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Article

Sand Substrate Thickness Regulates Growth Performance, Intestinal Antioxidant Defense, and Gut Microbiota in an Experimental Culture of Marsupenaeus japonicus

by
Xianyun Ren
1,2,
Kuangcheng Zhao
1,2,3,
Xueqiong Bian
1,2,4,
Shaoting Jia
1,2,
Ping Liu
1,2,
Jian Li
1,2,
Yuefeng Cai
1,2,3,* and
Jitao Li
1,2,*
1
State Key Laboratory of Mariculture Biobreeding and Sustainable Goods, Yellow Sea Fisheries Research Institute, Chinese Academy of Fishery Sciences, Qingdao 266071, China
2
Laboratory for Marine Fisheries Science and Food Production Processes, Qingdao Marine Science and Technology Center, Qingdao 266237, China
3
Jiangsu Key Laboratory of Marine Biotechnology, Jiangsu Ocean University, Lianyungang 222005, China
4
College of Fisheries and Life Science, Shanghai Ocean University, Shanghai 201306, China
*
Authors to whom correspondence should be addressed.
Animals 2026, 16(4), 586; https://doi.org/10.3390/ani16040586
Submission received: 8 December 2025 / Revised: 19 January 2026 / Accepted: 9 February 2026 / Published: 12 February 2026
(This article belongs to the Section Animal Physiology)
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Simple Summary

Shrimp farming often uses artificial ponds, but it is unclear how much sand should be placed at the bottom to best support shrimp health and growth. This study tested four different sand thicknesses in a 120-day farming experiment with kuruma shrimp to determine which condition was most suitable. We found that shrimp raised without sand grew more slowly and had much lower survival rates. In contrast, shrimp raised on a sand layer 10 cm thick grew faster, survived better, and showed healthier intestines. These shrimps also experienced less damage caused by internal stress and had fewer damaged intestinal cells. In addition, the community of helpful microorganisms living in the shrimp intestine was richer and more balanced when a 10 cm sand layer was used, while the absence of sand reduced this diversity. Very thick sand layers did not provide additional benefits and may increase health risks. Overall, this study shows that using an appropriate sand thickness, especially 10 cm, can improve shrimp growth, intestinal health, and long-term survival. These findings provide practical guidance for shrimp farmers and can help make shrimp production healthier, more efficient, and more sustainable.

Abstract

Kuruma shrimp (Marsupenaeus japonicus) exhibit natural sand-burrowing behavior, but the optimal sand substrate thickness for industrial farming remains unclear. This study evaluated the effects of different sand layer thicknesses on growth performance, intestinal health, oxidative status, and gut microbiota in Marsupenaeus japonicus. A 120-day controlled farming experiment was conducted using four sand substrate treatments: 0 cm (no sand), 5 cm, 10 cm, and 20 cm, with three replicate ponds per treatment. Growth indices, survival rate, intestinal histology, antioxidant enzyme activity, gene expression, and gut microbial composition were analyzed. Shrimp reared without sand showed markedly reduced growth and survival, increased intestinal damage, and higher oxidative stress. In contrast, shrimp cultured with a 10 cm sand layer exhibited improved growth and survival, lower intestinal oxidative damage and cell apoptosis, and healthier intestinal structure. This condition also supported a more diverse and stable intestinal microbial community and a lower abundance of opportunistic pathogenic bacteria compared with thinner or thicker sand layers. Overall, these results indicate that a sand substrate thickness of 10 cm provides the most favorable balance between growth, intestinal health, and microbial stability, offering practical guidance for optimizing kuruma shrimp aquaculture.

1. Introduction

Marsupenaeus japonicus (Decapoda: Penaeidae), commonly known as kuruma shrimp, is an economically important crustacean widely distributed throughout the Indo-West Pacific [1]. It is currently farmed in multiple Asian and European countries and accounts for more than 5% of global shrimp production [2]. China is the leading producer, with an annual output of 45,968 tons in 2024 [3]. Owing to its natural burrowing behavior, M. japonicus is traditionally cultivated using ecological farming approaches, and recent industrial systems increasingly incorporate sand substrates to improve production efficiency. Substrate quality is therefore a critical determinant of successful industrial farming of M. japonicus [4].
This issue is particularly relevant as high-density polyethylene (HDPE)-lined systems are increasingly replacing traditional earthen ponds, owing to their superior control of seepage, dike erosion, sediment intrusion, disease transmission, and harvesting efficiency. Given the inherently slow growth rate of M. japonicus, the integration of nursery and grow-out phases is commonly used to promote compensatory growth and shorten the culture period. In addition to substrate composition, sediment depth plays a crucial role in influencing shrimp survival and growth, as optimal conditions help maintain a stable microbial–shrimp equilibrium [5]. Burrowing behavior and sediment distribution have been well documented in earthworms [6], and soil depth preferences are known to vary among species in relation to organic matter distribution [7]. Considering the depth-dependent effects of sediment on feeding and burrowing behavior [8], the influence of sediment thickness on kuruma shrimp warrants systematic investigation.
Optimizing shrimp cultivation requires a thorough understanding of both physiological and behavioral responses. Kuruma shrimp exhibit inherent burrowing behavior and nocturnal feeding habits [4], which necessitate the provision of sandy substrates during the grow-out phase and the implementation of nighttime feeding protocols. Although substrate characteristics such as composition and particle size have been investigated [9], the sand thickness preferences of juvenile M. japonicus remain poorly characterized. We hypothesized that sediment depth influences survival and growth performance throughout the ontogeny of M. japonicus. To test this hypothesis, early juveniles were cultured under three sediment depths in a controlled grow-out experiment. This study elucidates the adaptive responses of kuruma shrimp to varying sand substrates and provides a scientific basis for optimizing healthy and efficient shrimp cultivation.

2. Material and Methods

2.1. Ethical Considerations

All animal procedures were conducted in strict accordance with the Guidelines for the Care and Use of Laboratory Animals of China. The experimental protocol was approved by the Institutional Animal Care and Use Committee (IACUC) of the Yellow Sea Fisheries Research Institute (Qingdao, China; Approval No. YSFRI2022026).

2.2. Animals Acquisition and Rearing Conditions

The experiment was carried out at the factory-based shrimp farming facilities of Changyi Haifeng Aquatic Breeding Co., Ltd., Weifang, China. Postlarvae of kuruma shrimp were obtained from wild broodstock originating from Penghu, Taiwan. The post-larvae had an initial average body length of 0.9 ± 0.1 cm and were nursery-reared for 28 days until reaching an average body length of 2.18 ± 0.12 cm and a body weight of 0.093 ± 0.01 g, after which they were transferred to the experimental culture system. A total of 12 culture ponds were used in this study. All ponds were rectangular cement tanks with an area of 8 m2 and a water depth of 1 m, each equipped with a drainage outlet located at the inner bottom side of the pond.

2.3. Culture System and Experimental Design

Four sand substrate treatments were established according to substrate thickness: 0 cm (non-sand control), 5 cm (low-sand group, LG), 10 cm (medium-sand group, MG), and 20 cm (thick-sand group, TG). Each treatment was conducted in triplicate, with 1400 shrimp stocked per pond, corresponding to an intensive culture density of 175 shrimp per square meter. All other environmental and management conditions were maintained consistently across treatments. The substrate consisted of a sandy bed composed of 70% sand (particle size 500–700 μm) and 30% soil (particle size 100–250 μm). Sundried beach sand was sieved through a 700 μm mesh and retained on a 500 μm mesh, while sun-dried pond soil was sieved through 250 μm and 100 μm meshes. The processed substrates were disinfected with bleaching powder at 60 ppm (available chlorine 20 ppm), thoroughly rinsed with clean water, and air-dried prior to use.
The experiment lasted for 120 days. When shrimp body length was less than 4.5 cm, they were fed four times daily at 06:00, 12:00, 18:00, and 24:00. Once body length exceeded 4.5 cm, feeding frequency was reduced to three times daily at 09:00, 18:00, and 24:00. During the first two months, shrimp were primarily fed frozen brine shrimp, whereas a formulated pellet diet containing 45% crude protein was used during the latter two months. Routine management procedures were performed daily at 08:00, including waste removal, partial water exchange, and the removal of molted shells. The daily water exchange volume was approximately 30% of the total culture water volume.

