1 Division of Pathology, Institute of Ophthalmology, University College London,
London, UK
2 Electron Microscopy, Institute of Ophthalmology, University College London,
London, UK
* Author for correspondence (e-mail: michael.cheetham{at}ucl.ac.uk )
Accepted 1 May 2002
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Summary |
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Key words: Rhodopsin, Aggresome, Proteasome, Chaperone, Glycosylation, Retinitis pigmentosa
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Introduction |
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Several of these mutations have been studied in detail at the cellular and
transgenic animal level. These studies suggest that rhodopsin mutations can be
divided into two major categories on the basis of the mechanisms of
pathogenesis. Mutations at the C-terminus of rhodopsin interfere with the
normal targeting of the protein to the photoreceptor outer segment
(Sung et al., 1991;
Sung et al., 1994
;
Tam et al., 2000
), whereas
mutations in the transmembrane, intradiscal or cytoplasmic domains result in
the misfolding of the protein (Kaushal and
Khorana, 1994
; Olsson et al.,
1992
; Roof et al.,
1994
; Sung et al.,
1991
; Sung et al.,
1993
). The mechanisms by which these misfolded and misrouted
proteins lead to photoreceptor death by apoptosis
(Portera Cailliau et al.,
1994
), however, remain unidentified.
In transfected cells, rhodopsin with mutations in the transmembrane,
intradiscal or cytoplasmic domains fails to translocate to the plasma membrane
and accumulates within the cell, in what has been described as the ER and
Golgi (Kaushal and Khorana,
1994; Sung et al.,
1991
; Sung et al.,
1993
). These mutant proteins trapped within the cell cannot form
the visual pigment with 11-cis-retinal
(Kaushal and Khorana, 1994
;
Sung et al., 1991
;
Sung et al., 1993
) and are
found in a complex with the ER-resident chaperones BiP and Grp94
(Anukanth and Khorana, 1994
),
supporting the notion that they are incorrectly folded. It appears, therefore,
that these mutations result in a protein that cannot translocate to the plasma
membrane in cultured cells or to the outer segment in photoreceptors because
it is misfolded.
The failure of rhodopsin to translocate to the outer segment per se does
not appear to be enough to cause retinitis pigmentosa, as heterozygous
rhodopsin knockout mice display little photoreceptor cell death
(Humphries et al., 1997).
Rather, it would appear that misfolded rhodopsin acquires a `gain of function'
that leads to cell death. Recent insights into the fate of misfolded proteins
in mammalian cells has suggested that there may be a common cellular response
to unfolded proteins, the formation of a specialised structure, the aggresome
(Kopito, 2000
). Aggresomes are
thought to result from a saturation of the normal proteolytic machinery by
misfolded proteins and accumulate as ubiquitinated inclusions near the
centrosome. Other characteristic features of aggresomes are the disruption of
the Golgi and intermediate filament networks and the recruitment of cellular
chaperones (Garcia-Mata et al.,
1999
; Johnston et al.,
1998
; Kopito,
2000
).
The presence of ubiquitinated, proteinacious inclusions within neurones is
associated with many forms of neurodegeneration, including Amyotrophic Lateral
Sclerosis, Alzheimer's disease, Parkinson's disease and several hereditary
diseases caused by expansions of polyglutamine repeats (e.g., Huntington's
disease and the spinocerebellar ataxias)
(Alves-Rodrigues et al., 1998;
Kaytor and Warren, 1999
;
Sherman and Goldberg, 2001
).
Studies of the inclusions formed in some of these diseases and the
pathogenetic basis of mutations that lead to some neurodegenerations have
suggested that they share features with aggresomes
(Johnston et al., 2000
;
Waelter et al., 2001
). Indeed,
a recent study of P23H mutant opsin expression in cell culture has shown that
mutant opsin aggregates colocalise, but do not co-aggregate, with other
proteins that form aggresomes, such as
F508 CFTR and TCR
subunits (Rajan et al., 2001
).
