J Biol Chem, Vol. 274, Issue 41, 29341-29351, October 8, 1999
Impaired Regulation of 3-Hydroxy-3-methylglutaryl-Coenzyme A
Reductase Degradation in Lovastatin-resistant Cells*
Tommer
Ravid
,
Rachel
Avner§,
Sylvie
Polak-Charcon¶,
Jerry
R.
Faust
, and
Joseph
Roitelman§**
From the Department of Clinical Biochemistry, Sackler
School of Medicine, Tel Aviv University, Ramat Aviv 69978, the
§ Institute of Lipid and Atherosclerosis Research, Sheba
Medical Center, Tel Hashomer 52621, the ¶ Institute of
Pathology, Sheba Medical Center, Tel Hashomer 52621, Israel, and
the Department of Cellular and Molecular Physiology, Tufts
University School of Medicine, Boston, Massachusetts 02111
 |
ABSTRACT |
L-90 cells were selected to grow in the
presence of serum lipoproteins and 90 µM lovastatin, an
inhibitor of 3-hydroxy-3-methylglutaryl-coenzyme A
reductase (HMGR). L-90 cells massively accumulate HMGR, a
result of >10-fold amplification of the gene and 40-fold rise in
mRNA, and also overexpress other enzymes of the mevalonate pathway. Western blot and promoter-luciferase analyses indicate that
transcriptional regulation of sterol-responsive genes by
25-hydroxycholesterol or mevalonate is normal. Yet, none of these genes
is regulated by lipoproteins, a result of severe impairment in the low
density lipoprotein receptor pathway. Moreover, L-90 cells do not
accelerate the degradation of HMGR or transfected HMGal chimera in
response to 25-hydroxycholesterol or mevalonate. This aberrant
phenotype persists when cells are grown without lovastatin for up
to 37 days. The inability to regulate HMGR degradation is not due to its overproduction since in LP-90 cells, which were selected for lovastatin resistance in lipoprotein-deficient serum, HMGR is overexpressed, yet its turnover is regulated normally. Also, the rapid
degradation of transfected 
subunit of T cell receptor is markedly
retarded in L-90 cells. These results show that in addition to gene
amplification and overexpression of cholesterogenic enzymes, statin
resistance can follow loss of regulated HMGR degradation.
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INTRODUCTION |
Mevalonate (MVA)1 is the
first committed precursor for biosynthesis of cholesterol and a
variety of essential nonsterol isoprenoids such as ubiquinone,
dolichol, isopentenyl-modified tRNAs, side chain of heme A, and the
farnesyl and geranylgeranyl moieties of many cellular proteins (1-4).
Cells must constantly synthesize MVA yet avoid accumulation of these
products to toxic levels. This problem is especially acute in mammalian
cells since cholesterol, the bulk end product of the MVA pathway, can
be obtained also by receptor-mediated endocytosis of low density
lipoproteins (LDL). Classical experiments by Brown and Goldstein (5)
and their co-workers have demonstrated that the increase in
intracellular levels of LDL-derived cholesterol triggers a feedback
response that down-regulates the expression of the LDL receptor (LDLR) and represses endogenous cholesterol synthesis. Conversely,
depletion of cholesterol leads to increased surface expression of LDLR
and enhanced rate of cholesterol synthesis (6).
The major regulatory step in the MVA pathway is the conversion of
HMG-CoA to MVA by the enzyme HMGR. Early experiments have shown that
cultured mammalian cells maintain low rate of endogenous MVA production
even in the presence of saturating amounts of sterols. HMGR activity
can be fully repressed only if the synthesis of MVA-derived nonsterol
is allowed, suggesting that HMGR activity (hence MVA synthesis) is
controlled through multivalent feedback regulation by sterol and
nonsterol MVA-derived metabolite(s) (7). These studies were greatly
advanced by the discovery of statins, potent HMGR inhibitors, such as
compactin and lovastatin (8, 9). When naive cells are exposed to
statins in their growth medium, they respond by drastically increasing
the amounts of HMGR to overcome the acute depletion in endogenously
synthesized sterol and nonsterol compounds (6, 10). This metabolic
control of HMGR is achieved through changes in the rate of HMGR gene
transcription (11-13), differential translational efficiency, and
stability of mRNA (14-18) and post-translationally by altering the
stability of the reductase protein (15, 19-22). Thus, when cells are
starved for sterols/MVA, the enzyme is stabilized and its half-life is prolonged, whereas abundance of sterols/MVA leads to its rapid degradation.
The mechanisms underlying the regulated degradation of HMGR are not yet
fully understood. Unlike other early enzymes in the MVA pathway, HMGR
is a resident glycoprotein of the endoplasmic reticulum (ER) with its
active site, contained within the C-terminal portion of the protein,
facing the cytosol (23). A highly conserved N-terminal hydrophobic
domain, which spans the membrane eight times, anchors the enzyme in the
ER (24). Deletion analyses have demonstrated that the membrane domain
is dispensable for catalytic activity, yet it is essential for the
regulated turnover of the enzyme (22, 25). Moreover, when fused to the
N termini of heterologous proteins, e.g. 
-galactosidase
of E. coli, the membrane domain confers ER localization and
sterol/MVA-regulated degradation onto this chimeric HMGal protein
(26-28). Although the precise structural determinants that alter
enzyme stability have not been fully elucidated (27, 29), the membrane
domain is believed to function as a sensor that monitors the levels of sterols and/or MVA-derived nonsterols in the ER membrane (29, 30).
Other key questions relate to the intracellular site(s) of degradation,
the role of potential trans-acting factor(s) that recognize
HMGR and tag it for rapid elimination when sterols/MVA are abundant,
and the protease(s) that actually destroys the HMGR protein. Current
evidence indicates that HMGR is degraded in the ER (31) in a process
that involves a yet unidentified short-lived protein(s) (31, 32).
Furthermore, HMGR degradation is severely inhibited by ALLN (33, 34)
and lactacystin (35) suggesting the participation of the 26 S
proteasome. Indeed, a mutation in the 26 S proteasome has been shown
to affect HMGR degradation in yeast (36). Although conjugation of
multi-ubiquitin chains to HMGR was demonstrated in yeast (37), it
remains to be established whether the ubiquitin pathway is involved
also in the degradation of mammalian HMGR.
To gain further insight into the regulated degradation of HMGR, we
sought to isolate cells that are defective in this process. In this
work, we describe a CHO-derived cell line, designated L-90, that was
selected for growth in the presence of serum lipoproteins and
increasing concentrations of lovastatin. Our results demonstrate that
L-90 cells survived the lovastatin selection by increasing the levels
of HMGR enzyme, in part by losing their ability to degrade HMGR in a
regulated manner. Analysis of L-90 cells should provide information on
cellular factors that are involved in HMGR turnover.
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EXPERIMENTAL PROCEDURES |
Materials--
DNA restriction and modification enzymes were
purchased from New England Biolabs (Beverly, MA) and MBI-Fermentas
(Vilnius, Lithuania). HighPrime random priming DNA labeling kit was
obtained from Roche Molecular Biochemicals. LipofectAMINETM
and Geneticin® (G418 sulfate) were obtained from Life Technologies, Inc. 25-Hydroxycholesterol was from Steraloids (Wilton, NH).
