Identification of intermediates in peroxidase catalytic cycle of a DNAzyme possessing heme

Article abstract of DOI:10.1246/bcsj.20190157

Heme in the ferric state (heme(Fe³⁺)) binds to G-quadruplex DNAs to form stable complexes that exhibit enhanced peroxidase activities. The complexes are considered DNAzymes possessing heme as a prosthetic group (heme-DNAzymes), and have been extensively investigated as promising catalysts for a variety of applications. On ESR and stopped-flow measurements, an iron(IV)oxo porphyrin π-cation radical known as Compound I was detected in reaction mixtures of heme-DNAzymes and hydrogen peroxide. This finding not only resolved the long-standing issue of the mechanism underlying the enhancement of the peroxidase activity of heme(Fe³⁺) in the scaffold of a G-quadruplex DNA, but also provided new insights as to the design of novel heme-DNAzymes.

Full text of DOI:10.1246/bcsj.20190157

Advance Publication Cover Page
Identification of Intermediates in Peroxidase Catalytic Cycle of a DNAzyme Possessing Heme
Ryosuke Shinomiya, Haruka Araki, Atsuya Momotake, Hiroaki Kotani,
Takahiko Kojima, and Yasuhiko Yamamoto*
Advance Publication on the web July 13, 2019
doi:10.1246/bcsj.20190157
© 2019 The Chemical Society of Japan
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Publication manuscripts.

Identification of Intermediates in Peroxidase Catalytic Cycle of a DNAzyme
Possessing Heme
Ryosuke Shinomiya,1 Haruka Araki,1 Atsuya Momotake,1 Hiroaki Kotani,1 Takahiko Kojima,1 and Yasuhiko
Yamamoto*1,2,3
1Department of Chemistry, University of Tsukuba, Tsukuba 305-8571
2 Tsukuba Research Center for Energy Materials Science (TREMS), University of Tsukuba, Tsukuba 305-8571
3 Life Science Center for Survival Dynamics, Tsukuba Advanced Research Alliance (TARA), University of Tsukuba, Tsukuba 305-8577
E-mail:yamamoto@chem.tsukuba.ac.jp
Yasuhiko Yamamoto
Dr. Yasuhiko Yamamoto received his Ph. D. in Chemistry from University of California, Davis, in 1986.
He is currently Professor of Chemistry at University of Tsukuba.
His research interests are primarily in the arears of bioinorganic chemistry and NMR spectroscopy.
Abstract
Heme in the ferric state (heme(Fe3+)) binds to G-
quadruplex DNAs to form stable complexes that exhibit
enhanced peroxidase activities. The complexes are considered
as DNAzymes possessing heme as a prosthetic group (heme-
DNAzymes), and have been extensively investigated as
promising catalysts for a variety of applications. On ESR and
stopped-flow measurements, an iron(IV)oxo porphyrin -cation
radical known as Compound I was detected in reaction mixtures
of heme-DNAzymes and hydrogen peroxide. This finding not
only resolved the long-standing issue of the mechanism
underlying the enhancement of the peroxidase activity of
heme(Fe3+) in the scaffold of a G-quadruplex DNA, but also
provided new insights as to the design of novel heme-
DNAzymes.
Keywords: DNA enzyme, Heme, Peroxidase activity.
Figure 1. Structures and numbering schemes for heme(Fe2+)
1. Introduction
(A), Molecular structure of the G-quartet (B), and
schematic representation of complexes between
heme(Fe2+) and all parallel-stranded tetrameric G-
quadruplex DNAs of d(TTAGGG) and d(TTAGGGA), (C)
and (C’), respectively, and the heme(Fe2+) coordination
structure of a carbon monoxide (CO) adduct of the
complex (D). In the complexes, CO binds to the heme Fe
atom on the side of the heme opposite to the G6 G-quartet,
and then a water molecule (H2O), sandwiched between the
heme and G6 G-quartet planes, is coordinated to the Fe
atom as another axial ligand, trans to the CO ligand.