2.4. Collection of Samples

After the initiation of the experiment, intestinal tissue samples of M. japonicus were collected on days 60, 90, and 120. On day 120, intestinal segments from six shrimp per replicate (n = 3 replicates per treatment) were collected and immediately fixed in 4% paraformaldehyde (Sangon Biotech, Shanghai, China). Of these, three biological replicates (one shrimp per replicate) were processed for histological analysis. To minimize the physiological variability associated with the molting cycle, shrimp were sampled during the intermolt stage (Stage C) [10]. For biochemical and molecular analyses, intestinal tissue samples weighing approximately 80–100 mg were excised, rapidly frozen, and stored at −80 °C until further use.

2.5. Basal Growth Index

At the end of the experimental period, the final body weight and body length of all shrimp were measured. Survival rate (SR), body length growth rate (BLGR), weight gain rate (WGR), and specific growth rate (SGR) were calculated using the following formulas:
SR (%) = (total shrimp − dead shrimp)/total shrimp × 100
BLGR (%) = (final body length − initial body length)/initial body length × 100
WGR (%) = (final weight − initial weight)/initial weight × 100
SGR (%/day) = (ln final weight − ln initial weight)/culture duration days × 100

2.6. Observations of Intestine Structure and Apoptosis

Intestinal histomorphological analysis was performed using established methods [11]. Briefly, intestinal samples were fixed in 4% paraformaldehyde for 24 h at 4 °C and then rinsed under running water for 8 h. Samples were dehydrated through a graded ethanol series (70%, 80%, 90%, and 100%), cleared in xylene, embedded in paraffin, and sectioned at a thickness of 4 µm using a Leica RM2016 microtome (Leica Microsystems, Wetzlar, Germany). Sections were stained with hematoxylin and eosin (H&E) and examined under a light microscope (Olympus, Tokyo, Japan). Histomorphometric parameters were quantified using the CaseViewer software (V.2.4, Budapest, Hungary). Unless otherwise stated, all chemical reagents were of analytical grade and supplied by Sinopharm Group Co., Ltd. (Shanghai, China).

2.7. Analysis of Antioxidant Enzyme Activity

Intestinal tissues stored at −80 °C were thawed on ice and homogenized (1:9, w/v) in phosphate-buffered saline (PBS; pH 7.2). The homogenates were centrifuged at 9300× g for 10 min, and the supernatants were collected for subsequent analyses. Total antioxidant capacity (T-AOC; Cat. No. A015-1-1) and glutathione peroxidase (GSH-PX; Cat. No. A005-1-2) activities were measured by colorimetric methods at 520 nm and 412 nm, respectively. Superoxide dismutase (SOD; Cat. No. A001-3-1) activity was determined using the WST-1 method at 450 nm. Catalase (CAT) activity was measured using a catalase assay kit (Cat. No. A007-1-1; ammonium molybdate method) at 405 nm. Malondialdehyde (MDA; Cat. No. A003-1-1) content was quantified using the thiobarbituric acid (TBA) method at 532 nm. All assay kits were purchased from Nanjing Jiancheng Bioengineering Institute (Nanjing, China). Protein concentrations were determined using the Bradford assay with bovine serum albumin as the standard [12]. All measurements were performed in triplicate using a SpectraMax M5 microplate reader (Molecular Devices, San Jose, CA, USA).

2.8. Gene Expression Analysis of Antioxidant Capacity and Energy (Carbohydrate, Lipid and Protein) Metabolism by Real-Time Quantitative PCR (qPCR)

Approximately 80–100 mg of intestinal tissue was weighed, and total RNA was extracted using RNA Isolater Total RNA Extraction Reagent (Vazyme Biotech Co., Ltd., Nanjing, China). RNA concentration and purity were assessed using a NanoDrop ND-2000 spectrophotometer (NanoDrop Technologies, Wilmington, DE, USA). Total RNA was reverse transcribed into complementary DNA (cDNA) using the PrimerScript™ One Step RT-PCR Kit (Vazyme Biotech Co., Ltd., Nanjing, China), following the manufacturer’s instructions. The transcription levels of genes related to antioxidant capacity (SOD, CAT, and heat shock protein 70, HSP70) and cellular immune defense (lysozyme, Lzm, and caspase-3) were quantified, with β-actin used as the internal reference gene. Primer sequences for all target genes are listed in Table 1. Amplification efficiencies were determined using a serial dilution of cDNA, and all primer pairs showed efficiencies ranging from 90% to 110% (Table 1).
Quantitative PCR was performed using a StepOnePlus™ Real-Time PCR System (Applied Biosystems, Foster City, CA, USA) with SYBR Premix Ex Taq II (Vazyme Biotech Co., Ltd., Nanjing, China). Each reaction was carried out in a 20 μL volume containing 10 μL of 2× SYBR Green I Master Mix, 1.0 μL of each gene-specific primer (10 μM), 6 μL of DEPC-treated water, and 2 μL of cDNA template [13]. The thermal cycling conditions were as follows: initial denaturation at 95 °C for 30 s; 45 cycles of denaturation at 95 °C for 5 s and annealing at 58 °C for 30 s; followed by melt curve analysis at 95 °C for 5 s, 60 °C for 60 s, heating to 95 °C, and cooling to 50 °C for 30 s. Relative gene expression levels were calculated using the 2−ΔΔCT method, as described by Livak and Schmittgen [14].

2.9. Gut Microbiota Profiling

Total genomic DNA of the hindgut microbiota from the control, LG, MG, and TG groups was extracted, and DNA quality and concentration were assessed prior to downstream analyses. The V3–V4 hypervariable region of the bacterial 16S rRNA gene was amplified using the universal primers 338F (5′-ACTCCTACGGGAGGCAGCA-3′) and 806R (5′-GGACTACHVGGGTWTCTAAT-3′). Amplified products were verified by 2% agarose gel electrophoresis, purified using a commercial gel extraction kit (Axygen Biosciences, Union City, CA, USA), and quantified with a Quantus™ Fluorometer (Promega, Madison, WI, USA). Sequencing libraries were constructed using the NEXTFLEX Rapid DNA-Seq Kit (Bioo Scientific Corp., Austin, TX, USA) and paired-end sequencing (2 × 300 bp) was performed on an Illumina MiSeq PE300 platform (Illumina, San Diego, CA, USA) according to standard protocols provided by Majorbio Bio-Pharm Technology (Shanghai, China). Raw sequencing data were processed, and operational taxonomic units (OTUs) were clustered at a 97% sequence similarity threshold using UPARSE (http://www.drive5.com/uparse/, accessed 15 January 2025). Alpha diversity indices and rarefaction curves were calculated using Mothur (http://www.mothur.org/, accessed 15 January 2025), while beta diversity analyses were conducted using QIIME (v2020.2.0). Data visualization and statistical plotting were performed using R software (v3.3.1).

2.10. Statistical Analysis

All statistical analyses were performed using IBM® SPSS Statistics Version 26.0 (IBM Corp., Armonk, NY, USA), with statistical significance set at p < 0.05. Quantitative data are presented as the mean ± standard error (SE) based on three biological replicates (n = 3). Prior to analysis, data were assessed for normality and homogeneity of variance. When these assumptions were satisfied, differences among groups were evaluated using one-way analysis of variance (ANOVA), followed by Duncan’s multiple range test for post hoc comparisons. When the assumptions of normality or homoscedasticity were not met, the nonparametric Kruskal–Wallis test was applied. Principal component analysis (PCA) was conducted using OriginPro 2021 software (OriginLab Corporation, Northampton, MA, USA) to explore relationships among variables across experimental groups.

3. Results

3.1. Growth Performance

As shown in Table 2, the growth rate, weight gain rate, and SGR of M. japonicus in the control group were consistently the lowest across all sampling periods compared with those in the sand substrate groups, with significant differences observed (p < 0.05). Compared with the MG and TG groups, shrimp in the LG group exhibited inferior growth performance. By day 120, the growth rate, weight gain rate, and SGR in the LG group were significantly lower than those in the MG and TG groups. Throughout the experimental period, shrimp cultured in the MG and TG groups maintained relatively high growth levels, with no significant differences in growth performance between these two groups. Survival rates in the control, LG, MG, and TG groups were 20.5%, 78.5%, 85.7%, and 87.3%, respectively. The control group showed the lowest survival rate, which was significantly lower than that of all sand substrate groups (p < 0.05). Survival in the LG group was significantly lower than that in the MG and TG groups, whereas the TG group exhibited the highest survival rate, although no significant difference was detected between the TG and MG groups.

