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. 2026 Sep 22;105(12):107812. doi: 10.1016/j.psj.2026.107812

Impact of GAstV-2 infection on intestinal microbiota and renal gene transcriptome in goslings

Junyang Fan a,b,c, Sai Mao a,b,c, Bingxue Yan d, Mengxiang Wang a, Jing Zhang a, Hongying Zhang a,b,c,⁎
PMCID: PMC13634751  PMID: 42777348

Abstract

Goose astrovirus-2 (GAstV-2) is the major etiological agent responsible for hyperuricemia and visceral gout in goslings, which causes substantial economic losses to the goose-raising industry. However, the complete pathogenic mechanisms underlying GAstV-2-induced hyperuricemia have not been fully elucidated. In this study, 1-day-old healthy Yangzhou goslings were orally inoculated with GAstV-2 to establish a viral infection model. The effects of GAstV-2 infection on growth performance, serum uric acid levels, cecal microbiota structure, and renal gene expression of goslings were systematically investigated. The results showed that GAstV-2 infection caused a cumulative mortality rate of 10%–30% in goslings and significantly impaired growth performance, as reflected by decreased average daily gain and average daily feed intake as well as an increased feed-to-gain ratio (P < 0.05). In addition, GAstV-2 challenge induced severe histopathological damage to the liver and kidney, disordered serum biochemical metabolism, and excessive release of pro-inflammatory cytokines, accompanied by a significant elevation in serum uric acid levels (P < 0.05). The 16S rRNA high‑throughput sequencing results confirmed that GAstV‑2 infection significantly decreased the abundances of uric acid‑degrading beneficial bacteria, including Paenibacillus polymyxa, Oscillospira guilliermondii, and Lactococcus lactis, while markedly increasing the abundance of Clostridium perfringens, which is closely associated with uric acid synthesis. Renal transcriptomic analysis confirmed that key genes involved in purine metabolism, including PRPS, IMPDH1, and NT5E, were significantly upregulated in the kidney tissues of infected goslings, which was closely correlated with the occurrence of hyperuricemia. In conclusion, GAstV-2 infection markedly inhibits growth, triggers hepatic and renal injury, increases serum uric acid and inflammatory levels, and remodels the intestinal microbiota composition and renal transcriptomic profiles in goslings. This study systematically explored the influences of GAstV-2 infection on growth performance, hepatorenal injury, and serum uric acid metabolism in goslings, and preliminarily revealed the potential underlying mechanism of GAstV-2-induced hyperuricemia by separately analyzing intestinal microbiome and renal transcriptome data. These findings provide fundamental data and a theoretical basis for further elucidating the pathogenic mechanism of GAstV-2 and developing targeted prevention and control strategies against gosling gout.

Keywords: Gosling gout, Uric acid metabolism, Transcriptomics, Gut microbiota

Introduction

In modern goose farming, gout has become a frequent metabolic disease that seriously threatens the health of goslings (Peng et al., 2023). Accumulating evidence has implicated goose astrovirus (GAstV) as the primary etiological agent driving gout outbreaks in goslings (Fu et al., 2022). Since its initial emergence in 2015, this pathogen has caused widespread outbreaks across various regions of China, with affected geese typically presenting characteristic visceral and articular urate deposition, and has been documented to yield infection rates of 70%-80% and mortality rates approaching 50% in severe cases (Ren et al., 2025). By 2022, GAstV had established itself as a dominant threat to China's goose industry, leading to substantial economic losses estimated at over 10 billion yuan (Chen et al., 2024).

GAstV is classified as a non-enveloped, single-stranded positive-sense RNA virus with a genomic size of approximately 7.2-7.3 kilobases, comprising two main genotypes (GAstV-1 and GAstV-2), of which the latter is the prevalent strain responsible for most recent outbreaks (Wei et al., 2024; Xu et al., 2023). The virus is primarily transmitted through eggs and via the fecal-oral route (Ren et al., 2023). Upon infecting goslings, the virus rapidly replicates in vital organs such as the liver and kidneys, causing tissue damage and extensive urate deposition, with diverse pathological manifestations. Hepatic lesions are characterized by narrowed or occluded hepatic sinusoids, hepatocyte swelling and rupture, accompanied by inflammatory cell infiltration(Yin et al., 2025). Renal damage is equally severe, with progressive necrosis and fibrosis of renal tubular epithelial cells leading to impaired uric acid excretion (Bi et al., 2024; Wang et al., 2023). Moreover, GAstV infection activates the innate immune response in goslings, triggering a cytokine storm that results in cellular apoptosis and tissue fibrosis(Bi et al., 2024; Hou et al., 2024). Related studies demonstrate that GAstV infection significantly upregulates xanthine oxidase (XOD) and adenosine deaminase (ADA) activities in gosling livers—key enzymes in purine metabolism that increase uric acid production—while concurrently downregulating the expression of the urate excretion transporter multidrug resistance-associated protein 4 (MRP4) in kidneys, collectively explaining the mechanism of suppressed uric acid excretion in infected goslings (Wu et al., 2020). Notably, the infection profoundly alters intestinal microbiota by expanding pro-inflammatory bacteria at the expense of beneficial species, and this dysbiosis accelerates disease progression by disrupting host metabolic pathways and further exacerbating uric acid accumulation (Li et al., 2024).

