During induction of the virulence program in the phytopathogenic fungus Ustilago maydis, the cell cycle is arrested on the plant surface and it is not resumed until the fungus enters the plant. The mechanism of this cell cycle arrest is unknown, but it is thought that it is necessary for the correct implementation of the virulence program. Here, we show that this arrest takes place in the G2 phase, as a result of an increase in the inhibitory phosphorylation of the catalytic subunit of the mitotic cyclin-dependent kinase Cdk1. Sequestration in the cytoplasm of the Cdc25 phosphatase seems to be one of the reasons for the increase in inhibitory phosphorylation. Strikingly, we also report the DNA-damage checkpoint kinase Chk1 appears to be involved in this process. Our results support the emerging idea that checkpoint kinases have roles other than in the DNA-damage response, by virtue of their ability to interact with the cell cycle machinery.
Developmental decisions often involve differentiation processes that need to reset the cell cycle for induction of a new morphogenetic program. This is probably also the case for induction of the virulence program in pathogenic fungi. It could therefore be assumed that in pathogenic fungi the control of the cell cycle, as well as morphogenesis, is linked somehow to the virulence program. The maize smut fungus Ustilago maydis is an excellent system to address the relationships between cell cycle, morphogenesis and pathogenicity (Perez-Martin et al., 2006; Steinberg and Perez-Martin, 2008). During the induction of virulence program in this fungus, an infectious dikaryotic hypha is produced as a result of the mating of a pair of compatible haploid budding cells. Therefore, the induction of the pathogenic program implies not only strong morphological changes (bud to hypha transition) but also genetic changes (haploid to dikaryotic transition). Accurate control of the cell cycle and morphogenesis is predicted during these transitions.
The formation of the infectious hypha in U. maydis depends on an intricate transcriptional program which primarily involves a transcriptional regulator called b-factor (Feldbrugge et al., 2004). The production of this master regulator is linked to the mating process that, after cell fusion, leads to the interaction of the two subunits composing the b-factor (bW and bE), one provided by each mating partner. This way, the mating of two compatible cells (i.e. carrying b-subunits able to dimerize) results in the formation of a dikaryotic cell, which undergoes a strong polar growth as well as an apparent cell cycle arrest on the plant surface. Eventually, the infectious hypha manages to enter the plant tissue, where it starts to proliferate, maintaining its dikaryotic status. The reasons for the apparent cell cycle arrest on the plant surface are unknown. It has been hypothesized that such a cell cycle adjustment would be required for a precise execution of the virulence program (Perez-Martin et al., 2006). In fact, the apparent cell cycle arrest of the infectious hypha on the plant surface observed in U. maydis seems to be more general, and it is also present in rust fungi such as Uromyces phaseoli (Heath and Heath, 1979).
Ectopic expression of compatible b-subunits in a haploid cell induces the formation of a monokaryotic filament that mimics its dikaryotic counterpart in all aspects of filamentous growth as well as apparent cell cycle arrest (Brachmann et al., 2001). Which mechanisms are used by the bW-bE heterodimer to induce the hyperpolarized growth and to arrest the cell cycle in the infectious hypha and at which stage this occurs are unknown, but these issues are currently being studied. For example, recent results from our laboratory indicated that the activation of hyperpolarized growth after bW-bE heterodimer formation is dependent on the U. maydis ortholog of the cyclin-dependent kinase Cdk5, a well-known regulator of neuron development in mammals (Alvarez-Tabares and Perez-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 takes place in the G2 phase as a consequence of an increase of inhibitory phosphorylation of the mitotic cyclin-dependent kinase. Interestingly, we also report that the DNA-damage checkpoint kinase Chk1 seems to be involved in this process.
G2 cell cycle arrest in U. maydis upon expression of compatible b-homeoproteins
To address the molecular mechanisms of the b-factor-dependent cell cycle arrest, we made use of the previously described (Brachmann et al., 2001) haploid U. maydis AB33 strain, which carry compatible (i.e. able to dimerize) bE and bW genes under the control of the regulatable nar1 promoter. As the nar1 promoter can be induced by growing cells in nitrate-containing medium (MM-NO3) and repressed in ammonium-containing medium (MM-NH4), b-factor-dependent infective filament formation can be elicited in AB33 by changing the nitrogen source. As a control we used a similar strain, AB34, which harbors incompatible bE and bW genes (i.e. unable to produce dimers) under the control of nar1 promoter (Brachmann et al., 2001) (Fig. 1A).