3.2. Analysis of Oxidative Stress

As shown in Table 3, T-AOC levels in M. japonicus varied significantly among sediment treatments. At day 90, T-AOC declined markedly in all groups, with the most pronounced decrease observed in the control group, resulting in significantly lower levels than those in the sand substrate groups. By day 120, T-AOC levels continued to decrease across all treatments; however, values in the MG and TG groups remained significantly higher than those in the control group. SOD activity increased by day 90 and subsequently decreased by day 120 in all groups. SOD activity was consistently highest in the control group throughout the experimental period. At day 120, SOD activity in the MG group was significantly lower than that in the control group but did not differ significantly from that in the TG group. CAT activity differed among sediment treatments. At day 60, CAT activity was significantly lower in all sand substrate groups than in the control group (p < 0.05). CAT activity increased by day 90 and declined by day 120 in the sand substrate groups, whereas it increased progressively in the control group throughout the culture period. GSH content decreased by day 90 and increased by day 120 in the control, LG, and TG groups. In contrast, GSH levels increased continuously in the MG group during the experiment, resulting in significantly higher values than those in the control group. At day 90, GSH-PX activity was significantly lower in the control group and significantly higher in the TG group compared with the other groups, while activities in the LG and MG groups decreased but without significant differences. By day 120, GSH-PX activity declined significantly in all groups, with the most pronounced decrease observed in the MG group, which differed significantly from the other treatments. MDA content, an indicator of intestinal oxidative damage, exhibited distinct temporal patterns among groups. At day 90, MDA content decreased significantly in the control group but increased slightly by day 120. In contrast, MDA content decreased progressively in all sand substrate groups throughout the experimental period. At day 60, MDA content was significantly lower in the TG group than in the other groups; however, this difference gradually diminished over time. By day 120, the lowest MDA content was observed in the MG group, which was significantly lower than that in all other groups.
As shown in Figure 1A, SOD expression was significantly higher in the TG group than in the other groups at days 60 and 90, whereas the highest expression level at day 120 was observed in the control group. Figure 1B shows that CAT expression was highest in the control group at days 60 and 90; at day 120, CAT expression peaked in the MG group but did not differ significantly from that in the TG group. Figure 1C demonstrates that Hsp70 expression remained relatively stable in the control group throughout the experiment, whereas sand substrate groups exhibited a biphasic pattern characterized by an initial decrease followed by an increase. Figure 1D shows consistently low Po expression in the control group, with peak expression observed in the TG group at days 60 and 90 and shifting to the LG group by day 120. As shown in Figure 1E,F, Lzm and caspase-3 expression in the sand substrate groups followed a U-shaped pattern, decreasing initially and increasing thereafter; at day 120, the highest expression levels were detected in the TG group, while the control group exhibited the lowest levels.

3.3. Changes of the Intestine Histological Structure of M. japonicus

After 120 days of culture, H&E staining revealed mild swelling and congestion of the intestinal wall in the control group (Figure 2). In contrast, the LG group exhibited pronounced intestinal wall thickening, which was primarily attributable to severe congestion and inflammatory infiltration. No obvious congestion or inflammation was observed in the MG or TG groups, and intestinal wall thickness in these groups was significantly reduced compared with that in the control and LG groups. Apoptosis in intestinal tissues was assessed using the TUNEL assay (apoptotic cells shown in green; Figure 3). The basal lamina exhibited swelling and fragmentation in response to differences in substrate thickness. Quantitative analysis showed that apoptotic cell numbers were lowest in the control group and highest in the LG group. With increasing substrate thickness, the number of apoptotic cells decreased significantly in the MG and TG groups.

3.4. Gut Microbiota

3.4.1. Diversity

The Ace and Chao indices were used to assess microbial abundance, whereas the Shannon and Simpson indices were used to evaluate microbial diversity. As shown in Table 4, the Ace and Chao indices in the control group were significantly lower than those in the sand substrate groups, indicating that the absence of a sand substrate reduced intestinal microbial abundance in M. japonicus. The Simpson diversity index was significantly higher in the MG group than in the LG and TG groups, suggesting that a sand substrate thickness of 10 cm was more conducive to enhancing intestinal microbial diversity in M. japonicus.
As illustrated in Figure 4, NMDS and PCoA analyses demonstrated a clear separation between the intestinal bacterial community of the control group and those of the sand substrate groups, whereas the sand substrate groups exhibited partial overlap. In terms of sample distribution, samples from the TG group were more tightly clustered, while samples from the other groups showed greater dispersion.

3.4.2. Key Species Responsible for Differences in Gut Microbiota Across Sand Thicknesses

At the phylum level, the predominant intestinal microbiota (relative abundance ≥ 1%) across all experimental groups of M. japonicus consisted mainly of Proteobacteria, Bacteroidetes, Actinobacteriota, and Firmicutes (Figure S1). The relative abundances of Proteobacteria and Firmicutes were lower in the sand substrate groups than in the control group, whereas Actinobacteriota were more abundant in the sand substrate groups. At the genus level, dominant taxa (relative abundance ≥ 1%) included Vibrio, Photobacterium, Ruegeria, and members of the families Rhodobacteraceae, Thiotrichaceae, and Flavobacteriaceae (Figure S2). Compared with the control group, the sand substrate groups exhibited higher proportions of Photobacterium and Actinomarinales, along with lower proportions of Rhodobacteraceae, Ruegeria, Thiotrichaceae, Enterococcus, and Motilimonas. The dominant taxa in each group were identified as Thiotrichaceae in the control group, Rhodobacteraceae in the LG group, Photobacterium in the MG group, and Vibrio in the TG group. Notably, Photobacterium in the LG group and Vibrio in the TG group each reached relative abundances of approximately 20%, which were significantly higher than those observed in the other groups.
A total of 219 amplicon sequence variants (ASVs) were shared among all four groups, with 283 ASVs shared between the control and LG groups, 277 between the control and MG groups, and 262 between the control and TG groups (Figure S3). The control group contained the highest number of unique ASVs (81), whereas the MG group had the lowest number (59). Pie chart analysis showed that the three most abundant taxa in the control and LG groups were Thiotrichaceae, Rhodobacteraceae, and Ruegeria. In the control and MG groups, the dominant taxa were Photobacterium, Rhodobacteraceae, and Thiotrichaceae, while the control and TG groups were dominated by Vibrio, Rhodobacteraceae, and Ruegeria. Across all groups, the three most prevalent taxa were Vibrio, Photobacterium, and Rhodobacteraceae.
The Kruskal–Wallis rank sum test results indicated that the relative abundance of Proteobacteria was significantly higher in the control group than in the sand substrate groups, whereas the abundances of Actinobacteriota, Chloroflexi, and Patescibacteria were significantly lower in the control group (Figures S4 and S5). Among the sand substrate groups, Proteobacteria abundance was highest in the MG group and lowest in the LG group. Actinobacteriota accounted for a substantial proportion of the microbiota in the sand substrate groups, indicating marked dominance, while Chloroflexi abundance was significantly higher in the LG group than in the other groups.
As shown in Figure S6, LEfSe analysis revealed that Pseudoalteromonadaceae were significantly enriched in the control group compared with the LG group, whereas Actinobacteriota were significantly enriched in the LG group. In comparisons between the control and MG groups, Alteromonadales were more enriched in the control group, while Photobacterium and Actinobacteriota were significantly enriched in the MG group. Similarly, comparisons between the control and TG groups showed significant enrichment of Pseudoalteromonadaceae in the control group and Actinobacteriota in the TG group.

3.4.3. Effect of Sand Thickness on the Function of Gut Microbiota

As shown in Figure 5, the predicted functions of the intestinal microbiota were predominantly associated with primary metabolic pathways across all experimental groups. Functions related to Genetic Information Processing, Environmental Information Processing, and Cellular Processes were present at lower relative abundances. Secondary pathway analysis further revealed that gene functions were mainly enriched in carbohydrate metabolism and amino acid metabolism within the Metabolic pathways category; replication and repair within Genetic Information Processing; membrane transport within Environmental Information Processing; and cellular community–prokaryotes within Cellular Processes. No significant differences in the relative abundance of predicted functional pathways were observed among the experimental groups, indicating a high degree of functional stability in the intestinal microbiota of M. japonicus and a minimal impact of sand substrate thickness on overall microbial functional potential.
Sediment plays a vital role in aquaculture by providing nutrients, regulating water quality, and serving as an ecological matrix that supports the survival and reproduction of diverse beneficial and benthic organisms. The behavior and growth of crustaceans are influenced by multiple sediment-related factors, including substrate presence, type, and thickness [15,16]. The present study demonstrates that the survival and growth of M. japonicus from the early juvenile to adult stages are strongly dependent on sand substrate thickness. In particular, shrimp reared under non-sand and low-sand conditions exhibited markedly reduced survival and growth performance. In natural aquatic systems, the sediment–water interface represents a zone of intense physical, chemical, and biological interactions, which directly influence the feeding and burrowing behaviors of benthic macroinvertebrates.

