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Vol. 16, Issue 2, 532-549, February 2005
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Department of Cell and Developmental Biology, University of North Carolina at Chapel Hill, Chapel Hill, NC 27599;
* Graduate Program in Pharmacology, Weill Medical College of Cornell University, New York, NY 10021
Submitted July 4, 2004;
Revised October 28, 2004;
Accepted November 11, 2004
Monitoring Editor: Keith Mostov
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSACKNOWLEDGMENTSREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSACKNOWLEDGMENTSREFERENCES |
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). Homologous genes have been characterized in Schizosaccharomyces pombe (kin1+) (Levin and Bishop, 1990), Drosophila melanogaster (dPAR-1) (Tomancak et al., 2000), and mammalian cells (mPARs: p78, EMK, MARK) (Drewes et al., 1995; Bohm et al., 1997). The mechanism of cell polarity regulation by Par-1 seems to differ depending on the organism and the cellular context. Some reports link Par-1 function to restricting localization of polarity determinants: P granules and PIE-1 in C. elegans (Kemphues et al., 1988; Tenenhaus et al., 1998); Oscar, Orb, BicD, and Egl in Drosophila oocyte (Tomancak et al., 2000; Huynh et al., 2001; Vaccari and Ephrussi, 2002); and Par-3 via 14-3-3 in Drosophila follicular epithelium (Benton and Johnston, 2003). Others find that Par-1 regulates stability, density, and/or organization of the microtubule cytoskeleton as in Drosophila and mammalian cells (Drewes et al., 1997; Shulman et al., 2000; Cox et al., 2001; Huynh et al., 2001; Doerflinger et al., 2003; Cohen et al., 2004).
In the budding yeast Saccharomyces cerevisiae, the Par-1 counterparts Kin1 and Kin2 have been isolated by homology to the kinase family of viral oncogenes (Levin et al., 1987). They belong to the Snf1 kinase family of the Ca2+/calmodulin-dependent kinase II (CaMK) group (Hanks et al., 1988). Kin1 and Kin2 are structurally similar serine/threonine kinases with an N-terminally located catalytic domain and C-terminal regulatory domain (Lamb et al., 1991; Donovan et al., 1994). The catalytic core and the C-terminal 42-amino acid tail are highly conserved between S. pombe, S. cerevisiae, C. elegans, D. melanogaster, and mammalian orthologues of Par-1. Therefore, S. cerevisiae, as a polarized organism highly amenable to genetic analysis, represents an advantageous model to study yeast Par-1 orthologues. However, little is known about the function of Kin1 and Kin2 in S. cerevisiae.
This work investigates the role of Kin1 and Kin2 proteins in polarized exocytosis in yeast. In S. cerevisiae, polarity is manifested by asymmetric growth of the bud, which requires polarized transport of Golgi-derived vesicles and their subsequent docking and fusion with the plasma membrane at the bud tip. Golgi-to-plasma membrane transport is dependent on the actin cytoskeleton and Rho/Rab GTPases, such as Rho3, Cdc42, and Sec4. Definition of the docking site and vesicle tethering is mediated by a multiprotein Exocyst complex, comprised of eight subunits: Sec3, Sec5, Sec6, Sec8, Sec10, Sec15, Exo70, and Exo84 (Guo et al., 1999). After docking, vesicle fusion with the plasma membrane involves correct pairing of one vesicle SNARE, Snc1/2, with two membrane t-SNAREs: Sso1/2 and Sec9 (Aalto et al., 1993; Protopopov et al., 1993; Brennwald et al., 1994). Although the aforementioned molecules constitute the core of the secretory machinery, polarized exocytosis is fine-tuned by many additional components.
Here, we report that yeast Par-1 orthologues regulate exocytosis. We show that Kin1 and Kin2 genetically interact with multiple components of the late exocytic machinery and physically associate with the t-SNARE Sec9 and the SNARE-binding protein Sro7. We demonstrate that Kin1 and Kin2 induce phosphorylation of Sec9 and its release from the plasma membrane, which, presumably, promotes incorporation of Sec9 into novel SNARE complexes. We further find that the conserved 42-amino acid tail of the yeast Par-1 ortholog plays a role in the autoinhibition of the kinase function in the secretory pathway of S. cerevisiae.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSACKNOWLEDGMENTSREFERENCES |
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-mercaptoethanol, Triton X-100, and protease inhibitors were obtained from Sigma-Aldrich (St. Louis, MO). Zymolase (100T) was purchased from Seikagaku (Tokyo, Japan). 125I-Protein A and [32P]ATP, 35S-TransLabel, [35S]methionine, and [32P]orthophosphate were obtained from PerkinElmer Life and Analytical Sciences (Boston, MA) and protein A-Sepharose CL-4B from Amersham Biosciences (Piscataway, NJ). Molecular weight markers and Tween 20 were from Bio-Rad (Hercules, CA). Restriction enzymes, calf intestinal phosphatase, and 
-phosphatase were purchased from New England Biolabs (Beverly, MA).
Yeast Genetic Techniques
Transformations were performed using the lithium acetate method (Becker and Guarente, 1991). Crosses of strains, tetrad dissection, and diploid sporulation were performed as described previously (Guthrie, 1991).
Antibodies
Rabbit antisera were raised against glutathione S-transferase (GST)-fusion proteins containing residues 852-1032 of the C terminus of Kin1 and containing residues 964-1121 of the C terminus of Kin2 (Cocalico Biologicals, Reamstown, PA). Antibodies were affinity purified as described previously (Lehman et al., 1999).
