC subunits of SCC-3 leads to a lack of pairing, impaired synapsis and an increase in
recombination intermediates (Severson et al., 2009; Lightfoot et al., 2011; Smolikov,
Eizinger, Hurlburt, et al., 2007; Pasierbek et al., 2003).
Once sister chromatid cohesion is established, it must be maintained throughout
the remainder of prophase I on the entire chromosomes, and from MI to MII at the
centromeres of chromosomes (Chan et al., 2003; Hagstrom 2002). In most organisms the
maintenance of centromeric cohesion is dependent on shugoshin (SGO-1) and protein
phosphatase 2A (PP2A), which work together to prevent the cleavage of centromeric
cohesion by separase (Peters, Tedeschi, and Schmitz 2008; Hagstrom 2002). However, in
C. elegans, there is a SGO-1 independent mechanism that relies on LAB-1 (a wormspecific protein that protects cohesion along the chromosome axis). It is possible that this
specific mechanism evolved due to the holocentric chromosomes found in C. elegans
meiosis. In C. elegans, the cohesion is localized across the arms of sister chromatids in
the bivalent, not at the centromere as in other organisms (Chan et al., 2003; Hagstrom
2002). While LAB-1 first localizes to the chromosomes in the TZ, it is extensively
localized to chromosome axis only at mid-pachytene (de Carvalho et al., 2008). LAB-1
protects REC-8 cohesion on the long arm of the bivalents late in prophase I (de Carvalho
et al., 2008). This allows the cohesion between the sister chromatids to remain intact
during MI division. Only when LAB-1 dissociates in MII can the cohesion between the
sister chromatids be cleaved allowing for their separation (de Carvalho et al., 2008).
Pairing
Once cohesion is established, the next step for the chromosomes is to find their
homologous partner in a process known as pairing. Cytologically, the onset of prophase
10 events is marked by the clustering of chromosomes to one side of the nucleus forming the
“meiotic bouquet” during the leptotene/zygotene stage in most organisms (Zickler 1999).
This transient structure results from the reorganization of chromosomes in the three
dimensional space of the nucleus to facilitate homologous pairing via the association with
telomeres with trans-membrane proteins that facilitate movement via cytoplasmic
interactions (Table 1-1) (Fridkin et al., 2008; Minn et al., 2009; Fridolfsson and Starr
2010; K. Zhou et al., 2009; Razafsky and Hodzic 2009). Homology searching occurs
between the homologous chromosomes as the telomeres are “shuffled” around in the
bouquet structure (Fridkin et al., 2008; Minn et al., 2009; Fridolfsson and Starr 2010; K.
Zhou et al., 2009; Razafsky and Hodzic 2009). The bouquet structure highlights the
dynamic nature of the morphological changes seen chromosomes throughout prophase
(L. Harper, Golubovskaya, and Cande 2004). While in most organisms, the telomeres are
attached to the nuclear membrane and brought into close proximity to facilitate
homologous pairing and chromosome alignment (Table 1-1). Unlike most organisms, C.
elegans uses pairing centers, located just proximal to the telomeres to facilitate pairing.
Since chromosomes are attached only on one side to the nuclear envelope and do not
cluster together, this is not defined as a bouquet, but is a bouquet-like structure (N. C.
Harper et al., 2011) (Phillips and Dernburg 2006). The dynamic movement of the
chromosomes is mediated by the cytoskeletal elements: actin, tubulin, and dynein (Table
1-1) (Sato et al., 2009; Baudrimont et al., 2010; Morimoto et al., 2012).
The bridge between the chromosomes, the nuclear membrane and the
cytoskeleton is established by SUN/KASH proteins that are conserved throughout
metazoans and have recently been identified in plants (Table 1-1) (Fridkin et al., 2008;
Minn et al., 2009; Fridolfsson and Starr 2010; K. Zhou et al., 2009; Razafsky and Hodzic
2009). SUN (Sad1/UNC-84 homology domain) proteins span the inner nuclear
membrane, interacting on the inner side of the membrane with either the nuclear lamina
or the telomeres depending on the organism (Figure 1-3) (Fridolfsson and Starr 2010; Z.
11 Zhou et al., 2012). KASH (Klarsicht/ANC-1/Syne-1 homology domain) proteins span
the outer nuclear membrane and bind to the SUN proteins in the perinuclear space of the
membrane (Figure 1-3). The KASH proteins interact with the cytoskeleton elements such
as actin or microtubules to facilitate movement of chromosomes inside the
nucleus(Fridkin et al., 2008; Minn et al., 2009; Fridolfsson and Starr 2010; K. Zhou et
al., 2009; Razafsky and Hodzic 2009).
In C. elegans, pairing centers (PCs) are cis acting regions of repetitive DNA
motifs that recruit the zinc finger proteins HIM-8, ZIM-1, ZIM-2, and ZIM-3. Together,
the PCs and these proteins are essential for accurate chromosome segregation (MacQueen
et al., 2005; Phillips et al., 2005). HIM-8 associates with the X chromosome while the
ZIMs associate with the autosomes (ZIM-1 with chromosomes II and III; ZIM-2 with V;
ZIM-3 with I and IV) (Phillips and Dernburg 2006). The fact that some ZIMs can
associate with more than one autosome suggests there must be additional mechanisms
that recognize homologous chromosomes (Phillips and Dernburg 2006; Zetka 2009; Rog
and Dernburg 2013). Once these associations are established on each of chromosomes in
a homologous pair, the actual movement of chromosomes can take place through
interactions with SUN/KASH nuclear trans-membrane proteins that link the
chromosomes to the cell cytoskeleton.
In C. elegans, SUN-1 and ZYG-12 are the SUN/KASH pair that interact with the
PCs although the exact nature of the interaction remains elusive (Minn et al., 2009; Zetka
2009; Phillips et al., 2009; Penkner et al., 2009). While ZYG-12 interacts with dynein
and microtubules anchoring the centrosome close to the nucleus (Malone et al., 2003).
SUN-1 interacts with the PCs inside the nucleus (Figure 1-3) (Sato et al., 2009; Penkner,
Tang, et al., 2007; Baudrimont et al., 2010). These two proteins interact and cytological
studies show that antibodies for both co-localize to the nuclear periphery. Baudrimont et
al., 2010 showed these SUN-1/ZYG-12 aggregates, representing a single chromosome of
a homologous pair, have a dynamic pattern of movement and coalesce into larger
12 patches, which correspond to the polarized chromosomes seen by DAPI staining, giving
the nucleus the crescent shape seen in the TZ. Not only are the aggregates polarized to
one side of the nucleus, but they are being “shuffled” around the patches in a highly
dynamic manner. These observations are in agreement with the proposed mechanism of
chromosome shuffling by SUN-KASH in the bouquet that facilitates homology searching
(Scherthan 2001; L. Harper, Golubovskaya, and Cande 2004; Hübner and Spector 2010).
Baudrimont et al., 2010 along with and Penkner et al., 2009 showed that posttranslational phosphorylation of SUN-1 was essential to this dynamic movement and that
PLK-2 (polo-like kinase 2) was responsible for the phosphorylation of SUN-1.
MacQueen and Villeneuve, 2002 had already shown that CHK-2 (checkpoint homolog 2)
was involved in pairing and polarization, not in DNA damage repair as in other
organisms. CHK-2’s exact molecular role remains unclear as its targets have not been
identified. SUN-1 phosphorylation allows it to interact with the PCs of the chromosomes
and for the polarization of the chromosomes to one side of the nuclear periphery (Penkner
et al., 2009; N. C. Harper et al., 2011; Labella et al., 2011). Depletion of CHK-2 or PLK2 leads to a lack of polarization and pairing. Null mutations of the ZIMs, HIM-8, or the
SUN-1/ZYG-1 complex result in non-homologous synapsis and/or a lack of pairing and
synapsis.
The current consensus in the field is that homologous pairs are linked to pSUN1/ZGY-12 complexes that are highly mobile due to their interaction with dynein and the
cytoskeleton. These can be seen as aggregates that form patches on one side of the
nucleus. It is within these patches that homology searching occurs with cytoplasmic
forces bringing together and forcing apart the homologous chromosomes. Only when the
correct homologs are in close proximity, can synapsis begin and overcome the
cytoplasmic forces, thus re-enforcing and stabilizing pairing. How exactly synapsis is
initiated remains unclear, but synapsis specific proteins are necessary to form functional
13 SUN-1 aggregates and allow for proper alignment so that synapsis can occur (Sato et al.,
2009; Penkner, Tang, et al., 2007; Baudrimont et al., 2010).
Synapsis
In most organisms, the synaptonemal complex (SC) is an evolutionarily
conserved tripartite protein structure connecting pairs of homologous chromosomes
during most prophase I stages and is required for the formation of most crossovers
(Figure 1-4A) (Page and Hawley 2004). In C. elegans, an absent or improperly formed
SC prevents crossover formation, resulting in missegregation of chromosomes (Figure 14B) (Colaiácovo et al., 2003). However, in other organisms, impaired formation of the
SC leads to a reduction in crossover formation (Page and Hawley 2004). The SC consists
of two primary structures, the lateral elements (LEs) and the central region (CR) (Figure
1-4A). While the structure and proteins are functionally conserved, there is little protein
sequence homology between most of the proteins that comprise the SC, making it
difficult to identify SC proteins across species (Page and Hawley 2004).
The lateral elements (Table 1-1) are found along the chromosome axis of each
homolog and associate with the chromatin of the sister chromatids (Page and Hawley
2004). The lateral elements all contain HORMA domain (Hop1, Rev7, Mad2) protein(s)
and assemble on the chromosomes prior to the start of pairing and just after the
establishment of cohesion (Tanneti et al., 2011; Zetka 2009; Page and Hawley 2004; Shin
et al., 2010; M. A. Handel and Schimenti 2010). HORMA domain proteins are the only
SC proteins to show evolutionarily conservation in protein sequence. Research shown
there is an intimate relationship between cohesion and the some of the lateral element
proteins in various organisms with the establishment of cohesion being necessary for the
localization of the lateral element proteins in many cases (Page and Hawley 2004). This
suggests an early establishment of a specific meiotic chromosome landscape, during
prophase just after cohesion establishment, on which additional protein structures will be
built (the rest of the SC) or that will facilitate other prophase events such as pairing, or
14 recombination (Tanneti et al., 2011; Zetka 2009; Page and Hawley 2004; Shin et al.,
2010; M. A. Handel and Schimenti 2010).
The central region proteins (Table 1-1) form the transverse connection between
the homologous chromosomes and thus connects the homologs together in close
proximity throughout prophase. One common feature across all CR proteins is the
presence of a coiled-coil domain, typically located near the center of the protein (Page
and Hawley 2004). Electron micrographs of mature SC structures suggest it is about 100
nm in diameter (Page and Hawley 2004). Drosophila and mammals have additional CR
proteins lacking coiled- coil domains and are essential to transverse filament
polymerization (their identity in each organism can be found in Table 1-1) (Page et al.,
2008; O. R. Davies, Maman, and Pellegrini 2012).
In C. elegans, SC width is about 90-125 mm (Figure 1-3B) (Page and Hawley
2004). The C. elegans lateral element proteins include HTP-1/2, HTP-3, and HIM-3
(Couteau 2005; Martinez-Perez 2005; Goodyer et al., 2008; Severson et al., 2009; Zetka,
Kawasaki, and Strome 1999) and there are four known CR proteins: SYP-1, SYP-2,
SYP-3, and SYP-4 (collectively known as SYPs) (Smolikov, Eizinger, Schild-Prufert, et
al., 2007; Smolikov, Schild-Prüfert, and Colaiácovo 2009; Colaiácovo et al., 2003;
Schild-Prüfert et al., 2011). HTP-3 is the first to assemble and assists in the loading of
HTP-1/2 and HIM-3 onto the chromosomes (Couteau 2005; Martinez-Perez 2005;
Goodyer et al., 2008; Severson et al., 2009; Zetka, Kawasaki, and Strome 1999).
Interestingly, HTP-3 and cohesion seem to be interdependent, essentially providing a
core chromatin landscape on which to build the SC and to promote meiotic
recombination and facilitate crossover formation (Goodyer et al., 2008). Once this
landscape is established, then HTP-1/2 and HIM-3 can be loaded onto the chromosomes
(Figure 1-4B). While HTP-1/2 does not require the presence of cohesion, HIM-3 does
require cohesion to load in mass (Couteau 2005; Martinez-Perez 2005; Zetka, Kawasaki,
and Strome 1999). These data suggest that replication, the establishment of sister
15 chromatid cohesion, pairing and early synapsis are not independent steps rather they are
integrated and dependent on one another to build a meiotic specific chromosome
foundation. It is on this foundation that the central region proteins, as well as those
proteins involved in recombination, will build a mature SC complex and initiate DSBs.
The four SYP proteins (SYP-1/2/3/4) all have the same basic structure, a coiledcoil domain with globular domains on either end. These proteins act in an interdependent
manner through head to head contact, building a transverse central region between
homologous chromosomes (Figure 1-4B) (Smolikov, Eizinger, Schild-Prufert, et al.,
2007; Smolikov, Schild-Prüfert, and Colaiácovo 2009; Colaiácovo et al., 2003; SchildPrüfert et al., 2011). If one SYP is missing, the CR does not form. The phenotypic
consequences of mutations in all four SYPs are identical: lack of synapsis and failure to
form crossovers (Smolikov, Eizinger, Hurlburt, et al., 2007; Smolikov, Schild-Prüfert,
and Colaiácovo 2009; Schild-Prüfert et al., 2011).
SYP polymerization initiates next to the PCs (near the telomeric region) of the
paired homologous chromosomes, but the exact molecular mechanism is unclear. CR
proteins appear to have a natural propensity to aggregate and form small polycomplexes
during meiosis just as the SC assembles and disassembles (Goldstein 1987). However,
these polycomplexes are also found in tissue culture cells where CRs are expressed
ectopically (Yang et al., 2006) and are frequently found in yeast meiotic mutants
(Voelkel-Meiman et al., 2012). In C. elegans, large polycomplex-like structures are
rarely observed upon SC assembly (Smolikov, Schild-Prüfert, and Colaiácovo 2008;
Bilgir et al., 2013; Sato et al., 2009)or disassembly (Clemons et al., 2013). However, CR
components are found to assemble between non-homologous chromosomes [for example
in htp-1/2 mutants (Couteau 2005; Martinez-Perez 2005; Couteau et al., 2004)] or sisters
[e.g., syp-3 truncation mutants(Smolikov, Eizinger, Hurlburt, et al., 2007; Smolikov,
Schild-Prüfert, and Colaiácovo 2008)]. By electron microscopy, polycomplexes are
reminiscent of SC structures and in most cases, they are not associated with DNA (Yuan,
16 Pelttari, and Brundell 1998; Goldstein 1987). Although polycomplexes can contain
multiple SC proteins, single components of the CR can form polycomplexes without the
aid of lateral element proteins(Yuan, Brundell, and Höög 1996). The natural propensity
of CR proteins to aggregate suggests there are molecular mechanisms to ensure that CR
polymerization only occurs in the context of the SC. Pathways regulating SC assembly to
prevent polycomplexes may be different between yeast meiosis and meiosis in other
organisms. When recombination or SC assembly is perturbed, the yeast CR protein Zip1
readily forms polycomplexs. On the contrary, none of the C. elegans CR proteins/SYPs
aggregate when some SC proteins are missing or recombination fails (Smolikov,
Eizinger, Hurlburt, et al., 2007; Smolikov, Schild-Prüfert, and Colaiácovo 2009; SchildPrüfert et al., 2011; Wojtasz et al., 2009). These findings raise the possibility that the
propensity for CR proteins to self-aggregate (to form polycomplexes) is more tightly
regulated in C. elegans meiosis.
The role of translational modifications in the morphogenesis of the SC is still
being determined. In yeast, it has been shown that SUMO (small ubiquitin like modifier)
has a role in building the central region of the SC (Voelkel-Meiman et al., 2013).
Furthermore, newly discovered yeast SC components, Ecm11 and Gmc2, demonstrate the
essential nature of SUMOylation to the formation of the SC. When Ecm11 was mutated,
not only did the SC not form, but polycomplex formation increased (Humphryes et al.,
2013). Additionally, Zip3 has been shown to be a SUMO E3 ligase and essential to SC
formation in yeast (Cheng 2006). These data suggest that translational modifications such
as SUMOylation may have a role in not only SC formation, but the prevention of
polycomplex formation. In addition to SUMOylation, phosphorylation has a role as well
in SC morphogenesis. In mammalian spermatogenesis studies, HORMAD1 and SMC3
were phosphorylated by ATM/ATR (serine/threonine kinases), which mediate
localization to unsynapsed regions between homologs (Fukuda et al., 2012), Furthermore,
it was shown that SYCP1, TEX12, and SYCE1 the mouse CR proteins, were
17 phosphorylated during SC disassembly by PLK1. While PLKs are present during
prophase, only PLK1 localized to the SC (Jordan, Karppinen, and Handel 2012).
Mechanisms that regulate SC assembly to prevent CR aggregation may therefore involve
post-translational modifications of CR proteins. In C. elegans, it is not clear if such
mechanisms exist and how CR proteins are post-translationally modified.
It is important to note that the initiation of synapsis in relationship to the initiation
of recombination varies among organisms. In yeast, mammals and plants synapsis is at
least partially dependent on recombination while in flies and worms synapsis formation is
completely independent of recombination (Bhalla and Dernburg 2008). This suggests that
two mechanisms exist for synapsis initiation: one that is dependent on recombination and
one that is independent of recombination (Bhalla and Dernburg 2008). Currently, it is not
clear what exactly initiates the polymerization of the CRs. However in C. elegans, once
the SYPs have started to polymerize at the pairing centers and homology is confirmed,
SC polymerization between the homologs proceeds in a highly processive manner. This
SC assembly is mediated by CRA-1, a tertatricopeptide (TPR) containing protein
(Smolikov, Schild-Prüfert, and Colaiácovo 2008). In C. elegans, CRA-1 is the only
protein identified so far that regulates the accurate assembly of the SYPs into a transverse
structure between the lateral elements and allows for the bypass the requirement of DSB
formation for SC assembly (Smolikov, Schild-Prüfert, and Colaiácovo 2008).
Given the essential nature of synapsis to the success or recombination and
crossover formation, it is not surprising that Bhalla and Dernberg, 2005 identified Pch2
(also known as TRIP13 in mammals), an AAA-ATPase, as a component in a synapsis
checkpoint. Recent work by Deshong et al., 2014 showed that PCH-2 was expressed
early in the germline prior to the entry of the nuclei into meiosis and once the nuclei were
in meiosis localized to the axis of the homologous chromosomes. In the absence of PCH2, early prophase events were accelerated and meiotic defects were observed. These
authors further show that recombination intermediates are inappropriately stabilized (they
18 did not provide a mechanism) and further hypothesized that PCH-2 is required for access
to the homologs by the DNA repair machinery (Deshong et al., 2014). Additionally,
when coupled with apoptotic mutants, the meiotic defects in repair became more severe,
suggesting there may be a issue with repair or detection of DNA damage (Deshong et al.,
2014). While Pch2 is essential in yeast and mammals for the recombination/pachytene
checkpoint, in C. elegans, it may dispensable for the recombination checkpoint,
functioning to regulate early prophase events. Therefore, it is now believed that PCH-2
introduces a kinetic barrier to slow the progression of prophase, allowing for completion
and control of fidelity of pairing, synapsis, and recombination (Deshong et al., 2014)
Meiotic recombination and crossover formation
Crossover is the exchange of genetic material, which promotes genetic diversity
within sexually reproducing organisms. The formation of crossovers (CO) is also
required for accurate segregation of chromosomes in MI, which further increases genetic
diversity. COs help provide tension to the chromosomes as they align on the MI spindle,
aiding in the proper bi-orientation relative to the spindle poles (Bhalla and Dernburg
2008; Kleckner 2006). In other organisms, multiple CO events can be cytologically
observed as chiasmata, the physical representation of a CO event (Bhalla and Dernburg
2008; Kleckner 2006). However, in C. elegans, there is one CO per chromosome pair due
to the tight regulation of CO formation: although there are multiple double strand breaks
(DSBs) formed over the length of the homologous pair and these DSBs are repaired as
non-crossovers (Zetka 2009; Bhalla et al., 2008; Martinez-Perez et al., 2008).
Given the importance of the formation of COs to segregation, several levels of
regulation are employed: crossover assurance, crossover interference, and crossover
homeostasis (Lui and Colaiácovo 2013). Crossover assurance maintains the one
obligatory crossover (minimum numbers of CO’s to provide enough tension between the
chromosomes on the spindle pole) (Bhalla and Dernburg 2008; Meneely et al., 2012;
Carlton, Farruggio, and Dernburg 2006; Nabeshima, Villeneuve, and Hillers 2004). In
19 most organisms, crossover interference is a mechanism by which COs are distributed
over a chromosome, so CO will not be close to each other (Lui and Colaiácovo 2013). In
C. elegans interference is complete, as only a single crossover is found per pair of
homologous chromosomes. In C. elegans, interference is complete to ensure that there is
one crossover per chromosome pair (Lui and Colaiácovo 2013). Crossover homeostasis
maintains the CO level regardless of the number of DSB that occur and is related to
crossover assurance. This suggests that minimum level of COs is necessary to provide
enough tension to correctly orient the chromosomes on the spindle (Mets and Meyer
2009; Bhalla and Dernburg 2008). While little is known regarding crossover homeostasis
in C. elegans, it is clear that crossover assurance and interference mechanisms are present
and functioning during meiosis (Libuda et al., 2013).
As previously mentioned, there is one CO per pair of homologous chromosomes
in C. elegans; the formation of many breaks suggests that crossover interference is
complete. Studies in C. elegans suggest that the SC may have a role in regulating of the
formation and spacing of CO by essentially defining the chromosome landscape for the
crossover machinery (Hillers and Villeneuve 2003; Nabeshima, Villeneuve, and Hillers
2004). Interestingly, COs in C. elegans tend to form in the terminal third of chromosomes
which gives an asymmetry to the chromosomes are they are remodeled into the bivalent
structure prior to the MI division (Lui and Colaiácovo 2013). More recent work indicates
that the lateral elements and early SC morphogenesis (coupled with early DSBs) may
influence which breaks become COs. This could occur through the density of chromatin
loops that is regulated by condensin I. This is hypothesized to give rise to an early
decision of which DSBs would become COs (Lui and Colaiácovo 2013). However, the
CO/NCO decision could be done later, after the assembly of cohesion, and regulated by
the density of CR proteins (Libuda et al., 2013). The ability of the SC to regulates
CO/NCO decision may stem from the tight connection between DSB formation and SC
formation: DSBs are introduced after SC formation because the axial element protein
20 HTP-3 allows for the recruitment of MRE-11 (a meiotic specific repair protein) which is
necessary for DSB formation. (Goodyer et al., 2008; Severson et al., 2009). Couteau and
Zekta, 2011, have shown that DSBs generated by exogenous DNA damaging agent are
repaired concurrently with SC axis separation in late prophase. This is also coupled with
a reduction in acetylation on histone 2A, allowed DSBs to be repaired as NCO.
Regardless of when the decision is made, it is clear that a chromosome landscape defined
cohesion, and a fully formed SC has a role in the decision process.
Thus far, I have described the importance of COs and their impact on segregation,
but not how COs are formed. Meiotic homologous recombination is the result of
preprogrammed DSBs initiated in leptotene/zygotene by a containing Spo11, a
topoisomerase II like protein as well as accessory proteins (Keeney and Giroux 1997).
DSBs are then resected by Mre11/Rad50/Xrs2 complex (MRE11/RAD50/NBS1 in
mammals) and by the COM-1 (completion of meiotic recombination) complex to expose
3’ ssDNA (Chin 2001; Penkner, Portik-Dobos, et al., 2007). RPA (replication protein A)
initially binds to the exposed ssDNA, but is replaced by the RecA homolog, Rad51,
which mediates the invasion of ssDNA into the homologous DNA (Chin 2001; Penkner,
Portik-Dobos, et al., 2007). The homologous DNA is then used as a template for
synthesis of DNA to fill in the missing DNA lost during resection (Bishop 2012;
Petalcorin et al., 2007; Takanami et al., 2003). At this point one of two paths can be
chosen, the DSB can be resolved as a CO or NCO (non crossover) (Figure 1-5). For those
DSBs that are selected to become COs, (Bhalla and Dernburg 2008). a double Holliday
junction is formed, then cleaved by a resolvase, leading to a CO (asymmetric cleavage) or
a NCO (symmetric cleavage). The remaining DSBs do not form a double Holliday
junction, and thus cannot form a CO, becoming NCOs (Figure 1-5) . This repair can lead
to gene conversion in cases in which new sequence are copies form the adjacent
chromosome containing allelic variation (Bishop 2012; Petalcorin et al., 2007; Takanami
et al., 2003). If all recombination pathways fail, NHEJ (non-homologous end joining)
21 will be initiated, leading to a lack of COs and segregation will be abnormal (Bishop 2012;
Petalcorin et al., 2007; Takanami et al., 2003).
In C. elegans, as in other organisms, DSB are formed at leptotene/zygotene where
SPO-11 is assisted by HIM-17 and HTP-3 and DSB repair continues until late pachytene.
HIM-17 is required for methylation of histone H3, which is suggested to alter the
chromatin state at DSB sites. HTP-3 is required for the recruitment of MRE-11, RAD-50
(radiation sensitive) and HIM-3 to prevent the exchange of DNA with the sister
chromatin (Couteau et al., 2004; Goodyer et al., 2008). DSB resection is mediated by
MRE-11/RAD-50 and COM-1. RAD-51 binds the ssDNA to stabilize it, and RAD-54
assists strand invasion (Bishop 2012). ZHP-3, MSH-4/HIM-14, MSH-5 (members of the
DNA mismatch repair family of proteins) as well as COSA-1 (crossover site associated
protein) are required for CO formation. These proteins are interdependent in promoting
Holliday junction formation and ensuring CO resolution (Bishop 2012; Yokoo et al.,
2012). Recently, the C. elegans’ resolvases have been discovered. HIM-6 is the scaffold
for nucleases XPF-1 (xeroderma pigmentosa group F), SLX-1 (synthetic lethal X) and
MUS-81 (mutagen sensitive) which function in redundant pathways to mediate cleavage
of Holliday junctions (Agostinho et al., 2013; Saito et al., 2013; O'Neil et al., 2013).
While GEN-1 can act as a resolvase, it is not dependent on the HIM-6 scaffold
(Agostinho et al., 2013; Saito et al., 2013; O'Neil et al., 2013). Additionally, RTEL-1
(regulator of telomere length) appears to regulate the number of COs by promoting an
alternative repair pathway, SDSA (synthesis dependent strand annealing) (Ward et al.,
2008; Garcia-Muse and Boulton 2007). In eukaryotes, while homologous recombination
is the common repair pathway of SPO-11 induced DSBs, if there are defects in the HR
pathways, NHEJ will be the repair mechanism of choice. NHEJ is not dependent on
synapsis or recombination and can take place in C. elegans if there is a lack of synapsis
or cohesion; however, this repair is error-prone leading to chromosomal fragmentation
22 and aggregation (Smolikov, Eizinger, Schild-Prufert, et al., 2007; Schild-Prüfert et al.,
2011; Bhalla et al., 2008; F. Wang et al., 2003; Dorsett 2007).
In many organisms, if recombination is not complete or DNA damage is present,
the pachytene checkpoint or DNA damage pathway will be activated, leading to repair
(in the germline), cell cycle arrest (only in the PMT) and in the most extreme cases,
apoptosis (in the late pachytene region of the germline (Gartner et al., 2000; Harrison and
Haber 2006; Hochwagen and Amon 2006). In C. elegans, DNA damage sensors, mrt-1,
hus-1 and clk-2 are implicated in checkpoint signaling (Eichinger and Jentsch 2011; A.
A. Davies, Neiss, and Ulrich 2010; Meier and Gartner 2006; Hochwagen and Amon
2006). mrt-1 and hus-1 are subunits of the highly conserved 9-1-1- complex that
associates with DSBs and facilitates protein-protein interactions between the signaling
pathway components (Hochwagen and Amon 2006). When DNA damage is sensed by
the 9-1-1 complex, then CEP-1(a p53 homolog) initiates the apoptotic pathway leading to
the removal of the damaged nuclei (Eichinger and Jentsch 2011; A. A. Davies, Neiss, and
Ulrich 2010; Meier and Gartner 2006; Hochwagen and Amon 2006). Additionally,
ATM-1 and ATL-1, two conserved DNA damage sensors, also have been implicated in
checkpoint signaling (Eichinger and Jentsch 2011; A. A. Davies, Neiss, and Ulrich 2010;
Meier and Gartner 2006). Despite having key roles in DNA damage signal transduction
in other organisms, CHK-2 has a different function in the C. elegans germ lines, it is
required for pairing (MacQueen 2001).
Between the synapsis checkpoint and pachytene checkpoint, nuclei defective in
synapsis and meiotic recombination are eliminated from the germline via apoptosis, as
previously stated, and subsequent engulfment by the sheath cells. However, synapsis or
DNA damage initiated apoptosis is CED-1 initiated leading to the activation of the core
apoptotic machinery(Gumienny et al., 1999; Gartner et al., 2000). However, it is unclear
how physiological apoptosis is initiated(Gumienny et al., 1999; Gartner et al., 2000).
Once a nucleus undergoes apoptosis, the sheath cell engulfs it, disconnecting it from the
23 common core of the rachis; it is then removed from the germline by the sheath
(Gumienny et al., 1999).
Late meiotic prophase I events
Following CO formation, chromosomes enter the diplotene and diakinesis stages
where they prepare for the meiotic divisions. It is here that SC disassembly, formation of
the cruciform structure (a bivalent structure with long and short arms) and the further
condensation of the chromosomes takes place. COs have a key role in these late meiotic
events and as previously mentioned, divide the chromosomes into a long arm and a short
arm, giving the bivalent an asymmetry. These arms become more evident as chromosome
condense in diakinesis (Figure 1-6).
In order for the meiotic divisions to occur, the SC must now be disassembled so
that chromosomes will be only connected via sister-chromatid cohesion and crossovers.
Disassembly in C. elegans takes place in three distinct steps. In the first step, the
asymmetric disassembly of the SC, the SYPs dissociate from the long arms while
remaining localized on the short arms. The lateral element proteins, HTP-1/2 are removed
from the short arms while remaining localized on the long arms (Martinez-Perez et al.,
2008). The HTP-3 and HIM-3 remain localized along the length of the chromosomes as
does REC-8 (Figure 1-6) (Martinez-Perez et al., 2008).
In the second step, the physical re-arrangement of the chromosome arms into the
distinctive cruciform, the resolution of the chromosomes occurs when the remaining
SYPs are depolymerized leaving both arms of the homologous pair free to rotate around
the chiasmata; however, the lateral elements remain along the chromosome axis and
rotate with the chromosome arms (Martinez-Perez et al., 2008). At this point only the
chiasmata and cohesion are holding the entire structure together (Figure 1-6) (MartinezPerez et al., 2008). This process is initiated as the nuclei enter diplotene and completes as
the nuclei migrate into the next stage, diakensis. The establishment of the cruciform
structure is critical to ensuring that the bivalent is correctly oriented on the spindle
24 (Ishiguro and Watanabe 2007; Watanabe 2005). Once the cruciform bivalent is formed,
the remaining SYPs localize to the interface between the homologous chromosomes,
which is referred to as the short arm (Martinez-Perez et al., 2008; de Carvalho et al.,
2008). HTP-1/2 and LAB-1 now localize to the long arms of the bivalent. HIM-3 and
HTP-3 are localized to both arms and cohesion is still intact along both arms of the
bivalent (Figure 1-6) (Martinez-Perez et al., 2008; de Carvalho et al., 2008).
The last step of SC disassembly the step-wise disassociation of all SC proteins.
(Figure 1-6). The SYPs dissociate from the short arms as the oocyte matures just prior to
fertilization. The lateral element proteins will disassociate from the bivalent only after
MI. The chromosomal passenger complex (CPC) consisting of AIR-2 (Aurora B kinase
homolog), CSC-1(Borealin) and BIR-1(Survivin) localizes to the short arms, where AIR2 phosphorylates REC-8, resulting in its cleavage and subsequent separation of the
homolog (Martinez-Perez et al., 2008; de Carvalho et al., 2008). The localization of the
CPC, specifically AIR-2 to the short arms is mediated by LAB-1 that functions in the
protection of REC-8 along the long arms of the bivalents (Martinez-Perez et al., 2008; de
Carvalho et al., 2008). Interestingly, SGO-1 that is typically active in other organisms in
preventing REC-8 cleavage, does not act in C. elegans meiosis (Kitajima et al., 2010;
Ishiguro and Watanabe 2007; Langegger, Hauf, and Kitajima 2007; Watanabe 2005).
All these steps must be properly executed as any deviation will result in problems
with segregation. If homologous chromosomes remain bound together by the SC proteins
or by the cohesion complex (Siomos et al., 2001; Rogers et al., 2002) mis-segregation
will occur during the MI divisions. While these steps can be observed cytologically, we
do not know the underlying mechanism by which they occur. Mutations in the worm
proteins LAB-1 and HTP-1/2 cause defects in the asymmetric disassembly, but also
exhibit upstream defects in chromosome pairing and synapsis.
Recent work by in our laboratory has uncovered another key player in SC
disassembly, akir-1(Clemons et al., 2013). AKIR-1 does not affect the assembly of the
25 SC or any other upstream events such as cohesion establishment or pairing, but appears
to have a distinct role in the removal of the SYPs from the chromosomes during the
disassembly process (Figure 1-7). Defects in the structure of the bivalent are observed
when SYPs are not properly removed in akir-1 mutant (Clemons et al., 2013).
Quantification of SYP signal intensity in mid diakinesis oocytes shows there is triple the
amount of SYP as compared to wild-type. As these oocytes mature, SYP dissociation is
delayed in late diakinesis oocytes(Figure 1-7). While the SYPs are affected, the lateral
elements are unaffected suggesting that AKIR-1 acts only on CR proteins in their
dissociation from the chromosomes and bivalents (Clemons et al., 2013). This has a
direct impact on the formation of the cruciform structure and the resolution of the short
arms. If the SYPs remain on the arms of the chromosomes and continue to connect the
homologs in a transverse fashion, the chromosomes are not free to rotate around the CO
during resolution. This was reflected in the abnormal bivalent structures of akir-1
mutants (Clemons et al., 2013). This work indicate that SC disassembly is a highly
regulated process; more work is necessary to determine the molecular players in this
process.
The meiotic divisions
In the meiotic divisions, it is critical for the proper segregation of the homolog
chromosomes to take place first (MI) and then for the sister chromatids to separate (MII).
In many organisms, once the SC is fully disassembled, the chiasmsa (the physical
manifestation of the CO) and cohesion hold the homologous chromosomes together(Page
and Hawley 2004). After MI, sister chromatids are bound only at their centromeres where
cohesion is protected by SGO-1/PP2A (Ishiguro and Watanabe 2007; Watanabe 2005). In
many eukaryotes, centromeres consist of repetitive regions, surrounded by
heterochromatin that mark where sister chromatids can bind to each other and remain
bound throughout prophase (Yamagishi et al., 2014). At the centromeres, the
nucleosomes are replaced with a centromere specific nucleosome, CENP-A which is
26 highly conserved from yeast to humans (Yamagishi et al., 2014). At the centromeres,
kinetochores are formed to allow the binding of the microtubules radiating from the
spindle to attach to the chromosomes to pull them towards the spindle, facilitating
separation of the homologous chromosomes in MI or the sister chromatids in MII
(Yamagishi et al., 2014). The orientation of the centromere/kinetochore is essential to
ensuring that the segregation of the chromosomes in MI and MII divisions if correct. As
the spindles are forming, the sister chromatids are bi-oriented, meaning that their
kinetochores are oriented in the same direction and will be pulled towards the same
spindle pole (Yamagishi et al., 2014). The tension caused by the chiasma assist in the
orientation of the sister chromatids (Ishiguro and Watanabe 2007; Watanabe 2005). As
the MI division progresses, the homologous chromosomes segregate. In the MII division,
the cohesion at the centromeres is cleaved and the kinetochores are now in a monopolar
attachment, allowing for the sisters to be pulled to the opposite poles, thus completing
meiosis (Ishiguro and Watanabe 2007; Watanabe 2005).
C. elegans chromosomes are holocentric and therefore, lack defined centromeres
to provide the proper orientation during the divisions, especially during the first division
where the homologs must separate. Kinetochore components, CENP-a, CENP-c, KNL-3
and KNL-1, form cup-like structures covering the long arms of the bivalents (Wignall et
al., 2003; Dumont, Oegema, and Desai 2010). A ring shaped protein structure, consisting
of the CPC and KLP-19/CEMP-F (motor proteins), encompasses the bivalent around the
short arms(Wignall et al., 2003; Dumont, Oegema, and Desai 2010). This represents the
kinetochore structure in C. elegans. These kinetochores orient and align the bivalent with
the homologs pointing towards the poles(Wignall et al., 2003; Dumont, Oegema, and
Desai 2010). Instead of the microtubules attaching to the kinetochores and pulling the
homologs apart, the CPC/KLP-19/CEMP-F ring promotes CLASP dependent
microtubule assembly, thereby pushing the homologs apart as the spindle poles are pulled
apart (Dumont, Oegema, and Desai 2010). During MII division, the kinetochores
27 assemble along the entire pole side of the sister chromatids as in mitosis and the process
repeats to separate the sister chromatids once cohesion is cleaved by separase after LAB1, HTP-3, and HIM-3 dissociate (de Carvalho et al., 2008; Maddox et al., 2004; Wignall
and Villeneuve 2009; Dumont, Oegema, and Desai 2010). If all the events of prophase I
have been completely successfully, the resulting gametes should have a full complement
of chromosomes that is exactly half the parental ploidy.