[3H]Acetate (81.2 mCi/mmol),
Expre35S35S protein labeling mix (>1000
Ci/mmol), and GeneScreen Plus membrane were from NEN Life Science
Products. [-32P]CTP and [-32P]UTP
(~800 Ci/mmol) were obtained from Amersham Pharmacia Biotech. Methionine- and cysteine-free MEM was purchased from ICN
Pharmaceuticals (Costa Mesa, CA). MEM and fetal calf serum were from
Biological Industries (Beit Haemek, Israel). Unless otherwise noted,
all other reagents were obtained from Sigma. Oligonucleotides
were synthesized by Biotechnology General, Inc. (Rehovot,
Israel). Mevalonolactone (Fluka, Buchs, Switzerland) was converted to
the sodium salt and made in 0.5 M K-HEPES, pH 7.2. Lovastatin, kindly provided by Merck, was converted to the sodium salt
and stored as a 20 mM stock solution in 0.5 M
K-HEPES, pH 7.2. Compactin was a generous gift from Dr. Robert Simoni,
Stanford University. Lipoprotein-deficient fetal calf serum (LPDS;
d 
1.25) and human LDL (d 1.019-1.063) were
prepared by ultracentrifugation, as described (38).
125I-LDL was prepared as described previously (38).
Plasmids--
pHMGS-Luc was prepared by inserting the promoter
region of HMG-CoA synthase (HMGS) gene, contained within a 596-base
pair HindIII fragment of pSynCAT-1 (39) (provided by G. Gil,
Medical College of Virginia), into pGL2-basic vector (Promega, Madison, WI). pHMGR-Luc was constructed by removing the chloramphenicol acetyltransferase sequence of pRedCAT-3 (provided by T. Osborne, University of California, Irvine) with SalI (12) and
replacing it with the 2.7-kb XhoI-SalI fragment
of pGL2-basic which contains luciferase and SV40 intron and
polyadenylation signal sequences. The following plasmids were used
without further modifications: reporter construct for rat
farnesyl-pyrophosphate synthase (FPPS) gene promoter pFPPS-Luc
(pFPPS-0.319 (40); provided by P. Edwards, UCLA), reporter construct
for human squalene synthase (SQS) gene promoter pSQS-Luc (pHSS1kb-Luc
(41); provided by I. Shechter, Uniformed Services University of the
Health Sciences), reporter construct for rat fatty-acid synthetase
(FAS) gene promoter pFAS-Luc (FAS-150, plasmid B (42); provided by T. Osborne), and expression vector of subunit of T cell receptor,
pTCR-Neo ((43) provided by R. Kopito, Stanford University).
pHMGal-Neo was prepared by inserting the 4.7-kb HMGal EcoRI
fragment of pSR-HMGal ((27) provided by R. Simoni, Stanford
University) into the EcoRI site of pcDNA3 (Invitrogen;
San Diego, CA). pGal-neo was constructed by ligating the 3.47-kb
-galactosidase fragment from pNASS (CLONTECH;
Palo Alto, CA) into the NotI site of pcDNA3. pRPS17 (17)
was provided by D. Peffley (Finch University of Health Sciences/the
Chicago Medical School), and full-length cDNA of human LDLR (44)
was provided by E. Leitersdorf (Hebrew University/Haddassah Medical School).
Antibodies--
Antibodies against HMGS, the cytoplasmic domain
of HMGR, and FPPS were kindly provided by P. Edwards. An anti-rat SQS
antibody was a gift from I. Shechter. Antibodies against the membrane
domain of HMGR were described previously (24). The
anti--galactosidase antibody was either a mouse monoclonal (Promega)
or a rabbit polyclonal antibody (Cappel Organon Teknika, West Chester,
PA). Protein A bacterial adsorbent and protein A-agarose were obtained
from Sigma and Bio-Rad, respectively.
Cells and Media--
CHO cells were grown at 37 °C in
5% CO2 atmosphere in a humidified incubator in FCS
("F") medium (MEM supplemented with 5% FCS, 2 mM
glutamine, 100U/ml penicillin, 100 µg/ml streptomycin). L-90 cells
were selected in this medium with increasing concentrations of
lovastatin (5, 7, 10, 15, 30, 45, 60, 90 µM) over a
4-month period. L-90 cells are continuously maintained in FCS medium
plus 90 µM lovastatin. To obtain LP-90 cells, CHO cells
were first adapted to grow for 2 months in "L" medium (MEM
supplemented with 5% LPDS, 2 mM glutamine, 100 units/ml
penicillin, 100 µg/ml streptomycin) and then selected, in the same
stepwise fashion, for lovastatin resistance in LPDS medium. LP-90 cells
are continuously maintained in LPDS medium plus 90 µM
lovastatin. In experiments where transfected cells were used, all media
also contained 250 µg/ml geneticin. For CHO cells, LPDS medium was
supplemented with 2 µM compactin and 100 µM
MVA (32). For L-90 and LP-90 cells, LPDS medium contained 90 µM lovastatin without MVA. Where indicated, the
respective LPDS medium was also supplemented with 2 µg/ml
25-hydroxycholesterol and 20 µg/ml cholesterol from a 200× stock
solution in absolute ethanol (LPDS + ST; "S") or with 20 mM MVA from a 2 M stock solution (LPDS + MVA;
"M").
DNA Transfections--
CHO and L-90 cells were transfected by
LipofectAMINE with 2 µg of each of the above-described promoter-Luc
plasmids together with 0.2 µg of Gal-Neo, according to the
manufacturer's instructions. Stable transfectants were selected in a
medium containing 750 µg/ml geneticin, and 300-500 resistant
colonies were pooled and expanded. The cells were maintained
continuously in 250 µg/ml of the antibiotic. Lines expressing HMGal
(pHMGal-Neo) or soluble -galactosidase (pGal-Neo) were
transfected with 2 µg of the indicated plasmids. Clones expressing
highest -galactosidase activity were isolated by limiting dilution.
Slot Hybridization of Genomic DNA--
Genomic DNA was adsorbed
onto GeneScreen Plus membrane using a slot blot filtration manifold.
Following UV cross-linking and prehybridization, the membrane was
hybridized overnight at 60 °C with 3 × 106 cpm/ml
of an ~1.4-kb 32P-labeled
EcoRI-PstI cDNA fragment of hamster HMGR
membrane-spanning region in 6× SSC, 5× Denhardt's solution, 0.5%
SDS, 100 µg/ml sheared denatured salmon sperm DNA. The membrane was
washed in 2× SSC, 0.1% SDS (30 min; 42 °C), 1× SSC, 0.1% SDS (60 min; 42 °C), 0.1× SSC, 0.1% SDS (40 min; 42 °C), and an
additional 30 min wash at 50 °C in the same solution. The membrane
was dried and exposed to x-ray film at 
80 °C with intensifying
screens. The radioactive probe was stripped (45), and the membrane was successively re-probed with 32P-labeled human LDLR and rat
glyceraldehyde-3-phosphate dehydrogenase cDNAs.
RNase Protection Assay--
To prepare an antisense RNA probe
for HMGR, we amplified a 206-base pair DNA fragment from the
cytoplasmic domain of Chinese hamster HMGR using pDGS6 plasmid as the
template and oligonucleotides 5'-CGGAATTCGTGCCTGGATGGTAAAGAG-3' ((46)
positions 1629-1648, including an EcoRI site), and
5'-CCGGTACCGGGTGTTTCAAGCCAGGCC-3' ((46) positions 1833-1814 plus a
KpnI site) as the forward and reverse primers, respectively.
The PCR product was digested with EcoRI and KpnI and ligated into BlueScript. The plasmid was linearized with
NotI and transcribed with T7 RNA polymerase in the presence
of [-32P]UTP using MaxiScriptTM kit
obtained from Ambion (Austin, TX). Similarly, 32P-labeled
antisense RNA probe for the S17 ribosomal protein was transcribed using
the AvaII-digested pRPS17 plasmid (17). Total cellular RNA
was extracted with RNAzol B reagent (Biotecx Laboratories, Houston, TX)
and hybridized overnight at 50 °C with gel-purified antisense
probes, according to the instructions provided by the manufacturer
(RPAIITM kit; Ambion). Following RNase digestion, the
protected fragments were analyzed by 8 M urea/PAGE and
autoradiography. After exposure, the protected bands were cut out of
the gel and quantified by liquid scintillation.
