Fungal cells are surrounded by an extracellular cell wall. This complex matrix of proteins and polysaccharides protects against adverse stresses and determines the shape of fungal cells. The polysaccharides of the fungal wall include 1,3-β-glucan and chitin, which are synthesized by membrane-bound synthases at the growing cell tip. A hallmark of filamentous fungi is the class V chitin synthase, which carries a myosin-motor domain. In the corn smut fungus Ustilago maydis, the myosin-chitin synthase Mcs1 moves to the plasma membrane in secretory vesicles, being delivered by kinesin-1 and myosin-5. The myosin domain of Mcs1 enhances polar secretion by tethering vesicles at the site of exocytosis. It remains elusive, however, how other cell-wall-forming enzymes are delivered and how their activity is coordinated post secretion. Here, we show that the U. maydis class VII chitin synthase and 1,3-β-glucan synthase travel in Mcs1-containing vesicles, and that their apical secretion depends on Mcs1. Once in the plasma membrane, anchorage requires enzyme activity, which suggests co-synthesis of chitin and 1,3-β-glucan polysaccharides at sites of exocytosis. Thus, delivery of cell-wall-forming enzymes in Mcs1 vesicles ensures local foci of fungal cell wall formation.
Fungi are one of the oldest and largest clades of microbes1,2. Up to 5 million species are thought to populate all habitats on Earth3. A hallmark of fungi is their cell wall. This extracellular meshwork of polysaccharides and glycoproteins shapes the fungal cell, thereby enabling substrate, soil or host-tissue exploration by polarized growth4. Moreover, the cell wall forms the environmental interface, which withstands abiotic and biotic stresses. As such, it is also crucial for pathogen recognition by mammalian and plant host cells5,6. The fungal cell wall consists mainly of 1,3-β-glucan and chitin, a polymer of N-acetylglucosamine7,8. Both sugars are synthesized at the apical growth region by membrane-bound 1,3-β-glucan synthase (GS) and chitin synthases (CHSs), respectively. Our knowledge about the mechanism of their apical delivery is fragmentary, but it is widely accepted that GS and CHSs travel in secretory vesicles9. Here, the vesicles fuse with the plasma membrane, thereby exposing their cargo, for synthesis of the extracellular cell wall. However, neither the detailed secretion pathways, nor the mechanism by which the activity of GSs and CHSs is coordinated during wall synthesis, is known.
Filamentous fungal genomes usually carry one or two genes encoding GS, whereas CHSs comprise large super-families, with up to 7–10 members10,
Here, we show that U. maydis class V CHS, class VII CHS and GS co-travel in the same secretory vesicle. Co-delivery is followed by co-secretion, which is dependent on Mcs1. Exocytosed enzymes remain closely positioned and stationary within the plasma membrane. This depends on their enzymatic activity and on the cell wall. Thus, co-localization of cell-wall-forming enzymes in Mcs1-containing vesicles ensures the establishment of local foci of coordinated cell wall synthesis.
Class VII CHSs have lost their myosin motor domain
In Aspergillus fumigatus, Aspergillus nidulans, Fusarium oxysporum, Magnaporthe oryzae and Neurospora crassa, class V and class VII carry a putative N-terminal MMD12,15,16,18,
Class VII CHS requires the motor domain of class V CHS for polar localization
Next, we asked why MMD is either absent from or truncated in class VII CHSs. In U. maydis, the MMD of Mcs1 mediates tethering of myosin5- and kinesin1-delivered secretory vesicles to the growth region17. This increases the pausing time of vesicles and thereby supports apical exocytosis17. We hypothesized that Chs6 is delivered in Mcs1-containing and Mcs1-tethered vesicles, which could have made Chs6 MMD redundant. To test this hypothesis, we investigated the localization of a fusion of a green fluorescent protein (GFP)-tag and Chs6 (GFP–Chs6), as well as the fusion protein GFP–MMD–Chs6, where the N-terminal MMD (aa 1–915) was fused to Chs6 (aa 59–1180; Fig. 1b; see Supplementary Table 1 for strain genotypes and Supplementary Table 2 for experimental strain usage). When expressed in the Δchs6 background, GFP–Chs6 localized to the growth region of yeast-like cells of U. maydis (Fig. 1c, left image; Fig. 1d). In the absence of Mcs1 (Δchs6Δmcs1 mutants), GFP–Chs6 lost this apical localization (Fig. 1c, middle image; Fig. 1d), despite its intracellular motility in vesicles (Fig. 1e, kymograph shows motility as diagonal lines). However, fusion of the MMD to Chs6 restored polar exocytosis and accumulation (Fig. 1c, right image; Fig. 1d). These results demonstrate that the MMD of Mcs1 is required for Chs6 positioning. Thus, we speculate that the MMD of class VII CHSs was removed gradually due to functional redundancy.
