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Timely cytokinesis/septation is essential for hyphal growth and conidiation in Aspergillus nidulans. Genetic analyses have identified that A. nidulans has components of the septum initiation network (SIN) pathway; one of these, SEPH, is a key player for early events during cytokinesis. However, little is known about how the SEPH kinase cascade is regulated by other components. Here, we demonstrate that the phosphoribosyl pyrophosphate synthetase family acts antagonistically against the SIN so that the downregulation of AnPRS family can bypass the requirements of the SIN for septum formation and conidiation. The transcription defect of the Anprs gene family accompanied with the reduction of AnPRS activity causes the formation of hyper-septation as well as the restoration of septation and conidiation in the absence of SEPH. Clearly, the timing and positioning of septation is related to AnPRS activity. Moreover, with the extensive yeast two-hybrid analysis and rescue combination experiments, it demonstrated that AnPRS members are able to form the heterodimers for functional interacting entities but they appear to contribute so unequally that Anprs1 mutant display relatively normal septation, but Anprs2 deletion is lethal. Thus, compared to in yeast, the AnPRS family may have a unique regulation mechanism during septation in filamentous fungi.
Cytokinesis is the process by which a cell splits its cytoplasm, accomplished by the contraction of a contractile actin ring, to produce two daughter cells. Accordingly, cytokinesis is the final step in cell division after the nuclear division of mitosis. For cell division to be successful, chromosome segregation, mitotic exit and cytokinesis must be executed in this order (Barr and Gruneberg, 2007; Sagona and Stenmark, 2010). Numerous studies have identified that mitotic exit requires the activation of the conserved signalling network, termed the mitotic exit network (MEN), in budding yeast and the septation initiation network (SIN) in fission yeast (Krapp et al., 2004; Barr and Gruneberg, 2007; Bedhomme et al., 2008; Brace et al., 2011; Meitinger et al., 2011). Obviously, to keep the co-ordination of mitosis and cytokinesis, it is crucial for the cell to carry out these events in the correct order and at the proper time. Although organisms of different kingdoms have developed unique mechanisms to execute cytokinesis, signals that trigger the onset of cytokinesis are evolutionarily conserved (McCollum and Gould, 2001; Baluska et al., 2006; Kim et al., 2006; Bedhomme et al., 2008). Thus, the use of simple eukaryotic microorganisms as a model will let us make rapid progress in understanding many cellular processes that are common in mammalian cells and plant cells. Unlike yeast, the filamentous fungus Aspergillus nidulans contains a mycelium of multinucleate cells that are partitioned by septa. During the germination in A. nidulans, the conidiospores undergo multiple rounds of nuclear division to produce eight or 16 nuclei in germlings, but they do not undergo septation until the cell reaches an appropriate size/volume, and then forms the first septum near the neck between spore and germ tube (Harris, 2001). Therefore, as a whole, inter-compartment development and mitosis in the mycelium becomes asynchronous (Harris, 2001). In addition, the septum does not subsequently disappear and daughter cells remain attached. Based on this characterization, the filamentous fungus A. nidulans is able to endure more defects in cytokinesis than single-cell yeast. Thus, A. nidulans is an excellent model organism for allowing an unambiguous identification of investigating the regulation features of cytokinesis (Gould and Simanis, 1997; Harris, 2001). Previous studies have reported that septum formation in A. nidulans requires the assembly of a septal band composed of a dynamic protein complex that is dependent upon a conserved protein kinase cascade (Harris, 2001; Kim et al., 2006). The serine/threonine protein kinase SEPH in A. nidulans, a Cdc7p orthologue from fission yeast, was first cloned in a screen for temperature-sensitive cytokinesis mutants. It has been identified that SEPH plays a central part in the initiation of septation prior to actin ring formation in A. nidulans (Bruno et al., 2001; Kim et al., 2006). Notably, downregulation of the SEPH kinase cascade would abolish septation, whereas hyper-activation would induce the formation of multiple septa (Bruno et al., 2001). Thus, SEPH is a positive regulator of the SIN which triggers cytokinesis in A. nidulans. However, little is known about how the SEPH kinase cascade is regulated by other components, or whether there exist any of the negative regulators that act antagonistically to others in the SIN. To gain insight into the regulatory mechanisms that underlie septation, 116 mutants that suppressed the defects of sepH in A. nidulans were isolated by UV mutation in this study. Furthermore, we found that the defects of the SEPH kinase cascade can be suppressed by the reduction of phosphoribosyl pyrophosphate (PRPP) synthetase activity.
