Developmental Regulation of DNA Replication in Drosophila melanogaster
By
Eugenia Agnes Park
B.S., Biochemistry
Washington State University
Pullman, WA, 1998
Submitted to the Department of Biology
in Partial Fulfillment of the Requirements for the Degree of
Doctor of Philosophy in Biology
At the
AMASSACHUSET-S INwTyIE
Massachusetts Institute of Technology
OF TECHNOLOGY
Cambridge, MA
SEP 13 2006
S gust 2006
LIBRARIES
U
© 2006 Eugenia Agnes Park. All rights reserved.
The author hereby grants to MIT permission to reproduce and to distribute
Publicly paper and electronic copies of this thesis document in whole or in part.
Signature of A uthor.................
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ARCHNES
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Department of Biology
August 17, 2006
Certified by.
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•..y
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............................................................................................
Terry L. Orr-Weaver
Professor of Biology
Thesis Supervisor
A ccepted by .............
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T
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Stephen Bell
Chair, Committee on Graduate Students
Department of Biology
Developmental Regulation of DNA Replication in Drosophila melanogaster
By
Eugenia Agnes Park
Submitted to the Department of Biology
in Partial Fulfillment of the Requirements for the Degree of
Doctor of Philosophy in Biology
ABSTRACT
In mitotic cell cycles, the genome must be replicated fully in each cell cycle to ensure the normal
complement of chromosomes. Failure to replicate chromosomes fully or a failure to limit
replication to once-per-cell-cycle may lead to aneuploidy and genomic instability. Variants of the
archetypal mitotic cell cycle, utilizing conserved cell cycle machinery, are employed during
metazoan development to achieve different aims. Endocycles, in which the cell cycle proceeds
without complete mitosis, generate polyploidy and are commonly employed to increase
metabolic capacity and cell size. D. melanogasterfollicle cell gene amplification, in which bidirectional replication occurs in the absence of detectable gap phases, serves to produce large
amounts of eggshell proteins and may also serve to regulate transcription. During D.
melanogasterembryogenesis, mitotic cell cycles, endocycles and cell cycle exit occur
concurrently. We undertook a screen to identify factors affecting developmentally regulated,
variant cell cycles during D. melanogasterembryogenesis. We identified a class of mutants with
apparently polyploid cells in normally diploid tissues indicating a failure to maintain mitotic
cycles. In this class of mutants, we identified and characterized new mutants in pavarottiand
tumbleweed, pav3CS3 and tumrn32a -2 .These mutants displayed phenotypic defects consistent with
failures in cytokinesis. In particular, tum3 2a-2o displayed multinucleate cells and abnormal
telophase spindles. We also describe the identification, cloning and characterization of the first
cyclinE mutant to undergo aberrant gene amplification, cyclinE1 36 . We observed a novel gene
amplification defect, dramatically increased replication fork progression in cyclinE'f36/cyclinEPs
and cyclinE1 36/cyclinEP28 follicle cells implicating CyclinE in the regulation of replication fork
speed.
Thesis Supervisor: Terry L. Orr-Weaver
Title: Professor of Biology
To my family... and T.C.
ACKNOWLEDGEMENTS
I would like to thank my advisor Terry Orr-Weaver for her guidance and support. Her love for
science and her integrity are inspiring. Thank you. Thanks also to the Orr-Weaver Lab, quite
frankly, THE GREATEST LAB ON THE PLANET and special thanks to Tamar Resnick with
whom I've had many helpful discussions. I would also like to thank my thesis committee for
guidance. Thanks to David MacAlpine, Cary Lai and Steve Bell for consistently helpful and
informative discussions and to my friends Eunice and Piyush who always had something
interesting to say about science. Last but not least, special thanks to my little sister June who has
been unflaggingly supportive.
TABLE OF CONTENTS
Chapter One:
Introduction:
The Developmental Regulation of DNA replication.........................................
-
Regulation of the mitotic cell cycle................................................................................8...
Regulation of the endocycle...........................................................................................9
Gene amplification....................................................... ................................................ 12
Cyclin/Cdk and regulationof the Pre-ReplicationComplex............................ ... 15
R eplication Initiation................................................................................
-
...............................
16
Eukaryotic origins of replication....................................................................................16
Epigenetic determinantsof metazoan origin activity.....................................18
The Pre-ReplicationComplex......................................................................................19
A molecular mechanismfor Mcm2-7 loading at the Pre-ReplicationComplex..............20
Mechanisms of Cyclin/Cdk regulationof the Pre-ReplicationComplex........................20
The transitionto replication........................................................................................24
R eplication Elongation...............................................................
-
...................
.............................................. 25
Replication Fork Progression......................................................................................26
Summ ary.................................................................................................... .............................. 27
R eferences ..................................................................... ..................................... ...................... 29
Chapter Two:
New mutants affecting developmentally regulated cell cycles during Drosophila
embryogenesis
..................................... 37
Sum mary.............................................................................................
38
Introduction ....................................................................................................................................
46
Resu lts............................................................................................................................................
-
A screenfor mutations affecting developmental cell cycle regulation.............................46
Class V mutants have large nuclei in the nervous system and epidermis................52
Characterizationand mapping of Ir8 and 3C157.......................................................55
Characterizationof 3C53 and 32a-20............................................................................58
3C53 and 32a-20 are alleles ofpavarottiand tumbleweed/RacGAP50..................61
D iscussion .................................................................................................................
-
. . ...........
75
The identificationof mutants that affect developmental regulationof the cell cycle........75
New mutants that affect mitotic cell cycles in the nervous system.................................76
tum3• 2a -2 disrupts cytokinesis and shows central spindle defects........................................77
M aterials and M ethods................................................................................................................
R eferences......................................................................................................................................8
79
1
Chapter Three:
The characterization of new cyclinE mutants that increase replication fork progression
during gene amplification
Sum m ary.............................................................................................
..................................... 85
Introduction............................................................................................... ............... ................. 86
Resu lts............................................................................................................................................9
1
- A new mutation in cyclinE that displays previously undescribeddefects in gene
amplification............................................................................
.................................... 91
- cyclinEJl36 /cyclinEPz8 and cyclinEIP6/cyclinEPz8have expanded amplified regions at
DAFC-66D, DAFC-30B and DAFC-34B.......................................................................94
- Increased replicationfork progression is not due to priorcell cycle defects........... 103
- Increasedreplicationfork progressionin cyclinEf36/cyclinEPz 8 follicle cells may reflect
increasedreplicationfork speed.....................................................................................110
- Polyteny is intact in cyclinE"36/cyclinEPz follicle cells.....................................115
- cyclinE'f36 displays a dominant defect in replicationfork progression.......... .... 118
- Double Parkedand the MCM complex localize as double bars in cyclinEf36/cyclinEPz8
................................ 122
follicle cells................................................................................
D iscussion......................................................................................
........................................ 126
M aterials and M ethods......................................................... ................................................. 134
References........................................................................
........................................
137
Chapter Four:
Conclusions and perspectives
Conclusions and perspectives...............................................................
References......................................................................................
.................................
........................................
143
152
CHAPTER ONE
Introduction
The Developmental Regulation of DNA Replication
DNA replication is developmentally regulated during Drosophilamelanogaster
development (Claycomb et al., 2002; Smith and Orr-Weaver, 1991). Endocycles, in
which alternating S and G phases produce polyploidy, and replication-based gene
amplification, are employed to increase tissue size and metabolic output. Drosophila
endocycles occur during embryogenesis, larval development and oogenesis and are
regulated by conserved replication factors. Gene amplification, in which successive
replication initiation events and elongation occur in the absence of detectable gap phases,
occurs during oogenesis. Drosophilafollicle cell gene amplification is employed to
produce high copy numbers of chorion, or eggshell, genes (Spradling, 1981). Like
endocycles, follicle cells employ conserved cell cycle regulators (Claycomb and OrrWeaver, 2005).
The following chapter reviews developmental regulation of endocycles and gene
amplification. In addition, current knowledge on mechanisms controlling DNA
replication initiation and elongation are reviewed.
Regulation of the mitotic cell cycle
The archetypal cell cycle consists of G 1, S, G2 and M phases and is regulated by
Cyclin/Cdk complexes consisting of a regulatory Cyclin subunit and a Cdk. Different
Cyclin/Cdk complexes act in different phases of the cell cycle. In S. cerevisiae,Cyclin
subunits confer phase-specificity on a single Cdk, Cdc28. The G1 Cyclins, the Clns, are
required for passage through START which signifies commitment to the cell cycle and Sphase entry. The G2 Cyclins, the Clbs, act in S, G2 and M phases. Degradation of the
Clbs is required for exit from mitosis (Reed, 1992). Unlike yeast, metazoans possess
multiple Cdks (Sherr and Roberts, 1999; Solomon, 1993). CyclinE/Cdk2 activity is
required for the G1/S transition. In mammals, CyclinA/Cdk2 regulates S-phase
progression. CyclinA/Cdkl and CyclinB/Cdkl activities are required for mitosis.
CyclinD/Cdk4 and CyclinD/Cdk6 mediate the convergence of growth factor signaling on
the cell cycle and are thought to indirectly regulate S-phase entry by potentiating
CyclinE/Cdk2 activity (Perez-Roger et al., 1999; Sherr and Roberts, 1999). Cyclin/Cdk is
regulated redundantly during the cell cycle. Mechanisms for regulating Cyclin/Cdk
activity include oscillatory Cyclin expression, regulatory phosphorylation, inhibition by
Cyclin/Cdk inhibitors (CKI) and the targeted degradation of Cyclins.
Regulation of the endocycle
Endocycles consist, minimally, of discrete S and G1 phases with one round of
DNA replication occurring per endocycle (Smith and Orr-Weaver, 1991). Some
endocycling tissues show vestiges of mitosis ranging from chromosome condensation to
nuclear divisions (Edgar and Orr-Weaver, 2001) and utilize mitotic machinery to regulate
Cyclin/Cdk activity. During mitotic cycles, the APC, an E3 ubiquitin ligase, marks
Cyclins for degradation by the 26S proteasome. Mutations in morula, an APC subunit,
lead to ectopic spindle formation and chromosome condensation in Drosophilanurse
cells undergoing endocycles, suggesting that mitotic regulators are expressed at low
levels in these cells (Kashevsky et al., 2002; Reed and Orr-Weaver, 1997). APC activity
may be required for establishing Drosophilaembryonic endocycles by clearing mitotic
Cyclins. Mutants in an APC coactivator, fizzy-related (fzr), fail to initiate embryonic
endocycles and fzr is developmentally regulated by Notch in a mitotic to endocycle
switch during oogenesis (Schaeffer et al., 2004; Sigrist and Lehner, 1997). In addition,
mitotic cyclin transcription is shut off during Drosophilaembryonic endocycles (Weiss et
al., 1998). Together, these observations suggest that endocycles arise from
downregulation of mitosis specific regulators and support the idea that endocycles are a
modification of the archetypal mitotic cell cycle.
Some endocycles show no vestiges of mitosis and may achieve oscillatory
Cyclin/Cdk activity through means that don't involve the APC or mitotic Cyclins. Cyclin
E/Cdk2 is important for mammalian endocycles. Double knockouts of the two murine
isoforms of CyclinE, CCNE1 and CCNE2, die midgestation due to defects in the placenta
without any apparent defects in embryos. Specifically, the trophoblast giant cells do not
attain normal levels of polyploidy. This may reflect the particular importance of
CyclinE/Cdk2 in regulating mammalian endocycles (Berthet et al., 2003; Geng et al.,
2003; Parisi et al., 2003). In Drosophila,CyclinE is required for embryonic endocycles
(Knoblich et al., 1994).
cyclinE transcription corresponds with S-phase in the Drosophilaendocycling
tissues and is key to developmental regulation of endocycles (Knoblich et al., 1994; Lilly
and Spradling, 1996). cyclinE is not expressed in intervening gap phases, suggesting that
endocycles are driven by pulses of cyclinE (Knoblich et al., 1994; Lilly and Spradling,
1996). Low CyclinE levels between pulses of cyclinE transcription are required for
replication. Ectopic expression of cyclinE in Drosophilalarval endocycling tissues
inhibits S-phase (Follette et al., 1998; Weiss et al., 1998).
Cyclical cyclinE transcription is important for endocycles and a biphasic
oscillator, consisting of the E2F1/Rbf transcription factor and the CyclinE/Cdk2 inhibitor
Dacapo, has been postulated to regulate cyclinE transcription (Edgar and Orr-Weaver,
2001). In the endocycling tissues of the Drosophilaembryo, the E2F1 transcription factor
regulates cyclinE expression as part of a G1/S transcriptional program (Asano and
Wharton, 1999; Duronio and O'Farrell, 1994; Royzman et al., 1997). A positive feedback
loop, in which CyclinE/Cdk2 hyperphosphorylates and inactivates Rbf, the E2F1
repressor, poses a mechanism for upregulating cyclinE. CyclinE activates expression of
the CyclinE/Cdk2 inhibitor dacapo, which encodes a CIP/KIP family member, thereby
inhibiting CyclinE/Cdk2 (de Nooij et al., 2000; Lane et al., 1996). Dacapo presents a
mechanism for downregulating CyclinE protein levels by inhibiting CyclinE/Cdk2
activity allowing accumulation of hypophosphorylated Rbf, which shuts off E2F1mediated cyclinE expression. Together, these regulatory loops may define a biphasic
oscillator for endocycles that ensures alternating S and G1 phases in endocycles (Edgar
and Orr-Weaver, 2001).
Both developmental signaling and growth signaling play key roles in endocycle
progression. During Drosophilaembryogenesis, endocycles occur in a precise pattern
corresponding to developmental stage (Smith and Orr-Weaver, 1991). Notch signaling is
involved in the mitotic to endocycle switch in Drosophilafollicle cell endocycles and
induces transcriptional changes in dacapo,fzr and string (Deng et al., 2001; Schaeffer et
al., 2004; Shcherbata et al., 2004). Drosophilalarval endocycles are inhibited by nutrient
deprivation (Britton and Edgar, 1998) and Ras and c-myc overexpression in mitotically
proliferating cells induce growth and hasten the G1/S transition, suggesting that growth
signaling also regulates the cell cycle (Johnston et al., 1999; Prober and Edgar, 2000). A
study of CIn3 translation in S. cerevisiae suggests another mechanism for coupling
growth to the endocycle. Cln3 mRNA carries a 5' untranslated open reading frame (ORF)
that reduces the efficiency of Cln3 translation, suggesting that Cln3 protein synthesis
reflects cellular ribosomal content (Polymenis and Schmidt, 1997). cyclinE mRNA
carries a number of 5' untranslated ORFs and mutations in eIF4A, a translation initiation
factor, confer defects in DNA replication (Galloni and Edgar, 1999). These results
suggest that cyclinE may act as a growth sensor in metazoan cells.
Gene amplification
Gene amplification is a replication-based method for increasing the output of a
gene. Replication based gene amplification occurs in Amphibians, Insects and the marine
ciliate Tetrahymena thermophila.Amphibians, Tetrahymena and Pterygotan insects
amplify rDNA through extrachromasomal mechanisms. Dipteran insects employ gene
amplification to increase the copy number of structural genes required for egg maturation
(Claycomb and Orr-Weaver, 2005).
The best-characterized example of gene amplification occurs in follicle cells
during Drosophilaoogenesis. During Drosophilafollicle cell gene amplification,
repeated replication initiation events, occurring without detectable gap phases, generate
high copy numbers of chorion genes required for eggshell formation (Spradling, 1981).
These replication initiation events generate multiple tandem replication forks that move
bi-directionally and generate an onion-skin structure (Fig. 1). Gene amplification occurs
at a handful of genomic sites known collectively as the DrosophilaAmplicons of follicle
Figure 1. Gene amplification in Drosophila follicle cells occurs by repeated replication
initiation events that generate tandem replication forks moving away from a central region.
Gene amplification is visualized by BrdU-incorporation at follicle cell amplicons. (A-C) BrdU
incorporation in single CantonS wildtype follicle cells in stage 10B (A), stage 11 (B) and stage
13 (C) egg chambers. In stage 10B egg chambers, follicle cells show 4-6 BrdU-labeled foci
corresponding to replication at follicle cell amplicons. In the example shown (A), 4 BrdUlabeled foci are evident and the major focus (arrow) corresponds to repeated replication initiation
events generating bi-directional, tandem replication forks at DAFC-66D. Replication initiation
and the generation of tandem replication forks in stage 10B are diagrammed to the right of (A)
with BrdU-labeling shown in pink. In stage 11 (B), only DAFC-66D (arrow) continues to gene
amplify. The last replication initiation events at DAFC-66D occur during this stage although
replication elongation continues through stages 12 and 13. By stage 13, BrdU-labeling reveals a
double bar structure at DAFC-66D (arrow). The double bar corresponds to BrdU incorporated by
opposing sets of tandem replication forks - diagrammed to the right of (C) with BrdU
incorporation in pink. Tandem replication forks have moved apart following cessation of
replication initiation events in stage 11 prior to the start of BrdU-labeling and incorporate BrdU
at distant sites.
stage 10B
0--
stage 11
~·lc~\·
~11"1~·
~I~
~c~s
CeLIIIII~
stage 13
IIIIILe
~cllrrr,
Cells, or DAFC. During gene amplification replication initiation events are
developmentally regulated: DAFC-66D undergoes replication initiation events in stages
10B and 11 and only replication elongation occurs from stage 12 on (Fig. 1). (Claycomb
et al., 2004; Claycomb et al., 2002). Replication elongation can be visualized
cytologically by BrdU-incorporation at elongating replication forks (Fig. 1) (Claycomb et
al., 2002).
Gene amplification is regulated by CyclinE. High CyclinE levels are thought to
restrict genomic replication (Calvi et al., 1998). DAFC escape this control and conserved
replication factors, components of the Pre-Replication Complex (Pre-RC) localize to the
amplicons (Asano and Wharton, 1999; Austin et al., 1999; Claycomb et al., 2002;
Royzman et al., 1999; Whittaker et al., 2000).
Cyclin/Cdk and regulation of the Pre-ReplicationComplex
Low Cyclin/Cdk activity in G of mitotic and endocycles is required for assembly
of the Pre-Replication Complex (Pre-RC) at origins of replication. The Pre-RC is an
assembly of the Origin Recognition Complex (ORC), Double Parked/Cdtl, Cdc6 and the
Mcm2-7 complex (Dutta and Bell, 1997). Regulated assembly of the Pre-RC in G
licenses origins of replication for firing in S-phase. Once-per-cell-cycle assembly
enforces once-per-cell-cycle genomic replication. Cyclin/Cdk phosphorylation of the PreRC regulates nuclear compartmentalization, chromatin association, protein stability, and
activity of complex members (Findeisen et al., 1999; Hendrickson et al., 1996; Ishimi et
al., 2000; Labib et al., 1999). Redundant, Cyclin/Cdk dependent mechanisms for
regulating Pre-RC assembly is supported by studies of re-replication in yeast in which
multiple Pre-RC components must be disrupted to achieve significant re-replication
(Gopalakrishnan et al., 2001; Nguyen et al., 2001). More recently, the direct binding of
Cyclins to replication factors (Clb2 to Cdc6 and Clb5 to Orc6) in S. cerevisiae have
presented a novel Cyclin-dependent mechanism for inhibiting Pre-RC assembly (Mimura
et al., 2004; Wilmes et al., 2004).
Gene amplification is different from the mitotic and endocycle in that there are no
detectable gap phases. CyclinE levels are constitutively high throughout the nucleus
(Calvi et al., 1998). In spite of this, the Pre-RC localizes to DAFC. Orc2, Orcl, Orc5,
DUP/Cdtl and Mcm2-7 localize to amplifying foci (Asano and Wharton, 1999; Austin et
al., 1999; Claycomb et al., 2002; Royzman et al., 1999; Whittaker et al., 2000) and
mutations in Pre-RC components Orc2, DUP/Cdtl, and Mcm6 result in reduced gene
amplification and corresponding thin eggshells (Landis et al., 1997; Schwed et al., 2002;
Whittaker et al., 2000). CyclinE/Cdk2 may be locally regulated or a molecular switch
may function to allow reiterative replication initiation. In addition, an ortholog of the Sphase kinase Dbf4/Cdc7, which is required for replication initiation in mitotic cells, may
be required for gene amplification. Mutants in chiffon, which shows homology to the
Cdc7 kinase cofactor Dbf4, displays reduced gene amplification (Landis and Tower,
1999).
