Quantification and evaluation of kinetic bio-catalytic pathway of horseradish peroxidase in an electron mediated reaction system and its applications in plant extracts

Article abstract of DOI:10.1016/j.saa.2012.10.024

The intermolecular coupling of 2,5-dimethoxyaniline (DMA) as mediated electron transfer reaction in presence of H₂O₂ and peroxidase in acetate buffer of pH 4.2 resulting green colored product having maximum absorption at λ_max = 740 nm was investigated by spectrophotometer. Under optimum conditions, linearity range for the quantification of H₂O₂ was 2.0-288.0 μM and for peroxidase were 0.59-9.46 and 0.443-9.46 nM by kinetic and fixed-time method, respectively. The catalytic efficiency and catalytic power were KeffD = 2.354 × 10⁵ M^-1 min^-1 and KpowD = 4.59 × 10^-4 min^-1, respectively. From the plot of d(1/D _o) vs d(1/V_o) and d(1/H_o) vs d(1/V _o), Michaelis-Menten constants for DMA and H₂O ₂were found that KmD = 1458 μM and KmH₂O₂ = 301 μM. Applicability of the method was tested for peroxidase activity in some plant extracts and compared with guaiacol/peroxidase system. Regarding superiority of the method, it is suggested that DMA/peroxidase system can be a better hydrogen donor for HRP assay than guaiacol system as evident from kinetic data.

Full text of DOI:10.1016/j.saa.2012.10.024

Spectrochimica Acta Part A: Molecular and Biomolecular Spectroscopy 102 (2013) 75–81
Contents lists available at SciVerse ScienceDirect
Spectrochimica Acta Part A: Molecular and
Biomolecular Spectroscopy
journal homepage: www.elsevier.com/locate/saa
Quantification and evaluation of kinetic bio-catalytic pathway of horseradish
peroxidase in an electron mediated reaction system and its applications in
plant extracts
⇑
Honnur Krishna, Padmarajaiah Nagaraja , Anantharaman Shivakumar, Nelligere A. Chamaraja,
Narayan Aradhana
Department of Studies in Chemistry, University of Mysore, Manasagangotri, Mysore 570 006, Karnataka, India
g r a p h i c a l a b s t r a c t
h i g h l i g h t s
"
"
"
"
2,5-Dimethoxyaniline is used as a
chromogenic co-substrate for
peroxidase assay.
A mathematical model has been
proposed for the evaluation of
catalytic parameters.
Peroxidase activity in some of crude
medicinal plant species has been
assessed.
Peroxidase activity has been
reported first time in T. fruticosum
and S. paniculatum.
a r t i c l e i n f o
a b s t r a c t
Article history:
Received 9 August 2012
Received in revised form 28 September 2012
Accepted 12 October 2012
Available online 24 October 2012
The intermolecular coupling of 2,5-dimethoxyaniline (DMA) as mediated electron transfer reaction in
presence of H2O2 and peroxidase in acetate buffer of pH 4.2 resulting green colored product having max-
imum absorption at kmax = 740 nm was investigated by spectrophotometer. Under optimum conditions,
linearity range for the quantification of H2O2 was 2.0–288.0 lM and for peroxidase were 0.59–9.46 and
0.443–9.46 nM by kinetic and fixed-time method, respectively. The catalytic efficiency and catalytic
power were KeDff = 2.354 ꢀ 105 Mꢁ1 minꢁ1 and KpDow = 4.59 ꢀ 10ꢁ4 minꢁ1, respectively. From the plot of
Keywords:
d(1/Do) vs d(1/Vo) and d(1/Ho) vs d(1/Vo), Michaelis–Menten constants for DMA and H2O2were found that
2,5-Dimethoxyaniline
Horseradish peroxidase assay
Mediated electron transfer system
Bio-catalyst
Mathematical model
Crude plant extracts
2O2
KDm = 1458 M and KHm = 301
l lM. Applicability of the method was tested for peroxidase activity in some
plant extracts and compared with guaiacol/peroxidase system. Regarding superiority of the method, it is
suggested that DMA/peroxidase system can be a better hydrogen donor for HRP assay than guaiacol sys-
tem as evident from kinetic data.
Ó 2012 Elsevier B.V. All rights reserved.
Introduction
Abbreviations: HRP, horseradish peroxidase; DMA, 2,5-dimethoxy aniline; 2,4-
2 O2
DMA, 2,4-dimethoxyaniline; KHm , Michaelis–Menten constant for H2O2; Vo, initial
Peroxidases (POD; EC 1.11.1.7) have dominated a prominent
position in biotechnology and associated research areas including
enzymology, biochemistry, medicine, genetics, physiology, histo-
and cytochemistry. Peroxidases are a class of enzymes extensively
distributed in higher plants, animals and microorganisms and are
associated with such catalytic activity as peroxidic, oxidative, and
rate of a reaction; [D]o, initial concentrations of DMA; [H2O2]o, initial concentration
2 O2
of H2O2; VHmax , maximum velocity for H2O2; KeDff and KGeff , catalytic efficiencies of
DMA method and guaiacol respectively; KDpow and KpGow, catalytic power of DMA
method and guaiacol.
