Mammalian Δ9 stearoyl-CoA desaturase 1 (SCD1) is a key enzyme in the biosynthesis of mono-unsaturated fatty acids in the endoplasmic reticulum (ER). It is a short-lived multispanning ER membrane protein, reported to be degraded by the ubiquitin-proteasome-independent pathway. We have examined SCD1 protein degradation using cultured mammalian cells. Exogenously expressed SCD1 in CHO-K1 cells was localized to the ER and turned over with a half-life of ∼3 hours. Unexpectedly, proteasome inhibitors increased the half-life of SCD1 to ∼6 hours. Endogenously expressed SCD1 in adipocyte-differentiated NIH 3T3-L1 cells was also rapidly degraded in a proteasome inhibitor-sensitive manner. In the presence of proteasome inhibitors, polyubiquitylated SCD1 accumulated in the ER and interacted with AAA-ATPase p97, which is involved in ER-associated degradation (ERAD). The 66-residue N-terminal segment carrying the PEST sequence is mainly responsible for SCD1 degradation and this segment induced instability in an otherwise stable ER membrane protein. Furthermore, SCD1 was degraded constitutively irrespective of the cellular levels of unsaturated fatty acids, which strictly regulate SCD1 gene expression. These findings indicate that the ubiquitin-proteasome-dependent ERAD system is also involved in constitutive SCD1 degradation.
The endoplasmic reticulum (ER) is the center of biosynthesis, folding and quality control of a variety of secretory proteins and secretory pathway-localized proteins, including membrane proteins (Sitia and Braakman, 2003). Perturbations that disrupt protein folding lead to an accumulation of unfolded proteins or protein aggregates in the ER, and eukaryotic cells have several distinct mechanisms to cope with these situations (Mori, 2000; Kaufman et al., 2002): (1) retention of unfolded proteins in the ER by the ER-Golgi retrieval pathway, (2) upregulation of ER chaperones, the so-called unfolded protein response, (3) transient repression of de novo protein biosynthesis to maintain the fidelity of protein folding (attenuation), and (4) removal of misfolded proteins by the proteasome-dependent ER-associated degradation (ERAD) pathway. In the process of protein elimination by proteasome-dependent ERAD, protein substrates are polyubiquitylated by the E2 ubiquitin-conjugating enzymes and E3 ubiquitin ligases, then retro-translocated to the cytosol by the concerted action of AAA-ATPase Cdc48p/p97 and the cofactor Npl4p-Ufd1p complex through the retrograde translocation channel, presumably including Sec61p, and finally degraded by proteasomes (Ellgaard and Helenius, 2003; Hampton, 2002). Recent experiments in yeast revealed that the ER membrane protein Ubx2p recruits the Cdc48p complex to ubiquitin ligases and ERAD substrates (Schuberth and Bachberger, 2005; Neuber et al., 2005). In mammals, the ER membrane protein VIMP recruits the p97-Ufd1-Npl4 complex to the putative dislocation channel protein derlin-1 (Ye et al., 2004; Lilley and Ploegh, 2004). The proteasome-dependent ERAD system is used not only for the removal of unfolded or unassembled membrane proteins, such as the ΔF508 mutant of the cystic fibrosis transmembrane conductance regulator or T-cell receptor α-subunit (TCRα), but also for degradation of the ER-resident normal proteins such as hydroxymethylglutaryl-CoA reductase (HMGR), Ubc6p, and yeast Δ9-fatty acid desaturase (Walter et al., 2001; Hampton, 2002; Braun et al., 2002).
Mammalian Δ9 stearoyl-CoA desaturase 1 (SCD1) is a short-lived integral membrane protein of the ER (Oshino and Sato, 1972). It is a key enzyme in the biosynthesis of mono-unsaturated fatty acids and catalyzes the introduction of the cis double bond at the Δ9 position of palmitoyl- or stearoyl-CoA to form palmitoleyl- or oleyl-CoA, respectively, by the oxygenase reaction, in which the reducing equivalents are provided by NADH via NADH-cytochrome b5 reductase and cytochrome b5 (Shimakata et al., 1972; Strittmatter et al., 1974). A key regulator of SCD1 gene expression is sterol-regulatory element-binding protein-1c (SREBP-1c), which mediates insulin-induced transcriptional activation of the SCD1 gene (Ntambi, 1999; Bene et al., 2001; Horton et al., 2002). SCD1 protein is induced 50- to 100-fold when dietary fat is restricted, whereas it rapidly decreases to undetectable levels with a half-life of 3-4 hours after the restricted-fat dietary regimen is stopped (Oshino and Sato, 1972).
In yeast, the abundance of yeast Δ9 fatty acid desaturase, Ole1p, in the ER is transiently regulated at the gene transcription level and by degradation of Ole1p. OLE1 is activated by the OLE pathway, in which transcription factors Spt23p and Mga2p are synthesized as ER-bound precursor proteins. Upon unsaturated fatty acid starvation, they are activated by a regulated ubiquitin-proteasome-dependent proteolysis, and translocated to the nucleus to activate OLE1 transcription (Rape et al., 2001). Furthermore, Ole1p is naturally short-lived and is degraded by ERAD (Braun et al., 2002).
