From the Department of Pathology, Harvard Medical School, Boston, Massachusetts 02115
Received for publication, October 3, 2002, and in revised form, December 11, 2002
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
We examined the effects of protein folding on
endoplasmic reticulum (ER)-to-cytosol transport (dislocation) by
exploiting the well-characterized dihydrofolate reductase (DHFR)
domain. DHFR retains the capacity to bind folate analogues in the lumen of microsomes and in the ER of intact cells, upon which it acquires a
conformation resistant to proteinase K digestion. Here we show that a
Class I major histocompatibility complex heavy chain fused to DHFR is
still recognized by the human cytomegalovirus-encoded glycoproteins US2
and US11, resulting in dislocation of the fusion protein from the ER
in vitro and in vivo. A folded state of the DHFR domain does not impair dislocation of Class I MHC heavy chains in vitro or in living cells. In fact, a slight acceleration
of the dislocation of DHFR heavy chain fusion was observed in
vitro in the presence of a folate analogue. These results suggest
that one or more of the channels used for dislocation can
accommodate polypeptides that contain a tightly folded domain of
considerable size. Our data raise the possibility that the Sec61
channel can be modified to accommodate a folded DHFR domain for
dislocation, but not for translocation into the ER, or that a channel
altogether distinct from Sec61 is used for dislocation.
Nascent polypeptides destined for secretion or membrane insertion
enter the ER1 via a protein
channel referred to as the translocon (1). N-Linked oligosaccharides are added cotranslationally, and the formation of
disulfide bonds may be initiated on the nascent polypeptide. Folding of
nascent chains is assisted by an elaborate system of chaperones that
includes calnexin, calreticulin, and oxidoreductases such as PDI and
ERp57. To exit the ER along the secretory pathway, proteins must pass
an ER quality control system. A failure to pass this checkpoint may
trigger protein degradation, which mostly takes place in the cytoplasm
(2). Terminally misfolded proteins or unassembled protein complexes,
which are dislocated from the ER to the cytosol, are degraded by the
proteasome (3). Upon arrival in the cytoplasm, glycoproteins may lose
their N-linked glycan in a reaction catalyzed by
N-glycanase, presumably prior to proteasomal degradation.
The human cytomegalovirus-encoded US2 and US11 glycoproteins mediate
the destruction of class I MHC heavy chains (HC) by catalyzing the
transfer of Class I HC from the ER to the cytosol, followed by
proteasomal degradation, all in a matter of a few minutes. US2 and US11
usurp the normal cellular degradation machinery to selectively
eliminate class I MHC as a means of immune evasion (4). p97, an
AAA ATPase, may mediate the extraction of ubiquitinated HC from
the ER in the case of US11-mediated degradation (5). However, because
ubiquitination takes place in the cytosol and the cytosolic tail of HC
is not directly ubiquitinated, ubiquitination cannot be the active
force behind HC dislocation but must be located downstream from some as
yet unidentified initiating events (6).
In a previous study we demonstrated that an EGFP-HC fusion protein
composed of EGFP fused to the N terminus of the Class I MHC luminal
domain is still subject to accelerated degradation when coexpressed
with US2 and US11 and yields similar deglycosylated breakdown
intermediates as seen for endogenous Class I HC. Surprisingly, EGFP
fluorescence was retained throughout the process, suggesting either
that dislocation occurred without a requirement for unfolding of the
GFP domain or that unfolding and refolding of GFP had to occur rapidly
(7).
Here we tested the effects of folding on ER dislocation by using the
well-characterized dihydrofolate reductase (DHFR) domain. DHFR is a
cytosolic enzyme that acquires a tightly folded conformation when bound
to folate analogues like methotrexate (MTX) or the hydrophobic
membrane-permeable derivative, trimetrexate (TMX) (8). DHFR, when
occupied by these ligands, acquires resistance to proteinase K
digestion, yielding a distinctive 21-kDa protease-resistant fragment.
In this capacity, DHFR can serve as a module that can block
translocation across a membrane when only unfolded polypeptides are
translocation competent. In this manner, fusion proteins using the DHFR
moiety have been used extensively to study transport of proteins into
mitochondria, a process strongly inhibited by MTX-stabilized DHFR (9).
Other studies using DHFR fusion proteins demonstrated that
chaperone-mediated import into lysosomes, import into chloroplasts, and
transport across the bacterial plasma membrane by the Sec machinery
require the unfolded conformation for the polypeptide substrate to be
translocated, because these processes are all compromised by a folded
DHFR domain (10-12). This situation, however, does not apply to all
cases of transport examined, because fully folded proteins can be
transported into mammalian peroxisomes and into Trypanosoma
brucei glycosomes (13, 14).
Secretory and most membrane proteins are inserted cotranslationally
into the ER and acquire their tertiary structure inside the ER lumen.
The translocation channel itself is believed capable of accommodating
polypeptide segments in an Here we show that a DHFR-HC fusion protein is recognized by US2 and
US11 for ER dislocation in vitro and in vivo.
DHFR retains the capacity to bind folate analogues in the lumen of
microsomes and in the ER of intact cells. Folding of the DHFR domain
does not impair dislocation of Class I MHC heavy chains in
vitro or in living cells. These results suggest that the
channel(s) used for dislocation can accommodate polypeptides that
contain a tightly folded domain of considerable size. Our data raise
the possibility that the Sec61 channel can be modified to accommodate a
folded DHFR domain for dislocation, but not for translocation into the ER, or that a channel distinct from Sec61 is used for dislocation.
Cell Lines and Antibodies--
U373-MG astrocytoma cells
(control), and US2 transfectants (US2+) were maintained as described
previously (17). DHFR-HC (U373DHFR-HC) cells and
US2-DHFR-HC (US2DHFR-HC) cells were maintained in
Dulbecco's modified Eagle's medium supplemented with 10% fetal calf
serum and 0.5 mg/ml G418 (Invitrogen, Frederick, MD). The cDNA Constructs and Transfection--
The DHFR-HC molecule
was cloned with the murine class I molecule H2-Kb signal
sequence at its N terminus and the class I heavy chain HLA-A2 allele
(amino acids 25-365) at the C terminus. The H2-Kb signal
sequence directs the chimeric molecule to the ER. The chimeric
constructs were inserted to pcDNA3.1 (Invitrogen, Carlsbad, CA).
