Skip to main content
NCBI home page
3 Biotech logoLink to 3 Biotech
. 2021 Jan 8;11(2):39. doi: 10.1007/s13205-020-02622-6

3-Methyl-2-benzothiazolinone hydrazone and 3-dimethylamino benzoic acid as substrates for the development of polyphenoloxidase and phenoloxidase activity by zymograms

Y García-Esquivel 1, Y Mercado-Flores 1, M A Anducho-Reyes 1, J Álvarez-Cervantes 1, E Aguirre-von Wobeser 2, A I Marina-Ramírez 3, A Téllez-Jurado 1,✉
PMCID: PMC7794263  PMID: 33479594

Abstract

In the present study, a sequential staining process of polyphenoloxidase and phenoloxidase enzymes was designed by the zymography technique. As a first step, electrophoresis was carried out under native conditions, and later, first staining was carried out with a revealing solution of 3-methyl-2-benzothiazoline hydrazone (MBTH)—3-dimethylamino benzoic acid (DMAB) that allowed the visualization of polyphenoloxidase enzymes, and later and using the same gel, we proceeded to the differential staining of phenoloxidase, adding a solution of H2O2. The technique was standardized using commercial enzymes of laccase (T. versicolor) and horseradish. The technique was used to identify polyphenoloxidases (laccases) and phenoloxidases (lignin peroxidase) of crude extracts obtained from the growth of the basidiomycete Lentinus strigosus on Pinus radiata. The technique showed great sensitivity to detect the different enzymatic activities (1.56 Activity Unit/mL minimum) in the same gel without interference between the enzymes and the solutions used. On the other hand, the efficiency of the technique was compared with the substrates that are commonly used for the detection of this type of activities such as 2,2′-Azino-bis(3-ethylbenzothiazoline-6-sulfonic acid) diammonium salt (ABTS) and guaiacol, observing greater sensitivity and minimal interference, so that the present method will allow in the same gel, and visualize polyphenoloxidase and phenoloxidase activities simultaneously facilitating expression studies.

Keywords: DMAB, Isoenzymes, Laccase, MBTH, Phenoloxidase, Zymography

Introduction

Laccases and peroxidases are widely distributed in plants and white-rot fungi; these enzymes are found as isoenzymatic systems that have been of great importance in genetic and physiological studies of different microorganisms (Janusz et al. 2013; Francoz et al. 2015). These enzymes catalyze oxidative reactions on phenolic, non-phenolic compounds, and similar molecules. The laccase catalytic cycle involves O2 as the final electron acceptor, where the substrate is oxidized by reducing O2 to H2O. Substrate + O2 → Oxidized substrate + H2O (Chen et al. 2012). Peroxidases carry out their catalytic cycle through the use of H2O2. It is made up of three consecutive reactions: (a) reaction of the active site of the enzyme with H2O2 reducing it to H20; this produces the oxidation of the iron protein, resulting in an intermediate compound named “Compound I” [Fe3+  + H2O2 → Fe4+  = OR′ (Compound I) + H2O]. (b) The protein called “Compound I” is reduced by a substrate molecule by donating an e−, giving rise to a radical of the substrate, and forming “Compound II” [Fe4+  = OR´ + Substrate → Fe4+  = OR′(Compound II) + Oxidized Substrate]. (c) Compound II is reduced by a donation of a second reducing substrate molecule; thus, the enzyme returns to its native state containing Fe3+ [Fe4+  = OR + Substrate → Fe3+  + H2O + Oxidized substrate] (Wong 2009). Given the above, laccases and peroxidases and general polyphenoloxidases and phenoloxidases act on a wide spectrum of compounds, so in determining and quantifying enzyme activity using spectrophotometric methods, it is essential to provide the appropriate conditions to favor the catalytic cycle of each of these enzymes as well as providing the appropriate substrates to identify each of the enzymatic activities studied.

The zymography technique is common to identify polyphenoloxidase isoenzymes such as laccase. Depending on the substrate used, these enzymes can generate colorful compounds resulting from their catalytic activity. In this way, substrates such as ABTS (Karp et al. 2012; Afreen et al. 2017; Khozani et al. 2020), catechol (Wang et al. 2016), and 2, 6-dimethoxyphenol (Bertrand et al. 2015) have been used; Iracheta-Cárdenas et al. 2016), guaiacol (Vantamuri and Kaliwal 2016),and O-tolidine (Kumar et al. 2017) mainly. On the other hand, for the visualization of peroxidases guaiacol (Wilkesman et al. 2014; Oliveira et al. 2017), O-dianisidine (Bouacem et al. 2018; Oloketuyi et al. 2020), TMB, ABTS have been used as substrates (Oloketuyi et al. 2020), O-toluidine (Jain et al. 2020), 4-Cl-1-Naphthol (Cilerdzic et al. 2017), and DMAB-MBTH (Zerva et al. 2017). No studies have been described that allow the visualization of phenoloxidase and polyphenoloxidase activities simultaneously, much less, using the same electrophoresis gel.

The present method is based on the oxidative coupling of MBTH and DMAB, a common reagent to determine glucose in a glucose oxidase–peroxidase system (Ngo and Lenhoff 1980). Furthermore, these substrates have been used to determine manganese peroxidase activity, which is a susceptible technique for detecting this enzyme spectrophotometrically (Castillo et al. 1994). The method described below has been proposed and tested only for peroxidase-coupled reactions using spectrophotometric techniques. Still, it has not been studied in detail in the determination of laccases and peroxidases using zymograms. This work aims to propose a method for the visualization of isoenzymes by the zymography technique that is based on the catalysis of the enzyme trapped in the acrylamide matrix on the substrate that diffuses through it. The oxidative coupled reaction of MBTH and DMAB with laccase and peroxidase is described sequentially; first, laccase activity is revealed in the substrate presence. Then, after the addition of H2O2, peroxidase or phenoloxidase activity is revealed. The described method was applied to extracts of Lentinus strigosus fungus grown on wood and filter paper, revealing differences in the zymographic profile of these enzymes.

