Mimicking peroxidase activity by a polymer-supported oxidovanadium(IV) Schiff base complex derived from salicylaldehyde and 1,3-diamino-2-hydroxypropane

Article abstract of DOI:10.1016/j.jinorgbio.2015.01.012

The polymer-supported oxidovanadium(IV) complex PS-[V^IVO(sal-dahp)] (2) (PS = chloromethylated polystyrene crosslinked with 5% divinylbenzene, and H₃sal-dahp = dibasic pentadentate ligand derived from salicylaldehyde and 1,3-diamino-2-hydroxypropane) was prepared from the corresponding monomeric oxidovanadium(IV) complex [V^IVO(Hsal-dahp)(DMSO)] (1), characterized and successfully used as catalyst for the peroxidase-like oxidation of pyrogallol. The oxidation of pyrogallol to purpurogallin with PS-[V^IVO(sal-dahp)] (2) was achieved under mild conditions at pH 7 buffered solution. Plausible intermediate species formed during peroxidase mimicking experiments are proposed, by studying the model complex [V^IVO(Hsal-dahp)(DMSO)] (1) by UV-visible and ⁵¹V NMR spectroscopies. The high peroxidase mimicking ability of polymer-supported complex 2, its stability in a wide pH range, the easy separation from the reaction media, and the reusability without considerable decrease in activity, suggest that this heterogeneous catalyst has high potential for application in sustainable industrial catalysis.

Full text of DOI:10.1016/j.jinorgbio.2015.01.012

Mimicking peroxidase activity by a polymer-supported oxidovanadium(IV)
Schiff base complex derived from salicylaldehyde and 1,3-diamino-2-
hydroxypropane
Mannar R. Maurya, Nikita Chaudhary, Fernando Avecilla, Isabel Cor-
reia
PII:
DOI:
Reference:
S0162-0134(15)00029-X
doi: 10.1016/j.jinorgbio.2015.01.012
JIB 9656
To appear in:
Journal of Inorganic Biochemistry
Please cite this article as: Mannar R. Maurya, Nikita Chaudhary, Fernando Avecilla, Is-
abel Correia, Mimicking peroxidase activity by a polymer-supported oxidovanadium(IV)
Schiff base complex derived from salicylaldehyde and 1,3-diamino-2-hydroxypropane,
Journal of Inorganic Biochemistry (2015), doi: 10.1016/j.jinorgbio.2015.01.012
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ACCEPTED MANUSCRIPT
Mimicking peroxidase activity by a polymer-supported oxidovanadium(IV)
Schiff base complex derived from salicylaldehyde and 1,3-diamino-2-
hydroxypropane
Mannar R. Mauryaa,*, Nikita Chaudharya, Fernando Avecillab, Isabel Correiac
a Department of Chemistry, Indian Institute of Technology Roorkee, Roorkee 247667, India
b Departamento de Química Fundamental, Universidade da Coruña, Campus de A Zapateira,
15071 A Coruña, Spain
c Centro de Química Estrutural, Instituto Superior Técnico, Universidade de Lisboa, Av. Rovisco
Pais 1, 1049-001 Lisbon, Portugal
Keywords: Vanadium; Polymer-supported; Crystal structure; Peroxidase mimetic catalyst;
Oxidation.
--------------------------------------------------------------
* Corresponding author. Tel.: +91 1332 285327; fax: +91 1332 273560.
E-mail address: rkmanfcy@iitr.ac.in (M.R. Maurya).
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Abstract
The polymer-supported oxidovanadium(IV) complex PS-[VIVO(sal-dahp)] (2) (PS =
chloromethylated polystyrene crosslinked with 5% divinylbenzene, and H3sal-dahp = dibasic
pentadentate ligand derived from salicylaldehyde and 1,3-diamino-2-hydroxypropane) was
prepared from the corresponding monomeric oxidovanadium (IV) complex [VIVO(Hsal-
dahp)(DMSO)] (1), characterized and successfully used as catalyst for the peroxidase-like
oxidation of pyrogallol. The oxidation of pyrogallol to purpurogallin with PS-[VIVO(sal-dahp)]
(2) was achieved under mild conditions at pH 7 buffered solution. Plausible intermediate species
formed during peroxidase mimicking experiments are proposed, by studying the model complex
[VIVO(Hsal-dahp)(DMSO)] (1) by UV-visible and 51V NMR spectroscopies. The high
peroxidase mimicking ability of polymer-supported complex 2, its stability in a wide pH range,
the easy separation from the reaction media, and the reusability without considerable decrease in
activity, suggest that this heterogeneous catalyst has high potential for application in sustainable
industrial catalysis.
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1. Introduction
Peroxidase enzymes have important applications in many physiological reactions [1–7].
Peroxidase-like activity of haloperoxidases has also been reported towards classical peroxidase
substrates using recombinant vanadium haloperoxidases [8]. However, the industrial application
of natural or recombinant enzymes is not economically viable due to difficult bulk production,
high costs, poor stability and very specific reaction conditions (such as pH and reaction
medium). Applications of heterogeneous vanadium catalysts can potentially overcome these
problems. The V2O5 nanowire [9] has been used as a heterogeneous catalyst mimicking
peroxidase activity. However, to the best of our knowledge no examples of peroxidase like
activity by polymer supported vanadium complexes have been reported so far.
The application of polymer supported vanadium complexes as catalysts has shown
tremendous growth over the past few years [10,11]. Vanadium complexes in the presence of
oxidants like H2O2 and tert-butylhydroperoxide (TBHP) are good oxidation catalysts as they
easily form intermediate species containing an electron rich peroxido group, able to transfer
oxygen to the organic substrate [12,13,14]. The polymer support mimics the protein mantle and
prevents the decomposition of the complexes which in turn enhances the turn over numbers.
