Abstract
The main objective of this study was to investigate the effects of commercial probiotic Bacillus sp. supplementation on seabream Sparus aurata larviculture under culture conditions. In this context, Bacillus was supplemented via rotifer feeding and water and its effects on pancreatic and intestinal enzyme activities as well as aquaculture parameters were evaluated during early life development. In the experimental group, as probiotic three Bacillus sp. spores were introduced via rotifer and larval culture tanks, while the larvae in control group did not feed any probiotic supplementation. At the end of the experiment on 40 days after hatching, the probiotic-supplemented group exhibited better growth performance and there were statistically differences in between groups of probiotic-treated and control regarding growth parameters (P < 0.01), despite insignificant survival rate (P > 0.05). In terms of enzymatic expressions, S. aurata larvae receiving probiotic supplementation through rotifers demonstrated noteworthy (P < 0.05) enhancements in specific activities of pancreatic and intestinal enzymes, except for amylase (P > 0.05), when compared to the control group. It is concluded that the administration of Bacillus sp. as probiotic bacteria through rotifer supplementation and water intake demonstrates significant positive impacts on both growth parameters and specific activities of main pancreatic and intestinal enzymes of seabream larvae.
This study is primarily focused on administration of three strains of Bacillus sp. (B. subtilis, B. licheniformis, and B. cereus) as a live food supplement in seabream, Sparus aurata, larval culture. It is revealed that supplementation of Bacillus sp. probiotics into rotifers causes marked enhancement not only in husbandry parameters such as both growth and survival rate but also in specific activities of pancreatic and intestinal enzymes in larvae of seabream.
Introduction
Over the past two decades, the identification of new aquatic pathogenic microorganisms in marine fish culture has become more difficult with increasing environmental sensitivity. The necessity of getting the products to the market as soon as possible, and the problems faced in maintaining culture conditions and nutrients have led scientists and producers to new searches due to the development of biotechnology. Especially, probiotics have been an important solution to overcome these bottlenecks. In the aquaculture sector, Probiotic bacteria have been extensively applied in many areas thorough two decades to enhance health, boost disease resistance, and improve feed efficiency in not only fish but also other aquatic organisms under aquaculture conditions. Acting primarily as biological control agents, they play a key role in promoting sustainable aquaculture practices for fish and other aquatic organisms by supporting overall health, disease prevention, and sustainability (Gatesoupe 1999, Verschuere et al. 2000, Suzer et al. 2008, Avella et al. 2010, Kuebutornye et al. 2019).
Probiotics are living microorganisms and influence the microbiota in many parts of the organism, even partially. Microorganisms found in the intestinal bacterial microbiota are basically divided into two groups: beneficial and harmful. Beneficial probiotic bacteria in the gastrointestinal tract are known to help prevent diseases caused by harmful microorganisms, vitamin production, and digestion of feeds. They are used as balancing supplements to keep environmental parameters at optimal levels (Gatesoupe 1999, Verschuere et al. 2000, Kuebutornye et al. 2019). Probiotic bacteria first colonize along the gastrointestinal tract. They regulate the intestinal microbiota and accelerate the growth of organisms by reducing intestinal problems. Benefits of probiotics on the well-being of the host are as follows: displacing pathogenic microorganisms through the release of occlusive substances, increasing the physicochemical parameters of water, strengthening the immune response of the host species, and improving the nutrition of the host by producing additional digestive enzymes (Gatesoupe 1999, Verschuere et al. 2000, Kuebutornye et al. 2019). Probiotic bacteria are used in solid and liquid forms in spores and/or live form in aquaculture. These bacteria can be added to live feeds, microdiets, pellet feeds culture media, and intraperitoneal injection. In larval rearing of some cultured fish, probiotic supplementation to live feeds is generally applied by adding live probiotic bacteria in liquid form to rotifer and Artemia enrichment media at specific concentrations. In this study, the effect of the administration of Bacillus sp. probiotic bacteria solely via rotifer and rearing water on husbandry performance and digestive physiology of S. aurata and early life development were investigated.
Materials and methods
Ethics statement
All animal care and handling procedures in the current study were conducted in accordance with the Guidelines for Experimental Animals approved by Ege University Center for Research on Laboratory Animals (2020-109; 28 October 2020).
Larval rearing and feeding
A flow chart illustrating the experimental procedures is presented in Fig. 1. Seabream larval culture was performed in triplicate in a green water recirculation system in 15 m3 cylindrical tanks at a density of 80 individuals ml−1. Until 40 days after hatching (DAH), larval rearing conditions and protocols were performed as described by Arığ et al. (2013).
Flow chart illustrating the experimental procedures. Within the scope of the experimental design, while no probiotic application was made to the control group, Bacillus prepared for the experimental group was supplemented to the live food and introduced to the larvae.
