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Originally published In Press as doi:10.1074/jbc.M403574200 on June 1, 2004

J. Biol. Chem., Vol. 279, Issue 34, 35644-35655, August 20, 2004
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Five Genes Involved in Biosynthesis of the Pyruvylated Gal{beta}1,3-Epitope in Schizosaccharomyces pombe N-Linked Glycans*

Ekaterina N. Andreishcheva{ddagger}§, Jeremy P. Kunkel{ddagger}, Trent R. Gemmill{ddagger}, and Robert B. Trimble{ddagger}||**

From the {ddagger}Wadsworth Center, New York State Department of Health, Albany, New York 12201-0509 and the ||Department of Biomedical Sciences, School of Public Health, State University of New York at Albany, Albany, New York 12201

Received for publication, March 31, 2004 , and in revised form, June 1, 2004.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
The N-linked galactomannans of Schizosaccharomyces pombe have pyruvylated Gal{beta}1,3-(PvGal) caps on a portion of the Gal{alpha}1,2-residues in their outer chains (Gemmill, T. R., and Trimble, R. B. (1998) Glycobiology 8, 1087–1095). PvGal biosynthesis was investigated by ethyl methanesulfonate mutagenesis of S. pombe, followed by the isolation of cells devoid of negatively charged N-glycans by Q-Sepharose exclusion and failure to bind human serum amyloid P component, which acts as a lectin for terminal PvGal residues. Mutant glycans were characterized by lectin binding, saccharide composition, exoglycosidase sensitivity, and NMR spectroscopy. Restoration of the cell surface negative charge by complementation with an S. pombe genomic library led to the identification of five genes involved in PvGal biosynthesis, which we designated pvg1–pvg5. Pvg1p may be a pyruvyltransferase, since NMR of pvg1– mutant N-glycans revealed the absence of only the pyruvyl moiety. Pvg2p–Pvg5p are crucial for attachment of the Gal{beta}1,3-residue that becomes pyruvylated. Pvg3p is predicted to be a member of the {beta}1,3-galactosyltransferase family, and Pvg3p-green fluorescent protein labeling was consistent with Golgi localization. Predicted Pvg1p and Pvg3p functions imply that Gal{beta}1,3-is added to the galactomannans and is then pyruvylated in situ, rather than by an en bloc addition of PvGal{beta}1,3-caps to the outer chain. Pvg4p-green fluorescent protein targeted to the nucleus, and its sequence contains a MADS-box DNA-binding and dimerization domain; however, it does not appear to solely control transcription of the other identified genes. Pvg2p and/or Pvg5p may contribute to an enzyme complex. Whereas a functional role for the PvGal epitope in S. pombe remains unclear, it is nonessential for either cell growth or mating under laboratory conditions.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Nearly all of the proteins found on the cell surfaces of almost all organisms are glycosylated. In most cases, the glycan moieties associated with these glycoproteins provide the surface with a net negative charge, which plays a significant role in the regulation of the cell influx/efflux processes and in cell-cell interactions such as cell aggregation and pathogenic adhesion. Whereas a wide range of acidic saccharides and saccharide modifications (e.g. muramic acid, hexuronic acids, sialic acids, succinate, lactate, and pyruvate) are found in bacterial polysaccharides (1–6), in general, sialylated and (less frequently) sulfated glycans are associated with vertebrate glycoproteins. In several yeast species (i.e. Saccharomyces cerevisiae, Candida albicans, Pichia holstii, and Pichia pastoris), phosphate groups present on extracellular glycans provide the negative cell surface charge (7–9). A notable exception is sialic acid {alpha}2,6- and {alpha}2,3-linked to penultimate {beta}-galactopyranosyl residues in glycans from two isolates of C. albicans (10, 11). In contrast to the charged groups summarized above, Schizosaccharomyces pombe N-linked polysaccharides contain pyruvic acid 4,6-ketallinked to Gal{beta}1,3-(PvGal)1 (12, 13). Pyruvate has only rarely been found in eukaryotic polysaccharides and glycoprotein oligosaccharides. Available examples include several species of green and red algae (14, 15), the nervous system of the sea slug Aplysia kurodai (16), and the marine sponge Microciona prolifera, in which PvGal is required for cellular recognition and adhesion (17). It is interesting to note the close structural similarity between the 4,6-ketal-linked pyruvate of the PvGal epitope and {alpha}2,6-linked sialic acid common in glycoproteins of higher eukaryotes (12, 13, 18).

Given that bacterial pyruvyltransferases have been identified in Xanthomonas sp. (19, 20), Rhizobium sp. (21, 22), and Bacillus anthracis (23), among other species, it was of interest to determine whether S. pombe has a genome-encoded functional analog of these proteins. However, initial data base searches could not identify any sequence homology between known bacterial pyruvyltransferases and any putative S. pombe proteins. Therefore, S. pombe strains devoid of negatively charged N-glycans were isolated following mutagenesis, and genetic complementation with an S. pombe genomic library was utilized to identify genes involved in the PvGal biosynthetic pathway.

Here we demonstrate that at least five different genes are required to synthesize the PvGal epitope. Although the inter-relationships between the encoded proteins remain to be completely established through future biochemical analyses, the results obtained suggest that PvGal{beta}1,3-synthesis is carried out by a coordinated enzymatic system in which the Gal{beta}1,3-residues are added to the S. pombe galactomannans and are then subsequently pyruvylated.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
General—Mallinckrodt Baker (Paris, KY) supplied 37% formaldehyde solution, 2-propanol, and anhydrous methanol. Pharmco (Brookfield, CT) provided anhydrous ethanol. Unless otherwise noted below, all other simple reagents were obtained from Sigma. Solvents, buffer components, and chemicals were ACS reagent grade or better. Sigma also supplied the antibiotics G418, kanamycin, and ampicillin. New England Biolabs (Beverly, MA) was the source for DNA restriction enzymes and T4 DNA ligase. All solutions were made using minimum 18-megaohm resistivity DDW produced by a Milli-QUF Plus Ultrapure water purification system (Millipore Corp., Bedford, MA) from distilled feedwater. DDW, buffers, and media solutions were sterilized by auto-claving. Standard molecular biology methods for bacteria and budding yeast were utilized (24, 25). Additional methods for fission yeast were also consulted (26, 27), including the Nurse Fission Yeast Handbook (available on the World Wide Web at www.sanger.ac.uk/PostGenomics/S_pombe/docs/nurse_lab_manual.pdf).

Cell Lines and Cell Growth—Fission yeast "wild-type" S. pombe strains (h– ura4–294) (ATCC 38436) and (h+ ura4+ leu1–32 ade6–704) (ATCC 96081) and S. cerevisiae strain (MAT{alpha} ura3–52 his3-{Delta}200 trp1-{Delta}901 lys2–801 suc2-{Delta}9 leu2–3,112) (SEY6210) (a kind gift from Dr. Scott Emr via Dr. Neta Dean) were used in this study. Cell growth media and growth conditions are described in the Supplemental Material.

Mutagenesis and Isolation of Mutants—"Wild-type" S. pombe (h– ura4–294) was mutagenized using ethyl methanesulfonate (Sigma). Following mutagenesis, cells were grown in YEDS liquid medium for 48 h (10–15 cell divisions) to allow phenotypic expression of the mutations. Mutant enrichment was achieved by repeated Q-Sepharose exclusion in DDW. Q-Sepharose Fast Flow (Amersham Biosciences) beads were prepared as described (12) and sterilized by suspension in 95% ethanol. Ethanol was removed by five repeated washes with DDW. Cells were collected by centrifugation in a 15-ml polypropylene centrifuge tube, washed three times with DDW, and resuspended in 10 ml of DDW. Approximately 1 ml of sterile Q-Sepharose beads (i.e. 2 ml of a 50% slurry) was added to the cell suspension, and the mixture was allowed to settle at room temperature for 5 min. The negatively charged cells bound to the beads and formed a fluffy precipitate that quickly settled to the bottom of the tube (Fig. 1A, wt). The supernatant containing unprecipitated cells was removed, and the procedure was repeated several times, until the cells remaining in the supernatant failed to bind and appeared to be uncharged (Fig. 1A, pvg–). The final cell suspension was grown overnight in YEDS. After dilution to ~100 cells/ml, 200-µl aliquots were spread on YEDS plates.



