Beneficial effects of SB on growth performance in whiteleg shrimp
First, we examined the effect of SB at 0%, 0.04%, 0.2% and 1% on the growth performance of whiteleg shrimp for 4 weeks in a laboratory setting. Growth performance metrics, including BW, BL, food consumption, and FCR of each group, were measured at the indicated times (Table S1). The control group and three treatment groups (SB 0.04%, SB 0.2% and SB 1%) had average initial BW of 3.26 g, 3.27 g, 3.29 g and 3.31 g, respectively (Table S1). No significant difference was observed among the four groups. The control group and three treatment groups (SB 0.04%, SB 0.2% and SB 1%) showed increased average BW (13.68%, 14.05%, 17.99% and 20.34%, respectively) at 2 weeks post treatment (2 weeks, Fig. 1A). There was statistical significance between the control and two treatment groups (SB 0.2% and SB 1%) (2 weeks, Fig. 1A). The control group and three treatment groups (SB 0.04%, SB 0.2% and SB 1%) also showed increased average BW (33.5%, 36.25%, 47.23%, and 54.43%, respectively) at 4 weeks post treatment (4 weeks, Fig. 1A). There was statistical significance between the control and two treatment groups (SB 0.2% and SB 1%) (4 weeks, Fig. 1A).
The control group and three treatment groups (SB 0.04%, SB 0.2%, and SB 1%) had average initial BL of 65.75 mm, 65.85 mm, 65.68 mm, and 66.11 mm, respectively (Table S1). No significant difference was observed among the four groups. The control group and three treatment groups (SB 0.04%, SB 0.2% and SB 1%) showed increased average BL (2.90%, 13.20%, 4.82% and 5.53% respectively) 2 weeks post treatment (2 weeks, Fig. 1B). There was statistical significance between the control and two treatment groups (SB 0.2% and SB 1%) (2 weeks, Fig. 1B). The control group and three treatment groups (SB 0.04%, SB 0.2% and SB 1%) showed increased average BL (6.63%, 7.45%, 12.30%, and 13.37%, respectively) at 4 weeks post treatment (4 weeks, Fig. 1B). There was statistical significance between the control and two treatment groups (SB 0.2% and SB 1%) (4 weeks, Fig. 1B). Representative photographs of each shrimp group are shown (4 weeks, Fig. 1C).
Furthermore, the control group and three treatment groups (SB 0.04%, SB 0.2% and SB 1%) had an average FCR of 1.76, 1.77, 1.53, and 1.42, respectively, 2 weeks post treatment (2 weeks, Fig. 1D). There was statistical significance between the control and two treatment groups (SB 0.2% and SB 1%) (2 weeks, Fig. 1D). Likewise, the control group and three treatment groups (SB 0.04%, SB 0.2% and SB 1%) had an average FCR of 2, 1.84, 1.61, and 1.59, respectively, 4 weeks post treatment (4 weeks, Fig. 1D). Statistical significance was found between the control and two treatment groups (SB 0.2% and SB 1%) (4 weeks, Fig. 1D). Overall, the above data indicated that SB dose-dependently promoted the growth performance of shrimp and that the minimum effective dose was 0.2%.
Since water quality is important for shrimp farming, next, water quality metrics were monitored for 28 days in a laboratory setting (Table S2). The average water temperature of the four shrimp groups was 28.11–28.55 °C. Their average salinity was 15.17–15.96% and their average pH value was 7.96–8.56. In addition, the average dissolved oxygen was 9.36–9.67 mg/mL and the average nitrite was 0.35–0.66 mg/mL. The average ammonia nitrogen of the three shrimp groups was 0.12–0.29 mg/mL.
Finally, we assessed the effect of SB on shrimp growth over 135 days in a field trial (Table 1). The data showed that the initial BW of shrimp in the control and two treatment groups (SB 0.2% and SB 1%) were 0.38 g, 0.37 g and 0.37 g, respectively (Table 1). No significant difference was observed in the initial BW among the four groups (Table 1). The control and treatment groups (SB 0.2% and SB 1%) showed increased average BW to 22.38 g, 25.51 g and 26.29 g, respectively, 135 days post treatment (Table 1). Statistical significance between the control and two treatment groups (SB 0.2% and SB 1%) was found in the final BW (Table 1).
