The rabbit experiments for antibody preparation in the study were carried out in accordance with protocols approved by the Animal Care and Welfare Committee at Shandong University School of Life Sciences (SYDWLL-2021-53).

The healthy shrimp M. japonicus (8–10 g) were purchased from a shrimp farm near Qingdao, Shandong Province, China. A shrimp culture system in the laboratory was used to cultivate the shrimp (40). For tissue distribution analysis, different organs, including the heart, hepatopancreas, gill, intestines, and stomach, were dissected from shrimp for protein and total RNA extraction. Vibrio anguillarum (10⁸ CFU) was fed into the mouthparts of shrimp as the immune challenge, except during the fluorescent immunocytochemistry assay. In the fluorescent immunocytochemistry assay, V. anguillarum (10⁶ CFU) was injected into the shrimp. PBS (140 mM NaCl, 2.7 mM KCl, 10 mM Na₂HPO₄, 1.8 mM KH₂PO₄, pH 7.4) was fed or injected into the control shrimp. Tissues were collected at different time points (3, 6, 12, 24, and 48 h) after bacterial infection for time-course expression analysis. The hemolymph was collected from the ventral sinus from three shrimp using a syringe containing an equal volume of precooled anticoagulant buffer (0.45 M NaCl, 10 mM KCl, 10 mM EDTA, and 10 mM HEPES, pH 7.45). The hemocytes were immediately harvested by centrifugation at 600×g for 8 min (4°C) from the experimental and control shrimp.

Different organs (heart, hepatopancreas, gills, stomach, and intestines) and hemocytes from three shrimp were homogenized in the Trizol reagent (TransGen Biotech, Beijing, China), and total RNA was extracted according to the manufacturer’s protocol. cDNA was synthesized using 5 μg of the total RNA via a SMART cDNA synthesis kit (Clontech, Beijing, China) following manufacturer’s instructions. The obtained cDNA was kept in a −20°C freezer until use.

The sequence of Nos was obtained from transcriptome sequencing in our laboratory. The amino acid sequence was predicted using the ExPASy tool (http://www.expasy.org). The online BLAST program (http://blast.ncbi.nlm.nih.gov/) was used to determine the similarity of NOS with other NOS proteins. The Simple Modular Architecture Research Tool (SMART) (http://smart.embl-heidelberg.de/) was employed to predict the domain architecture of NOS. Sequence alignment of Nos with other Nos genes was performed using MEGA 5.0 and GeneDoc software (http://www.flu.org.cn/en/download-47.html). The phylogenetic tree of NOS proteins was constructed using MEGA 5.0 with the neighbor-joining method (41).

Semiquantitative reverse transcription PCR (RT-PCR) was performed with cDNA obtained from different tissues as the template using primers Nos-RT-F and Nos-RT-R ( Table 1 ) for tissue distribution analysis. The PCR reaction protocol was as follows: 94°C for 3 min; 94°C for 30 s, 54°C for 30 s, 72°C for 30 s (35 cycles); and 72°C for 10 min. The ATCB (β-actin) gene, as an internal control, was amplified using primers β-actin-RT-Fand β-actin-RT-R ( Table 1 ).

Quantitative real-time PCR (qPCR) was used for time-course expression analysis of Nos and AMP genes in shrimp upon V. anguillarum challenge with Nos RT primers and antimicrobial peptide RT primers, and β-actin gene was used as an internal control ( Table 1 ). After reverse transcription of RNA to cDNA, the cDNA was used as the template for a qPCR reaction comprising 95°C for 3 min; 40 cycles of 95°C for 15 s and 59°C for 50 s, followed by melting from 72°C to 95°C. The reaction mix contained 5 μl of 2× Premix Ex Taq, 1 μl of diluted cDNA, and 2 μl (1 mM) each of a pair of primers. Each experiment was performed in triplicate. The qPCR data were analyzed using the 2^−△△Ct method (42). Student’s t-test was used to assess whether the differences between the date were significant (p < 0.05) and One-way ANOVA was used for multiple comparisons.

