Self Assembly of Coiled-Coil Peptide-Porphyrin Complexes
We are interested in the controlled assembly of photoelectronic materials using peptides as scaffolds and porphyrins as the conducting material. We describe the integration of a peptide-based polymer strategy with the ability of designed basic peptides to bind anionic porphyrins in order to create regulated photoelectronically active biomaterials. We have described our peptide system in earlier work, which demonstrates the ability of a peptide to form filamentous materials made up of self-assembling coiled-coil structures. We have modified this peptide system to include lysine residues appropriately positioned to specifically bind meso-tetrakis(4-sulfonatophenyl)porphine (TPPS(4)), a porphyrin that contains four negatively charged sulfonate groups at neutral pH. We measure the binding of TPPS(4) to our peptide using UV--visible and fluorescence spectroscopies to follow the porphyrin signature. We determine the concomitant acquisition of helical secondary structure in the peptide upon TPPS(4) binding using circular dichroism spectropolarimetry. This binding fosters polymerization of the peptide, as shown by absorbance extinction effects in the peptide CD spectra. The morphologies of the peptide/porphyrin complexes, as imaged by atomic force microscopy, are consistent with the coiled-coil polymers that we had characterized earlier, except that the heights are slightly higher, consistent with porphyrin binding. Evidence for exciton coupling in the copolymers is shown by red-shifting in the UV--visible data, however, the coupling is weak based on a lack of fluorescence quenching in fluorescence experiments.
Self Assembly of Coiled-Coil Peptide-Porphyrin Complexes
Bashkim Kokona, Andrew M. Kim, R. Claire Roden, Joshua P. Daniels,
Brian J. Pepe-Mooney, Brian C. Kovaric, Julio C. de Paula,
Karl A. Johnson, and
Department of Biology, Haverford College, 370 Lancaster Avenue, Haverford, Pennsylvania 19041
Received January 14, 2009; Revised Manuscript Received March 13, 2009
We are interested in the controlled assembly of photoelectronic materials using peptides as scaffolds and porphyrins
as the conducting material. We describe the integration of a peptide-based polymer strategy with the ability of
designed basic peptides to bind anionic porphyrins in order to create regulated photoelectronically active
biomaterials. We have described our peptide system in earlier work, which demonstrates the ability of a peptide
to form ﬁlamentous materials made up of self-assembling coiled-coil structures. We have modiﬁed this peptide
system to include lysine residues appropriately positioned to speciﬁcally bind meso-tetrakis(4-sulfonatophe-
), a porphyrin that contains four negatively charged sulfonate groups at neutral pH. We measure
the binding of TPPS
to our peptide using UV-visible and ﬂuorescence spectroscopies to follow the porphyrin
signature. We determine the concomitant acquisition of helical secondary structure in the peptide upon TPPS
binding using circular dichroism spectropolarimetry. This binding fosters polymerization of the peptide, as shown
by absorbance extinction effects in the peptide CD spectra. The morphologies of the peptide/porphyrin complexes,
as imaged by atomic force microscopy, are consistent with the coiled-coil polymers that we had characterized
earlier, except that the heights are slightly higher, consistent with porphyrin binding. Evidence for exciton coupling
in the copolymers is shown by red-shifting in the UV-visible data, however, the coupling is weak based on a
lack of ﬂuorescence quenching in ﬂuorescence experiments.
Protein self-assembly has been recognized as a powerful
paradigm for the creation of biomaterials that can be engineered
with atomic precision and can be regulated by environmental
conditions. The focus here is on imparting photoelectronic activity
onto peptide-based structural scaffolds. The use of porphyrins and
hemes as photoelectronic modules has been extensively studied
and several reviews have been published describing these efforts.
The electronic properties of porphyrins have also been studied
bound to other biomacromolecules, such as DNA,
with an eye to developing biomaterials with
photoelectronic properties. The advantage in using protein-based
scaffolds is the ability to regulate assembly and disassembly of
biomaterials by using the diverse functionality afforded by both
natural and unnatural amino acids.