Thesis outline
The focus of this research is to understand the molecular mechanisms of SC
morphogenesis in the context of the akir-1 mutant background. To accomplish this, an
RNAi screen was conducted over two lesser studied chromosomes in the akir-1 mutant
background to identify genes that had a role in SC disassembly. This work is described in
Chapter 2. The results of the RNAi screen lead to the discovery of a SC assembly defect
as the result of the knockdown of a CSN/COP9 signalosome subunit, csn-5. In Chapter 3,
the novel role for the CSN/COP9 signalosome in meiotic prophase I events is described
and examined. Chapter 4 is a review of the methods used in these investigations. Chapter
5 is a summary of this work and the future studies that may be undertaken to extend the
analyses started in this thesis.
28 Figure 1-1: Events in meiosis. Progression through meiotic prophase I events ensures
proper segregation at the meiotic divisions. Maternal homolog in red and paternal
homolog in blue at left. Meiotic synthesis occurs and cohesion landscape is established
along the sister chromatids prior to entry into prophase I. Prophase I has five stages:
Leptotene, Zygotene, Pachytene, Diplotene and Diakinesis. Each stage has a specific set
of associated events. Leptotene and Zygotene are where pairing and initiation of synapsis
and meiotic recombination take place. As chromosomes move into pachytene, synapsis
and recombination complete. Crossovers are formed. In the next stages, diplotene and
diakinesis, bivalent resultion and desynapsis take place to prepare the chromosomes for
meiotic divisions. In mid diakinesis cellularization occurs. In meiotic division I (MI), the
homologous chromosomes are segregated. In meiotic division II (MII), the sister
chromosomes are separated as cohesion between them is terminated.
29 Figure 1-2: Organization of the C. elegans gonad. A) A schematic representation of
one of the gonadal arms. The somatic tissues are represented in green and the remaining
germline (the nuclei) are represented by the blue circles, crescents and the chromosomes.
Each of the stages of meiosis are depicted: pre-meiotic tip (PMT), the transition zone
(TZ) where leptotene and zygotene take place, pachytene, diplotene, and diakinesis. B) A
micrograph image of one wild-type gonadal arm stained with DAPI (gray-scaled). The
oocyte most proximal in the gonad and closest to the spermatheca is referred to as the
diakinesis -1 or D -1 oocyte. The third gonad from the spermatheca is referred to as the
diakinesis -3 or D-3 oocyte. The image is projection of a three-dimensional data stack
taken at 60X magnification. Scale bar is 10µm. C) A schematic representation of a cross
section of a gonadal arm in pachytene depicting the hollow tubular nature of the gonad,
the rachis.
30 Figure 1-3: A representation of the SUN-KASH proteins in C. elegans. SUN-KASH
proteins for a trans-membrane bridge that facilitates chromosome movement during the
pairing of homologous chromosomes in leptotene and zygotene. Pairing centers interact
with the SUN-1 protein that spans the inner nuclear membrane. The KASH protein ZYG12 spans the outer nuclear membrane and binds to the SUN-1. During pairing, each
chromosome is bound to an individual SUN-KASH complex that facilitates chromosome
movement via the interaction of the KASH protein with dynein.
31 Figure 1-4: A representation of the general structure of the synaptonemal complex.
A) A general schematic representation of the conserved structure of the synaptonemal
complex. The homologous chromosomes are in blue and pink. The lateral elements are in
green and the central region proteins are in red, forming the transverse elements that bind
the homologs together in synapsis. B) A representation of the C. elegans complex. The
homologous chromosomes are represented in blue and pink. The lateral elements HTP-3
and HIM-3 are shown in green and are the first to localize to the chromosome axis. HTP1/2 and LAB-1 lateral elements are in gold and are the next to localize along the
chromosomal axis. The four central regions proteins, the SYPs, are presented in red.
They are last to localize to the SC and their polymerization is interdependent, meaning all
four SYPs must be present for polymerization to occur.
32 Figure 1-5: A simplified diagram of meiotic recombination. SPO-11 and its associated
proteins initiate double strand breaks (DSBs) on one of the sister chromatids. RAD-51
localizes to the resected ends of the break. Next strand invasion and synthesis occurs. If
double Holliday junction is formed and cleaved by resolvases asymmetrically, a
crossover will be formed. COSA-1, a crossover specific protein will localize to the
crossovers. Subsequently, if the Holliday junction is cleaved symmetrically, a noncrossover will be formed. Alternatively, non-crossovers may be formed if the invading
strand dissociated and re-anneals via synthesis dependent strand annealing (SDSA)
mechanism.
33 Figure 1-6: C. elegans specific progression through the events of meiotic prophase.
In C. elegans, synapsis occurs prior to recombination. De-synapsis occurs in a step-wise
process: 1) the asymmetric de-synapsis with SYPs relegated to the short arms and
HTP1/2 and LAB-1 remaining on the long arms. HTP-3 and HIM-3 remain on both arms.
2) Resolution of the bivalent and formation of the cruciform structure. The SYPs on the
short arms dissociate allowing for the movement of the chromosomes around the
chiasmata to form the bivalent cruciform structure. The asymmetry of the chromosomes
is now realized in the long and short arms of the bivalent. 3) Step wise removal of the SC
proteins as the oocyte matures and is about to be fertilized. The SYPs are localized to the
short arms of the bivalent, between the homologous pairs of chromosomes. LAB-1 and
HTP-1/2 are localized to the long arms, protecting the cohesion between the sister
chromatids in the MI division.
34 Figure 1-7: Synaptonemal complex disassembly and bivalent defects in akir-1
mutants. Micrographs of SYP-1(red) and DAPI(blue) stained wild-type (A ,B) and akir1 mutant oocytes (C, D). Images are projections of a three-dimensional data stack. Scale
bar is 2µm. AKIR-1have abnormal bivalent structure in D-1 oocytes and SYP-1
aggregates present. Wild-type D -1 oocytes have a compact bivalent structure and no
SYP-1 present.
Table 1-1: Conservation of meiotic factors
A.
S.
thaliana
cerevisae
#chromosome
10
32
(2n)
Chromosome
Movement
SUN/KASH
component
35 C.
D.
elegans melanogaster
12
SUN-1/
ZYG-12
Microtubules/
dynein
M.
muscalsus
8
40
N/A
SUN-1/2
Microtubules
Microtubules
SUN1/2
Mps3/
Csm4
Movement
mediator
N/A
Actin
Pairing
mechanism
Bouquet
Bouquet
Pairing
Center
Centromere
Bouquet
ZYP1
Zip1
SYP1/2/3/4
CONA,
C(3)G,
Corolla
SYCP1,
SYCE1/2/
3, TEX12
ASY1
Red1,
Hop1
HTP1/2/3,
LAB-1,
HIM-3
C(2)M, ORD
SYCP2/3,
HORMA
D1/2
6
6
22-24
Synapsis
Central
region
Lateral
elements
Crossovers
9
90
/meiosis
Modified from Lui and Colaiacovo, 2013
36 CHAPTER 2
RNAi SCREEN FOR ENHANCERS AND SUPPRESSORS OF MEIOTIC DEFECTS
OF AKIR-1 MUTANTS
Introduction
I performed an enhancer/suppressor screen aimed to identify proteins required for
proper SC disassembly and meiotic chromosome segregation utilizing a newly discovered
SC mutant, akir-1. AKIRINs [also known as FBI1 (mammalian AKIRIN2), MIGHTY
(mammalian AKIRIN1), subolesn (ticks), and bhringi (flies)] are highly conserved
proteins (Macqueen and Johnston 2009)with no previous known role in meiosis. Meiotic
roles for these proteins can be suggested based on their high expression in rat and fly
testis as well as the C. elegans gonad (Komiya et al., 2008; Chintapalli, Wang, and Dow
2007; Rebecca Hunt-Newbury 2007). C. elegans and D. melanogaster have one copy of
the gene while vertebrates have two. None have been found in bacteria, yeast or plants
(Goto et al., 2007). The biological functions for AKIRIN are diverse: tumor
proliferation(Komiya et al., 2008), myoblast differentiation (Salerno et al., 2009), innate
immune response (Goto et al., 2007), and muscle development (Nowak et al., 2012).
AKIRIN is thought to have also a role in transcription through interactions with
transcription factors and chromatin remodelers (Goto et al., 2007; Nowak et al., 2012).
Akirin was isolated as a gene involved in innate immune response in Drosophila
(Goto et al., 2007). It had been established that AKIRIN2 (then known as FBI1) was an
NF-kB interacting protein and transcriptional activator in mammalian cell lines{(D. K.
Lee et al., 2005). Both of these authors demonstrated that AKIRINS were nuclear
proteins and that the nuclear localization signal is located on the C-terminal end.
However, Goto et al., 2007 was the first to show that lack of AKIRIN leads to embryonic
lethality in both Drosophila and mammals. Furthermore, these authors confirmed
AKIRIN’s role in embryonic development by examining a transgenic mouse line:
37 akirin1-/- mice did not display embryonic lethality in contrast akirin2-/- did, dying before
embryonic day 9.5. The cause for this lethality was not determined (Goto et al., 2007).
akirin was recovered by a RNAi screen in Drosophila S2 cells aimed at isolating
components of innate immunity response signaling pathways; mainly in the Imd and Toll
pathways (Goto et al., 2007). It was determined that Akirin acted in parallel with NF-kB
in transcription of key immune genes required for defense against Gram-negative bacteria
(Goto et al., 2007) and suggested that AKIRIN has a direct a transcriptional function in
the immune response {Komiya:2008id, (Goto et al., 2007). Akirin’s function in
transcriptional regulation in development in Drosophila was confirmed by Nowak, et al.,
2013. Here, akirin mutants were shown to display a wide range of muscle development
defects, which resulted from differential regulation of myogenesis. Specifically, Akirin
genetically and physically interacts with Twist and the Brahma complex to regulate
embryonic myogenesis. Homozygote embryos die as a result of abnormal development of
somatic musculature (Nowak et al., 2012). While this work established a mechanism for
developmental defects in AKIRIN mutants, our lab demonstrated a role for AKIRIN in
meiosis in worms.
C. elegans akir-1 mutants have a defects in the disassembly of the SC consisting
of the aggregation of the central region protein SYP-1 (discussed in Chapter 1) which
leads to an abnormal bivalent structure (Clemons et al., 2013). In addition, chromosomes
fail to fully condense and these defects are independent of the condensin complex.
Furthermore, meiotic divisions are delayed and lagging chromosomes are observed in
both meiotic divisions. Two alleles were studied, akir-1(rj1) and akir-1(gk528) (Clemons
et al., 2013). The rj1 allele is a point mutation, a histadine to proline substitution at
position 190, and was isolated from a forward screen (Clemons et al., 2013). The gk528
deletion allele removes 42% of the protein including the promoter region and this mutant
obtained from the C. elegans Gene Knockout Consortium (Oklahoma Medical Research
Foundation). Both alleles exhibit a reduction in the number of eggs laid (rj1- a 5 fold
38 reduction and gk528- 3 fold reduction) and embryonic lethality (rj1-21% and gk258- 16%
increase) (Clemons et al., 2013).
AKIR-1 is proposed to regulate the removal of SYPs from the long arm of the
bivalent in a way that prevents the accumulation of the unloaded SYPs back on the short
arm of the bivalent at diakinesis (Clemons et al., 2013). This would allow for the proper
resolution of the chromosome arms to for a normal bivalent structure (Clemons et al.,
2013). In akir-1 mutants, the unloaded SYPs from the long arm aggregate on the short
arms, preventing proper resolution, distorting the shape of the bivalent and generating a
gap between the sisters in the bivalent. As the oocyte matures and SYPs are removed
from the short arm, this gap remains generating an abnormal bivalent structure (Clemons
et al., 2013).
Not much is known about the mechanisms of SC disassembly, due to the fact that
only few genes have been identified to act in this process. The screen I performed was
undertaken with the intent to identify additional genes promoting proper SC disassembly
via their genetic interaction with akir-1. We used a RNA interference (RNAi) feeding
protocol to conduct the screen, a common tool to knock down gene function via the
degradation of mRNA (R. Kamath 2003). This screen involved comparing the effect of
the RNAi of clones from Chromosome II and IV on the number of progeny produced in
the akir-1 mutant background as compared to the wild-type background. Clones that
demonstrated an effect were then subjected to cytological analyses to determine if the
affect on embryonic lethality was due to defects in chromosome structure or SC in
prophase I.
The advantages of using RNAi are several fold. First, clone identity is linked
directly to the phenotype. Secondly, RNAi can create a partial loss of function which
helpful when studying genes with multiple functions, by allowing the by-pass of early
developmental defects and examination of phenotypes in gonadal tissues. We examined
the additive effects of having two proteins depleted from meiotic pathways. As
39 previously mentioned the akir-1 background had a higher level of embryonic lethality,
therefore any RNAi clone that enhanced or suppressed that lethality was subjected to
further analyses. A feeding library of about 16,757 clones was obtained for this study (R.
Kamath 2003). This library had been utilized in other screens in a high throughput
manner to investigate gene function on a genome wide level in C. elegans (R. S. Kamath
et al., 2003; A. Fraser, Kamath, and Zipperlen 2000).
Results
An RNAi screen identified potential genes that interact with akir-1
This RNAi enhancer/suppressor screen was designed to recover genes that
interact with akir-1 in SC disassembly. Because akir-1 mutants already had an increased
embryonic lethality due to SC and bivalent structure defects, we used that lethality as a
means to measure interaction with akir-1. From data available, we assumed that akir-1
only acted in meiosis and the reduced fecundity was due the meiotic defects observed.
The RNAi clones examined enhanced, suppressed or had no affect on the lethality of
akir-1 mutant background. We sought clones affecting the akir-1 background more than
the wild-type background. These clones should define genes that interact with akir-1
directly or act in parallel pathways of SC disassembly.
We chose to screen clones from chromosomes II and IV because they were
previously understudied in other RNAi screens (Simmer et al., 2003; R. S. Kamath et al.,
2003; S. S. Lee et al., 2002) In total, 6,247 clones were examined in the preliminary
screen, 3,552 from chromosome II and 2,695 from chromosome IV. This encompassed
approximately 37% of the C. elegans genome. The screen was conducted in two phases.
The first phase, a preliminary screen of all clones on chromosomes II or IV, relied on a
visual assessment of F1 progeny. Clones were scored as having a more (suppression of
lethality) or less (enhancement of lethality) progeny as compared to the empty vector
controls for the akir-1 mutant and wild-type backgrounds. Any clone that passed the
preliminary screen was then evaluated in the second phase, a secondary screen where the
40 clones were evaluated for reproducibility of the lethality observed in the preliminary
screen and F1 progeny were quantified. Clones that passed the secondary screen were
then selected for cytological analyses (Figure 2-1).
For the chromosome II preliminary screen, a liquid culture protocol was utilized
to facilitate a high throughput analysis of clones. This protocol was developed and used
in to evaluate gene knockdown in a high-throughput fashion (Lehner, Tischler, and Fraser
2006). One change that we made was to expose L1 larvae to RNAi, as opposed to a later
developmental stage, to ensure the maximum amount of exposure to the RNAi plasmids
to the F1 offspring. While the feeding protocol does facilitate a high throughput of
screening, it is the least efficient method of introducing RNAi to the worm and can result
in lower penetrance of phenotypes when compared other methods of introducing RNAi
plasmids such as injection or soaking (R. S. Kamath et al., 2001).
Initially, only 1,152 RNAi clones were cultured for the chromosome II
preliminary screen, L1 worms of the akir-1(rj1) and wild-type were placed into the
media, grown to adults and the number of F1 progeny were scored visually for an
increase or decrease in progeny as compared to the controls (wild-type and the pL4440
empty vector). This pilot study was conducted to test the efficacy of the liquid protocol
and assess the screen design. We recovered eight clones that demonstrated differences in
embryonic lethality. However, we had to abandon the liquid protocol after 1,152 clones
as when we started the next round screening, we experienced difficulty with lethality of
the wild-type P0 (parental generation) worms and those that survived had greatly
reduced fecundity. We tested reagents, media, and consumables (the 96-well plates used
in the screen) to determine what could cause a high level of lethality in the wild-type
controls. After determining that there was a problem with the laboratory distilled water
several sources of water from the Biology Building were tested. None would allow for
the growth of worms in liquid culture. Although the water in the Biology Building is
filtered and purified, we cannot control all the trace chemicals that might be found in it.
41 While this is not a problem for the NGM plate cultures, which we frequently used for
general worm culture, because we were using a liquid culture methodology for this RNAi
screen, it could have been a factor. It is known throughout different labs in the Biology
Department that the water quality is variable throughout the year (personal
communication with Malone and Logsdon Labs).
While the akir-1(rj1) worms appeared to not be affected, the increase in lethality
across the majority of wild-type controls meant we could not properly assess the changes
in embryonic lethality as we lacked the proper wild-type controls. It could be that the
akir-1 mutants are not as sensitive to the water issue compared to the wild-type. This
sensitivity could be related to AKIR-1’s role in the immune response (discovered in other
organisms, but not studies in C. elegans). Although both wild-type and akir-1(rj1) strains
are isogenic, (akir-1(rj1) was outcrossed to wild-type 6 times), there are not completely
identical due to: 1) possible mutations linked to akir-1(rj1) that were not cleared by
outcrossing and 2) the propagation of these strains for a year since the outcross that may
lead to accumulation of mutations. Therefore, it could be that akir-1 worms contain a
linked unknown gene mutation that infers resistance or that the wild-type contain
mutation in a gene that results in sensitivity to environmental conditions. At this point,
the underlying cause is undetermined. For this screen it was essential to know if clones
were interacting only with the akir-1 mutant background. The wild-type controls were
critical in determining if a clone interacted with akir-1 or if it affected both backgrounds.
We did not rescreen the original 1,152 clones as we had no reason to believe the pilot
data to be problematic.
I completed the preliminary screen on the remaining 2,400 clones from
chromosome II using a traditional plate methodology, which is more established method
with less variability. While we retained the bulk of the screen protocol, the only major
change using individual NGM plates instead of a 96-well plate liquid format. While this
method was more labor intensive, the results were reproducible and we had no issues
42 with increased lethality in the wild-type controls. The clones were visually screened for
increases or decreases in embryonic lethality as compared to the controls. Out of the
3,552 clones examined, 229 demonstrated a change in embryonic lethality (Table A-1).
For chromosome IV preliminary screen, we used the traditional plate methodology on the
akir-1(gk528) mutant and the wild-type background. RNAi clones were seeded onto
plates, L1 larvae were placed on the plates, and number F1 progeny were scored visually.
Of the 2695 clones examined, 207 demonstrated a change in embryonic lethality (Table
A-2).
For the secondary screen, the 229 clones from chromosome II and the 207 from
chromosome IV were screened again in triplicate using a traditional plate method for
both the akir-1 mutants (rj-1 allele for Chromosome II and gk528 allele for Chromosome
IV) and wild-type backgrounds and their F1 progeny were quantified (Tables A-1 and A2). This secondary screen allowed us to quantify the number of F1 progeny produced for
each clone. The average and standard deviation were calculated for each genotype . We
then normalized (See Methods in Chapter 4 for detailed description of normalization
analyses, including example calculation). the averages to the pL4440 empty vector
controls for each genotype. Initially, we normalized the backgrounds to each individual
set of experiments and their controls (Table A-3). To determine if there was a change in
embryonic lethality in the respective akir-1 background, we took the ratio of the
normalized average for the akir-1 background to the normalized average for the wild-type
background. This would provide a value by which we could measure embryonic lethality
in the akir-1 background and use the value to select clones for cytological analysis. We
had anticipated using this ratio as a threshold that would potentially link cytological
defects with a reduction in F1 progeny. Since we are comparing the two backgrounds to
look for a change in lethality, the empty vector control ratio should equal 1.0. Clones that
were close to 1.0 would be weak enhancers or suppressors and therefore unlikely to yield
a cytological defect. We set an arbitrary threshold of 0.90 for enhancers and 1.10 for
43 suppressors. Any clone that fell below or above those thresholds could be considered for
cytological analyses although we chose the most severe enhancers and suppressors (Table
A-3). However, once we completed the entire screen, we observed variation between the
controls. To eliminate this variation, we averaged the controls and then normalized the
clone averages to the new control values (Tables A-1 and A-2).
Cytological analyses of chromosome II clones
Out of the 229 clones that were quantified from the Chromosome II screen, 52
were selected for cytological analysis (Table A-3). We selected clones with ratios
representing the most severe enhancers and suppressors in order to assess if the ratio of
progeny could be associated with a prophase I defect detectable by cytology.
Additionally, we included clones with ratios closer to the control ratio of 1 to verify that
these clones would not have cytological defects. This would allow us to more precisely
determine the threshold at which we could detect cytological defects. In the future, this
would allow us to select clones based on their ratio. The RNAi experiments were carried
out on both akir-1(rj1) mutant and wild-type backgrounds and SYP-1 immunostaining
was preformed. Gonads were analyzed by fluorescence microscopy and defects in SYP-1
staining and/or bivalent structure were scored. We define SC defects as any abnormal
SYP-1 localization that occurs between the initial localization of SYP-1 in the TZ to the
disassociation in D -1 oocyte. This includes: aggregation upon assembly, lack of
localization, premature removal, and aberrant removal (both arms retained, removed form
both, diffused or aggregated). Bivalent defects are defined as univalents, chromosome
clumps, lack of chromosome condensation, fragments or any other defect that results in a
lack of condensed cruciform structure as observed in wild-type diakinesis oocytes. While
we examined the entire gonad for defects such as alteration of the length of the various
stages, defects in SC staining from TZ-late pachytene, or gross morphological defects in
the entire gonad, quantitative analysis was done on the D -1 oocytes as this was where the
akir-1 mutant phenotype was most prominent. All 52 clones were examined in the akir 44 1(rj1) background. Of the 52 clones analyzed, two: F10G7.4 (ratio=0.5) and a novel open
reading frame (ORF), ZC329.9 (ratio=0.88) demonstrated bivalent defects on the akir1(rj1) background. Both of these were considered to be enhancers of the akir-1 mutant
phenotype and we validated the clones by sequencing. The remaining clones did not
demonstrate any SC or bivalent defects in D -1 oocytes (Table 2A-4 and A-5).
F10G7.4
We identified unc-85 (F10G7.4) as having a potential interaction, enhancing the
embryonic lethality, on the akir-1(rj-1) mutant background (Table A-5). 75% of D-1
oocytes in akir-1(rj1);unc-85(RNAi) had a bivalent defect indicated by the presence of
univalents (Figure 2-2D) when compared to the akir-1(rj1),pL4440(RNAi) that had no
univalents (Figure 2-2B) (akir-1(rj1);unc-85(RNAi) n=32, akir-1(rj1),pL4440(RNAi);
p<0.0005 Fishers Exact Test). Univalents were not seen in the unc-85(RNAi) (Figure 23C’). SYP-1 was not preformed during the first RNAi experiments and since the
chromosome defects appeared to be highly penetrant, we decided to focus on the bivalent
defects rather than possible SC defects.
unc-85(ok2125) is a deletion mutants that removes the entire gene from the
promoter region to the 3’UTR, rendering this a null allele (provided by the C. elegans
knockout consortium) . Gonads were stained for SYP-1 and 56% of the D-1 oocytes had
a comparable phenotype to akir-1(rj1);unc-85(RNAi) (Figure 2-3C’). As we were focused
on the bivalent defects observed, we did not perform SYP-1 staining at this time. The
distance between univalents were measured and quantified (Figure 2-2E). There was an
8-fold difference in the distance between the homologs in unc-85(ok2125) (n=32) and
those in wild-type (n=32) (p<0.0005, Mann Whitney Test).
We outcrossed the unc-85(ok2125) strain six times and selected two clones for
analysis. Both clones were scored for SC and bivalent defects. None of the D -1 oocytes
(n=20) examined had any SC or bivalent defects (Figure2-4A”-C” and A’’’-C’’’). One
clone was crossed into the akir-1(gk528) background to create a double mutant and
45 prepared for analysis. Only 13% of the D -1 oocytes had univalents (n=15) while the
other 87% had bivalent defects as seen in akir-1 mutants. A second analysis found 30%
gonads with a bivalent defect with univalent present (n=17). The remaining 70% had
SYP-1 aggregates as observed in akir-1 mutant background. 102 gonads were examined
over five different experiments and no SC defect was identified. Therefore, the unc85(ok2125);akir-1(528) double mutants has a bivalent defect albeit with incomplete
penetrance.
ZC239.6
We identified ZC239.6, a novel ORF, as an enhancer to the reduced fecundity of
akir-1(rj1) mutant background. 80% of D -1 oocytes for akir-1(rj1);ZC239.6(RNAi) had
an absence of bivalent structure with chromosomes having condensation defects. These
defects had a rather stretched out or diffuse appearance (Figure 2-4C’’) when compared
to akir-1(rj1);pL4440(RNAi), where six bivalents could be clearly identified and their
structure was normal (Figure 2-4C) ( akir-1(rj1);ZC239.6(RNAi) n=29, akir1(rj1);pL4440(RNAi) n=37; p<0.0005 Fishers Exact Test). akir-1(rj1);ZC239.6(RNAi)
also demonstrated a wild-type SYP-1 localization pattern in D-1 oocytes (n=15) (Figure
2-4A’’-C’’). ZC239.6(RNAi) had six bivalents with no bivalent defects (n=18). SYP-1
(n=5) staining was normal when compared to that of wild-type (n=32).
No deletion mutant is available. We preformed additional RNAi experiments to
confirm our findings, but we were unable to replicate the abnormal bivalent observed
earlier (Figure 2-4B’’’ and C’’’) with 100% of D -1 oocytes showing an akir-1 mutant
phenotype (n=17). Whole worms were examined and again, no bivalent defects were
observed (n=13). After studying 84 gonads, no SC defect was observed and we are
currently unable to replicate the initial diffuse bivalent defect observed.
Cytological analyses of chromosome IV clones
Out of the 2,695 clones that were preliminarily screened, 207 clones were
quantified for the chromosome IV secondary screen. As we were unable to link the
46 changes in embryonic lethality (akir-1 mutant/wild-type ratio) to cytological defects in
the chromosome II screen, we took a different approach to chromosome IV. Gene
ontology terms (GO Terms) were used to identify those clones involved in reproduction.
Each RNAi clone is identified by a unique sequence identifier, which was used to query
WormBase and build a GO term summary for each clone. Those clones whose GO term
summary included: reproduction, meiosis, oogenesis, meiotic chromosome segregation,
regulation of meiosis, gamete generation, or oocyte development were selected for
cytology. This narrowed the candidates to 25 clones, which were selected for cytological
analyses (Table A-6 and Table A-7). Of these 25 clones, 4 demonstrated either SC or
bivalent defects: B0547.1(ratio=0.15), C09G4.3(ratio=0.23), Y66H1B.2(ratio=0.30), and
F38H4.9(ratio=1.20) in the akir-1(gk528) mutant background.
F38H4.9
The C. elegans homolog of the catalytic subunit of protein phosphatase 2A
(PP2A), a highly conserved protein that regulates chromosome segregation, let92(F38H4.9), is a suppressor of the akir-1 mutant embryonic lethality. akir-1(gk528);let92(RNAi) (n=26) and akir-1(gk528);pL4440(RNAi ) (n=37) had six bivalents with wildtype bivalent structure (Figure 2-5). SYP-1 aggregates were present in both the double
and single mutants. let-92(RNAi) had six wild-type bivalents (n=19) and wild-type SYP-1
staining (n=6).
A deletion mutant let-92(ok1537) removes the promoter and 35% of the coding
region (exons 1 and 2) rendering this allele likely null. However, this strain was not
balanced and we determined that the strain was heterozygous (let-92(ok1537)/+). A total
of 80 worms were singled out onto individual plates and allowed to reproduce. PCR was
performed to verify which worms were homozygous for the deletion. These homozygotes
were then crossed with the nT1 balancer chromosome to generate a balanced strain: let92(ok1537);nT1. Given that this mutant is homozygous larval lethal, we decided to focus
on other candidates.
47 Y66H1B.2
Y66H1B.2 codes for fln-1, the orthologs to human filamin A and is required for
normal fertility (DeMaso et al., 2011). fln-1 was identified as an enhancer, increasing
embryonic lethality in the akir-1(gk528) mutant background. akir-1(gk528);fln-1(RNAi)
displayed a wild-type bivalent structure in 55% of D -1 oocytes (Figure 2-6C-F)(akir1(gk528);fln-1(RNAi) n=54; akir-1(gk528);pL4440(RNAi) n=37; p<0.0005, Fishers Exact
Test). However, these mutants retain the SYP-1 aggregates as seen in akir-1 mutants
(Figure 2-5) (akir-1(gk528);fln-1(RNAi) n=16). fln-1(RNAi) mutants showed no unusual
SC (n=14) or bivalent (n=21) defects.
A deletion mutant, Y66H1B.2(ok2611), was obtained and validated via PCR. As
with the fln-1(RNAi), the deletion allele removes 23% of the N-terminal coding region.
No SC or bivalent defects were observed in Y66H1B.2(ok2611) (Figure 2-6G-H). The
double mutant akir-1(gk528);Y66H1B.2(ok2611) was generated. 75% of the D-1 oocytes
examined had aggregates and bivalent defects seen in akir-1 mutants (n=25) and were not
statistically different from the akir-1 background (Figure 2-6 I-J)(p=0.31)
C09G4.3
C09G4.3 codes for cks-1, a cyclin dependent kinase regulatory subunit, which is
active in cell cycle regulation. akir-1(gk528);cks-1(RNAi) produced 78% of D -1 oocytes
that lacked six distinct bivalents and had clumped chromosomes compared to akir1(gk528), pL4440(RNAi) controls (Figure 2-7) (akir-1(gk528);cks-1(RNAi) n=18; akir1(gk528), pL4440(RNAi) n=37, p<0.0005 Fishers Exact Test). In wild-type D -3 oocytes,
chromosomes have started to condense with the six bivalents becoming distinct and
bivalent remodeling being initiated as the oocytes mature (Chan, Severson, and Meyer
2004a). In akir-1(gk528);cks-1(RNAi) 78 % of D -3 oocytes, the chromosome displays no
bivalent structure (Figure 2-7G) (akir-1(gk528);cks-1(RNAi) n=18; akir-1(gk528),
pL4440(RNAi) n=37, p<0.0005 Fishers Exact Test). csk-1(RNAi) D -1 oocytes also
48 displayed bivalent defects. While the bivalent structure was more condensed and
bivalents could be determined, 41% of oocytes had more than six DAPI bodies or a
possible fragment (Figure 2-7F) (csk-1(RNAi) n=29, pL4440(RNAi) n=43; p<0.0005
Fishers Exact Test).
In addition to the bivalent defects, akir-1(gk528);cks-1(RNAi) had an SC defect as
well. 44% of the D-1 oocytes had SYP-1 localization to the chromosomes and no SYP-1
aggregates (Figure 2-7) (akir-1(gk528);cks-1(RNAi) n=16; akir-1(gk528), pL4440(RNAi)
n=19, p<0.005 Fishers Exact Test). In csk-1(RNAi) there is no SYP-1 localization to the
D -1 bivalents which is comparable to wild-type worms (akir-1(gk528);cks-1(RNAi)
n=16, csk-1(RNAi) n=21, p<0.5 Fishers Exact Test). A deletion mutant, cks-1(tm1916) is
available from the National BioResource Project in Japan, but was not ordered.
B0547.1
csn-5(B0547.1) is a CSN/COP9 signalosome subunit which contains isopeptidase
catalytic core and has been shown to function in the regulation of the CULLIN E3 ligases
(Wei and Deng 2003). Previous studies by Smith et al., 2002 and Orsborn et al., 2007
have shown that csn-5(RNAi) results in smaller gonads, a lack of oocytes and sterility.
However, these authors did not examine prophase I events in these studies.
In akir-1(gk528);csn-5(RNAi) mutants, 80% of the D -1 oocytes displayed SYP-1
aggregation (n=21) and bivalent defects (n=24) as seen in the akir-1 mutant background.
The remaining 20% (n=5) of the oocytes displayed wild-type bivalents. SYP-1 staining
was inconclusive. csn-5(RNAi) in this experiment not only had oocytes, but there were no
SC or bivalent defects in the oocytes. csn-5(RNAi) was also performed on a SYP-3::GFP
background and found to have SYP aggregates in pachytene region of the 37% of the
gonads examined (csn-5(RNAi) n=27, pL4440(RNAi) n=42; p<.0005 Fisher’s Exact
Test). Finding SYP-3 aggregates in early prophase I indicates defects in SC assembly.
Lack of SC assembly, such as syp-1(me17) mutants, leads to defects in pairing,
recombination and crossover formation all of which lead to segregations errors and
49 lethality. When coupled with the embryonic lethality already observed in the akir-1
background, csn-5 knockdown lead to an increase in embryonic lethality and an enhancer
phenotype.
Given that previous studies have indicated a lack of oocytes in csn-5(RNAi), we
concluded that protein knockdown using a RNAi feeding protocol was not sufficient to
induce the previously highly penetrant phenotypes seen in the Smith et al., 2002 and
Orsborn et al., 2007. These authors used the injection method of introducing RNAi into
the gonad. A deletion mutant was available, csn-5(ok1064). This mutant was obtained
and the results of analyses are contained in Chapter 4.
Discussion
F10G7.4
F10G7.4 was identified as an enhancer and codes for UNC-85, one of the two C.
elegans homologs of Asf1 (anti-silencing factor 1). Asf1 is a known histone chaperone
and chromatin remodeler (Ransom, Dennehey, and Tyler 2010; Donham, Scorgie, and
Churchill 2011; Polo 2014)and expressed in the germline (GRIGSBY and FINGER
2008). Originally identified through studies in yeast, Asf1 mutants show defects in DNA
replication and repair as well and transcription (Ransom, Dennehey, and Tyler 2010).
Asf1 is highly conserved and facilitates chromatin disassembly ahead of replication fork
and reassembly of chromatin after replication is complete (Ransom, Dennehey, and Tyler
2010). The reassembly of chromatin after replication and repair is due to an interaction
between Asf1 and CAF-1(chromatin assembly factor 1) where CAF-1 is recruited to the
newly synthesized DNA and Asf1 chaperones H3 histones to CAF-1, which incorporates
them into chromatin (Ransom, Dennehey, and Tyler 2010). Asf1 is considered a nuclear
chaperone of histones. Grigsby et al., 2008 showed that unc-85 is not only highly
expressed in replicating somatic cells, but also throughout the entire C. elegans gonad
and localized to the sites of DNA replication and repair, which is consistent with known
50 Asf1 functions. Grigsby et al., 2008 also identified unc-85 as having defects in postembryonic division, which were attributed to replication defects.
Given that there are two Asf1 homologs in C. elegans, unc-85 and asfl-1, with
functional redundancy in germline replication (Grigsby et al., 2009), it would be prudent
to examine the interaction of asfl-1 on the akir-1 mutant background. Furthermore, while
unc-85 and asfl-1 single mutants do not exhibit germline defects, the unc-85;asfl-1
double mutant does have smaller gonads with abnormal morphology, lack of bivalents
(DAPI body clumps) and a increase in apoptosis (Grigsby et al., 2009). This data
suggests that akir-1;unc-85 act similarly to unc-85;asfl-1 and may hint to a role for akir-1
in modifying chromatin structure, possibly as a nuclear chaperone. As only the DAPI
bodies were examined, the size and morphology of akir-1;unc-85 double mutants should
be examined further. Additionally, apoptosis should be examined. If the defects of the
akir-1;unc-85 double mutants are similar to that of the unc-85;asfl-1 double mutants, this
may suggest a role for AKIR-1 in chromatin modifications which would warrant further
investigation. unc-85 and asfl-1 may act in upstream of akir-1 as the unc-85; asfl-1
double mutant affects the germline prior to SC disassembly, were akir-1’s function is
restricted to SC disassembly. However, it is possible that the akir-1;unc-85 double
mutant will be also affecting upstream meiotic events similarly to unc-85;asfl-1.
Examination of the RNAi clone F10G7.3/unc-85 from the Ahringer library shows
that asfl-1 is a secondary target of the clone (the genes share a 79% homology in the
nucleotide sequence) (Grigsby and Finger 2008). It is likely both genes were targeted and
the bivalent defect we observed could have be the result of the knockdown of both genes
in the akir-1 mutant background. If both genes are indeed targeted, I would also expect a
reduction in the wild-type background as well. However, since that was not observed, it
could be that the depletion of both unc-85 and asfl-1 may have a greater affect in the
akir-1 background, leading it to enhancement of lethality, due the bivalent defects.