The template of dsRNA synthesis was obtained by RT-PCR amplification using primers Nos-Fi and Nos-Ri linked to the T7 promoter ( Table 1 ). The dsGfp (green fluorescent protein) served as the control and was amplified using primers Gfp-Fi and Gfp-Ri ( Table 1 ). DsRNA was synthesized using T7 RNA polymerase (Fermentas, Burlington, Canada) with an NTP mixture. For the RNA interference assay, the 50 µg of dsRNA was injected into the muscle of the penultimate segment of shrimp, and the same dose was injected again at 12 h. The efficiency of RNA interference was assessed at 48 h using qPCR. V. anguillarum was fed into mouthparts of shrimp at 48 h after the first injection.

The number of bacteria was detected at 6 h after the interference experiment or NOS inhibitor (^NG-monomethyl-l-arginine (L-NMMA)) injection. The stomach and intestines were dissected from three shrimp and homogenized in PBS. The obtained homogenate was diluted by 400- and 200-fold. Samples (50 μl) of the 400- or 200-fold diluted homogenate were placed on a plate containing Luria-Bertani (LB) medium for bacterial culture. The bacterial colonies were counted after incubation at 37°C overnight. The experiment was performed three times.

The method for bacterial load quantification using 16S rRNA genes mainly follows previous reports (43, 44). Briefly, for preparation of DNA sample of real-time PCR standards: Total microbiome DNA was extracted from shrimp gut samples using Genomic DNA Purufication Kit (Toyobo, Japan) and used for amplifying the target 16S rRNA genes by PCR via bacterial universal primers (891F and 1003R) (45) on a thermal Cycler T960 (Heal Force, Shanghai, China). The PCR product was then purified using SanPrep Column PCR Product Purification Kit (Sangon Biotech, Shanghai, China) and was quantified spectrophotometrically. A series of dilutions were then prepared, which were used as templates for quantitative real-time PCR with the primers (891F and 1003R). The cycle threshold (46) and the quantity of the template were used to make the standard curve for 16S rRNA gene quantification. The conditions of the qPCR assays of 16S rRNA genes were the same as those of the PCR described above. The copy number of 16S rRNA gene was calculated by following a previous method (43).

The gut samples from at least nine shrimp were collected and randomly divided to three samples of equal wet weight prior to DNA extraction. Total microbiome DNA was extracted using the above method for qPCR analysis with primers (891F and 1003R). The qPCR was performed to calculate 16S rRNA gene copies in shrimp using the standard curve. The qPCR procedure of 16S rRNA gene was the same as the ordinary qPCR described above, with the following modification, the PCR procedure consisted of an initial incubation at 95°C for 10 min, followed by 40 cycles of 95°C for 10 s and 60°C for 50 s, followed by a melting period from 65°C to 95°C. All the qPCRs were done in triplicate for both the standards and the microbial DNA samples.

The intestines from three shrimp were homogenized in radioimmunoprecipitation assay (RIPA) lysis buffer (Beyotime, Shanghai, China). The homogenate was centrifuged at 11,000×g for 10 min (4°C), and the supernatant was collected for further analysis. The supernatant was mixed with 3 × loading buffer (10% glycerin, 2% SDS, 0.1% bromophenol blue, 14.4 mM 2-mercaptoethanol, 50 mM Tris-HCl, pH 6.8) and incubated at 100°C for 10 min to denature the proteins fully. The sample was then separated using a 10% SDS-PAGE gel and transferred to a nitrocellulose membrane (0.45 μM) with transfer buffer (20% ethyl alcohol, 0.037% SDS, 25 mM Tris, 20 mM glycerin).

