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Article

Isolation and Mechanistic Characterization of Pediococcus pentosaceus WQ-30 from Kimchi for Efficient In Vitro Purine Nucleoside Degradation Relevant to Hyperuricemia

by
Qi Wu
1,
Yibin Wang
1,
Zifu Ni
1,*,
Zhongke Sun
1,
Siyuan Bai
2 and
Le Wang
1,2,*
1
School of Biological Engineering, Institute of Biomass Science and Engineering, Henan University of Technology, Zhengzhou 450001, China
2
Institute for Complexity Science, Henan University of Technology, Zhengzhou 450001, China
*
Authors to whom correspondence should be addressed.
Foods 2026, 15(5), 816; https://doi.org/10.3390/foods15050816
Submission received: 29 January 2026 / Revised: 16 February 2026 / Accepted: 20 February 2026 / Published: 27 February 2026
(This article belongs to the Section Food Biotechnology)
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Abstract

Hyperuricemia (HUA) is a metabolic syndrome caused by elevated levels of uric acid (UA) serum, posing a significant threat to human health. Lactic acid bacteria degrade or adsorb UA precursors such as purine nucleosides and metabolites. By inhibiting intestinal nucleoside absorption, UA synthesis is reduced and HUA alleviated. A total of 60 fermented food samples and 20 soil samples were collected for screening. Strains were selected based on their inosine and guanosine degradation efficiency, and all degradation assays were performed in triplicate (n = 3). We isolated a strain that efficiently degrades inosine and guanosine at rates of 93.99% and 98.88%, respectively. This strain was identified as Pediococcus pentosaceus (P. pentosaceus) via 16S rDNA sequencing and named WQ-30. Whole-genome assembly yielded one chromosome and one plasmid, with 1705 coding sequences. The key gene rihC, encoding a nucleoside hydrolase, was identified through gene functional annotation. Heterologous expression and purification confirmed that RihC was approximately 36 kDa. Recombinant RihC exhibited optimal nucleoside hydrolase activity at pH 7 and 37 °C. This study provides a promising strain for functional food development and a mechanistic basis for the application of P. pentosaceus with purine nucleoside degradation and UA-lowering activities.

Graphical Abstract

1. Introduction

Hyperuricemia (HUA) is a metabolic disorder characterized by abnormally elevated blood uric acid (UA) levels due to purine metabolism dysfunction [1]. Disrupted purine metabolism or excessive dietary purine rapidly raises serum urate and reshapes the gut microbiota [2,3]. With the increasing prevalence of poor lifestyles, unhealthy diets and metabolic syndrome [4], the incidence of HUA has been gradually rising, posing a serious threat to human health. Currently, the primary treatment and preventive measures for HUA involve the use of xanthine oxidase (XOD) inhibitors and uricosuric agents [5]. Yet chronic use is expensive [6] and carries serious side effects [7]. Hence, a safe [8], natural and affordable HUA remedy is urgently needed [9]. To fill that gap, this study focuses on the functional strain Pediococcus pentosaceus (P. pentosaceus) WQ-30, which exhibits superior UA-lowering and purine nucleoside degradation activities. This strain may serve as a potential candidate for the development of probiotic interventions against HUA.
Studies have revealed that probiotics can regulate gut microbiota [10], degrade exogenous purine nucleosides, and inhibit XOD activity, thereby effectively lowering UA levels and alleviating HUA [11,12]. For instance, Lactobacillus acidophilus F02 exhibited a 94% degradation rate toward inosine in vitro [13]. Pediococcus acidilactici GR-5 reduced serum UA levels by 52.17% in vivo [14]. Similarly, Pediococcus acidilactici SWU-HX39 decreased serum UA by 49.8% and inhibited XOD activity by 46.76% in vivo [15]. Although purine-degrading and UA-lowering strains have gained increasing attention [16,17], only a limited number of probiotics have been well validated in vivo or show potential clinical applications for HUA alleviation [18]. It is therefore necessary to screen for highly efficient UA-lowering strains. Compared with other P. pentosaceus strains widely used in food fermentation and probiotic products [19], the strain WQ-30 investigated in this study exhibits nucleoside degradation activity, which further expands the application potential of this strain in food and health applications.
Current studies show that probiotics degrade nucleosides via two main enzymatic systems: purine nucleoside phosphorylase and nucleoside hydrolase [20]. The former converts inosine and guanosine into hypoxanthine, guanine, and ribose-1-phosphate via phosphorolysis, while the latter catalyzes the hydrolysis of nucleosides into bases and ribose [21]. Although studies on nucleoside hydrolase systems have been reported in lactic acid bacteria [22], the corresponding mechanisms in P. pentosaceus remain unclear. Specifically, key pathway gaps remain to be elucidated in its regulatory mechanism, including transcriptional regulators, signal induction pathways, and crosstalk with purine catabolism. Meanwhile, the degradation characteristics, coding gene sequences, catalytic properties, and regulatory mechanisms of its nucleoside hydrolases have not been fully elucidated [23]. Moreover, few studies have directly linked P. pentosaceus’ metabolic functions to its UA-lowering activity, leading to unclear specificity and targeting. Additionally, wild-type strains generally exhibit low activity of key enzymes, and heterologous expression systems for key enzymes have not been established. These engineering limitations impede functional verification and strain optimization, restricting the strain’s application potential and failing to meet industrial demands for efficient functional strains. Thus, constructing high-efficiency expression vectors via genetic engineering to improve enzyme activity and expression has become an important research direction in this field [24].
Against this backdrop, in-depth exploration of key genes and clarification of the specific purine nucleoside degradation pathway in P. pentosaceus have emerged as a critical breakthrough. That helps advance research in this field from basic exploration to practical application. Additionally, overcoming the activity limitations of wild-type strains and constructing high-efficiency functional strains via technical means is another key breakthrough. Based on the above analysis, we propose the hypothesis that the nucleoside hydrolase encoded by rihC is functionally responsible for inosine and guanosine degradation in P. pentosaceus. It aims to screen and evaluate the UA-lowering strain P. pentosaceus WQ-30, identify and annotate the key gene rihC by whole-genome analysis, and verify its function through heterologous expression in vitro, thereby providing theoretical support and technical pathways for developing high-activity UA-lowering probiotic resources and low-purine foods.