Isolation of mutants that suppress loss of function of SEPH during cytokinesis
Previous studies have identified that SEPH from A. nidulans, as a homologue of serine–threonine kinase Cdc7p in fission yeast, plays an important role during cytokinesis. To further understand the regulation mechanism in which SEPH is involved during cytokinesis, we sought the extragenic mutations that suppress the loss function of SEPH. According to a previous finding that septation defects of the SEPH mutant at a restrictive temperature of 42°C resulted in failed conidiation, the abolished conidiation will be linked to septation defects (Liu and Morris, 2000; Liu et al., 2003). Hence, we used conidiation as an indicator of septation capability. To visually screen for conidiation, a chartreuse colour (chaA1) in the GQ1 strain (sepH1, pyrG89, chaA1) was seen as a colour of conidiospores after mutagenesis (Fig. 1A). Among the 116 independent mutants that restored conidiation at a restrictive temperature of 42°C, three categories were classified based on colony phenotypes at both 30°C and 42°C respectively (Fig. 1B). Mutations in class I, to which most of mutations belonged, had a robust conidiation capacity at both temperatures. In class II, the colonies showed almost normal conidiation with a chartreuse colour at 30°C. Whereas, for an unknown reason, all mutants of class II at 42°C had light brown conidiospores. In class III, the colonies displayed similar colony phenotypes but had a reduced colony size compared to those in class II at 42°C; by contrast, at 30°C, class III had a significantly reduced growth rate compared to that of wild type. Theoretically, because 30°C is a permission temperature for sepH1, the defects at this temperature must be caused by UV-induced suppressor mutations. Since a mutation embedding a growth defect phenotype can be easily cloned, the strain S110 (suppressor of sepH, No. 110), which has a remarkable growth defect at 30°C, was selected for further study. To firstly determine whether the suppressor belonged to the intragenic or the extragenic mutations for sepH, S110 was crossed to the wild-type strain R153. Among them (n = 306), two kinds of genotypes in progeny were the same as in the parent strains GQ1 (sepH1, n = 69) and S110 (sepH1, s110; n = 79), and the other two types belonged to wild type (n = 84) and a new type referred to as sin110 (n = 74). Consequently, the ratio of the four different types of colonies in the progeny was about 1:1:1:1. This genetic analysis result suggests that the mutation site of S110 belongs to the extragenic mutation, which must be located at a genetic locus other than the sepH region. To further examine the phenotype caused by this mutation, the spores of Sin110, S110 and wild type were inoculated and cultured at 30 and 42°C separately (Fig. 1C). S110 and Sin110 strains showed slightly reduced colony size compared to the control R153 strain but both exhibited conidiation at 42°C. Notably, it seems that Sin110 produced more spores than did S110, indicating that the mutation in Sin110 caused an antagonistical phenotype compared to sepH mutant.
Sin110 mutation restored septation as well as conidiation in the absence of SEPH
To further detect whether the restored conidiation was concomitant with the restoration of septation, we observed the mycelium phenotypes of GQ1, S110 and Sin110 stained with 4,6-diamidino-2-phenylindole (DAPI) and Calcofluor white (CFW). Upon the germination of the conidium, the formation of septum took place at the ‘neck’ site in very short germlings from both S110 and Sin110 strains at 30°C (Fig. 2C and E). In comparison, almost no detectable septa were found in similar size of wild-type germlings. In addition, three-nuclei basal cell was common to be seen in germlings in wild-type strain and the average nucleus number (n = 100) in wild type was 3.15 ± 0.79 in the basal cell. In comparison, in Sin110 strain, four- or five-nucleus basal cell was common to be seen (n = 100) and the average nucleus number was 4.04 ± 0.82 nuclei (n = 100), indicating disturbed nuclear positioning or the aberrant formation of delocalized septa. Normally, septum formation requires the assembly of a septal band, which is accomplished by the contraction of actin ring. A previous study has identified SEPH as a central player in the initiation of septation prior to actin ring formation in A. nidulans (Bruno et al., 2001). To further check if suppression by Sin110 also restores the actin rings that are absent in the sepH mutant, an immunostaining experiment for actin was carried out. As hypothesized, double mutants of sin110 and sepH were capable of developing a clear actin ring at the septum site (Fig. 2G and H).
Moreover, as opposed to the sepH1 mutant GQ1 with a completely stopped septation under the restrictive temperature conditions, Sin110 caused hyper-septation compared to that of S110. When cultured at 42°C, the growth-retarded phenotype in Sin110 had been relatively relieved than that cultured at 30°C but Sin110 mutant still had a reduced hyphal growth rate with a shorter germling tube than that of wild type. Consequently, the distance between septa in Sin110 was 11.76 ± 4.11 (μm) instead of 23.6 ± 3.21 (μm) in wild type under the same cultural medium (n = 140) so that Sin110 had almost twice number of septa than wild type did for the same length of hyphae but there was no significant difference in the number of nuclei/compartment between them. Thus, according to the average distance between septa, we concluded that Sin110 mutant had a hyper-septation phenotype.
