Glucose induces rapid changes in the secretome of Saccharomyces cerevisiae
© Giardina et al.; licensee BioMed Central Ltd. 2014
Received: 12 September 2013
Accepted: 31 January 2014
Published: 12 February 2014
Protein secretion is a fundamental process in all living cells. Proteins can either be secreted via the classical or non-classical pathways. In Saccharomyces cerevisiae, gluconeogenic enzymes are in the extracellular fraction/periplasm when cells are grown in media containing low glucose. Following a transfer of cells to high glucose media, their levels in the extracellular fraction are reduced rapidly. We hypothesized that changes in the secretome were not restricted to gluconeogenic enzymes. The goal of the current study was to use a proteomic approach to identify extracellular proteins whose levels changed when cells were transferred from low to high glucose media.
We performed two iTRAQ experiments and identified 347 proteins that were present in the extracellular fraction including metabolic enzymes, proteins involved in oxidative stress, protein folding, and proteins with unknown functions. Most of these proteins did not contain typical ER-Golgi signal sequences. Moreover, levels of many of these proteins decreased upon a transfer of cells from media containing low to high glucose media. Using an extraction procedure and Western blotting, we confirmed that the metabolic enzymes (glyceraldehyde-3-phosphate dehydrogenase, 3-phosphoglycerate kinase, glucose-6-phosphate dehydrogenase, pyruvate decarboxylase), proteins involved in oxidative stress (superoxide dismutase and thioredoxin), and heat shock proteins (Ssa1p, Hsc82p, and Hsp104p) were in the extracellular fraction during growth in low glucose and that the levels of these extracellular proteins were reduced when cells were transferred to media containing high glucose. These proteins were associated with membranes in vesicle-enriched fraction. We also showed that small vesicles were present in the extracellular fraction in cells grown in low glucose. Following a transfer from low to high glucose media for 30 minutes, 98% of these vesicles disappeared from the extracellular fraction.
Our data indicate that transferring cells from low to high glucose media induces a rapid decline in levels of a large number of extracellular proteins and the disappearance of small vesicles from the extracellular fraction. Therefore, we conclude that the secretome undergoes dynamic changes during transition from glucose-deficient to glucose-rich media. Most of these extracellular proteins do not contain typical ER signal sequences, suggesting that they are secreted via the non-classical pathway.
Protein secretion is an important process for both prokaryotic and eukaryotic cells. Secretory proteins include growth factors, inflammatory cytokines, coagulation factors, extracellular matrix proteins, proteases, and protease inhibitors[1–9]. As such, they participate in various physiological processes such as immune defense, blood coagulation, cell growth, cell differentiation, and proliferation[1–5, 7–9]. For fungi that have thick cell walls, secretory proteins are involved in the formation and maintenance of cell walls, cell separation, and nutrient scavenging[10–17]. Proteins that are secreted from viruses, bacteria, fungi, and parasites are important for pathogen-host interactions[18–25]. Furthermore, secretory proteins also play crucial roles in cancer angiogenesis, differentiation, invasion, and metastasis[6, 7, 26–29].
Proteins can be secreted from cells via the classical and non-classical pathways[2, 3, 8, 9, 23, 30, 31]. In the classical pathway, proteins that contain a specific N-terminal signal sequence are translocated into the ER, transported to the Golgi and then secreted by secretory vesicles[30, 32, 33]. Accumulating evidence indicates that a large number of signal-less proteins can also be secreted through the non-classical pathway[1, 2, 4, 5, 10, 21, 23, 31],, including metabolic enzymes, chaperones, translation factors, and transcriptional regulators[5, 8, 9, 11, 20–24, 31, 34–37]. Glyceraldehyde-3-phosphate dehydrogenase (GAPDH) is a glycolytic enzyme and has been shown by immuno-transmission electron microscopy (immuno-TEM) to be present on the surface of Candida albicans and Saccharomyces cerevisiae. Using the immuno-TEM technique, another glycolytic enzyme, enolase, was also found on the surface of Listeria monocytogenes. In systemic infection by Candida albicans, enolase and GAPDH are secreted[11, 14, 16, 38]. GAPDH on the surface of Candida albicans is enzymatically active and binds to the host proteins fibronectin and laminin[14, 16, 38]. Furthermore, GAPDH and Hsp70 have been recently identified as major antigens in sera from patients infected with Echinostoma friedi.
Gluconeogenic enzymes have been identified in multiple secretomic studies from bacteria and parasites[18, 22–24, 37]. Fructose-1,6-bisphosphatase is found in the secretomes from Bacillus anthracis and Clonorchis sinensis, while isocitrate lyase is detected in the secretome in Bacillus anthracis. Moreover, malate dehydrogenase is present in the secretomes from Bacillus anthracis, Clonorchis sinensis, and Schistosoma mansoni. Phosphoenolpyruvate carboxykinase is also detectable in the secretomes from Clonorchis sinensis, Echinostoma caproni, and Schistosoma mansoni[22, 23, 37].