2. Material and Methods

2.1. Materials

Sample sources: A total of 60 naturally fermented pickle samples were collected in Zhengzhou, Henan Province, China, starting from 10 October 2024, with a sampling duration of approximately one month. These samples included homemade pickles and homemade-like pickles purchased from local supermarkets, which were mainly made from white radish and Chinese cabbage via spontaneous anaerobic fermentation without inoculation of exogenous strains. These fermented products were naturally dominated by lactic acid bacteria, mainly including Lactobacillus and Pediococcus. All samples were fermented to maturity and stored at a low temperature after collection.
A total of 20 soil samples were collected in Zhengzhou, Henan Province, China, starting from 10 October 2024, with a sampling duration of approximately one month. The samples were obtained from local vegetable planting areas in the Huanghuai Plain, which is dominated by cinnamon soil with stable physicochemical properties and serves as an important natural habitat for lactic acid bacteria. Surface soil at 0–20 cm depth was collected using a sterile sampler. After removing impurities, the soil was placed into sterile sampling bags and stored immediately at low temperature for the subsequent isolation and screening of lactic acid bacteria, including Lactobacillus, Pediococcus and Weissella.
Culture media: Man, Rogosa and Sharpe (MRS) Agar medium (g/L): beef extract 10.0, peptone 10.0, yeast extract 5.0, glucose 20.0, sodium acetate 5.0, diammonium citrate 2.0, K2HPO4 2.0, MgSO4 0.58, MnSO4 0.25, Tween-80 1.0, and agar 20.0 (for solid medium). Calcium carbonate was added to form transparent zones for the preliminary screening of lactic acid bacteria. pH 6.2 ± 0.2; 121 °C, 15 min. LB medium (g/L): tryptone 10.0, yeast extract 5.0, NaCl 10.0. pH 7.0 ± 0.2; 121 °C, 15 min. Nutrient solution (g/L): Arabinogalactan 1 g, pectin 2 g, xylan 1 g, potato starch 3 g, glucose 0.4 g, yeast extract 3 g, peptone 1 g, mucin 4 g, L-cysteine 0.5 g. Simulated gastric fluid (SGF): 3.33 g/dL pepsin, pH 2 or 3. Simulated intestinal fluid (SIF) (g/L): NaHCO3 12.5, bile salts 6.0, pancreatin 0.9, pH 6.8. Fasted state: SGF (pH 2) and SIF without nutrient supplementation. Fed state: SGF (pH 3) and SIF supplemented with nutrients.
Reagents: Beef extract, peptone, yeast extract, tryptone, agar, porcine bile salt: Beijing Aoboxing Biotechnology Co., Ltd., Beijing, China, all biochemical reagents. Glucose, MgSO4, NaHCO3: Tianjin Kemiou Chemical Reagent Co., Ltd., Tianjin, China, all analytical-grade. Sodium acetate, diammonium citrate, K2HPO4, MnSO4, NaCl: Tianjin Tianli Chemical Reagent Co., Ltd., Tianjin, China, all analytical-grade. Tween-80: Tianjin Kemiou Chemical Reagent Co., Ltd., Tianjin, China, chemically pure. Pepsin (≥250 95 U mg−1), trypsin (≥10,000 U mg−1): Shanghai Shifeng Biotechnology Co., Ltd., Shanghai, China, biochemical reagents, NaOH, HCl, H3PO4: Sinopharm Chemical Reagent, Shanghai, China, all analytical-grade. Potassium phosphate, phenolphthalein, sterile defibrinated sheep blood: Hope Bio-Technology, Qingdao, China. L-cysteine: Tianjin Huasheng Chemical Reagent Co., Ltd., Tianjin, China, Mucin: Shanghai Shifeng Biotechnology Co., Ltd., Xylan: Shanghai Yuanye Bio-Technology Co., Ltd., Shanghai, China, biochemical reagent. Arabinogalactan and pectin: Xinxiang Sanwei Disinfectant Preparations Co., Ltd., Xinxiang, China, analytical-grade. Xanthine, allopurinol, XOD, inosine, guanosine, and isopropyl β-D-1-thiogalactopyranoside (IPTG): Shanghai Macklin Biochemical Co., Ltd., Shanghai, China, biochemical reagent. DNA extraction kit, plasmid extraction kit, purification kit: Sangon Biotech (Shanghai) Co., Ltd., Shanghai, China.
Equipment: LDZX-50KBS vertical autoclave (Shen’an Medical Device, Shanghai, China). TG20KR-D high-speed refrigerated centrifuge (Thermo Fisher Scientific, Waltham, MA, USA). Multiskan FC microplate reader (Hao xing Biotechnology, Xi’an, China). LC-20A HPLC system (Shimadzu, Kyoto, Japan). Universal Hood II gel documentation system (Bio-Rad, Hercules, CA, USA). JY600C electrophoresis apparatus (Junyi Oriental, Beijing, China). XP Cycler PCR thermal cycler (Bioer, Hangzhou, China). Ni-NTA affinity chromatography column (Sangon Biotech, Shanghai, China).
Strains: Staphylococcus epidermidis, E. coli, and P. pentosaceus were obtained from the laboratory stock collection.

2.2. Screening and Identification of Lactic Acid Bacterial Strains with Purine Nucleoside Degradation and UA-Lowering Activity

Using a sterile loop, pick 0.1 g of test sample and mix it with 9 mL of sterile water. Incubate the mixture at 37 °C for 20 min, and then take 100 μL of the sample and prepare serial dilution solutions (10−1 to 10−6) with sterile water. Pipette 200 μL of each dilution and spread them separately on the MRS solid medium containing 1.0% calcium carbonate, followed by anaerobic incubation at a constant temperature of 37 °C for 48 h. Single colonies were isolated via the streak plate method and purified repeatedly until morphologically uniform colonies were obtained. Gram staining was conducted on the colonies, and their growth curves were determined.
According to preliminary experiments and the relevant literature [13], the bacterial strain was inoculated in MRS broth and incubated at 37 °C to the logarithmic growth phase. An inosine–guanosine neutral potassium phosphate solution was prepared and stored in a 4 °C refrigerator for later use. Pipette 2 mL of shaken bacterial culture, centrifuge at 4000 r/min for 10 min, and wash the precipitate with PBS three times. The cells were adjusted to a uniform concentration of 1 × 109 CFU/mL and resuspended in 750 μL of the inosine–guanosine solution. After incubating in a 37 °C incubator for 1 h, the reaction was terminated by heating in a 100 °C water bath for 5 min. Centrifuge at 8000 r/min and 4 °C for 10 min, take the supernatant, filter through a 0.22 μm sterile membrane, and perform HPLC analysis. Identify inosine and guanosine retention times via the external standard method. Plot a standard curve with the concentration (x-axis) and peak area (y-axis). Chromatographic conditions: Liquid chromatograph LC-20A, reverse-phase column Agilent-C18 (4.6 mm × 250 mm), mobile phase V (water): V (methanol): 90:10, flow rate: 1 mL/min, detection wavelength: 254 nm, column temperature: 30 °C. Genomic DNA of the strains was extracted using a bacterial genomic DNA extraction kit. PCR amplification was performed using primers 27F (5′-AGAGTTTGATCCTGGCTCAG-3′) and 1492R (5′-TACGGCTACCTTGTTACGACTT-3′). The reaction conditions were as follows: 95 °C 3 min, 95 °C 15 s, 54 °C 15 s, 72 °C 23 s, 30 cycles, 72 °C 5 min. The PCR products were sent for sequencing. The obtained 16S rDNA sequences were subjected to NCBI BLAST+version 2.17.0 analysis on the NCBI website https://www.ncbi.nlm.nih.gov/#!/landingpage (accessed on 18 December 2024) to compare the homology with the sequencing information of known strains. MEGA 7.0 was used to perform multiple sequence alignment analysis of the sequences of the strains selected in this study and the reference strain sequences retrieved from the NCBI database.

