From the Department of Biological Sciences, Stanford University, Stanford, California 94305-5020
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ABSTRACT |
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The steady-state level of the resident
endoplasmic reticulum protein, 3-hydroxy-3-methylglutaryl-coenzyme A
reductase (HMGR), is regulated, in part, by accelerated degradation in
response to excess sterols or mevalonate. Previous studies of a
chimeric protein (HM-Gal) composed of the membrane domain of HMGR fused to Escherichia coli The endoplasmic reticulum resident membrane protein,
3-hydroxy-3-methylglutaryl-CoA reductase
(HMGR)1 (EC 1.1.1.34;
GenBankTM accession number M12705), is the rate-limiting
enzyme in the cholesterol biosynthetic pathway, catalyzing the
conversion of 3-hydroxy-3-methylglutaryl-CoA to mevalonate, which is
further metabolized to many downstream isoprenoid products as well as cholesterol (1). These downstream metabolites participate in feedback
regulation of HMGR to reduce its protein levels when sufficient
cholesterol is obtained through exogenous sources, and conversely, flow
through the mevalonate pathway is increased by up-regulating synthesis
and extending the protein half-life of HMGR when intracellular sterol
levels are low (2-5). This phenomenon of regulated HMGR degradation
has been a focus of study for many years; however, the mechanism of
regulation and the identity of the proteolytic components responsible
for degradation remain largely unknown.
The topology of HMGR, which is a 97-kDa glycoprotein, consists of two
major domains: a transmembrane domain that spans the endoplasmic
reticulum membrane eight times and the C-terminal catalytic domain,
which contains the active site and resides in the cytosol (6-8).
Structural studies of the Pseudomonas mevalonii HMGR
counterpart have revealed that the catalytic domain, in which key
residues involved in substrate recognition and catalysis are conserved
between bacteria and mammals, requires dimerization of two HMGR
molecules through this domain to form the substrate-binding and active
site (9). In addition, radiation inactivation studies of HMGR from rat
liver have demonstrated that the enzymatically active form of HMGR is a
dimer (26, 27).
The membrane domain of HMGR has been of interest due to work that has
shown that determinants for regulated degradation reside in this region
(8, 10). Experiments in which the cytosolic domain of HMGR was replaced
with the soluble Escherichia coli We have now generated several different HMGR fusion proteins with the
cytosolic domain replaced with various heterologous proteins. We have
discovered that there is an apparent correlation between the oligomeric
state of the heterologous cytosolic protein and resulting stability of
the fusion protein, with monomeric fusion proteins being degraded
relatively rapidly and oligomeric fusion proteins degraded slowly.
Based on these findings, we have developed a hypothesis that the
oligomeric state of HMGR, determined at least in part by interactions
through the cytosolic catalytic domain, plays a role in determination
of the degradation rate.
In this paper, we have tested this hypothesis using an "inducible
oligomerization" system that utilizes FK506-binding protein 12 (FKBP)
and the synthetic dimeric drug AP1510 (13, 14). The drug FK506 is a
well characterized immunosuppressant that binds to FKBPs, mimicking an
unknown endogenous ligand and activating the calcineurin signaling
pathway (15, 16). AP1510 is a nontoxic, cell-permeable compound that
consists essentially of two FK506 derivatives fused together through a
linker region and thus has the capability of dimerizing two
FKBP-containing proteins in vivo (14). We have constructed a
fusion protein called HM-3FKBP, which consists of three tandem repeats
of FKBP fused to the C terminus of the membrane domain of HMGR. Using
the dimeric AP1510 drug, we manipulated the oligomeric state of
HM-3FKBP in vivo and studied the degradation phenotype both
as a monomer and an oligomer. Additionally, we have also fused three
FKBP sequences to the C terminus of one of our "fast" monomeric
constructs, HM-Hyg, and tested whether AP1510 can stabilize its fast
degradation through oligomerization.
If the oligomeric state of HMGR is important for stability and
degradation, we predict that the level of HMGR expression should influence the relative proportion of monomers and dimers and thus should also influence the degradation rate. We have varied the in
vivo expression levels of HMGR by expression from an inducible promoter system, and the results show that the protein is more rapidly
degraded at lower concentrations. These findings further support our
hypothesis that oligomerization of HMGR molecules through the cytosolic
domain stabilizes the protein and that the monomeric state results in
faster degradation.
Materials and Reagents--
DNA purification kits were obtained
from QIAGEN Inc. Restriction enzymes and molecular biology reagents
were purchased from Life Technologies, Inc. and New England Biolabs
Inc. Transfection and cell culture reagents were obtained from Life
Technologies, Inc. The Tet-OffTM Expression System was
purchased from CLONTECH. All chemicals unless
otherwise noted were obtained from Sigma. FK506 was obtained from
Gerald Crabtree (Stanford University) and Fujisawa, Inc. AP1510 was
generously provided by Ariad Pharmaceuticals. Compactin was the kind
gift from Akira Endo (Tokyo Noko University). Anti-HA monoclonal
antibodies were purchased from Roche Molecular Biochemicals. Anti- Cell Culture--
Stably transfected CHO-K1 cell lines were
maintained in minimal essential medium supplemented with nonessential
amino acids, 5% fetal calf serum (MEM-FCS medium), and 0.25 mg/ml
active G418 in a humidified 5% CO2 incubator at 37 °C
and passaged every 3 days. Stable Tet-Off UT-2 cells were maintained
under the same conditions, except that MEM-FCS medium was replaced with
MEM-FBS medium, which contained 5% Tet System approved fetal bovine
serum (CLONTECH) instead of 5% fetal calf serum
and also 1 mM sodium mevalonate and 0.1 mg/ml active G418.
