Mutanase Enzyme from Paracoccus mutanolyticus RSP02: Characterization and Application as a Biocontrol Agent

Abstract

Mutanases are enzymes that have the ability to cleave α-1,3 linkages in glucan polymer. In the present investigation, mutanase enzyme purified from the culture filtrate of Paracoccus mutanolyticus was evaluated for Streptococcal biofilm degradation and antimicrobial activity against pathogenic fungi along with enzyme kinetics, activation energies, pH and thermal stability. Biochemical and molecular characterization depicted that the enzyme showed optimum activity at pH 5.5 and at 50 °C. It displayed Michaelis–Menten behaviour with a K_m of 1.263 ± 0.03 (mg/ml), V_max of 2.712 ± 0.15 U/mg protein. Thermal stability studies denoted that it required 55.46 and 135.43 kJ mol⁻¹ of energy for activation and deactivation in the temperature range of 30–50 °C and 50–70 °C respectively. Mutanase activity was enhanced ~ 50 and 75% by Fe²⁺ and EDTA, respectively, while presence of Hg²⁺ and Mn²⁺ inhibit > 90% of its activity. This enzyme has a molecular mass of 138 kDa and showed monomeric nature by Zymography. Scanning electron microscopy analysis of mutanase treated Streptococcal cells revealed cleavage of linkages among the cells and complete separation of cells, indicating its potential in dentistry as an anticaries agent in the prophylaxis and therapy of dental caries. In addition, antifungal activity of mutanase against Colletotrichum capsici MTCC 10147 and Cladosporium cladosporioide MTCC 7371 revealed that the enzyme has potential towards biological control of phytopathogens which could be used as an alternative bio-control agent against chemical pesticides in the future.

Electronic supplementary material

The online version of this article (10.1007/s12088-019-00821-1) contains supplementary material, which is available to authorized users.

Introduction

α-1,3-Glucanases (EC 3.2.1.59), also called mutanases, are enzymes capable of hydrolyzing glycosidic bonds of α-(1,3)-D-glucan present in exopolysaccharide (EPS) matrix produced by oral Streptococci and cell wall of fungi [1]. The α-(1,3)-D-glucan produced by oral Streptococci is water insoluble in nature and harbour other acid producing microbial strains (Lactobacillus sp.) responsible for demineralization of tooth enamel during the process of caries development [2]. Mutanases were shown to have a potential role in preventing dental caries because of their ability to degrade water insoluble α-(1,3)-D-glucans i.e. mutans produced by cariogenic Streptococci. In addition to its medical application to treat dental caries, mutanases were also used to break the cell wall component, especially α-(1,3)-D-glucan of pathogenic fungi and was shown to have application in agriculture field as a biocontrol agent [3].

Mutanases producing microorganisms have been mainly isolated from soil source, however recently Waiter et al. [4] reported mutanases in gut microbiome of insect Diaperis boleti. Mutanases isolated from both fungi and bacteria are known to hydrolyze α-(1,3)-D-glucan, but differ in their catalytic activity from source to source. Fungal mutanases from Trichoderma, [1] and Aspergillus, [5] were shown to have exolytic activity on mutan and produce glucose as an end product, while mutanases from bacterial source (Paracoccus [6] Paenibacillus [2], Bacillus [7], and Streptomyces [8–10]) cleave the mutan and produces nigero-oligomers indicating its endolytic nature [11]. This endolytic activity plays a significant role in the biofilm degradation hence, bacterial mutanases are the preferred source towards prevention of dental caries [6, 12] and reports are also available on bacterial mutanases that indicates their potential to degrade the biofilm formed by Streptococci by in vitro studies [6–8]. In the previous study, authors have reported a novel strain, Paracoccus mutanolyticus RSP02, for mutanase production. Since this strain is newly reported for the production of mutanase, the present investigation mainly focused on the enzyme characterization with respect to its kinetic properties, thermal stability and evaluation of its application in biofilm degradation and biological control of plant pathogens. The present study will assist to find out the optimum conditions of mutanase in terms of pH, temperature and compatibility with metal ions to make use of its efficiency towards dental plaque degradation/reduction and biological control of fungal plant pathogens.