2.3. Characteristics of the P. pentosaceus Strain with Purine Nucleoside Degradation and UA-Lowering Activity

2.3.1. Surface Hydrophobicity Analysis of the P. pentosaceus Strain

The cell surface hydrophobicity of WQ-30 was determined according to the method described by Reuben et al. [25]. Cells were harvested by centrifugation at 4000× g for 10 min and resuspended in PBS to an optical density at 600 nm (OD600) of 0.4 (A0), and 1 mL of chloroform was mixed with 3 mL of bacterial suspension. After vortexing at 3000 rpm for 10 s, the mixture was incubated statically at room temperature for 1 h. The absorbance (Ax) of the upper aqueous phase was measured. A negative control was included using sterile PBS to account for spontaneous sedimentation during the assay. All measurements were performed in triplicate and calculated according to Formula (1):
H y d r o p h o b i c i t y   ( % ) = A 0 − A X A 0 × 100

2.3.2. Autoaggregation Ability Analysis of the P. pentosaceus Strain

The autoaggregation ability was determined according to the method described by Reuben et al. [26]. Bacterial suspensions were prepared as described in Section 2.3.1 and adjusted to OD600 = 0.4 (A0). After vortexing at 3000 rpm for 10 s, 4 mL of the suspension was incubated statically at room temperature for 3 h. The OD600 of the upper suspension was then measured (At). A negative control was included using sterile PBS to account for spontaneous sedimentation during the assay. All measurements were performed in triplicate and calculated according to Formula (2):
A u t o a g g r e g a t i o n % = A 0 − A t A 0 × 100

2.3.3. Acid and Salt Tolerance of the P. pentosaceus Strain

According to a previous method with slight modifications [27], bacterial suspensions were prepared as described in Section 2.3.1, and resuspended in PBS. Inoculate into MRS with pH values of 2.5, 3.0, 3.5, and 4 at an inoculum size of 5%. After 3 h cultivation, OD600 was determined. Inoculate the bacterial solution in MRS containing 0.3% and 0.5% bile salts. The OD600 were measured at 1 h, 3 h, and 5 h, with normal MRS as the control.

2.3.4. Simulated Gastrointestinal Fluid Tolerance Test of the P. pentosaceus Strain

According to a previous method with slight modifications [28], the bacterial suspension was prepared as described in Section 2.3.1. Here, 1 mL of the suspension was mixed with 9 mL of SGF (fed and fasted states) and incubated with shaking. After 3 h, 1 mL was transferred to 9 mL SIF (fed and fasted states) and incubated for another 3 h. After serial dilution, the suspension was spread onto agar plates and incubated under anaerobic conditions at 37 °C for 48 h. Tolerance to simulated gastrointestinal fluids was evaluated based on the colony counts. A control group was treated with sterile PBS instead of simulated gastrointestinal fluids and incubated under the same conditions.

2.3.5. XOD Activity Inhibition Assay of the P. pentosaceus Strain

To clarify whether the XOD inhibitory effect was derived from live bacterial cells, extracellular metabolites or intracellular components, three samples were prepared: cell-free supernatant (CFS), resting bacterial suspension, and cell-free extract (CFE), according to a previous method with slight modifications [29].
CFS: This step involved centrifuging 10 mL of log-phase lactic acid bacteria suspension at 6000 rpm for 10 min. The collected supernatant was designated as the MRS fermentation CFS.
Preparation of bacterial suspension: The cell pellet was washed three times with sterile PBS, resuspended to OD600 = 1.0, and incubated at 37 °C for 12 h.
CFE: A log-phase cell suspension was sonicated on ice for 10 min (300 W, 5 s on, 5 s off). After centrifugation at 8000 rpm for 5 min, the supernatant was filtered through a 0.22 μm filter to obtain CFE. XOD inhibition rates of CFS, bacterial suspension, and CFE were determined. A reaction mixture containing 1.5 mL xanthine solution (0.20 g/L), 1.5 mL sample, and 2 mL XOD solution (0.20 U/mL, pre-incubated at 37 °C for 20 min) was monitored at 295 nm using a microplate reader. The linearity of the assay at 295 nm was validated, and corresponding background controls were included. In the control group, XOD solution was replaced with sterile PBS buffer. Background absorbance (without samples or enzymes) was subtracted to eliminate non-specific interference, including potential absorption from bacterial metabolites, from xanthine absorbance at 295 nm. All samples were assayed in triplicate, with allopurinol as the positive control. The calculation using Formula (3) for the XOD inhibitory rate is as follows:
I n h i b i t i o n   R a t e % = ( 1 − C − D A − B ) × 100
where A is the absorbance of the solution containing XOD but no sample; B is the absorbance of the solution containing neither XOD nor sample; C is the absorbance of the solution containing both XOD and sample; and D is the absorbance of the solution containing the sample but no XOD.

2.3.6. Safety Evaluation of the P. pentosaceus Strain

Hemolysis of the P. pentosaceus Strain
According to a previous method with slight modifications [30], the test strains were inoculated on sterile defibrinated sheep blood agar plates. Using Staphylococcus epidermidis as the control, the strains were spot-inoculated at the center of the plate. The test lactic acid bacteria were spot-inoculated evenly three times around the control strain. After incubation at 37 °C for 48 h, the hemolysis of lactic acid bacteria and the control strain was observed.
Antibiotic Resistance of the P. pentosaceus Strain
According to a previous method with slight modifications [31], the antibiotic susceptibility of candidate strains was determined using the disk diffusion method (Table S1). The bacterial suspension in the logarithmic growth phase was serially diluted, and 100 μL of it was pipetted and spread onto MRS solid medium. After the surface of the plate dried, antibiotic susceptibility disks were placed at the center of each Petri dish. The plates were incubated under anaerobic conditions at 37 °C for 48 h. The diameter of the inhibition zone around each disk was measured individually to determine the antibiotic susceptibility.

2.4. Whole-Genome Sequencing and Genome Annotation of the P. pentosaceus Strain

The DNA of the bacterial strain culture was extracted by Sangon Biotech (Shanghai) Co., Ltd., and the complete genome map was determined. For the raw data generated by sequencing, data statistics, quality assessment, and quality trimming were performed using Fast QC 0.11.2. Quality-controlled Nanopore reads were assembled and error-corrected with Canu v1.3 to eliminate clipping artefacts and small indels. The circular genome map was drawn with CGView v1.0 to display genes on both strands, clusters of orthologous groups (COG) functional categories, GC content and genome size. Functional annotation was performed on the protein-coding sequences, and functional genes related to UA metabolism were screened out. Prokka 1.10 software was used to predict rRNA, tRNA, gene structures in the bacterial genome, open reading frames, and protein-coding genes. The sequencing data were aligned to 11 databases, including the Kyoto Encyclopedia of Genes and Genomes (KEGG) and Gene Ontology (GO), to complete gene functional annotation. Pan-genome analyses were performed using Roary and LS-BSR according to the standard bioinformatics pipeline.

2.5. Sequence Analysis of RihC Protein for Purine Nucleoside Degradation and UA-Lowering Activity in P. pentosaceus WQ-30

The protein sequence of the key gene rihC was retrieved from the NCBI database. Based on the obtained protein sequence, we further conducted secondary structure modeling using the SWISS-MODEL server https://swissmodel.expasy.org/interactive (accessed on 20 June 2025). To further verify the structural characteristics of the protein sequence encoded by the key gene rihC, we conducted a detailed sequence alignment search on the Protein Data Bank (PDB) website https://www.rcsb.org/ (accessed on 20 June 2025). After completing the above steps, we used MEGA 7.0 software to conduct an in-depth sequence alignment analysis of the protein sequence encoded by the key gene rihC and its aligned sequences. To present the results of the protein sequence alignment more intuitively, we further performed protein sequence alignment using the ESPript 3.0 tool https://espript.ibcp.fr/ESPript/ESPript/ (accessed on 20 June 2025).