Class VII and class V CHSs are co-delivered
The role of Mcs1 in Chs6 exocytosis suggested that both could be co-delivered in the same vesicle. Indeed, both fluorescent proteins Mcs1–GFP3 and Chs6–GFP3 covered similar regions of the growing cell tip (Fig. 2a; plasma membrane labelled by mCherry-Sso1, ref. 26). In apical regions of the cell that were photobleached to reduce signal interference, a treatment harmless to the cell17, Chs6–GFP3 moved rapidly in a bidirectional manner, often pausing before turning direction (Supplementary Video 1 and Supplementary Fig. 8a). This pattern was indistinguishable from Mcs1–GFP3 (Supplementary Fig. 8a and Supplementary Video 2). Indeed, the velocity, flux and pausing time of the fluorescent signals for Mcs1–GFP3 and Chs6–GFP3 were not significantly different (Fig. 2b). This is consistent with the hypothesis that Chs6 and Mcs1 travel in the same vesicle. To test this, we co-visualized red-fluorescent mCherry3–Mcs1 and Chs6–GFP3. Indeed, ∼90% of the mCherry3–Mcs1 signals co-travelled with Chs6–GFP3 (Fig. 2c,d; Supplementary Fig. 8b and Supplementary Video 3). This reinforces the idea that Chs6 and Mcs1 are co-delivered to the growth region in the same carrier.
Class VII and class V CHSs are co-secreted
Mcs1-carrying vesicles fuse with the plasma membrane, thereby allowing chitin synthesis into the cell wall. If Mcs1 and Chs6 are co-delivered, both enzymes should be exocytosed simultaneously. Indeed, mCherry3-Mcs1 co-localized with Chs6–GFP3 in the apical growth region ‘cap’, where cell wall formation occurs (Fig. 2e, co-localization results appear in yellow; Fig. 2f, arrowheads, Supplementary Fig. 9). We gathered quantitative data on CHS secretion rates by monitoring the reappearance of the GFP signal following photobleaching. This recovery was considered to represent the delivery and exocytosis of unbleached fluorescent signals17. We expressed Mcs1–GFP3 and Chs6–GFP3, respectively, in cells that carry the plasma membrane marker mCherry–Sso1. At 5 min after photobleaching the growth region, signals reappeared (Fig. 2g, arrowheads; the mCherry-Sso1 image was taken from the ‘Pre’ image). These fluorescent signals overlapped with the mCherry–Sso1 signal (Supplementary Fig. 10), confirming that fluorescent Chs6–GFP3 has been inserted into the plasma membrane. We determined the number of newly secreted GFP signals in the plasma membrane. We did not find significant differences in secretion rates for Mcs1–GFP3 and Chs6–GFP3 at the apical growth region or in the mother cell (Fig. 2h), where reduced rates of secretion have been reported17. Moreover, in cells co-expressing mCherry3–Mcs1 and Chs6–GFP3, both signals appear together after photobleaching (Fig. 2i). Taken together, these data are consistent with the notion that Mcs1 and Chs6 are exocytosed together.
Next, we asked for further evidence that Mcs1 is required for secretion of Chs6. We generated a conditional mutant, replacing the mcs1 native promoter with the repressible crg promoter27. In the presence of arabinose, mCherry3–Mcs1 was expressed (Supplementary Fig. 11, Mcs1↑, co-localization with Chs6–GFP3 results in yellow signals), whereas mCherry3–Mcs1 was depleted in glucose-containing medium. Consistent with a role of Mcs1 in Chs6 secretion, low Mcs1 levels resulted in reduced Chs6–GFP3 signals at the growth region (Fig. 2j). Instead, Chs6–GFP3 signals accumulated in the cytoplasm (Supplementary Fig. 11, Mcs1↓; arrowheads). The secretion rate of Chs6–GFP3 at buds was consistently and significantly reduced (Fig. 2k). Taken together, our results support a model in which Mcs1 and Chs6 co-travel in the same secretory vesicle, and this results in co-secretion at sites of cell wall formation. Tethering of these vesicles is performed by the MMD of Mcs1, rendering redundant the MMD of Chs6. This could explain the gradual loss of the MMD in class VII CHSs.
GS delivery and secretion requires Mcs1
We further characterized our CHS mutants by ultrastructural analyses. This revealed that Δchs6 mutant cells form significantly thicker cell walls than wild-type cells (Fig. 3a,b). This suggests that Δchs6 cells reinforce their cell wall. Consistently, Δchs6 cells were more resistant to treatment with cell-wall-digesting enzymes than wild-type cells (Fig. 3c,d). Defects in the fungal cell wall are known to activate the cell-wall integrity pathway, which results in reinforcement of the cell wall due to increased activity of CHSs or GS (refs. 28,
Secretory vesicles contain various numbers of CHSs and GS
Our live-cell imaging experiments suggested that Mcs1, Chs6 and Gsc1 are delivered in the same vesicle. We gathered further evidence for this by ultrastructural investigation of secretory vesicles, purified from strains AB33Ch3Mcs1_Chs6G3 and AB33Ch3Mcs1_G3Gsc1, following protocols published for S. cerevisiae31. In epifluorescent microscopy, these vesicles combined red and green fluorescence, confirming that they contain all three enzymes (Fig. 4a). We then performed immuno-gold labelling, using commercial antibodies raised against mCherry and GFP. The chemically fixed vesicle preparation was homogeneous, and antibodies exclusively labelled the membranes (Supplementary Fig. 13). These experiments confirmed that Mcs1, Chs6 and Gsc1 co-localized in the same vesicles (Fig. 4b). These vesicles had a diameter of ∼35 nm (Fig. 4c; mean ± standard deviation: 36.81 ± 4.27 nm, n = 16, for vesicles containing Mcs1 and Chs6; 34.04 ± 4.14 nm, n = 21, for vesicles containing Mcs1 and Gsc1). This size is concordant with the dimensions of neuronal secretory vesicles (30–60 nm; refs 32,33). We found variation in the number of Mcs1, Chs6 and Gsc1 per vesicle (1–4 signals; Fig. 4b). To address this quantitatively, we measured the fluorescence intensity of moving Mcs1–GFP3, Chs6–GFP3 and GFP3–Gsc1 in living cells and compared these intensities with fluorescent nuclear pores. Each nuclear pore contains sixteen Nup107–GFP nucleoporin fusion proteins and thus has a uniform fluorescent signal intensity (Fig. 4d; arrowheads). This allows them to be used as an internal calibration standard for measuring protein numbers in living cells of U. maydis34,35. This approach revealed that secretory vesicles usually carry one or two Mcs1 and Chs6 molecules, but more than two Gsc1 molecules co-travel in the vesicles (Fig. 4e; number of Gsc1 significantly different from Mcs1 and Chs6 at P < 0.0001, Mann–Whitney test).
