From the Department of Microbiology and Immunology,
University of Oklahoma Health Sciences Center, Oklahoma City,
Oklahoma 73190, the § Department of Cell Biology, Scripps
Research Institute, La Jolla, California 92037, the
Biota
Structural Biology Laboratory, St Vincent's Institute of Medical
Research, Fitzroy, Victoria 3065, Australia, the ** Departments of
Medical Biochemistry and Genetics, of Chemistry, and of
Biochemistry and Biophysics, Texas A&M University, College Station,
Texas 77843-1114, and the ¶ Protein Crystallography
Laboratory, St. Vincent's Institute of Medical Research, Fitzroy,
Victoria 3065, Australia
Received for publication, October 29, 2000
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ABSTRACT |
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Perfringolysin O (PFO), a member of the
cholesterol-dependent cytolysin family of pore-forming
toxins, forms large oligomeric complexes comprising up to 50 monomers.
In the present study, a disulfide bridge was introduced between
cysteine-substituted serine 190 of transmembrane hairpin 1 (TMH1) and
cysteine-substituted glycine 57 of domain 2 of PFO. The resulting
disulfide-trapped mutant (PFOC190-C57) was devoid
of hemolytic activity and could not insert either of its transmembrane
Perfringolysin O (PFO)1
is a member of the large family (1) of
cholesterol-dependent cytolysins (CDCs). Two defining
characteristics of CDC cytolysis are the formation of large
homo-oligomeric structures (2, 3) on sterol-containing membranes, and
the absolute dependence of pore formation on the presence of
cholesterol, or closely related sterols, in the membrane (4, 5). These
toxins are produced as soluble monomers that ultimately form the large homo-oligomeric pore-forming structure on sterol-containing membranes. The only crystal structure of a soluble, monomeric form of a CDC was
recently solved by Rossjohn et al. (6) for perfringolysin O from Clostridium perfringens. The structure of the
membrane-bound oligomeric structure of a CDC is not presently
available; however, we have used multiple fluorescence techniques to
identify the residues in domain 3 that form two The timing of oligomer assembly and of the insertion of individual
transmembrane domains is controversial. Rossjohn et al. (6)
originally proposed that the CDCs form a prepore complex on the
membrane prior to its insertion into the membrane, similar to other
unrelated We have now trapped the PFO oligomer in a prepore state by the
introduction of a disulfide bridge into the structure of the soluble
PFO monomer. In the present study, a disulfide bridge was introduced
between cysteine-substituted serine 190 of transmembrane hairpin 1 (TMH1) and glycine 57 of domain 2. In the oxidized state, the same
large oligomeric prepore complex described by Shepard et al.
(13) was found to form on the membrane surface. Furthermore, the
initial rate of PFO binding to the membrane, mediated by domain 4 (Fig.
1), was not altered by the presence or absence of the disulfide link in
PFO (14). However, the disulfide-linked PFO oligomers were unable to
insert their TMHs or form pores in the membrane. Thus, the
disulfide-trapped PFO allowed us to convert the membrane-bound monomer
into a prepore complex and then (at our discretion) to synchronize the
conversion of the prepore to pore complex by the addition of reducing
reagent. These studies provide the first direct evidence that the
prepore facilitates the membrane insertion of the large transmembrane
Design of the Disulfide in PFO--
Potential sites for cysteine
residue pairs that might form disulfide bonds were selected using the
method of Hazes and Dijkstra (15). Briefly, potential residue pairs
were initially selected on the basis of appropriate C Preparation of the PFO Derivatives and Their Modification with
Fluorescent Probes--
The cysteine-less gene for PFO encoding
PFOC459A that was cloned in the expression vector pTrcHisA
(Invitrogen, Carlsbad, CA) (8) was used as the template in PCR
mutagenesis to create the double mutant PFOC190-C57. Four
additional mutants of PFOC190-C57 were generated in which a
cysteine was substituted for amino acid residues Ala215,
Val202, Lys288, or Ile303.
Ala215 and Val202 reside in TMH1, whereas
Lys288 and Ile303 reside in TMH2. PFO
derivatives were expressed and purified as before (7, 8), except that
the fractions were stored in the absence of DTT to maintain the
disulfide bond. Labeling of the cysteine derivatives,
PFOC190-C57/C215, PFOC190-C57/C202,
PFOC190-C57/C288, and PFOC190-C57/C303, with
IANBD or IATR was performed as previously described (7, 8), except that
3 M guanidine hydrochloride and DTT were not used in the
labeling procedure. The relative hemolytic activity of the NBD-labeled
and unlabeled proteins were determined as described previously (8). The
extent of dye reaction with the PFOC190-C57 disulfide
mutant was less than 9%, thereby indicating that >90% of the
sulfhydryls were in disulfide bonds. The extent of labeling of the
triple mutants, PFOC190-C57/C215,
PFOC190-C57/C202, PFOC190-C57/C288, and
PFOC190-C57/C303, was determined as before (8) and
indicated that 100% of the cysteines not in disulfide bonds were
labeled with IANBD or IATR (data not shown).
Preparation of Liposomes--
All phospholipids were obtained
from Avanti Polar Lipids (Alabaster, AL). Cholesterol was obtained from
Steraloids (Wilton, NH). Liposomes were prepared by extrusion using
1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine and
cholesterol at a ratio of 45:55 mol% as before (8).
