All experimental procedures involving giant freshwater prawns (M. rosenbergii) were reviewed and approved by the Laboratory Animals Research Center, University of Phayao, and the Animal Ethics Committee of the University of Phayao, Thailand (Approval No. 63 01 04 010; approved on 4 August 2020). The study was conducted in accordance with institutional guidelines for the care and use of aquatic animals in research and relevant standards for animal welfare, biosafety, and humane experimental practice.

Healthy adult prawns were acclimatized before the experiment and maintained under controlled water quality conditions to minimize stress. Animal handling, dietary intervention, bacterial challenge with A. hydrophila, hemolymph collection, tissue sampling, and survival monitoring were performed only by trained personnel using standardized procedures. The number of prawns used was limited to the minimum required to achieve the study objectives, and the sample size was determined before experimentation using G*Power software. During the challenge experiment, animals were monitored regularly for abnormal behavior, morbidity, and mortality. All efforts were made to reduce pain, distress, and unnecessary handling throughout the study. Moribund prawns and dead animals were promptly removed from the tanks, and biological waste and bacterial cultures were handled and disposed of in accordance with institutional biosafety procedures.

The study was conducted from March to May 2021 at the laboratory and aquaculture research facilities of the University of Phayao, Phayao, Thailand. Healthy adult M. rosenbergii were obtained from a local commercial farm in Chiang Rai, Thailand, and transported to the laboratory for acclimatization before the feeding trial and bacterial challenge experiment.

This study evaluated the effects of partial replacement of fishmeal with fermented L. leucocephala on immune responses and resistance to A. hydrophila infection in adult M. rosenbergii. Three dietary treatments were prepared: a control diet, a diet in which 10% of fishmeal was replaced with fermented L. leucocephala, and a diet in which 20% of fishmeal was replaced with fermented L. leucocephala. After acclimatization, prawns were randomly allocated to dietary groups and fed for 2 weeks before challenge with A. hydrophila. Hemolymph, hematopoietic tissue, and intestinal samples were collected at predetermined time points for immunological, histological, and molecular analyses. A separate survival experiment was conducted using three independent replicate tanks per treatment group.

Healthy M. rosenbergii weighing 25–30 g were obtained from a local farm in Chiang Rai, Thailand, and maintained in the laboratory under acclimatized conditions for 7 days before the start of the experiment. Adult prawns were selected because the present study primarily focused on immune status and immune-related responses following dietary supplementation and bacterial challenge. In addition, their larger body size facilitated the collection of hemolymph for immunological and molecular analyses. During acclimatization, prawns were fed a commercial diet three times daily at 07:00, 13:00, and 19:00 h.

The A. hydrophila culture and the protocol for determining the median lethal dose in M. rosenbergii were described previously [20]. The stock culture of L. plantarum TISTR1284 was stored in the laboratory at −80°C in de Man, Rogosa, and Sharpe medium (HiMedia Laboratories Pvt. Ltd., Mumbai, India) containing 20% glycerol. The thawed stock culture was cultivated in fresh medium and incubated at 37°C for 48 h. The culture was harvested, and the optical density at 600 nm was adjusted to 0.1 using 0.85% sterile NaCl solution to obtain an inoculum concentration of 1.5 × 10⁸ CFU/mL.

L. leucocephala leaf tips were collected, washed, and air-dried. Fermentation was performed using L. plantarum TISTR1284 at an inoculum level of 1 × 10⁶ CFU/g plant material following a modified method described previously [21]. The plant material was packed in airtight containers and incubated under anaerobic conditions at room temperature (25°C–30°C) for 30 days. After fermentation, samples were oven-dried and ground into a fine powder for diet formulation. Amino acid composition of fermented and non-fermented samples was analyzed by Central Laboratory (Thailand) Co., Ltd., Bangkok, Thailand, using high-performance liquid chromatography, and the results were expressed as mg/100 g dry weight (Table 1).

Antioxidant and phytochemical activities were analyzed as shown in Table 2. Fermentation increased most essential and non-essential amino acids while reducing mimosine content, indicating improved nutritional quality of L. leucocephala leaves (Table 1).

Three diets were prepared for this study. The control diet was formulated using fishmeal as the primary protein source, whereas the two experimental diets incorporated fermented L. leucocephala powder to replace 10% or 20% of fishmeal, designated as the 10% L. leucocephala and 20% L. leucocephala diets, respectively. The dry ingredients were homogenized and subsequently combined with 30% water to form a uniform mash. This mixture was processed into pellets using an extruder (EXT15HP3V03; Siam Farm Services Co., Ltd., Bangkok, Thailand) fitted with a 2-mm die. The pellets were dried at 4°C under forced-air circulation until the moisture content was reduced to approximately 10%. Once dried, all diets were sealed and stored at −20°C until further use.

The ingredient composition of the control and experimental diets is summarized in Table 3. The experimental diets were formulated to evaluate the practical replacement of fishmeal with fermented L. leucocephala powder and were not designed to be strictly isonitrogenous or isolipidic.

Following diet preparation, pellet samples were analyzed for amino acid and fatty acid profiles by Central Laboratory (Thailand) Co., Ltd., and the results are presented in Tables 4 and 5.

Proximate composition analysis of the experimental diets, including crude protein, crude lipid, ash, moisture, crude fiber, and carbohydrate contents, was conducted according to standard procedures described by the Association of Official Analytical Chemists [22]. Crude protein was determined using the Kjeldahl method, crude lipid by Soxhlet extraction, moisture by oven drying, and ash by combustion in a muffle furnace. Carbohydrate content was calculated by difference using the following formula:

% Carbohydrate = 100 − (% Moisture + % Fat + % Ash + % Crude fiber + % Protein)

The results of proximate composition analysis are shown in Table 3.