Replication Initiation
Eukaryotic origins of replication
Genetic screens for autonomously replicating sequences (ARS) led to the
identification of sequence-conserved origins of replication in S. cerevisiae (Stinchcomb
et al., 1979; Struhl et al., 1979). Origins of replication bind ORC, which nucleates PreRC assembly thereby defining sites of replication initiation. S. cerevisiaeorigins of
replication are atypical for eukaryotes in that they consist of several well-defined
sequence elements (10-20 bp) spread over an approximate 200 bp interval. The Aelement consists of the 11 bp ARS consensus sequence (ACS) and is necessary but not
sufficient for origin activity. B-elements contribute to origin activity to varying degrees
but as a group, are essential (Dutta and Bell, 1997). Like S. cerevisiae, S. pombe and
Yarrowia lipolytica origins demonstrate ARS activity but unlike S. cerevisiae, they lack
consensus sequences (Vernis et al., 1997; Vernis et al., 1999). These origins are AT rich
and approximately 1 kb in size and S. pombe Orc4p carries a specialized binding domain
that recognizes AT rich sequences (Chuang and Kelly, 1999).
Metazoan replicators are more complicated. Generally, yeast replicators
encompass a single, preferred replication initiation site or origin of replication. Metazoan
origins may encompass several replication initiation sites without a predominant
preferred origin of replication and range in size from 1 to 50 kb (Bielinsky and Gerbi,
2001). Comparisons of these replicators have not yielded consensus sequences or
conserved sequence features. Metazoa show developmental plasticity and cell-type
specificity in origin usage and this may indicate the epigenetic nature of metazoan origins
(Blow, 2001). In Drosophilagene amplification at DAFC-66D, the ORC binding sites are
sequence defined at ACE3 (Amplification Control Element 3) and orif3 (Austin et al.,
1999).
Epigenetic determinantsof metazoan origin activity
Chromatin structure, including covalent modification of DNA and modifications
of chromatin packaging proteins, regulate origin usage. CpG methylation inhibits
replication initiation: Methylated DNA does not bind ORC in Xenopus egg extracts, CpG
islands are correlated with metazoan origins, and methylase deficient cell lines show less
localized replication (Delgado et al., 1998; Gilbert, 2004; Harvey and Newport, 2003a;
Rein et al., 1999). In addition, chromatin structural changes such as histone acetylation
affect replication initiation. In Drosophila,during follicle cell gene amplification,
mutations in the histone deacetyltransferase Rpd-3 result in nuclear localization of ORC
and genomic replication, suggesting that histone acetylation promotes origin usage.
Consistent with this, tethering Rpd-3 or polycomb to gene amplifying origins of
replication reduces replication while tethering the acetyltransferase Chameau increases
replication (Aggarwal and Calvi, 2004).
Transcription may promote an open chromatin configuration that is favorable for
replication initiation. Numerous transcription factors affect Drosophilafollicle cell gene
amplification. Mutants in E2F1, E2F2, DP, Rbfl, Myb, Mipl20 and Mipl30 lead to gene
amplification defects (Beall et al., 2002; Bosco et al., 2001; Cayirlioglu et al., 2003;
Royzman et al., 1999). Mutants in E2F2, Myb and Mipl30 disrupt origin specification in
Drosophilafollicle cells. These mutants undergo genomic replication rather than gene
amplification (Beall et al., 2004; Beall et al., 2002; Cayirlioglu et al., 2001; Cayirlioglu et
al., 2003). Transcription factors may impinge on gene amplification by recruiting
chromatin re-modeling factors, which influence origin usage (Aggarwal and Calvi, 2004;
Bosco et al., 2001).
Topological factors may also affect replication initiation. Eukaryotic replication
origins are distributed in intergenic regions (Wyrick et al., 2001). Active transcription
may generate negative supercoils in intergenic regions, which may play a role in origin
specification. Interestingly, DrosophilaORC has been shown to bind preferentially to
negatively supercoiled DNA (Remus et al., 2004).
The Pre-ReplicationComplex
The order of Pre-Replication Complex assembly was determined by
immunodepletion experiments in Xenopus and through experiments utilizing
temperature-sensitive alleles in yeast. In summary, ORC is required for Cdc6 and
DUP/Cdtl binding to origins, which are required, in turn, for loading of the Mcm2-7
complex, the putative replicative helicase (Maiorano et al., 2000; Tanaka et al., 1997).
Pre-RC assembly is regulated by Cyclins by Cyclin/Cdk phosphorylation and inhibitory
binding by Cyclins (Mimura et al., 2004; Wilmes et al., 2004). The Pre-RC is conserved
in yeast, flies, mammals, Xenopus and plants, suggesting that mechanisms for replication
are conserved across eukaryotes. In addition, Pre-RC components are necessary for
mitotic cell cycles, endocycles and Drosophilafollicle cell gene amplification indicating
conserved mechanisms across different modes of DNA replication.
A molecular mechanismfor Mcm2-7 loading at the Pre-ReplicationComplex
In S. cerevisiae, ORC has been shown to bind DNA cooperatively with Cdc6 in
an ATP-dependent manner (Speck et al., 2005). In Xenopus, Cdc6 is known to stabilize
ORC binding to chromatin (Harvey and Newport, 2003b). In Xenopus, DUP/Cdtl
localizes to origins after Cdc6 (Tsuyama et al., 2005). Cdc6 ATPase activity is ORC- and
origin DNA-dependent and functions at a step prior to ORC ATP hydrolysis, which is
required for Mcm2-7 loading (Bowers et al., 2004). Loss of Cdc6 ATPase activity
stabilizes DUP/Cdtl at origins and prevents Mcm2-7 loading. As is the case in S.
cerevisiae, ORC and Cdc6 ATPase activities are required for Mcm2-7 loading (Harvey
and Newport, 2003b). These observations suggest a molecular machine that loads Mcm27 onto replication origins in an orderly manner in the following scheme: ORC and Cdc6
bind cooperatively to origins. DUP/Cdtl localizes after Cdc6 binding. ORC-Cdc6
cooperative binding triggers Cdc6 ATPase activity. ORC ATPase activity soon follows
and Cdc6 ATPase activity either directly or indirectly results in re-modeling of the
complex accompanied by Mcm2-7 loading and DUP/Cdtl release (Fig. 2).
Mechanisms of Cyclin/Cdk regulationof the Pre-ReplicationComplex
Cyclin/Cdk regulation of the Pre-RC is varied and redundant and regulates
nuclear compartmentalization, chromatin binding, catalytic activity and protein stability.
In Xenopus and Drosophila,high Cyclin/Cdk activity inhibits ORC binding to chromatin
(Findeisen et al., 1999; Remus et al., 2005). In mammals, Cdk phosphorylation of Cdc6
at N-terminal sites exposes a nuclear export signal leading to nuclear export in S phase
Figure 2. Replication initiation.
In mitotic and endocycles, Pre-Replication Complex assembly at origins of replication is
restricted to G1 during a period of low Cyclin/Cdk (CDK) activity. Regulation of PreReplication Complex assembly is key to once-per-cell-cycle control of DNA replication. (A)
Pre-Replication Complex assembly begins with cooperative ORC and Cdc6 binding to origin
DNA. DUP/Cdtl localizes after Cdc6. Sequential ATP hydrolysis by Cdc6, then ORC, leads to
Mcm2-7 loading and completion of the Pre-Replication Complex (B). (C) At replication
initiation, McmlO localizes. S-phase kinases Dbf4/Cdc7 (DDK) and Cyclin/Cdk (CDK) are
required for localization of Cdc45, which subsequently travels with the replication fork. Sld3 is
required for Cdc45 localization during replication initiation.
A. Cdc6 and ORC cooperative binding
;/0,
Cdc6 ATPase, ORC ATPase
B. The Pre-Replication Complex
~KLJX~
DDK, CDK
C. Generation of the functional helicase
Duj
sZ74ýý
(Delmolino et al., 2001). In addition, Cdc6 degradation is signaled by ubiquitylation by
the SCF/Cdc4 E3 ligase in S. cerevisiae(Perkins et al., 2001) and in vitro experiments
indicate that Cdc6 degradation occurs following Cyclin/Cdk phosphorylation of Cdc6
(Elsasser et al., 1999). Pre-RC assembly is also regulated by Cyclins binding to ORC and
Cdc6. Clb5 binds to and inhibits Orc6 and Clb2 binds to and inhibits Cdc6 in S.
cerevisiae(Mimura et al., 2004; Wilmes et al., 2004).
Cyclin/Cdk regulation of DUP/Cdtl affects chromatin binding and protein
stability. CyclinA/Cdk phosphorylation has been shown to inhibit chromatin binding of
human DUP/Cdtl in vitro and inhibition of Cdkl activity in murine cells leads to
accumulation of dephosphorylated DUP/Cdtl onto chromatin (Sugimoto et al., 2004). In
addition to chromatin localization, Cyclin/Cdk regulates DUP/Cdtl abundance.
CyclinA/Cdk2 phosphorylaton of human DUP/Cdtl has been shown to promote
DUP/Cdtl binding to the Skp2 F-box protein, a cofactor for the SCF E3 ligase, which has
been shown to target DUP/Cdtl for proteolysis (Nishitani et al., 2006; Sugimoto et al.,
2004). In Drosophila,consensus Cdk phosphorylation sites in the N-terminus of
DUP/Cdtl are required for cell cycle dependent degradation and DUP/Cdtl degradation
appears to be dependent on CyclinE/Cdk2 phosphorylation (Thomer et al., 2004).
Cyclin/Cdk regulates chromatin binding of the MCM complex. Phosphorylation
of Xenopus Mcm4 by CyclinB/Cdkl reduces its affinity for chromatin (Hendrickson et al,
1996). There is also evidence that Cdk2 phosphorylation of Mcm2 primes this subunit for
phosphorylation by other kinases regulating chromatin association (Montagnoli et al.,
2006).
The transitionto replication
Following formation of the Pre-RC, the transition to replication requires the
loading of multiple factors that unwind DNA and localize DNA polymerases (Fig. 2)
(Pacek et al., 2006; Wohlschlegel et al., 2002). These include McmlO, Cdc45/Sld3, th
GINS complex and Dpbl1/Sld2. In Xenopus and S. cerevisiae,Mcml0O binds to origins in
an Mcm2-7 dependent manner. Cdc45 localization, in turn, is dependent on Mcml0O
(Sawyer et al., 2004; Wohlschlegel et al., 2002) and, in yeast, on Sld3 with which it
forms a complex (Kamimura et al., 2001; Kanemaki and Labib, 2006; Nakajima and
Masukata, 2002). Two other protein complexes, localize to origins at replication. These
are the GINS complex and Dpbl 1/Sld2. This assembly of complexes is required for
recruiting DNA polymerases. Mcml0O and Cdc45 are primarily responsible for
localization of DNApola-primase (Mimura et al., 2000; Ricke and Bielinsky, 2004;
Uchiyama et al., 2001; Zou and Stillman, 2000). Recruitment of DNApolE to origins
requires the GINS complex and Dpbl 1/Sld2 (Takayama et al., 2003).
Two S-phase kinases regulate replication initiation: Cyclin/Cdk and Dbf4/Cdc7.
In Xenopus, Dbf4/Cdc7 associates with chromatin in an Mcm2-7 dependent manner prior
to Cdc45 localization (Jares and Blow, 2000; Jares et al., 2004). Sequential kinase
activity appears to be important for DNA replication. In a Xenopus cell-free system,
exposure of chromatin to Dbf4/Cdc7 and Cyclin/Cdk2 promoted efficient DNA
replication but exposure of chromatin to these kinases in the reverse order did not
(Walter, 2000). Cyclin/Cdk phosphorylation may promote initiation by promoting
Dpbl 1/Sld2 assembly. Mutation of all the potential Cdk phosphorylation sites of Sld2 has
been shown to inhibit complex assembly and replication (Tak et al., 2006).
Replication Elongation
Several of the proteins involved in the transition to replication and in replication
fork biogenesis travel with the replication fork. Mcml0, Cdc45, and the GINS complex
have been identified at replication forks in Xenopus and S. cerevisiae (Aparicio et al.,
1997; Calzada et al., 2005; Gambus et al., 2006; Pacek et al., 2006). Mutants in S.
cerevisiae McmlO0 and Cdc45 show stalled replication forks (Merchant et al., 1997;
Tercero et al., 2000). Recently, a Cdc45/Mcm2-7/GINS (CMG) complex associated with
helicase activity was purified from Drosophilaextracts suggesting that this complex
comprises the replication fork helicase (Moyer et al., 2006).
Three DNA polymerases localize to the replication fork: DNApola-primase,
DNApol8 and DNApolE (Garg and Burgers, 2005). DNA pola-primase possesses RNA
and DNA polymerase activities and mediates priming at initiation to start leading strand
synthesis and travels with the replication fork to prime lagging strand synthesis.
DNApolI is thought to mediate Okazaki fragment maturation during lagging strand
synthesis. DNApols is thought to perform leading strand synthesis. In addition, RPA,
RFC, and PCNA are loaded. RPA consists of three subunits and is functionally
homologous to E. coli SSB and binds to single-stranded DNA. PCNA is a trimeric sliding
clamp that increases the processivity of DNA polymerases. RFC loads PCNA.
Replication Fork Progression
Replication Fork Progression (RFP) is regulated by DNA secondary structure and
chromatin bound proteins. Differences in Replication Fork Speed (RFS) have been
observed for cells employing different modes of DNA replication. In Drosophiladiploid
cells, replication forks move at ~2.6 kb/min (Blumenthal et al., 1974). The RFS for
Drosophilapolytene larval salivary glands has been measured at ~300 bp/min
(Steinemann, 1981). During gene amplification, replication forks move at about ~50-100
bp/min (Spradling and Leys, 1988). These differences may be due to chromatin structure
(for example, the persistence of cohesins on polytene chromosomes) or topological
factors. Multiple, tandem replication forks during gene amplification may generate
significant superhelical strain. At DAFC-66D, -~5 replication forks moving in the same
direction are spaced -~10kb apart.
RFS and RFP are also regulated by protein factors at the replication fork.
Helicases are common targets for modulating RFP. Replication fork progression at
normal speeds through S. cerevisiae telomeric and subtelomeric sequences requires the
Rrm3p helicase, suggesting that cellular helicases have specialized functions (Ivessa et
al., 2002). Not surprisingly, replication fork pausing mechanisms often target the
helicase. During prokaryotic replication termination, the trans-acting factors Tus in E.
coli and RTP in B. subtilis bind to and inhibit the replicative helicase (Bussiere and
Bastia, 1999). In a proof-of-principle, a protein inhibitor of the replicative helicase was
shown to reduce replication fork speed In E. coli, (Skarstad and Wold, 1995). Helicase
activity is regulated by Cyclin/Cdk phosphorylation (Ishimi et al., 2000), and this may be
an important mechanism for regulating replication fork speed.
Regulation of replication fork progession may be important for gene expression.
Polar intergenic Replication Fork Barriers (RFB), block replication fork progression
opposing transcription at the rDNA locus of S. cerevisiae, S. pombe, mouse and Xenopus
(Rothstein et al., 2000). In E. coli, head-on transcription severely inhibits replication fork
progression while co-directional transcription does not, revealing the importance of
coordinated transcription and replication (Mirkin and Mirkin, 2005). The interplay
between DNA replication machinery and transcriptional machinery is not necessarily
direct. Transcription factors may affect nucleosome arrangement or recruit chromatin remodeling machinery (Bosco et al., 2001). During Drosophilafollicle cell gene
amplification, mutants in transcription factors affect replication initiation (Beall et al.,
2004; Beall et al., 2002; Bosco et al., 2001; Royzman et al., 1999). Some of these
mutants may affect replication fork progression.
Summary
The mechanisms regulating DNA replication are conserved between mitotic cell
cycles, endocycles and Drosophilafollicle cell gene amplification. A period of low
Cyclin/Cdk, a G1 phase, in which the Pre-RC can assemble is conserved between mitotic
and endocycles. The Pre-RC is conserved during Drosophilafollicle cell gene
amplification. Mechanisms for regulating replication initiation are conserved between
different modes of DNA replication. Aspects of replication fork progression are likely to
be conserved. Mcm2-7, Cdc45 and PCNA travel with the replication forks in mitotic cells
(Claycomb et al., 2002; Loebel et al., 2000). These proteins co-localize with replication
forks during gene amplification. In addition, DUP/Cdtl, co-localizes with replication
forks during gene amplification. It has been postulated that DUP/Cdtl is required at the
replication forks to maintain Mcm2-7 at slow-moving replication forks (Claycomb et al.,
2002).
We performed a screen to identify developmental regulators of mitotic and
endocycles during Drosophilaembryogenesis. Three cell cycle phenomena - cell cycle
exit, mitotic cell cycles and endocycles - occur concurrently during Drosophila
embryogenesis. We took advantage of this period to screen for developmental regulators
of S-phase using PCNA, as an S-phase marker. We identified a class of mutants that
displayed large nuclei in normally diploid tissues. In this class, we identified new
mutations in pavarotti and tumbleweed, pav3C53 and tum32a-20, which are required for
cytokinesis. In addition, we characterized a new cyclinE mutant, cyclinElf36 , displaying
increased replication fork progression during gene amplification. This previously
undescribed gene amplification defect implicates CyclinE in regulating replication fork
progression and raises the intriguing possibility that replication fork progression is plastic
during gene amplification and may be subject to developmental regulation.
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Vernis, L., A. Abbas, M. Chasles, C.M. Gaillardin, C. Brun, J.A. Huberman, and P.
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Short DNA fragments without sequence similarity are initiation sites for
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CHAPTER Two
New mutants affecting developmentally regulated cell cycles during
Drosophila embryogenesis
Eugenia A. Park and Terry L. Orr-Weaver
Whitehead Institute and Dept. of Biology, Massachusetts Institute of Technology,
Cambridge, MA 02142
SUMMARY
During Drosophilaembryogenesis, cell cycle exit, mitotic cell cycles and endocycles all
occur within a narrow time span. The developmental patterning of these cell cycles are
well characterized and afford a unique opportunity to study developmental regulation of
the cell cycle. We screened 300 EMS mutants for developmentally uncoordinated
replication by an in situ hybridization assay for PCNA transcription, a marker of the G1/S
transition. We identified 30 mutants that may reflect functions in developmental
signaling, G1/S transcription and DNA replication. We further characterized a class of
mutants displaying large, apparently polyploid nuclei in normally diploid cells. 3C157,
Ir8 and 2k32 - of which 3C157 and Ir8 are allelic - displayed large and diffuse DNA
masses in isolated cells of the nervous system. 32a-20 (formerly 32a) and 3C53 displayed
large nuclei in the nervous system and epidermis. We cloned these mutants and identified
new alleles of tumbleweed, tum3 2a-20 , and pavarotti, pavC3 53. These mutants have defects
consistent with blocks to cytokinesis.
INTRODUCTION
In metazoans, divergent cell cycles must be regulated accurately throughout
development to build and maintain a viable organism. The G1/S/G2/M cell cycle is only
one of multiple cell cycles. Variants of the archetypal cell cycle are utilized in different
developmental contexts to achieve different aims. The early embryonic divisions of
insects, marine invertebrates and amphibians are rapid S/M cycles. These cycles allow for
speedy embryogenesis, potentially important for organisms with exposed and vulnerable
embryos. Endocycles are variants of the archetypal cell cycle that lack complete mitoses
but consist of discrete S and G1 phases with one round of DNA replication occurring per
endocycle (Smith and Orr-Weaver, 1991). These cycles generate polyploidy and are
thought to be a strategy for increasing growth and metabolic capacity without the large
scale cytoskeletal rearrangements required by mitosis (Edgar and Orr-Weaver, 2001).
