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Department of MIP, Louisiana State University Health Sciences Center School of Dentistry, 1100 Florida Avenue, Box F8-130, New Orleans, LA 70119, USA
Correspondence
Glen E. Palmer
gpalme{at}lsuhsc.edu
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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mutant was unaffected in either yeast–hypha or chlamydospore differentiation. Furthermore, the atg9 mutant survived within and killed a mouse macrophage-like cell line as efficiently as control strains. The data suggest that autophagy plays little or no role in C. albicans differentiation or during interaction with host cells.
-mannosidase; API, aminopeptidase I; CPY, carboxypeptidase Y; DIC, differential interference contrast; SAP, secreted aspartyl protease |
INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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). In the model yeast Saccharomyces cerevisiae, autophagy is critical for survival during nitrogen starvation, and to complete the differentiation process of sporulation (Takeshige et al., 1992; Tsukada & Ohsumi, 1993). The molecular mechanism behind autophagy has been intensively studied in yeast and mammalian cells, and many of the components that are required have been identified (Abeliovich & Klionsky, 2001; Klionsky, 2005). A portion of cytoplasmic material is bound in a double-membrane structure known as an autophagosome, which fuses with the vacuolar membrane to release a single-membrane-bound compartment within the vacuole lumen. Degradation of this material by the hydrolytic enzymes within the vacuole recycles cellular building blocks such as amino acids, which can be used in the synthesis of new macromolecules. Thus autophagy is typically considered a non-specific means to degrade ‘bulk’ cytoplasmic material. However, recent studies have revealed that under some conditions certain cargo is preferentially targeted for sequestration within the autophagosome (Nair & Klionsky, 2005). For example, in S. cerevisiae under nutrient-replete conditions (i.e. when autophagy is repressed), the vacuolar hydrolases aminopeptidase I (API) and -mannosidase (AMS) are delivered from cytoplasm to vacuole via the cvt (cytoplasm to vacuole trafficking) pathway. The cvt pathway closely resembles autophagy, and involves sequestration of the cargo proteins within a double-membrane vesicle, which subsequently fuses to the vacuole to release its contents within the lumen. The vesicles formed are smaller than autophagosomes and appear to specifically deliver the API and AMS hydrolases. Interestingly, the cvt and autophagy pathways rely on much of the same cellular machinery, and many mutants defective in autophagy are also blocked in the cvt pathway (Abeliovich & Klionsky, 2001; Thumm et al., 1994; Tsukada & Ohsumi, 1993). Furthermore, under starvation conditions API and AMS are delivered to the vacuole in the autophagosome, underscoring the close relationship between these pathways.
Candida albicans is a commensal organism of some mammalian species including humans, where it commonly resides on mucosal surfaces. Under conditions of host immunosuppression C. albicans can invade host tissues to cause a diverse range of diseases. Infection of mucosal surfaces is common, with HIV patients being particularly susceptible to oral and oesophageal candidiasis (de Repentigny et al., 2004), while neutropenic patients are at risk of disseminated disease, with a high mortality rate (Maertens et al., 2001).
Given the importance of autophagy in a diverse array of eukaryotic cellular processes, we decided to establish the role of autophagy in C. albicans survival and differentiation within the host. To date the role the vacuole or indeed autophagy plays during Candida–host interaction has not been established. We hypothesized that autophagy is required for C. albicans adaptation and differentiation, two properties critical for survival within and infection of the mammalian host.
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METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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) were performed within the DS gene program (Accelrys).
Strain construction.
