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Article

Serum Uric Acid-Reducing Effect and Intestinal Mucosal Barrier-Repairing Function of Limosilactobacillus reuteri MBHC10138

by
Jinhua Cheng
1,2,3,*,†,
Youjin Lee
2,4,5,†,
Joo-Hyung Cho
2,3 and
Joo-Won Suh
2,3,*
1
School of Pharmacy and Food Engineering, Wuyi University, Jiangmen 529000, China
2
Myongji Bioefficacy Research Center, Myongji University, Yongin 17058, Republic of Korea
3
R&D Research Center, MBHC Co., Ltd., Yongin 17058, Republic of Korea
4
Institute for Future Earth, Pusan National University, Busan 46241, Republic of Korea
5
Department of Biological Sciences, Pusan National University, Busan 46241, Republic of Korea
*
Authors to whom correspondence should be addressed.
†
These authors contributed equally to this work.
Microorganisms 2026, 14(2), 384; https://doi.org/10.3390/microorganisms14020384
Submission received: 28 November 2025 / Revised: 1 February 2026 / Accepted: 2 February 2026 / Published: 5 February 2026
(This article belongs to the Special Issue Advances in Host Gut Microbiota—2nd Edition)
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Abstract

Hyperuricemia is a metabolic disorder characterized by elevated serum uric acid levels and is increasingly linked to alterations in intestinal mucosal condition and gut microbiota composition. Probiotics have been proposed as safe, non-pharmacological approaches for managing hyperuricemia, but strain-specific evidence remains limited. This study aimed to evaluate the anti-hyperuricemic potential of Limosilactobacillus reuteri MBHC10138, isolated from human breast milk, and to examine its association with purine metabolism–related parameters, renal morphological features, intestinal barrier-associated markers, and gut microbiota composition. In vitro, MBHC10138 effectively degraded purine nucleosides that are metabolized into uric acid, suggesting its potential to reduce uric acid production in the host. In a mouse model of diet- and oxonate-induced hyperuricemia, oral administration of MBHC10138 significantly lowered serum uric acid levels to a level comparable with allopurinol treatment, while improving renal morphology. Histological and molecular analyses demonstrated restoration of the tight junction proteins zonula occludens-1 and occludin, indicative of enhanced intestinal barrier integrity. Furthermore, MBHC10138 administration modulated the gut microbiota by restoring microbial α-diversity and significantly increasing the relative abundances of the Clostridia vadinBB60 group and Oscillospiraceae, taxa associated with butyrate production, compared with the allopurinol-treated group. Collectively, these findings indicate that MBHC10138 exerts dual actions against hyperuricemia and intestinal barrier dysfunction through the regulation of purine metabolism, promotion of renal urate excretion, and modulation of gut microbial composition. MBHC10138 may thus represent a promising probiotic candidate for the prevention and adjunctive management of hyperuricemia-related metabolic disorders.

1. Introduction

Excessive intake of purine-rich foods and alcohol has contributed to the global rise in hyperuricemia, a major metabolic disorder characterized by elevated serum uric acid (SUA) [1]. Hyperuricemia develops when uric acid production exceeds renal or intestinal excretion capacity, resulting in uric acid (UA) accumulation and monosodium-urate crystal formation [2,3,4]. Chronic hyperuricemia is also linked to cardiovascular and cerebrovascular diseases [5,6], and its prevalence has steadily increased in both sexes over the past decade, particularly among men [7].
Because hyperuricemia is often asymptomatic, pharmacologic intervention is limited to symptomatic or severe cases. Allopurinol (ALLO), a xanthine oxidase inhibitor, is the most widely prescribed urate-lowering agent; however, it can cause severe hypersensitivity reactions, including Stevens–Johnson syndrome and other severe cutaneous adverse reactions (SCAR), particularly in individuals carrying the HLA-B5801 allele, which is highly prevalent among Asian populations [8,9,10]. Other urate-lowering agents also exhibit unpredictable toxicity, occasionally leading to fatal outcomes [11]. Therefore, developing safe, non-pharmacological strategies for the prevention and management of hyperuricemia is urgently needed.
Uric acid homeostasis involves both renal and intestinal excretion. While renal mechanisms are well characterized, intestinal pathways remain less understood. The intestine excretes approximately 30% of total body UA through ATP-binding cassette subfamily G member 2 (ABCG2)-mediated transport, acting as an extra-renal route of urate elimination [12,13]. Impaired intestinal excretion or transporter dysfunction can lead to elevated SUA levels and hyperuricemia [14]. Moreover, hyperuricemia has been reported to exacerbate intestinal pathologies, including colon cancer progression [15]. The gut microbiota is closely involved in uric acid metabolism [16,17]. Microbial dysbiosis can disrupt mucosal immunity and compromise the intestinal barrier, allowing translocation of bacteria, cytokines, and lipopolysaccharides into circulation and aggravating systemic inflammation [18,19]. Disruption of the intestinal mucosal barrier, particularly the loss of tight junction proteins such as ZO-1, occludin, and claudin-1, has been implicated in increased intestinal permeability and the progression of systemic metabolic and inflammatory disorders [20,21].
Probiotics can beneficially modulate the gut environment by restoring microbiota diversity, producing health-promoting metabolites, and repairing the intestinal mucosal barrier [22,23]. Among them, Limosilactobacillus reuteri is a well-characterized probiotic species that has garnered significant interest for its ability to colonize the gastrointestinal tract of diverse hosts and exert multifaceted health benefits. These benefits include modulation of the gut microbiota, reinforcement of the intestinal epithelial barrier, and immunoregulatory activities [24,25,26]. Notably, certain strains of L. reuteri have demonstrated efficacy in ameliorating metabolic disorders, such as obesity [27,28] and hyperlipidemia [29], through interactions with host metabolism. However, its specific role and mechanistic involvement in purine metabolism and hyperuricemia remain largely unexplored.
In the screening process for anti-hyperuricemic probiotics, L. reuteri MBHC10138 was selected based on its potent purine nucleoside–degrading activity in vitro [30]. This property is of particular interest because nucleosides can be converted to uric acid in vivo, contributing to hyperuricemia. Thus, interventions that reduce uric acid generation and/or enhance its elimination may effectively lower serum uric acid levels. Building on this premise and the established link between gut microbiota dysbiosis and hyperuricemia, we hypothesized that L. reuteri MBHC10138, could alleviate hyperuricemia by modulating key pathways involved in uric acid production and excretion. Therefore, the objectives of this study were to: (1) evaluate the anti-hyperuricemic efficacy of L. reuteri MBHC10138 in a mouse model induced by a high-purine diet and potassium oxonate; and (2) investigate its potential to restore intestinal barrier function and gut microbial composition disrupted by hyperuricemia.

