Lpx1p links glucose-induced calcium signaling and plasma membrane H+-ATPase activation in Saccharomyces cerevisiae cells

Lpx1p links glucose-induced calcium signaling and plasma membrane H+-ATPase activation in... Abstract In yeast, as in other eukaryotes, calcium plays an essential role in signaling transduction to regulate different processes. Many pieces of evidence suggest that glucose-induced activation of plasma membrane H+-ATPase, essential for yeast physiology, is related to calcium signaling. Until now, no protein that could be regulated by calcium in this context has been identified. Lpx1p, a serine-protease that is also involved in the glucose-induced activation of the plasma membrane H+-ATPase, could be a candidate to respond to intracellular calcium signaling involved in this process. In this work, by using different approaches, we obtained many pieces of evidence suggesting that the requirement of calcium signaling for activation of the plasma membrane H+-ATPase is due to its requirement for activation of Lpx1p. According to the current model, activation of Lpx1p would cause hydrolysis of an acetylated tubulin that maintains the plasma membrane H+-ATPase in an inactive state. Therefore, after its activation, Lpx1p would hydrolyze the acetylated tubulin making the plasma membrane H+-ATPase accessible for phosphorylation by at least one protein kinase. calcium signaling, plasma membrane H+-ATPase, Lpx1, Saccharomyces cerevisiae INTRODUCTION Plasma membrane H+-ATPase is an essential enzyme for yeast physiology since by pumping protons out of cells it creates the electrochemical potential that allows cells to accumulate nutrients against their concentration gradients and to control intracellular pH (Goffeau and Slayman 1981). This enzyme is regulated both at the transcriptional (Capieaux et al.1989; Rao, Drummond-Barbosa and Slayman 1993) and at the post-translational (Serrano 1993) levels on glucose addition, this sugar being the most important trigger for its regulation. Originally, it was proposed that H+-ATPase post-translational activation would be caused by phosphorylation catalyzed by protein kinases that would have as targets two phosphorylation sites at the plasma membrane H+-ATPase C-terminal tail (Portillo 2000). According to this model, phosphorylation of residue Ser-899 would be responsible for a decrease in Km for ATP, and phosphorylation of Thr-912 would lead to an increase of Vmax related to ATP hydrolysis (Portillo, Eraso and Serrano 1991). Later, it was demonstrated that the second phosphorylation site includes another serine residue (Ser-911) that would also be associated with glucose addition (Lecchi et al.2005, 2007). Moreover, it was confirmed that phosphorylation of both residues (Ser-911 and Thr-912) seems to be connected to the activation process (Mazón, Eraso and Portillo 2015). Although it has been shown that glucose-induced activation of the plasma membrane H+-ATPase is caused by phosphorylation, the search for protein kinases involved in this process led, up to now, to the identification of Ptk2p as a unique protein kinase involved with the phosphorylation of the Ser-899 residue at the C-terminal tail (Goossens et al.2000; Eraso, Mazón and Portillo 2006). Recently, a wide screening of protein kinases present in yeast cells reconfirmed the essential involvement of Ptk2p in the glucose-induced activation of the plasma membrane H+-ATPase. Thus, these data indicate that the current model suggesting the existence of two phosphorylation sites (and another protein kinase indeed involved in this process, besides Ptk2p) seems to be incorrect or, at least, incomplete (Pereira et al.2015). Indeed, new elements have been introduced in this scenario confirming the complexity of the mechanism responsible for plasma membrane H+-ATPase post-translational activation. Over the years, many results have suggested the existence of a clear relationship between calcium signaling and plasma membrane H+-ATPase activation (Coccetti et al.1998; Tisi et al.2002, 2004; Trópia et al.2006; Pereira et al.2008; Groppi et al.2011; Bouillet et al.2012). Saccharomyces cerevisiae as a eukaryotic organism can use calcium-signaling pathways to control different cellular processes (Anraku, Ohya and Iida 1991; Paidhungat and Garrett 1997; Denis and Cyert 2002; Serrano et al.2002). In the cytosol of yeast cells, the free Ca2+ concentration is strictly regulated (Miseta et al.1999b) by the action of a variety of transporters, channels, pumps and co-transporters (Cyert and Philpott 2013). Ca2+ in yeast is retained mainly in the vacuole (Dunn, Gable and Beeler 1994; Miseta et al.1999a) and the vacuolar pool is maintained mostly by the action of Pmc1p (a Ca2+-ATPase) and Vcx1p (a Ca2+/H+ exchanger). The release of calcium stored in vacuoles is made through the calcium channel Yvc1p, which has its activity controlled by different cellular stimuli (Cunningham 2011). Apparently, the connection between calcium signaling and H+-ATPase activation starts when glucose-induced phosphatidylinositol-4,5-bisphosphate hydrolysis occurs. Mediated by phospholipase C (encoded by the PLC1 gene), this hydrolysis produces two intracellular messengers: diacylglycerol and inositol 1,4,5-trisphosphate (IP3) (York et al.1999; Shears 2000). This Plc1p activation would be initiated in response to a stimulus from protein G (Gpa2p), which in turn would be activated in response to glucose uptake and its phosphorylation by sugar kinases (Bouillet et al.2012). In turn, the level of IP3 is regulated by its phosphorylation by Arg82p, an inositol kinase, generating two types of inositol tetrakisphosphate—I(1,3,4,5)P4 and I(1,4,5,6)P4—and an inositol pentaphosphate, I(1,3,4,5,6)P5 (Saiardi et al.1999; Odom et al.2000). A possible relationship between IP3 and the vacuolar Ca2+ channel, Yvc1p, was demonstrated previously, suggesting that this channel could participate in a mechanism involved in intracellular calcium level control in response to glucose addition. IP3 would interact directly or indirectly with Yvc1p, regulating the intensity of calcium signaling in the cytosol (Bouillet et al.2012). In arg82Δ yeast strains, where there is no conversion of IP3 into IP4 and/or IP5, glucose-induced increase of IP3, H+-ATPase activation and calcium signaling are more pronounced (Tisi et al.2004). At low external calcium concentration, the vacuolar calcium channel Yvc1p’s activity seems to become more important for proper glucose-induced calcium signaling and plasma membrane H+-ATPase activation. Moreover, there are many pieces of evidence suggesting that the intensity of the intracellular calcium signal, in these conditions, is strongly dependent on the action of Yvc1p (Bouillet et al.2012) and that there is a connection between the IP3 signal, Yvc1 activity, calcium signaling and glucose-induced activation of the plasma membrane H+-ATPase in yeast cells (Tisi et al.2004; Trópia et al.2006). Nevertheless, since no IP3 receptor homolog has been identified in the S. cerevisiae genome (Wera, Bergsma and Thevelein 2001), the existence of such a transduction pathway is still a matter of controversy (Bouillet et al.2012). On the other hand, some data suggest the involvement of a multifunctional enzyme in glucose-induced activation of the plasma membrane H+-ATPase (Campetelli et al.2013). By this model, glucose addition would trigger activation of proteolytic activity of Lpx1p, which degrades the tubulin bound to the inactive form of plasma membrane H+-ATPase at its C-terminal tail. This degradation releases the C-terminal tail leading to the exposure of sites that are targets of phosphorylation, causing H+-ATPase activation (Campetelli et al.2005, 2013). Considering these two sets of evidence for H+-ATPase activation, calcium involvement and Lpx1p dependence, and the apparent difficulty in finding other protein kinase(s), mainly the calcium-dependent ones, involved in enzyme phosphorylation/activation, we explored a possible relationship between calcium signaling and Lpx1p activation in connection with glucose-induced regulation of the plasma membrane H+-ATPase. Here, we demonstrated that calcium signaling is indeed directly related to Lpx1p proteolytic action, and this activation process is clearly related to the plasma membrane H+-ATPase. MATERIAL AND METHODS Strains and growth conditions Saccharomyces cerevisiae strains shown in Table 1 were grown in media containing 2% peptone and 1% yeast extract (YP) supplemented with appropriate carbon sources or in SD medium—0.67% yeast nitrogen base without amino acids (Difco, Detroit, MI, USA) supplemented with drop-out amino-acids (leucine, tryptophan, histidine and methionine) and nucleotide (uracil) and 2% glucose. Cells were grown in a rotatory incubator shaker - New Brunswick Model 25 (GMI, Ramsey, MN, USA) at 200 rpm and 30°C until the end of the logarithmic phase (OD600nm ∼ 2.0). Alternatively, and when requested, yeast cells were grown in induction medium (SD medium supplemented with 8% galactose). Table 1. Saccharomyces cerevisiae strains used in this study. Strain  Genotype  Source  PJ69-2A  MATa trp1-901 leu2-3, 112 ura3-52 his3-200 gal4Δ gal80Δ LYS2::GAL1-HIS3 GAL2-ADE2 met2::GAL7-lacZ  James Caffrey (Saiardi et al.2000)  PJ69-4A1  PJ69-2A arg82::KanMX2  James Caffrey (Saiardi et al.2000)  LBCM 1630  PJ69-2A yvc1::KanMX2  Bouillet et al. (2012)  LBCM 1713  LBCM 1630; lpx1::LEU2  This work  LBCM 1714  PJ69-4A1; lpx1::LEU2  This work  BY4741  MATa his3Δ1 leu2Δ0 lis2Δ0 ura3Δ0  EUROSCARF  BY4741 lpx1Δ  BY4741; lpx1::KanMX4  EUROSCARF  BY4741 ptk2Δ  BY4741; ptk2::KanMX4  EUROSCARF  LBCM 1738  BY4741 lpx1Δ + pYES2/CT-EV  This work  LBCM 1739  BY4741 lpx1Δ + pYES2/CT-LPX1  This work  LBCM 1764  BY4741 lpx1Δ + pYES2/CT-LPX1-MOD  This work  Strain  Genotype  Source  PJ69-2A  MATa trp1-901 leu2-3, 112 ura3-52 his3-200 gal4Δ gal80Δ LYS2::GAL1-HIS3 GAL2-ADE2 met2::GAL7-lacZ  James Caffrey (Saiardi et al.2000)  PJ69-4A1  PJ69-2A arg82::KanMX2  James Caffrey (Saiardi et al.2000)  LBCM 1630  PJ69-2A yvc1::KanMX2  Bouillet et al. (2012)  LBCM 1713  LBCM 1630; lpx1::LEU2  This work  LBCM 1714  PJ69-4A1; lpx1::LEU2  This work  BY4741  MATa his3Δ1 leu2Δ0 lis2Δ0 ura3Δ0  EUROSCARF  BY4741 lpx1Δ  BY4741; lpx1::KanMX4  EUROSCARF  BY4741 ptk2Δ  BY4741; ptk2::KanMX4  EUROSCARF  LBCM 1738  BY4741 lpx1Δ + pYES2/CT-EV  This work  LBCM 1739  BY4741 lpx1Δ + pYES2/CT-LPX1  This work  LBCM 1764  BY4741 lpx1Δ + pYES2/CT-LPX1-MOD  This work  View Large Table 1. Saccharomyces cerevisiae strains used in this study. Strain  Genotype  Source  PJ69-2A  MATa trp1-901 leu2-3, 112 ura3-52 his3-200 gal4Δ gal80Δ LYS2::GAL1-HIS3 GAL2-ADE2 met2::GAL7-lacZ  James Caffrey (Saiardi et al.2000)  PJ69-4A1  PJ69-2A arg82::KanMX2  James Caffrey (Saiardi et al.2000)  LBCM 1630  PJ69-2A yvc1::KanMX2  Bouillet et al. (2012)  LBCM 1713  LBCM 1630; lpx1::LEU2  This work  LBCM 1714  PJ69-4A1; lpx1::LEU2  This work  BY4741  MATa his3Δ1 leu2Δ0 lis2Δ0 ura3Δ0  EUROSCARF  BY4741 lpx1Δ  BY4741; lpx1::KanMX4  EUROSCARF  BY4741 ptk2Δ  BY4741; ptk2::KanMX4  EUROSCARF  LBCM 1738  BY4741 lpx1Δ + pYES2/CT-EV  This work  LBCM 1739  BY4741 lpx1Δ + pYES2/CT-LPX1  This work  LBCM 1764  BY4741 lpx1Δ + pYES2/CT-LPX1-MOD  This work  Strain  Genotype  Source  PJ69-2A  MATa trp1-901 leu2-3, 112 ura3-52 his3-200 gal4Δ gal80Δ LYS2::GAL1-HIS3 GAL2-ADE2 met2::GAL7-lacZ  James Caffrey (Saiardi et al.2000)  PJ69-4A1  PJ69-2A arg82::KanMX2  James Caffrey (Saiardi et al.2000)  LBCM 1630  PJ69-2A yvc1::KanMX2  Bouillet et al. (2012)  LBCM 1713  LBCM 1630; lpx1::LEU2  This work  LBCM 1714  PJ69-4A1; lpx1::LEU2  This work  BY4741  MATa his3Δ1 leu2Δ0 lis2Δ0 ura3Δ0  EUROSCARF  BY4741 lpx1Δ  BY4741; lpx1::KanMX4  EUROSCARF  BY4741 ptk2Δ  BY4741; ptk2::KanMX4  EUROSCARF  LBCM 1738  BY4741 lpx1Δ + pYES2/CT-EV  This work  LBCM 1739  BY4741 lpx1Δ + pYES2/CT-LPX1  This work  LBCM 1764  BY4741 lpx1Δ + pYES2/CT-LPX1-MOD  This work  View Large Molecular biology methods and in silico analysis Yeast cells were transformed by using the lithium acetate protocol (Gietz et al.1995). A pVTU-apoaequorin plasmid (pVTU-AEQ) was generated by inserting in pVTU XhoI–PstI sites the XhoI–PstI-digested fragment obtained by PCR on pYX212-AEQ (Tisi et al.2002) using the oligonucleotides 5΄-TTTCTCGAGAATCTATAACTACAAAAAACACATACAGGAA-3΄ and 5΄-TAACTGCAGGCCCTAGGATCCATGGTGAA-3΄. The LPX1 gene was obtained as previously described (Campetelli et al.2013) and then inserted into the pYES2/CT plasmid (Invitrogen, Carlsbad, CA, USA) allowing inducible expression by galactose of recombinant proteins fused with a 6× His-tag at the C-terminus. For this purpose, plasmid and amplified LPX1 gene were previously digested with KpnI and XbaI enzymes (see Supplementary Material 1) (Fermentas, Waltham, MA, USA). Digested plasmid was treated with shrimp alkaline phosphatase (SAP) (Fermentas) for 1 h at 37°C prior to ligation. The SAP-treated plasmid was used for the T4 ligase reaction (Sigma-Aldrich, St Louis, MO, USA) with LPX1 insert. This ligation reaction was performed at 23°C for 1 h, and Escherichia coli strain TOP10 was used for plasmid replication. BY4741 lpx1Δ cells were transformed with pYES2/CT + LPX1 (Gietz et al.1995) and selected in SD medium with 2.5% agar—from here on denominated as pYES2/CT-LPX1. The presence of the gene insertion into the plasmid and correct transformation in bacteria and yeast were confirmed by PCR (forward primer: 5΄-AAAGGTACCATGGAACAGAACAGGTCC-3΄; reverse primer: 5΄-TTTTCTAGATTACAGTTTTTGTTTAGTCG-3΄). BY4741 lpx1Δ cells were transformed with an empty vector (here denominated as pYES2/CT-EV) as well, to be used as the control treatment. In order to find regions in Lpx1p protein structure that could interact with Ca2+, the WebFEATURE 4.0 server (Liang et al.2003; Wu, Liang and Altman 2008) was run using a deposited structure (crystal form II; trigonal) of Lpx1p from S. cerevisiae (PDB access number: 2y6u) (Thoms et al.2011) with settings defined to search for all functional sites possibly present in this protein (Supplementary Material 2). To verify the influence of the C-terminal region of Lpx1p that would interact with Ca2+ in Lpx1p activation, this region was removed by treating the LPX1 gene with PvuII enzyme that cuts at 909 bp after the initiation codon. The modified gene was cloned into the pYES2/CT plasmid (see Supplementary Material 3). Digested LPX1 gene and pYES2/CT plasmid were then treated with KpnI. Digested plasmid was used for the T4 ligase reaction (Sigma-Aldrich) with modified LPX1 insert. This ligation reaction was performed at 23°C overnight and E. coli strain TOP10 was used for plasmid replication. BY4741 lpx1Δ cells were transformed with pYES2/CT + modified LPX1 (Gietz et al.1995) and selected in SD medium with 2.5% agar—from here on denominated as pYES2/CT-LPX1-MOD. The presence of the gene insertion into the plasmid and correct transformation in bacteria and yeast were confirmed by PCR (forward primer: 5΄-AAAGGTACCATGGAACAGAACAGGTCC-3΄; reverse primer: 5΄-GCGTGAATGTAAGCGTGAC-3΄), by comparing different amplicon sizes between pYES2/CT-LPX1 and pYES2/CT-LPX1-MOD. Measurement of cytosolic free calcium by bioluminescence assay By using a standard method, cytosolic free calcium was measured by using the aequorin-based bioluminescence assay (Tisi et al.2002; Brandão 2014; Tisi, Martegani and Brandão 2015). Yeast strains containing the apoaequorin-expressing plasmid pVTU-AEQ were grown in rich medium (YP with 2% glucose) until they entered the exponential phase. Cells were harvested and washed by filtration with sterile water and resuspended in MES/Tris 0.1 M (pH 6.5). After 90 min of incubation at room temperature, cells were loaded with 50 μM coelenterazine (Sigma-Aldrich, St. Louis, MO, USA) for 20 min. For removal of excess coelenterazine, cells were washed twice by centrifugation (2000 g, 5 min) with MES/Tris 0.1 M pH 6.5. Glucose-induced aequorin luminescence was measured in a Lumat LB 9507 luminometer (Berthold Technologies, Bad Wildbad, Germany) at intervals of 10 s for 1 min before and for at least 10 min after addition of 100 mM glucose (final concentration). Lpx1p His-tag expression and purification To obtain cells for Lpx1p His-tag extraction, the lpx1Δ mutant transformed with pYES2/CT-LPX1, and pYES2/CT-LPX1-MOD plasmids were grown in SD medium for 24 h. Cells were collected, washed three times by centrifugation (2000 g, 5 min) with sterile water and resuspended in induction medium (SD medium supplemented with 8% galactose). After overnight incubation in a rotatory incubator shaker (New Brunswick Model 25) at 200 rpm and 30°C, cells were harvested by vacuum filtration on glass fiber filters, immediately frozen in liquid nitrogen and stored until use. Cells were resuspended in lysis buffer (50 mM Na3PO4, 300 mM NaCl and 5 mM β-mercaptoethanol), disrupted by vigorous shaking with glass beads (with 90 s intervals on ice for five times) and then centrifuged at 2000 g for 10 min. The resulting supernatant (crude extract) was incubated with HIS-Select nickel affinity gel (Sigma-Aldrich, St. Louis, MO, USA) for 1 h at 4°C with gentle agitation. Unbound proteins were removed by discarding supernatant after centrifugation at 1000 g for 5 min. Proteins bound to resin were eluted with elution buffer (50 mM Na3PO4, 300 mM NaCl, 5 mM β-mercaptoethanol). Induction and protein purification were performed with pYES2/CT-EV strain, to be used as the control treatment. Aliquots (50 μg) of protein fractions were separated by SDS-PAGE using 12% (w/v) polyacrylamide gels. Gels were stained with silver staining and images were captured by Image Scanner software (Thermo Fisher Scientific, Waltham, MA, USA). The molecular masses of the proteins were calculated by interpolating their specific relative migration distances (Rf) in a curve prepared with correspondent values of appropriated molecular mass markers (Supplementary Material 4). Western blotting assays were performed with eluates obtained from cell extracts of the lpx1Δ mutant transformed with pYES2/CT-EV, pYES2/CT-LPX1 and pYES2/CT-LPX1-MOD constructions. Protein content was determined by using the Lowry method (Lowry et al.1951). Next, the proteins were transferred to nitrocellulose membranes (Thermo Fisher Scientific, Waltham, MA, USA) according to the recommendations of the Bio-Rad Transference Kit (Bio-Rad Laboratories, Hercules, CA, USA). Membranes were blocked for 1 h at room temperature, using PBS containing 0.1% Tween-20 and 5% skimmed milk powder. Furthermore, membranes were incubated with mouse monoclonal primary antibody anti-His (Invitrogen, Carlsbad, CA, USA). Subsequently, membranes were washed and incubated with secondary antibody horseradish peroxidase-goat anti-mouse IgG (H + L) (Invitrogen, Carlsbad, CA, USA). Immunolabeling was visualized using WESTAR Nova 2.0 (Cyanagen, Bologna, Italy) as