Electron Microscopy--
The disulfide-trapped mutant (900 nM) (± dithiothreitol) in buffer (20 mM HEPES
(pH 7.4), 40 mM NaCl ± 1 mM DTT) in
12-µl droplets were coated with 0.5-1 µl of a 0.5 mg/ml lipid
mixture containing ~60 mol% cholesterol (Sigma) and 40 mol%
1,2-dioleoyl-sn-glycero-3-phosphocholine in chloroform.
After incubation at room temperature for 10 min, the PFO oligomeric
rings were transferred to carbon support films that coated electron
microscope grids and then negatively stained with 1% (w/v) uranyl
acetate as described previously (16). Electron micrographs were
recorded (magnification, ×40,000) with a Philips electron microscope
model 208S.
Fluorescence Measurements--
All fluorescence measurements
were performed in a SLM-8100 photon counting spectrofluorimeter (SLM
Instruments) as described previously (8). An excitation wavelength of
470 nm was used for NBD, and the emission intensity was measured
between 500 and 600 nm. The bandpass was 4 nm for all experiments.
Emission scans of NBD-labeled residues in TMH1 (Cys215 or
Cys202) or TMH2 (Cys288 or Cys303)
were recorded in both the presence and absence of 10 mM DTT in 2 ml of buffer A (50 mM HEPES (pH 7.5), 100 mM NaCl) at 37 °C. Excess liposomes were added to
monomeric NBD-labeled PFOC190-C57 derivatives (185 nM) and allowed to incubate for 30 min at 37 °C to
ensure that oligomerization and insertion into the membrane were
complete before the intensity measurements were made.
Fluorescence resonance energy transfer (FRET)-based kinetic experiments
were carried out with only the following changes to our previous
procedures (17). The donor dye was NBD, and the acceptor dye was
tetramethylrhodamine (final PFO concentration was 190 nM).
The donor-labeled PFOC190-C57/C215 was mixed at a 1:1 molar
ratio with acceptor-labeled PFOC190-C57/C215 in 2 ml of
buffer A in the presence or absence of 10 mM DTT at 37 °C. The excitation wavelength was set at 470 nm, and the donor emission intensity was measured every 20 s at 540 nm. After
30 s, excess liposomes were injected into the mixture and the
emission intensity was monitored for an additional 1170 s. The
liposomes had been titrated as described previously to ensure that an
excess of liposomes was used to quantitatively bind and insert the PFO (8).
Hemolysis Measurement--
Hemolysis was measured on the SLM
8100 at 37 °C by following the decrease in right-angle light
scattering of erythrocytes as they lysed, using the procedures
described previously by Harris et al. (17).
Gel Electrophoresis--
Denaturing agarose gel electrophoresis
was performed as described elsewhere (13). Briefly, in all samples in
which PFO was incubated with liposomes, the liposomes (45 mol%
1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine, 55 mol%
cholesterol) were incubated with PFO for 30 min at 37 °C. Oligomeric
complexes were solubilized with SDS sample buffer and separated on
1.5% SeaPlaque agarose (FMC, Rockland, ME) in SDS gel reservoir buffer
(18). Samples were loaded and the gel was run at 100 V for 3 h.
Gels were fixed in 10% (v/v) acetic acid, 30% (v/v) methanol
overnight, then dried in an Easy Breeze gel dryer (Hoefer, San
Francisco, CA). The dried gel was stained with Coomassie Brilliant Blue
R and then destained to visualize the protein bands.
Hemolytic Activity of the Disulfide-trapped PFO--
The
substitution of cysteines for amino acid residues Ser190
and Gly57, and the subsequent formation of the disulfide
bridge between TMH1 and domain 2, rendered PFO hemolytically inactive
(Table I). If the disulfide bridge alone
were responsible for the lack of activity of the double
mutant, then the activity would be restored when the disulfide bridge
was reduced. As predicted, once reducing agent was added to the sample,
the hemolytic activity was restored to nearly wild-type levels (Table
I). Similarly, the NBD-labeled and unlabeled derivatives of
PFOC190-C57 (PFOC190-C57/C215,
PFOC190-C57/C202, PFOC190-C57/C288, and
PFOC190-C57/C303) exhibited less than 6% of the hemolytic
activity of the parent toxin PFOC459A in a nonreducing
environment (Table I).
We have previously determined the kinetics of erythrocyte hemolysis by
PFO by monitoring the decrease in right-angle light scattering that
accompanies hemolysis (17). In their studies, Harris et al.
(17) showed that PFO-dependent hemolysis exhibited a lag
period prior to the onset of hemolysis. When pre-reduced PFOC190-C57 was injected into a stirred suspension of
erythrocytes, the same hemolysis kinetics were observed. Hemolysis
began after about 100 s and reached completion by ~370 s after
the injection of the toxin (Fig. 2). In
contrast, no hemolysis was detected when the disulfide-trapped mutant
PFOC190-C57 was examined by this method in the absence of
DTT. However, when DTT was added to the disulfide-trapped mutant
370 s after it had been mixed with the erythrocytes, hemolysis
began almost immediately and was complete within 60 s. The lysis
of the erythrocytes was nearly 7 times faster than when the
disulfide-trapped mutant was reduced with dithiothreitol prior to
adding the toxin to the erythrocytes. Therefore, even though
PFOC190-C57 was hemolytically inactive before reduction of
the disulfide, it was clear that the cytolytic mechanism had progressed
to a stage that largely eliminated the typical lag period. The
rate-limiting step in hemolysis therefore must occur prior to the stage
at which the oxidized PFOC190-C57 derivative is trapped. We
therefore characterized the oligomeric state of the disulfide-trapped
mutant and examined the environment of the two transmembrane domains in
the disulfide-trapped state.
