Using infrared spectroscopy of cyanylated cysteine to map the membrane binding structure and orientation of the hybrid antimicrobial peptide CM15

Article abstract of DOI:10.1021/bi200903p

The synthetic antimicrobial peptide CM15, a hybrid of N-terminal sequences from cecropin and melittin peptides, has been shown to be extremely potent. Its mechanism of action has been thought to involve pore formation based on prior site-directed spin labeling studies. This study examines four single-site β-thiocyanatoalanine variants of CM15 in which the artificial amino acid side chain acts as a vibrational reporter of its local environment through the frequency and line shape of the unique CN stretching band in the infrared spectrum. Circular dichroism experiments indicate that the placements of the artificial side chain have only small perturbative effects on the membrane-bound secondary structure of the CM15 peptide. All variant peptides were placed in buffer solution, in contact with dodecylphosphatidylcholine micelles, and in contact with vesicles formed from Escherichia coli polar lipid extract. At each site, the CN stretching band reports a different behavior. Time-dependent attenuated total reflectance infrared spectra were also collected for each variant as it was allowed to remodel the E. coli lipid vesicles. The results of these experiments agree with the previously proposed formation of toroidal pores, in which each peptide finds itself in an increasingly homogeneous and curved local environment without apparent peptide-peptide interactions. This work also demonstrates the excellent sensitivity of the SCN stretching vibration to small changes in the peptide-lipid interfacial structure.

Full text of DOI:10.1021/bi200903p

Article
pubs.acs.org/biochemistry
Using Infrared Spectroscopy of Cyanylated Cysteine To Map the
Membrane Binding Structure and Orientation of the Hybrid
Antimicrobial Peptide CM15
Katherine N. Alfieri, Alice R. Vienneau, and Casey H. Londergan*
Department of Chemistry, Haverford College, Haverford, Pennsylvania 19041-1392, United States
ABSTRACT: The synthetic antimicrobial peptide CM15, a
hybrid of N-terminal sequences from cecropin and melittin
peptides, has been shown to be extremely potent. Its
mechanism of action has been thought to involve pore
formation based on prior site-directed spin labeling studies.
This study examines four single-site β-thiocyanatoalanine
variants of CM15 in which the artificial amino acid side
chain acts as a vibrational reporter of its local environment
through the frequency and line shape of the unique CN
stretching band in the infrared spectrum. Circular dichroism
experiments indicate that the placements of the artificial side
chain have only small perturbative effects on the membrane-bound secondary structure of the CM15 peptide. All variant peptides
were placed in buffer solution, in contact with dodecylphosphatidylcholine micelles, and in contact with vesicles formed from
Escherichia coli polar lipid extract. At each site, the CN stretching band reports a different behavior. Time-dependent attenuated
total reflectance infrared spectra were also collected for each variant as it was allowed to remodel the E. coli lipid vesicles. The
results of these experiments agree with the previously proposed formation of toroidal pores, in which each peptide finds itself in
an increasingly homogeneous and curved local environment without apparent peptide−peptide interactions. This work also
demonstrates the excellent sensitivity of the SCN stretching vibration to small changes in the peptide−lipid interfacial structure.
haracterizing the structure, structural distribution, and
membrane binding geometry of peripheral membrane
lipid systems (or other cellular structures) during periods
relevant to their antimicrobial activity.
C
proteins (PMPs) is a challenge that requires new experimental
approaches. Recent advances in crystallography using surfac-
tants1 and solid state NMR of oriented samples2 have led to
new insights into the structure and function of a number of
such proteins and peptides. However, a hallmark of these
studies is that the experimental approach used depends to a
large extent on the lipid and protein system of interest without
a single, unified approach that can be applied to proteins of any
size in arbitrary lipid systems. EPR spectroscopy of site-directed
spin-labels has this sought-after flexibility when applied to
PMPs,3 with documented limitations because of the size and
chemical nature of the most typical spin-label4 and the need for
external solution- or lipid-phase paramagnetic species to
address directly the extent of membrane burial.3
AMPs that disrupt target membranes have been proposed to
do so by first binding to the surface of the bilayer.7 This initial
binding event is postulated to result in expansion of the outer
leaflet of the lipid bilayer, after the peptide has reached a critical
concentration, followed by membrane thinning. While the
events that follow initial binding depend greatly on the
particular AMP, several models of membrane disruption,
leading to cell death from membrane destruction or
permeabilization, have been proposed.8 One general mecha-
nism for the action of antimicrobial peptides is the formation of
pores. The formation of tight pores mediated by peptide
aggregation is termed the barrel-stave model, while the
formation of wide pores, in which peptides are separated by
intervening phospholipids and a strong local curvature is
induced in the vicinity of the pore with the headgroups knitting
together the two leaflets, is called the toroidal pore model.8 An
alternative proposed mechanism of AMP membrane disruption
is the detergent-like disintegration mechanism, in which the
peptides permeate the lipid bilayer after reaching a critical
concentration and remove pieces of the bilayer without
translocating into the hydrophobic core.8,9 Analytical techni-
Antimicrobial peptides (AMPs) constitute a subset of
membrane-active species generating much recent interest
because of their possible use as antibiotic agents in an era of
global overuse of antibiotic drugs.5 The relative simplicity of
their sequences belies the complexity of their function, and with
a few notable exceptions, the mechanism of action of many of
these peptides has still not been clearly determined.6 This
challenge has been caused largely by a paucity of direct
experimental techniques that can reveal the membrane-bound
structure of such peptides in contact with their target native
Received: June 10, 2011
Revised: November 15, 2011
Published: November 21, 2011
© 2011 American Chemical Society
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ques with many different native length and time scales have
been used to provide evidence (which is often indirect) of each
of these mechanisms in varying AMP/lipid systems.
on CM15 allows them to “snorkel” into DPC’s polar headgroup
region in a fashion analogous to the snorkeling observed for
charged or hydrophobic residues at lipid surfaces.18 Most
recently, we used two central residues of CM15 to demonstrate
the ability of cyanylated cysteine (C*) to act as a site-specific
infrared reporter of membrane binding.19 This work aims to
characterize further CM15’s interactions with lipid−water
surfaces in model and physiological lipid systems using this
novel vibrational probe group, with ramifications for refining
both the membrane-bound structure and remodeling activity of
CM15 and understanding the particular sensitivity of
cyanylated cysteine in a membrane-bound context.
CM15 is a 15-residue antimicrobial peptide that is a hybrid of
the first seven residues of cecropin A and residues 2−9 of
mellitin.10,11 Cecropin A is a naturally occurring, 37-residue,
helical antimicrobial peptide found in the silk moth Hyalophora
cecropia, and mellitin is a 26-residue peptide found in the
venom of the honey bee Apis mellifera. While mellitin has
potent, broad-spectrum antimicrobial activity, it is extremely
hemolytic, causing damage to host red blood cells. In contrast,
cecropin A shows low cytotoxicity but has lower antimicrobial
activity.8 The CM15 hybrid was first created by Andreu,11 who
combined the favorable characteristics of each AMP. Although
a number of cecropin−mellitin hybrids have been synthesized,
CM15 is the shortest sequence displaying both broad-spectrum
antimicrobial activity and low cytotoxicity.11 Its short length
facilitates its solid-phase synthesis, making CM15 an ideal AMP
for detailed studies of its action on model membranes.
