The materials used for the study include FSI (20210820, Shandong Bozhou R&B Food Sales Co., Ltd., China); L.f (GDMCC 1.1796), Lactobacillus acidophilus (L.a; GDMCC 1.208) and Lactobacillus plantarum (L.p; GDMCC 1.140) (Guangdong Province Microbial Strain Conservation Center, China); Inulin (food grade) (China Food & Moon Health Technology Co.); Glucose, Papain, Glutathione Peroxidase (GSH-PX) assay kit (Colorimetric method), Superoxide Dismutase (SOD) assay kit (WST-1 method), Lactate dehydrogenase (LDH) assay kit, total protein quantitative assay kit (Thomas Brilliant Blue method) and DNS reagent (Shanghai Yuanye Biotechnology Co., Ltd., China); MRS broth, MRS medium (with agar), and MRS broth (without glucose) (Guangdong Haibo Microbial Technology Co., China).

An appropriate amount of degreased FSI powder was weighed with 120 mesh and the PFSI was extracted using an ultrasonic-microwave assisted method with the parameters of microwave power 500 W, ultrasonic power 270 W, extraction time 21 min, and material-liquid ratio 1:95. It was cooled at room temperature and centrifuged for 10 min (8,000 rpm/min). The supernatant was transferred and concentrated to obtain the polysaccharide concentrate. The polysaccharide concentrates were then deproteinated using the Sevage method, papain, and TCA method, respectively, and the polysaccharide retention ratio (Y₁) and protein removal ratio (Y₂) were analyzed. Ultimately the overall results were evaluated by an overall score (0.5 Y₁+ 0.5 Y₂), which was optimized to select the best deproteination process corresponding to various deproteination methods. The polysaccharide concentrates were obtained and placed in molecular weight cut-off (MWCO) 3,500 Da dialysis bags and were treated in running water for 48 h. To the solution of polysaccharide supernatant, 4 times the volume of anhydrous ethanol was added and refrigerated at 4°C overnight. This was followed by washing the lower layer of the precipitate with anhydrous ethanol several times and it was desiccated under decompression. The concentration of the dried products was set at 100 mg/mL and eluted with 200 mL of distilled water, and 0.1, 0.2, and 0.3 mol/L of sodium chloride solutions separately at a filtration rate of 1 mL/min. The solvents were collected, and the polysaccharides were purified, whereas the polysaccharide concentrates without deproteinization were prepared with the above purification operation as PFSI-1. The determination of total saccharide content was conducted by the phenol-sulfuric acid method (Ren et al., 2019). It was calculated that the polysaccharide yield of PFSI-1 after purification was 56.62 ± 1.77%. The determination of protein content was referenced to the instructions of the Kemas Brilliant Blue kit. The extraction, isolation, and purification process of different PFSI are illustrated in Figure 1. The yield of polysaccharides was measured as follows:

Where Y is the yield of polysaccharide (%), V is the total volume of sample solution (mL), C is the concentration of polysaccharide (mg/mL), D is the dilution factor, and m is the mass of the weighed sample (g).

The above polysaccharide sample solution was prepared as 10 mg/mL, and distilled water was used as the control for scanning at a full wavelength within 200 ~ 600 nm.

The fully dried KBr and polysaccharide samples were blended and ground with an agate mortar in a sufficiently dry environment at a mass ratio of 100: 1–200: 1. The slices were pressed and scanned with an infrared spectrometer in the range of 400–4,000 cm⁻¹.

The PFSI was identified based on the Molisch reaction precipitation method (Pawar et al., 2018) and the 3,5-dinitrosalicylic acid (DNS) method according to the standard procedure reported by Alexander et al. (2011) and McCleary and McGeough (2015).

The MRS culture was sterilized at 121°C for 15 min, then the plates were inverted, and the L.f (L.a or L.p) lyophilized powder was inoculated into the MRS modified medium plates in a sterile environment, incubated at 37°C until 36 h. After two generations of incubation, a single L.f (L.a or L.p) colony was picked for two generations of liquid culture, and subsequently, a liquid bacterial suspension was fabricated.

The medium was prepared to contain dissolved polysaccharides at a mass concentration of 2.5 g/L using various purification levels of PFSI, inulin, and glucose, as the carbohydrate source, with ultrasonic-assisted dissolution. After sterilization at 121°C and chilling to ambient temperature, the insoluble substances were removed by filtration through a membrane. The inoculum was 5% 1.5×10⁸ CFU/ml L.f suspension and cultivated at 37°C for 36 h. The OD₆₀₀ was measured at 0 h and 36 h. The positive controls were inulin and glucose, and the negative control was the glycose-free medium. The samples were tested three times simultaneously. The biomass was measured by centrifuging the suspension at 3,500 r/min for 10 min, after which the bacteria were gathered and washed twice with phosphate buffer. Using phosphate buffer as a blank control, the absorbance value of the suspension was assessed at 600 nm, which indicated the biomass by absorbance value.

