JBC Avanti Polar Lipids

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Originally published In Press as doi:10.1074/jbc.M404544200 on September 8, 2004

J. Biol. Chem., Vol. 279, Issue 47, 49138-49150, November 19, 2004
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
279/47/49138    most recent
M404544200v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow reprints & permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Hung, G.-C.
Right arrow Articles by Chiang, H.-L.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Hung, G.-C.
Right arrow Articles by Chiang, H.-L.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg   Add to Reddit   Add to Technorati  
What's this?

Degradation of the Gluconeogenic Enzymes Fructose-1,6-bisphosphatase and Malate Dehydrogenase Is Mediated by Distinct Proteolytic Pathways and Signaling Events*

Guo-Chiuan Hung{ddagger}, C. Randell Brown§, Allison B. Wolfe, Jingjing Liu, and Hui-Ling Chiang

From the Department of Cellular and Molecular Physiology, Penn State University College of Medicine, Hershey, Pennsylvania 17033

Received for publication, April 23, 2004 , and in revised form, August 25, 2004.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
The key gluconeogenic enzyme fructose-1,6-bisphosphatase (FBPase) is subjected to catabolite inactivation and degradation when glucose-starved cells are replenished with fresh glucose. In various studies, the proteasome and the vacuole have each been reported to be the major site of FBPase degradation. Because different growth conditions were used in these studies, we examined whether variations in growth conditions could alter the site of FBPase degradation. Here, we demonstrated that FBPase was degraded outside the vacuole (most likely in the proteasome), when glucose was added to cells that were grown in low glucose media for a short period of time. By contrast, cells that were grown in the same low glucose media for longer periods of time degraded FBPase in the vacuole in response to glucose. Another gluconeogenic enzyme malate dehydrogenase (MDH2) showed the same degradation characteristics as FBPase in that the short term starvation of cells led to a non-vacuolar degradation, whereas long term starvation resulted in the vacuolar degradation of this protein. The N-terminal proline is required for the degradation of FBPase and MDH2 for both the vacuolar and non-vacuolar proteolytic pathways. The cAMP signaling pathway and the phosphorylation of glucose were needed for the vacuolar-dependent degradation of FBPase and MDH2. By contrast, the cAMP-dependent signaling pathway was not involved in the non-vacuolar degradation of these proteins, although the phosphorylation of glucose was required.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Gluconeogenesis is a highly regulated cellular process by which yeast can produce glucose from alternative nutrient sources such as acetate, amino acids, lactate, and glycerol. When exogenous glucose is available, however, many gluconeogenic enzymes are transcriptionally repressed (1–4) and the existing enzymes are subjected to rapid inactivation and proteolytic degradation (5–9). Extensive studies have been conducted on the catabolite repression of gluconeogenic enzymes at the transcriptional level (10, 11). Through these studies, a general model of the associated signaling events has been established. Upon glucose depletion, Mig1p, a Cys2-His2 zinc finger DNA-binding protein is phosphorylated and localized in the cytoplasm. However, with the addition of exogenous glucose, Mig1p is dephosphorylated and imported into the nucleus, in a process that is facilitated by the nuclear transport receptor Msn5p (12, 13). Dephosphorylation is mediated by the inactivation of the Snf1-Snf4 protein kinase (14–16) and the activation of the Glc7-Reg1 protein phosphatase (17, 18). Once in the nucleus, Mig1p binds to the glucose-regulated promoters and recruits the general co-repressor complex Cyc8p-Tup1p to repress gene transcription (19, 20).

In contrast to the well characterized events that regulate signaling in transcriptional repression, our knowledge of the mechanisms that lead to the inactivation and degradation of gluconeogenic enzymes is not as well developed. FBPase1 has been the most extensively studied substrate for this process. After glucose-starved cells are replenished with fresh glucose, FBPase is phosphorylated, inactivated, and then degraded (5, 9, 21–23). To date, the site of FBPase degradation has been the subject of considerable controversy. For example, it has been reported that FBPase degradation is a proteasome-dependent process (24–29) where the proteolysis of FBPase depends on polyubiquitination and the function of the 26 S proteasome. The N-terminal proline residue in FBPase is essential for proteasome degradation, because mutation of the proline residue inhibits FBPase degradation (25). However, mutation of the Ser-11 phosphorylation site did not affect the degradation of FBPase in the proteasome, suggesting that phosphorylation is not required for this pathway of degradation (25). In contrast to these results, our laboratory and others have established that FBPase degradation depends upon the proper function of the vacuole (23, 30–41). In this vacuole-dependent degradation pathway, FBPase is first imported into intermediate transport vesicles known as Vid vesicles (39). This import requires the action of a number of proteins, including Ssa2p (32), cyclophilin A (33), and Vid22p (34). The loaded Vid vesicles then traffic to the vacuole and deliver FBPase for degradation, via a process that resembles heterotypic vesicle fusion. This event is dependent upon Vid24p, a small GTPase, Ypt7p, and v-SNARE proteins on Vid vesicles, whereas the t-SNARE Vam3p is required on the vacuole (36). The type I phosphatase Glc7p and its regulator Reg1p have also been shown to be required for the fusion of Vid vesicles with the vacuole (41).

In addition to FBPase, the cytosolic enzyme malate dehydrogenase (MDH2) is subjected to rapid degradation when glucose-starved cells are replenished with fresh glucose (42, 43). Mutagenesis studies have shown that the first 12 amino acids at the N terminus in MDH2 ({Delta}nMDH2) are essential for degradation. More recently, an interaction between MDH2 and FBPase has been demonstrated using the yeast two-hybrid assay and surface plasmon resonance analysis (44). Again, the N-terminal 12 amino acids are important for MDH2-FBPase interaction, because FBPase interacted with {Delta}nMDH2 with a much lower affinity. These results suggest that the first 12 amino acids of MDH2 mediate both interactions and the proper targeting of MDH2 for degradation.

