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J. Biol. Chem., Vol. 281, Issue 30, 21445-21457, July 28, 2006
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1


12
From the
Liver Diseases Branch, NIDDK, National Institutes of Health, Bethesda, Maryland 20892 and the
Department of Biology, McGill University, Montreal, Quebec H3A 1B1, Canada
Received for publication, November 30, 2005 , and in revised form, May 3, 2006.
| ABSTRACT |
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| INTRODUCTION |
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Two mammalian proteins with heme transport activity have recently been described. The receptor for feline leukemia virus type C is a transporter required for the development of erythroid cells, and the human ortholog of this transporter can facilitate the export of intracellular heme (5). Shayeghi and colleagues (6) reported the identification of a murine heme transporter, HCP1, expressed on the apical surface of duodenal epithelial cells. This transporter has similarity to bacterial metal-tetracycline transporters, but no clear ortholog has been identified in non-mammalian species. Free-living worms such as Caenorhabditis elegans and parasitic helminths have an absolute requirement for dietary heme transport, as these organisms completely lack the capacity to synthesize heme de novo (7). Heme transporters in these organisms have yet to be identified.
Candida albicans is a dimorphic fungus that is part of the commensal flora of the human gastrointestinal tract and is frequently associated with mucocutaneous infections. In patients with compromised immune systems, Candida species can disseminate hematogenously and cause life-threatening infections. This fungus efficiently uses heme as a nutritional source of iron and exhibits strategies to acquire heme from host erythrocytes. Candida spp. secrete hemolytic factors that lead to the release of erythrocyte hemoglobin (8) and hemoglobin binds to the cell wall of both yeast and hyphal forms of the fungus (9). Recently, a cell surface heme-binding protein, Rbt5, was shown to be involved in heme-iron utilization (10). Although a plasma membrane transport system for heme has not been identified, intracellular degradation of heme through the heme oxygenase encoded by CaHMX1 is required for hemeiron utilization (11, 12).
Although C. albicans and Saccharomyces cerevisiae express very similar systems of iron uptake (13), they differ in their capacity to use heme as a nutritional source of iron. Both species utilize reductive systems of iron uptake and express plasma membrane metalloreductases of the FRE family coupled to a ferrous iron transporter complex that consists of a permease (Ftr1) and a multicopper oxidase (Fet3). Both species also utilize non-reductive systems consisting of siderophore-iron transporters of the ARN/SIT family. Heme uptake in C. albicans occurs independently of the reductive system and submicromolar concentrations of heme or hemoglobin will support vigorous growth of a strain lacking high affinity ferrous iron transport (12). Similar concentrations of heme and hemoglobin do not stimulate growth of a S. cerevisiae strain lacking ferrous uptake (15).
We exploited this difference in heme utilization to identify genes from C. albicans that, when expressed in S. cerevisiae, would enhance heme uptake activity. We identified a family of fungal-specific proteins that are required for the transport of flavin adenine dinucleotide (FAD) into the endoplasmic reticulum (ER),2 where it is required for oxidative protein folding.
| EXPERIMENTAL PROCEDURES |
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strain. The same procedure was used to delete FLC2 in the flc1
strain transformed with plasmid pMET3-FLC1. Clones containing the correctly targeted deletion cassette were plated on 5-fluoroorotic acid-containing plates and 5-fluoroorotic acid-resistant clones were selected. The TET-FLC1flc2
flc3
strain was constructed in R1158 (MATa his3-1 leu2-0 met15-0 URA3::CMV-tTA) (20). The promoter replacement of the FLC1 gene was made as described (20) using primers YPL221W-upRP188 and YPL221W-downRP188. The complete coding sequence for FLC2 was replaced by the nurseothricin resistance gene cassette natMX (21). The complete coding sequence for FLC3 was replaced by the URA3 gene from plasmid Yep352, using primers YGL139W-up (URA3) and YGL139W-down (URA3). All constructs and strains generated were verified by extensive PCR analysis and selection on the appropriate media. The fet3
and fet3
hmx1
strains were constructed as described (22, 23). Rich medium (YPD) and synthetic defined medium (S.D.) were prepared as described (24). Defined iron media were prepared as described (25) using yeast nitrogen base without iron and 1 mM ferrozine, an iron chelator, in addition to the indicated hemin supplement. Lee-Buckley-Campbell media used to induce hyphal growth of C. albicans strains was made as described (26) with addition of 50 µM bathophenanthroline disulfonate and 1 µM hemin. Milk-Tween agar media was prepared as described (27). To create hypoxic conditions, cultures were incubated in an anaerobic chamber using a BBL-GasPak (BD Biosciences) for 5 h. Plasmids and LibrariesA C. albicans genomic DNA library was constructed in a S. cerevisiae 2-µm vector and was a kind gift from J. Berman. The S. cerevisiae genomic DNA library in 2-µm vector YEp13 was obtained from ATCC. The CaFLC1 open reading frame (ORF) was amplified using primers CA_BMST_F and ca_NCSM_R, digested with SacI and NcoI, and ligated into the SacI and NcoI sites of the vector YIpDCE51 (23) to construct plasmid YIpDCE1-CaFLC1. This plasmid was linearized with StuI for integration at ADE2. Plasmids pCaFLC1, pFLC1, pFLC2, and pFLC3 were constructed using vector pYX242 and the PCR-amplified ORFs of the corresponding genes. For pCaFLC1, primers CA_BMST_F and ca_NCSM_R were used and the library clone containing CaFLC1 was the template. The PCR product was digested with BamHI and SmaI and inserted into the same sites of the vector. Primer pair YPL221-F and YPL221-R was used to amplify FLC1, YAL053-F and YAL053-R for FLC2, and YGL139-F and YGL139-R for FLC3. Genomic DNA from S. cerevisiae YPH499 was used as the template to amplify FLC1, FLC2, and FLC3. FLC1 was inserted into EcoRI/SmaI sites, FLC2 into HindII/SmaI, and FLC3 into NcoI/ApaI of pYX242.