TPC of fermented and non-fermented L. leucocephala leaf powder and experimental diets was analyzed using the Folin–Ciocalteu colorimetric method with modifications [23]. Briefly, 100 µL of each extract was mixed with 125 µL of Folin–Ciocalteu reagent, followed by the addition of 300 µL of 20% sodium carbonate solution. The reaction mixture was adjusted to a final volume of 1 mL with double-distilled water and incubated at 25–30°C in the dark for 2 h. Absorbance was measured at 760 nm using a UV–Vis spectrophotometer. Gallic acid was used as the calibration standard, and the results were expressed as mg GAE/g extract.

TFC of fermented and non-fermented L. leucocephala leaf powder and experimental diets was determined using the aluminum chloride colorimetric method [23]. Briefly, 250 µL of extract was mixed with 75 µL of 5% sodium nitrite solution and incubated for 6 min. Then, 150 µL of 10% aluminum chloride solution was added, and the mixture was incubated for 5 min. Subsequently, 500 µL of 1 M sodium hydroxide was added, and the volume was adjusted to 2.5 mL with distilled water. Absorbance was measured at 510 nm using a UV–Vis spectrophotometer. Catechin was used as the reference standard, and the results were expressed as mg CE/g extract.

DPPH radical scavenging activity: Fermented and non-fermented L. leucocephala leaf powder and experimental diets were evaluated using the DPPH radical scavenging assay [23]. DPPH solution was prepared in ethanol and protected from light before use. The extracts were diluted in ethanol at different concentrations (0–20 mg/mL). Twenty microliters of sample solution was combined with 180 µL of DPPH solution in a 96-well microplate, then incubated in the dark at 37°C for 30 min. Absorbance was read at 540 nm using a microplate reader. Ascorbic acid and Trolox were used as positive controls, whereas DPPH solution without sample was used as the negative control. Radical scavenging activity was expressed as percentage inhibition, and IC₅₀ values were obtained by nonlinear regression analysis using GraphPad Prism version 10.5.0.774 (GraphPad Software, Boston, MA, USA).

ABTS radical scavenging activity: ABTS radical scavenging activity of fermented and non-fermented L. leucocephala leaf powder and experimental diets was determined using a previously reported method with modifications [23]. Briefly, 20 µL of extract at various concentrations (0–20 mg/mL) was mixed with 2.0 mL of diluted ABTS working solution. The ABTS radical cation was generated by mixing 7.5 mM ABTS stock solution with 2.5 mM potassium persulfate in the dark at 25°C–30°C for 16 h. The reaction mixtures were incubated for 5 min at 25°C, and absorbance was measured at 734 nm using a UV–Vis spectrophotometer (Optizen POP, Mecasys Co., Ltd., Daejeon, Korea). Percentage inhibition and IC₅₀ values were calculated using dose-response curves generated with GraphPad Prism software.

The reducing ability of fermented and non-fermented L. leucocephala leaf powder and experimental diets was determined using the FRAP assay [24]. The assay is based on the reduction of ferric ions to ferrous ions, forming a blue Fe²⁺/TPTZ complex. The extract was mixed with freshly prepared FRAP reagent and incubated at 25°C–30°C for 30 min. Absorbance was recorded at 593 nm using a UV–Vis spectrophotometer. Antioxidant activity was expressed as mg FeSO₄/g extract.

To evaluate whether replacing fishmeal protein with fermented L. leucocephala leaves affects the immune system of prawns, adult prawns were selected because they possess a fully developed immune system, allowing consistent and reliable evaluation of diet-induced immune responses. For the experimental challenge and immune assessment, prawns were acclimatized for 1 week, during which they were fed the control diet three times daily at 07:00, 13:00, and 19:00 h. They were then randomly divided into three groups (n = 70 per group) and maintained in plastic tanks (44 × 62 × 34 cm) containing 45 L of aerated water, with 15 prawns per tank.

Sample size determination was performed using G*Power software version 3.1.9.7 prior to experimentation to ensure adequate statistical power to detect differences among treatment groups. Group 1 received the basic diet and served as the control group. Group 2 received the basic diet with 10% fishmeal replaced by fermented L. leucocephala. Group 3 received the basic diet with 20% fishmeal replaced by fermented L. leucocephala. Prawns were fed three times daily at 07:00, 13:00, and 19:00 h for 2 weeks.

After the 2-week feeding period, prawns were injected with A. hydrophila at a dose corresponding to the previously reported median lethal dose for M. rosenbergii (8.91 × 10⁵ CFU/mL) [20] and maintained under the same feeding regimen and environmental conditions for 1 week. At each sampling time point (6, 12, 24, 48, 72, 96, and 120 h post-infection), five prawns were randomly selected from each treatment group for collection of hemolymph, hematopoietic tissue, and intestinal samples for immunological analyses. This experiment was conducted using a single replicate per treatment group. For the survival experiment, prawns were divided into three groups as described above, with n = 15 per group. This experiment was conducted using three independent replicate tanks per treatment group to observe and record mortality.

Water quality parameters were monitored throughout the experimental period. Water temperature was maintained at 27°C–29°C, DO above 5 mg/L, and pH at 7.5–8.0. Total ammonia and nitrite concentrations were maintained below 0.1 mg/L and 0.5 mg/L, respectively, which are within acceptable ranges for M. rosenbergii culture [25]. To maintain water quality, approximately 30%–50% of the water was replaced every 3 days, and the filtration system was operated continuously throughout the experiment.

Hemolymph (500 µL) was withdrawn from the heart of M. rosenbergii into a 1-mL syringe containing cold Alsever’s solution and transferred to a 1.5-mL microcentrifuge tube. A drop of mixed hemolymph was placed on a hemocytometer, and total hemocyte count was determined under a light microscope (Olympus CX22; Olympus Corporation, Tokyo, Japan). Hemocyte counting was performed by an investigator blinded to the treatment groups.