Double-stable Tet-Off UT-2 cells were maintained under the same
conditions as described above, except that the medium did not contain
sodium mevalonate. The double-stable Tet-Off CHO-AA8 cell line was
maintained under the same conditions as described above, except that
the medium also contained 0.1 mg/ml hygromycin B.
Plasmid Construction--
pMKIT HM-Hyg was constructed by PCR of
the E. coli hygromycin phosphotransferase (hph)
gene from a plasmid template, with restriction sites introduced in the
5' and 3' PCR primers, followed by restriction enzyme digestion and
ligation into the pMKIT plasmid vector containing the HMGR membrane
domain sequence. The maximum amount of PCR sequence was replaced with
native sequence using naturally occurring restriction sites to reduce
errors introduced by PCR, and the remaining unreplaced sequence was
subjected to DNA sequencing to verify accuracy. pMKIT HM-Gal deletion
mutants were generated by PCR of the 3' sequence of the lacZ
gene, using PCR primers to create deletions that corresponded to loss
of the C-terminal 10 or 20 amino acids from the Generation of Stable CHO-K1 Cell Lines--
CHO-K1 cells were
plated in 60-mm dishes at 60-80% confluency and transfected by
lipofection using LipofectAMINE PLUS (Life Technologies, Inc.)
according to the manufacturer's instructions. Cells were split the
following day onto multiple 10-cm dishes and fed selection medium
containing 1-2 mg/ml active G418 for 9-14 days. Resistant colonies
were either pooled or cylinder-cloned and analyzed by radiolabeling to
identify high expressing populations. The HM-eGFP stable cell line was
selected for high expressers by fluorescence-activated cell sorting.
Generation of Double-stable Tet-Off UT-2 Cell Line--
The
pTet-Off regulator plasmid DNA (CLONTECH) was
transfected into UT-2 cells with the same method as described above.
The stable Tet-Off cells were selected with 1 mg/ml G418 for 14 days, and the resistant colonies were pooled. After the stable Tet-Off cells
were obtained, the pooled stable Tet-Off cells were transfected with
pTRE HMGR recombinant plasmid DNA with the same method as described
above, and the double-stable cells were selected with minimal essential
medium supplemented with 5% lipid poor serum (MEM-LPS medium) prepared
by the method of Rothblat et al. (18) for 14 days. The
resistant cells were cylinder-cloned and analyzed by radiolabeling to
identify high expressing populations.
Generation of Double-stable Tet-Off CHO-AA8 Cell Line--
pTRE
HM-Gal DNA and pTK Hyg DNA (CLONTECH) were
cotransfected into commercially purchased stable Tet-Off CHO-AA8 cells
(CLONTECH) in a ratio of 20:1 with the same method
as described above. Transfectants were selected with 0.6 mg/ml
hygromycin B for 14 days, and resistant colonies with the highest
Pulse-Chase Analysis--
Cells were grown in 60-mm dishes to
70-80% confluency in MEM-FCS or MEM-FBS medium. The following day,
the medium was changed to MEM-LPS medium containing 10 µM
compactin and 100 µM sodium mevalonate. After 20 h,
cells were starved for 1 h in methionine/cysteine-free minimal
essential medium supplemented with 10 µM compactin and 100 µM sodium mevalonate and labeled for 0.5 h in
methionine/cysteine-free minimal essential medium containing 10 µM compactin, 100 µM sodium mevalonate, and
100 µCi of Tran35S-label per plate. Cells were chased in
MEM-LPS medium supplemented with 10 µM compactin, 100 µM sodium mevalonate, 2 mM methionine, and 2 mM cysteine out to various time points. At each chase time point, cells were collected by washing three times in ice-cold phosphate buffered saline, followed by lysis in ice-cold solubilization buffer (50 mM Tris-HCl (pH 7.5), 150 mM NaCl,
1% Nonidet P-40, 0.5% sodium deoxycholate, 0.1% SDS, 2 mM phenylmethylsulfonyl fluoride, 0.1 mM
leupeptin, 2 µg/ml calpain inhibitor I, and 100 mM
dithiothreitol). Lysates were centrifuged at 16,000 × g for 15 min at 4 °C to remove insoluble materials, and
supernatants were collected. 10-µl aliquots were reserved for boiling
trichloroacetic acid precipitation and scintillation counting. The
remaining supernatants were processed for immunoprecipitation with the
appropriate antisera. Antigen-antibody complexes were precipitated with
protein A-Sepharose (Amersham Pharmacia Biotech), washed twice in
solubilization buffer and once in 10 mM Tris-HCl (pH 7.5)
and 0.1% Nonidet P-40, and then solubilized in sample buffer (62.5 mM Tris-HCl (pH 6.8), 8 M urea, 15% SDS, 20%
glycerol, 0.25% bromphenol blue, and 25 mg/ml dithiothreitol) for 30 min at 37 °C. Samples normalized for equal trichloroacetic
acid-precipitable counts were loaded on 5-15% SDS-polyacrylamide
gradient gels for electrophoresis. The gels were then fixed in a
solution containing 45% methanol and 10% acetic acid, impregnated
with Enlightning (NEN Life Science Products), dried, and exposed to
x-ray film. Bands were quantitated using a PhosphorImager
(Bio-Rad).
