3942
Research Article
Differences in endoplasmic-reticulum quality control
determine the cellular response to disease-associated
mutants of proteolipid protein
Peristera Roboti, Eileithyia Swanton* and Stephen High
Faculty of Life Sciences, The University of Manchester, Oxford Road, Manchester M13 9PT, UK
*Author for correspondence ([email protected])
Journal of Cell Science
Accepted 1 September 2009
Journal of Cell Science 122, 3942-3953 Published by The Company of Biologists 2009
doi:10.1242/jcs.055160
Summary
Missense mutations in human PLP1, the gene encoding myelin
proteolipid protein (PLP), cause dysmyelinating PelizaeusMerzbacher disease of varying severity. Although disease
pathology has been linked to retention of misfolded PLP in the
endoplasmic reticulum (ER) and induction of the unfolded
protein response (UPR), the molecular mechanisms that govern
phenotypic heterogeneity remain poorly understood. To address
this issue, we examined the cellular response to missense
mutants of PLP that are associated with distinct disease
phenotypes. We found that the mild-disease-associated mutants,
W162L and G245A, were cleared from the ER comparatively
quickly via proteasomal degradation and/or ER exit. By
Introduction
A number of clinically important human diseases are known to be
caused by protein misfolding at the endoplasmic reticulum (ER).
The ER is a major site for the synthesis of membrane and secretory
proteins in eukaryotic cells, and nascent polypeptides are
translocated across the ER membrane as extended chains, which
then typically fold and oligomerise with the assistance of ER
chaperones. Correctly folded and assembled proteins are packaged
into transport vesicles for delivery to the Golgi complex, whereas
those that have not acquired the proper conformation are normally
retained in the ER by a quality-control system that helps to ensure
that misfolded or misassembled proteins are not deployed within
the cell. Persistently misfolded proteins are normally
retrotranslocated into the cytosol and degraded by proteasomes via
a process known as ER-associated degradation (ERAD) (Anelli and
Sitia, 2008; Vembar and Brodsky, 2008). Situations that disrupt
protein folding lead to the production of potentially toxic misfolded
proteins in the ER; hence, eukaryotic cells employ a defence
mechanism that maintains a balance between protein synthesis,
folding and degradation, known as the unfolded protein response
(UPR) (Ron and Walter, 2007; Schroder and Kaufman, 2005). The
UPR can be initiated by three transmembrane stress sensors, IRE1,
ATF6 and PERK, that are activated by the accumulation of
misfolded proteins in the ER. These form distinct branches of the
UPR that function to reduce the overall rate of protein synthesis
and upregulate the folding and degradation capacity of the ER,
thereby reducing levels of misfolded protein in the ER, and
potentially restoring homeostasis (Friedlander et al., 2000; Ng et
al., 2000). However, if the cell fails to re-establish ER and cellular
homeostasis, prolonged UPR signalling can induce programmed
cell death (Szegezdi et al., 2006).
contrast, the more ‘aggressive’ A242V mutant, which causes
severe disease, was significantly more stable, accumulated at
the ER and resulted in a specific activation of the UPR. On the
basis of these findings, we propose that the rate at which mutant
PLP proteins are cleared from the ER modulates disease
severity by determining the extent to which the UPR is activated.
Supplementary material available online at
http://jcs.biologists.org/cgi/content/full/122/21/3942/DC1
Key words: ER-associated degradation, ER stress, PelizaeusMerzbacher disease, Protein misfolding, Unfolded protein response
A variety of pathological conditions are associated with the
production of misfolded proteins at the ER. Some of these diseases,
such as cystic fibrosis, are caused by a loss of function of the relevant
protein, whereas others, such as retinitis pigmentosa, are due to a
toxic gain of function of the misfolded mutant protein (Aridor and
Balch, 1999; Rutishauser and Spiess, 2002). As a result,
considerable progress has been made towards understanding the
molecular mechanisms that mediate ER retention and ERAD of
mutant proteins (Hebert and Molinari, 2007). Interestingly, there is
a number of gain-of-function ER folding diseases in which different
mutations in the same gene result in distinct disease phenotypes
(Nelis et al., 1999). Little is known about whether variations in the
ability of the ER quality-control machinery to deal with different
mutant forms of the same protein contribute to such variations.
In order to address this issue, we examined the cellular response
to different disease-associated mutants of proteolipid protein (PLP).
PLP is a polytopic membrane protein that, together with its splice
variant DM20, represents the major protein constituents of central
nervous system (CNS) myelin. Mutations in the gene encoding PLP
(PLP1) cause Pelizaeus-Merzbacher disease (PMD), an X-linked
recessive leukodystrophy characterised by dysmyelination of the
CNS (Koeppen and Robitaille, 2002; Yool et al., 2000). A variety
of PLP missense mutations, duplications and deletions have been
identified, and these cause a wide spectrum of disease phenotypes
(Garbern, 2007). Because missense mutations usually cause more
severe phenotypes than null mutations (Boison and Stoffel, 1994;
Klugmann et al., 1997), it is believed that disease pathology is
largely induced by the presence of the mutant protein, rather than
by the absence of a functional polypeptide (or loss of function).
Most of the missense mutations studied to date disrupt the trafficking
of PLP to the cell surface (Gow et al., 1994b; Gow and Lazzarini,
Journal of Cell Science
PLP quality control
1996; Woodward, 2008), and it is generally believed that the
pathogenic mechanism underlying PMD is the accumulation of
misfolded PLP in the ER leading to activation of the UPR and
oligodendrocyte apoptosis (Gow and Lazzarini, 1996; Gow et al.,
1998; Southwood et al., 2002). However, the disease caused by
missense mutations can range in phenotypic severity from mild
PMD and late-onset spastic paraplegia type 2 (SPG2), which is
characterised by hypomyelination and thinning of the myelin sheath
(Hodes et al., 1997; Kobayashi et al., 1994), to severe connatal PMD
associated with widespread oligodendrocyte apoptosis and a virtual
absence of compact myelin (Gencic et al., 1989; Hudson et al.,
1989). The critical factors that determine whether a particular
mutation of the gene encoding PLP gives rise to a mild disease with
hypomyelination or a severe disease with oligodendrocyte apoptosis
are currently poorly understood (Dhaunchak and Nave, 2007; Gow
and Lazzarini, 1996; Gow et al., 1998; Kramer-Albers et al., 2006;
Southwood and Gow, 2001).