Defect of Sin110 can be rescued by phosphoribosyl pyrophosphate synthetase 1 clone (Anprs1)
To further identify which mutant gene is involved in the aforementioned suppression of sepH in septation, a complementation test was performed using an A. nidulans genomic DNA library bearing an AMA1 plasmid vector (Aleksenko and Clutterbuck, 1997). As shown in Fig. 3A–C, one of the clones that can rescue the defect of Sin110 was obtained. After sequencing and blasting, this clone included part of AN6710.4 and the full length of AN6711.4 located on A. nidulans chromosome I from the genomic sequence 69087 to 73189 nt, which is referred to as strain T2. AN6711.4 encodes the putative protein phosphoribosyl pyrophosphate synthetase, called Prs1 in yeast, which is an important protein phosphatase that is present in several metabolic pathways (Hove-Jensen, 1988). To further confirm the function of the Prs1 homologue in A. nidulans (AnPRS1) as well as to eliminate the effect of the fragment in AN6710.4 during the complementation test, the open reading frame (ORF) of the Anprs1 gene was cloned to the AMA1 vector to make a plasmid called pAMA1-AnPRS1. After transformation to Sin110, AnPRS1 could consistently rescue the defects of Sin110 to some extent, suggesting that the expression of the extra copy of Anprs can partly rescue the defect of Sin110. Furthermore, using microscopy, as shown in Fig. 3D and E, relatively normal septation patterns were restored in strain T2, and even resulted in hyper-septation phenotype at both 30°C and 42°C. According to these complementation data, it is indicated that the defect of Sin110 is possibly related to the mutation of the Anprs1 gene. However, after three independent sequencing assays, the genomic sequence of a whole gene of Anprs1 in Sin110 mutant was exactly the same as that in the wild type (Fig. S3). We next wondered if Sin110 may cause mutation in other members of the Anprs family. Unexpectedly, no mutation was found in whole gene regions of Anprs2 or Anprs3 in the Sin110 mutant (Fig. S3). That suggests that the septation defect suppression of sepH was not due to the mutation of the Anprs gene family directly, but possibly to other reasons that affected the function of AnPRS.
Expression and characterization of a functional GFP–AnPRS1 fusion under the control of the alcA promoter
To further confirm and test how AnPRS1 functions in A. nidulans during cytokinesis, we used a conditional strain in which the Anprs1 gene was under the control of the inducible/repressible alcA promoter. As shown in Fig. 4A, homologous integration of the Anprs1 fragment in the plasmid into the genomic Anprs1 locus generated two copies of Anprs1: a truncated Anprs1 gene with its own promoter and a gfp–Anprs1 fusion, called ZGA01, under the control of the alcA promoter. As shown in Fig. 4B, the strain ZGA01 had the integration at the desired site. When ZGA01 was grown on a non-repressing medium (i.e. with glycerol as the sole carbon source), it displayed a number of phenotypic similarities to that of the wild type, including hyphal growth rate, colony size and septation (Fig. 4C). However, when grown on a repressing medium that contained glucose, ZGA01 induced the phenotype of hyper-septation in germlings or hyphae, accompanied with a slower growth rate compared to the control strain WJA01 (Fig. 4C, E and F), indicating that this conditional mutant produced a consistent phenotype with the sin110 mutation. Meanwhile, GFP–AnPRS1 exhibited a cellular localization pattern in mature cells. Occasionally, GFP–AnPRS1 appeared at the predicted septation site possibly prior to a detectable septum formation because in matured septa there was no detectable GFP–AnPRS1 accumulation found (Fig. 4D). This suggests that AnPRS1 may function in the cytosol and septation sites. Moreover, the micrograph examined by transmission electron microscopy (TEM) showed that, when the expression of AnPRS1 was turned off, ZGA01 showed the aberrant formation of delocalized septa compared to the wild type (Fig. 4G), which indicated that AnPRS1 may play an important role in the timing and positioning of septum formation.
Anprs1 full ORF deletion mutants display normal timing of cytokinesis
Based on the above growth phenotype of Anprs1 regulated by the alcA promoter under the repressed condition, we hypothesized that Anprs1 is not an essential gene in A. nidulans. We next made a full-length deletion mutant of Anprs1 (Fig. 5A and B) to further support our observations of the effect of AnPRS1 on septation in the conditional strain. Surprisingly, it seemed as if this whole-gene deletion mutant, referred to as ZGA02, did not show the detectable septation defect that was observed in the alcA(p)–Anprs1 conditional strain ZGA01 under the repressed condition (Fig. 5C and D). We wondered whether the truncated Anprs1 fragment in the conditional strain caused this difference. To test this possibility, a C-terminal truncated deletion variant of Anprs1 (Anprs1ΔC), which only had a truncated fragment of the same length of that in Anprs1 in the conditional strain ZGA01, was successfully made by DNA double joint homologue integration of fusion PCR products (Fig. 5E and F); we refer to this strain as ZGA03. Perhaps most interestingly, the Anprs1ΔC mutant caused a premature and hyper-septation phenotype (Fig. 5G) that was consistent with strains Sin110 and ZGA01 under the repressed condition. These data indicate that the existence of a truncated fragment of AnPRS1 may have the dominant negative function or have the competitive inhibition for the AnPRS family under normal circumstances. To test this hypothesis, we made another double mutant ZGA04 by crossing Anprs1ΔC mutant ZGA03 with sepH mutant GQ1. As a result, Anprs1ΔC mutant could not completely suppress the defect of sepH in septation. As shown in Fig. S4, after cultured in liquid medium at 42°C for 20 h, ZGA04 was unable to form the septum but showed some of more chitin desposition at the bases of branches than wild type had. It may indicate that the septation occurred but abolished during the development of the contractile ring to the mature septum formation, suggesting Anprs1ΔC had weaker ability to suppress sepH mutation than sin110 did.