The yeast Saccharomyces cerevisiae is an excellent model system to study global changes in transcription, translation, and protein levels in response to environmental changes such as the availability of nutrients[39–44]. Although yeast can utilize both fermentable and non-fermentable carbon sources, glucose is the preferred carbon source. It has profound effects in gene transcription, translation, mRNA stabilities, and enzymatic activities[45–50]. For example, glucose leads to inactivation of gluconeogenic enzymes, mitochondrial proteins, and enzymes involved in the metabolism of acetate, maltose, glycerol, and galactose[51–55]. Inactivation of gluconeogenic enzymes during glucose re-feeding prevents energy futile cycles that are detrimental to cells. The key gluconeogenic enzyme fructose-1,6-bisphosphatase (Fbp1p) has been used extensively to study glucose-induced degradation[53, 56–59]. Fbp1p is either ubiquitinated and degraded in the proteasome[60, 61], or phosphorylated and degraded in the vacuole[53, 56–59]. Importantly, the site of Fbp1p degradation is dependent on the duration of starvation. For the vacuole pathway, gluconeogenic enzymes including Fbp1p, malate dehydrogenase (Mdh2p), isocitrate lyase (Icl1p), phosphoenolpyruvate carboxykinase (Pck1p), and malate synthase (Mls1p) were in the extracellular fraction during growth in low glucose. Furthermore, their levels in the extracellular fraction were reduced following a transfer of cells to media containing high glucose. This decrease is dependent on the presence of glucose in the media. When cells were transferred to media without glucose, these proteins did not decrease levels in the extracellular fraction. We hypothesized that changes in the secretome induced by glucose were not limited to gluconeogenic enzymes. The goals of this study were to use the iTRAQ approach to test our hypothesis and to identify proteins in the secretome whose levels changed upon a transfer of cells from low to high glucose media.
Here, we report the identification of 347 extracellular proteins from two independent iTRAQ experiments. This included metabolic enzymes and proteins involved in oxidative stress, translation, protein folding, and proteins with unknown functions. Most of these proteins did not contain the N-terminal ER signal sequence. Many of these identified proteins are also commonly found in secretomic studies from bacteria, fungi, parasites, and human cells[19, 20, 28, 35]. Using an extraction procedure and Western blotting, we confirmed that metabolic enzymes (glyceraldehyde-3-phosphate dehydrogenase, 3-phosphoglycerate kinase, glucose-6-phosphate dehydrogenase, pyruvate decarboxylase), proteins involved in oxidative stress (superoxide dismutase and thioredoxin), and heat shock proteins (Ssa1p, Hsc82p, and Hsp104p) were present in the extracellular fraction in cells grown in low glucose. The extracellular level of these proteins was rapidly reduced following a transfer of cells from low to high glucose media. Furthermore, we performed TEM studies and observed numerous small vesicles in total extracts isolated from cells grown in low glucose. Following a shift of cells to media containing high glucose for 30 min, most of these vesicles disappeared. We conclude that the secretome undergoes dynamic changes during transition from glucose-deficient to glucose-rich media.
Experimental conditions to study the effects of glucose on protein levels
To further determine whether or not changes in Lia1p and Fbp1p levels were due to the presence of glucose in the media, wild-type cells expressing Lia1p-GFP were grown in YPKG and transferred to YPD or YP (without added glucose) for 0, 2, and 4 hours (Figure 1B). Again, when cells were transferred from YPKG to YPD, levels of Lia1p increased and levels of Fbp1p decreased (Figure 1B, left panels). However, when cells were transferred from YPKG to YP, Lia1p levels did not increase and Fbp1p levels did not decrease (Figure 1B, right panels). Therefore, the presence of 2% glucose in the YPD media is critical in up-regulating Lia1p and down-regulating Fbp1p.
We next examined whether or not cells grown in YPKG media for 3 days were able to re-grow in YPD media containing 2% glucose (Figure 1C). Wild-type cells were grown in YPKG for 3d to OD600 = 4-5/ml. Cells were diluted to OD600 = 0.2/ml in YPD containing 2% glucose or YP media without glucose for 0–8 hours and changes in cell densities were measured at OD600 using a spectrophotometer. When cells were diluted in YPD, cells increased their densities after a lag. Likewise, when cells were diluted in YPKG, cells densities also increased after a lag. Hence, cells are able to resume growth when diluted in YPD or YPKG. In contrast, there was little increase in density when cells were cultured in YP media without glucose, suggesting that the presence of glucose in the media is required for cells to re-grow.
Fbp1p is re-distributed from the periplasm/extracellular fraction to the cytoplasm when cells are transferred from YPKG to YPD
Extraction of extracellular proteins
We next used an extraction procedure to determine whether or not this protocol could detect a decline in Fbp1p levels in the extracellular fraction. This protocol utilizes the combination of reducing agents such as β-mercaptoethanol (βME) or dithiothreitol (DTT) and high pH to release proteins in the extracellular fraction[13, 63]. This method of extraction has been utilized to study the secretion of mammalian galectin-1 expressed in Saccharomyces cerevisiae. A similar method has been used to identify proteins associated with the cell wall in Candida albicans[16, 38].
