The Cholesterol-Dependent Cytolysin Signature Motif: A
Critical Element in the Allosteric Pathway that Couples
Membrane Binding to Pore Assembly
Kelley J. Dowd, Rodney K. Tweten*
Department of Microbiology and Immunology, The University of Oklahoma Health Sciences Center, Oklahoma City, Oklahoma, United States of America
The cholesterol-dependent cytolysins (CDCs) constitute a family of pore-forming toxins that contribute to the pathogenesis
of a large number of Gram-positive bacterial pathogens.The most highly conserved region in the primary structure of the
CDCs is the signature undecapeptide sequence (ECTGLAWEWWR). The CDC pore forming mechanism is highly sensitive to
changes in its structure, yet its contribution to the molecular mechanism of the CDCs has remained enigmatic. Using a
combination of fluorescence spectroscopic methods we provide evidence that shows the undecapeptide motif of the
archetype CDC, perfringolysin O (PFO), is a key structural element in the allosteric coupling of the cholesterol-mediated
membrane binding in domain 4 (D4) to distal structural changes in domain 3 (D3) that are required for the formation of the
oligomeric pore complex. Loss of the undecapeptide function prevents all measurable D3 structural transitions, the
intermolecular interaction of membrane bound monomers and the assembly of the oligomeric pore complex. We further
show that this pathway does not exist in intermedilysin (ILY), a CDC that exhibits a divergent undecapeptide and that has
evolved to use human CD59 rather than cholesterol as its receptor. These studies show for the first time that the
undecapeptide of the cholesterol-binding CDCs forms a critical element of the allosteric pathway that controls the assembly
of the pore complex.
Citation: Dowd KJ, Tweten RK (2012) The Cholesterol-Dependent Cytolysin Signature Motif: A Critical Element in the Allosteric Pathway that Couples Membrane
Binding to Pore Assembly. PLoS Pathog 8(7): e1002787. doi:10.1371/journal.ppat.1002787
Editor: Theresa M. Koehler, The University of Texas-Houston Medical School, United States of America
Received April 6, 2012; Accepted May 19, 2012; Published July 5, 2012
Copyright: ? 2012 Dowd, Tweten. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits
unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: These studies were supported by a grant to RKT from the National Institutes of Health, NIAID (AI037657). The funders had no role in study design, data
collection and analysis, decision to publish, or preparation of the manuscript.
Competing Interests: The authors have declared that no competing interests exist.
* E-mail: Rodney-Tweten@ouhsc.edu
The cholesterol-dependent-cytolysin (CDC) family of toxins
consists of over 25 members that are produced by many different
species of Gram-positive bacterial pathogens  and contribute
in various ways to the pathogenesis of these organisms [2,3,4,5].
Members of this family exhibit high levels of homology in their
primary structures (40–70%) and in the crystal structures of their
soluble monomers [6,7,8,9]. The region within the CDC primary
structure that exhibits the highest degree of sequence identity is
an 11-residue peptide known as the undecapeptide or trypto-
phan-rich motif, which is located near the C-terminus of the
molecule in domain 4 (D4) (Fig. 1). The undecapeptide
(ECTGLAWEWWR) is the signature motif for the CDCs 
and so proteins exhibiting this peptide sequence have a high
probability of belonging to the CDC family. The pore forming
mechanism of the CDCs that use cholesterol as their receptoris
highly sensitive to changes in the primary structure of the
undecapeptide [6,10,11,12,13,14,15]. These studies suggest that
the undecapeptide plays an important role in the CDC pore-
forming mechanism, yet since Iwamoto et al.  began studying
the effects of chemically altering the undecapeptide in 1987 its
contribution to the pore forming mechanism of the CDCs has
The undecapeptide is located at the tip of D4 of the CDC
structure, as shown in the structure of the CDC produced by
Clostridium perfringens, perfringolysin O (PFO) (Fig. 1). D4 also
contains the cholesterol recognition/binding motif (CRM) and two
other short loops (L2 and L3) near the undecapeptide (reviewed in
). Upon recognition of membrane cholesterol by the
CRM,loops L2 and L3 insert into the membrane. These
interactions anchor the monomers in a perpendicular orienta-
tionto the membrane surface where the tip of D4 is anchored to
the membrane surface and the top of D3 resides about 113 A˚
above the membrane surface [18,19,20]. Although the sidechains
of several residues of loops L2 and L3 and the undecapeptide
insert into and anchor the monomers to the membrane they do
not penetrate deeply into the bilayer core [19,21].
It had been generally accepted in the field that the undecapep-
tide motif wasthe CRM of the CDCs, although this function had
never been demonstrated unambiguously. An early study by
Iwamoto et al.  showed that chemical modification of the
undecapeptide cysteine caused independent defects in both
binding and pore formation. Since that time it has been shown
that mutation of many of the undecapeptide residues often affects
both binding and pore formation [6,10,11,12,13,14,15]. We
recently showed, however, that the CRMresides in the nearby
D4 loop L1 (Fig. 1) and is comprised of a threonine-leucine pair
that is strictly conserved in all known CDCs . Upon
cholesterol binding by the CRM the nearby loops L2 and L3
and the conserved undecapeptide insert into the bilayer surface
PLoS Pathogens | www.plospathogens.org1July 2012 | Volume 8 | Issue 7 | e1002787
and anchor the monomer in a perpendicular orientation to the
membrane surface [19,21,23,24]. Membrane binding in conjunc-
tion with monomer-monomer interactions  initiates and drives
a dramatic series of secondary and tertiary structural changes in
D3, which is about 60 A˚distant from the tip of D4 (Fig. 1). These
structural changes are necessary for the assembly of the membrane
bound monomers into the large oligomeric pore complex
[23,24,25,26,27]. Soluble monomers of PFO do not exhibit these
D3 structural changes, even at the high concentrations required
for crystallization of the protein : membrane binding is
required to initiate the structural changes in D3 [20,25].
As indicated above, the pore-forming mechanism of PFO-like
CDCs is highly sensitive to mutations in the undecapeptide
[6,10,11,12,13,14,15]. Furthermore, the conformational changes
in the PFO undecapeptide, reflected by the membrane insertion of
its tryptophan residues, are conformationally coupled to the
structural changes in TMH1 required for the formation of the b-
barrel pore . This observation suggests that the undecapeptide
of PFO is involved in the allosteric coupling of membrane binding
to the initiation of the D3 structural changes that are necessary for
monomer-monomer interaction and the formation of the oligo-
meric b-barrel pore complex.