Mammalian SCD1 is degraded by a proteasome-independent pathway (Ozols, 1997). Mziaut et al. (Mziaut et al., 2000) demonstrated that SCD1 fused to the N-terminus of green fluorescent protein (GFP), when expressed in CHO-K1 cells, was correctly targeted to the ER and, in contrast to GFP, became unstable in the ER and degraded with a half-life of a few hours. Mziaut et al. also demonstrated that deletion of the 26- or 44-residue N-terminal segment from the SCD1-GFP fusion protein led to stabilization of the truncated proteins. In turn, the 1 to 33-residue N-terminal segment transplanted to the N-terminus of the GFP rendered the reporter extremely short-lived. They thus concluded that the ∼30-residue N-terminal segment of SCD1 constitutes a motif responsible for the rapid degradation of SCD1. In a recent report, Ozols and collaborators characterized the protease responsible for SCD1 degradation using an in vitro assay system with SCD1-induced rat liver microsomes (Ozols, 1997; Heinemann and Ozols, 1998), and purified a plasminogen (PLG)-like protease as the SCD1-specific protease (Heinemann et al., 2003b). SCD1 protease is presumably in the 90 kDa proform, and its conversion to a series of smaller proteins markedly increases its proteolytic activity (Heinemann et al., 2003b). In vitro degradation of SCD1 is not inhibited by a wide variety of protease inhibitors; lysosomotrophic agents (chloroquine, leupeptin and pepstatin A), the calpain inhibitor N-acetyl-leucyl-methionyl, and the proteasome inhibitor lactacystin do not inhibit the in vitro degradation of SCD1 in isolated microsomal membranes (Ozols, 1997). The responsible protease appears to be a serine or thiol protease, because SCD1 is sensitive to dithiothreitol, diisopropyl phosphofluoridate, and leupeptin (Heinemann et al., 2003a; Heinemann et al., 2003b). In spite of these findings, however, there is limited information on SCD1 degradation in vivo.
SCD1 is a typical PEST sequence-rich membrane protein (Rogers et al., 1986). Generally, PEST sequence-containing proteins are targets of rapid protein degradation (Rechsteiner and Rogers, 1996). Although the pathways (or enzymes) responsible for degrading PEST proteins remain controversial, a considerable body of evidence supports the idea that PEST sequences target proteins for degradation by the ubiquitin-proteasome system (Rechsteiner and Rogers, 1996). In light of the apparently distinct mechanisms for degradation of SCD proteins between yeast and mammalian cells, we used cultured mammalian cells to examine whether SCD1 is degraded exclusively by the ubiquitin-independent pathway, or whether the ubiquitin-dependent pathway is also involved in parallel.
Here, we demonstrate that exogenously expressed SCD1 in CHO-K1 cells, as well as the endogenously expressed form in adipocyte-differentiated 3T3-L1 cells, was degraded rapidly with a half life of less than 3.5 hours, and this degradation was significantly inhibited by the proteasome inhibitors, MG132, and epoxomicin (P<0.01). Consistent with these findings, the polyubiquitylated SCD1 intermediates accumulated in the ER in the presence of MG132, and interacted with AAA-ATPase p97, which is required for the retro-translocation of the ERAD (Ellgaard and Helenius, 2003), thus indicating involvement of the ERAD pathway in controlling cellular levels of mammalian SCD1, as is the case for yeast Ole1p (Braun et al., 2002). Furthermore, we demonstrated that the 66-residue N-terminal segment of SCD1 containing the PEST sequences has an important role in proteasomal degradation of ER-localizing proteins.
SCD1 is degraded by a proteasome-dependent pathway
Using a cell-free assay system, Ozols and collaborators demonstrated that SCD1 protein is degraded through a proteasome-independent pathway, although in vivo evidence has been lacking (Ozols, 1997; Heimemann and Ozols, 1998). We, therefore, addressed this point in cultured mammalian cells. CHO-K1 cells transiently expressing HA-tagged SCD1 were subjected to pulse-chase analysis with [35S]methionine, followed by immunoprecipitation using an anti-HA antiserum. In the absence of the proteasome inhibitor MG132, SCD1 protein was degraded with a half life of ∼3 hours (Fig. 1A). In the presence of MG132, the half-life of SCD1 increased to ∼6 hours. This effect was also observed with another proteasome inhibitor, epoxomicin (Fig. 1B). Similarly, exogenously expressed SCD1 was degraded rapidly in HeLa cells, and MG132 inhibited the degradation (Fig. 1A). The same results were obtained in HEK-293 cells (data not shown). As a control, the degradation of TCRα, a typical ERAD substrate (Huppa and Ploegh, 1997), was also efficiently inhibited by MG132 (t1/2<2 hours in the absence of the inhibitor, and t1/2>8 hours in the presence of the inhibitor; Fig. 1A). Heinemann et al. (Heinemann et al., 2003a) demonstrated that SCD1 degradation is inhibited by leupeptin in vitro. By contrast, however, in our study the stability of SCD1 in vivo was not affected by an even higher concentration (100 μM) of leupeptin (Fig. 1B). These results suggest that proteasomes are involved in the degradation of SCD1 protein in cultured cells.
Endogenously expressed SCD1 protein is also degraded by a proteasome-dependent pathway
To exclude the possibility that the rapid degradation of SCD1 and its inhibition by proteasome inhibitors were due to exogenous overexpression of SCD1 in CHO-K1 cells, we analyzed the metabolic stability of endogenously expressed SCD1 protein in adipocyte-differentiated NIH 3T3-L1 cells (Green and Meuth, 1974). NIH 3T3-L1 cells expressed SCD1 during differentiation into adipocytes (Fig. 2A). The adipocyte-differentiated cells were subjected to pulse labeling with [35S]methionine and chase reactions, followed by immunoprecipitation with anti-SCD1 antiserum (Fig. 2B). The endogenously expressed SCD1 was degraded with a half-life of ∼3.5 hours, and MG132 effectively inhibited this degradation (t1/2∼7.5 hours; Fig. 2B). Together, these results indicate that proteasomes are involved in the degradation of endogenous SCD1 protein.
