The Plant Journal (2003) 34, 269–281
ER quality control can lead to retrograde transport from the
ER lumen to the cytosol and the nucleoplasm in plants
Federica Brandizzi1,, Sally Hanton2, Luis L. Pinto daSilva2, Petra Boevink3, David Evans1, Karl Oparka3,
Jürgen Denecke2,y and Chris Hawes1,y
1
Research School of Biological and Molecular Sciences, Oxford Brookes University, Gipsy Lane, Oxford OX3 0BP, UK,
2
Centre for Plant Sciences, Leeds Institute for Plant Biotechnology and Agriculture, School of Biology, University of
Leeds, Leeds LS2 9JT, UK, and
3
Scottish Crop Research Institute, Dundee DD2 5DA, UK
Received 8 August 2002; revised 17 December 2002; accepted 24 January 2003.
For correspondence (fax þ44 1865 483955; e-mail [email protected]).
y
Both laboratories contributed equally.
Summary
Quality control in the secretory pathway is a fundamental step in preventing deleterious effects that may
arise by the release of malfolded proteins into the cell or apoplast. Our aims were to visualise and analyse
the disposal route followed by aberrant proteins within a plant cell in vivo using fluorescent protein technology. A green fluorescent protein (GFP) fusion was detected in the cytosol and the nucleoplasm in spite
of the presence of an N-terminal secretory signal peptide. In contrast to secreted GFP, the fusion protein
was retained in the cells where it was degraded slowly, albeit at a rate much higher than that of the
endoplasmic reticulum (ER)-retained derivative GFP-HDEL. The fusion protein could not be stabilised by
inhibitors of transport or the cytosolic proteasome. However, the protein is a strong lumenal binding
protein (BiP) ligand. Complete signal peptide processing even after long-term expression in virus-infected
leaves rules out the possibility that the documented accumulation in the cytosol and nucleoplasm is
because of the bypassing of the translocation pores. The data are consistent with the hypothesis that
the fusion protein is disposed off from the ER via a retrograde translocation back to the cytosol. Moreover,
accumulation in the nucleoplasm was shown to be microtubule dependent unlike the well-documented
diffusion of cytosolically expressed GFP into the nucleoplasm. The apparent active transport of the GFP
fusion into the nucleoplasm may indicate an as yet undiscovered feature of the ER-associated degradation
(ERAD) pathway and explain the insensitivity to degradation by proteasome inhibitors.
Keywords: GFP, ERAD, quality control, secretory pathway.
Introduction
As nascent polypeptides are translocated into the endoplasmic reticulum (ER) lumen through the Sec61 pore
complex (Brodsky, 1998), the signal peptide is cleaved
and the proteins are likely to attain the final conformation
only after the import is complete (Gething and Sambrook,
1992; Hartl, 1996). Malfolded and incompletely assembled
proteins emerge as by-products in the ER. Several errors,
such as the incorrect incorporation of amino acids or premature termination of translation, can give rise to mutant or
truncated proteins. A variety of quality control mechanisms
operate to prevent their release into the cell or the apoplast
where they could be detrimental to the physiology of the
ß 2003 Blackwell Publishing Ltd
plant. This process also leads to their disposal so that
valuable amino acids can be recycled.
Degradation of newly synthesised malfolded or unassembled proteins in the secretory pathway is an essential
function of the quality control system in the ER so that only
correctly folded, processed, and completely assembled
proteins exit from this compartment for further transport.
One strategy for protein quality control in the secretory
pathway involves retention in the ER followed by ER-associated degradation (ERAD; Hiller et al., 1996; Ivessa et al.,
1999; Ward et al., 1995). Retrograde transport is ATP dependent and requires the luminal ER chaperone BiP (Brodsky
269
270 Federica Brandizzi et al.
et al., 1999; Pempler et al., 1997) and the ER membrane
translocation pore protein Sec61p (Brodsky et al., 1999;
Huges et al., 1997; Wiertz et al., 1996).
The ERAD pathway in yeast and animal cells has mainly
been demonstrated indirectly via inhibition of degradation
by the cytosolic proteasome (Amshoff et al., 1999; Hiller
et al., 1996; Huges et al., 1997; McCracken et al., 2000;
Pempler et al., 1997; Simpson et al., 1999; Wiertz et al.,
1996). More importantly, it is still unclear how permanently
malfolded proteins are distinguished from folding intermediates that may yet assume the correct conformation.
More recently, it has been demonstrated that the ERAD
route may not require full unfolding of the ERAD substrate,
as a green fluorescent protein (GFP) could be tagged onto
an ERAD substrate and followed from the ER to the cytosol
(Fiebiger et al., 2002). In other words, retrograde translocation has different properties compared to the classic forward translocation, as predicted from genetic studies (Zhou
and Schekman, 1999).
In plants, little is known about the mechanisms and
proteolytic systems that mediate protein quality control
in the secretory system. A suggestion of the existence of
a protein retrograde transport and degradation pathway in
plant cells has arisen from the analysis of the location of the
ricin catalytic A subunit in tobacco protoplasts (Frigerio
et al., 1998). It has also been demonstrated that the ricin
A chain in tobacco protoplasts is subject to de-glycosylation and degradation after ER retrotranslocation (Di Cola
et al., 2001). However, there is currently no evidence of a
malfolded protein being targeted from the ER to the cytosol
for degradation, although it is known that malfolded luminal proteins can be targeted to the vacuole via a Golgimediated pathway (Coleman et al., 1996; Pueyo et al.,
1995). Some other malfolded proteins are degraded via a
brefeldin A and heat shock-independent pathway in a so far
unidentified cellular compartment (Pedrazzini et al., 1997).
Here, we show that it is possible to visualise the ERAD
route of malfolded proteins, subjected to the quality control
processes in vivo, using chimeras of calreticulin subunits
and GFP. We demonstrate that the transport of malfolded
proteins, after retrograde exit from the ER, is cytoskeleton
dependent and results in accumulation within the nucleoplasm. The slow degradation is possibly because of this
unusual transport route and leads to relatively easy detection of ERAD activity.
