Metagenomic Analysis Reveals Adaptive Responses of Intestinal Microbial Community in Penaeus vannamei to Hypersaline Conditions

Abstract

The intestinal microbiota plays a vital role in host health and environmental adaptation. However, the response of the gut microbial community in Penaeus vannamei to hypersaline conditions remains poorly understood. In this study, we used metagenomic sequencing to compare the structural and functional profiles of intestinal bacteria in shrimp reared in the L-, M- and H-salinity groups. Alpha-diversity increased significantly with salinity, and PCoA revealed clear separation of microbial communities among groups. Core species analysis showed that five of the seven shared core taxa belonged to Vibrio. Microbial source tracking indicated that the proportion of environmentally derived bacteria increased with salinity. Co-occurrence networks under M and H salinities were more complex but maintained stability comparable to L. Notably, the low-salinity group was enriched with potential pathogens (e.g., Vibrio, Chryseobacterium) and infection-related functions. Functional analysis revealed that the high-salt H group exhibited enrichment of enzymes such as proline dehydrogenase (PutB), glutamate-cysteine ligase (GshA), and methyltransferases (HpnR). These enzymes interconnect compatible solutes including L-proline, L-glutamate, betaine, dimethylglycine, and glutathione, playing a crucial role in enhancing microbial osmoprotection. Furthermore, shared functions across salinities were associated with energy metabolism, protein synthesis, osmoprotection, and antioxidation. These findings, for the first time, simultaneously reveal the potential pathogenic characteristics of the L-salinity group and the adaptation mechanisms of the H-salinity group to hypersaline environments from both structural and functional perspectives of shrimp intestinal microbiota, providing insights for health management in high-salinity aquaculture.

1. Introduction

Penaeus vannamei, commonly known as the Pacific white shrimp, is native to the tropical Pacific coast of South America. Because of its rapid growth rate, high feed efficiency, strong stress tolerance, and broad salinity adaptability, this species has been extensively cultured worldwide and holds substantial economic value [1,2]. Despite these advantages, frequent outbreaks of bacterial diseases have seriously constrained the sustainable development of the P. vannamei aquaculture industry, resulting in significant economic losses. Among these diseases, acute hepatopancreatic necrosis disease (AHPND), hepatopancreatic necrosis syndrome (HPNS), and white feces syndrome (WFS) are the most prevalent and destructive [1,3,4,5]. Accumulating evidence further indicates that imbalances in the intestinal microbiota—particularly disruptions of core microbial taxa—are closely associated with the onset and progression of bacterial infections in P. vannamei [6,7,8].

The intestinal microbiota, as a vital symbiotic system of the host, plays crucial roles in nutrient absorption, physiological regulation, immune defense, and the maintenance of osmotic balance [9,10,11,12]. In aquatic animals, gut microbial communities are shaped by a complex interplay of both biotic and abiotic factors. However, previous studies have predominantly focused on dietary composition, trophic level, and biotic influences, such as developmental stage and pathogen exposure [4,13,14]. In contrast, growing evidence indicates that salinity represents a key environmental driver of intestinal microbial diversity and community structure [15,16] and consequently exerts a significant influence on host health [1,4,17]. Beyond shaping community composition, salinity can also profoundly remodel gut microbial metabolic pathways, including carbohydrate, lipid, and amino acid metabolism; energy conversion [12,17,18,19]; DNA replication and transcription; protein synthesis and modification [20]; and the biosynthesis of osmoprotectants (e.g., proline, glutamate, and betaine) and antioxidants (e.g., glutathione and peroxidases) [12,17,21,22]. Despite these advances, existing studies on the intestinal microbiota of bivalves and crustaceans have largely concentrated on low-salinity (<35‰) or brackish water conditions [17,23,24,25,26]. By comparison, research conducted under high-salinity environments remains limited and has relied almost exclusively on 16S rRNA gene sequencing [27,28], which primarily enables robust characterization of microbial community structure. As a result, functional investigations of these microbial communities based on metagenomic have been reported scarcely.

In northern coastal China, the use of primary salt-evaporation ponds for P. vannamei culture has enhanced the integrated utilization of coastal salt fields along the Yellow and Bohai Seas. These ponds, commonly referred to in China as “Dawangzi” (large salt-pan ponds), are characterized by extensive farming areas and relatively low shrimp stocking densities. However, the persistently high salinity in these ponds often leads to reduced shrimp yields, thereby constraining economic returns. The gut microbiota plays a pivotal role in shrimp growth, immune function, and disease resistance; yet, the ecological and functional responses of intestinal microbial communities to high-salinity stress remain poorly understood. To address this knowledge gap, metagenomic sequencing was employed in the present study, for the first time, to comprehensively characterize the structural and functional profiles of the intestinal bacterial communities of P. vannamei under high-salinity conditions. Specifically, this study aimed to: (i) elucidate variations in gut bacterial community structure under hypersaline conditions; (ii) explore the relationships between gut microbial features and host health under hypersaline stress; and (iii) uncover the adaptive mechanisms by which intestinal bacteria respond to hypersaline environments. Overall, this study provides new insights into the regulatory effects of salinity on the gut microbiota of P. vannamei and offers a scientific basis for promoting healthy and sustainable shrimp aquaculture under high-salinity conditions.

