From the Department of Biological Sciences, Columbia
University, New York, New York 10027 and the ¶ Departments of
Biochemistry & Molecular Biology and Chemistry, University of
Massachusetts, Amherst, Massachusetts 01003
Received for publication, June 17, 2002, and in revised form, October 25, 2002
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ABSTRACT |
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The SecA ATPase drives the processive
translocation of the N terminus of secreted proteins through the
cytoplasmic membrane in eubacteria via cycles of binding and release
from the SecYEG translocon coupled to ATP turnover. SecA forms a
physiological dimer with a dissociation constant that has previously
been shown to vary with temperature and ionic strength. We now present
data showing that the oligomeric state of SecA in solution is altered by ligands that it interacts with during protein translocation. Analytical ultracentrifugation, chemical cross-linking, and
fluorescence anisotropy measurements show that the physiological dimer
of SecA is monomerized by long-chain phospholipid analogues. Addition of wild-type but not mutant signal sequence peptide to these SecA monomers redimerizes the protein. Physiological dimers of SecA do not
change their oligomeric state when they bind signal sequence peptide in
the compact, low temperature conformational state but polymerize when
they bind the peptide in the domain-dissociated, high-temperature
conformational state that interacts with SecYEG. This last result shows
that, at least under some conditions, signal peptide interactions drive
formation of new intermolecular contacts distinct from those
stabilizing the physiological dimer. The observations that signal
peptides promote conformationally specific oligomerization of SecA
while phospholipids promote subunit dissociation suggest that the
oligomeric state of SecA could change dynamically during the protein
translocation reaction. Cycles of SecA subunit recruitment and
dissociation could potentially be employed to achieve processivity in
polypeptide transport.
The SecA translocation ATPase mediates preprotein translocation
through the cytoplasmic membrane of eubacteria via cycles of binding
and release from the SecYEG translocon (1-5) coupled to its own ATPase
cycle (6-8) (reviewed in Ref. 9). Although the default translocation
pathway is believed to involve an initial phase powered by the ATPase
activity of SecA and a later phase powered by a transmembrane
electrochemical potential coupled to SecYEG, the ATPase activity of
SecA can mediate the translocation of an entire preprotein in the
absence of a transmembrane potential (7). This observation has led to
the conclusion that the SecA ATPase can mediate the processive
translocation of polypeptide chains (10).
Each cycle of ATP binding and hydrolysis by SecA is believed to result
in the translocation of about 40 residues of preprotein (7, 11). There
is substantial evidence that SecA interacts with both the N-terminal
signal sequence (6, 12-21) that targets preproteins for export from
the cytoplasm as well as the mature region of the preprotein (22, 23).
These binding interactions presumably allow SecA to transfer
polypeptide segments to SecYEG. However, the details of the preprotein
binding and transfer reactions are not understood, so there is little
information on how processivity is achieved in preprotein translocation.
Achieving efficient processive translocation is likely to involve
complex interactions between SecA and the preprotein. Experimental evidence for such complexity comes from studies on the interaction of
synthetic signal sequence peptides with SecA in different states. The
binding of such peptides inhibits the ATPase activity of a 64-kDa
N-terminal fragment of SecA with elevated basal activity (19, 21) but
stimulates the lipid-activated ATPase activity of intact SecA (17, 18).
The different functional consequences of signal peptide interactions in
these two assays suggest that SecA interacts with preproteins
differently at different stages of its ATP-driven conformational
reaction cycle.
Acidic phospholipids are required for efficient SecA-mediated
preprotein translocation both in vivo (24) and in
vitro (25, 26). SecA inserts into phospholipid monolayers in a
reaction that is enhanced by the presence of acidic phospholipids (27). The ability of SecA to interact with the hydrocarbon region of phospholipids in bilayer membranes is also supported by several experiments conducted using vesicles containing acidic phospholipids (26, 28-30). Bilayer destabilizing lipids increase hydrocarbon exposure (29) and accelerate the rate of both preprotein translocation (26) and a conformational change that can be induced in SecA by
interaction with vesicles (30). Moreover, SecA can be labeled by lipids
containing photoactivatable groups in their hydrocarbon moieties (31,
32) when such probes are incorporated into pure lipid vesicles,
although experiments of this kind also indicate that SecA becomes
shielded from such interactions when it binds to SecYEG (14, 32).
SecA is believed to form a physiological dimer based on the
preponderance of the dimeric form in hydrodynamic assays in
vitro (33-35). The monomer-dimer equilibrium is sensitive to
temperature and the ionic environment, and the dimer has a tendency to
form higher order oligomers as protein concentration is increased (34, 36). Detailed analysis of hydrodynamic data indicates that two different forms of the SecA dimer are present in solution under some
circumstances, differing either in their conformation or in the nature
of their intersubunit interface (34). Different studies have come to
differing conclusions regarding the oligomeric state of SecYEG in the
active translocation complex, some supporting its functioning as a
monomer (37) but others supporting its functioning as a dimer (38, 39)
or a tetramer (40). The possibility that SecYEG has a different
oligomeric organization from SecA, combined with the general complexity
of the processive protein translocation reaction, has raised the
possibility that SecA could change its oligomeric state during the
functional translocation cycle (9, 34, 37, 38, 40). However, little
evidence has been presented to support this possibility.
The present work shows that phospholipid and signal peptide ligands
alter the complex equilibria between monomeric and oligomeric forms of
SecA, supporting the possibility that changes in the oligomeric state
of SecA could play a functional role in the protein translocation reaction.
Buffers and Reagents--
KET buffer contains 50 mM
KCl, 1.0 mM Na-EDTA, 25 mM Tris-Cl, pH 7.6. Phospholipid analogues were purchased from Avanti Polar Lipids
(Alabaster, AL) and used without further purification.
Protein Purification--
Escherichia coli SecA
variants were purified as described (41). Strains overexpressing the
PrlD suppressor mutants were obtained from D. B. Oliver
of Wesleyan University.
Signal Peptide Synthesis and Purification--
The
KRR-LamB signal peptide
(+H3N-MMITLRKRRKLPLAVAVAAGVMSAQAMA-COO Tryptophan Fluorescence Anisotropy
Measurements--
Excitation-corrected emission spectra were measured
as described using 297 nm excitation (41). Total fluorescence was
monitored at 340 nm, whereas the anisotropy values were averaged in a
window from 320 to 380 nm to improve the signal to noise ratio of the data. The anisotropy curves changed uniformly throughout this spectral
region and showed no fine structure. Whereas background subtraction was
generally performed using spectra collected from a pure buffer sample,
a fluorescent contaminant in the dicaproyl phospholipid stocks required
protein-free solutions at equivalent phospholipid concentrations to be
used for background subtraction of samples containing concentrations of
these species in excess of 1 mM (i.e. for the
fluorescence experiments summarized in Table II).
Critical Micelle Concentration (CMC) and Micelle Size
Measurements--
Detergent CMCs were determined using in
situ elastic light-scattering measurements in the fluorimeter.
