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Article
Dynamic loading and redistribution of the Mcm2-7
helicase complex through the cell cycle
Sara K Powell
†
, Heather K MacAlpine
†
, Joseph A Prinz, Yulong Li, Jason A Belsky & David M MacAlpine
*
Abstract
Eukaryotic replication origins are defined by the ORC-dependent
loading of the Mcm2-7helicase complex onto chromatin in G1.
Paradoxically, there is a vast excess of Mcm2-7relative to ORC
assembled onto chromatin in G1. These excess Mcm2-7complexes
exhibit little co-localization with ORC or replication foci and can
function as dormant origins. We dissected the mechanisms regu-
lating the assembly and distribution of the Mcm2-7complex in the
Drosophila genome. We found that in the absence of cyclin E/Cdk2
activity, there was a 10-fold decrease in chromatin-associated
Mcm2-7relative to the levels found at the G1/S transition. The
minimal amounts of Mcm2-7loaded in the absence of cyclin
E/Cdk2activity were strictly localized to ORC binding sites. In
contrast, cyclin E/Cdk2activity was required for maximal loading
of Mcm2-7and a dramatic genome-wide reorganization of the
distribution of Mcm2-7that is shaped by active transcription.
Thus, increasing cyclin E/Cdk2activity over the course of G1is not
only critical for Mcm2-7loading, but also for the distribution of
the Mcm2-7helicase prior to S-phase entry.
Keywords cell cycle; chromatin; DNA replication; Mcm2-7
Subject Categories Cell Cycle; Chromatin, Epigenetics, Genomics &
Functional Genomics; DNA Replication, Repair & Recombination
DOI 10.15252/embj.201488307 |Received 26 February 2014 | Revised 27
November 2014 | Accepted 2December 2014 | Published online 2January 2015
The EMBO Journal (2015)34:531–543
Introduction
The duplication of a eukaryotic genome within the confines of
S-phase is a remarkable event. First, the sheer scale of the process—
tens to thousands of million base pairs of DNA—needs to be precisely
copied once and only once within just a few hours. In addition, DNA
replication forks must initiate within and progress through diverse
local chromatin environments including accessible euchromatin
and repressive heterochromatin. And finally, the process has to be
dynamic and capable of responding to environmental and develop-
mental cues. Thus, with every cell cycle, thousands of DNA replica-
tion start sites (origins) must be selected and activated in a regulated
and coordinated manner to ensure that the entire genome is faithfully
duplicated (reviewed in Masai et al,2010).
Each potential origin of replication is marked by the origin recog-
nition complex (ORC) (Bell & Stillman, 1992; Rao & Stillman, 1995).
In G1, the Mcm2-7 complex, the replicative helicase, is loaded as a
double hexamer at ORC binding sites in an ORC-, Cdc6-, and Cdt1-
dependent manner to form the pre-replicative complex (pre-RC)
(Evrin et al, 2009; Remus et al, 2009; Gambus et al, 2011). The
assembly of the pre-RC in G1 “licenses” the origin for potential
activation in the subsequent S-phase. As a cell enters S-phase,
cyclin-dependent kinase (CDK) and Dbf4-dependent kinase (DDK)
activate the Mcm2-7 helicase by the recruitment of Cdc45 and the
GINS complex to form the CMG (Cdc45, Mcm2-7, and GINS) holo-
helicase complex (reviewed in Tanaka & Araki, 2013). After initia-
tion of DNA replication, the Mcm2-7 helicase, as part of the CMG
complex, travels with and unwinds the DNA ahead of the replica-
tion fork (Aparicio et al, 1997; Labib, 2000; Pacek & Walter, 2004;
Pacek et al, 2006; Sekedat et al, 2010).
In theory, a double hexamer of Mcm2-7 loaded at each origin
should be sufficient to replicate the genome, with a single hexamer
of Mcm2-7 traveling with each bidirectional DNA replication fork.
However, nuclear Mcm2-7 protein levels are in vast excess relative
to ORC or the number of replication origins (Burkhart et al, 1995;
Lei et al, 1996; Donovan et al, 1997; Mahbubani et al, 1997;
Edwards et al, 2002). In vitro, multiple double hexamers of Mcm2-7
are able to be loaded at ORC binding sites by reiterative rounds of
ATP hydrolysis (Bowers et al, 2004; Evrin et al, 2013). Electron
microscopy revealed that these double hexamers are distributed
throughout the DNA template (Evrin et al, 2009; Remus et al,
2009), suggesting the ability of the Mcm2-7 complex to passively
translocate away from ORC binding sites. Similarly, in vivo immuno-
fluorescence and chromatin association studies have revealed that
Mcm2-7 are broadly distributed throughout the nucleus and exhibit
little co-localization with ORC (Madine et al, 1995; Krude et al,
1996; Romanowski et al, 1996b; Ritzi et al, 1998; Edwards et al,
2002; Harvey & Newport, 2003). Together, these seemingly paradox-
ical observations regarding the location and quantity of Mcm2-7
have been termed the “MCM Paradox” (Takahashi et al, 2005).
Although the mechanisms regulating the loading of multiple
Mcm2-7 complexes and their distribution throughout the genome
are unclear, increasing data suggest that the excess Mcm2-7 are
Pharmacology and Cancer Biology, Duke University Medical Center, Durham, NC, USA
*Corresponding author. Tel: +1 919 681 6077; E-mail: david.macalpine@duke.edu
†
These authors contributed equally
ª2015 The Authors The EMBO Journal Vol 34 |No 4|2015 531
Published online: January 2, 2015
important for maintaining genome stability during replicative stress.
Mcm2-7 levels can be depleted by more than 90% with little, if any,
impact on progression through S-phase (Woodward et al, 2006;
Crevel et al, 2007; Ge et al, 2007; Ibarra et al, 2008). However, in
the absence of the full complement of Mcm2-7, there is a marked
reduction in the utilization of dormant origins and an increase in
cell death when cells encounter replicative stress during S-phase
(Crevel & Cotterill, 1991; Woodward et al, 2006; Ge et al, 2007;
Ibarra et al, 2008). Thus, the full complement of Mcm2-7 is critical
for the activation of dormant replication origins and function to
preserve genome integrity during replicative stress.
We set out to examine this “MCM Paradox” from a biochemical
and genome-wide perspective by using chromatin association assays
and genome-wide chromatin immunoprecipitation (ChIP) to quan-
tify the loading and distribution of Mcm2-7 at different points in the
cell cycle of Drosophila Kc cells. Specific questions we set out to
address included: (i) How does Mcm2-7 loading onto chromatin
progress during G1? (ii) Do Mcm2-7 complexes load adjacent to
ORC? (iii) Where are Mcm2-7 distributed throughout the genome?
and (iv) Do other chromosomal features (e.g. transcription units)
impact the distribution of Mcm2-7?
Results
Mcm2-7loading is regulated by cyclin E/Cdk2activity
Mcm2-7 chromatin association is dynamic throughout the cell cycle.
