Lacticaseibacillus rhamnosus alleviates hyperuricemia by restricting intestinal nucleoside absorption

Abstract

Hyperuricemia is a growing metabolic disorder, whereas current microbiota-based strategies primarily focus on urate degradation or excretion, leaving upstream regulation of nucleoside precursors largely unexplored. Here, we isolated a nucleoside-degrading probiotic strain, Lacticaseibacillus rhamnosus (L. rhamnosus), from traditional fermented dairy products and demonstrated its urate-lowering effect in a hyperuricemia rat model. The strain significantly reduced serum urate levels without altering renal urate transporter expression, suggesting a mechanism independent of renal excretion. Multi-omics and Caco-2 cell assays revealed that L. rhamnosus expresses intracellular nucleoside hydrolases that convert guanosine and inosine into poorly absorbed purine bases, thereby limiting intestinal nucleoside uptake and reducing hepatic substrate supply for urate synthesis. Two key hydrolases were functionally validated by gene cloning and enzymatic assays. In addition, L. rhamnosus reshaped gut microbial composition, modulated host metabolic pathways, and alleviated systemic inflammation. Collectively, this study identifies a previously uncharacterized substrate-restriction mechanism by which probiotics alleviate hyperuricemia, providing novel insight into microbiota-based strategies for dietary and therapeutic intervention in purine metabolism.

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1. Introduction

Hyperuricemia is a prevalent metabolic disorder worldwide, with a steadily increasing incidence.¹ It has become a significant risk factor for various chronic diseases, including gout, diabetes, chronic kidney disease, and cardiovascular conditions.^2,3 Current clinical interventions primarily rely on pharmacological agents that inhibit uric acid synthesis (e.g., allopurinol, febuxostat) or promote urate excretion (e.g., benzbromarone, diuretics).^4,5 However, these medications are often associated with considerable side effects and poor patient compliance, limiting their long-term safety and widespread application.⁶ Therefore, the development of safer and more sustainable urate-lowering strategies has become a research priority.

In humans, uric acid is the final product of purine metabolism, with its regulation involving multiple processes including hepatic synthesis, intestinal absorption, and renal and intestinal excretion.⁷ Historically, research has primarily focused on renal excretion, aiming to reduce serum urate levels by enhancing renal clearance.⁸ Recently, however, growing evidence has highlighted the role of gut microbiota in regulating host urate metabolism.⁹ Certain lactic acid bacteria have been shown to lower serum urate by inhibiting hepatic xanthine oxidase (XOD) activity or modulating the expression of intestinal urate transporters such as ABCG2.¹⁰ Moreover, some gut microbes can degrade uric acid or its intermediate metabolites, thereby influencing host urate homeostasis.^11,12 In contrast, the metabolic behavior of upstream purine precursors—namely nucleosides such as guanosine and inosine in the gut, as well as their relationship with serum urate levels, remains poorly understood.

Previous studies have demonstrated that nucleosides like guanosine and inosine can be rapidly converted into uric acid in the liver and thus represent important precursors for hyperuricemia.¹³ Notably, nucleosides and their degraded purine bases may exhibit distinct intestinal transport characteristics, which could contribute to differences in their systemic bioavailability and urate burden.¹⁴ This raises the hypothesis that certain gut bacteria capable of degrading nucleosides into less absorbable purine bases before absorption might limit the substrate availability for hepatic urate synthesis, thereby exerting a “substrate-restriction” urate-lowering effect. While some lactic acid bacteria have been suggested to possess nucleoside-degrading activity, the specific enzymatic machinery, metabolic pathways, and their in vivo relevance remain largely uncharacterized.

In this study, we isolated a strain of Lacticaseibacillus rhamnosus (L. rhamnosus) with nucleoside hydrolase activity from traditional fermented dairy products. Using a hyperuricemia animal model, combined with intestinal epithelial transport models, multi-omics approaches, and functional validation, we systematically investigated whether this strain could lower serum urate levels by degrading nucleosides, reducing their intestinal absorption, and limiting the hepatic availability of urate precursors. This work not only uncovers a microbial mechanism distinct from previously described urate degradation or excretion pathways, but also provides new insights into probiotic-based strategies for managing hyperuricemia and expanding their functional repertoire.

2. Materials and methods

2.1. Strain cultivation

L. rhamnosus strains were obtained from Wahaha Group Co., Ltd (Hangzhou, China). The strains were cultured in de Man–Rogosa–Sharpe (MRS) broth under standard culture conditions at 37 °C for 12 h. After incubation, bacterial suspensions were centrifuged at 5000 rpm for 10 min at 4 °C, and the supernatants were discarded. Pellets were washed twice with sterile phosphate-buffered saline (PBS) and resuspended in PBS. Cell concentrations were determined by plate counting and adjusted to the required levels. All preparations were freshly made and verified via 16S rRNA gene sequencing prior to use.

2.2. Rat model

All animal procedures were approved by the Laboratory Animal Ethics Committee of Dr Can Biotechnology (Zhejiang, China; Approval No. 2023DRK0726). Eight-week-old male Sprague-Dawley (SD) rats (200–230 g) were obtained from Hangzhou Haowo Biotechnology and housed under specific pathogen-free (SPF) conditions with ad libitum access to food and water. After a one-week acclimatization, rats were randomly assigned to experimental groups, and body weight and food intake were recorded regularly.