We measured the DNA content of these strains under induction and non-induction conditions using fluorescence-activated cell sorting (FACS) analysis and found that cells expressing compatible b-proteins accumulated with a 2C DNA content (Fig. 1B). To specifically ascribe the cell cycle stage at which arrest takes place, we introduced two cytological markers that helped us to distinguish G2 from early mitosis in a strain expressing compatible b-proteins: an α-tubulin-GFP protein fusion allows the detection of the microtubule cytoskeleton (Steinberg et al., 2001) and a Cut11-RFP fusion protein labels the nuclear envelope (Pérez-Martín, 2009). During the G2 phase, the nuclear envelope is present and the microtubules form a cytoplasmic network (Fig. 1C, inset arrow); however, once cells enter mitosis, the nuclear envelope is disassembled 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 nuclear envelope and a defined cytoplasmic array of microtubules (Fig. 1C, arrows), all of which, together with the 2C DNA content, is consistent with a G2 cell cycle arrest during the formation of the infective filament.
G2 cell cycle arrest is dependent on the inhibitory phosphorylation of Cdk1
G2-M transition in U. maydis is enabled by the activity of two CDK complexes: Cdk1-Clb1 and Cdk1-Clb2 (Garcia-Muse et al., 2004; Sgarlata and Perez-Martin, 2005a). To determine the kinase activity associated 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 high affinity (Ducommun and Beach, 1990) and that we previously proved to be a convenient assay for characterizing the U. maydis Cdk1 protein (Garcia-Muse et al., 2004). Cell lysates obtained from AB33 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 for histone H1 kinase activity (Fig. 2A). Consistently with the inability of the hyphae to enter mitosis, Cdk1-associated kinase activity was downregulated by more than 90% after induction of the expression of compatible b-factor genes (Fig. 2B).
Downregulation of CDK activity can be produced in different ways in U. maydis (Perez-Martin et al., 2006), one of these, which is particularly active during the control of G2-M transition, is the phosphorylation of the Tyr15 residue in Cdk1 (Sgarlata and Perez-Martin, 2005a). Therefore, we analyzed levels of Cdk1 inhibitory phosphorylation upon induction of b-factor genes. For this, we used a specific antibody raised against the phosphorylated human Cdc2-Y15-P peptide, which recognizes the Tyr15-phosphorylated form of U. maydis Cdk1 (Sgarlata and Perez-Martin, 2005a). Tyr15P-Cdk1 levels increased eightfold when compatible b-proteins were expressed (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 are replaced with residues that cannot be phosphorylated (Thr14 to Ala and Tyr15 to Phe, respectively) (Sgarlata and Perez-Martin, 2005a). As continuous expression of this allele produces a deleterious effect in the cell (Sgarlata and Perez-Martin, 2005a), we decided to introduce an ectopic copy of this mutant allele under the same promoter 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 as the cell produces b-proteins. As a control, we constructed an AB33 strain carrying an ectopic copy of a wild-type cdk1 allele under control of the nar1 promoter. We used 3×Myc-tagged versions of both the wild-type and mutant ectopic Cdk1 to discriminate between the levels of protein produced by the endogenous locus and the ectopically expressed alleles (Fig. 3A). We found that in the strain expressing the Cdk1 version refractory to inhibitory phosphorylation (AB33 cdk1AF), the Cdk1-associated kinase activity remained high. By contrast, in AB33 or in the control strain expressing a Cdk1 version that can be phosphorylated (AB33cdk1), the kinase activity was downregulated after b-factor expression (Fig. 3B). Consistently, in the presence of Cdk1AF mutant protein, the b-factor-dependent filaments carried several nuclei, indicating that no cell cycle arrest was taking place (Fig. 3C,D). Expression of wild-type Cdk1 produced filaments that were indistinguishable from those obtained with the AB33 control strain. Taken together, these results indicate that inhibitory phosphorylation of Cdk1 has a major role during the b-factor-induced G2 arrest in U. maydis.
Cdk1 inhibitory phosphorylation during b-factor-induced cell cycle arrest depends on Wee1
Cdk1 inhibitory phosphorylation during vegetative growth depends on 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 described strain (AB31) (Brachmann et al., 2001) that expresses compatible b-factor genes under control of the crg1 promoter, which is induced in growth medium containing arabinose as a carbon source and repressed when glucose is present (Bottin et al., 1996). We introduced the conditional allele wee1nar1 in this strain (Fig. 4A) (Sgarlata and Perez-Martin, 2005a). This allele enables the expression of wee1 in cells incubated in minimal medium plus nitrate as a nitrogen source (MM-NO3) as well as the downregulation of wee1 expression in complete medium (CM, Fig. 4B).