Protein design efforts have focused largely on taking advan-
tage of side-chain functional groups to coordinate the metal ions
bound to various porphyrin structures;
we instead take advan-
tage of noncovalent binding between porphyrin pendant groups
and amino acid functional groups as this would allow greater
ﬂexibility to use different metalated porphyrins without com-
promising design strategies. We have focused on studies
involving peptide interactions with meso-tetrakis(4-sulfonatophe-
) by taking advantage of electrostatic
interactions with the sulfonate groups.
have been demonstrated between TPPS
and peptides or proteins
containing basic amino acids in aqueous solution. These include
studies of general binding to proteins such as tubulin,
It is well-known that TPPS
itself can aggregate at low pH (pH
< 2.0), forming either J aggregates or, at higher porphyrin
concentration, H aggregates. The structural difference in these
aggregates derives from different stacking geometries between
porphyrin pairs, in which the J aggregates are thought to form an
array of slipped face-to-face stacking interactions.
of J aggregates reveal that these linear arrays appear as nanorods
in the micrometer scale range;
little is known about large scale
structural properties of H aggregates. These structures have been
well characterized by spectroscopic methods. Monomeric TPPS
has a Soret absorption band at 413 nm at neutral pH. This band is
red-shifted to 434 nm in its diacid form at low pH. J aggregates
are characterized by a spectroscopic signature containing a strongly
red-shifted Soret band at 490 nm (J-band) and a second band at
706 nm (Q
H aggregates, in contrast, are characterized
instead by a blue-shifted Soret band.
Thus, most of the work involving protein/TPPS
has been done under these low pH conditions where the
porphyrins can self-assemble.
The focus in the work at low
pH has been on how protein binding might regulate the
J-aggregate assembly and modulate the relevant electronic
transitions. However, at such low pH, many peptides and
proteins are susceptible to unfolding, thus, interactions at neutral
pH are more appropriate for biomaterials design to maintain
the integrity of protein structure.
Several papers have
described charged peptide-porphyrin interactions
these papers explored the ability to couple protein
folding and self-assembly with porphyrin interaction. We
became interested in such coupling of protein folding upon
porphyrin interaction as a mechanism to create biomaterials
whose self-assembly is driven, at least in part, by protein
We have demonstrated that TPPS
bind noncovalently to a peptide through interactions between
the porphyrin sulfonate groups and appropriately spaced lysine
* To whom correspondence should be addressed. Tel.: (610) 896-4205.
Fax: (610) 896-4963. E-mail: firstname.lastname@example.org.
Current Address: Department of Chemistry, Lewis and Clark College,
0615 SW Palatine Hill Rd., Portland, OR 97219.
Biomacromolecules 2009, 10, 1454–14591454
10.1021/bm9000553 CCC: $40.75 2009 American Chemical Society
Published on Web 04/17/2009
residues in the peptide.
The peptide, Cp3K-N (see sequence
in Table 1), contains three lysine residues that are spaced i,i +
4 with respect to one another. This spacing would juxtapose
the lysines to interact with three sulfonate groups upon
acquisition of a helical structure (Figure 1). Circular dichroism,
UV-visible spectroscopy, and analytical ultracentrifugation
experiments from this earlier work demonstrated that the
peptide/porphyrin interaction was speciﬁc and thus presented
an attractive model for precise engineering of photoelectronically
Toward the goal of developing peptide scaffolds for photo-
electronic functionalization, we created a coiled-coil based
polymerization strategy that could be combined simply with a
porphyrin-binding module as described above and as illustrated
in Figure 1.
Coiled coils are protein structural motifs that
take advantage of amphipathic R-helices to drive precise helix-
pairing interactions. Most natural and designed coiled coils are
hydrophobic along one continuous face of the helix, resulting
in well-deﬁned assemblies. However, we modiﬁed a sequence
that normally creates an amphipathic helix to favor offset helix
This was done by inserting two alanines
between two 14-residue amphipathic sequences, placing out of
phase the hydrophobic residues in the second half of the
sequence by 210° with respect to the ﬁrst half. Thus, helix
pairing would necessitate a staggered structure that could
polymerize to form ﬁlaments and ﬁbrils in the 10
range. In the past several years, such redesign strategies using
coiled coils to form nanoscale biomaterials have been realized
and attempts to build functionality and larger-scale structural
features are well underway.
We then re-engineered the polymerizing peptide (CpA)
described in our earlier work
to incorporate the lysines at the
same relative positions as that described for Cp3K-N.
new peptide, which we call Cp3K (Table 1), thus contains
elements of CpA and Cp3K-N with some additional modiﬁca-
tions; a model of the presumed interaction of Cp3K with TPPS
is shown in Figure 1. In unpublished work, we found that
replacing the cysteines in CpA with isoleucines (CpA-I; Table
1) enhances the hydrophobic interactions between helices and
fosters self-assembly at lower peptide and salt concentrations.