2. Materials and Methods

2.1. Sample Collection and Measurement of Physicochemical Parameters

Sampling was conducted in July 2021 in Bohai Seafoods Co., Ltd., Binzhou, Shandong Province, China (38.03–38.06° N, 117.56–118.01° E). Intestinal samples of P. vannamei were collected from three ponds (i.e., L, M and H) with different salinity levels: 31.56 ± 0.85 psu (Practical Salinity Unit), 39.62 ± 1.23 psu, and 51.23 ± 0.46 psu, with areas of L (106.7 ha), M (133 ha), and H (167 ha). The stocking density was consistently maintained at 150,000 individuals per hectare. Each pond included three sampling sites, and five shrimp were collected at each site [29]. In addition, 1 L of water was sampled at a depth of 0.5 m at each site. All shrimp originated from the same batch, were reared for the same duration, and had comparable average body weights. Shrimp intestines were dissected using sterilized surgical instruments and immediately transferred into a sterile 2 mL centrifuge tube. All samples were flash-frozen in liquid nitrogen and stored at −80 °C until genomic DNA extraction. Some water samples were directly analyzed for total nitrogen (TN) and total phosphorus (TP), whereas others were filtered through 0.45 μm glass fiber membranes using a vacuum filtration pump. The resulting filtrates were used to determine total ammonia nitrogen (NH₄⁺-N), nitrate nitrogen (NO₃⁻-N), nitrite nitrogen (NO₂⁻-N), sulfide, and orthophosphate (PO₄³⁻-P). All parameters were quantified using an automatic discrete analyzer (Model CleverChem 380, DeChem-Tech, Hamburg, Germany). In addition, some water samples were filtered through pre-combusted (450 °C for 4 h) Whatman GF/F filters, and the filtrates were analyzed for dissolved organic carbon (DOC) with a TOC analyzer (Multi N/C 2100s, Analytik Jena, Jena, Germany). In situ measurements of temperature, pH, DO (dissolved oxygen), and salinity were obtained using a YSI handheld multi-parameter instrument(Model YSI ProPlus, YSI Incorporated, St. Petersburg, FL, USA).

2.2. DNA Extraction, Library Construction, and Sequencing

Total intestinal DNA was extracted using the QIAamp DNA Microbiome Kit (Qiagen, Germantown, MD, USA) following the manufacturer’s instructions. DNA concentration was quantified with a Quantus Fluorometer (PicoGreen, Eugene, OR, USA), and integrity was verified by agarose gel electrophoresis. DNA samples with concentrations > 10 ng µL⁻¹ were transferred into sterile centrifuge tubes and stored at −80 °C until further analysis. Metagenomic libraries were prepared using the NEXTFLEX^® Rapid DNA-Seq Kit (Bioo Scientific, Austin, TX, USA). Briefly, genomic DNA was randomly sheared, and ~400 bp fragments were selected for end repair and adapter ligation. The ligated products were then PCR-amplified and purified to obtain the final libraries. For each sample, triplicate PCR systems were set up for amplification with the following conditions: initial denaturation at 98 °C for 2 min, 10 cycles at 98 °C for 30 s, 65 °C for 30 s, 72 °C for 60 s, and final extension at 72 °C for 4 min. Then the triplicate amplifications of each sample were pooled and purified with a PCR fragment purification kit from Takara (Kusatsu, Japan). Library concentration was determined using a Qubit fluorometer and quantitative real-time PCR (qPCR), while fragment size distribution was assessed using a bioanalyzer. The qualified libraries were sequenced on the Illumina NovaSeq 6000 platform by Majorbio Bio-pharm Technology Co., Ltd. (Shanghai, China). The raw sequencing data for each sample all exceed 14 Gbp. Raw sequence data were submitted to the NCBI Sequence Read Archive (SRA) database under accession number PRJNA1356844.

2.3. Metagenomic Data Processing

Raw reads of sequencing data were processed using Fastp (v0.20.0) for quality control to obtain high-quality clean reads. The steps included: (a) removal of adapter sequences from both 3′ and 5′ ends; (b) elimination of reads shorter than 50 bp; and (c) exclusion of reads with an average Phred quality score < 20 and containing ambiguous bases (N). High-quality reads were then aligned to the P. vannamei reference genome (https://www.ncbi.nlm.nih.gov/datasets/genome/GCF_042767895.1/, accessed on 8 August 2025) using BWA (v0.7.17) to remove host-derived sequences, yielding clean reads for downstream analysis. De novo assembly was performed with MEGAHIT (v1.1.2), and contigs ≥ 300 bp were retained. Open reading frames (ORFs) were predicted using Prodigal (v2.6.3), with ORFs shorter than 100 nt filtered out. The predicted gene sequences were clustered using CD-HIT (v4.6.1) with thresholds of 90% sequence identity and 90% coverage, and the longest sequence from each cluster was selected as the representative to construct a non-redundant gene catalog. Subsequently, SOAPaligner (v2.21) was used to map clean reads from each sample to the non-redundant gene catalog (95% identity threshold), and read counts for each gene were calculated. Genes with fewer than two mapped reads were removed, resulting in a final set of non-redundant unigenes for taxonomic and functional annotation. Taxonomic annotation was performed using DIAMOND (v0.8.35) by aligning unigenes against the NCBI NR database. Based on the alignment results and gene abundance profiles, the relative abundances of taxa at various taxonomic ranks (domain, kingdom, phylum, class, order, family, genus, and species) were calculated to generate community composition profiles. For functional annotation, the amino acid sequences of unigenes were aligned against the KEGG database (version 94.2) using DIAMOND (v0.8.35), identifying corresponding KEGG Orthology (KO) terms, KEGG Enzymes (EC numbers), and KEGG Pathways. Functional abundances were quantified by integrating annotation and gene abundance data to support downstream function analysis.

2.4. Bioinformatic and Statistical Analysis

Based on the results of normality and homogeneity of variance tests, one-way analysis of variance (ANOVA) or the Kruskal–Wallis test was applied to evaluate differences in physicochemical parameters, alpha-diversity indices, taxonomic abundance, and functional profiles among groups. Statistical significance was set at p < 0.05. LEfSe (Linear Discriminant Analysis Effect Size) was employed to identify group-specific microbial taxa and functional biomarkers [30]. Principal coordinate analysis (PCoA) and redundancy analysis (RDA) were performed in R (v4.4.0) using the vegan package. The statistical significance of PCoA was assessed by PERMANOVA, and that of RDA was evaluated using the envfit test. Core microbiota were defined as taxa present in all samples with a relative abundance ≥ 0.5% [31]. The origins of intestinal bacteria were traced using FEAST [32]. At the species level, significant correlations (Spearman’s |r| > 0.8, p < 0.001) were used to construct bacterial co-occurrence networks [33], which were visualized with Gephi (v0.10.1). Network complexity was quantified using the igraph package (v1.3.4) in R, including metrics such as node number, edge number, positive/negative correlation ratio, average degree, network density, and modularity. Modularity was used to assess the degree of network partitioning into distinct modules [34].