51 Although we initially saw a similar bivalent defect in the unc-85(ok2125), this
defect was lost upon outcrossing and is consistent with the lack of defects observed in
unc-85(RNAi). It is possible there could have been a secondary mutation in the unc85(ok2125) background that contributed to premature separation. While the unc85(ok2125);akir-1(528) did not have a bivalent defect and the RNAi clone F10G7.3 also
targets asfl-1. If the asfl-1;akir-1 double mutant shows a bivalent defect then it could
indicate a possible interaction between akir-1 and both unc-85 and asfl-1. If the bivalent
defect is recapitulated, then this would support the argument that unc-85 and asfl-1 are
redundant and as well as indicating an interaction with akir-1.
F38H4.9
let-92(F38H4.9) is a does not affect akir-1 mutant lethality (ratio=1.2) but was
selected based on the GO terms. let-92 the C. elegans homolog of the catalytic subunit of
protein phosphatase 2A (PP2A), a highly conserved protein that regulates chromosome
segregation. PP2A is a heterodimeric complex consisting of a catalytic subunit, a scaffold
subunit and a regulatory subunit which specifies localization (Schindler 2011). In both
mitosis and meiosis, PP2A, through its interaction with shugoshin (SGO), is essential in
maintaining sister chromatid cohesion throughout cell division (B. Xiong and Gerton
2010). Cohesion sister chromatids is established during S phase, and is maintained
throughout prophase by SGO and PP2A. During metaphase in most organisms, cohesion
is removed from the arms of the chromosomes, but retained at the centromeres to
maintain the connection between the chromatids (B. Xiong and Gerton 2010). However,
in C. elegans LAB-1 replaces SGO-1as the “protector” of cohesion. It is not known if
LAB-1 interacts with PP2A (de Carvalho et al., 2008). The loss of a functional PP2A
complex results in premature separation of chromatids and defects in chromosome
segregation. Mutants of let-92 are known to be homozygous lethal.
Nolt et al., 2011 showed that in yeast, PP2A mutants, specifically, PP2ACdc55 had
various prophase I defects in pre-meiotic DNA replication and homologous
52 recombination. In C. elegans, mutants with defects in recombination lead to altered
numbers of DAPI bodies in D -1 oocytes (Lui and Colaiácovo 2013). Given the known
mitotic defects for depletion of PP2A and the defects in prophase I (Nolt et al., 2011), a
chromosome defects was with expected numerous (7-24) DAPI bodies, clumping of
chromatin, or a combination of these chromosome alterations. The fact that we saw no
defects in either of the RNAi experiments suggests the RNAi failed. At most, there could
have been a hypomorphic situation generated with the knockdown of let-92, again
leading to chromosome defects. The lethality ratio was only 1.2 suggesting that there was
no affect or a very weak suppression. This is likely due to the affect of let-92 on both
backgrounds which would have produced a ratio that appears to be normal or close to
normal, despite being homozygous lethal in deletion alleles (Figure 2-1B).
Y66H1B.2
C. elegans has two filamin genes: fln-1, which is highly conserved and fln-2,
which is poorly conserved. fln-1 has three isoforms with fln-1a being the full-length
transcript (DeMaso et al., 2011). Filamin has roles in maintaining the actin cytoskeleton,
acting as a scaffold for signaling components, and as a molecular sensor for mechanical
forces (Kovacevic, Orozco, and Cram 2013). In C. elegans, FLN-1 is thought to initiate
the release of calcium as an oocyte enters the spermatheca, activating a signaling cascade
that transduces the signal for the spermatheca to contract (Kovacevic, Orozco, and Cram
2013).
We did not test if Y66H1B.2(ok2611) is a null allele; however, Kovacevic and
Cram, 2010, did not observe any defects in brood size. Trans-heterozygotes with the
Y66H1B(tm545) null allele were not significantly different from wild-type (Kovacevic
and Cram 2010). These data suggests this deletion does not affect fertility. fln-1 is
essential to signaling to the somatic gonad tissue and potentially the germline that
ovulation is occurring as oocytes are passing through the spermatheca (Kovacevic,
Orozco, and Cram 2013). While the Y66H1B.2(ok2611) is likely not null and does not
53 appear to have an affect on ovulation, it could be that we would not see an affect with this
allele on the akir-1 background. This could be tested by quantifying the F1 progeny of
the double mutants using Y66H1B.2(ok2611) and akir-1(gk528) as controls. However,
the fln-1(RNAi) knockdown is likely to affect ovulation. Mutations that result in defective
ovulation lead to an endomitotic oocyte (EMO) (Iwasaki et al., 1996). This could be why
we see an enhancement of the embryonic lethality in the akir-1 background. This could
be tested by examining the akir-1(gk528);fln-1(RNAi) mutants for the presence of the
EMO. While the bivalents in the akir-1(gk528);fln-1(RNAi) mutants appeared wild-type,
their length and structure were not examined or quantified. This should be done with the
results being compared to akir-1 mutants to test for significant differences. Alternatively,
the Y66H1B(tm545) allele has been demonstrated to be null and could be used to generate
the Y66H1B(tm545);akir-1(gk528) double mutant and evaluated for SC and bivalent
defects.
C09G4.3
Cks1 was identified as a Cdc28 (also known as CDK-1) binding partner and is
involved in cell cycle regulation in S. cerevisiae (Hadwiger et al., 1989) and is conserved
(Krishnan, Nair, and Pillai 2010). Previously, cks-1(RNAi) has been shown to inhibit the
meiotic divisions, specifically the first meiotic division (Polinko and Strome 2000).
These authors also noted, that in MI chromosomes were not condensed, they appeared
“ragged” when compared to wild-type and failed to decondense and enter interphase
(Polinko and Strome 2000).
The bivalent defects observed in the cks-1(RNAi) oocytes in diakinesis are
reminiscent of those caused by defects in DNA damage/repair mutations such as rad-51
nulls and mre-11(iow1) where instead of six well structured bivalents, there were
clumps or aggregations of chromosomes found in the D-1 oocyte (Martin et al., 2005;
Yin and Smolikove 2013). As the primary target of Cks1 is Cdk1, it is likely that a
depletion of CKS-1 leads to inactivation of CDK-1 in C. elegans. Cdc28/Cdk1 depletion
54 has been shown to mitotic catastrophe leading to a lack of chromosome stability and
uneven chromosome condensation (Kitazono and Kron 2002; Blank et al., 2006).
Furthermore, it has been shown in vitro that CDK mediates recruitment of condensin I
(Kimura et al., 1998). Given the roles the condensins have in preparing chromosomes for
meiotic divisions(Chan, Severson, and Meyer 2004a; Csankovszki et al., 2009), the
bivalent defect we observe in both akir-1(gk528);cks-1(RNAi) and csk-1(RNAi) are could
be due to two factors. The first would be a lack of activation of the DNA checkpoint due
to the lack of an active CKS-1/CDK-1 complex. The second would be the lack of
chromosome condensation as a result of uneven chromosome condensation due to
recruitment of only condensin I, but not condensin II to the meiotic chromosomes.
Cdk1 has also been implicated in the regulation of Aurora B kinase, Ipl1, by
direct phosphorylation in yeast (Zimniak et al., 2012). This directly implicates Cdk1 in
SC disassembly, as Ipl1 is known to coordinate SC disassembly and crossover formation
in yeast (Jordan et al., 2009). In C. elegans, AIR-2 (the Ipl1 homolog) localizes to the
bivalents as they mature in late diakinesis (Kaitna et al., 2002; Rogers et al., 2002). AIR2 facilitates the removal of cohesion in MI and MII divisions. This raises the question
whether CDK-1 is acting directly on AIR-2 to remove cohesion or if CDK-1/CKS-1
interacts with AKIR-1, to regulate SC disassembly in C. elegans.
These data indicate a role of cks-1 in prophase I events in C. elegans, including
DNA damage repair, chromosome condensation, and SC disassembly. As no SYP-1
aggregates are observed, CKS-1’s role is either upstream in preventing the unloading of
SYP-1 or in the turnover of unloaded SYP-1 by the proteasome pathway. The deletion
mutant cks-1(tm1916) should be obtained and examined to see if these observations can
be replicated, however, this mutant is listed as having a lethal/sterile phenotype. It maybe
that we are able to observe these SC and bivalent defects only because the RNAi
generates a hypomorphic situation where there is partial, not complete knockdown of
CKS-1. Given our observations and the literature support for function, cks-1 appears to
55 have a function in the akir-1 background and should be further examined to determine the
extent of that interaction.
Evaluation of the screen design
This RNAi screen was designed to isolate genes involved in SC disassembly via
the enhancement and suppression of the akir-1 mutant phenotypes; SYP-1
aggregation/delayed disassembly and bivalent structure defects in diakinesis. However,
as the screen was implemented prior to the completion of mapping, identification and
characterization of the akir-1(rj1) allele, the assumption was made that the gene in
question was meiotic in nature and evaluation of larval progeny would be sufficient to
determine changes in embryonic lethality on the akir-1 background. However, this is not
the case, as akir-1 has since been shown to have developmental defects as well (S.
Smolikove, N. Balukoff, and R. Bowman personal communication). This leads to a
problem in discerning changes in lethality due to meiotic defects versus developmental
defects. In the preliminary and secondary screens, we used viable F1 progeny as a means
to detect changes in lethality. If AKIR-1 had solely functioned in meiosis, this would
have been a feasible assay for meiotic interactions. However, with the revelation of a
mitotic function, this complicated the screen as we could no longer link the reduction in
viable progeny to only meiosis, but to developmental defects as well. Taken together, we
were detecting three different classes of enhancement of the akir-1 lethality phenotype:
one class due to only meiotic interactions, a second class due to only developmental
interactions, and a third class due to both meiotic and developmental interactions
combined. This meant we could no longer use the variability of F1 progeny to determine
enhancement or suppression of lethality as an assay in the screen. This likely explains
why, we had such a low hit rate when we examined the chromosome II clones
cytologically.
A solution would be to conduct fecundity studies to evaluate clones to determine
which defects are present. A reduction in the number of eggs laid would represent a
56 meiotic defect while a reduction of viable larval progeny could indicate a developmental
defect. Increased embryonic lethality could be attributed both to meiotic and
developmental defects. Though this approach is time consuming, it would require high
reagent costs. A better approach would be to use GO Terms as done in the chromosome
IV screen, to narrow the number of clones to be examined to a manageable number. Of
the 25 clones selected by this method, csn-5 and cks-1 demonstrated alterations in
embryonic lethality in the akir-1 and wild-type backgrounds that could be directly
connected to aberrant meiotic prophase I events. Thus far, only 37% of the genome has
been screened for akir-1 interactors and is possible that more exist. Completion of this
screen should involve a candidate approach either in the initial selection of clones for the
preliminary screen or to identify clones for cytological analyses after the preliminary
screen.
57 Figure 2-1: A schematic representation of the RNAi enhancer/suppressor screen.
A) The basic methodology of the screen. Description is provided in text. B) The expected
outcomes of the screen hits on the wild-type and akir-1 backgrounds. Clones with no
affect or are embryonic lethal in both backgrounds will show no interaction. Enhancers or
suppressors will only show an affect on the akir-1 mutant background.
58 Figure 2-2: Bivalent defects in unc-85(RNAi) and quantification of defects. A-D)
Micrographs of DAPI stained oocytes in D-1 of indicated genotypes. Wild-type, akir1(rj1) and unc-85(RNAi) all have wild-type bivalent structure. akir-1(rj1);unc-85 has
univalent present in diakinesis oocytes. E) the average number of univalents in D-1
oocytes in wild-type and unc-85(ok2125).
59 Figure 2-3: Bivalent defects in unc-85(RNAi) and unc-85(ok2125).
A-C’’’) Micrographs of SYP-1(red) and DAPI(blue and gray scale) D-3 and D-1 oocytes
of indicated genotypes. unc-85(ok2125) has wild-type bivalent structure. unc85(ok2125);akir-1gk(528) double mutant did not have univalent. Images are projects
through a three- dimensional data stack. Scale bar 2µm.
60 Figure 2-4: Bivalent defects in ZC239.6 mutants. A-C’’’) Micrographs of SYP-1 (red)
and DAPI(blue) stained oocytes in D-1 of indicated genotypes. Akir1(rj1);ZC239.6(RNAi) lacked bivalent structure. This defect was no replicable. Images
are projects through a three-dimensional data stack. Scale bar 2µm.
61 Figure 2-5: The lack of SC and bivalent defects in let-29(RNAi). A-H) Micrographs of
SYP-1 (red) and DAPI (blue) stained oocytes in D-1 of indicated genotypes. let-92(RNAi)
had no defects. Images are projects through a three-dimensional data stack. Scale bar
2µm.
62 Figure 2-6: Y66H1B.2(RNAi) and Y66H1B.2(ok2611) mutants. A-J) Micrographs of
SYP-1 (red) and DAPI(blue) stained oocytes in D-1 of indicated genotypes.
Y66H1B.2(RNAi) and Y66H1B.2(ok2611) mutants have no defects. Images are projects
through a three-dimensional data stack. Scale bar 2µm.
63 Figure 2-7: SC and bivalent defects in C09G4.3(RNAi) mutants. A-I) Micrographs of
SYP-1 (red) and DAPI (blue) stained oocytes in D-1 of indicated genotypes.
C09G4.3(RNAi) mutants have bivalent and SC defects. Images are projects through a
three-dimensional data stack. Scale bar 2µm.
64 Table 2-1: Potential Candidate genes from RNAi screen
Gene
Name
Seq ID
Function
Chromosome
Akir-1/wt
ratio of F1
progeny*
Screen
Phenotype
unc-85
F10G7.3
II
0.5
Enhancer
Novel
cks-1
let-92
ZK239.6
C09G4.3
F38H4.9
Chromatin
remodeler
unknown
CDK subunit
PP2A catalytic
subunit
II
IV
IV
0.88
0.43
1.63
Enhancer
Enhancer
Suppressor
fln-1
Y66h1B.2
IV
0.19
Enhancer
csn-5
B0547.1
Filamin A
CSN/COP9
IV
0.09
Enhancer
signalosome
subunit
* See Chapter 2 text for description of ratio and Chapter 4 for calculation method
65 CHAPTER 3
A NOVEL ROLE FOR THE CSN/COP9 SIGNALOSOME IN SC ASSEMBLY
Introduction
The following data has incorporated into a manuscript, submitted and is currently
under peer review. Additional authors include Martha Dean, Nathan Balukoff and Sarit
Smolikove from the Department of Biology at the University of Iowa. I was involved in
all aspects of experimental design for this study and carried out data collection with the
assistance of an undergraduate student, Martha Dean (PCR validation of crosses,
performed crosses, and IHC preparations). Our lab research assistant, Nathan Balukoff
assisted with western analyses, preparation and data collection for Figure 3-7, 3-8, 3-17,
and Table 3-7. Sarit Smolikove outcrossed the csn mutant strains and made Figure 3-7.
Accurate segregation of meiotic chromosomes is essential to the formation of
gametes. We identify the CSN/COP9 signalosome as having a role in chromosome
synapsis. In CSN/COP9 signalosome mutants, SC proteins aggregate and fail to assemble
properly on homologs, leading to defects in pairing, repair of meiotic recombination
intermediates and crossover formation. We also discovered defects in MAPK signaling in
CSN/COP9 mutants, which leads to a severe reduction in oocyte maturation.
CSN/COP9 Signalosome
The CSN/COP9 signalosome is a highly conserved protein complex originally
described in Aradidopsis as a repressor of photomorphogenesis (Chamovitz and Segal
2001). The complex is comprised of eight subunits which are remarkably similar to the
lid complex of the 26S proteasome and the complex is involved in protein modification
(Figure 3-1) (Bech-Otschir, Seeger, and Dubiel 2002; Wei and Deng 2003). Seven
CSN/COP9 signalosome subunits have been identified in C. elegans. Five subunits
(CSN-1,2,3,4, and 7) contain a PCI (proteasome, COP9 signalosome, initiation factor 3)
domain and two (CSN-5 and CSN-6) contain MPN (Mpr1-Pad1-N-terminal) domains
66 (Wei and Deng 2003). The PCI domains are thought to facilitate protein-protein
interactions and may also have nucleic acid binding properties (Kim et al., 2001). The
CSN-5 MPN domain contains a JAMM (Jab1/MPN/Mov34) motif, which includes the
metalloisopeptidase catalytic activity, which can cleave ubiquitin and ubiquitin-like posttranslational modifiers (such as NED-8/Rub1) (Figure 3-1) (Cope et al., 2002; Gusmaroli
et al., 2007; Wolf, Zhou, and Wee 2003; Merlet et al., 2009). The CSN-6 MPN domain
lacks the JAMM motif and thus the metalloisopeptidase activity (Peng, Serino, and Deng
2001; Pick et al., 2012). The signalosome is involved in the regulation of protein function
via multiple pathways, but most studies have been carried out in the context of the
ubiquitin pathway via the CULLIN-RING E3 ubiquitin ligases (CRLs) (Figure 3-1)
(Bech-Otschir, Seeger, and Dubiel 2002; Wolf, Zhou, and Wee 2003; Choo et al., 2011).
CULLIN RING E3 ligases (CRLs) are the most well studied of the CSN/COP9
substrates and have known roles in mediating the ubiquitin/proteasome pathway and
promoting localization and/or protein interactions through post-translational
monoubiquitination (Petroski and Deshaies 2005; Chiba and Tanaka 2004). The current
literature supports a model where the CSN/COP9 signalosome destabilizes the CRL
complexes by deneddylation which inhibits CRL activity (Wei, Serino, and Deng 2008;
Stratmann and Gusmaroli 2012). However, null signalosome mutants do not show
stabilization of CRL (Wei, Serino, and Deng 2008; Stratmann and Gusmaroli 2012). This
suggests that successive cycles of neddylation and deneddylation are required for CRL
function. If the CSN/COP9 signalosome is rendered non-functional by mutation, for
example as in the csn mutants, the CRLs will also be rendered non-functional. The
CSN/COP9 signalosome regulates cell cycle, gene expression, and DNA damage repair,
through mechanisms that do not necessarily involve CRLs (Wei, Serino, and Deng 2008;
Tian et al., 2010; Stratmann and Gusmaroli 2012). In Drosophila, the CSN/COP9
signalosome has been implicated in the disruption of oocyte axis formation via regulation
of gurken proteins (Doronkin, Djagaeva, and Beckendorf 2002).
67 Null mutants of the CSN/COP9 signalosome generated in other model organisms
(yeast and Drosophila) have shown that the loss of one subunit renders it inactive and
leads to lethality (Mundt, Liu, and Carr 2002; Stuttmann, Parker, and Noël 2009;
Doronkin, Djagaeva, and Beckendorf 2003; Kotiguda et al., 2012). CSN-5 (also known
as Jab1) is the only subunit in those organisms which has been shown to act outside the
holocomplex in such cellular activities as nuclear export, degradation, and protein
stabilization (Kotiguda et al., 2012; Yoshida et al., 2010). These activities fall into two
major cellular functions, cellular proliferation and apoptosis (Shackleford and Claret
2010). The CSN-5 subunit of CSN/COP9 signalosome in C. elegans has also been
implicated in muscle development (R. K. Miller et al., 2009), and the regulation of
germline P-granule component, GLH-1, through interactions with KGB-1, a member of
the JNK kinase family (Smith et al., 2002; Orsborn et al., 2007). While CSN-5 RNAi has
been shown to reduce the size of gonads in C. elegans (Smith et al., 2002; Orsborn et al.,
2007), a role for CSN-5 in the germline has not been molecularly defined.
The work described here indicates a novel role in C. elegans for the CSN/COP9
signalosome in meiotic prophase I events that are critical for the formation of functional
gametes. Mutations in signalosome components specifically affect SC assembly and
oocyte maturation. In csn mutants SYP-1 formed aggregates (polycomplex-like
structures) and they persisted throughout meiotic prophase I. Additionally, we observed
reduced chromosomal pairing throughout meiotic prophase as well as disruption in
meiotic recombination and crossover formation. We also found an increase in apoptosis,
likely due to the disruption of events earlier in pachytene. Oocyte maturation also is
disrupted, leading to a severe reduction in the number of oocytes in diakinesis which
renders the worms sterile. Our working model is that the CSN/COP9 complex regulates
SC morphogenesis by inhibiting SYP DNA-independent self-assembly. Without
CSN/COP9 function SC morphogenesis is perturbed (leading to SYP aggregate
formation) as are downstream events (e.g., pairing and recombination), which are
68 dependent on proper SC formation. We also present data indicating, the CSN
signalosome influences oocyte maturation by permitting meiotic progression via
MAPK/MPK-1 activation.
Results
csn mutants exhibit defects in SC morphogenesis and meiotic progression
We have identified csn-5 as a gene involved in SC morphogenesis via an RNAi
suppressor/enhancer screen of a mutant (akir-1) exhibiting aberrant SC aggregation.
Previous studies utilizing RNAi methodology to examine the role of the CSN complex
genes in C. elegans demonstrated that csn-5 was required for normal gonad morphology.
csn-5(RNAi) resulted in formation of short gonads and down-regulation of the P-granule
component GLH-1 (Smith et al., 2002; Orsborn et al., 2007). However, SC
morphogenesis, chromosome dynamics, and meiotic recombination in meiotic prophase I
were not examined in these studies. Here we focused our studies on the function of the
CSN/COP9 signalosome in these meiotic processes.
Two deletion alleles were analyzed in this study: csn-2(tm2823) and csn5(ok1064). The csn-2(tm2823) allele is missing most of exon 3 which results in deletion
of 28% of the coding region and the expected introduction of deletion of the proteinprotein interaction domain. The csn-5(ok1064) allele is missing exons 1, 2, and 3 which
results in deletion of 64% of the coding region (including the MPN catalytic domain)
and creates a frame shift (Figure 3-2A). We conclude that both csn alleles are likely to be
null. The csn-2(tm2823) and csn-5(ok1064) alleles will be referred here collectively as
csn mutants.
In wild-type nuclei, SC initiates assembly at the transition zone
(leptotene/zygotene) when SC proteins load on chromosomes (Figure 3-2C); the SC is
fully assembled at early pachytene. SC disassembly is initiated at the end of pachytene
and CR disassembly is complete by the end of diakinesis. To determine if SC
morphogenesis was affected in csn mutants, we performed immunohistochemical
69 analyses using antibodies against SYP-1, SYP-4, HIM-3, and HTP-3 (Zetka, Kawasaki,
and Strome 1999; Goodyer et al., 2008; Smolikov, Schild-Prüfert, and Colaiácovo 2009;
Clemons et al., 2013) (Figures 3-2 through 3-6). In both csn mutants, we observed
smaller, morphologically different gonads compared to wild-type (Figure 3-3), as
previously published for csn-5(RNAi) experiments (Smith et al., 2002; Orsborn et al.,
2007). The nuclei in the csn mutant gonads were unevenly spaced throughout the gonad.
There also appeared to be no distinct rachis (central canal) as in wild-type worms. The
chromosomes of csn mutants clustered to one side of the nucleus (a polarized
organization) as found in the wild-type transition zone (leptotene/zygotene) nuclei which
are indicative of meiotic entry (Zetka 2009). Unlike wild-type, some polarized nuclei
were found throughout the gonad in the csn mutants intermixed with nuclei with a more
normal dispersed chromosomal organization. The persistence of polarized chromosomes
has been observed previously in synapsis defective mutants such as syp-1(me17)
(Smolikov, Eizinger, Hurlburt, et al., 2007). In addition to the persistent polarized
chromosome organization, we also determined the mitotic/meiotic boundary using
antibodies for SC proteins, HIM-3 and HTP-3, since these LE proteins localize to
chromosomes axes upon the transition from mitosis to meiosis. This localization occurred
concurrently with polarization of chromosomal organization and did not show any
defects in the germline of csn mutants. These data indicate: 1) the transition from mitosis
to meiosis took place in the csn mutants and 2) the localization of lateral element proteins
of the SC was not perturbed in csn mutants (Figures 3-5 and 3-5). Thus, although gonads
are smaller in csn mutants and have fewer nuclei, meiotic entry has occurred and SC
assembly has initiated.
In contrast to the pattern of localization of lateral element proteins in the csn
mutants, the CR protein SYP-1 showed an aberrant pattern of localization. SYP-1 protein
aggregates (polycomplex-like structures) were found in the transition-like zone at the
distal end of the gonad and through the late-pachytene-like zone in the proximal end of
70 the gonad. These occurred in 100% of the mutant gonads examined (wild-type n= 26,
csn-2 n=37, csn-5 n=34; p<0.0005; Fishers Exact Test) for both csn mutants (Figure 3-2
B-F’’). CR/SYP aggregates were 4-fold wider than a typical SC (width of wild-type SC0.22µm±0.23 n=25, width of SC aggregate- csn-2 0.83µm±0.23 , n=70 and csn-5
0.86µm ±0.31, n=90 p<<0.001 Mann Whitney test) and typically there was one
aggregate per nucleus (csn-2 1.12, n=62 and csn-5 1.13, n=82). While some nuclei
contained a single SYP-1 aggregate with no additional SYP localization, most nuclei
contained partially assembled linear SC in addition to the aggregates. As there are
currently four SYP proteins, we examined the localization of SYP-4 to identify if the
aggregation defects were specific to SYP-1 or generally affect all the CR components
(Figure 3-6). SYP-4 also forms persistent aggregates suggesting the defects observed in
the csn mutants are not specific to SYP-1.
P-granules are germline RNA storage compartments that are composed of
mRNAs and proteins; these include the GLH-1 and PGL-1 proteins that are important for
P-granule function. A recent paper by Bilgir et al., 2013 described failure in SC assembly
in pgl-1mutants. Since GLH-1 is known to be regulated by CSN-5 (Orsborn et al., 2007),
SYP aggregation could potentially be induced by a reduction in function of GLH-1 (and
the consequent P-granule defects). To test this hypothesis we examined SYP-1
localization in GLH-1 and KGB-1 mutants. KGB-1 binds GLH-1 and promotes its
degradation, while CSN-5 promotes GLH-1 stabilization by competing with KGB-1 for
binding to GLH-1 (Smith et al., 2002; Orsborn et al., 2007). If CSN-5 influenced SC
through its role in P-granule function, then glh-1 mutants should show identical
phenotypes (SC aggregation) to csn-5 mutants. kgb-1 mutants should show an opposing
phenotype to that of glh-1 mutants. It is hard to discern what is the opposite phenotype of
SC aggregation; we presume it is a form of destabilize CR proteins, which may lead to
lack of SC. We did not observe any changes in SC structure, including aggregation, in
71 kgb-1 and glh-1 mutants (Figure 3-7). We concluded from this, that P-granule
destabilization is likely not the cause of SYP aggregation in csn mutants.
csn-2 and csn-5 mutants similarly affect CR assembly
Having determined both csn mutants display SYP aggregation, we asked if the
defects in SC assembly are indistinguishable in both of our mutants. Not all nuclei within
the transition-like zone and pachytene-like zone had aggregates. The gonads were divided
into six zones sized as in Colaiacovo, 2003 (Colaiácovo et al., 2003), Figure 3-8A) and
were scored for the percent of nuclei with aggregates in each zone. Each zone represents
a size unit (36µMx36µM window) organized sequentially (zone 1 being the distal premeiotic tips (PMT) and zone 6 the proximal late pachytene region). This division into
zones was performed according to the standard protocol for quantitative analysis of early
to mid-meiotic events in the C. elegans germline, (e.g., RAD-51 analysis, also see
Chapter 4). While the csn mutant gonads were smaller, we saw all the hallmarks that
meiosis had initiated (polarized nuclei and lateral element immunostaining) and that
pachytene stage was initiated (the depolarization of most nuclei). Therefore, we divided
the mutant gonad into zones in the same manner as the wild-type. By keeping the size
unit the same, we got the best approximation of the zones in the mutants in order to
compare them to wild-type. We measured nuclei for the presence or absence of SYP-1
aggregates (Figure 3-8B-D and 3-9 wild-type: 0% nuclei with aggregates; n=98; csn-2:
65% nuclei with aggregates, n=57; csn-5: 58% with aggregates, n=50; p<0.0005 for
pairwise comparisons between wild-type and each csn mutant; Fishers Exact Test). The
mutants were not different from each other in the fraction of nuclei with aggregates
although both were significantly different from wild-type. Furthermore, there was no
difference in the size of the aggregates between the csn mutants or number of aggregates
per nucleus (p>>0.05 Mann Whitney test). The early appearance of SYP aggregates as
SC assembly initiates (zone 2-3) indicates that the primary defect observed in csn
72 mutants is in SC assembly. The increase in the percent of nuclei with aggregates from
Zone 3 to 5 may reflect additional defects in maintenance of the SC.
SYP-1 aggregation could result from over-expressing SYPs (Smolikov, SchildPrüfert, and Colaiácovo 2009). To address this point; we performed western analysis to
determine the level of SYP-1 in the csn mutants. Both csn mutants had a reduced level of
SYP-1 compared to wild-type (Figure 3-10A, csn-2 34% and csn-5 80% of wild-type).
We also utilized a cytology-based assay to quantify the amount of nuclear SYP-1 protein
in csn nuclei compared to wild-type. We observed a 2-fold decrease in nuclear SYP-1 in
the csn mutants compared to wild-type (Figure 3-10B). There was no difference between
the csn mutants (wild-type n =26, csn-2 n= 36 and csn-5 n= 44; p<0.05 for pairwise
comparisons between wild-type and each csn mutant; Mann Whitney test). These data
indicate that the SYP-1 aggregates are not the result of detectable over-expression of
SYP-1.
csn-2 and csn-5 are required for gonad proliferation and fertility
The overall length of the gonads of csn mutants is shorter than observed in wildtype (Table 3-1), which could result from reduced proliferation of mitotic cells. If mitotic
proliferation (prior to meiotic entry) was affected, the size of the mitotic zone (i.e., the
PMT) would be shorter in csn mutants and there would be few nuclei over all as is seen
in the csn mutants. As nuclei enter meiosis, they acquire a polarized configuration of
chromosomes indicating meiosis was initiated. We used this polarization to measure the
length of the PMT of gonads for each genotype. In the csn mutants, the PMT region was
60% and 77% shorter than in wild-type (Figure 3-11A n=10 for each strain p<0.0005 for
wild-type vs csn-2; p=0.005 for wild-type vs csn-5 and p<0.05 for csn-2 vs csn-5; Mann
Whitney Test). An interpretation of these data is that the transition from mitosis to
meiosis occurs earlier in these mutants compared to wild-type.
When nuclei move to diakinesis, the final stage of meiotic prophase I, wild-type
gonads contain an average of 8.1±1.1 (n=75 gonads) diakinesis nuclei. These diakinesis
73 nuclei are also referred to as oocytes, although the cellularization process occurs only
towards the end of diakinesis. Unlike wild-type, most gonads of csn homozygotes lacked
diakinesis nuclei/oocytes (Figure 3-11B p<0.0005 for each pairwise comparison with
wild type; Mann Whitney Test). In csn-2 mutants, only about 25% of the gonads (n= 125
gonads) had diakinesis nuclei, and average of 0.8±0.76 per gonad (Table 3-2). In csn-5
mutants, 65% of the gonads had diakinesis nuclei with an average of 1.23± 0.98 per
gonad (n=100 gonads). We performed an egg lay assay to determine the number of eggs
laid and their viability. For the csn mutants, no eggs were laid in a three-day period (csn2 n=11 and csn-5 n=13); in contrast wild-type worms had an average of 247±16 eggs laid
per worm (n=3, Table 3-3).
csn-2 and csn-5 are required for pairing stabilization
Pairing interactions between homologous chromosomes are initiated in a SCindependent manner at specific chromosomal sites (pairing centers). The term “pairing
stabilization” describes pairing interactions that spread outside the pairing centers and
lead to the persistence of homolog association throughout pachytene. In syp mutants, loci
distant from the pairing centers exhibit a very low level of pairing throughout meiosis
(Zetka 2009). Since the data indicated that csn mutants lack a fully functional SC; we
expected to find that pairing stabilization had been compromised in the csn mutants,
similar to syp mutants. To test this, we used a 5S ribosomal RNA locus near the central
region of chromosome V to analyze pairing interactions between homologous
chromosomes by fluorescence in situ hybridization (FISH). The gonads were divided into
six zones and were scored for the percent of nuclei with paired 5S loci in each zone
((MacQueen 2002), Figure 3-12 and 3-13, Tables 3-4 and 3-5).
In zone 1, as expected, wild-type and syp-1(me17) control nuclei, as well as the
csn mutants, showed little to no homologous pairing, fewer than 15% of 5S loci had
paired chromosomes. As the nuclei progress through meiotic prophase I, wild-type
chromosomes initiated pairing and maintained high levels of pairing through the
74 pachytene zones. In syp-1(me17) because there is no SC formed mutant chromosomes
remain unpaired throughout the germline (Figures 3-12 and 3-13). csn-2 mutants pairing
levels never exceeded 20% of the 5S loci paired in any zone, indicating the majority of
the chromosomes were unpaired. Overall, csn-2 and syp-1(me17) mutants exhibit similar
pairing defects throughout meiotic prophase I (Figure 3-12 and 3-13). In contrast to csn-2
mutants, csn-5 mutants initiated pairing similarly to wild-type. Since the transition from
mitosis to meiosis is occurring earlier in csn-5 mutants, pairing initiates at zone 2,
compared to zone 3 in wild-type (Figure 3-12). In zone 4, csn-5 mutants showed a
reduction in the percent of nuclei with paired chromosomes, but the levels were
intermediate between the ones observed in wild-type and syp-1 or csn-2 mutants. The
percent of nuclei with paired chromosomes for csn-5 mutant remained higher than csn-2
mutant for zones 5 and 6. These and other differences between the two csn mutants are
discussed below (Discussion section). When taken together these findings are consistent
with a view where defects in SC assembly perturb pairing stabilization. These data also
indicate that SYP assembled in a linear manner on chromosomes in csn mutants (Figure
3-13 B-D blue) does not support pairing stabilization and these SC-like structures likely
represent non-homologous synapsis and/or SYP assembly between sisters.
Meiotic recombination and crossover formation are perturbed in csn mutants
In mutants that do not form the SC, events downstream of pairing and SC
assembly such as meiotic recombination are perturbed (Zetka 2009). We expected the csn
mutations would have a similar effect on recombination. RAD-51 is a strand exchange
protein used as an indirect marker for DSB formation and subsequent repair in C. elegans
(Colaiácovo et al., 2003). The gonads were divided into zones as previously described
and the numbers of RAD-51 foci per nucleus were counted.
Mitotic nuclei in zones 1 and 2 showed very low levels of RAD-51 both in wildtype and csn-2 mutants. csn-5 mutants exhibit slightly increased levels of RAD-51 foci in
mitotic nuclei (p<0.01 Mann Whitney Test). RAD-51 foci levels increased at the
75 entrance to meiosis in all genotypes tested, as expected from the induction of meiotic
DSBs (Figure 3-14 zone 1: wild-type n=159, csn-2 n= 60, csn-5 n=46; Table 3-6
p<0.0005 wild-type vs csn-5 and csn-2 vs csn-5; Mann Whitney Test). The increase in
RAD-51 foci/nucleus occurred earlier in csn mutants, likely due to the fact meiotic entry
occurred earlier (Figure 3-14; zone 2: wild-type n=209, csn-2 n= 70, csn-5 n= 49; Table
3-6 p<0.0005 wild-type vs csn-5 and p<0.005 csn-2 vs csn-5; Mann Whitney Test).
Despite the similarity of RAD-51 localization patterns in the distal part of the germline,
the overall levels of RAD-51 foci in early prophase were approximately 2-fold increased
in csn mutants compared to that of wild-type (Figure 3-14 zone 3: wild-type n=199, csn-2
n=61, csn-5 n=62; p<0.0005 for wild-type vs csn mutants; Mann Whitney Test; zone 4:
wild-type n=177, csn-2 n= 56, csn-5 n=47; p<0.0005 wild-type vs csn mutants; zone 5:
wild-type n=174, csn-2 n= 69, csn-5 n=55; p<0.0005 wild-type vs csn mutants; Mann
Whitney Test). The levels of RAD-51 foci in the csn mutants remain higher than wildtype in zones 4-6, indicating the repair of DSBs was affected (Figure 3-14 and 3-15, e.g.,
zone 6: wild-type n= 142, csn-2 n= 47, csn-5 n= 49; p<0.0005 wild-type vs csn-5 and
csn-2 vs csn-5; p<0.05 wild-type vs csn-2; Mann Whitney Test). In late pachytene (zone
6), we observed a difference between csn-2 and csn-5 mutants, with csn-5 mutants
maintained RAD-51 foci (zone 6) at high levels, while they decreased in csn-2 mutants.
In C. elegans, one obligatory crossover is observed per chromosome pair
(Colaiácovo et al., 2003), that results in six crossovers per nucleus. COSA-1, a conserved
cyclin related protein, that localizes to crossovers and can be used to monitor the number
of crossovers per nucleus (Figure 3-15 and 3-16). A reduction in COSA-1 foci indicates a
defect in crossover formation. We tested the csn mutants to determine if crossover
formation was affected using a GFP-tagged COSA-1, crossed into each mutant strain.