The initial BL of shrimp in the control and two treatment groups (SB 0.2% and SB 1%) were 28.58 mm, 28.02 mm, and 28.14 mm, respectively (Table 1). No significant difference was observed among the three groups (Table 1). Similarly, the control and treatment groups (SB 0.2% and SB 1%) had average final BL of 118.59 mm, 136.54 mm, and 142.66 mm, respectively (Table 1). Statistical significance was found between the control and two treatment groups (SB 0.2% and SB 1%) (Table 1). Overall, the above data indicated that SB dose-dependently promoted the growth performance of shrimp in farm trials.
Water quality metrics in this trial were monitored for 135 days (Table S3). The average water temperature of the water housing the four shrimp groups was 29–30 °C. The average salinity was 20% and the average pH value was 7.89–8.39. In addition, the average dissolved oxygen was 9.32–9.77 mg/mL and the average nitrite was 0. 53–0.8 mg/mL. The average ammonia nitrogen for the three shrimp groups was 0.45–0.49 mg/mL.
SB protects against V. parahaemolyticus infection in shrimp
Next, we tested the effect of SB on shrimp in protecting against vibriosis (Fig. S2B) based on survival, bacterial counts, and hepatopancreatic pathology. As expected, the NIC group had 100% survival (NIC, Fig. 2A). However, the IC group had a survival rate of 75% (IC, Fig. 2A). In contrast, SB dose-dependently increased this survival rate (SB, Fig. 2A). Furthermore, we examined hepatopancreatic pathology in the 5 shrimp groups. First, gross examination data showed that the IC shrimp had a yellowish to whitish hepatopancreas compared to the black hepatopancreas in the NIC shrimp (top, IC vs. NIC, Fig. 2B). However, SB treatment improved the hepatopancreatic pathology based on its color (top, SB, Fig. 2B). Next, the IC group had a shortened gut compared to the NIC group. In sharp contrast, SB had a longer gut than IC group (IC vs. SB, Fig. 2B). Accordingly, hepatopancreatic morphology data indicated that SB treatment improved the hepatopancreatic pathology (middle, SB, Fig. 2B). In parallel, microscopic examination showed that shrimp without V. parahaemolyticus challenge had a normal architecture of hepatopancreas with a high number of B, F, and R cells (top, NIC, Fig. 2C). In contrast, V. parahaemolyticus profoundly damaged the hepatopancreatic structure and reduced the number of B, F, and R cells in the shrimp hepatopancreas (top, IC, Fig. 2C). However, SB treatment dose-dependently reduced hepatopancreatic lesions and restored the number of B, F, and R cells in the shrimp hepatopancreas (bottom, SB, Fig. 2C). We also determined the percentage of healthy cells of shrimp hepatopancreas (right, Fig. 2C). Finally, we determined the bacterial counts in different organs of five shrimp groups (SB, Fig. 2D). First, few bacteria in the hepatopancreas of the NIC shrimp grew on TCBS plates (NIC, Fig. 2D). As expected, the hepatopancreas of IC shrimp showed the most bacteria on TCBS plates (IC, Fig. 2D). However, SB dose-dependently reduced the bacteria in the hepatopancreas of the infected shrimp (SB, Fig. 2D). Likewise, SB dose-dependently reduced bacterial counts in the muscle (SB, Fig. 2D), stomach (SB, Fig. 2D), and gut (SB, Fig. 2D) of the infected shrimp. Taken together, SB effectively rescued shrimp from vibriosis.
Protective effects of SB on survival rate, hepatopancreatic structure and bacterial counts in different organs of shrimp following V. parahaemolyticus infection. (A–D) Five groups of shrimp were fed with control diets and diets containing 0.04%, 0.2% and 1% SB (Groups 3–5), followed by infection with V. parahaemolyticus. The survival rate (A), digestive tract (B), hepatopancreatic structure (left, C), percentage of healthy hepatopancreatic cells (right, C), and bacterial load (D) in the gut (GU), stomach (ST), muscle (MS), and hepatopancreas (HP) of the un-medicated non-infected controls (NIC), un-medicated infected controls (IC) and SB-treated shrimp groups (0.04%, 0.2% and 1% SB) were measured. Blue arrowhead, yellow arrowhead, and black arrowhead indicate B, F and R cells of hepatopancreas.