A small family of CDCs, typified by Streptococcus intermedius
intermedilysin (ILY) use human CD59 as their receptor, rather
than cholesterol [29,30,31]. The D3 structural changes in ILY can
be initiated by binding to human CD59 in membranes that are
largely, though not completely depleted of cholesterol . ILY
still requires a CRM-mediated membrane interaction with
cholesterol to maintain its anchor to the membrane surface (it
disengages from CD59 during prepore to pore conversion
[22,33]), but it remains unclear if cholesterol binding also
participates in initiation of the D3 structural changes necessary
for assembly of the oligomer pore complex. Interestingly, in
contrast to the CDCs that use cholesterol as their receptor, the
pore forming mechanism of ILY is comparatively insensitive to
mutations within its undecapeptide , which suggests that it may
not play as significant of a role in the pore forming mechanism of
In the present study we performed a detailed molecular analysis
of a point mutation in the undecapeptide of PFO that reduces its
pore-forming activity 100-fold, whereasthe analogous mutation
has no significant effect on the mechanism of ILY . In PFO this
mutant blocks all measurable structural transitions in D3 and
prevents the stable interaction of membrane-bound monomers.
We further show that the effect of this mutation on the activity of
PFO is similar to that observed for cholesterol bound native ILY in
the absence of CD59. These results show that the undecapeptide
of PFO is a critical structure within the allosteric pathway of PFO
that couples cholesterol binding to the initiation of structural
changes within D3, which lead to the formation of the b-barrel
pore. We further show that this pathway appears to be missing in
the CD59-binding ILY, so that assembly of its pore complex is
initiated by its interaction with CD59 rather than cholesterol.
Cytolytic activity of PFO mutated at Arg-468
Arg-468 is the last residue of the PFO undecapeptide
(ECTGLAWEWWR), as well as in the ILY undecapeptide
(GATGLAWEPWR). Substitution of the PFO undecapeptide at
this residue with alanine decreases its hemolytic activity 100-fold
(Table 1), whereas substitution of the analogous residue in ILY has
little effect on the activity . A series of mutants were generated
for Arg-468 of PFO to examine the effects of size, length and
charge of the residue atposition 468 on the hemolytic activity of
PFO (Table 1). Neither conservative nor non-conservative
substitutions were tolerated: all substitutions decreased hemolytic
activity $100-fold. Based on the crystal structure of PFO the only
intramolecular contacts established by Arg-468 are hydrogen
bonds between its sidechain NH1 and the CRM carbonyls
(Fig. 1B). Interestingly, this contact is lost in the ILY monomer
(Fig. 1C), which presumably results from differences in its
undecapeptide structure . We selected the PFOR468Amutant
for further studies into the defect(s) induced by substitution of the
Arg-468 residue on the PFO pore-forming mechanism.
We have shown that a conserved Thr-Leu pair in Loop 1, and
not the undecapeptide, is responsible for CDC binding to
membrane cholesterol , yet mutations within the conserved
undecapeptide were often observed to affect binding [6,13,34]. To
confirm that the loss of hemolytic activity by the PFOR468Amutant
was not due solely to a defect in binding we examined the ability of
the mutant to bind to human RBCs by flow cytometry. In order to
prevent cell lysis at high concentrations of toxin, derivatives of
native PFO and PFOR468Awere generatedin which an engineered
disulfide was introduced between residuesThr-319 in b4 and Val-
334 in b5 that prevent the rotation of b5 away from b4 in domain
3 (PFOb4b5and PFOR468ANb4b5). The engineered disulfide there-
forepreventsthe formation of a functional pore by blocking the
intermolecular interaction of b1 of one monomer with b4 of
another monomer . A third cysteine was substituted at residue
Asp-30 in both mutants, which is at the amino terminus of PFO, so
that specific fluorescent probes could be introduced into these
The CDCs are a large family of pathogenesis-associated
pore-forming toxins that are expressed by many Gram-
positive pathogens. The conserved undecapeptide motif
of the CDCs has been regarded as the signature peptide
sequence for these toxins, yet its function has remained
obscure. The studies herein show that the undecapeptide
forms a critical structural element in the allosteric pathway
that couples membrane binding to cholesterol to the
initiation of distal structural changes, which are required
for the assembly of the pore forming complex. These
studies provide the first insight into the function of this
highly conserved sequence and show that through
evolution this pathway is missing in the CD59-binding
Table 1. Cytolytic activity of PFO derivatives with mutations
in the Arg-468 residue.
Toxin EC50(M)% WT
Cytolytic activity of PFO and Arg-468 mutants is shown as the effective
concentration (EC50) of toxin required for 50% lysis of human erythrocytes
under standard assay conditions (see Materials and Methods for details).
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mutants. This mutation does not affectthe structure of PFO or its
At the highest concentration of PFOR468Awe observed about a
50% decrease in binding to hRBCs compared to PFO, although at
lower concentrations this difference was greater (Fig. 2A). The
decrease in binding, however, doesn’t account for the 100-fold
decrease in cytolytic activity. PFO follows an ordered series of
coupled conformational changes that are initiated by binding
[19,20,23,25], therefore the major defect induced by the
PFOR468Amutation affects an event after binding, which then
prevents formation of the pore complex.