Plasmid Construction and Generation of Mutants
Constructs were generated by polymerase chain reaction (PCR) amplification by using primers (QIAGEN Operon, Alameda, CA) incorporating appropriate restriction sites. Primers were designed to incorporate 
1000 base pairs upstream of the open reading frame (ORF) and 250 base pairs downstream of the stop codon into the construct. SalI/EcoR1 sites were used to subclone full-length KIN1 into the pRS426 vector. To generate KIN2 constructs, KIN2 (1-1147), kin2-CT (521-1147), KIN2-NT (1-526), kin2-KD (K128M), KIN2-
42 (1-1106), and 800 base pairs of the sequence upstream of the ORF were introduced into pRS426 as a NotI/EcoR1 segment, and subsequently, wild-type and mutant KIN2 ORF sequences were subcloned into the same vector by using EcoR1/XhoI sites. Fusion PCR technique was used to generate KIN2-NT, KIN2-42, and kin2-KD. For the fusion reaction, segments with overlapping sequences generated in the first round of the amplification were resolved on 1.5% low-temperature agarose gel, cut out, and used as templates in the second round of the PCR amplification. Catalytic domains of SNF1, HSL1, GIN4, KCC4, YPL141C, KIN4, and YPL150W were mapped by BLAST sequence alignment with KIN2. NotI/XhoI sites were used to subclone HSL1 (1-462), GIN4 (1-432), KCC4 (1-437), YPL141C (1-387), and YPL150W (1-426) into pRS426 vector. NotI/SalI sites were used to subclone SNF1 (1-432) and KIN4 (1-366) into the pRS426 vector. Galactose-inducible KIN1 and KIN2 constructs were generated by insertion of PCR-amplified fragments behind the GAL promoter of GAL/HIS/CEN (BamH1/SalI sites used) and GAL/LEU/CEN (BamH1/XhoI sites used) vectors, respectively. Generation of high copy SEC9 construct and Sec9-CT (402-651) GST-fusion protein were described elsewhere (Rice et al., 1997). Sec9-NT1 (1-168) and Sec9-NT2 (166-401) GST-fusion proteins were obtained by subcloning Sec9-NT1 and Sec9-NT2 into the BamH1/SalI and BamH1/EcoR1 sites, respectively, of the pGEX4T1 vector (Amersham Biosciences). The Sec9-NT2-S315A mutant was created by fusion PCR via mutation-incorporating primers. All mutants were verified by sequencing. Generation of GST-KIN2 kinase domain (1-520) construct was obtained by subcloning a PCR-generated fragment containing kin2-NT (1-520) into the BamHI-XhoI sites of the pGEX4T1 vector. The C-terminal Kin2 fragments kin2-CT (523-1147) and kin2-CT42 (523-1106) were placed under the control of a T7 promoter for transcription and translation by PCR amplification. The upstream oligo was designed for translation beginning at residue 532 in the KIN2 coding sequence. The downstream oligo was designed to anneal 40 base pairs distal to the stop codon of KIN2. PCR products for coupled translation/transcription were generated using templates containing either KIN2 or KIN2-42 mutant. Translation of each PCR product resulted in a radiolabeled protein of the expected size on SDS-PAGE.
Suppression Assays
Late sec mutant cells were transformed with high copy plasmids (e.g., vector, KIN1, and KIN2) or a galactose-inducible Kin2 construct (vector and GALKIN2)and grown on selective medium, picked into microtiter plates, and replica plated onto YP-D medium (YP supplemented with 2% glucose) or YP-Gal (YP supplemented with 1% galactose). Transformants were tested for growth at permissive (25°C) and restrictive (14, 33, 35, and 37°C) temperatures.
Two-Hybrid Analysis
ORFs of kin2-CT (521-1147) and kin2-CT42 (521-1106) were subcloned as N-terminal fusion fragments into NcoI/BamH1 sites of pAS1-CYH2 GAL4-binding domain vector (BD). ORFs of KIN2 (1-1147), KIN2-NT (1-526), kin2-CT (521-1147), KIN2-FL42 (1-1106), and kin2-CT42 (521-1147) were inserted as N-terminal fusion fragments into NcoI/BamH1 sites of pACT2 GAL4-activation domain vector (AD). Constructs were transformed into the PJ694
strain containing GAL4-inducible HIS3 and ADE reporter genes. Construct expression was verified by Western blotting. Transformants expressing interacting proteins gained the ability to grow on -His and -Ade media. As a control empty BD and AD vectors were analyzed for the interaction with each AD and BD fusion construct, respectively. Only signals detected in the absence of control background were considered positive.
In Vitro Binding Assays
PCR generated fragments containing kin2-CT (523-1147) and kin2-CT42 (523-1106) were placed under control of a T7 promoter and in vitro transcribed and translated using a coupled reticulocyte lysate transcription/translation system (TnT; Promega, Madison, WI) in the presence of [35S]methionine. The radiolabeled proteins were diluted in binding buffer (10 mM HEPES-KOH, pH 7.4., 140 mM KCl, 2 mM MgCl2, 0.5% Triton) with GST-Kin2 kinase domain (present at 1 µM) for 1 h at 4°C. Supernatant and pellet fractions were separated and run on a 7% gel, dried, and exposed to film. Binding of radiolabeled proteins was quantified using the PhosphorImager screen and STORM ImageQuant software (Amersham Biosciences).
Construction of Deletion Strains
Gene disruption was performed via homologous recombination. Initially, flanking sequences upstream and downstream of the target ORF were introduced into the integration vector, respectively, upstream and downstream of the coding sequence of the appropriate selection marker. Plasmids were linearized before the transformation. The KIN1 gene was disrupted by the insertion of the LEU2 marker via pRS305 LEU2 vector. KIN2 gene was disrupted by the insertion of the HIS3 marker via pRS303 HIS3 vector. The genotype of the yeast strain transformed with these constructs was MAT a/; ura3-52/ura3-52; leu2-3112/leu2-3112; his3-200/his3-200. Diploids were selected on either -His or -Leu medium and sporulated. Tetrads were dissected and analyzed for the presence of the selective markers and the viability at different temperatures. Disruption was confirmed by PCR and immunoblotting with -Kin1 and -Kin2 antibodies. Crossing of the single kin1 and kin2 deletion strains generated a strain with the double deletion kin1, kin2, which was analyzed in a similar manner. The SNF1 gene was disrupted using a purified PCR product containing SNF1 sequences flanking the Kanamycin gene (snf1::KanR), which was transformed into the wild-type strain and in the strain homozygous for the KIN1 deletion and heterozygous for the KIN2 deletion: MAT a/a; kin1::LEU2/kin1::LEU2; KIN2/kin2::HIS3; leu2-3112/leu2-3112; his3-200/his3-200; ura3-52/ura3-52 to create the triple deletion mutant kin1, kin2, snf1. YPD-G418 plates were used for the selection of the KanR-containing transformants. Diploids were sporulated, and tetrads were analyzed for marker distribution. Generation of the triple mutant kin1, kin2, snf1 was confirmed by PCR.
Subcellular Fractionation
Procedure was described in detail previously (Lehman et al., 1999). Three strains were used: vector only (pRS426), KIN1 on high copy vector, and KIN2 on high copy vector. Cells were washed, spheroplasted, lysed, and divided into two pools, which were subjected to the differential treatment with or without Triton X-100 and centrifuged at 30,000 x g. Supernatant and pellet fractions were isolated and normalized for further analysis. Samples boiled in SDS sample buffer were subjected to 7% SDS-PAGE and blotted with affinity-purified antibodies to Kin1, Kin2, and Sso1/2. Signal was quantified using the PhosphorImager screen and STORM ImageQuant software (Amersham Biosciences).