To construct epitope-tagged genes, a triple copy of the HA tag was amplified by PCR using primers 221-cHA-F and 221-cHA-R for FLC1 and in vivo recombination in yeast was used to insert this PCR product into the C terminus of FLC1 in plasmid pYX-FLC1 cut with SmaI. The resulting plasmid, pYX-FLC1-HA, was subsequently rescued from yeast and amplified in Escherichia coli. The same method was used to make plasmids expressing pYX-FLC2-HA and pYX-FLC3-HA. Primers 221-cHA-R and YAL053-cHA-F were used for pYX-FLC2-HA. Primers 221-cHA-R and YGL139-cHA-F were used for pYX-FLC3-HA. Functionality of carboxyl terminus HA-tagged versions of FLC1, FLC2, and FLC3 was confirmed by complementation of the flc1
flc2
pMET3-FLC1 strain on media supplemented with methionine.
To make plasmid pRS416-FLC1-HA, the promoter region of FLC1 was PCR amplified from genomic DNA using primers prom-YPL221-F and prom-YPL221-R and digested with KpnI and EcoRI. This KpnI/EcoRI fragment and EcoRI/SmaI fragment from pYX-FLC1-HA were subsequently ligated into Kpn/EcoRI and EcoRI/SmaI sites of pRS426. KpnI/NotI fragment was cut from this plasmid and ligated into Kpn/NotI sites of pRS416. To make plasmid pRS415-FLC2-HA, a PstI/BamHI fragment from the FLC2 library clone containing the FLC2 upstream sequences and a BamHI/SacII fragment of the FLC2 ORF from plasmid pYX-HUF2-HA were ligated into the PstI/SacII sites of pRS415. To make plasmid pRS413-FLC3-HA, a HindIII/EcoRI fragment containing the promoter region of FLC3 from the FLC3 library clone and the EcoRI/SmaI fragment from pYX-FLC3-HA were ligated into HindIII/SmaI sites of pRS415. Then the ApaI/SmaI fragment of this plasmid was ligated into the same sites of pRS413. To make plasmid pMET3-FLC1, the EcoRI/SmaI fragment of the FLC1 gene from pYX-FLC1-HA was inserted into vector pRS313-MET3 cut with the same restriction enzymes.