SB increases probiotics and reduces pathogenic bacteria in shrimp gut
To explore the mechanism through which SB affects growth performance and resistance to Vibrio in whiteleg shrimp, the gut microbiota of shrimp were analyzed using 16S rRNA next-generation sequencing (NGS) analysis. The number of sequences, operational taxonomic units (OTUs), and diversity indices in the gut digesta of control shrimp and shrimp fed with SB (1%) are summarized in Table S4. Rarefaction curves showed that the number of sequences from 2 shrimp groups were enough to reveal the major OTUs (Fig. S3A). The gut microbiota of control shrimp (CTR) aged 126 days, had higher diversity than SB-fed shrimp as evidenced by Shannon and Chao1 diversity indices (Table S4). Detailed changes at the phylum, class, order, family, and genus levels under SB treatment are shown in Table S5 and Fig. S2B–E. Nine genera of probiotics and 10 genera of pathogens were identified from shrimp guts (Genus, Table S5). Among them, SB increased 6 beneficial bacterial genera, Lactobacillus, Megasphaera, Bifidobacterium, Prevotella, Ruminococcus and Collinsella (Fig. 3A). In contrast, SB decreased 6 harmful bacterial genera, Vibrio, Photobacterium, Pseudoalteromonas, Planctomicrobium, Tenacibaculum and Corynebacterium_1 (Fig. 3B). Next, we used selective medium plates to analyze the change in the number of Lactobacillus species and Vibrio species in the intestinal digesta of 4 shrimp groups 28 days post treatment. We found that SB significantly increased the proportion of Lactobacillus species in the shrimp digesta in a dose-dependent fashion using the lactic acid bacteria screening medium, MRS (Fig. 3C). However, SB significantly decreased the proportion of Vibrio species in the shrimp digesta in a dose-dependent manner using the Vibrio screening medium, TCBS (Fig. 3D).
SB increases 6 probiotic genera and decreases 6 genera of pathogenic bacteria in shrimp guts. (A–B) The whole gut of 114-day-old shrimp, 9 animals a group, from 4 groups (Fig. 1) were collected to obtain gut bacteria DNA, followed by bacterial 16S rRNA NGS analysis. The proportion of probiotics (A) and pathogens (B) of the gut microbiota were analyzed at the genus level. (C–D) The same gut bacteria from Fig. 3A and B were plated on MRS plates and TCBS plates for bacterial counting (top). The count of Lactobacillus (LAB, C) and Vibrio species (D) in each group was re-plotted into histograms (bottom). (E–F) Primers specific for L. johnsonii (E) and Vibrio parahaemolyticus (F) were used to quantify the abundance of both species in the shrimp gut microbiota (Fig. 3A and B) using PCR (left). The fold change of both species was re-plotted into histograms (right). DNA marker (M) and amplicon of gut bacterial DNA of shrimp fed with SB (0.04%, 0.2%, and 1%), water (NC) and DNA of L. johnsonii or Vibrio parahaemolyticus (PC). Data from 3 repeats are presented as the mean ± SD. One-way ANOVA test was used for statistical analysis of differences between groups and P (*) < 0.05, P (**) < 0.01, and P (***) < 0.001 are considered statistically significant.
Furthermore, we used MALDI-TOF MS to identify Lactobacillus species as well as Vibrio species in the intestinal digesta of control whiteleg shrimp (Fig. S4). MS profiles of bacterial proteins were searched against an in-house database using Biotyper 3.1 as published28. The data showed the dominant presence of L. johnsonii which was more prevalent than L. reuteri in the gut digesta of control shrimp (Table 3). We also discovered the dominant presence of V. parahaemolyticus, which as more prevalent than one unidentified Vibrio spp. in the shrimp digesta (Table 3). Hereafter, L. johnsonii, L. reuteri, V. parahaemolyticus, and/or Vibrio spp. were used in the study.
Finally, we confirmed the presence of L. johnsoni and V. parahaemolyticus in the gut digesta of the four shrimp groups from Fig. 1 using semi-quantitative polymerase chain reaction (PCR) with a primer pair of 16S rRNA of and irgB, respectively. We demonstrated that SB significantly increased the proportion of Lactobacillus species in the shrimp digesta in a dose-dependent manner (Figs. 3E and S8A). On the contrary, SB significantly decreased the proportion of Vibrio species in the shrimp digesta in a dose-dependent manner using the Vibrio screening medium, TCBS (Figs. 3F and S8B).
