Journal of Cell Science
Fission yeast nucleolar protein Dnt1 regulates G2/M transition and
cytokinesis by downregulating Wee1 kinase
Zhi-yong Yu, Meng-ting Zhang, Gao-yuan Wang, Dan Xu, Daniel Keifenheim, Alejandro Franco, Jose Cansado,
Hirohisa Masuda, Nick Rhind, Yamei Wang and Quan-wen Jin
There was an error published in J. Cell Sci. 126, 4995-5004.
One source of funding was accidentally omitted. The correct funding section is as shown below.
This work was supported by the National Institutes of Health [grant number GM-069957 to N.R.]; MEC European Regional
Development Fund co-funding from the EU [grant number BFU2011-22517 to J.C.]; the National Natural Science Foundation of China
[grant number 31171298 to Q.W.J.]; Key Project of the Chinese Ministry of Education [grant number 108076 to Q.W.J.]; the 111 Project
of Education of China [grant number B06016]. H.M. is supported by Cancer Research UK (grant to Takashi Toda, Laboratory of Cell
Regulation, Cancer Research UK London Research Institute, Lincoln’s Inn Fields, London, UK). Deposited in PMC for release after
We apologise to the readers for this error.
? 2014. Published by The Company of Biologists Ltd | Journal of Cell Science (2014) 127, 259 doi:10.1242/jcs.146225
Journal of Cell Science
Fission yeast nucleolar protein Dnt1 regulates G2/M
transition and cytokinesis by downregulating Wee1
Zhi-yong Yu1, Meng-ting Zhang1, Gao-yuan Wang1, Dan Xu1, Daniel Keifenheim2, Alejandro Franco3,
Jose Cansado3, Hirohisa Masuda4, Nick Rhind2, Yamei Wang1,* and Quan-wen Jin1,*
1State Key Laboratory of Cellular Stress Biology, School of Life Sciences, Xiamen University, Xiamen 361102, Fujian, China
2Department of Biochemistry and Molecular Pharmacology, University of Massachusetts Medical School, 364 Plantation St., Worcester, MA 01605,
3The Yeast Physiology Group, Department of Genetics and Microbiology, Facultad de Biologı ´a, Universidad de Murcia, 30071 Murcia, Spain
4Laboratory of Cell Regulation, Cancer Research UK London Research Institute, Lincoln’s Inn Fields Laboratories, 44 Lincoln’s Inn Fields, London
WC2A 3LY, UK
*Authors for correspondence (email@example.com; firstname.lastname@example.org)
Accepted 5 August 2013
Journal of Cell Science 126, 4995–5004
? 2013. Published by The Company of Biologists Ltd
Cytokinesis involves temporally and spatially coordinated action of the cell cycle, cytoskeletal and membrane systems to achieve
separation of daughter cells. The septation initiation network (SIN) and mitotic exit network (MEN) signaling pathways regulate
cytokinesis and mitotic exit in the yeasts Schizosaccharomyces pombe and Saccharomyces cerevisiae, respectively. Previously, we have
shown that in fission yeast, the nucleolar protein Dnt1 negatively regulates the SIN pathway in a manner that is independent of the
Cdc14-family phosphatase Clp1/Flp1, but how Dnt1 modulates this pathway has remained elusive. By contrast, it is clear that its
budding yeast relative, Net1/Cfi1, regulates the homologous MEN signaling pathway by sequestering Cdc14 phosphatase in the
nucleolus before mitotic exit. In this study, we show that dnt1+positively regulates G2/M transition during the cell cycle. By conducting
epistasis analyses to measure cell length at septation in double mutant (for dnt1 and genes involved in G2/M control) cells, we found a
link between dnt1+and wee1+. Furthermore, we showed that elevated protein levels of the mitotic inhibitor Wee1 kinase and the
corresponding attenuation in Cdk1 activity is responsible for the rescuing effect of dnt1D on SIN mutants. Finally, our data also suggest
that Dnt1 modulates Wee1 activity in parallel with SCF-mediated Wee1 degradation. Therefore, this study reveals an unexpected
missing link between the nucleolar protein Dnt1 and the SIN signaling pathway, which is mediated by the Cdk1 regulator Wee1 kinase.
Our findings also define a novel mode of regulation of Wee1 and Cdk1, which is important for integration of the signals controlling the
SIN pathway in fission yeast.
Key words: Fission yeast, Nucleolus, Dnt1, Wee1, G2/M transition, Cytokinesis
Eukaryotic cells use a highly conserved mechanism to control
entry into mitosis, which depends on the activation of cyclin-
dependent kinase (CDK), a key enzyme consisting of the protein
kinase Cdk1 and its regulatory subunit, cyclin B (Morgan, 1997;
Nurse, 1990). The highly activated Cdk1 drives entry into mitosis
by phosphorylating a wide range of targets (Lindqvist et al., 2009;
Ubersax et al., 2003). In the fission yeast Schizosaccharomyces
pombe, the machinery regulating entry into mitosis has been
thoroughly characterized (Nurse, 1994). The activity of the
fission yeast Cdk1 (encoded by the cdc2+gene) oscillates
throughout the cell cycle, peaking as cells enter mitosis (Morgan,
1997). During interphase, Cdk1 activity is held in check by the
inhibitory phosphorylation on conserved tyrosine 15 (Tyr15) in
the ATP binding domain by Mik1 and Wee1 kinases (Berry and
Gould, 1996; Lundgren et al., 1991; Russell and Nurse, 1987b),
and this modification interferes with the ability of Cdk1 to
transfer phosphate to the target substrate (Morgan, 1997). Upon
the G2/M transition and mitotic onset, a sharp reversal of this
phosphorylation event is triggered by the Cdc25 family of
phosphatases, which act as a major rate-limiting step in the
activation of Cdk1 (Millar et al., 1991; Nilsson and Hoffmann,
2000; Russell and Nurse, 1986).