These observations inspired us to design vanadium complexes covalently linked to Merrifield
resin (chloromethylated polystyrene) to obtain biomimetic heterogeneous catalysts. Depending
on the nature of ligands and metal complexes different strategies were implemented for the resin
modification [10,11,15].
Smith et al. [16] prepared an oxidovanadium(IV) complex [VIVO(Hsal-dahp)]n (H3sal-dahp =
dibasic pentadentate ligand derived from salicylaldehyde and 1,3-diamino-2-hydroxypropane) in
which the ligand is similar to salen [17] but has an additional free hydroxyl group which can be
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used for immobilization on e.g. polystyrene. Despite showing good catalytic properties complex
[VIVO(Hsal-dahp)]n could not be immobilized on chloromethylated polystyrene due to its
polymeric form, consisting of ---(L)V=O---(L)V=O--- chains. However, the complex can easily
be converted into monomeric species using polar solvents like DMSO or DMF, which can then
be immobilized on chloromethylated polystyrene.
In this contribution, we report the isolation of monomeric [VIVO(Hsal-dahp)(DMSO)]
complex, its immobilization on chloromethylated polystyrene and its characterization. The
polymer supported complex (abbreviated as PS-[VIVO(sal-dahp)]) shows good peroxidase-like
activity in aqueous medium. The analogous complex [VVO(hap-dahp)] (derived from o-
hydroxyacetophenone and 1,3-diaminohydroxypropane) has been encapsulated in the nanopores
of zeolite-Y and applied as catalyst for oxidation reactions [18].
2. Experimental Section
2.1. Materials and methods
V2O5, (Loba Chemie, India), 1,3-diamino-2-hydroxypropane (Aldrich, U.S.A.),
salicylaldehyde (Sisco Research, India) and 30% aqueous H2O2 (Qualigens, India) were used as
obtained. A gift sample of chloromethylated polystyrene [18.9 % Cl (5.35 mmol Cl per gram of
resin)] cross-linked with 5 % divinylbenzene (PS-Cl) was obtained from Thermax Limited, Pune,
India. All other chemicals and solvents used were of AR grade.
Vanadium content in PS-[VIVO(sal-dahp)] (2) was determined by inductively coupled
plasma-mass spectrometry (ICP-MS). The sample was prepared by taking 0.020 g of 2 and
treating it with 6 mL of nitric acid-hydrochloric acid (1 : 1) mixture on a hot plate. After
complete leaching the residue was dissolved in deionized-distilled water, filtered and diluted to
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50 mL. Elemental analyses (C, H, N and S) of the complexes were obtained in an Elementar
model Vario-EL-III. Thermal stability of neat, as well as, polymer-supported complexes was
studied using a Perkin-Elmer (Pyris Diamond) thermogravimetric analyzer under oxygen
atmosphere (flow rate: 200 ml/ min, heating rate: 10 oC/ min and temperature range: 35-1000 oC)
using alumina powder as reference. IR spectra with 2 cm–1 resolution were recorded as KBr
pellets on a Nicolet 1100 FT-IR spectrometer. Electronic absorption spectra of ligand and neat
complex were recorded in methanol using a Shimadzu 1601 UV-visible (UV-vis)
spectrophotometer. The spectrum of polymer-supported complex was recorded in Nujol by
layering the sample mull inside a cuvette and keeping the reference cuvette layered with Nujol.
EPR spectra were recorded with a Bruker ESP 300E X-band spectrometer coupled to a Bruker
ER041 X-band frequency meter (9.45 GHz) and the spin Hamiltonian parameters were obtained
by simulation of the spectra with the computer program of Rockenbauer and Korecz [19].
Complex 1 was dissolved in DMF (p.a. grade) (3 mM), previously degassed by passing N2 for 10
min, and the solution was immediately frozen in liquid nitrogen. The EPR spectrum was
measured at 77 K. The EPR spectrum of solid polymer-supported complex 2 was measured at
room temperature in quartz tubes. 51V NMR spectra of ca. 3 mM solutions of complex 1 in DMF
(p.a. grade) (5% D2O added) were recorded on a Bruker Avance III 400 MHz instrument. 51V
chemical shifts were referenced relative to neat VVOCl3 as external standard.
Atomic force microscopy (AFM) imaging was performed on a scanning probe
microscope from NTEGRA (NT-MDT) in a semi-contact mode. The morphology and elemental
analysis of both the choloromethylated polystyrene and polymer supported complex were
obtained by a FEI-Quanta 200 FEG field emission scanning electron microscope (FE-SEM)
coupled with an energy dispersive X-ray analyses (EDAX) accessory. The sample was coated
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with a thin film of gold to prevent surface charging and to protect the surface material from
thermal damage by electron beam. A uniform thickness of about 0.1 mm was maintained. A
constant voltage of 20 kV was applied so that only the outer surface was accessible.
2.2. Synthesis
[VIVO(acac)2] [20] and H3sal-dahp (I) [16] were prepared according to methods reported in the
literature.
2.2.1. [VIVO(Hsal-dahp)(DMSO)] (1)
The polymeric complex [VIVO(Hsal-dahp)]n was prepared by reaction of VOSO4·5H2O
(0.506 g, 2.00 mmol) and H3sal-dahp (0.596 g, 2.00 mmol) in methanol as reported in the
literature [16]. An alternative method was also used and is given here. To a stirred solution of
H3sal-dahp (0.596 g, 2.00 mmol) in methanol (10 mL) was added a solution of [VIVO(acac)2]
(0.530 g, 2.00 mmol) in methanol (10 mL) and the reaction mixture was refluxed for 1 h, after
which a light orange solid was formed. The solid was filtered, washed with methanol and dried
in vacuum over silica gel. Yield: 0.580 g (80 %).