A key requirement for all of these cell cycles is the restriction of Cyclin/Cdk
activity. A window of low Cyclin/Cdk activity is required for assembly of pre-replication
complexes at replication origins (Hua et al., 1997). During G1/S/G2/M cycles,
Cyclin/Cdk activity remains high throughout the cell cycles except for a window in early
G1 following downregulation of Cyclins A and B and preceding upregulation of CyclinE.
During the early embryonic S/M cycles of Xenopus, nuclear compartmentalization
restricts Cyclin/Cdk activity (Blow and Laskey, 1988). Some endocycling tissues show
vestiges of mitosis ranging from chromosome condensation to nuclear envelope
breakdown (Edgar and Orr-Weaver, 2001) and utilize mitotic machinery to achieve low
Cyclin/Cdk activity. Mutations in morula, an APC subunit, lead to ectopic spindle
formation and chromosome condensation in Drosophilanurse cells suggesting that
regulators of mitosis are expressed at low levels in these cells (Kashevsky et al., 2002;
Reed and Orr-Weaver, 1997). To modulate activity, Cyclin/Cdk complexes are regulated
on many levels including oscillatory Cyclin expression, regulatory phosphorylation,
inhibition by Cyclin/Cdk inhibitors (CKI), and through targeted degradation of Cyclins.
Developmental signaling plays a key role in regulating different cell cycles. In
Drosophila,Notch signaling mediates a mitotic to endocycle switch in ovarian follicle
cells, and no less than 36 pattern formation genes are involved in the developmental
regulation of mitotic cell cycles during embryogenesis (Deng et al., 2001; Edgar et al.,
1994; Keller Larkin et al., 1999; Schaeffer et al., 2004; Shcherbata et al., 2004).
Drosophilaembryogenesis provides an elegant example of developmental regulation of
the cell cycle. Cell cycle exit and three different cell cycles - S/M, S/G2/M and S/G occur dynamically in a 6 hour time span (Fig. 1). Cycles 1-13 consist of rapidly
alternating S and M phases and are nuclear divisions that occur more or less
synchronously in a common cytoplasm. Cycles 14-16, the postblastoderm divisions,
consist of S, G2 and M phases. These divisions (S/G2/M) occur in mitotic domains in
which cells differentiating into the same tissue undergo mitosis synchronously and at the
same developmental time (Foe et al., 1993). Following cycle 16, the embryonic epidermis
exits the cell cycle while cells of the developing nervous system continue to undergo
mitotic cycles. Also following cycle 16, the developing larval tissues initiate endocycles
consisting of S and G phases and continue these through embryogenesis and larval
development resulting in highly polytene tissues. Embryonic endocycles occur in
spatiotemporal domains reminiscent of the mitotic domains of the postblastoderm
divisions (Fig. 2) (Smith and Orr-Weaver, 1991).
Figure 1. Diagram of variant cell cycles utilized during embryogenesis.
Following fertilization, cell cycles 1-13 are syncitial divisions that occur synchronously
in a common cytoplasm. These rapid S/M cycles correspond to stages 1-8 of embryonic
development. By cycle 14, cellularization is complete and a gap phase is added
coincident with a requirement for zygotic string/CDC25transcription. Cycles 14-16 (the
postblastoderm divisions) consist of S/G2/M phases and occur in mitotic domains in
which groups of cells differentiating together undergo mitoses at different times. The
dorsal epidermis undergoes cycles 14, 15, and 16 in stages 9, 10 and 11 respectively.
Following cycle 16, the epidermis exits the cell cycle in stages 11 and 12 (not shown).
The developing nervous system continues to cycle mitotically while the endodomains, or
developing larval tissues, initiate S/G endocycles in stage 11. Shown in the image is a
stage 12 embryo (anterior is to the left) in situ hybridized with antisense riboprobe
against PCNA, which is expressed at G1/S, to visualize the nervous system (NS) and the
endodomains (ENDO).
1-13
Syncytial Divisions
M
4
14-16
Postblastoderm
Divisions
G2
M!
S
M
G2
17
Endo Cell
Cycle
17
Mitotic
Cell Cycle
I
CNS
ENDO
\\
Figure 2. Embryonic endocycles in the larval tissues occur in developmentally
regulated spatiotemporal domains.
(A-F) Endocycle domains are shown in color at the developmental stage in which they
undergo DNA replication (figure taken from Smith & Orr-Weaver, 1991). Gray shading
indicates larval tissues not undergoing DNA replication in a given developmental stage.
(A) As the germband begins to retract in stage 11, the salivary gland undergoes the first
detectable endocycle S-phase 3 hours after the last mitosis (green). (B) During germband
retraction in stage 12, the anterior and posterior midgut (am and pm) undergo DNA
replication (red) along with isolated large nuclei between am and pm (red dots). (C)
During dorsal closure in stage 13, DNA replication continues in the anterior and posterior
midgut (red). The hingut undergoes DNA replication (hg, yellow) and shortly thereafter
the Malpighian tubules initiate DNA replication 4.5 hours after the last mitosis (mt,
purple). (D) During head involution in stage 14, replication continues in the anterior and
posterior of the midgut (red), hindgut (yellow) and Malpighian tubules (purple). In
addition, a 2 nd round of DNA replication initiates in a stripe in the central region of the
midgut (mg, blue). (E) In stage 15 with the first constriction of the sac-like central
midgut, the stripe of replicating tissue expands anteriorly and posteriorly (blue) and a
group of dorsally located cells extending along the posterior part of the embryo
undergoes DNA replication (pink). (F) In stage 16 as the midgut convolutes, replication
extends throughout the midgut (blue) and continues in the dorsally located cells (pink).
A
P
vi
ml
A
L
~KJ~j LI)I
F
dc
IF
~CCLL/LI
Transcriptional regulation is key to developmental control of the cell cycle. The
mitotic domains of the postblastoderm divisions arise from the zygotic transcription of
string/CDC25phosphatase, an activator of Cdkl that regulates the G2/M transition
(Edgar et al., 1994; Edgar and O'Farrell, 1990). String (stg) transcription occurs in pulses
anticipating mitoses in each mitotic domain. Epidermal cell cycle exit requires
developmentally coordinated changes in the transcriptional regulation of three genes.
cyclinE transcription is downregulated. fizzy-related (fzr), an APC co-activator related to
S. cerevisiae Cdhl, is transcriptionally upregulated. dacapo (dap), a CIP/KIP family
CyclinE/Cdk2 inhibitor, is also transcriptionally upregulated (de Nooij et al., 1996;
Knoblich et al., 1994; Lane et al., 1996; Sigrist and Lehner, 1997). Perturbation of any of
these transcriptional regulatory events leads to a complete additional round of cell
division. Embryonic endocycles coincide with pulses of cyclinE andfzr transcription
(Knoblich et al., 1994; Sigrist and Lehner, 1997). Transcription of cyclinE andfzr occurs
in a spatiotemporal pattern mirroring endocycle S-phases and expression of both genes is
required for endocycle S-phase (Knoblich et al., 1994; Sigrist and Lehner, 1997). The
periodic transcription of cyclinE is clearly important for endocycles, as continuous
ectopic expression of cyclinE inhibits S-phases (Sauer et al., 1995). fzr transcription is
probably required to reduce mitotic Cyclin/Cdk activity (Reed and Orr-Weaver, 1997;
Sigrist and Lehner, 1997). The complex developmental control of cell cycle regulators is
almost certainly due to complex promoters that contain modular cis-elements. The
cyclinE and string promoters span 10 kb and 15 kb respectively (Edgar et al., 1994; Jones
et al., 2000). Tissue-specific elements have been defined in the promoters of cyclinE,
string and dacapo and multiple developmental cues are likely to converge on these
promoters (Deng et al., 2001; Edgar et al., 1994; Jones et al., 2000; Meyer et al., 2002;
Schaeffer et al., 2004).
In addition to stg, cyclinE,fzr, and dap, an E2F regulated G1/S transcriptional
program is developmentally regulated during embryogenesis (Asano and Wharton, 1999;
Duronio and O'Farrell, 1994; Duronio and O'Farrell, 1995; Royzman et al., 1997;
Whittaker et al., 2000). This program precedes S-phase in the endodomains.
Developmentally coordinated E2F mediated transcription is required for normal
endocycle S-phase (Duronio et al., 1998; Royzman et al., 1997). This program includes
PCNA (DNA polymerase processivity factor), RNR1 and RNR2 (ribonucleotide reductase
subunits), Double Parked(a replication initiation factor), Orcl (a component of the
replication initiator complex) and cyclinE (Asano and Wharton, 1999; Duronio and
O'Farrell, 1994; Royzman et al., 1997; Whittaker et al., 2000).
To better understand the relationships between variant cell cycles during
development, we undertook a screen to identify regulators of mitotic cycles and
endocycles during embryogenesis. We screened 300 EMS-mutagenized 3 "dchromosome
lines (Moore et al., 1998) and recovered 26 mutants falling into 5 phenotypic classes
reflecting functions in transcriptional regulation and developmental regulation of Sphase. In addition, we identified a class of mutants in which normally diploid cells
became polyploid, suggesting functions that distinguish mitotic cycles from endocycles
during development. Amongst these, we cloned mutations in pavarotti, encoding a
kinesin-like protein, and tumbleweed, encoding a Rho-family GAP.
RESULTS
A screenfor mutations affecting developmental cell cycle regulation
To recover mutations altering developmental cell cycle regulation during
embryogenesis, we looked for alterations in the pattern of expression of PCNA, a
component of the G1/S transcriptional program. PCNA is expressed in the mitotically
proliferating nervous system and endocycling larval gut, allowing examination of both
tissues. Previously, Irena Royzman and Allyson Whittaker screened 700 EMS
mutagenized 3rd chromosome lines, from a collection of mutagenized lines generated in
the laboratory of Ruth Lehmann (Moore et al., 1998), for aberrant PCNA expression in
the interest of identifying developmental regulators of G1/S transcription (Moore et al.,
1998; Royzman et al., 1997). We were primarily interested in identifying developmental
regulators of DNA replication using PCNA expression as a marker of DNA replication
and screened an additional 300 lines from the collection. Embryos were collected from
heterozygous flies, aged 8-15 hr, and in situ hybridized using a PCNA riboprobe as
described previously (Royzman et al., 1997). Heterozygous embryos carried a Ubx-lacZ
transgene. Homozygous embryos were detected by absence of lacZ expression, as
detected by in situ hybridization with a lacZ riboprobe. We recovered 26 mutants falling
into 5 phenotypic classes (Table 1, Fig. 3). In addition, we included three mutants
isolated by Irena Royzman and Allyson Whittaker as they fit our phenotypic classes.
Class I consisted of 6 mutants that failed to downregulate PCNA mRNA in the
anterior and posterior midgut in stage 14 embryos (Table 1, Fig. 3D). Mutants in cyclinE
and dup display a block to replication and fail to downregulate the G1/S transcriptional
program in the endodomains (Sauer et al., 1995; Whittaker et al., 2000). Class I mutants
Table 1. Mutants recovered from a screen for regulators of developmental cell cycle
specificity.
In situ hybridization with PCNA antisense riboprobe of EMS-mutagenized homozygous
mutant embryos identified 30 alleles falling into five phenotypic classes. Class V mutants
showed large cells in the CNS. Of these, 3C53, 3C157, Ir8, 2k32 and 32a (marked with
v5 displayed large nuclei corresponding
are mapped to the
2 nd
to large cells. (II) and (III) indicate that alleles
and 3 rd chromosomes respectively. (?) indicate that the allele has
not been mapped to a chromosome, * indicates mutants belonging to two phenotypic
classes. ** were isolated by I. Royzman and A. Whittaker. Additional phenotypic
descriptions are given under comments. Italics indicate description of PCNA expression
in tissues abbreviated as follows: AM/PM - anterior and posterior midgut, MG - central
midgut, HG - hindgut, MT - Malpighian tubules, ENDO - all endodomains (AM/PM,
MG, HG and MT), CNS - central nervous system, VNC - ventral nerve cord.
Class I
Ectopic PCNA expression
in endodomains.
Class II
Small brain and aberrant
PCNA expression in
endodomains.
Class III
Reduced PCNA expression
in CNS. No PCNA
expression in MG.
Class IV
Reduced PCNA expression.
Class V
Large cells CNS
Allele No.
3C22 (III)
3C37 (III)
3C92 (III)
3C225 (III)
3C240 (III)
3C241 (?) *
3Cll (III)
3C41 (III)
3C137 (III)
3C187 (III)
3C44 (III)
3C83 (III)
3C87 (III)
3C165 (III) *
3C168 (III)
3C249 (III)
3C115 (III)
3C118 (III)
3C148 (III)
3C]74 (?)
3C175 (?)
3C210 (III) *
3C143 (III)
3C23 (III)
V3C53 (III)
V'3C157 (III)
3C165 (III) *
3C207 (?)
3C210 (III) *
3C241 (?) *
VIr8 (III) **
'2k32 (III) **
1v32a (II) **
Comments
AM/PM in stage 14.
AM/PM in stage 14.
AM/PM in stage 14. Reduced MG.
AM/PM in subset of stage 14. Reduced MG.
AM/PM in stage 14. No MG.
AM/PM in stage 14. Large cells CNS.
AM/PM in stage 14. No HG or MT.
HG and MT domains small.
MG in all stages. No HG or MT.
HG and MT domains small.
AM/PM in stage 14.
AM/PM in stage 14.
AM/PM in stage 14.
AM/PM in stage 14. Large cells CNS.
AM/PM in subset of stage 14.
AM/PM in stage 14.
AM/PM low
ENDO low. CNS low.
ENDO low.
ENDO low.
AM/PM, MG low. Large cells CNS.
Large cells VNC. VNC disorganized.
AM/PM in stage 14. Large cells CNS.
AM/PM, MG low. Large cells CNS.
AM/PM in stage 14. Large cells CNS.
Figure 3. PCNA expression phenotypes of select mutants recovered from a screen
for regulators of developmental cell cycle specificity.
Mutant embryos were in situ hybridized with PCNA and lacZ antisense riboprobes.
Homozygotes were detected by absence of lacZ expression from an Ubx-lacZ transgene
on a balancer chromosome. (A, B) CantonS wildtype (WT) stage 13 and 14 embryos are
shown. (A) In stage 13, PCNA expression is seen in the nervous system (*), anterior and
posterior midgut (arrowheads), hindgut and Malpighian tubules (arrows) corresponding
to DNA replication in these tissues. (B) In stage 14, PCNA expression is evident in a
stripe in the central midgut (**) corresponding to 2 nd round DNA replication in this
tissue. PCNA expression in the anterior and posterior midgut is downregulated
(arrowheads). (C, D) Class I mutant 3C37 stage 13 and 14 embryos are shown. (C) In
stage 13, PCNA is expressed appropriately in the anterior and posterior midgut, hindgut
and Malpighian tubules. (D) In stage 14, PCNA inappropriately persists in the anterior
and posterior midgut (arrowheads). (E, F) Class II mutant 3C11 stage 13 and 14 embryos
are shown. Class II mutants display small brains (*) suggestive of proliferative defects.
(E) In stage 13, PCNA is expressed in anterior and posterior midgut (arrowheads) but the
hindgut and Malpighian tubules are not evident (arrows). (F) In stage 14, PCNA
expression persists inappropriately in anterior and posterior central midgut (arrowheads).
(G, H) Class III mutant 3C44 stage 13 and embryo with terminal phenotype are shown.
(G) In stage 13, 3C44 displays reduced PCNA expression in the nervous system (*)
relative to wildtype. (H) In stage 14, homozygotes fail to express PCNA coinciding with
the 2 nd round of replication in the central midgut (**).
o
(j
efd
Q
(with the exception of 3C241, which has not been mapped to a chromosome), are on the
3"
chromosome whereas cyclinE and dup are located on the 2 nd chromosome. Although
CyclinE and DUP may directly participate in G1/S transcript downregulation, we favor
the model that completion of S phase signals this event and that CyclinE and DUP act
indirectly by blocking S phase progression. We expect that Class I mutants either block
DNA replication or affect components of an active, cell-cycle dependent signaling
mechanism for G1/S transcript downregulation. These mutants may affect replication
factors.
Class II mutants had small brains and failed to downregulate transcripts in the
midgut (Table 1, Fig. 3E-F). These mutants affected both the nervous system and
endodomains, suggesting conserved function between mitotic cycles and endocycles.
3C41 and 3C187 specifically affected ectoderm-derived tissues - the central nervous
system, hindgut and Malpighian tubules. The hindgut and Malpighian tubules are
ectoderm derived unlike the midgut, which is mesoderm derived (Skaer, 1993). 3C41 and
3C187 may be ectoderm specific whereas 3C11 and 3C137, which affect both the central
nervous system and the midgut, are more likely to affect cell cycle regulators.
Class III mutants displayed reduced PCNA expression in the nervous system and a
failure to downregulate PCNA expression in the anterior and posterior midgut (Table 1,
Fig. 3G-H). Failure to downregulate PCNA in the midgut is reminiscent of cyclinE and
dup mutants in which replication is blocked (Sauer et al., 1995; Whittaker et al., 2000).
Again, neither cyclinE nor dup are on the 3 r chromosome. All Class III mutants map to
the 3 rd chromosome. We expect that Class III mutants block DNA replication. Like Class
I mutants, Class III mutants may affect replication factors.
Class IV consists of 7 mutants that displayed reduced PCNA expression in the
endodomains or in the case of 3C118, in both endodomains and the nervous system
(Table 1). This phenotype suggests defects in G1/S transcription. These mutants may
encode transcription factors.
Class V mutants displayed large cells in the nervous system (Table 1, Fig. 4). 10
mutants displayed this phenotype. Three belonged to other classes besides Class V. These
mutants are described further in the following section.
Class V mutants have large nuclei in the nervous system and epidermis
We were interested in examining how mitotic cycles and endocycles are
distinctively regulated in a developmental context. Ten Class V mutants displayed large
cells in the nervous system in our primary screen. Since cell size is linked to ploidy
(Edgar and Orr-Weaver, 2001), we expected these mutants might enumerate the
developmental mechanisms by which mitotic cycles are converted to endocycles in either
functional or pathological ways.
We conducted a secondary screen on Class V mutants to examine DNA content in
the large cells. We stained 8-15 hr embryos from each mutant line with the DNA dye
DAPI and confirmed large nuclei in the nervous system of 5 of these mutants (Table 1,
Fig. 4). The remaining mutants in this class may have large cells reflective of increased
cell size and not increased ploidy. 3C53 and 32a displayed large nuclei throughout the
nervous system while Ir8, 3C157 and 2k32 displayed large, diffuse DNA masses in
isolated cells in the nervous system (Fig 4). We conducted complementation tests on the
Figure 4. Class V mutants display large, apparently polyploid nuclei in normally
diploid cells.
Class V embryos (stages 12-14) were DAPI stained and examined for large nuclei. Of the
ten mutants initially identified as having large cells, five mutants displayed large nuclei in
the ventral nerve cord of the central nervous system as shown here. (A) CantonS wildtype
control, (B) 3C53, (C) 32a, (D) 2k32, (E) 3C157, and (F) Ir8. Scale bar is 5 ptm. (B, C)
3C53 and 32a showed large nuclei (arrowheads) throughout the nervous system. (D-F)
2k32, 3C157 and Ir8 showed large, diffuse DNA masses (arrowheads) in a subset of
nuclei.
3"
chromosome mutants, 3C53, Ir8, 3C157 and 2k32 and determined Ir8 and 3C157 to
be allelic.
