From the Department of Biology, Massachusetts Institute of Technology, Cambridge, Massachusetts 02139
Received for publication, September 15, 2000, and in revised form, March 20, 2001
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Little is known about the conformations of newly
synthesized polypeptide chains as they emerge from the large ribosomal
subunit, or how these conformations compare with those populated
immediately after dilution of polypeptide chains out of denaturant
in vitro. Both in vivo and in
vitro, partially folded intermediates of the tailspike protein
from Salmonella typhimurium phage P22 can be trapped in the
cold. A subset of monoclonal antibodies raised against tailspike
recognize partially folded intermediates, whereas other antibodies
recognize only later intermediates and/or the native state. We have
used a pair of monoclonal antibodies to probe the conformational
features of full-length, newly synthesized tailspike chains recovered
on ribosomes from phage-infected cells. The antibody that recognizes
early intermediates in vitro also recognizes the
ribosome-bound intermediates. Surprisingly, the antibody that did not
recognize early in vitro intermediates did recognize
ribosome-bound tailspike chains translated in vivo. Thus,
the newly synthesized, ribosome-bound tailspike chains display structured epitopes not detected upon dilution of tailspike chains from
denaturant. As opposed to the random ensemble first populated when
polypeptide chains are diluted out of denaturant, folding in
vivo from the ribosome may begin with polypeptide conformations already directed toward the productive folding and assembly pathway.
Although a subset of globular proteins can refold correctly
in vitro, many other proteins fail to fold in the test tube
(1, 2). For the limited set of proteins whose folding has been investigated both in vivo and in vitro,
polypeptide chains fold more efficiently in their native cellular
environments (3, 4). Part of the difference between folding in
vitro and in vivo is due to the action of molecular
chaperones; these protein complexes increase the yield of successfully
folded protein for some polypeptide sequences (5, 6). Yet there are
many proteins whose folding appears to be unaided by the presence of
molecular chaperones, particularly in vivo (7-10).
The N-terminal domain(s) of some proteins are able to fold
co-translationally to native-like conformations (8, 11, 12); many other
proteins are unable to fold correctly in the absence of C-terminal
domains (13-15). These results suggest that additional cellular
components may be required for the successful folding of these
polypeptide chains in vivo. The recent discovery that the
ribosome-associated trigger factor may cooperate with the molecular
chaperone DnaK to assist in the folding of some polypeptide chains (16,
17) suggests that additional parts of the cellular folding machinery
may lie very close to the ribosome (18) and may involve the ribosome
itself (19, 20).
What is the conformational state of nascent polypeptide chains not
bound by molecular chaperones? During polypeptide synthesis, after
amino acids are added to the growing polypeptide chain at the
aminoacyltransferase center within the ribosome, they pass through a
tunnel, emerging at the exit site on the back side of the large
ribosomal subunit (21-23). Newly synthesized polypeptide chains must
be held in some conformation in order to protect the nascent chain from
proteolytic digestion or unproductive associations with other nascent
or unfolded chains. This is true even during co-translational folding,
which still appears to require the presence of a complete folding
domain (50-200 amino acids) before folding can commence (10).
The experimental difficulties involved in studying a dynamically
growing nascent polypeptide chain, attached to a large (2-4 MDa)
ribosome, have limited studies of the conformations of nascent chains
on actively translating ribosomes. Most investigations have focused on
the interactions of nascent chains with chaperones (24, 25), or
co-translational folding of nascent polypeptide chains to native or
native-like states, detected by immunogold labeling (26), enzymatic
activity (3, 8), or protection from protease digestion (13, 27). Many
of these conformational studies have used stalled ribosomes, generated
either chemically (28) or with truncated mRNA messages (12) and/or
synchronized cell-free translation systems (13, 27), to trap and
populate discrete ribosome-nascent chain species. Investigations using physiological translation systems have been rare (8).
In this paper, we examine the conformation of ribosome-bound, nascent
polypeptide chains of the tailspike protein from Salmonella typhimurium phage P22. Tailspike is a homotrimeric protein with 666 amino acid residues per chain, arranged in an extended structure that includes a large central parallel
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-helix domain
(Fig. 1) (29, 30). Near the C terminus,
the three separate amino acid chains wrap around one another to form an
interdigitated triangular
-prism domain (31). Past folding studies
have demonstrated that trimerization, and the subsequent
interdigitation of the C-terminal domain, is crucial for correct
formation of the native structure (32). Tailspike is unusual in that it
represents one of the few proteins whose folding and assembly has been
studied extensively both in vitro and in vivo
(32-37). Because the SDS-resistant native state is distinguishable
from its folding and assembly intermediates by SDS or native gel
electrophoresis, the folding and assembly process can be monitored in
crude extracts (33, 38). During folding in vivo, partially
folded monomeric and trimeric intermediates have been identified (34).
In vitro intermediates have been characterized by Seckler
and co-workers (32, 39), using fluorescence and circular dichroism
spectroscopy. Both monomeric and trimeric intermediates have been
identified, corresponding to the in vivo intermediates (32).
Fig. 2 shows a schematic diagram for the
pathway of tailspike folding and assembly after release of polypeptide
chains from ribosomes or dilution of chains out of denaturant.
View larger version (50K):
[in a new window]
Fig. 1.
Ribbon diagram of the crystal
structure of P22 tailspike (29). The N-terminal head-binding
domain was crystallized separately (30). The three polypeptide chains
of the monomer are depicted in red, blue, and green.
Ovals indicate the approximate epitope locations for the
monoclonal antibodies used in this study; for clarity, the positions of
the epitopes are indicated only for the red polypeptide
chain. The dorsal fin, -helix, and interdigitated domains are
labeled.
View larger version (17K):
[in a new window]
Fig. 2.
The folding pathway of released P22 tailspike
chains (63). Unfolded polypeptide chains organize first into
structured monomeric intermediates (M), which can either proceed down
the refolding pathway or convert into aggregation-prone
monomers. All intermediates can be trapped by cold incubation of
in vitro refolding mixtures or newly lysed cells from
in vivo assays. Folding proceeds through a trimeric
intermediate termed the protrimer (Pt), which migrates as a
discrete band on native polyacrylamide gels (64), observed during both
in vivo folding and in vitro refolding studies
(34, 53). Formation of the native trimer (T) is slow
(t1/2 ~5 min in vivo; >24 h in
vitro at 10 °C). The native trimer is resistant to unfolding by
SDS and therefore appears as a distinct band on SDS-polyacrylamide
gels. The folding and aggregation intermediates are sensitive to
SDS-unfolding and therefore co-migrate during SDS-PAGE.
Unlike the folding of many small globular proteins, tailspike folding exhibits Arrhenius behavior (i.e. linear dependence of the folding rate on temperature) (39). The temperature dependence is quite steep, resulting in an activation energy of ~165 kJ/mol (39). Although the reasons for non-Arrhenius behavior during folding are still under debate (40, 41), for the tailspike it appears that the stabilities and kinetics of the phases of folding are equally affected by temperature. A critical feature of this behavior is that tailspike intermediate populations can be trapped in the cold and remain competent for further folding for many hours (37).