Hyperuricemia was induced using a high-purine diet (20% yeast extract) combined with oral administration of potassium oxonate (300 mg kg⁻¹ day⁻¹) for 3 weeks.¹⁵ Control rats received a standard diet and PBS. Treatment groups were orally administered 200 μL of L. rhamnosus suspension (1.5 × 10⁸ CFU mL⁻¹) or allopurinol (20 mg kg⁻¹) daily, while vehicle controls were given PBS. All histopathological evaluations were performed by an investigator blinded to the treatment groups to avoid subjective bias.

2.3. Cell lines and culture conditions

Caco-2 cells were purchased from the Cell Bank of the Chinese Academy of Sciences (Shanghai, China) and cultured in high-glucose Dulbecco's Modified Eagle Medium (DMEM; Gibco, USA) supplemented with 20% fetal bovine serum (FBS), 100 μg mL⁻¹ penicillin, and 100 μg mL⁻¹ streptomycin. Cells were maintained in a humidified atmosphere at 37 °C with 5% CO₂. For intestinal barrier modeling, cells were seeded onto Transwell inserts and cultured for 21 days, with daily medium replacement, to allow for monolayer differentiation. Differentiation and barrier integrity were evaluated by measuring transepithelial electrical resistance (TEER) and alkaline phosphatase (ALP) activity. Only monolayers with TEER values exceeding 500 Ω cm² and showing stable ALP activity were considered fully differentiated and used for subsequent transport assays.¹⁶

2.4. Quantification of uric acid excretion

After an overnight fast, blood was collected via the tail vein, while urine and feces were obtained from rats housed in individual metabolic cages. Serum was separated by centrifuging blood at 6000 rpm for 15 min at 4 °C. Serum uric acid levels were analyzed using an automated biochemical analyzer (IDEXX Laboratories, Westbrook, USA). Urinary uric acid was measured with a commercial assay kit (Nanjing Jiancheng Bioengineering Institute, China). Fecal uric acid concentrations were assessed using high-performance liquid chromatography (HPLC).

2.5. Histological analysis

Liver and kidney tissues were fixed in 4% (v/v) paraformaldehyde at 4 °C overnight, dehydrated through a graded ethanol series, and embedded in paraffin. Sections (5–8 μm) were cut, dewaxed, rehydrated, and stained with hematoxylin and eosin (H&E). Images were acquired using an Olympus BX53 microscope (Olympus, Tokyo, Japan).

2.6. Measurement of uric acid-associated enzymes and inflammatory markers

Hepatic enzyme activities, including XOD, adenosine deaminase (ADA), and purine nucleoside phosphorylase (PNP), were quantified using corresponding commercial kits following the manufacturers’ instructions.

Serum concentrations of pro-inflammatory cytokines and related markers—including interleukin-1β (IL-1β), interleukin-6 (IL-6), tumor necrosis factor-α (TNF-α), lipopolysaccharide (LPS), and malondialdehyde (MDA)—were analyzed via enzyme-linked immunosorbent assay (ELISA) kits, following the manufacturers’ instructions. Serum creatinine (Cr) and blood urea nitrogen (BUN) levels were analyzed using an automated biochemical analyzer (IDEXX Laboratories, Westbrook, USA).

2.7. Quantitative real-time PCR (qPCR)

Total RNA from tissues and cells was isolated using TRIzol reagent (Invitrogen, Waltham, MA, USA) following the manufacturer's protocol. A total of 500 ng RNA per sample was reverse-transcribed into complementary DNA (cDNA) using the RevertAid First Strand cDNA Synthesis Kit (Thermo Fisher Scientific, Waltham, MA, USA). Quantitative PCR was conducted with PowerUp SYBR Green Master Mix (Thermo Fisher Scientific) on a CFX real-time PCR system (Bio-Rad, Hercules, CA, USA). Primer sequences were synthesized by Tsingke Biotechnology (Nanjing, China) and are listed in Table S1.

2.8. Nucleoside degradation assay

L. rhamnosus suspensions at a density of 1.5 × 10⁸ CFU mL⁻¹ were centrifuged to remove the MRS broth and washed twice with PBS. The bacterial pellets were then resuspended in 750 μL of sodium phosphate buffer (0.1 M, pH 7.4) containing inosine, guanosine, guanine, hypoxanthine, and xanthine as substrates. After incubation, an equal volume of 5% (v/v) trifluoroacetic acid was added to terminate the reaction.¹⁷ The resulting mixtures were passed through 0.22 μm membrane filters and analyzed by HPLC to determine substrate concentrations. Degradation rate (%) was calculated based on the decrease in substrate concentration relative to the initial value.

2.9. Untargeted metabolomics analysis

Serum samples were collected, frozen, lyophilized, and homogenized with extraction solvent (methanol : water = 4 : 1, v/v) containing L-2-chlorophenylalanine as an internal standard. After ultrasonic extraction and centrifugation, supernatants were collected for LC-MS/MS analysis. Chromatographic separation and mass spectrometry were performed using a UHPLC-Q Exactive HF-X system (Thermo Fisher Scientific) equipped with an ACQUITY HSS T3 column. Data were acquired in both positive and negative ionization modes under a data-dependent acquisition strategy. Raw files were processed using Progenesis QI for peak alignment, normalization, and compound identification by matching MS/MS spectra against public databases (HMDB, Metlin) and an in-house library. Further analysis was conducted on the Majorbio Cloud Platform (mmajorbio.com). Differential metabolites were identified based on variable importance in projection (VIP) values >1.0 from PLS-DA and p < 0.05 from Student's t-test. Data were log₂-transformed and Pareto-scaled before statistical analysis.

