© 2015. Published by The Company of Biologists Ltd | Journal of Cell Science (2015) 128, 4171-4182 doi:10.1242/jcs.172429
RESEARCH ARTICLE
A new mitochondrial pool of cyclin E, regulated by Drp1, is linked to
cell-density-dependent cell proliferation
ABSTRACT
The regulation and function of the crucial cell cycle regulator cyclin E
(CycE) remains elusive. Unlike other cyclins, CycE can be uniquely
controlled by mitochondrial energetics, the exact mechanism being
unclear. Using mammalian cells (in vitro) and Drosophila (in vivo)
model systems in parallel, we show that CycE can be directly
regulated by mitochondria through its recruitment to the organelle.
Active mitochondrial bioenergetics maintains a distinct mitochondrial
pool of CycE (mtCycE) lacking a key phosphorylation required for
its degradation. Loss of the mitochondrial fission protein dynaminrelated protein 1 (Drp1, SwissProt O00429 in humans) augments
mitochondrial respiration and elevates the mtCycE pool allowing
CycE deregulation, cell cycle alterations and enrichment of stem cell
markers. Such CycE deregulation after Drp1 loss attenuates cell
proliferation in low-cell-density environments. However, in high-celldensity environments, elevated MEK–ERK signaling in the absence
of Drp1 releases mtCycE to support escape of contact inhibition and
maintain aberrant cell proliferation. Such Drp1-driven regulation of
CycE recruitment to mitochondria might be a mechanism to modulate
CycE degradation during normal developmental processes as well as
in tumorigenic events.
KEY WORDS: Cyclin E, Drp1, Mitochondria, Cell cycle,
Cell proliferation
INTRODUCTION
Spatiotemporal regulation of the cell cycle is crucial in various
developmental processes, with cell cycle deregulation leading to
cell proliferative disorders like tumorigenesis. As mitotic cells move
through the growth (G1) phase to the DNA synthesis (S) phase
followed by a short growth phase and mitosis (G2-M), they are
governed by specific cyclins at each phase (Lodish et al., 2000).
Cyclin E (CycE, SwissProt P24864 and O96020 in humans)
governs the transition from G1 to S phase by initiating DNA
replication (Hwang and Clurman, 2005). In a regulated mitotic
cycle, CycE abundance and activity is at a maximum during the G1S transition and progressively decreases through the S phase to a
minimum at the G2 and M phases. CycE binds and activates its
partner kinase cyclin-dependent kinase 2 (CDK2) to phosphorylate
downstream targets in the cell nucleus to initiate DNA synthesis
(Siu et al., 2011). In mouse models, CycE is dispensable for the
1
Department of Genetics, University of Alabama at Birmingham, Birmingham,
2
AL 35294, USA. Department of Chemistry, Georgia State University, Atlanta,
3
GA 30303, USA. Department of Cell Development and Integrative Biology,
4
University of Alabama at Birmingham, Birmingham, AL 35294, USA. Department of
Pathology, University of Alabama at Birmingham, Birmingham, AL 35294, USA.
*Author for correspondence ([email protected])
Received 9 April 2015; Accepted 30 September 2015
development of the embryo proper but is indispensable for
development of polyploidy, release from quiescence and
oncogenic transformation, all of which require S phase entry
(Geng et al., 2003). These functions of CycE have been found to be
CDK2 independent (Geng et al., 2007), the details of which still
remain to be investigated. Moreover, CDK2-independent CycE
localization to the centrosome might be linked to S phase entry
(Matsumoto and Maller, 2004). Apart from CDK2, CycE also binds
to CDK1, although this interaction remains less characterized
(Welcker and Clurman, 2005).
The CycE abundance is precisely regulated to restrict its activity
to the G1-S phase transition. This is achieved as active CycE is
phosphorylated and subsequently ubiquitylated to target it for
degradation (Siu et al., 2011). Deregulation of CycE abundance
and/or activity leads to deregulation of cell proliferation:
constitutively elevated CycE throughout the cell cycle causes
DNA damage and chromosome misalignment, ultimately leading to
genomic instability and aneuploidy (Spruck et al., 1999; Hwang and
Clurman, 2005). CycE overexpression has been noted in various
tumors, which can arise from loss of function of components
responsible for CycE degradation (Davis et al., 2014). Although
elevated nuclear CycE levels are used as markers for tumorigenicity
in various cancer types (Hwang and Clurman, 2005; Donnellan and
Chetty, 1999), overabundance of an uncharacterized cytosolic pool
of CycE has been noted in cancer tissues (Donnellan and Chetty,
1999). Specific low-molecular-weight forms of CycE have been
proposed to be the direct cause behind CycE-driven tumorigenicity
(Bedrosian et al., 2004; Bales et al., 2005).
The understanding of CycE regulation and function remains
incomplete and shrouded with controversies. Interestingly, CycE
stability requires ATP production from mitochondria in mammalian
cells and in Drosophila melanogaster (Mandal et al., 2010, 2005).
Enhancement of mitochondrial ATP production during the G1 to S
transition in normal proliferating fibroblasts likely stabilizes CycE
during this period of the cell cycle (Mitra et al., 2009, 2013).
Mitochondrial ATP-synthesizing capability can be directly or
indirectly influenced by a certain group of proteins that directly
modulate the ability of mitochondria to undergo fusion or fission
events with each other (Westermann, 2010; Galloway et al., 2012).
Reduction of the levels or activity of the key mitochondrial fission
protein, dynamin-related protein 1 (Drp1, SwissProt O00429 in
humans) leads to elevation of CycE levels in various mammalian
cells and in Drosophila (Mitra et al., 2012, 2009; Qian et al., 2012).
The emerging model suggests that cell cycle regulators regulate
Drp1 (Taguchi et al., 2007; Kashatus et al., 2011; Horn et al., 2011),
which, in turn, further regulates CycE levels in proliferating cells
(Mitra, 2013). We have previously shown that although loss of Drp1
deregulates CycE in various cell contexts, it promotes aberrant cell
proliferation only in the presence of EGFR signaling (Mitra et al.,
2012). Using mammalian cells and Drosophila model systems in
4171
Journal of Cell Science
Danitra J. Parker1, Archana Iyer2, Shikha Shah1, Aida Moran1, Anita B. Hjelmeland3, Malay Kumar Basu4,
Runhua Liu1 and Kasturi Mitra1,*
parallel, we report that a new mitochondrial pool of CycE, which
can be modulated by Drp1, likely through regulation of
mitochondrial energetics, is linked to control of cell proliferation
in a cell-density-dependent manner.
RESULTS
Detection of a new mitochondria-associated pool of CycE in
mammalian cells and in Drosophila
The canonical function of CycE is in the nucleus, but CycE might
re-locate to the centrosomes to promote centrosome duplication (Siu
et al., 2011) and to the Golgi membrane to be ubiquitylated (Lu and
Pfeffer, 2013). Here, we investigated whether the mitochondrial
effect on CycE (Mandal et al., 2010, 2005; Mitra, 2013; Qian et al.,
2012) can be mediated by recruitment of the molecule to
mitochondria. Immortalized mouse embryonic fibroblasts (MEFs)
had a distinct cytosolic pool of CycE (Fig. 1A, upper panel), a
fraction of which colocalized with the mitochondrial marker Tom20 (arrows in Fig. 1A, lower panel). This indicated the presence of a
mitochondrial CycE pool, referred to as mtCycE hereafter. Multiple
washes with PBS buffer before cell fixation reduced the level of the
mtCycE pool, dramatically indicating its labile and transient nature
(data not shown). We also detected the mtCycE pool in cancer cells
(Fig. S1A,B). Next, we quantified the mtCycE pool in individual
MEFs by colocalization analyses of CycE with Tom-20. Linear
regression analyses of the mtCycE versus the non-mtCycE pool
revealed a strong correlation between the two CycE pools,
suggesting that the two pools influence each other and possibly
interact (Fig. 1B). We further confirmed the presence of the new
mtCycE pool in crude mitochondrial fractions from MEFs, which
were devoid of the nuclear marker Sox-2 and the cytosolic marker
actin, and strongly enriched for the mitochondrial marker Tom20
and modestly for mitochondria-associated endoplasmic reticulum
(MAM) marker protein disulfide isomerase (PDI; also known as
P4HB) (Fig. 1C). Although the non-mitochondrial fraction harbors
the characteristic doublet bands of CycE, we consistently detected
the lower band of CycE in the mitochondrial fraction. The
specificity of the widely used anti-CycE antibody was confirmed
in cells transfected with CycE small interfering RNA (siRNA) to
reduce its protein level by only 20% to avoid affecting cell
proliferation (Fig. S1C). CycE after binding to CDK2 gets autophosphorylated at degrons, which is required for the subsequent
ubiquitylation and degradation of CycE (Siu et al., 2011).
Therefore, CDK2-dependent phosphorylation of CycE reflects the
active CycE pool that eventually gets degraded to keep CycE levels
regulated. We found that the active pool of CycE phosphorylated at
T62 [ p-(T62)CycE; the N-terminal degron] mostly resided in the
nucleus (Fig. 1D,E). In addition, colocalization of the transfected
mitochondrial reporter mito-RFP with p-(T62)CycE was
significantly lower than that with CycE in individual cells
(Fig. 1E,F). These data suggest that the mtCycE pool is devoid of
the CDK2-dependent T62 phosphorylation and thus deemed
inactive. However, it remains to be examined whether there are
other phosphorylated sites on mtCycE.