⇑
Corresponding author. Tel.: +91 821 2412653; fax: +91 821 2421263.
E-mail address: profpn58@yahoo.com (P. Nagaraja).
1386-1425/$ - see front matter Ó 2012 Elsevier B.V. All rights reserved.
http://dx.doi.org/10.1016/j.saa.2012.10.024

76
H. Krishna et al. / Spectrochimica Acta Part A: Molecular and Biomolecular Spectroscopy 102 (2013) 75–81
hydroxylation [1]. Peroxidases are involved in a wide range of
physiological processes related to plant growth and development
[2]. Peroxidases are very commonly used in the construction of
biosensors and other analytical applications such as ELISA (HRP
activity in some of the crude plant tissues had a strong absorption
of violet-colored species at kmax = 540 nm. Even though the result-
ing intermolecular coupling of both the derivatives of dimethoxy
aniline (2,4-DMA and 2,5-DMA) are same radical species but may
be due to the stabilization of the meta position of methoxy group
in the excited state of 2,5-DMA derivative that makes it to absorb
at higher wavelength region might have resulted in different col-
ored species [26]. The main advantage of the proposed method
over the reference method is that the solubility of 2,4-DMA is in
alcohol and 2,5-DMA is in dil. HCl (0.112 N) wherein alcohol med-
ium inhibition of peroxidase activity take place to a little extent.
Moreover, in the present investigation absorption at longer wave-
length enables it to avoid the background interference caused by
the biological constituents. Therefore the present method has an
advantage over the reported method. Also, it is for the first time
we have reported peroxidase activity in plant sample such as Tal-
inum fruticosum and Santalum paniculatum.
is probably the most common enzyme used as
a reporter
(enzyme-labeled antibody) in enzyme immunoassays) and also
used for developing convenient and quick methods for the deter-
mination and quantification of hydrogen peroxide in both biologi-
cal and industrial samples [3]. In modern organic synthesis, uses of
purified peroxidases, immobilized in various matrices or peroxi-
dase rich microorganisms provide a better way of downstream
processing for product quality and recovery and has become an
important emerging application of this enzyme [4]. Gene gun
delivery system and electroporation study inside the living cells
for the time measurements of intracellular dissolved oxygen con-
centration are also its applications as sensors [5]. Peroxidase has
also been used for analytical applications in diagnostic kits, for
the quantification of clinically important biomarkers such as uric
acid, glucose, cholesterol, and lactose [6]. It is also used in the opti-
cal measurement of reactive oxygen species (ROS) [7] which is very
important since it can act as part of signal transduction pathway
from cell wall loosening and growth by elongation as well as
cross-linking of cell wall components. The balance between cleav-
age and cross-linking of cell wall is associated with ROS action and
with H2O2 and ascorbate concentration [8].
Many methods have been reported for the determination of
H2O2 or glucose using peroxidase activity such as fluorescence
[9], chemiluminescence [10], electrochemical [11], magneto elastic
sensors [12], amperometric [13], flow injection analysis [14],
potentiometric assay [15], and coulometric biosensor techniques
[16]. The instruments used in these are either very expensive or less
versatile. The selectivity of the luminescence is poor. One of the
drawbacks of electrochemical sensors is the interference by oxida-
tion or reduction of other compounds at the working potential and
also, electro-analytical technique needs several steps to immobilize
the enzyme on a solid support, which may reduce the enzyme activ-
ity (15%) resulting in the waste of expensive biocatalyst, and it is
also a time consuming process [7]. Spectrophotometric method is
considered to be the most convenient analytical technique, because
of its inherent simplicity, low cost, and wide availability in most
quality control laboratories. Some of the co-substrates commonly
used for the enzymatic determination of peroxidase activity and
H2O2 include phenol with 4-aminoantipyrine [17], o-dianisidine,
3,30,5,50-tetramethyl benzidine (TMB) and o-phenylenediamine
[18], 2,20-azino-bis(3-ethylbenzthiazoline-6-sulfonic acid) (ABTS)
[19], pyrocatechol and aniline [20], MBTH with DBZ [21], gold nano-
particles with chitosan [22], silica sol–gel hybrid membranes [23]
and 2,4-dimethoxyaniline (2,4-DMA) [24]. But these reagents have
some limitations such as carcinogenicity and mutagenicity of
o-dianisidine, and solubility of DBZ, 2,4-DMA and TMB in water is
quiet difficult. Poor sensitivity is the common disadvantage in case
of 4-aminoantipyrine, also auto oxidation of phenol and TMB in
presence of atmospheric oxygen is the main drawback in using
these reagents. 2-Hydroxy-1-naphthaldehyde thiosemicarbazon
(HNT) [9] is soluble in dimethylformamide, which itself will
interfere with the determination of H2O2 assay and derivatives of
thiosemicarbazon are highly toxic substances.