Metabolic Labeling, Immunoprecipitation, and
Immunoblotting--
On Day 0, CHO (8 × 105) or L-90
(1.2 × 106) cells were plated in FCS medium in 60-mm
dishes. On Day 1, the cells were refed with LPDS medium containing 2 µM compactin plus 100 µM MVA (CHO) or 90 µM lovastatin (L-90). On Day 2, the medium was aspirated, and the cells were starved for 1 h in 1 ml of starvation medium (methionine- and cysteine-free MEM supplemented with 5% LPDS and 2 µM compactin, 100 µM MVA (CHO) or 90 µM lovastatin (L-90 and LP-90)). The cells were
pulse-labeled for 30 min in 0.5 ml of starvation medium containing
100-150 µCi of Expre35S35S protein labeling
mix and chased in LPDS medium supplemented with 2 mM
methionine. Unless otherwise noted (e.g. Fig. 11), L-90 and
LP-90 cells were chased in LPDS medium containing 90 µM
lovastatin. CHO cells were chased in LPDS medium containing 2 µM compactin plus 100 µM MVA. Where
indicated, sterols (2 µg/ml 25-hydroxycholesterol plus 20 µg/ml
cholesterol) or MVA (20 mM) were added to the chase medium.
The cells were lysed, processed for immunoprecipitation with the
indicated antibodies, and samples were resolved by 5-15% gradient
SDS-PAGE, as described previously (32). Because L-90 and LP-90 cells
contain high levels of enzyme, only 10-13% of lysate volume was used
for immunoprecipitating HMGR from these samples. This allowed
comparable studies with CHO cells and ensured that HMGR was
precipitated quantitatively (data not shown). For immunoblot analyses,
cells were plated as described above. Twenty four h prior to analysis,
the cells received fresh FCS, LPDS, LPDS + ST, or LPDS + MVA media.
After lysis and estimation of protein content, samples containing equal
amounts of protein were separated by SDS-PAGE and transferred to
Optitran BAS-83 reinforced nitrocellulose membranes (Schleicher & Schuell). The membranes were probed with the appropriate primary
antibodies, followed by horseradish peroxidase-conjugated secondary
antibodies and enhanced chemiluminescence reaction (32).
Electron Microscopy--
Cells were fixed in 2.5%
glutaraldehyde in 0.1 M cacodylate buffer (pH 7.2) for
1 h at room temperature. Fixed cells were scraped off the dish and
post-fixed in 2% OsO4 in cacodylate buffer for 2 h.
After dehydration in graded ethanol solutions, the cells were embedded
in Epon. Ultra-thin sections were contrasted with uranyl acetate and
lead citrate and examined with a Jeol-1200 EX II electron microscope.
Degradation of 125I-LDL--
On Day 0, 1 × 105 CHO cells or 1.5 × 105
L-90 cells were plated in a 24-well dish in FCS medium. On
Day 1, the cells were refed with LPDS medium containing 2 µM compactin plus 100 µM MVA (CHO) or 90 µM lovastatin (L-90), in the absence or presence of 2 µg/ml 25-hydroxycholesterol and 20 µg/ml cholesterol. On Day 2, quadruplicate wells received 125I-LDL at 10 µg/ml
(250-650 cpm/ng) in the absence or presence of 400 µg/ml unlabeled
LDL. Following incubation at 37 °C for 5 h, the amount of
trichloroacetic acid-soluble 125I radioactivity was
determined, as described (38).
Measurements of Lipid Synthesis and Content--
CHO and L-90
cells were set on Day 0 in their respective FCS medium. On Day 1, the
cells were washed extensively to remove lovastatin, and LPDS medium
(without lovastatin) was added to both cell types. On Day 2, all dishes
were washed, and triplicate plates received FCS medium with or without
90 µM lovastatin, as indicated in Table I, and the cells
were pulse-labeled with 50 µCi of [3H]acetate. Cells
were harvested, and 3H-labeled lipids were separated by
thin layer chromatography and quantified, as described (47). Cellular
cholesterol content was quantified by gas-liquid chromatography.
Enzymatic and Other Assays--
On Day 0, 1 × 105 CHO cells or 1.5 × 105
L-90 cells were plated in a 24-well dish in their
respective FCS media. On Day 1, replicate wells were washed with PBS,
and the cells were refed either with fresh FCS, LPDS, LPDS + ST, or
LPDS + MVA media (see above). On Day 2, the cells were washed with PBS
and processed for measurements of luciferase activity, according to the
manufacturer's instructions (Promega). The hydrolysis of
o-nitrophenyl -D-galactopyranoside by
-galactosidase was measured in digitonin-permeabilized cells, as
described previously (26). Protein was measured either according to
Lowry et al. (48) or with the protein microassay reagent (Bio-Rad). Luciferase-specific activity was calculated as light units/mg cell protein. -Galactosidase-specific activity was
calculated as A420/mg cell protein/h. In both
cases, the results are expressed as percent of the specific activity in
LPDS-treated cells ("100%") in the same experiment. X-ray films
were scanned and quantified using a Model GS-690 imaging densitometer
and the MultiAnalyst software package from Bio-Rad.
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RESULTS |
CHO cells were selected for growth in increasing concentrations of
lovastatin in the presence of serum lipoproteins (FCS
medium). After each incremental increase in lovastatin concentration
>90% of the cells died. Those that survived 90 µM
lovastatin are designated L-90 cells and have been maintained at this
concentration since June 1994. Fig. 1
shows that L-90 cells are highly resistant to lovastatin
(IC50 
240 µM), unlike naive CHO cells
whose growth is sensitive to low concentrations of the drug
(IC50 8 µM). L-90 cells remain highly
resistant to lovastatin (IC50 180-200 µM)
even after culture in the absence of the drug for over 100 days (data
not shown).
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Fig. 1.
Effect of lovastatin on growth of CHO and
L-90 cells. On Day 0, 5 × 105 CHO ( ) or
7 × 105 L-90 ( ) cells were plated in
24-well dishes in FCS medium without lovastatin. On Day 1, triplicate
wells received fresh medium plus lovastatin at the indicated
concentrations. The cells were refed on Day 3. On Day 5, the wells were
washed with PBS; the cells were dissolved in 0.5 ml of 1 N
NaOH, and protein content was measured. The results, given as "%"
of protein content in wells receiving no lovastatin (100% = 35-60
µg/well), are the average of two experiments.
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L-90 cells are resistant to lovastatin, at least in part, due to
elevated activity of the cholesterogenic pathway, as shown in Table
I. When lipid synthesis and cellular
content were measured, the rate of [3H]acetate
incorporation into sterols was >120-fold higher in L-90 cells than in
CHO cells. Moreover, whereas addition of 90 µM lovastatin totally abolished sterol synthesis in CHO cells, L-90 cells
incorporated acetate into sterols in amounts comparable to sterol
synthesis in uninhibited CHO cells (Table I). These results are
consistent with the notion that L-90 cells have elevated activity of
HMGR of which >85% can be blocked by lovastatin. Interestingly, L-90 cells synthesize ~5-fold more fatty acids than CHO cells (Table I).
The basis for this finding is unclear (see "Discussion"), but it
was also noted in other statin-resistant cell lines (see Table II in
Ref. 10). Compared with CHO cells, L-90 cells also contain reduced
(70-80%) content of total cholesterol that was mostly in the free
form (Table I).
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Table I
Lipid metabolism in CHO and L-90 cells
Cellular lipid synthesis and content were determined as described under
"Experimental Procedures."
