The juvenile P. vannamei were randomly distributed into two groups (in triplicate) in culture tanks (56 cm × 41 cm × 32.5 cm, water depth: 30 cm). Each tank was equipped with an oxygenation device and fitted with a nylon cloth filter of 500 mesh size. The stocking density was set at 30 shrimp per tank. The animals were fed 3–5% of their live body weight for three times (morning, noon, and evening) a day with commercialized feed, primarily composed of fish meal, yeast, peanuts, soybean meal, and others. The commercial feed containing crude protein 43%, crude fat 6%, ash 16%, and water 12%. The experiment lasted for 30 days.

The aquaculture water was prepared by diluting seawater with tap water at a ratio of 11:1.4, maintaining a salinity of ~ 13‰. No water exchange occurred throughout the cultivation process.

Each treatment group were as follows: the CK group did not add any microbial preparations and was managed routinely, the PB group indicated that B. licheniformis was added to the water every 5 days during the aquaculture process at a dosage of 5 × 10⁴ CFU/mL.

At the end of the trial, the shrimp and the environment were sampled. Six shrimp were randomly taken from each tank. The shrimp were placed in a −20°C refrigerator to freeze for 20 min to death and then transferred to a 4°C refrigerator for cold storage. Then they were soaked in 75% alcohol for 30 s and rinsed with sterile water. In a clean bench, the shrimp intestinal tracts were extracted using sterile tweezers and packed into sterile centrifuge tubes, sealed, and stored at −80°C.

A sterile sampler was used to take a sample of 500 mL at a water depth of 10 cm in each culture tank, which was first filtered with a 500-mesh nylon cloth for primary filtration, and after the primary filtration, it was filtered with a 0.22 μm filter membrane for secondary filtration, and after filtration, the membrane was put into a 50 mL sterile centrifuge tube and stored at −80 °C.

Subsequently, the shrimp intestinal tract samples and the water samples were subjected to high-throughput sequencing.

After extracting the DNA of the samples by CTAB, the quality and purity of DNA were assessed through 1% agarose gel electrophoresis. The concentration of DNA was diluted to 1 ng·μL⁻¹ to serve as a template for PCR amplification. The PCR reaction was performed using Phusion^® High-Fidelity PCR Master Mix with GC Buffer, along with high-performance, high-fidelity enzymes, and the prokaryotic universal primers 341F and 806R. The primers and sequences utilized for PCR amplification were as described below:

341F (5'-CCTAYGGGRBGCASCAG-3')

806R (5'-GGACTACNNGGGTATCTAAT-3')

The amplified products were analyzed using 1% agarose gel electrophoresis to determine the sizes of the amplified bands. Subsequently, the PCR products were purified using the Agencourt AMPure XP nucleic acid purification kit.

The library was constructed using the Ion Plus Fragment Library Kit 48rxns from Thermo Fisher. After library construction, quantification was performed using Qubit, and libraries that passed quality control were sequenced on the Thermo Fisher Life Ion S5™ system.

The original sequences were analyzed using the QIIME software, with end-splicing accomplished by FLASH. Preliminary quality control was conducted through UPARSE, removing low-quality sequences, followed by clustering of all effective tags into OTUs and species annotation of representative OTU sequences. For the top 20 genera in abundance, heatmaps were generated using R language. The QIIME package was used to calculate alpha diversity for each sample, and principal component analysis (PCA) and other graphical representations were generated using the VennDiagram package.

The experimental data were presented as the mean ± standard error. The independent-samples t-test and one-way analysis of variance (ANOVA) were used to identify significant differences using SPSS software. A level of P < 0.05 was accepted as statistically significant. The data processing was conducted using Excel 2016. Graphs were generated using Origin 2018 software.

The growth parameters of shrimp were measured (Table 1). The data clearly indicated that B. licheniformis significantly enhanced growth indicators (P < 0.05), including shrimp length, weight, survival rate, and overall yield compared to the CK group. FCR represents the feed required to cultivate 1 kg of shrimp, and serves as a crucial efficiency indicator. A lower FCR indicates better feed conversion. As shown in the table, the PB group's FCR was significantly lower than the CK group's. In summary, B. licheniformis promoted shrimp growth and improved feed conversion.

