Article
pubs.acs.org/JACS
Control of Heme Coordination and Catalytic Activity by
Conformational Changes in Peptide−Amphiphile Assemblies
Lee A. Solomon,† Jacob B. Kronenberg,‡ and H. Christopher Fry*,†
†
Argonne National Laboratory, 9700 South Cass Avenue, Argonne, Illinois 60439, United States
Illinois Math and Science Academy, 1500 West Sullivan Road, Aurora, Illinois 60506, United States
‡
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S Supporting Information
*
ABSTRACT: Self-assembling peptide materials have gained
significant attention, due to well-demonstrated applications,
but they are functionally underutilized. To advance their utility,
we use noncovalent interactions to incorporate the biological
cofactor heme-B for catalysis. Heme-proteins achieve differing
functions through structural and coordinative variations. Here,
we replicate this phenomenon by highlighting changes in heme
reactivity as a function of coordination, sequence, and morphology (micelles versus fibers) in a series of simple peptide
amphiphiles with the sequence c16-xyL3K3-CO2H where c16 is a palmitoyl moiety and xy represents the heme binding region:
AA, AH, HH, and MH. The morphology of this peptide series is characterized using transmission electron and atomic force
microscopies as well as dynamic light scattering. Within this small library of peptide constructs, we show that three
spectroscopically (UV/visible and electron paramagnetic resonance) distinct heme environments were generated:
noncoordinated/embedded high-spin, five-coordinate high-spin, and six-coordinate low-spin. The resulting material’s functional
dependence on sequence and supramolecular morphology is highlighted 2-fold. First, the heme active site binds carbon
monoxide in both micelles and fibers, demonstrating that the heme active site in both morphologies is accessible to small
molecules for catalysis. Second, peroxidase activity was observed in heme-containing micelles yet was significantly reduced in
heme-containing fibers. We briefly discuss the implications these findings have in the production of functional, self-assembling
peptide materials.
■
INTRODUCTION
Controlling catalytic activity through the organized assembly of
molecules is at the heart of many biological processes and
remains a challenge in the field of de novo protein design and
nanoscale architectures.1−9 Natural proteins achieve this
through a chain of amino acids that fold into a threedimensional structure that in turn governs molecular
organization and activity. Achieving complex biologically
relevant reactivity in synthetic materials remains a challenge
because of the need to simultaneously balance properties that
lead to supramolecular assembly while maintaining precise
molecular control in the catalytic active site. Peptide
amphiphiles (PAs), a class of supramolecular biomaterials,
provide a simple solution to this problem.10−12 The peptide
serves as a scaffold that typically includes a recognition,
structural, and functional site. The structural region guides
assembly, while the functional site, typically a modified or
unnatural amino acid or sequence of amino acids, is used in
catalysis. 2,13−23 The peptide, however, can be further
programmed to incorporate added functions, ultimately
generating a protein-like catalytic material. In our efforts to
design biologically inspired materials, we have engineered metal
binding sites into peptide amphiphiles to generate functional
supramolecular assemblies.24,25 Transition metal binding
peptides, heme binding amyloid-β aggregates, and hemoprotein
assemblies have demonstrated catalytic properties.5,26−28 In this
© 2017 American Chemical Society
work, we incorporate the naturally occurring cofactor heme-B,
inspired by the diversity of heme enzymes, to elevate peptide
amphiphiles to a new level of sophistication. We demonstrate a
system where the supramolecular structure alone can control
heme coordination and reactivity. As a result, we present a
peptide-based self-assembling material that functions like a
natural protein.
Heme-B (Fe-protoporphyrin IX) has an impressively diverse
functional library in nature. Uncoordinated, it is toxic due to its
ability to produce reactive oxygen species.29,30 However, when
associated with a protein, function can be focused toward
important metabolic activities. This is due to the heme’s
immediate coordination environment, which is significantly
influenced by the protein structure.31,32 For example, nitrophorin coordinates heme with a single histidine and, due to its
structure, functions as a nitric oxide carrying protein found in
insects, whereas neuroglobin uses a bis-histidine coordination
to bind oxygen in the brain, Figure 1.33−36 Cytochrome c
employs one histidine and one methionine ligand to carry out
high potential electron transfer.37−41 At present, no artificial
material is able to associate with a single cofactor and carry out
such diverse array of functions, but achieving this level of
control would add new dimensions to material applications.
Received: February 17, 2017
Published: May 15, 2017
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are well understood in natural systems, but translating them to
materials and other technologies has proven difficult.42−44
In this work, we not only demonstrate these long-range
conformational changes (i.e., micelle to fiber transition) in
peptide−amphiphile assemblies, but also use them to control
reactivity of our noncovalently bound cofactor. First, we
highlight our ability to vary the heme-binding site through a
single “mutation” within the primary sequence that changes the
coordination environment around the heme, Figure 1. Second,
we present striking evidence that the supramolecular assembly,
micelles versus fibers, significantly influences the heme
coordination mode. Third, we highlight the material’s ability
to bind carbon monoxide, which serves as a redox inactive
surrogate to other biologically relevant gases like O2 and NO
and thus confirms heme active site accessibility for catalysis.
Finally and most intriguingly, we demonstrate our ability to
dramatically reduce peroxidase activity solely on the basis of the
peptide amphiphile supramolecular structure, micelles versus
fibers. These discoveries emphasize the robustness of the
peptide amphiphile in developing next generation, functional,
biomolecular materials.
Figure 1. Protein structure, active site, and design inspiration for
functional heme peptide amphiphiles. Cartoon depictions of crystal
structures for nitrophorin (PDB ID 1ERX), neuroglobin (PDB ID
2VRY), and cytochrome c (PDB ID 3CYT). Details of the primary
coordination sphere highlighting no-coordination, single histidine, bishistidine, and histidine-methionine. Idealized binding of the designed
heme-binding peptide amphiphiles in a β-sheet conformation with
their abbreviated names and sequence. Color coding for PA molecules:
gray, palmitoyl/c16; yellow, alanine; orange, methionine; red,
histidine; green, leucine; and blue, lysine.