Results
previous observations that small deletions or N-terminal
fusions to plant calreticulin cause severe malfolding, induction of the ER chaperone BiP and lack of secretion (Crofts
and Denecke, unpublished). Therefore, a secreted derivative of GFP was fused to various calreticulin domains, and
chimeric proteins were tested for increased ER retention
and vacuolar or cytoplasmic location. Figure 1 shows a
comparison, in Nicotiana tabacum leaf protoplasts, of the
behaviour of a GFP fusion protein containing the P-region
of calreticulin (Michalak et al., 1992) fused to the C-terminus
of GFP (sGFP-P) and sGFP-HDEL, a well-documented ER
marker. As expected, the latter accumulated in the mobile
elements of the ER and the nuclear envelope (Figure 1a,
arrowheads). In contrast, sGFP-P showed much weaker
fluorescence and was only faintly detected in the ER,
nuclear envelope, the cytosol and the nucleoplasm
(Figure 1b). Visualising GFP in the protoplasts expressing
sGFP-P was very difficult because of a very low signal to
noise ratio of the GFP in these cells.
We also expressed sGFP-HDEL and sGFP-P in the epidermal cells of Nicotiana benthamiana leaves using the viral
vector pTXS.P3C2 (Boevink et al., 1996), thus permitting
expression to be followed for longer periods of time. Figure 1(c) shows that sGFP-HDEL labels the nuclear envelope
and the classical cortical ER network. In contrast, sGFP-P
labels not only the nuclear envelope and the ER, but also the
cytosol and the nucleoplasm (Figure 1d,e,g). This pattern
was not altered in cells subjected to cold shock for 30 min or
brefeldin A (100 mg ml1) for 3 h (data not shown). Also in
tobacco protoplasts, the amount of cellular sGFP-P could
not be increased by treatment with brefeldin A or inhibition
of ER export via co-expression with Sec12 (Phillipson et al.,
2001), suggesting that the protein was not transported to
distal locations of the secretory pathway and then degraded
(data not shown). The presence in the cytosol and the
nucleoplasm was confirmed by double labelling with propidium iodide (compare Figure 1e,f). Even though thesGFP-P
hybrid did not contain a C-terminal HDEL motif, no significant levels of secretion could be detected (Figure 1g,h).
Under the same experimental conditions, secreted GFP
lacking the HDEL retention signal accumulated in the apoplast (data not shown; also see Boevink et al., 1999), while
free GFP (cGFP) resulted in diffuse fluorescence in the
cytosol and the nucleoplasm but not in the ER (Figure 8).
Thus, we can conclude that the calreticulin P-domain deviates the GFP fusion from the secretory pathway, even though
it is fused to the C-terminus of the protein and should be
synthesised only after signal peptide-mediated co-translational translocation (Vitale et al., 1993) has been initiated.
The calreticulin P-domain deviates GFP from the
secretory pathway to the cytosol and the nucleoplasm
Cytosolic and nuclear localisation of sGFP-P is not a
result of bypassing translocation into the ER lumen
To generate vital markers suitable for the visualisation of
quality control routes in plant cells, we took advantage of
One possible explanation for the above result could be
that high expression levels of sGFP-P lead to significant
ß Blackwell Publishing Ltd, The Plant Journal, (2003), 34, 269–281
GFP illuminates ER quality control
271
Figure 1. Retention and retrograde transport of GFP fusion proteins.
Tobacco protoplasts expressing sGFP-HDEL and sGFP-P were analysed by confocal microscopy 72 h after DNA transfer.
(a) The fluorescence of sGFP-HDEL accumulates in the ER (empty arrowhead) and in the nuclear envelope (full arrowhead, scale bar ¼ 10 mm).
(b) The fluorescence because of sGFP-P is weak in comparison to sGFP-HDEL. However, it is detectable in the ER, nuclear envelope, cytosol and nucleoplasm (full
arrowhead, scale bar ¼ 10 mm). Imaging of sGFP-P required on average five times higher laser intensity with respect to that of sGFP-HDEL for the same pinhole
aperture. The empty arrowhead indicates autofluorescent chloroplasts.
(c–i) Imaging differently targeted GFPs in virus-transformed Nicotiana bentamiana leaf epidermal cells.
(c) Ten days post virus infection expression of PVX.sGFP-HDEL is located in the ER (empty arrowhead) and the nuclear envelope (full arrowhead, scale
bar ¼ 20 mm).
(d) Expression of PVX.sGFP-P is located in the cortical ER (empty arrowhead) and the cytoplasm (full arrowhead, scale bar ¼ 10 mm).
(e) An optical section through the middle of an epidermal cell shows that sGFP-P also accumulates in the nucleoplasm (full arrowhead) and the cytoplasm. Note
that small vacuoles (empty arrowhead) can be seen in negative contrast and the apoplast is delimited by two layers of cytoplasmic fluorescence (star, scale
bar ¼ 10 mm).
(f) Double labelling the nuclei with propidium iodide (50 mg l1, 10 min) after an RNase treatment (50 mg l1, 5 min) in the same cells as in (e) confirms their
identity (full arrowhead) in sGFP-P-expressing cells.
(g,h) Trichomes expressing PVX.sGFP-P and double labelled with FM64 dye (Molecular Probes, California), which stains the plasma membrane (red, arrowhead),
confirms the absence of the apoplastic location of sGFP-P (scale bar ¼ 15 mm).
bypassing of the translocation pores in the rough ER membrane. If so, then this would be accompanied by a lack of
signal peptide cleavage. Such an event can easily be
detected, as the presence of a signal peptide is known to
cause an increase in the molecular weight of the nascent
secretory protein produced in vitro in the absence of microsomes (Denecke et al., 1990). We thus generated a similar
GFP fusion protein that lacked an N-terminal signal peptide (cytosolic GFP-P or cGFP-P) and compared its molecular weight with that of sGFP-P after production in
protoplasts, in virus-infected leaves and in Escherichia coli
(Figure 2).