Characterizationand mapping of Ir8 and 3C157
Ir8 and 3C157 displayed isolated large and diffuse DNA masses in the vicinity of
the ventral nerve cord (Fig. 4E-F). Ir8 displayed a more pronounced phenotype than
3C157 suggesting Ir8 is the stronger allele. To determine if these diffuse DNA masses
displayed characteristics of mitosis, we performed tubulin and phosphohistone H3
antibody staining (Fig. 5). Although we were not able to discern mitotic spindles with
tubulin antibody staining, we found that the large and diffuse DNA masses were
phosphohistone H3 positive (Fig. 5). Phosphohistone H3 localized to irregularly shaped
and irregularly condensed DNA masses indicating that in Ir8 and 3C157, phosphohistone
H3 does not strictly coincide with condensed DNA as observed in normal mitoses. These
observations suggest that Ir8 and 3C157 disrupt functions in chromatin dynamics,
perhaps chromosome condensation.
The large nuclei phenotype may be due to defects specific to nervous system
development or may be a consequence of the timing of the onset of zygotic regulation of
the cell cycle. The central nervous system develops via lineage specification of cells
generated by asymmetric divisions of neuroblasts. These divisions result in the generation
of both neurons and glial cells that provide support functions (Goodman and Doe, 1993).
To determine if Ir8 and 3C157 affect specific lineages, we performed ELAV antibody
staining on Ir8 and 3C157 embryos. ELAV is a pan-neuronal marker (Robinow and
White, 1988). The pattern of ELAV antibody staining did not strictly correspond with
Figure 5. Ir8 and 3C157 polyploid nuclei retain aspects of mitosis.
(A-I) Large, diffuse DNA masses in 3C157 stage 12 embryos are phosphohistone H3
positive. Arrowheads indicate diffuse, poorly condensed regions that are phosphohistone
H3 positive. The first column (A, D, G) shows anti-phosphohistone H3 staining. The
second column (B, E, H) shows DAPI staining. The third column (C, F, I) shows merged
channels with anti-phosphohistone H3 in red and DAPI in blue. Scale bar is 10 [tm.
phospho H3
3C157
3C]57
DNA
phospho H3
DNA
large nuclei - some large nuclei were ELAV positive while others were not - suggesting
polyploidy in neuronal and non-neuronal cells (Fig. 6). We favor the explanation that the
large nuclei in Ir8 and 3C157 are due to misregulation of the cell cycle and that the
nervous system specificity of the defects are due to perseverance of the maternal
contribution.
To determine the lethal stage of Ir8/3C157transheterozygotes, we placed Ir8 and
3C157 in trans to a balancer carrying an actin-GFPtransgene. We crossed these strains
and collected embryos over a 24 hour period examining 25 larvae for transgene
expression. All larvae were GFP positive suggesting that Ir8/3C157transheterozygotes
are embryonic lethal or that larvae fail to hatch.
We deficiency mapped Ir8 and 3C157 with 70 deficiencies deleting portions of
the 3" chromosome but failed to identify a deficiency uncovering the large nuclei
phenotype. We undertook meiotic mapping and mapped the large nuclei phenotype to the
genetic interval between h and cu.
Characterizationof 3C53 and 32a-20
3C53 and 32a-20, a recombinant of 32a, affected cells more or less uniformly
throughout the nervous system suggesting a general defect in mitotic cell cycles. These
mutants also appeared to have large cells in the epidermis. This phenotype suggested
either (1) defects during cycles 14-16 of the epidermis or (2) unrestricted DNA
replication in the period following cycle 16 when epidermal cells exit the cell cycle. We
eliminated unrestricted DNA replication as a potential mechanism by examining BrdUlabeled stage 12 embryos. Mutations that affect cell cycle exit result in abundant
Figure 6. Some large DNA masses in Ir8 homozygous embryos do not stain for
ELAV - a neuronal marker.
Stage 13 embryos were stained with monoclonal rat anti-ELAV (Developmental
Hybridoma Studies Bank) and DAPI. The ventral nerve cord at mid-embryo are shown
and the scale bar in panel (A) is 5 lpm. Sibling embryos of genotypes Ir8/CyO or
CyOICyO were used as control. (A-C) anti-ELAV and DAPI staining of a sibling control
embryo. (D-F) anti-ELAV and DAPI staining of an Ir8 homozygous embryo. The ventral
nerve cord as defined by ELAV staining is disorganized in Ir8 (D) but not in a sibling
control (A). Large DNA masses are evident at the periphery of the ventral nerve cord
(arrow and arrowhead). Arrowheads in panels D, E, F point to a large DNA mass that is
seemingly ELAV negative while arrows in panels D, E, F point to a large DNA mass that
is seemingly ELAV positive.
59
replication in the dorsal epidermis at this stage (Lane et al., 1996; Sigrist and Lehner,
1997). Both 3C53 and 32a-20 exited the cell cycle appropriately, suggesting that the
large nuclei in the epidermis were due to defects during cycles 14-16 (Fig. 7).
In order to examine mitoses, we performed tubulin antibody staining on 3C53 and
32a-20 homozygotes. We detected abnormal mitotic spindles associated with polyploid
nuclei in the vicinity of the ventral nerve cord of stage 12 embryos (Fig. 8). Spindles in
3C53 homozygotes appeared depleted of astral microtubules and displayed intensely
stained clumps of tubulin at spindle poles. In addition, some nuclei lacked spindles but
were decorated with tubulin clumps. 32a-20 homozyotes displayed less severe spindle
defects than 3C53. Spindles were more intact albeit morphologically abnormal - fat and
carrying tubulin clumps. The spindle defects suggested that 3C53 and 32a-20 might
affect determinants of spindle morphology.
3C53 and 32a-20 are alleles of pavarottiand tumbleweedlRacGAP50C respectively
Next, we deficiency mapped 3C53 and 32a-20. 3C53 was placed in trans to a
collection of 70 deficiencies deleting segments of the 3" chromosome. Of these,
Df(3L)GN24 and Df(3L)GN50, failed to complement the large nuclei phenotype. These
overlapping deficiencies delete the polytene salivary gland cytological intervals 63F47;64C13-15 and 63E1-2;64B17 respectively. We examined the candidate genes falling
into this region and given the spindle defects seen in 3C53 homozygotes, identified
pavarotti (pav), a kinesin-like protein previously demonstrated to play a role in
cytokinesis, as a good candidate for 3C53 (Adams et al., 1998). We obtained an allele of
pavarotti,pavB200, and determined that this allele fails to complement both lethality and
Figure 7. The dorsal epidermis of 32a and 3C53 homozygous mutant embryos exits
the cell cycle successfully.
In stage 12 of embryonic development, the dorsal epidermis exits from the cell cycle
following the postblastoderm mitotic divisions. Mutants in fzr and dap fail to exit the cell
cycle as indicated by robust BrdU-labeling of dorsal epidermal cells (Lane et al., 1996;
Sigrist and Lehner, 1997). 32a (B) and 3C53 (D) homozygous mutant embryos and
sibling control embryos (A and C) were labeled with BrdU. Stage 12 sibling control
embryos (A and C) are not labeled with BrdU in the dorsal epidermis (yellow arrows).
BrdU-labeled cells to either side of non-labeled domains are in the peripheral nervous
system, which are undergoing mitotic divisions. 32a and 3C53 homozygous embryos (B
and D respectively) were identified by the presence of large nuclei in the nervous system
and epidermis. Yellow arrows in B and D point to dorsal epidermal domains, which are
not BrdU-labeled, indicating successful cell cycle exit in the dorsal epidermis.
Figure 8. 3C53 and 32a-20 homozygotes display abnormal mitotic spindles
associated with polyploid nuclei in the nervous system.
Images in the first column (A, D, G, J, M) are stained with anti-tubulin. Images in the
second column (B, E, H, K, N) are stained with DAPI. Images in the third column (C, F,
J, L, O) show anti-tubulin in red and DAPI in blue. The scale bar shown in panel A is 5
[tm. (A-C) Spindle in nervous system of CantonS wildtype stage 12 embryo is shown.
(D-J) Spindles in nervous system of stage 12 3C53 embryos are shown. (D-F) 3C53
homozygote shows a spindle with depleted astral and interpolar microtubules. (G-J) 3C53
homozygote shows disorganized clumps of microtubules associated with a nucleus. (J-0)
Fat and wide spindles in 32a-20 homozygote stage 12 nervous system.
CantonS
3C53
32a-20
the large nuclei phenotype in trans to 3C53. Therefore, 3C53 is an allele of pavarottiand
3 3
.
henceforth we designate the allele pavCS
We meiotically mapped 32a to the genetic interval between the genes pr and cu
and recovered a recombinant chromosome, which we designated 32a-20. We tested 50
deficiencies in this region and identified Df(2R)CX1 as the sole deficiency producing the
large nuclei phenotype in trans to 32a-20. We identified a candidate interval containing
approximately 100 genes and further limited the candidate interval by male
recombinational mapping of the mutation (Preston and Engels, 1996; Preston et al.,
1996). We were able to map 32a-20 to an interval containing about 10 genes. Among
these genes, tumbleweed (tum), formerly known as racGAP50C,a Rho family GTPase
required for cytokinesis (Zavortink et al., 2005), was an attractive candidate. We
sequenced the coding region of tum and identified a Leu to His substitution in a
conserved residue in the GAP catalytic domain box II (Fig. 9). We expect that the LA64H
substitution is the critical mutation in our allele, which we designated tumrn32a20
Pav, a kinesin-like protein, and Tum, a Rho family GAP, are conserved proteins
with homologs in C. elegans, humans and mice. Pav and Tum, and their orthologs in
other organisms, interact directly, co-localize throughout the cell cycle, are functionally
interdependent and are required for cytokinesis (Kuriyama et al 2002, Hirose et al. 2001,
Mishima et al. 2002, Jantsch-Plunger et al. 2000, Raich et al. 1998, Somma et al. 2002,
Adams et al, 1998, Zavortink and Saint, 2005). Both Pav and Tum are required for the
formation of the central spindle, a bundled microtubule structure that arises in anaphase
and persists through telophase. The central spindle is required for successful cytokinesis
in mammalian cells, C. elegans and Drosophila(Straight and Field, 2000).
Figure 9. Tum32a 2 0 carries a L464H substitution in a conserved residue in the GAP
catalytic domain.
Tum is a 625 residue protein that has coiled-coil (blue box) and Cysteine-rich domains
(green box). Tum interacts with Pav and Pbl, and binding domains are at the N-terminus
(Somers and Saint 2003). Tum has a GAP catalytic domain characteristic of Rho family
GAP.
tum 3 2a-20
is a new mutant that carries an L464H substitution in box II of the GAP
catalytic domain (red box). Catalytic dead transgenic constructs carry small deletions in
GAP domain box I. The two other known Tum mutants are truncated due to nonsense
mutations. tumDHS is truncated at residue 195 while tumAR 2 is truncated at residue 470
(Jones and Bejsovec 2005).
?,0
Itt
cri
S
Ho
C)
c6
I
HL
CTr
I
I
Li Li
I
To determine if pav3C53 and tum 32 a-20 block cytokinesis, we stained embryos with
antibodies directed against the nuclear and cell membranes and a DNA dye. We observed
binucleate cells at high frequency in the dorsal epidermis of tumrn32a -20 stage 12 embryos
(Fig. 10D-I). In this stage, the dorsal epidermis has just completed the last postblastoderm
division and is exiting from the cell cycle. Binucleate cells were not observed in the
dorsal epidermis of pav3CS3. We did, however, observe large DNA masses suggestive of
polyploidy (Fig. O1N). Cytokinesis defects have been described in pav mutants
previously (Adams et al., 1998). Our inability to identify binucleate cells could indicate a
defect prior to cytokinesis or an early block to cytokinesis in which multinucleate cells
subsequently fuse. It is possible that multinucleate cells are most abundant following the
initial cytokinesis defect. We were unable to examine the earliest postblastoderm division
affected by pav3CS3 with certainty. Although heterozygotes were positively marked with a
balancer carrying Ubx-lacZ, we found that transgene expression could not be detected
with confidence until stage 12.
We believe that tum 3 2a-20 embryos first exhibit a cytokinesis defect in the dorsal
epidermis in cycle 15. This assertion is based on the presence of multipolar spindles in
dorsal epidermal cells undergoing cycle 16 (Fig. 11D-F). In addition to multipolar
spindles, we observed abnormal central spindles in telophase cells (Fig. 11J-O). We
observed a telophase cell with no central spindle (Fig. 11J-L) and a single, poorly defined
central spindle (Fig. 11M-O). We did not detect these defects in 48 spindles examined in
control embryos. In a quantification of 31 and 48 spindles in tum 32a -20 and heterozygous
sibling embryos respectively, we found the two abnormal anaphase/telophase spindles
(Fig. 11J-O) occurring at a frequency of 6% in turn32a-20 , whereas 21% of spindles in
heterozygous sibling embryos showed central spindles in telophase. However, we
examined few late anaphase/telophase spindles. These results, however, are consistent
with defects in central spindle assembly preceding a block to cytokinesis in tum"32 a-2
Figure 10. tumrn ""2 2o and pav3 cs3 display defects consistent with disrupted cytokinesis.
Stage 12 embryos stained with anti-lamin, anti-phosphoTyrosine, and YOYO-1 DNA
dye. Dorsal epidermal cells were imaged. The first column (A, D, G, I, L), shows antilamin staining. The second column (B, E, H, K, M), shows anti-phosphoTyrosine
staining. The third column (C, F, I, L, N), shows merged channels with anti-lamin in
blue, anti-phosphoTyrosine in red and YOYO-1 in green. The first row (A-C), shows
dorsal epidermal cells in yw control embryos. Single nuclei (green) shown in (C) are
enveloped by nuclear lamin shown in (A) and phosphoTyrosine shown in (B). (D-I)
32
turn
-2 °
embryos have binucleate cells in the dorsal epidermis. Arrowhead in (F) points
to the junction between two closely appositioned nuclei where nuclear lamin is localized
in (D) but arrowhead in (E) points to the same junction and no phosphoTyrosine is
evident. However, phosphoTyrosine in (E), envelopes both nuclei, indicating that this is a
binucleate cell. Arrowheads in (G-I) point to a similar junction between two nuclei.
Arrow in (I) points to a large DNA mass that may have arisen from nuclear fusion
following a failure in cytokinesis. (J-N) pav3C 53 display large DNA masses, arrow in (N),
which may result from cytokinesis failure. We also observed small nuclei, arrowhead in
(L) suggestive of aneuploidy.
yell//ow white
,,,3
2a-20
aV3C53
Figure 11. tum
r
""
display spindle defects in the dorsal epidermis.
All images were taken in the dorsal epidermis of stage 11/12 embryos. anti-tubulin is in
red and YOYO-1 DNA dye is in green. (A-C) yw control metaphase spindle. (D-F)
tum3 2a -20
multipolar spindle in metaphase. (G-I) yw control telophase cell with a tightly-
bundled central spindle evident between separating DNA masses. (J-L) shows an unusual
spindle in tum 32a-20 . Distance between nuclei suggest that this cell may be in telophase cell
and has failed to assemble a central spindle. (M-O) A central spindle with poor
microtubule bundling in turn32a-20 . (A, D, G, J, M) anti-tubulin. (B, E, H, K) YOYO-1
DNA dye. (C, F, I, L) merge.
Tubulin
yellow white
yellow white
t'/ ?32a-20
1
DNA
Tubulin
DNA
DISCUSSION
The identificationof mutants that affect developmental regulationof the cell cycle
In our screen for regulators of cell cycle specificity during development, we
identified 30 mutants falling into five phenotypic classes. We were particularly interested
in identifying factors that distinguish mitotic cycles from endocycles in this
developmental context.
Transcriptional regulation is clearly a motif in developmental cell cycle
specification. The Notch signaling pathway is known to impinge on the transcription of
key regulators during the mitotic to endocycle switch of follicle cells during Drosophila
oogenesis. string and dacapo are downregulated while fizzy-related is upregulated (Deng
et al., 2001; Edgar et al., 1994; Jones et al., 2000; Meyer et al., 2002; Schaeffer et al.,
2004). The E2F transcription factor is known to regulate cyclinE transcription during
embryogenesis but regulation is tissue-specific and other transcription factors are likely
to play variable roles in different tissues undergoing different types of cell cycle
(Royzman et al., 1997; Sauer et al., 1995). Class III and IV mutants displayed reduced
GI/S transcripts in the CNS, endodomains or both and may prove useful in identifying
additional transcription factors responsive to developmental signaling.
Class I and III mutants failed to downregulate PCNA expression in the
endodomains. This defect is reminiscent of mutants in cyclinE and dup, which display an
early S-phase block (Knoblich et al., 1994; Whittaker et al., 2000). We excluded cyclinE
and dup as candidates because these genes are located on the
2 nd
chromosome whereas
Class I and III mutants (with the exception of 3C241) map to the 3 rd chromosome.
Although Cdk2 maps to the third chromosome, hemizygous mutants in Cdk2 are larval
lethal and display no embryonic defects, suggesting that Class I and III mutants are not
likely mutants in Cdk2 (Lane et al. 2000). Class I and III mutants may affect replication
factors based on their phenotypic similarity to mutants in cyclinE and dup. These mutants
may be useful for examining G1/S transcript downregulation. cyclinE and dup mutants
fail to downregulate G1/S transcripts in the endodomains, suggesting that S-phase
progression is required for G1/S transcript downregulation. CyclinE and DUP are both
critical for replication initiation - an immediate early block to S-phase - leaving the
question as to the timing of transcript downregulation with respect to S-phase open. Class
I and III mutants may block S-phase at later points.
Unlike Class III mutants, Class I mutants specifically affect the endodomains.
Developmentally regulated fzr and cyclinE expression are required for endocycles
(Knoblich et al., 1994; Sigrist and Lehner, 1997). In addition, transcription of mitotic
cyclins must be turned off (Weiss et al., 1998). Class I mutants may affect developmental
regulation of transcriptional events required for endocycles. Alternatively, the CNS may
display normal G1/S transcription due to the perdurance of maternal products in this
tissue.
New mutants that affect mitotic cell cycles in the nervous system
We believed that Class IV mutants that showed signs of polyploidy in normally
diploid tissues would either enumerate the ways in which the canonical cell cycle is
modified to yield endocycles or elucidate the developmental signals that maintain mitotic
cell cycles at a time during which endocycles initiate. Two mutants in this class, 3C53
and 32a-20 (formerly 32a), were identified to be alleles of pavarotti, a MKLP-1 family
member, and tumbleweed, a Rho family GTPase activating protein. 2k32, 1r8 and 3C157
(of which 1r8 and 3C157 are allelic), displayed isolated large nuclei in the nervous
system.
The large isolated nuclei seen in 2k32, Ir8 and 3C157 suggested two possibilities:
the mutants affect regulation of cell division specific to nervous system development; or
the mutants affect mitotic cell cycle regulators and the maternal contribution is sufficient
to drive cell division until late embryonic development when only the nervous system is
proliferating. Examination of Ir8 and 3C157 indicated that large nuclei occur in both
neuronal and non-neuronal cells. This observation indicates a lack of cell type specificity.
We observed phosphohistone H3 localization to poorly condensed DNA, suggesting that
Ir8 and 3C157 may affect chromatin dynamics. We favor the explanation that Ir8 and
3C157 disrupt mitotic functions and that any potential tissue and cell type specificity is
due to the perdurance of maternal products in conjunction with the developmental timing
of cell divisions in the nervous system (Salzberg et al., 1994).
tum 3 2a20
disrupts cytokinesis and shows central spindle defects
We observed multinucleate cells in tum
32a
-20 indicating
a block to cytokinesis (Fig.