2.6. Extraction and Verification of the Key Gene rihC

PET-28a was selected as the expression vector, and the genome of strain WQ-30, which has strong abilities to degrade inosine and guanosine as well as high tolerance, was used as the template. PCR amplification was performed using the primers RihC-F1: 5′-ATGTCAACAAAGATTATCATGGACACTGACCCTGG-3′ and RihC-R1: 5′-TTATTTAGGTTGGTCGATGGCTGCCACG-3′, while homologous recombination gene amplification was conducted with the primers RihC-F2: 5′-CAGCAAATGGGTCGCGGATCCATGTCAACAAAGATTATCATGGACACTGAC-3′ and RihC-R2: 5′-ACGGAGCTCGAATTCGGATCCTTATTTAGGTTGGTCGATGGCTGCC-3′. The reaction conditions were as follows: 95 °C 3 min, 95 °C 15 s, 55 °C 15 s, 72 °C 23 min, 30 cycles, 72 °C 5 min. Plasmid pET28a was extracted using a plasmid extraction kit and subjected to single-enzyme digestion with BamH I. After being transformed into E. coli DH5α for enzyme digestion verification, it was introduced into E. coli BL21(DE3) competent cells, followed by colony PCR identification. The successfully identified single colonies were then inoculated in LB medium containing kanamycin (Kan) and cultured at 37 °C for 12 h. Finally, the degradation rates of the constructed strains were determined using inosine and guanosine as substrates.
The constructed strain was cultured with IPTG induction. The strain was inoculated into LB medium containing Kan (50 μg/mL) and cultured at 37 °C for 4 h. Then, IPTG was added, and the culture was continued at 37 °C for 20 h. The cells were collected for ultrasonic disruption, and the recombinant protein was detected by SDS-PAGE. The purified enzyme was collected and purified using nickel column affinity chromatography. The ability of crude enzyme and purified enzyme to degrade inosine and guanosine was determined with inosine and guanosine as substrates. Using the non-constructed strain as the control, the degradation rates were determined according to the method described in Section 2.2. The purified enzyme activity was measured at 20–60 °C (pH 7.0) and pH 4–9, respectively, to investigate its optimal temperature and pH.

3. Results and Discussion

3.1. Screening and Probiotic Properties of Strains

In this study, a total of 398 strains were isolated and identified from fermented pickles and soil, with 198 lactic acid bacterial strains being screened. Inosine and guanosine degradation rates for some strains are shown in Table S2. Finally, one strain of lactic acid bacteria with high degradation efficiency was selected, exhibiting degradation rates of 93.99% for inosine and 98.88% for guanosine. Compared with the 94.26% for inosine and 60% for guanosine degradation reported for Lactobacillus acidophilus F02 [13], our strain showed superior activity. This may be attributed to the high substrate affinity and catalytic efficiency of key nucleoside hydrolases in WQ-30. In addition, the efficient purine metabolic pathway of this strain may also contribute to its superior guanosine degradation capacity compared with previously reported lactic acid bacteria. Collectively, the strain’s exceptional guanosine degradation capacity addresses the inadequate guanosine metabolism in previously reported lactic acid bacteria, offering a valuable resource for targeted UA biodegradation strategies.
On MRS solid medium, the strain formed small, milky-white colonies with smooth, rounded, and opaque edges. The colonies exhibited a convex center that was easily picked up with an inoculation loop (Figure 1a). Gram-staining images of the colonies are shown in Figure S1. These morphological characteristics align with the typical features of P. pentosaceus as reported in the literature [32]. The growth pattern can be observed through the growth curve (Figure 1b). The strain grew slowly during 0–2 h, followed by rapid proliferation in the 2–12 h exponential growth phase, when the fermentation broth pH dropped sharply to 3.91 at 12 h. After 12 h, the strain entered the stationary phase with slowed growth due to nutrient depletion and accumulation of acidic substances, and the broth pH continued to decrease slowly.
16S rDNA sequencing combined with NCBI BLAST homology alignment revealed 99% similarity between the strain and P. pentosaceus. Therefore, the strain was identified as belonging to the Pediococcus genus and named P. pentosaceus WQ-30. Additionally, a phylogenetic tree was constructed using MEGA software (Figure 1c). Phylogenetic analysis placed WQ-30 within the P. pentosaceus clade, confirming its species identity. While grouping with P. pentosaceus-type strains, WQ-30’s basal phylogenetic position implies potential genetic and functional divergence. Collectively, WQ-30 exhibited robust acidogenic and rapid-growth traits, highlighting its potential as a food fermentation starter.
Surface hydrophobicity and autoaggregation of WQ-30 determine its initial contact with host cells, offering a rapid indicator of epithelial adhesion [33]. WQ-30 exhibited 45.7% hydrophobicity and 40.3% autoaggregation, indicating modest adhesive potential (Figure 2a). This characteristic is crucial for its colonization in the host gastrointestinal tract, thereby laying a foundation for its UA-lowering function in vivo.
After 3 h in 0.3% and 0.5% bile salts, WQ-30 showed no significant difference in survival rate (OD600) compared to the control group (Figure 2b). This demonstrates that WQ-30 has good bile salt tolerance and can effectively adapt to variations in bile salt concentration. After 3 h at a pH above 4.0, the OD600 of the strains remained around 0.4. After 3 h at pH values of 2.5 and 3.0, cultures stabilized at an OD600 of approximately 0.2, demonstrating certain acid tolerance (Figure 2c). Similarly, the P. pentosaceus eLab 60WB strain was able to survive and grow in low-pH MRS medium after 4 h [34]. These acid and bile salt tolerance properties, together with its adhesion potential, confirm that WQ-30 can withstand the harsh gastrointestinal environment.
The viable count of WQ-30 in fasted SGF was approximately 1 × 102 CFU/mL, while that in fed SIF reached 1 × 106 CFU/mL (Figure 2d). The gastric environment exhibits pH fluctuations between 1.5 and 3.0 due to dietary influences. When bacterial strains pass through intestinal fluid, bile salts and trypsin exert inhibitory effects on them [35]. Therefore, the excellent tolerance of WQ-30 to gastrointestinal fluids ensures its survival during passage through the gastrointestinal tract, which is a prerequisite for probiotics to exert their functions.
Further in vitro XOD inhibition assays were conducted on WQ-30 and its extracts (CFE and CFS). WQ-30 showed a 40% inhibition rate at 10 min of reaction (Figure 3a), while WQ-30 CFE and CFS both peaked in inhibitory activity at 8 min (Figure 3b,c). These results indicate that both WQ-30 and its extracts exhibit XOD inhibitory activity. XOD is the key enzyme catalyzing the oxidation of xanthine to UA, and inhibiting XOD activity can reduce UA production. Consistent with related findings [22], WQ-30 also efficiently degraded inosine and guanosine, with degradation rates of 93.99% and 98.88%, respectively. As core purine nucleosides, the degradation of inosine reduces the production of hypoxanthine, thereby cutting off the upstream source of xanthine. Notably, all these results were obtained in vitro. Although nucleoside degradation and XOD inhibition may theoretically contribute to UA lowering by reducing the substrate supply and blocking key catalytic steps, the potential synergism and actual effects of these two pathways under in vivo conditions remain to be further verified. Therefore, WQ-30 and its extracts exhibit XOD inhibitory activity and purine nucleoside degradation capacity in vitro, supporting their potential UA-lowering properties. This effect may be attributed to their metabolites or cellular components, which could bind to XOD and block its catalytic reaction.
When lactic acid bacteria are ingested by humans or animals, their hemolytic activity should be evaluated to ensure safety. Compared with Staphylococcus aureus and E. coli, WQ-30 exhibited γ-hemolysis (non-hemolysis) (Figure S2). Probiotics exhibiting γ-hemolysis are widely recognized as safe. Antibiotic susceptibility testing is a prerequisite for safe strain application. As shown in Table S3, WQ-30 exhibited differential sensitivity to various antibiotics, with high sensitivity to penicillin, ampicillin, tetracycline, erythromycin, and chloramphenicol (inhibition zone > 17 mm) and low sensitivity to ciprofloxacin and lincomycin. This is consistent with the antibiotic susceptibility profile characteristics of common lactic acid bacteria [36].
Taken together, probiotic characterization and safety assessment confirm that WQ-30 can effectively withstand the gastrointestinal environment and maintain sufficient viability. In vitro assays revealed that WQ-30 possesses both efficient purine nucleoside degradation and XOD inhibitory activities, which may act as complementary mechanisms underlying its uric acid-lowering potential. This comprehensive evaluation demonstrates that WQ-30 combines favorable gastrointestinal adaptability, reliable safety, and promising functional activity, making it a highly promising candidate for developing functional foods targeting HUA.WQ-30 outperforms existing strains with targeted UA-lowering capacity via robust inosine/guanosine degradation and stable probiotic traits. Its functional advantages may be associated with the regulation or expression differences in specific metabolic genes. To clarify the molecular basis of this characteristic, subsequent whole-genome analysis will be conducted to identify core functional genes.

