4130 Research Article

IntroductionDevelopmental decisions often involve differentiation processes thatneed to reset the cell cycle for induction of a new morphogeneticprogram. This is probably also the case for induction of the virulenceprogram in pathogenic fungi. It could therefore be assumed that inpathogenic fungi the control of the cell cycle, as well asmorphogenesis, is linked somehow to the virulence program. Themaize smut fungus Ustilago maydis is an excellent system to addressthe relationships between cell cycle, morphogenesis andpathogenicity (Perez-Martin et al., 2006; Steinberg and Perez-Martin, 2008). During the induction of virulence program in thisfungus, an infectious dikaryotic hypha is produced as a result ofthe mating of a pair of compatible haploid budding cells. Therefore,the induction of the pathogenic program implies not only strongmorphological changes (bud to hypha transition) but also geneticchanges (haploid to dikaryotic transition). Accurate control of thecell cycle and morphogenesis is predicted during these transitions.

The formation of the infectious hypha in U. maydis depends onan intricate transcriptional program which primarily involves atranscriptional regulator called b-factor (Feldbrugge et al., 2004).The production of this master regulator is linked to the matingprocess that, after cell fusion, leads to the interaction of the twosubunits composing the b-factor (bW and bE), one provided by eachmating partner. This way, the mating of two compatible cells (i.e.carrying b-subunits able to dimerize) results in the formation of adikaryotic cell, which undergoes a strong polar growth as well asan apparent cell cycle arrest on the plant surface. Eventually, theinfectious hypha manages to enter the plant tissue, where it startsto proliferate, maintaining its dikaryotic status. The reasons for theapparent cell cycle arrest on the plant surface are unknown. It hasbeen hypothesized that such a cell cycle adjustment would berequired for a precise execution of the virulence program (Perez-Martin et al., 2006). In fact, the apparent cell cycle arrest of theinfectious hypha on the plant surface observed in U. maydis seems

to be more general, and it is also present in rust fungi such asUromyces phaseoli (Heath and Heath, 1979).

Ectopic expression of compatible b-subunits in a haploid cellinduces the formation of a monokaryotic filament that mimics itsdikaryotic counterpart in all aspects of filamentous growth as wellas apparent cell cycle arrest (Brachmann et al., 2001). Whichmechanisms are used by the bW-bE heterodimer to induce thehyperpolarized growth and to arrest the cell cycle in the infectioushypha and at which stage this occurs are unknown, but these issuesare currently being studied. For example, recent results from ourlaboratory indicated that the activation of hyperpolarized growthafter bW-bE heterodimer formation is dependent on the U. maydisortholog of the cyclin-dependent kinase Cdk5, a well-knownregulator of neuron development in mammals (Alvarez-Tabares andPerez-Martin, 2008; Castillo-Lluva et al., 2007; Flor-Parra et al.,2007).

Here, we address some of the questions related to the b-factor-induced cell cycle arrest, showing that such cell cycle arrest takesplace in the G2 phase as a consequence of an increase of inhibitoryphosphorylation of the mitotic cyclin-dependent kinase.Interestingly, we also report that the DNA-damage checkpointkinase Chk1 seems to be involved in this process.

ResultsG2 cell cycle arrest in U. maydis upon expression ofcompatible b-homeoproteinsTo address the molecular mechanisms of the b-factor-dependent cellcycle arrest, we made use of the previously described (Brachmannet al., 2001) haploid U. maydis AB33 strain, which carry compatible(i.e. able to dimerize) bE and bW genes under the control of theregulatable nar1 promoter. As the nar1 promoter can be inducedby growing cells in nitrate-containing medium (MM-NO3) andrepressed in ammonium-containing medium (MM-NH4), b-factor-dependent infective filament formation can be elicited in AB33 by

During induction of the virulence program in thephytopathogenic fungus Ustilago maydis, the cell cycle isarrested on the plant surface and it is not resumed until thefungus enters the plant. The mechanism of this cell cycle arrestis unknown, but it is thought that it is necessary for the correctimplementation of the virulence program. Here, we show thatthis arrest takes place in the G2 phase, as a result of an increasein the inhibitory phosphorylation of the catalytic subunit of themitotic cyclin-dependent kinase Cdk1. Sequestration in thecytoplasm of the Cdc25 phosphatase seems to be one of thereasons for the increase in inhibitory phosphorylation.

Strikingly, we also report the DNA-damage checkpoint kinaseChk1 appears to be involved in this process. Our resultssupport the emerging idea that checkpoint kinases have rolesother than in the DNA-damage response, by virtue of theirability to interact with the cell cycle machinery.

Supplementary material available online athttp://jcs.biologists.org/cgi/content/full/122/22/4130/DC1

Key words: Cell cycle, Ustilago maydis, Chk1, Phytopathogenicfungi

Summary

A role for the DNA-damage checkpoint kinase Chk1 inthe virulence program of the fungus Ustilago maydisNatalia Mielnichuk, Cecilia Sgarlata and José Pérez-Martín*Departamento de Biotecnología Microbiana, Centro Nacional de Biotecnología CSIC, Campus de Cantoblanco-UAM, 28049 Madrid, Spain*Author for correspondence (jperez@cnb.csic.es)

Accepted 10 September 2009Journal of Cell Science 122, 4130-4140 Published by The Company of Biologists 2009doi:10.1242/jcs.052233

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changing the nitrogen source. As a control we used a similar strain,AB34, which harbors incompatible bE and bW genes (i.e. unableto produce dimers) under the control of nar1 promoter (Brachmannet al., 2001) (Fig. 1A).