ß Blackwell Publishing Ltd, The Plant Journal, (2003), 34, 269–281
Expression of sGFP-P and cGFP-P in E. coli clearly showed
an additional higher molecular weight band when the
signal peptide was present (Figure 2, compare lanes 5 and
6). Cytosolic GFP-P was produced at much higher levels
(data not shown), and the sample was diluted 10-fold compared to the E. coli extract from sGFP-P producers to allow
comparison of the molecular weights.The data areconsistent
with the generally lower capacity of periplasmic protein
productioninE. colicomparedtothatofcytosolicproduction.
The additional band is because of incomplete signal peptide
processing, hence providing a molecular weight marker
to compare it with the proteins expressed in planta.
272 Federica Brandizzi et al.
Figure 2. Cellular location of sGFP-P is not because of the bypassing of
translocation into the ER.
Cellular extracts of protoplasts transformed with sGFP-P (lane 2), of leaves
virally transformed with sGFP-P (lane 3), cGFP-P (i.e. lacking the signal
peptide, lane 4), of Escherichia coli expressing cGFP-P (lane 5) and sGFP-P
(lane 6) were analysed by SDS–PAGE and blotting with GFP antiserum. The
presence of one band in all the lanes with the exception of the sGFP-P
expressed in E. coli indicates cleavage of the sporamin signal peptide and
therefore absence of protein mistargeting. Lane 1 ¼ negative control.
In contrast, sGFP-P and cGFP-P produced in virusinfected leaves have an identical mobility on SDS–PAGE
(Figure 2, compare lanes 3 and 4), which is also similar to
that of sGFP-P produced in tobacco leaf protoplasts (lane 2).
No higher molecular weight form containing a signal peptide could be detected, indicating faithful translocation into
the ER, regardless of the method of gene transfer.
The data confirm that signal peptide processing in planta
is generally complete and difficult to bypass (Denecke et al.,
1990). This is a strong evidence against the bypassing of
translocation, together with the fact that the onset of translocation and probably signal peptide processing is likely to
be completed well before the P-domain emerges from the
ribosome. Therefore, we propose that sGFP-P first enters
the ER lumen at least partially, undergoes efficient signal
peptide processing by signal peptidase, and is later relocated to the cytosol and the nucleoplasm.
sGFP-P associates with BiP and is degraded
To obtain evidence for malfolding of the C-terminal portion
of the fusion protein, we tested if sGFP-P can interact with
BiP in an ATP-dependent fashion. Virus-infected leaf discs
were subject to in vivo labelling followed by extraction and
immunoprecipitation with anti-BiP serum (Figure 3). Plants
expressing sGFP-P exhibited higher synthesis rates for BiP
compared to the control (Figure 3, compare lanes 1 and 2),
indicating ER stress possibly because of the presence of
malfolded protein. Furthermore, an abundant protein of
lower molecular weight co-precipitated with BiP. This protein was clearly absent in the control lane in which in vivo
labelling of leaf discs from untransformed plants revealed
only BiP in the immunoprecipitate.
To confirm that the band corresponds to sGFP-P, pellets
were washed with ATP (3 mM) in BiP release buffer. Figure 3
shows that the majority of the co-precipitating protein was
Figure 3. Anti-BiP immunoprecipitation of extracts of leaves transformed
with PVX.sGFP-P.
Discs of untransformed and PVX.sGFP-P-transformed leaves of Nicotiana
benthamiana were labelled for 3 h with 35S-methionine and 35S-cysteine.
After immunoprecipitation, proteins were analysed by SDS–PAGE and
fluorography. Extracts of labelled leaves, untransformed (lane 1) and transformed with PVX.sGFP-P (lane 2) were immunoprecipitated with anti-BiP
(a-BiP). The precipitate was then washed with ATP (þATP, lane 3 (untransformed) to lane 4 (transformed)), and ATP washes were immunoprecipitated
with maize anti-calreticulin (a-Cal, lane 5 (untransformed) to lane 6 (transformed)].
removed from BiP (lane 4) but it could be re-precipitated
from the supernatant with anti-maize calreticulin antibodies (Figure 3, lane 6). Thus, the result confirmed the
identity of the co-precipitated protein as the GFP fusion
carrying the P region of calreticulin.
To test whether the interaction with BiP was because of
GFP itself or the fused P-domain, we compared cytosolic
GFP with secreted GFP (sGFP), sGFP-HDEL and sGFP-P
(Figure 4). To avoid the variation inherent to virus infections, the efficiency of which is difficult to control or quantify, as well as differences in the physiology of individual
leaves which could influence BiP levels, we used transient
expression in tobacco leaf protoplasts as a model system.
All experimental parameters were thus the same, except for
the transfected plasmid.
Protoplasts were transfected and metabolically pulselabelled (P) followed by a 10 h chase period (C), and cell
extracts and culture medium were immunoprecipitated
with anti-GFP serum. Figure 4 shows that sGFP-P co-precipitates much more BiP than the three controls (panel a).
Line scans are shown for sGFP-HDEL and sGFP-P to illustrate that the differences are quantitative (panel b).
Figure 4 also shows that sGFP-P is slowly degraded, but
much more rapidly than sGFP-HDEL whose half-life is well
beyond the entire chase period. Quantification of the signal
from phosphorimaging revealed that after the chase,
labelled sGFP-P dropped to 6.7% during the chase period.
This corresponded to an estimated half-life of 2.5 h. It can
be concluded that sGFP-P is degraded because it does not
chase into the medium (data not shown). In contrast, the
appearance of sGFP in the medium is easily detected after
the chase (not shown).
To confirm these results, we co-expressed sGFP-HDEL
and sGFP-P in the same protoplast suspension to rule out
variance in the pulse-chase experiments. The line scan
reveals the signal for sGFP-HDEL, sGFP-P and BiP, and
ß Blackwell Publishing Ltd, The Plant Journal, (2003), 34, 269–281
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273
Figure 4. Evidence showing that sGFP-P constitutes a typical BiP-ligand with strong ATPdependent affinity compared to other GFP derivatives.