Characterization of the Prepore Complex by Agarose Gel
Electrophoresis and Electron Microscopy--
As shown in Fig. 2,
disulfide locking TMH1 to domain 2 of PFO trapped the PFO in a
structural state that did not allow pore formation to occur, but did
prime the system for hemolysis. Although the structural nature of this
trapped state was initially unclear, we suspected that it was trapped
in a prepore complex. Therefore, we employed a SDS-agarose gel
electrophoresis (SDS-AGE) system, previously described by Shepard
et al. (13), to directly examine the distribution of PFO
oligomers under nonreducing and reducing conditions. As expected, in
the absence of liposomes, PFO and PFOC190-C57 (unreduced or
reduced) migrate as a monomer in the SDS-agarose gel system (Fig.
3, lanes 1-3). In
the presence of cholesterol-containing liposomes under nonreducing
conditions, PFOC190-C57 (Fig. 3, lane
5) formed large oligomeric complexes comparable to the
parent toxin PFOC459A (lanes 4 and
6) and to those formed under conditions in which the
disulfide of PFOC190-C57 was reduced (lane
7). Although oxidized PFOC190-C57 formed the
same-sized oligomer as the cysteine-less parent toxin PFOC459A, it apparently was more susceptible to
dissociation by SDS than if it was reduced and allowed to insert
(compare lane 5 with lane 7). If a chemical cross-linker is first used to stabilize
this complex, we do not observe the dissociation of the oxidized
PFOC190-C57 oligomer into monomers (data not shown). The
oligomer of the reduced form of PFOC190-C57 appeared to be
as stable to SDS as the cysteine-less parent toxin PFOC459A. These data suggest that the formation of the
transmembrane
The oligomeric complexes formed by PFOC190-C57 were also
examined by electron microscopy (EM) after their formation on lipid
monolayers in the presence (Fig.
4A) or absence (Fig.
4B) of DTT. Primarily rings and some arcs were observed by
EM and are typical of the structures typically seen by EM when
membranes are treated with PFO (e.g. Ref. 13). Furthermore,
a comparison of the structures present in the presence or absence of
DTT reveals that the oxidation state of the disulfide of
PFOC190-C57 has little effect on the overall topography of
the PFO oligomers as seen by EM.
Oligomerization Rates of Reduced and Unreduced
PFOC190-C57/C215 Determined by FRET--
The extent of
FRET between a donor dye and an acceptor dye depends upon, among other
things, the distance between the dyes. FRET is therefore an excellent
technique for detecting the proximity of two proteins if one
polypeptide is labeled with a donor dye and the other is labeled with
an appropriate acceptor dye. We have previously used this approach to
monitor the oligomerization of PFO monomers (17). Furthermore, since
fluorescence is a nondestructive technique, one can monitor the signal
continuously and hence determine the kinetics of association.
The rate of oligomerization of the disulfide-trapped PFO was
examined by FRET between donor NBD-labeled
PFOC190-C57/C215 and acceptor tetramethylrhodamine-labeled
PFOC190-C57/C215 when cholesterol-containing liposomes were
added to a 1:1 mixture of these proteins. The existence of FRET, and
hence the close approach of donor- and acceptor-labeled subunits, can
be detected by the reduction in donor emission intensity as excitation
energy is transferred to the acceptor. As shown in Fig.
5, the introduction of liposomes to a
mixture of donor- and acceptor-labeled PFO subunits causes a decrease
in NBD intensity, indicating that the subunits have associated. The
decrease in NBD intensity is due to FRET because no decrease in NBD
intensity was observed when the sample lacked acceptor dyes (Fig. 5).
Most important, the rate and extent of FRET were virtually identical
whether or not the disulfide was reduced. Hence, the introduction of
the disulfide did not appreciably affect the rate of PFO
oligomerization, thereby demonstrating that the rate-limiting step in
the process that leads to oligomer formation is not significantly
altered by the presence of the disulfide bond.
Membrane Insertion of TMH1 and TMH2 in the Disulfide-trapped
Complex--
Since hemolysis was prevented (Table I, Fig. 2), but the
formation of the oligomer was unaffected by the introduction of the
C190-C57 disulfide (Fig. 5), we presumed that the disulfide bond
prevented the insertion of at least TMH1 because the disulfide bond
covalently links residues 190 of TMH1 and 57 of domain 2 (Fig. 1).
Although it was easy to envision that the disulfide bridge could
prevent TMH1 from assuming the conformation necessary for proper
insertion into the bilayer, it was not clear whether inhibiting the
movement of TMH1 would also prevent TMH2 from moving into the membrane.
In contrast to TMH1, which is partially buried in the core of the PFO
molecule, TMH2 is largely exposed to the solvent (6) and hence this
domain, unrestricted by a disulfide, might partially or fully insert
into the membrane independent of TMH1.