Complex [VIVO(sal-dahp)]n was dissolved in a minimum amount of hot DMSO, filtered
and left for slow evaporation at room temperature. An orange solid slowly precipitated which
was filtered and dried in a vacuum desiccator over silica gel. Anal. Calc. for C19H22N2O5SV
([VIVO(Hsal-dahp)(DMSO)]) (441.39): C, 51.70; H, 5.02; N, 6.34; S, 7.26%. Found: C, 52.13;
H, 4.92; N, 6.50; S, 6.94%.
Light orange crystals of [VIVO(Hsal-dahp)(DMSO)]·H2O (1·H2O) suitable for X-ray
diffraction study was grown from DMSO, which were filtered and dried at room temperature.
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2.2.2. PS-[VIVO(sal-dahp)] (2)
Choloromethylated polystyrene beads (1.0 g) were suspended in DMF (20 mL) and left
for 2 h. A solution of [VIVO(Hsal-dahp)(DMSO)] (1.0 g, 2.3 mmol) in DMF (30 mL) was added
to the above suspension of beads. After adding triethylamine (3 mL) and K2CO3 (0.069 g, 0.50
o
mmol) the resulting mixture was heated at 90 C for 2 days with slow mechanical stirring.
Finally, the beads were filtered off, washed with DMF, followed by hot methanol and dried at 80
oC. Recovery yield: 94%. Vanadium content found by ICP-MS: 0.145 mmol g–1 of resin.
2.3. Peroxidase mimetic activity: catalytic oxidation of pyrogallol
Catalyst PS-[VIVO(sal-dahp)] was suspended in acetonitrile for 2 h before starting the
reaction. The peroxidase mimetic activity was determined using UV-Vis spectrophotometry by
monitoring the absorbance increase at 420 nm (due to the formation of purpurogallin, ɛ = 2640
M–1 cm–1) for up to 2 h [9,28]. The reaction was carried out in 3 mL of phosphate buffer (1M, pH
7). The reaction was started with the addition of 1 mL of 0.25×10–1 M solution of 30% H2O2 to 1
mL of 0.25×10–1 M pyrogallol solution and catalyst PS-[VIVO(sal-dahp)] (0.007 g). The steady
state kinetics were done by varying the concentration of pyrogallol (0.2–1.4 mM) while keeping
a fixed concentration of H2O2 (0.25×10–6 M) and catalyst PS-[VIVO(sal-dahp)] (0.007 g). The
reaction was carried out in 2 mL phosphate buffer (1 M, pH 7) and monitored
spectrophotometrically by measuring the formation of purpurogallin at 420 nm (ɛ = 2640 M–1
cm–1) in time scan mode. The initial reaction rates were fitted to Michaelis-Menten equation and
calculated. Lineaweaver-Burk linearizations were performed using Sigma-Plot software (Systat
Software Inc.) with enzyme kinetics module to determine kinetic parameters.
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2.4. X-Ray crystal structure determination
Three-dimensional X-ray data were collected on a Bruker Kappa Apex CCD
diffractometer at low temperature for 1·H2O by the - scan method. Reflections were measured
from a hemisphere of data collected from frames each of them covering 0.3º in . Of the 18878
reflections measured, all were corrected for Lorentz and polarization effects and for absorption
by multi-scan methods based on symmetry-equivalent and repeated reflections. A total of 2200
independent reflections exceeded the significance level (F/F) > 4.0. Complex scattering
factors were taken from the program package SHELXTL [21]. The structure was solved by
direct method and refined by full matrix least-squares on F2. Hydrogen atoms were left to refine
freely. Refinements were done with allowance for thermal anisotropy of all non-hydrogen atoms.
Further details of the crystal structure determination are given in Table 1. A final difference
Fourier map showed no residual density outside: 0.412 and –0.368 e.Å–3. A weighting scheme w
2
= 1/[σ2(Fo ) + (0.038600 P)2 + 1.162400 P], where P = (|Fo|2 + 2|Fc|2)/3, was used in the latter
stages of refinement.
<<Table 1>>
3. Results and discussion
Reaction between [VIVO(acac)2] (Hacac = acetylacetone) and tribasic pentadentate Schiff
base H3sal-dahp (I) in methanol leads to the formation of an oxidovanadium(IV) complex where
the ligand acts as dibasic tetradentate and coordinates through the phenolic oxygen atoms and the
azomethine nitrogen atoms. The hydroxyl oxygen of the ligand is not involved in the metal
coordination. The complex is polymeric in nature, having the formula [VIVO(Hsal-dahp)]n in the
solid state, with polymeric chains of V=O----V=O [16]. Dissolving [VIVO(Hsal-dahp)]n in hot
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o
(80 C) DMSO and leaving it for crystallization at room temperature results in breaking of the
V=O----V=O chains, yielding the monomeric species [VIVO(Hsal-dahp)(DMSO)] (1); Scheme 1.
Reaction of chloromethylpolystyrene cross-linked with 5% divinylbenzene (PS–Cl) with
complex 1 in DMF, in the presence of triethylamine, resulted in the formation of the
corresponding polymer-grafted oxidovanadium(IV) complex, PS–[VIVO(sal-hdap)] (2); Scheme
1. Complexes were characterized by spectroscopic techniques (IR, UV-vis and EPR),
thermogravimetry, AFM and FE–SEM as well as EDAX studies.
<<Scheme 1>>
3.1. TGA studies
The thermal stability of neat (1) and polymer-supported (2) complexes was studied by
thermogravimetric analysis. The neat complex [VIVO(Hsal-dahp)(DMSO)] (1) is stable up to ca.