■
RESULTS AND DISCUSSION
Design. The peptide−amphiphiles were designed to
emulate naturally occurring heme active sites by reproducing
the coordination environments shown in Figure 1. The
peptides follow the simple design, c16-xyL3K3-CO2H. We
have employed our rational design strategy from our previous
study where c16 is a palmitoyl moiety that was included to
promote hydrophobic collapse, the first step in peptide
amphiphile assembly. The positively charged trilysine (K3)
headgroup is introduced as a pH trigger to induce fiber
assembly. In other words, raising the pH close to the pKa of the
lysine residues reduces electrostatic repulsion, allowing
individual peptide molecules to come within van der Waals
distances. Three leucine amino acids (L3) serve as a β-sheet
structural motif that assists in the formation of long-aspect ratio
nanofibers. The heme binding site is represented by x and y,
where y is typically histidine (H), the most common hemebinding amino acid in nature, and x represents the site
employed to vary the coordination state: A, M, or H (Figure 1).
As a result, the binding site xy is generated (AA, AH, HH, or
In addition to coordination, neuroglobin, cytochrome c, and
hemoglobin further control heme-function through large-scale
conformational changes.42−44 For example, cytochrome c in its
native state is an electron transfer protein, but it is also
overexpressed in cancerous cells where a conformational
change leads to a functional change from an electron transfer
protein to a peroxidase. Korendovych et al. were able to
redesign a natural protein with existing conformational changes
to engineer a switchable eliminase.45 Similarly, Grosset et al.
engineered allosteric rearrangements in a de novo protein,
using heme as a redox-switch, but were unable to couple that to
a function.46 The triggers and effects of long-range interactions
Figure 2. Characterization of supramolecular morphologies in different aqueous solutions. Atomic force micrographs (2 μm × 2 μm) of HH
assemblies in HEPES (A) without and (B) with hemin and in 10 mM NH4OH (C) without and (D) with hemin. The height profiles to the right of
the micrographs are measurements of individual micelles or fibers without (black) and with hemin (red).
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drive the micelle to fiber transition (Figure S5). AFM highlights
that when heme is coordinated/embedded in the peptide fibers
(10:1 peptide:heme), all xyHeme peptide assemblies have the
same height profile, 7−8 nm (Figure S3).
A relationship between morphology and secondary structure
is also noted. In HEPES buffer where micelles are observed,
FT-IR analysis indicates that AH and HH lack a well-defined
secondary structure as observed by the broad vibrational
modes. MH exhibits a β-sheet component with vibrational
modes at 1630 and 1681 cm−1. The presence of β-sheets in MH
is somewhat anomalous as β-sheets are often linked to fiber
formation. This relationship between β-sheets and fibers is
observed in AA in HEPES buffer where a high degree of βsheet content is observed with an intense amide I vibration at
1630 cm−1 and a less intense band at 1681 cm−1, Figure S6a.58
We then added heme (at a ratio of 10 peptides:1 heme to
ensure binding) to see if it has an effect on structure. Upon
addition of heme, no significant change in secondary structure
was observed for any of the peptide assemblies (Figure S6b)
consistent with the lack of change in observed morphology.
At pH 10.5, we detect the formation of β-sheets by FTIR and
CD (Figures S6C and S7, respectively) with or without heme
(10 peptide:1 heme, Figure S5D). The pKa of lysine is close to
10.5 and becomes neutralized, allowing neighboring molecules
to interact and form β-sheets. The FTIR indicates that all xy
assemblies, both apo and holo, at higher pH yield amide I
vibrations (1630 and 1681 cm−1) consistent with β-sheets. We
note that the CD spectra do not yield signature β-sheet spectra
with a minimum at 218 nm. The red-shifted spectra (λmin =
218−229 nm) (Figure S7) are attributed to a superhelical twist
within the fiber construct, and the observed variations in
superhelicity (i.e., degree of red shifting) are influenced by
changes in the peptide sequence.59 Therefore, in the xy peptide
series, a lack of well-defined secondary structure at neutral pH
in HEPES buffer directly correlates to spherical micelle
structures, whereas β-sheet formation at high pH directly
relates to long aspect ratio fibers in bundled networks. In both
micelles and fibers, the overall structure is largely unaffected by
the addition of heme. As a result, this peptide series allows the
examination of how both the primary amino acid sequence of
the peptides and the supramolecular assembly influence the
heme cofactor binding and function.
Ferric Heme Binding. To investigate how PA sequence
and morphology influence heme coordination environment, we
employed electronic absorption (EA) spectroscopy and
electron paramagnetic resonance (EPR) spectroscopy that
together help describe the mechanism of heme insertion, the
ligand environment, and the spin state of the metal-centered
cofactor, Figure 4 and Table 1.
Heme Coordinated to Micelle Characterization
(HEPES, pH 7). For spectroscopic characterization, the
peptide:heme stoichiometry was maintained at 10:1 to ensure
complete heme coordination. In HHHeme micelles at neutral
pH, the electronic absorption spectrum yields signature Soret
(λmax = 413 nm) and Q-band (λmax = 535, 560 nm) values
consistent with bis-histidine axial coordination to heme, in
agreement with the spectrum for many bis-histidine coordinating proteins including neuroglobin (Figure 4A, Table 1).60 The
EPR spectrum represents a purely S = 1/2, low-spin, type II
(rhombic) spin configuration (Figure 4C, Table 1) and is
characteristic of many low-spin, bis-histidine coordinated heme
proteins including neuroglobin.61−64 AAHeme (fibers at neutral
pH) does not offer a histidine containing coordination site and
MH) and is further denoted as xy (e.g., AH) in the apo state
(uncoordinated heme) or xyHeme (e.g., AHHeme) in the holo
state (coordinated heme). For the initial design visualization,
we assumed the formation of parallel β-sheets, typical for
amphiphiles (Figure 1). AHHeme was designed to have axial
ligation similar to that of the β-sheet-rich nitrophorin with a
single histidine available for coordination (Figure 1). Other
examples of single histidine coordinated hemes are horseradish
peroxidase as well as the peptide-heme microperoxidases that
will be discussed later.47−50 The second peptide in our set is
HHHeme; the bis-histidine coordination is similar to neuroglobin, cytochrome b proteins (Figure 1),51 and a number of de
novo designed, α-helical bundle peptides.52−56 Third, MHHeme
was designed to offer a His-Met axial ligation similar to that
found in cytochrome c. Finally, we employ AAHeme as an
uncoordinating control peptide to monitor any background
heme activity. These peptides produce a modest library that
highlights the ability to tune heme-coordination and function
within a peptide−amphiphile material through simple alterations in the primary coordination environment.
