Production, Purification, and Characterization of a Novel Exopolysaccharide from Probiotic Lactobacillus amylovorus: MTCC 8129

Abstract

Extracellular polysaccharides (EPS) produced by Lactic Acid Bacteria have an individual effect on the flavour and consistency of novel food materials, as well as potential therapeutic applications. The purpose of this study was to create, improve, and characterise EPS from Lactobacillus amylovorus MTCC 8129. FTIR examination showed the compound's composition (acetyl group, hydroxy group, ring structure) as well as the numerous interlinks between sugar residues, which were then validated by Nuclear Magnetic Resonance Spectroscopy. Thermogravimetric examination showed that the EPS exhibited resistance to heat at a temperature of 640 °C, with antioxidant levels ranging from 70 to 85% and emulsification activity above 50%. Furthermore, it has 180% water holding capacity and 140% oil holding capacity. Based on these findings, it seems that the EPS that was reviewed might potentially be an advantageous addition to the food processing industry.

Introduction

Food companies throughout the globe are continually looking for natural value-added substances or additives that improve functionality and biological activity. Furthermore, people are looking for healthier eating alternatives [1]. Probiotic Exopolysaccharides (EPS) is an important factor that is essential for improving the sensory aspects of food items, including texture and features. EPS have also received great interest for their prospective and pharmacological uses because of their biodegradability and lack of toxicity [2]. Lactic acid bacteria (LAB) are well recognised for their use in the dairy industry due to their technical and functional food purposes [3]. Moreover, microorganisms from extreme surroundings are also extensively recorded as the origins of various applications in the food sector, including antioxidant characteristics, emulsion stabilization, conduct, and others [4].

Although lacking in flavour, the extended duration of EPS was reported to improve the taste of milk products made with LAB when consumed [5]. EPS, when present in the gastrointestinal system for a certain duration, enhances the population of probiotic bacteria in the gut [6]. Contrary to EPS produced by other microorganisms, EPS obtained from LAB has many health advantages, such as its impact on cholesterol levels and its activity [7].

Lactic acid bacteria have a broad ability to create exopolysaccharides. Scientists worldwide are doing research on the production, purification, and characterization of exopolysaccharides from known strains of these bacteria [8]. Numerous studies have been authored about the examination, division, recognition, formation, genetic and metabolic manipulation, and practical characteristics of exopolysaccharides (EPS) generated by lactic acid bacteria (LAB) [9]. The properties of EPS produced from LAB are influenced by the specific strain, culture conditions, and composition of the medium [10]. The structure and characteristics of EPS are influenced not only by the quantity of the polymer generated but also by other parameters, such as the molecule's capacity to form bonds and its molecular configuration [11]. The majority of LABs generate a limited quantity of EPS, typically less than 100 mg/L, when cultured on a complicated medium. Utilizing elaborate mediums, including yeast extract and beef extract, is not economically viable for large-scale manufacturing because of their high cost and their interference with the precipitation of EPS using alcohols or acetone [2, 12].

In this work, EPS produced by Lactobacillus amylovorus MTCC 8129 was extracted from the culture broth using the acetone precipitation method. Additionally, the chemical composition was determined using FTIR, one-dimensional (1D) NMR, and TGA techniques. An in-depth analysis was conducted on the EPS, focusing on its structure and functioning, which uncovered its potential as a distinctive antioxidant, emulsifier, and flocculating agent. This finding has significant potential for future biotechnological uses.

Materials and Methods

Bacterial Strains and Culture Conditions

Lactobacillus amylovorus MTCC 8129 was purchased from MTCC (IMTECH Chandigarh) and cultured in MRS broth (Himedia) at 37 ºC for 24–48 h.

Screening Test for EPS Production

Test Tube Method

The tube method for EPS screening was performed as described by Christensen et al. [13], Test tubes containing MRS media and 1% L.amylovorus MTCC 8129 after inoculation were then kept for 24 h at 37 °C. Biofilm that is developed on the surfaces and sides of the polystyrene test tubes was stained with crystal violet for one hour, followed by the removal of the stain via double washing with phosphate-buffered saline (PBS). After that, PBS is used to wash the stained polystyrene test tube twice to remove the stain. After the test tube method was performed, the visible film on the sides and bottom of the tube visible film implied the formation of biofilm [13].