We measured the DNA content of these strains under inductionand non-induction conditions using fluorescence-activated cellsorting (FACS) analysis and found that cells expressing compatibleb-proteins accumulated with a 2C DNA content (Fig. 1B). Tospecifically ascribe the cell cycle stage at which arrest takes place,we introduced two cytological markers that helped us to distinguishG2 from early mitosis in a strain expressing compatible b-proteins:an -tubulin-GFP protein fusion allows the detection of themicrotubule cytoskeleton (Steinberg et al., 2001) and a Cut11-RFPfusion protein labels the nuclear envelope (Pérez-Martín, 2009).During the G2 phase, the nuclear envelope is present and themicrotubules form a cytoplasmic network (Fig. 1C, inset arrow);however, once cells enter mitosis, the nuclear envelope isdisassembled and the microtubules are concentrated in the spindle(Fig. 1C, inset, arrowhead) (Steinberg et al., 2001; Straube et al.,2005). Filaments produced after expression of compatible b-proteins showed a single nucleus surrounded by an intact nuclearenvelope and a defined cytoplasmic array of microtubules (Fig. 1C,arrows), all of which, together with the 2C DNA content, isconsistent with a G2 cell cycle arrest during the formation of theinfective filament.

G2 cell cycle arrest is dependent on the inhibitoryphosphorylation of Cdk1G2-M transition in U. maydis is enabled by the activity of two CDKcomplexes: Cdk1-Clb1 and Cdk1-Clb2 (Garcia-Muse et al., 2004;

Sgarlata and Perez-Martin, 2005a). To determine the kinase activityassociated with Cdk1 during b-factor-dependent filament formation,we took advantage of the Schizosaccharomyces pombe protein Suc1,which is known to bind specifically to mitotic CDKs with highaffinity (Ducommun and Beach, 1990) and that we previouslyproved to be a convenient assay for characterizing the U. maydisCdk1 protein (Garcia-Muse et al., 2004). Cell lysates obtained fromAB33 and AB34 strains growing in induction (MM-NO3) and non-induction (MM-NH4) conditions were incubated with Suc1-conjugated Sepharose beads and purified fractions assayed forhistone H1 kinase activity (Fig. 2A). Consistently with the inabilityof the hyphae to enter mitosis, Cdk1-associated kinase activity wasdownregulated by more than 90% after induction of the expressionof compatible b-factor genes (Fig. 2B).

Downregulation of CDK activity can be produced in differentways in U. maydis (Perez-Martin et al., 2006), one of these, whichis particularly active during the control of G2-M transition, is thephosphorylation of the Tyr15 residue in Cdk1 (Sgarlata and Perez-Martin, 2005a). Therefore, we analyzed levels of Cdk1 inhibitoryphosphorylation upon induction of b-factor genes. For this, we useda specific antibody raised against the phosphorylated human Cdc2-Y15-P peptide, which recognizes the Tyr15-phosphorylated formof U. maydis Cdk1 (Sgarlata and Perez-Martin, 2005a). Tyr15P-Cdk1levels increased eightfold when compatible b-proteins wereexpressed (Fig. 2C,D).

To address the importance of this phosphorylation during the b-factor-induced cell cycle arrest, we took advantage of the cdk1AF

allele, in which the inhibitory phosphorylation sites in Cdk1 arereplaced with residues that cannot be phosphorylated (Thr14 to Alaand Tyr15 to Phe, respectively) (Sgarlata and Perez-Martin, 2005a).

Fig. 1. Expression of b-homeoproteins in Ustilago maydis produces a G2 cell cycle arrest. (A)Scheme of the cassettes expressing compatible (AB33) or non-compatible (AB34) b-factor genes. Only the compatible pair is able to form the heterodimer. (B)FACS analysis of the DNA content of AB34 and AB33 strainsgrowing in inducing (MM-NO3) and non-inducing (MM-NH4) conditions. The period of incubation in testing medium is indicated (hours). (C)Cell images ofAB33-derived cells carrying Tub1-GFP and Cut11-RFP fusions to detect the microtubule cytoskeleton (arrows) and the nuclear envelope growing in conditions thatinduce the expression of the b-factor genes (MM-NO3). Different stages during the production of the infective hyphae are shown. The inset shows the same straingrowing in non-inducing conditions (MM-NH4). Two cells are shown, one in G2 phase (the nuclear envelope is present and the microtubules are forming acytoplasmic network, arrow) and the other is in the middle of mitosis (there is no nuclear envelope signal and the microtubules are concentrated in the spindle,arrowhead). Scale bars: 10m.

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As continuous expression of this allele produces a deleterious effectin the cell (Sgarlata and Perez-Martin, 2005a), we decided tointroduce an ectopic copy of this mutant allele under the samepromoter as the b-factor genes (i.e. nar1 promoter) in AB33 cells,in such a way that production of Cdk1AF occurs at the same time asthe cell produces b-proteins. As a control, we constructed an AB33strain carrying an ectopic copy of a wild-type cdk1 allele under controlof the nar1 promoter. We used 3�Myc-tagged versions of both thewild-type and mutant ectopic Cdk1 to discriminate between the levels

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of protein produced by the endogenous locus and the ectopicallyexpressed alleles (Fig. 3A). We found that in the strain expressingthe Cdk1 version refractory to inhibitory phosphorylation (AB33cdk1AF), the Cdk1-associated kinase activity remained high. Bycontrast, in AB33 or in the control strain expressing a Cdk1 versionthat can be phosphorylated (AB33cdk1), the kinase activity wasdownregulated after b-factor expression (Fig. 3B). Consistently, inthe presence of Cdk1AF mutant protein, the b-factor-dependentfilaments carried several nuclei, indicating that no cell cycle arrest