The gonads were divided into zones, as previously indicated. Wild-type nuclei (n=88) in
zone 6 had an average of 5.9 (± 0.03) bright COSA-1 foci, indicating crossovers had been
properly formed (Figure 3-15, 8% of nuclei with less than 6 foci). However, in the csn-2
76 mutant, 95% of the nuclei (n=111) in zone 6 had less than six foci (Figure 3-14 1±0.16
foci per nucleus, p<0.0005; Mann Whitney Test). In the csn-5 mutant, we observed a
wider distribution of the number of COSA-1 foci observed, with 82% of the nuclei with
less than six foci (an average of 2.8 ±0.21 foci per nucleus, p<0.0005; Mann Whitney
Test). The average numbers of COSA-1 foci was significantly different between the two
csn mutants (p<0.0005; Mann Whitney Test). These data suggest a role for CSN/COP9 in
crossover formation.
Apoptosis is increased in csn mutants and dpMPK-1 levels are reduced
Lack of synapsis (Bhalla 2005) or an accumulation of DSBs (Colaiácovo et al.,
2003) results in increased apoptosis at the late pachytene due to the activation of synapsis
and pachytene checkpoints (Zetka 2009). CED-1, is expressed during the process of
engulfment; a mechanism of clearing apoptotic corpses from the germline in late
pachytene. Thus, ced-1::GFP, exclusively surrounds apoptotic nuclei and is frequently
used as a marker to detect apoptosis.
Both csn mutants had lower average numbers of nuclei in late pachytene (Figure
3-17 wild-type average = 52.2, n=25 gonads; csn-2 = 26.7, n=19; and csn-5= 30.9, n=18;
p< 0.0005 for comparison of wild-type and csn mutants; Mann Whitney Test). However,
only csn-2 had a significant increase in apoptosis (4-fold) while csn-5 had apoptotic
levels similar to wild-type (Figure 3-17 wild-type average =2.96, n=25 gonads ; csn-2 =
8.2, n=19; and csn-5= 3.3, n=18; p< 0.0005 comparison of wild-type vs csn-2 mutant and
csn-2 vs csn-5 mutants; Mann Whitney Test). When normalized for the number of nuclei
in late pachytene, both mutants showed increased apoptosis, but csn-2 mutants showed a
larger increase.
There are two apoptotic checkpoints in C. elegans meiosis that are activated by
unsynapsed chromosomes: the synapsis checkpoint mediated by PCH-2 (Bhalla 2005)
and the meiotic recombination checkpoint mediated by CEP-1 (Rutkowski et al., 2011).
We investigated whether removing these two genes could bypass the DNA
77 damage/synapsis checkpoint leading to apoptosis in the csn mutants. pch-2(tm1458);csn2(tm2823) and pch-2(tm1458);csn-5(ok1064) double mutants were generated and cep1(RNAi) was performed in the csn mutants. Overall gonad length, number of oocytes in
diakinesis, and the number of nuclei containing aggregates were measured. If increased
apoptosis in late pachytene was the reason for the severe reductions in oocyte numbers,
then bypassing the checkpoint function should increase the numbers of gonad nuclei:
nuclei that were destined to die and be engulfed by the sheet cells, do not get removed
from the germline. This increase in nuclei numbers should increase the overall length of
the gonad. We observed no change in overall gonad length (Table 3-1) between the single
csn mutants and the corresponding double mutants, nor any increase in oocytes in
diakinesis in young adults (one day post-L4, Table 3-2). Since clearing apoptotic corpses
may take time, we scored the same genotypes two days later, giving the opportunity for
accumulation of cells (in double mutants) otherwise destined for apoptosis (csn single
mutants). In csn-5 mutants overall gonad length was decreased with age, possibly due to
the defects in mitotic proliferation. However, both csn-5; pch-2 and csn-5; cep-1 mutants
showed increased gonadal size (double the size) compared to csn-5 single mutants ( three
days post-L4, Table 3-1). These data indicate that both the synapsis and the DNA
damage checkpoint are activated in csn-5 mutants.
In csn mutants, two types of meiotic nuclei are found: nuclei with partially
assembled SC and nuclei with one SYP aggregate. We measured the percent of nuclei
with aggregates in double mutants with perturbed apoptotic machinery compared to
single mutants. csn-5(ok1064);pch-2(tm1458), csn-2(tm2823);cep-1(RNAi), and csn5(ok1064);cep-1(RNAi) double mutants exhibited a decrease (2-fold) in the number of
nuclei with SYP-1 aggregates in late pachytene compared to the respective single csn
mutant (Figure 3-18 p<0.0005 comparison of wild-type vs csn mutant; Fishers Exact
Test, see Table 3-7). Therefore, nuclei with partially assembled SC /non-aggregated were
more frequently found in the double mutants and these were the nuclei more likely to be
78 lost by apoptosis. These data indicate partially assembled SC/non-aggregated nuclei
activate the synapsis checkpoint in csn-5 mutants and are more likely to be eliminated by
apoptosis.
Given the known physical interaction between CSN-5 and MPK-1, a MAPK
signaling protein (Li 2004; Zhong and Sternberg 2006), and the substantial pachytene
arrest (defects in progression from pachytene to diplotene) observed in csn mutants, we
examined whether MAPK signaling was disrupted in csn mutants. Phosphorylated MPK1 (dpMPK-1), the active form of MPK-1, is found in two distinct regions of the germline:
late pachytene and late diakinesis. The increase of dpMPK-1 in pachytene serves as a
signal for pachytene progression. The diakinesis dpMPK-1 is required for maturation of
oocytes (Di Agostino et al., 2002; Sasagawa et al., 2007). Nuclei of mpk-1 null mutants
completely arrest at mid-pachytene and no oocytes are observed (M.-H. Lee et al., 2007).
However, when only MPK-1 phosphorylation is eliminated (let-60 mutants) limited
pachytene arrest occurs and oocyte numbers are severally reduced (Church, Guan, and
Lambie 1995). Using an antibody for phosphorylated MPK-1, we stained wild-type and
csn mutant gonads. None of the csn-2 mutants examined had dpMPK-1 staining in late
pachytene (n=32) while only 3% of the csn-5 mutants (n=27) had dpMPK-1 staining. In
contrast, 82% of wild-type (n=75) and 75% of syp-1(me17) (n=32) gonads had dpMPK-1
staining (Figure 3-19A p<0.0005 wild-type vs csn mutants; Fishers Exact Test). MPK-1
has two isoforms in C. elegans, MPK-1A (43.1 kD) that is mostly somatic and MPK-1B
(50.6 kD) is germline specific (Rutkowski et al., 2011; M.-H. Lee et al., 2007). Western
analyses were conducted and the intensity of the bands was quantified and normalized to
the tubulin controls (Figure 3-19B). In wild-type, both isoforms were detected with
MPK-1A having an average normalized intensity of 2.27. The MPK-1A band was
detected in both csn mutants (csn-2= 0.99 and csn-5=0.96 normalized intensities),
although it was 2-fold lower in both mutants. The germline MPK-1B had an intensity of
9.91, but was not detected in either csn mutant (Figure 3-19B). These data indicate that
79 csn mutants lead to reduced MAPK/MPK-1 signaling which almost completely blocks
pachytene exit and severely reduces oocyte numbers. These data are consistent with the
observation that removing apoptosis checkpoints (pch-2 or cep-1) could not increase
oocytes number in csn mutants: even if more nuclei survived apoptosis, they could still
not exit pachytene arrest in the absence of dpMPK-1.
Taken together, these data indicate the CSN/COP9 signalosome has multiple roles
in meiosis: the signalosome affects the number of germline nuclei, SC assembly and
stabilization, recombination, MAPK signaling and the promotion of pachytene exit.
Discussion
CSN/COP9 is required for meiotic prophase I events
The CSN/COP9 complex has diverse and well-documented somatic functions, yet
the understanding of its role in meiosis is limited (Wei and Deng 2003). Studies of the C.
elegans and D. melanogaster CSN/COP9 indicate it plays a critical role in the regulation
of Vasa/P-granule proteins in the germline (Doronkin, Djagaeva, and Beckendorf 2003;
Smith et al., 2002; Orsborn et al., 2007). Here, we have shown CSN/COP9 has a
previously unknown meiotic function: it is essential for proper SC assembly, independent
of its P-granule role in the germline. We demonstrated that events following SC assembly
(e.g., stabilization of homolog pairing interactions and the repair of meiotic DSBs) are
perturbed as well. Both csn mutants show similar, but not identical effects on these
processes. In the absence of CSN/COP9, the SYPs (CR proteins) aggregate. In C.
elegans, stabilization of pairing interactions is absolutely dependent on SC formation and
independent of DSB formation and repair (MacQueen 2002). Therefore, it is reasonable
to propose that the pairing defects observed in CSN/COP9 mutants stem from defects in
SC formation. The limited amount of SC that assembles on chromosomes in csn mutants
cannot support the stabilization of pairing interactions (FISH analysis). In the absence of
stabilized pairing interactions, unresolved recombination intermediates (marked by RAD51) accumulate, presumably because homologs do not pair. This leads to a severe
80 reduction in crossover formation (marked by COSA-1 foci) in csn mutants and an
elevation of apoptosis. The magnitude of these phenotypes in csn-2 mutants closely
resembles that of syp null mutants, supporting our model that the later meiotic defects
(pairing and recombination) stem from inability to form a functional SC. The aberrant SC
and the meiotic recombination defects that follow both contribute to apoptosis in the csn
mutants. The decrease in the fraction of nuclei with aggregated SYP-1 when apoptosis is
bypassed in csn mutants indicates non-aggregated nuclei are preferentially selected for
elimination, perhaps as a result of physiological apoptosis. Alternatively, the nonaggregated may accumulate a higher level of recombination intermediates. We did not
immunostain for both aggregation and RAD-51, but our data show that there is an over
all increase in average number of RAD-51 foci per nucleus. Both synapsis checkpoint
and DNA damage checkpoint contributes to the elimination of these nuclei with nonaggregated forms of SC.
CSN/COP9 is required for normal levels of germline proliferation
Consistent with previous studies in C. elegans utilizing RNAi (Smith et al., 2002;
Orsborn et al., 2007), the two csn mutants have a reduced gonad size. We suggest this
reduction is due to a proliferation defect, as the number of mitotically dividing nuclei in
the pre-meiotic tip is reduced in the csn mutants and therefore there are overall fewer
nuclei in the gonad as seen in this study and others (Smith et al., 2002; Orsborn et al.,
2007). Proliferation defect is defined as any hindering to the proper progression of the
cell cycle (longer mitotic stages, arrest), as opposed to a reduction in nuclei number due
to increased apoptosis. Genes involved in apoptosis are not expressed in the PMT
(Gartner et al., 2000). However, reduction in nuclei is often referred to as proliferation
defects (Gartner et al., 2000). Although, proliferation defects were not specifically
examined, there is support in the literature for such a role which is discussed here. Given
the observations made in this thesis, the future studies of the signalosome’s role in
prophase I will include examination of the PMT for proliferation defects. While we know
81 that the csn mutants affect proliferation, we do not know if that affect is a direct
interaction of the CSN/COP9 signalosome on proliferation or an indirect interaction
through the ubiquitin/proteasome pathway (involving the CRLs).
There are two mechanisms that govern proliferation in the PMT zone: GLP-1/
Notch signaling and CYE-1/CDK-2 pathway. In the first mechanism, signaling has a key
role in maintaining the PMT by preventing the entry into meiosis. In the second
mechanism, the cell cycle has a critical role in maintaining a stem cell population in the
PMT through mitotic division. The cell cycle involving CYE-1/CDK-2 acts
independently of the GLP-1 pathway, but data suggest these are redundant pathways for
ensuring proper proliferation in the PMT. Regulation of these two mechanisms is
essential to maintaining the proliferative and regulating the entrance into meiosis.
GLP-1, a Notch receptor, is essential to mitotic proliferation in the PMT
(Crittenden et al., 2003). GLP-1 initiates a signaling cascade in the distal tip cell (DTC)
that specifies cell fate in the early germline and represses the entry into meiosis; nulls
result in a germline tumor [uncontrolled mitotic proliferation without meiotic entry
(Pepper et al., 2003)]. In order for mitotic proliferation to take place, GLP-1/Notch
signaling levels must remain high although they do decrease at the boundary of PMT and
TZ zones as meiosis is initiated (Crittenden et al., 2003; Pepper et al., 2003).
Bioinformatic predictions of interactions between GLP-1, CSN-2 and CUL-3 could
provide a mechanism for how csn mutants demonstrate a reduction in germline
proliferation (Zhong and Sternberg 2006). CRLs act in the ubiquitin/proteasome pathway
or by translational modifications (Figure 3-1), therefore CUL-3 could potentially
modulate the expression of GLP-1 or suppress Notch signaling, reducing proliferation.
HannB et al., 2011 called the signalosome “a kinase complex” which gives it an
additional role outside of the proteasome degradation pathway and perhaps a role in
signal transduction. This alternative mechanism, by which the CSN/COP9 signalosome,
through CSN-2, could directly interact with GLP-1/ Notch signaling altering its function
82 and allowing for proliferation defects. Given that several kinases, for example CK2,
PKD, AKT, and inositol 1,3,4-triphosphate 5/6 kinase are known to associate with the
signalosome, it is possible that the transduction of signaling could be altered (Uhle et al.,
2003; Haun et al., 2012; Sun, Wilson, and Majerus 2002). While these mechanisms are
certainly a possibility, the current literature supports a model by which proliferation is
affected in a more indirect way, via the CULLIN RING E3 ligases.
The second mechanism by which csn mutants could influence proliferation is by
modulating the cell cycle. Fox et al., 2011 demonstrated that the proliferative mitotic cell
cycle in the PMT of the worm gonad is very different from somatic cells. 57% of the cell
cycle is spend in S-phase, 39% is spend in G2 phase and 4% in M and G1 phases. The
total cell cycle time is 6.5-8 hours. The mitotic germline nucleus essentially replicates its
DNA and prepares for replication again.
Two important cell cycle regulators, Cyclin E and cyclin dependent kinase
(CDK), are active in germline proliferation in the PMT zone. These two proteins are
critical in the proximal end of the PMT where cells enter meiosis and where GLP-1 is not
highly expressed. Cyclin E and CDK function to maintain the inhibition of meiosis entry.
In C. elegans, CYE-1 (cyclin E) and CDK-2 have been shown to regulate proliferative
cell fate and the cell cycle. Both proteins are expressed throughout the PMT and
depletion of either protein results in cell cycle arrest (Fox et al., 2011).
CUL-2 has been implicated in proliferation. The loss of CUL-2LRR-1 results in cell
cycle arrest as well as defect in meiotic entry and SC assembly (Burger et al., 2013).
Specifically, CUL-2LRR-1 counteracts the ATL1/DNA replication checkpoint. It also
affects the size of the PMT and acts in the Notch signaling pathway, although it is unclear
how. CUL-2 is not the only CULLIN that is implicated in cell cycle regulation. In
mitotic cells, CUL-4 negatively regulates CDT-1 during S-phase in embryonic cells and
prevents aberrant re-initiation of replication during (Zhong et al., 2003; KIM and Kipreos
2007; Wei, Serino, and Deng 2008). This preserves genome stability in mitotically
83 replicating cells. Since cells in the proliferative zone spend the majority of their cycle in
S- phase, CUL-4 could regulate the rapid re-initiation of replication in these cells as well.
Interestingly, Drosophila csn4 and csn5 mutants cannot stabilize Cyclin E leading
to defects in cell cycle progression of mitotically proliferating germline nuclei (Doronkin,
Djagaeva, and Beckendorf 2003). Both csn 4 and csn5 demonstrated a dominate genetic
interaction indicating that the lack of stabilization was likely due to the signalosome
holoenzyme rather than csn5 acting independently. Furthermore, mutations in both genes
led to an increase in neddylated cullin1 (Doronkin, Djagaeva, and Beckendorf 2003).
This suggests the affect of the CSN/COP9 signalosome may be indirect, acting through
the CRLs to either stabilize substrates through monoubquitination or polyubiqutination
leading to degradation by the proteasome. This argument is supported by the fact these
authors also found dominate genetic interactions between cullin1 and csn4 or csn5.
Additionally, they observed a difference in the cellular localization of the neddylated
cullin1. In the CSN mutants, neddylated culin1 was found in the cytoplasm, not the
nucleus, they suggest that the signalosome may also regulate the shuttling of CRL’s
which could affect degredation of cell cycle regulators (Doronkin, Djagaeva, and
Beckendorf 2003).
Fox et al., 2011 also examined glp-1;cye-1(RNAi) double mutants to determine if
knocking out both pathways would allow nuclei to prematurely enter meiosis. They saw a
premature entry into meiosis and a spatial movement of meiotic entry toward the distal
end of the germline, which is similar to that seen in the csn mutants. Given the known
and possible interactions of the CSN/COP9 signalosome, likely an indirect affect through
the CRLs, with the DTC/GLP-1 signaling and CYE/CDK-2 pathway, these data suggests
a conserved function of the CSN/COP9 signalosome in pre-meiotic germline
proliferation. Future studies should include an examination of these pathways to
determine if the lack of nuclei is truly a result of proliferation defects.
84 CSN/COP9 is required for meiotic progression and pachytene exit
Once nuclei of csn mutants enter meiosis, chromosomes cluster to one side of the
nucleus as in wild-type; unlike wild-type however, a portion of these nuclei do not reacquire the normal dispersed chromosomal organization as they progress through meiosis
(Figure 3- 2 D’-F’, D’’-F’’). This phenotype of persistent polarized chromosome
organization is reminiscent of syp null mutants during meiotic progression. This finding,
together with the observation that csn mutants do initiate meiotic recombination and form
some crossovers, is consistent with meiotic progression from leptotene to pachytene in
these mutants. However, unlike syp mutants, csn mutants produce very few
diakinesis/oocyte nuclei. Reduced oocyte production could be attributed to the decrease
in the numbers of nuclei, yet the effect on oocyte production is greater than expected
from the reduction in number of pachytene nuclei destined to be oocytes. We found
proximal MAPK signaling (dpMPK-1) is reduced in csn mutants. MAPK signaling is
essential for pachytene progression (M.-H. Lee et al., 2007) and so we infer that reduced
dpMPK-1 levels are likely the primary contributor to the severe reduction in oocyte
numbers in csn mutants. As synapsis defective mutants (e.g., syp-1) still exit pachytene
and form oocytes in comparable levels to wild-type, the lack of MAPK signaling in the
csn mutants defines another function for the CSN/COP9 complex and is not a secondary
effect of the synapsis defects. Since CSN-5 physically interacts with MPK-1 (Li 2004;
Zhong and Sternberg 2006), the absence of MPK-1 phosphorylation may be due to the
absence of this interaction.
The absence of proximal MAPK signaling may indicate a problem not only with
the nuclei themselves, but with in the somatic sheath that surrounds the gonad (starting at
the transition zone and extending all the way to the spermatheca). MAPK signaling is
activated by the major sperm protein (MSP) that binds to receptors on the sheath cells
(M. A. Miller 2003). Current literature supports a model by which the gonadal sheath
promotes the transduction of the MAPK signal from the spermatheca towards the distal
85 gonad and facilitates oocyte maturation (McCarter et al., 1997; M. A. Miller 2003). Laser
ablation of the primordial sheath cells (SS cells) in larvae at the L2/L3 molt or at mid-L4
resulted in an intact gonad with no sheath (McCarter et al., 1997). However, the lack of
an intact sheath resulted in a lack of proliferation, and a defect in pachytene exit,
reminiscent of the phenotypes observed in the csn mutants. The lack of a central rachis in
csn mutants also suggests there is a problem with gonadal organization and perhaps the
transmission of MAPK and other critical signals to the csn mutant nuclei. Oocytes and
sheath cells are bound together by gap junctions, which are particularly evident in the
proximal gonad (D. H. Hall et al., 1999). These junctions place the oocytes in close
enough proximity to the sheath cells to allow for transduction of MAPK signals (or
others) through various receptors such as VAB-1/Ephrin (M. A. Miller 2003; Kuwabara
2003). These data indicate an essential function of cell-cell interactions in the germline to
promote oocyte maturation. Given what appears to be a reduction in cell-cell contact (the
absence of a rachis and disruption of gonadal morphology) between some nuclei to the
gonadal sheath, it is possible that there is a disruption of signaling mechanisms in the csn
mutants. However, it is unclear at this time what role the CSN/COP9 signalosome has in
the MAPK signaling pathway. Sasagawa et al., 2007, demonstrated CUL-2 and CUL-5
lack dpMPK-1 signaling; however these authors failed to discern a mechanism by which
the signalosome or CRLs influenced dpMPK-1 activation.
Our data are consistent with a dual meiotic role for the CSN/COP9 signalosome:
1) promoting early prophase SC assembly required for homolog pairing and meiotic
recombination, and 2) a second function in mid-prophase, involving interaction with
MAPK signaling promote pachytene exit. Previously studied germline proliferation and
progression mutants (leading to short gonads) were not reported to have defects in SC
assembly (Smith et al., 2002; Orsborn et al., 2007) and support our model that the
proliferation and SC function of the CSN/COP9 complex are separated. The role of the
CSN/COP9 signalosome in meiotic progression seems to be conserved, as in Drosophila
86 csn4null, csn5null and csn8null mutants which arrest at the pachytene-diplotene transition
(Oren-Giladi et al., 2008; Oron et al., 2007; Oron et al., 2002). csn8null germline clones
resulted in stage 7 arrest (Oren-Giladi et al., 2008) while csn4null clones arrested in stages
5-6 (Oron et al., 2002) In the csn8null clones cullin neddylation was impaired indicating
the oogenesis defect maybe a indirect affect of CSN/COP9 signalosome via CRL
regulation. Additionally, oocyte polarity was severely altered in csn4 and csn5 mutants.
Oskar (posterior fate), bicoid (anterior fate) and gurken (anteroposterior and dorsoventral
axis) were all examined in the the csn4 and csn5 mutants. csn4nulls had normal oskar
staining, but lacked bicoid and gurken. csn5null animals did not stain for any of these three
proteins (Oron et al., 2002). This could indicate a role for the signalosome in
transcription as well. Gurken transcription occurs in stage 7 as the nucleus to an anterior
dorsal position. It was suggested that the lack of polarity is what arrests the oocytes in
stages 5-7. Again, it is not clear is this a direct or indirect affect of the signalosome.
How CSN/COP9 regulates chromosome synapsis
In C. elegans, CR/SYPs assembly can be misregulated in certain meiotic mutants
without forming aggregates (Couteau 2005; Martinez-Perez 2005; Smolikov, Eizinger,
Hurlburt, et al., 2007; Smolikov, Schild-Prüfert, and Colaiácovo 2008). These aberrant
forms of SC assembly appear to be fully formed SCs that assembled in the wrong
chromosomal context as in HIM-3 mutants (Zetka 1995). The situation found in the csn
mutants is different: in addition to aberrant SC assembly (short stretches) ~50% of nuclei
contain one bright SYP aggregate not in contact with DNA. In C. elegans, lack of any of
the SC proteins results in elimination of the SYPs without their aggregation (SchildPrüfert et al., 2011), indicating that mechanisms exist to remove SYPs not bound to
DNA. csn mutants are therefore, likely perturbed in mechanisms designed to clear
aggregated SYPs, and assemble a “SC-like” structure which is invisible to the
degradation machinery. In yeast, SC proteins are undergoing rapid turnover (the SC
assembled continually)(Voelkel-Meiman et al., 2012). If a similar rapid exchange of
87 SYPs occurs during C. elegans meiosis, SC assembly defects (problems in SC assembly
upon meiotic entry) and SC stabilization defects (throughout pachytene) are related, due
to continuous assembly of SC protein occurring throughout prophase.
There are three known examples of polycomplex-like structures in C. elegans
mutants: cra-1;spo-11 double mutants(Smolikov, Schild-Prüfert, and Colaiácovo 2008),
pgl-1 mutants at 25oC and higher (Bilgir et al., 2013) and dynein mutants, in early
prophase (Sato et al., 2009). cra-1;spo-11 double mutant nuclei remained polarized
throughout pachytene as in syp-1 mutant and much like what we see in the csn mutants;
however, unlike the csn mutants, they displayed a low level of normal SYP-1 staining in
the transition zone. This indicated that the early stages of SC assembly were likely intact.
SYP-1 aggregates were observed in pachytene in these double mutants, which
corresponded to the initiation of DSB and suggested that the absence of spo-11, SYP-1
will aggregate. This showed CRA-1 likely has a role in regulating SC assembly but after
the pairing stabilization. In the absence of CRA-1, DSB are required for full SC assembly
(Smolikov, Schild-Prüfert, and Colaiácovo 2008). Another mutant, PGL-1 a P-granule
component that helps promote stable pairing, also displayed aggregation of SYP proteins
in response to excessive heat (above 26.5C) (Bilgir et al., 2013). However, this
phenotype was also observed in wild-type worms at the higher temperatures as well,
indicating that SC assembly is temperature sensitive. These authors suggest that this
maybe means by which C. elegans adapts to heat stress and increases population fitness
(Bilgir et al., 2013). The last group of mutants known to have SYP aggregation are the
dynein mutants (Sato et al., 2009). Dynein is essential to the pairing and synapsis of
homologous chromosomes in C. elegans. Interactions between the pairing centers on the
homologous chromosomes, the nuclear envelope and the cytoskeleton generate tension
between chromosomes, which in turn promotes SC polymerization. In the absence of this
tension, the SYP will aggregate but only in early pachytene (Sato et al., 2009). Pairing
eventually does occur, but is delayed. However, synapsis does seem to be abnormal.
88 Although CSN/COP9 could interact with one of these pathways, our analysis thus
far is consistent with a different function; aggregates in csn mutants are found in the
presence of DSBs (unlike cra-1), at normal growth temperatures [20oC, unlike pgl-1
mutants], and throughout the germline (unlike dynein mutants). Therefore, we propose
that CSN/COP9 participates in SC assembly in a novel manner (Figure 3-19B). The
signalosome role is not merely due to promoting SYP degradation, since csn mutants do
not show increase in SYP-1 levels.
Pathways of SC assembly involve post-translational modifications made to SC
proteins. It is conceivable these modifications could facilitate CR protein association with
chromosomes and prevent their aggregation in the absence of binding to DNA. In yeast, it
was shown SUMOylation promotes lateral element (Watts and Hoffmann 2011) and CR
(Voelkel-Meiman et al., 2013) assembly. Mouse SC assembly is regulated by
phosphorylation of lateral element proteins (Fukuda et al., 2012). In C. elegans, SYP-1
and SYP-2 appear to be post-translationally modified; however, precise identities of these
modifications are not yet known (Schild-Prüfert et al., 2011). All four SYPs contain
many potential sites for phosphorylation, ubiquitination, and SUMOylation (Table 3-8).
In C. elegans, an evolutionarily conserved ubiquitin/sumo modifier (Lake and Hawley
2013) is linked to SC disassembly, but is not required for SC assembly (Bhalla, Wynne,
and Jantsch 2008).
The discovery that the CSN/COP9 complex is required to prevent SYP
aggregation raises the question of whether CSN/COP9 is involved in post-translational
modification of the SYPs to prevent their aggregation. SYP aggregates contain all four
SYP proteins, hence one aggregation-prone SYP may lead to the capture of all SYPs. The
CSN/COP9 complex’s activity in deneddylation has been well documented, but it also
possesses Ser/Thr kinase and deubiquitination associated activities (Wei and Deng 2003).
Thus, one role of the CSN/COP9 signalosome could be phosphorylating or removing
Nedd8 or ubiquitin from SYPs and thereby precluding SYP self-assembly when not
89 bound to DNA. In the absence of such modification, SYPs would aggregate in the
absence of DNA binding.
CSN/COP9 may repress the aggregation of SYP proteins indirectly, by inhibiting
CRL monoubiquitination of SYPs, which promotes SC assembly. Alternatively,
CSN/COP9 might directly modify the SYPs. CSN/COP9 subunits interact with multiple
proteins (Kato and Yoneda-Kato 2009) other than CRLs(Mergner and Schwechheimer
2014). Furthermore, CSN/COP9 function is not limited to deneddylation, indicating these
additional functions of the CSN/COP9 complex (deubiquitination and phosphorylation)
are possible alternative modes of action.
The CSN subunits and their roles in meiotic events
The CSN/COP9 complex is composed of seven to eight subunits, depending on
the organism (Figure 3-1) (Wei and Deng 2003). Recent structural studies suggest the
signalosome holoenzyme consists of two asymmetrical subcomplexes; one containing
CSN1/2/3/8 and the other containing CSN4/5/6/7 with the two conserved subunits, CSN2
and CSN5 located on the periphery of their respective structures (Sharon et al., 2009).
The subcomplexes are held together by a single link between CSN1 and CSN6 (Sharon et
al., 2009). Mutant analyses have indicated the loss of any one subunit leads to
signalosome disassembly (Busch et al., 2007; Serino and Deng 2003). The CSN5 subunit
contains the deneddylation activity, but cannot function in deneddylation outside the
holoenzyme (Cope and Deshaies 2003). Smaller sub-complexes, with variable subunit
composition have been isolated as well: CSN4-7 and CSN1/2/3/8 Arabidopsis and
Drosophila (Sharon et al., 2009; Oron et al., 2002; Kotiguda et al., 2012) and CSN-4-56-8 in mammals (Tomoda 2005). However, there is no direct evidence for biological
function of these subcomplexes and there are no crystal structures to confirm their
conformation. The csn mutant phenotypes are not identical, suggesting functions outside
the CSN/COP9 signalosome for individual subunits. For example: Drosophila csn4, csn5
and csn8 mutations cause larval lethality, but they die in different larval stages
90 (Doronkin, Djagaeva, and Beckendorf 2002; Oren-Giladi et al., 2008; Cope and Deshaies
2003; Oron et al., 2002). Both csn4null and csn5null died as 3rd instar larvae (Cope and
Deshaies 2003), but csn4null succumbed to molting defects while csn5null died as a result
of developing melanotic tumors (Oren-Giladi et al., 2008). Csn8null lived past the 3rd
instar stage, but interestingly had molting defects and melanotic tumors (Oren-Giladi et
al., 2008). It is also worth nothing that the 3rd instar stages of csn8null were much smaller
than wild-type, indicating a problem with growth. Oron et al., 2007 suggests that later in
development, CSN/COP9 signalosome may be affecting transcriptional regulation as well
as hormonal pathways, leading to the defects observed. These authors also noted that
csn4 and csn5 were found as stable monomers outside the signalosome complex. It has
been shown that CSN4/5/6/7 can exist as a subcomplex in Drosophila. This could be
evidence of biological role for this subcomplex later in development.
S. pombe csn1 and csn2 mutants show defects in meiotic entry and meiotic
recombination, while mutants in the other subunits have no clear meiotic phenotypes
(Mundt, Liu, and Carr 2002). csn1 and csn2 both display defects in meiotic S phase,
specifically the incomplete replication of chromosomes and other genomic instability
which resulted in the activation of the DNA checkpoint (Mundt et al., 1999). However,
examination of csn4 and csn5 mutants failed to detect any sensitivity to radiation or
problems with S phase replication (Mundt, Liu, and Carr 2002). It is also worth noting
that with all csn mutants examined, there was hyperneddylation of Pcu1, the S. pombe
homolog of cullin1 (Mundt, Liu, and Carr 2002).
CSN2 (mammals) and CSN5 (Arabidopsis, Drosophila, mammalian cell lines) are
the only subunits shown to act outside CSN/COP9 in vivo (Wei and Deng 2003). CSN5
binds numerous substrates and has been proposed to promote apoptosis or cell
proliferation outside the CSN/COP9 complex(Wei, Serino, and Deng 2008; Chamovitz
2009). In mammals, a truncated CSN2 isoform (referred to as ALIEN) acts in
nucleosome assembly (Eckey et al., 2007).We have shown that both csn-2 and csn-5
91 mutants lead to defects in SC assembly, a reduction in pairing, and increase in DSB
repair defects in meiotic recombination. However, the magnitude of these effects varies:
the csn-2 mutant almost mimics a syp null mutant, while the csn-5 mutant shows milder
pairing stabilization defects and double the numbers of crossovers compared to the csn-2
mutant. The SC is driving the stabilization of early prophase pairing interactions (zone 23), while later events (zone 6) are also promoted by the stabilizing role of crossovers,
which are higher in the csn-5 mutant. Both mutants show similar SC defects; however,
csn-5 mutant aggregation is suppressed by the pch-2 mutant while the aggregation in csn2 mutants is not. This suggests the SC assembly defects may not be identical on the
molecular level in these two mutant backgrounds, although both csn-2 and csn-5
aggregated SCs are indistinguishable by cytological methods. The mis-assembled SC of
the csn-5 mutant is different from that of the csn-2 mutant in two aspects. First, it can
promote basal levels of pairing stabilization in early prophase, and second, this deficient
stabilization can trigger the synapsis checkpoint.
Another major difference between the two csn mutants is in meiotic
recombination. Both RAD-51 foci and crossover levels are 2-fold higher in late
pachytene of the csn-5 mutant compared to the csn-2 mutant. The higher crossover level
could be the result of better pairing stabilization in the csn-5 mutants. The larger
increased level of recombination intermediates in csn-5 compared to csn-2 mutants is less
easily explained. It is possible the intermediate pairing levels delay repair via alternative
modes of repair (sister chromatid recombination), resulting in an increase in overall
recombination intermediates.
Taken together, our data suggest that the residual SC structure in the csn-5 mutant
is more “functional” compared to the one in csn-2 mutant. It is possible the milder effects
of csn-5 are due to it not being a null allele. This seems unlikely since the csn-5(ok1064)
allele contains a large deletion which removes its catalytic MPN domain and ~60% of the
predicted coding sequence. An alternative explanation for the distinct phenotypes is that
92 one of the proteins acts solely in the CSN/COP9 signalosome, while the other acts both in
the CSN/COP9 signalosome and in an additional CSN-based sub-complex (or individual
subunit). The only prior known function of CSN-2 outside CSN/COP9 is in a context of a
C-terminally truncated protein isoform (Bech-Otschir et al., 2001). However, C. elegans
has only one protein isoform of CSN-2, making this mode of action unlikely. CSN-5 has
known functions outside the CSN/COP9 complex. Therefore, we favor a model in which
CSN-5 acts both in and outside the CSN/COP9 signalosome. In addition, CSN-5 could
also directly affect meiotic recombination, independent of its role in pairing stabilization.
In Drosophila, the csn5 mutant lethal phenotype is attributed to the activation of the
germline DNA damage checkpoint (Doronkin, Djagaeva, and Beckendorf 2002). In
somatic tissues, CSN-5 interacts with multiple proteins, some of which are involved in
DSB repair/signaling such as p53 (Bech-Otschir et al., 2001), c-Jun (Tsuge et al., 2011)
or Rad1 [9-1-1 complex (Huang et al., 2007)]. This may explain why csn-5 mutants have
slightly increased RAD-51 foci levels in the mitotic zone as well as in late pachytene
nuclei. Alternatively, the increase in recombination intermediates in both csn mutants
could be due to inefficient homologous repair (HR). Moss et al., 2010 demonstrated in
fission yeast that Ddb1-Cul4Cbt2 regulated nucleotide synthesis was required for efficient
HR to occur. These authors proposed that Ddb1-Cul4Cbt2 mediated Spd1 degradation via
ubquitination, which lead to ribonucleotide reductase (RNR) activation. This allowed for
completion of gap repair across resected ssDNA at break foci (Moss et al., 2010). If
Ddb1-Cul4Cbt2 were rendered nonfunctional by a mutated signalosome complex then HR
would likely fail leading to the increase in RAD-51 foci and lack of crossover formation.
CUL-4 has numerous functions in DDR depending on the adaptor attached to the
CULLIN subunit (Hannß and Dubiel 2011; Jackson and Xiong 2009).Given that an intact
CSN-5 subunit is required for the regulation of CRLs, this could explain the increase in
the RAD-51 foci in csn-5 mutant.
93 Another possible mechanism could involve the ubiquitintion of histones H3 and
H4 which is mediated buy Cul4-DD4-ROC1 in mammalian cells (HeLa) under going
mitosis (H. Wang et al., 2006). The ubiquitination of H3 and H4 leads to the release of
nucleosomes around the damage foci and allows for the recruitment of repair proteins (H.
Wang et al., 2006). Again, the lack of a functional Cul4-DD4-ROC1 complex as a result
of a non-functional CSN/COP9 signalosome could have a profound impact on DNA
damage repair. It is also worth noting that in csn-5 mutants, we saw an increase of RAD51 foci in zones 1 and 2 where meiotic recombination had yet to be initiated. These foci
could be residual DNA damage that is unrepaired due to a lack of a functional CUL-4
complex.