2. Materials and Methods

2.1. Degradation of Purine Compounds by MBHC10138

2.1.1. Incubation in Nucleoside Solution

The purine-degrading ability of L. reuteri MBHC10138 was evaluated with minor modifications of a previously described method [31]. Live cells, cell-lysate debris, and cell-lysate supernatant were tested to determine their ability to degrade purine nucleosides. MBHC10138 was cultured in de Man–Rogosa–Sharpe (MRS; Difco, Sparks, MD, USA) medium at 37 °C for 48 h under anaerobic conditions. Cultures were centrifuged (4000× g, 10 min, 4 °C), and the pellets were washed three times with 0.85% NaCl. The washed pellet was used as the live-cell preparation.
To prepare the lysate fractions, cells were suspended in 3 mL of 1 mM lysozyme and incubated for 3 h. The lysed material was freeze-dried, weighed, and resuspended in 0.85% NaCl (10 mg/mL), then sonicated. The lysate was centrifuged (1500× g, 10 min); the supernatant contained soluble cytosolic components, and the pellet represented the membrane fraction [32]. Each sample—live cells (0.05 g), lysate debris (10 mg), or lysate supernatant (10 mg)—was mixed with 750 μL of 1 mM inosine, guanosine, or adenosine solution and incubated at 37 °C for 2 h with shaking (120 rpm). After incubation, mixtures were centrifuged (4000× g, 10 min, 4 °C), and 270 μL of the supernatant was collected. To terminate reactions, 30 μL of 0.1 M HClO4 was added. Samples were filtered (0.22 μm) and analyzed by HPLC.
To evaluate the degradation capacity of Limosilactobacillus reuteri strain MBHC10138 in comparison with other strains, three L. reuteri strains: DSM 17938, KACC 11452, and MBHC10668 were employed for live bacterial assays. L. reuteri DSM 17938 was obtained from a commercial BioGaia product (Stockholm, Sweden) and confirmed by 16S rRNA sequencing. L. reuteri KACC 11452 was acquired from the Korean Agricultural Culture Collection, and L. reuteri MBHC10668 was isolated from infant feces. All strains were cultured anaerobically in MRS medium at 37 °C for 48 h, adjusted to a density of 108 CFU/mL, and subsequently assessed for purine degradation using the method described above.

2.1.2. HPLC Analysis

HPLC analysis followed a modified method [33]. A Waters 1525 binary pump with a 2498 photodiode-array detector and a YMC ODS C18 column (250 × 4.6 mm, 5 μm; YMC, Kyoto, Japan) was used. The column temperature was maintained at 37 °C [31]. The mobile phase consisted of 0.1 mM NaClO4 and 0.187 M H3PO4 in deionized water, filtered and degassed ultrasonically for 20 min. Isocratic elution was performed at 1 mL/min, and nucleosides were detected at 254 nm. Retention times were 10.2 min (adenosine), 16.6 min (guanosine), and 21.2 min (inosine); peaks were quantified against authentic standards.

2.1.3. Utilization of Purine Compounds for Growth

Growth utilization of purine nucleosides by MBHC10138 was assessed using a chemically defined (DM) medium [34] containing purine nucleotides, nucleosides, or bases. MBHC10138 (2 × 108 CFU/mL) was inoculated into DM supplemented with 400 μM purine nucleosides and incubated anaerobically at 37 °C for 8 h. Growth was monitored at OD600 (Tecan Infinite M200 Pro, Grödig, Austria). DM without purine nucleosides served as the control. Experiments were performed in triplicate.

2.2. Hyperuricemia Model

2.2.1. Animal Experiment and Treatment

Seven-week-old male C57BL/6 mice (RaonBio, Yongin, Gyeonggi, Republic of Korea) were housed (4 mice per cage) under controlled conditions (22 ± 2 °C, 55 ± 5% humidity, 12 h light/dark cycle) with free access to food and water. All animal procedures were approved by the Institutional Animal Care and Use Committee of Myongji University (MJIACUC-2021001) and conformed to the NIH Guide for the Care and Use of Laboratory Animals.
After 1 week of acclimatization, mice were randomly assigned to the experimental groups using a random number–based allocation method. Group allocation was performed to ensure comparable baseline body weights across groups (n = 8 per group): (a) control (Con), (b) hyperuricemia (HU), (c) allopurinol (ALLO), and (d) MBHC10138 (3 × 108 CFU/mouse/day) and (e) MBHC10138 (3 × 109 CFU/mouse/day). To induce hyperuricemia, mice in groups b–e received intraperitoneal injections of potassium oxonate (300 mg/kg body weight/day) dissolved in 3 g/L carboxymethylcellulose sodium (CMC-Na), while Con mice received vehicle only. Potassium oxonate was administered at a dose of 300 mg/kg body weight/day based on previous studies demonstrating that this dosage effectively inhibits uricase activity and reliably induces hyperuricemia in mice without causing overt systemic toxicity [35]. Group Con was fed a standard diet ANI-93 M (RaonBio, Yongin, Gyeonggi, Republic of Korea), whereas groups b–e were fed a high-purine diet (HPD), containing 57% of yeast extract in ANI-93 M.
MBHC10138 cultures were freshly prepared daily in MRS broth (24 h, 37 °C), washed twice with 0.85% NaCl, and resuspended for oral administration (3 × 108, 3 × 109 CFU/mouse/day) for 21 days. ALLO was orally administered (5 mg/kg body weight/day) according to the previous study [33,36]. Control and HU groups received 0.85% NaCl vehicle. Body weight was recorded every 3 days. Successful establishment of hyperuricemia was verified by a significant increase in serum uric acid levels in the HU group compared with the control group.