Materials and methods
Instrumentation
A Jasco model UVIDEC-610 ultraviolet–visible (UV–Vis) spectro-
photometer with 1.0-cm matched cells was used for all absorbance
measurements. A water bath shaker (NSW 133, New Delhi, India)
was used to maintain constant temperature for color development.
All pH measurements and adjustments were done by a digital pH
meter (model EQ-614, Equip-tronics, Mumbai, India).
Chemical reagents and their preparation
H2O2 stock solution (1.0%, v/v) was prepared by diluting the
commercial reagent (30%, v/v, E. Merck, Mumbai, India), and its
concentration was standardized by titration with secondary stan-
dard, KMnO4. DMA was purchased from Cica-Reagent (Kanto
Chemical CO., INC) Tokyo. DMA (58.75 mM) was prepared by dis-
solving 45 mg in 3.2 mL of dil. HCl (0.112 N) and diluting to 5 mL
with distilled water. Peroxidase (100 U/mg) was purchased from
Himedia (Mumbai, India), and the solution was prepared by dis-
solving 2 mg in 10 mL of 0.1 mol/L KH2PO4/NaOH buffer (pH 6.0).
Double distilled water was used throughout the experiment. All
reagents used were of analytical grade unless stated otherwise.
Sample collection and preparation for the determination of peroxidase
activity in crude plant extract samples
As a source of peroxidase, leaf portion from plants of Boerhavia
diffusa (Punarnava), Cissus quadrangularis (Veldt Grape or Devil’s
Backbone), S. paniculatum (Sandalwood), T. fruticosum (Waterleaf),
Prosopis cineraria (Velvet Mesquite), were collected from local agri-
cultural fields and their extracts were prepared as reported in our
previous study [24] and used. The accuracy of the proposed meth-
od was also simultaneously evaluated by comparing with guaiacol
assay based on spectrophotometric method.
General experimental protocol for the assay
In order to overcome some of the above drawbacks, we
attempted a new approach for the quantification of HRP and
H2O2 based on spectrophotometric method by using DMA as a
chromogenic probe. In our earlier study, the same reagent has been
used for the quantification of glucose in human serum [25]. Here a
new kinetic mathematical model has been used to evaluate the
Michaelis–Menten constants of the substrates and the results
obtained were compared with the previously reported 2,4-DMA
method [24]. In 2,4-DMA method the measurement of peroxidase
Experimental protocol conducted for the quantification of hydrogen
peroxide
The linearity for the assay of H2O2 was examined in a 3 mL of
the reaction mixture containing 1958
idase in a 100 mM acetic acid/sodium acetate buffer of pH 4.2. The
H2O2 concentrations used were in the range of 2–1152 M. The
lM DMA and 4.73 nM perox-
l
change in the absorbance was continuously recorded at 740 nm,
at 1 min interval. The slope of the graph gave the initial rate reac-

H. Krishna et al. / Spectrochimica Acta Part A: Molecular and Biomolecular Spectroscopy 102 (2013) 75–81
77
for maximum absorption of the colored product was 740 nm. The
result is graphically presented as the inset in Fig. 1.
Evaluation of kinetic constants for DMA and H2O2
Separate experiment for each H2O2 concentration was per-
formed with varying concentrations of DMA. The H2O2 and DMA
concentration ranges of 25–407 and 489.5–1958 lM, respectively
in the final volume of 3 ml were used for each kinetic study. The
pH and temperature were kept constant. The results are shown
in Figs. 2 and 3.
Assuming the initial rates (Vo), a general equation for the mech-
anism in the forward direction is given as a function of all substrate
concentrations:
2O2
1
1
KmH
1
KmD
1
¼
þ
:
þ
:
ð1Þ
ð2Þ
ð3Þ
ð4Þ
Vo Vmax Vmax ½H2O2ꢂo Vmax ½Dꢂo
Rewriting the above equation as
Fig. 1. (A) Calibration graph for the quantification of H2O2
absorption spectrum of H2O2: 1958 M DMA + 4.73 nM peroxidase in 100 mM
acetate/acetic acid at pH 4.2 with varying concentrations of H2O2 (75, 100 and
125 M) and the corresponding reagent blank containing neither H2O2 nor enzyme
showed negligible absorption. Spectrum was recorded after incubating reaction
mixture for 5 min at 30 °C.