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  		FIG. 1.
PvGal epitope expression assays. A, Q-Sepharose binding assay of S. pombe cells. Negatively charged pvg+ wild-type cells adhere tenaciously to positively charged beads, whereas uncharged pvg– mutant cells remain indifferent to them. B, SAP staining assay of fixed cells. SAP staining of wild-type cells is diagnostic for the presence of pyruvylated terminal Gal{beta}1,3-. wt, wild-type positive control; pvg1––pvg5–, pvg– mutant strains; sc, S. cerevisiae negative control.

 
SAP Staining of Fixed Cells—Biotinylation of SAP (Sigma) and purification of the product were performed as described (28), with minor modifications. Briefly, the commercial SAP solution (containing 1 mg of SAP) was added to a pre-equilibrated Centricon YM-10 Mr 10,000 cut-off centrifugal filter (Amicon, Beverly, MA), washed twice by centrifugation with 1-ml additions of TBSC (10 mM Tris-Cl, 140 mM NaCl, 2 mM CaCl2, pH 8.0) to remove EDTA and concentrated to ~500 µl. Next, a fresh 11-mM solution of the BASHSE (Sigma) biotinylation reagent was produced by dissolving 0.3 mg of BASHSE in 50 µl of TBSC. A 0.1-volume aliquot of the BASHSE solution was added to the SAP solution in a microcentrifuge tube, yielding a ~550-µl reaction mixture (~1 mM BASHSE, ~2 mg/ml SAP). The biotinylation reaction was incubated for 18 h at room temperature. The mixture was washed twice by centrifugation in the filter with 1-ml additions of TBSC (with 0.1% NaN3 as a preservative). The biotinylated SAP product, concentrated to ~200 µl (~5.0 mg/ml) and stored at 4 °C, was used without further characterization.

Colonies chosen for screening were grown overnight in 1-ml YEDS aliquots. Cells were collected by centrifugation, washed twice with TBS (10 mM Tris-Cl, 140 mM NaCl, pH 8.0), and fixed with 400 µl of 10% formalin (i.e. 1:10 dilution of commercial formaldehyde solution in TBS) for 60 min at room temperature. Fixed cells were washed twice with TBSC. For each assay, a 0.2-µl aliquot of the 5 mg/ml biotinylated-SAP solution was diluted with 49.8 µl of TBSC. The cells were resuspended in 50 µl of this solution (containing 1 µg of biotinylated SAP) and incubated at room temperature for 30 min. Cells were then washed twice with TBSC. For each assay, a 0.5-µl aliquot of 100 units/ml alkaline phosphatase-avidin D (Vector Laboratories, Burlingame, CA) was diluted with 49.5 µl of TBSC. The cells were resuspended in 50 µl of this solution (containing 50 milliunits of alkaline phosphatase-avidin D) and incubated at room temperature for 30 min. Cells were washed three times with TBSC, and aliquots of each preparation containing approximately equal numbers of cells were transferred (spotted) onto Whatman 3MM chromatography paper (Whatman, Maidstone, Kent, UK) and air-dried. Color development after the addition of Western Blue alkaline phosphatase substrate (Promega, Madison, WI) indicated SAP binding (Fig. 1B).

Lectin Blotting—Cells were grown overnight in 1-ml YEDS aliquots, collected by centrifugation, washed with DDW, and resuspended in 400 µl of lectin blotting buffer (50 mM sodium citrate, 100 mM 2-mercaptoethanol, 1% SDS, pH 5.5). Suspensions were heated in a 100 °C water bath for 5 min and pelleted by centrifugation. Soluble glycoproteins were precipitated from the decanted supernatant by the addition of ice-cold acetone to 80% (29). Glycoproteins were resuspended in 200 µl of lectin blotting buffer/mg of protein, denatured by boiling for an additional 5 min, and loaded at 600 ng/lane onto 8% SDS-PAGE gels. Sample preparation; SDS-PAGE; electroblotting to Immobilon-P transfer membranes (Millipore); and lectin blotting with concanavalin A (for {alpha}-linked Man), GSL/BSL I-B4 (for {alpha}-linked Gal), and PNA (for {beta}-linked Gal) were performed according to standard protocols from the suppliers. Bio-Rad was the source for PAGE supplies. Vector Laboratories supplied biotinylated concanavalin A lectin and biotinylated GSL/BSL I-B4 isolectin. Biotinylated PNA lectin was obtained from Roche Applied Science.

Compositional Analysis—Acid hydrolysis of the N-linked polysaccharides was performed as described (12, 28). Monosaccharides were separated by HPAEC on a Dionex (Sunnyvale, CA) CarboPac PA1 4 x 250-mm analytical column isocratically with 16 mM NaOH and were monitored by PAD using a Dionex ED50 Electrochemical Detector.

Depyruvylation and Exoglycosidase Digestions—Wild-type galactomannans were depyruvylated by mild acid hydrolysis as described (12, 28). Earlier work showed that Gal{beta}1,3-in the wild-type S. pombe N-linked galactomannans represented about 10% of the total Gal (28). To confirm the presence of unsubstituted Gal{beta}1,3-residues in the pvg1– mutant, we incubated ~3-µmol Man equivalents (~50 nmol of total oligosaccharide) of wild-type and mutant N-linked glycans overnight with 20 milliunits of green coffee bean {alpha}-galactosidase (Sigma) in 40 µl of digestion buffer (50 mM sodium citrate, 5 mM CaCl2, pH 6.0) or with 300 milliunits of Xanthomonas manihotis {beta}-galactosidase (expressed in Escherichia coli) (New England Biolabs) in 40 µl of digestion buffer (50 mM sodium citrate, pH 4.5). Following precipitation with ice-cold 60% methanol (29), supernatants were evaporated under vacuum, and Gal release was measured by HPAEC-PAD. To ensure that Gal released by {beta}-galactosidase would be observable by HPAEC-PAD, we optimized the amounts of total injected N-glycans so that release of only 10% of Gal by {alpha}-galactosidase (~100 nmol) yielded a prominent peak.

Oligosaccharide Preparation and NMR Spectroscopy—S. pombe cells were grown to early stationary phase in YEDS. Following disruption by vortexing with 0.5-mm acid-washed glass beads, cell debris was removed by centrifugation. Nonglycosylated proteins were precipitated by 60% saturation with solid ammonium sulfate. After centrifugation, the glycoprotein-containing supernatant fraction was equilibrated by dialysis with deglycosylation buffer (50 mM sodium citrate, pH 5.5), and oligosaccharides were released from the glycoproteins with endo-{beta}-N-acetylglucosaminidase H (30) (expressed in E. coli). Deglycosylated proteins were removed by solvent precipitation (28, 29). The released N-glycans were sized on an extra fine mesh Bio-Gel P-4 (Bio-Rad) column, and the large galactomannans eluting at the void volume were examined further. Samples were prepared, and one- and two-dimensional NMR spectra were recorded, processed, assigned, and analyzed as described (28, 31).

