Protein Synthesis and Quality Control at the Endoplasmic Reticulum
Organization of translocon complexes
in ER membranes
A.V. Nikonov* and G. Kreibich*†1
*Department of Cell Biology, New York University School of Medicine, New York, NY 10016, U.S.A., and †Kaplan Comprehensive Cancer Center,
New York University School of Medicine, New York, NY 10016, U.S.A.
Abstract
Protein translocation in the ER (endoplasmic reticulum) and N-glycosylation are fundamental processes
essential for the normal functioning of eukaryotic cells. They are the initial steps in the intracellular
pathway that are followed by secretory proteins and membrane proteins of the endomembrane system
and the plasma membrane. The translocation and concurrent N-glycosylation of these proteins take place
on a large molecular machine, the TC (translocon complex), which is associated with membrane-bound
polysomes. Segregation of TCs into a differentiated domain of the ER, the rough ER, may increase the
efficiency of protein synthesis on membrane-bound polysomes. Our research is concerned with the assembly,
functional organization and dynamics of the TCs in the ER, and their contribution to the functioning and
the morphological appearance of this organelle. We hypothesize that the TCs form higher-order structures
defining the rough domain of the ER. These structures, which are immobilized or diffuse slowly in the plain of
the ER membrane, may be formed and stabilized by mRNAs interconnecting the TCs, by cytoskeletal elements
and/or by hypothetical proteins that form links between the TCs. We have established the M3/18 cell line,
which expresses the GFP (green fluorescent protein)–Dad1 fusion protein quantitatively and functionally
incorporated into the OST (oligosaccharyltransferase). GFP–Dad1 can be used as a reporter molecule for the
lateral mobility of the TCs since the OST is tightly associated with the complex. As determined by FRAP
(fluorescence recovery after photobleaching), the lateral mobility of GFP–Dad1-tagged TCs was much more
restricted than expected from the estimated size of the TC and can be affected by the functional state of
the TCs. Currently, we are studying the possible involvement of cytoskeletal elements in the organization
of the TCs. Our data suggest that microtubules also play a role in the immobilization of the TCs.
Components of the TC (translocon
complex)
The RER [rough ER (endoplasmic reticulum)] is the site
where proteins made on membrane-bound polysomes are
inserted into or translocated across the ER membrane.
Here, a complex molecular machinery effects the signalsequence-mediated targeting, co-translational translocation
and processing of nascent polypeptide chains [1]. Present
evidence suggests that the proteins concerned with these
functions form an oligomeric structure, the TC, which has
Sec61p at its core [2,3] (Figure 1). Our research on the TC had
focused primarily on the structure and function of the OST
(oligosaccharyltransferase). This complex has been identified
and purified from various species [4–6]. Ribophorin I and
II were originally described in our laboratory as the first
membrane proteins specific to the RER [7]. From previous
findings concerning the near-neighbour relationship of the
ribophorins with other components of the TC, it is very likely
that the OST is intimately associated with membrane-bound
Key words: endoplasmic reticulum, fluorescence recovery after photobleaching (FRAP), lateral
mobility, microtubule, oligosaccharyltransferase, translocon complex.
Abbreviations used: ER, endoplasmic reticulum; FRAP, fluorescence recovery after photobleaching; LBR, lamin B receptor; OST, oligosaccharyltransferase; RER, rough ER; TC, translocon complex;
TMD, transmembrane domain; GFP, green fluorescent protein.
1
To whom correspondence should be addressed (e-mail [email protected]).
ribosomes and Sec61p [8–10]. It had been shown that when
rough microsomes are solubilized with certain non-ionic
detergents, membrane-bound polysomes co-sediment with
the TC, including the OST [7–9,11]. Also, the ribophorins can
be chemically cross-linked to the 60 S ribosomal subunit [12],
and affinity-purified polyclonal and anti-peptide antibodies
against ribophorin I and II inhibit the translocation of nascent
polypeptide chains [13].
