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Genes to Cells (2009) 14, 861-870. doi:10.1111/j.1365-2443.2009.01316.x
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Lag-phase autophagy in the methylotrophic yeast Pichia pastoris

 Shun-ichi Yamashita1, Hiroya Yurimoto1, Dai Murakami1, Mari Yoshikawa1, Masahide Oku1 and Yasuyoshi Sakai1,2,*

1 Division of Applied Life Sciences, Graduate School of Agriculture, Kyoto University, Kyoto 606-8502, Japan
2 CREST, Japan Science and Technology Agency, 5, Sanbancho, Chiyoda-ku, Tokyo 102-0075, Japan


   Abstract
Top
 Abstract
Introduction
 Results
 Discussion
 Experimental procedures
 References
 
When microbes sense environmental changes, they often temporarily attenuate cell growth to adapt to the new situations, showing a lag phase. In this study, we report that the methylotrophic yeast, Pichia pastoris, induced autophagy during the lag phase after the cells were shifted from glucose to methanol medium. Through the autophagic process at least two proteins, aminopeptidase I precursor and cytosolic aldehyde dehydrogenase, were found to be transported into the vacuole, which was dependent on PpAtg11 and PpAtg17, respectively. Notably, PpAtg1 and PpAtg17 were required for early exit from the lag phase during the methanol adaptation. In accordance, phosphorylation states of elongation initiation factor 2{alpha} indicated reductions of intracellular amino-acid pools in the atg mutant strains. Together, these data demonstrate the importance of amino acid recycling by autophagy during a cell-remodeling process.


   Introduction
Top
 Abstract
 Introduction
Results
 Discussion
 Experimental procedures
 References
 
Environmental changes often induce reorganizations of intracellular components in microbes. Such reorganization process has extensively been studied using the methylotrophic yeast, Pichia pastoris. After transferred to methanol medium, this organism undergoes a massive proliferation of peroxisomes while inducing syntheses of more than 10 unique enzymes for methanol metabolism, including peroxisomal alcohol oxidase (Aox). The strong methanol induction of protein synthesis in P. pastoris has been utilized as an ideal heterogeneous gene expression system (Cereghino & Cregg 2000).

Shifting P. pastoris wild-type cells from synthetic glucose medium to synthetic methanol medium brings about a delay of cell growth (See Results). This lag phase is thought to be a period during which the cells remodel their metabolism and/or intracellular structures, as elucidated by pioneering studies on diauxic growth (Monod 1942; Roseman & Meadow 1990). Here, we report an autophagic activity in the lag phase.

Autophagy is defined as degradation system for cytosolic components after they are transported into the lysosome/vacuole. It is conserved from yeast to higher eukaryotes, and is involved in a wide variety of physiological processes, including adaptation to neonatal starvation (Kuma et al. 2004) and cellular defense against invading bacteria (Nakagawa et al. 2004).

In P. pastoris, several types of autophagic pathways have been described; for example, pexophagy is the selective transport of peroxisomes, and the cytoplasm-to-vacuole (Cvt) pathway selectively transports precursor aminopeptidase I (prApeI) (Sakai et al. 1998; Farre et al. 2007). These pathways are observed under different metabolic conditions: pexophagy is induced when peroxisomes become dispensable for cellular metabolism (for instance, when the carbon source of the culture is shifted from methanol to other ones), and the Cvt pathway is active during vegetative growth. Both of these pathways require PpAtg11. This molecule and its Saccharomyces cerevisiae ortholog can bind to receptor proteins for several specific protein complexes or organelles and act as scaffolds for the recruitment of other Atg proteins (Kim et al. 2001; Yorimitsu & Klionsky 2005). In contrast, under starvation conditions, Atg17 functions in the recruitment of the Atg proteins to induce bulk autophagy, termed macroautophagy (Kabeya et al. 2005; Cheong et al. 2008; Kawamata et al. 2008). In this study, we show that both PpAtg11 and PpAtg17 are involved in the autophagy induced during the lag phase. The two proteins contributed to the autophagy differently in terms of their cargo specificities and physiological impact, as now reported below.