Genetic Methods and Plasmid Construction—An S. pombe genomic library (32) (a kind gift from Dr. Tony Carr) was used for complementation of mutation and restoration of wild-type phenotype. Library plasmids recovered from pvg– mutants with restored wild-type phenotypes were sequenced using appropriate primers (Table SI, Supplemental Material) to determine their S. pombe genetic content. S. pombe pvg genes were identified through Washington University BLAST (33, 34) comparison of these sequences against the annotated S. pombe GeneDB gene data base (35, 36) (available on the World Wide Web at www.genedb.org/genedb/pombe/index.jsp). All plasmids in this study were derived from the low copy S. pombe shuttle vector pUR19 (ars1+, ura4+) (ATCC 77298) (32) or the high copy S. pombe shuttle vector pSP2 (ars1+, S. cerevisiae URA3) (ATCC 77498) (37). Details of cloning and expression plasmid construction are available in the Supplemental Material. The expression plasmids pUR-(1–5) GFPc and pSP-(1–5) GFPc were transformed into the respective pvg1––pvg5– mutants and tested for complementation of mutation and restoration of wild-type phenotype as described above. The cellular location of the expressed Pvg(1–5) p-GFP fusion proteins was determined by fluorescence microscopy.

A G418-resistant h+ strain was designed by random integration of an EcoRI–AscI kanMX6-GFP cassette into the S. pombe (h+ ura4+ leu1–32 ade6–704) strain. Briefly, the pFA6a-kanMX6-P3nmt1-GFP plasmid (38) (a kind gift from Dr. Jürg Bähler) was digested with EcoRI and AscI, and a 3.5-kbp fragment containing the kanMX6 marker (39), together with the (S65T) GFP gene (40) under transcriptional control of the nmt1 promoter (41), was isolated using the QIAquick Gel Extraction Kit (Qiagen, Valencia, CA). The cassette was transformed into the (h+ ura4+ leu1–32 ade6–704) strain and replica-plated on YEDS/G418 plates. G418-resistant, GFP-expressing h+ colonies were used for mutagenesis and subsequent mating analysis.

Transcription Analysis by End Point RT-PCR—End point RT-PCR was performed to determine whether transcription of pvg1–pvg5 was significantly affected in any of the mutants and whether Pvg4p, a putative transcription factor (see "Results"), was involved in regulation of one or more of pvg1–pvg3 and pvg5. Briefly, nucleic acids were isolated from wild-type and pvg– mutant cells grown to early stationary phase in YEDS. Following DNase I (Invitrogen) digestion to remove genomic DNA, cDNA was synthesized from mRNA transcripts using the SuperScript First-Strand Synthesis System for RT-PCR (Invitrogen), utilizing SuperScript II RT and oligo(dT)12–18 primer. PCR was performed with Platinum TaqDNA polymerase (Carlsbad, CA) and appropriate gene-specific primers for pvg1–pvg5 and actin (act1) control (Table I). Difficult amplification of pvg3 due to secondary structure effects necessitated the use of the forward primer pvg3-TMD, which did not incorporate the transmembrane domain coding sequence. Both positive controls (no DNase I) containing genomic DNA and negative controls (no SuperScript II RT) were utilized.


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  		TABLE I
Oligonucleotide primers for RT-PCR

 
Mating Analysis—Mating analysis was performed by crossing selected pvg– mutants of the (h– ura4–294) strain with the G418-resistant (h+ ura4+ leu1–32 ade6–704) strain on malt extract medium plates. Inversely, several pvg– mutant strains were derived from the G418-resistant (h+ ura4+ leu1–32 ade6–704) strain as described above for (h– ura4–294) and were crossed with the wild-type (h– ura4–294) strain. Following 3 days of incubation at room temperature, plates were inspected by light microscopy for the formation of crescent-shaped zygotic asci. Cell mixtures from each cross were then replated on CSM-Leu/G418 medium. Plates were examined for growth after 2 days of incubation at 33 °C. Wild-type (pvg+) cells of both mating types were used as controls.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Isolation and Characterization of Mutants Defective in Synthesis of Negatively Charged N-Linked Polysaccharides—Since no phosphate or sulfate has previously been detected in S. pombe N-linked glycans (12, 13), pyruvate appears to be the only negatively charged functional group on the cell surface. Based on this assumption, we isolated glycosylation mutants using the inability to bind to Q-Sepharose beads as the primary screen (Fig. 1A; c.f. wt and pvg–). Those mutants with highly underglycosylated or undergalactosylated galactomannans, which are known to be highly osmosensitive (42, 43), were effectively excluded from further examination by repeated DDW washes. We applied biotinylated SAP staining of fixed cells to further screen mutant colonies for the absence of pyruvylated Gal (Fig. 1B). By a fortuitous coincidence, serum amyloid P component exhibits calcium-dependent lectin-like affinity for the PvGal epitope, an apparently nonphysiological ligand (44, 45). About 70 colonies, unable to bind to Q-Sepharose beads and unreactive with SAP, were obtained from five separate ethyl methanesulfonate mutagenesis trials. We performed lectin blot analyses (not shown), with biotinylated concanavalin A and biotinylated GSL/BSL I-B4, to roughly determine the composition of the glycans present on the glycoproteins of the SAP-negative strains. The mutant strains used for further investigation had overall levels of mannosylation and {alpha}-galactosylation similar to the wild-type strain, suggesting that the mutations only affected the relatively infrequent PvGal modification found in S. pombe N-linked glycans.

Transformation of these strains with an S. pombe genomic library (32) resulted in classification into five different groups, each complementable by one of five plasmids. Partial sequencing of their respective genomic inserts revealed that all five plasmids contained open reading frames predicted to encode proteins of unknown functions (Table II). When cloned into pUR-KAN (Fig. S1, Supplemental Material) for expression, each coding sequence, with its upstream promoter region, allowed full complementation of only its corresponding complementation group and restored expression of pyruvylated N-linked glycans, as determined by Q-Sepharose binding and SAP staining of fixed cells. We designated the five genes pvg1–pvg5 and registered these names with the S. pombe GeneDB gene data base (35, 36) (available on the World Wide Web at www.genedb.org/genedb/pombe/index.jsp).


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  		TABLE II
Identity of pvg1–pvg5 genes, putative functions, and bioinformatic results

 
Whereas ~98% of the large wild-type galactomannans are negatively charged (12, 13), the glycans from the five pvg– strains were totally incapable of binding Q-Sepharose beads. This suggested a complete, or nearly complete, lack of negatively charged functionalities on these glycans. Compositional analysis, by acid hydrolysis and HPAEC-PAD, of the large N-linked glycans from representatives of each of the five complementation groups showed that all had monosaccharide compositions similar to those of wild-type cells (not shown). This corroborated lectin blot results and confirmed that neither the disruption of the pvg gene products nor artifacts from the random mutagenesis dramatically impacted the metabolic pathways responsible for the bulk of N-linked glycosylation in these mutants and that the lack of glycan negative charge did not result from a complete Gal deficiency.

Wild-type S. pombe galactomannans are very large; the "average" composition of Endo H-released galactomannans is Pv5–6Gal52–57Man62–64GlcNAc (13, 28). Approximately 80% of the Man residues of the outer chain poly-Man{alpha}1,6-backbone are occupied by Gal{alpha}1,2-side chains, of which about 10% are capped by PvGal{beta}1,3- (Fig. 2) (13, 28). Unsubstituted {beta}1,3-linked Gal is not found in the wild type. Therefore, about 8% of outer chain extension Man residues in toto are substituted by PvGal{beta}1,3Gal{alpha}1,2-, and all pyruvate in S. pombe galactomannans is associated with this appendage.



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  		FIG. 2.
Composite structure of the S. pombe galactomannan outer chain extension. Schematic of pvg+ wild-type outer chain extension showing residue nomenclature (blue) and pvg– mutant terminations (red). 6M, 6-monosubstituted Man{alpha}1,6; 2,6M, 2,6-disubstituted Man{alpha}1,6; tG, terminal Gal{alpha}1,2; 3G, 3-substituted Gal{alpha}1,2; {beta}G, Gal{beta}1,3.