Although previous studies have deciphered the infection mechanisms of GAstV-2, the molecular pathways responsible for elevated uric acid and alterations in the gut microbiota after infection remain unclear. Against this backdrop, the present study established a gosling gout model via oral GAstV-2 inoculation. Renal transcriptome sequencing and gut microbial composition analysis were applied to preliminarily explore the mechanism underlying GAstV-2-induced hyperuricemia in goslings.

Materials and methods

Experimental animals and virus strain

A total of 125 1-day-old healthy Yangzhou goslings were obtained from Zhengzhou Ruixiang Poultry Co., Ltd. (Zhengzhou, China). Five goslings were randomly selected, euthanized by intravenous injection (i.v.) of 20 mg/kg sodium pentobarbital, and subjected to gross necropsy. No visceral abnormalities were observed. Fecal swabs from the remaining 120 goslings were tested by reverse transcription-polymerase chain reaction (RT-PCR; primers listed in Supplementary Table 1), and all were negative for GAstV-2, indicating that the flock was free of infection and suitable for the study. The challenge inoculum was the GAstV-2 strain CH-HNXX-2021, which was archived in our laboratory and previously characterized (Zhang et al., 2025). After reactivation in embryonated goose eggs and confirmation by RT-PCR, the stock virus titer was determined to be 101.03 EID₅₀·mL⁻¹, equivalent to 1 × 109.28 genome copies/mL. The complete genome of this strain has been deposited in GenBank under accession number PX495085.1.

Animal grouping and infection

A total of 120 goslings were randomly divided into four groups (n = 30/group; 3 replicate pens, 10 goslings/pen) and reared in separate cages for 14 days under controlled conditions (26–30°C, 75–85% relative humidity [RH]). On the day of challenge, the CH-HNXX-2021 stock virus (10¹·⁰³ EID₅₀/mL) was serially diluted 10-fold in sterile phosphate-buffered saline (PBS). Goslings in the H-GAstV, M-GAstV, and L-GAstV groups were orally gavaged with 0.2 mL of the diluted virus (10⁻¹, 10⁻², and 10⁻³ of the original stock titer, respectively), while goslings in the Control group were orally gavaged with 0.2 mL of sterile PBS. Clinical signs, mortality, and gross lesions were recorded daily for 14 days to evaluate the dose-response effects of GAstV-2 on gout development. All experimental procedures were approved by the Animal Care and Use Committee of Henan Agricultural University (Approval NO. HNND20250211337) and were performed in accordance with the relevant animal ethical guidelines.

Growth performance and morbidity

Throughout the 14-day experimental period, goslings had ad libitum access to a basal diet and clean drinking water; the formulation and nutrient levels of the basal diet are presented in Supplementary Table 2. The individual body weight (BW) was recorded for each gosling, and feed residues in each pen were weighed daily before feeding. Based on these data, the average daily feed intake (ADFI = total feed intake per pen / number of experimental days), average daily gain (ADG = total weight gain per gosling / number of experimental days), and feed-to-gain ratio (F/G = ADFI / ADG) were calculated. Daily mortality was recorded, and the survival rate was calculated over the experimental period to assess the effects of different GAstV-2 challenge doses on gosling growth performance and health status.

Necropsy and sample collection

At the end of the experiment (day 14), six goslings were randomly selected from each group. Blood samples were collected via cardiac puncture, and serum was separated by centrifugation at 3,000 rpm for 15 min at 4°C. The goslings were then euthanized by intravenous injection of 20 mg/kg sodium pentobarbital, and gross lesions of the liver, kidneys, spleen, and other visceral organs were immediately examined and photographed. Within 10 min after euthanasia, liver and kidney tissue samples were collected under sterile conditions for histopathological examination and viral load determination. Meanwhile, cecal contents were aseptically harvested, snap-frozen in liquid nitrogen, and stored at −80°C for subsequent intestinal microbiota analysis.

H&E staining and histopathology

Liver and kidney tissue samples were fixed in 4% paraformaldehyde solution (BBI Life Sciences, Shanghai, China) for 24 h at room temperature. Fixed samples were then dehydrated through a graded ethanol series, cleared in xylene, and embedded in paraffin wax. Paraffin sections (4-5 μm thick) were stained with hematoxylin and eosin (H&E) using a commercial staining kit (Nanjing Jiancheng Bioengineering Institute, Nanjing, China) following the manufacturer’s instructions. Stained sections were examined under a light microscope (Olympus, Tokyo, Japan), and histopathological lesions (e.g., inflammatory cell infiltration, cell necrosis, and inflammatory foci) were recorded and photographed.

Serum biochemistry and inflammatory cytokine analysis

Serum concentrations of albumin (ALB), globulin (GLB), alanine aminotransferase (ALT), aspartate aminotransferase (AST), creatinine (CRE), and blood urea nitrogen (BUN) were determined using an automatic biochemical analyzer (Beckman Coulter, Brea, CA, USA). The levels of TNF-α (No. MM-093802), IL-1β (No. MM-36910O1), and IL-6 (No. MM-052102) in the serum were quantified in triplicate using commercial enzyme-linked immunosorbent assay (ELISA) kits (Jiangsu Meimian Industrial Co., Ltd., Yancheng, China) following the manufacturer’s protocols to ensure assay accuracy and reproducibility.