Dimerization Experiments--
Cells were treated with no drug or
with 200 nM AP1510 or 400 nM FK506 in MEM-LPS
medium supplemented with 100 µM sodium mevalonate and 10 µM compactin for 20 h prior to pulse-chase analysis
as described above. Drug concentrations were maintained throughout the
experiment in starvation, labeling, and chase media.
Experiments of Regulating HMGR and HM-Gal Expression
Levels--
Double-stable Tet-Off UT-2 cells were treated either with
no doxycycline or with various concentrations of doxycycline in MEM-LPS
medium containing 100 µM sodium mevalonate and 10 µM compactin for 20 h prior to pulse-chase analysis
as described above. Drug concentrations were maintained throughout the
experiment in starvation, labeling, and chase media. Double-stable
Tet-Off CHO-AA8 cells were treated either with no doxycycline or with
various concentrations of doxycycline along with no other reagent or
with 2.5 µM 25-hydroxycholesterol or 4 µg/ml calpain
inhibitor I (N-acetyl-leucyl-leucyl-norleucinal) (Calbiochem-Novabiochem Corporation) in MEM-LPS medium supplemented with 100 µM sodium mevalonate and 10 µM
compactin for 20 h prior to Enzyme Activity Assays--
Monomeric HMGR Membrane Domain Fusion Proteins Are Subject to Rapid
Degradation, whereas Oligomeric Fusion Proteins Are More
Stable--
We have now generated several chimeric proteins consisting
of the membrane domain of HMGR fused to various heterologous proteins. In the course of analyzing these fusion proteins, we observed that some
chimeric proteins are degraded rapidly, whereas others are stable and
exhibit normal regulated degradation characteristic of HMGR despite the
fact that all constructs have the identical membrane domain from HMGR.
We have found that the degradation phenotype correlates with the
oligomeric state of the cytosolic domain, with monomeric proteins being
degraded rapidly and dimeric and higher order proteins degraded
normally. In addition to HM-Gal (HM designates the HMGR membrane domain
and is followed by a designation for the heterologous cytosolic
protein), another construct we have made is HM-Hyg, which is composed
of the HMGR membrane domain fused to the hygromycin phosphotransferase
protein, which confers resistance to the antibiotic hygromycin B. We
have also constructed HM-XGPRT, which consists of the membrane domain
fused to the bacterial xanthine guanosylphosphoribosyltransferase,
which enables cells to utilize xanthine as an alternate source for
purines. Additionally, we have created HM-eGFP (with the membrane
domain fused to enhanced green fluorescent protein) and HM-3HA (with
three HA epitope tags attached to the end of the membrane domain). The
results of degradation studies by pulse-chase analysis of these
proteins in stably transfected cells are shown in Table
I.
Fusion proteins with normal degradation phenotypes include HM-Gal and
HM-XGPRT, whereas HM-Hyg, HM-3HA, and HM-eGFP are degraded relatively
rapidly (Table I). Consideration of the oligomeric state of these
heterologous proteins fused to the membrane domain reveals that
We assayed the heterologous cytosolic domains of the chimeras for
functional activity as evidence of oligomeric assembly in vivo while fused to the HMGR membrane domain. We detected enzyme activity for HM-Gal (12) and HM-XGPRT (data not shown), indicating they
are present as tetramers and trimers, respectively. HM-Hyg is capable
of conferring resistance to hygromycin B in transfected CHO cells (data
not shown), suggesting that HM-Hyg is capable of folding correctly and
that, most likely, the rapid degradation observed in this construct is
not due to misfolding. Similarly, HM-eGFP is also correctly folded as
evidenced by detectable GFP fluorescence in transfected cells (data not
shown). Based on the correlation between the oligomeric state and
degradation phenotype, we hypothesize that interaction between the
membrane domains induced through the cytosolic domain is required for
normal stability and sterol-regulated degradation to occur.