Oligomerization of PFOR468A
Upon membrane binding PFO monomers oligomerize into
large SDS-resistant prepore complexes containing approximately
36 monomers . An oligomerization assay was performed with
the PFOR468Amutant. Due to the lower binding affinity observed
in Fig. 2A we increased the concentration of human red blood cells
(hRBCs) to ensure complete binding of PFOR468A. The concen-
tration of hRBCs in the binding studies in Fig. 2A was maintained
at 46106/ml whereas in the oligomerization assay shown in
Fig. 2B the concentration hRBCs ranged from 2.56107to
Figure 1. The molecular structures of PFO and ILY. Shown in A is a ribbon representation of the crystal structure of PFO . The domain 3 b5
strand and associated a-helix (a1) that swing away from b4 are highlighted in red. The locations of Asn-197 (at the D2–D3 interface) and Val-322
(buried under the loop formed by a1b5) are shown in space-filled atoms. The twin a-helical bundles in D3 (cyan) extend into the twin transmembrane
b-strands (TMHs). The conserved undecapeptide is shown in blue in D4. In panel B an enlarged view of the conserved undecapeptide loop and the
CRM containing loop L1 of PFO is shown. In Panel C the analogous structures are shown for ILY as are shown in panel B for PFO.All structures were
derived from the crystal structures of PFO and ILY [6,7]). In panel D we show the structural changes in PFO as it makes the transition from the bound
monomer state to the membrane embedded oligomer. The membrane embedded monomer structure is based on the 3D reconstruction of the
pneumolysin pore fitted with the PFO crystal structure . D3 breaks its contacts with D2 and swings out in order to extend the a-helical bundles (in
cyan) into the twin TMHs. This transition also repositions Asn-197 from the D2–D3 interface to a solvent exposed position within the lumen of the
membrane pore. Prior to or simultaneously with the disruption of the D2–D3 interface the a1b5 loop (red) swings away from b4 thus exposing the
edge b4 (as well as exposing Val-322 to the solvent), which can then pair with b1 of a second monomer. Upon transition to the pore the oligomeric
complex undergoes a 40 A˚vertical collapse to insert the b-barrel pore into the bilayer . After b5 breaks contact with b4 the location of a1b5 loop
is not known, its position in the model is for illustrative purposes only. All structures were generated using VMD .
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2.56108/ml, which wasapproximately 6–60 fold higher than the
concentration used in the flow cytometry assay. After the toxins
were allowed to bind, the samples were solubilized with SDS
without heating and separated by SDS-agarose gel electrophoresis
(SDS-AGE), which separates the monomer and oligomer forms
. Native PFO readily formed SDS-resistant oligomers at all
concentrations of RBCs (Fig. 2B) whereas PFOR468Adid not form
detectable levels of SDS-resistant oligomers (Fig. 2B). Therefore,
the major defect in the PFOR468Apore-forming mechanism
follows binding and prevents the formation of an SDS-stable
Rotation of b5 away from b4
Several structural transitions in domain 3 are initiated by
membrane binding, which are required for oligomerization and
pore formation [25,26,27]. One of these structural transitions is
the rotation of b-strand 5 (b5) away from the adjacent b-strand 4
(b4) of the core b-sheet in domain 3 (Fig. 1A), whichcontributes to
the formation of the SDS-resistant prepore oligomer .
Rotation of b5 away from b4 allows the formation of edge-on
hydrogen bonds between the peptide backbones of b4 and b1 of
two membrane-bound monomers .
The disruption of the b4/b5 interaction can be followed
spectroscopically using the environmentally sensitive fluorescent
probe, NBD (7-nitrobenz-2-oza-1,3 diazole) , which is
positioned on the sulfhydryl of a cysteine substituted for Val-322
in b4 (Fig. 1A). Val-322 is buried under the residues of b5 and so a
probe positioned here is in a hydrophobic pocket. The fluores-
cence emission of NBD is quenched by water, therefore as b5
rotates away from b4 the NBD positioned in b4 moves from a
nonpolar to polar environment, which results in a decrease in its
fluorescence emission intensity as it is exposed to the aqueous
milieu . The PFOR468ANV322C-NBDmutant exhibited virtually
no change in the NBD emission compared to functional
PFOV322C-NBD(Fig. 3). These results show that the rotation of
b5 away from b4 does not occur in membrane bound PFOR468A.
Monomer-monomer interaction of PFOR468A
The studies above show that PFOR468Adoes not form SDS-
resistant oligomers, which is likely due to the loss of the
intermolecular b1–b4 interaction of monomers. This observation,
however, did not rule out the possibility that PFOR468Amonomers
could still form a SDS-sensitive oligomer. To determine whether
PFOR468Aformed SDS-sensitive oligomers the PFOR468Amono-
mer association was examined using fluorescence resonance
energy transfer (FRET). A cysteine was substituted for the amino
terminal Asp-30 and labeled with either donor fluorophore (D)
(Alexa Fluor 488) or acceptor fluorophore (A) (Alexa Fluor 568). A
mixture containing a 4:1 molar ratio of A-labeled PFOR468Aor
unlabeled PFOR468A(U) to D-labeled toxin was incubated with
membranes and fluorescence emission intensity of D was
When membrane-bound PFO monomers associate to form the
prepore oligomer the distance (R0) between D and Afluorescent
dyes on the monomers decreases, which results in the FRET-
dependent quenching of the D emission (R0is typically,10 nm)
. As expected, we observed an A-dependent quenching of the
D emission for functional PFO as it oligomerized [35,37], whereas
nochangein the donor fluorescence
PFOR468A(Fig. 4).This result shows that the PFOR468Amonomers
do not interact, or only form transient interactions that cannot be
detected by FRET. FRET requires the donor and acceptor pair be
at a fixed distance during the lifetime of the donor emission, which
for Alexa-488 is approximately 4 ns . Therefore the PFOR468A
monomers are, at most, only interacting briefly within a timeframe
that is shorter than the fluorescence lifetime of the Alexa dye.
The status of the domain 3 TMHs in PFOR468A
The D2–D3 interface is disrupted in order to extend the D3 a-
helical bundle into transmembrane b-hairpin 1 (TMH1) ,
which together with TMH2 ultimately contribute to the formation
of the membrane spanning b-barrel pore [26,27]. First, the a-
helical bundle that forms TMH1 must break its interaction with
D2 to unravel and form the extended b-hairpin structure, which
eventually inserts into the bilayer as part of the b-barrel pore .
Disruption of the TMH1 contact with D2 can be measured by
Figure 2. Binding and oligomerization of PFO and PFOR468A. (A) Binding of PFOb4b5and PFOR468ANb4b5to human RBCs (46106/ml in a final
volume of 0.5 ml) was measured by flow cytometry. The disulfide locked b4b5 versions of each protein  were used to prevent the lysis of the RBCs
during flow cytometry. (B). Oligomerization of PFO and PFOR468A(both toxins were maintained at 440 nM)on human RBCs (concentrations ranged
from 2.56107/ml to 2.56108/ml in a final volume of 40 ml) was determined using SDS-agarose gel electrophoresis (SDS-AGE) and the proteins were
detected with anti-PFO antibody after transfer to nitrocellulose paper. The analyses are representative of at least 3 experiments.