Strains used in this study are described in Table 1. Gene deletion strains were constructed by the PCR-based approach described by Wilson et al. (1999), using the ura3 his1 arg4 strain BWP17, kindly provided by Dr A. Mitchell (Columbia University). atg9 : : ARG4 and atg9 : : HIS1 deletion cassettes were amplified by PCR using pARGSpe and pGEMHIS1 plasmids, respectively, as template with primers ATG9DISF and ATG9DISR (Table 2). BWP17 was first transformed with atg9 : : ARG4 to generate heterozygote strains BAA1 and BAA4. Each heterozygote strain was then transformed with the atg9 : : HIS1 cassette to generate the double deletion strains BAA1H1 and BAA4H4. Correct integration of either cassette was confirmed at each step by PCR analysis using primer pairs ATG9DETF and ATG9DETR, ATG9AMPR and ARG4DET2, or ATG9AMPF and HIS1F1268 (Table 2). Southern blot analysis was also performed using an ATG9-specific probe to the 3'-UTR of ATG9, amplified using primers ATG9PBF and ATG9PBR (Table 2). Correct gene deletion resulted in the replacement of 2858 bp of the 2859 bp ATG9 ORF, with either the 2822 nt HIS1 or 2161 nt ARG4 encoding cassettes. Finally a wild-type copy of ATG9 including 5' and 3' flanking sequences was introduced to the deletion strains on pLA2, to produce a prototrophic ‘reconstituted’ strain. Prototrophic deletion strains were produced by transforming the deletion strains with plasmid vector alone (pLUX). Either plasmid was digested with NheI prior to transformation to target integration into (and reconstitution of) the URA3 loci. The presence/absence of ATG9 in the prototrophic deletion/reconstituted strains was confirmed by amplification using the ATG9DETF/R primer pair.
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), and BAA1H1 and BAA4H4 (atg9/atg9) using the PCR-based tagging approach described by Gerami-Nejad et al. (2001). Primers LAP4GFPF and LAP4GFPR (Table 2
) were designed to amplify a GFP : : URA3 cassette from plasmid pGFPURA3 (Gerami-Nejad et al., 2001) with 78 bp of homology on either end to direct integration at the 3' end of the C. albicans LAP41 gene to form an in-frame LAP41–GFP fusion. The amplified cassette was transformed into the above strains; transformant colonies were selected, and screened for correct integration by PCR. Genomic DNA was prepared from transformants and PCR-amplified using primers LAP4DETR and URA3-5 (Table 2); a product of 470 bp confirmed the expected integration event.
DNA manipulations.
PCR was performed using standard reagents. Plasmids pGEMURA3, pGEMHIS1 and pRSARGSpe (Wilson et al., 1999) were provided by Dr A. Mitchell (Columbia University). Plasmid pLUX (Ramon & Fonzi, 2003), was provided by Dr W. Fonzi (Georgetown University). Plasmids pLA2 and pLA9 were made by amplifying the ATG9 ORF with 693 bp of 5'-UTR and 214 nt 3'-UTR from genomic DNA with ATG9AMPF and ATG9AMPR (Table 2) to incorporate BamHI sites. The resulting product was cloned into the BamHI site of pLUX.
Growth conditions.
Strains were routinely grown on YPD (1 % yeast extract, 2 % Bacto Peptone, 2 % glucose) at 30 °C, supplemented with uridine (25 µg ml–1) when necessary (Guthrie & Fink, 1991). For growth curves, overnight cultures were subcultured to 20 ml fresh YPD medium to OD600 0.2 and incubated at 30 °C with shaking. OD600 was determined from samples taken hourly. Transformants were selected on minimal media [6.75 g l–1 yeast nitrogen base plus ammonium sulfate and without amino acids, 2 % glucose, 2 % Bacto agar (YNB)] supplemented with the appropriate auxotrophic requirements, as described for S. cerevisiae (Burke et al., 2000), except for uridine, which was added at 25 µg ml–1.
Phenotypic assays.