Mcm2-7 are loaded onto chromatin in G1, travel ahead of the repli-
cation fork in S-phase, and are removed from the chromatin as
S-phase progresses (reviewed in Masai et al, 2010). In order to
examine the dynamics of Mcm2-7 chromatin association, we devel-
oped approaches to arrest Drosophila Kc cells at specific points in
the cell cycle when Mcm2-7 would presumably be associated or not
associated with ORC at potential origins of replication. We were
particularly interested in identifying a defined cell cycle point imme-
diately before Mcm2-7 helicase activation and its movement with
the DNA replication fork.
We reasoned that inhibition of cyclin E/Cdk2 kinase activity
would be sufficient to arrest cells in G1 immediately prior to origin
activation. We first depleted cyclin E and Cdk2 separately using
RNAi in Drosophila Kc cells. In Drosophila, cyclin E is the regula-
tory subunit of the cyclin-dependent kinase 2 (Cdk2), both of
which, like their mammalian homologs, function together to drive
cells out of G1 and into S-phase (Dulic et al, 1992; Ohtsubo &
Roberts, 1993; Knoblich et al, 1994). We also overexpressed Dacapo,
a p27 homolog and potent inhibitor of cyclin E/Cdk2 kinase
activity (Lane et al, 1996), from a copper-responsive metallothion-
ein promoter. As controls, cells were arrested in early G1 by RNAi
depletion of Dup/Cdt1, a key replication licensing factor required
for Mcm2-7 loading (Whittaker et al, 2000) and at the G1/S transi-
tion by treatment with 1 mM hydroxyurea (HU). Dup/Cdt1 RNAi,
cyclin E RNAi, Cdk2 RNAi, Dacapo overexpression (+Dacapo), and
HU treatment all resulted in a cell cycle arrest with the majority
(70–85%) of cells having a 2C DNA content by flow cytometry
(Fig 1A). In contrast, only 30–40% of the cells from an asynchro-
nous population of Drosophila cells exhibit 2C DNA content
(Supplementary Fig S1D).
We next assessed the relative amounts of nuclear chromatin-
associated Mcm2-7 in each of these conditions by chromatin
fractionation (Fig 1B). A polyclonal antibody specific to Drosophila
Orc2 (Austin et al, 1999) and a monoclonal antibody (AS1.1) that
recognizes all six Mcm2-7 subunits (Chen et al, 2007) were used to
assess the chromatin-associated and whole cell extract levels of ORC
and Mcm2-7. As expected, cells lacking the licensing factor, Dup/
Cdt1, exhibited no detectable chromatin-associated Mcm2-7 (Fig 1B,
lane 1). The background levels of Mcm2-7 chromatin association in
the Dup/Cdt1-depleted cells were similar to the levels observed in
G2-arrested cells where the presence of geminin prevents pre-RC
formation (McGarry & Kirschner, 1998) (Supplementary Fig S1B). In
contrast, cells arrested at the G1/S transition by HU treatment had a
robust accumulation of nuclear Mcm2-7 (Fig 1B, lane 5). Interest-
ingly, cyclin E and Cdk2 RNAi cells exhibited an intermediate
phenotype (Fig 1B, lanes 2 and 3) with considerably less nuclear
chromatin-associated Mcm2-7 than HU-arrested cells. We observed
a slight increase in the amount of chromatin-associated Mcm2-7 in
+Dacapo cells (Fig 1B, lane 4) over both cyclin E and Cdk2 RNAi
cells. However, the amount of chromatin-bound Mcm2-7 in +Dacapo
cells was still markedly reduced compared to levels seen in
HU-treated cells. In contrast to the cell cycle fluctuations in chroma-
tin-associated Mcm2-7 levels, ORC remained constant for each
condition surveyed. Importantly, the cell cycle differences observed
in Mcm2-7 chromatin association were not simply due to changes in
protein levels as the levels of ORC and Mcm2-7 remained constant
in the non-nuclear cytoplasmic fraction. The chromatin-associated
Mcm2-7/ORC ratio was calculated for each condition, normalized to
the level in HU to obtain a fold difference, and plotted on a log
10
scale (Fig 1C). A 40-fold difference in chromatin-associated Mcm2-7
was observed between Dup/Cdt1 RNAi and HU and 9-fold to 15-fold
differences in loading were observed between HU and cells with
impaired cyclin E/Cdk2 kinase activity (cyclin E RNAi, Cdk2 RNAi,
and +Dacapo). Together, our results suggest that while minimal
loading of the Mcm2-7 complex can occur in the absence of cyclin
E/Cdk2 activity, the loading of the full complement of Mcm2-7
requires cyclin E/Cdk2 kinase activity.
Maximal loading of Mcm2-7coincides with entry into S-phase
Our results suggest that Mcm2-7 levels continue to increase during
G1 concomitant with increasing Cdk2 kinase activity prior to entry
into S-phase. However, it remained possible that the maximal
Mcm2-7 loading we observed was perhaps a consequence of replica-
tive stress resulting from HU treatment and the activation of
dormant replication origins (Anglana et al, 2003). Thus, we wanted
to assess Mcm2-7 loading without perturbing S-phase or activating
the intra-S-phase checkpoint.
To monitor pre-RC assembly as cells synchronously enter
S-phase from G1, we induced overexpression of Dacapo to block
cyclin E/Cdk2 kinase activity and prevent the full complement of
Mcm2-7 from being assembled on the chromatin in G1. We then
released the cells from the Dacapo arrest by removing the inducer
(Cu
2+
) of Dacapo expression. Entry into the next cell cycle was
prevented by the addition of the mitotic inhibitor, colcemid, to the
medium. As cells progressed into S-phase, we monitored DNA
content, Dacapo protein levels, and the chromatin-associated levels
of Mcm2-7 and ORC (Fig 2). We found that most cells entered
The EMBO Journal Vol 34 |No 4|2015 ª2015 The Authors
The EMBO Journal Transcription shapes the distribution of Mcm2-7Sara K Powell et al
532
Published online: January 2, 2015
S-phase by 3 h and had completed DNA replication by 9 h. Entry
into S-phase at 3 h coincided with a decrease in Dacapo expression
and a marked increase (~9-fold) in Mcm2-7 association with the
chromatin. The increase in Mcm2-7 chromatin association at the
onset of S-phase entry was transient as Mcm2-7 were likely removed
from chromatin with passage of the DNA replication fork (Kuipers
et al, 2011). Together, these results demonstrate that the cyclin E/
Cdk2 kinase-dependent loading of the full complement of Mcm2-7 is
a regulated process that occurs during a normal cell cycle.