Materials and methods

Zymography assays

For the zymography tests, acrylamide–bis acrylamide gels were prepared for native conditions (without SDS and b-mercaptoethanol) as described by Laemmli (1970). Two commercial enzymes were used (Trametes versicolor laccase and horseradish peroxidase) (Sigma-Aldrich). 100 AU (Activity Unit) of each enzyme were taken and resuspended in phosphate buffer (pH 6.5) and acetate buffer (pH 5.5) for the enzymes laccase and peroxidase, respectively. The enzymes were injected into an electrophoresis gel consisting of a concentration gel (5%) and a separation gel (12%). The electrophoresis was carried out in a Mini Vertical Gel System EC 120 kit (Bio-Rad). Electrophoresis was carried out at a constant 100 V for 90 min at 4 °C. Once the electrophoresis stage was finished, the gels were washed with deionized water and were subjected to the corresponding staining processes.

Preparation of the DMAB–MBTH developer solution in acetate buffer and phosphate buffer

The DMAB–MBTH system was prepared according to Ngo and Lenhoff (1980) with some modifications. The complex was dissolved in two different buffers to measure the activities of the commercial enzymes tested (Lcc and horseradish). To measure Lcc activity, the complex was dissolved in phosphate buffer (100 mM, pH 6.5). To measure peroxidase activity, an acetate buffer (50 mM, pH 5.5) was used to solubilize the DMAB–MBTH complex. In both cases, the complex final concentration was 2.5 mM for DMAB and 1 mM for MBTH.

Zymogram using DMAB–MBTH for laccase and peroxidase revelation

After electrophoresis, each gel was immersed in distilled water and washed for 5 min three times. Subsequently, 100 mL of phosphate buffer (100 mM, pH 6.5) or acetate buffer (50 mM, pH 5.5) was added depending on the enzyme system to be revealed. The gels were kept stirred for 20 min at 50 rpm. After this time, each gel was immersed in the corresponding developer solution DMAB–MBTH in phosphate buffer and DMAB–MBTH in acetate buffer. In both cases, the laccase activity is the first to appear due to the purple band’s presence in the acrylamide gel. After developing the Lcc activity, the polyacrylamide matrix was kept under stirring (50 rpm) in the same solution for 20 min. After this time, it proceeded to specific staining for peroxidase by adding to the developer solution of each gel, H2O2, at different concentrations (10–20 mM) until the appearance of purple bands. The developer solution was removed to stop the reaction, and three washes were carried out with distilled water.

Laccase activity by means of the MBTH–DMAB reaction

Lcc activity was assayed on a spectrophotometer (Thermo Scientific), using MBTH and DMAB as substrates. The substrates solution consisted of 0.5 mL of 0.033 mM MBTH solution in 50 mM acetates buffer, pH 6; 0.5 mL of 1.6 mM DMAB solution in 50 mM acetate buffer, pH 6 and 0.5 mL of enzyme extract. For the blank, water was added instead of enzyme extract. The reaction was incubated at room temperature for 30 min, and the absorbance at 590 nm was measured. Under the conditions tested, the enzymatic activity was defined as the amount of enzyme necessary to catalyze the reaction of 1 µmol of product per minute per mL, using a molar extinction coefficient of 43,600 M/cm.

Determination of Lcc and peroxidase activities

To determine the enzymatic activity of Lcc in liquid, two methods were used, the first with ABTS according to the method reported by Li et al. (1998) and guaiacol according to the procedure described by Hemeda and Klein (1991).

Propagation and conservation of L. strigosus

The Lentinus strigosus strain used in this work was isolated from the Huasteca hidalguense (29º 48′ 06.35″ N, 98º 44′ 45.97″ W), Mexico, and later identified by molecular methods (data not shown). The L. strigosus strain was propagated on PDA (Potato-Dextrose Agar) plates added to 1% with yeast extract. They were incubated for 7 days at 28 °C. For their conservation, the plates invaded by the fungus were stored at 4 °C until use.

Preinoculum preparation

To prepare the preinoculum, was used the method described by Quintanar et al. (2012) with some modifications. Two plates invaded by L. strigosus were added to an Erlenmeyer flask for every 100 mL of sterile water, which were cut into squares of approximately 1 cm2. The flask was kept at 150 rpm at 28 °C for 24 h to facilitate detachment of the mycelium from the agar. The supernatant with the mycelium in suspension was used for the inoculum of the corresponding media.

Solid-state fermentation in Pinus radiata residues and filter paper

Two tests were carried out, one with P. radiata chips using the method described by Quintanar et al. (2012) and the other with filter paper (Navarro et al. 2014). For the first test, 2 g of chips of P. radiata with particle size 12 (0.661 in) were sterilized at 15 lb pressure for 20 min, and then inoculated with the L. strigosus preinoculum adjusting humidity to 75%. In the second test, filter paper (previously sterilized) with particle size 12 (0.661 in) was inoculated with the L. strigosus preinoculum, adjusting to a humidity percentage of 75%. For both cases, sterile plastic trays of 7 cm of diameter were used. The inoculated substrates were kept at 28 °C for 9 days. All tests were performed in triplicate.

Obtaining of crude enzymatic extracts

The enzyme extract was obtained on days 3, 5, and 9. For each gram of substrate, 10 mL of acetate buffer (50 mM, pH 6) was added. The fermentation was immersed in the buffer for 30 min, and with stirring at 150 rpm. Subsequently, the supernatant was centrifuged at 2500×g for 40 min. The enzymatic extract was subjected to isoelectric focusing using a Rotofor Cell kit (Bio-Rad), a solution of ampholytes in a pH range of 3–10 was used. To the fractions obtained from the extracts of culture in wood and filter paper, Lcc activity was determined, using ABTS as substrate. The fractions with Lcc activity were*** subjected to electrophoresis under native conditions to subsequently carry out the stain with the developer solution MBTH–DMAB in phosphate buffer. The wood extract fractions were subjected to electrophoresis for determining Lcc activity with MBTH–DMAB, lignin peroxidase using MBTH–DMAB, and MnP using phenol red.

Determination of Lcc activity in solution with ABTS and MBTH–DMAB

Lcc activity was tested using ABTS (5 mM) in acetate buffer (50 mM, pH 6) as substrate, following the protocol described by More et al. (2011). The Enzyme Activity Unit (AU) was defined as the amount of enzyme needed to oxidize 1 μmol ABTS per minute, using a molar extinction coefficient of 36,000 M/cm. The same fractions were tested using MBTH–DMAB as substrate.