Summary
We therefore, propose the main germline function of the CSN/COP9 signalosome is to
promote SC assembly and pairing stabilization (Figure 3-20A). While CSN-2 and CSN-5
both promote synapsis by suppression of aggregate formation via their function in
CSN/COP9, we suggest CSN-5 also acts outside the CSN/COP9 signalosome in
antagonizing SC stabilization. A mechanism for the negative regulation of pairing
stabilization may require the inhibition of PCH-2 (Deshong et al., 2014). A dual function
of CSN-5 in SC assembly (positive, part of CSN/COP9 complex) and pairing
stabilization (negative and outside the CSN/COP9 complex) may explain why the meiotic
defects of csn-5 are milder than ones observed for csn-2 mutants. However, it is unclear
at the time if the signalosome is acting directly in the mechanisms of SC assembly or if it
is acting indirectly through the CRLs (Figure 3-20B). Given that many csn mutant
phenotypes have been subsequently linked to hyperneddylation of CRLs, it is likely that
CRLs are involved. Alternatively, it could be that in specific cases, such as the absence of
MAPK signaling in the proximal end of the gonad, the signalosome could be directly
involved by facilitating signal transduction or providing a platform for protein-protein
interactions.
94 Figure 3-1: CSN/COP9 signalosome structure and function. The CSN/COP9
signalosome consists of eight subunits and its structure has been biochemical determined.
CSN5 (in bright red) is the catalytic core of the holoenzyme. CSN2 (blue) is required for
holoenzyme integrity. The signalosome is thought to stabilize substrates through
phosphorylation. The most studies aspect of the signalosome is its regulation of the
Cullin E3 ligases via deneddylation which deactivates the ligases. Deactivation of the
ligase will prevent mono-ubiquitation that leads to substrate localization or substrate
interaction. Non-functional ligases can also lack of degradation of substrates as they are
not poly-ubiquitinated and channeled to the 26S proteasome.
95 Figure 3-2: SC central element assembly defects in csn mutants. A) CSN alleles used
in this study: csn-2(tm2823) and csn-5(1064), black rectangles represent exons, black
lines introns, gray areas represent UTR regions, and pink rectangles the deletion in each
allele. B-F’’) Micrographs of SYP-1 (red) and DAPI (blue) stained wild-type (B-F), csn2 (B’-F’)and csn-5 (B’’-F’’) mutants nuclei representing the various stages of the C.
elegans gonad. Images are projections through half of a three-dimensional data stacks.
Scale bar is 2µm. PMT=pre-meiotic tip, TZ= transition zone, EP= early pachytene,
MP=mid pachytene, LP= late pachytene. SYP-1 aggregates appear in the TZ-like stage of
the gonad and persist through the LP-like stage.
96 Figure 3-3: csn mutants have smaller gonads and lack of oocytes progressing
through diakinesis. SYP-1 aggregates appear in the TZ-like stage of the gonad and
persist through the LP-like stage. A) Whole gonad from wild-type (B-F), csn-2 (B’-F’)
and csn-5 (B’’-F’’) mutants SYP-1 and DAPI stained. Images are projections through
half of a three-dimensional data stacks. Scale bar is 2µm. PMT=pre-meiotic tip, TZ=
transition zone, EP= early pachytene, MP=mid pachytene, LP= late pachytene. Scale Bar
40 µm B) SYP-1 (grayscale) staining only of gonads showing aggregation throughout the
gonad, starting at transition zone.
97 Figure 3-4: HTP-3 not aggregate in csn mutants. Micrographs of HTP-3 (red) and
DAPI (blue) stained wild-type, csn-2(tm2823) and csn-5(ok1064) nuclei representing the
various stages of the C. elegans gonad. Images are projections through three-dimensional
data stacks. Scale bar is 2µm. PMT=pre-meiotic tip, TZ= transition zone, EP= early
pachytene, MP=mid pachytene, LP= late pachytene. HTP-3 localization is not affected in
csn mutants.
98 Figure 3-5: HIM-3 does not aggregate in csn mutants. Micrographs of HIM-3 (red)
and DAPI (blue) stained wild-type, csn-2(tm2823) and csn-5(ok1064) nuclei representing
the various stages of the C. elegans gonad. Images are projections through threedimensional data stacks. Scale bar is 2µm. PMT=pre-meiotic tip, TZ= transition zone,
EP= early pachytene, MP=mid pachytene, LP= late pachytene. HIM-3 localization is not
affected in the csn mutants.
99 Figure 3-6: SYP-4 aggregates in csn mutants. A-E’) Micrographs of SPY-4 (green)
and DAPI (blue) stained wild-type, and csn-5(ok1064) nuclei representing the various
stages of the C. elegans gonad. Images are projections through three-dimensional data
stacks. Scale bar is 2µm. PMT=pre-meiotic tip, TZ= transition zone, EP= early
pachytene, MP=mid pachytene, LP= late pachytene. Aggregation affects all SYP-4 and
likely all SYPs.
100 Figure 3-7: P-granule components kgb-1(um3) and glh-1(gk100) mutants do not
have SYP-1 aggregation phenotype. A-C’) Micrographs of SYP-1(red) and DAPI(blue)
stained wild-type (A-C), kgb-1(um3)(A’-C’), and glh-1(gk100) (B’’-C’’) mutant nuclei
representing the various stages of the C. elegans gonad. Images are projections through
three-dimensional data stacks. Scale bar is 2µm. EP=early pachytene, MP=mid
pachytene, LP=late pachytene. kgb-1(um3) and glh-1(gk100) are temperature sensitive
alleles. Worms cultured at 26C do not exhibit SYP-1 aggregation. P-granules do not
appear to be involved in the aggregation phenotype.
101 Figure 3-8: Quantification of the SYP-1 aggregates. A) Schematic representation of
the zones of the C. elegans gonad. PMT=pre-meiotic tip, TZ= transition zone, EP= early
pachytene, MP=mid pachytene, LP= late pachytene. B-C) Quantification of SYP-1
aggregates in zones of the gonad. B) Percent of nuclei with: no SYP-1 (black), linear SC
(blue), aggregated SYP-1 (red) and other (yellow), zones as in A, n nuclei scored
(ordered by zone number): wild type: 60, 63, 49, 55, 42 41, csn-2: 119, 120, 147, 148,
108, csn-5: 136, 179, 166, 171, 139, 64. In collaboration with S. Smolikove and N.
Balukoff
102 Figure 3-9: Representative images of nuclei and aggregation phenotypes.
Micrographs of SYP-1(red) and DAPI(blue) stained csn-2 mutant nuclei representing the
aggregation types observed. Images are projections through three-dimensional data stack.
In collaboration with S. Smolikove and N. Balukoff
103 Figure 3-10: Quantification of SYP-1 protein in csn mutants. A) Western analysis
confirming the reduction of expression of SYP-1 in csn mutants. Normalization values
(α-SYP-1/α-TUB) shown are the average of 2 different experiments. Normalized
intensities: wild-type 0.72±0.26, csn-2 0.24±0.20 and csn-5 0.58±0.11. B) Quantification
of the amount of SYP-1 in the nuclei measured by the number of pixels per nucleus and
in the cytosolic portion of the oocyte. Reduced nuclear SYP-1 localization in csn
mutants. *pMW<0.0005-nuclear average and **pMW<0.0005-cytosolic average.
104 Table 3-1: Average gonad lengths from PMT to diplotene and p-values for pairwise
comparisons between the single mutant control and the double mutants.
Gonad Length, PMT to diplotene
p-value MW
One Day old
Adults
wild-type
csn-2(tm2823)
Average
(mM)
307
193.9
Standard
error
10.23
2.99
% of csn
mutant
NA
NA
wt vs csn
mutant
csn-2 vs
csn-5
csn-5(ok1064)
176.5
4.17
NA
1.57E-04
pch-2(tm1458)
311.3
3.53
NA
pch2(tm1458);csn2(tm2823)
187.5
11.56
97
1.57E-04
pch2(tm1458);csn5(ok1064)
185.9
12.56
105
1.57E-04
cep-1(RNAi)
372.1
8.58
NA
csn2(tm2823);cep1(RNAi)
190.3
7.38
98
1.57E-04
csn5(ok1064);cep1(RNAi)
185.4
8.7
105
1.57E-04
0.545
Three day old
Adults
wild-type
csn-5(ok1064)
Average
(mM)
327.5
124.6
Standard
error
4.5
4.5
% of csn
mutant
NA
NA
wt vs csn
mutant
csn-2 vs
csn-5
pch2(tm1458);csn5(ok1064)
214.3
19.77
172
1.50E-04
csn5(ok1064);cep-1
212.7
9.88
171
1.50E-04
1.57E-04
0.008
0.88
NA
Note: 1 day adult (24 hours post L4) is the standard age for examining meiotic events.
n=10 for each genotype studied, NA= Data not available
105 Figure 3-11. Quantification of the lack of oocytes. A) Relative sizes of the pre-meiotic
tips for wild-type and the csn mutants. Number of rows is indicative of length of PMT.
The size of the mitotic zone is reduced in csn mutants. B) Quantification of the number of
gonads that contained oocytes in diakinesis for the csn mutants, *pMW<0.0005 and
**pMW<0.005.
106 Table 3-2: The average number of oocytes in diakinesis for csn mutants and csn mutant,
apoptosis checkpoint double mutants
wild-type
csn-2(tm2823)
csn-5(ok1064)
syp-1(me17)
pch-2(tm1458)
pch-2(tm1458);csn-2(tm2823)
pch-2(tm1458);csn-5(ok1064)
cep-1(RNAi)
csn-2(tm2823);cep-1(RNAi)
csn-5(ok1064);cep-1(RNAi)
ced-4(RNAi)
csn-2(tm2823);ced-4(RNAi)
csn-5(ok1064);ced-4(RNAi)
n = the number of gonads examined
Average number of
oocytes per gonad
8.4
Standard
Error
0.15
n
50
0.37
0.06
125
1.16
0.098
100
9.15
0.25
17
10.92
0.28
50
0.24
0.07
50
0.16
0.06
49
14.35
0.35
20
0.4
0.15
20
0.25
0.09
20
14.5
0.41
20
0.35
0.13
20
0.35
0.10
20
107 Table 3-3: The average number of eggs laid for csn mutants and apoptosis checkpoint
double mutants.
wild-type
Average number of
eggs laid per worm
247
Standard
Error
16
n
3
0
0
13
0
0
11
148
22
3
185
42
2
0
0
3
0
0
3
NA
NA
NA
0
0
3
0
0
3
NA
NA
NA
0
0
3
0
0
3
csn-2(tm2823)
csn-5(ok1064)
syp-1(me17)
pch-2(tm1458)
pch-2(tm1458);csn-2(tm2823)
pch-2(tm1458);csn-5(ok1064)
cep-1(RNAi)
csn-2(tm2823);cep-1(RNAi)
csn-5(ok1064);cep-1(RNAi)
ced-4(RNAi)
csn-2(tm2823);ced-4(RNAi)
csn-5(ok1064);ced-4(RNAi)
n = the number of worms examined
NA= Data not available
108 Figure 3-12. Pairing stabilization is affected in csn mutants. A) Analysis of pairing
stabilization between wild-type, syp-1(me17), csn-2, and csn-5 mutants. A schematic
representation of the timing of meiotic stages relative to the zones in the C. elegans
gonad. zone 1= pre-meiotic tip, zone 2 and 3= transition from mitosis to meiosis, zone 46 = pachytene. The black arrow represents the movement of nuclei through the stages
(zones) of meiosis. csn mutants show defects in pairing stabilization.
109 Figure 3-13: High magnification micrographs of individual nuclei from FISH
analysis. Images are projections through three-dimensional data stacks. 5S FISH probe
foci are in green and DAPI stained chromosomes are in blue. zone 4= early pachytene,
zone 5 = mid pachytene, zone 6= late pachytene. Scale bar is 2µm.
110 Table 3-4: Number of nuclei counted for FISH analyses
wild-type
syp-1
csn-2
csn-5
Zone 1
73
41
47
78
Zone 2
87
49
28
104
Zone 3
74
38
45
103
Zone 4
97
42
35
97
Zone 5
97
40
49
106
Zone 6
81
22
33
65
Total Nuclei
509
232
237
553
111 Table 3-5: p-values calculated by Fishers Exact Test for all pairwise comparisons of
FISH data
wt vs
csn-2
wt vs
csn-5
csn-2
vs csn-5
wt vs
syp-1
csn-2
vs syp-1
csn-5 vs
syp-1
Zone 1
3.78E-01
1.79E-02
2.47E-01
3.49E-01
1.00E+00
3.75E-01
Zone 2
5.93E-01
9.07E-08
7.54E-03
2.52E-01
1.00E+00
1.10E-03
Zone 3
2.92E-05
1.00E+00
8.06E-06
3.03E-06
4.98E-01
6.23E-07
Zone 4
9.15E-07
6.87E-08
3.57E-01
2.16E-06
7.75E-01
6.71E-01
Zone 5
6.25E-10
6.31E-07
2.49E-02
1.60E-12
2.12E-01
3.76E-04
Zone 6
2.51E-08
2.84E-02
2.23E-04
1.81E-05
6.74E-01
9.94E-03
112 Figure 3-14: Accumulation of recombination intermediates in csn mutants Analysis
of RAD-51 foci in wild-type compared to csn mutants. Position along the x-axis refers to
the zone in the gonad (Figure2-7A). Schematic representation of the timing of meiotic
stages relative to zones scored in the csn mutants compared to wild-type. RAD-51 foci
accumulate upon entrance to meiosis in csn mutants.
113 Table 3-6: p-values and total number of nuclei counted per zone for RAD-51 analyses
p-values Mann Whitney Test
wt vs
wt vs
csn-2
csn-2
csn-5
vs csn-5
Number of nuclei counted per
zone
wild-type
csn-2
csn-5
Zone 1
0.307
< 2e-06
0.0002
159
60
46
Zone 2
0.131
<0.000004
0.0094
209
70
49
Zone 3
< 2e-06
< 2e-06
0.0022
199
61
62
Zone 4
Zone 5
< 2e-06
< 2e-06
< 2e-06
< 2e-06
0.0322
0.8092
177
174
56
69
47
55
Zone 6
0.023764
< 2e-06
7.82E-12
142
47
49
114 Figure 3-15: Reduced crossover formation in csn mutants. Quantitative analysis of
COSA-1 foci in zone 6 of the wild-type and zone 6-like section of the csn mutants color
code for number on COSA-1 is at right. Number of crossovers marked by COSA-1 is
reduced,
115 Figure 3-16: High magnification micrographs from RAD-51 and COSA-1 analyses.
Micrograph images of COSA-1 foci (green), chromosomes DAPI (blue), and SYP-1 (red)
in wild-type and csn mutants. E) Micrograph images of RAD-51 foci (red), COSA-1 foci
(green), and chromosomes DAPI (blue) in wild-type and csn mutants. Images are
projections through three-dimensional data stacks. Scale bar is 2µm.
116 Figure 3-17. Apoptosis and MPK-1 expression are altered in csn mutants.
Quantification of the number of nuclei with CED-1::GFP present in late pachytene of the
C. elegans gonad. Red bars represent the total number of apoptotic nuclei in the late
pachytene region. The blue bars represent the total number of nuclei in the late pachytene
region. Apoptosis is increased in csn-2 mutants, but not in csn-5 mutants, *pMW<0.0005.
There is also a reduction of overall nuclei in the late pachytene region of the gonad in
both csn mutants, **pMW<0.0005.
117 Figure 3-18: Analysis of SYP-1 aggregate phenotype in csn mutants and apoptosis
checkpoint double mutants. Bypassing the apoptotic checkpoints reduces the number of
nuclei with aggregates.
118 Table 3-7: p-values and total number of nuclei counted in late pachytene zone for
apoptotic analyses.
Number of
nuclei
counted
p-value FET
single vs
csn mutant
wild-type
164
csn-2(tm2823)
173
9.15E-48
csn-5(ok1064)
164
1.70E-59
pch-2(tm1458)
171
pch-2(tm1458);csn-2(tm2823)
198
328
pch-2(tm1458);csn-5(ok1064)
cep-1(RNAi)
502
csn-2(tm2823);cep-1(RNAi)
336
180
csn-5(ok1064);ced-1(RNAi)
csn-2 vs
csn-5
0.0191
1.83E-47
2.91E-43
0.0230
7.09E-47
1.02E-42
0.239
119 Figure 3-19: Quantification of dpMPK-1 expression in csn mutants.
A) Quantification of pdMPK-1via cytological analyses. csn mutants lack MPK-1 staining
in late pachytene and in diakinesis, *pFET<0.0005 B) Western analysis confirming the
lack of expression of MPK-1B in csn mutants. MPK-1A is mostly somatic and MPK-1B
is germline specific. Normalization values (α-MPK-1/α-TUB) shown are the average of
3 different experiments. Normalized intensities: wild-type 2.27±1.03, csn-2 0.96±0.42
and csn-5 0.99±0.09.
120 Table 3-8: Predicted post-translational modification sites for SYP proteins
Protein
Size
(AA)
Molecular
Weight
(kD)
Total
Phosphorylation sites1
Total
Ubiquitination
sites2
Total
SUMOylation
sites3
SYP-1
489
56.6
462
16
6
SYP-2
213
23.7
191
8
2
SYP-3
224
25.8
186
4
0
SYP-4
605
67.3
403
15
2
1
Generated with GPS2.0 phosphorylation prediction software
2
Generated with UbiPred: Prediction of ubiquitination sites software
3
Generated with GPS-SUMO sumoylation site prediction software
121 Figure 3-20: Model for CSN/COP9 signalosome function in meiotic prophase
I. A) The proposed model for CSN/COP9 signalosome function in prophase
events. The signalosome holocomplex promotes synapsis while the CSN-5
subunit (acting in an independent manner) antagonizes synapsis. This accounts for
the differences in severity of defects between the csn mutant alleles. Additionally,
the holocomplex regulates MAPK signaling and the initiation of disassembly.
Panel A made by Sarit Smolikove. B) A non-functional CSN/COP9 signalosome
holocomplex could have either not stabilize SYP or a SYP interacting
substrate (a direct interaction). Alternatively, non-functional holoenzyme could
alter the regulation of CULLIN E3 ligases leading to a breakdown in the
ubiquitin/proteasome pathway. This would be an indirect interaction that could
affect a substrate that regulates polycomplex formation or in the case of
monoubiquitinaion, the SYPs themselves.
122 CHAPTER 4
METHODS AND MATERIALS
C. elegans strains and culture conditions
Most C. elegans strains were cultured under standard conditions at 20°C on petri
dishes containing nematode growth media (NGM) and HP50 E. coli as a food source
(Brenner 1974). Several strains (in bold) were maintained at 15°C and cultured for
experiments at 26°C. N2 Bristol worms were utilized as the wild-type background. The
following mutations and chromosome rearrangements were used and are described in
Wormbase:
LGI: asfl-1(ok2060), cep-1(ep347), csn-2(tm2823), glh-1(gk100), ned-8(gk3086),
hT2[bli-4(e937) qIs48]
LGII: akir-1(gk528),, akir-1(rj1), pch-2(tm1458), unc-85(ok2125)
LGIV: csn-5(ok1064), kgb-1(um3),let-92(ok1537), nT1[qIs50], uba-1(ok1374)
LGV: syp-1(me17)
The following transgenic lines were used: meIs8(GFP::COSA-1), smIs34 [ced-1p::ced1::GFP + rol-6(su1006)], iowEX15[ned-8;Pdor1::dsRED], iowEx13[uba-1; Pdor1::dsRed]
Some strains were provided by the Caenorhabditis Genetics Center (CGC) [which
is funded by NIH Office of Research Infrastructure Programs (P40 OD010440)], the C.
elegans Reverse Genetics Core Facility at UBC, which is part of the International C.
elegans Gene Knockout Consortium, and National Bioresource Project for the
Experimental Animal “Nematode C. elegans”, Japan.
PCR validation of strains
Genomic DNA was isolated from homozygous adult worms for each strain (Table
4-1) and PCR amplified with primers designed to flank the corresponding deletion in
each allele strain. Primers and PCR conditions are listed in Table 4-2. PCR products were
then gel electrophoresed on a 1% agarose gel at 100-125V for 15-30 minutes depending
123 on the expected size of the PCR products. Ethidum Bromide was used to detect the PCR
products in the agarose gels. Gels were imaged on a FOTO/Analyst Express-PC systems
feature FOTO/Analyst PC Image software (Fotodyne).
RNAi screen for identification genes enhancing or suppressing akir-1 phenotype
A RNAi screen was conducted on the akir-1 background (akir-1(rj1) for
chromosome II and akir-1(gk528) for chromosome IV) to discover genes affecting SC
morphogenesis. RNAi bacterial clones from the Ahringer C. elegans RNAi library (R.
Kamath 2003) for chromosomes I and IV were selected for analysis. The clones were
grown in Luria Broth (LB) media + 50ug/ml ampicillin (AMP) at 37°C for 12-16 hours.
The clone bacterial cultures were then seeded onto NGM + 50ug/ml ampicillin AMP + 1
mM isopropyl-B-D-1-thiogalactopyranoside (IPTG) plates. The pL4440 vector used to
generate the RNAi clones (Timmons and Fire 1998)was used as an empty vector control.
Liquid worm culture was conducted for part of the screen. Bacterial cultures were
inoculated into 500ul of LB+50ug/ml AMP was placed in 96-well, deep well plates and
grown at 37°C in a shaker incubator (300 rpm) overnight. The next day IPTG was added
to a final concentration of 1mM. The cultures were then placed back in the incubator to
shake for and additional hour. These plates were then spun at 4000g for 20 minutes to
concentrate the RNAi bacteria. The bacteria were resuspended in NGM media with
50ug/ml AMP and 1uM IPTG. 75uL of bacteria was aliquoted into 96-well flat bottomed
plates. Synchronized L1 (15-20) worms of the appropriate genotype were seeded into the
plates and allowed to culture for 36-40 hours at 20°C shaking at 300rpm.
For traditional plate culture, synchronized cultures of L1 larvae were generated
for the akir-1 mutant and wild-type strains. akir-1 mutant or wild-type L1 larvae (P0
generation) were placed on the NGM plated seeded with RNAi bacteria and were
cultured for about 46 hours (2.5 days) at 20°C until they reached adulthood and started to
lay eggs. The F1 generation was visually scored at L1 stage on a stereomicroscope for a
124 reduction or increase in viable progeny as compared to the controls and clones exhibiting
a change in lethality were selected for replication and further analysis.
We then conducted a fecundity study on clones that passed the initial screen
to determine if the change in viability of F1 progeny was due to meiotic or developmental
defects. To determine the fecundity, single L4 akir-1 mutant or wild-type worm was
placed on seeded NGM plate and allowed to lay eggs for a 15 hour period. The worm was
then moved to a fresh NGM plate and again allowed to lay eggs. This was repeated for a
three-day period and each experiment was preformed in triplicate (n=3). Eggs were
counted for each genotype examined then allowed to hatch. L1 larvae were counted and
permitted to grow to adults, which were again counted. We then could account for which
clones were causing meiotic defects (a reduction in the number of eggs laid) verses the
clones that were causing developmental defects (larval arrest). Clones demonstrating
meiotic defects (a reduction in the number of eggs laid) were selected for cytological
screening.
Quantification of F1 progeny for RNAi screen
The progeny for each control (pL4440 empty vector on the akir-1 mutant
and wild-type backgrounds) and the experimental clone on each background were scored
for each of the three replicate plates. The average number of progeny were calculated. for
each experimental clone the average of the experimental clone progeny was divided by
the average of the control progeny. This was done in both backgrounds to
normalize for any variability between plates.
For example, wild-type experimental F1 progeny average was 300 worms and
wild-type control F1 progeny was 300 worms. To normalize divide the experimental F1
average by the control F1 Average (300/300=1). Do the same for the akir-1 mutants:
akir-1 experimental F1 average was 50 and akir-1 control F1 average was 150. Divide
the experimental by the control (50/150=0.3). Then to determine the enhancement or
suppression ratio the normalized akir-1 value was divided by the normalized wild-type
125 value (0.3/1=0.3) for the final ratio. Under our threshold criteria, this value would be
considered an enhancer of the lethality observed in the akir-1 mutant background.
RNAi feeding protocols
RNAi clones are ground 6 -12 hours in LB+ampicillin (50ug/ml). Cultures are
then seeded onto IPTG plates (see above) and left to grow overnight (R. Kamath 2003).
Either synchronized L1 or L4 larvae were placed on the plates, left to develop to adults.
These adults were subjected to cytological analyses or their F1 progeny scored for
viability.
Microscopy
Adult hermaphrodites, 20 hour post-L4 stage, were dissected to release gonads.
DAPI and immunostaining was performed as described in (Colaiácovo et al., 2003). For
transgenic lines utilizing GFP fusions, fixation was in methanol for 1-5 minutes, then
washed and prepared for microscopy as in (Colaiácovo et al., 2003). Whole mount
worms were prepared by Carnoy’s Fixation. Anti-bodies were used at the following
dilutions: α-SYP-1, 1:500; α-SYP-4, 1:500 (M Colaiacovo) ; α-HIM-3, 1:500 (M
Zetka); α-HTP-3, 1:500 (M Zetka ); α-RAD-51 1:10,000 (ModEncode); α-dpMPK-1
1:500 (Sigma). The secondary antibodies used were: Alexa Fluor 488 α-mouse, Alexa
Fluor 488 α-rabbit Alexa Fluor 555 α-rabbit, Alexa Fluor 568 α-goat, Alexa Fluor 568
α–guinea pig (Invitrogen), and DyLight 594 α-goat (Jackson Immunochemicals, West
Grove, PA).
The images were acquired using the DeltaVision wide-field fluorescence
microscope system (Applied Precision) with Olympus 100x/1.40- or 60x numerical
aperture lenses. Optical sections were collected at 0.20-um increments with a
coolSNAPHQ camera (Photometrics) and deconvolved with softWoRx software (Applied
Precision). Gonadal and nuclei images are projections halfway through three-dimensional
data stacks (Multiple 0.2-µm slices/stack), except of where full projections are indicated,
126 and were prepared using softWoRx Explorer 1.3.0 software (Applied Precision) or
FIJI(Chen et al., 2012).
Quantification of Immunostained Foci
Strains selected for study were evaluated for SC morphology by SYP-1
immunostaining. Those strains showing defects in SC morphology were replicated and
the defect penetrance was quantified. Wild-type worms were simultaneously stained and
used as controls. Statistical comparisons between genotypes were performed using the
Fishers Exact Test, 95% confidence interval.
SYP aggregates were defined as SYP signals with width larger than that of wildtype SC. When measured, even the smallest aggregates were larger than the larger SC
width measured and above the average SC with plus 2 standard deviations. Quantitative
analysis of the intensity of SYP-1 signals was performed using FIJI (Chen et al., 2012).
This was performed under guided model option with a freehand polygon section in all Zstacks of a particular SYP signal and to multiple gonads from each genetic background.
We set a threshold of 250 for the maximum grey value being measured, to ensure
that over-exposed images were not included in the analysis. To obtain SYP-1 signal
intensity, we subtracted the background of the same image from the SYP-1 signal
intensity. We also normalized the SYP-1 signal intensity to the sum of all the background
controls also accounting for area size of the aggregates. Statistical comparisons between
genotypes were performed using the two-tailed Mann–Whitney test, 95% confidence
interval.
Quantification of RAD-51 foci was performed for all six zones composing the
premeiotic tip to late pachytene regions of the germline as in Colaiacovo et al.,2003. The
total number of nuclei (n) were scored per zone from three gonads each for wild-type,
csn-2 and csn-5 mutants. Statistical comparisons between genotypes were performed
using the two-tailed Mann–Whitney test, 95% confidence interval. Quantification of
GFP::COSA-1 was carried out as in (Yokoo et al., 2012) with zone 6 selected to be
127 analyzed. The total number of nuclei (n) were scored in zone 6 for 5 gonads. Statistical
comparisons between genotypes were performed using the two-tailed Mann–Whitney
test, 95% confidence interval.
Fluorescence in situ Hybridization (FISH)
The 5S FISH probe was generated as in (MacQueen 2002) from a PCR fragment
generated by amplifying C. elegans genomic DNA with the 5′TACTTGGATCGGAGACGGCC-3′ and 5′-CTAACTGGACTCAACGTTGC-3′ primers.
Fragments were labeled with fluorescein-12-dCTP (PerkinElmer, Waltham, MA).
Homologous pairing was monitored quantitatively as in (MacQueen 2002). The total
number of nuclei scored per zone (n) from three gonads each for wild-type, csn-2 and
csn-5 mutants. Statistical comparisons between genotypes were performed using the
Fishers Exact Test, 95% confidence interval.
Western Analyses
For each strain selected for analysis, 200 L4 homozygote larvae were picked and
aged to adults. Worms were washed twice in M9 buffer to remove as much bacteria as
possible. Gonadal proteins were extracted from these worms by boiling for 5 minutes in
1X Laemmli buffer, separated on 4-20% Tris-MOPS gradient gels (GenScript Express
Plus PAGE gels), and then transferred onto a nitrocellulose membrane. After transfer,
membranes were stained with Ponceau Stain to determine the efficiency of protein
transfer. Membranes were then rinsed with 1X PBST (PBS +1% Tween 20) and then
blocked with 5% milk for 1 hour. The membranes were then incubated with primary
antibodies (diluted in 1% Bovine Serum Albumin (BSA)) and incubated overnight at
4°C. The next day, membranes were then washed 3x10 minutes in 1X PBST, incubated
2-3 hours with secondary antibodies diluted in 5% milk, washed again 3X10 minutes
with 1X PBST then detected using an chemiluminescence kit (Bioexpress-WesternBright
ECL kit). Membranes were exposed to x-ray film (RPI-CLASSIC X-Ray film) which
was developed using an x-ray film developer (Protec).
128 Primary antibodies used in this study were mouse α-dpMPK-1, (1:1000) (Sigma)
and guinea pig α-SYP-1, (1:1000) (K Nabeshima). Secondary antibodies used were αmouse antibody or α-guinea pig conjugated to horseradish peroxidase (HRP), (1:10,000)
(COMPANY NAME). Mouse α-tubulin, (1:000)(DSHB) was used as a loading control.
Quantification was done on FIJI(Chen et al., 2012). Statistical comparisons between
genotypes were performed using the Fishers Exact Test, 95% confidence interval.
Apoptotic analyses
The csn mutants were introgressed to ced-1::GFP strain (smIs34 [ced-1p::ced1::GFP + rol-6(su1006)]) to assess apoptotic levels as per(Z. Zhou, Hartwieg, and
Horvitz 2001). Images were taken at 60x and nuclei which displayed CED-1::GFP
localization were counted as well as the total number of nuclei in the bend region (late
pachytene). 10 different gonads were quantified. Statistical comparisons between
genotypes were performed using the two-tailed Mann–Whitney test, 95% confidence
interval.
Table 4-1: Strains used in this study
Genes used
Allele Genotype
this study
fln-1
Y66H1B.2(ok2611) IV.
akir-1
129 Strain name
Reference
RB1877
Kovacevic
and Gram
2010
Clemons et al
2013
Clemons et al
2013
Grisby et al
2009
akir-1(gk528)/hT2[bli-4(e937) let?(q782] I,III
akir-1(rj1)/hT2[bli-4(e937) let?(q782] I,III
C03D6.5(ok2060) I.
SSM7
CE1255
SSM34
This study
SSM68
This Study
glh-1
cep-1(ep347) I.
csn-2(tm2823) I/hT2[bli-4(e937)
let-?(q782) qIs48](I;III).
csn-5(ok1064)
IV/nT1[qIs51](IV;V).
glh-1(gk100) I.
VC178
kgb-1
kgb-1(um3) IV.
KB3
let-92
let-92(ok1537)
IV/nT1[qIs51](IV;V).
pch-2(tm1458) II.
SSM42
Smith et all
2002
Smith et al
2002
This Study
AV307
unc-85
syp-1(me17) V/nT1[unc-?(n754)
let-? qIs50] (IV;V).
unc-85(ok2125) II.
Transgenic
lines
GFP::COSA
-1
CED1::GFP
meIs8 [pie-1p::GFP::cosa-1 + unc- CU1546
119(+)] II
smIs34 [ced-1p::ced-1::GFP + rol- AV630
6(su1006)]
akir-1
asfl-1
cep-1
csn-2
csn-5
pch-2
syp-1
SSM1
RB1662
CA388
SMM11
Balla and
Denberg 2005
MacQueen et
al 2002
Grisby et al
2009
Yokoo et all
2012
Wang et al
2010
Table 4-1continued:
Double mutants generated
csn-5;pch-2
csn-5(ok1064)
IV/nT1[qIs51](IV;V);pch2(tm1458) II
csn-5;CED-1::GFP csn-5(ok1064)
IV/nT1[qIs51](IV;V);
smIs34 [ced-1p::ced1::GFP + rol-6(su1006)]
csn-5;GFP::COSA- csn-5(ok1064)
1
IV/nT1[qIs51](IV;V);meIs
8 [pie-1p::GFP::cosa-1 +
unc-119(+)] II
csn-5;akir-1
csn-5(ok1064)
IV/nT1[qIs51](IV;V);akir1(gk528)
csn-5;cep-1
csn-5(ok1064)
IV/nT1[qIs51](IV;V);cep1(ep347) I.
csn-2;pch-2
csn-2(tm2823) I/hT2[bli4(e937) let-?(q782) qIs48]
(I;III); pch-2(tm1458) II
csn-2;CED-1::GFP csn-2(tm2823) I/hT2[bli4(e937) let-?(q782) qIs48]
(I;III); smIs34 [ced1p::ced-1::GFP + rol6(su1006)]
csn-2;GFP::COSA- csn-2(tm2823) I/hT2[bli1
4(e937) let-?(q782) qIs48]
(I;III);meIs8 [pie1p::GFP::cosa-1 + unc119(+)] II
akir-1;unc-85
akir-1(gk528)/hT2[bli4(e937) let-?(q782] I,III;
unc-85(ok2125) II.
akir-1;fln-1
akir-1(gk528)/hT2[bli4(e937) let-?(q782]
I,III;Y66H1B.2(ok2611)
IV.
130 SSM96
This study
SSM107
This study
SSM115
This study
SSM100
This study
This study
SSM98
This study
SSM108
This study
SSM54
This study
SSM29
This study
SSM43
This study
Table 4-2: Primer sequences and PCR conditions for validation of deletion strains
WildDeletion
type
Product
Conditions
Allele
Primer Sequence product
Csn-5
F GCAACAAGCTGG
1600
600
54/2:00
(ok1064)
ATAACCAGTC
R GACAGAAGTGTC
GATGGACATG
csn-2
F GATAGTGGCTCA
1200
900
54/2:00
(tm2628)
GAACCAGACG
R TACCATAACCAT
CATTGCGGCTG
akir-1
F GTCGATGTGTGG
2000
1000
56-58/2:00
(gk528)
CCTCCCGCTG
R CAAGAAATCCGT
CTGAGAAATGAG
TTC
pch-2
F ACTTTGATTTTC
997
434
54/2:00
(tm1428)
GCGGGACC
R AGCTGAAGATGA
GAAGTGCATG
glh-1
F GCGGAGAGAGA
953
370
54/1:30
(gk100)
AACAATAGTTCG
R ATCAAATTTACA
ACGAGGGAAGA
A
kgbF CGGCAAAATCAA
1510
252
52/1:30
1(um3)
TTTCCTGA
R CAAAAATGAATC
CGCCACTC
cep-1
F GAGAAATTGTCC
2800
340
55/3:00
(ep347)
CGTTCCCG
R TCAAAGCACCGC
ATTTCTCA
unc-85
F GAAACATTCACG
2180
480
59/2:00
(ok2125)
AGGGTACGCG
R CTGGTGTCTGAC
GAATGACCACA
let-92
F CCGTATCAACGT
1100
250
56/2:00
(ok1537)
TTGCCTAACAAT
R AGGATCGAGTAA
CACTGCTCC
fln-1
F TGGACGTCCTGG
2500
250
58/2:30
(ok2611)
GAAGAACT
R CTCTCCAGCTTC
ACGTGGTG
131 132 CHAPTER 5
CONCLUSIONS AND FUTURE DIRECTIONS
While the relationship between maternal age and aneuploidy in mammals has
been recognized for some time, only with recent advances have we been able to
determine that up to 20% of oocytes are aneuploid and that the majority of trisomic
fetuses are the result of maternal MI defects (Hassold and Hunt 2001; Hassold and Hunt
2009; Hunt and Hassold 2008). Despite our growing abilities to detect the outcome of
aneuploidy in humans, we still lack a firm understanding of molecular mechanisms that
give rise to maternal MI defects. However, progress is being made in elucidating the
mechanisms of aneuploidy in model organisms such as C. elegans. Our lab has already
made a contribution to the understanding of the regulation of SC disassembly through the
discovery of akir-1. This gene was identified and characterized, showing defects in SYP1 localization and bivalent structure (Clemons et al., 2013). However, questions remain
as to the molecular cause for these defects in akir-1 mutants.