10). Our observations are consistent with prior work indicating that tum is required for
cytokinesis (Zavortink et al., 2005). In addition, we detected central spindle defects in
tum
3 2 -2
a-20: turn
a 0
32
displayed weak central spindles (Fig. 11) and the frequency of central
spindles in tum32a-20 was reduced. Similar central spindle defects were observed for
tumrn"",
an allele that is truncated due to a nonsense mutation at Arg195 and lacking the
GAP catalytic domains (Fig. 10) (Zavortink et al., 2005). The phenotypic similarity
between tum32a-20 and tumDHr
5 suggests
compromised catalytic activity in turn32a- 20. I
addition, the location of the substitution mutations in the GAP catalytic domain of tum"32a
20
may portend loss of GAP catalytic activity. Tum catalytic activity appears to be
required for cytokinesis. Transgenic expression of Tum catalytically dead mutants
carrying small deletions known to abolish catalytic activity, Tum AVE and TurnmAYRL, failed
to rescue the central spindle defect of tum"DHS (Zavortink et al., 2005).
Unlike the two known loss-of-function alleles, tumrnDHS and turnm2 (Fig. 10),
tum 32a-20 is
a full-length protein and unlike the catalytically dead transgenic rescue
constructs, TurnmIE and TumAYRL (Fig. 10), we expect that tum 32 a-20 retains physiological
expression patterns. The fact that Tum32 -20 is a full-length protein is important because
Tum affects Pav protein stability (Zavortink et al., 2005). Tum 32~ 02 is more likely than
TumDH 15 and TumAr 2 to retain native structure. In addition, the deletions in TumDH 15 and
TumAR2 may remove as yet undefined regulatory domains or disrupt protein structure.
tum3 2a-2 0
is potentially useful for dissecting the different roles of Tum -
stabilization of Pav and catalytic activity. Zavortink and colleagues report that only the
Pav interaction domain and not GAP catalytic activity is required for Pav stabilization
(Zavortink et al., 2005). However, they stabilized Pav levels by overexpressing the
catalytically dead constructs, TurnmIE and TurnAYRL, and the effects on Pav stability may be
indirect.
MATERIALS AND METHODS
Genetic screen
2 nd
chromosome mutants were isolated by Irena Royzman and Allyson Whittaker
(Royzman et al., 1997). 3" chromosome mutants were provided by R. Lehmann (Moore
et al., 1998). The genetic screen was carried out as described previously (Royzman et al.,
1997).
Cytological analysis and microscopy
In situ hybridization was carried out with digoxigenin-labeled RNA probes
exactly as described previously (Royzman et al., 1997). Antibody stainings were
performed as described previously (Dej et al., 2004). YL1/2 and YOL1/34 (Sera Lab)
anti-tubulin antibodies were used. Anti-phosphohistone H3 (Upstate Biotechnology) was
used at 1:250. Anti-DmOLamin (Gruenbaum et al., 1988) was used at 1:200 and antiphosphoTyrosine (Zymed) was used at 1:100 with methanol fixation. DNA stains were
performed with DAPI at 10 [tg/mL or YOYO-1 at 1:2000 (Molecular Probes). AntiDm0Lamin was obtained from Paul Fischer (SUNY at Stony Brook) and all secondary
antibodies were from Jackson Immunoresearch.
Images of multinucleate cells and epidermal spindle defects were collected using
a Zeiss Axiovert 100 M Meta confocal microscope with LSM510 software. A Zeiss
Axiophot microscope with Plan-Neofluar 20x, Plan-NeoFluar 25x Inn Korr, or PlanNeoFluar 40x oil objectives were used to collect images with a SPOT RT CCD camera
and software.
Mapping and Sequencing
Chromosome II and III deficiency kits and pavB 20 0 (Adams et al., 1998) were from
the Bloomington Stock Center. We performed male recombinational mapping (Chen et
al., 1998) using EP2383, EP2423,EP2041, EP2407, EP2054 and EP993.
Sequencing of tum 32a-20 was performed as described previously (Dej et al., 2004).
DNA from 10 embryos was purified and exons were sequenced on both strands and
compared to the Drosophilagenome sequence Release 2.
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CHAPTER THREE
The characterization of new cyclinE mutants that increase replication
fork progression during gene amplification
Eugenia A. Park, David M.MacAlpine and Terry L. Orr-Weaver
Whitehead Institute and Dept. of Biology, Massachusetts Institute of Technology,
Cambridge, MA 02142
Eugenia A. Park performed all of the immunolocalization, microscopy, flow cytometry and
quantitative real-time PCR for this work. For Drosophila2L tiling array experiments, Eugenia
Park prepared probes and hybridized arrays and assisted in scanning arrays. David MacAlpine
scanned arrays and analyzed data.
SUMMARY
We have identified the first cyclinE mutants to display aberrant gene
amplification during Drosophilaoogenesis. cyclinEuz36/cyclinEPzs and
cyclinE'f6 /cyclinEP28 are the first mutants to display increased replication fork
progression in gene amplification, demonstrating an absence of CyclinE independent
Replication Fork Barriers (RFBs) during this process. The rate of oogenesis is normal in
cyclinEl'f6 /cyclinEP 8 ovaries, suggesting that increased replication fork progression is due
to increased replication fork speed. These results implicate CyclinE in the regulation of
replication fork speed. Double Parked (DUP) and the Mcm2-7 complex showed
localization to the double bar cytological feature, corresponding to bi-directional
replication forks, leaving open the possibility that CyclinE/Cdk2 regulation of DUP and
Mcm2-7 may affect replication fork progression.
INTRODUCTION
The regulation of replication fork progression is important for varied biological
processes such as DNA damage checkpoint responses, gene expression and replication
termination (Rothstein et al., 2000). Replication fork progression is regulated, in part, by
factors found at the replication fork. PCNA, the DNA polymerase processivity factor, is a
key component of the active replication fork. It is known to interact with two of three
polymerases that localize to the replication fork, DNApol8 and DNApols (Garg and
Burgers, 2005). Additionally, biochemical approaches to determining components at
stalled eukaryotic replication forks have identified DNApola-primase, Mcm2-7, Cdc45,
Mcml0, GINS, Mrcl and Tofl proteins among others at stalled replication forks
(Calzada et al., 2005; Pacek et al., 2006). DNA polct is involved in lagging strand
synthesis during replication elongation. The Mcm2-7 complex is the putative replicative
helicase that travels with replication forks (Aparicio et al., 1997; Labib et al., 2000).
Cdc45, Mcml 0 and GINS are three other replication factors (Aparicio et al., 1999;
Gambus et al., 2006; Merchant et al., 1997; Moyer et al., 2006; Seki et al., 2006;
Takayama et al., 2003; Tercero et al., 2000; Wohlschlegel et al., 2002). Mrcl and Tofl
act as sensors and effectors in replication checkpoint pathways. They stabilize stalled
forks during checkpoint responses and regulate replication fork progression during
normal DNA replication (Szyjka et al., 2005; Tourriere et al., 2005).
Replicative helicases generally promote replication fork progression. In E. coli, a
protein inhibitor of the replicative helicase reduces replication fork speed (Skarstad and
Wold, 1995). Replication fork progression at normal speeds through S. cerevisiae
telomeric and subtelomeric sequences requires the Rrm3p helicase (Ivessa et al., 2002).
In addition, replication fork pausing mechanisms often target the helicase. During
prokaryotic replication termination, the trans-acting factors Tus in E. coli and RTP in B.
subtilis bind to and inhibit replicative helicase (Bussiere and Bastia, 1999). Replication
fork progression may be regulated at the level of competing helicases. At the S.
cerevisiae rDNA locus, the DNA helicase Piflp maintains replication fork pausing while
Rrm3p promotes replication fork progression (Ivessa et al., 2000). Helicases are also
regulated by phosphorylation. The Mcm2-7, the putative eukaryotic replicative helicase,
is a multisubunit complex that is phosphorylated at multiple sites on Mcm2 by S-phase
kinases Cdc7 and Cdk2 and ATR kinases (Montagnoli et al., 2006). The Cdc7
phosphorylation sites negatively regulate chromatin association of Mcm2, and
phosphorylations at these sites are stimulated by Cdk2 phosphorylation, suggesting that
sequential phosphorylation by multiple kinases modulates Mcm2-7 activity (Montagnoli
et al., 2006). CyclinB/Cdkl phosphorylation of Mcm4 restricts chromatin association
(Hendrickson et al., 1996). CyclinA/Cdk2 phosphorylation has been shown to restrict
Mcm4,6,7 subcomplex helicase activity (Ishimi et al., 2000). In addition, p27Kipl, the
CyclinE/Cdk2 inhibitor, binds to Mcm7 and inhibits initiation of DNA replication
(Nallamshetty et al., 2005).
Replication fork progression is also regulated by factors that do not localize to
replication forks. Replication fork barriers (RFBs) have been identified in prokaryotes,
yeast and metazoans (Rothstein et al., 2000). RFBs in prokaryotes and yeast consist of
DNA sequence elements that bind trans-acting factors to impose a block to replication
fork progression. Replication terminator sequences in E. coli and B. subtilis, Ter, bind
Tus and RTP respectively. In prokaryotic replication termination, RFBs colocalize
replication forks with replication termination machinery such as topoisomerase, which is
important for the decatenation of circular chromosomes (Zechiedrich and Cozzarelli,
1995). In S. pombe, the Rtsl sequence element binds the Swil/Swi3 complex, Tofl and
Mrcl homologs, and acts as a polar RFB to ensure the directional replication of the mat]
locus (Lee et al., 2004). This directional replication induces strand-specific mating type
switching by inducing an imprint in the next round of replication (Dalgaard and Klar
2000, 2006). Swil/Swi3 also binds at the imprint site and pauses replication forks
(Dalgaard and Klar, 2000; Lee et al., 2004). Like Tofl and Mrcl, Swi 1 and Swi3 are
required for stabilization of stalled forks and activate replication checkpoint signaling
(Noguchi et al., 2004). Not all barriers to replication fork movement are protein/DNA
units. Homopurine/homopyrimidine sequences have been shown to pause replication
forks thereby slowing replication fork progression (Rao, 1994; Rao, 1996; Rao et al.,
1988).
The regulation of replication fork progression may be important for gene
expression. RFBs are found in the rDNA loci of S. cerevisiae, S. pombe, mouse, Xenopus
and humans (Rothstein et al., 2000). In S. cerevisiae, S. pombe, mouse and Xenopus,
polar intergenic RFBs block replication fork progression opposing transcription (Brewer
and Fangman, 1988; Brewer et al., 1992; Gerber et al., 1997; Hernandez et al., 1993;
Kobayashi et al., 1992; Linskens and Huberman, 1988; Lopez-estrano et al., 1998;
Lucchini and Sogo, 1994; Sanchez et al., 1998; Wiesendanger et al., 1994). In E. coli,
head-on transcription severely inhibits replication fork progression while co-directional
transcription does not, revealing the importance of coordinated transcription and
replication (Mirkin and Mirkin, 2005). The Tetrahymena rDNA locus shows a co-
directional bias in replication fork pausing, suggesting that even co-directional
transcription and replication may be refractory to replication fork progression
(MacAlpine et al., 1997). This may be due to nucleosomal rearrangements required for
transcription.
Follicle cell gene amplification during Drosophilaoogenesis is a useful system
for studying DNA replication. During gene amplification, replication occurs from defined
origins of replication utilizing conserved cell cycle machinery. The visualization of
replication factor localization and DNA synthesis by BrdU incorporation is easy to
perform. In addition, replication initiation events occur from well-defined, isolated
origins of replication, allowing for the examination of replication elongation by
cytological methods.
During gene amplification, multiple, tandem replication forks move outwards
from origins of replication at five genomic loci known as the DrosophilaAmplicons of
Follicle Cells - DAFC-7F,DAFC-30B, DAFC-34B, DAFC-62D and DAFC-66D (J. Kim
and T. Orr-Weaver, unpublished results), (Claycomb et al., 2004; Spradling, 1981). These
replication forks amplify genomic regions containing eggshell constituents to high copy
numbers (Claycomb et al., 2002; Delidakis and Kafatos, 1989; Spradling, 1981) and gene
amplification is required for eggshell formation. The study of female-sterile mutants
affecting gene amplification and eggshell formationindicate that cell cycle machinery is
conserved during gene amplification. Females mutant for mcm6, dup/Cdtl, orc2, and
chiffon, which shows homology to dbf4, a specificity factor for the S-phase kinase Cdc7,
produce eggs with thin eggshells due to reduced gene amplification (Landis et al., 1997;
Landis and Tower, 1999; Schwed et al., 2002; Whittaker et al., 2000). In addition,
localization studies show that replication factors Orcl, Orc2, Orc5, DUP/Cdtl, Cdc45,
the MCM complex and PCNA localize to amplicons (Asano and Wharton, 1999; Austin
et al., 1999; Claycomb et al., 2002; Loebel et al., 2000; Royzman et al., 1999; Whittaker
et al., 2000).
The examination of female-sterile mutants, have identified genes required for
replication initiation, amongst these are components of the Pre-Replication Complex and
a putative co-factor for an S-phase kinase, chiffon. In addition, the cis-acting elements
which regulate the location of replication initation events, by binding the ORC Complex,
have been identified (Austin et al., 1999). Although the cis-acting elements, replication
factors and regulatory kinases regulating replication initiation during gene amplification
are characterized, little is known about the factors regulating replication elongation
during gene amplification. Cis-acting elements regulating replication elongation have not
been defined but such elements may be important for coordinating replication and
transcription during gene amplification. No genetic studies unmasking replication factors
involved in elongation have been forthcoming, possibly due to pleiotrophic effects on
replication initiation.
We have characterized the first mutants known to increase replication fork
progression during gene amplification. In addition, these are the first mutants in cyclinE
28
to display gene amplification defects. cyclinElf36/cyc1inEPzs and cyclinElf36/cycinEP
follicle cells display increased replication fork progression, demonstrating the lack of
stringent RFBs and implicating CyclinE in the control of replication fork progression.
RESULTS
A new mutation in cyclinE that displayspreviously undescribeddefects in gene
amplification
cyclinE'116 is an EMS allele that was isolated from a screen for regulators of the
G 1/S transcriptional program (Royzman et al., 1999; Royzman et al., 1997; Whittaker et
al., 2000). This allele is lethal in trans to Df(3L)TE35DI that deletes cyclinE and
1 3 6 is
cyclinEPzS, a P-element allele. In trans to cyclinEP
z,' a P-element allele, cyclinEf
sterile. In trans to cyclinEP28, a hypomorphic EMS allele, cyclinE*36 is semi-sterile.
6 /cyclinEPz
cyclinEfP
8
and cyclinE' 36a/cyclinEP28 mutant follicle cells display cytological
defects consistent with increased replication fork progression. The double bar structure is
seen in follicle cells of BrdU-labeled stage 12 and 13 egg chambers and corresponds to
gene amplification at DAFC-66D (Calvi and Spradling, 2001; Claycomb et al., 2002).
There is strong evidence that the double bar structure (Fig. 1B) corresponds to tandem
replication forks moving bi-directionally away from a central origin of replication
(Claycomb et al., 2002). The gap distance between bars reflects the distance traveled by
replication forks after termination of replication initiation events in stage 11 but prior to
the start of BrdU-labeling (Claycomb et al., 2002). cyclinEPf36cyclinEPz8 and
13 6/cyclinEP28
cyclinE•
mutant follicle cells had double bars in stage 13 egg chambers that
appeared to have increased gap distances relative to a CantonS wildtype control (Fig.
lA). We confirmed that the double bars corresponded to gene amplification at DAFC66D in cyclinEf36 /cyclinEZ8 mutant follicle cells by performing BrdU/FISH co-labeling
using a DAFC-66D specific FISH probe (Fig. 1C).
Figure 1. Double bars corresponding to replication at DAFC-66D are distantly
P
spaced in cyclinEf'/cyclinE"
mutant follicle cells.
(A) cyclinEf36/cyclinEPZ8 follicle cells have BrdU-labeled double bars with larger gap
distances than those seen in CantonSfollicle cells. Stacked confocal images of BrdUlabeled double bars in stage 13 egg chambers were deconvolved (left column) and
rendered into 3D projections (right column). The CantonSfollicle cell double bar has a
gap distance of 300 nm while the cyclinElf 6/cyclinEPZ8 follicle cell double bar has a gap
distance of 700 nm. (B) The double bar corresponds to bi-directional replication
occurring at DAFC-66D. Multiple, tandem replication forks incorporate BrdU (red shade)
as they move outwards. The gap distance (bracketed line) is the distance between the
BrdU-labeled bars. (C) The double bars seen in cyclinEf3l6/cyclinEP 8 (red) correspond to
DAFC-66D as visualized by FISH using a DAFC-66D specific probe (green). The scale
bar is 2 Rm.
I
Iil
~----~c
~---21
~L---~311
~s~-L,
c~s~
To confirm that gap distances were increased in cyclinEl3 6/cyclinEPz 8 , we
deconvolved stacked confocal images and rendered 3D projections to accurately measure
the distance between bars (Fig. lA). We measured gap distances in 10 double bars each
for cyclinE13 6/cyclinEPz 8 and an OregonR wildtype control (Table 1). The average gap
distance for OregonR follicle cells was 0.38 t 0.06 Rm. For cyclinEJ'6/cyclinEPz
8 follicle
cells, the average gap distance was 0.54 _ 0.13 tm. OregonR follicle cells showed gap
distances ranging from 0.30 to 0.50 [m while cyclinEP36/cyclinEP 8 mutant follicle cells
showed gap distances ranging from 0.30 to 0.80 [tm. This phenotype is mosaic. Within
one egg chamber, we found one double bar with a gap distance of 0.30 and another with a
gap distance of 0.70. We conclude that the gap distances between bars is increased in
cyclinE'f36/cyclinEPz8 mutant follicle cells relative to OregonR follicle cells. This
phenotype is consistent with increased replication fork progression.
8 and cyclinE'13
6/cyclinEPz 8 have expanded amplified regions at
cyclinE'-*6/cyclinEPz
DAFC-66D, DAFC-30B and DAFC-34B
We expected that increased replication fork progression would coincide with an
expanded amplified region at DAFC-66D. We performed quantitative real-time PCR on
genomic DNA purified from cyclinEP36/cyclinEPZ8 stage 13 egg chambers and determined
fold amplification at DAFC-66D in 10 kb intervals (Fig. 2A). Our results showed that the
6/cyclinEPzS follicle cells extends 100 kb to
amplified region at DAFC-66D in cyclinEf3
either side of the amplified maximum (Fig. 2A), whereas in OregonR wildtype control
follicle cells, the amplified region extends 50 kb to either side of the amplified maximum
(Fig. 2A). For cyclinE'36 cyclinEP28 follicle cells, we observed that the amplified region at
Table 1. GAP distances in cyclinE''I6cyclinEra and CantonS follicle cell double bars.
Gap distances were measured for 10 BrdU-labeled double bars in CantonS and
cyclinElf36/cyclinEP8 follicle cells. Distances were measured from 3D projections of
deconvolved confocal images and are given in pm. The mean gap distance of ten double
bars is given with standard deviation as error.
Double bar
1
2
3
4
5
6
7
8
9
10
Mean
CantonS
0.3
0.5
0.4
0.4
0.4
0.4
0.4
0.3
0.3
0.4
0.38 t 0.06
8
cyclinElf36/cyclinEzP
0.8
0.6
0.6
0.3
0.7
0.6
0.3
0.5
0.3
0.7
0.54 ± 0.13
Figure 2. The DAFC-66D amplified region is increased two-fold in
cyclinEl6 I/cyclinEP'Z
and cyclinE'6/cyclinEP2 8 mutant follicle cells.
Quantitative real-time PCR was used to determine fold amplification at DAFC-66D. Fold
amplification (Y-axis) was determined at 10 kb intervals surrounding the amplified
maximum at 0 kb (X-axis). (A) Amplification profiles were determined for
cyclinE'P6 /cyclinEPZ8 (blue and yellow) and OregonR (pink) stage 13 egg chambers. Two
independent DNA preps (trial 1 and trial 2) were used for cyclinEl36/cyclinEPz8 (blue and
6 /cyclinEPz 8
yellow). (B) Partial amplification profiles were determined for cyclinE /P
(blue), cyclinE'f 36/cyclinEP2 8 (orange) and OregonR (pink) stage 13 egg chambers. In both
(A) and (B), the amplified region is doubled in cyclinEln 6 /cyclinEPz
8
and
cyclinE'f6 /cyclinEP28 stage 13 egg chambers relative to the wildtype control. Errors are
the standard deviation of the sample.