We also examined whether or not cells that were extracted retained the ability to internalize exogenous molecules into the cells. FM 4–64 (FM) is a vital dye that is internalized into live cells and actively transported to the vacuole via the endocytic pathway. Wild-type cells were grown in YPKG for 3d (t = 0) or transferred to glucose for 30 min (t = 30). Cells were treated with or without Triton X-100 and then incubated with FM for 3 hours. The same amounts of cells were also subjected to the extraction protocol and treated in the absence or presence of Triton X-100 followed by incubation with FM for 3 hours. The distribution of FM was then observed using fluorescence microscopy (Figure 3E-3L). In t = 0 and t = 30 non-extracted cells, this dye was transported to the vacuole membrane as ring-like circles (Figure 3E and3G). In contrast, when these cells were treated with Triton X-100 and then incubated with FM, this dye was not on the vacuole membrane (Figure 3F and3H). In t = 0 and t = 30 cells that were extracted and then incubated with FM, this dye was also transported correctly to the vacuole membrane (Figure 3I and3K). However, when extracted cells were treated with Triton X-100 and incubated with FM, this dye was not on the vacuole membrane (Figure 3J and3L). Thus, FM is targeted correctly to the vacuole membrane in both non-extracted and extracted cells. Furthermore, the endocytic pathway is functional in cells that were either grown in YPKG or transferred to YPD for 30 min.
Identification of extracellular proteins using iTRAQ
The findings that multiple gluconeogenic enzymes showed reduced levels in the extracellular fraction after a transfer of cells to YPD prompted us to hypothesize that changes in the secretome induced by glucose were not limited to gluconeogenic enzymes. To identify other extracellular proteins that changed levels in cells that were transferred from low to high glucose media, we used the iTRAQ technique that we previously employed to identify proteins in the total proteome that change their relative amounts under the same conditions. Because a significant reduction in protein levels in the extracellular fraction was observed at the t = 30 min time point for gluconeogenic enzymes, our iTRAQ experiments were performed using wild-type cells that were transferred to YPD for 30 min. For these iTRAQ experiments, gluconeogenic enzymes that were shown to be in the extracellular space were considered as positive controls that should be identified in the extracellular fraction. Their levels should decrease in the extracellular fraction at the t = 30 min time point. Additional controls for extracellular proteins included isoforms of glyceraldehyde-3-phosphate dehydrogenase Tdh1p, Tdh2p, and Tdh3p that have been shown to be on the cell-surface of Saccharomyces cerevisiae[11, 38, 67]. Ssa1p and Ssa2p are on the cell surface of Saccharomyces cerevisiae as indicated by immunofluorescence microscopy and served as additional controls for extracellular proteins. Invertase Suc2p and the constitutively expressed acid phosphatase Pho3p are secreted into the periplasm[68, 69] and should also be identified in the extracellular fraction.
Overall, we have confidently identified 318 extracellular proteins in iTRAQ1 and 392 extracellular proteins in ITRAQ2 with an estimated local False Discovery Rate (FDR) of less than 5%. Proteins from which only a single peptide had been identified were manually inspected and only those that were identified in both experiments were included. (Proteins that were identified with a single peptide with 95% confidence in only one of the two iTRAQ experiments were not included). After removing those single-peptide proteins, a total of 347 extracellular proteins remained including 264 extracellular proteins from iTRAQ1 and 324 proteins from iTRAQ2. The name of each identified protein with their accession number and the number of peptides identified with at least 95% confidence are listed in Additional file1: Table S1. In general, both iTRAQ1 and iTRAQ2 identified similar sets of proteins, but iTRAQ2 identified additional proteins that nonetheless belonged to the same biological functions. For instance, there are four Ssa proteins in the Hsp70 family in Saccharomyces cerevisiae. Three of the four Ssa proteins (Ssa1p, Ssa2p, and Ssa4p) were identified in iTRAQ1, whereas all four Ssa proteins were identified in iTRAQ2. As seen in the Venn diagram (Figure 5B), 241 identified proteins were seen in both iTRAQ experiments, whereas 23 proteins were uniquely identified in iTRAQ1 and 83 proteins were uniquely identified in iTRAQ2.