Excitation and emission wavelengths were set to 297 and 298 nm,
respectively, and the excitation-corrected 90° light scattering
signal was measured as a function of detergent concentration (in KET
buffer) using vertical excitation and emission polarizers with a 30-s
averaging time for digital photon-counting and 4-nm slits. Linear
regressions were used to fit the pre- and post-CMC regions in the plot
of light scattering versus detergent concentration, and the
CMC was calculated as the point at which these curves intersect. The
molecular masses of the lyso-MPG and lyso-myristoylphosphatidylcholine
(lyso-MPC) micelles in KET buffer were determined to be 35 and 65 kDa,
respectively, from static light scattering and refractive index
measurements performed using Dawn EOS and Optilab detectors (Wyatt
Inc., Santa Barbara, CA). The details of these experiments will be
published elsewhere.
Analytical Ultracentrifugation Measurements--
Sedimentation
velocity experiments were performed at 20 °C in KET buffer in a
Beckman XL-A centrifuge using an 8-slot rotor at 20,000 rpm.
Double-sector Epon centerpieces were used with 420 µl in the sample
cell and 440 µl of buffer in the reference cell. Absorbance
measurements at 280 nm were taken in 0.002-cm radial steps. Absorbance
and refractive index scans were measured from each cell every 8 min
over the course of 14 h. Absorbance data were analyzed with the
program SEDFIT using the continuous distribution c(S) and c(M) Lamm
equation model (42, 43). The partial specific volume of the protein was
assumed to be 0.734 ml/g, and the density of the solvent was calculated
to be 0.998148 g/ml (44). An ensemble of ~90 scans was used for the
final refinement of each c(M) distribution plot. The meniscus was
identified by manual inspection and refined during fitting, and the
data range was truncated near the bottom of the plateau region in the
ensemble of sedimentation curves. The value of the frictional ratio
f/f0 was initially assumed to be 1.2 and refined by the program (Table I). S values between ~0.5 and ~18
were considered and divided into 1000 steps. Maximum entropy
regularization was used assuming a confidence level of 0.9. All fits
yielded a root mean square deviation below 0.1% for the entire
ensemble of scans (light gray traces in Fig.
1A).
Evaluation of the Effect of Micelle Binding on Analytical
Ultracentrifugation Results--
Data from the sedimentation velocity
runs were analyzed using the program SEDFIT (42, 43) as described above
assuming 8 evenly spaced values for the overall partial specific volume of the hydrodynamic particle between 0.734 and 0.905 cm3/g.
The first value represents the partial specific volume of the protein,
whereas the second value represents an upper limit for the partial
specific volume of the lyso-MPC and lyso-MPG micelles given the fact
that they are more dense than D2O buffer (as evidenced by
their observed sedimentation rather than floatation in this environment). The resulting plot of molecular mass versus
the assumed partial specific volume of the hydrodynamic particle was empirically fit to a third order polynomial. This equation was used to
calculate the total molecular mass of the particle for any given value
of its overall partial specific volume
( Cross-linking Experiments--
50 µM (20 °C) or
46 µM (37 °C) SecA (monomer concentration) was
preincubated for 10 min in cross-linking buffer (10 mM KCl, 20 mM MgOAc2, 50 mM
triethanolamine, pH 7.5) with signal peptides (at a 100 µM concentration) and/or phospholipids (at a 2 mM concentration). Cross-linking was initiated by the
addition of 0.1% glutaraldehyde for 5 (20 °C) or 3 (37 °C) min
and stopped by adding 2× SDS gel loading buffer and boiling for 3 min.
Cross-linked proteins were analyzed by 4% SDS-PAGE and stained with
Coomassie Blue.
Long-chain Phospholipid Analogues Monomerize the Physiological
Dimer of SecA--
Analytical ultracentrifugation (Fig.
1, A-C),
fluorescence anisotropy (Fig. 1D), and glutaraldehyde
cross-linking (Fig. 2A) experiments show that the physiological dimer of SecA is monomerized by
the long-chain phospholipid analogues lyso-MPG or lyso-MPC. Fluorescence anisotropy experiments (Fig. 1D) show that SecA
remains a dimer in the presence of equivalent concentrations of
short-chain phospholipid analogues with the same head group structures
(dicaproylphosphatidylglycerol (DCPG) and dicaproylphosphatidylcholine
(DCPC)).
Absorbance scans from sedimentation velocity experiments conducted
either in the absence or presence of 150 µM lyso-MPG
(Fig. 1A) show an obvious reduction in the sedimentation
coefficient of SecA in the presence of this phospholipid analogue. The
program SEDFIT was used to fit these data and data from an equivalent experiment conducted in the presence of 150 µM
lyso-MPC using the continuous distribution c(S) and c(M)
Lamm equation model (42, 43), with results summarized in Table
I. The c(M) mass distributions inferred
from these analyses (Fig. 1B) indicate that the molecular
mass of SecA is reduced by approximately half in the presence of either
phospholipid analogue, suggesting that they monomerize the
physiological dimer of SecA. Equivalent hydrodynamic results are
obtained in the presence of a 300 or 500 µM concentration of either phospholipid analogue (data not shown).
Several details of these sedimentation velocity experiments deserve
comment. The experiment conducted in the absence of phospholipid confirms that 1 µM E. coli SecA is present
primarily in the form of a dimer (33, 34) at room temperature in a
buffer containing 50 mM KCl, 1 mM EDTA, 25 mM Tris-Cl, pH 7.5 (Fig. 1B and Table I).
Consistent with previous observations (33), a high frictional ratio is
observed for this dimer (Table I). Sedimentation velocity analyses of
the kind used here give time-averaged molecular masses for hydrodynamic
species undergoing rapid equilibration at the sedimentation boundary
(42, 43). Therefore, the minority population of SecA monomer observed
in Fig. 1B indicates that a small fraction of the SecA
molecules are incapable of forming the physiological dimer, probably
because of N-terminal proteolysis (36). The molecular mass of the
nonexchanging SecA monomer is slightly underestimated in Fig.
1B because a single frictional ratio must be assumed in analyzing the entire ensemble of hydrodynamic species (because of
software limitations), and the high value required to model the
sedimentation of the dimer apparently overestimates the frictional ratio of the monomer.
Sedimentation velocity experiments conducted on pure phospholipid
samples and monitored using refractive index measurements show that the
sedimentation of both lyso-MPG and lyso-MPC micelles is
substantially slower than that of the SecA monomer under these conditions (data not shown). Therefore, an approximately constant micelle concentration is present in the protein-containing regions of
the cell during the sedimentation velocity experiments. Because the
phospholipid species have no absorbance at 280 nm, they do not
contribute to the optical absorbance profiles used to determine the
molecular mass distribution of the protein. Therefore, the hydrodynamic
properties of the protein-phospholipid complex can be assessed
directly from the sedimentation absorbance profiles even in the
presence of the micelles.