When AB31-derived wee1 conditional cells were grown in repressive conditions (CM) and in the presence of arabinose, multinucleated filaments were observed (Fig. 4C). In similar conditions, AB31 cells had a single nucleus, indicating that Wee1 is necessary for b-factor-induced cell cycle arrest. Consistently, the inability to arrest the cell cycle in the absence of Wee1 correlated with a dramatic decrease in the level of Tyr15-P Cdk1, as well as a high level of Cdk1-associated kinase activity (Fig. 4D,E).
b-factor-dependent cell cycle arrest is not solely mediated by transcriptional regulation of wee1
We analyzed the levels of wee1 mRNA in AB33 cells after b-factor induction and found a sevenfold increase with respect to the AB34 control strain (Fig. 5A and supplementary material Fig. S1). Since overexpression of wee1 produces a G2 cell cycle arrest (Sgarlata and Perez-Martin, 2005a), we wondered whether this upregulation was responsible for b-factor-induced cell cycle arrest. To answer this question, we exchanged the endogenous wee1 native promoter with the constitutively expressed scp promoter in AB33 cells (Fig. 5B). This mutant allele, wee1scp, produces a low and constitutive level of wee1 mRNA without any appreciable defect in cell cycle regulation in normal conditions (Sgarlata and Perez-Martin, 2005a). In AB33 wee1scp cells, wee1 mRNA level was maintained at a low level upon b-factor induction (Fig. 5C) However, we found a minor effect in cell cycle arrest: less than 25% of filaments showed more than one nucleus (Fig. 5D,E), indicating that the observed transcriptional upregulation of wee1 is not the only reason by which cell cycle arrests during b-factor induction.
Ectopic cdc25 overexpression impairs the b-factor-induced cell cycle arrest
The level of inhibitory phosphorylation of Cdk1 also depends on the activity of the Cdc25 phosphatase, which reverses the CDK inhibition (Sgarlata and Perez-Martin, 2005b). Therefore, we analyzed the levels of cdc25 mRNA during b-factor-induced filamentation. However, we found that it was not affected by the presence of an active b-heterodimer (Fig. 6A). In spite of this result, we wondered whether it was possible to release the cell cycle arrest by overexpressing cdc25. To achieve this, we introduced in a strain carrying compatible b-factor genes under control of the nar1 promoter (AB33 cells), an ectopic copy of cdc25 under the control of the dik6 promoter, which is strongly induced by the b-proteins (Brachmann et al., 2001) (Fig. 6B). In the resulting strain, induction of b-factor genes resulted in a dramatic increase in cdc25 mRNA levels (Fig. 6C). Interestingly, the filaments expressing high levels of cdc25 had more than one nucleus in a substantial proportion (64% of the filaments) (Fig. 6D). Furthermore, the cells expressing cdc25 under control of the dik6 promoter showed a low level of inhibitory phosphorylation of Cdk1 (Fig. 6E).
Altogether, the results obtained with Cdc25 and Wee1 indicated that a similar regulatory scheme controlling G2-M transition via CDK inhibitory phosphorylation during vegetative growth is also involved in b-factor-induced cell cycle arrest.
Cdc25 seems to be retained in the cytoplasm during b-factor-induced filamentation
The observation that overexpression of cdc25 resulted in a multinucleated filament can be explained if b-factor-induced cell cycle arrest relies on an inhibition of Cdc25. One way to control Cdc25 activity in U. maydis cells is to regulate its subcellular localization (Mielnichuk and Perez-Martin, 2008). We explored this possibility by monitoring Cdc25 nucleus to cytoplasm distribution in cells expressing GFP-tagged Cdc25 from the genomic locus. To quantitatively evaluate changes in localization, fluorescence intensities of the Cdc25 signal were determined for a circular region corresponding to the nucleus (N) and a comparable area in the cytoplasm (C), as described (Mielnichuk and Perez-Martin, 2008). We found that in inductive conditions (MM-NO3) the ratio N/C in control cells (AB34 cells) was about 1.1, whereas this ratio in AB33 cells was around 0.2 (Fig. 7A).