Overall, the TC is composed of more than 25 polypeptides
with a total of at least 60 TMDs (transmembrane domains),
and it has a molecular mass of about 850 kDa [2,3]. The best
evidence for the size and composition of the TC comes from
recent structural studies on membrane-bound ribosomes
isolated under conditions that preserve their interaction with
the TC. The overall diameter of the TC is about 125 Å
[10]. An active TC carries a ribosome (250 Å in diameter),
which has a molecular mass of about 5000 kDa and may
also be associated with cytosolic factors [2,14–16]. This
would increase the mass to 7000 kDa, with an additional
500 kDa contributed by each 1 kb of mRNA. If active TCs
organized into a polysome synthesizing a protein of 20 kDa,
the structure will have the molecular mass of more than
36 600 kDa and have at least 305 TMDs. The molecular mass
and number of TMDs will increase proportionally with the
size of the protein encoded by an mRNA. Such a large
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Figure 1 Stages of TC assembly
The individual subunits of Sec61p or the OST (A) are rather quickly integrated into oligomeric structures (B), which are
assembled into the TC (C). The TCs are recruited into a polysome (D). Some of the membrane-bound polysomes may be
segregated in differentiated domains of the RER (E). Since the diameter of a ribosome (≈250 Å) is significantly larger
than that of the TC (125 Å) [10], RER-specific proteins (solid ellipses) associated with TCs could provide physical interaction
between adjacent TCs to form large molecular arrays. Attachment of microtubules via ER membrane proteins such as
CLIMP-63 [17] could also restrain the lateral mobility of the TC.
structure with TMDs interacting with the lipid bilayer, and
domains exposed to the cytosol and the ER lumen, may
be subjected to considerable friction that would impede its
lateral diffusion.
FRAP (fluorescence recovery after
photobleaching) as a tool for studying the
functional organization of the TCs
The fluid mosaic membrane model that was proposed about
30 years ago emphasized the free mobility of membrane
proteins in the plane of the membrane [18]. Since then it has
been found that many membrane proteins are not completely
free to diffuse in the plane of the lipid bilayer. Such proteins
may be retarded in their lateral diffusion or completely
immobilized through interactions with cytoskeletal elements,
with extracellular matrix proteins or by becoming a part of
large oligomeric arrays. These insights were gained at least in
part through biophysical methods including FRAP [19,20].
In FRAP analysis, a small area of a cell that contains a fluorescent probe, such as GFP (green fluorescent protein),
is exposed to irreversible photobleaching radiation by an
intense laser flash. The fluorescence recovery of the bleached
area, through exchange of bleached for non-bleached molecules, is recorded with an attenuated laser beam. The rate of
recovery is used to calculate an effective diffusion constant
(Deff ) for the mobile fraction of the GFP-tagged structure
[20]. The lateral diffusion of integral membrane proteins in
the lipid bilayer decreases with the natural logarithm of the
radius of the diffusing molecule [21]:
Deff = k(ln c/a − 0.577)
(1)
where Deff is the diffusion constant, k and c are constants
that take into account the membrane bilayer thickness, its
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viscosity and the viscosity of the surrounding medium, and a
is the radius of the TMDs of the diffusing membrane protein
complex. Edidin and co-workers [22] used this equation to
calculate the size of the GFP-tagged H2Ld or TAP1 complex.
We have recently described M3/18 cells expressing a GFP–
Dad1 fusion protein that is functionally integrated into the
OST as one of the subunits of the complex [23]. We have demonstrated that the mobility of GFP–Dad1-tagged active TCs
is about seven times lower than that of the highly mobile
LBR (lamin B receptor)–GFP construct [23]. These results
provided the first direct evidence that the lateral mobility of
TCs in ER membranes of the living cells is severely impaired.
Using eqn (1) and assuming that friction is only caused by
interactions of TMDs with the lipid bilayer, we calculated
that the TCs would be organized into large rafts occupying
an area of approx. 5.7 × 102 µm2 . Considering the state and
morphology of the RER in these cells, one has to assume that,
besides the physical size of the rafts, other mechanisms are
expected to play a role in restricting the mobility of TCs.