One possibility that is consistent with the phenotypic variation
observed in PMD patients is that different mutations might have
distinct effects on the ability of the quality-control machinery to
deal with the mutant proteins, thereby producing distinct cellular
responses. To test this hypothesis, we examined the quality control
and associated cellular responses to naturally occurring missense
mutations in similar regions of PLP, which produce a markedly
different phenotype. In particular, we focused on two neighbouring
mutations within the fourth transmembrane (TM) domain of PLP,
one that causes severe PMD and a second that is associated with
mild disease. We find that, although both mutant proteins are
strongly retained in the ER and subjected to ERAD, the efficiency
with which they are degraded and the subsequent cellular response
elicited are quite different. Thus, whereas the A242V mutant protein
associated with severe PMD is relatively resistant to degradation
and induces a robust ER-stress response, the mild-disease-associated
G245A mutant is rapidly cleared from the ER and fails to activate
the UPR. Hence, our studies reveal differential sensitivity of the
ER-associated quality-control and degradation machineries to
different mutant forms of the same protein. These findings have
implications for our understanding of the mechanisms that determine
the severity of PMD, and potentially other gain-of-function proteinfolding diseases.
Results
Retention of disease-associated PLP mutants by the ER
quality-control system
In order to examine how the ER quality-control machinery deals
with different mutant forms of PLP associated with distinct disease
phenotypes, we focused on missense mutations within the TM
domains of the polypeptide, because these might be expected to
induce comparable folding defects. Genotype-phenotype correlation
studies have shown that about 80% of the TM-domain mutations
in the PLP1 gene lead to severe PMD (Cailloux et al., 2000), with
one well-studied example being the substitution of a valine for an
alanine (A242V; msd mutation) in the fourth TM domain (Gencic
and Hudson, 1990; Yamamoto et al., 1998). By marked contrast,
mutation of a neighbouring glycine to alanine (G245A), and of a
tryptophan to leucine (W162L) in the third TM domain, both
produce a milder phenotype (Hubner et al., 2005) (see Fig. 1).
By using an established heterologous expression system
(Swanton et al., 2003), we first tested whether the ER quality-control
machinery is able to recognise and retain the mutant forms of PLP.
Stable HeLa-cell lines expressing the different forms of PLP with
3943
Fig. 1. Locations of the PLP missense mutations under study. A twodimensional model of PLP topology illustrating the four TM domains,
cytosolic N- and C-termini, palmitoylation sites (squiggles), and disulphide
bonds (straight lines). The relative positions of the missense mutations that
have been examined in this study are indicated. The A242V (msd) mutation
associated with severe PMD is in red, whereas the W162L and G245A
mutations associated with mild PMD are in black. The Myc-His6 (mh) epitope
tag at the C-terminus of the recombinant protein is also indicated.
a C-terminal Myc-His6 (mh) tag under the control of a tetracyclineinducible promoter were generated (Fig. 1) and the subcellular
distribution of the proteins examined using immunofluorescence
microscopy. Much of the wild-type (wt)-PLPmh colocalised with
concanavalin A (ConA) bound to cell-surface glycoproteins (Fig.
2A, top row) and was visible on numerous microvilli protruding
from the cell surface (Fig. 2A, insets). Very little colocalisation with
the ER chaperone calnexin was observed (Fig. 2A, centre row),
indicating that the vast majority of the wt protein was properly
folded and had passed the ER quality control. A further pool of wtPLPmh was seen in lysosome-associated membrane protein 1
(Lamp1)-containing late endosomal and/or lysosomal structures
(Fig. 2A, bottom row), most probably representing protein that had
been internalised from the cell surface (Gow et al., 1994a; Simons
et al., 2002). Importantly, a very similar subcellular distribution of
wt-PLP is seen in cultured oligodendrocytes (Dhaunchak and Nave,
2007; Kramer-Albers et al., 2006; Simons et al., 2002; Trajkovic
et al., 2006), demonstrating that this HeLa-cell expression system
provides a suitable model for studying the folding and quality control
of PLP, consistent with previous studies, including our own
(Sinoway et al., 1994; Swanton et al., 2003; Swanton et al., 2005).
In contrast to the wt protein, A242V-PLPmh distribution was
restricted to a reticular network that strongly colocalised with
calnexin (Fig. 2B, centre row), consistent with this mutant being
misfolded and retained by the ER quality-control machinery (Gow
and Lazzarini, 1996; Kramer-Albers et al., 2006; Swanton et al.,
2003). A characteristic feature of HeLa cells expressing A242VPLPmh was the appearance of vacuolar-like structures (Fig. 2B,
insets). The periphery of these structures contained both mutant
PLPmh and calnexin, resembling the distended ER that has been
observed under conditions of chronic ER stress (Alvarez et al., 1999;
Dalal et al., 2004; Lass et al., 2008; Lin et al., 1999). The milddisease-associated mutants W162L and G245A were also present
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3944
Fig. 2. Distinct subcellular distribution of wt and PMD-causing mutant PLP proteins. HeLa cells were induced to express (A) wt-, (B) A242V-, (C) W162L- or (D)
G245A-PLPmh, fixed, permeabilised, and probed with Myc-specific antibodies (green in merge) and antibodies against either the ER-resident protein calnexin or
the late-endosomal and lysosomal protein Lamp1 (red in merge). Alternatively, fixed wt-PLPmh- or mutant-PLPmh-expressing cells were stained with
fluorescently modified ConA to selectively label cell-surface glycoproteins (red in merge) before permeabilisation and immunostaining with anti-Myc antibody.
Colocalisation of PLPmh with specific organelle or plasma-membrane markers appears yellow. Insets (dashed boxes) show magnified areas. Scale bars: 10mm.
in the ER, as evidenced by their colocalisation with calnexin (Fig.
2C,D, centre rows). In addition, a fraction of W162L-PLPmh
seemed to be exported from the ER, and could be seen at the cell
surface and also within Lamp1-positive structures (Fig. 2C, top and
bottom rows). G245A-PLPmh, by contrast, showed no overlap with
the late-endosomal and lysosomal marker (Fig. 2D, bottom row),
and was not obvious at the plasma membrane (Fig. 2D, top row),
suggesting that, similar to the severe-disease-associated A242V
mutant, G245A-PLP is primarily retained in the ER.
In order to provide a quantitative estimate of the amount of each
PLPmh variant reaching the plasma membrane, cells were treated
with a membrane-impermeable biotinylation reagent. Biotinylated
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PLP quality control
3945
Fig. 3. Analysis of cell-surface expression of wt and mutant PLP.