4. Discussion

4.1. Effects of Different Sand Laying Thicknesses on the Shrimp Growth Performances

The effects of sediment on crustacean growth were evaluated over a 120-day culture period, during which clear differences in growth performance were observed among experimental groups. Shrimp in the control group exhibited the lowest growth at day 60, while growth performance in the LG group declined noticeably by day 90, particularly with respect to weight gain rate. In contrast, shrimp in the MG and TG groups maintained relatively stable and higher growth rates throughout the culture period. M. japonicus undergoes frequent molting during metamorphosis, juvenile growth, and reproduction [15], and molting frequency is strongly influenced by environmental conditions, with increased molting generally associated with enhanced growth [17]. At day 90, molting activity was lowest in the control group, whereas shrimp in the MG and TG groups exhibited higher molting frequencies, which likely contributed to their superior growth performance. Poor growth in the control and LG groups may therefore be attributed to the absence of sand or insufficient sand thickness, indicating that a 5 cm sand layer is inadequate to support optimal growth. By day 120, shrimp in the MG and TG groups clearly outperformed those in the other groups. Previous studies have suggested that a 10 cm sand layer reduces unnecessary movement, conserves energy, and promotes growth in M. japonicus [18]. The lack of significant growth differences between the MG and TG groups further suggests that the growth benefits of increasing sand thickness reach a plateau at approximately 10 cm.
Survival data further supported the importance of appropriate sand substrate thickness. The control group exhibited the lowest survival rate (20.5%), whereas survival rates in the LG, MG, and TG groups were markedly higher, at 78.5%, 85.7%, and 87.3%, respectively, with no significant difference between the MG and TG groups. Daily observations indicated that substantial mortality occurred predominantly in the control group, particularly after day 90, when shrimp appeared severely worn, consistent with previous findings [19]. In sand-free conditions, limited burrowing behavior forced shrimp to interact more frequently with hard surfaces, resulting in increased mechanical damage to appendages. In addition, sand-free culture conditions likely elevated physiological stress and compromised immune function, predisposing shrimp to appendage infections and mortality. The low survival rate in the control group was also associated with intensified aggressive interactions. Under high-density culture conditions, competition for space and food was more pronounced in the absence of sand, leading to increased aggression. Consistent with previous reports, the addition of sediment has been shown to significantly improve growth performance in penaeid shrimp compared with bottomless culture systems [10]. Given the inherently slow growth of M. japonicus and its lower growth and survival rates relative to other shrimp species, even under pond culture conditions [20], optimizing sand thickness is particularly important for industrial production. In the present study, both the MG and TG groups exhibited favorable growth and survival outcomes; however, from a practical perspective, a 10 cm sand layer represents a more cost-effective and labor-efficient option for industrial M. japonicus culture.

4.2. Effects of Different Sand Thicknesses on the Antioxidant Defense System

ROS are inevitable byproducts of aerobic metabolism, and environmental stress can elevate ROS levels in crustaceans, leading to oxidative stress when antioxidant defenses are insufficient [20]. Antioxidant enzymes and nonenzymatic antioxidants play critical roles in scavenging free radicals and protecting cellular integrity [21]. In the present study, T-AOC activity declined over time in all groups; however, values in the sand substrate groups remained significantly higher than those in the control group during the mid to late culture stages, indicating improved overall antioxidant capacity. In contrast, SOD activity was consistently higher in the control group and peaked at day 90, suggesting increased oxidative challenge under sand-free conditions. CAT activity in the sand substrate groups exhibited a pattern similar to that of SOD, with the TG group showing the highest activity at the end of the culture period. Notably, GSH levels in the TG group were higher than those in the LG and MG groups at day 120, implying greater oxidative stress under excessive sand thickness. GSH-PX is a key antioxidant enzyme involved in protecting cells and DNA from oxidative damage [22]. Although GSH-PX activity declined from day 60 to day 120 in all groups, the MG group exhibited the lowest activity among the sand substrate treatments, suggesting that oxidative stress was relatively alleviated under a 10 cm sand substrate. Consistently, MDA levels, a widely used indicator of lipid peroxidation and oxidative stress in aquatic organisms [23], were markedly lower in the MG group than in the other groups during long-term culture, indicating reduced intestinal lipid peroxidation [24].
Sand thickness also influenced the expression of intestinal immune- and antioxidant-related genes in M. japonicus. Among the sand substrate groups, the TG group showed higher expression levels of SOD, CAT, and Hsp70 during the mid to late culture stages, suggesting increased oxidative stress under excessive sand thickness. PO, a key component of the crustacean immune system involved in host recognition, defense, and melanization [25], exhibited low expression in the control group, further indicating compromised immune function under sand-free conditions. Lzm, an important component of the nonspecific immune system [26], and caspase-3, a key regulator of oxidative-stress-induced apoptosis [27], displayed U-shaped expression patterns in the sand substrate groups, decreasing initially and increasing by day 120. Notably, caspase-3 expression was lowest in the MG group, indicating reduced intestinal cell apoptosis under a 10 cm sand substrate during long-term culture. These results suggest that an intermediate sand thickness provides a more favorable oxidative and immune environment for M. japonicus by balancing antioxidant defense and minimizing intestinal cellular damage.

4.3. Effects of Different Sand Thicknesses on Apoptosis

Environmental stress can disrupt intracellular ROS homeostasis, leading to alterations in cellular components and biosynthetic processes and ultimately triggering apoptosis [28]. TUNEL analysis showed that, among the sand substrate groups, the LG group exhibited the highest number of apoptotic cells, followed by the MG and TG groups. These findings indicate that increasing sand substrate thickness effectively reduced oxidative damage in the shrimp intestine and, consequently, decreased apoptotic cell numbers. Unexpectedly, the control group displayed significantly fewer apoptotic cells than the sand substrate groups. This observation may reflect a selection effect, whereby only individuals with stronger stress tolerance survived under sand-free conditions, resulting in an apparently lower level of apoptosis among the remaining shrimp.
Excessive ROS not only induces apoptosis but also compromises intestinal tissue integrity [29]. Environmental stress that damages tissues, cells, and organelles can disrupt normal metabolic processes [30]. Histological examination of intestinal tissues at day 120 revealed swelling and congestion of the intestinal wall in both the control and LG groups, with more severe congestion and wall thickening observed in the LG group. Such inflammatory responses were associated with pronounced thickening of the intestinal wall. In contrast, no obvious congestion or inflammation was detected in the MG or TG groups, and intestinal wall thickness in these groups was markedly reduced compared with that in the control and LG groups. The observed edema and wall thickening were likely attributable to inflammation-induced vascular dilation, cellular swelling, and widening of intercellular spaces. Examination of the intestinal basal lamina further showed that it remained smooth and intact in the control, MG, and TG groups but appeared swollen and disrupted in the LG group. Given that normal intestinal morphology is essential for efficient digestion and nutrient absorption [31], these structural alterations suggest that long-term culture under insufficient sand thickness can impair intestinal integrity and cellular function in M. japonicus.