Fig. 2. b-factor-induced Cdk1 inactivation in U. maydis.(A)Kinase activity of the Cdk1 complexes purified usingSuc1 beads obtained from AB33 and AB34 cells growingin inducing and non-inducing conditions for 8 hours. Theupper panel shows an immunoblot using anti-PSTAIREantibody, which detects Cdk1 and Cdk5 kinases in U.maydis; only Cdk1 binds to Suc1 beads. The Suc1 proteinprecipitates were used for in vitro phosphorylation ofhistone H1 (lower panel, autoradiograph of SDS-PAGE).(B)Quantification of Cdk1 kinase activity after b-factorexpression. Kinase activity obtained from AB34 cellsgrowing in non-induction conditions (MM-NH4) was usedas reference (100% of activity). Means ± s.d. are shown.(C)Levels of Cdk1 inhibitory phosphorylation after b-factor induction. Protein extracts from the indicatedstrains growing in inducing conditions (MM-NO3) for theindicated time (hours) were separated by SDS-PAGE.Immunoblots were incubated successively with anantibody that recognizes the Cdk1 phosphorylated form(Cdc2-Y15P) and anti-PSTAIRE. (D)Levels of Cdk1phosphorylation were determined by quantifying the levelof antibody signal using ChemiDoc (Bio-Rad). Signalfrom the phosphopeptide-specific antibodies wasnormalized to the amount of phosphorylation of controlstrain (AB34) at time zero. Differences in loading ofsamples were corrected by dividing each phosphopeptide-specific antibody signal by the Cdk1 (PSTAIRE) antibodysignal. Means ± s.d. are shown.

Fig. 3. Cdk1 inhibitory phosphorylation is required for b-factor-induced cell cycle arrest in U. maydis. (A)StrainsAB33 Pnar1:cdk1-myc and AB33 Pnar1:cdk1AF-myc, inaddition to the endogenous cdk1 wild-type allele, carry anectopic extra copy of a cdk1-myc or cdk1AF-myc alleleunder the control of nar1 promoter, in such a way thatconditions inducing b-factor gene expression also inducedthe production of the Myc-tagged Cdk1 proteins. Panelshows western blot using anti-PSTAIRE antibody of thedifferent strains, after 8 hours in inducing conditions(MM-NO3). Note the levels of Cdk-Myc and Cdk1AF-Mycproteins with respect to endogenous levels of wild-typeCdk1. (B)Kinase activity of the indicated strains. Activityof the control strain AB34 growing in MM-NO3 was usedas reference (100% activity). (C,D)Cell images of AB33(control) and AB33-derived strains expressing cdk1-myc(cdk1) or the non-phosphorylatable cdk1AF-myc (cdk1AF)alleles, incubated for 8 (C) and 24 (D) hours in inducingconditions (MM-NO3). Cells were stained with DAPI todetect nuclei. Scale bars: 20m.

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was taking place (Fig. 3C,D). Expression of wild-type Cdk1 producedfilaments that were indistinguishable from those obtained with theAB33 control strain. Taken together, these results indicate thatinhibitory phosphorylation of Cdk1 has a major role during the b-factor-induced G2 arrest in U. maydis.

Cdk1 inhibitory phosphorylation during b-factor-induced cellcycle arrest depends on Wee1Cdk1 inhibitory phosphorylation during vegetative growth dependson the activity of the essential Wee1 kinase (Sgarlata and Perez-Martin, 2005a). To address whether Wee1 is required for b-factor-induced cell cycle arrest, we took advantage of previously describedstrain (AB31) (Brachmann et al., 2001) that expresses compatibleb-factor genes under control of the crg1 promoter, which is inducedin growth medium containing arabinose as a carbon source andrepressed when glucose is present (Bottin et al., 1996). Weintroduced the conditional allele wee1nar1 in this strain (Fig. 4A)(Sgarlata and Perez-Martin, 2005a). This allele enables theexpression of wee1 in cells incubated in minimal medium plusnitrate as a nitrogen source (MM-NO3) as well as the downregulationof wee1 expression in complete medium (CM, Fig. 4B).

When AB31-derived wee1 conditional cells were grown inrepressive conditions (CM) and in the presence of arabinose,multinucleated filaments were observed (Fig. 4C). In similarconditions, AB31 cells had a single nucleus, indicating that Wee1is necessary for b-factor-induced cell cycle arrest. Consistently, theinability to arrest the cell cycle in the absence of Wee1 correlatedwith a dramatic decrease in the level of Tyr15-P Cdk1, as well asa high level of Cdk1-associated kinase activity (Fig. 4D,E).

b-factor-dependent cell cycle arrest is not solely mediated bytranscriptional regulation of wee1We analyzed the levels of wee1 mRNA in AB33 cells after b-factorinduction and found a sevenfold increase with respect to the AB34control strain (Fig. 5A and supplementary material Fig. S1). Sinceoverexpression of wee1 produces a G2 cell cycle arrest (Sgarlataand Perez-Martin, 2005a), we wondered whether this upregulationwas responsible for b-factor-induced cell cycle arrest. To answerthis question, we exchanged the endogenous wee1 native promoterwith the constitutively expressed scp promoter in AB33 cells (Fig.5B). This mutant allele, wee1scp, produces a low and constitutivelevel of wee1 mRNA without any appreciable defect in cell cycleregulation in normal conditions (Sgarlata and Perez-Martin, 2005a).In AB33 wee1scp cells, wee1 mRNA level was maintained at a lowlevel upon b-factor induction (Fig. 5C) However, we found a minoreffect in cell cycle arrest: less than 25% of filaments showed morethan one nucleus (Fig. 5D,E), indicating that the observedtranscriptional upregulation of wee1 is not the only reason by whichcell cycle arrests during b-factor induction.