Determination of lignin peroxidase (LiP) activity in solution

The enzymatic activity of LiP was tested in a spectrophotometer (Thermo Scientific), following the protocol described by Ngo and Lenhoff (1980) with the following modifications: the reaction mixture (1510 µL) consisted of 500 µL of DMAB (5 mM) in phosphate buffer (50 mM, pH 6.5), 500 µL of MBTH (0.1 mM), in phosphate buffer (50 mM, pH 6.5), and 500 µL of crude enzyme extract. The reaction was initiated by the addition of 10 µL of H202 (4 mM). The oxidation reaction was measured by monitoring the increase in absorbance at 590 nm. The initial absorbance was determined by adding H202 and the final absorbance after 30 min. The AU was defined as the amount of enzyme needed to catalyze the reaction of 1 µmol of product per minute per mL, using a molar extinction coefficient of 43,600 M/cm.

Determination of manganese peroxidase (MnP) activity in solution

MnP activity was determined using phenol red as substrate, by spectrophotometry (Thermo Scientific), following the protocol described by Kuwahara et al. (1984). The AU was defined as the amount of enzyme needed to oxidize 1 μmol of Phenol Red per minute.

Results

Development of Lcc and peroxidase enzymes with DMAB–MBTH

The study of enzyme complexes using zymograms is based on the fact that when the gel is immersed in the developer solution, the substrates and other reagents diffuse through the acrylamide matrix, where they are transformed by the catalytic action of the enzyme that is analyzed and that it is retained in the polymer matrix.

To visualize the commercial laccase activity of T. versicolor with the MBTH–DMAB complex in gel, different dilutions of the enzyme were made, starting with 100 AU/mL to estimate the detection limit of the system. The system being a visual technique, the concentration of enzyme applied per well of the gel was not considered. It was observed that the detection limit is approximately 1.56 AU/mL, where unstable bands of activity are generated. With this enzymatic activity, the bands generated disappeared after 10 min. Therefore, to maintain a stable band, the enzymatic activity must be around 6.25 AU/mL.

Figures 1 and 2 show the zymogram for the commercial enzymes T. versicolor Lcc and horseradish peroxidase revealed after incubation in MBTH–DMAB solution in phosphate buffer (Fig. 1) and acetate buffer (Fig. 2). In both cases, as part of the Lcc enzyme catalysis, purple bands were observed after 10 min of incubation. So far, the coupled oxidation of MBTH and DMAB has been reported exclusively for peroxidase reactions and under chemical conditions; however, the present study demonstrates the ability of Lcc to oxidize MBTH in the absence of H2O2 and Mn+. This non-phenolic type compound, like other substrates, is not specific for peroxidases; given its MBTH structure, it can be oxidized by the Lcc enzyme. In the oxidative coupling reaction, in the first stay, MBTH loses two electrons and a proton to form an electrophilic intermediate, which is an oxidized form of this compound. In a second electrophilic substitution reaction, this intermediate reacts with DMAB forming a colored complex. Therefore, without the presence of this intermediate, there is no color formation. The positive reaction in samples with Lcc activity allows the oxidation of MBTH and can be attributed to this enzyme. This is the first study that evidences MBTH and DMAB as a substrate in the determination of Lcc enzymes.

Fig. 1.

Fig. 1

Open in a new tab

Sequential staining of an electrophoretic gel to detect laccase and peroxidase activity in phosphate buffer. a Detection of commercial laccase of T. versicolor revealed with DMAB–MBTH solution. b Staining specific for commercial horseradish peroxidase, started by adding H2O2 to the developer solution at a final concentration of 13 mM once the laccase activity was visualized

Fig. 2.

Fig. 2

Open in a new tab

Sequential staining of an electrophoretic gel to detect laccase and peroxidase activity in acetate buffer. a Detection of commercial laccase of T. versicolor revealed with DMAB–MBTH solution. b Specific staining of commercial horseradish peroxidase initiated by the addition of H2O2 to a final concentration of 13 mM once laccase activity was visualized

For peroxidases, the assay is based on the oxidative coupling of MBTH and DMAB in the presence of H202; this compound was added at different concentrations, determining that at a final concentration of 13 mM, a positive reaction is observed (Figs. 1b, 2). Under the conditions tested, substrate oxidation for peroxidase action is immediate, generating well-defined deep purple bands. Furthermore, it was observed that during the reaction, the color development is stable.

Considering that in plants and fungi, a mixture of laccases and peroxidases can be found in the extract to be analyzed; it is recommended that the Lcc activity be revealed in the first instance. After the enzyme positive reaction, the gel should be left to stand for 10–20 min. This step is essential to prevent other enzymes from interfering with the peroxidase activity present in the extracts, which initiate their reaction by adding H2O2.

Revealed in ABTS compared to MBTH–DMAB

Oxidation of ABTS by the Lcc produces bright green bands (Fig. 3). In comparing the use of ABTS and MBTH–DMAB, it is evident that both substrates allow the detection of well-defined bands (Figs. 1a, 2, 3). These results demonstrate that, like other compounds used in the determination of Lcc-type enzymes, MBTH–DMAB is a substrate that, after being oxidized, generates a coloration that allows its use in zymography techniques.

Fig. 3.

Fig. 3

Open in a new tab

Zymogram of the commercial laccase of T. versicolor using ABTS dissolved in phosphate buffer as a developer solution

Revealed in guaiacol compared to MBTH–DMAB

Guaiacol has been used to determine peroxidases by spectrophotometric methods, and like other compounds, it can be oxidized by Lcc. In this case, the Lcc activity was revealed after 20 min of incubation; it should be noted that compared to what was observed in ABTS and MBTH–DMAB, the reaction time and appearance of the Lcc bands increased, observing a faint color coloration reddish (Fig. 4a). Peroxidase activity was revealed after the addition of H202 at a final concentration of 10 mM. However, despite being the same enzyme concentration, a marked effect is observed in the color and intensity of the bands formed as a consequence of the substrate type. It is evident that, when using guaiacol as a peroxidase developer substrate, no defined bands were observed in the acrylamide matrix (Fig. 4b), representing a disadvantage in the elaboration of zymograms for the determination of these enzymes, in particular, indicating that the use of Guaiacol is less sensitive than the other substrates.

Fig. 4.