Figure 3. Disruption of the b4b5 interface of PFO and PFOR468A.
A cysteine was substituted for Val-322, located in the D3 b4 strand. Each
derivative was labeled with NBD and incubated in the presence (dashed
line) and absence (solid line) of human erythrocyte ghost membranes.
The fluorescence emission intensity of NBD was measured from 500–
600 nm. The data are representative of 3 experiments.
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placing a NBD probe on a cysteine substituted for Asn-
197inTMH1 (Fig. 1). Asn-197 resides at this interface and
undergoes a nonpolar to polar transition as the a-helical bundle
breaks contact with D2 and unravels to form the extended b-
hairpin . The subsequent insertion of the b-barrel pore can be
followed by placing a NBD probe on cysteine-substituted Ala-215
in TMH1, which undergoes a polar to nonpolar transition as its
sidechain inserts into the bilayer core .
As expected, the fluorescence emission of the NBD probe on
cysteine substituted Asn-197 in native PFO decreases to less than
25% of its initial value as the a-helical bundle disengages from its
interface with D2 (Fig. 5A, left panel). Also, as expected, the
fluorescence emission of the NBD probe located at position 215 in
TMH1 of PFO increases as it makes the transition from its polar
environment in the soluble monomer to its membrane embedded
position in the b-barrel pore (Fig. 6A, left panel). In contrast, little
change was detected in the fluorescence emission of the NBD
probe at both locations in membrane bound PFOR468ANN197C-NBD,
showing that TMH1 did not disengage from its interface with D2
(Fig. 5A, right panel) and insert into the membrane (Fig. 6A, right
Native PFO drives the membrane insertion of the
As shown above, the membrane-bound monomers of PFOR468A
do not interact and the D3 structural transitions that lead to the
insertion of the b-barrel pore do not occur in PFOR468A: in essence
the monomers remain inert after binding. Therefore, we next
determined whether functional PFO could form chimeric oligo-
mers with PFOR468Aand drive these structural transitions.
The same experiments were performed as in Figs. 5A and 6A
except that a 4:1 ratio of unlabeled PFO or PFOR468Awas mixed
with the labeled species prior to their addition to the liposomes. As
expected, the relative emission intensity of the NBD probe
was similar when each fluorescence species was mixed with a
4-molar excess of the unlabeled homologous protein. For
PFON197C-NBDcompare the left panels of Fig. 5A and Fig. 5B
and for PFOA215C-NBDcompare the left panels of Fig. 6A and
Fig. 6B. Similarly, no change was observed in the NBD emission
for PFOR468ANN197C-NBD(compare Fig. 5A, right panel to Fig. 5B
center panel) and for PFOR468ANA215C-NBD(compare Fig. 6A, right
panel to Fig. 6B center panel) when they were mixed with a
4-molar excess of unlabeled PFOR468A.
However, when a 4-molar excess of unlabeled PFO was mixed
with the NBD-labeled species of PFOR468Ait drove the disruption
of the D2–D3 interface and insertion of the b-barrel pore. We
observed the expected decrease in the fluorescence emission of the
NBD probe located at the D2–D3 interface (compare the right and
left panel of Fig. 5), as b5 swings away form b4. Also, the relative
emission intensity increased as the probe located at position 215
inserted into the bilayer (compare the right and left panels in
Fig. 6). Furthermore, the change in the emission intensity of the
NBDin PFOR468ANA215C-NBDwhen mixed with a 4-molar excess of
PFO was quantitatively similar to that observed for PFOA215C-NBD
alone or mixed with the unlabeled PFO. Therefore, nearly all of
the PFOR468ANA215C-NBDTMHs were converted to a membrane-
These results show that functional PFO can form sufficient
intermolecular contacts with PFOR468Ato efficiently drive the
Figure 4. FRET-detected monomer association of PFO and
PFOR468A. A cysteine was substituted for Asp-30, located in domain 1
and the derivatives were labeled with Alexa Fluor 488 (donor, D) or
Alexa Fluor 568 (acceptor, A).A 4:1 molar ratio of A-labeled PFOR468A
(dashed line) or unlabeled PFOR468A(U; solid line) to D-labeled toxin was
incubated in the presence of human erythrocyte ghost membranes and
fluorescence emission intensity of D was measured.
Figure 5. Disruption of the D2/D3 interface in PFO and PFOR468A. (A) A cysteine was substituted for TMH1 residue Asn-197, which is located
within the D2/D3 interface. Each derivative was labeled with NBD and incubated in the presence (dashed line) and absence (solid line) of human
erythrocyte ghost membranes. If TMH1 breaks its contact with D2 then Asn-197 moves from a buried, nonpolar location at the interface with D2 to
the lumen of the pore. An NBD positioned at this location will therefore undergo a nonpolar to polar transition, which results in the quenching of the
fluorescence emission. (B) Unlabeled native PFO or PFOR468Awere mixed at a 4:1 molar ratio with PFON197C-NBDand PFOR468ANN197C-NBDderivatives. The
fluorescence emission intensity of NBD was measured from 500 to 600 nm. These data are representative of 3 experiments.
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disruption of its domain 2–3 interface and the membrane insertion
of its b-barrel. Hence, PFOR468Ais competent to undergo the
necessary D3 structural changes and insert its TMHs into the
membrane, but is unable to initiate these changes because it is
missing the allosteric signal that is initiated by membrane binding.
These data also indicate that the rate of binding of the PFOR468A
monomers to the membrane surface is not significantly different
from that of the native PFO monomers, otherwise the PFO
monomers would preferentially interact with each other before
interacting with PFOR468A, which would have resulted in a less
efficient conversion of the PFOR468Amonomers to an inserted
Structural features of cholesterol-bound ILY
Our previous studies suggested that cholesterol binding by ILY
was not necessary to trigger the D3 structural changes that are
necessary for the formation of the oligomeric complex : it
appeared that CD59 binding, not cholesterol binding, initiated the
D3 structural changes. Subsequent studies showed that the ILY
CRM must initiate a cholesterol-dependent interaction to trigger
the membrane insertion of loops L1–L3, which is necessary to
anchor ILY to the membrane when it disengages from CD59
during prepore to pore conversion . Therefore, if ILY could
bind directly to cholesterol, in the absence of CD59, we predict
that this interaction alone would not trigger the formation of the
pore complex, as control of this process has been transferred to the
CD59-binding site .