We next sought to detect the new mtCycE pool in vivo in the
Drosophila follicle cell layer where we have previously
demonstrated specific mitochondrial regulation of CycE (Mitra
et al., 2012). The follicle cell layer is the epithelial cell layer
encapsulating the Drosophila egg chambers. Using an antibody
against the Drosophila CycE (DmCycE), we detected a distinct pool
of DmCycE colocalizing strongly with the mitochondrial marker
ATP-B (the ATP synthase β subunit) in the terminally differentiated
follicle cell layer (Fig. 1G). The early follicle cells, after
4172
Journal of Cell Science (2015) 128, 4171-4182 doi:10.1242/jcs.172429
differentiating from the lineage-specific stem cells, undergo
mitotic divisions during developmental stages 1 through 6. After
stage 6, the follicle cells exit the mitotic cycle to terminally
differentiate into the epithelial cell layer, which is further patterned
into various cell types (Klusza and Deng, 2011). We have
previously reported differential mitochondrial regulation in the
mitotic follicle cells and the differentiated-patterned main body
follicle cells (MBCs) and posterior follicle cells (PFCs) (Mitra et al.,
2012). Here, we found that the mtDmCycE pool was significantly
higher in the MBCs than the PFCs or the mitotic follicle cells
(Fig. 1H; Fig. S2A), suggesting that the mtDmCycE pool is
developmentally regulated in the Drosophila follicle cell layer.
Our novel observation of the existence of the mtCycE pool
(revealed by two distinct antibodies against mammalian and
DmCycE) likely underlies the mechanism behind a direct
mitochondrial regulation of CycE. Based on the focal
organization of mtCycE (Fig. 1A) that was identified in a cell
fraction with modest enrichment of a MAM marker (Fig. 1C), we
speculate that the mtCycE pool could reside at contact sites between
mitochondria and endoplasmic reticulum.
An increase in the mtCycE pool caused by Drp1 loss
deregulates CycE
The levels of mtCycE in the various cell types in the Drosophila
follicle cell layer (Fig. 1H) negatively correlate with the previously
reported status of Drp1-driven mitochondrial fission (Mitra et al.,
2012), indicating that reduced Drp1 activity might elevate the
mtCycE pool. We tested this possibility in MEFs obtained from the
DRP1-knockout (DRP1-KO) embryos and thereafter immortalized
with the SV-40T antigen (Ishihara et al., 2009). Comparison of the
CycE and Tom-20 colocalization between the wild-type (WT) and
the DRP1-KO MEFs revealed a significantly elevated mtCycE pool
in the absence of Drp1 (Fig. 2A,B). Introduction of Drp1–GFP into
the DRP1-KO MEFs reduced the mtCycE pool when compared to
introduction of the EGFP vector (Fig. 2C), thus confirming that the
levels of Drp1 regulate the levels of the mtCycE pool. We further
validated the effect of Drp1 loss on the mtCycE pool in the
Drosophila follicle cell layer by generating Drp1 functionally null
clones to compare DmCycE localization between the clones and the
background WT follicle cells. We have previously shown that Drp1null follicle cell clones harbor hyperfused mitochondrial clusters
(Mitra et al., 2012). Here, we found that the majority of the DmCycE
pool localized to the mitochondrial clusters in the Drp1-null cells in
the differentiated MBC region (arrows in Fig. 2D) or in early mitotic
stages (Fig. S2B), confirming our observation in the MEFs.
We next isolated mitochondria from the WT and DRP1-KO
MEFs to confirm the increase in the mtCycE pool upon Drp1 loss.
The crude mitochondrial fractions from the DRP1-KO MEFs had
significant levels of actin (Fig. 2E), which might be of functional
relevance, as recently reported (Li et al., 2015). However, the MAM
marker, PDI, was more enriched in the WT mitochondrial fraction
than the DRP1-KO mitochondrial fraction. The mitochondrial
fractions from the DRP1-KO MEFs indeed harbored higher levels of
the low-molecular-weight band of CycE although the amount of the
non-mitochondrial upper bands of CycE was comparable between
the WT and DRP1-KO (Fig. 2F). Although, the mtCycE pool was
devoid of the CDK2-dependent T62 phosphorylation in both WT
and DRP1-KO cells (Fig. 2G), the non-mitochondrial active p-(T62)
CycE pool was higher in the DRP1-KO cells, which likely resulted
from higher CDK2 levels in those fractions (Fig. 2G). Furthermore,
a distinct CDK2 pool was also detected in the mitochondrial fraction
from both WT and DRP1-KO MEFs (Fig. 2G), with the latter
Journal of Cell Science
RESEARCH ARTICLE
Journal of Cell Science (2015) 128, 4171-4182 doi:10.1242/jcs.172429
Fig. 1. Identification of mtCycE. (A) Confocal micrographs of CycE and Tom-20 immunocytochemistry in MEFs (upper panels; N, nucleus). Boxed regions are
magnified in the lower panels showing colocalization of CycE and Tom20 (merged yellow pixels, arrows). (B) Linear regression analysis showing correlation of
the colocalized and non-colocalized CycE signal (with Tom20) quantified from confocal micrographs of individual cells (dots), and no correlation between the
colocalized and non-colocalized Tom20 signal (with CycE); R2 regression coefficients and P-values (Pearson’s correlation two-sided test) are given.
(C) Immunoblot showing a distinct CycE band in crude mitochondrial fractions (Mito). (D) Low-resolution confocal micrograph of immunocytochemistry of HSP-60
(SwissProt P10809 in humans) and p(T62)CycE revealing the latter is localized in cell nucleus housing DNA. (E) High-resolution confocal micrograph of
immunocytochemistry of CycE and p-(T62)CycE in a representative mitoRFP-expressing cell (*). (F) Quantification of colocalization of mitoRFP signal with CycE
or p-(T62)CycE in individual cells, showing the latter does not have a prominent mitochondrial pool. Results are mean±s.e.m. (n>30). (G) Immunohistochemistry
in Drosophila ovarioles revealing colocalization of DmCycE and ATP-B in the differentiated follicle cell (FC) layer (merged yellow pixels). (H) Quantification
showing that there is a higher fraction of the colocalized DmCycE pool (with ATP-B) in differentiated MBCs when compared to that of the mitotic follicle cells or
differentiated PFCs. Results are mean±s.e.m. (n>10). *P≤0.05 (Student’s t-test). Scale bars: 10 µm (A,E,G), 40 µm (D).
showing an extra CDK2 band (Fig. S1D). Our data demonstrates
that loss of Drp1 leads to enhancement of the mtCycE pool devoid
of the p-T62 degron (inactive) with an associated increase in the
amount of non-mitochondrial p-(T62)CycE (active).
Next, we investigated whether the enhanced mtCycE pool in the
DRP1-KO MEFs is associated with aberrantly high CycE levels
throughout the cell cycle. We detected only modestly higher CycE
levels in the asynchronously proliferating DRP1-KO MEFs when
compared to the WT. However, DRP1-KO MEFs synchronized in
S phase or in G2-M phase had significantly higher levels of CycE
than similarly synchronized WT MEFs (Fig. 2H). This finding is
consistent with observations in mitotic breast cancer cells (Qian
et al., 2012) and suggests that Drp1 loss reduces degradation of
CycE throughout the cell cycle. Inhibition of CycE degradation in
mitosis can sustain aberrantly higher levels of the mitotic cyclin
cyclin B (Keck et al., 2007). Indeed, we observed that cyclin B
4173
Journal of Cell Science
RESEARCH ARTICLE
RESEARCH ARTICLE
Journal of Cell Science (2015) 128, 4171-4182 doi:10.1242/jcs.172429
levels were higher in the mitotic DRP1-KO MEFs in comparison to
the WT cells, whereas cyclin A levels were not substantially
different between them (Fig. 2H; Fig. S1E).
Taken together, our results suggest that Drp1 loss elevates the
mtCycE pool and deregulates CycE by maintaining higher levels of
CycE throughout the cell cycle. As the mtCycE pool lacks the p-T62
degron, our results imply that elevation of mtCycE pool in the
absence of Drp1 impairs CycE degradation, which is the major
cause of CycE deregulation (Siu et al., 2011).
The mtCycE pool is modulated by mitochondrial
bioenergetics that is governed by Drp1
CycE is actively degraded in Drosophila harboring a mutation in
Cytochrome c oxidase subunit V that lowers mitochondrial ATP
production (Mandal et al., 2010, 2005). Moreover, we have
previously found that the level of canonical nuclear CycE pool is
reduced upon mitochondrial depolarization (Mitra et al., 2009).