Fig. 4

Open in a new tab

Sequential staining of an electroporesis gel to visualize laccase and peroxidase activity. a Zymogram of the commercial laccase of T. versicolor using guaiacol dissolved in phosphate buffer as a developer solution. b Differential staining of the electrophoresis gel to visualize horseradish peroxidase activity by adding H2O2 as a developer solution once laccase activity was visualized

Zymographic profile of Lentinus strigosus during wood and filter paper degradation

The assay based on spectrophotometric and electrophoretic methods for determining Lcc activity from L. strigosus extracts cultivated in P. radiata residues was measured on days 3, 5, and 9 of culture. Table 1 shows the results for Lcc and LiP activity using ABTS and MBTH–DMAB as substrate. Due to ABTS oxidation, color formation confirms Lcc activity; the fractions used using this substrate have activity values that range between 41.20 and 504.81 AU/mL. These same sample fractions present activity values between 1.18 and 3.11 AU/mL used MBTH–DMAB as substrate. Se observa una disminución considerable en la detección de la actividad enzimática.

Table 1.

Lcc and LiP activity for fractions obtained from the L. strigosus culture on P. radiata residues

Day Fraction ABTS Guaiacol MBTH–DMAB MBTH–DMAB
Laccase (AU/mL) Laccase (AU/mL) Laccase (AU/mL) LiP (AU/mL)
3 10 310.370 ± 1.786 152.032 ± 2.421 1.977 ± 0.064 2.185 ± 0.092
11 504.815 ± 5.702 216.650 ± 1.281 2.842 ± 0.073 2.863 ± 0.106
12 303.148 ± 5.675 98.672 ± 1.204 3.112 ± 0.342 3.827 ± 0.213
5 10 132.315 ± 1.530 32.784 ± 1.119 2.658 ± 0.106 0.636 ± 0.021
11 460.463 ± 1.370 104.230 ± 2.534 2.822 ± 0.056 1.118 ± 0.052
12 99.444 ± 2.546 21.946 ± 1.439 1.181 ± 0.109 2.086 ± 0.244
9 12 41.204 ± 1.580 16.342 ± 1.329 1.889 ± 0.074 1.952 ± 0.044
13 194.352 ± 3.195 32.409 ± 2.438 2.419 ± 0.074 2.434 ± 0.123
18 66.667 ± 2.406 11.305 ± 1.097 1.899 ± 0.332 1.106 ± 0.024
Open in a new tab

The differences observed for the same type of samples are subject to the determination conditions, including the incubation time, the type of substrate, the molar extinction coefficient, and the reaction volume. The LiP activity presented values of 1.10 to 3.82 AU/mL in DMAB–MBTH. On the other hand, under the conditions tested, MnP activity was not detected. Table 2 shows the Lcc activity for the fractions obtained from filter paper culture extracts; it is observed that when using this substrate, the Lcc activity presented activity values between 4.53 and 78.88 AU/mL.

Table 2.

Lcc activity for fractions obtained from the L. strigosus culture on filter paper

Day Fraction ABTS Guaiacol
Laccase (AU/mL) Laccase (AU/mL)
3 11 9.074 ± 0.699 3.124 ± 0.050
12 8.981 ± 1.123 5.231 ± 0.342
13 78.889 ± 1.667 51.098 ± 2.341
5 11 4.259 ± 0.424 1.932 ± 0.021
12 24.444 ± 1.211 14.182 ± 0.976
14 5.833 ± 0.481 1.360 ± 001
9 12 14.630 ± 0.699 3.162 ± 0.030
13 4.537 ± 0.321 0.921 ± 0.001
Open in a new tab

A zymogram was performed from the spectrophotometrically analyzed samples to determine the enzymatic profile of L. strigosus during its growth on P. radiata and filter paper. This test was performed to differentiate between hydrolytic and lignocellulolytic activities. For the extracts obtained from the culture in P. radiata, during the staining process, bands with Lcc activity were revealed on days 3, 5, and 9, observing a different isoenzymatic profile for each day of culture. The Lcc bands began to develop after the 10 min incubation; after 15 min, all the bands corresponding to the Lcc activity presented in the zymogram were revealed (Fig. 5). In this condition, the lowest Lcc activity quantified using ABTS as substrate was 41.20 AU/mL, and the highest was 504.81 UA/mL, as it is the substrate where the highest Lcc activity was quantified. In all cases, we observed the formation of wide bands. After the positive reaction of bands with Lcc activity, the gel was kept in the same solution for an additional 10 min. After this time, 10 µL of H202 was added, revealing for day 3, an enzyme with peroxidase activity present in all the fractions analyzed for this day (Fig. 5). The peroxidase activity quantified on day 3 using MBTH–DMAB as substrate was 3 AU/mL. In the analyzed sample corresponding to day five, no bands with peroxidase activity were observed. Finally, peroxidase activity was detected on day 9, which corresponded to 1.10–2.43 AU/mL and was detected again in the corresponding gel (Fig. 5). Despite the low amount of peroxidase activity quantified spectrophotometrically, the development process favored the detection and visualization of the bands corresponding to peroxidase activity, results suggest that the applied method is sensitive to low amounts of enzyme.

Fig. 5.

Fig. 5

Open in a new tab

Visualization of laccase and peroxidase activities by the zymography technique. The different fractions separated by their pI from the crude enzyme extract obtained from the growth of L. strigosus on P. radiata residues at different times are shown. Lanes 1–3: fractions obtained after 3 days of culture. Lanes 4–6: fractions obtained after 5 days of culture. Lanes 7–9: fractions obtained after 9 days of culture

To compare the enzymatic profile of L. strigosus on days 3, 5, and 9 on filter paper, the extracts obtained were analyzed by zymography. In this case, after 15 min of incubation, the bands with Lcc activity were revealed, observing a decrease in the isoenzymes present in this condition compared to that obtained in the zymogram from culture extracts in P. radiata. It should be noted that, on filter paper, the Lcc activity using ABTS as a substrate ranged between 4.53 and 78.88 AU/mL. The zymogram for filter paper extracts is shown in Fig. 6. Again, as observed in the development of peroxidase activity, it is observed that, at low amounts of Lcc activity, detection and visualization of bands are possible, confirming the sensitivity of the staining process. Following the development methodology, the gel was kept in the same solution for an additional 10 min. Subsequently, 10 µL of H2O2 was added, keeping the gel in agitation. In this case, no bands with peroxidase activity were revealed. The staining method described in the present work demonstrates a mixture of enzymes with Lcc and peroxidase activity during the degradation of P. radiata residues. The fungus understudy secretes at least two isoenzymes with Lcc activity and a peroxidase (Fig. 6). The extracts analyzed from filter paper showed a decrease in the secreted isoenzymes, observing that, under this condition, L. strigosus only secretes enzymes with Lcc activity and no peroxidases.