3.2. Whole-Genome Sequence Analysis of the P. pentosaceus Strain

To characterize the metabolic pathway by which WQ-30 degrades inosine and guanosine, whole-genome sequencing was undertaken. As shown in Figure 4, assembly of the high-quality filtered reads for WQ-30 yielded one chromosome and one plasmid (Figure S3), with lengths of 1,795,115 bp and 6052 bp, and G+C contents of 37.33% and 40.98%, respectively. The genome contained 1705 coding sequences (CDSs), 15 rRNAs, and 55 tRNAs, with protein-coding genes accounting for approximately 92.81% of the genome. Compared to other strains of P. pentosaceus [37], WQ-30 exhibits significant differences in plasmid size and GC content, which may be closely related to its unique metabolic functions. Notably, WQ-30 demonstrates remarkable UA-lowering activity. Although plasmid-related genes may be speculated to contribute to this phenotype, such a functional link remains hypothetical and requires further experimental validation. Further functional validation and mechanistic studies could provide deeper insights into the genetic basis of its UA-lowering activity and the role of these genes in UA-lowering activity. Similar to Lactobacillus plantarum, SQ001 was found to harbor enzyme-coding genes involved in nucleotide hydrolysis [38]. WQ-30 may utilize distinct enzyme systems to mediate purine nucleoside degradation. Therefore, comprehensive functional annotation of the WQ-30 genome will enable us to uncover the strain-specific genes and metabolic pathways that drive its distinctive traits.
Functional annotation of the WQ-30 proteome against the COG database assigned 1287 genes to 20 functional categories, accounting for 75.48% of all predicted coding sequences (Figure 5). This high annotation ratio indicates that the WQ-30 genome encodes an extensive functional repertoire, providing a solid basis for survival and metabolism across diverse environments. Among these, 123 genes were annotated for carbohydrate transport and metabolism, 159 genes for general function prediction, and 143 genes for translation, ribosomal structure, and biogenesis. Additionally, 92 genes were associated with amino acid transport and metabolism, and 122 genes were annotated for transcription functions. These highly represented metabolic pathways are all essential for bacterial metabolism, indicating that WQ-30 possesses broad metabolic capabilities and adaptability. Compared to other known strains of P. pentosaceus [39], WQ-30 exhibits some unique features in gene functional annotation. For example, it carries substantially more carbohydrate transport and metabolism genes than P. pentosaceus CECT 8330 [40], a trait linked to its adaptation to carbohydrate-rich fermentation conditions. Additionally, its markedly higher number of transcription-related genes suggests enhanced flexibility in gene expression regulation, collectively bolstering its survival across diverse environments. These findings establish a framework for WQ-30 metabolism and provide a reference for subsequent gene mining and pathway analysis.
The protein-coding gene sequences of WQ-30 were compared and analyzed through the GO database, resulting in the annotation of 13,642 genes (Figure 6). A total of 832 genes were annotated to biological processes, 188 to cellular metabolic processes, and 354 to molecular functions, with the remaining genes assigned to other GO categories. In molecular function annotation, the pathway with the highest functional abundance is catalytic activity, and cellular and metabolic processes are the predominant biological processes represented. Notably, WQ-30 encodes a large number of catalytic activity-related genes, which are closely associated with inosine and guanosine degradation, and these genes encode metabolic enzymes that underpin the strain’s capacity for purine nucleoside metabolism, thereby supporting the hypothesis that WQ-30 harbors specialized pathways for inosine and guanosine catabolism.
The protein-coding gene sequences of WQ-30 were aligned using the KEGG database (Figure 7). A total of 1042 coding genes were annotated, primarily categorized into cellular processes (38 genes), environmental information processing (104 genes), genetic information processing (164 genes), metabolism (536 genes), and organismal systems (17 genes). Genes involved in metabolism are the most abundant, indicating that the strain possesses an extensive suite of metabolic genes closely linked to its survival and adaptability. This is followed by genetic information processing, where nucleotide metabolism is directly associated with inosine and guanosine metabolism. Through KEGG analysis, specific coding genes participating in inosine and guanosine metabolism can be identified. These genes are predicted to function in various biological processes, including nucleotide synthesis, degradation, and transport. Specifically, we identified genes encoding purine-metabolic enzymes such as ribonucleoside hydrolase, which play crucial roles in the degradation of inosine and guanosine.
Overall, whole-genome sequencing and functional annotation of WQ-30 identified abundant metabolism-related genes, with enrichment in purine metabolism and catalytic activity. These genomic features provide solid support for the strain’s excellent inosine/guanosine degradation capacity and UA-lowering potential. The identification of enzyme-encoding genes involved in the purine nucleoside degradation pathway has initially established a link between genomic characteristics and phenotypic advantages. Combined with the previously characterized probiotic traits, these genomic data lay a solid foundation for subsequent identification of core functional genes via comparative genomics and elucidation of the UA-lowering mechanism.

3.3. Comparative and Pan-Genome Analysis of Eight P. pentosaceus Strains

Comparative genomic and pan-genomic analyses were performed on eight strains of P. pentosaceus (Figure 8a). The numbers of genes identified in P. pentosaceus WQ-30, SL 4, JQI 7, SRCM 102736, SRCM 102734, FDAARGOS 1011, FDAARGOS 1009 and FDAARGOS 1134 were 69, 93, 19, 78, 86, 56, 30, and 60, respectively. Most strain-specific genes are hypothetical proteins. The pan-genome analysis revealed the presence of 1361 core genes and 499 specific genes. The heatmap provides a binary view of gene presence, with color coding that directly distinguishes present versus absent genes (Figure 8b), facilitating the identification of UA-lowering activity-related genes among strains. This is crucial for screening potential UA-lowering microbial resources and offers new insights into the mechanisms underlying this activity. Furthermore, by comparing the conservation and variability of these genes among different strains, we can further explore their expression and function under various environmental conditions, as well as their potential for practical applications. Collectively, these analyses confirm distinct genetic diversity across the eight P. pentosaceus strains, with core and strain-specific genes establishing a framework for investigating functional specialization linked to UA-lowering traits.