In addition, the lysines at all other positions and all of the
glutamates in our original design were replaced with glutamines
to avoid unwanted electrostatic interactions. The work described
here shows our efforts to characterize the photoelectronic and
structural properties of Cp3K/TPPS
Molecular Modeling. The starting molecular model for the coiled-
coil polymer was taken from our published model for the 30-residue
The model for the Cp3K-N interaction with TPPS
was then overlaid onto the CpA polymer model in the following way
(see Table 1 for the peptide sequences). Using InsightII 2000 (Biosym
Technologies, San Diego), the backbone atoms from the central 10
amino acid residues in the Cp3K-N peptide (docked with TPPS
superimposed onto the equivalent set of atoms within the ﬁrst two
heptads of CpA. This allowed the TPPS
to be properly oriented relative
to the polymerizing module of CpA. The original Cp3K-N structure
was then deleted, leaving only the TPPS
porphyrin docked onto the
CpA sequence. The appropriate residues in CpA were then mutated to
lysine residues and these side chains were then manually adjusted to
be consistent with their conformations in the original Cp3K-N/TPPS
model. This exercise was repeated two more times to give a three-
module repeating structure as shown in Figure 1.
Peptide Synthesis and Puriﬁcation. The synthesis of Cp3K was
carried out on an Applied Biosystems (Foster City, CA) 433A peptide
synthesizer using standard FMOC chemistry and using PAL resin
(Advanced ChemTech, Louisville, KY), providing an amide at the
carboxy-terminus. The peptides were acetylated at the amino terminus
prior to TFA cleavage. Peptides were puriﬁed using RP-HPLC with a
mobile phase of water and acetonitrile and 0.1% TFA on a Varian
ProStar (Varian, Inc., Palo Alto, CA) system equipped with a Varian
Dynamax semipreparative C18 column. Peptide was collected and
lyophilized for long-term storage. The peptide identity was conﬁrmed
using MALDI-TOF mass spectrometry, yielding a molecular mass of
3611.8 Da (theoretical MW ) 3612.2 Da).
Solution Preparation. Lyophilized peptide was dissolved in Milli-
Q-ﬁltered water to make stock solutions and used within a few weeks
to avoid problems with unwanted aggregation. Peptide stock solution
concentrations (1-2 mM) were determined by using a modiﬁed
ninhydrin procedure from Rosen.
Porphyrin stock solutions (<1 mM)
were prepared by dissolving TPPS
(Frontier Scientiﬁc Inc.) in 6 mM
NaOH and measuring the concentration of this solution using ε
5.33 × 10
Circular Dichroism Spectropolarimetry. CD data were collected
on an Aviv Associates (Lakewood, NJ) model 202-01 CD spectropo-
larimeter. Wavelength scans were performed at 25 °C in the range of
198-250 nm for the peptide bands and 350-550 nm for the porphyrin
bands using a wavelength step size of 1.0 nm. Data points represent
3 s of averaging time. All samples were measured in 10 mM Tris-
HCl, pH 7.8, unless stated otherwise.
UV-Visible Spectroscopy. UV-visible data were collected using
a 1 mm cuvette on a Lambda 25 UV/vis spectrophotometer (Perkin-
Elmer Instruments, Waltham, MA). Wavelength scans from 350 to 800
nm were performed at room temperature, using a wavelength step size
of 1.0 nm. Samples were prepared in 10 mM Tris-HCl, pH 7.8.
Fluorescence Spectroscopy. Fluorescence spectroscopy was per-
formed usinga1cmcuvette on a Fluorolog-2 spectroﬂuorometer (Spex
Industries, Inc., Edison, NJ) under the control of the dedicated SPEX
DM3000F software. Fluorescence scans were performed at room
temperature in the range of 500 to 800 nm using increments of 1.0
nm, an integration time of 1 s, and an excitation wavelength of 413
nm, which is at the absorption peak maximum for free porphyrin.
Samples were prepared in 10 mM Tris-HCl, pH 7.6.
Fluorescence Microscopy. Fluorescence microscopy was performed
on an Axiophot microscope (Carl Zeiss Inc., Thornwood NY) equipped
with a 100 W mercury excitation source and using a D436/20 excitation
ﬁlter, a D455DCLP dichroic (Chroma Technology, Rockingham, VT),
and a 590LP emission ﬁlter (XF3016, Omega Optical, Brattleboro VT).