PO activity in hemolymph was measured according to a previously described protocol with modifications [26]. Briefly, 100 µL of hemolymph was withdrawn from the heart into a 1-mL syringe containing 900 µL of cold Tris-buffered saline-I (50 mM Tris, 210 mM NaCl, 5 mM KCl, and 2.5 mM MgCl₂, pH 7.5). The hemolymph was centrifuged at 5000 × g for 15 min at 4°C to separate hemocytes, and the resulting hemolymph supernatant was collected for PO activity analysis. Ten microliters of hemolymph were incubated with 190 µL of 5 mM L-DOPA (D9628; Sigma-Aldrich, St. Louis, MO, USA) dissolved in 50 mM Tris-HCl, pH 7.5, for 20 min at 25°C to develop dopachrome. The dopachrome reaction was measured at 490 nm using a VersaMax™ Microplate Reader (Molecular Devices, San Jose, CA, USA).

Hematopoietic tissue was collected, fixed in Davison’s fixative for 24 h, and then washed with 70% ethanol. The tissue was processed using an automatic sample preparation system (Tissue-Tek® VIP™ 5 Jr.; Sakura Finetek Japan Co., Ltd., Tokyo, Japan) and embedded in paraffin. Paraffin blocks were sectioned at 5 µm, and the sections were placed on glass slides. Tissue sections were stained with Mayer’s hematoxylin (Cat. No. 05-06002/L; Bio-Optica Milano SpA, Milan, Italy) and eosin Y plus alcoholic solution (Cat. No. 05-11007/L; Bio-Optica Milano SpA). Hematopoietic tissue sections were examined under a light microscope (Nikon Upright Microscope Eclipse Ni-U; Nikon Corporation, Tokyo, Japan) at 40× magnification. Prophase, metaphase, anaphase, and telophase in hematopoietic cell nuclei were counted and averaged. Mitotic cell quantification was performed by an investigator blinded to the treatment groups.

Total RNA was isolated from various tissues of M. rosenbergii using Tri Reagent® (Cat. No. TR118; Molecular Research Center, Inc., Cincinnati, OH, USA) according to the manufacturer’s protocol. RNA quality and quantity were assessed by measuring absorbance at 260 and 280 nm using NanoDrop One (Thermo Fisher Scientific, Waltham, MA, USA). One microgram of RNA was subjected to cDNA synthesis using ReverTra Ace™ quantitative (q)PCR RT Master Mix with gDNA Remover (TOYOBO Co., Ltd., Osaka, Japan) according to the manufacturer’s protocol. The cDNA was stored at −20°C until further analysis.

The cDNA samples were analyzed for relative expression levels of the genes anti-lipopolysaccharide factor (ALF), Crustacean hematopoietic factor (CHF), proPO, Relish, TRAF6, Dorsal, IMD, HSP70, and Cu/Zn-SOD in hemocytes and hematopoietic tissue using QIAquant 96 5plex (Qiagen, Hilden, Germany). Amplification was performed in a 96-well plate with a 20-µL fluorescent qPCR reaction mixture. The reaction mixture contained 1 µL of cDNA, 10 µL of SensiFAST™ SYBR® No-ROX kit (BIO-98050; Meridian Bioscience, Cincinnati, OH, USA), 0.4 µL each of forward and reverse primers (10 µM/µL), and 8.2 µL of sterile dH₂O. All qRT-PCR primers are listed in Table 6[27–32].

The thermal qRT-PCR cycle profile followed the manufacturer’s protocol: one cycle of enzyme activation at 95°C for 2 min, followed by 40 cycles of denaturation at 95°C for 5 s and annealing/extension at 60°C for 20 s. Sterile dH₂O served as the negative control. EF-1α was selected as the reference gene for normalization. After completion of the qRT-PCR program, data were analyzed using QIAquant96 software. The baseline was set automatically by the software to maintain consistency. The comparative CT (2−ΔΔCT) method was used to analyze gene expression levels [33].

Data are presented as mean ± standard deviation. Statistical analyses were performed using GraphPad Prism software version 10.5.0.774 (GraphPad Software, Boston, MA, USA). Before statistical analysis, data normality was assessed using the Shapiro–Wilk test, and homogeneity of variance was evaluated using the Brown–Forsythe test. Data that satisfied parametric assumptions were analyzed using one-way analysis of variance, followed by Tukey’s multiple comparisons test. When assumptions of normality or homogeneity of variance were not met, non-parametric analysis was performed using the Kruskal–Wallis test followed by Dunn’s multiple comparison test. Survival data were analyzed using the Kaplan–Meier method. Differences were considered statistically significant at p < 0.05.

Fermentation of L. leucocephala significantly altered the amino acid profile. Most essential amino acids increased after fermentation, including lysine (+24.6%), leucine (+23.5%), phenylalanine (+28.7%), and valine (+16.3%). In contrast, the toxic non-protein amino acid mimosine was markedly reduced by 42.2%, indicating detoxification during fermentation (Table 1). Overall, the total essential amino acid content was higher in the fermented material than in the non-fermented material.

The inclusion of fermented L. leucocephala meal gradually altered the amino acid composition of the diets in a dose-dependent manner (Table 4). Most essential amino acids, including lysine, methionine, leucine, and threonine, showed a decreasing trend as the inclusion level increased from 0% to 20%. Lysine content decreased from 2711.91 mg/100 g in the control diet to 2132.84 mg/100 g in the 20% replacement group. A similar pattern was observed for methionine and threonine. Non-essential amino acids also showed a general decreasing trend with increasing inclusion levels, particularly glutamic acid and aspartic acid. Overall, dietary amino acid profiles were moderately affected by higher inclusion levels of fermented leaf meal.