Mcs1, Gsc1 and Chs6 concentrate in stationary focal points of cell wall formation
The co-delivery of Gsc1, Chs6 and Mcs1 in the same vesicle raises the possibility that these enzymes, once exocytosed into plasma membrane, cooperate in wall formation. To investigate this, we observed the ‘behaviour’ of mCherry3–Mcs1, Chs6–GFP3 and GFP3–Gsc1 after insertion into the apical plasma membrane of U. maydis cells. We found that exocytosed Mcs1 and Chs6 or Mcs1 and Gsc1 continued to co-localize and remain stationary in the plasma membrane (Fig. 5a). Secreted mCherry3–Mcs1, Chs6–GFP3 or GFP3–Gsc1 usually co-localized for several minutes (Fig. 5b), suggesting that diffusion in the plasma membrane is restricted. This finding is surprising, as the fluorescent syntaxin GFP–Sso1 diffuses into photobleached regions (Supplementary Fig. 14). Thus, cell-wall-forming enzymes are actively immobilized after exocytosis. We tested if the cytoskeleton restricts diffusion of secreted Mcs1–GFP3 and GFP3–Gsc1 by counting newly secreted fluorescent signals in photobleached growth regions in control cells and mutants that were truncated in the MMD of Mcs1 (ref. 26). In addition, we depolymerized the cytoskeleton with latrunculin A and benomyl, which disassemble F-actin and microtubules in U. maydis, respectively36. In control cells, ∼80% of all Mcs1 signals are stationary and non-diffusive (Fig. 5c, Control; Supplementary Video 5). This number did not change significantly when the MMD was truncated or the cytoskeleton was disrupted (Fig. 5c, LatA + Ben; no significant difference, P = 0.422, Kruskal–Wallis test). Thus, we conclude that the cytoskeleton has no obvious role in anchoring secreted cell-wall-forming enzymes. Next, we asked whether the enzymes are physically linked to the wall and therefore restricted in diffusion within the plasma membrane. We removed the wall with cell-wall-degrading enzymes and investigated Mcs1–GFP3 in protoplasts (Fig. 5d). In the absence of the wall, most Mcs1–GFP3 showed random motility, and the number of non-diffusive Mcs1–GFP3 signals was reduced significantly (Fig. 5c,e, ‘No cell wall’; P < 0.0001, Student's t-test; Supplementary Video 5). This raised the possibility that synthetic enzymes are anchored via their newly formed polysaccharide chains. We tested this by observing newly secreted Mcs1–GFP3 and GFP3–Gsc1 in the presence of the CHS inhibitor nikkomycin Z (ref. 37) and the GS inhibitor caspofungin38. Indeed, simultaneous treatment with both inhibitors increased the diffusive motion of secreted Mcs1–GFP3 (Fig. 5c,f, ‘NikkoZ+CSG’; Supplementary Video 5) and GFP3–Gsc1 (Fig. 5g, ‘NikkoZ+CSG’). Interestingly, treatment with either nikkomycin Z or caspofungin alone had an intermediate effect on GFP3–Mcs1 mobility (Fig. 5c, ‘NikkoZ’ and ‘CSG’). On the other hand, blocking CHS activity with nikkomycin Z increased the diffusion of Gsc1 (Fig. 5g, ‘NikkoZ’). These results suggest that immobilization of one enzyme depends, in part, on the activity of the other enzymes, suggesting their cooperation during cell wall synthesis. Taken together, these results support a model in which CHSs and GS are co-delivered in the same secretory vesicle, to ensure co-exocytosis and the formation of local foci of cell wall synthesis activity.
Cell wall synthases co-travel in small secretory vesicles in hyphae
U. maydis is a dimorphic fungus: it grows as yeast-like cells and also as tip-growing elongate hyphae. The results described so far were derived from an investigation of the yeast-like growth form, and we set out to extend our study to hyphal cells. Consistent with the extension of the hyphal tip, fluorescent Mcs1, Chs6 and Gsc1 concentrated at the growing hyphal apex (Fig. 6a). After photobleaching, rapidly moving vesicles carrying mCherry3–Mcs1/Chs6–GFP3 and mCherry3–Mcs1/GFP3–Gsc1 were observed (Fig. 6b; Supplementary Fig. 15 and Supplementary Video 6). A small portion of Chs6 and Gsc1 travelled independently of Mcs1 (Fig. 6c), which confirms the results described for yeast-like cells and suggests additional Mcs1-independent secretion pathways. Next, we purified vesicles from hyphal cells of mCherry3–Mcs1/Chs6–GFP3 and mCherry3–Mcs1/GFP3–Gsc1 co-expressing strains. In contrast to the preparation from yeast-like cells, we did not chemically ‘fix’ these vesicles. This improved the quality of the electron micrographs and revealed a largely homogeneous population of small vesicles (diameter 23.40 ± 3.15 nm, n = 30). Immuno-gold labelling indicated that only a small number of these vesicles contain Mcs1. A portion of these Mcs1-positive vesicles was either paired with Chs6 or with Gsc1 (Fig. 6d). Thus, in U. maydis, co-delivery of cell wall synthases in the same vesicle is characteristic of hyphae as well as yeast-like cells.