To make plasmid pVAN1, the ORF of VAN1 was amplified by PCR from the VAN1 library clone using primers Van1-pr-F and Van1-pr-R and digested with XbaI and SmaI. This fragment was inserted into XbaI/SmaI sites of YEp351. The plasmid pKTR4 was made by inserting the KTR4 gene into the PstI/HindIII sites of YEp351. KTR4 was amplified by PCR using primers KTR4-pr-f and KTR4-pr-r from the KTR4 library clone. Plasmid pPTC1 was made by inserting the PTC1 gene into the XbaI/SmaI sites of YEp351 using primers PTC1+pr-F and PTC1+pr-R from the PTC1 library clone. Plasmid pHOC1 was made by inserting HOC1 into XhoI/SacI sites of pRS426 using primers Hoc1-t-F and Hoc1-t-R and genomic DNA of S. cerevisiae YPH499 as the template. To make plasmid pMNN9, the MNN9 gene was inserted into the XbaI/ClaI sites of pRS426 using primers MnnI-t-F and MnnI-t-R and genomic DNA as template. Plasmid pMET3-CaFLC1 was constructed by subcloning the SmaI/BamHI fragment of pCaFLC1 with CaFLC1 ORF into vector pCaEXP (28). Vector was digested with PstI, treated with Klenow fragment, and then digested with BamHI. Plasmid was linearized with StuI and integrated into the RP10 locus of the C. albicans genome. Plasmids pGAL-FLC1 (pBG1800-YPL221W) and pGAL1-FAD1 (pBG1805-YDL044C) were obtained from the YEAST ORF collection (Open Biosystems). Functionality of the cloned genes was confirmed by sequencing and/or phenotypic analysis. Plasmid pJS401 with UPRE-LacZ was a gift from D. Griffith (29). Immunofluorescence, Fractionation, and Western BlottingThe strain CRY (MATa his3 ura3 leu2 trp1 ade2) (gift from E. Cabib) was transformed with centromeric plasmids containing HA-tagged FLC genes and transformants were grown to mid-log phase and the cells were prepared for immunofluorescence microscopy as described (30). Subcellular fractionation was performed as described (31) with the following modifications. Cells were disrupted with glass beads, and unbroken cells were removed by centrifugation at 500 x g for 2 min. Cell lysate was applied to the top of 20-60% continuous sucrose gradient. Samples were centrifuged at 28,500 x g for 17 h and 0.9-ml fractions were collected. Lysates and gradients were prepared with EDTA or 2 mM MgCl2. Western blotting was performed using a 1:4000 dilution of HA.11 (Covance) as the primary antibody followed by 1:3000 dilution of horseradish peroxidase-conjugated sheep anti-mouse antibody (Amersham Biosciences). Antibody to Dpm1, porin, and Vps10 were purchased from Molecular Probes and used according to the manufacturer's manual. Anti-Gas1p antibody was used at 1:5000 dilution (32). Antibody was detected using enhanced chemiluminescence (Amersham Biosciences).
Radiolabeling and ImmunoprecipitationsMetabolic labeling and immunoprecipitation was performed as described (33). Briefly, cells were grown overnight in SD (-ura) with 2% raffinose as the carbon source and 0.2% galactose. Glucose was added to 2% and cells were incubated 24 h more. Cells were pulsed-labeled with L-[35S]methionine for 12 min and chased with unlabeled cysteine and methionine for the indicated number of minutes. Immunoprecipitation was performed using a monoclonal anti-CPY antibody (Molecular Probes). The immunoprecipitate was analyzed by SDS-PAGE and phosphorimaging.
Electron MicroscopyS. cerevisiae R1158 and the triple mutant TET-FLC1flc2
flc3
strains were grown to an A600 of 0.2 in liquid YPD at 30 °C with shaking. Cultures were divided, and doxycycline was added to a final concentration of 10 µg/ml; the other half did not receive any additions and was used as control. Subcultures were incubated an additional 9 h. Cells were pelleted and washed twice with water and then fixed in 1.5% (w/v) potassium permanganate for 20 min at room temperature with occasional mixing. The cells were then washed with water, incubated in 0.5% sodium periodate for 20 min at room temperature, washed with water, incubated for 15 min in 1% ammonium chloride, washed with water again, and left overnight in 1% uranyl acetate at 4 °C. The next day cells were washed several times with water, dehydrated in an ethanol/water series, and finally embedded in Epon 812 resin. Ultrathin sections were viewed with a JEOL 2000 electron microscope.
AssaysNorthern blot analyses were performed as described (25). Probes for CaFLC1 and ACT1 were prepared from PCR products corresponding to the open reading frame. Heme uptake was measured as described (34) with the following modifications. Yeast were grown in SD medium overnight and then diluted to an absorbance of 0.1 and incubated 5 h. Washed cells were suspended in phosphate-buffered saline containing 5% glucose, 0.05% Tween 80, and 0.5% bovine serum albumin and preincubated for 10 min at 37 °C. 55Fe-hemin was added at a final concentration of 2.3 µM to 0.1 ml of cell suspension with a final absorbance of 1.0 and incubated 60 min at 37 or 4 °C. To stop the reaction, 20 µl of 1 mM cold hemin was added and cell suspensions were transferred to microfilter plates on ice. The cells were washed 6-7 times with buffer without glucose. Accumulation of 55Fe was measured by scintillation counting. Heme uptake was reported as the difference between 55Fe accumulation at 37 and 0 °C. 55Fe-hemin was synthesized as described (35) from protoporphyrin IX (Porphyrin Products, Logan, UT) and 55FeCl3 (PerkinElmer Life Sciences).