Antioxidant and phytochemical properties of experimental diets supplemented with fermented L. leucocephala

The antioxidant activity and phytochemical contents of diets supplemented with fermented L. leucocephala are shown in Table 7. Antioxidant capacity and phytochemical content increased dose-dependently with increasing levels of fermented L. leucocephala supplementation. The 20% fermented L. leucocephala diet showed the highest antioxidant activity in both DPPH and ABTS radical scavenging assays, as evidenced by the lowest IC₅₀ values (1612.57 ± 19.87 and 305.38 ± 17.19 µg/mL, respectively). In contrast, the control diet showed the lowest activity. Likewise, FRAP values were significantly higher with increasing supplementation levels, ranging from 6.02 ± 0.50 mg FeSO₄/g extract in the control diet to 27.78 ± 2.23 mg FeSO₄/g extract in the 20% supplementation group. Phytochemical analysis also showed higher TPC and TFC after supplementation with fermented L. leucocephala. The highest TPC (11.62 ± 0.50 mg GAE/g extract) and TFC (1.364 ± 0.4 mg CE/g extract) values were observed in the 20% supplementation group compared with the control diet. Overall, the results showed that fermented L. leucocephala improved the antioxidant activity and phytochemical profiles of the experimental diets, particularly at the 20% supplementation level.

Hemocyte proliferation: The groups of M. rosenbergii fed diets supplemented with 10% and 20% fermented L. leucocephala had more circulating hemocytes than the control group and showed a statistically significant increase in circulating hemocytes during 6–72 h after infection with A. hydrophila (Figure 1A). At 96 h, only the 10% L. leucocephala group (3.22 × 10⁶ ± 1.94 × 10⁵ cells/mL) had significantly higher hemocyte circulation than the control (2.58 × 10⁶ ± 8.7 × 10⁴ cells/mL) and 20% L. leucocephala groups (2.54 × 10⁶ ± 1.77 × 10⁵ cells/mL) (p < 0.05, Figure 1A).

Hematopoietic cell division was significantly greater in the 20% L. leucocephala group than in the control group at 0 h (p < 0.001), 12–24 h (p < 0.05), and 72–120 h (p < 0.01) post-infection (Figure 1B). Although the 10% L. leucocephala group showed a trend toward a higher number of dividing cells, the difference was not statistically significant compared with the 20% L. leucocephala group and the control group (Figure 1B). Hematopoietic cells undergoing cell division are shown in Figure 1C and Supplementary Figure S1. Dividing hematopoietic cells were identified by nuclear membrane disappearance and chromosome condensation during prophase (Figure 1C, red arrow), metaphase (Figure 1C, white arrow), anaphase (Figure 1C, black arrow), and telophase (Figure 1C, blue arrow).

The expression of CHF in hemocytes at all time points was greater in the 10% and 20% L. leucocephala groups than in the control group (Figure 1D). However, a significant increase in CHF expression was observed in the 20% L. leucocephala group at 12 h (2.92 ± 1.00, p < 0.05), 24 h (7.90 ± 1.00, p < 0.001), 72 h (2.86 ± 0.36, p < 0.05), and 96 h (1.72 ± 0.37, p < 0.05) post-infection compared with the control group. In the 10% L. leucocephala group, CHF expression (5.83 ± 0.56) was significantly increased at 24 h (p < 0.01) compared with the control group (1.00 ± 0.15). However, no statistically significant difference in hemocyte CHF expression was observed between the 10% and 20% L. leucocephala groups.

The expression of CHF in the hematopoietic tissue of the 10% L. leucocephala group was significantly greater at 0 h (3.05 ± 0.60, p < 0.05), 24 h (42.24 ± 6.15, p < 0.01), and 72 h (3.46 ± 1.10, p < 0.05) post-infection compared with the control group (Figure 1E). The expression of CHF in the 20% L. leucocephala group tended to be higher than that in the control group at 0–120 h post-infection, with significant increases observed only at 12–24 h and 72 h (p < 0.05) (Figure 1E). However, CHF expression in the 20% L. leucocephala group was consistently lower than that in the 10% L. leucocephala group. PO activity: PO activity in the hemolymph of the three treatment groups of M. rosenbergii during A. hydrophila infection is shown in Figure 2A. Although M. rosenbergii receiving 10% and 20% L. leucocephala showed a trend toward increased PO activity during 0–96 h post-infection, significantly higher PO activity than that in the control group was observed only at 48 h (p < 0.01).

In addition, mRNA expression of proPO in hemocytes significantly increased during 0–72 h post-infection (Figure 2B). The expression of proPO in the 20% L. leucocephala group was significantly higher than that in the control group at 0–6 h (p < 0.01) and 24–72 h (p < 0.05) post-infection. In the 10% L. leucocephala group, proPO expression was elevated at 0 h but did not differ significantly from that in the control group; however, it became significantly higher at 12 h post-infection compared with the control group (p < 0.05) (Figure 2B).

The mRNA expression levels of TRAF6 and Dorsal in hemocytes were significantly lower in the 10% and 20% L. leucocephala groups than in the control group during 6–48 h post-infection (Figure 3A and B). At 72 h, TRAF6 expression remained downregulated in both treatment groups; however, the differences were no longer statistically significant compared with the control group. TRAF6 expression subsequently returned to levels comparable to those of the control group at 96–120 h post-infection (Figure 3A). Similarly, Dorsal mRNA expression was downregulated in the 10% and 20% L. leucocephala groups during 6–72 h and then returned to near control levels at 96–120 h post-infection (Figure 3B).

In hematopoietic tissue, TRAF6 expression did not differ significantly among the three groups throughout the experimental period (Figure 3C). In contrast, Dorsal expression in hematopoietic tissue was significantly decreased at 6 h in the 10% L. leucocephala group (0.21 ± 0.04) compared with the 20% L. leucocephala group (0.91 ± 0.19) and the control group (1.04 ± 0.34) (p < 0.01, Figure 3D). Although a general trend toward reduced Dorsal expression was observed at other time points, significant reductions were detected only in the 20% L. leucocephala group at 12 h (0.60 ± 0.08) and 72 h (0.64 ± 0.12), and in the 10% L. leucocephala group at 96 h (0.46 ± 0.10), compared with the control group (1.09 ± 0.49) (Figure 3D).