However, the c(M) distribution (Fig. 1B) calculated for the
complex depends directly on the overall partial specific volume of the
hydrodynamic particle ( Evaluation of the Concentration Dependence of Monomerization Using
Fluorescence Anisotropy Spectroscopy--
Steady-state fluorescence
anisotropy measurements offer a convenient means to monitor the
oligomerization state of a fluorophore-containing protein because of
their sensitivity to changes in rotational correlation time (48). The
intrinsic tryptophan fluorescence of E. coli SecA can
therefore be used to assess the concentration dependence of the
monomerization reaction induced by either lyso-MPG or lyso-MPC
(triangles in Fig. 1D). A steep quenching in
relative total fluorescence (lower panel in Fig.
1D) is observed in the concentration range from 25 to 150 µM when either phospholipid analogue is titrated onto
SecA, coinciding approximately with the CMCs of these micelle-forming
lyso-lipids (~64 µM for lyso-MPG and 68 µM for lyso-MPC). The observed fluorescence quenching
indicates that a protein conformational change occurs upon phospholipid binding (26, 28-30). Because quenching decreases the excited-state lifetime of the fluorophore ensemble, it tends to cause a small increase in fluorescence anisotropy in the absence of a change in the
rotational diffusion coefficient of the protein. Instead, a 30%
decrease in anisotropy closely parallels the major change in relative
total fluorescence in both the lyso-MPG and lyso-MPC titrations
(upper panel in Fig. 1D), indicating that a
substantial increase in the rotational diffusion coefficient of the
protein accompanies the conformational transition. Based on the results of the sedimentation velocity experiments, most of this increase in
rotational diffusion rate is attributable to monomerization of the SecA dimer.
A second apparent binding interaction is observed exclusively in the
titration with the lyso-MPG micelles in the concentration range from
150 to 200 µM, causing an additional 20% quenching in
relative total fluorescence (lower panel in Fig.
1D). However, this second binding event produces a small
increase in anisotropy (upper panel in Fig. 1D),
which could be caused either by the reduction in the lifetime of the
fluorophore ensemble because of the quenching and/or by a slight
reduction in the molecular rotation rate because of the binding of a
second lyso-MPG micelle to the protein.
Phospholipid-induced Monomerization Is Also Observed in Chemical
Cross-linking Experiments--
In the absence of phospholipids,
exposure of SecA to 0.1% glutaraldehyde for 5 min at 20 °C
yields primarily a covalent dimer when samples are analyzed using
SDS-PAGE (lane 5 in Fig. 2A). Addition of either
lyso-MPC or lyso-MPG reduces the cross-linking of the protomers and
results in the protein running predominantly as a monomer even after
glutaraldehyde exposure (lanes 6 and 9 in Fig.
2A).
Similar Total Fluorescence Changes during the Phospholipid-induced
Monomerization and the Endothermic Transition of SecA--
Thermal
titrations can be used to induce an ATP-modulated endothermic
conformational transition in E. coli SecA (8, 28, 49, 50)
(Fig. 1E), which produces an increase in the mobility of the
Evaluation of Different Phospholipid and Detergent Species for the
Ability to Monomerize SecA--
To characterize the chemical features
of the phospholipids responsible for monomerizing the physiological
dimer of SecA, fluorescence anisotropy titrations were conducted with a
variety of different phospholipid and detergent species (Table II). All
of the long-chain lyso-lipid species that were tested trigger
dissociation of the physiological dimer of SecA, with the concentration
producing 50% monomerization being consistently at or slightly below
the CMC of the phospholipid species except in the case of
lyso-palmitoyl-PC. In contrast, the short-chain diacylphospholipids
DCPG and DCPC fail to induce monomerization of SecA at concentrations
either below or above their CMCs. Whereas the nonionic detergents
Synthetic Signal Peptide Redimerizes Phospholipid-monomerized
SecA--
A synthetic analogue of the signal sequence of the LamB
outer membrane protein from E. coli has been used as model
preprotein substrate in a variety of biochemical and biophysical
studies (19). The "KRR-LamB" synthetic signal peptide contains
three extra residues (KRR) compared with the original LamB sequence to
enhance its solubility, but a modification like this has been demonstrated not to perturb LamB signal sequence function in
vivo (13). Sedimentation velocity experiments show that SecA
molecules monomerized by the binding of lyso-MPG are redimerized in the presence of 25 µM wild-type KRR-LamB signal peptide (Fig.
3A and Table I). A small
population of higher order oligomers is also observed in this
experiment, but the dominant hydrodynamic species is a dimer (Fig.
3A and Table I).
A significant increase in molecular mass is also observed when 25 µM wild-type KRR-LamB signal peptide is added to SecA
molecules monomerized by interaction with lyso-MPC (Table I). In this
case, the apparent molecular mass of the complex formed in the presence of the signal peptide is slightly lower than that expected for a
protein dimer (~165 kDa versus slightly greater than 200 kDa). This observation suggests that the redimerization of SecA may be
incomplete under these conditions. Higher concentrations of signal
peptide yield a complex of the same molecular mass (data not shown),
indicating that the lack of complete redimerization in lyso-MPC is not
because of incomplete saturation of the signal peptide binding site.
Therefore, the equilibrium constant for the redimerization of the
SecA-signal peptide complex with lyso-MPC appears to be lower than that
for the equivalent complex with lyso-MPG.
When the concentration dependence of the redimerization of
lyso-MPG-bound SecA by the wild-type KRR-LamB signal peptide is assessed using tryptophan fluorescence anisotropy spectroscopy, a
sigmoidal change in fluorescence anisotropy is observed with a
Kd of 14 µM (Fig. 3B) and a
Hill coefficient of 2 (Fig. 3C), consistent with the
occurrence of a cooperative dimerization reaction involving the binding
of one signal peptide per monomer. The
As opposed to the large fluorescence quenching that is observed upon
phospholipid-induced monomerization of SecA, only a minimal increase in
total fluorescence is observed upon redimerization by the wild-type
KRR-LamB peptide (Fig. 3B). Moreover, the anisotropy level
of the redimerized protein is lower than that of the physiological dimer prior to phospholipid exposure. Because the binding of the KRR-LamB peptide to the physiological dimer of SecA produces only minimal changes in either total Trp fluorescence (data not shown) or
anisotropy (Fig. 4B, below),
both of these observations suggest that the conformation of the
KRR-LamB-bound SecA dimer formed in the presence of lyso-MPG might
differ from that of the original physiological protein dimer.
