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Kin1 and Kin2 are Saccharomyces cerevisiae counterparts of Par-1, the Caenorhabditis elegans kinase essential for the establishment of polarity in the one cell embryo. Here, we present evidence for a novel link between Kin1, Kin2, and the secretory machinery of the budding yeast. We isolated KIN1 and KIN2 as suppressors of a mutant form of Rho3, a Rho-GTPase acting in polarized trafficking. Genetic analysis suggests that KIN1 and KIN2 act downstream of the Rab-GTPase Sec4, its exchange factor Sec2, and several components of the vesicle tethering complex, the Exocyst. We show that Kin1 and Kin2 physically interact with the t-SNARE Sec9 and the Lgl homologue Sro7, proteins acting at the final stage of exocytosis. Structural analysis of Kin2 reveals that its catalytic activity is essential for its function in the secretory pathway and implicates the conserved 42-amino acid tail at the carboxy terminal of the kinase in autoinhibition. Finally, we find that Kin1 and Kin2 induce phosphorylation of t-SNARE Sec9 in vivo and stimulate its release from the plasma membrane. In summary, we report the finding that yeast Par-1 counterparts are associated with and regulate the function of the exocytic apparatus via phosphorylation of Sec9.
Par-1 has been implicated in providing intracellular polarization cues in various biological systems. Initially, Par-1 has been described in Caenorhabditis elegans as one of the “partitioning” genes essential for anterior-posterior axis specification and asymmetry of zygote division (Kemphues et al., 1988 ). 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.
Yeast cells were grown in either YP medium (1% bacto-yeast extract, 2% bactopeptone; Difco, Detroit, MI), S medium (0.67% yeast nitrogen base; Difco), or SC medium (0.67% yeast nitrogen base with dropout selection-appropriate amino acid nutrients) supplemented with 2% glucose or 1% galactose. Sorbitol, sodium azide, sodium fluoride, N-ethylmaleimide, β-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 λ-phosphatase were purchased from New England Biolabs (Beverly, MA).
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).
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 ).
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-Δ42 (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-Δ42, 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-CTΔ42 (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-Δ42 mutant. Translation of each PCR product resulted in a radiolabeled protein of the expected size on SDS-PAGE.
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.
ORFs of kin2-CT (521-1147) and kin2-CTΔ42 (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-FLΔ42 (1-1106), and kin2-CTΔ42 (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 PJ694α 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.
PCR generated fragments containing kin2-CT (523-1147) and kin2-CTΔ42 (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).
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/α; ura3-52/ura3-52; leu2-3112/leu2-3112; his3-Δ200/his3-Δ200. 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 α-Kin1 and α-Kin2 antibodies. Crossing of the single kin1Δ and kin2Δ deletion strains generated a strain with the double deletion kin1Δ, kin2Δ, which was analyzed in a similar manner. The SNF1 gene was disrupted using a purified PCR product containing SNF1 sequences flanking the Kanamycin gene (snf1Δ::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; kin1Δ::LEU2/kin1Δ::LEU2; KIN2/kin2Δ::HIS3; leu2-3112/leu2-3112; his3-Δ200/his3-Δ200; ura3-52/ura3-52 to create the triple deletion mutant kin1Δ, kin2Δ, snf1Δ. 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 kin1Δ, kin2Δ, snf1Δ was confirmed by PCR.
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 × 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).
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- Δ200; 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 ).
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 β-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.
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 5× boiling buffer (5% SDS, 50 mM Tris, pH 8.0, 25 mM EDTA) and diluted 20× 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 α-Kin1, α-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 α-Kin1 or affinity-purified α-Sec9 antibodies. Samples were boiled and resolved on 7% SDS-PAGE, and a 35S signal was detected by autoradiography.
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 1× protease inhibitor mix). Lysates were centrifuged, combined with an equal volume of 2× 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 20× 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 2× restriction buffer 3 and 2 μl/tube of calf intestinal phosphatase (CIP) for 30 min at 37°C; 3) incubated with 2× buffer 3 only for 30 min at 37°C (mock control for CIP); 4) incubated 1:1 with 2× λ-phosphatase buffer and λ-phosphatase (1.5 μl/tube) for 30 min at 30°C; and 5) incubated with 2× λ-phosphatase buffer only for 30 min at 30°C (mock control for λ-phosphatase). Samples were boiled, separated on an 8% gel, and immunoblotted with α-Sec9 antibody.