3. Results

3.1. Microbial Diversity Analysis

The alpha-diversity analysis revealed the effects of salinity on the intestinal bacterial communities (Figure 1a,b). Both the Chao (R² = 0.47, p = 0.042) and Shannon (R² = 0.47, p = 0.042) indices increased significantly with rising salinity (Figure 1a,b). The PCoA demonstrated a clear separation of gut samples among the L, M, and H salinity groups (PERMANOVA, R² = 0.382, p = 0.005) (Figure 1c).

3.2. Correlation Analysis of Environmental Factors

Salinity showed an increasing trend across the three ponds (L, M, and H) and differed significantly among them (p < 0.05). The physicochemical parameters of the water varied along the salinity gradient (

Table 1. Physical and chemical properties in water of different salinities (Mean ± S.E.).

Parameters	L	M	H
DOC (mg/L)	13.73 ± 0.97 b	16.18 ± 0.25 c	12.05 ± 0.91 a
TAN (mg/L)	0.91 ± 0.03 c	0.75 ± 0.06 b	0.2 ± 0.03 a
Nitrite (mg/L)	0.014 ± 0.001 a	0.037 ± 0.003 b	0.054 ± 0.006 c
Nitrate (mg/L)	0.033 ± 0.015 a	0.087 ± 0.015 b	0.040 ± 0.016 a
TN (mg/L)	8.29 ± 0.42 b	8.34 ± 0.17 b	6.10 ± 0.26 a
Sulfide (mg/L)	0.060 ± 0.000 b	0.050 ± 0.000 a	0.047 ± 0.006 a
TP (mg/L)	0.15 ± 0.06 a	0.23 ± 0.14 a	0.06 ± 0.0 a
SRP (mg/L)	0.02 ± 0.01 a	0.01 ± 0.01 a	0.01 ± 0.01 a
Salinity (psu)	31.56 ± 0.85 a	39.62 ± 1.23 b	51.23 ± 0.46 c
pH	8.07 ± 0.15 a	8.23 ± 0.06 a	8.43 ± 0.06 b
DO (mg/L)	7.03 ± 0.26 a	8.3 ± 0.17 b	7.33 ± 0.13 a
Temp (°C)	28.57 ± 0.25 a	28.9 ± 0.1 a	28.8 ± 0.1 a

Note: L: low-salinity pond; M: medium-salinity pond; H: high-salinity pond; DOC: dissolved organic carbon; TAN: total ammonia nitrogen; Nitrite: nitrite nitrogen; Nitrate: nitrate nitrogen; TN: total nitrogen; TP: total phosphorus; SRP: soluble reactive phosphorus; DO: dissolved oxygen; Temp: temperature; Within a row, different letters denote significant differences (p < 0.05), and the same letter indicates no significant difference.; W Data are expressed as mean ± standard error, n = 3.

). Total ammonia nitrogen (TAN) and sulfide concentrations decreased with increasing salinity, with significantly higher levels observed at salinity L compared to the others (p < 0.05). In contrast, nitrite and pH increased with salinity, reaching the highest values at salinity H, which were significantly greater than those at salinity L and M (p < 0.05). The salinity M exhibited the highest concentrations of dissolved organic carbon (DOC), nitrate, total nitrogen (TN), total phosphorus (TP), dissolved oxygen (DO), and temperature (Temp), among which, DOC, nitrate, and DO were significantly higher than those of other salinity groups (p < 0.05), and TN was significantly higher than that of salinity H. Eight chemical and four physical factors were included in the redundancy analysis (Figure 2). Among them, DOC (R² = 0.69, p = 0.0023), TN (R² = 0.56, p = 0.032), TP (R² = 0.88, p = 0.02), and salinity (R² = 0.61, p = 0.046) were significantly correlated with the intestinal bacterial community structure of P. vannamei. These results indicate that TP, DOC, TN, and salinity are the key environmental factors shaping the bacterial composition in the shrimp intestine.

3.3. Taxonomic Composition and Variations

To further characterize the structural features and differences in intestinal bacterial communities under different salinities, taxonomic composition and comparative analysis were performed at both the phylum and genus levels (Figure 3). At the phylum level, Proteobacteria was the most dominant taxon in the shrimp gut, with an average relative abundance of 58.70%. Furthermore, the relative abundance of Proteobacteria decreased with increasing salinity (62.24%, 58.68%, and 55.19%), whereas Cyanobacteria increased (5.57%, 6.61%, and 20.67%). In addition to Proteobacteria and Cyanobacteria, Bacteroidetes and Actinobacteria were also abundant. Gemmatimonadetes was significantly enriched at salinity H (0.00%, 0.00%, and 0.02%, p < 0.05). At the species level, the dominant intestinal bacterial species included Vibrio sp. Hep-1b-8 (19.71%), Vibrio brasiliensis (4.54%), Arthrospira platensis (2.08%), and Vibrio campbellii (2.04%). The relative abundance of Vibrio sp. Hep-1b-8 decreased with salinity (27.55%, 20.73%, and 10.85%), while A. platensis (0.73%, 0.76%, and 4.76%), V. campbellii (1.28%, 1.75%, and 3.10%), Oceaniradius stylonematis (0.87%, 1.37%, and 2.11%), and Arthrospira sp. PLM2.Bin9 (0.38%, 0.50%, and 2.46%) showed increasing trends. The abundances of Vibrio sp. Hep-1b-8 and V. brasiliensis were significantly lower at salinity H compared with the salinity L (p < 0.05).