Serum UA, XOD levels, and viral load assays

Serum uric acid (UA) concentration and xanthine oxidase (XOD) activity were determined using commercial assay kits (Nanjing Jiancheng Bioengineering Institute, Nanjing, China) following the manufacturer’s instructions, with each sample assayed in three technical replicates to ensure data reliability. Total RNA was isolated from liver and kidney tissues using an RNA isolation kit (Omega Bio-tek, Norcross, GA, USA), and complementary DNA (cDNA) was synthesized by reverse transcription using the HiScript II First-Strand cDNA Synthesis Kit (Vazyme Biotech Co., Ltd., Nanjing, China). Viral load was quantified by quantitative reverse transcription polymerase chain reaction (qRT-PCR) with three technical replicates per reaction, and GAstV-2-specific primer sequences for this assay are listed in Supplementary Table S1.

Cecal microbiota composition analysis

Total microbial DNA was isolated from cecal contents using the E.Z.N.A.® Soil DNA Kit (Omega Bio-tek, Norcross, GA, USA) following the manufacturer’s instructions. DNA concentration and purity were determined using a NanoDrop 2000 spectrophotometer (Thermo Scientific) by measuring A260/A280 and A260/A230 absorbance ratios. DNA integrity was verified by 1% agarose gel electrophoresis. The V3-V4 hypervariable regions of the bacterial 16S rRNA gene were amplified using primers 338F (5′-ACTCCTACGGGAGGCAGCAG-3′) and 806R (5′-GGACTACHVGGGTWTCTAAT-3′) in a GeneAmp 9700 thermal cycler (Applied Biosystems, Foster City, CA, USA). The PCR reaction system (50 μL final volume) consisted of: 25 μL 2 × Taq PCR MasterMix, 1 μL each primer (10 μmol/L), 2 μL DNA template, and sterile ddH₂O to a final volume of 50 μL; PCR conditions: 95°C for 5 min, followed by 30 cycles of 95°C for 30 s, 55°C for 30 s, and 72°C for 45 s, with a final extension at 72°C for 10 min. PCR amplicons were purified using the AxyPrep DNA Gel Extraction Kit (Axygen Biosciences, Union City, CA, USA), and sequencing libraries were generated using the NEXTflex™ Rapid DNA-Seq Kit (Bioo Scientific, Austin, TX, USA). Library sequencing was performed on an Illumina MiSeq platform with 2 × 300 bp paired-end reads. Raw reads were quality-filtered using fastp v0.20.0 to remove low-quality reads and adapter sequences, merged using FLASH v1.2.7, and clustered into operational taxonomic units (OTUs) at a 97% sequence similarity cutoff using QIIME. Taxonomic assignment of OTUs was performed using the RDP Classifier v2.11 (70% confidence cutoff) based on the SILVA 16S rRNA database v138.2.

RNA extraction and transcriptome analysis

Total RNA was isolated from 50 mg of renal tissue using 1 mL of TRIzol reagent (Invitrogen, Carlsbad, CA, USA). Briefly, tissue was homogenized in TRIzol, followed by phase separation with 200 μL chloroform per 1 mL TRIzol. The mixture was centrifuged at 12000 × g for 15 min at 4°C, and the aqueous phase was carefully transferred to a new sterile tube. An equal volume of isopropanol was added to the aqueous phase to precipitate RNA; the resulting RNA pellet was washed twice with 75% RNase-free ethanol and then dissolved in RNase-free water. RNA quality was assessed using a NanoDrop spectrophotometer (Thermo Scientific, Wilmington, DE, USA); only samples with A260/A280 ratios of 1.8-2.0 and A260/A230 ratios ≥ 1.8 were used for subsequent analyses. Ribosomal RNA (rRNA) was depleted using the Illumina Ribo-Zero Plus Kit (Illumina, San Diego, CA, USA), and the remaining mRNA was fragmented into approximately 200 nucleotides (nt) using fragmentation buffer. Double-stranded (ds) cDNA was synthesized from fragmented mRNA by reverse transcription. After end repair, A-tailing, adapter ligation, and 12-cycle PCR amplification, cDNA libraries were generated. Library concentration was quantified using a Qubit fluorometer (Thermo Scientific), and library size was verified by agarose gel electrophoresis (main peak: 300-350 bp). Qualified libraries were pooled at equimolar ratios and sequenced on an Illumina NovaSeq 6000 S4 flow cell with 2 × 150 bp paired-end reads, generating approximately 40 M raw reads per sample. Raw reads were quality-trimmed using Trimmomatic v0.39 to remove low-quality bases (Phred score < 20) and adapter sequences. Clean reads were aligned to the reference genome (Anser cygnoides; GenBank assembly: GCF_000971095.1) using HISAT2 v2.2.1, and transcript assembly and quantification were performed using StringTie v2.2.1. Differentially expressed genes (DEGs) were identified using DESeq2 v1.34.0 with the following thresholds: |log2 fold change (FC)| ≥ 1 and P < 0.05. Gene Ontology (GO) and Kyoto Encyclopedia of Genes and Genomes (KEGG) enrichment analyses were performed to annotate DEG functions. Key DEGs were validated by qRT-PCR with three biological and three technical replicates per sample.

Statistical analysis

All experimental data were analyzed using GraphPad Prism 9 software (GraphPad Software, LLC, San Diego, CA, USA) and expressed as mean ± standard error of the mean (SEM). The Shapiro‑Wilk test was used to assess data normality, and Levene’s test was applied for homogeneity‑of‑variance evaluation. Two‑tailed Student’s t‑test was used for two‑group comparisons. One‑way analysis of variance (ANOVA) followed by Tukey’s post‑hoc test was employed for multiple‑group comparisons when normality and homogeneity‑of‑variance assumptions were met; non‑parametric tests were performed if these assumptions were violated. In RNA-seq differential expression analysis and gut microbiota LEfSe analysis, the Benjamini-Hochberg false-discovery rate (FDR) procedure was applied to the calculated P-values for multiple-testing correction to reduce false-positive bias resulting from multiple comparisons.