Heme, i.e., an iron-protoporphyrin IX complex (Figure 1A),
in the ferric state (heme(Fe3+)) binds to G-quadruplex DNAs to
form stable complexes that exhibit enhanced peroxidase
activities.1-5 These complexes are considered as DNAzymes
possessing heme (heme-DNAzyme) as a cofactor, and have been
extensively investigated as promising catalysts for a variety of
applications. Despite the exponential increase in the number of
papers on heme-DNAzyme applications published in recent
years,6-21 their structure-function relationship has largely
remained unknown.4,22-24 A G-quadruplex DNA is stabilized
by a unique structural motif, known as the G-quartet, which is
formed through a cyclic and coplanar association of four guanine
bases through Hoogsteen hydrogen bonds (Figure 1B).25 We
demonstrated that heme(Fe3+) (or heme(Fe2+)) binds selectively
to the 3’-terminal G-quartet (G6 G-quartet) of an all parallel-
stranded tetrameric G-quadruplex DNA formed from a single
repeat sequence of the human telomere, (d(TTAGGG))4 (6mer),
to form a complex (heme(Fe3+)-6mer or heme(Fe2+)-6mer
complex) (Figure 1C).26-32 The heme(Fe3+)-6mer complex has
been reported to exhibit peroxidase activity32 as well as various
spectroscopic properties remarkably similar to those of the
oxygen storage hemoprotein myoglobin26-28, 30, 31 (Mb). Hence
the heme(Fe3+)-6mer complex could be considered as an
excellent model for elucidating the structure-function
relationships in heme-DNAzymes.
It is often assumed that the peroxidase activity of a heme-
DNAzyme is elicited through a reaction mechanism similar to
that of peroxidases such as horseradish peroxidase (HRP)
(Scheme 1A). The assumption is not indisputable, because

mixtures of the complexes and H2O2, which is characteristic of
47
Compound I.46,
The UV-vis absorption signatures of
Compound I48, 49 could also be detected in the spectrum of a
mixture of heme(Fe3+)-6mer/A complex and H2O2, recorded
through stopped-flow experiments.
These results clearly
demonstrated generation of Compound I in the catalytic cycle of
the heme-DNAzyme. This finding provides the fundamental
basis of the structure-function relationship in heme-DNAzymes.
2. Experimental
Heme-DNA Complex Preparation.
DNA sequences,
d(TTAGGG) and d(TTAGGGA), purified with a C-18 Sep-Pak
cartridge, were purchased from Tsukuba Oligo Service Co.
The oligonucleotides were obtained by ethanol precipitation and
then desalted with Microcon YM-3 (Millipore, Bedford, MA).
The concentration of each oligonucleotide was determined
spectrophotometrically using the absorbance at 260 nm (molar
extinction coefficients λ260 = 6.89 × 104 and 8.43 × 104 cm-1M-1
Scheme 1. Catalytic cycle of horseradish peroxidase
(HRP)43 (A) and schematic representation of the active site of
HRP44 (B). In (B), amino acid residues involved in the “push-
pull mechanism”33, 42 are shown.
for
d(TTAGGG)
and
d(TTAGGGA),
respectively).
functional groups involved in the catalytic cycle of a heme-
DNAzyme ought to be different from those of HRP. The
catalytic cycle of HRP as well as of cytochrome P450 proceeds
through the iron(IV)oxo porphyrin π-cation radical intermediate
known as Compound I formed through heterolytic O-O bond
cleavage of Fe-bound hydrogen peroxide (H2O2), and this
process is controlled through the so-called “push-pull”
mechanism (Scheme 1B).33-44 The “push” effect in HRP is
provided by the electron-donating effect of the proximal
histidine (His170), which possibly contributes to stabilization of
higher oxidation states of the heme Fe atom during the
catalysis.33-37 On the other hand, during the catalytic cycle of
HRP, the distal His (His42) accepts a proton from the proximal
oxygen atom of Fe-bound H2O2 and transfers it to the distal
oxygen one to generate leaving H2O. Furthermore, in concert
with the action of His42, a nearby cationic amino acid side chain
(Arg38) facilitates heterolytic O-O bond cleavage through
stabilization of the developing negative charge on the distal
oxygen atom during the bond cleavage (Scheme 1B). Thus, the
“pull” mechanism is provided by the combination of His42 and
Arg38. 38-42
Heme(Fe3+) was purchased from Sigma-Aldrich Co.
Preparation of the heme(Fe3+)-DNA complexes was carried out
by mixing 5 - 20 µM heme(Fe3+) and 10 - 40 µM DNA as
described previously.31
Measurement of UV-Vis Absorption and Amplex Red
Oxidation Kinetics.
recorded at 298 K with a Beckman DU 640 spectrophotometer
using 50 mM potassium phosphate buffer, pH 6.80, containing
300 mM KCl as the solvent. Horseradish peroxidase (HRP)
was purchased as a lyophilized powder from Sigma-Aldrich Co.
UV-Vis absorption spectra were
and used without further purification.