Both Wee1 kinase and Cdc25 phosphatase are in turn tightly
regulated to provide an accurate control of the mitotic onset, and
a variety of factors and signaling pathways converge upon Wee1
and Cdc25 to govern the G2/M transition (Callegari and Kelly,
2007; Calonge et al., 2010; Kuntz and O’Connell, 2009; Lo ´pez-
Avile ´s et al., 2005; O’Connell et al., 2000; Suda et al., 2000).
response (SR), the cell-geometry sensing (CGS) and the
G2 DNA-damage-checkpoint pathways, all regulate Tyr15
phosphorylation through Wee1 and Cdc25 (Callegari and Kelly,
2007; Martin and Berthelot-Grosjean, 2009; Millar et al., 1995;
Moseley et al., 2009; Petersen and Nurse, 2007; Rhind et al.,
1997; Russell and Nurse, 1987a; Shiozaki et al., 1998). The SR
pathway connects the nutrient-responding TOR pathway with the
recruitment of Polo kinase (Plo1) to the spindle pole body (SPB)
Journal of Cell Science
and Cdk1 activation, and is responsible for nutritional modulation
of mitotic entry (Petersen and Hagan, 2005; Petersen and Nurse,
2007). The CGS pathway controls mitotic entry in response to
cell geometry through two related inhibitory kinases of Wee1,
Cdr1/Nim1 and Cdr2, and it involves a spatial gradient of the
protein kinase Pom1 along the long axis of the cell, coupling cell
length to G2/M transition (Hachet et al., 2011; Martin and
Berthelot-Grosjean, 2009; Moseley et al., 2009). Finally, the G2
DNA damage delays entry into mitosis primarily by inhibiting
Cdc25, but it also has a secondary effect through the activation of
Mik1 (Rhind et al., 1997; Rhind and Russell, 2001).
Wee1 has been shown to be destabilized during G2 and M phases
of the cell cycle in diverse organisms ranging from yeasts to humans,
and therefore degradation is thought to be a major mechanism
triggering the decrease in Wee1 levels as cells enter mitosis (Aligue
et al., 1997; Kellogg, 2003; McGowan and Russell, 1995; Sia et al.,
1998; Watanabe et al., 2005). It has been demonstrated that a
multicomponent E3 (ubiquitin ligase), the SCF (Skp1–Cdc53/Cullin-
1–F-box) complex, is responsible for Wee1 degradation at the onset
of M-phase in budding yeast, frog egg extracts and human somatic
2004). However, whether SCF-mediated Wee1 degradation is
conserved in fission yeast awaits direct evidence, although it has
been shown that skp1 mutants display cell elongation and G2-delay
phenotypes (Lehmann et al., 2004).
In fission yeast, a spindle pole body (SPB)-based regulatory
network, called the septation initiation network (SIN), triggers
exit from mitosis and cytokinesis. The SIN is homologous to
the mitotic exit network (MEN) in budding yeast, although the
two pathwayshave diverged
Balasubramanian et al., 2004; Gould and Simanis, 1997;
Guertin et al., 2002a; Krapp and Simanis, 2008). The SIN
signaling cascade is initiated by activation of the GTPase Spg1
and promoted by three protein kinases and their associated
subunits: Cdc7, Sid1-Cdc14 and Sid2-Mob1. Assembly of SIN
signaling components and their regulators occurs at the SPB on a
platform built by the SIN scaffolding components Cdc11 and
Sid4, which recruit all the members of the SIN to the SPBs. The
SIN controls the final stages of cell division, including
actomyosin ring contraction and formation of the division
septum. Loss-of-function mutations in sin genes result in
elongated multinucleate cells as a result of multiple rounds of
nuclear division, and cell growth in the absence of cell division.
SIN signaling needs to coordinate cytokinesis with completion
of chromosome segregation and is negatively regulated by Cdk1
activity. Previous studies have shown that the transition from a
symmetrical to an asymmetrical distribution of Cdc7 kinase on
the SPBs during anaphase, the initial association of Sid1 kinase
with only the new SPB at anaphase B, and maximum SIN
activation, are all dependent upon Cdk1 inhibition (Chang et al.,
2001; Dischinger et al., 2008; Guertin et al., 2000).
Previously, in a genetic screen for suppressors of the
cytokinesis checkpoint defect in SIN mutants, we identified the
nucleolar protein Dnt1 as an inhibitor of SIN signaling (Jin et al.,
2007). Although the amino acid sequence of Dnt1 shows weak
similarity to that of budding yeast nucleolar proteins Net1/Cfi1, it
seems that Dnt1 functions in a distinct way from Net1/Cfi1. Our
study revealed that, unlike Net1/Cfi1, which regulates the MEN
through theCdc14 phosphatase,
independently of Clp1/Flp1 (hereafter referred to as Clp1), the
fission yeast homologue of Cdc14 (Jin et al., 2007). However, a
infunction (reviewed by
detailed mechanism explaining how Dnt1 antagonizes the SIN
signaling in fission yeast remained unclear.
In this work, we show that S. pombe cells display a delay in
G2/M transition during the cell cycle in the absence of dnt1+, and
this delay correlated with persistent Tyr15phosphorylation on
Cdk1. Genetic and biochemical analyses revealed that the G2/M
transition defect in dnt1D cells is due to elevated activity and
protein level of the mitotic inhibitor Wee1 kinase, because
removal of Wee1 completely abolished the rescuing effect of
dnt1D on SIN mutants. Finally, our data also suggest that Dnt1
modulates Wee1 activity in parallel with SCF-mediated Wee1
degradation. Thus, in fission yeast Dnt1 negatively regulates SIN
signaling by modulating the protein levels of Wee1 and therefore
the activity of Cdk1. The above findings reveal the existence of a
novel mode of control of SIN by a nucleolar protein.