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To examine whether lovastatin resistance and the increased
cholesterogenic activity in L-90 cells were afforded by elevated amounts of HMGR, the steady state levels of the enzyme were determined. Equal amounts of protein from CHO and L-90 cells were immunoblotted with anti-HMGR antibodies and, as shown in Fig.
2A, L-90 cells indeed
accumulated massive amounts of HMGR. Densitometric quantification demonstrated that L-90 cells contained ~50-fold more HMGR than CHO
cells when both lines were incubated in LPDS medium (Fig. 2A) and over 1000-fold more enzyme when the cells were
maintained in medium containing lipoproteins (FCS medium; data not
shown). Furthermore, electron microscopy (Fig. 2B) revealed
the presence of extensive proliferation of smooth membranes and
formation of crystalloid ER (49) to accommodate the large amounts of
HMGR in these cells. The increased levels of HMGR in L-90 cells
were, in part, the result of HMGR gene amplification, as determined by
slot blot hybridization of genomic DNA with a 32P-labeled
cDNA probe for HMGR (Fig. 2C). We estimated that
L-90 cells contain 10-30-fold more copies of the gene for HMGR
than CHO cells (Fig. 2C). Neither the gene for LDLR (Fig.
2C) nor for HMGS (data not shown) was amplified.
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Fig. 2.
HMGR is overexpressed in L-90 cells.
A, immunoblot analysis. CHO or L-90 cells were
incubated for 24 h in LPDS medium that was supplemented with 2 µM compactin and 100 µM MVA (CHO cells) or
with 90 µM lovastatin (no MVA added; L-90 cells).
The cells were lysed, as described (32), and 50 µg of postnuclear
supernatant proteins were analyzed by immunoblotting, as described
under "Experimental Procedures." B, crystalloid ER is
present in L-90 cells. L-90 cells maintained in FCS medium
plus 90 µM lovastatin were processed for electron
microscopy, as described under "Experimental Procedures." Note the
packed extended tubules of smooth membrane comprising the crystalloid
ER. Inset, cross-section of crystalloid ER. No such
structures were observed in CHO cells (data not shown).
Bar = 1 µm. C, the gene for HMGR, but not
for LDLR, is amplified in L-90 cells. Genomic DNA was isolated
from CHO and L-90 cells, and the indicated amounts were adsorbed
onto a GeneScreen Plus membrane using a slot blot filtration manifold.
The membrane was successively hybridized with 32P-labeled
cDNA probes for HMGR, LDLR, and
glyceraldehyde-3-phosphate dehydrogenase, as
described under "Experimental Procedures."
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We also determined whether enzymes of cholesterol biosynthesis other
than HMGR were elevated and properly regulated in L-90 cells
(Fig. 3). Cells were incubated for
24 h with (FCS) or without (LPDS) serum lipoproteins or in LPDS
medium supplemented with sterols or MVA, and equal amounts of cell
lysates were analyzed by immunoblotting using antibodies specific to
HMGS, HMGR, FPPS, and SQS. Clearly, relative to CHO cells in FCS, the
levels of all examined proteins were highly elevated in L-90
cells (Fig. 3, e.g. compare lanes 1 and
5). However, unlike CHO cells, the amounts of these enzymes
did not increase further when L-90 cells were switched to LPDS
medium (Fig. 3, compare lanes 1 and 2 with lanes 5 and 6). Moreover, their steady state
levels in L-90 cells decreased only moderately when sterols or
MVA were also included in the LPDS medium (Fig. 3, compare lanes
3 and 4 with lanes 7 and 8). This
is in a sharp contrast to the response of CHO cells where such
additions prevented the LPDS-induced up-regulation of the enzymes.
Quantitatively, if a value of 100% was assigned to the level of a
protein in LPDS-treated cells (Fig. 3, lanes 2 and
6), then addition of sterols to CHO and L-90 cells,
respectively (lanes 3 and 7), reduced the levels
of HMGS to 5 and 60%; HMGR declined to 10 and 50% (see also Fig.
9A), and FPPS decreased to 15 and 80%.
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Fig. 3.
Steady state levels and regulation of enzymes
in the MVA pathway. On Day 0, CHO or L-90 cells were plated
in FCS medium without or with 90 µM lovastatin,
respectively. On Day 2, the cells were fed with the same fresh medium
(designated F, lanes 1 and 5) or
switched to LPDS medium (designated L, lanes 2 and 6), LPDS plus sterols (2 µg/ml 25-hydroxycholesterol + 20 µg/ml cholesterol, designated S, lanes 3 and
7), or LPDS plus MVA (20 mM, designated
M, lanes 4 and 8). The LPDS media for
CHO cells contained 2 µM compactin and 100 µM MVA. The media for L-90 cells included 90 µM lovastatin. After additional 24 h incubation, the
cells were lysed, and nuclei-free extracts were prepared. Seventy-five
µg of protein from each treatment were analyzed by 5-15% SDS-PAGE
and immunoblotting, using antibodies against HMGS, FPPS, HMGR
cytoplasmic domain, or SQS, as described in Fig. 2.
Arrowheads indicate the relevant antigens.
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Since cholesterol biosynthetic enzymes are known to be coordinately
regulated at the transcriptional level, we examined whether the blunted
response of these enzymes in L-90 cells might be the consequence
of defect(s) in common transcriptional regulatory factor(s), such as
SREBPs (50, 51), or merely a manifestation of their relatively long and
unregulated half-life (except for HMGR, see below). To this end, we
created a series of stably transfected CHO and L-90 cell lines
that expressed firefly luciferase under the control of promoters of
several sterol-regulated genes. These included HMGS, HMGR, FPPS, SQS,
LDLR, and FAS. Luciferase-specific activities were measured in cells
that were incubated for 24 h in FCS, LPDS, LPDS plus sterols, or
LPDS plus MVA, and the results were normalized to LPDS-treated cells
(Fig. 4). When CHO cells were switched
from FCS to LPDS medium, luciferase activity was induced about 4-fold
when luciferase was expressed from the promoters of HMGS, FPPS, or SQS
genes. When transcribed from the HMGR or LDLR promoters, the removal of
lipoproteins resulted in approximately 2-fold increase in luciferase
activity and only in a 40% rise when the enzyme was driven by the FAS
promoter (Fig. 4). As expected, addition of sterols completely
abolished this up-regulation, resulting in 4-5-fold differences in
luciferase activities between the LPDS-induced and sterol-repressed
states. This magnitude was comparable to the degree of repression
reported in previous studies (12, 40-42). Addition of excess MVA only
partially prevented the induction of luciferase upon transfer to LPDS
medium (approximately 2-fold difference; Fig. 4). Remarkably, in
L-90 cells all tested reporter constructs were completely
refractory to the up-regulation induced by the omission of serum
lipoproteins, but the regulation by sterols added in ethanol or by MVA
appeared normal (Fig. 4). Yet, we noted some small but consistent
differences between CHO and L-90 cells with respect to the degree
of luciferase repression by sterols (e.g. HMGS, SQS, LDLR;
Fig. 4). These differences may be related to the genomic integration
site(s) and/or subtleties in sterol accessibility to the regulatory
machinery of these promoters. In contrast to the
lipoprotein-dependent response, we concluded that the
non-lipoprotein-mediated transcriptional regulation of transfected
genes was intact in L-90 cells. This indicates that there was
neither a defect in general transcriptional regulation of
sterol-sensitive genes nor any hindrance in the accessibility of
exogenously added sterols or MVA to the intracellular pool(s) that are
involved in this regulation. Hence, the attenuated response to sterols
or MVA in the steady levels of endogenous HMGS, FPPS, and SQS enzymes,
observed in Fig. 3, may be attributed to their relatively slow and
unregulated turnover rates (52). Moreover, under such conditions HMGR
appeared as a short-lived protein in CHO cells but not in L-90
cells (see below).