2.10. Microbial DNA extraction and 16S rRNA sequencing

Genomic DNA was isolated from rat fecal samples using the TIANamp Fecal DNA Kit (Tiangen Biotech, Beijing, China) according to the manufacturer's protocol. DNA concentration and purity were assessed using a Qubit Fluorometer (Thermo Fisher Scientific, USA). The V4 region of the 16S rRNA gene was amplified using primers 515F and 806R. PCR reactions were carried out on a Bio-Rad thermal cycler, and products were confirmed by 2% agarose gel electrophoresis. Amplicons were purified using the Agarose Gel DNA Purification Kit (Tiangen Biotech) and quantified with the Qubit system. Equimolar amplicons were pooled and sequenced on an Illumina MiSeq platform (Illumina, USA). Sequence data were analyzed and classified taxonomically using the NovoMagic cloud platform (magic.novogene.com).

2.11. RNA sequencing and transcriptome analysis

After nucleoside treatment, total RNA from L. rhamnosus was extracted using TRIzol reagent (Invitrogen, USA). RNA quality and quantity were assessed using a Bioanalyzer and NanoDrop spectrophotometer. Transcriptome libraries were prepared using the Illumina® Stranded mRNA Prep Kit and sequenced on either the Illumina NovaSeq X Plus or MGI DNBSEQ-T7 platform (PE150). Raw reads were trimmed with fastp, aligned to the reference genome using HISAT2, and assembled with StringTie. Differential expression and functional analyses were performed via the Majorbio Cloud Platform (mmajorbio.com).

2.12. Cloning, expression, and activity assay of nucleoside-degrading enzymes

Two nucleoside-degrading enzyme genes from L. rhamnosus were synthesized by Tsingke Biotechnology (Nanjing, China) based on transcriptomic analysis. The genes were cloned into the pET-28a vector using the ClonExpress® Ultra One Step Cloning Kit (Vazyme), and the recombinant plasmids were transformed into E. coli Trans5α competent cells. For protein expression, E. coli BL21 harboring the recombinant plasmids was cultured in LB medium supplemented with 50 μg mL⁻¹ kanamycin. When the OD₆₀₀ reached ∼0.6, expression was induced with 0.1 mM IPTG at 30 °C for 8 h. Cells were collected, washed, lysed by ultrasonication, and recombinant enzymes were purified via Ni-affinity chromatography. Protein concentrations were determined using a BCA Protein Assay Kit (Beyotime, Shanghai, China) according to the manufacturer's instructions.

The biochemical properties of INR1 and INR2 were further characterized. Enzymatic activity was quantified by HPLC based measurement of substrate conversion and expressed as U mg⁻¹ protein (1 U = 1 μmol substrate converted per min). Temperature dependence was determined between 20 °C and 70 °C at pH 7.0. The pH dependence was determined by measuring enzymatic activity in 50 mM buffers individually adjusted to pH 3.0, 4.0, 5.0, 6.0, 7.0, 8.0, and 9.0. Sodium citrate–phosphate buffer was used for acidic pH, phosphate buffer for neutral pH, and Tris–HCl buffer for alkaline pH. All buffers were adjusted to the target pH at 25 °C using a calibrated pH meter. Reactions contained 1 mM substrate and were incubated for 10 min at 37 °C; residual activity was normalized to the maximal value. All assays were performed in triplicate.

2.13. Transport assays across Caco-2 monolayers

After cell monolayers were fully differentiated, Transwell-based transport assays were performed. Guanosine, inosine, guanine, or hypoxanthine (0.45 mL in HBSS) was added to the apical chamber, while 1.2 mL of HBSS was added to the basolateral chamber. The plates were incubated at 37 °C with shaking (100 rpm) for 2 h. At 30 min intervals, 50 μL of apical medium was collected and replaced with fresh HBSS. At the end of incubation, samples from the basolateral chamber were collected to determine the amount of uric acid transported across the monolayer, representing apical-to-basolateral absorption. Concentrations of nucleosides and their corresponding bases on both sides were quantified by HPLC.

The apparent permeability coefficient (P_app, cm s⁻¹) was calculated according to eqn (1):

where dQ/dt is the rate of substrate appearance in the basolateral chamber (μmol s⁻¹), A is the membrane surface area (cm²), and C₀ is the initial concentration (μM) in the apical chamber.

2.14. HPLC analysis of nucleosides and uric acid-related metabolites

Quantification of inosine, guanosine, hypoxanthine, guanine, and uric acid was performed using a HPLC system (Waters 2690) equipped with a UV detector set at 254 nm. Separation was achieved on an Agilent ZORBAX SB-C18 column (4.6 × 250 mm, 5 μm) at 30 °C. The mobile phase consisted of 25 mM KH₂SO₄ (pH 3.5) containing 5% (v/v) methanol, delivered at a flow rate of 1.0 mL min⁻¹. Sample injection volume was 20 μL. Metabolites were identified by comparing retention times with authentic standards and quantified using standard curves generated for each compound.

2.15. Statistical analysis

Statistical analyses were performed using GraphPad Prism (v9.0). Data are presented as mean ± standard error of the mean (SEM). For two-group comparisons, unpaired two-tailed Student's t-tests or nonparametric tests were used based on data distribution. For multiple-group comparisons, one-way ANOVA followed by appropriate post hoc tests was applied. Normality was assessed using the Shapiro–Wilk test, and homogeneity of variance was verified using Levene's test. Tukey's multiple-comparison test was applied when the ANOVA results were significant. Differences were considered statistically significant at p < 0.05. Each biological replicate represents one individual rat, and all measurements (including serum, urine, and fecal uric acid) were performed in technical triplicates to minimize analytical variation. The number of biological replicates (n) is indicated in the respective figure legends.