Therefore, we investigated whether maintenance of the mtCycE
pool is directly or indirectly mediated by mitochondrial energetics
and how Drp1 could regulate it. Mitochondrial oxygen
consumption is coupled to mitochondrial ATP synthesis by the
mitochondrial transmembrane potential maintained by the electron
4174
transport chain (ETC), while ATP is synthesized utilizing the
transmembrane potential (Nicholls and Ferguson, 2013). Inhibition
of mitochondrial ATP synthase (by treatment with oligomycin) or
disruption of the mitochondrial transmembrane potential (by
treatment with the protonophore CCCP) reduced the mtCycE pool
significantly within 3 h in both the WT and the DRP1-KO MEFs
(Fig. 3A,B). The release of mtCycE happened without any gross
change in mitochondrial shape, and mtCycE was removed from the
mitochondrial tubules in the DRP1-KO MEFs, but it remained
associated with swollen mitochondrial structures arising after
bioenergetic perturbations (arrows in Fig. 3A, lower panel,
compared with Fig. 2A, lower panel). Interestingly, DRP1-KO
MEFs that had an elevated mtCycE pool (Fig. 2A,B) also had
increased spare respiratory capacity as revealed by the enhanced
stimulation of the oxygen consumption rate (OCR) upon CCCP
treatment in the DRP1-KO MEFs, which have a basal OCR
comparable to the WT MEFs (Fig. 3C). Such boosted mitochondrial
energetics in the absence of DRP1 has also been found in other
mouse and human cells (Serasinghe et al., 2015; Kashatus et al.,
2015). Furthermore, we found that the enhanced spare respiratory
capacity between the DRP1-KO and WT MEFs was abrogated after
2 h of serum depletion, whereas both WT and DRP1-KO MEFs had
Journal of Cell Science
Fig. 2. Drp1 regulates CycE levels by modulating the mtCycE pool. (A) CycE and Tom-20 immunocytochemistry in WT and DRP1-KO MEFs.
(B) Quantification of Pearson’s colocalization coefficient (R) between CycE and Tom-20 revealing an elevated mtCycE pool in DRP1-KO MEFs. (C) Quantification
of Pearson’s colocalization coefficient (R) in DRP1-KO MEFs expressing EGFP or Drp1–EGFP demonstrating that Drp1 expression lowers the mtCycE pool.
Results are mean±s.e.m. (n>30). *P≤0.05 (Student’s t-test). (D) DmCycE and ATP-B immunohistochemistry in Drosophila ovarioles revealing a CycE pool
associated with mitochondrial clusters in Drp-null clone (arrows) as opposed to dispersed CycE in cytosol in the WT background in the follicle cell (FC) layer.
White lines demarcate clonal boundaries. (E) Immunoblot of the mitochondria-enriched fraction demonstrating enrichment of Tom20, PDI and reduced actin in the
WT mitochondrial fractions (mito) with the DRP1-KO mitochondrial fractions having less PDI enrichment and still retaining actin. 20 µg protein was loaded in each
lane. (F) Immunoblot demonstrating an elevated mtCycE pool in DRP1-KO MEFs. 20 µg protein was loaded in each lane. (G) Immunoblot demonstrating elevated
levels of p-(T62)CycE in the non-mitochondrial fraction (F1) of DRP1-KO MEFs in comparison to the WT; CDK2 is present in both the mitochondrial and nonmitochondrial fraction in WT and DRP1-KO MEFs. 20 µg protein was loaded in each lane. (H) Immunoblot of asynchronous (Asynch) and S- or G2-M-phasesynchronized WT and DRP1-KO MEFs showing elevated cyclin E and cyclin B levels in S and G2-M phases in DRP1-KO in comparison to the WT. There were
comparable cyclin A levels between WT and DRP1-KO. Tom-20 levels served as loading control. Numbers denote densitometric quantification of the bands as
first normalized by the loading control and then by the WT asynchronous values. Scale bars: 10 µm.
RESEARCH ARTICLE
Journal of Cell Science (2015) 128, 4171-4182 doi:10.1242/jcs.172429
enhanced spare respiratory capacity in the presence of serum
(Fig. 3D). We conclude that serum-dependent bolstered electron
transport chain activity (indicated by enhanced spare respiratory
capacity) in the DRP1-KO MEFs likely underlies the increase in the
mtCycE pool observed in these cells.
Our results demonstrate that the mtCycE pool is directly or
indirectly modulated by mitochondrial bioenergetics that is
governed by Drp1. Perturbation of mitochondrial bioenergetics
induced by physiological means, by drugs or by pathologic
mutations might release mtCycE and make it susceptible to
degradation, as observed in Drosophila harboring mitochondrial
mutants (Mandal et al., 2010, 2005).
Deregulation of CycE driven by Drp1 loss reduces G1 phase
and prolongs S phase, leading to enrichment of stem and
progenitor cell markers
Given that Drp1 loss elevates the potentially active CycE pool
(Fig. 2G), maintains higher levels of CycE throughout the cell cycle
(Fig. 2H) and recapitulates key CycE overexpression phenotypes
(Mitra, 2013; Qian et al., 2012), we wanted to investigate whether
Drp1 loss causes any of the cell cycle aberrations seen upon CycE
overexpression. CycE overexpression advances S phase entry,
consequently reducing G1 length, but slows S phase progression
after entry, consequently increasing S phase length (Hwang and
Clurman, 2005). We used four different cell cycle analysis
approaches to compare various aspects of cell cycle status between
the WT and the DRP1-KO MEFs (Fig. 4A). Flow cytometry
analyses of DNA content indicated that the DRP1-KO MEFs have
∼15% more cells in S phase in comparison to the WT MEFs (Fig.
S3B). This observation was confirmed by quantifying the proportion
of cells with PCNA–GFP in DNA replication foci, which is seen in
the S phase, and the proportion of cells with diffused nuclear
PCNA–GFP, which is seen in cells in any other cell cycle phase
(Leonhardt et al., 2000) (Fig. 4B). Monitoring the alteration in
PCNA–GFP foci distribution as the cells progress through S phase
(Leonhardt et al., 2000) (Fig. S3C, right panel) revealed higher
numbers of DRP1-KO MEFs only in the early S phase (Fig. 4B). We
did not observe any difference in the G1 phase using a flow
cytometry assay (Fig. S3B). However, with a FUCCI cell cycle
reporter, which is based on Cdt-1- and Geminin-degron-tagged
fluorescent proteins marking G1 as red, G1-S as yellow and S, G2
and M as green (Sakaue-Sawano et al., 2008), we found that the
number of cells in G1 or G1-S was 50% or lower in the DRP1-KO
MEFs compared to the WT MEFs, suggesting progression through
G1 is different in DRP1-KO MEFs (Fig. 4C). We ignored the green
population to avoid the ambiguity between the S, G2 and M status of
these cells. We also used Aurora-B–GFP to identify the mid-body
proper or remnants to be able to quantify cells having undergone
cytokinesis to commence G1 (Fig. S3C, left panel) and found that
DRP1-KO MEFs had 22% fewer cells in early G1 when compared to
WT. Therefore, the combination of the various cell cycle analyses
suggest that loss of Drp1 causes cells to spend longer time in early
S phase, likely due to early entry into S phase.
Stem cells characteristically have constitutively enhanced cyclin
E activity (Orford and Scadden, 2008), a long S phase and short G1
phase, which is reversed during the process of differentiation (Singh
and Dalton, 2009). Given that DRP1-KO MEFs have the above
properties, we compared the expression of the stem cell marker Sox2 between the WT and the DRP1-KO MEFs. Intriguingly, we found
that the DRP1-KO MEFs had higher levels of Sox-2 (Fig. 4D,E),
which showed a trend towards a significant reduction within 24 h of
re-introduction of Drp1 (Fig. S4A). We reasoned that if the higher
expression of Sox-2 in the DRP1-KO MEFs were dictated by the
altered cell cycle status, which has been proposed to be a factor for
4175
Journal of Cell Science
Fig. 3. Drp1 modulates the mtCycE-pool, likely through bioenergetics control. (A) CycE and Tom-20 immunocytochemistry of oligomycin-treated MEFs
showing that there was loss of mtCycE from the tubular mitochondria, but that it was retained in swollen mitochondrial structures in DRP1-KO MEFs treated with
oligomycin (arrows). (B) Quantification of the Pearson’s colocalization coefficient (R) between CycE and Tom-20 showing increased mtCycE in DRP1-KO MEFs,
whereas treatment with oligomycin (10 µg/ml) or CCCP (10 µM) lowered the level of the mtCycE pool both in WT and DRP1-KO MEFs. (C) Assessment of the
oxygen consumption rate (OCR) by employing extracellullar flux analysis showing comparable basal OCR between WT and DRP1-KO cells and elevated
maximal respiratory capacity (stimulated by CCCP induced depolarization) in DRP1-KO MEFs. A CCCP dose–response curve is shown in Fig. S3A.
(D) Comparison of spare respiratory capacity reflected by the fold increase in OCR with CCCP, in the presence or absence of serum in the medium showing that
the elevated spare respiratory capacity of the DRP1-KO cells is serum dependent. Results are mean±s.e.m. (n>30 for B, and n=4 for C and D). *,#P≤0.05
(Student’s t-test). Scale bar: 10 µm.
RESEARCH ARTICLE
Journal of Cell Science (2015) 128, 4171-4182 doi:10.1242/jcs.172429
maintaining stemness (Singh and Dalton, 2009), loss of Drp1 in
other cells would cause an increase in other relevant stem or
progenitor cell markers. We chose to downregulate Drp1 in an
ovarian cancer cell line (A2370) and a glioblastoma cell line
(U87MG) using Drp1-specific short hairpin RNAs (shRNAs).