In the present study, water pH gradually decreased in both the CK and PB groups. Initially, there was no significant difference between the two groups. However, in the later stages, the pH in the PB group was significantly lower than that in the CK group (P < 0.05). A reduction in pH over time can be attributed to microbial processes such as organic matter degradation, nitrification, and the resultant increase in carbonic acid (Ebeling et al., 2006). Nimrat et al. (2012) found that the addition of Bacillus probiotics to Penaeus vannamei resulted in a significant reduction in pH. Hassan et al. (2022) observed a significant decrease in the pH of the ponds after probiotics treatment (P < 0.05) compared to pre-treatment levels. These reports were consistent with the results of this study.

In the later stages of aquaculture, the concentrations of ammonia and nitrite in the water of both the CK and PB groups increased considerably. This was attributed to the accumulation of shrimp excreta and uneaten feed in the culture tanks. The proteins and amino acids in these substances contributed to the elevated levels of organic nitrogen in the water, thereby stimulating the production of microbial metabolic byproducts, including NO2−-N, NO3−-N, and NH4+-N (Jackson et al., 2003; Zhou and Boyd, 2015). However, the concentrations of ammonia and nitrite in the PB group were significantly lower than those in the CK group (P < 0.05). Microorganisms play diverse ecological roles as producers, consumers, and decomposers, and participate in various nitrogen cycling processes, such as nitrogen fixation (Yousuf et al., 2017), nitrification (Rout et al., 2017), ammonification (Hui et al., 2019), and denitrification (Verbaendert et al., 2011), etc. The microbial preparations used in this experiment were Bacillus strains. Monier et al. (2023) found that the application of Bacillus species probiotics (B. subtilis and B. licheniformis) to shrimp aquaculture lowered total ammonia nitrogen (TAN) and NH₃ in the pond water; Nimrat et al. (2020) found that the addition of Bacillus probiotics in the treatment group was associated with remarkable reductions in ammonia-nitrogen and nitrite-nitrogen concentrations in the water than the control group; Wang et al. (2022) similarly found that adding Bacillus to the P. vannamei culture process significantly reduced ammonia and nitrite contents in the water; These findings illustrated that microecological preparations dominated by Bacillus bacteria played a significant role in reducing ammonia and nitrite in water, consistent with this study's results.

This study indicated that compared to the CK group, B. licheniformis significantly improved shrimp growth indicators, including length, weight, survival rate, and yield (P < 0.05). Furthermore, the feed conversion ratio was significantly reduced (P < 0.05). B. licheniformis can promote intestinal health in shrimp, enhance the immune system, directly secrete digestive enzymes, and increase the activity of shrimp digestive enzymes, thereby improving feed utilization and promoting shrimp growth and survival. This has been confirmed in many studies and practices. For instance, Mirbakhsh et al. (2021) found that Bacillus subtilis administration improved growth and enhanced the immune response of Penaeus vannamei during the early hatchery period, thus increasing resistance to bacterial pathogens. Ranjit Kumar et al. (2013) found that dietary supplementation with Bacillus licheniformis inhibited pathogenic bacteria in the gut of Macrobrachium rosenbergii, enhanced growth and feed efficiency, and improved immune response, thereby increasing protection against bacterial infections. Ziaei-Nejad et al. (2006) also concluded that the use of Bacillus at different growth stages of shrimp promoted digestive enzyme activity, and enhanced survival rates and yield. These findings support the idea that B. licheniformis promotes shrimp growth and improves survival rates.

The results indicated that B. licheniformis increased the number of OTUs, and the Chao1, Shannon, and Simpson indices of the microbial communities in both the aquaculture water and shrimp intestinal tracts, compared to the CK group. This suggested that B. licheniformis enhanced bacterial diversity within shrimp farming systems. As a probiotic, B. licheniformis, has the ability to improve the aquaculture water quality, promote nutrient cycling and other roles, thus increasing bacterial diversity. The diversity of microbial communities plays a crucial role in maintaining ecological functions, low diversity may indicate poor functional stability of microbial communities, increasing the risk of disease in organisms. The diversity indices of aquaculture water are positively correlated with shrimp health, and variations in intestinal bacterial structure and diversity distinguish between healthy and diseased shrimp. Numerous studies confirmed this: Wu et al. (2016) analyzed the intestinal bacterial community of healthy and diseased shrimp by high-throughput sequencing, the results showed that the bacterial diversity was significantly lower in diseased shrimps than in healthy ones. Yao et al. (2018) and Dai et al. (2018) found that the bacterial Shannon index in the intestinal tract of diseased shrimp was significantly lower than that of healthy shrimp. DU Shi-cong et al. (2019) found that planktonic bacteria in Penaeus vannamei culture ponds before the disease emergence exhibited a significantly lower Shannon index than healthy ponds and suggested that a decrease in Shannon index could be a sign of divergence in shrimp health. Reyes et al. (2022) compared the microbiome of Penaeus vannamei with high and low survival in shrimp hatchery tanks, the Shannon diversity index was significantly lower at the low-survival tanks.