Alcian blue binding was measured as described previously (36-38). Briefly, the stain (0.1%) was prepared in 0.02 N HCl and the suspension was centrifuged to eliminate insoluble precipitates. The cells were grown for 48 h in S.D. (-Met), then diluted to an absorbance of 0.2 and incubated 2 days in SD with or without methionine. An absorbance of 1.0 of cells was applied to microfilter plates. Cells were washed two times with 0.2 ml of 0.9% NaCl and then supplemented with 0.2 ml of Alcian blue solution. The mixture was maintained statically at room temperature for 10-15 min and then washed twice with 2 ml of 0.02 N HCl.
-Galactosidase assays were performed as described (39).
FAD uptake assays were performed using transport-competent membranes in permeabilized cells, and were prepared essentially as described (40, 41) with the following modifications. Cells from 50-100 ml of culture were harvested and suspended at a density of 50 A600 units/ml in 100 mM Tris-HCl, pH 9.4, and 10 mM dithiothreitol and incubated 5 min at room temperature. Cells were centrifuged and re-suspended in 1 ml of 0.75 x SC with or without 1 mM methionine, 0.5% glucose, 0.7 M sorbitol, and 10 mM Tris-HCl, pH 7.5, with 0.5 mg/ml zymolyase T-100. After 30 min incubation at 30 °C, spheroplasts were collected and resuspended in 0.75 x SCM containing 0.7 M sorbitol and 1% glucose with or without 1 mM methionine. Cells were allowed to recover at 30 °C for 20 min, then washed with lysis buffer (0.4 M sorbitol, 20 mM HEPES, pH 6.8, 150 mM potassium acetate, 2 mM magnesium acetate). Cells were resuspended in lysis buffer at 200 A600 units/ml. Aliquots of suspensions were slowly frozen over liquid nitrogen for 60 min and stored at -80 °C. Uptake of FAD was measured as described (42). Permeabilized yeast cells were thawed quickly and washed three times with ice-cold reaction buffer (20 mM HEPES, pH 6.8, 150 mM potassium acetate, 250 mM sorbitol, 5 mM magnesium acetate) and then resuspended in the same buffer. Protein concentrations in the samples were normalized using the bicinchoninic acid method (Pierce). Membranes were incubated in 1 mM FAD at 30 °C for the indicated period of time. Uptake was terminated by the addition of 1 ml of ice-cold reaction buffer and samples were quickly centrifuged 16,000 x g for 1 min. Membranes were washed twice in reaction buffer and solubilized in 2% Triton X-100. FAD content was measured fluorometrically at an excitation wavelength of 450 nm and an emission wavelength of 530 nm on an ISS PC fluorescent spectrophotometer. Fluorescence of vesicles incubated in the absence of FAD served as a background control.
| RESULTS |
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strain (Fig. 1B). Using a C. albicans genomic library cloned into a S. cerevisiae high copy expression vector, we identified a plasmid that stimulated the growth of the fet3
strain only in the presence of hemin. An analysis of the nucleotide sequence of this clone indicated the presence of two C.albicans genes, orf19.2501 and orf19.2503. Subsequent subcloning of these genes revealed that only orf19.2501 conferred the utilization of hemin as an iron source in the S. cerevisiae fet3
strain (Fig. 1B). This gene was termed CaFLC1 for flavin carrier 1 after subsequent characterization revealed a role in FAD transport. To determine whether CaFLC1 stimulated the uptake of intact hemin or whether it stimulated the extracellular release of iron from hemin, we expressed CaFLC1 in a strain from which both the intracellular heme degrading enzyme, HMX1 (22), and FET3 had been deleted. Expression of CaFLC1 in a strain lacking Hmx1p resulted in slower growth on hemin when compared with the congenic strain expressing Hmx1p, suggesting that CaFLC1 expression led to the uptake of intact hemin and that intracellular degradation of this hemin was needed for growth.
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We examined the cellular localization of CaFlc1p by constructing a strain in which a triple HA tag was introduced to the carboxyl terminus of FLC1 and the fusion gene under the control of the MET3 promoter was integrated at the RP10 locus. By indirect immunofluorescence, Flc1-HA was localized exclusively to punctate, intracellular vesicles, with no detectable signal on the plasma membrane or vacuolar membranes (Fig. 3, A and B).
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We questioned whether overexpression of S. cerevisiae FLC1, FLC2, or FLC3 in S. cerevisiae could also confer growth on hemin in a manner similar to that of CaFLC1. The ORFs corresponding to FLC1, -2, and -3 were cloned into a high copy vector under the control of the strong TPI1 promoter and the resulting plasmids were transformed into the fet3
strain. Transformants were grown in iron-depleted medium, then plated in serial dilutions on iron-depleted medium containing hemin. Overexpression of both FLC1 and FLC2, but not FLC3, could stimulate the growth of the fet3
strain when hemin constituted the sole source of iron (Fig. 4), whereas overexpression had no effect on growth in the fet3
strain in the absence of heme (data not shown), suggesting that the FLC genes may function similarly in S. cerevisiae and C. albicans.As S. cerevisiae is a well established and convenient model for studying eukaryotic gene function, we examined the function of the FLC genes endogenous to S. cerevisiae.