 
Lectin blot analyses (not shown) of purified glycoproteins from the wild-type and five mutant strains, using concanavalin A and GSL/BSL I-B4, indicated almost equivalent glycoprotein glycan profiles for all. PNA, which has a 10-fold higher affinity for {beta}-linked Gal than for {alpha}-linked Gal (51), showed a significantly greater staining of pvg1– glycoproteins than of wild-type or pvg2––pvg5– glycoproteins, indicative of the presence of terminal Gal{beta}1,3- (Fig. 3). However, incubation of pvg1– glycoproteins with {beta}-galactosidase did not decrease PNA staining (not shown). To confirm this unexpected result, we prepared depyruvylated Gal{beta}1,3-containing glycans by exposing wild-type galactomannans to mild acid hydrolysis. Digestion of these glycans with {beta}-galactosidase yielded no appreciable Gal release, as determined by HPAEC-PAD monosaccharide analysis (not shown) and NMR oligosaccharide analysis (not shown). These results, taken together with those of PNA lectin blotting, imply that the Gal{beta}1,3-residues are inaccessible to this enzyme.



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  		FIG. 3.
PNA lectin blot analysis of purified glycoproteins from pvg+ wild-type and pvg– mutant strains. Significant increases in staining of pvg1– glycoproteins (arrows) are indicative of the absence of Pv and consequent exposure of terminal Gal{beta}1,3-. wt, wild type; pvg1––pvg5–, pvg– mutant strains.

 
Since the pvg1––pvg5– mutant galactomannans were observed to be roughly as abundant in Gal as wild-type galactomannans, the absence of the negative charge could be predicted to arise from either of two mutant structural phenotypes, one missing the terminal pyruvyl group but retaining the terminal Gal and the other devoid of the entire (relatively infrequent) PvGal{beta}1,3-cap. NMR analysis of wild-type and pvg1––pvg5– galactomannans confirmed the presence of both structural expectations (Fig. 4). Wild-type and pvg1– glycans exhibited the 4.67-ppm anomeric proton signal of Gal{beta}1,3-({beta}G) (shifted upfield to 4.64 ppm in pvg1–) and the 4.30-ppm C4-H resonance of 3-substituted Gal{alpha}1,2-(3G) (shifted upfield to 4.25 ppm in pvg1–). However, the signal for pyruvate protons (Pv) at 1.54 ppm was absent in the pvg1– glycans. The glycans from pvg2––pvg5– mutants displayed neither of these particular proton signals and are thus devoid of the entire PvGal{beta}1,3-cap.



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  		FIG. 4.
NMR analysis of pvg+ wild-type and pvg– mutant galactomannans. The purified N-linked galactomannans from wild-type S. pombe and from each of the pvg– strains were examined by one-dimensional 1H NMR spectroscopy. A, diagnostic proton signals from C1-H of 2,6-disubstituted Man{alpha}1,6- (2,6M) at 5.21 ppm, C1-H of terminal Gal{alpha}1,2- (tG) at 5.09 ppm, C1-H of 6-monosubstituted Man{alpha}1,6- (6M) at 4.88 ppm, C1-H of Gal{beta}1,3- ({beta}G) at 4.67 ppm (shifted upfield to 4.64 ppm in the absence of pyruvate deshielding), C4-H of 3-substituted Gal{alpha}1,2- (3G) at 4.30 ppm (shifted upfield to 4.25 ppm in the absence of pyruvate deshielding), and Pv at 1.54 ppm (36). Only wild-type galactomannans contain Pv. Only wild-type and pvg1–-galactomannans exhibit the 3G and {beta}G signals. See Fig. 2 for a schematic of galactomannan outer chain extension. B, detail of the 3G section of a pvg1–-galactomannan two-dimensional DQF-COSY-NMR spectrum, confirming connectivity between C4-H and C2-H of 3G at 4.30 ppm and 4.03 ppm, respectively. C, detail of the {beta}G section of a pvg1–-galactomannan two-dimensional COSY spectrum, confirming connectivity between C1-H and C2-H of {beta}G at 4.67 and 3.67 ppm, respectively.

 
All five pvg sequences were determined from the S. pombe gene data base to specify relatively small deduced proteins (Table II). All five genes lack introns, a somewhat surprising result, considering that 43% of S. pombe genes contain introns, and such genes have an average of ~2.2 introns/gene (35, 36). By comparison, only 5% of S. cerevisiae genes contain introns, and such genes have an average of ~1.1 introns/gene (35, 36). A collection of bioinformatic algorithms were queried with the deduced protein sequences of the five pvg genes. The predicted (unmodified) molecular weights and isoelectric points, cellular locations, transmembrane domains and topology, and functions of each of the encoded proteins are presented with their genome data base annotations (Table II). With the exception of Pvg4p, the proteins are predicted to have Type II-oriented single-pass transmembrane domains. Such proteins have a cytoplasmic N terminus and a C terminus within the lumen of a secretory compartment or within the periplasm for a yeast plasma membrane protein. Several neural net glycosylation prediction algorithms were also canvassed (Table III), although they were not trained specifically with yeast proteins.


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  		TABLE III
Predicted glycosylation of Pvg1–Pvg5 proteins

 
To determine subcellular localization of Pvg1p–Pvg5p, we expressed C-terminal GFP fusions of each protein in the respective pvg1––pvg5– mutants as described above and in the Supplemental Material. The transformants exhibited growth rates similar to that of wild-type cells. Each of the Pvg(1–5)p-GFP fusion proteins complemented its corresponding lesion, as determined by Q-Sepharose binding and SAP staining of fixed cells. Images showing the localization of some of these proteins were obtained (Fig. 5) and are discussed below. However, low expression levels from the pUR-based plasmids or possibly post-translational processing events (e.g. misfolding of GFP) prevented sufficient fluorescence for localization of Pvg2p-GFP or Pvg5p-GFP.



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  		FIG. 5.
Expression and localization of Pvg1p–Pvg4p C-terminal GFP-fusion proteins. Pvg1p-GFP exhibits a pericellular location; Pvg2p-GFP has a polar location; Pvg3p-GFP has a cytoplasmic location; and Pvg4p-GFP has a nuclear location. Expression was in pUR19, except for Pvg2p-GFP, which was in pSP2. The Pvg5p-GFP fusion expressed from both pUR19 and pSP2 successfully complemented the pvg5– mutation but failed to exhibit fluorescence.

 
To examine the effect of high copy expression of the C-terminal GFP-tagged proteins, we constructed pSP2-based expression plasmids as detailed in the Supplemental Material. The pSP-(1–3,5)GFPc transformants grew about half as fast as did their pUR-(1–3,5)GFPc counterparts. The pSP-4GFPc transformants were extremely slow growing and appeared to experience difficulties with cell division, giving rise to elongated cells. Three of the Type II transmembrane constructs (Pvg(1,2,5)p-GFP) successfully complemented their corresponding lesions. The Pvg3p-GFP fusion construct, specifying a probable {beta}1,3-galactosyltransferase, as well as the extremely slow growing pSP-4GFPc transformant, failed to restore charge. With the exception of pSP-2GFPc, we were unable to obtain detectable GFP signal from the pSP-GFPc transformants. Thus, overexpression of Pvg-GFP fusion proteins from pSP2-based plasmids led in most cases to cellular disruption and misfolding of the Pvg and/or GFP portion.