Dnt1 positively regulates G2/M transition in the cell cycle
While studying dnt1+for its role in cytokinesis, we made several
observations that indicate that dnt1+might also regulate the cell
cycle progression. First, the rescuing effect of dnt1D on
temperature-sensitive SIN mutants cdc14-118, sid2-250 and
actomyosin ring formation mutant cdc8-110 was sensitive to
protein level or activity of Cdc25, because the enhanced growth at
30˚C or 33˚C of double mutants dnt1D cdc14-118, dnt1D sid2-25
and dnt1D cdc8-110 compared with single mutants cdc14-118,
sid2-250 or cdc8-110, was abolished when higher levels of Cdc25
were induced by a gain-of-function version of cdc25+(i.e. nmt1-
cdc25-D1) (Daga and Jimenez,1999;Talladaetal.,2007)(Fig. 1A
and supplementary material Fig. S1). Second, dnt1D deletion was
synthetically sick with the temperature-sensitive mutant of
cdc25-22 (Fig. 1B). Third, dnt1D deletion was synthetic lethal
with cdc25D in a cdc2-3w genetic background, which is
largely insensitive to Cdc25 (Fantes, 1981). When cdc2-3w
cdc25D::ura4+cells were crossed with the dnt1D::kanRcdc2-3w
mutant, triple mutant spores germinated to form microcolonies
with elongated branched cells that stopped dividing after several
cell divisions (Fig. 1C). Fourth, we also noticed that dnt1D cells
divided at a significantly longer length than wild-type cells, and
this was also true for cdc25-22 dnt1D cells when compared with
cdc25-22 cells (Fig. 1D,E). In S. pombe, there is a direct
correlation between cell length at septation and the timing of
mitotic commitment, and a longer cell length at division indicates
mitotic delay (Mitchison and Nurse, 1985). Thus, the above data
suggest that Dnt1 mightbe a Cdc25-independentpositive regulator
of G2/M transition in fission yeast.
Next, we examined whether levels of Cdc2 itself and its
phosphorylation at Tyr15(pY15) were affected by the absence of
dnt1+. We found that whereas Cdc2 protein level remained
unchanged in dnt1D cells, Cdc2 phosphorylation at Tyr15was
significantly increased in these cells (Fig. 2A,B). To more
directly monitor the levels of Cdc2 phosphorylation at Tyr15
during the cell cycle, we performed a cdc25-22 block-and-release
assay. Cells synchronized in G2 by the temperature-sensitive
cdc25-22 mutation were released to permissive temperature to
allow mitotic entry. As shown in Fig. 2C, the kinetics of Cdc2
dephosphorylation at Tyr15in cdc25-22 dnt1D cells was notably
delayed compared with control cells. The maximum percentage
of binucleated (entering mitosis) and septated cells was also
reduced for an extended period in the cdc25-22 dnt1D mutant,
confirming a cell cycle defect at G2. Moreover, the duration of
Journal of Cell Science 126 (21) 4996
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Cdc2 Tyr15dephosphorylation during mitosis in dnt1D cells was
extended compared with control cells (Fig. 2C). Taken together,
these results suggest that, in fission yeast, Dnt1 positively
regulates both G2/M transition and mitotic exit during the cell
cycle by affecting the phosphorylation status of Cdc2 kinase.
Dnt1 regulation of the G2/M transition acts through Wee1
Our genetic analyses suggested that the cell cycle defect at G2/M
in dnt1D cells was not dependent on Cdc25. To dissect how Dnt1
Fig. 2. Dnt1 positively regulates G2/M transition during the cell cycle by
affecting the phosphorylation status of Cdc2 kinase. (A) The protein level
of Cdc2 is not altered in dnt1D cells. Protein samples were prepared from wild
type without GFP-Atb2, wild type with GFP-Atb2 and dnt1D GFP-Atb2
strains. Immunoblotting was performed with anti-PSTAIR (top) and anti-GFP
(bottom) antibodies to detect total Cdc2 and Atb2, respectively. The levels of
Cdc2 were normalized to those of GFP-Atb2 in each strain. n53; P.0.05.
(B) Phosphorylation of Cdc2 on Tyr15(Y15P) is increased in dnt1D cells.
Immunoblotting was performed with anti-phospho-Cdc2 (Tyr15) (top) and
anti-PSTAIR (bottom) antibodies to detect phosphorylated Cdc2 (Y15P) and
total Cdc2, respectively. The levels of phosphorylated Cdc2 (Y15P) were
normalized to those of total Cdc2 in each strain. n53; *P,0.05. (C) G2/M
transition and cell cycle progression is delayed in dnt1D cells. Wild-type
(cdc25-22) or dnt1D mutant (cdc25-22 dnt1D) cells were synchronized at
36˚C for 3.5 hours and then released from the growth arrest by transfer back
to 25˚C. Aliquots were taken at different time intervals and were either fixed
and stained with calcofluor and DAPI, or subjected to immunoblotting with
anti-phospho-Cdc2 (Tyr15) and anti-PSTAIR antibodies to detect
phosphorylated Cdc2 (Y15P) and total Cdc2, respectively (middle). The
percentages of binucleated and septated cells were counted for each time
point (top). The levels of phosphorylated Cdc2 (Y15P) were normalized to
those of total Cdc2 at each time point, with the relative ratio at time 0 set as
Fig. 1. Characterization of cell-cycle-progression defects in dnt1D cells.
(A) The suppressive effect of dnt1D on the temperature-sensitive SIN mutant cdc14-
118 is counteracted by increased levels of Cdc25 in a gain-of-function version of
cdc25+(nmt1-cdc25-D1). Serial dilutions (10-fold) of the indicated strains were
spotted on EMM with or without thiamine and incubated at the indicated
temperatures. (B) Negative genetic interaction between dnt1D and cdc25-22. Serial
dilutions (10-fold) of the indicated strains were spotted on YE plates and incubated
for 3–5 days at the indicated temperatures before being photographed. (C) dnt1D
deletion is synthetic lethal with cdc25D in a cdc2-3w genetic background. Normal-
looking four-spore asci obtained on sporulating ME plates for 2 days after a cross
between dnt1D::kanRcdc2-3w and cdc25D::ura4+cdc2-3w strains was dissected
using a micromanipulator. Colonies formed from germinated spores were replicated
on plates with YE plus G418 (selecting for dnt1D::kanR) and EMM without uracil
(selecting for cdc25D::ura4+) (left). Microcolonies corresponding to triple mutant
the right. (D) Live-cell images of calcofluor-stained wild-type, dnt1D, cdc25-22 and
dnt1D cdc25-22 cells grown in EMM at 25˚C. Scale bar: 10 mm. (E) Quantification
of cell lengths at cell division. Cells were grown in EMM at 25˚C and n.200 cells
average lengths of septated cells and standard deviation (s.d.) within the cell
population for each genotype strain. The numbers in each plot correspond to the
mean values of cell length. ***P,0.001.