Monomeric HM-Gal Deletion Mutants Are Rapidly
Degraded--
As a first test of our hypothesis, we
constructed deletion mutants in which the C-terminal 10-20 amino acids
of the Induced Oligomerization Results in Stabilization of HMGR Membrane
Domain Fusion Proteins--
In the experiments described above,
it is possible the deletions in HM-Gal protein could result in the
proteins being recognized as misfolded and degraded by a process
unrelated to normal HMGR degradation even though the mutant
We constructed a new fusion protein with three tandem copies of FKBP
fused to the HMGR membrane domain, called HM-3FKBP. Initial experiments
with chimeras containing only one FKBP gave ambiguous results, and more
striking results were obtained with three copies of FKBP as reported
previously (14). However, with three copies of the ligand-binding
protein, there can be higher order structures formed in addition to
dimers. We stably expressed HM-3FKBP in CHO-K1 cells and measured its
half-life by pulse-chase analysis in both the presence and absence of
sterols. As our hypothesis predicts for a monomeric fusion construct,
the half-life of HM-3FKBP was rapid, <3 h even in the absence of
sterols, compared with the half-life of the endogenous HMGR control,
~9 h (Fig. 4A, No Drug lanes; and Table I). Also shown in Fig.
4A, treatment of cells with 200 nM AP1510
significantly stabilized HM-3FKBP compared with cells receiving no drug
and cells treated with the monomeric form of the drug (FK506). In fact,
in the cells receiving AP1510, the half-life of HM-3FKBP was ~7.5 h,
which is near the normal half-life of HMGR in the absence of sterols.
The control testing the effects of FK506 is critical since it is often
the case that ligand binding can stabilize a protein. As shown in Fig.
4A, FK506 did appear to have a small effect, but could not
stabilize HM-3FKBP to the same extent as AP1510. In three repetitions
of this experiment, the greatest effect of FK506 was observed in the
experiment shown here, and in one experiment, there was no effect.
We also examined the effect of AP1510 on the degradation rate of
HM-3FKBP in the presence of a regulatory sterol, 2.5 µM
25-hydroxycholesterol. In the presence of sterols, we detected only a
minor increase in degradation rate (Fig. 4, A and
B, compare No Drug lanes) since the
half-life was short even in the absence of sterol treatment. However,
AP1510 treatment was capable of moderately stabilizing sterol-accelerated degradation as well (Fig. 4B). We feel
that it is striking that the half-life of HM-3FKBP when measured in the
presence of AP1510 is very similar to that of HMGR in both the absence
and presence of sterols, suggesting that this artificial system is able
to mimic the normal physiological situation.
To be sure that the effect of AP1510 we observed is due to
oligomerization through the FKBP domain, we tested the protein HM-3HA,
which is identical to HM-3FKBP except that it is missing the three FKBP
domains. When we subjected HM-3HA-transfected cells to the same drug
regimen as described above for HM-3FKBP, we saw that AP1510 treatment
no longer had any effect on degradation (Fig.
5, A and B). These
results indicate that stabilization of HM-3FKBP by AP1510 is mediated
through the FKBP domains.
If the stabilization we observed in HM-3FKBP is due specifically to the
dimerizer AP1510 and not to other artifactual reasons involved in the
drug treatment, we would predict that addition of an excess of the
monomeric form of the drug (FK506) should be able to competitively
abolish the stabilization through competition for FKBP-binding sites.
Fig. 6 shows that treatment of cells with AP1510 plus a 5-fold molar excess of FK506 resulted in the loss of
stabilization of HM-3FKBP induced by AP1510. These results also support
our conclusion that AP1510 is able to oligomerize HM-3FKBP and that the
resulting oligomers stabilize degradation of the membrane domain.
AP1510 Can Also Stabilize HM-Hyg-3FKBP--
In an effort to
determine whether our other previously studied monomeric constructs
were degraded rapidly due to the monomeric state of the membrane
domain, we fused three copies of FKBP to the C terminus of one of them,
HM-Hyg, and studied its degradation using the AP1510 system. Cells
stably transfected with this HM-Hyg-3FKBP construct were treated with
no drug or with 200 nM AP1510 or 400 nM FK506
and subjected to pulse-chase analysis in the absence or presence of 2.5 µM 25-hydroxycholesterol. Our results again show a
significant stabilization of HM-Hyg-3FKBP in both the presence and
absence of sterols when the cytosolic domains are oligomerized by
AP1510 (Fig. 7A).
As a control, we also subjected cells stably transfected with HM-Hyg,
which lacks the FKBP domains, to the same drug regimen and pulse-chase
analysis. As shown in Fig. 7B, AP1510 treatment had no
effect on HM-Hyg degradation.
HMGR Concentration Can Affect Its Rate of Degradation--
Our
work described above presents the following: 1) the correlation of
degradation rates with the oligomeric structure of the cytosolic domain
of various HM-X proteins, 2) mutations in the
oligomerization domain of HM-Gal resulting in monomer formation and
rapid degradation, and 3) the AP1510-induced oligomerization of
HM-3FKBP and HM-Hyg-3FKBP. These findings all suggest that oligomerization of HMGR is required for its stability. If this hypothesis is correct and since oligomerization is a
concentration-dependent process, we predict that changes in
HMGR concentration inside cells should change the ratio of oligomers to
monomers and, as a result, also the HMGR degradation rate.