To determine whether either TMH in a disulfide-trapped PFO molecule
could insert into the membrane, four derivatives of
PFOC190-C57 were generated in which a cysteine was
substituted for amino acid residues located either in TMH1 at amino
acids Val202 or Ala215 or in TMH2 at amino
acids Lys288 or Ile303. All four residues have
been previously shown to face the membrane in TMH1 or TMH2 (7, 8) and
hence are excellent indicators of the transmembrane insertion of the
two TMHs. These mutants were modified with NBD, an environmentally
sensitive fluorescent dye that we have used previously to map the
orientation of these residues in the membrane (7, 8). Since the
emission of NBD is strongly quenched by water, the fluorescence
intensity increases significantly when the dye at these locations in
the TMHs moves from an aqueous to a nonaqueous environment (7, 8,
13).
The fluorescence emission spectrum of each mutant was determined
spectroscopically before and after incubation with liposomes in the
absence or presence of reducing reagent (Fig.
6). In three cases, the fluorescence
intensities of NBD in the PFOC190-C57 mutants that had been
incubated with liposomes in the absence of DTT were nearly the same as
those of the respective soluble monomers (Fig. 6). For the V202C
mutant, which has the dye located at the tip of TMH1 near the
Thus, neither TMH1 nor TMH2 were able to interact normally with the
membrane in oxidized PFOC190-C57 because neither TMH1 nor
TMH2 inserted into the membrane in the disulfide-trapped PFO prior to
reduction of the Cys190-Cys57 disulfide bridge.
These data also suggest that the insertion of both TMHs may proceed in
a concerted fashion and that TMH2 apparently cannot insert independent
of TMH1.
Insertion Kinetics of TMHs--
Pore formation by PFO requires its
binding to the membrane, oligomerization, and TMH insertion into the
bilayer. The rate-limiting step in this process is unknown, but could
occur in any of these three stages of pore formation. Since pore
formation is blocked in the oxidized PFOC190-C57 protein
(Fig. 2, Table I) without significantly altering either prepore complex
assembly (Figs. 3 and 4) or the interaction of domain 4 with the
membrane (14), binding and oligomerization can be examined in the
absence of insertion. Thus, the kinetics of binding and oligomerization
can be measured both with and without insertion to determine whether
insertion slows the process, i.e. is the rate-limiting step
in the process. The rate of domain 4 exposure to the membrane is the
same for reduced and oxidized PFOC190-C57 (14), and the
rate of PFO oligomerization is also unaffected by the presence or
absence of the disulfide bond (Fig. 5). These results suggest that
insertion is not rate-limiting.
To determine directly whether TMH insertion or a previous step is
rate-limiting, we have taken advantage of the disulfide bond to arrest
the pore formation process prior to insertion. As noted above, oxidized
PFOC190-C57 will form a prepore complex, but will not
insert its TMHs. Thus, by allowing the disulfide-trapped PFO to bind
and assemble into prepore complexes on the membrane, we can then
measure the intrinsic rate of TMH insertion in these arrested complexes
simply by adding DTT. This approach allows us to synchronize TMH
insertion in the sample since most or all of the TMHs will already be
in prepore complexes and will be poised to insert into the bilayer.
Membrane insertion of the TMHs was monitored by
the increase in fluorescence intensity over time of NBD
located at the membrane-facing residues C215 in TMH1
(PFOC190-C57/C215) or C288 located within TMH2
(PFOC190-C57/C288) (Fig. 7).
In these experiments, liposomes were injected after 30 s into a
stirred solution of the NBD-labeled PFO in the presence or absence of
reducing agent. For samples that were not pre-reduced prior to the
addition of liposomes, DTT was injected into the unreduced samples at
either 120 or 370 s after the addition of the liposomes. Those
samples that had not been pre-reduced were therefore able to form
prepore complexes prior to the addition of DTT, but were not able to
form pores.
The rate of insertion of the TMHs that were trapped on the membrane in
the prepore state for either 120 or 370 s after the addition of
liposomes proceeded much faster than that of the pre-reduced sample
(Fig. 7). Therefore, the extent of the rapid insertion of the TMHs
appears to reflect the extent of prepore formation because the rate of
formation of the prepore oligomer by oxidized PFOC190-C57
is nearly identical to that by reduced PFOC190-C57. In
fact, the observed rate of insertion, although rapid, includes the rate
of mixing of the DTT and the rate at which the disulfide is reduced.
Therefore, the actual rate of insertion of the two TMHs from the
prepore complex is even faster than the observed rate. Not
surprisingly, after the initial rapid rate of insertion of the
The concept of a prepore as an intermediate state for pore-forming
toxins was first proposed by Walker et al. (19) for the pore-forming toxin The large size of the assembled CDC pore has led to speculation as to
the mechanism by which the CDC polypeptides form their large
transmembrane -hairpins (TMHs) into the membrane unless the disulfide was reduced.
Both the size of the oligomer formed on the membrane and its rate of
formation were unaffected by the oxidation state of the
Cys190-Cys57 disulfide bond; thus, the
disulfide-trapped PFO was assembled into a prepore complex on the
membrane. The conversion of this prepore to the pore complex was
achieved by reducing the C190-C57 disulfide bond.
PFOC190-C57 that was allowed to form the prepore prior to
the reduction of the disulfide exhibited a dramatic increase in the
rate of PFO-dependent hemolysis and the membrane insertion
of its TMHs when compared with toxin that had the disulfide reduced
prior mixing the toxin with membranes. Therefore, the rate-limiting
step in pore formation is prepore assembly, not TMH insertion. These
data demonstrate that the prepore is a legitimate intermediate during
the insertion of the large transmembrane
-sheet of the PFO oligomer.