Furthermore, based on intergroup comparison analysis and LEfSe analysis, biomarker species were identified for each salinity group by selecting taxa that simultaneously exhibited high relative abundance and high LDA scores. At salinity L, seven biomarker species were identified: Vibrio sp. Hep-1b-8 (27.55%, 20.73%, and 10.85%), Vibrio cholerae (0.88%, 0.53%, and 0.30%), Sinobacterium caligoides (0.44%, 0.18%, and 0.29%), Chryseobacterium sp. NEB161 (0.41%, 0.32%, and 0.00%), Peptoniphilus harei (0.44%, 0.10%, and 0.16%), Vibrio anguillarum (0.40%, 0.15%, and 0.06%), and Chryseobacterium sp. CECT 9293 (0.28%, 0.19%, and 0.02%). At salinity M, three biomarker species were detected: Anoxybacter fermentans (0.04%, 0.58%, and 0.00%), Chryseobacterium indoltheticum (0.04%, 0.42%, and 0.00%), and Fusobacterium sp. IOR10 (0.00%, 0.31%, and 0.00%). At salinity H, two biomarker species were identified: Bacillus glennii (0.08%, 0.10%, and 0.23%) and Roseibacterium elongatum (0.01%, 0.04%, and 0.28%).

3.4. Core Species Analysis

Core species in the intestinal bacterial communities across the three salinity groups were identified based on species occurrence frequency and relative abundance (Figure 4). The numbers of core bacterial species in the L, M, and H groups were 29, 20, and 22, accounting for 2.93%, 1.31%, and 1.10% of the total species, respectively, showing a decreasing trend with increasing salinity. Similarly, the total relative abundance of core species declined with salinity, from 53.00% at salinity L to 47.40% at salinity M and 37.88% at salinity H. At salinity L, core bacteria species were primarily affiliated with Proteobacteria (17, 42.95%), Actinobacteria (3, 3.16%), and Cyanobacteria (3, 1.98%); At salinity M, they were mainly associated with Proteobacteria (12, 39.32%), Bacteroidetes (4, 2.76%), and Cyanobacteria (2, 1.69%); At salinity H, core taxa were dominated by Proteobacteria (11, 23.88%) and Cyanobacteria (10, 13.39%). Across all three salinity groups, seven core species were shared: Vibrio sp. Hep-1b-8 (19.71%), V. brasiliensis (4.54%), A. platensis (2.08%), V. campbellii (2.04%), Vibrio parahaemolyticus (1.70%), Pseudomonas syringae (0.89%), and Vibrio rotiferianus (0.56%). The total relative abundances of these seven shared core species were 35.74%, 35.80%, and 23.00% in the L, M, and H groups, respectively. In addition, the relative abundance of V. sp. Hep-1b-8 showed a significantly positive correlation with TAN, TN, and Sulfide (p < 0.05), and a significantly negative correlation with Nitrite, Salinity, and pH (p < 0.05). The relative abundance of A. platensis was significantly negatively correlated with Sulfide (p < 0.05). The relative abundance of V. campbellii exhibited a significantly positive correlation with Salinity (p < 0.05) and a significantly negative correlation with TAN (p < 0.05).

3.5. Source Analysis of Intestinal Microbiota

Source tracking analysis was employed to assess the proportion of environmental-derived bacteria in the intestine of shrimp (Figure 5). The number (974, 1494, and 1955) and proportion (3.07%, 4.70%, and 6.39%) of shared species among water, sediment, and gut increased with rising salinity. The total relative abundance of these shared species varied greatly across habitats at salinity L and M, whereas differences among habitats were smaller at salinity H. Moreover, although the dominant phyla of shared species differed substantially among habitats under L and M salinities, these differences were less pronounced under H salinity. These results suggest that the shrimp intestine under high salinity conditions harbors a greater proportion of bacteria derived from environments, including water and sediment. To verify this, microbial source tracking analysis was conducted. The analysis confirmed that, relative to the low- and medium-salinity groups, the high-salinity H group contained more water-derived and sediment-derived bacteria in the shrimp intestine. Specifically, the total contribution of environmental bacteria (water and sediment) to the shrimp intestinal microbiota showed a marked increase with rising salinity, reaching 22.6%, 25.9%, and 42.4% in the L, M, and H groups, respectively.

3.6. Co-Occurrence Network Analysis

Co-occurrence network and robustness analysis revealed that the M and H groups exhibited slightly higher values in total links, portion of negative links, average degree, and network density compared with the L group (

Table 2. Topological properties of bacterial networks in shrimp intestine of different salinities.

Network Attributes	L	M	H
Nodes	165	164	174
Total links	2356	3272	3443
Portion of positive links (%)	61.29	56.82	58.93
Portion of negative links (%)	38.71	43.18	41.07
Average degree	28.56	39.90	39.57
network density	0.17	0.24	0.23
Modularity	0.787	0.580	0.699

Note: L: low-salinity pond; M: medium-salinity pond; H: high-salinity pond.

; Figure 6). However, it showed a marginally lower modularity. The robustness values were comparable among the three salinity groups, indicating no substantial differences in overall network stability.

3.7. Metagenomic Insights into Microbial Functional Potential

To elucidate the functional implications of microbial community variation, metagenomic sequencing were conducted. Sequencing of the nine samples yielded 134.36 Gbp of raw reads, enabling the prediction of 224,912 genes (Supplementary Table S1). After redundancy removal, nonredundant genes (unigenes) were obtained. These unigenes were functionally annotated against the KEGG Pathway, KEGG Orthology (KO), and KEGG Enzyme databases (Figure 7).