Supramolecular Characterization. To test the hypothesis
that supramolecular assembly has potential to effect the
coordination environment surrounding the heme, it was crucial
to analyze the morphology under various conditions. We find
by transmission electron microscopy (TEM), atomic force
microscopy (AFM), and dynamic light scattering (DLS) that
the peptide−amphiphiles in HEPES buffer at pH 7 yield
spherical micelles, Figure S1 (with the exception of AA). AFM
shows that the spheres are ∼3−7 nm in diameter in both the
presence and the absence of heme (at a ratio of 10 peptides:1
heme, Figures 2A,B and S1). Furthermore, DLS experiments
suggest ∼7 nm diameter micelles in close agreement with the
microscopy data (Figure S2). The spherical micelle formation is
attributed to the large palmitoyl (c16) tail of the peptide−
amphiphiles undergoing hydrophobic collapse in concert with
electrostatic repulsion of the polar headgroup lysine residues.
AAHeme was the only peptide to deviate from the spherical
assembly where AFM, TEM, and DLS all showed fibers at
neutral pH, Figure S1E. We presume that the lack of a bulky
histidine residue at the aliphatic interior of the assembly
eliminates steric repulsion, thus allowing the formation of fibers
to occur.
In 10 mM ammonium hydroxide at pH 10.5, the amine
group on lysine is neutralized, thus decreasing the effect of
electrostatic repulsion resulting in a micelle to fiber transformation (Figure 3)57 that was observed in all peptides studied
as indicated by TEM and AFM (Figures 2C−D, S3, and S4). In
addition, we employed HEPES at pH 10.5 (outside of the
buffer range) to emphasize that HEPES, as a molecule, does not
prevent the formation of fibers and that pH alone is adequate to
Figure 3. A cartoon representation of the spherical micelle formation
in HEPES buffer at pH 7 and long aspect ratio nanofibers in 10 mM
NH4OH at pH 10.5.
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Table 1. Summation of Heme Coordination Characterization by Electronic Absorption Spectroscopy, Electron Paramagnetic
Resonance Spectroscopy, and Binding Constant Analysis of the PAs, xyHeme, in Fiber (10 mM NH4OH, pH 10.5) and Micelle
(HEPES, pH 7.0) Morphologiesa
a
EAS and EPR values of proteins with targeted coordination environments are listed for comparison. H.S. = high spin, L.S. = low spin.
therefore yields a spectrum comparable to that of free heme,
Figure 4A, Table 1. Free heme in aqueous solutions readily
forms aggregates yielding dramatically blue-shifted visible
spectra from solubilized heme (Table 1).65,66 Furthermore,
AAHeme yielded an entirely S = 5/2, high-spin EPR spectrum
(Figure 4C) similar to that of free heme (Figure S8). In
AAHeme, we suggest that heme aggregation is broken up and the
molecule is solubilized (note: heme axial ligation is fulfilled by
the coordination of a water molecule or hydroxide anion) but
not coordinated by the peptide producing both the red shift in
the EA spectrum (in relationship to aggregated heme) and the
signature high-spin spectrum from EPR, Table 1.
Despite having one less histidine in its sequence than
HHHeme, AHHeme micelles yield a bis-histidine heme coordination as the observed EPR spectrum is predominantly low-spin
with g-values identical to those of HHHeme (Figure 4C). On the
other hand, MHHeme micelles yielded a mixture of coordination
states: a predominantly high spin, S = 5/2, EPR spectrum and
an observable low-spin, bis-histidine contribution (Figure 4C).
Consistent with the EPR data, the EA spectra for AHHeme and
MHHeme micelles suggest a mixture of uncoordinated and bishistidine coordinated states (Figure 4A). This observation is
consistent with the Soret and Q-band position and intensities,
which represent averages between the spectra for bis-histidine
heme coordination in HHHeme and embedded heme in AAHeme.
The presence of bis-histidine axial ligation in AHHeme and
MHHeme micelles is due to the greater degree of flexibility
within the micelle assembly as compared to the rigid structure
of the β-sheet fibers. Thus, bis-histidine coordination in
micelles occurs with any peptide−amphiphile in our series
that has a histidine, for example, xHHeme.
Heme Coordinated to Fibers Characterization (pH
10.5). Next, we explore how a rigid supramolecular structure
affects the heme-binding site. In fibers at high pH, the
electronic absorption spectra indicate unique Soret and Q
bands for each peptide, suggesting a variation in coordination
environment from one sample to the next (Figure 4B and
Table 1). Heme coordinated to HHHeme fibers exhibits a
predominant signal attributed to bis-histidine coordination
environment, similar to that of micelles, as indicated by the EA
and EPR spectra (Figure 4B and D and Table 1). Note:
Impurities of free heme are also detected, but the EAS is
consistent with low-spin heme. Because AAHeme at neutral and
high pH offers the same morphology, no significant
spectroscopic changes were observed (Figure 4B and D and
Table 1). We again suggest a lack of axial coordination to the
peptide while the heme is fixed/solubilized in the matrix of the
assembled fiber. Interestingly, MHHeme fibers yield spectra
similar to those observed for AAHeme fibers, suggesting that
heme is not coordinated but embedded in the peptide
Figure 4. Characterization of ferric heme coordination to different
supramolecular constructs. Electronic absorption spectroscopy of
heme (100 μM) bound to peptide (1 mM) at (A) pH 7.0 and (B)
pH 10.5. EPR spectroscopy of heme (1 mM) bound to peptide (10
mM) at (C) pH 7.0 and (D) pH 10.5. Vertical lines and labels mark
the high-spin (dashed lines) and low-spin (solid lines) states. AAHeme,
blue; AHHeme, red; HHHeme, green; and MHHeme, purple.
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then is available, the heme molecule will coordinate. The
binding constant analysis determines which coordination site is
the strongest, but most importantly highlights the significant
influence the rigid secondary structure has over heme binding
affinity.
The EA spectra and EPR data found for heme coordination
to the xyHeme series highlight a strong link between peptide
sequence, molecular ordering, morphology, and heme coordination. HHHeme highlights our ability to design a peptide that
maintains bis-histidine coordination when converting from
micelles to fibers. MHHeme indicates that the bulky methionine
side chain upon the ordering/formation of β-sheet rich fibers
effectively blocks the histidine residues available for heme
coordination in a micelle. Finally, AHHeme highlights a change
in coordination environment concomitant with a morphological
shift from (1) micelles yielding low-spin, bis-histidine
coordination to (2) high density β-sheet containing fibers
providing high-spin, single histidine axial ligation. These pHdependent changes serve as highly programmable features for
the development of functional heme peptide materials.