Finally, the PFO TMHs do not appear to insert independently, but
instead their insertion is coupled.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-hairpins that span
the membrane in the inserted oligomeric PFO complex (7, 8). The
corresponding residues were found to exist as six short
-helices in
the crystal structure of the soluble PFO monomer (6). The
-helical
to
-strand transition in the secondary structure together with the contribution of two
-hairpins per monomer to the transmembrane
-sheet are so far unique to PFO and perhaps the other CDCs.
-barrel pore-forming toxins. In the prepore model (9),
individual membrane-spanning
-hairpins of each monomer are brought
together to form an oligomeric prepore that coordinates the insertion
of the pre-
-hairpins into the membrane as a single membrane-spanning
-barrel. This model has been shown to be used by toxins that form
small oligomeric prepore complexes such as Aeromonas
hydrophila aerolysin, Staphylococcus aureus
-hemolysin, and Clostridium septicum
-toxin (9-12).
In support of this model for the CDCs, we have recently shown (13) that PFO oligomerization can be uncoupled from the membrane insertion of the
transmembrane
-hairpins by lowering the temperature. Shepard et al. (13) also demonstrated that PFO insertion into planar bilayers resulted in the formation of discrete, large channels. Based
on these findings, PFO was proposed to form a prepore complex prior to
the insertion of its transmembrane domain.
-sheet of the CDCs into the bilayer.
EXPERIMENTAL PROCEDURES
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ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-C
distances. Sulfur positions were generated for these residues, and a
check was made to determine whether certain stereochemical criteria
were obeyed. An important criterion was that the
-3 angles of a
potential bridge should not deviate by more than 30° from observed
preferences. Selected pairs were subjected to energy minimization, and
energetically favorable conformations were chosen (less than 10 kcal/mol). This method will be successful if main-chain conformations
are very similar between wild-type and mutant. Calculations were based on the crystal structure of PFO
(6).2 The model had been
refined to an R-factor of 0.211 (Rfree of 0.268) at 2.2 Å resolution with good
stereochemistry. The best candidate for engineering a disulfide bridge
between domain 2 and TMH1 was the residue pair of G57 and S190 (Fig.
1). The two residues were predicted to
form a bond that complied with the stereochemical (
-3 approximately
57.0°) and conformational energy (7.9 kcal/mol) criteria.
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Fig. 1.
Location of the disulfide bond in the
structure of PFO. The location of the S190C-G57C disulfide is
shown in a ribbon representation of the -carbon backbone
of the crystal structure of the PFO monomer. The PFO domains 1 though 4 are designated as D1-D4. Residues Cys57 and
Cys190, which form the disulfide bridge, are shown as
space-filled atoms. The
-helices that comprise TMH1 and TMH2 are
shown in black and are labeled as such.
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
Relative hemolytic activity of cysteine-substituted mutants of PFO
before and after derivation with IANBD under reducing and non-reducing
conditions
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Fig. 2.
Comparison of the hemolytic activity of the
disulfide-trapped mutant under reduced and nonreduced states. The
kinetics of erythrocyte hemolysis by PFOC190-C57 was
monitored by the decrease in right-angle light scattering (17) of an
erythrocyte suspension at 37 °C in the presence
(Pre-reduced) or absence (Unreduced) of 10 mM DTT. PFOC190-C57 was exposed to 10 mM DTT for 20 min at 4 °C to completely reduce the
disulfide bond prior to being added to the pre-reduced sample. When
PFOC190-C57 (100 ng) was injected at 30 s into the
stirred erythrocyte suspension under reducing conditions
(Pre-reduced), hemolysis reached completion within 370 s of injection. However, in the absence of reducing agent
(Unreduced), no detectable hemolysis occurred. 370 s
after the addition of the oxidized PFOC190-C57, the
injection of 10 mM DTT into the unreduced sample resulted
in a nearly immediate decrease in the light scattering that was
complete within 60 s.
-barrel additionally stabilizes the oligomer to
dissociation by SDS.
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Fig. 3.
SDS-AGE analysis of PFO prepore
oligomers. The formation of oligomer by the disulfide-trapped
mutant was monitored by SDS-agarose electrophoresis.
PFOC459A (PT or parent toxin = cysteineless
PFOC459A) and the disulfide mutant (DS = PFOC190-C57) were able to form oligomers in the presence
(lanes 4-7) of phosphatidylcholine-cholesterol
liposomes under either reducing (lane 6) or
nonreducing (lanes 4 and 5)
conditions. In the absence of liposomes, both PFOC459A and
PFOC190-C57, oxidized or reduced, migrated as monomers
(lanes 1-3). Each sample was solubilized with
SDS sample buffer and separated by SDS-AGE.
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Fig. 4.
Electron micrographs of reduced and unreduced
PFOC190-C57. The oligomers formed by
PFOC190-C57 with (A) and without (B)
reducing reagent were examined by electron microscopy as described
under "Experimental Procedures." The bar represents a
distance of 100 nm.
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Fig. 5.
Oligomerization rates of
PFOC190-C57/C215 in its reduced and oxidized states.
