Download ER storage diseases: a role for ERGIC

JCS ePress online publication date 30 May 2006
2532
Research Article
ER storage diseases: a role for ERGIC-53 in
controlling the formation and shape of Russell bodies
Laura Mattioli1, Tiziana Anelli2,3, Claudio Fagioli2, Carlo Tacchetti1, Roberto Sitia2,3,*,‡ and Caterina Valetti1,‡
1
MicroSCoBiO Research Center and IFOM Center of Cell Oncology and Ultrastructure, Department of Experimental Medicine, University of
Genova, Italy
2
DiBiT-San Raffaele Scientific Institute and 3Università Vita-Salute San Raffaele, Via Olgettina 58, 20132 Milano, Italy
*Author for correspondence (e-mail: [email protected])
‡
These authors contributed equally to this work
Journal of Cell Science
Accepted 13 March 2006
Journal of Cell Science 119, 2532-2541 Published by The Company of Biologists 2006
doi:10.1242/jcs.02977
Summary
Owing to the impossibility of reaching the Golgi for
␮ chains
secretion or the cytosol for degradation, mutant Ig-␮
␮⌬CH1) accumulate as
that lack the first constant domain (␮
detergent-insoluble aggregates in dilated endoplasmic
reticulum cisternae, called Russell bodies. The presence of
similar structures hallmarks many ER storage diseases, but
their pathogenic role(s) remain obscure. Exploiting
inducible cellular systems, we show here that Russell bodies
form when the synthesis of ␮⌬CH1 exceeds the degradation
capacity. Condensation occurs in different sub-cellular
locations, depending on the interacting molecules present
in the host cell: if Ig light chains are co-expressed,
␮⌬CH1-light
detergent-insoluble
chain
oligomers
accumulate in large ribosome-coated structures (rough
Key words: ER associated degradation, ERGIC-53, ER storage
diseases, Protein aggregation, Russell bodies
Introduction
Secretory and membrane proteins fold and assemble in the
endoplasmic reticulum (ER), under the assistance of a vast
array of specialized chaperones and enzymes. The ER exerts a
stringent quality control on its protein products. Proteins that
fail to attain their native state are initially retained in the ER
and eventually dispatched to the cytosol for proteasomal
degradation (Ellgaard et al., 1999; Goldberg, 2003; Sitia and
Braakman, 2003). This ER-associated degradation (ERAD)
process is responsible for maintaining homeostasis in the ER
lumen, and implies the recognition of misfolded or orphan
proteins and their retro-translocation or dislocation across
the ER membrane. Trimming of N-glycans is important to
discriminate between unfolded and terminally misfolded
glycoproteins, which explains how newly made proteins are
given sufficient time to attain their three dimensional structure,
a task that can take hours for secretory IgM and other complex
molecules (Tsai et al., 2002; Ellgaard and Helenius, 2003;
Hebert et al., 2005).
Pathology offers us many examples of disorders in the ER
protein factory. Many genetic diseases, e.g. cystic fibrosis, are
due to loss of function because otherwise functional proteins
do not pass ER quality control, and are hence degraded. In
other cases, the problem is the accumulation of mutant proteins
in the ER, which can cause stress and cytotoxicity. An example
of these disorders, collectively known as ER storage diseases
(ERSD), is the hepatopathy caused by the accumulation of
mutant ␣1 anti-trypsin (Liu et al., 1997; Carlson et al., 1989).
Since not all patients harboring a mutant protein develop liver
disease, cellular defense mechanisms must be operating. How
these aggregates exert their toxicity on some cells is unknown.
Russell bodies (RB) are dilated cisternae of the ER
containing large amounts of mutant immunoglobulins (Ig). RB
are frequently detected in multiple myelomas, probably
reflecting the intrinsic tendency of Ig genes to undergo somatic
mutation. Cells with RB are called Mott cells (Weiss et al.,
1984; Alanen et al., 1985). Although they contain massive
protein aggregates in the ER, Mott cells remain viable (Weiss
et al., 1984; Alanen et al., 1985) and undergo mitosis (Valetti
et al., 1991), indicating that, unlike hepatocytes harboring
mutant ␣1 anti-trypsin, myeloma cells can cope with the
presence of RB. We could induce RB formation in lymphoid
and non-lymphoid cells by over-expressing a mutant Ig-␮
heavy chain that lacks the first constant domain (␮⌬CH1)
provided that light chains (L) were also synthesized (Valetti et
al., 1991). RB are induced also by other Ig isotypes, when the
CH1 domain is missing (Kaloff and Haas, 1995). Their
presence correlates with the accumulation of detergentinsoluble Ig, which can neither be transported to the Golgi nor
be dispatched to the cytosol for degradation (Schweitzer et al.,
1991; Valetti et al., 1991).
During Ig assembly, the CH1 domain pairs with the constant
domain of light chains (CL). In the absence of light (L) chains,
the CH1 domain strongly binds the immunoglobulin-binding
protein (BiP/grp78), an ER-resident chaperone of the hsp70
family (Munro and Pelham, 1986). This results in retention of
Russell bodies). In absence of light chains, instead,
aggregation occurs in smooth tubular vesicles and is
controlled by N-glycan-dependent interactions with ERGolgi intermediate compartment 53 (ERGIC-53). In cells
containing smooth Russell bodies, ERGIC-53 co-localizes
with ␮⌬CH1 aggregates in a Ca2+-dependent fashion. Our
findings identify a novel ERGIC-53 substrate, and indicate
that interactions with light chains or ERGIC-53 seed
␮⌬CH1 condensation in different stations of the early
secretory pathway.
Journal of Cell Science
Smooth and rough Russell bodies
heavy (H) chains in the ER, until assembly with L chains
displaces BiP (Hendershot et al., 1987; Hendershot, 1990).
Although formation of H2L2 subunits is sufficient for
secretion of ‘monomeric’ Ig (IgG, IgD and IgE), IgM and
IgA must be further assembled into covalent polymers,
often containing J chains (Hendershot and Sitia, 2004).
Polymerization is developmentally regulated, offering an
additional means to control IgM secretion at the posttranslational level (Sitia et al., 1990). B lymphocytes are unable
to form polymers and as a result they degrade virtually all
secretory IgM and IgA they produce, mostly by the ERAD
pathway (Sitia et al., 1990; Brewer et al., 1994; Guenzi et al.,
1994; Mancini et al., 2000; van Anken et al., 2003). Owing to
the inefficiency of polymerization, some IgM is degraded also
by plasma cells (Sitia et al., 1987; Fra et al., 1993; Fagioli et
al., 2001).