160 oC. Thereafter, the decomposition of the organic part occurs in three consecutive exothermic
o
steps and the final residue (21.0% at ca. 500 C) corresponds to V2O5. The theoretical value
(20.5%) is close to the experimentally obtained one. The thermal decomposition of PS-
[VIVO(sal-dahp)] (2) occurs in three steps. The weight loss of 4.8%, observed in the 100-200 oC
temperature range, may be due to loss of trapped solvent and water molecules. The weight loss in
o
the 200-400 C temperature range is possibly due to partial collapse of the complex moiety.
Major decomposition occurs between 400-500 oC, which is tentatively attributed to
decomposition of the major part of the polymer residue along with decomposition of the
remaining part of the organic ligand. Further weight loss of 1.8% was observed between 500-900
oC. The final residue of 0.9% is due to V2O5 and the vanadium content calculated from this value
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is 0.17 mmol g–1 of resin, which is roughly in agreement with the vanadium content obtained
from ICP-MS (0.145 mmol g–1 of resin) and EDAX (0.127 mmol g–1 of resin).
3.2. Structure description of [VIVO(Hsal-dahp)(DMSO)]·H2O (1·H2O).
An orange prism of [VIVO(Hsal-dahp)(DMSO)]·H2O (1·H2O) crystallizes in a
centrosymmetric orthorhombic space group, Pnma. One half water molecule and one half
[VIVO(Hsal-dahp)(DMSO)] complex are present in the asymmetric unit and grow through x, -
y+1/2, z (#1) symmetry transformation for the generation of equivalent atoms. The complex
adopts a six-coordinated structure with distorted octahedral geometry. The phenolic oxygen and
the imine nitrogen atoms of the ligand coordinate the vanadium center while the hydroxyl group
does not participate in the coordination. One DMSO molecule and one oxido group complete the
coordination sphere. The equatorial plane therefore comprises the [N(1), O(1), N(1A), O(1A)]
donor atom set, and the VVO center stands above this plane by 0.3455(12) Å.
Fig. 1 shows an ORTEP representation of 1·H2O and Fig. 2 shows the crystal packing in
which the complex forms an antiparallel dimer through - interactions between phenol rings
and C=N bonds of neighbouring molecules. Table 2 presents selected bond lengths and angles.
The distances between centroids are: dc1–c2 = 3.457 Å [c1 (C7AB-N1AB), c2 (C1AF-C2AF-
C3AF-C4AF-C5AF-C6AF)], dc3–c4 = 3.457 Å [c3 (C7AF-N1AF), c4 (C1AB-C2AB-C3AB-
C4AB-C5AB-C6AB)]. Intermolecular hydrogen bonds between water molecules and complexes
are present in the crystal packing (see Table 3). An intramolecular hydrogen bond is present
between the hydroxyl group and the coordinated DMSO molecule.
<<Figures 1 and 2>> <<Tables 2 and 3>>
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3.3. IR spectral studies
IR spectra of the ligand (H3sal-dahp), [VIVO(Hsal-dahp)(DMSO)], PS-Cl and PS-
[VIVO(sal-dahp)] are shown in supporting information (SI, Fig. SI-1A and 1B) and selected
bands and its tentative assignment are presented in Table 4. The ligand spectrum displays a broad
band at 3390 cm–1 due to free –OH groups. The bonding of hydroxyl oxygen to PS-Cl could not
be ascertained by IR spectroscopy as PS-[VIVO(sal-dahp)] (2) also exhibits a weak broad band at
3400 cm–1 due to trapped residual water in the polymer matrix. The band at 1607 cm–1, due to the
azomethine group, shifts to lower wavenumbers in both complexes, corroborating the
coordination of the nitrogen to the vanadium center. Neat complex 1 exhibits a sharp band at ca.
950 cm–1 due to monomeric ν(V=O) stretching mode, while the supported complex presents this
band at 965 cm–1. Other characteristic bands due to polystyrene are also present in the spectrum
of the supported complex (see SI).
<<Table 4>>
3.4. Electronic spectral studies
Electronic absorption spectra of a similar ligand (H2hap-dahp), its vanadium complex and
zeolite-Y encapsulated complex have been described in the literature [18]. The spectral data for
the ligand H3sal-dahp (I) and its neat as well as polymer-grafted vanadium complex are
presented in Table 4; Fig. 3 (A and B) presents the electronic absorption spectra. All ligand
bands appear in the spectra of the complexes with shifts in their positions. Bands due to d–d
transitions for [VIVO(Hsal-dahp)(DMSO)] (1) become clearer in DMF and have max at 488 (ɛ/
M–1 cm–1 = 98) and 748 nm (ɛ/ M–1 cm–1 = 7). The polymer-supported complex PS-[VIVO(sal-
dahp)] (2) exhibits a ligand to metal charge transfer (LMCT) band at 386 nm (Fig. 3B). In
addition, one d–d band at 515 nm could also be identified.
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<<Figures 3 >>
3.5. EPR spectral study
The 1st derivative EPR spectra of a “frozen” (77 K) solution of [VIVO(Hsal-
dahp)(DMSO)] (1) in DMF (4.7×10–3 M) and of solid PS-[VIVO(sal-dahp)] (2) at room
temperature are compared and presented in Fig. 4. The spectra of both complexes are well
resolved. The spectrum of 2 is characteristic of magnetically diluted VIVO-complexes and the
well-resolved EPR hyperfine features indicate that the vanadium(IV) centers are well dispersed
in the polymer matrix. Both spectra were simulated and the obtained spin Hamiltonian
parameters g// and A// (Table 5) are similar for both compounds confirming a similar donor set for
the neat and the PS-complex. The A// parameters are also in agreement with the value estimated
using the additivity relationship proposed by Wüthrich [22] and Chasteen [23] (with estimated
accuracy of ±3×10–4 cm–1), which is 161×10–4 cm–1, considering the equatorial donor set {2 ×
Nimine, 2 × Ophenolate } [17, 22, 23]. The predicted contributions to the parallel hyperfine coupling
constant for the donor groups under consideration are the same. EPR does not help in the
assignment of the apical donor, since the unpaired electron probably occupies a dxy orbital, which
has its lobes in the equatorial plane. Therefore, a molecule of DMSO might be present in the
apical position of the PS-complex, as it is in the neat complex.