FRET was used to examine the rate of association of
PFOC190-C57/C215 monomers in the absence (dashed
line) or the presence (solid line) of
DTT. NBD- and tetramethylrhodamine-labeled PFOC190-C57/C215
were mixed at a 1:1 molar ratio in the fluorimeter cuvette and NBD
emission at 520 nm was monitored every 20 s. After 30 s,
liposomes were injected to initiate membrane oligomerization of the
toxin. The rate of oligomerization was monitored by the decrease in
donor emission as a function of time. In both cases, any
non-FRET-dependent change in intensity of the donor was
subtracted from the experimental results. F0 is
the initial NBD intensity prior to liposome addition, and F
is the NBD emission intensity at time t. To illustrate the
acceptor dependence of the FRET, the acceptor-labeled toxin was
replaced with unlabeled PFOC190-C57/C215 in one sample
containing oxidized PFOC190-C57/C215 (dotted
line). The data were normalized to between 0 and 1 to
compare the rate of oligomerization of each species of PFO.
-turn
of the hairpin, the NBD appeared to enter a somewhat more nonpolar
environment, but not the environment observed when the protein is
allowed to fully insert. It therefore appears that a probe at the other
end of TMH1 from the disulfide bond may interact to some extent with
the bilayer, but cannot insert properly into the membrane. However,
upon reduction of the disulfide, the probe at residue 202, as well at
the other residues, fully entered the membrane because the fluorescence intensities of the NBD dyes located at these four positions increased to approximately the same levels as reported previously for these residues as they enter the membrane (7, 8).
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Fig. 6.
Membrane insertion of the TMHs of
disulfide-trapped PFOC190-C57 under reducing and
nonreducing conditions. The insertion of TMH1 or TMH2 in the
disulfide-trapped PFOC190-C57 oligomer was monitored by the
change in fluorescence intensity of an NBD dye attached to a cysteine
residue substituted in PFOC190-C57 for Ala215
or Val202 in TMH1, or Lys288 or
Ile303 in TMH2. The fluorescence intensity of each
NBD-labeled PFO mutant was determined as a monomer in solution
(solid line), and also as the membrane-bound
toxin after the addition of phosphatidylcholine-cholesterol liposomes
in the presence (dotted line) and absence
(dashed line) of 10 mM DTT. A large
increase in the emission intensity of the NBD at positions 202, 215, 288, and 303 reflects the insertion of the transmembrane -hairpins
into the nonpolar environment of the membrane (7, 8).
View larger version (17K):
[in a new window]
Fig. 7.
Insertion kinetics of the
PFOC190-C57 transmembrane
-sheet. The rates of insertion of the
transmembrane
-sheets of the disulfide-trapped PFO were followed by
monitoring the increase in fluorescence intensity of NBD covalently
attached to residue 215 in TMH1 (PFOC190-C57/C215-NBD)
(upper panel) or in TMH2 at residue 288 (PFOC190-C57/C288-NBD) (lower panel).
Residues 215 and 288 both face the membrane in the assembled pore (7,
8) and are excellent indicators of the insertion of TMH1 and THM2,
respectively, into the bilayer. Liposomes were injected at 30 s
into a stirred mixture at 37 °C that contained nonreduced PFO
derivative (dashed or dotted line) or
PFO derivative that had been previously reduced with 10 mM
DTT (solid line). The unreduced samples were then
reduced by the addition of DTT at either 130 s (dashed
line) or 370 s (dotted line)
after the addition of liposomes.
-sheet of the prepore complex at 120 or 370 s, the rate of
insertion then decreased to a rate comparable to that of the pre-reduced sample. Therefore, the fraction of the PFO that had been
converted to the prepore complex rapidly inserted its TMHs into the
membrane, while the fraction that had not yet formed the prepore was
much slower to insert its TMHs because it had not yet fully
oligomerized into the insertion-competent prepore stage. These data
demonstrate that TMH insertion occurs more rapidly than prepore complex
assembly, and hence that the rate of prepore formation dictates the
observed rate of pore formation. These results are therefore consistent
both with the hemolysis kinetics (Fig. 2) and with the faster rate of
pore formation observed when disulfide-arrested prepore complexes are
released by DTT (14).
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-hemolysin from Staphylococcus aureus.
The prepore presumably facilitates the insertion of the membrane
spanning
-barrel, although this has not been directly demonstrated.
By quantitatively interrupting the pore formation process and
synchronizing PFO in its prepore state, we have been able to determine
that the assembly of the prepore complex is the rate-limiting step in
cytolysis and that, once the prepore is formed, the insertion of the
transmembrane
-sheet is rapid. This observation is not only
interesting in terms of defining a role of the prepore, it is
fascinating that it occurs in the CDCs, which form channels that are
substantially larger than most other pore-forming toxins and may be
composed of up to 50 monomers (reviewed in Ref. 20). The central cavity
of the CDC pore is ~25 nm in diameter, sufficiently large to permit
the passage of large macromolecules such as proteins (reviewed in Refs.
21 and 22) and even DNA (23).
-sheet (6, 13, 24). Two models have been proposed to
explain the insertion of the CDC
-sheet, one based on the prepore
mechanism that has been shown to mediate the formation of small pores
by several toxins (9-12) and the other on the gradual enlargement of a
small oligomer and pore into a large oligomer and pore by the
sequential addition of monomers to the inserted complex (24). However,
recent studies by Shepard et al. (13) showed that the
processes of PFO oligomerization and the insertion of the transmembrane
-sheet could be uncoupled, suggesting that a prepore mechanism might
be the modus operandi for the CDCs. The results of the current studies
strongly support the formation of a prepore as the penultimate step in
the insertion of the transmembrane
-sheet of PFO (see Fig.