We also examined Mcm2-7 nuclear localization by immunofluo-
rescence in an asynchronous cell population (Supplementary Fig
S2). In order to visualize chromatin-associated Mcm2-7 in the
nucleus, it is necessary to remove the cytoplasmic and non-
chromatin-associated Mcm2-7 population (which are the majority)
by permeabilizing and washing the cells prior to fixation (Claycomb
et al, 2002). Unfortunately, the permeabilization prior to fixation
makes it impossible to monitor non-chromatin-associated factors
such as cyclin E or Cdk2 levels. In the absence of a suitable cell
cycle marker for cells in G1 or early S-phase, we instead used mono-
methylation of lysine 20 on histone H4 (H4K20me1) as a marker of
cells in late S-phase or G2/M. H4K20me1 is a well-characterized cell
cycle-regulated chromatin mark that is generated by PR-Set7/Set8 in
very late S-phase and persists on the chromatin up until mitosis
(Abbas et al, 2010; Oda et al, 2010) (Supplementary Fig S2A). Thus,
cells in late S-phase and G2/M will be identifiable by elevated levels
of H4K20me1 and, in contrast, cells in G1 and early to mid-S-phase
will be marked by very low levels of H4K20me1. We found that
H4K20me1 and Mcm2-7 staining cell populations were mutually
exclusive with very little overlap consistent with Mcm2-7 being
removed from chromatin by the end of S-phase (Supplementary Fig
+ Dacapo HU
Cdk2
RNAi
Cyclin E
RNAi
Dup/Cdt1
RNAi
Counts
Nuclear
chromatin
fraction
Mcm2-7
Orc2
Mcm2-7
Orc2
Cytoplasmic
fraction
*
Dup/Cdt1 RNAi
Cyclin E RNAi
Cdk2 RNAi
+Dacapo
HU
A
BC
2C 4C 2C 4C 2C 4C 2C 4C 2C 4C
Dup/Cdt1 RNAi
Cyclin E RNAi
Cdk2 RNAi
+ Dacapo
HU
1
10
100
Normalized Mcm2-7/ORC ratio (log 10)
1 2 3 4 5
Figure 1. Cyclin E/Cdk2activity modulates Mcm2-7loading during G1.
A FACS profiles of DNA content for cells arrested at different points in the cell cycle by Dup/Cdt1RNAi, cyclin E RNAi, Cdk2RNAi, Dacapo overexpression (+Dacapo), and
1mM HU.
B Analysis of Mcm2-7nuclear chromatin association at different points in the cell cycle. Nuclear chromatin fractions and cytoplasmic extracts were assayed for Orc2
and Mcm2-7by Western blot. A non-specific band is indicated by an asterisk (*).
C Quantification of the ratio of chromatin-bound Mcm2-7relative to Orc2(log
10
scale) for a minimum of 11 replicates (mean SD).
Mcm2-7
Orc2
Mcm2-7
Orc2
Dacapo
Hours post-Dacapo release
0 1 3 5 7 9 11 13
Nuclear
chromatin
fraction
Cytoplasmic
fraction
Figure 2. The full complement of Mcm2-7is loaded during an
unperturbed G1/S transition.
Cell cycle analysis by FACS of cells arrested in G1by Dacapo overexpression
followed by release back into the cell cycle for 13 h (top). Western blot analysis of
Mcm2-7and Orc2for the nuclear chromatin and cytoplasmic fractions of
Mcm2-7, Orc2, and Dacapo.
ª2015 The Authors The EMBO Journal Vol 34 |No 4|2015
Sara K Powell et al Transcription shapes the distribution of Mcm2-7The EMBO Journal
533
Published online: January 2, 2015
S2B). The remaining H4K20me1-negative cells displayed a range of
Mcm2-7 staining from barely detectable to a robust signal, presum-
ably representing cells in early G1 (low cyclin E/Cdk2 activity) and
cells in late G1/early S-phase (high cyclin E/Cdk2 activity), respec-
tively (Supplementary Fig S2C). Thus, even in a dividing population
of cells, we can clearly detect multiple stages of Mcm2-7 loading in
G1 and early S-phase cells. These findings also parallel an increase in
chromatin-associated Mcm2-7 from late M phase into and peaking at
the G1/S transition (Kuipers et al,2011;Symeonidouet al, 2013).
The cyclin E/Cdk2kinase-dependent phase of Mcm2-7loading
requires the canonical pre-RC assembly pathway
It is well documented that ORC, Cdc6, and Cdt1 are able to load
multiple Mcm2-7 double hexamers onto DNA templates in vitro
(Bowers et al, 2004; Evrin et al, 2009; Remus et al, 2009). In vivo,
the situation is more complex with ORC, Cdc6, and Cdt1 dynami-
cally associating with chromatin (McNairn et al, 2005; Xouri et al,
2007; Sonneville et al, 2012) and ATP hydrolysis driving both the
loading and release of the Mcm2-7 helicase complex (Frigola et al,
2013). It was evident from our biochemical studies that Dup/Cdt1
was required to load minimal amounts of Mcm2-7 in a cyclin
E-independent manner (Fig 1). However, it remained unclear
whether the full complement of Mcm2-7 loading we observed,
which is stimulated by cyclin E/Cdk2 kinase activity, was dependent
on the canonical pre-RC assembly pathway (e.g. Cdc6 and
Dup/Cdt1).
To determine whether the full complement of Mcm2-7 loading
we observed in HU-arrested cells is dependent on the established
pre-RC assembly pathway, we depleted cells of Cdc6 and Dup/Cdt1
immediately after the cyclin E/Cdk2-independent pre-RC assembly
step (Fig 3A). Specifically, we arrested cells in early G1 by RNAi
depletion of cyclin E for 24 h followed by treatment with either
control (pUC) or Cdc6 and Dup/Cdt1 dsRNA for 24 h. Cyclin E was
subsequently restored by overexpression of RNAi-resistant FLAG-
cyclin E. To assess Mcm2-7 loading at a static point in the cell cycle
immediately after entry into S-phase, the cells were released into
medium containing 1 mM HU. As expected, the majority of cells
from each condition arrested with a 2C DNA content (Fig 3B).
We found that cells depleted of Dup/Cdt1 and Cdc6 were unable
to recruit additional Mcm2-7 when cyclin E expression and progres-
sion through G1 were restored by FLAG-cyclin E overexpression
Kc FLAG-cyclin E-
inducible cells
Cyclin E RNAi
pUC RNAi Dup/Cdt1 &
Cdc6 RNAi
Induce FLAG-cyclin E & 1 mM HU
HU
Cyclin E RNAi
Dup/Cdt1
RNAi pUC RNAi Dup/Cdt1 & Cdc6 RNAi
Counts
2C 4C 2C 4C 2C 4C 2C 4C 2C 4C 2C 4C
+FLAG-E +FLAG-E
Mcm2-7
Orc2
Cyclin E RNAi
pUC RNAi
Dup/Cdt1 RNAi
FLAG-cyclin E
Cdc6 RNAi
1 mM HU
Mcm2-7
Orc2
Dup/Cdt1
Cyclin E
1 2 3 4 5 6
B
AC
-+
++
++
-+
+-
--
-+
--
-+
++
--
-+
--+-
+-
-+
+-
++
Cytoplasmic
fraction
Nuclear
chromatin
fraction
Figure 3. The cyclin E/Cdk2kinase-dependent phase of Mcm2-7loading requires the canonical pre-RC assembly pathway.