The interaction between CM15 and lipids has been studied
by several groups since CM15’s introduction. Feix and co-
workers used EPR-SDSL and LUVs formed from Escherichia
coli polar lipid extract to model CM15’s action on microbes.8,12
They generated six different single-cysteine mutants of CM15
to attach the spin-label at different sites and used the spin-labels
to map the peptide’s membrane immersion depth as a function
of residue site. They concluded that the labels farthest toward
the hydrophobic face of the peptide are positioned approx-
imately 12.5 Å below the membrane surface, while the labels on
the hydrophilic face reside ∼2.5 Å above the surface. These
results suggest that CM15 initially binds with the helical axis ∼5
Å below the membrane surface, in agreement with the initial
docking step in the two-state peptide−bilayer interaction model
proposed by Huang.13 The Feix group also explored CM15’s
ability to form pores in E. coli and Pseudomonas aeruginosa
membranes via osmoprotection, which determines the size of
transmembrane channels. These studies suggested that CM15
forms pores with diameters of 2.2−3.8 nm.14 Further EPR-
SDSL studies suggested that the pores formed are not
consistent with the tight “barrel-stave” pore model, because
strong spin−spin coupling, which would be evidence of
peptide−peptide contacts, was not observed.15
The β-thiocyano group of C* leads to a CN stretching band
that appears in the infrared absorption spectrum between 2153
and 2164 cm−1 in biologically relevant environments.19−26 This
band, which is moderately strong and can be observed clearly in
25 μm thick aqueous samples at approximately 1 mM, is
sensitive to its local environment in two ways. The frequency
depends on water exposure (via a blue shift due to weak
hydrogen bonds donated by water) and the local electrostatic
environment (via a red shift with increasing solvent polar-
ity).22,26 The line shape in homogeneous environments is most
typically a Voigt profile that varies from narrow and more
Lorentzian in character to broad and Gaussian. This line shape
trend has been interpreted as evidence that C* can report on
solvent dynamics through a motional narrowing mechanism in
solvents with faster femto- to picosecond dipolar reorientation
dynamics.26 The CN band of C* has been used to map
electrostatic potential in enzyme active sites,21,22,24 at a binding
interface between two structured proteins,25 and also to
document site-specific order−disorder transitions in both
model peptides27 and intrinsically disordered proteins.20
A similar nitrile-derivatized amino acid, p-cyanophenylala-
nine,28 was used as a probe of membrane binding in peptides
bound to membranes29 and to the interior surface of reverse
micelles.30 The CN stretching band of this side chain exhibits a
red shift qualitatively similar to that of C* upon burial in the
membrane and the resulting local exclusion of water. Two
peptides derived from natural sequences, of which CM15 is
one, were used recently in contact with DPC micelles to
demonstrate that the CN stretching band of C* is also a
sensitive probe of the membrane exposure of labeled side
chains.19 C* is projected to be a widely useful probe of
membrane−protein interactions, because it can be incorporated
into sequences of arbitrary size via site-directed mutagenesis to
cysteine followed by chemical ligation of the CN group at the
free thiol. Here C* is used to map the membrane-bound
structure of CM15, a short peptide, but its usefulness is not
limited to such short sequences.
Bastos et al.16 studied the interactions between CM15 and
model LUVs formed from DMPC using FTIR centered on the
lipid headgroups. By following the lipid carbonyl stretch as a
function of peptide concentration, they reported that the lipid
remained ordered in the presence of an increasing level of
peptide. The group also concluded that the peptide becomes
sequestered in localized pores, leaving the bulk lipid
unperturbed. This report of pore formation by CM15 agrees
with the conclusions made by the Feix group about the
peptide’s mode of membrane perturbation.
Because of its relatively well-characterized initial orientation
when bound to model membranes, CM15 has recently been
used as a model system to develop new analytical techniques
designed for general use in membrane-bound peptides and
proteins. Zangger et al.17 employed NMR paramagnetic
relaxation techniques to explore CM15’s orientation and
immersion depth in DPC micelles. They found that the
hydrophobic side of the helix interacts with the hydrophobic
core of the micelle, while the hydrophilic side of the helix is
closer to the micelle surface but remains immersed in micellar
samples. Additionally, the length of the charged lysine residues
Two centrally located residues in CM15’s sequence were
previously identified as sites for introduction of the C*
vibrational probe group. One of these residues (Ala10) is
known to be at least partially solvent-exposed, and the other
(Ile8) is deeply buried in contact with multiple lipid
interfaces.8,12 In addition, for this study, one position at each
end of the peptide sequence was chosen for attachment of the
probe for examination of peptide−lipid interactions and local
structure and dynamics near the sequence termini. The last
residue on either end of the sequence was avoided because the
absolute termini help to cap the helix-forming sequence and
have major effects on the overall helical propensity when
binding to membranes.14 To avoid the absolute C-terminus and
possible capping effects, the second-to-last residue, Val14, was
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replaced with C*. When Lee et al. compared the sequences of
10 different cecropins, they found that all 10 contained Trp2
and seven contained Lys3.31 Because these two residues are
highly conserved among cecropins, the fourth N-terminal
position, Leu4, was selected as the site closest to the N-
terminus for introduction of the vibrational probe with minimal
perturbation of the bound structure. Table 1 shows the chosen
liquid chromatography (HPLC) solvents were purchased from
Pharmco-AAPER. Sodium phosphate monobasic monohydrate
(NaH2PO4·H2O), sodium phosphate dibasic (Na2HPO4),
MOPS [3-(N-morpholino)propanesulfonic acid], and KCl
were purchased from J. T. Baker Chemical Co., Fisher
Chemical, and Aldrich, respectively. 5,5′-Dithiobis(2-nitro-
benzoic acid) (DTNB) was purchased from Acros, and
NaCN and D,L-dithiothreitol (DTT) were purchased from
Aldrich. All previously mentioned purchased items were used as
received. Dodecylphosphocholine (DPC) and E. coli bacterial
polar lipid extract (BPL) were purchased from Avanti Polar
Lipids dissolved in chloroform. The BPL has a 67:23.2:9.8
(weight percent) phosphatidylethanolamine (PE):phosphati-
dylglycerol (PG):cardiolipin (CL) ratio. All aqueous solutions
were prepared with doubly deionized, degassed, Milli-Q quality
water.