8).
View larger version (56K):
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Fig. 8.
Hypothesized prepore mechanism of PFO.
Hypothesized insertion mechanism of PFO based on the available
experimental data. In panel A, soluble PFO
monomers are shown binding the membrane. Domain 4 has recently been
shown to interact first with the membrane surface (14), and the highly
conserved undecapeptide in domain 4 is shown in cyan in
panel A. A cholesterol-rich region in the
membrane is shown in red. In panel B,
the monomers form an oligomer and in panels C and
D, the -helices of the monomers are shown to extend into
two
-hairpins (TMH1 in purple and TMH2 in
green) to form the
-sheet of the prepore. In
panels B and C, the transmembrane
-hairpins are still shown in their
-helical structures; however;
in panel C, domain 3 is predicted to begin to
twist clockwise so that both TMHs can begin to unfold from their
-helical structure and likely enter into a disordered intermediate
structure (data not shown). The disulfide bridge mutant would be stuck
in the state shown in panel B or C.
The timing of the TMHs transition into
-hairpins remains unknown,
but we propose, as shown in panel D, that a
-sheet may be formed prior to its insertion into the membrane. In
panel E, the prepore is then converted to the
pore complex by the insertion of the
-sheet into the membrane. The
prepore structure and pore structure remain hypothetical at this time,
except it has been established that TMH1 and TMH2 span the bilayer as
amphipathic
-hairpins (7, 8). To illustrate the hypothesized
structural changes, only a section of the oligomer is shown, but up to
50 monomers participate in the formation of the entire ring-shaped,
pore-forming oligomeric complex.
By introducing a disulfide bridge between residues 190 and 57 to impede the movement of TMH1, we found that we could abrogate PFO-dependent hemolysis. Yet, if the disulfide bridge was reduced after the PFO had been incubated with erythrocytes, hemolysis proceeded at a significantly faster rate than if the PFO had been reduced prior to its addition to the erythrocytes. The typical lag period that we observed when the disulfide of this mutant was reduced prior to the addition of the erythrocytes, and which had been previously observed for PFO-dependent hemolysis (17), was nearly abolished when the disulfide-trapped PFO was preincubated with the erythrocytes prior to reduction of the disulfide bridge. This observation suggested that the disulfide trapped PFO had accumulated at a step that primed it for insertion and pore formation on the erythrocytes, presumably a prepore complex.
Analysis of the disulfide-trapped mutant by electron microscopy and by
SDS-AGE revealed that PFOC190-C57 formed the same oligomers
on liposome membranes whether or not the disulfide was oxidized
(hemolytically inactive) or reduced (fully hemolytic). The oxidized and
reduced oligomer exhibited similar mobilities in SDS-AGE gels (Fig. 3),
although the disulfide-trapped prepore appeared to be somewhat less
stable to the SDS. Therefore, the interaction of the transmembrane
-hairpins in the pore complex helps stabilize the oligomer since the
inserted form of PFOC190-C57 exhibited the same stability
as the cysteine-less parent toxin PFOC459A. The stability
of the oligomeric complex of various pore-forming toxins to
dissociation by SDS has been well documented for the heptameric pore
complexes of S. aureus
-hemolysin, C. septicum
-toxin, and A. hydrophila aerolysin (9-12) Additionally,
the rate of oligomerization, as determined by FRET, of the oxidized and
reduced forms of PFOC190-C57 was unaffected by the presence
of the disulfide bond. Therefore, changes in the oligomerization rate
or the extent of oligomer formation could not account for the
cytolytically inactive phenotype exhibited by oxidized
PFOC190-C57.
PFO is novel in that it contributes two transmembrane -hairpins per
monomer to the formation of the membrane-spanning
-sheet (7, 8),
whereas other pore-forming toxins such as S. aureus
-hemolysin and anthrax protective antigen utilize a single TMH per
monomer (11, 25). PFO is also unique in that each membrane-spanning
-hairpin in the PFO pore complex exists originally as three short
-helices in domain 3 of the soluble monomer (7, 8). In the soluble
monomer, the TMH1
-helices have a large percentage of their
structure buried against domain 2 and the core
-sheet of domain 3, whereas the
-helices that comprise TMH2 are mostly solvent-exposed
(6). Therefore, it was possible that TMH2 could have partially or
wholly inserted into the bilayer even though the movement of TMH1 was
restricted by the disulfide bond between Cys190 and
Cys57 in oxidized PFOC190-C57. We had
previously proposed that the domain 3 core
-sheet must both move
away from its domain 2 and domain 3 contacts and twist for both TMHs to
unravel from their
-helical structure in the monomer and form the
extended transmembrane
-hairpins (7). By restricting the movement of
TMH1 with a disulfide bond, the required movement of the domain 3 core
-sheet may also have been limited. NBD probes located on
membrane-facing residues either near the predicted
-turn in TMH2 or
within one of its membrane-spanning
-strands demonstrated that TMH2
also did not enter the membrane. Therefore, the disulfide bridge
between TMH1 and domain 2 prevented the insertion of both TMH1 and TMH2
into the membrane. These observations indicate that the PFO TMHs do not
insert independently into the membrane and that their insertion is
likely coupled by hydrogen bonding between the hairpins and one or more
prerequisite conformational changes in domain 3.