Ectopic cdc25 overexpression impairs the b-factor-induced cellcycle arrestThe level of inhibitory phosphorylation of Cdk1 also depends onthe activity of the Cdc25 phosphatase, which reverses the CDKinhibition (Sgarlata and Perez-Martin, 2005b). Therefore, weanalyzed the levels of cdc25 mRNA during b-factor-inducedfilamentation. However, we found that it was not affected by thepresence of an active b-heterodimer (Fig. 6A). In spite of this result,we wondered whether it was possible to release the cell cycle arrestby overexpressing cdc25. To achieve this, we introduced in a straincarrying compatible b-factor genes under control of the nar1

promoter (AB33 cells), an ectopic copy of cdc25 under the controlof the dik6 promoter, which is strongly induced by the b-proteins(Brachmann et al., 2001) (Fig. 6B). In the resulting strain, inductionof b-factor genes resulted in a dramatic increase in cdc25 mRNA

Fig. 4. Wee1 is required for b-factor-dependent cell cycle arrest. (A)Schemeshowing the constructions inserted in the strains used. AB31 carriescompatible b-factor genes under the control of the crg1 promoter that can beinduced by arabinose as a carbon source and repressed in the presence ofglucose. In AB31wee1nar1, the endogenous promoter of wee1 was exchangedwith the regulatable nar1 promoter. (B)Levels of wee1 mRNA in the indicatedstrains when cells were grown in conditions for b-factor gene expression (1%arabinose) and conditions for nar1 expression (MM-NO3) or repression (CM,complete medium carrying casamino acids and NH4 as nitrogen source).(C)Cell images of AB31 (control) and AB31-derived strain carrying aconditional wee1nar1 allele (wee1OFF) after 8 and 24 hours of incubation underconditions of expression of b-factor genes and repression of wee1 (completemedium plus arabinose). Cells were stained with DAPI to detect nuclei.(D)Levels of Cdk1 phosphorylation of AB31 and AB31 wee1nar1 cellsgrowing for the indicated time in complete medium plus arabinose as carbonsource. Quantification was as above. (E)Quantification of Cdk1 kinaseactivity after b-factor expression in the indicated strains. Kinase activityobtained from control AB32 cells growing in induction conditions (MM-NO3)was used as reference (100% activity). Means ± s.d. are shown.

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levels (Fig. 6C). Interestingly, the filaments expressing high levelsof cdc25 had more than one nucleus in a substantial proportion(64% of the filaments) (Fig. 6D). Furthermore, the cells expressingcdc25 under control of the dik6 promoter showed a low level ofinhibitory phosphorylation of Cdk1 (Fig. 6E).

Altogether, the results obtained with Cdc25 and Wee1 indicatedthat a similar regulatory scheme controlling G2-M transition viaCDK inhibitory phosphorylation during vegetative growth is alsoinvolved in b-factor-induced cell cycle arrest.

Cdc25 seems to be retained in the cytoplasm during b-factor-induced filamentationThe observation that overexpression of cdc25 resulted in amultinucleated filament can be explained if b-factor-induced cellcycle arrest relies on an inhibition of Cdc25. One way to controlCdc25 activity in U. maydis cells is to regulate its subcellularlocalization (Mielnichuk and Perez-Martin, 2008). We explored thispossibility by monitoring Cdc25 nucleus to cytoplasm distributionin cells expressing GFP-tagged Cdc25 from the genomic locus. Toquantitatively evaluate changes in localization, fluorescenceintensities of the Cdc25 signal were determined for a circular regioncorresponding to the nucleus (N) and a comparable area in thecytoplasm (C), as described (Mielnichuk and Perez-Martin, 2008).We found that in inductive conditions (MM-NO3) the ratio N/C incontrol cells (AB34 cells) was about 1.1, whereas this ratio in AB33cells was around 0.2 (Fig. 7A).

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Previous work from our laboratory showed that retention ofCdc25 in the cytoplasm of U. maydis depends on the essential 14-3-3 protein Bmh1 (Mielnichuk and Perez-Martin, 2008). Toaddress whether 14-3-3 was required for b-factor-induced cell cyclearrest, we expressed compatible b-factor genes in conditions ofdownregulation of bmh1 expression (Fig. 7B). We found that inthese conditions, morphologically aberrant filaments were producedthat contained several nuclei each (Fig. 7D, middle panel). Since14-3-3 proteins are extremely pleiotropic (van Hemert et al., 2001),it is hard to unequivocally ascribe the effects observed infilamentation after bmh1 downregulation to the inability to arrestcell cycle. To link this directly to a defect in the ability to retainCdc25 in the cytoplasm, we took advantage of a cdc25 mutant allele(cdc25AAA, Fig. 7C), which was impaired in the interaction withBmh1 and therefore accumulates in the cell nucleus (Mielnichukand Perez-Martin, 2008). We found that a strain carrying thecdc25AAA allele and expressing compatible b-proteins resulted in ahigh percentage of filaments carrying more than one nucleus (Fig.7D, bottom panel). Collectively, these data suggest that custody ofCdc25 in the cytoplasm is part of the mechanism by which b-factor-dependent cell cycle arrest occurs.