Mechanisms that affect the lateral
mobility of TCs
The restricted mobility of the TCs was suggested by previous
findings. (i) In hepatocytes, which have a well-differentiated
RER and smooth ER, a sharp transition between these
two continuous membrane domains is observed. (ii) The
morphology of the RER (flattened cisternae) and the smooth
ER (contorted tubules) is strikingly different. (iii) In grazing
sections of the RER, membrane-bound polysomes are
arranged in regular figures that would not be expected if
TCs carrying them were freely mobile. (iv) Large aggregates
of polysomes containing components of the TC can be
isolated from rough microsomes solubilized with non-ionic
Protein Synthesis and Quality Control at the Endoplasmic Reticulum
detergents. On electron micrographs, the detergent-insoluble
microsomal residues were found to consist of curved membrane remnants with ribosomes attached to a proteinaceous
fibrillar meshwork that was insensitive to RNase treatment
[7,24].
Biochemical evidence [7,11], as well as our FRAP results
[23], suggested that membrane-bound polysomes and thus
TCs may be segregated in a RER-specific domain. By direct
or indirect interactions among TCs they may form large rafts,
which may provide a more efficient environment for protein
synthesis on membrane-bound polysomes. In pancreatic
AR42J cells, we have shown that the induction of amylase
synthesis and secretion by dexamethasone results in a striking
rearrangement of the RER from a tubulo-vesicular configuration into stacks of cisternae without an increase in the
number of membrane-bound ribosomes [25]. Morphological
research dating back to the early work by Palade [26,27]
described particles on the surface of ER membranes that are
arranged in characteristic patterns, such as circles, spirals,
loops or hairpins, which have been later recognized as
membrane-bound polysomes. Christensen’s laboratory has
shown that large mRNAs encoding thyroglobulin (330 kDa)
or collagen (150 kDa) generate ribosome arrangements in the
form of spirals or double rows that contain more than 30
ribosomes [28]. It was found that the spacing between the
centres of ribosomes along the mRNA was quite constant
(about 250 Å) and independent of the type of the translated
mRNA. The regular spacing between ribosomes requires
that during translation the rate of initiation and elongation
remains constant. Irregularities in these processes, including
‘ribosome pausing’ during translation [29], would result
in prominent gaps and clustering of ribosomes, unless the
TCs are part of a rigid, raft-like structure. Furthermore,
the average actual length of the mRNA segments that
interconnect adjacent ribosomes is about double the length
than would be needed to interconnect adjacent ribosomes.
Therefore, the distance between ribosomes, which is constant,
is not defined by the length of the interconnecting mRNA
segment or the diameter of the ribosome [28]. The authors
proposed a model in which ‘. . . adjacent strands of mRNA in
large bound polysomes would be maintained at a relatively
constant distance from one another by spacing mechanisms,
perhaps involving membrane proteins of the RER’. The
diameter of the TC is about 125 Å [3,10]. Since the distance
between the ribosomes is much larger than the diameter of
the TC [28], linker proteins would be required if TCs form
raft-like arrays (Figures 1 and 2B).
The lateral mobility of TCs is affected by
depolymerization of microtubules
Another possible explanation for the low lateral mobility
of GFP–Dad1 in M3/18 cells is that the active TCs are
partially immobilized by interaction of membrane-bound
ribosomes with cytosolic components including cytoskeletal
elements. It has been shown that the injected dextran particles
of the size of ribosomes are restrained in their diffusion
Figure 2 The lateral mobility of TCs can be affected by
mechanisms involving cytoskeletal elements, such as
microtubules, or proteins that bridge adjacent TCs
(A) Schematic representation of membrane-bound polysomes together
with microtubules that are linked to the ER membrane via a microtubuleanchoring protein such as CLIMP-63. Two ribosomes that are not part of
the polysomes are also shown. (B) Model depicting the immobilization
of TCs associated with membrane-bound polysomes by cross-bridges
interconnecting adjacent TCs. The black circles represent active TCs, while
grey circles interconnected by mRNAs indicate the ribosomes. Crosshatched circles depict inactive TCs that do not carry ribosomes. Linker
proteins are represented by dark grey ellipses.