Previous work from our laboratory showed that retention of Cdc25 in the cytoplasm of U. maydis depends on the essential 14-3-3 protein Bmh1 (Mielnichuk and Perez-Martin, 2008). To address whether 14-3-3 was required for b-factor-induced cell cycle arrest, we expressed compatible b-factor genes in conditions of downregulation of bmh1 expression (Fig. 7B). We found that in these conditions, morphologically aberrant filaments were produced that contained several nuclei each (Fig. 7D, middle panel). Since 14-3-3 proteins are extremely pleiotropic (van Hemert et al., 2001), it is hard to unequivocally ascribe the effects observed in filamentation after bmh1 downregulation to the inability to arrest cell cycle. To link this directly to a defect in the ability to retain Cdc25 in the cytoplasm, we took advantage of a cdc25 mutant allele (cdc25AAA, Fig. 7C), which was impaired in the interaction with Bmh1 and therefore accumulates in the cell nucleus (Mielnichuk and Perez-Martin, 2008). We found that a strain carrying the cdc25AAA allele and expressing compatible b-proteins resulted in a high percentage of filaments carrying more than one nucleus (Fig. 7D, bottom panel). Collectively, these data suggest that custody of Cdc25 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 arrest
Cytoplasmic retention of Cdc25 by 14-3-3 proteins requires the previous phosphorylation of target sites in Cdc25 (van Hemert et al., 2001). The DNA-damage checkpoint kinase Chk1 is one of the described kinases able to phosphorylate Cdc25 (Karlsson-Rosenthal and Millar, 2006). Interestingly, the mutations carried in the cdc25AAA allele affect the ability of the DNA-damage checkpoint kinase 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 in Chk1 and expressing compatible b-proteins were impaired in their ability to arrest the cell cycle (Fig. 8A,B).
Even when the b-factor-dependent filament produced in AB33 cells mimics its dikaryotic counterpart in all aspects of filamentous growth, we sought to analyze the consequences of the absence of chk1 in the formation of the infective filament in more `native' conditions. Crosses of compatible haploid wild-type cells on charcoal-containing plates (Banuett and Herskowitz, 1989) resulted in the formation of cell-cycle-arrested dikaryotic hyphae (Flor-Parra et al., 2006). Therefore, we crossed compatible wild-type and Δchk1 strains on charcoal-containing plates. Wild-type crosses led to a filamentous appearance of the colony, whereas Δchk1 mutants were strongly attenuated in filament formation (Fig. 8C,D). We also analyzed the nuclear composition of filaments in these crosses. To distinguish the b-factor-induced filaments from the cell population background (frequently enriched in aberrant elongated cells) the haploid strains we used expressed a GFP fusion to a nuclear localization 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 two nuclei, whereas Δchk1 filaments frequently carried more than two nuclei (Fig. 8E,F), which was consistent with a defect in the ability to arrest entry into mitosis during the formation of the infective hyphae.
Chk1 is transiently activated during b-factor-induced cell cycle arrest
The above results indicated that during induction of the pathogenic development in U. maydis, a G2 cell cycle arrest takes place in which the checkpoint kinase Chk1 has a role. Chk1 is a protein that is produced in normal conditions in an inactive form, which is activated after phosphorylation by upstream kinases (Chen and Sanchez, 2004). Therefore, we examined whether Chk1 was activated during the formation of the b-factor-dependent filament. In U. maydis cells, as in other organisms, Chk1 activation can be easily monitored by the accumulation of a GFP-tagged Chk1 protein into the nucleus (Perez-Martin, 2009). When AB33-derived cells carrying 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 nucleus throughout the incubation period (Fig. 9A). Interestingly, AB33-derived long filaments seemed not to accumulate any nuclear GFP signal. To quantify this apparently transient response, we measured cells (n=80 in two independent experiments) of different length and plotted this against the presence or not of a nuclear GFP signal (Fig. 9B), confirming that only shorter filaments (i.e. early stages) showed nuclear accumulation of Chk1.
There are several possible explanations for this behavior. One possibility is that Chk1 is degraded at later stages. A different explanation is that even when the nuclear membrane is present during filament formation (see Fig. 1C), it becomes permeable at later stages. And finally, it is possible that Chk1 is transiently activated during b-factor-dependent filamentation. To address these possibilities, we monitored the decrease in the mobility of a Myc-tagged protein during electrophoresis as a surrogate marker for their phosphorylation-dependent activation (Perez-Martin, 2009). First, we observed that the overall levels of Chk1 were not affected by the incubation period, excluding degradation of Chk1 as a possible explanation (Fig. 9C). Interestingly, the electrophoretic shift was maximum after 3 hours of induction but decreased to negligible after 9 hours, supporting the explanation that Chk1 is transiently activated during the formation of the b-factor-dependent filament.