Fig. 6.

Fig. 6

Open in a new tab

Visualization of laccase and peroxidase activities by the zymography technique. The different fractions separated by their pI from the crude enzyme extract obtained from the growth of L. strigosus on filter paper at different times are shown. Lanes 1–3: fractions obtained after 3 days of culture. Lanes 4–6: fractions obtained after 5 days of culture. Lanes 7–9: fractions obtained after 9 days of culture

Discussion

In the biochemical study using zymograms to analyze enzymatic systems made up of laccases and peroxidases, a key element is the staining process. These enzymes carry out oxidation reactions and can act on the same compound complicating their detection by zymography techniques. The staining system for laccases and peroxidases is based on the use of redox compounds, whose physical and chemical properties change as their oxidation state changes. Specifically, for detecting bands in a gel of activity, this characteristic must necessarily be associated with a color change.

In the determination of laccases, one of the most studied and used substrates is ABTS, due to its stability and redox chemistry are known. The colorimetric test is based on cationic radicals formation from ABTS, which exhibits a color change (Putter and Becker 1983). The oxidation of this substrate by the enzyme produces a bright green color (Fig. 3). Lcc zymograms have also been revealed with catecol 2,6-Dimethoxyphenol, P-anisidine, and O-toludine (Téllez-Téllez et al. 2012). MBTH, as other substrates are not specific for peroxidases, its use to detect the catalytic activity of laccase has already been studied. Castillo et al. (1994), assuming that laccases and peroxidases are usually present in white-rot fungi cultures, analyzed the interference of Lcc activity in a reaction mixture designed for the determination of MnP by a method spectrophotometric. These authors designed a reaction mixture using MBTH–DMAB as a substrate for the determination of MnP (80.07 mM MBTH, 0.99 mM DMAB, 0.3 mM MnSO4, 17 nM MnP, and 0.05 mM H2O2 in lactic acid–succinate buffer), reported that under the tested conditions, when using up to 1520 units of Lcc activity/mL, this activity is little detected, presenting low interference with its method to determine MnP (Castillo et al. 1994). It was observed in the present work that low concentrations of T. versicolor and L. strigosus laccase enzymes could be detected differentially from the LiP and MnP enzymes. Furthermore, it was shown that Lcc could oxidize MBTH and initiate the coupled oxidation of this compound and DMAB. Although laccases and peroxidases have different catalytic requirements, being relatively nonspecific on reducing substrates, they can oxidize aromatic compounds with high redox potential; thus, both enzymes have the ability to act on the same substrate, and MBTH is not the exception.

The quantification of peroxidases has been used as H+ donor substrates; veratryl alcohol, guaiacol, and O-dianisidine (Haemmerli et al. 1987; Suzuki et al. 2006). Even though veratryl alcohol is one of the most widely used substrates, a disadvantage is that it is oxidized to veratraldehyde, a colorless compound that does not allow the detection of bands in a zymogram. In contrast to veratrilic alcohol, guaiacol is oxidized to a red tetraguaiacol complex (Tonami et al. 2004). In general, employing spectrophotometric techniques, guaiacol has been widely used in determining these enzymes; however, there are reports in which it is shown that guaiacol can inhibit the activity of peroxidases in concentrations of 24 mM and 40 mM (Halpin et al. 1989). In the present study, the concentration of guaiacol in the developer solution was 5 mM, a lower concentration than that reported to inhibit this type of enzyme. However, after incubation of horseradish peroxidase in guaiacol solution and the addition of 10 mM H2O2, a stable coloration was not observed, avoiding the formation of well-defined bands (Fig. 4b), representing a disadvantage the use of guaiacol for this particular technique, that is based on the development of colorful bands due to the activity of enzymes trapped in a polyacrylamide matrix.

The zymograms of plant peroxidases have been revealed using 2, 3, 3′, 5, 5′-tetramethylbenzidine (TMB), ABTS, and O-diasidine as substrates (Oloketuyi et al. 2020). After evaluating these three substrates, it was reported that TMB allows the detection of bands with peroxidase activity; however, it is mentioned that the staining buffer and the solution to stop the reaction affect the physical characteristics of the gels, which change dimension and do not allow a comparison of activity bands. In addition to reporting that after treatment with sulfuric acid of samples stained with TMB, a negative effect is observed in detecting bands, so it is suggested to omit this step. It has been described that the use of ABTS in the development of peroxidase bands is less effective, given that under the same experimental conditions, eight bands of activity were identified when using TMB as a substrate. In contrast, with ABTS, only six bands were visualized (Oloketuyi et al. 2020). The use of O-dianisidine has been reported in the study of the induction of peroxidase activity in the Lulo fruit (Solanum quitoense L.) as a response to the pathogen causing actracnosis; in this case, a revealing solution of O-dianisidine (1.8 mM) in phosphate buffer (100 mM, pH 6.5) and H2O2 (1.2%) was used to reveal peroxidase bands in electrophoresis gels of crude extracts of Lulo (Higuer et al. 2009). In addition to the low solubility in water these compounds, one of the main disadvantages of O-diadisidine is that, like other compounds such as benzidine, O-toluidine, and O-toludin, which are mutagenic and carcinogenic (Liu and Gibson 1997).

In the detection of peroxidase activity by electrophoretic techniques, the products can be in the non-visible spectrum, inhibit enzyme activity, or have low sensitivity (Haemmerli et al. 1987; Halpin et al. 1989), making detection difficult of these enzymes in zymogram studies. In the determination of peroxidase enzymes using MBTH and DBAM as a substrate for spectrophotometric methods, it has been reported that peroxidase can be determined in pM amounts in solution, this being a sensitive chromogenic essay for this type of enzyme (Ngo and Lehoff 1980; Castillo et al. 1994). We observed that when using the MBTH–DMAB system as a revealing solution no interference was observed, and laccase and peroxidase activity could be visualized in the same zymogram (Figs. 1b, 2b), compared to what was observed in guaiacol as a revealing solution where a lower sensitivity is observed for peroxidase activity (Fig. 4a). The MBTH–DMAB system for peroxidases is based on the coupled oxidation of these compounds, after which a dark purple color is produced, described as an indamine dye (Ngo et al. 1980). Oxidation of MBTH by FeCl3, followed by reaction with DMAB, in methanol solution giving a deep blue color has been reported (Sawicki et al. 1961). In this case, the oxidation of MBTH was chemically generated; the same reaction mechanism is suggested during the oxidation of this compound mediated peroxidase-type enzymes (Ngo and Lehoff 1980; Castillo et al. 1994). Similarly, in the reaction of the Lcc with the MBTH–DMAB system after the catalytic action, we observed the formation of intense purple color in both cases, so the same chain reaction probably follows.