To address these questions, I undertook a RNAi screen on the akir-1 mutant
background with the aim of recovering genes that interact in the akir-1 pathway and
novel genes that function in meiosis. While the screen only covered 37% of the worm
genome, six candidates were identified (Table 2-1)_. One of these candidates, csn-5, has
been a focus of my studies. Of the other candidates, two merit further study: unc-85 and
cks-1. RNAi of both of these genes in the akir-1 background lead to distinct bivalent
defects: unc-85 with a presence of univalents in D -1 oocytes and cks-1 with chromosome
condensation and SC defects. While unc-85 or its homolog Asf1 have been not
implicated in SC morphogenesis or in bivalent resolution, they have been implicated in
meiosis as the unc-85;asfl-1 double mutant has a disorganized gonadal morphology
(GRIGSBY and FINGER 2008). This suggests a role in meiosis and should be further
examined to define that role. Furthermore, the downstream target of CKS-1, CDK-1 has
133 been implicated in meiotic events in yeast including, homologous recombination, SC
disassembly and crossover formation .To determine the roles each of these genes have in
meiosis, additional analyses should be initiated including pairing analysis (FISH), meiotic
recombination analysis (RAD-51 IHC), and crossover formation (COSA-1) analysis
(each described in Chapter 4). In addition, I would also examine condensin and cohesion
through cytological analysis to determine if their localization is perturbed. Biochemical
analyses such as western analyses and immunoprecipitation should be conducted to
determine protein levels and protein interactions. The later experiments, especially the
cytology with cohesion and condensin specific antibodies, will give a better indication if
these particular proteins are disturbed in these mutants, resulting in the defects observed.
Additionally, the generation of double mutants of unc-85;akir-1 (already done), asfl1;unc-85, and cks-1;akir-1 or cdk-1;akir-1 will allow for the examination of the possible
interactions. These analyses will allow us to answer the question: do unc-85/asfl-1 or cks1 act in a pathway with akir-1 to modulate SC disassembly or do they have other novel
roles in SC morphogenesis, meiotic recombination and crossover formation.
A role for CSN/COP9 signalosome in meiotic prophase I
The original aim of the RNAi screen was to identify genes involved in SC
disassembly and I identified two interesting candidates. In addition, I recovered csn-5, a
gene encoding for an CSN/COP9 signalosome subunit, which was involved in SC
assembly. Of the six candidates listed in Table 2-1 , csn-5 was interesting in that it was
defective in assembly, not disassembly. While the screen was designed to detect genes
affecting disassembly, we recovered a gene that affected assembly. csn-5(RNAi)
increased the lethality of the akir-1 mutant background resulting in an enhancer
phenotype (ratio =0.15). The assembly defects in csn-5 mutants caused an increase in
apoptosis, but isn’t the primary reason for the lack of oocytes that we observed. Because
we did not count the number of oocytes in the csn-5(RNAi) experiments (but noted there
were more than 1), I suspect that the reduction of MAPK signaling may be the primary
134 reason for the increase of lethality. It could be that MAPK signaling does not only
facilitate the exit from pachytene but also may initiate the SC disassembly process.
Therefore, despite the screen being designed for detection of disassembly
mutants, it actually detected assembly mutants too. I used two mutants, csn-5(ok1064)
and csn-2(tm2823) to characterize how the signalosome impacted meiotic prophase I
events, which are discussed in Chapter 3. Meiotic prophase events are affected in both
these mutants; however, not to the same extent. The fact that csn-2 mutants have more
severe defects suggests a model where csn-2 is functioning within the signalosome
holocomplex, while the csn-5 is operating outside and separate from the holocomplex, in
an antagonistic manner (Figure 3-20). The data described in Chapter 3 supports this
model.
An alternative hypothesis would be that csn-2 is acting outside the signalosome
either on its own or in a subcomplex of the signalosome. Mammals have two isoforms of
CSN-2, both with different functions, CSN-2 in the holocomplex and ALIEN outside the
complex (Wei and Deng 2003). C. elegans only has one isoform, and therefore, we
hypothesize it can only function in the holocomplex. Given that the other subunits of the
signalosome can form sub-complexes, it is a possibility that csn-2 is acting outside the
holocomplex as part of a sub-complex. As yet, none of the sub-complexes found in
mammalian cell lines, Drosophila, or Arabidopsis have displayed a biological function
(Wei, Serino, and Deng 2008).
Therefore, this is a less likely explanation for the differences observed in the csn
mutants. In order to determine if the csn mutants are functioning in or outside the
holocomplex, characterization of different signalosome subunits will be necessary. My
current data suggests that csn-2 and csn-5 function in different pathways. By replicating
the analyses described in Chapter 3 with other csn mutants, we will be able to tell if these
subunits are acting as part of the entire complex or individually. This will allow us to
discern a possible mechanism for action and design the appropriate experiments to test
135 that mechanism. An allele for another CSN/COP9 subunit (csn-6) is available and it is
currently under examination in the laboratory by another graduate student who is using
the same analyses as were conducted on csn-2 and csn -5 mutants. If csn-6 data align
with csn-2 then this strengthens the argument that csn-5 is potentially acting outside the
signalosome. However, if csn-6 data aligns with csn-5 then the second model would be
supported, where csn-2 is acting outside the complex. Currently, no other csn deletion
mutants are available to study. RNAi could be used to examine the role of the other
subunits. The data show that csn-5(RNAi) is not highly penetrant by the feeding protocol;
using other RNAi for knockdown of the other subunits lead to a similar outcome .
Therefore it may be challenging to discern the function of the individual subunits or the
signalosome as a whole. However, introduction of RNAi by injection has shown high
penetrance and could be considered as an alternative for double mutant analyses.
Currently, transgenic tools such as extrachromosomal arrays are not feasible in
the study of meiosis as they are not expressed in the germline (Boulin and Hobert 2011).
Only low-copy integrated transgene can be expressed and inherited(Boulin and Hobert
2011). However, integration is not targeted by homologous recombination and therefore,
integration can occur at any place in the genome leading to lethality depending on where
the array integrates(Boulin and Hobert 2011). A better method for examine signalosome
function maybe to generate csn mutants using the CRISPR system which allows for
precise gene editing using Cas9 nuclease to generate a break in DNA targeted by a single
short RNA and then repair by either HR or NHEJ (Waaijers et al., 2013; Tzur et al.,
2013). This allows for the generation of insertions, deletions, or other mutations, which
can knockdown or knockout genes in a heritable manner. Such a tool could also be used
for genome engineering and facilitating the generation of tagged proteins by fusing the
reporters directly into the gene. Not only could precise knockouts be made for each of the
signalosome subunits, epitope tagged fusions could also be generated, eliminating the
need to design antibodies specific to the gene. This could allow for the biochemical assay
136 of CSN/COP9 signalosome function, which I currently cannot accomplish due to a lack
of sufficient amounts of antibodies for csn-5 and a lack of antibodies for csn-2. Currently,
we do not know if csn-5 and csn-2 are acting within the context of the entire signalosome
holocomplex or if they are acting independently outside the complex. Coimmunoprecipitation experiments with following by western analyses will allow for
examination of the complex and subunits. It is worth noting that there is evidence that
subcomplexes do exist. This would have to be done in conjunction with the examination
of other csn subunits to determine the extent of complex association.
The role of the CSN/COP9 signalosome in MAPK signaling
The differences observed between the csn-5 and csn-2 in pairing, recombination,
and crossover formation suggested they might differ in the lack of MAPK signaling in the
proximal gonad. Both the cytological and western analyses show the two alleles
completely lacked dpMPK-1 in the germline, rendering both mutants sterile. These data
support a function for both alleles in the signalosome holocomplex in MAPK signaling. It
is known that the signalosome can associate with kinases (Wei, Serino, and Deng 2008)
and the signalosome is facilitating signal transduction through an interaction with
kinases. To date, there is no in vivo evidence confirming this role. Determining the
interactions between the signalosome and the known members of the MAPK pathway is
essential to determine the role of the signalosome in the MAPK pathway. This could be
facilitated by yeast 2-hybrid or co-immunopreciptation experiments to determine
interactions with MAPK signaling pathway components such as MPK-1.
Dissecting the role of the signalosome in MAPK signaling would not only be
novel, but allow for the examination of the role of MAPK signaling in the exit from
pachytene. While csn-5 was identified as an enhancer in the RNAi screen, the
enhancement of lethality was not directly due to an interaction with akir-1, but rather a
lack of oocytes as the result of the absence of MAPK signaling. It could be that the
signalosome acts upstream of akir-1 in the initiation of disassembly. At this time, we
137 cannot study the affects of the signalosome on SC disassembly as so few oocytes advance
to that point.
Determining the substrates of the signalosome regulating SC formation
There are also pre-meiotic proliferation defects in the csn mutants as evidenced by
the shorter PMTs and the over lack of nuclei in the gonads. Given the possible
involvement of the CULLIN E3 ligases (CRLs) in PMT proliferation, this raises the
question: what substrates does the signalosome act upon and how are they modified.
The CSN/COP9 signalosome has been shown to directly interact with the CRLs,
regulating their function through cycles of neddylation and deneddylation. CRLs have
two functions: monoubiquitination of substrates, which facilitates localization and protein
interactions (Zhong et al., 2003; Bosu and Kipreos 2008; Chiba and Tanaka 2004). It
could be that CRLs are acting in this manner, where the substrate could be either the SYP
proteins themselves or another protein that interacts with the SYPS to ensure proper
polymerization and loading onto the lateral elements in forming the SC. In contrast, it
could be there is a potential regulatory protein that inhibits SYP polymerization outside
the context of SC assembly. If this protein relies on monoubiquitination for interaction
with the SYPs, this interaction could be abolished due to a non-functional CRL.
The majority of studies have focused on the signalosome-CRL-proteasome
pathways (Cope and Deshaies 2003; Wei and Deng 2003). It is possible that in the
absence of functional signalosome, the CRL-proteasome pathway is disrupted, leading to
the defects in SC assembly. It could be that there are key proteins that need to be tightly
regulated through degradation to prevent SYP polycomplex formation. Disruption of this
regulation would lead to SYP polycomplex formation presumably leaving not enough SC
assembled on chromosomes to stabilize the pairing interaction.
The signalosome could directly interact with the SYPs. As previously discussed,
the signalosome associates with several kinases, and there could be a direct interaction
with the SYPs. Alternatively, the SYPs could be neddylated as the CRLs are or
138 ubiquitinated by the CRLs. It could be through these associations, that SYPs are directly
modified. It will be key to determine how the SYPs are regulated in order to define the
role of the signalosome in preventing their aggregation during assembly. Ectopic
expression of these proteins leads to polycomplex formation, therefore there should be a
mechanism by which they are prevented from aggregating. There are numerous sites for
phosphorylation, sumoylation, and ubiquitination on all of the SYP proteins (Table 3-8),
yet very little is currently known about if these post-transcriptional modifications are
present and if present how they affect the function of the SYP proteins.
There are six CRLs found in C. elegans. In addition, we have initiated studies to
examine how the depletion of NED-8 and UBA-1 will impact SC morphogenesis.
Depletion of NED-8 should render all the CRLs non-functional. The CRLs appear to
have redundant functions, therefore, knockout of NED-8 should render all CRLS nonfunctional and eliminate the redundancy issue. Knocking out NED-8, UBA-1 or the
CRLs would likely lead to proliferation issues, given the roles that CRLs have in
proliferation. However, given the importance of the CRLs-proteasome pathway in
development, the worms would likely display a larval lethal phenotype. We already know
that ned-8 and uba-1 deletion strains are larval lethal. UBA-1 is the only ubiquitinactivating enzyme in C. elegans, therefore depletion should knockout all ubiquitin
pathways which results in lethality. To bypass the somatic affects of these proteins and to
enable study in the germline, we generated somatic rescue lines. I designed extra
chromosomal arrays for the wild-type ORFs and regulatory regions and these arrays
were injected into the ned-8 and uba-1 deletion mutants by Carlos Carvalho (University
of Saskatchewan). The expression of the arrays in the somatic tissues allowed for the
bypass of the larval lethality while allowing for the retention of the knockout of the gene
of interest in the gonad as the arrays will not be expressed in that tissue, thus allowing us
to determine if the CRLs are involved in the SC assembly. If we see aggregation of the
SYPs in either of the rescue lines would indicate a role in SC assembly and therefore
139 indicate to us if the CSN/COP9 signalosome is interacting indirectly through the CRLs,
although epistatic analyses with csn mutant will be needed to confirm this is indeed the
pathway by which SC assembly is regulated. An absence of aggregates would suggest
that the signalosome has a more direct influence on SC assembly. This work is being
completed by Nathan Balukoff.
In conclusion, my genetic screening approach led to identification of six potential
genes acting in SC assembly/disassembly and/or bivalent structure. My study of one of
these genes lead to identification of a novel complex acting in SC assembly in meiosis:
the CSN/COP9 signalosome. The csn mutants, csn-2 and csn-5 have roles in pairing
stabilization, meiotic recombination, and crossover formation. They abolish MAPK
signaling, leading to pachytene arrest, which results in sterility. While this study defines a
role for the signalosome in meiotic prophase I events, the exact mechanisms by which the
signalosome functions in this role are still unclear.
140 APPENDIX A
RNAI SCREEN METADATA
Table A-1: Chromosome II RNAi clones that were quantified
Wild-type
akir-1(rj1)
Average
Average
Chrom II
WT/
# of
stdev
# of
stdev
Clone ID
94.9
progeny
progeny
B0454.3
146.50
9.09
1.36
0.63
1.54
C08F1.7
126.10
27.22
2.27
2.80
1.33
C18A3.5
222.00
20.30
2.06
1.23
2.34
C32D5.12
314.33
191.0
5.50
1.50
3.31
6
C41H7.1
89.92
25.92
1.81
0.64
0.95
C52E2.I
128.57
67.00
1.58
0.52
1.35
D2062.2
108.11
39.80
1.67
0.58
1.14
F22E5.2
100.00
27.24
1.33
0.58
1.05
F36H5.4
118.93
50.10
1.92
0.72
1.25
F42G2.2
134.00
27.62
2.19
1.56
1.41
F42G2.4
50.60
12.02
1.00
0.00
0.53
F42G2.6
134.97
49.66
1.89
1.02
1.42
F45C12.14 126.31
27.34
1.50
0.25
1.33
F45C12.2
123.18
31.23
1.93
1.22
1.30
F59H6.2
103.75
39.05
1.04
0.32
1.09
H41C03.2
221.58
48.96
4.42
2.92
2.33
K12H6.12
137.75
79.77
2.53
1.72
1.45
T08E11.4
150.83
5.64
2.17
0.29
1.59
B0047.2
168.08
41.90
3.83
1.04
1.77
C06A1.1
39.85
26.41
0.89
0.19
0.42
C17C3.8
300.00
130.2
9.31
0.76
3.16
2
C33F10.12 341.08
159.3
9.92
4.08
3.59
8
C40A11.2
89.55
35.45
2.89
2.99
0.94
C52E2.G
79.56
44.31
2.44
2.50
0.84
F08G2.2
55.39
29.13
1.86
2.44
0.58
F16G10.6
128.80
23.34
3.61
2.99
1.36
F16G10.7
95.58
83.00
2.62
2.26
1.01
F28C6.9
180.33
28.04
5.67
4.31
1.90
F29A7.3
109.66
38.08
3.33
4.04
1.16
F36H5.9
137.13
33.12
4.25
4.34
1.45
F45C12.12
83.25
32.25
1.94
1.29
0.88
0.10
0.17
0.16
0.42
akir1(rj1)
/WT
0.1
0.1
0.1
0.1
0.14
0.12
0.13
0.10
0.15
0.17
0.08
0.14
0.11
0.15
0.08
0.34
0.19
0.16
0.29
0.07
0.71
0.1
0.1
0.1
0.1
0.1
0.1
0.1
0.1
0.1
0.1
0.1
0.1
0.1
0.1
0.2
0.2
0.2
0.75
0.2
0.22
0.19
0.14
0.27
0.20
0.43
0.25
0.32
0.15
0.2
0.2
0.2
0.2
0.2
0.2
0.2
0.2
0.2
akir-1/
13.18
Table A-1: continued F45C12.5
65.71
F58E1.2
103.89
F59H6.12
59.75
K05F1.4
177.75
T08E11.2
113.45
T23B7.1
470.33
Y59C2A.3
ZK250.5
B0047.5
C17G10.5
C25H3.10
C33C12.4
C49D10.1
EEED8.1
F11G11.13
F59E12.11
F59E12.4
K05F1.6
R52.5
Y47G7B.2
ZK546.13
AH6.9
B0281.2
C25H3.1
C32D5.3
C34F11.3
C40A11.7
C40A11.9
C41C4.8
E02H1.1
F07F6.1
F09C12.7
F26G1.5
F40H7.9
F43C1137.
F
F58F12.1
F59E12.12
K07D4.3
R05F9.6
141 2.05
2.83
1.55
3.92
2.89
11.17
1.17
3.18
0.64
1.66
2.01
5.09
0.69
1.09
0.63
1.87
1.20
4.96
0.16
0.21
0.12
0.30
0.22
0.85
0.2
0.2
0.2
0.2
0.2
0.2
103.47
101.35
126.30
180.80
242.67
166.56
48.72
121.53
206.94
94.33
241.44
196.00
126.25
131.11
247.67
88.67
180.72
259.31
118.67
198.67
22.11
144.67
32.59
95.89
90.31
147.89
300.83
52.00
84.06
15.50
32.33
44.27
11.35
15.41
185.8
0
24.81
43.24
4.36
79.02
57.87
16.50
27.13
37.63
66.94
8.96
68.69
4.58
35.20
33.72
47.08
76.64
71.04
74.80
48.68
64.07
12.63
23.29
17.75
12.17
14.57
37.27
96.60
12.00
26.04
3.44
2.89
5.00
7.08
11.17
5.92
1.75
5.17
9.17
3.67
11.67
8.38
5.72
6.33
10.72
5.33
11.22
14.69
6.17
12.17
1.28
7.75
1.89
5.44
5.33
7.78
18.13
2.84
4.22
2.69
1.02
2.00
1.23
6.93
4.35
0.90
4.19
9.67
2.02
6.60
2.06
3.69
3.69
2.63
0.33
6.01
5.75
2.47
1.59
0.25
5.45
1.84
1.71
0.58
2.06
8.56
2.91
3.02
1.09
1.07
1.33
1.91
2.56
1.76
0.51
1.28
2.18
0.99
2.54
2.07
1.33
1.38
2.61
0.93
1.90
2.73
1.25
2.09
0.23
1.52
0.34
1.01
0.95
1.56
3.17
0.55
0.89
0.26
0.22
0.38
0.54
0.85
0.45
0.13
0.39
0.70
0.28
0.89
0.64
0.43
0.48
0.81
0.40
0.85
1.11
0.47
0.92
0.10
0.59
0.14
0.41
0.40
0.59
1.38
0.22
0.32
0.2
0.2
0.3
0.3
0.3
0.3
0.3
0.3
0.3
0.3
0.3
0.3
0.3
0.3
0.3
0.4
0.4
0.4
0.4
0.4
0.4
0.4
0.4
0.4
0.4
0.4
0.4
0.4
0.4
89.00
161.67
88.36
75.11
30.16
94.38
15.07
21.72
4.75
10.00
5.14
3.89
2.70
1.00
3.89
0.96
0.94
1.70
0.93
0.79
0.36
0.76
0.39
0.30
0.4
0.4
0.4
0.4
Table A-1: continued C07E3.5
107.58
C08F1.5
59.52
C08H9.5
153.83
C17G10.6
206.76
F10G7.4
100.89
F22E5.13
124.08
Y62F5A.E
79.78
ZK177.8
103.67
F10G7.3
100.00
C17C3.4
225.61
C25H3.7
208.50
C33C12.5
51.44
EEED8.6
74.67
F12A10.4
81.00
F33G12.5
107.33
F58E1.6
51.81
F59E12.10 120.94
K12H6.6
43.83
ZK546.7
80.00
B0454.6
49.33
F22E5.12
112.75
F59B10.9
119.64
K05F1.7
245.50
K09F6.6
126.92
R11F4.2
83.17
T13H5.5
46.17
Y49F6C.4
110.72
ZK673.9
138.11
C27D6.6
172.13
C30B5.7
233.50
C49D10.9
112.33
C52E2.F
90.50
F35C5.3
79.89
F56D1.1
360.00
K05F1.8
294.33
M176.6
130.00
Y38F1A.1
163.53
ZK1062.4
140.33
ZK177.10
96.17
B0304.5
120.89
24.89
8.27
64.87
71.54
34.80
7.89
22.74
14.64
34.80
92.68
15.22
35.18
28.10
45.00
71.11
78.02
95.78
36.85
4.00
8.11
34.07
25.11
30.33
16.06
87.40
13.59
28.02
5.87
24.43
51.80
51.73
14.01
15.78
91.93
54.08
67.02
29.25
48.25
50.45
38.72
7.67
4.50
10.17
14.12
7.45
8.00
5.17
7.00
7.45
18.97
18.43
4.22
6.44
6.83
8.33
4.33
10.42
3.44
6.28
4.83
10.33
11.11
23.97
12.44
8.58
4.50
10.94
13.22
18.53
24.42
13.25
9.56
8.61
42.44
32.83
14.83
17.56
15.00
10.56
14.44
8.08
1.32
7.22
7.02
1.70
9.54
3.44
3.28
1.70
8.69
14.59
0.69
2.68
3.75
10.10
5.39
11.02
2.91
4.33
2.75
5.53
3.98
10.23
8.58
6.61
2.78
3.73
1.02
0.95
11.32
5.85
7.50
6.88
10.49
10.76
0.76
11.57
9.67
3.86
6.32
1.13
0.63
1.62
2.18
1.06
1.31
0.84
1.09
1.05
2.38
2.20
0.54
0.79
0.85
1.13
0.55
1.27
0.46
0.84
0.52
1.19
1.26
2.59
1.34
0.88
0.49
1.17
1.46
1.81
2.46
1.18
0.95
0.84
3.79
3.10
1.37
1.72
1.48
1.01
1.27
142 0.58
0.34
0.77
1.07
0.57
0.61
0.39
0.53
0.57
1.44
1.40
0.32
0.49
0.52
0.63
0.33
0.79
0.26
0.48
0.37
0.78
0.84
1.82
0.94
0.65
0.34
0.83
1.00
1.41
1.85
1.01
0.73
0.65
3.22
2.49
1.13
1.33
1.14
0.80
1.10
0.5
0.5
0.5
0.5
0.5
0.5
0.5
0.5
0.5
0.6
0.6
0.6
0.6
0.6
0.6
0.6
0.6
0.6
0.6
0.7
0.7
0.7
0.7
0.7
0.7
0.7
0.7
0.7
0.8
0.8
0.8
0.8
0.8
0.8
0.8
0.8
0.8
0.8
0.8
0.9
Table A-1: continued C33F10.2
235.25
F12A10.5
136.56
F22E5.14
99.67
F40H3.3
82.93
F54D5.4
90.40
K09F6.3
27.33
K10B2.2
261.67
R12C12.4
W03C9.5
ZK546.16
ZK546.6
AH6.1
C09G5.4
C17G10.7
C32D5.10
EEED8.8
91.00
94.03
218.90
72.89
64.73
107.36
100.00
218.50
376.89
K10B2.5
T010D4.11
T06D8.9
T09A5.5
Y53C12A.
4
ZC239.6
EEED8.13
F31D5.2
K07D4.8
R12C12.6
T07A11.6
C34C6.6
F56D3.2
T12C9.4
T25E4.2
Y53C1B.2
C09F9.4
C49D10.2
C17G3.10
C24H12.6
F07F6.4
F12E12.B
67.33
63.06
136.52
52.33
80.29
10.93
22.55
22.30
32.07
24.19
11.72
146.4
9
17.06
13.13
20.94
25.45
54.37
14.14
56.79
51.12
186.0
2
26.10
23.01
7.17
9.02
9.32
169.00
64.67
104.33
79.33
116.00
113.14
39.67
109.47
103.17
183.31
80.00
66.00
47.33
131.17
97.78
65.50
64.33
74.00
24.19
42.50
25.40
33.94
17.78
18.35
65.36
14.19
47.62
17.37
8.54
19.01
36.03
5.85
28.54
9.53
143 30.64
17.67
12.56
10.33
11.28
3.33
33.42
1.88
10.07
2.34
7.02
6.42
3.18
6.83
2.48
1.44
1.05
0.87
0.95
0.29
2.76
2.32
1.34
0.95
0.78
0.86
0.25
2.54
0.9
0.9
0.9
0.9
0.9
0.9
0.9
11.50
11.11
27.29
9.56
9.39
15.28
13.33
31.75
51.44
3.04
1.71
9.36
8.26
2.34
10.67
5.69
7.63
32.72
0.96
0.99
2.31
0.77
0.68
1.13
1.05
2.30
3.97
0.87
0.84
2.07
0.73
0.71
1.16
1.01
2.41
3.90
0.9
0.9
0.9
0.9
1.0
1.0
1.0
1.0
1.0
8.94
8.60
19.61
7.47
11.00
2.86
3.31
8.18
5.62
2.65
0.71
0.66
1.44
0.55
0.85
0.68
0.65
1.49
0.57
0.83
1.0
1.0
1.0
1.0
1.0
24.30
9.72
16.10
11.98
17.61
17.06
6.43
18.60
16.56
30.78
13.69
12.00
8.61
25.33
18.81
12.33
12.67
15.00
0.54
4.50
9.21
2.37
13.39
6.23
9.72
15.72
10.01
3.39
2.78
4.32
12.22
4.19
4.51
5.36
1.78
0.68
1.10
0.84
1.22
1.19
0.42
1.15
1.09
1.93
0.84
0.70
0.50
1.38
1.03
0.69
0.68
1.84
0.74
1.22
0.91
1.34
1.29
0.49
1.41
1.26
2.34
1.04
0.91
0.65
1.92
1.43
0.94
0.96
1.0
1.1
1.1
1.1
1.1
1.1
1.2
1.2
1.2
1.2
1.2
1.3
1.3
1.4
1.4
1.4
1.4
Table A-1: continued F43C1137. 100.61
H
M01D1.8
100.33
W10G11.1
32.80
3
Y57A10A.
62.44
A
ZK177.6
75.00
C01F1.2
50.44
D2013.5
73.53
F19H8.1
137.61
W07E6.1
94.00
C06C3.4
3.69
Y53C12A.
47.50
1
T05A7.5
121.89
ZK546.16
147.53
F09E5.3
80.00
F19B10.6
79.89
Y74E4A.A
40.67
B0286.2
52.66
C01B12.8
106.67
C32B5.15
106.67
F18A12.5
79.76
F42A8.1
42.15
R05F9.4
77.42
Y49F6B.M
28.67
Y54C5B.1
70.33
Y49F6B.R
23.58
C32D5.6
110.78
C41C4.4
95.00
T02G5.5
70.11
T25D3.3
105.56
ZK1290.8
102.11
C08E3.10
104.11
F56D3.1
83.67
T02H6.1
93.33
Y27F2A.B
40.00
T09A5.11
22.89
F31D5.3
58.20
F44G4.8
95.25
144 18.81
19.56
13.48
1.06
1.48
1.4
3.84
2.88
19.22
6.58
5.51
2.57
1.06
0.35
1.46
0.50
1.4
1.4
9.05
11.71
6.44
0.66
0.89
1.4
30.51
22.19
11.21
32.59
12.00
3.74
3.75
14.17
10.28
15.34
28.62
20.00
0.83
10.40
3.82
9.17
7.96
7.05
9.01
0.14
5.89
0.79
0.53
0.77
1.45
0.99
0.04
0.50
1.07
0.78
1.16
2.17
1.52
0.06
0.79
1.4
1.5
1.5
1.5
1.5
1.6
1.6
7.57
37.38
52.57
39.12
20.23
30.04
9.28
9.28
9.75
0.88
7.23
4.16
32.89
20.22
12.08
23.70
5.32
9.62
17.93
3.89
30.68
8.82
7.21
8.22
21.38
37.23
28.78
33.89
20.11
19.67
10.43
13.89
29.11
29.78
22.00
11.86
21.08
7.78
19.85
6.89
37.50
32.31
23.33
35.83
33.78
35.50
29.25
31.83
13.64
8.36
22.50
36.93
6.70
7.71
13.04
3.46
5.76
10.78
14.34
15.05
13.11
6.60
13.00
0.38
7.44
2.15
13.70
3.26
4.16
21.46
11.54
17.33
3.63
8.61
4.88
6.44
1.74
12.74
1.28
1.55
0.84
0.84
0.43
0.55
1.12
1.12
0.84
0.44
0.82
0.30
0.74
0.25
1.17
1.00
0.74
1.11
1.08
1.10
0.88
0.98
0.42
0.24
0.61
1.00
2.18
2.57
1.53
1.49
0.79
1.05
2.21
2.26
1.67
0.90
1.60
0.59
1.51
0.52
2.85
2.45
1.77
2.72
2.56
2.69
2.22
2.42
1.03
0.63
1.71
2.80
1.7
1.7
1.8
1.8
1.8
1.9
2.0
2.0
2.0
2.0
2.0
2.0
2.0
2.1
2.4
2.4
2.4
2.4
2.4
2.5
2.5
2.5
2.5
2.6
2.8
2.8
Table A-1: continued W05H5.4
91.60
ZK970.4
56.59
ZK131.4
73.58
C18E9.5
71.78
C50D2.2
108.78
H20J04.B
34.84
T02H7.3
81.22
C08B11.3
95.78
K01C8.9
2.22
K07E8.10
98.89
F39E9.2
54.11
C17A2.6
38.67
K12D12.1
9.22
C46E10.4
17.78
Y110A2A1
78.83
898.D
C18H9.3
76.00
T01D1.6
95.00
ZC101.2
9.82
W03H9.4
12.33
K01C8.6
9.86
R07C3.7
75.00
Y48C3A.1
54.75
ZK430.1
42.00
T19D12.6
86.67
Y46G5.K
48.10
K
C54C9.1
13.56
W03C9.3
28.77
B0286.4
6.13
W10G11.1
28.00
C17F4.4
27.44
D2013.7
0.78
D2085.1
11.61
ZK131.7
0.69
F54A331.
0.78
E
ZK430.2
25.78
D2085.3
0.77
F57C2.3
49.33
145 14.21
10.53
20.84
2.04
4.67
14.56
17.06
7.31
2.41
6.74
76.53
9.02
11.97
23.86
27.19
35.50
21.92
30.62
31.00
47.17
15.00
34.83
43.31
1.03
46.33
26.58
19.22
4.98
10.80
47.80
1.89
5.94
7.18
13.47
0.76
3.21
23.26
24.67
0.45
32.65
8.84
1.84
4.06
2.88
14.51
0.97
0.60
0.78
0.76
1.15
0.37
0.86
1.01
0.02
1.04
0.57
0.41
0.10
0.19
0.83
2.69
1.66
2.32
2.35
3.58
1.14
2.64
3.29
0.08
3.52
2.02
1.46
0.38
0.82
3.63
2.8
2.8
3.0
3.1
3.1
3.1
3.1
3.3
3.3
3.4
3.5
3.6
3.9
4.4
4.4
30.18
10.93
0.55
14.98
9.97
30.05
20.25
3.46
24.89
11.35
47.17
60.00
6.30
8.48
7.11
58.50
42.39
32.71
69.50
38.70
8.10
4.58
3.80
2.20
5.58
28.54
17.17
18.63
37.01
7.47
0.80
1.00
0.10
0.13
0.10
0.79
0.58
0.44
0.91
0.51
3.58
4.55
0.48
0.64
0.54
4.44
3.22
2.48
5.27
2.94
4.5
4.5
4.6
5.0
5.2
5.6
5.6
5.6
5.8
5.8
22.33
13.55
8.55
15.38
21.20
0.38
7.06
0.10
0.19
11.11
25.05
5.58
26.67
32.22
0.93
15.30
1.08
1.30
2.01
5.58
4.58
6.11
3.67
0.12
9.73
0.14
0.26
0.14
0.30
0.06
0.30
0.29
0.01
0.12
0.01
0.01
0.84
1.90
0.42
2.02
2.44
0.07
1.16
0.08
0.10
5.9
6.3
6.6
6.9
8.5
8.6
9.5
11.3
12.0
8.57
0.25
12.10
45.96
1.53
119.83
11.58
1.10
138.9
0
0.27
0.01
0.52
3.49
0.12
9.09
12.8
14.4
17.5
Table A-1: continued Y25C1A.5
0.28
W01D2.1
0.62
C26D10.2
0.25
R53.4
0.85
Y110A2A5
0.47
4.I
F44F4.2
1.17
F22B5.2
0.67
146 0.25
0.13
0.08
0.13
0.21
0.83
2.22
1.08
3.72
2.12
0.29
1.42
0.14
2.41
0.56
0.00
0.01
0.00
0.01
0.00
0.06
0.17
0.08
0.28
0.16
21.6
25.9
31.2
31.5
32.3
0.52
0.29
6.03
5.15
6.47
1.69
0.01
0.01
0.46
0.39
37.2
55.6
Table A-2: Chromosome IV RNAi clones that were quantified
Wild-type
akir-1(gk528)
Average
Average
Chrom IV
WT/ akir# of
stdev
# of
stdev
Clone ID
133.2 1/29
progeny
progeny
Y11D7A.4
215.91
49.42 2.80
2.40
1.62 0.10
Y38C1AB.e 344.57
167.69 5.54
1.44
2.59 0.19
T19E7.2
172.03
23.16 3.06
1.15
1.29 0.11
F55G1.10
215.86
73.57 4.03
1.64
1.62 0.14
C25A8.1
251.90
29.49 4.76
3.65
1.89 0.16
E04A4.8
164.40
87.58 3.38
1.29
1.23 0.12
M03D4.1
129.71
35.42 2.72
1.55
0.97 0.09
Y17G98.e
196.76
57.80 4.37
1.17
1.48 0.15
C46G7.1
42.57
66.40 1.06
0.10
0.32 0.04
C01F6.8
108.63
20.23 2.69
1.25
0.82 0.09
F54E12.4
105.00
20.22 2.67
1.15
0.79 0.09
F58F6.3
295.50
52.19 7.54
4.59
2.22 0.26
F18F11.4
274.07
68.12 7.59
1.97
2.06 0.26
T09A12.4
213.44
113.02 6.05
6.76
1.60 0.21
ZK180.4
143.90
63.77 4.18
2.26
1.08 0.14
K08B4.1
186.73
17.32 5.44
1.48
1.40 0.19
F33D4.2
126.44
124.02 3.89
4.86
0.95 0.13
F58E2.9
178.05
19.35 5.60
1.82
1.34 0.19
F36A4.7
69.59
12.29 2.26
0.42
0.52 0.08
B0547.1
225.77
65.01 7.37
0.90
1.69 0.25
T07A9.11
156.74
25.13 5.14
4.36
1.18 0.18
H23C24.c
161.38
93.13 5.31
2.15
1.21 0.18
D2096.3
292.78
127.26 10.06
3.27
2.20 0.35
F08B4.5
230.69
83.07 7.93
3.49
1.73 0.27
C31H1.3
323.28
97.21 11.28
2.54
2.43 0.39
C44B12.7
210.47
29.87 8.15
3.72
1.58 0.28
C06G3.8
165.29
52.05 6.43
2.00
1.24 0.22
C28C12.8
152.00
28.58 5.99
1.79
1.14 0.21
Y57G11C.1
158.60
55.25 6.27
2.15
1.19 0.22
6
B0546.3
242.81
22.39 9.91
1.17
1.82 0.34
W02A2.7
82.62
54.47 3.46
1.92
0.62 0.12
F49C12.12 157.52
25.85 6.68
7.07
1.18 0.23
K08B4.2
207.47
49.49 9.05
2.86
1.56 0.31
C28C12.2
190.95
165.46 8.52
1.84
1.43 0.29
M02B7.1
245.46
53.99 11.03
3.31
1.84 0.38
147 akir1(gk528)
/WT
0.06
0.07
0.08
0.09
0.09
0.09
0.10
0.10
0.11
0.11
0.12
0.12
0.13
0.13
0.13
0.13
0.14
0.14
0.15
0.15
0.15
0.15
0.16
0.16
0.16
0.18
0.18
0.18
0.18
0.19
0.19
0.19
0.20
0.21
0.21
Table A-2: continued W03B1.1
259.48
C33D9.2
190.78
Y55H10A.2 194.57
K02B2.5
183.80
F38A1.8
112.93
Y41D4A_34
25.14
57.D
C31H1.7
183.14
T22D1.10
232.00
F41H10.7
146.92
C09G4.3
65.22
Y52D5A.C 196.24
T26A8.1
206.37
C06E4.2
178.08
K11H12.7
260.75
K07F5.13
128.19
E04A4.5
192.43
K07H8.9
177.55
C24D10.2
208.25
C43G2.5
225.97
T11B7.1
187.74
ZK381.4
211.77
F38A5.5
164.37
Y73B6A.a
247.16
F54E12.1
66.57
F44E8.2
245.15
H35B03.2
250.99
C33H5.7
227.89
Y66H1B.2
60.57
C02B10.5
147.73
F45E4.3
222.22
T07A9.10
129.00
F55G11.5
127.38
B0212.3
196.01
Y62E10A.