Results

Effect of GAstV-2 infection on mortality and visceral gout lesions of goslings

GAstV-2 infection induced dose-dependent mortality and visceral gout lesions in goslings. As shown in Fig. 1A, no mortality was observed in the control group, indicating that the rearing environment remained free of exogenous pathogenic contamination throughout the study. The high-dose GAstV group (H-GAstV) exhibited a cumulative mortality rate of 30%, whereas the medium-dose (M-GAstV) and low-dose (L-GAstV) groups showed mortality rates of 20% and 10%, respectively. At the termination of the experiment, all surviving goslings were euthanized for necropsy. Macroscopic examination revealed that control goslings possessed liver and kidney tissues with normal architecture and no grossly visible lesions. In contrast, GAstV-2-infected goslings displayed dose-dependent visceral gout lesions: the liver was firm and friable, studded with pinhead-sized hemorrhages, and white urate crystal deposits were additionally evident on the hepatic capsule in the H-GAstV group. Marked pathological changes were also observed in the kidneys: the L-GAstV group showed only scattered petechial hemorrhages; the M-GAstV group presented with dark red, moderately enlarged kidneys featuring coalescing hemorrhagic foci; and the H-GAstV group exhibited severe, dark purple renal swelling accompanied by extensive subcapsular patchy hemorrhages that obscured the corticomedullary junction (Fig. 1B) . Collectively, these findings demonstrate that GAstV-2 infection is capable of inducing mortality in goslings and elicits characteristic gout-like pathological lesions in a dose-dependent manner.

Fig. 1.

Fig. 1

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Effects of GAstV-2 infection on gosling mortality and organ damage (A) gosling mortality following GAstV-2 infection (B) impact of GAstV-2 infection on gosling organ damage.

Effect of GAstV-2 infection on growth performance of goslings

As shown in Table 1, GAstV infection adversely affected gosling growth. At 7 and 14 days post - infection, the control goslings weighed 133.93g and 246.53g, with an ADG of 11.86g. In contrast, H-GAstV group goslings weighed only 96.87g at 7 days, a 27.67% decrease from the control (P <0.05). At 14 days post-infection, the H -GAstV, M-GAstV and L-GAstV groups weighed 175.50g, 192.47g and 201.63g, showing significant weight reductions of 28.81%, 21.92% and 18.21%, respectively (P <0.01). Moreover, H-GAstV and M-GAstV groups exhibited lower ADG and ADFI along with higher F/G (P <0.05). These findings indicate that GAstV-2 infection impaired gosling growth by suppressing ADG, reducing ADFI and lowering feed efficiency.

Table 1.

The Impact of GAstV-2 Infection on the growth performance of goslings.

Group Initial BW (g) 7d BW (g) 14d BW (g) ADG (g) ADFI (g) F/G
Control 80.43 133.93a 246.53a 11.86a 30.67a 2.59a
H-GAstV 79.41 96.87b 175.50b 6.82 b 24.39b 3.59bc
M-GAstV 79.97 99.77 192.47bc 8.02 bc 27.25b 3.40b
L- GAstV 80.13 101.17 201.63c 8.66 c 26.01b 3.01ac
SEM 0.48 5.59 8.26 0.58 0.74 0.12
P-value 0.989 0.027 <0.01 <0.01 <0.01 <0.01
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Note: within a column, data points marked with different superscript letters differ significantly (P < 0.05), whereas those sharing the same superscript letter or with no letter do not differ significantly (P > 0.05).

Effect of GAstV-2 infection on hepatic and renal histological structure in goslings

To investigate the impact of GAstV-2 infection on the histoarchitecture of the liver and kidney in goslings, paraffin-embedded sections were stained with hematoxylin-eosin (H&E). In control goslings, the hepatic parenchyma was unremarkable: hepatocyte cords radiated regularly from central veins, and no pathological changes were detected. In contrast, GAstV-2 infected goslings exhibited distinct dose-dependent hepatic lesions. The predominant alteration was marked steatosis, characterized by the accumulation of discrete round lipid droplets that displaced the nuclei to the peripheral region of the cells. Hydropic degeneration was also prominent, resulting in hepatomegaly with pale, loosely arranged cytoplasm and a honeycomb-like morphology. In the high-dose group, foci of coagulative necrosis were further observed, indicating severe hepatocellular damage (Fig. 2A). Similarly, renal tissues from control goslings maintained intact glomeruli and neatly arranged tubular epithelial cells, with no interstitial fibrosis or inflammatory cell infiltration; following GAstV-2 challenge, the kidneys presented a clear dose-related pathological profile, with infected goslings showing extensive inflammatory cell infiltration and obvious necrosis of tubular epithelial cells, while significant swelling of tubular epithelial cells in the medium- and high-dose groups further highlighted virus-induced nephrotoxicity (Fig. 2B). These histopathological findings confirm that GAstV-2 exerts significant hepatotoxic and nephrotoxic effects in goslings.

Fig. 2.