Table 1. Amino Acid Sequences of Synthesized CM15
a
Variants
peptide
sequence
unmodified CM15
L4C*
I8C*
A10C*
V14C*
Ac-KWKLFKKIGAVLKVL-NH2
Ac-KWKC*FKKIGAVLKVL-NH2
Ac-KWKLFKKC*GAVLKVL-NH2
Ac-KWKLFKKIGC*VLKVL-NH2
Ac-KWKLFKKIGAVLKC*L-NH2
Peptide Synthesis and Purification. All peptides were
synthesized on a 0.1 mmol scale on an Applied Biosystems ABI
443A synthesizer using standard 9-fluorenylmethoxycarbonyl
(Fmoc) chemistry. For each peptide, the C-terminal amino
acid, all β-branched amino acids, and each single residue
preceding a β-branched amino acid were double-coupled. A
PAL resin solid-phase support was used to furnish a C-terminal
carboxamide upon cleavage, and acetic anhydride was used for
N-acetylation. The purity of peptides was verified by reverse-
phase HPLC using an analytical-scale Microsorb 100-5 C18
250 mm × 4.6 mm column (Varian, Inc.) with a flow rate of 1
mL/min and a 0 to 100% or a 20 to 70% gradient of acetonitrile
in water with 0.1% trifluoroacetic acid as the modifier. The
identity of peptides was confirmed via MALDI-MS performed
at the Wistar Institute (Philadelphia, PA).
a
C* represents cyanylated cysteine.
Cyanylation of Peptides. Each cleaved, lyophilized
peptide was treated for 20 min with 0.01 M HCl and
relyophilized to remove remaining trifluoroacetate ions. The
HCl-treated peptides were redissolved in 50 mM sodium
phosphate buffer (pH ∼7.5) and treated with 100× DTT to
yield the free thiol at the single-cysteine side chain. DTT was
separated from the peptides using size-exclusion chromatog-
raphy [28 cm × 10 mm column of Sephadex G-10 equilibrated
in 50 mM sodium phosphate buffer (pH ∼7)], and the reduced
peptides with buffer salts were lyophilized. The reduced
peptides were redissolved in 50 mM sodium phosphate buffer
(pH ∼6.5) and treated with 5 molar equiv of DTNB dissolved
in 200 mM sodium phosphate buffer (pH 7.0) for 20 min to
form a mixed disulfide at the cysteine, turning the solution
yellow. The solution was then treated with 50 equiv of NaCN
dissolved in 50 mM sodium phosphate buffer (pH 6.5) to yield
a thiocyanate at the cysteine, turning the solution orange. The
cyanylated peptide was isolated using the 28 cm × 10 mm
column of Sephadex G-10 equilibrated in either 20 mM sodium
phosphate buffer (pH ∼7) (for DPC experiments) or a 20 mM
MOPS/100 mM KCl buffer solution (pH 6.5) (for BPL
experiments). Peptide-containing fractions were concentrated
∼5-fold using a Savant Speed Vac SCV100 centrifugal vacuum
device, and the presence of the nitrile moiety on the peptide
was verified using infrared spectroscopy. The final sample
concentration was 2 mM in peptide.
Figure 1. Helical wheel diagram for CM15 at the lipid−solvent
interface based on previous studies,8,12,15 with sites chosen for
substitution with cyanylated cysteine colored red.
variant sequences. Figure 1 is a helical wheel diagram of
membrane-bound CM15 based on previous EPR studies by the
Feix group,8,12,15 showing the sites where C* was introduced as
well as the residue positions in relation to the lipid−solvent
interface.12,15
The four chosen single-C* variants of CM15 are used here to
examine its structure in three different environments: aqueous
buffer solution, DPC micelles, and bacterial polar lipid extract
from E. coli cells (BPL). The aqueous samples are used to show
the sensitivity of the probe group to the peptide environment in
the absence of lipid. The micelle-bound samples are used to
produce CD spectra free of any spectral warping from scattered
light and to indicate the basic propensities of side chains to be
buried at a generic phospholipid interface. The E. coli lipids
present an environment in which CM15 adopts an active
structure that porates the lipid vesicles. The spectrum of each
side chain label is examined over time during this process to
provide a site-specific probe of each residue’s microenviron-
ment in the vesicle-bound structure while membrane
remodeling occurs.
EXPERIMENTAL PROCEDURES
■
Preparation of Lipid-Containing Samples. DPC and
BPL were dried down from chloroform stock solutions under a
stream of nitrogen gas. For DPC samples, the resulting lipid
film was hydrated with the speed-vacuumed peptide solution,
resulting in samples that were approximately 2 mM peptide in
100 mM DPC. For BPL samples, BPL was further dried for ∼1
Materials. Fmoc-labeled amino acids and all peptide
synthesis reagents were purchased from Advanced Chem
Tech, except N,N-diisopropylethylamine (DIEA) (Pharmco-
AAPER) and acetic anhydride (Aldrich). Peptide cleavage
reagents were purchased from Aldrich, and high-performance
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h under vacuum and the resulting film was hydrated with a 20
mM MOPS/100 mM KCl buffer (pH 6.5). We prepared large
unilamellar vesicles (LUVs) by freezing and thawing the
hydrated BPL solution five times, followed by extrusion 13
times through 100 nm polycarbonate membrane filters, using a
Mini-Extruder (Avanti Polar Lipids). The speed-vacuumed
peptide solution (30 μL) was added to the extruded BPL
solution, resulting in samples that were approximately 1−2 mM
peptide in 50 mM BPL.
(2)
RESULTS AND DISCUSSION
■
Peptide Synthesis and Modification. All cleaved pep-
tides were nearly analytically pure according to analytical
HPLC, so repeated size-exclusion chromatography associated
with chemical ligation at cysteine was the only purification
used. Cyanylation of cysteine residues followed published
procedures with nearly quantitative yield according to infrared
results showing a clear SCN stretching band. It is worth noting
again that although this labeling methodology is applied here in
the context of short, synthetic peptides, the general technique is
expected to be of use in proteins of arbitrary size with an
expression system that allows placement of a single cysteine
residue at sites of interest. Indeed, the preparation of samples in
this study is in some ways slightly more difficult than isolating
single-C* variants of larger proteins reported elsewhere20,22,24
because of the lack of commercially available size-exclusion
membranes effective for small peptides (<3000 Da) and the
resulting requirement for more selective size-exclusion
chromatography.
Far-UV Circular Dichroism. CD spectra were recorded
between 190 and 250 nm on an Aviv model 410
spectropolarimeter. Speed-vacuumed, cyanylated peptide sam-
ples were diluted 100-fold, yielding a solution of 20−30 μM
peptide in 1−2 mM sodium phosphate buffer. For the native
sequence, the lyophilized peptide was dissolved in 5 mM
sodium phosphate buffer (pH 6.5−7.0), yielding a peptide
concentration of ∼30 μM. These samples were analyzed in a 1
mm quartz cell. For spectra in DPC, dried DPC was hydrated
with the 2−3 mM peptide solution, resulting in a DPC
concentration of 100 mM, and samples were analyzed without
dilution in a 0.1 mm quartz demountable cell (Starna Cells).