   Results
Top
 Abstract
 Introduction
 Results
Discussion
 Experimental procedures
 References
 
Induction of autophagy during lag phase after carbon-source shift

When wild-type P. pastoris cells were transferred from synthetic glucose medium to synthetic methanol medium, they showed an ca. 6-h lag phase and subsequently proceeded to a growing (log) phase (Fig. 1A). Since the lag phase was not evident when the cells were transferred to methanol medium supplied with 20 different amino acids each at 0.1 mg/mL concentration (Fig. 1A), we speculate that the cells in the lag phase may be under amino acid starvation condition, although the cells are endowed with a complete set of pathways for amino acid syntheses.


Figure 1
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  		Figure 1  Induction of autophagy after shifting the carbon source to methanol. (A) Growth curves of a P. pastoris wild-type strain SY305 after transfer from glucose to methanol medium. The cells were grown in synthetic glucose medium, and transferred to either synthetic methanol medium that contains no amino acids (SM, open circles) or methanol medium supplemented with 0.1 mg/mL amino acids (SM + amino acids, filled circles). The OD600 value is plotted in the log scale. (B) Immunoblot analysis detecting 3xMyc-PpAtg8 expressed in the wild-type strain, SY201. The upper panel shows data obtained from SDS–PAGE and the lower panel shows data from SDS–PAGE with 6 M Urea. The numbers indicate time after transfer to methanol medium (hours). (C) Localization of YFP-PpAtg8. The left column represents the signal pattern of YFP-PpAtg8, the middle column shows fluorescence from YFP-PpAtg8 (green) and FM4-64 (red) staining the vacuolar membrane, and the right column represents the corresponding brightfield images (DIC). The time points of microscopic observation after the transfer to methanol medium are indicated at the top of the panels. Bar, 2 µM. (D) Immunoblot detection of YFP-PpAtg8 at the indicated time points after transfer to methanol medium. The band positions corresponding to YFP-PpAtg8 and its processed form (YFP*) are indicated by arrows.

 
As the expression levels and lipidation of ScAtg8 are increased in response to nutrient starvation so that macroautophagy can be induced (Kirisako et al. 1999; Ichimura et al. 2000), we examined changes in PpAtg8 during the lag phase. Similar to the nitrogen-starvation case, the expression levels of 3xMyc-PpAtg8 and the amount of its lipidated form increased after the carbon-source shift (Fig. 1B).

Next, Yellow Fluorescent Protein (YFP)-tagged PpAtg8 was utilized as a marker of autophagic membrane flux, as this protein is localized on autophagic membranes (Kirisako et al. 1999; Kabeya et al. 2000). Fluorescence microscope showed that some of the YFP signal showed a diffuse pattern inside the vacuole 3 h after the carbon-source shift, in addition to showing a dot pattern in the cytoplasm. After 6 h, the signal was localized exclusively inside the vacuole (Fig. 1C). This result suggests that the transport of PpAtg8 into the vacuole, a hallmark of autophagy, occurs in the lag phase after the carbon-source shift.

The transport of PpAtg8 was investigated in several mutant strains (Fig. 2A). Atg1 is a protein kinase required for autophagic processes, and Ypt7 is involved in fusion of the autophagic and vacuolar membranes. In Ppatg1{Delta} and Ppypt7{Delta} strains, the transport of YFP-PpAtg8 into the vacuole was not observed; instead multiple dots of YFP-PpAtg8 signal were observed in the cytoplasm of Ppypt7{Delta} cells. These data support the notion that the observed trafficking of YFP-PpAtg8 represents autophagy.


Figure 2
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  		Figure 2  Transport of YFP-PpAtg8 into the vacuole in Ppatg mutants after shifting the carbon source to methanol. (A) The denoted atg and ypt7 mutants harboring YFP-PpAtg8 were cultured and subjected to fluorescence microscope as described in Fig. 1C. Bar, 2 µM. (B) Immunoblot detection of YFP-PpAtg8 and its processed form was carried out as described in Fig. 1D.

 
Biochemical detection of YFP derived from YFP-PpAtg8 also indicated autophagic activity (Fig. 1D). When a fluorescent protein-tagged Atg8 is transported into the vacuole via autophagy and exposed to vacuolar proteinases, it is rapidly subjected to digestion, leaving behind a "free form" of the fluorescent protein that is structurally more resistant to the proteinases (Yorimitsu et al. 2006). Hence detecting the free form of the fluorescent protein can be a measure of autophagic activity. In our experimental system, the free YFP form was detected from the samples acquired 3 and 6 h after the carbon-source shift (Fig. 1D).