180.33
M
VZK822L.1 105.96
F11E6.5
163.51
Y38C1BA.c 163.03
C55F2.2
156.64
T06C10.2
230.96
148 62.68
40.47
49.94
21.13
62.02
11.75
8.66
9.02
8.56
5.39
1.41
1.71
2.45
5.52
1.29
1.95
1.43
1.46
1.38
0.85
0.41
0.30
0.31
0.30
0.19
0.21
0.21
0.21
0.21
0.22
23.38
1.20
0.20
0.19
0.04
0.22
31.60
14.18
22.05
45.52
23.05
48.78
51.55
9.74
14.19
12.60
38.74
39.45
30.49
114.55
68.61
44.11
29.24
18.31
62.57
66.73
26.50
37.62
51.23
39.58
44.94
36.29
24.60
8.76
11.33
7.21
3.23
9.73
10.28
8.95
13.33
6.88
10.35
9.93
11.87
12.93
10.79
12.38
9.61
14.74
4.02
15.06
16.14
14.67
4.00
9.81
14.93
9.06
8.99
13.97
0.84
2.04
4.11
2.04
2.24
7.86
1.66
1.99
3.45
1.00
3.97
2.24
0.78
3.12
3.83
1.69
3.09
0.60
0.95
4.35
4.73
1.49
6.00
2.24
2.19
8.04
2.87
1.37
1.74
1.10
0.49
1.47
1.55
1.34
1.96
0.96
1.44
1.33
1.56
1.70
1.41
1.59
1.23
1.86
0.50
1.84
1.88
1.71
0.45
1.11
1.67
0.97
0.96
1.47
0.30
0.39
0.25
0.11
0.34
0.35
0.31
0.46
0.24
0.36
0.34
0.41
0.45
0.37
0.43
0.33
0.51
0.14
0.52
0.56
0.51
0.14
0.34
0.51
0.31
0.31
0.48
0.22
0.22
0.23
0.23
0.23
0.23
0.23
0.23
0.25
0.25
0.26
0.26
0.26
0.26
0.27
0.27
0.27
0.28
0.28
0.30
0.30
0.30
0.30
0.31
0.32
0.32
0.33
34.03
12.88
1.63
1.35
0.44
0.33
19.48
32.86
28.54
41.00
34.22
7.58
11.83
11.81
11.55
17.06
5.50
3.99
1.80
2.19
1.52
0.80
1.23
1.22
1.18
1.73
0.26
0.41
0.41
0.40
0.59
0.33
0.33
0.33
0.34
0.34
Table A-2: continued C18H7.4
181.62
Y38F2A_57
197.98
43.i
JC8.3
169.03
Y41D4A_31
238.02
92.b
C31H1.8
203.84
C42D4.8
126.80
F20D12.1
159.04
F41A4.2
127.15
C49H3.11
44.22
K07H8.5
185.50
Y73F8A.q
166.56
Y38F2A_61
163.90
26.F
W09G12.3
172.92
F42C5.1
72.60
C06A12.4
171.90
C52D10.3
250.99
K08D12.h
96.32
C48A7.1
158.33
Y37E11A_9
200.00
3.F
F37C4.6
206.28
Y41E3.10
193.81
JC8.5
128.60
C47E12.1
140.76
Y45F10A.5 227.10
B0564.1
184.69
F26D10.3
42.81
Y66H1A.4
147.19
Y104H12D.
164.20
b
Y45F10D.1
135.90
2
C44B12.1
103.29
Y59E9_122.
250.95
A
Y45F10A.6 188.17
Y116A8B.5 200.94
T22D1.9
164.11
149 20.13
13.42
7.30
1.36
0.46
0.34
26.10
14.92
1.68
1.49
0.51
0.35
18.73
12.99
1.64
1.27
0.45
0.35
104.62 18.33
5.41
1.79
0.63
0.35
65.44
46.88
13.75
32.25
49.26
68.88
42.17
15.79
10.21
12.83
10.33
3.60
15.31
14.15
7.52
1.82
3.82
3.61
1.81
2.31
1.45
1.53
0.95
1.19
0.95
0.33
1.39
1.25
0.54
0.35
0.44
0.36
0.12
0.53
0.49
0.36
0.37
0.37
0.37
0.37
0.38
0.39
56.80
13.99
2.43
1.23
0.48
0.39
13.37
54.68
14.99
37.30
6.34
60.55
14.80
6.37
15.30
22.52
8.70
14.55
3.73
4.27
4.36
2.89
3.01
4.31
1.30
0.55
1.29
1.88
0.72
1.19
0.51
0.22
0.53
0.78
0.30
0.50
0.39
0.40
0.41
0.41
0.41
0.42
51.07
18.47
2.07
1.50
0.64
0.42
41.34
35.21
10.50
7.24
59.78
46.79
40.58
34.12
19.52
18.60
12.42
14.12
22.85
18.86
4.39
15.50
7.75
1.26
3.00
11.30
2.58
9.82
5.73
1.17
1.55
1.46
0.97
1.06
1.70
1.39
0.32
1.11
0.67
0.64
0.43
0.49
0.79
0.65
0.15
0.53
0.43
0.44
0.44
0.46
0.46
0.47
0.47
0.48
15.58
17.50
0.77
1.23
0.60
0.49
36.20
14.61
1.09
1.02
0.50
0.49
14.79
11.23
2.35
0.78
0.39
0.50
89.73
28.37
4.94
1.88
0.98
0.52
10.92
40.99
62.07
21.51
23.16
18.96
3.05
1.28
6.36
1.41
1.51
1.23
0.74
0.80
0.65
0.53
0.53
0.53
Table A-2: continued Y45F10B.1
164.20
2
Y57G11C.5 218.02
C55C3.5
78.43
F02H6.7
250.31
Y11D7A.9
85.26
K08F11.4
132.67
Y38F2A_61
149.83
26.1
Y46C8_97.
167.57
A
Y41D4B_93
187.11
0.C
F09E8.6
230.80
Y105C5A.C 187.41
Y67D8A_38
184.89
4.6
Y62E10A.e 129.85
Y57G11C.1
157.37
2
H02112.1
92.24
C04G6.3
190.22
Y7A9B.1
196.23
Y45F10B.3 155.96
F20C5.1
105.05
R07H5.1
113.20
Y37A1B.7
171.03
R05A10.5
263.80
K03D3.3
150.89
T25B9.6
124.57
Y116A8A.7 191.39
K09B11.1
157.71
AC8.1
80.71
Y73F8A.G 200.07
Y73F8A.A
182.82
Y41D4A_30
76.80
73.A
K03D3.4
186.16
Y55F3A_74
179.29
7.b
Y40H7A.4
208.67
H25K10.3
158.09
150 31.30
19.01
6.56
1.23
0.66
0.53
22.33
26.24
44.38
1.67
19.51
25.64
9.30
29.67
10.17
15.88
2.22
6.37
7.09
2.68
6.21
1.64
0.59
1.88
0.64
1.00
0.88
0.32
1.02
0.35
0.55
0.54
0.54
0.54
0.55
0.55
18.62
18.14
8.13
1.12
0.63
0.56
16.40
20.46
2.42
1.26
0.71
0.56
25.03
23.02
4.82
1.40
0.79
0.57
37.83
26.89
28.95
23.87
7.13
5.46
1.73
1.41
1.00
0.82
0.58
0.59
13.99
23.68
4.51
1.39
0.82
0.59
45.68
16.74
4.15
0.97
0.58
0.59
35.61
20.48
4.76
1.18
0.71
0.60
39.96
57.02
41.14
54.65
72.02
27.37
39.65
85.99
40.08
39.06
12.95
12.64
22.23
54.18
17.54
12.24
25.28
26.17
20.86
14.13
15.46
23.40
36.38
21.02
17.37
26.79
22.14
11.37
28.56
26.42
3.79
6.70
7.22
3.31
2.64
6.24
3.08
11.37
0.69
0.74
1.56
3.33
1.52
9.40
5.75
0.69
1.43
1.47
1.17
0.79
0.85
1.28
1.98
1.13
0.94
1.44
1.18
0.61
1.50
1.37
0.42
0.87
0.90
0.72
0.49
0.53
0.81
1.25
0.72
0.60
0.92
0.76
0.39
0.98
0.91
0.61
0.61
0.61
0.61
0.62
0.63
0.63
0.63
0.64
0.64
0.64
0.64
0.65
0.66
0.66
16.86
11.24
1.56
0.58
0.39
0.67
8.03
27.66
1.95
1.40
0.95
0.68
24.38
27.70
2.02
1.35
0.96
0.71
2.40
26.07
32.72
24.99
7.17
1.96
1.57
1.19
1.13
0.86
0.72
0.73
Table A-2: continued Y4C6A.h
163.15
T28F3.6
183.43
Y41E3.11
203.17
Y51H4A.1
192.47
Y38F2A_57
22.89
43.F
Y43D4A.b
131.05
Y105C5A.J 160.06
H02112.6
108.62
R09E10.5
145.90
K03H6.1
136.85
M18.7
140.44
Y105C5A.
159.78
H
F13E9.3
113.56
F25H8.1
104.08
H02112.7
75.43
Y67D8A_38
167.36
2.b
Y37A1B.8
170.35
F11A10.3
47.41
F13E9.1
129.68
Y76B12C_6
123.82
7.C
B0564.4
169.17
Y69A2A_29
156.19
91.e
M18.5
83.48
F08B4.1
20.72
T04A11.11 112.70
T04A11.3
120.87
T27E7.3
138.48
Y69A2A_17
150.01
88.d
K10D11.1
96.07
Y41D4A_27
19.02
68.A
R102.7
134.19
F58G6.4
131.31
F44D12.6
17.78
C18H7.8
167.93
151 15.76
27.36
32.95
40.17
25.87
29.10
32.85
31.12
4.16
3.72
9.12
7.61
1.22
1.38
1.53
1.44
0.89
1.00
1.13
1.07
0.73
0.73
0.74
0.74
35.61
3.81
4.87
0.17
0.13
0.76
3.04
36.11
9.01
42.37
24.23
17.76
22.26
28.52
20.18
27.21
26.80
27.64
5.25
3.78
7.28
9.77
7.11
7.82
0.98
1.20
0.82
1.10
1.03
1.05
0.77
0.98
0.70
0.94
0.92
0.95
0.78
0.82
0.85
0.86
0.90
0.90
23.04
31.45
6.12
1.20
1.08
0.90
17.34
15.52
14.04
22.66
20.87
15.16
20.44
11.55
7.66
0.85
0.78
0.57
0.78
0.72
0.52
0.92
0.92
0.92
50.36
33.80
4.92
1.26
1.17
0.93
45.15
4.47
24.37
34.67
9.76
26.93
2.36
3.03
10.38
1.28
0.36
0.97
1.20
0.34
0.93
0.93
0.95
0.95
16.76
26.06
4.51
0.93
0.90
0.97
13.76
35.94
10.94
1.27
1.24
0.98
17.86
34.36
14.98
1.17
1.18
1.01
10.52
4.62
17.40
11.56
40.10
18.52
4.60
25.30
27.18
31.54
6.11
1.84
3.75
10.24
2.84
0.63
0.16
0.85
0.91
1.04
0.64
0.16
0.87
0.94
1.09
1.02
1.02
1.03
1.03
1.05
24.68
34.33
5.96
1.13
1.18
1.05
15.59
22.08
2.77
0.72
0.76
1.06
6.15
4.38
0.58
0.14
0.15
1.06
49.85
10.75
20.58
31.19
30.93
31.63
4.63
43.85
4.49
10.06
1.17
7.77
1.01
0.99
0.13
1.26
1.07
1.09
0.16
1.51
1.06
1.11
1.19
1.20
Table A-2: continued F38H4.9
6.15
Y9C9A_54.
178.38
C
H01G02.2
110.67
C32H11.7
107.62
M117.2
49.34
C42C1.5
92.02
K07F5.7
119.21
F40F11.2
44.01
F58G6.1
83.64
C47E12.4
54.38
ZK593.2
67.03
T25B9.7
92.57
M7.1
48.50
F32B6.7
107.52
Y55F3A_75
19.32
0.E
F28D1.1
55.43
Y55H10A.1 2.76
Y69E1A.3
100.59
F12F6.6
8.46
Y45F10D.8 39.71
Y116A8C.3
4.76
5
F42C5.10
15.37
Y41D4A_30
2.05
73.b
AC7.1
4.14
C47E12.5
1.47
F42C5.8
10.84
F38E11.5
5.33
Y41D4A_34
1.38
57.A
F49C12.8
1.29
152 2.73
1.61
0.44
0.05
0.06
1.20
29.83
46.69
2.80
1.34
1.61
1.20
12.22
18.48
9.42
37.03
19.53
12.25
11.67
6.46
21.37
14.99
9.18
33.97
29.03
28.39
13.78
25.76
33.48
12.54
24.36
15.84
20.24
28.11
14.98
33.79
9.02
6.38
2.52
7.54
4.19
3.34
4.32
1.45
0.34
6.55
4.31
5.98
0.83
0.81
0.37
0.69
0.89
0.33
0.63
0.41
0.50
0.69
0.36
0.81
1.00
0.98
0.48
0.89
1.15
0.43
0.84
0.55
0.70
0.97
0.52
1.17
1.20
1.21
1.28
1.29
1.29
1.31
1.34
1.34
1.39
1.39
1.42
1.44
5.32
6.38
2.77
0.15
0.22
1.52
5.24
1.72
29.16
3.98
12.83
18.60
0.93
35.47
3.62
17.64
4.59
0.12
6.88
1.77
6.96
0.42
0.02
0.76
0.06
0.30
0.64
0.03
1.22
0.12
0.61
1.54
1.55
1.62
1.97
2.04
4.30
2.22
1.84
0.04
0.08
2.14
1.72
8.08
5.83
0.12
0.28
2.41
0.71
1.13
0.23
0.02
0.04
2.54
0.92
0.56
10.23
7.51
2.51
1.14
10.89
5.41
2.36
0.31
6.45
2.56
0.03
0.01
0.08
0.04
0.09
0.04
0.38
0.19
2.78
3.56
4.61
4.66
0.54
1.56
0.76
0.01
0.05
5.19
0.49
2.28
1.44
0.01
0.08
8.14
153 Table A-3: Clones selected for cytological analysis from Chromosome II RNAi screen
Wild-type
akir-1(rj1)
Chrom II
Average# of
Average# of
akirstdev
stdev
Clone ID
progeny
progeny
1(rj1)/WT*
C18A3.5
222.00
20.30
2.06
1.23
0.07
B0454.3
146.50
9.09
1.36
0.63
0.07
F59H6.2
103.75
39.05
1.04
0.32
0.07
F45C12.14
126.31
27.34
1.50
0.25
0.09
F22E5.2
100.00
27.24
1.33
0.58
0.10
C52E2.I
128.57
67.00
1.58
0.52
0.1
K12H6.12
137.75
79.77
2.53
1.72
0.1
F42G2.6
134.97
49.66
1.89
1.02
0.10
T08E11.4
150.83
5.64
2.17
0.29
0.10
D2062.2
108.11
39.80
1.67
0.58
0.11
F45C12.2
123.18
31.23
1.93
1.22
0.11
F36H5.4
118.93
50.10
1.92
0.72
0.12
F42G2.2
134.00
27.62
2.19
1.56
0.12
C32D5.12
314.33
191.06
5.50
1.50
0.13
C08F1.7
126.10
27.22
2.27
2.80
0.13
F42G2.4
50.60
12.02
1.00
0.00
0.14
H41C03.2
221.58
48.96
4.42
2.92
0.14
B0047.2
168.08
41.90
3.83
1.04
0.16
F45C12.12
83.25
32.25
1.94
1.29
0.17
T23B7.1
470.33
185.80
11.17
5.09
0.17
T08E11.2
113.45
15.41
2.89
2.01
0.18
F59H6.12
59.75
44.27
1.55
0.64
0.19
F28C6.9
180.33
28.04
5.67
4.31
0.2
EEED8.1
121.53
37.63
5.17
4.19
0.3
E02H1.1
95.89
12.17
5.44
1.71
0.4
C34F11.3
198.67
64.07
12.17
1.59
0.4
C08H9.5
153.83
64.87
10.17
7.22
0.5
ZK177.8
103.67
14.64
7.00
3.28
0.5
F10G7.4
100.89
34.80
7.45
1.70
0.50
C07E3.5
107.58
24.89
7.67
8.08
0.5
F52C6.4
463.3
53.45
39.3
15.3
0.6
*Normalized to the pL4440 control for each individual set of experiments
Table A-3: continued
Wild-type
akir-1(rj1)
Chrom II
Average# of
Average# of
akirstdev
stdev
Clone ID
progeny
progeny
1(rj1)/WT*
K07D4.5
129.7
36.02
12.0
12.2
0.6
T10D4.4
452.7
71.84
46.0
22.6
0.7
ZK250.5
331.3
164.00
35.3
10.3
0.7
ZK1240.6
430.7
28.73
49.7
6.0
0.8
ZC239.6
169.0
73.99
24.3
15.1
1.0
R12C12.6
116.00
33.94
17.61
2.37
1.1
C16C4.4
387.3
74.87
79.0
46.1
1.4
F19H8.1
137.61
32.59
28.62
7.05
1.50
C40D2.3
425.3
67.42
100.0
29.5
1.6
C41C4.4
95.00
23.70
32.31
3.26
2.4
T25D3.3
105.56
9.62
35.83
21.46
2.4
C32D5.6
110.78
12.08
37.50
13.70
2.4
F44G4.8
95.25
37.23
36.93
12.74
2.8
W05H5.4
91.60
14.21
35.50
1.89
2.8
C50D2.2
108.78
4.67
47.17
0.76
3.1
F39E9.2
54.11
76.53
26.58
8.84
3.54
C18H9.3
76.00
30.18
47.17
8.10
4.47
T01D1.6
95.00
10.93
60.00
4.58
4.50
Y48C3A.1
54.75
20.25
42.39
17.17
5.57
R07C3.7
75.00
30.05
58.50
28.54
5.62
T19D12.6
86.67
24.89
69.50
37.01
5.77
*Normalized to the pL4440 control for each individual set of experiments
154 155 Table A-4: Chromosome II cytological defect analyses-wild-type
Chrom II
Clone ID
C18A3.5
B0454.3
F59H6.2
F45C12.14
F22E5.2
C52E2.I
K12H6.12
F42G2.6
T08E11.4
D2062.2
F45C12.2
F36H5.4
F42G2.2
C32D5.12
C08F1.7
F42G2.4
H41C03.2
B0047.2
F45C12.12
T23B7.1
T08E11.2
F59H6.12
F28C6.9
EEED8.1
E02H1.1
C34F11.3
C08H9.5
ZK177.8
F10G7.3
F10G7.4
CF* (# of gonads
examined)
Not done
Not done
Not done
Not done
Not done
not done
not done
Not done
Not done
Not done
Not done
Not done
Not done
Not done
Not done
Not done
Not done
Not done
Not done
Not done
Not done
Not done
Not done
not done
Not done
not done
not done
not done
not done
not done
SYP-1(#of gonads
examined)
4
Not done
Not done
Not done
Not done
Not done
Not done
Not done
Not done
Not done
Not done
Not done
Not done
Not done
Not done
Not done
Not done
Not done
Not done
Not done
Not done
Not done
15
Not done
Not done
Not done
Not done
Not done
61
52
C07E3.5
Not done
Not done
F52C6.4
not done
31
K07D4.5
not done
18
* Carnoy’s Fixation=whole worm prep
NA= not available
SC defect
NO
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
NO
N/A
N/A
N/A
N/A
N/A
NO
weak
pheno
N/A
NO
NO
156 Table A-4: continued
Chrom II
CF* (# of gonads SYP-1(#of gonads
Clone ID
examined)
examined)
T10D4.4
not done
32
ZK250.5
not done
19
ZK1240.6
not done
0
ZC239.6
not done
41
R12C12.6
not done
Not done
C16C4.4
not done
32
F19H8.1
not done
Not done
C40D2.3
not done
39
C41C4.4
Not done
Not done
T25D3.3
Not done
Not done
C32D5.6
Not done
Not done
F44G4.8
Not done
Not done
W05H5.4
Not done
Not done
C50D2.2
Not done
Not done
F39E9.2
Not done
Not done
C18H9.3
Not done
Not done
T01D1.6
not done
Not done
Y48C3A.1
Not done
Not done
R07C3.7
Not done
Not done
T19D12.6
Not done
Not done
* Carnoy’s Fixation=whole worm prep
NA= not available
SC defect
NO
NO
N/A
NO
N/A
NO
N/A
NO
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
157 Table A-5: Chromosome II cytological defect analyses-akir-1(rj1)
Chrom II
Clone ID
C18A3.5
B0454.3
F59H6.2
F45C12.14
F22E5.2
C52E2.I
K12H6.12
F42G2.6
T08E11.4
D2062.2
F45C12.2
F36H5.4
F42G2.2
C32D5.12
C08F1.7
F42G2.4
H41C03.2
B0047.2
F45C12.12
T23B7.1
T08E11.2
F59H6.12
F28C6.9
EEED8.1
E02H1.1
C34F11.3
C08H9.5
ZK177.8
F10G7.3
CF* (# of
gonads
examined)
Slides prepped
Slides prepped
Slides prepped
Slides prepped
Slides prepped
10
10
Slides prepped
Slides prepped
Slides prepped
Slides prepped
Slides prepped
Slides prepped
18
Slides prepped
Slides prepped
Slides prepped
Slides prepped
Slides prepped
Slides prepped
Slides prepped
Slides prepped
Not done
10
Not done
10
10
10
Not done
SYP-1(#of
gonads
examined)
52
6
4
36
36
25
26
47
0
11
50
9
11
17
34
23
0
51
31
0
0
0
35
25
9
20
21
23
85
F10G7.4
Not done
41
C07E3.5
Not done
15
F52C6.4
Not done
42
K07D4.5
Not done
52
* Carnoy’s Fixation=whole worm prep
SC Defect
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
Bivalent
Defect
Number
of exps
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
False
Positive?
NO
NO
NO
NO
2
2
2
2
2
3
3
2
1
1
2
1
1
1
1
1
1
1
1
1
1
1
1
3
1
3
3
3
5
4
1
2
2
158 Table A-5: continued
Chrom II
Clone ID
T10D4.4
ZK250.5
ZK1240.6
ZC239.6
R12C12.6
C16C4.4
F19H8.1
C40D2.3
C41C4.4
T25D3.3
C32D5.6
F44G4.8
W05H5.4
C50D2.2
F39E9.2
CF* (# of
gonads
examined)
Not done
Not done
Not done
13
SYP-1(#of
gonads
examined)
76
68
24
84
10
32
Not done
58
10
7
Not done
53
Not done
13
Not done
5
Not done
14
Not done
22
Not done
2
Not done
31
Slides
15
prepped
C18H9.3
Slides
16
prepped
T01D1.6
10
7
Y48C3A.1
Slides
0
prepped
R07C3.7
Slides
5
prepped
T19D12.6
Slides
12
prepped
* Carnoy’s Fixation=whole worm prep
Bivalent
Defect
Number
of exps
5
2
2
5
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
False
Positive??
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
NO
1
NO
NO
NO
NO
3
1
NO
NO
2
NO
NO
2
SC Defect
NO
NO
NO
NO
3
2
3
2
1
1
1
1
1
1
1
Table A-6: Clones selected for cytological analysis from Chromosome IV
RNAi screen
Wild-type
akir-1(gk528)
akirChrom IV Average# of
Average# of
stdev
stdev
1(gk528)
Clone ID
progeny
progeny
/WT*
F55G1.10
215.86
73.57
4.03
1.64
0.09
F54E12.4
105.00
20.22
2.67
1.15
0.12
B0547.1
225.77
65.01
7.37
0.90
0.15
C28C12.8
152.00
28.58
5.99
1.79
0.18
C28C12.2
190.95
165.46
8.52
1.84
0.21
T22D1.10
232.00
14.18
11.33
2.04
0.22
C09G4.3
65.22
45.52
3.23
2.04
0.23
ZK381.4
211.77
68.61
12.38
3.83
0.27
F38A5.5
164.37
44.11
9.61
1.69
0.27
Y66H1B.2
60.57
37.62
4.00
1.49
0.30
F20D12.1
159.04
13.75
12.83
3.82
0.37
C44B12.1
103.29
14.79
11.23
2.35
0.50
T22D1.9
164.11
62.07
18.96
6.36
0.53
C55C3.5
78.43
26.24
9.30
6.37
0.54
F20C5.1
105.05
72.02
14.13
2.64
0.62
H02112.6
108.62
9.01
20.18
7.28
0.85
H02112.7
75.43
14.04
15.16
7.66
0.92
M18.5
83.48
10.52
18.52
6.11
1.02
F08B4.1
20.72
4.62
4.60
1.84
1.02
F38H4.9
6.15
2.73
1.61
0.44
1.20
M117.2
49.34
9.42
13.78
2.52
1.28
M7.1
48.50
9.18
14.98
4.31
1.42
C47E12.5
1.47
0.56
1.14
0.31
3.56
F38E11.5
5.33
7.51
5.41
2.56
4.66
F49C12.8
1.29
0.49
2.28
1.44
8.14
* Normalized to pL4440 controls on each background
159 160 Table A-7: GO Terms for chromosome IV RNAi clones selected for
cytological analyses
Gene Public
Name
Sequence Name
(Gene)
ama-1
C02B10.5
C44B12.1
C55C3.5
cks-1
csn-5
F36A4.7
C02B10.5
C44B12.1
C55C3.5
C09G4.3
B0547.1
csn-5
B0547.1
csn-5
csn-5
csn-5
csr-1
csr-1
B0547.1
B0547.1
B0547.1
F20D12.1
F20D12.1
reproduction
reproduction
reproduction
oogenesis
meiosis
cytokinesis
DEAD/H-box RNA
helicase binding
gonad development
oogenesis
reproduction
gamete generation
P granule
csr-1
F20D12.1
reproduction
ddb-1
M18.5
gamete generation
ddb-1
dic-1
dic-1
E04A4.5
egl-19
elo-2
exos-4.1
F38A1.8
F38E11.5
F38E11.5
F49C12.12
M18.5
F08B4.1
F08B4.1
E04A4.5
C48A7.1
F11E6.5
B0564.1
F38A1.8
F38E11.5
F38E11.5
F49C12.12
reproduction
oogenesis
reproduction
reproduction
reproduction
reproduction
gonad development
reproduction
oocyte development
reproduction
reproduction
fat-6
VZK822L.1
reproduction
fln-1
Y66H1B.2
meiosis
fln-1
Y66H1B.2
reproduction
H35B03.2
his-55
H35B03.2
F54E12.1
reproduction
reproduction
GO Term
Table A-7: continued his-58
his-61
hlh-12
hlh-12
hsp-1
icl-1
ilys-4
itr-1
itr-1
F54E12.4
F55G1.10
C28C12.8
C28C12.8
F26D10.3
C01F6.8
C55F2.2
F33D4.2
F33D4.2
itr-1
F33D4.2
JC8.5
lag-1
lag-1
let-70
let-70
let-92
let-92
JC8.5
K08B4.1
K08B4.1
M7.1
M7.1
F38H4.9
F38H4.9
let-92
F38H4.9
let-92
let-92
mesp-1
mesp-1
F38H4.9
F38H4.9
C28C12.2
C28C12.2
mesp-1
C28C12.2
mex-5
mig-32
npp-1
npp-1
npp-1
npp-1
nspb-3
par-5
par-5
pgl-1
pgl-1
pgl-1
pgl-1
pgl-1
W02A2.7
F11A10.3
K07F5.13
K07F5.13
K07F5.13
K07F5.13
F38A5.5
M117.2
M117.2
ZK381.4
ZK381.4
ZK381.4
ZK381.4
ZK381.4
Reproduction
reproduction
regulation of meiosis
reproduction
reproduction
reproduction
reproduction
oocyte maturation
reproduction
reproductive process in a
multicellular organism
reproduction
cell division
reproduction
meiosis
reproduction
centrosome separation
cytokinesis
DEAD/H-box RNA
helicase binding
meiotic spindle disassembly
reproduction
cell division
cytokinesis
meiotic chromosome
segregation
P granule
reproduction
cell division
cytokinesis
oogenesis
reproduction
reproduction
meiosis
reproduction
gamete generation
oogenesis
P granule
P granule organization
reproduction
161 Table A-7: continued pme-3
F20C5.1
pme-3
F20C5.1
pole-2
pyp-1
rpc-1
rpl-12
rpl-18
rpl-20
rpn-1
rpn-7
rps-18
rps-2
rps-24
rps-8
ruvb-2
ruvb-2
ruvb-2
sar-1
sars-1
sec-24.1
skn-1
srz-23
swd-2.2
tag-335
tag-49
F08B4.5
C47E12.4
C42D4.8
JC8.3
Y45F10D.12
E04A4.8
T22D1.9
F49C12.8
Y57G11C.16
C49H3.11
T07A9.11
F42C5.8
T22D1.10
T22D1.10
T22D1.10
ZK180.4
C47E12.1
F12F6.6
T19E7.2
F58E2.9
C33H5.7
C42C1.5
AC7.1
Meiosis
meiotic chromosome
segregation
reproduction
reproduction
reproduction
reproduction
reproduction
reproduction
reproduction
reproduction
reproduction
reproduction
reproduction
reproduction
cell division
cytokinesis
reproduction
reproduction
reproduction
reproduction
reproduction
reproduction
reproduction
reproduction
reproduction
tin-9.2
B0564.1
gonad development
uaf-2
Y116A8C.35
reproduction
uba-1
C47E12.5
oogenesis
uba-1
C47E12.5
reproduction
vha-19
Y66H1A.4
yars-1
zen-4
Y55H10A.1
Y66H1A.4
K08F11.4
M03D4.1
reproduction
reproduction
reproduction
cell division
zen-4
M03D4.1
Cytokinesis
162 163 APPENDIX B
THE ROLE OF AKIRIN IN DROSOPHILA OOGENESIS
Introduction
The role of meiosis is to produce gametes and mis-segregation of chromosomes
during meiosis can lead to aneuploidy. The role that AKIRIN has in SC disassembly was
discussed in Chapter 1 and 2. The hypothesis of this project was that the SC defects that
were observed in akir-1 mutants in C. elegans are evolutionarily conserved.
Mutants exhibiting SC disassembly defects have been previously identified in
yeast [Cdc5(Sourirajan and Lichten 2008)and Ipl1/AirB (Jordan et al., 2009)] and C.
elegans [LAB-1(de Carvalho et al., 2008), HTP-1/2 (Martinez-Perez 2005) and AKIR-1
(Clemons et al., 2013)]. In yeast, the targets of Ipl1/AirB and Cdc5 were not identified
and therefore their function in the regulation of SC disassembly is unknown. LAB-1 and
HTP1/2 all affect not just SC disassembly, but also have affects on the formation of
crossovers, which suggests, that their affects on SC disassembly may not be entirely
direct. AKIR-1 only affects SC disassembly (and not crossover formation) and hence, is
likely a component involved in disassembly. Furthermore, studies in Drosophila have
uncovered many more proteins involved in SC disassembly: Hus1, Nhk1, Mad/BubR1,
Ord, Incenp, and DCap-g (Peretz et al., 2009; Ivanovska 2005; Malmanche et al., 2007;
S. E. Bickel, Orr-Weaver, and Balicky 2002; Resnick et al., 2009). This what? makes the
study of SC disassembly in Drosophila ideal as we can utilize the known mutants to
conduct epistatic analyses to determine if AKIRIN has a role in SC disassembly and if so
what is it.
As in C. elegans, the follicles in Drosophila ovarioles are arranged in a temporal
and spatial organization that corresponds to the events in prophase I (Figure B-4). In the
germarium (leptotene/zygotene), homologous pairing and SC assembly is established in
numerous proto-oocytes. As these proto-oocytes mature, the SC is disassembled on the
majority with only one oocyte reaching maturity, while the rest will develop into nurse
164 cells in the follicle. Pachytene starts during Stage 2 and proceeds until Stage 10. Late
prophase events, including SC disassembly begins during Stages 5-7 and the SC is fully
disassembled by Stage 10 (McKim, Jang, and Manheim 2002; Page 2001).
Methods
Fly Strains and culture
Fly stocks were maintained at 21C on standard cornmeal-agar-yeast food.
Females selected for analysis were fed wet yeast daily and aged for 3-4 days prior to
dissection and ovary analyses. The yw stock was used as the wild-type control in
experiments except where indicated. akirinKG01343, akirinEY08097, and akirin DG08309 were
obtained from the Bloomington Drosophila Stock Center. akirin EY08097-e4 and akirin
EY08097-e5
were generated as described in Nowak et al,. 2013. akirin EY08097-e82 was
generated as described in Goto et al., 2008 (Table B-1). The TRiP.HMS01010 RNAi
stock was obtained from the Bloomington Stock Center and generated by the TRiP RNAi
Project at Harvard Medical School. The two maternal GAL4 lines were provided by Tina
Tootle.
RT-PCR
Five to ten pair of ovaries were collected per genotype to be tested, placed in
500uL of Trizol (Invitrogen), and mechanically dissociated in a 1.5mL tube by a pestle.
A standard chloroform/isopropanol extraction was conducted to recover the total mRNA
from the Trizol solution. The resulting pellet was washed in 75% ethanol, subjected to
DNAse I treatment to remove genomic DNA and stored at -80C. RT-PCR was
conducted using a SuperScript III 1-Step RT-PCR kit (Invitrogen) as per manufacturers
instructions. Primers and PCR conditions can be found in Table B2. Gel electrophoresis
was carried out to examine the products.
165 Cytology
Five to ten pairs of ovaries were collected per genotype to be analyzed. Upon
dissection, ovaries were placed in room temperature 200 uL of Grace’s Media (Lonza) in
a 1.5mL tube. The ovaries were allowed to settled to the bottom of the tube and the
majority of Grace’s Media was removed. 4% paraformaldehyde fix was prepared (in
Grace’s Media), 300uL was added to the tube containing the ovaries, and the tube was
placed on a rotator for 10 minutes at room temperature. The ovaries were washed six
times for ten minutes each wash in antibody wash (1X PBS+ 0.01% Triton X-100+ 0.1%
BSA). Primary antibodies were diluted in antibody wash: mouse α-C(3)G, 1:500 (Scott
Hawley) and rabbit α-Akirin, 1:500 (Jean-Marc Reichhart). 300uL of primary antibody
was added to the tube and the ovaries were incubated overnight at 4C on rotator. Ovaries
were then washed six times for ten minutes each wash in antibody wash. Secondary
antibodies used were AlexaFluor (Invitrogen) goat a-mouse 568 or 555, 1:1000 and
AlexaFluor (Invitrogen) goat a-rabbit 568 or 555, 1:1000. The ovaries were incubated
for 2-4 hours at room temperature on a rotator. After incubation, the ovaries were washed
six times for ten minutes each wash in antibody wash and then stained with DAPI,
1:10,1000 for ? minutes. A final wash with 1XPBS was conducted to remove excess
DAPI and the ovaries were either stored in 1xPBS at 4C or mounted onto slides
immediately. Microscopy and image acquisition was carried out as described in Chapter
4. Follicle staging was done in accordance with Spradling, 1993.
Results
Establishing an Akirin model system
We obtained six strains for this study, three of which are P-element insertions and
three of which are excision lines generated by Nowak et al., 2012 and Goto et al,, 2008
(Table B-1). Of these six, the three P-element insertions are in the first intron of the
akirin gene: akirinKG01343, akirinEY08097, and akirin DG08309 (Figure B-1A). akirinEY08097
and akirinKG01343 are homozygous viable stocks while akirin DG08309 is a balanced
166 heterozygote stock (Table B-1). These stocks shall be referred to as akirinKG, akirinEY,
and akirinDG. The remaining three strains are imprecise excision lines generated by the
removal of the P-element in akirinEY08097. akirin EY08097-e4 removes the first exon of the
akirin gene and part of the first intron as well. akirin EY08097-e5 is a result of an imprecise
excision which reinserted into the 5’ end of exon 2 (Nowak et al., 2012). akirin EY08097-e82
has not been mapped and the extent of the excision is not known; however, it is
homozygous lethal (Figure B-1A and B). I will refer to these stocks as akirinEx4, akirinEx5,
and akirinEx82. We used y1,w67c23 as a control as our study stocks were generated in that
background.
We knew from previous studies that some akirin mutants were homozygous lethal
(Goto et al., 2007; Nowak et al., 2012) and therefore needed a viable homozygous model
for this study. Trans-heterogygote combinations of the six alleles were generated to
determine if any allelic combination could be used as a model system for study (Figure
B-1B). When akirinKG, akirinEY and akirinEx82 were crossed with any other allele, they
yielded viable progeny. akirinKG and akirinEY are both homozygous viable, but akirinEx82
is not (FigureB-1B). This indicates that akirinKG and akirinEY are likely not null mutants.
akirinDG, akirinEx4, and akirinEx5 are all homozygous lethal and when crossed with each
other leads to lethality, indicating that these alleles are likely null.
I conducted reverse-transcription PCR (RT-PCR) to determine if akirin is
transcribe in the akirinEY and akirinKG alleles. Primers were designed to detect each of the
five known transcripts (Figure B-1C; Tables B-2 and B-3). I tested the presence of akirin
transcripts in the carcasses and ovaries of F1 flies from wild-type, akirinEY and
akirinKG/DG transheterozygotes (Figure B-1D). Wild-type and akirinEY had strong bands
indicating the presence of the mRNA in both the carcass and ovary; however, akirinKG/DG
had a visually reduced amount of transcript (Figure B-1D). These data were not
quantified and the precise amount of reduction is not known. rps17 (ribosome subunit)
was used as a control. Given that the akirinDG allele is null, these data indicate a basal
167 amount of akirin transcription in the akirinKG allele; it is likely a hypomorph. akirinEx4,
akirinEx5 and akirinEx85 were not tested.