In this study, we have further investigated the fate of mutant rhodopsin in
vitro, identifying the formation of ubiquitinated mutant opsin aggresomes and
delineating mechanisms of mutant opsin quality control and degradation. The
relevance to RP is further highlighted by the observation that mutant opsin
can recruit the normal protein to aggresomes.
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Materials and Methods |
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Cell culture
COS-7 cells were grown in DMEM/F12 with Glutamax-I+10% heat inactivated FBS
and 50 µg/ml gentamicin with an atmosphere of 5% CO2 at
37°C. 24 hours after seeding glass eight-well chamber slides with
2x104 cells per well, the cells were transfected with 0.2
µg DNA per well with 2 µl of plus reagent and 1 µl of lipofectamine
according to the manufacturers instructions. For 35 mm dishes, the
transfection mix was scaled up and cells were transfected 24 hours after
seeding with 106 COS-7 cells. For Golgi disruption experiments,
brefeldin A (BFA) was added to the cells to a final concentration of 20
µg/ml for 15 minutes prior to fixation. For proteasome inhibition, cells
were treated with MG-132 at 5 µM for the indicated time and a parallel
transfection of F508 CFTR-GFP was used as a positive control for
aggresome formation via proteasome inhibition. N-linked glycosylation was
inhibited by the addition of tunicamycin to a final concentration of 0.8
µg/ml for the indicated treatment time; the efficiency of inhibition was
confirmed by PNGase F treatment of samples. The incidence of aggresome
formation was estimated by scoring 400 transfected cells from four separate
experiments for the presence of aggresomes by an observer that was blind to
the cell transfection status.
Immunofluorescence
The following fixation conditions were used for each antibody: for
vimentin, Hsc70, ß-COP and c-myc ubiquitin staining, COS-7 cells were
fixed in 100% methanol at -20°C for 6 minutes then washed twice in PBS;
for BiP (Grp78) staining, cells were fixed in 3.7% formaldehyde in PBS for 20
minutes then washed three times in PBS for 5 minutes then permeabilised in
0.05% Triton X-100 for 5 minutes followed by two 5 minute washes in PBS. For
calnexin staining, cells were fixed in 3% formaldehyde/0.1% glutaraldehyde in
0.08 M sodium cacodylate-HCl buffer, pH 7.4 for 20 minutes followed by two 5
minute washes in PBS and one 5 minute wash in 50 mM NH4Cl in PBS,
followed by one rinse in PBS. Cells were permeabilised in 0.1% Triton X-100
and 0.05% SDS for 4 minutes followed by three 5 minute washes in PBS.
Following fixation and permeabilisation, cells were blocked for 1 hour at
22°C with 10% appropriate normal serum, prior to incubation with
antibodies. The primary antibodies were incubated for 1 hour at 22°C,
followed by three 5 minute washes in PBS, followed by a 1 hour incubation at
22°C with conjugated secondary antibodies and then washed with four washes
for five minutes in PBS; DAPI was used at 1:5000 in PBS for 5 minutes in the
third wash. The titres of antibodies used were: mouse anti-ß-COP 1:50
dilution in PBS+10% normal goat serum (NGS) followed by goat anti-mouse Cy3
1:500 in PBS+10% NGS; mouse anti-vimentin 1:10 dilution in PBS + 10% NGS
followed by goat anti-mouse Cy3 1:500 in PBS+10% NGS; mouse anti-Hsc70(BRM22)
1:50 dilution in PBS+10% NGS followed by goat anti-mouse Cy3 1:200 in PBS+10%
NGS; anti-c-myc hybridoma supernatant, clone 9E10, was used neat followed by
goat anti-mouse Cy3 1:500 in PBS+10% NGS; ERGIC-53 1:1000 dilution of mouse
ascites in PBS+10% NGS followed by goat anti-mouse Cy3 1:500 in PBS+10% NGS;
rabbit anti-calnexin 1:100 in PBS+10% normal donkey serum (NDS) followed by
donkey anti-rabbit Cy3 1:200 in PBS+10% NDS; goat anti-BiP 1:40 in PBS+10% NDS
followed by donkey anti-goat Cy3 1:200 in PBS+10% NDS. For dual labelling,
opsin was visualised with mAb 1D4 conjugated to FITC, at a concentration of
2.5 µg/ml in PBS+10% normal mouse serum and added after primary and
secondary antibodies had bound and after three 5 minute washes in PBS.