Biofilm Detection Using a Microtiter Plate Assay (MtP)

Biofilm production can be widely quantified using a microtiter plate assay in which 1% glucose was added to BHI and the bacterial suspension concentration was adjusted to 1 × 10⁸/ml (0.5 McFarland). The microplates containing bacteria were incubated at 37 °C. The isolates showed biofilm formation on the sides of 96 well plates that were dyed with 150 µl of crystal violet for 15 min and the cells were discarded in the well by washing twice with PBS pH 7.2 and drying at 60 °C for 1 h. The wells can be fixed before crystal violet staining by either drying them at 60 °C for an hour or submerging them in 150 µl of methanol for 20 min [14]. The crystal violet stain was then removed by rinsing the well twice with PBS. After the wells had been air-dried, methanol was used to dye the coated walls of the well. This is subsequently measured at 590 nm using a plate reader. Wells containing sterile BHI enriched with 1% glucose served as the negative controls in triplicate The experimental results were analysed by computing the mean ± standard deviation (SD) and doing an analysis of variance (ANOVA) to ascertain statistically significant distinctions. [15].

Method Using Congo Red Agar (CRA)

Lactobacillus amylovorus MTCC 8129 was screened for EPS production by culturing on CRA medium [16]. The CRA medium consists of 37 g/L BHI agar, 0.8 g/L Congo Red, and 36 g/L sucrose. After incubating at 37 °C for 24 h, the colony that exhibited varying colors was used to distinguish between biofilm producers and non-producers. Colonies that are black and have a dry crystalline texture are indicative of biofilm production, while colonies that remain pink are not biofilm producers [17].

Phenotype Characterization of EPS

EPS production was confirmed using the ruthenium red agar method. A stock solution of 10% ruthenium red filter sterilized was added to MRS agar to a final concentration of 0.08%. The bacterial cultures were then inoculated on the ruthenium red agar plate and incubated at 37 °C for 24 h [18].

Extraction and Purification of EPS

To produce EPS, bacterial cells obtained from 72hgrown MRS cultures were treated with 4% TCA for 30 min. Subsequently, it was centrifuged at 5000 × g (20 min) at 4 °C to precipitate the proteins. The precipitate was then added to an equal volume of chilled acetone and incubated at 4 °C overnight. The EPS was recovered after centrifugation at 12,000 × g for 20 min at 4 °C. The EPS pellet was subjected to dialysis using a 5 kDa membrane filter and lyophilized.

The lyophilized EPS was purified according to the modified protocol of Dabour and LaPointe (2005). 1% of the lyophilized EPS powder was extracted twice with an equal volume of phenol–chloroform-isoamyl alcohol (25:24:1 [v/v/v]) and precipitated overnight with an equal volume of acetone. The precipitated EPS was dissolved in water, dialyzed using a 5 kDa membrane for 24 h with two changes of water, and subsequently lyophilized using a lyophilizer (LYODEL).

Biochemical Characterization of EPS

The overall carbohydrate content of EPS was examined using the phenol–sulphuric acid approach [17], while the protein content was analyzed through Bradford's assay. The Barker Summerson assay [19] was utilized to quantify the lactic acid and the DNS (Dinitro Salicylic Acid) method [20] was employed to estimate the quantity of reducing sugars present in the exopolysaccharide.

Structural Characterization of Purified EPS

Fourier Transform Spectroscopy (FTIR)

Analysis of the functional groups present in the EPS sample was carried out using FTIR spectroscopy [21]. EPS was made into pellets by pressing with KBR at a 1:90 ratio and scanned at a range of 4000–400 cm⁻¹ with a 4 cm⁻¹ resolution [22].

Nuclear Magnetic Resonance (NMR)

NMR analysis was performed using 1D techniques at 60 °C after dissolving the samples in H₂O. Chemical shifts obtained in the aqueous samples were expressed in the form of ppm with reference to ¹H signals, respectively [23].

Thermogravimetric Analysis (TGA)

The EPS thermal stability was performed using TGA analysis. 5 mg of the sample was added into an aluminum sample holder in the microprocessor-driven temperature-control unit with an increasing temperature of 25 to 600 °C and a 10 °C gradient per minute at 50 min⁻¹ under N₂ gas flow [23].