Electrochemistry. Redox behavior is another aspect where
the binding site exerts control over heme in both natural and de
novo proteins.32,50,52,53,70−72 Here, we demonstrate the ability
to change the redox behavior of heme in our peptide materials
through sequence and structure. In micelles, where the peptides
exhibit similar coordination environments, AHHeme, HHHeme,
and MHHeme exhibit similar midpoint potentials (EM) versus
SHE: −315, −270, and −312 mV, respectively (Figure S10).
When we analyzed the peptide fibers, the trend changed, such
that AHHeme, HHHeme, and MHHeme all exhibit very different EM
values versus SHE: −655, −333, and −442 mV, respectively
(Figure S10). We attribute these results to the established
variation in coordination environment between the micelle and
β-sheet-rich fibrous morphologies. For example, there is
negligible change in coordination state between HHHeme in
micelles versus fibers, which extends to a minimal change in EM,
but AHHeme yields a dramatic change that we attribute to
different coordination modes, bis-histidine in micelles versus
single-histidine in the β-sheet fibers.
Gas Binding. Heme proteins often bind and transport small
molecules like water, dioxygen (O2), or nitric oxide (NO) using
coordination bonds. Each of these small molecules plays a vital
role in signaling and catalysis, and when combined with the
well-established properties of peptide amphiphiles produces
multivariate functional materials that could be used in
vasodilation (NO),73 neurotransmission (NO and CO),74,75
and O2 delivery or activation.76,77 We have chosen to explore
carbon monoxide (CO) gas binding because CO serves as a
redox inactive surrogate to these gases binding to ferrous (Fe2+)
heme where (1) it offers major insight into small molecule
accessibility of the xyHeme active site relevant to enzymatic
activity and (2) provides information on how the peptide
sequence can affect gas binding.78 The CO adduct was obtained
by first chemically reducing the heme molecule in the
peptide:heme assembly with a small amount of sodium
dithionite in a nitrogen box. The sample then was transferred
to a cuvette equipped with a rubber septum. Finally, the sample
was sparged with CO for 30 s. UV/visible spectroscopy yields
similar spectra when CO is coordinated to a histidine axially
ligated ferrous heme, Figure S11. Therefore, infrared spectroscopy is employed because the heme binding pocket and ligand
coordination directly influence the vibrational frequency of CO
(νCO) either through enhanced backbonding or through ligand
assembly. This observation is further corroborated by the
purely high-spin EPR spectrum and is in opposition to the
mixture of high- and low-spin heme observed in MHHeme
micelles. It should be noted that cytochrome c, which
coordinates heme through histidine-methionine ligation, yields
a low-spin, type I, highly anisotropic low-spin (HALS)
spectrum with a g value of ∼3.3, further suggesting that we
did not achieve the desired histidine-methionine coordination
in MHHeme (Figure 1).67 We attribute the lack of heme
coordination to the increased molecular ordering (i.e., β-sheets)
within the assembly where the bulky methionine residue
sterically blocks heme access to the histidine coordination site.
Finally, heme coordinated to AHHeme fibers yields an EA
spectrum indicative of coordinated heme but not typical of bishistidine axial ligation as it is blue-shifted to a value similar to
that of nitrophorin, which possesses single histidine axial
ligation (Figure 4B and Table 1). The EPR spectrum shows a
predominantly high-spin species as expected for single histidine
axially coordinated heme like nitrophorin.64,68 The value
observed at gz = 3.71 is typical of a low-spin type I (HALS)
spectrum and is observed in our control experiment where
heme is analyzed in the presence of lysine (Figure S8).63 We
conclude that the observed spectrum for heme coordinated to
AH Heme fibers is predominantly a high-spin spectrum
representative of a single histidine-coordinated heme with
some propensity for unresolved low-spin states.
Binding Constant Analysis. Peptide-to-heme stoichiometry and binding constants (Table 1) obtained from a series of
titration experiments indicate a dependence on the flexible
micelle structures when compared to the rigid fiber structures,
Figure S9. We employed a binding analysis method typically
used to analyze heme or transition metal binding to de novo
designed proteins, eq S1.69 We modified the equation to yield a
value n that represents the minimum number of peptides
required to bind one heme. The results for micelles suggest a
3:1 (peptide:heme) stoichiometry, while fibers yield a 6:1
stoichiometry (Figure S9). Within the micelle construct, the
lack of secondary structure yields a more flexible peptide
environment, allowing for the more favorable bis-histidine
coordination. As a result, the highest binding affinity (lowest
Kd) is simply the peptide with more available histidines,
HHHeme. MHHeme in micelles yields a less favorable bis-histidine
coordination environment than AHHeme due to steric crowding
from the bulky thioether at the heme-binding site.
In contrast to the favorable bis-histidine coordination
environment in micelles, the β-sheet-rich fibers offer a rigid
structure that contributes significantly to heme coordination
and binding affinity. While the HHHeme binding affinity is
similar to that found for micelles, AHHeme binding affinity
actually increases (lower Kd) despite the morphologically
induced change to a single-histidine coordination environment.
This increase in binding affinity is promoted by the rigid fiber
structure and consequently the nonbulky alanine in the “distal”
position of the heme-binding pocket. As a result, the opposite
trend is observed in MHHeme, where the steric crowding of the
bulky methionine that was observed in micelles is amplified by
the rigid system and is reflected in the decrease in heme
binding affinity (increased Kd). Despite the lack of a histidine
containing coordination site, AA Heme does produce a
“solubilization” curve, suggesting heme incorporation within
the fiber construct. This helps to explain the overall mechanism
of heme insertion such that the amphiphilic micelle or fiber first
encapsulates the hydrophobic heme molecule. If a binding site
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molecular construct immediately surrounding the heme
molecule yielding a distribution of states. We focus our
discussion on the major contributing peak. In the case of
HHHeme, we assume one of the histidine ligands becomes
uncoordinated, allowing CO binding to occur, yielding a distal
histidine similar to that found in myoglobin. Again, the value
obtained for HHHeme, νCO = 1961 cm−1, is higher than values
obtained for myoglobin, but it is slightly lower than the value
obtained for the micelle conformation. We suggest that the
slight increase in heme−CO back-bonding within the fiber
assembly is due to the presence of a distal histidine in a more
rigid environment. When we produce a similar coordination
environment to the distal-site mutants of myoglobin (e.g., His
→ Ala), we observe an increase in stretching frequency for
AHHeme, νCO = 1971 cm−1, slightly greater than that found in
the micelle structure for AHHeme. This is likely due to the lack
of secondary structure in the micelles allowing uncoordinated
histidine residues to interact with the heme−CO complex in
the pocket, yielding a slightly lower νCO. As a result, the
HHHeme CO frequency in fibers is 10 cm−1 lower than that
found for AHHeme. We suggest that in the β-sheet enriched
fibrous assemblies, CO serves as an excellent probe of the heme
active site, indicating enhanced back-bonding due to a distal
histidine in the case of HHHeme and reduced back-bonding due
to the aliphatic alanine in the case of AHHeme.