B. licheniformis altered the microbial diversity, bacterial abundance, and community structure in the aquaculture water and the shrimp intestinal tracts, however, the impacts differed between the two environments. In the present study, B. licheniformis resulted in changes in the OTUs of bacteria in the aquaculture water and the shrimp intestinal tracts. Notably, in the shrimp intestinal tract samples, the S_PB group shared more OTUs with the S_CK group, indicating a lesser impact of B. licheniformis on the shrimp intestinal tract. Compared to the aquaculture water, the microbial community in the shrimp intestines was more stable. Luo et al. (2006) demonstrated that external environmental factors could influence the microbial community structure in the shrimp intestinal tract. However, as shrimp cultivation progressed to a certain stage, the intestinal bacterial community became relatively stable and mature, becoming less susceptible to environmental influences. This finding aligned with the results of this study.

B. licheniformis increased the relative abundance of beneficial bacterial genera. It can enhance synergistic interactions among microorganisms, such as stimulating the growth of other probiotics through the production of bioactive substances, or by establishing symbiotic relationships to participate together in certain biochemical processes. In the culture water samples, it was found that after the addition of B. licheniformis, the relative abundance of the genera Bacillus, Gemmobacter, and Paracoccus was higher than in the CK group. Bacillus can inhibit the growth of pathogenic bacteria, as a biological barrier to compete with pathogenic bacteria for nutrients (Tran et al., 2023); The genera Gemmobacter (Liu et al., 2021) and Paracoccus (Si et al., 2020) have strong denitrification capabilities and effectively decompose organic matter in water, belonging to the beneficial bacteria in shrimp aquaculture process.

B. licheniformis reduced the relative abundance of potential pathogens. As a probiotic, B. licheniformis competes with potential pathogens for nutrients and living space, and it can produce a broad spectrum of antimicrobial substances. The present study found that B. licheniformis, decreased the relative abundance of Flavobacterium, a potential pathogen, in shrimp intestinal tracts (Dai et al., 2018). It was observed that Vibrio counts in the culture water significantly decreased following the addition of B. licheniformis (P < 0.05), during the cultivation period between day 10 and day 30. Vibrio, a prevalent pathogen in aquatic animal farming environments, is a primary pathogen during the shrimp larval stage and can lead to shrimp mortality (Zhou et al., 2012). The increase of pathogenic bacteria can lead to the occurrence of disease, which has been confirmed in many studies: Rahardjo et al. (2023) investigated microbial composition in rearing media of Penaeus vannamei infected by the white feces disease (WFD) and healthy Penaeus vannamei. They found that Vibrio vulnificus dominated in WFD-infected shrimp pond water. Oxley et al. (2002), Abraham et al. (2004) and Phung et al. (2020) found introducing Bacillus into shrimp cultivation significantly reduced the number of Vibrio. These findings were consistent with the idea that B. licheniformis could increase the relative abundance of beneficial bacterial genera and decrease the relative abundance of potential pathogens.

Some studies have identified the genera such as Rhodobacter, Pseudomonas, and Paracoccus are the dominant bacterial groups in aquaculture water (Wu, 2016). In the present study, these genera were detectable in the aquaculture water, but at relatively low percentages. The genus Aurantimicrobium was found to be the most predominant in the aquaculture water, accounting for the largest proportion of the bacterial community. Numerous studies have found that Gammaproteobacteria is the absolutely dominant bacterial phylum in shrimp intestines, with Vibrio and Aeromonas being the absolutely dominant genera (Beardsley et al., 2011; Liu et al., 2010). However, in this study, the dominant bacterial genera in shrimp intestines were Gemmobacter, Fluviicola, and Aurantimicrobium. It is hypothesized that the aforementioned changes might be related to the fact that the present study was conducted in a small-scale farming setting. Microorganisms respond sensitively to environmental changes, and shifts in microbial genera are influenced by variations in the environment, with specific genera changing across different aquaculture environments (Quero et al., 2020). For example, Al-Masqari et al. (2022) cultured Penaeus vannamei under different temperature conditions and found that temperature changes significantly influenced the composition of bacterial communities. Furthermore, variations in probiotic species, strains, and application dosages can lead to differences in microbial community composition.