Why is ERAD unable to cope with certain mutant proteins?
A simple explanation would be that their rate of synthesis
exceeds the capacity of degradation. However, not all
transport-incompetent Ig mutants cause RB biogenesis when
degradation is inhibited. Therefore, additional features must be
present for condensation-aggregation in the ER to take place.
One possibility is that certain molecular interactions favor the
formation of small aggregates that seed RB formation. Indeed,
both assembly with L chains and inter-subunit interactions via
cysteine 575 in the ␮s tailpiece, are important for efficient RB
formation (Valetti et al., 1991). Therefore, the RB generated
by mutant IgM somehow reflect abortive attempts to form
secretion-competent polymers. Also the dislocation of ERAD
substrates to the cytosol is likely to play an important role in
RB biogenesis. In order to negotiate transport across the ER
membrane, ERAD substrates must be unassembled and
partially unfolded, with reduction of inter-chain disulfide
bonds preceding dislocation (Fagioli et al., 2001). Soluble
degradation substrates must be captured and concentrated in
the vicinity of the retro-translocation machinery. Mechanisms
must therefore be operating that couple disassembly and partial
unfolding to the insertion of the substrate into the dislocon
channels. The partially unfolded molecules generated in the
process are at high risk for aggregating (Kopito and Sitia,
2000). In addition, the strong tendency of H chains lacking the
CH1 domain to form insoluble aggregates in the ER can be
related to the fact that they bind less BiP than wild-type
molecules, thus enjoying less chaperoning.
In view of the biological and clinical interest of RB, we have
investigated their biogenesis in lymphoid and non-lymphoid
cells. Our results show that accumulation-aggregation ensues
when the production of ␮⌬CH1 exceeds the capacity of the
ERAD system. In addition, we present evidence indicating that
condensation-aggregation can take place in two different subregions of the early secretory pathway, depending on the
interacting molecules present in the host cell. When L chains
are present, aggregation takes place in the rough ER. In their
absence, instead, ␮⌬CH1 aggregate in smooth tubular vesicles
in association with ERGIC-53, a marker of the ER-Golgi
intermediate compartment (ERGIC) (Schweizer et al., 1991).
Results
Induction of RB-like structures in the absence of L
chains
We have previously described a stable myeloma transfectant
2533
that expresses a mutant ␮ chain lacking the first constant
domain (␮⌬CH1), but fails to form RB (N[␮⌬CH1]).
Expression of L chains readily induced the appearance of RB,
suggesting that assembly with L chains is important for RB
formation, possibly by cross-linking polymers in the ER
through their unpaired CL domains [see Valetti et al. (Valetti et
al., 1991) and below]. Another possibility is that assembly with
L chains stabilizes mutant H chains (Sitia et al., 1987; Fagioli
et al., 2001), thus slowing down ERAD. In this way, ␮⌬CH1
could reach a threshold concentration in the ER, or within a
specialized sub-compartment(s) thereof, sufficient to seed
condensation.
To determine whether RB could be induced in the absence
of L chains, we modulated the ratio between synthesis and
degradation. First, we increased the rate of ␮⌬CH1 production
by exposing cells to sodium butyrate. At the concentration of
1 mM, sodium butyrate approximately doubled transgene
expression without significantly inducing apoptosis, as assayed
by metabolic pulse assays and annexin V-propidium iodide
staining (our unpublished results). As shown in Fig. 1A,
untreated N[␮⌬CH1] and N[␮1] cells (the latter expressing the
wild-type form of the secretory ␮ chains, ␮s) display a diffuse
staining of the ER (Valetti et al., 1991). These clones have
presumably reached an equilibrium between synthesis and
degradation of the orphan ␮ chains. After 40 hours of sodium
butyrate treatment, 30-50% of N[␮⌬CH1] cells developed RBlike structures, of varying in size and number. In contrast,
N[␮1] did not form RB despite increased ␮ synthesis as
identified by a brighter immunofluorescent staining pattern
(Fig. 1A) and in western blots (Fig. 1B). ␮s and ␮⌬CH1
differed also with respect to their detergent solubility (Fig. 1B).
Whereas virtually all wild-type ␮s chains were detergent
soluble (s), half or more ␮⌬CH1 accumulated in the insoluble
fraction (i).
Therefore, RB can form also without L chains. However,
morphological differences were evident in myeloma
transfectants depending on the presence or absence of L
chains. Whereas L chain-containing RB tended to be fewer in
number and more circular (see J[␮⌬CH1] cells, Fig. 1A), a
constant feature in N[␮⌬CH1] cells was that the structures
containing mutant ␮ chains were smaller and less regular,
often polygonal (Fig. 1A, NaBut and Fig. 1C). Their size
ranged from 0.2 to 0.6 ␮m, with a mean value of 0.4 ␮m (Fig.
1E), hence much smaller that L chain containing RB in
J[␮⌬CH1] (average 1.3 ␮m). Cryo-EM analyses with anti-␮
(Fig. 1D) confirmed that in both cases the structures contained
abundant ␮⌬CH1 chains.
Besides confirming the size and shape differences observed
by immunofluorescent staining, resin embedding electron
microscopy (EM) studies (Fig. 1C) revealed that in cells
expressing L chains, RB displayed ribosomes all around their
surface. In contrast, the structures present in N[␮⌬CH1] cells
had only occasional ribosomes, often concentrated in discrete
areas. We will refer to these latter structures as smooth RB
(sRB).
Inhibiting degradation favors smooth RB formation
To test whether a decrease in degradation could also enhance
RB formation, we utilized proteasome inhibitors. First, we
confirmed that ␮⌬CH1 chains are degraded by proteasomes,
by chasing pulse-labeled cells with or without MG132 (data
Journal of Cell Science
2534
Journal of Cell Science 119 (12)
Fig. 1. RB-like structures form in the absence of L
chains when ␮⌬CH1 synthesis is increased.
(A) Induction of RB in myeloma transfectants.