This study provides evidence for a central role of fungal myosin–CHSs in focal point secretion of cell-wall-forming enzymes. We integrated fluorescent tags into the native loci of two CHSs and a GS in U. maydis, allowing us to visualize native levels of these cell-wall-forming enzymes. We used these strains to show, both by live-cell imaging and ultrastructural studies, that the Mcs1, Chs6 and Gsc1 co-travel in secretory vesicles. Consistent with a central role of the MMD of Mcs1, we demonstrate that secretion of Chs6 depends largely on Mcs1 activity. On the other hand, exocytosis of Gsc1 is less dependent on Mcs1. A significant proportion of Gsc1 neither co-localizes with Mcs1 nor requires Mcs1 for exocytosis. This suggests the existence of an Mcs1-independent secretion pathway for GS. However, this putative pathway is not sufficient to reinforce the cell wall when chs6 and mcs1 are deleted. This suggests that the Mcs1-dependent delivery of cell-wall-forming enzymes is the dominant pathway. U. maydis contains four polar-localized CHSs (ref. 11), and we speculate that these enzymes are delivered in Mcs1 vesicles, but experimental evidence to support this is missing. However, it is important to note that our ultrastructural results, as well as low rates of anterograde delivery of Mcs1 vesicles17 suggest that cell wall synthase-containing carriers represent a minor population of all secretory vesicles.
While co-delivery of CHSs and GS in the same vesicle is an efficient way of ensuring focused co-exocytosis and coordinated activity of cell-wall-forming enzymes, it may not be the only way of constructing a fungal cell wall. Studies of the localization of cell wall synthases in the Spitzenkörper of N. crassa suggest that several CHSs and GS travel in distinct subpopulations of secretory vesicles20,39,
How common is this myosin–CHS dependent secretion pathway? Studies in U. maydis suggest that the class V myosin–CHS supports polar exocytosis by tethering vesicles to apical F-actin17. We demonstrate here that this pathway also delivers other cell-wall-forming enzymes, including class VII CHS and GS. We propose that this co-delivery renders the MMD of class CHS VII redundant in U. maydis. If so, the loss of this MMD in other fungi suggests co-delivery of cell-wall-forming enzymes is of general importance in filamentous fungi. In support of this, we found a single myosin–CHS with a highly conserved MMD in ascomycete and basidiomycete genomes. Even the ancient cryptomycete fungus R. allomyces carries a highly conserved myosin–CHS (ref. 43,44). Thus, we can speculate that this is, indeed, an ancestral pathway. However, variations on this theme are likely, as the genomes of the zygomycete Mucor circinelloides and chytridiomycete Spizellomyces punctatus carry numerous myosin–CHSs, suggesting multiple secretion pathways. More studies are needed to investigate this possibility. On the other hand, ascomycete fungi appear to use the largely reduced MMD of class VII CHSs to bind F-actin16,45, possibly involving a core set of conserved amino acids in their truncated MMD (Supplementary Fig. 16; ∼53% identical; PFAM). Although this reduced MMD cannot replace its counterpart in class V CHSs (ref. 46), it seems that both CHSs cooperate in the secretion of wall-forming enzymes in A. nidulans47. In essence, fungal cell wall formation involves an ancestral myosin–CHS-dependent secretion pathway. Cell wall synthases are co-delivered in myosin–CHS-containing transport vesicles (Fig. 6e) and, after co-secretion, cooperate in cell wall formation. We consider it likely that the delivery of such ‘cell wall factories’, consisting of multiple cell wall synthases, is a widespread mechanism found in all classes of fungi, including filamentous ascomycetes. As such, this pathway is pivotal to polarized fungal growth, and essential for colonizing the environment and invading animal and plant hosts.
Strains, plasmids and expression constructs
The U. maydis strains AB33, SG200, FB1ΔChs6, FB1ΔChs6ΔMcs1, SG200ΔChs6, AB33_Mcs1G3, AB33mCh3Mcs1, SG200G3Mcs1ΔMM, SG200G3Mcs1, FB1Chs6G3, AB33GSso1 and FB2N107G have been described previously11,17,21,26,48,
The pnChs6G3 plasmid contains the U. maydis class VII CHS gene, fused to three copies of the egfp gene. This plasmid was constructed by replacing the phleomycin resistance cassette from plasmid pChs6G3 (ref. 17) with the hygromycin phospho-transferase gene resistance cassette (hygR), by standard procedures.