The chromogenic
substrate, Amplex Red (10-acethyl-3,7-dihydroxyphenoxazine),
was purchased from Sigma-Aldrich, and the reaction was
monitored by following the appearance of the oxidized product,
7-hydroxyphenoxazin-3-one (Resorufin) through the following
reaction, which absorbs light at ~570 nm.32
We found, through characterization of the peroxidase
activities of complexes possessing a series of chemically-
modified hemes, that the complex monotonously increases with
increasing electron density of the heme Fe atom, supporting the
“push” mechanism in the catalytic cycle of the complex.32
Furthermore, the “pull” mechanism in the catalytic cycle of the
complex was also confirmed by comparative studies on
complexes of heme(Fe3+) with all parallel-stranded monomeric
G-quadruplex DNAs formed from inosine(I)-containing
sequences, i.e., d(TAGGGTGGGTTGGGTGIG) DNA(18mer)
and d(TAGGGTGGGTTGGGTGIGA) DNA(18mer/A).45 We
found that heme(Fe3+) binds to the 3’-terminal G-quartet of the
G-quadruplex DNAs to form complexes exhibiting peroxidase
activities, and that the activity of the heme(Fe3+)-
DNA(18mer/A) complex was greater than that of the
heme(Fe3+)-DNA(18mer) one, indicating that the 3’-terminal A,
i.e., A19, of the sequence acts as a general acid-base catalyst that
promotes the catalytic reaction of the heme(Fe3+)-
DNA(18mer/A) complex.
Kinetic studies were performed on a Beckman DU640
spectrometer. (d(TTAGGG))4 and (d(TTAGGGA))4 in 50 mM
potassium phosphate buffer, pH 6.80, were heat-denatured at
363 K for 5 min., followed by cooling to 298 K. Amplex Red
was dissolved in dimethylformamide to prepare a 10 mM stock
solution. Heme(Fe3+) and then Amplex Red were added to 0.1
µM and 50 µM, respectively, to 10 µM DNA in 50 mM
potassium phosphate buffer, pH 6.80. To initiate the oxidation
reactions, 5 – 200 mM hydrogen peroxide (H2O2) were added to
the solution mixture. For the kinetic measurements of the HRP
system, 1 – 100 M H2O2 was added to the solution mixture of
0.01 µM protein and 50 µM Amplex Red in 50 mM potassium
phosphate buffer, pH 6.80, to initiate the reactions. The initial
slope (R0) of the time evolution of 570-nm absorbance due to
Resorufin was used as an index for the peroxidase activities of
the complexes and HRP.
In this study, we attempted to capture short-lived
intermediates in the peroxidase catalytic cycle of the
heme(Fe3+)-6mer complex and a related complex composed of
heme(Fe3+) and all parallel-stranded tetrameric G-quadruplexes,
(d(TTAGGGA))4, (6mer/A), and (heme(Fe3+)-6mer/A complex)
through ESR spectroscopy and stopped-flow experiments. We
could detect a convex peak at g = ~2 in the ESR spectra of frozen
Electron Spin Resonance (ESR) Spectroscopy. ESR
spectra were recorded on a Bruker BioSpin X-band spectrometer
(EMXPlus9.5/2.7) with a liquid helium transfer system under
non-saturating microwave power conditions (4.0 mW) operating
at 9.394 GHz. The magnitude of the modulation was chosen to
optimize the resolution and the signal-to-noise (S/N) ratio of the

Figure 2. The time evolution of the oxidation reaction of Amplex Red (10-acethyl-3,7-dihydroxyphenoxazine) with the
heme(Fe3+)-6mer/A complex and H2O2, monitored by following the appearance of the oxidized product, 7-hydroxyphenoxazin-
3-one (Resorufin) (A). Time-evolution of 570-nm absorbance due to Resorufin produced on the reactions of the heme(Fe3+)-
6mer and heme(Fe3+)-6mer/A complexes is shown in the inset. Samples comprised 1.0 μM heme(Fe3+), 10 μM DNA, 50 μM
Amplex Red, and 5 mM H2O2 in 50 mM potassium phosphate buffer, pH 6.80, at 298 K. The values of 0.05 ± 0.01 and 0.37 ±
0.04 μM/s were obtained from the initial slopes (R0) for the heme(Fe3+)-6mer and heme(Fe3+)-6mer/A complexes, respectively.