BglII Secretion Assays
Secretion assays examining the effect of multicopy KIN2 were performed on sec15-1 mutant cells (ura3-52; sec15-1) transformed with KIN2 on a high copy vector or vector only (pRS426). Cells were grown overnight at 25°C and then shifted to 36°C for 1 h. Cells were then processed for BglII secretion as described previously (Adamo et al., 1999). Secretion assays on GAL induced KIN1 were performed on sec1-1 mutant cells (GAL+, ura3-52; leu2-3112; his3- 200; sec1-1) containing vector only (GAL/LEU2) or galactose inducible KIN1 on GAL/LEU2integrative vector as well as a wild-type control strain. Cells were grown overnight to an OD599 = 0.5 in YP-raffinose (3%) medium at 25°C and then induced with 1% galactose for 2 h before shifting the cells to 33°C for 2 h. Cells were then processed for BglII secretion as described previously (Adamo et al., 1999).
Native Immunoprecipitation
Cells from strains containing either high copy KIN1, KIN2, or vector alone were grown O/N in SC-D medium to OD599 = 0.5 and then shifted for 2 h to grow in YP-D medium at 25°C to OD599 = 1 to the total of 266 OD units. Subsequently, cells were harvested, washed in ice-cold 10:20:20 buffer (10 mM Tris, 20 mM sodium azide, 20 mM sodium fluoride), and spheroplasted in 17.8 ml of spheroplast buffer (100 mM Tris, 20 mM sodium azide, 20 mM sodium fluoride, 1.2 M sorbitol, 21 mM -mercaptoethanol, 0.1 mg/ml 100T Zymolyase) for 30 min at 37°C. Spheroplasts were lysed in 8.8 ml of ice-cold lysis buffer (20 mM HEPES-KOH, 150 mM KCl, 0.5% IGEPAL) with protease inhibitors (PIC): 2 mM 4-(2-aminoethyl)benzenesulfonyl fluoride, 1 mM phenylmethylsulfonyl fluoride, 20 µM Pepstatin A, 2 µg/ml leupeptin, aprotinin, and antipain. The insoluble material was pelleted by 10-min spin in a 4°C Microfuge, and the supernatant was used to set up immunoprecipitation (IP) reactions (1 ml/1 IP). Saturating amounts of affinity purified antibodies to Kin1 and Kin2 were incubated with the lysate for 1.5 h on ice. In parallel, control IPs were set up with equal amounts of purified preimmune IgG. Next, protein A-Sepharose was added to the lysates (60 µl of 1:1 suspension per IP), and tubes were placed on a nutator for 1 h. Beads were washed four times with lysis buffer and boiled in 100 µl of sample buffer. Samples were subjected to SDS-PAGE, transferred to the nitrocellulose membrane, and immunoblotted with polyclonal antibodies to Kin1, Kin2, Sec9, and Sro7 followed by I125-protein A secondary.
Dithiosuccinnimidylpropionate (DSP) Cross-linking
Strains containing high copy SEC9 and high copy KIN1 were grown overnight in SD (S medium with 2% glucose). Six OD units of cells were labeled with [35S]methionine and cysteine in 4 ml for 1 h at 30°C. Cross-linking and immunoprecipitation were performed as described previously (Lehman et al., 1999). Labeled cells were washed in phosphate-buffered saline (PBS)-azide, spheroplasted in 1 ml of spheroplast buffer, and lysed in 300 µl of PBS with PIC. The lysate was divided into two pools, one treated with the chemical cross-linker DSP dissolved in dimethyl sulfoxide (DMSO), and the control pool was treated with DMSO only. The cross-linker was quenched by ammonium acetate, samples were boiled in 5x boiling buffer (5% SDS, 50 mM Tris, pH 8.0, 25 mM EDTA) and diluted 20x with IP buffer (10 mM Tris, pH 8, 150 mM NaCl, 0.5% Tween 20, 0.1 mM EDTA). Cell lysates were then subjected to two rounds of IPs. First, samples from both pools were incubated with affinity-purified antibodies to -Kin1, -Sec9, and preimmune IgG overnight at 4°C. Immune complexes were pooled via protein A-Sepharose. For the second round of IPs beads were resuspended in the reducing boiling buffer (1% SDS, 10 mM Tris, pH 8.0, 5 mM EDTA, 0.1 mM dithiothreitol [DTT]), boiled, diluted with IP buffer, and supernatants were subjected to IPs with either affinity-purified -Kin1 or affinity-purified -Sec9 antibodies. Samples were boiled and resolved on 7% SDS-PAGE, and a 35S signal was detected by autoradiography.
Nonnative Immunoprecipitation and Phosphatase Treatment
Yeast strains containing high copy SEC9 and either inducible KIN2 on GAL/LEU/CEN integrative vector or GAL/LEU/CEN empty vector were grown in SC medium with 3% raffinose overnight and induced with 1% galactose for 4 h at 25°C. Then, 100 OD units/each were harvested, washed, spheroplasted in Eppendorf tubes (5 OD/tube), and lysed in 40 µl/tube TEAE/sorbitol (10 mM triethanolamine, 1 mM EDTA, pH 7.2, 0.8 M sorbitol with 1x protease inhibitor mix). Lysates were centrifuged, combined with an equal volume of 2x boiling buffer (10 mM Tris, pH 8.0, 25 mM EDTA pH 8.0, 1% SDS), boiled at 95°C for 5 min, and diluted 20x with IP buffer (10 mM Tris, pH 8.0, 150 mM NaCl, 0.1 mM EDTA, 0.5% Tween 20). Combined supernatants were distributed into Eppendorf tubes (1.4 ml/tube) and immunoprecipitated with antisera to Sec9 (6 µl/tube) overnight on ice. Immune complexes were pooled down with protein A-Sepharose (65 µl/tube) for 2 h at 4°C and washed with IP buffer five times. Combined beads were distributed into Eppendorf tubes (25-µl bed volume/tube), which were subjected to five treatments: 1) boiled immediately; 2) incubated 1:1 with 2x restriction buffer 3 and 2 µl/tube of calf intestinal phosphatase (CIP) for 30 min at 37°C; 3) incubated with 2x buffer 3 only for 30 min at 37°C (mock control for CIP); 4) incubated 1:1 with 2x -phosphatase buffer and -phosphatase (1.5 µl/tube) for 30 min at 30°C; and 5) incubated with 2x -phosphatase buffer only for 30 min at 30°C (mock control for -phosphatase). Samples were boiled, separated on an 8% gel, and immunoblotted with -Sec9 antibody.