After blocking with 5% skimmed milk (in TBS, 10 mM Tris HCl, pH 7.5, 150 mM NaCl) for 1 h, the membrane was incubated with rabbit polyclonal antibodies (ERK antiserum, 1:200 dilution; β-actin antiserum, 1:500 dilution, phospho-ERK1-T202/Y204 + ERK2-T185/Y187, 1:5,000 dilution) for 1 h at room temperature, and then incubated at 4°C overnight. The ERK and β-actin antibodies were obtained by immunizing rabbits with recombinant proteins in the laboratory. Phospho-ERK1-T202/Y204 + ERK2-T185/Y187 rabbit antibodies were purchased from Abclone (AP0472, Wuhan, China). Histone-3 poly-clonal antibodies were purchased from Abclone (A2348, Wuhan, China). The membrane was washed three times with TBST (10 mM Tris HCl, pH 7.5, 150 mM NaCl, 0.02% Tween 20), 10 min each time, to remove nonspecifically bound antiserum. The membrane was then incubated with alkaline phosphatase-conjugated goat anti-rabbit IgG (ZB2308 ZSGB-Bio, Beijing, China) at room temperature for 3 h. The unbound antiserum was removed by washing three times with TBST for 10 min. TBS was then used to wash the membrane three times for 5 min each time. Thereafter, Substrate color developing solution, NBT (A610379, BBI)/5-bromo-4-chloro-3-indolyl phosphate (BCIP) (A610072, BBI) was added to visualize the signal in the dark at room temperature.

NO is easily soluble in water and generates nitrite; therefore, the concentration of nitrite in shrimp tissues was determined using the Gress Reagent colorimetric method (47). The stomach and intestine tissues were obtained from three shrimp, weighed, and placed into a glass homogenizer containing nitric oxide detection reagents (Beyotime). After fully homogenizing the tissues on ice, the homogenate was centrifuged at 4°C, 13,000×g for 10 min, and the supernatant was retained. The supernatant was tested for its nitrite concentration using an NO assay kit (Beyotime). According to the manufacturer, the NaNO₂ standard was used to construct a standard curve of NO.

For this experiment, the shrimp were divided into four groups. The L-NMMA (an inhibitor for NOS) solution (diluted in water) was injected into shrimp as pretreatment at the penultimate segment of the shrimp. The concentration of L-NMMA in shrimp of the different groups was 1.5, 3, and 6 µM. The other group of shrimp was injected with the same amount of ddH₂O as a control. The pretreated shrimp were used in the subsequent experiments, i.e., detection of the NO concentration at 3 h; detection of the number of bacteria at 6 h; immune challenge as described above; and detection of the number of bacteria at 6 h.

The experimental shrimp were randomly divided into four groups. Different doses of sodium nitroprusside (SNP, a kind of NO donor) were injected into three groups of shrimp, with the final concentration in each group being 5, 10, and 20 μM per shrimp. The control group was injected with the same amount of ddH₂O. The bacteria load at 6 h was then detected.

We first performed an inhibitor toxicity test. Shrimp were randomly divided into four groups and each group contained 10 shrimp. Then, L-NMMA was injected into shrimp at different concentrations (0, 1.5, 3, and 6 μM). Shrimp injected with the same amount of ddH₂O were used as the control. At 48 h postinhibitor treatment, shrimp survival was calculated. Next, 40 healthy shrimp were randomly divided into two groups; one group was injected with 3 μM L-NMMA and orally administrated with V. anguillarum (10⁸ CFU), and the control group was injected with ddH₂O and then challenged by the same amount of bacteria. The dead shrimp were monitored every 12 h, and the survival rate were calculated using GraphPad Prism 5.0 software (GraphPad Inc, La Jolla, CA, USA).

Survival rate of NO-donor, SNP-treated shrimp challenged by V. anguillarum was also analyzed. The toxicity test of SNP was firstly carried out as the method described above using three different concentrations (5, 10, and 20 μM). Healthy shrimp were then randomly divided into two groups (25 shrimp each), and the survival rate of shrimp with 10 μM SNP injection following bacterial challenge was analyzed, using the same amount of ddH₂O injection as the control.