Plots of the R0s for the heme(Fe3+)-6mer and heme(Fe3+)-6mer/A complexes (B), and HRP (C), as a reference, against the
concentration of H2O2 ([H2O2]). The HRP sample comprised 0.05 μM protein, 50 μM Amplex Red, and 1-100 μM H2O2 in 50
mM potassium phosphate buffer, pH 6.80, at 298 K.
observed spectrum (modulation amplitude, 5−20 G; modulation
frequency, 100 kHz).
relative to the corresponding parameters of the heme(Fe3+)-
6mer/A complex, respectively. As compared with the values
obtained for HRP, the Km(H2O2)s and kcats of the complexes were
significantly larger and smaller, respectively (Table 1).
For ESR samples of heme-DNA complexes, 0.1 mM heme
and 0.1 mM DNA were mixed in 50 mM potassium phosphate
buffer, pH 6.80, and 300 mM KCl, and then 15 mM H2O2 was
added. The mixed solution was immediately frozen for the
measurement at 5 K under a He atmosphere.
Stopped-Flow Experiments. Formation of Compound I
was monitored by stopped-flow experiments performed on a
UNISOKU USP-SFM-CRD10 stopped-flow spectrometer
ESR Spectra. ESR spectra of the heme(Fe3+)-6mer and
heme(Fe3+)-6mer/A complexes were measured in 50 mM
potassium phosphate buffer (pH 6.80) at 5 K (Figure 3A and B)
In the spectra, axial signals at g = 5.95 and g║ = 2.00 were
observed, and these signals were markedly similar to those
reported for the acidic met-forms of native Mb and its variants.52
In addition, a rhombic high-spin type signal was also observed
at g = 6.33 in the spectrum of the heme(Fe3+)-6mer complex
(Figure 3A), whereas such a signal was not observed in the
spectrum of the heme(Fe3+)-6mer/A complex (Figure 3B).
Furthermore, unidentified broad peaks were observed near g = 2
in the spectra of the heme(Fe3+)-6mer and heme(Fe3+)-6mer/A
complexes (Figure 3A and B).
equipped with a multichannel photodiode array, a 150-W xenon
51
lamp, and a double-mixing apparatus at 277 K.50,
For
measurements of the heme-DNA complex systems, 5 μM heme
and 10 μM DNA in 50 mM potassium phosphate buffer (pH
6.80) and 300 mM KCl were mixed with 15 mM H2O2. On the
other hand, 5 μM HRP was mixed with 1 mM H2O2 in 50 mM
potassium phosphate buffer, pH 6.80.
The spectra of samples prepared by freesing mixtures of the
complex ([heme(Fe3+)-6mer complex] or [heme(Fe3+)-6mer/A
complex] = 0.1 mM) and [H2O2] = 15 mM in phosphate buffer
solutions, pH 6.80 (the interval between mixing and freezing
being ~3 s), exhibited convex peaks at g ~ 2 (Figures 3A’ and
B’), which were similar to the spectral characteristics of
Compound I46,47, 53-57, i.e., an iron(IV)oxo porphyrin π-cation
radical complex, as observed for the HRP sample similarly
prepared from a solution mixture of [HRP] = 0.1 mM and [H2O2]
= 0.2 mM (see Figure S1 in the Supporting Information). In
addition, a signal at g ~ 6, derived from an unreacted ferric high-
spin species, and one at g ~ 4.3 due to degraded heme57 were
observed, as seen in Figure 3A’ and B’, and the rhombic high-
spin type signal at g = 6.33 was observed as a shoulder of a peak
at g = 5.94 in the spectrum of the heme(Fe3+)-6mer complex
(Figure 3A’). On the other hand, the convex peak at g ~ 2 due
3. Results
Kinetics Measurements. The peroxidase activities of the
heme(Fe3+)-6mer and heme(Fe3+)-6mer/A complexes were
evaluated using Amplex Red as a substrate, and the initial slope
(R0) of the time evolution of 570-nm absorbance due to the
oxidized product, 7-hydroxyphenoxazin-3-one (Resorufin), was
used as an index for the activity, as reported previously32, 45
(Figure 2A). The peroxidase activity of HRP, as a reference,
was similarly measured and evaluated.
The R0s of the
complexes and HRP in the presence of various H2O2
concentrations ([H2O2]) at 298 K were determined using the
Michaelis-Menten equation to yield Michaelis constants for
H2O2 (Km(H2O2)) and catalytic rate constants (kcat) (Figure 2B
and C, and Table 1). The Km(H2O2) and kcat of the heme(Fe3+)-
6mer complex were larger and smaller by factors of ~3 and ~1/3
Table 1. Steady state kinetics parameters for H2O2 oxidation of Amplex Red catalyzed by the heme(Fe3+)-6mer and heme(Fe3+)-
6mer/A complexes, and HRP in 50 mM potassium phosphate buffer, pH 6.80, 300 mM KCl, at 298 K.