N[␮⌬CH1], N[␮1] and J[␮⌬CH1] cells were treated
with or without 1 mM sodium butyrate (NaBut) for 40
hours, fixed and stained with fluorescent anti-␮
antibodies. Note that RB-like structures are induced by
NaBut treatment in N[␮⌬CH1] but not in N[␮1]. As
previously reported (Valetti et al., 1991), RB are present
also in untreated, ␭ producing J[␮⌬CH1] cells; bar, 5
␮m. (B) The presence of RB correlates with
accumulation of detergent-insoluble ␮⌬CH1 chains.
Cells were lysed in TX100 and equal amounts of soluble
(s) and insoluble (i) fractions resolved under reducing
conditions, blotted and decorated with anti-␮.
(C,D) Different RB morphology in myeloma cells is
dependent on L chain synthesis. J[␮⌬CH1] and
N[␮⌬CH1] cells, the latter treated with 1 mM NaBut for
40 hours, were processed for both Epon embedding (C)
and cryo-sectioning (D). Ultrathin cryo-sections were
immuno-gold labeled using anti-␮ antibodies and
protein A-gold (15 nm). Arrowheads point to
membrane-associated ribosomes. Bar, 100 nm. (E) The
table summarizes the RB diameters (expressed in ␮m0
in N[␮⌬CH1] cells treated with 1 mM NaBut for 40 hours, and in untreated J[␮⌬CH1] cells. Measurements were carried out on electron
microscope images. 60 and 30 RB were measured in N[␮⌬CH1] and J[␮⌬CH1] cells respectively, each from two independent preparations.
not shown) as described for ␮s (Mancini et al., 2000; Fagioli
et al., 2001). In the course of these experiments, we observed
that myeloma cells are extremely sensitive to proteasome
inhibitors, the vast majority of them undergoing apoptosis after
6 hours of treatment or less (Cenci et al., 2006). To circumvent
this problem, we combined treatments. As shown in Fig. 2,
adding MG132 for the last 2 hours of a 24-hour treatment with
sodium butyrate, doubled the number of cells containing sRB
in N[␮⌬CH1] cells, indicating that aggregation-prone ␮⌬CH1
chains condense in the ER in a detergent insoluble state when
their synthesis exceeds disposal.
Induction of RB in non-lymphoid cells
In view of the possibility that sodium butyrate exerted
additional effects, we generated inducible HeLa cells
(H[␮⌬CH1]) in which ␮⌬CH1 synthesis is driven by a Tet-Off
promoter (Gossen and Bujard, 1992). These cells formed RBlike structures within a few days after removal of tetracycline
(Fig. 3A), in correlation with the accumulation of detergentinsoluble ␮⌬CH1 chains (Fig. 3B). In the first 2 days of
induction, when few ␮⌬CH1 are detectable in the Triton X100 (TX100) soluble fraction (Fig. 3B), a weak reticular and
peri-Golgi staining was evident by immunofluorescence
microscopy. At day 4, small, bright dots became evident in 2040% of ␮⌬CH1 producing cells (see inset). These dots, usually
one in each cell, probably represent the first morphological
sign of ␮⌬CH1 aggregation. With time, they increased
progressively in size, and this correlated with an increase in the
insoluble pellet (Fig. 3B). These structures, present in 50-70%
of cells after day 5, were irregular in shape (see also below)
and were generally positioned close to the microtubuleorganizing center (MTOC) area, as shown by their proximity
to ␥-tubulin (see Fig. 3C).
Dose-response experiments confirmed that a critical ␮⌬CH1
concentration must be reached for triggering condensationaggregation in the ER (summarized in Fig. 3D). At
concentrations of tetracycline of 0.25 ng/ml, when synthesis of
the transgene was minimal, the majority of ␮⌬CH1 remained
in the soluble fraction. At 0.12 ng/ml, insoluble ␮⌬CH1 chains
became preponderant, and increased progressively at lower
tetracycline concentrations. The soluble fraction remained
essentially constant, possibly reflecting the chaperoning power
of the cell.
Fig. 2. Pharmacological inhibition of proteasomal degradation favors
sRB formation. N[␮⌬CH1] cells were cultured for 24 hours with or
without 1 mM sodium butyrate (NaBut). Aliquots of cells treated
with NaBut were exposed to MG132 for the last 2 hours in culture.
The histogram depicts the percentage of cells containing ␮⌬CH1
sRB structures and represents the average of two independent
experiments.
Journal of Cell Science
Smooth and rough Russell bodies
2535
Fig. 3. Induction of sRB in non-lymphoid
cells. (A) Time-dependent formation of
sRB in H[␮⌬CH1] induced to synthesize
␮⌬CH1 chains. After culture in the
absence of tetracycline for the indicated
periods, cells were harvested, fixed and
stained with fluorescent anti-␮; bar 5 ␮m.
The inset in the center panel (day 4)
shows a magnification of a cell containing
a small ␮-positive dot. (B) sRB formation
correlates with accumulation of
detergent-insoluble ␮⌬CH1 chains in
H[␮⌬CH1]. Aliquots of detergent soluble
(s) and insoluble (i) material from cells
treated as in A were analyzed by western
blotting as described in legend to Fig. 1.
(C) sRB localize preferentially in the
vicinity of the MTOC. H[␮⌬CH1] cells
cultured for 5 days without tetracycline
were simultaneously stained with anti-␥tubulin and anti-␮ antibodies; bar 5 ␮m.
(D) Dose-dependent formation of sRBs
imply a threshold concentration for
␮⌬CH1 condensation. H[␮⌬CH1] cells
were cultured for 5 days in medium
supplemented with decreasing doses of
tetracycline as indicated. ␮⌬CH1 chains
were quantified in extracts containing total (TOT, lysis in SDS) or TX100 soluble (SOL) and insoluble (INS) proteins and expressed as arbitrary
units. At the tetracycline concentration of 0.25 ng/ml or more, most ␮⌬CH1 proteins are found in the soluble fraction. At 0.12 ng/ml detergentinsoluble chains become preponderant and continue to increase at lower concentrations, when more ␮⌬CH1 chains are synthesized.
Distinct sites of accumulation in the presence or
absence of L chain in the inducible H[␮⌬CH1] clone
The co-expression of mutant ␮ and ␭ chains dramatically
influenced the shape of RB structures in non-lymphoid
cells also (Fig. 4). Instead of the irregularly shaped
structures concentrated around the MTOC area (Fig. 4a),
numerous spherical vacuoles, rather regular in shape and
dispersed throughout the cell, accumulated in ␭ producing
cells (Fig. 4b). These structures were stained by Ac38
anti-idiotypic antibodies (Reth et al., 1979; Valetti et al.,
1991), confirming proper pairing between the ␮ and ␭
variable (V) domains (Fig. 4b). As in myeloma
transfectants (compare Fig. 1, J[␮⌬CH1] cells), these
structures contained an electron dense material
surrounded by ribosome-coated membranes (Fig. 4h). In
contrast, smaller vacuoles and tubules, generally devoid
of ribosomes, accumulated in cells expressing ␮⌬CH1
chains alone (Fig. 4g).