Using two distinct shRNAs against Drp1, we indeed saw an increase
in the level of the well-studied stem and progenitor cell marker
Aldh1 in the ovarian cancer cell line (Flesken-Nikitin et al., 2013;
Landen et al., 2010) (Fig. 4F) and of Sox-2 in the glioblastoma cell
line (Ying et al., 2011; Eyler et al., 2011) (Fig. S4B). This data is
consistent with the report that reduction of Drp1 activity prevents
cell differentiation in embryonic stem cells (Todd et al., 2010).
However, in Drosophila, the Drp1-null adult follicle cell stem cells
can successfully differentiate into follicle cells (expressing the
lineage specific marker, FasIII) that fail to undergo terminal
differentiation in certain signaling contexts (Mitra et al., 2012).
Our data demonstrate that loss of Drp1, which leads to CycE
deregulation due to aberrant elevation of the mtCycE pool, leads to
alterations in the cell cycle that might promote the expression of
stem and progenitor cell markers.
Deregulation of CycE-driven by Drp1 loss attenuates cell
proliferation in low-cell-density environments and supports
aberrant cell proliferation in high-cell-density environments
Given that Drp1 loss leads to CycE deregulation and stem cell
marker enrichment, both of which can modulate the rate of
cell proliferation, we performed detailed investigations of the
cell proliferation status of the DRP1-KO MEFs. A growth curve
analysis revealed that, although the WT MEFs slowed down
proliferation at a high cell density, the DRP1-KO MEFs maintained
4176
cell proliferation in high-cell-density cultures (Fig. 5A).
Furthermore, comparison of colony forming assays revealed that
the DRP1-KO MEFs proliferated slower than the WT when there
were fewer cells in the colonies (low density), whereas they formed
bigger colonies when the colonies grew larger with higher cell
numbers (high density) (Figs 5B and 6C,D). Interestingly, the
DRP1-KO MEFs had an inner core with a surrounding lightly
stained region of cells in comparison to a uniform colony in the WT
(Fig. 5B), indicating a possible alteration of cell properties during
the growth of the DRP1-KO MEF colonies. The DRP1-KO MEFs
grew by more than tenfold, from low to high cell densities, in
comparison to the twofold change for the WT MEFs (Fig. 5C). In
addition, the DRP1-KO MEFs proliferated further with addition of
fresh medium whereas the WT MEFs remained contact inhibited in
high-cell-density cultures (Fig. 5C), confirming the loss of contact
inhibition in the DRP1-KO MEFs. Flow-cytometry-based cell cycle
analyses indicated that distribution of DRP1-KO population in G1
and S phases (as compared to the WT) showed the opposite trend
when seeded in low versus high cell densities (Fig. S3E) (cell
numbers are given in the Materials and Methods).
We reasoned that the mitotic stage of the Drosophila follicle cell
layer development, where the WT follicle cells are mitotically
active, might be similar to the low-cell-density environment of the
in vitro MEF culture. Supporting this notion, we found that Drp1null mitotic follicle cells in Drp1-null clones underwent fewer
divisions than the WT follicle cells (Fig. 5E). In addition, 27% of
the mitotic DRP1-null cells had detectable Geminin expression
(which marks S, G2 and M cells) in comparison to only 17% of the
WT background cells. This indicates that cell cycle aberration might
underlie the slower proliferation of the Drp1-null mitotic follicle
Journal of Cell Science
Fig. 4. Effect of Drp1 loss on cell cycle regulation and stem cell marker expression. (A) Schematic representation of approaches used for cell cycle analyses:
flow cytometry of DNA content (Fl. Cyto); Aurora B in mid-body (ArB); FUCCI reporter expression (FUCCI), PCNA distribution (PCNA Dist.). (B) Quantification of
the percentage of cells in S phase based on nuclear PCNA–GFP distribution, showing that DRP1-KO MEFs have a higher early S phase cell population in
comparison to the WT. Results are mean±s.e.m. (n=3). (C) Quantification of cells in G1 and G1-S phase based on expression of FUCCI reporter showing that
DRP1-KO MEFs have fewer cells in G1 and G1-S in comparison to the WT population (expressed as percentage of WT, dashed line). Results are mean±s.e.m.
(n=3). (D) Immunoblot for the stem cell marker Sox-2 showing elevated Sox-2 levels in DRP1-KO MEFs. Actin levels serve as loading controls.
(E) Immunocytochemistry for Sox-2 confirming a greater number of DRP1-KO MEFs positive for Sox-2 in the nucleus. (F) Immunoblot for Drp1 and the stem cell
marker Aldh1 in A2780 ovarian cancer cell line after expression of control (Cnt) or two distinct Drp1 shRNAs, demonstrating that reduction of Drp1 leads to
increased expression of Aldh1. Actin levels serve as controls. *P≤0.05 (Student’s t-test). Scale bar: 50 µm.
RESEARCH ARTICLE
Journal of Cell Science (2015) 128, 4171-4182 doi:10.1242/jcs.172429
cells, which was possibly reflected in our previous observation of
Drp1-null mitotic clones having higher phospho-histone 3 levels
and elevated BrdU incorporation (Mitra et al., 2012). The high-celldensity environment of the contact-inhibited WT MEFs and
aberrantly proliferating DRP1-KO MEFs might be reflected in the
post-mitotic differentiated Drosophila follicle cell monolayer layer
where the WT follicle cells have exited mitosis and Drp1-null clones
aberrantly proliferate by employing local EGFR signaling in the
PFC region; Drp1-null MBCs lacking EGFR activation can exit the
mitotic cycle and enter differentiation successfully (Mitra et al.,
2012; Fig. 5D). As expected, the Drp1-null PFCs continue to
express Geminin, signifying mitotically cycling cells (Fig. 5F).
Taken together, our data suggest that Drp1 loss attenuates the cell
proliferation rate in low-cell-density environments and certain
signals in the high-cell-density environments might switch the cell
properties to allow them to escape contact inhibition of cell
proliferation.
mtCycE is released by enhanced MEK–ERK signaling in the
absence of Drp1 to maintain aberrant cell proliferation in
high-cell-density environments
Drp1-null PFCs in the Drosophila follicle cell layer have elevated
levels of phosphorylated ERK [known as Rolled in Drosophila, and
ERK1/2 (MAPK3 and MAPK1) in mammals] (Fig. S4D), possibly
due to activation of the downstream components of the EGFR
signaling pathway (Yarden and Shilo, 2007). The DRP1-KO MEFs
also showed higher levels of phosphorylated ERK ( p-ERK) in
comparison to the WT MEFs, but only when seeded at high cell
densities (Fig. 6A), further highlighting the similarity between the
effects of Drp1 loss in MEFs at high cell density and in Drosophila
PFCs. In the high-cell-density DRP1-KO MEF population, p-ERK
was markedly enhanced in a subset of cells and was mostly
intracellular, as opposed to being at the plasma membrane as in the
WT-MEFs (Fig. 6B).
ERK is phosphorylated by MEK proteins (also known as
MAPK2K proteins) and the phosphorylation is abrogated with
MEK inhibitors (Akinleye et al., 2013). We chose the widely used
MEK inhibitor PD98059 to investigate the effect of abrogation of
ERK phosphorylation on cell proliferation in the presence or
absence of Drp1. We first confirmed that PD98059 lowers the pERK signal both in WT and DRP1-KO MEFs (Fig. S4E). The
DRP1-KO MEFs were increasingly sensitive to MEK inhibition
within a dose range of 0 to 10 μM PD98059 (Fig. 6C) compared to
the WT MEFs in a colony-forming assay. Quantification of the
Crystal Violet staining (Fig. 6D) and colony diameter (Fig. 6E) after
treatment with 5 μM PD98059 shows that PD98059 reduced cell
4177
Journal of Cell Science
Fig. 5. Effect of Drp1 loss on cell proliferation. (A) Growth curve analysis showing that DRP1-KO cells proliferate at high cell densities when WT cell
proliferation is attenuated. Results are mean±s.e.m. (n=3). (B) Colony assay of WT and DRP1-KO MEFs at low (7 days) and high cell densities (10 days).
Representative images of Crystal-Violet-stained colonies depicting cell densities (insets, magnified in Fig. S3D) and high-cell-density colony morphology (to the
right) are also shown. (C) Quantification of the colony assay from B normalized to low-cell-density values (dashed line) showing the massive increase in DRP1-KO
proliferation from low to high cell density, which is further increased with fresh medium. Results are mean±s.e.m. (n=3). (D) Cartoon depicting cell proliferation
properties of Drp1-null clones introduced in the Drosophila follicle cell layer, which develops through a mitotic phase that transitions to the post-mitotic
differentiated phase. Drp1-null PFCs maintain aberrant cell proliferation and form multilayers as we previously reported (Mitra et al., 2012). (E) Quantification of
the number of cell divisions in Drp1-null or control WT clones (each point) introduced in the Drosophila follicle cell layer showing that Drp1-null clones undergo
fewer cell divisions in the mitotic stage of development. (F) Drp1-null clones in the Drosophila follicle cell layer identified by loss of UbiGFP showing that Drp1-null
PFC clone expresses Geminin, signifying active mitosis, whereas Drp1-null MBC clones (*) do not express Geminin, signifying a post-mitotic differentiated status
similar to the WT UbiGFP-positive follicle cells. White lines demarcate the clonal boundaries. *P≤0.05 (Student’s t-test). Scale bar: 10 µm.