Immunofluorescence was visualised with a Zeiss LSM 510 laser scanning confocal
microscope. The following excitation/emission conditions were used in separate
channels with either the x40 or x63 oil immersion objective: DAPI
364/475-525 nm; FITC/GFP 488/505-530 nm; Cy3 543/560 nm.
Electron microscopy
Cells were fixed overnight in Karnovsky's fixative (3% glutaraldehyde, 1%
paraformaldehyde in 0.08 M sodium cacodylate-HCl buffer, pH 7.4) then rinsed
three times in PBS and incubated in a 1% aqueous osmium tetroxide solution for
1 hour at room temperature. Following three rinses in distilled water, cells
were dehydrated by 10 minute immersions in 50%, 70%, 90% and 4x100%
ethanol. Finally, the wells containing dehydrated cells were filled with
araldite resin and placed in a 60°C oven to harden overnight. Sections for
examination by transmission electron microscopy were cut using a Leica
ultracut S microtome and diamond knife, mounted on copper grids and contrasted
with lead citrate. Images were viewed using a JEOL 1010 TEM operating at 80 kV
and photographs recorded onto Kodak electron image film.
Preparation of cell extracts
24 hours after transfection, cells were treated with 5 µM MG-132 for 16
hours or 0.8 µg/ml Tunicamycin for 16 hours in DMEM/F12 with Glutamax-I+10%
FBS without antibiotics. Cells were washed twice in 4°C PBS and incubated
in 290 µl PBS/1% n-Dodecyl-ß-D-Maltoside plus 10 µl of protease
inhibitor cocktail and 166 µM MG-132 for 30 minutes on ice. Cell lysates
were transferred to 1.5 ml microcentifuge tubes on ice and homogenised by
passage through a 23G needle five times. 200 µl of cell lysate was
centrifuged at 17,500 g for 15 minutes at 4°C. Pellets
were solublised in 50 µl 10 mM Tris·HCl pH 7.5, 1% SDS+2 µl
protease inhibitor cocktail for 5 minutes at 22°C. 150 µl of RIPA
buffer (50 mM Tris-HCl pH 8, 150 mM NaCl, 1 mM EDTA, 1% NP-40, 0.1% SDS, 0.05%
sodium deoxycholate) was added, and the pellets were then sonicated for three
5 second bursts. For SDS PAGE, an equal volume of 2x modified Laemmli
sample buffer (1xsample buffer, 0.125 M Tris-HCl, pH 6.8, 5% SDS, 15%
glycerol, 10% 2-mercaptoethanol 0.012% bromophenol blue) was added to the
soluble and insoluble fraction. For immunoblotting, cell fractions were
normalised for total protein and separated by 10% SDS-PAGE and electroblotted
onto nitrocellulose. For immunodetection of opsin, mAb 1D4 was used at a
concentration of 0.5 µg/ml and goat anti-mouse HRP (Pierce) was used at
1:50,000 in PBS+5% Marvel (Premier Brands), 0.1% Tween-20. The
chemiluminescent detection reagent ECL Plus (Amersham Pharmacia Biotech) was
used to detect immobilised antigens according to the manufacturer's
instructions. The electrophoretic mobility of different opsin glycoforms was
determined empirically using PNGase F and Endo H; 15 µg protein was
digested with PNGase F (1500 units) or Endo H (1500 units) for 2 hours at
37°C in 1xG7 (PNGase) or 1xG5 (Endo H) buffer (NEB) before
resolving by SDS-PAGE as described above.