The DHFR-HC was transfected into control and US2+ cells using LipofectAMINE (Invitrogen, Frederick, MD) and the standard protocol provided by the manufacturer.
In Vitro Transcription, Translation, and Degradation
Assay--
pcDNA3.1-DHFR-HC was linearized with SspI for
in vitro transcription. Transcription and translation were
performed as described previously (18). In vitro degradation
assays were carried out by translating the indicated mRNA in rabbit
reticulocyte lysates (RRLs) in the presence of microsomes as described
(18). Microsomes were spun down (20,000 × g, 15 min)
and resuspended in 10 µl of homogenization buffer (10 mM
HEPES, pH 7.4, 1 mM EDTA, 0.25 M sucrose)
containing 1 mg/ml RNase (Roche Molecular Biochemicals) with or without
0.1 mM TMX and incubated for 30 min at 30 °C. 15 µl of
Flexi RRL in the presence or absence of TMX containing 50 µM of the proteasome inhibitor ZLLL (MG132) were added,
and aliquots were withdrawn at the indicated time points. A microsomal pellet was obtained by centrifugation (20,000 × g, 15 min). Reducing sample buffer was added to the pellet, then it was
analyzed using SDS-PAGE.
In Vitro TMX Binding Experiments--
DHFR-HC mRNA was
translated in the presence of control microsomes. Microsomes were
resuspended in 10 µl of homogenization buffer and incubated for 30 min at 30 °C with or without TMX. To measure the extent to which
carryover of TMX with the pellet fraction occurred, unlabeled
microsomes were treated in a similar fashion. Microsomes were spun down
and washed once with 1 ml of cold homogenization buffer. Microsome
pellets were resuspended, combined, and lysed together in
homogenization buffer containing 1% Triton X-100 with or without
proteinase K, as indicated.
Metabolic Labeling of Cells, Pulse-chase Analysis,
Immunoprecipitation, and EndoH Digestion--
Cells were detached by
trypsin treatment, followed by starvation in methionine/cysteine-free
Dulbecco's modified Eagle's medium for 1 h. Cells were
metabolically labeled with 500 µCi/ml of
[35S]methionine/cysteine (1200 Ci/mmol, PerkinElmer Life
Sciences, Boston, MA) at 37 °C for the times indicated. Pulse-chase
experiments, cell lysis, and immunoprecipitation were performed as
described previously (19). The immunoprecipitates were analyzed by
SDS-PAGE followed by fluorography. Folate analogues were added during
the starvation, pulse, and chase periods. TMX (kind gift from
Medimmune, Gaithersburg, MD) was used at 0.1 mM, and MTX
(Aldrich, WI) was used at 10 µM. EndoH (New England
BioLabs, Beverly, MA) digestions were performed according to the
manufacturer's instructions.
Subcellular Fractionation--
Subcellular fractionation of
metabolically labeled US2DHFR-HC cells was
performed as described previously (17).
In Vivo TMX Binding Experiments--
One million
US2DHFR-HC cells were pulse-labeled for 15 min at 37 °C
with or without TMX. To measure the extent to which TMX binding to DHFR
occurred post-lysis rather than in the ER, the same number of unlabeled
U373 cells was exposed to TMX in the same manner. These cells were
washed twice with cold PBS, and the unlabeled cells were resuspended
and combined with pulse-labeled cells. The mixed unlabeled and
pulse-labeled cells were spun down and lysed together in 1 ml of
Nonidet P-40 0.5% lysis buffer. DHFR-HC was immunoprecipitated, and
resistance to proteinase K was assessed, as indicated. Carryover of TMX
from the unlabeled cells should then be able to stabilize
35S-labeled DHFR-HC not exposed to TMX in the course of
metabolic labeling. To achieve maximal binding of TMX to the DHFR
moiety, 0.1 mM TMX was added directly to lysates where
indicated. DHFR-HC was immunoprecipitated and one-half of the sample
was kept on ice, while the other half of each sample was treated with
proteinase K as described below. TMX binding to DHFR-HC was calculated
as the ratio between the 21-kDa digestion product and the intact DHFR-HC. The maximal binding control, in which TMX was added to lysates, was used to normalize for percentage of binding.
Proteinase K Digestion--
Microsomes were lysed in 30 µl of
homogenization buffer containing 1% Triton X-100. Proteinase K was
added to a final concentration of 100 µg/ml. Digestion was performed
on ice for 30 min. Phenylmethylsulfonyl fluoride (2 mM
final concentration) was added to inactivate proteinase K. Reducing
sample buffer was added, and samples were boiled and immediately loaded
onto the gel. Digestion of In Vitro Translated DHFR-HC Retains Folding Capacity in
Microsomes--
We generated a polypeptide composed of the
H-2Kb signal sequence, followed by DHFR fused in-frame to
the N terminus of HLA-A2 heavy chain. We first verified that DHFR, a
cytosolic enzyme, can fold properly in the lumen of the ER, where both
redox potential and calcium concentrations differ significantly from
those in the cytosol. The H-2Kb-DHFR-HC construct was
translated in vitro in the presence of microsomes derived
from U373 cells (Fig. 1). Following
isolation of the microsomal fraction by centrifugation, we added
trimetrexate (TMX), a membrane-permeable folate analogue, to the
microsomes. Microsomes were then extensively washed to eliminate traces
of unbound drug, lysed in 1% Triton X-100, and subjected to proteinase K digestion. The generation of a 21-kDa DHFR fragment resistant to
proteinase K digestion is indicative of binding of the folate analogue
and stabilization of the tightly folded conformation of DHFR. Inclusion
of TMX renders the DHFR portion of the DHFR-HC fusion resistant to
proteinase K digestion, as judged by the appearance of a 21-kDa
polypeptide only when microsomes were preincubated with TMX (Fig. 1,
lanes 2 and 3). To exclude the possibility of binding of TMX to DHFR after lysis, rather than in intact microsomes, we incubated a separate preparation of microsomes with TMX, washed the
pellet, and lysed these microsomes together with DHFR-HC-containing microsomes that had not been exposed to TMX. Digestion of the DHFR-HC
fusion by proteinase K was complete in this case (Fig. 1, lane
4), excluding the possibility of carryover of the drug. Hence,
DHFR-HC inserted in the ER folds properly and acquires a tight
conformation when bound to a folate analogue.