Purification of the Recombinant Proteins
All GST-fusion proteins were produced, purified, and their protein concentrations estimated as described previously (Rossi et al., 1997).
In Vitro Kinase Assays
Yeast strains containing high copy KIN1, KIN2, or empty vector were grown on SC-D (SC medium with 2% glucose) overnight and switched to YP-D medium for 2.5 h. Cells were harvested (33 OD units/1 IP reaction), washed in 10:20:20 buffer, spheroplasted, lysed, and subjected to native immunoprecipitation with antibodies to Kin1, Kin2, or preimmune serum as described above. Immune complexes were pulled down with protein A-Sepharose (60 µl of 1:1 slurry/1 IP), beads were washed twice with lysis buffer and twice with PK buffer (50 mM Tris, pH 7.5, 5 mM MgCl2), and combined and distributed into Eppendorf tubes. For one kinase reaction (total volume of 50 µl), we used 25 µl of 1:1 bead slurry, 1.5 µM recombinant protein, 1x PKi buffer (50 mM Tris, pH 7.5, 5 mM MgCl2, 1 mM DTT), 0.1 mM cold ATP, and 1 µl of [32P]ATP. The kinase reaction was incubated at 30°C for 30 min, spun, and the supernatant containing recombinant proteins was boiled in an equal volume of sample buffer. Samples were subjected to 15% SDS-PAGE, and gels were dried and exposed to film.
In Vivo Kinase Assays
Cells containing multicopy SEC9 and GAL-KIN2 (GAL-KIN2/CEN/LEU2) or empty vector control (GAL/CEN/LEU2) were grown O/N in S medium with 3% raffinose to an OD599 of 0.5. Next day the cells were split in half and shifted into fresh medium containing either S with 2% raffinose or YP-low phosphate with 2% raffinose for 1 h at 27°C. Two ODs of the culture grown in S with 2% raffinose were then induced with 1% galactose and labeled with [35S]methionine (0.28 µCi/ml) for 4 h at 27°C. 2 ODs of the culture growing in YP-low phosphate with 2% raffinose were simultaneously induced with 1% galactose in the presence of [32P]orthophosphate (10 µCi/µl) for 4 h at 27°C. At the end of the incubation, the cultures were spun in glass tubes for 5 min at 25°C. All samples were washed in Tris (10 mM), sodium azide (20 mM), and sodium fluoride (20 mM) and then spheroplasted in spheroplast buffer (100 mM Tris, 1.2 M sorbitol, 10 mM sodium azide, 0.015% -mercaptoethanol, 0.1 mg/ml Zymolyse 100T) for 30 min at 37°C. Subsequently, samples were spun in Eppendorf tubes for 5 min at 2000 rpm. The pellets were lysed in 75 µl of ice-cold 1x PBS and then boiled immediately in 2x boiling buffer (2% SDS, 20 mM Tris, pH 8, 10 mM EDTA) for 5 min at 95°C. Samples were diluted with 1 ml of IP buffer (10 mM Tris, pH 8, 150 mM NaCl, 0.5% Tween 20, 0.1 mM EDTA), and microfuged for 10 min at 4°C. Supernatants were then removed, further diluted in IP buffer, and used for immunoprecipitations with respective antibodies for 1 h on ice. For [32P]orthophosphate-labeled cells, RNAse A (DNase, protease free; Sigma-Aldrich) was added to 30 µg/ml (1 µl of enzyme/ml IP buffer) to reduce background on gels from radiolabeled RNA contaminants. Immune complexes were pulled down with protein A-Sepharose, washed with IP buffer, and IP buffer with 2 M urea and 1% betamercaptoethanol before boiling in 100 µl of SB. Samples were run on SDS polyacrylamide gels, dried, and exposed to film.
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSACKNOWLEDGMENTSREFERENCES |
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; Robinson et al., 1999). To isolate candidate downstream effectors of RHO3 function in exocytosis, we performed a genetic screen for dosage suppressors of the slow growth phenotype of a RHO3 deletion strain (rho3). We constructed a strain containing the RHO3 coding sequence under the control of the glucose-repressible GAL promoter as the only source of Rho3 in the cell. This strain, which grows normally on galactose-containing media but extremely slowly in glucose-containing media, was transformed with a yeast genomic library prepared in a multicopy vector, and transformants were selected for growth on glucose-containing medium. Plasmids from colonies growing on glucose were isolated, retested for suppression, and then analyzed by sequencing. From this analysis, we identified 25 suppressing plasmids containing overlapping parts of six distinct loci. Five of the six loci had genes previously isolated as dosage suppressors of a rho3 mutant: BEM1 (1 isolate), SRO9 (1 isolate), SEC4 (2 isolates), SSO2 (3 isolates), and RHO3 itself (17 isolates) (Imai et al., 1996; Adamo et al., 1999). Subcloning of the various regions in the sixth locus identified the suppressing gene to be coincident with the KIN1 gene (Figure 1A). Although several suppressors of rho3 (SEC4, SRO7, SRO77, SEC9, and SSO2) are known components of the late secretory machinery, the function of KIN1 is not known. The KIN1 open reading frame alone was sufficient to suppress rho3, and thus we identified KIN1 as a novel dosage suppressor of Rho3.
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Mutations in RHO3 affect both actin organization and post-Golgi vesicle transport (Imai et al., 1996; Adamo et al., 1999). The rho3-V51 cold-sensitive mutant has a secretory defect in the absence of functional and structural perturbations of the actin cytoskeleton; hence, this mutant affects a function of Rho3 in exocytosis that is independent of actin (Adamo et al., 1999). To determine whether Kin1 functions specifically in the secretory pathway downstream of Rho3, we assessed the ability of KIN1 to suppress the rho3-V51 mutant. rho3-V51 is viable at 25°C, but not at 14°C. Figure 1B shows that overexpression of KIN1 resulted in restoration of growth of the rho3-V51 mutant at the restrictive temperature (14°C). In addition, we have previously reported that the growth defect of cdc42-6, the secretion-impaired mutant of another Rho GTPase, is specifically suppressed by introduction of KIN1 on the multicopy plasmid (Adamo et al., 2001). This suppression was specific to cdc42-6, because KIN1 failed to suppress more pleiotropically defective alleles of CDC42 such as cdc42-1.