Glutaraldehyde cross-linking experiments support the conclusion that
signal peptide binding induces redimerization of SecA molecules
monomerized by either lyso-MPG or lyso-MPC, based on the increase in
the level of dimer observed when cross-linking is conducted in the
presence of the wild-type KRR-LamB signal peptide but not the Conformationally Specific Polymerization of SecA Dimers by
Synthetic Signal Peptide--
Addition of 25 µM
wild-type KRR-LamB signal peptide to a solution containing
physiological dimers of SecA at 20 °C produces only a small increase
in the apparent molecular mass of the protein, concomitant with an
increase in the width of the molecular mass distribution, as determined
by sedimentation velocity measurements (Fig. 4A). Therefore,
signal peptide binding to SecA dimers in the compact conformational
ground state induces a modest tendency to self-associate (that is also
observed in the glutaraldehyde cross-linking experiment in lane
4 in Fig. 2A) but without producing a significant shift
of the population into the form of higher order oligomers. Consistent
with this conclusion, at most minor changes in total Trp fluorescence
(lower panel in Fig. 4B and additional data not
shown) or anisotropy (upper panel in Fig. 4B) are
observed when 25 µM wild-type KRR-LamB signal peptide is
added to SecA at 24 °C (Fig. 4B). However, a precipitous
decrease is observed in the Perin plot (48) of reciprocal anisotropy versus temperature when such samples undergo thermal
titration (upper panel in Fig. 4B). This increase
in Trp fluorescence anisotropy coincides with the onset of the
endothermic conformational transition as detected by quenching of total
fluorescence (lower panel in Fig. 4B). Based on
several lines of evidence, this anisotropy change corresponds to the
formation of higher order oligomers by the SecA dimer, indicating that
signal peptide induces protein polymerization when it binds to the
domain-dissociated conformation adopted by SecA at temperatures above
that of the endothermic transition.
The first line of evidence supporting the conformationally specific
polymerization of SecA derives from the detailed properties of the
observed fluorescence changes. The change in total fluorescence observed in Fig. 4B derives almost entirely from the protein
conformational change that occurs during the endothermic transition.
When signal peptide is added to SecA samples after first heating them
to a temperature high enough to induce the endothermic transition, the
strong increase in anisotropy occurs in the absence of any significant
change in total fluorescence (data not shown), establishing that the
anisotropy change does not derive from an alteration in fluorescence
lifetime and must instead derive from an increase in the rotational
correlation time of the Trp ensemble (48). Theoretically, this increase
could be caused either by a protein conformational change leading to
slower rotation of the physiological dimer or by formation of higher
order oligomers of SecA, which would rotate more slowly than the
physiological dimer. However, the large change in rotational
correlation time required to produce the observed 60% increase in
anisotropy seems unlikely to derive from a conformational change and
much more likely to derive from higher order oligomerization. This
conclusion is supported by the observation of detectable turbidity
indicative of light scattering in the samples exposed to elevated
temperature in the presence of signal peptide but not in those exposed
to the same temperature in the absence of signal peptide (data not
shown). Light scattering requires the presence of particles that are
large compared with the wavelength of visible light and therefore
indicates the formation of higher order protein complexes. This
conclusion is also supported by cross-linking experiments, which yield
primarily dimer when WT SecA is exposed to glutaraldehyde at 37 °C
in the absence of signal peptide but high molecular weight species that
do not enter the gel when cross-linking is performed in the presence of
the wild-type KRR-LamB signal peptide at the same temperature
(lanes 2-4 in Fig. 2B). Finally, the
polymerization of domain-dissociated SecA in the presence of signal
peptide is also supported by analytical ultracentrifugation
experiments, which show continued steady progression of the
sedimentation boundary in SecA samples at 37 °C but rapid clearance
of the protein from the sample cell at the same temperature in the
presence of signal peptide (data not show), indicating the formation of
large protein complexes.
When thermal titration of SecA is conducted in the presence of an
equivalent concentration of the KRR-LamB-
Because SecA retains a dimeric structure during the endothermic
transition (41), the polymerization reaction indicates that an
additional intersubunit interface is likely to be formed between protomers when they bind signal peptide in the domain-dissociated conformation. The formation of an equivalent interface between phospholipid-bound SecA monomers could be responsible for their signal
peptide-dependent dimerization given the evidence discussed above for the related conformational properties of the SecA protomer following either phospholipid-induced monomerization or
endothermic domain dissociation combined with the evidence
for possible conformational differences between the physiological dimer
and signal peptide-redimerized SecA.
The Effect of prlD Mutations in SecA on Signal Peptide-induced
Polymerization--
PrlD mutations in SecA are selected based on their
ability to suppress secretion defects caused by mutations in the signal sequence of a preprotein in vivo (52, 53). These alleles
generally facilitate the endothermic conformational transition of SecA
and shift its Tm to lower temperature (50). When
thermal titrations are conducted on a series of prlD alleles
of SecA in the presence of 25 µM wild-type KRR-LamB
signal peptide, the onset of the polymerization reaction moves to lower
temperature, tracking the onset of the endothermic conformational
transition in each allele as monitored by total fluorescence
spectroscopy (Fig. 4C). This correspondence establishes that
the polymerization reaction is controlled by the conformational state
of SecA rather than deriving from the thermodynamic properties of the
signal peptide itself. When thermal titrations of this set of SecA
variants are conducted in the presence of the KRR-LamB- Lyso-lipids Reverse the Signal Peptide-induced Polymerization of
SecA--
To verify that the signal peptide-induced polymerization
does not represent some form of irreversible aggregation, the ability of different lipid species to reverse the polymerization was evaluated using steady-state tryptophan fluorescence anisotropy experiments (Fig.
4D). After heating a SecA sample to a temperature high
enough to induce the endothermic conformational transition and domain dissociation, the addition of wild-type KRR-LamB signal peptide causes
a rapid increase in anisotropy indicating polymerization (arrows on the left in Fig. 4D).
Addition of a 250 µM sub-CMC concentration of DCPG
produces no change in Trp anisotropy (arrow on the
right in the upper trace in Fig. 4D).
However, addition of an equal concentration of lyso-MPG, which is above
its CMC, reduces the anisotropy to a similar level to that observed
prior to signal peptide addition (arrow on the
right in the lower trace in Fig. 4D),
consistent with reversion to a dimeric form. Thus, dissociation of the
signal peptide-induced polymers is achieved by introduction of a
phospholipid species at a concentration that induces monomerization of
the physiological dimer of SecA, but not by the introduction of an
equivalent concentration of a phospholipid species with the same head
group structure and similar hydrocarbon content that is not capable of
monomerizing SecA. Additional experiments show that the anisotropy
level of signal peptide-induced SecA polymers is reduced by the
introduction of lyso-MPC at a concentration above its CMC that induces
monomerization of SecA but not by an equivalent concentration of DCPC
that does not induce monomerization (data not shown). However, whereas
lyso-MPC fully reverses modest signal peptide-induced anisotropy
changes (reflecting lower degrees of polymerization), it only partially
reverses stronger anisotropy changes of the kind shown in Fig.
4D. Therefore, the lyso-PG species seems to be somewhat more
effective that the lyso-PC species in mediating SecA subunit
dissociation under these conditions.