Chk1 is required for full virulence
U. maydis infection of maize results in anthocyanin pigment production by the plant and the formation of tumors that are filled with proliferating fungal cells, which eventually differentiate into black teliospores (Banuett and Herskowitz, 1996). To address the consequences of a defective cell cycle arrest during corn infection by U. maydis, we inoculated maize plants by stem injection with mixtures of wild-type and Δchk1 mutant strains. Both crosses were able to infect plants; however, the mutant strains were less efficient (42% of the plants showed no symptoms) and the severity of the symptoms was minor (Fig. 10A). Interestingly, plants infected with the Δchk1 mutant rarely showed large tumors (Fig. 10C), and even in these rare occasions, no teliospores were found, suggesting a role of Chk1 beyond the initial steps of infection (i.e. infective filament formation).
To address whether the mutant hyphae proliferate inside the plant, symptomatic leaves obtained from both mutant and wild-type crosses were sampled after 1 week of inoculation and Chlorazole Black E staining was used to visualize invading hyphae (Brachmann et al., 2003). Septated hyphae grown through epidermal cells were observed in plants inoculated with compatible wild-type strains. When material obtained from mutant crosses was analyzed, mutant hyphae were observed, although they showed a more branched appearance with shorter cell compartments with lobed tips (Fig. 10B).
Here, we showed that formation of an active bW-bE heterodimer in Ustilago maydis results in a sustained cell cycle arrest in G2 phase. This cell cycle arrest is a consequence of the accumulation of phosphorylated inactive forms of Cdk1, the mitotic CDK in this fungus. Inability to phosphorylate this protein, either by downregulation of the kinase Wee1, or by expression of a Cdk1 allele that is refractory to inhibitory phosphorylation, resulted in filaments that were not arrested in the cell cycle. We tried to determine which molecular mechanisms were responsible for such accumulation 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 with 14-3-3, which is helped by the upregulation of wee1, results in an increase in the inhibitory phosphorylation of Cdk1, which inhibits the G2-M transition during the formation of infective hyphae.
A surprising result was the realization that Chk1, a well-known regulator involved in the DNA-damage response, has roles in this cell cycle arrest. Furthermore, we found that Chk1 was transiently activated during formation of the infective hypha. Transient activation of Chk1 is compatible with the proposed effect on the cell cycle of these checkpoint systems, which appear to be more devoted to induce a transient arrest (thus providing time to solve the problems) rather than a permanent arrest (Toettcher et al., 2009). It could also be that additional elements are required to sustain the observed cell cycle arrest, and that Chk1 activation is just the trigger. It is unclear how the expression of b-factor genes is linked to Chk1 activation. We found no increase in either mRNA or protein levels of Chk1 upon b-factor induction (not shown). As Chk1 activation is linked to DNA damage in vegetative cells (Perez-Martin, 2009), we analyzed whether there was DNA damage associated with the induction of the infective hyphae by looking for the formation of Rad51 foci as a reporter (Kojic et al., 2008) (supplementary material Fig. S2). We found no proof of massive DNA damage (i.e. no accumulation of Rad51 foci were observed); however, we cannot discard the idea that limited DNA damage is responsible for Chk1 activation. It is interesting to note that one of the first genes described in which transcription was directly activated by b-proteins encoded a putative DNA polymerase-β, which is one member of the X family of DNA polymerases that are involved in a number of DNA repair processes (Brachmann et al., 2001; Ramadan et al., 2004). Unfortunately, no role has been determined so far for this factor in U. maydis.
An interesting question that has not yet been addressed is the reason for such a cell cycle arrest. We believe that this specific cell cycle arrest in U. maydis has a mechanistic role. In U. maydis, the G2 phase is characterized by polar formation of a bud, which requires the rearrangement of the cytoskeleton and involves specialized sets of motors, such as cytoplasmic dynein (Steinberg et al., 2001; Straube et al., 2001), which support the polar extension of the cell. In other words, in the G2 phase, the cytoskeletal growth machinery is set up to support polar growth. Assuming that the formation of the infective hyphae is based on a similar mechanism as polar bud growth, a prolonged G2 phase is best suited to support tip growth during this stage.