The kinetics of YFP fluorescence detection in the vacuole by microscope (Fig. 1C) was slower than that of free-YFP detection by immunoblot (Fig. 1D). Since the Atg8 expression is induced gradually after the carbon-source shift (Fig. 1B) and maturation of YFP moiety takes some period, it is possible that a fraction of YFP-PpAtg8 is transported into the vacuole before it becomes fluorescent. Considering the difference of the kinetics between the two assays, these data indicate that an autophagic activity is induced during the lag phase; thus the observed phenomenon is hereafter designated lag-phase autophagy (LPA).

PpAtg11 and PpAtg17 involvements in LPA

To investigate the molecular requirements of LPA, we observed the localization of YFP-PpAtg8 in several atg mutant strains. As in the wild-type strain (Fig. 1C), the YFP-PpAtg8 fluorescence showed a diffuse pattern in the vacuole of Ppatg11{Delta} and Ppatg17{Delta} cells 6 h after the carbon-source shift (Fig. 2A). In contrast, in Ppatg11{Delta} Ppatg17{Delta} double-knockout cells, the YFP-PpAtg8 signal was not observed in the vacuole. These results suggest that either of these two proteins may be required for LPA. In line with this, the YFP-PpAtg8 signal was not observed in the vacuole in Ppatg1{Delta} cells, in which both the PpAtg11- and PpAtg17-driven processes are thought to be impaired.

We also examined the existence of free YFP form derived from YFP-PpAtg8 in these mutants. In lysates from both Ppatg11{Delta} and Ppatg17{Delta} cells, the free-form YFP was detected, although not to as great an extent as from the wild-type sample. In Ppatg1{Delta} cells, the free YFP was not detected (Fig. 2B). These results demonstrate the contributions of PpAtg1, PpAtg11 and PpAtg17 to LPA.

PpAtg11-dependent transport of aminopeptidase I (Ape1) precursor into the vacuole during LPA

The involvement of PpAtg11 in LPA suggests a selective cargo transport into the vacuole. In S. cerevisiae, prApeI is a majority cargo of the Cvt pathway and is processed into a mature form inside the vacuole (Harding et al. 1995). To test whether LPA delivers P. pastoris prApe1 (PpprApe1) to the vacuole, we expressed Cyan Fluorescent Protein (CFP)-PpprApe1.

We investigated the localization of CFP-PpprApe1 after the carbon-source shift to methanol. In wild-type and Ppatg17{Delta} cells, CFP-PpprApe1 fluorescence localized to the lumen of the vacuole, and in addition, to a dot structure in the cytoplasm. In contrast, in Ppatg1{Delta} and Ppatg11{Delta} cells, the CFP-PpprApe1 signal localized only to the dot in the cytoplasm (Fig. 3A). This result shows that the transport of PpprApe1 into the vacuole during LPA is dependent on PpAtg1 and PpAtg11. Interestingly, the CFP-PpprApe1 dots 3 h after the carbon-source shift were less frequently associated to the vacuolar surface in Ppatg11{Delta} than in the other strains. This tendency is also observed as for the YFP-PpAtg8 localization (Fig. 2A) and may reflect the fact that PpAtg11 is found on the vacuolar surface (Oku et al. 2006).


Figure 3
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  		Figure 3  Transport of CFP-PpprApe1 to the vacuole after shifting the carbon source to methanol. (A) Localization pattern of CFP-PpprApe1 in the wild type (WT) and denoted atg mutants 3 and 6 h after the medium shift. As for the wild-type strain, the localization in glucose medium is also shown. The CFP signal is presented alone (left) or merged with the red FM-4-64 signal (middle). The corresponding brightfield images are also shown (right). Bar, 2 µM. (B) Immunoblot analysis of cell lysates from the strains used in (A). Lysates were acquired 3 h after the cells were transferred to methanol medium. The band positions corresponding to CFP-PpprApe1 (full length, shown as FL) and its processed form (truncated form, shown as TR) are indicated.

 
The PpprApeI transport was confirmed by immunoblot analysis detecting a band corresponding to CFP plus Ape1 propeptide processed inside the vacuole (Shintani et al. 2002; Suzuki et al. 2002). This band was detected in the samples from wild-type and Ppatg17{Delta} cells, but not in the samples from Ppatg1{Delta} or Ppatg11{Delta} cells (Fig. 3B).