per the manufacturer’s instructions. Images were captured by Image Scanner software (Amershan Biosciences, Bath, UK). Measurement of proteolytic Lpx1p activity Calcium dependence of the proteolytic activity of the wild-type and lpx1Δ cells was assayed by using azocasein (Thermo Fisher Scientific, Waltham, MA, USA) as substrate (Charney and Tomarelli 1947). Briefly, samples of total membranes containing 50–100 μg protein were added to a reaction mixture (100 mM Tris–HCl, pH 8.0, 0.5% azocasein), with or without the addition of 1 mM calcium, in a final volume of 500 μL. The mixture was kept for 1 h at 40°C, and after this time, the reaction was stopped by adding 250 μL of 10% w/v trichloroacetic acid. The mixture was centrifuged (2000 g, 5 min) to remove coagulated proteins and 500 μL of supernatant was neutralized with 500 μL 5 M KOH, and the absorbance at 428 nm was measured using a BioMate 3 spectrophotometer (Thermo Fisher Scientific, Waltham, MA, USA). A change in absorbance of 0.01 (428 nm) per minute, resulting from azocasein hydrolysis, was defined as one arbitrary unit (AU). The control was obtained by mixing trichloroacetic acid with the azocasein solution prior to the addition of the protein sample (Secades and Guijarro 1999). Protein content was determined by using the Lowry method (Lowry et al.1951). An additional Lpx1p activity evaluation was performed with eluates obtained from the Ni-affinity column from lpx1Δ mutant transformed with pYES2/CT-LPX1. Samples containing 100 μg protein were dissolved in non-reducing Laemmli buffer (62.5 mM Tris–HCl, pH 6.8, 10% w/v glycerol, 0.001% w/v bromophenol blue) and run at 100 V at 4°C in 8% (w/v) polyacrylamide gel. After electrophoresis, the corresponding Lpx1p bands were excised from the gel and washed three times in 100 mM Tris–HCl buffer, pH 8.0, with gentle agitation, in order to remove excess SDS from the running buffer. Then, they were added to a reaction mixture (100 mM Tris–HCl, pH 8.0, 0.5% azocasein), containing 100 μM calcium, in a final volume of 500 μL and the assay was performed as described previously in this work. A negative control was obtained by using the corresponding material from lpx1Δ mutant transformed with pYES2/CT-EV. Measurement of plasma membrane H+-ATPase activity In vivo activation of plasma membrane H+-ATPase was detected by using two different standard approaches: indirectly, measuring glucose-induced proton-pumping activity, or directly, in purified plasma membranes by following ATP hydrolysis. In the first case, around 500 mg (wet weight) of cells grown in YP medium (with 2% glucose) was resuspended in 100 mM Tris–HCl buffer pH 4.5 containing 100 mM KCl and incubated in a vessel in a total volume of 5.0 mL. Changes in pH suspension were recorded before and after addition of 100 mM glucose (final concentration). Calibration pulses of 100 nmol HCl were also added and used as reference in calculations of proton pumping. The maximal rate of proton pumping was calculated from the slope of the line indicating the pH variation in the medium, as described previously, and indicated in mmol H+ h−1 g−1 cell (Souza, Tropia and Brandao 2001). To measure proton-pumping activity from lpx1Δ or yvc1Δ mutants transformed with pYES2/CT-EV and pYES2/CT-LPX1 vectors, cells grown in SD medium were washed twice by centrifugation (2000 g, 5 min) with 100 mM Tris–HCl buffer pH 4.5 containing 100 mM KCl. After washing, cells were incubated in the same buffer containing 100 mM galactose (final concentration) prior to proceeding with the assay to promote the induction of the LPX1 gene. After incubation with 100 mM galactose for different times (10, 30 or 50 min), cells were washed twice by centrifugation (2000 g, 5 min) with 100 mM Tris–HCl buffer pH 4.5 containing 100 mM KCl, and then used to measure in vivo proton-pumping activity as described (Souza, Tropia and Brandao 2001). Cells without incubation with 100 mM galactose were used as controls. In the second approach, to measure directly the H+-ATPase activity, lpx1Δ mutant transformed with pYES2/CT-EV and pYES2/CT-LPX1 vectors was collected after overnight incubation in induction medium (SD medium supplemented with 8% galactose) in a rotatory incubator shaker (New Brunswick Model 25) at 200 rpm and 30°C, prior to proceeding with plasma membrane purification. Cells were washed twice by centrifugation (2000 g, 5 min) with 100 mM MES/Tris buffer (pH 6.5), collected on glass fiber filters by vacuum filtration, immediately frozen in liquid nitrogen and stored until use. Cells were disrupted in 100 mM Tris buffer (supplemented with 330 mM sorbitol and 5 mM β-mercaptoethanol), by vigorous shaking with glass beads, with 90 s intervals on ice five times and centrifuged at 2000 g for 10 min. Purified plasma membranes were obtained essentially as described before (dos Passos et al.1992), with resuspension of plasma membranes in glycerol buffer (20% w/v glycerol, 10 mM Tris, 1 mM β-mercaptoethanol). Protein content was determined by using the Lowry method (Lowry et al.1951). ATPase activity was determined as described previously (dos Passos et al.1992). On the other hand, and to study a direct connection between calcium availability, Lpx1p activity and plasma membrane H+-ATPase regulation, we tried to reconstitute in vitro the plasma membrane H+-ATPase activation. First, to extract plasma membranes, wild-type cells were resuspended in 100 mM MES/Tris buffer (pH 6.5) at a density of 150 mg mL−1 (wet mass) and incubated in a shaking water bath at 30°C. After 4 h, samples (glucose-starved cells) were collected on glass fiber filters by vacuum filtration and immediately frozen in liquid nitrogen and stored until use. The procedures used to obtain plasma membranes were performed as described previously (dos Passos et al.1992), without addition of EDTA in lysis and glycerol buffers. Briefly, to obtain a membrane-free extract (MFE), cells were suspended in 100 mM Tris buffer without protease inhibitors (supplemented with 330 mM sorbitol and 5 mM β-mercaptoethanol), disrupted by vigorous shaking with glass beads with 90 s intervals on ice five times, and centrifuged at 2000 g for 10 min. The supernatant was then centrifuged at 100 000 g for 30 min. After ultracentrifugation, a new supernatant yielded membrane-free extracts (from here on denominated as MFE) from all strains; these as well as HIS-Select nickel affinity gel eluates resulting from the application of crude extract of lpx1Δ mutant cells transformed with pYES2/CT-EV, pYES2/CT-LPX1 and pYES2/CT-LPX1-MOD were subsequently submitted to overnight dialysis at 4°C with incubation buffer (50 mM HEPES, 10 mM sodium azide, 5 mM ammonium molybdate). This procedure was necessary in order to remove any interfering compound for in vitro H+-ATPase activity assay. Protein content was determined by using the Lowry method (Lowry et al.1951). In vitro H+-ATPase activation was performed by resuspending plasma membrane samples (40 μg protein) obtained from glucose-starved wild-type cells in an incubation buffer (50 mM HEPES, 10 mM sodium azide, 5 mM ammonium molybdate). Then, 100 μM Ca2+ and/or MFE (50 μg protein) was added (or not) according to the different tests performed, to a final volume of 250 μL (more details in Results section). In all samples, 1 mM MgCl2 (final concentration) was also added. The samples were incubated for 30 min at 30°C in a water bath, for in vitro H+-ATPase activation. Different calcium concentrations (100 μM, 1 and 5 mM), different times of incubation (10, 30 and 60 min) and different MFE protein concentrations (50, 100, 200 and 400 μg) for in vitro H+-ATPase activation were tested. Sodium orthovanadate of 50 μM was added as well to verify H+-ATPase activity inhibition. MFE and HIS-Select nickel affinity gel eluates were assayed to check for any ATPase activity as control. All results are expressed as percentages relative to their related controls. To assay H+-ATPase activity, the reactions were started with concentrated ATP solution (Sigma-Aldrich) to obtain 2 mM as final concentration. ATPase activity was determined as described previously (dos Passos et al.1992) and appropriate controls were performed for each assay (see Results section for specific details). Statistical analysis The experiments were performed at least three times. Standard deviations were indicated in each figure or table. Statistical analyses were performed by using Student's t-test or ANOVA. Differences were considered statistically significant when the P value was <0.05. RESULTS Lpx1p influences H+-ATPase activation, but not intracellular calcium concentration It was previously suggested that in glucose-induced activation of yeast plasma membrane H+-ATPase, calcium signaling is a critical factor (Souza, Tropia and Brandao 2001; Tisi et al.2004; Trópia et al.2006; Pereira et al.2008; Bouillet et al.2012). However, in the corresponding signal transduction pathway a component that actually would respond to the intracellular calcium signal was never found. Since the Lpx1p protein also participates in this process (Campetelli et al.2013), we wondered whether this protease could be a possible candidate that would respond to the calcium signal, thereby explaining its function in plasma membrane H+-ATPase glucose-induced activation. Indeed, as demonstrated in Fig. 1A, plasma membrane H+-ATPase glucose-induced activation (measured indirectly by proton-pumping rate) was reduced in strains presenting deletion in the LPX1 gene. Comparable results (i.e. reduced H+-ATPase glucose-induced activation) were also observed in yvc1Δ strain (absence of Yvc1p reduces glucose-induced calcium signaling by preventing calcium release from the vacuole, which reduces H+-ATPase activation). On the other hand, deletion of ARG82 causes a clear stronger glucose-induced activation of the plasma membrane H+-ATPase compared with wild-type cells, since these cells present high IP3 levels in cytoplasm, which enhances calcium release from vacuole leading to an increase in the H+-ATPase activity. However, when LPX1 deletion was also included in this arg82Δ mutant, H+-ATPase activity was reduced as observed in the cell presenting only the lpx1Δ mutation. The same standard was also observed in a strain presenting deletions in the YVC1 and LPX1 genes, i.e. H+-ATPase activity showed comparable low levels to yvc1Δ cells. The same was observed in the double mutant yvc1Δ lpx1Δ (Fig. 1A). Figure 1. View largeDownload slide Effects of different gene deletions on H+-ATPase proton-pumping activity and intracellular calcium signaling in S. cerevisiae cells (genetic background PJ69-2A). (A) Proton-pumping activity was measured with around 500 mg of cells, after addition of 100 mM glucose (final concentration). (B) Cytosolic free calcium was measured by using a standard aequorin-based bioluminescence assay after addition of 100 mM glucose (final concentration), indicated by a striped arrow at ‘0’ s, in yeast strains containing apoaequorin-expressing plasmid pVTU-AEQ and loaded with 50 μM coelenterazine. Different letters indicate that mean values are statistically different (P < 0.05). RLUs/s: relative luminescence units per second. Figure 1. View largeDownload slide Effects of different gene deletions on H+-ATPase proton-pumping activity and intracellular calcium signaling in S. cerevisiae cells (genetic background PJ69-2A). (A) Proton-pumping activity was measured with around 500 mg of cells, after addition of 100 mM glucose (final concentration). (B) Cytosolic free calcium was measured by using a standard aequorin-based bioluminescence assay after addition of 100 mM glucose (final concentration), indicated by a striped arrow at ‘0’ s, in yeast strains containing apoaequorin-expressing plasmid pVTU-AEQ and loaded with 50 μM coelenterazine. Different letters indicate that mean values are statistically different (P < 0.05). RLUs/s: relative luminescence units per second. In the same set of strains and in similar conditions, the influence of LPX1 deletion in calcium signaling was assessed (Fig. 1B). Initially, it was observed that the strain with single LPX1 deletion showed calcium signaling comparable to wild-type cells. When LPX1 was deleted in combination with ARG82, there was a stronger intracellular calcium signal, as observed in arg82Δ mutant cells. Calcium signaling was reduced in yvc1Δ cells and in the double mutant yvc1Δ lpx1Δ strain. Therefore, the results shown in Fig. 1 suggest that Lpx1p is not necessary for generation of calcium signal on glucose treatment; however, it is required for calcium-mediated activation of the H+-ATPase. Thus, Lpx1p should work downstream of IP3 and calcium signaling and, consequently, can form a link between the calcium signal and the H+-ATPase function. Moreover, this hypothesis was also reinforced by the fact that a database search, using Lpx1p crystal structure, indicated the presence of a potential region for calcium interaction with a high-confidence prediction (95% precision performance) located at its C-terminus (Supplementary Material 2), suggesting that this domain would be an EF-hand calcium-binding domain. Overexpression of LPX1 in an lpx1Δ mutant restores normal proton-pumping rate and increases H+-ATPase activity An lpx1Δ mutant yeast strain expressing the LPX1 gene fused with a His-tag at the 3΄-extremity in a galactose-inducible vector (pYES2/CT-LPX1) was generated to verify the potential connection between calcium signaling and Lpx1p activation. As control, the lpx1Δ yeast strain was transformed with the corresponding empty vector (pYES2/CT-EV). In order to check the suitability of these constructions, transformant yeasts were submitted to galactose induction for different incubation times (10, 30 and 50 min for pYES2/CT-LPX1 and 50 min for pYES2/CT-EV). After induction, glucose-induced extracellular acidification was performed. As shown in Fig. 2A, cells expressing the LPX1 gene presented an increase in proton-pumping rates after galactose induction (P < 0.05), when compared with non-induced cells. After 10, 30 and 50 min, proton-pumping rate was increased to 58.0%, 87.3% and 112.4%, respectively, compared with cells without galactose induction. Proton-pumping activity after 50 min induction in pYES2/CT-LPX1 cells was comparable to the wild-type strain (P > 0.05). Cells transformed with the empty plasmid did not exhibit any increase in proton-pumping rate after 50 min galactose induction, showing the same rate as found in the lpx1Δ strain (P > 0.05). Interestingly, when the yvc1Δ  mutant was also transformed with such constructions and exposed to galactose, there was no increase in the H+-pumping rate even after 50 min, with the activity remaining comparable to that registered for the yvc1Δ  mutant (Fig. 2A). These results suggest that the activity of the calcium channel Yvc1 is not controlled by Lpx1p. Figure 2. View largeDownload slide Plasma membrane proton-pumping rate and ATPase activity in lpx1Δ mutant transformed with pYES2/CT-EV and pYES2/CT-LPX1 vectors after induction by galactose in different times. Wild-type strain (BY4741) and the corresponding lpx1Δ and yvc1Δ mutants were used as controls. (A) Proton-pumping rate was measured with around 500 mg of cells after induction for different times with 100 mM galactose. Cells without incubation with 100 mM galactose were used as controls. (B) To measure ATPase activity from purified plasma membrane H+-ATPase, cells were collected after overnight induction with galactose, and membranes were obtained as described previously with a standard method. Different letters and asterisk indicate that mean values are statistically different (P < 0.05). Figure 2. View largeDownload slide Plasma membrane proton-pumping rate and ATPase activity in lpx1Δ mutant transformed with pYES2/CT-EV and pYES2/CT-LPX1 vectors after induction by galactose in different times. Wild-type strain (BY4741) and the corresponding lpx1Δ and yvc1Δ mutants were used as controls. (A) Proton-pumping rate was measured with around 500 mg of cells after induction for different times with 100 mM galactose. Cells without incubation with 100 mM galactose were used as controls. (B) To measure ATPase activity from purified plasma membrane H+-ATPase, cells were collected after overnight induction with galactose, and membranes were obtained as described previously with a standard method. Different letters and asterisk indicate that mean values are statistically different (P < 0.05). In addition, to check the role of Lxp1p in H+-ATPase activation, ATPase activity was measured in purified plasma membranes of the lpx1Δ yeast strains transformed with pYES2/CT empty vector (pYES2/CT-EV) and the one containing the LPX1 gene (pYES2/CT-LPX1). After overnight galactose induction, plasma membranes from strain pYES2/CT-LPX1 showed higher activity (P < 0.05) than purified plasma membranes from the control strain, pYES2/CT-EV (Fig. 2B). Proteolytic activity of Lpx1 Initially, crude extracts of lpx1Δ strains transformed with pYES2/CT-EV (empty vector, used as control), pYES2/CT-LPX1 (vector containing LPX1 whole gene) and pYES2/CT-LPX1-MOD (vector containing a modified LPX1 gene, where the binding calcium domain was removed) were prepared and applied to a nickel column, to perform Lpx1p (or Lpx1p-MOD) His-tag purification. In order to confirm the presence of His-tagged Lpx1 proteins, a western blotting assay was performed (Fig. 3A) with eluates obtained from lpx1Δ cells transformed with pYES2/CT-EV, pYES2/CT-LPX1 and pYES2/CT-LPX1-MOD (without calcium-binding domain). The results demonstrated that the eluates from the nickel column recovered from lpx1Δ cells presented specific bands only in extracts obtained from lpx1Δ cells transformed with pYES2/CT-LPX1 and pYES2/CT-LPX1-MOD. Furthermore, no band was detected in the extract obtained from the strain transformed with the empty vector. Additionally, reduction in molecular mass of the Lpx1p from 49 to 41 kDa was confirmed in a western blotting experiment that used extracts obtained from lpx1Δ cells transformed from pYES2/CT-LPX1 and pYES2/CT-LPX1-MOD, respectively. These results were supported by differences in their specific relative migration distances (Rf) (Supplementary Material 4). Figure 3. View largeDownload slide Lpx1p His-Tag expression and proteolytic activity. Cells were incubated overnight in induction medium, collected and disrupted in lysis buffer. Resulting supernatant (crude extract) was incubated with HIS-Select nickel affinity gel (Sigma-Aldrich). Unbound proteins were removed by centrifugation. Column-bound proteins were recovered with elution buffer. (A) Western blotting assay using anti-His antibodies in samples of nickel-column eluates obtained from lpx1Δ strain transformed with the empty vector pYES2/CT-EV and vectors carrying pYES2/CT-LPX1 or