Collectively, the overall data demonstrated that SB regulated the intestinal flora of shrimp mainly through augmenting probiotics and reducing pathogens.
SB inhibits the growth of V. parahaemolyticus via an increase of L. johnsonii growth and its antimicrobial metabolites
To tease out the mechanism through which SB promoted probiotics and inhibited pathogenic bacteria, we first tested the in vitro effects of SB on growth of L. johnsonii. The MAC experiments indicated that a negative control, 30 µg/mL Amp, completely inhibited the growth of L. johnsonii in MRS medium (Amp, Fig. 4A). In sharp contrast, a positive control, 0.1% peptone, increased the growth of L. johnsonii (Peptone, Fig. 4A). Of note, SB at 0.5 µg/mL and more significantly promoted the growth of L. johnsonii in a dose-dependent manner (SB, Fig. 4A). Next, we assessed the in vitro effects of SB on growth of V. parahaemolyticus. The MIC experiments showed that a positive control, 30 µg/mL Amp, completely inhibited the growth of V. parahaemolyticus in TSB medium (Amp, Fig. 4B). However, SB at 200 µg/mL and more significantly inhibited the growth of V. parahaemolyticus in a dose-dependent manner (SB, Fig. 4B). The data suggested that SB directly promoted growth of probiotics, which antagonized pathogens.
SB suppresses Vibrio growth via up-regulation of Lactobacillus growth and its antimicrobial metabolites. (A) L. johnsonii (LJ) was cultured in MRS medium containing Amp, peptone, and SB at the indicated dosages at 37 °C under anaerobic conditions for 11 h. Bacterial growth rate (%) is shown. (B) The same procedure as (A) was performed except that V. parahaemolyticus (VP) was grown in TSB under aerobic conditions for 6 h. (C) Antagonism of L. reuteri (LR) and LJ toward pathogens. VP (1st column, top) and Vibrio spp. (VS) (2nd column, top) were spread on TSA and incubated with MRS agar (NC1), a paper disc containing 150 µl water (NC2) and 0.125 µg Chl (PC). Conversely, LJ (3rd column, top) and LR (4th column, top) were spread on MRS agar plates and incubated with TSA (NC3), and a paper disc containing 150 µl water (NC4) and 0.125 µg Chl (PC). Their inhibition zones are shown. (D) Antagonism of the metabolites of LJ toward pathogens. VP (1st column, left) and VS (2nd column, left) were spread on TSA and incubated with 150 µl MRS medium (NC1), 150 µl water (NC2), and a hole containing 0.125 µg Chl (PC), and 150 µl supernatant of LJ (LJSN) and LR (LRSN). Their inhibition zones are shown. (E) SB up-regulated 5 antimicrobial metabolites of LJ as characterized in Fig. S5. (F) Antagonism of the mixture of 5 metabolites of SB-treated LJ toward VP. VP was spread on TSA and incubated with 150 µl MRS medium (NC1), 150 µl water (NC2), 0.125 µg Chl 150 µl (PC), the supernatant of LJ (LJSN) and SB-treated LJ (LJSN + SB), and the mixture of five metabolites (5AM) present in the supernatant of LJ (5AM) and SB-treated LJ (5AM + SB). Their inhibition zones are shown. Data from 3 repeats are presented as the mean ± SD. One-way ANOVA test was used for statistical analysis of differences between groups and P (*) < 0.05, P (**) < 0.01, and P (***) < 0.001 are considered statistically significant. Scale bar = 10 mm.