3. Results

3.1. L. rhamnosus alleviates hyperuricemia by reducing serum urate and modulating hepatic urate metabolism

Recent studies have highlighted that certain gut microbes may alleviate hyperuricemia by degrading uric acid or its precursors.¹⁸ To test this hypothesis, we isolated a strain of L. rhamnosus from traditional fermented dairy products, which exhibited the ability to degrade guanosine and inosine, two key precursors of uric acid (Fig. 1A). We then administered this strain orally to hyperuricemic rats to evaluate its in vivo effects (Fig. 1B). After intervention, the relative abundance of the bacterium significantly increased in both feces and colonic mucosa (p < 0.05; Fig. 1C), without affecting body weight or food intake (p > 0.05; Fig. S1A and S1B).

Fig. 1 L. rhamnosus alleviates hyperuricemia and modulates hepatic purine metabolism in rats. (A) In vitro degradation efficiency of L. rhamnosus against uric acid, guanosine, inosine, guanine, hypoxanthine, and xanthine (n = 3 independent replicates). (B) Schematic illustration of the experimental design involving oral administration of L. rhamnosus in hyperuricemic rats. (C) Relative abundance of L. rhamnosus in feces and colonic mucosa, determined by RNA quantification (n = 8 per group). (D–F) Uric acid levels in serum (D), urine (E), and feces (F) following L. rhamnosus intervention. Color gradients in the heatmaps indicate urate concentrations, with blue representing lower levels and red representing higher levels. The color scales correspond to the following ranges: 50–356 μM (serum), 72–200 μM μg g⁻¹ (urine), and 2.27–16 μg g⁻¹ (feces). Each biological replicate represents one individual rat, and each measurement was performed in technical triplicate (n = 8 biological replicates per group). (G) Hepatic enzyme activities of xanthine oxidase (XOD), adenosine deaminase (ADA), and purine nucleoside phosphorylase (PNP) following L. rhamnosus intervention. (n = 8 per group). Data are presented as mean ± SEM. Statistical significance was determined by one-way ANOVA followed by Tukey's post hoc test. *p < 0.05, **p < 0.01, ***p < 0.001.

We next measured uric acid levels in biological samples. Oral administration of L. rhamnosus significantly reduced serum uric acid levels compared to the hyperuricemic diet group (p < 0.05; Fig. 1D), while uric acid levels in urine and feces remained unchanged (p > 0.05; Fig. 1E and F). Given that uric acid metabolism is primarily regulated in the liver through various metabolic enzymes,¹⁹ we further assessed hepatic enzyme activities (Fig. 1G). The activities of XOD and ADA were significantly decreased following bacterial intervention (p < 0.05), whereas PNP activity showed no significant change (p > 0.05).

These results demonstrate that L. rhamnosus administration significantly reduced serum uric acid levels and inhibited hepatic XOD and ADA activities, without affecting urate excretion.

3.2. L. rhamnosus alleviates tissue injury and inflammation without affecting renal urate transport

Chronic hyperuricemia can disrupt systemic metabolism and lead to organ damage.²⁰ We first assessed pathological changes in the kidney and liver. H&E staining revealed that hyperuricemic rats exhibited marked tubular epithelial cell detachment (black arrows) and inflammatory cell infiltration (yellow arrows) in the kidney (Fig. 2A). Interestingly, allopurinol intervention was associated with aggravated renal pathology, whereas L. rhamnosus administration alleviated renal injury. Consistently, the renal index was significantly increased in the allopurinol group (p < 0.05; Fig. 2B), whereas no significant changes were observed in the L. rhamnosus group (p > 0.05).

Fig. 2 L. rhamnosus alleviates tissue injury and inflammation without affecting renal function. (A) Representative H&E staining of kidney tissues from hyperuricemic rats showing tubular injury and inflammation following L. rhamnosus intervention (black arrows, tubular epithelial cell detachment; yellow arrows, inflammatory cell infiltration). (B) Renal index of rats following L. rhamnosus administration (n = 8 per group). (C) Representative H&E staining of liver tissues from hyperuricemic rats following L. rhamnosus intervention (black arrows, vacuolar degeneration; yellow arrows, inflammatory cell infiltration). (D) Hepatic index of rats following L. rhamnosus administration (n = 8 per group). (E) Levels of oxidative stress and endotoxin-related markers, including malondialdehyde (MDA), serum lipopolysaccharide (LPS), creatinine (Cr), and blood urea nitrogen (BUN), following L. rhamnosus administration (n = 8 per group). (F) Serum inflammatory cytokines, including IL-1β, IL-6, IL-10, and TNF-α, following L. rhamnosus administration (n = 8 per group). Data are presented as mean ± SEM. Statistical significance was determined by one-way ANOVA followed by Tukey's post hoc test. *p < 0.05, **p < 0.01, ***p < 0.001.

Liver histology further showed vacuolar degeneration (black arrows) and inflammatory infiltration (yellow arrows) in hyperuricemic rats (Fig. 2C), which were aggravated by allopurinol administration. In contrast, L. rhamnosus treatment appeared to moderately improve hepatic morphology, although the extent of histological recovery was limited. Nevertheless, this morphological improvement was not reflected in hepatic index values, which remained unchanged across treatment groups (p > 0.05; Fig. 2D).

We next measured oxidative stress and endotoxin-related markers (Fig. 2E). L. rhamnosus intervention significantly reduced MDA levels in kidney tissue and serum LPS levels (p < 0.05), suggesting its potential antioxidant and barrier-protective effects. Nevertheless, renal function markers (serum Cr and BUN) did not change significantly (p > 0.05).