4.4. Effects of Different Sand Thickness on the Intestinal Bacteria

The intestinal microbiota plays a crucial role in crustacean health and homeostasis by influencing community diversity and function, host immunity, metabolism, and growth [32,33]. Previous studies have shown that substrate type affects hindgut microbial diversity in Urechis unicinctus larvae and alters crustacean gut microbiota, thereby modulating host metabolism and disease resistance [34,35]. Consistent with these findings, sand substrate thickness in the present study influenced the taxonomic composition, core microbial groups, and interspecific interactions of the intestinal microbiota of M. japonicus [36,37]. The absence of sand reduced microbial diversity and altered community structure, which may compromise functional stability and increase disease risk [38]. Notably, a 10 cm sand substrate was associated with improved intestinal microbial diversity, as reflected by a higher Simpson index in the MG group, indicating a more favorable intestinal environment.
Across all experimental groups, Proteobacteria dominated the intestinal microbiota, suggesting that this phylum represents a core component of the M. japonicus gut microbiome [39]. Proteobacteria are widespread in aquatic ecosystems and contribute to nutrient cycling, organic matter mineralization, nitrogen and phosphorus removal, and organic degradation [40]. Given their prevalence in pond sediments, sediment-associated Proteobacteria may influence the intestinal microbiota through direct shrimp–sediment interactions [41,42], while increased abundance during later culture stages may also reflect the use of formulated feeds [43]. High Proteobacteria abundance has been associated with healthy shrimp intestines [44], and the elevated proportion observed in the MG group suggests that a 10 cm sand substrate may promote intestinal health. Bacteroidota, which contribute to intestinal mucosal immune homeostasis and host energy metabolism, were also widely distributed across groups, whereas Actinobacteriota and Chloroflexi were detected at lower but notable abundances. Actinobacteriota are involved in the degradation of polysaccharides and proteins, attenuation of inflammation, and enhancement of host resistance [45], and their abundance has been positively associated with normal shrimp growth [46]. The significantly lower abundance of Actinobacteriota in the control group corresponded with its poorer growth performance, supporting this relationship. The presence of anaerobic Chloroflexi, some of which possess photosynthetic capacity [47], further suggests that aquaculture water microbiota may contribute to shaping the intestinal microbial community under certain conditions.
Distinct dominant taxa were observed among experimental groups. Thiotrichaceae predominated in the control group, Rhodobacteraceae in the LG group, Photobacterium in the MG group, and Vibrio in the TG group. Notably, the relative abundances of Photobacterium in the MG group and Vibrio in the TG group each reached approximately 20%, exceeding those in the other groups. Photobacterium is commonly detected in mariculture systems, where it participates in organic matter degradation [48], produces antibacterial compounds [49], and contributes to host defense by inhibiting pathogenic Vibrio species in the shrimp intestine [50]. The enrichment of Photobacterium in the MG group may therefore be associated with enhanced resistance to pathogen invasion. In contrast, Vibrio is a ubiquitous genus in shrimp aquaculture, and elevated intestinal abundance is often considered an early warning indicator of disease outbreaks [51]. The higher abundance of Vibrio in the TG group suggests that excessive sand thickness may increase disease risk in cultured M. japonicus. Previous studies have demonstrated that shrimp intestinal microbiota shares greater similarity with pond sediment communities than with rearing water [52], supporting the notion that sediment-derived microorganisms contributed to the increased Vibrio abundance observed in the TG group. In practical aquaculture settings, excessive sand thickness has been associated with pond bottom odor and water quality deterioration. It is therefore plausible that a 20 cm sand substrate allows shrimp to burrow more deeply, facilitating the transfer of sediment-associated opportunistic pathogens into the intestine. Despite these compositional differences, functional prediction analyses indicated that intestinal microbiota across all groups were primarily associated with metabolic pathways, particularly carbohydrate and amino acid metabolism. Consistent with findings in Procambarus clarkii [53], no significant differences in predicted microbial functions were observed among groups, suggesting that the intestinal microbiota of M. japonicus maintains functional stability across different sand thicknesses.

5. Conclusions

In this study, four sand substrate thicknesses were evaluated to determine their effects on the growth performance and intestinal health of M. japonicus. The results demonstrated that shrimp reared in a sand-free environment exhibited significantly reduced growth performance, whereas survival and growth rates were highest at a sand thickness of 10 cm. Analyses of intestinal immune responses and antioxidant capacity indicated that decreasing sand thickness increased oxidative stress in shrimp, while the 10 cm sand substrate imposed the least environmental stress by the end of the culture period. Intestinal histological observations and TUNEL assays further showed that insufficient sand thickness increased susceptibility to intestinal tissue damage and apoptosis. In addition, gut microbiota composition was clearly influenced by sand substrate thickness. Although the presence of sand enhanced intestinal microbial abundance and diversity, excessive sand thickness was associated with an increased abundance of Vibrio. Taken together, these findings indicate that an intermediate sand thickness provides the most favorable balance between growth performance, intestinal health, and microbial stability. Therefore, considering both biological outcomes and practical production efficiency, a sand substrate thickness of 10 cm is recommended for industrial culture of M. japonicus.

Supplementary Materials

The following supporting information can be downloaded at: https://www.mdpi.com/article/10.3390/ani16040586/s1, Figure S1: Top 10 bacterial phyla in M. japonicus under sand thickness gradients. (A) Relative abundance, (B) control vs LG (Welch’s t-test), (C) control vs MG (Welch’s t-test), (D) NG vs TG (Welch’s t-test); Figure S2: Dominant bacterial genera (top 10) in M. japonicus across sand treatments. (A) Relative abundance, (B–D) Pairwise comparisons: control vs LG/MG/TG (Welch’s t-test); Figure S3: Core microbiota analysis in M. japonicus intestine. (A) Venn diagram, (B) Species distribution; Figure S4: Phylum-level taxa differential analysis (Kruskal-Wallis) in M. japonicus intestine; Figure S5: Abundance distribution of key phyla: Proteobacteria, Actinobacteriota, Chloroflexi, Patescibacteria (box plots); Figure S6: Linear discriminant analysis (LDA) effect size (LEfSe) bar plot.

Author Contributions

X.R.: Writing—original draft, Supervision. K.Z.: Data curation, Methodology. X.B.: Investigation, Validation. S.J.: Writing—review and editing. P.L.: Formal analysis, Visualization. J.L. (Jian Li): Formal analysis, Visualization. Y.C.: Writing—review and editing, Project administration. J.L. (Jitao Li): Writing—review and editing, Project administration. All authors have read and agreed to the published version of the manuscript.

Funding

This work was supported by the Key R&D Program of Shandong Province, China [grant number 2023LZGC020]; the Shandong Key R&D Program (Competitive Innovation Platform) [grant number 2024CXPT071-1]; the basic scientific research operating funds for national key laboratories [grant number BRESG-JB202517]; the China Agriculture Research System of MOF and MARA [grant number CARS-48]; and the Central Public-interest Scientific Institution Basal Research Fund, CAFS [grant number 2023TD50].

Institutional Review Board Statement

Under approval number YSFRI-2025033, this study was conducted with the approval of the committee at the Yellow Sea Fisheries Research Institute.

Informed Consent Statement

Not applicable.

Data Availability Statement

The data supporting the findings of this study are available from the corresponding author upon reasonable request.

Conflicts of Interest

We confirm that we have no financial or non-financial interests that could be perceived as potential conflicts of interest regarding the research presented in this study.