(lM). (B) Inset shows the
2O2
½H2O2ꢂo ½H2O2ꢂo KmH
½H2O2ꢂo
KDm ½H2O2ꢂ
l
o
¼
þ
:
þ
:
Vo
Vmax
Vmax ½H2O2ꢂo Vmax ½Dꢂo
l
Differentiating the above equation we get
ꢀ
ꢁ
ꢀ
ꢁ
1
Vo
KDm ꢃ ½H2O2ꢂ
1
½Dꢂo
o
½H2O2ꢂo ꢃ d
¼ 0 þ
ꢃ d
Vmax
tion, which was then plotted against the concentration of H2O2 to
obtain a calibration graph. The linearity for the assay of H2O2 was
found to be in the range of 2–288 lM. The calibration graph for the
Or
ꢀ
ꢁ
ꢀ
ꢁ
1
Vo
KmD
Vmax
1
½Dꢂo
quantification of H2O2 is shown in Fig. 1.
d
¼ 0 þ
ꢃ d
Experimental protocol conducted for the quantification of peroxidase
and protein
The change in absorbance with respect to time for the quantifi-
cation of peroxidase was measured continuously for 5 min in a
3 mL reaction mixture containing 1958
H2O2, in 100 mM acetic acid/sodium acetate buffer of pH 4.2 and
100 L of varying concentrations of peroxidase enzyme against
lM DMA and 72 lM
l
the corresponding control containing all the reagents except per-
oxidase. The linearity for fixed-time method was evaluated by
incubating the reaction mixture as described above for 10 min at
30 °C and measuring the absorbance of the colored solution. The
calibration graph for the peroxidase assay was found to be in the
range of 0.59–9.46 nM and 0.443–9.46 nM by kinetic and fixed
time method, respectively.
Total protein concentration was determined in triplicate by
Lowry et al. [27] method using bovine serum albumin as a
standard.
Fig. 2. Kinetic behavior of two substrate reactions for pure HRP (4.73 nM). Plot of
differentiation of co-substrate (DMA)–velocity relationship, according to the Eq. (4).
Unit of enzyme
One unit of enzyme is defined as that amount of enzyme, which
utilizes 1 lmol of H2O2 for the intermolecular coupling of 1 lmol
of DMA to form green colored coupled product per min under
the standard assay conditions.
Results and discussion
Absorption spectrum of hydrogen peroxide
The absorption spectrum of the colored solution produced at
(75, 100 and 125 lM) concentrations of H2O2 was measured by
the recommended general experimental protocol in a final 3 mL
of reaction mixture and the spectrum was recorded at a scan rate
of 2 nm/s after incubating the reaction mixture for 5 min at 30 °C
on a spectrophotometer in the wavelength range 400–800 nm
against the corresponding reagent blank. The optimum wavelength
Fig. 3. Kinetic behavior of two substrate reactions for pure HRP (4.73 nM). Plot of
differentiation of substrate (H2O2)–velocity relationship, according to the Eq. (7).

78
H. Krishna et al. / Spectrochimica Acta Part A: Molecular and Biomolecular Spectroscopy 102 (2013) 75–81
By plotting a graph of d(1/Vo) vs d(1/Do) we get
Effect of pH and concentration of buffer solutions
The effect of pH on the enzyme activity was studied by using
different buffers of concentrations ranging from 0.1 M to 2.0 M
such as acetic acid/sodium acetate buffer (pH 3.6–5.6), citric
acid/potassium citrate buffer (pH 3.6–5.6), KH2PO4/NaOH buffer
(pH 6.0–8.0), and KH2PO4/K2HPO4 buffer (pH 6.0–7.5). The activity
was maximum at pH 4.2 in a 100 mM acetic acid/sodium acetate
buffer solution as evident from Supplementary material, Fig. S5,
in the final 3 mL of the reaction mixture. Hence, acetate buffer of
pH 4.2 was selected for further assay procedures.
KmD
Vmax
Slope ¼
and Intercept ¼ 0
From Lineweaver–Burk plot of rate vs concentration of 2,5-DMA
for horseradish peroxidase (figure not shown, concentration of 2,5-
DMA range used between 40 and 2608
ity for DMA, was found to be 0.0390
l
M), VDmax, maximum veloc-
l
M minꢁ1 and from the plot of
d(1/Vo) vs d(1/[D]o) (Fig. 2) a mean slope value for each fixed con-
centration of H2O2 and varying concentrations of Do was calculated
and used for the evaluation of KDm and was found to be
KDm = 1458
lM.