Fig. 2

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Histopathological examination of liver and kidney damage in GAstV-2- infected goslings (A) Liver tissue sections demonstrating histological changes following GAstV-2 infection, with green arrows indicating inflammatory cell infiltration, blue arrows indicating hepatocyte steatosis (fatty degeneration), red arrows indicating hepatocyte vacuolar degeneration, and yellow arrows indicating hepatocyte necrosis. (B) Kidney tissue sections showing histological alterations following GAstV-2 infection, with green arrows indicating inflammatory cell infiltration, red arrows indicating renal tubular epithelial cell necrosis, and yellow arrows indicating renal tubular epithelial cell swelling.

Effect of GAstV-2 infection on serum biochemical parameters and inflammatory factor levels in goslings

The effects of GAstV-2 infection on serum biochemical indices of goslings are shown in Table 2. Compared with the Control group, the H-GAstV group showed significantly increased levels of ALT, AST, ALB, GLB, BUN, and CRE (P < 0.05), with the elevation of ALT and AST being highly significant (P < 0.01), indicating that H-GAstV infection caused severe liver damage in goslings and may affect renal metabolic function. BUN levels in the M-GAstV and L-GAstV groups were also significantly increased, while no significant changes were observed in other biochemical parameters.

Table 2.

Effects of GAstV-2 infection on biochemical indices of serum in goslings.

Group ALT (U/L) AST (U/L) ALB (g/L) GLB (g/L) BUN (mg/dL) CRE (umol/L)
Control 17.36a 46.71a 16.66a 22.92a 1.68a 1.92a
H-GAstV 35.01b 76.53b 21.64b 34.96b 3.19b 4.27b
M-GAstV 23.35ac 57.20ac 20.76 32.02 2.97b 3.1
L- GAstV 19.69a 56.03ac 19.5 31.14 2.93b 3.05
SEM 2.45 3.66 0.74 1.66 0.16 0.26
P-value 0.0016 0.0033 0.0222 0.0467 0.0002 0.0008
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Note: within a column, data points marked with different superscript letters differ significantly (P < 0.05), whereas those sharing the same superscript letter or with no letter do not differ significantly (P > 0.05).

The effects of GAstV-2 infection on serum inflammatory levels in goslings are shown in Fig. 3. Compared with the Control group, IL-1β, IL-6, and TNF-α levels in the blood of goslings infected with different doses of GAstV-2 were significantly elevated (P < 0.01). Moreover, these inflammatory cytokine levels increased with higher infection doses, indicating that GAstV-2 infection induces a strong inflammatory response in the body, with the intensity of the inflammatory response closely related to the infection dose.

Fig. 3.

Fig. 3

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Inflammatory cytokine levels in the serum of GAstV-2-infected goslings. (A) IL-1β level. (B) TNF-α level .(C) IL-6 level.

Effects of GAstV-2 infection on serum uric acid, XOD activity, and viral load in goslings

The effects of GAstV-2 infection on serum uric acid (UA), xanthine oxidase (XOD) levels, and tissue viral loads in goslings are illustrated in Fig. 4. Compared with the control group, GAstV-2 infection extremely significantly elevated the serum UA and XOD levels in goslings (P < 0.01), exhibiting a dose-dependent increasing trend (Fig. 4A-B), which suggests that the virus can disrupt the host's purine metabolism process and exacerbate oxidative stress responses. Meanwhile, the viral RNA copy numbers in the liver and kidney tissues of goslings were also extremely significantly increased (P < 0.01), with the viral load peaking in the high-dose infection group (Fig. 4C-D). Collectively, these findings indicate that GAstV‑2 infection can induce gosling mortality, reduce growth performance, trigger serum biochemical disorders, hepatic‑renal tissue damage and systemic inflammatory responses, and the high‑dose infection group exhibited the most serious pathophysiological impairments. Therefore, to elucidate the pathogenic mechanism underlying GAstV‑2‑induced gosling gout, we selected samples from the Control and H-GAstV for cecal microbiota analysis and renal transcriptome sequencing.

Fig. 4.

Fig. 4

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Effects of GAstV-2 infection on serum uric acid, XOD activity, and viral load in goslings.(A) Serum UA concentration. (B) Serum XOD concentration. (C) Viral load of GAstV in liver tissues. (D) Viral load of GAstV in kidney tissues.

Effects of GAstV-2 infection on cecal microbiota in goslings

The impact of GAstV-2 infection on the cecal microbiota of goslings is shown in Fig. 5. α-diversity indices revealed decreased microbial richness and evenness in GAstV-2 infected groups, while β-diversity analysis demonstrated distinct clustering between infected and Control groups (Fig. 5A-B). At the phylum level, GAstV-2 infection upregulated the relative abundance of Bacillota and Actinomycetota, while downregulating Bacteroidota and Pseudomonadota compared to the Control group (Fig. 5C). Species-level analysis identified increased proportions of Enterococcus cecorum and Pseudarthrobacter phenanthrenivorans, alongside reduced levels of Phocaeicola dorei and Corynebacterium bovis in infected goslings (Fig. 5D). LefSe analysis further differentiated microbial signatures: Streptococcus parasanguinis, Trueperella bernardiae, Paenibacillus polymyxa, Oscillospira guilliermondii and Lactococcus lactis were enriched in the Control group, whereas Veillonella sp. oral taxon 158, Enterococcus sp. MES2, and Clostridium perfringens dominated in HGAstV-infected groups (Fig. 5E). Functional prediction via PICRUSt2 highlighted significant upregulation of the penicillin and cephalosporin biosynthesis pathway, alongside downregulation of tetracycline biosynthesis, cyanoamino acid metabolism,polyketide sugar unit biosynthesis, Alzheimer's disease-related pathways, NOD-like receptor signaling, polycyclic aromatic hydrocarbon degradation, steroid hormone biosynthesis, and apoptosis pathways (Fig. 5F).