The single-chain and the protrimer intermediates trapped in the cold can be separated by electrophoresis through non-denaturing acrylamide gels run in the cold (33). Unlike the microsecond and millisecond lifetimes for the folding intermediates of many small globular proteins, tailspike folding intermediates persist for several hours (32). Warming cold-trapped intermediates permits their continued folding to the native trimer (37). Cold-trapping therefore provides a method for temporarily stabilizing transient folding intermediates, permitting their investigation with more time-consuming experimental methods, without altering the tailspike folding pathway (34).
The tailspike single-chain partially folded intermediate is thermolabile, and temperature-induced destabilization of this intermediate causes tailspike folding to partition from the productive pathway to the aggregation pathway (33, 38). Although off-pathway aggregation occurs both in vivo and in vitro, the intermediates in vivo are less labile, requiring higher temperatures to initiate aggregation. Overexpression of GroEL/S does not alter the yield of native tailspike nor does it rescue tailspike folding mutants (42, 43). In addition, newly synthesized wild type tailspike chains do not appear to be associated with DnaK (44) or the GroE complex (43). Despite the similarities between the in vitro and in vivo folding pathways, tailspike folding in vivo always results in higher yields of folded protein. These inconsistencies prompted us to examine other participants in protein folding in vivo; specifically, whether the translation machinery itself can stabilize early intermediates in vivo.
A set of monoclonal antibodies isolated against native tailspike recognizes at least seven different classes of epitopes on the native structure (45).1 Three of these antibodies were used previously to monitor the formation of intermediates and the native state during tailspike refolding in vitro (46). The epitopes recognized by these antibodies appeared at different stages during refolding. One of the antibodies, 236-3, in addition to recognizing in vitro refolding intermediates, also detected tailspike polypeptide chains in the presence of immunoprecipitated ribosomes from in vitro translation mixtures. However, two other antibodies, which recognized epitopes present only on later intermediates and the native trimer during refolding in vitro, did not detect tailspike polypeptide chains in the presence of the immunoprecipitated ribosomes. These results suggest the ribosome-associated tailspike chains contained significant, but not yet native-like, levels of structural information (46).
The methods used previously to analyze in vivo intermediates
of tailspike folding (33) disrupted the ribosomal structures just prior
to analysis. In order to determine the conformation of the nascent
tailspike polypeptide chain while still attached to the ribosome, we
characterized newly synthesized tailspike polypeptide chains using cell
preparations from which intact ribosomes were recovered. Below we
report the adoption of conformational motifs in newly synthesized,
ribosome-bound tailspike polypeptide chains, prior to their release
into the cytoplasm, and we compare these conformations with those
adopted by tailspike polypeptide chains at the onset of in
vitro refolding.
![]() |
EXPERIMENTAL PROCEDURES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
In Vitro Refolding-- Tailspike was purified from P22-infected Salmonella cultures (see description below) as described previously (47). Protein concentrations were determined based on an absorbance value of 1.0 at 278 nm for a 1 mg/ml solution (48). Tailspike was denatured and refolded as described previously (32, 46). Briefly, 1 mg/ml tailspike was unfolded in 5 M urea, 63 mM sodium phosphate, 0.6 mM EDTA, pH 2.9, for 30 min at 25 °C. Aliquots of the unfolded protein were refolded at various time points (up to 24 h) by 50-fold dilution into 50 mM sodium phosphate, 1 mM EDTA, 0.5 mM dithiothreitol, pH 7.0. Refolding was performed at 10 °C and, as judged by SDS-PAGE,2 resulted in ~75% of denatured chains recovering the native conformation after 24 h, consistent with previous results (32, 46). The remaining ~25% of denatured chains form non-native aggregates. After the final sample was diluted into refolding buffer, all samples were then placed on wet ice and diluted to a tailspike concentration conducive to the competition ELISA (i.e. a concentration where complete refolding of the protein would result in 70% antibody recognition, in order to stay within the sensitive range of the assay; see below) (46). Samples were then subjected to the competition ELISA (described below) but with an initial antigen-antibody incubation at 4 °C for only 10 min; loading and incubation of these mixtures in 96-well plates was also performed at 4 °C. Plate data were analyzed as described below and compared with the antibody recognition for an equivalent concentration of native (never unfolded) tailspike.
Cell Growth and Tailspike Expression--
S.
typhimurium strain DB7136(su) was grown
and infected with phage P22 as described previously (47). Briefly,
cells were grown at 30 °C to a density of 2 × 108
cells/ml and infected with a P22 variant,
5
amN114/13
amH101
(amber mutations in genes 5 and 13 prevent
formation of full-length coat and lysis proteins, respectively, thereby
preventing phage head assembly and bacterial cell lysis). As a control
for tailspike production, cells were infected with a P22 variant
containing the aforementioned amber mutations in genes 5 and
13 as well as an amber mutation in the tailspike,
9
amN110, which produces only a
small N-terminal fragment of tailspike shown previously to be
unrecognized by the anti-tailspike monoclonal antibodies (45). Infected
cells were cultured for 90 min at 30 °C. Cultures were chilled
quickly on ice, and ice cubes of buffer R (50 mM Tris, pH
7.5, 10 mM MgCl2, 150 mM KCl) were
added to speed chilling. Cells were collected by centrifugation at
4000 × g and resuspended in buffer R containing 80 mM octyl glucopyranoside, to promote cell lysis. Cells were
then frozen in liquid nitrogen and stored at
80 °C.
Isolation of Ribosome-Nascent Chain Complexes-- All steps were performed on ice or at 4 °C, in order to halt tailspike polypeptide chain maturation (37). Cells were defrosted for 3 min in a cold water bath, until just thawed; this freeze/thaw cycle was sufficient to effect cell lysis. Lysates were spun for 90 s at 14,000 rpm in a microcentrifuge, and the supernatants were loaded onto 16-ml 10-30% linear sucrose gradients with a 1-ml 60% sucrose cushion. The gradients were spun at 150,000 × g for 7.5 h and separated into 30 0.58-ml fractions. The absorbance at 260 nm was used to measure the level of nucleic acid in each fraction, and a small portion of some fractions was also subjected to SDS-PAGE, to determine which fractions contained tailspike and ribosomal proteins.
RNase Digestion of Lysates-- To determine the role of RNA-containing structures, cell cultures were split in two. One-half was treated with 50 µg/ml RNase and 50 mM EDTA (final conditions) to destabilize and digest the ribosomal particles.
Electron Microscopy of Ribosome Particles-- Ribosome-containing fractions were examined using transmission electron microscopy. Fractions were dialyzed against buffer R to lower the sucrose concentration, applied to carbon-coated glow discharged 400 mesh copper grids, negatively stained with 3% uranyl acetate, blotted dry, and examined in the JEOL 1200 EX electron microscope.