PCoA based on enzyme functional annotations revealed significant differentiation in the intestinal bacterial communities among the L, M, and H salinities (PERMANOVA, R² = 0.3189, p = 0.009). In the salinity L, enriched orthologous genes from the KEGG Orthology database included K12827, K11967, K19676, K15014, K02991, K10960, K22380, K06546, K01408, and K06183, enriched enzymes from the KEGG Enzyme database included EC 2.7.10.1, 2.3.2.31, 3.4.21.46, 4.4.1.13, 2.6.1.64, 1.3.1.83, 1.3.1.111, 2.7.8.29, and 5.4.99.19, and the major enriched function pathways at level 3 from the KEGG Pathway database were Salmonella infection and Staphylococcus aureus infection. In the M-salinity group, enriched orthologous genes included K11904, K07497, K17589, K01744, K03500, K00951, K20965, K00962, K00860, and K03584, enriched enzymes included EC 2.7.1.25, 2.1.1.176, 2.1.1.185, 2.7.7.8, 5.3.1.13, 2.7.7.77, 3.1.3.100, 1.7.1.17, and 2.3.1.274, and the dominant KEGG Pathway level 3 functions included Cyanoamino acid metabolism, Fatty acid metabolism, Sulfur metabolism, Base excision repair, and Fatty acid degradation. In the H-salinity group, enriched orthologous genes included K04599, K22790, K04955, K06904, K07012, K01501, K02407, K03496, K02705, and K00104, enriched enzymes included EC 2.1.1.-, 3.2.1.20, 6.3.2.2, 2.3.3.1, 4.2.3.1, 3.5.1.88, and 1.5.5.2, and the key enriched KEGG Pathway level 3 function was Phosphonate and phosphinate metabolism. Based on occurrence frequency and relative abundance, the shared top 10 orthologous genes, enzymes, and function pathways at level 3 across different salinities were identified and visualized in a heatmap. The shared top 10 orthologous genes included K16506, K16726, K10595, K05030, K11904, K01469, K00412, K00134, K03283, and K01880. The shared top 10 enzymes were EC 2.7.11.1, 2.3.2.26, 2.3.2.27, 3.4.19.12, 3.6.4.12, 2.7.7.6, 3.6.4.13, 3.5.2.9, 2.4.1.-, and 1.11.1.7. The shared top 10 function pathway at level 3 included Metabolic pathways, Biosynthesis of secondary metabolites, Microbial metabolism in diverse environments, Carbon metabolism, Biosynthesis of amino acids, Ribosome, Glycolysis/Gluconeogenesis, Protein processing in endoplasmic reticulum, RNA transport, and Thermogenesis.

4. Discussion

The relationship between the intestinal microbiota and host health has received considerable attention [4,9,12]. In the present study, we demonstrate that salinity exerts a significant influence on both the structural and functional composition of intestinal bacterial communities in P. vannamei. Principal coordinates analysis (PCoA) revealed a clear separation of samples across the three salinity treatments (L, M, and H), confirming salinity as a key environmental factor shaping intestinal microbial composition. Furthermore, redundancy analysis (RDA) showed that salinity, together with other physicochemical variables—including total phosphorus (TP), dissolved organic carbon (DOC), and total nitrogen (TN)—collectively influences the structure of the intestinal microbiota in P. vannamei. From an ecological perspective, community diversity is closely associated with ecosystem stability. Higher diversity generally provides greater functional redundancy and versatility, thereby enhancing the capacity of microbial communities to maintain stability under environmental disturbances [35,36]. Consistent with this ecological framework, the α-diversity of intestinal bacteria (as reflected by the Chao and Shannon indices) increased with salinity in the present study, in agreement with previous reports [17,37]. However, in contrast to these findings, other studies have reported higher bacterial diversity in the gut of freshwater shrimp compared with marine shrimp [38]. Such discrepancies among studies may stem from differences in salinity ranges, diversity metrics (e.g., weighted versus unweighted indices), and sequencing depth [1], all of which can substantially influence diversity estimates. Notably, the observed increase in α-diversity with rising salinity in this study may reflect host-mediated adjustments in intestinal selective pressures under high-salinity conditions. These adjustments could facilitate the colonization of environmentally derived, salt-adapted microorganisms, thereby contributing to enhanced functional stability of the intestinal ecosystem [4,39]. Supporting this interpretation, source-tracking analysis revealed that the proportion of intestinal bacteria originating from environmental sources (water and sediment) increased with salinity.

The phylum Proteobacteria exhibit strong environmental adaptability and play essential roles in nutrient acquisition and osmotic regulation [40,41]. Within this phylum, members of the family Rhodobacteraceae are able to synthesize vitamin B₁₂, which is essential for shrimp growth [42], while simultaneously inhibiting pathogen proliferation through the production of tropodithietic acid (TDA) [43]. In addition, certain Gammaproteobacteria can synthesize compatible solutes, such as betaine and ectoine, thereby enhancing microbial tolerance to osmotic stress under fluctuating salinity conditions [40]. Similarly, the phylum Bacteroidetes harbors a wide array of carbohydrate-active enzymes involved in the degradation of dietary polysaccharides and carbohydrate metabolism [44,45], thereby partially compensating for the host’s limited capacity for carbohydrate utilization [46,47]. By contrast, members of the phylum Cyanobacteria, in addition to serving as a potential nutritional source, may also function as probiotic microorganisms that participate in intestinal metabolic processes [48]. Notably, some cyanobacteria possess trk-type K⁺ transporter genes (trkG) [41] or synthesize osmoprotectants such as betaine [49,50], both of which contribute to the maintenance of osmotic balance. Furthermore, within the phylum Actinobacteria, members of the family Cellulomonadaceae secrete a broad spectrum of hydrolytic enzymes, including amylases, xylanases, and cellulases [43]. Moreover, certain genera, such as Actinotalea, are capable of producing short-chain fatty acids (e.g., acetate), which exhibit antagonistic activity against pathogenic Vibrio species in shrimp [51,52]. Collectively, these functional attributes underscore the potential contributions of dominant bacterial taxa to host nutrition, intestinal health, and environmental adaptation. Consistent with these functional roles, the intestinal bacterial community of P. vannamei in the present study was dominated by Proteobacteria, Bacteroidetes, Cyanobacteria, and Actinobacteria, in agreement with previous reports [17,24,29,44].

In the present study, the L-salinity treatment was characterized by an enrichment of several potentially pathogenic bacteria, including Vibrio sp. Hep-1b-8, V. brasiliensis, V. anguillarum, V. cholerae, Chryseobacterium sp. NEB161, and Chryseobacterium sp. CECT 9293. Notably, numerous members of the genus Vibrio have been widely documented as causative agents of diseases in marine animals [17,53,54]. For example, V. brasiliensis has been associated with black gill disease in P. vannamei [55], whereas V. cholerae is of particular concern due to its recognized zoonotic potential [56,57]. Similarly, several species within the genus Chryseobacterium, such as C. chaponense, C. gleum, and C. indologenes, are known to cause infections in economically important fish species, including Atlantic salmon, rainbow trout, and Nile tilapia [58,59,60,61]. Taken together, the enrichment of these taxa indicates that P. vannamei reared under L-salinity conditions may be exposed to an elevated risk of bacterial infection. Consequently, under suboptimal management practices, such microbial shifts could pose a substantial threat not only to shrimp health but also to the overall stability of the culture system.