Consistent with the presence of ribosomes on their
membranes, rough RB (rRB) containing ␭ chains were
stained by anti-PDI (data not shown), but lacked ERGIC53 (Fig. 4e). In contrast, sRB containing ␮⌬CH1 alone
were intensely stained by anti-ERGIC-53 antibodies
(Fig. 4d), suggesting that in the absence of L chains,
condensation occurs in distal regions of the early
secretory compartments, including ERGIC.
How could the presence of ␭ chains induce
condensation in rRB? In the absence of CH1, the constant
domain of L chains, CL, does not find its natural
counterpart. Like most Ig domains, CL tends to form
dimers (Leitzgen et al., 1997). This could favor aggregate
Fig. 4. The presence or absence of L chains dictates the site of RB
accumulation. HeLa Tet-Off cells were transiently transfected with
␮⌬CH1 expression vectors alone (a,d,g) or in combination with wildtype ␭ (L; b,e,h) or with a mutant ␭ lacking the constant domain (VL;
c,f). ␮⌬CH1 appears red, whereas Ac38 and ERGIC-53 are green; bar 5
␮m. g and h are imagines of resin-embedded EM sections. In cells
expressing L chains, numerous ribosomes are uniformly distributed on
the RB membrane. Electron density of RB content demonstrates high
protein density. Bars, 200 nm.
2536
Journal of Cell Science 119 (12)
Journal of Cell Science
formation, by linking different (␮⌬CH1-␭)n polymers in a
three-dimensional mesh (Valetti et al., 1991). To verify this
model, we deleted the CL domain from murine ␭ chains. These
truncated variable domains (VL) assembled with ␮⌬CH1
chains, as demonstrated by Ac38 staining (Fig. 4c), but did not
alter the distribution of ␮⌬CH1, which accumulated within
ERGIC-53-positive sRB (Fig. 4f). Therefore, the unpaired CL
domains seem to play an important role in favoring rRB
formation.
ERGIC-53 is enriched in sRB, in the absence of L
chains
After 7 days in culture without tetracycline, the majority of the
ERGIC-53 staining was concentrated in the region occupied by
␮⌬CH1 (Fig. 5e). This clearly contrasted with the distribution
observed in cells cultured in the presence of tetracycline (Fig.
5a). At day 3 of induction, an intermediate phenotype was
observed, indicating that ERGIC-53 was recruited in the
␮⌬CH1-containing structures (Fig. 5c). The distribution of
GM-130 was not altered, although sRB were generally
adjacent to the Golgi complex (Fig. 5b,d,f).
We further characterized sRB by EM. In the absence of L
chains, ␮⌬CH1 accumulated in tubular structures, somehow
reminiscent of a dish of spaghetti or a ball of yarn (Fig. 5g-i).
The interlaced tubules were generally devoid of ribosomes
(Fig. 5i-j) and intensely stained by anti-␮ and anti-ERGIC-53
antibodies (Fig. 5g and h, respectively). Many examples of
continuity between these tubules and normal ER cisternae were
present (see Fig. 5j).
Altogether, these results indicated that ␮⌬CH1 can
accumulate in different compartments of the early exocytic
route. When L chains are present, mutant proteins condense in
the rough ER, whereas in their absence aggregation occurs in
tubular vesicles enriched in ERGIC-53. L chain expression
superimposes a more regular shape to rRB. An important role
is played by the unpaired CL domain, which could trigger
condensation by linking ␮⌬CH1 polymers or perhaps favoring
polymerization in the ER (see Fig. 4).
ERGIC-53 is important for sRB biogenesis and binds
␮⌬CH1 in a Ca2+-dependent manner
To determine whether the co-localization of ␮⌬CH1 and
ERGIC-53 reflected specific interactions between the two
molecules, we exploited the finding that glycoprotein binding
to the lectin requires Ca2+ (Itin et al., 1996; AppenzellerHerzog et al., 2004). Thus, H[␮⌬CH1] cells were incubated
with thapsigargin or cyclopiazonic acid (CPA), two drugs
known to induce Ca2+ release from the ER in an irreversible or
reversible manner, respectively (Doutheil et al., 1997). When
Ca2+ levels were lowered from intracellular stores with CPA
(Fig. 6A, top right panel), or thapsigargin (not shown), ␮⌬CH1
remained confined to sRB, whereas ERGIC-53 regained its
normal dispersed distribution (Fig. 6A, top right). ERGIC-53
re-localized into sRB upon removal of CPA and culture in
Fig. 5. In the absence of L chains, ␮⌬CH1 condense primarily in tubular vesicles and saccules enriched in ERGIC-53. (a-f) H[␮⌬CH1] cells
were cultured with (a,b) or without tetracycline for 3 (c,d) or 7 days (e,f) and stained with antibodies against ERGIC-53 (a,c,e in green),
GM130 (b,d,f; green) or ␮ (c-f; red). Note that ERGIC-53 abandons its canonical distribution (a) to co-localize with ␮⌬CH1-containing sRB
upon tetracycline removal (panels c,e yellow staining). ␮⌬CH1 accumulates close to GM130, but does not overlap with it (d,f); bar, 5 ␮m.
(g,h) After 7 days of culture without tetracycline, cells were processed for cryo-EM with anti-␮ (g) or anti-ERGIC-53 (h, G1/93) antibodies and
revealed with protein-A coupled to 15 nm gold particles. ␮⌬CH1 chains accumulate in saccules enriched in ERGIC-53. Bars, 100 nm.
(i,j) H[␮⌬CH1] cells were kept for 7 days without tetracycline and processed for resin embedding and EM. Continuity between rough ER and
␮⌬CH1-containing sRB are evident in j (see arrow; the arrowheads point to regions rich in ribosomes). Bars, 100 nm.