Resistance to temperature stress was determined on YPD agar at 37 and 42 °C, and osmotic stress on YPD agar plus 2.5 M glycerol or 1.5 M NaCl. Secreted aspartyl protease (SAP) secretion was examined on BSA+YE agar (Crandall & Edwards, 1987). Carboxypeptidase Y (CPY) activity was measured using a colorimetric assay as previously reported (Palmer et al., 2003). Resistance to nitrogen starvation was determined using an assay similar to that of Noda et al. (2000). Each strain was grown in YNB broth for 48 h at 30 °C. Cells were washed twice in SD–N and 107 cells resuspended in 2 ml SD–N medium (0.17 % yeast nitrogen base without ammonium sulfate or amino acids, 2 % glucose). Samples taken at intervals were plated to YPD agar, and viability determined as c.f.u. after 2 days at 30 °C. Accumulation of autophagic bodies within the vacuole was assayed by transferring cells grown overnight in YPD to SD–N medium in the presence of 1 mM PMSF (Noda et al., 2000). PMSF inhibits the breakdown of the delivered autophagic bodies. After 6–24 h at 30 °C, cells were examined by differential interference contrast (DIC) microscopy. Filamentation on M199 and 10 % fetal calf serum (FCS) agar was performed as previously described (Palmer & Sturtevant, 2004). Cells from overnight cultures were also induced to filament in 10 % FCS (in distilled H2O) at 37 °C after inoculation at 106 cells ml–1. Chlamydospores were induced on cornstarch-Tween agar as previously described (Palmer et al., 2004). Sensitivity to H2O2 and rapamycin was determined by measuring growth in YPD medium supplemented with the appropriate compound. Approximately 1000 cells from an overnight culture were inoculated to 200 µl growth medium within the wells of a 96-well plate. After 48 h incubation at 30 °C (250 r.p.m.), growth was measured at OD600 using a plate reader.
Phagocytic assays.
The murine macrophage cell line, J774A.1 (ATCC TIB-67) was grown according to ATTC instructions in D-MEM, high glucose, 4 mM glutamine, 10 % fetal bovine serum at 37 °C under 5 % CO2. The same culture conditions were used for incubations with C. albicans. C. albicans strains were incubated with J774A.1 cells as described by Lorenz et al. (2004). Briefly, J774.A1 cells were seeded overnight in 12-well plates (2x105 per well) on 18 mm coverslips and incubated at 37 °C under 5 % CO2. C. albicans strains were grown overnight at 30 °C, washed and incubated with J774A.1 cells at an m.o.i. of 2.0 (4x105 per well) unless otherwise noted.
Macrophage survival.
C. albicans and J774A.1 cells were incubated for 1, 5 and 24 h and then wells were washed and incubated with 0.2 µM calcein AM (final concentration) (LIVE/DEAD Viability/Cytotoxicity Kit, Molecular Probes). C. albicans cells were simultaneously stained with calcofluor (0.225 µM). Coverslips were removed from wells and observed under a fluorescence microscope. Macrophages that fluoresced green were viable. Macrophage survival was quantified by counting at least four fields for each well. Results are presented as the mean number of macrophages observed per field.
C. albicans survival.
This was assessed using an end-point dilution assay as described by Rocha et al. (2001). Macrophages were seeded overnight in 96-well plates at 5x103 per well. C. albicans (50 µl) were added to the first column of cells (150 µl), then serially diluted 1 : 4 for six columns so that the resulting m.o.i. were between 2 and 1.9x10–3. The plates were incubated for 48 h. Controls were wells with C. albicans but no macrophages. The lowest dilution of the control well where it was possible to discriminate distinct colonies was counted. The same dilution was counted for the C. albicans plus macrophage well. Results are presented as (number of colonies in the presence of macrophages/number of colonies in the absence of macrophages)x100. Each experiment was set up in quadruplicate and P values determined using the unpaired Student's t-test.
Vacuole morphology and API localization
Vacuole morphology was visualized using the fluorescent dye FM4-64, as reported (Palmer et al., 2003; Vida & Emr, 1995). In order to localize the API–GFP fusion protein, cells were grown into the exponential phase in YPD medium, stained with FM4-64, washed twice in distilled water, and visualized using an Olympus BX51 fluorescence microscope. All cells were applied to polylysine-coated slides prior to viewing.
Western blot analysis
Western blot analysis was performed basically as described previously (Palmer & Sturtevant, 2004). Cell extracts of C. albicans were prepared by lysis with glass beads in the presence of a protease inhibitor cocktail (Sigma). Lysates were microfuged at 13 000 r.p.m. to yield supernatant and pellet fractions. The pellet was resuspended in 50 µl SDS-PAGE sample buffer. Protein lysates (pellet and supernatant fractions) prepared from equivalent numbers of cells were loaded per lane in 12 % SDS-PAGE gels. Gels were transferred to PROTRAN (Schleicher and Schuell) and blotted with a rabbit polyclonal anti-GFP antibody (Anaspec). Equivalent loading of SDS-PAGE gels was confirmed by staining with Coomassie blue, and membranes were stained with Ponceau Red (Sigma) following the manufacturer's directions. A horseradish-peroxidase-conjugated goat anti-rabbit secondary antibody (Rockland) was used for detection with reagents supplied by Pierce.