A
70
60
50
40
-.0
PC
Is
wo
10
0
-80
-100
D
-40
OregontR -- cyclinE, I 6/cyclinEPz
-C1
-60
-20
0
20
Distance (kb)
8 trialI
40
60
80
100
cyclinE/3 6/CyClinEPZ 8 trial2
rr\
3u
45
40
a 35
o 30
S25
20
4 15
10
5
0
'0
10
-i
OregoniR
20
-.-
30
cyclin
40
50
60O
Distance (kb)
6 /cyclnEPz 8
70
80
-- cyclinE 69/cyclinE
90
28
100
DAFC-66D extends 100 kb to the right of the amplified maximum (Fig. 2B). We also
observed that while the amplified maximum in cyclinE'l6/cyclinEPz8 and
P 2 8 follicle cells is comparable to OregonR follicle cells (25-30 fold
cyclinE'9 6/cyclinE
amplified), suggesting that replication initiation events are intact in these cyclinE
mutants. The slope of drop-off in flanking regions may be gentler in cyclinE'f36 /cyclinEPz 8
2 8 follicle cells (Fig. 2B), indicating that replication forks are
and cyclinE'~6/cyclinEP
farther apart and that the rate of replication initiation events is reduced in
cyclinE'f36/cyclinEPz8 and cyclinE'f36/cyclinEP28 follicle cells. To examine additional
amplicons, we performed genomic DNA microarray experiments using the Drosophila
2L tiling array with sequences from Chromosome 2L. In addition, this array carried a 250
kb genomic segment surrounding DAFC-66D on Chromosome 3. DAFC-30B and DAFC34B are located on Chromosome 2L. We hybridized the Drosophila2L tiling array
(MacAlpine et al., 2004) with probe made from flow-sorted 16C follicle cell nuclei. 2C
embryonic DNA was used to synthesize a reference probe. Array experiments were
performed in triplicate and dyes swapped in one experiment. Results were Loess
normalized and plotted as the log base 2 ratio of enrichment on the Y-axis and
chromosomal position of the array feature on the X-axis (Fig. 3A).
We examined array features corresponding to DAFC-30B, DAFC-34B and DAFC66D. Plots were smoothened by taking the moving average of 15 continguous features
and overlaid (Fig. 3B). Our results indicate that amplified regions are broader at DAFC6/cyclinEPz
30B and DAFC-34B in cyclinEfP
8 follicle
cells relative to OregonR (Fig. 3A
and B). We found that in OregonR follicle cells DAFC-30B was amplified over a 160 kb
region (Fig. 3B). In cyclinE' 6/cyclinEPz8 mutant follicle cells, DAFC-30B was amplified
Figure 3. Amplified regions at DAFC-30B and DAFC-34B are increased two-fold in
6 cyclinE"P
cyclinE'J3
and cyclinE'P6 IcyclinEP 28 mutant follicle cells.
Drosophila2L tiled arrays were hybridized with labeled follicle cell 16C DNA and
labeled embryonic 2C DNA as reference. (A) Scatter plot for the length of chromosome
2L. X-axis is chromosome position and Y-axis is the log 2 ratio of enrichment. Peaks
corresponding to gene amplification at DAFC-30B and DAFC-34B are broader in
cyclinEJP6/cyclinEP 8 follicle cells than in OregonR follicle cells. Peaks are significant to
P.0.01. (B) High resolution plots for OregonR (solid line) and cyclinEf36/cyclinEPz 8
(dashed line) at DAFC-30B, DAFC-34B and DAFC-66D, were smoothened by finding
the moving average of 15 contiguous data points.
100
R
~
·
r=
·N
h;l
O
01
·
8
9
·
·
17
Z
00
vr
vr
~ci
0j
U,
I$)
rcl
0
-e
.
U,
0
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9'E
O'W
'Z 0Z
5'1
0"1
e.*
0;
0
o~
c
:c
It
.
. .
I
. .
I
-
-
I', Il,.
•1
-
m
I
i
I
0
V
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OOp
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5 Julumq3)laU21
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*p1
over a 300 kb region (Fig. 3B). In OregonR, DAFC-34B was amplified over a 100 kb
region (Fig. 3B). In cyclinEf-6/cyclinEPzs, DAFC-34B was amplified over a 200 kb region
(Fig. 3B). The amplified regions for DAFC-30B and DAFC-34B were doubled in
cyclinEf36/cyclinEPz8 relative to the wildtype control. This is consistent with the doubling
in amplified region seen for DAFC-66D as assayed by quantitative real-time PCR (Fig.
2). The Drosophila2L tiling array included a 250 kb region surrounding DAFC-66D and
for OregonR, the amplified region was about 120 kb (Fig. 3B). We were not able to
detect the non-amplified endpoints for cyclinEf3 6/cyclinEP8 but the expanded amplified
region appears to be greater than 250 kb (Fig. 3B). The increased amplification levels
seen in regions flanking the maximum are consistent with an expanded amplified region
relative to OregonR (Fig. 3B).
Our microarray experiments indicated that in OregonR follicle cells, DAFC-30B
is amplified over a 160 kb region and that DAFC-66D is amplified over a 120 kb region
(Fig. 3B). This differs from the approximate 100 kb amplified regions for DAFC-30B and
DAFC-66D detected by quantitative real time PCR (Claycomb et al., 2004; Claycomb et
al., 2002). The expanded amplified regions that we observed for DAFC-30B and DAFC66D suggest that microarrays are more sensitive for detecting lower copy number DNA.
Alternatively, the smoothing algorithm used in the high resolution plots (Fig. 3B) may
affect the boundaries of the amplified regions. Maximum amplification levels at DAFC66D determined by the tiled microarray differed significantly from values obtained by
quantitative real-time PCR. As assayed by tiled microarray, DAFC-66D is amplified 8fold in both cyclinE'f6/cyclinEPz8 and OregonR follicle cells (Fig. 3B). whereas
quantitative real-time PCR indicates about 30-fold amplification in both of these
102
populations (Fig. 2B). These differing values may reflect the limitations of the tiled
microarray for quantifying high copy number DNA.
Given the results from quantitative real-time PCR and 2L tiling array experiments,
we conclude that amplified regions for DAFC-66D, DAFC-30B and DAFC-34B are
expanded in cyclinEn36/cyclinEPZ8 and cyclinE'36/cyclinEP28 mutant follicle cells. These
results are consistent with increased replication fork progression.
Increased replicationfork progression is not due to prior cell cycle defects.
Follicle cells undergo developmentally regulated cell cycles during egg chamber
development. During stages 1-6, follicle cells proliferate mitotically. In stages 7-9,
follicle cells undergo endocycles. In stage 10A, follicle cells exit endocycles and in stage
10B, follicle cells initiate synchronous gene amplification. Perturbations to these prior
cell cycle events could explain the increased replication fork progression that we
8
observed in cyclinEf3 6/IcyclinEPz
and cyclinE~f36 /cyclinEP28 mutant follicle cells.
Premature initiation of gene amplification would give replication forks more time to
travel.
To determine if follicle cell endocycles were perturbed in cyclinE36/IcyclinEPZ8
females, we performed flow cytometry on DAPI-stained follicle cell nuclei isolated from
whole ovaries. Normally, follicle cells undergo three rounds of DNA replication
generating a ploidy of 16C. A final endocycle generating 32C ploidy is presumed to
occur in a subset of cells, possibly the dorsal anterior cells (Lilly and Spradling, 1996).
Follicle cell nuclei isolated from whole ovaries and stained with DAPI generate a flow
cytometry profile with five peaks corresponding to ploidies of 2C, 4C, 8C, 16C and 32C
103
(Fig. 4A). cyclinEf36 /cyclinEPz 8 mutant follicle cells undergo three endocycles generating
4C, 8C and 16C peaks, but may fail to undergo a complete fourth endocycle to generate a
32C peak (Fig. 4A). Alternatively, a 32C peak may be obscured in
cyclinElf36/cyclinEPz8 follicle cells (Fig. 4A).
In OregonR, most follicle cells are 16C, as evidenced by the prominent 16C peak
relative to the 2C, 4C and 8C peaks (Fig. 4A). In cyclinE-f 6/cyclinEPz8 , a greater
proportion of follicle cells are 2C, 4C and 8C, as indicated by the greater size of these
peaks relative to the 16C peak (Fig. 4A). Also in cyclinEf36/cyclinEPz8 , a larger fraction of
follicle cells have intermediate ploidies indicative of a larger fraction of cells undergoing
S-phase (Fig. 4A). This is evident from the enlarged areas between 4C and 8C, 8C and
16C peaks and also between the 16C peak and a possibly obscured 32C peak (Fig. 4A).
We conclude that endocycles are delayed in cyclinElf 6/cyclinEPz
8
follicle cells, possibly
due to prolonged S-phases.
6/cyclinEPZ8 follicle cells replicated genomic DNA to the
To determine if cyclinEJn
same extent as OregonR follicle cells, we calculated DNA indices for
cyclinEf36/cyclinEPz8 and OregonR follicle cell nuclei analyzed by flow cytometry (Table
2). The DNA index is a ratio of the mean fluorescence of a population of cells stained
with a stoichiometrically binding DNA dye to that of a reference population stained in the
same tube. The DNA index normalizes for sample-to-sample differences in staining. We
determined DNA indices for 4C, 8C and 16C peaks (and also the 32C peak in OregonR).
6/cyclinEPz 8 follicle cell nuclei.
in reference to the 2C peaks for OregonR and cyclinEfP
The DNA indices calculated for cyclinEf36/cyclinEPz8 follicle cell nuclei were not
significantly different from those calculated for OregonR follicle cell nuclei (Table 2).
104
Figure 4. The expanded amplified region at DAFC-66D is not due earlier cell cycle
defects.
(A) Follicle cell endocyles are delayed and have prolonged S-phases in
cyclinE'f36/cyclinEPz8 females. Follicle cell nuclei were isolated from OregonR and
cyclinE'f36 /cyclinEPz 8 ovaries, stained with DAPI and their ploidies determined by flow
cytometry. Fluorescence intensity is given on X-axis on a logarithmic scale. Y-axis
indicates cell count. In OregonR, 2C, 4C, 8C, 16C and 32C ploidies are evident in five
peaks. Note that the majority of cells have a ploidy of 16C as evidenced by the
prominence of this peak. In cyclinEf36 /cyclinEPZ8 , a greater proportion of cells are 2C, 4C,
and 8C as evidenced by the prominence of these peaks relative to the 16C peak. The 32C
peak may be obscured by S-phase nuclei. S-phase cells (the population between peaks)
are enriched. (B) Confocal images running the depth of the nucleus were collected and
stacked to produce 2D images used to quantify BrdU-foci in StlOB. X-axis is the number
of BrdU foci/nucleus. (gen) indicates genomic replication. Y-axis is the percentage of
cells falling into the categories given on the X-axis. The distributions of follicle cells for
cyclinE'136/cyclinEPZ 8 (magenta) and CantonS (teal) are indistinguishable (p=0.46),
suggesting that synchronous gene amplification initiates properly in cyclinEP6/cyclinEPz 8
follicle cells. (C) Quantitative real-time PCR of stage 10B (blue), 11 (pink), and 13
(yellow) cyclinEf36/cyclinEPz 8 follicle cells in a region flanking the DAFC-66D maximum
indicates that replication forks have not progressed beyond 40 kb in stage 10B and 11 egg
chambers (arrowhead). Thus, replication forks travel 60 kb between stages 11 and 13 (*).
X-axis is distance from amplification maximum in kb. Y-axis is fold amplification.
105
JS
S
uo!qu;U!Idunm PIOa
Table 2. DNA indices for OregonR and cyclinE'f36cyclinEP' follicle cell nuclei.
DNA indices were tabulated from three independent experiments. Purified follicle cell
nuclei were DAPI-stained and analyzed by flow cytometry to determine ploidies (Fig.
2A). Mean absorbance/fluorescence was determined for 2C, 4C, 8C, 16C and 32C peaks.
The DNA index was calculated by finding the ratio of mean fluorescence of 4C, 8C, 16C
and 32C peaks to the mean absorbance/fluorescence of the 2C peak (4C/2C, 8C/2C,
16C/2C, 32C/2C). Each DNA index (4C/2C, 8C/2C, 16C/2C, 32C/2C) was determined in
three independent experiments. Mean DNA index and standard deviation as error from
three independent determinations of DNA index is given. The 32C/2C DNA index was
not determined (ND) for cyclinElf36 /cyclinEPz8 due to the absence of a clear 32C peak
(Fig. 3A).
107
4C/2C
OregonR
1.71 ± 0.03
cyclinEf3 6 /cyclinEPZ8
1.81 ± 0.02
8C/2C
16C/2C
2.87 ± 0.11
5.08 ± 0.13
2.96 ± 0.10
5.10 ±_0.12
32C/2C
8.67 ± 0.66
ND
We conclude that cyclinE"6/cyclinEPs8follicle cells replicate DNA fully during the
endocycles.
To determine if cyclinElf 36/cyclinEPz8 follicle cells exit endocycles with the
appropriate developmental timing, BrdU-labeled nuclei were quantified in stage 10A egg
chambers by which time endocycles normally are complete and follicle cell nuclei do not
display nuclear BrdU-labeling (Calvi et al., 1998). In cyclinElf36/cyclinEPz8 , 2.8% of 531
follicle cell nuclei in nine egg chambers displayed nuclear BrdU-labeling whereas in
OregonR, 2.7'% of 732 follicle cell nuclei in ten egg chambers displayed nuclear BrdUlabeling. In spite of endocycle delays, cyclinE'f3 6 /cyclinEPz8 follicle cells replicate their
DNA fully as indicated by DNA indices (Table 2) and terminate endocycles with the
appropriate developmental timing in stage 10A as indicated by our quantification of
nuclear BrdU-labeling.
To determine if synchronous gene amplification initiates at the appropriate
developmental time and not earlier (which might result in increased replication fork
progression), we quantified BrdU foci in stage 10B egg chambers in cyclinE'f3 6/cyclinEPz8
and OregonR follicle cells (Fig. 4B). We counted 394 cyclinEhf 36/cyclinEPz8 follicle cells
in 6 egg chambers and 510 OregonR follicle cells in 8 egg chambers.
36/cyclinEP•
8 follicle cells displayed 4.8 BrdU foci/nucleus on average while
cyclinE~f
OregonR follicle cells displayed 4.7 BrdU foci/nucleus (Fig. 4B). We applied the
Wilcoxon Rank Sum test to determine if the two distributions (Fig. 4B) were significantly
different, and determined a P-value of 0.426, indicating that cyclinE1f36/cyclinEPz 8 follicle
cells are not significantly different from OregonR follicle cells in the number of BrdU
109
foci/nucleus in stage 10B. We conclude that synchronous gene amplification initiates
8
with the appropriate developmental timing in cyclinE1P6/cyclinEPz
follicle cells.
Gene amplification at DAFC-66D initiates during follicle cell endocycles. Gene
amplification during endocycles results in about 4 fold amplification by stage 10A (Calvi
et al., 1998). Therefore, abnormal gene amplification occurring during endocycle Sphases in cyclinEf36/cyclinEPz 8 follicle cells (Fig. 4A) may account for increased
replication fork progression in cyclinEP36/cyclinEPz8 follicle cells. We determined DAFC66D amplification profiles for cyclinElf6 /cyclinEPz 8 follicle cells in stage 10B,11 and 13
egg chambers to measure replication fork progression after the onset of synchronous gene
amplification in stage 10B. We found that replication forks travel 40 to 60 kb between
stages 11 and 13 (Fig. 4C). Previous work indicates that in OregonR, replication forks
travel 10 to 20 kb between stages 11 and 13 (Claycomb et al., 2002). We conclude that
gene amplification occurring during endocycles is not responsible for increased
replication fork progression in cyclinEf3 6/cyclinEP" 8 follicle cells.
Increasedreplicationfork progression in cyclinE/f36cyclinEP8 follicle cells may reflect
increasedreplicationfork speed
Increased replication fork progression may result from developmental delays
during oogenesis. Normally, replication elongation occurs during stages 10B, 11, 12 and
13 (Claycomb et al., 2002). If these stages are prolonged, replication forks may travel
farther given more time. We assayed egg-laying rates to determine the time required for
oogenesis in cyclinEf36/cyclinEPZ8 females. Females generally have two ovaries each
containing 16 egg-producing units known as ovarioles. We dissected and counted
110
ovarioles in cyclinE'f3 6 /cyclinEPz 8 and heterozygous sibling ovaries. 798 ovarioles in 52
1 36
cyclinE/cyclinE]
/cyclinEPz
8
ovaries and 492 ovarioles in 32 heterozygous sibling ovaries were
counted. We determined that cyclinE'f36 /cyclinEPz8 females have 15.3 t 1.8 ovarioles per
ovary while heterozygous sibling females have 15.4 _ 2.1 ovarioles per ovary where the
errors are the standard deviations of the samples. We concluded that cyclinE1f36/cyclinEPz8
ovaries have the same number of ovarioles per ovary as heterozygous sibling ovaries. To
approximate the duration of oogenesis, we assayed the number of eggs produced in a 24hour period at maximum fecundity (Fig. 5A). Newly hatched females were collected over
a period of 24 hours, and 30 females were placed in an egg collection chamber with
OregonR males. Eggs were collected over a 24-hour period and counted for 12 days. The
rate of egg production reached a maximum around day 7 for both cyclinE'f36/cyclinEPZ 8
and heterozygous sibling control females (Fig. 5A). Both cyclinEf3 6/cyclinEP8 and
heterozygous sibling females produced about 20 eggs in a 24-hour period at maximum
fecundity (Fig. 5A). Since each female has about 30 ovarioles (15 ovarioles per ovary),
this measure indicated that an ovariole produces an egg in approximately 36 hours in both
cyclinE'f36/cyclinEPz 8 and control females. This measure is intermediate to the 12-hour
duration calculated using a similar method and the minimal 63.5 hours calculated by Lin
and Spradling (Lin and Spradling, 1993; Spradling, 1993). Based on the results that (1)
cyclinE'f36/cyclinEPz8 and control females are comparable in terms of the number of
ovarioles per ovary and (2) that egg-laying rates are comparable over an extended period
(Fig. 5A), we concluded that the time required for egg chamber maturation is not
dramatically different in cyclinEf3 6/cyclinEPr8 females relative to a sibling control.
111
Figure 5. Increased replication fork progression is not due to developmental delays
in oogenesis.
(A) Egg-laying rates were determined by quantifying the number of eggs laid by 30
females in a 24-hour period. Per female egg-laying rate is on the Y-axis as a function of
fly age in days on X-axis. Egg-laying rates for cyclinEf36/cyclinEPz8 females (blue) and
sibling females (pink) peak at day 6 at about 20 eggs/day. (B) Egg chamber stages were
quantified between at day 7. Apparently apoptotic egg chambers with pyknotic nuclei, are
given as "A" on X-axis. Frequency of occurrence is on Y-axis. 673 cyclinEuf3 6/cyclinEPZ8
egg chambers (teal) and 494 sibling egg chambers (magenta) were counted with the
standard deviation in egg chamber frequency in 4 ovaries as error.