Although ILY does not bind significantly to cholesterol-rich cell
membranes that lack human CD59 , we unexpectedly
discovered that it binds well to cholesterol-rich liposomes, even
better than PFO (Fig. 7A). Furthermore, this binding is dependent
on the CRM, as a CRM knockout (ILYDM) lacksdetectable
binding to liposomes (Fig. 7A). Therefore, does this CRM-
mediated binding trigger the D3 structural changes like PFO and
formation of a b-barrel pore? To address this question we first
generated cholesterol-rich liposomes with entrapped 5(6)-carboxy-
fluorescein (CF) and then treated them with PFO or ILY. The
fluorescence emission of the concentrated liposome-trapped dye is
quenched, but if the dye is released from the liposome its
fluorescence emission increases upon dilution as it is released from
the liposome [39,40,41]. PFO exhibited a dose-dependent release
Figure 6. TMH insertion in PFO and PFOR468A. A cysteine was substituted for Ala-215 in PFO and PFOR468A, which is located in TMH1.The
sidechain of Ala-215 is in an aqueous environment in the soluble monomer, but enters the membrane upon formation of the membrane spanning b-
barrel . Therefore an NBD probe positioned at this site undergoes a polar to nonpolar transition that is detected by an increase in the fluorescence
emission of the probe.(A) Each NBD-labeled derivative was incubated in the presence (dashed line) and absence (solid line) of human erythrocyte
ghost membranes. (B) Unlabeled native PFO or PFOR468Awere mixed in a 4:1 molar ratio with PFOA215C-NBDand PFOR468ANA215C-NBDderivatives. The
fluorescence emission for all experiments wasrelative to the maximum emission change observed for NBD-labeled PFO. The fluorescence emission
intensity of NBD was measured from 500–600 nm. The data are representative of 3 experiments.
Figure 7. ILY binding and pore formation on cholesterol-rich
liposomes. (A) Binding of PFO, ILY and ILYDMto cholesterol-rich POPC
liposomes was measured by SPR. The data is representative of 3
experiments. (B) Pore formation on liposomes was measured as the
emission intensity of CF increased upon dilution as pores are formed in
the liposomes. The change in the emission intensity of CF over time in
an untreated sample was subtracted from the experimental data. The
data are representative of at least 3 analyses. ILYDMcontains glycine
substitutions for the ILY CRM residues Thr-517 and Leu-518, which
knocks out CRM-dependent binding to cholesterol-rich membranes
.(C) To measure the insertion of the b-barrel pore a cysteine was
substitutedand modified with NBDfor TMH1 residue Ala-215 of PFO or
its analog, His-242 in ILY. Each derivative was incubated in the presence
(dashed line) and absence (solid line) of cholesterol-rich liposomes. As
the soluble monomer binds to and forms a pore in the membrane the
NBD probe positioned in TMH1 makes the transition from a polar
environment in the soluble monomer (solid line) to the nonpolar
environment of the membrane (dashed line), which is reflected by an
increase in the NBD fluorescence emission .
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of the dye as evidenced by the increased emission of the dye as the
concentration of PFO was increased. Although about twice as
much ILY as PFO is bound to the liposomes, the ILY released less
than 6% of dye released by PFO at the highest concentrations
(Fig. 7B). We confirmed that the b-barrel of ILY was not inserting
by measuring the insertion of TMH1. A NBD probe was position
in TMH1 at cysteine-substituted His-242, which is a membrane
facing residue in the b-barrel .Consistent with the lack of pore
formation, ILY did not insert its b-barrel into the liposomal
membranes (Fig. 7C).
Although pores were not forming, it was possible that the D3
b4–b5 interactionwas disrupted upon ILY binding to cholesterol-
rich liposomes. The disruption of the b4–b5 interaction is detected
by a decrease in the emission intensity of an NBD probe positioned
in b4 as b5 rotates away from b4 its exposes the probe to the
aqueous milieu thereby quenching its emission. This transition did
not occur in the liposome bound ILY (Fig. 8), although it does
occur in ILY bound to human CD59 containing cell membranes
.Collectively these results suggest that while ILY can bind to
cholesterol-rich POPC liposomes, like PFOR468Aits binding does
not trigger the D3 structural transitions necessary to initiate the
formation of the oligomeric pore complex.
The PFO pore forming mechanism is highly sensitive to
changes in the undecapeptide structure [6,13,16], but until now
the molecular basis for its role in the CDC mechanism has been
elusive. The studies herein show that mutation of the undecapep-
tide arginine residue uncouples membrane binding from the D3
structural transitions, which are necessary for the assembly of the
pore complex.This mutation blocks all detectable structural
changes in D3and prevents the stable interaction of the
membrane-bound monomers. In essence, the structure of mem-
brane bound monomers of this mutant appears relatively
unchanged from that of the soluble monomer. Hence, for the
first time these studies demonstrate a function for the conserved
undecapeptide, which forms a critical structural elementin the
allosteric pathway that couples membrane binding to the D3
structural changes that lead to pore formation. The studies also
show that binding initiates changes through this allosteric pathway
that allow monomer-monomer interaction, but it is the monomer-
monomer interactions that subsequently drive the major D3
structural transitions that are required for formation of the
oligomeric pore complex.Furthermore, these studies show that this
pathway has been lost in a CD59-binding CDC, which was a
necessary evolutionary step towards transferring control of this
process from the cholesterol-binding site to the CD59-binding site.
These studies show that mutation of Arg-468 disrupts the
allosteric signal that couples binding to the D3 structural changes
that lead to the formation of the oligomeric pore complex and that
functional derivatives of PFO can drive the major structural
changes in D3, which are necessary for membrane insertion of the
TMHs of PFOR468A. Hotze et al. showed that monomer-
monomer contact could drive the major structural transitions in
D3 in a mutant of PFO that was trapped in a prepore complex.