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RESULTS |
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; Noda et al., 2000). Deletion of ATG9 results in complete abrogation of autophagy and cvt pathways, leading to a block in API and AMS delivery, sensitivity to nitrogen starvation and an inability to complete sporulation (Lang et al., 2000; Noda et al., 2000). BLAST searches to the C. albicans genome sequence database (www.candidagenome.org/) identified a single ATG9 orthologue, predicted to encode a protein of 952 amino acids, sharing 32 % identity and 47.6 % similarity with S. cerevisiae Atg9p. A Kyte–Doolittle plot of the predicted C. albicans Atg9p sequence suggests six to eight transmembrane domains, which is in good agreement with the five to eight predicted transmembrane domains of the S. cerevisiae protein (Lang et al., 2000; Noda et al., 2000). In order to investigate the importance of autophagy (and the related cvt pathway) in the pathogen C. albicans, we constructed a strain in which both ATG9 alleles were deleted. Correct genotype was checked by both PCR detection and Southern blot analysis. The deletion mutant was readily produced, indicating that this gene is not essential for viability. A wild-type copy of ATG9 including 5' and 3' flanking sequences was then reintroduced to the deletion strain to produce a ‘reconstituted’ strain.
The atg9 mutant had a growth rate comparable to wild-type and reconstituted strains in liquid YPD culture, and was not sensitive to osmotic, temperature or oxidative stresses (data not shown). Levels of vacuolar CPY activity were also determined to be comparable between mutant and control strains, suggesting that trafficking from the Golgi apparatus to the vacuole was not affected in the atg9 mutant (data not shown), and the mutant strain was unaffected in SAP activity (data not shown). The mutant strain was not sensitive to the autophagy-inducing drug rapamycin, or the replication checkpoint inhibitor caffeine (data not shown). Vacuole morphology was analysed using the fluorescent dye FM4-64 (Vida & Emr, 1995). The atg9 mutant was observed to have an intact vacuole morphology, similar to that of ATG9+ control strains (data not shown).
C. albicans atg9 is defective in autophagy
We examined resistance to nitrogen starvation by measuring viability (as c.f.u.) after transfer to medium lacking a nitrogen source (SD–N) (Fig. 1a). Wild-type and reconstituted strains underwent two to three further cell divisions after shifting to SD–N medium, and then maintained viability for the duration of the experiment (30 days). However, the atg9 strain did not continue to divide after the shift to SD–N and c.f.u. steadily declined to 0, clearly demonstrating that atg9 cells are sensitive to nitrogen starvation. We next induced the formation of autophagosomes in SD–N media containing 1 mM PMSF. PMSF is a serine protease inhibitor which blocks the action of vacuolar PrB, an enzyme required for degradation of autophagic bodies within the vacuole (Noda et al., 2000). Under these conditions wild-type and ATG9 reconstituted strains accumulate autophagic bodies within the vacuole, giving the vacuole a granular appearance (Fig. 1b). As expected, the atg9 strain did not accumulate autophagic bodies under these conditions, confirming that our atg9 mutant is defective at an early step of autophagy, possibly autophagosome formation.
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is defective in cvt trafficking), to generate a C-terminal Lap41–GFP fusion, in ATG9+ and atg9 backgrounds. Tagged strains were grown under nutrient-replete conditions (YPD) to the exponential phase of growth, conditions where cvt trafficking occurs but not autophagy, and observed by fluorescent microscopy (Fig. 2a). ATG9+ cells had fluorescence in three distinct patterns: 1, single intense spot outside the vacuole; 2, single intense spot within vacuole lumen; 3, diffuse staining of the vacuole lumen. These patterns resemble API–GFP localization in S. cerevisiae (Suzuki et al., 2002) and correspond nicely with the cvt trafficking events: 1, oligomerized API–GFP precursor within the cytoplasm, or the membrane-bound cvt vesicle prior to vacuolar delivery; 2, cvt body released into the vacuole; 3, mature API–GFP released into the vacuole lumen following degradation of cvt vesicle (Wang & Klionsky, 2003). The distribution of API–GFP in the atg9 background mutant was altered (Table 3). No diffuse staining of the vacuole lumen was detected, and an increased proportion of cells had a single spot distribution. Furthermore, the spots were of increased intensity in the atg9 mutant as compared to the ATG9+ control strains, suggesting an accumulation of API–GFP in the oligomeric cytoplasmic cvt complex.