To test this prediction, the HMGR cDNA was cloned into the Tet-Off
Expression System (see "Experimental Procedures") and transfected into UT-2 cells, which lack endogenous HMGR. Using the Tet-Off system,
the level of expression can be modulated by various concentrations of
doxycycline. The half-life of HMGR at the various HMGR concentrations was determined with pulse-chase assays. As shown in Fig.
8, the half-life of HMGR was dramatically
decreased in cells treated with doxycycline to reduce the steady-state
levels of HMGR. When cells were treated with 0.25 ng/ml doxycycline,
the half-life of HMGR was reduced to ~2.5 h compared with the normal
half-life of ~7.5 h in untreated cells. When the expression level of
HMGR was further suppressed with a higher concentration of doxycycline (0.50 ng/ml), the basal half-life of HMGR declined further to ~1.9 h.
These results also support our hypothesis that the rates of degradation
of HMGR degradation are determined, in part at least, by the
equilibrium between oligomeric and monomeric states.
We have considered the membrane domain of HMGR necessary and
sufficient for regulated degradation based on initial studies with
HM-Gal, a model protein that mirrors full-length HMGR in all aspects of
degradation while lacking the native cytosolic catalytic domain
entirely (12, 25). Our current studies of various chimeras of HMGR with
the cytosolic domain replaced with different heterologous proteins
suggest that this assumption may not be entirely correct, and with
HM-Gal, we had studied a chimera that retained an additional aspect of
HMGR important for degradation. We now propose that, in addition to the
membrane domain that is clearly required, interaction through the
cytosolic domain of HMGR is important for normal regulated degradation
to occur, most likely by bringing key elements of the membrane domain
into close proximity. We believe that dimerization of the membrane
domain, or at least close apposition, that is promoted through
interactions in the cytosolic domain is an additional requirement for
HMGR stability.
To test our hypothesis, we used three approaches. 1) We mutated a known
oligomeric, chimeric protein (HM-Gal) that exhibits normal regulated
degradation in order to convert it into a monomeric protein. 2) We used
an inducible oligomerization system to generate oligomeric proteins
with drugs in vivo. 3) We examined the effect of altering
the levels of HMGR with a regulated expression system to determine the
concentration dependence of the degradation rate. All three approaches
support the hypothesis that oligomerization of HMGR through the
catalytic cytosolic domain determines the half-life of the protein.
It is of note that in many of our monomeric constructs, although the
basal half-life is fast, addition of sterols in some cases is still
capable of further accelerating degradation. There are a few
possibilities to explain this. The first is that sterols induce an
additional effect either directly or indirectly on the membrane domain
to facilitate recognition of HMGR as a substrate for proteolysis. This
effect could be independent of the oligomeric state of the membrane
domain and would be less striking in proteins that have an extremely
fast half-life, as they are already being rapidly degraded. Another
possibility is that the membrane domains are capable of weak
interactions without the cytosolic domain, so there are some weak
dimers present even between the presumed monomeric constructs, and
addition of sterols results in a more complete dissociation of these
dimers to monomers. This may explain the variable half-lives observed
in the different fast monomeric proteins. It is possible that
differences in the extent of interaction in the membrane domain
permitted by the cytosolic component (perhaps due to varying degrees of
steric hindrance) are reflected in the differences in the resulting
equilibrium between monomers and dimers. Certainly these possibilities
are not mutually exclusive, and the explanation could be some
combination of the above. However, the simplest model would be to
propose that sterols bind directly to the membrane domain of HMGR and
that this binding can induce dissociation of dimers within the membrane
domain and/or a conformational change that exposes a degradation signal
or cleavage site to target HMGR for proteolysis.
HMGR exists as a dimer, as indicated by structural studies of bacterial
HMGR (9) and also by radiation inactivation studies that have
identified the functionally active form of HMGR as a mass that is
equivalent to two HMGR molecules (26, 27). We speculate that
dimerization of the cytosolic domain is important in promoting
dimerization of the membrane domain and that the oligomeric state of
the membrane domain affects its degradation such that monomers are more
susceptible to degradation and dimers are more stable. Dimerization of
the membrane domain may slow or prevent recognition of degradation
signals, resulting in a stable complex, whereas conformational changes
and/or dissociation into monomers normally induced by sterols may serve
to expose these signals and to promote recognition by the proteolytic machinery.
There is also correlative evidence that the membrane domain of HMGR has
a direct sterol-sensing/binding function based primarily on mutagenesis
studies demonstrating that mutations within the membrane domain can
render HMGR insensitive to sterols (25). In addition, other proteins
involved in cholesterol sensing (sterol regulatory element binding
protein (SREBP) cleavage-activating protein (28)) and transport
(NPC1 gene product (29, 30)) share sequence homology in the
putative sterol-binding region of the HMGR membrane domain.