CO vibrational analysis for MHHeme is similar to that for
AAHeme. This is consistent with the comparable electronic
absorption spectra (Figure S11). While MHHeme was able to
coordinate heme in micelles, the CO data are consistent with
the lack of ferric heme binding to MH fibers. That is, the heme
embeds itself in the fiber but without axial ligation to histidine.
When comparing the CO vibrational spectra for AAHeme and
MHHeme, a slight increase in the contribution of νCO = 1964
cm−1 is noticeable that may suggest some axial histidine
coordination.
The CO binding studies clearly highlight that the heme
binding PAs are capable of binding gases like CO. Furthermore,
detailed vibrational analysis highlights and supports our claims
of controlling the coordination environment through careful
sequence design and morphology. These results provide a basic
understanding and some guiding principles for designing hemebinding peptides that can catalyze or transport and release gas
on the basis of environmental triggers.
Supramolecular Control of Enzymatic Activity. Many
natural and synthetic heme proteins display peroxidase activity,
a natural reaction that catalyzes the oxidation of high potential
substrates using hydrogen peroxide (H2O2).31 We acknowledge
the existence of other peptide:heme complexes, like microperoxidase (MP-11),79 de novo designed heme binding
peptides,80−83 and even the demonstration of an active
amyloid-β peptide,28 as well as heme solubilized micelles that
yield peroxidase activity,84−88 but these systems do not probe
the effects of supramolecular morphology. Here, we employed
heme-peroxidase catalysis as a benchmark reaction to
investigate specifically how supramolecular morphology in our
PA−heme assemblies can control catalytic reactivity.89 We have
chosen the established protocol where H2O2 activated by the
xyHeme assemblies reacts with the colorless molecule 3,3′,5,5′tetramethylbenzidine (TMB) to yield the single electron
oxidation product, TMB•+ (λmax = 652 nm) and water.90 All
of these assays were carried out with 30 μM peptide, 1 μM
heme in HEPES buffer at pH 7. Peptide assemblies were first
prepared in either pH 7 HEPES (micelles) or 10.5 NH4OH
coordination enhancement/disruption as indicated in the wide
range of observed stretching frequencies, νCO = 1949−1971
cm−1, Figure 5 and Table 2.78
Figure 5. Carbon monoxide vibrational analysis probing enzymatic
capability and molecular structure. Infrared spectroscopy of heme−CO
binding in the CO stretching frequency region at pH = 7 and pH =
10.5. AAHeme, blue; AHHeme, red; HHHeme, green; and MHHeme, purple.
Hemin, 100 μM; peptide, 1 mM.
Table 2. Summary of CO Vibrational Analysisa
pH = 7
peptide
AAHeme
AHHeme
HHHeme
MHHeme
νmaj (%)
−1
1964 cm
(57%)
1967 cm−1
(87%)
1963 cm−1
(96%)
1969 cm−1
(80%)
pH = 10.5
νmin (%)
−1
1948 cm
(43%)
1948 cm−1
(13%)
1974 cm−1
(4%)
1953 cm−1
(20%)
νmaj (%)
−1
1965 cm
(53%)
1971 cm−1
(58%)
1961 cm−1
(86%)
1964 cm−1
(63%)
νmin (%)
1948 cm−1
(47%)
1955 cm−1
(42%)
1977 cm−1
(14%)
1951 cm−1
(37%)
a
The major (νmaj) and minor (νmin) vibrational frequencies and their
corresponding percent contributions are derived from Gaussian peak
fitting (Figure S12).
For micelles at pH = 7, AHHeme, νCO = 1967 cm−1, HHHeme,
νCO = 1963 cm−1, and MHHeme, νCO = 1969 cm−1 yield similar
values (Figure 5, Table 2) due to their similar coordination
environments consistent with EAS and EPR. For CO to bind in
the micelles, one histidine must dissociate, resulting in a distalhistidine ligand. The observed vibrational frequencies are
notably higher than those obtained for the analogous
coordination environment of myoglobin at neutral pH78 (νCO
= 1947 cm−1) likely because the micelle assembly lacks the
more sophisticated structure of a fully folded protein.
Mutations to various residues in the myoglobin active site
result in an observed decrease in CO-heme backbonding,
evidenced by an increase in stretching frequencies,78 νCO =
1965−1971 cm−1, consistent with the measured values here. In
the case of AAHeme where fibers are formed regardless of
environment and no discernible coordination to the peptide is
observed, two vibrational states were found, νCO = 1948 and
1964 cm−1, with the former yielding the most intense peak,
which is consistent with a five-coordinate heme−CO with no
axial ligation to the peptide.
Interestingly, with xyHeme fibers, which greatly influence the
coordination environment of ferric heme, we observe different
CO vibrational frequencies for each assembly (Figure 5). In
general, the signals are broader or split when compared to the
data at pH 7. This is most likely due to slight variations in the
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Table 3. Michaelis−Menten Parameters for the c16-xyL3K3-CO2H Peptide Series Comparing Micelles and Fibersa
a
Free heme and MP-11 parameters are included for comparison. The values determined are from TMB oxidation with respect to varying the H2O2
concentration.
(fibers) with a 30-fold excess of peptide to heme (1.5 mM
peptide, 50 μM heme). This 30:1 peptide:heme ratio (as
opposed to the 10:1 peptide:heme used in material characterization) was employed to ensure that, upon a 50-fold dilution
into HEPES, the heme would stay coordinated to the assembly.
As a result, the observed peroxidase reactivity is generated by
the heme bound to the assembly and not free heme, which
displays peroxidase activity in the absence of peptide. The
dilution process does not alter the peptide micelle and fiber
structures (Figure S13), and heme remains coordinated (Figure
S14). We also monitored free heme activity as well as
microperoxidase to generate comparisons to an unprotected
heme molecule and an optimized peroxidase.