The steps of an isoenzymatic study include four stages: (a) extraction of proteins, (b) separation by electrophoresis, (c) visualization of isoenzymes using a revealing solution, and (d) analysis of results. Once the protein molecules have been separated, it is necessary to visualize them, staining or revealing a crucial step in isoenzyme detection and interpretation of results. This contribution improves the staining process of Lcc-type isoenzymes and peroxidase in zymography studies. After applying the method developed in the present work to commercial type enzymes (Lcc and horseradish peroxidase), we demonstrated the ability of Lcc to catalyze the coupled oxidation of MBTH and DMAB, and we improved the development of peroxidase enzymes by treating samples of these enzymes with the MBTH–DMAB developer solution. The results obtained for the fungal extracts analyzed to demonstrate the usefulness of this method to perform selective staining, from extracts containing a mixture of enzymes with Lcc and peroxidase activity, allowing a differential analysis associated with the catalytic requirements of each enzyme, and the location in situ of the same. On the other hand, the simultaneous study of laccases and peroxidases is favored using the same staining process; sequentially, each activity is revealed. Conditions that make this technique a sensitive and stable method for detecting this type of enzyme by zymography.

Laccases and peroxidases have been the subject of genetic and physiological studies; given their functions, a mixture of these enzymes is generally found in extracts of plants and fungi. The peroxidases and laccases present in plants participate in the formation and development of roots, lignin formation, biosynthesis of bethaline, and phenolic compounds such as phenylpropanoids (Francoz et al. 2015). Due to their dual hydroxylic and peroxidative cycles, peroxidases can produce ROS and oxidize aromatic compounds associated with the hardening or loosening of the cell wall (Berthet et al. 2012). Likewise, Lcc and peroxidases participate in defense mechanisms against pathogens or mechanical damage, electron transport, and auxins oxidation (Sawicki et al. 1961). In contrast, in basidiomycete fungi and some ascomycetes, the degradation of lignin is attributed to four main enzymes: LiP, Manganese peroxidase, Versatile peroxidase, and Lcc (Kuwahara et al. 1984).

Conditions for the staining of enzyme systems formed by laccases and peroxidases

Laccases and peroxidases are present in plants and fungi; consequently, we will generally find a mixture of both enzymes (Janusz et al. 2013; Francoz et al. 2015). Although plants and fungi perform totally different functions, it is necessary to point out that laccase and peroxidases are involved in closely related functions in each of these organisms. Given the function of laccases and peroxidases present in plants and fungi, it is necessary to implement a development method aimed at the simultaneous study of both enzymes, which allows the presence or absence of these to be related under different physiological stages. The above implies using a developer solution with a substrate for which both enzymes show affinity, with an immediate reaction that generates a stable color and is easy to visualize. The specific staining for each isoenzyme will be given by the catalytic requirements of the enzyme being analyzed. To improve staining, the development solution must be made up of substrates and soluble reagents, which improves the diffusion of these inside the gel, favoring the substrate availability for the enzyme trapped in the acrylamide matrix. The staining process must not damage the sample or alter the gel dimension to be analyzed.

Selective development for laccases and/or peroxidases in the presence of MBTH–DMAB

Considering enzymatic systems sources to be analyzed, a staining method is provided that allows the specific staining of laccases and/or peroxidases from samples containing both enzymatic activities. The staining process reacts with laccases and/or peroxidases, favoring the simultaneous study of these enzymes involved in closely related functions. In the developer solution, the MBTH and DMAB substrates are transformed by the effect of Lcc and peroxidase activity, followed by the precipitation of the dye in areas of activity, with the formation of intense purple color, a necessary characteristic in the visualization of bands for the detection of isoenzymes by zymography, showing that both enzymes have an affinity for the same substrate. The positive reaction for Lcc and peroxidase using the same developer solution reduces the expense of reagents, time, and favors the study of both enzymes. The staining process can be applied selectively or differentially, given the catalytic requirements of Lcc and peroxidase. The highest concentrations used in the present study were 5 mM for DMAB and 1 mM for MBTH, for which both compounds were soluble in pH 5.5 acetate buffer and pH 6.5 phosphate buffer. The staining process does not interfere with the gel dimension, and no negative effect was observed on the samples analyzed.

Conclusion

Laccase-like peroxidase catalyzes the oxidation reaction of MBTH, forming the intermediate that reacts with DMAB. The implemented method improves the staining and visualization process of isozymes of the Lcc and peroxidase-type analyzed by zymography, being useful in analyzing extracts containing a mixture of enzymes with Lcc and peroxidase activity. The method can detect at least 1.57 AU/mL, but a minimum of 6 AU/mL is needed to obtain stable bands. It was possible to detect a crude extract's enzymatic profile using the sequential staining proposed in this work in the same gel, representing a great advantage for studies of secretion of lignocellulosic enzyme systems.

Acknowledgements

This work was carried out thanks to the support of the National Council of Science and Technology (CONACyT) of Mexico.

Author contributions

YG-E: fomal analysis, methodology, investigation, and writing—original draft. YM-F: validation, conceptualization, and supervisión. MAA-R: software, formal analysis, and data curation. JÁ-C: visualization and supervisión. EA-vW: validation, formal analysis, and supervisión. AIM-R: conceptualisation and supervisión. AT-J: conceptualization, writing review and editing, resources, project administration, and supervision.

Compliance with ethical standards

Conflict of interest

The authors declare that they have no conflict of interest in the publication.

Ethical approval

This paper does not contain any studies with human participants or animals performed by any of the authors.