Localization of Flc Proteins to the Endoplasmic ReticulumBecause deletion and overexpression of CaFLC1 was correlated with changes in heme uptake activity in C. albicans and because the predicted amino acid sequences of the FLC genes suggested a polytopic integral membrane protein, we initially hypothesized that the FLC genes were directly involved in heme uptake on the cell surface. Although CaFlc1p was expressed in intracellular vesicles and not detected on the plasma membrane, other transporters involved in metal uptake in yeast are expressed on intracellular vesicles (30, 46). We therefore examined the cellular localization of the Flc proteins in S. cerevisiae. Overexpression of Flc1p from a strong promoter resulted in the detection of Flc1p on membranes throughout the cell (data not shown). Therefore, we constructed centromeric plasmids containing FLC1, -2, or 3 with a carboxyl-terminal triple HA tag in which each gene was expressed from its endogenous promoter. Functionality of the tagged alleles was confirmed by complementation of the flc1
flc2
strain (see Fig. 6). Strains transformed with these plasmids were examined by indirect immunofluorescence, and Flc1p-HA was detected at the periphery of the cell and in a ring-like intracellular structure (Fig. 5, A and B). This intracellular structure co-localized with the nucleus by 4',6-diamidino-2-phenylindole staining, and this pattern of peripheral and perinuclear signal is typical of the yeast ER, and typically does not indicate plasma membrane localization (47-49). These data suggested that Flc1p was localized to the ER; however, localization of Flc2p and Flc3p could not be confirmed by this method, as the fluorescent signal was not detectable.
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strain or FLC1 in the flc2
strain failed, and we considered the possibility that FLC1 and FLC2 might be synthetically lethal. To test this, we cloned the FLC1 ORF into a low copy plasmid under the control of the methionine-regulatable MET3 promoter, and transformed the resulting plasmid, pMET-FLC1, into the flc1
strain. FLC2 was successfully deleted in this strain when it was maintained in methioninefree medium and Flc1p was expressed from the plasmid. We tested whether deletion of both FLC1 and FLC2 was lethal by plating the congenic strains containing pMET-FLC1 on either medium without methionine (on which plasmid-born Flc1p is expressed) or on medium with methionine (on which plasmidborn Flc1p is not expressed at significant levels). Although all strains grew equally well on medium without methionine (Fig. 6A), the flc1
flc2
strain failed to grow on medium with methionine (Fig. 6B), indicating that FLC1 and FLC2 were synthetically lethal and together encoded an essential function in yeast. Notably, the addition of hemin in various concentrations to plates containing methionine did not rescue the lethality of the flc1
flc2
strain. Similarly, the addition of long chain fatty acids and ergosterol, the products of essential heme-dependent biosynthetic pathways, did not rescue the lethality of the flc1
flc2
strain. These results again suggested that the transport of heme was not the essential function of Flc1p and Flc2p and that the essential function of the FLC genes was unknown.
Impaired Cell Wall Integrity in Strains Deleted for FLC1 and FLC2To gain clues as to the essential function of Flc1p and Flc2p, we tested various compounds for their effects on the growth of FLC-deleted strains. Addition of 1 M sorbitol to media containing methionine resulted in a partial restoration of growth to the flc1
flc2
strain (Fig. 7B). Osmotic stabilizers such as sorbitol can rescue the growth of strains with defective cell walls. Glucosamine, which can be used by the cell for the synthesis of chitin, glycosylphosphatidylinositol anchors, and N-glycosylated proteins (52), also partially rescued the growth of the flc1
flc2
strain (Fig. 7C). Chitin is an essential carbohydrate component of the cell wall (53). Calcofluor White is a fluorescent dye that binds to chitin, and mutant strains with defects in cell wall synthesis frequently exhibit sensitivity to this agent as well as increased deposition of chitin in the cell wall (53). The flc2
strain exhibited sensitivity to Calcofluor White (Fig. 7D) and another chitin-binding dye, Congo Red (data not shown). Furthermore, microscopic examination of the flc1
flc2
strain stained with Calcofluor White revealed increased chitin deposition in the cell wall (Fig. 7E), especially at the bud neck, the site of maximal cell wall synthesis and chitin deposition, and bud scars. This increase in staining was not present in the flc1
flc2
strain or the wild type strain when Flc1p was overexpressed (Fig. 7, F and G). These results all suggested that the flc1
flc2
strain was impaired in some aspect of cell wall biosynthesis.