2.2.2. Serum Biochemical Analysis

On day 21, mice were anesthetized with 3% isoflurane, and blood was collected via cardiac puncture. Samples were kept on ice for 1 h and centrifuged (2000× g, 15 min, 4 °C) to obtain serum. UA, creatinine, and blood urea nitrogen (BUN) were measured using a biochemical analyzer (FUJIFILM DRI-CHEM NX500i, Kanagawa, Japan). Serum xanthine oxidase (XO) activity was measured using a commercial colorimetric enzymatic activity assay kit (Xanthine Oxidase Activity Assay Kit, Sigma-Aldrich, St. Louis, MO, USA; MAK078), according to the manufacturer’s instructions. This assay quantifies XO activity based on the enzymatic conversion of xanthine to uric acid and hydrogen peroxide, which is detected via a colorimetric reaction. XO activity was expressed as U/L.

2.2.3. Monitoring and Euthanasia

Mice were monitored daily for weight, activity, posture, and coat condition. Animals showing ≥20% weight loss or abnormal behavior (score ≥ 3) were humanely euthanized. Euthanasia was performed under 3% isoflurane anesthesia followed by cervical dislocation.

2.2.4. Histopathological Evaluation

Histological assessments were performed by investigators who were blinded to the treatment groups to minimize observer bias. Kidney and intestinal tissues (ileum) were fixed in 4% paraformaldehyde (24 h, 4 °C) and embedded in paraffin. Sections (5 μm) were stained with hematoxylin and eosin (H&E) for kidney evaluation and Alcian blue (pH 2.5) followed by nuclear fast red counterstaining to visualize intestinal acidic mucins and the underlying tissue architecture simultaneously. Glomerular tuft area was quantified in CaseViewer (3DHISTECH, Budapest, Hungary) using the closed-polygon tool; 25 fields per sample were analyzed. Goblet cells were quantified by counting Alcian blue–positive cells per intact crypt. A total of 28 (Control), 30 (HU), 22 (ALLO), and 19 (MBHC10138) crypts were analyzed. Results were plotted in GraphPad Prism v8.

2.2.5. Fecal Microbiota Analysis

Intestinal fecal contents were collected, immediately frozen at −80 °C, and processed using the Exgene™ Stool DNA Mini Kit (GeneAll, Seoul, Republic of Korea) according to the manufacturer’s protocol. The V3–V4 region of the bacterial 16S rRNA gene was amplified with primers 341F/805R [37], and taxonomic profiling was performed using the EzBioCloud Microbial Taxonomic Profiling platform (Sanigen, Anyang, Republic of Korea).

2.2.6. Tissue Collection and RNA Extraction

Frozen kidney and intestinal samples were ground under liquid nitrogen. Total RNA was isolated using the Takara RNA Isolation Kit (Takara, Dalian, China) following the manufacturer’s protocol [36].
RNA integrity was verified by 1% agarose gel electrophoresis, and concentration and purity were measured with an ND-1000 spectrophotometer (NanoDrop, Thermo Fisher, Waltham, MA, USA).

2.2.7. Quantitative Real-Time PCR (qRT-PCR)

Total RNA was isolated with an RNA isolation kit (Takara, Dalian, China). Genomic DNA was removed and cDNA was synthesized using the gDNA Eraser Takara PrimeScript RT Kit (Takara, Dalian, China) according to the manufacturer’s instructions. SYBR Premix Ex Taq (Takara) was used for qPCR (10 μL reaction volume) on a Roche LightCycler® system (Roche, Basle, Switzerland). The cycling conditions were 95 °C for 3 s and 60 °C for 20 s, 45 cycles. Primer sequences followed published references [38,39]. Molecular analyses, including qRT-PCR and ELISA, were conducted with investigators blinded to group allocation during sample processing and data analysis. Primer sequences are summarized in Table 1. Relative gene expression levels were calculated using the 2−ΔΔCt method, with GAPDH used as the housekeeping gene for normalization.

2.2.8. Tight Junction Protein (TJP) Expression

Intestinal tissues were homogenized (100 mg in 2 mL PBS) and analyzed for protein content using the Pierce™ BCA Protein Assay Kit (Thermo Fisher Scientific, USA). Zonula occludens-1 (ZO-1) levels were measured with an ELISA kit (LSBio, Newark, CA, USA; LS-F24053-1), and occludin levels with an ELISA kit (Antibodies-online, Pottstown, PA, USA; ABIN6720584) according to the manufacturer’s protocols.

2.3. Statistical Analysis

All data are expressed as mean ± SD. Normality was tested using the Shapiro–Wilk test. For normally distributed data, one-way ANOVA followed by Tukey’s post hoc test was applied. All analyses were performed in GraphPad Prism v8.0 and SPSS v19. A p value < 0.05 was considered statistically significant.