(A)HeLa cells expressing wt or mutant PLPmh were left untreated
(–) or were treated (+) with membrane-impermeant sulpho-NHSSS-biotin. Biotinylated proteins were precipitated using
NeutrAvidin beads (beads, ‘B’) and analysed by immunoblotting
with anti-Myc or anti-TfR antibodies. The biotinylation levels of
wt and mutant PLPmh were compared with 20% total cell lysates
(total, ‘T’). (B)The levels of monomeric and oligomeric PLPmh
on immunoblots were quantified by densitometry. Biotinylated
PLPmh was expressed as a percentage of total PLPmh and
normalised to the equivalent TfR levels. Histogram shows mean ±
s.e.m. from three independent experiments (*P<0.05, **P<0.01,
t-test).
(i.e. cell surface) PLPmh was isolated with NeutrAvidin beads, and
detected by immunoblotting with anti-Myc antibody (Fig. 3A, top
panel). Labelling of plasma-membrane-localised transferrin receptor
(TfR) was used to control for loading and biotinylation efficiency
in each experiment (Fig. 3A, bottom panel). A considerable fraction
of wt-PLPmh was biotinylated and could be bound to NeutrAvidin
beads following treatment of cells with sulpho-NHS-SS-biotin (Fig.
3A, cf. lanes 2 and 4). A significant amount of biotinylated W162LPLPmh was also detected (Fig. 3A, lane 16), consistent with the
subcellular distribution observed by immunofluorescence
microscopy (Fig. 2C). By contrast, virtually no A242V- and even
less G245A-PLPmh was detected in NeutrAvidin-bound fraction
(Fig. 3A, lanes 8 and 12), providing further evidence that these
mutants do not reach the cell surface efficiently (Fig. 3B). The ability
of A242V-PLPmh, but not the wt or mild-disease-associated
mutants, to form SDS-resistant oligomers (Swanton et al., 2005)
was also clearly apparent (Fig. 3A, cf. lanes 1, 5, 9 and 13) and is
discussed in more detail later.
Together, these results are consistent with the hypothesis that
disease-associated mutations perturb the folding of PLP, thereby
preventing authentic trafficking of the protein to the plasma
membrane. Interestingly, the stringency with which the different
mutants are retained by the ER quality-control machinery seems to
vary. Hence, a fraction of W162L-PLPmh clearly exits the ER,
progressing to the plasma membrane and lysosomal compartments.
This suggests that the folding defect induced by this mutation might
be comparatively subtle. By contrast, the A242V and G245A
mutants are primarily retained in the ER. Because these two
mutations produce very different disease phenotypes, they represent
a particularly useful model that allows us to examine how the ability
of the ER quality-control system to deal with different mutant forms
of the same protein might contribute to disease severity.
Degradation of PLP mutants occurs at different rates and via
distinct pathways
The underlying pathogenic mechanism of missense mutations in
PLP is suggested to be the accumulation of the mutant protein in
the ER, which leads to activation of the UPR and oligodendrocyte
apoptosis (Gow and Lazzarini, 1996; Gow et al., 1998; Southwood
and Gow, 2001; Southwood et al., 2002). The extent of PLP
accumulation in the ER is likely to be controlled, at least in part,
by the rate at which the polypeptide is degraded. Thus, the
efficiency with which different mutant proteins are degraded could
be an important factor determining disease pathology. Whereas
plasma-membrane proteins can be endocytosed and degraded in
lysosomes (Luzio et al., 2007), aberrant membrane proteins retained
by the ER quality-control machinery are typically retrotranslocated
and degraded by the ubiquitin-proteasome system in the cytosol
(Vembar and Brodsky, 2008). In order to examine both the route
and rate of degradation of the different forms of PLP, pulse-chase
studies were performed in the presence of well-defined proteasome
or lysosomal-protease inhibitors (Fig. 4A). These experiments
revealed some striking differences in the pathways leading to
degradation of the different PLPmh proteins.
As previously observed, wt-PLPmh is relatively stable in cultured
cells (Kramer-Albers et al., 2006; Swanton et al., 2003), and was
turned over with a half-life of ~6.8 hours (Fig. 4A, wt). Treatment
of cells with a combination of the lysosomal inhibitors leupeptin
and pepstatin A blocked the degradation of the wt protein completely
(Fig. 4A, wt). This suggests that the lysosome is the major site for
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3946
Fig. 4. Pulse-chase analysis of wt- and mutant-PLP degradation. (A)HeLa cells were left uninduced (–) or induced to express wt-, W162L-, A242V- or G245APLPmh. Cells were treated for 1 hour with DMSO, Z-LLF-CHO (10mM) or a mixture of leupeptin (100mM) and pepstatin A (1mg/ml), pulse-labelled for a further
hour with [35S]methionine/cysteine, and then chased with unlabelled methionine/cysteine up to 8 hours in the continued presence of the compounds. [35S]-labelled
PLPmh was recovered by immunoprecipitation with anti-Myc, resolved by SDS-PAGE and visualised by phosphorimaging. The radioactivity of the wt, W162L,
A242V (monomer and dimer) and G245A-PLPmh bands at each time point was quantified by phosphorimaging and expressed as a percentage of the corresponding
value at 0-hour chase. The graphs show the mean ± s.e.m. from at least three independent experiments (*P<0.05, **P<0.01, ***P<0.001, t-test). (B)HeLa cells
were left uninduced (–) or induced to express A242V-PLPmh for 2 or 20 hours, and then subjected to pulse-chase analysis as described in Fig. 4A. [35S]-labelled
monomeric and dimeric A242V-PLPmh was quantified by phosphorimaging as described in Fig. 4A. The graph shows the mean ± s.e.m. from at least three
independent experiments.
degradation of wt-PLPmh, consistent with its lysosomal and plasmamembrane localisation observed by immunofluorescence
microscopy (Fig. 2A). Indeed, accumulation of wt-PLPmh in
Lamp1-positive structures was clearly visible in inhibitor-treated
cells (supplementary material Fig. S1A). The proteasome inhibitor
Z-LLF-CHO also caused some inhibition of degradation of wt-
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PLP quality control
PLPmh (Fig. 4A, wt), reflecting either a minor fraction of the wt
protein that misfolds and is degraded by ERAD or the inhibition
of some lysosomal proteases by Z-LLF-CHO (Lee and Goldberg,
1998) (cf. supplementary material Fig. S1A).
We then examined the degradation of the different mutant forms
of PLPmh. W162L-PLPmh was considerably less stable than the
wt protein, having a half-life of ~4.4 hours (Fig. 4A, W162L). Its
degradation was reduced by the lysosomal-protease inhibitors
leupeptin and pepstatin A, and even more strongly by Z-LLF-CHO
(Fig. 4A, W162L). Lactacystin, a second proteasomal inhibitor,
which is chemically unrelated to Z-LLF-CHO, also significantly
stabilised W162L-PLPmh (supplementary material Fig. S2).