A previous study on PpprApe1 dynamics in the P. pastoris Cvt pathway reported that loss of PpAtg26 reduces the efficiency of the transport into the vacuole, in contrast to the case of S. cerevisiae, in which Atg26 is dispensable for the Cvt pathway (Cao & Klionsky 2007; Farre et al. 2007). In accordance with that study, Ppatg26{Delta} cells showed a partial defect in CFP-PpprApe1 transport during LPA, as shown by microscopic (Fig. 3A) and immunoblot (Fig. 3B) analyses. These results indicate that the PpAtg11-dependent LPA of PpprApe1 utilizes PpAtg26 as a facilitator.

PpAld6 transport to the vacuole during LPA is dependent on PpAtg17

We next aimed to investigate the function of PpAtg17 in LPA. Although S. cerevisiae Atg17 functions in bulk transport of cytoplasmic components for accomplishing macroautophagy, another report identified cytosolic aldehyde dehydrogenase Ald6 as a "preferential" cargo of the macroautophagic pathway (Onodera & Ohsumi 2004). Hence, we examined whether PpAld6 was transported to the vacuole through LPA. For this purpose, we expressed PpAld6-CFP in a strain devoid of two vacuolar proteases (pep4{Delta}prb1{Delta}) (Gleeson et al. 1998) where accumulation of autophagic bodies is prominent. Several atg mutant strains were subsequently generated from this strain.

Six hours after the carbon-source shift, the PpAld6-CFP signal was observed inside the vacuole of the pep4{Delta}prb1{Delta} strain (Fig. 4A). The fluorescence of PpAld6-CFP in the vacuoles often formed patchy patterns beneath the vacuolar surface, and colocalized with the signals from FM 4-64 dye (Fig. 4A). As a control, CFP is expressed alone under regulation of a strong GAP (glyceraldehyde-3-phosphate dehydrogenase) promoter in the same proteinase-deficient strain. Fluorescence microscope of the CFP after the carbon-source shift to methanol gave much weaker signal of the fluorescence inside the vacuole (Fig. 4A). This result suggests the preferential transport of PpAld6 by LPA.


Figure 4
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  		Figure 4  Transport of PpAld6-CFP to the vacuole after shifting the carbon source to methanol. (A) Localization of CFP or PpAld6-CFP 6 h after transfer to methanol medium in wild type (WT) and the designated atg mutants derived from a vacuolar protease-deficient strain, SMD1163. (B) Electron microscopy of the wild-type (WT) strain SY305 was carried out 6 h after the shift from glucose to methanol medium (upper image) or to nitrogen starvation [SD(-N)] medium (lower image) in the presence of 1 mM PMSF. Autophagic bodies are indicated by arrows. Bar, 1 µM. (C) PpAld6-expressing cells of the denoted atg mutant strains with intact vacuolar proteases were acquired at the indicated time points after the medium shift, and subjected to immunoblot analysis. The band positions corresponding to PpAld6-CFP (FL) and its processed form (TR) are indicated.

 
The detailed morphology of the intra-vacuolar structures formed in the process of LPA was examined by electron microscope (Fig. 4B). Wild-type cells were transferred to synthetic methanol medium (for the observation of LPA) or nitrogen-starved glucose medium (for the observation of macroautophagy) in the presence of PMSF to avoid the degradation of the intra-vacuolar structures. Although the size of the structures in LPA was similar to that of macroautophagic bodies observed under nitrogen-starvation condition (200–300 nm diameter), most of the intra-vacuolar structures in LPA were found in a cluster near the limiting membrane of the vacuole, in contrast to the scattered pattern of macroautophagic bodies.

The PpAld6 signal inside the vacuole was observed when PpATG11 was furthermore deleted from the parental pep4{Delta}prb1{Delta} strain, but not when PpATG1 or PpATG17 was deleted (Fig. 4A), showing that PpAld6 is transported to the vacuole dependent on PpAtg1 and PpAtg17.

The dependence of PpAld6-CFP transport on PpAtg17 was confirmed by immunoblot analysis detecting free CFP (processed in the vacuole) in lysate samples from strains harboring PpAld6-CFP and intact vacuolar proteases (Fig. 4C). Before the carbon-source shift (0 h), no samples showed free CFP. In contrast, the free form was detected in samples from wild-type and Ppatg11{Delta} cells obtained after 6-h shift to methanol medium, but not from Ppatg1{Delta} or Ppatg17{Delta} cells. These results indicate that PpAtg17-dependent LPA delivers PpAld6 to the vacuole.