<<Figure 4>> <<Table 5>>
3.6. Field emission-scanning electron microscope (FE-SEM) and energy dispersive X-ray
analysis (EDAX) studies
Field emission scanning electron micrographs (FE-SEM) of chloromethylated
polystyrene (PS-Cl) beads and PS-[VIVO(sal-dahp)] beads (2) were recorded and their images are
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shown in Fig. 5 (a and b). Only small morphological changes were observed among plain
polymer beads (PS-Cl) and polymer-anchored complex. However, a considerable increase in the
diameter of beads was noticed. After complex anchoring EDAX (Fig. 5 (a’ and b’)) analysis
shows 0.13+0.03 mmol g–1 of resin of average vanadium content on the surface, which is in
agreement with the value obtained with ICP-MS (0.145 mmol g–1).
<<Figure 5>>
3.7. Atomic force microscopic (AFM) study
The morphological changes of PS-Cl (Fig. 6) after anchoring of complex were also
studied by measuring the surface roughness using AFM. The surface roughness and mean height
for PS-Cl were 7.28 nm and 72.8 nm, respectively. Upon reaction of PS-Cl with [VIVO(Hsal-
dahp)(DMSO)], the surface roughness and mean height decreased to 1.92 nm and 13.8 nm,
respectively. This shows that the complex was anchored in the bead’s pores, thereby reducing its
porosity [24–26].
The morphology of the PS-complex 2 was also evaluated after 4 catalytic cycles, showing
small increase in the surface roughness and mean height (2.45 nm and 18.16 nm, respectively),
and thus suggests only minor morphological changes. These observations suggest low leaching
levels after 4 oxidation cycles.
<<Figure 6>>
3.8. Peroxidase mimetic activity of polymer supported complex
Peroxidase mimetic activity of polymer-supported complex PS–[VIVO(sal-dahp)] (2) was
investigated by using the typical peroxidase substrate, pyrogallol, [27] in the presence of H2O2.
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The peroxidase-like oxidation of pyrogallol (Scheme 2) yields purpurogallin, whose formation
can be monitored by UV-visible spectroscopy.
<<Scheme 2>>
Preliminary experiments were carried out using:
(i)
1 mL of 0.25×10–1 M pyrogallol in 3 mL of phosphate buffer (1M, pH 7);
1 mL of 0.25×10–1 M pyrogallol in 3 mL of phosphate buffer (1M, pH 7) and 1
mL of 0.25×10–1 M solution of 30% H2O2 ;
(ii)
(iii) 1 mL of 0.25×10–1 M pyrogallol in 3 mL phosphate buffer (1M, pH 7), 1 mL of
0.25×10–1 M solution of 30% H2O2 and different amounts of previously swollen
catalyst.
Reactions were followed by UV-visible spectrophotometry after 2 min of their
preparation by measuring the formation of the oxidation product at 420 nm. The transparent
pyrogallol solution turns yellow [28] in the presence of H2O2 showing a very weak absorption at
ca. 420 nm, due to the formation of purpurogallin; Fig. 7.
Solutions of set (iii) gradually changed to pink and then to reddish-brown along with the
appearance of a band at 420 nm, which was not present in the catalyst spectrum, indicating that
pyrogallol can be oxidized by H2O2 in the absence/presence of catalyst, with the oxidation extent
being different in the two cases. Furthermore, slight variations in color, along with different
absorption intensities at 420 nm, could be observed for samples containing different catalyst
amounts; Fig. 7. The new band at 420 nm is due to the formation of purpurogallin, which is
absent in the pyrogallol spectrum [29, 30]. The monitoring of the intensity of this band for the
sample containing 0.007 g of catalyst showed maximum measurable absorbance within 2 h; Fig.
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8. Further reaction optimization for maximizing pyrogallol oxidation was done by changing
catalyst and oxidant amounts, and pH of the reaction.
<<Figures 7 and 8>>
The effect of catalyst’s amount was evaluated by taking five different amounts (0.005–
0.020 g) under the above reaction conditions, i.e. 1 mL of 0.25×10–1 M pyrogallol in 3 mL
phosphate buffer (pH 7, 1 M), 1 mL of 0.25×10–1 M solution of 30% H2O2 and reaction
o
temperature 25 C. The plot in Fig. 9 shows that the oxidation of pyrogallol increased
considerably when using 0.005 g of catalyst (compared to the uncatalyzed reaction). Increasing
the catalyst’s amount to 0.007 g slightly improved the activity, while only minimal gains were
attained with further increase (to 0.010 g, 0.015 g or 0.020 g). However, the overall reaction time
decreased; e.g. the assay with 0.020 g of catalyst showed the activity equivalent to the assay with
0.007 g catalyst within 1.5 h (data not shown). Therefore, a compromise between reaction time
and catalyst amount was assumed, and other reaction conditions were optimized considering
0.007 g of catalyst.
The dependence of the pyrogallol oxidation on different amounts of oxidant was also
evaluated. Thus, different H2O2 concentrations (from 2.5 × 10–7 to 0.03 M) were considered in
each set having a fixed amount of pyrogallol solution (1 mL of 0.25×10–1 M), catalyst (0.007 g)
and pH 7 phosphate buffer (3 mL, 1 M). The reaction was monitored at 25 oC for 2 h and Fig. 10
shows the plots of absorbance at λmax = 420 nm at different concentrations of H2O2 with time.