In the current study, we attempted to resolve three questions: 1) Is FBPase degraded by both the proteasome and vacuole and if so, what conditions determine the site of FBPase degradation?; 2) Is FBPase degradation unique, or do other cytosolic enzymes share the same degradation characteristics?; and 3) If FBPase can be targeted and degraded by distinct proteolytic pathways, what are the signals that control this process? To address these questions, we investigated whether variations in growth conditions could contribute to differences in FBPase degradation. We showed that the degradation of FBPase progressively shifted from a non-vacuolar proteolytic pathway to a vacuolar pathway when cells were starved for longer periods of time in glucose-limited medium. We also showed that MDH2 exhibited the same degradation characteristics as FBPase in that it was degraded in the non-vacuole pathway when cells were starved for the short term, but it was degraded in the vacuole when cells were starved for longer periods of time. The N-terminal proline was required for the degradation of FBPase and MDH2, whether cells were subjected to short term or long term starvation conditions. However, the cAMP-dependent signaling pathway was exclusively involved in the vacuolar degradation of FBPase and MDH2. This signaling pathway is required for the phosphorylation of FBPase as well as the fusion of Vid vesicles with the vacuole. These results suggest that the cAMP signaling pathway plays multiple roles in the vacuole-dependent degradation of FBPase in response to glucose.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Yeast Strains, Culture Conditions, and Reagents—Yeast strains used in this study are listed in Table I. Media utilized in experiments included YPD (1% yeast extract (US Biological), 2% peptone (US Biological), and 2% glucose) and YPKG (1% yeast extract, 2% peptone, 1% potassium acetate, and 0.5% glucose). Yeast transformation was carried out using the lithium acetate/Tris EDTA protocol. For the induction of gluconeogenic enzymes, cells were grown at 30 °C in an environmental shaker (300 rpm) in YPKG for 1–3 days. Cell density was measured, and cells were shifted to YPD for 0–2 h to induce degradation of the gluconeogenic enzymes. For these experiments, aliquots of cells (1 A600) were used for each time point. Rabbit anti-FBPase antibodies were produced as described previously (30, 31). The {Delta}pep4{Delta}prb1{Delta}prc1 strain (45) was obtained from Dr. A. Kornberg (Stanford University). Anti-MDH2 antibodies were obtained from Dr. McAlister-Henn (University of Texas, Houston, TX) (43). Horseradish peroxidase-conjugated anti-rabbit antibodies and horseradish peroxidase-conjugated anti-mouse antibodies were purchased from Amersham Biosciences. Western blots were visualized using the enhanced chemiluminescence (ECL) kit (PerkinElmer Life Sciences).


View this table:
[in this window]
[in a new window]
  		TABLE I
Yeast strains used in this study

 
Construction of FBPase-GFP and MDH2-GFP—We utilized a PCR-based method developed by Longtine et al. (46). In brief, a set of bacterial plasmids bearing an enhanced GFP with selectable nutrient marker (HIS5+ or TRP1) was used. PCR primers were designed according to the published protocol with minor modifications (46). To maximize yeast transformation efficiency, the sequences corresponding to the genomic DNA in PCR primers were increased to 50 or 60 nucleotides in length. The location of reverse primers was also shifted further downstream to increase transformation efficiency. A BLAST search was performed on the selected reverse primer sequences to ensure their specificity. The PCR primer sequences used in this study are listed in Table II. The PCR mix contained 100 pmol of each primer, 2 mM MgCl2, 250 µM dNTPs, and 5 units of Taq polymerase (Qiagen). The thermocycling conditions were as follows: after the initial denaturation at 94 °C for 5 min, 30–35 cycles of 30 s at 94 °C, 30 s at 55 °C and 2 min 30 s at 72 °C. The PCR products were purified and concentrated using Promega PCR Cleanup kit (Madison, WI). Genomic integration was confirmed by PCR. To examine the localization of FBPase-GFP and MDH2-GFP, cells were grown in YPKG for 1 or 3 days. Cells were then shifted to YPD media for 0–2 h and examined by fluorescence microscopy using a Zeiss fluorescence microscope and Axiovision software.


View this table:
[in this window]
[in a new window]
  		TABLE II
PCR primers used in this study

 
Plasmids and Mutagenesis—The FBPase open reading frame was cloned into a pYES vector. The PCR product was fused with a V5 epitope and polyhistidine (His6) (pYES2.1, Invitrogen) with a 2µ origin and URA3 for auxotrophic selection (pB189). To produce the FBPase-V5–6xHis construct containing LEU2 as a nutritional marker, a 2.2-kb fragment was first obtained from YEP351 by restriction enzyme digestion using PstI and SnaBI. This fragment was then ligated into the PstI and HpaI sites replacing URA3 and the 2µ origin of pB189 (pB284). To obtain a 3-kb FBPase fragment containing the 1-kb 5' region, the open reading frame, and the 1-kb 3' region (pB263), PCR primers were designed (Table II) and the PCR products were cloned into the PCR2.1 Topo vector (Invitrogen). pB287 containing the 5'-promoter region of FBPase fused with His6, was constructed by ligating a HpaI/BstBI fragment into HpaI/ClaI site of pB189. Mutagenesis of FBPase to change the first proline to serine (pB288) and the 11th serine to alanine (pB289) was carried out using the QuikChange XL site-directed mutagenesis kit (Stratagene, La Jolla, CA). This was based on pB287 using the PCR primers listed in Table II. PCR primers were synthesized and purified by Integrated DNA Technologies (Coralville, IA), and FBPase constructs were confirmed by sequencing. Prior to transformation into {Delta}fbp1 cells, constructs were linearized by HindIII digestion.

MDH2 truncations and site-directed mutagenesis were also carried out using the QuikChange XL site-directed mutagenesis kit and the PCR primers listed in Table II. Plasmids bearing the wild type (pBS-MDH2) and the {Delta}nMDH2 genes were obtained from Dr. McAlister-Henn (47). Site-directed mutagenesis was performed using the wild type MDH2 gene as the template. Mutations were confirmed by DNA sequencing (Core facility, Penn State College of Medicine). Various mutated forms of MDH2 were introduced to a {Delta}mdh2::HIS3 cell by integration of the linearized DNA digested by SnaBI (47).

Differential Centrifugation, in Vitro Vid Vesicle/Vacuole Fusion, and FBPase Phosphorylation—Differential centrifugation and in vitro Vid vesicle/vacuole fusion were performed as described previously (36). FBPase phosphorylation was conducted as described before (23).


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
FBPase Is Degraded by Two Distinct Proteolytic Pathways— We have shown previously that FBPase is degraded in the vacuole when glucose-starved cells are shifted to glucose containing media (30–41). However, a proteasome-dependent degradation of this same enzyme has also been demonstrated by the Wolf group (24–29). The reason for this discrepancy is unclear, although different experimental conditions are used in the two laboratories. To address this issue, we investigated whether variations in growth conditions could result in different sites of FBPase degradation. In our previous studies, we used {Delta}pep4 cells to demonstrate that FBPase degradation occurs in the vacuole. The PEP4 gene encodes for proteinase A, a major proteinase of the vacuole (48, 49). However, other proteinases are still present in the vacuole of this mutant, leading to an incomplete block of FBPase degradation. Therefore, this partial inhibition of degradation in the {Delta}pep4 strain may have contributed to the controversy in this field. To overcome this problem, we used a mutant strain harboring triple deletions of the PEP4, PRB1, and PRC1 genes that encode proteinases A, B, and C, respectively. A defect in FBPase degradation in this mutant would suggest that degradation occurs in the vacuole, whereas normal FBPase degradation in this mutant would indicate an extravacuolar site of degradation.