PpAtg1 and PpAtg17 are needed for optimal Aox synthesis and early exit from lag phase

To search for the physiological roles of LPA, we compared the growth curves of wild-type cells and atg mutants after the carbon-source shift. Intriguingly, the lag phases of the Ppatg1{Delta} and Ppatg17{Delta} strains were clearly prolonged (Fig. 5A). Under the tested condition, wild-type cells proceeded to log phase 6 h after the carbon-source shift, whereas Ppatg1{Delta} and Ppatg17{Delta} cells 18 h after the shift (Fig. 5A). In contrast, PpATG11 deletion had no effect on the length of the lag phase (Fig. 5A). These results suggest that PpAtg1 and PpAtg17 are required for early exit from the lag phase.


Figure 5
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  		Figure 5  Kinetics of methanol adaptation in atg mutants. (A) Growth curves of the wild-type (SY305) and denoted atg mutants (SY306–308) after the medium shift were plotted as in Fig. 1A. (B) Immunoblot analysis of Aox was carried out as for the samples from the strains used in (A) acquired at the indicated time points after the medium shift (hours).

 
When the Ppatg17{Delta} cells were transferred to methanol medium supplied with amino acids, they did not undergo a detectable lag phase and showed a similar growth profile to that of wild type (Fig. 5A), suggesting that the prolonged lag phase in the Ppatg17{Delta} strain results from shortage of amino acid pool.

Next we investigated the kinetics of protein synthesis required for growth on methanol. Aox, the primary enzyme for methanol metabolism, is dramatically induced at the transcriptional level, and is indispensable for growth on methanol (Ellis et al. 1985). We examined the abundance of this protein as a typical reporter for protein synthesis after carbon-source shift. Aox synthesis was remarkably delayed in Ppatg1{Delta} and Ppatg17{Delta} cells, but was induced normally in Ppatg11{Delta} cells (Fig. 5B), suggesting that PpAtg1 and PpAtg17 are required for optimal Aox synthesis. Taken together with the growth profiles, these results strongly suggest that PpAtg1 and PpAtg17 are required for providing a sufficient amino acid supply for the cells to proceed into the log phase.

Phosphorylation of eIF2{alpha} is enhanced in LPA-deficient strains

The idea that LPA contributes to the supply of amino acids led us to speculate that mutants defective in LPA may undergo more severe amino acid starvation at the lag phase. To test this notion, we estimated the activity of Gcn2. This protein phosphorylates eukaryotic initiation factor 2{alpha} (eIF2{alpha}), a component of the ternary complex essential for global translation in eukaryotic cells. The activity of Gcn2 is up-regulated by the levels of free charged tRNAs, which increase in response to a reduction of the cytosolic amino acid pool. Thus, the phosphorylation state of eIF2{alpha} is regarded as a reporter of amino acid starvation. Therefore, we examined the phosphorylated form of eIF2{alpha} (P-eIF2{alpha}) to determine whether the carbon-source shift to methanol causes amino acid starvation.

In a wild-type strain, the intensity of the P-eIF2{alpha} signal increased and peaked at 6 h after the carbon-source shift (Fig. 6). This result suggests an amino acid starvation at the lag phase after the carbon-source shift.


Figure 6
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  		Figure 6  Phosphorylation of eIF2{alpha} after shifting the carbon source to methanol. Immunoblot analysis of phosphorylated eIF2{alpha} (P-eIF2{alpha}) was carried out as for the samples from the same strains used in Fig. 5. The numbers represent time points of sample acquisition after the carbon-source shift (hours).

 
Furthermore immunoblot analysis indicated that the intensities of the P-eIF2{alpha} signals in Ppatg1{Delta} and Ppatg17{Delta} samples remained above the wild-type level (Fig. 6). In addition, it is of note that even in lysates from the Ppatg11{Delta} strain, which showed almost no growth phenotypes under these conditions, the P-eIF2{alpha} signal intensity was higher than that from the wild-type strain. These results support the notion that LPA is required for sufficient amino acid supply that ensures optimal protein synthesis for cells to transit from the lag phase to the growth phase.