112
A 25
20
F
,10
5
0
d4
d5
d6
d7
d8
d9 dl0
Day of egg collection
-.- cyclinEIAP6/cyciAnEFZ8
B
dll
d12
-&-sibling control
0.25
0.2
0.15
S0.10.05
0
'2/3 4
5
6
7
8
9 10A 10B 11
12
-0.05
Egg chamber stage
z
Scycycc6/ CYC n£
8
Msibling control
13
14
A
To assess whether there are stage-specific delays in egg chamber development,
we quantified egg chambers in cyclinEf36/cyclinEPz8 and heterozygous sibling ovaries
(Fig. 5B) at maximum fecundity (d7 in Fig. 5A). We counted 673 egg chambers in 4
cyclinE'f6 /cyclinEPz8 ovaries and 494 egg chambers in 4 sibling control ovaries. We did
not observe dramatic differences in the frequency of stage 2 through stage 5 egg
chambers between cyclinEa36/cyclinEPz8 and sibling control females (Fig. 5B). During
these stages, follicle cells are undergoing mitotic divisions. In addition, we did not
observe dramatic differences in the frequencies of stage 7 and 8 egg chambers during
which follicle cells are undergoing endocycles (Fig. 5B). We did see an appreciable
difference in the frequency of stage 9 egg chambers during which follicle cells are
undergoing their final endocycles (Fig. 5B). Stage 9 egg chambers occurred at a
frequency of 6% in cyclinEPf36cyclinEPz8and 2% in sibling control ovaries. Interestingly,
enrichment of stage 9 egg chambers in cyclinEf36 /cyclinEPz ovaries corresponded to
increased S-phase length in the terminal endocycles as detected by ploidy analysis of
follicle cells (Fig. 4A) and may reflect a developmental delay coupled to a cell cycle
defect. The frequencies of gene amplification stages (10B, 11, 12 and 13) were similar in
cyclinE16/IcyclinEPz8 and control ovaries (Fig. 5B). We saw a slight elevation in the
frequency of stage 10B egg chambers in cyclinEf3 6/cyclinEPz8 ovaries, suggestive of
increased time spent in this stage. We do not think that increased replication fork
progression in cyclinEf36/cyclinEPz8
follicle cells is due to increased duration of stage
10B, since we have shown that replication forks progress about 60 kb between stages 11
and 13 (Fig. 4C). We conclude that the developmental timing of oogenesis is not altered
8 females. Our results suggest
significantly in cyclinEf36/cyclinEýP
that increased
114
replication fork progression in cyclinE'f3 6/cyclinEP 8 is not due to developmental delays in
oogenesis.
Polyteny is intact in cyclinElf 36 /cyclinEPz8 follicle cells
Our results suggest that increased replication fork progression in
cyclinE'f3 6/cyclinEPZ8 follicle cells is due to increased replication fork speed and not due to
developmental delays that prolong the period of gene amplification. Polyteny affects
replication fork speed (Steinemann, 1981). As such, we thought that increased replication
fork progression in cyclinE'f3 6/cyclinEPr8 and cyclinE'f36/cyclinEP28 follicle cells might be
due to defects in polytene chromosome structure. To determine if polyteny is maintained
in cyclinE1f36/cyclinEP
8
follicle cells, we performed BrdU/FISH co-labeling experiments
using FISH probes directed against DAFC-66D (Fig. 6A-D). We found that polytene
chromosome structure is largely intact in cyclinE1f
36/cyclinEPz8
mutant follicle. In general,
in cyclinEf 361/cyclinEPz 8 follicle cells, we saw one DAFC-66D FISH focus per cell that
looked qualitatively similar to DAFC-66Dfoci in the sibling control (Fig. 6A and C).
This observation suggested that polyteny is maintained for the 3 rd chromosome in
cyclinE'f36/cyclinEPz8 follicle cells.
We quantified BrdU and DAFC-66D FISH foci in 424 sibling control and 290
cyclinE'f36/cyclinEP=8 follicle cells to determine if the frequency of follicle cells displaying
multiple DAFC-66D FISH foci or no FISH foci, both consistent with loss of polyteny, is
increased in cyclinE'f36/cyclinEPz 8 follicle cells (Fig. 6E-F). In all cases, we found that
DAFC-66D FISH foci were coincident with BrdU-labeled foci. In all cases with one
DAFC-66D FISH focus/nucleus, the DAFC-66D FISH focus was coincident with the
115
Figure 6. Polytene chromosome structure is intact in cyclinE'1 6/cyclinEýa follicle
cells.
BrdU/FISH co-labeling was performed on cyclinElf 36/cyclinEPz 8 and sibling control
ovaries using a DAFC-66D specific FISH probe. (A-D) Projections of stacked confocal
images of follicle cells in stage 12 egg chambers. (A, C) DAFC-66D FISH. (B, D)
DAFC-66D FISH in green and BrdU in red. (E and F) Stacked confocal images were
used to quantify the number of BrdU-labeled foci/nucleus (X-axis) with number of nuclei
falling into each category on the Y-axis. The number of nuclei showing one, two and no
FISH foci in each category on the X-axis are shown in red, blue and yellow, respectively.
116
Q
ci 40,
o0
am
W
00
r0o
z"..
,•.
--
8
C
'-O
q
.tiamN .-iaqmnN
0
oý
00
0
t-
0
0D
W
0
a
p!alJN .jIaq•lnN
0
-D
emN
0
55-
(
largest BrdU-*labeled focus in the nucleus (Fig. 6A-D). In cyclinE'f3 6/cyclinEPz 8 follicle
cells, 2.9% (8/290) of follicle cells in stage 12 egg chambers showed two DAFC-66D
FISH foci and 0.3% (1/290) showed no DAFC-66D FISH foci (Fig. 6F). In sibling
control follicle cells, 1.2% (5/424) of follicle cells in stage 12 egg chambers showed two
DAFC-66D foci and 0.2% (1/424) showed no DAFC-66D FISH foci (Fig. 6E).
Although the occurrence of two FISH foci/nucleus is higher in
cyclinElf3 6 /cyclinEPz
8
follicle cells (2.9%) than sibling control follicle cells (1.2%), we
believe that polyteny is intact in cyclinE'f3 6/cyclinEPz8 follicle cells. 6 of the 8 nuclei
displaying two BrdU FISH foci/nucleus were found in one of six cyclinEf3 6/cyclinEPz 8
egg chambers examined. We cannot, however, exclude the possibility that increased
replication fork progression is due to aberrant polytene chromosome structure.
cyclinE'ý 6 displays a dominant defect in replicationfork progression
We sequenced the coding region of cyclinE'136 and identified a G to E substitution
mutation (Fig. 7A). This amino acid substitution is not present in an isogenic line. The G
to E substitution occurs in the 5 th residue N-terminal to the MRAIL hydrophobic patch,
suggesting that the mutation may affect CyclinE/Cdk2 substrate recognition (Furstenthal
et al., 2001; Schulman et al., 1998).
We wished to determine whether increased replication fork progression was a
gain-of-function or loss-of-function phenotype. We performed MPM2 antibody staining
on cyclinElf3 6/cyclinEPz 8 mutant follicle cells. Localization of MPM2 to subnuclear foci is
known to reflect CyclinE activity (Calvi et al., 1998). We were unable to detect any
differences in intensities of subnuclear foci between cyclinEf36 /cyclinEPz8 and OregonR
118
Figure 7. cyclinE' l
6
displays a dominant phenotype.
(A) cyclinE'f36 is a G to E substitution mutant in a non-conserved residue 5 residues
removed from the MRAIL substrate recognition motif. (B) Quantitative real-time PCR at
DAFC-66D was performed on stage 13 egg chamber DNA as described previously. Xaxis is distance from amplification maximum in kb. Y-axis is fold amplification.
Df(2L)TE35D1/+ (brown) did not show a pronounced broadening of the amplified region
suggesting that increased replication fork progression is not a loss-of-function phenotype.
This was true of cyclinE 2 8/+ (pink), cyclinEPZB/+ (blue). cyclinEf36/+ (yellow) showed a
slight broadening of the amplified region relative to other cyclinE mutant heterozygotes
and OregonR (black), indicating that cyclinE*36 is a dominant mutant.
119
A
.*•9*
*
*
.*
*.***
*
•******9*
C.elegans
D. rerio
M- musculus
THVMERHPHL QPKMRAILLD WLIEVCEVYK
TRVMERHPNL QPKMRAILLD WLMEVCEVYK
D. melanogaster
ISMLEQHPGL QPRMRAILLD WLIEVCEVYK
EHFLQRHPLL QARMRAVLLD WLMEVCEVYK
Gly to Glu
B
50
45
40
a 35
30
20
;
15
10
5
0
'10
20
30
- cyclin£E28/CyO
-+"OregonR
40
50
60
Distance (kb)
cyclin£AEf 6/CyO
-- cyclinEPz 8/CyO
70
80
90
100
-- DJ2L)7£35DI/CyO
controls, suggesting that any changes in CyclinE activity are beyond the limit of detection
by this assay (data not shown).
To determine if increased replication fork progression is recessive or dominant,
we performed quantitative real time PCR on stage 13 egg chamber DNA to establish
amplification profiles for cyclinE'f36/+, cyclinEPz8 /+ and cyclinEP28 /+egg chambers (Fig.
7B). To determine the loss-of-function phenotype, we examined Df(2L)TE35D1/+, which
lacks one copy of cyclinE. Df(2L)TE35D1/+ stage 13 egg chambers displayed a small
increase in fold amplification relative to OregonR stage 13 egg chambers at 20 kb, 30 kb,
40 kb and 50 kb from the amplification maximum (Fig. 7B). However, the differences
were small (about 5 units fold amplification at any point). cyclinEP8s/+ displayed an
amplification profile very similar to Df(2L)TE35D1/+ and cyclinEP28/+ follicle cells
displayed an amplification profile very similar to OregonR follicle cells (Fig. 7B).
cyclinE'f36/+ follicle cells, however, displayed considerable differences in fold
amplification relative to OregonR at 20 kb, 30 kb and 40 kb (about 15 units fold
amplification difference at each point - Fig. 7B). At 50 kb, the difference dropped down
to about 7 units fold amplification and at 60 kb to about 5 units fold amplification. In
cyclinE'f3 6/+, the slope of the drop-off in amplification levels has a noticeable shoulder
near the amplification maximum (Fig. 7B), indicating that the last replication forks
initiated from DAFC-66D travel farther in cyclinE'f36/+ follicle cells relative to all of the
other heterozygous mutants described and the OregonR control. At 25-fold amplification
levels, OregonR and cyclinEP28/+ replication forks have traveled about 10 kb.
Df(2L)TE35DI and cyclinEP8 /+ replication forks have traveled about 15 kb. cyclinE1f36/+
have traveled about 30 kb (Fig. 7B). Thus, the phenotype of increased replication fork
121
progression is dominant in cyclinE'36 /+. We cannot distinguish whether this dominant
effect is a dominant loss-of-function or a dominant gain-of-function phenotype.
Double Parkedand the MCM complex localize as double bars in cyclinEP6/IcyclinEPZ 8
follicle cells
The MCM complex localizes to replication forks and is the presumptive
replicative helicase in eukaryotes (Aparicio et al., 1997; Calzada et al., 2005; Gambus et
al., 2006; Ishimi, 1997; Labib et al., 2000; Moyer et al., 2006). During gene
amplification, MCM subunits and Double Parked (DUP), the DrosophilaCdtl homolog,
localize to replication forks (Claycomb et al., 2002). DUP is a known target of
CyclinE/Cdk2 (Thomer et al., 2004) and DUP is required for MCM complex localization
to replication forks (Claycomb). DUP and MCM chromatin localization is regulated by
Cyclin/Cdk phosphorylation (Hendrickson, Thomer, Masai?) (Masai et al., 2000;
Montagnoli et al., 2006). To determine if increased replication fork progression in
cyclinEf36/cyclinEPZ8 follicle cells is due to altered chromatin localization of DUP and
MCM complex, we performed in situ localization studies to examine DUP and MCM
complex localization to replication forks.
We observed localization of DUP and MCM subunits to double bars at reduced
levels (Fig. 8). In cyclinE36/IcyclinEPz 8 follicle cells, we saw MCM localized, at reduced
levels, to double bars with increased gap distances, relative to an OregonR wildtype
control (Fig. 8K-N). MCM double bars with increased gap distances in
cyclinEjf36/cyclinEPf8 follicle cells are consistent with increased replication fork
progression. In addition, in cyclinElf36/cyclinEPZ 8 follicle cells, we saw DUP localized, at
122
Figure 8. Dup and MCM subunits localize to double bars in cyclinE' 6 /cyclinEra
follicle cells.
cyclinEJ6 I/cyclinEPz8 and OregonR ovaries were stained with anti-Dup (A-F) or antiMcm2-7 (G-N). (A-F) confocal imaging of Dup double bars showing single confocal
images and projections, in which single confocal images taken at different depths are
stacked on top of each other, for representative double bars. Single images and
projections are marked in figure. (G-N) show anti-Mcm2-7 in red and YOYO-1 in green
with single images and projections as marked. Arrows in (A-N) point to double bars.
Scale bar is 1 [tm
123
OregonR
8
36/cyc2nEPz
cyc1njEJcI
DUP
OregonR
MCM
P1,0jection
P z8
cychnE/jEf 6/cyc/n£E
reduced levels, to double bars. Although gap distances in these double bars appeared to
be slightly increased in cyclinEl3 6/cyclinEPz 8 follicle cells, gap distances were not
increased as dramatically as seen for MCM double bars in cyclinEft36/cyclinEPz8 follicle
cells. These observations may indicate that DUP becomes uncoupled from replication
forks and that this uncoupling correlates with increased replication fork progression in
cyclinE'f36/cyclinEP8 follicle cells. Alternatively, these MCM and DUP localization
phenomena may occur in different subsets of cells. We found MCM and DUP double
bars in a small fraction of follicle cells examined and these subsets might not overlap. In
addition, measurement of gap distances in BrdU-labeled double bars indicates a range of
3 6/cyclinEPz8 follicle cells (Table 1). Given
replication fork progression defects in cyclinE'f
the generally reduced localization of MCM and DUP to double bars in
cyclinE'fS6/cyclinEPz8 follicle cells, we conclude that increased replication fork
progression is not due to increased localization of DUP and the MCM complex to
replication forks. In addition, we conclude that MCM complex localization to double bars
with increased gap distances is consistent with increased replication fork progression.
125
DISCUSSION
We have characterized the first mutants known to affect replication fork
progression during gene amplification. cyclinEl3 6/cyclinEPz8 and cyclinE'~36/cyclinEP28
follicle cells displayed increased replication fork progression as evidenced by increased
gap distances in BrdU-labeled double bars and expanded amplified regions for DAFC30B, DAFC-34B, and DAFC-66D as assayed by quantitative real-time PCR and
comparative genomic hybridization (CGH) of the Drosophila2L tiling array. These
mutants result in a dramatic two-fold increase in replication fork progression at DAFC30B, DAFC-34B and DAFC-66D.
Our results demonstrate that gene amplification is not regulated by Replication
Fork Barriers (RFBs). We cannot exclude the possibility that CyclinE is essential to sitespecific RFB function during gene amplification. CyclinE may regulate RFB function by
regulating trans-acting factors or by modulating replication fork sensitivity to sitespecific RFBs by regulating replication fork components.
Mutations in replication factors that compromise replication initiation during gene
amplification result in thin eggshells (Landis et al., 1997; Landis and Tower, 1999;
Schwed et al., 2002; Whittaker et al., 2000). Replication fork progression has not been
examined in any mutants affecting gene amplification replication initiation and it is
questionable whether examining replication elongation in the context of impaired
replication initiation is informative or even possible. The cyclinE' 6 allele is unusual in
that it is highly specific to replication elongation. In cyclinElf6/cyclinEPz 8 and
cyclinE1 f36/cyclinEP28 follicle cells, DAFC-66D is amplified (at maximum) to levels
126
comparable to wildtype (Fig. 2B) and as we have shown, the cell cycle events leading up
to gene amplification are minimally perturbed in cyclinEfl 6/cyclinEPz8 follicle cells.
The functional significance of actively restricting replication fork progression
during gene amplification is yet unclear. We examined the eggshells of eggs laid by
cyclinE'f•6 cyclinEP'z females by Scanning Electron Microscopy. Follicle cell footprints,
dorsal appendages, micropyle and operculum looked completely normal, indicating that
increased replication fork progression does not disrupt eggshell formation (Fig. 9).
Although we were unable to detect eggshell defects, cyclinEP6/cyclinE•
'8
and
cyclinEf36/cyclinEP28 females were sterile and semi-sterile respectively, suggesting that
limiting replication fork progression during follicle cell gene amplification may be
important for egg viability. Although we cannot exclude the possibility that reduced
2 8 females is due to
viability of eggs laid by cyclinEf3l/cyclinEPz8 and cyclinE~6/cyclinE"
effects in other tissues, an intriguing possibility is that misregulated replication fork
progression disrupts the expression of genes important for egg viability. In wildtype, at
DAFC-66D, replication forks travel 20 kb at most between stages 11 and 13 (Claycomb
et al., 2002). In cyclinE3'6/cyclinEPz 8 , replication forks at DAFC-66D travel 60 kb
between stages 11 and 13 (Fig. 4C). These active replication forks may impede gene
expression during late stages.
An expedient explanation for increased replication fork progression is that
developmental delays increase the time spent in gene amplification. The prevailing view
regarding replication fork progression during gene amplification is that bi-directional
replication occurs until follicle cells slough off in stage 14 of egg chamber development
(Claycomb and Orr-Weaver, 2005; Spradling and Leys, 1988). The primary locus for
127
Figure 9. Embryos laid by cyclinE'/ 61cyclinE e females have normal looking
eggshells.
Embryos were collected from cyclinE3 6I/cyclinEPz8 and sibling control females and
visualized by scanning electron microscopy. 6 embryos were examined for each sample
and representative images are shown. Scale bar is 100 jim. Follicle cell footprints
(arrow), dorsal appendages (arrowhead) and micropyle (*) look normal in eggs laid by
6 /cyclinEPz 8 relative to those laid by a sibling females.
cyclinE**
128
regulation is thought to be replication initiation, which is under developmental control.
Following replication initiation events, slow-moving replication forks travel at about 50
to 100 bp/min (Claycomb et al., 2002; Spradling and Leys, 1988). In this model, the
elapsed time between replication initiation events of gene amplification (occurring
largely in stages 10B and 11 at DAFC-66D) and follicle cell removal in stage 14,
determines the span of gene amplification.
We did not detect any delays in oogenesis in cyclinE'16/cyclinEPz8 ovaries,
suggesting that increased replication fork progression in cyclinEJP6 /cyclinEPz8 follicle
cells is not due to a prolonged period between the onset of gene amplification and the
sloughing off of follicle cells in stage 14 egg chambers.
Increased replication fork progression might result from disrupted polytene
structure. We conclude that increased replication fork progression in cyclinElf36/cyclinEPz8
36
P 28 follicle cells is not due to disrupted polyteny. Replication fork
and cyclinE'1
/cyclinE
speed is, in part, dependent on higher order DNA structure and increased replication fork
speed in cyclinE3 6 . Polyteny affects replication fork speed. Measurements of replication
fork speed indicate that while replication forks move at ~2.6 kb/min in Drosophila
diploid cells, replication fork speed is an order of magnitude lower in Drosophila
polytene larval salivary glands at ~300 bp/min. This order of magnitude difference may
be due to protein/DNA complexes specific to polytene chromosomes. Prior to gene
amplification, follicle cells undergo endocycles resulting in polytene chromosomes,
which are maintained during gene amplification (Calvi and Spradling, 2001). The
polytene structure of follicle cell chromosomes could serve as a major determinant of
replication fork progression during gene amplification.