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flc2
Synthetic LethalityTo further investigate the essential function of Flc1p and Flc2p, we screened a yeast genomic library for genes that, when expressed from a high copy plasmid, could rescue the lethality and permit the growth of the flc1
flc2
strain on methionine-containing medium. Plasmids identified in this screen were isolated and retransformed into the flc1
flc2
strain to confirm the suppressor phenotype. Plasmids were then sequenced and the individual ORFs within each plasmid were subcloned into the high copy vector and retested. The genes identified in this screen are presented in Table 2. In addition to FLC1 and FLC2, we determined that both FLC3 and CaFLC1 rescued the lethality of the flc1
flc2
strain, which indicated that each of these genes encoded a protein of similar function.
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flc2
strain. Mannoproteins form the outer layer of the yeast cell wall and two
-1,6-mannan polymerase complexes (M-Pol I and M-Pol II) operate in the Golgi apparatus to attach the long mannan backbone found on certain glycoproteins (56). We tested and found that, in addition to VAN1, overexpression of HOC1, encoding a component of M-Pol II (55), could also suppress the lethality of the flc1
flc2
strain. However, overexpression of MNN9, encoding a component of both M-Pol I and M-Pol II, did not. In a separate screen for S. cerevisiae genes that could facilitate the uptake of hemin when overexpressed, we identified KTR4, a gene that exhibits homology to the KTR family of
-1,2-mannosyltransferases involved in O-linked glycosylation and N-linked outer chain mannosylation (57). Overexpression of KTR4 also suppressed the lethality of the flc1
flc2
strain. MSG5, a protein phosphatase involved in signaling through the cell integrity pathway (58), and PTC1, a protein phosphatase involved in signaling through the mitogen-activated protein kinase osmosensing cascade (59) were also identified as suppressors. Strains with defects in cell wall biogenesis have been reported to become dependent on stress-response signaling pathways to remain viable (60, 61). Overexpression of these phosphatases could alter the activity of stress-response signaling pathways and permit the continued growth of the flc1
flc2
strain. APM2 encodes an adaptor-like protein that may be involved in protein or vesicular trafficking, and may suggest that alterations in these processes facilitate heme uptake or contribute to the lethality of the flc1
flc2
strain. Taken together, the genes identified in the suppressor screen suggested that the lethality of the flc1
flc2
strain could be rescued by an increase in the activity of outer chain mannosyltransferases.
Impaired Synthesis of Cell Wall Components in the flc1
flc2
StrainIn yeast, subsequent to the attachment of the long
-1,6-mannose polymers by M-Pol I and II, branches are added by additional mannosyltransferases, and phosphomannose residues are attached at some of these branches (56). Negatively charged phosphomannose residues can be detected in the cell wall by staining with the cationic dye Alcian blue (36), and we tested the congenic wild type, flc1
, flc2
, and flc1
flc2
strains for the presence of phosphomannose residues in the cell wall by Alcian blue staining (Fig. 8A). The flc1
strain exhibited a modest decrease in staining when compared with the wild type, whereas the flc1
flc2
strain exhibited a more dramatic decrease in staining, indicating that the loss of FLC gene expression led to a decrease in the phosphomannose content of the cell wall.
We examined the cell wall of a FLC-depleted strain by transmission electron microscopy. A strain in which FLC2 and FLC3 were deleted and FLC1 was controlled by the tetracycline-regulatable promoter (20) and the congenic parent strain were grown in the presence of doxycycline to shut off FLC1 expression. Electron microscopic analysis revealed that the FLC-depleted strain exhibited a thickened cell wall and an accumulation of amorphous material in the developing bud (Fig. 8B). In vitro synthesis of
-1,6-D-glucan, a major component of the yeast cell wall, was also impaired in this strain (data not shown). These data indicated that in the absence of FLC gene expression, yeast cells exhibit defects in both the mannoprotein component and the glucan component of the cell wall.
Impaired FAD Transport and Oxidative Protein Folding in the flc1
flc2
StrainBoth N-linked core glycosylation of and disulfide bond formation in proteins within the secretory pathway occur in the ER and these processes are highly conserved among eukaryotes (53, 62). Core N-glycosylation precedes and is required for outer chain mannosylation in the Golgi. Disulfide bond formation requires the oxidizing environment of the ER and proteins lacking properly formed disulfide bridges are retained in the ER. We examined the flc1
flc2
strain for defects in early steps of N-glycosylation and oxidative protein folding by following the maturation of carboxypeptidase Y (CPY). Native CPY contains five disulfide bonds and undergoes N-glycosylation in the ER and Golgi before undergoing proteolytic processing to its mature form in the vacuole (63). Defects in N-glycosylation do not prevent the sorting of CPY to the vacuole, whereas failure to form disulfide bonds causes CPY to accumulate in the ER. Using strains bearing a copy of FLC1 under the control of the GAL1,10 promoter, CPY maturation was monitored using pulse-chase and immunoprecipitation (Fig. 8C). In wild type cells, most of the ER form of CPY was processed after 5 min and none of the ER form was detectable after 10 min of chase. In contrast, in the flc1
flc2
strain most of the ER form of CPY was still present after 5 min and the ER form remained readily detectable after 10 min of chase. The electrophoretic mobilities of the ER and Golgi forms of CPY were similar to those of the wild type, however, suggesting that although exit from the ER was delayed, core N-glycosylation was intact.