In our initial experiments, we confirmed that FBPase degradation is dependent on vacuole proteinases when cells were grown under our previously established growth condition (YPKG for 48 h, Fig. 1A). Next, however, we examined whether variations in growth conditions may result in different degradation patterns. Wild type and {Delta}pep4{Delta}prb1{Delta}prc1 cells were grown in glucose-limited media for various periods of time, shifted to fresh glucose for 0–2 h, and examined for FBPase degradation. As expected, the control wild type cells degraded FBPase when cultured under all growth conditions (Fig. 1B). However, a progressive reduction of FBPase degradation was observed for the {Delta}pep4{Delta}prb1{Delta}prc1 mutant, depending on how long this strain had been cultured in glucose limited media (Fig. 1A). FBPase was rapidly degraded when this mutant strain was grown under low glucose conditions (YPKG) for 24 h. Following 36 h of starvation, however, FBPase degradation was somewhat delayed. The degradation of FBPase was further retarded following starvation for 48 h, whereas FBPase degradation was almost completely blocked when this strain was starved for 72 h. Therefore, the rate of FBPase degradation in this {Delta}pep4{Delta}prb1{Delta}prc1 mutant is dependent on the duration of glucose starvation. These results suggest that FBPase is degraded predominantly by a non-vacuolar pathway following 24 h of starvation. By contrast, a retarded degradation of FBPase in the {Delta}pep4{Delta}prb1{Delta}prc1 mutant following a 72-h starvation period suggests that FBPase is degraded primarily in the vacuole under this circumstance. Owing to these dramatic differences in FBPase degradation in 1-day- versus 3-day-starved cells, all subsequent experiments were conducted using cells that were starved for these two time periods.



View larger version (18K):
[in this window]
[in a new window]
  		FIG. 1.
FBPase degradation shifts from a non-vacuolar to a vacuolar pathway depending on the duration of glucose starvation. A and B, wild type and {Delta}pep4{Delta}prb1{Delta}prc1 mutants were grown in YPKG for 24, 36, 48, and 72 h and then shifted to glucose for 0 and 2 h. Cell lysates were obtained and examined for FBPase levels in the {Delta}pep4{Delta}prb1{Delta}prc1 strain (A) and wild type cell (B).

 
To further confirm that FBPase was degraded in the vacuole when 3-day-starved cells were shifted to glucose, we performed localization studies using GFP-tagged FBPase. The {Delta}pep4-{Delta}prb1{Delta}prc1 mutant exhibited high background fluorescence under both short term and long term starvation conditions and as such was not suitable for these studies. In contrast, the {Delta}pep4 cell displayed low background fluorescence when transformed with an FBPase-GFP construct. When FBPase-GFP distribution was examined in the {Delta}pep4 cell, a low but detectable fluorescence signal was observed in the cytosol at t = 0 in cells starved for 1 or 3 days (Fig. 2A). However, the FBPase-GFP signal disappeared rapidly following a shift of 1-day-starved cells to glucose, suggesting that FBPase-GFP was degraded in this cell. By contrast, a detectable signal of FBPase-GFP was present in the vacuole of {Delta}pep4 cells that were starved for 3 days and then shifted to glucose (Fig, 2A). This study further confirmed our previous indirect immunofluorescence analysis (30), whereby FBPase was localized to the vacuole in {Delta}pep4 cells in response to glucose.



View larger version (41K):
[in this window]
[in a new window]
  		FIG. 2.
FBPase is targeted to the vacuole when 3-day-starved {Delta}pep4 cells are shifted to glucose. A, {Delta}pep4 cells transformed with FBPase-GFP were grown in YPKG for 1 and 3 days and then shifted to glucose for 0 and 2 h. Cells were examined using Nomarski microscopy, and the GFP was visualized under fluorescence microscopy. B, {Delta}pdr5 cells were grown in YPKG for 1 and 3 days and then shifted to glucose in the absence or presence of MG132 for 0 and 2 h. FBPase degradation was examined. The 3-day-starved cells expressed FBPase as a truncated form. The levels of the 25-kDa forms of FBPase at t = 0 and t = 2 h are shown.

 
The proteasome is a major cytosolic site of protein degradation (50, 51). Furthermore, it has been reported that FBPase is degraded in the proteasome (24–29). Therefore, we next determined whether the proteasome was responsible for FBPase degradation in 1-day-starved cells. For these experiments, we used a {Delta}pdr5 cell that is sensitive to the effects of the proteasome inhibitor MG132 (52). This strain was chosen because this inhibitor is ineffective in most wild type strains. When 1-day-starved {Delta}pdr5 cells were shifted to glucose in the absence of MG132, FBPase was rapidly degraded (Fig. 2B). However, the addition of MG132 to these same cells retarded FBPase degradation (Fig. 2B), suggesting that the proteasome plays some role in FBPase degradation, at least under 1-day starvation condition. Unfortunately, results for FBPase degradation in cells starved for 3 days were difficult to interpret. Under this condition, FBPase was expressed as a 25-kDa truncated form. Interestingly, when we examined a number of proteasome mutant strains, the majority of the FBPase was also present as a truncated form. Although the significance of this is not clear at the present time (see "Discussion"), truncated FBPase was stable following a glucose shift in these cells (Fig. 2B).

For the vacuolar-dependent pathway, FBPase is associated with intermediate transport vesicles (Vid vesicles) before being targeted to the vacuole (39). It has not been established, however, whether these vesicles play a role in proteasome-mediated degradation. Furthermore, if FBPase is targeted for degradation in two proteolytic pathways, FBPase distribution within cells may be different under different growth conditions. Wild type cells were starved for 1 or 3 days and then shifted to glucose for 30 min. Organelles were fractionated by differential centrifugation followed by separation on a sucrose density gradient. Using this protocol, we showed previously that soluble proteins were in lighter density fractions, whereas Vid vesicles were in heavier density fractions (36, 39). In 1-day-starved cells, most of the FBPase was found in light density fractions (Fig. 3A). By contrast, in 3-day-starved cells, FBPase exhibited a broader distribution with significant amounts of FBPase extending to heavier density fractions (Fig. 3A). The presence of multiple FBPase peaks may represent various stages of Vid vesicle maturation in wild type cells, and this may correlate with different functions of Vid vesicles. Regardless, our fractionation results indicate that FBPase distribution in response to glucose stimulation can change depending on starvation conditions.



View larger version (25K):
[in this window]
[in a new window]
  		FIG. 3.
Vam3p and Ypt7p are required for FBPase degradation in 3-day-starved cells, but not 1-day-starved cells. A, 1- and 3-day-starved wild type cells were shifted to glucose for 30 min and lysed. Total extracts were subjected to differential centrifugation and sucrose density gradient as described (36). Each fraction from the sucrose gradient separation was collected, processed, and solubilized in SDS sample buffer. FBPase distribution was examined by Western blotting with anti-FBPase antibodies. B, {Delta}vam3 and {Delta}ypt7 mutants were grown in YPKG for 1 and 3 days. These cells were then shifted to glucose for 0 and 2 h and examined for FBPase degradation.