RESEARCH ARTICLE
Journal of Cell Science (2015) 128, 4171-4182 doi:10.1242/jcs.172429
proliferation of the DRP1-KO cells in high cell densities, whereas
the WT cells were affected mostly at low cell densities. Consistent
with this, PD98059 treatment also reduced the Sox-2 signal in the
DRP1-KO cells at high cell densities (Fig. S4C). Next, we tested
whether MEK inhibition with PD98059 would attenuate the
aberrant cell proliferation of the p-ERK-positive Drp1-null PFCs
in the differentiated follicle cell layer in Drosophila. We found that
the presence of 100 μM PD98059 in the food rescued the aberrant
cell proliferation in 37% (n=41) of Drp1-null PFC clones as they
formed a single layer of differentiated cells in comparison to 13%
(n=41) with the DMSO control (Fig. 6G). Therefore, the data
suggest that enhanced MEK–ERK signaling in the absence of Drp1
allows the cells to escape contact inhibition to maintain aberrant cell
proliferation in high-cell-density environments both in vitro and in
vivo.
Next, we investigated how the mtCycE pool is linked to the cell
proliferation status in the presence and absence of Drp1. Our data
thus far show that DRP1-KO MEFs with higher mtCycE proliferate
slower than the WT MEFs at low cell densities. Interestingly, we
noted that DmCycE did not colocalize with the mitochondrial
marker in the aberrantly proliferating Drp1-null PFCs in
Drosophila (Fig. S2C). Therefore, we hypothesized that the
increase in the mtCycE pool in the absence of Drp1 is lowered or
released by the elevated ERK signaling at high cell densities to
support aberrant cell proliferation. To test this hypothesis, we
investigated the effects of cell density and MEK inhibition on
mtCycE in growing colonies. The cells in the periphery of the
growing colonies do not contact each other and are in a low-celldensity environment, whereas the cells within the colony are in a
4178
high-cell-density contact-inhibited environment (Gumbiner and
Kim, 2014). In the WT MEFs the mtCycE pool was reduced in the
contact-inhibited (high cell density) compared to proliferating state
(low cell density) (Fig. 6F). Moreover, PD98059 reduced the
mtCycE pool of the WT cells in low cell densities as it reduced cell
proliferation. As expected, PD98059 did not affect the mtCycE
pool of the contact-inhibited WT MEFs as it did not affect their
proliferation. Therefore, we conclude that mtCycE in the WT cells
is positively linked to their cell proliferation status. However, the
elevated mtCycE pool in the DRP1-KO MEFs was associated with
lower cell proliferation at low cell densities and the mtCycE pool
was lowered to the levels of the WT cells at the high cell densities
where the DRP1-KO cells proliferated aberrantly (Fig. 6F).
Furthermore, PD98059 prevented the loss of mtCycE in the
DRP1-KO MEFs at high cell densities as it attenuated their
proliferation (Fig. 6F). PD98059, as expected, did not affect the
mtCycE levels of the DRP1-KO cells in low cell densities as it did
not affect their proliferation. This suggests that aberrantly elevated
mtCycE in the DRP1-KO MEFs is inhibitory towards cell
proliferation in low-cell-density environments and is released in
high-cell-density environments in a MEK–ERK-dependent manner
to allow aberrant cell proliferation.
DISCUSSION
Here, we provide the first evidence of a direct mitochondrial control
of CycE mediated by its recruitment to the organelle. Our findings
are based on the comparison of CycE properties in the presence or
absence of the mitochondrial fission protein Drp1 in MEFs (in vitro)
and the Drosophila follicle cell layer (in vivo). Our data support a
Journal of Cell Science
Fig. 6. Effect of MEK–ERK signaling and cell density on mtCycE and cell proliferation in the presence and absence of Drp1. (A) Immunoblot showing
elevated p-ERK levels in DRP1-KO MEFs only at high cell densities as compared to the WT. Total ERK (t-ERK) remains mostly unchanged.
(B) Immunocytochemistry for p-ERK in MEFs at high cell densities showing elevated intracellular p-ERK in a subset of DRP1-KO MEFs as opposed to p-ERK in
the cell periphery (likely cell membrane) in WT (high-magnification images of the boxed regions are shown on the right). (C) Quantification of the colony assay in
the presence of the MEK inhibitor PD98059 (PD) showing greater inhibition of DRP1-KO cell proliferation in a dose-dependent manner compared to that of WT.
(D) Quantification of colony assays in the presence or absence of PD98059 at low (7 days) and high cell densities (10 days) showing that PD98059 affects
WT MEFs at low cell densities and DRP1-KO MEFs only at high cell densities; see Fig. S3D for representative cell densities. (E) Quantification of colony diameters
from colony assays demonstrating PD98059 does not significantly affect WT or DRP1-KO MEFs at low cell densities but affects DRP1-KO MEFs at high cell
densities. (F) Quantification of the effect of PD98059 and cell density on the fraction of CycE colocalized with Tom-20 showing that mtCycE is lowered at high
cell densities both in WT and DRP1-KO MEFs, and that PD98059 lowers the mtCycE pool in WT MEFs at low cell densities but prevents loss of mtCycE in
DRP1-KO MEFs at high cell densities. Results in D–F are mean±s.e.m. (n=3 for D,E; n>30 for F). (G) Effect of PD98059 on cell proliferation of the Drp1-null PFCs
in the Drosophila follicle cell (FC) layer as identified by lack of UbiGFP. In the presence of DMSO, 87% of clones showed the multilayer arrangement of aberrantly
proliferating Drp1-null PFCs with only 13% showing lack of this phenotype; in the presence of 100 µM PD98059 37% of the Drp1-null clones lacked the phenotype
(n=41). White lines demarcate the clonal boundaries. *,#P≤0.05 (Student’s t-test). Scale bars: 5 µm.
model whereby a certain level of mtCycE, lacking the N-terminal
phospho-degron (T62) and maintained by Drp1, likely through
modulation of mitochondrial bioenergetics, is positively linked to
cell proliferation, with deregulation of such an mtCycE pool
deregulating cell proliferation (Fig. 7). We speculate that the
mtCycE pool might be under local bioenergetic control at the
contact sites between mitochondria and endoplasmic reticulum as
crosstalk between these organelles modulates mitochondrial
bioenergetics (Cárdenas et al., 2010). Correlation of the mtCycE
pool with the non-mtCycE pool suggests that part of the total CycE
pool might get recruited to the mitochondria in a controllable
fashion. We propose that the mitochondrial recruitment of CycE
upon enhancement of the bioenergetic capability and its controlled
release upon lowering of mitochondrial bioenergetics might be part
of the normal life cycle of CycE. Our data suggest that
bioenergetics-driven mitochondrial docking of CycE might
prevent phosphorylation of the N-terminal degron and thus
prevent degradation of the molecule. Indeed, loss of
mitochondrial bioenergetics, which would release mtCycE, leads
to active degradation of the molecule (Mandal et al., 2005).
Therefore, we hypothesized that increasing the steady-state
mtCycE-pool would prevent degradation and deregulate CycE.
Indeed, loss of Drp1 boosts mitochondrial energetics and enhances
the potentially degradation-resistant mtCycE pool to maintain
elevated levels of the molecule throughout the cell cycle. Such
deregulation of CycE attenuates cell proliferation in low-celldensity environments but supports aberrant cell proliferation in
high-cell-density environments because the mtCycE pool
undergoes MEK–ERK-dependent release (Fig. 7).
It remains to be examined whether reduction of the excess
mtCycE pool is sufficient to allow aberrant cell proliferation in the
absence of Drp1 in high-cell-density environments or whether
release of the excess mtCycE in the absence of Drp1 actively
promotes S phase entry at high cell densities. Nonetheless, our
current model implies that certain differentiated states maintained
with reduced Drp1 would allow mitochondria to act as a reservoir of
inactive CycE that could be released by undue MEK–ERK
activation allowing aberrant cell proliferation and possibly
initiating tumorigenic events. This concept explains why the
Drosophila MBCs with elevated mtCycE (Fig. 1F) and reduced
Drp1 activity only differentiate in the absence of EGFR–MEK–
ERK signaling (Klusza and Deng, 2011; Mitra et al., 2012), which
would otherwise release the mtCycE pool to aberrantly maintain
cell proliferation and prevent differentiation. By contrast, the
Journal of Cell Science (2015) 128, 4171-4182 doi:10.1242/jcs.172429
Drosophila PFCs having lower mtCycE (Fig. 1F) and higher
Drp1 activity differentiate under the influence of EGFR–MEK–
ERK signaling (Klusza and Deng, 2011; Mitra et al., 2012).
MEK–ERK signaling might release mtCycE by altering
mitochondrial bioenergetics or by directly impacting upon the
mitochondrial recruitment of CycE as p-ERK can re-localize to
mitochondria (Rasola et al., 2010; Horbinski and Chu, 2005).