References

  1. Kawato, S.; Nishitsuji, K.; Arimoto, A.; Hisata, K.; Kawamitsu, M.; Nozaki, R.; Kondo, H.; Shinzato, C.; Ohira, T.; Satoh, N.; et al. Genome and transcriptome assemblies of the kuruma shrimp, Marsupenaeus japonicus. G3 2021, 11, jkab268. [Google Scholar] [CrossRef] [PubMed]
  2. Zhong, S.; Mao, Y.; Wang, J.; Liu, M.; Zhang, M.; Su, Y. Transcriptome analysis of Kuruma shrimp (Marsupenaeus japonicus) hepatopancreas in response to white spot syndrome virus (WSSV) under experimental infection. Fish Shellfish Immunol. 2017, 70, 710–719. [Google Scholar] [CrossRef] [PubMed]
  3. Ministry of Agriculture of the People’s Republic of China. China Agriculture Yearbook; China Agricultural Press: Beijing, China, 2025. [Google Scholar]
  4. Anand, P.S.; Aravind, R.; Balasubramanian, C.P.; Francis, B.; Rajan, R.V.; Vinay, T.N.; Kumar, S.; Sudheer, N.; Antony, J.; Rajamanickam, S.; et al. Reproductive and nursery performance of Kuruma shrimp Penaeus (Marsupenaeus) japonicus Form II: Effect of sandy bottom and light intensity in the rearing system. Aquac. Int. 2024, 32, 8009–8033. [Google Scholar] [CrossRef]
  5. Rajeev, R.; Adithya, K.K.; Kiran, G.S.; Selvin, J. Healthy microbiome: A key to successful and sustainable shrimp aquaculture. Rev. Aquac. 2021, 13, 238–258. [Google Scholar] [CrossRef]
  6. Capowiez, Y.; Bottinelli, N.; Sammartino, S.; Michel, E.; Jouquet, P. Morphological and functional characterisation of the burrow systems of six earthworm species (Lumbricidae). Biol. Fertil. Soils 2015, 51, 869–877. [Google Scholar] [CrossRef]
  7. Capowiez, Y.; Gilbert, F.; Vallat, A.; Poggiale, J.C.; Bonzom, J.M. Depth distribution of soil organic matter and burrowing activity of earthworms-mesocosm study using X-ray tomography and luminophores. Biol. Fertil. Soils 2021, 57, 337–346. [Google Scholar] [CrossRef]
  8. Zhao, J.; Liu, X.; Liao, X.; He, Z.; Chen, X.; Sun, C.; Ni, Z. Transcriptome analysis reveals the effects of sand substrate removal and body colour change of kuruma shrimp, Marsupenaeus japonicus. Aquac. Res. 2021, 52, 577–588. [Google Scholar] [CrossRef]
  9. Du, Y.; Song, A.; Chu, L.; Shan, H. The application of different bottom substrates in Penaeus japonicus culture: Impact on growth, environmental factors, the microbial community and the hepatopancreas transcriptome. Aquaculture 2025, 598, 742067. [Google Scholar] [CrossRef]
  10. Gao, Y.; Zhang, X.; Wei, J.; Sun, X.; Yuan, J.; Li, F.; Xiang, J. Whole transcriptome analysis provides insights into molecular mechanisms for molting in Litopenaeus vannamei. PLoS ONE 2015, 10, e0144350. [Google Scholar] [CrossRef]
  11. Zhao, K.; Jia, S.; Wang, J.; Bian, X.; Chen, S.; Li, J.; Liu, P.; Li, J.; Cai, Y.; Ren, X. Effects of sand substrate removal on the intestinal antioxidant and metabolism in Marsupenaeus japonicus. Fish Shellfish Immunol. 2024, 152, 109786. [Google Scholar] [CrossRef]
  12. Bradford, M.M. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal. Biochem. 1976, 72, 248–254. [Google Scholar] [CrossRef] [PubMed]
  13. Zhang, X.; Li, M.; Tao, X.; Yang, Y.; Sun, P.; Jin, M.; Zhou, Q.; Jiao, L. Effects of dietary montmorillonite supplementation on the growth performance, antioxidant capacity, intestinal barrier and microbiota composition in Marsupenaeus japonicus. Aquaculture 2022, 557, 738330. [Google Scholar] [CrossRef]
  14. Livak, K.J.; Schmittgen, T.D. Analysis of relative gene expression data using real-time quantitative PCR and the 2−ΔΔCT method. Methods 2001, 25, 402–408. [Google Scholar] [CrossRef] [PubMed]
  15. Jee, J.H.; Koo, J.G.; Keum, Y.H.; Kang, J.S.; Park, K.H.; Kang, J.C. Effects of diet and sediment type on survival, growth and moulting of juvenile tiger crab, Orithyia sinica L. Aquac. Res. 2007, 38, 26–35. [Google Scholar] [CrossRef]
  16. Zhang, X.; Yuan, J.; Sun, Y.; Li, S.; Gao, Y.I.; Yu, Y.; Liu, C.; Wang, Q.; Lv, X.; Zhang, X.; et al. Penaeid shrimp genome provides insights into benthic adaptation and frequent molting. Nat. Commun. 2019, 10, 356. [Google Scholar] [CrossRef]
  17. Ding, L.; Yu, J.; Peng, X.; Yang, G.; Du, T.; Tang, Q.; Yi, S. Changes in molting frequency and expression patterns of molting-related genes in Macrobrachium rosenbergii with exogenous calcium supplement in water. Aquaculture 2024, 586, 740761. [Google Scholar] [CrossRef]
  18. Anand, P.S.; Balasubramanian, C.P.; Christina, L.; Kumar, S.; Biswas, G.; De, D.; Ghoshal, T.; Vijayan, K.K. Substrate based black tiger shrimp, Penaeus monodon culture: Stocking density, aeration and their effect on growth performance, water quality and periphyton development. Aquaculture 2019, 507, 411–418. [Google Scholar] [CrossRef]
  19. Türkmen, G. Pond Culture of Penaeus semisulcatus and Marsupenaeus japonicus (Decapoda, Penaeidae) on the West coast of Turkey. Turk. J. Fish. Aquat. Sci. 2007, 7, 7–11. [Google Scholar]
  20. Zheng, C.; Zhao, Q.; Li, E.; Zhao, D.; Sun, S. Role of hypoxia in the behaviour, physiology, immunity and response mechanisms of crustaceans: A review. Rev. Aquac. 2022, 14, 676–687. [Google Scholar] [CrossRef]
  21. Hu, X.; Ma, W.; Zhang, D.; Tian, Z.; Yang, Y.; Huang, Y.; Hong, Y. Application of natural antioxidants as feed additives in aquaculture: A review. Biology 2025, 14, 87. [Google Scholar] [CrossRef]
  22. Fan, J.; Li, B.; Hong, Q.; Yan, Z.; Yang, X.; Lu, K.; Chen, G.; Wang, L.; Chen, Y. A glutathione peroxidase gene from Litopenaeus vannamei is involved in oxidative stress responses and pathogen infection resistance. Int. J. Mol. Sci. 2022, 23, 567. [Google Scholar] [CrossRef] [PubMed]
  23. Siddique, Q.; Xie, X.; Yin, F. An integrated metabolomics and proteomics analysis reveals copper-zinc alloy surface contact-mediated metabolic disruption, lipid peroxidation, and osmotic imbalance of Cryptocaryon irritans tomonts. Aquaculture 2025, 595, 741713. [Google Scholar] [CrossRef]
  24. Demirci-Çekiç, S.; Özkan, G.; Avan, A.N.; Uzunboy, S.; Çapanoğlu, E.; Apak, R. Biomarkers of oxidative stress and antioxidant defense. J. Pharm. Biomed. 2022, 209, 114477. [Google Scholar] [CrossRef] [PubMed]
  25. Zdybicka-Barabas, A.; Stączek, S.; Kunat-Budzyńska, M.; Cytryńska, M. Innate Immunity in Insects: The Lights and Shadows of Phenoloxidase System Activation. Int. J. Mol. Sci. 2025, 26, 1320. [Google Scholar] [CrossRef]
  26. Xu, Y.; Xu, W.; Hu, X.; Su, H.; Wen, G.; Yang, K.; Cao, Y. Toxicity of the microcystin-producing cyanobacteria Microcystis aeruginosa to shrimp Litopenaeus vannamei. Ecotoxicology 2022, 31, 1403–1412. [Google Scholar] [CrossRef]
  27. Chen, Z.; Tayyab, M.; Yao, D.; Aweya, J.J.; Zheng, Z.; Zhao, X.; Lin, Z.; Zhang, Y. Cell death in crustacean immune defense. Rev. Aquacult. 2025, 17, e12976. [Google Scholar] [CrossRef]
  28. Ali, M.J.; Tao, Y.; Li, Y.; Sayouh, M.A.; Lu, S.; Qiang, J.; Xu, P. Modulation of chronic hypoxia on ovarian structure, oxidative stress, and apoptosis in female Nile Tilapia (Oreochromis niloticus). Aquaculture 2024, 590, 741081. [Google Scholar] [CrossRef]