The defects of Sin110 can be rescued by AnPRS members
According to all above results, there is a high probability that when the whole gene of Anprs1 was deleted, other members of the same family rescued the function. Genomic information analysis indicated that there were three members of Anprs in A. nidulans: Anprs1 (AN6711.4), Anprs2 (AN1965.4) and Anprs3 (AN3169.4). As previously reported, Prs's predicted function is to catalyse the biosynthesis of PRPP from ribose-5-phosphate and ATP, the whole process being called PRPP synthetase activity (Hove-Jensen, 2004). As such we suspect that the abnormal hyper-septation phenotypes displayed in Anprs1 mutant strains may have some connections with their PRPP synthetase activity. To prove our hypothesis, biochemical assays to test PRPP synthetase activity were performed according to standard protocols described in Experimental procedures. The results indicated that measurable PRPP synthetase activities in extracts from Sin110 and the alcA(p)–Anprs1 conditional strain (ZGA01) under the repressed condition, as well as from the Anprs1ΔC truncated deletion mutant (ZGA03), were substantially lower than that from the same background wild-type strain (WJA01) (Fig. 6A). Consequently, the relative AnPRS enzyme activities in mutant strains ZGA01 and ZGA03 were reduced to 67 ± 3% and 68 ± 5%, respectively, compared to the wild-type strain. Notably, Sin110 caused a mostly severe defect resulting in only 44 ± 5% of the retained relative AnPRS enzyme activity compared to the wild type. Accordingly, normal cytokinesis seems to depend on a normal level of PRPP synthetase activity. Because the above data have verified that no mutation was found in Anprs genes in Sin110, we wondered whether the decreased AnPRS activity in Sin110 was related to the transcription of the Anprs family. As expected, by real-time qPCR, three members of the Anprs family in A. nidulans were dramatically downregulated in Sin110 so that the mRNA levels of Anprs1, Anprs2 and Anprs3 were 12 ± 8%, 39 ± 5% and 28 ± 9%, respectively, compared to the wild-type strain WJA01 (Fig. 6B), suggesting that the mutation of sin110 was mostly related to the reduced transcription of the Anprs gene family. In response to these results, we cloned ORFs of Anprs1, Anprs2 and Anprs3 into the AMA1 vector to make the plasmids pAMA1-AnPRS1, pAMA1-AnPRS2 and pAMA1-AnPRS3 respectively. As shown in Fig. 6C, the colonies transformed by a single kind of Anprs plasmid exhibited only partially rescue of colony size and conidiation. Moreover, it seems that the rescue efficiency was different in the colonies transformed by Anprs1 alone, Anprs2 alone or Anprs3 alone. Among them, Anprs2 induced a stronger rescue than either of Anprs1 or Anprs3. In comparison, the defects of Sin110 in growth and septation can be dramatically rescued when transformed by a mixture of three different Anprs plasmids at DNA amount ratio 1:1:1, resulting in the transformants having almost the same colony size and conidiation as the wild type, indicating that the defects of cytokinesis in Sin110 can be completely rescued by the expression of the extra copies of the Anprs family. Thus, AnPRS1 may need AnPRS2 or AnPRS3 to form an isoenzyme complex to function. To confirm this hypothesis, next we constructed the different members of Anprs family into the Gal4 DNA binding domain and the GAL4 activation domain separately. All combinations (AnPRS1 and AnPRS2; AnPRS1 and AnPRS3; AnPRS2 and AnPRS3) showed a robust growth in high-stringency media (Fig. 6D), indicating that the reporter genes (histidine, adenine prototroph and beta-galactosidase) could be activated. None of those yeast cells transfected by single pGBKT7-Anprs or pGADKT7-Anprs showed growth under the high-stringency media, suggesting that none of the bait and prey plasmids had detectable auto-activation. These results demonstrate that AnPRS1, AnPRS2 and AnPRS3 are really able to form the heterodimeric complexes of AnPRS polypeptides.