DHFR-HC Dislocates in Vitro in a US2- or US11-dependent
Manner--
We next examined the ability of DHFR-HC to undergo
US2/US11-mediated degradation. In previous work we established an
in vitro system that recapitulates US2/US11-mediated
degradation. The hallmark of this in vitro reaction is the
disappearance of glycosylated HLA-A2 HC from the pellet fraction in a
US2- or US11-dependent manner. Over time, we observe the
appearance of a soluble deglycosylated intermediate released from the
microsomal fraction, when incubation is carried out in the presence of
the proteasome inhibitor ZLLL (18). We applied the same in
vitro system to DHFR-HC. DHFR-HC mRNA was translated in the
presence of microsomes derived from control or US2- or US11-expressing
cells (Fig. 2A). In all three types of microsomes some of the glycosylated DHFR-HC was detected in
the supernatants, probably due to inefficient separation of the pellet
and supernatant fractions by the conditions of centrifugation used.
Therefore, as an indicator of dislocation we considered both the loss
of glycosylated DHFR-HC from the pellet fraction and the appearance of
the deglycosylated intermediate over time in the supernatant fraction.
We did not observe the appearance of a deglycosylated intermediate in
the supernatants of U373-derived microsomes (Fig. 2A,
Supernatants, lanes 1 and 2). However,
when DHFR-HC was translated into microsomes derived from US2+ and US11+ cells, a polypeptide with higher mobility appeared at the 60-min chase
point. This polypeptide corresponds to deglycosylated DHFR-HC and is
indicative of a US2/US11-dependent dislocation reaction (Fig. 2A, Supernatants, lanes 5,
6, 9, and 10). As for HLA-A2 HC (18),
US2+ microsomes support better the dislocation of DHFR-HC than US11+
microsomes, as measured by disappearance from the microsome fraction
(Fig. 2B) and accumulation of the deglycosylated
intermediate in the supernatants (Fig. 2A,
Supernatants, lanes 6 and 8 versus lanes 10 and 12).
We then measured whether formation of a tightly folded domain inside
the ER lumen would impede dislocation. Microsomes were incubated at
30 °C with TMX 30 min prior to the chase to ensure efficient binding
of the drug to the DHFR-HC protein. We did not detect inhibition of
dislocation of DHFR-HC from microsomes treated with TMX. In fact, we
consistently observed a slight but reproducible increase in the amount
of deglycosylated intermediate released from US2+ microsomes but not
for US11+ microsomes (Fig. 2A, Supernatants, lane 8 versus 6). We examined a more
detailed time course for dislocation of DHFR-HC from US2+ microsomes.
At all chase points, the deglycosylated intermediate was more abundant
for microsomes treated with TMX when compared with controls (Fig.
2C, lanes 6 versus 2 and 7 versus 3). We conclude that the compactly folded DHFR moiety
does not impede dislocation of the DHFR-HC fusion.
DHFR-HC Is Stable in U373 Cells and Largely Retained within the
ER--
Unassembled class I MHC heavy chains are recognized by the
quality control machinery and are subject to dislocation followed by
proteasome-dependent degradation (20, 21). It was therefore necessary to verify that DHFR-HC does not dislocate spontaneously in
control cells. U373 cells were transfected with DHFR-HC, and the
stability of the fusion protein was monitored by pulse-chase analysis.
We also assessed intracellular transport of the fusion protein by
performing EndoH digestion. DHFR-HC was stable in control cells even
after 3 h of chase (Fig. 3,
lanes 1, 3, and 5). While endogenous
class I MHC acquired EndoH resistance at 90 min. of chase, the DHFR-HC
fusion remains EndoH sensitive even after 3 h of chase (Fig. 3,
lanes 2, 4, and 6). Inclusion of TMX
affected neither stability nor EndoH sensitivity of the protein (Fig.
3, lanes 7-12). We conclude that the DHFR-HC is relatively
stable and fails to leave the ER and that the stability of DHFR-HC is not affected by DHFR ligands.
TMX Binds DHFR-HC in the ER Lumen--
To demonstrate binding of
TMX to the DHFR domain of DHFR-HC fusion inside the ER lumen,
US2DHFR-HC cells were pulsed for 15 min. in the presence or
in the absence of TMX followed by immunoprecipitation and treatment
with proteinase K. Under these conditions, radiolabeled DHFR-HC
corresponds to ER-disposed protein. To control for binding of TMX to
the DHFR moiety post-lysis and to assess the efficiency of TMX binding,
we performed the experiment as outlined in Fig. 4A. Pulse-labeled cells not
exposed to TMX were washed and mixed with the same number of unlabeled
cells incubated with TMX. If binding of TMX to the DHFR-HC fusion would
occur post-lysis by carryover of the drug, one should expect to see
stabilization of labeled fusion protein obtained from cells not exposed
to TMX (Fig. 4A, lanes 1 and 2). As a
negative control, 35S-labeled US2DHFR-HC cells
were mixed with unlabeled cells not exposed to TMX (Fig. 4A,
lanes 3 and 4). Upon proteinase K digestion of
anti-heavy chain immunoprecipitates exposed post-lysis to TMX, a 21-kDa
band appeared, indicating some carryover of TMX (Fig. 4B,
lane 2). In addition, we included a control for maximal
binding of TMX to the DHFR moiety (Fig. 4A, lanes
7 and 8). US2DHFR-HC cells were
pulse-labeled in the presence of TMX, washed, and lysed. We then added
TMX directly to the lysates to obtain maximal exposure of the DHFR
domain to TMX. Under these conditions DHFR was stabilized significantly
better (Fig. 4B, lane 8). We quantified the
binding of each condition by calculating the ratio of the 21-kDa
digestion product over the intact DHFR-HC. The ratio for maximal
binding, in which TMX was added to lysates, was defined as 100%
binding. Binding of TMX to DHFR-HC was 86.5% of the maximal binding
observed in a detergent lysate and at least 3-fold higher than binding post-lysis (Fig. 4C). These results indicate that
binding of TMX to the DHFR domain of DHFR-HC fusion occurs
pre-lysis inside the ER lumen.
Rate of Dislocation of DHFR-HC in Cells Is Not Affected by DHFR
Folding--
We performed pulse-chase experiments to monitor the
dislocation of DHFR-HC coexpressed with US2. We saw that
pulse-labeled DHFR-HC expressed in US2DHFR-HC disappeared
completely within 1 h of chase (not shown). In the presence of
proteasome inhibitor, glycosylated DHFR-HC was converted to the
deglycosylated intermediate (Fig.