All GST-fusion proteins were produced, purified, and their protein concentrations estimated as described previously (Rossi et al., 1997 ).
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, 1× 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.
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% β-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 1× PBS and then boiled immediately in 2× 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.
Rho3 is a member of the Rho family of GTPases that exhibits multiple genetic and physical interactions with components of the exocytic machinery and is itself required for efficient exocytosis (Adamo et al., 1999 ; 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 (rho3Δ). 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 rho3Δ 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 rho3Δ (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 rho3Δ, and thus we identified KIN1 as a novel dosage suppressor of Rho3.
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.
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.
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 sro7/77Δ, 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.
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-Δ42 mutant, with a deletion of the conserved 42 amino acid C-terminal tail (Figure 3A).
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-Δ42, 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-Δ42 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-Δ42 (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-Δ42, 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-CTΔ42 (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-CTΔ42 (regulatory domain with the deletion of the C-terminal 42 amino acids) and Kin2-Δ42 (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-CTΔ42 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-Δ42 (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-Δ42, 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-CTΔ42 (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.
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, kin1Δ and kin2Δ single disruptant strains were viable. The double disruptant kin1Δ, kin2Δ progeny, obtained by crossing of the kin1Δ strain with kin2Δ, demonstrate normal growth at all temperatures tested (25, 14, and 37°C). The single, kin1Δ and kin2Δ, and double kin1Δ, kin2Δ 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.
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, kin1Δ, kin2Δ, snf1Δ. kin1Δ, kin2Δ, snf1Δ was created by substitution of SNF1 with the KanR (Kanamycin) gene sequence in the context of the diploid strain homozygous for the KIN1 deletion (kin1Δ) and heterozygous for the KIN2 deletion (kin2Δ). A triple disruptant was obtained via sporulation and tetrad analysis. The triple mutant kin1Δ, kin2Δ, snf1Δ as well as the double kin1Δ, snf1Δ, and kin2Δ, snf1Δ mutants displayed slow growth phenotype inherent to the SNF1 deletion alone (Celenza and Carlson, 1984 ). The kin1Δ, kin2Δ, snf1Δ triple mutant did not display any synthetic defect in growth or secretion (as verified by invertase secretion assays; our unpublished data).
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 × 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.
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).
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 α-Kin2, α-Sso2, α-Snc1, α-Sro7, and α-Sec4 antibodies, respectively. This experiment revealed that the α-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.
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: λ-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).
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 [γ-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 (Figure7B,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 (Figure7B,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.
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 × 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.
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 nonpermissive temperature of 33°C. Suppression by GAL-KIN1 is lost when the cells are grown on noninducing YPD media. We examined the ability of galactose induced sec1-1 transformants to secrete BglII after a shift to restrictive temperature. As shown in Figure 9C, we find that control sec1-1 cells containing an empty GAL vector show an accumulation of internal BglII. In contrast cells containing GAL-KIN1 show a dramatic suppression of the secretory defect to levels of internal BglII found in wild-type yeast cells. Therefore, the same conditions of Kin1 overexpression that result in a reduction of Sec9 levels on the membrane also yield an overall “gain of function” in the secretory pathway.
Based on these data, we suggest that the positive effect of Kin1 and Kin2 on the secretory pathway may be mediated through regulation of the plasma membrane t-SNARE Sec9. Consistent with this assumption, overexpression of SEC9 suppresses growth defects of a number of late sec mutants that also are suppressed by introduction of multicopy KIN1 and KIN2. High copy SEC9 is known to restore the viability of rho3Δ, sec4-P48, sec1-1, sec3-2, sec8-9, sec9-4, and sec15-1 (Lehman et al., 1999 ). When we compared the relative suppression capabilities of high copy KIN1, KIN2, and SEC9, we found that KIN1 and KIN2 largely mimic the SEC9 suppression profile and exert an effect either equal or less potent in comparison with that of SEC9 (Table 4). The only mutant that KIN1 and KIN2, but not SEC9, are capable of suppressing is sec2-41, the significance of which remains to be addressed. Together, our results are consistent with Kin1 and Kin2 acting in the secretory pathway by positively regulating Sec9 function.