Effect of reagent co-substrate on the enzyme activity
Again rewriting Eq. (1) we get
Solutions containing DMA concentrations ranging from 326 to
3912 lM were added to reaction mixture containing 120.0 lM
2O2
½Dꢂo
½Dꢂo KHm
½Dꢂo
KDm ½Dꢂo
H2O2, 4.73 nM peroxidase, and 100 mM acetate/acetic acid buffer
of pH 4.0 in a final 3 mL total volume, for the study of co-substrate
effect on enzyme assay. As shown in Supplementary material,
Fig. S6, rate of reaction increased, with increase in concentration
¼
þ
:
þ
:
ð5Þ
ð6Þ
ð7Þ
Vo
Vmax Vmax ½H2O2ꢂo Vmax ½Dꢂo
Again differentiating the above equation we get
ꢀ
ꢁ
ꢀ
ꢁ
2O2
1
Vo
KHm ꢃ ½Dꢂo
1
of DMA up to 1958
in enzyme activity was observed. Therefore, 1958
l
M, beyond which no considerable increase
M DMA was
½Dꢂo ꢃ d
¼ 0 þ
ꢃ d
l
Vmax
½H2O2ꢂo
chosen as optimum concentration of co-substrate in the final
3 mL of the reaction mixture.
Or
ꢀ
ꢁ
ꢀ
1
ꢁ
2O2
1
Vo
KmH
d
¼ 0 þ
ꢃ d
Effect of temperature on the sensitivity of enzyme assay
The temperature effect on the enzyme assay was studied by
adding solutions containing 1958 lM DMA, 72.0 lM H2O2, and
Vmax
½H2O2ꢂo
Again by plotting a graph of d(1/Vo) vs d(1/[H2O2]o) we get
4.73 nM peroxidase in a 100 mM acetate/acetic acid buffer of pH
4.2 and incubating the reaction mixture for 10 min at temperatures
in the range 0–100 °C in a total of 3 mL volume. A proportional in-
crease in the absorbance values of color formed was observed with
increase in temperature up to 30 °C as evident from Fig. 5 and the
values decreased thereafter. Therefore, 30 °C was chosen as opti-
mum temperature for the assay.
2O2
KmH
Slope ¼
and Intercept ¼ 0
Vmax
Similarly, from Lineweaver–Burk plot of rate vs concentration of
H2O2 for horseradish peroxidase (Fig. 4, concentration of H2O2
range used between 2 and 1152
l
M), VHm2aOx2 was found to be
0.1382
l
M minꢁ1 and from the plot of d(1/Vo) vs d(1/[H2O2]o)
Fig. 3, a mean slope value for each fixed concentration of DMA
and varying concentration of H2O2 was calculated and used for
Calibration graphs for the assay
2O2
2O2
the evaluation of KHm and found that KmH = 301
lM.
The calibration curve for H2O2 assay was carried out under the
optimized experimental condition which is shown in Fig. 1. The re-
sults showed that there was an increase in the rate of reaction with
Optimization of the experimental conditions for maximum activity of
enzyme assay
increase in the concentration of H2O2 up to 290
mained constant thereafter. The linearity for the calibration graph
was observed in the range of 2–288 M (figure not shown). The
detection limits, quantification limits and molar extinction co-effi-
cient for the quantification of H2O2 were done by fixed time meth-
lM H2O2, and re-
Optimizations of experimental conditions parameters such as
effect of substrates, co-substrates, different buffer concentrations
of pH 3.0–8.0, temperature and incubation period, which affect en-
zyme assay, have been studied.
l
od and found to be 0.55 lM (sample/noise ratio, S/N ratio as a
general definition it is the reciprocal value of co-efficient of varia-
tion, [S/N] = 0.4), 1.83
l
M and 0.58 ꢀ 104 L/mol/cm, respectively.
Fig. 5. Effect of temperature on the sensitivity of enzyme assay containing 1958
DMA + 72
MꢃH2O2 + 4.73 nM peroxidase in 100 mM acetate/acetic acid at pH 4.2
in a final 3 mL of the reaction mixture.
lM
Fig. 4. (A) Lineweaver–Burk plots for horseradish peroxidase by the proposed
method. (B) Inset shows the value of 1/V at the higher concentration of H2O2.
l

H. Krishna et al. / Spectrochimica Acta Part A: Molecular and Biomolecular Spectroscopy 102 (2013) 75–81
79
0.0292 ꢀ 103 minꢁ1 and 0.0970
l
Mꢁ1 minꢁ1, respectively. The cat-
alytic efficiency (KDeff = 1/slope ꢀ [E]o, where slope = 897.87 minꢁ1
ꢄ
ꢅ
and [E]o = 4.73 nM) and catalytic power KDpow ¼ VmH2ax2 =KmH2O2 of
O
the proposed method are KDeff = 2.354 ꢀ 105 Mꢁ1 minꢁ1 and
KDpow = 4.59 ꢀ 10ꢁ4
, respectively and by guaiacol method are
KGeff = 1.78 ꢀ 104 Mꢁ1 minꢁ1 and KpGow = 8.44 ꢀ 10ꢁ5 minꢁ1, respec-
tively which show that proposed method is 13 times more efficient
than the guaiacol method. The lower value of KmH
2O2
which was
301 lM for DMA/HRP, may be due to a stronger affinity of active
site of HRP in presence of DMA to that of H2O2 molecules which
signifies the extent of selectivity and specificity of the proposed
reaction. Also, these observations suggest that DMA can be a better
proton donor than guaiacol for the HRP assay.