5B). To test whether TMX
affects the dislocation rate in live cells, US2DHFR-HC
cells were pulse-labeled for 5 min and chased for the indicated times
in the absence (Fig. 5A) or presence (Fig. 5B) of
proteasome inhibitors. In the absence of proteasome inhibitors, DHFR-HC
was mostly degraded after 20 min of chase (Fig. 5A,
lanes 1-3). In the presence of the proteasome inhibitor ZL3VS, we observed the conversion of DHFR-HC to the
deglycosylated intermediate (Fig. 5B, lanes
10-12). As a control, we treated cells with MTX, for which its
hydrophilicity should exclude it from the ER, to account for possible
effects downstream of dislocation, which might include the inability of
folded DHFR to be targeted for proteasomal degradation as efficiently
as DHFR not exposed to MTX (22). We did not observe any effect of TMX
or MTX on the dislocation rate of DHFR-HC, regardless of the presence
of proteasome inhibitors (Fig. 5A, lanes 4-9 and
Fig. 5B, lanes 13-18). If extremely rapid
unfolding occurred at 37 °C, then perhaps a chase at room
temperature might slow down the process. Even at room temperature,
DHFR-HC was dislocated at a rate similar to that seen for the
endogenous HC. We did not detect any effect on dislocation upon
inclusion of TMX (not shown). These data are consistent with the
hypothesis that complete unfolding is not required for dislocation.
In keeping with our earlier observations, inclusion of TMX did not
change the subcellular localization of the DHFR-HC fusion. The
deglycosylated fusion product fractionated with the cytosolic compartment, irrespective of the folded state of the DHFR moiety (Fig.
5C, lane 8 versus 6). We
conclude that the DHFR-HC fusion behaves like the endogenous Class I
heavy chains as far as US2-mediated dislocation is concerned.
Importantly, the folded state of the DHFR domain does not interfere
with ER dislocation.
DHFR-HC Is Dislocated in a Folded State--
To exclude the
possibility that rapid unfolding in the ER precedes dislocation despite
the binding of TMX, we examined the resistance of the DHFR-HC fusion
protein to proteinase K when chased in the presence or absence of TMX.
US2DHFR-HC cells were pulse-labeled for 15 min in the
presence of TMX, washed twice with PBS, and chased for 60 min to ensure
complete conversion of glycosylated DHFR-HC to the deglycosylated
intermediate, a reaction that occurs in the cytoplasm. Following TMX
removal, cells were chased in the presence (Fig.
6A, lanes 11 and
12) or absence (lanes 9 and 10) of
TMX. DHFR-HC was immunoprecipitated and digested with proteinase K. To
achieve maximal binding of TMX to the DHFR moiety, TMX was included
throughout the pulse and chase as well as added into the lysis buffer
(Fig. 6A, lanes 13 and 14). This level
of binding was defined as 100% binding. Resistance of DHFR-HC
to proteinase K when TMX was present during the pulse and chase was
72% of maximal binding (Fig. 6B), 62% when TMX was present
only during the pulse period, and removed by washing prior to the
60-min chase. We also measured the binding of TMX to DHFR-HC during the
pulse. US2DHFR-HC cells were pulse-labeled for 15 min in
the presence of TMX. Cells were washed twice with PBS, lysed, and
subjected to immunoprecipitation and proteinase K digestion (Fig.
6A, lanes 3 and 4). Addition of TMX
during the pulse stabilized DHFR-HC to 75% of level seen when TMX was
added to the lysate (Fig. 6A, lanes 5 and
6, and Fig. 6B). Neither ER-resident DHFR-HC nor
the deglycosylated intermediate were stabilized when TMX was omitted
(Fig. 6A, lanes 1 and 2, and
7 and 8). The similar levels of stabilization of
DHFR-HC conferred by TMX in all three conditions indicates that binding
of TMX to the DHFR domain of DHFR-HC fusion during the pulse accounts
for most of the binding observed at the end of the chase period when
TMX is removed by washing. Therefore, TMX binds to DHFR in the ER
and interacts stably with DHFR-HC in the course of dislocation.
The US2-mediated dislocation of Class I MHC heavy chains from the
ER involves a viral product that interacts with the correctly folded
protein, as inferred from the crystal structure of the US2-HC· In the course of dislocation the Class I HC undergoes disulfide
bond reduction prior to deglycosylation, suggesting an unfolding step
to assure smooth threading of the substrate through the translocon and
the proteasome (17). We have shown that an EGFP-HC fusion is also a
substrate for US2- and US11-dependent dislocation (7). The
results of these experiments suggest the possibility that folded,
fluorescent EGFP can be transported across the ER membrane. The EGFP-HC
model, however, does not allow control over the folded state of EGFP.
It is difficult to exclude the formal possibility that partial
unfolding of the EGFP-HC fusion is required for dislocation. Partial
unfolding might be followed by rapid refolding, thus accounting for
fluorescent EGFP-HC species in the cytosol.
To address the necessity of unfolding for proteins to be dislocated
from the ER, we chose DHFR as a fusion partner to Class I HC. In the
presence of folate analogues DHFR acquires a tightly folded, highly
protease-resistant conformation. Thus, the inclusion or omission of
these small molecules allows control over the stability of the DHFR
moiety. Furthermore, DHFR fusions have been used extensively to address
questions pertaining to protein translocation. The best studied
examples concern proteins destined for import into mitochondria and
peroxisomes. DHFR fused to a mitochondrial signal peptide fails to be
imported when the compactly folded state is imposed on DHFR by
inclusion of folate analogues (9). Curiously, DHFR targeted to the TAT
translocation system of chloroplasts is still translocated across the
thylakoid despite the presence of MTX (26). This supports the notion
that the machinery responsible for TAT-mediated translocation
apparently can transport proteins in a fully folded state. Therefore,
assaying DHFR transport in the presence or absence of folate analogues
allows an assessment of how folding affects the translocation of a
DHFR-containing reporter substrate.
The DHFR moiety, however, has not been used as a fusion partner to
study export from a membrane-delimited compartment, mainly owing to
limitations of membrane permeability of folate analogues such as MTX.