There are examples in other systems that suggest that the
oligomerization state of membrane proteins may change in response to
changes in the membrane lipid composition. One example is the UDP-GlcNAc:dolichol-P GlcNAc-1-phosphotransferase in the endoplasmic reticulum, which is responsible for the committed step of
dolichol-linked oligosaccharide synthesis. UDP-GlcNAc:dolichol-P
GlcNAc-1-phosphotransferase is detectable as both monomers and dimers,
and it has been reported that the activity of this enzyme varies
depending upon local phospholipid composition (32, 33). Dan and Lehrman
(34) have suggested that different phospholipid concentrations may
alter the ratio of monomers to dimers and, in this manner, regulate
activity. Another example of membrane lipids regulating oligomerization has been reported in work with transcobalamin receptor II (35, 36),
which is normally present in the plasma membrane. Bose et
al. (37) have suggested that transcobalamin receptor II can transition between monomeric and dimeric forms depending on the membrane cholesterol content. The authors were able to show that cholesterol depletion in native intestinal plasma membranes or its
enrichment in microsomal membranes resulted in the in situ conversion of the dimeric to the monomeric form or of the monomeric to
the dimeric form, respectively.
In summary, the model we currently favor in light of our work is shown
in Fig. 9. The simplest hypothesis would
involve a direct sterol-sensing function by the membrane domain that
would initiate dissociation of the membrane domains and not necessarily of the cytosolic domains. This sterol-bound monomeric form of the
membrane domain would then be the prime substrate for proteolysis. Our
current work is aimed at demonstrating a sterol-induced change in the
interaction between membrane domains using cross-linking experiments.
-galactosidase, as a replacement of
the normal HMGR cytosolic domain, have shown that the regulated
degradation of this chimeric protein, HM-Gal, is identical to that of
HMGR (Chun, K. T., Bar-Nun, S., and Simoni, R. D. (1990)
J. Biol. Chem. 265, 22004-22010; Skalnik, D. G.,
Narita, H., Kent, C., and Simoni, R. D. (1988) J. Biol.
Chem. 263, 6836-6841). Since the cytosolic domain can be
replaced with
-galactosidase without effect on regulated
degradation, it has been assumed that the cytosolic domain was not
important to this process and also that the membrane domain of HMGR was
both necessary and sufficient for regulated degradation. In contrast to
our previous results with HM-Gal, we observed in this study that
replacement of the cytosolic domain of HMGR with various heterologous
proteins can have an effect on the regulated degradation, and the
effect correlates with the oligomeric state of the replacement
cytosolic protein. Chimeric proteins that are oligomeric in structure
are relatively stable, and those that are monomeric are unstable. To
test the hypothesis that the oligomeric state of the cytosolic domain
of HMGR influences degradation, we use an "inducible" system for
altering the oligomeric state of a protein in vivo. Using a
chimeric protein that contains the membrane domain of HMGR fused to
three copies of FK506-binding protein 12, we were able to induce
oligomerization by addition of a "double-headed" FK506-like
"dimerizer" drug (AP1510) and to monitor the degradation rate of
both the monomeric form and the drug-induced oligomeric form of the
protein. We show that this chimeric protein, HM-3FKBP, is unstable in
the monomeric state and is stabilized by AP1510-induced
oligomerization. We also examined the degradation rate of HMGR as a
function of concentrations within the cell. HMGR is a functional dimer;
therefore, its oligomeric state and, we predict, its degradation rate
should be concentration-dependent. We observed that it is
degraded more rapidly at lower concentrations.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-galactosidase enzyme
showed that the resulting chimera (termed HM-Gal), when stably
transfected into Chinese hamster ovary (CHO) cells, exhibits a
half-life that mirrors that of HMGR and is similarly subject to
sterol-accelerated degradation (11, 12, 31). Based on this observation,
we have assumed that the cytosolic domain was not important and that
the membrane domain was both necessary and sufficient to confer
regulated degradation.
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-galactosidase monoclonal antibodies were purchased from Promega. Sodium mevalonate was converted from
DL-mevalonolactone as described by Brown and co-workers
(17). Digitonin was purchased from Fluka.
-galactosidase
protein. The PCR products were ligated into pMKIT HM-Gal and verified
by DNA sequencing. pCMV HM-eGFP was created by ligation of the HMGR membrane domain sequence (EcoRI/PstI fragment)
into the commercially purchased pCMV-N1 eGFP vector
(CLONTECH) in frame and upstream of the eGFP coding
sequence. FKBP-containing constructs were generated by PCR of triple
copies of FKBP and ligation of the PCR product into pMKIT HM or pMKIT
HM-Hyg to create pMKIT HM-3FKBP and pMKIT HM-Hyg-3FKBP, respectively.
The stop codon was removed from the end of the coding sequence of Hyg
by PCR prior to FKBP insertion. All constructs were verified by DNA
sequencing. pTRE HMGR was produced by ligation of full-length HMGR
cDNA sequences (EcoRI/BamHI fragment) into
the commercially purchased pTRE-Off vector
(CLONTECH). pTRE HM-Gal was generated by insertion
of full-length HM-Gal cDNA sequences
(EcoRI/XbaI fragment) into the pTRE-Off vector.