The catalyst showed good response towards different H2O2 concentrations and the minimum
H2O2 concentration used, 0.25×10–6 M, showed a high activity profile.
<<Figures 9 and 10>>
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The peroxidase activity was also evaluated in buffer solutions of five different pH values.
The maximum measurable absorbance at λmax = 420 nm, independent of time, is shown in Fig.
11. The reaction rate was fastest at pH 8 for which maximum activity was reached within 20
min. Activity was comparatively slower at pH 7, for which maximum absorption (same value as
observed for pH 8) was obtained after 2h. For pH values 5, 6 and 9 the reaction became
extremely slow and the maximum absorbance at λmax = 420 nm was only ca. 0.25. Despite
showing maximum activity and higher reaction rate at pH 8, pH 7 was considered optimal, since
it is physiologically relevant and also environmentally similar to natural peroxidases in
biological systems. Moreover, no metal leaching was observed from the polymer-supported
complex at different pH values making it a versatile catalyst for different reaction environments.
<<Figure 11>>
Additionally, catalyst 2 is recyclable up to 4 cycles (after washing with acetonitrile and
drying) exhibiting roughly the same activity. In order to rule out the possibility of the peroxidase
activity being due to free leached vanadyl ([VO]2+) species, instead of the polymer-anchored
complex PS-[VIVO(sal-dahp)], after the catalytic runs at different pH values the reaction
mixtures were analyzed by ICP-MS. No vanadium content was found demonstrating that the
peroxidase mimetic activity is due to the bound PS-[VIVO(sal-dahp)].
A control experiment with VOSO4 (0.15 mM) (under the optimized reaction conditions)
showed only a small increase in absorbance at 420 nm demonstrating the catalytic efficiency of
the supported catalyst and the fact that the pyrogallol oxidation is exclusively due to the complex
PS-[VIVO(sal-dahp)], as there is no leaching of free vanadyl species (as shown by ICP-MS) [see
Fig. SI-6]. However, it should be pointed out that at pH 7 the aqueous vanadium speciation
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–
involves the formation of hydroxide species, such as (VO)2(OH)5 , and particularly precipitation
of insoluble VO(OH)2, which can account for the poor activity of VOSO4.
For better understanding the peroxidase mimetic activity of PS-[VIVO(sal-dahp)] kinetic
parameters such as Vmax, KM, Kcat were determined (where Vmax is the maximum rate, KM is the
Michaelis-Menten constant and Kcat is the turn over frequency). For this purpose, several
o
experiments were conducted in phosphate buffer of pH 7 (1 M) at 25 C by varying the
concentration of pyrogallol from 0.2 mM to 1.2 mM while H2O2 and catalyst concentrations were
kept constant in all experiments (Fig. 12(A)). The reactions were carried out for 5 minutes and
initial reaction rates were calculated by measuring the absorbance at 420 nm in kinetic mode.
According to typical enzymatic kinetic assay, the reaction rates were fitted to the Michaelis-
Menten Eq. (1):
(1)
Where Vi is the initial reaction rate, Vmax is the maximum reaction rate, [S] is the substrate
concentration and KM is the Michaelis-Menten constant.
It was observed that for catalyst PS-[VIVO(sal-dahp)] Vi follows saturation kinetics with
respect to substrate concentration [S]. The rate of reaction is first-order for low substrate
concentrations [S] i.e. a linear plot is obtained for Vi versus [S]. At sufficient high [S], the catalyst
becomes saturated with the substrate and the rate of reaction is zero-order. Thus, PS-[VIVO(sal-
dahp)] follows a Michaelis-Menten behavior towards pyrogallol.
The Lineweaver-Burk plot was employed to calculate the kinetic parameters (Fig. 12(B)).
The Vmax obtained was 9.2×10–1 µM min–1 and the turn over frequency (kcat) determined was
3.40×105 min–1, using Eq. (2).
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….(2)
<<Figure 12>>
Thus, we can conclude that PS-[VIVO(sal-dahp)] is a suitable supported catalyst to
perform the oxidation of pyrogallol in biological conditions (pH 7, aqueous medium), mimicking
peroxidase activity and therefore can be employed to perform environment friendly
heterogeneous catalytic process.
3.9. Reactivity of [VIVO(Hsal-dahp)(DMSO)] towards H2O2 and possible reaction mechanism
The stability of complex [VIVO(Hsal-dahp)(DMSO)] was evaluated by suspending it in a
mixture of acetonitrile (5 mL) and pH 8 buffer (3 mL). After 2 h under stirring the complex was
recovered, dried and its IR spectrum recorded. No significant changes were observed by
comparing its spectrum to that of the fresh complex, evidencing the complex’s stability. Both
spectra are included in supplemental information (Fig. SI-5).
The reactivity of [VIVO(Hsal-dahp)(DMSO)] towards H2O2 was studied by UV-vis
absorption and 51V NMR spectroscopies. The spectral changes in the absorption spectra of
[VIVO(Hsal-dahp)(DMSO)] (1) during the drop wise addition of a H2O2 solution (50 mM) shows
the band at 361 nm slowly disappearing. Simultaneously, two new bands appear at 310 and 398
nm, along with two isosbestic points at 340 and 395 nm (see supplemental information, Fig. SI-
3). In order to observe changes in the d–d bands, higher concentration was required and therefore
DMF was used. The drop wise addition of H2O2 (50 mM) to [VIVO(Hsal-dahp)(DMSO)] in DMF
(5 mL of ca. 1.75 × 10–3 M solution) resulted in the slow but complete disappearance of both d-d
bands at 488 and 748 nm; Fig. 13. These spectral changes indicate oxidation of V(IV) center to
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V(V) and plausible generation of monooxidoperoxidovanadium(V) species [VVO(O2)(Hsal-
dahp)(DMSO)]–. Now, addition of 3 drops of 0.25×10–1 M pyrogallol solution to the above
mixture resulted in the appearance of both d–d bands (Fig. 13) confirming the regeneration of
original oxidovanadium(IV) complex.