Par-1 is a key regulator of polarity-axis formation in a number of organisms. This includes the anterior-posterior axis of the C. elegans zygote and the Drosophila oocyte and the apico-basal axis of polarized epithelia in Drosophila and mammalian cells. Par-1 has been proposed to regulate cell fate determination and polarized cell morphology by positioning polarity markers, modulating activity of signaling pathways such as Wnt (canonical) and phencyclidine (planar cell polarity) and reorganizing the microtubule cytoskeleton (Kemphues et al., 1988 ; Drewes et al., 1997 ; Tenenhaus et al., 1998 ; Shulman et al., 2000 ; Cox et al., 2001 ; Huynh et al., 2001 ; Sun et al., 2001 ; Vaccari and Ephrussi, 2002 ; Benton and Johnston, 2003 ; Doerflinger et al., 2003 ; Cohen et al., 2004 ).
This is the first report linking Par-1 to regulation of exocytosis. Here, we show that the yeast Par-1 counterparts Kin1 and Kin2 exhibit multiple genetic and physical interactions with components of the exocytic machinery and thus function in the post-Golgi secretory pathway in S. cerevisiae. We demonstrate that both KIN1 and KIN2 suppress the growth abnormality of rho3Δ, rho3-V51, sec1, sec2, sec3, sec4, sec10, and sec15, mutants of genes important for polarized vesicle transport and docking. However, we observed that neither KIN1 nor KIN2 suppress sec9-4 and sro7Δ, sro77Δ, which are mutants of proteins involved in SNARE complex assembly and fusion. Thus, our genetic data position Kin1 and Kin2 function between the vesicle docking and fusion stages of exocytosis and identify these kinases as downstream effectors of the small GTPases: Rho3, Cdc42, Sec4, and several components of the vesicle tethering machinery. Moreover, these structurally homologous proteins display functional redundancy as well, because they show an identical pattern of suppression of the late secretory mutants. Consistent with genetic evidence, the analysis of physical associations of Kin1 and Kin2 show that these proteins exist in a complex with proteins necessary for fusion of exocytic vesicles with the plasma membrane. We demonstrate that Kin1 and Kin2 coimmunoprecipitate with the t-SNARE Sec9, and the homologue of the Drosophila tumor suppressor lethal giant larvae, Sro7, which has been previously reported to bind to Sec9 (Lehman et al., 1999 ). Based on physical and genetic data, we propose that Kin1 and Kin2 transmit a signal from the upstream-acting GTPases and the docking complex to the SNARE proteins to ensure correct vesicle fusion.
How conserved is the role of Par-1 in exocytosis? A high degree of functional conservation was observed for proteins constituting the Golgi-to-cell surface vesicle delivery machinery, including the Exocyst complex and the SNARE proteins (Kee et al., 1997 ; Rossi et al., 1997 ). Therefore, potentially, a link between Par-1 and the polarized exocytic machinery might represent a universal feature. Remarkably, a collaboration with Dr. Anne Muesch (Cornell Medical College, Ithaca, NY) revealed that mammalian Par-1 (EMK1) exists in a protein complex with the basolateral t-SNARE Syntaxin 4 and the Exocyst components in Madin-Darby canine kidney (MDCK) cells (our unpublished data, personal communication), suggesting that Par-1 is involved in the regulation of exocytosis in polarized mammalian cells as well. Nevertheless, there are some differences between the two systems. In MDCK cells, EMK1 interacts with the t-SNARE Syntaxin 4 but not with the mammalian equivalent of Sec9, SNAP25. On the other hand, our group did not detect any association between Kin1/Kin2 and Sso1/2, the yeast counterparts of the mammalian syntaxins. Thus, although Par-1 function in exocytosis is likely to be conserved, the precise molecular mechanism of signal transduction by Par-1 might differ from species to species.