-galactosidase activity were isolated using fluorescence-activated
cell sorting (10).
-galactosidase activity assays.
-Galactosidase activity assays
were performed as described previously (12). Briefly, cells seeded in
triplicate in 24-well dishes were permeabilized with 50 µg/ml
digitonin and incubated at 37 °C with a 1 mg/ml concentration of the
colorimetric enzyme substrate
o-nitrophenyl-
-D-galactopyranoside until
color change was visually detectable. Reactions were stopped by
addition of 1 M Na2CO3.
Concentration of cleaved product was quantitated using a Beckman DU-64
spectrophotometer. Specific activity values were calculated by
normalization to time of incubation and total protein, which was
determined on duplicate plates by the method of Lowry et al.
(19).
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
Half-lives of HMGR membrane domain fusion proteins
-galactosidase is enzymatically active as a tetramer; xanthine
guanosylphosphoribosyltransferase is a trimer (15, 20); and hygromycin
phosphotransferase (21), green fluorescent protein, and the HA epitope
tag are monomers, suggesting that the oligomeric state of the cytosolic
domain may influence degradation of the membrane domain.
-galactosidase enzyme in HM-Gal were removed since it has
been shown that these residues are critical for tetramerization of
-galactosidase and formation of the active enzyme (22, 23). Removal
of these amino acid residues should result in the monomerization of the cytosolic region of HM-Gal. Transfected CHO cell lines expressing either HM-Gal
10 (missing the C-terminal 10 amino acids) or
HM-Gal
20 (missing the C-terminal 20 amino acids) were analyzed by
pulse-chase analysis to measure half-lives. As shown in Fig.
1, both deletion mutants, in contrast to
full-length HM-Gal, were degraded abnormally fast (Fig. 1, compare
A and B with C). The half-life of
HM-Gal was ~9 h, and the presence of sterols reduced it to ~5 h.
For both HM-Gal
10 and HM-Gal
20, the half-lives were <2 h, and
there was no acceleration by sterols. To demonstrate that our deletion mutants were no longer assembled into tetramers, we measured
-galactosidase activity in these cell lines, as
-galactosidase
activity is known to be dependent upon tetramerization of the enzyme.
The results in Fig. 2 support the
conclusion that tetramerization of
-galactosidase in the deletion
mutants has been abolished since HM-Gal
10 and HM-Gal
20 cell lines
have lost
-galactosidase activity. In these experiments, the
HM-Gal/HM-Gal
10/HM-Gal
20
-galactosidase protein ratio was
1:0.8:0.2 (data not shown).
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Fig. 1.
Monomerization of HM-Gal through deletions of
the C-terminal 10-20 amino acids results in rapid degradation.
Deletion of the C-terminal 10 amino acids (called HM-Gal 10) or 20 amino acids (called HM-Gal
20) from HM-Gal resulted in significantly
faster turnover compared with intact HM-Gal. Cells were stably
transfected with each construct and subjected to pulse-chase analysis
as described under "Experimental Procedures." A, results
of pulse-chase experiment in cells stably transfected with HM-Gal
10.
Cells were metabolically labeled and chased in the absence (NA
lanes) or presence (+ Sterols lanes) of 2.5 µM 25-hydroxycholesterol for the indicated times and then
subjected to immunoprecipitation with anti-
-galactosidase
antibodies, separation by SDS-polyacrylamide gel electrophoresis, and
autoradiography. Bands were quantitated using a PhosphorImager.
B, results of pulse-chase experiment in HM-Gal
20 cells
under the same conditions as described for A. C,
results of control wild-type (wt) HM-Gal pulse-chase
experiment. Experimental procedures were the same as described for
A.
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Fig. 2.
HM-Gal deletion mutants lose
-galactosidase activity.
-Galactosidase
activity was measured in each of the three cell lines as well as in a
control vector-only transfected cell line as described under
"Experimental Procedures." Cells were permeabilized with 50 µg/ml
digitonin and incubated with 1 mg/ml
o-nitrophenyl-
-D-galactopyranoside at
37 °C until color change was visually detectable, and then cleaved
product was quantified using a Beckman DU-64 spectrophotometer.
Specific activity is reported in units of
A420/(hours of incubation × milligram of
protein). wt, wild-type.
-galactosidase is attached to the membrane domain of HMGR. A better
test of our hypothesis would be to study the same HM-X
chimeric protein both as a monomer and a dimer in the same cell line
and to compare the degradation phenotypes in each of these states. This
was made possible by taking advantage of the previously described
inducible dimerization system utilizing FKBP and synthetic
"dimerizer" drugs based on FK506 (14, 16, 24). Treatment of cells
expressing chimeric proteins including FKBP with the dimerizer drug
AP1510, which is composed of two FK506 derivatives fused
together, resulted in dimerization of the FKBP-containing fusion
proteins. A schematic of this system is shown in Fig.
3.
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Fig. 3.