Journal of Cell Science
Smooth and rough Russell bodies
2537
Fig. 6. ␮⌬CH1 binds to ERGIC-53 in a Ca2+-dependent
manner. (A) Ca2+-dependent co-localization of ERGIC53 and ␮⌬CH1. After 5 days of culture without
tetracycline, H[␮⌬CH1] cells were treated for 2 hours
with or without 50 ␮M CPA (top right and left panels,
respectively), a drug that reversibly depletes
intracellular Ca2+ stores. CPA was then washed out and
cells were cultured in Ca2+-containing medium for 15
and 45 minutes (bottom left and right panels,
respectively) before fixation and co-staining with anti-␮
(red) and anti-ERGIC-53 (green); bar 5 ␮m.
(B) ERGIC-53 is recruited to the insoluble fraction in
cells expressing ␮⌬CH1. Detergent-soluble (s) and
insoluble (i) fractions were obtained from cells treated
with or without tetracycline as indicated. In order to
preserve Ca2+-dependent interactions, EDTA was
omitted from lysis buffer, and 1 mM CaCl2 added (lanes
3-6). An aliquot of cells was lysed in the presence of
EDTA (lanes 1-2). Aliquots were resolved under
reducing conditions, and blotted. The filter was
sequentially decorated with anti-ERGIC-53 and anti-␮.
(C) Ca2+-dependent co-immunoprecipitation of ERGIC53 and ␮⌬CH1. H[␮⌬CH1] cells were cultured without
tetracycline for 2 days and then transfected with
ERGIC-53 wt, the KKAA or the lectin-deficient mutant
N156A. After a further 40 hours in culture without
tetracycline, cells were treated with 50 ␮M CPA for 2
hours to deplete intracellular Ca2+ stores (lanes 3-4), and cross-linked with DSP. Equal aliquots of the lysates were immunoprecipitated with
anti-␮ or anti-ERGIC-53, as indicated. Immunoprecipitates were resolved by SDS-PAGE, transferred to nitrocellulose and probed with antiMyc, in order to detect co-precipitated exogenous ERGIC-53. (D) The amount of ERGIC-53 co-immunoprecipitated with ␮⌬CH1 was
quantified and expressed as the percentage relative to total immunoprecipitable material (IP with anti-ERGIC-53). Average of two independent
experiments.
Ca2+-containing medium for 15 and 45 minutes (Fig. 6A,
bottom left and right). These findings suggest that sRB do not
represent a dead end station, but are still connected with
vesicular traffic.
Further evidence for a Ca2+-dependent interaction between
␮⌬CH1 and ERGIC-53 was obtained by biochemical studies.
In non-induced H[␮⌬CH1] cells, virtually all ERGIC-53 was
detergent-soluble (Fig. 6B, lanes 5 and 6). After tetracycline
removal, when cells accumulated abundant detergentinsoluble ␮⌬CH1 chains, a considerable fraction of ERGIC53 was present in the insoluble fraction (lanes 3 and 4).
However, if cells were lysed in the absence of Ca2+, ERGIC53 was found exclusively in the soluble fraction (lanes 1 and
2). Therefore, the sub-cellular localization of ERGIC-53 is
altered by the presence of ␮⌬CH1 aggregates in a Ca2+dependent fashion.
To further investigate the interactions between ␮⌬CH1
and ERGIC-53 we performed cross-linking and coimmunoprecipitation studies (Fig. 6C,D). Anti-␮ antibodies
co-precipitated a considerable fraction of ERGIC-53 wt and the
localization-defective KKAA mutant (Fig. 6C, lane 1) but only
traces of the binding-deficient N156A mutant. The interaction
was clearly inhibited when intracellular Ca2+ stores were
depleted (Fig. 6C, compare lanes 1 and 3). Treatment with CPA
significantly reduced the amount of ERGIC-53 that could be
precipitated with anti-␮ antibodies in cells expressing wt or
KKAA molecules, but had no effect on the binding-deficient
N156A mutant (Fig. 6D), confirming that ERGIC-53 binds
␮⌬CH1 chains in a Ca2+-dependent way via its lectin domain
(Appenzeller et al., 1999). Despite our efforts, we failed to co-
immunoprecipitate endogenous ERGIC-53 with anti-␮
antibodies. This may reflect technical limitations, and also the
transient nature of the interactions between ERGIC-53 and
substrates. Over-expressed ERGIC-53 molecules could be
more easily co-immunoprecipitated because of their
localization in the ER (L.M. and C.V., unpublished results),
where cargo capture prevails. Nonetheless, the Ca2+-dependent
delocalization assays shown in Fig. 6A, are consistent with an
interaction of ␮⌬CH1 with endogenous ERGIC-53.
Mannose trimming of ␮⌬CH1 is required for sRB
biogenesis
The above data established that in the absence of L chains
␮⌬CH1 chains condense in sRB in association with ERGIC53. If interactions with this lectin molecule played an active
role in condensation, perturbing N-glycan processing could
have an effect on sRB formation in cells lacking L chains. In
view of the role of mannose residues in targeting misfolded
proteins to degradation (Helenius and Aebi, 2004) these
experiments could also shed light on the relationships between
ERAD and RB formation. We have previously demonstrated
that the mannosidase I inhibitor kifunensine prevents
degradation/dislocation of orphan wild-type ␮s (Fagioli and
Sitia, 2001) and ␮⌬CH1 chains (our unpublished data), as
efficiently as proteasome inhibitors. Therefore, if the
concentration of an aggregation prone molecule were sufficient
to determine condensation, kifunensine should favor the
biogenesis of RB in both myeloma and HeLa transfectants, by
causing accumulation of ␮⌬CH1 chains in the ER. If in
contrast, glycan-dependent interactions with intracellular
Journal of Cell Science
2538
Journal of Cell Science 119 (12)
Fig. 7. Mannose trimming is required for
␮⌬CH1 condensation and efficient sRB
formation. (A,B) Kifunensine causes
accumulation of ␮⌬CH1 but no sRB formation
in myeloma transfectants. N[␮⌬CH1] were
treated for 40 hours with sodium butyrate
(NaBut, 1 mM) or with kifunensine (Kif, 2
␮g/ml), to increase the expression or prevent the
degradation of ␮⌬CH1, respectively. Cells were
then fixed and processed for
immunofluorescence microscopy with anti-␮ (A,
bar 5 ␮m) or lysed in TX100. Equal amounts of
soluble (s) and insoluble (i) lysate fractions were
analyzed by western blotting (B). (C,D)
Kifunensine causes accumulation of detergentsoluble ␮⌬CH1 in the ER of HeLa cells. Cells
were cultured for 5 days without tetracycline,
with or without 2 ␮g/ml kifunensine, and
processed for immunofluorescence microscopy
(C, bar 5 ␮m) or western blotting (D).