Microorganisms that are consistently present in the host are defined as the core microbiota [62,63,64]. The presence of a core microbiome has been widely documented across diverse host systems, including mammals, insects, and fish [65,66,67,68], and is generally regarded as a key component in maintaining the structural and functional stability of the intestinal microbial community [69,70]. Previous studies have shown that, although core species constitute only a small proportion of the total intestinal microbiota in shrimp, they typically exhibit high relative abundance, which tends to increase with salinity [4]. In the present study, core species accounted for less than 5% of the total community but contributed more than 35% of the overall relative abundance, which is consistent with previous observations. However, in contrast to earlier findings, both the proportion and the overall relative abundance of core species declined with increasing salinity. This discrepancy may be attributable to differences in the salinity ranges examined. At the same time, this pattern suggests that high-salinity conditions impose substantial environmental stress on the shrimp gut ecosystem, thereby markedly reshaping the composition of core microbial members. Among these core taxa, vibrios are widely distributed in marine environments and are characterized by remarkable metabolic versatility. Specifically, they can utilize a broad range of organic carbon sources, including chitin, alginate, agar, cellobiose, and fructose [71]. For example, certain strains of V. parahaemolyticus, V. fluvialis, V. mimicus, and V. alginolyticus are capable of hydrolyzing starch, proteins, lipids, gelatin, lecithin, and chitin [71,72]. Moreover, vibrios exhibit extremely short generation times; notably, the doubling time of V. parahaemolyticus can be as brief as approximately 10 min [71]. In addition, vibrios have been shown to respond rapidly and intensively to episodic nutrient inputs, such as organic matter derived from algal blooms or iron supplied by desert dust deposition [71]. Consequently, they play a pivotal role in marine carbon cycling and biogeochemical processes. Meanwhile, vibrios are capable of producing a range of digestive enzymes and play a significant role in host nutrient metabolism [29]. For example, Vibrio spp. have been shown to produce α-amylase and lipase in sea bass larvae [73], and V. parahaemolyticus can secrete extracellular proteases [74], all of which enhance host energy harvest. However, as previously discussed, many Vibrio species are opportunistic pathogens capable of inducing diseases in marine animals [17,53,54,55,56,57]. Pathogenic Vibrios exhibit a significant negative correlation with beneficial bacteria. Probiotics (i.e., Lactobacillus and Rhodovulum, etc.) colonizing the digestive tract of fish and shrimp demonstrate antimicrobial properties that can inhibit pathogens including Vibrio [17]. Furthermore, infection by Vibrio may promote the proliferation of other potential pathogens in the host intestine, such as Photobacterium, Sphingomonas, Atopostipes, Staphylococcus, and Acinetobacter. These pathogens compete with beneficial bacteria for nutrients and colonization sites, thereby reducing intestinal microbial diversity and destabilizing the community structure, which, in turn, weakens the host’s resistance to pathogenic bacteria [75,76]. Generally, disease may occur when an imbalance in the structure and function appears in intestinal microbiota, especially, between those pathogenic bacteria including Vibrio and beneficial bacteria. It has been found that elevated abundance of Vibrio is closely associated with disease progression in shrimp, lobster, and crab [17]. Additionally, the dynamics of vibrio populations are regulated by various environmental parameters, particularly temperature, salinity, and dissolved organic carbon [71]. Therefore, rational management of vibrios in shrimp aquaculture is essential. This can be achieved through measures such as appropriate feeding to control the organic load in water strictly, probiotic supplementation to enhance gut resilience and occupy ecological niches, and regular monitoring of pathogen loads, thereby maintaining vibrio abundance within a reasonable range. In summary, an optimal balance should be struck between utilizing the metabolic functions of vibrios and preventing their disease risks, ultimately achieving both ecological stability of the culture system and healthy shrimp cultivation.

Microbial co-occurrence networks are widely used to elucidate complex intra-community interactions and to identify potential keystone taxa [77]. In general, high network modularity indicates the formation of distinct subcommunities and reflects pronounced ecological niche differentiation [78]. In the present study, the reduced modularity observed in the M- and H-salinity networks suggests a partial loss of niche differentiation under high-salinity conditions compared with the L-salinity group. In such networks, nodes and links represent microbial taxa and their interactions, respectively, encompassing both negative relationships (e.g., competition, parasitism, and predation) and positive associations (e.g., mutualism) [77,79]. Previous studies have reported that increasing salinity in shrimp gut ecosystems tends to increase the number of nodes and connections, while simultaneously reducing overall network complexity and the proportion of negative correlations [4]. Consistent with these reports, the M- and H-salinity groups in the present study exhibited higher numbers of nodes and links than the L-salinity group. However, in contrast to earlier findings, both network complexity and the proportion of negative interactions were also higher under elevated salinity in this study. Moreover, network robustness did not differ significantly among salinity treatments. From an ecological perspective, networks characterized by higher connectivity and greater structural complexity are generally considered more stable and more resistant to external disturbances [80]. In addition, an increased proportion of negative interactions may further enhance network stability by limiting excessive dominance of individual taxa and promoting competitive balance [81]. Taken together, these results indicate that the shrimp gut microbiota under high-salinity conditions maintains strong structural stability and adaptive capacity despite increased environmental stress.