Therefore we used TMX, a lipophilic DHFR inhibitor in use as an
anti-protozoan drug (27), to impose tight folding of an
ER-localized DHFR domain. The affinity of TMX for murine DHFR is
~1-10 µM. We used a concentration of 100 µM to ensure efficient (over 90%) binding. At this
concentration, TMX interacted with and induced stable folding of
DHFR-HC expressed in microsomes, as assessed by resistance to
proteinase K (Fig. 1).
First we tested the effects of folate analogues on the dislocation of
DHFR-HC in vitro, using an assay that recapitulates US2- and
US11-mediated dislocation. We expected that if unfolding of the
dislocation substrate is required, enforcing a folded conformation of
the DHFR domain should slow down dislocation or even abrogate it, as
shown for import of DHFR fusions into mitochondria. In fact, for US2+
microsomes but not for US11+ microsomes we observed, if anything, an
enhancement of dislocation, as judged by greater accumulation of the
diagnostic soluble deglycosylated intermediates at the early time
points of chase (Fig. 2C). These data suggest potentiation
rather than inhibition of dislocation by the presence of a tightly
folded DHFR moiety. Because acceleration was seen only in US2+ and not
in US11+ microsomes, it is unlikely that this effect is due to events
downstream of dislocation, such as activation of peptide
N-glycanase or other factors involved in the
generation of the soluble intermediate. This selectivity, observed when
DHFR ligands are included for US2-mediated dislocation, could therefore
be attributed to the different mechanisms utilized by US2 and US11 to
degrade class I HC. For example, US2 dislocates the HC in a manner that
is rather independent of the HC cytosolic tail sequence, whereas
US11-mediated dislocation requires a complete HC cytoplasmic tail (18).
Furthermore, a tailless US2 is incapable of dislocating HC, whereas
tailless US11 retains the capacity to dislocate the HC (18). Therefore,
it is conceivable that folding of the DHFR domain in the ER lumen
facilitates the interaction with US2 either directly or indirectly.
Nonetheless, for neither US2- nor US11-mediated dislocation does the
folding of the luminal DHFR domain inhibit the transfer of the DHFR-HC
fusion protein from the membrane fraction to the soluble fraction.
Next, we verified these results in live cells. The DHFR-HC was stably
transfected into control and US2+ astrocytoma cells. We found that
DHFR-HC is highly stable in control U373 cells and is retained largely
in the ER (Fig. 3). In vivo binding of TMX to DHFR in the ER
was assessed by measuring proteinase K resistance and found to be as
efficient as binding to solubilized DHFR-HC (Fig. 4). Control
experiments excluded carryover and post-lysis binding of the drug as an
explanation for the observed proteinase K resistance. In US2+ cells,
the DHFR-HC was dislocated rapidly, with kinetics similar to those seen
for endogenous HC (Fig. 5). We could not show a significant
acceleration of dislocation upon DHFR folding, as we observed in
vitro. This observation is most likely attributable to the more
rapid kinetics of dislocation in vivo. Regardless, both in
TMX- or MTX-treated cells, dislocation occurs at the same rate. To
establish that dislocation had in fact occurred in the manner seen for
the endogenous HC, we also verified the transport of DHFR-HC from the
ER membrane to the cytoplasm by subcellular fractionation. Following a
15-min pulse label, glycosylated DHFR-HC and glycosylated endogenous
heavy chain were retrieved from the pellet fraction. After a 60-min chase in the presence of proteasome inhibitor, the deglycosylated DHFR-HC fusion protein accumulated mainly in the cytosolic fraction, regardless of the inclusion of TMX (Fig. 5C), and similar to
the behavior of endogenous heavy chains. Therefore, tight folding of
the DHFR moiety did not impose any constraints on dislocation. To
exclude the remote possibility of rapid unfolding of the DHFR moiety,
an unlikely event in view of the demonstrated binding of a folate
analogue, and dislocation as an unfolded polypeptide, we compared the
degree of binding of TMX to DHFR when cells are pulse-labeled in the
presence of TMX and chased in the presence or absence TMX. The
stabilization of DHFR in both cases was similar and comparable to the
binding of TMX to DHFR during the chase period (Fig. 6). We conclude,
therefore, that the vast majority of DHFR-HC molecules dislocate as a
protein that retains TMX binding.
Proteins can be translocated across membranes in a fully folded state,
while the membrane compartment in question maintains a gradient of low
mw solutes. Perhaps the most striking example for this phenomenon is
the peroxisomal import of colloidal gold particles conjugated to
proteins bearing the peroxisomal targeting signal (13). Therefore,
dislocation from the ER into the cytoplasm may be regulated in a way
that prevents leakage of other ER contents into the cytosol, yet can
accommodate folded proteins.
Several lines of biochemical and genetic evidence support the
involvement of the Sec61 translocon as the channel through which proteins are exported back to cytosol. In yeast, degradation of misfolded ER proteins is tightly coupled to the unfolded protein response. In a genetic screen that exploited this coupling, several alleles of Sec61 were reported to significantly inhibit the dislocation of CPY*, a misfolded ER protein, with minimal disturbance of ER import
(28). These alleles, however, were found to support ER degradation of
Ubc6 (29) and the mouse class I MHC allele
H-2Kb.2 These
findings suggest the existence of alternative routes for dislocation
that may not involve Sec61. An alternative mechanism for ER dislocation
might involve polyubiquitination of the substrate and extraction from
the lipid bilayer in an ATP-dependent manner. Although this
might be a plausible mechanism for Ubc6, which is tail-anchored to the
ER membrane with most of the protein facing the cytoplasm, it is more
difficult to envisage such a mechanism for a classic type I membrane
proteins like class I MHC products. Moreover, K Biochemical studies demonstrate that dislocated HC associates with
Sec61 The example of the mechanosensitive channel MscL shows how adjustment
of the packing angle of the transmembrane helical segments can result
in a change from an almost completely sealed pore to a cannel with a
pore size of over 25 Å (31). This type of rearrangement might be used
more generally to control permeability across a biological membrane,
such as the ER. The maximum pore size seen for the modestly sized MscL
channel might be significantly smaller than that for the multisubunit
assemblies such as Sec61, for which changes in pore size could be even
more dramatic. A recent study by Helenius and coworkers (32)
demonstrated that nascent polypeptides can fold partially within the
cavity of the translocon pore when forced to fold in close proximity to
surface of the ER membrane. Conformational changes of this type might
reconcile our findings with the prevailing view that proteins are
dislocated through the Sec61 translocon, or through any other channel,
in a largely unfolded state. Although tightly folded DHFR cannot be
inserted into the ER lumen through the Sec61 translocon, the spatial
restrictions for dislocation from the ER might be different.