 
To further establish that FBPase was degraded by two distinct proteolytic pathways, we used the {Delta}ypt7 and {Delta}vam3 mutants. Both Ypt7p and Vam3p have been shown to play important roles in many known trafficking pathways leading to the vacuole (53–60). These include the sorting of carboxy peptidase Y to the vacuole (53), the targeting of autophagosomes to the vacuole, the cytosol to vacuole targeting pathway (54–56), and homotypic vacuole fusion (57–60). In addition, for the vacuole-dependent FBPase degradation pathway, the last step requires the participation of Ypt7p and Vam3p (36). Because Ypt7p and Vam3p are specific for the vacuolar trafficking pathways, we used these mutants to distinguish between the vacuolar versus non-vacuolar degradation of FBPase. The {Delta}ypt7 and {Delta}vam3 mutants were glucose-starved for 1 or 3 days, shifted to fresh glucose, and examined for FBPase degradation (Fig. 3B). FBPase was degraded in both mutants following a 1-day starvation. However, FBPase degradation was impaired when these same mutants were grown under glucose starvation conditions for 3 days (Fig. 3B). These results support the conclusion that short term starvation leads to a non-vacuolar degradation of FBPase, whereas long term starvation leads to FBPase degradation in the vacuole.

MDH2 and FBPase Show Similar Degradation Characteristics—Glucose-induced degradation of MDH2 has been described previously (42, 43). However, whether this protein is degraded in the vacuole or proteasome has not been addressed. Therefore, we next determined whether MDH2 showed similar degradation characteristics as FBPase. Wild type cells and triple vacuolar mutants were glucose-starved for 1 and 3 days, shifted to glucose for various periods of time, and examined for the degradation of MDH2 (Fig. 4, A and B). Glucose addition induced a rapid degradation of MDH2 when the {Delta}pep4{Delta}prb1{Delta}prc1 cells were starved for 1 day (Fig. 4A). However, MDH2 degradation was impaired when this mutant was starved for 3 days (Fig. 4A). As expected, the control wild type cells degraded MDH2 whether cells were starved for 1 or 3 days (Fig. 4B).



View larger version (45K):
[in this window]
[in a new window]
  		FIG. 4.
The length of glucose starvation determines the pattern of MDH degradation in the {Delta}pep4{Delta}prb1{Delta}prc1 mutant. The {Delta}pep4{Delta}prb1{Delta}prc1 (A) and wild type (B) strains were grown in YPKG for 24, 36, 48, and 72 h. Cells were then transferred to medium containing fresh glucose for 2 h. Cell lysates were obtained and blotted with anti-MDH2 antibodies. C, {Delta}pep4 cells expressing MDH2-GFP were grown in YPKG for 1 and 3 days. They were then shifted to glucose for 2 h, and MDH2-GFP was visualized using fluorescence microscopy.

 
To further confirm that MDH2 was targeted to the vacuole in 3-day-starved cells, we examined the distribution of MDH2-GFP in the {Delta}pep4 cell. MDH2-GFP was detected in the cytosol at t = 0 in the {Delta}pep4 cell starved for either 1 or 3 days (Fig. 4C). However, the MDH2-GFP signal disappeared when 1-day-starved cells were shifted to glucose for 2 h, suggesting that MDH2-GFP was degraded. In contrast, most of the fluorescence signal was found in the vacuole when 3-day-starved {Delta}pep4 cells were shifted to glucose for 2 h (Fig. 4C).

We next determined whether MDH2 exhibited similar localization characteristics as FBPase. To address this question, 1- and 3-day-starved cells were glucose-shifted and fractionated as described above and the fractions were blotted with MDH2 antibodies. MDH2 exhibited a similar distribution as FBPase (Fig. 5A). In 1-day-starved cells, the majority of MDH2 was distributed in lighter fractions. In 3-day-starved cells, however, high levels of this protein were found in lighter fractions, whereas smaller amounts of MDH2 were also detected in heavier fractions (Fig. 5A). Therefore, FBPase and MDH2 appeared to co-localize, whether they were targeted to the vacuolar or to the non-vacuolar degradation pathways.



View larger version (25K):
[in this window]
[in a new window]
  		FIG. 5.
Ypt7p and Vam3p are required for MDH2 degradation when long term starved cells are shifted to glucose. A, wild type cells were starved for 1 and 3 days in YPKG and refed with fresh glucose for 30 min. Cell lysates were subjected to differential centrifugation and sucrose density gradients as described (36). The distribution of MDH2 was examined by Western blotting of the fractions from the sucrose gradient. B, {Delta}vam3 and {Delta}ypt7 cells were grown in YPKG for 1 and 3 days, shifted to glucose, and examined for MDH2 degradation.

 
If long term starved cells utilize Vid vesicles to deliver MDH2 to the vacuole for degradation, then the {Delta}ypt7 and {Delta}vam3 mutants should block the degradation of MDH2 in response to glucose. Fig. 5B shows that MDH2 degradation was normal for 1-day-starved {Delta}ypt7 cells and {Delta}vam3 cells. By contrast, the degradation of MDH2 was retarded in {Delta}ypt7 and {Delta}vam3 cells that were starved for 3 days (Fig. 5B). Therefore, MDH2 shows the same degradation characteristics as FBPase, in that a shift of short term starved cells to glucose leads to a non-vacuolar degradation of MDH2. By contrast, a shift of long term starved cells to glucose leads to a predominantly vacuolar-dependent degradation of MDH2.

Vacuolar and Proteasome Degradation Pathways Share Common Components—A number of VID and GID genes have been characterized. VID genes were originally isolated during a search for genes involved in the vacuolar degradation of FBPase (40), whereas GID genes were identified in a search for genes involved in the degradation of FBPase via the proteasome (25). Interestingly, a recent genome-wide search has also identified a number of VID genes, including VID24, VID28, and VID30, that are also involved in the proteasomal-dependent degradation of FBPase (28). This suggests that common components exist for both proteolytic pathways. To confirm the dual roles of these VID and GID genes, we examined FBPase and MDH2 degradation in vid and gid mutants that were starved for 1 and 3 days. As is shown in Fig. 6, {Delta}gid2 and {Delta}gid8 mutants blocked FBPase degradation in 1-day-starved cells, a result that is consistent with earlier reports (28). However, FBPase degradation was also blocked in these strains when they were starved for 3 days. Likewise, the {Delta}vid24, {Delta}vid28, and {Delta}vid30 mutants blocked FBPase degradation under both starvation conditions (Fig. 6A). In a similar manner, these mutants were defective in MDH2 degradation, whether they were starved for 1 or 3 days (Fig. 6B). Thus, the proteasome and vacuole degradation pathways share common components for the degradation of two cargo proteins, FBPase and MDH2.