Glutaraldehyde cross-linking experiments yield similar results, showing
reversal of the signal peptide-induced polymerization of SecA by lipids
specifically when they are used under conditions that induce
monomerization of the physiological dimer. A 2 mM concentration of lyso-MPG that is above the threshold required to
induce monomerization produces a strong increase in the amount of
monomeric SecA entering the gel, whereas an equivalent concentration of
DCPG that does not induce monomerization has little effect on the
signal peptide-induced polymerization (lanes 5 and
6 in Fig. 2B). The failure to obtain
glutaraldehyde cross-linked dimers in this experiment in the presence
of lyso-MPG and signal peptide, even though such dimers were observed
after glutaraldehyde cross-linking in the presence of these ligands at
20 °C (lane 7 in Fig. 2A), must be
attributable to the different conditions used in this experiment
(i.e. either the higher temperature or the shorter cross-linking time). However, the efficient cross-linking of the physiological dimer under identical solution conditions (lane 3 in Fig. 2B) provides further evidence that there are
differences in the structure of the intersubunit interface in the SecA
dimer when both lyso-MPG and signal peptide are bound.
The reversibility of the signal peptide-induced polymerization
of SecA makes it unlikely that this phenomenon is caused by protein
aggregation. The data showing that this reversal only occurs when
phospholipid analogues are used under conditions that produce
monomerization of the physiological dimer of SecA reinforce this conclusion.
The SecA translocation ATPase is know to have complex oligomeric
behavior in aqueous solutions (33-36). Although a dimer is the
predominant species present in purified SecA samples in the absence of
other proteins, the protein has a tendency to form higher order
oligomers or dissociate into monomers that depends on protein
concentration, temperature, and the composition of the buffer. In this
paper, we show that ligands that will be encountered by SecA in
the course of protein translocation strongly modulate its
oligomeric behavior.
The protein translocation reaction is believed to be initiated by the
binding of SecA to the N-terminal signal peptide that targets proteins
for export from the cytoplasm (6, 9, 12-21). Because the signal
peptide is cleaved by a protease with a periplasmic active site, at
least its C-terminal region must be translocated across the cytoplasmic
membrane, a process that presumably occurs when SecA binds to SecYEG.
The data presented in this paper show that signal peptide binding tends
to produce oligomerization of SecA in a manner that is modulated by the
conformational state of the protein, with the tendency being weak in
the conformational ground state but strongly enhanced in the high
temperature domain-dissociated state (8, 28, 41, 49, 50) of the
physiological dimer or after monomerization by lyso-lipids. In this
context, changes in oligomeric interactions could occur at specific
steps in the conformational reaction cycle of the SecA-SecYEG complex
(1-9) in a manner controlled by the presence or absence of the
preprotein transport substrate. The results presented in this paper
therefore raise the possibility that ligand-triggered changes in
oligomeric interactions between SecA protomers may play a role in
the complex dynamics of the preprotein translocation reaction (9,
10).
SecA appears to be shielded from the hydrophobic region of the bilayer
when stably inserted into SecYEG (14, 32). However, it does interact
directly with phospholipid bilayers including their hydrophobic cores
in the absence of SecYEG (27). The ability of SecA to interact with the
hydrocarbon region of phospholipids in bilayer membranes is also
supported by several experiments conducted using vesicles containing
acidic phospholipids (26, 28-30), and these interactions are enhanced
(28) in the high temperature domain-dissociated conformation that is
the product of the endothermic conformational transition in SecA.
Because preprotein translocation requires cycles of SecA insertion and retraction from SecYEG (1-5), SecA will not be stably inserted into
SecYEG at all stages of the productive translocation cycle. When it is
retracted from SecYEG, SecA could interact directly with the
phospholipid bilayer, which would then have the opportunity to drive
conformationally specific changes in SecA that could contribute to a
carefully controlled progression of the overall conformational reaction
cycle. Such interactions could account for the acceleration in the rate
of preprotein translocation in the presence of bilayer destabilizing
lipids (26), which increase the solvent exposure of the hydrocarbon
moieties of the phospholipids (29).
The monomerization of SecA by certain phospholipid and detergent
species that is established in this paper is of uncertain physiological
relevance. We believe that monomerization is mediated by the
interaction of SecA with hydrophobic moieties exposed on the surface of
the micelles formed by the active species. Because DCPG and DCPC fail
to induce monomerization of SecA even at super-CMC up to 35 mM, the phospholipid head groups cannot be responsible for
inducing the monomerization of SecA. Whereas lauryl dimethylamineoxide induces monomerization of SecA, two other detergents with equivalent 12-carbon aliphatic chains ( Based on two lines of evidence, the active species responsible for
subunit dissociation in SecA are likely to be micelles and/or
proto-micellar aggregates. First, there is a strong correlation between
the CMC of the amphiphile and the concentration required to produce
monomerization of SecA (Table II), which is readily explained if some
kind of micellar aggregate is the active species. Second, the apparent
cooperativity of the monomerization reaction varies from ~3 to ~8
for the different active species (Table II). This number gives the
minimum number of amphiphile molecules bound during the reaction (54),
so the data indicate that more than one amphiphile molecule per monomer
is required to drive monomerization in all cases. Whereas these data do
not exclude the possibility that a small but variable number of
molecules could drive the monomerization of SecA for the different
amphiphiles, they could also be explained based on variations in the
apparent cooperativity of the micellization reaction (55-57) for the
different amphiphiles if micelle formation is involved in driving
monomerization. In the case of a micelle containing a large number of
monomers (as is the case for all of the species examined here), an
ideal micellization process would involve a very high degree of
cooperativity associated with the formation of micelles starting at the
CMC. However, high sensitivity titration calorimetry studies show that
real micellization reactions show very substantial deviations from
ideality (55-59), including the formation of protomicellar aggregates
of heterogeneous structure at concentrations in the vicinity of the
nominal CMC (i.e. both below and above it). Because of these
species, the apparent cooperativity of the micellization process is
generally considerably lower than the aggregation number of the micelle (55-57) and can be in the range observed for the apparent
cooperativity of the monomerization of SecA by the different
amphiphiles (Table II). Monomerization of SecA by protomicellar
aggregates formed at concentrations below the CMC could therefore
explain the results observed with lyso-lauryl-PC and lauryl
dimethylamineoxide, which trigger the dissociation reaction with
different apparent cooperativities at slightly sub-CMC concentrations.
The calorimetric studies indicate that the protomicellar aggregates
formed near the CMC expose more hydrocarbon to the aqueous environment
than the fully formed micelles present at limiting concentrations above
the CMC (55, 57, 58). The likelihood that such species are involved in
inducing the monomerization of SecA by some of the amphiphiles suggests
that surface exposure of hydrocarbon groups on the aggregates could be
an important molecular parameter in mediating the effect. In this
context, it is noteworthy that the nonionic detergents that fail to
induce monomerization have comparatively larger head groups that will tend to shield the hydrocarbon chains more thoroughly in the
corresponding micelles, giving further support to the hypothesis that
the level of static or dynamic exposure of hydrophobic moieties on the
surface of the micelle could potentially be a critical parameter in
determining the efficacy of the species in driving monomerization of SecA.