Since the rate and extent of oligomer formation were largely unaffected by the oxidation state of PFOC190-C57 and the inability of TMHs to insert into the membrane, PFOC190-C57 was effectively trapped on the membrane surface in a fully oligomerized, uninserted state that, by definition, corresponded to a prepore complex. In this state, neither of the TMHs in the disulfide-trapped prepore complex were embedded in the membrane, even though both TMHs were capable of normal insertion into the membrane upon reduction of the Cys190-Cys57 disulfide bond. Therefore, the disulfide linkage effectively allowed us to synchronize the majority of the molecules into a single state, the prepore. By allowing the oxidized PFOC190-C57 mutant to form a prepore complex on liposomal membranes prior to reduction, we could dramatically increase the rate of membrane insertion of both TMH1 and TMH2 (Fig. 7). This dramatic increase in the rate of insertion of the TMHs also corresponded nicely with the increased rates of hemolysis of erythrocytes (Fig. 2) and of pore formation (14) that were also observed when oxidized prepore complexes were allowed to form prior to DTT addition. Thus, TMH insertion, pore formation, and hemolysis proceeded rapidly upon reduction of the disulfide bond because the majority of the PFO molecules were already assembled into the prepore state and therefore primed for synchronous insertion into the membrane.
These results are consistent with the prepore model (Fig. 8) because
the insertion of the transmembrane -sheet should occur more rapidly
if the prepore complex is first allowed to form on the membrane before
insertion is initiated. It had been suggested previously that a
significant energy barrier might prevent the simultaneous insertion of
the large
-sheet of the CDCs, and that the synchronized insertion of
so many
-hairpins would make insertion of the CDCs via a
prepore-based mechanism unlikely (24). However, by arresting the pore
formation process after the assembly of the oxidized
PFOC190-C57 prepore complex and then synchronizing the
insertion of the transmembrane
-hairpins, upon addition of reducing
agent, we have shown that quite the opposite effect is observed;
formation of the prepore significantly enhances the rate of insertion
of the transmembrane
-sheet. Since the membrane insertion of the
large
-sheet of the prepore complex is far more rapid if the prepore
is first allowed to assemble, the rate-limiting step in cytolysis is a step involved in the formation of the prepore complex, not the insertion of the transmembrane
-hairpins.
PFO pore formation therefore involves the assembly of a prepore complex
that, once an insertion-competent size is reached, can initiate
membrane insertion of the membrane-spanning -sheet. The exact size
of an insertion competent prepore complex is currently unclear,
although based on SDS-AGE, EM, and planar bilayer analyses shown here
and elsewhere (13), it must be significantly larger than the heptameric
structures observed for pore-forming toxins such as
-hemolysin (11),
aerolysin (26, 27) and anthrax protective antigen (25). Shepard
et al. (13) observed that the channels formed by PFO in
planar bilayers exhibited conductances of 4-6 nanosiemens, which are
20-40 times larger than that observed for the smaller toxin pores (28,
29). However, whether or not a complete ring must form prior to
membrane insertion of the transmembrane
-sheet remains to be
determined unambiguously.
By synchronizing the insertion of the prepore -hairpins, we have
been able to provide direct evidence that prepore complex formation
facilitates the insertion of the transmembrane
-sheet for a
pore-forming toxin. Therefore, the prepore state is an important intermediate in the assembly and insertion of the transmembrane
-sheet of the CDCs. In addition, the data support the remarkable concept that the PFO oligomer complex can possibly coordinate the
simultaneous insertion of up to 100
-hairpins to form the transmembrane
-sheet. Therefore, the prepore mechanism is
sufficiently flexible to accommodate the insertion of both large (PFO
and the CDCs) and small (
-hemolysin, anthrax protective antigen)
membrane-spanning
-sheets.
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FOOTNOTES |
---|
* This work was supported by National Institutes of Health Grant AI37657 (to R. K. T.) and by the Robert A. Welch Foundation (to A. E. J.).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.
To whom correspondence should be addressed: Dept. of
Microbiology and Immunology, BMSB, Rm. 1053, 940 Stanton L. Young
Blvd., University of Oklahoma Health Sciences Center, Oklahoma City, OK
73190. Tel.: 405-271-2133; Fax: 405-271-3117; E-mail:
rod-tweten@ouhsc.edu.
Published, JBC Papers in Press, December 1, 2000, DOI 10.1074/jbc.M009865200
2 J. Rossjohn, M. W. Parker, and R. K. Tweten, unpublished data.
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ABBREVIATIONS |
---|
The abbreviations used are:
PFO, perfringolysin
O;
TMH, transmembrane -hairpin;
IANBD, N,N'-dimethyl-N-(iodoacetyl)-N'-(7-nitrobenz-2-oxa-1,3
diazolyl)ethylenediamine;
IATR, tetramethylrhodamine-5(and
6)-iodoacetamide;
FRET, fluorescence resonance energy transfer;
AGE, agarose gel electrophoresis;
DTT, dithiothreitol;
NBD, N'-(7-nitrobenz-2-oxa-1,3-diazolyl)ethylenediamine;
CDC, cholesterol-dependent cytolysins;
EM, electron
microscopy;
PFOC190-C57/C303, an example of the
nomenclature for a derivative of PFOC459A (the
cysteine-less derivative of PFO) in which residues 190 and 57 have been
substituted with cysteines that form a disulfide (the residue after the
slash indicates a residue that has been replaced by a cysteine that is
not in a disulfide and therefore has a free sulfhydryl group).
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REFERENCES |
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