Accumulation of ubiquitylated SCD1 in the presence of MG132
We then examined whether SCD1 is modified by polyubiquitin chains, using immunoprecipitation. CHO-K1 cells transiently transfected with empty plasmid or plasmids harboring SCD1-HA cDNA were cultured in the presence or absence of MG132. They were lysed in buffer containing SDS and the cell lysates were subjected to immunoprecipitation using anti-HA serum and subsequent immunoblot analysis with anti-polyubiquitin monoclonal antibody (Fig. 3A, lanes 1-5). Polyubiquitylated SCD1-HA was detected in the presence of MG132 (Fig. 3A, lane 5). Immunoblot analysis with anti-HA antiserum revealed that the steady state level of SCD1-HA increased in the presence of MG132 (Fig. 3A, compare lanes 9 and 10). However, only a faint band of polyubiquitylated SCD1-HA was detected with anti-HA antiserum (Fig. 3A, lower panel); detection of ubiquitin-modified proteins at steady state seemed to be difficult, as reported previously (Liao et al., 1998). The same results were obtained for endogenously expressed SCD1 in differentiated 3T3-L1 cells (Fig. 3B); a polyubiquitylated band for the immunoprecipitate with anti-SCD1 peptide antibodies was detected only for cells grown in the presence of MG132 (lane 6), but not in its absence (lane 5; DMSO). The immunoprecipitates with control IgGs did not have such signals, irrespective of the presence or absence of MG132 (Fig. 3B, lanes 1-3), indicating that this ubiquitylation was not due to exogenous overproduction of SCD1 protein. To further confirm the polyubiquitylation of SCD1, SCD1-HA and Myc-tagged ubiquitin were coexpressed in the presence of MG132 in CHO-K1 cells, and the cell lysates were subjected to immunoprecipitation using anti-Myc IgGs, which were then subjected to immunoblotting with anti-HA antibodies (Fig. 3C). Again, the polyubiquitylated bands were detected for the immunoprecipitates processed with anti-HA antibodies (Fig. 3C, lane 4). The lysates prepared from the cells that were transfected with either empty plasmid or plasmids harboring SCD1-HA cDNA or Myc-ubiquitin cDNA did not show any ubiquitylation, even in the presence of MG132 (Fig. 3C, lanes 1-3). Taken together, these results clearly indicate that SCD1 protein is degraded via a ubiquitin-proteasome pathway.
Ubiquitinated SCD1 accumulates in the ER and interacts with p97
Typical proteasome-dependent ERAD substrates are ubiquitylated on the ER and retro-translocated from the ER to the cytoplasm by the p97-Ufd1-Npl4 complex (Meyer et al., 2002; Ye et al., 2003). We therefore examined whether SCD1 degradation depends on the ERAD system. CHO-K1 cells transfected with either pcDNA3.1 or plasmids harboring SCD1-HA cDNA were cultured in the presence or absence of MG132 and fractionated into supernatant and microsomal fractions, which were subjected to immunoprecipitation using anti-HA antiserum. Immunoblot analysis of the immunoprecipitates with anti-polyubiquitin monoclonal antibody revealed that the polyubiquitylated SCD1-HA was enriched in the microsomal fraction, but not in the supernatant fraction (Fig. 4A, compare lanes 11 and 12). By contrast, the polyubiquitylated smear was undetectable in cells transfected with empty plasmid, irrespective of the presence or absence of MG132 (Fig. 4A, lanes 5-8), or detected only weakly in the microsomes of cells expressing SCD1-HA in the absence of MG132 (lane 10). Furthermore, recovery of SCD1-HA in the ER fraction clearly increased in the presence of MG132 (compare lanes 23 and 25), confirming the results in Fig. 3A.
Of note, p97 co-immunoprecipitated with SCD1-HA from the microsomal fraction of the SCD1-HA-expressing cells that were grown in the presence of MG132 (Fig. 4A, lane 38), whereas the signal was not detectable in the microsomes of cells grown under all the other conditions (Fig. 4A, lanes 32, 34, and 36). Approximately 1.3% of the total amount of p97 was recovered to the ER of the cells expressing SCD1-HA (Fig. 4B). We cannot, however, rigorously rule out the trivial possibility that p97 became associated with SCD1-HA after membrane solubilization. Taken together, these results strongly suggest that SCD1 is degraded by proteasome-dependent ERAD.
The N-terminal segment is important for SCD1 degradation by the ubiquitin-proteasome system
The rapid turnover of the SCD1-GFP fusion protein in CHO-K1 cells is compromised by the deletion of either the 27- or 45-residue N-terminal segment of SCD1 (Mziaut et al., 2000). Furthermore, appending the 1 to 33-residue N-terminal segment of SCD1 onto the N-terminus of GFP destabilizes the fusion protein, although it is not targeted to the ER but remains in the cytoplasm, indicating that the ∼30-residue N-terminal segment fused to a reporter protein functions as a rapid degradation signal, even if the fusion protein remains in the cytoplasm (Mziaut et al., 2000). A web-based algorithm PEST-Find (http://www.at.embnet.org/embnet/tools/bio/PESTfind) revealed PEST sequences in the N-terminus of SCD1 (Fig. 5A), suggesting that the N-terminal region has an important role in proteasome-dependent SCD1 degradation. We examined this hypothesis using a 66-amino acid N-terminal-truncated SCD1 mutant (Δ66 SCD1). When expressed in CHO-K1 cells, Δ66 SCD1 (Fig. 5B) localized to the ER, just like the wild-type protein (Fig. 5C). In pulse-chase experiments, Δ66 SCD1 was stable compared with wild-type SCD1 (Fig. 5D, compare left and right panels), confirming the previous report by Mziaut et al. (Mziaut et al., 2000). In addition, the residual degradation reaction of Δ66 SCD1 was insensitive to epoxomicin (Fig. 5D, right), indicating that the N-terminal segment of SCD1 is important for proteasome-dependent degradation of SCD1.