Because the coding sequence of KIN2 is similar to that of KIN1 (51% identity), we asked whether they shared a common function downstream of Rho3 and Cdc42 by determining the ability of high-copy KIN2 to suppress the growth defects associated with rho3-V51 and cdc42-6 mutant strains. As shown in Table 1, we found that KIN2 strongly suppressed the growth defect of these mutants at their respective restrictive temperatures (14°C for rho3V-51 and 32°C for cdc42-6), and the potency of KIN2 was identical to that of KIN1. The fact that both kinases specifically suppress secretory-defective alleles of Rho3 and Cdc42 suggests that both KIN1 and KIN2 act in the secretory pathway downstream of the Rho GTPases.
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KIN1 and KIN2 Exhibit Genetic Interactions with Components of the Late Secretory Machinery
The ability of KIN1 and KIN2 to suppress the cdc42-6 and rho3-V51 alleles suggests a possible role for these kinases in the regulation of exocytosis. To further explore this idea, we examined the ability of multicopy KIN1 and KIN2 to suppress the temperature-sensitive growth defect of a number of secretory (sec) mutants. An example of the suppression analysis, summarized in Table 2, is shown in Figure 2. Elevated dosage of KIN1 and KIN2 restored the growth of the late sec mutants sec4-P48 and sec15-1 to wild-type levels at 14 and 35°C, respectively (Figure 2). sec4-P48 is a cold-sensitive effector domain mutant of the Rab GTPase SEC4 (Brennwald et al., 1994), whereas sec15-1 is a temperature-sensitive mutant of one of the components of the Exocyst complex, Sec15 (TerBush and Novick, 1995). We examined the secretory defect in sec15-1 cells with multicopy KIN2 compared with control sec15-1 cells containing empty vector after a shift to the restrictive temperature. This analysis demonstrated that although unsuppressed sec15-1 cells are found to accumulate 36% of BglII internally, the accumulation is reduced to wild-type levels in sec15-1 cells containing high-copy KIN2 where only 18% of BglII is found internally. Therefore, the suppression of the growth defect was found to correlate closely to suppression of the secretion defect in these cells. Suppression analysis demonstrated that KIN1 and KIN2 also suppress the growth defect of a number of other late secretory mutants. These include the temperature-sensitive mutants of two additional components of the Exocyst complex; Sec3 (sec3-2) and Sec10 (sec10-2), which were rescued at the nonpermissive temperature of 35°C by introduction of KIN1 and KIN2 on high copy. Also, we show that expression of multicopy KIN1 and KIN2 restores the growth of sec1-1 at 33°C and sec2-41 at 33°C. sec1-1 is a mutant of SEC1, which is involved in SNARE assembly (Carr et al., 1999), and sec2-41 is a mutant in SEC2, the nucleotide exchange factor for Sec4 (Walch-Solimena et al., 1997). Thus, KIN1 and KIN2 exhibit strong genetic interactions with multiple components of the exocytic machinery. The suppression profile of KIN1 was identical to that of KIN2, providing additional evidence in favor of the functional redundancy of the two kinases (Table 4). The ability of KIN1 and KIN2 to suppress several late sec mutant genes indicates that they function downstream of these proteins at the later stage of exocytosis.
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However, we found that KIN1 and KIN2 do not suppress the temperature-sensitive phenotype of the sec9-4 mutant (Table 2), a mutant of the t-SNARE Sec9 that is defective in SNARE complex assembly and that is required for vesicle fusion with the plasma membrane (Rossi et al., 1997), nor do they suppress the growth defect of sro7/77, a mutant with a double disruption of genes encoding Sec9-binding proteins Sro7 and Sro77 (Table 4). These observations place Kin1 and Kin2 function downstream of polarized vesicle delivery and upstream of the terminal fusion event.
Analysis of the Structural Requirements for KIN1 and KIN2 Function in the Secretory Pathway
To elucidate the nature of Kin1 and Kin2 function in the secretory pathway, we determined the minimal domain requirement that confers suppression. Kin1 and Kin2 contain a kinase domain at the N terminus of the protein and a regulatory domain at the C terminus (Figure 3A). The kinase domain as well as the 42-amino acid stretch on the extreme carboxy terminus are highly conserved between Kin1 and Kin2 and a number of their orthologues from other species. Because Kin1 and Kin2 proteins display structural and functional redundancy, we focused on Kin2. Mutant KIN2 constructs with deletions of the kinase domain or the 42 amino acid C-terminal tail were designed to assess the significance of these domains for Kin2 function in the secretory pathway. The following mutants were generated: KIN2-NT, lacking the regulatory C-terminal domain of the protein; kin2-CT, lacking the catalytic N-terminal domain; kin2-KD, the kinase-dead mutant, where a single critical Lys128 residue in the second catalytic domain (conserved residue mapped by kinase sequences alignment; Hanks et al., 1988) was mutated to a Met; and finally the KIN2-42 mutant, with a deletion of the conserved 42 amino acid C-terminal tail (Figure 3A).
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Function was assayed by analysis of the suppression properties of the KIN2 mutants expressed at high copy. The suppression of the mutant phenotype of several late sec genes was tested, including sec15-1 and sec4-P48, sec1-1, sec2-41, and sec10-2 (Figure 3B). Wild-type KIN2 on a multicopy plasmid behaved as reported above (Figure 2 and Table 2), restoring the viability of these mutants at certain restrictive temperatures. The kinase-inactive KIN2 mutants kin2-CT, lacking the entire kinase domain, and kin2-KD, the kinase-dead mutant, failed to suppress the growth defect of all sec mutants tested (Figure 3B). These data demonstrate that the catalytic activity of Kin2 is critical for its function in the secretory pathway.