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Fig. 4.
Promoter activities and regulation of
sterol-sensitive genes. CHO (open bars) and L-90
(closed bars) cells were stably transfected with the
indicated promoter-luciferase DNA constructs and 300-500 resistant
colonies of each were pooled. Cells were initially plated in FCS medium
and then switched to the indicated media for additional 24 h
incubation, as described in Fig. 3. The LPDS media for CHO cells
contained 2 µM compactin and 100 µM MVA,
and all media for L-90 cells included 90 µM
lovastatin. Luciferase-specific activities (light units/mg cell
protein) were determined in triplicate wells and normalized relative to
cells incubated in LPDS (100%). The results are mean ± S.E. of
five to nine independent experiments.
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The aberrant regulation of L-90 cells by serum lipoproteins could
result from defects in receptor-mediated uptake of LDL. In
Table II we quantified the output of the
LDLR pathway by measuring the rate of 125I-LDL degradation
at 37 °C. The results clearly show that, when cultured under induced
conditions, L-90 cells degraded 125I-LDL at only 35%
the rate measured in CHO cells. Moreover, whereas exogenous sterols
down-regulated LDL degradation by >80% in CHO cells, the lower
activity of the LDLR pathway in L-90 cells appeared resistant to
suppression by saturating concentrations of sterols and decreased by
less than 40% (Table II). Thus, the inability of L-90 cells to
activate the transcription of sterol-responsive genes upon removal of
serum lipoproteins stemmed from the attenuated activity of their LDLR
pathway.
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Table II
Impaired 125I-LDL degradation in L-90 cells
Degradation of 125I-LDL at 37 °C was determined as described
under "Experimental Procedures." The results, corrected for
nonspecific LDL degradation, are the mean ± S.E. of 10 separate
experiments.
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The decreased steady state levels of HMGR in sterols- or MVA-treated
L-90 cells could have resulted from slower protein translation. Therefore, we measured the rate of HMGR synthesis in cells maintained under different metabolic conditions (FCS, LPDS, LPDS plus sterols, or
LPDS plus MVA) (Fig. 5). The
incorporation of radiolabeled amino acids was linear for up to 40 min
(data not shown), and HMGR was immunoprecipitated from equal amounts of
labeled trichloroacetic acid-precipitable material. The amounts of
labeled HMGR rose ~4-fold when CHO cells were transferred from FCS-
to LPDS-containing medium, and this increase was prevented when cells
also received sterols or MVA (Fig. 5, lanes 1-4). When
incubated in LPDS medium, the rate of HMGR synthesis was nearly 30-fold
higher in L-90 cells compared with CHO cells (Fig. 5, compare
lanes 2 and 6). This was expected in light of the
multiple copies of HMGR gene in these cells (see Fig. 2C).
However, there was again no significant change (<20% increase) in
HMGR synthesis when L-90 cells were switched from FCS to LPDS
(compare lanes 5 and 6). Nevertheless, in the presence of MVA or sterols there was an ~1.5- and ~2-fold decrease in HMGR synthetic rate, respectively (Fig. 5, lanes 7 and
8).
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Fig. 5.
Effect of lipoproteins, MVA, or sterols on
HMGR synthesis rate. On Day 0, CHO or L-90 cells were
plated in 60-mm dishes in FCS medium without or with 90 µM lovastatin, respectively. On Day 2, the cells were fed
with either FCS (F), LPDS (L), LPDS + MVA
(M), or LPDS + sterols (S) media, as specified in
Figs. 3 and 4. The LPDS media for CHO cells contained 2 µM compactin and 100 µM MVA, and all media
for L-90 cells included 90 µM lovastatin. After
24 h incubation, the cells were switched for 1 h to
methionine- and cysteine-deficient media, containing the same
supplements, and then labeled for 30 min with 100 µCi of
Expre35S35S protein labeling mix. HMGR was
immunoprecipitated from all lysates containing 5.5 × 107 dpm of trichloroacetic acid-insoluble 35S
counts. The gel was exposed to the film at 80 °C for 5 h
(CHO) or 1.5 h (L-90).
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The rates of HMGR synthesis in CHO and L-90 cells directly
correlated with the steady state levels of reductase mRNA, as
determined by RNase protection assays (Fig.
6). 32P-Labeled antisense RNA
probes for Chinese hamster HMGR and S17 ribosomal protein were
hybridized with total RNA from CHO and L-90 cells grown under
different metabolic conditions, digested with RNase, and analyzed by
PAGE. As shown, L-90 cells contained ~40-fold more HMGR
mRNA compared with CHO cells (Fig. 6B), a result consistent with multiple copies of HMGR gene in L-90 cells. Fig. 6 also demonstrates that in both cell lines HMGR mRNA levels
declined to 40-50 and ~65% upon addition of sterols and MVA,
respectively, when compared with cells maintained in LPDS medium (Fig.
6A, lanes 5 and 6 and 10 and
11). However, contrary to CHO cells where FCS caused >50%
decline, in L-90 cells HMGR mRNA levels were hardly affected
by serum lipoproteins (Fig. 6A, compare lanes 3 and 4 with lanes 8 and 9, respectively). These results were in full accordance with the data
shown in Fig. 4 (HMGR panel) and demonstrated that the transcription of
the multiple copies of the endogenous HMGR gene is regulated normally
in L-90 cells.
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Fig. 6.
Transcriptional regulation of HMGR gene.
CHO and L-90 cells were set for the experiment, as described in
Fig. 3. Total RNA was extracted, and 5 µg were hybridized, either
singly or simultaneously, with 2 × 105 cpm of
32P-labeled antisense RNA probes for HMGR and S17 ribosomal
protein, as described under "Experimental Procedures." Protected
fragments were analyzed on 8% polyacrylamide, 8 M urea
gel. The migration of size markers, free probes, and protected
fragments is indicated. A, lanes 1 and
13, free S17 and HMGR probes, respectively; lanes
2 and 12, singly protected fragments of S17 protein and
HMGR, (probes for HMGR or S17 protein, respectively, were not included
in these samples); lanes 3-6 and 8-11, both
probes were simultaneously hybridized to RNA from CHO and L-90
cells incubated in FCS (F), LPDS (L), LPDS + MVA
(M), and LPDS + sterols (S) media, respectively,
as specified in Fig. 3. In lane 7, both probes were
hybridized to yeast tRNA. To assess properly the degree of regulation,
the HMGR probe used for hybridizing L-90 RNA was synthesized in
the presence of a 30-fold molar excess of unlabeled UTP. Following
autoradiography, the protected bands were cut out of the gel and
counted. Because HMGR probes were of different specific radioactivity,
percent regulation was determined separately for each cell type by
calculating the ratio of cpm in HMGR band/cpm in S17 band and
normalizing it to the value obtained in LPDS-treated cells.
B, RNase protection assay was performed on RNA
isolated from LPDS-treated CHO (lane 1) and L-90
(lane 2) cells, as described above, with the exception that
the HMGR probe was of the same specific radioactivity. "Fold"
difference was calculated from the ratio of cpm in HMGR band/cpm in S17
band for each sample, setting the value in CHO cells as 1. In both
panels, the quantitative results are the mean of seven experiments done
on five separate RNA preparations.
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A major post-translational mode for controlling HMGR levels is achieved
through regulated degradation of the enzyme. Therefore, we next
examined whether the turnover of HMGR was regulated in the same manner
in CHO and L-90 cells. Cells, maintained in LPDS medium, were
pulsed with [35S]methionine/cysteine and chased in the
absence or presence of excess MVA or sterols, and at various time
points HMGR was immunoprecipitated (Fig.