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Results |
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To confirm that the mutant protein was not present in the Golgi we used brefeldin A (BFA) to disrupt the Golgi apparatus. Treatment with BFA dispersed the Golgi in all the cells and dispersed the wild-type opsin trafficking through the Golgi but had no effect on the mutant opsin inclusion (Fig. 2). Treatment with BFA for periods up to 8 hours did not reduce the incidence of aggregate formation, suggesting that transport of opsin from the ER to Golgi is not required for the formation of these aggregates. Therefore, the mutant protein was not trapped within the Golgi and instead appeared to be associated with a distinct structure located close to the Golgi.
|
Mutant opsin forms ubiquitinated aggresome-like structures
As the mutant opsin appeared to form an intracellular inclusion that had
some of the characteristics of an aggresome
(Kopito, 2000), we
investigated whether they shared other features with aggresomes.
Cotransfection of P23H opsin (Fig.
3) or K296E opsin (data not shown) with c-myc-tagged ubiquitin
resulted in the recruitment of ubiquitin to the opsin inclusions, suggesting
that the structures are ubiquitinated. There was no association of ubiquitin
with the wild-type protein, the ubiquitin retained its diffuse pan-cellular
staining pattern and did not colocalise with the opsin at any stage of its
biogenesis (data not shown). The presence of opsin inclusions also led to the
disruption of the intermediate filament network as shown by vimentin staining
(Fig. 3). Vimentin is
relocalised from its normal distribution to a bright ring surrounding the
inclusion, a feature seen in many aggresome-like structures
(Kopito, 2000
). Examination of
the colocalisation of molecular chaperones with inclusions demonstrated that
the cytoplasmic chaperone Hsc70 (of the Hsp70 family) is recruited to the
inclusions. By contrast, the ER-resident Hsp70, BiP, is predominantly excluded
from the inclusions but does occasionally localise at the periphery of the
inclusion forming a ring around the inclusion (data not shown), a pattern that
was also seen with other ER markers. Collectively, these data suggest that the
mutant opsin inclusions are aggresomes.
|
Ultrastructure of opsin aggresomes
Electron microscopy of COS-7 cells containing mutant opsins confirmed the
localisation of the aggregate close to the microtubule organising centre
(MTOC) and the disruption of the Golgi and intermediate filament networks
(Fig. 4). The juxtanuclear
structures were also surrounded by mitochondria. These mutant opsin
structures, therefore, display most of the ultrastructural hallmarks of
aggresomes. The membrane content of the structures, however, was much higher
than that observed in aggresomes (Heath et
al., 2001; Johnston et al.,
1998
; Waelter et al.,
2001
) formed from other proteins, and in places the structure
appears to be surrounded by membranes (arrowheads in
Fig. 4). This may correspond to
the `ring' of ER staining that was observed in some cells and could reflect
the highly hydrophobic nature of the opsin transmembrane domains acting to
recruit lipid vesicles as well as cellular chaperones.
|
Co-expression of wild-type and mutant opsin results in recruitment of
the normal protein to the mutant opsin aggresome
We investigated whether mutant opsin aggresome formation could influence
the processing of the normal wild-type protein in cell culture. COS-7 cells
were co-transfected with the wild-type protein tagged at its C-terminus with
GFP and untagged wild-type or untagged P23H mutant protein. To make a clearer
distinction between normal trafficking through the Golgi and inclusion
formation, the cells were treated with BFA prior to fixation and analysis of
wild-type opsin-GFP localisation. Co-transfection of normal wild-type
opsin-GFP with an excess of WT untagged opsin did not lead to an increase in
inclusion formation above that seen with the GFP-tagged protein alone. By
contrast, co-transfecting wild-type opsin-GFP with untagged P23H opsin led to
a significant increase in the incidence of wild-type opsin aggresomes
(Fig. 5). In the presence of an
excess of untagged P23H opsin, the incidence of cells with aggresomes
containing the WT opsin-GFP approached the level of aggresome formation seen
with untagged P23H opsin alone (between 35 and 50% at 24 hours). This suggests
that when both wild-type and mutant proteins are expressed, as in ADRP, the
mutant opsin aggresome can recruit the normal wild-type protein.
|
Degradation of mutant opsin is via the ubiquitin-proteasome pathway
and the proteasome is required for efficient retrotranslocation
As aggresome formation is generally enhanced by treatment with proteasome
inhibitors, we examined the effect of the proteasome inhibitor MG-132 on opsin
aggresome formation. In contrast to the expression of F508-CFTR where
proteasome inhibition leads to aggresome formation in 100% of transfected
cells (Johnston et al., 1998
),
treatment with MG-132 for 8 or 14 hours did not have a marked effect on the
incidence of aggresome formation by P23H, K296E or wild-type opsin as assessed
by aggresome counts (data not shown). Morphologically, the most pronounced
effect of proteasome inhibition was to increase the intensity of ER staining
of mutant opsin as demonstrated by colocalisation with BiP
(Fig. 6) and calnexin (data not
shown). In untreated cells (Fig.