Radiolabeling Nascent Polypeptide Chains-- To radiolabel the newly synthesized polypeptide chains, cell cultures were pulsed for 3 min or 90 s with 3.0 µCi/ml culture 14C-labeled amino acids (PerkinElmer Life Sciences) after 90 min of P22 infection. Cultures were then chased with a large volume of ice-cold unlabeled casamino acids, while simultaneously chilling the entire culture on ice, occasionally with the addition of buffer R ice cubes. Cultures were then centrifuged, resuspended, and flash-frozen as described above.
Competition ELISA Assay--
Mouse anti-tailspike monoclonal
antibodies were a generous gift from the lab of Dr. Michel Goldberg
(Institut Pasteur); for this study, monoclonal antibodies 175-3 and
51-2 were used. These antibodies have been shown to recognize epitopes
on the native tailspike trimer corresponding to a central region of the
-helix domain (
(
-helix)) and a portion of the
-coil domain
that includes Glu-405 (
(Glu-405)), respectively (Fig. 1)
(45).1 An ELISA competition assay was used to detect the
presence of tailspike epitopes; this was performed as described (49)
with the following modifications. Briefly, aliquots of purified
tailspike or refolding reaction were mixed with 0.04 µg/ml (final
concentration) anti-native tailspike monoclonal antibody diluted in
PBS/Tween (PBS buffer with 0.05% (w/v) Tween 20) and incubated
overnight at 4 °C. Mixtures were then loaded, in triplicate, into
wells of a microtiter plate coated with 1.0 µg/ml tailspike and
incubated at room temperature (22 °C). After 30 min, wells were
washed three times with PBS/Tween and incubated for 30 min with
alkaline phosphatase-labeled goat anti-mouse IgG (Southern
Biotechnology Associates). The plate was again washed three times,
incubated with p-nitrophenyl phosphate solution (2 mg/ml
p-nitrophenyl phosphate (Sigma) in 1 M
ethanolamine, 1 mM MgSO4, pH 9.8), and
monitored for absorbance at 405 nm. The amount of
absorbance corresponded to the amount of tailspike antibody not
bound to components of the gradient fraction. Control samples with no
antigen present permitted calculation of the percent of monoclonal
antibody recognizing tailspike antigen in the refolding reactions. It
should be noted that, under the conditions used here, the assay
operated under parameters that perturbed the equilibrium established in
the antigen/antibody mixtures, i.e. in some situations, more
than 10% of the unbound antibody adhered to the well of the microtiter
plate (50). Hence, data used to construct the standard curve shown in
Fig. 3 cannot be used to construct a Scatchard or Klotz plot for
determination of an accurate dissociation constant (50). For percent
recognition values >70%, percent recognition is much less sensitive
to additional amounts of native tailspike antigen present. Therefore,
in Figs. 8 and 9, percent recognition values >90% have been depicted
as a flat line at 100% recognition.
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Epitope Development during in Vitro Refolding--
At 10 °C,
tailspike refolds very slowly (t1/2 ~12 h) (32),
and intermediates populated during refolding can be trapped for many
hours by further chilling to 0 °C (37). Our assay for the
conformation of these intermediates used the binding of monoclonal
antibodies directed against tailspike epitopes (46). Since the
refolding tailspike polypeptide chains were transient intermediates
trapped by cold incubation, we were concerned about surface
denaturation of these intermediates during conventional enzyme-linked
immunosorbent assays. In such assays, the proteins are adsorbed to the
wells of a plastic microtiter plate. To circumvent this step, we used a
competition ELISA, previously developed as a method to detect
conformations of folding intermediates (51). This assay involves
incubating the antibody and antigen in solution, rather than attaching
the antigen to the wells of a plastic plate, thereby avoiding surface
binding or denaturation problems. This modification permits assaying
for conformation-sensitive epitopes under physiological conditions.
Excess, unbound antibody was then detected in a classical ELISA
measurement, which permitted calculation of the amount of antibody
bound to antigen in solution (51). Binding curves for both monoclonal
antibodies to native tailspike are shown in Fig.
3. Under the conditions used for this study, these assays were both sensitive and precise within the dynamic
range of 0-70% antibody recognition (see "Experimental Procedures" for details), and the experiments described below were
designed to stay within this range.
|
We used two different monoclonal antibodies, one that recognizes a
central portion of the -helix domain (
(
-helix)) (Fig. 1), and
the other that recognizes a portion of the
-helix domain located
just after the dorsal fin loop domain, containing residue Glu-405
(
(Glu-405)) (Fig. 1) (45, 46).1 Antibody binding to
in vitro tailspike refolding intermediates was performed
using the conditions of Goldberg and co-workers (46). As shown in
Fig 4, we detected
time-dependent changes in monoclonal antibody binding over
the course of the refolding reaction (24 h). For the
(
-helix)
monoclonal antibody, at the earliest refolding time points the presence
of the central
-helix epitope was minimal (Fig. 4). The binding of
this antibody to early intermediates of refolding tailspike was similar
to that reported by Friguet et al. (46) for three other
tailspike monoclonal antibodies. Binding of the
(
-helix) antibody
increases with time, reaching a maximum at the end of refolding.
|
For (Glu-405) antibody recognition of in vitro refolding
intermediates, we observed quite different results (Fig. 4). At the
earliest time points for refolding, there was significant antibody
recognition of the Glu-405-containing epitope. Indeed,
(Glu-405)
antibody recognition reached a maximum at time points shown previously
by Seckler and co-workers (52) to contain a large percentage of
protrimer intermediate, instead of native state. Recognition of
intermediates exceeded recognition of an equivalent concentration of
native, never unfolded tailspike (120 versus 100%
recognition), indicating that the
(Glu-405) antibody has more
affinity for these non-native tailspike chains than for the native
structure. In addition, whereas only ~75% of tailspike chains have
reached the native state after 24 h of refolding,
(Glu-405)
antibody binding was >75% of native tailspike binding at 24 h,
also demonstrating affinity of this antibody for the ~25% of
tailspike polypeptide chains adopting non-native conformations.
It is possible that binding of a monoclonal antibody could induce or
stabilize a conformation of the ensemble of refolding tailspike
polypeptide chains and therefore not accurately represent the tailspike
conformational distribution at a given time point during refolding.
However, the (
-helix) antibody did not bind to tailspike chains
at the earliest refolding time points, indicating that this antibody
was incapable of inducing structure via binding for the early tailspike
refolding intermediates. The
(Glu-405) antibody, which did bind to
early tailspike refolding intermediates, may possibly induce structure
formation; nevertheless, we did observe time-dependent
changes in antibody binding during refolding, indicating that induced
structure binding is not the only source of antibody recognition.
Preparation of Ribosome Structures-- At later stages of P22 infection of Salmonella, transcription and translation of host cell genes are inhibited. The gene 5 coat protein and the gene 9 tailspike protein are the two major proteins synthesized within infected cells. For these experiments, we used a phage strain carrying a nonsense mutation in gene 5. This prevents the synthesis of complete coat polypeptide chains and therefore the assembly of high molecular weight, rapidly sedimenting phage capsids. In addition, in these cells, the gene 9 tailspike polypeptide chain is the single major phage-encoded translation product.