3. Results

3.1. Evaluation of Purine Compounds Degradation

3.1.1. Degradation of Purine Compounds by L. reuteri MBHC10138

To evaluate the potential anti-hyperuricemic activity, the adenosine-, guanosine-, and inosine-degrading abilities of L. reuteri MBHC10138 were assessed. When live cells were applied, MBHC10138 exhibited nearly complete (≈100%) degradation of all three purine nucleosides (Figure 1A–C). When lysate debris was used, the strain degraded 80.0% of both adenosine and guanosine (Figure 1A,B) and 46.7% of inosine (Figure 1C). The lysate supernatant degraded 43.5%, 49.5%, and 52.0% of adenosine, guanosine, and inosine, respectively (Figure 1A–C). The HPLC chromatogram of degradation of guanosine and inosine were shown in Figure S1. These findings indicate that both cytosolic and cell wall components of MBHC10138 possess purine-nucleoside-degrading activity.
To compare the degradation capacity of L. reuteri strain MBHC10138 with other strains under a more stringent condition, the bacterial concentration was lowered to 108 CFU/mL and the reaction time was reduced to 15 min. At equivalent bacterial concentrations, strain MBHC10138 demonstrated significantly greater purine degradation activity compared to the other L. reuteri strains tested. MBHC10138 degraded guanosine, inosine, and adenosine by 69%, 82.5%, and 96.3%, respectively. Under the same conditions, DSM 17938 degraded these substrates by 58.6%, 60%, and 19.3%; KACC 11452 by 5%, 6%, and 18%; and MBHC10668 by 20%, 23.5% and 30.6%, respectively (Figure S2).

3.1.2. Utilization of Purine Compounds for Bacterial Growth

To determine whether purine compounds support bacterial growth, MBHC10138 was cultured in a chemically defined medium containing various purine nucleotides, nucleosides, or bases. Growth was enhanced in all purine-supplemented media compared with the control medium lacking purine compounds. After 8 h of incubation, growth increased by 15.3%, 25.7%, and 28.4% in adenine-, guanine-, and IMP-supplemented media, respectively, compared with the control (Figure 2). These results demonstrate that MBHC10138 can utilize purine compounds as growth substrates.

3.2. Animal Experiment

3.2.1. Effect of L. reuteri MBHC10138 on Body Weight and Serum Uric Acid Levels

Hyperuricemia was induced by a high-purine diet and intraperitoneal injection of potassium oxonate, and MBHC10138 was administered daily for 21 days. Body weight did not differ significantly among groups (Figure 3A).
The serum uric acid (SUA) level of the HU group reached 71.4 μmol/L, representing a 66% increase relative to the control (Con, 43.1 μmol/L), confirming successful model establishment. The serum UA levels of the ALLO, MBHC10138 (3 × 108 CFU/mL) and MBHC10138 (3 × 109 CFU/mL) groups were 26.7 μmol/L, 24.5 μmol/L and 16.6 μmol/L, respectively, all of which were significantly lower than those in the HU group. However there was no significant difference among Allopurinol, MBHC10138 (3 × 108 CFU/day) and MBHC10138 (3 × 109 CFU/day) group (p < 0.0001, Figure 3B).
Serum xanthine oxidase (XO) activity, a key enzyme converting hypoxanthine to uric acid, was 16.4 U/L in the HU group, slightly elevated compared with the Con group. XO activity was markedly reduced in the ALLO group (5.6 U/L) and further decreased in the MBHC10138 (3 × 109 CFU/day) group (3.4 U/L), consistent with the reduction in SUA (Figure 3C).

3.2.2. Histopathological and Biochemical Analysis of Kidney Tissues

Approximately 60–70% of systemic uric acid is excreted via the kidneys; thus, sustained hyperuricemia may induce glomerular and tubular alterations [40]. H&E-stained sections revealed pronounced structural changes in the HU group, including enlarged glomeruli (Figure 4A). The ALLO- and MBHC10138-treated groups maintained glomerular morphology comparable to the Con group. Quantitative analysis showed that the glomerular tuft area increased by 46% in the HU group, 20.5% in the ALLO group, and 4.8% in the MBHC10138 group relative to Con. Compared with HU, MBHC10138 reduced glomerular hypertrophy by 27.2%, suggesting protection against renal structural damage (Figure 4B).
Serum creatinine and BUN levels were elevated in the HU group, indicating renal stress, and also increased slightly in the ALLO group (Figure 4C,D). In contrast, the MBHC10138 group exhibited significantly lower creatinine (13.6 μmol/L vs. 18.0 μmol/L in HU) and BUN (18.86 mg/dL vs. 24.62 mg/dL in HU) levels, implying no evidence of renal functional impairment by these markers.

3.2.3. Expression of Renal Urate Transporters

The basolateral membranes of proximal tubular cells contain organic anion transporters (OAT1 and OAT3), which are essential for urate excretion [41]. Dysfunction of OAT1, OAT3, or URAT1 contributes to hyperuricemia [42]. The mRNA levels of mOAT1 (0.8) and mOAT3 (0.9) were slightly reduced in HU compared with Con (Figure 4E,F), whereas their expression was significantly upregulated in the MBHC10138 group (1.4-fold each).
URAT1 expression was elevated in the HU group (1.2-fold) relative to Con but decreased to 0.5 in the ALLO group and 0.93 in the MBHC10138 group (Figure 4G). Taken together, the observed reduction in serum uric acid, coupled with the modulation of renal transporter gene expression (upregulation of OAT1/OAT3 and downregulation of URAT1), suggests that MBHC10138 may promote renal urate excretion, potentially through influencing these transporter pathways.

3.2.4. Intestinal Histology and Tight Junction Protein Expression

Histological analysis was performed using Alcian Blue (pH 2.5) followed by Nuclear Fast Red counterstaining to concurrently visualize intestinal acidic mucins and the underlying tissue architecture. The HU and ALLO groups exhibited marked mucosal deterioration, characterized by villous surface irregularities and significant submucosal edema. In contrast, the MBHC10138-treated group maintained mucosal morphology and acidic mucin distribution comparable to the Control group, demonstrating a protective effect on the intestinal barrier (Figure 5A).
The mRNA levels of tight junction proteins ZO-1 and occludin, crucial for intestinal barrier integrity [43,44], were significantly decreased in the HU group—particularly occludin (p < 0.0001)—indicating increased intestinal permeability [45]. Both ZO-1 and occludin expression were significantly restored in the MBHC10138 group (p < 0.01; Figure 5B,C).
ELISA analysis confirmed these trends at the protein level. ZO-1 and occludin expression were reduced in HU and ALLO groups but restored in MBHC10138-treated mice (HU vs. MBHC10138, p < 0.01; Figure 5D,E). Collectively, these findings suggest that MBHC10138 is associated with improvement of intestinal barrier–related histological and molecular markers under hyperuricemic conditions.
Consistent with these histological observations, quantitative analysis showed that the number of Alcian blue–positive goblet cells per crypt was significantly reduced in the HU group, whereas MBHC10138 treatment significantly increased goblet cell counts compared with the HU group, indicating partial restoration of mucus-producing epithelial cells under hyperuricemic conditions (Figure 5F).