A Schematic of the experiment.
B FACS profiles of DNA content for each cell population assayed.
C Western blot analysis for Orc2and Mcm2-7in the nuclear chromatin fraction and Orc2, Mcm2-7, Dup/Cdt1, and cyclin E in the cytoplasmic fraction for each
condition assayed.
The EMBO Journal Vol 34 |No 4|2015 ª2015 The Authors
The EMBO Journal Transcription shapes the distribution of Mcm2-7Sara K Powell et al
534
Published online: January 2, 2015
(Fig 3C, lanes 4 and 5). In contrast, control RNAi (pUC) cells were
able to load additional Mcm2-7 when driven through G1 by FLAG-
cyclin E overexpression (Fig 3C, lane 3). The increase of chromatin-
associated Mcm2-7 was not due to a difference in FLAG-cyclin E
overexpression. These results indicate that Dup/Cdt1 and Cdc6 are
needed to load the full complement of Mcm2-7 onto chromatin
throughout G1 in a regulated manner.
Genome-wide distribution of Mcm2-7is determined by
cyclin E/Cdk2activity and local transcription
Earlier immunofluorescence-based studies noted that Mcm2-7 do
not strictly co-localize with ORC in the nucleus during S-phase
(Madine et al, 1995; Krude et al, 1996; Romanowski et al, 1996b).
The dichotomy in Mcm2-7 chromatin association that we observed
between cells arrested by cyclin E RNAi and cells arrested at
the G1/S transition by HU treatment prompted us to investigate
the genome-wide distribution of Mcm2-7 relative to ORC.
Specifically, we used chromatin immunoprecipitation to address
where Mcm2-7 was localized in relation to ORC in cyclin E- and
HU-arrested cells.
We used genome-wide mapping experiments to localize ORC in
asynchronous cells (MacAlpine et al, 2010; Roy et al, 2010)
(Fig 4A) and Mcm2-7 in cyclin E RNAi (Fig 4B) treated cells. We
identified 5,135 ORC peaks and 3,792 Mcm2-7 peaks throughout
the Drosophila Kc genome. The concordance between ORC and
Mcm2-7 in cyclin E RNAi treated cells was >93%, consistent with
the critical role of ORC in pre-RC assembly (Fig 4C). Not surpris-
ingly, we found that ORC and Mcm2-7 were specifically enriched
at early activating origins of DNA replication mapped by the
incorporation of a nucleotide analog, BrdU, during an HU arrest
(Supplementary Fig S3).
Strikingly, we found a very different pattern of Mcm2-7 localiza-
tion at the G1/S transition during an HU arrest (Fig 5A). In contrast
to the tight co-localization with ORC observed in the cyclin
E-depleted cells, we observed a “binary” pattern of Mcm2-7 localiza-
tion across the genome. Specifically, we observed broad chromo-
somal regions containing Mcm2-7 signal punctuated by the absence
of Mcm2-7 localization. The dramatic change in Mcm2-7 was not
due to a global change in chromatin configuration in the HU-
arrested cells because ORC binding remained indistinguishable
between the asynchronous and HU-arrested cells (Supplementary
Fig S4). Together with our biochemical results, these data suggest
that the full complement of Mcm2-7 we observed at the G1/S transi-
tion has re-distributed from ORC binding sites. We do not believe
that Mcm2-7 are completely coating the DNA, but rather that we are
observing the likelihood of detecting Mcm2-7 signal within specific
genomic features (see below).
The binary distribution of Mcm2-7 across the genome prompted
us to investigate genomic features that may be associated with the
broad regions of high or low Mcm2-7 levels along the chromosome.
The Mcm2-7 localization pattern relative to annotated genomic
features suggested that Mcm2-7 may be displaced by actively tran-
scribed genes (Fig 5A). To quantitatively assess the Mcm2-7 distri-
bution relative to transcription units, we generated histograms of
Mcm2-7 enrichment for transcribed and non-transcribed genes
(Fig 5B) and found a bimodal pattern of Mcm2-7 enrichment.
Specifically, active genes had no or very little Mcm2-7 signal,
whereas inactive or non-transcribed genes exhibited an elevated
Mcm2-7 signal (P<1.02 ×10
257
;t=40.16). We also considered
1627 284
3508
ORC Mcm2-7
C
1.00 2.25 3.50 4.75 6.00
Chromosome 2L (Mb)
-20246
Mcm2-7 CycE RNAi
B
1.00 2.25 3.50 4.75 6.00
Chromosome 2L (Mb)
-20246
ORC
A
Figure 4. Mcm2-7loading is restricted to ORC binding sites in the absence of cyclin E.
A Genome-wide analysis of ORC localization by ChIP-chip. ORC enrichment from asynchronous cells is depicted for a 5-Mb section of chromosome 2L.
B Genome-wide analysis of Mcm2-7localization in early G1by ChIP-chip. Mcm2-7enrichment from cyclin E RNAi-depleted cells is depicted for a 5-Mb section of
chromosome 2L.
C Venn diagram depicting the overlap between ORC and Mcm2-7peaks.
ª2015 The Authors The EMBO Journal Vol 34 |No 4|2015
Sara K Powell et al Transcription shapes the distribution of Mcm2-7The EMBO Journal
535
Published online: January 2, 2015
ED
CB
A
Early
Late
0
3
03
0
05 30 35 0
0
050100
Percent of Meta Gene
Mcm2-7 HU Enrichment
Mcm2-7 Late S-Phase ChIP Signal
Mcm2-7 Late S-Phase ChIP Signal
Figure 5. The Mcm2-7chromatin distribution in G1is dependent on cyclin E/Cdk2activity and the transcription machinery.
A Genome-wide analysis of Mcm2-7localization at the G1/S transition by ChIP-chip. Mcm2-7enrichment from HU-arrested cells is depicted for a 5-Mb section of
chromosome 2L. Inset: transcribed (green) and non-transcribed (red) genes are indicated above with genes on the positive strand on the top and those on the
negative on the bottom.
B Bimodal distribution of Mcm2-7enrichment over transcribed and non-transcribed genes. Histogram showing the distribution of probe scores found within
transcribed (green) and non-transcribed (red) genes.
C“Meta”-gene analysis of Mcm2-7enrichment for different deciles of gene expression and their aggregated probe intensities.
D Mcm2-7is displaced from chromatin by DNA replication. Genome-wide analysis of Mcm2-7localization in late S-phase (6h post HU release) by ChIP-chip. Mcm2-7
enrichment is depicted for a 10-Mb section of chromosome 2L (filled gray), replication timing profile (black line) and early (yellow) and late (purple) replication timing
domains.
E Box-plots representing late S-phase Mcm2-7ChIP signal found within early (246) or late (167) replication domains.