Functional Characteristics of the EPS

Antioxidant Activity Assay

DPPH free radical scavenging activity was performed as described by Shimada et al. [24]. 200 μl of EPS sample with concentrations ranging from (0.2, 0.5, 1, 2, and 5 mg/ mL⁻¹) was mixed with 0.2 mM 20 μl DPPH dissolved in ethanol. The EPS-DPPH mixture was incubated at a dark condition at ambient temperature for 1 h and then the mixture was centrifuged (5000 × g for 10 min), and the OD was measured at 517 nm using a Microplate reader (Biotek). The equation for calculating DPPH RSA.

$$\left[\left(\text{A1}-\text{A2}\right)/\text{A1}\right]\times 100$$

where A1 is the value of the absorbance of a DPPH solution without a sample. A2 is the absorbance of EPS. Positive control- ascorbic acid [25].

ABTS scavenging activities were done according to Nitha et al. [26]. ABTS was diluted till it reached an absorbance of ~ 0.75 at 734 nm in phosphate buffer—pH 7.40. 180 µl of EPS with concentrations ranging from (0.2, 0.5, 1.0, 2.0, and 5.0 mg/mL ⁻¹) were mixed into 20 ul of ABTS solution and allowed to react at 30 °C for 5 min and OD 732 nm was measured using a microplate reader (Biotek). The equation for calculating ABTS RSA.

$$\left[\left(\text{A1}-\text{A2}\right)/\text{A1}\right]\times 100$$

A1 is the absorbance value of the ABTS radical solution without the sample in this example, and A2 is the absorbance value of the EPS solution. The positive control was ascorbic acid.

H₂O₂ scavenging activity was performed as described previously, with minor changes. 50 µL of EPS with concentrations ranging from (0.2, 0.5, 1.0, 2.0, and 5.0 mg/ mL⁻¹) was mixed with 120 µL of phosphate buffer (0.1 M, pH 7.40) and 30 µL of H₂O₂ solution (40 mM). After vigorously shaking the test mixture, it was incubated at 30 °C for 10 min. After this, the OD 230 nm was measured using a microplate reader (Biotek). The H₂O₂ scavenging activity was determined using the following equation.

$$\left[1-\left(\text{A2}-\text{A3}\right)/\text{A1}\right]\times 100,$$

A1 is the absorption of the control MilliQ water, A2 is the absorption of the specimen, and A3 is the absorption of the specimen without H₂O₂ solution. The positive control was ascorbic acid [25].

Ferric Reducing Antioxidant Power (FRAP) Assay

The FRAP assay was conducted to determine the capability of the extracts to decrease Fe³⁺ . The technique utilized was based on Oyaizu's (1986) method, with slight modifications. The experiment involved combining 50 µL of samples with different concentrations, 50 µL of distilled water, and 50 µL of 1 percent potassium ferricyanide, then incubating the mixture at 50 °C for 20 min. Afterward, 50 µL of 10% TCA and 0.1% ferric chloride were added, and the absorbance was measured at 700 nm. A rise in absorbance indicated an increase in the reducing power of the mixture. To serve as a positive control, ascorbic acid was used at a concentration of 1 mg/mL. The outcomes were expressed as a graph with the obtained IC50 values [25].

Analysis of Emulsifying Activity

The emulsifying efficiency of the EPS produced by L.amylovorus MTCC 8129 was assessed using the Cooper and Goldenberg method [26]. Food-grade vegetable oils (coconut, almond oil, groundnut oil, olive oil, and sesame oil) were rapidly mixed for 2 min in a 1 mg mL⁻¹ aqueous phase containing the EPS (oil: EPS in a 3:2, v/v ratio). At 24-h intervals, the height of all three layers, including the oil, emulsion, and aqueous layers, was monitored. [(volume of the emulsion layer × total volume) *100] was used to determine the emulsification index (E). EPS xanthan gum, along with the synthetic surfactant Tween 20 (SRL), was used as the test standard [27].