3.4. Analysis of Potential Virulence Factors and Antibiotic Resistance Genes

Virulence factors refer to the properties that enable microorganisms to establish themselves within a specific host species and enhance their potential to cause disease. In the genome of WQ-30, there is one adhesion-related gene, one immunomodulatory gene, and there are eight genes encoding stress proteins (Table S4). These genes endow WQ-30 with certain adhesion and stress response capabilities, but do not imply significant pathogenicity. Preliminary hemolytic tests confirmed the lack of expression of these virulence genes, consistent with functional validation results of bacterial virulence factors reported in other studies. For example, when studying Enterococcus faecalis, it was found that its virulence factors are mainly associated with adhesion, invasion, and evasion of host immune defenses [41]. Despite harboring adhesion and immunomodulatory genes, WQ-30 exhibits limited virulence expression, which may be linked to its ecological niche and host adaptability. The WQ-30 genome harbors genes encoding resistance to fluoroquinolones, clindamycin, and related antimicrobials (Table S5).
This study’s analysis demonstrates that the genes encoding antibiotic resistance proteins in this strain are chromosomally located. Therefore, these genes are unlikely to be transferred or disseminated to other pathogenic bacteria. At the molecular level, this proves that WQ-30 lacks the capacity to transmit or receive antibiotic resistance genes, which is further supported by prior drug susceptibility tests. Through characterization of the strain’s properties, the results confirm that the strain exhibits good safety and maintains certain sensitivity to antibiotics. These characteristics indicate that the P. pentosaceus strain not only performs excellently under laboratory conditions but also possesses potential for practical application in real-world environments. These safety data lay a critical foundation for subsequent exploration of its purine nucleoside degradation and UA-lowering associated genes.

3.5. Genes for Purine Nucleoside Degradation and UA-Lowering Activity in P. pentosaceus WQ-30

This study conducted whole-genome sequencing and annotation of WQ-30 to explore the molecular mechanisms underlying its UA-lowering activity. Given that purine metabolism constitutes a complex process multifactorial [42], we further analyzed relevant pathways using the KEGG database (Figure S4).
As a critical junction in the purine metabolic pathway, inosine and guanosine metabolism was the focus of our analysis. Whole-genome analysis of WQ-30 identified key genes involved in these pathways, thereby clarifying the molecular mechanism underlying its UA-lowering activity. As shown in Table 1, we characterized the roles of several key enzymes in the degradation pathways of inosine and guanosine. The ribonucleoside hydrolase gene rihC (EC:3.2.2.-) encodes an enzyme that efficiently hydrolyzes inosine and guanosine to yield the corresponding bases and ribose. Additionally, the adenosine deaminase encoded by the add gene can convert adenosine to inosine, which is subsequently involved in inosine degradation. These two genes synergistically block the direct purine nucleoside-to-UA conversion pathway. Meanwhile, hpt directly converts hypoxanthine to inosine monophosphate (IMP) while purH converts other intermediates to IMP via de novo purine synthesis. Together, they channel free purine bases into the purine nucleotide synthesis cycle, reducing their conversion to UA and enabling purine recycling. This process lowers UA production by reducing substrates at the source and intercepting intermediates, forming the core molecular basis for the strain’s UA-lowering activity.
These enzymes play crucial roles in the purine metabolic pathway, and their activities are finely regulated by the intracellular metabolic state. Their synergistic action endows WQ-30 with functional nucleoside degradation pathways, thereby maintaining purine metabolic homeostasis. Notably, the rihC gene stands out as the most critical functional determinant, and this central role provides a rationale for its subsequent selection in heterologous expression studies. While previous studies have reported the role of other microorganisms in UA-lowering activity [43], our research has revealed several novel genes associated with this trait, providing new clues for future investigations. These newly discovered elements may play pivotal roles in UA-lowering activity processes, offering promising targets for developing more efficient biodegradation strategies. To further elucidate its function, subsequent studies will focus on characterizing the RihC hydrolase from P. pentosaceus.

3.6. Characteristics of the RihC Hydrolase from P. pentosaceus

Nucleoside hydrolases degrade ribonucleosides by catalyzing the hydrolysis of β-N-glycosidic bonds, yielding free bases and ribose. Through genome annotation and screening, we identified a key gene responsible for inosine and guanosine degradation, designated rihC. Multiple amino acid sequence alignment was performed on RihC nucleoside hydrolase with 8QND [44], 3G5I [45], 1MAS [46], and 1EZR [47]. This analysis revealed that residues critical for β-N-glycosidic bond cleavage (His232, Asp10, Asp14, Asp15, Asp233, Asn165) are highly conserved across nucleoside hydrolases (Figure 9). Consistent with nucleoside hydrolase catalytic mechanisms, His232 acts as a proton donor, activating the leaving group via protonation of substrate base-specific sites (purine N7/pyrimidine N3) to drive bond cleavage. Asp residues (Asp10/Asp14/Asp15) coordinate Ca2+ via their side-chain carboxyl groups to activate water molecules and stabilize the catalytic microenvironment, forming a Ca2+-Asp residue synergistic catalytic core. Asn165 stabilizes the oxocarbenium ion transition state, lowering the glycosidic bond cleavage energy barrier. These conserved residues, as core active site units of RihC, collectively mediate specific β-N-glycosidic bond cleavage. It is further confirmed that RihC has the ability to degrade inosine and guanosine. Furthermore, we determined the crystal structure of RihC. The asymmetric unit of the crystal contains four nearly identical protein subunits and belongs to space group P21, indicating that the protein exists as a tetramer in the crystal. We further modeled the tertiary structure of RihC [44] (Figure S5). These structural features not only confirm the function of RihC as a nucleoside hydrolase but also reveal its unique structural characteristics, which are crucial for understanding its role in UA metabolism.
To thoroughly investigate the role of the rihC gene in WQ-30’s UA-lowering activity, we performed overexpression validation. PCR amplification yielded the full-length 910 bp rihC gene (Figure S6), further confirming the presence of the rihC fragment in the strain. As shown in Figure S7, the single-colony PCR verification profile demonstrated successful amplification of the target band using RihC-F1/RihC-R1 primers. The rihC fragment was successfully ligated with plasmid pET-28a. Restriction digestion of the extracted recombinant plasmid yielded a band corresponding to the expected size, confirming correct assembly (Figure S8). The verified plasmid was then transformed into E. coli BL21(DE3). SDS-PAGE analysis showed a clear RihC band at approximately 36 kDa (Figure 10a), consistent with literature reports and confirming successful expression [48]. In Figure 10b, the reduced band intensity indicated effective removal of contaminating proteins during purification, thereby enhancing the purity of the target protein.
HPLC assays showed that the constructed strain achieved degradation rates of 99.67% for inosine and 84.33% for guanosine. Compared with the control strain, these degradation rates exhibited statistically significant differences (Figure 10c), confirming successful integration of the rihC gene fragment into E. coli BL21 (DE3). This functional validation via heterologous expression not only confirms the catalytic activity of RihC but also supports its physiological role in purine nucleoside degradation in native P. pentosaceus WQ-30. Additionally, the Bacillus parametracis YC06 strain can efficiently degrade inosine and guanosine. Its whole-genome sequencing revealed that the degradation mechanism involves multiple key enzyme genes [49]. These findings indicate that introducing specific enzyme genes through genetic engineering can also confer purine nucleoside degradation capability. The degradation rates of inosine and guanosine by the crude enzyme and purified enzyme exhibited significant differences compared with those of the control group (Figure 10d).
Since variations in temperature and pH are inevitable in complex fermentation environments, higher tolerance can reduce the cost of fermentation condition control. In this study, the degradation rates of inosine and guanosine were used to reflect the activity variation trend of the enzyme under different conditions. These support the analysis of RihC’s enzymatic properties. The effect of different pH values on the activity of the purified enzyme was determined (Figure 10e). As the pH increased from 4 to 7, the activity of the enzyme gradually increased, and the degradation rates of inosine and guanosine also progressively rose. The enzyme exhibited optimal activity at pH 7. Subsequently, as the pH increased from 7 to 9, the enzyme’s activity gradually declined. At pH 4 and 9, the enzyme retained certain activity, indicating its tolerance to both acidic and alkaline conditions. The relative activity of the enzyme was measured at 20–60 °C (Figure 10f). The enzyme activity gradually increased with rising temperature, and the degradation rates of inosine and guanosine also progressively rose. Furthermore, the enzyme still retains a certain level of activity at 20 °C. The highest activity was observed at 37 °C, followed by a rapid decline with further temperature increases, until complete inactivation occurred at 60 °C. In this study, we focused on temperature and pH effects on recombinant RihC activity, as these are critical for fermentation performance [50,51]. The enzyme showed good adaptability to pH/temperature fluctuations; its high tolerance reduces fermentation control costs, supporting potential industrial or probiotic applications.
In summary, this study clarified that the RihC hydrolase encoded by the rihC gene in P. pentosaceus WQ-30 mediates purine nucleoside degradation through conserved active sites and a tetrameric structure. Its heterologous expression and enzymatic characterization not only confirmed the gene’s core function in UA lowering but also provided support for its application in practical fermentation environments. Subsequent detailed enzymatic studies will be conducted on the RihC hydrolase.