The EMBO Journal Vol 34 |No 4|2015 ª2015 The Authors
The EMBO Journal Transcription shapes the distribution of Mcm2-7Sara K Powell et al
536
Published online: January 2, 2015
that the bimodal distribution of Mcm2-7 enrichment between active
and inactive genes might be due to the activation of early origins
that are enriched near actively transcribed genes. However, we
found that the bimodal distribution for Mcm2-7 was not dependent
on early origin activity, but instead was a specific feature of anno-
tated transcripts (Supplementary Fig S5).
The enrichment of Mcm2-7 signal at non-transcribed genes was
indistinguishable from intergenic levels (Supplementary Fig S6),
suggesting that active transcription was responsible for displacing
Mcm2-7 from chromatin. We reasoned that if active transcription
was sufficient to displace Mcm2-7 from chromatin, then in the
absence of additional Mcm2-7 loading, we should expect that both
low and high transcription levels should be able to displace Mcm2-7
from chromatin. To address this, we binned the 14,594 genes into
deciles based on their relative gene expression levels and plotted the
Mcm2-7 enrichment relative to a “meta”-gene body (Fig 5C). We
found that Mcm2-7 levels were depleted over the entire transcrip-
tion unit for actively transcribed genes. The seven most expressed
deciles (green and gray lines) exhibited the same degree of Mcm2-7
depletion, suggesting that any amount of transcription is sufficient
to displace Mcm2-7 from the chromatin.
We sought to directly test the requirement for active transcription
in the displacement of the Mcm2-7 complex from gene bodies. As
we only observe differences in Mcm2-7 distribution during an HU
arrest when further pre-RC assembly is blocked, we would need to
specifically alter transcription during the progression from G1 to S,
which would likely interfere with the transcription of key cell
cycle-regulated genes (e.g. E2F-regulated genes). As an alternative
approach, we examined the genome-wide localization of Mcm2-7 in
two different Drosophila cell lines (Kc167 and S2) treated with HU.
Although much of the transcriptome is the same between these cell
lines (Cherbas et al, 2011), we identified approximately 100 genes
that were significantly transcribed in one cell line and off in the
other. As before, we found that the Mcm2-7 complex was excluded
from actively transcribed genes in both cell lines (Supplementary
Fig S7). However, when we specifically examined those genes that
were only transcribed in one cell line, we observed a significant
(Mann–Whitney–Wilcoxon test; P<7×10
5
) decrease in Mcm2-7
occupancy that was specific for the transcribed cell line. We
conclude that transcription is required to displace the Mcm2-7
complexes from gene bodies. Together, these results suggest that
individual Mcm2-7 complexes can be displaced by active transcrip-
tion and that after the G1/S transition, they cannot be re-established
or translocate into these regions.
The Mcm2-7 helicase complex is inherently dynamic throughout
the cell cycle; it is loaded onto chromatin, travels with the replica-
tion fork, and is rapidly removed from the chromatin by passage of
the replication fork (Madine et al, 1995; Krude et al, 1996;
Romanowski et al, 1996a; Kuipers et al, 2011). Our biochemical and
genomic data indicate that the full complement of Mcm2-7 is loaded
onto chromatin in late G1 and redistributes throughout non-tran-
scribed genes and intergenic sequences by the G1/S transition. If this
broad distribution of chromatin-associated Mcm2-7 at the G1/S tran-
sition represents the true biological distribution of the Mcm2-7
complex, then we would expect that these Mcm2-7 complexes would
be displaced by replication during S-phase. To test this hypothesis,
we arrested cells at the G1/S transition by treatment with 1 mM HU
and then released them synchronously into S-phase. We surveyed
the genome-wide distribution of Mcm2-7 near the end of S-phase
(6 h after HU release) (Fig 5D, dark gray). We found that Mcm2-7
had been displaced from early replicating regions of the genome and
only remained at those sequences copied in late S-phase (Fig 5D,
black line and purple segments). To quantify these results, we used
our prior segmentation of the genome into early (yellow) and late
(purple) replicating domains (Lubelsky et al, 2014) and compared it
to the distribution of Mcm2-7 at the end of S-phase. We found a
significant enrichment of Mcm2-7 associated with late replicating
domains relative to early domains (Fig 5E, P<2.37 ×10
68
,
t=18.22). Together, these data argue that the broad distribution of
Mcm2-7 we observe at the G1/S transition represents the full and
functional complement of Mcm2-7 on the chromatin.
Discussion
In order to ensure the fidelity of the DNA replication program and
genome integrity, there are many more Mcm2-7 helicase complexes
loaded onto DNA than potential start sites marked by ORC. We have
examined this “MCM Paradox” using both genomic and biochemical
approaches to understand the mechanisms by which the full
complement of excess Mcm2-7 are loaded and distributed through-
out the genome to preserve genomic integrity. We found that in
early G1, minimal levels of Mcm2-7 were loaded onto chromatin at
ORC binding sites independent of cyclin E/Cdk2 activity (Fig 6).
Concurrent with and dependent on increasing cyclin E/Cdk2 activity
during G1, we observed an increase in Mcm2-7 loading that culmi-
nated in a 10-fold higher level of chromatin-associated Mcm2-7 at
the G1/S transition. Both the cyclin E/Cdk2 activity-independent
Mcm2-7 loading and cyclin E/Cdk2 activity-dependent Mcm2-7
loading were dependent on the canonical pre-RC assembly pathway.
Strikingly, the full complement of Mcm2-7 when assembled onto
chromatin was not restricted to sequences immediately adjacent to
ORC binding sites, but rather distributed throughout the genome
and shaped by active transcription.
Precise regulation of cyclin E/Cdk2kinase activity is critical for
pre-RC assembly and genomic stability
Pre-RC assembly begins in telophase (Dimitrova et al, 1999), contin-
ues through G1, and culminates just prior to entry into S-phase
(Symeonidou et al, 2013). The precipitous drop in CDK activity as
cells exit mitosis likely leads to the limited amounts of Mcm2-7 being
assembled onto chromatin that we detect specifically at ORC binding
sites. As the cells progress into G1, increasing levels of Cdk2 activity
contribute to cyclin E/Cdk2-dependent Mcm2-7 loading. Indeed, a
gradual increase of Cdk2 activity occurs following exit from mitosis
in cycling human cells (Spencer et al, 2013). Conversely, at some
point, increasing CDK activity leads to the inhibition of pre-RC
assembly and ultimately origin activation (reviewed in Masai et al,
2010). In S. cerevisiae, CDK activity targeted toward pre-RC compo-
nents directly impairs pre-RC assembly (Drury et al, 1997; Elsasser
et al, 1999; Nguyen et al, 2001); however, in higher eukaryotes, the
evidence for a direct role is less clear. In human cell culture, phos-
phorylation of Cdt1 by CDK actively promotes its degradation by the
SCF-Skp2 E3 ligase (Liu et al, 2004; Nishitani et al, 2006). However,
this appears to be a minor pathway as the bulk of Cdt1 is degraded in
ª2015 The Authors The EMBO Journal Vol 34 |No 4|2015
Sara K Powell et al Transcription shapes the distribution of Mcm2-7The EMBO Journal
537
Published online: January 2, 2015
S-phase by PCNA-coupled Cul4-Ddb1 destruction (Arias & Walter,
2006; Jin et al, 2006; Senga et al, 2006).