Dnt1 in cell cycle control 4997
Journal of Cell Science
facilitates the G2/M progression, we conducted epistasis
experiments measuring the cell length at division of double
mutants for dnt1D and a series of genes involved in G2/M
transition control (Fig. 3B and supplementary material Fig. S2).
Mostly these double mutants either showed additive effect on cell
length or divided at an intermediate length. Notably, no increase
in cell size at division was observed in cdc2-1w dnt1D, wee1-50
dnt1D and wee1D dnt1D double mutant cells compared with the
cdc2-1w, wee1-50 and wee1D single mutants, respectively
(Fig. 3A,B). Cell length measurements also revealed that the
longer size at division in the dnt1D mutant was not dependent on
enhanced activity of Clp1 phosphatase or Pom1 kinase, which
have been shown to be involved in negatively regulating Cdc25
and Cdr1/2 activity respectively (supplementary material Fig. S2)
(Martin and Berthelot-Grosjean, 2009; Moseley and Nurse, 2009;
Wolfe and Gould, 2004; Wolfe et al., 2006). These findings
suggest that Wee1 is required for Dnt1-mediated cell cycle
control during mitotic commitment.
Previous studies have shown that in fission yeast, G2 DNA-
damage-checkpoint activation causes G2/M delay and sustained
Wee1-mediated Tyr15phosphorylation on Cdc2 until completion
of repair (O’Connell et al., 1997; Rhind et al., 1997). Moreover,
deletion of the ribosome-associated RACK1 (receptor of
activated C kinase) ortholog Cpc2 leads to a Wee1-dependent
cell cycle delay at the G2/M transition (Nu ´n ˜ez et al., 2010). To
investigate the possible functional connection between Dnt1 and
the function of either the G2 DNA-damage checkpoint or Cpc2,
we examined the cell length of double mutants between dnt1D
and G2 DNA-damage-checkpoint mutants (rad3D, chk1D, cds1D
and tel1D) and cpc2D. As shown in supplementary material Fig.
S3A, all double mutants divided at an intermediate cell length,
suggesting that Dnt1 does not regulate these pathways.
Consistent with these observations, we found that G2 DNA-
damage-checkpoint mutants did not counteract the suppressive
effect of dnt1D on SIN mutant cdc14-118. Moreover, cpc2D
further elongated the length of dnt1D cells and enhanced the
rescue of the cdc14-118 mutant by dnt1D, showing an additive
effect (supplementary material Fig. S3B,C, and data not shown).
Recently, we have shown that during metaphase Dnt1 interacts
with and inhibits Dma1 (Wang et al., 2012). Previous studies
have shown that Dma1 localizes to the SPB through interaction
with the SIN scaffold protein Sid4 and it ubiquitylates Sid4
to prevent recruitment of the Polo-like kinase and the SIN
activator Plo1 to SPBs during a mitotic checkpoint arrest
(Guertin et al., 2002b; Johnson and Gould, 2011). It is also
known that the SPB component Cut12 is functionally linked with
Fig. 3. Dnt1 controls G2/M transition by negatively regulating Wee1 protein levels. (A) Live-cell images of calcofluor-stained wee1-50, wee1-50 dnt1D, wee1D
and wee1D dnt1D cells. Cells were first grown in YE at 25˚C and then shifted to 37˚C for 4 hours (for wee1-50 and wee1-50 dnt1D) or grown in EMM at 25˚C
(for wee1D and wee1D dnt1D) before calcofluor staining and photography. Scale bar: 10 mm. (B) Quantification of cell lengths at cell division. Cells were grown in
either YE at 25˚C and then shifted to 37˚C for 4 hours (for wee1-50 and wee1-50 dnt1D) or grown in EMM at 25˚C (for wee1D, wee1D dnt1D, cdc2-1w and cdc2-1w
dnt1D).n.200cellswith septawere measuredfor each strain. (C,D)Wee1protein levelsareelevatedindnt1D cells.(C)Wild-type strains without orwith3HA-6His-
to detect Wee1. Cdc2 was used as loading control. (D) Wee1 luciferase activity was analyzed using Wee1-R.luciferase and Ade4-F.luciferase assay system in wild-
type and dnt1D cells. The schematic depiction of Wee1-R.luciferase and Ade4-F.luciferase constructs is shown (top). Cells were collected from cultures grown in YE
at 30˚C. Wee1-R.luciferase and Ade4-F.luciferase activity was measured and the relative luminescence was quantified with ratio between Wee1-R.luciferase and
Ade4-F.luciferase activity in wild-type cells being set as 1.00 (bottom). n53; **P,0.01. (E) wee1+mRNA level is unchanged in dnt1D cells. Total RNA was
extracted from wild-type and dnt1D cells and subject toRT-PCR. PCR products wereresolvedin PAGE gels andthen silverstained (top). Thelevelsof wee1+mRNA
were normalized to those of act1+(internal control) and the relative ratio in wild-type cells was set as 1.00 (bottom). n53; P.0.05.
Journal of Cell Science 126 (21) 4998
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Plo1. The gain-of-function mutant cut12-S11 promotes Plo1
recruitment to the G2 phase SPB, boosts global Plo1 activity and
drives advanced mitotic commitment (Bridge et al., 1998;
Grallert et al., 2013). To examine whether Cut12 or Dma1-
regulated Plo1 is involved in Dnt1 modulation of G2/M
transition, we first measured the cell length of double mutants
between dnt1D and cut12-S11 or dma1D, and found that neither
of these mutants could reverse the longer size at division of
dnt1D cells (supplementary material Fig. S4A). Moreover,
neither cut12-S11 nor dma1D could rescue the synthetic
lethality of dnt1D cdc25-22 double mutant (supplementary
material Fig. S4B). Therefore, the G2/M transition defects in
dnt1D cells are not linked to Cut12- or Dma1-controlled Plo1.