Materials and Methods

Extraction of Mutan and Mutanase

The α-(1,3)-D-glucan (mutan) substrate used in this study was produced and extracted from Streptococcus mutans MTCC 497 as described by Buddana et al. [13], briefly, S. mutans (MTCC 497) strain was grown in brain heart infusion (BHI) medium supplemented with 5% sucrose at 37 °C at an agitation speed of 150 rpm. After 72 h of incubation, the cell mass was separated by centrifugation at 15,000g and the pellet was treated with 2 N KOH for 2 h at 95 °C for the extraction of cell associated glucans. The alkali mixture was centrifuged at 15,000g for 30 min and supernatant was collected and neutralized to pH 7.0 with glacial acetic acid for the precipitation of mutan. The precipitated mutan was collected and washed thrice with deionized water, lyophilized and stored at room temperature until further use. While mutanase was produced from P. mutanolyticus using α-(1-3)-D-glucan supplemented medium consisting of (g/L) inositol—10, peptone—5, yeast extract—3, KH₂PO₄—2, NH₄Cl—2, KNO₃—2 and MgSO₄7H₂O—0.3 as reported earlier by Buddana et al. [6]. The produced extracellular enzyme was concentrated and purified using ion exchange chromatography. The purified mutanase was used in the study for characterization at a concentration of 1 mg/ml (2.59 U/mg). The protein markers used for native PAGE were procured from M/s Genei (Bangalore, India).

Zymography

For zymography, the mutanase was produced and purified using ammonium sulphate precipitation followed by DEAE cellulose anion exchange chromatography as reported earlier [6] and used in this study. Zymography study was carried out by using non- denaturing (native) PAGE with 8% polyacrylamide. The substrate [α-(1-3)-D-glucan] (50 mg/ml) was initially dispersed in deionized water using sonication process and was loaded to the separating gel. A 4% stacking gel was used for preparation of wells. Mutanase samples both reduced (treated with β-mercaptoethanol) and native (untreated) were loaded into wells along with two standard native proteins (catalase, 240 kDa) and bovine serum albumin (67 kDa). Electrophoresis was carried out at a potential of 100 volts and at a current of 20 mA at 4 °C. Later the gel was made into two equal parts, where one of it was used for visualization of protein bands by 0.025% Coomassie Brilliant Blue R 250 dye and the other part for mutanase enzyme activity by Congo red staining [6]. For mutanase activity, the gel, after electrophoresis was incubated in 0.2 M sodium acetate buffer (pH 5.5) initially for a period of 12 h. The gel was then stained with 0.25% Congo red solution for 30 min and then washed with 1 M NaCl solution for visualization of mutanolytic activity.

Enzyme Characterization

Effect of Temperature

The effect of temperature on the mutanase activity was studied by incubating the enzyme–substrate complex (1.0 ml of purified enzyme solution was reacted with 1.0 ml of 0.2% substrate) for 30 min at various temperatures ranging from 30 to 70 °C with an interval of 5 °C while, for stability studies, the mutanase was subjected to predetermined temperatures (30–70 °C with an interval of 5 °C) for 60 min and enzyme activity was performed at 50 °C (20 ml reaction volume). Aliquots were withdrawn at regular intervals of 10, 20, 30, 40, 50 and 60 min and the enzyme assay was carried out according to Michael Somogyi method [14, 15]. One unit of mutanase activity (U) is defined as the amount of enzyme that catalysed for the release of reducing sugars equivalent to 1.0 μmol/min under the defined conditions. The activation (E_a) and deactivation (E_d) energies were determined according to Arrhenius law. The activation energy was calculated from the slope of linear plot between enzyme activity Ln(V) versus 1/^oK, for the temperature range of 30–50 °C, whereas deactivation energy was calculated from the slopes of thermal stability studies between 55 and 70 °C.