First, a sterile syringe with 800 μl of prechilled anticoagulant buffer and 4% paraformaldehyde mixed solution (1:1) was used to collect an equal volume hemolymph from ventral sinus of three shrimp. After mixing, the solution was transferred to a precooled centrifuge tube and placed on ice for 10 min. The mixture was centrifuged at 600×g at 4°C for 5 min, and the supernatant was discarded. The pellet was gently suspended in 400 μl PBS, and then the suspension was dropped onto a cationic glass slide and incubated in the dark for 1 h. The unattached material was removed using six washes with PBS for 6 min each time. The glass slides were then treated with 0.2% Triton X-100 (in PBS) for 5 min to enhance the permeability of the cell membrane. Afterwards, the glass slides were washed with PBS for 5 min six times. The hemocytes were blocked in 3% BSA at 37°C for 30 min, and then incubated with antidorsal antibodies (1:20, diluted in 3% BSA) at 4°C overnight in the dark. The antidorsal antibodies were removed by washing six times with PBS. The cells on the slides were blocked with 3% BSA at 37°C for 5 min, and then incubated with Alexa fluor 488-conjugated anti-rabbit secondary antibodies (1:500 in 3% BSA) for 2 h at 37°C in the dark. The slides were washed with PBS six times for 5 min each and then incubated with DAPI (1:1,000 diluted in PBS) at room temperature for 10 min. After removing the DAPI by washing with PBS, the cells were covered with a coverslip and observed under an Olympus BX51 fluorescence microscope (Shinjuku-ku, Tokyo, Japan).

Data are presented as the mean ± the standard deviation (SD) of at least three replicates. Significance differences for paired comparisons were analyzed using Student’s t-test, asterisks in the figures indicate statistical significance (^* p < 0.05, ^** p < 0.01, and ^*** p < 0.001). One-way ANOVA was used for multiple comparisons; the different lowercase letters indicate significant differences (p < 0.05) in the ANOVA analysis. Densitometry analyses of Western blot bands were based on three independent replicates using ImageJ software (National Institutes of Health, https://imagej.nih.gov/ij/download.html). Shrimp survival rate was calculated, and the survival curves are presented as Kaplan-Meier plots and the statistically using a log-rank test. All statistical analysis was produced using GraphPad 5.0 data view software.

The cDNA sequence of Nos was obtained by intestinal transcriptome sequencing of M. japonicus. The domain architecture of the NOS comprises a NOS domain (AA 54–416), a flavodoxin–1 domain (AA 466–658), a FAD-binding–1 domain (AA 720–949), and an NAD-binding-1 domain (AA 981–1093) ( Figure 1A ). Phylogenetic analysis showed that the shrimp NOS was clustered with invertebrate NOS, which are distinct from vertebrate cNOS and iNOS ( Supplementary Figure S1 ).

The tissue distribution of Nos in normal shrimp was analyzed using qPCR, which showed that Nos is mainly distributed in the gastrointestinal tract, including the stomach and intestines and the nervous system ( Figure 1B ). It could also be detected in hemocytes and the epithelium ( Figure 1B ). The mRNA expression patterns of Nos in the stomach and intestines were measured after bacterial challenge. In the stomach, Nos increased significantly at 3 and 24 h after bacterial challenge and then returned to normal levels ( Figure 1C ). In the intestines, significant upregulation of Nos occurred at 6 to 12 h after pathogen challenge ( Figure 1D ). We also tested the concentration of nitrite (to represent the NO concentration in solution) in the shrimp gastrointestinal tract after feeding with V. anguillarum. The results showed that the concentration of nitrite in the stomach and intestines increased significantly after 3 to 6 h of bacterial stimulation ( Figures 1E, F ). These results suggested that NOS might be involved in antibacterial immunity in the gastrointestinal tract of shrimp.

To analyze the function of NOS in the antibacterial response, Nos RNA interference (RNAi) was performed, and the nitrite concentration and bacterial load in the gastrointestinal tract were analyzed. After knockdown of Nos ( Figures 2A, D ), the shrimp were infected orally with V. anguillarum. The concentration of nitrite in the stomach and intestines decreased compared with that in the dsGfp injection group ( Figures 2B, E ). Correspondingly, the bacterial load in the tested tissues increased by about 5.3- and 4.2-fold in the Nos-knockdown group compared with that in the control group ( Figures 2C, F ). These results suggested that Nos plays a role in the homeostasis of the microbiota in the shrimp gastrointestinal tract.