1 and Fig. 8,
untreated), the level of mutant opsin staining in the ER was relatively weak
compared with the staining of the aggregates. Following treatment with MG-132,
there was little change in the pattern of wild-type opsin staining but a
dramatic increase in the intensity of ER staining of the mutant opsins was
observed in all cells (Fig.
6).
|
|
The increase in mutant opsin ER staining correlated with an increase in the
level of mutant protein, as judged by western blotting
(Fig. 7). Following MG-132
treatment there was little change in the amount or the composition of the WT
opsin in either the n-Dodecyl-ß-D-Maltoside (DM) soluble or insoluble
fractions. By contrast, the total amount of P23H and K296E opsin is
dramatically increased after incubation with the proteasome inhibitor
(Fig. 7). In the case of the
P23H mutant, this increase was predominantly in the insoluble fraction. DM
solubilisation affords maximal stability for folded opsin but does not simply
produce soluble and aggregated fractions. DM solubilisation probably
fractionates between folded or nearly folded opsin and opsin folding
intermediates, small aggregates and larger aggregates. The major opsin
component in the DM-soluble fraction of untreated cells for the wildtype and
both mutants corresponded to the mature form of the protein, which is Endo H
insensitive. The major opsin component in the DM-insoluble fraction of
untreated cells had the same electrophoretic mobility as PNGase-F-digested
opsin, suggesting that it was deglycosylated. Deglycosylation is a hallmark of
retrotranslocation from the ER. Treatment with MG-132 led to an increase in
the Endo-H-sensitive glycoforms of mutant opsin in the DM soluble and DM
insoluble fractions (data not shown). In addition to the deglycosylated
protein, the insoluble fraction consisted mainly of opsin dimer and high
molecular weight species that migrated as a smear that extended to the top of
the gel, a feature that is characteristic of ubiquitinated proteins
(Ward et al., 1995). This
smear may correspond to the variable number of ubiquitin molecules that are
added to individual opsin molecules during the process of polyubiquitination,
although opsin is prone to self-association and aggregation during SDS-PAGE.
The relatively low level of P23H opsin in the untreated insoluble fraction
most probably reflects the insolubility of the opsin aggresomes in the
DM-insoluble pellets and their subsequent failure to enter the SDS-PAGE gel.
These data suggest that a functioning proteasome is required for the
degradation of the mutant opsin by ER-associated degradation (ERAD) and that
the proteasome also plays a role in the retrotranslocation of the mutant
protein from the ER.
|
Glycosylation is required for mutant opsin ERAD but not quality
control
We examined the effect of tunicamycin, which blocks the synthesis of
precursor oligosaccharides required for N-linked glycosylation, on normal and
mutant opsin processing and quality control in vitro. As previously described
(Kaushal et al., 1994),
tunicamycin did not affect the transport of the wild-type protein to the
plasma membrane and did not lead to the retention of the protein within the
cell; the small amount of intra-cellular staining observed corresponded to
protein trafficking through the Golgi (Fig.
8). The processing of the P23H and K296E mutant proteins, in
contrast, was dramatically altered by tunicamycin. Similar to the situation
with MG-132 treatment, tunicamycin led to an increase in the intensity of ER
staining in all cells compared with the untreated cells, as shown by
colocalisation with BiP (data not shown) and calnexin
(Fig. 8). The incidence of
aggresome formation, however, did not appear to change significantly during
either 8 or 14 hours of incubation with tunicamycin.