Next, we performed antagonistic tests between two Lactobacillus species, L. johnsonii (LJ) and L. reuteri (LR), and two Vibrio species, V. parahaemolyticus (VP) and Vibrio spp. (VS) using disc diffusion assays. As expected, two negative controls, MRS agar and water, could not inhibit V. parahaemolyticus as characterized by lack of inhibition zone (NC1 and NC2, VP (1st column), Fig. 4C). In contrast, a positive control (PC), a disc containing 0.125 µg Chl, inhibited the growth of V. parahaemolyticus as shown by inhibition zones (PC, VP (1st column), Fig. 4C). Likewise, a disc containing L. johnsonii and L. reuteri significantly inhibited V. parahaemolyticus as inhibition zones (LJ and LR, VP (1st column), Fig. 4C). We noticed that L. johnsonii exhibited slightly better inhibition of V. parahaemolyticus than L. reuteri and the positive control, a disc containing 0.125 µg Chl (LJ, LR and PC, VP (1st column), Fig. 4C). In parallel, we conducted antagonistic tests between two Lactobacillus species and Vibrio spp. Using disc diffusion assays. L. johnsonii exhibited slightly better inhibition of Vibrio spp. Than L. reuteri and the positive control, a disc containing 0.125 µg Chl (LJ, LR, and PC, VS (2nd column), Fig. 4C).
Furthermore, disc diffusion assays were applied to test the action of V. parahaemolyticus and Vibrio spp. On two Lactobacillus species. As expected, two negative controls, TSA and water vehicle, failed to inhibit L. johnsonii as characterized by the lack of an inhibition zone (NC3 and NC4, LJ (3rd column), Fig. 4C). In contrast, a positive control (PC), a disc containing 0.125 µg Chl, inhibited growth of L. johnsonii as shown by inhibition zones (PC, LJ (3rd column), Fig. 4C). Likewise, discs containing V. parahaemolyticus and Vibrio spp. failed to inhibit the growth of L. johnsonii based on inhibition zones (VP and VS, LJ (3rd column), Fig. 4C). In parallel, we found that the positive control, a disc containing 0.125 µg Chl, could inhibit the growth of L. reuteri (PC, LR (4th column), Fig. 4C). However, no antagonism of two Vibrio species, V. parahaemolyticus and Vibrio spp., and two negative controls TSA and water, toward L. reuteri was seen (VP, VS, NC3 and NC4, LR (4th column), Fig. 4C). The overall data revealed antagonism of Lactobacillus species toward Vibrio species and not vice versa.
To further understand the mechanism by which Lactobacillus inhibited Vibrio, agar well diffusion assays were performed to measure the antagonism between the supernatants of L. johnsonii and L. reuteri and Vibrio species. As expected, two negative controls, MRS medium and water, failed to inhibit V. parahaemolyticus as characterized by lack of an inhibition zone (NC1 and NC2, VP, Fig. 4D). In contrast, a positive control (PC), 0.125 µg Chl, inhibited the growth of V. parahaemolyticus as shown by inhibition zones (PC, VP, Fig. 4D). Likewise, the cell-free supernatant of L. johnsonii (LJSN) and L. reuteri (LRSN) significantly inhibited V. parahaemolyticus as shown by inhibition zones (LJSN and LRSN, VP, Fig. 4D). The supernatant of L. johnsonii had a slightly more potent inhibition of V. parahaemolyticus than that of L. reuteri and the positive control, 0.125 µg Chl (LJSN, LRSN, and PC, VP, Fig. 4D). In parallel, we performed antagonistic tests between the supernatant of two Lactobacillus species and Vibrio spp. using agar well diffusion assays. The supernatant of L. johnsonii had a slightly more potent inhibition of Vibrio spp. than that of L. reuteri and the positive control, 0.125 µg Chl (LJSN, LRSN, and PC, VS, Fig. 4D). Furthermore, we tried to identify the antibacterial metabolites present in the supernatant of L. johnsonii using LC–MS/MS. Consequently, five metabolites of L. johnsonii, LA, AA, PA, 3-HPA, and BA, were identified from the supernatant of L. johnsonii (CTR, Fig. 4E) compared to their standards (Fig. S5). Furthermore, LC–MS/MS data revealed that SB significantly up-regulated the above metabolites in the supernatant of L. johnsonii at the indicated time (Fig. 4E and Table S6) as well as in the gut microbiota (Fig. 5E and Table S7). We also evaluated the antimicrobial effects of the 5 metabolites on V. parahaemolyticus using agar well diffusion assays. No inhibition of V. parahaemolyticus growth was observed in a disc containing MRS medium and distilled water but inhibition was observed for Chl (NC1 and NC2 vs. PC, Fig. 4F), whilst the supernatant of L. johnsonii inhibited V. parahaemolyticus growth (LJSN, Fig. 4F) to a lesser degree than that of L. johnsonii with SB treatment (LJSN + SB, Fig. 4F). We also evaluated the antimicrobial potency of the 5 metabolites, in combination, at the dosage that equaled their quantity in the supernatant of L. johnsonii. A mixture of the 5 metabolites showed significant inhibition of V. parahaemolyticus growth (5AM, Fig. 4F). In addition, we tested the antimicrobial potency of a mixture of the 5 metabolites at the dosage that equaled their quantity in the L. johnsonii supernatant with SB treatment for 7 h. This mixture showed significant inhibition of V. parahaemolyticus growth (5AM + SB, Fig. 4F). Obviously, a mixture of the 5 metabolites (5AM + SB) and supernatant of SB-treated L. johnsonii (LJSN + SB) corresponding to their amount in SB treatment groups, had superior inhibition of V. parahaemolyticus compared to their amount in control groups (5AM and LJSN) (Fig. 4F). Overall, SB antagonized growth of V. parahaemolyticus through up-regulation of the antimicrobial metabolites from L. johnsonii.