Characterization of pvg1– Mutant—A sequence similarity search by standard protein-protein BLAST (33, 55) showed that the predicted amino acid sequence of Pvg1p shares an ~300–350-aa stretch of about 30% identity and about 45% similarity with nearly 20 identified or predicted pyruvyltransferases. These include the PssK "exopolysaccharide polymerization" proteins of Rhizobium leguminosarum bv. trifolii and R. leguminosarum bv. viciae (56) and the Streptococcus thermophilus EpsL protein (57), the latter including a shorter 118-aa portion with 39% identity and 56% similarity. Among the reported sequences were a number of "polysaccharide polymerization" and "exopolysaccharide biosynthesis" proteins, including those from Nostoc sp., Leuconostoc mesenteroides, Oenococcus oeni, Burkholderia fungorum, Actinobacillus pleuropneumoniae, and Bacillus subtilis. Reverse position-specific (RPS) BLAST (58) confirmed the presence of a ~300-aa conserved pyruvyltransferase (pfam4230) domain (Pfam protein family data base) (50). Of great interest and with the highest scoring standard BLAST sequence homology to Pvg1p (including a shorter 128-aa stretch with 43% identity and 57% similarity) is a "possible galactosyltransferase" from Cryptosporidium parvum. This relatively large (827-aa) predicted protein appears to be bifunctional, with an N-terminal galactosyltransferase (pfam01762) domain similar to that of Pvg3p and the identified C-terminal pyruvyltransferase domain.

Both one- and two-dimensional NMR analysis show the presence of unsubstituted Gal{beta}1,3-residues in N-linked galactomannans of the pvg1– mutant (Fig. 4), indicating that 1) PvGal epitope synthesis occurs by subsequent transfer of the pyruvyl group to Gal{beta}1,3-residues already present on the N-linked galactomannan chains and does not require the formation of some PvGal-containing precursor, and 2) Pvg1p is the only protein among the five discovered that is required for pyruvate addition and is putatively a pyruvyltransferase. A functionally active C-terminal GFP fusion of Pvg1p appears to localize to the cell periphery, where it may act just prior to glycoprotein secretion and incorporation into the cell wall (Fig. 5A).

The S. pombe GeneDB annotation for pvg1 indicates that there is no S. cerevisiae analog (Table II), as would be expected from the absence of pyruvate in S. cerevisiae glycoprotein mannans. Other bioinformatic results support both a cell membrane location and a pyruvyltransferase function for Pvg1p (Table II). A salient bioinformatic prediction is a cluster of O-linked mucin-type glycans immediately adjacent to the transmembrane domain (Table III). Such clusters are purported to impart rigidity to the polypeptide backbone, thereby extending terminal domains from the membrane (59).

Characterization of pvg2– Mutant—Standard protein-protein BLAST revealed that the deduced sequence of Pvg2p possesses a 159-aa middle section having 26% identity and 42% similarity with another putative S. pombe protein (from SPAC2F3.01), an S. cerevisiae Sur1p homolog. Budding yeast Sur1p transfers Man to inositolphosphorylceramide (60) and possesses three transmembrane domains, one near the N terminus and two close to one another and near the C terminus. Whereas a definite Sur1p homolog has not been identified in fission yeast, S. pombe is predicted to have two other highly homologous proteins with significant similarity to Sur1p, and budding yeast has one other homologous protein. However, unlike Sur1p, Pvg2p is not predicted to have C-terminal transmembrane domains (Table II).

Three iterations of position-specific iterated (PSI) BLAST (55) and a search for short, nearly exact matches revealed lesser similarities with a few glycosyltransferases, including yeast Och1p. Och1p is the {alpha}1,6-mannosyltransferase, with homologs in both S. cerevisiae and S. pombe, responsible for the first step of outer chain elongation of N-linked oligosaccharides (61). An N-terminal transmembrane domain is predicted for budding yeast Och1p but not for the fission yeast homolog. RPS BLAST also revealed the presence of a conserved Och1p-related domain, which encompasses Och1- and Sur1-like proteins, as well as S. cerevisiae Hoc1p (homologous to Och1p) (62). Hoc1p is a component of one of two mannan polymerase complexes involved in outer chain {alpha}1,6-Man elongation (63, 64).

Sur1p, Och1p, and Hoc1p all possess the tripeptide DXD sequence that coordinates an Mn2+ ion, which plays a central role in catalytic utilization of a nucleotide saccharide donor in most, but not all, glycosyltransferases (65–67). However, when aligned against Sur1- and Och1-like proteins, Pvg2p shows a noticeable absence of the expected DXD motif, having SRS instead (aa 187–189). Indeed, of the five predicted Pvg proteins that we identified, only Pvg3p exhibits the DXD sequence.

The S. pombe GeneDB annotation for pvg2 indicates that it is possibly S. pombe-specific, with no apparent orthologs (Table II). The role of Pvg2p remains unknown, although NMR results show that it is essential for Gal{beta}1,3-addition (Fig. 4). A C-terminal GFP fusion was localized primarily to the cell poles, consistent with deposition of newly synthesized galactomannans in growing cells (Fig. 5B). Paradoxically, other bioinformatic results predict localization of Pvg2p to the membrane of a secretory compartment (i.e. Golgi and endoplasmic reticulum), in apparent contradiction to empirical results (Table II). Pvg2p is also ascribed a ligase function of unknown specificity.

Characterization of pvg3– Mutant—Although standard protein-protein BLAST of the predicted amino acid sequence for Pvg3p indicated some homology to known and putative glycosyltransferases as well as to other proteins, it was indecisive. Three iterations of PSI BLAST clarified a strong homology to several hundred {beta}1,3-glycosyltransferases, particularly with many glycoprotein (UDP-Gal: {beta}-GlcNAc) {beta}1,3-galactosyltransferases from humans, Old World hominoids, and a variety of mammals and higher eukaryotes. RPS BLAST confirmed that pvg3 encodes a protein with a conserved galactosyltransferase (pfam01762) domain of the glycosyltransferase family 31 (GT31) in the Carbohydrate Active Enzyme data base (66). The GT31 family of over 200 structurally related inverting {beta}1,3-glycosyltransferases, which contain a "GT-A" or "SpsA" Rossmann-like fold (66–68), is predominantly composed of (UDP-Gal: {beta}-GlcNAc) {beta}1,3-galactosyltransferases from species ranging from yeasts to plants to humans (69, 70). As mentioned earlier, of special interest is the presence in the GT31 family of a putative bifunctional enzyme from C. parvum, with an N-terminal galactosyltransferase domain similar to that of Pvg3p, and a C-terminal pyruvyltransferase (pfam04230) domain similar to that of Pvg1p described above.

Pvg3p is the only member of the GT31 family predicted to be present in S. pombe. This corresponds with the finding that galactomannans purified from pvg3– cells lack both pyruvate and {beta}1,3-linked Gal (Fig. 4), since the PvGal epitope contains the only {beta}-linked Gal. The expressed Pvg3p-GFP fusion showed punctate labeling typical for localization within organelles of the cytoplasmic compartment (Fig. 5C), a pattern also seen for S. pombe Golgi-resident Gma12p, an {alpha}1,2-galactosyltransferase (71). These data, along with the homology to known {beta}1,3-galactosyltransferases, strongly imply that Pvg3p is the {beta}1,3-galactosyltransferase that adds Gal to the outer chain Gal{alpha}1,2-residues of galactomannan.

Other bioinformatic results support both a secretory compartment location and a galactosyltransferase function for Pvg3p (Table II).

Characterization of pvg4– Mutant—Standard protein-protein BLAST of the deduced Pvg4p amino acid sequence indicated strong homology to over 100 MADS-box transcription factors, particularly homeotic plant proteins. The MADS-box domain is an ancient and conserved contiguous sequence of ~56 amino acids found, generally N-terminally, in transcriptional regulation proteins in species ranging from yeasts to plants to humans (72). RPS BLAST confirmed the presence in Pvg4p of a MADS-box (pfam00319) domain.