but still mobile. The high protein concentration in the
cytoplasm does not, however, explain the immobilization
of ribosome-size particles by itself, since the particles are
still mobile in protein solutions with a protein concentration
comparable with that found in the cytoplasm [30,31]. The
slow diffusion of TCs observed in our previous FRAP
experiments may therefore be caused, at least in part,
by interaction of membrane-bound ribosomes with the
cytoskeletal elements. The ER is known to form an elaborate
network of tubules and cisternae that are thought to
be stabilized by microtubules [32]. Recent ultrastructural
reconstructions of the three-dimensional arrangement of
cellular organelles showed in fact that microtubules are in
very close proximity to the ER [33]. Moreover, it has been
shown that the RER-specific membrane protein CLIMP63, which itself can form oligomers, apparently tethers
microtubules to the ER [17]. There are two possibilities
with regard to how cytoskeletal elements may interact with
TCs. (a) They are forming a grid-like structure parallel
to the plane of the ER membrane. Even if they are not
directly attached to ER membranes, they may provide
fence-like barriers for the diffusion of membrane-bound
ribosomes. (b) Cytoskeletal elements are attached to the ER
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Figure 3 Treatment of the cells with vinblastin results in
increased lateral mobility of the TCs
M3/18 cells were grown at 39.5◦ C and left untreated (1) or treated with
(2) cycloheximide (80 µM; 30 min), (3) puromycin (100 µM; 30 min),
(4) lantrunculin B (10 µM; 2 h), (5) taxol (5 µM; 4 h) or (6) vinblastin
(10 µM; 2 h). FRAP experiments and data analyses were carried out as
described previously [23] except that a heated stage (39.5◦ C) was used.
causes depolymerization of microtubules, resulted in a faster
diffusion of the TCs. Since we did not detect any inhibition
of protein synthesis in vinblastin-treated cells, the observed
increases in the lateral mobility of the TCs strongly suggest
that microtubules in close proximity to the ER may affect the
mobility of membrane-bound polysomes.
This work was supported by a grant from the American Cancer
Society (RPG-92-009-06-CB) and A.V.N. was supported by a NIH
grant in Dermatology (5T32 ARO 7190).
References
membranes via receptors or adaptors, thus forming a post-like
arrangement perpendicular to the ER membrane. In this arrangement, cytoskeletal elements may affect predominantly
polysomal assemblies that get entangled via their mRNAs.
Both possible arrangements of microtubules with regard to
the plane of the ER membrane are illustrated in Figure 2(A).
To test whether cytoskeletal elements interfere with the lateral mobility of TCs we have performed FRAP experiments
on M3/18 cells that were treated with drugs that affect the
state of the cytoskeleton (Figure 3). The experiments were
performed as described previously [23], except that they
were done on a heated stage (39.5◦ C) and not at room
temperature. Under these conditions the lateral mobility of
GFP–Dad1 was somewhat higher than previously reported.
However, we also observed a higher lateral mobility for
LBR–GFP (Deff = 0.395 µm2 /s) than that observed at room
temperature (0.330 µm2 /s) [23]. The new Deff value for LBR–
GFP is in good agreement with results reported by others
[34]. The new set of data confirmed our early findings that
the lateral mobility of the TCs is severely restricted in control
cells (Figure 3, bar 1), and addition of cycloheximide to
the growth medium had no significant effect on the their
diffusion (Figure 3, bar 2). In contrast, treatment of the
cell with puromycin, which causes termination of protein
synthesis and disassembly of polysomes, resulted in a 3fold increase in the diffusion rate of the TCs (Figure 3,
bar 3). Treatment of the cells with lantrunculin B, a drug
that causes disassembly of actin microfilaments, and taxol,
which stabilizes microtubules, had no significant effect on the
lateral mobility of TCs (Figure 3, bars 4 and 5 respectively).
However, treatment of the cells with vinblastin, which
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1 Sabatini, D.D. and Adesnik, M. (1995) in The Metabolic Basis of Inherited