7 and Table
III). In LPDS medium HMGR turned over
with a half-life of 12.5 h in CHO cells. Upon addition of MVA or
sterols the rate of reductase degradation was accelerated more than
4-fold, decreasing its t1/2 to <3 h (Fig. 7B and Table III). Strikingly, neither MVA nor sterols had
any significant effect on the rate of HMGR degradation in L-90
cells (Fig. 7, Table III L-90 "on"). In multiple experiments the
half-life of HMGR in L-90 cells averaged 12 h, with only a
marginal (15-30%) effect by MVA (t1/2 = 11.3 h) or sterols (t1/2 = 11 h) (Fig. 7B
and Table III L-90 "on"). Importantly, we have verified that
the amount of antibody used was sufficient for the quantitative and
complete precipitation of HMGR from L-90 cell lysates (see
"Experimental Procedures"). Moreover, the presence of lovastatin in
the chase medium did not alter the immunogenic properties of the enzyme (data not shown).
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Fig. 7.
Degradation HMGR is not regulated in L-90
cells. A, CHO or L-90 cells were plated in FCS or
FCS plus 90 µM lovastatin medium, respectively. On Day 1, the cells were switched to LPDS medium supplemented with 2 µM compactin, 100 µM MVA (CHO cells) or 90 µM lovastatin (L-90 cells). On Day 3, the cells
were starved for 1 h in a methionine- and cysteine-deficient
medium. Lovastatin was present in the starvation medium of L-90
cells. The cells were pulse-labeled for 30 min with 110 µCi (370 mCi/ml) of Expre35S35S protein-labeling
reagent. The cells were chase in above LPDS medium that was
supplemented with 2 mM methionine and either MVA (20 mM) or sterols (2 µg/ml 25-hydroxycholesterol plus 20 µg/ml cholesterol). At the indicated time points cells were
solubilized, and HMGR was immunoprecipitated using anti-HMGR membrane
domain antibodies followed by SDS-PAGE and fluorography. The gel
containing CHO samples was exposed for 8 h. Samples of L-90
cells were exposed for 1.5 h. B, densitometric analysis
of gels. The results are the mean of five independent representative
experiments. Half-life values are also given in Table III. , no
addition; , MVA;  , sterols.
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Table III
Effect of lovastatin on HMGR and HMGal half-life in L-90 cells
Degradation rates of HMGR and HMGal were determined in CHO-HMGal cells,
in L-90-HMGal cells that were continuously maintained in 90 µM lovastatin (L-90 "on"), and in L-90 cells that
were grown without lovastatin for 2, 5, 9, 16, 23, and 37 days (L-90
"off"). Pulse-chase experiments were performed as described
under "Experimental Procedures" and in Figs. 7 and 11. L-90
"off" cells were assayed without lovastatin. The half-lives of HMGR
and HMGal in L-90 "off" cells were very similar between 2 and 37 days (see text) and thus were averaged. In each experiment, "fold
acceleration by sterols" was calculated by dividing the protein's
t1/2 in the absence of sterols () by its
t1/2 in the presence of sterols (+). The results are
the mean ± S.E. of 11-16 separate experiments.
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The obvious lack of regulated HMGR degradation in L-90 cells
could have been the consequence of alterations in either the enzyme
molecule or the cellular machinery that operates in its degradation. To
distinguish between these possibilities, CHO and L-90 cells were
transfected with the plasmid pHMGal-Neo that encodes the chimeric
protein HMGal (26) under the control of the constitutive immediate-early promoter of human cytomegalovirus. Stable
G418-resistant colonies were pooled and assayed for expression and
regulation of -galactosidase activity. The results, normalized to
the activity in LPDS medium (100%), are shown in Fig.
8A. Consistent with
numerous previous reports (see Ref. 32 and references therein),
HMGal activity in CHO cells dropped to 13-30% of control
values when the cells were incubated either with lipoproteins (FCS),
MVA, or sterols in ethanol. This is due to a 3-6-fold acceleration in
the rate of HMGal turnover (32). None of these additions had any
appreciable effect on HMGal activity in L-90 cells (Fig. 8A). That this persistent activity resulted from a highly
stabilized HMGal protein was directly demonstrated in a pulse-chase
experiment where HMGal and HMGR were sequentially immunoprecipitated
from the same labeled samples (Fig. 8B). In CHO cells the
degradation of both HMGal and HMGR was accelerated to the same extent
following the addition of MVA or sterols, whereas neither agents had
any significant effect on the half-life of HMGR or HMGal in L-90
cells (Fig. 8B).
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Fig. 8.
Degradation of HMGal is not regulated in L-90
cells. A, on Day 0, pools of HMGal-transfected CHO
(open bars) and L-90 (closed bars) cells
were plated in FCS medium without or with 90 µM
lovastatin, respectively. On Day 1, the cells were fed with FCS, LPDS,
LPDS + MVA, or LPDS + sterols, as detailed in Fig. 3. After 24 h,
the cells were permeabilized with digitonin and
-galactosidase-specific activity was measured, as described under
"Experimental Procedures." The results
(A420/h/mg protein) were normalized relative to
cells incubated in LPDS and are the mean ± S.E. of four
experiments, each done on quadruplicate wells. B,
transfected CHO and L-90 cells, expressing highest
HMGal activity, were cloned by limiting dilution. The cells
were pulse-labeled and chased in the indicated media, as described in
Fig. 7. HMGal and HMGR were sequentially immunoprecipitated with
anti--galactosidase and anti-HMGR antibodies, respectively. Similar
results were obtained with three independent clones. Half-life values
are also given in Table III.
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The loss of regulated HMGR degradation in L-90 cells could have
been the outcome of selecting cells for lovastatin resistance in the
presence of serum lipoproteins (see below). Alternatively, it was
conceivable that this aberrant phenotype resulted from the massive
overproduction of HMGR protein that could have saturated a
rate-limiting step and/or depleted critical trans-acting
factor(s) that operates in reductase degradation. The latter
possibility was directly tested through analyzing yet another HMGR
overexpressing cell line, designated LP-90. Unlike L-90, LP-90
cells were gradually adapted to grow in 90 µM lovastatin
in a medium that lacks lipoproteins (LPDS medium; see "Experimental
Procedures"). Western blot analysis (Fig.
9A) demonstrated that LP-90
cells overproduce HMGR to the same extent as L-90 cells (Fig.
9A, compare lanes 6 and 10). However, unlike L-90 cells, it was evident that the levels of HMGR in
LP-90 cells were tightly regulated by serum lipoproteins, MVA, or
sterols, in a similar fashion as in CHO cells (Fig. 9A). A
direct examination of degradation rate by pulse-chase analysis (Fig.
9B) demonstrated that the half-life of HMGR in LP-90 cells
decreased from 12 to 3 h or to 1.5 h upon addition of MVA or
sterols, respectively (mean results of three experiments).
These changes in reductase turnover were very similar to the
effects observed in parental CHO cells and in clear contrast
to L-90 cells (Fig. 9B). Thus, the finding that LP-90
cells were capable of degrading large amounts of HMGR with
t1/2 of less than 2 h demonstrated
that overexpression of HMGR itself is not likely to cause
impaired regulation of HMGR degradation.
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Fig. 9.