Digital images were captured using an Orca ER cooled CCD camera
(Hamamatsu Photonics, Bridgewater, NJ) and a Dell computer (Dell
Inc., Round Rock, TX) running Windows XP (Microsoft Corp., Seattle,
WA) and Metamorph (Molecular Devices, Downingtown, PA) software.
Images were converted from 12-bit ﬁles into an 8-bit RGB format and
cropped and composited using Photoshop (Adobe Systems Inc., San
Analytical Ultracentrifugation. Ultracentrifugation experiments
were performed using a Beckman model ProteomeLab XL-A analytical
ultracentrifuge (Palo Alto, CA) equipped with an An-60 Ti rotor.
Velocity experiments used two-channel Epon charcoal-ﬁlled center-
pieces with 12 mm path length containing 435 µL of sample and 450
µL of buffer reference. Samples were prepared in 10 mM Tris-HCl,
pH 7.8 and 0.1 M NaCl. Sedimentation velocities of peptide boundaries
Table 1. Peptide Sequences
Heptad positions denoted with the letters a-g.
Peptide-Based Biomaterials Biomacromolecules, Vol. 10, No. 6, 2009 1455
were assayed at 30000 rpm at a temperature of 25 °C. Absorbance
data were collected using a radial step size of 0.003 cm and delay time
of 0 s. Data were analyzed using the DCDT+ program version 2.0.9
28802 (John Philo, Thousand Oaks, Ca). Partial speciﬁc volume,
density, and viscosity were calculated using SednTerp v. 1.08.
Atomic Force Microscopy. AFM data were collected in tapping
mode using a Bioscope atomic force microscope (Digital Instruments,
Santa Barbara, CA). Samples were prepared in 10 mM Tris-HCl, pH
7.8 at peptide/porphyrin ratios required for complex formation and
incubated on freshly cleaved mica (SPI Supplies, West Chester, PA)
surface for 10 min at room temperature in a constant humidity chamber.
Samples were rinsed brieﬂy in Milli-Q water and allowed to dry under
Results and Discussion
The Cp3K peptide used in this study contains both a protein
polymerization domain and a TPPS
binding domain as de-
scribed in the Introduction (sequence is shown in Table 1). We
ﬁrst tested biophysical features of the Cp3K peptide to compare
its ability to self-assemble to our earlier published work.
with earlier designs, Cp3K remains largely unfolded at neutral
pH in the absence of NaCl as judged by the lack of helix content
as measured by circular dichroism experiments (Figure 2A).
However, there is a signiﬁcantly higher helix content as
compared to CpA (Table 1), our original design, as judged by
an increase in signal intensity at 222 nm and a red-shift of the
higher energy band toward 208 nm. One contribution to the
enhanced helix content probably arises from weak association
of the peptides through the enhanced hydrophobic core, in which
we replaced the original cysteines in two a heptad positions
with isoleucines. This is supported by the ﬁnding that CpA-I
(Table 1), a related peptide with isoleucine at all a positions,
also shows increased helix content relative to CpA. We had
shown previously that polymerization due to coiled-coil as-
sembly can be induced by the presence of high salt concentra-
Thus, we looked at the sensitivity of self-assembly in
the presence of NaCl for Cp3K in comparison to CpA and
CpA-I (Figure 2B). As expected, Cp3K becomes more helical
as a function of added salt and, as predicted based on the
enhanced hydrophobic core, the approach to full helix content
occurs at much lower NaCl concentrations (1 M) as compared
to CpA (3 M). This increased sensitivity to NaCl is similar to
that observed for CpA-I (also around 1 M). Finally, direct
evidence for Cp3K self-assembly in the presence of NaCl was
tested by sedimentation velocity analytical ultracentrifugation
(SV). Analysis of SV data collected for Cp3K peptide at 100
µM in 0.1 M NaCl at 25 °C reveals a heterogeneous solution
Figure 1. Model of Cp3K coiled-coil polymer interacting with TPPS
. The model contains three coiled-coil peptide units (shown in ribbon form)
whose backbone representation is taken from the structure that we described for the CpA polymerizing model.
The view emphasizes the
overlapping nature of the helices with respect to one another. At the sites of overlap, isoleucine and leucine residues at the a and d heptad
positions, respectively (side chains not shown), are projecting toward one another between the helices to provide optimal hydrophobic packing
interactions for a dimeric coiled coil. Three lysine residues are shown interacting with three of the four sulfonate groups of each TPPS
and is highlighted in ball-and-stick representation.