Here we show that PFO can drive these changes in PFOR468A,
which is trapped at a much earlier stage where the monomers
cannot interact with each other. Our data suggest that membrane
binding is allosterically coupled to structural changes in PFO,
which facilitate monomer-monomer interaction, but alone this
allosteric pathway does not drive the major D3 structural
transitions (i.e., disengagement of D2–3 interface and the b4–b5
interaction): these changes are driven by the subsequent interac-
tion of monomers.
Soluble monomers of PFO do not interact and form oligomers,
even at the high concentrations required for crystallization [7,28].
Therefore, PFO membrane binding must initiate structural
changes in the monomers that facilitate their interaction. The
monomer-monomer interactions then drive the major conforma-
tional changes within domain 3, whichare required for the
formation of the b-barrel pore . In this way PFO ensures an
efficient assembly of the oligomeric pore complex on the target
membrane. Membrane-bound PFOR468Amonomers did not
appear to form interactions that were of sufficient duration to be
detected by FRET. For FRET to occur the donor and acceptor
fluorophores must be at a fixed distance for a time that is equal to
or greater than the half-life of the donor fluorescence emission,
which is approximately 4 nsec for the Alexa-488 dye .
Therefore, the mutation of Arg-468 appears to prevent the
changes in the monomer structure that allows monomers to
initially interact and form stable contacts that then drive the D3
structural changes. This mutation results in a membrane-bound
monomer that appears to retain the structure of the soluble
monomer, which cannot form any detectable intermolecular
In the crystal structure of PFO the only contacts made by Arg-
468 are hydrogen bonds between the NH1 of its guanidinium
group and the backbone carbonyls of the CRM Thr-Leu pair
(Fig. 1). Therefore, its substitution with alanine only prevents the
formation of these two hydrogen bonds. This contact is interesting
because it hydrogen bonds with the CRM, and may help stabilize
the CRM structure in PFO. Hence, this contact may explain why
mutation or chemical modification of the undecapeptide affects
binding, as well as assembly of the oligomeric pore complex of
PFO [6,13,16]. We cannot know with certainty, however, that this
contact is essential to the allosteric pathway, only that the
substitution of Arg-468 disrupts the allosteric pathway that couples
membrane binding to the formation of the pore complex. The
crystal structures of the cholesterol-binding CDCs PFO ,
suilysin (SLY)  and anthrolysin O (ALO)  have revealed
that the undecapeptide 3D structure is highly variable: no two
undecapeptides 3D structures have been shown to be the same
[7,9,43], even though their undecapeptide primary structures are
identical. It is possible that these differences are due to an inherent
flexibility of the undecapeptide and/or crystal contacts that affect
the structural arrangement of the undecapeptides in the crystals.
Hence, the conformational coupling of binding to the D3
structural changes may proceed through different undecapeptide
mediated contacts in the CDCs. Alternatively, if the structure of
Figure 8. The b4–b5 interaction in cholesterol bound ILY.
Cysteines were substituted for Val-322 in b4 of PFO or the analogous
Val-349 in b4 of ILY and modified with NBD. Upon membrane binding
b5 rotates away from b4, thus, the probe makes a transition from a
nonpolar environment in the soluble monomer (solid line) to a polar
environment in the membrane oligomer (dashed line), which results in
the quenching of the NBD fluorescence . The data are represen-
tative of 3 experiments.
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the undecapeptide is flexible, as is suggested by the crystal
structures, then membrane binding may lock it into a specific
conformation that transmits the allosteric signal to D3, which
cannot be achieved in PFOA468A.
Functional PFO can drive D3 conformational changes in
PFOR468Aand the membrane insertion of itsTMHs in chimeric
oligomers comprised of both proteins. Therefore, PFOR468Ais
structurally competent to form a pore, but lacks the conforma-
tional signal that initiates the necessary changes in its structure that
facilitate the formation of stable intermolecular contacts. The fact
that native PFO can drive these structural changes in PFOR468A
indicates that it can establish a sufficient number of contacts with
the PFOR468Amonomers to drive these conformational changes in
the latter. No stable intermolecular interactions of the PFOR468A
monomers alone were detected by FRET showing that they do not
interact, or that the interactions are transient and only exist on a
timescale that cannot be detected by FRET. The ability of
functional PFO derivatives to interact with PFOR468Aindicates
that at least one of the surfaces of PFOR468Ais accessible to the
functional PFO derivatives, which allows the functional PFO
derivatives to dock with PFOR468A.This interaction allows the
functional PFO derivatives to establish contact with andsubse-
quently drive the structural changes in PFOR468AD3 that are
necessary for the formation of the oligomeric pore complex.
It is clear thatPFOR468Abindingwas also affected by the Arg-468
to alanine mutation. If Arg-468 does make contact with the CRM
carbonyls, as suggested by the PFO crystal structure, then it is
possible that this substitution partially destabilized the CRM
structure thereby affected binding. However, avidity may be a
more important factor that contributes tothe difference in binding
of wildtype PFO and PFOR468A. Wildtype PFO and all mutants
thereof generated to date still form membrane oligomers (most are
represented herein). Oligomerizationis an important component of
the binding interaction due to the avidity of the oligomeric
complex versus the binding affinity of a single monomer.
Oligomerization of PFO begins shortlyafter binding [37,44], thus
binding of wildtype PFO and its derivatives reflects the avidity of
the oligomeric complex rather than the affinity of single
monomer.PFOR468Ais the first mutant that has been described,
whichis trapped in a monomer state on the membrane.Hence,
therelatively poorbinding exhibited by PFOR468Amay actually
reflect the true binding interaction of native PFO monomers in the
absence of oligomerization. It is also important to note that in the
experiments in which functional PFO was used to drive the
structural transitions in PFOR468A
quantitative conversion of these transitions with a 4:1 molar ratio
of functional PFO to PFOR468A. If PFOR468Amonomers bound
the membrane with a significantly lower affinity than native PFO
monomers then the probability of the native PFO monomers
interacting with the PFOR468Amonomers would be decreased and
therefore it would be unlikely that we would have observed the
near quantitative conversion of the PFOR468Amonomers to a
The CD59-binding CDCs, ILY , vaginolysin (VLY) 
and lectinolysin (LLY) [45,46] exhibit undecapeptides with
significant changes to their primary structures, most notably a
proline substitution for the second conserved tryptophan (consen-
EKTGLVWEPWR; LLY, EKTGLVWEPWR). Unlike PFO,
ILY does not maintain the hydrogen bond contacts between
Arg-495 and the CRM (Fig. 1). This may be one of contacts in the
cholesterol-dependent allosteric pathway that was disrupted
during the evolution of the CD59-binding site, which was
necessary to transfer of control of the D3 structural changes from
that we obtained near
the cholesterol-binding site to the CD59-binding site. Consistent
with this scenario is the observation that substitution of the
analogous arginine residue in ILY has little effect on the ILY pore-
forming mechanism . We have shown herein that when ILY
binds to cholesterol in the absence of CD59 it remains largely inert
on the membrane, similar to what we observed for PFOR468A.