How does cyclin E/Cdk2 activity promote the loading of the full
Mcm2-7 complement? Prior experiments performed on quiescent
mammalian cells re-entering the cell cycle suggested that cyclin E/
Cdk2 kinase activity stabilizes Cdc6 and Cdc7, two factors critical
for pre-RC assembly and initiation, respectively (Mailand & Diffley,
2005; Chuang et al, 2009). In contrast, cycling cells become compe-
tent to assemble the pre-RC immediately after the metaphase to
anaphase transition (Clijsters et al, 2013). Following exit from
anaphase, Cdc6 is degraded by APC/C-Cdh1; however, there is a
short window between late G1 and S where Cdc6 protein may accu-
mulate before being degraded in S-phase by a PIP-box and Crl4-Cdt2
mediated mechanism (Clijsters & Wolthuis, 2014). This second
wave of Cdc6 expression in late G1 is likely dependent on cyclin E/
Cdk2 activity and E2f-regulated gene expression and may serve to
promote loading the full complement of Mcm2-7 on the chromatin.
Alternatively, the Mcm2-7 complexes, which are loaded proximal to
the ORC binding sites in a cyclin E- and Cdk2-independent manner,
may serve to promote additional Mcm2-7 loading upon exposure to
increasing cyclin E/Cdk2 kinase activity prior to S-phase by a direct
Mcm2-7–cyclin E–Cdt1 interaction (Geng et al, 2007). The transition
from a cyclin E/Cdk2 kinase-independent mode to cyclin E/Cdk2
kinase-dependent mode of Mcm2-7 loading would facilitate a feed
forward mechanism to amplify Mcm2-7 loading as G1 progresses.
The precise regulation of cyclin E/Cdk2 kinase activity during G1
is critical for pre-RC assembly and genome stability. Cyclin E/Cdk2
kinase activity is required to load the full Mcm2-7 complement, yet
deregulation of cyclin E/Cdk2 kinase activity results in a shortening
of G1, a lengthening of S-phase, and genomic instability (Ohtsubo &
Roberts, 1993; Resnitzky et al, 1994; Wimmel et al, 1994; Spruck
et al, 1999). The genomic instability and lengthening of S-phase
appear to be due to a defect in pre-RC assembly (Ekholm-Reed et al,
2004). We speculate that deregulation of cyclin E/Cdk2 kinase activ-
ity would result in the cells transitioning from mitosis being immedi-
ately exposed to high levels of cyclin E/Cdk2 kinase activity
instead of the gradual increase that normally occurs over the course
of G1 (Spencer et al, 2013). Thus, insufficient pre-RCs would be
Cyclin E/Cdk2 Activity
transcribed non-transcribed
ORC
2
3
4
56
7
Cdc6
Dup/Cdt1
Mcm2-7
Figure 6. Model of pre-RC assembly and redistribution of Mcm2-7in G1.
Mcm2-7is loaded at ORC binding sites immediately after entry into G1.AsG1progresses, increasing cyclin E/Cdk2activity promotes the loading of additional Mcm2-7
complexes resulting in the full complement of Mcm2-7being loaded by the end of G1. All Mcm2-7loading is dependent on Cdc6and Cdt1. Prior to or coinciding with the entry
into S-phase, the full complement of Mcm2-7redistributes along the chromosomes and is displaced from transcribed genes.
The EMBO Journal Vol 34 |No 4|2015 ª2015 The Authors
The EMBO Journal Transcription shapes the distribution of Mcm2-7Sara K Powell et al
538
Published online: January 2, 2015
assembled on chromatin as the cyclin E-deregulated cells rapidly
enter S-phase and may not properly redistribute throughout the
genome. In vivo, mutations that impact Mcm2-7 levels and pre-RC
assembly have been linked to genome instability and tumorigenesis
(Pruitt et al, 2007; Shima et al, 2007; Chuang et al, 2010).
Genome-wide redistribution of Mcm2-7
Our chromatin immunoprecipitation studies of Mcm2-7 localization
provide the first genome-wide view of the Mcm2-7 distribution in a
higher eukaryote and also reveal a dramatic reorganization of
Mcm2-7 during late G1. In the absence of cyclin E/Cdk2 kinase
activity (early G1), we find that there is near perfect concordance
between ORC and Mcm2-7 peaks. A similar distribution of overlap-
ping ORC and Mcm2-7 peaks is also observed in S. cerevisiae
(Wyrick et al, 2001; Eaton et al, 2010). Not surprisingly, these
genome-wide datasets from S. cerevisiae reinforce the essential role of
ORC in pre-RC assembly and suggest that Mcm2-7 loading is a local
phenomenon restricted to the adjacent sequences. In contrast, in
Drosophila, we see a dramatic change in the distribution of Mcm2-7
that occurs by the G1/S transition. Not only are more Mcm2-7 loaded
onto the chromatin, but they are also broadly enriched along the
entire chromosome and not restricted to ORC binding sites.
The distribution of the full Mcm2-7 complement along the
genome is shaped by active transcription. Mcm2-7 signal is enriched
in both intergenic and non-transcribed sequences, but depleted from
actively transcribed transcripts. The displacement of Mcm2-7 is not
localized to exons, but rather occurs over the entirety of the tran-
script suggesting that local sequence bias (e.g. increased GC content
in exons) is not a contributing factor. We do not envision that the
full complement of Mcm2-7 is coating these intergenic and non-
transcribed regions, but rather represent an increased probability of
finding an Mcm2-7 complex in static regions of the genome relative
to those that are actively undergoing transcription.
We propose that Mcm2-7 residing in transcribed regions are
displaced by the passage of RNA Pol II during transcription. The
displacement of Mcm2-7 during transcription is analogous to the
removal of Mcm2-7 from inactive origins during passage of the
replication fork in S-phase (Madine et al, 1995; Krude et al, 1996;
Romanowski et al, 1996a; Kuipers et al, 2011). The bimodal distri-
bution of Mcm2-7 at intergenic and non-transcribed genes relative
to active genes may only be established after the transition into
S-phase. Prior to S-phase and the cessation of pre-RC assembly,
Mcm2-7 are likely in a cycle of loading, translocation, and subse-
quent eviction in transcribed regions; however, once pre-RC assem-
bly is inhibited they are no longer able to re-occupy transcribed
regions following eviction by active transcription.