FTIR Spectroscopy. For spectra in buffer and 100 mM
DPC, cyanylated peptide samples were placed between two
windows of a 22 μm CaF2 BioCell (BioTools, Jupiter, FL)
inside a Biojack temperature circulating jacket held at 25 °C.
Spectra were recorded at 2 cm−1 resolution (1024 scans) using
a Bruker Optics Vertex 70 FTIR spectrometer with a
photovoltaic HgCdTe detector. A spectrum of buffer solution
or 100 mM DPC in buffer solution was subtracted from the raw
spectrum, and further baseline correction was accomplished by
fitting the baseline outside the region from 2145 to 2180 cm−1
to a polynomial and subtracting the fit.
Far-UV Circular Dichroism. Figure 2 presents CD spectra
of all CM15 variants, including the non-cysteine-containing
ATR-IR was used for peptides in 50 mM BPL because of the
opacity of the samples and the additional advantage of
selectively viewing species that cease to be well-suspended in
solution. After being supplemented with the peptide to the
extruded BPL solution and shaken slightly (to homogenize
them), samples were immediately deposited manually onto a
zinc selenide crystal in an ATR trough cell (Pike Technologies,
Madison, WI) that was then sealed to maintain hydration.
Spectra were recorded at 2 cm−1 resolution (1024 scans) on the
same FTIR spectrometer described above. A spectrum of 100
mM BPL in 20 mM MOPS/100 mM KCl buffer was subtracted
from the raw spectrum, and baseline correction was
accomplished as described above. Spectra of C* analogs in
BPL were acquired approximately every half-hour for 12 h, and
changes in the spectra over time were documented.
Infrared Data Analysis. Line shapes for the CN stretching
bands were analyzed with no assumption about the symmetry
of the line shape, in constrast to prior analyses of the same band
in more homogeneous samples. The mode (most probable, or
maximal) frequency was directly drawn from the data. Mean
CN stretching frequencies were calculated using eq 1 for the
first central moment of the distribution:
Figure 2. Far-UV circular dichroism for all CM15 peptides in aqueous
buffer and in a DPC micelle solution. C* locations are marked on each
plot.
original sequence, in aqueous buffer solution (diluted for
solubility reasons) and in contact with DPC micelles (at the
same high peptide concentrations as infrared measurements).
Because of excessive light scattering and resulting wavelength-
dependent spectral warping, CD spectra were not acquired for
the cyanylated mutants in E. coli lipids. In the micellar CD
(1)
where ω is the frequency in wavenumbers and I(ω) is the
absorbance as a function of frequency. The variance for each
CN stretching band was calculated as the second central
moment of the distribution:
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Figure 3. Infrared CN stretching absorption bands for C*-labeled CM15 variants in an aqueous buffer solution, DPC micelles, and a BPL solution
after the samples had been exposed for 20 min. The nonscaled maximal absorbance of all samples was between 200 and 800 micro-optical density
units.
experiments, the global secondary structure of CM15 in the
lipid-bound state is not greatly perturbed by the placement of
the artificial side chain at the four chosen sites. The DPC-
bound CD spectra are nearly sumperimposable for all peptides
except V14C*. Each spectrum exhibits significant α-helical
character with a strong minimum at 208 nm and evidence of a
weaker negative feature at 222 nm [yielding R values32 of 0.67
(V14C*) to 0.72 (WT) for all DPC-bound peptides]. For such
a short peptide, determination of the percent helicity via fitting
to standard helical and random coil spectra would not be
particularly useful because the number of chromophores
participating in the amide excitons is small, and in the presence
of a lipid, it is likely that the bound structural distribution of
CM15 does not follow a simple two-state helix−coil structural
distribution. In the case of V14C*, the shape of the spectrum
remains similar to that of the other peptides, but there is a
decrease in the molar ellipticity measured for the peptide in
contact with DPC micelles. This is likely due to partial
unfolding of the peptide at the C-terminus, which is apparently
increased by the presence of the less hydrophobic C* in place
of the more hydrophobic valine. This possibility is further
discussed below in the context of IR results at the same site. A
lack of complete quantitative agreement between all DPC-
bound CD spectra indicates that there is a small perturbative
influence of each C* placement on CM15’s DPC-bound
structure; this is not particularly surprising in the context of
recent measurements of C*’s effect on the helical stability of
alanine-repeat peptides.27 Whether these secondary structural
changes affect the orientation of the peptide with respect to the
micelle cannot be determined from CD spectra, which report
only on the relationships between backbone amide groups of
the peptide.
native peptide in DPC, the spectra in aqueous buffer are all
significantly altered as compared to that of the native sequence.
All buffer samples were highly visually transparent and
displayed no macroscopic evidence of aggregation, so the
assumption is that no interpeptide aggregation occurred in any
of these samples. The native sequence displays an aqueous CD
spectrum with a minimum at 201 nm and a very weak higher-
wavelength feature near 222 nm, which could be interpreted as
a mainly random coil spectrum with some small residual signal
from helical structures. The aqueous CD spectra for each
mutant exhibit two minima at varying wavelengths and MREs,
indicating that each peptide has a different latent helical
propensity in an aqueous environment. The spectra with lower-
wavelength minima generally display weaker negative high-
wavelength features, suggesting that this group of mutants
might be described as having different helical propensities lying
along a general helix−random coil reaction coordinate.
Interestingly, the L4C* and A10C* peptides appear to be
more helical in buffer solution than the native sequence. The
CD results in buffer indicate that the introduction of the
vibrational probe at different sites causes steric changes that
lead to a different average global secondary structure in
aqueous buffer for the C*-containing variants, with some
residual helical propensity remaining for all modified sequences
as evidenced by the weak, high-wavelength negative feature in
each of the aqueous buffer spectra. C* is not expected to be a
conservative mutation at any of the chosen sites, but the basic
helical propensity and mostly helical DPC-bound structure of
the peptide are maintained in all C* variant peptides.
A complementary observation of secondary structure would
be the infrared amide I absorption band for each peptide. One
particular advantage of nitriles as vibrational probes is that CN
stretching vibrations are clearly visible in H2O solutions, which
were used for all samples here. Collection of amide I signals
Although the CD spectra for the cysteine mutants in DPC do
not show large changes compared to the CD spectrum of the
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a
Table 2. Analysis of CN Stretching Bands from Infrared Data in Figure 3
buffer
mode
DPC
mode
E. coli lipids
⟨ω⟩
σ
⟨ω⟩
σ
⟨ω⟩
mode
σ
L4C*
I8C*
A10C*
V14C*
2161.6
2162.8
2161.2
2162.8
2161.5
2163.5
2160.6
2163.5
4.7
4.2
7.3
5.3
2159.0
2158.2
2162.1
2159.8
2159.6
2159.6
2160.1
2158.7
5.0
6.0
8.4
4.3
2157.6
2160.2
2160.9
2160.5
2157.7
2157.7
2161.5
2159.0
5.2
5.4
8.6
5.4
a
All units are cm−1.