To further assess systemic inflammation, we quantified several inflammatory cytokines in serum (Fig. 2F). L. rhamnosus treatment significantly decreased the levels of pro-inflammatory cytokines IL-1β and IL-6 (p < 0.05), while increasing the anti-inflammatory cytokine IL-10 (p < 0.05). TNF-α levels remained unchanged (p > 0.05).

Given that renal excretion is a major route for uric acid elimination,²¹ we analyzed the expression of urate transporters in the kidney. L. rhamnosus administration did not significantly alter the expression of the reabsorption-related transporters GLUT9 (Fig. S2A) and URAT1 (Fig. S2B), nor did it affect the excretion-related transporters ABCG2 (Fig. S2C) and OAT1 (Fig. S2D) (p > 0.05).

Collectively, these findings highlight the systemic anti-inflammatory and tissue-protective potential of L. rhamnosus in hyperuricemic conditions.

3.3. L. rhamnosus modulates gut microbial diversity and composition in hyperuricemic rats

Probiotic intake is known to reshape gut microbial communities and may impact host health.²² To explore this, we analyzed the gut microbiota of hyperuricemic rats after L. rhamnosus intervention. Compared to the normal diet group, hyperuricemic rats exhibited a reduced number of operational taxonomic units (OTUs), whereas L. rhamnosus treatment reversed this trend and increased OTU richness (Fig. S3A).

Analysis of alpha diversity further revealed that L. rhamnosus significantly elevated the Chao1 and ACE indices (p < 0.05), indicating enhanced microbial richness (Fig. 3A). However, no significant changes were observed in Shannon or Simpson indices (p > 0.05), suggesting that overall diversity and evenness remained relatively stable. Venn diagram analysis demonstrated distinct shifts in microbial composition following L. rhamnosus treatment (Fig. 3B). Principal component analysis (PCA) further showed that the microbial profiles of L. rhamnosus-treated rats were positioned between those of the normal diet and hyperuricemic diet groups (Fig. 3C), indicating a partial restoration of gut microbial structure.

Fig. 3 L. rhamnosus reshapes gut microbial composition and restores richness in hyperuricemic rats. (A) Alpha diversity indices including Chao1, ACE, Shannon, and Simpson following L. rhamnosus intervention (n = 8 per group). (B) Venn diagram of shared and unique operational taxonomic units (OTUs) among groups (n = 8 per group). (C) Principal component analysis (PCA) of gut microbial communities (n = 8 per group). (D) Firmicutes-to-Bacteroidetes (F/B) ratio at the phylum level (n = 8 per group). (E) Differential taxa enriched in the L. rhamnosus group identified by LEfSe analysis (LDA score >4, p < 0.05; n = 8 per group). (F) Heatmap showing the relative abundance of representative genera across groups (n = 8 per group). Data are presented as mean ± SEM. Statistical significance was determined by one-way ANOVA or T-test as appropriate. *p < 0.05, **p < 0.01, ***p < 0.001.

At the phylum level, L. rhamnosus significantly increased the Firmicutes-to-Bacteroidetes ratio (p < 0.05; Fig. 3D). LEfSe analysis identified several differentially enriched genera and species in the L. rhamnosus group (Fig. 3E), including beneficial taxa such as Lactobacillus johnsonii, Romboutsia ilealis, and Faecalitalea (Fig. 3F and Fig. S3B). These genera have been previously associated with anti-inflammatory effects, gut barrier integrity, and enhanced fermentation capacity, which may contribute to the observed urate-lowering effects.^23–25 In contrast, Oscillibacter, a genus often linked with gut dysbiosis and elevated serum urate levels,²⁶ was significantly reduced after L. rhamnosus administration (Fig. S3C). This suggests that the probiotic may selectively suppress potentially harmful bacteria while promoting beneficial species.

Taken together, these results indicate that L. rhamnosus improves microbial richness and selectively modulates key gut taxa, thereby contributing to the restoration of gut homeostasis in hyperuricemic rats.

3.4. L. rhamnosus reshapes host metabolic profiles through bile acid and lipid-related pathways

Chronic hyperuricemia is often accompanied by metabolic disturbances.²⁷ To investigate whether L. rhamnosus modulates host metabolism, we performed untargeted metabolomic profiling in hyperuricemic rats treated with L. rhamnosus.

PCA revealed clear separation between the metabolic profiles of the normal diet group and the hyperuricemia diet group, indicating that hyperuricemia substantially disrupted host metabolic patterns (Fig. 4A). Notably, the L. rhamnosus group showed a metabolic distribution that tended to lie between the normal diet and hyperuricemia diet groups in the PCA plot, suggesting a partial restoration of metabolic status. Upset diagram analysis showed that a total of 945 metabolites were shared among the three groups, while 11 and 6 unique overlapping metabolites were identified between the L. rhamnosus group and the normal diet or hyperuricemia diet group, respectively (Fig. 4B).

Fig. 4 L. rhamnosus modulates host metabolic profiles via bile acid and lipid-related pathways. (A) Principal component analysis (PCA) showing distinct separation of serum metabolomic profiles among groups (n = 6 per group). (B) Upset plot showing the number of shared and unique metabolites across the three groups (n = 6 per group). (C) Volcano plot displaying differential metabolites between L. rhamnosus and hyperuricemic diet groups. Metabolites with VIP > 1 and p < 0.05 are considered significant (n = 6 per group). (D) Heatmap of representative differential metabolites involved in bile acid and lipid metabolism (n = 6 per group). (E) KEGG pathway enrichment analysis of differential metabolites between L. rhamnosus and hyperuricemic diet groups, highlighting pathways related to lipid metabolism, bile acid biosynthesis, and signal transduction (n = 6 per group). (F) Correlation heatmap between key differential metabolites and representative bacterial species. Spearman correlation was calculated, and only significant associations are shown (p < 0.05; n = 6 per group). Data are presented as mean ± SEM. Statistical significance was determined by one-way ANOVA followed by Tukey's post hoc test. *p < 0.05, **p < 0.01, ***p < 0.001.