Fluorescence quenching experiments indicate a diameter for the Sec61
pore of up to 60 Å during active protein translocation (33). A dilated
pore of that size is sufficient to accommodate a fully folded DHFR
moiety, the smallest cross-section of which is estimated to be 40 Å.
The translocon, upon engagement of a dislocation substrate, may acquire a dilated conformation through which even folded proteins can be
efficiently dislocated. Accordingly, we propose that proteins can
dislocate in a partially folded state with the retention of considerable tertiary structure. This obviates the need for complete unfolding in the ER prior to dislocation and might explain how toxins
that travel from the ER to the cytoplasm retain their toxicity (34,
35). Because folded proteins can apparently be exported from, but not
imported into the ER, the dislocation process cannot be described as ER
translocation in reverse, but must require a distinct conformation of
the known translocon or different channels altogether.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-helical configuration. However,
completion of folding is thought to take place in the ER lumen (15).
Consistent with these findings, the translocation of a hybrid between
the presecretory protein, preprocecropin A, and DHFR (ppcecDHFR) into
mammalian microsomes is signal peptide-dependent and will
proceed post-translationally. Although membrane insertion and signal
peptide removal of the preprocecropin moiety did not require unfolding
of the DHFR domain, completion of translocation required unfolded DHFR,
because translocation was blocked by inclusion of MTX. When MTX is
present, and hence the DHFR portion of the ppcecDHFR is compactly
folded, ppcecDHFR is susceptible to proteinase K digestion when
translated in the presence of microsomes. It was concluded that folded
MTX-occupied DHFR is unable to fit into the Sec61 aqueous pore to allow
import into the ER (16).
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
HC serum
was generated by immunizing rabbits with a bacterially expressed
luminal fragment of HLA-A2 and HLA-B27 heavy chain.
HC immunoprecipitates was done in a
similar fashion. Immunoprecipitates were washed twice with NET buffer
(Nonidet P-40 0.5%, NaCl 150 mM, EDTA 5 mM,
Tris (pH 7.4) 50 mM), 30 µl of homogenization buffer
containing 1% Triton X-100 was added, and proteinase K was added to a
final concentration of 100 µg/ml at room temperature for 45 min.
Phenylmethylsulfonyl fluoride (2 mM) was added, and an
equal volume of 2-fold concentrated reducing sample buffer was added.
Samples were boiled and immediately loaded onto the gel.
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
View larger version (57K):
[in a new window]
Fig. 1.
The DHFR-HC fusion is inserted into human
astrocytoma microsomes and binds TMX within the lumen of the
microsomes. DHFR-HC mRNA was translated in the presence of
U373 cells microsomes. Microsomes were isolated by centrifugation and
incubated in the absence (lanes 1, 2, and
4) or in the presence of TMX (lane 3). Microsomes
were isolated by centrifugation and washed twice to remove unbound TMX.
Unlabeled microsomes treated with TMX and washed in the same manner
were mixed with the labeled microsomes that were not exposed to TMX
(lane 4). Microsomes were lysed in 1% Triton X-100 and
digested with proteinase K (lanes 2-4). Only DHFR-HC that
was incubated with TMX was protected from proteinase K digestion as
indicated by the 21-kDa polypeptide (lane 3). There is no
indication for post-lysis binding of TMX to DHFR-HC (lane
4).
View larger version (48K):
[in a new window]
Fig. 2.
DHFR-HC is subject to US2- and US11-mediated
degradation in vitro. Dislocation is not
inhibited by a folded DHFR. A, DHFR-HC mRNA was
translated into microsomes derived from control (lanes
1-4), US2-transfected (lanes 5-8), or
US11-transfected (lanes 9-12) U373 cells for 45 min.
Microsomes were isolated by centrifugation and were resuspended in
homogenization buffer containing RNase in the presence or absence of
TMX. Following 30-min incubation with TMX, Flexi RRL containing 50 µM ZLLL was added to initiate dislocation. Aliquots were
withdrawn at the indicated times. Microsomes and supernatants were
analyzed by SDS-PAGE (12%). B, autoradiograms were
quantitated with ALPHAIMAGER software (Alpha Innotech, San Leandro,
CA). Band intensities in the pellet fractions were plotted as
percentages of DHFR-HC remaining relative to intensity at 0-min chase
point. C, DHFR-HC mRNA was translated into microsomes
derived from US2-transfected U373 cells for 45 min. Samples were
treated with TMX as in A. Aliquots were withdrawn at the
indicated times, and supernatants were analyzed by SDS-PAGE
(12%).
View larger version (35K):
[in a new window]
Fig. 3.
DHFR-HC is stable in U373 cells, and it is
largely retained within the ER. U373DHFR-HC cells were
pulse-labeled with [35S]methionine for 15 min and chased
up to 180 min. Cells were lysed in 1% SDS then diluted to 0.07% SDS
with Nonidet P-40 lysis mix followed by immunoprecipitation with
anti-class I heavy chain serum ( HC). Half of the immunoprecipitates
were digested with EndoH. Samples were analyzed by SDS-PAGE (12%). TMX
was added 1 h before the metabolic labeling throughout the chase
(lanes 7-12).
View larger version (39K):
[in a new window]
Fig. 4.
TMX binds DHFR-HC within the ER lumen.
A, US2DHFR-HC cells were pulse-labeled with
[35S]methionine for 15 min in the absence (lanes
1-4) or in the presence (lanes 5-8) of TMX. Cells
were washed with cold PBS and mixed with an equal number of unlabeled
cells incubated in the presence (lanes 1 and 2)
or in the absence (lanes 3 and 4) of TMX. Cells
were lysed in Nonidet P-40 lysis buffer. TMX was added directly to
lysates (lanes 7 and 8). Lysates were
immunoprecipitated with anti-class I heavy chain serum ( HC). Half of
the immunoprecipitates were digested with proteinase K. B,
samples were analyzed by SDS-PAGE (12%). C, autoradiograms
were quantitated with ALPHAIMAGER software (Alpha Innotech, San
Leandro, CA). Binding of TMX to DHFR-HC was calculated from the ratio
of the band intensity of proteinase K digested DHFR-HC (lanes
2, 4, 6, and 8) relative to
intact DHFR-HC (upper band, lanes 1,
3, 5, and 7), respectively. Binding
was plotted as percentages of the ratio derived for maximal binding
(21-kDa fragment in lane 8 relative to DHFR-HC in lane
7).