Analysis of a kinase-dead mutant of Kin1 and Kin2 in yeast showed that the catalytic activity of these kinases is essential for suppression of sec mutants and thus for function in exocytosis. Therefore, we focused on finding a target and downstream effector of Kin1 and Kin2 in the secretory pathway. We assessed the ability of these kinases to phosphorylate a number of components of the late exocytic machinery, including the post-Golgi SNAREs. We found that the t-SNARE Sec9 is the only protein out of those tested that is phosphorylated in vivo upon overexpression of Kin1 and Kin2. Although we cannot exclude the possibility that another, as yet untested, component of the exocytic machinery is the target of Kin1/2 function in this pathway, the genetic and biochemical evidence provided here strongly suggests that Sec9 is the ultimate target of Kin1/2 action on exocytosis. Despite the fact that Sec9 can serve as a direct substrate of Kin1 and Kin2 in vitro, analysis of a phospho-mutant of Sec9 revealed that in vivo Kin1/Kin2-mediated phosphorylation of Sec9 is likely to be indirect. It is presently unclear which protein kinase transmits the signal from Kin1/Kin2 to Sec9.
How is the t-SNARE Sec9 regulated by yeast Par-1 orthologues? Recent studies demonstrated that SNARE protein stability, localization, and assembly into functional complexes are amenable to regulation by phosphorylation and dephosphorylation as is the final process of membrane fusion itself (Foster et al., 1998 ; Cabaniols et al., 1999 ; Peters et al., 1999 ; Risinger and Bennett, 1999 ; Chung et al., 2000 ; Kataoka et al., 2000 ; Lin and Scheller, 2000 ; Verona et al., 2000 ; Marash and Gerst, 2001 ; Pombo et al., 2001 ; Nagy et al., 2002 , 2004 ; Polgar et al., 2003 ; Tian et al., 2003 ). Here, we show that Kin1 and Kin2 alter the intracellular distribution of Sec9. At steady state, Sec9 is distributed approximately equally between the cytosol and the membrane. We observed that expression of Kin1 and Kin2 results in depletion of the membrane pool of Sec9 and induces phosphorylation of cytosolic Sec9. Thus, yeast Par-1 orthologues either release membrane-bound Sec9 or prevent its attachment to the membrane by yet unknown mechanism involving phosphorylation of the t-SNARE. Our data suggest that Kin1 and Kin2 transmit a signal to the secretory pathway by increasing the level and, possibly, activity of Sec9 in the cytosol. Consistent with this, overexpression of KIN1 and KIN2 produced the same effect as overexpression of SEC9 with respect to restoration of the growth of a number of sec mutants. The only mutant that we found to be rescued by KIN1 and KIN2, but not by SEC9 overexpression, is sec2, encoding an exchange factor for Sec4. It can be either due to Kin1/Kin2 having an additional function independent of regulation of Sec9, or alternatively, Sec9 might require Kin1/Kin2-induced phosphorylation to gain suppression of sec2.
The increase in the proportion of Sec9 in the cytosol induced by Kin1 and Kin2 suggests a mechanism by which their action promotes exocytosis. Previous work has suggested that Sec9 represents the limiting component in the formation of plasma membrane SNARE complexes and is present in wild-type cells in molar amounts that are 5- to 10-fold lower than the other two SNAREs it forms complexes with: Sso1/2 and Snc1/2 (Brennwald et al., 1994 ). Therefore increasing amounts of Sec9 available for formation of SNARE complexes either by increasing overall Sec9 protein levels (i.e., increasing SEC9 dosage) or increasing the proportion of free Sec9 available to form SNARE complexes would be expected to have strong positive effect on exocytic activity. However, it is known that SNARE complex assembly occurs on the plasma membrane. How does the Kin1/2-mediated release of Sec9 from the membrane into the cytosol promote exocytosis?
It is likely that Kin1 and Kin2 phosphorylate and release from the membrane free, unassembled Sec9 for two reasons. First, we have not detected the interaction between Kin1/Kin2 and two other exocytic SNAREs: the t-SNARE Sso and the v-SNARE Snc. Second, we have not detected a dramatic difference in the amount of Sec9 present in SNARE complexes as a consequence of Kin1 or Kin2 activity (our unpublished observation), and thus this pathway does not affect steady-state amounts of SNARE complexes present. Nevertheless, it cannot be ruled out that Kin1 and Kin2 phosphorylate a fraction of Sec9 in “old”, inactive SNARE complexes and initiate its release, priming Sec9 for a new round of vesicle fusion.