We next analyzed differential metabolites between the L. rhamnosus group and the hyperuricemia diet group. As shown in the volcano plot (Fig. 4C), 105 metabolites were significantly altered, including 40 upregulated and 65 downregulated compounds. Based on VIP values (VIP > 1), key altered metabolites were identified, including bile acids (e.g., glycocholic acid, cholic acid, taurochenodeoxycholic acid), flavonoids (e.g., baicalin, liquiritigenin), and lipid-related compounds (e.g., sphingosine), all of which were significantly downregulated in the L. rhamnosus group (Fig. 4D). These metabolites may play critical roles in the anti-hyperuricemic effects of L. rhamnosus.

To further investigate functional pathways involved, we performed KEGG pathway enrichment analysis (Fig. 4E). Differential metabolites were significantly enriched in pathways related to lipid metabolism, bile acid biosynthesis, and signaling transduction, including cholesterol metabolism, primary bile acid biosynthesis, phospholipase D signaling pathway, fat digestion and absorption, and sphingolipid signaling pathway. These changes suggest that L. rhamnosus may modulate urate metabolism and inflammation via regulation of lipid and bile acid pathways.

Finally, we performed a correlation analysis between significantly altered metabolites and species-level gut microbiota abundance (Fig. 4F). The resulting heatmap showed that several key metabolites—including cholic acid, enalkiren, cytarabine, and ubiquinone-2—were significantly correlated with beneficial microbes such as Lactobacillus johnsonii, Lactobacillus murinus, and Ruminococcus torques (p < 0.05). These findings suggest that gut microbiota may influence host metabolic state through metabolite-mediated mechanisms, thereby contributing to the alleviation of hyperuricemia.

3.5. L. rhamnosus degrades nucleosides through intracellular nucleoside hydrolases

While changes in gut microbiota and their metabolites are important for urate metabolism, L. rhamnosus may additionally exert a direct hypouricemic effect through its intrinsic nucleoside-degrading activity. In our initial screening, this strain was identified based on its ability to degrade guanosine and inosine, prompting further investigation into its degradation mechanism.

To evaluate this, we performed nucleoside degradation assays using live bacteria, bacterial lysates, and culture supernatants. The results showed that both live L. rhamnosus and its lysates significantly degraded guanosine and inosine (p < 0.05), whereas the supernatant exhibited no such activity (p > 0.05; Fig. 5A). Notably, live bacteria exhibited higher degradation efficiency than lysates (Fig. S4A and S4B), suggesting that intracellular enzymes are primarily responsible. Substrate-specific assays confirmed that the degradation efficiency was independent of the substrate concentration (Fig. 5B and C).

Fig. 5 L. rhamnosus degrades nucleosides via intracellular nucleoside hydrolases. (A) In vitro degradation rate of live L. rhamnosus, bacterial lysates, and culture supernatants against inosine and guanosine (n = 3 independent replicates). (B and C) Inosine (B) and guanosine (C) degradation by L. rhamnosus at different substrate concentrations (n = 3). (D) KEGG pathway enrichment of differentially expressed genes after nucleotide exposure based on transcriptomic analysis. (E) Volcano plot of differentially expressed genes involved in nucleoside metabolism, highlighting two upregulated candidate hydrolases. (F) SDS-PAGE analysis of recombinant proteins expressed in E. coli corresponding to the two candidate nucleoside hydrolases. (G and H) Enzymatic activities of purified hydrolases in degrading inosine to hypoxanthine (G) and guanosine to guanine (H) (n = 3). Data are presented as mean ± SEM. Statistical significance was determined by one-way ANOVA followed by Tukey's post hoc test. *p < 0.05, **p < 0.01, **p < 0.001.

To further elucidate the molecular mechanism, transcriptomic analysis was conducted during guanosine and inosine degradation. KEGG enrichment revealed upregulation of genes involved in nucleotide metabolism (Fig. 5D). Among the differentially expressed genes, two candidates encoding putative nucleoside hydrolases were significantly elevated (Fig. 5E). These genes were cloned and heterologously expressed in E. coli, and SDS-PAGE analysis confirmed successful expression of recombinant proteins with molecular weights between 35 and 40 kDa (Fig. 5F).

Functional assays demonstrated that these two enzymes catalyzed the hydrolysis of inosine to hypoxanthine (Fig. 5G) and guanosine to guanine (Fig. 5H), leading to a reduction in nucleoside levels. Finally, a dose-dependent nucleoside degradation assay using live L. rhamnosus further confirmed that the transformation efficiency increased with bacterial concentration (Fig. S4C and S4D).

To further verify the catalytic characteristics of the recombinant enzymes, specific activities and enzymatic profiles of INR1 and INR2 were analyzed (Fig. S5). Both enzymes exhibited measurable hydrolytic activity toward their respective substrates, with INR1 showing slightly higher activity than INR2 (p < 0.05). Temperature-dependent assays revealed that both enzymes exhibited the highest activity at 37 °C, while pH-dependent assays showed an optimal catalytic range around pH 6.5–7.0. These results indicate that INR1 and INR2 are mesophilic, neutral nucleoside hydrolases consistent with the physiological environment of L. rhamnosus, supporting their functional role in nucleoside degradation under intestinal conditions.