The SIN, which includes the main component Cdc7p and the GTPase Spg1p, is emerging as a primary regulatory pathway to control cytokinesis in fission yeast Schizosaccharomyces pombe (Sawin, 2000; McCollum and Gould, 2001; Krapp and Simanis, 2008). In comparison, a functionally similar group of proteins comprise the MEN in budding yeast Saccharomyces cerevisiae (de Bettignies and Johnston, 2003; Brace et al., 2011; Meitinger et al., 2011). Meanwhile, several lines of evidence indicate that A. nidulans has components of the SIN–MEN pathway (sepH → sepL → sidB), one of which, sepH, is required for early events during cytokinesis (Bruno et al., 2001; Kim et al., 2009; Si et al., 2010). The cytokinesis in fungi can be viewed as a three-stage process: (i) selection of a division site, (ii) orderly assembly of protein complexes and (iii) dynamic events that lead to a constriction of the contractile ring and to septum construction (Simanis, 2003; Walther and Wendland, 2003). Nevertheless, in the filamentous fungus A. nidulans, previous studies have indicated that actin ring formation, as an initial event of cytokinesis, occurs after entering mitosis. SEPH, a putative Cdc7p orthologue, probably functions upstream of actin ring formation during cytokinesis. Thus, it suggests that possibly SEPH is required during early stages of septation. Further studies have indicated that SEPH is required for construction of the actin ring, and the deletion of sepH has been shown to result in a viable strain that fails to septate at any temperature (Bruno et al., 2001; Sharpless and Harris, 2002; Westfall and Momany, 2002). Therefore, SEPH, as a major factor in catalysing one of the initial events in cytokinesis, might have to work together with multiple other protein complexes to regulate cytokinesis. In this study, by using a combination of forward and reversed genetics techniques, the results presented here reveal that there exist protein regulators that act antagonistically towards components of the SIN in the filamentous fungus A. nidulans. Furthermore, we found that the defects of the SEPH kinase cascade in septation can be suppressed by the mutation of sin110 which is caused by the transcription defects of the Anprs gene family.
Suppressors of SEPH for septum formation in A. nidulans
Through UV mutagenesis, 116 independent mutants were obtained that could restore cytokinesis to a certain extent in the absence of sepH. Among them, three different classes were delineated based on colony phenotypes, but all showed septation capabilities to varying extents in 42°C (Fig. 1B). In class I, the mutants had a robust conidiation capacity in the absence of sepH, but at permission temperature 30°C they did not show the growth defect phenotype. Thus, they were difficult to clone. In comparison, class II and especially class III mutants could not only recover cytokinesis but they also displayed a slow-growth rate phenotype in the presence of sepH. This suggests that these genes may also have other functions than that of a negative regulator of cytokinesis; some (if any) of these functions may belong to the recovery mutation of sepH (i.e. the intragenic mutation of sepH). To exclude this possibility, the experiment of back-crossing mutants with wild type was carried out, and the results clearly indicate that most mutants had the extragenic mutation (Fig. S1). Notably, when sepH was defective, the functional mutations of these components could suppress the failed cytokinesis. Thus, our data clearly provide direct evidence for the existence of the antagonizing components of the SIN during cytokinesis. Moreover, a prior suppressor screen has identified smoA and smoB as suppressors of SIN mutations (Kim et al., 2006). Based on published information combined with ours, we found that both of sin110 and smoA/smoB mutations caused the reduced hyphal growth of colony and could suppress sepH mutations during conidiation, but the hyphal wavy morphology phenotype, along with the high temperature-sensitive characterization in smoA/smoB mutants, indicated that smoA and smoB could not belong to PRS family. Solving this puzzle needs the results for further cloning of smoA and smoB in the future.
The possible relationship between AnPRS and SEPH
It has been reported that in yeast, Prs functions not only in phospholipid metabolism but also in the MAPK-relied, cell wall integrity pathway by interacting with major components of this pathway (Vavassori et al., 2005). Additionally, Prs3 interacts with the yeast orthologue of GSK3 (glycogen synthase kinase 3) and Rim11, a serine/threonine kinase involved in several signalling pathways (Kleineidam et al., 2009). Most notably, our data (Figs 3 and 4) indicate that the decline of AnPRS1 expression suppresses the defect of sepH, and the reduction of PRPP synthetase activity causes both premature and hyper-septation, which is consistent with the phenotype of overexpression of SEPH in yeast (Bruno et al., 2001). These findings clearly show that the AnPRS family is involved in the function of co-regulating cytokinesis with SEPH in A. nidulans, which has not previously been reported. We propose a possible explanation of the relationship between AnPRS and SEPH during cytokinesis in A. nidulans as follows. Because predicted function of AnPRS is to catalyse the biosynthesis of PRPP from ribose-5-phosphate and ATP. However, SEPH, a serine–threonine protein kinase, also needs ATP to function. Thus, AnPRS1 and SEPH may competitively bind ATP, resulting in the proper timing of cytokinesis. In addition, there is the possibility that SEPH may act as a protein kinase and AnPRS1 as a phosphatase to regulate the phosphate and de-phosphate reactions of the same substrate (not just for ATP alone). Consequently, this would explain why low enzyme activity of AnPRS induces the hyper-septation phenotype as well as the suppression of the SEPH defect. In addition, in S. pombe, central players of SIN are cascaded kinases of GTPase Spg1p, Cdc7p, Sid1p and Sid2p (Krapp et al., 2004). Spg1p accumulates in its GTP bound form, which allows recruitment of the protein kinase Cdc7p. These kinases and their associated proteins exhibit important function to trigger septation (Sohrmann et al., 1998). To further answer whether the sin110 mutation specifically suppresses SEPH or other SIN-cascaded kinase mutations in A. nidulans, we deleted Spg1p homologue in A. nidulans (AN7206.4) in the background of sin110 mutant. As a result, double mutations showed a robust conidiation (G. Zhong and L. Lu, unpubl. data) and septation phenotype (Fig. S4). This suggests Sin110 was capable of suppressing mutations of other SIN components in addition to SEPH.