Figure 4. (A−D) Infrared CN stretching absorption bands for C*-labeled CM15 variants (with the location of the label marked on each plot) as a
function of time in a BPL solution. (E) Average frequencies and (F) peak variances for all CN bands as a function of time.
from these samples would necessitate the use of D2O as the
solvent, thus the current reliance on CD spectra to estimate the
peptides’ secondary structure.
were aggregated. A possible exception is A10C*; this is
discussed below.
Time-dependent infrared spectra in the same spectral region
for each labeled peptide in contact with E. coli lipids over a
period of 7 h after initial mixing are presented in Figure 4A−D.
The mean frequencies and variances of all CN bands as a
function of time are shown in Figure 4E,F. The most obvious
trend in panels A−D of Figure 4 is an increase in the band’s
signal intensity with time. Because the absorbance of the band
is proportional to the sample concentration near the surface of
the ATR crystal, the increase in the CN band’s absorbance with
time indicates that the surface-localized peptide concentrations
increased continuously over the 7 h time period. An increase in
signal intensity is most likely due to separation of the peptide−
Infrared Spectroscopy. Figure 3 shows the infrared
absorption spectra in the nitrile stretching region for L4C*,
I8C*, A10C*, and V14C* in aqueous buffer, DPC micelles,
and E. coli lipid vesicles. The mean and mode frequencies and
the calculated variances for each of the CN stretching bands in
Figure 3 are listed in Table 2. In Figure 3, there is little variation
in the frequency or line width of the aqueous samples,
suggesting that the artificial side chain in each sample senses a
similar aqueous environment without the exclusion of water
and resulting red shift that might be expected if the peptides
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lipid complex from the solution, which would lead to a higher
concentration of the peptide near the horizontal ATR crystal.
This possible cause of the increase in signal intensity is
supported by the observation of visible precipitates upon
removal of the sample from the IR spectrometer. These
precipitates are likely due to the penetration and remodeling of
lipid vesicles by CM15 during the waiting period. It is
important to note that over several hours, the ATR signal
from the peptide increases by a factor of only ∼2 with the
formation of these insoluble species, which are certainly not
peptide aggregrates but are instead the end product of vesicles
destroyed by CM15 and thus mainly composed of lipids [as
evidenced by much greater IR absorbance in the CH stretching
region (data not shown)].
A detailed inspection of the infrared results from each probe
site begins in the center of the peptide and moves outward.
A10C*. Although alanine is a hydrophobic residue, it has
been reported that Ala10 is positioned on the solution-exposed
side of the amphipathic α-helix formed when CM15 is in the
membrane-bound state. EPR-SDSL studies showed that a long
spin-labeled side chain positioned at position 10 was solvent-
exposed when the peptide was bound to E. coli lipids.8,12 In
contrast, a recent NMR study concluded that residues on the
hydrophilic side of CM15, including the short native Ala10,
remain buried in DPC micelles but are closer to the micelle−
solution surface than residues on the hydrophobic face of the
helix.17 Because Ala10 was characterized as a solvent-exposed
residue by EPR, A10C* was previously chosen as a control for
the examination of cyanylated cysteine’s sensitivity to
membrane burial versus solvent exposure.19 If C* is sensitive
to membrane burial and the position 10 is solvent-exposed,
then there should be no change in the CN stretching band of
A10C* in a lipid versus aqueous environment.
Solvent-dependent studies of methyl thiocyanate (MeSCN)
and other studies of C* in water-solvated peptides showed that
the maximal infrared absorption of the aliphatic thiocyanate
nitrile stretching band occurs at 2163 cm−1 in aqueous
environments.23,26,27 Thus, it was expected that A10C*’s CN
stretch in buffer, DPC, and E. coli lipids would be centered at
2163 cm−1. In Figure 3A and Table 2, the mean CN stretching
frequencies for A10C* are 2161.2 (buffer), 2162.1 (DPC), and
2160.9 cm−1 (E. coli lipids). The frequency modes are 2160.6
(buffer), 2160.1 (DPC), and 2161.5 cm−1 (E. coli lipids). The
mean and mode frequencies of A10C*’s CN stretch do not
differ to a large extent among the three environments. This
result indicates that A10C* is in a similar solvent environment
in all three samples, an observation that agrees qualitatively
with the EPR observation of solvent exposure of the spin-label
in place of A10.8,12,15
However, the CN frequency is significantly lower than 2163
cm−1 in all three spectra, suggesting that the environment of C*
is not fully aqueous regardless of membrane exposure.
Bischak20 examined MeSCN’s CN stretch at varying
water:THF ratios and showed that the CN stretch occurs
near 2161 cm−1 in an 80% water/20% THF mixture; if this
result is used as a benchmark, then the A10C* side chain is still
in a predominantly solvent-exposed environment under all
conditions, consistent with the wheel diagram in Figure 1.
CD data (Figure 2D) indicate that A10C* is at least partly
helical (R = 1.02, with an MRE lower than in the bound form)
in aqueous buffer. Because of the frequent occurrence of
hydrophobic residues in the sequence, the peptide is likely at
least partially collapsed, leading to interactions between the
artificial side chain and hydrophobic or partly water-excluded
regions of the peptide. When compared to the mean CN
frequencies of the other cyanylated peptides in buffer, A10C*’s
mean frequency is the lowest. While all of the other label sites
are adjacent to a lysine, position 10 is surrounded by two
hydrophobic residues on either side. Thus, C* is located in a
more hydrophobic part of the peptide in A10C* than in the
other single-C* variants. The CN frequency lower than 2163
cm−1 could be explained through interactions with hydrophobic
nearest-neighbor side chains in the collapsed and/or folded
aqueous peptide.
For the lipid-bound samples, where A10C* is folded akin to
the native sequence, cyanylated cysteine’s rotational freedom
about the β-carbon and S atom may also cause the solvent-
exposed nitrile group to come into transient contact with the
membrane−solvent surface, possibly leading to membrane-
bound A10C*’s slightly red-shifted mean frequency compared
to the value of 2163 cm−1. Similar frequencies (near 2159−
2161 cm−1) for this probe group have been recently observed
by Webb et al. along a well-structured protein binding interface:
the interpretation from that system suggested that partially
water-exposed sites can exhibit such frequencies if their local
electrostatic environments are suitably perturbed by directional
electric fields,25 which is a distinct possibility near the polar
headgroup region and the lipid−water interface. At this
particular site in CM15, a clear explanation for the slightly
low frequency of this mostly solvent-exposed probe in the
membrane-bound form of the peptide is currently elusive.