As expected, treatment with tunicamycin prevented the formation of the mature diglycoslyated form of opsin and resulted in the accumulation of non-glycosylated opsin as seen by western blotting (Fig. 9). Although treatment with tunicamycin did not have a major effect on the level of the wild-type protein, the steady state levels of the mutant proteins increased considerably, particularly in the insoluble fraction (data not shown). These observations show that the targeting of mutant opsin for ERAD is dependent on N-linked glycosylation. ER retention of mutant opsin, however, is not affected by tunicamycin and reveals a glycan-independent quality control mechanism that prevents mutant opsin escaping the ER.
|
9-cis-retinal promotes the targeting of P23H mutant opsin to the
plasma membrane
Previous studies have suggested that the addition of 11-cis-retinal and
9-cis-retinal to mutant-opsin-expressing cells can improve the folding of
mutant opsins (Li et al.,
1998a). Therefore, we examined the effect of 9-cis-retinal on
mutant opsin processing and aggresome formation in P23H-opsin-expressing
cells. The 9-cis-retinal increased the level of mutant opsin as assessed by
western blotting and in particular the mature form of the protein in the
soluble fraction, suggesting efficient transit through the Golgi apparatus
(Fig. 10). This was confirmed
by immunocytochemical analysis of opsin localisation
(Fig. 10), as an increase in
plasma membrane staining with P23H could be observed. However, the incubation
of 9-cis-retinal did not lead to a significant decrease in the formation of
aggresomes over the period of the treatment time. The addition of
9-cis-retinal to K296E opsin expressing cells had no effect on the processing
of the mutant opsin (data not shown), as would be expected as the mutation has
removed the site of retinal attachment.
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Discussion |
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The correct folding of rhodopsin appears to be exquisitely sensitive to
changes in the primary amino acid sequence, as judged by the large number of
single amino acid substitutions that lead to ADRP (OMIM:
http://www.ncbi.nlm.nih.gov/enterz/dispomim.cgi?id=180380
). At present, the reasons for this sensitivity are not clear even though the
structure of rhodopsin has been solved
(Palczewski et al., 2000), but
may relate to the hydrophobicity of the seven transmembrane domains and the
need for their membrane insertion, coupled to their arrangement in the fully
folded protein. It is also possible that folding of the protein is subjected
to particularly tight quality control in order to prevent the incorporation of
improperly folded protein into the outer segment, as this could lead to an
unacceptable level of `noise' through constitutive activity.
Glycoslyation plays pivotal roles in protein folding, oligomerisation,
quality control, sorting and transport
(Helenius and Aebi, 2001).
Rhodopsin is glycosylated at asparagine residues 2 and 15 in the ER, and the
glycan groups are modified in the Golgi on the way to the outer segment in
photoreceptors or the plasma membrane in COS-7 cells. The importance of
glycosylation in the folding and the processing of opsin has been studied
extensively but has generated apparently conflicting data. Inhibition of
glycoslyation with tunicamycin, which blocks the synthesis of precursor
oligosaccharides required for N-linked glycoslyation, has been used to study
rhodopsin biogenesis in vivo and in vitro. Treatment of isolated Xenopus
laevis and Rana pipiens retinas with tunicamycin disrupts normal
outer segment disc assembly but has no effect on the intracellular transport
of rhodopsin (Fliesler et al.,
1985
; Fliesler and Basinger,
1985
). Similarly, tunicamycin does not prevent the transport of
rhodopsin to the plasma membrane in cell culture
(Kaushal et al., 1994
), and
deglycosylated rhodopsin retains its spectral properties
(Kaushal et al., 1994
;
Prasad et al., 1992
). By
contrast, site-directed mutagenesis of the asparagine residues required for
N-linked glycosylation leads to the accumulation of Drosophila
rhodopsin (Rh1) protein within the ER and retinal degeneration
(Katanosaka et al., 1998
;
O'Tousa, 1992
;
Webel et al., 2000
). However,
such changes in the primary amino acid sequence may also disrupt protein
folding and lead to the protein being retained within the ER because of
misfolding, as seen with mutations in the tri-peptide consensus sequence for
N-linked glycosylation in bovine opsin
(Kaushal et al., 1994