  29. Zhao, M.; Yao, D.; Li, S.; Zhang, Y.; Aweya, J.J. Effects of ammonia on shrimp physiology and immunity: A review. Rev. Aquacult. 2025, 12, 2194–2211. [Google Scholar] [CrossRef]
  30. Wang, P.; Liu, H.; Zhao, S.; Yu, S.; Xie, S.; Hua, S.; Yan, B.; Xing, C.; Gao, H. Hypoxia stress affects the physiological responses, apoptosis and innate immunity of Kuruma shrimp, Marsupenaeus japonicus. Fish Shellfish Immunol. 2022, 122, 206–214. [Google Scholar] [CrossRef]
  31. Guo, J.; Chen, Y.; Zhang, Y.; Zhang, R.; Inaba, K.; Osato, T.; Zhao, X.; Han, Y.; Ren, T. Oxygen nanobubble-induced hyperoxia: Effects on growth, digestive enzyme activity, intestinal morphology, and biochemical parameters in kuruma prawn (Penaeus japonicus). Aquac. Rep. 2025, 43, 102882. [Google Scholar] [CrossRef]
  32. Chen, H.; Pan, J.; Wang, Y.; Qiao, Y.; Han, F.; Xu, C.; Farhadi, A.; Li, E. Growth, health status and gut microbiota of the scalloped spiny lobster (Panulirus homarus) at different salinities. Aquaculture 2023, 562, 738779. [Google Scholar] [CrossRef]
  33. Medina-Félix, D.; Garibay-Valdez, E.; Vargas-Albores, F.; Martínez-Porchas, M. Fish disease and intestinal microbiota: A close and indivisible relationship. Rev. Aquac. 2022, 15, 820–839. [Google Scholar] [CrossRef]
  34. Lynn, D.J.; Benson, S.C.; Lynn, M.A.; Pulendran, B. Modulation of immune responses to vaccination by the microbiota: Implications and potential mechanisms. Nat. Rev. Immunol. 2022, 22, 33–46. [Google Scholar] [CrossRef] [PubMed]
  35. Chen, W.; Zhang, S.; Sun, Y.; Tian, B.; Song, L.; Xu, Y.; Liu, T. Effects of substrate on the physiological characteristics and intestinal microbiota of Echiura worm (Urechis unicinctus) juveniles. Aquaculture 2021, 530, 735710. [Google Scholar] [CrossRef]
  36. Sun, S.; Yang, M.; Fu, H.; Ge, X.; Zou, J. Altered intestinal microbiota induced by chronic hypoxia drives the effects on lipid metabolism and the immune response of oriental river prawn Macrobrachium nipponense. Aquaculture 2020, 526, 735431. [Google Scholar] [CrossRef]
  37. Sha, H.; Lu, J.; Chen, J.; Xiong, J. A meta-analysis study of the robustness and universality of gut microbiota–shrimp diseases relationship. Environ. Microbiol. 2022, 24, 3924–3938. [Google Scholar] [CrossRef]
  38. Gao, C.H.; Zhang, S.; Wei, M.Y.; Ding, Q.S.; Ma, D.N.; Li, J.; Wen, C.; Li, H.; Zhao, Z.-Z.; Wang, C.-H.; et al. Effects of shrimp pond effluent on functional traits and functional diversity of mangroves in Zhangjiang Estuary. Environ. Pollut. 2022, 297, 118762. [Google Scholar] [CrossRef]
  39. Zhou, L.; Qu, Y.; Qin, J.G.; Chen, L.; Han, F.; Li, E. Deep insight into bacterial community characterization and relationship in the pond water, sediment and the gut of shrimp (Penaeus japonicus). Aquaculture 2021, 539, 736658. [Google Scholar] [CrossRef]
  40. Song, M.; Pan, L.; Zhang, M.; Huang, F.; Gao, S.; Tian, C. Changes of water, sediment, and intestinal bacterial communities in Penaeus japonicus cultivation and their impacts on shrimp physiological health. Aquac. Int. 2020, 28, 1847–1865. [Google Scholar] [CrossRef]
  41. Cornejo-Granados, F.; Lopez-Zavala, A.A.; Gallardo-Becerra, L.; Mendoza-Vargas, A.; Sánchez, F.; Vichido, R.; Brieba, L.G.; Viana, M.T.; Sotelo-Mundo, R.R.; Ochoa-Leyva, A. Microbiome of Pacific Whiteleg shrimp reveals differential bacterial community composition between Wild, Aquacultured and AHPND/EMS outbreak conditions. Sci. Rep. 2017, 7, 11783. [Google Scholar] [CrossRef]
  42. Sun, Y.; Han, W.; Liu, J.; Huang, X.; Zhou, W.; Zhang, J.; Cheng, Y. Bacterial community compositions of crab intestine, surrounding water, and sediment in two different feeding modes of Eriocheir sinensis. Sci. Rep. 2020, 16, 100236. [Google Scholar] [CrossRef]
  43. Duan, Y.; Wang, Y.; Liu, Q.; Dong, H.; Li, H.; Xiong, D.; Zhang, J. Changes in the intestine microbial, digestion and immunity of Litopenaeus vannamei in response to dietary resistant starch. Sci. Rep. 2019, 9, 6464. [Google Scholar] [CrossRef] [PubMed]
  44. Tao, J.H.; Duan, J.A.; Jiang, S.; Feng, N.N.; Qiu, W.Q.; Ling, Y. Polysaccharides from Chrysanthemum morifolium Ramat ameliorate colitis rats by modulating the intestinal microbiota community. Oncotarget 2017, 8, 80790–80803. [Google Scholar] [CrossRef] [PubMed]
  45. Kesarcodi-Watson, A.; Kaspar, H.; Lategan, M.J.; Gibson, L. Probiotics in aquaculture: The need, principles and mechanisms of action and screening processes. Aquaculture 2008, 274, 1–14. [Google Scholar] [CrossRef]
  46. Fan, L.; Li, Q.X. Characteristics of intestinal microbiota in the Pacific white shrimp Litopenaeus vannamei differing growth performances in the marine cultured environment. Aquaculture 2019, 505, 450–461. [Google Scholar] [CrossRef]
  47. Fan, L.; Wang, Z.; Chen, M.; Qu, Y.; Li, J.; Zhou, A.; Xie, S.; Zeng, F.; Zou, J. Microbiota comparison of Pacific white shrimp intestine and sediment at freshwater and marine cultured environment. Sci. Total Environ. 2019, 657, 1194–1204. [Google Scholar] [CrossRef]
  48. Gouife, M.; Chen, S.; Huang, K.; Nawaz, M.; Jin, S.; Ma, R.; Wang, Y.; Xue, L.; Xie, J. Photobacterium damselae subsp. damselae in mariculture. Aquac. Int. 2022, 30, 1453–1480. [Google Scholar] [CrossRef]
  49. Zhu, T.; Xu, L.; Yang, L.; Zhao, P.; Zhu, X.; Wu, Z.; Song, Z. Microbiologically influenced corrosion inhibition of carbon steel by a novel bacterium, Photobacterium sp., in simulated marine environment. Environ. Res. 2025, 264, 120298. [Google Scholar]
  50. Dai, W.; Yu, W.; Xuan, L.; Tao, Z.; Xiong, J. Integrating molecular and ecological approaches to identify potential polymicrobial pathogens over a shrimp disease progression. Appl. Microbiol. Biotechnol. 2018, 102, 3755–3764. [Google Scholar] [CrossRef]
  51. Xiong, J.; Wang, K.; Wu, J.; Qiuqian, L.; Yang, K.; Qian, Y.; Zhang, D. Changes in intestinal bacterial communities are closely associated with shrimp disease severity. Appl. Microbiol. Biotechnol. 2015, 99, 6911–6919. [Google Scholar] [CrossRef]
  52. Li, H.; Gu, S.; Wang, L.; Shi, W.; Jiang, Q.; Wan, X. Dynamic changes of environment and gut microbial community of Litopenaeus vannamei in greenhouse farming and potential mechanism of gut microbial community construction. Fishes 2024, 9, 155. [Google Scholar] [CrossRef]
  53. Wan, R.; Zhang, C.; Tang, Y.; Zhu, J.; Yang, N.; Su, S. Effects of Different Sources of Culture Substrate on the Growth and Immune Performance of the Red Swamp Crayfish (Procambarus clarkii). Int. J. Mol. Sci. 2023, 24, 14098. [Google Scholar] [CrossRef]
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Figure 1. Immunity and antioxidant gene expression in M. japonicus intestine under sand thickness variations. (A) SOD gene, (B) CAT gene, (C) Hsp70 gene, (D) Po gene, (E) Lzm gene, (F) Caspase-3 gene. a–d The values in the same row sharing different superscript letters are significantly different.