However, the environment of A10C* is nearly the same in the
absence of lipid, in contrast to all of the other label sites. Our
diagnosis of predominant solvent exposure of the A10C* side
chain agrees with prior EPR results and disagrees with the
NMR assignment of A10 as buried in DPC micelles. The
disagreement with NMR could be due simply to the greater
length of the artificial C* side chain (approximately 3−5 Å
from the N atom to the backbone) or the spin-labeled MTSL
side chain in EPR (∼6−7 Å in length33) in place of the much
shorter A10 (approximately 2.1 Å from the methyl H atoms to
the backbone).
Edelstein et al.27 reported that the CN stretching band of
cyanylated cysteine in alanine-repeat helical peptides is weakly
sensitive to changes in the local secondary structure of the
peptides: a more helical local secondary structure leads to a
broader CN line width. This change in line width with
secondary structure formation was also observed in experiments
in which C* was placed in a domain of the measles virus
nucleoprotein known to form a helix when bound to a
physiological partner.20 Here, the line shape analysis in Table 2
indicates that A10C*’s CN stretching band is broadened in E.
coli lipids as compared to aqueous buffer. The variance of the
CN stretching band is 8.6 cm−1 in lipids versus 7.3 cm−1 in
buffer. Because A10C* adopts a helical structure in lipids but is
less structured in aqueous environments, the line width
broadening upon membrane binding may be a result of the
peptides forming a helical secondary structure while the probe
remains mainly solvent-exposed.
When A10C* was exposed to E. coli lipids over time (Figure
4A), the frequency and line width did not change appreciably
(Figure 4E,F). The probe at position 10 retains the same
degree of solvent exposure even after contact with bacterial
lipids for several hours and noticeable changes in the lipid
morphology. Thus, the parallel orientation of CM15 versus the
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membrane surface does not vary as CM15 acts on its lipid
environment.
4E), from 2160.2 cm−1 after incubation for 20 min in the E. coli
lipid solution to 2158.3 cm−1 after 7 h, a shift of −1.9 cm−1.
The gradual red shifting of the mean frequency suggests that
the probe is exposed to a less H-bonding environment over
time. The second trend in Figure 4F is a decrease in variance
over time, from 5.4 cm−1 after 20 min to 2.5 cm−1 after 7 h, as a
small high-frequency shoulder vanishes from the spectrum. The
I8C* probe’s local environment becomes increasingly homoge-
neous and less water-exposed with time.
L4C*. This site is buried in the membrane when CM15 is
bound to E. coli lipids, with a burial depth similar to that of
residue 8 according to EPR.12 As expected, the CN stretching
band in the DPC and lipid environments for L4C* (Figure 3C)
is red-shifted versus that of the aqueous spectrum. The mean
frequency exhibits a red shift of 2.6 cm−1 in DPC and a red shift
of 4.0 cm−1 in E. coli lipids, while the mode shifts by 1.9 cm−1 in
DPC and 3.8 cm−1 in E. coli lipids versus buffer. These results
agree with the prior conclusion that L4C* is buried when
CM15 is membrane-bound and also suggest a greater burial
depth in anionic BPL than in DPC. The variance of L4C*’s CN
stretching band is nearly the same among the three L4C*
samples.
ATR-IR spectra of L4C* following exposure to BPL (Figure
4C) exhibit very little change over time. Both the mean
frequency and variance of L4C*’s CN stretch remain relatively
constant over time (Figure 4E,F), with a higher degree of
scatter in the calculated variance than for other variants. The
mean frequency was 2157.6 cm−1 after incubation in the E. coli
lipid solution for 20 min and 2157.5 cm−1 after 7 h. The local
environment of the L4C* probe does not change over time, in
contrast to that of I8C*, which also exhibits a buried probe
group. A model explaining how pore formation might influence
C*’s local environment, and thus the probe signals at the ends
versus the middle of the peptide, is discussed below.
V14C*. According to the conclusions of previous EPR-
SDSL studies, residue 14 of CM15 is only slightly buried on
average.8,12 NMR experiments indicated that Val14 is
positioned on the hydrophilic side of CM15’s amphipathic α-
helix, buried slightly inside, but close to the polar surface of the
DPC micelle.17 The V14C* CN stretching band (Figure 3D)
clearly shifts to the red when the peptide is in DPC and E. coli
lipids, compared to the aqueous spectrum. The mean frequency
moves by −3.0 cm−1 in DPC and −2.3 cm−1 in BPL, while the
mode shifts by −4.8 cm−1 in DPC and −4.5 cm−1 in E. coli
lipids versus buffer. This probe is at least partly buried in the
membrane when bound to both DPC and BPL, agreeing
qualitatively with the EPR conclusion of a near-interfacial depth
at this site.12 In contrast to magnetic experiments, however, the
V14C* IR probe group reports the label’s distribution of
environments rather than just the average.
I8C*. Feix et al.8,12,15 determined that residue 8 in CM15’s
sequence is deeply membrane-buried in the presence of E. coli
lipids. NMR experiments in DPC micelles agreed, with Ile8
immersed in the hydrophobic core of the micelle.17 Solvent-
dependent studies showed that the thiocyanate CN infrared
absorption exhibits a red shift of up to 10 cm−1 in non-
hydrogen-bonding, polar solvents such as THF and
DMSO.19,23,26,34 If THF is a good model for the environment
below the membrane interface, largely water-excluded but with
substantial dipolar character, then I8C*’s CN stretch is
expected to shift to the red by a significant amount in DPC
and E. coli lipids versus that in aqueous solution.
As expected, the I8C* CN stretching band in the DPC and
lipid environments (Figure 3B) is red-shifted compared to that
of the aqueous spectrum. Table 2 shows that the mean
frequency exhibits a red shift of 4.6 cm−1 in DPC and a red shift
of 2.6 cm−1 in E. coli lipids, while the mode shifts by 3.9 cm−1 in
DPC and 5.8 cm−1 in E. coli lipids versus buffer. These results
indicate that I8C* is buried when CM15 is membrane-bound
and agree qualitatively with those of prior EPR-SDSL and
NMR studies. The changes in mean and mode are a
quantitative indication that the line shape in E. coli lipids is
skewed with a substantial shoulder at higher frequencies. The
CN stretching band in E. coli lipids is also noticeably narrower
than either the aqueous or DPC-bound bands.
The line width of methyl thiocyanate’s CN stretch narrows
significantly in THF and alkane solvents compared to more
dipolar solvents, such as water.19,26 This phenomenon may
result from a smaller inhomogeneous frequency distribution
imposed by the near-zero dipoles of the nonaqueous solvent
molecules. I8C*’s spectra in buffer, DPC, and E. coli lipids
indicate that the peptide’s CN stretching band narrows in E. coli
lipids only. This result could be explained by peptide−lipid
headgroup electrostatics. Feix measured CM15’s membrane
binding affinity in PE/PG versus E. coli lipids, which are
composed of the anionic phospholipids PG and CL, as well as
the zwitterionic phospholipid PE. They determined that CM15
had a greater binding affinity for the more anionic E. coli
lipids.35 CM15 should also have a greater binding affinity for E.
coli lipids versus DPC, which is zwitterionic. The electrostatic
attraction between the cationic, lysine-rich peptide and the
anionic lipid headgroups would cause the peptide to be buried
more uniformly in the vesicles, with residue 8 farther on
average from the lipid−solvent interface than in the DPC
samples. If the I8C* probe is buried more uniformly in the
lipids and more excluded from the solution and the headgroups
(which interact strongly with lysine), then it would be exposed
to a more homogeneous, less dipolar environment, resulting in
a more narrow distribution of CN stretching frequencies.