Effect of pH

In order to determine the optimum pH for mutanase activity, enzyme–substrate complex (1.0 ml of purified enzyme solution and 1.0 ml of 0.2% substrate dispersed in different pH buffers) was incubated for 30 min in different pH buffers ranging from 4.0 to 10.0. Buffers with various pH range used include, sodium citrate (pH 4.0 and pH 5.0), sodium acetate (pH 5.5), phosphate (pH 6.0 and pH 7.0), Tris-HCl (pH 8.0), glycine–NaOH (pH 9.0) and bicarbonate (pH 10.0). For stability studies, initially the enzyme solution was incubated at various pH buffers (4.0–10.0) for 6 h at room temperature without substrate. Enzyme aliquots were withdrawn at regular intervals and residual enzyme activity was estimated.

Effect of Metal Ions

The effect of metal ions on the mutanase activity was estimated by supplementing different metal ions such as sodium (Na⁺), potassium (K⁺), ammonium (NH₄⁺), calcium (Ca²⁺), magnesium (Mg²⁺), barium (Ba²⁺), copper (Cu²⁺), iron (Fe²⁺, Fe³⁺), cobalt (Co²⁺), mercury (Hg²⁺), manganese (Mn²⁺), tin (Sn²⁺) and zinc (Zn²⁺) at a concentration of 1 mM. Initially, the enzyme was incubated in respective metal solutions for 60 min along with control (without any metal ion) and later the enzyme activity was performed at 50 °C for 30 min and the reducing sugar equivalents were quantified using Michael Somogyi method. The residual activity was calculated against the control.

Effect of Chemical Agents

The effect of chemical agents such as EDTA, phenylmethylsulfonyl fluoride (PMSF), iodoacetic acid (IAA), phenanthroline, β-mercaptoethanol, Triton X-100, para-aminobenzoic acid (PABA) and imidazole on the mutanase activity was evaluated by incubating the enzyme in a specific chemical agent at a concentration of 1 mM for 60 min. After incubation, the enzyme assay was performed and residual activity was calculated against control.

Enzyme Kinetics

The enzyme kinetic parameters such as K_m and V_max were evaluated by using different concentrations (1–10 mg/ml at an interval of 1 mg/ml of substrate). The enzyme activity was performed at 50 °C by incubating for 30 min for each substrate concentration, without changing the enzyme concentration. The Michaelis–Menten parameters (K_m and V_max) were calculated using Lineweaver–Burk plot.

Biofilm Degradation Studies

For biofilm degradation studies, Streptococcus mutans was grown in brain heart infusion (BHI) medium supplement with sucrose as carbon source. After 48 h of incubation at 37 °C the cells were centrifuged at 10,000g for 10 min. The cells were then washed with 50 mM phosphate buffer (pH 7.2) for removal of medium components. Then the cells were divided into two parts; one part was treated with mutanase enzyme (~ 1 mg/ml having 2.59 U/mg), and the other with sodium acetate buffer (control) for 6 h. After incubation, the cells were then washed with Millipore water, fixed with 3.5% gluteraldehyde and used for scanning electron microscopy (SEM) analysis.

Antifungal Activity

Well-plate method was followed for antifungal activities and for measuring the zone of inhibition by mutanase against selected test organisms. The test organisms used in the study include human pathogens Aspergillus fumigatus MTCC 12039, Candida albicans and plant pathogens Cladosporium cladosporioide MTCC 7371, Colletotrichum capsici MTCC 10147 for determination of antifungal activity. Initially fungal strains were grown in Czapek dox broth. Purified mutanase was diluted with 0.2 M sodium acetate buffer (pH 5.5) to get final concentration of 100 μg/ml. Medium and the Petri plates were initially sterilized at 121 °C for 15 min. Under sterilized environment, the agar medium was poured into plates. After solidification, 60 µl of test inoculum was spread on the plates using sterile spreader and wells were made with sterile cork borer. In each well, 100 µl of test (mutanase), control (buffer) and standard (cycloheximide) were placed. The plates were first incubated for 30 min at 4 °C to allow the compounds to diffuse into agar, and then subsequently incubated for 48 h at room temperature for antifungal activity. Zone diameters were expressed in mm using calibrated scale.