The 66-residue N-terminal segment of SCD1 destabilizes other ER-localized membrane proteins
To further examine the function of the N-terminal region of SCD1, we constructed fusion proteins in which the 66-residue N-terminal segment was fused to the N-terminus of EGFP, SCD66-EGFP. When expressed in CHO-K1 cells, EGFP and SCD66-EGFP both localized to the cytoplasm (Fig. 6A) and degraded slowly, and both degradations were epoxomicin-insensitive (Fig. 6B). Analysis of the stability of the EGFP constructs including SCD33-EGFP by FACScan confirmed these results (Fig. 6C). These observations were in marked contrast to the findings of Mziaut et al. (Mziaut et al., 2000), who reported that the fusion construct in which the 1 to 33-residue N-terminal segment of SCD1 was fused to the N-terminus of EGFP (SCD33-EGFP), and was extremely short-lived (t1/2 ∼1.5 hour) in CHO-K1 cells. We have no adequate explanation for this discrepancy. It should be noted, however, that the expression efficiency as assessed by flow cytometry was 40% for EGFP and 15% for SCD1wt-EGFP: similar levels to those reported by Mziaut et al. (Mziaut et al., 2000); 45.8% for the former and 14.4% for the latter). By contrast, the expression efficiency was distinctly different for SDC33-EGFP: 35% in the present study, but 6.2% in the previous report.
We next examined whether the ER localization signal destabilizes the SCD66-appended substrates. For this purpose, EGFP was fused to the C terminus of cytochrome b5, the C-terminal tail anchored ER membrane protein, to construct b5-EGFP. Furthermore, SCD66 was fused to the N-terminus of b5-EGFP to create SCD66-b5-EGFP. When expressed in CHO-K1 cells, SCD66-b5-GFP and b5-GFP both co-localized with the ER marker protein calnexin (Fig. 7A). Analysis of the metabolic stability of these fusion proteins by [35S]methionine pulse-chase experiments, indicated that the turnover rate of b5-EGFP was relatively slow (Fig. 7B, left panel). By contrast, SCD66-b5-GFP was rapidly degraded with a half life of 4 hours, and the degradation was significantly inhibited by epoxomicin (Fig. 7B, right panel) (P<0.05). Taken together, these results indicated that the 66-residue N-terminal segment of SCD1 functions as a degradation signal on the ER.
SCD1 degradation is not affected by cellular levels of unsaturated fatty acids or sterols
SCD1 gene transcription is regulated by unsaturated fatty acids, and exogenous polyunsaturated fatty acids repress SCD1 gene transcription (Ntambi, 1995; Waters and Ntambi, 1996). Furthermore, the SCD1 gene is SREBP-responsive gene that is involved in cholesterol homeostasis (e.g. HMG-CoA synthase, HMGR, squalen synthase or low density lipoprotein receptor), fatty acid synthesis (e.g. fatty acid synthase, acetyl-CoA carboxylase) and fatty acid desaturation (Shimano, 2001), and transcription of these genes is regulated by cellular sterols; transcription is repressed by excess sterols, and induced by sterol depletion (Tabor et al., 1999). In particular, cellular sterol levels exert feed-back regulation on HMGR in the ER through multiple mechanisms; high sterol levels shut-down transcription of the HMGR gene concomitant with accelerated protein degradation (Goldstein and Brown, 1990). The HMGR gene should also be regulated by unsaturated fatty acids, given that SREBP isoforms are down-regulated by unsaturated fatty acids (Hannah et al., 2001). The degradation of HMGR is accelerated by the sterol-induced binding of its sterol-sensing domain to Insig-1 (Sever et al., 2003). The apparent similarity of the regulation of gene expression between the HMGR and SCD1 genes led us to examine whether SCD1 degradation is regulated by cellular levels of sterols or polyunsaturated fatty acids. For this purpose, the SCD1-HA-transfected HEK-293 cells (CHO-K1 cells could not be used in this experiment because medium containing linoleic acid must be used for the culture) were cultured in the presence of either lipid-depleted fetal bovine serum (FBS) or untreated FBS, and the turnover rate of SCD1-HA was examined after pulse-chase analysis using [35S]methionine (Fig. 8). Real-time PCR revealed that gene transcription for HMGR, low density lipoprotein receptor and SCD1 was induced in response to the presence of delipidated FBS, whereas transcription of the glyceraldehyde 3-phosphate dehydrogenase (GAPDH) gene, which is unrelated to lipid biosynthesis, was not affected (Fig. 8A). The turnover of SCD1, however, was not altered in the presence of delipidated FBS (Fig. 8B,C). Similarly, overexpression of Insig-1 or replenishment of polyunsaturated fatty acids in cultured CHO-K1 cells did not significantly affect the SCD1 degradation rate (data not shown).
These results suggested that rapid degradation of SCD1 is not affected by either polyunsaturated fatty acids or sterols, and proceeds constitutively. Taken together, the data demonstrated that SCD1 is a naturally short-lived protein that is degraded constitutively by the ubiquitin-proteasome pathway. Thus, cellular levels of SCD1 seem to be determined by the SCD1 gene transcription rate and SCD1 mRNA stability (Ntambi, 1995; Sessler et al., 1996).