Overall, IMD expression was elevated in the 10% and 20% L. leucocephala groups, although expression patterns varied across time points (Figure 4A). At 0 h, IMD expression in hemocytes did not differ significantly between the 10% L. leucocephala group and the control group. However, the 20% L. leucocephala group (7.25 ± 1.74) showed significantly higher IMD expression than both the 10% L. leucocephala group (1.46 ± 0.29) (p < 0.05) and the control group (0.99 ± 0.6) (p < 0.01) (Figure 4A). During A. hydrophila infection, IMD expression remained upregulated in the 10% L. leucocephala group, although the increase was not statistically significant compared with the control group. In contrast, the 20% L. leucocephala group (2.20 ± 0.01) exhibited significantly higher IMD expression at 6 h post-infection than the control group (1.14 ± 0.63) (p < 0.05) (Figure 4A). IMD expression further increased in both treatment groups during 12–24 h post-infection, with the highest expression observed at 12 h (Figure 4A). Thereafter, although IMD expression remained higher than that in the control group, statistically significant differences were observed only at 72 h in both the 10% and 20% L. leucocephala groups (Figure 4A).

Relish expression was significantly increased in the 20% L. leucocephala group at 0 h (1.40 ± 0.08) compared with the control group (1.01 ± 0.17) (p < 0.05) (Figure 4B). At 6 h after bacterial challenge, Relish expression was significantly elevated in both the 10% (11.64 ± 5.49) and 20% L. leucocephala groups (12.69 ± 4.93) compared with the control group (1.13 ± 0.67) (p < 0.05) (Figure 4B). At 12 h, significantly higher Relish expression was observed only in the 20% L. leucocephala group (1.77 ± 0.10) compared with the 10% L. leucocephala group (1.23 ± 0.11) and the control group (1.04 ± 0.35) (p < 0.05) (Figure 4B). At 24 h, Relish expression was significantly upregulated in the 10% L. leucocephala group compared with the control group (p < 0.05) (Figure 4B). Although slight increases in Relish expression were observed in both treatment groups during 48–96 h post-infection, the differences were not statistically significant. At 120 h, Relish expression was significantly higher in the 20% L. leucocephala group (3.19 ± 1.04) than in the control group (1.00 ± 0.17) (p < 0.05) (Figure 4B).

HSP70 expression was significantly higher in the 20% L. leucocephala group than in the 10% L. leucocephala and control groups throughout the 0–120 h post-infection period (p < 0.001) (Figure 4C). HSP70 expression was highest at 0 h and gradually decreased until 48 h post-infection. Thereafter, expression gradually increased during 72–96 h, and by the end of the experimental period, no significant differences in HSP70 expression were observed among the groups.

Cu/Zn-SOD mRNA expression was significantly upregulated in the 20% L. leucocephala group compared with the 10% L. leucocephala and control groups during 0–12 h and at 120 h post-infection (p < 0.001) (Figure 4D). At 24 h post-infection, Cu/Zn-SOD expression was markedly elevated in both the 10% (50.44 ± 29.87) and 20% L. leucocephala groups (40.48 ± 26.46) compared with the control group (0.92 ± 0.17), although no significant difference was observed between the two treatment groups. During 72–120 h post-infection, both L. leucocephala-supplemented groups showed a trend toward increased Cu/Zn-SOD expression relative to the control group.

In addition, the marked upregulation of Cu/Zn-SOD expression in the 20% L. leucocephala group was associated with significantly lower ABTS and DPPH IC₅₀ values than in the control group, indicating enhanced radical-scavenging activity. This group also exhibited higher FRAP values and increased TPC and TFC compared with the control group (Table 7). These findings suggest that elevated antioxidant gene expression corresponded with improved antioxidant capacity and phytochemical content.

ALF mRNA expression was generally upregulated in the 10% and 20% L. leucocephala groups compared with the control group throughout the 0–120 h post-infection period (Figure 4E). The 20% L. leucocephala group exhibited significantly higher ALF expression than both the 10% L. leucocephala and control groups at 0 h (p < 0.01), 6 h (p < 0.01), 24 h (p < 0.05 and p < 0.01, respectively), and 120 h post-infection (p < 0.01) (Figure 4E). In contrast, ALF expression in the 10% L. leucocephala group (2.55 ± 0.32) was significantly higher than that in the control group (1.00 ± 0.12) only at 24 h post-infection (p < 0.05) (Figure 4E). In contrast to ALF, C-lectin expression was generally lower in the 10% and 20% L. leucocephala groups than in the control group at all time points. However, significant reductions were observed only during 6–12 h and 72–96 h post-infection (Figure 4F).

The histology of M. rosenbergii intestines showed normal structures in all groups (Figure 5A and Supplementary Figure S2). Alterations in epithelial cell height (Figure 5B), microvilli height (Figure 5C), and muscular wall thickness (Figure 5D) were not observed.

Overall, IMD expression was higher in the 20% L. leucocephala group than in the control and 10% L. leucocephala groups throughout the experimental period (Figure 6A). Compared with the control group, IMD expression in the 20% L. leucocephala group was significantly increased at 6 h (p < 0.05), 24 h (p < 0.001), and 48 and 96 h post-infection (p < 0.01), and at 120 h post-infection (p < 0.01). However, IMD expression in the 20% L. leucocephala group did not differ significantly from that in the 10% L. leucocephala group. In the 10% L. leucocephala group, IMD expression was significantly higher than that in the control group at 48 h (p < 0.05) and 120 h post-infection (p < 0.01) (Figure 6A).