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flc2
strain. Oxidative protein folding in the ER is driven by a protein relay that transfers oxidizing equivalents to folding proteins, and the essential, ER luminal protein Ero1p initiates this process (64). Ero1p is an FAD-dependent enzyme, and Ero1p activity is highly sensitive to changes in free FAD levels. FAD is briskly transported into the ER, but the gene(s) encoding this transport activity have not been identified in any organism. Because ero1-1 mutants can be rescued by overexpression of FAD synthetase (65), we tested and observed that overexpression of this enzyme also partially rescued the lethality of the flc1
flc2
strain (Fig. 9A), and hypothesized that the FLC genes might be involved in the transport of FAD into the ER.
We measured the transport of FAD into microsomes by growing congenic FLC deletion strains in medium with or without methionine supplementation for 5 h, then incubating permeabilized cells in FAD and measuring retained FAD by fluorescence spectrophotometry. After 5 h in methionine-containing medium, the flc1
flc2
culture was still increasing in density, albeit very slowly, and the cells retained a trace amount of Flc1p expression from the methionine promoter (data not shown). We found that, whereas wild type cells exhibited a robust uptake of FAD, the flc1
flc2
strain exhibited very low uptake of FAD when grown in methionine-containing medium (Fig. 9B, left panel). Transport of FAD into wild type microsomes was time dependent and reached a maximum at 10 min (Fig. 9B, right panel), which was similar to the FAD transport kinetics identified in rat liver microsomes (42). We measured only a small amount of time-dependent FAD transport into microsomes from the flc1
flc2
strain. Similar to rat liver microsomes, FAD uptake into wild type microsomes was inhibited by the anion transport inhibitor 4,4'-diisothiocyanostilbene-2,2'-disulfonic acid, and insensitive to EDTA (data not shown). This loss of FAD transport activity was apparent after only5hof methionine-induced Flc1p shut-off, whereas other phenotypes of the flc1
flc2
strain were observed after 16-24 h after shut-off, suggesting that the loss of FAD transport might be a direct consequence of Flc protein depletion.
Activation of the Unfolded Protein Response in the flc1
flc2
StrainDefects in FAD-dependent disulfide bond formation in the ER can lead to misfolding, aggregation, and retention of proteins within the ER. Yeast cells respond to the accumulation of misfolded proteins in the ER by activating the unfolded protein response (UPR), a signal transduction pathway between the ER and the nucleus (66). Activation of the UPR leads to the transcription of genes containing a UPR response element (UPRE) in their promoter regions. We addressed whether oxidative protein folding defects in strains lacking FLC genes would activate the UPR by transforming congenic FLC deletion strains with a UPRE-LacZ reporter plasmid (29) and measuring
-galactosidase activity in media with and without methionine (Fig. 9C). The flc1
flc2
strain exhibited approximately five times the activity of the wild type strain when Flc1p was not expressed, indicating the accumulation of misfolded proteins in the ER.