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). This can be followed as a shift in molecular mass on a Western blot (Klionsky et al., 1992; Suzuki et al., 2002). In order to further confirm the cvt trafficking defects, we immunodetected the API–GFP fusion protein in ATG9+ and atg9 backgrounds, using a polyclonal anti-GFP antibody. The predicted molecular mass of GFP is 26.9 kDa and that of C. albicans Lap41p 56.7 kDa (prior to proteolytic maturation). In ATG9+ strains, delivery of API–GFP to the vacuole would expose the GFP tag to the proteolytically active vacuole lumen, which is liable to have resulted in its cleavage and/or degradation. This may account for the inability to detect the API–GFP fusion species by Western blotting (Fig. 2b), and the fairly low level of fluorescence detected within the vacuole lumen. Unfortunately the anti-GFP antibody recognized a peptide in the minus GFP negative control (lane 1, Fig. 2b), of around 25 kDa. A similar non-specific band has been described in S. cerevisiae (Suzuki et al., 2002) and may have obscured the detection of peptide species of a similar size, including any GFP that is cleaved upon vacuolar delivery. However, a species of 75–80 kDa, corresponding to the predicted API–GFP molecular mass was detected in the atg9 background, indicating accumulation of API–GFP in the atg9 mutant. This species was detected in the pelletable fraction of the cell extracts, but not the soluble fraction. This suggests that the API–GFP has accumulated as part of a large complex before vacuole delivery, perhaps either the highly oligomerized precursor API (cvt body), or even within a cvt vesicle (Yorimitsu & Klionsky, 2005). These data clearly demonstrated that, similar to the S. cerevisiae atg9 mutant, our C. albicans atg9 mutant has a defect in cvt trafficking to the vacuole.
Autophagy is not required for C. albicans differentiation
In the model eukaryote S. cerevisiae, mutants defective in autophagy are unable to complete sporulation (Takeshige et al., 1992; Tsukada & Ohsumi, 1993). Moreover, autophagy has been intimately associated with a wide range of differentiation and development events in higher eukaryotes. We therefore assessed the consequences of a defect in autophagy on two major differentiation processes of C. albicans. Yeast–hypha differentiation was examined on solid and in liquid M199 and FCS media. In each case the atg9 mutant produced filaments that were indistinguishable from the ATG9+ control strains (data not shown), indicating that under in vitro conditions autophagy is not essential for yeast–hypha differentiation. Similarly, when induced on cornstarch-Tween agar, the atg9 mutant produced chlamydospores with similar kinetics and abundance as the ATG9+ control strains (data not shown). Together these results suggest that autophagy is not required for two major differentiation events in C. albicans.
Autophagy is not required for C. albicans survival within or killing of a macrophage cell line
Previous studies using two C. albicans mutants with defects in vacuole biogenesis demonstrated that some vacuolar function(s) are required for survival within, and killing of a mouse macrophage-like cell line (J774A.1) (Palmer et al., 2005). We considered that while not required for yeast–hypha differentiation or chlamydospore formation, autophagy may be important for normal interaction with host cells or survival within host tissues. To test this we analysed the outcome of atg9 interaction with the J774A.1 cell line, by determining macrophage and C. albicans survival rates. Following co-culture, the macrophages rapidly phagocytosed wild-type C. albicans. This was followed by filamentation of C. albicans within the macrophage, which resulted in stretching of the J774A.1 cells, followed by macrophage lysis. The atg9 mutant killed the J774A.1 cells as efficiently as ATG9+ strains (data not shown), indicating that autophagy is not required for killing of the macrophage cell line. C. albicans survival of the macrophage challenge was determined using an end-point dilution assay (Rocha et al., 2001). We found that the atg9 cells exhibited similar susceptibility to the J774A.1 interaction as ATG9+ control strains (data not shown), indicating that this pathway does not aid C. albicans survival within the J774A.1 cells.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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mutant strain constructed within this study was blocked in autophagy and the closely related cvt pathway, and was sensitive to nitrogen starvation. However, the atg9 mutant had a normal growth rate, vacuolar CPY activity, resistance to osmotic and temperature stress, and vacuole morphology. These findings suggest that the atg9 mutant does not have a generalized defect in vacuolar function (Palmer et al., 2005), but a specific defect in cvt/autophagy trafficking.