The same process as in 2.7 was adopted. The carbohydrate source for preparing the MRS medium was dissolved by ultrasonication at 1.25, 2.5, 5.0, and 10 g/L to obtain four groups of optimal carbon sources with different concentrations. OD₆₀₀ was measured after 36 h of incubation of L.f , L.a, and L.p. The negative control was a glycose-free MRS medium (concentration of 0 g/L). Each sample was tested three times simultaneously.

The same process as in 2.7 was used to attain the optimal carbohydrate source. As the carbohydrate sources for the formulation of MRS medium, ultra-sonication to solubilization, which were 1.25, 2.5, 5.0, and 10 g/L, yielded four groups of different concentrations of the optimum carbohydrate sources. The OD₆₀₀ was measured at 0 h, 6, 12, 18, 24, 30, and 36 h. The negative control group was glycose-free MRS medium. Each sample was tested three times simultaneously.

The fermentation solutions were cultured in the optimal carbohydrate source at 0 h, 6, 12, 18, 24, 30, and 36 h by centrifugation at 6,000 r/min for 10 min, and then the supernatant fermentation broth was withdrawn and set aside. The method described in McCleary and McGeough (2015) was referred to and modified, the standard curve was plotted, and the regression equation y = 0.8319x−0.0828 was calculated with associated coefficient R² = 0.9998. To determine the reducing saccharide content in the fermentation medium using the DNS method, 1 mL of each was taken in the proportion of DNS reagent: the supernatant (V/V) = 1:1, and left to react fully in a boiling water bath for 5 min, taken out and placed in an ice bath to cool down rapidly. The absorbance was measured at 540 nm. Distilled water was applied as a blank control group and the optimum concentration of carbohydrate source of 10 g/L was adopted as the experimental group and the negative control group was glycose-free MRS medium. Each sample was tested three times simultaneously.

The measurement was determined using the kit instructions, by taking 50 uL of supernatant fermentation broth obtained above and kneading it with 3 mL of Kemas Brilliant Blue col-or solution and letting it stand for 10 min to determine the OD value at 595 nm. Distilled water was used as a blank control group and 0.524 g/L of the standard was considered a positive control group. Each sample was tested three times in parallel. The following formula was used for the calculation.

In the formula, OD₀ refers to the absorbance value of the blank group at 595 nm, OD₁ refers to the absorbance value of the positive control group at 595 nm, OD₂ refers to the absorbance value of the sample to be measured at 595 nm, C₁ refers to the concentration of the standard protein, and D refers to the dilution multiple.

The fermentation supernatant was obtained above. The lactate content was determined by the p-hydroxyphenyl method detailed in Meng et al. (2017), and the standard curve was plotted to obtain the regression equation y = 0.0554x−0.0278 with the correlation coefficient R² = 0.9997. Each sample was tested three times simultaneously.

LDH, SOD, and GSH-Px dynamics were measured in the fermentation solution obtained above. Sequentially to the instructions for the experiment, the simultaneous tests were repeated three times.

Data are shown as mean ± SEM, and statistical analysis was performed using Origin 2021, Graphpad Prism 8, and EXCEL 2019. To compare the differences between multiple groups, a one-way analysis of variance (ANOVA) was performed in this software. P < 0.05 indicates statistical significance.

The absence of obvious absorption peaks at 260 nm and 280 nm in Figure 5 indicates that PFSI-2, PFSI-3, and PFSI-4 do not contain impurity components such as nucleic acids and proteins, but the absorption peak is obvious around 260 nm in PFSI-1 which is not purified by deproteinization.

The IR spectra of PFSI-1, PFSI-2, PFSI-3, and PFSI-4 are illustrated in Figure 6. The peaks in the range of 3,600~3,000 cm⁻¹, 2,900 cm⁻¹, 1,600~1,400 cm⁻¹, and 1,200~700 cm⁻¹ are all typical polysaccharide absorption peaks. Among them, the absorption peaks in the range of 3,600~3,000 cm⁻¹ suggest the possible presence of polysaccharides. The broad and strong absorption peak is the O-H functionality absorption peak, where the absorption peak is created by the hydroxyl group on the glycan chain. The waveforms of the peaks are wide and the wave numbers move to lower frequencies because of the presence of a large number of hydroxyl groups on the polysaccharide chain and the formation of a hydrogen bond within or between the molecules. The fainter absorption peak around 2,900 cm⁻¹ is caused by the C-H stretching vibration. The more pointed absorption peaks around 1,600 cm⁻¹ are (carboxyl or acid amine groups) C=O stretching vibrations and N-H variable angle vibrations, denoting absorption peaks caused by asymmetric carbonyl stretching. In combination with the UV spectrum, the absorption peaks occurred due to the stretching vibration of the carboxylate ion (–COO–) in the samples. The absorption peaks at around 1,400–1,200 cm⁻¹ are attributed to the stretching vibration of C=C. The larger absorption peak between 1,200 and 1,000 cm⁻¹ arises from the C-OH stretching vibration (Xiang et al., 2014). In this way, the characteristic absorption peaks at around 800 cm⁻¹ are not only characteristic of α-type differential isomers but also caused by the stretching vibration of C=O on the pyranose ring, signifying that the glycan chain of the sample contains a pyranose ring structure, confirming the presence of a-D-pyranose (C₁-H) as well as α and β-configuration glycosidic bonds. From the analysis of each absorption peak before 1,000 cm⁻¹, it was initially inferred that the tested samples were saccharide structures. The results of the IR spectra show that PFSI-1, PFSI-2, PFSI-3, and PFSI-4 have the functional groups of polysaccharides as typical polysaccharides.