(panel a) Pulse chase of 70% 35S-methionine
and 30% 35S-cysteine-labelled tobacco protoplasts transformed with DNA encoding for cytosolic GFP (cGFP), secreted GFP (sGFP), ERretained GFP (sGFP-HDEL) and secreted GFP
fused to the P region of calreticulin (sGFP-P).
sGFP co-precipitates BiP but is degraded during
the chase period.
(panel b) Quantification of pulse-chased sGFPHDEL and sGFP-P immunoprecipitation, confirming the instability of sGFP-P.
(panel c) Line scans of pulse-chase samples
from an experiment in which sGFP-P and
sGFP-HDEL were co-expressed in the same protoplasts. Note the much faster degradation of
sGFP-P compared to that of sGFP-HDEL.
again illustrates the much shorter half-life of sGFP-P compared to that of sGFP-HDEL (Figure 4c).
The weak association of BiP with sGFP-HDEL may indicate that GFP folds very slowly. The apparent increase in the
amount of co-precipitating BiP with sGFP-HDEL after the
chase (Figure 4b) indicates that labelled BiP remains stable
during the chase period and can be associated with
de novo synthesised (cold) sGFP-HDEL. This is not surprising as the total amount of sGFP-HDEL increases significantly during the experimental period. In conclusion, the
data clearly suggest that sGFP-P has increased BiP-binding
properties and is also degraded more quickly than the GFP
controls.
GFP-P does not produce the typical degradation
intermediate shown by secreted GFP
The results from Figure 4 indicate the presence of a degradation intermediate of lower molecular weight for sGFP and
sGFP-HDEL. This intermediate was identical in molecular
ß Blackwell Publishing Ltd, The Plant Journal, (2003), 34, 269–281
weight in both cases, although the precursors differed in
the presence of the HDEL motif. We concluded that GFP was
most likely proteolytically cleaved near its C-terminus,
yielding identical proteins regardless of the presence of
the HDEL motif. This band of lower molecular weight could
also be seen in Western blots, but was only detected in the
cells (Figure 5b) and not in the medium (Figure 5a). It
should be noted that the P-domain was also fused to the
C-terminus as the HDEL peptide, but interestingly, this Cterminal processing intermediate was not observed for
sGFP-P (Figure 5). The absence of the typical intermediate
suggested that degradation of this fusion protein occurred
in a different manner.
The ratio between precursor and degradation intermediate was shifted towards the precursor when the HDEL
motif was present. Given the proteolytically benign environment of the ER lumen, we postulated that the degradation product was a result of ER export, and occurred
downstream of the ER in the secretory pathway, not in
the medium.
274 Federica Brandizzi et al.
the soluble fraction, which also contains vacuolar contents.
The results are consistent with the hypothesis that a portion
of sGFP-HDEL is transported out of the ER and may explain
the generally low fluorescence of secreted GFP in the
apoplast (Batoko et al., 2000; Boevink et al., 1999). Figure 6
again demonstrates the stability of sGFP-P, which is generally lower than that of sGFP-HDEL.
Degradation of sGFP-P is not because of a
clasto-lactacystin b-lactone-sensitive machinery
Figure 5. Despite the absence of a retention/retrieval signal, the sGFP-P
fusion is not secreted.
Tobacco protoplasts were electroporated in the absence of foreign DNA or
with DNA encoding for cytosolic GFP (cGFP), secreted GFP (sGFP), ERtargeted GFP fused to HDEL (sGFP-HDEL) or to the P region of maize
calreticulin (sGFP-P). Proteins were detected by SDS–PAGE (10%) blotting
with GFP antiserum.
(a) Immunodetection of proteins in the protoplast medium. The efficiency of
secretion of sGFP is high compared to that of the other constructs as
detected in blots on 4 concentrated medium. Despite the absence of a
retention/retrieval signal, sGFP-P is not secreted (arrowhead).
(b) Immunoblots of cellular extracts show bands with expected gel mobility
and partial degradation products of sGFP and sGFP-HDEL, but not of sGFP-P
(arrowhead).
To provide further evidence for this, we conducted pulsechase labelling of sGFP-HDEL-producing cells and separated cells into soluble and microsomal fractions via an
established osmotic shock procedure (Denecke et al., 1992).
Figure 6 shows that the degradation intermediate was only
detected after a prolonged chase (14 h) but exclusively in
Figure 6. Proteasomal inhibition with clasto-lactacystin b-lactone has no
detectable effect on sGFP-P.
Protoplasts co-electroporated with sGFP-P and sGFP-HDEL were treated
with the proteasome inhibitor clasto-lactacystin b-lactone (100 mM) after 1 h
pulse. Samples were then chased for 3 and 14 h. Chased protoplasts were
subjected to osmotic shock. sGFP-P and the lower molecular size band
present in sGFP-HDEL lane were not stabilised by the inhibitor. –¼ control;
bL ¼ clasto-lactacystin b-lactone; S ¼ soluble fraction; P ¼ pellet.
Recent work has shown that the ricin A chain exploits ER
retrotranslocation before being degraded in the protoplast
cytosol upon de-glycosylation (Di Cola et al., 2001). The
degradation is partially reduced by the proteasome inhibitor, clasto-lactacystin b-lactone. On the basis of these studies, it has been suggested that ricin A may be degraded
by the proteasome, even if the protein is not necessarily
malfolded.
We pulsed co-electroporated protoplasts with sGFP-P
and sGFP-HDEL as control and chased them at 3 and
14 h in the absence or presence of 100 mM clasto-lactacystin
b-lactone. sGFP-HDEL appeared in the soluble and the
pellet fractions. The presence of this protein in the soluble
fraction might be because of the release of the luminal
content of the endomembranes during preparation of the
samples.
The sGFP-P fusion did not appear to be stabilised by the
proteasome inhibitor (Figure 6). An instability of the sGFP-P
in the cytosol might indicate the lack of increase in the
soluble and pellet ratio over the time course of the experiment.