pYES2/CT-LPX1-MOD. Arrows indicate specific bands. (B) Proteolytic activity of total membranes from wild-type and lpx1Δ mutant was assessed by using azocasein as substrate. A protein fraction was added to a reaction mixture (100 mM Tris–HCl, pH 8.0, 0.5% azocasein) with or without addition of calcium. One enzymatic activity unit was defined as a change in absorbance of 0.01 (428 nm) per minute in these conditions. (C) Lpx1p activity was evaluated in excised protein bands obtained from SDS-PAGE gels after electrophoresis of eluates of the lpx1Δ mutant transformed from pYES2/CT-EV and pYES2/CT-LPX1 vectors. Samples (100 μg of protein) were dissolved in non-reducing Laemmli buffer and run in non-reducing conditions at 4°C. Lpx1p bands (indicated by arrows in A) and the corresponding region obtained from the control system (indicated by a circle in A) were excised from gel and added to a reaction mixture (100 mM Tris–HCl, pH 8.0, 0.5% azocasein), with or without addition of calcium. One enzymatic activity unit was defined as a change in absorbance of 0.01 (428 nm) per minute in these conditions. In all cases, results are expressed as specific activity (as mg protein−1). Different letters and asterisk indicate that mean values are statistically different (P < 0.05). Figure 3. View largeDownload slide Lpx1p His-Tag expression and proteolytic activity. Cells were incubated overnight in induction medium, collected and disrupted in lysis buffer. Resulting supernatant (crude extract) was incubated with HIS-Select nickel affinity gel (Sigma-Aldrich). Unbound proteins were removed by centrifugation. Column-bound proteins were recovered with elution buffer. (A) Western blotting assay using anti-His antibodies in samples of nickel-column eluates obtained from lpx1Δ strain transformed with the empty vector pYES2/CT-EV and vectors carrying pYES2/CT-LPX1 or pYES2/CT-LPX1-MOD. Arrows indicate specific bands. (B) Proteolytic activity of total membranes from wild-type and lpx1Δ mutant was assessed by using azocasein as substrate. A protein fraction was added to a reaction mixture (100 mM Tris–HCl, pH 8.0, 0.5% azocasein) with or without addition of calcium. One enzymatic activity unit was defined as a change in absorbance of 0.01 (428 nm) per minute in these conditions. (C) Lpx1p activity was evaluated in excised protein bands obtained from SDS-PAGE gels after electrophoresis of eluates of the lpx1Δ mutant transformed from pYES2/CT-EV and pYES2/CT-LPX1 vectors. Samples (100 μg of protein) were dissolved in non-reducing Laemmli buffer and run in non-reducing conditions at 4°C. Lpx1p bands (indicated by arrows in A) and the corresponding region obtained from the control system (indicated by a circle in A) were excised from gel and added to a reaction mixture (100 mM Tris–HCl, pH 8.0, 0.5% azocasein), with or without addition of calcium. One enzymatic activity unit was defined as a change in absorbance of 0.01 (428 nm) per minute in these conditions. In all cases, results are expressed as specific activity (as mg protein−1). Different letters and asterisk indicate that mean values are statistically different (P < 0.05). The proteolytic activity of Lpx1 and its Ca2+ dependence was demonstrated by two different approaches: first, we isolate total membranes from wild-type and lpx1Δ cells and the activity was tested using a non-specific substrate (azocasein). As demonstrated in Fig. 3B, the presence of 1 mM calcium increases the proteolytic specific activity (AU/mg protein−1) in samples from wild-type cells; on the other hand, this effect was not observed in membrane samples obtained from the lpx1Δ mutant cells. Another experiment was performed to confirm the proteolytic activity of Lpx1. The corresponding protein bands obtained from PAGE gels of eluate from the lpx1Δ mutant transformed with pYES2/CT-LPX1 vector showed proteolytic activity clearly affected by the presence of 100 μM calcium. Excised gel fractions obtained from the strain transformed with an empty vector showed no proteolytic activity, even in the presence of calcium (Fig. 3C). In vitro plasma membrane H+-ATPase activation In order to confirm the connection between calcium signaling, Lpx1p activity and H+-ATPase activation, an in vitro system to reconstitute H+-ATPase was used. Initially, purified plasma membranes and dialyzed MFE obtained from glucose-starved wild-type cells were prepared. In these conditions, plasma membrane H+-ATPase is in a non-activated state (probably with acetylated tubulins bounded to the H+-ATPase C-terminal tail as described by Campetelli et al. (2013)). MFE provides the in vitro system with all elements required for H+-ATPase activation, including Ptk2p (required for H+-ATPase phosphorylation) and Lpx1p (necessary for acetylated tubulin proteolysis). To achieve the most suitable conditions for this in vitro activation assay, different calcium concentrations (100 μM, 1 and 5 mM), different times of incubation (10, 30 and 60 min) and different MFE protein concentrations (50, 100, 200 and 400 μg) for in vitro H+-ATPase activation were tested to define the most suitable conditions (data not shown). Thus, adequate conditions for the assay were as follows: 30 min of incubation with 50 μg of protein from MFE and with the addition of 100 μM calcium. When purified plasma membranes obtained from the wild-type glucose-starved cells were incubated only with (i) dialyzed MFE from the same strain or (ii) only calcium, there was no H+-ATPase activation in comparison with the activity detected when only purified plasma membrane from wild-type glucose-starved cells was used (Fig. 4A). However, when purified plasma membranes were combined with both MFE and calcium, H+-ATPase activation was observed (47.5% increase in activity, P < 0.05). Additionally, activity was strongly reduced (77.4% reduced, P < 0.05) when 50 μM sodium orthovanadate, a specific plasma membrane H+-ATPase inhibitor, was added. This seems to confirm that ATP hydrolytic activity measure in this assay was due to plasma membrane H+-ATPase (Fig. 4A). Figure 4. View largeDownload slide H+-ATPase in vitro activation assay. (A) Effect of calcium, membrane-free extract (MFE) and sodium orthovanadate addition in H+-ATPase in vitro activation assay. A method to evaluate in vitro H+-ATPase activation was established in this work. Purified plasma membranes and dialyzed MFE from BY4741 cells were used. Plasma membranes from glucose-starved wild-type cells were added to incubation buffer. Calcium and/or MFE was added accordingly to the different assays performed. To verify H+-ATPase activity inhibition, 50 μM of sodium orthovanadate (specific H+-ATPase inhibitor) was added. (B) Effect of Lpx1p and Ptk2p on H+-ATPase in vitro activation. Plasma membranes from glucose-starved wild-type cells were added to incubation buffer. MFE from lpx1Δ strain (lacking Lpx1p, but containing Ptk2p) or ptk2Δ cells (missing Ptk2p, but expressing Lpx1p) and calcium were added according to the different assays performed. (C) Plasma membranes from glucose-starved wild-type cells were added to incubation buffer with MFE from lpx1Δ or ptk2Δ strains. Eluates resulting from the application of cell extracts of lpx1Δ mutant (transformed with pYES2/CT-LPX1 or pYES2/CT-LPX1-MOD vectors) to a Ni-affinity column, with or without calcium addition, were used to evaluate the H+-ATPase in vitro activation. All results are expressed as a percentage relative to wild-type control assay with plasma membranes only. Asterisks indicate that mean values are statistically different from those seen in each relative control used (P < 0.05). Figure 4. View largeDownload slide H+-ATPase in vitro activation assay. (A) Effect of calcium, membrane-free extract (MFE) and sodium orthovanadate addition in H+-ATPase in vitro activation assay. A method to evaluate in vitro H+-ATPase activation was established in this work. Purified plasma membranes and dialyzed MFE from BY4741 cells were used. Plasma membranes from glucose-starved wild-type cells were added to incubation buffer. Calcium and/or MFE was added accordingly to the different assays performed. To verify H+-ATPase activity inhibition, 50 μM of sodium orthovanadate (specific H+-ATPase inhibitor) was added. (B) Effect of Lpx1p and Ptk2p on H+-ATPase in vitro activation. Plasma membranes from glucose-starved wild-type cells were added to incubation buffer. MFE from lpx1Δ strain (lacking Lpx1p, but containing Ptk2p) or ptk2Δ cells (missing Ptk2p, but expressing Lpx1p) and calcium were added according to the different assays performed. (C) Plasma membranes from glucose-starved wild-type cells were added to incubation buffer with MFE from lpx1Δ or ptk2Δ strains. Eluates resulting from the application of cell extracts of lpx1Δ mutant (transformed with pYES2/CT-LPX1 or pYES2/CT-LPX1-MOD vectors) to a Ni-affinity column, with or without calcium addition, were used to evaluate the H+-ATPase in vitro activation. All results are expressed as a percentage relative to wild-type control assay with plasma membranes only. Asterisks indicate that mean values are statistically different from those seen in each relative control used (P < 0.05). All in vitro plasma membrane H+-ATPase activity rates measured in these assays ranged between 0.10 and 0.28 μmol Pi min−1 mg−1 protein. There was no ATPase activity detected when the assay was performed with only MFE or eluates from all strains used in in vitro activation (data not shown). Then, to confirm the connection between calcium signaling, Lpx1p activity and Ptk2p (a protein kinase) in the context of the phosphorylation/activation of H+-ATPase, this in vitro H+-ATPase activation was tested using purified membranes obtained from wild-type cells and different MFEs prepared from ptk2Δ and lpx1Δ mutant strains. As can be seen in Fig. 4B, the addition of MFE obtained from lpx1Δ alone or in combination with 100 μM calcium does not result in any increase of the H+-ATPase activity (P > 0.05). When using MFE from ptk2Δ mutants, a slight increase (11.8%, P < 0.05) was verified on addition of 100 μM calcium. Moreover, there was no increase in H+-ATPase activity when the Ni-column eluate obtained from the lpx1Δ strain transformed with an empty vector (pYES2/CT-EV, used as control) was assessed, even with calcium addition combined with MFE from lpx1Δ or ptk2Δ strains (data not shown). Also, there was no increase (P > 0.05) in H+-ATPase activity when Ni-column eluate obtained from the lpx1Δ strain transformed with pYES2/CT-LPX1 vector was associated with MFE prepared from the ptk2Δ mutant, with or without 100 μM calcium addition (Fig. 4C, first two columns at left side). At the same time, when the Ni-column eluates obtained from the lpx1Δ strain transformed pYES2/CT-LPX1 vector was added to the incubation system, in association with MFE from lpx1Δ strain, a clear increase (57.3%; P < 0.05) in H+-ATPase activity was only observed when 100 μM calcium was present (central two columns in Fig 4C). On the other hand, with Ni-column eluates obtained from the lpx1Δ strain transformed with pYES2/CT-LPX1-MOD vector, any increase of the H+-ATPase activity was observed in association with MFE from lpx1Δ even in the presence of 100 μM calcium (Fig. 4C, two last columns at right side). DISCUSSION In the past 20 years, a large amount of evidence has suggested that glucose-induced activation of plasma membrane H+-ATPase is clearly dependent on calcium metabolism (dos Passos et al.1992; Brandão et al.1994; Coccetti et al.1998; Souza, Tropia and Brandao 2001; Trópia et al.2006; Pereira et al.2008; Bouillet et al.2012). Nevertheless, until now, a Ca2+-dependent protein kinase that could be involved in H+-ATPase activation through phosphorylation has never been identified. In fact, it was demonstrated that protein kinase Ptk2p would be the enzyme responsible for glucose-induced phosphorylation of residue Ser-899, located at the H+-ATPase C-terminal tail, leading to a Km reduction of the ATPase (Goossens et al.2000; Pereira et al.2015), but this kinase does not seem to belong to any calcium responsive class of protein kinases. Moreover, neither a second protein kinase (which would be responsible for the phosphorylation of Ser-911/Thr-912 residues) nor any other Ca2+-dependent protein has ever been described as being involved in this activation process. Therefore, to overcome this apparent contradiction, it seems necessary to us to develop different strategies to demonstrate how glucose-induced calcium signaling would be connected to the plasma membrane H+-ATPase activation process. Interestingly, it was already proposed that a serine protease is related to H+-ATPase activation. These pieces of evidence suggest that activation of the H+-ATPase would require a glucose-induced hydrolysis of an acetylated tubulin to liberate its C-terminal tail, making it available for phosphorylation and activation (Campetelli et al.2013). This proteolytic action is exerted by a serine protease encoded by the LPX1 gene (Campetelli et al.2005, 2013), and, as a new component of this elaborated pathway, it could be investigated as a possible candidate that would respond to the calcium signal, thereby explaining its function in the glucose-induced activation of plasma membrane H+-ATPase. We also found that Lpx1p presents a potential region for calcium interaction (an EF-hand, calcium-binding domain), apparently located at its C-terminus, when a search with WebFEATURE mas made. WebFEATURE allows scanning for different functional sites in proteins using predicted 3D molecular structure (Liang et al.2003; Wu, Liang and Altman 2008). The EF-hand motif is the most common calcium-binding motif found in proteins. In a large number of proteins, this motif does indeed bind calcium (or, in some cases, magnesium) and these proteins exert diverse functions such as calcium buffering in cytosol, signal transduction between cellular compartments and muscle contraction (Lewit-Bentley and Réty 2000). Thus, this predicted that an EF-hand calcium-binding domain in the structure of Lpx1p could be responsible for calcium interaction and possibly regulate its activity. Additionally, Thoms et al. (2011) found, also in a search for functional sites in the Lpx1p structure, that the cap domain of this protein shows similarity to calmodulin, a well-known intracellular protein target of Ca2+ in eukaryotes. According to these authors, this cap covers the Lpx1p active site, and its N-terminal loop shows characteristics indicating high flexibility, which could suggest that this loop might act as a lid that can regulate access to the active site (Thoms et al.2011). Curiously, it seems that Lpx1p also exhibits acyl hydrolase and phospholipase A activities and can be found in peroxisomes (Thoms et al.2008); it also shows an ambiguous distribution (Huh et al.2003). Therefore, Lpx1p can be considered a good candidate to form a link between calcium signaling and glucose-induced plasma membrane H+-ATPase activation. This possibility was initially investigated by using strains with single or combined deletions in different genes: ARG82, YVC1 and LPX1. Cells with deletion in ARG82 (which encodes an inositol kinase responsible for the phosphorylation of IP3) have an increase in both glucose-induced calcium signaling and H+-ATPase activation (Tisi et al.2004; Trópia et al.2006; Pereira et al.2008). This increase suggests that H+-ATPase activation is dependent on calcium signaling. As demonstrated here, deletion of LPX1 alone or in combination with ARG82 led to a reduction in the glucose-induced activation of the plasma membrane H+-ATPase. Nevertheless, in the lpx1Δ mutant the calcium signaling was comparable to wild-type cells. Additionally, in the double mutant arg82Δ lpx1Δ, glucose-induced calcium signaling was also higher, as in the single mutant arg82Δ, suggesting that Lpx1p is not directly involved in calcium signaling. These results were also confirmed when calcium signaling and proton-pumping activity were measured in yvc1Δ and yvc1Δ lpx1Δ strains. An alternative explanation for these results would be the existence of a signal transduction pathway, in which a glucose-induced calcium signal would be responsible for protease Lpx1p activation. According to a model suggested before (Campetelli et al.2013), this protease hydrolyzes an acetylated tubulin bound to the H+-ATPase C-terminal tail, making this region accessible to protein kinases. Indeed, here we demonstrated that the lpx1Δ strain expressing the LPX1 gene inserted in a galactose-inducible vector (pYES2/CT-LPX1) showed progressive increase in the in vivo H+-ATPase activity, confirming that Lpx1p is indeed essential to H+-ATPase activation. The results of a similar experiment done with the yvc1Δ  mutant transformed with the same vector also suggest that the calcium channel Yvc1 is not controlled by Lpx1p. These results, combined with proteolytic activity assays, indicate that Lpx1p proteolytic activity could be the target of calcium signaling, making the C-terminal tail of the H+-ATPase accessible to phosphorylation performed by at least one protein kinase (Ptk2p). To verify the direct connection between calcium signaling, Lpx1p activity and H+-ATPase activation, it was necessary to establish an in vitro system by which it would be possible to reconstitute the plasma membrane H+-ATPase activation. Thus, when purified plasma membrane (isolated from wild-type cells) was incubated with MFE from lpx1Δ or ptk2Δ strains, in the presence of 100 μM calcium, H+-ATPase activation was absent or had a slight increase, respectively. This suggests once again that these two proteins (Lpx1 and Ptk2) are necessary to trigger a proper activation. Moreover, calcium and Lpx1p addition (obtained from the eluate of a Ni-affinity column of a cell extract of the lpx1Δ mutant transformed with a pYES2/CT-LPX1 vector), combined with MFE from lpx1Δ cells (which gives other components necessary for activation, but not Lpx1p), resulted in an increase of H+-ATPase activity only when 100 μM calcium is also present. However, in similar conditions when the MFE from ptk2Δ strain was used, no increase in H+-ATPase activity was observed, even in the presence of 100 μM calcium. These results reinforce the importance of Ptk2p in the activation process of H+-ATPase. Finally, when modified Lpx1p protein (without the C-terminal tail portion that potentially interacts with Ca2+) was used in combination with MFE from lpx1Δ cells and calcium, no increase in the H+-ATPase activity was observed. Altogether, these approaches suggest that most probably glucose-induced calcium signaling is connected to plasma membrane H+-ATPase activation through a calcium-induced activation of the serine protease Lpx1p. Thus, with the results presented here, the following mechanism can be proposed: on glucose addition, Ca2+ binds to Lpx1p, leading to its activation making possible the degradation of the acetylated tubulin bound to the plasma membrane H+-ATPase. Therefore, the H+-ATPase C-terminal tail would be accessible to phosphorylation performed by at least one protein kinase (Ptk2p) leading to the activation of the plasma membrane H+-ATPase (Fig. 5). Figure 5. View largeDownload slide Lpx1p calcium binding and H+-ATPase activation. (A) Proposed mechanism by which Lpx1p is possibly regulated by calcium availability. In the presence of glucose, calcium is released into cytoplasm and binds to Lxp1p. (B) After calcium binding, Lxp1p is able to degrade acetylated tubulin bound to plasma membrane H+-ATPase releasing its C-terminal tail. This would make the H+-ATPase C-terminus phosphorylation sites accessible to at least one protein kinase (Ptk2p) making possible the enzyme activation. Figure 5. View largeDownload slide Lpx1p calcium binding and H+-ATPase activation. (A) Proposed mechanism by which Lpx1p is possibly regulated by calcium availability. In the presence of glucose, calcium is released into cytoplasm and binds to Lxp1p. (B) After calcium binding, Lxp1p is able to degrade acetylated tubulin bound to plasma membrane H+-ATPase releasing its C-terminal tail. This would make the H+-ATPase C-terminus phosphorylation sites accessible to at least one protein kinase (Ptk2p) making possible the enzyme activation. Therefore, taking together the data from our previous work (dos Passos et al.1992; Brandão et al.1994; Coccetti et al.1998; Souza, Tropia and Brandao 2001; Trópia et al.2006; Pereira et al.2008; Groppi et al.2011; Bouillet et al.2012) and the new results shown in this work, we suggest the existence of a signal transduction pathway with two branches, by which glucose addition controls calcium availability in the cytosol with a direct consequence for plasma membrane H+-ATPase activation (Fig. 6). In the first branch, glucose uptake and its subsequent phosphorylation generate a signal (probably sugar phosphates) that would stimulate G protein Gpa2p to interact with and/or activate phospholipase C (Plc1p). Then, Plc1p would hydrolyze phosphatidylinositol-4,5-bisphosphate, generating diacylglycerol and IP3. IP3 would interact, directly or indirectly, with Yvc1p regulating the intensity of calcium signaling in the cytosol (Bouillet et al.2012). In the second branch, devoted to the control of Pmc1p Ca2+-ATPase activity, the glucose sensor Snf3p (Özcan and Johnston 1999) could also detect sugar phosphates (Dlugai et al.2001), and in some way transduce this signal leading to an increase in Pmc1p activity (Souza, Tropia and Brandao 2001; Trópia et al.2006; Pereira et al.2008). The balance between these two branches would be responsible for the transient nature of calcium signaling. This calcium signaling seems to be responsible for activation of Lpx1p, which hydrolyzes an acetylated tubulin bound to plasma membrane H+-ATPase, allowing thus the H+-ATPase C-terminal tail to be released, enabling phosphorylation of C-terminal sites of the H+-ATPase and in this way its activation. Figure 6. View largeDownload slide Working model: glucose-induced activation of plasma membrane H+-ATPase in yeast cells. A signal transduction pathway with two branches, resulting in intracellular calcium-signal generation, is triggered by internalization followed by phosphorylation of glucose, which generates a signal (maybe relative amounts of glucose-6-P and/or glucose-1-P) that would stimulate the complex constituted of the G protein, Gpa2p, and phospholipase C eliciting the activation of the phospholipase C. Then, phosphatidylinositol-4,5-bisphosphate hydrolysis would generate diacylglycerol and IP3. In the first branch, IP3 would act directly or indirectly on the vacuolar calcium channel Yvc1p leading to an increase in the intracellular calcium signal. Besides that, in a second branch, the C-terminal tail of the glucose sensor Snf3p controlling the activity of the vacuolar Ca2+-ATPase, Pmc1p, would detect the signal. The final intensity of the calcium signal would be the result of the partial contribution of each branch of this system. This calcium signaling seems to be responsible for activation of Lpx1p, which hydrolyzes an acetylated tubulin bound to plasma membrane H+-ATPase, allowing thus the H+-ATPase C-terminal tail to be released, enabling phosphorylation of C-terminal sites of the H+-ATPase and in this way its activation. Figure 6. View largeDownload slide Working model: glucose-induced activation of plasma membrane H+-ATPase in yeast cells. A signal transduction pathway with two branches, resulting in intracellular calcium-signal generation, is triggered by internalization followed by phosphorylation of glucose, which generates a signal (maybe relative amounts of glucose-6-P and/or glucose-1-P) that would stimulate the complex constituted of the G protein, Gpa2p, and phospholipase C eliciting the activation of the phospholipase C. Then, phosphatidylinositol-4,5-bisphosphate hydrolysis would generate diacylglycerol and IP3. In the first branch, IP3 would act directly or indirectly on the vacuolar calcium channel Yvc1p leading to an increase in the intracellular calcium signal. Besides that, in a second branch, the C-terminal tail of the glucose sensor Snf3p controlling the activity of the vacuolar Ca2+-ATPase, Pmc1p, would detect the signal. The final intensity of the calcium signal would be the result of the partial contribution of each branch of this system. This calcium signaling seems to be responsible for activation of Lpx1p, which hydrolyzes an acetylated tubulin bound to plasma membrane H+-ATPase, allowing thus the H+-ATPase C-terminal tail to be released, enabling phosphorylation of C-terminal sites of the H+-ATPase and in this way its activation. Beyond representing a clear advance in our understanding of how glucose-induced calcium signaling and plasma membrane H+-ATPase activation would be connected, these findings can help in the elaboration of new strategies to find out the identity of all protein kinases that are involved in plasma membrane H+-ATPase phosphorylation. Perhaps, the fact that a proteolytic degradation of acetylated tubulin precludes and/or occurs in parallel with phosphorylation could help in conceiving new approaches to identify those protein kinases in addition to Ptk2p. SUPPLEMENTARY DATA Supplementary data are available at FEMSYR online. Acknowledgements We are also grateful to Dr James Caffrey from National Institute of Environmental Health Sciences, USA, for the strains used in this work and to Marco Vanoni (Università di Milano-Bicocca, Milan, Italy) for pVTU-AEQ and pYX212-AEQ plasmids. FUNDING This work was partially financed by grants from Universidade Federal de Ouro Preto, from Fundação de Amparo a Pesquisa do Estado de Minas Gerais (FAPEMIG), Process CBB 824/06; from Conselho Nacional de Desenvolvimento–CNPq, Process 304815/2012-3 (research fellowship to RLB), Process 475672/08-5 (research grant) and CAPES, Process 2041/2012 (PhD fellowship to FFO), Process PNPD/2013 (research fellowships to RHSD), Process PNPD 2755/2011 (research fellowships to FGS), Process BEX 11122/13-7 (PhD Sandwich fellowship to DDC). Conflict of Interest. None declared. REFERENCES Anraku Y, Ohya Y, Iida H. Cell cycle control by calcium and calmodulin in Saccharomyces cerevisiae. Biochim Biophys Acta  1991; 1093: 169– 77. Google Scholar CrossRef Search ADS PubMed  Bouillet LE, Cardoso AS, Perovano E et al.   The involvement of calcium carriers and of the vacuole in the glucose-induced calcium signaling and activation of the plasma membrane H+-ATPase in Saccharomyces cerevisiae cells. Cell Calcium  2012; 51: 72– 81. Google Scholar CrossRef Search ADS PubMed  Brandão RL, de Magalhães-Rocha NM, Alijo R et al.   Possible involvement of a phosphatidylinositol-type signaling pathway in glucose-induced activation of plasma membrane H+-ATPase and cellular proton extrusion in the yeast Saccharomyces cerevisiae. Biochim Biophys Acta  1994; 1223: 117– 24. Google Scholar CrossRef Search ADS PubMed  Brandão RL. The relationship between glucose-induced calcium signaling and activation of the plasma membrane H+-ATPase in Saccharomyces cerevisiae cells. In: Nakamura S (ed.). Handbook of H+-ATPases . Singapore: Pan Stanford Publishing Pte Ltd, 2014, 431– 48. Google Scholar CrossRef Search ADS   Campetelli AN, Monesterolo NE, Previtali G et al.   Activation of H+-ATPase by glucose in Saccharomyces cerevisiae involves a membrane serine protease. Biochim Biophys Acta  2013; 1830: 3593– 603. Google Scholar CrossRef Search ADS PubMed  Campetelli AN, Previtali G, Arce CA et al.   Activation of the plasma membrane H+-ATPase of Saccharomyces cerevisiae by glucose is mediated by dissociation of the H+-ATPase–acetylated tubulin complex. FEBS J  2005; 272: 5742– 52. Google Scholar CrossRef Search ADS PubMed  Capieaux E, Vignais ML, Sentenac A et al.   The yeast H+-ATPase gene is controlled by the promoter binding factor TUF. J Biol Chem  1989; 264: 7437– 46. Google Scholar PubMed  Charney J, Tomarelli RM. A colorimetric method for the determination of the proteolytic activity of duodenal juice. J Biol Chem  1947; 171: 501– 5. Google Scholar PubMed  Coccetti P, Tisi R, Martegani E et al.   The PLC1 encoded phospholipase C in the yeast Saccharomyces cerevisiae is essential for glucose-induced phosphatidylinositol turnover and activation of plasma membrane H+-ATPase. Biochim Biophys Acta  1998; 1405: 147– 54. Google Scholar CrossRef Search ADS PubMed  Cunningham KW. Acidic calcium stores of Saccharomyces cerevisiae. Cell Calcium  2011; 50: 129– 38. Google Scholar CrossRef Search ADS PubMed  Cyert MS, Philpott CC. Regulation of cation balance in Saccharomyces cerevisiae. Genetics  2013; 193: 677– 713. Google Scholar CrossRef Search ADS PubMed  Denis V, Cyert MS. Internal Ca2+ release in yeast is triggered by hypertonic shock and mediated by a TRP channel homologue. J Cell Biol  2002; 156: 29– 34. Google Scholar CrossRef Search ADS PubMed  Dlugai S, Hippler S, Wieczorke R et al.   Glucose-dependent and -independent signalling functions of the yeast glucose sensor Snf3. FEBS Lett  2001; 505: 389– 92. Google Scholar CrossRef Search ADS PubMed  dos Passos JB, Vanhalewyn M, Brandao RL et al.   Glucose-induced activation of plasma membrane H+-ATPase in mutants of the yeast Saccharomyces cerevisiae affected in cAMP metabolism, cAMP-dependent protein phosphorylation and the initiation of glycolysis. Biochim Biophys Acta  1992; 1136: 57– 67. Google Scholar CrossRef Search ADS PubMed  Dunn T, Gable K, Beeler T. Regulation of cellular Ca2+ by yeast vacuoles. J Biol Chem  1994; 269: 7273– 8. Google Scholar PubMed  Eraso P, Mazón MJ, Portillo F. Yeast protein kinase Ptk2 localizes at the plasma membrane and phosphorylates in vitro the C-terminal peptide of the H+-ATPase. Biochim Biophys Acta  2006; 1758: 164– 70. Google Scholar CrossRef Search ADS PubMed  Gietz RD, Schiestl RH, Willems AR et al.   Studies on the transformation of intact yeast cells by the LiAc/SS-DNA/PEG procedure. Yeast  1995; 11: 355– 60. Google Scholar CrossRef Search ADS PubMed  Goffeau A, Slayman CW. The proton-translocating ATPase of the fungal plasma membrane. Biochim Biophys Acta  1981; 639: 197– 223. Google Scholar CrossRef Search ADS PubMed  Goossens A, de La Fuente N, Forment J et al.   Regulation of yeast H+-ATPase by protein kinases belonging to a family dedicated to activation of plasma membrane transporters. Mol Cell Biol  2000; 20: 7654– 61. Google Scholar CrossRef Search ADS PubMed  Groppi S, Belotti F, Brandão RL et al.   Glucose-induced calcium influx in budding yeast involves a novel calcium transport system and can activate calcineurin. Cell Calcium  2011; 49: 376– 86. Google Scholar CrossRef Search ADS PubMed  Huh W-K, Falvo JV, Gerke LC et al.   Global analysis of protein localization in budding yeast. Nature  2003; 425: 686– 91. Google Scholar CrossRef Search ADS PubMed  Lecchi S, Allen KE, Pardo JP et al.   Conformational changes of yeast plasma membrane H+-ATPase during activation by glucose: role of threonine-912 in the carboxy-terminal tail. Biochemistry  2005; 44: 16624– 32. Google Scholar CrossRef Search ADS PubMed  Lecchi S, Nelson CJ, Allen KE et al.   Tandem phosphorylation of Ser-911 and Thr-912 at the C terminus of yeast plasma membrane H+-ATPase leads to glucose-dependent activation. J Biol Chem  2007; 282: 35471– 81. Google Scholar CrossRef Search ADS PubMed  Lewit-Bentley A, Réty S. EF-hand calcium-binding proteins. Curr Opin Struct Biol  2000; 10: 637– 43. Google Scholar CrossRef Search ADS PubMed  Liang MP, Banatao DR, Klein TE et al.   WebFEATURE: an interactive web tool for identifying and visualizing functional sites on macromolecular structures. Nucleic Acids Res  2003; 31: 3324– 7. Google Scholar CrossRef Search ADS PubMed  Lowry OH, Rosebrough NJ, Farr AL et al.   Protein measurement with the Folin phenol reagent. J Biol Chem  1951; 193: 265– 75. Google Scholar PubMed  Mazón MJ, Eraso P, Portillo F. Specific phosphoantibodies reveal two phosphorylation sites in yeast Pma1 in response to glucose. FEMS Yeast Res  2015; 15: fov030. Google Scholar CrossRef Search ADS PubMed  Miseta A, Fu L, Kellermayer R et al.   The Golgi apparatus plays a significant role in the maintenance of Ca2+ homeostasis in the vps33Δ vacuolar biogenesis mutant of Saccharomyces cerevisiae. J Biol Chem  1999a; 274: 5939– 47. Google Scholar CrossRef Search ADS   Miseta A, Kellermayer R, Aiello DP et al.   The vacuolar Ca2+/H+ exchanger Vcx1p/Hum1p tightly controls cytosolic Ca2+ levels in S. cerevisiae. FEBS Lett  1999b; 451: 132– 6. Google Scholar CrossRef Search ADS   Odom AR, Stahlberg A, Wente SR et al.   A role for nuclear inositol 1,4,5-trisphosphate kinase in transcriptional control. Science  2000; 287: 2026– 9. Google Scholar CrossRef Search ADS PubMed  Özcan S, Johnston M. Function and regulation of yeast hexose transporters. Microbiol Mol Biol R  1999; 63: 554– 69. Paidhungat M, Garrett S. A homolog of mammalian, voltage-gated calcium channels mediates yeast pheromone-stimulated Ca2+ uptake and exacerbates the cdc1 (Ts) growth defect. Mol Cell Biol  1997; 17: 6339– 47. Google Scholar CrossRef Search ADS PubMed  Pereira MB, Tisi R, Fietto LG et al.   Carbonyl cyanide m-chlorophenylhydrazone induced calcium signaling and activation of plasma membrane H+-ATPase in the yeast Saccharomyces cerevisiae. FEMS Yeast Res  2008; 8: 622– 30. Google Scholar CrossRef Search ADS PubMed  Pereira RR, Castanheira D, Teixeira JA et al.   Detailed search for protein kinase(s) involved in plasma membrane H+-ATPase activity regulation of yeast cells. FEMS Yeast Res  2015; 15: fov003. Google Scholar CrossRef Search ADS PubMed  Portillo F. Regulation of plasma membrane H+-ATPase in fungi and plants. Biochim Biophys Acta  2000; 1469: 31– 42. Google Scholar CrossRef Search ADS PubMed  Portillo F, Eraso P, Serrano R. Analysis of the regulatory domain of yeast plasma membrane H+-ATPase by directed mutagenesis and intragenic suppression. FEBS Lett  1991; 287: 71– 4. Google Scholar CrossRef Search ADS PubMed  Rao R, Drummond-Barbosa D, Slayman CW. Transcriptional regulation by glucose of the yeast PMA1 gene encoding the plasma membrane H+-ATPase. Yeast  1993; 9: 1075– 84. Google Scholar CrossRef Search ADS PubMed  Saiardi A, Caffrey JJ, Snyder SH et al.   Inositol polyphosphate multikinase (ArgRIII) determines nuclear mRNA export in Saccharomyces cerevisiae. FEBS Lett  2000; 468: 28– 32. Google Scholar CrossRef Search ADS PubMed  Saiardi A, Erdjument-Bromage H, Snowman AM et al.   Synthesis of diphosphoinositol pentakisphosphate by a newly identified family of higher inositol polyphosphate kinases. Curr Biol  1999; 9: 1323– 6. Google Scholar CrossRef Search ADS PubMed  Secades P, Guijarro JA. Purification and characterization of an extracellular protease from the fish pathogen Yersinia ruckeri and effect of culture conditions on production. Appl Environ Microb  1999; 65: 3969– 75. Serrano R. Structure, function and regulation of plasma membrane H+-ATPase. FEBS Lett  1993; 325: 108– 11. Google Scholar CrossRef Search ADS PubMed  Serrano R, Ruiz A, Bernal D et al.   The transcriptional response to alkaline pH in Saccharomyces cerevisiae: evidence for calcium-mediated signalling. Mol Microbiol  2002; 46: 1319– 33. Google Scholar CrossRef Search ADS PubMed  Shears SB. Transcriptional regulation: a new dominion for inositol phosphate signaling? Bioessays  2000; 22: 786– 9. Google Scholar CrossRef Search ADS PubMed  Souza M, Tropia M, Brandao R. New aspects of the glucose activation of the H+-ATPase in the yeast Saccharomyces cerevisiae. Microbiology  2001; 147: 2849– 55. Google Scholar CrossRef Search ADS PubMed  Thoms S, Debelyy MO, Nau K et al.   Lpx1p is a peroxisomal lipase required for normal peroxisome morphology. FEBS J  2008; 275: 504– 14. Google Scholar CrossRef Search ADS PubMed  Thoms S, Hofhuis J, Thöing C et al.   The unusual extended C-terminal helix of the peroxisomal α/β-hydrolase Lpx1 is involved in dimer contacts but dispensable for dimerization. J Struct Biol  2011; 175: 362– 71. Google Scholar CrossRef Search ADS PubMed  Tisi R, Baldassa S, Belotti F et al.   Phospholipase C is required for glucose-induced calcium influx in budding yeast. FEBS Lett  2002; 520: 133– 8. Google Scholar CrossRef Search ADS PubMed  Tisi R, Belotti F, Wera S et al.   Evidence for inositol triphosphate as a second messenger for glucose-induced calcium signalling in budding yeast. Curr Genet  2004; 45: 83– 9. Google Scholar CrossRef Search ADS PubMed  Tisi R, Martegani E, Brandão RL. Monitoring yeast intracellular Ca2+ levels using an in vivo bioluminescence assay. Cold Spring Harb Protoc  2015; 2015: 210– 3. Google Scholar PubMed  Trópia M, Cardoso A, Tisi R et al.   Calcium signaling and sugar-induced activation of plasma membrane H+-ATPase in Saccharomyces cerevisiae cells. Biochem Biophys Res Commun  2006; 343: 1234– 43. Google Scholar CrossRef Search ADS PubMed  Wera S, Bergsma JCT, Thevelein JM. Phosphoinositides in yeast: genetically tractable signalling. FEMS Yeast Res  2001; 1: 9– 13. Google Scholar CrossRef Search ADS PubMed  Wu S, Liang MP, Altman RB. The SeqFEATURE library of 3D functional site models: comparison to existing methods and applications to protein function annotation. Genome Biol  2008; 9: R8. Google Scholar CrossRef Search ADS PubMed  York JD, Odom AR, Murphy R et al.   A phospholipase C-dependent inositol polyphosphate kinase pathway required for efficient messenger RNA export. Science  1999; 285: 96– 100. Google Scholar CrossRef Search ADS PubMed  © FEMS 2017. All rights reserved. For permissions, please e-mail: journals.permissions@oup.com http://www.deepdyve.com/assets/images/DeepDyve-Logo-lg.png FEMS Yeast Research Oxford University Press