Fig. 5.

Fig. 5

Fig. 5

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Cecal microbial composition in GAstV-2-infected goslings. (A) α-diversity analysis. (B) β-diversity analysis. (C) Phylum - level composition analysis. (D) Species - level composition analysis. (E) LEfSe analysis of differential microbes. (F) PICRUSt2 - based functional prediction of gut microbiota.

Effects of GAstV-2 infection on renal gene expression in goslings

To investigate the molecular mechanisms underlying GAstV-2 induced gout in goslings, renal transcriptomic analysis was performed on GAstV-2 infected and Control groups. PCA revealed distinct separation in gene expression profiles between GAstV-2 infected and Control groups (Fig. 6A). Using thresholds of |log2FC| ≥1 and P-value < 0.05, 2,319 significantly upregulated genes and 2,144 downregulated genes were identified (Fig. 6B-C). GO and KEGG enrichment analyses (ClusterProfiler) demonstrated that differentially expressed genes (DEGs) were primarily enriched in 265 biological processes (BP), 49 cellular components (CC), and 170 molecular functions (MF). DEG-encoded proteins localized predominantly to the extracellular region, cell junctions, and extracellular space, exhibited catalytic activity, ion binding, and metal ion binding, and were implicated in small molecule/lipid metabolic processes and cell cycle regulation (Fig. 6D). The KEGG analysis identified 181 pathways categorized as Metabolism (99), Environmental Information Processing (31), Organismal Systems (19), Cellular Processes (18), and Genetic Information Processing (14), with no enrichment observed in Human Diseases or Drug Development-related pathways (Fig. 6E). The top 20 enriched pathways included Cell cycle, FoxO signaling pathway, Toll-like receptor signaling pathway, Carbon metabolism, and p53 signaling pathway (Fig. 6F). Notably, uric acid is the end‑product of purine metabolism. Although purine metabolism was not among the top‑20 globally enriched KEGG pathways, we performed targeted analysis on differentially expressed genes annotated to this pathway. The results demonstrated that GAstV‑2 infection exhibits significant correlations with the up‑regulation of key genes encoding PRPS (EC 2.7.6.1) , IMPDH1 (EC 1.1.1.205) and NT5E (EC 3.1.3.5) within the purine metabolic pathway, which directly drives excessive uric acid production and subsequent renal accumulation in infected goslings (Fig. 6G). To verify the accuracy of the transcriptomic sequencing results, we performed quantitative real-time reverse transcription polymerase chain reaction (qRT‑PCR) to validate six key genes in the uric acid (UA) synthesis pathway, including ectonucleoside triphosphate diphosphohydrolase 5 (ENTPD5, EC 3.6.1.6), ribonucleotide reductase regulatory subunit M2 (RRM2, EC 1.17.4.1), phosphodiesterase 3A (PDE3A, EC 3.1.4.17), guanine deaminase (GDA, EC 3.5.4.3), inosine-5′-monophosphate dehydrogenase 1 (IMPDH1, EC 1.1.1.205) and ecto-5′-nucleotidase (NT5E, EC 3.1.3.5). The primer sequences for all these genes are detailed in Supplementary Table 1. As shown in Fig. 6H, the qRT‑PCR results were highly consistent with the transcriptomic sequencing data, further indicating the accuracy and reliability of the transcriptomic sequencing results in this study.

Fig. 6.

Fig. 6

Fig. 6

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Renal gene expression profiles in GAstV-2-infected goslings. (A) Beta diversity analysis. (B) Heatmap of identified genes. (C) Volcano plot of identified genes. (D) Top 10 GO functional enrichment analyses of DEGs. (E) KEGG enrichment analysis and classification of DEGs. (F) Top 20 KEGG enrichment analyses of DEGs. (G) DEGs-based enrichment analysis of the Purine Metabolism pathway. The red color indicates upregulated genes, while the blue color indicates downregulated genes. (H) qRT - PCR validation of DEGs.

Discussion

GAstV-2 has become an important pathogenic factor for avian gout. To explore the pathogenesis of avian gout caused by GAstV-2, this study analyzed the impact of GAstV‑2 infection on goslings from multiple perspectives. The results showed that GAstV-2 infection significantly inhibits the growth performance of goslings, which is manifested as slow weight gain, reduced feed intake, and increased feed-to-gain ratio. The higher the infection dose, the more obvious the inhibitory effect, which is consistent with the research of Anping Wang, et al (Wang et al., 2023). The detection results of uric acid and XOD showed that after GAstV-2 infection, the level of uric acid and the activity of XOD were significantly increased, indicating that GAstV-2 infection can affect the uric acid generation pathway and promote the increase of uric acid.