The slower degradation of sGFP-HDEL and the appearance of the lower molecular weight form was also not
influenced by the presence of the proteasome inhibitor.
sGFP-P first accumulates in the ER and then in the
nucleoplasm
It is not possible to show that the microsomal sGFP-P
chases out into a cytosolic form. Such classical pulse-chase
result is only possible if the chase product is at least as
stable or more stable than the precursor, but this does not
seem to be the case. However, the complete signal peptide
processing, as well as the typical ER fluorescence pattern,
indicates that sGFP-P enters the ER and folds sufficiently
to support fluorescence. To completely rule out that sGFP-P
in the ER/nuclear envelope and the cytosol/nucleoplasm
derived from two different pools exhibiting different targeting routes, we monitored fluorescence distribution in leaf
epidermal cells as a function of time. sGFP-HDEL accumulated in the nuclear envelope and in the ER 3 days after
virus infection (Figure 7a). The location of the protein
fusion did not change with prolonged virus infection
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275
Figure 7. Accumulation of sGFP-P in the cytoplasm and nucleoplasm is time dependent.
With sGFP-HDEL, fluorescence can be seen in
both the ER and the cross-sections of the
nucleus, clearly in the nuclear envelope 3 days
(a) and 3 weeks (b) post-infection. With epidermal cells expressing sGFP-P, 3 days post-infection fluorescence can clearly be seen in the
nuclear envelope (c); however, 2 weeks postinfection fluorescence accumulates in the cytoplasm and nucleoplasm masking residual fluorescence in the nuclear envelope (d). Relative
fluorescent intensity in the nuclear envelope
compared to the nucleoplasm can easily be
seen via line scans across the images.
(Figure 7b). Figure 7(c) shows that 3 days after virus infection, sGFP-P was primarily detected in the ER and the
nuclear envelope. However, prolonged incubation for
2 weeks resulted in the accumulation of sGFP-P in the nucleoplasm with lower levels detected in the nuclear envelope
(Figure 7d). The data suggested that the location of sGFP-P
ß Blackwell Publishing Ltd, The Plant Journal, (2003), 34, 269–281
shifted from the ER to the cytosol and the nucleoplasm with
time. This result argues against the presence of two alternative pools of the fusion protein and is consistent with our
hypothesis of retrograde ER transport of sGFP-P.
It is worth noting that the protoplasts and the epidermal cells appear to exhibit different levels of sGFP-P
276 Federica Brandizzi et al.
Figure 8. Photobleaching analysis of the fluorescence of cells transformed with PVX.cGFP, PVX.sGFP-P and PVX.sGFP-C.
The fluorescence of the nucleus of an untreated epidermal cell transformed with PVX.cGFP, releasing cytoplasmic GFP (a, arrowhead; scale bar ¼ 10 mm) is not
appreciably quenched in 1 min (b), resulting in a little difference in the fluorescence intensity in the recovery period (c). Parallel results obtained with the control
in which the nuclei of colchicine-treated cells (d, arrowhead, scale bar ¼ 10 mm) do not show appreciable loss of fluorescence after photobleaching (e) and
therefore recovery (f). The fluorescence of a nucleus of an untreated epidermal PVX.sGFP-P-transformed cell (g, arrowhead; scale bar ¼ 10 mm) shows little
quenching after 1 min of photobleaching (compare (h) with recovery image (i), taken 30 sec later). A nucleus of a cell transformed with PVX.sGFP-P treated with
colchicine (j, arrowhead; scale bar ¼ 10 mm) was photobleached for 1 min (k) and does not show the recovery of fluorescence within the nucleoplasm after 1 min
(l) or even 5 min (not shown), but recovery can be seen in the nuclear envelope. (m–o) Epidermal cells transformed with PVX.sGFP-C showed accumulation of
GFP in the ER, cytosol and nucleoplasm, similar to those transformed with PVX.sGFP-P. Photobleaching the nuclei of untreated epidermal cells (m, arrowhead,
scale bar ¼ 10 mm) did not induce an appreciable loss of fluorescence after 1 min of photobleaching (n), and no recovery was therefore detectable within 5 min
(o). (p–r) Photobleaching the nuclei of colchicine-treated cells expressing PVX.sGFP-C (p, arrowhead, scale bar ¼ 10 mm). After photobleaching for 1 min, the
nucleus of cells expressing sGFP-C in the presence of colchicine showed loss of fluorescence (q) and no recovery even after 5 min (r). White horizontal line
indicates the transect along which pixel intensity was measured and plotted as arbitrary fluorescence units in the overlaid graph.
ß Blackwell Publishing Ltd, The Plant Journal, (2003), 34, 269–281
GFP illuminates ER quality control
fluorescence, with the protein showing increased stability
in the leaves. This may reflect differences in the species
used for the two expression systems or a different metabolic state between the cells in planta and the protoplasts.
The higher detection of sGFP-P in the leaves may therefore
be because of either a slower degradation of sGFP-P or a
quicker saturation of the degradation machinery compared
to the protoplasts.
Nuclear targeting of sGFP-P is active and microtubule
dependent
Even though cytoplasmic detection of sGFP-P could be
explained by the ERAD pathway, the location in the nucleoplasm was unexpected. This could be because of the fusion
protein diffusing through the nuclear pores thus bypassing
the proteasome. To test this, we monitored fluorescence
recovery of the nucleus after photobleaching (FRAP;
Figure 8) and investigated the dynamics of the process
by comparing the movement of cytosolic GFP and sGFPP. Both cytosolic GFP and sGFP-P were difficult to bleach in
the nucleoplasm because of the rapid recovery of fluorescence, presumably because of rapid diffusion/transport of
the proteins back into the nucleus (Figure 8a–c,g–i).