Disease (Scriver, C.R., Beaudet, A.L., Sly, W.S. and Valle, D., eds.),
pp. 459–553, McGraw-Hill, New York
2 Rapoport, T.A., Jungnickel, B. and Kutay, U. (1996) Annu. Rev. Biochem.
65, 271–303
3 Johnson, A.E. and van Waes, M.A. (1999) Annu. Rev. Cell Dev. Biol. 15,
799–842
4 Kelleher, D.J., Kreibich, G. and Gilmore, R. (1992) Cell 69, 55–65
5 Kelleher, D.J. and Gilmore, R. (1994) J. Biol. Chem. 269, 12908–12917
6 Knauer, R. and Lehle, L. (1999) Biochim. Biophys. Acta 1426, 259–273
7 Kreibich, G., Ulrich, B.L. and Sabatini, D.D. (1978) J. Cell Biol. 77, 464–487
8 Görlich, D., Prehn, S., Hartmann, E., Kalies, K.U. and Rapoport, T.A. (1992)
Cell 71, 489–503
9 Wang, L. and Dobberstein, B. (1999) FEBS Lett. 457, 316–322
10 Menetret, J., Neuhof, A., Morgan, D.G., Plath, K., Radermacher, M.,
Rapoport, T.A. and Akey, C.W. (2000) Mol. Cell 6, 1219–1232
11 Marcantonio, E.E., Amar-Costesec, A. and Kreibich, G. (1984) J. Cell Biol.
99, 2254–2259
12 Kreibich, G., Freienstein, C.M., Pereyra, B.N., Ulrich, B.L. and Sabatini, D.D.
(1978) J. Cell Biol. 77, 488–506
13 Yu, Y.H., Sabatini, D.D. and Kreibich, G. (1990) J. Cell Biol. 111,
1335–1342
14 Walter, P. and Johnson, A.E. (1994) Annu. Rev. Cell Biol. 10, 87–119
15 Wiedmann, B., Sakai, H., Davis, T.A. and Wiedmann, M. (1994) Nature
(London) 370, 434–440
16 McCallum, C.D., Do, H., Johnson, A.E. and Frydman, J. (2000) J. Cell Biol.
149, 591–602
17 Klopfenstein, D.R., Kappeler, F. and Hauri, H.P. (1998) EMBO J. 17,
6168–6177
18 Singer, S.J. and Nicholson, G.L. (1972) Science 175, 720–731
19 Jacobson, K., Sheets, E.D. and Simson, R. (1995) Science 268, 1441–1442
20 Lippincott-Schwartz, J., Presley, J.F., Zaal, K.J., Hirschberg, K., Miller, C.D.
and Ellenberg, J. (1999) Methods Cell Biol. 58, 261–281
21 Vaz, W.L., Criado, M., Madeira, V.M., Schoellmann, G. and Jovin, T.M.
(1982) Biochemistry 21, 5608–5612
22 Marguet, D., Spiliotis, E.T., Pentcheva, T., Lebowitz, M., Schneck, J. and
Edidin, M. (1999) Immunity 11, 231–240
23 Nikonov, A.V., Snapp, E., Lippincott-Schwartz, J. and Kreibich, G. (2002)
J. Cell Biol. 158, 497–506
24 Marcantonio, E.E., Grebenau, R.C., Sabatini, D.D. and Kreibich, G. (1982)
Eur. J. Biochem. 124, 217–222
25 Rajasekaran, A.K., Morimoto, T., Hanzel, D.K., Rodriguez-Boulan, E. and
Kreibich, G. (1993) J. Cell Sci. 105, 333–345
26 Palade, G.E. (1956) J. Biophys. Biochem. Cytol. 2, 171–200
27 Palade, G.E. (1955) J. Biophys. Biochem. Cytol. 1, 59–61
28 Christensen, A.K. and Bourne, C.M. (1999) Anat. Rec. 255, 116–129
29 Wolin, S.L. and Walter, P. (1993) J. Cell Biol. 121, 1211–1219
30 Luby-Phelps, K., Castle, P.E., Taylor, D.L. and Lanni, F. (1987) Proc. Natl.
Acad. Sci. U.S.A. 84, 4910–4913
31 Seksek, O., Biwersi, J. and Verkman, A.S. (1997) J. Cell Biol. 138, 131–142
32 Terasaki, M., Chen, L.B. and Fujiwara, K. (1986) J. Cell Biol. 103,
1557–1568
33 Marsh, B.J., Mastronarde, D.N., Buttle, K.F., Howell, K.E. and McIntosh, J.R.
(2001) Proc. Natl. Acad. Sci. U.S.A. 98, 2399–2406
34 Ellenberg, J., Siggia, E.D., Moreira, J.E., Smith, C.L., Presley, J.F., Worman,
H.J. and Lippincott-Schwartz, J. (1997) J. Cell Biol. 138, 1193–1206
Received 9 June 2003