3.6. L. rhamnosus degrades nucleosides and modulates their intestinal absorption and conversion to urate

To further validate whether L. rhamnosus exerts nucleoside-degrading activity in vivo, we measured the concentrations of nucleosides and purines in the jejunal contents. Oral administration of L. rhamnosus significantly reduced the levels of guanosine and inosine, while increasing the concentrations of their respective degradation products, guanine and hypoxanthine (p < 0.05; Fig. 6A). These results further support the in vivo activity of L. rhamnosus in degrading nucleosides via its intracellular hydrolases.

Fig. 6 L. rhamnosus degrades nucleosides and modulates their intestinal absorption and conversion to urate. (A) Concentrations of nucleosides (guanosine, inosine) and their degradation products (guanine, hypoxanthine) in the jejunal contents of hyperuricemic rats following L. rhamnosus intervention (n = 8 per group). (B) Uric acid production from guanosine, inosine, guanine, and hypoxanthine in Caco-2 Transwell model after treatment with or without L. rhamnosus (n = 6 independent experiments). (C) Translocation rates of inosine and hypoxanthine across Caco-2 monolayers (n = 6). (D) Apparent permeability coefficients (P_app, apparent permeability coefficient, cm s⁻¹) of inosine and hypoxanthine in Caco-2 monolayers (n = 6). (E) Translocation rates of guanosine and guanine across Caco-2 monolayers (n = 6). (F) Apparent permeability coefficients (P_app, apparent permeability coefficient, cm s⁻¹) of guanosine and guanine in Caco-2 monolayers (n = 6). Data are presented as mean ± SEM. Statistical significance was determined by one-way ANOVA followed by Tukey's post hoc test. *p < 0.05, **p < 0.01, **p < 0.001.

We then employed a Caco-2 Transwell model to evaluate whether this degradation affects uric acid production. After treatment with L. rhamnosus, the uric acid levels derived from guanosine and inosine were significantly reduced (p < 0.05), whereas guanine and hypoxanthine did not induce a comparable increase in urate production (p > 0.05; Fig. 6B). These findings suggest that the microbial conversion of nucleosides into purine bases may attenuate their contribution to systemic urate levels.

Given previous reports indicating distinct transport properties between nucleosides and purines, we hypothesized that their differential transmembrane absorption may account for the observed variation in urate production.¹⁴ To test this, we compared the permeability and P_app of guanosine, inosine, guanine, and hypoxanthine using the Caco-2 model. The results showed that nucleosides exhibited significantly higher translocation rates and P_app values than purines (p < 0.05; Fig. 6C–F), indicating that nucleosides are more efficiently absorbed by intestinal epithelial cells.

Taken together, these results reveal a microbial mechanism by which L. rhamnosus reduces uric acid production through enzymatic degradation of dietary nucleosides, thereby preventing their efficient absorption and conversion into uric acid. This mode of action highlights the potential of L. rhamnosus to modulate substrate accessibility at the intestinal level, thereby contributing to its urate-lowering effect.

4. Discussion

Hyperuricemia is an increasingly prevalent metabolic disorder globally, posing significant challenges to public healthcare systems.²⁸ Traditional urate-lowering medications, such as allopurinol and febuxostat, have limitations, including side effects and varying individual efficacy, driving the search for safer and more effective interventions.^29,30 Recently, gut microbiota has emerged as an essential factor in regulating host purine metabolism and urate homeostasis, placing probiotics at the forefront of novel therapeutic approaches.^13,31 However, previous probiotic strategies primarily focused on direct uric acid degradation, purine base breakdown, or enhancement of renal and intestinal urate excretion.^32,33 Despite these advances, the role of dietary nucleosides in urate metabolism has remained underexplored. In this study, we propose and validate an alternative mechanism: L. rhamnosus effectively reduces serum urate levels by enzymatically degrading dietary nucleosides, such as guanosine and inosine, thereby decreasing their intestinal absorption and subsequent availability for hepatic urate synthesis.

Our research identified two key intracellular nucleoside hydrolases in L. rhamnosus, confirming their efficient enzymatic activity toward guanosine and inosine via recombinant protein expression and in vitro assays. Although microbial nucleoside hydrolases have been previously identified, their physiological roles in host purine metabolism, particularly urate synthesis regulation, remain poorly characterized.^34,35 Unlike uricase (direct urate degradation) or guanine deaminase (purine base degradation), our findings highlight nucleoside hydrolase-mediated degradation as an upstream intervention strategy, effectively reducing urate precursors before systemic absorption. This provides a novel microbial target and expands the existing conceptual framework of probiotic functionality.

Transwell assays using Caco-2 cells revealed that nucleosides have substantially higher transepithelial permeability than purine bases (guanine and hypoxanthine). This is explained by their distinct transport routes: nucleosides utilize active transporters such as CNTs and ENTs,^36–38 while purine bases primarily rely on passive diffusion.³⁹ Consequently, enzymatic conversion of nucleosides into purine bases profoundly changes their absorption kinetics and lowers the systemic availability of urate precursors. Furthermore, CNT2 (SLC28A2) and ENT1/2 (SLC29A1/2) exhibit substrate specificity toward inosine and guanosine, and are known to be modulated by intestinal metabolites such as short-chain fatty acids (SCFAs) and bile acids. Therefore, L. rhamnosus induced remodeling of the gut environment may also indirectly influence nucleoside uptake by altering transporter expression or activity, further contributing to reduced urate production. This substrate-level intervention represents a distinctive probiotic mechanism, in contrast to conventional strategies targeting urate degradation or enhanced excretion.