The pnG3Gsc1 plasmid contains a triple egfp tag fused to the gene for the catalytic subunit of GS. Fragments of 1,011 bp from the gsc1 5′ upstream region, 1,032 bp of the gsc1 promoter, and 1,006 bp open reading frame (ORF) of gsc1 were amplified by PCR from genomic DNA of U. maydis strain 521. A 717 bp fragment from the egfp ORF and 2,798 bp of the hygR cassette were amplified from the plasmid pHrpl25G (ref. 53). This was done using primers YH166, YH269, YH270, YH273, YH421, YH422, YH423, YH424, YH425 and YH426 (Supplementary Table 3). Ligation of all fragments was carried out by in vivo recombination in S. cerevisiae, following published protocols52. Two additional copies of egfp were introduced into a BsrGI restriction site of the resulting plasmid pnGGsc1 using standard ligation techniques.
The pnGChs6 plasmid contains the U. maydis class VII CHS gene, fused to a single copy of the egfp gene. The fusion construct is expressed under the chs6 native promoter. It was constructed from a DNA fragment of 2,220 bp, covering the chs6 promoter, 717 bp from the egfp ORF, 3,543 bp from the chs6 ORF and 1,201 bp from the chs6 terminator amplified by PCR either with genomic DNA of U. maydis strain 521 or plasmid pn3GMcs1 (ref. 17) as templates, and using primers MU213, MU214, MU215, SK179, MU216, MU217, MU211 and MU218 (Supplementary Table 3). The products were recombined into the BamHI and HindIII digested plasmid pNEBcbx-yeast34 in S. cerevisiae, resulting in pnGChs6.
The pnGM-Chs6 plasmid contains the MMD of the U. maydis class V CHS gene, fused to the class VII CHS gene and to a single copy of the egfp gene. The fusion construct is expressed under the chs6 native promoter. It was constructed from a DNA fragment of 2,220 bp, covering the chs6 promoter, 717 bp from the egfp ORF, 2,745 bp from the mcs1 N-terminal ORF, 3,369 bp from the chs6 C-terminal ORF and 1,201 bp of the chs6 terminator. The fragments were amplified by PCR either with genomic DNA of U. maydis strain 521 or plasmid pn3GMcs1 (ref. 17) as templates, and using primers SK50, SK434, SK179, MU209, MU210 and MU211 (Supplementary Table 3). The products were recombined into the BamHI and HindIII digested plasmid pNEBcbx-yeast34 in S. cerevisiae, resulting in pnGM-Chs6.
The pPcmCh3Mcs1 plasmid allows expression of the fusion gene mcherry3-mcs1 under the conditional crg-promoter27. It was generated by replacing the 958 bp fragment of the mcs1 promoter in plasmid pmCh3Mcs1 (ref. 17) with a 3,525 bp fragment containing the crg1 promoter, which was obtained from pCcrgKin1rigor (ref. 17) using primers YH461 and YH462. The plasmid was generated using in vivo recombination in S. cerevisiae, following published protocols52.
To generate strains AB33Chs6G3 and AB33mCh3Mcs1_Chs6G3, plasmid pChs6G3 was digested with BglI and integrated into the chs6 locus of strains AB33 (ref. 48) and AB33mCh3Mcs1 (ref. 17), respectively. Strain AB33Mcs1G3 was generated by homologous integration of plasmid pMcs1G3 (ref. 17) into strain AB33. Strains AB33G3Gsc1 and AB33Ch3Mcs1_G3Gsc1 were generated by linearization and integration of plasmid pG3Gsc1 into the gsc1 locus of strains AB33 or AB33mCh3Mcs1 (ref. 17). AB33Mcs1G3_ChSso1 and AB33Chs6G3_ChSso1 were generated by ectopic integration of plasmid pomChSso1 (ref. 26), linearized with PciI, into the genome of strains AB33Mcs1G3 and AB33Chs6G3. Strain FB1ΔChs6GChs6 was generated by ectopic integration of plasmid pnGChs6, linearized with HpaI, into the genome of FB1ΔChs6 (ref. 21). FB1Chs6G3_rCh3Mcs1 and AB33G3Gsc1_rCh3Mcs1contain plasmid pPcmCh3Mcs1, digested with SapI and integrated into the mcs1 locus of strains FB1Chs6G3 (ref. 17) and AB33G3Gsc1. Strains FB1ΔChs6ΔMcs1GChs6 and FB1ΔChs6ΔMcs1GChs6MMD were generated by ectopic integration of plasmids pnGChs6 or pnGM-Chs6 linearized with HpaI into the genome of FB1ΔChs6ΔMcs1 (ref. 11).
U. maydis liquid cultures were grown for 8–12 h in complete medium (CM, ref. 54) containing 1% (wt/vol) glucose (CMglucose) with shaking at 200 revolutions per minute (r.p.m.) at 28 °C. To induce hyphal growth, cells were washed and transferred to nitrate minimal medium (NM, ref. 48) supplemented with 1% (wt/vol) glucose (NMglucose). Cells were grown under these conditions for 5–10 h at 200 r.p.m. at 28 °C. Strains FB1Chs6G3_rCh3Mcs1 and AB33G3Gsc1_rCh3Mcs1 were grown in complete medium containing 1% (wt/vol) arabinose, which induces gene expression from the crg1 promoter27. To repress expression of mcs1, cells were transferred into CMglucose for 12 h at 28 °C, with shaking at 200 r.p.m.