Comparison of functions of proteins identified in iTRAQ1 and iTRAQ2 to the yeast ORFs
iTRAQ1 (n = 264)
iTRAQ2 (n = 324)
Genome (n = 6334)
Small molecule metabolic process
Cellular nitrogen compound metabolic process
Response to stress
Cellular amino acid metabolic process
Carbohydrate metabolic process
Generation of precursor metabolites and energy
Cofactor metabolic process
Cellular component assembly
Lipid metabolic process
Cellular protein modification process
DNA metabolic process
Biological process unknown
Sulfur compound metabolic process
Cell wall organization or biogenesis
tRNA metabolic process
Macromolecular complex assembly
Anatomical structure development
Anatomical structure formation in morphogenesis
Ribonucleoprotein complex assembly
Nucleobase-containing compound catabolic process
Protein complex assembly
Cytoskeleton-dependent intracellular transport
Plasma membrane organization
Classification of extracellular proteins identified by iTRAQ
According to our iTRAQ data, a large number of extracellular proteins that we have identified were involved in the metabolisms of carbohydrates, amino acids, lipids, and nucleotides (Additional file1: Table S1). For the metabolism of carbohydrates, enzymes involved in glycolysis/gluconeogenesis, the pentose pathway, glycogen biosynthesis, the TCA cycle, and alcohol production were identified as present in the extracellular fraction (Additional file1: Table S1). The secretion of these enzymes in response to different stimuli has been reported in many studies; for example, glycolytic enzymes are secreted as immunogens during infection by Candida albicans[16, 38, 70], and GAPDH and enolase may be secreted during invasion by pathogens, as they bind to various mammalian proteins such as lysozyme, fibronectin, actin, myosin, and plasmin[16, 38, 70]. Elevated levels of glycolytic enzymes are secreted from various cancer cells[7, 26–29, 71]. Furthermore, enolase, GAPDH, 3-phosphoglycerate kinase, alcohol dehydrogenase, pyruvate kinase, fructose-1,6-bisphosphate aldolase, phosphoglycerate mutase and triose phosphate isomerase are commonly identified in the secretomes from bacteria, fungi, parasites and mammalian cells[6, 7, 22–24, 27, 37, 70]. In this current study, we have also identified enzymes involved in glycolysis, the pentose pathway, glycogen biosynthesis, alcohol metabolism, as were a large number of proteins involved in the metabolisms of amino acids and purines/pyrimidines.
Anti-oxidant proteins are important for detoxifying potentially damaging reactive oxygen and nitrogen intermediates. The anti-oxidant proteins superoxide dismutase and thioredoxin peroxidase have been reported in various secretomic studies[20, 27, 31, 36, 72]. Many of these anti-oxidant proteins were identified in our study. Heat shock proteins Hsp70, Hsp90, and Hsp97 constituted the largest protein family in the secretome from adult worms of Schistosoma japonicum. The immunodominant 47-kDa antigen in Candida albicans was the breakdown product of Hsp90 and was detected in patients with systemic candidiasis[38, 74]. The Hsp70s play important roles in protein folding, protein translocation across membranes, gene regulation, and protection from heat shock[75, 76]. Multiple protein-folding ATPase, heat shock proteins, and co-chaperones were also identified in the current study. Protein disulfide isomerases and cyclophilins may act with heat shock proteins to assist in protein folding and are frequently observed in many secretomic studies from different species[6, 26, 27, 36]. The current study also identified proline isomerase, proline rotamase, and protein disulfide isomerase.
Validation of extracellular proteins identified by iTRAQ
We next confirmed that proteins that are known to be present in the extracellular fraction/cell-surface were indeed distributed in the extracellular fraction. Scw4p and Scw11p are soluble cell wall proteins, and the majority of Scw4p and Scw11p was found in the extracellular fraction (Figure 6). We then determined the distribution of enzymes involved in carbohydrate metabolism. Tdh1p is a cell-surface protein[11, 38, 67], and was in the extracellular fraction (Figure 6). Pgk1p is 3-phosphoglycerate kinase that transfers high-energy phosphoryl groups from 1,3-bisphosphoglycerate to ADP during glycolysis and the reverse reaction during gluconeogenesis. Zwf1p is glucose-6-phosphate dehydrogenase that catalyzes the first and irreversible step of the pentose phosphate pathway. Pdc1p is the major pyruvate decarboxylase that decarboxylates pyruvate to acetaldehyde for ethanol production. Consistent with our iTRAQ results, Pgk1p, Pdc1p, and Zwf1p were detectable in the extracellular fraction (Figure 6). We also examined the distribution of proteins involved in oxidative stress. Sod1p is the copper-zinc superoxide dismutase that detoxifies superoxide, while Trx1p is thioredoxin that protects cells against oxidative and reductive stress. Both Sod1p and Trx1p were in the extracellular fraction (Figure 6). Finally, we examined the distribution of heat shock proteins. As mentioned above, Ssa1p is a cell-surface protein and was in the extracellular fraction (Figure 6). Hsc82p is required for folding of nascent polypeptides as well as for the refolding of denatured proteins. Hsp104p refolds and reactivates previously denatured and aggregated proteins. Hsc82p and Hsp104p were also present in the extracellular fraction (Figure 6). In contrast, Vps34p, Tor1p, and Tpk1p were distributed mostly in the intracellular fraction (Figure 6). Thus, proteins participate in many different biological functions including those involved in glycolysis/gluconeogenesis, the pentose pathway, alcohol production, oxidative stress, and protein folding are all present in the extracellular faction.