Chk1 is involved in the b-factor-dependent cell cycle arrestCytoplasmic retention of Cdc25 by 14-3-3 proteins requires theprevious phosphorylation of target sites in Cdc25 (van Hemert etal., 2001). The DNA-damage checkpoint kinase Chk1 is one of the

Fig. 5. Transcriptional regulation of wee1 has a minor role duringb-factor-dependent cell cycle arrest. (A)Northern analysis of wee1mRNA levels in strains carrying compatible (AB33) orincompatible (AB34) b-factor genes and grown for the indicatedtime (hours) in inducing (MM-NO3) or repressing (MM-NH4)conditions for b-factor expression. Ribosomal RNA was used asloading control. (B)Scheme showing the genes relevant for theexperiments showed in panels C-E. The scp promoter transcribesat a low and constitutive level of expression. (C)Levels of wee1mRNA in the indicated strains when cells were grown inconditions for b expression (MMNO3) for the indicated time(hours). (D)Images of the AB33-derived strain carrying thewee1scp allele incubated for 8 hours in induction conditions. Cellswere stained with DAPI to detect nuclei. (E)Quantification offilaments carrying single nucleus after 24 hours of incubation.Mean and s.d. are shown.

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described kinases able to phosphorylate Cdc25 (Karlsson-Rosenthaland Millar, 2006). Interestingly, the mutations carried in thecdc25AAA allele affect the ability of the DNA-damage checkpointkinase Chk1 to arrest the cell cycle in U. maydis (Perez-Martin,2009). Therefore, we analyzed whether Chk1 was required for b-factor-induced cell cycle arrest. To our surprise, cells defective inChk1 and expressing compatible b-proteins were impaired in theirability to arrest the cell cycle (Fig. 8A,B).

Even when the b-factor-dependent filament produced in AB33cells mimics its dikaryotic counterpart in all aspects of filamentousgrowth, we sought to analyze the consequences of the absence ofchk1 in the formation of the infective filament in more ‘native’conditions. Crosses of compatible haploid wild-type cells oncharcoal-containing plates (Banuett and Herskowitz, 1989) resultedin the formation of cell-cycle-arrested dikaryotic hyphae (Flor-Parraet al., 2006). Therefore, we crossed compatible wild-type and chk1strains on charcoal-containing plates. Wild-type crosses led to afilamentous appearance of the colony, whereas chk1 mutants werestrongly attenuated in filament formation (Fig. 8C,D). We alsoanalyzed the nuclear composition of filaments in these crosses. Todistinguish the b-factor-induced filaments from the cell populationbackground (frequently enriched in aberrant elongated cells) thehaploid strains we used expressed a GFP fusion to a nuclearlocalization signal under control of the dik6 promoter. In this way,only cells resulting from mating and therefore expressing the b-factor-dependent program produced a fluorescent nuclear signal.We found that almost all cells expressing the b-factor carried twonuclei, whereas chk1 filaments frequently carried more than twonuclei (Fig. 8E,F), which was consistent with a defect in the abilityto arrest entry into mitosis during the formation of the infectivehyphae.

Chk1 is transiently activated during b-factor-induced cell cyclearrestThe above results indicated that during induction of the pathogenicdevelopment in U. maydis, a G2 cell cycle arrest takes place inwhich the checkpoint kinase Chk1 has a role. Chk1 is a protein thatis produced in normal conditions in an inactive form, which isactivated after phosphorylation by upstream kinases (Chen andSanchez, 2004). Therefore, we examined whether Chk1 wasactivated during the formation of the b-factor-dependent filament.In U. maydis cells, as in other organisms, Chk1 activation can beeasily monitored by the accumulation of a GFP-tagged Chk1 proteininto the nucleus (Perez-Martin, 2009). When AB33-derived cellscarrying a Chk1-GFP fusion were induced to produce filaments,we observed a clear accumulation of the GFP signal in the nucleus.Control AB34 cells showed no accumulation of GFP in the nucleusthroughout the incubation period (Fig. 9A). Interestingly, AB33-derived long filaments seemed not to accumulate any nuclear GFPsignal. To quantify this apparently transient response, we measuredcells (n80 in two independent experiments) of different length andplotted this against the presence or not of a nuclear GFP signal (Fig.9B), confirming that only shorter filaments (i.e. early stages) showednuclear accumulation of Chk1.

There are several possible explanations for this behavior. Onepossibility is that Chk1 is degraded at later stages. A differentexplanation is that even when the nuclear membrane is presentduring filament formation (see Fig. 1C), it becomes permeable atlater stages. And finally, it is possible that Chk1 is transientlyactivated during b-factor-dependent filamentation. To address thesepossibilities, we monitored the decrease in the mobility of a Myc-tagged protein during electrophoresis as a surrogate marker for theirphosphorylation-dependent activation (Perez-Martin, 2009). First,

Fig. 6. Expression of cdc25 under controlof the dik6 promoter. (A)Northern analysisof cdc25 mRNA levels in strains carryingcompatible (AB33) or incompatible(AB34) b-factor genes and growing for theindicated time (hours) in inducing (MM-NO3) or repressing (MM-NH4) conditionsfor b-factor expression. Ribosomal RNAwas used as loading control. (B)Schemeshowing the constructions inserted in thedifferent strains. AB33 Pdik6:cdc25 carriesan additional ectopic copy of the cdc25gene under the control of the dik6promoter that is activated by the presenceof a compatible b-heterodimer.(C)Northern analysis of cdc25 levels in thecontrol strain (AB33) and the straincarrying the ectopic copy of cdc25 underdik6 control growing in non-inducing(MM-NH4) and inducing (MM-NO3)conditions for the time indicated (hours).(D)Images of an AB33-derived straincarrying an ectopic copy of cdc25 underthe control of the b-factor-induced dik6promoter after 8 and 24 hours of growth inconditions for expression of b-factor genes(MM-NO3). Cells were stained with DAPIto detect nuclei. (E)Levels of Cdk1phosphorylation of AB33 and AB33Pdik6:cdc25 cells growing for the indicatedtime in MM-NO3. Quantification was asabove. Means ± s.d. are shown.