To prepare ribosomes from P22-infected Salmonella cultures,
we took advantage of the natural lytic effects of phage infection, caused by the production of endogenous lysozyme (see "Experimental Procedures" for details). Infected cells were incubated long enough to synthesize late-stage phage proteins. The infected cells were concentrated by centrifugation and then lysed by a single freeze/thaw cycle, allowing lysozyme to escape the membrane barrier and digest bacterial cell walls. The lysate supernatant was fractionated by
sedimentation through a linear 10-30% sucrose gradient. All manipulations were performed on ice or at 4 °C, in order to trap and
stabilize transient tailspike folding intermediates (34, 37, 53).
Sedimentation efficiently separated ribosomal structures from other
cellular components. As shown in Fig.
5a, absorbance peaks corresponding to nucleic acids were
separated from the slowly sedimenting nucleic acids (tRNAs, degraded
mRNAs, DNA fragments). The sharpest peaks correspond to 50 S and
30 S ribosomal subunits. The 70 S position, where intact ribosomes
would be expected, corresponds to the leading shoulder seen in
fractions 16-19. The absorbing species that sedimented faster than
ribosomes were presumably polysomes. Digesting the lysates with RNase
prior to sedimentation disrupted the larger 260 nm absorbing
structures, consistent with their identification as ribosomes and
ribosomal subunits (Fig. 5b).
|
We examined more carefully the structures sedimenting through the sucrose gradients by analyzing fractions by SDS-PAGE and detecting host and phage proteins by silver staining. Fig. 5c shows the distribution of proteins within several representative fractions as follows: top fractions contained many soluble proteins; a band corresponding to the 60-kDa protein of GroEL appeared at the expected sedimentation coefficient of 21 S; and low molecular weight bands corresponding to the individual ribosomal proteins appeared at 30, 50, and 70 S. A band corresponding to the full-length SDS-resistant tailspike trimer appeared near the top of the gradient, where native tailspike is known to sediment (54). In addition, a species corresponding to the apparent molecular weight of SDS-denatured tailspike chains was found throughout the ribosome-containing fractions, including those expected to contain polyribosomes. The mobility of these chains indicates that they had not yet reached the SDS-resistant native state at the time of cell lysis.
Fig. 5d shows the distribution of proteins through a sucrose gradient from a lysate digested with RNase prior to centrifugation. The small ribosomal proteins were absent from the middle of the gradient, confirming that the ribosomal subunits had been dissociated by the treatment. In addition, the band corresponding to tailspike chains was also absent from these fractions.
To confirm that the central sucrose gradient fractions contain intact
ribosome particles, electron micrographs (Fig.
6a) were taken of negatively
stained aliquots from ribosome fractions expected to contain small
polysomal particles (indicated by the arrow in Fig.
5a). These images show the presence of intact 70 S
ribosomes, and also many small polysomes. There are few other species
present in the image fields, demonstrating the efficacy of the sucrose gradient preparation procedure. Confirming that RNase digestion destroys intact ribosomal structures, a corresponding fraction from an
RNase-digested lysate (see arrow in Fig. 5b)
shows no ribosome-like structures but only an occasional phage particle from the initial infection (Fig. 6b).
|
Localization of Newly Synthesized Tailspike Polypeptides with Ribosomes-- As noted above, the ribosome-associated tailspike band did not migrate as native trimer in the unheated SDS gel (Fig. 5c), indicating that these tailspike chains had not yet reached the native state at the time of cell lysis. The tailspike polypeptide chains found in the ribosomal fractions could therefore represent one or more non-native states of tailspike as follows: newly synthesized chains associated with the ribosomes, released chains interacting nonspecifically with ribosomes, or other rapidly sedimenting forms of the tailspike, such as intracellular aggregates or inclusion body precursors (38).
To determine whether the tailspike chains present in the ribosome
fractions were newly synthesized or arising from aggregates of
increasing size, newly synthesized polypeptides were radiolabeled by
pulsing cultures for 3 min with 14C-labeled amino acids.
Cultures were then chased with cold casamino acids and immediately
cooled on ice, lysed, and separated through sucrose gradients;
fractions from these gradients were separated on SDS-polyacrylamide
gels. Quantitation of the phosphorimage from these gels shows that the
tailspike chains in the ribosome fractions were labeled during the
pulse (Fig. 7a). A significant fraction of the label was in chains corresponding to full-length tailspike chains. These presumably correspond to newly synthesized chains, completed or almost completed, but not yet released from the
ribosome. Full-length newly synthesized chains were also detected with
shorter (90 s) pulses. Partial translation products may be present but
would not be expected to form discrete electrophoretic species. This
distribution of translated chains in vivo differs somewhat
from the results found for many proteins labeled using in
vitro translation systems, where partial translation products represent a much larger fraction of the translated chains, and the rate
of translation is much slower (13, 55).
|
As would be expected for nascent chains, the tailspike polypeptide chains no longer sedimented in the ribosome fractions when the ribosomes were digested with RNase (Fig. 7a). Longer chase times (20 min) also resulted in loss of the radiolabeled tailspike chains from the ribosome fractions (data not shown), indicating that the radiolabeled tailspike polypeptide chains were indeed newly synthesized and chased off of the ribosomes. Fig. 7a shows that the fractions containing the largest amounts of ribosome-bound tailspike chains were associated with the 30 S and small polysome fractions. The association with the 30 S subunits was unexpected, given that the exit site for newly synthesized polypeptide chains has been localized on the 50 S subunit (21, 26).
It is important to consider whether the ribosome-bound tailspike intermediate is a major populated (possibly obligate) folding intermediate or only a minor intermediate populated in a parallel pathway to native trimer formation. We detected the ribosome-bound chains using routine silver staining (Fig. 5c). Quantitation of the ribosome-associated tailspike bands indicates that 10-20% of all intermediate-conformation tailspike chains are located in ribosome-containing fractions, and this localization is RNase-sensitive (Fig. 5d). This result suggests that these chains do represent a major component of maturing chains in vivo.
Newly synthesized tailspike chains can proceed through two well defined pathways as follows: (a) the productive pathway, through the protrimer to the native trimer, or (b) the inclusion body pathway, through higher order multimeric species to the aggregated state (38, 56). Under the growth conditions used for these experiments, >90% of newly synthesized tailspike chains mature to native trimer (34, 60). In the experiment shown in Fig. 7c, we quantitated the amount of radiolabeled native tailspike trimer present before and after RNase digestion of the cell lysate. Native trimer was well resolved in the native gels from partially folded or assembled intermediates (Fig. 5c) (57). The amount of radiolabeled trimer present near the top of the gradient increased upon RNase digestion. Presumably, these additional radiolabeled trimers arose from ribosome-associated tailspike chains released by RNase digestion, which were then able to associate into native trimers. Our results therefore indicate that 1) a significant fraction of newly synthesized tailspike chains populate the ribosome-bound intermediate, and 2) the ribosome-bound polypeptide chains were capable of maturing to the native trimer structure.