Anti-pathogenic mechanism of 5 antimicrobial metabolites produced by L. johnsonii. (A) The IC50 of each antimicrobial metabolite for V. parahaemolyticus (VP) was determined. (B–C) V. parahaemolyticus (VP) (1 × 106 CFU/mL) were grown in TSB containing vehicle (NC), 30 µg/ml ampicillin (Amp), or each antimicrobial metabolite at the dosage that equaled their quantity in the supernatant of L. johnsonii for 90 min, followed by PI staining. The bacteria were divided into 2 aliquots. One aliquot of the bacteria was analyzed using flow cytometry (B). The other aliquot was analyzed with fluorescent microscopy (left, C). The percentage (%) of PI-positive (dead) cells was determined (right, C). (D) V. parahaemolyticus (VP) was treated with 30 µg/ml ampicillin and a mixture of the 5 antimicrobial metabolites which equaled the composition of the metabolites in the supernatant of SB-treated L. johnsonii, for the indicated times. The bacteria, V. parahaemolyticus were analyzed using TEM. Their representative images are shown (left). Arrowheads in yellow, red, white, and blue indicate separation of the cytoplasmic and outer membranes, distorted outer membrane, empty cells, and membrane discontinuity, respectively. Their death (%) was quantified and re-plotted into histograms (right). Scale bar = 1 µm. (E) Shrimp (86 days old) were fed with a standard diet (CTR) and a diet containing SB at the indicated dosages for 4 weeks. Their gut digesta were subjected to LC–MS/MS analysis, followed by identification and quantification of 5 antimicrobial metabolites (µg/g of digesta) as shown in Fig. S6 and Table S7. Data from 3 repeats are presented as the mean ± SD. One-way ANOVA test was used for statistical analysis of differences between groups and P (*) < 0.05, P (**) < 0.01, and P (***) < 0.001 are considered statistically significant.
Antimicrobial metabolites of L. johnsonii suppress V. parahaemolyticus via membrane destruction
To investigate the antimicrobial mode of action of these metabolites produced by Lactobacillus, we assessed the bactericidal activities of individual metabolites toward V. parahaemolyticus. The IC50 values of the 5 compounds against V. parahaemolyticus in ascending order were: 3-HPA (0.37 µg/mL) < PA (4.70 µg/mL) < BA (4.86 µg/mL) < AA (5.30 µg/mL) < LA (14 µg/mL) (Fig. 5A). Next, we investigated cell death of V. parahaemolyticus using a flow cytometer and a microscope. Flow cytometry data showed that no cell death of V. parahaemolyticus before treatment with a mixture of the 5 metabolites was shown by propidium iodide (PI) staining (0 min, Fig. 5B). However, Amp induced death in V. parahaemolyticus (Amp, Fig. 5B). Consistently, a mixture of the 5 metabolites at the dosage that equaled their quantity in the supernatant of L. johnsonii caused a higher death of V. parahaemolyticus over time (VP, Fig. 5B). The above data was confirmed by fluorescent microscopy (5AM, Fig. 5C). Transmission electron microscopy (TEM) showed that, in the absence of the mixture of the 5 metabolites, the control bacteria had intact membranes and regular cytoplasm (0 min, Fig. 5D). In marked contrast, 30 min treatment with the mixture increased the percentage of damaged/dead cells in V. parahaemolyticus to 32.33% (30 min, Fig. 5D). Likewise, this percentage increased to 75.33% after 90 min treatment (90 min, Fig. 5D). We also confirmed the presence of the 5 metabolites in the digesta of control and SB-fed shrimp (Table S7 and Fig. 5E). The LC–MS/MS data showed that the amount of the 5 metabolites per gram in control shrimp guts in descending order was: LA (18.06 µg) > AA (17.13 µg) > 3-HPA (3.14 µg) > PA (0.20 µg) > BA (0.06 µg) (CTR, Fig. 5E). In contrast, SB dose-dependently increased the amount of these 5 metabolites. The amount of the 5 metabolites per gram of 1% SB-fed shrimp digesta in descending order was: AA (46.29 µg) > LA (34.53 µg) > 3-HPA (11.85 µg) > PA (0.28 µg) > BA (0.18 µg) (SB, Fig. 5E). Overall, SB up-modulated the level of 3-HPA, AA, and BA to a greater extent than LA and PA in shrimp guts. Overall, SB significantly escalated the level of these metabolites in shrimp.