Relish expression (Figure 6B) was generally higher in the 20% L. leucocephala group than in the control and 10% L. leucocephala groups throughout the experimental period. Specifically, Relish expression in the 20% L. leucocephala group was significantly elevated (p < 0.01) at 0 h (2.34 ± 0.34) compared with the control (1.04 ± 0.32) and 10% L. leucocephala groups (1.02 ± 0.35). Significant increases were also observed at 6 h (2.37 ± 0.77) compared with the control group (1.01 ± 0.17) and 10% L. leucocephala group (1.28 ± 0.21), at 48 h (2.29 ± 0.64) compared with both the control (1.01 ± 0.19) and 10% L. leucocephala groups (1.28 ± 0.21) (p < 0.01 and p < 0.05, respectively), and at 120 h (2.74 ± 0.50) compared with the control group (1.06 ± 0.43) and 10% L. leucocephala group (1.28 ± 0.62) (p < 0.01) post-infection. Although Relish expression in the 10% L. leucocephala group tended to be higher than that in the control group, significant differences were observed only at 24 h (2.25 ± 0.66) and 72 h (1.42 ± 0.25) post-infection (p < 0.05) (Figure 6B).

HSP70 expression (Figure 6C) was generally higher in the 20% L. leucocephala group than in the control and 10% L. leucocephala groups throughout the experimental period. At 0 h, HSP70 expression in the 20% L. leucocephala group (3.62 ± 0.84) was significantly higher than that in the control (1.03 ± 0.31) and 10% L. leucocephala groups (1.46 ± 0.67) (p < 0.001 and p < 0.01, respectively). At 6 h post-infection, HSP70 expression in the 20% L. leucocephala group (2.59 ± 0.98) was significantly higher than that in the control group (1.00 ± 0.14) (p < 0.05). Markedly elevated expression was also observed at 12 h (53.11 ± 20.19) and 24 h (16.77 ± 6.19) (p < 0.001), as well as at 96 h (7.33 ± 5.00) and 120 h (6.97 ± 1.03) post-infection compared with the control group (p < 0.001).

In the 10% L. leucocephala group, HSP70 expression tended to be higher than that in the control group, although significant differences were observed only at 6 h (4.36 ± 0.85) and 120 h (5.83 ± 1.73) post-infection (p < 0.001). Notably, at 6 h post-infection, HSP70 expression was higher in the 10% L. leucocephala group than in the 20% L. leucocephala group.

Cu/Zn-SOD expression (Figure 6D) was generally higher in the 20% L. leucocephala group than in the control and 10% L. leucocephala groups throughout the experimental period. Significant upregulation in the 20% L. leucocephala group was observed at 0 h (7.02 ± 2.93) compared with the control group (1.07 ± 0.49) (p < 0.001) and the 10% L. leucocephala group (3.04 ± 0.30) (p < 0.01). At 6 h post-infection, Cu/Zn-SOD expression increased further to 15.32 ± 2.38 and remained significantly higher than in the control group (p < 0.001). Significant increases were also observed at 24 h (3.25 ± 1.33) compared with both the control and 10% L. leucocephala groups (p < 0.01 and p < 0.05, respectively), and at 48 h (3.79 ± 2.28) compared with the control group (1.06 ± 0.40) (p < 0.05).

In the 10% L. leucocephala group, Cu/Zn-SOD expression also tended to be higher than that in the control group, although the increase was less pronounced than that observed in the 20% L. leucocephala group. Significant differences between the 10% L. leucocephala and control groups were detected only at 6, 24, and 96 h post-infection (p < 0.01).

ALF expression (Figure 6E) was generally higher in the 10% and 20% L. leucocephala groups than in the control group. Significant upregulation was observed at 0, 12, 48, 72, and 120 h post-infection in both treatment groups compared with the control group, although no significant differences were detected between the 10% and 20% L. leucocephala groups at these time points. At 6 h post-infection, ALF expression in the 20% L. leucocephala group (18.44 ± 16.15) also showed an increasing trend; however, the difference was not statistically significant compared with the control group. In contrast, the 10% L. leucocephala group (23.01 ± 4.31) exhibited significantly higher ALF expression than the control group (1.89 ± 1.86) at this time point.

C-lectin expression (Figure 6F) at 0 h was significantly lower in the 10% L. leucocephala (0.16 ± 0.07) and 20% L. leucocephala groups (0.24 ± 0.17) than in the control group (1.04 ± 0.33). However, at 12 h and 48 h post-infection, C-lectin expression was markedly elevated in the 20% L. leucocephala group, reaching 52.56 ± 10.57 and 11.24 ± 3.54, respectively, which were significantly higher than those in the control group (1.03 ± 0.26 and 1.04 ± 0.37, respectively) and the 10% L. leucocephala group (7.60 ± 2.58 and 1.81 ± 1.58, respectively) (p < 0.001).

C-lectin expression in the 10% L. leucocephala group also showed an increasing trend during the 12–96 h post-infection period, similar to that observed in the 20% L. leucocephala group. At 72 h, expression in the 10% L. leucocephala group (30.26 ± 6.47) was significantly higher than that in the control group (1.01 ± 0.20) and the 20% L. leucocephala group (6.27 ± 2.26) (p < 0.001). After 72 h post-infection, C-lectin expression in the 20% L. leucocephala group gradually decreased, resulting in a significant difference compared with the 10% L. leucocephala group (p < 0.05).

The effect of 10% and 20% fermented L. leucocephala in fishmeal replacement on the survival of adult M. rosenbergii during A. hydrophila infection is shown in Figure 7. Survival of M. rosenbergii tended to be higher in the 10% and 20% L. leucocephala groups than in the control group; however, the differences were not statistically significant (p = 0.167 and p = 0.204, respectively). No dose-dependent effect was observed.