Laser-based epifluorescence microscopy
Cells were placed onto a 2% (wt/vol) agar cushion and observed using an IX81 motorized inverted microscope (Olympus), equipped with a PlanApo ×100/1.45 oil TIRF objective (Olympus) and a VS-LMS4 Laser-Merge-System with solid-state lasers (488 nm/70 mW and 561 nm/70 mW, Visitron System). For photobleaching experiments, a 405 nm/60 mW diode laser was used, which was attenuated by an ND 0.6 filter, resulting in 15 mW output power, coupled into the light path by an OSI-IX 71 adaptor (Visitron System) and controlled by a UGA-40 controller (Rapp OptoElectronic) and VisiFRAP 2D FRAP control software for Meta Series 7.5.x (Visitron System). Simultaneous observation of red and green fluorescent protein fluorescence was achieved with a Dual-View Microimager (Photometrics) equipped with a dual line beamsplitter (z491/561, Chroma), an emission beamsplitter (565 DCXR, Chroma), an ET-Bandpass 525/50 (Chroma) and a BrightLine HC 617/73 (Samrock). Images were acquired using a Photometrics CoolSNAP HQ2 camera (Roper Scientific). All parts of the system were under the control of the software package MetaMorph (MDS Analytical Technologies). Samples were observed for no longer than 10 min, to prevent oxygen depletion. All image processing was performed in MetaMorph.
All statistical analyses were performed using the software Prism5 (GraphPad). Before comparison, data sets were tested for normal distribution using a Shapiro–Wilk normality test. When normally distributed (P < 0.05), they were further analysed using unpaired two-tailed Student's t-testing, with Welch's correction, to account for potential differences in the variances of the data sets. The means ± standard error of such normally distributed data are shown as bar charts in the figures.
In cases where at least one data set was non-normally distributed (P > 0.05 in Shapiro–Wilk tests), we used non-parametric Mann–Whitney testing. Multiple data sets, where at least one data set was non-normal, were compared using non-parametric Kruskal–Wallis testing. Analysis of non-normal data sets is represented by whisker plots (= box plots), which show 25th/75th percentiles as blue lines, the median as a red line, and the minimum and maximum values by the ends of the whiskers. All values that are incorporated into the main body of the manuscript are given as means ± standard deviation unless stated otherwise.
Quantitative analysis of fluorescent intensities and motility
All measurements were carried out in 14-bit images using the software MetaMorph. Motility measurements of Mcs1G3 and Chs6G3 were determined in strains AB33Mcs1G3 and AB33Chs6G3. For velocity, frequency and pausing time measurements of Mcs1G3 and Chs6G3, an image series of 150 frames at 200 ms of the 488 nm observation laser were taken after photobleaching of the whole bud by a 75 ms light pulse using a 405 nm laser (60 mW) at 100% laser power (beam diameter 30 pixels). Velocities, frequencies and pausing times of Mcs1G3 and Chs6G3 signals were measured in kymographs using MetaMorph. Intensity measurements of Chs6G3 and G3Gsc1 were determined in strains FB1Chs6G3_rCh3Mcs1 and AB33G3Gsc1_rCh3Mcs1 under mCh3Mcs1 induced or repressed conditions. Pictures were taken at an exposure time of 200 ms of the 488 and 561 nm observation lasers. Analysis of the signal intensities of Chs6G3 and G3Gsc1 at the growth region was performed by defining an area of interest in the apical growth regions, followed by measuring the average intensities at the plasma membrane using MetaMorph. All measured values were corrected for background and the mean average values were calculated.
Quantitative analysis of co-motility
Co-motility of Ch3Mcs1 with Chs6G3 or G3Gsc1 was analysed in the yeast-like and hyphal growth forms of strains AB33Ch3Mcs1_Chs6G3 and AB33Ch3Mcs1_G3Gsc1. Cells were placed onto 2% (wt/vol) agar cushions and the entire bud or 20 µm of the hyphal tip was photobleached with a 75 ms light pulse from a 405 nm laser (60 mW) at 100% laser power (beam diameter 30 pixels). After a 5 s pause, image series of 75 frames at 150 ms and binning 2 were taken using both the 488 nm and the 562 nm observation lasers at 50% output power. Co-motility was analysed in kymographs using MetaMorph.
FRAP-based secretion assays
Mcs1G3, Chs6G3 and G3Gsc1 secretion rates were determined essentially as described in ref. 17. In brief, cells of strains AB33Mcs1G3_ChSso1, AB33Chs6G3_ChSso1, FB1Chs6G3_rCh3Mcs1 and AB33G3Gsc1_rCh3Mcs1 were observed by live-cell microscopy. Reference images were taken at 70% output power of the 488 nm and 100% output power of the 561 nm observation lasers at an exposure time of 250 ms followed by a 100 ms light pulse using a 405 nm laser (60 mW) at 100% laser power (beam diameter 30 pixels) to bleach the growth region or one flank of the mother cell. An image was acquired directly after laser treatment to confirm successful photobleaching. Subsequently, an image series of 75 frames at 5 min after photobleaching was acquired, using 70% of the 488 nm and 100% of the 561 nm observation lasers at an exposure time of 250 ms. Stable insertions of fluorescent signals at the cell periphery, confirmed in kymographs, were considered exocytosed enzymes. The number of inserted signals per 1 µm plasma membrane and per 5 min after photobleaching was determined. To determine the position of secreted Chs6G3 relative to the mChSso1-labelled plasma membrane, intensity profiles of inserted signals were taken and overlaid with mChSso1 images, taken prebleaching. Intensity values of 14 secreted signals were measured and averaged.