This observation of homogeneity of burial in I8C* highlights
a central difference of C* versus EPR spin-labels near the
membrane interface: the frequency variation (or the line shape)
of C* should be able to report directly on the distribution of
environments sampled by a specific side chain, rather than just
reporting the average exposure to lipid- or solution-phase
paramagnetic species that yields a single depth number in EPR
experiments.
The CN stretching band of V14C* narrows slightly in DPC
as compared to that in buffer. The variance of the DPC
spectrum (4.3 cm−1) is smaller than that of the buffer spectrum
(5.3 cm−1). On the other hand, the broadening observed for
V14C*’s CN stretch in E. coli lipids suggests a greater
heterogeneity in the probe’s environment when bound to
BPL. The frequency range covered by the BPL spectrum for
V14C* indicates that the probe is able to sample both aqueous
and substantially nonaqueous environments, and thus, the line
broadening mechanism previously observed due to structural
formation around a fully aqueous C* probe group20,27 is not
expected to be relevant at this particular site. The V14C*
peptide’s CD spectrum exhibits a lower MRE in contact with
Figure 4B exhibits two trends in the I8C* peptide’s CN
stretch over the course of exposure to BPL over 7 h. The first
trend is a decrease in the band’s mean frequency, shown
quantitatively in the plot of mean frequency versus time (Figure
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DPC micelles: this might be due to local fraying at the C-
terminus of the helix caused by the presence of the probe.
Some fraying of the C-terminus of this short peptide is also
anticipated regardless of the artificial C* residue, because
helical peptides tend to unfold more easily at the C-terminus.36
Either fraying at the C-terminal end of the CM15 helix or the
inherent rotational flexibility of the C* side chain allows the
V14C* probe group to sample a greater diversity of local
environments than any of the other probe groups located at
different sites in the bound CM15 peptides. Figure 5 shows the
mean frequencies and variances (Figure 4E,F) suggest only
small, if any, changes over time in the orientation of CM15 with
respect to the water−lipid interface that are consistent with the
toroidal pore model of AMP membrane perturbation.7
Figure 4A shows that the A10C* probe group remains in a
similar, solvent-exposed environment during exposure to E. coli
lipids throughout a 7 h time period. If CM15 were disrupting
the membrane via a detergent-like carpet mechanism, the
interfacial orientation of individual peptides would be expected
to be more random and disordered. The detergent mechanism
would likely cause the A10C* probe to be solvent-exposed in
some peptides and lipid-buried in others, resulting in broad and
time-dependent CN band IR spectra as peptide-coated,
micellelike particles were created and released. This is clearly
not the case in Figure 4A.
Feix et al.8,12 reported that CM15 initially binds parallel to
the membrane surface with a solvent-exposed spin-label at
position 10. If CM15 formed pores in the bilayer, it would
insert itself perpendicular to the surface as the outer lipid leaflet
expanded. If CM15 were simply to reorient itself with respect
to the bilayer axis, then residue 10 would remain solvent-
exposed, as our experiments indicate it does. A lack of evidence
of vibrational coupling in the IR spectra, which agrees with the
previously observed lack of label−label coupling in EPR
experiments,15 supports the EPR-based suggestion that the
pores formed are toroidal with lipid headgroups intervening
between peptides. In such a scenario, even if the peptide were
inserted across the bilayer, the A10C* probe would still be
partially exposed to the polar headgroups of the lipid and partly
to the solvent, just as it is when the peptide is surface-bound.
Figure 6 depicts the proposed orientations of all labeling sites
with respect to the lipid bilayer before and after pore formation.
Figure 4B suggests that residue 8 of CM15 becomes slightly
more water-excluded after incubation for hours in the BPL
vesicle solution. CM15, as a whole, is most likely not becoming
immersed more deeply in the bilayer, according to A10C*’s CN
stretch. The greater burial of the probe at label position 8 while
the probe in position 10 remains in the same relative position
with respect to the membrane interface could be due to
changes in membrane curvature, consistent with pore
formation. The walls of a pore should have a higher degree
of local curvature than the vesicle surface, caused by the
insertion of polar headgroups along the pore’s surface. If the
position 8 coincided with the portion of the pore surface with
the highest degree of curvature, then L8C* would be buried
more uniformly in the hydrophobic interior compared to its
Figure 5. CN stretching band for V14C* in a BPL solution after
exposure for 20 min, fit to two Gaussians with variable widths (w) and
frequencies (xc).
CN stretch after exposure to lipid for 20 min, fit to two
Gaussians of similar width, one centered at 2164.1 cm−1 and
one at 2157.2 cm−1. This fit provides evidence of two possible
frequency subpopulations resulting from C* in solvent-exposed
and lipid-buried environments.
When V14C* is exposed to BPL over time, its mean CN
frequency (Figure 4D,E) jumps from 2161.5 cm−1 after lipid
exposure for 20 min to 2159.4 cm−1 after 1 h and then remains
relatively constant over the remainder of the 7 h period.
Although the variance data are scattered, the CN stretching
band’s line width does not change systematically with time.
These results suggest that with a possible change during the
first hour, position 14 does not become increasingly buried or
solvent-exposed with time and remains accessible to both the
solvent and the membrane.
Pore Formation by CM15. Figure 4 largely agrees with a
previous proposal for CM15’s mechanism of membrane
disruption. The trends in the C* nitrile stretching bands’
Figure 6. Cartoon representation of the positioning of CM15 during pore formation in BPL bilayers, with IR label sites indicated. Graphical
distances do not correspond to any physical scale. Legend: light blue, aqueous solution; dark blue, polar headgroups; yellow, hydrophobic lipid
chains; red cylinder, CM15 helix.
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original environment at the unperturbed bilayer surface. Figure
6 suggests how the local environment of residue 8 after pore
formation could become more alkane as the result of
membrane curvature, while the solvent exposure of residue
10 remains unchanged.
can be placed in largely arbitrary positions along a membrane-
active domain. Because C* does weakly perturb helical folding
and the secondary structures of CM15 variant peptides here are
not quantitatively identical, careful consideration should be
applied in the placement of this probe group.