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we observed that the overall levels of Chk1 were not affected bythe incubation period, excluding degradation of Chk1 as a possibleexplanation (Fig. 9C). Interestingly, the electrophoretic shift wasmaximum after 3 hours of induction but decreased to negligibleafter 9 hours, supporting the explanation that Chk1 is transientlyactivated during the formation of the b-factor-dependent filament.

Chk1 is required for full virulenceU. maydis infection of maize results in anthocyanin pigmentproduction by the plant and the formation of tumors that are filledwith proliferating fungal cells, which eventually differentiate intoblack teliospores (Banuett and Herskowitz, 1996). To address theconsequences of a defective cell cycle arrest during corn infectionby U. maydis, we inoculated maize plants by stem injection withmixtures of wild-type and chk1 mutant strains. Both crosses wereable to infect plants; however, the mutant strains were less efficient(42% of the plants showed no symptoms) and the severity of thesymptoms was minor (Fig. 10A). Interestingly, plants infected withthe chk1 mutant rarely showed large tumors (Fig. 10C), and evenin these rare occasions, no teliospores were found, suggesting arole of Chk1 beyond the initial steps of infection (i.e. infectivefilament formation).

To address whether the mutant hyphae proliferate inside the plant,symptomatic leaves obtained from both mutant and wild-type crosses

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were sampled after 1 week of inoculation and Chlorazole Black Estaining was used to visualize invading hyphae (Brachmann et al.,2003). Septated hyphae grown through epidermal cells were observedin plants inoculated with compatible wild-type strains. When materialobtained from mutant crosses was analyzed, mutant hyphae wereobserved, although they showed a more branched appearance withshorter cell compartments with lobed tips (Fig. 10B).

DiscussionHere, we showed that formation of an active bW-bE heterodimerin Ustilago maydis results in a sustained cell cycle arrest in G2phase. This cell cycle arrest is a consequence of the accumulationof phosphorylated inactive forms of Cdk1, the mitotic CDK in thisfungus. Inability to phosphorylate this protein, either bydownregulation of the kinase Wee1, or by expression of a Cdk1allele that is refractory to inhibitory phosphorylation, resulted infilaments that were not arrested in the cell cycle. We tried todetermine which molecular mechanisms were responsible for suchaccumulation of phosphorylated forms of Cdk1. Taken together,our results support the idea that upon induction of b-factor genes,the prevention of nuclear accumulation of Cdc25 by interaction with14-3-3, which is helped by the upregulation of wee1, results in anincrease in the inhibitory phosphorylation of Cdk1, which inhibitsthe G2-M transition during the formation of infective hyphae.

Fig. 7. Cytoplasmic retention of Cdc25 is required for b-factor-induced cell cycle arrest in U. maydis. (A)Subcellular localization of Cdc25 in AB34 (non-compatible b-factor genes) and AB33 (compatible b-factor genes) derivate cells carrying an endogenous functional Cdc25-3GFP fusion. Cells were incubatedunder inducing conditions (MM-NO3) for 8 hours. Graph indicates the quantification of nucleus to cytoplasm ratio of GFP fluorescence signal. Quantification offluorescence signal was performed by measuring pixel intensity in the nuclei and in an equivalent area of the cytoplasm. Mean and s.d. are shown. (B)Schemeshowing the construction carried by the strain that express b proteins under conditions of down-regulation of the 14-3-3 Bmh1 protein. The bmh1crg1 allele isrepressed in the presence of glucose and transcribed when arabinose was the only carbon source. (C)Schematic representation of the cdc25AAA allele showing theamino acid changes introduced to void the 14-3-3 binding sites. The phosphatase catalytic domain (PPase) and the putative nuclear localization signal (NLS) ofCdc25 are also showed. (D)Cell images of AB33 (control) and AB33-derived strain carrying either a conditional bmh1crg1 allele (bmh1OFF) or the cdc25AAA allelethat has an impaired interaction with the 14-3-3 protein Bmh1 (cdc25AAA), after 8 hours of incubation in induction conditions for expression of b-factor genes (MM-NO3, bmh1crg1 was repressed in these conditions because the carbon source was glucose). Cells were stained with DAPI to detect nuclei.

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A surprising result was the realization that Chk1, a well-knownregulator involved in the DNA-damage response, has roles in thiscell cycle arrest. Furthermore, we found that Chk1 was transientlyactivated during formation of the infective hypha. Transientactivation of Chk1 is compatible with the proposed effect on thecell cycle of these checkpoint systems, which appear to be moredevoted to induce a transient arrest (thus providing time to solvethe problems) rather than a permanent arrest (Toettcher et al., 2009).It could also be that additional elements are required to sustain theobserved cell cycle arrest, and that Chk1 activation is just the trigger.It is unclear how the expression of b-factor genes is linked to Chk1activation. We found no increase in either mRNA or protein levelsof Chk1 upon b-factor induction (not shown). As Chk1 activationis linked to DNA damage in vegetative cells (Perez-Martin, 2009),we analyzed whether there was DNA damage associated with theinduction of the infective hyphae by looking for the formation ofRad51 foci as a reporter (Kojic et al., 2008) (supplementarymaterial Fig. S2). We found no proof of massive DNA damage (i.e.

no accumulation of Rad51 foci were observed); however, we cannotdiscard the idea that limited DNA damage is responsible for Chk1activation. It is interesting to note that one of the first genes describedin which transcription was directly activated by b-proteins encodeda putative DNA polymerase-, which is one member of the X familyof DNA polymerases that are involved in a number of DNA repairprocesses (Brachmann et al., 2001; Ramadan et al., 2004).Unfortunately, no role has been determined so far for this factor inU. maydis.