Kma (mM)
163 ± 2
60 ± 7
0.04 ± 0.01
Kcata (s−1)
18 ± 1
51 ± 2
1100 ± 40
Kcat/Km (mM−1 s −1
)
Heme(Fe3+)-6mer complex
Heme(Fe3+)-6mer/A complex
HRP
0.11
0.85
28000
a Obtained by fitting of the plots shown in Figure 2B and C to the Michaelis-Menten equation.

In contrast, the addition of a large excess of H2O2 to the
heme(Fe3+)-6mer/A complex resulted in an apparent biphasic
change in the Soret band, such that the Soret band initially
exhibited a rapid decrease and then a gradual one (Figure 4C).
As shown in Figure 4D, the spectrum of the heme(Fe3+)-6mer/A
complex was markedly altered by the addition of H2O2, and a
drastic decrease in the Soret band and an increase at ~350 nm in
the spectrum recorded after the H2O2 addition are similar to the
UV-Vis spectral signatures of Compound I48, 49 (see also Figure
S2C and D in the Supporting Information).
4. Discussion
Peroxidase Activity. Analysis of the [H2O2]-dependent R0
change observed for the heme(Fe3+)-6mer and heme(Fe3+)-
6mer/A complexes using the Michaelis-Menten equation yielded
Km(H2O2)s and kcats of the complexes that were significantly
larger and smaller than the corresponding ones of HRP,
respectively (Table 1).
The difference in the Km(H2O2)
between the complex and HRP could be to some extent due to
their distinct heme coordination structures. We found, through
NMR characterization of a carbon monoxide (CO) adduct of the
heme(Fe2+)-6mer complex, that a water molecule sandwiched
between the heme and G6 G-quartet planes is coordinated to the
Fe atom as the other axial ligand (axial H2Oint), trans to axial CO,
and is hydrogen-bonded to the carbonyl oxygen atoms of G6
bases (Figure 1D).31 Upon the oxidation of the heme Fe center
in the CO adduct of the heme(Fe2+)-6mer complex to heme(Fe3+),
the Fe-bound CO in the complex is thought to be replaced by
H2O (axial H2Oext), while axial H2Oint is retained, and, as a result,
a bis-H2O-coordinated heme(Fe3+), i.e., heme(Fe3+) possessing
H2Oint and H2Oext as axial ligands, in the heme(Fe3+)-DNA
complex is formed under low pH conditions.32 Therefore
binding of H2O2 to the heme Fe3+ center of the complex proceeds
through a ligand replacement reaction, as opposed to the binding
of H2O2 to a penta-coordinated heme(Fe3+) of resting state
HRP44, 58, 59 through a simple second-order reaction. Hence the
H2O2 binding affinity of the heme(Fe3+)-6mer complex is
intrinsically lower compared with that of resting state HRP, as
reflected on the finding that the Km(H2O2) of the complex is
considerably larger than that of HRP. The peroxidase activities
of the complexes could be interpreted on the basis of the push-
pull mechanism,32, 45 as in the case of the catalytic cycle of HRP
(Scheme 1A). We found that the donor strength of the axial
H2Oint ligand in the heme(Fe3+)-6mer complex is markedly
weaker than that of the proximal His in Mb.32 Furthermore, the
imidazolate character of the axial His170 in HRP was shown to
be enhanced through the formation of a strong hydrogen bond
with Asp247 (Scheme 1B), and hence the donor strength of the
His170 in HRP is even stronger than that of the proximal His in
Mb.43, 58, 60 Therefore, the push mechanism due to the axial
His170, together with the pull one due to both His42 and Arg38,
in HRP contributes significantly to the increase in the activity
(Scheme 1B). Consequently, the low peroxidase activity of the
heme(Fe3+)-6mer complex is due to the weak donating ability of
the axial H2Oint, in addition to the absence of functional groups
capable of furnishing the pull mechanism in the complex. Hence,
the ~3-fold increase in the kcat on the addition of an extra A at
the 3’-terminal of the sequence indicated that the 3’-terminal A
of the heme(Fe3+)-6mer/A complex acts as a general acid-base
catalyst that promotes Compound I formation through its pull
mechanism (Scheme 2).