Kifunensine was re-added every 24 hours
(Tokunaga et al., 2000). In both lymphoid and
non-lymphoid cells, kifunensine caused
accumulation of abundant ␮⌬CH1, which was
mainly in the soluble fraction when mannose
trimming was prevented. As expected, ␮⌬CH1
chains present in kifunensine-treated cells
display slower electrophoretic mobility. (E,F) The association between ERGIC-53 and ␮⌬CH1 is inhibited by kifunensine. H[␮⌬CH1] cells
were cultured in the absence of tetracycline for 2 days, with or without kifunensine (Kif, 2 ␮g/ml, lanes 5-6) and then transfected with ERGIC53 wt. After further 40 hours of culture, cells were treated with 50 ␮M CPA for 2 hours (lanes 3-4) before co-immunoprecipitation and
quantification, as described in the legend to Fig. 6C. Kifunensine was present for the entire duration of the experiment.
lectins were important for condensation, mannosidase
inhibitors could prevent sRB formation.
In a first series of experiments, N[␮⌬CH1] cells were treated
with either sodium butyrate or kifunensine for 40 hours. As
expected, both treatments considerably increased the amount
of ␮⌬CH1 found in the cells lysates (Fig. 7A,B lanes 3-6).
However, neither the formation of sRB structures (Fig. 7A) nor
the accumulation of detergent-insoluble molecules (Fig. 7B
lanes 5 and 6) was induced by kifunensine.
Also in H[␮⌬CH1] cells, kifunensine prevented sRB
formation (Fig. 7C). After 5 days of tetracycline removal, less
than 10% of the cells in which mannosidase I was inhibited
with kifunensine contained sRB, compared with over 60% in
controls. Most cells displayed a diffuse, ER staining pattern
when decorated with anti-␮ (Fig. 7C). In both kifunensinetreated myeloma and HeLa cells ␮⌬CH1 chains displayed
slower mobility because of their higher mannose content and
accumulated largely in the detergent-soluble fraction (panels B
and D).
Inhibition of mannose trimming also inhibits the
interactions between ␮⌬CH1 and ERGIC-53 (Fig. 7E,F).
When cells were treated with kifunensine the quantity of
ERGIC-53 co-precipitated with anti-␮ antibodies was
significantly reduced (Fig. 7E, lane 5) to a level similar to that
obtained with CPA (see lane 3) as quantified in Fig. 7F and
Fig. 6D.
Kifunensine had minor if any effects on rRB formation in
cells synthesizing L chains (data not shown), indicating that
mannose removal was not essential for detergent insolubility
per se. Altogether, these data suggest a crucial role for mannose
processing in determining the fate of aggregation-prone
␮⌬CH1 chains, and imply that, in the absence of L chains, a
key factor in this process could be ERGIC-53.
Discussion
ER storage disorders (ERSD) are a recently defined class of
diseases characterized by the accumulation of aberrant
proteins in the ER. They are ‘conformational diseases’
(Kopito and Ron, 2000), a definition that nicely pinpoints the
pathogenic role of proteins that owing to mutations or lack of
cofactors assume aberrant conformations. These are
recognized by the ER quality control system, and cannot
proceed to the Golgi. If these transport-incompetent molecules
are not efficiently routed to the cytosol for proteasomal
degradation (dislocated), accumulation in the ER lumen
inevitably ensues. Not all proteins that fail to reach the Golgi,
however, accumulate in the ER. Why do certain proteins, such
as for instance, PiZ␣1AT or ␮⌬CH1, fail to dislocate to the
cytosol? ERAD could be inefficient in their capture,
unfolding, dislocon insertion, and/or delivery to the cytosol,
but it seems clear that these proteins have an intrinsic tendency
to form oligomers, probably too big to negotiate export
(Carrell and Lomas, 1997).
Our results reveal important facets of the mechanisms that
determine aggregation of ␮⌬CH1 chains and hence RB
formation. First of all, by manipulating the rate of synthesis
and degradation of the condensation-prone ␮⌬CH1 chains, we
could demonstrate that RB formation ensues when the
synthesis exceeds the degradative capacity. In both lymphoid
and non-lymphoid cells, detergent-insoluble ␮⌬CH1 chains
accumulate when synthesis is increased (Figs 1, 3) or
degradation inhibited (Fig. 2). It seems therefore that cells can
Journal of Cell Science
Smooth and rough Russell bodies
cope with limited amounts of mutant ␮ chains. This is evident
in HeLa cells, where intracellular detergent-soluble ␮⌬CH1
reach a plateau (Fig. 3B,D), only insoluble chains increasing
with time. Since CH1 domains of all Ig classes bind BiP with
high affinity, formation of detergent-insoluble aggregates by
mutants that lack them could reflect reduced interactions with
the main ER chaperone.
The threshold for aggregation varies for different cell types
and also depends on interacting molecules. For example,
the fraction of insoluble ␮⌬CH1 is much higher in cells
synthesizing L chains. The latter have notable consequences
also on the shape and location of RB. In their absence,
␮⌬CH1 condensation occurs in irregular masses of tubular
vesicles and vacuoles surrounded by smooth membranes.
These sRB are enriched in ERGIC-53, a lectin involved in
intracellular glycoprotein transport (Itin et al., 1996; BenTekaya et al., 2005). In contrast, in the presence of L chains,
RB adopt circular and regular shape, contain PDI and are
surrounded by a membrane rich in ribosomes (Figs 1, 4),
suggesting that ␮⌬CH1 condensation occurs near the site of
synthesis.
How do L chains favor rRB formation? Two non-mutually
exclusive mechanisms can be envisaged. First, the association
with L could slow down ␮⌬CH1 degradation, imposing a
further requirement for unfolding-dissociation before
dislocation (Fagioli et al., 2001). This would increase
concentration in the ER lumen, indirectly promoting
condensation-aggregation. Second, because of the tendency of
most Ig constant domains to form dimers, the unpaired CL
associated to ␮⌬CH1 via VL-VH interactions could link
different polymers, thus promoting condensation (Valetti et al.,
1991). In favor of the latter model is the observation that L
chains lacking the CL do not favor rRB formation. Despite
proper VH-VL pairing, demonstrated by reactivity with antiidiotypic antibodies, condensation-aggregation occurred in
ERGIC-53-positive sRB, as in the absence of L chains.