This requirement would echo the activity of bilayer-destabilizing
phospholipids in stimulating the conformational reaction cycle of SecA
(26, 29). Therefore, the micelle-induced monomerization of SecA
characterized in this article might reflect what happens to SecA when
interacting with destabilized regions of the phospholipid bilayer
between successive rounds of insertion into SecYEG. Thus, the ability
of certain phospholipid and detergent micelles to drive subunit
dissociation of SecA could reflect the ability of phospholipid
membranes in the correct microenvironment to drive changes in the
interactions between SecA protomers at a specific stage of its
conformational reaction cycle when engaged with SecYEG. If this
hypothesis is correct, oligomeric interactions of SecA could be
reciprocally controlled by signal peptide interaction and phospholipid
interaction at different stages of the translocation cycle.
Each cycle of ATP-dependent binding and release of SecA
from the SecYEG translocon is believed to drive the translocation of
about 40 residues of preprotein (11). Therefore, to achieve processive
translocation of an entire preprotein, there must be some mechanism by
which the translocation of each successive 40-residue polypeptide
segment is efficiently coordinated (10). It is possible that a single
SecA dimer could re-bind to C-terminal segments of a translocating
preprotein molecule following delivery of an N-terminal segment of the
same preprotein to SecYEG. In this case, the probability of rebinding
to a proximal segment of the preprotein could be enhanced by keeping
SecA partially bound to SecYEG between the pumping cycles. However, the
data presented in this paper raise the possibility that processivity
could be achieved using a different mechanism. The observation of
conformationally specific, signal peptide-dependent higher
order oligomerization of SecA suggests that a SecA protomer with bound
preprotein could recruit additional SecA molecules to mediate
translocation of the C-terminal segments of the same preprotein
molecule. Subunit recruitment would likely be temporally coupled to the
delivery of the currently bound preprotein segment to the translocon
because entry into the high temperature domain dissociated conformation
where subunit recruitment occurs is likely to gate the binding of SecA
to SecYEG (20, 50, 60). The phospholipid-induced monomerization of SecA
reported in this paper could then mediate subunit release and recycling.
The tandem motor domains in SecA bear some sequence homology (61) and
strong structural homology (8, 41) to those in ATP-dependent superfamily I and II helicases, with the
closest relationship being observed with the DEAD-box family of RNA
helicases (62, 63). These enzymes mediate the processive unwinding of nucleic acid duplexes, and two competing models have been advanced to
explain how processivity is achieved in this reaction. The "inchworm" model proposes that unwinding is mediated by the
unidirectional translation of a helicase protomer along a single strand
of a nucleic acid polymer, which pushes the duplex open at its leading edge (64, 65). The "rolling" model proposes that unwinding is
mediated when one protomer bound to a single-stranded segment of the
nucleic acid polymer recruits a second protomer to bind to the upstream
segment of the strand in an equivalent manner, thereby stabilizing it
in the single-stranded state (66-68). The first model assumes that a
helicase monomer is the functionally active species, whereas the second
model assumes that oligomerization plays a fundamental role in
mediating processivity. Proponents of the inchworm model cite a 1-base
displacement of a bound oligonucleotide observed in comparing
nucleotide-free and ATP-bound structures of the PcrA superfamily I
helicase (64). They also point out that no consistent pattern of
oligomerization has been observed in the existing helicase crystal
structures, including multiple representatives from both superfamilies
I and II (65, 69). Proponents of the rolling model cite the fact that
both the Rep (68) and NS3 (70) helicases have both been shown to
dimerize upon binding specific substrate DNA structures and therefore
in a conformationally specific manner. The results presented in this paper suggest that SecA could achieve processivity in polypeptide transport using a subunit recruitment mechanism similar to that invoked
in the rolling model for the mechanism of the structurally homologous
ATP-dependent helicases.
While this paper was under review, Or et al. (71) reported
related results. In that work, chemical cross-linking and fluorescence resonance energy transfer results were presented showing that the
physiological dimer of E. coli SecA is dissociated by
interaction with phospholipids and some analogues. These results are
mostly consistent with the results reported here. However, Or et
al. (71) also presented chemical cross-linking data suggesting
that SecA monomerizes upon binding the KRR-LamB signal peptide,
directly contrary to the results reported here. This discrepancy could possibly reflect differences in the behavior of SecA in the
cross-linking buffer used by these authors, although consistent
behavior is observed in our fluorescence and cross-linking experiments
despite the fact that there are greater differences in the compositions of the buffers used in these experiments. On the other hand, the intersubunit cross-linking efficiency is very low in the SecA dimer in
the experiments reported by Or et al. (71), and its reduction in the presence of signal peptide could potentially be
explained by effects unrelated to a change in the oligomeric state of
SecA. For instance, a protein conformational change upon signal peptide
binding could reduce the efficiency of the cross-linking reaction even
in the absence of dissociation of the physiological dimer.
Or et al. (71) also reported isolation of a mutant form of
E. coli SecA with a strongly reduced tendency to form the
physiological dimer. Based on the fact that this variant retains a
small fraction of the activity exhibited by the wild-type protein in an
in vitro preprotein translocation assay, they argue that
SecA is likely to function fundamentally as a monomer and to use an
inchworm mechanism to mediate processivity in preprotein transport
(71). However, they do not characterize the quantitative change in the equilibrium constant for dimerization so that some degree of
dimerization of this variant is still possible at specific stages of
the transport reaction. Furthermore, it is unclear whether the in
vitro assay employed in their studies would be sensitive to
defects in the processivity of the SecA-mediated component of the
preprotein translocation reaction, which is coupled jointly to the
ATPase activity of SecA and also to the proton-motive force (7) (in the
absence of uncoupling reagents that block the formation of transmembrane electrochemical potential gradients). Most importantly, the monomeric variant of SecA exhibits only a few percent of the protein translocation mediated by wild-type dimers under equivalent conditions with wild-type SecYEG, suggesting to us that the reduced ability to form the physiological dimer could be causing a severe defect in translocation activity. In this context, we believe that
additional studies will be required to determine whether subunit
recruitment plays a role in mediating processivity in preprotein transport.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS and METHODS
RESULTS
DISCUSSION
REFERENCES
MATERIALS and METHODS
TOP
ABSTRACT
INTRODUCTION
MATERIALS and METHODS
RESULTS
DISCUSSION
REFERENCES
)
and the
78 variant
(+H3N-MMITLRKRRKLPVAAGVMSAQAMA-COO
)
were synthesized and purified as described (13). The
concentration of signal peptide in a concentrated stock solution in
water was determined using quantitative amino acid analysis (conducted
at the W. M. Keck Biopolymer Facility at Yale University). Some
wild-type signal peptide preparations had a tendency to generate a
precipitate when diluted into KET buffer, complicating fluorescence
anisotropy measurements. However, these preparations produced
equivalent redimerization of lyso-myristoylphosphatidylglycerol
(lyso-MPG)1-bound SecA
monomers in sedimentation velocity assays where the precipitate is
rapidly cleared from the cell so that it does not interfere with quantitation.
overall), as calculated from the assumed
values for the phospholipid-to-protein mass ratio in the complex
(R) and the partial specific volume of the phospholipid
(
PL), as shown in Equation 1.