LP-90 cells overexpress HMGR yet normally
regulate its degradation. A, CHO or L-90 cells
were plated in FCS medium without or with 90 µM
lovastatin, respectively. LP-90 cells were plated in LPDS medium with
90 µM lovastatin. On the following day, the cells were
fed with FCS (F; lanes 1, 5 and 9),
LPDS (L; lanes 2, 6, and 10), LPDS + MVA (M; lanes 3, 7, and 11), or LPDS + sterols (S; lanes 4, 8, and 12) media,
as described under "Experimental Procedures" and in Fig. 3. The
LPDS media for CHO cells contained 2 µM compactin and 100 µM MVA, and all media of L-90 and LP-90 cells
contain 90 µM lovastatin. Following 24 h incubation,
the cells were lysed, and 75 µg (CHO cells, lanes 1-4) or
15 µg (L-90 and LP-90 cells, lanes 5-12) of
post-nuclear cell extracts were analyzed by immunoblotting using
anti-HMGR antibodies. B, on Day 0, CHO, L-90, or
LP-90 cells were plated in FCS (CHO), FCS + 90 µM lovastatin (L-90), or LPDS + 90 µM lovastatin (LP-90) media, respectively. On
Day 1, all cells were refed with LPDS medium, as described under
"Experimental Procedures" and in Fig. 7. On Day 2, the cells were
pulse-labeled and chase in LPDS medium supplemented with the indicated
additions, as detailed in Fig. 7. HMGR was immunoprecipitated with
anti-HMGR antibodies and analyzed by SDS-PAGE.
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Based on the results presented in Fig. 8, we concluded that
the loss of regulated degradation of HMGR in L-90 cells
is due to changes in trans-acting factor(s)
required for accelerated turnover of the enzyme. This
factor(s) might be involved in metabolic cues that signal for
HMGR destabilization and/or in tagging the enzyme for
degradation and/or in the terminal proteolytic elimination of
the protein. We reasoned that metabolic cues might be
specific for degradation of HMGR, whereas tagging mechanisms
and/or cellular proteolytic systems could be generally
involved and possibly utilized in the ER degradation of other membrane
proteins. To address directly this question and further focus on the
mechanism(s) altered in L-90 cells, CHO and L-90 cells were
stably transfected with the plasmid pTCR-Neo that encodes the subunit of the T cell receptor (TCR; Ref. 43). It has been
previously shown in a variety of cell lines that a singly expressed
TCR is degraded rapidly and constitutively. Indeed, for many years
it served as a prime model for protein degradation in the ER (53-55).
Recent studies, however, have demonstrated that TCR is actually
degraded by the proteasome in the cytosol or at the cytoplasmic face of
the ER (43, 56). Nevertheless, TCR is co-translationally inserted
and glycosylated in the ER before its reverse translocation to the
cytoplasm for deglycosylation and proteolysis (43, 56). The
intracellular fate of TCR was monitored in CHO and in L-90
cells by pulse-chase and immunoprecipitation with specific monoclonal
antibodies (Fig. 10). TCR was
synthesized in both cell types as a 
40-kDa peptide N-glycosidase F-sensitive protein (Fig. 10A),
indicating that TCR is initially inserted and core-glycosylated in
the ER. We consistently observed higher levels of expression in
L-90 cells (Fig. 10A; compare lanes 1 and
2 with lanes 3 and 4, respectively). As shown in
Fig. 10B, TCR was rapidly degraded in CHO cells with a
half-life of ~1/2 h, a value consistent with previous reports
(55). Remarkably, degradation of TCR was significantly retarded in
L-90 cells, and high levels of the protein persisted for over
2 h after synthesis. Appreciable amounts could also be observed by
5 h of chase (Fig. 10B). From this and similar
experiments, the half-life of TCR in L-90 cells was determined
to be 2.5-3 h, over 5-fold longer than in CHO cells. Addition of
sterols during the chase had no effect on the half-life of TCR in
either cell lines (data not shown).
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Fig. 10.
Degradation of TCR is severely retarded in L-90 cells. A, duplicate
dishes of pooled populations of TCR-transfected CHO and L-90
cells were pulse-labeled for 15 min with
Expre35S35S protein-labeling reagent, as
described in Fig. 7A. At the end of the pulse, the cells
were lysed, and TCR was immunoprecipitated. Bound TCR was eluted
and either mock-digested (lanes 1 and 3) or
digested with 1000 units of peptide N-glycosidase F
(PNGase F) (lanes 2 and 4), according
to the manufacturer's instructions. Digestion products were separated
on 7.5% SDS-PAGE. B, cells were pulse-labeled for 15 min
and chased in unlabeled medium for the indicated times, and TCR was
immunoprecipitated. Arrows denote the fully glycosylated
species of TCR. Asterisk denotes partially glycosylated
or de-glycosylated species of the protein (43). In panel B,
there is an additional band that migrates slightly faster than this
species, and it co-migrates with the fully de-glycosylated TCR in
A. Half-life of TCR (see "Results") was determined
from the rate of decay of the slower migrating species (upper
band).
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The failure of L-90 cells to accelerate the turnover of HMGR in
response to exogenous MVA or sterols was manifested in cells that were
continuously maintained at 90 µM lovastatin. Therefore, it was of interest to investigate whether this aberrant phenotype is
stable and whether it depended on the presence of the inhibitor. For
that, HMGal-expressing L-90 cells were grown in a lovastatin-free FCS medium for varying periods, up to 37 days, and the degradation in
these L-90 "off" cells of HMGR and HMGal was monitored
by pulse-chase and immunoprecipitation. In each experiment, we also
measured in parallel the basal as well as sterol-accelerated turnover
of HMGR and HMGal in the parental CHO cells and in L-90 cells
that remained "on" lovastatin. The results of these experiments are illustrated in Fig. 11 and summarized
in Table III. As soon as 2 days after removing lovastatin, the
half-life of HMGR dropped from 12 h in L-90 "on" cells
to 4.6 h in L-90 "off" cells. This fast degradation
rate remained relatively constant (S.E. ± 0.3 h) throughout the
ensuing 35 days in lovastatin-free medium. It should be noted that the
half-life of 12.1 ± 0.9 h for HMGR in L-90 "on"
cells is very similar to that in the parental CHO cells (12.5 ± 0.7 h; Table III, see also Fig. 7). Nevertheless, whereas addition
of sterols to CHO cells accelerated reductase degradation nearly 5-fold
(t1/2 = 2.7 h), only a marginal effect of
sterols (15-35%) was observed in L-90 "on" (t1/2 = 11.0 h) as well as in L-90 "off" cells (t1/2 = 3.6 h; Table III).
This minimal effect of sterols was evident in L-90 cells after
removing lovastatin either for 2 days (sterols, 4.2 h; +sterols,
3.1 h), 9 days (sterols, 4.9 h; +sterols, 4.0 h), or 37 days (sterols, 4.2 h; +sterols, 3.0 h). In all cases,
the turnover of HMGal paralleled that of HMGR (Table III),
demonstrating that the effect of lovastatin on HMGR degradation rate is
not the result of its direct binding to the active site of the enzyme. Importantly, the degradation of TCR remained retarded in L-90 "off" cells (t1/2 2 h in cells that were
off lovastatin for 19 days as compared with 2.5 h in L-90 "on" cells and 0.5 h in CHO cells). Moreover, within 24 h
after re-challenging L-90 "off" cells with 90 µM lovastatin, HMGR re-acquired its extended half-life
(as in L-90 "on" cells), again without any appreciable effect
of sterols on reductase degradation (data not shown). These results
demonstrated that the loss of regulation of HMGR degradation is a
stable phenotype of L-90 cells that does not depend on the
presence of lovastatin.
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Fig. 11.
Rapid and unregulated degradation of HMGR in
L-90 "off" cells. HMGal-expressing L-90 cells were grown
for 3 days in FCS medium with (+, lanes 1-9) or without
(, lanes 10-18) 90 µM lovastatin. On
the 4th day, the cells were switched to LPDS medium with or without 90 µM lovastatin, as indicated. On Day 5, the cells were
pulse-labeled and chased in indicated media in the absence (lanes
1-4 and 10-13) or presence (lanes 6-9 and
15-18) of sterols, as described under "Experimental
Procedures" and in Figs. 7 and 8. HMGR and HMGal were
immunoprecipitated and analyzed by SDS-PAGE and fluorography. Not shown
in this figure are CHO-HMGal cells that were analyzed in parallel.