The other major phage-encoded structural protein synthesized during late-stage infection is the 22-kDa amber fragment of the coat (gene 5) protein (58). This amber fragment is unable to assemble into phage capsids and aggregates within cells.3 The 14C-labeled amber fragment is present throughout the gradient, presumably representing the aggregating species (Fig. 7b). Although digestion of the lysate with RNase eliminated the tailspike band present in the ribosome fractions, it had no affect on the coat protein amber fragment band (Fig. 7b). Similarly, the distribution of radiolabeled GroEL throughout the gradient was the same for both undigested and RNase-digested lysates (Fig. 7c).
Probing the Conformation of Newly Synthesized Tailspike Chains with Antibodies-- The newly synthesized tailspike polypeptide chains found associated with the ribosomes were at or near their complete length of 666 amino acids. Since the ribosomal exit tunnel for newly synthesized polypeptides protects only 30-40 residues, most of the tailspike polypeptide chain must be outside the ribosome. To investigate the conformation of the polypeptide chain during the ribosome-associated stage of maturation, we determined the binding of the two monoclonal antibodies described above.
Fig. 8a shows the results of
assaying gradient fractions for the presence of epitopes recognized by
the (
-helix) antibody. Top fractions showed high levels of
antibody binding (>90%), saturating the sensitivity range for this
assay (i.e. >70% antibody recognition; see "Experimental
Procedures" for details). The antigen recognized in these fractions
presumably represents the accumulation of native tailspike trimer. In
addition, there is a significant level (with respect to the in
vitro reaction) of tailspike
-helix epitopes detected in the
ribosome-containing region of the gradient, including fractions
corresponding to small polyribosomes.
|
ELISA competition assays with the (
-helix) antibody were also
performed on samples that were digested with RNase (Fig.
8b). As expected, the large amount of free tailspike antigen
at the top of the gradient remained. The antigen levels in the polysome regions were reduced, as expected for nascent chains, but not eliminated. Subtracting the RNase-digested background binding from the
antibody binding to the undigested lysate (Fig. 8c) yielded a positive peak centered over the 70 S small polysome position and a
negative peak centered over the 30 S position. The positive peak
confirms the presence of RNase-sensitive antibody binding in these fractions.
The development of the conformational epitope in vivo is in direct contrast to the in vitro results shown in Fig. 3, where we observed negligible epitope development immediately after dilution of tailspike chains from denaturant (Fig. 4). We do not yet know if these tailspike chains represent a single ribosome-bound conformation or an ensemble of intermediate states associated with the ribosome. What is clear, however, is that these newly synthesized, ribosome-bound tailspike chains adopt conformations not significantly populated by the early in vitro refolding intermediates.
The negative peak, although small, appeared consistently through
multiple repetitions of the assay and corresponds to a significant difference in antibody recognition for 3-4 separate fractions. The
negative peak indicates that there was increased antibody binding to
30 S fractions for the RNase-digested lysate versus the
untreated lysate. The increase in antibody binding did not arise from
RNase digestion of cellular components, as a control infection with a
P22 variant containing a nonsense mutant in the tailspike gene
9 (see "Experimental Procedures" for details) exhibited no increase in antibody binding (data not shown). From the absorbance at 260 nm, SDS-polyacrylamide gel analysis, and pulse-chase
experiments, it is clear that there were fewer 30 S subunits and newly
synthesized tailspike chains present in these fractions after RNase
digestion. Since these particles were not further dissociated, they may
represent a class of 30 S-tailspike polypeptide complexes that were
refractory to the RNase digestion. The increase in antigenicity
suggests that the small population of 30 S subunits present after
RNase digestion contained structured tailspike epitopes that were
better recognized by the (
-helix) monoclonal antibody. Our
results are unable to distinguish whether the additional antibody
binding was due to increased affinity between the antibody and antigen (for example, due to a rearrangement of the epitope to a conformation with more native-like structure) or increased accessibility of the
antibody to the antigen epitope due to the RNase digestion procedure.
The background binding seen in the middle to lower gradient fractions
for the RNase-digested lysate (Fig. 8b) may represent nonspecific interactions with host cell components or with other forms
of the tailspike chains. This region of the gradient contains the
following two classes of tailspike chains that would not be sensitive
to the RNase digestion: (a) input phage particles from the
initial infection whose tailspike chains are assembled onto the phage
particle, a form which is highly antigenic (59), and (b)
aggregated tailspike chains that are intermediates to the inclusion
body state (56). In fact, the (
-helix) antibody used in this
assay was not reactive with multimeric tailspike aggregation
intermediates (60). In addition, we observed intact phage particles in
electron micrographs of RNase-digested gradient fractions (Fig. 6),
suggesting residual phage particles from the initial infection are the
cause of the background binding.
The conformations of the tailspike nascent chains were also probed with
the (Glu-405) monoclonal antibody (Fig. 1). Binding of this antibody
to lysate gradient fractions yielded results similar to the binding of
the
(
-helix) antibody. Antibody binding corresponded to the
distribution of native trimer in top gradient fractions (again,
overloading the sensitivity range of the binding assay) and fractions
containing 70 S ribosomal and polysomal particles (Fig.
9). As the
(Glu-405) antibody also
recognized the earliest in vitro refolding intermediates
(Fig. 4), it is perhaps not surprising that newly synthesized,
ribosome-associated tailspike chains are also recognized.
|
As a control for antibody binding to components other than newly
synthesized tailspike chains, we also infected cells with a phage
containing a nonsense mutant in the tailspike gene 9 (see "Experimental Procedures" for details). The lysate from this
infection was fractionated through a sucrose gradient and assayed for
(Glu-405) antibody binding. There was a significant reduction in the
level of antibody binding throughout the gradient for this lysate (Fig. 9), as would be expected for an antibody detecting only tailspike folding intermediates and native trimer in the wild type lysate. Background binding presumably represents binding to residual input phage and nonspecific association with other cellular components.
The specific tailspike structural epitopes detected on the newly
synthesized polypeptides represent conformations developed while the
chains were still associated with ribosome structures. By virtue of
their ribosome-bound state, these polypeptide chains by definition
represent an earlier state of the polypeptide chain maturation than for
released chains isolated from the cytoplasm. These conformations were
clearly different than those for the early intermediates formed during
in vitro refolding.
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The tailspike polypeptide chains synthesized by phage-infected Salmonella cells in these experiments were translated, folded, and assembled at 30 °C. Under these conditions, there was no significant loss to aggregation (Fig. 5d, and data not shown), and the majority of the polypeptide chains reached the native state. However, the majority of tailspike polypeptides refolding from denaturant at 30 °C are unable to reach the native state and instead formed inactive high molecular weight aggregates (56), even though these chains refold efficiently in vitro at lower temperatures (4). Despite systematic searches, no clear evidence has been found for a role for prokaryotic molecular chaperones in tailspike folding in vivo (43, 44). An alternative hypothesis is that the translational machinery itself participates in the early stages of chain folding.