Fig. 6. Calibration graph for the quantification of HRP by kinetic method (j) and
fixed time (+) method.
Discussion of proposed reaction pathway for the intermolecular
coupling of DMA in response to the enzymatic activity
The relative standard deviation was 1.68% (n = 6) for the determi-
nation of 72 lM H2O2. The linear ranges for the quantification of
HRP were 0.59–9.46 nM and 0.443–9.46 nM by the kinetic and
fixed time methods, respectively, as shown in Fig. 6. The limit of
detection and quantification for HRP were 0.0361 nM and
0.12 nM, respectively. The linearity range for HRP assay by some
of the reported analytical methods were 0.0227–1.136 nM for
chemiluminescence [10], 5.4 ꢀ 10ꢁ4 nM to 0.1088 nM for electro-
chemical [11] method and by spectrophotometric method its line-
arity range were 0.294–4.71 nM and 0.15–2.0 nM for MBTH-DBZ
[21] method and 0.2022–1.13 nM and 0.0079–0.756 nM for 2,4-
DMA [24] method, by kinetics and fixed time method respectively.
The probable reaction mechanism involved is based on the per-
oxidase mediated self coupling of activated DMA which is similar
to our earlier report on 2,4-DMA [24]. Here, 2,5-dimethoxy aniline
acts as a mediator for the electron transfer in this peroxidase cat-
alyzed assay system. The above mentioned reaction is analogous
to HRP-catalyzed synthesis of polyaniline as suggested by Caramy-
shev et al. [28]. The colored product was confirmed by spectral
analysis as 2,4-dimethoxy aniline derivative and from that we
can presume that 2,5-dimethoxy aniline also undergoes in a simi-
lar way. Even though the resulting intermolecular coupling of both
the derivatives of dimethoxy aniline (2,4-DMA and 2,5-DMA) are
same radical species but may be due to the stabilization of the
meta position of methoxy group in the excited state of 2,5-DMA
derivative that makes it to absorb at higher wavelength region
might have resulted in different colored species [26]. The peroxi-
dase-catalyzed reaction of DMA is as shown in Scheme 1. The free
radical is released by the oxidation of H2O2 through a ferryl inter-
Evaluation of kinetic parameters for the enzyme assay
Kinetic parameters for the enzyme assay were studied under the
optimized experimental conditions. A Lineweaver–Burk plot was
used for the evaluation of Michaelis–Menten constants of H2O2
mediate (FeIV = O-porphyrin
p-cation radical) of the peroxidase
[29]. Under the reaction condition examined, DMA loses one elec-
tron and one proton upon enzymatic oxidation in the presence of
concentration ranging from 2 to 1152
equation obtain for the straight line was
V is minꢁ1) with a correlation coefficient of 0.9889 (n = 6), as shown
l
M. The linear regression
1
897:87
¼
þ 7:234 (unit of
V
CH2
O
2
H2O2, forming the
p-cation radical [30]. This p-cation radical
in Fig. 4. Also inset of Fig. 4 (B) shows the value of 1/V at the higher
undergoes coupling with another radical, forming an intense
green-colored product showing strong absorption at 740 nm.
2O2
concentration of H2O2. The values of KH
and VHm2aOx2 for the perox-
m
idase enzyme from the Lineweaver–Burk plot were found to be
301 M and 0.1382 EU/min, respectively, which are lower when
compared to our earlier work [24] that had the values of 1733
l
lM
Application of the proposed assay for the assessment of peroxidase
activity in crude plant extracts
and 0.3552 EU/min, respectively. The catalytic constant
ꢂ
ꢃ
ꢂ
ꢃ
H
O
2
2
Vmax
KcaOt
Kcat
¼
and specificity constant
for peroxidase
KHm2
½Eꢂo
2
The plant samples selected for the study are very useful in the
field of medicine and used especially in Ayurveda, the indigenous
substrates hydrogen peroxide and DMA was found to be
NH
NH2
OCH3
OCH3
+
Peroxidase
1.
H2O2
H3CO
H3CO
DMA
NH
imine radical
OCH3
OCH3
OCH3
OCH3
OCH3
OCH3
OCH3
OCH3
N
N
N
N
2.
2
OCH3
imine radical
OCH3
delocalised coupled
product
Scheme 1. Proposed reaction pathway for the formation of green-colored product of 2,5-dimethoxyaniline.

80
H. Krishna et al. / Spectrochimica Acta Part A: Molecular and Biomolecular Spectroscopy 102 (2013) 75–81
Table 1
Assessment of peroxidase activity in crude plant extracts.