Complementation Analyses
Given that the akirinKG could be used to generating a viable trans-heterozygote
model, we wanted to determine if Akirin was necessary for female fertility in Drosophila.
I conducted a series of complementation crosses to determine which trans-heterozygotes
would demonstrate an increase in lethality as we had seen in our studies of AKIRIN in C.
elegans. I crossed the akirinKG P0 (parental generation) females with P0 males of the
other akirin mutant alleles and wild-type controls (as well as the reciprocal crosses when
applicable). F1 trans-heterozygous progeny for the akirin mutants and controls were
obtained and quantified for each genotype generated, was calculated the percent of
expected F1 adults for each series of crosses. (Figure B-2 A-D and Table B-4).
In the first series of crosses, I tested only akirinKG, akirinDG and akirinEY alleles.
The akirinKG homozygous strain demonstrated a 3-fold reduction in the number of F1
progeny (n=75) as did the trans-heterozygotes akirinKG/EY (2.5-fold reduction n=54) and
akirinKG/DG (1.6-fold reduction n=106) when compared to the trans-heterozygote controls
(Table B-4 p<0.0005, Fishers Exact Test). akirinDG is homozygous lethal and produced
no F1 progeny which was statistically different from the trans-heterozygote control
akirinDG/+ (n=324) (Table B-4 p<0.0005). However, when crossed to the akirinKG allele,
akirinDK/KG F1 progeny were produced (n=217), but were not significantly different from
the control akirinDG/+ (n=324) (Figure B-2 and Table B-4, p=0.06 Fishers Exact Test).
In the second series of crosses, akirinKG and akirinDG females were crossed to the
excision alleles (reciprocal crosses set up where applicable) as well as to wild-type as to
controls (FigureB-2B-D). Of the three excision lines tested, only akirinEx5 had a
significant reduction (2-2.5 fold) in F1 progeny when crossed with akirinKG (including
the reciprocal cross) (akirinKG/Ex5 n=72 akirinEx5/KG n=88 p<0.0005 Fishers Exact test).
akirinKG crosses to the other two excision alleles were not significantly different than the
168 controls (Table B-4). Crosses of akirinDG to akirinEx5 and akirinEx4 produced no F1
progeny confirming these three alleles were likely nulls. However, the crosses to
akirinEx82 produced F1 adults (akirinDG/Ex82 n=3 and akirinEx82/DG n=37 p<0.0005 Fishers
Exact test). Taken together, these data indicated that akirinKG and akirinEx82 are likely
hypomorphic alleles and could be used to generate a viable model to study akirin in
Drosophila as they demonstrate a reduction in F1 progeny.
While these F1 adults were viable, we needed to determine if the females were
fertile or sterile. In addition to the complementation analyses, I conducted a 48-hour
fecundity study on F1 females that carried the akirinKG allele as well as one of the null
alleles (FigureB-3 and Table B-5). The akirinEx5/KG F1 females had a 6-fold decrease in
eggs hatched compared to the trans-heterozygous controls. Both the akirinKG/DG and
akirinKG/Ex5 F1 females had a 1.75-fold decrease in the number of eggs hatched. These
indicated that while F1 females with all three genotypes are fertile, akirinEx5/KG females
have significantly reduced fertility (Table B-5 p<0.0005 Fishers Exact test). These data,
along with the complementation data, indicated that the akirinEX5/KG trans-heterozygote
would be a viable model for this study. Additionally, akirinKG/Ex5 and akirinKG/DG would
possibly be good alternative models.
Akirin localization in the Drosophila germline
If Akirin acts in the germline, we expect it to localize to the germline. If Akirin’s
function is conserved than this localization should be nuclear, restricted to meiotic nuclei
of the germline and to late prophase I nuclei. There were several unsuccessful attempts to
generate AKIR-1 antibody in worms, therefore, the germline localization of Akirin was
never examined and this work in Drosophila could provide new insights about the
germline localization of Akirin. We generated an antibody for Akirin (using the peptide
sequence SNRFAKDSTEPSP (AA56-68) as immunogen) and used this antibody to test
localization in the germline.
169 Although, akirinE5/KG is likely the better model for the study of Akirin based on
our complementation and fecundity data, I preformed Akirin immunostaining on
akirinKG/DG F1 females as I had established that stock first. I quantified the number of
oocytes with no Akirin, Akirin dispersed, and Akirin in the nucleus (Figure B-6 and
Table B-6). In wild-type, no Akirin was found in germarium or in the early stage
follicles. At Stage 6, 28% of the oocytes had Akirin staining on the nucleus while in
Stage 7, 53% of oocyte nuclei had staining. In Stages 8-10, 100% of the wild-type
oocytes had a dispersed pattern of Akirin staining. (Figure B-5A and B-6A, Table B-6).
In contrast, in akirinKG/DG follicles, 100% of oocyte nuclei from the germarium to Stage 7
had no Akirin localization. Akirin localized to the nucleus much later, in Stage 8 where
85% of the nuclei had staining. In stages 9 and 10, Akirin localized to 90% and 100% of
the nuclei respectively (Figure B-5A and B, Table B-6) and remains localized on the
nucleus through Stage 10B. The difference localization patterns between Stages 6-10 are
statistically significant (Table B-6 p< 0.0005 Fishers Exact test).
A role for Akirin in Drosophila SC disassembly
My previous data established that Akirin is required for fertility in Drosophila
females. The next step was to determine if SC assembly or disassembly was affected in
these mutants. Therefore, the focus of our immunostaining analyses was on the
localization of C(3)G, a SC central region protein from the germarium to Stage 10. It is
worth noting that the SC disassembled at two different stages in Drosophila: in the protooocytes located in the germarium and in Stages 4-7 in the maturing egg. The second
disassembly event is variable between the different genetic backgrounds (Kim McKim,
Eric Joyce, members of the Orr-Weaver lab, personal communication). We included both
SC disassembly events in our analyses although we are interested in the later disassembly
stages. Although, akirinE5/KG is likely the better model system for the study of Akirin
based on our complementation and fecundity data, I preformed C(3)G immunostaining on
akirinKG/DG F1 females as I had established that stock first.
170 I examined the follicles to detect a CR defect using C(3)G immunostaining in
both wild-type and the akirinKG/DG mutant. The stage in which the SC disassembles
depends on genetic background. In many wild-type backgrounds, the SC starts to
disassemble in Stages 4-5 (McKim, Jang, and Manheim 2002; Page 2001). In our wildtype background, SC disassembly occurs later in stages 5-7. C(3)G localized to the
homologous chromosomes in the proto-oocytes in the germarium as expected in 100% of
the germariums examined. In Stage 2-5 follicles, C(3)G localized to the chromosomes in
100% of the nuclei examined. At Stage 6, when the SC should start to disassemble,
C(3)G staining along the chromosome axis was observed in 93% of the oocyte nuclei
(Figure B-7A and E). In Stage 7, 80% of the nuclei had a fragmented C(3)G pattern of
staining and 20% of nuclei had a diffuse pattern of staining. This indicates that SC
disassembly had started. However in Stage 8, 90% of the nuclei retained the fragmented
C(3)G staining, indicating that SC disassembly was not complete. Nuclei in Stages 9 and
10 also retained fragmented staining (Figure B-7A and Table B-7).
In the trans-hetrozygote controls, akirinDG/+ and akirinKG/+, C(3)G localization
along the axis is observed in 100% of the oocyte nuclei from the germarium to Stage 4
for both controls. In Stage 5, both controls had C(3)G staining on the chromosome axis
(84% and 93% Figure B-7B and C, Table B-7). At Stage 6, fragmented staining pattern
was observed in 66% of akirinDG/+ nuclei and 59% of akirinKG/+ nuclei. However, it is in
Stage 7 that the controls differ with akirinKG/+ retaining the fragmented staining in 90% of
the nuclei and retaining the fragmentation pattern until Stage 9 (Figure B-7C). In
contrast, 66% of akirinDG/+ nuclei had a dispersed C(3)G staining pattern and Stages 8-10
had 100% of their nuclei having the dispersed staining. (Figure B-7B Table B-7).
The akirinKG/DG mutant had a C(3)G staining pattern similar to the akirinKG/+
control. The germarium through Stage 4, 100% of the nuclei had C(3) on their axis. 88%
of Stage 5 nuclei had axis staining while at Stage 6 only 50% of nuclei had axis staining.
The remaining 50% had either fragmented or dispersed staining patterns (Figure B-7D).
171 Stage 7 had 29% of the nuclei showing an axial staining pattern while 71% had
fragmented or diffuse staining (n=7). In Stage 8, 66% of the nuclei had a fragmented
staining and in Stages 9 and 10, 100% of the nuclei had a dispersed staining pattern
(Figure B-7D and Table B-7). The only stage that was significantly different between the
akirinKG/DG mutant and the control was Stage 6 (p<0.05 for all pairwise comparisons,
Fishers Exact test).
I also performed C(3)G staining on akirinKG/Ex82 and controls (Figure B-8 and
Tables B-8 and B-9), although we later determined that this trans-heterozygote was not
an ideal model system. However, upon examination of the wild-type quantification and
that of the akirinEY/Ex82 that should have been similar to the control as it should express
Akirin due to the hypomorphic nature of akirinEx82 and the fact that akirinEY is not a null
allele, I saw that that the staining pattern was statistically different (Table B-8). When I
compared all the control trans-heterozygotes C(3)G staining patterns (Table B-9), they
were all statistically different. This led me to believe there was variability in the staining
of C(3)G and therefore, I pooled all the wild-type control data together, then analyzed the
two trans-heterozygote mutants (Figure B-9). When data analyzed this way, the only
statistically different pairwise comparison of Stages 5-7 where akirinDG/KG and
akirinKG/Ex82 at Stage 6 (p<0.0005 Fishers Exact test).
I also conducted an RNAi experiment using a UAS RNAi stock line from the
TRiP collection (Harvard Medical School) crossed to two different germline specific
GAL4 stock lines, maternal alpha Gal4(2) and maternal alpha Gal4(3). F1 progeny were
scored and there was no reduction in progeny numbers (Table B-10). This leads us to
propose that the RNAi would not be a viable model for studying Akirin’s role in meiosis.
Discussion
Akirin is required for fertility in Drosophila females
Goto et al., 2008 and Nowak et al., 2013 previously demonstrated that akirin null
alleles were lethal which made it necessary to find a viable akirin mutant to discern the
172 role of Akirin in meiotic prophase I events in Drosophila. The complementation analyses
allowed us to determine which of the akirin mutant strains in a trans-heterozygous cross
would not only produce F1 progeny, but also F1 females that were fertile. Out of the six
stocks that were examined, three are null, akirinDG, akirinEx4 and akirinEx5. These are not
only homozygote lethal, but produce no F1 progeny when crossed to each other. Two of
the other alleles, akirinKG and akirinEx82, are candidates for such a viable model system.
akirinKG is a hypomorphic allele as demonstrated by RT-PCR (reduction of
mRNA product in akirinKD/DG) and viability in the complementation studies. When
crossed to any of the null alleles, akirinKG trans-heterozygotes are viable regardless of the
direction of the cross (Figure B-1, B-2 and Table B-4); however only akirinKG/Ex5 mutants
exhibit a reduction in F1 progeny and F1 fertile females. Both the akirinKG and
akirinKG/Ex5 mutants would be good candidates to study Akirin. Akirin levels in F1
progeny of both genotypes should be quantified by qRT-PCR and western analyses using
yw as an appropriate control as both of these stocks were generated in that background.
While the akrinEx82 mutant is homozygous lethal, when crossed to the akirinDG
null mutant, it produced F1 progeny, but only when akirinEx82 females are used. This
indicated there maybe a maternal contribution requirement for Akirin function. However,
this is not seen with the akirinKG allele. This observation could be confirmed with
additional experiments including crosses with the other null alleles, akirinEx4 and
akirinEx5. RT-PCR on viable F1 trans-heterozygote progeny could be used to determine if
there is a reduction in transcription. Furthermore, the excision of the P-element was not
mapped.
The akirinEY allele is a P-element insertion, which is supposed to disrupt akirin,
but the homozygous viability and lack of reduction in mRNA product show that this
allele is not likely mutated. This could be due to the central location of the P-element in
the intron where it might be removed due to splicing during RNA processing.
173 Regardless, this allele produced mRNA products comparable to wild-type (not
quantified) and therefore should not be considered as a mutant.
Overall, these data from the complementation studies and F1 fertility studies
indicate there is a requirement for Akirin in Drosophila female fertility. While we have
identified at least two viable allelic combinations which should be further validated, these
should not be the only mutants considered. We also examined the use of RNAi under the
control of UAS-Gal4 system (Duffy 2002). RNAi transgenic stocks have been generated
with UAS control elements promoting their expression when mated to tissue specific
Gal4 producing lines. In this case, we wanted to deplete Akirin in the germline and used
germline specific Gal4 lines. However, we as we did not observe a reduction in progeny
numbers, suggesting that we did not activate the UAS controlled transcription of the
RNAi vector. As we now have at least two viable systems, we did not purse the RNAi
system. Alternatively, we could construct a UAS controlled rescue vector and generate a
somatic rescue stock from one of the null allele stocks. When crossed to a somatic Gal4
expressing stock, Akirin would be expressed in the somatic tissues, bypassing the
developmental defects, yet retain Akirin depletion in the germline. Regardless of the
system developed, Akirin levels in F1 progeny of both genotypes should be quantified by
qPCR and western analyses using yw as an appropriate control.
Akirin depletion leads to meiotic defects
I initiated the cytological analyses prior to the completion of the complementation
analyses. As a result, we did not have all the appropriate allelic combination available for
examination, only the akirinKG/DG trans-heterozygote was examined. This combination is
an adequate model system as it demonstrated a reduction in F1 progeny and had F1
female fertility. In this system, we examined Akirin and C(3)G localization, that later
being used to detect defects in SC disassembly.
174 The change in Akirin localization between wild-type and the akirinKG/DG mutant
from the cytoplasm to the nucleus in the later stages of pachytene (Stage 8-10) suggest
that Akirin is retained in the nucleus and perhaps on the chromosomes. This could be
confirmed through cytology with co-localization of nucleus specific proteins or
chromosome specific markers. Additionally, co-immunoprecipitation and western
analyses using antibodies for known nuclear protein as well as chromatin associated
proteins would provide data on whether Akirin is specifically interacting with other
proteins or chromatin. This experiment should be conducted with additional genetic
backgrounds such as akirinKG and akirinKG/Ex5 mutants to confirm that the change in the
staining is the result of depletion of Akirin. The implication of the change in localization
is not immediately clear as we do not know if or what proteins Akirin interacts with
physically or genetically. If the C. elegans candidates in this thesis are shown to interact
with Akirin, they could be come candidates for additional studies to determine the impact
of the change of localization on meiotic events in Drosophila.
The aim of this study was to determine if depletion of Akirin lead to meiotic
defects, specifically defects in SC disassembly. Unlike C. elegan,s where SC disassembly
has been extensively studied, relatively few studies have examined SC disassembly in
Drosophila, although SC disassembly mutants have been identified (Peretz et al., 2009;
Ivanovska 2005; Malmanche et al., 2007; S. E. Bickel, Orr-Weaver, and Balicky 2002;
Resnick et al., 2009). We used C(3)G as a marker for SC morphogenesis as it is the major
CR protein and is required for full synapsis and is likely the first to be removed as the SC
is disassembled (Page 2001). Three distinct stages of C(3)G localization were seen: the
first is on the chromosome axis, indicating that SC is formed; the second is fragments of
C(3)G along the chromosome axis, indicating that SC disassembly has initiated, and
lastly, a diffuse localization in the nucleus, indicating that C(3)G has been removed from
the chromosomes, but not yet removed from the nucleus (scoring system for SC defects
adapted from Resnick et al., 2009). Previous studies had shown that the SC disassembly
175 should be completed by Stage 6 and that C(3)G should no longer localize to the
chromosomes after Stage 6 in w1118 background (Page 2001). However, Resnick et al,,
2009 showed that C(3)G remains on the nucleus in a fragmented stage until Stage 8 in
OregonR flies. Within the field it is not clear exactly when the SC is fully disassembled
(Kim McKim, Eric Joyce, members of the Orr-Weaver lab, personal communication).
The initial stages of disassembly start at Stages 5-6, but the completion [the diffuse of
C(3)G throughout the nucleus and no residual fragments on chromosomes] has been
observed from Stage 6-8. Therefore, the exact timing of SC disassembly is not clear and
is likely one reason we were not able to detect a stronger SC defect in the akirinKG/DG
mutants. The akirinKG/Ex82 was not an appropriate allelic combination as there is likely not
a reduction in the function of Akirin as evidenced by the lack of reduction in F1 progeny.
Despite not having a clearly defined stages that represent the initiation and completion of
SC disassembly, in akirinKG/DG mutants had over 80% of Stage 5 nuclei with C(3)G on
the axis of the chromosomes indicating that SC disassembly had not been initiated. While
this is significantly different than our wild-type data, this was the only stage at which
there was a significant difference. To resolve if there is truly a change in the initiation of
SC disassembly, the experiment should be replicated with akirinKG/DG, akirinKG/Ex5,
akirinKG/+, akirinDG/+ and yw controls. This replication of experiments will account for
any differences in the controls and use both of the stronger trans-heterozygote system.
The reciprocal crosses should also be set up to confirm there is no difference in the
paternal/maternal contribution of Akirin on fertility.
176 Figure B-1: Identifying potential trans-heterozygote combinations to model akirin
deficiency in Drosophila. A) akirin alleles used in this study: akirinKG01343,
akirinEY08097, akirin DG08309, akirin EY08097-e4, akirin EY08097-e5, akirin EY08097-e82 B) Viability
of trans-heterozygote combinations from complementation study. Lethal combinations
in red. C) Schematic of known mRNA transcripts (FLYBASE). Red arrows mark Akirin
primer location, size 1300bp. D) Gel electrophoresis of RTPCR of Akirin in ovaries and
carcasses of wild-type, akirin EY08097, and akirinKG01343/DG08309. akirinKG01343/DG08309 has a
reduction in mRNA. NC=Not completed
Table B-1: Strains used in the Drosophila Akirin project
Alleles Used in
Genotype
Study
y1 w67c23; P{SUPorAkirinKG01343
P}akirinKG01343 ry506
y1 w67c23;
08309
P{wHy}akirinDG08309/TM3, Sb1
AkirinDG
Ser1
08097
AkirinEY
y1 w67c23; P{EPgy2}akirinEY08097
w;Akir4/TM3, P{GAL4-twi.G}2.3,
AkirinEY08097-e4
P{UAS-2xEGFP}AH2.3, Sb1 Ser1
AkirinEY08097-e5
w;Akir5/TM3, Df dlacZ
akirin-e82
hs GAL4/CyO; Aki 82 Excision/Tm6b
Control
HMS01010
RNAi
mat
alphaGAL4(2)
mat
alphaGAL4(3)
y1 w67c23
y1 sc* v1;
P{TRiP.HMS01010}attP2/TM3, Sb1
w[*]; P{w[+mC]=matalpha4-GALVP16}V2H (Ch 2)
w[*];: P{w[+mC]=matalpha4-GALVP16}V2H (Ch3)
177 Source
Status
Bloomington
hypomorph
Bloomington
possible null
Bloomington viable
Nowalk et al,
null
2012
Nowalk et al,
null
2012
Goto et al,
2008
178 Table B-2: Primers used for RT-PCR analyses
Lab
Primer Name
Primer (5’-3’)
ID#
509
DMakiRD-F
GCACACCGCCACTGCAGACATA
510
DMakiRD-R
TGCCAATTCTGTCGGTCTGTTATG
511
DMakiRC-F
TCTGGCATTTGTCACGTAAAAGAG
DMakiRABCD512
GATTATTGCCTCGATTCCAAATGATG
R
513
DMakiRAD-F
TAAATAAAGCGCGTGTGAAGTGGT
514
DMakiRE-R
TCCAAATGATGGATGATTCACGTC
DMakiRABDE515
CATTTTTGCTAAGAAGCGAGACGA
F
516
DMaki-F
AGCAGTACGATGCCTTTGTCAAGT
517
DMaki-R
GGTTTGTTTTGCGTTTGCTTTAGT
519
DMrpS17-F
ATCGAGAAGTACTACACTCGCCTGA
518
DMrpS17-R
TCCTGCAACTTGATGGAGATACC
Table B-3: Conditions for RT-PCR and amplicon length
Primer
Primer
Temp
forward
reverse
amplicon RA
513
512
55
amplicon RB
515
520
57
amplicon RC
511
517
55
amplicon RD
513
510
57
amplicon RE
515
514
55
amplicon akirin
517
515
57
amplicon
519
518
57
RPS17
mRNA
size (bp)
120
180
1000
1200
180
1300
Ext.
time
1:00
1:00
1:30
1:30
1:00
1:40
177
1:00
179 Figure B-2: Akirin is required for fertility. A-D) Complementation analyses for
various trans-heterozygote combinations. *p<0.0005 Fishers Exact test
No standard deviation was calculated as most of these were results of one experiment.
180 Table B-4: Number of trans-heterozygote F1 adult flies expected and p-values Fishers
Exact test for comparisons to the trans-heterozygote controls
trans % Flies
Total
Total
stdv
n
pFET
heterozygote
expected
Observed Expected
combinations
KG/KG
50.5
73
144.5
1.1
73
<0.0005
DG/DG
0.0
0
271.0
0.0
0
<0.0005
EY/EY
73.9
17
23.0
17
0.38
KG/EY
57.4
54
94.0
54
<0.0005
DG/EY
116.0
58
50.0
58
0.06
DG/KG
62.4
217
347.5
14.4
217
0.06
KG/DG
89.1
106
119.0
106 <0.0005
KG/WT
148.5
459
309.0
276.9 459
WT/KG
113.7
191
168.0
191
DG/WT
77.9
324
416.0
39.8
324
WT/DG
78.5
153
195.0
153
EY/WT
103.6
86
83.0
86
KG/ex82
62.1
77
124.0
12.3
77
0.69
ex82/KG
146.2
38
26.0
38
0.57
DG/ex82
2.2
3
134.0
1.7
3
<0.0005
ex82/DG
60.7
37
61.0
37
<0.0005
KG/WT
67.4
95
141.0
95
WT/KG
102.3
175
171.0
175
DG/WT
82.7
163
197.0
163
WT/DG
93.0
185
199.0
185
ex82/WT
123.3
143
116.0
143
WT/Ex82
93.3
194
208.0
194
KG/ex5
51.4
72
140.0
9.5
72
<0.0005
ex5/KG
44.7
88
197.0
6.9
88
<0.0005
DG/ex5
0.0
0
101.0
0.0
0
<0.0005
ex5/DG
0.0
0
165.0
0.0
0
<0.0005
KG/WT
115.2
303
263.0
30.4
303
WT/KG
124.3
251
202.0
69.3
251
DG/WT
80.2
207
258.0
10.4
207
WT/DG
85.9
201
234.0
34.4
201
ex5/WT
113.1
259
229.0
8.0
259
WT/ex5
69.3
142
205.0
59.4
142
KG/ex4
75.4
49
65.0
49
0.019
ex4/KG
103.6
58
56.0
58
0.54
DG/ex4
0.0
0
147.0
0
<0.0005
ex4/DG
0.0
0
115.0
0
<0.0005
KG/WT
137.9
91
66.0
91
DG/WT
85.7
72
84.0
72
ex4/WT
88.5
77
87.0
77
WT/ex4
126.2
130
103.0
130
n= number of flies counted
181 Figure B-3: The number of hatched eggs is reduced in akirin trans-heterozygote
models. akirinKG01343/DG08309 , akirin EY08097-e5/KG01343, and akirin KG01343/ EY08097-e5, all
have a significant reduction in the number of eggs hatched. *p<0.00005 and **p<0.0005
Fishers Exact test for all pairwise comparisons.
182 Table B-5: Results of fecundity study for selected trans-heterozygote model systems and
p-values Fishers Exact test
p-values FET
F1
% Embryos to WT
to DG TO KG TO EX5 TO EX4
Genotypes
n
Hatched
(48Hrs)
DG/KG
98
81.7
0.004
1
0.385
KG/DG
105
52.5
0.152
0.253
7.8E-10
EX5/KG
1.0E-14
1.4E-21 1.20E115
11.5
17
KG/EX5
176
53
0.1971
1.7E-09 0.0001
EX4/KG
189
76.8
0.0443
1
0.0010
KG/EX4
302
94
8.1E-08
0.59
0.31
DG/WT
294
82
WT/DG
128
61
KG/WT
63
91
WT/KG
107
76
EX5/WT
129
69
WT/EX5
126
78.5
EX4/WT
89
93.3
WT/EX4
45
89
WT
103
63
n=number of embryos counted
183 Figure B-4: A schematic representation of Drosophila oogenesis in prophase I.
The SC is shown in red and the oocyte in blue. Only the germarium and stages 2-7
represent prophase I events. In the 2a region of the germarium, the germline cells
differentiate and the SC forms in proto-oocytes. As the proto-oocytes mature, the SC
disassembles on all but one, which will become the oocyte in Stage 2-14. This oocyte
retains its SC along the axis of the chromosomes until Stage 4 when disassembly starts
and the chromosomes start to form the karyosome. By Stage 5, SC proteins start to
dissociate from the chromosomes and disassembly is complete by Stage 7.
184 Figure B-5: Immunostaining of Drosophila follicles with Akirin specific antibodies.
A) Akirin is in red/grayscale and chromatin is stained with DAPI in blue. In Stages 4-5
wild-type follicles, Akirin is found in the nucleus and is associate with the chromosomes
in the oocytes. In later stages, it is diffuse throughout the oocytes. In the
akirinKG01343/DG08309 mutant, the Akirin localization is specific to the nucleus with the
lack of diffuse staining in Stages 4-10. Scale bar is 40uM. B) Stage comparison between
wild-type and akirinKG01343/DG08309 mutant showing the lack of staining in the oocytes.
Scale bar is 40uM
185 Figure B-6: Quantification of Akirin staining. A) Quantification of wild-type staining.
Akirin localizes in Stage 6 to the nucleus and remains dispersed throughout the oocyte
until Stage 10. B) Quantification of akirinKG01343/DG08309 mutant Akirin localization.
Akirin appears to localize to the nucleus in Stage 8 and remains there throughout the
remaining stages. Stages 8, 9, and 10 are statistically significant with a p<0.00005 Fishers
Exact test (Table B-6).
186 Table B-6: Akirin quantification, n values and p-values Fishers Exact test
Akirin in
Akirin
No
Wild-type
n
nucleus
dispersed
Akirin
Germ
0
0
24
24
st3
0
0
24
24
st4
0
0
22
22
st5
0
0
20
20
st6
6
0
19
25
st7
9
4
10
23
st8
1
27
0
28
st9
0
18
0
18
st10
0
21
0
21
Akirin in
Akirin
nucleus
dispersed
Germ
0
0
st3
0
0
st4
0
0
st5
0
0
st6
0
0
st7
0
0
st8
17
0
st9
9
0
st10
10
0
n=number of ovarioles examined
KG/DG
No
Akirin
21
21
21
19
20
21
3
1
0
n
21
21
21
19
20
21
20
10
10
FETp-value
1
1
1
1
1.50E-07
1.87E-15
2.21E-59
2.21E-59
2.21E-59
187 Figure B-7: Quantification of C(3)G staining. A) wild-type B) akirin DG0830/+ C)
akirinKG01343/+ D) akirinKG01343/DG08309. Blue represents C(3)G localization on the
chromosome axis, green represents dispersed C(3)G in the nucleus and red represents
fragmented C(3)G on the chromosomes. n and p-values for each mutant can be found in
Table B7. akirinKG01343/DG08309 is statistically different from the heterozygous controls,
p<0.005 FET, but is marginally different from wild-type, p=0.03. E) Micrographs of SC
disassembly in Stage 4 and 5 nuclei immunostained with C(3)G in red and chromatin in
DAPI blue. Stage 4 wild-type and akirinKG01343/DG08309 demonstrates C(3)G is on the
chromosome axis. Stage 5 wild-type is still on the axis but akirinKG01343/DG08309
demonstrates the fragmented appearance. Scale bar is 5 uM.
Table B-7: Quantification of C(3)G, n values and p-values for Fishers Exact test
C3G
No
C3G
C3G
N
WT
on
C3
fragment dispersed
value
Axis
G
Germ
16
0
0
0
16
st4
14
0
0
0
14
st5
5
0
0
0
5
st6
14
1
0
0
15
st7
0
4
1
0
5
st8
0
9
1
0
10
stg 9
0
4
1
0
5
stg 10
0
1
0
0
1
p-value FET
to
to
C3G
C3G
C3G
No
N
to
DG/KG
DG/
KG/
on Axis fragment dispersed C3G value WT
wt
wt
Germ
20
0
0
0
20
1
1
1
st4
18
0
0
0
18
1
1
1
st5
15
2
0
0
17
0.99 0.09
1
0.000
st6
10
4
5
0
19
0.03 0.008
68
st7
0
2
5
0
7
0.12 0.99
0.31
st8
0
8
4
0
12
0.32 0.011 0.64
stg 9
0
0
3
0
3
0.07
1
0.19
stg 10
0
0
6
0
6
0.14
1
1
C3G
C3G
C3G
No
N
DG/WT
on Axis fragment dispersed C3G value
Germ
20
0
0
0
20
st4
17
0
0
0
17
st5
7
5
0
0
13
st6
0
6
3
0
9
st7
0
3
5
0
8
st8
0
1
9
0
10
stg 9
0
0
2
0
2
stg 10
0
0
0
0
0
C3G
C3G
C3G
No
N
KG/wt
on Axis fragment dispersed C3G value
Germ
20
0
0
0
20
st4
17
0
0
0
17
st5
14
1
0
0
15
st6
1
10
6
0
17
st7
0
9
1
0
10
st8
0
8
2
0
10
stg 9
0
2
1
0
3
stg 10
0
0
1
0
1
n=number of ovarioles examined
188 189 Figure B-8: Quantification of C(3)G staining for akirin EY08097-e82 models systems.
Blue represents C(3)G localization on the chromosome axis, green represents dispersed
C(3)G in the nucleus and red represents fragmented C(3)G on the chromosomes. A) wild-­‐
type with balancers on chromosome 3, B) akirinEY08097 / EY08097-­‐e82 mutant, C) akirinKG01343/ EY08097-­‐e82 Both stocks in A and B express Akirin but have statistically different C(3)G staining patterns (Table B8 and B9) 190 Table B-8: Quantification of C(3)G staining for akirin EY08097-e82 systems
WT
C3G
C3G
C3G
No
n
(balancers)
on
fragment dispersed C3G
Axis
Germ
36
0
0
0
36
st3
24
0
0
0
24
st4
28
5
0
0
33
st5
15
14
0
0
29
st6
0
21
4
0
25
st7
0
9
18
0
25
st8
0
2
12
0
14
stg 9
0
0
8
0
8
stg 10
0
0
4
0
4
EY/ex 82
C3G
fragment
C3G
dispersed
No
C3G
n
pFET
to WT
Germ
st3
st4
st5
st6
st7
C3G
on
Axis
27
21
19
3
0
0
0
0
2
19
15
23
0
0
0
0
0
0
0
0
0
0
0
0
27
21
21
22
15
23
st8
0
15
0
0
15
stg 9
stg 10
0
0
4
2
3
0
0
0
7
2
0.99
1
0.69
0.02
0.27
0.0000
79
0.0000
017
0.025
0.06
KG/ex 82
C3G
fragment
C3G
dispersed
No
C3G
n
pFET
to WT
Germ
st3
st4
st5
C3G
on
Axis
37
34
34
33
0
0
0
1
0
0
0
0
0
0
0
0
37
34
34
34
st6
8
19
1
0
28
st7
st8
stg 9
stg 10
1
0
0
0
11
12
2
1
6
13
8
7
0
0
0
0
18
25
10
8
1
1
0.02
0.0000
37
0.0000
9
0.04
0.044
0.477
1
n=number of ovarioles examined
191 Table B-9: Comparison between wild-type and Akirin expressing backgrounds for C(3)G
staining.
WT
C3G on
C3G
C3G
No
n
(balancers)
Axis
fragmented dispersed C3G value
Germ
24
0
0
0
24
st3
28
5
0
0
33
st4
15
14
0
0
29
st5
0
21
4
0
25
st6
0
9
18
0
25
st7
0
2
12
0
14
st8
0
0
8
0
8
stg 9
0
0
4
0
4
stg 10
0
0
0
0
0
EY/ex 82
Germ
st3
st4
st5
st6
st7
st8
C3G on
Axis
21
19
3
0
0
0
0
C3G
fragmented
0
2
19
15
23
15
4
C3G
dispersed
0
0
0
0
0
0
3
No
C3G
0
0
0
0
0
0
0
n
value
21
21
22
15
23
15
7
stg 9
stg 10
0
0
2
0
0
0
0
0
2
0
pFET to
WT
0.99
1
0.69
0.02
0.27
0.000079
0.000001
7
0.025
0.06
C3G
dispersed
0
0
0
0
1
1
1
0
No
C3G
0
0
0
0
0
0
0
0
n
value
16
14
5
15
5
10
5
1
pFET to
WT
1
0.69
0.06
6.46E-10
4.12E-07
0.017
0.047
1
wt (YW)
C3G on
C3G
Axis
fragmented
Germ
16
0
st4
14
0
st5
5
0
st6
14
1
st7
0
4
st8
0
9
stg 9
0
4
stg 10
0
1
n=number of ovarioles examined
192 Figure B-9: Comparison of pooled wild-type samples, akirinDG/KG and akirinKG/Ex82.
Due to the variation observed between the wild-type controls in the C(3)G
immunostianing experiments, all wild-type controls were pooled and then compared to
the trans-heterozygotes to determine if there was an SC defect in prophase I. There are
weak defects in stages 5-7 where C(3)G remains on the axis of the homologous
chromosomes later than wild-type, but only Stage 5 is significantly different. P<0.0005
Fishers Exact test.
Table B-10: Quantification of RNAi experiment
F1 progeny
n value
% of totalobs
w/y sc v;w/w;mat a
136
48.056
gal4(3)/TM3Sb
w/y sc v;w/w;mat a
147
51.94
gal4(3)/HMS01010
Total
283
%of total
expected
50
FET p value
p=0.7
50
w/y sc v;mat a gal4
(2)/w;w/TM3Sb
w/y sc v;mat a gal4
(2)/w;w/HMS01010
Total
147
47.72
50
161
52.27
50
w/y sc v;w;w/TM3 sb
w/y sc
v;w;w/HMS01010
Total
86
84
50.588
49.41
50
50
170
w;w;wt/gal4(3)
274
100
100
w;gal4 (2)/w;w
138
100
100
225
100
100
w;w;w
n=number of flies counted
193 p=0.7
308
p=0.5
194 APPENDIX C
EXPERIMENTS PERFORMED FOR CLEMONS ET AL., 2013
For our paper, Clemons et al., 2013, I performed the following work: prepared slides for
immunostaining and collected microscopy data for Table 4 and Figure 5. This paper
characterized the role of akir-1 in SC disassembly (summaries of major conclusions of
this paper can be found in Chapters 1 and 2). In akir-1 mutants, there are two major
defects observed: 1) aggregation of SYP proteins in diakinesis -1 oocyte and 2) bivalent
structural defects which include an abnormal bivalent structure and an increase in length
of the e bivalents. My work focused on the increased length defects. In akir-1 mutants,
we found that the diakinesis bivalents were twice as long as compared to wild-type
bivalents (p<0.0001 Mann Whitney test). We wanted to determine if this specific defect
in length was the outcome of earlier SC disassembly defects or if this defect was
independent of SC function.
Previous studies had shown that C. elegans chromosome condensation in
diakinesis is regulated by the condensin complexes consisting of subunits CAPG-2, DPY28, and HCP-6 (Chan, Severson, and Meyer 2004b; Csankovszki et al., 2009). It was also
shown that null mutants of these subunits result in the lack of bivalent structure.
Therefore, in order to determine if AKIR-1 was interacting with DPY-28 (condensin I/IDC
subunit) or HCP-6 (condensin II) or in an independent pathway, we conducted RNAi
experiments on capg-2(tm1833), dpy(s939) and hcp-6(mr17) using akir-1(RNAi). Data on
dpy(s939) and hcp-6(mr17) were included in the paper. We found that akir-1(RNAi)
yielded bivalents with increased length as was seen in akir-1(gk528) mutants. However,
the dpy(s939) and hcp-6(mr17) showed no such increase ( Table 4, Clemons et al.,,,,, ,
2013) Furthermore, akir-1(RNAi);hcp-6(mr17) and akir-1(RNAi); dpy(s939) showed an
additive affect indicating that akir-1 was working in a spate pathway (Table 4).
Additionally, neither of the single condensin subunit mutants demonstrated SYP-1
aggregation in diakinesis -1 (Figure 5, Clemons et al., 2013). There was no aggregation
195 of the SC defects observed in the double mutants (Figure 5). Therefore, we concluded
that AKIR-1 has a separate affect on SC disassembly, independent of that of the
condensins.
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