Analysis of Water-Holding Capacity (WHC) and Oil-Holding Capacity (OHC)

The capacity of holding water for L.amylovorus MTCC 8129 EPS was determined by adding 500 mg of EPS to a centrifuge tube along with 10 mL of distilled water. The supernatant was discarded after centrifuging, and the weight of the tube before and after osmosis was observed by simple weighting. WHC was determined using the following equation:

$$\%\text{WHC}=\left(\text{Weight of water bound}/\text{Initial sample weight}\right)\times 100$$

The OHC of EPS was performed according to Wang and Kinsella [28]. 500 mg of EPS was mixed with 10 mL of sunflower oil in a cyclomixer. The mixture was then left to stand at 37 °C for half an hour, with intermittent shaking every 10 min. Centrifuged the mixture for 25 min at 3,200 rpm, and the pellet was weighed. OHC was determined using the equation below.:

$$\%\text{OHC}=\left(\text{Weight of oil bound}/\text{Initial sample weight}\right)\times 100$$

As standard, Xanthan gum was used. All tests were carried out in triplicate. The experimental results were analysed by computing the mean ± standard deviation (SD) and doing an analysis of variance (ANOVA) to ascertain statistically significant distinctions.

Results

Screening Test for EPS Production

The Tube Method is a qualitative test for identifying biofilm produced at the air and liquid interfaces. When compared to control test tubes, L. amylovorus MTCC 8129 cultivated test tubes exhibited more crystal violet dye binding, suggesting biofilm formation (Fig. 1) [13]. Similarly, the microtiter plate (MtP) assay for L.amylovorus MTCC 8129 revealed OD values greater than the blank, indicating biofilm formation. Using the cutoff value, isolates may be categorized as biofilm producers or not (Fig. 2) [14]. The experimental results were analysed by computing the mean ± standard deviation (SD) and doing an analysis of variance (ANOVA) to ascertain statistically significant distinctions.

Fig. 1.

The Tube Method test for identifying bacteria that generate biofilm as a result of the presence of a visible film. L.amylovorus MTCC 8129 was showed more film compare to the blank

Fig. 2.

The microtiter plate (MtP) test assesses biofilm production using a plate reader. L.amylovorus MTCC 8129 has OD values higher for the higher inoculum

Conformation of EPS Production

Congo red staining of L.amylovorus MTCC 8129 showed colonies displaying dark red colonies with a dry, crystalline consistency after 72 h of growth on BHI agar. All these assays confirmed that L.amylovorus MTCC 8129 produced biofilm [29].

The absorption of ruthenium red by the cell wall of bacteria causes EPS-producing strains to become pink, while non-EPS-producing strains stay white. This is due to the existence of EPS, which prevents the bacterial cell wall from absorbing the stain present in the medium, resulting in white colonies. The results showed that the strain grew white colonies on ruthenium red-containing MRS media.

EPS Extraction and Purification

Total EPS from L.amylovorus MTCC 8129 was extracted using the acetone precipitation method and the EPS yield after purification was 610 mg/L [30].

Biochemical Characterization of EPS

L.amylovorus MTCC 8129 produced EPS with a total carbohydrate content of 90.23%. The Bradford test and UV spectrum readings at 595 nm showed no response, indicating the absence of nucleic acid or protein. The production of lactic acid by L.amylovorus MTCC 8129 was found to be 33%, respectively. Carbohydrates are responsible for growth, viability, and lactic acid production [31, 32]. L.amylovorus MTCC 8129 consumed 64% of the reducing sugars tested. The consumption of sugars during microbial development might explain the reduction in overall sugar content. All tests were carried out in triplicate and the experimental results were analysed by computing the mean ± standard deviation (SD) (Fig. 3, 4).

Fig. 3.

The technique of Congo red agar is utilized to detect the exopolysaccharides (EPSs) generated by L.amylovorus MTCC 8129 while being cultivated in a medium consisting of 10% sucrose, which are then released into the culture medium. The formation of biofilm of EPSs by L.amylovorus MTCC 8129 is indicated by the presence of black colonies

Fig. 4.