The liver is the main site of uric acid synthesis and can regulate uric acid generation through key enzymes such as xanthine oxidase (Lou et al., 2023). Studies have shown that liver damage can increase uric‑acid production by disturbing the activities of pivotal purine‑metabolizing enzymes. Meanwhile, liver injury can also elevate serum uric acid and aggravate gout‑like symptoms via releasing pro‑inflammatory factors and impairing the body’s antioxidant capacity (Lou et al., 2023; Tang et al., 2024; Yi et al., 2020). The kidney is the core organ of uric acid excretion. Approximately 70% of uric acid is excreted through the complex process of glomerular filtration, renal tubular reabsorption, and secretion. Studies have shown that hyperuricemia and kidney health have a complex bidirectional relationship. Elevated uric acid levels can lead to the deposition of uric acid crystals in joints (causing gouty arthritis) and kidneys (causing uric acid nephropathy), while impaired renal function can further reduce uric acid excretion, forming a vicious cycle (Wang et al., 2025; Zheng and Sun, 2022). In the GAstV‑2‑infected gosling model, obvious hepatic fatty degeneration, cellular vacuolar degeneration, necrosis and inflammatory cell infiltration were observed in liver tissues, together with severe inflammatory infiltration and necrosis in kidney tissues. Therefore, both enhanced uric acid production induced by hepatic lesion and impaired uric acid excretion caused by renal damage jointly contribute to hyperuricemia after GAstV‑2 infection in goslings.

This study also evaluated the effects of GAstV‑2 infection on serum biochemical profiles of goslings. ALT and AST are critical biomarkers of hepatocyte injury; their abnormal elevation can trigger purine metabolic disturbance and indirectly accelerate uric acid production (Visaria et al., 2022; Xie et al., 2022). Globulins represent a group of plasma proteins, and their elevated levels are closely associated with hepatic inflammatory responses and autoimmune-mediated liver injury (Lauletta et al., 2016; Yeong et al., 2016). Previous studies using gouty‑arthritis mouse models have demonstrated that monosodium urate (MSU) crystal deposition induces dysregulated expression of renal urate transporters (e.g., URAT1, GLUT9), further disturbing the metabolism of blood urea nitrogen and creatinine (Guo et al., 2023; Wang et al., 2024; Zhao et al., 2017). In the gosling gout model established in the present study, GAstV‑2 infection significantly increasing GLB, ALT, AST, CRE and BUN. These serum biochemical alterations are consistent with the hepatic and renal inflammatory infiltration and necrosis observed in histopathological sections, suggesting concurrent hepatic parenchymal damage and renal functional impairment, which collectively aggravate hyperuricemia in GAstV‑2‑infected goslings.

Gut microbiota play a pivotal regulatory role in host uric acid synthesis and metabolism. Paenibacillus polymyxa can protect renal function and facilitate uric acid excretion. Previous studies have demonstrated that dietary supplementation with 1.5 mg/kg Paenibacillus polymyxa significantly reduces serum uric acid and creatinine levels in broilers (Zhou et al., 2024). As a core fermentative genus in the gut, Oscillospira guilliermondii ferments dietary fiber to produce acetate, propionate and butyrate. Among these metabolites, butyrate up-regulates the expression of the urate transporter ABCG2 and promotes intestinal uric acid excretion (Le Roy et al., 2020; Ren et al., 2024). Lactococcus lactis alleviates intestinal inflammatory damage by modulating the TLR/NF-κB signaling pathway and suppressing the release of pro-inflammatory cytokines; additionally, it degrades purine nucleosides and inhibits xanthine oxidase (XOD) activity, thereby decreasing uric acid biosynthesis (Fu et al., 2024; Zhou et al., 2026). Clostridium perfringens secretes virulence factors such as β2-toxin to disrupt intestinal barrier integrity and induce intestinal inflammation and cell apoptosis (Gao et al., 2020). Its infection markedly elevates the contents of xanthine, uric acid and urate in duck spleen tissues (Zeng et al., 2025). Our experimental results showed that GAstV‑2 infection significantly reduced the relative abundances of uric acid metabolism‑beneficial bacteria, including Paenibacillus polymyxa, Oscillospira guilliermondii, and Lactococcus lactis, in the cecum of goslings, while markedly increasing the abundance of Clostridium perfringens, a bacterium that facilitates uric acid production. Based on these findings, we speculate that GAstV-2-driven alterations in the intestinal microbial community may contribute to elevated host XOD activity, suppressed expression of the urate transporter ABCG2, and activation of systemic inflammatory cascades. These microbial perturbations may ultimately induce hepatic and renal functional disorders, disrupt host uric acid homeostasis, and contribute to uric acid metabolic disorders.