To further clarify the role of NOS in the gastrointestinal homeostasis of shrimp, L-NMMA was used to inhibit NOS activity in shrimp. The toxic effect of L-NMMA in shrimp was first analyzed by calculating the survival rate of shrimp after L-NMMA injection. Three different concentrations of L-NMMA (1.5, 3, and 6 μM/shrimp) were used in the experiments, following a previously reported method (48). The results showed that three concentrations of the inhibitor used in the experiment did not reduce the viability of the shrimp ( Supplementary Figure S2A ). The shrimp were then injected with three different concentrations of L-NMMA without further bacterial infection, and the nitrite concentration was analyzed in the shrimp gastrointestinal tract. The results showed that the concentrations of nitrite in gastrointestinal tract declined significantly in the inhibitor-injected shrimp in a dose-dependent manner ( Figures 3A, B ). The bacterial load increased significantly by 9.87 times in the stomach and 30.37 times in the intestines, of the 6μM/shrimp injection group compared with that in the control group ( Figures 3C, E ). Next, after injecting shrimp with L-NMMA, the shrimp were infected orally with V. anguillarum, and the bacterial number was analyzed. The results showed that the number of bacteria in the 6-μM L-NMMA injection group increased by 5.01 times in the stomach and 15.63 times in the intestines compared with that of the control group ( Figures 3D, F ). To confirm the above results, bacterial loads were also analyzed by detecting 16S rRNA gene copies via qPCR ( Supplementary Figure S3 ). The results showed that after being injected with 3 μM L-NMMA, the 16S rRNA gene copies were increased significantly compared with the control group with or without bacterial infection, in the stomach ( Figure 3G ) and intestines ( Figure 3H ). The survival rate of shrimp injected with 3 μM L-NMMA per shrimp was also analyzed. The results showed that the survival rate of the shrimp was reduced significantly after treatment with L-NMMA and bacteria compared with the control ( Figure 3I ). These results suggested that NOS plays an important role against bacterial invasion and might be involved in the maintenance of microbiota homeostasis in the shrimp gastrointestinal tract.

To confirm the role of NO in shrimp gut microbiota homeostasis regulation, the NO-donor, SNP injection was performed and bacterial loads were analyzed. We first performed toxicity test of the shrimp treated with different concentrations of SNP (5, 10, and 20 μM); the results showed that the different doses of the inhibitor used in the experiment did not reduce the viability of the shrimp ( Supplementary Figure S2B ). To determine whether NO can resist bacterial infection, uninfected shrimp were injected with the NO-donor, SNP, and the number of bacteria was analyzed 6 h later. The results showed that the number of bacteria decreased after SNP treatment ( Figures 4A, B ). The bacterial loads were also analyzed by detecting the 16S rRNA gene copies in SNP-treated shrimp challenged by bacteria via qPCR. The results showed that the 16S rRNA genes were decreased significantly both in the stomach and intestines of the shrimp ( Figures 4C, D ). The survival rate of SNP-treated shrimp challenged by V. anguillarum was increased significantly after treatment with SNP compared with the control ( Figure 4E ). The results suggest that the NO-donor treatment can increase NO level and further induce the expression of antibacterial effectors against infected bacteria in shrimp.

To analyze the possible antibacterial mechanism of NOS in shrimp, we analyzed the mRNA expression of AMPs, including antilipopolysaccharide factors (ALFs) and Crustins in Nos-knockdown shrimp challenged by V. anguilarum. The results showed that the expression of Alf-c1, e1, and Crus I-1, Crus I-2, and Crus I-5 was decreased significantly ( Figure 5A ). We also analyzed AMP expression in the stomach of Nos-RNAi-shrimp without bacterial challenge; the result showed that the expression levels of four Alfs (Alf-c2, Alf-d2, Alf-e1 and Alf-e2), and two CrusIs (CrusI-1 and CrusI-5) were also significantly decreased in the stomach of Nos-RNAi-shrimp ( Supplementary Figure S4A ). The AMP expression in the stomach of the L-NMMA-treated group challenged by bacteria was further analyzed; the results showed that the expression levels of seven Alfs and three CrusIs were decreased in the shrimp ( Figure 5B ). Similarly, the AMP expression in the stomach of the L-NMMA-treated shrimp without bacterial challenge was also analyzed, and similar results were obtained ( Supplementary Figure S4B ). All the results suggested that NOS might have constitutive expression in the stomach for NO production, and the NO in the stomach of shrimp with or without bacterial challenge acted as a signaling molecule to promote AMP expression.