Loading next page...
 
/lp/ou_press/lpx1p-links-glucose-induced-calcium-signaling-and-plasma-membrane-h-jDpuc3pItj
Publisher
Blackwell
Copyright
© FEMS 2017. All rights reserved. For permissions, please e-mail: journals.permissions@oup.com
ISSN
1567-1356
eISSN
1567-1364
D.O.I.
10.1093/femsyr/fox088
Publisher site
See Article on Publisher Site

Abstract

Abstract In yeast, as in other eukaryotes, calcium plays an essential role in signaling transduction to regulate different processes. Many pieces of evidence suggest that glucose-induced activation of plasma membrane H+-ATPase, essential for yeast physiology, is related to calcium signaling. Until now, no protein that could be regulated by calcium in this context has been identified. Lpx1p, a serine-protease that is also involved in the glucose-induced activation of the plasma membrane H+-ATPase, could be a candidate to respond to intracellular calcium signaling involved in this process. In this work, by using different approaches, we obtained many pieces of evidence suggesting that the requirement of calcium signaling for activation of the plasma membrane H+-ATPase is due to its requirement for activation of Lpx1p. According to the current model, activation of Lpx1p would cause hydrolysis of an acetylated tubulin that maintains the plasma membrane H+-ATPase in an inactive state. Therefore, after its activation, Lpx1p would hydrolyze the acetylated tubulin making the plasma membrane H+-ATPase accessible for phosphorylation by at least one protein kinase. calcium signaling, plasma membrane H+-ATPase, Lpx1, Saccharomyces cerevisiae INTRODUCTION Plasma membrane H+-ATPase is an essential enzyme for yeast physiology since by pumping protons out of cells it creates the electrochemical potential that allows cells to accumulate nutrients against their concentration gradients and to control intracellular pH (Goffeau and Slayman 1981). This enzyme is regulated both at the transcriptional (Capieaux et al.1989; Rao, Drummond-Barbosa and Slayman 1993) and at the post-translational (Serrano 1993) levels on glucose addition, this sugar being the most important trigger for its regulation. Originally, it was proposed that H+-ATPase post-translational activation would be caused by phosphorylation catalyzed by protein kinases that would have as targets two phosphorylation sites at the plasma membrane H+-ATPase C-terminal tail (Portillo 2000). According to this model, phosphorylation of residue Ser-899 would be responsible for a decrease in Km for ATP, and phosphorylation of Thr-912 would lead to an increase of Vmax related to ATP hydrolysis (Portillo, Eraso and Serrano 1991). Later, it was demonstrated that the second phosphorylation site includes another serine residue (Ser-911) that would also be associated with glucose addition (Lecchi et al.2005, 2007). Moreover, it was confirmed that phosphorylation of both residues (Ser-911 and Thr-912) seems to be connected to the activation process (Mazón, Eraso and Portillo 2015). Although it has been shown that glucose-induced activation of the plasma membrane H+-ATPase is caused by phosphorylation, the search for protein kinases involved in this process led, up to now, to the identification of Ptk2p as a unique protein kinase involved with the phosphorylation of the Ser-899 residue at the C-terminal tail (Goossens et al.2000; Eraso, Mazón and Portillo 2006). Recently, a wide screening of protein kinases present in yeast cells reconfirmed the essential involvement of Ptk2p in the glucose-induced activation of the plasma membrane H+-ATPase. Thus, these data indicate that the current model suggesting the existence of two phosphorylation sites (and another protein kinase indeed involved in this process, besides Ptk2p) seems to be incorrect or, at least, incomplete (Pereira et al.2015). Indeed, new elements have been introduced in this scenario confirming the complexity of the mechanism responsible for plasma membrane H+-ATPase post-translational activation. Over the years, many results have suggested the existence of a clear relationship between calcium signaling and plasma membrane H+-ATPase activation (Coccetti et al.1998; Tisi et al.2002, 2004; Trópia et al.2006; Pereira et al.2008; Groppi et al.2011; Bouillet et al.2012). Saccharomyces cerevisiae as a eukaryotic organism can use calcium-signaling pathways to control different cellular processes (Anraku, Ohya and Iida 1991; Paidhungat and Garrett 1997; Denis and Cyert 2002; Serrano et al.2002). In the cytosol of yeast cells, the free Ca2+ concentration is strictly regulated (Miseta et al.1999b) by the action of a variety of transporters, channels, pumps and co-transporters (Cyert and Philpott 2013). Ca2+ in yeast is retained mainly in the vacuole (Dunn, Gable and Beeler 1994; Miseta et al.1999a) and the vacuolar pool is maintained mostly by the action of Pmc1p (a Ca2+-ATPase) and Vcx1p (a Ca2+/H+ exchanger). The release of calcium stored in vacuoles is made through the calcium channel Yvc1p, which has its activity controlled by different cellular stimuli (Cunningham 2011). Apparently, the connection between calcium signaling and H+-ATPase activation starts when glucose-induced phosphatidylinositol-4,5-bisphosphate hydrolysis occurs. Mediated by phospholipase C (encoded by the PLC1 gene), this hydrolysis produces two intracellular messengers: diacylglycerol and inositol 1,4,5-trisphosphate (IP3) (York et al.1999; Shears 2000). This Plc1p activation would be initiated in response to a stimulus from protein G (Gpa2p), which in turn would be activated in response to glucose uptake and its phosphorylation by sugar kinases (Bouillet et al.2012). In turn, the level of IP3 is regulated by its phosphorylation by Arg82p, an inositol kinase, generating two types of inositol tetrakisphosphate—I(1,3,4,5)P4 and I(1,4,5,6)P4—and an inositol pentaphosphate, I(1,3,4,5,6)P5 (Saiardi et al.1999; Odom et al.2000). A possible relationship between IP3 and the vacuolar Ca2+ channel, Yvc1p, was demonstrated previously, suggesting that this channel could participate in a mechanism involved in intracellular calcium level control in response to glucose addition. IP3 would interact directly or indirectly with Yvc1p, regulating the intensity of calcium signaling in the cytosol (Bouillet et al.2012). In arg82Δ yeast strains, where there is no conversion of IP3 into IP4 and/or IP5, glucose-induced increase of IP3, H+-ATPase activation and calcium signaling are more pronounced (Tisi et al.2004). At low external calcium concentration, the vacuolar calcium channel Yvc1p’s activity seems to become more important for proper glucose-induced calcium signaling and plasma membrane H+-ATPase activation. Moreover, there are many pieces of evidence suggesting that the intensity of the intracellular calcium signal, in these conditions, is strongly dependent on the action of Yvc1p (Bouillet et al.2012) and that there is a connection between the IP3 signal, Yvc1 activity, calcium signaling and glucose-induced activation of the plasma membrane H+-ATPase in yeast cells (Tisi et al.2004; Trópia et al.2006). Nevertheless, since no IP3 receptor homolog has been identified in the S. cerevisiae genome (Wera, Bergsma and Thevelein 2001), the existence of such a transduction pathway is still a matter of controversy (Bouillet et al.2012). On the other hand, some data suggest the involvement of a multifunctional enzyme in glucose-induced activation of the plasma membrane H+-ATPase (Campetelli et al.2013). By this model, glucose addition would trigger activation of proteolytic activity of Lpx1p, which degrades the tubulin bound to the inactive form of plasma membrane H+-ATPase at its C-terminal tail. This degradation releases the C-terminal tail leading to the exposure of sites that are targets of phosphorylation, causing H+-ATPase activation (Campetelli et al.2005, 2013). Considering these two sets of evidence for H+-ATPase activation, calcium involvement and Lpx1p dependence, and the apparent difficulty in finding other protein kinase(s), mainly the calcium-dependent ones, involved in enzyme phosphorylation/activation, we explored a possible relationship between calcium signaling and Lpx1p activation in connection with glucose-induced regulation of the plasma membrane H+-ATPase. Here, we demonstrated that calcium signaling is indeed directly related to Lpx1p proteolytic action, and this activation process is clearly related to the plasma membrane H+-ATPase. MATERIAL AND METHODS Strains and growth conditions Saccharomyces cerevisiae strains shown in Table 1 were grown in media containing 2% peptone and 1% yeast extract (YP) supplemented with appropriate carbon sources or in SD medium—0.67% yeast nitrogen base without amino acids (Difco, Detroit, MI, USA) supplemented with drop-out amino-acids (leucine, tryptophan, histidine and methionine) and nucleotide (uracil) and 2% glucose. Cells were grown in a rotatory incubator shaker - New Brunswick Model 25 (GMI, Ramsey, MN, USA) at 200 rpm and 30°C until the end of the logarithmic phase (OD600nm ∼ 2.0). Alternatively, and when requested, yeast cells were grown in induction medium (SD medium supplemented with 8% galactose). Table 1. Saccharomyces cerevisiae strains used in this study. Strain  Genotype  Source  PJ69-2A  MATa trp1-901 leu2-3, 112 ura3-52 his3-200 gal4Δ gal80Δ LYS2::GAL1-HIS3 GAL2-ADE2 met2::GAL7-lacZ  James Caffrey (Saiardi et al.2000)  PJ69-4A1  PJ69-2A arg82::KanMX2  James Caffrey (Saiardi et al.2000)  LBCM 1630  PJ69-2A yvc1::KanMX2  Bouillet et al. (2012)  LBCM 1713  LBCM 1630; lpx1::LEU2  This work  LBCM 1714  PJ69-4A1; lpx1::LEU2  This work  BY4741  MATa his3Δ1 leu2Δ0 lis2Δ0 ura3Δ0  EUROSCARF  BY4741 lpx1Δ  BY4741; lpx1::KanMX4  EUROSCARF  BY4741 ptk2Δ  BY4741; ptk2::KanMX4  EUROSCARF  LBCM 1738  BY4741 lpx1Δ + pYES2/CT-EV  This work  LBCM 1739  BY4741 lpx1Δ + pYES2/CT-LPX1  This work  LBCM 1764  BY4741 lpx1Δ + pYES2/CT-LPX1-MOD  This work  Strain  Genotype  Source  PJ69-2A  MATa trp1-901 leu2-3, 112 ura3-52 his3-200 gal4Δ gal80Δ LYS2::GAL1-HIS3 GAL2-ADE2 met2::GAL7-lacZ  James Caffrey (Saiardi et al.2000)  PJ69-4A1  PJ69-2A arg82::KanMX2  James Caffrey (Saiardi et al.2000)  LBCM 1630  PJ69-2A yvc1::KanMX2  Bouillet et al. (2012)  LBCM 1713  LBCM 1630; lpx1::LEU2  This work  LBCM 1714  PJ69-4A1; lpx1::LEU2  This work  BY4741  MATa his3Δ1 leu2Δ0 lis2Δ0 ura3Δ0  EUROSCARF  BY4741 lpx1Δ  BY4741; lpx1::KanMX4  EUROSCARF  BY4741 ptk2Δ  BY4741; ptk2::KanMX4  EUROSCARF  LBCM 1738  BY4741 lpx1Δ + pYES2/CT-EV  This work  LBCM 1739  BY4741 lpx1Δ + pYES2/CT-LPX1  This work  LBCM 1764  BY4741 lpx1Δ + pYES2/CT-LPX1-MOD  This work  View Large Table 1. Saccharomyces cerevisiae strains used in this study. Strain  Genotype  Source  PJ69-2A  MATa trp1-901 leu2-3, 112 ura3-52 his3-200 gal4Δ gal80Δ LYS2::GAL1-HIS3 GAL2-ADE2 met2::GAL7-lacZ  James Caffrey (Saiardi et al.2000)  PJ69-4A1  PJ69-2A arg82::KanMX2  James Caffrey (Saiardi et al.2000)  LBCM 1630  PJ69-2A yvc1::KanMX2  Bouillet et al. (2012)  LBCM 1713  LBCM 1630; lpx1::LEU2  This work  LBCM 1714  PJ69-4A1; lpx1::LEU2  This work  BY4741  MATa his3Δ1 leu2Δ0 lis2Δ0 ura3Δ0  EUROSCARF  BY4741 lpx1Δ  BY4741; lpx1::KanMX4  EUROSCARF  BY4741 ptk2Δ  BY4741; ptk2::KanMX4  EUROSCARF  LBCM 1738  BY4741 lpx1Δ + pYES2/CT-EV  This work  LBCM 1739  BY4741 lpx1Δ + pYES2/CT-LPX1  This work  LBCM 1764  BY4741 lpx1Δ + pYES2/CT-LPX1-MOD  This work  Strain  Genotype  Source  PJ69-2A  MATa trp1-901 leu2-3, 112 ura3-52 his3-200 gal4Δ gal80Δ LYS2::GAL1-HIS3 GAL2-ADE2 met2::GAL7-lacZ  James Caffrey (Saiardi et al.2000)  PJ69-4A1  PJ69-2A arg82::KanMX2  James Caffrey (Saiardi et al.2000)  LBCM 1630  PJ69-2A yvc1::KanMX2  Bouillet et al. (2012)  LBCM 1713  LBCM 1630; lpx1::LEU2  This work  LBCM 1714  PJ69-4A1; lpx1::LEU2  This work  BY4741  MATa his3Δ1 leu2Δ0 lis2Δ0 ura3Δ0  EUROSCARF  BY4741 lpx1Δ  BY4741; lpx1::KanMX4  EUROSCARF  BY4741 ptk2Δ  BY4741; ptk2::KanMX4  EUROSCARF  LBCM 1738  BY4741 lpx1Δ + pYES2/CT-EV  This work  LBCM 1739  BY4741 lpx1Δ + pYES2/CT-LPX1  This work  LBCM 1764  BY4741 lpx1Δ + pYES2/CT-LPX1-MOD  This work  View Large Molecular biology methods and in silico analysis Yeast cells were transformed by using the lithium acetate protocol (Gietz et al.1995). A pVTU-apoaequorin plasmid (pVTU-AEQ) was generated by inserting in pVTU XhoI–PstI sites the XhoI–PstI-digested fragment obtained by PCR on pYX212-AEQ (Tisi et al.2002) using the oligonucleotides 5΄-TTTCTCGAGAATCTATAACTACAAAAAACACATACAGGAA-3΄ and 5΄-TAACTGCAGGCCCTAGGATCCATGGTGAA-3΄. The LPX1 gene was obtained as previously described (Campetelli et al.2013) and then inserted into the pYES2/CT plasmid (Invitrogen, Carlsbad, CA, USA) allowing inducible expression by galactose of recombinant proteins fused with a 6× His-tag at the C-terminus. For this purpose, plasmid and amplified LPX1 gene were previously digested with KpnI and XbaI enzymes (see Supplementary Material 1) (Fermentas, Waltham, MA, USA). Digested plasmid was treated with shrimp alkaline phosphatase (SAP) (Fermentas) for 1 h at 37°C prior to ligation. The SAP-treated plasmid was used for the T4 ligase reaction (Sigma-Aldrich, St Louis, MO, USA) with LPX1 insert. This ligation reaction was performed at 23°C for 1 h, and Escherichia coli strain TOP10 was used for plasmid replication. BY4741 lpx1Δ cells were transformed with pYES2/CT + LPX1 (Gietz et al.1995) and selected in SD medium with 2.5% agar—from here on denominated as pYES2/CT-LPX1. The presence of the gene insertion into the plasmid and correct transformation in bacteria and yeast were confirmed by PCR (forward primer: 5΄-AAAGGTACCATGGAACAGAACAGGTCC-3΄; reverse primer: 5΄-TTTTCTAGATTACAGTTTTTGTTTAGTCG-3΄). BY4741 lpx1Δ cells were transformed with an empty vector (here denominated as pYES2/CT-EV) as well, to be used as the control treatment. In order to find regions in Lpx1p protein structure that could interact with Ca2+, the WebFEATURE 4.0 server (Liang et al.2003; Wu, Liang and Altman 2008) was run using a deposited structure (crystal form II; trigonal) of Lpx1p from S. cerevisiae (PDB access number: 2y6u) (Thoms et al.2011) with settings defined to search for all functional sites possibly present in this protein (Supplementary Material 2). To verify the influence of the C-terminal region of Lpx1p that would interact with Ca2+ in Lpx1p activation, this region was removed by treating the LPX1 gene with PvuII enzyme that cuts at 909 bp after the initiation codon. The modified gene was cloned into the pYES2/CT plasmid (see Supplementary Material 3). Digested LPX1 gene and pYES2/CT plasmid were then treated with KpnI. Digested plasmid was used for the T4 ligase reaction (Sigma-Aldrich) with modified LPX1 insert. This ligation reaction was performed at 23°C overnight and E. coli strain TOP10 was used for plasmid replication. BY4741 lpx1Δ cells were transformed with pYES2/CT + modified LPX1 (Gietz et al.1995) and selected in SD medium with 2.5% agar—from here on denominated as pYES2/CT-LPX1-MOD. The presence of the gene insertion into the plasmid and correct transformation in bacteria and yeast were confirmed by PCR (forward primer: 5΄-AAAGGTACCATGGAACAGAACAGGTCC-3΄; reverse primer: 5΄-GCGTGAATGTAAGCGTGAC-3΄), by comparing different amplicon sizes between pYES2/CT-LPX1 and pYES2/CT-LPX1-MOD. Measurement of cytosolic free calcium by bioluminescence assay By using a standard method, cytosolic free calcium was measured by using the aequorin-based bioluminescence assay (Tisi et al.2002; Brandão 2014; Tisi, Martegani and Brandão 2015). Yeast strains containing the apoaequorin-expressing plasmid pVTU-AEQ were grown in rich medium (YP with 2% glucose) until they entered the exponential phase. Cells were harvested and washed by filtration with sterile water and resuspended in MES/Tris 0.1 M (pH 6.5). After 90 min of incubation at room temperature, cells were loaded with 50 μM coelenterazine (Sigma-Aldrich, St. Louis, MO, USA) for 20 min. For removal of excess coelenterazine, cells were washed twice by centrifugation (2000 g, 5 min) with MES/Tris 0.1 M pH 6.5. Glucose-induced aequorin luminescence was measured in a Lumat LB 9507 luminometer (Berthold Technologies, Bad Wildbad, Germany) at intervals of 10 s for 1 min before and for at least 10 min after addition of 100 mM glucose (final concentration). Lpx1p His-tag expression and purification To obtain cells for Lpx1p His-tag extraction, the lpx1Δ mutant transformed with pYES2/CT-LPX1, and pYES2/CT-LPX1-MOD plasmids were grown in SD medium for 24 h. Cells were collected, washed three times by centrifugation (2000 g, 5 min) with sterile water and resuspended in induction medium (SD medium supplemented with 8% galactose). After overnight incubation in a rotatory incubator shaker (New Brunswick Model 25) at 200 rpm and 30°C, cells were harvested by vacuum filtration on glass fiber filters, immediately frozen in liquid nitrogen and stored until use. Cells were resuspended in lysis buffer (50 mM Na3PO4, 300 mM NaCl and 5 mM β-mercaptoethanol), disrupted by vigorous shaking with glass beads (with 90 s intervals on ice for five times) and then centrifuged at 2000 g for 10 min. The resulting supernatant (crude extract) was incubated with HIS-Select nickel affinity gel (Sigma-Aldrich, St. Louis, MO, USA) for 1 h at 4°C with gentle agitation. Unbound proteins were removed by discarding supernatant after centrifugation at 1000 g for 5 min. Proteins bound to resin were eluted with elution buffer (50 mM Na3PO4, 300 mM NaCl, 5 mM β-mercaptoethanol). Induction and protein purification were performed with pYES2/CT-EV strain, to be used as the control treatment. Aliquots (50 μg) of protein fractions were separated by SDS-PAGE using 12% (w/v) polyacrylamide gels. Gels were stained with silver staining and images were captured by Image Scanner software (Thermo Fisher Scientific, Waltham, MA, USA). The molecular masses of the proteins were calculated by interpolating their specific relative migration distances (Rf) in a curve prepared with correspondent values of appropriated molecular mass markers (Supplementary Material 4). Western blotting assays were performed with eluates obtained from cell extracts of the lpx1Δ mutant transformed with pYES2/CT-EV, pYES2/CT-LPX1 and pYES2/CT-LPX1-MOD constructions. Protein content was determined by using the Lowry method (Lowry et al.1951). Next, the proteins were transferred to nitrocellulose membranes (Thermo Fisher Scientific, Waltham, MA, USA) according to the recommendations of the Bio-Rad Transference Kit (Bio-Rad Laboratories, Hercules, CA, USA). Membranes were blocked for 1 h at room temperature, using PBS containing 0.1% Tween-20 and 5% skimmed milk powder. Furthermore, membranes were incubated with mouse monoclonal primary antibody anti-His (Invitrogen, Carlsbad, CA, USA). Subsequently, membranes were washed and incubated with secondary antibody horseradish peroxidase-goat anti-mouse IgG (H + L) (Invitrogen, Carlsbad, CA, USA). Immunolabeling was visualized using WESTAR Nova 2.0 (Cyanagen, Bologna, Italy) as per the manufacturer’s instructions. Images were captured by Image Scanner software (Amershan Biosciences, Bath, UK). Measurement of proteolytic Lpx1p activity Calcium dependence of the proteolytic activity