   Discussion
Top
 Abstract
 Introduction
 Results
 Discussion
Experimental procedures
 References
 
After P. pastoris cells are switched to synthetic methanol culture, they undergo a lag phase during which they reconstitute intracellular organizations for methanol metabolism. Since the lag phase was not evident in case of methanol culture with sufficient amino acid supply (Figs 1A and 5A), the lag phase in the synthetic methanol culture is thought to result from the shortage of amino acid pool. Here we discover and characterize autophagy induced in the lag phase (LPA). Deleting any of the tested PpATG genes (PpATG1, PpATG11 and PpATG17) increased the phosphorylation of eIF2, implying lower levels of intracellular amino acid pool in the gene-disrupted cells (Fig. 6). Furthermore, PpAtg1 and PpAtg17 are found to be necessary for the optimal synthesis of Aox and early exit from the lag phase (Fig. 5A). Taken together these findings suggest that the physiological role of LPA is to recycle amino acids for the efficient synthesis of proteins necessary for the new metabolism. A previous study showed the role of amino acid recycling by autophagy in maintaining cell viability during nitrogen starvation (Onodera & Ohsumi 2005). Moreover, physiological roles of autophagy in the amino acid supply or efficient protein synthesis have been elucidated in several steps of mammalian development (Kuma et al. 2004; Tsukamoto et al. 2008). Our current results have physiological relevance to these pioneering studies.

In the process of LPA characterization we found that the cytosolic precursor form of aminopeptidase I (PpPrApeI) was transported to the vacuole dependent on PpAtg11 (Fig. 3). Previously, the transport of PrApeI through Cvt pathway was found to be a constitutive process in a growing phase (Wang & Klionsky 2003). In contrast, most of the fluorescence from CFP-PpprApe1 was found in the cytoplasm in glucose culture and localized inside the vacuole after the shift to methanol culture (Fig. 3A), indicating induction of PpprApeI transport. The molecular basis underlying this induction is yet to be elucidated, but it is of note that the transcript level of S. cerevisiae ATG11 is greatly enhanced when the cells are transferred from fermentative to respiratory medium (Roberts & Hudson 2006). Similar inductions of autophagic components may give rise to the PpprApeI transport.

We also found that PpAld6 is preferentially transported through LPA (Fig. 4). We detected bulk transport of cytosolic components into the vacuole during LPA by expressing cytosolic CFP, but the extent of the transport was much lower than that of PpAld6-CFP transport (Fig. 4A). Furthermore, the morphology of intra-vacuolar structures observed during LPA was distinct from that of macroautophagic bodies observed during bulk transport (Fig. 4B). Intriguingly, deletion of PpALD6 shortened the lag phase after methanol shift (data not shown), which implies the physiological importance of PpAld6 degradation under this condition. Thus, although the main physiological role of LPA seems to be the amino acid recycling for the new metabolism, it is also plausible to assume that the LPA functions in removing a specific set of proteins that cause the growth attenuation after the carbon-source shift.


   Experimental procedures
Top
 Abstract
 Introduction
 Results
 Discussion
 Experimental procedures
References
 
Strains and media

The strains used in this study are listed in Table 1. The strains were grown in YPD (1% yeast extract, 2% peptone, 2% glucose), SD (0.67% yeast nitrogen base without amino acids, 2% glucose, supplemented with the appropriate amino acids), or SM (0.67% yeast nitrogen base without amino acids, 0.8% methanol, supplemented with the appropriate amino acids). Methanol medium supplemented with amino acids contain 0.1 mg/mL of 20 different L-amino acids.


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  		Table 1  Pichia pastoris strains used in this study
 
Plasmids

The oligonucleotide primers used in this study are listed in Table 2. The plasmid expressing YFP-PpAtg8 (pSAP115) has been described previously (Mukaiyama et al. 2004). For the expression vector harboring the PpACT1 promoter, the upstream region of PpACT1 was amplified by PCR using P. pastoris PPY12 genomic DNA as a template and oligonucleotide primers, ACT1P-Fw-Nde and ACT1P-Rv-Eco. Subsequently, the fragment was digested by NdeI and EcoRI and ligated into pIB1 (Sears et al. 1998) to generate pSY001. For the expression of N-terminal and C-terminal CFP tagged constructs, CFP fragments were amplified by PCR using pECFP-N1 (Clontech, Mountain View, CA) as a template and the primer sets, N-CFP-Fw-Eco, N-CFP-Rv-Bam and C-CFP-Fw-Hind, C-CFP-Rv-Hind. Subsequently, the fragments for N-terminal and C-terminal tagging were digested and ligated into the EcoRI and BamHI sites of pSY001 and the HindIII site of pIB1, yielding pSY003 and pSY004, respectively. For the expression of CFP-PpprAPE1, the PpprAPE1 open reading frame was amplified by PCR using P. pastoris PPY12 genomic DNA as a template and the primer set, APE1-Fw-Bgl and APE1-Rv-Xho, digested with BglII and XhoI and ligated into the BamHI and XhoI sites of pSY003, yielding pSY413. For expressing CFP alone, the CFP-coding DNA fragment was cloned into pIB2 (Sears et al. 1998), which gives pSY004. For the expression of PpALD6-CFP, PpALD6 was amplified by PCR using PPY12 genomic DNA as a template and the primer set, RPPA05532-Up-Fw-Bgl and RPPA05532-Rv-Xho. The fragment containing the promoter region of PpALD6 was digested with BglII and XhoI and ligated into pSY004 at the BamHI and XhoI sites, yielding pSY417.