From a functional perspective, the metabolic capacity of the gut microbiota exhibited pronounced adaptation to salinity gradients. Notably, the PCoA based on predicted bacterial functions was consistent with that based on community structure, indicating that functional differentiation closely paralleled taxonomic shifts across different salinity conditions. Accumulating evidence suggests that intestinal microorganisms play critical roles in regulating host immune function. For instance, immune developmental defects observed in germ-free animals can be partially restored through colonization with gut microbiota [82]. In this context, previous studies have demonstrated that the potentially pathogenic bacterium V. parahaemolyticus can enhance its tolerance to salinity stress by reducing energy expenditure and strengthening oxidative stress resistance [83]. Moreover, salinity stress has been shown to regulate the expression of the pirA gene in V. parahaemolyticus, thereby modulating its virulence [84]. Furthermore, simultaneous exposure to salinity stress and V. parahaemolyticus challenge can disrupt gut microbial functions, leading to pronounced alterations in lipopolysaccharide biosynthesis pathways [85]. In addition, toxin secretion systems of pathogenic V. parahaemolyticus mediate both intra- and interspecific competition, which may further shape shrimp gut microbial diversity and community composition [86]. Consistent with these mechanisms, the L-salinity group in the present study was enriched in pathways associated with bacterial infections (e.g., Salmonella and Staphylococcus aureus infections). Correspondingly, shrimp reared under L-salinity conditions harbored multiple opportunistic pathogens, predominantly belonging to the genera Vibrio and Chryseobacterium. In contrast, no potential pathogenic taxa or disease-associated functional pathways were detected in the M- and H-salinity groups. Taken together, these results suggest that high-salinity environments may suppress the proliferation of opportunistic pathogens, thereby providing shrimp with a natural microbial defense barrier and ultimately reducing disease risk.

In this study, the fatty acid metabolism pathway, amino acid metabolism pathway, aspA (K01744, encoding aspartate ammonia lyase), recO (K03584, encoding recombination mediator protein), the base excision repair pathway, rsmB/RsmB (K03500/EC 2.1.1.176, encoding 16S rRNA methyltransferase), RlmB (EC 2.1.1.185, encoding 23S rRNA methyltransferase), and pnp/Pnp (K00962/EC 2.7.7.8, encoding polyribonucleotide nucleotidyltransferase, PNPase) were significantly enriched in the M-salinity group. Notably, previous studies have demonstrated that certain bacterial taxa can promote shrimp growth and enhance immunity and disease resistance through the production of essential nutrients, particularly fatty acids and amino acids [87]. In this context, short-chain fatty acids (SCFAs), which are primarily generated via the fermentation of indigestible carbohydrates (e.g., dietary fiber) in the gut, play crucial roles in competitive exclusion, immune modulation, and the activation of host defense responses [88]. Moreover, SCFAs have been shown to regulate the expression of ion transporter genes, thereby modulating epithelial permeability and ion absorption [12]. Similarly, amino acids are indispensable for animal growth [89,90] and are also critically involved in immune function [55,91,92], antioxidant defense [93,94], and osmotic regulation [95,96]. Importantly, L-aspartate functions as a precursor for compatible solutes such as ectoine and 5-hydroxyectoine [21] and undergoes reversible deamination catalyzed by aspartate ammonia lyase [97], highlighting its role in microbial osmoadaptation. In addition, the RecO protein participates not only in DNA repair processes [98] but also in maintaining lactate dehydrogenase stability under salt stress, thereby enhancing cellular tolerance to high-salinity conditions [99]. Consistent with this, the base excision repair pathway plays a fundamental role in correcting DNA damage induced by oxidation, alkylation, and deamination [100]. Furthermore, RsmB and RlmB are involved in rRNA methylation, which is essential for maintaining ribosomal integrity and translational fidelity [101,102,103,104]. Meanwhile, polynucleotide phosphorylase (PNPase) contributes to multiple cellular processes, including RNA turnover, DNA repair, virulence regulation, and biofilm formation [105,106,107,108]. Taken together, the enrichment of these functional pathways and genes suggests that the shrimp gut microbiota under M salinity conditions may enhance microbial adaptability to saline environments through coordinated mechanisms. These include optimizing fatty acid and amino acid metabolism, strengthening DNA repair and RNA processing systems, and maintaining ribosomal stability and translational accuracy, thereby collectively supporting microbial resilience and host–microbe homeostasis under salinity stress.

In the H-salinity group, glcD (K00104, encoding the D subunit of glycolate oxidase), MalZ (EC 3.2.1.20, α-glucosidase), GltA (EC 2.3.3.1, citrate synthase), PutB (EC 1.5.5.2, proline dehydrogenase), GshA (EC 6.3.2.2, glutamate–cysteine ligase), and HpnR (EC 2.1.1.-, methyltransferase) were significantly enriched. Glycolate oxidase catalyzes the oxidation of glycolate to glyoxylate, thereby linking glycolate metabolism with central energy metabolism [109]. Accordingly, the enrichment of these metabolic enzymes suggests an increased capacity for energy acquisition under high-salinity conditions. In parallel, osmotic regulation emerges as another key functional feature of the H-salinity microbiota. L-proline and L-glutamate function as major compatible solutes that support cellular growth and division under osmotic and other environmental stresses [21]. Moreover, glycine undergoes sequential methylation via various methyltransferases to generate betaine and dimethylglycine, which act not only as effective osmoprotectants but also as important precursors for glutathione biosynthesis [21,102]. Notably, glutathione can be directly utilized by halophilic bacteria such as Vibrio to enhance resistance to osmotic stress and to function as a key intracellular reductant for scavenging reactive oxygens, thereby maintaining redox homeostasis [21,110]. In this context, proline dehydrogenase catalyzes the oxidation of proline to pyrroline-5-carboxylate (P5C), which is subsequently converted to glutamate, thus linking proline metabolism with both osmotic balance and energy metabolism [111]. Meanwhile, glutamate–cysteine ligase, the rate-limiting enzyme in glutathione biosynthesis, functionally integrates glutamate, glycine, and glutathione metabolism [21,110]. Taken together, the enrichment of these genes and enzymes in the H-salinity group indicates a coordinated enhancement of microbial functions related to energy acquisition, compatible solute synthesis, and antioxidant defense. Consequently, the gut microbiota under high-salinity conditions exhibit strengthened energy metabolism, improved osmotic regulation, and elevated oxidative stress resistance, collectively reflecting their functional adaptation to hypersaline environments.