The mechanism(s) by which the Mcm2-7 enrichment transitions
from ORC-specific localization to a broad and distributed pattern
along the chromosome is not clear. We envision at least two possi-
bilities—translocation along the DNA away from ORC or the loading
of Mcm2-7 at distal sites by chromosomal looping. In vitro reconsti-
tution of Mcm2-7 loading on circular DNA templates revealed
Mcm2-7 double hexamers distributed randomly on the template irre-
spective of the location of the ACS or ORC binding site (Evrin et al,
2009; Remus et al, 2009). Thus, in vitro, Mcm2-7 double hexamers
can be loaded and are free to translocate along the dsDNA template.
However, it is very difficult to imagine how, in vivo, Mcm2-7
complexes could translocate along the DNA given the chromatin
obstacles such as nucleosomes, DNA binding proteins, and active
transcription. Nucleosomes would have to be displaced and re-
assembled throughout the genome requiring ATP-dependent chro-
matin remodeling activities and histone chaperones (Alabert &
Groth, 2012). Alternatively, the loading of Mcm2-7 at sites distal to
ORC may be achieved by chromatin looping mediated by cohesin
complexes that are close to ORC in Drosophila (MacAlpine et al,
2010). In mammalian cells, cohesin has been shown to be required
for looping chromatin at replication factories (Guillou et al, 2010).
Defining an origin of replication
The search for sequence-based replicators, similar to the ARS
element in S. cerevisiae, has been a holy grail for the mammalian
replication field. In vitro, any sequence can be replicated in Xenopus
extracts and plasmid-based assays have also exhibited very promis-
cuous replication (Krysan & Calos, 1991). Recently, the analysis of
mammalian replication intermediates by next-generation sequencing
identified G4-quadruplex sequences as potential replicators (Cayrou
et al, 2011; Besnard et al, 2012; Valton et al, 2014). Despite the
identification of G4 quadruplex structures as origins of replication,
there is little concordance between datasets, indicating that these
degenerate structures, which occur every few kilobases, are not
sufficient for origin specification. Work from the Hamlin group
using two-dimensional gel electrophoresis demonstrated that many
genomic fragments have the potential to harbor inefficient replica-
tion origins (Mesner et al, 2006). We propose that the broad distri-
bution of the full complement of Mcm2-7 we observe contributes to
the apparent promiscuity of metazoan origin selection. In vitro
experiments in Xenopus have shown that ORC is not required for
initiation after pre-RC assembly (Hua & Newport, 1998; Rowles
et al, 1999). Thus, those Mcm2-7 helicases located distally from
ORC would still have the potential to be activated and could likely
function as dormant origins in the presence of replicative stress.
In our prior studies, we have demonstrated that early origins of
replication, which are resistant to HU, are enriched for ORC binding
sites (MacAlpine et al, 2010). These ORC binding sites also serve as
focal points for the cyclin E-independent Mcm2-7 loading. Together,
these data suggest that Mcm2-7 loaded proximal to ORC binding sites
have an increased likelihood of activation during early S-phase. In
S. cerevisiae, Cdc45 specifically associates with early activating origins
in G1 of the cell cycle (Aparicio et al, 1999). Thus, during early G1 in
Drosophila, Cdc45 may specifically associate with the few Mcm2-7
complexes loaded proximal to ORC binding sites, increasing the likeli-
hood of activation of this subset of pre-RCs during entry into S-phase.
Materials and Methods
Drosophila cell culture
Cells were cultured in 150-mm plates at an approximate density of
1×10
6
cells/ml in Schneider’s Insect Cell Medium (Invitrogen)
supplemented with 10% heat-inactivated FBS (Hyclone) and 1%
penicillin/streptomycin/glutamine (Invitrogen). Dacapo was overex-
pressed for 48 h in the presence of 500 lM copper sulfate.
FLAG-tagged cyclin E was overexpressed in the presence of 500 lM
ª2015 The Authors The EMBO Journal Vol 34 |No 4|2015
Sara K Powell et al Transcription shapes the distribution of Mcm2-7The EMBO Journal
539
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copper sulfate in the medium. To arrest at G1/S, cells were incu-
bated with 1 mM HU for 24 h (unless otherwise noted). G2-arrested
cells were treated with 3% DMSO and assayed after 48 h.
dsRNA synthesis and RNAi
Primers were designed with a 50T7 sequence (TTAATACGACTC-
ACTATAGGGAGA) to amplify a 500–900 bp region in the gene of
interest. The region was amplified using standard PCR followed by
gel extraction (Qiagen Qiaquick gel extraction) and the PCR product
was subsequently used as a template for dsRNA production (Pro-
mega T7 Ribomax express large scale expression). The dsRNA was
purified by phenol/chloroform extraction followed by ethanol
precipitation. Primers used for each target: Dup/Cdt1 dsRNA, F:
50ctatcagtatcaagaacaggcg, R: 50tgctttccaccagactg; cyclin E dsRNA,
F: 50gccatccgtcacataagca, R: 50atcgtggaagcaagcagac; Cdk2 dsRNA, F:
50tgtggccctcaaaaagattc, R: 50gaagtaagcgtgctgcagtg; Cdc6 dsRNA, F:
50gccacagcacctgatcagtccttcgcg, R: 50gcagtttacaagaaactatgcacc; pUC
dsRNA, F: 50agctcactcaaaggcggtaa, R: 50gcctacatacctcgctctgc.
Cells were washed with serum-free Schneider’s insect medium
(Invitrogen), resuspended to 1 ×10
6
cells/ml in serum-free insect
medium, and then replated. dsRNA (15 lg/1 ×10
6
cells/ml) was
added to the medium and incubated for 1 h at 25°C. Medium with
2×serum was added to make the total medium 1×in respect to
serum. Cyclin E, Cdk2 and pUC RNAi cells were assessed 48 h after
RNAi, Dup/Cdt1 and Cdc6 RNAi cells were assessed after 24 h.
FACS preparation and analysis
Cells were harvested by centrifugation, washed with ice-cold 1×
PBS, resuspended in residual 1×PBS, and fixed overnight at 4°C
with ice-cold 100% ethanol. Next, the cells were centrifuged, the
supernatant removed, and washed with 1×PBS—1% fetal bovine
serum. The cells were resuspended in a final volume of 800 ll1×
PBS—1% fetal bovine serum with 50 mg/ml of propidium iodide
and 0.1 mg/ml RNaseA. The cells were incubated at 37°C for 1 h.
Equal numbers of cells per condition were assayed using a FACS-
Canto machine running BD FACSDiva software and R bioconductor
flowCore package to generate histograms.
Chromatin fractionation
Fractionation of the chromatin was optimized for Drosophila cell
lines from established protocols (Hancock, 1974; Ritzi et al, 1998).
Cells were harvested and then washed with ice-cold 1×PBS twice.