MADS-box transcription factors bind to A/T-rich cis-acting regulatory elements, which are classified into two general lineages (73–75). Members of the serum response factor (SRF)-like Type I subtype bind to the canonical "CArG" sequence 5'-CC(A/T)6GG-3', whereas those of the myocyte-specific enhancer factor 2 (MEF2)-like Type II subtype bind to the DNA sequence 5'-YTA(T/A)4TAR-3'. Three iterations of PSI BLAST returned Pvg4p homology to over 1000 MADS-box proteins in the MEF2-like Type II subfamily. A search for short, nearly exact matches confirmed this as the strongest affiliation, particularly with the four human MEF2A–D (76) and Drosophila melanogaster D-MEF2 (77). The predicted protein sequence of the Pvg4p MADS-box domain (aa 2–57) shows 55–60% identity and 73–75% similarity with these five members. Indeed, aa 2–45 of these six sequences are 68–70% identical and 79–83% similar (Fig. 6).



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  		FIG. 6.
Multiple sequence alignment of the 56-aa MADS-box domains of human myocyte-specific enhancer factors 2A–D, D. melanogaster D-MEF2, and S. pombe Pvg4p. The alignment, generated by ClustalW 1.8 (78), shows a strong homology in {alpha}-helix I and the first portion of {beta}-strand I, implying that Pvg4p binds to a MEF2-like Type II MADS-box-binding sequence. Weak homology in the remainder of {beta}-strands I and II indicates that Pvg4p has a distinctive dimerization area. On the C-terminal side of the MADS-box (not shown), {alpha}-helix II of Pvg4p may bind distinct accessory factors.

 
MADS-box transcription factors homodimerize and can in some cases form heterodimers with other MADS-box transcription factors. These dimers associate with DNA target sequences and accessory protein factors to assemble into multicomponent regulatory complexes. X-ray crystal structures for complexes of the MADS-box domain proteins SRF, Mcm1p, and MEF2A indicate a helix-strand-turn-strand-helix structure for the regulatory portion of each transcription factor (79–81). After an N-terminal sequence, the first {alpha}-helix is the major determinant of DNA-binding specificity, whereas the {beta}-strands are necessary for dimerization. Beyond the MADS-box, the second {alpha}-helix is also involved in dimerization and is critical for the specific recruitment of other factors (72–75, 79–81). Alignment of the 56-aa MADS-box portions of Pvg4p and five MEF2 MADS-box transcription factors indicates that Pvg4p may bind to a MEF2-like Type II promoter sequence while having distinctive dimerization and accessory factor binding areas (Fig. 6). A search of the 300-bp regions immediately upstream of the START codons for each of the pvg genes, using the public noncommercial version of TRANSFAC 6.0 (data base on eukaryotic transcription factors, their genomic binding sites, and DNA binding profiles) and the program Patch 1.0 (pattern search for transcription factor binding sites) (available on the World Wide Web at www.gene-regulation.com), revealed that only two of them, pvg1 and pvg5, contain sequences with homology to known MEF2-like promoters. These were 5'-CTAAAT(T)TAG-3' for pvg1 and 5'-(G)TTAAATAG-3' for pvg5, centered 172 and 107 bp upstream from their respective START codons.

There are two other MADS-box transcription factor genes predicted to be in S. pombe, one (SPBC19G7.06) with no S. cerevisiae ortholog and of unknown function and the other the pheromone receptor Map1p, involved in mating-type determination (82, 83). The latter gene is a functional homolog of the S. cerevisiae transcription factor Mcm1p (79, 84) and is also homologous to the S. cerevisiae Arg80p (85). However, both Mcm1p and Arg80p belong to the SRF-like Type I MADS-box subfamily (75, 85). The other S. cerevisiae MADS-box transcription factors are encoded by SMP1 and RLM1, which belong to the MEF2-like Type II MADS-box subfamily (75). Rlm1p is activated by phosphorylation in the (Mpk1/Slt2) mitogen-activated protein kinase pathway (86, 87). Of particular relevance, most if not all of the target genes for Rlm1p are involved in cell wall biosynthesis and remodeling during growth and differentiation and as a result of cell integrity signaling induced by environmental stresses (88, 89). Similarly, Smp1p is phosphorylated in the stress-activated (Hog1) mitogen-activated protein kinase pathway (90). However, whereas the phosphorylation sites are similar for Rlm1p and Smp1p, and whereas the two proteins are known to heterodimerize, they do not appear to have overlapping functions (86, 88, 90).

Three PSI BLAST iterations of the non-MADS-box remainder of Pvg4p (i.e. aa 57–372) indicated very strong homology to dozens of DNA-directed RNA polymerase II (large subunit) proteins. A search of this portion for short, nearly exact matches revealed a midprotein sequence (aa 158–193) extremely rich in proline, histidine, and aromatic amino acids. Analysis of this sequence indicates several regions that might serve as ligand areas for proteins containing SH3 (Src homology 3) or WW (tryptophan-tryptophan) domains (91). Proteins containing these proline-rich motifs are involved or postulated as adaptors between membrane signal receptors and cytoplasmic or cytoskeletal factors, acting as mediators of intracellular signal transduction (91).

A functionally active Pvg4p-GFP fusion protein showed strong general nuclear localization with no apparent preferential labeling of nuclear substructures (Fig. 5D). One-dimensional NMR of N-linked glycans purified from pvg4– cells indicated that Pvg4p function is required for Gal{beta}1,3-addition (Fig. 4).

Other bioinformatic results confirm both a nuclear location and a transcription regulation function for Pvg4p (Table II). A striking prediction is the presence of nearly 20 O-linked {beta}-GlcNAc sites (Table III). Many of these are putative "yin-yang" sites for the dynamic reciprocity between O-{beta}-GlcNAc attachment and O-phosphorylation (92). Yin-yang sites have been shown to be involved as control nodes for cellular regulation in multicellular eukaryotes (93, 94) in, for example, the MADS-box transcription factor SRF (95). However, direct evidence of O-{beta}-GlcNAc-modified proteins in simple eukaryotes remains elusive (93, 94).

Characterization of pvg5– Mutant—Standard protein-protein BLAST of the predicted Pvg5p amino acid sequence provided no insight into the possible function of Pvg5p in PvGal synthesis. Neither three iterations of PSI BLAST nor a search for short, nearly exact matches contributed helpful information. However, an interesting observation is that a C-terminal 151-aa portion of Pvg5p shows 20% identity and 39% similarity with a C-terminal section of Pvg1p. Nevertheless, these proteins do not possess overlapping biosynthetic functions, since the pvg5 gene failed to complement the pvg1– mutant phenotype, and vice versa. Based on NMR data, Pvg5p is essential for Gal{beta}1,3-addition, whereas Pvg1p is required for terminal pyruvylation.

As in the case of pvg2, the S. pombe GeneDB annotation for pvg5 indicates no apparent orthologs for the possibly S. pombe-specific gene (Table II). Other bioinformatic results suggest a cytoplasmic or mitochondrial location for Pvg5p (Table II), although the former location is inconsistent with the predicted Type II transmembrane domain. A transferase or oxidoreductase function is also proposed.

Transcription Analysis by End Point RT-PCR—End point RT-PCR of transcripts of each of the five pvg genes (and act1 control) in each of the five pvg– mutant strains indicated that the translatable regions of each gene were transcribed at a level and size comparable with that of the wild-type strain (Fig. 7). Most notably, Pvg4p does not appear to solely control transcription of any of the other four genes (pvg1–pvg3 and pvg5). Therefore, despite the presence of potential MEF2-like Type II MADS-box-binding promoter sequences upstream from pvg1 and pvg5, expression of the transcripts appears unaffected. Negative controls (no RT) indicated no genomic DNA contamination of the mRNA samples (not shown).