Multicopy expression of KIN2-NT, lacking the entire C-terminal domain, and KIN2-42, lacking the 42 amino acid C-terminal tail, rescued the growth phenotype of all late sec mutants tested: sec15-1, sec4-P48, sec1-1, sec2-41, and sec10-2 (Figure 3B). Thus, the regulatory domain of Kin2 is functionally dispensable. Furthermore, we observed that the KIN2-NT and KIN2-42 mutants at high copy gain the ability to suppress several secretory mutants: sec1-1, sec2-41, and sec10-2, at temperatures at which the wild-type KIN2 failed to suppress (Figure 3B). The sec1-1 and sec2-41 temperature-sensitive mutants are suppressed at 33°C and the sec10-2 mutant at 35°C in a comparable manner by KIN2, KIN2-NT, and KIN2-42 (our unpublished data). However, KIN2 fails to suppress sec1-1 at 35°C and sec2-41 and sec10-2 at 37°C, whereas the KIN2-NT mutant with the deletion of the regulatory domain is able to rescue these mutants at the more restrictive temperatures. Remarkably, the truncation of the distal 42 amino acids at the C terminus of KIN2, in KIN2-42, is sufficient to phenocopy the gain of function observed by KIN2-NT. Both mutant forms of KIN2 are capable of suppressing sec1-1 at 35°C and sec2-41 and sec10-2 at 37°C (Figure 3B). These data demonstrate that the C-terminal nonkinase domain of Kin2 acts as a negative regulator of Kin2 function in the secretory pathway and that the conserved 42-amino acid tail is essential for this negative regulatory function.
The greater potency of the Kin2 constructs lacking the distal C-terminal sequence might reflect the acquisition of the catalytically "active" protein conformation in the absence of the putatively inhibitory C-terminal tail. Possibly, in a dormant state the wild-type kinase exists in a closed conformation, with the tail bound to the catalytic core, hindering its activity, until the "ON" regulatory event relieves this autoinhibition (Figure 3C). This hypothesis presupposes the presence of a direct physical interaction between the tail of Kin2 and its kinase domain. To test whether this intramolecular interaction takes place, we used a yeast two-hybrid analysis. We created the following constructs: Kin2-CT and Kin2-CT42 (encoding the regulatory domain with and without the C-terminal tail region, respectively) as GAL4 binding domain fusions and Kin2 (full-length protein), Kin2-NT (kinase domain), Kin2-CT (regulatory domain), Kin2-CT42 (regulatory domain with the deletion of the C-terminal 42 amino acids) and Kin2-42 (full-length protein with the deletion of C-terminal 42 amino acids) as GAL4 activation domain fusions. All constructs in activation and binding domain fusions were expressed at comparable levels as verified by Western blot analysis (our unpublished data). We found that the C-terminal regulatory domain of Kin2 (Kin2-CT in GAL4BD) binds to the catalytic N-terminal domain of Kin2 (Kin2-NT in GAL4AD) (Table 3). Moreover, this interaction is mediated by the conserved 42 amino acid tail, because it is abolished by truncation of the tail region in the C-terminal domain of Kin2: Kin2-CT42 does not bind to Kin2-NT. Kin2-CT does not interact with itself, which is consistent with the regulatory domain interacting with the kinase domain only. In addition, although Kin2-CT fails to interact with the full-length Kin2, it shows interaction with the C-terminally truncated Kin2 Kin2-42 (lacking the distal 42 amino acids). This result may signify that Kin2-CT associates with Kin2 in a presumably "open" or active conformation, as in Kin2-42, but not with Kin2 in a closed conformation, as in full-length kinase. Thus, yeast two-hybrid analysis revealed that the regulatory domain of Kin2 binds to its catalytic domain and that the 42-amino acid tail is a prerequisite for this interaction. To further support the possible interaction of the Kin1/2 kinase domain with the C-terminal domain, we examined the ability of the domains to interact in vitro. We made use of a recombinant GST-fusion of the kinase domain and in vitro-translated C-terminal domain fragments containing either the intact C terminus, Kin2-CT (523-1147), or an identical domain lacking the conserved C-terminal 42 amino acids, Kin2-CT42 (523-1106). The results shown in Figure 3C demonstrate that these domains do in fact interact and that this interaction requires the C-teminal 42 residues of the Kin2. Together, the in vitro binding data and the two-hybrid analysis strongly suggest that the C-terminal regulatory domain of Kin2 physically interacts with the N-terminal kinase domain to mediate an autoinhibitory regulation of the kinase. Furthermore, we show that the highly conserved 42-amino acid tail of Kin2 is critical for the both the physical interaction between these domains and for the negative regulatory effect of the C-terminal domain as demonstrated by the effects on suppression of the late sec mutants. Together, this strongly suggests that C-terminal domain of Kin1/2 functions as an autoinhibitory domain and that this autoinhibition requires the highly conserved C-terminal 42 amino acids. This provides the first mechanistic insight into to the function of this highly conserved 42-residue sequence at the C terminus of all Par-1 family kinases.
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The Kinase Domain of SNF1, but Not Other Kinases of the CaMK Group, Show Suppression Properties of the Kin1 and Kin2 Kinases
It was previously reported that deletion of either KIN1 or KIN2 is neither lethal nor deleterious for cell growth (Lamb et al., 1991; Donovan et al., 1994). Because our data demonstrated functional redundancy of KIN1 and KIN2, we proceeded to analyze whether the presence of at least one of these genes is required for cell viability. We created strains carrying single or double disruptions of these genes by homologous recombination. Consistent with the previous reports, kin1 and kin2 single disruptant strains were viable. The double disruptant kin1, kin2 progeny, obtained by crossing of the kin1 strain with kin2, demonstrate normal growth at all temperatures tested (25, 14, and 37°C). The single, kin1 and kin2, and double kin1, kin2 disruptants did not exhibit any significant defects in growth or secretion, as confirmed by invertase secretion assays (our unpublished data).
KIN1 and KIN2 orthologues in S. pombe, C. elegans, D. melanogaster, and mammalian cells display pronounced gene disruption phenotypes. Hence, it is likely that in S. cerevisiae anther molecule(s) acts to substitute the compromised function of Kin1 and Kin2. Kin1 and Kin2 belong to the CaMK protein kinase group, members of which share significant sequence similarity in the catalytic domain. Therefore, we examined whether other kinases of this group display functional redundancy with Kin1 and Kin2. BLAST search was used to identify the closest homologues of Kin1 and Kin2, and, as a result, we considered seven proteins of the CaMK group for further analysis, including Snf1, Hsl1, Gin4, Kcc4, Ypl141c, Kin4, and Ypl150w. To determine whether these proteins act in the secretory pathway, we tested their ability to rescue the growth defect of sec2-41, sec10-2, and sec15-1. Because several members of the CaMK group, such as Hsl1, Gin4, and Kcc4 bear a sequence at the distal carboxyl tail almost identical to that of Kin1 and Kin2, we hypothesized that they might be subjected to a similar mode of autoregulation as Kin2. To overcome this possible autoinhibition, we generated deletion constructs of SNF1 (encoding amino acids 1-432), HSL1 (1-462), GIN4 (1-432), KCC4 (1-437), YPL141C (1-387), KIN4 (1-366), and YPL150W (1-426), which lack the regulatory domain while preserving intact all regions important for catalytic activity based on sequence alignment. These constructs were introduced on a multicopy plasmid into sec15-1, sec10-2, and sec2-2 mutants and incubated at permissive and nonpermissive temperatures. We show that the catalytic domain of SNF1 (SNF1-NT) suppresses the growth defect of sec15-1, sec10-2, and sec2-2 mutants (Figure 4). Furthermore, the suppression of sec mutants by SNF1-NT is comparable to that of KIN2-NT. By contrast, constructs encoding the catalytic domains of Hsl1, Gin4, Kcc4, Ypl141C, Kin4, and Ypl150W failed to restore growth of the sec mutants at restrictive temperatures to any significant degree. Our data demonstrate that Snf1, which is structurally closer to Kin1 and Kin2 than any of the other seven members of the CaMK group, is the only kinase, out of those tested that may function redundantly with Kin1 and Kin2 in exocytosis.