Exogenous expression of plasminogen does not influence SCD1 protein degradation in cultured cells
SCD1 is degraded by plasminogen (PLG)-like protein in rat liver microsomes (Heinemann et al., 2003b). Immunoblot analysis using anti-human PLG antibodies, which gave strong signals for Chinese hamster liver extracts or its microsomes, or recombinant human PLG-flag, revealed that PLG was scarcely expressed in HeLa cells, CHO-K1 cells or HEK-293 cells (Fig. 9A). We therefore examined whether exogenously expressed PLG influences SCD1 degradation in cultured cells. SCD1-HA and flag-tagged human PLG (PLG-flag) were co-expressed in CHO-K1 cells, and the steady-state level and turnover rate of SCD1-HA were both measured. The expressed PLG-flag localized mainly in P1, composed of mitochondria and heavy microsomes, and P2, composed of microsomes, indicating that significant amounts of the expressed PLG-flag localized within the secretory organelles (Fig. 9B). We then examined if the steady-state level of SCD1-HA is influenced by co-expression with increasing amounts of PLG-flag. There were no clear dose-dependent decreases, however, in SCD1-HA (Fig. 9C). Similarly, [35S]methionine pulse-chase experiments revealed that SCD1-HA degradation was not influenced by co-expression with PLG-flag (Fig. 9D).
Knockdown of PLG does not affect SCD1 protein degradation in HepG2 cells
In contrast to the cells examined above, we found that PLG was expressed to a detectable level in HepG2 cells (Fig. 9E). Therefore, we examined the effect of PLG-knockdown on SCD1 turnover in HepG2 cells. Exogenously expressed SCD1-HA was degraded in control HepG2 cells with a half-life similar to those in HeLa cells or CHO-K1 cells. Unexpectedly, degradation rate of SCD1-HA was not affected even after ∼93% depletion of PLG (Fig. 9F,G). Together, these results indicate that PLG is not involved in SCD1 degradation, at least in cultured cells.
Ozols and collaborators developed an in vitro assay for SCD1 degradation using microsomes isolated from SCD1-induced rat liver, and demonstrated that the degradation is rapid (half-life of ∼5 hours), selective for SCD1 and proceeds by a non-proteasome pathway; the reaction is not inhibited by either lysosomal protease inhibitors, calpain inhibitors or proteasome inhibitors (Ozols, 1997; Heinemann and Ozols, 1998). Using this system, they purified a ∼90 kDa microsomal endopeptidase as the SCD1 protease, which exhibited serine or thiol protease characteristics, as it was inhibited by diisopropyl phosphofluoridate, dithiothreitol or leupeptin (Heinemann et al., 2003a). The ∼90 kDa form seems to be a proform, because it is degraded to smaller-size fragments during purification, concomitant with an increase in its activity (Heinemann et al., 2003b). The protease was identified as a microsomal form of PLG, because the reaction is inhibited by a plasmin-specific inhibitor, Bdellin (Heinemann et al., 2003b). In this regard, in vitro degradation of SCD1 in liver microsomes isolated from PLG-deficient mice was markedly compromised (Heinemann et al., 2003b).
To our knowledge, however, there are no in vivo data on the cellular stability of SCD1; even data on the effects of proteasome inhibitors have been lacking. Therefore, the question has remained as to whether the microsomal form of plasminogen is the enzyme solely responsible for the rapid degradation in vivo. In this study, we examined the cellular stability of SCD1 using cultured mammalian cells and demonstrated that there is a ubiquitin-proteasome-dependent pathway for SCD1 degradation. The degradation of SCD1 was inhibited by proteasome inhibitors, and as such, the level of microsomal SCD1 was increased. Furthermore, polyubiquitylated SCD1 accumulated in the ER in the presence of proteasome inhibitors and interacted with AAA-ATPase p97, which is essential for proteasome-dependent ERAD. These results suggest that SCD1 is degraded by proteasome-dependent ERAD.
We addressed the involvement of a PLG-like protease in the SCD1 degradation in cultured cells. The reported sequence of SCD1 protease spans the entire PLG sequence except for the 68-residue N-terminal segment of the secreted form (Heinemann et al., 2003b), although neither the topogenic sequence nor the membrane topology have been reported. Consistent with this notion, knockout of the PLG gene influenced SCD1 degradation as examined by an in vitro degradation assay (Heinemann et al., 2003b). We, therefore, exogenously expressed human PLG in CHO-K1 cells and examined whether the expression affected the cellular stability of SCD1. PLG was expressed in the cells and was found to be localized in the secretory organelles. There was no significant stimulation, however, of SCD1 degradation, although it remains unknown whether the expressed PLG assumed the correct topology in the ER. In CHO-K1 cells and other cultured mammalian cells including HeLa and HEK-293, immunoblotting revealed no significant expression of PLG; nevertheless SCD1 was rapidly degraded. Conversely, we found that PLG was expressed to a detectable level in HepG2 cells and exogenously expressed SCD1-HA was degraded with a half-life similar to that in CHO-K1 or HeLa cells, and SCD1-HA degradation was not affected by PLG knockdown. Taken together, we considered that the ubiquitin-proteasome system is mainly responsible for the rapid SCD1 degradation in cultured mammalian cells, although these results do not necessarily exclude a PLG-dependent degradation pathway.
We demonstrated that the 66-residue N-terminal segment of SCD1 is rich in PEST sequences and is important for proteasome-dependent degradation. Deletion of the 66-residue N-terminal segment made SCD1 more stable than the wild-type enzyme, but the degradation was not completely arrested by this manipulation and the mutant (Δ66SCD1) was degraded at a slower rate in a proteasome inhibitor-resistant manner, suggesting that Δ66 SCD1 was degraded by some non-proteasomal pathway.
Similarly, several membrane proteins are degraded by a non-proteasomal pathway, e.g. a type I transmembrane protein thyroperoxidase and polytopic membrane protein voltage sensor mutant of the Shaker K+-channel subunit (Fayadat et al., 2000; Myers et al., 2004). The functional division, however, of the proteasomal and non-proteasomal degradation pathways in the ER remains unknown.