4. Conclusions

This study isolated and identified the P. pentosaceus WQ-30 strain, which exhibits degradation activity toward inosine and guanosine. Through in-depth characterization, whole-genome sequencing, and functional annotation, we identified rihC as the potential key gene mediating purine nucleoside degradation and UA-lowering activity. Under in vitro experimental conditions, the rihC engineered strain exhibits certain UA-lowering efficacy, which further validates the role of this gene and its corresponding enzyme in UA-lowering function. Taken together, these findings establish a clear and integrated relationship among the strain, its key gene, the encoded enzyme, and the UA-lowering function.
Notably, this study has certain limitations. On the one hand, the validation and analysis of UA-lowering activity were based on in vitro experiments, lacking supporting data from in vivo animal studies and human clinical trials. On the other hand, the application of this strain in real food matrices has not been evaluated. Its survival stability, functional activity, and fermentation adaptability in food systems remain unclear, which cannot directly support its subsequent industrial application.
Future research will focus on four aspects: First, 30 mice with HUA will be used to verify the in vivo UA-lowering efficacy. Main endpoints include serum uric acid, liver XOD activity, and hepatic/renal function indexes. Second, the application adaptability of the strain in actual food matrices will be explored, and industrial fermentation processes will be optimized to promote its large-scale production and transformation. Third, long-term safety evaluations will be carried out in accordance with regulatory standards for functional foods, and compliance-related data will be improved to remove obstacles for commercialization. Fourth, with the rihC gene as the core, gene editing and enzyme engineering will be adopted to enhance the UA efficacy and application potential of the strain. Through the above research, the UA research system of P. pentosaceus WQ-30 will be gradually improved, providing theoretical and technical support for the application of lactic acid bacterial resources in the field of UA-lowering functional foods.

Supplementary Materials

The following supporting information can be downloaded at https://www.mdpi.com/article/10.3390/foods15050816/s1, Figure S1: Gram staining image; Figure S2: Hemolysis image; Figure S3: Plasmid map of WQ-30; Figure S4: Inosine and guanosine metabolic pathways; Figure S5: The tertiary structure of RihC; Figure S6: PCR amplification bands of the rihC gene; Figure S7: Colony PCR amplification results; Figure S8: Single enzyme digestion analysis of homologous recombination plasmid; Table S1: Types, potencies of antimicrobial discs and interpretive criteria for inhibition zone diameters; Table S2: Degradation rates of inosine and guanosine of different strains; Table S3: Sensitivity of strains to antibiotics; Table S4: Potential virulence factors of the strain; Table S5: Identification of antibiotic resistance-related genes in lactic acid bacteria strains.

Author Contributions

Q.W.: writing—review and editing, writing—original draft, visualization, software. Y.W.: investigation, data curation, conceptualization. Z.N.: writing—review and editing, writing—original draft, validation. Z.S.: writing—review and editing, project administration. S.B.: data curation, conceptualization. L.W.: writing—review and editing, investigation, project administration, data curation. All authors have read and agreed to the published version of the manuscript.

Funding

This work was supported by the National Natural Science Foundation of China (21978070); the Youth Found of the Natural Science Foundation of Henan Province (252300423562); Major Science and Technology Projects in Henan Province (251100110300); the Tuoxin Talent Cultivation Program of Henan University of Technology (2025); and the Program for the Top Young Talents of Henan Association for Science and Technology (2023).

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

The original contributions presented in this study are included in the article/Supplementary Materials. Further inquiries can be directed to the corresponding authors.

Conflicts of Interest

The authors declare no competing interests.