Michaelis−Menten kinetic analysis was performed to
compare the peroxidase reactivity of micelles and fibers with
respect to varying peroxide concentration (Figures S15 and
16), and the results are summarized in Table 3. We also
measured the kinetics with respect to varying TMB
concentration to indicate that our reported values are under
saturated TMB conditions (Figure S17). Bleaching of the heme
was not observed during the course of the experiments (Figure
S18,19). The heme binding micelles, AHHeme and MHHeme,
exhibit similar degrees of peroxidase activity, whereas their
fibrous assemblies display a weak, baseline level of reactivity
(Figure 6A,B). AHHeme was found to have the most dramatic
difference between the two morphologies with the turnover
number (kcat) being 16 times greater in micelles. Because of the
relatively high KM in micelles versus fibers, the overall catalytic
efficiency (kcat/KM) was found to be only 5 times greater in
AHHeme micelles. We found a similar trend with MHHeme in
which the turnover number (kcat) for micelles was 6 times
greater than fibers. Turnover for HHHeme in micellar form was
only 3 times greater than its fiber counterpart. In fact, when
compared to the other peptides, HHHeme micelles exhibited a
decrease in turnover while the fibers increased (Figures 6A,B
and S16). This increase in kcat for HHHeme fibers may be
attributed to a higher redox potential that is directly associated
with its preferred low-spin, bis-histidine axial ligation. While
AHHeme, HHHeme, and MHHeme micelles coordinate heme
through bis-histidine axial ligation with comparable redox
potentials, their difference in reaction rates may be attributed to
the heme affinity. That is, HHHeme binds heme with the greatest
affinity (Table 1) and shows the slowest rate of reaction among
the micelles. For peroxidase activity to function properly, H2O2
must first react with the ferric heme before generating the active
oxo-ferryl intermediate, compound I. If the bis-histidine
coordination is strong enough, the reactivity would be slower
because one of the histidines would have to dissociate prior to
Figure 6. Testing of peroxidase activity at pH 7 exhibits dependence
on sequence and supramolecular morphology. Peroxidase activity, as
seen by the oxidation of TMB, mediated by peptide−amphiphile series
in either a (A) micelle or (B) fiber morphology. Heme activity with no
peptide is represented by the dashed black line. Reaction conditions:
peptide = 30 μM, heme = 1 μM, H2O2 = 6 mM, and TMB = 300 μM,
HEPES buffer pH 7. (C) Representative Michaelis−Menten analysis
for AHHeme, in micelles (solid red line) and fibers (dashed red line).
Additional Michaelis−Menten analyses are available in Figure S15.
(D) Total TMB oxidized upon completion of the peroxidase reaction.
Solid bars represent micelles, and hashed bars represent fibers. Color
code: blue, AA; red, AH; green, HH; purple, MH; and black, free
heme.
reacting with H2O2. This binding affinity-based argument may
hold for comparing the peptide micelles, but the fiber
assemblies display a lower and baseline level of peroxidase
reactivity with lower kcat and Km values. We therefore attribute
the lowered activity specifically to the change in morphology
from micelles to fibers, which, depending on the peptide
sequence (e.g., AH), may involve a change in coordination
environment that leads to a dramatic change in overall
reactivity.
For comparison, we measured the peroxidase activity of
heme alone under our reaction conditions to highlight the
effect of the peptide on peroxidase activity. Qualitatively, the
peptide micelles enhance the catalytic ability of heme, while
fibers decrease the activity (Figure 6A and B). Michaelis
Menten kinetic analysis indicated that kcat/KM for free heme is
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sickle cell disease, ischemia reperfusion, and malaria).30 When
that micelle material reaches the target, this stimuli responsive
material can react with the environment, triggering a change in
morphology from micelles to fibers, all while sequestering and
thus detoxifying heme by limiting its ability to generate reactive
oxygen species. Furthermore, the well-established regenerative
properties of PAs can be employed to provide a scaffold and
stimulate the growth of healthy tissue.10
greater than that found for the peptide micelles, and kcat was
lower suggesting that the heme molecule does not catalyze
TMB over long periods of time and is evident in our analysis of
total TMB oxidation (see below). This suggests that, while
seemingly faster, the overall catalytic performance was weak.
We also compared the peptide micelles with heme solubilized
in surfactant micelles (heme in 1 wt % Tween 20, 10 nm
micelles and 10 wt % SDS, 3 nm micelles) to highlight the
peptides role in catalytic performance. Peroxidase activity of
heme solubilized in surfactant micelles leads to slower TMB
product formation than peptide micelles, Figure S20. In
addition, inconsistent generation of both one- and two-electron
TMB oxidation products in Tween and SDS micelles
containing heme was not comparable to the results found for
our peptide assemblies, suggesting that the heme coordinating
ability of the peptide micelles plays an impactful role in
generating controlled peroxidase activity. In contrast, peroxidase activity of microperoxidase-11 (an isolated heme-peptide
segment from peroxidase) under our reaction conditions
yielded a kcat/Km value ∼4000 times greater and a kcat value
∼500 times greater than the baseline activity of the fibers
highlighting the significantly decreased catalytic ability of the
fiber construct.
We measured the final product concentration to compare the
total amount of TMB oxidized (Figure 6D). We acidified our
samples to yield the final two-electron oxidation product,
TMBox, and measured the absorbance at 450 nm (ε = 59 000
M−1 cm−1). Our results are consistent with the kinetic data in
that AHHeme and MHHeme were the best performers yielding
117 and 110 μM of TMBox, respectively. HHHeme produced 76
μM of TMBox. The fiber assemblies yielded a baseline amount
of TMBox (10−25 μM) similar to that of our control example
of heme (no peptide) in HEPES. HHHeme fibers represent an
exception to this trend yielding 52 μM of TMBox. Consistent
with the Michaelis−Menten kinetics, HHHeme fibers exhibit
more activity than do the other fibrous assemblies. Therefore,
our peptides demonstrate slow peroxidase activity as micelles,
but when analyzed for peroxidase activity as fibers, they
typically exhibit baseline reactivity similar to that found for our
control sample with just heme. The most dramatic change in
reactivity is within the AHHeme system in which the micelle
heme peroxidase activity is enhanced, whereas the fibers
significantly limit the reactivity most notably characterized by
the 16-fold decrease in turnover number when comparing
micelles to fibers.