Wee1 protein is upregulated in dnt1D cells
Our epistasis analyses suggested that the G2/M delay in dnt1D
cells most likely acts through Wee1 to promote maintenance of
Cdc2 Tyr15phosphorylation. This Cdc2 inhibitory effect could be
exerted by increasing Wee1 activity and/or Wee1 abundance. To
gain insight into the mechanism by which Wee1 inhibits Cdc2
when dnt1+is absent, we determined Wee1 protein levels in the
wild-type and dnt1D strains expressing a genomic version of
HA3His6-Wee1 (Raleigh and O’Connell, 2000). Although we
occasionally observed an increase of HA3His6-Wee1 levels in
dnt1D cells compared with wild-type cells (Fig. 3C), we
experienced some technical difficulty with the reproducibility
of these assays. Therefore, we turned to a newly developed
strategy to monitor protein levels by tagging Wee1 with
luciferase from thesea pansy
R.luciferase) (Matthews et al., 1977), while using firefly-
control (see Fig. 3D, top panel). By using this strategy we
found a reproducibly significant increase of Wee1 protein levels
in dnt1D cells compared with wild-type cells (Fig. 3D, bottom
panel). Therefore, Dnt1 negatively affects the abundance of
Wee1 kinase, which is a negative regulator of Cdk1.
To understand how Wee1 protein is affected by Dnt1, we
examined the mRNA levels of the wee1+gene in dnt1D cells by
RT-PCR. As shown in Fig. 3E, our results showed that the
increase in Wee1 protein levels observed in dnt1D cells did not
result from enhanced expression of wee1+mRNA. Thus, it is
possible that Dnt1 might influence the stability and/or turnover of
Wee1 protein. The observation that Wee1 is upregulated in dnt1D
cells led us to explore the possibility that protein levels of various
Cdk1 regulators, such as Cdc25, Cdc13 (cyclin B), Pom1, Cdr1
or Cdr2 might be altered in this mutant. However, none of these
proteins was affected by deletion of dnt1+(supplementary
material Fig. S5 and data not shown).
as an internal
Elevated Wee1 levels are responsible for the rescue of
compromised SIN mutants by dnt1D
We addressed the functional significance of the negative
regulation of Wee1 by Dnt1 during activation of SIN signaling
by exploring how wee1+affects the rescue of SIN mutants by
dnt1D. Maximal SIN activation depends upon Cdk1 inhibition
and thus SIN mutants are very sensitive to high Cdk1 activity.
Thus, it is plausible that the low Cdk1 activity promoted by
elevated Wee1 levels in dnt1D cells might be sufficient to
maintain the SIN signaling activity to certain degree. To test this
possibility, we first examined whether elevated Wee1 levels
alone could rescue SIN mutants. Intriguingly, we found that a
slight elevation of Wee1 levels by using a P81nmt1-GFP-Wee1
construct rescued the SIN mutant cdc14-118 to a similar extent as
dnt1D (Fig. 4A). Next, we introduced the wee1D mutation into
single SIN mutants (cdc14-118 and sid2-250) or double mutants
with dnt1D (dnt1D cdc14-118 and dnt1D sid2-250), and
monitored either growth at different temperatures or the
accumulation of multinucleated cells after a shift to 37˚C in
liquid cultures (Fig. 4B,C). Strikingly, wee1D completely
abolished rescue of SIN mutants by dnt1D, as indicated by the
similar temperature sensitivity and rate of multinucleated cells
between SIN and wee1D dnt1D SIN strains (Fig. 4B,C). We also
observed that the wee1D completely abolished rescue of the
Fig. 4. Rescue of compromised SIN mutants by dnt1D is abolished by
deletion of wee1+. (A) Slightly elevated Wee1 levels rescue a cdc14-118
mutant. Serial dilutions (10-fold) of the indicated strains were spotted on
EMM with or without thiamine after being grown in YE and then washed by
EMM liquid and incubated at the indicated temperatures. (B,C) Rescue of SIN
mutants by dnt1D is dependent on the presence of wee1+. (B) Serial dilutions
(10-fold) of the indicated strains were spotted on YE plates and incubated for
3–5 days at the indicated temperatures before being photographed. (C) Single,
double and triple mutant cells with indicated genotypes were first grown in
YE at 25˚C, and then they were shifted to 37˚C (for cdc14-118 mutants; left)
or 33˚C (for sid2-250 mutants; right). Cells were sampled every 30 minutes
for a period of 6 hours, fixed and stained with DAPI. n.200 cells were
counted at each time point and the frequencies of cells with multiple nuclei
(as indication of cytokinetic defects) were quantified.
Dnt1 in cell cycle control4999
Journal of Cell Science
(supplementary material Fig. S6). Furthermore, rescue of SIN
mutants by dnt1D was not affected by either dma1D or cut12-S11
(supplementary material Fig. S7), suggesting that Dma1- or
Cut12-controlled Plo1 is not involved in SIN regulation by Dnt1.
These results allow us to conclude that rescue of compromised
SIN mutants by dnt1D is solely dependent on elevated Wee1
levels and the concomitant attenuation of Cdk1 activity.
Dnt1 functions in parallel with SCF to regulate Wee1
The finding that the absence of dnt1+leads to increased Wee1
levels prompted us to investigate the possible mechanism
involved in this alteration. We anticipated that Dnt1 might
influence the turn-over rate of Wee1. It has been shown that the
SCF complex is responsible for Wee1 degradation at the onset of
M phase in budding yeast, frog and human cells (Kaiser et al.,
1998; Michael and Newport, 1998; Watanabe et al., 2004).