Results and Discussion

Zymography

Zymography studies of purified mutanase indicates a clear hydrolytic zone (in lanes 3 and 4) at same place for both native and reduced (β-mercaptoethanol treated) enzyme after Congo red treatment. Occurrence of hydrolytic zones for both proteins indicated that the mutanase produced by P. mutanolyticus is a monomer. This observation is in corroboration with the reported mutanases from Paenibacillus curdlanolyticus MP-1 sp [14]. In contrast, fungal mutanase from Trichoderma asperellum CECT 20539 is a dimer [3], whereas mutanase from Streptomyces chartreusis and Trichoderma harzianum CCM-F470 is a tetramer [8, 16]. The Coomassie Brilliant Blue treated gel also showed protein bands (in lanes 1 and 2) at the same position similar to that noticed with Congo red treated gel indicating the observed clear zone is due to the mutan degradation by the loaded mutanase (Fig. 1). Otsuka et al. [17] working with chimeric glucanase also reported the zymography of the enzyme using mutan agar well-plate diffusion assay while PAGE based zymographic analysis of mutanase is being reported for the first time in this study.

Fig. 1.

Native PAGE and zymography of mutanase enzyme produced from P. mutanolyticus. M1: catalase 240 kDa; M2 and M3: bovine serum albumin 67 kDa; L1 and L3: native mutanase enzyme; L2 and L4: reduced mutanase enzyme; L3 and L4: hydrolytic zone corresponding to protein bands on native gel indicating the mutanase activity

Enzyme Characterization

Effect of Temperature

Influence of temperature on the activity and stability of mutanase enzyme was studied at a range of 30–70 °C. The results demonstrate that temperature plays a vital role on catalytic behaviour of this enzyme (Fig. 2). This can be confirmed based on the data that optimized activity was observed at 50 °C (Fig. 2a) and any variation in reaction temperature resulted in drastic reduction in substrate catalysis. Besides being the temperature of 50ºC which provides better stability for the enzyme (Fig. 2b) conflicting reports were observed in literature with regard to temperature maxima for mutanases. Pleszczynska et al. [18] reported an optimum temperature range of 40–45 °C for mutanase produced by P. curdlanolyticus MP-1, whereas Matsuda et al. [7] observed it at 50 °C for Bacillus circulans HU-MI mutanase. This difference in results might be due to the specific action of the mutanase enzyme among different species. Further analysis of temperature dependent mutanase activity profile revealed that mutanase from P. mutanolyticus has its catalytic function at 50 °C and thereafter sharply declines with further increase in temperature up to 70 °C (Fig. 2a). Only 50% of enzyme activity was retained when temperature was increased from 50 to 60 °C and upon further increase in reaction temperature to 70 °C only 2.9% of residual activity was observed which may be attributed to protein denaturation. Temperature based enzyme stability studies indicated that this mutanase was stable up to 50 °C for 60 min. However, at 55 °C, the enzyme lost 35% of activity within 20 min and almost 90% of activity at 65 °C (Fig. 2b). In general, it was reported that the thermo stability of different bacterial mutanases was in the range of 40–50 °C [19]. Suyotha et al. [20] reported that the mutanase from P. glycanilyticus FH11 was stable for only 10 min at 50 °C, whereas Pleszczynska et al. [18] observed that P. curdlanolyticus MP-1 mutanase was stable for 60 min at 45 °C. These data further reveal that mutanase isolated from P. mutanolyticus in the present study has better application potential at industrial level due to its higher stability compared to reports available in the literature [18, 20].

Fig. 2.