Similar experiments were performed on L-90 cells that were
maintained without lovastatin for 2-37 days. Half-life values of HMGR
and HMGal are summarized in Table III.
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DISCUSSION |
Mevalonate is an essential molecule for life of all eukaryotic
cells. Therefore, exposing naive cells to inhibitory concentrations of
statins causes elevation in HMGR levels in an effort to overcome the
blockage in endogenous MVA production. Unless MVA is added to the
growth medium, prolonged incubations with these drugs lead to cell
death. This homeostatic response has been exploited to select several
cell lines that are resistant to the cytotoxic effects of these potent
HMGR inhibitors (10, 49, 57, 58). In most cases, selection for statin
resistance has been achieved with cells that were initially adapted to
grow in a cholesterol-deficient medium. Under such conditions, the sole
source of the cell for cholesterol was through its endogenous synthesis
in the MVA pathway. Upon addition of statins, first at very low
concentrations, only cells that could develop higher HMGR levels have
survived, and resistant lines have been established by iterating this
step at increasing drug concentrations. The best studied of these lines is UT-1, a variant of CHO cells selected to grow in 40 µM
compactin (49). UT-1 cells survive at such high compactin
concentrations due to overexpression of HMGR (~2% of total cell
protein) resulting from an ~15-fold amplification of the gene (49,
59). Importantly, transcriptional control of HMGS, HMGR, and LDLR
genes, as well as sterol-regulated turnover of HMGR protein, is normal
in UT-1 cells (49, 59, 60).
Although gene amplification is frequently encountered in acquired drug
resistance (61), it is not the sole mechanism that allows cell growth
at high statin concentrations. Skalnik et al. (57) have
shown that in ML-100 cells, which grow in 100 µM
compactin, neither the gene for HMGR is amplified nor are HMGR mRNA
levels elevated. Sinensky et al. (62) reported that
lovastatin resistance of the MX1 cell line appears to be the result of
a mutation in HMGR that lowers its Km value toward
HMG-CoA. Finally, a point mutation in the promoter of the HMGR gene
confers lovastatin resistance to the archaebacterium Haloferax
vulcanii (63).
In the current study, we show that L-90 cells withstand
lovastatin toxicity by elevating the levels of HMGS and by the massive accumulation of HMGR. Concomitant with the overexpression of HMGR, a
membrane-bound enzyme, there is an impressive proliferation of smooth
ER and formation of crystalloid ER, similar to the situation observed
in UT-1 cells (49). However, L-90 cells are unique in that they
achieve accumulation of HMGR not only by gene amplification and rise in
mRNA levels but also by loss of regulated turnover of the protein.
In addition, compared with parental CHO cells, L-90 cells have
reduced content of cholesterol. This may account for the highly
elevated levels of FPPS and SQS in L-90 cells, presumably in an
effort to compensate for their chronic deprivation in MVA-derived
isoprenoids and sterols. Inasmuch as the transcription of many, if not
all, genes of the MVA pathway is regulated by SREBPs (30), the finding
that L-90 cells simultaneously overexpress HMGS, HMGR, FPPS and
SQS suggests that the proteolytic cleavage/activation of the ER
precursors for these transcription factors occurs in these cells at a
much higher rate, and/or that the mature SREBPs reside in the nuclei of
L-90 cell for a longer time. This is corroborated by the finding
that fatty acids synthesis is also accelerated in L-90 cells, in
agreement with the involvement of SREBPs in the transcription of
acetyl-CoA carboxylase, FAS, and stearoyl-CoA desaturase (30).
The slow and stepwise selection of cells for drug resistance may
cause the accumulation of multiple genetic lesions, leading to a
complex phenotype. Indeed, L-90 cells stand out from similar statin-resistant cells because of two important features that make them
especially interesting. First, L-90 cells have lost their ability
to degrade HMGR in a regulated fashion. Hence, in these cells reductase
appears to be regulated solely at the transcriptional level, by
controlling amounts of mRNA. Second, L-90 cells exhibit a
severe impairment in their capacity to respond to changes in the
availability of cholesterol carried in serum lipoproteins. We show that
this attenuated response is the result of reduced activity of the LDLR
pathway, which resembles the defective LDL-mediated regulation of
cholesterogenic enzymes in cells from individuals afflicted with
familial hypercholesterolemia (5). In experiments to be presented
elsewhere, we demonstrate that the reduced LDLR activity is due to
fewer surface receptors rather than lower LDL binding
affinity.2 Decreased activity
and abnormal regulation of LDLR have been also reported in the
compactin-resistant CR200 cells (58).
Cells have evolved a highly complex mechanism to get rid of excess HMGR
molecules when products of the MVA pathway are in abundance. Although
the molecular details of this mechanism are still vastly unknown, it is
firmly established that the membrane domain of HMGR is the
cis-acting element in the regulated degradation of the
enzyme. The results presented here clearly demonstrate that, similar to
endogenous reductase, the degradation of transfected HMGal in
L-90 cells is not accelerated by addition of exogenous sterols or
MVA. Thus, the fault in the regulation of degradation is a
trans-acting property of the cells and not a cis
defect in HMGR protein. This loss of regulated turnover is a stable
phenotype since after 37 days in culture without lovastatin (~30
generations) such L-90 "off" cells could not accelerate
the degradation HMGR or HMGal when challenged with excess sterols (see
below). It is unlikely that sterols or MVA fail to reach their target
intracellular compartments/factors since L-90 cells manifest
sterol-sensitive transcriptional regulation of HMGR and other
responsive genes. Also, L-90 cells display normal sterol
stimulation of acyl-CoA:cholesterol acyltransferase activity as well as
[14C]MVA-dependent synthesis of cholesterol
and prenylated proteins.2 These results indicate that in
L-90 cells sterols reach the ER and are capable of regulating ER
events such as the proteolytic activation of SREBPs and cholesterol
esterification. Yet, in L-90 cells sterols fail to accelerate
HMGR degradation, another ER-associated process (31). It is equally
unlikely that the apparent loss of regulated HMGR degradation in
L-90 cells is due to the overproduction of HMGR protein that
might have overwhelmed this complex degradative process. Indeed, we
show that comparably massive amounts of HMGR do not impede the capacity
of LP-90 cells to accelerate the turnover of the enzyme when challenged
with MVA or sterols. This is in full agreement with observations in
other statin-resistant lines, such as UT-1, C100, or CR200 cells, which
overproduce HMGR to a similar or even greater extent, yet regulate
normally the turnover of the enzyme (19, 35, 58).
What might have caused the loss of regulation of HMGR turnover in
L-90 cells? Unlike other statin-resistant lines (LP-90, UT-1, and
C100), L-90 cells were gradually adapted to grow in increasing
lovastatin concentrations in the presence of serum lipoproteins.
Studies by Nakanishi et al. (15) and Roitelman and Simoni
(32) demonstrated that sterol-accelerated degradation of HMGR will not
ensue unless MVA-derived nonsterols are synthesized in the cells. Yet,
MVA can stimulate HMGR degradation even when endogenous sterol
synthesis is specifically inhibited (32). The notion that rapid HMGR
degradation requires both sterol and nonsterol metabolic signals is
also supported by the differential sensitivity of these signals to
perturbation of cellular Ca2+ stores (32). Therefore, under
normal growth conditions, when LDL-cholesterol is abundan
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ABSTRACT
INTRODUCTION