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strain, and determining whether FAD could inhibit growth of this strain on medium containing hemin as the source of iron (Fig. 10). Whereas CaFlc1p could stimulate the growth of the fet3
strain on hemin (Fig. 10A), the addition of excess FAD completely blocked the stimulatory effects of CaFlc1p (Fig. 10B). FAD had no effect on the slow growth of the fet3
strain in the absence of CaFlc1p. These data are consistent with a mechanism in which heme is a low affinity substrate for an FAD transporter encoded by FLC family members. | DISCUSSION |
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flc2
strain, and thereby supply the essential function lacking in this strain, they are all strongly predicted to have similar, if not identical, biochemical activities. The Role of the FLC Genes in FAD TransportAmple biochemical evidence supports the existence of a robust FAD transport system localized to the membranes of the ER. In yeast, FAD rapidly enters purified microsomes, where it can directly bind to and induce the oxidation of luminal proteins (see below) (65, 67). Biochemical characterization of FAD transport in rat liver microsomes suggests a similar type of transport mechanism in which uptake is initially rapid, then plateaus. FAD transport is bidirectional, as an equally rapid FAD efflux system is also present. Neither process requires exogenous sources of energy and both are inhibited by anion transport inhibitors (42). The FAD transport activities measured in these yeast studies exhibits kinetic characteristics nearly identical to those reported for the mammalian microsomes. These kinetic observations also raise the possibility that microsomal FAD transport may represent facilitated diffusion or a carrier-mediated transport process in which the transporter permits free cytosolic FAD to equilibrate within the lumen of the ER. This is supported by our observation that overexpression of Flc1p did not result in higher levels of microsomal FAD accumulation in wild type cells (Fig. 9B). Although a mitochondrial carrier for FAD (Flx1p) has been identified in yeast (68), genes encoding a microsomal FAD carrier have not been identified previously in any eukaryote. Because the loss of FAD transport in our studies occurred within5hof methionine-induced Flc1p shut-off, and because this phenotype preceded the appearance of other phenotypes by several hours, loss of FAD transport may be a direct result of FLC gene deletion. The Flc proteins do not exhibit homology to any known family of carrier proteins, but members of this family exhibit significant sequence conservation and marked similarity in their hydrophobicity profiles. Depending on the prediction program used, each family member is predicted to have 9-10 transmembrane domains, and thus could function directly as an FAD transporter. Alternatively, the Flc proteins could indirectly affect FAD transport through interactions with an unidentified downstream partner.
The Role of FAD in Oxidative Protein FoldingProtein disulfide formation is a crucial step in protein folding and is required by many proteins that traverse the secretory pathway. A protein relay delivers oxidizing equivalents to folding proteins in the ER. This process is initiated by the essential ER luminal protein Ero1p (69, 70), which catalyzes a disulfide exchange to oxidize protein-disulfide isomerase (PDI) (71). Protein-disulfide isomerase can then catalyze the oxidation and rearrangement of disulfide bonds in folding proteins (72). Ero1p is an FAD-binding protein (65), and the activity of Ero1p is highly dependent on free FAD levels (65). Depletion of cellular riboflavin (and subsequently FAD) leads to a profound defect in oxidative protein folding, and mutations in Ero1p can be rescued by increased synthesis of FAD through overexpression of FAD synthetase. Two other proteins involved in oxidative protein folding, Erv1p and Fmo1p, also require FAD for activity (64).
Loss of FAD transport activity at the ER membrane, as occurred in the FLC-deleted strains, would be predicted to result in depletion of intraluminal FAD and loss of Ero1p activity. In the absence of active Ero1p, misfolded proteins would be expected to accumulate and trigger the UPR. Accordingly, deletion of the FLC genes resulted in activation of a UPR reporter construct (Fig. 9C). Loss of Ero1p function, which is essential in yeast, also explains the synthetic lethality of the flc1
flc2
strain. Loss of Ero1p function would ultimately lead to depletion of active forms of many proteins within the secretory pathway, and likely accounts for the pleiotropic phenotypes observed in the flc1
flc2
strain, including morphologically aberrant cell walls with loss of
-1,6-glucan synthase activity and depletion of phosphomannoproteins. Depletion of a Schizosaccharomyces pombe FLC orthologue, PKD2, also resulted in altered cell morphology and reduced glucan levels (73). Strains carrying mutations in PDI1 have also been reported to exhibit phenotypes associated with cell wall defects, such as sensitivity to caffeine and zymolyase (14).
We observed that the processing of CPY in the ER, which requires the formation of five disulfide bonds, was slightly delayed in the flc1
flc2
strain in which expression of GALFLC1 was repressed by glucose (Fig. 8C). Strains lacking Ero1p or Pdi1p typically exhibit more pronounced delays in the processing of CPY, suggesting that the ER is not completely depleted of FAD and there is residual Ero1p activity after deletion of FLC1 and FLC2. This could be explained by trace levels of expression of pGAL-FLC1 or native FLC3, or by a process in which the cells stop dividing before the ER becomes fully depleted of FAD.
Members of the FLC family of genes are present in essentially every fungal genome examined to date (45). Yet no clear homologues to these genes are present in other eukaryotic genomes, despite the kinetically similar FAD transport process in mammals and clear conservation of the FAD-dependent oxidative protein folding machinery in higher eukaryotes. One possibility is that sequence homology has fallen below a level that is detected in conventional blast searches. Further investigation will be required to identify FAD transporters in these organisms.
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1 Both authors are supported by the Intramural Research Program of the NIDDK, National Institutes of Health. ![]()
2 To whom correspondence should be addressed: Bldg. 10, Rm. 9B-16, 10 Center Dr., MSC 1800, Bethesda, MD 20892-1800. Tel.: 301-435-4018; Fax: 301-402-0491; E-mail: carolinep{at}intra.niddk.nih.gov.