<<Figure 13>>
These observations were corroborated by 51V NMR experiments, which showed that after
addition of H2O2 (22 l, 50 mM) to 0.5 ml of the complex solution in DMF (1.75×10–3 M) two
new peaks appear at −540 ppm and −600 ppm (Fig. 14). The peak at −540 ppm is in the expected
range for cis-dioxovanadate complexes with N,O-ligands, [31,32,33] and the one at −600 ppm in
the expected range for monooxidoperoxidovanadium(V) species [31,33]. The cis-dioxido species
is probably a dimer of the type [(VVO2)2(Hsal-dahp)], for which resonances in the same chemical
shift region have been observed [31]. Also, the 51V NMR spectrum of a solution containing only
the complex, which was left under air for 24 h showed the presence of the same peak (see SI-4).
With incremental additions of H2O2 both peaks increase. Addition of 20 L of pyrogallol
solution (0.25×10–1 M) to a solution containing the [VIVO(Hsal-dahp)(DMSO)] complex
(1.75×10–3 M) and excess of H2O2 resulted in the disappearance of the resonance at −540 ppm,
an increase in the intensity of the resonance at −600 ppm and the appearance of a new peak at
−622 ppm (Fig. 14). These observations suggest the regeneration of the [VIVO(Hsal-
dahp)(DMSO)] complex and the partial formation of an inorganic peroxovanadate complex, in
agreement with the UV-Vis absorption spectra.
<<Figure 14>>
During the catalytic cycle the vanadium center is in contact with either H2O2 followed by
pyrogallol or vice-versa. Therefore, the reactivity of [VIVO(Hsal-dahp)(DMSO)] (1) with H2O2
19

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in the presence of pyrogallol was also studied. Changes observed by UV-Vis spectroscopy are
shown in Fig. 15. The concentration of complex 1 in acetonitrile was selected in order to have
the 488 nm band clearly visible. Thus, drop wise addition of pyrogallol (0.25×10–1 M in
acetonitrile) to complex 1 (0.27×10–3 M, 13 ml) resulted in the generation of a new band at ca.
670 nm. Simultaneously, the intensity of the 488 nm band gradually decreased and finally
disappeared showing the formation of a new species, probably involving coordination of
pyrogallol to the vanadium(IV) center. At this stage, the drop wise addition of a solution of H2O2
(50 mM H2O2 solution in acetonitrile) to the above mixture slowly regenerated the d–d band at
488 nm along with the flattening of the newly generated band at ca.670 nm band (Fig. 15).
Coordination of pyrogallol to the vanadium center was further corroborated by an ESI-
MS study in which pyrogallol was added to [VIVO(Hsal-dahp)(DMSO)] (1) in acetonitrile. The
peak at m/z = 566 is assignable to the species [VIVO(Hsal-dahp)(PG)K2H]+. The respective ESI-
MS spectrum is included in Fig. SI–7.
The lower solubility of complex 1 in acetonitrile precluded its study by 51V NMR in this
solvent. However, the study carried out with DMF in which the H2O2 solution was added to a
solution containing [VIVO(Hsal-dahp)(DMSO)] (1.75×10–3 M) and pyrogallol (0.3 mol
equivalents), the only resonance observed was the one at −600 ppm, confirming the regeneration
of VIVO complex, in the catalytic cycle.
<<Figure 15>>
All these observations suggest the detachment of pyrogallol from the vanadium center
followed by its oxidation to purpurogallin after addition of H2O2 and regeneration of the VIV
species. These observations suggest that sequential addition of substrate and oxidant do not play
any specific role and the transfer of oxygen to pyrogallol takes place from peroxidovanadium(V)
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species. Further, the metal center in the catalyst restores its oxidation state after the catalytic
reaction.
51V NMR spectra measured 72 h after dissolution of [VIVO(Hsal-dahp)(DMSO)]
(1.75×10–3 M) in DMF showed the presence of the −540 ppm peak, confirming the assignment of
this resonance to the oxidation product, the dimeric cis-dioxido complex: [(VVO2)2(Hsal-dahp)]
(see supplemental information, Fig. SI–4). For solutions containing H2O2 and pyrogallol the
same resonance was present in the NMR spectra, as well as resonances at higher field (−525 and
−530 ppm) corresponding probably to decomposition products. The large bandwidth of the
resonances suggests the presence of organic moieties around the vanadium atom.
Peroxidovanadate species usually decompose rapidly with time and its resonance appears at
lower field.
Based on all these mechanistic studies, Fig. 16 has been proposed for the catalytic cycle
involved in the oxidation of pyrogallol in aqueous medium by [VIVO(Hsal-dahp)(DMSO)].
<<Figure 16>>
4. Conclusions
A mononuclear complex [VIVO(Hsal-dahp)(DMSO)] has been prepared and structurally
characterized. The polymer-supported complex PS-[VIVO(sal-dahp)] was synthesized by reacting
[VOIV(Hsal-dahp)(DMSO)] with chloromethylated polystyrene cross-linked with 5%
divinylbenzene in DMF. The supported complex was successfully applied as peroxidase mimic
in aqueous medium. The oxidation of pyrogallol to purpurogallin has been achieved under mild
reaction conditions at pH 7 buffered solution. Sequential addition of substrate and oxidant do not
play any specific role during the catalytic reaction and oxygen transfer to pyrogallol takes place
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from a monoperoxidovanadium(V) species, identified in the 51V NMR studies. The catalytic
metal center restores its oxidation state after the catalytic reaction. Catalyst PS-[VIVO(sal-dahp)]
showed good stability over a wide pH range. Its high activity at pH 7 and recyclability without
considerable decrease in activity support its application as catalyst for greener processes in
industry.