In addition, the glycolysis/gluconeogenesis pathways, along with petB (K00412, encoding the cytochrome b subunit of ubiquinol–cytochrome c reductase) and gapA (K00134, encoding glyceraldehyde-3-phosphate dehydrogenase), were shared across all salinity groups. Glycolysis and gluconeogenesis jointly maintain cellular glucose homeostasis [112]. Meanwhile, the cytochrome b subunit, a core component of electron transport chain complex III, plays a central role in oxidative phosphorylation [113], whereas glyceraldehyde-3-phosphate dehydrogenase catalyzes a key energy-yielding step in glycolysis [114]. In parallel, glycine functions as a precursor for key osmoprotectants such as betaine and for the antioxidant glutathione [12,21]. Collectively, the conservation of these pathways and genes across salinity groups indicates that gut microbial communities maintain a high level of metabolic activity related to energy production under different salinity conditions. Moreover, glyS1 (K01880, encoding glycyl-tRNA synthetase), Mcm8 (EC 3.6.4.12, DNA helicase), DbpA (EC 3.6.4.13, RNA helicase), and RpoZ (EC 2.7.7.6, RNA polymerase subunit omega), as well as the amino acid biosynthesis and ribosome pathways, were consistently shared among the three salinity ponds. The co-occurrence of these genes and pathways suggests that gut microbial communities retain an active capacity for transcription, translation, and protein synthesis across varying salinity regimes. In addition, oplAH/OplAH (K01469/EC 3.5.2.9, encoding 5-oxoprolinase) and Vpo1 (EC 1.11.1.7, peroxidase) were also conserved among all groups. 5-Oxoprolinase catalyzes the hydrolysis of 5-oxo-L-proline to L-glutamate, thereby linking metabolic pathways associated with osmoprotectants and antioxidants, including proline, aspartate, and glutathione [21,110]. Concurrently, peroxidases contribute to the detoxification of reactive oxygen species (ROS), playing a crucial role in alleviating oxidative stress [115]. Overall, the shared functional profiles across salinity gradients indicate that shrimp gut microbial communities maintain homeostasis through coordinated energy metabolism, sustained protein synthesis, and effective osmoprotective and antioxidant mechanisms. Consequently, these findings provide both theoretical and practical insights for optimizing shrimp aquaculture strategies in high-salinity pond systems and for the development of novel probiotic applications.

Previous studies have shown that variations in salinity can enhance microbial energy expenditure and metabolic pathways to regulate osmoadaptation of intestinal bacteria, thereby affecting both community function and bacterial survival [17,116,117,118,119,120]. Consistent with our findings, a study on P. monodon demonstrated that under high-salinity stress—compared to its optimal growth salinity—the functional profiles of its intestinal microbiota showed significant upregulation in metabolic pathways related to amino acids and carbohydrates [25]. However, this study did not observe the marked enhancement in antioxidant stress-related functional pathways found in present research. In addition, integrated analysis of the gut microbiota and intestinal transcriptome of P. monodon revealed elevated expression of NADH dehydrogenase and NADH cytochrome reductase, who are involved in multiple energy and metabolic pathways, under high salinity [121]. Enriched metabolic activity may supply the additional energy required for osmotic adjustment following salinity changes [122]. In this study, shrimp gut microbiota exhibited distinct response patterns under different levels of salinity stress. The M-salinity group appeared to enhance adaptability through multiple mechanisms, whereas the H-salinity group, exposed to stronger osmotic pressure, showed enrichment of functions associated with osmotic adjustment. Despite these differences, the shared functional characteristics across salinity gradients suggest that microorganisms consistently require elevated energy metabolism and protein synthesis to sustain normal growth and reproduction while simultaneously allocating part of their energy to deal with osmotic and oxidative stresses, which align with previous findings [17,116,117,118,119,120]. In addition, host-microbiome interactions play a crucial role in salinity tolerance [123], with microbiome responses to salinity stress being highly species-specific [124]. In this study, the high-salinity group exhibited enrichment of B. glennii and R. elongatum, both of which are reported to produce catalase and oxidase under salinities as high as 50‰ [125,126], and the host shrimp also needs to enhance energy metabolism for osmotic regulation [124]. Therefore, the bacteria enriched in the high-salinity group may contribute to enhanced host adaptation to high-salinity environments. The present findings that shrimp gut microbiota exhibit distinct alterations in their metabolic functions across varying salt conditions suggest that gut microbiota may establish functional complementarity with the host, thereby enhancing adaptation of shrimp to hypersaline aquaculture environments.

This study systematically elucidated the structural characteristics and adaptive mechanisms of the intestinal bacterial community of P. vannamei under hypersaline conditions. Nonetheless, several limitations should be acknowledged. The present work relied primarily on metagenomic and correlation-based analysis and therefore lacked experimental validation of key metabolic pathways and functional genes. To address this limitation, future studies should integrate quantitative PCR, metatranscriptomic, metabolomic, and isotopic tracing approaches to clarify the underlying regulatory mechanisms. Overall, this study provides a comprehensive framework for understanding the adaptive strategies of shrimp gut microbiota in hypersaline environments. However, further integrative multi-omics analyses combined with targeted experimental validation remain essential to substantiate and refine these adaptive mechanisms, thereby supporting the optimization of healthy and sustainable aquaculture practices in hypersaline systems.

5. Conclusions

In summary, this study systematically characterized the structural and functional features of the intestinal bacterial community of P. vannamei under hypersaline aquaculture conditions, revealing significant differences across salinity gradients. Bacterial alpha-diversity increased with salinity, accompanied by distinct shifts in community composition and functional profiles. Proteobacteria was the dominant phylum, with Vibrio sp. Hep-1b-8 and V. brasiliensis as the predominant species. The L-salinity group was enriched with potential opportunistic pathogens (e.g., Vibrio and Chryseobacterium) and bacterial infection-related functional pathways, while the proportion of environmentally derived bacteria increased with salinity. The M- and H-salinity groups exhibited higher network complexity but comparable stability to the L-salinity group, indicating enhanced adaptation to high-salinity environments. Functional enrichment in the H-salinity group highlighted strengthened osmotic regulation capabilities. The bacterial functions shared across salinities were primarily associated with energy metabolism, protein synthesis, osmoprotection, and antioxidant defense. Overall, these findings advance the understanding of how shrimp intestinal bacterial communities adapt structurally and functionally to hypersaline conditions, providing valuable insights for developing health-oriented and sustainable aquaculture strategies for P. vannamei in high-salinity environments.