We also analysed the influence of the microtubule-destabilising drug colchicine (1 mM for 30 min) on the transport of the proteins into the nucleus. Colchicine did not
prevent the nuclear localisation of cytoplasmic GFP after
photobleaching (Figure 8d–f), indicating that this molecule
diffused rapidly into the nucleus in a microtubule-independent fashion. The same results were obtained after treatment with another plant-specific microtubule inhibitor,
oryzalin (10 mg ml1 for 30 min; data not shown). However,
in sGFP-P-expressing cells treated with colchicine (Figure 8j), the nucleoplasm was bleached but a significant
amount of fluorescence persisted in the nuclear envelope
(Figure 8k). Although fluorescence subsequently recovered
fully in the nuclear envelope well within 1 min after bleaching (Figure 8l), reflecting rapid diffusion from the ER lumen,
fluorescence in the nucleoplasm did not recover under
these conditions.
These results suggested that nuclear targeting of sGFP-P
from the cytosol, but not cytosolic GFP itself, was microtubule dependent and illustrated that the two molecules
reached the nucleoplasm via different pathways. It could
thus be ruled out that the nuclear targeting of sGFP-P was
because of an sGFP-intrinsic nuclear localisation mechanism.
A putative nuclear localisation signal has been identified
within the P coding region of maize calreticulin (Napier
et al., 1994). To test if this region was responsible for our
observations, we carried out similar experiments on an
ER-targeted GFP fused to the C region of the maize calreticulin (sGFP-C). The latter region did not contain a nuclear
ß Blackwell Publishing Ltd, The Plant Journal, (2003), 34, 269–281
277
targeting sequence, but fusion of this domain caused a
similar distribution of fluorescence compared to that of
sGFP-P and included the ER, the nuclear envelope, the
cytoplasm and the nucleoplasm (Figure 8m–r). In FRAP
experiments in the absence of colchicine, bleaching was
partial, followed by a quick recovery (Figure 8m–o). The
fluorescence in the nuclei of colchicine-treated cells transformed with sGFP-C did not recover after photobleaching
(Figure 8p–r). Thus, we concluded that the nuclear targeting of sGFP-P and sGFP-C was independent of any known
motif and might be a novel feature of the ERAD pathway.
Discussion
Visualisation of the ERAD pathway with a GFP fusion
Quality control by the ER is proposed to occur via two
different mechanisms, the disposal by the lytic vacuole
(Hong et al., 1996) and the ERAD pathway (Ward et al.,
1995). Evidence for the former pathway arose from the
generation of an invertase fusion protein containing a Cterminal malfolded portion (Hong et al., 1996). It could be
argued that this fusion protein mimics a partially unfolded
protein as it is composed of a folded portion (invertase) and
an additional malfolded portion that is recognised by the
quality control machinery. This disposal route was shown
to be dependent on the vacuolar sorting receptor VPS10,
which could recognise the incompletely folded protein
either directly, or perhaps via interaction with an adapter
molecule that becomes a VPS10 ligand once it is associated
with an incorrectly folded protein. It is also argued that such
a mechanism would be energetically economic compared
to ERAD, as the unfolding of the entire molecule (including
the folded invertase) would cost more energy. In contrast,
the proteins that are completely unfolded may follow the
ERAD pathway as they remain translocation competent.
The GFP fusion protein used in this study appeared to be
sufficiently folded to support fluorescence but yet permitted us to visualise the ERAD pathway. The fact the
ER-targeted GFP constructs without H/KDEL retention signals have been shown to be secreted (Batoko et al., 2000;
Boevink et al., 1999) suggests that a retention mechanism
must be in operation to explain ER staining because of
sGFP-P. This is most likely because of the association of
sGFP-P with BiP as demonstrated biochemically. As proposed for rice prolamins (Li et al., 1993) and for assemblydefective form of the bean vacuolar storage protein phaseolin (Pedrazzini et al., 1997), proteins can thus be retained
in a signal-independent manner within the ER through
prolonged interaction with other proteins that contain an
HDEL signal. In the case of sGFP-P, the candidate would be
the molecular chaperone BiP, as suggested by the biochemical evidence presented here.
278 Federica Brandizzi et al.
Biochemical and genetic analyses have indicated that
luminal proteins were selected for proteolysis and exported
to the cytosol by the ERAD process. This required initial
retention of the substrate protein in the ER by BiP (Brodsky
and McCracken, 1999; Pempler et al., 1997) and wasfollowed,
sometimes after a considerable lag phase, by rapid degradation without the detection ofdetectableproteolyticintermediates in the cytosol (Hiller et al., 1996; Ivessa et al., 1999;
Sommer and Wolf, 1997). The dynamics of sGFP-P, namely
entering the ER lumen, an ATP-dependent interaction with
BiP as well as a distribution in the ER, the cytosol and the
nucleoplasm, showed features similar to those exhibited by
ERAD substrates. Moreover, the distribution of the fluorescencebecauseofsGFP-PexpressionwasnotalteredbyBFAor
cold treatment, as would be expected if a transport between
ER and Golgi along the secretory pathway prior to the release
into the cytosol was involved (Boevink et al., 1999). Our
results therefore indicate in vivo evidence of the occurrence
of an ER retrograde transport of a protein.
The aberrant location of sGFP-P is not because of bypassing the translocation process itself, but occurs after cleavage of the signal peptide. As higher molecular weight
forms of the constructs containing the signal peptide were
not detected, it is likely that the protein enters the translocation machinery, although it does not show whether
translocation is completed before retrograde transport is
initiated. Most importantly, the fact that sGFP-P is first
detected in the ER and later in the cytosol and nucleoplasm
rules out the possibility that one pool of sGFP-P enters the
ER while the other bypasses translocation. If the latter were
the case, deviation from the secretory pathway should be
independent of time.