L. leucocephala leaves are widely recognized as a rich source of protein and essential minerals, making them a promising alternative feed ingredient for livestock and aquaculture species in tropical regions. Their high protein content is a key nutritional advantage, along with the presence of 17 flavonoids known for biological activities, including antioxidant and anti-inflammatory effects [14]. In the present study, L. plantarum TISTR1284 was used to improve the bioavailability and functional properties of L. leucocephala leaves. Fermentation is known to enhance food safety and nutritional quality by producing organic acids, including lactic, acetic, propionic, and formic acids, which help eliminate harmful bacteria and toxins while preserving sensory qualities [17]. However, the present experimental design does not allow separation of the individual contributions of fermentation, plant substitution, and probiotic effects; therefore, the outcomes observed here should be interpreted as the combined effect of the fermented ingredient rather than being attributed to a single factor.

A. hydrophila, a pathogenic Gram-negative bacterium, poses a serious threat to Macrobrachium nipponense and M. rosenbergii [34, 35]. Plant extracts from Withania somnifera and Melaleuca cajuputi have been shown to enhance immune responses and increase shrimp survival during A. hydrophila infection [36, 37]. L. leucocephala seed protein has also been shown to enhance bacterial disease resistance in the catfish Clarias gariepinus [12]. In the present study, dietary supplementation with fermented L. leucocephala leaves enhanced immune parameters and improved survival of M. rosenbergii following A. hydrophila challenge.

Crustacean resistance to pathogens relies primarily on the innate immune system. Hemocytes are crucial cells that mediate physiological responses to infection and environmental stress. Probiotics and medicinal plant extracts have been applied to enhance immune parameters in M. rosenbergii [38, 39]. In the present study, increased circulating hemocytes and a higher number of mitotic cells in hematopoietic tissue were observed when fermented L. leucocephala leaves were added to the diet of M. rosenbergii. This response may contribute to accelerated maturation of hemocyte precursors in hematopoietic tissue, thereby maintaining hemocyte population and functionality during bacterial challenge [40].

L. leucocephala contains apigenin, kaempferol, and juglanin, which induce the transcriptional activity of pHRE-Luc before promoting erythropoietin expression and stimulating red blood cell production [14]. Upregulation of CHF is a biological indicator associated with hemocyte and hematopoietic cell proliferation in M. rosenbergii [27]. The increase in hematopoietic proliferation, reflected by CHF expression in hemocytes and hematopoietic tissue of the 10% and 20% L. leucocephala groups, suggests that fermented L. leucocephala promotes hematopoietic activity and hemocyte homeostasis during bacterial infection.

PO is an important enzyme that promotes humoral defense mechanisms and the elimination of pathogens in invertebrates [41]. Plant extracts have been shown to induce PO activity and improve immunocompetence against pathogens in crustaceans [38, 42, 43]. In the present study, M. rosenbergii fed fermented L. leucocephala showed increased PO activity in hemolymph and significant upregulation of proPO. Antimicrobial and antioxidative activities induced by phytochemicals in L. leucocephala have been previously reported [44]. These findings suggest that key bioactive properties of L. leucocephala were retained after fermentation, although the specific compounds responsible for the observed effects were not directly quantified in the present study.

TRAF6 is a highly expressed gene that acts as a crucial signal transducer triggering downstream cascades mediated by TNF receptors and the IL-1 receptor/Toll-like receptor superfamily during immune responses [45]. In Fenneropenaeus penicillatus, higher TRAF6 expression was correlated with peroxinectin expression after infection by white spot syndrome virus and Vibrio alginolyticus [46]. TRAF6-like transcription was upregulated after Staphylococcus aureus and Edwardsiella ictaluri challenge in Procambarus clarkii [47]. Knockdown of TRAF6-like inhibited downstream effector genes, including Dorsal [47]. TRAF6 regulates immune responses during A. hydrophila infection because mannose-binding lectin and crustin are decreased after TRAF6 silencing [30].

In the present study, TRAF6 expression decreased within 48 h, and Dorsal expression declined at specific time points in the 10% and 20% groups. These findings indicate modulation of inflammatory signaling; however, bacterial burden or clearance was not quantified in this study. Therefore, improved survival should not be directly interpreted as increased bacterial clearance. Flavonoids in L. leucocephala have been reported to reduce pro-inflammatory cytokine secretion in RAW 264.7 macrophages [14], which may partly explain the moderated inflammatory profile observed here.

Two major pathways are involved in shrimp immunity: the Toll pathway and the IMD pathway. These pathways share downstream signaling components, and crosstalk between them contributes to protection against bacterial infection [48]. In the present study, C-lectin was downregulated, whereas IMD-related genes (IMD and Relish) and effector/stress genes (HSP70, Cu/Zn-SOD, and ALF) were upregulated, particularly in the 20% group. This pattern suggests coordinated immune modulation rather than simple immune suppression. Such modulation may reflect regulated immune adjustment to dietary bioactive compounds, as previously observed in crustaceans exposed to functional feed additives and plant-derived extracts [49, 50].

However, the precise molecular mechanism underlying the activation of the immune pathway by fermented L. leucocephala remains unclear and warrants further mechanistic investigation. In particular, this study did not identify the specific bioactive compounds responsible for modulating the IMD pathway, although phytochemicals such as phenolics have been reported to influence immune responses in C. gariepinus [51]. Nevertheless, the exact molecular mediators and their direct interactions with IMD pathway components remain unclear and require targeted metabolomic and functional studies.

In addition to immunomodulation, L. leucocephala has been associated with gut health and nutrient utilization [52]. Replacing 25%–50% of fishmeal with L. leucocephala leaves in Nile tilapia diets did not adversely affect fish health, although some growth retardation was observed [11]. Improvements in digestibility and nutrient uptake have also been reported when L. leucocephala leaves are fermented with beneficial gut bacteria. For example, Bacillus subtilis and Bacillus circulans enhanced nutritional value and growth performance in Labeo rohita when included in diets containing 30%–40% L. leucocephala [53]. In crustaceans, host-associated probiotics such as Lactococcus lactis have shown similar benefits in M. rosenbergii, improving both growth and immune function [39].