of the wild-type and lpx1Δ cells was assayed by using azocasein (Thermo Fisher Scientific, Waltham, MA, USA) as substrate (Charney and Tomarelli 1947). Briefly, samples of total membranes containing 50–100 μg protein were added to a reaction mixture (100 mM Tris–HCl, pH 8.0, 0.5% azocasein), with or without the addition of 1 mM calcium, in a final volume of 500 μL. The mixture was kept for 1 h at 40°C, and after this time, the reaction was stopped by adding 250 μL of 10% w/v trichloroacetic acid. The mixture was centrifuged (2000 g, 5 min) to remove coagulated proteins and 500 μL of supernatant was neutralized with 500 μL 5 M KOH, and the absorbance at 428 nm was measured using a BioMate 3 spectrophotometer (Thermo Fisher Scientific, Waltham, MA, USA). A change in absorbance of 0.01 (428 nm) per minute, resulting from azocasein hydrolysis, was defined as one arbitrary unit (AU). The control was obtained by mixing trichloroacetic acid with the azocasein solution prior to the addition of the protein sample (Secades and Guijarro 1999). Protein content was determined by using the Lowry method (Lowry et al.1951). An additional Lpx1p activity evaluation was performed with eluates obtained from the Ni-affinity column from lpx1Δ mutant transformed with pYES2/CT-LPX1. Samples containing 100 μg protein were dissolved in non-reducing Laemmli buffer (62.5 mM Tris–HCl, pH 6.8, 10% w/v glycerol, 0.001% w/v bromophenol blue) and run at 100 V at 4°C in 8% (w/v) polyacrylamide gel. After electrophoresis, the corresponding Lpx1p bands were excised from the gel and washed three times in 100 mM Tris–HCl buffer, pH 8.0, with gentle agitation, in order to remove excess SDS from the running buffer. Then, they were added to a reaction mixture (100 mM Tris–HCl, pH 8.0, 0.5% azocasein), containing 100 μM calcium, in a final volume of 500 μL and the assay was performed as described previously in this work. A negative control was obtained by using the corresponding material from lpx1Δ mutant transformed with pYES2/CT-EV. Measurement of plasma membrane H+-ATPase activity In vivo activation of plasma membrane H+-ATPase was detected by using two different standard approaches: indirectly, measuring glucose-induced proton-pumping activity, or directly, in purified plasma membranes by following ATP hydrolysis. In the first case, around 500 mg (wet weight) of cells grown in YP medium (with 2% glucose) was resuspended in 100 mM Tris–HCl buffer pH 4.5 containing 100 mM KCl and incubated in a vessel in a total volume of 5.0 mL. Changes in pH suspension were recorded before and after addition of 100 mM glucose (final concentration). Calibration pulses of 100 nmol HCl were also added and used as reference in calculations of proton pumping. The maximal rate of proton pumping was calculated from the slope of the line indicating the pH variation in the medium, as described previously, and indicated in mmol H+ h−1 g−1 cell (Souza, Tropia and Brandao 2001). To measure proton-pumping activity from lpx1Δ or yvc1Δ mutants transformed with pYES2/CT-EV and pYES2/CT-LPX1 vectors, cells grown in SD medium were washed twice by centrifugation (2000 g, 5 min) with 100 mM Tris–HCl buffer pH 4.5 containing 100 mM KCl. After washing, cells were incubated in the same buffer containing 100 mM galactose (final concentration) prior to proceeding with the assay to promote the induction of the LPX1 gene. After incubation with 100 mM galactose for different times (10, 30 or 50 min), cells were washed twice by centrifugation (2000 g, 5 min) with 100 mM Tris–HCl buffer pH 4.5 containing 100 mM KCl, and then used to measure in vivo proton-pumping activity as described (Souza, Tropia and Brandao 2001). Cells without incubation with 100 mM galactose were used as controls. In the second approach, to measure directly the H+-ATPase activity, lpx1Δ mutant transformed with pYES2/CT-EV and pYES2/CT-LPX1 vectors was collected after overnight incubation in induction medium (SD medium supplemented with 8% galactose) in a rotatory incubator shaker (New Brunswick Model 25) at 200 rpm and 30°C, prior to proceeding with plasma membrane purification. Cells were washed twice by centrifugation (2000 g, 5 min) with 100 mM MES/Tris buffer (pH 6.5), collected on glass fiber filters by vacuum filtration, immediately frozen in liquid nitrogen and stored until use. Cells were disrupted in 100 mM Tris buffer (supplemented with 330 mM sorbitol and 5 mM β-mercaptoethanol), by vigorous shaking with glass beads, with 90 s intervals on ice five times and centrifuged at 2000 g for 10 min. Purified plasma membranes were obtained essentially as described before (dos Passos et al.1992), with resuspension of plasma membranes in glycerol buffer (20% w/v glycerol, 10 mM Tris, 1 mM β-mercaptoethanol). Protein content was determined by using the Lowry method (Lowry et al.1951). ATPase activity was determined as described previously (dos Passos et al.1992). On the other hand, and to study a direct connection between calcium availability, Lpx1p activity and plasma membrane H+-ATPase regulation, we tried to reconstitute in vitro the plasma membrane H+-ATPase activation. First, to extract plasma membranes, wild-type cells were resuspended in 100 mM MES/Tris buffer (pH 6.5) at a density of 150 mg mL−1 (wet mass) and incubated in a shaking water bath at 30°C. After 4 h, samples (glucose-starved cells) were collected on glass fiber filters by vacuum filtration and immediately frozen in liquid nitrogen and stored until use. The procedures used to obtain plasma membranes were performed as described previously (dos Passos et al.1992), without addition of EDTA in lysis and glycerol buffers. Briefly, to obtain a membrane-free extract (MFE), cells were suspended in 100 mM Tris buffer without protease inhibitors (supplemented with 330 mM sorbitol and 5 mM β-mercaptoethanol), disrupted by vigorous shaking with glass beads with 90 s intervals on ice five times, and centrifuged at 2000 g for 10 min. The supernatant was then centrifuged at 100 000 g for 30 min. After ultracentrifugation, a new supernatant yielded membrane-free extracts (from here on denominated as MFE) from all strains; these as well as HIS-Select nickel affinity gel eluates resulting from the application of crude extract of lpx1Δ mutant cells transformed with pYES2/CT-EV, pYES2/CT-LPX1 and pYES2/CT-LPX1-MOD were subsequently submitted to overnight dialysis at 4°C with incubation buffer (50 mM HEPES, 10 mM sodium azide, 5 mM ammonium molybdate). This procedure was necessary in order to remove any interfering compound for in vitro H+-ATPase activity assay. Protein content was determined by using the Lowry method (Lowry et al.1951). In vitro H+-ATPase activation was performed by resuspending plasma membrane samples (40 μg protein) obtained from glucose-starved wild-type cells in an incubation buffer (50 mM HEPES, 10 mM sodium azide, 5 mM ammonium molybdate). Then, 100 μM Ca2+ and/or MFE (50 μg protein) was added (or not) according to the different tests performed, to a final volume of 250 μL (more details in Results section). In all samples, 1 mM MgCl2 (final concentration) was also added. The samples were incubated for 30 min at 30°C in a water bath, for in vitro H+-ATPase activation. Different calcium concentrations (100 μM, 1 and 5 mM), different times of incubation (10, 30 and 60 min) and different MFE protein concentrations (50, 100, 200 and 400 μg) for in vitro H+-ATPase activation were tested. Sodium orthovanadate of 50 μM was added as well to verify H+-ATPase activity inhibition. MFE and HIS-Select nickel affinity gel eluates were assayed to check for any ATPase activity as control. All results are expressed as percentages relative to their related controls. To assay H+-ATPase activity, the reactions were started with concentrated ATP solution (Sigma-Aldrich) to obtain 2 mM as final concentration. ATPase activity was determined as described previously (dos Passos et al.1992) and appropriate controls were performed for each assay (see Results section for specific details). Statistical analysis The experiments were performed at least three times. Standard deviations were indicated in each figure or table. Statistical analyses were performed by using Student's t-test or ANOVA. Differences were considered statistically significant when the P value was <0.05. RESULTS Lpx1p influences H+-ATPase activation, but not intracellular calcium concentration It was previously suggested that in glucose-induced activation of yeast plasma membrane H+-ATPase, calcium signaling is a critical factor (Souza, Tropia and Brandao 2001; Tisi et al.2004; Trópia et al.2006; Pereira et al.2008; Bouillet et al.2012). However, in the corresponding signal transduction pathway a component that actually would respond to the intracellular calcium signal was never found. Since the Lpx1p protein also participates in this process (Campetelli et al.2013), we wondered whether this protease could be a possible candidate that would respond to the calcium signal, thereby explaining its function in plasma membrane H+-ATPase glucose-induced activation. Indeed, as demonstrated in Fig. 1A, plasma membrane H+-ATPase glucose-induced activation (measured indirectly by proton-pumping rate) was reduced in strains presenting deletion in the LPX1 gene. Comparable results (i.e. reduced H+-ATPase glucose-induced activation) were also observed in yvc1Δ strain (absence of Yvc1p reduces glucose-induced calcium signaling by preventing calcium release from the vacuole, which reduces H+-ATPase activation). On the other hand, deletion of ARG82 causes a clear stronger glucose-induced activation of the plasma membrane H+-ATPase compared with wild-type cells, since these cells present high IP3 levels in cytoplasm, which enhances calcium release from vacuole leading to an increase in the H+-ATPase activity. However, when LPX1 deletion was also included in this arg82Δ mutant, H+-ATPase activity was reduced as observed in the cell presenting only the lpx1Δ mutation. The same standard was also observed in a strain presenting deletions in the YVC1 and LPX1 genes, i.e. H+-ATPase activity showed comparable low levels to yvc1Δ cells. The same was observed in the double mutant yvc1Δ lpx1Δ (Fig. 1A). Figure 1. View largeDownload slide Effects of different gene deletions on H+-ATPase proton-pumping activity and intracellular calcium signaling in S. cerevisiae cells (genetic background PJ69-2A). (A) Proton-pumping activity was measured with around 500 mg of cells, after addition of 100 mM glucose (final concentration). (B) Cytosolic free calcium was measured by using a standard aequorin-based bioluminescence assay after addition of 100 mM glucose (final concentration), indicated by a striped arrow at ‘0’ s, in yeast strains containing apoaequorin-expressing plasmid pVTU-AEQ and loaded with 50 μM coelenterazine. Different letters indicate that mean values are statistically different (P < 0.05). RLUs/s: relative luminescence units per second. Figure 1. View largeDownload slide Effects of different gene deletions on H+-ATPase proton-pumping activity and intracellular calcium signaling in S. cerevisiae cells (genetic background PJ69-2A). (A) Proton-pumping activity was measured with around 500 mg of cells, after addition of 100 mM glucose (final concentration). (B) Cytosolic free calcium was measured by using a standard aequorin-based bioluminescence assay after addition of 100 mM glucose (final concentration), indicated by a striped arrow at ‘0’ s, in yeast strains containing apoaequorin-expressing plasmid pVTU-AEQ and loaded with 50 μM coelenterazine. Different letters indicate that mean values are statistically different (P < 0.05). RLUs/s: relative luminescence units per second. In the same set of strains and in similar conditions, the influence of LPX1 deletion in calcium signaling was assessed (Fig. 1B). Initially, it was observed that the strain with single LPX1 deletion showed calcium signaling comparable to wild-type cells. When LPX1 was deleted in combination with ARG82, there was a stronger intracellular calcium signal, as observed in arg82Δ mutant cells. Calcium signaling was reduced in yvc1Δ cells and in the double mutant yvc1Δ lpx1Δ strain. Therefore, the results shown in Fig. 1 suggest that Lpx1p is not necessary for generation of calcium signal on glucose treatment; however, it is required for calcium-mediated activation of the H+-ATPase. Thus, Lpx1p should work downstream of IP3 and calcium signaling and, consequently, can form a link between the calcium signal and the H+-ATPase function. Moreover, this hypothesis was also reinforced by the fact that a database search, using Lpx1p crystal structure, indicated the presence of a potential region for calcium interaction with a high-confidence prediction (95% precision performance) located at its C-terminus (Supplementary Material 2), suggesting that this domain would be an EF-hand calcium-binding domain. Overexpression of LPX1 in an lpx1Δ mutant restores normal proton-pumping rate and increases H+-ATPase activity An lpx1Δ mutant yeast strain expressing the LPX1 gene fused with a His-tag at the 3΄-extremity in a galactose-inducible vector (pYES2/CT-LPX1) was generated to verify the potential connection between calcium signaling and Lpx1p activation. As control, the lpx1Δ yeast strain was transformed with the corresponding empty vector (pYES2/CT-EV). In order to check the suitability of these constructions, transformant yeasts were submitted to galactose induction for different incubation times (10, 30 and 50 min for pYES2/CT-LPX1 and 50 min for pYES2/CT-EV). After induction, glucose-induced extracellular acidification was performed. As shown in Fig. 2A, cells expressing the LPX1 gene presented an increase in proton-pumping rates after galactose induction (P < 0.05), when compared with non-induced cells. After 10, 30 and 50 min, proton-pumping rate was increased to 58.0%, 87.3% and 112.4%, respectively, compared with cells without galactose induction. Proton-pumping activity after 50 min induction in pYES2/CT-LPX1 cells was comparable to the wild-type strain (P > 0.05). Cells transformed with the empty plasmid did not exhibit any increase in proton-pumping rate after 50 min galactose induction, showing the same rate as found in the lpx1Δ strain (P > 0.05). Interestingly, when the yvc1Δ  mutant was also transformed with such constructions and exposed to galactose, there was no increase in the H+-pumping rate even after 50 min, with the activity remaining comparable to that registered for the yvc1Δ  mutant (Fig. 2A). These results suggest that the activity of the calcium channel Yvc1 is not controlled by Lpx1p. Figure 2. View largeDownload slide Plasma membrane proton-pumping rate and ATPase activity in lpx1Δ mutant transformed with pYES2/CT-EV and pYES2/CT-LPX1 vectors after induction by galactose in different times. Wild-type strain (BY4741) and the corresponding lpx1Δ and yvc1Δ mutants were used as controls. (A) Proton-pumping rate was measured with around 500 mg of cells after induction for different times with 100 mM galactose. Cells without incubation with 100 mM galactose were used as controls. (B) To measure ATPase activity from purified plasma membrane H+-ATPase, cells were collected after overnight induction with galactose, and membranes were obtained as described previously with a standard method. Different letters and asterisk indicate that mean values are statistically different (P < 0.05). Figure 2. View largeDownload slide Plasma membrane proton-pumping rate and ATPase activity in lpx1Δ mutant transformed with pYES2/CT-EV and pYES2/CT-LPX1 vectors after induction by galactose in different times. Wild-type strain (BY4741) and the corresponding lpx1Δ and yvc1Δ mutants were used as controls. (A) Proton-pumping rate was measured with around 500 mg of cells after induction for different times with 100 mM galactose. Cells without incubation with 100 mM galactose were used as controls. (B) To measure ATPase activity from purified plasma membrane H+-ATPase, cells were collected after overnight induction with galactose, and membranes were obtained as described previously with a standard method. Different letters and asterisk indicate that mean values are statistically different (P < 0.05). In addition, to check the role of Lxp1p in H+-ATPase activation, ATPase activity was measured in purified plasma membranes of the lpx1Δ yeast strains transformed with pYES2/CT empty vector (pYES2/CT-EV) and the one containing the LPX1 gene (pYES2/CT-LPX1). After overnight galactose induction, plasma membranes from strain pYES2/CT-LPX1 showed higher activity (P < 0.05) than purified plasma membranes from the control strain, pYES2/CT-EV (Fig. 2B). Proteolytic activity of Lpx1 Initially, crude extracts of lpx1Δ strains transformed with pYES2/CT-EV (empty vector, used as control), pYES2/CT-LPX1 (vector containing LPX1 whole gene) and pYES2/CT-LPX1-MOD (vector containing a modified LPX1 gene, where the binding calcium domain was removed) were prepared and applied to a nickel column, to perform Lpx1p (or Lpx1p-MOD) His-tag purification. In order to confirm the presence of His-tagged Lpx1 proteins, a western blotting assay was performed (Fig. 3A) with eluates obtained from lpx1Δ cells transformed with pYES2/CT-EV, pYES2/CT-LPX1 and pYES2/CT-LPX1-MOD (without calcium-binding domain). The results demonstrated that the eluates from the nickel column recovered from lpx1Δ cells presented specific bands only in extracts obtained from lpx1Δ cells transformed with pYES2/CT-LPX1 and pYES2/CT-LPX1-MOD. Furthermore, no band was detected in the extract obtained from the strain transformed with the empty vector. Additionally, reduction in molecular mass of the Lpx1p from 49 to 41 kDa was confirmed in a western blotting experiment that used extracts obtained from lpx1Δ cells transformed from pYES2/CT-LPX1 and pYES2/CT-LPX1-MOD, respectively. These results were supported by differences in their specific relative migration distances (Rf) (Supplementary Material 4). Figure 3. View largeDownload slide Lpx1p His-Tag expression and proteolytic activity. Cells were incubated overnight in induction medium, collected and disrupted in lysis buffer. Resulting supernatant (crude extract) was incubated with HIS-Select nickel affinity gel (Sigma-Aldrich). Unbound proteins were removed by centrifugation. Column-bound proteins were recovered with elution buffer. (A) Western blotting assay using anti-His antibodies in samples of nickel-column eluates obtained from lpx1Δ strain transformed with the empty vector pYES2/CT-EV and vectors carrying pYES2/CT-LPX1 or pYES2/CT-LPX1-MOD. Arrows indicate specific bands. (B) Proteolytic activity of total membranes from wild-type and lpx1Δ mutant was assessed by using azocasein as substrate. A protein fraction was added to a reaction mixture (100 mM Tris–HCl, pH 8.0, 0.5% azocasein) with or without addition of calcium. One enzymatic activity unit was defined as a change in absorbance of 0.01 (428 nm) per minute in these conditions. (C) Lpx1p activity was evaluated in excised protein bands obtained from SDS-PAGE gels after electrophoresis of eluates of the lpx1Δ mutant transformed