Acknowledgements
Authors (MRM and NC) are thankful to the Council of Scientific and Industrial Research,
New Delhi, India for financial support. I. Correia thanks Fundação para a Ciência e Tecnologia
for Investigador FCT contract and UID/QUI/00100/2013.
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Tables
Table 1
Crystal data and structure refinement for [VIVO(sal-dahp)(DMSO)]·H2O (1·H2O).
1·H2O
Formula
Formula weight
T, K
C19H24N2O6SV
459.40
100(2)
Wavelength, Å
Crystal system
Space group
a/Å
0.71073
Orthorhombic
Pnma
16.9139(5)
15.8319(4)
7.3289(2)
1962.53(9)
4
b/Å
c/Å
V/Å3
Z
F000
956
Dcalc/g cm-3
/mm-1
1.555
0.652
2.73 to 28.32
/ (º)
Rint
0.0376
Crystal size/ mm3
Goodness-of-fit on F2
R1[I>2(I)] a
0.22 × 0.19 × 0.15
1.063
0.0284
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Rw (all data) b
Largest differences peak and hole (eÅ-3)
0.0780
0.412 and –0.368
2
2 2
aR1 = Fo - Fc/Fo. bRw(F2) = [w(Fo -Fc2)2]/[w(Fo ) ]1/2
Table 2
Selected bond lengths [Å] and angles [°] for [VIVO(Hsal-dahp)(DMSO)]·H2O (1·H2O).
Bond lengths
V(1)-O(1)
V(1)-O(2)
V(1)-N(1)
V(1)-O(1M)
N(1)-C(7)
N(1)-C(8)
1·H2O
1.9571(10)
1.6025(15)
2.1064(12)
2.3177(14)
1.280(2)
1.4647(18)
Bond angles
O(2)-V(1)-O(1)
O(2)-V(1)-N(1)
O(1)-V(1)-N(1)
O(2)-V(1)-O(1M)
O(1)-V(1)-O(1M)
N(1)-V(1)-O(1M)
1·H2O
104.55(5)
95.31(5)
87.89(4)
167.21(6)
84.78(4)
76.03(4)
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Table 3
Hydrogen bonds parameters for [VIVO(Hsal-dahp)(DMSO)]·H2O (1·H2O).
D-H...A
d(D-H)
0.88(4)
0.81(5)
0.81(5)
0.80(3)
d(H...A)
1.91(4)
2.51(4)
2.51(4)
2.11(3)
d(D...A)
2.793(2)
3.195(2)
3.195(2)
2.902(2)
<(DHA)
175(3)
143(2)
143(2)
172(3)
O(1W)-H(2WA)...O(3)#2
O(1W)-H(1WA)...O(1)#3
O(1W)-H(1WA)...O(1)#4
O(3)-H(3O)...O(1M)
Symmetry transformations used to generate equivalent atoms:
#1 x,-y+1/2,z #2 x+1/2,y,-z+5/2 #3 x,-y+1/2,z+1
#4 x,y,z+1
Table 4
Selected IR (cm–1) and electronic absorption spectral data (nm) of ligand and complexes.
Compounds
ν(OH)
ν(C=N)
ν(V=O)
λmax/nm (ɛ/M–1 cm–1)
H3sal-dahp (I)
3390 (br)a
1607
-
215 (4.4×104), 256 (2.3×104), 282
(0.6×104), 316 (0.8×104), 402 (2.2×103) b
232 (4.4×104), 270 (2.2×104), 361
(8.2×103) b
[VIVO(Hsal-dahp)(DMSO)] (1)
3386
1597
1604
950
965
488 (98), 748 (7) c
PS-[VIVO(sal-dahp)] (2)
3400 (br) a
280, 386, 515 d
a br = broad; b in MeOH, c in DMF; d in Nujol
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Table 5
Spin Hamiltonian parameters obtained from simulation of the EPR spectra of complexes 1 and 2.
Complex
[VIVO(Hsal-dahp)(DMSO)] (1) 1.983
PS-[VIVO(sal-dahp)] (2)
g┴
A┴×104 cm–1
53.9
g//
A//×104 cm–1
158.9
1.956
1.951
1.981
57.3
160.9
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Figure legends/ Scheme captions
Fig. 1. ORTEP representation of [VIVO(Hsal-dahp)(DMSO)]H2O (1H2O) showing the atom
labeling scheme. The ligand coordinates through the two imine-N and two phenolate-O atoms,
with the hydroxyl-O atom protonated and not taking part in coordination. The thermal ellipsoids
are drawn at 30% probability level.
Fig. 2. ORTEP representation of the crystal packing in [VIVO(Hsal-dahp)(DMSO)]H2O (1H2O)
evidencing the - interactions between phenol rings and C=N bonds of neighbour molecules
Fig. 3. Electronic absorption spectra of (a): H3sal-dahp (I) and [VIVO(Hsal-dahp)(DMSO)] (1)
recorded in methanol. Inset shows spectrum of 1 in DMF in the visible region; (b): PS-
[VOIV(sal-dahp)] (2) recorded in Nujol®.
Fig. 4. 1st derivative X-band EPR spectra of [VOIV(Hsal-dahp)(DMSO)] (1) (a) recorded from
DMF solution at 77 K, and of PS-[VOIV(sal-dahp)] (2) (b) recorded from neat solid at room
temperature.
Fig. 5. Field emission scanning electron micrographs of PS-Cl (a) and PS-[VIVO(sal-dahp)] (b)
and the energy dispersive x-ray analysis graphs of PS-Cl (a’) and PS-[VIVO(sal-dahp)] (b’).
29