Many heme-containing proteins and peptide assemblies
display peroxidase activity. In fact, myoglobin has been
demonstrated to yield peroxidase activity,91 and, as mentioned
earlier, cytochrome c in the presence of cancerous cells has
been proposed to change from an electron transfer protein to a
peroxidase.41 Here, we demonstrate a similar phenomenon in
that, with the same peptide (AHHeme), we can modulate the
reactivity just by changing the morphology from micelles to
fibers, a potentially useful mechanism in a peptide amphiphile
material. We acknowledge that our peptide micelles do not
make great peroxidases, as indicated by our comparison with
microperoxidase (i.e., the turnover frequency, kcat, and catalytic
efficiency, kcat/KM, greatly exceed the values found for our
peptide micelles). We suggest here that significantly limiting
peroxidase reactivity of heme is a potential route to detoxify
free heme in medicinal applications. In other words, we can
imagine beginning with or delivering the apo-micelle to treat
diseases that result in excess of free heme or hemolysis (e.g.,
■
CONCLUSION
We have clearly demonstrated the ability to control heme
coordination and function through peptide sequence design
and supramolecular structure. Morphological control is
exhibited through changes in buffer choice and pH; for
example, micelle and fiber assemblies can be formed. We have
highlighted that different heme coordination environments with
varying affinities are observed depending on the morphology
and primary sequence/designed-binding site: AHHeme complexes where the micelle conformation yields low-spin, bishistidine ligation but the fiber yields high-spin, single-histidine
coordination. With regard to eliciting function, we have
highlighted the ability of the material to coordinate the redox
inactive surrogate small molecule, carbon monoxide, which
highlights both the active site accessibility of the heme and the
ability of the heme to bind and potentially transport this
molecule crucial to neurotransmission, and vasodilation.
Finally, we have found a strong influence of supramolecular
morphology on peroxidase activity where micelles exhibit
enhanced catalytic activity over fibers composed of the same
peptide that exhibit baseline catalytic activity. The catalytic
activity in micelles can be further tuned through the primary
sequence, with AHHeme and MHHeme displaying the highest
catalytic efficiencies in the presented series. The function of the
heme is crucial to producing advanced peptide materials and
will be explored in future work where studies on more complex
assemblies are underway.
This peptide amphiphile system provides multiple avenues
with which to control potential enzymatic activity that can
ultimately be translated to the material’s functional properties.
Sequence can be used to modulate the enzymatic rate, while
gross structure can act as an on/off switch allowing for tuning
of the reactivity as a function of environment. These results
significantly impact molecular design strategies for functional
peptide materials where we have discovered that supramolecular structure plays an essential role dictating heme
function. For example, we will investigate fibrous structures that
have potential use in anti-inflammatory signaling where the
peptide assembly could be employed to sequester and break
down toxic free-heme, resulting from sustained injuries (similar
to the protein heme oxygenase I) while simultaneously
exploiting known peptide amphiphile technologies that
promote healthy tissue regeneration.92 Overall, we have
demonstrated a new means of functionalizing current
peptide−amphiphile technologies through the empirical design
of peptides with engineered conformational changes that
influence metal cofactor active sites resulting in controlled,
protein-like, peptide materials.
■
MATERIALS AND METHODS
Peptide Synthesis, Purification, and Characterization. The
synthetic procedure for c16-AHL3K3-CO2H has been reported in our
previous studies.24 The syntheses of c16-AAL3K3-CO2H, c16HHL3K3-CO2H, and c16-MHL3K3-CO2H, cleavage from the resin,
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and RP-HPLC purification followed the same strategy as the
previously reported peptide. MALDI-TOF MS (Bruker UltrafleXtreme
MALDI-TOF) was used to identify the peptides; c16-AAL3K3-CO2H:
calcd for C58H111N11O10 + [H+], 1122.85; found, 1122.933 m/z. c16AHL3K3-CO2H: calcd for C61H113N13O10 + [H+], 1188.87; found,
1189.006 m/z. c16-HHL3K3-CO2H: calcd for C64H115N15O10 + [H+],
1254.90; found, 1255.023 m/z. c16-MHL3K3-CO2H: calcd for
C63H117N13O10S + [H+], 1247.88; found, 1248.864 m/z, Figure S21.
Sample Preparation. Each peptide (3−4 mg) was dissolved in
nanopure water (Millipore A10) to obtain a 1 wt % solution, c16AAL3K3-CO2H (1 wt %, 8.9 mM), c16-AHL3K3-CO2H (1 wt %, 8.4
mM), c16-HHL3K3-CO2H (1 wt %, 7.9 mM), and c16-MHL3K3CO2H (1 wt %, 8.0 mM). Hemin (Porcine, Sigma-Aldrich) was
dissolved in DMSO (Sigma-Aldrich) to achieve a 10 mM stock
solution. Note: Hemin/DMSO stock solutions were always made to
ensure that the final DMSO concentration in the sample was less than
1% (v/v). Typically, 38 μL of a 1 wt % stock solution was dissolved in
260 μL of either HEPES (50 mM HEPES, 100 mM NaCl, pH 7.0) or
10 mM NH4OH, pH 10.5, to yield a 1 mM sample. The samples were
then heated to 65 °C for 10 min and cooled back to room temperature
to ensure formation of the supramolecular assembly. After the sample
was cooled, 3 μL of the 10 mM hemin stock solution was added to the
sample to yield 100 μM hemin. The samples were again heated to 65
°C for 10 min and cooled back to room temperature to ensure
complete heme coordination. Titration experiments analyzed samples
that contained 50 μM heme in either HEPES buffer or 10 mM
NH4OH. Preassembled peptide was added to individual solutions
containing heme such that the peptide concentration ranged from 0 to
1000 μM in 50 μM increments. The samples were equilibrated at
room temperature for 1 h prior to UV/visible measurements. The
experimental data were fit using a modified equation (see Supporting
Information)69 to analyze for binding stoichiometry (n) as well as
binding constant (Kd).
Microscopy. Scanning electron micrographs were obtained with a
JEOL 7500 field emission scanning electron microscope equipped with
a transmission electron detector. Samples of varying concentrations
were diluted 100-fold in water and drop cast onto a 400 mesh copper
grid with a carbon support film (Ted Pella). After 1 min, the excess
solution was wicked away and the sample was air-dried.
Atomic force microscopy (AFM) images were obtained with a
Veeco MultiMode 8 scanning probe microscope equipped with a
silicon nitride tip for imaging soft-materials. The sample was prepared
by drop casting 100 μL of a 200 μM (peptide) sample on freshly
cleaved mica (Ted Pella) and allowed to incubate for 20 min. The
excess sample was wicked away with filter paper, and the sample was
dried prior to measurements.
Secondary Structure Analysis. To analyze secondary structural
formation in the absence of heme, circular dichroism spectroscopy
(Jasco, Inc. J-815) was employed to analyze the typical n−π*
transitions found for a β-sheet assembly. Samples were prepared by
diluting the 1 mM peptide samples described in the previous section 5fold to yield a 200 μM sample. Additional secondary structural
characterization was achieved with infrared spectroscopy (Thermo
Scientific, Nicolet 6700 FT-IR spectrophotometer). Ten microliters of
the samples described in the previous section were dropcast onto a 32
mm CaF2 plate (Sigma-Aldrich) and were air-dried. The thin films
were aligned in the spectrophotometer, and the amide I vibrations in
the region from 1500−1800 cm−1 were analyzed.