from pYES2/CT-EV and pYES2/CT-LPX1 vectors. Samples (100 μg of protein) were dissolved in non-reducing Laemmli buffer and run in non-reducing conditions at 4°C. Lpx1p bands (indicated by arrows in A) and the corresponding region obtained from the control system (indicated by a circle in A) were excised from gel and added to a reaction mixture (100 mM Tris–HCl, pH 8.0, 0.5% azocasein), with or without addition of calcium. One enzymatic activity unit was defined as a change in absorbance of 0.01 (428 nm) per minute in these conditions. In all cases, results are expressed as specific activity (as mg protein−1). Different letters and asterisk indicate that mean values are statistically different (P < 0.05). Figure 3. View largeDownload slide Lpx1p His-Tag expression and proteolytic activity. Cells were incubated overnight in induction medium, collected and disrupted in lysis buffer. Resulting supernatant (crude extract) was incubated with HIS-Select nickel affinity gel (Sigma-Aldrich). Unbound proteins were removed by centrifugation. Column-bound proteins were recovered with elution buffer. (A) Western blotting assay using anti-His antibodies in samples of nickel-column eluates obtained from lpx1Δ strain transformed with the empty vector pYES2/CT-EV and vectors carrying pYES2/CT-LPX1 or pYES2/CT-LPX1-MOD. Arrows indicate specific bands. (B) Proteolytic activity of total membranes from wild-type and lpx1Δ mutant was assessed by using azocasein as substrate. A protein fraction was added to a reaction mixture (100 mM Tris–HCl, pH 8.0, 0.5% azocasein) with or without addition of calcium. One enzymatic activity unit was defined as a change in absorbance of 0.01 (428 nm) per minute in these conditions. (C) Lpx1p activity was evaluated in excised protein bands obtained from SDS-PAGE gels after electrophoresis of eluates of the lpx1Δ mutant transformed from pYES2/CT-EV and pYES2/CT-LPX1 vectors. Samples (100 μg of protein) were dissolved in non-reducing Laemmli buffer and run in non-reducing conditions at 4°C. Lpx1p bands (indicated by arrows in A) and the corresponding region obtained from the control system (indicated by a circle in A) were excised from gel and added to a reaction mixture (100 mM Tris–HCl, pH 8.0, 0.5% azocasein), with or without addition of calcium. One enzymatic activity unit was defined as a change in absorbance of 0.01 (428 nm) per minute in these conditions. In all cases, results are expressed as specific activity (as mg protein−1). Different letters and asterisk indicate that mean values are statistically different (P < 0.05). The proteolytic activity of Lpx1 and its Ca2+ dependence was demonstrated by two different approaches: first, we isolate total membranes from wild-type and lpx1Δ cells and the activity was tested using a non-specific substrate (azocasein). As demonstrated in Fig. 3B, the presence of 1 mM calcium increases the proteolytic specific activity (AU/mg protein−1) in samples from wild-type cells; on the other hand, this effect was not observed in membrane samples obtained from the lpx1Δ mutant cells. Another experiment was performed to confirm the proteolytic activity of Lpx1. The corresponding protein bands obtained from PAGE gels of eluate from the lpx1Δ mutant transformed with pYES2/CT-LPX1 vector showed proteolytic activity clearly affected by the presence of 100 μM calcium. Excised gel fractions obtained from the strain transformed with an empty vector showed no proteolytic activity, even in the presence of calcium (Fig. 3C). In vitro plasma membrane H+-ATPase activation In order to confirm the connection between calcium signaling, Lpx1p activity and H+-ATPase activation, an in vitro system to reconstitute H+-ATPase was used. Initially, purified plasma membranes and dialyzed MFE obtained from glucose-starved wild-type cells were prepared. In these conditions, plasma membrane H+-ATPase is in a non-activated state (probably with acetylated tubulins bounded to the H+-ATPase C-terminal tail as described by Campetelli et al. (2013)). MFE provides the in vitro system with all elements required for H+-ATPase activation, including Ptk2p (required for H+-ATPase phosphorylation) and Lpx1p (necessary for acetylated tubulin proteolysis). To achieve the most suitable conditions for this in vitro activation assay, different calcium concentrations (100 μM, 1 and 5 mM), different times of incubation (10, 30 and 60 min) and different MFE protein concentrations (50, 100, 200 and 400 μg) for in vitro H+-ATPase activation were tested to define the most suitable conditions (data not shown). Thus, adequate conditions for the assay were as follows: 30 min of incubation with 50 μg of protein from MFE and with the addition of 100 μM calcium. When purified plasma membranes obtained from the wild-type glucose-starved cells were incubated only with (i) dialyzed MFE from the same strain or (ii) only calcium, there was no H+-ATPase activation in comparison with the activity detected when only purified plasma membrane from wild-type glucose-starved cells was used (Fig. 4A). However, when purified plasma membranes were combined with both MFE and calcium, H+-ATPase activation was observed (47.5% increase in activity, P < 0.05). Additionally, activity was strongly reduced (77.4% reduced, P < 0.05) when 50 μM sodium orthovanadate, a specific plasma membrane H+-ATPase inhibitor, was added. This seems to confirm that ATP hydrolytic activity measure in this assay was due to plasma membrane H+-ATPase (Fig. 4A). Figure 4. View largeDownload slide H+-ATPase in vitro activation assay. (A) Effect of calcium, membrane-free extract (MFE) and sodium orthovanadate addition in H+-ATPase in vitro activation assay. A method to evaluate in vitro H+-ATPase activation was established in this work. Purified plasma membranes and dialyzed MFE from BY4741 cells were used. Plasma membranes from glucose-starved wild-type cells were added to incubation buffer. Calcium and/or MFE was added accordingly to the different assays performed. To verify H+-ATPase activity inhibition, 50 μM of sodium orthovanadate (specific H+-ATPase inhibitor) was added. (B) Effect of Lpx1p and Ptk2p on H+-ATPase in vitro activation. Plasma membranes from glucose-starved wild-type cells were added to incubation buffer. MFE from lpx1Δ strain (lacking Lpx1p, but containing Ptk2p) or ptk2Δ cells (missing Ptk2p, but expressing Lpx1p) and calcium were added according to the different assays performed. (C) Plasma membranes from glucose-starved wild-type cells were added to incubation buffer with MFE from lpx1Δ or ptk2Δ strains. Eluates resulting from the application of cell extracts of lpx1Δ mutant (transformed with pYES2/CT-LPX1 or pYES2/CT-LPX1-MOD vectors) to a Ni-affinity column, with or without calcium addition, were used to evaluate the H+-ATPase in vitro activation. All results are expressed as a percentage relative to wild-type control assay with plasma membranes only. Asterisks indicate that mean values are statistically different from those seen in each relative control used (P < 0.05). Figure 4. View largeDownload slide H+-ATPase in vitro activation assay. (A) Effect of calcium, membrane-free extract (MFE) and sodium orthovanadate addition in H+-ATPase in vitro activation assay. A method to evaluate in vitro H+-ATPase activation was established in this work. Purified plasma membranes and dialyzed MFE from BY4741 cells were used. Plasma membranes from glucose-starved wild-type cells were added to incubation buffer. Calcium and/or MFE was added accordingly to the different assays performed. To verify H+-ATPase activity inhibition, 50 μM of sodium orthovanadate (specific H+-ATPase inhibitor) was added. (B) Effect of Lpx1p and Ptk2p on H+-ATPase in vitro activation. Plasma membranes from glucose-starved wild-type cells were added to incubation buffer. MFE from lpx1Δ strain (lacking Lpx1p, but containing Ptk2p) or ptk2Δ cells (missing Ptk2p, but expressing Lpx1p) and calcium were added according to the different assays performed. (C) Plasma membranes from glucose-starved wild-type cells were added to incubation buffer with MFE from lpx1Δ or ptk2Δ strains. Eluates resulting from the application of cell extracts of lpx1Δ mutant (transformed with pYES2/CT-LPX1 or pYES2/CT-LPX1-MOD vectors) to a Ni-affinity column, with or without calcium addition, were used to evaluate the H+-ATPase in vitro activation. All results are expressed as a percentage relative to wild-type control assay with plasma membranes only. Asterisks indicate that mean values are statistically different from those seen in each relative control used (P < 0.05). All in vitro plasma membrane H+-ATPase activity rates measured in these assays ranged between 0.10 and 0.28 μmol Pi min−1 mg−1 protein. There was no ATPase activity detected when the assay was performed with only MFE or eluates from all strains used in in vitro activation (data not shown). Then, to confirm the connection between calcium signaling, Lpx1p activity and Ptk2p (a protein kinase) in the context of the phosphorylation/activation of H+-ATPase, this in vitro H+-ATPase activation was tested using purified membranes obtained from wild-type cells and different MFEs prepared from ptk2Δ and lpx1Δ mutant strains. As can be seen in Fig. 4B, the addition of MFE obtained from lpx1Δ alone or in combination with 100 μM calcium does not result in any increase of the H+-ATPase activity (P > 0.05). When using MFE from ptk2Δ mutants, a slight increase (11.8%, P < 0.05) was verified on addition of 100 μM calcium. Moreover, there was no increase in H+-ATPase activity when the Ni-column eluate obtained from the lpx1Δ strain transformed with an empty vector (pYES2/CT-EV, used as control) was assessed, even with calcium addition combined with MFE from lpx1Δ or ptk2Δ strains (data not shown). Also, there was no increase (P > 0.05) in H+-ATPase activity when Ni-column eluate obtained from the lpx1Δ strain transformed with pYES2/CT-LPX1 vector was associated with MFE prepared from the ptk2Δ mutant, with or without 100 μM calcium addition (Fig. 4C, first two columns at left side). At the same time, when the Ni-column eluates obtained from the lpx1Δ strain transformed pYES2/CT-LPX1 vector was added to the incubation system, in association with MFE from lpx1Δ strain, a clear increase (57.3%; P < 0.05) in H+-ATPase activity was only observed when 100 μM calcium was present (central two columns in Fig 4C). On the other hand, with Ni-column eluates obtained from the lpx1Δ strain transformed with pYES2/CT-LPX1-MOD vector, any increase of the H+-ATPase activity was observed in association with MFE from lpx1Δ even in the presence of 100 μM calcium (Fig. 4C, two last columns at right side). DISCUSSION In the past 20 years, a large amount of evidence has suggested that glucose-induced activation of plasma membrane H+-ATPase is clearly dependent on calcium metabolism (dos Passos et al.1992; Brandão et al.1994; Coccetti et al.1998; Souza, Tropia and Brandao 2001; Trópia et al.2006; Pereira et al.2008; Bouillet et al.2012). Nevertheless, until now, a Ca2+-dependent protein kinase that could be involved in H+-ATPase activation through phosphorylation has never been identified. In fact, it was demonstrated that protein kinase Ptk2p would be the enzyme responsible for glucose-induced phosphorylation of residue Ser-899, located at the H+-ATPase C-terminal tail, leading to a Km reduction of the ATPase (Goossens et al.2000; Pereira et al.2015), but this kinase does not seem to belong to any calcium responsive class of protein kinases. Moreover, neither a second protein kinase (which would be responsible for the phosphorylation of Ser-911/Thr-912 residues) nor any other Ca2+-dependent protein has ever been described as being involved in this activation process. Therefore, to overcome this apparent contradiction, it seems necessary to us to develop different strategies to demonstrate how glucose-induced calcium signaling would be connected to the plasma membrane H+-ATPase activation process. Interestingly, it was already proposed that a serine protease is related to H+-ATPase activation. These pieces of evidence suggest that activation of the H+-ATPase would require a glucose-induced hydrolysis of an acetylated tubulin to liberate its C-terminal tail, making it available for phosphorylation and activation (Campetelli et al.2013). This proteolytic action is exerted by a serine protease encoded by the LPX1 gene (Campetelli et al.2005, 2013), and, as a new component of this elaborated pathway, it could be investigated as a possible candidate that would respond to the calcium signal, thereby explaining its function in the glucose-induced activation of plasma membrane H+-ATPase. We also found that Lpx1p presents a potential region for calcium interaction (an EF-hand, calcium-binding domain), apparently located at its C-terminus, when a search with WebFEATURE mas made. WebFEATURE allows scanning for different functional sites in proteins using predicted 3D molecular structure (Liang et al.2003; Wu, Liang and Altman 2008). The EF-hand motif is the most common calcium-binding motif found in proteins. In a large number of proteins, this motif does indeed bind calcium (or, in some cases, magnesium) and these proteins exert diverse functions such as calcium buffering in cytosol, signal transduction between cellular compartments and muscle contraction (Lewit-Bentley and Réty 2000). Thus, this predicted that an EF-hand calcium-binding domain in the structure of Lpx1p could be responsible for calcium interaction and possibly regulate its activity. Additionally, Thoms et al. (2011) found, also in a search for functional sites in the Lpx1p structure, that the cap domain of this protein shows similarity to calmodulin, a well-known intracellular protein target of Ca2+ in eukaryotes. According to these authors, this cap covers the Lpx1p active site, and its N-terminal loop shows characteristics indicating high flexibility, which could suggest that this loop might act as a lid that can regulate access to the active site (Thoms et al.2011). Curiously, it seems that Lpx1p also exhibits acyl hydrolase and phospholipase A activities and can be found in peroxisomes (Thoms et al.2008); it also shows an ambiguous distribution (Huh et al.2003). Therefore, Lpx1p can be considered a good candidate to form a link between calcium signaling and glucose-induced plasma membrane H+-ATPase activation. This possibility was initially investigated by using strains with single or combined deletions in different genes: ARG82, YVC1 and LPX1. Cells with deletion in ARG82 (which encodes an inositol kinase responsible for the phosphorylation of IP3) have an increase in both glucose-induced calcium signaling and H+-ATPase activation (Tisi et al.2004; Trópia et al.2006; Pereira et al.2008). This increase suggests that H+-ATPase activation is dependent on calcium signaling. As demonstrated here, deletion of LPX1 alone or in combination with ARG82 led to a reduction in the glucose-induced activation of the plasma membrane H+-ATPase. Nevertheless, in the lpx1Δ mutant the calcium signaling was comparable to wild-type cells. Additionally, in the double mutant arg82Δ lpx1Δ, glucose-induced calcium signaling was also higher, as in the single mutant arg82Δ, suggesting that Lpx1p is not directly involved in calcium signaling. These results were also confirmed when calcium signaling and proton-pumping activity were measured in yvc1Δ and yvc1Δ lpx1Δ strains. An alternative explanation for these results would be the existence of a signal transduction pathway, in which a glucose-induced calcium signal would be responsible for protease Lpx1p activation. According to a model suggested before (Campetelli et al.2013), this protease hydrolyzes an acetylated tubulin bound to the H+-ATPase C-terminal tail, making this region accessible to protein kinases. Indeed, here we demonstrated that the lpx1Δ strain expressing the LPX1 gene inserted in a galactose-inducible vector (pYES2/CT-LPX1) showed progressive increase in the in vivo H+-ATPase activity, confirming that Lpx1p is indeed essential to H+-ATPase activation. The results of a similar experiment done with the yvc1Δ  mutant transformed with the same vector also suggest that the calcium channel Yvc1 is not controlled by Lpx1p. These results, combined with proteolytic activity assays, indicate that Lpx1p proteolytic activity could be the target of calcium signaling, making the C-terminal tail of the H+-ATPase accessible to phosphorylation performed by at least one protein kinase (Ptk2p). To verify the direct connection between calcium signaling, Lpx1p activity and H+-ATPase activation, it was necessary to establish an in vitro system by which it would be possible to reconstitute the plasma membrane H+-ATPase activation. Thus, when purified plasma membrane (isolated from wild-type cells) was incubated with MFE from lpx1Δ or ptk2Δ strains, in the presence of 100 μM calcium, H+-ATPase activation was absent or had a slight increase, respectively. This suggests once again that these two proteins (Lpx1 and Ptk2) are necessary to trigger a proper activation. Moreover, calcium and Lpx1p addition (obtained from the eluate of a Ni-affinity column of a cell extract of the lpx1Δ mutant transformed with a pYES2/CT-LPX1 vector), combined with MFE from lpx1Δ cells (which gives other components necessary for activation, but not Lpx1p), resulted in an increase of H+-ATPase activity only when 100 μM calcium is also present. However, in similar conditions when the MFE from ptk2Δ strain was used, no increase in H+-ATPase activity was observed, even in the presence of 100 μM calcium. These results reinforce the importance of Ptk2p in the activation process of H+-ATPase. Finally, when modified Lpx1p protein (without the C-terminal tail portion that potentially interacts with Ca2+) was used in combination with MFE from lpx1Δ cells and calcium, no increase in the H+-ATPase activity was observed. Altogether, these approaches suggest that most probably glucose-induced calcium signaling is connected to plasma membrane H+-ATPase activation through a calcium-induced activation of the serine protease Lpx1p. Thus, with the results presented here, the following mechanism can be proposed: on glucose addition, Ca2+ binds to Lpx1p, leading to its activation making possible the degradation of the acetylated tubulin bound to the plasma membrane H+-ATPase. Therefore, the H+-ATPase C-terminal tail would be accessible to phosphorylation performed by at least one protein kinase (Ptk2p) leading to the activation of the plasma membrane H+-ATPase (Fig. 5). Figure 5. View largeDownload slide Lpx1p calcium binding and H+-ATPase activation. (A) Proposed mechanism by which Lpx1p is possibly regulated by calcium availability. In the presence of glucose, calcium is released into cytoplasm and binds to Lxp1p. (B) After calcium binding, Lxp1p is able to degrade acetylated tubulin bound to plasma membrane H+-ATPase releasing its C-terminal tail. This would make the H+-ATPase C-terminus phosphorylation sites accessible to at least one protein kinase (Ptk2p) making possible the enzyme activation. Figure 5. View largeDownload slide Lpx1p calcium