The ruthenium red stain on the bacterial cell wall is pink distinguishing the EPS producing strains, white colonies from those that do not produce EPS

Bioanalytical Characterization of EPS

Analysis of the Structure

Fourier Transform Spectroscopy (FTIR)

The presence of functional groups in the EPS produced by L.amylovorus MTCC 8129 exhibited a broad O–H stretching at 3300.25 cm⁻¹ indicating the vibration of sugars that are responsible for EPS water solubility [33]. The C–C was observed at 1635.06 cm⁻¹ (Fig. 5). The absorption bands around 1220.94–1012.62 cm⁻¹ represent the stretching vibrations of alcohols, acids, esters, and ether groups (C–O) [33]. The sharp C-O vibration was seen at 1012.62 cm⁻¹. Because of the presence of numerous sugar chains (polysaccharides), the absorption of about 1200–800 cm^-1 was deemed the fingerprint region [34]. Absorption bands detected at 1220.94 cm⁻¹ [35] reflected stretching frequencies of C-O alcohols, acids, esters, etc. FTIR was used to investigate changes in the functional group of EPS [36]. In addition to the functional groups previously described in the structure, the FT-IR spectra revealed a more complicated structure for EPS with distinct functional groups [37].

Fig. 5.

FTIR spectrum of EPS from L.amylovorus MTCC 8129

Nuclear Magnetic Resonance (NMR)

The polysaccharide ¹H NMR spectrum may be divided into three major regions: the anomeric region (δ_H 4.5–5.5), the ring proton region (δ_H 3.1–4.5), and the alkyl region (δ_H 1.2–2.3). Signals between δ5.2 and δ4.5 ppm in this spectrum represent the anomeric protons of the EPS's monosaccharide components [23].

These signals are frequently employed as indications to distinguish between complicated carbohydrate compounds. Protons connected to C2—C6 were ascribed to the signals in the spectrum (Fig. 6) between δ4.4 and δ3.2 ppm. Because of overlapping chemical changes, these signals were poorly resolved. The pattern of these signals resembled the spectrum of a reference sample [38] (Fig. 7, 8).

Fig. 6.

NMR of EPS from L.amylovorus MTCC 8129

Fig. 7.

TGA of EPS from L.amylovorus MTCC 8129

Fig. 8.

In vitro antioxidant capability of the EPS produced by Lactobacillus amylovorus MTCC 8129 where a DPPH radical scavenging activity, b ABTS radical scavenging activity, c H₂O₂ radical scavenging activity and d FRAP activity

Three signals in the anomeric region of the ¹H NMR spectrum corresponded to a trisaccharide repeating unit [38]. According to the carbohydrate research database (www.glyco.ac.ru), the chemical shift 5.2 (¹H, s) corresponds to the anomeric proton of α-D-mannose, 5.0 α-D-glucose, and 4.89 β -D-glucose. According to Maeda et al. (2004), chemical shifts δ5.14 were attributed to α-hex pyranosyl residue, while δ4.49, δ4.7, and δ4.44 were ascribed to pyranose ring forms in α-anomeric configuration [38]. The resonances at δ_H 5.341, 5.322, and 5.2 ppm are characteristic of anomeric proton in α -anomeric residues, while the signals at δ H 4.908, 4.408, and 4.450 ppm are typical of β-anomeric protons (Hallack et al., 2010). It was also said that the signal at 5.2 corresponds to α-D-mannose and that the signal at 4.9 corresponds to β-D-glucose [31].

Thermogravimetric Analysis (TGA)

Thermal analysis displays the thermal decomposition curves of EPS, revealing the two decomposition peaks in the curves. This demonstrates the complexities of EPS thermal deterioration. The initial breakdown stage of EPS occurs from 200 °C to 480 °C, with a maximal disintegration rate at 230 °C accounting for 14.4% of weight loss. This is sometimes referred to as dehydration induced by EPS moisture absorption. The second line of deterioration occurs between 500 °C and 679 °C, with an increase at 640.7 °C and a weight loss of 56.72%. The greatest weight loss occurs at about 650 °C during EPS thermal breakdown. At this stage, polysaccharide and protein chains are depolymerized, and sugar units are dehydrated [31].

Functional Properties of the EPS

In Vitro Antioxidant Activity Determination

The antioxidant potential of the EPS was evaluated by their scavenging ability by eliminating the free radicals in the DPPH, H₂O₂, ABTS, and FRAP by keeping ascorbic acid as standard. The EPS produced by L.amylovorus MTCC 8129 exhibited significant scavenging activity with a low conc of 0.5 mg/mL⁻¹. The experiments were conducted three times, and the results were evaluated by calculating the mean ± standard deviation (SD). The maximal activity levels of DPPH, H₂O₂, ABTS, and FRAP-mediated methods were 81, 72.43, 73.76, and 70%, respectively [32] (Table 1, 2).