Validation of the t30/t0 ratios
We next used the t30a/t0a and t30b/t0b ratios from iTRAQ1 and the t30c/t0c ratio from iTRAQ2 to identify proteins that changed levels following a transfer to YPD (Additional file2: Table S2). The Protein Pilot Program calculates a p-value for each protein t30/t0 ratio based on the mean of the t30/t0 ratios of all the peptides belonging to that protein, and the scatter of those peptide ratios. We highlighted in bold the protein t30/t0 ratio whose calculated p-value was less than 0.05 (Additional file2: Table S2). Also highlighted (Additional file2: Table S2) are a total of 230 proteins that contained at least one of the three t30/t0 ratios having p-values smaller than 0.05. Because gluconeogenic enzymes decrease their levels after cells are transferred to YPD, they should have reduced t30/t0 ratios. Consistent with this idea, the t30/t0 ratios for these gluconeogenic enzymes were reduced. In a similar manner, significantly decreased t30/t0 ratios were observed for enzymes involved in glycolysis, the pentose pathway, alcohol production, glycogen biosynthesis, and the TCA cycle. Moreover, proteins involved in oxidative stress, in protein folding, and in other functions also exhibited a significant reduction in t30/t0 ratios (Additional file2: Table S2).
Small Vesicles disappear in response to glucose re-feeding
In this study, we used wild-type cells that were grown in YNB-based or YPKG-based media to examine glucose effects in up-regulating Lia1p and down-regulating Fbp1p. Amounts of Lia1p and Fbp1p were low in cells grown in YNB-based media and were higher in cells grown in YPKG. Adding glucose directly to the existing YPKG media produced a slow down-regulation of Fbp1p, whereas up-regulation of Lia1p was not observed. One possible explanation for these results is that metabolites or wastes present in the existing media prevented a fast increase in Lia1p levels and a fast decrease in Fbp1p levels. In contrast, changing the culture media from YPKG to YPD produced a faster down-regulation of Fbp1p and a faster up-regulation of Lia1p. Therefore, the YPKG to YPD condition is appropriate to study glucose effects in regulating protein levels. Down-regulation of gluconeogenic enzymes under this experimental condition is caused by degradation of pre-existing proteins in the vacuole. In contrast, when cells were grown in the presence of high glucose for a prolonged period of time, gluconeogenic enzymes are either not expressed or expressed at low levels[90–94]. Low expression levels of these enzymes are likely a result from low rates of protein synthesis. Therefore, the underlying mechanisms for catabolite inactivation of gluconeogenic enzymes in cells that are transferred from YPKG to YPD are fundamentally different from low rates of synthesis of these enzymes in cells that are continuously grown in the presence of high glucose.
It is interesting that proteins that are involved in so many diverse functions are secreted from cells. The secretion of these metabolic enzymes may be dependent on the needs of cells. For example, metabolic enzymes may be secreted when the needs inside the cells are decreased. If this is true, the extracellular space might be used as a storage place for these proteins. It is possible that secreted proteins have functions outside the cells similar to what they normally perform inside the cells. Alternatively, these proteins may have functions in the extracellular fraction unrelated to their functions in metabolism. For example, GAPDH and enolase bind to components of the extracellular matrix and are involved in the invasion of Candida albicans into host tissues[14, 38].
Our TEM studies indicated that small vesicles were present in the extracellular fraction in cells grown in YPKG and that they disappeared following a shift to YPD. A rapid disappearance of small vesicles from the extracellular fraction at the t = 30 time point may be caused by internalization into the cytoplasm or release into the culture media. The presence of extracellular vesicles in the culture media in Saccharomyces cerevisiae has been documented by Oliveira et al.. Interestingly, 82 proteins identified in extracellular vesicles isolated from the culture media overlapped with the proteins that we identified in iTRAQ1 (Figure 10C) and 87 proteins overlapped with iTRAQ2 (Figure 10D). In our study, cells were harvested by a low speed centrifugation and the culture media was discarded following centrifugation. The extraction procedure was then performed on cells that were pelleted. Therefore, the proteins that we identified are unlikely to be derived from the culture media.
Using the extraction procedure and Western blotting, we confirmed that extracellular proteins including Tdh1p, Pgk1p, Zwf1p, Pdc1p, Sod1p, Trx1p, Ssa1p, Hsc82p, and Hsp104p were in the extracellular fraction in cells grown in YPKG. All of these proteins showed reduced t30/t0 ratios in the iTRAQ experiments, suggesting that these proteins are cleared from the extracellular fraction at the t = 30 min time point, and the extraction protocol and Western blotting confirmed that levels of these extracellular proteins decreased following a transfer of cells to YPD for 30 min. These extracellular proteins were associated with membranes in the vesicle-enriched fraction. However, when membranes were disrupted by SDS, these proteins were in the 200,000 × g supernatant fraction, suggesting that the distribution of these proteins in the vesicle-enriched fraction is dependent on membrane integrity. There are several possible paths that might be involved in the observed decreases in extracellular proteins and vesicles upon a shift of cells to high glucose, such as internalization into the cytoplasm, degradation in the extracellular space, or release into the culture media. Future experiments will be needed to elucidate the molecular mechanisms responsible for the decline of protein levels in the extracellular fraction and the disappearance of small vesicles from the extracellular fraction.