The atg9 mutant has no detectable defect in either chlamydospore or yeast–hypha differentiation, a result which was surprising given the importance of this pathway in a diverse array of eukaryotic biological processes, including S. cerevisiae sporulation. Microarray studies by Lorenz et al. (2004) demonstrated that upon phagocytosis by J774A.1 cells, C. albicans undergoes a starvation-like transcriptional response, suggesting that the phagolysosome is a nutritionally poor environment. Vacuolar proteases including API, CPY and PrB are upregulated following macrophage ingestion, suggesting that vacuole-mediated proteolysis may be important for survival following phagocytosis. However, our atg9 mutant was not deficient in its ability to survive within or kill a mouse macrophage-like cell line, suggesting that autophagy may not be required for C. albicans–macrophage interaction. This prompted us to conduct a small-scale pilot study to test the virulence of the atg9 mutant in the well-defined mouse model of haematogenously disseminated candidiasis. Only three mice were infected per C. albicans strain, but our results (mean survival time and fungal burden in the kidneys) clearly suggested that the atg9 mutant is fully virulent (data not shown). Taken together, these results indicate that autophagy plays little or no role in C. albicans differentiation or pathogenesis. It remains possible that autophagy is important for the long-term survival of C. albicans as a commensal organism, or to reside on a peripheral host site such as skin, or perhaps within the environment.
Previously we found a C. albicans vps11 mutant to be defective in vacuole biogenesis and to exhibit a range of pleiotropic phenotypes, including a reduced growth rate, sensitivity to a range of stresses and to nitrogen starvation, and reduced vacuolar and secreted protease activities (Palmer et al., 2003, 2005). Moreover, this mutant is unable to undergo yeast–hypha differentiation and was unable to kill the J774A.1 macrophage cell line. Work with a partially functional vps11 allele revealed that restoring a normal growth rate, stress resistance, and SAP and vacuolar hydrolase activities was not sufficient to restore normal filamentation or macrophage killing. Both vps11 mutants have highly fragmented vacuole morphology and are sensitive to nitrogen starvation, suggesting either of these vacuolar functions to be important in yeast–hypha differentiation and macrophage killing. As outlined above, the major role of the vacuole during nitrogen starvation is through the process of autophagy. The results presented here clearly demonstrate that vacuolar functions relating to autophagy are of little consequence in either yeast–hypha differentiation or interaction with macrophages.
Gow & Gooday (1982, 1984) originally observed the vacuole to undergo a dynamic expansion during C. albicans germ-tube emergence from a parental yeast cell. Furthermore, during apical extension of the germ-tube, asymmetrical division of the protoplasm yields subapical compartments composed almost entirely of vacuole, whereas the protoplasm migrates at the hyphal tip (Barelle et al., 2003; Gow & Gooday, 1984). After a delay, the highly vacuolated compartments regenerate cytoplasm and the vacuoles recede. This pattern of vacuole inheritance also seems to influence cell cycle progression and branching frequency of the hyphae (Barelle et al., 2003; Veses et al., 2005). At present the mechanism by which vacuole expansion is mediated, and its importance during host interaction, are unknown. Our results further strengthen the hypothesis that the major role the vacuole plays in yeast–hypha differentiation relates to its morphology, and perhaps its physical expansion to generate ‘empty’ hyphal compartments. Moreover, our results eliminate the autophagic and cvt trafficking pathways as mediating germ-tube induced vacuole expansion.
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ACKNOWLEDGEMENTS |
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Edited by: D Sanglard
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Received 22 August 2006;
revised 23 September 2006;
accepted 26 September 2006.
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