Table 1.

The FTIR frequency range and functional groups present in the EPS sample

Vegetable oils	Emulsification Index
	Lactobacillus amylovorus MTCC 8129 EPS	Synthetic tween 20	EPS biopolymer Xanthan Gum
Coconut	54.01 ± 0.25	12.00 ± 0.15	55.11 ± 0.25
Almond	32.66 ± 0.25	6.66 ± 0.13	51.14 ± 0.15
Olive	52.66 ± 1.20	17.33 ± 0.24	72.56 ± 0.54
Groundnut	32.00 ± 0.97	26.66 ± 1.25	51.58 ± 0.47
Sesame	35.33 ± 0.65	23.40 ± 0.54	74.22 ± 0.26

Table 2.

The ¹H NMR Spectrum Formula, structure and biological properties present in the EPS sample

S. no.	Formula	Structure	Biological properties	References
ALPHA-D-MANNOSE	C6H12O6		Mannose plays a critical role in the process of glycosylation, which is a kind of post-translational modification that occurs in proteins. It plays a role in the process of attaching sugar molecules to proteins, which affects their function and stability	[31]
ALPHA-D-GLUCOSE	C6H12O6		The oxidation of glucose yields carboxylic acids like pyruvic acid	[38]
BETA-D-GLUCOSE	C6H12O6		The oxidation of glucose yields carboxylic acids like pyruvic acid	[38]

Emulsifying Activity Study

The emulsification capability of L. amylovorus MTCC 8129 EPS was compared to that of a synthetic surfactant, Tween 20, and a commercially available bacterial biopolymer, Xanthan gum. The mean ± standard deviation (SD) was used to assess the experimental data from all triplicate testing. EPS was discovered to be capable of stabilizing different oils made from vegetables by forming a hydrophobic phase (Table 3). For all of the studied vegetable oils, the EPS generated by L.amylovorus MTCC 8129 was shown to be more effective than the synthetic surfactant employed as a control. L.amylovorus MTCC 8129 EPS was shown to have higher emulsifying activity than commercial bacterial EPS xanthan gum. Only olive oil and sesame oil had lower emulsifying actions. L.amylovorus MTCC 8129 EPS formed a stable emulsion with five different food-grade vegetable oils. (Coconut 54 ± 0.25%, almond oil 32.66 ± 0.25%, olive oil 52.66 ± 1.20%, groundnut oil 32 ± 0.97%, and sesame 35.33 ± 0.65%) [25].

Table 3.

The emulsifying ability of Lactobacillus amylovorus MTCC 8129 EPS in various vegetable oils was compared to commercial EPS biopolymer and synthetic emulsifiers

S.no.	Wavelength cm−1	Functional Group	References
1	3300.25 cm−1	Hydroxyl group (O–H)	[33]
2	2924.08 cm−1	Methylene group -CH2-	[33]
3	1635.06 cm−1 and 1552.23 cm−1	C=O Carbonyl group	[34]
4	1220.94 cm−1	C–O–C and C–O–H bonds	[36]
5	1012.62 cm−1	Carbohydrate C–O stretching	[36]
6	804.31 cm−1	β-glycoside	[37]

Water-Holding and Oil-Holding Capacity Determination

L.amylovorus MTCC 8129 EPS has a water retention capacity (WHC) of 180%. The ANOVA table revealed statistically significant differences (p < 0.05) when compared to the control, xanthan gum, which demonstrated stronger stability and affinity with water, with an oil holding capacity of 163.3%. Oil-holding capacity (OHC) allows the carbohydrate polymer chains to be permeable. The OHC of the L. amylovorus MTCC 8129 EPS was found to have more capacity than commercial norms [32].