An interesting question that has not yet been addressed is thereason for such a cell cycle arrest. We believe that this specific cellcycle arrest in U. maydis has a mechanistic role. In U. maydis, theG2 phase is characterized by polar formation of a bud, whichrequires the rearrangement of the cytoskeleton and involvesspecialized sets of motors, such as cytoplasmic dynein (Steinberget al., 2001; Straube et al., 2001), which support the polar extensionof the cell. In other words, in the G2 phase, the cytoskeletal growthmachinery is set up to support polar growth. Assuming that the

Fig. 8. Chk1 is required for b-factor-induced cell cycle arrest. (A)Cell images of AB33 (control) and a derivate strain lacking the chk1 gene incubated for 8 hoursin inducing conditions (MM-NO3). Cells were stained with DAPI to detect nuclei. (B)Percentage of cells producing mononucleated filaments (i.e. cell-cyclearrested) after 24 hours of incubation; means ± s.d. are shown. (C)Scheme of the formation of white filamentous dikaryon. a1b1 cells are indicated in white, a2b2cells are indicated in dark gray, and the dikaryon is shown in light gray. Filamentous growth is indicated by thin lines rising from the colony. (D)Crosses of controlstrains FB1 x FB2 and chk1 mutant strains in charcoal-containing agar plates. Note the gray appearance of mutant cross indicating impairment in filamentformation. (E)Compatible strains carrying a NLS-GFP fusion under control of the b-factor-dependent dik6 promoter were scrapped from agar surface, mounted onmicroscopy slides and epifluorescence was observed. Upper image shows DIC images of cells and bottom image shows fluorescence in GFP channel.(F)Quantification of the nuclear content of filaments obtained from charcoal plates.

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formation of the infective hyphae is based on a similar mechanismas polar bud growth, a prolonged G2 phase is best suited to supporttip growth during this stage.

An additional interesting question that emanates from our resultsconcerns the additional roles that Chk1 might have during thepathogenic process, when the fungus is growing inside the plant.A simple answer would be related to the ability of fungal cells todeal with DNA damage occurring during the proliferation insidethe plant in response to the plant defense system (i.e. in the formof reactive oxygen species), because mutants defective in Chk1function were extremely sensitive to DNA-damage agents (Pérez-Martín, 2009). However, we believe that this is not the case, becausefungal cells defective in Brh2, the BRCA2 homolog, were able toinfect and complete the life cycle at similar levels as wild-type cells,despite being extremely sensitive to DNA-damage agents (Kojic etal., 2002). We favor alternative explanations related to regulationof the fungal cell cycle inside the plant. U. maydis proliferates insidethe plant as a dikaryotic hyphae. In most dikaryotic Basidiomycetes,the proper distribution of the two genetically distinct nuclei during

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mitotic cell division is ensured by the formation of auxiliary clampcells, where one nucleus is entrapped and therefore mitosis occursin two distinct cell compartments (Casselton, 2002). In this particularmanner of division, an accurate control of the cell cycle is predicted.Interestingly, work performed in Coprinopsis cinerea andSchizophyllum commune indicated a role of homeoproteins fromthe b-factor family in this process (Brown and Casselton, 2001). Itis tempting to speculate that Chk1 has a role in this process; itsabsence might therefore affect the ability of the dikaryotic cells todivide properly and therefore proliferation might be affected.

It has become increasingly clear that elements from the DNA-damage response cascade can be used, even in the absence ofapparent DNA damage, to modulate cell cycle progression duringdevelopmental processes such as midblastula transition inDrosophila embryos (Sibon et al., 1997) or in the asynchronousdivision at two-cell-stage C. elegans embryos (Brauchle et al., 2003).The surprising finding that a protein involved in DNA damageresponses has a role in a fungal developmental process mirrors theseprevious results. In addition, our results reinforce the emerging idea

Fig. 9. Chk1 is transiently activated during b-factor filamentation. (A)Chk1 islocalized transiently in the nucleus during b-factor-dependent filamentation.Images of an AB33-derived strain carrying a Chk1-3GFP fusion grown inMM-NO3 for 3 (top right panel), 6 (middle right panel) and 9 hours (bottomright panel). Control strain AB34 carrying a Chk1-3GFP fusion was grown inMM-NO3 for 3 hours (left panel). All cells are shown at the samemagnification. (B)Distribution of cells showing nuclear GFP accumulation infunction of cell length. Quantification is the result of measurement of twoindependent experiments, counting 80 cells in each. Means ± s.d. are shown.(C)In vivo phosphorylation of Chk1 during b-factor-dependent filamentation.AB33- and AB34-derived cells carrying an endogenous Chk1-3�Myc allelewere incubated in MM-NO3 for the indicated time (in hours). Protein extractswere immunoprecipitated with a commercial anti-Myc antibody andimmunoprecipitates were subjected to SDS-PAGE and immunoblotted withanti-Myc antibody.