We demonstrated here that the 66-residue N-terminal segment of SCD1 containing PEST sequences (SCD66) made a stable ER membrane protein, b5-EGFP, susceptible to proteasome-dependent degradation. By contrast, it failed to stimulate the degradation of cytoplasm-localizing EGFP. These results indicated that SCD66 is not a general degradation signal, but functions as a specific degradation signal in the ER membrane. In the ER, there might be a mechanism that recognizes the SCD66 segment and leads to proteasome-mediated degradation.
HMGR is a polytopic ER membrane protein and a rate-limiting enzyme of the cholesterol biosynthesis pathway. It is well established that its cellular level is regulated by at least two independent control systems that respond to cellular levels of sterols or unsaturated fatty acids. One involves sterol repression of the HMGR gene through regulated cleavage of the ER membrane-bound transcription factor SREBP, the second involves sterol-induced rapid degradation of HMGR, the regulated degradation via the ERAD system.
In spite of the similar gene regulation between HMGR and SCD1, the SCD1 half-life in cultured cells remained unaffected by feeding unsaturated fatty acids or culture in the presence of lipid-depleted FBS, albeit expression of the SCD1 gene responded properly. We thus concluded that SCD1 is naturally short-lived and is constitutively degraded within the ER by the ubiquitin-proteasome system, as is also the case for the yeast SCD, Ole1p (Braun et al., 2002).
Materials and Methods
The following reagents were obtained from the companies shown in parentheses. MG132 (Sigma Chemical Co., St Louis, MO, USA), epoxomicin and leupeptin (Peptide Institute, Osaka, Japan), an expression vector in mammalian cells, pcDNA3.1 (Invitrogen, Carlsbad, CA, USA), rabbit anti-hemagglutinin (HA) antiserum (Covance, Rapids, MI, USA), rabbit anti-calnexin polyclonal antibodies (Stress Gen, Victoria, Canada), mouse anti-Myc antibody (Upstate, Waltham, MA, USA), mouse anti-polyubiquitin monoclonal antibody (Medical & Biological Laboratories, Aichi, Japan), mouse anti-p97 monoclonal antibody (Progen Biotechnik, Heidelberg, Germany), goat anti-human PLG antibodies (Cedarlane Laboratories, Ontario, Canada), protein A-Sepharose (Amersham Biosciences, Piscataway, NJ, USA), and Pfu turbo DNA polymerase (Stratagene, La Jolla, CA, USA). Antibodies against SCD1 were raised in rabbits against a synthetic 15-mer peptide corresponding to the C-terminal 15 amino acid residues of rat SCD1. Antibodies against rat Tim23 and EGFP were as described by Ishihara and Mihara (Ishihara and Mihara, 1998) and Miyazaki et al. (Miyazaki et al., 2001), respectively.
Vector and cDNA
To construct pCMV SCD1-HA and pCMV Δ66SCD1-HA, the cDNA fragments of rat SCD1 were amplified by polymerase chain reaction (PCR) using SCD1 cDNA (Mihara, 1990) as the template and the following oligonucleotides as the primers, and cloned between the HindIII and XbaI sites in pRcCMV-HA (Ukaji et al., 2002). SCD1-wt-F: CACAAGCTTCCCACCATGCCGGCCCACATGCTC; SCD1-wt-R: GTGTCTAGAGCTACTCTTGTGGCTCCC; SCD1-Δ66-F: CACAAGCTTCCCACCATGAAGCTGGAGTACGTCTGGAGG.
To construct pCMV b5-EGFP, the cDNA fragment was amplified by PCR using rat cytochrome b5 cDNA (Mitoma and Ito, 1992) as the template and the following oligonucleotides as the primers, and cloned between the HindIII and BamHI sites in EGFP-N1 (Clontech, Palo Alto, CA, USA). b5-F: CACAAGCTTCCCACCATGGCCGAGCAGTCA; b5-R: GCTTGGATCCCGATCTTCTGCCATGTAGAGGC.
To construct pCMV SCD66-b5-EGFP, cDNA fragments encoding the 66-residue N-terminal segment (SCD66) and cytochrome b5 were generated by PCR, and then connected by overlapping PCR. The obtained PCR fragment was cloned between the HindIII and BamHI sites in EGFP-N1 (Clontech). To construct pCMV TCRα-3xflag, mouse TCRα cDNA was amplified by PCR using pHDS58 (Saito et al., 1984) as the template and the following oligonucleotides as the primers, and cloned between the EcoRI and BamHI sites in p3xFLAG-CMV-14 (Sigma). TCR-F: GAGCGAATTCCCACCATGCTCCTGGCACTCCTCCCA; TCR-R: CCGGGATCCACTGGACCACAGCCTCAGCGT.
To construct SCD1-wt-enhanced GFP (EGFP), SCD66-EGFP and SCD33-EGFP were amplified by PCR using SCD1 cDNA as a template, and SCD1-wt-F, SCD1-wt-GR; SCD1-wt-F, SCD1-66-R; and SCD1-wt-F, SCD1-33-R as the primers, and the obtained cDNA fragment was cloned between HindIII and AgeI sites in EGFP-N1 (Clontech). SCD1-wt-GR: GGCGACCGGTGGCCGGCTACTCTTGTGGCTCCCATC; SCD1-66-R: GGCGACCGGTGGCGCGGGCGGGGGCCCCTCCTCATC; SCD1-33-R: GGCGACCGGTGGCTTCTCTCGTCCATTCTGCAG.