Co-localization before and after secretion
mCh3Mcs1 and Chs6G3 or G3Gsc1 co-localizations were determined by photobleaching the growth region in cells of strains AB33Ch3Mcs1_Chs6G3 and AB33Ch3Mcs1_G3Gsc1, placed on a 2% (wt/vol) agar cushion, using a 100 ms 405 nm laser light pulse (60 mW; beam diameter 30 pixels). For co-localization during delivery to the plasma membrane, an image series of 75 frames was taken 2 s after bleaching using 70% of the 488 nm and 80% of the 561 nm observation lasers at an exposure time of 150 ms. For co-localization, after secretion to the plasma membrane, an image series of 75 frames was taken 5, 7 or 10 min after bleaching using 70% of the 488 nm and 80% of the 561 nm observation lasers at an exposure time of 200 ms. Stable insertion of individual signals was confirmed in kymographs using MetaMorph. The percentage of inserted Chs6G3 or G3Gsc1 signals that co-localized with mCh3Mcs1 after 5, 7 or 10 min was determined.
Protoplast formation was carried out following previously described procedures55. Strains SG200, SG200ΔChs6 and FB1ΔMcs1ΔChs6 were grown overnight in YEPSlight liquid media (0.4% (wt/vol) yeast extract, 0.4% (wt/vol) peptone and 2% (wt/vol) sucrose), reaching an optical density at 600 nm of 0.6–0.8. Cells were centrifuged in a Heraeus Biofuge Stratos benchtop centrifuge (Kendro Laboratory Products) at 3,000 r.p.m. for 10 min. The cell pellet was washed by resuspending in sodium citrate sorbitol buffer (SCS, 20 mM sodium citrate, pH 5.81, 1 M sorbitol) and subsequent centrifugation. The resulting sediment was resuspended in SCS containing 12.5 mg ml−1 novozyme lytic enzymes (Novo Nordisk) and kept at room temperature for 10–15 min. Subsequently, the degree of protoplast formation was assessed by light microscopy. Cells were considered ‘protoplasts’ when the cells lost their ‘cigar-shaped’ appearance and became partially or fully rounded.
Secretory vesicle purification
Secretory vesicles were purified from the yeast and hyphal growth forms of strains AB33mCh3Mcs1_Chs6G3 and AB33mCh3Mcs1_G3Gsc1, following published procedures31. For purification from yeasts, cells were grown overnight in YEPSlight liquid medium. For hyphal cells the strains were grown over night in CM (ref. 12) containing 1% (wt/vol) glucose (CMglucose), with shaking at 200 revolutions per minute (r.p.m.) at 28 °C. To induce hyphal growth, cells were washed and transferred to nitrate minimal medium (NM, ref. 48) supplemented with 1% (wt/vol) glucose (NMglucose). Cells were grown under these conditions for 6 h at 200 r.p.m. at 28 °C. Protoplasts were prepared as previously described55. Protoplasts were sedimented by centrifugation and resuspended in 100 mM PIPES buffer (pH 7.2), containing 0.8 M sorbitol, 1 mM EDTA adjusted to pH 7.2 and protease inhibitor cocktail (Sigma-Aldrich). The protoplasts were disrupted in a Dounce homogenizer on ice and the cell extract was centrifuged at 10,000g for 10 min, using a Heraeus Biofuge Stratos benchtop centrifuge. The supernatant was collected and ultracentrifuged at 100,000g for 1 h, using a TLA 120.1 rotor and Optima MAX Ultracentrifuge (Beckman Coulter). After removal of the supernatant, the clear microsomal pellet was resuspended in 40 µl PIPES buffer.
Determining protein numbers in secretory vesicles
Numbers of G3Gsc1, Chs6G3 and Mcs1G3 molecules in moving fluorescent dots were estimated in cells of strains AB33Mcs1G3, AB33Chs6G3 and AB33G3Gsc1. To determine the number of proteins in a moving signal, short image series were captured. Moving signals were identified and their fluorescent signal intensity, corrected for the adjacent background, was determined in the first frame of the image sequence. This fluorescent intensity was compared to the fluorescent intensity of individual fluorescent nuclear pores in strain FB2N107G (ref. 56), following published protocols34,35.
Liquid cultures of SG200 and SG200ΔChs6 were grown overnight, cells were collected by centrifugation at 4,000g for 5 min in a 15 ml Falcon tube, and the pellet was fixed in 2% (vol/vol) glutaraldehyde and 2% (vol/vol) formaldehyde in 0.1 M PIPES buffer pH 7.2, overnight. Samples were post-fixed in 2% (wt/vol) potassium permanganate in dH2O, dehydrated through increasing concentrations of ethanol (50–100%) and embedded in Durcupan resin (Sigma Aldrich). Ultrathin sections (80 nm) were collected on pioloform-coated EM copper grids (Agar Scientific), contrasted with lead citrate and examined using a JEOL JEM 1400 transmission electron microscope operated at 120 kV. Images were taken with a digital camera (ES 100 W charge coupled device, Gatan). To quantify the cell wall thickness, sections were sampled using systematic uniform random procedures and images were analysed using MetaMorph. Measurements were made from profiles showing a visible plasma membrane, to ensure that a medium section was chosen.