The molecular mass of the protein in the complex was then
calculated based on the assumed phospholipid-to-protein mass ratio, yielding data of the kind shown in Fig. 1C.
(Eq. 1)
RESULTS
TOP
ABSTRACT
INTRODUCTION
MATERIALS and METHODS
RESULTS
DISCUSSION
REFERENCES
View larger version (25K):
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Fig. 1.
Long-chain phospholipid analogues monomerize
the physiological dimer of SecA. The interaction of wild-type
E. coli SecA with various water-soluble phospholipid
analogues is characterized in KET buffer using analytical
ultracentrifugation and tryptophan fluorescence spectroscopy.
A, four representative absorbance scans from equivalent time
points are shown from sedimentation velocity experiments conducted at
20 °C on 1.0 µM SecA either in the absence
(upper panel) or presence (lower panel) of 150 µM lyso-MPG. The gray traces at the
bottom of each panel show the curve fitting residuals for
the complete ensemble of ~90 absorbance scans used for molecular mass
calculations. B, molecular mass distribution c(M) profiles
were calculated by SEDFIT (42, 43) from sedimentation velocity
experiments like those shown in panel A conducted either in
the absence of phospholipid ( ) or in the presence of 150 µM lyso-MPG (···) or 150 µM lyso-MPC
(- - -). The analysis assumes a partial specific volume of 0.734 cm3/g for all hydrodynamic species. Equivalent results were
obtained when the lyso-lipid concentrations were increased to 300 or
500 µM (data not shown). C, systematic
analysis of the effect of phospholipid binding to SecA on the molecular
mass of the protein as inferred from the sedimentation velocity data.
Because neither the partial specific volume of the phospholipid
(
PL, expressed in cm3/g) nor
the amount of phospholipid bound to SecA is known with certainty, the
analysis was performed assuming all reasonable values for these
parameters. The graph shows the mass of the protein
component of the protein-detergent complex excluding the mass of the
bound lipid. See "Materials and Methods" for details of the
analysis. The curves for which the partial specific volume of the lipid
is not indicated in the figure assumed values of 0.758, 0.783, 0.807, and 0.832 cm3/g. Solid and dashed
lines are used in alternation to facilitate visualization.
D, titrations of 0.25 µM SecA with lyso-MPG
(
), lyso-MPC (
), DCPG (
), or DCPC (
) were conducted at
24 °C and monitored by tryptophan fluorescence spectroscopy.
Steady-state anisotropy is shown in the top panel, whereas
relative total fluorescence at 330 nm is shown in the bottom
panel. E, thermal titrations of 0.25 µM
SecA were conducted in the absence (
) or presence of a 1.0 mM concentration of lyso-MPG (···), lyso-MPC
(- - -), DCPG (-·-·-), or DCPC (-··-··-). Reciprocal
steady-state anisotropy is shown in the top panel, and
relative total fluorescence at 330 nm is shown in the bottom
panel. Reciprocal anisotropy is plotted here because of the linear
relationship between this parameter and temperature for a particle of
constant size and shape as described by the Perin equation (48).
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Fig. 2.
Ligand-dependent changes in
chemical cross-linking of SecA. The effects of phospholipid
analogues (at 2 mM) and signal peptides (at 100 µM) on the oligomeric state of SecA were analyzed using
glutaraldehyde (GA) cross-linking experiments evaluated by
SDS-polyacrylamide gel electrophoresis. See "Materials and Methods"
for experimental details. A, cross-linking conducted for 5 min at 20 °C. B, cross-linking conducted for 3 min at
37 °C. The cross-linking time was reduced in this experiment because
a larger amount of background cross-linking was observed as temperature
was increased.
Hydrodynamic properties of SecA in the presence of model translocation
ligands
overall), which
will change depending on how much phospholipid is attached to the
protein molecule. To estimate the influence of this effect on the
inferred molecular mass distributions, the sedimentation velocity data
from the experiments conducted in the presence of phospholipid micelles
were reanalyzed assuming varying lipid to protein ratios in the
hydrodynamic particles (shown explicitly for lyso-MPG in Fig.
1C and summarized for both phospholipid species in Table I).
Because the partial specific volumes of these phospholipid species are
not known, this analysis was performed assuming all reasonable values
for this parameter. Most phospholipids and detergents have partial
specific volumes in the range between 0.85 and 0.89 cm3/g
(45-47), but the analysis was performed assuming a considerably broader range of values up to the experimental upper limit of 0.905 cm3/g (see "Materials and Methods") and down to 0.734 cm3/g (i.e. equivalent to the partial specific
volume of the protein). This analysis shows that the sedimentation
velocity data for SecA in the presence of 150 µM lyso-MPG
are inconsistent with the hydrodynamic particle containing anything
larger than a SecA monomer (Fig. 1C). Moreover, the
calculated molecular mass closely matches that of the SecA monomer for
the binding of amounts of lipid ranging from a single lyso-MPG monomer
to a complete micelle (which has a molecular mass of 35 kDa) assuming a
partial specific volume in the range from 0.85 to 0.90 cm3/g. An equivalent analysis of the sedimentation data
obtained in the presence of lyso-MPC yields a similar conclusion (Table II). However, in this case, assuming that
a small number of lyso-MPC monomers are bound to the protein gives a
molecular mass estimate slightly higher than that of the SecA monomer,
whereas a closer match is obtained assuming that one micelle (which has
a molecular mass of 65 kDa) is bound to the protein molecule.
Efficacy of different detergent species in inducing monomerization of
SecA
-helical wing domain in the protomer but not dissociation of the
physiological dimer (41). This transition does produce a decrease in
tryptophan anisotropy (i.e. increase in reciprocal anisotropy in the Perin plot (48) in the upper panel of Fig. 1E), but the magnitude of this anisotropy change is
significantly smaller than that produced by the phospholipid-induced
monomerization reaction (upper panels in Fig. 1,
D and E). Nonetheless, the change in relative
total fluorescence that takes place during the endothermic conformational transition (20, 28, 50) (lower panel in Fig. 1E) is very similar to the change that takes place during
the phospholipid-induced monomerization (lower panel in Fig.
1D), suggesting that there may be similarities
in the conformational changes that take place within the SecA protomer
in the two cases. Consistent with this possibility,
phospholipid-monomerized SecA does not experience any additional change
in relative total fluorescence upon subsequent thermal titration (Fig.
1E). Dissection of the contribution of the individual Trp
residues in SecA to the fluorescence changes shown in Fig.
1D2 also supports
the conclusion that a related conformational change takes place in the
protomer during phospholipid-induced monomerization and during the
endothermic conformational transition, even though the physiological
SecA dimer does not monomerize in the latter case.