Importantly, although it may seem contradictory that elevation of unassembled SNARE levels in the cytosol at the expense of the membrane-bound pool promotes vesicle fusion, we hypothesize that Sec9 is recruited from the cytosolic rather than the membrane pool into new SNARE complexes. Under this assumption, the more efficient release of Sec9 from the membrane induced by Kin1 and Kin2 phosphorylation replenishes the pool of assembly-competent Sec9 near sites of recruitment. Consequently, increased cycling of Sec9 would promote complex assembly and vesicle fusion. Thus, we think that Kin1 and Kin2 promote secretion by increasing the pool of Sec9 available for incorporation into new, active SNARE complexes.
It is also possible that Kin1 and Kin2 help to spatially direct SNARE complex formation by locally priming Sec9 for assembly. In S. cerevisiae fusion of vesicles is restricted to the tip of the bud. Nevertheless, the t-SNARE Sec9 is not polarized, in fact, it localizes ubiquitously in the cytoplasm and homogenously over the entire perimeter of the plasma membrane (Brennwald et al., 1994 ). Distribution of the other t-SNARE, Sso, is also nonpolar (Brennwald et al., 1994 ). Therefore, other molecules are required to restrict formation of fusion-competent SNARE complexes to the bud tip. Exogenous Kin1 and Kin2 localize ubiquitously in the cell (our unpublished observation). However, based on our genetic data, these kinases act downstream of molecules directing polarized vesicle transport and docking, such as Rho3, Cdc42, Sec4, and the Exocyst complex. Thus, Kin1 and Kin2 are likely to be activated and regulate Sec9 in a polarized manner, possibly by phosphorylating and releasing Sec9 from the membrane locally, near the sites of fusion.
The C-terminal 42 amino acid tail sequence (termed “KA1” domain) is highly conserved throughout Par-1 orthologues from yeast to mammalian cells. Nevertheless, the role of this conserved segment remained unknown. Our data show that deletion of these 42 amino acids results in the gain of function of the kinase in the secretory pathway. Kin2 constructs with a deletion of the tail display stronger suppression of the growth defect of sec mutants relative to the wild-type kinase, suggesting that the tail plays an inhibitory role. Binding of the regulatory domain of the kinase to its catalytic core is a known mechanism of kinase autoinhibition (Hu et al., 1994 ; Tu et al., 1997 ; Tu and Wigler, 1999 ; Tan et al., 2001 ). Inhibition can happen via two scenarios: in cis, with one molecule circling on itself (as shown in Figure 3D), and in trans, mediated through head-to-tail dimer formation. In a “closed” configuration an inhibitory segment inactivates a kinase by blocking either the ATP- or the substrate-binding sites. Consistent with this mechanism of autoinhibition, we show that catalytic (N-terminal) and regulatory (C-terminal) domains of Kin2 directly interact with each other and that this interaction requires an intact 42 amino acid tail sequence. Hence, we provide functional and physical evidence for the inhibitory role of the C-terminal tail. Considering a remarkable degree of conservation of the tail sequence between Par-1 orthologues, it is likely that the autoinhibitory function of the tail is conserved as well.
In summary, we report three novel findings. First, we demonstrate that yeast Par-1 counterparts are associated with and seem to positively regulate the function of the exocytic apparatus. Second, we show that Kin1 and Kin2 induce phosphorylation of Sec9 and its release from the plasma membrane to the cytosol, promoting its recycling and/or availability for incorporation into newly formed SNARE complexes. Third, we find that the conserved 42-amino acid tail of the yeast Par-1 orthologues plays a role in autoinhibition.
This suggests an interesting possibility that the role of Par-1 in cell polarity establishment in a variety of organisms might be tied to the control of Golgi-to-plasma membrane vesicle targeting. It is possible to envision that the primary function of Par-1 is to regulate polarized secretion, which in turn is necessary for proper localization of polarity determinants and microtubule capture/stabilization, processes that require Par-1 in a number of species. It remains to be seen how essential and conserved is the function of Par-1 in exocytosis.
We thank Drs. Anne Muesch, Enrique Rodriguez-Boulan, and Guido Wendel for invaluable advice and critical comments on the manuscript. This work was supported by The National Institutes of Health grant GM-54712 (to P. B.) and The Tri-Institutional Training Program in Vision Research grant T32EY007138 (to M. E.).
Article published online ahead of print in MBC in Press on December 1, 2004 (http://www.molbiolcell.org/cgi/doi/10.1091/mbc.E04-07-0549).