View larger version (67K):
[in a new window]
Fig. 5.
US2-mediated dislocation of DHFR-HC in live
cells is not affected by DHFR folding. US2DHFR-HC
cells were pulse-labeled with [35S]methionine for 5 min
and chased up to 20 min in the absence (A) or in the
presence (B) of the proteasome inhibitor, ZL3VS.
TMX (lanes 4-6 and 13-15) or MTX (lanes
7-9 and 16-18) were added 1 h prior to metabolic
labeling throughout the chase. Cells were lysed in 1% SDS, diluted to
0.07% SDS with Nonidet P-40 lysis mix, and then followed by
immunoprecipitation with anti-class I heavy chain serum. The
immunoprecipitates were analyzed by SDS-PAGE (12%). C,
US2DHFR-HC cells were pulse-labeled with
[35S]methionine for 15 min and chased for 60 min in the
presence of the proteasome inhibitor, ZL3VS. TMX
(lanes 3, 4, 7, and 8) was
added 1 h prior to metabolic labeling throughout the chase. Cells
were homogenized and subjected to fractionation as described under
"Experimental Procedures." The DHFR-HC and endogenous HC were
recovered from the 100-kg pellet (p) and the 100-kg
supernatant (s) by immunoprecipitation with anti-class I
heavy chain serum. The immunoprecipitates were analyzed by SDS-PAGE
(12%).
View larger version (39K):
[in a new window]
Fig. 6.
DHFR-HC dislocates as a folded protein.
A, US2DHFR-HC cells were pulse-labeled with
[35S]methionine for 15 min in the absence (lanes
1-6) or in the presence (lanes 7-14) of
ZL3VS. Cells were washed twice with cold PBS and chased in
the presence (lanes 11-14) or in the absence (lanes
7-10) of TMX. Cells were lysed in Nonidet P-40 lysis buffer. TMX
was added directly to lysates (lanes 5 and 6, and
13 and 14). Lysates were immunoprecipitated with
anti-class I heavy chain serum ( HC). Half of the immunoprecipitates
were digested with proteinase K, and samples were analyzed by SDS-PAGE
(12%). B, autoradiograms were quantitated with ALPHAIMAGER
software (Alpha Innotech, San Leandro, CA). Binding of TMX to DHFR-HC
was calculated from the ratio of the band intensity of proteinase
K-digested DHFR-HC (lanes 2, 4,
6, 8, 10, 12, and
14) relative to intact DHFR-HC (upper band,
lanes 1, 3, 5, 7,
9, 11, and 13), respectively. Binding
was plotted as percentages of the ratio derived for maximal binding
(21-kDa fragment in lane 6 is relative to DHFR-HC
in lane 5 for pulse and 21-kDa fragment in lane
14 is relative to DHFR-HC in lane 13 for chase).
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
2m complex (23). This interaction diverts the
HC and US2 itself with remarkably rapid kinetics (half-life of 3-5
min) back to the cytosol for degradation by the proteasome. When the proteasome is blocked, the HC accumulates in the cytosol as a deglycosylated intermediate (24, 25). The ability of the class I
molecule to fold and interact with US2 and the extraordinarily rapid
rate of dislocation thus make the US2-Class I MHC interaction a
suitable model to study spatial and conformational constraints on dislocation.
R mutations that
remove all lysines from the class I cytosolic tail do not prevent
US11-mediated dislocation of HC, whereas these mutants obviously cannot
be ubiquitinated in their cytoplasmic tail (6).
in US2+ cells or by dithiothreitol treatment in control cells
(24). In addition, elegant cross-linking experiments show that Sec61
interacts directly with ApoB100 destined for ER degradation. In fact,
ApoB100 lingers as a nascent protein until assembly with lipids is
completed, and degradation is initiated on the nascent chain (30). This
model is different from what we propose for US2-mediated degradation.
US2 recognizes fully folded class I MHC as indicated by
coimmunoprecipitation of US2 and Class I heavy chains with W6/32, an
antibody that recognizes fully assembled MHC (24). Furthermore,
Daudi-derived microsomes, which lack the MHC light chain
2m, do not support dislocation in vitro.
Dislocation ensues, however, when microsomes are reconstituted with
2m.3 These
findings suggest that dislocation of Class I heavy chains is initiated
by US2 for Class I molecules that must have left the translocon
already. Either the translocon is re-engaged subsequently, or an
altogether different channel is used for dislocation. The mechanism by
which Class I is recruited to such channels remains to be identified.
![]() |
FOOTNOTES |
---|
* This work was supported in part by National Institutes of Health Grant 5R37-AI33456.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 Dorot Foundation fellowship.
§ To whom correspondence should be addressed: Dept. of Pathology, Harvard Medical School, 200 Longwood Ave. Armenise Bldg., Rm. 137, Boston, MA 02115. Tel.: 617-432-4777; Fax: 617-432-4775; E-mail: hidde_ploegh@hms.harvard.edu.
Published, JBC Papers in Press, December 12, 2002, DOI 10.1074/jbc.M210158200
2 R. Casagrande and H. L. Ploegh, unpublished data.
3 M. Furman, unpublished observations.
![]() |
ABBREVIATIONS |
---|
The abbreviations used are:
ER, endoplasmic
reticulum;
MHC, major histocompatibility complex;
HC, class I MHC heavy
chain;
DHFR, dihydrofolate reductase;
MTX, methotrexate;
TMX, trimetrexate;
EGFP, enhanced green fluorescent protein;
ppcecDHFR, hybrid between preprocecropin A and DHFR;
RRL, rabbit reticulocyte
lysate;
EndoH, endoglycosidase H;
PBS, phosphate-buffered saline;
ZLLL, N-benzyloxycarbonyl-Leu-Leu-Leu-aldehyde;
ZL3VS, N
-benzyloxycarbonyl-Leu-Leu-Leu-vinyl
sulfone.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. | Heinrich, S. U., Mothes, W., Brunner, J., and Rapoport, T. A. (2000) Cell 102, 233-244[Medline] [Order article via Infotrieve] |
2. | Tsai, B., Ye, Y., and Rapoport, T. A. (2002) Nat. Rev. Mol. Cell. Biol. 3, 246-255[CrossRef][Medline] [Order article via Infotrieve] |
3. | Huppa, J. B., and Ploegh, H. L. (1997) Immunity 7, 113-122[Medline] [Order article via Infotrieve] |
4. | Wiertz, E. J., Jones, T. R., Sun, L., Bogyo, M., Geuze, H. J., and Ploegh, H. L. (1996) Cell 84, 769-779[Medline] [Order article via Infotrieve] |
5. | Ye, Y., Meyer, H. H., and Rapoport, T. A. (2001) Nature 414, 652-656[CrossRef][Medline] [Order article via Infotrieve] |
6. |
Shamu, C. E.,
Story, C. M.,
Rapoport, T. A.,
and Ploegh, H. L.