Therefore, we first examined whether SCF is also required for
Wee1 degradation in fission yeast. Wee1 protein stability was
measured with Wee1-R.luciferase activity in the 26S proteasome
subunit mutant mts3-1 (Seeger et al., 1996), and temperature-
sensitive mutant skp1-A7, which carries a mutation (T454C) in
the major SCF subunit Skp1 (Lehmann et al., 2004). We found
that Wee1 protein was more abundant in mts3-1 and skp1-A7
cells than in wild-type cells at the restrictive temperature
(Fig. 5A), suggestingthat the SCF-mediated
dependent pathway is responsible for Wee1 proteolysis in
To further explore whether Dnt1 influences SCF-mediated
Wee1 degradation, we measured Wee1-R.luciferase activity and
cell length at division in skp1-A7 dnt1D double mutant cells. We
observed that dnt1D further enhanced Wee1 protein stability and
cell elongation in the skp1-A7 mutant (Fig. 5A,B). These data
suggest that Dnt1 probably does not directly affect SCF-mediated
We also noticed that the elevated Wee1 protein level caused by
skp1-A7 mutation could rescue SIN mutants to a similar degree as
dnt1D, and the triple mutant skp1-A7 dnt1D cdc14-118 showed
an even lower rate of accumulation of multinucleate cells
(Fig. 5C,D). It is noteworthy that the skp1-A7 mutation did not
rescue cdc14-118 more effectively than dnt1D, although the
skp1-A7 mutant had increased levels of Wee1. This is probably
due to the thermosensitivity of the skp1-A7 mutation, which
could make cdc14-118 cells sicker. Nonetheless, these data are
consistent with the idea that Dnt1 works in parallel with SCF to
regulate the rate of Wee1 protein turn-over.
Previously, we identified the nucleolar protein Dnt1 as an
inhibitor of the SIN signaling in fission yeast (Jin et al., 2007),
but its exact role and mechanism of action in the regulation of
SIN remained unclear. Here, we show that deletion of dnt1+leads
to an increased abundance of the Cdk1 inhibitor Wee1 kinase and
thus suppresses Cdk1 activity and causes a G2/M transition
delay. The ability of dnt1D to lower Cdk1 activity through Wee1
is responsible for the rescue of SIN mutants, because deletion of
the wee1+gene completely abolished elevated SIN signaling
caused by dnt1D. Hence, we have uncovered a novel mechanism
by which Dnt1 controls SIN signaling during late mitosis through
its effect on Cdk1 activity.
Negative correlation between SIN signaling and Cdk1
Initiation of cytokinesis is the final crucial event of the cell cycle,
and it must be tightly coordinated with completion of
chromosome segregation. In fission yeast, the timing of
cytokinesis is regulated by the SIN signaling pathway. Studies
performed with this organism have lent support to the notion that
SIN signaling is negatively regulated by Cdk1 activity, and that
maximum SIN activation depends upon Cdk1 inactivation. For
example, an early study showed that inactivation of Cdk1 by
cdc2-33 mutation promotes cytokinesis and septum formation
even in metaphase-arrested cells induced by overexpression of
the spindle assembly checkpoint protein Mad2 (He et al., 1997).
Subsequently, another study confirmed that loss of Cdc2 activity
in these metaphase-arrested cells was sufficient to allow SIN
Fig. 5. Dnt1 functions in parallel with SCF to regulate Wee1 degradation.
(A) Wee1 luciferase activity was analyzed using Wee1-R.luciferase and
Ade4-F.luciferase assay system in wild-type and mutant cells with indicated
genotypes, as described in Fig. 3D. Cells were collected from cultures grown
in YE at either 25˚C or 37˚C for 4 hours. ***P,0.001, *P,0.05.
(B) Quantification of cell lengths at cell division. Cells were first grown in YE
at 25˚C and then shifted to 37˚C for 4 hours before being imaged. n.200 cells
with septa were measured for each strain. ***P,0.001. (C) Serial dilutions
(10-fold) of the indicated strains were performed as in Fig. 4B. (D) The
frequencies of cells containing multiple nuclei (as an indication of cytokinetic
defects) were quantified in single, double and triple mutants with indicated
genotypes, as described in Fig. 4C.
Journal of Cell Science 126 (21) 5000
Journal of Cell Science
kinase complex Sid1–Cdc14 to localize at one SPB and to
promote cytokinesis onset (Guertin et al., 2000). More recently, it
has been shown that inactivation of Cdk1 using an analogue-
sensitive mutant allele cdc2-as promotes the asymmetrical
distribution of SIN proteins (Sid1 and Cdc7) to the spindle
poles, and the recruitment of the most downstream SIN
component Mob1 and b-(1,3) glucan synthase to the contractile
ring (Dischinger et al., 2008). Conversely, high Cdk1 activity
caused by increased expression of nondegradable Cdc13 (cyclin
B) blocked cells in anaphase and prevented SIN kinase Cdc7
from becoming asymmetrically localized to one SPB during
anaphase B; it also blocked all subsequent relocalization events
in the SIN and therefore inhibited septation (Chang et al., 2001;
Yamano et al., 1996).
The negative-feedback control of cytokinesis by Cdk1 activity
is not restricted to fission yeast and is conserved in all eukaryotes
from yeasts to vertebrates. In another model organism, budding
yeast S. cerevisiae, inactivation of Cdk1 in cells arrested by
expression of a non-degradable CLB2 promotes mitotic exit and
cytokinesis (Ghiara et al., 1991). Inactivation of Cdk1 is also
required for localization of the vesicle fusion machinery to the
bud neck during mitotic exit and cytokinesis (VerPlank and Li,
2005). Recently, it has been shown that Cdk1 is asymmetrically
localized to mother SPBs during early anaphase and inhibits
Cdc15 (orthologous to S. pombe Cdc7) and thus MEN signaling
activation (Ko ¨nig et al., 2010). In dividing cultured human cells,
inhibition of Cdk1 by selective small-molecule inhibitors also
caused premature cytokinesis, indicating that Cdk1 activity is
necessary and sufficient for maintaining the mitotic state of the
cells and preventing premature cytokinesis (Vassilev et al.,
2006). However, the detailed mechanism involved in this control
is not entirely understood.