Effect of temperature on the mutanase enzyme from Paracoccus mutanolyticus RSP02, activity (a) and stability (b)

Activation and deactivation energies for P. mutanolyticus mutanase were calculated based on catalytic functionality of enzyme at different temperatures (30–70 °C). An Arrhenius plot was obtained between Ln(V) and reciprocal of absolute temperature (1/^oK), whose slope equals − E_a/RT is used for calculating activation energy (E_a), whereas deactivation energy of mutanase was calculated by plotting the log of K_d as a function of the inverse of the absolute temperature. The plot represented linear from in the temperature range from 30 to 50 °C suggesting the Arrhenius plot is a reliable model to represent the effect of temperature on catalytic activity. From the plot, the calculated activation and deactivation energies were found to be 55.46 and 135.43 kJ mol⁻¹ respectively.

Effect of pH

The effect of pH on enzymatic activity and stability was investigated at a pH range from 4.0 to 10.0 using α-(1-3)-D-glucan as substrate. It was clear from the observations that purified mutanase from P. mutanolyticus exhibited optimal catalytic activity in the pH range of 5.5–7.0 (Fig. 3). The mutanase activity was drastically reduced at acidic (4.0 and 5.0) and at alkaline pH ranges (7.5–10.0). Similar results were reported for P. curdlanolyticus MP-1 mutanase where maximum activity was noticed in the pH range of 5.5–6.5 by Pleszczynska et al. [18], while Takehara et al. [8] working with S. chartreusis F2 purified mutanase observed optimized activity in the pH range of 5.5–6.0. Likewise, little higher optimum pH range (7.5–8.5) was recorded for mutanase of B. circulans WL-12 [21]. The data over a pH range of 4–10 on the other hand indicated that this enzyme showed little variation in activity even incubating for 6 h (data not shown) suggesting that mutanase of the present study is highly stable in above pH range similar to that of other bacterial mutanases [5, 18, 21, 22].

Fig. 3.

Effect of pH on the mutanase enzyme from Paracoccus mutanolyticus RSP02

Effect of Metal Ions

Metal ions are known to bind to the enzymes and act as cofactors which may influence the enzyme activity [23]. Metal ions will alter the protein conformational changes or by influencing the availability of the enzyme to the substrate. The effect of different metal ions on mutanase activity was evaluated and the results were tabulated in Table 1. Among the tested metal ions, mutanase activity was stimulated by Fe²⁺ (149%), Cu²⁺ (132%) and Ba²⁺ (112%), whereas metal ions such as Fe³⁺ inhibited it up to 70% and Hg²⁺, Mn²⁺ inhibited up to 90%. It is interesting to note that a 23% enhancement of mutanase activity was reported by Pleszczynska et al. [18] in presence of Ca²⁺ while in the present study, supplementation of Ca²⁺ to the reaction mixture showed little variation in catalytic activity of mutanase isolated from P. mutanolyticus. Supplementation of Hg²⁺ ion resulted in complete loss of activity with respect to mutanase from P. curdlanolyticus [18] as well as from fungal strain, T. harzianum [1], whereas only 20% activity was noticed with mutanase from P. mutanolyticus (Table 1). It is interesting to note that iron did not show any impact on mutanases of other bacterial strain Paenibacillus MP-1, or fungal mutanase from T. harzianum [1, 18], while it showed approximately 50% higher activity in presence of iron in case of P. mutanolyticus mutanase.

Table 1.

Effect of metal ions on the mutanase activity from Paracoccus mutanolyticus RSP02

Metal ion	Relative activity (%)
Control	100.00 ± 0.40
Fe2+	149.81 ± 1.86
Cu2+	132.03 ± 0.87
Ba2+	112.56 ± 0.78
Co2+	106.26 ± 1.06
Ca2+	97.01 ± 1.45
Mg2+	91.50 ± 1.76
Na+	86.12 ± 0.64
K+	81.04 ± 0.94
Zn2+	74.00 ± 0.89
Sn2+	67.48 ± 0.17
NH4+	66.16 ± 1.01
Fe3+	29.71 ± 0.34
Hg2+	18.99 ± 0.50
Mn2+	13.91 ± 0.47