Figure 3. ESR spectra of the heme(Fe3+)-6mer complex (A)
and heme(Fe3+)-6mer/A one (B) at 5K. Spectra recorded for
samples prepared in 50 mM potassium phosphate buffer, pH
6.80, by addition of a 150-fold stoichiometric excess of H2O2
to the heme(Fe3+)-6mer complex in the absence (A’) and
presence of the substrate, Amplex Red (A’’), and that
recorded for the heme(Fe3+)-6mer/A complex after addition
of
a 150-fold stoichiometric excess of H2O2 to the
heme(Fe3+)-6mer/A complex (B’). In (A’) and (B’), the Y-
gains of signals near g = 2 are expanded by a factor of ~2.5
and convex peaks at g = ~2 due to Compound I are indicated
by downward pointing arrows.
to Compound I was not detected even though H2O2 was added
to the heme(Fe3+)-6mer complex in the presence of the substrate,
Amplex Red, and only signals due to a ferric high-spin species
and degraded heme were observed (Figure 3A’’).
Stopped-flow Experiments. A stopped-flow apparatus50,51
was used to detect short-lived intermediates in the reaction of the
heme(Fe3+)-6mer or heme(Fe3+)-6mer/A complex with H2O2 at
pH 6.8 and 277 K. The absorption maximum of the Soret band
of the heme(Fe3+)-6mer/A complex, i.e., 406 nm, was red-shifted
by 3 nm relative to that of the heme(Fe3+)-6mer one, i.e., 403 nm
(Figure 4A), indicating that the heme environment in the
heme(Fe3+)-6mer complex is affected by the addition of the extra
A at the 3’-terminal of the DNA, i.e., A7. Upon addition of a
large excess of H2O2 to the aqueous solution containing the
heme(Fe3+)-6mer complex, a gradual decrease in the Soret band
was observed (Figure 4B). Since photodegradation of the
complex during stopped-flow measurements is essentially
negligible (Figure S2A and B in the Supporting Information), the
decrease in the Soret band could be due to the formation of
Compound I, although UV-Vis spectral signatures of Compound
I48, 49 were not apparent (see below).
ESR Detection of Compound I. In the ESR spectrum of the
heme(Fe3+)-6mer complex at 5 K, an axial high-spin signal with
g = 5.95 and g║ = 2.00, and a rhombic high-spin-type signal

0.8
0.6
Heme(Fe3+)-6mer complex
1.0
0.5
0
A
C
B
D
0.5 ms
8 s
Heme(Fe3+)-6mer/A complex
Heme(Fe3+)-6mer complex
0.4
0.2
0
300
400
500
600
700
300
400
500
600
700
Wavelength (nm)
Wavelength (nm)
0.8
0.6
0.8
0.6
Heme(Fe3+)-6mer/A complex
Heme(Fe3+)-6mer/A complex
0.5 ms
2 s
0.5 ms
2 s
0.4
0.2
0
0.4
0.2
0
ൈ 20
300
400
500
Wavelength (nm)
600
700
300
400
500
Wavelength (nm)
600
700
Figure 4. UV-Vis absorption spectra of the heme(Fe3+)-6mer (black) and heme(Fe3+)-6mer/A complexes (red) in 300 mM KCl,
50 mM potassium phosphate buffer, pH 6.80, at 298 K (A), and absorption spectral changes in the course of the reactions, for
time spans of ~0.5 ms to 8 s, of the heme(Fe3+)-6mer complex with a 1500-fold stoichiometric excess of H2O2 in 50 mM
potassium phosphate buffer, pH 6.80, and 300 mM KCl at 277 K, monitored with a time interval of 40 ms with a stopped-flow
spectrometer (B). In (B), the spectra recorded with an interval of 800 ms are shown. Absorption spectral changes in the
course of the reactions, for time spans of ~0.5 ms to 2 s, of the heme(Fe3+)-6mer/A complex with a 1500-fold stoichiometric
excess of H2O2 in 50 mM potassium phosphate buffer, pH 6.80, and 300 mM KCl at 277 K, monitored with a time interval of
40 ms with a stopped-flow spectrometer (C). In (C), the spectra recorded with an interval of 200 ms are shown, and the Y-
gains of the regions 500 – 650 nm are expanded by a factor of ~20.
at ~0.5 ms (blue) and 2 s (red) after the addition of H2O2, extracted from (C) are compared (D).
The spectra of the heme(Fe3+)-6mer/A complex, recorded
with g = 6.33 were observed (Figure 3A). The spectrum of the
complex at 298 K previously reported26 was axial, and hence a
species with a rhombic symmetry of the d electron system was
not detected. Therefore, the axial high-spin signals observed in
the spectra at 5 K and 298 K could be due to bis-H2O-
coordinated heme(Fe3+) in the complex, with axial symmetry of
the d electron system. In addition, the rhombic high-spin
species detected in the spectrum of the complex at 5 K might be
due to the presence of two (or more) orientations of the axial
between the porphyrin -cation radical (S = 1/2) and the iron
center (S = 1).61, 62 The newly-emerged convex peaks at g ~ 2
in the spectra of the frozen samples of mixtures of the complexes
and H2O2 exhibited such ESR characteristics of Compound I
(Figure 3 A’ and B’), supporting Compound I formation in the
heme(Fe3+)-6mer and heme(Fe3+)-6mer/A complex systems.