Schematic of the effects of a dimerizer on
membrane domain fusion and FKBP fusion constructs. A,
chemical structure of the dimerizer AP1510; B, model of
AP1510 action. FKBP normally exists as a monomer; therefore, we assume
that in the absence of a dimerizer, HM-FKBP is primarily monomeric.
Addition of a dimerizer will induce dimerization of the FKBP domains,
which in turn brings together the membrane domains. ER,
endoplasmic reticulum.
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Fig. 4.
Oligomerization of the membrane domain
through three FKBP domains stabilizes HM-3FKBP degradation in the
presence of a dimerizer. Cells stably transfected with HM-3FKBP
were pretreated with no dimerizer (No Drug lanes) or with
200 nM AP1510 (+ AP1510 lanes) or 400 nM FK506 (+ FK506 lanes) for 20 h in
MEM-LPS medium prior to pulse-chase analysis. AP1510 and FK506 were
maintained throughout the starvation, labeling, and chase periods as
described under "Experimental Procedures." Cells were chased in the
absence (A) or presence (B) of 2.5 µM 25-hydroxycholesterol. Cells were lysed,
immunoprecipitated with anti-HA antibodies, separated by
SDS-polyacrylamide gel electrophoresis, and subjected to
autoradiography.
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Fig. 5.
Control HM-3HA cells, which lack FKBP
domains, are not stabilized by a dimerizer. Control HM-3HA cells
(no FKBP domains) were treated identically with drugs and subjected to
pulse-chase analysis as described for Fig. 6. Cells were chased in the
absence (A) or presence (B) of 2.5 µM 25-hydroxycholesterol prior to harvesting,
immunoprecipitation, and SDS-polyacrylamide gel electrophoresis.
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Fig. 6.
Excess FK506 treatment can abolish
stabilization induced by the dimerizer AP1510 in HM-3FKBP cells.
Cells were pretreated with 200 nM AP1510, 200 nM AP1510 + 2 µM FK506, or 2 µM
FK506 for 20 h prior to the pulse-chase experiments. Drugs were
maintained throughout the entire experiment as described under
"Experimental Procedures." Degradation was measured in the absence
(A) and presence (B) of 2.5 µM
25-hydroxycholesterol.
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Fig. 7.
HM-Hyg-3FKBP degradation is also stabilized
by the dimerizer AP1510. A, cells stably transfected
with HM-Hyg-3FKBP were pretreated with no drug or with 200 nM AP1510 or 400 nM FK506 in MEM-LPS medium for
20 h prior to pulse-chase experiments carried out as described
under "Experimental Procedures." Drugs were maintained throughout
the experiment, and cells were chased either in the absence (top
panel) or presence (bottom panel) of 2.5 µM 25-hydroxycholesterol. B, control HM-Hyg
cells were subjected to the same drug treatment and experimental
conditions as described for A.
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Fig. 8.
HMGR concentration impacts on the degradation
rate, with a faster degradation rate correlated with a lower expression
level. Double-stable Tet-Off UT-2 cells were pretreated with no
doxycycline (Dox) or with 0.25 or 0.50 ng/ml doxycycline for
20 h in MEM-LPS medium prior to pulse-chase analysis. Doxycycline
was maintained throughout the entire pulse-chase periods as described
under "Experimental Procedures." At various chase time points,
cells were collected, lysed, immunoprecipitated with anti-HMGR
antibodies against the HMGR membrane domain, separated by
SDS-polyacrylamide gel electrophoresis, and subject to
autoradiography.
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
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Fig. 9.
Schematic of how degradation of HMGR is
initiated by sterol treatment. We propose that the membrane domain
of HMGR senses sterol levels in the endoplasmic reticulum
(ER) by direct binding to cholesterol. The binding of
sterols initiates dissociation of the membrane domain, whereas the
catalytic domain remains associated. Monomerization of the membrane
domain exposes a degradation signal, and the monomer now becomes
susceptible to degradation.
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ACKNOWLEDGEMENT |
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We thank Gerald Crabtree for many helpful discussions about the use of dimerizer technology.
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FOOTNOTES |
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* This work was supported in part by National Institutes of Health Grant HL 26502.The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Supported by a fellowship from the American Heart Association.
§ Present address: Chugai Pharmaceutical Co., Ltd., Central Research Labs, 41-8 Takada-3-chome, Toshima-ku, Tokyo 171-8545, Japan.
¶ To whom correspondence should be addressed. Tel.: 650-725-4817; Fax: 650-725-5807; E-mail: rdsimoni{at}leland.stanford.edu.
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ABBREVIATIONS |
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The abbreviations used are: HMGR, 3-hydroxy-3-methylglutaryl-CoA reductase; CHO, Chinese hamster ovary; FKBP, FK506-binding protein 12; Hyg, hygromycin; HA, hemagglutinin; MEM, minimal essential medium; FCS, fetal calf serum; FBS, fetal bovine serum; PCR, polymerase chain reaction; eGFP, enhanced green fluorescent protein; LPS, lipid-poor serum; XGPRT, xanthine guanosylphosphoribosyl- transferase.
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