Thus, the data obtained from all model organisms studied to
date point to a universal requirement for inactivation of CDK1 to
promote cytokinesis. This control mechanism is regarded as the
ideal coupling between nuclear and cell division. However, so
far, only two budding yeast proteins involved in mitotic exit and
cytokinesis, CDC15 and MOB1, have been confirmed to be
phosphorylated by Cdk1 (Jaspersen and Morgan, 2000; Ko ¨nig
et al., 2010). Whether homologues of these proteins in other
organisms (such as Cdc7 and Mob1 in fission yeast, or Mst2 and
Mob1 in Drosophila and vertebrates (reviewed by Doxsey et al.,
2005) are also phosphorylated by Cdk1 remains to be
demonstrated. Further studies will be required to identify and
define the complete repertoire of Cdk1 substrates influencing the
timing of cytokinesis in fission yeast and higher eukaryotes.
Nucleolar-protein-mediated control of Cdk1 activity and
Dnt1 is not the first nucleolar protein identified in fission yeast to
be involved in regulating both Cdk1 activity and SIN signaling.
Another nucleolar protein, the Cdc14 phosphatase family
member Clp1, is a well-studied example of such regulation.
Atthe end ofmitosis,Clp1
phosphorylation of Cdc25 and destabilizes and inactivates
Cdc25 by promoting recognition of Cdc25 by two ubiquitin E3
ligases, the anaphase-promoting complex/cyclosome (APC/C)
(RING-type) and Pub1/Pub2 (HECT-type) (Esteban et al., 2004;
Esteban et al., 2008; Wolfe and Gould, 2004). Thus, stabilized
Cdc25 allows clp1D cells to delay mitotic Cdk1 inactivation at
the end of mitosis and advance into G2 (Wolfe and Gould, 2004).
Moreover, SIN signaling is probably antagonized by persistent
Cdk1 activity. In addition to its role in affecting timely Cdk1
inactivation, Clp1 also directly contributes to robust and
successful cytokinesis, particularly when the cell division
machinery (such as the actomyosin ring) is slightly perturbed
or damaged (Mishra et al., 2004). A recent study revealed that
Clp1 is tethered at the contractile ring (CR) through its
association with the anillin-related protein Mid1, and regulates
CR protein dynamics and mobility by dephosphorylating CR
components Cdc15 and possibly Myo2 (myosin II heavy chain)
(Clifford et al., 2008).
Cdc14-family phosphatases are conserved in all eukaryotes
examined so far, but have been best studied in both budding and
fission yeasts. The negative regulation of Cdk1 at late mitosis
represents a conserved feature of Cdc14 phosphatase family
members (for reviews see D’Amours and Amon, 2004; Krapp
et al., 2004; Mocciaro and Schiebel, 2010; Stegmeier and Amon,
2004). Budding yeast ScCdc14 and fission yeast Clp1 are
regulated in part by their cell-cycle-dependent changes in
localization, with both proteins thought to be sequestered and
inactivated in the nucleolus during interphase. The activity of
ScCdc14 is controlled by its association with the competitive
inhibitor Cfi1/Net1, and only during anaphase is ScCdc14
released from its inhibitor to spread into the nucleus and
cytoplasm, allowing the dephosphorylation of its substrates
(reviewed by Mocciaro and Schiebel, 2010; Stegmeier and
Amon, 2004). In contrast to ScCdc14, both Clp1 and mammalian
Cdc14B are released from the nucleolus upon entry into mitosis
(Cho et al., 2005; Cueille et al., 2001; Nalepa and Harper, 2004;
Trautmann et al., 2001), but how this process is regulated is
currently unknown. Although the amino acid sequence of Dnt1
shows weak similarity to that of budding yeast Net1/Cfi1 and it
has been speculated that Dnt1 might function as a nucleolar
inhibitor of Clp1 (Jin et al., 2007), the present study demonstrates
that Dnt1 actually influences Wee1 stability and thus Cdk1
activity. Therefore, there are at least two parallel pathways
emerging from the nucleolus which converge at Cdk1 to regulate
SIN signaling: one including both Dnt1 and Wee1, and another
involving Clp1-controlled Cdc25 (Fig. 6).
Our findings also suggest that SCF-mediated degradation of
Wee1 is probably conserved in fission yeast, although the direct
evidence of Wee1 ubiquitylation by SCF is currently lacking. The
elucidation of the specific mechanism by which Dnt1 controls
Fig. 6. Model depicting how Dnt1- and Clp1-dependent modulation of
Wee1 and Cdc25, respectively, converge at Cdk1 to regulate SIN
signaling. Signaling flows in wild-type cells (A) and dnt1D or skp1-A7
mutants (B) are summarized.
Dnt1 in cell cycle control5001
Journal of Cell Science
degradation of Wee1 remains a mystery and merits further
Materials and Methods
General yeast strains, media and growth conditions
Fission yeast strains used in this study were constructed by standard techniques
(Moreno et al., 1991) and are listed in supplementary material Table S1. S. pombe
strains were grown in rich medium [yeast extract (YE)] or Edinburgh minimal
medium (EMM) with appropriate supplements (Moreno et al., 1991). EMM with
5 mg/ml thiamine was used to repress expression from the nmt1 promoter. YE
containing 100 mg G418 (Sigma) or nourseothiricin (clonNAT) (Werner
BioAgents) per liter was used for selecting kanRand natRcells, respectively.
For serial-dilution drop tests for growth, three serial 10-fold dilutions were made
and 5 ml of each was spotted on plates with the starting cell number of 104. Cells
were pre-grown in liquid YE or EMM at 25˚C until they reached exponential phase
and were then spotted onto YE or EMM plates at the indicated temperatures and
incubated for 3 to 5 days before scanning. In block-and-release experiments
performed with cdc25-22 mutant strains, the cells were grown in YES medium to
an A600of 0.2 at 25˚C (permissive temperature), shifted to 36˚C for 3.5 hours, and
then released from the growth arrest by transfer back to 25˚C.