(1999)
J. Cell Biol.
147,
45-58 |
7. |
Fiebiger, E.,
Story, C.,
Ploegh, H. L.,
and Tortorella, D.
(2002)
EMBO J.
21,
1041-1053 |
8. | Sasso, S. P., Gilli, R. M., Sari, J. C., Rimet, O. S., and Briand, C. M. (1994) Biochim. Biophys. Acta 1207, 74-79[Medline] [Order article via Infotrieve] |
9. | Eilers, M., and Schatz, G. (1986) Nature 322, 228-232[Medline] [Order article via Infotrieve] |
10. |
Salvador, N.,
Aguado, C.,
Horst, M.,
and Knecht, E.
(2000)
J. Biol. Chem.
275,
27447-27456 |
11. | Endo, T., Kawakami, M., Goto, A., America, T., Weisbeek, P., and Nakai, M. (1994) Eur. J. Biochem. 225, 403-409[Abstract] |
12. | Arkowitz, R. A., Joly, J. C., and Wickner, W. (1993) EMBO J. 12, 243-253[Abstract] |
13. | Walton, P. A., Hill, P. E., and Subramani, S. (1995) Mol. Biol. Cell 6, 675-683[Abstract] |
14. | Hausler, T., Stierhof, Y. D., Wirtz, E., and Clayton, C. (1996) J. Cell Biol. 132, 311-324[Abstract] |
15. | Mingarro, I. I., Nilsson, I. I., Whitley, P., and von Heijne, G. (2000) BMC Cell. Biol. 1, 3[CrossRef][Medline] [Order article via Infotrieve] |
16. | Schlenstedt, G., Zimmermann, M., and Zimmermann, R. (1994) FEBS Lett. 340, 139-144[CrossRef][Medline] [Order article via Infotrieve] |
17. |
Tortorella, D.,
Story, C. M.,
Huppa, J. B.,
Wiertz, E. J.,
Jones, T. R.,
Bacik, I.,
Bennink, J. R.,
Yewdell, J. W.,
and Ploegh, H. L.
(1998)
J. Cell Biol.
142,
365-376 |
18. |
Furman, M. H.,
Ploegh, H. L.,
and Tortorella, D.
(2002)
J. Biol. Chem.
277,
3258-3267 |
19. |
Rehm, A.,
Stern, P.,
Ploegh, H. L.,
and Tortorella, D.
(2001)
EMBO J.
20,
1573-1582 |
20. | Raposo, G., van Santen, H. M., Leijendekker, R., Geuze, H. J., and Ploegh, H. L. (1995) J. Cell Biol. 131, 1403-1419[Abstract] |
21. |
Hughes, E. A.,
Hammond, C.,
and Cresswell, P.
(1997)
Proc. Natl. Acad. Sci. U. S. A.
94,
1896-1901 |
22. |
Johnston, J. A.,
Johnson, E. S.,
Waller, P. R.,
and Varshavsky, A.
(1995)
J. Biol. Chem.
270,
8172-8178 |
23. |
Gewurz, B. E.,
Gaudet, R.,
Tortorella, D.,
Wang, E. W.,
Ploegh, H. L.,
and Wiley, D. C.
(2001)
Proc. Natl. Acad. Sci. U. S. A.
98,
6794-6799 |
24. | Wiertz, E. J., Tortorella, D., Bogyo, M., Yu, J., Mothes, W., Jones, T. R., Rapoport, T. A., and Ploegh, H. L. (1996) Nature 384, 432-438[CrossRef][Medline] [Order article via Infotrieve] |
25. | Jones, T. R., and Sun, L. (1997) J. Virol. 71, 2970-2979[Abstract] |
26. |
Hynds, P. J.,
Robinson, D.,
and Robinson, C.
(1998)
J. Biol. Chem.
273,
34868-34874 |
27. | Piper, J. R., Johnson, C. A., Krauth, C. A., Carter, R. L., Hosmer, C. A., Queener, S. F., Borotz, S. E., and Pfefferkorn, E. R. (1996) J. Med. Chem. 39, 1271-1280[CrossRef][Medline] [Order article via Infotrieve] |
28. | Zhou, M., and Schekman, R. (1999) Mol. Cell 4, 925-934[Medline] [Order article via Infotrieve] |
29. |
Walter, J.,
Urban, J.,
Volkwein, C.,
and Sommer, T.
(2001)
EMBO J.
20,
3124-3131 |
30. |
Pariyarath, R.,
Wang, H.,
Aitchison, J. D.,
Ginsberg, H. N.,
Welch, W. J.,
Johnson, A. E.,
and Fisher, E. A.
(2001)
J. Biol. Chem.
276,
541-550 |
31. | Perozo, E., Cortes, D. M., Sompornpisut, P., Kloda, A., and Martinac, B. (2002) Nature 418, 942-948[CrossRef][Medline] [Order article via Infotrieve] |
32. | Kowarik, M., Kung, S., Martoglio, B., and Helenius, A. (2002) Mol. Cell 10, 769-778[Medline] [Order article via Infotrieve] |
33. | Hamman, B. D., Chen, J. C., Johnson, E. E., and Johnson, A. E. (1997) Cell 89, 535-544[Medline] [Order article via Infotrieve] |
34. |
Rapak, A.,
Falnes, P. O.,
and Olsnes, S.
(1997)
Proc. Natl. Acad. Sci. U. S. A.
94,
3783-3788 |
35. | Tsai, B., Rodighiero, C., Lencer, W. I., and Rapoport, T. A. (2001) Cell 104, 937-948[Medline] [Order article via Infotrieve] |