There is also significant evidence suggesting malfolding
of sGFP-P or portions of it. First, sGFP-P is a much stronger
BiP ligand than sGFP-HDEL itself. It is also degraded faster
than sGFP-HDEL, albeit slowly compared to the findings on
the ricin A chain. We have also shown that sGFP and sGFPHDEL are proteolytically cleaved into a defined degradation
intermediate, also seen in A. thaliana plants expressing
secreted GFP and GFP-HDEL (Zheng, Hawes and Moore,
unpublished), which is not seen for sGFP-P. This suggests
that sGFP-P is not degraded in the same way. We propose
that this GFP fusion highlights the ERAD pathway in plant
cells. Perhaps, the small size of GFP allows retrograde
translocation in spite of folding into a conformation that
supports partial fluorescence, a possibility recently suggested by Fiebiger et al. (2002).
An unexpected result was the time-dependent location of
sGFP-P in the nucleoplasm. This appeared to be in a manner distinct from that of the well-known nuclear transport of
cytosolic GFP. It may be this unexpected targeting step that
prevents the protein from degrading quickly in the proteasome, which would explain the lack of inhibition by lactacystin.
Our results indicate that different pathways for protein
degradation may exist in protoplasts. Different sensitivities
to degradation pathways verified for sGFP-P and ricin A
may depend on the exposure to different signals for degradation. For example, de-glycosylation occurring for ricin A
may be an essential prerequisite for proteasome-mediated
degradation. However, it is possible that sGFP-P and ricin A
may share similar signals for retrotranslocation, but not for
degradation. The fact that sGFP-P fusion does not seem to
be a candidate for proteasomal degradation could explain
the longer half-life of this protein in comparison to ricin A.
Relevance of the nuclear targeting of
an ERAD substrate
The unexpected nuclear localisation of the fusion protein
prompted us to investigate the dynamics of sGFP-P by
FRAP experiments. The nuclear localisation of free GFP
could be explained by its small size that was below the
exclusion limit for passive transport through the nuclear
pore complex. In our experiments, free GFP was localised
only in the cytosol and nucleoplasm. Reits et al. (1997)
proved by FRAP experiments that, in mammalian cells, free
GFP movement to the nucleus follows passive diffusion
dynamics. Our FRAP experiments confirm these data, as
photobleaching the nuclei of cells expressing free GFP in
the presence and absence of the microtubule-destabilising
agents did not alter the rapid rate of diffusion of the GFP to
the nucleus.
Photobleaching the nuclei of cells transformed with
PVX.sGFP-P and treated with microtubule inhibitors
induced a non-recoverable loss of fluorescence in the
nucleoplasm, although the fluorescence because of
sGFP-P quickly recovered in the nuclear envelope. The
recovery in the nuclear envelope after photobleaching
would be expected to occur because the synthesis pathway
starts in the ER/nuclear envelope, followed by diffusion
within the ER lumen. However, the complete inhibition of
protein movement towards the nucleoplasm by colchicine
indicates that the transport of the malfolded GFP fusion to
the nucleus is microtubule mediated, and occurs in a manner different from that of cytosolic GFP. Moreover, this
movement does not seem to depend on the nuclear targeting sequence present on the sGFP-P itself. In FRAP experiments on PVX.sGFP-C expressing cells, which present a
similar cellular distribution pattern to sGFP-P producers,
despite the absence of a putative nuclear localisation signal
in the C-region of calreticulin, showed that in the absence of
microtubules there was no recovery of bleached nucleoplasm fluorescence.
ERAD substrates require cytosolic or nucleoplasmic factors such as proteasomes to mediate their degradation
(Ivessa et al., 1999). Proteasomes interact continuously
with cytosolic and nuclear proteins and perform quality
ß Blackwell Publishing Ltd, The Plant Journal, (2003), 34, 269–281
GFP illuminates ER quality control
control to degrade proteins that are either tagged by ubiquitination or otherwise already in an appropriate conformation for degradation (Reits et al., 1997). Little data exist
on the location of the proteosome in plants. They have been
located in the cytoplasm of potato suspension cells by
direct immunofluorescence, are possibly associated with
elements of the cytoskeleton and may also reside in the
nuclei (Vallon and Kull, 1994).
Reichel and Beachy (2000) have demonstrated that inhibition of proteasomal activity results in the location of a
tobacco mosaic virus movement protein–GFP fusion on the
ER. These data indicate a location of proteasomes similar
to that known for animal cells (Arcangeletti et al., 2000;
Palmer et al., 1994; Rivett, 1998). Reits et al. (1997) have
showed the occurrence of an active transport of the proteasome into the nuclei. It is therefore reasonable to presume
that in the system described here sGFP-P and sGFP-C move
towards the nuclei along the proteasome pathway. A possible saturation of the proteasome machinery because of
overexpression could justify the visualisation of the GFP
fusions in our experiments.
Conclusions
We have illuminated a retrograde transport pathway from
the ER lumen to the cytosol using a GFP-tagging approach.
In addition, our data demonstrate that the ERAD pathway
may involve transport of malfolded proteins to the nucleoplasm. In combination with the biochemical assays for
signal peptide processing and interaction with the ER chaperone BiP, this system will be invaluable to elucidate the
various steps controlling the disposal pathway for malfolded proteins.
Experimental procedures
Constructs
Amplification products were generated by using Taq/Pwo (HYBAID
Ltd, Ashford, UK). All DNA manipulations were performed according to the established procedures (Sambrook et al., 1989). E. coli
XL1-Blue strain (Stratagene, La Jolla, USA) was used for the
amplification of all the plasmids with the exception of pET23b(þ)
vector (Novagen, Madison, USA), which was amplified in
BL21(DE3).
Virus vector. Potato virus X (pTXS.P3C2) was chosen as an
expression vector (Boevink et al., 1996), and the wild-type GFP
was used. The following constructs were cloned as ClaI–NsiI
fragments under the transcriptional control of duplicated
subgenomic RNA promoters: cytosolic GFP (cGFP, i.e. without
any signal); secreted GFP (sGFP, i.e. a fusion of the sporamin
signal peptide to GFP at the N-terminus); GFP targeted to the ER
by the sporamin signal peptide and retained/retrieved by HDEL
(sGFP-HDEL); GFP targeted to the ER by the sporamin signal
peptide and C-terminally fused in-frame to aa 185–297 of the P
ß Blackwell Publishing Ltd, The Plant Journal, (2003), 34, 269–281
279
region of maize calreticulin (sGFP-P); untargeted GFP and Cterminally fused in-frame to aa 185–297 of the P region of maize
calreticulin (GFP-P); GFP targeted to the ER by the sporamin signal
peptide and C-terminally fused in-frame to aa 298–392 of the C
region of maize calreticulin (sGFP-C) (Napier et al., 1994).