Accumulation of the mutant protein was observed in the ER of ZLLF-CHO-treated cells, and in the late endosomes and/or
lysosomes of cells treated with leupeptin and pepstatin A
(supplementary material Fig. S1B). These data suggest that both
proteasomal and lysosomal pathways are involved in the
degradation of W162L-PLPmh. By contrast, the severe-diseaseassociated A242V mutant was more slowly degraded, with a halflife of ~5.6 hours (Fig. 4A, A242V). Degradation of A242V-PLPmh
was almost entirely inhibited by Z-LLF-CHO, but unaffected by
lysosomal-protease inhibitors (Fig. 4A, A242V). Together with the
ER retention observed by immunofluorescence microscopy (Fig.
2B), this indicates that A242V-PLPmh is degraded via the ERAD
pathway. Similarly, degradation of the mild-disease-associated
G245A mutant was proteasome dependent but unaffected by
inhibition of lysosomal proteases (Fig. 4A, G245A), suggesting
that, similar to the A242V variant, it is also degraded by ERAD.
We found that vacuolar-like structures surrounded by ER membrane
were observed after treatment both of A242V-PLPmh- and G245APLPmh-expressing cells with Z-LLF-CHO (supplementary material
Fig. S1C,D, insets), presumably in response to the inhibition of
proteasomal degradation and increased accumulation of misfolded
mutant PLPmh species in the ER. The role of the proteasome in
the degradation of both of these PLP mutants was further supported
by the accumulation of their polyubiquitylated forms in cells treated
with proteasome inhibitor (supplementary material Fig. S3, square
brackets).
Interestingly, the G245A mutant of PLPmh was degraded
significantly more rapidly than the A242V mutant, with a half-life
of ~3.6 hours (Fig. 4A, G245A). This observation suggested that
the severe-disease-associated A242V variant is more resistant to
ERAD than is the G245A mutant that causes mild PMD. Consistent
with this interpretation, in comparison with A242V-PLPmh,
considerably less radiolabelled G245A-PLPmh was produced during
the 1-hour pulse-labelling period (Fig. 4A, cf. A242V and G245A,
DMSO panels, 0-hour chase). Furthermore, proteasome inhibition
dramatically increased the amount of the radiolabelled G245A
mutant generated during this time (Fig. 4A, G245A, cf. DMSO and
Z-LLF-CHO panels, 0-hour chase), suggesting that rapid
degradation of G245A-PLPmh limits the accumulation of this
mutant protein. We considered the possibility that a simple
accumulation of A242V-PLPmh in the ER could impede the ERAD
pathway and repeated the pulse-chase assay using cells that
contained different levels of total A242V mutant. The rate of
degradation remained constant irrespective of whether cells were
induced to express A242V-PLPmh for 2 hours or 20 hours prior to
pulse-labelling (Fig. 4B). We therefore conclude that the difference
in the rates of degradation of A242V and G245A-PLPmh reflects
a difference in the ability of the ERAD machinery to dispose of
these two mutant proteins.
When taken together, our results (summarised in Table 1)
demonstrate that all three PLP mutants studied are degraded via the
proteasome, reflecting their status as ERAD substrates. However,
for the W162L mutant, ER retention is less stringent and lysosomal
degradation also plays an important role in protein turnover.
Significantly, the two PLP mutants that are associated with mild
PMD are more efficiently cleared from the ER than the severedisease-associated A242V mutant. Of particular interest is the
observation that the A242V and G245A mutants, both of which are
tightly ER-retained, are degraded at different rates, with the milddisease-associated mutant being removed significantly more rapidly
than the mutant linked to severe PMD.
Distinct physical properties of PLP mutants
In order to investigate the molecular basis for the difference in PLP
degradation rates, we examined the physical properties of the mutant
proteins. We have previously shown that a key feature of diseaseassociated PLP mutants is their tendency to rapidly assemble into
SDS-resistant homo-oligomers (Swanton et al., 2005). This
behaviour was apparent in the pulse-chase studies performed here
(Fig. 4 and supplementary material Fig. S4), with SDS-resistant
dimers of the severe-disease-associated mutant A242V clearly
evident immediately after the 1-hour pulse-labelling period (Fig.
4A, A242V, DMSO panel, 0-hour chase). By contrast, the milddisease-associated mutants, particularly the G245A variant, had a
very modest tendency to form stable oligomers (Fig. 4A, W162L
and G245A, DMSO panels). In both cases, dimer formation was
accentuated in the presence of proteasome inhibitors (supplementary
material Fig. S4, W162L and G245A, cf. DMSO and Z-LLFCHO/Lactacystin panels), suggesting that these mutants can form
oligomers if given sufficient opportunity. Hence, there seems to be
a direct correlation between the rapid formation of SDS-resistant
oligomers and the stability of the PLP mutants.
Misfolded proteins, including some mutant PLP, can form
aberrant disulphide bonds (Dhaunchak and Nave, 2007) and might
also be prone to aggregation, either of which could potentially
impede ERAD (Hosokawa et al., 2006). However, no obvious
correlation between the extent of disulphide bonding and the
degradation rates of the different mutants was seen (supplementary
material Fig. S5A), and the sedimentation behaviour of the different
mutants was similar (supplementary material Fig. S5B). These
Table 1. Summary of the localisation and degradation characteristics of the PLPmh missense mutants
Localisation
PLPmh
Disease severity
ER
Late endosomes
and/or lysosomes
wt
W162L
A242V
G245A
–
Mild
Severe
Mild
–
+
+
+
+
+
–
–
Degradation
% cell-surface PLPmh
Approximate
half-life (hours)
Proteosomes
Lysosomes
14.9±2.4
7.5±0.4
0.8±0.1
4.3±2.1
6.8
4.4
5.6
3.6
–
+
+
+
+
+
–
–
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Fig. 5. Expression of A242V-PLP activates IRE1 signalling. (A)HeLa cells were induced to express G245A-PLPmh by using 1mg/ml doxycycline, whereas
expression of A242V-PLPmh was induced with 0.05mg/ml doxycycline (A242VlowDOX). Cells were treated with Z-LLF-CHO (10mM) for 1 hour, pulse-labelled
with [35S]methionine/cysteine in the presence of Z-LLF-CHO for the times indicated, lysed and lysates immunoprecipitated with anti-Myc. Immunoprecipitated
[35S]-labelled PLPmh was quantified by phosphorimaging and expressed as a percentage of the amount of protein synthesised after 10 minutes in order to
normalise differences in cell number. The graph shows the mean from two independent experiments. (B)HeLa cells were induced to express wt-, A242VlowDOX-,
G245A- or W162L-PLPmh for increasing periods of time (0-48 hours). Treatment of uninduced cells with 10 mM DTT for 2 hours served as a positive control to
trigger Xbp1-mRNA splicing. PLPmh was detected by immunoblotting RIPA-solubilised cell lysates with anti-Myc. Tubulin was assessed as a protein loading
control. Alternatively, total RNA was isolated from cells and Xbp1-mRNA splicing was determined by RT-PCR. Unspliced (u) and spliced (s) Xbp1 mRNA are
indicated. GAPDH served as a cDNA loading control. ATF6 immunoblots were performed by analysing RIPA lysates prepared from a second independent
experiment. Treatment of uninduced cells with 2 mM DTT for 2 hours served as a positive control to trigger cleavage of the full-length ATF6 protein (p90 ATF6).