The AMP expression in the intestines of Nos-RNAi- or L-NMMA-treated shrimp was further analyzed ( Figure 6 ). The expression levels of four Alfs (Alf-b1, Alf-c1, Alf-d2, and Alf-e1) and three CrusIs (CrusI-1, CrusI-2, and CrusI-5) were significantly decreased in the intestines of Nos-RNAi-shrimp with bacterial challenge ( Figure 6A ). We also analyzed AMP expression in intestines of Nos-RNAi-shrimp without bacterial challenge; the result showed that the expression levels of several Alfs were decreased in the shrimp ( Supplementary Figure S5A ). In the L-NMMA-treated shrimp challenged by bacteria, expression levels of Alf-b1, Alf-c2, Alf-d1, Alf-d2, Alf-e1, and Alf-e2, CrusI-1, CrusI-2, and CrusI-5 were declined significantly in the intestines of the shrimp compared with the control group ( Figure 6B ), and similar results were obtained in the intestines of the shrimp without bacterial challenge ( Supplementary Figure S5B ). The results suggested that NOS possessed constitutive expression in intestines for NO production, and the NO in intestines of the shrimp acted as a signaling molecule to promote AMP expression.

All above results suggested that the NOS possessed constitutive and inducible characteristics in gastrointestinal tract of shrimp, the NO generated by NOS in shrimp with or without bacterial challenge act as a signaling molecule to regulate AMP expression, and the AMPs expressed in the gastrointestinal tract were responsible for regulation of microbiota homeostasis in shrimp.

AMP expression is regulated by Toll, IMD, JAK-STAT pathways, or FOXO signaling in shrimp (34, 49, 50), and the transcription factors (including dorsal, relish, or STAT) of the pathways were phosphorylated and translocated into nucleus to promote AMP expression in response to pathogen infection. Bacterial infection upregulates NOS and promotes the activation of the NF-κB-like transcription factor in Drosophila (11). The level of phosphorylated ERK (p-ERK) increased after NO donor treatment in human breast MCF7 cells (51), and p-ERK promotes the translocation of the nuclear factor kappa B (NF-κB)-like transcription factor, dorsal, into the nucleus to regulate the expression of AMPs (52). To analyze how the NOS regulated AMP expression in shrimp, we firstly detected the ERK phosphorylation after knockdown of Nos or NOS inhibitor L-NMMA treatment in shrimp. The results showed that the level of p-ERK was significantly decreased in the intestines of Nos-RNAi shrimp ( Figures 7A, B ) and L-NMMA-treated shrimp ( Figures 7C, D ) after bacterial challenge for 1 h compared with that in the control group. However, no significant decrease was detected in the shrimp without bacterial challenge ( Figures 7A, D ). The results suggested that the increased NO level in shrimp challenged by bacteria is responsible for phosphorylation of ERK.

To analyze if the phosphorylated ERK is related with dorsal phosphorylation and nuclear translocation, the subcellular distribution of dorsal was analyzed by Western blotting and fluorescent immunocytochemical assays. The translocation of dorsal into the nucleus decreased significantly in the intestines of L-NMMA-treated shrimp challenged by V. anguillarum compared with that in the control group ( Figures 8A, B ). Similar results were obtained in hemocytes of L-NMMA-treated shrimp challenged by bacteria analyzed by fluorescent immunocytochemical assay ( Figures 8C, D ). These results suggested that the activated ERK by NO signaling phosphorylated Dorsal and induced nuclear translocation of the transcription factor and then positively regulated expression of AMPs in shrimp.