These data suggest that through evolution ILY has lost the
allosteric pathway that couples cholesterol binding to the D3
structural changes in order to transfer control of the assembly of
the oligomeric complex to the CD59-binding site [33,42,46].
Recently others have proposed that the membrane attack
complex/perforin (MACPF) family of proteins may exhibit a
CDC-like pore forming mechanism [47,48,49,50,51]. This pro-
posal is based on the presence of a conserved protein fold that is
similar to D3 of the CDCs , which we have shown forms the b-
barrel pore structure of the CDCs [26,27]. The MACPF proteins
play important roles in immune defense as well as in the
pathogenesis of eukaryotic pathogens such as Toxoplasma gondii
and Plasmodium falciparum [53,54,55]. These proteins exhibit
little sequence homology with the CDCs and do not exhibit an
undecapeptide motif. It is possible, however, that they will
alsoexhibit an analogous allosteric mechanism to regulate the
assembly of their pore complex.
These studies provide the first evidence that shows the
conserved undecapeptide plays an integral role in the allosteric
coupling of cholesterol-mediated membrane binding to distal
structural changes, which are necessary for the monomer-
monomer interactions that drive the assembly of the b-barrel pore.
Materials and Methods
Bacterial strains, plasmids and chemicals
The genes for native ILY and PFO were cloned into pTrcHisA
(Invitrogen) as described previously [27,42]. All mutations were
made in native ILY (naturally cysteine-less) or the cysteine-less
PFO derivative (PFOC459A) backgrounds. The various CDCs and
their derivatives are summarized in Table 2. All chemicals and
enzymes were obtained from Sigma, VWR and Research
Organics. All fluorescent probes were obtained from Molecular
Probes (Invitrogen). Polyclonal anti-PFO antibody was affinity
purified from hyperimmune rabbit serum. Secondary antibody
goat anti-rabbit-HRP was obtained from BioRad. Sterols were
obtained from Steraloids and lipids were obtained from Avanti
Generation and purification of toxin derivatives
PCR QuikChange mutagenesis (Stratagene) was used to make
the various amino acid substitutions in native ILY or PFOC459A
and DNA sequences of the PFO and ILY mutants were
determined by the Oklahoma Medical Research Foundation Core
DNA Sequencing Facility. The expression and purification of
recombinant toxins and derivatives inEscherichia coliBL21 DE3
were carried out as previously described [27,56]. Purified protein
was stored in HBS [100 nM NaCl, 50 mM HEPES; (pH 7.5)],
50 mMtris(2-carboxyethyl)phosphine (TCEP) and 10% (vol/vol)
glycerol at 280uC.
Modification of cysteine-substituted toxin derivatives
with fluorescent probes
The labeling of PFO, PFOR468Aand ILY cysteine-containing
derivatives with IANBD [iodoacetamido-N,N9-dimethyl-N-)7-ni-
trobenz-2-oxa-1,3-diazolyl)ethylene-diamine; Molecular Probes]
was carried out as previously described [27,42]. Toxin derivatives
were labeled using a 20-fold molar excess of the probe overnight at
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room temperature (22uC).The labeling reactions for the PFOV322C
derivatives also contained 3 M guanidine hydrochloride to
increase the efficiency of labeling.
Following the modification with the probes the mixtures were
passed over a Sephadex G-50 column equilibrated in HBS. The
labeled samples were made 10% (vol/vol)in glycerol and stored at
280uC. Proteins were typically labeled at an efficiency of 80–
Hemolytic activity of toxins and derivatives
The cytolytic activity of the toxins and their derivatives on
human red blood cells (hRBCs) was measured as previously
described  except that the procedure was adapted to a
microtiter plate format. Briefly, fresh human RBCs (hRBCs) were
washed and suspended to 5% in phosphate buffered saline (PBS).
The PFO and its derivatives were serially diluted in 2-fold steps in
a microtiter plate at a final volume of 50 ml per well to which 50 ml
of a 5% suspension of hRBCs was added and incubated for 1 hour
at 37uC. After incubation, unlysed RBCs were removed by
centrifugation of the plate at 34006g for 10 min. The EC50for
hemolysis (effective concentration of toxin for 50% hemolysis)was
determined by quantifying hemoglobin release by measuring the
absorbance of the supernatantat 540 nm using a DU640B
choline (POPC) and cholesterol at a ratio of 45:55 mol% were
prepared as previously described . Carboxyfluorescein-contain-
ing liposomes were made by adding CF [5(6)-carboxyfluorescein]to
the cholesterol/lipid mixture in HBS at a concentration of 50 mM
before extruding . After extrusion, the encapsulated liposomes
were then passed over a Sephadex G-50 column in HBS pH 7.5 to
separate unencapsulated CF from liposomes.
Surface plasmon resonance analysis of ILY and PFO
Surface plasmon resonance (SPR) was performed with a
BIAcore 3000 system (Oklahoma Medical Research Foundation)
using an L1 sensor chip (Biacore) as previously described .
Binding analysis was performed as previously described  with
the following modification: nine consecutive 10 ml injections of the
toxins and their derivatives (100 ng per injection) in HBS were
passed over the liposome-coated chip at a flow rate of 10 ml/min.
Liposome release assay
The pore forming activity of PFO and ILY and ILY DM was
measured by incubating serial dilutions of toxin with 100 ml of a
1:1000 dilution of carboxyfluorescein (CF)-containing liposomes in
HBS for 1 h at 37uC. Samples were read on a Victor3V Wallac
1420 Multilabel counter (Perkin Elmer) using wavelength settings
optimized for high count fluorescein detection.