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  		Table 2  Oligonucleotide primers used in this study
 
Growth conditions for the induction of autophagy-related pathways

Pichia pastoris cells which were complemented for auxotrophy by transformation with PpARG4 and PpHIS4 vectors were grown to stationary phase on YPD medium; subsequently this culture was transferred to fresh YPD medium and grown to mid-log phase. The cells were transferred to SD medium and grown to early log phase (approximately 5 h). The cells were harvested by centrifugation at room temperature and washed with yeast nitrogen base at once. Subsequently, the cells were resuspended in SM or SD-N, and were cultured in a rotary shaker at 28 °C. Cells for microscope and biochemical analysis were sampled at each time point after medium shift.

Electron microscope

Cells were subjected to rapid freezing and freeze-substitution fixation, and observed as previously reported (Kirisako et al. 1999).

Fluorescent microscope

To label the vacuolar membrane, cells were transferred to SD medium with 0.93 µg/mL FM4-64 (Molecular Probes, Eugene, OR) and were grown to early log phase (approximately 5 h). The cells were shifted to SD-N or SM medium and cultured at 28 °C. Fluorescent microscope was carried out as described previously (Yamashita et al. 2006).

Biochemical methods

AntiGFP polyclonal antibody was purchased (Molecular probes). SDS–PAGE gels containing 6 M Urea treated with AG 501-X8 resin were used to detect the lipidated form of PpAtg8. Cell lysates were prepared at each time point after methanol adaptation by Multi-beads shocker (Yasui Kikai, Ohsaka, Japan) with lysis buffer containing 50 mM Tris–HCl, pH7.5, 1 mM EDTA, 1 mM PMSF, protease inhibitor cocktail (Complete EDTA free, Roche Diagnostics, Mannheim, Germany), and phosphatase inhibitor (PhosSTOP, Roche Diagnostics). Lysate concentrations were calculated by the Bradford method (Bio-Rad, Hercules, CA), and equal amounts of lysates were denatured by boiling with SDS sample buffer and loaded onto the gel. After blotting, the blot was blocked for more than 30 min using 5% skim milk in TBS-T buffer (containing, 2.42 g Tris, 8 g NaCl, 0.1% Tween-20, pH 7.6). Blots were incubated with antiGFP antibody, antiP-eIF2a antibody, or antiAox antibody (1 : 3000, 1 : 1000, or 1 : 10 000 dilutions, respectively) in TBS-T buffer for 60 min with gentle shaking. Blots were rinsed in TBS-T three times, and incubated with antiRabbit IgG HRP conjugate (1 : 10 000 dilution) in TBS-T for 30 min. After incuba-tion, blots were rinsed in TBS-T four times, and immunoreactive bands were detected using Western Lightning Chemiluminescence Reagent Plus (Perkin-Elmer Life Sciences, Waltham, MA). For quantitative analysis of protein signals, the bands were analyzed by densitometry using a CS Analyzer (ATTO and Rise Corporation, Tokyo, Japan).


   Acknowledgements
 
The author thanks Dr Yoshinori Ohsumi (NIBB, Japan) for assisting with the EM study, and Dr Benjamin S. Glick for the gift of P. pastoris plasmids. This study was supported by Grant-in-Aid for Scientific Research on Priority Areas 18076002 from the Japan Society for the Promotion of Science.


   Footnotes
 
Communicated by: Akihiko Nakano

* Correspondence: ysakai{at}kais.kyoto-u.ac.jp


   References
Top
 Abstract
 Introduction
 Results
 Discussion
 Experimental procedures
 References
 
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Received: 22 December 2008
Accepted: 24 April 2009





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