View larger version (26K):
[in this window]
[in a new window]
  		FIG. 6.
Common components function in the vacuolar and proteasomal proteolytic pathways. Wild type, {Delta}gid2, {Delta}gid8, {Delta}vid24, {Delta}vid28, and {Delta}vid30 mutants were starved for 1 and 3 days and then shifted to fresh glucose. The degradation profiles of FBPase (A) and MDH2 (B) were examined. For MDH2 blots, note the presence of a minor, nonspecific band. This band was also present in blots from {Delta}mdh2 cell lysates.

 
The N-terminal Proline Is Required for MDH2 and FBPase Degradation—The sequence required for the glucose-induced degradation of MDH2 has been mapped to the N-terminal 12 amino acids (42). Because this protein can be degraded in the vacuole or proteasome, we examined whether this sequence plays a role in one or both of these pathways. Strains expressing MDH2 lacking the first 12 amino acids ({Delta}n) were examined for MDH2 degradation in 1- and 3-day-starved cells. To our surprise, the {Delta}nMDH2 strain was defective for degradation in both 1- and 3-day-starved cells (Fig. 7, A and B). To further define the sequence required for MDH2 degradation, we created small truncations within the N-terminal 12 amino acids. The deletion of residues 9–12 (EQDS) did not inhibit the degradation of MDH2 in either 1- or 3-day-starved cells. Similarly, the deletion of residues 5–8 (TPSI) did not block MDH2 degradation in either 1- or 3-day-starved cells. However, the deletion of the N-terminal 1–4 residues (PHSV) inhibited the degradation of MDH2 in both 1- and 3-day-starved cells. These results suggest that the N-terminal 4 amino acid residues of MDH2 contain the critical sequence for both vacuolar and non-vacuolar degradation pathways.



View larger version (15K):
[in this window]
[in a new window]
  		FIG. 7.
The N-terminal domain is necessary for the degradation of MDH2. A, {Delta}mdh2 cells were transformed with plasmids containing either wild type or various MDH2 truncations. Cells expressing wild type, {Delta}1–4, {Delta}5–8, {Delta}9–11, and {Delta}1–12 ({Delta}n) MDH2 were grown in YPKG for 1 (A) and 3 days (B). Cells were shifted to glucose for 0 and 2 h, and MDH2 degradation was examined.

 
Sequence comparison of FBPase and MDH2 indicates that both proteins contain a proline at their N terminus. This proline residue was reported to be important for the degradation of FBPase in the proteasome, because a proline to serine substitution blocked its degradation (25). To further define the sequences required for MDH2 degradation, we produced the same proline to serine substitution on MDH2 and examined the degradation of the P1S mutation (Fig. 8A). When cells expressing P1S were starved for 1 day and then shifted to glucose, this mutant blocked MDH2 degradation. Interestingly, P1S also inhibited MDH2 degradation when cells were starved for 3 days (Fig. 8A).



View larger version (23K):
[in this window]
[in a new window]
  		FIG. 8.
The N-terminal proline is required for the degradation of MDH2 and FBPase. A, site-directed mutagenesis was performed to produce the N-terminal proline to serine (P1S) substitution and a serine to alanine mutation at position 3 (S3A) in MDH2. The P1S and S3A mutants were confirmed by DNA sequencing and then transformed into {Delta}mdh2 cells. Cells were grown in YPKG for 1 and 3 days. Cells were shifted to glucose and examined for MDH2 degradation. B, site-directed mutagenesis was used to produce the P1S and S11A mutations in FBPase as described under "Experimental Procedures." These mutations were confirmed by DNA sequencing and transformed into the {Delta}fbp1 cells. Cells were grown in YPKG for 1 and 3 days, and glucose was added for 2 h. FBPase degradation was examined.

 
MDH2 is phosphorylated following a glucose shift (43). Phosphorylation was not observed in {Delta}nMDH2 (43), suggesting that the first 12 amino acids are required for the phosphorylation. However, the exact phosphorylation sites have not been mapped. Because a serine residue at position 3 was within the critical 1–4 sequence, we also mutated this residue to determine whether MDH2 degradation was affected. The S3A mutation did not block MDH2 degradation in either 1- or 3-day-starved cells (Fig. 8A). Therefore, the sequence required for both the vacuolar and non-vacuolar degradation of MDH2 has been mapped to the proline residue at the N terminus of the protein.

The above results prompted us to re-examine the role that the N-terminal proline plays in FBPase degradation. As mentioned above, this residue is required for FBPase degradation via the proteasome pathway (25). However, whether the same proline residue is required for the vacuole degradation pathway has not been determined. We produced the P1S mutation and examined the degradation of FBPase in short term- and long term starved cells (Fig. 8B). The P1S mutation inhibited FBPase degradation in 1-day-starved cells. This P1S mutation also blocked FBPase degradation in 3-day-starved cells. Therefore, the same proline reside is required for both non-vacuolar and vacuolar proteolysis pathways.

FBPase is phosphorylated by the cAMP-dependent pathway when cells are shifted to glucose, and this phosphorylation has been mapped to the serine 11 residue (21, 22). Therefore, we determined whether phosphorylation of FBPase at serine 11 was required for its degradation. We produced an FBPase mutation that substitutes serine for alanine at position 11. This S11A mutation did not block FBPase degradation in 1-day-starved cells, suggesting that phosphorylation at this site is not required for the FBPase degradation in short term starved cells (Fig. 8B). This is consistent with the report that the S11A mutation had no effect on FBPase degradation in the proteasome (27). Likewise, the S11A mutation exhibited normal FBPase degradation in 3-day-starved cells (Fig. 8B), suggesting that phosphorylation at this site is not required for FBPase degradation in the vacuole.

Glucose Phosphorylation Is Required for FBPase Degradation—Because FBPase and MDH2 each can be degraded by two distinct proteolytic pathways, different signaling events may be required to mediate their degradation in the proteasome or the vacuole. For the proteasome-dependent pathway, glucose phosphorylation by hexokinases is required (27). The cAMP-dependent pathway also appears to be required for FBPase degradation, although this has been a matter of debate. For example, it was reported that the cAMP pathway is not involved in FBPase degradation in the proteasome (27). In contrast, a Ras2-dependent degradation of FBPase has been described (23). To clarify these issues, we next examined the role that these signaling pathways played in FBPase degradation under different growth conditions.