Using the iTRAQ technique, we identified 347 proteins that were present in the extracellular fraction. The majority of these extracellular proteins do not contain ER signal sequences, suggesting that they are secreted via a non-classical pathway. Using the extraction procedure and Western blotting, we confirmed that enzymes involved in glycolysis/gluconeogenesis, the pentose pathway, alcohol production, oxidative stress, and protein folding were in the extracellular fraction. Furthermore, their levels were all reduced in response to changes to glucose enriched media. Our TEM studies indicated that there was a rapid disappearance of small vesicles from the extracellular fraction at the t = 30 min time point. We also observed a rapid decline in protein levels for a large number of extracellular proteins upon a transfer of cells to YPD for 30 min. We suggest that the secretome undergoes dynamic changes during transition from glucose-deficient to glucose-rich media. The plasticity of the secretome may be required to maintain the function and integrity of the cell depending on its physiological or nutritional states.
Cell culture, media and antibodies
Wild-type (BY4742, MATα his3Δ1 leu2Δ0 lys2Δ0 ura3Δ0) and wild-type cells expressing GFP tagged proteins were inoculated from -70°C frozen stock into 2 ml YPD (1% yeast extract, 2% peptone, and 2% glucose) in sterile glass tubes at 30°C in an environmental shaker at 250–300 rpm for 16 hours. Cell densities were measured using a Beckman DU640B spectrophotometer. Cells (50 μl) were diluted to OD600 = 0.01/ml in 10 ml YNB (0.67% yeast nitrogen base with amino acids and 2% glucose) or in 10 ml YPKG (1% yeast extract, 2% peptone, 1% potassium acetate, and 0.5% glucose), in sterile glass tubes for 72 hours at 30°C until cell density reached OD600 = 4-5/ml. The YNB media was supplemented with 0.1 ml of the 100 × of solution containing 200 mg uracil, 200 mg L-histidine, 300 mg L-leucine, and 300 mg L-lysine. Cells containing OD600 = 10/ml were aliquoted and used for each time point. In experiment I, 2% glucose was added directly to the existing YNB culture and incubated for 0, 2, and 4 hours at 30°C. In experiment II, cells were pelleted by a low speed centrifugation at 3,000 × g for 10 min and resuspended in fresh YNB-based media containing 2% glucose for 0, 2, and 4 hours at 30°C. In experiment III, 2% glucose was added directly to the existing YPKG culture for 0, 2, and 4 hours at 30°C. In experiment IV, cells were pelleted and resuspended in YPD containing 2% glucose for 0, 2, and 4 hours at 30°C. Cells grown in YPKG also pelleted and resuspended in YP (without added glucose) for 0, 2, and 4 hours at 30°C. Cells (OD600 = 10/ml) were harvested at each time point and processed. Levels of Lia1p and Fbp1p, and Tpi1p were examined by Western blotting. For re-growth experiments, wild-type cells were grown in 10 ml YPKG for 3d at 30°C in an environmental shaker until OD600 = 4-5/ml. Cells were pelleted and diluted to OD600 = 0.2/ml in 10 ml YPD (1% yeast extract, 2% peptone, 2% glucose), 10 ml YPKG (1% yeast extract, 2% peptone, 1% potassium acetate, and 0.5% glucose) or 10 ml YP (1% yeast extract, 2% peptone) at 30°C. Cell densities were measured at OD600 for 0, 2, 4, 6, and 8 hours using a Beckman DU640B spectrophotometer.
Extraction and Western blotting
Extraction of extracellular proteins was performed as described[2, 64, 77]. Wild-type cells and wild-type cells expressing GFP tagged proteins were inoculated from -70°C frozen stock into 2 ml YPD culture in sterile glass tubes in an environmental shaker at 250–300 rpm at 30°C for 16 hours. Cells (50 μl) were cultured into 10 ml of YPKG for 72 hours at 30°C. Cell densities were measured and cells (OD600 = 10/ml) were harvested (t = 0) or transferred to YPD containing glucose in an environmental shaker at 250–300 rpm at 30°C for 15, and 30 min (t = 15 and t = 30). At each time point, 10 mM NaN3 (final concentration) was added to stop the reactions. Cells were harvested by a low speed centrifugation at 3,000 × g for 10 minutes. Following centrifugation, the culture media was removed and cell pellets were resuspended in 100 μl of extraction buffer that contained 0.1 M Tris pH 9.4 and 10 mM β-mercaptoethanol. Samples were placed in a 37°C water bath for 15 min at 200 rpm. Following the extraction procedure, cells were pelleted at 3,000 × g for 10 minutes. The extracellular proteins (100 μl) that were released into the supernatants were precipitated in 15% trichloroacetic acid (TCA), washed, and solubilized in SDS-PAGE sample buffer. The remaining cell-associated fraction was lysed and solubilized in SDS-PAGE sample buffer. The distribution of intracellular and extracellular proteins was examined by Western blotting using Western Lighting Plus ECL kit (Perkin Elmer). Wild-type cells expressing GFP tagged proteins were purchased from Invitrogen. Anti-GFP antibodies were purchased from Abcam. Anti-TPI antibodies were from Proteintech Group.