Effect of Chemical Agents

The effect of various chemical agents on the mutanase activity was studied by incubating the enzyme for one hour at 1.0 mM concentration and performing the assay. The data suggested that EDTA enhanced the mutanase activity 76% confirming that the enzyme is not a metalloenzyme (Table 2). This result is in contrast with observed 66% reduction in activity of Paenibacillus MP-1 mutanase by EDTA [18]. Phenyl methyl sulfonyl fluoride (PMSF), para-aminobenzoic acid (PABA) and imidazole inhibited the enzyme activity by 37, 33 and 21% respectively. Similar kind of inhibition of mutanase activity was also observed by Pleszczynska et al. [18] with Paenibacillus MP-1. The observed partial reduction of mutanase activity (37%) in presence of PMSF does suggest that P. mutanolyticus mutanase may have serine amino acid in its catalytic domain. However, chemical agents such as iodoacetic acid, phenanthroline, β-mercaptoethanol, Tween 80 and Triton X-100 do not show any effect on the enzyme activity. An identical study by Pleszczynska et al. [18] revealed total enzyme inhibition with p-chloromercuribenzoate (PCMB), whereas EDTA and iodoacetamide reduced the enzyme activity by 66 and 43% respectively. On the other hand, non-ionic detergents like Tween 20 and 80 induce the enzyme activity by 116 and 52% respectively.

Table 2.

Effect of chemical agents on the mutanase activity from Paracoccus mutanolyticus RSP02

Chemical agent	Residual activity (%)
Control	100.00 ± 0.40
EDTA	175.96 ± 2.01
IAA	103.10 ± 1.06
Phenanthroline	100.00 ± 0.50
SDS	100.00 ± 0.89
β-Mercaptoethanol	100.00 ± 0.94
Triton-X	100.00 ± 0.87
Tween-80	96.12 ± 0.78
Urea	86.82 ± 1.45
Imidazole	78.29 ± 0.17
PABA	66.66 ± 1.01
PMSF	62.79 ± 0.64

Enzyme Kinetics

A steady-state mutanase kinetic study was performed using different concentration of α-(1-3)-D-glucan as substrate material and constant quantity of mutanase at 50 °C. These data exhibited Michaelis–Menten kinetics (Fig. 4). A Lineweaver–Burk plot was plotted between (1/S) substrate concentration versus enzyme activity (1/V) for calculating V_max and K_m (Fig. 4). The K_m and V_max values of purified mutanase were estimated as 1.263 ± 0.03 (mg) and 2.712 ± 0.15 U/mg protein respectively. Similar study with Paenibacillus MP-1 mutanase revealed a very high K_m (11.6 mg) [18], whereas mutanase from Pseudomonas sp. had a K_m of 80 mM with anhydroglucose units for insoluble glucan from S. mutans [24]. However, the data presented here is comparable with the data reported by Meyer and Phaff [21] where the authors used highly purified colloidal glucan from Schizosaccharomyces pombe.

Fig. 4.

Michelin–Menten hyperbola and Lineweaver–Burk plot of mutanase enzyme from Paracoccus mutanolyticus RSP02

Biofilm Degradation Application

The application of mutanase on streptococcal biofilm degradation studies was performed by treating actively growing S. mutans with purified mutanase from P. mutanolyticus and observing under scanning electron microscopy (SEM). SEM studies in the present study revealed that the S. mutans cells were compact in nature and engrossed with network like threads interconnecting with other cells indicating the production of exo-polysaccharides [α-(1-3)-glucans] (Fig. 5a). Whereas, complex biofilm forming cells when treated with purified mutanase, resulted in appearance of individual cells or loss of connectivity among cells (Fig. 5b) revealing the biofilm degradation capability of mutanase enzyme. This is a direct and first visual report on mutanase based biofilm degradation, however, similar biofilm degradation pattern on glass plate has been reported [6, 25].

Fig. 5.