Furthermore, Compound I was not formed even though H2O2
was added to the heme(Fe3+)-6mer complex in the presence of
Amplex Red (Figure 3A’’), demonstrating that Compound I was
rapidly consumed through oxidation of the substrate. Thus, on
the basis of these ESR spectra, the formation of Compound I as
an intermediate in the peroxidase catalytic cycles of the
complexes was confirmed.
H2Oext
,
with respect to the heme(Fe3+), which possibly
influences the axial symmetry of the high-spin ferric center. In
contrast to the case of the heme(Fe3+)-6mer complex, the
spectrum of the heme(Fe3+)-6mer/A complex at 5 K did not
exhibit
a
rhombic high-spin-type signal, and hence the
Stopped-flow Detection of Compound I.
Since
orientation of the axial H2Oext, with respect to the heme(Fe3+),
might be fixed through its interaction with the 3’-terminal A, i.e.,
A7, in the complex. Furthermore, the broad peaks near g = 2
in the ESR spectra of the complexes (Figures 3A and B) are
likely to be due to the presence of unidentified minor species in
the samples, and similar broad ones near g = 2 could also be
detected in the spectrum of a complex between heme and a G-
quadruplex heme-binding DNA aptamer.3
Compound I exhibits a characteristic absorption spectrum,48, 49
the stopped-flow technique has frequently been used to detect
this short-lived intermediate in the peroxidase catalytic cycle.
The absorption spectral signatures of Compound I were clearly
observed in the spectrum of the heme(Fe3+)-6mer/A complex
(Figure 4C), although it was not apparent in the spectra of the
heme(Fe3+)-6mer one (Figure 4B).
In the spectrum of
Compound I, an increase at ~350 nm, and small absorption
maxima at ~580 and ~650 nm are observed, and these features
have been recognized as its UV-Vis spectral signatures48, 49
Besides these spectral signatures of Compound I, a decrease in
the Soret band is likely to be most prominent among the spectral
An asymmetric signal, with a line width of ~15 G, at g = 1.995
has been reported to be an ESR spectral signature of Compound
I.46 The broadening of the Compound I signal is due to short
electron spin relaxation times through spin-exchange interaction
.

Scheme 2. “Pull mechanism” possibly provided by A7 base of the heme(Fe3+)-6mer/A complex, which facilitates heterolytic
O-O bond cleavage of Fe-bound hydrogen peroxide (H2O2). A similar mechanism has been proposed previously.22-24 The
axial H2O molecule is sandwiched between the heme and G-quartet planes31, 32 and the hydrogen bond between Fe-bound
H2O and the nearby A7 base45 is added to the proposed mechanism as a dotted line.
changes observed upon the conversion of a ferric high-spin heme
complex to Compound I (see Figure S2D in the Supporting
Information). Hence, the rapid decrease in the Soret band
observed in Figure 4B might be considered as an indicator of the
formation of Compound I, although bleaching of the complex by
H2O2 cannot be ruled out. The observation of an ESR spectral
signature of Compound I, i.e., a convex peak at g ~ 2, in the
spectrum of a sample prepared by freezing a mixture of the
complex and H2O2 within ~3 s (Figure 3A’) supported this
interpretation. As far as the sensitivities of the techniques for
detecting Compound I are concerned, the ESR technique, which
relies on the observation of its characteristic signal at g ~ 2,
would be advantageous over the UV-Vis one, which is possibly
hampered by spectral overlap between a ferric high-spin heme
complex and Compound I, particularly in cases where only a
trace amount of Compound I is formed in samples, such as the
case shown in Figure 4B.
Although the molecular mechanism responsible for the
enhancement of the intrinsic peroxidase activity of heme(Fe3+)
through interaction with DNA has yet to be fully elucidated, it
has been shown that binding to heme(Fe3+) to a folded DNA is
not sufficient to activate heme(Fe3+).2 We found that, in the
complex, a water molecule (H2O), sandwiched between the
heme and G-quartet planes, is coordinated to the Fe center as an
32
axial ligand.31,
Theoretical study indicated that the
polarization of the H2O molecule is considerably greater than
that of an ordinary one,31 and hence the axial H2O in the complex
is possibly capable of being an electron-donating ligand which
increases the Lewis basicity of the heme Fe atom to activate Fe-
bound H2O2 for the reaction.