To facilitate the global use of C* to understand more
complicated membrane-bound proteins from a site-specific
point of view, there is a clear need for more systematic
experimental work to clarify the exact reporting ability of the
CN stretching band in the broad contexts of lipid bilayers and
the lipid−solution interface, each of which presents extremely
heterogeneous environments with respect to local polarity,
charge, and dynamics. In this study, a few residues near the
solution−lipid boundary were investigated. Before meaningful
conclusions can be drawn in much more complicated systems,
there is a need for a clear and systematic study of the
dependence of the CN stretching band’s frequency and line
shape on both its depth in the bilayer and its response through
the headgroup and interfacial regions. This band’s response to
its structural environment has been shown by a number of
recent studies involving enzymatic active sites and binding
interfaces to be quite complicated,20−22,25 with site-specific
responses that do not appear to vary systematically solely on
the basis of either H-bonding or electrostatic factors according
to either experiment or increasingly sophisticated
theory.23,39−41 Therefore, a thorough study of the response of
this particular probe group to lipid-bound and lipid-influenced
environments must be an experimental priority in the future.
We can already see that its complicated and sensitive response
may lead to its wide, and likely quite illuminating, use in many
different possible systems.
Panels C and D of Figure 4 indicate that the environments
around the probe groups at positions 4 and 14 do not change
appreciably with time. This lack of change is also consistent
with toroidal pore formation as schematically represented in
Figure 6, in which the environment of the peptide’s ends
remains nearly the same while the curvature near the center of
the peptide changes slightly as a result of pore formation.
The results in Figure 4 say nothing about the relative
orientation or registry between peptides or the orientation of
individual peptides with respect to the bilayer plane or the axes
of the pore. Presumably, multiple peptides are involved in the
formation of each pore, but they do not interact closely enough
for the relatively short-range C* probe groups to report any
clear peptide−peptide interactions. Spin-labels, which have a
much larger length scale for observable label−label interactions,
also failed to show any peptide−peptide interactions within 20
Å,15 and this was used previously to rule out barrel-stave
poration as CM15’s mechanism of action. Toroidal pore
formation was proposed on the basis of experiments using
peptide-bound spin-labels at primarily position 4:15 this study
adds a more comprehensive map of the lipid exposure of the
rest of the peptide.
Our infrared results could be used to validate recent
molecular dynamics simulations of pore formation by
antimicrobial peptides, which tend to exhibit a pronounced
lack of registry in the peptides even as pores form and
expand.37,38 Mixed quantum mechanical/molecular mechanics
simulations could also be useful in further interpretation of the
frequencies observed in Figures 3 and 4 for the probe CN
vibration. One possible issue in this work mentioned by a
reviewer is a disconnect in the time scale; according to stopped-
flow experiments, the initial time scale for formation of the first
pores created by amphipathic peptides can be seconds, rather
than minutes or hours (and according to some MD simulations,
it can be as fast as tens or hundreds of nanoseconds), so it is
possible that there is already some formation of pores at the
earliest time points in Figure 4. This would mean that any time
evolution observed in Figure 4 is mostly due to evolution of the
more macroscopic lipid structure and small resulting changes in
the peptide’s bound environment. Even in the event that pore
formation is not the process revealed in the Figure 4 time
dependence, all of the IR data are still consistent with toroidal
pore formation and maintenance of a mostly helical secondary
structure, and the data clearly do not agree with either barrel-
stave pore formation or surfactant-like disintegration.
CONCLUSIONS
■
C* is a generally useful probe group for evaluating protein−
membrane contacts in PMPs. In the case of the interaction of
CM15 with BPL, C* is able to show in a site-specific way how
the peptide interacts with its target membrane interface. The
data from single-C* variants are consistent with toroidal pore
formation and clearly rule out detergent-like disintegration as
the mechanism of lipid disruption.
This study demonstrates several unique features of the C*
probe for revealing protein−membrane interactions. C*’s
frequency and line shape are clearly sensitive to small changes
in the local environment around the probe group, including
varying levels of membrane binding and water exposure. Unlike
most nonvibrational site-specific probes, the CN band of C* is
able to report on the local structural distrbution, which is
important in peptides and proteins that can interact with a
variety of lipid, solvent, and peptide-based partners in the
complex environment of a lipid−water interface. In the case of
CM15, this ability to report on the heterogeneity of the probe’s
local environment leads to functional conclusions, among
which is the confirmation of the toroidal pore model for the
mechanism of action.
Further interpretation of our data might become possible in
light of additional systematic study of the C* probe in contact
with lipids. In particular, most of the interpretations here are
predicated on the C* frequency being largely dependent on the
presence or absence of water molecules around the CN moiety.
A systematic study of the depth dependence of C*, as it passes
out of the solvent, through the polar headgroups, and into the
alkane interior of the bilayer, will be crucial to the general
ability of C* to report on the orientation and surface structure
Methodological Ramifications. From this study, a few
general obervations can be made regarding the possible general
use of this infrared probe strategy for mapping the structural
relationship between a membrane protein and its heteroge-
neous environment. With just four sites in a small peptide, it is
clear in this context that the C* probe moiety can be a very
sensitive reporter of its local environment. Both the frequency
and line shape of the CN stretching band show significant
variations at the four sites of interest in CM15/lipid samples,
and new structural conclusions can be drawn from these
variations. There is only a small perturbation of the lipid-bound
structure as a result of the artificial amino acid substitution; as
long as this residue is not used to replace charged residues, it
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AUTHOR INFORMATION
■
Corresponding Author
*Telephone: (610) 896-1217. Fax: (610) 896-4963. E-mail:
clonderg@haverford.edu.
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Funding
K.N.A. acknowledges a summer fellowship made possible by a
grant from the Howard Hughes Medical Institute to Haverford
College. A.R.V. acknowledges a scholarship from the Arnold
and Mabel Beckman Foundation. C.H.L. acknowledges a
Cottrell College Science Award from Research Corp., a New
Faculty Start-Up Award from the Dreyfus Foundation, and
Grant R15-GM088749 from the National Institute of General
Medical Sciences.
ACKNOWLEDGMENTS
■
Dr. Jimmy Feix is thanked by K.N.A. for important suggestions
at a formative stage of this work.
ABBREVIATIONS
■
AMPs, antimicrobial peptides; ATR, attenuated total reflec-
tance; BPL, bacterial polar lipids, extracted from E. coli; C*, β-
thiocyanatoalanine or cyanylated cysteine; CD, far-UV circular
dichroism; CM15, sequence hybrid of residues 1−7 of cecropin
and residues 2−9 of mellitin; DPC, dodecylphosphatidylcho-
line; DMPC, 1,2-dimyristoyl-sn-glycero-3-phosphocholine;
DOPC, 1,2-dioleoyl-sn-glycero-3-phosphocholine; EPR, elec-
tron paramagnetic resonance; FTIR, Fourier transform infrared
spectroscopy; LUVs, GUVs, and SUVs, large, giant, and small
unilamellar vesicles, respectively; MTSL, S-(2,2,5,5-tetramethyl-
2,5-dihydro-1H-pyrrol-3-yl)methyl methanesulfonothioate;
NMR, nuclear magnetic resonance; PMPs, peripheral mem-
brane proteins; SDSL, site-directed spin labeling.
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