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Figure 1. Screening of strain Pediococcus pentosaceus (P. pentosaceus) WQ-30. (a) Morphology of WQ-30 on MRS agar. (b) Growth production curves of WQ-30. (c) Acid production curves of WQ-30. (d) Phylogenetic tree of WQ-30. Error bars represent mean ± SD (n = 3).
Figure 1. Screening of strain Pediococcus pentosaceus (P. pentosaceus) WQ-30. (a) Morphology of WQ-30 on MRS agar. (b) Growth production curves of WQ-30. (c) Acid production curves of WQ-30. (d) Phylogenetic tree of WQ-30. Error bars represent mean ± SD (n = 3).
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Figure 2. Characterization of strain P. pentosaceus WQ-30. (a) Hydrophobicity and autoaggregation of WQ-30. (b) Bile salt tolerance of WQ-30. (c) Acid tolerance of WQ-30. (d) Gastrointestinal tolerance of WQ-30. Error bars represent mean ± SD (n = 3). The P-value indicates statistical significance, with “ns” denoting no significant difference. p < 0.001 is considered to indicate a significant difference (** p < 0.01, **** p < 0.0001, # p < 0.05, ## p < 0.01, #### p < 0.0001).
Figure 2. Characterization of strain P. pentosaceus WQ-30. (a) Hydrophobicity and autoaggregation of WQ-30. (b) Bile salt tolerance of WQ-30. (c) Acid tolerance of WQ-30. (d) Gastrointestinal tolerance of WQ-30. Error bars represent mean ± SD (n = 3). The P-value indicates statistical significance, with “ns” denoting no significant difference. p < 0.001 is considered to indicate a significant difference (** p < 0.01, **** p < 0.0001, # p < 0.05, ## p < 0.01, #### p < 0.0001).
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Figure 3. XOD inhibition rate of WQ-30. (a) XOD inhibition rate of WQ-30 cell suspension. (b) XOD inhibition rate of WQ-30 CFE. (c) XOD inhibition rate of WQ-30 CFS. Error bars represent mean ± SD (n = 3).
Figure 3. XOD inhibition rate of WQ-30. (a) XOD inhibition rate of WQ-30 cell suspension. (b) XOD inhibition rate of WQ-30 CFE. (c) XOD inhibition rate of WQ-30 CFS. Error bars represent mean ± SD (n = 3).
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Figure 4. The chromosomal map of the WQ-30 gene.
Figure 4. The chromosomal map of the WQ-30 gene.
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Figure 5. Completed whole-genome sequencing of WQ-30 COG annotation.
Figure 5. Completed whole-genome sequencing of WQ-30 COG annotation.
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Figure 6. Completed whole-genome sequencing of WQ-30 GO annotation.
Figure 6. Completed whole-genome sequencing of WQ-30 GO annotation.
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Figure 7. Completed whole-genome sequencing of WQ-30 KEGG annotation.
Figure 7. Completed whole-genome sequencing of WQ-30 KEGG annotation.
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Figure 8. Comparative analysis of gene annotations for 8 P. pentosaceus strains. (a) The Venn diagram illustrates the genomes of P. pentosaceus strains, showing different numbers of unique genes and shared genes. (b) Through comparison with the genomes of these P. pentosaceus strains, the pangenome exhibits a set of core genes and accessory genes. Dark green indicates the presence of a gene cluster, and white indicates its absence. The large dark green block represents the core genome, while the variable regions represent the accessory genome.
Figure 8. Comparative analysis of gene annotations for 8 P. pentosaceus strains. (a) The Venn diagram illustrates the genomes of P. pentosaceus strains, showing different numbers of unique genes and shared genes. (b) Through comparison with the genomes of these P. pentosaceus strains, the pangenome exhibits a set of core genes and accessory genes. Dark green indicates the presence of a gene cluster, and white indicates its absence. The large dark green block represents the core genome, while the variable regions represent the accessory genome.
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Figure 9. Multiple sequence alignment of the RihC protein. Note: 8QND, Hydrolase from Limosilactobacillus reuter; 3G5I, Hydrolase from Escherichia coli K-12; 1MAS, Hydrolase from Crithidia fasciculata; 1EZR, Hydrolase from Leishmania major. The key residues in the RihC active site are marked with *. The identical amino acid residues are shown in red color. The blue boxes delineate the conserved secondary structural elements in the aligned sequences. Special positions such as insertions, deletions, or non-standard residues are marked with #. The secondary structure elements (α-helices and β-strands) of P. pentosaceus RihC are indicated above the alignment.
Figure 9. Multiple sequence alignment of the RihC protein. Note: 8QND, Hydrolase from Limosilactobacillus reuter; 3G5I, Hydrolase from Escherichia coli K-12; 1MAS, Hydrolase from Crithidia fasciculata; 1EZR, Hydrolase from Leishmania major. The key residues in the RihC active site are marked with *. The identical amino acid residues are shown in red color. The blue boxes delineate the conserved secondary structural elements in the aligned sequences. Special positions such as insertions, deletions, or non-standard residues are marked with #. The secondary structure elements (α-helices and β-strands) of P. pentosaceus RihC are indicated above the alignment.
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Figure 10. Characteristics of RihC hydrolase. (a) SDS-PAGE analysis of expression products. (b) SDS-PAGE analysis of the purified enzyme. (c) The degradation rates of inosine and guanosine by the engineered strain and the control strain. (d) Degradation rates of inosine and guanosine by crude enzyme and purified enzyme. (e) The effect of pH on the activity of the purified enzyme. (f) The effect of temperature on the activity of the purified enzyme. The boxed of (a,b) region indicates the position of the target protein band. Error bars represent mean ± SD (n = 3). The p-value indicates statistical significance, with “ns” denoting no significant difference. P < 0.001 is considered to indicate a significant difference. (* p < 0.05, ** p < 0.01, *** p < 0.001, **** p < 0.0001, ### p < 0.001, #### p < 0.0001).
Figure 10. Characteristics of RihC hydrolase. (a) SDS-PAGE analysis of expression products. (b) SDS-PAGE analysis of the purified enzyme. (c) The degradation rates of inosine and guanosine by the engineered strain and the control strain. (d) Degradation rates of inosine and guanosine by crude enzyme and purified enzyme. (e) The effect of pH on the activity of the purified enzyme. (f) The effect of temperature on the activity of the purified enzyme. The boxed of (a,b) region indicates the position of the target protein band. Error bars represent mean ± SD (n = 3). The p-value indicates statistical significance, with “ns” denoting no significant difference. P < 0.001 is considered to indicate a significant difference. (* p < 0.05, ** p < 0.01, *** p < 0.001, **** p < 0.0001, ### p < 0.001, #### p < 0.0001).
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Table 1. Analysis of genes related to UA metabolism.
Table 1. Analysis of genes related to UA metabolism.
Gene IDGene NameProductEC Anno
Chrom1_000191rihCribonucleoside hydrolase RihCEC:3.2.2.-
Chrom1_000557addadenosine deaminaseEC:3.5.4.4
Chrom1_001407purHbifunctional phosphoribosyl aminoimidazole carboxamide formyl transferase/IMP cyclo hydrolaseEC:2.1.2.3 EC:3.5.4.10
Chrom1_001495hpthypoxanthine phosphoribosyl transferaseEC:2.4.2.8
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Wu, Q.; Wang, Y.; Ni, Z.; Sun, Z.; Bai, S.; Wang, L. Isolation and Mechanistic Characterization of Pediococcus pentosaceus WQ-30 from Kimchi for Efficient In Vitro Purine Nucleoside Degradation Relevant to Hyperuricemia. Foods 2026, 15, 816. https://doi.org/10.3390/foods15050816

AMA Style

Wu Q, Wang Y, Ni Z, Sun Z, Bai S, Wang L. Isolation and Mechanistic Characterization of Pediococcus pentosaceus WQ-30 from Kimchi for Efficient In Vitro Purine Nucleoside Degradation Relevant to Hyperuricemia. Foods. 2026; 15(5):816. https://doi.org/10.3390/foods15050816

Chicago/Turabian Style

Wu, Qi, Yibin Wang, Zifu Ni, Zhongke Sun, Siyuan Bai, and Le Wang. 2026. "Isolation and Mechanistic Characterization of Pediococcus pentosaceus WQ-30 from Kimchi for Efficient In Vitro Purine Nucleoside Degradation Relevant to Hyperuricemia" Foods 15, no. 5: 816. https://doi.org/10.3390/foods15050816

APA Style

Wu, Q., Wang, Y., Ni, Z., Sun, Z., Bai, S., & Wang, L. (2026). Isolation and Mechanistic Characterization of Pediococcus pentosaceus WQ-30 from Kimchi for Efficient In Vitro Purine Nucleoside Degradation Relevant to Hyperuricemia. Foods, 15(5), 816. https://doi.org/10.3390/foods15050816

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