Protoplast expression. For N. tabacum protoplasts, expression
of mGFP5 was used, which is a GFP with a codon usage optimised
for plant nuclear expression (Haseloff et al., 1997). Substituting the
wild-type GFP with mGFP5 produced the constructs described
above, with the exception of sGFP-C, which was not used in
protoplast expression. The constructs were amplified and
cloned as ClaI–BamHI fragments into pLL4, a novel 35S based
plant expression vector in which the NcoI codon spanning the ATG
initiation codon was replaced by ClaI. The plasmid contained a
spacer in between the promoter and the 30 untranslated end of the
nopaline synthase gene, which ended with a BamHI site.
Escherichia coli expression. For E. coli expression, the coding
regions of GFP-P and sGFP-P previously used for virus expression
were cloned as an NheI–SacI fragment into an expression vector
pET23b(þ).
Expression systems
Virus expression. The run-off transcripts and inoculation on
leaves of N. benthamiana were produced as described (Boevink
et al., 1996). For all the experiments described here, plants 10–
14 days after transformation were used.
Protoplast transient expression. Tobacco leaf protoplasts
were prepared, and electroporation experiments were
performed as described previously (Denecke and Vitale, 1995).
Protoplasts were harvested for protein analysis 24 h after
electroporation. For microscopy, cell suspensions were cultured
for 24, 48, or 72 h.
Escherichia coli expression. The GFP-P and sGFP-P constructs
were cloned in the pET-23b(þ) vector, and protein production was
induced in BL21(DE3) lysogens. Positive clones were selected for
low-scale protein production. A single colony was inoculated
initially into 5 ml of Luria-Bertani (LB) containing ampicillin
(100 ml ml1) and further expanded into a 50 ml shaker culture
in 250 ml flasks. The cells were incubated with shaking at 378C until
OD600 was approximately 0.9. Protein production was induced by
the addition of 1 mM IPTG and further incubation of the culture for
2.5 h at 378C. Cells were then pelleted leaving an equal volume of
culture medium with the cells, before storage at 708C. Protein gel
analysis was performed after 1 day.
Protein extraction and gel blot analysis
For immunoprecipitation, leaf discs were homogenised in ice-cold
buffer composed of 200 mM Tris–HCl (pH 8), 300 mM NaCl, 1%
Triton X-100 and 1 mM EDTA (Crofts et al., 1998). Harvesting of
protoplasts and culture medium was performed as described
(Denecke and Vitale, 1995).
Protoplast and E. coli cellular extracts were obtained by repetitively pipetting the cells in an ice-cold buffer containing 0.1 M Tris–
HCl (pH 7.8), 0.2 M NaCl, 1 mM EDTA, 2% b-mercaptoethanol and
0.2% Triton X-100 (Pedrazzini et al., 1997). Protein analysis of the
protoplast culture medium was performed after concentration
with aqueous ammonium sulphate solution (final concentration
280 Federica Brandizzi et al.
60%) in the presence of BSA as a carrier (200 mg per 600 ml of
medium) and re-suspending the protein precipitate in a buffer
containing 25 mM Tris and 5 mM EDTA. An equal volume of
protein extract was added to 2 SDS loading dye (Crofts et al.,
1998), prior to boiling and gel loading. Protein analysis was performed by SDS–PAGE on 10–12% acrilamide gels. Immunodetection of GFP in protein gel blots was performed using the enhanced
chemiluminescence (ECL) system (Amersham, Life Sciences, UK),
with rabbit polyclonal antiserum raised against GFP (Molecular
Probes, Inc., USA) at 1 : 3000 dilution.
Immunoprecipitation and pulse chase
For sGFP-P immunoprecipitation, untransformed and transformed
leaf disks of the same size were labelled for 3 h with 100 mCi ml1
Pro-mix (Amersham Life Science, UK; containing 70% 35S-methionine and 30% 35S-cysteine) in MS medium. Homogenisation of the
leaf tissue and subsequent immunoprecipitation were performed
essentially as described (Crofts et al., 1998) with a purified anti-BiP
serum. ATP washes of the immunoprecipitates were performed as
described previously (Vitale et al., 1993). Re-immunoprecipitation
was performed with anti-maize calreticulin. Proteins were analysed by SDS–PAGE (12% gel), and labelled proteins were visualised with a phosphorimager SI. Pulse-chase experiments used a
1 h labelling and chase buffer containing 10 mM methionine and
5 mM cysteine.
Clasto-lactacystin b-lactone (Calbiochem, CN Biosciences, UK)
was used as the proteasome inhibitor at a concentration of 100 mM
(stock solution 20 mM in DMSO) after a 1 h pulse. Samples were
then chased at different time points. Chased protoplasts were
pelleted, and proteins were extracted in four times volume of
Tris–EDTA buffer (1 mM EDTA, 1 mM Tris).
Confocal microscopy
Leaf segments were imaged with a Zeiss LSM 410 confocal microscope using a 488 nm Argon ion laser plus a narrow bandpass filter
set (BP 505–515 nm) and photobleached using the region of interest software. For propidium iodide observations, a 543 nm Helium
ion laser with the LP 570 nm emission filter configuration was
used. The analysis of the fluorescence intensity was performed
with the line measurement software package of the microscope.
Acknowledgements
The BBSRC is acknowledged for a grant (P08503) supporting this
research, and we thank Drs J. Boyce and R. Napier for allowing us
the use of the calreticulin DNA and antibodies. Luis L. Pinto daSilva
is indebted to the British Council and the Brazilian CNPQ for
supporting a scientific exchange to the University of Leeds.
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