3.2.5. Intestinal Microbial Diversity

Gut microbiota composition was analyzed at both the phylum and family levels. The predominant phyla were Bacteroidetes and Firmicutes (Figure 6A). In the Con group, their proportions were 30.88% and 51.44%, respectively. In HU, Bacteroidetes increased to 44.15% while Firmicutes decreased to 35.32%. The ALLO group displayed 62.07% and 29.66%, whereas the MBHC10138 group showed 44.47% and 50.71%, respectively.
At the family level, MBHC10138 increased most taxa compared to HU, except for Akkermansiaceae (Figure 6B). The Firmicutes/Bacteroidetes ratio was markedly lower in HU and further decreased by ALLO treatment, whereas MBHC10138 partially restored this ratio (Figure 6C). The relative abundances of the Clostridia vadinBB60 group and Oscillospiraceae tended to increase in MBHC10138-treated mice compared with the HU group, while showing a significant increase compared with the allopurinol-treated group (Figure 6D).
Alpha diversity, assessed by the Shannon index, was reduced in HU but significantly increased in the MBHC10138 group, indicating restoration of microbial diversity (Figure 6E).

4. Discussion

Hyperuricemia is a metabolic disorder characterized by elevated serum UA levels, primarily resulting from excessive intake of purine-rich foods such as meat, seafood, and alcohol [1]. The global incidence of hyperuricemia and gout has steadily increased, including among individuals under 30 years of age [46]. Although asymptomatic hyperuricemia often remains untreated, persistent elevation of UA can lead to gout, which is characterized by recurrent inflammatory attacks caused by urate crystal deposition in joints when SUA exceeds approximately 7.0 mg/dL (≈416 µmol/L).
Conventional pharmacotherapies, including allopurinol and febuxostat (xanthine oxidase inhibitors) and benzbromarone (a uricosuric agent), effectively reduce SUA levels but are often limited by adverse effects such as severe cutaneous adverse reactions (SCAR), cardiovascular complications, and hepatotoxicity [47,48]. URC102, a selective URAT1 inhibitor currently under clinical evaluation, has shown potential in lowering SUA [49]. However, safer, non-pharmacological strategies remain desirable. In this context, probiotic supplementation has been reported to lower SUA and prevent hyperuricemia. Lactobacillus brevis DM9216 isolated from Chinese sauerkraut degraded purine nucleosides and reduced SUA levels in vivo [50]. Probiotics offer an accessible, non-prescription alternative that can complement or substitute pharmacological interventions.
In the present study, L. reuteri MBHC10138, isolated from human breast milk, demonstrated potent purine-nucleoside-degrading capacity and significantly reduced SUA levels in a potassium oxonate–induced hyperuricemia mouse model. The ability of MBHC10138 to utilize purine compounds for growth suggests that it can metabolize host-derived nucleosides, thereby reducing their absorption. This mechanism resembles that of Lactobacillus strain PA-3, which degrades hypoxanthine, inosine, and IMP to prevent hyperuricemia [34]. MBHC10138 showed enhanced growth in purine-supplemented media, particularly with IMP, supporting this metabolic utilization pathway.
HPLC analysis revealed that live MBHC10138 cells exhibited the strongest purine degradation activity, approximately twice as high as the lysate supernatant, indicating that enzymatic activity is primarily associated with cell membrane components. This finding aligns with reports that Lactococcus lactis possesses membrane-associated nucleoside transport systems (BmpA–NupABC and UriP) responsible for purine uptake and degradation [51]. To definitively distinguish between the contributions of live metabolic activity, structural components, and potential medium-derived factors, future studies will incorporate critical controls, including heat-killed bacterial preparations and vehicle-only (conditioned medium) groups. Furthermore, nucleoside hydrolases or phosphorylases may be involved in the reaction of cell debris and cell lysates group. Further work should be focused on the investigation of these enzymes.
In the present study, hyperuricemia mice model was made by using uricase inhibitor and high purine diet, which was widely used in the hyperuricemia study [52]. Allopurinol was administered at a dose of 5 mg/kg/day as a positive control. This dose was selected based on its established efficacy in comparable murine hyperuricemia models, where it reliably inhibits xanthine oxidase and reduces serum urate levels. Notably, even at this moderate dose, allopurinol elicited a significant hypouricemic effect, confirming the sensitivity and validity of our experimental model. The observed efficacy of the probiotic strain MBHC10138, which achieved a comparable reduction in serum uric acid, is therefore particularly noteworthy. It suggests that MBHC10138 may possess a potency similar to a clinically relevant dose of allopurinol, operating through a distinct, microbiome-mediated mechanism rather than direct enzyme inhibition. Future studies employing a broader dose range of allopurinol (e.g., from sub-therapeutic to supra-therapeutic levels) would help to more precisely calibrate the comparative potency between pharmacological and probiotic interventions.
In vivo, MBHC10138 administration substantially decreased SUA and serum XO levels in hyperuricemic mice. XO catalyzes the oxidation of hypoxanthine and xanthine to UA while generating reactive oxygen species that burden cardiovascular and renal systems [53]. Hence, suppression of XO activity by MBHC10138 likely contributed to the observed SUA reduction. Interestingly, administration of MBHC10138 at dose 3 × 108 CFU/day or 3 × 109 CFU/day both decreased SUA and serum XO levels in hyperuricemic mice, but no significant difference between the doses. Notably, a dose of 1 × 109 CFU/day is commonly employed in many animal studies of probiotic interventions [54], we selected the robustly effective high dose (3 × 109 CFU/day) for subsequent mechanistic experiments to ensure a clear signal. An alternative explanation could be that gut colonization dynamics or saturation of the relevant metabolic pathways limit further dose-dependent responses, although this hypothesis requires experimental validation in future studies. Future studies should include a broader dose–response evaluation, along with detailed toxicological assessments, to define the optimal therapeutic window of MBHC10138.