The ribosomes from P22-infected Salmonella were cold-trapped and recovered in association with newly synthesized tailspike polypeptide chains. Chilling in vivo folding or in vitro refolding tailspike has been shown previously to trap the transient conformations of folding intermediates (32, 34, 37), due to the unusually strong temperature dependence of tailspike folding (39). Surprisingly, the ribosome-associated tailspike polypeptides recovered from the pulse-labeling experiments were full-length, within the accuracy of the SDS gel fractionation (Fig. 5c). This indicates that while translation of the tailspike polypeptide may proceed quickly (~20 amino acids/s for bacterial cells), the tailspike polypeptide chain then spends the first several minutes of its existence in close association with the ribosome. These tailspike chains were not "dead ends," as pulse-chase experiments showed maturation of the labeled chains to native tailspike (data not shown), and separation from the ribosome after RNase digestion resulted in increased yields of native trimer (Fig. 7c). The experiments reported here do not distinguish between chains still tethered due to slow termination or terminated chains still bound to the ribosome. In either case, we would expect more than 600 amino acids of the newly synthesized chains to have emerged from the exit site but remain at the ribosome surface.
Conformations of Nascent Tailspike Polypeptide Chains--
The
newly synthesized, full-length tailspike polypeptide chains isolated on
Salmonella ribosomes provided an opportunity to study the
conformation of a transiently cold-trapped nascent polypeptide chain
from a physiologically relevant translation system. Two monoclonal
antibodies to tailspike, each of which recognizes a distinct structural
epitope in the parallel -helix of the native protein,1
specifically recognized the ensemble of ribosome-associated tailspike polypeptide chains (Figs. 8 and 9). This indicates that the epitopes recognized by these antibodies were already present on the newly synthesized tailspike chains. Yet during in vitro refolding
experiments, the
(
-helix) epitope was not formed at early
refolding times. Thus the ribosome-bound tailspike chains appear to
adopt a conformation with structure in the central region
-helix
domain, not detected for early in vitro refolding species.
The simplest explanation for the more structured conformations of newly synthesized ribosome-bound tailspike chains (versus refolding intermediates in vitro) is that the nascent tailspike polypeptide chains can achieve a more structured conformation through direct interactions with sites on the ribosome and/or because of the vectorial nature of polypeptide synthesis in vivo. We are currently performing experiments to fully characterize the tailspike-ribosome interaction and to determine its effect on tailspike chain folding.
What fraction of the newly synthesized, ribosome-bound tailspike
polypeptide chains adopt the recognizable -helix epitope? If one
assumes that the affinity of the
(
-helix) antibody for the
ribosome-bound, transient intermediate is equal to the affinity for
native tailspike, then the concentration of epitope-containing chains
in these fractions was ~10 pM (Fig. 3). This
concentration of tailspike chains was greater than the concentration of
radiolabeled, ribosome-bound tailspike chains observed in the
pulse-labeling experiments (~1 pM; Fig. 7; comparison to
radiolabeled tailspike standards). An ~10-fold higher concentration
of epitope versus radiolabeled chains indicates that,
indeed, there were significant differences between the affinity of the
(
-helix) antibody for the ribosome-bound, transient conformation,
and native tailspike. Without an accurate measure of the affinity of
this antibody for the intermediate conformation, we are currently
unable to determine accurately the fraction of radiolabeled chains that
adopt the
-helix epitope. Adoption of the
-helix epitope
conformation prior to ribosome release therefore may not be an obligate
intermediate in the tailspike folding pathway in vivo;
however, these results are consistent with a model where the majority
of the radiolabeled tailspike chains adopt the
-helix epitope prior
to dissociation from the ribosome.
In addition, it appears that the ribosome-associated conformation
persists in tailspike chains isolated in association with the 30 S
ribosomal subunit (Fig. 8), suggesting that an area of the ribosome
possibly influencing nascent polypeptide structure could be localized
at least in part on the 30 S subunit (Fig. 10). The persistence of detectable
tailspike conformations on 30 S subunits provides evidence that the
observed tailspike conformations do not arise due to ribosome-bound
chaperone interactions, because the ribosome-associated chaperone
trigger factor has been shown to interact only with the 50 S subunit
(61).
|
The increase in tailspike monoclonal antibody binding to the RNase-digested 30 S fractions occurred despite the destruction of >90% of the 30 S particles by RNase and the corresponding loss of the majority of associated, newly synthesized tailspike chains (Figs. 5 and 7a). Therefore, the increase in observed antigenicity arose from a very small fraction of remaining tailspike chains, suggesting that digestion with RNase may reveal a class of 30 S-tailspike complexes that contains more reactive tailspike antibody epitopes. Possibly, digestion with RNase alters the structure of some tailspike-containing 30 S subunits in a way that also alters the conformation of the tailspike chain, increases its accessibility to the antibody, or both. The altered tailspike conformation may then protect the remaining 30 S subunits from further RNase digestion.
These findings have interesting implications for the folding of
tailspike chains in vivo. Recognition of the newly
synthesized tailspike chains by (
-helix) antibodies suggests that
the
-helix may form early during in vivo folding. This
correlates well with results from in vitro refolding
experiments, which indicate organization of the
-helix domain occurs
early during refolding (32), and in fact, a sequence corresponding to
the
-helix domain alone can adopt a
-helix structure in
vitro (52). Formation of rungs of the
-helix domain while the
polypeptide chain is still attached to the ribosome could provide
crucial chain organization that predisposes the released chains for
folding to the native structure, instead of aggregating. Association of
newly synthesized tailspike chains with a specific site or sites on the
ribosome, possibly located on the 30 S subunit, could prime the
polypeptide chains for further steps of folding and assembly, in
particular the formation of a conformation capable of efficient trimer
formation and C-terminal strand interdigitation (Fig. 1). Certainly the
tailspike folding intermediates trapped on the ribosome (the earliest
in vivo folding intermediates trapped for tailspike) appear
to have additional structural development versus that seen
for the earliest in vitro refolding intermediates (Fig.
4).
Roles of the Ribosome in Nascent Chain Maturation-- The presence of full-length, newly synthesized polypeptide chains on the ribosome, even after cell lysis and sucrose gradient centrifugation, suggests that these chains were relatively tightly bound by the ribosome. Such associations have not been described previously. Our experiments suggest that this association is a physiological complex, as these chains do eventually mature to native tailspike trimers. We are currently performing experiments to determine if the ribosome itself contains sites that recognize and bind partially folded polypeptide chains. Such interactions with the ribosome may protect the newly synthesized polypeptide from degradation or unproductive interactions, without relying on additional host cell systems, such as the chaperone networks (62).
As noted above, when ribosomal particles were disrupted by normal lysate manipulations or RNase digestion, significant amounts of full-length, newly synthesized tailspike polypeptide chain remained preferentially associated with remaining 30 S particles (Fig. 8c). This was surprising, given the accepted view that newly synthesized polypeptide chains emerge from the ribosome via an exit site on the back side of the 50 S subunit. One interpretation of this result is that upon dissociation of the active ribosomes into 30 S and 50 S subunits, a population of nascent tailspike polypeptides bound to 70 S ribosomes remain preferentially associated with the 30 S subunits, presumably through specific interactions with the 30 S subunit or some intermediary (Fig. 10). In the intact ribosome, wrapping around to reach the 30 S subunit clearly requires extension of the polypeptide chain beyond the ~30 amino acids necessary to reach the outer surface of the ribosome. Since the isolated nascent chains are full-length (more than 200 nm if extended), the surface of the 30 S subunit would be readily accessible to the nascent chain.