Cells were resuspended in hypotonic buffer [10 mM HEPES-KOH
(pH 7.5), 20 mM KCl, 0.25 mM EDTA, and a Roche protease mini
tab], kept on ice for 4 min then centrifuged for 4 min at 700 gat
4°C. The pellet was resuspended in lysis buffer [10 mM HEPES-KOH
(pH 7.5), 70 mM NaCl, 20 mM KCl, 5 mM MgCl
2
, 2 mM CaCl
2
,
0.5% NP-40, and a Roche protease inhibitor tab] and underlayed
with 1 volume of 30% sucrose. The lysate was centrifuged for
3 min at 3,000 gat 4°C. The supernatant was set aside as the whole
cell lysate sample. The pellet was resuspended in lysis buffer and
centrifuged with a sucrose cushion. The pellet was washed with low
salt buffer [20 mM HEPES/KOH (pH 7.5), 0.5 mM MgCl
2
, 0.3 M
sucrose, KCl, and Roche protease inhibitor tab] and centrifuged at
4°C for 3 min at 3,000 g. The pellet was washed again in low salt
buffer and resuspended in 2×SDS +b-ME loading buffer. To verify
that the proteins associated with the nuclear pellet were soluble,
they were washed in high salt buffer (480 mM HEPES/KOH (pH
7.5), 0.5 mM MgCl
2
, 0.3 M sucrose, KCl, and Roche protease inhibi-
tor tab) (Supplementary Fig S8A). Alternatively, to solubilize the
chromatin-bound fraction, pellets were resuspended in low salt
buffer supplemented with 5 mM CaCl
2
and treated with 45 U of
micrococcal nuclease for 10 min prior to quenching with 5 mM
EGTA (Supplementary Fig S8B).
Immunofluorescence
For double labeling of Mcm2-7 and H4K20me1, cells were treated
with 0.5% Triton X-100 in PBS for 1 min, washed, fixed with 4%
paraformaldehyde in PBS for 10 min, and blocked with 3% BSA in
PBS-T (0.1% Triton in PBS). Cells were stained with monoclonal
Mcm2-7 antibody (AS1.1, 1:100) and anti-H4K20me1 antibody
(ab9051, 1:1,000), followed by secondary detection with Alexa Fluor
568 goat anti-mouse and Alexa Fluor 633 goat anti-rabbit antibodies
(1:500). Images were acquired with Zeiss Axio Imager wide field
fluorescence microscope, and the median intensities of individual
cells (n=277) from each channel were measured by ImageJ. The
plot of H4K20me1 signal versus Mcm2-7 signal was generated in R.
Western blot analysis
Mcm2-7 monoclonal mouse antibody (AS1.1) was used at 1:100
(Chen et al, 2007), Orc2 rabbit polyclonal antibody was used at
1:3,000 (Austin et al, 1999), Dacapo (NP-1) used at 1:1,000 (Iowa
Hybridoma Bank), Dup/Cdt1 guinea pig raised antibody was used at
1:5,000, and cyclin E antibody raised in rabbits was used at 1:500
(d-300 Santa Cruz sc-33748). Secondary antibodies: goat anti-mouse
IRDye 800CW IgG (LiCor), goat anti-rabbit Alexa Fluor 680 IgG (Invi-
trogen), and goat anti-guinea pig IRDye 800 (Rockland Immuno-
chemicals), were all used at 1:10,000. Westerns were visualized and
quantified using LiCor infrared technology and gray scaled in ImageJ.
ChIP-chip sample preparation
Samples were prepared as described previously (MacAlpine et al,
2010). Mcm2-7 (AS1.1) antibody was used at a 1/25 dilution. All
experiments were performed in duplicate.
ChIP-Chip and ChIP-Seq analysis
Within-replicate probe intensities were determined and between-
slide intensities normalized via the R (R Development Core Team,
2008) package limma (Smyth, 2004). Replicated probe intensities
were determined via MA2C (Song et al, 2007). Subsequently, all
analyses were done in R using the combined replicate probe intensi-
ties. Gene bodies were obtained from (Graveley et al, 2011). Mcm2-7
ChIP-seq and corresponding input samples were performed on
HU-arrested Kc and S2 cells. log2 RPKM ratios of Mcm ChIP-seq to
input sample were calculated over all Drosophila gene models from
the 5.12 annotation release, excluding any genes overlapping a Kc
or S2 early origin. Genes were then subsequently subsetted into
untranscribed (<1 RPKM) and transcribed (>4 RPKM) groups
based on expression data from Cherbas et al (2011). log2 ratios of
The EMBO Journal Vol 34 |No 4|2015 ª2015 The Authors
The EMBO Journal Transcription shapes the distribution of Mcm2-7Sara K Powell et al
540
Published online: January 2, 2015
Mcm2-7 ChIP-seq to input samples were then compared between
the cell lines for four gene classes: untranscribed genes in both Kc
and S2 (4,593 genes), transcribed genes in both Kc and S2 (4,804
genes), untranscribed in Kc and transcribed in S2 (91 genes), and
transcribed in Kc and untranscribed in S2 (128 genes). The Mann–
Whitney–Wilcoxon Test was used to evaluate differences in the log2
ratio distributions between Kc and S2.
FLAG-cyclin E stable transfection
SLIC cloning (Li & Elledge, 2012) was used to introduce a FLAG
fusion tag in frame at either the N-terminus or C-terminus of cyclin
E cDNA (clone LD22682) in the pMK-CTAP plasmid (pMK-CTAP
was a gift from Artavanis laboratory). Briefly, the plasmid was
digested with a single restriction site enzyme, N term =XhoI, C
term =SpeI. The 50ends were treated with T4 DNA polymerase
(NEB) for 45 min at room temperature. Approximately 150 ng of
vector in a 1:1 ratio was annealed to FLAG primers in 1×ligation
mix (Invitrogen) at 37°C for 30 min. The reactions were trans-
formed into max efficiency DH5acells (Invitrogen). Effectene trans-
fection reagent kit (Qiagen) protocol was followed for suspension
cells in 100-mm dish, with the only exception being that 150 ll
Effectene reagent was added. Cells were plated at 1 ×10
6
cells/ml
and incubated at 25°C for 2 days. Hygromycin B (Sigma) was then
added to 0.125 lg/ml.
Data access
All genomic data are publicly available at NCBI GEO and SRA data
repositories with the following accessions numbers: GSE17282,
GSE17283, GSE41349, GSE63915, SRP050273.
Supplementary information for this article is available online:
http://emboj.embopress.org
Acknowledgements
We thank Bob Duronio, Don Fox, and MacAlpine laboratory members for criti-
cal comments on the manuscript. Don Rio for the Dacapo construct, Steve Bell
for Mcm2-7antibody, and the Artavanis laboratory for pMK plasmid. This work
was supported by an American Cancer Society Research Scholar Grant
(120222-RSG-11-048-01-DMC) and the National Institutes of Health
(1R01GM104097-01A1).
Author contributions
SKP and DMM designed the experiments and project. SKP performed the
biochemical experiments, HKM performed the genome-wide experiments, JAP
and JAB executed the bioinformatic analysis, and YL performed the immuno-
fluorescence experiments. SKP, HKM, and DMM wrote the manuscript.
Conflict of interest
The authors declare that they have no conflict of interest.
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