Figure 2. Circular dichroism analysis of Cp3K, CpA, and CpA-I. (A)
Circular dichroism spectra were taken of 100 µM each of CpA and
Cp3K in 10 mM Tris-HCl, pH 7.8 and measured at 25 °C. (B) NaCl
concentration dependence as monitored at 222 nm for each peptide
under the same conditions as that reported in (A).
1456 Biomacromolecules, Vol. 10, No. 6, 2009 Kokona et al.
with an average molecular weight distribution in the range of
300 kDa (based on calculations using the best-ﬁt values of s
of 5.7 × 10
S, and D
value of 1.54 × 10
indicating a signiﬁcant degree of polymerization comparable
in size to that observed for our other polymerizing peptides
(R.F., unpublished data). AFM images conﬁrmed the presence
of polymers for Cp3K in the presence of 0.1 M NaCl but these
were quite heterogeneous in their morphology (Figure S1 in
Supporting Information), with a broad distribution of heights
centered on 2.2 nm.
Interestingly, upon long-term storage of Cp3K stock solutions,
there was a conversion to β-sheet ﬁbrils (based on CD and AFM
experiments; see Figures S2 and S3 in Supporting Information).
These β-sheet ﬁbrils are more uniform in their lateral dimensions
and quite distinct in their morphology from that observed for
the R-helical polymers. We suspect that this propensity to form
β-sheet ﬁbrils is a consequence of the number of glutamines
that we engineered into the sequence; such polyglutamine
sequences are known to foster β-sheet ﬁbril self-assembly and
their structures have been characterized by AFM.
alternative structure was helpful as a control in showing
speciﬁcity of TPPS
binding to the R-helical ﬁlaments (see
Cp3K contains three lysines in the ﬁrst two hydrophobic
heptad repeats, all spaced i,i + 4 with respect to one another.
Based on modeling studies, this spacing facilitates the engage-
ment of three sulfonate groups on an anionic porphyrin, TPPS
as illustrated in Figure 1. This lysine distribution is identical to
that in Cp3K-N, which we had shown previously to become
helical upon tight binding of TPPS
We looked at whether
could induce helix content in Cp3K. CD experiments
with Cp3K suggest that, upon increasing amounts of TPPS
the CD spectrum is converted from that representing a largely
unfolded structure into an R-helical structure, predicted to
contain approximately 91% helix content
). There is
no contribution to CD in this region of free porphyrin alone.
Stoichiometries beyond 1:0.5 Cp3K/TPPS
result in a red-
shifting of both the 208 and 222 bands, suggestive of formation
of polymers due to effects on band extinction caused by light
There is also an induced TPPS
Soret CD band
upon peptide binding, suggesting that asymmetry has been
imposed on the TPPS
electronic transitions (Figure 3 inset, cyan
line). The intensity of this induced porphyrin band is consistent
with that reported elsewhere for a similar structural model
This CD band is absent in porphyrin by itself (Figure
3 inset, black line). TPPS
cannot bind to Cp3K when the
peptide forms cross-β-sheet ﬁbrils (Figure 3 inset, purple line),
providing additional support that proper juxtaposition of the
three lysines in an R-helical conformation is critical for tight
binding of TPPS
. The Soret band (normally at 413 nm) is split,
resulting in a positive component at 400 nm and a negative
component at 424 nm. The absolute magnitudes of these two
components are approximately equal in intensity to one another,
suggesting that the porphyrins are binding to the peptide in a
single conformation. This effect was also observed in binding
to our original model system, Cp3K-N, except that the order of
the positive and negative bands is reversed upon binding Cp3K-
We attempted SV experiments on this complex to determine
the size of the polymers but the samples precipitated in the time
frame of the experiments (6-9 h).
We also studied the effect of the addition of Cp3K on the TPPS
absorbance bands. At a pH of 7.8, TPPS
exists in its base form
as a monomer with its Soret peak at 413 nm (Figure 4). Upon
addition of substoichiometric amounts of Cp3K, we saw a loss in
intensity of the 413 nm peak; however, as the peptide concentration
exceeds the porphyrin concentration, a new red-shifted peak appears
at 425 nm, suggesting evidence for peptide binding
between bound porphyrins. Red-shifting upon
binding was also observed for the porphyrin Q
-bands (Figure 4
inset), providing further support for exciton coupling of porphyrins.