Discussion

The production of L.amylovorus MTCC 8129 was conducted in this investigation. The presence of EPS was verified using both quantitative and qualitative analysis [39]. Analysis of the total sugar/carbohydrate tests indicated that the EPS generated consisted of a variety of sugars. The formation of lactic acid and organic acids has been shown to impede microbial growth either directly in their dissociated state or indirectly by releasing protons into the media. Lactic acid is a metabolic waste product generated during the fermentation of bacterial carbohydrates. Lactic acid bacteria (LAB) were found to restrict the proliferation of harmful microorganisms during the process of food fermentation [40]. EPS is a probable heteropolysaccharide consisting of diverse neutral sugars connected by different types of bonds, together with certain sugar units [41]. The ability of EPS to endure high temperatures may be elucidated by its three-stage thermal disintegration, with the first stage including the loss of water. At this temperature, the second phase occurs as a result of the comparatively elevated glucose level and the presence of its sugar units, leading to the anticipated thermal decomposition of the fractions. These studies have shown that heat degradation occurs in two stages, like the EPS process [42]. The subsequent stage is assessing the thermal stability of EPS by comparing it to the highest temperature at which EPS degradation occurs. The hypothesised composition of the EPS indicates a C6 backbone with a galactose sidechain, and the EPS exhibits a Man/Gal ratio [43]. The properties of bacterial EPS may be used in the food industry to improve the structural texture of food as emulsifiers, gelling agents, suspending agents, and stabilisers. Moreover, probiotic bacteria are non-pathogenic, making them appropriate for a wide range of uses in food companies [44]. A heteropolysaccharide produced by L.amylovorus MTCC 8129, can serve as a suspending agent and gelling substance in the food industry [45] [46].

As a result, this inquiry looked at the effectiveness of the EPS. In antioxidant tests such as ABTS, DPPH, H₂O₂, and FRAP for antioxidant capabilities, the L.amylovorus MTCC 8129 EPS demonstrated improved effectiveness with increasing doses, which is equivalent to results from four Auriculariales for polysaccharides. The DPPH and ABTS assays revealed that a concentration of 0.5–2.5 mg/mL of EPS resulted in 70–80% scavenging activity. [47, 48]. The efficacy of H₂O₂ as an antioxidant was shown to be more pronounced at larger concentrations. At a radical scavenging activity of 72%, all approaches exhibited a plateau in activity as the concentration of EPS increased. This suggests that there is a saturation point based on the previously provided EPS concentration values. At a concentration of 2.5 mg/mL, the EPS exhibited its highest activity, resulting in an antioxidant activity of 76–78%. [48]. Moreover, the presence of antioxidants in a food product is very advantageous since these compounds effectively inhibit the oxidation of vitamins and other nutrients, thereby extending the product's shelf life during preservation. The antioxidant activity has significant promise for use as an antioxidant agent and functional supplement within the food sector. Consequently, substances with enhanced antioxidant potential are used as food additives to serve as antioxidants [32].

The emulsification activity experiment demonstrates that the EPS derived from L.amylovorus MTCC 8129 has the ability to emulsify the molecular structure of fatty acids found in both food and food-grade vegetable oils [49]. The findings validate that the emulsifying capability of L.amylovorus MTCC 8129 EPS surpasses that of the commercially available EPS, xanthan gum. The spatial durability of EPS is accountable for the stabilisation of emulsion as a result of its broad creation of interconnected networks. The emulsification feature is crucial in the food, dairy, beverage, and meat processing businesses. The findings indicate that the EPS derived from L.amylovorus MTCC 8129 has the ability to effectively stabilize emulsions of both oil-in-water and water-in-oil nature, making it a promising ingredient for use in diverse sectors of the food industry. The meal stabilised with EPS acquires thickening properties as a result of the presence of large macromolecular structures in the water-based medium. [50].

In our study, the EPS produced by L.amylovorus MTCC 8129 demonstrated superior activity (140–180%) compared to both WHC and OHC in this report [51]. The low molecular weight and low porosity result in the low WHC of EPS. According to Levine and Slade, the crispiness of the chips can be highly improved while using a low percentage of WHC [52]. This suggests that the current study can be highly applicable in food baking to maintain texture under low moisture conditions [53]. This research marks the initial documentation of the bacterial L.amylovorus MTCC 8129 EPS investigation that aims to showcase the possibilities of utilization for food-related purposes [54].

Conclusions

The current study concludes that the EPS produced by L. amylovorous MTCC 8129 can be used for different food technological applications as it possesses antioxidant properties and an emulsifying nature. This is the first report of L. amylovorous MTCC 8129 for EPS generation and food industry application.

Author’s Contribution

JM planned and carried out the experiments. KNR conceived the study, helped in the preparation of the manuscript and assisted in the interpretation of the results. All authors provided critical feedback and approved the final manuscript.