References

  1. Afreen S, Shamsi TN, Baig MA, Ahmad N, Fatima S, Qureshi MI, Hassan MI, Fatma T. A novel multicopper oxidase (laccase) from cyanobacteria: purification, characterization with potential in the decolorization of anthraquinonic dye. PLoS ONE. 2017;12(4):e0175144. doi: 10.1371/journal.pone.0175144. [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Berthet S, Thevenin J, Baratiny D, Demont-Caulet N, Debeaujon I, Bidzinski P, Lapierre C. Role of plant laccases in lignin polymerization. Adv Bot Res Academic Press. 2012;61:145–172. doi: 10.1016/B978-0-12-416023-1.00005-7. [DOI] [Google Scholar]
  3. Bertrand B, Martínez-Morales F, Tinoco-Valencia R, Rojas S, Acosta-Urdapilleta L, Trejo-Hernández MR. Biochemical and molecular characterization of laccase isoforms produced by the white-rot fungus Trametes versicolor under submerged culture conditions. J Mol Catal B Enzym. 2015;122:339–347. doi: 10.1016/j.molcatb.2015.10.009. [DOI] [Google Scholar]
  4. Bouacem K, Rekik H, Jaouadi NZ, Zenati B, Kourdali S, El Hattab M, Badis A, Annane R, Beja S, Hacene H, Bouanane-Darenfed A, Jauaadi B. Purification and characterization of two novel peroxidases from the dye-decolorizing fungus Bjerkandera adusta strain CX-9. Int J Biol Macromol. 2018;106:636–646. doi: 10.1016/j.ijbiomac.2017.08.061. [DOI] [PubMed] [Google Scholar]
  5. Castillo MD, Stenstrom J, Ander P. Determination of manganese peroxidase activity with 3-methyl-2-benzothiazolinone hydrazone and 3-(dimethylamino) benzoic acid. Anal Biochem. 1994;218(2):399–404. doi: 10.1006/abio.1994.1198. [DOI] [PubMed] [Google Scholar]
  6. Chen YR, Sarkanen S, Wang YY (2012) Lignin-degrading enzyme activities. In: Biomass conversion. Humana Press, Totowa, NJ. pp 251–268. Doi: 10.1007/978-1-61779-956-3_21 [DOI] [PubMed]
  7. Cilerdzic J, Galic M, Vukojevic J, Brceski I, Stajic M. Potential of selected fungal species to degrade wheat straw, the most abundant plant raw material in Europe. BMC Plant Biol. 2017;17(2):75–81. doi: 10.1186/s12870-017-1196-y. [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. Francoz E, Ranocha P, Nguyen-Kim H, Jamet E, Burlat V, Dunand C. Roles of cell wall peroxidases in plant development. Phytochemistry. 2015;112:15–21. doi: 10.1016/j.phytochem.2014.07.020. [DOI] [PubMed] [Google Scholar]
  9. Haemmerli SD, Schoemaker HE, Schmidt HW, Leisola MS. Oxidation of veratryl alcohol by the lignin peroxidase of Phanerochaete chrysosporium Involvement of activated oxygen. FEBS Lett. 1987;220(1):149–154. doi: 10.1016/0014-5793(87)80893-1. [DOI] [Google Scholar]
  10. Halpin B, Pressey R, Jen J, Mondy N. Purification and characterization of peroxidase isoenzymes from green peas (Pisum sativum) J Food Sci. 1989;54(3):644–649. doi: 10.1111/j.1365-2621.1989.tb04672.x. [DOI] [Google Scholar]
  11. Hemeda HM, Klein BP. Inactivation and regeneration of peroxidase activity in vegetable extracts treated with antioxidants. J Food Sci. 1991;56(1):68–71. doi: 10.1111/j.1365-2621.1991.tb07977.x. [DOI] [Google Scholar]
  12. Higuera BL, Martínez S, Caicedo O. Inducción de actividad peroxidasa y de fenoles totales como respuesta del fruto de lulo (Solanum quitoense l.) al patógeno causal de la antracnosis. Rev Col Quim. 2009;38(1):25–42. [Google Scholar]
  13. Iracheta-Cárdenas MM, Rocha-Peña MA, Galán-Wong LJ, Arévalo-Niño K, Tovar-Herrera OE. A Pycnoporus sanguineus laccase for denim bleaching and its comparison with an enzymatic commercial formulation. J Environ Manage. 2016;177:93–100. doi: 10.1016/j.jenvman.2016.04.008. [DOI] [PubMed] [Google Scholar]
  14. Jain KK, Kumar A, Shankar A, Pandey D, Chaudhary B, Sharma KK. De novo transcriptome assembly and protein profiling of copper-induced lignocellulolytic fungus Ganoderma lucidum MDU-7 reveals genes involved in lignocellulose degradation and terpenoid biosynthetic pathways. Genomics. 2020;112(1):84–198. doi: 10.1016/j.ygeno.2019.01.012. [DOI] [PubMed] [Google Scholar]
  15. Janusz G, Kucharzyk KH, Pawlik A, Staszczak M, Paszczynski AJ. Fungal laccase, manganese peroxidase and lignin peroxidase: gene expression and regulation. Enzyme Microb Technol. 2013;52(1):1–12. doi: 10.1016/j.enzmictec.2012.10.003. [DOI] [PubMed] [Google Scholar]
  16. Karp SG, Faraco V, Amore A, Birolo L, Giangrande C, Soccol VT, Pandey A, Soccol CR. Characterization of laccase isoforms produced by Pleurotus ostreatus in solid state fermentation of sugarcane bagasse. Bioresour Technol. 2012;114:735–739. doi: 10.1016/j.biortech.2012.03.058. [DOI] [PubMed] [Google Scholar]
  17. Khozani MA, Emtiazi G, Aghaei SS, Ghasemi SM, Zolfaghari MR. Application of fungal laccase for heavy metals precipitation using tannin as a natural mediator. Int J Environ Sci Technol. 2020 doi: 10.1007/s13762-020-02992-7. [DOI] [Google Scholar]
  18. Kumar A, Singh D, Sharma KK, Arora S, Singh AK, Gill SS, Singhal B. Gel-based purification and biochemical study of laccase isozymes from Ganoderma sp. and its role in enhanced cotton callogenesis. Front Microbiol. 2017;8:674. doi: 10.3389/fmicb.2017.00674. [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Kuwahara M, Glenn JK, Morgan MA, Gold MH. Separation and characterization of two extracelluar H2O2-dependent oxidases from ligninolytic cultures of Phanerochaete chrysosporium. FEBS Lett. 1984;169(2):247–250. doi: 10.1016/j.heliyon.2020.e03170. [DOI] [Google Scholar]