The role of N-glycan processing in RB formation
The data presented in Figs 1-3 are consistent with a simple
model, in which RB form when the production of a transportincompetent molecule exceeds the degradation capacity.
However, the results obtained with kifunensine, an inhibitor
of ER mannosidase I that blocks ␮ chain degradation (Fagioli
and Sitia, 2001) imply that a mere increase in concentration
is not sufficient to cause RB formation. Despite kifunensinetreated NSO and HeLa transfectants accumulating abundant
␮⌬CH1, neither formed sRB (Fig. 7). By contrast,
kifunensine did not interfere with RB formation in cells
expressing L chains, suggesting an important role for
mannose trimming in triggering ␮⌬CH1 condensation in
sRB.
Interactions between ␮⌬CH1 and ERGIC-53
ERGIC-53 is a Ca2+-dependent lectin thought to concentrate
selected secretory glycoproteins in forward transport vesicles
budding from the ER (Itin et al., 1996; Appenzeller-Herzog
et al., 2004; Ben-Tekaya et al., 2005). At steady state,
ERGIC-53 is enriched in the ERGIC, owing to the presence
of a KKFF motif in its cytosolic tail. Our results identify
␮⌬CH1 as a novel substrate of ERGIC-53. They also indicate
that interactions between ␮⌬CH1 and ERGIC-53 play an
2539
important role in sRB biogenesis in the absence of L chains.
ERGIC-53 looses its normal distribution and re-localizes in
sRB, in association with ␮⌬CH1. Part of ERGIC-53 is found
in the detergent-insoluble fraction and can be cross-linked to
␮⌬CH1 in a Ca2+ dependent manner. Ca2+ was necessary for
association, and the N156A mutant was inactive, implying
specific interactions through the lectin domain. The
observation that kifunensine inhibited binding of ␮⌬CH1
chains to ERGIC-53 was somehow unexpected, since another
inhibitor of ER mannosidases (deoxymannojiirimycin) did
not prevent binding of cathepsin Z (Appenzeller et al., 1999).
Persistence of the terminal mannose may favor binding of
␮⌬CH1 to calnexin and calreticulin (Mancini et al., 2000)
(L.M., C.V. and R.S., unpublished results), thus preventing
interactions with ERGIC-53. Another intriguing possibility is
that subtle structural features that are dependent on the
interactions between N-glycans and the surrounding
polypeptide backbone determine the specificity of binding
(Appenzeller-Herzog, 2005; Cals et al., 1996; de Lalla et al.,
1998).
In the absence of L chains, therefore, ␮⌬CH1 chains
condense in tubular vesicles, in association with ERGIC-53.
This network is largely devoid of ribosomes and contains a
dense material intensely stained by anti-␮ antibodies. Some of
these tubules are clearly continuous with ER cisternae.
Condensation of ␮⌬CH1 could take place at the junctions,
which often appear as if constricted by a ring. These tubular
vesicles could correspond to ER exit sites, in which ERGIC53 and other molecular complexes concentrate cargo for Golgi
transport. It is also possible that junctions or sRB correspond
to sites of the early secretory pathway specialized in
dislocating ERAD substrates across the ER membrane for
cytosolic delivery (Kamhi-Nesher et al., 2001).
Protein-protein interactions dictate the shape and
localization of RB
Our results are consistent with an active role of ERGIC-53 in
seeding ␮⌬CH1 aggregation in a sugar- and lectin-dependent
way. Replacing Cys575 or treating cells with 2 mercaptoethanol inhibits RB formation (Valetti et al., 1991), indicating
that formation of disulfide bonds linking different [␮⌬CH1]2
dimers is essential for efficient condensation. As discussed
above, L chains could mediate the association of polymers, or
favor their formation in the ER (Bornemann et al., 1995; Cals
et al., 1996). Likewise, ERGIC-53 could seed condensation,
perhaps exploiting its property to form hexamers. The results
presented in Fig. 6 indicate that once formed, detergentinsoluble complexes do not require the continuous interaction
with ERGIC-53. They also imply that these aggregates are not
sequestered but remain in dynamic contact with the exocytic
pathway, since ERGIC-53 can re-localize in sRB upon readdition of Ca2+.
In conclusion, our findings reveal that protein aggregation in
the ER depends on several cooperating events: abundant
synthesis and reduced degradation of proteins that have an
intrinsic tendency to form aggregates because of reduced
chaperone binding and interactivity with ancillary proteins
that favor their concentration and seed condensation. The
equilibrium between synthesis, processing, and degradation of
aberrant species is subject to delicate and continuous
adjustments, operated by the ER quality control machinery.
2540
Journal of Cell Science 119 (12)
Materials and Methods
Reagents
Anti-␥-tubulin
monoclonal
antibodies,
cyclopiazonic
acid,
dithiobis(succinimidylpropionate, geneticin, N-ethylmaleimide, sodium butyrate,
tetracycline, thapsigargin and Triton X-100, were from Sigma-Aldrich;
dithiobis(succinimidylpropionate) (DSP) was from Pierce, hygromycin B from
Invitrogen and kifunensine from Toronto Research Chemicals Inc.
Polyclonal anti ERGIC-53 (Spatuzza et al., 2004) was a kind gift from Stefano
Bonatti (University of Naples, Italy). Polyclonal anti-mouse ␮ was purchased from
Zymed; HRP anti-␭, HRP anti-mouse IgG, TRITC anti-␮ were from Southern
Biotechnology Associates; HRP anti-rabbit was from Dako; FITC anti-mouse ␥
chains were from Jackson Laboratories. Anti-Myc (9E10) and anti-ERGIC-53
(G1/93) mAbs were described elsewhere (Evan et al., 1985; Schweizer et al., 1988),
and anti-GM-130 was purchased from Transduction Labs.
Cell lines
NSO (Cowan et al., 1974), J558L (Oi et al., 1983) and the derived stable
transfectants N[␮1], N[␮⌬CH1] and J[␮⌬CH1] were obtained and maintained as
described previously (Sitia et al., 1987; Sitia et al., 1990; Valetti et al., 1991). HeLa
Tet-Off (Clontech Laboratories) were maintained in DMEM as recommended by
the supplier.