Recently, Ras-driven MEK–ERK signaling has been shown to
activate Drp1-dependent mitochondrial fission (Serasinghe et al.,
2015; Kashatus et al., 2015) in line with our previous finding that
EGFR signaling (mediated by downstream activation of MEK–
ERK signaling) promotes mitochondrial fission (Mitra et al., 2012).
We have further shown that loss of Drp1-driven mitochondrial
fission aberrantly enhances EGFR signaling to support aberrant cell
proliferation (Mitra et al., 2012). Consistent with this, our current
findings show that Drp1 loss maintains ERK-dependent cell
proliferation only at high cell densities. Activation of the Ras–
Raf–MEK–ERK pathway brought about by oncogenic Ras causes
CycE-dependent cell transformation (Geng et al., 2003, 2007) only
when CycE is susceptible to degradation (Minella et al., 2005). Our
study, based on DRP1-KO MEFs that are immortalized (using SV40T antigens) (Ishihara et al., 2009) and a Drosophila model
system, predicts that lowering of Drp1 activity would prevent
oncogenic Ras-driven transformation at low cell densities because
Ras-driven MEK–ERK signaling would fail to release mtCycE for
degradation. Indeed, recent reports attempting to transform cells in
the absence of Drp1 (using combinations of transforming agents;
Kashatus et al., 2015; Serasinghe et al., 2015) have demonstrated
that Ras-driven transformation is attenuated when Drp1 is lowered.
However, Ras-driven MEK–ERK signaling in cells with reduced
Drp1 at high cell densities would release mtCycE to support
aberrant cell proliferation and possibly lead to cell transformation.
Our findings clearly point towards involvement of contact
inhibition pathways in modulating the interplay of Drp1-driven
mitochondrial fission and the EGFR–Ras–MEK–ERK pathway.
Cell–cell contact signals through various pathways impinge on the
Hippo pathway (McClatchey and Yap, 2012). Interestingly, Drp1
loss in the Drosophila follicle cell layer closely phenocopies loss of
Hippo signaling in the increase in CycE levels and maintenance of
aberrant cell proliferation in defined cell contexts (Meignin et al.,
2007; Mitra et al., 2012). Moreover, recent studies indicate that
various aspects of the Hippo pathway involve mitochondria (Sing
et al., 2014; Ohsawa et al., 2012; Del Re et al., 2014; Nagaraj et al.,
2012). Therefore, directed studies to investigate the interplay
Fig. 7. Schematic diagram depicting the proposed role of mtCycE. Our proposed model for the interplay between mitogenic and contact inhibition stimuli in
controlling the mtCycE pool to regulate cell proliferation, as revealed by the comparison of the data from the WT and DRP1-KO MEFs, as well as Drp1-null and WT
background cells in the Drosophila follicle cell (FC) layer.
4179
Journal of Cell Science
RESEARCH ARTICLE
RESEARCH ARTICLE
between Drp1 and components of EGFR–Ras–MEK–ERK and
Hippo pathways will facilitate understanding of the exact role of
mitochondrial fission in cell proliferation and tumorigenesis.
MATERIALS AND METHODS
Materials
Biochemicals were obtained from Fisher Biochemicals or Sigma. Other
materials used were as follows: oligomycin, CCCP, aphidicolin, nocodazole
and PD98059 (Sigma); Drp1 shRNA (Dharmacon); Premo FUCCI Cell
Cycle Sensor (Life Technologies); Flouromount G (SouthernBiotech);
FuGENE® HD Transfection Reagent (Promega); Vectashield (Vectorlabs);
Micro BCA Protein Assay Kit, RIPA buffer (Pierce); Luminata Forte
Western HRP substrate (Millipore); 4% paraformaldehyde aqueous solution
and Triton X-100 (FisherScientific); Labtek chambers (Nalgene Nunc
International); Millicell EZ slides (Millipore); DMEM (GIBCO); Graces
Medium w/L-Glut (Bioworld); Nutri-fly BF (Genesee Scientific);
Oligofectamine (Invitrogen); and CycE and Allstar siRNA (Qiagen).
Primary antibodies were against: 44/42 MAPK (ERK1/2) (western
blotting, 1:1000; immunofluorescence, 1:100) phosphorylated p44/42
MAPK (western blotting, 1:1000; immunofluorescence, 1:100) CycE1
(HE12) (western blotting, 1:1000; immunofluorescence 1:100) Cyclin A
(BF683) (1:300) Cyclin B1 (1:1000) p-CycE1(T62) (western blotting, 1:500;
immunofluorescence 1:100) and CDK2 (1:10000) (Cell Signaling); actin
AB-5 (1:10000; BD Biociences), Tom20 (western blotting, 1:10000;
immunofluorescence, 1:200) and DmCycE (d-300; 1:100) (Santa Cruz
Biotechnology), Sox2 (western blotting, 1:1000; immunofluorescence,
1:300; BD Bioscience), HSP-60 (1:200; BD Bioscience), ATP-B (1:100;
Abcam) and Geminin (1:100; a gift from Mary Lilly, Section of Gamete
Development, NICHD, Bethesda, USA). Secondary antibodies were from
Jackson ImmunoResearch Laboratories.
Cell culture, transfection and transduction
Cells used were immortalized WT and DRP1-KO MEFs (a gift from
Katsuyoshi Mihara, Dept. of Molecular Biology, Kyushu University,
Fukuoka, Japan) the U87MG glioblastoma cell line, cells isolated from the
GBM xenograft line D456MG, the A2780 ovarian cancer cell line and the
DU145 prostate cancer cell line. All cells were maintained in Dulbecco’s
modified Eagle’s medium (DMEM) with 10% fetal bovine serum (FBS) using
standard cell culture techniques. 1×106 and 1.8×105 MEFs were seeded per
well of a 12-well plate to obtain high and low cell densities, respectively. Cells
were harvested for immunoblot analyses 16 to 24 h post seeding.
Cells were transfected using FuGENE® with 500 ng of DNA per well of
an eight-well Labtek chambers and analyzed after 24–48 h, or using
Oligofectamine with 120 pmols of CycE siRNA and analyzed after 72 h.
For transduction of the FUCCI cell cycle probe, the manufacturer’s protocol
was followed after optimization. The transduction of the DRP1 shRNAs (5′AATAAGTTGGAGTAAAGTAGC-3′ and 5′-TTCAATAACCTCACAATCTCG-3′) or scrambled-shRNA-expressing lentiviral particles were
performed following standard methods and cells were harvested after 72 h.
Journal of Cell Science (2015) 128, 4171-4182 doi:10.1242/jcs.172429
mitochondrial fraction 2 (F2) as the supernatant. F1 and F2 were pooled to
obtain the total non-mitochondrial fraction. Protein estimation was
performed using a standard micro BCA kit.
For immunoblot analyses, asynchronous or synchronized cells (extracted
in Laemmli buffer), or mitochondrial and non-mitochondrial lysates
(extracted in RIPA buffer) were run on 12% polyacrylamide gels and
transferred onto PVDF membranes following standard techniques. After
blocking with fat-free milk, membranes were probed with relevant primary
antibodies followed by horseradish peroxidase (HRP)-tagged secondary
antibodies. The signal was detected using a chemiluminescence system.
Drosophila melanogaster clone generation
Previously published Drosophila stocks (Mitra et al., 2012) were
maintained and crosses performed at room temperature in vials containing
NutriFly food. Drp1-null or control FRT40A clones were generated by
crossing males of the genotype hsflp; ubiquitin nls GFP (UbiGFP) FRT40A/
CyO with virgin females of the genotype drp1KG03815 FRT40A/CyO or
FRT40A/CyO. The reciprocal cross with the drp1KG03815 FRT40A/CyO
was avoided as it yielded fewer progeny. The progenies selected against
CyO with genotype hsFLP; drp1KG03815FRT40A/ubiGFP FRT40A were
collected within 5 days of eclosion. The adult flies were heat pulsed in a
38°C water bath for 1 h to generate follicle cell clones and were maintained
in food with granulated yeast for the following 8 days and the ovaries were
dissected thereafter. For the experiment using PD98059, the heat-shocked
flies were maintained in food containing 100 µM PD98059 or DMSO
control for 8 days and the ovaries were dissected thereafter. To quantify the
clonal divisions, the total number of nuclei in each clone was counted from
confocal micrographs and expressed as Log (base 2) values.
Immunocytochemistry
Cells were seeded in eight-well Millicell chambers or glass-bottom Matek
dishes (for colony assay), fixed using fresh 4% paraformaldehyde and
permeabilized with freshly prepared 0.1% Triton X-100. Importantly, we
avoided any PBS wash before fixation as that decreased the mtCycE pool.
Fixed and permeabilized cells were blocked in 1% BSA in PBS (5% BSA
for CycE staining) followed by incubation with primary and subsequently
Cy3- or Cy5-tagged secondary antibodies with PBS washes after each
incubation. Finally, slide chambers were mounted with Fluoromount G with
Hoechst 33342 (10 µg/ml) to stain DNA and analyzed using confocal
microscopy.
Immunohistochemistry
Drp1-null mosaic Drosophila ovaries were isolated and immediately fixed
in fresh 4% paraformaldehyde and then dissected into ovarioles, which were
then permeabilized in PBS with Triton X-100 (0.5%). Only longer
permeabilization (30 min) allowed us to identify the mtCycE pool.