  20. Laemmli UK. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature. 1970;227(5259):680–685. doi: 10.1038/227680a0. [DOI] [PubMed] [Google Scholar]
  21. Li K, Helm RF, Eriksson KEL (1998) Mechanistic studies of the oxidation of a non‐phenolic lignin model compound by the laccase/1‐hydroxybenzotriazole redox system. Biotechnol Appl Bioc 27(3):239–243. https://iubmb.onlinelibrary.wiley.com/doi/abs/. Doi: 10.1111/j.1470-8744.1998.tb00500.x
  22. Liu EH, Gibson DM. Visualization of peroxidase isozymes with eugenol, a noncarcinogenic substrate. Anal Biochem. 1997;79(1–2):597–601. doi: 10.1016/0003-2697(77)90439-0. [DOI] [PubMed] [Google Scholar]
  23. More SS, Renuka PS, Pruthvi K, Swetha M, Malini S, Veena SM. Isolation, purification, and characterization of fungal laccase from Pleurotus sp. Enzyme Res. 2011;2011:1–7. doi: 10.4061/2011/248735. [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. Navarro D, Rosso MN, Haon M, Olivé C, Bonnin E, Lesage-Meessen L, Chevret L, Coutinho PM, Henrissat B, Berrin JG. Fast solubilization of recalcitrant cellulosic biomass by the basidiomycete fungus Laetisaria arvalis involves successive secretion of oxidative and hydrolytic enzymes. Biotechnol Biofuels. 2014;7(1):1–14. doi: 10.1186/s13068-014-0143-5. [DOI] [PMC free article] [PubMed] [Google Scholar]
  25. Ngo TT, Lenhoff HM. A sensitive and versatile chromogenic assay for peroxidase and peroxidase-coupled reactions. Anal Biochem. 1980;105(1):389–397. doi: 10.1016/0003-2697(80)90475-3. [DOI] [PubMed] [Google Scholar]
  26. Oliveira HP, Silva RG, Oliveira JT, Sousa DO, Pereira ML, Souza PF, Soares AA, Gomes VM, Mosteiro-Moreira ACO, Moreno FBMB, Vasconcelos IM. A novel peroxidase purified from Marsdenia megalantha latex inhibits phytopathogenic fungi mediated by cell membrane permeabilization. Int J Biol Macromol. 2017;96:743–753. doi: 10.1016/j.ijbiomac.2016.12.083. [DOI] [PubMed] [Google Scholar]
  27. Oloketuyi S, Annovi G, De Marco A. Peroxidase zymograms obtained by agarose native gel electrophoresis have unmet resolution and completeness. Int J Biol Macromol. 2020;156(1):869–873. doi: 10.1016/j.ijbiomac.2020.04.058. [DOI] [PubMed] [Google Scholar]
  28. Pütter J, Becker R (1983) Peroxidase. In: Methods of enzymatic analysis. Vol 3. HU Bergmeyer (Ed.). pp 286–293.
  29. Quintanar-Gómez S, Arana-Cuenca A, Mercado-Flores Y, Gracida-Rodríguez, JN, y Téllez-Jurado A (2012) Effect of particle size and aeration on the biological delignification of corn straw using Trametes sp. 44. BioResource. 7(1):327–344. https://ojs.cnr.ncsu.edu/index.php/BioRes/article/view/BioRes_07_1_0327_QuintanrG_CFRJ_Particle_Size_Aeration_Bio_Delignification
  30. Sawicki E, Stanley TW, Hauser TR, Elbert W, Nee JL. Mass spectra of aliphatic thiols and sulfides. Anal Chem. 1961;33(6):702–722. doi: 10.1021/ac60174a018. [DOI] [Google Scholar]
  31. Suzuki T, Honda Y, Mukasa Y, Kim SJ. Characterization of peroxidase in buckwheat seed. Phytochemistry. 2006;67(3):219–224. doi: 10.1016/j.phytochem.2005.11.014. [DOI] [PubMed] [Google Scholar]
  32. Téllez-Téllez M, Sánchez C, Díaz R, Díaz-Godínez G (2012) Zymogram patterns of extracellular laccases of Pleurotus species grown on non-inducer agar medium. Rev Mex Ing Quim 11(3):383–388. http://www.scielo.org.mx/scielo.php?script=sci_arttext&pid=S1665-27382012000300003&lng=es&tlng=en
  33. Tonami H, Uyama Nagahata R, Kobayashi S. Guaiacol oxidation products in the enzyme-activity assay reaction by horseradish peroxidase catalysis. Chem Lett. 2004;33(7):796–797. doi: 10.1246/cl.2004.796. [DOI] [Google Scholar]
  34. Vantamuri AB, Kaliwal BB. Purification and characterization of laccase from Marasmius species BBKAV79 and effective decolorization of selected textile dyes. 3 Biotech. 2016;6(2):1–10. doi: 10.1007/s13205-016-0504-9. [DOI] [PMC free article] [PubMed] [Google Scholar]
  35. Wang KF, Hu JH, Guo C, Liu CZ. Scale-up laccase production from Trametes versicolor stimulated by vanillic acid. Bioproc Biosyst Eng. 2016;39(7):1041–1049. doi: 10.1007/s00449-016-1582-0. [DOI] [PubMed] [Google Scholar]
  36. Wilkesman J, Castro D, Contreras LM, Kurz L. Guaiacol peroxidase zymography for the undergraduate laboratory. Biochem Mol Biol Edu. 2014;42(5):420–426. doi: 10.1002/bmb.20809. [DOI] [PubMed] [Google Scholar]
  37. Wong DW. Structure and action mechanism of ligninolytic enzymes. Appl Biochem Biotechnol. 2009;157(2):174–209. doi: 10.1007/s12010-008-8279-z. [DOI] [PubMed] [Google Scholar]
  38. Zerva A, Zervakis GI, Christakopoulos P, Topakas E. Degradation of olive mill wastewater by the induced extracellular ligninolytic enzymes of two wood-rot fungi. J Environ Manage. 2017;203:791–798. doi: 10.1016/j.jenvman.2016.02.042. [DOI] [PubMed] [Google Scholar]

Articles from 3 Biotech are provided here courtesy of Springer

RESOURCES