binding and H+-ATPase activation. (A) Proposed mechanism by which Lpx1p is possibly regulated by calcium availability. In the presence of glucose, calcium is released into cytoplasm and binds to Lxp1p. (B) After calcium binding, Lxp1p is able to degrade acetylated tubulin bound to plasma membrane H+-ATPase releasing its C-terminal tail. This would make the H+-ATPase C-terminus phosphorylation sites accessible to at least one protein kinase (Ptk2p) making possible the enzyme activation. Therefore, taking together the data from our previous work (dos Passos et al.1992; Brandão et al.1994; Coccetti et al.1998; Souza, Tropia and Brandao 2001; Trópia et al.2006; Pereira et al.2008; Groppi et al.2011; Bouillet et al.2012) and the new results shown in this work, we suggest the existence of a signal transduction pathway with two branches, by which glucose addition controls calcium availability in the cytosol with a direct consequence for plasma membrane H+-ATPase activation (Fig. 6). In the first branch, glucose uptake and its subsequent phosphorylation generate a signal (probably sugar phosphates) that would stimulate G protein Gpa2p to interact with and/or activate phospholipase C (Plc1p). Then, Plc1p would hydrolyze phosphatidylinositol-4,5-bisphosphate, generating diacylglycerol and IP3. IP3 would interact, directly or indirectly, with Yvc1p regulating the intensity of calcium signaling in the cytosol (Bouillet et al.2012). In the second branch, devoted to the control of Pmc1p Ca2+-ATPase activity, the glucose sensor Snf3p (Özcan and Johnston 1999) could also detect sugar phosphates (Dlugai et al.2001), and in some way transduce this signal leading to an increase in Pmc1p activity (Souza, Tropia and Brandao 2001; Trópia et al.2006; Pereira et al.2008). The balance between these two branches would be responsible for the transient nature of calcium signaling. This calcium signaling seems to be responsible for activation of Lpx1p, which hydrolyzes an acetylated tubulin bound to plasma membrane H+-ATPase, allowing thus the H+-ATPase C-terminal tail to be released, enabling phosphorylation of C-terminal sites of the H+-ATPase and in this way its activation. Figure 6. View largeDownload slide Working model: glucose-induced activation of plasma membrane H+-ATPase in yeast cells. A signal transduction pathway with two branches, resulting in intracellular calcium-signal generation, is triggered by internalization followed by phosphorylation of glucose, which generates a signal (maybe relative amounts of glucose-6-P and/or glucose-1-P) that would stimulate the complex constituted of the G protein, Gpa2p, and phospholipase C eliciting the activation of the phospholipase C. Then, phosphatidylinositol-4,5-bisphosphate hydrolysis would generate diacylglycerol and IP3. In the first branch, IP3 would act directly or indirectly on the vacuolar calcium channel Yvc1p leading to an increase in the intracellular calcium signal. Besides that, in a second branch, the C-terminal tail of the glucose sensor Snf3p controlling the activity of the vacuolar Ca2+-ATPase, Pmc1p, would detect the signal. The final intensity of the calcium signal would be the result of the partial contribution of each branch of this system. This calcium signaling seems to be responsible for activation of Lpx1p, which hydrolyzes an acetylated tubulin bound to plasma membrane H+-ATPase, allowing thus the H+-ATPase C-terminal tail to be released, enabling phosphorylation of C-terminal sites of the H+-ATPase and in this way its activation. Figure 6. View largeDownload slide Working model: glucose-induced activation of plasma membrane H+-ATPase in yeast cells. A signal transduction pathway with two branches, resulting in intracellular calcium-signal generation, is triggered by internalization followed by phosphorylation of glucose, which generates a signal (maybe relative amounts of glucose-6-P and/or glucose-1-P) that would stimulate the complex constituted of the G protein, Gpa2p, and phospholipase C eliciting the activation of the phospholipase C. Then, phosphatidylinositol-4,5-bisphosphate hydrolysis would generate diacylglycerol and IP3. In the first branch, IP3 would act directly or indirectly on the vacuolar calcium channel Yvc1p leading to an increase in the intracellular calcium signal. Besides that, in a second branch, the C-terminal tail of the glucose sensor Snf3p controlling the activity of the vacuolar Ca2+-ATPase, Pmc1p, would detect the signal. The final intensity of the calcium signal would be the result of the partial contribution of each branch of this system. This calcium signaling seems to be responsible for activation of Lpx1p, which hydrolyzes an acetylated tubulin bound to plasma membrane H+-ATPase, allowing thus the H+-ATPase C-terminal tail to be released, enabling phosphorylation of C-terminal sites of the H+-ATPase and in this way its activation. Beyond representing a clear advance in our understanding of how glucose-induced calcium signaling and plasma membrane H+-ATPase activation would be connected, these findings can help in the elaboration of new strategies to find out the identity of all protein kinases that are involved in plasma membrane H+-ATPase phosphorylation. Perhaps, the fact that a proteolytic degradation of acetylated tubulin precludes and/or occurs in parallel with phosphorylation could help in conceiving new approaches to identify those protein kinases in addition to Ptk2p. SUPPLEMENTARY DATA Supplementary data are available at FEMSYR online. Acknowledgements We are also grateful to Dr James Caffrey from National Institute of Environmental Health Sciences, USA, for the strains used in this work and to Marco Vanoni (Università di Milano-Bicocca, Milan, Italy) for pVTU-AEQ and pYX212-AEQ plasmids. FUNDING This work was partially financed by grants from Universidade Federal de Ouro Preto, from Fundação de Amparo a Pesquisa do Estado de Minas Gerais (FAPEMIG), Process CBB 824/06; from Conselho Nacional de Desenvolvimento–CNPq, Process 304815/2012-3 (research fellowship to RLB), Process 475672/08-5 (research grant) and CAPES, Process 2041/2012 (PhD fellowship to FFO), Process PNPD/2013 (research fellowships to RHSD), Process PNPD 2755/2011 (research fellowships to FGS), Process BEX 11122/13-7 (PhD Sandwich fellowship to DDC). Conflict of Interest. None declared. REFERENCES Anraku Y, Ohya Y, Iida H. Cell cycle control by calcium and calmodulin in Saccharomyces cerevisiae. Biochim Biophys Acta  1991; 1093: 169– 77. Google Scholar CrossRef Search ADS PubMed  Bouillet LE, Cardoso AS, Perovano E et al.   The involvement of calcium carriers and of the vacuole in the glucose-induced calcium signaling and activation of the plasma membrane H+-ATPase in Saccharomyces cerevisiae cells. Cell Calcium  2012; 51: 72– 81. Google Scholar CrossRef Search ADS PubMed  Brandão RL, de Magalhães-Rocha NM, Alijo R et al.   Possible involvement of a phosphatidylinositol-type signaling pathway in glucose-induced activation of plasma membrane H+-ATPase and cellular proton extrusion in the yeast Saccharomyces cerevisiae. Biochim Biophys Acta  1994; 1223: 117– 24. Google Scholar CrossRef Search ADS PubMed  Brandão RL. The relationship between glucose-induced calcium signaling and activation of the plasma membrane H+-ATPase in Saccharomyces cerevisiae cells. In: Nakamura S (ed.). Handbook of H+-ATPases . Singapore: Pan Stanford Publishing Pte Ltd, 2014, 431– 48. Google Scholar CrossRef Search ADS   Campetelli AN, Monesterolo NE, Previtali G et al.   Activation of H+-ATPase by glucose in Saccharomyces cerevisiae involves a membrane serine protease. Biochim Biophys Acta  2013; 1830: 3593– 603. Google Scholar CrossRef Search ADS PubMed  Campetelli AN, Previtali G, Arce CA et al.   Activation of the plasma membrane H+-ATPase of Saccharomyces cerevisiae by glucose is mediated by dissociation of the H+-ATPase–acetylated tubulin complex. FEBS J  2005; 272: 5742– 52. Google Scholar CrossRef Search ADS PubMed  Capieaux E, Vignais ML, Sentenac A et al.   The yeast H+-ATPase gene is controlled by the promoter binding factor TUF. J Biol Chem  1989; 264: 7437– 46. Google Scholar PubMed  Charney J, Tomarelli RM. A colorimetric method for the determination of the proteolytic activity of duodenal juice. J Biol Chem  1947; 171: 501– 5. Google Scholar PubMed  Coccetti P, Tisi R, Martegani E et al.   The PLC1 encoded phospholipase C in the yeast Saccharomyces cerevisiae is essential for glucose-induced phosphatidylinositol turnover and activation of plasma membrane H+-ATPase. Biochim Biophys Acta  1998; 1405: 147– 54. Google Scholar CrossRef Search ADS PubMed  Cunningham KW. Acidic calcium stores of Saccharomyces cerevisiae. Cell Calcium  2011; 50: 129– 38. Google Scholar CrossRef Search ADS PubMed  Cyert MS, Philpott CC. Regulation of cation balance in Saccharomyces cerevisiae. Genetics  2013; 193: 677– 713. Google Scholar CrossRef Search ADS PubMed  Denis V, Cyert MS. Internal Ca2+ release in yeast is triggered by hypertonic shock and mediated by a TRP channel homologue. J Cell Biol  2002; 156: 29– 34. Google Scholar CrossRef Search ADS PubMed  Dlugai S, Hippler S, Wieczorke R et al.   Glucose-dependent and -independent signalling functions of the yeast glucose sensor Snf3. FEBS Lett  2001; 505: 389– 92. Google Scholar CrossRef Search ADS PubMed  dos Passos JB, Vanhalewyn M, Brandao RL et al.   Glucose-induced activation of plasma membrane H+-ATPase in mutants of the yeast Saccharomyces cerevisiae affected in cAMP metabolism, cAMP-dependent protein phosphorylation and the initiation of glycolysis. Biochim Biophys Acta  1992; 1136: 57– 67. Google Scholar CrossRef Search ADS PubMed  Dunn T, Gable K, Beeler T. Regulation of cellular Ca2+ by yeast vacuoles. J Biol Chem  1994; 269: 7273– 8. Google Scholar PubMed  Eraso P, Mazón MJ, Portillo F. Yeast protein kinase Ptk2 localizes at the plasma membrane and phosphorylates in vitro the C-terminal peptide of the H+-ATPase. Biochim Biophys Acta  2006; 1758: 164– 70. Google Scholar CrossRef Search ADS PubMed  Gietz RD, Schiestl RH, Willems AR et al.   Studies on the transformation of intact yeast cells by the LiAc/SS-DNA/PEG procedure. Yeast  1995; 11: 355– 60. Google Scholar CrossRef Search ADS PubMed  Goffeau A, Slayman CW. The proton-translocating ATPase of the fungal plasma membrane. Biochim Biophys Acta  1981; 639: 197– 223. Google Scholar CrossRef Search ADS PubMed  Goossens A, de La Fuente N, Forment J et al.   Regulation of yeast H+-ATPase by protein kinases belonging to a family dedicated to activation of plasma membrane transporters. Mol Cell Biol  2000; 20: 7654– 61. Google Scholar CrossRef Search ADS PubMed  Groppi S, Belotti F, Brandão RL et al.   Glucose-induced calcium influx in budding yeast involves a novel calcium transport system and can activate calcineurin. Cell Calcium  2011; 49: 376– 86. Google Scholar CrossRef Search ADS PubMed  Huh W-K, Falvo JV, Gerke LC et al.   Global analysis of protein localization in budding yeast. Nature  2003; 425: 686– 91. Google Scholar CrossRef Search ADS PubMed  Lecchi S, Allen KE, Pardo JP et al.   Conformational changes of yeast plasma membrane H+-ATPase during activation by glucose: role of threonine-912 in the carboxy-terminal tail. Biochemistry  2005; 44: 16624– 32. Google Scholar CrossRef Search ADS PubMed  Lecchi S, Nelson CJ, Allen KE et al.   Tandem phosphorylation of Ser-911 and Thr-912 at the C terminus of yeast plasma membrane H+-ATPase leads to glucose-dependent activation. J Biol Chem  2007; 282: 35471– 81. Google Scholar CrossRef Search ADS PubMed  Lewit-Bentley A, Réty S. EF-hand calcium-binding proteins. Curr Opin Struct Biol  2000; 10: 637– 43. Google Scholar CrossRef Search ADS PubMed  Liang MP, Banatao DR, Klein TE et al.   WebFEATURE: an interactive web tool for identifying and visualizing functional sites on macromolecular structures. Nucleic Acids Res  2003; 31: 3324– 7. Google Scholar CrossRef Search ADS PubMed  Lowry OH, Rosebrough NJ, Farr AL et al.   Protein measurement with the Folin phenol reagent. J Biol Chem  1951; 193: 265– 75. Google Scholar PubMed  Mazón MJ, Eraso P, Portillo F. Specific phosphoantibodies reveal two phosphorylation sites in yeast Pma1 in response to glucose. FEMS Yeast Res  2015; 15: fov030. Google Scholar CrossRef Search ADS PubMed  Miseta A, Fu L, Kellermayer R et al.   The Golgi apparatus plays a significant role in the maintenance of Ca2+ homeostasis in the vps33Δ vacuolar biogenesis mutant of Saccharomyces cerevisiae. J Biol Chem  1999a; 274: 5939– 47. Google Scholar CrossRef Search ADS   Miseta A, Kellermayer R, Aiello DP et al.   The vacuolar Ca2+/H+ exchanger Vcx1p/Hum1p tightly controls cytosolic Ca2+ levels in S. cerevisiae. FEBS Lett  1999b; 451: 132– 6. Google Scholar CrossRef Search ADS   Odom AR, Stahlberg A, Wente SR et al.   A role for nuclear inositol 1,4,5-trisphosphate kinase in transcriptional control. Science  2000; 287: 2026– 9. Google Scholar CrossRef Search ADS PubMed  Özcan S, Johnston M. Function and regulation of yeast hexose transporters. Microbiol Mol Biol R  1999; 63: 554– 69. Paidhungat M, Garrett S. A homolog of mammalian, voltage-gated calcium channels mediates yeast pheromone-stimulated Ca2+ uptake and exacerbates the cdc1 (Ts) growth defect. Mol Cell Biol  1997; 17: 6339– 47. Google Scholar CrossRef Search ADS PubMed  Pereira MB, Tisi R, Fietto LG et al.   Carbonyl cyanide m-chlorophenylhydrazone induced calcium signaling and activation of plasma membrane H+-ATPase in the yeast Saccharomyces cerevisiae. FEMS Yeast Res  2008; 8: 622– 30. Google Scholar CrossRef Search ADS PubMed  Pereira RR, Castanheira D, Teixeira JA et al.   Detailed search for protein kinase(s) involved in plasma membrane H+-ATPase activity regulation of yeast cells. FEMS Yeast Res  2015; 15: fov003. Google Scholar CrossRef Search ADS PubMed  Portillo F. Regulation of plasma membrane H+-ATPase in fungi and plants. Biochim Biophys Acta  2000; 1469: 31– 42. Google Scholar CrossRef Search ADS PubMed  Portillo F, Eraso P, Serrano R. Analysis of the regulatory domain of yeast plasma membrane H+-ATPase by directed mutagenesis and intragenic suppression. FEBS Lett  1991; 287: 71– 4. Google Scholar CrossRef Search ADS PubMed  Rao R, Drummond-Barbosa D, Slayman CW. Transcriptional regulation by glucose of the yeast PMA1 gene encoding the plasma membrane H+-ATPase. Yeast  1993; 9: 1075– 84. Google Scholar CrossRef Search ADS PubMed  Saiardi A, Caffrey JJ, Snyder SH et al.   Inositol polyphosphate multikinase (ArgRIII) determines nuclear mRNA export in Saccharomyces cerevisiae. FEBS Lett  2000; 468: 28– 32. Google Scholar CrossRef Search ADS PubMed  Saiardi A, Erdjument-Bromage H, Snowman AM et al.   Synthesis of diphosphoinositol pentakisphosphate by a newly identified family of higher inositol polyphosphate kinases. Curr Biol  1999; 9: 1323– 6. Google Scholar CrossRef Search ADS PubMed  Secades P, Guijarro JA. Purification and characterization of an extracellular protease from the fish pathogen Yersinia ruckeri and effect of culture conditions on production. Appl Environ Microb  1999; 65: 3969– 75. Serrano R. Structure, function and regulation of plasma membrane H+-ATPase. FEBS Lett  1993; 325: 108– 11. Google Scholar CrossRef Search ADS PubMed  Serrano R, Ruiz A, Bernal D et al.   The transcriptional response to alkaline pH in Saccharomyces cerevisiae: evidence for calcium-mediated signalling. Mol Microbiol  2002; 46: 1319– 33. Google Scholar CrossRef Search ADS PubMed  Shears SB. Transcriptional regulation: a new dominion for inositol phosphate signaling? Bioessays  2000; 22: 786– 9. Google Scholar CrossRef Search ADS PubMed  Souza M, Tropia M, Brandao R. New aspects of the glucose activation of the H+-ATPase in the yeast Saccharomyces cerevisiae. Microbiology  2001; 147: 2849– 55. Google Scholar CrossRef Search ADS PubMed  Thoms S, Debelyy MO, Nau K et al.   Lpx1p is a peroxisomal lipase required for normal peroxisome morphology. FEBS J  2008; 275: 504– 14. Google Scholar CrossRef Search ADS PubMed  Thoms S, Hofhuis J, Thöing C et al.   The unusual extended C-terminal helix of the peroxisomal α/β-hydrolase Lpx1 is involved in dimer contacts but dispensable for dimerization. J Struct Biol  2011; 175: 362– 71. Google Scholar CrossRef Search ADS PubMed  Tisi R, Baldassa S, Belotti F et al.   Phospholipase C is required for glucose-induced calcium influx in budding yeast. FEBS Lett  2002; 520: 133– 8. Google Scholar CrossRef Search ADS PubMed  Tisi R, Belotti F, Wera S et al.   Evidence for inositol triphosphate as a second messenger for glucose-induced calcium signalling in budding yeast. Curr Genet  2004; 45: 83– 9. Google Scholar CrossRef Search ADS PubMed  Tisi R, Martegani E, Brandão RL. Monitoring yeast intracellular Ca2+ levels using an in vivo bioluminescence assay. Cold Spring Harb Protoc  2015; 2015: 210– 3. Google Scholar PubMed  Trópia M, Cardoso A, Tisi R et al.   Calcium signaling and sugar-induced activation of plasma membrane H+-ATPase in Saccharomyces cerevisiae cells. Biochem Biophys Res Commun  2006; 343: 1234– 43. Google Scholar CrossRef Search ADS PubMed  Wera S, Bergsma JCT, Thevelein JM. Phosphoinositides in yeast: genetically tractable signalling. FEMS Yeast Res  2001; 1: 9– 13. Google Scholar CrossRef Search ADS PubMed  Wu S, Liang MP, Altman RB. The SeqFEATURE library of 3D functional site models: comparison to existing methods and applications to protein function annotation. Genome Biol  2008; 9: R8. Google Scholar CrossRef Search ADS PubMed  York JD, Odom AR, Murphy R et al.   A phospholipase C-dependent inositol polyphosphate kinase pathway required for efficient messenger RNA export. Science  1999; 285: 96– 100. Google Scholar CrossRef Search ADS PubMed  © FEMS 2017. All rights reserved. For permissions, please e-mail: journals.permissions@oup.com

Journal

FEMS Yeast ResearchOxford University Press

Published: Feb 1, 2018

There are no references for this article.

You’re reading a free preview. Subscribe to read the entire article.


DeepDyve is your
personal research library

It’s your single place to instantly
discover and read the research
that matters to you.

Enjoy affordable access to
over 18 million articles from more than
15,000 peer-reviewed journals.

All for just $49/month

Explore the DeepDyve Library

Search

Query the DeepDyve database, plus search all of PubMed and Google Scholar seamlessly

Organize

Save any article or search result from DeepDyve, PubMed, and Google Scholar... all in one place.

Access

Get unlimited, online access to over 18 million full-text articles from more than 15,000 scientific journals.

Your journals are on DeepDyve

Read from thousands of the leading scholarly journals from SpringerNature, Elsevier, Wiley-Blackwell, Oxford University Press and more.

All the latest content is available, no embargo periods.

See the journals in your area

DeepDyve

Freelancer

DeepDyve

Pro

Price

FREE

$49/month
$360/year

Save searches from
Google Scholar,
PubMed

Create lists to
organize your research

Export lists, citations

Read DeepDyve articles

Abstract access only

Unlimited access to over
18 million full-text articles

Print

20 pages / month

PDF Discount

20% off