Specificity of the anti-ATF6 antibody was confirmed using a commercial ATF6-transfected cell lysate (Lysate, ‘L’; Imgenex). (C)A242V-PLPmh-expressing HeLa
cells were induced with increasing concentrations of doxycycline (0.01-1mg/ml) for 24 hours. Cell lysates were analysed by western blotting with anti-Myc or antitubulin, and Xbp1-mRNA splicing was determined by RT-PCR.
results suggest that neither variable formation of aberrant oxidation
products nor variable aggregation of the mutant proteins underlies
the differences in the rates of degradation observed.
Differential activation of the UPR by PLP mutants
The UPR is believed to modulate disease severity in PMD
(Southwood et al., 2002), and we therefore compared the magnitude
of the UPR induced by expression of the disease-associated mutant
forms of PLP. In this context, we focussed on the two primarily
ER-retained mutants, A242V and G245A, because they are degraded
via ERAD at different rates and are associated with diseases of
varying severity. For such comparative analyses, it was important
to ensure that the rate of synthesis of the two proteins was similar.
This was achieved by lowering the concentration of doxycycline
used to induce expression of A242V-PLPmh (Fig. 5A).
We first examined the ER-stress response to the synthesis of these
mutant forms of PLP by monitoring the IRE1-XBP1 branch of the
UPR (Yoshida et al., 2001). Treatment of cells with the reducing
agent dithiothreitol (DTT), a potent inducer of the UPR, resulted
in the appearance of spliced Xbp1 mRNA (Fig. 5B, Xbp1 panel;
cf. lanes 1 and 2, 9 and 10, 17 and 18, 25 and 26). By contrast,
Xbp1-mRNA splicing was not visible in cells expressing wtPLPmh, demonstrating that synthesis of the properly folded protein
did not induce the UPR over a 48-hour period (Fig. 5B, Xbp1 panel,
lanes 2-8). Xbp1-mRNA splicing was measured at several different
times following induction of mutant PLPmh expression, because
UPR signalling can diminish with time (Lin et al., 2007; Rutkowski
et al., 2006). The severe-disease-associated A242V-PLPmh
accumulated rapidly in cells following addition of doxycycline (Fig.
5B, Myc panel, lanes 10-16). Spliced Xbp1 mRNA was clearly
Journal of Cell Science
PLP quality control
visible after 8 hours (Fig. 5B, Xbp1 panel, lane 12, see arrow),
demonstrating that expression of this mutant protein induces a rapid
ER-stress response. Interestingly, prolonged expression of A242VPLPmh at low levels of induction resulted in attenuation of Xbp1mRNA splicing (Fig. 5B, Xbp1 panel, lanes 14-16), consistent with
observations that IRE1 activation is reduced during persistent ER
stress (Lin et al., 2007; Rutkowski et al., 2006). Despite having a
similar rate of synthesis to A242V-PLPmh (Fig. 5A), less G245APLPmh accumulated in doxycycline-induced cells (Fig. 5B, Myc
panel, cf. lanes 10-16 and 18-24), reflecting the more efficient
degradation of this mild-disease-associated mutant. No spliced Xbp1
mRNA could be detected in cells expressing G245A-PLPmh, even
after 48 hours of continual synthesis of the mutant protein (Fig. 5B,
Xbp1 panel, lanes 18-24). This is an important finding because it
suggests that efficient removal of G245A from the ER by ERAD
protects the cells by preventing the activation of the UPR. Likewise,
expression of W162L-PLPmh, which is effectively cleared from
the ER by a combination of ERAD and export, failed to induce
Xbp1-mRNA splicing (Fig. 5B, Xbp1 panel, lanes 26-32). Thus,
there is a clear correlation between the stability, and hence load, of
the different mutant forms of PLP in the ER and their ability to
induce the IRE1-XBP1 arm of the ER-stress response. The
dependence of IRE1 activation on misfolded protein load was
confirmed by varying the steady-state level of A242V-PLPmh using
different doxycycline concentrations (Fig. 5C). As predicted, the
extent of Xbp1-mRNA splicing increased in proportion to total levels
of the mutant protein (Fig. 5C), and this splicing was maintained
when the mutant was expressed at high levels (supplementary
material Fig. S6).
We also examined activation of a second UPR sensor, ATF6, a
90-kDa transmembrane protein that is proteolytically activated under
conditions of ER stress. The ATF6 cleavage product is unstable,
and therefore ATF6 activation is often measured by following loss
3949
of the full-length protein (DuRose et al., 2006; Haze et al., 1999;
Rutkowski et al., 2006). Induction of the UPR with DTT resulted
in the disappearance of virtually all full-length ATF6 (Fig. 5B, ATF6
panel, cf. lanes 1 and 2, 9 and 10, 17 and 18, 25 and 26),
demonstrating efficient ATF6 cleavage. By contrast, no loss of p90
ATF6 was observed upon induction of wt-, G245A- or W162LPLPmh expression (Fig. 5B, ATF6 panel, lanes 2-8, 18-24, 26-32).
This suggests that synthesis of these forms of PLP did not stimulate
ATF6 cleavage, and is consistent with their inability to activate
IRE1. Expression of A242V-PLPmh also failed to induce observable
cleavage of p90 ATF6 (Fig. 5B, ATF6 panel, lanes 10-16), despite
causing activation of IRE1-XBP1 (Fig. 5B, Xbp1 panel, lanes 12
and 13). Because measuring the disappearance of full-length ATF6
might not be sufficiently sensitive to detect a small degree of ATF6
cleavage, we increased A242V-PLPmh expression using a higher
doxycycline concentration. However, no evidence of ATF6 cleavage
was seen even upon prolonged high-level expression of the mutant
protein, which induced robust Xbp1 splicing (supplementary
material Fig. S6, cf. Xbp1 and ATF6 panels). Therefore, we
conclude that expression of A242V-PLPmh does not efficiently
activate ATF6 in this system.