Permeabilizied ovarioles were blocked in 2% BSA in PBS with Triton
X-100 followed by incubation with primary antibodies and then conjugated
secondary antibodies with PBS Triton X-100 washes after each incubation.
Ovarioles were stained with Hoechst 33342 and mounted with Vectashield
and analyzed using confocal microscopy.
Cell synchronization and flow cytometry
Mitochondria isolation and immunoblotting
Mitochondria were isolated from MEFs by modifying a published protocol
(Frezza et al., 2007). Briefly, cells were lifted from the plate, centrifuged and
then homogenized. The homogenate was centrifuged at 600 g to obtain the
non-mitochondrial fraction 1 (F1). Supernatant, containing mitochondria,
was centrifuged at 10,000 g to obtain the mitochondrial pellet and the non-
4180
Colony assays
300 cells were seeded in each 35-mm tissue culture dish and allowed to grow
for 7 (low cell densities) to 10 days (high cell densities) without replenishing
the medium. PD98059 or DMSO was added in fresh medium after 24 h of
seeding as required. Colonies were stained using Crystal Violet solution
(0.25%) and quantified by measuring Crystal Violet absorbance on a BioTek Powerwave HT multiplate reader with Gen 2.05 software or by
measuring colony diameters using Carl Zeiss’s Zen software. For the
purpose of immunostaining, the growing colonies were replenished with
fresh medium 24 h before fixation to avoid the confounding influence of
nutrient or growth factor deprivation.
Microscopy
High-resolution fluorescence microscopy was performed using the LSM
700 laser scanning confocal microscope (Carl Zeiss). Optical slices were
Journal of Cell Science
Cells were synchronized in S phase using aphidicolin (2.5 µg/ml, 16 h) and
in the G2-M phase using nocodazole (1 µg/ml, 16 h). Propidium iodide
staining (50 µg/ml, 1 h) was performed following our previous protocol
(Mitra et al., 2009) and stained cells were analyzed by flow cytometry at the
Center for Aids Research Flow Core Facility, UAB using a Becton
Dickinson LSRII flowcytometer. Data were acquired using software Becton
Dickinson FACSDiva ver. 8.0 and analyzed using Becton Dickinson Verity
House ModFit software.
acquired with a 1.2 N.A. 40× Plan Neofluar oil objective using the
appropriate microscope settings. Image processing and colocalization
analyses were performed on 30 or more cells using the Zen Black
software (Carl Zeiss). Colocalization was expressed as a Pearson’s
coefficient (R) or as a fraction of the colocalized pool, as appropriate. For
regression analysis, the total colocalized and non-colocalized pools were
obtained by multiplying the mean colocalized or non-colocalized signal by
the number of pixels in each category. For quantification of signals of
p-ERK or Sox-2, more than 300 cells were included.
Oxygen consumption
Oxygen consumption rates (OCRs) were obtained using the Seahorse
Extracellular Flux Analyzer (Seahorse Biosceince). 60,000 cells were
seeded in each 24-well chamber and measurements obtained after 16–20 h.
Three readings were obtained from each well for basal or CCCP-stimulated
oxygen consumption. The optimal dose of CCCP (5 µM or 1 µM in the
presence or absence of serum, respectively) was used after proper titration
(Fig. S3A).
Statistical analyses
Student’s t-test was used for assessing statistical significance of the
difference in mean values. Linear regression analyses and its statistical
significance was performed using R (version 3.1.2). The R2 goodness of fit
was calculated by fitting the linear model of the data and the P-value was
obtained using Pearson’s correlation two-sided test.
Acknowledgements
We acknowledge K. Mihara for sharing the immortalized DRP1-KO and WT MEFs;
M. Lilly for sharing Geminin antibody; Kristin Spraggins and Onna Marie Baldwin for
help with data analyses; Marion Spell for the help in the Flowcytometry analyses;
Michael Sack, NHLBI, for allowing access to Seahorse extracellular flux analyzer
and Richa Rikhy for valuable discussions.
Competing interests
The authors declare no competing or financial interests.
Author contributions
K.M. designed experiments; D.P., A.I. and A.H. performed experiments; K.M.
performed Seahorse XF experiments; D.P., A.I., K.M., S.S., A.M. and M.K.B.
performed analyses; K.M. wrote the manuscript.
Funding
K.M. is supported by the National Institutes of Health [grant number R21ES025662].
Deposited in PMC for release after 12 months.
Supplementary information
Supplementary information available online at
http://jcs.biologists.org/lookup/suppl/doi:10.1242/jcs.172429/-/DC1
References
Akinleye, A., Furqan, M., Mukhi, N., Ravella, P. and Liu, D. (2013). MEK and the
inhibitors: from bench to bedside. J. Hematol. Oncol. 6, 27.
Bales, E., Mills, L., Milam, N., McGahren-Murray, M., Bandyopadhyay, D., Chen,
D., Reed, J. A., Timchenko, N., van den Oord, J. J., Bar-Eli, M. et al. (2005).
The low molecular weight CycE isoforms augment angiogenesis and metastasis
of human melanoma cells in vivo. Cancer Res. 65, 692-697.
Bedrosian, I., Lu, K. H., Verschraegen, C. and Keyomarsi, K. (2004). Cyclin E
deregulation alters the biologic properties of ovarian cancer cells. Oncogene 23,
2648-2657.
Cá rdenas, C., Miller, R. A., Smith, I., Bui, T., Molgó , J., Mü ller, M., Vais, H.,
Cheung, K.-H., Yang, J., Parker, I. et al. (2010). Essential regulation of cell
bioenergetics by constitutive InsP3 receptor Ca2+ transfer to mitochondria. Cell
142, 270-283.
Davis, R. J., Welcker, M. and Clurman, B. E. (2014). Tumor suppression by the
Fbw7 ubiquitin ligase: mechanisms and opportunities. Cancer Cell 26, 455-464.
Del Re, D. P., Matsuda, T., Zhai, P., Maejima, Y., Jain, M. R., Liu, T., Li, H., Hsu,
C.-P. and Sadoshima, J. (2014). Mst1 promotes cardiac myocyte apoptosis
through phosphorylation and inhibition of Bcl-xL. Mol. Cell 54, 639-650.
Donnellan, R. and Chetty, R. (1999). CycE in human cancers. FASEB J. 13,
773-780.
Eyler, C. E., Wu, Q., Yan, K., MacSwords, J. M., Chandler-Militello, D., Misuraca,
K. L., Lathia, J. D., Forrester, M. T., Lee, J., Stamler, J. S. et al. (2011). Glioma
Journal of Cell Science (2015) 128, 4171-4182 doi:10.1242/jcs.172429
stem cell proliferation and tumor growth are promoted by nitric oxide synthase-2.
Cell 146, 53-66.
Flesken-Nikitin, A., Hwang, C.-I., Cheng, C.-Y., Michurina, T. V., Enikolopov, G.
and Nikitin, A. Y. (2013). Ovarian surface epithelium at the junction area contains
a cancer-prone stem cell niche. Nature 495, 241-245.
Frezza, C., Cipolat, S. and Scorrano, L. (2007). Organelle isolation: functional
mitochondria from mouse liver, muscle and cultured fibroblasts. Nat. Protoc. 2,
287-295.
Galloway, C. A., Lee, H. and Yoon, Y. (2012). Mitochondrial morphology—
emerging role in bioenergetics. Free Radic. Biol. Med. 53, 2218-2228.
Geng, Y., Yu, Q., Sicinska, E., Das, M., Schneider, J. E., Bhattacharya, S.,
Rideout, W. M., Bronson, R. T., Gardner, H. and Sicinski, P. (2003). Cyclin E
ablation in the mouse. Cell 114, 431-443.
Geng, Y., Lee, Y.-M., Welcker, M., Swanger, J., Zagozdzon, A., Winer, J. D.,
Roberts, J. M., Kaldis, P., Clurman, B. E. and Sicinski, P. (2007). Kinaseindependent function of cyclin E. Mol. Cell 25, 127-139.
Gumbiner, B. M. and Kim, N.-G. (2014). The Hippo-YAP signaling pathway and
contact inhibition of growth. J. Cell Sci. 127, 709-717.
Horbinski, C. and Chu, C. T. (2005). Kinase signaling cascades in the
mitochondrion: a matter of life or death. Free Radic. Biol. Med. 38, 2-11.
Horn, S. R., Thomenius, M. J., Johnson, E. S., Freel, C. D., Wu, J. Q., Coloff,
J. L., Yang, C.-S., Tang, W., An, J., Ilkayeva, O. R. et al. (2011). Regulation of
mitochondrial morphology by APC/CCdh1-mediated control of Drp1 stability. Mol.
Biol. Cell 22, 1207-1216.
Hwang, H. C. and Clurman, B. E. (2005). Cyclin E in normal and neoplastic cell
cycles. Oncogene 24, 2776-2786.
Ishihara, N., Nomura, M., Jofuku, A., Kato, H., Suzuki, S. O., Masuda, K., Otera,
H., Nakanishi, Y., Nonaka, I., Goto, Y.-i. et al. (2009). Mitochondrial fission factor
Drp1 is essential for embryonic development and synapse formation in mice. Nat.