An additional interesting question that emanates from our results concerns the additional roles that Chk1 might have during the pathogenic process, when the fungus is growing inside the plant. A simple answer would be related to the ability of fungal cells to deal with DNA damage occurring during the proliferation inside the plant in response to the plant defense system (i.e. in the form of reactive oxygen species), because mutants defective in Chk1 function were extremely sensitive to DNA-damage agents (Pérez-Martín, 2009). However, we believe that this is not the case, because fungal cells defective in Brh2, the BRCA2 homolog, were able to infect and complete the life cycle at similar levels as wild-type cells, despite being extremely sensitive to DNA-damage agents (Kojic et al., 2002). We favor alternative explanations related to regulation of the fungal cell cycle inside the plant. U. maydis proliferates inside the plant as a dikaryotic hyphae. In most dikaryotic Basidiomycetes, the proper distribution of the two genetically distinct nuclei during mitotic cell division is ensured by the formation of auxiliary clamp cells, where one nucleus is entrapped and therefore mitosis occurs in two distinct cell compartments (Casselton, 2002). In this particular manner of division, an accurate control of the cell cycle is predicted. Interestingly, work performed in Coprinopsis cinerea and Schizophyllum commune indicated a role of homeoproteins from the b-factor family in this process (Brown and Casselton, 2001). It is tempting to speculate that Chk1 has a role in this process; its absence might therefore affect the ability of the dikaryotic cells to divide 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 of apparent DNA damage, to modulate cell cycle progression during developmental processes such as midblastula transition in Drosophila embryos (Sibon et al., 1997) or in the asynchronous division at two-cell-stage C. elegans embryos (Brauchle et al., 2003). The surprising finding that a protein involved in DNA damage responses has a role in a fungal developmental process mirrors these previous results. In addition, our results reinforce the emerging idea that checkpoint kinases might have roles other than in the DNA-damage response, since they can interact with the cell cycle machinery. This has been proved by the involvement of Chk2-like kinases in the pathway linking circadian and cell cycles, in both mammals and fungi (Gery et al., 2006; Pregueiro et al., 2006).
Materials and Methods
Strains and growth conditions
Ustilago maydis strains are listed in supplementary material Table S1 and are derived from FB1 and FB2 backgrounds (Banuett and Herskowitz, 1989). Media were prepared as described (Holliday, 1974). Controlled expression of genes under the crg1 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 analysis
U. 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 described protocols were followed (Garcia-Muse et al., 2004). All quantification was done using a 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 secondary anti-rabbit antibody conjugated to horseradish peroxidase and immunoreactive proteins were visualized using a chemiluminescent substrate. The chemiluminescent signal was analyzed using ChemiDoc XCS+ (Molecular Imager, Bio-Rad).
Plasmid and strain constructions
To construct the different strains, transformation of U. maydis protoplasts with the indicated 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) (a gift from William K. Holloman, Cornell University, NY). To express cdk1 alleles under nar1 promoter control and wee1 under nar1 or scp promoter control, previously described vectors were used (Sgarlata and Perez-Martin, 2005a). Plasmids to insert the cdc25AAA allele and the conditional bmh1crg1 allele have been described (Mielnichuk and Perez-Martin, 2008). Disruption and tagging of chk1 was performed as described (Perez-Martin, 2009).
To express cdc25 under dik6 promoter control, we exchanged the crg1 promoter from pRU11-Cdc25 plasmid (Sgarlata and Perez-Martin, 2005b) with the dik6 promoter 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 promoter from plasmid pnGFP (Straube et al., 2001) (a gift from Gero Steinberg, University of Exeter, Exeter, UK) was exchanged with the dik6 promoter from the plasmid pCLB1dik6 (Flor-Parra et al., 2006) resulting in the plasmid pDik6-NLSGFP.
Samples were mounted on microscope slides and visualized in a Nikon eclipse 90i microscope equipped with a Hamamatsu ORCA-ER CCD camera. All the images in this study are single planes. Standard DAPI, GFP and Rhodamine filter sets were used for epifluorescence analysis. The software used with the microscope was MetaMorph 7.1 (Universal Imaging, Downingtown, PA). Images were further processed 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 and Perez-Martin, 2008). Briefly, the intensity of the nuclear and cytoplasmic signal was determined by measuring pixel intensity in the nucleus and of an equivalent area in the cytoplasm, and the ratio was determined. 20 cells were quantified for each experiment. To quantify Rad51-GFP foci formation, we followed the procedures described previously (Kojic et al., 2008).
Plant infection and charcoal assays were described previously (Flor-Parra et al., 2007). Staining of infected plant samples with Chlorazole Black E was done as described previously (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 inspiring discussions. N.M. was supported by JAE. This work was supported by a Grant from the Spanish Government (BIO2008-04054).