When glucose is added to cells, this compound is converted to glucose 6-phosphate via the actions of hexokinase 1, hexokinase 2, and glucokinase 1 (61–63). To determine the role that glucose phosphorylation plays in FBPase degradation, we examined various deletion mutants lacking these genes. The deletion of individual genes did not affect the degradation of FBPase or MDH2 (Fig. 9, A and B), suggesting that these genes have redundant functions. Consistent with this notion, a triple mutant harboring deletions of HXK1, HXK2, and GLK1 inhibited FBPase degradation in 1- and 3-day-starved cells (Fig. 9A). The same mutants also blocked MDH2 degradation in 1- and 3-day-starved cells (Fig. 9B). Therefore, glucose phosphorylation by these kinases is required for FBPase and MDH2 degradation in the proteasome and the vacuole.



View larger version (21K):
[in this window]
[in a new window]
  		FIG. 9.
Phosphorylation of glucose is required for the degradation of FBPase and MDH2. Mutants harboring single or triple deletions of the HXK1, HXK2, and GLK1 genes were grown in YPKG in 1 and 3 days. Cells were shifted to glucose and examined for the degradation of FBPase (A) and MDH2 (B).

 
The cAMP-dependent Signaling Pathway Is Involved in Vacuolar Degradation of FBPase and MDH2—The addition of glucose to cells that have been grown in media containing non-fermentable carbon sources causes a rapid increase in cAMP (64–66). Members of the cAMP-mediated pathway include the G protein-coupled receptor Gpr1p and the heterotrimeric G protein Gpa2p. These proteins stimulate cAMP synthesis by activating adenylate cyclases. The intracellular G proteins Ras1p and Ras2p also regulate the activity of adenylate cyclases (64–66). Increased cAMP then stimulates a cAMP-dependent kinase to phosphorylate target proteins. When FBPase and MDH2 degradation was examined in {Delta}gpr1, {Delta}gpa2, and {Delta}ras2 strains, they were degraded normally in 1-day-starved cells (Fig. 10, A and B). In contrast, when these cells were starved for 3 days, the degradation of FBPase and MDH2 was retarded (Fig. 10, A and B). Therefore, the cAMP-dependent signaling pathway appears to play some role in the degradation of FBPase in the vacuole, whereas it is not required for degradation that occurs outside the vacuole.



View larger version (18K):
[in this window]
[in a new window]
  		FIG. 10.
The c-AMP-dependent signaling pathway is exclusively involved in the vacuole-dependent degradation of FBPase and MDH2. Strains with deletions of GPA2, GPR1, and RAS2 were glucose-starved for 1 and 3 days. They were then shifted to glucose for 2 h and examined for the degradation of FBPase (A) and MDH2 (B).

 
The cAMP Pathway Mediates FBPase Phosphorylation and Vid Vesicle-Vacuole Fusion—To more clearly define the role that members of the cAMP pathway play in the process of FBPase degradation, we attempted to determine the site of action of these proteins. Mutation of the S11 FBPase residue does not affect FBPase degradation (Fig. 8B), indicating that phosphorylation at this site is not required for the glucose-induced degradation of this protein. However, whether other residues can be phosphorylated by protein kinase A has not been determined. To address this question, we determined whether the S11A FBPase mutant protein was phosphorylated in response to glucose. Cells expressing wild type and S11A FBPase were glucose-shifted, and FBPase phosphorylation was examined (Fig. 11A). For these experiments, similar amounts of FBPase were precipitated as detected by Western blotting with anti-FBPase antibodies (not shown). In wild type cells, FBPase phosphorylation was low at t = 0, and increased at t = 30 min. This phosphorylation was severely reduced in the protein kinase A mutant that harbors mutations in the TPK1, TPK2, TPK3, and BCY1 genes, suggesting that phosphorylation is dependent on protein kinase A. Interesting, the S11A-FBPase was phosphorylated in response to glucose at t = 30 min. Therefore, serine 11 is apparently not the only phosphorylation site in the FBPase protein.



View larger version (19K):
[in this window]
[in a new window]
  		FIG. 11.
The cAMP pathway is required for FBPase phosphorylation and Vid vesicle/vacuole fusion. A, FBPase phosphorylation was determined in cells expressing wild type FBPase, S11A-FBPase, and in the tpkw1{Delta}tpk2{Delta}tpk3{Delta}bcy1 mutant. Results were quantitated using a PhosphorImager and densitometry. B, the {Delta}vid24 and {Delta}ras2 mutants were grown in glucose-limited media and then shifted to glucose for 30 min. Cell lysates were subjected to differential centrifugation whereby FBPase distribution in the high speed pellet (P) and high speed supernatant (S) was examined. C, Vid vesicles, vacuoles, and cytosol were isolated from wild type and the {Delta}ras2 mutants as described (36). These components were combined as indicated and tested for the activation of alkaline phosphatase.

 
Next, we examined the distribution of FBPase in mutants that were shifted to glucose for 30 min. If the cAMP pathway is involved in FBPase import into Vid vesicles, then FBPase should accumulate in the cytosol in these mutants. However, if this signaling pathway is involved in Vid vesicle/vacuole fusion, FBPase should accumulate in Vid vesicle fractions. When FBPase distribution was examined, a significant portion of FBPase was detected in the Vid vesicle pellet fraction in the {Delta}ras2 mutant as well as in the cytosolic fraction (Fig. 11B). The distribution was similar to that seen in the {Delta}vid24 mutant that blocks fusion of Vid vesicles with the vacuole (36). Similar FBPase distribution patterns were obtained with the {Delta}gpr1 and {Delta}gpa2 mutants (not shown). Because FBPase accumulated in Vid vesicle fractions, these results suggest that the cAMP signaling pathway is involved in the downstream fusion of Vid vesicles with the vacuole.

To verify a role for cAMP signaling in Vid vesicle/vacuole fusion, we utilized an in vitro fusion assay previously developed in our laboratory (36). For these experiments, Vid vesicles, vacuoles, and cytosol were isolated from wild type and {Delta}ras2 mutant cells that had been shifted to glucose for 30 min (36). Mutant and wild type components were combined as indicated in Fig. 11C, and fusion was measured in terms of the activation of alkaline phosphatase. When cytosol, vesicles, or vacuoles were isolated from the {Delta}ras2 mutant, alkaline phosphatase activity was low (Fig. 11C), indicating that one or all of these components were defective. To identify the site of this defect, various combinations of mutant and wild type components were tested. Alkaline phosphatase activity was low when mutant vesicles were mixed with wild type vacuoles and cytosol. Likewise, when {Delta}ras2 vacuoles were combined with wild type vesicles and cytosol, alkaline phosphatase activity was also low. In contrast, when the cytosol from {Delta}ras2 mutant was used, the fusion activity was high. Thus, the {Delta}ras2 mutant contained defective Vid vesicles and defective vacuoles, but normal cytosol. Taken together, these results suggest that the cAMP signaling pathway is required for multiple events in the FBPase degradation process. These include the phosphorylation of FBPase, as well as the fusion of Vid vesicles with the vacuole. Furthermore, both the vesicles and the vacuoles appear to be the site of action for this signaling pathway.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
The site of FBPase degradation in response to glucose has been a matter of debate. FBPase has been reported to be ubiquitinated and degraded in the proteasome (24–29). However, FBPase also appears to be degraded in the vacuole via a Vid vesicle-dependent process (30–41). In addition to these different conclusions regarding the site of degradation, conflicting results were reported concerning whether the cAMP signaling pathway is involved in FBPase degradation. One report has indicated that this signaling pathway is not required for FBPase degradation (27). However, a Ras2-dependent increase in intracellular cAMP levels was also shown to be important for FBPase degradation (23). Because different growth conditions were used to induce FBPase expression in these laboratories, one explanation for this discrepancy is that growth conditions of cells determine the site of FBPase degradation. In this current study, we demonstrated that variations in growth conditions indeed led to different mechanisms of FBPase degradation. Here, we provided evidence showing that at least one of the factors that determines the fate of FBPase is the duration of the starvation condition, even in cells that were grown in the same glucose limited culture media. The addition of glucose to cells that underwent short term starvation (1d) resulted in a non-vacuolar degradation of FBPase. By contrast, the addition of glucose to long term (3-day)-starved cells led to the degradation of FBPase in the vacuole.