130
Increased replication fork progression is a dominant phenotype conferred by
cyclinE1 36 . This mutant allele carries a Gly to Glu substitution five amino acids removed
from the MRAIL hydrophobic patch. The MRAIL hydrophobic patch interacts with
cognate RXL sequences and is important for Cyclin/Cdk substrate specificity (Loog and
Morgan, 2005), inhibition of Cyclin/Cdk by CIP/KIP inhibitors (Wohlschlegel et al.,
2001) and chromatin localization of Clb5 in S. cerevisiaeand CyclinE in Xenopus via
interaction with RXL motifs in Orc6 and Cdc6, respectively (Furstenthal et al., 2001;
Wilmes et al., 2004). The proximity of the Gly to Glu substitution found in cyclinE f36 to
the MRAIL hydrophobic patch suggests that this mutation disrupts CyclinE/Cdk2 or
CyclinE binding to replication factors. We are unable to predict, however, whether the
Gly to Glu substitution increases or decreases CyclinE activity and we did not detect
appreciable changes in CyclinE protein levels by in situ antibody staining of
8 and cyclinE'6/cyclinEP
28 ovaries (data not shown).
cyclinEf~6/cyclinEPz
CyclinE may directly regulate replication fork progression by covalently
modifying replication fork components. In particular, we thought that the cyclinElf36
phenotype of increased replication fork progression might be due to altered
CyclinE/Cdk2 regulation of DUP/Cdtl and the Mcm2-7 complex. Both DUP/Cdtl and
Mcm2-7 are known to localize to double bars at DAFC-66D (Claycomb et al., 2002). In
cyclinE'f36 /cyclinEPz 8 follicle cells, we observed DUP/Cdtl and Mcm2-7 localizing to
double bars at reduced levels (Fig. 8).
Although reduced localization to double bars is counterintuitive to increased
replication fork progression, these proteins may be localized in excess and CyclinE/Cdk2
regulatory phosphorylation is complex. MCM subunits exist as multiple phosphoisoforms
131
that are cell cycle dependent (Young and Tye, 1997), Cyclin/Cdk phosphorylation is most
clearly established for Mcm2 and Mcm4. While hyper-phosphorylation by CyclinB/Cdkl
reduces chromatin affinity in both and Xenopus and human cells (Fujita et al., 1998;
Hendrickson et al., 1996), complete dephosphorylation abolishes chromatin binding in
Xenopus (Pereverzeva et al., 2000). Phosphorylation of Mcm4 by CyclinA/Cdk2 has been
shown to inhibit Mcm4,6,7 helicase activity in human cells (Ishimi et al., 2000). The
effects on Mcm2-7 localization may be downstream of DUP/Cdtl localization.
DUP/Cdtl has previously been shown to be required for Mcm2-7 localization during
gene amplification (Claycomb et al., 2002). Ectopic overexpression of a DUP
phosphoacceptor mutant dominantly inhibits replication elongation at DAFC-66D
(Thomer et al., 2004), suggesting that DUP phosphorylation by CyclinE/Cdk2 promotes
replication elongation. Thus, reduced CyclinE activity would be expected to reduce
replication fork progression, suggesting that CyclinEf '36 has increased activity. The
1
possibility remains that increased replication fork progression in cyclinEV
6
mutants is
due to altered CyclinE/Cdk2 regulation of DUP/Cdtl and MCM2-7 complex at the
replication fork.
Increased replication fork progression seen in cyclinEV36/cyclinEPz 8 and
28
follicle cells demonstrates plasticity in replication fork speed during
cyclinE1 0/cyclinEP
gene amplification. This plasticity may be functional. Previously, amplified regions for
DAFC-30B (Claycomb et al., 2004), DAFC-34B (J. Kim and T. Orr-Weaver, unpublished
results) and DAFC-66D (Claycomb et al., 2004; Claycomb et al., 2002; Spradling and
Leys, 1988) were determined to be ~100 kb in span with replication forks predicted to
travel -50 kb to either side of confined replication initiation sites. Our results by
132
comparative genomic hybridization of the Drosophila2L tiling array indicate greater
variability with DAFC-30B amplifying a 160 kb region, DAFC-34B amplifying a 100 kb
region and DAFC-66D amplifying a 120 kb region. Replication fork speed may be
regulated variably at the different amplicons perhaps to meet the varying demands of
transcription at these amplicons. In addition, replication fork speed may be regulated
variably in different subsets of cells in response to developmental cues.
133
MATERIALS AND METHODS
Quantitative Real-Time PCR
Quantitative realtime PCR was performed using primer sets spanning 50 kb on either side
of ACE3 (denoted as distance 0) at 10-kb intervals and primers to a nonamplified
intergenic region on chromosome arm 3R (located approximately 25 kb upstream of the
DNApola locus) as described previously (Claycomb et al., 2002). Primer sets spanning
60 - 100 kb on either side of ACE3 were generated as described previously (Claycomb et
al., 2002) and supplied by GeneLink. In Fig. 2, three ten-fold dilutions of stage 1-8 ovary
DNA were used as standard. Relative fluorescence was measured for each sample in
relation to the standard curves and standard deviations of triplicate reactions were
calculated by the ABI Prism 7000 software. Fold amplification was calculated by
dividing the relative fluorescence of the amplicon site by the relative fluorescence for the
nonamplified intergenic region near DNApola. Error is the standard deviation of the ratio
A/C = (FA/FC)*{[(SA/FA)A2 + (SC/FC)A2]}. A, amplicon locus; C, control locus; FA,
relative fluorescence for amplicon locus, FC, relative fluorescence of control locus; SA,
standard deviation for amplicon locus; SC, standard deviation for control locus.
Flow cytometry and DNA index
Follicle cell nuclei were isolated as described previously and stained with DAPI (Lilly
and Spradling, 1996). Flow-sorting and ploidy analyses were performed on a MoFlo flow
cytometer (Dako, Fort Collins, CO) at the M.I.T. Flow Cytometry Core Facility. An
Argon Ion laser was tuned to multi-line UV with a 450/22 bp filter. 16C nuclei were
collected for DNA microarray experiments. For the calculation of DNA indices, ploidy
analyses were performed in three separate experiments. 2C, 4C, 8C, 16C, and 32C peaks
134
were gated and the mean of total fluorescence intensity calculated. DNA indices were
found by dividing the mean of total fluorescence for the 4C, 8C, 16C, or 32C peaks by
the mean of total fluorescence for the 2C peak within the same sample. Error is standard
deviation.
Drosophila2L tiling array
Drosophila2L tiling arrays were kindly provided by David MacAlpine and Steven Bell
(M.I.T., Cambridge, MA). DNA from 16C follicle cells and embryonic DNA for
reference was purified as previously described (Claycomb et al., 2004). DNA was
labeled, slides hybridized and data analyzed as previously described (MacAlpine et al.,
2004). To calculate p-values, total data points were used to model the normal distribution.
Experiments were performed in triplicate with one dye-swap.
Sequencing
Sequencing was performed as described previously using genomic DNA from 8 embryos
for each sequencing run (Dej et al., 2004), cyclinE coding regions were sequenced with
two-fold coverage and the mutation was sequenced with four-fold coverage. An isogenic
strain was sequenced in parallel and the Gly to Glu substitution was not present in this
strain.
Cytology
Anti-DUP and anti-MCM2-7 antibody staining was performed on whole ovaries as
previously described (Claycomb et al., 2002). YOYO-1 (Molecular Probes) was used to
135
stain DNA at 1:2000. BrdU/FISH co-labeling was performed as previously described
(Claycomb et al., 2004). Imaging was performed using a Zeiss Axiovert 100 M Meta
confocal microscope with LSM510 software. Excitation of YOYO-1 and rhodamine dyes
used the 488 and 543 nm lasers respectively. Deconvolution was carried out using
Huygens2.3-professional (Scientific Volume Imaging). Rendering and analysis of threedimensional data was carried out using the MeasurementPro module of Imaris3 Surpass
3.2 (Bitplane).
Scanning Electron Microscopy
SEM was performed as previously described (Claycomb et al., 2004).
136
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141
CHAPTER FOUR
Conclusions and perspectives
142
CONCLUSIONS & PERSPECTIVES
We observed increased replication fork progression in cyclinE' 6 mutants, which
correlates with reduced DUP and Mcm2-7 co-localization to sites of replication initiation
(data not shown) and to replication forks. We favor the model that CyclinE/Cdk2
regulates DUP and Mcm2-7 at the replication fork to mediate fork progression. Changed
chromatin configuration and topological parameters are alternative explanations. In the
following chapter, we note the alternative models and delineate the reasons why we have
excluded them. To conclude the chapter, we discuss CyclinE/Cdk2 regulation of DUP
and Mcm2-7 and propose future experiments investigating a possible role for
CyclinE/Cdk2 in regulating replication fork progression via DUP.
Chromatin structure
FISH with DAFC-66D and DAFC-7F specific probes, and BrdU-labeled foci
indicated that follicle cell polytene chromosome structure, defined previously by these
methods (Calvi and Spradling, 2001), is maintained in cyclinEJP6/cyclinEPZ
8 follicle
cells.
However, altered chromatin structure may not manifest as loss of polyteny. Cohesins,
histone modifications, or other protein complexes defining chromatin structure may be
disrupted in cyclinEf36 mutants.
We performed anti-acetylhistone H4 (Lys5) (Upstate Biotechnology), staining on
6 /cyclinEPz
cyclinEfn3
8
28 ovaries. Anti-acetylhistone H4 (Lys 5)
and cyclinEf36/cyclinEP
localizes to DAFC-66D during gene amplification (G. Bosco, unpublished results).
Acetylated histone H4 increases gene amplification (Aggarwal and Calvi, 2004),
suggesting that this histone modification promotes replication, perhaps by promoting an
143
open chromatin configuration (Aggarwal and Calvi, 2004). Anti-acetylhistone H4 (Lys5)
6/cyclinEPzs and cyclinE'f3 6/cyclinE•2 8 follicle cells, similar to
localized to foci in cyclinE'13
a wildtype control, suggesting that replication fork progression defects are not due to
alterations in H4 acetylation (data not shown). We were not able to detect altered
chromatin structure by in situ antibody staining with anti-acetylhistone H4 (Lys5).
Additional antibodies specific for histone modifications are commercially
available and several have been tested for specific localization in follicle cells (G. Bosco,
unpublished results). Many of these produced pan-nuclear staining in the follicle cells.
More interesting localization patterns may be uncovered by washing the samples with
high salt buffer to remove non-chromatin bound fractions. This approach was taken to
unmask specific Mcm2-7 localization to follicle cell amplicons (Claycomb et al., 2002).
Cohesin localization to follicle cell chromosomes has not been characterized.
However, antibodies to Drosophilacohesin subunits Rad21 (Lee et al., 2005) and Smcl
(Dorsett et al., 2005; Thomas et al., 2005) exist and cytological studies may prove an
expedient route to examine cohesin localization in cyclinElf36/cyclinEP8 and
cyclinE'f 36/cyclinEP2 8 follicle cells.
Increaseddistance between replicationforks in cyclinE'36/cyclinEPz8 and
cyclinE'f36/cyclinE P28follicle cells
Increased replication fork progression may be due to topological factors, such as
the helical strain generated by multiple, tandem replication forks. In OregonR wildtype
follicle cells at DAFC-66D, replication forks are ~10 kb apart (Spradling and Leys,
1988). This inter-replication fork distance is calculated by determining the number of
144
replication initiation events (equal to the number of replication forks moving tandemly in
one direction) and dividing the interval amplified by these amplification forks by the
number of replication forks. For example, DAFC-66D is amplified ~30 fold by our
measure, consistent with the results of J. Claycomb and colleagues (Claycomb et al.,
2002). ~30 fold amplification corresponds to 5 replication initiation events and therefore
5 replication forks moving in one direction spread over a 50 kb interval, which indicates
replication fork spacing of -~10 kb in wildtype. Since the amplified region is doubled at
DAFC-66D in cyclinEf36/cyclinEPzs and cyclinEf'6 /cyclinEP28 follicle cells, and
amplification levels at the maximum are not diminished, this suggests that replication
forks are spaced ~20 kb apart. However, these values are approximate since replication
forks may not be distributed uniformly.
CyclinE/Cdk2 is required for replication initiation (Knoblich et al., 1994). As
such, we would like to note that a potential mechanism for increased replication fork
progression seen in cyclinEIf36 mutants would result from a defect in replication initiation.
Decreased replication initiation rates would allow replication forks to travel farther
between origin firings. The resulting increased distance between replication forks could
relieve superhelical strain allowing replication forks to travel faster. We do not think this
is a likely mechanism given that fold amplification is not decreased at DAFC-66D in
cyclinE"36 mutants. In addition, in wildtype, relatively small differences are seen in
replication fork progression among the follicle cell amplicons. Replication forks progress
~50 kb for DAFC-30B, DAFC-62D and DAFC-66D although these amplicons are
amplified 4-, 3.5- and 30-fold respectively (Claycomb et al., 2004; Claycomb et al.,
2002). If the topological strain generated by tandem replication forks affects replication
145
fork speed, we would expect a correlation between the number of replication initiation
events and replication fork progression.
Regulation of DUP and Mcm2-7 at replicationforks
Increased replication fork progression in cyclinEJP6/cyclinEP8 and
cyclinE'f36/cyclinEP28 follicle cells may be due to aberrant CyclinE/Cdk2 regulation of the
protein machinery at the replication fork. Both DUP and Mcm2-7 localize to double bars
(Claycomb et al., 2002). During gene amplification, Mcm2-7 localization to double bars
requires unimpaired DUP function, suggesting that DUP (which is required along with
Cdc6 for Mcm2-7 loading at the Pre-Replication Complex) is involved in maintaining
Mcm2-7 at slow-moving replication forks (Claycomb et al., 2002; Spradling and Leys,
1988).
Thomer et al. have described gene amplification defects for DUP mutants
incapable of undergoing CyclinE/Cdk2 phosphorylation (Thomer et al., 2004).
Overexpression of an N-terminal deletion (C-DUP) greatly increases BrdU-incorporation
at gene amplifying foci, whereas overexpression of an N-terminal phosphomutant lacking
N-terminal Cdk2 phosphorylation consensus sequences (DUP 10(A)) inhibits gene
amplification completely. Of particular interest, overexpression of DUP 10(A) inhibited
gene amplification at DAFC-66D during developmental stages when DAFC-66D is
undergoing replication elongation only, suggesting that CyclinE/Cdk2 is required for
replication fork progression. Amplification parameters (maximum amplification and
amplification profile) have not been determined under conditions of C-DUP
overexpression and it is unclear whether increased BrdU-incorporation at gene
146
amplifying foci is due to increased replication initiation or increased replication fork
progression. C-DUP lacks potential ubiquitylation sites regulating DUP degradation
(Nishitani et al., 2006; Senga et al., 2006). Thomer et al. have shown that CyclinE/Cdk2
activity is required for DUP degradation, and that the N-terminus is necessary and
sufficient for DUP degradation in follicle cells, suggesting that CyclinE/Cdk2
phosphorylation of DUP at the N-terminus promotes DUP degradation (Thomer et al.,
2004). DUP localizes to replication forks during gene amplification and failures to
degrade DUP may have consequences at replication forks. Although DUP is required for
Mcm2-7 localization to double bars (Claycomb et al., 2002), this observation may reflect
an earlier requirement for DUP in loading the MCM complex, perhaps during replication
initiation. The role of DUP, at replication forks during gene amplification, is not yet
clear. We observed reduced localization of DUP to double bars, indicating that DUP
localization to double bars is not limiting for replication fork progression.
We also observed reduced localization of Mcm2-7 to double bars in
cyclinEf36/cyclinEPz
8 follicle
cells. Given that helicases generally promote replication fork
progression, our observation suggests that Mcm2-7 localization is not limiting for
replication fork progression. In general, Cyclin/Cdk activity inhibits Mcm2-7 chromatin
localization (Findeisen et al., 1999; Fujita et al., 1998; Hendrickson et al., 1996) and
reduced localization of Mcm2-7 to double bars may indicate that cyclinEf3 6 is a gain-offunction mutant. We do not think this is the case. Normally, nuclear CyclinE levels are
high during gene amplification. Reduced Mcm2-7 localization at double bars in
cyclinE•36/cyclinEPz8 follicle cells does not reflect total Mcm2-7 chromatin localization,
which may more accurately reflect CyclinE/Cdk2 activity. Total chromatin localization
147
6 /cyclinEPz 8 follicle cells. We saw nuclear Mcm2-7
may, in fact, be increased in cyclinEf31
localization in a small fraction of cyclinEl 36
/cyclinEP
s follicle
cells (Fig. 1A). Normally,
Mcm2-7 localizes to foci corresponding to gene amplification at DAFC-66D (CantonSin
Fig. 1A). Increased ectopic replication seen in cyclinEf36/cyclinE•z8 and
3 6/cyclinEP2
8 follicle cells (Fig. 1B) is consistent with more widespread Mcm2-7
cyclinE"f
localization. Calvi et al. postulated that high nuclear CyclinE levels are important for
restricting genomic DNA replication during gene amplification (Calvi et al., 1998). If
CyclinE inhibits nuclear Mcm2-7 chromatin localization and replication fork progression
at follicle cell amplicons, our results are consistent with reduced CyclinE activity in
cyclinEI36 mutant follicle cells.
Future Experiments
Our results suggest that cyclinEf36 is a loss-of-function allele. We would like to
confirm that CyclinE/Cdk2 activity is reduced in cyclinE'f36 mutants. CyclinE/Cdk2
activity could be determined by performing in vitro kinase assays using H1 as a target.
We will immunoprecipitate kinase using anti-DUP which has been shown to pull down
functional Cdk2 (Thomer et al., 2004). Alternatively it may be necessary to
immunoprecipitate from lines expressing UAS:6Xmyc:CDK2 using anti-Myc, given that
CyclinE'f36 may interefere with DUP binding.
Overexpression of the N-terminal truncation of DUP (C-DUP) leads to increased
gene amplification. It has not been determined whether this effect is due to increased
replication initiation or elongation. We would be interested to know whether replication
fork progression is increased in these mutants, and if so, what the effect of C-DUP
148
Figure 1. cyclinE"/IcyclinEvafollicle cells display nuclear Mcm2-7 localization
consistent with more BrdU-labeled sites throughout nucleus.
PanelA: cyclinE'P6 /cyclinEPz8 follicle cells (D-F) displayed nuclear Mcm2-7 localization
as visualized by anti-Mcm2-7 staining (A, D) and DNA staining with YOYO-1 (B, E) in
stage 12 egg chambers. In a CantonS wildtype control (A-C), Mcm2-7 localizes to a
single major focus corresponding to replication at DAFC-66D. (C, F) merge with antiMcm2-7 in red and YOYO in green. Antibody staining was performed as in Chapter 3.
Panel B: Confocal images of BrdU-incorporation in stages 11 and 13 in CantonS,
2 8 follicle cells. Imaging was performed as
cyclinElf36/cyclinEPz8 and cyclinE'36/cyclinEP
described in Chapter 3. (C) cyclinE'f36/cyclinEPz 8 and (E) cyclinEf36/cyclinEP28 follicle
cells display punctate BrdU-labeled spots throughout the nucleus in stage 11 correlating
with more widespread Mcm2-7 localization in cyclinEf36/cyclinE"8 follicle cells. (A)
CantonS control with 1 BrdU focus/nucleus. (B, D, F), In stage 13 egg chambers,
28
CantonS, cyclinE'I36/cyclinEPZS and cyclinEf36/cyclinEP
follicle cells display BrdU-
labeled double bars corresponding to replication elongation at DAFC-66D.
149
Mcm2-7
merge
DNA
CantonS
cyc/inEI
P
6/cychnE
Z8
B
stage 11
CantonS
28
cycn/i '1 6/cycalnE'
stage 13
6/cyclinEPz 8 or cyclinE*
3 6/cyclinEP28 background. These
expression is in a cyclinEf3
experiments would begin to address whether CyclinE regulates replication forks through
DUP. C-DUP lacks potential ubiquitylation sites regulating DUP degradation (Arias and
Walter, 2006; Nishitani et al., 2006; Senga et al., 2006) and increased gene amplification
may be due to persistence of DUP in the cell. In addition, the N-terminus may carry
other, as yet unmapped, regulatory domains. A Cdk phosphomimetic DUP mutant would
be informative for determining how CyclinE regulates DUP because it would more
specifically address CyclinE regulation of DUP.
151
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