From Table 4, it can be seen that the Molisch reaction was positive (+), indicating a violet ring appears at the junction of the two liquids as well as the presence of polysaccharides in the samples. The DNS reactions of PFSI-1, PFSI-2, PFSI-3, and PFSI-4 were positive (+), indicating the products of the reaction being brownish red and the presence of reducing polysaccharide in the samples.

As demonstrated in Figure 7, each concentration of carbohydrate sources at 2.5 g/L had increased bacteriophage biomass vs. the blank control (P < 0.05). On the other hand, PFSI-4 was found with more bacteriophage biomass than PFSI-1, PFSI-2, and PFSI-3 (P < 0.05), which may be correlated with the molecular weight of polysaccharides, monosaccharide composition, and structure. When compared with glucose, the bacteriophage biomass grew with no significant difference (P > 0.05), but the aggregated amount was more than that of glucose. In conclusion, PFSI-4 can serve as the best available carbohydrate source for consequential studies.

The biomass of L.f , L.a, and L.p was found to increase significantly (P < 0.05) with increasing PFSI-4 concentration by the 2-fold dilution method. From Figure 8, it can be seen that to some extent it has a prebiotic function on Lactobacillus at 36 h, but the biomass, after acting on L.f, was more than that of L.a and L.p. This may suggest a selective and different intensity of role on various species of Lactobacillus. In a subsequent study, the viability associated with the action of PFSI-4 on L.f will be continued to be studied.

The results are shown in Figure 9, indicating that from 0 h to 30 h, PFSI-4 significantly accelerated the proliferation of L.f quantitatively (P < 0.05). Each concentration of PFSI-4 increased in the logarithmic growth period from 0 to 30 h, while the growth rate started decelerating or even decaying from 30 to 36 h. In summary, PFSI-4 could shorten the delay period of L.f culture, making it enter the logarithmic phase earlier, and accelerate the growth rate of logarithmic phase bacteria.

The impact of PFSI-4 on L.f toward the utilization of reducing saccharides is plotted in Figure 10. The cultures with or without the addition of PFSI-4 had a temporal effect from 0 to 18 h. When L.f gradually entered the logarithmic phase, L.f growth was accelerated, and utilization of reducing saccharide was quickened. From 18 h to 30 h, as the L.f growth rate gradually leveled off, reducing saccharide utilization also gradually made no significant difference (p > 0.05). At 36 h, although the growth rate of L.f had gradually slowed down, reducing saccharide utilization continued to increase, though the rate at which it was increasing dropped. In the above, the reducing saccharide utilization in the experimental group with the addition of PFSI-4 was significantly higher than that in the blank control group, indicating that the addition of PFSI-4 could promote the absorption and utilization of reducing saccharides by L.f (P < 0.05) and facilitate its growth.

The effect of PFSI-4 on protein synthesis is shown in Figure 11. The protein content at all stages of growth was higher than that without PFSI-4, indicating that PFSI-4 could promote L.f protein synthesis (P < 0.05) and improve the physiological activeness of the organism.

Producing acid ability is an essential physiological characteristic of lactobacilli, and lactate is the major metabolite of organoleptic acid in L.f . The changes in lactate content in the fermentation broth reflected the growth status of the bacteria, and the lactate synthesis velocity was enhanced when the bacteria grew faster. From Figure 12, we can conclude that the lactate concentration gradually increased with the increase of fermentation time with or without the addition of PFSI-4, but the lactate content of each culture section with the addition of PFSI-4 was always superior to that without the addition of PFSI-4, which means that PFSI-4 promoted the growth of L.f and increased the lactate content (P < 0.05).

As shown in Figure 13A, LDH catalyzed the conversion of pyruvate to lactate under anaerobic conditions, and the LDH vitality was markedly elevated by the additive PFSI-4 vs. the non-additive carbohydrate source. At 24 h fermentation time, the LDH viability of the experimentally administered group reached approximately the maximum of 68.19 U/mL, and the LDH viability of the blank control group reached approximately the maximum of 45.33 U/mL. PFSI-4 remarkably boosts the viability of LDH (P < 0.05).

From Figures 13B,C, it was observed that SOD and GSH-Px vitality changes achieved maximum vitality at about 18 h. The SOD and GSH-Px vitality of the experimental group was superior to that of the control group at all measured time points. SOD and GSH-Px have been identified as critical antioxidant enzymes in cells, which scale off excess free radicals and defend cells from damage and assume a vital role in the oxidative and antioxidant balance of the organism. The level of SOD and GSH-Px activity under anaerobic conditions represents the strength of the physiological activity of the cells. The addition of PFSI-4 increased the activity of LDH, SOD, and GSH-Px, indicating that PFSI-4 could strengthen the physiological activity of L.f (P < 0.05) and stimulate its growth.