Construction of epitope-tagged strains carrying Ade4-F.luciferase and
Wee1-R.luciferase and luciferase-activity assay
Firefly and Renilla Luciferase lacking start codons were cloned from the pGL3 and
pRL-TK plasmids (Promega) into pFA6a plasmid carrying either kanRor natR
selection marker. These plasmids were used to add C-terminal epitope tags to the
endogenous locus to create Ade4-F.luciferase (kanR) and Wee1-R.luciferase (natR)
alleles using PCR-based gene targeting (Ba ¨hler et al., 1998). Candidate
transformants by homologous recombination were selected by growth on
selective plates containing G418 or ClonNAT and subsequently confirmed by
Western blot analyses
Yeast cell extracts were prepared in lysis buffer (120 mM Tris-HCl, pH 6.8, 4%
SDS, 20% glycerol, 8M urea, 0.6 M b-mercaptoethanol) by glass bead disruption
using FastPrep homogenizer (MP Biomedical), and denatured in 26SDS sample
buffer (100 mM Tris-HCl, pH 6.8, 4% SDS, 20% glycerol, 0.2% Bromophenol
Blue, 0.6 M b-mercaptoethanol) before being resolved on 8–12% SDS-PAGE gels.
For detection, mouse monoclonal anti-GFP (Roche), rabbit polyclonal anti-Myc
(GeneScript), rat monoclonal anti-HA (Roche) and rabbit polyclonal anti-Cdc2
(Tyr15) (sc-7989-R, Santa Cruz Biotechnology) were used as the primary
antibodies (1:1000–1:2000 dilutions). Cdc2 or GFP-atb2 was detected using
rabbit polyclonal anti-PSTAIRE (sc-53, Santa Cruz Biotechnology) or mouse
monoclonal anti-GFP (Roche), respectively, as loading controls (1:1000 dilution).
Goat anti-mouse, anti-rabbit or anti-rat conjugated to horseradish peroxidase
(Pierce) was used as the secondary antibody at 1:10,000 dilution. Membranes were
developed with ECL western blotting reagents (Pierce). Protein levels were
quantified using ImageJ software (National Institutes of Health, Bethesda, MD)
with Cdc2 or GFP-atb2 as a loading control.
Luciferase activity assay in yeast cell lysate
Yeast strains carrying both Ade4-F.luciferase (kanR) and Wee1-R.luciferase (natR)
were used. Exponential phase cultures were pre-grown in liquid YE at 30˚C, 25˚C
or shifted from 25˚C to 37˚C for 4 hours, and 1–106107cells were collected and
washed once with sterile water. Cell lysates were prepared in 100–200 ml cold 16
passive Lysis Buffer (PLB) by glass bead disruption using FastPrep homogenizer.
Firefly and Renilla luciferase activities in cell lysates were measured using the
Dual-Reporter Assay System (Promega) following instructions provided by the
RT-PCR and silver staining
Total RNA was isolated from yeast cells grown in YES liquid medium to an A600
of ,0.4 and disrupted with FastPrep homogenizer by using the TRI Reagent
Solution (TRIZOL) (TaKaRa Bio) and following the manufacturer’s instructions.
RT was performed with Prime-Script RT reagent Kit (TaKaRa) at 37˚C for
15 minutes followed by treatment at 85˚C for 5 seconds. Primer pairs to detect
wee1+and act1+transcripts by PCR were as follows: wee1-2189 bp-F, 59- GT-
GAATACATTGCGCGTAAG-39 and wee1-2540 bp-R, 59-CTACGCATTTCAA-
CCCAGC-39 (amplifying a 400 bp fragment); act1-F, 59-GTATGCCTCTGGTC-
GTACCAC-39 and act1-R, 59-CAATTTCACGTTCGGCGGTAG-39 (amplifying a
200 bp fragment). PCR products were then resolved by 5% Tris-acetate-EDTA
PAGE (TAE-PAGE). The gels were stained with 0.25% AgNO3and scanned and
images were processed with Adobe Photoshop software and the intensities of PCR
products were quantified using ImageJ software with act1+transcript as an internal
GFP-fusion proteins were observed in cells after fixation with cold methanol or in
live cells. For DAPI (49, 6-diamidino-2-phenylindole, Sigma) staining of nuclei,
cells were fixed with cold methanol, washed in PBS and resuspended in PBS plus
1 mg/ml DAPI. To determine cell size at division the yeast strains were grown in
either EMM2 or YES medium to an A600of ,0.5 and treated with Calcofluor
white (CW, Sigma) at 50 mg/ml final concentration, which specifically stains cell
wall and septum. A minimum of 100 septated cells were scored for each mutant.
Photomicrographs were obtained using a Nikon 80i fluorescence microscope
coupled to a cooled CCD camera (Hamamatsu, ORCA-ER) and image processing,
analysis and cell length measurement were carried out using Element software
(Nikon) and Adobe Photoshop.
All experiments were repeated at least three times with similar results. In order to
determine statistical significance of our data, two tailed Student’s t-tests were
performed, and P,0.05 was considered statistically significant.
Balasubramanian, Dannel McCollum, Jianhua Liu, Paul Russell,
Iain Hagan, Takashi Toda, Janni Petersen, James Moseley and the
Yeast Genetic Resource Center Japan (YGRC) for providing yeast
strains or plasmids.
Y.W. and Q.W.J. conceived the project and designed the
experiments; Z.Y.Y., M.T.Z., G.Y.W., D.X., D.K., A.F., J.C. and
H.M. performed the experiments; Z.Y.Y., J.C., H.M., N.R., Y.W. and
Q.W.J. analyzed data; Y.W. and Q.W.J. wrote the paper.
This work was supported by the National Institutes of Health [grant
number GM-069957 to N.R.]; MEC European Regional Development
Fund co-fundingfrom the EU [grant number BFU2011-22517 to J.C.];
the National Natural Science Foundation of China [grant number
31171298 to Q.W.J.]; Key Project of the Chinese Ministry of
Education [grant number 108076 to Q.W.J.]; the 111 Project of
Education of China [number B06016]. Deposited in PMC for release
after 12 months.
Supplementary material available online at
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