Heme Coordination. Electronic absorption spectroscopy (Cary
50 UV spectrometers) was employed to monitor the key π−π*
transitions typical of porphyrin derived molecules. The 1 mM peptide/
0.1 mM hemin samples described earlier were transferred to a quartz
cuvette with a 0.1 cm path length window (Starna Cells, Inc.) and
analyzed from 300−800 nm.
X-band continuous wave EPR experiments were carried out using a
Bruker ELEXSYS E580 spectrometer operating in the X-band (9.4
GHz) and equipped with an Oxford CF935 helium flow cryostat with
an ITC-5025 temperature controller. Samples for EPR (same
preparation as described in the Sample Preparation) were concentrated to 1 mM hemin and 10 mM peptide with a 10 000 molecular
weight cutoff spin diafiltration system (EMD Millipore Inc., Amicon
Ultra-0.5 Centrifugal Filter Unit with Ultracel-10 membrane). All
experiments were performed at 10 K with modulation amplitude set to
1 and power set to 10 mW.
Electrochemistry. The samples were placed in a spectroelectrochemical cell (1 mm quartz) equipped with a platinum mesh
working electrode, platinum wire auxiliary electrode, and a Ag/AgCl
reference electrode (Basi, Inc.). The samples were electrochemically
reduced over a range from +200 to −700 mV versus SHE. Each
applied voltage setting was allowed to equilibrate for a minimum of 20
min prior to UV/visible spectral acquisition (PerkinElmer, Lambda
950, UV/vis/NIR spectrophotometer). Midpoint potential analysis
was achieved by fitting a standard Boltzmann curve to the obtained
data (OriginPro 9.1).
Carbon Monoxide Binding. The heme ferrous state was obtained
through chemical reduction by adding 5 μL of a concentrated sodium
dithionite (Sigma-Aldrich) solution (100 mg/mL) into a preassembled
1000 μM peptide/100 μM hemin solution (300 μL) in an eppendorf
tube. All samples were equilibrated and handled in an inert, nitrogen
atmosphere (Plas Laboratories Inc. 830 Series Compact Glove Box).
Carbon monoxide (99.99%, Airgas) was added directly through the
solution in the eppendorf tube for 30 s. Ten microliters of the solution
was dropcast onto a CaF2 plate where CO(g) was gently blown over
the droplet resulting in a thin film of the PAHeme material. The samples
were than analyzed by FTIR spectroscopy (Thermo Scientific, Nicolet
6700 FT-IR spectrophotometer). The samples were stable against
oxidation during the course of the experiments. All obtained data were
fit to a double Gaussian peak distribution due to the pronounced
shoulders in some of the spectra.
Peroxidase Activity Assay. Peroxidase activity was monitored
using a Varian Cary 50 spectrophotometer and the kinetics software
package. All kinetics were performed in a solution of HEPES buffer,
pH 7 (50 mM HEPES 100 mM NaCl). The following stock solutions
were used: TMB (10 mg/mL, 41.6 mM in DMSO), hydrogen
peroxide (100 mM diluted in HEPES); H2O2 stock concentration was
standardized by the method of Klassen et al.93 and by UV/vis (ε230 =
72.8 M−1 cm−1). Stock solutions of 50 μM heme and 1500 μM PA in
HEPES (micelles) or 20 mM NH4OH (fibers) were equilibrated
overnight in the dark. In disposable plastic cuvettes, 2 mL of HEPES
buffer was added. TMB was added at a concentration of 300 μM.
H2O2 was varied from 1.0 to 10 mM. To initiate the reaction, 40 μL of
the heme stock solution was added and mixed via pipetting to yield 1
μM heme and 30 μM PA. The generation of the single electron
oxidation product of TMB was monitored (ε652 = 39 000 M−1 cm−1)
over a 5 min period collecting data points every 10 s. Initial velocities
(vo, μM s−1) were determined by fitting the linear region of the
kinetics (Figure S14). The initial velocity was then plotted versus
peroxide concentration (Figure S15). Michaelis−Menten curves were
fit using Origin 9.1 software to the equation (v0 = kCat[E]0[S]/(KM +
[S])). The total amount of oxidized TMB was calculated after allowing
the reaction to go to completion for 1 h. The reaction then was diluted
10-fold in 1 M HCl to yield the final, yellow TMB oxidation product.
The absorption was monitored at 450 nm to determine the total
amount of TMB oxidized, ε450 = 59 000 M−1 cm−1. Microperoxidase11 (Sigma-Aldrich) and free heme were prepared at 50 μM in HEPES.
SDS (10 wt % in HEPES, pH 7, diameter = 3 nm) and TWEEN 20 (1
wt % in HEPES, pH 7, diameter = 10 nm) micelles were characterized
by DLS (data not shown) to confirm correct micelle diameter. Heme
was added to a final concentration of 50 μM. The control solutions
were aged overnight in the dark to be consistent with the sample
preparation of our peptides. Because of its high degree of reactivity, 10
μL of the MP-11 stock (250 nM final concentration) was added to
initiate the reaction of the control experiment.
■
ASSOCIATED CONTENT
S Supporting Information
*
The Supporting Information is available free of charge on the
ACS Publications website at DOI: 10.1021/jacs.7b01588.
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AFM, DLS, SEM, FTIR, CD, EPR, binding constant
analysis, electrochemistry, CO binding, dilution stability,
Michaelis Menten plots, heme bleaching studies, and
mass spectroscopy analysis including Figures S1−S21
and eq S1 (PDF)
AUTHOR INFORMATION
Corresponding Author
*hfry@anl.gov
ORCID
Lee A. Solomon: 0000-0003-1471-9510
H. Christopher Fry: 0000-0001-8343-5189
Notes
The authors declare no competing financial interest.
■
ACKNOWLEDGMENTS
We wish to thank Tijana Rajh for helpful discussions related to
the manuscript in addition to her assistance with EPR. J.K.
participated in this work through the Illinois Mathematics and
Science Academy’s Student Inquiry and Research (SIR)
program supported in part through the State of Illinois
Education Assistance Fund. This work was performed at the
Center for Nanoscale Materials, a U.S. Department of Energy
Office of Science User Facility under contract no. DE-AC0206CH11357.
■
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