The AnPRS family may act as a heterodimer to exert the biological activity
Phosphoribosyl pyrophosphate synthetase (PRS) catalyses the biosynthesis of PRPP from ribose-5-phosphate and ATP (Khorana et al., 1958; Roessler et al., 1991). In the S. cerevisiae genome, there are five unlinked genes (PRS1–PRS5) capable of encoding the PRS enzyme (PRS; ATP: d-ribose-5-pyrophosphotransferase; EC 188.8.131.52). None of the PRS genes is essential, but the contributions of the PRS gene products do not appear to be equal in S. cerevisiae (Vavassori et al., 2005). Nevertheless, loss of either Prs1 or Prs3 has far-reaching consequences for metabolism, ranging from altered chitin synthesis and constitutive activation of the cell integrity pathway to an apparent disturbance in phospholipid metabolism. Moreover, an extensive yeast two-hybrid (Y2H) analysis and deletion combination experiments demonstrated that viable minimal subunits exist as two interacting functional entities (Prs1/Prs3, Prs2/Prs5 or Prs4/Prs5 in wild type) which seem to be capable of compensating for each other because, in the absence of one entity or one of its components, the yeast cells still survive (Hove-Jensen, 2004). This clearly suggests that Prs activity in S. cerevisiae is carried out by heterodimeric complexes of Prs polypeptides. Based on homologue analysis, we found three predicted PRS genes in A. nidulans: Anprs1 (AN6711.4), Anprs2 (AN3169.4) and Anprs3 (AN1965.4). According to the protein sequence analysis of identity and homology in alignment by DNAStar software, AnPRS1 is a putative homologue of yeast Prs1 whereas AnPRS3 is most likely a homologue of Prs5 (identity number 45.54%) and not of Prs3 (identity number 36.45%) as expected, in yeast. We hypothesized that AnPRS2 may have an important comprehensive function as a partner of Prs2, Prs3 and Prs4 in yeast. In fact, the deletion of Anprs2 in A. nidulans resulted in cell death, indicating that AnPRS2 may be a central subunit functioning as an interactive AnPRS heterodimer to exert its biology activity which is completely different from the homologue in budding yeast (Fig. S2). Nevertheless, our data (Fig. 5C and D) indicate that the whole Anprs1 gene deletion mutant ZGA02 did not show any detectable defects while a C-terminal truncated deletion variant of Anprs1 (Anprs1ΔC) ZGA03 caused defects in septation and AnPRS enzyme activity. Moreover, Anprs1ΔC had weaker ability to suppress sepH mutation than Sin110 did (Fig. S4) possibly due to the difference between Anprs1ΔC and sin110 mutants in retained AnPRS enzyme activity (Fig. 6). This suggests that the existence of a truncated fragment of Anprs1 may disturb the normal function of Anprs2 or Anprs3, which may function similarly to a dominant-negative mutant. Furthermore, Y2H experimental data in Fig. 6 demonstrate that AnPRS members in A. nidulans are able to form the heterodimers for interacting functional entities AnPRS1/AnPRS2, AnPRS2/AnPRS3 and AnPRS1/AnPRS3. Perhaps, in the absence of AnPRS1, functional AnPRS2/AnPRS3 may be capable of compensating for the function of the AnPRS1 complex with AnPRS2 or AnPRS3. In contrast, when a truncated Anprs1ΔC existed, it is able to induce a competitive inhibition with other members of AnPRS to form a functional AnPRS complex. Moreover, we found that the contributions of the Anprs gene products do not appear to be equal in A. nidulans, such that Anprs1 was not essential (Fig. 5C and D), but deletion of Anprs2 resulted in cell death (Fig. S2), indicating that the AnPRS family may have a unique mechanism in filamentous fungi that is completely different from single-cell yeast. Future studies on AnPRS function as a heterodimer should clarify the interaction details in the AnPRS family.
Strains, media, culture conditions, plasmids and transformation
A list of A. nidulans strains used in this study is provided in Supporting information (Table S1). The media MM (minimal media), YAG (yeast + agar + glucose media), YUU (YAG + uridine + uracil), YUUK (YUU + KCl) and MMGPR (MM + glycerol + pyrodoxine + riboflavin) are described in previous references (Kafer, 1977; Wang et al., 2009). MMGTPR: MMGPR with 6.25 mM threonine. Growth conditions, crosses and induction conditions for alcA(p)-driven expression were as previously described (Wang et al., 2006). Expression of tagged genes under the control of the alcA promoter was regulated by different carbon sources: repression on glucose, derepression on glycerol and induction on threonine. Standard DNA transformation procedures were used for A. nidulans (Osmani et al., 1988; May, 1989). The plasmids used are listed in Table S3.