In order to examine the consequences of UPR activation by
mutant PLP, we focused on the transcription factor CHOP, which
is upregulated by ER stress and is a key mediator of ER-stressinduced apoptosis (Oyadomari and Mori, 2004). Levels of CHOP
mRNA, assessed by reverse-transcriptase PCR (RT-PCR), were
low under basal conditions and increased markedly upon treatment
of cells with DTT (Fig. 6A, cf. lanes 1 and 2, 9 and 10, 17 and
18, 25 and 26, 33 and 34). The abundance of CHOP mRNA was
not altered by expression of wt-PLPmh, or of the G245A and
W162L variants (Fig. 6A, lanes 2-8, 18-24 and 26-32), providing
further evidence that the rapidly cleared mutant proteins do not
induce the UPR. By contrast, synthesis of the severe-disease-
Fig. 6. Expression of A242V-PLP upregulates CHOP expression
and decreases cell viability. (A)HeLa cells were induced to
express wt-, A242V-, G245A- or W162L-PLPmh for increasing
periods of time (0-48 hours). Total RNA was isolated and levels
of CHOP mRNA determined by RT-PCR. Treatment of
uninduced cells with 5 mM DTT for 4 hours served as a positive
control to induce CHOP mRNA levels. GAPDH served as a
cDNA loading control. (B)HeLa cells were induced to express
wt-, A242V-, G245A- or W162L-PLPmh for the indicated
periods of time. Cell viability was measured via the MTT assay,
and expressed relative to uninduced cells. The graph shows the
mean ± s.e.m. from three experiments.
Journal of Cell Science
3950
Journal of Cell Science 122 (21)
associated A242V mutant at a similar rate caused a dramatic
increase in CHOP mRNA within 8 hours (Fig. 6A, lanes 10-16).
As observed for IRE1-XBP1, an even greater upregulation of
CHOP mRNA was provoked by increased levels of A242VPLPmh (Fig. 6A, lanes 34-40). Because CHOP is generally
associated with proapoptotic UPR signalling (Oyadomari and
Mori, 2004), we next examined the effect of the different mutant
proteins on cell viability. In line with their inability to induce any
of the UPR markers tested, wt-PLPmh and the mild-diseaseassociated W162L and G245A mutants had little effect on cell
viability (Fig. 6B). Cells also tolerated similar rates of A242VPLPmh synthesis, showing only a very marginal decrease in
viability after 48 hours (Fig. 6B). However, inducing higher
expression levels of this mutant resulted in a clear loss of cell
viability (Fig. 6B), consistent with the idea that a threshold of
CHOP expression is required to promote apoptosis (Rutkowski et
al., 2006). These findings further highlight the correlation between
load of misfolded protein in the ER, UPR signalling and cell fate.
Together, these results suggest that the rate at which mutant
proteins are cleared from the ER is an important factor in
determining their ability to induce the UPR. Consistent with a role
of the UPR in determining PMD pathology, the two mutant forms
of PLP that cause mild PMD are degraded comparatively quickly
and fail to activate the UPR, whereas the more stable severe-diseaseassociated variant rapidly induces an ER-stress response, resulting
in upregulation of the UPR target gene CHOP.
Discussion
It is believed that the pathology of PMD, and many other gain-offunction ER folding diseases, is caused by activation of the UPR,
which leads to programmed cell death. Thus, factors that govern
the extent and duration of UPR activation in response to expression
of different mutant proteins are likely to play an important role in
determining disease pathology. However, little is known about the
nature of these factors, and the molecular basis for the variation in
the severity of PMD remains unclear. In order to address this issue,
we used a stable inducible HeLa-cell system to describe the
fundamental cellular events associated with expression of three
different mutant forms of PLP associated with diseases of varying
severity. Together, our results suggest that the rate at which different
mutant proteins are cleared from the ER determines their ability to
activate an ER-stress response and might thus play an important
role in determining disease pathology. Hence, whereas the more
aggressive A242V mutant has an inherently slower turnover and
induces ER stress, the mild-disease-associated W162L and G245A
variants are more rapidly degraded and fail to activate the UPR.
Immunofluorescence microscopy and quantitative cell-surface
biotinylation revealed that all three mutant forms of PLP were
subjected to some degree of ER retention. Of particular interest was
the observation that both the A242V and G245A mutants were
almost exclusively ER localised. These mutations are associated
with severe and mild forms of the disease, respectively, and
therefore ER retention alone cannot explain the differences in
phenotypic severity. A fraction of the W162L mutant was able to
pass the ER quality control, reaching the cell surface and late
endosomes and/or lysosomes. By contrast, a GFP-tagged version
of the W162L mutant transiently expressed in COS-7 cells was
found to accumulate in a perinuclear compartment (Koizume et al.,
2006). This apparent discrepancy might be due to the general
tendency of PLP to be retained intracellularly in COS-7 cells
(Kramer-Albers et al., 2006; Thomson et al., 1997). Importantly,
trafficking of wt-PLP in the inducible HeLa cells used here mimics
that seen in oligodendroglial cells (Dhaunchak and Nave, 2007;
Kramer-Albers et al., 2006), with very little, if any, wt-PLP
detectable in the ER. Thus, we are confident that inducible
expression of PLP in HeLa cells provides a valuable system with
which to analyse the cellular processing of mutant forms of this
protein, and that our results reveal fundamental principles about
this process.
Significantly, the A242V and G245A mutants were both retained
in the ER and degraded via ERAD, yet they produce distinct disease
phenotypes. This observation provided us with a useful model with
which to examine whether variations in the ability of the ER qualitycontrol machinery to deal with different mutant forms of the same
protein could underlie differences in cellular pathology. Strikingly,
the two mutant proteins were turned over with very different
kinetics. Although both were degraded via ERAD, G245A-PLP was
degraded significantly faster than the A242V mutant. As a result,
A242V-PLP accumulated at the ER to a greater extent. Importantly,
when synthesised at comparable rates, only the more stable A242V
mutant induced the UPR. These results suggest that the efficiency
of ERAD is an important factor in determining the extent of UPR
activation caused by expression of different mutant forms of the
same protein (see model in Fig. 7).