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  		FIG. 7.
End point RT-PCR transcript expression analysis in pvg+ wild-type and pvg1––pvg5– mutant S. pombe. Nucleic acids were isolated from wild-type and mutant cells. Following DNase I digestion to destroy genomic DNA, RT of mRNA with oligo(dT) primer and PCR with gene-specific primers was performed. In each pvg– mutant, each of the five pvg genes (and act1 control) is transcribed untruncated, at wild-type level. wt, wild-type cells; pvg1––pvg5–, mutant cells; pvg1–pvg5, pvg transcripts; act1, actin control transcript. Appropriate positive controls (no DNase) and negative controls (no RT) were utilized (not shown). See Table I for expected product sizes.

 
Mating Analysis—It has been reported that S. pombe strains greatly deficient in surface Gal are incapable of sexual conjugation (43). To test whether the PvGal epitope plays any role in the sexual behavior of S. pombe cells, we spot-mixed pvg– mutant strains of both mating types ((h– ura4–294) and G418-resistant (h+ ura4+ leu1–32 ade6–704)), defective in synthesis of pyruvylated galactomannans, on malt extract medium plates. After 3 days of incubation at room temperature, each cross-pattern ((h– pvg+) x (h+ pvg+), (h– pvg–) x (h+ pvg+), (h– pvg+) x (h+ pvg–), (h– pvg–) x (h+ pvg–)) showed an almost equal number of crescent-shaped zygotic asci. Cell mixtures from each cross were replated on CSM-Leu/G418 medium. After 2 days of incubation at 33 °C, each cross-pattern showed an almost equal number of colonies (Fig. 8). Microscopic examination indicated these colonies to be prototrophic haploid crossover recombinants; diploid cells were not observed. No colonies were observed in the control (uncrossed) quadrants of the plates. Therefore, neither the terminal pyruvate (i.e. pvg1– mutant) nor the complete PvGal epitope (i.e. pvg2––pvg5– mutants) of cell wall N-glycans appears to play any significant role in growth or mating (i.e. conjugation, meiosis, and sporulation). Interestingly, however, pvg1– cells with exposed Gal{beta}1,3-residues showed an increased propensity to agglutinate at late stationary phase of growth in liquid medium.



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  		FIG. 8.
Mating analysis on CSM-Leu/G418 plates. Mutant pvg– strains of both h– and h+ mating types ((h– ura4–294) and G418-resistant (h+ ura4+ leu1–32 ade6–704)), defective in synthesis of pyruvylated galactomannans, were mixed on malt extract medium plates and incubated at room temperature for 3 days. Cell mixtures from each cross were replated on CSM-Leu/G418 plates. Wild-type (pvg+) cells of both h– and h+ mating types were the controls. Each cross-pattern showed an almost equal number of colonies. No colonies were observed in the control (uncrossed) quadrants of the plates. 1, (h– pvg+) x (h+ pvg+); 2, (h– pvg+); 3, (h– pvg–) x (h+ pvg+); 4, (h+ pvg+); 5, (h– pvg+) x (h+ pvg–); 6, (h– pvg–); 7, (h– pvg–) x (h+ pvg–); 8, (h+ pvg–).

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
We have identified in S. pombe at least five genes that contribute proteins that participate in the cell surface PvGal epitope synthesis pathway. Whereas we cannot at this point propose a complete mechanism for PvGal biosynthesis, the data are most consistent with a nearly quantitative addition of pyruvate directly to Gal{beta}1,3-residues already present on the N-linked galactomannan chain rather than by a presynthesis of PvGal-containing intermediates and transfer of PvGal to terminal outer chain Gal{alpha}1,2-residues. Since unsubstituted {beta}1,3-linked Gal was found in pvg1– glycans (Fig. 4), we suggest, based on sequence homology, that Pvg1p is a pyruvyltransferase. Of the four additional proteins that we found to be required for Gal{beta}1,3-addition (Pvg2p–Pvg5p), only the one containing the putative galactosyltransferase domain, Pvg3p, seems a likely candidate to be the required {beta}1,3-galactosyltransferase. The fact that a predicted Carbohydrate Active Enzyme data base GT31-family protein acts in the PvGal production pathway is particularly interesting, because the S. pombe Gal{beta}1,3Gal epitope shares structural similarity with the mammalian glycoprotein glycan structures (Gal{beta}1,3GlcNAc, Gal{beta}1,3GalNAc, GalNAc{beta}1,3Gal, GlcNAc{beta}1,3Gal, and GlcNAc{beta}1,3GalNAc), which are synthesized by other GT31 enzymes (66).

Whereas ascribing a regulatory role for Pvg4p, a predicted MADS-box transcription factor, in the transcription of pvg1–pvg3 and pvg5 seemed reasonable, the PvGal synthesis system does not appear to be solely under its control. No obvious canonical SRF-like Type I or MEF2-like Type II cis-acting regulatory elements for MADS-box binding were discovered in the promoter regions of the pvg2 and pvg3 genes, although each transcription factor possesses a distinct binding specificity. Potential MEF2-like Type II sequences were identified in the pvg1 and pvg5 promoters. However, expression analysis by end point RT-PCR, particularly the finding that transcription of pvg1–pvg3 and pvg5 is not significantly affected in the pvg4– mutant, does not clarify the situation. MADS-box-containing proteins specifically recruit other factors into multicomponent heteromeric regulatory complexes. It is reasonable to postulate that one or more transcriptional modulators that interact with Pvg4p as well as other Pvg4p target genes remain yet to be identified.

The functions of gene products Pvg2p and Pvg5p remain enigmatic. The annotated S. pombe genome suggests no apparent orthologs for pvg2 and pvg5, which may be S. pombe-specific (Table II). Both genes are predicted to encode transmembrane proteins, with Pvg2p experimentally located to the plasma membrane at the cell poles but Pvg5p not yet localized. Whereas bioinformatic algorithms predict a Golgi or endoplasmic reticulum location for Pvg2p, a cytoplasmic or mitochondrial location is proposed for Pvg5p (Table II). If Pvg2p and/or Pvg5p is located in the periplasmic space, one or both may interact with Pvg1p or with each other and have a role, perhaps as lectins, within a plasma membrane-bound enzyme complex that adds the terminal pyruvate to galactomannans. However, this does not fully explain why the absence of either protein results in the lack of the entire PvGal epitope and not merely the terminal pyruvate. Both or either of the two proteins may also coordinate at the plasma membrane directly or indirectly with the Pvg4p MADS-box-containing transcription factor, as part of a cell wall integrity signal transduction process. If Pvg2p and/or Pvg5p is indeed Golgi-resident, one or both may interact with the Pvg3p {beta}1,3-galactosyltransferase in a manner akin to that of members of the S. cerevisiae mannosyltransferase activity complexes (63, 64).

The Bähler group (96) has used whole genome DNA microarrays to determine which S. pombe genes are involved in a core environmental stress response (i.e. genes with significantly changed expression in response to free radical, heavy metal, thermal, osmotic, or redox stress). None of the pvg genes showed significant changes during these stresses. Similarly, the same group has also identified genes having significantly altered expression during sexual differentiation of S. pombe (i.e. conjugation, meiosis, and sporulation). The PvGal epitope synthesis genes pvg2, -3, and -5 (and possibly also pvg1) were determined to be induced "early" in sexual differentiation (97). It is important to note that the RNA for the RT-PCR experiments that we have reported here was isolated from cultures in early stationary phase and not from cultures under the nutrient-depleted conditions required to instigate sexual differentiation. The Bähler group's findings are especially interesting in light of subsequent findings that S. pombe-specific genes such as pvg1, -2, and -5, as compared with genes that are well conserved across species, are disproportionately induced during sexual development (98).