Identification of active compounds of SB with an ability to promote growth of Lactobacillus
Finally, we tried to identify the active compounds from SB based on L. johnsonii growth. Using a bioactivity-directed fractionation and isolation strategy, we first obtained the crude extract (CE) of SB and then partitioned the CE into water and butanol (BuOH) fractions (Fig. S7A). The CE, BuOH fraction, and water fraction were incubated with L. johnsonii to assess their bioactivities. As a result, the CE and BuOH fraction but not water fraction promoted the growth of L. johnsonii (Fig. 6A and B and Table 4). Next, we purified 4 compounds from the BuOH fraction of SB, palmitic acid, linoleic acid, linolenic acid, and stearic acid, which were characterized using GC-GC/MS (Fig. S7B and Table 5). Meanwhile, we evaluated the MAC values of these 4 compounds based on L. johnsonii growth We found that linolenic acid had higher growth-promoting activity toward L. johnsonii than linoleic acid and stearic acid (Fig. 6C and Table 5). In contrast, palmitic acid showed no growth promotion activity toward L. johnsonii (Fig. 6C and Table 5). Of note, the overall data demonstrated that SB promoted the growth performance and reduced vibriosis in shrimp via regulation of gut microbiota, i.e., increased probiotics and reduced pathogens (Fig. 6D).
Identification of active compounds of SB with an ability to boost Lactobacillus growth. (A) The crude extract (CE) of SB was fractionated into butanol (BuOH) and water (H2O) fractions based on the bioactivity-directed fractionation and isolation procedure (Fig. S7). L. johnsonii (LJ) was cultured in MRS medium (CTR) and the medium containing ampicillin (Amp, 30 µg/ml) and the BuOH fraction at 0.125, 0.25, 0.5,1 and 2 µg/ml at 37 °C under anaerobic conditions for 10 h. The growth rate (%) of bacteria was obtained from the ratio of the OD600 of the treatment group to that of the control group multiplied by 100%. (B) The same procedure as (A) was conducted except that MRS medium (CTR) and the medium containing Amp, H2O fraction at 0.125, 0.25, 0.5, 1 and 2 µg/ml was used. (C) The same procedure as (A) was conducted except that palmitic acid, stearic acid, linoleic acid and linolenic acid at the indicated dosages were used. (D) In eubiosis, probiotics and pathogens are balanced in gut microbiota of healthy shrimp. In dysbiosis, probiotics are outcompeted by pathogens in shrimp gut and, in turn, this imbalance of microbiota causes host diseases. SB can promote the growth of probiotics and the production of their antimicrobial metabolites, leading to decreased pathogens. As a result, SB can enhance gut health, host health, and growth performance as well as rectify vibriosis and other disease in shrimp.
In conclusion, SB was able to augment the growth performance and host defense to Vibrio in whiteleg shrimp. This augmentation arose from SB-mediated regulation of gut microbiota, i.e., the promotion of probiotics and their antimicrobial metabolites, which counteracted the growth of pathogens. Accordingly, three active ingredients of SB were characterized for increased probiotic growth.
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- Source: https://www.nature.com/articles/s41598-024-62436-x