Hyperuricemia is also associated with renal dysfunction. The kidneys excrete nearly two-thirds of body urate, and excessive UA accumulation causes glomerular hyperfiltration, tubular overload, and urate-crystal-mediated inflammation [55,56]. Glomerular hypertrophy is a compensatory adaptation to sustain filtration area but eventually compresses renal capillaries [57]. In the present study, MBHC10138 significantly reduced glomerular tuft hypertrophy observed in hyperuricemic mice, restoring renal morphology and maintaining normal structure comparable with controls. In addition, MBHC10138 normalized serum creatinine and blood urea nitrogen levels, indicating nephroprotection and absence of toxicity.
Renal UA handling depends on balanced tubular secretion and reabsorption mediated by specific urate transporters. OAT1 and OAT3 on the basolateral membrane of proximal tubular cells facilitate urate excretion, whereas URAT1 on the apical membrane mediates reabsorption [42,58,59,60]. In hyperuricemia, OAT1 and OAT3 expression is typically downregulated [38], while URAT1 is upregulated, promoting UA retention. In this study, MBHC10138 significantly upregulated mOAT1 and mOAT3 and downregulated mURAT1, suggesting its potential on regulating urate excretion and reabsorption. This dual modulation explains its overall SUA-lowering effect. However, future studies directly measuring transporter protein levels, membrane localization, and functional urate flux are required to confirm this mechanism.
The intestine also plays a critical role in UA elimination, accounting for approximately 30% of total excretion [61]. Increased intestinal permeability during hyperuricemia impairs this function and promotes systemic inflammation [45]. Tight junction proteins, including ZO-1 and occludin, regulate epithelial barrier integrity and are disrupted by inflammatory cytokines [62]. Our findings demonstrated that MBHC10138 restored TJP expression and intestinal architecture damaged by hyperuricemia. Alcian-blue staining confirmed increased goblet cell numbers in the intestinal crypts, preservation of villus structure and absence of submucosal edema in MBHC10138-treated mice. These effects resemble those of Lactobacillus spp., which enhance TJP expression via strain-specific Toll-like receptor-2 signaling [63]. Thus, increased ZO-1 and occludin levels in the MBHC10138 group indicate improved intestinal barrier function and reduced permeability. To visually corroborate the observed upregulation of tight junction gene expression and the improvement in barrier function, future investigations should include immunofluorescence microscopy to assess the distribution and organization of key proteins (e.g., claudin-1, occludin) at the epithelial cell junctions.
It is well documented that intestinal barrier integrity is dynamically governed by the turnover of tight junction (TJ) proteins, whose internalization and degradation are tightly controlled by specific signaling cascades. For instance, the mycotoxin deoxynivalenol (DON) impairs the barrier by selectively activating the MAPK (p38 and ERK) pathways, leading to the upregulation of the E3 ubiquitin ligase Nedd4-2 and subsequent clathrin-mediated endocytosis and lysosomal degradation of TJ proteins such as claudin-1 and occludin [64]. Conversely, protective agents like EPA and DHA have been shown to counteract this damage by inhibiting the DON-induced MAPK activation and stabilizing TJ proteins at the membrane, partly through a PPARγ-dependent pathway [65]. Although our model is distinct from DON intoxication, the shared endpoint—TJ protein homeostasis—suggests that MBHC10138 may exert its protective effect by modulating similar regulatory hubs, such as the MAPK signaling or nuclear receptor pathways, to stabilize TJ complexes and prevent their aberrant internalization. Future studies directly assessing the activity of these specific pathways in response to MBHC10138 treatment will be crucial to validate this hypothesis and delineate its unique mechanism of action.
Gut microbial dysbiosis has been implicated in hyperuricemia progression [16,66,67]. In hyperuricemic mice, taxa expressing the allantoinase gene are diminished, disturbing urate metabolism [16]. Probiotics can restore microbial balance and promote production of short-chain fatty acids (SCFAs), which alleviate intestinal inflammation and strengthen the mucosal barrier [22]. MBHC10138 modulated the intestinal microbiota by increasing the abundance of group Clostridia vadinBB60 group and Oscillospiraceae, both of which produce butyrate—an SCFA that fuels epithelial cells and supports immune tolerance [68,69,70,71]. The enrichment of butyrate-producing-associated taxa, notably Clostridia vadinBB60 and Oscillospiraceae, suggests a potential shift in gut microbial metabolism that may favor butyrate generation. As butyrate has been implicated in metabolic and anti-inflammatory benefits [69], this could represent one plausible pathway contributing to the observed effects, although direct SCFA measurement is required for confirmation.
However, we must point out that the methodological consideration in this study for using the 16S rRNA gene V3–V4 region for microbiota profiling. While this approach robustly captures broad shifts in community structure and diversity, its resolution for discriminating closely related taxa at the species or strain level is limited. Therefore, the taxonomic assignments and subsequent ecological interpretations presented here are most confident at the family and genus levels. We have focused our discussion on trends observed at these higher taxonomic ranks and acknowledge that finer resolution would be required to unequivocally identify specific effector strains or to fully resolve the composition within certain heterogeneous families (e.g., Lachnospiraceae, Ruminococcaceae). Nonetheless, the consistent and significant shifts observed at these levels provide reliable evidence for the substantial impact of MBHC10138 on the gut ecosystem in the context of hyperuricemia.
Finally, MBHC10138 administration increased α-diversity compared with HU mice, indicating recovery of microbial richness and composition [72]. These microbial alterations may synergize with MBHC10138’s biochemical effects to reduce SUA levels. Collectively, these findings demonstrate that L. reuteri MBHC10138 attenuates hyperuricemia by degrading purine nucleosides and modulating uric acid metabolism, while concurrently restoring intestinal barrier function and reshaping the gut microbiota. Therefore, MBHC10138 represents a promising probiotic candidate for the prevention and management of hyperuricemia and associated metabolic disorders.