To construct flag-tagged plasminogen (PLG-flag), human plasminogen was amplified by PCR using a human kidney cDNA library (Clontech) as a template, with PLG-F and PLG-R as the primers, and the obtained cDNA fragment was cloned between the HindIII and XbaI sites in p3xFLAG-CMV-14 (Sigma). PLG-F: GAGCAAGCTTCCACCATGGAACATAAGGAAGTGGTTCTT; PLG-R: CTGTCTAGAATTATTTCTCATCACTCCCTCAATCCAAGT.
To construct Myc-tagged ubiquitin, human ubiquitin was amplified by PCR a using a human kidney cDNA library (Clontech) as a template, and ubi-F and ubi-R as the primers, and the obtained cDNA fragment was cloned between the EcoRI and KpnI sites in pCMV-Myc (Clontech). ubi-F: GGACGAATTCGGATGCAGATCTTCGTGAAGACTCTGACT; ubi-R: CCTTGGTACCTTACCCACCTCTGAGACGGAGCACCAGGT
Cell culture and transfection
CHO-K1 cells were cultured in F-12 medium supplemented with 10% fetal bovine serum (FBS) and 10 μg/ml gentamycin (Invitrogen). HEK-293 and HeLa cells were cultured in Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% FBS and 10 μg/ml gentamycin. HepG2 cells were cultured in minimum essential medium (MEM) supplemented with 1% non essential amino acids, 1 mM sodium pyruvate, 2 mM glutamine and 10% FBS. Liposome-mediated transient transfection was performed using lipofectamine (Invitrogen) according to the manufacturer's instructions. The cells were cultured at 37°C under 5% CO2 after transfection. NIH 3T3-L1 cells were cultured in DMEM supplemented with 10% FBS and 10 μg/ml gentamycin. For differentiation of cells into adipocytes, confluent preadipocyte monolayers were incubated for 48 hours in DMEM containing 10% FBS and differentiation cocktail consisting of 115 μg/ml methylisobutylxanthine (Sigma), 390 ng/ml dexamethasone (Sigma), and 10 μM insulin (Sigma). After 48 hours culture, the cells were maintained in DMEM containing 10% FBS and 10 μM insulin, changing the medium every 2 days. The cell morphology was monitored daily for the appearance of cytoplasmic lipid droplets using a phase-contrast microscope and oil Red O staining.
Small interference RNAs (siRNAs) targeting EGFP (control) and PLG were chemically synthesized: EGFP sense: 5′-CUACAACAGCCACAACGUCTT-3′; EGFP antisense: 5′-GACGUUGUGGCUGUUGUAGTT-3′; PLG sense: 5′-CCGCAAUCCUGACGGAAAATT-3′; PLG antisense: 5′-UUUUCCGUCAGGAUUGCGGTT-3′.
HepG2 cells were transfected with siRNA using Lipofectamine 2000 reagent (Invitrogen). After 12 hours, the cells were transfected with pCMV SCD1-HA using Fugene 6 (Roche Diagnostics, Penzberg, Germany). After 12 hours, the cells were transfected with siRNA as described above, cultured for 24 hours, and then subjected to pulse-chase experiments.
Cells on coverslips were fixed with 4% paraformaldehyde at room temperature for 20 minutes and permeabilized with 0.1% Triton X-100. They were incubated with 1% bovine serum albumin in phosphate-buffered saline (PBS) for 20 minutes, and then incubated with primary antibodies for 1 hour at room temperature. After several washes in PBS, the coverslips were incubated with fluorescein isothiocyanate-conjugated goat anti-mouse antibodies (BioSource, Camarillo, CA, USA) or Texas-Red-conjugated goat anti-rabbit antibodies (Cappel, Durham, NC, USA) for 1 hour at room temperature, followed by washing in PBS. Fluorescent images were taken using a confocal laser microscope Radiance 2000 (BioRad, Hercules, CA) and analyzed.
Cells (in a 3.5 cm dish) were incubated for 1 hour in cysteine- and methionine-free medium. They were then pulse-labeled for 30 minutes with 1.85 MBq 35S-labeling mix (Perkin Elmer, Wellesley, MA, USA) in 1 ml culture medium in the absence or presence of proteasome inhibitors. The labeling medium was removed, and the cells were subjected to a chase reaction in normal medium. At the indicated time points, cells were washed with ice-cold PBS, lysed in 50 μl solubilization buffer [50 mM Tris-HCl buffer (pH 7.5) containing 2% sodium dodecyl sulfate (SDS)] and denatured at 95°C for 5 minutes. The mixture was then diluted 20-fold with immunoprecipitation buffer [50 mM Tris-HCl buffer (pH 7.5) containing 150 mM NaCl, and 1% Triton X-100]. The lysates were clarified by brief centrifugation. The cell lysates (150 μg) were incubated with specific antibodies and protein A-Sepharose at 4°C for 5 hours. Protein A-Sepharose was collected by centrifugation, washed with the washing buffer [50 mM Tris-HCl buffer (pH 7.5) containing 150 mM NaCl, 1% Triton X-100, and 0.2% SDS], and the antigen was extracted with SDS-polyacrylamide gel electrophoresis (PAGE) loading buffer [62.5 mM Tris-HCl buffer (pH 6.8) containing 2% SDS, 5% β-mercaptoethanol and 10% glycerol]. The immunoprecipitates were resolved by SDS-PAGE and analyzed using a BAS 2500 image analyzer (Fuji Photo Film, Tokyo, Japan). Band intensities were quantified using Image Gauge software (Fuji Photo Film).