Trpan blue staining and FM4-64 uptake
Wild-type cells were grown in 10 ml YPKG for 3d at 30°C in an environmental shaker. Cells were either harvested at t = 0 min or transferred to YPD for 30 min at 30°C. Cells were resuspended in 200 μl PBS buffer containing 140 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 1.8 mM KH2PO4, pH 7.4 and incubated in the absence or presence of 2% Triton X-100 for 30 min. This was followed by incubation with 20 μl of 0.4% trypan blue in PBS for 30 min at room temperature. The same amounts of t = 0 and t = 30 cells were subjected to the extraction procedure and pelleted. Extracted cells were resuspended in 200 μl PBS and incubated in the absence or presence of 2% Triton X-100 for 30 min. Treated and non-treated cells were then incubated with 20 μl of 0.4% trypan blue for 30 min. Cells were observed using a Zeiss microscope.
For FM uptake experiments, wild-type cells were grown in 10 ml YPKG for 3d at 30°C. Cells were either harvested at t = 0 min or transferred to YPD media for 30 min (t = 30) at 30°C. Half of the cells were pelleted, resuspended in 200 μl PBS, and incubated with 1 μl FM (16 mg/ml) for 3 hours at 30°C. Another half of the cells were extracted, resuspended in 200 μl PBS and then incubated with 1 μl FM for 3 hours at 30°C. The distribution of FM was visualized using a Zeiss fluorescence microscope.
iTRAQ labelling and data analysis
Two TRAQ labelling procedures were performed following the instructions from Applied Biosystems with minor modifications. Wild-type yeast cells were grown in YPKG and either harvested (t = 0 min) or the same amounts of cells (OD600 =10/ml) were transferred to media containing 2% glucose for 30 min (t = 30 min). Extracellular proteins were extracted as described above and then precipitated in 15% TCA. Protein concentrations were determined using Bio-Rad DC assay. Extracted proteins were reduced with 5 mM of tris(2-carboxyethyl) phosphine at 60°C for 60 min, and alkylated with 4 mM iodoacetamide for 30 min at room temperature in the dark. Samples were digested with 10 μg of trypsin at 48°C for 16 hours. Peptides were labelled with iTRAQ tags, combined, dried and resuspended in 500 μl of 10 mM ammonium formate, pH 3.6, in 20% acetonitrile/80% water. Labelled peptides were subjected to SCX separation by loading onto a passivated Waters 600E HPLC system with a 4.6 × 250 mm PolySULFOETHYL AspartamideTM column (PolyLC, Columbia MD) followed by elution at a flow rate of 1 ml per minute with a 1 hour gradient going from 10 mM→666 mM ammonium formate in 20% acetonitrile, pH 3.6. After evaporating and resuspending the resulting 15 SCX fractions 3 times in a SpeedVac (final resuspension in 9 ml of 2% acetonitrile, 0.1% TFA), reverse phase C18 nanoflow-LC separation of the 15 resuspended SCX fractions was accomplished by loading each fraction onto a Chromolith CapRod column (150 × 0.1 mm, Merck) from a 5 ml sample loop on an LC-Tempo MALDI separation/plate spotting system at 2 ml per minute in 6.8% acetonitrile, 0.1% TFA, then eluting with a 50 minute gradient from 6.8% to 79% acetonitrile, 0.1% TFA at a flow rate of 2.5 ml per minute. A 3 ml per minute flow of MALDI matrix solution was added in-line post-column (7 mg/ml recrystallized CHCA (a-cyano-hydroxycinnamic acid), 2 mg/ml ammonium phosphate, 0.1% trifluoroacetic acid, 80% acetonitrile), and the resulting eluant was automatically spotted onto a stainless steel MALDI target plate every 6 seconds (0.55 μl per spot), for a total of 370 spots per original SCX fraction. Each target plate was analyzed in a data-dependent manner on either an ABI 5800 MALDI TOF-TOF or an ABI 4800 MALDI TOF-TOF. The MS spectra for an entire 2D-LD experiment were thus taken from 5500 MALDI spots, averaging 500 laser shots per spot at laser power 3400. In a data-dependent manner, 10,000-15,000 MS/MS spectra were taken of the strongest example of each unique m/z mass observed across each target plate, using up to 2500 laser shots per MS/MS spectrum at laser power 4100, with CID gas at 1.2 to 1.3 × 10-6 Torr.