Plant samples (common name)
Activity (units)a
Proteina
(mg)
Specific activity
(units/mg)
Relative catalytic power
Relative catalytic efficiency
ꢂ
ꢃ
ꢂ
ꢃ
KDeff
KGeff
KDpow
KGpow
DMA
50.32 1.2
Guaiacol
DMA
Guaiacol
Boerhavia diffusa (Punarnava)
Cissus quadrangularis (Devil’s
Backbone)
62.40 1.7 110.40 1.0
0.455 0.565
3.779 4.019
8.40
17.97
1.84
4.265
116.60 1.1 124.00 1.5
30.85 1.2
Prosopis cineraria (Banni leaf)
Santalum paniculatum (Sandalwood)
Talinum fruticosum (Waterleaf)
124.26 1.0 130.77 1.2 311.30 1.2
0.385 0.420
0.123 0.147
0.199 0.186
10.16
3.96
2.85
10.20
12.28
2.85
36.50 1.8
5.45 1.3
43.40 1.3 294.96 1.5
5.12 1.6 27.34 1.1
a
Mean (n = 3 replicate determinations) R.S.D.
medicine of India. The assessment of peroxidase activity in crude
plant tissue samples was performed carefully by analyzing the ef-
fect of different buffers of pH 4.0–8.0 and considering buffer to tis-
sue ratio from 5:1 to 15:1 mL/g. The maximum peroxidase activity
was observed in the buffer solution of potassium dihydrogen phos-
phate/sodium hydroxide at pH 6.0 and highest specific activity was
observed at 10:1 mL/g ratio. Hence, the extraction of the crude ex-
tracts was carried out using the buffer solution of potassium dihy-
drogen phosphate/sodium hydroxide at pH of 6.0 and at 10:1 mL/g
ratio for getting a better crude extract of the system.
Analysis of the plant extracts obtained from P. cineraria and C.
quadrangularis showed high peroxidase activity as determined by
the proposed method and reported guaiacol method with high spe-
cific activity whereas T. fruticosum had less peroxidase activity and
the results are shown in Table 1. Plant extracts of B. diffusa, S. pan-
iculatum, and P. cineraria had the highest content of proteins. The
catalytic efficiency of all plant samples was analyzed by perform-
ing a control containing all the reagents including crude plant ex-
tract except H2O2 to avoid possible polyphenol oxidases
interference with the peroxidase enzyme which could oxidize
2,5-DMA. The relative catalytic efficiency of S. paniculatum and P.
cineraria was high compared to B. diffusa, and T. fruticosum extracts.
C. quadrangularis and P. cineraria had high catalytic power com-
pared to other plant extracts when examined under the assay
conditions.
tion of biosensors. The relative catalytic efficiency and relative cat-
alytic power of the DMA/peroxidase system is higher than unity
with reference to guaiacol/peroxidase system for all crude extracts
examined, which prove higher detection rate of the DMA/
peroxidase system thereby claiming advantage of the proposed
method. Regarding the superiority of the proposed method, it is
suggested that DMA/peroxidase system can be a better hydrogen
donor for the HRP assay than guaiacol system as evident from
the kinetic data. Furthermore, it is for the first time we have re-
ported peroxidase activity in plant sample such as T. fruticosum
and S. paniculatum.
Acknowledgment
One of the authors, Honnur Krishna would like to thank Univer-
sity of Mysore, Mysore, Karnataka, India (SC/ST special cell) for
financial support for the research work and for providing the re-
search laboratory facilities.
Appendix A. Supplementary material
Supplementary data associated with this article can be found, in
the online version, at http://dx.doi.org/10.1016/j.saa.2012.10.024.
References
Conclusion
[1] J.R. Whitaker, Principles of Enzymology for the Food Sciences, third ed., Marcel
Dekker, New York, 1985. p. 592.
[2] S. Shigeoka, T. Ishikawa, M. Tamoi, Y. Miyagawa, T. Takeda, Y. Yabuta, K.
Yoshimura, J. Exp. Bot. 53 (2002) 1305–1319.
[3] T.A. Sergeyeva, N.V. Lavrik, A.E. Rachkov, Z.I. Kazantseva, S.A. Piletsky, A.V.
El’skaya, Anal. Chim. Acta 391 (1999) 289–297.
[4] K. Ikehata, I.D. Buchanan, M.A. Pickard, D.W. Smith, Bioresour. Technol. 96
(2005) 1758–1770.
[5] Y.E.L. Koo, Y. Cao, R. Kopelman, S.M. Koo, M. Brasuel, M.A. Philbert, Anal. Chem.
76 (2004) 2498–2505.
The reagents required for the assay are inexpensive, requires in
small quantity, thereby making the assay affordable. The higher
apparent molar absorptivity, lower values of detection limits and
RSD for HRP are the claims for the superiority of the method. Evi-
dence from the kinetic study also explains that the lesser values
2O2
(KmH = 301 M and VmH2aOx2 = 0.1382 EU minꢁ1) for the peroxidase
l
enzyme from the Lineweaver–Burk plot, indicate that there is a
stronger affinity of active site of DMA to that of H2O2 molecules
which signifies selectivity and specificity of the proposed reaction.