). Our
data show that the major effect of inhibiting N-linked glycosylation in COS-7
cells is to prevent the degradation of the mutant protein and lead to its
retention in the ER, whereas glycosylation does not appear to be required for
the folding and transport of the normal protein. These results are consistent
with the major role of N-linked glycosylation of rhodopsin in the early
secretory pathway being the quality control and degradation of misfolded
protein. The glycan moieties of rhodopsin may be required for the recognition
of misfolded opsin by ER-resident lectin chaperones involved in ERAD, such as
calnexin and calreticulin (Helenius and
Aebi, 2001
), or the recently discovered ER degradation enhancing
-mannosidase-like protein, EDEM
(Hosokawa et al., 2001
). It
appears, however, that the N-linked glycan moieties of rhodopsin are not
required for the non-glycosylated protein to be recognised as misfolded as the
mutant protein is retained within the ER and prevented from exiting to the
Golgi. Therefore, the quality control of opsin folding is not dependent on the
recognition of the misfolded protein by ER-resident lectin chaperones, such as
calnexin and calreticulin, but may involve other chaperones such as BiP and
Grp94, which have previously been shown to bind mutant opsins
(Anukanth and Khorana, 1994
).
The major role of rhodopsin glycosylation in the early secretory pathway
appears to be the facilitation of misfolded protein degradation. Later in the
secretory pathway, however, N-linked glycosylation may play other pivotal
roles in post-Golgi protein sorting, targeting to the rod outer segment, disk
assembly (Fliesler et al.,
1985
; Fliesler and Basinger,
1985
) or even phototransduction
(Kaushal et al., 1994
).
The misfolded opsin is recognised and bound by the ERAD-associated lectins,
then the protein is destined for retrotranslocation and degradation by the
proteasome. Both luminal and transmembrane proteins retained in the ER are now
known to be retrotranslocated into the cytosol in a process that involves ER
chaperones and components of the protein import machinery
(Plemper and Wolf, 1999). Once
exposed to the cytosolic milieu, retrotranslocated proteins are degraded by
the proteasome, in most cases following polyubiquitination. There is growing
evidence that both the ubiquitin-conjugating machinery and proteasomes may be
associated with the cytosolic face of the ER membrane and that they could be
functionally coupled to the process of retrotranslocation
(Hirsch and Ploegh, 2000
). In
the case of CFTR, inhibition of the proteasome does not inhibit
retrotranslocation but instead stimulates aggresome formation by increasing
the amount of undegraded misfolded CFTR in the cytosol
(Johnston et al., 1998
). By
contrast, inhibition of the proteasome appears to increase the amount of
mutant opsin that is retained within the ER, suggesting that the
retrotranslocation of mutant opsin is coupled to the function of the
proteasome, as has been described for several other proteins
(Chillaron and Haas, 2000
;
Mancini et al., 2000
;
Mayer et al., 1998
). Another
difference between mutant opsin and CFTR is the efficiency of degradation of
the misfolded protein.
F508CFTR is degraded very effectively by the
proteasome, and it is only on proteasome inhibition that the mutant protein
aggresomes are readily seen in all cells expressing the mutant protein. Mutant
opsin does not appear to be efficiently degraded by the proteasome and forms
aggresomes spontaneously in the absence of proteasome inhibition. The
incidence of aggresome formation varies between mutations and most probably
reflects the rate that the proteins achieve a native or quasi-native state and
their misfolding rate.
Li et al. demonstrated that the addition of 11-cis-retinal and
9-cis-retinal to mutant-opsin-expressing cells could improve the folding of
T17M mutant opsin but not a mutation in the C-terminus of the opsin
(Li et al., 1998a). Incubation
of P23H-expressing cells with 9-cis-retinal increased the level of the mature
form of the protein in the soluble fraction, suggesting efficient transit
through the Golgi apparatus, and this correlated with an increase in plasma
membrane staining. However, the incubation with 9-cis-retinal overnight did
not lead to a significant decrease in the formation of aggresomes during the
treatment time. These data suggest that, although the ligand can be used to
promote mutant opsin folding and prevent its degradation, once an opsin
aggresome has formed treatment with ligand cannot disaggregate it.