Fig. 10. Virulence of chk1 cells. (A)Quantification of symptoms in maizeplants after infection with wild-type (control) or chk1 mutant crosses.(B)Images of Chlorazole-Black-E-stained control and chk1 hyphae growinginside the plant tissue 1 week after infection. Scale bar: 15m. (C)Images ofinfected plants. Images show typical large tumors found in wild-type crossesand the small tumors found in mutant crosses (left). When male flowers wereinfected, wild-type crosses produce hypertrofied anthers, whereas limiteddeformation is observed in mutant infections (right).

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that checkpoint kinases might have roles other than in the DNA-damage response, since they can interact with the cell cyclemachinery. This has been proved by the involvement of Chk2-likekinases in the pathway linking circadian and cell cycles, in bothmammals and fungi (Gery et al., 2006; Pregueiro et al., 2006).

Materials and MethodsStrains and growth conditionsUstilago maydis strains are listed in supplementary material Table S1 and are derivedfrom FB1 and FB2 backgrounds (Banuett and Herskowitz, 1989). Media wereprepared as described (Holliday, 1974). Controlled expression of genes under thecrg1 and nar1 promoters was performed as described previously (Brachmann et al.,2001; Garcia-Muse et al., 2004). FACS analyses were described previously (Garcia-Muse et al., 2003).

DNA, RNA and protein analysisU. maydis DNA isolation was performed as previously described (Tsukuda et al.,1988). RNA isolation and northern analysis were performed as described previously(Alvarez-Tabares and Perez-Martin, 2008; Castillo-Lluva and Perez-Martin, 2005).Protein extraction and western blotting were also performed as described previously(Alvarez-Tabares and Perez-Martin, 2008; Garrido et al., 2004; Pérez-Martín, 2009).To purify Cdk1 complexes and analyze their kinase activity, previously describedprotocols were followed (Garcia-Muse et al., 2004). All quantification was done usinga Phosphorimager (Molecular Dynamics). To detect the phosphorylated and non-phosphorylated forms of Cdk1, commercial antibodies were used, as described(Sgarlata and Perez-Martin, 2005a). Primary antibody was followed by a secondaryanti-rabbit antibody conjugated to horseradish peroxidase and immunoreactiveproteins were visualized using a chemiluminescent substrate. The chemiluminescentsignal was analyzed using ChemiDoc XCS+ (Molecular Imager, Bio-Rad).

Plasmid and strain constructionsTo construct the different strains, transformation of U. maydis protoplasts with theindicated constructions was performed as described previously (Tsukuda et al., 1988).All fluorescent protein fusions were already described: GFP-Tub1 (Steinberg et al.,2001); Cut11-RFP (Perez-Martin, 2009); Cdc25-3GFP (Mielnichuk and Perez-Martin, 2008); Chk1-3GFP (Perez-Martin, 2009); GFP-Rad51 (Kojic et al., 2008) (agift from William K. Holloman, Cornell University, NY). To express cdk1 allelesunder nar1 promoter control and wee1 under nar1 or scp promoter control, previouslydescribed vectors were used (Sgarlata and Perez-Martin, 2005a). Plasmids to insertthe cdc25AAA allele and the conditional bmh1crg1 allele have been described(Mielnichuk and Perez-Martin, 2008). Disruption and tagging of chk1 was performedas described (Perez-Martin, 2009).

To express cdc25 under dik6 promoter control, we exchanged the crg1 promoterfrom pRU11-Cdc25 plasmid (Sgarlata and Perez-Martin, 2005b) with the dik6promoter from the plasmid pCLB1dik6 (Flor-Parra et al., 2006) resulting in the pDik6-Cdc25 plasmid. To express NLS-GFP under dik6 promoter control, the otef2 promoterfrom plasmid pnGFP (Straube et al., 2001) (a gift from Gero Steinberg, Universityof Exeter, Exeter, UK) was exchanged with the dik6 promoter from the plasmidpCLB1dik6 (Flor-Parra et al., 2006) resulting in the plasmid pDik6-NLSGFP.

MicroscopySamples were mounted on microscope slides and visualized in a Nikon eclipse 90imicroscope equipped with a Hamamatsu ORCA-ER CCD camera. All the images inthis study are single planes. Standard DAPI, GFP and Rhodamine filter sets wereused for epifluorescence analysis. The software used with the microscope wasMetaMorph 7.1 (Universal Imaging, Downingtown, PA). Images were furtherprocessed with Adobe Photoshop 8.0.

To quantify the ratio of the nuclear intensity (N) to the cytoplasmic intensity (C)of Cdc25-GFP signal procedures already described were followed (Mielnichuk andPerez-Martin, 2008). Briefly, the intensity of the nuclear and cytoplasmic signal wasdetermined by measuring pixel intensity in the nucleus and of an equivalent area inthe cytoplasm, and the ratio was determined. 20 cells were quantified for eachexperiment. To quantify Rad51-GFP foci formation, we followed the proceduresdescribed previously (Kojic et al., 2008).

Infection assaysPlant infection and charcoal assays were described previously (Flor-Parra et al., 2007).Staining of infected plant samples with Chlorazole Black E was done as describedpreviously (Brachmann et al., 2003).

We thank W. K. Holloman (Cornell University, Ithaca, NY) and G.Steinberg (University of Exeter, Exeter, UK) for providing plasmids.Prof. Holloman is also thanked for critical reading and inspiringdiscussions. N.M. was supported by JAE. This work was supported bya Grant from the Spanish Government (BIO2008-04054).

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