The adoption of structured conformations in a nascent polypeptide chain
reported here might play a general role in successful protein folding
in vivo. While tailspike polypeptide chains dwell in
association with the ribosome for an unusually long time (possibly for
reasons due to viral infection; see above), moderate slowing of
translation, termination, and/or polypeptide chain release may permit
the development of conformational preferences for a range of newly
synthesized chains. For the large fraction of polypeptide chains that
have no interaction with molecular chaperones during chain maturation
and folding, interactions with specific sites on the ribosome and/or
within the nascent chain itself could provide early folding assistance.
Early ribosome-bound folding may be particularly useful during the
folding of elongated protein structures that need protection from the
cellular environment not provided by the nascent chain folding back on
itself. Finally, formation of a partially native nascent polypeptide
chain could be of particular importance for the successful folding and
assembly of multimeric protein complexes, encouraging the development
of structured, assembly-competent monomer chain conformations prior to
nascent chain release and association.
![]() |
ACKNOWLEDGEMENTS |
---|
We thank Cameron Haase-Pettingell for technical advice and assistance. We also thank Stephen Raso for reviewing the manuscript and Michel Goldberg and Lisa Djavadi-Ohaniance (Institut Pasteur) for invaluable assistance with the ELISA competition assay and generosity in supplying the tailspike monoclonal antibodies. In addition, we thank Madhulika Jain for help with the in vitro refolding experiments and Patricia Reilly and the MIT Biomedical Electron Microscopy Laboratory (recipient of National Institutes of Health Shared Instrumentation Grant S10 RR05734) for assistance with the electron microscope.
![]() |
FOOTNOTES |
---|
* This work was supported in part by Grant GM17980 from the National Institutes of Health.The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Supported by Postdoctoral Fellowship GM19715 from the National
Institutes of Health. To whom correspondence should be addressed: Dept.
of Biology 68-340D, Massachusetts Institute of Technology, Cambridge,
MA 02139. Tel.: 617-253-3545; Fax: 617-252-1843; E-mail: plclark@mit.edu.
Published, JBC Papers in Press, April 23, 2001, DOI 10.1074/jbc.M008490200
1 M. Jain, J. King, and P. L. Clark, manuscript in preparation.
3 C. A. Haase-Pettingell and J. King, unpublished results.
![]() |
ABBREVIATIONS |
---|
The abbreviations used are: PAGE, polyacrylamide gel electrophoresis; ELISA, enzyme-linked immunosorbent assay; PBS, phosphate-buffered saline.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. | Anfinsen, C. B. (1973) Science 181, 223-230[Medline] [Order article via Infotrieve] |
2. |
Lorimer, G. H.
(1996)
FASEB J.
10,
5-9 |
3. | Fedorov, A. N., and Baldwin, T. O. (1999) J. Mol. Biol. 294, 579-586[CrossRef][Medline] [Order article via Infotrieve] |
4. |
Seckler, R.,
Fuchs, A.,
King, J.,
and Jaenicke, R.
(1989)
J. Biol. Chem.
264,
11750-11753 |
5. | Skowyra, D., Georgopoulos, C., and Zylicz, M. (1990) Cell 62, 939-944[Medline] [Order article via Infotrieve] |
6. | Langer, T., Lu, C., Echols, H., Flanagan, J., Hayer, M. K., and Hartl, F. U. (1992) Nature 356, 683-689[CrossRef][Medline] [Order article via Infotrieve] |
7. | Rospert, S., Looser, R., Dubaquie, Y., Matouschek, A., Glick, B. S., and Schatz, G. (1996) EMBO J. 15, 764-774[Abstract] |
8. | Nicola, A. V., Chen, W., and Helenius, A. (1999) Nat. Cell Biol. 1, 341-345[CrossRef][Medline] [Order article via Infotrieve] |
9. |
Wickner, S.,
Maurizi, M. R.,
and Gottesman, S.
(1999)
Science
286,
1888-1893 |
10. | Feldman, D. E., and Frydman, J. (2000) Curr. Opin. Struct. Biol. 10, 26-33[CrossRef][Medline] [Order article via Infotrieve] |
11. | Gilmore, R., Coffey, M. C., Leone, G., McLure, K., and Lee, P. W. (1996) EMBO J. 15, 2651-2658[Abstract] |
12. |
Komar, A. A.,
Kommer, A.,
Krasheninnikov, I. A.,
and Spirin, A. S.
(1997)
J. Biol. Chem.
272,
10646-10651 |
13. | Netzer, W. J., and Hartl, F. U. (1997) Nature 388, 343-349[CrossRef][Medline] [Order article via Infotrieve] |
14. | Miller, S., Schuler, B., and Seckler, R. (1998) 37, 9160-9168 |
15. | Neira, J. L., and Fersht, A. R. (1999) J. Mol. Biol. 285, 1309-1333[CrossRef][Medline] [Order article via Infotrieve] |
16. | Deuerling, E., Schulze-Specking, A., Tomoyasu, T., Mogk, A., and Bukau, B. (1999) Nature 400, 693-696[CrossRef][Medline] [Order article via Infotrieve] |
17. | Teter, S. A., Houry, W. A., Ang, D., Tradler, T., Rockabrand, D., Fischer, G., Blum, P., Georgopoulos, C., and Hartl, F. U. (1999) Cell 97, 755-765[Medline] [Order article via Infotrieve] |
18. |
Caldas, T.,
Laalami, S.,
and Richarme, G.
(2000)
J. Biol. Chem.
275,
855-860 |
19. | Kudlicki, W., Coffman, A., Kramer, G., and Hardesty, B. (1997) Folding Des. 2, 101-108[Medline] [Order article via Infotrieve] |
20. |
Pal, S.,
Chandra, S.,
Chowdhury, S.,
Sarkar, D.,
Ghosh, A. N.,
and Gupta, C. D.
(1999)
J. Biol. Chem.
274,
32771-32777 |
21. |
Cate, J. H.,
Yusupov, M. M.,
Yusupova, G. Z.,
Earnest, T. N.,
and Noller, H. F.
(1999)
Science
285,
2095-2104 |
22. |
Ban, N.,
Nissen, P.,
Hansen, J.,
Moore, P. B.,
and Steitz, T. A.
(2000)
Science
289,
905-920 |
23. |
Nissen, P.,
Hansen, J.,
Ban, N.,
Moore, P. B.,
and Steitz, T. A.
(2000)
Science
289,
920-930 |
24. | Frydman, J., Nimmesgern, E., Ohtsuka, K., and Hartl, F. U. (1994) Nature 370, 111-117[CrossRef][Medline] [Order article via Infotrieve] |
25. |
Pfund, C.,
Lopez-Hoyo, N.,
Ziegelhoffer, T.,
Schilke, B. A.,
Lopez-Buesa, P.,
Walter, W. A.,
Wiedmann, M.,
and Craig, E. A.