The intensely green colored chromogenic product obtained by the
coupling of DMA is stable with high molar extinction co-efficient.
This method can be adopted in the routine analysis for the quanti-
fication of peroxidase activity in crude extracts of plant samples
and also be used for quantifying any other substrate which is enzy-
matically converted into H2O2. The main advantage of the pro-
posed method over the 2,4-DMA method is that the solubility of
2,4-DMA is in alcohol and 2,5-DMA is in dil HCl (0.112 N) wherein
alcohol medium inhibition of peroxidase activity will take place.
Moreover, in the present investigation absorption at longer wave-
length enables it to avoid the background interference caused by
the biological constituents [25]. Even though some of the tech-
niques claim better sensitivity than this proposed method, but they
require costly instruments and involve multiple steps in prepara-
[6] E. Agostini, J.H. Andez-Ruiz, M.B. Arnao, S.R. Milrad, H.A. Tigier, M. Acosta,
Biotechnol. Appl. Biochem. 35 (2002) 1–7.
[7] A.K. Poulsen, A.M. Scharff-Poulsen, L.F. Olsen, Anal. Biochem. 366 (2007) 29–
36.
[8] F. Passardi, C. Penel, C. Dunand, Trends Plant Sci. 9 (2004) 534–540.
[9] B. Tang, Y. Wang, Spectrochim. Acta A 59 (2003) 2867–2874.
[10] X. Yang, Y. Guo, Z. Mei, Anal. Biochem. 393 (2009) 56–61.
[11] K. Jiao, W. Sun, Microchem. J. 72 (2002) 123–130.
[12] P. Panga, Y. Zhanga, S. Ge, Q. Cai, S. Yao, C.A. Grimes, Sensor. Actuat. B Chem.
136 (2009) 310–314.
[13] F. Gao, R. Yuan, Y. Chai, S. Chen, S. Cao, M. Tang, J. Biochem. Biophys. Meth. 70
(2007) 407–413.
[14] I. da cruz Vieira, O. Fatibello-Filho, Analyst 123 (1998) 1809–1812.
[15] F. Deyhimi, F. Nami, J. Mol. Catal. B Enzym. 68 (2011) 162–167.
[16] F. Vianello, L. Zennaro, A. Rigo, Biosens. Bioelectron. 22 (2007) 2694–2699.
[17] B. Tang, Y. Wang, L. Ma, Anal. Bioanal. Chem. 378 (2004) 523–528.
[18] A.V. Kireyko, I.A. Veselova, T.N. Shekhovtsova, Russ. J. Biochem. 32 (2006) 71–
77.
[19] Q. Huang, W.J. Weber, Environ. Sci. Technol. 39 (2005) 6029–6036.
[20] A.M. Rad, H. Ghourchian, A.A. Moosavi-Movahedi, J. Hong, K. Nazari, Anal.
Biochem. 362 (2007) 38–43.
[21] P. Nagaraja, A. Shivakumar, A.K. Shrestha, Anal. Biochem. 395 (2009) 231–236.

H. Krishna et al. / Spectrochimica Acta Part A: Molecular and Biomolecular Spectroscopy 102 (2013) 75–81
81
[22] Q. Xu, C. Mao, N.N. Liu, J.J. Zhu, J. Sheng, Biosens. Bioelectron. 22 (2006) 768–
773.
[27] O.H. Lowry, N.J. Rosebrough, A.L. Farr, R.J. Randall, J. Biol. Chem. 193 (1951)
265–275.
[23] W. Li, R. Yuan, Y. Chai, L. Zhou, S. Chen, N. Li, J. Biochem. Biophys. Meth. 70
(2008) 830–837.
[24] A. Shivakumar, R. Dinesh, Honnur Krishna, P. Nagaraja, Enzym. Microb.
Technol. 47 (2010) 243–248.
[25] P. Nagaraja, Honnur Krishna, A.. Shivakumar, A.K. Shrestha, Clin. Biochem. 45
(2012) 139–143.
[28] A.V. Caramyshev, V.M. Lobachov, D.V. Selivanov, E.V. Sheval, A.K. Vorobiev,
O.N. Katasova, V.Y. Polyakov, A.A. Makarov, I.Y. Sakharov, Biomacromolecules
8 (2007) 2549–2555.
[29] C. Regalado, B.E. Garcia-Almendárez, M.A. Duarte-Vázquez, Phytochem. Rev. 3
(2004) 243–256.
[30] J. Huang, H.B. Dunford, Can. J. Chem. 68 (1990) 2159–2163.
[26] M. Fogiel, The Organic Chemistry Problem Solver, Research and Education
Association, 1, 1978, p. 1025.