S. pombe mutants of Gal epimerase gps2 and UDP-Gal transporter gms1 are both viable but are osmosensitive with aberrant cell morphology (42, 43), indicating that the presence of Gal, and hence the PvGal epitope, is not essential. However, some possible recognition function for the epitope cannot be discounted. It was tempting to speculate that PvGal and/or the Gal{beta}1,3Gal epitope is involved in the mating process. Mutant pvg1– cells do have an increased tendency, as compared with the pvg2––pvg5– mutants, to agglutinate when reaching late stationary phase, which is consistent with the observations that galactose or wild-type galactomannans, but not mannose, glucose, or galactose-deficient {Delta}gms1 mannans, inhibit calcium-dependent nonsexual flocculation (99, 100). In addition, mannose-dependent co-flocculation with Pediococcus damnosus, which occurs at high levels for S. cerevisiae but at very low frequency for wild-type S. pombe (due to shielding by terminal Gal and PvGal), was greatly enhanced in the galactose-deficient {Delta}gms1 S. pombe (101). Although galactose has been reported to promote sexual flocculation (100) and conjugation (43), we demonstrated that cells totally devoid of pyrvuate as well as those lacking the complete PvGal epitope mate with frequencies comparable with that of wild-type cells, at least under laboratory conditions. We hypothesize that there is functional discrimination between the penultimate Gal{alpha}1,2-residue and the terminal Gal{beta}1,3-residue, with the presence of penultimate Gal{alpha}1,2-being necessary for sexual flocculation and with terminal Gal{beta}1,3-involvement in nonsexual flocculation behavior. The addition of Pv to Gal{beta}1,3Gal{alpha}1,2-thus impedes nonsexual flocculation.

In other organisms pyruvylated saccharides appear to have various functional roles. Pyruvylation of the secreted exopolysaccharides of Xanthomonas sp. contributes to the physical and chemical properties of xanthan gum (1, 19). In Rhizobium sp. and Sinorhizobium sp., acidic exopolysaccharides are necessary for symbiotic nodulation of root hairs for nitrogen fixation (22, 102). Decreases in pyruvic acid content correlate with ineffective nodule formation (103), whereas increases in pyruvyltransferase activity correlate with an increased level of Pv-Gal and with root hair attachment (104). Lactic acid bacteria also secrete exopolysaccharides, and, in S. thermophilus, disruption of the putative pyruvyltransferase EpsL in ropy strains results in nonviability (57). In apparent contradiction, nonropy strains do not possess EpsL but exhibit typical cell growth. A widely used strategy among bacteria (e.g. B. anthracis) may be the addition of pyruvate to peptidoglycan-associated polysaccharides, which appears to facilitate the anchorage of several cell surface proteins by noncovalent binding to an S-layer homology domain (23). However, Bacillus licheniformis polysaccharide contains pyruvylated Gal, but it is not covered by an S-layer lattice (105).

We do not expect to have identified all of the genes involved in or affecting PvGal synthesis. In order to select against gross glycosylation deformities, we intentionally and effectively excluded osmosensitive cells by the lack of an osmolytic support (e.g. mannitol and KCl) in the mutant enrichment process. Mechanistic links appear to remain absent. In bacterial polysaccharide synthesis, cytoplasmic phosphoenolpyruvate is employed as the high energy pyruvate donor to produce ketal-linked pyruvylated monosaccharides in the saccharide repeating units of the pyrophosphoprenyl-lipid-anchored precursors (20, 21, 106–109). However, in S. pombe, as in all eukaryotic cells, glycoprotein glycan production is a highly compartmentalized process, in which attachment of terminal monosaccharides occurs in the lumen of the Golgi apparatus. We were unable to identify reports of either pyruvic acid or energetically favorable pyruvate intermediates in the lumens of the endoplasmic reticulum or Golgi. Indeed, any such compounds would probably require specific transporters to enable them to traverse the membrane from the cytoplasm. Conversion of pyruvate into a high energy intermediate within the Golgi would almost certainly require several unique luminal proteins. Given the mutants identified to date and the structures of their N-linked glycans (Figs. 2 and 4), it seems reasonable to predict that there should be at least one pyruvate transporter and pyruvate-intermediate processing enzyme in the Golgi apparatus. Alternately, based on our evidence that Pvg1p is located at the plasma membrane, pyruvylation of terminal Gal{beta}1,3-in the outer chain extensions may occur there or in the periplasm, where there may be a role for Pvg2p and/or Pvg5p.

Experiments to identify the relationships among the identified Pvg proteins and to further elucidate the mechanisms of PvGal epitope synthesis are ongoing. These include but are not limited to the search for possible additional genes and proteins; efforts to knock out, individually and in various combinations, the five identified pvg genes; in situ expression of GFP fusion or epitope-tagged proteins; cloning, purification, and further biochemical characterization of the proteins; development of assays to determine their biochemical and biological functions; and site-directed mutagenesis to examine structure-function associations.


   FOOTNOTES
 
* This work was supported in part by National Institutes of Health, United States Public Health Service, Grant GM23900 (to R. B. T.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

The on-line version of this article (available at http://www.jbc.org) contains one additional figure and table. Back

§ Present address: Laboratory of Bacterial Toxins, Center for Biologics Evaluation and Research, Federal Drug Administration, 8800 Rockville Pike, Bethesda, MD 20892. Back

These authors contributed equally to this work. Back

** To whom correspondence should be addressed: Wadsworth Center, C-547, New York State Department of Health, P.O. Box 509, Albany, NY 12201-0509. Tel.: 518-474-5511; Fax: 518-402-5381; E-mail: trimble{at}wadsworth.org.

1 The abbreviations used are: PvGal, 4,6-pyruvylated galactose, 4–6-O-[(R)-(1-carboxyethylidine)]-Gal{beta}1,3-; Pv, pyruvate; GFP, green fluorescent protein; DDW, distilled deionized water; YEDS, yeast extract dextrose supplements medium; CSM, complete sythetic medium; SAP, human serum amyloid P component; BASHSE, biotinamidohexanoic 3-sulfo-N-hydroxysuccinimide ester; TBSC, Tris-buffered saline with calcium; TBS, Tris-buffered saline; GSL/BSL I-B4, Griffonia (Bandeiraea) simplificolia lectin I-B4 isolectin; PNA, peanut agglutinin; HPAEC, high pH anion exchange chromatography; PAD, pulsed amperometric detection; RT, reverse transcription; RPS, reverse position-specific; aa, amino acid(s); PSI, position-specific iterated; GT31, glycosyltransferase family 31; SRF, serum response factor; MEF2, myocyte-specific enhancer factor 2. Back


   ACKNOWLEDGMENTS
 
We are indebted to Dr. Tony Carr for graciously providing the S. pombe genomic library and for helpful advice on plasmid recovery, and we gratefully acknowledge the gift of pFA6a-kanMX6 plasmids from Dr. Jürg Bähler. We acknowledge use of the Wadsworth Center Structural Biology NMR Facility and of the Wadsworth Center Advanced Light Microscopy Core Facility. We thank Dr. Alexei Mikhailov for performing the light microscopy in Fig. 1A and Dr. Alexey Khodjakov for performing the fluorescence microscopy in Fig. 5. Oligonucleotide synthesis and DNA sequencing were executed in the Wadsworth Center Molecular Genetics Core. We express our gratitude to Dr. John Cipollo for valuable suggestions during the course of this work. We thank Frank Abbruscato for help with molecular techniques and Catherine Lubowski for excellent technical assistance.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

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J. Cell Sci., November 1, 2004; 117(23): 5623 - 5632.
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