In contrast, porphyrin binding to Cp3K-N caused a loss of the 413
nm Soret band with concomitant increase in a blue-shifted peak at
403 nm (Figure 6), as shown in our earlier work.
This blue shift
is also likely a consequence of changes in the electronic structure
of the bound porphyrin, but may also be inﬂuenced by H-aggregate
Unlike the Cp3K-N/TPPS
binding behavior, as judged by the presence of an isosbestic point
in UV-visible titration data,
the Cp3K and TPPS
shows multiple-state behavior.
If there is strong exciton coupling of TPPS
upon addition of
Cp3K, we might expect a signiﬁcant quenching in the ﬂuorescence
of the porphyrins bound in the self-assembling polymer. We
measured the ﬂuorescence of TPPS
upon excitation at 413 nm
for the porphyrin by itself or in complex with Cp3K (Figure 5).
Excitation at 413 nm resulted in ﬂuorescence bands for TPPS
Figure 3. Induction of helix content in Cp3K upon addition of TPPS
Samples were prepared using 20 µM Cp3K in 10 mM Tris-HCl, pH
7.8 and measured at 25 °C. Inset: CD region of TPPS
and absence of 50 µM Cp3K; cyan: 1:2 Cp3K/TPPS
; black: TPPS
alone, purple: 1:2 Cp3K/TPPS
with Cp3K in a β-sheet conformation.
Figure 4. Effect of Cp3K binding on TPPS
UV-visible absorbance spectra were taken of 20 µM TPPS
with increasing Cp3K concentrations (inset: porphyrin Q
Samples were prepared in 10 mM Tris-HCl, pH 7.8, and measured
at 25 °C.
Peptide-Based Biomaterials Biomacromolecules, Vol. 10, No. 6, 2009 1457
644 and 705 nm. We observed a slight increase and red-shift in
the emission bands upon titration with Cp3K. The red-shifting is
most likely due to polymerization effects, as seen in the UV-visible
experiment. The separation between the bands also becomes more
distinctive. The major band is shifted toward 655 nm and the second
band toward 720 nm. The lack of quenching suggests that there is
no signiﬁcant buildup of exciton coupling between porphyrins in
the polymeric structure.
Interestingly, the emission spectra of
in the presence of Cp3K-N and Cp3K are almost identical
(Figure 6), providing additional support for minimal exciton
Because we observed minimal quenching of ﬂuorescence
emission, we realized that we could probe directly for TPPS
binding to the Cp3K ﬁlaments by using ﬂuorescence microscopy.
Fluorescence microscopy of a 1:1 ratio of TPPS
to Cp3K showed
evidence of large scale aggregation with what look like highly
tangled ﬁlaments (Figure 7). We suspect that this is what happens
when the samples precipitate; we are unlikely to see smaller
aggregates due to relatively poor sensitivity of this technique. The
smallest polymers that we observed in the ﬁeld presented in Figure
7 are in the range of 0.1-0.3 µm, consistent with the size of the
polymers observed in AFM images (see below). In the absence of
Cp3K, no ﬂuorescence of TPPS
was observed (data not shown).
To probe the morphology of the Cp3K/TPPS
polymers at higher
resolution, we used AFM imaging. Samples for AFM were
prepared from the same samples used for the ﬂuorescence
microscopy, using a 1:1 ratio of peptide to porphyrin. There is
signiﬁcant heterogeneity in polymer length and some branching is
seen, possibly through porphyrin-porphyrin interactions between
polymers (Figure 8). Polymer height measurements indicate an
average height of 3.52 ( 0.2 nm. Cp3K and CpA polymers in the
presence of NaCl have an average height of 2.2 ( 0.6 (determined
from an analysis of Figure S1) and 2.9 ( 0.6 nm, respectively.
Thus, the difference in height between the polymers in NaCl versus
polymers in the presence of TPPS
is suggestive of porphyrin
Our goal in this work was to create photoelectronically active
biomaterials with atomic level precision of assembly using a coiled-
coil polymerizing peptide system and TPPS
, an anionic porphyrin.
We show that assembly at the nanoscale mimics the assembly that
we observed previously for a soluble peptide/porphyrin system,
involving charged interactions between lysine side chains on the
peptide and sulfonate groups on the porphyrin.