Scanning electron microscopy of biofilm degradation studies by mutanase enzyme. aS. mutans cells grown in BHI supplemented with sucrose, arrows indicating the cell to cell interconnection with water insoluble exo-polysaccharides. b Mutanase treated cells, showing clevage of cells from biofilm aggregations

Antifungal Activity

Biological control of plants from phytopathogenic fungal disease is one of the most important challenges in agriculture. Around 85% of plant diseases are caused by fungal or fungal-like organisms. Structurally, fungal cell walls are composed of branched 1,3-β-D-glucans cross-linked to chitin in the core region, whereas outer layers are embedded with 1,3-α-D-glucans [26]. Currently, chemical fungicides are extensively adopted in control of plant diseases, however extensive usage of chemical fungicides has its own drawbacks due to their highly toxic nature, development of resistance, presence of fungicide residues in food products and pollution of environment [27]. Application of biological agents such as enzymes as an alternative strategy to control plant diseases is one of the environmental friendly approaches.

To evaluate the bio-control potentiality of P. mutanolyticus mutanase, in the present study, antifungal activity of mutanase enzyme was determined against four fungal pathogens (Colletotrichum capsici MTCC 10147, Aspergillus fumigatus MTCC 12039, Cladosporium cladosporioide MTCC 7371 and Candida albicans). From the Table 3 it is evident that mutanase enzyme exhibited significant antifungal activity against all the tested strains. Among the tested organisms, the enzyme showed better antifungal activity against C. albicans, C. capsici and C. cladosporoides than the A. fumigatus. This may be due to the presence of 1,3-α-D-glucans in outer layer of their cell walls, whereas in A. fumigatus it was embedded in lower side. Mutanase antifungal activity with respect to A. fumigatus is 44% lower than the standard cycloheximide. Comparative analysis of mutanase activity in C. capsici with standard drug revealed that the antifungal activity is comparable to that of cycloheximide. Whereas in C. cladosporioide, mutanase exhibited 35% higher antifungal activity compared with standard. In C. albicans, the standard drug cycloheximide did not show any antifungal activity, whereas mutanase exhibited significantly higher activity (Table 3). This application will synergize the anticariogenic potential of mutanases as C. albicans exhibits synergistic activity in the formation of dental caries along with S. mutans [28]. Only a single report on antifungal property of mutanase was available in the literature. Ait-Lahsen et al. [29] performed antifungal activity of exo-α-1,3-glucanase isolated from T. harzianum against different fungal pathogens like A. niger, Fusarium oxysporum, Penicillium aurantiogriseum etc., by tube method and observed 70% of antifungal activity against P. aurantiogriseum at 270 µg/mL of enzyme concentration.

Table 3.

Antifungal activity of purified mutanase enzyme from P. mutanolyticus against different pathogenic fungi

Organism	Zone of growth inhibition in mm
	Mutanase	Cycloheximide
Colletotrichum capsici	18.3 ± 0.6	20.0 ± 0.3
Cladosporium cladosporoides	18.4 ± 0.6	11.8 ± 0.2
Candida albicans	19.7 ± 0.7	00
Aspergillus fumigatus	12.1 ± 0.5	21.6 ± 0.4

Conclusions

An endolytic mutanase isolated from P. mutanolyticus RSP-02 was purified and characterized and evaluated for breakdown of complex water insoluble α-(1-3)-D-glucan polymer of cariogenic Streptococci. This enzyme has low K_m value with respect to other bacterial mutanases, a very high deactivation energy (135.43 kJ mol⁻¹) and low activation energy (55.46 kJ mol⁻¹), thermal and pH stability, broader pH range activity profile, ability to degrade insoluble mutan suggests its potential application in the field of dentistry as an active constituent of oral hygiene products. Antifungal activity of mutanase enzyme against fungal phytopathogens (Colletotrichum capsici MTCC 10147 and Cladosporium cladosporioide MTCC 7371) revealed the application of mutanase enzyme as an alternative bio-control agent against chemical fungicide in the control of plant pathogens.

Electronic supplementary material

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