Immunoprecipitiation for detection polyubiquitylated SCD1
Cells were transfected with pcDNA3.1 or the plasmid harboring SCD1-HA cDNA and grown for 48 hours. They were incubated for 5 hours in the presence or absence of 50 μM MG132, washed with ice-cold PBS, lysed in 100 μl solubilization buffer [50 mM Tris-HCl buffer (pH 7.5) containing 2% SDS and 5 mM N-ethylmaleimide (NEM)], and incubated at 95°C for 10 minutes. The mixture was then diluted 10-fold with immunoprecipitation buffer [50 mM Tris-HCl buffer (pH 7.5) containing 150 mM NaCl, 1% Triton X-100 and 5 mM NEM]. The lysates were clarified by brief centrifugation. The supernatants (800 μg) were incubated with specific antibodies and Protein A-Sepharose at 4°C for 5 hours. Protein A-Sepharose was collected by centrifugation, washed with washing buffer [50 mM Tris-HCl buffer (pH 7.5) containing 150 mM NaCl, 1% Triton X-100, 0.2% SDS and 5 mM NEM], and the antigen was extracted with SDS-PAGE loading buffer. The immunoprecipitates were analyzed by SDS-PAGE and subsequent immunoblotting using specific antibodies.
Cell fractionation and immunoprecipitation
Cells transfected with pcDNA3.1 or the plasmid harboring SCD1-HA cDNA and grown for 48 hours were incubated for 5 hours in the presence or absence of 50 μM MG132. They were washed with PBS and collected by centrifugation at 700 g for 5 minutes. The cells were washed once with homogenization buffer [10 mM Hepes-KOH buffer (pH 7.5) containing 0.25 M sucrose and 5 mM NEM], homogenized in 1 ml homogenization buffer by passing through a 27-gauge needle 20 times, and then centrifuged at 1,000 g for 10 minutes to obtain a post-nuclear supernatant. The post-nuclear supernatant (400 μg) was centrifuged at 100,000 g for 30 minutes to separate the cytosolic and membrane fractions, which were lysed with the lysis buffer [20 mM Hepes-KOH buffer (pH 7.5) containing 150 mM NaCl, 2 mM MgCl2, 1% Triton X-100 and 5 mM NEM] at 4°C for 1 hour. The lysates were clarified by centrifugation and incubated with specific antibodies and protein A-Sepharose at 4°C for 5 hours. The immunoprecipitates were analyzed by SDS-PAGE and subsequent immunoblotting using specific antibodies.
Real-time quantitative PCR
Total RNA was extracted from HEK-293 cells using an RNeasy Mini Kit (Qiagen, Hilden, Germany). First-strand cDNA was synthesized by a SuperScript III First-Strand Synthesis System for RT-PCR (Invitrogen) according to the manufacturer's instructions. Primers for real-time quantitative PCR were designed using Primer Express software (Applied Biosystems, Foster City, CA. USA). The sequence of primer sets for human β-actin were: 5′ TGT CCC CCA ACT TGA GAT GTA TG 3′ (actin forward) and 5′ CCT CAT TTT TAA GGT GTG CAC TTT T 3′ (actin reverse); for human SCD: 5′ GAG GTA CTA CAA ACC TGG CTT GCT G 3′ (SCD forward) and 5′ CCA CTC TTG TAG TTT CCA TCT CCG G 3′ (SCD reverse); for human HMGR: 5′ TAC CAT GTC AGG GGT ACG TC 3′ (HMGR forward) and 5′ CAA GCC TAG AGA CAT AAT CAT C 3′ (HMGR reverse); for human low density lipoprotein receptor (LDLR): 5′ CAA TGT CTC ACC AAG CTC TG 3′ (LDLR forward) and 5′ TCT GTC TCG AGG GGT AGC TG 3′ (LDLR reverse); for human GAPDH: 5′ TGG AGT CCA CTG GCG TCT TC 3′ (GAPDH forward) and 5′ TTC ACA CCC ATG ACC AAC ATG 3′ (GADPH reverse). Real-time quantitative PCR was performed using QuantiTect SYBR Green PCR (Qiagen) and analyzed with an ABI PRISM 7000 Sequence Detection System (Applied Biosystems).
Preparation of delipidated FBS
Fetal bovine serum was delipidated using the method of Hannah et al. (Hannah et al., 2001). Serum (50 ml) was mixed with 40 ml of n-butanol and 60 ml of isopropyl ether at room temperature for 60 minutes, followed by 20 minutes incubation on ice. The mixture was centrifuged at 2,000 g for 5 minutes at room temperature and the aqueous phase was re-extracted with 20 ml of isopropyl ether. The aqueous phase was lyophilized and reconstituted in 20 ml of distilled water, which was then dialyzed against PBS and stored in aliquots at -20°C. Delipidated FBS contained no detectable free fatty acids compared with untreated FBS (787 μM) as determined by `Free fatty acids, Half-microtest' (Roche Diagnostics).
The cells were transfected with N-terminal segments of SCD1 and EGFP fusions and grown for 48 hours. The cells were treated with 10 μg/ml cycloheximide for 0, 2 and 4 hours. They were then washed with PBS and collected by centrifugation at 700 g for 5 minutes. A population of 10,000 cells was then analyzed for EGFP fluorescence intensity using a FACScan flow cytometer (Becton Dickinson, Franklin Lakes, NJ, USA).
We thank Takeshi Watanabe (RIKEN, RCAI, Research Unit for Immune Surveillance) for providing us with mouse TCRα subunit cDNA. We also thank J. L. Degen (Children's Hospital Research Foundation, Cincinnati, OH, USA) for the information on PLG knockout mice. This work was supported by grants from the ministry of Education, Science and Evolutional Science and Technology, andSpecially Promoted Research from the Ministry of Education, Science, and Culture of Japan, and Takeda Science Foundation to K.M.
- Accepted February 21, 2006.
- © The Company of Biologists Limited 2006