For 1-day-starved cells, FBPase is most likely degraded in the proteasome. Consistent with this observation, the addition of proteasome inhibitors to otherwise wild type cells inhibited FBPase degradation in 1-day-starved cells. However, caution should be taken when using proteasome inhibitors or proteasome mutants, because they may cause indirect effects. In our studies, long term starvation in several proteasome mutants led to the expression of FBPase as a 25-kDa truncated form. Interestingly, this truncated form of FBPase was not degraded in these proteasome mutants following glucose refeeding (not shown). In a similar manner, the truncated form of FBPase was not degraded when 3-day-starved {Delta}pdr5 cells were transferred to glucose in the presence of the proteasome inhibitor. Although the reason for this inhibition is not clear at the present time, one explanation is that the truncated FBPase lacks critical domains necessary for degradation to occur. As a result, this protein remains stable. Another possibility is that the use of proteasome mutants or proteasome inhibitors changed the ubiquitin pool within cells. A reduction in free ubiquitin levels can potentially impair the functions of conjugating enzymes to ubiquitinate substrate proteins. In our previous studies, we showed that at least one of the ubiquitin-conjugating enzymes, Ubc1p, is required for Vid vesicle formation and function in the vacuole pathway (38). Therefore, when the free ubiquitin pool is reduced, Ubc1p activity may be affected. This can lead to an FBPase import defect and a subsequent degradation defect.

Owing to the limitations of experiments with proteasome mutants or inhibitors, we believed that more compelling evidence can be obtained through the examination of FBPase degradation in {Delta}pep4{Delta}prb1{Delta}prc1 mutants deficient in major vacuole proteinases. When these mutant strains were grown under long term starvation conditions, FBPase degradation in response to glucose was severely retarded, suggesting that FBPase degradation occurs in the vacuole. Furthermore, FBPase-GFP was found in the vacuole when 3-day-starved {Delta}pep4 cells were shifted to glucose, indicating that FBPase was targeted to the vacuole under this condition. Additional support for the vacuole-dependent degradation of FBPase came from studies with the {Delta}ypt7 and {Delta}vam3 mutants. These strains also blocked FBPase degradation in 3-day-starved cells, but not in 1-day-starved cells. Based upon these observations, we conclude that long term starvation leads to the degradation of FBPase in the vacuole.

In this study, we identified MDH2 as another substrate for both degradation pathways. Both FBPase and MDH2 were degraded in the vacuole in long term starved cells. Likewise, they were degraded in the non-vacuolar pathway in short term starved cells. It is intriguing that both proteins required the N-terminal proline residue for the vacuolar and the non-vacuolar degradation pathways. Accordingly, the P1S mutation on both proteins inhibited their degradation in short term as well as long term starved cells. For the proteasomal degradation pathway, the N-terminal proline is necessary for the ubiquitination of FBPase (25). However, the vacuolar degradation pathway is unlikely to utilize this system, because FBPase ubiquitination was not observed under the vacuolar degradation conditions (38). FBPase is known to interact with MDH2 under glucose starvation conditions (44). Therefore, we determined whether the degradation defect seen in the P1S-FBPase and P1S-MDH2 mutants resulted from a reduction in their interactions. We compared the interactions between the P1S-FBPase with the wild type MDH2 to the wild type counterparts. Unfortunately, we did not observe any reduction in the FBPase-MDH2 interaction when the P1S-FBPase mutant was used. In a similar manner, the interaction between P1S-MDH2 and wild type FBPase was not reduced. Furthermore, the interactions remained the same, before or after glucose shift (not shown). Therefore, the N-terminal proline is not required for the FBPase-MDH2 interaction. We suspect that the N-terminal proline serves as a determinant by binding to an as yet unidentified factor or factors. If this scenario is true, it will be important to identify these factors and to determine how they function in the vacuolar pathway.

Finally, we showed that signaling pathways play important roles in determining whether cells degrade FBPase and MDH2 in the vacuole or in the proteasome. Phosphorylation of glucose by hexokinases and glucokinase is required for FBPase and MDH2 degradation, not only for the vacuole pathway, but also for the proteasome pathway. In contrast, the cAMP-dependent signaling pathway is exclusively involved in vacuole-dependent degradation. Deletions of components of the cAMP signaling pathway, including GPA2, GPR1, and RAS2, all inhibited FBPase and MDH2 degradation in long term starved cells. By contrast, the deletion of these genes had no effects on FBPase and MDH2 degradation in short term starved cells. Although FBPase is known to be phosphorylated on serine 11 residue (21, 22), the S11A mutation in FBPase did not affect degradation in short term or long term starved cells. Furthermore, when this mutant was examined, it was phosphorylated in response to glucose at a level similar to wild type FBPase. Therefore, serine 11 is not the only site for FBPase phosphorylation.

In the {Delta}ras2 mutant, a significant amount of FBPase was associated with Vid vesicles, suggesting that Ras2 is involved in the fusion of Vid vesicles with the vacuole. This is consistent with the results that both vesicles and vacuole were defective in fusion in the {Delta}ras2 mutant. Our results suggest that the cAMP signaling pathway plays multiple roles in the vacuole degradation pathway. It is required for the phosphorylation of FBPase, but the serine 11 residue is not the only target for phosphorylation. This signaling pathway is also required for the proper function of Vid vesicles and the vacuole. Further experiments will be needed to identify the downstream effectors regulated by the cAMP signaling pathway and to identify factors that recognize the N-terminal proline residue.