-octylglucoside,
-dodecylmaltoside, and
C12E8 (octaethylene glycol dodecyl ether) also
fail to trigger monomerization at concentrations either below or above
their CMCs, lauryl dimethylamineoxide does so at a concentration slightly below its CMC.
View larger version (21K):
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Fig. 3.
Synthetic signal peptide redimerizes
phospholipid-monomerized SecA. The experiments were conducted on
wild-type E. coli SecA in KET buffer either in the presence
or absence of the synthetic KRR-LamB signal sequence peptide (13).
A, molecular mass c(M) distribution profiles were calculated
by SEDFIT (42, 43) from sedimentation velocity experiments on 1.0 µM SecA in the presence of 150 µM lyso-MPG
either without (···) or with 25 µM wild-type
KRR-LamB signal peptide (-·-·-). The experiments were performed at
20 °C. Equivalent results were obtained when the lyso-lipid
concentration was increased to 300 µM (data not shown).
B, after monomerizing 2.0 µM SecA by exposure
to 250 µM lyso-MPG, titrations of either wild-type
KRR-LamB signal peptide (squares) or the 78 mutant
peptide (circles) were monitored using tryptophan
fluorescence spectroscopy. Steady-state anisotropy is shown in the
top panel, whereas relative total fluorescence at 330 nm is
shown in the bottom panel. The arrows indicate
the values observed for the physiological dimer of SecA prior to the
addition of lyso-MPG. C, Hill plot for the titration of
wild-type KRR-LamB signal peptide presented in panel B based
on the steady-state anisotropy data.
78 variant of the KRR-LamB
signal peptide contains a 4-residue deletion that has been observed to
severely impair the function of the corresponding signal sequence
in vivo (51). The specificity of the signal peptide-induced
redimerization is demonstrated by the fact that an equivalent titration
of lyso-MPG-monomerized SecA with the KRR-LamB-
78 mutant signal
peptide produces only minor changes in fluorescence anisotropy (Fig.
3B).
View larger version (24K):
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Fig. 4.
Synthetic signal peptide induces
polymerization of the high temperature domain-dissociated conformation
of SecA. Experiments were performed on E. coli SecA in
KET buffer. A, molecular mass distribution profiles were
calculated by SEDFIT from sedimentation velocity experiments conducted
on 1.0 µM wild-type SecA either in the absence ( ) or
presence of 25 µM wild-type KRR-LamB signal
peptide (···). The experiments were performed at
20 °C. B, thermal titrations of 0.25 µM
wild-type SecA were monitored using tryptophan fluorescence
spectroscopy in the absence of signal peptide (
) or in the presence
of a 25 µM concentration of either wild-type KRR-LamB
signal peptide (···) or the
78 mutant peptide (-·-·-).
Perin plots (48) of reciprocal anisotropy are shown in the top
panel, whereas plots of relative total fluorescence at 330 nm are
shown in the bottom panel. C, thermal titrations
were conducted in the same manner as in panel B on wild-type
SecA (
) as well as the Y134C (---), A373V (-·-·-), and A507V
(-··-··-) prlD suppressor mutants (52, 53) in the
presence of 25 µM wild-type KRR-LamB signal peptide.
D, steady-state tryptophan fluorescence anisotropy data
showing dissociation of signal peptide-induced SecA polymers
specifically by phospholipids producing monomerization of the
physiological dimer (i.e. by lyso-MPG above its CMC but not
by DCPG). The A373V variant of SecA was heated to 37.5 °C to induce
domain dissociation and allowed to equilibrate. This variant was used
because the lower temperature of the endothermic transition gives
efficient signal peptide-induced polymerization at a reduced
temperature. Addition of wild-type KRR-LamB signal peptide
(arrows on the left) causes a rapid increase in anisotropy
reflecting protein polymerization. After 5 min, phospholipids were
added at the indicated concentrations (arrows on the right).
To facilitate visualization of the results, an arbitrary offset has
been added to the data from the DCPG experiment but not the lyso-MPG
experiment. There were minimal changes in the total fluorescence during
these experiments other than a small amount of photooxidation (data not
shown).
78
variant (lanes 6-11 in Fig. 2A).
78 mutant signal peptide,
polymerization is still observed but only at substantially higher
temperatures than with the wild-type signal peptide (Fig. 4B). There is a latent kinetic component in thermal
titrations of this kind, so that the higher temperature at which the
major change in anisotropy is observed could reflect slower and less efficient polymerization of SecA by the mutant compared with wild-type signal peptide. The ability of SecA to mediate processive preprotein transport suggests that its binding site for transport substrate is
capable of recognizing and binding a great diversity of polypeptide sequences. In this context, some degree of interaction between SecA and
a mutated signal peptide is not surprising. Tighter binding of
wild-type signal sequences to the polypeptide transport site of SecA
could help ensure efficient initiation of preprotein translocation.
78 mutant
signal peptide, the prlD alleles also shift the
polymerization reaction to lower temperature so that its thermal
dependence more closely resembles that of wild-type SecA with wild-type
signal peptide (data not shown). Therefore, the prlD alleles
enhance the efficiency of the interaction of SecA with a defective
signal peptide in this in vitro assay.
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS and METHODS
RESULTS
DISCUSSION
REFERENCES
-dodecylmaltoside and
C12E8) do not, indicating that having a
hydrocarbon chain of a given length is not sufficient for an amphiphile
to be active. Whereas this pattern could be attributable to a complex
interplay of requirements for head group and hydrocarbon structures, it
could also be explained by requirements for the physiochemical
properties of the micelle formed by the amphiphile.
![]() |
ACKNOWLEDGEMENTS |
---|
We thank D. B. Oliver for providing the prlD mutants of SecA, J. E. Gouaux and R. Olson for access to the analytical ultracentrifuge and advice on its use, and M. Crawford of Yale University for performing quantitative amino acid analyses.
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FOOTNOTES |
---|
* This work was supported by National Institutes of Health Grants GM58549 (to J. F. H.) and GM34962 (to L. M. G.), a startup grant from Columbia University (to J. F. H.), and a long-term postdoctoral fellowship from the European Molecular Biology Organization (to J. B.). The Columbia analytical ultracentrifuge facility was supported by National Institutes of Health shared equipment Grant S10RR12848.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.
§ These authors contributed equally to the results reported in this paper.
To whom correspondence should be addressed. Tel.:
212-854-5443; Fax: 212-865-8246; E-mail:
hunt@sid.bio.columbia.edu.
Published, JBC Papers in Press, October 27, 2002, DOI 10.1074/jbc.M205992200
2 J. J. Fak, A. Itkin, D. D. Nicolae, C. M. Golsaz, and J. F. Hunt, manuscript in preparation.
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ABBREVIATIONS |
---|
The abbreviations used are: lyso-MPG, lyso-myristoylphosphatidylglycerol; lyso-MPC, lyso-myristoylphosphatidylcholine; CMC, critical micelle concentration; DCPG, dicaproylphosphatidylglycerol; DCPC, dicaproylphosphatidylcholine.
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REFERENCES |
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