As stated earlier,
the advantage of developing this particular binding strategy is to
allow us to use different metalated porphyrins without having to
Figure 5. Effect of Cp3K binding on TPPS
conditions include 5 µm porphyrin in 10 mM Tris-HCl, pH 7.6.
Figure 6. Comparison of the absorbance and ﬂuorescence spectra
of 5 µM TPPS
saturated with 40 µM Cp3K or Cp3K-N in 10 mM
Tris-HCl, pH 7.6.
Figure 7. Fluorescence microscopy of Cp3K/TPPS
at a 1:1 ratio (10 µM each) in 10 mM Tris-HCl, pH 7.6,
were deposited on a freshly cleaved mica surface and incubated in
a humidiﬁer box for 10 min at room temperature, rinsed brieﬂy, and
then allowed to dry prior to being imaged.
Figure 8. AFM topography image of Cp3K/TPPS
polymers. The AFM
image was collected on the sample that was prepared for ﬂuores-
cence microscopy (see Figure 7).
1458 Biomacromolecules, Vol. 10, No. 6, 2009 Kokona et al.
change the design strategy. Because metalation of porphyrins affects
their redox properties,
it offers a long-term goal for implementing
asymmetric addition strategies to create intrinsic voltage potentials
across our peptide polymers.
Addition of the porphyrin does not disrupt the designed
polymerizing potential of the peptide, as judged by comparing the
secondary structure and the polymer morphology to our earlier
published work; in fact, it contributes a helix-stabilizing force.
Exciton coupling appears to be complex, with UV-visible
spectroscopic data supporting such coupling. However, the extent
of coupling is weak as evidenced by little ﬂuorescence quenching.
While we have successfully created photoelectronically active
biomaterials, the ﬂuorescence microscopy and AFM imaging
suggest a signiﬁcant degree of heterogeneity in polymer length and
tangling; this was not entirely unanticipated since we observed a
similar degree of heterogeneity in coiled-coil assembly in our earlier
We argued that the heterogeneity was due in part to the
small scale of the ﬁlament widths; ﬁlaments that aggregate laterally
show greater persistence length and overall increased homogeneity
in ﬁbril width.
Although we intend to pursue rudimentary
conductive properties of these materials, we are currently probing
environmental conditions to favor the formation of larger, and more
regular, ﬁbril structures using our peptide/porphyrin system.
J-aggregate interactions by the addition of coiled-
coil forming peptides may result in forming ﬁbril structures with
more homogeneous characteristics.
Such studies are currently
Acknowledgment. We wish to thank Walter Smith and Al
Schwab for helpful advice in interpreting the data. We also thank
Meg Twomey for able technical assistance and Joe Cammisa
for generating the graphics image of the Cp3K/TPPS
This research was supported by grants from NSF (CHE-0616615
and MCB-0516025 to R.F.), the David and Lucile Packard
Foundation to J.C.d.P., K.A.J., and R.F., and grants to Haverford
College from the Arnold and Mabel Beckman Foundation and
the HHMI Undergraduate Science Education Program to support
Supporting Information Available. Atomic force micros-
copy image of Cp3K in 0.1 M NaCl and circular dichroism
spectrum and atomic force microscopy image of Cp3K peptide
forming cross-β-sheet ﬁbrils. This material is available free of
charge via the Internet at http://pubs.acs.org.
References and Notes
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Peptide-Based Biomaterials Biomacromolecules, Vol. 10, No. 6, 2009 1459
Self Assembly of Coiled-Coil Peptide-
Bashkim Kokona, Andrew M. Kim, R. Claire Roden, Joshua P. Daniels, Brian J. Pepe-
Mooney, Brian C. Kovaric, Julio C. de Paula,
Karl A. Johnson, and Robert Fairman*
Department of Biology, Haverford College, 370 Lancaster Ave, Haverford, PA 19041
Current Address: Department of Chemistry, Lewis and Clark College, 0615 SW
Palatine Hill Rd., Portland, OR 97219
Figure S1. Tapping-mode AFM images of 100 µM Cp3K in 10 mM Tris-HCl, pH 7.8
and 0.1 M NaCl.
Figure S2. CD spectra of Cp3K in α-helix and β-sheet conformations. Cp3K
concentration is 100 µM in 10mM Tris-HCl, pH 7.8 prepared with fresh stock solution
and aged stock solution.
Figure S3. Tapping-mode AFM image of 100 µM Cp3K in cross-β-sheet fibril