   FOOTNOTES
 
* This work was supported by Public Health Services Grant RO1GM59480 from the National Institutes of Health (NIH) (to H.-L. C.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

{ddagger} Present address: Bldg. 8, Rm. 403, Laboratory of Biochemistry and Genetics, NIDDK/NIH, Bethesda, MD 20892-0830. Back

§ To whom correspondence should be addressed: Dept. of Cellular and Molecular Physiology, Penn State University College of Medicine, 500 University Dr., Hershey, PA 17033. Tel.: 717-531-0859; Fax: 717-531-7667; E-mail: crb13{at}psu.edu.

1 The abbreviations used are: FBPase, fructose-1,6-bisphosphatase; MDH2, malate dehydrogenase; GFP, green fluorescence protein. Back


   ACKNOWLEDGMENTS
 
We thank Dr. A. Kornberg (Stanford University) for the {Delta}pep4{Delta}prb1{Delta}prc1 strain. We thank Dr. McAlister-Henn (University of Texas, Houston, TX) for the wild type and {Delta}nMDH2 plasmids, the {Delta}mdh2strain, and anti-MDH2 antibodies. We thank Dr. Prieto (Instituto de Agroquimica y Tecnologia de los Alimentos, Spain) for the {Delta}hxk1{Delta}hxk2{Delta}glk2 strain. We also thank Dr. Broach (Princeton University) for the tpk1w1{Delta}tpk2{Delta}tpk3{Delta}bcy1mutant strain. Primers used in this study were synthesized in the Core Facility at Penn State University College of Medicine. DNA was sequenced in the Core Facility at Penn State College of Medicine.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Gancedo, J. M. (1998) Microbiol. Mol. Biol. Rev. 62, 334-361[Abstract/Free Full Text]
  2. Carlson, M. (1999) Curr. Opin. Microbiol. 2, 202-207[CrossRef][Medline] [Order article via Infotrieve]
  3. Rolland, F., Winderickx, J., and Thevelein, J. M. (2002) FEMS Yeast Res. 2, 183-201[Medline] [Order article via Infotrieve]
  4. Schuller, H. J. (2003) Curr. Genet. 43, 139-160[Medline] [Order article via Infotrieve]
  5. Holzer, H. (1989) Revis. Biol. Celular. 21, 305-319[Medline] [Order article via Infotrieve]
  6. Gamo, F. J., Navas, M. A., Blazquez, M. A., Gancedo, C., and Gancedo, J. M. (1994) Eur. J. Biochem. 222, 879-884[Medline] [Order article via Infotrieve]
  7. Holzer, H., and Purwin, C. (1986) Biomed. Biochim. Acta 45, 1657-1663[Medline] [Order article via Infotrieve]
  8. Foy, J. J., and Bhattacharjee, J. K. (1978) J. Bacteriol. 136, 647-656[Abstract/Free Full Text]
  9. Gancedo, C. (1971) J. Bacteriol. 107, 401-405[Abstract/Free Full Text]
  10. Zaragoza, O., Lindley, C., and Gancedo, J. M. (1999) J. Bacteriol. 181, 2640-2642[Abstract/Free Full Text]
  11. Zaragoza, O., and Gancedo, J. M. (2001) FEBS Lett. 506, 262-266[CrossRef][Medline] [Order article via Infotrieve]
  12. De Vit, M. J., Waddle, J. A., and Johnston, M. (1997) Mol. Biol. Cell. 8, 1603-1618[Abstract]
  13. De Vit, M. J., and Johnston, M. (1999) Curr. Biol. 9, 1231-1241[CrossRef][Medline] [Order article via Infotrieve]
  14. Ostling, J., and Ronne, H. (1998) Eur. J. Biochem. 252, 162-168[Medline] [Order article via Infotrieve]
  15. Treitel, M. A., Kuchin, S., and Carlson, M. (1998) Mol. Cell. Biol. 18, 6273-6280[Abstract/Free Full Text]
  16. Smith, F. C., Davies, S. P., Wilson, W. A., Carling, D., and Hardie, D. G. (1999) FEBS Lett. 453, 219-223[CrossRef][Medline] [Order article via Infotrieve]
  17. Ludin, K., Jiang, R., and Carlson, M. (1998) Proc. Natl. Acad. Sci. U. S. A. 95, 6245-6250[Abstract/Free Full Text]
  18. Sanz, P., Alms, G. R., Haystead, J., and Carlson, M. (2000) Mol. Cell. Biol. 20, 1321-1328[Abstract/Free Full Text]
  19. Johnston, M. (1999) Trends Genet. 15, 29-33[CrossRef][Medline] [Order article via Infotrieve]
  20. Papamichos-Chronakis, M., Gligoris, T., and Tzamarias, D. (2004) EMBO J. 5, 368-372[CrossRef]
  21. Mazon, M. J., Gancedo, J. M., and Gancedo, C. (1982) J. Biol. Chem. 257, 1128-1130[Abstract/Free Full Text]
  22. Rittenhouse, J., Moberly, L., and Marcus, F. (1987) J. Biol. Chem. 262, 10114-10119[Abstract/Free Full Text]
  23. Jiang, Y., Davis, C., and Broach, J. R. (1998) EMBO. J. 17, 6942-6951[CrossRef][Medline] [Order article via Infotrieve]
  24. Schork, S. M., Bee, G., Thumm, M., and Wolf, D. H. (1994) FEBS Lett. 349, 270-274[CrossRef][Medline] [Order article via Infotrieve]
  25. Hammerle, M., Bauer, J., Rose, M., Szallies, A., Thumm, M., Dusterhus, S., Mecke, D., Entian, K. D., and Wolf, D. H. (1998) J. Biol. Chem. 273, 25000-25005[Abstract/Free Full Text]
  26. Schule, T., Rose, M., Entian, K. D., Thumm, M., and Wolf, D. H. (2000) EMBO J. 19, 2161-2167[CrossRef][Medline] [Order article via Infotrieve]
  27. Horak, J., Regelmann, J., and Wolf, D. H. (2002) J. Biol. Chem. 277, 8248-8254[Abstract/Free Full Text]
  28. Regelmann, J., Schule, T., Josupeit, F. S., Horak, J., Rose, M., Entian, K. D., Thumm, M., and Wolf, D. H. (2003) Mol. Biol. Cell 14, 1652-1663[Abstract/Free Full Text]
  29. Schork, S. M., Thumm, M., and Wolf, D. H. (1995) J. Biol. Chem. 270, 26446-26450[Abstract/Free Full Text]
  30. Chiang, H.-L., and Schekman, R. (1991) Nature 350, 313-318[CrossRef][Medline] [Order article via Infotrieve]
  31. Chiang, H.-L., Schekman, R., and Hamamoto, S. (1996) J. Biol. Chem. 271, 9934-9941[Abstract/Free Full Text]
  32. Brown, C. R., McCann, J. A., and Chiang, H.-L. (2000) J. Cell Biol. 150, 65-76