For immuno-gold labelling, purified secretory vesicles from strains AB33Ch3Mcs1_Chs6G3 and AB33Ch3Mcs1_G3Gsc1 were fixed in 1% formaldehyde (in 0.1 M PIPES, pH 7.2) and adhered to pioloform-coated 100-mesh copper EM grids (Agar Scientific). Alternatively, vesicles from hyphal cells were adhered to EM grids without prior chemical fixation. After washing with PBS (pH 7.2), grids were incubated with 0.5% (wt/vol) fish skin gelatine (Sigma Aldrich) in PBS for 10 min, followed by incubation with a rabbit anti-mCherry antibody (MBL, PM005) for 30 min. After washing with PBS (3 × 5 min), the grids were incubated with 5 nm protein A gold (BBI Solutions) for 20 min. This was followed by fixation in 1% (vol/vol) glutaraldehyde/PBS for 5 min and additional incubation in 0.05% (wt/vol) glycine/PBS for 5 min. After washing with PBS, the primary rabbit anti-GFP antibody (Abcam, ab6556) was applied for 30 min followed by subsequent washes in PBS (3 × 5 min) before incubating with 10 nm protein A gold (BBI Solutions). All antibodies and protein A gold were diluted with 0.5% (wt/vol) fish skin gelatine in PBS (pH 7.2). Finally, the samples were washed in PBS (6 × 5 min) and distilled water (10 × 1 min) before contrasting in a 1:9 mixture of 2% (wt/vol) uranyl acetate: 2% (wt/vol) methylcellulose for 10 min. The grids were then air-dried and analysed using a JEOL JEM 1400 electron microscope. The specificity of the gold signal was verified by applying the protein A gold alone. The average diameter of labelled vesicles was estimated from digital images.
Wheat germ agglutinin labelling of chitin
Fluorescein isothiocyanate (FITC)-conjugated wheat germ agglutinin stain was performed as described previously11,57. Stained cells of strains SG200 and SG200ΔChs6 were placed onto 2% (wt/vol) agar cushions, and images of cells were taken using 40% of the 488 nm observation laser at an exposure time of 50 ms. Measurement of wheat germ agglutinin stain intensity at the growth region was performed by determining an area of interest at the apical growth regions, using digital images. This was followed by measuring the average intensities at the cell wall using MetaMorph. All measured values were corrected for background and the mean average values were calculated.
For all inhibitor experiments in liquid culture, logarithmically growing cells of strains SG200G3Mcs1 and SG200G3Mcs1ΔMM were incubated for 30 min with benomyl at 30 µM (stock: 30 mM in DMSO; Fluka), latrunculin A at 10 µM (stock: 20 mM in DMSO; provided by K. Tenney, University of California) or nikkomycin Z at 5 µM (stock: 5 mg in mQH2O; Sigma-Aldrich) and caspofungin 10 µgml−1 (stock: 10 mgml−1 in mQH2O; provided by N. J. Talbot, University of Exeter). Control cells were treated with respective amounts of the solvent DMSO used in the inhibitor assays. Cells were placed onto a 2% (wt/vol) agar cushion containing the same concentration of their respective inhibitors and observed immediately. For measurements of plasma membrane insertion of individual signals of G3Mcs1 and G3Mcs1ΔMM, image series of 100 frames were taken 5 min after photobleaching, using 70% of the 488 nm observation laser at an exposure time of 200 ms. Stable insertion of individual signals was confirmed in kymographs using MetaMorph. The number of stationary and non-stationary signals per 1 µm plasma membrane and per 5 min was determined.
Orthologues of class V and class VII CHSs were identified using a BLAST search at NCBI (http://blast.ncbi.nlm.nih.gov/Blast.cgi), using the predicted amino-acid sequence of U. maydis Mcs1 (NCBI accession no. XP_011389642.1) and Chs6 (NCBI accession no. XP_011389509.1) as bait. Predicted CHS sequences for S. punctatus were obtained from the Broad Institute (https://www.broadinstitute.org/annotation/genome/FGI_Blast/Blast.html). All accession numbers for the predicted protein sequences are provided in Fig. 1a, Supplementary Fig. 1a and Supplementary Fig. 16. The degree of sequence identity and similarity between proteins was determined by using EMBOSS needle (http://www.ebi.ac.uk/Tools/psa/emboss_needle/). The protein domain predictions were done in PFAM (http://pfam.sanger.ac.uk/search) and SMART (http://smart.embl-heidelberg.de/). Sequence alignments were done using ClustalOmega (http://www.ebi.ac.uk/Tools/msa/clustalo/). Maximum likelihood phylogenetic trees were calculated in MEGA 5.2 (http://www.megasoftware.net/mega_beta.php; ref. 58), using Bootstrap testing and 1,000 replications.
The authors thank U. Fuchs, S. Milne and P. Splatt for technical support. M.M.-U. thanks N.J. Talbot for financial support. G.S. acknowledges J. Stajich for fruitful discussions. This work was supported by the Biotechnology & Biological Sciences Research Council (grants BB/H019774/1 and BB/I020667/1 to G.S.).
Motility of Chs6-GFP3 in U. Maydis.
Motility of Mcs1-GFP3 and Chs6-GFP3 in a yeast-like cell of U. maydis.
Co-motility of mCherry3-Mcs1 and Chs6-GFP3 in a yeast-like cell of U. maydis.
Co-motility of mCherry3-Mcs1 and GFP3-Gsc1 in a yeast-like cell of U. maydis.
Diffusive motion of Mcs1-GFP3 in control cells, cell wall-less protoplasts and in the presence of the CHS inhibitor nikkomycin Z and the GS inhibitor caspofungin.
Co-motility of mCherry3-Mcs1 and GFP3-Gsc1 in a hyphal cell of U. maydis.
About this article
Nature Communications (2017)