5. Conclusions

L. reuteri MBHC10138 effectively reduced serum uric acid and xanthine oxidase levels by degrading and utilizing purine compounds. The strain modulated renal urate transporters, restored glomerular morphology, and enhanced urate excretion, thereby improving kidney function. Furthermore, MBHC10138 strengthened intestinal barrier integrity by up-regulating tight junction proteins and increasing the abundance of Clostridia vadinBB60 group species associated with butyrate production. It also restored gut microbial diversity and composition disrupted by hyperuricemia. Collectively, these findings suggest that MBHC10138 is a promising probiotic candidate for the prevention and management of hyperuricemia.

Supplementary Materials

The following supporting information can be downloaded at: https://www.mdpi.com/article/10.3390/microorganisms14020384/s1, Figure S1: HPLC chromatogram of nucleoside degradation by MBHC10138; Figure S2: Comparison of nucleoside degradation by L. reuteri MBHC10138 and other L. reuteri strains.

Author Contributions

Conceptualization, Y.L. and J.C.; Data curation, Y.L. and J.-H.C.; Formal analysis, Y.L. and J.C.; Funding acquisition, J.C. and J.-W.S.; Investigation, Y.L. and J.-H.C.; Writing—original draft, J.C. and Y.L.; Writing—review & editing, J.C., J.-H.C. and J.-W.S. All authors have read and agreed to the published version of the manuscript.

Funding

This research was funded by the Start-up Fund for Scientific Research of High-level Talents of Wuyi University (Grant NO. 510170040617) and the Research Capacity Enhancement Project for Key Construction Disciplines of Guangdong Province (Grant No. 2025ZDJS058). This research was also supported by the Global-Learning & Academic research institution for Master’s·PhD students, and Postdocs (LAMP) Program of the National Research Foundation of Korea (NRF) grant funded by the Ministry of Education (No. RS-2023-00301938), by the Basic Science Research Program through the NRF funded by the Ministry of Education (No. RS-2025-02663016). This study was supported by R & D budget of MBHC Co. Ltd., Korea (MBHC20258825-1).

Institutional Review Board Statement

All protocols for the animal experiment were approved by the Institutional Animal Care and Committee (IACUC) of Myongji University (MJIACUC-2021001, 18 March 2021) and thus conducted by the NIH guide for the Care and Use of Laboratory Animals.

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

Authors Jinhua Cheng, Joo-Hyung Cho and Joo-Won Suh were employed by the company MBHC Co., Ltd. The remaining authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