Although viable LAB counts in the final pelleted feed and during storage were not quantified, the present findings suggest that the beneficial effects observed in M. rosenbergii were likely mediated not only by live probiotic cells but also by fermentation-derived paraprobiotics and postbiotics acting synergistically with enhanced plant bioactive compounds. Paraprobiotics, including inactivated microbial cells and cell-wall components such as peptidoglycans, teichoic acids, and surface proteins, are known to interact with crustacean pattern-recognition receptors, thereby stimulating hemocyte proliferation and activating downstream immune pathways, including the IMD–Relish signaling cascade observed in the present study. Concurrently, fermentation by L. plantarum may generate beneficial postbiotic metabolites, including organic acids, bacteriocins, and bioactive intracellular compounds, which may further contribute to immune modulation and disease resistance.

The present findings suggest that L. plantarum fermentation may help mitigate one of the major limitations associated with plant protein utilization, namely ANFs. As shown in Table 1, fermentation reduced mimosine levels by 42.18%, decreasing from 92.46 to 53.46 mg/100 g. Mimosine is a toxic non-protein amino acid known to impair nutrient utilization and physiological performance in animals. Furthermore, the incorporation of fermented L. leucocephala enhanced the phytochemical and antioxidant properties of the diets, as evidenced by increased TPC and TFC, higher FRAP values, and improved radical-scavenging activity, as indicated by lower DPPH and ABTS IC₅₀ values in the 20% inclusion group. Collectively, these findings suggest that fermented diets may function as promising functional feeds enriched with detoxified plant matrixes, fermentation-derived metabolites, and microbial-derived bioactive components that collectively support innate immune responses in prawns.

One limitation of the present study is that the experimental diets were not strictly formulated to be isonitrogenous. As shown in Table 2, crude protein content decreased from 44.63% in the control diet to 36.75% in the 20% replacement group, representing a substantial nutritional variation that may have influenced the physiological and immunological responses of M. rosenbergii. Therefore, the effects cannot be attributed exclusively to the bioactive properties of fermented L. leucocephala, as differences in dietary protein levels may also have contributed to the observed responses. In addition, crystalline amino acids such as lysine and methionine were not supplemented to achieve precise amino acid balancing among the experimental diets. Consequently, variations in essential amino acid composition resulting from the inclusion of fermented L. leucocephala may have further influenced immune and physiological responses. Future studies using strictly isonitrogenous, isolipidic, and amino acid-balanced diets are required to better distinguish the specific functional effects of fermented L. leucocephala from potential nutritional confounding factors.

Another limitation is that the feeding trial duration was relatively short compared with conventional aquaculture nutrition studies, and growth performance parameters such as weight gain, specific growth rate, and feed conversion ratio were not evaluated. In addition, pellet water stability and nutrient leaching rates were not quantitatively assessed. Since feed stability in water may influence nutrient availability, feed intake, and feeding efficiency in prawns, these parameters should be evaluated in future studies to better characterize the physicochemical quality and practical applicability of the experimental diets under aquaculture conditions. Therefore, the long-term nutritional suitability and commercial applicability of the diets could not be fully determined in the present study. Furthermore, the current experimental design does not allow precise determination of the optimal inclusion level or dose-dependent immune responses associated with fermented L. leucocephala supplementation.

The present study also has limitations related to fermentation characterization and microbial analysis. Post-fermentation LAB viability, pH, lactic acid concentration, titratable acidity, and detailed biochemical characteristics of the fermented product were not directly quantified. In addition, the microbial composition of the fermented material and viability of L. plantarum in the final feed and during storage were not evaluated. Consequently, microbial dynamics and fermentation-derived metabolites associated with the fermented product could not be fully characterized. Therefore, the relative contributions of live probiotic cells, postbiotic metabolites, and paraprobiotic effects remain unclear. Moreover, microbial safety analyses, including assessments of pathogen and mycotoxin contamination, were not performed directly, although fermentation was conducted under sealed, anaerobic conditions to minimize contamination risk. Future studies integrating comprehensive fermentation profiling, microbial safety analyses, metabolite characterization, and gut microbiota analysis would provide stronger mechanistic insight into host–microbe interactions and the functional properties of fermented L. leucocephala.

Antioxidant capacity was not directly measured in hemolymph or tissues of M. rosenbergii. The present study primarily evaluated antioxidative responses through immune- and oxidative stress-related gene expression, particularly Cu/Zn-SOD expression in hemocytes. Therefore, the antioxidative effects of fermented L. leucocephala supplementation should be interpreted as indirect molecular evidence rather than direct biochemical confirmation of antioxidant activity.

Several limitations were also associated with the bacterial challenge experiment. Unchallenged control prawns were not sampled at all post-challenge time points (6–120 h), limiting the ability to distinguish the direct effects of dietary supplementation from diet × infection interaction effects. In addition, bacterial load in hemolymph or tissues was not quantified. Therefore, the improved survival and immune responses observed in the treatment groups cannot be directly interpreted as evidence of enhanced bacterial clearance capacity. Correlation analyses between immune parameters and survival outcomes were also not performed, preventing detailed evaluation of the quantitative relationship between immune modulation and disease resistance. Future studies incorporating time-matched unchallenged controls, quantification of bacterial load, and integrated multivariate statistical analyses would strengthen the mechanistic interpretation of immune responses and disease resistance in M. rosenbergii.

Despite these limitations, the consistent enhancement of several immune parameters and improved resistance to A. hydrophila observed in the treatment groups suggest that fermented L. leucocephala may contribute beneficial immunomodulatory effects in prawns. Importantly, no adverse effects on intestinal morphology were detected, supporting the safety of the dietary inclusion levels used in the present study. Nevertheless, the findings should be interpreted with caution, and further long-term studies using rigorously controlled diet formulations and comprehensive physiological, microbiological, and biochemical evaluations are warranted to clarify the specific contribution of fermented L. leucocephala to immune modulation and disease resistance in prawn aquaculture.