The burden of degrading opsin could weigh heavily on a rod photoreceptor
cell. Normally, the outer segment and the vast quantities of rhodopsin that it
contains are degraded by the retinal pigment epithelium (RPE). Given that
opsin corresponds to at least 30% of the protein produced by a rod cell, even
if only a small percentage of this misfolded and needed to be degraded it
could represent a significant problem for the ER-resident chaperones that
perform the quality control and ERAD, as well as the ubiquitin-proteasome
machinery. This could compromise the normal biosynthetic activity of secretory
pathway in the inner segment and lead to the shortening of outer segments that
has been observed in animal models with the P23H mutation
(Machida et al., 2000). The
rate of mutant opsin misfolding would determine the effect on normal protein
biosynthesis and thereby outer segment length. The presence of mutant protein
also appears to affect the processing of the normal protein
(Colley et al., 1995
;
Wu et al., 1998
), and dominant
mutations in the rhodopsin gene, Nina-E, of Drosophila exert a
dominant-negative effect on the biosynthesis of the normal rhodopsin protein
(Colley et al., 1995
;
Kurada and O'Tousa, 1995
). Our
data show that mutant opsin aggresomes can recruit the normal protein. It may
be that the smaller aggregates of mutant protein that are the precursors of
aggresomes and are destined for degradation also recruit the wild-type
protein, leading to the degradation of the normal protein. In photoreceptors
in vivo, opsin aggresomes may form only rarely as the photoreceptor has a
protein folding machinery that is finely tuned for opsin synthesis, including
specialised chaperones (Chapple et al.,
2001
). Aggregates of electron-dense rhodopsin have been observed
in both human ADRP and animal models of rhodopsin ADRP
(Bunt-Milam et al., 1983
;
Li et al., 1998b
), suggesting
their potential importance to the disease process. The formation of opsin
aggresomes, however, in human and animal models of RP remains to proven. We
propose that aggresome formation could occur as a stochastic event with a
frequency that is inversely proportional to the mutant protein's ability to
fold correctly. This `risk' could in turn be further modified by cellular
stress, which would compromise the cellular chaperone machinery and could
explain the apparent `one-hit' nature of photoreceptor cell death in rhodopsin
RP (Clarke et al., 2001
). The
formation of an aggresome within a photoreceptor could lead rapidly to the
death of the photoreceptor, as opposed to its dysfunction.
The formation of opsin aggresomes may have important consequences for
photoreceptor viability. The presence of protein aggregates has been shown to
compromise the ubiquitin-proteasome machinery
(Bence et al., 2001), and this
could be detrimental to several cellular pathways. Similarly, the disruption
of the cytoskeleton could alter intracellular protein and organelle transport.
The recruitment of cellular chaperones to the protein aggregates could expose
the cells to environmental stress and may stimulate an unfolded protein
response, which could lead to the downstream activation of caspases
(Nakagawa et al., 2000
). We
have shown that mutant opsin aggregates can recruit the normal protein, and it
is possible that the aggregates recruit other proteins. A recent study has
shown that opsin aggregates appear to be specific, and although they
colocalise with other protein aggregates, such as CFTR and TCR
subunits, they do not co-aggregate (Rajan
et al., 2001
). Nevertheless, the recruitment of specific
opsin-binding proteins [e.g. Tctex (Tai et
al., 1999
) and other C-termini-binding sorting factors
(Tam et al., 2000
) or
phototransduction components] to opsin aggresomes could also influence the
targeting of the normal protein to the outer segment and compromise
photoreceptor viability. Cell-culture-based systems will be of value in
determining some of the consequences of opsin misfolding and aggresome
formation and how these events can be manipulated. It is vital, however, to
determine the fate of misfolded opsin in photoreceptors in vivo and determine
the precise cellular consequences for photoreceptors of rhodopsin misfolding
in order to develop novel therapies for ADRP.
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Acknowledgments |
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References |
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