The precise mechanism through which the proximal ER-stress
sensors are activated is not yet clear, but for IRE1 might involve
direct binding of misfolded proteins to the lumenal domain of the
protein (Credle et al., 2005; Zhou et al., 2006). Thus, it is formally
possible that the difference in IRE1 induction by the two ER-retained
mutant proteins reflects variations in their ability to bind to IRE1.
Nevertheless, when the steady-state level of A242V-PLP was
decreased sufficiently, this mutant also failed to induce Xbp1-mRNA
splicing (Fig. 5B,C). This supports our suggestion that the extent
of UPR activation reflects differences in the stability of mutant
polypeptides, and thus the level to which they accumulate in the
ER, and not some other, as-yet poorly defined, property of particular
Fig. 7. Schematic for the postulated molecular
mechanism underlying mild and severe PMD
resulting from different PLP missense mutations.
Journal of Cell Science
PLP quality control
mutant proteins. Recent work indicates that the three stress sensors
in mammalian cells (IRE1, ATF6 and PERK) possess distinct
sensitivity to different forms of ER stress (DuRose et al., 2006).
Such a model would be consistent with our observation that
expression of A242V-PLPmh failed to induce efficient cleavage of
ATF6, despite causing robust activation of IRE1 and induction of
CHOP mRNA (Fig. 6A and supplementary material Fig. S6).
Because the PERK pathway plays a dominant role in CHOP
upregulation during ER stress (Harding et al., 2000; Oyadomari
and Mori, 2004), the increase in CHOP seen here most likely reflects
activation of PERK, suggesting that expression of A242V-PLPmh
activates both IRE1 and PERK.
The half-life of A242V-PLP reported here (5.5 hours) is longer
than that observed in immortalized oligodendrocytes (3 hours)
(Kramer-Albers et al., 2006). This might be due to differences in
the expression systems used, the timecourse over which degradation
was followed and/or intrinsic differences in the ERAD capacity of
the cells. Nevertheless, regardless of the absolute rate of A242VPLP degradation, our findings clearly support a model in which
variation in the relative stability of different mutant forms of PLP
in the ER underlies their ability to induce ER stress. This correlation
between mutant-protein stability and activation of the UPR extended
to the W162L mutant, which was also degraded comparatively
rapidly and failed to induce UPR signalling. In this case, a
considerable fraction of the mutant protein escaped the ER qualitycontrol system and was directed to lysosomes, either via post-ER
quality-control systems (Arvan et al., 2002) or following the
normal trafficking pathway of the wt protein (Simons et al., 2002).
In either case, forward transport of W162L-PLP probably
contributed to the effective clearance of this mutant protein from
the ER and its failure to induce ER stress. Similarly, a recent study
showed that a proportion of the I186T (rsh) mutant of PLP, which
also causes mild disease, exits the ER in immortalized
oligodendrocytes and is degraded by proteasome- and lysosomedependent pathways (Kramer-Albers et al., 2006). In contrast to the
W162L mutant studied here, Kramer-Albers et al. found that the
half-life of I186T-PLP was comparable to that of the severe-diseaseassociated mutant A242V-PLP. This difference in stability of the
two ‘mild’ mutants relative to A242V-PLP probably reflects
variations in their trafficking efficiency and/or partitioning between
proteasomal and lysosomal degradation pathways. In any event, the
results of Kramer-Albers et al. support our hypothesis that the ability
of certain mutant proteins to partially escape ER quality control
reduces their capacity to activate the UPR, therefore disrupting cell
function (Fig. 7).
The ‘set point’ that determines whether cells survive or die
when faced with ER stress varies between cell types, and there
is some evidence that myelinating cells might have a distinct
response to UPR activation (Lin and Popko, 2009; Southwood
et al., 2002). For example, CHOP, which is generally considered
to promote apoptosis, seems to have a protective effect in
oligodendrocytes (Southwood et al., 2002). As a result, the
ultimate consequences of UPR activation in HeLa cells might
not be directly comparable to those in oligodendrocytes. However,
expression of A242V-PLPmh in HeLa cells was accompanied by
induction of CHOP, as seen in oligodendrocytes from mice and
humans harbouring PLP1 mutations (Southwood et al., 2002),
suggesting that this early response to expression of mutant PLP
is conserved. In the case of HeLa cells, high-level A242V-PLPmh
expression was also correlated with an increase in cell death,
although further studies using oligodendrocytes will be needed
3951
to fully elucidate the consequences of UPR activation in these
specialised cells.
Because the UPR is believed to modulate the pathology of PMD,
these results provide insight into the molecular mechanisms that
govern the severity of this disease. Our data suggest that the extent
of UPR activation is determined, at least in part, by the rate at which
the mutant protein is cleared from the ER. Removal from the ER
might occur via forward trafficking, or through effective ERAD,
either of which seems to minimise UPR signalling. These findings
might also be relevant to other ER folding diseases whose pathology
is dictated by the UPR.
We previously established a direct correlation between the extent
of mutant PLP oligomerisation and the severity of PMD (Swanton
et al., 2005), and our current data support the proposal that there
is a link between the stability of PLP mutants and the rate of homooligomerisation (Fig. 7). The observation that the A242V and
G245A mutant proteins are degraded with different efficiencies and
exhibit distinct oligomeric properties suggests that these missense
mutations produce distinct effects on the protein structure, despite
being located in the same regions of the polypeptide. Although the
high-resolution structure of PLP has not yet been resolved,
modelling the fourth TM domain of PLP predicts that the two
mutations produce similar structural alterations (supplementary
material Fig. S7). Because the presence of b-branched valine
residues in TM helices can facilitate helix-helix interactions by
providing a conformationally restricted interface (Curran and
Engelman, 2003), we speculate that the A242V substitution might
specifically increase the dimerisation interface of this mutant, thus
increasing the propensity of A242V-PLP to form dimers.
In conclusion, our studies provide evidence for a link between
the stability of different mutant forms of PLP, their ability to induce
the UPR and the severity of the disease that they cause. Thus, we
suggest that the efficiency of ERAD is an important factor governing
the extent of UPR activation in response to expression of different
mutant proteins. These findings have implications for other gainof-function ER folding diseases in which the UPR modulates disease
pathology, and suggest that strategies aimed at facilitating the
removal of mutant proteins from the ER continue to have
considerable therapeutic potential (Powers et al., 2009).