Cell Biol. 11, 958-966.
Kashatus, D. F., Lim, K.-H., Brady, D. C., Pershing, N. L. K., Cox, A. D. and
Counter, C. M. (2011). RALA and RALBP1 regulate mitochondrial fission at
mitosis. Nat. Cell Biol. 13, 1108-1115.
Kashatus, J. A., Nascimento, A., Myers, L. J., Sher, A., Byrne, F. L., Hoehn,
K. L., Counter, C. M. and Kashatus, D. F. (2015). Erk2 phosphorylation of Drp1
promotes mitochondrial fission and MAPK-driven tumor growth. Mol. Cell 57,
537-551.
Keck, J. M., Summers, M. K., Tedesco, D., Ekholm-Reed, S., Chuang, L.-C.,
Jackson, P. K. and Reed, S. I. (2007). Cyclin E overexpression impairs
progression through mitosis by inhibiting APCCdh1. J. Cell Biol. 178, 371-385.
Klusza, S. and Deng, W.-M. (2011). At the crossroads of differentiation and
proliferation: precise control of cell-cycle changes by multiple signaling pathways
in Drosophila follicle cells. Bioessays 33, 124-134.
Landen, C. N., Jr., Goodman, B., Katre, A. A., Steg, A. D., Nick, A. M., Stone,
R. L., Miller, L. D., Mejia, P. V., Jennings, N. B., Gershenson, D. M. et al. (2010).
Targeting aldehyde dehydrogenase cancer stem cells in ovarian cancer. Mol.
Cancer Ther. 9, 3186-3199.
Leonhardt, H., Rahn, H.-P., Weinzierl, P., Sporbert, A., Cremer, T., Zink, D. and
Cardoso, M. C. (2000). Dynamics of DNA replication factories in living cells.
J. Cell Biol. 149, 271-280.
Li, S., Xu, S., Roelofs, B. A., Boyman, L., Lederer, W. J., Sesaki, H. and
Karbowski, M. (2015). Transient assembly of F-actin on the outer mitochondrial
membrane contributes to mitochondrial fission. J. Cell Biol. 208, 109-123.
Lodish, H., Berk, A., Zipursky, S. L., Matsudaira, P., Baltimore, D. and Darnell, J.
(2000). Section 13.1, Overview of the Cell Cycle and Its Control. In: Molecular Cell
Biology, 4th edn. New York: W. H. Freeman.
Lu, A. and Pfeffer, S. R. (2013). Golgi-associated RhoBTB3 targets cyclin E for
ubiquitylation and promotes cell cycle progression. J. Cell Biol. 203, 233-250.
Mandal, S., Guptan, P., Owusu-Ansah, E. and Banerjee, U. (2005). Mitochondrial
regulation of cell cycle progression during development as revealed by the
tenured mutation in Drosophila. Dev. Cell 9, 843-854.
Mandal, S., Freije, W. A., Guptan, P. and Banerjee, U. (2010). Metabolic control of
G1-S transition: cyclin E degradation by p53-induced activation of the ubiquitinproteasome system. J. Cell Biol. 188, 473-479.
Matsumoto, Y. and Maller, J. L. (2004). A centrosomal localization signal in cyclin E
required for Cdk2-independent S phase entry. Science 306, 885-888.
McClatchey, A. I. and Yap, A. S. (2012). Contact inhibition (of proliferation) redux.
Curr. Opin. Cell Biol. 24, 685-694.
Meignin, C., Alvarez-Garcia, I., Davis, I. and Palacios, I. M. (2007). The salvadorwarts-hippo pathway is required for epithelial proliferation and axis specification in
Drosophila. Curr. Biol. 17, 1871-1878.
Minella, A. C., Welcker, M. and Clurman, B. E. (2005). Ras activity regulates cyclin
E degradation by the Fbw7 pathway. Proc. Natl. Acad. Sci. USA 102, 9649-9654.
Mitra, K. (2013). Mitochondrial fission-fusion as an emerging key regulator of cell
proliferation and differentiation. Bioessays 35, 955-964.
Mitra, K., Wunder, C., Roysam, B., Lin, G. and Lippincott-Schwartz, J. (2009).
A hyperfused mitochondrial state achieved at G1-S regulates cyclin E buildup and
entry into S phase. Proc. Natl. Acad. Sci. USA 106, 11960-11965.
4181
Journal of Cell Science
RESEARCH ARTICLE
RESEARCH ARTICLE
Sing, A., Tsatskis, Y., Fabian, L., Hester, I., Rosenfeld, R., Serricchio, M., Yau,
N., Bietenhader, M., Shanbhag, R., Jurisicova, A. et al. (2014). The atypical
cadherin fat directly regulates mitochondrial function and metabolic state. Cell
158, 1293-1308.
Singh, A. M. and Dalton, S. (2009). The cell cycle and Myc intersect with
mechanisms that regulate pluripotency and reprogramming. Cell Stem Cell 5,
141-149.
Siu, K. T., Rosner, M. R. and Minella, A. C. (2011). An integrated view of cyclin E
function and regulation. Cell Cycle 11, 57-64.
Spruck, C. H., Won, K.-A. and Reed, S. I. (1999). Deregulated cyclin E induces
chromosome instability. Nature 401, 297-300.
Taguchi, N., Ishihara, N., Jofuku, A., Oka, T. and Mihara, K. (2007). Mitotic
phosphorylation of dynamin-related GTPase Drp1 participates in mitochondrial
fission. J. Biol. Chem. 282, 11521-11529.
Todd, L. R., Damin, M. N., Gomathinayagam, R., Horn, S. R., Means, A. R. and
Sankar, U. (2010). Growth factor erv1-like modulates Drp1 to preserve
mitochondrial dynamics and function in mouse embryonic stem cells. Mol. Biol.
Cell 21, 1225-1236.
Welcker, M. and Clurman, B. (2005). Cell cycle: how cyclin E got its groove back.
Curr. Biol. 15, R810-R812.
Westermann, B. (2010). Mitochondrial fusion and fission in cell life and death. Nat.
Rev. Mol. Cell Biol. 11, 872-884.
Yarden, Y. and Shilo, B.-Z. (2007). SnapShot: EGFR signaling pathway. Cell 131,
1018.e1-1018.e2.
Ying, M., Wang, S., Sang, Y., Sun, P., Lal, B., Goodwin, C. R., Guerrero-Cazares,
H., Quinones-Hinojosa, A., Laterra, J. and Xia, S. (2011). Regulation of
glioblastoma stem cells by retinoic acid: role for Notch pathway inhibition.
Oncogene 30, 3454-3467.
Journal of Cell Science
Mitra, K., Rikhy, R., Lilly, M. and Lippincott-Schwartz, J. (2012). DRP1dependent mitochondrial fission initiates follicle cell differentiation during
Drosophila oogenesis. J. Cell Biol. 197, 487-497.
Nagaraj, R., Gururaja-Rao, S., Jones, K. T., Slattery, M., Negre, N., Braas, D.,
Christofk, H., White, K. P., Mann, R. and Banerjee, U. (2012). Control of
mitochondrial structure and function by the Yorkie/YAP oncogenic pathway.
Genes Dev. 26, 2027-2037.
Nicholls, D. G. and Ferguson, S. (2013). Bioenergetics, 4th edn. Academic Press.
Ohsawa, S., Sato, Y., Enomoto, M., Nakamura, M., Betsumiya, A. and Igaki, T.
(2012). Mitochondrial defect drives non-autonomous tumour progression through
Hippo signalling in Drosophila. Nature 490, 547-551.
Orford, K. W. and Scadden, D.T. (2008). Deconstructing stem cell self renewal:
genetic insights into cell cycle regulation. Nat. Rev. Gen. 9, 115-128.
Qian, W., Choi, S., Gibson, G. A., Watkins, S. C., Bakkenist, C. J. and Houten,
B. V. (2012). Mitochondrial hyperfusion induced by loss of the fission protein Drp1
causes ATM-dependent G2/M arrest and aneuploidy through DNA replication
stress. J. Cell Sci. 125, 5745-5757.
Rasola, A., Sciacovelli, M., Chiara, F., Pantic, B., Brusilow, W. S. and Bernardi,
P. (2010). Activation of mitochondrial ERK protects cancer cells from death
through inhibition of the permeability transition. Proc. Natl. Acad. Sci. USA 107,
726-731.
Sakaue-Sawano, A., Kurokawa, H., Morimura, T., Hanyu, A., Hama, H., Osawa,
H., Kashiwagi, S., Fukami, K., Miyata, T., Miyoshi, H. et al. (2008). Visualizing
spatiotemporal dynamics of multicellular cell-cycle progression. Cell 132,
487-498.
Serasinghe, M. N., Wieder, S. Y., Renault, T. T., Elkholi, R., Asciolla, J. J.,
Yao, J. L., Jabado, O., Hoehn, K., Kageyama, Y., Sesaki, H. et al. (2015).
Mitochondrial Division is requisite to RAS-induced transformation and targeted by
oncogenic MAPK pathway inhibitors. Mol. Cell 57, 521-536.
Journal of Cell Science (2015) 128, 4171-4182 doi:10.1242/jcs.172429
4182