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To test whether the function of at least one of these genes, KIN1, KIN2, or SNF1, is essential for cell viability, we generated the strain with a triple disruption, kin1, kin2, snf1. kin1, kin2, snf1 was created by substitution of SNF1 with the KanR (Kanamycin) gene sequence in the context of the diploid strain homozygous for the KIN1 deletion (kin1) and heterozygous for the KIN2 deletion (kin2). A triple disruptant was obtained via sporulation and tetrad analysis. The triple mutant kin1, kin2, snf1 as well as the double kin1, snf1, and kin2, snf1 mutants displayed slow growth phenotype inherent to the SNF1 deletion alone (Celenza and Carlson, 1984). The kin1, kin2, snf1 triple mutant did not display any synthetic defect in growth or secretion (as verified by invertase secretion assays; our unpublished data).
Kin1 and Kin2 Physically Associate with Components of the Late Exocytic Machinery
Because genetic data place Kin1 and Kin2 function to the late secretory pathway, we analyzed whether these proteins physically associate with any of the components of the exocytic machinery.
Initially, we generated antibodies to detect Kin1 and Kin2 and characterized their localization in budding yeast. Antibodies were raised against a region in the C-terminal domain of Kin1 and Kin2. Affinity-purified antisera were tested by Western blot analyses on samples containing either high copy KIN1 and KIN2 or empty vector (Figure 5A). Kin1 and Kin2 antisera specifically recognized each protein and did not cross-react with the other gene product, with both proteins running on SDS-PAGE at 145 kDa as predicted by the estimated molecular weight (Lamb et al., 1991; Donovan et al., 1994). Next, we analyzed the intracellular distribution of Kin1 and Kin2 by cell fractionation. Lysates from cells overexpressing Kin1 or Kin2 were treated with or without Triton X-100 and centrifuged at 30,000 x g. Supernatant and pellet fractions obtained were analyzed by SDS-PAGE and Western blot with anti-Kin1, anti-Kin2, and anti-Sso1/2 (as an internal control) antibodies (Figure 5B). We determined that 70% of both Kin1 and Kin2 occur in the supernatant or cytosolic fraction (precisely 71.5% of Kin1 and 73.7% of Kin2 as an average of 3 or more experiments), and the remaining 30% is found in the pellet or membrane-bound fraction. Therefore, consistent with the previous report (Tibbetts et al., 1994), Kin1 and Kin2 partition to both cytosolic and membrane-associated pools in S. cerevisiae.
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Genetic data suggest that Kin1 and Kin2 act downstream of Rho3, Cdc42, Sec4, and components of the Exocyst complex but upstream of the t-SNARE Sec9 and the Sec9-binding protein Sro7. Therefore, we hypothesized that Kin1 and Kin2 interact and function together with proteins important for the final stages of exocytosis, such as the SNAREs. To test this, we used the candidate approach to search for Kin1- and Kin2-interacting partners. First, we observed that antibodies against both Kin1 and Kin2 bring down the t-SNARE Sec9 when both genes are expressed at high copy (Figure 6A). Sec9 immunoprecipitation with Kin1 and Kin2 antibodies was detected with the endogenous levels of Kin1 and Kin2 as well (our unpublished data). To confirm that Kin1 exists in a protein complex with Sec9 by a different method, we used a chemical cross-linking procedure. Cells from strains overexpressing both KIN1 and SEC9 were labeled with [35S]methionine for 1 h and then lysed. Lysed cells were treated with the chemical cross-linker DSP and subjected to two rounds of immunoprecipitation. In the first round, samples were divided into two pools and incubated with affinity-purified antibodies either against Kin1 or against Sec9, and immune complexes formed were pulled down via protein A-Sepharose. In the second round, each of the two pools was subjected to a denaturing immunoprecipitation with either anti-Sec9 or anti-Kin1 antibodies. Our results show that in the presence of the cross-linker Kin1 coimmunoprecipitates Sec9, and in its turn, Sec9 pulls down Kin1 (Figure 6B).
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To identify other potential Kin1- and Kin2-interacting partners, we performed a series of native immunoprecipitation experiments testing for the association of Kin1 and Kin2 with proteins acting in the secretory pathway. Namely, we examined whether antibodies against Kin2 can pull down two other exocytic SNAREs, Sso and Snc, the Sec9-interacting protein Sro7, and the Rab GTPase Sec4. Kin2 was immunoprecipitated from the cell lysates carrying multicopy KIN2, and the sample was analyzed by Western blotting with -Kin2, -Sso2, -Snc1, -Sro7, and -Sec4 antibodies, respectively. This experiment revealed that the -Kin2 antibody brings down significant amounts of Sro7, a homologue of a tumor suppressor protein lethal giant larvae (Lgl), but not other proteins tested (Figure 6C). As expected, Sro7 also coimmunoprecipitates with anti-Kin1 antibodies (our unpublished data). Therefore, Kin1 and Kin2 associate with t-SNARE Sec9 and the Sec9-binding protein Sro7.
To determine whether association of Kin1 and Kin2 with Sec9 and Sro7 is a result of a direct binding between these proteins, we used a yeast two-hybrid analysis. Kin1 and Kin2 in both GAL4 binding and GAL4 activation domain fusions did not show interaction with either Sec9 or Sro7 in GAL4 activation and GAL4 binding fusions, respectively (our unpublished data). This indicates that the interaction of Kin1 and Kin2 with Sec9 and Sro7 is not direct but occurs via other intermediaries in the complex.