Beyond its direct enzymatic degradation of nucleosides, L. rhamnosus appears to exert a broader regulatory effect on the intestinal ecosystem. The enrichment of commensal taxa such as Lactobacillus johnsonii, Romboutsia ilealis, and Faecalitalea, together with the suppression of potentially harmful genera like Oscillibacter, suggests a shift toward a more anti-inflammatory microbial configuration.⁴⁰ Such compositional restructuring likely influences host metabolic homeostasis through the production of microbial metabolites that interact with bile acid and sphingolipid pathways. These pathways are key regulators of inflammation, oxidative balance, and purine turnover, and their downregulation following probiotic treatment implies that L. rhamnosus may alleviate hyperuricemia not only by reducing nucleoside availability but also by restoring host metabolic signaling through microbiota–derived intermediates.

These findings suggest that microbiota restructuring and metabolic modulation likely operate in tandem with the enzymatic degradation of nucleosides. Although causality remains to be fully clarified, it is conceivable that the initial degradation of nucleosides by L. rhamnosus alters intestinal substrate availability, which in turn reshapes the surrounding microbial community and metabolic environment. The resulting shifts in bile acid⁴¹ and lipid signaling⁴² may feed back to the host, suppressing hepatic urate synthesis and inflammatory responses through a bidirectional regulatory network. Moreover, microbial metabolites such as SCFAs and secondary bile acids could further influence intestinal or hepatic expression of nucleoside transporters and xanthine oxidase, extending the probiotic's regulatory reach to multiple metabolic nodes.

From a broader systems perspective, the modulation of bile acid and lipid pathways observed here may also participate in urate regulation through metabolic cross-talk. Bile acids function as endocrine-like signals that activate FXR and TGR5, both of which are known to downregulate hepatic xanthine oxidase and attenuate oxidative stress. Likewise, sphingolipid metabolism is closely linked to inflammatory tone and redox balance, two key determinants of urate accumulation and tissue injury.^43–45 Hence, L. rhamnosus induced remodeling of these pathways may synergize with its nucleoside-degrading capacity, establishing a multilayered mechanism that integrates microbial, metabolic, and enzymatic regulation of urate homeostasis.

Additionally, L. rhamnosus demonstrated notable protective effects against hyperuricemia-induced renal injury and systemic inflammation, independent of changes in renal urate transporter expression (GLUT9, URAT1, ABCG2, OAT1). This contrasts markedly with traditional medications like allopurinol, which may cause adverse effects due to their direct enzyme inhibition mechanisms.^46,47 The probiotic-mediated protection appears more likely derived from early intervention in precursor substrate availability and systemic inflammation regulation, highlighting potential broader physiological benefits beyond urate control.

However, this study has several limitations. First, animal models do not fully replicate the complexity of human urate metabolism, necessitating further clinical validation. Second, while the differential intestinal absorption of nucleosides versus purine bases has been clearly demonstrated, specific intestinal transporter proteins and signaling pathways involved require further elucidation. Additionally, long-term safety and efficacy of L. rhamnosus administration on host microbiota and metabolism remain to be thoroughly evaluated.

Overall, this study elucidates a novel microbial mechanism whereby L. rhamnosus enzymatically degrades nucleoside precursors, reducing their intestinal absorption and subsequent hepatic conversion to urate. Moreover, the concurrent modulation of gut microbiota and host metabolome, particularly through bile acid and sphingolipid pathways, suggests a cooperative systems-level mechanism that underlies its urate-lowering effects. This finding establishes a substrate-level intervention paradigm and provides theoretical and translational insights into microbiome-based strategies for hyperuricemia management.

5. Conclusions

This study demonstrates that L. rhamnosus alleviates hyperuricemia through a substrate-restriction mechanism by degrading dietary nucleosides and limiting their contribution to urate synthesis. Beyond direct enzymatic activity, the probiotic also reshaped gut microbiota, modulated host metabolism, and alleviated renal injury and systemic inflammation. These findings expand the functional repertoire of probiotics and underscore the potential of food-derived microbes as safe and sustainable interventions for hyperuricemia and related metabolic disorders.

Author contributions

P. C. performed the majority of the experiments, curated and analyzed the data, and wrote the original draft. H. C. contributed to study design, method optimization, and data analysis. S. C. assisted with method optimization and project supervision. X. Y., and J. C. contributed to study design and supervision. X. Y. contributed to funding acquisition and manuscript editing. J. C. contributed to method optimization, data curation, and manuscript revision. All authors reviewed and approved the final version of the manuscript.

Conflicts of interest

The authors declare no competing interests.

Data availability

The 16S rRNA, transcriptomic, and metabolomic datasets generated in this study have been deposited in public repositories. The 16S rRNA and transcriptome data are available at the NCBI Sequence Read Archive under BioProjects PRJNA1291202, and PRJNA1291240, and the metabolomics data are available at the MetaboLights repository under accession number MTBLS12735 (https://www.ebi.ac.uk/metabolights/MTBLS12735).

Supplementary information (SI) is available. See DOI: https://doi.org/10.1039/d5fo03882k.

Acknowledgements

This work was financially supported by the “Pioneer” and “Leading Goose” R&D Program of Zhejiang (2022C02017), Zhejiang Agricultural Science and Technology Plan (2025SNJF007), the Key R&D Program of Shandong Province, China (2023TZXD025), and the Key Research and Development Program of Zhejiang Province (2023C02058).

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