Mutagenesis and screening for septation revertants
Strain GQ1 bearing the chaA1 mutation was used for mutagenesis. Approximately 107 conidia were suspended in 20 ml of sterile distilled water. They were irradiated with ultraviolet light at a dosage of 8000 mW cm−2 for 65 s with agitation using CL-1000 (Ultra-Violet Products, Unit1). The irradiation rendered 10% viability. Mutagenized spores were plated on the YUU medium. After being incubated at 42°C for 3 days, the colonies showing the chartreuse colour were picked as conidiating revertant candidates for further analysis.
Complementation test of the sin110 mutation
The complement test of sin110 was performed using the pRG3–AMA–NotI genomic DNA library as follows (Yelton et al., 1984; Osherov et al., 2000). Transformants were selected for restoration of pyrimidine prototrophy on the YAG medium. The isolates showing the rescue phenotypes of sin110 defects were selected for further study. Next, plasmids were extracted from these rescued Aspergillus clones and then transformed to competent Escherichia coli as previously described (Rasmussen et al., 1990). After purifying the plasmids from the positive clones, we retransformed these purified plasmids separately to the Sin110 strain to confirm the rescued phenotypes. Then, the genomic insert in pRG3–AMA–NotI was end-sequenced using vector-specific primers and blasted by the A. nidulans genome database (Aspergillus Sequencing Project, Broad Institute of MIT and Harvard).
Tagging of AnPRS1 with GFP
To generate an alcA(p)–gfp–Anprs1 fusion construct, a 1133 bp fragment of Anprs1 was amplified from TN02A7 genomic DNA with primer Rec-Anprs1-5′ (NotI site included) and primer Rec-Anprs1-3′ (XbaI site included) (Table S2). The 1133 bp amplified DNA fragment was cloned into the corresponding sites of pLB01, yielding pLB-Anprs1 5′ (Liu et al., 2003). This plasmid was transformed into TN02A7. Homologous recombination of this plasmid into the Anprs1 locus should result in an N-terminal GFP fusion of the entire Anprs1 gene under control of the alcA promoter and a fragment of Anprs1 under its own promoter. The transformant, which formed the normal colonies under the inducing condition but showed slow-growth defects at 30°C under the repressing conditions, was subjected to diagnosis PCR analysis using a forward primer (GFP upstream) designed to recognize the gfp sequence, and a reverse primer (Anprs1 downstream) designed to recognize the Anprs1-3′ sequence.
Transmission electron microscopy
Transmission electron microscopy examination was carried out mainly as described in previous research (Horiuchi et al., 1999; Ichinomiya et al., 2005). In brief, for sample preparation, germlings grown on YAG plates for 10 h were fixed firstly for 5 h in 0.5% glutaraldehyde with 0.1 M phosphate buffer (pH 7.0), and then washed by 0.1 M phosphate buffer (pH 7.0). Next the sample was fixed in a secondary fixation including 2% osmium tetroxide for 2 h in 0.1 M phosphate buffer. After dehydration with a graded ethanol series, specimens were embedded in epoxy resin. The sections, cut on an ultramicrotome with a glass knife, were stained with uranyl acetate and lead citrate and observed by TEM (Hitachi H-7650).
Constructions of gene replacement strains
A strain containing the Anprs1 null mutation was created by double joint PCR (Yu et al., 2004). The Aspergillus fumigatus pyrG gene in plasmid pXDRFP4 was used as a selectable nutritional marker for fungal transformation. The linearized DNA fragment 1 which included a sequence of about 563 bp that corresponded to the regions immediately upstream of the Anprs1 start codon was amplified with the primers 5′Anprs1-For and 5′Anprs1-Rev+Tail (Table S2). Linearized DNA fragment 2 including a sequence of about 801 bp that corresponded to the regions immediately downstream of the Anprs1 stop codon was amplified with primers 3′Anprs1-For+Tail and 3′Anprs1-Rev (Table S2). Lastly, purified linearized DNA fragments 1 and 2 plus the pyrG gene were mixed and used in a fusion PCR with primers Nested-Forward and Nested-Reverse. The final fusion PCR products were purified and used to transform A. nidulans strains TN02A7. A similar strategy was used to construct the truncated strain by using primers 5′For-Anprs1 and 5′Rev-Anprs1 for the 5′ region, 3′For-Anprs1 and 3′Rev-Anprs1 for the 3′ region, and 5′Nest-Anprs1 and 3′Nest-Anprs1 for the fusion product. The final 2190 bp Anprs1 5′-AfpyrG-Anprs1 cassette was purified and used to transform A. nidulans strains TN02A7. The Anprs2 deletion strain and sin110/ΔspgA double mutant strain were constructed by using the similar strategy and transformed to TN02A7 and Sin110-1 strains respectively. Oligonucleotides used in this study are listed in Table S2.