The peroxidase activity of heme-DNAzymes has been
assumed to be elicited through a reaction mechanism essentially
identical to the HRP peroxidation cycle.12, 24 The proposed
24
mechanism for heme-DNAzymes12,
has been supported
In contrast, as manifested in the enhanced peroxidase activity
of the heme(Fe3+)-6mer/A complex relative to that of the
heme(Fe3+)-6mer one (Table 1), the “pull” mechanism provided
by the 3’-terminal A accelerated the Compound I formation so
that an appreciable amount of Compound I was rapidly formed
in the heme(Fe3+)-6mer/A complex system upon the addition of
H2O2 (Figure 4C). As described above, since the conversion of
a ferric high-spin heme complex to Compound I results in a
considerable decrease in the Soret band (Figure S2D in the
Supporting Information), the time scale of the decrease in the
Soret band of the system is thought to be accelerated with
increasing rate of Compound I formation. Consequently, the
rapid decrease in the Soret band of the heme(Fe3+)-6mer/A
complex system, relative to that of the heme(Fe3+)-6mer
complex counterpart, also supported enhancement of the rate of
Compound I formation through the “pull” mechanism provided
by the 3’-terminal A, which ensured the formation of Compound
I enough to observe its characteristic absorption spectrum
(Figure 3B).
Peroxidase Catalytic Cycle of Heme-DNAzymes. The
peroxidase activity of heme(Fe3+) is dramatically enhanced
when it is bound to the G-quartet. 1-8, 32, 45 The enhancement of
the catalytic activity of heme(Fe3+) through its interaction with
nucleic acids is likely to be relevant to primordial chemistry as
well as to current biology. In addition, elucidation of the
structure-function relationship of heme-DNAzymes not only
leads to the discovery of a novel mechanism for control of the
heme(Fe3+) reactivity in the scaffold of nucleic acids, but also
paves a way to the creation of novel heme-DNAzymes.
indirectly by the findings that demonstrated that the “push-pull”
mechanism is operative in the heme-DNAzyme peroxidation
32, 45
cycle.23,
In this study, using ESR spectroscopy and
stopped-flow measurements, we could detect Compound I
formed in the heme(Fe3+)-6mer and heme(Fe3+)-6mer/A
complex systems, providing direct evidence of the catalytic
mechanism of the complexes. Hence the mechanism for the
heme-DNAzyme peroxidation cycle was proved to be identical
to that for the HRP one. This finding will allow us to design
and enhance the catalytic activity of heme-DNAzymes with
reference to the HRP peroxidation mechanism.
5. Concluding Remarks
We could detect a high valent iron(IV)oxo porphyrin π-
cation radical species known as Compound I formed in the
peroxidase catalytic cycle of heme-DNAzymes composed of
heme, heme(Fe3+), and all parallel-stranded tetrameric G-
quadruplex DNAs related to a single repeat sequence of the
human telomere, i.e., d(TTAGGG) and d(TTAGGGA). This
finding confirmed that the peroxidase activity of heme-
DNAzymes is realized through Compound I, as in the case of
heme enzymes such as HRP. Since functional groups involved
in the peroxidation cycle of the heme-DNAzymes are different
from those of HRP, elucidation of the structure-function
relationship in the heme-DNAzymes will lead to discovery of a
novel mechanism for control of the heme(Fe3+) reactivity, which
will contribute to exploration of the biological versatility of
heme as
a prosthetic group of nucleic acid catalysts.
Furthermore, since heme is believed to be an ancient
compound,63 the results presented here provide a new insight

64
into the concept of
a
primordial “RNA world”
by
Suzuki, Y. Yamamoto, J. Porphyr. Phthalocyanines 2014,
18, 741.
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Di, T. Shibata, H. Tai, A. Suzuki, S. Neya, Biochemistry
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demonstrating the redox-catalyzing ability of heme in the
scaffold of nucleic acids, which will expand the repertories of
nucleic acid catalysts.
Acknowledgement
32. R. Shinomiya, Y. Katahira, H. Araki, T. Shibata, A.
Momotake, S. Yanagisawa, T. Ogura, A. Suzuki, S. Neya,
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This work was financially supported by JSPS KAKENHI (No.
16KT0048 to Y.Y. and 17H03027 to T.K.), and the Bilateral
Open Partnership Joint Research Project (No. BBD29011 to
Y.Y.).
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