Although the liver is the primary organ for uric‑acid synthesis, our study revealed that GAstV‑2 exhibits strong renal tropism, with higher viral loads in kidney tissues than in other organs. This observation is consistent with the findings reported by Bi et al. (Bi et al., 2024). Accordingly, we hypothesize that the kidney may play a more critical role than the liver in the pathogenesis of gosling gout. To decipher the molecular mechanism underlying GAstV-2-induced gout in goslings, we therefore performed transcriptomic analysis on the kidney, the major target organ of GAstV-2.The results showed that differentially expressed genes following GAstV‑2 infection were enriched in pathways including FoxO, Toll‑like receptor, PPAR, and fructose‑mannose metabolism, suggesting that viral infection activates renal inflammatory, immune and metabolic networks of the host and contributes to the pathological injury of gout (Kang et al., 2017; Kim et al., 2022; Li et al., 2022). PRPS is an isoform of phosphoribosyl pyrophosphate synthetase (PRS), whose core function is to catalyze the generation of phosphoribosyl pyrophosphate (PRPP)—the initiator and rate-limiting substrate for the de novo synthesis of purine nucleotides (Ahmed et al., 1999). Studies have demonstrated that the upregulated gene or protein expression of PRPS markedly increases PRPP production, thereby accelerating the de novo synthesis of purine nucleotides, and this process ultimately leads to excessive uric acid generation (Zhang et al., 2022). IMPDH1 is also a key rate-limiting enzyme in the de novo purine nucleotide synthesis pathway, which can catalyze the oxidation of inosine monophosphate (IMP) to xanthosine monophosphate (XMP). Research has shown that inhibition of IMPDH1 with mizoribine reduces the production of uric acid precursors during de novo purine nucleotide synthesis, and combined with the regulatory mechanism of uric acid excretion, it may consequently decrease the uric acid level in the organism (Myers et al., 2024). NT5E, also known as ecto-5′-nucleotidase, can catalyze the hydrolysis of adenosine monophosphate (AMP) to produce adenosine. Adenosine is then catalyzed by adenosine deaminase (ADA, corresponding to EC 3.5.4.6 mentioned above) to form inosine, which is further metabolized to hypoxanthine. Hypoxanthine is ultimately catalyzed by xanthine oxidase (XOD) to generate xanthine—a uric acid precursor—and thus participates in uric acid synthesis. Therefore, the observed mRNA up‑regulation of NT5E theoretically may indirectly promote subsequent metabolic flux toward uric acid generation by increasing adenosine supply, thereby potentially participating in the regulation of uric acid levels (Duarte et al., 2006; Savio et al., 2021). In this study, transcriptome analysis suggested that GAstV‑2 infection exhibits significant correlations with the upregulated expression of PRPS, IMPDH1 and NT5E in renal tissues. We thus speculate that GAstV-2 may synergistically drive the disorder of uric acid generation through the simultaneous activation of the expression of these three key metabolic enzymes in the kidney via both the de novo purine nucleotide synthesis and purine salvage pathways. Specifically, the upregulated expression of PRPS can increase the supply of PRPP, the initial substrate for purine nucleotide synthesis, and provide the material basis for uric acid generation; the elevated expression of IMPDH1 can accelerate the conversion of IMP to XMP in the de novo purine nucleotide synthesis pathway, further increasing the reserve of intermediate products for uric acid generation; while the upregulated expression of NT5E can provide more substrate sources for XOD-catalyzed uric acid generation indirectly by promoting the hydrolysis of AMP to adenosine. The synergistic effect of the three enzymes disrupts the dynamic balance between uric acid generation and metabolism in goslings, leading to an abnormal increase in uric acid levels, which in turn exacerbates the inflammatory injury and dysfunction of renal tissues, and ultimately participates in the pathological process of GAstV-2-induced gout in goslings.

Conclusion

In summary, this study established a GAstV-2 infection model in goslings. The results showed that GAstV-2 infection dose-dependently retarded growth, triggered hepatorenal histological damage and systemic inflammation, and increased serum uric acid and xanthine oxidase levels. Microbiome analysis demonstrated that GAstV-2 infection disrupted cecal microbial homeostasis by significantly reducing the abundance of uric acid-degrading beneficial bacteria including Paenibacillus polymyxa, Oscillospira guilliermondii and Lactococcus lactis, while enriching the uric acid synthesis-related pathogen Clostridium perfringens. Renal transcriptome analysis further revealed that GAstV-2 infection significantly upregulated the key purine metabolism genes PRPS, IMPDH1, and NT5E, thereby activating purine synthesis and promoting excessive uric acid accumulation. Collectively, these findings clarify the integrated mechanism linking intestinal microbiota dysbiosis and renal purine metabolic disorder in GAstV-2-induced hyperuricemia and gout, providing fundamental evidence for the pathogenesis research and targeted prevention of gosling gout.

Author contribution

Junyang Fan substantially participated in the research design, comprehensive experimental implementation, sample processing, raw data collection, statistical analysis, and result interpretation. He independently drafted the original manuscript, approved the final version for publication, and takes full public accountability for all aspects of this work, ensuring that any issues regarding the accuracy or integrity of the research are appropriately investigated and resolved. Sai Mao contributed to the conception and design of the research framework, guided data analysis and result interpretation, and critically revised the key intellectual content of the manuscript. She approved the final manuscript and accepts full responsibility for the accuracy and integrity of the reported findings. Bingxue Yan, Mengxiang Wang, and Jing Zhang substantially participated in experimental implementation, sample pretreatment, data collection and organization, as well as preliminary data analysis and result interpretation. They revised the key academic content of the manuscript, approved the final version, and assume public responsibility for their respective contributions. As the sole corresponding author, Hongying Zhang oversaw the overall research design, project administration, and funding acquisition. She critically reviewed and finalized the manuscript for academic rigor and accuracy, approved the final published version, and takes overall accountability for resolving any research-related accuracy or integrity concerns. All authors have read and approved the final manuscript.

Disclosures

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

Acknowledgments

This research was financially supported by the Joint Fund for Science and Technology Research of Henan Province (252301420064), the Natural Science Foundation of Henan Province (252300421660), and the Key Research and Development Program of Henan Province (221111111300). The authors sincerely thank Mingfan Yang and Yue Jin for providing experimental sites and equipment that facilitated the smooth completion of this study.

Footnotes

Supplementary material associated with this article can be found, in the online version, at doi:10.1016/j.psj.2026.107812.

Appendix. Supplementary materials

mmc1.docx (22.6KB, docx)
mmc2.docx (17KB, docx)

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