Journal of Cell Science
Plasmids and vectors
cDNA encoding ␮⌬CH1 was prepared by RT-PCR from total RNA isolated from
J[␮⌬CH1] cells. The first strand cDNA was synthesized by incubation with a ␮specific primer (GGTTAGTTTGCATACACAGAG) and Moloney murine leukemia
virus reverse transcriptase RNase H Minus Point Mutant (Promega) for 1 hour at
42°C. The resulting cDNA was used for PCR amplification with primers containing
unique restriction sites: EcoRI for the forward primer (GGAATTCGCACACAGGACCTCACC) and XbaI for the reverse primer (AAATCTAGACTGGTTGAGCGCTAGCATGG). PCR products were digested and cloned in pTRE (Clontech
Laboratories Inc.) and in pCDNA3.1(+) (Invitrogen, Life Technologies).
pcDNA3.1 encoding ␭1 chain was described previously (Fagioli et al., 2001).
The truncated variant lacking the constant domain (VL), was obtained by
PCR amplification with primers containing unique restriction sites, EcoRI
(GAATTCATGGCCTGGATTTCACTT) and XbaI (CTCGAGCTATAGGACAGTCAGTTTGGT) for the forward and reverse primers, respectively. The latter
contains a stop codon before the XbaI site. PCR products were cloned in
pCDNA3.1(+).
Vectors encoding Myc-tagged, glycosylated human wild-type and mutant
ERGIC-53-KKAA and ERGIC-53-N156A (Appenzeller et al., 1999) were kind
gifts from Hans-Peter Hauri (Biozentrum, University of Basel, Switzerland).
Establishment of Tet-inducible HeLa cells
HeLa Tet-Off cells were transfected with pTRE-␮⌬CH1 (50 ␮g) and pTK-Hyg (5
␮g, Clontech Laboratories). Clones were grown in the presence of 3 ␮g/ml
tetracycline, 400 ␮g/ml hygromycin and 100 ␮g/ml geneticin. Hygromycinresistant clones were screened by immunofluorescence 72 hours after tetracycline
removal.
Cell lysis and western blotting
Cells were washed and lysed at the concentration of 1⫻104 cells/␮l in buffer A
(0.2% TX100, 50 mM Tris-HCl pH 7.5), 150 mM NaCl, 5 mM EDTA, 1 mM Nethylmaleimide and a cocktail of protease inhibitors (Roche). The TX100-insoluble
fraction was separated by centrifugation at 3,400 g for 10 minutes, washed twice in
buffer A and further solubilized in lysis buffer B (1% SDS, 50 mM Tris-HCl pH
7.5) for 10 minutes at room temperature (RT), diluted in 50 mM Tris-HCl pH 7.5,
0.2% TX100, to keep the volume of the soluble and insoluble fractions equal, and
sonicated for 10 seconds. Immunoprecipitation and western blotting were performed
as previously described (Valetti et al., 1991). EDTA was omitted from buffer A, and
1 mM CaCl2 added in co-immunoprecipitation assays involving ERGIC-53. The
intensity of the relevant bands was quantified by the software package, IPLab
spectrum V3.2 (Scanalytics, Rockville, MD).
Chemical cross-linking
Cells were washed three times with Hanks and incubated for 10 minutes at RT with
1 mM DSP on a rocking platform. The reaction was quenched by rinsing the cells
twice with 50 mM Tris-HCl pH 7.5 followed by incubation for 5 minutes at RT with
the same buffer.
Immunofluorescence microscopy
Lymphoid cells were incubated on poly-L-lysine-coated coverslips for 20 minutes
at RT whereas HeLa Tet-Off cells were grown directly on 10-mm2 coverslips. Cells
were fixed in 3% paraformaldehyde and permeabilized with 0.1% TX100; for ␥tubulin staining, cells were fixed in methanol at –20°C. In cells expressing ␮⌬CH1,
pre-adsorbed ␥-chain-specific rabbit anti-mouse antibodies were used to avoid
cross-reactions. Samples were inspected on an Olympus inverted fluorescence
microscope (model IX70). Images were recorded with a digital CCD camera (model
C4742-95, Hamamatsu), using the IPLab spectrum V3.2 software package
(Scanalytics) and processed with Adobe Photoshop 7.0 (Adobe Systems).
Electron microscopy
For cryosectioning, cells were fixed with 2% paraformaldehyde and 0.2%
glutaraldehyde in PBS for 2 hours, washed with PBS containing 20 mM glycine,
scraped off the dish, centrifuged and embedded in 12% gelatin in PBS. Small blocks
of embedded cells were incubated overnight with 2.3 M sucrose at 4°C, mounted
on aluminum pins and frozen in liquid nitrogen. 60 nm ultrathin cryosections were
cut at –120°C, using a cryo-ultramicrotome (Leica-Ultracut EM FCS), and picked
up with 1% methylcellulose in 1.15 M sucrose. Cryosections were then incubated
with primary antibodies and revealed with protein A gold, according to previously
described protocols (Slot et al., 1989).
For Epon embedding, cells were fixed with 0.1 M cacodylate buffer, pH 7.4,
containing 2.5% glutaraldehyde at RT for 10 minutes. After washing with 0.1 M
cacodylate buffer pH 7.4, and post-fixing for 10 minutes at RT with 0.1 M
cacodylate buffer, containing 1% OsO4 w/v. Cells were then washed with 0.1 M
cacodylate buffer, and processed for Epon embedding (Polybed 812, Polysciences).
Sections were analyzed with a Philips CM-10 electron microscope (Einhdoven, the
Netherlands).
This work is dedicated to the memory of César Milstein, who raised
our interest in Russell bodies. We thank S. Bonatti (University of
Naples), C. E. Grossi (University of Genoa), H.-P. Hauri (University
of Basel) and M. Otsu (DiBiT-HSR, Milan), for providing excellent
reagents and helpful suggestions; K. Cortese, M. C. Gagliani and M.
Bono for help with electron microscopy. This work was supported
through grants from the Associazione Italiana per la Ricerca sul
Cancro (AIRC), Ministero della Sanità, MIUR (CoFin and Center of
Excellence in Physiopathology of Cell Differentiation) and Telethon.
Electron microscopy studies were performed at the Telethon Facility
for Electron Microscopy (Grant no. GTF03001). T.A. was a recipient
of a fellowship from the Federazione Italiana Ricerca sul Cancro.
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