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Figure 1. Effects of live, lysate debris, and lysate supernatant of L. reuteri MBHC10138 on purine nucleosides. Data are presented as medians with interquartile ranges. (A) Adenosine degradation; (B) Guanosine degradation; (C) Inosine degradation. Data are presented as mean ± SD. Statistical analysis was performed using one-way ANOVA followed by Dunnett’s multiple comparisons test. Different letters (a–c) indicate significant differences (p < 0.05).
Figure 1. Effects of live, lysate debris, and lysate supernatant of L. reuteri MBHC10138 on purine nucleosides. Data are presented as medians with interquartile ranges. (A) Adenosine degradation; (B) Guanosine degradation; (C) Inosine degradation. Data are presented as mean ± SD. Statistical analysis was performed using one-way ANOVA followed by Dunnett’s multiple comparisons test. Different letters (a–c) indicate significant differences (p < 0.05).
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Figure 2. Growth of L. reuteri MBHC10138 in defined media supplemented with purine nucleotides, nucleosides, or bases. Data are presented as medians with interquartile ranges. Statistical analysis was performed using one-way ANOVA followed by Dunnett’s multiple comparisons test. * p < 0.05, ** p < 0.01, *** p < 0.001, **** p < 0.0001 vs. control.
Figure 2. Growth of L. reuteri MBHC10138 in defined media supplemented with purine nucleotides, nucleosides, or bases. Data are presented as medians with interquartile ranges. Statistical analysis was performed using one-way ANOVA followed by Dunnett’s multiple comparisons test. * p < 0.05, ** p < 0.01, *** p < 0.001, **** p < 0.0001 vs. control.
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Figure 3. Effects of L. reuteri MBHC10138 on body weight, serum uric acid, and xanthine oxidase activity. (A) Body weights; (B) Serum UA levels; (C) Serum xanthine oxidase activity. Data are presented as mean ± SD. Statistical differences were analyzed by one-way ANOVA followed by Tukey’s post hoc test. Different letters (a–c) indicate significant differences (p < 0.05).
Figure 3. Effects of L. reuteri MBHC10138 on body weight, serum uric acid, and xanthine oxidase activity. (A) Body weights; (B) Serum UA levels; (C) Serum xanthine oxidase activity. Data are presented as mean ± SD. Statistical differences were analyzed by one-way ANOVA followed by Tukey’s post hoc test. Different letters (a–c) indicate significant differences (p < 0.05).
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Figure 4. Protective effects of L. reuteri MBHC10138 on renal histology and function. (A) H&E staining (15×, scale bar = 100 μm). (B) Glomerular tuft area. (C) Serum creatinine. (D) Blood urea nitrogen. (E–G) Relative mRNA expression of mOAT1, mOAT3, and mURAT1. Data represent mean ± SD (n = 8). Statistical analysis was performed by one-way ANOVA followed by Tukey’s post hoc test. Different letters indicate significant differences (p < 0.05). Arrows indicate glomerulus. Scale bar: magnifications of 15× = 100 µm, and 50× = 20 µm.
Figure 4. Protective effects of L. reuteri MBHC10138 on renal histology and function. (A) H&E staining (15×, scale bar = 100 μm). (B) Glomerular tuft area. (C) Serum creatinine. (D) Blood urea nitrogen. (E–G) Relative mRNA expression of mOAT1, mOAT3, and mURAT1. Data represent mean ± SD (n = 8). Statistical analysis was performed by one-way ANOVA followed by Tukey’s post hoc test. Different letters indicate significant differences (p < 0.05). Arrows indicate glomerulus. Scale bar: magnifications of 15× = 100 µm, and 50× = 20 µm.
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Figure 5. Alcian blue staining and TJP expression in the intestine. (A) Intestine histopathology, magnification of 5× alcian blue staining. (B) The mRNA expression of ZO-1 in mice intestines. (C) The mRNA expression of Occludin in the intestine. (D) ELISA analyzed protein expression levels of intestine ZO-1. (E) ELISA analyzed protein expression levels of intestine occludin. (F) Quantification of Alcian blue–positive goblet cells per crypt (representative section). Numbers in parentheses indicate the total crypts analyzed (Control, n = 28; HU, n = 30; ALLO, n = 22; MBHC10138, n = 19). Scale bar: magnification of 5× = 200 μm, 15× = 100 μm. All data are shown as mean ± SD. Statistical differences among groups were analyzed by one-way ANOVA followed by Tukey’s post hoc test. Means with different superscript letters (a–c) are significantly different at p < 0.05.
Figure 5. Alcian blue staining and TJP expression in the intestine. (A) Intestine histopathology, magnification of 5× alcian blue staining. (B) The mRNA expression of ZO-1 in mice intestines. (C) The mRNA expression of Occludin in the intestine. (D) ELISA analyzed protein expression levels of intestine ZO-1. (E) ELISA analyzed protein expression levels of intestine occludin. (F) Quantification of Alcian blue–positive goblet cells per crypt (representative section). Numbers in parentheses indicate the total crypts analyzed (Control, n = 28; HU, n = 30; ALLO, n = 22; MBHC10138, n = 19). Scale bar: magnification of 5× = 200 μm, 15× = 100 μm. All data are shown as mean ± SD. Statistical differences among groups were analyzed by one-way ANOVA followed by Tukey’s post hoc test. Means with different superscript letters (a–c) are significantly different at p < 0.05.
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Figure 6. Effects of L. reuteri MBHC10138 on intestinal microbiota composition. (A) Phylum-level composition. (B) Family-level composition. (C) Firmicutes/Bacteroidetes ratio. (D) Relative abundance of Clostridia vadinBB60 group and Oscillospiraceae. (E) Shannon index (alpha diversity). Data are expressed as mean ± SD. One-way ANOVA followed by Tukey’s post hoc test; different letters indicate significant differences (p < 0.05).
Figure 6. Effects of L. reuteri MBHC10138 on intestinal microbiota composition. (A) Phylum-level composition. (B) Family-level composition. (C) Firmicutes/Bacteroidetes ratio. (D) Relative abundance of Clostridia vadinBB60 group and Oscillospiraceae. (E) Shannon index (alpha diversity). Data are expressed as mean ± SD. One-way ANOVA followed by Tukey’s post hoc test; different letters indicate significant differences (p < 0.05).
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Table 1. Primer sequences for qRT-PCR.
Table 1. Primer sequences for qRT-PCR.
GenePrimerSequence (5′-3′)Amplicon Length (bp)
OAT1ForwardGAGCAGAGGAAAGCAGAAGC123
ReverseCCCTTTAGTGCTGTGTGACG
OAT3ForwardTACAGTTGTCCGTGTCTGCT153
ReverseCTTCCTCCTTCTTGCCGTTG
URAT1ForwardAGGTCCTGACAGGTTCTGT200
ReverseCTCTGCCTTCCTCCTGTTGA
OccludinForwardCACACAGGACATGCCTCCAC189
ReverseGGCTGCCTGAAGTCATCCAC
ZO-1ForwardACAGGCCATTACGAGCCTCT107
ReverseGGAGGCTGTGGTTTGGTAGC
GAPDHForwardGGCACAGTCAAGGCTGGAATG100
ReverseATGGTGGTGAAGACGCCAGTA
OAT1, organic anion transporter 1; OAT3, organic anion transporter 3; URAT1, solute carrier family 22; ZO-1, zonula occludens-1; GAPDH, glyceraldehyde 3-phosphate dehydrogenase.
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Cheng, J.; Lee, Y.; Cho, J.-H.; Suh, J.-W. Serum Uric Acid-Reducing Effect and Intestinal Mucosal Barrier-Repairing Function of Limosilactobacillus reuteri MBHC10138. Microorganisms 2026, 14, 384. https://doi.org/10.3390/microorganisms14020384

AMA Style

Cheng J, Lee Y, Cho J-H, Suh J-W. Serum Uric Acid-Reducing Effect and Intestinal Mucosal Barrier-Repairing Function of Limosilactobacillus reuteri MBHC10138. Microorganisms. 2026; 14(2):384. https://doi.org/10.3390/microorganisms14020384

Chicago/Turabian Style

Cheng, Jinhua, Youjin Lee, Joo-Hyung Cho, and Joo-Won Suh. 2026. "Serum Uric Acid-Reducing Effect and Intestinal Mucosal Barrier-Repairing Function of Limosilactobacillus reuteri MBHC10138" Microorganisms 14, no. 2: 384. https://doi.org/10.3390/microorganisms14020384

APA Style

Cheng, J., Lee, Y., Cho, J.-H., & Suh, J.-W. (2026). Serum Uric Acid-Reducing Effect and Intestinal Mucosal Barrier-Repairing Function of Limosilactobacillus reuteri MBHC10138. Microorganisms, 14(2), 384. https://doi.org/10.3390/microorganisms14020384

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