Figure 1. Immunity and antioxidant gene expression in M. japonicus intestine under sand thickness variations. (A) SOD gene, (B) CAT gene, (C) Hsp70 gene, (D) Po gene, (E) Lzm gene, (F) Caspase-3 gene. a–d The values in the same row sharing different superscript letters are significantly different.
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Figure 2. Intestinal histopathology. (A) control group, (B) LG group, (C) MG group, (D) TG group. Abbreviations: ML, muscular layer; a, intestinal wall thickness.
Figure 2. Intestinal histopathology. (A) control group, (B) LG group, (C) MG group, (D) TG group. Abbreviations: ML, muscular layer; a, intestinal wall thickness.
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Figure 3. Gut epithelial apoptosis in M. japonicus.
Figure 3. Gut epithelial apoptosis in M. japonicus.
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Figure 4. Beta diversity analysis of intestinal microbiota in M. japonicus. (A) Nonmetric multidimensional scaling (NMDS); (B) Principal coordinates analysis (PCoA).
Figure 4. Beta diversity analysis of intestinal microbiota in M. japonicus. (A) Nonmetric multidimensional scaling (NMDS); (B) Principal coordinates analysis (PCoA).
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Figure 5. KEGG functional prediction of intestinal microbiota in M. japonicus under sand thickness treatments: (A) control group, (B) LG group, (C) MG group, (D) TG group.
Figure 5. KEGG functional prediction of intestinal microbiota in M. japonicus under sand thickness treatments: (A) control group, (B) LG group, (C) MG group, (D) TG group.
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Table 1. Primers used for real-time quantitative PCR.
Table 1. Primers used for real-time quantitative PCR.
Gene NameForward Primer (5′-3′)Reverse Primer (5′-3′)Amplification Efficiencies (%)
SODTCAATCCTCTCCCACACACACAGACAGGCAGAGCAGT96
CATGCCTCAGGAGAACGTTGTTGTGAGCTTGTCTGAGAGTGGG92
Hsp70ACAAGTCCATCAACCCCGATGGTGAAGGTCTGAGTCTGCT90
LzmCAGTAGTGGCTGTGTCATGCAGGTATGCACGGACAGTCTC110
Caspase-3CTCTCACGACGCCTACATTTCCCTGTTGTTCCTGTTT97
β-actinTCCACGAGACCACATACAACCACTTCCTGAACGATTGA100
Table 2. Growth performance of cultured M. japonicus under varying sand substrate thicknesses.
Table 2. Growth performance of cultured M. japonicus under varying sand substrate thicknesses.
Growth Indicators (%)ControlLGMGTG
30 d Length growth rate61.11 ± 0.82 a86.90 ± 1.74 b95.69 ± 2.42 c91.29 ± 2.27 bc
60 d Length growth rate128.94 ± 2.95 a155.08 ± 4.31 b155.16 ± 3.10 b160.18 ± 0.76 b
90 d Length growth rate184.98 ± 1.15 a197.02 ± 1.25 b204.98 ± 1.18 c203.17 ± 0.88 c
120 d Length growth rate216.05 ± 2.54 a227.17 ± 0.96 b237.31 ± 0.56 c237.48 ± 3.34 c
30 d Weight gain rate381.36 ± 14.70 a633.51 ± 47.90 b802.69 ± 14.95 c718.64 ± 19.77 d
60 d Weight gain rate1308.96 ± 13.69 a1836.92 ± 108.47 b1910.57 ± 67.67 bc2030.82 ± 33.45 c
90 d Weight gain rate2205.56 ± 52.46 a3448.03 ± 191.60 b3832.80 ± 85.41 c3607.89 ± 124.69 bc
120 d Weight gain rate4534.59 ± 60.57 a4772.76 ± 59.81 b5148.21 ± 18.50 c5318.46 ± 114.30 c
30 d Specific growth rate5.31 ± 0.01 a6.64 ± 0.22 b7.33 ± 0.06 c7.05 ± 0.008 c
60 d Specific growth rate4.41 ± 0.02 a4.94 ± 0.09 b5.00 ± 0.06 b5.10 ± 0.03 b
90 d Specific growth rate3.49 ± 0.03 a3.96 ± 0.06 b4.08 ± 0.02 c4.01 ± 0.04 bc
120 d Specific growth rate3.20 ± 0.01 a3.24 ± 0.01 b3.30 ± 0.003 c3.33 ± 0.02 c
Survival rate20.57 ± 2.63 a78.54 ± 1.26 b85.71 ± 0.91 c87.32 ± 0.56 c
a–d The values in the same row sharing different superscript letters are significantly different.
Table 3. Intestinal immunity and antioxidant parameters in M. japonicus under sand thickness gradients.
Table 3. Intestinal immunity and antioxidant parameters in M. japonicus under sand thickness gradients.
Parameters (%)ControlLGMGTG
60 d T-AOC activity1.71 ± 0.02 a1.71 ± 0.03 a1.56 ± 0.04 b1.28 ± 0.03 c
90 d T-AOC activity0.42 ± 0.03 a0.67 ± 0.02 b0.53 ± 0.03 c0.68 ± 0.02 b
120 d T-AOC activity0.41 ± 0.01 a0.38 ± 0.02 b0.45 ± 0.008 c0.46 ± 0.008 c
60 d SOD activity29.52 ± 0.48 a25.04 ± 0.35 b23.31 ± 0.68 c27.05 ± 3.38 d
90 d SOD activity43.44 ± 0.44 a32.37 ± 0.40 b34.42 ± 0.49 c33.48 ± 0.60 d
120 d SOD activity30.24 ± 0.48 a29.55 ± 0.98 ab28.11 ± 0.39 c28.61 ± 0.25 bc
60 d CAT activity0.22 ± 0.007 a0.09 ± 0.007 b0.16 ± 0.006 c0.17 ± 0.004 c
90 d CAT activity0.25 ± 0.007 a0.21 ± 0.005 b0.29 ± 0.004 c0.32 ± 0.005 d
120 d CAT activity0.27 ± 0.007 a0.15 ± 0.002 b0.26 ± 0.009 a0.31 ± 0.004 c
60 d GSH content29.47 ± 0.73 a31.42 ± 0.57 b25.76 ± 1.15 c27.88 ± 0.83 a
90 d GSH content16.62 ± 0.72 a21.46 ± 0.97 b31.46 ± 0.67 c22.15 ± 0.97 b
120 d GSH content24.37 ± 0.94 a22.01 ± 1.09 b41.22 ± 0.75 c52.36 ± 0.90 d
60 d GSH-PX activity34.92 ± 0.58 a37.17 ± 0.71 b38.11 ± 1.01 b35.62 ± 0.41 a
90 d GSH-PX activity18.12 ± 0.95 a34.99 ± 0.87 b33.92 ± 0.71 b45.93 ± 0.34 c
120 d GSH-PX activity13.21 ± 0.73 a19.42 ± 0.62 b8.51 ± 0.66 c14.61 ± 1.03 a
60 d MDA content1.75 ± 0.03 a1.76 ± 0.03 a1.70 ± 0.04 b1.39 ± 0.02 c
90 d MDA content0.39 ± 0.05 a0.50 ± 0.03 b0.79 ± 0.03 c0.57 ± 0.04 d
120 d MDA content0.46 ± 0.02 a0.41 ± 0.02 b0.32 ± 0.02 c0.37 ± 0.03 b
a–d The values in the same row sharing different superscript letters are significantly different.
Table 4. Intestinal microbiota alpha diversity in M. japonicus across sand thicknesses.
Table 4. Intestinal microbiota alpha diversity in M. japonicus across sand thicknesses.
GroupsShannon IndexSimpson IndexACE IndexChao Index
Control4.05 ± 0.16 a0.07 ± 0.002 a417.18 ± 13.30 a419.68 ± 14.99 a
LG4.38 ± 0.18 b0.05 ± 0.002 b546.56 ± 14.99 b546.81 ± 12.66 b
MG3.99 ± 0.15 a0.09 ± 0.003 c441.81 ± 18.32 c442.39 ± 17.46 c
TG4.37 ± 0.12 b0.04 ± 0.003 d512.59 ± 18.14 d513.96 ± 17.15 d
a–d The values in the same row sharing different superscript letters are significantly different.
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Ren, X.; Zhao, K.; Bian, X.; Jia, S.; Liu, P.; Li, J.; Cai, Y.; Li, J. Sand Substrate Thickness Regulates Growth Performance, Intestinal Antioxidant Defense, and Gut Microbiota in an Experimental Culture of Marsupenaeus japonicus. Animals 2026, 16, 586. https://doi.org/10.3390/ani16040586

AMA Style

Ren X, Zhao K, Bian X, Jia S, Liu P, Li J, Cai Y, Li J. Sand Substrate Thickness Regulates Growth Performance, Intestinal Antioxidant Defense, and Gut Microbiota in an Experimental Culture of Marsupenaeus japonicus. Animals. 2026; 16(4):586. https://doi.org/10.3390/ani16040586

Chicago/Turabian Style

Ren, Xianyun, Kuangcheng Zhao, Xueqiong Bian, Shaoting Jia, Ping Liu, Jian Li, Yuefeng Cai, and Jitao Li. 2026. "Sand Substrate Thickness Regulates Growth Performance, Intestinal Antioxidant Defense, and Gut Microbiota in an Experimental Culture of Marsupenaeus japonicus" Animals 16, no. 4: 586. https://doi.org/10.3390/ani16040586

APA Style

Ren, X., Zhao, K., Bian, X., Jia, S., Liu, P., Li, J., Cai, Y., & Li, J. (2026). Sand Substrate Thickness Regulates Growth Performance, Intestinal Antioxidant Defense, and Gut Microbiota in an Experimental Culture of Marsupenaeus japonicus. Animals, 16(4), 586. https://doi.org/10.3390/ani16040586

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