Protein identification was accomplished using the Paragon Algorithm as implemented in ProteinPilotTM 4.0 software (ABSciex). Search parameters were set as cysteine alkylation: iodoacetamide, ID focus: biological modifications, search effort: thorough. Combined spectra were searched against the species-specific (S. cerevisiae) NCBI RefSeq database concatenated with a reversed “decoy” version of the same database plus 156 common lab contaminants as described by Keller et al.. To reduce the potential number of false positive protein identifications, the Paragon Algorithm search results were further filtered through the use of a very stringent Local False Discovery Rate (FDR) estimation calculated from the Proteomics System Performance Evaluation Pipeline (PSPEP)[98, 99]. We accepted protein IDs with an estimated local False Discovery Rate (FDR) estimate of <5%. A total of 318 extracellular proteins were identified in iTRAQ1 (Additional files3 and4) and 392 extracellular proteins were identified in iTRAQ2 (Additional files5 and6) using these criteria for acceptance. We next removed proteins that were identified by only a single confidently identified peptide in only one of the two iTRAQ experiments, (proteins that were identified with a single peptide with 95% confidence, but in both experiments, were included). From the identifiers (gi numbers) obtained from searching against the NCBI RefSeq database, we used the Saccharomyces Genome Database (Stanford University) and the Gene Ontology Slim Mapper to search for the function for each protein. BioVenn analysis was used to compare proteins identified from different studies.
Transmission electron microscopy
Immuno-TEM was performed as described previously[77, 101]. Wild-type cells were grown in 2 ml YPKG for 3d and harvested (t = 0 min) or transferred to YPD for 30 min and harvested (t = 30 min). The cells were fixed with 3% para-formaldehyde and 0.2% glutaraldehyde for 16 hours. Cells were then washed by centrifugation at 4,500 × g sequentially in 1 ml of 0.5 M sorbitol in 0.08 M potassium phosphate buffer (pH 6.7), 0.25 M sorbitol in 0.08 M potassium phosphate buffer (pH 6.7), and 0.08 M potassium phosphate buffer (pH 6.7). Cells were resuspended in 1 ml of sodium metaperiodate (1% w/v), incubated at room temperature for 20 min, re-suspended in 1 ml of 50 mM ammonium chloride, and then incubated for 15 min at room temperature. Cells were centrifuged at 4,500 × g and dehydrated by 5 min incubation in 1 ml 50% cold ethanol, 70% cold ethanol and 80% cold ethanol followed by 10 min incubation in 85% cold ethanol, 90% cold ethanol, 95% cold ethanol and 100% cold ethanol. Finally, 1 ml 100% ethanol was added to the cell pellet and incubated for 5 min at room temperature. After dehydration, cells were sequentially incubated in 1 ml LR White 2:1, ethanol:LR White at room temperature rotator for 2 hours, 1:1 ethanol:LR White resin at room temperature rotator overnight, 1:2 ethanol:LR White resin at room temperature rotator for 2 hours and 100% resin at room temperature rotator for 2 hours three times. Cells were transferred to gelatin capsules, dried and cut into 10 nm thin sections. Sections were placed onto grids and incubated with affinity purified Fbp1p antibodies followed by goat anti-rabbit antibodies conjugated with 10 nm gold particles (Ted Pella) and viewed on a JEOL JEM-1400 electron microscope.
For negative staining and TEM, wild-type cells were grown in 200 ml of YPKG in sterile glass flasks for 72 hours and split into equal aliquots based on cell density. Cells (100 ml) were transferred to YPD media containing 2% glucose for 30 minutes (t = 30 min). Cells (100 ml) from the t = 0 and t = 30 time points were harvested and resuspended in 4 ml extraction buffer containing 0.1 M Tris pH 9.4 and 10 mM β-mercaptoethanol at 37°C for 15 minutes. Total extracts from the t = 0 and t = 30 cells were centrifuged at 3,000 × g for 10 minutes at 4°C. The 3,000 × g supernatant was further centrifuged at 200,000 × g for 2 hours at 4°C. The resulting 200,000 × g pellets were resuspended in 30 μl of PBS. Samples were fixed with 3% para-formaldehyde and 0.2% glutaraldehyde for two hours at 4°C. Nickel grids were incubated with 10 μl of poly-lysine buffer for 5 minutes and washed with distilled water for 3 minutes. Total extracts (10 μl) were incubated with poly-lysine coated grids for 30 minutes. The grids were blotted with 0.1% BSA in PBS buffer. The grids were washed once with PBS buffer and twice with distilled water. The grids were then blotted with Whatman filter papers and stained with 2% uranyl acetate for 10 minutes in the dark. The grids were dried by blotting with Whatman filter papers and samples were viewed on a JEOL JEM-1400 electron microscope.
We thank the Mass Spectrometry and Proteomics Facility at Penn State College of Medicine for performing 2D-LC/MALDI-MS/MS. TEM studies were performed at the Electron Microscopy Core Facility at Penn State College of Medicine. This work was supported by a Penn State Bridge Fund to Hui-Ling Chiang.
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