Two-fold serial dilutions of PFOb4b5and PFOR468ANb4b5labeled
with Alexa-488 were incubated with washed human RBCs (16106
cells) in PBS for 30 min at 4uC.Samples were then brought to a
final volume of 500 ml with cold PBS and analyzed by a
FACSCalibur flow cytometer (University of Oklahoma Health
Sciences Center), gating on live cells. The emission wavelength
was 530 nm and the excitation was 488 nm with a bandpass of
30 nm. The disulfide locked b4b5 versions of PFO and PFOR468A
have cysteines substituted for residues Thr-319 and Val-334 ,
which forms a disulfide that prevents b5 from rotating away from
b4. This disulfide prevents the lysis, but not binding to the RBCs
during flow cytometry .
The geometric mean fluorescence of RBCs alone was subtract-
ed from the experimental data for both PFO derivatives and the
net fluorescence was graphed using GraphPad Prism.
Table 2. PFO and its derivatives used herein.
Toxin or its derivativeDescription
PFO Recombinant native PFO that contains a alanine substitution for the native cysteine (Cys-459) in the
PFO substituted at Arg-468 with alanine
PFO with an engineered disulfide between b-strands 4 and 5 at cysteine substituted Thr-319 in b4 and Val-334 in
The analogous mutation to PFOb4b5
PFO with an engineered cysteine for Val-322. Val-322 is located in b4 and is buried under the a1b5 loop. It
undergoes a nonpolar to polar transition as the a1b5 loop rotates away from b4 
The analogous mutation toPFOV322C
Cysteine-substituted Asn-197. Asn-197 undergoes a nonpolar to polar transition upon disruption of the D2–D3
The analogous mutation toPFON197
Cysteine-substituted Ala-215 in TMH1. Ala-215 faces the membrane in the b-barrel pore 
The analogous mutation to PFOA215C
ILY Recombinant native ILY
Cholesterol binding site knockout by glycine substitution for the Thr-Leu pair of the CRM in ILY
The analogous mutation in ILY to PFOA215C
The analogous mutation in ILY to PFOV322C
A summary of the CDCs and their derivatives used in the present study is shown.
Allosteric Control of Pore Assembly
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SDS-agarose gel separation of PFO monomer and
SDS-agarose gel electrophoresis was performed as previously
described . Briefly, samples were incubated with different
concentrations of washed hRBCs, for 30 min at 37uC.Toxins were
maintained at 440 nM and the hRBCs concentrations ranged
from 2.56107/ml to 2.56108/ml in a final volume of 40 ml.
Samples were solubilized with SDS sample buffer and the
complexes were analyzed on a 1.5% SDS-agarose gel (100 V,
120 min) and then transferred to nitrocellulose membranes.
Protein bands were identified using rabbit anti-PFO antibody
followed by horseradish peroxidase tagged goat anti-rabbit
secondary IgG. The bands were visualized using a chemilumines-
cent substrate (ECL Western Blotting Detection Reagents,
Amersham/GE Healthcare) and autoradiography.
Human erythrocyte ghost membrane preparation
Human erythrocyte (hRBC) ghost membranes were prepared as
previously described with some modifications [27,42]. After
hypotonic lysis of the hRBCs for 15 min at 4uC in lysis buffer
[5 mM sodium phosphate (monobasic), pH 7.5, 1 mM EDTA],
cytoplasmic constituents were separated from the membranes by
dialysis with 2 L of the lysis buffer by recirculation through a
Vivaflow 200 0.2 mm PES cassette (Sartorius Stedim Biotech).
Membrane protein content was quantified using the Bradford
method (Bio-Rad Protein Assay, Bio-Rad Laboratories, Inc.) as
previously described .
All fluorescence intensity measurements were performed using a
Fluorolog-3 Spectrofluorometer with the fluorescence software
(Horiba JobinYvon). NBD measurements were made using the
following settings: an excitation wavelength of 480 nm and an
emission wavelength of 540 nm with a bandpass of 5 nm.
Emission intensity was scanned between 500 and 600 nm at a
resolution of 1 nm with an integration time of 0.1 sec. In a typical
experiment, labeled and unlabeled samples containing 10 mg total
toxin each were incubated with hRBC ghost membranes
(equivalent to 300 mg of membrane protein) or 20 ml liposomes
in HBS for 15 min at 37uC before taking spectral measurements.
For all experiments the fluorescence intensity of the unlabeled
samples was subtracted from that of the fluorescent probe-labeled
samples in order to control for the intrinsic fluorescence of the
sample in the absence of the probe.
Fluorescence resonance energy transfer (FRET)
FRET analysis was performed as previously described  with
the following changes. The PFO and PFOR468Aderivatives were
labeled with either Alexa Fluor 488 (donor, D) or Alexa Fluor 568
(acceptor, A).Parallel samples were prepared containing 10 mg of
D-labeled toxin mixed with a 4-fold molar excess of either A-
labeled toxin or unlabeled (U) toxin in a total volume of 2 ml. To
correct for light scattering and direct excitation of the acceptor, a
sample was prepared in parallel in which unlabeled PFO or
PFOR468A(U) replaced the donor-labeled PFO to create the UA
sample, therefore net DA=DA-UA. The samples were mixed in
the presence of hRBC ghost membranes (equivalent to 300 mg of
membrane protein) for 15 minutes at 37uC and the donor
emission intensity measured from 500 nm to 600 nm. The donor
emission intensity of samples in which unlabeled PFO derivatives
replaced donor-labeled PFO derivatives was measured and
subtracted from the donor-labeled samples to control for any
intrinsic fluorescence of the toxin or direct excitation of the
We would like to acknowledge the technical assistance of P. Parrish.
Conceived and designed the experiments: KJD RKT. Performed the
experiments: KJD. Analyzed the data: KJD RKT. Contributed reagents/
materials/analysis tools: KJD RKT. Wrote the paper: KJD RKT.
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Allosteric Control of Pore Assembly
PLoS Pathogens | www.plospathogens.org11July 2012 | Volume 8 | Issue 7 | e1002787