The physical association of Kin1 and Kin2 with the SNARE machinery supports the hypothesis that these Par-1 counterparts play a role in exocytosis at a stage between vesicle docking site recognition and fusion with the plasma membrane.
Kin1 and Kin2 Induce Phosphorylation of Sec9 In Vivo and Its Release from the Plasma Membrane to the Cytosol
Next, we focused on finding a downstream target of Kin1 and Kin2 function in the exocytic pathway. We observed that the interaction partner of Kin1 and Kin2, the t-SNARE Sec9, undergoes a size shift upon transient overexpression of the catalytically active Kin2 kinase (Figure 7A). Overexpression of Kin1 gave a similar shift in Sec9 mobility but induction of the kinase-dead mutants of Kin1 and Kin2 failed to induce any detectable shift in Sec9 (our unpublished data). To test whether this shift is indeed the result of phosphorylation, we analyzed the effect of phosphatase treatment on the mobility of immunoprecipitated Sec9 protein after Kin2 induction. Cells carrying either vector alone or CEN/GAL KIN2 were incubated in galactose-containing medium for 4 h to induce Kin2 expression, and lysates obtained from these cells were subjected to immunoprecipitation with anti-Sec9 antibodies. Sec9-containing immune complexes were subsequently treated with two different phosphatases: -phosphatase or CIP. Phosphatase treatment, but not mock control treatment, abolished the Kin2-dependent size shift of Sec9, demonstrating that the change in Sec9 mobility in response to Kin2 induction is indeed due to phosphorylation (Figure 7A).
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We next examined whether Sec9 is phosphorylated directly by Kin1/Kin2 in vitro. We made use of recombinant Sec9 protein as a substrate in an in vitro kinase reaction and looked for [
-32P]ATP incorporation in the presence of immunoprecipitated Kin1/Kin2 proteins bound to protein A-Sepharose beads. The data shown represent phosphorylation reactions in the presence of Kin2; however, we found virtually identical results for immunoprecipitated Kin1 in these assays. The recombinant Sec9 protein was initially divided into three domains, Sec9-NT1 (amino acids 1-168), Sec9-NT2 (amino acids 166-401), and Sec9-CT (corresponding to the SNAP25 domain amino acids 401-651), each of which were fused to GST, expressed in bacteria, and purified as described previously (Rossi et al., 1997). This analysis demonstrated that the Sec9-NT2 protein turned out to be an excellent substrate for Kin2 (Figure 7B, 1), with phosphoacceptor activity significantly greater than that of casein, which was previously identified as a test substrate of Kin1 and Kin2 in vitro (Lamb et al., 1991; Donovan et al., 1994). We subsequently mapped the site of Sec9 phosphorylation by Kin2 to serine 315 by sequential deletion and mutagenesis of serine or threonine residues to alanine. In particular, we found that the substitution of serine 315 to alanine abolished the ability of Kin2 to phosphorylate Sec9-NT2 in vitro (Figure 7B, 2). We used the same strategy to map a significantly weaker in vitro phosphoacceptor site in the SNAP-25 domain of Sec9 to serine 632 (our unpublished data). We next determined the effect of the mutation of these sites on the Kin2-induced phosphorylation of Sec9 in vivo. Surprisingly, we found that the Sec9-S315A as well as Sec9-S315A, S632A proteins were identical to wild-type Sec9 in the Kin2 induced phosphorylation as judged by a mobility shift (Figure 7C). Therefore, the major phosphoacceptor sites on Sec9 phosphorylated by Kin2 in vitro are not responsible for Kin2-induced phosphorylation of Sec9 in vivo. This result indicates that in vivo Kin1/Kin2 are unlikely to directly phosphorylate Sec9, but rather this phosphorylation occurs as a downstream effect of Kin1/Kin2 induction, presumably by direct or indirect activation of a kinase, which in turn phosphorylates Sec9 at a site or sites other than serine 315.
To assess the specificity of the catalytic activity of Kin1 and Kin2, we examined whether other components of the late exocytic machinery are phosphorylated by these kinases. In vivo experiments were performed on cells carrying either vector alone or CEN/GAL KIN2, radioactively labeled with [32P]orthophosphate during a 4-h induction of Kin2 in galactose-containing medium. Lysates from these cells were immunoprecipitated with antibodies against Sec9, Sso1/2, Sro7, and a number of components of the Exocyst complex and a subset of small GTPases involved in secretion. An identical set of strains was simultaneously labeled with [35S]methionine to control for the presence of the proteins in the lysates examined. Proteins were separated on SDS-PAGE, and their phosphorylation state was determined by autoradiography. Out of all proteins tested, only Sec9 displayed an increased level of [32P]orthophosphate incorporation in cells overexpressing Kin2 (Figure 8). Thus, Kin2 specifically induces the phosphorylation of Sec9. These data allow us to hypothesize that Kin1 and Kin2 act in the secretory pathway by regulating the phosphorylation of the t-SNARE Sec9.
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To address the functional significance of the Sec9 phosphorylation induced by Kin1 and Kin2, we tested whether expression of these kinases affects the subcellular localization of Sec9. We analyzed the distribution of Sec9 into pellet and supernatant fractions after centrifugation at 30,000 x g in galactose-induced and uninduced cells carrying CEN/GAL KIN1. In uninduced cells (as well as in cells carrying empty CEN/GAL vector; our unpublished data) 50-60% of Sec9 is cytosolic and partitions into the supernatant fraction, whereas the rest is membrane bound (the Triton-sensitive pellet fraction) (Figure 9). Interestingly, in cells expressing CEN/GAL KIN1 the proportion of the cytosolic Sec9 is increased relative to the membrane-bound Sec9 (Figure 9). Averaging three independent experiments, induction of Kin1 expression resulted in reproducible elevation of the cytosolic Sec9 levels to 70-75% of the total Sec9 pool. As expected, Kin1 does not alter distribution of Sso1/2 under identical conditions (Figure 9). Furthermore, Sec9 undergoes a Kin1-mediated mobility shift exclusively in the cytosolic but not the membrane fraction. As expected, induction of KIN2 had the same effect on Sec9 distribution (our unpublished data). Thus, overexpression of Kin1 or Kin2 results in release of a fraction of Sec9 from the plasma membrane into the cytosol.
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To determine the effect of GAL-induced Kin1 overexpression on overall growth and secretory function, we examined the ability of sec1-1 cells transformed with a GAL-KIN1 construct to grow and secrete the periplasmic protein BglII. As shown in Figure 9B, we find that galactose-induced expression of Kin1 protein results in dramatic suppression of the growth defect at the </