Microscopy and image processing
Several sterile glass coverslips were placed on the bottom of Petri dishes, and gently overlaid with liquid media containing conidia. Strains were grown on the coverslips at related temperature prior to observation under microscope. The GFP–AnPRS1 signal was observed in live cells by placing the coverslips on a glass slide. DNA and chitin were stained using DAPI and CFW (Sigma Aldrich, St Louis), respectively, after the cells had been fixed with 4% paraformaldehyde (Polyscience, Warrington, PA) (Harris et al., 1994). For immunofluorescent detection of actin, the growing hyphae were fixed in PBS (pH 7.4) with 4% paraformaldehyde at room temperature for 30 min. After three washes in PBS, cells were permeabilized with 0.1% Triton X-100 in PBS for 15 min, washed three times with PBS and blocked in blocking buffer (1% bovine serum albumin in PBS) at room temperature for 30 min. Cells were then stained with antiactin mouse monoclonal (MP, 1:200) and Alexa Fluor 546 rabbit antiactin mouse IgG (Invitrogen, 1:400) as primary and secondary antibodies respectively. Differential interference contrast (DIC) images of the cells were collected with a Zeiss Axio Imager A1 microscope (Zeiss, Jena, Germany). These images were then collected and analysed with a Sensicam QE cooled digital camera system (Cooke Corporation, Germany) with MetaMorph/MetaFluor combination software package (Universal Imaging, West Chester, PA) and the results were assembled in Adobe Photoshop 7.0 (Adobe, San Jose, CA).
RNA isolation and quantitative RT-PCR analysis
The mycelia were cultured for 2 days in related media, and were then pulverized to a fine powder in the presence of liquid nitrogen. Total RNA was isolated using Trizol (Invitrogen, 15596-025) following manufactory instruction. One hundred milligrams of mycelia per sample was used as the starting material for the determination of total RNA. Reverse transcription polymerase chain reaction (RT-PCR) was carried out using SuperScript™III First Strand Synthesis System (Invitrogen, 18080-051), and then cDNA was used for real-time analysis. For real-time reverse transcription quantitative PCR (RT-qPCR), independent assays were performed using SYBR Premix Ex TaqTM (TaKaRa, DRR041A) with three biological replicates, and expression levels were normalized to actin mRNA level. The 2ΔCT method was used to determine the change in expression.
Assay of AnPRS activity
Phosphoribosyl pyrophosphate synthetase activity was assayed as following a modified version of Hove-Jensen (2004). Briefly, conidial spores were inoculated in the liquid medium and shaken at 200 r.p.m. at 37°C for 48 h. The mycelia were then resuspended in 1000 μl of extraction buffer (50 mM KH2PO4/K2HPO4 at pH 7.5, 10% glycerol, 0.1% Triton X-100, 5 mM MgCl2, 1 mM EDTA, 1 mM EGTA, 1 mM PMSF and 5 mM DTT) and disrupted by MP Fast-Prep 24, clarified by centrifugation (10000 g for 10 min at 4°C). Measurements were performed in microplates by mixing 10 μl of the supernatant of the enzyme extract with 100–200 μl of measuring buffer (KH2PO4/K2HPO4 at pH 7.5, 5 mM MgCl2, 3.75 mM R5P, 2 mM ATP, 3.75 mM phosphoenolpyruvate, 0.2 mM NADH, 1.5 U myokinase, 3 U pyruvate kinase and 1.5 U lactate dehydrogenase). The oxidation of NADH was measured by monitoring absorption at 340 nm using a Multiskan Spectrum (Thermo Electron Corporation, Waltham, MA, USA) at room temperature. The specific activity of AnPRS is expressed as μmol min−1 (mg of NADH oxidized)−1, using the molar extinction coefficient of 6220 M−1 for NADH. The soluble protein content of the supernatant was determined using a dye binding assay (Bradford, 1976).
Interaction analysis by yeast two-hybrid analysis
Saccharomyces cerevisiae strain AH109 (Clontech, Palo Alto, CA) was used as the host for the two-hybrid interaction experiments. The full length of cDNA of Anprs2 (AN1965.4) and Anprs3 (AN3169.4) was placed in frame with the DNA binding domain of GAL4 by polymerase chain reaction (PCR) amplification from the UniZAP A. nidulans cDNA library, and subcloned into the pGBKT7 vector (Clotech, Palo Alto, CA). Full-length cDNA of Anprs1 and Anprs3 were used for the activation domains in the pGADT7 vector. The histidine and adenine prototrophy and β-galactosidase were performed according to the Clontech Yeast Protocols Handbook (Clotech, Palo Alto, CA).
This work was financially supported by the National Natural Science Foundation of China to L. L. (NSFC31070031, 30770031) and Natural Science Foundation of the Jiangsu Higher Education Institutions of China (Grant No. 11KJA180005) and the Priority Academic Program Development (PAPD) of Jiangsu Higher Education Institutions, and the Research and Innovation Project for College Graduates of Jiangsu Province (CXZZ11_0885). A. nidulans strain TN02A7 was a gift of B.R. Oakley (Ohio State University, Columbus, Ohio); A. nidulans strain AJM68 was a gift of Christopher J. Staiger (Purdue University, West Lafayette); Plasmid pLB01 was a gift from Dr Bo Liu (University of California, Davis); Plasmid pXDRFP4 and pRG3–AMA–NotI genomic DNA library and UniZAP A. nidulans cDNA library were from FGSC (http://www.fgsc.net).