Materials and Methods
Reagents and antibodies
Media and reagents were from Lonza (Wokingham, UK), Z-LLF-CHO from
Calbiochem, protease-inhibitor cocktail from Sigma, and leupeptin, pepstatin A and
lactacystin from BIOMOL (Exeter, UK). Antibodies were from Upstate (Myc epitope),
Sigma (Myc epitope and calnexin), Developmental Studies Hybridoma Bank,
University of Iowa (Lamp1), BIOMOL (polyubiquitin conjugates, clone FK2), Zymed
(TfR) and Imgenex (ATF6). Anti-a-tubulin was kindly provided by Keith Gull
(University of Oxford, UK). Fluorophore- and HRP-conjugated secondary antibodies
were from Molecular Probes (Paisley, UK) and Sigma, respectively.
Molecular cloning and DNA manipulations
The cDNA encoding wt-PLPmh (Swanton et al., 2003) was subcloned into the NotI
and EcoRV sites of the tetracycline-responsive expression vector pTRE2hyg
(Clontech), and QuikChange site-directed mutagenesis (Stratagene) was used to
generate the W162L or G245A point mutations.
Cell culture
Inducible HeLa-cell lines expressing W162L and G245A-PLPmh were generated by
transfection of HeLa Tet-On cells (Clontech) with the relevant pTRE2hyg plasmids
using Lipofectamine 2000 (Invitrogen). Stable transfectants were selected by
resistance to 400 mg/ml hygromycin B. Clonal cell lines were chosen based on the
level of inducible expression, and maintained in DMEM containing 10% foetal bovine
serum and 2 mM L-glutamine, plus 100 mg/ml geneticin and 100 mg/ml hygromycin
B at alternate passages. Expression was induced by 1 mg/ml doxycycline and analyses
were conducted after 20-24 hours, unless stated otherwise.
3952
Journal of Cell Science 122 (21)
Immunofluorescence microscopy
Cells grown on coverslips were fixed with 3% formaldehyde and 0.2% glutaraldehyde
in PBS for 30 minutes. After unreacted fixative was quenched, cells were
permeabilised with 0.1% Triton X-100 and 0.05% SDS in PBS for 4 minutes, then
incubated for 60 minutes with primary antibodies diluted into PBS. Cells were washed
with PBS and incubated with the appropriate fluorophore-conjugated secondary
antibodies diluted in PBS for another 60 minutes. After washing with PBS, coverslips
were mounted with Mowiol and analysed using an Olympus BX60 upright microscope
driven by MetaMorph software (Universal Imaging Corporation).
Biotinylation of cell-surface proteins
Cell-surface proteins were biotinylated using 0.25 mg/ml sulpho-NHS-SS-biotin
(Pierce) in PBS for 30 minutes at 4°C. Unreacted biotin reagent was quenched by
adding ice-cold Tris-HCl, pH 7.6, to 50 mM. Cells were washed with PBS, lysed
in RIPA buffer (10 mM Tris-HCl, pH 7.6, 150 mM NaCl, 1% NP-40, 0.1% SDS,
0.2% deoxycholate) containing protease-inhibitor cocktail and incubated at 4°C for
1 hour. Insoluble material was removed by centrifugation at 5000 g for 10 minutes
at 4°C. Biotinylated conjugates were affinity-purified with NeutrAvidin resin
(Pierce) at room temperature for 60 minutes, resolved by SDS-PAGE and detected
by immunoblotting.
Journal of Cell Science
Pulse-labelling/chase analysis
Cells were incubated in methionine/cysteine-free DMEM for 20 minutes, then pulselabelled in the same medium containing 20 mCi/ml EasyTag [35S]methionine/cysteine
(PerkinElmer) for the indicated periods of time. After pulse-labelling, cells were rinsed
twice with PBS, then incubated with normal growth medium containing 4 mM nonradioactive methionine and cysteine for up to 8 hours (chase). At the indicated times
of pulse (pulse-labelling analysis) or chase (pulse-chase analysis), cells were rinsed
twice with ice-cold PBS and lysed in immunoprecipitation (IP) buffer (10 mM TrisHCl, pH 7.6, 140 mM NaCl, 1 mM EDTA, 1% Triton X-100) containing proteaseinhibitor cocktail at 4°C for 1 hour. After centrifugation at 5000 g for 10 minutes at
4°C, clarified cell lysates were processed for denaturing IP as described (Swanton
et al., 2003). Radiolabelled proteins were visualised by SDS-PAGE and
phosphorimaging. [35S]-labelled PLPmh was quantified using AIDA software, and
GraphPad Prism 4 software was used for data and statistical analysis. The mean values
from the pulse-labelling studies were fitted with one-phase-exponential-association
curves according to the formula y  ymax ⫻ [1–exp(–kx)] (where krate constant and
xtime), whereas the averaged data from the pulse-chase studies were fitted with
one-phase-exponential-decay curves according to the formula y  100 ⫻ exp(–kx).
The decay rate constant (k), which is representative of the rate of protein degradation,
was determined for each curve and the half-life calculated (t1/20.69/k).
RT-PCR analysis of Xbp1-mRNA splicing and CHOP-mRNA
induction
Cells were harvested in Trizol (Invitrogen) and total RNA was extracted according to
the manufacturer’s instructions. RNA (1 mg) was reverse-transcribed into singlestranded cDNA using the AMV-RT system (Roche), and the cDNA used as template for
PCR amplification of Xbp1, CHOP or GAPDH mRNA. Primers used for RT-PCR
analysis included: Xbp1, 5⬘-ACAGCGCTTGGGGATGGATG-3⬘ and 5⬘-TGACTGGGTCCAAGTTGTCC-3⬘; CHOP, 5⬘-CCCTGCTTCTCTGGCTTGGCTGAC-3⬘
and 5⬘-GCTCCCAATTGTTCATGCTTGGTGC-3⬘; and GAPDH, 5⬘-GACCCCTTCATTGACCTCAACTACATG-3⬘ and 5⬘-GTCCACCACCCTGTTGCTGTAGCC-3⬘. PCR products were resolved on 2% agarose gels.
Cell-viability assay
Cell viability was assessed by measuring their ability to metabolise 3-(4,5dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT; Sigma). 500 ml of 2.5
mg/ml MTT was added to each well of a 24-well plate and cells incubated at 37°C
for 2 hours. The media was discarded and reduced crystals of MTT solubilised with
500 ml DMSO. 150 ml of DMSO solution was transferred to a well of a 96-well plate
so as to create triplicates of each sample, and absorbance at 570 nm was measured
with an automatic plate reader.
This work was supported by The Wellcome Trust. We thank Martin
Lowe, Craig McKibbin and Jim Warwicker for critical comments on
this manuscript and helpful discussions. Deposited in PMC for release
after 6 months.
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