(1998)
EMBO J.
17,
3981-3989 |
26. | Bernabeu, C., and Lake, J. A. (1982) Proc. Natl. Acad. Sci. U. S. A. 79, 3111-3115[Abstract] |
27. | Frydman, J., Erdjument-Bromage, H., Tempst, P., and Hartl, F. U. (1999) Nat. Struct. Biol. 6, 697-705[CrossRef][Medline] [Order article via Infotrieve] |
28. | Kudlicki, W., Odom, O. W., Kramer, G., and Hardesty, B. (1994) J. Mol. Biol. 244, 319-331[CrossRef][Medline] [Order article via Infotrieve] |
29. | Steinbacher, S., Seckler, R., Miller, S., Steipe, B., Huber, R., and Reinemer, P. (1994) Science 265, 383-386[Medline] [Order article via Infotrieve] |
30. | Steinbacher, S., Miller, S., Baxa, U., Budisa, N., Weintraub, A., Seckler, R., and Huber, R. (1997) J. Mol. Biol. 267, 865-880[CrossRef][Medline] [Order article via Infotrieve] |
31. | Kreisberg, J. F., Betts, S. D., and King, J. (2000) Protein Sci. 9, 2338-2343[Abstract] |
32. | Fuchs, A., Seiderer, C., and Seckler, R. (1991) Biochemistry 30, 6598-6604[Medline] [Order article via Infotrieve] |
33. |
Goldenberg, D.,
Berget, P.,
and King, J.
(1982)
J. Biol. Chem.
257,
7864-7871 |
34. | Goldenberg, D., and King, J. (1982) Proc. Natl. Acad. Sci. U. S. A. 79, 3403-3407[Abstract] |
35. |
Yu, M. H.,
and King, J.
(1988)
J. Biol. Chem.
263,
1424-1431 |
36. | King, J., Fane, B., Haase-Pettingell, C., Mitraki, A., Villafane, R., and Yu, M.-H. (1990) in Protein Folding: Deciphering the Second Half of the Genetic Code (Gierasch, L. M. , and King, J., eds) , pp. 225-239, American Association for the Advancement of Science, Washington, D. C. |
37. |
Betts, S. D.,
and King, J.
(1998)
Protein Sci.
7,
1516-1523 |
38. |
Haase-Pettingell, C. A.,
and King, J.
(1988)
J. Biol. Chem.
263,
4977-4983 |
39. |
Danner, M.,
and Seckler, R.
(1993)
Protein Sci.
2,
1869-1881 |
40. | Chan, H. S., and Dill, K. A. (1998) Proteins Struct. Funct. Genet. 30, 2-33[CrossRef][Medline] [Order article via Infotrieve] |
41. | Karplus, M. (1997) Folding Des. 2, S69-S75[Medline] [Order article via Infotrieve] |
42. |
Brunschier, R.,
Danner, M.,
and Seckler, R.
(1993)
J. Biol. Chem.
268,
2767-2772 |
43. |
Gordon, C. L.,
Sather, S. K.,
Casjens, S.,
and King, J.
(1994)
J. Biol. Chem.
269,
27941-27951 |
44. |
Sather, S. K.,
and King, J.
(1994)
J. Biol. Chem.
269,
25268-25276 |
45. |
Friguet, B.,
Djavadi-Ohaniance, L.,
Haase-Pettingell, C. A.,
King, J.,
and Goldberg, M. E.
(1990)
J. Biol. Chem.
265,
10347-10351 |
46. |
Friguet, B.,
Djavadi-Ohaniance, L.,
King, J.,
and Goldberg, M. E.
(1994)
J. Biol. Chem.
269,
15945-15949 |
47. | King, J., and Yu, M.-H. (1986) Methods Enzymol. 131, 250-266[Medline] [Order article via Infotrieve] |
48. | Sauer, R. T., Krovatin, W., Poteete, A. R., and Berget, P. B. (1982) 21, 5811-5815 |
49. | Friguet, B., Djavadi-Ohaniance, L., and Goldberg, M. E. (1989) in Protein Structure: A Practical Approach (Creighton, T. E., ed) , pp. 287-310, Oxford University Press, Oxford, UK |
50. | Friguet, B., Chaffotte, A. F., Djavadi-Ohaniance, L., and Goldberg, M. E. (1985) J. Immunol. Methods 77, 305-319[CrossRef][Medline] [Order article via Infotrieve] |
51. | Friguet, B., Djavadi-Ohaniance, L., and Goldberg, M. E. (1984) Mol. Immunol. 21, 673-677[CrossRef][Medline] [Order article via Infotrieve] |
52. | Danner, M., Fuchs, A., Miller, S., and Seckler, R. (1993) Eur. J. Biochem. 215, 653-661[Abstract] |
53. | Speed, M. A., Wang, D. I. C., and King, J. (1996) Nat. Biotech. 14, 1283-1287[Medline] [Order article via Infotrieve] |
54. | Bazinet, C., Benbasat, J., King, J., Carazo, J. M., and Carrascosa, J. L. (1988) Biochemistry 27, 1849-1856[Medline] [Order article via Infotrieve] |
55. |
Fulton, A. B.,
and L'Ecuyer, T.
(1993)
J. Cell Sci.
105,
867-871 |
56. |
Speed, M. A.,
Wang, D. I.,
and King, J.
(1995)
Protein Sci.
4,
900-908 |
57. | Goldenberg, D. P., Smith, D. H., and King, J. (1983) Proc. Natl. Acad. Sci. U. S. A. 80, 7060-7064[Abstract] |
58. | Botstein, D., Waddell, C. H., and King, J. (1973) J. Mol. Biol. 80, 669-695[Medline] [Order article via Infotrieve] |
59. | Shea, R. G. (1977) Ph.D. thesis , Massachusetts Institute of Technology, Cambridge |
60. |
Speed, M. A.,
Morshead, T.,
Wang, D. I.,
and King, J.
(1997)
Protein Sci.
6,
99-108 |
61. | Lill, R., Crooke, E., Guthrie, B., and Wickner, W. (1988) Cell 54, 1013-1018[Medline] [Order article via Infotrieve] |
62. | Campbell, A. M. (1996) in Escherichia coli and Salmonella (Neidhardt, F. C. , Curtiss, R. , Ingraham, J. L. , Lin, E. C. C. , Low, K. B. , Magasanik, B. , Reznikoff, W. S. , Riley, M. , Schaechter, M. , and Umbarger, H. E., eds), 2nd Ed. , pp. 2325-2338, ASM Press, Washington, D. C. |
63. |
King, J.,
Haase-Pettingell, C.,
Robinson, A. S.,
Speed, M.,
and Mitraki, A.
(1996)
FASEB J.
10,
57-66 |
64. | Goldenberg, D. P. (1989) in Protein Structure: A Practical Approach (Creighton, T. E., ed) , pp. 225-250, IRL Press, Oxford University Press |