After mouth opening at 3 DAH, initial feeding was exogenously begun with rotifers (30% Brachionus plicatilis and 70% Brachionus rotundiformis) and enrichment was maintained by algae and Selco Sparkle (INVE, Belgium). Larval feeding was done with 0.60, 0.40, and 0.30 g/106 rotifers on days 3, 4, and 5. Selco Spresso (INVE, Belgium) was used for enrichment and larvae were fed with an average density of 10 individuals ml−1 rotifer. Larvae were fed only rotifer for the first 20 days and then, feeding was continued with Artemia nauplii. Artemia was fed in two different grades as nauplii and enriched metanauplii of two different sizes (∼480 and 550 µm) in uniformly increasing amounts per ml between 20 and 40 DAH. Larval feeding regime was performed according to Arığ et al. (2013).
Experimental design
Bactosafe® (BernAqua, Belgium) was used as a probiotic in triplicated experiments. This product contains B. subtilis, B. licheniformis, B. cereus, Pediococcus acidilactic i, and a growth medium.
Larvae of seabream were classified into two experimental groups within each treatment, with a cylindrical rearing tank containing about 12 ± 1 × 105 individuals and triplicates per group. In the feeding regime, the control group (referred to as Control) did not receive any probiotics, while the experimental group (referred to as Pro) received probiotics through rotifers and added to rearing water until 20 DAH.
Preparation of probiotic microorganism
Bactosafe® was incubated before being introduced into the live feed according to the manufacturer’s instructions at 10 g l−1. For this purpose, the product was incubated in disinfected seawater (10 g l−1) at a temperature of 33°C for 6–12 h with moderate aeration and periodically checked. The incubation of bacterial spores was determined by periodic pH controls and microscopic observations. Bactosafe® (10 g m−3) was added to the rotifer culture tank media on day 0. The same treatment was applied to the enriched rotifer. With the start of larval feeding, the probiotic was transferred to the larvae with live feed for the first 20 days. Thus, it was thought that it would be easier for the probiotic bacteria to settle in the gastrointestinal tract of the larvae.
Sampling and dissection
Larval sampling and isolation of gastrointestinal section were performed as described by Arığ et al. (2013). Around 30 individuals per group of larvae were sampled in triplicate at 7-day intervals for determination of growth performance. The following formulae were used to determine the Specific Growth Rate (SGR) and survival rate (SR), which are markers for growth performance was calculated by following formulae:
where IBW denotes the initial body weight of fish (mg), FBW denotes the final body weight of fish (mg), and Δt denotes the time interval (days). Survival rate is given by
To analyse of selected digestive enzymes, groups of larvae were sampled in triplicated at 7-day intervals from each tank at the same time (08:00) and depth (30-50cm). For this analysis, dissected gastrointestinal tract on a glass slide at 0°C was used.
Analytical procedures
Analytical procedures and homogenization processes were applied according to Suzer et al. (2013). Specific activities of trypsin and chymotrypsin were assessed using Nα-Benzoyl-dl-arginine-p-nitroanilide and Benzoyl-l-tyrosine ethyl ester as substrates, respectively, following the method described by Tseng et al. (1982) and Worthington (1982). Amylase and lipase activity were assessed using starch and β-naphthylcaprylate as substrates, respectively, following the protocols described by Métais and Bieth (1968) for amylase and the modified method of McKellar and Cholette (1986) as adapted by Versaw et al. (1989) for lipase. An enzyme of the brush border membrane and alkaline phosphatase (AP) activity were analyzed by using p-nitrophenyl phosphate MgCl2 as a substrate, following Bessey et al. (1946). Intestinal enzymes, aminopeptidase N (AN), and leucine–alanine peptidase (LAP) were assessed by utilizing l-leucyl-β-naphthylamide and leucine–alanine as the substrate, respectively (Porteous and Clark 1965, Nicholson and Kim 1975). Enzyme activities were measured in micromoles of substrate (U) hydrolyzed per minute at specific temperatures: at 25°C for trypsin and chymotrypsin and at 37°C for other enzymes and then expressed as specific activity (mU/mg protein−1), where the protein concentration was determined using the Bradford method (Bradford 1976).
Statistical analysis
Data were presented as mean ± SD, with a sample size of n = 30 for larval growth and n = 5 for enzymatic analysis. The Levene test was employed to assess the homogeneity of variance across all datasets. Survival data were analyzed using Fisher’s chi-square test, while both pancreatic and intestinal enzymes, as well as larval growth data, underwent analysis via one-way analysis of variance followed by Newman–Keul’s multiple range test for post-hoc analysis. Additionally, Pearson product–moment correlation tests were conducted to explore relationships between variables, and the results were depicted as a heatmap colored correlation matrix. A significance level of P ≤ 0.05 and P ≤ 0.01 was employed for all analyses except for survival, which was tested at a significance level of 0.05. All statistical analyses were evaluated by SPSS 25.0 software.
Results and discussion
Growth
The growth of S. aurata larvae was monitored in all groups for 40 days as presented in Table 1. Initially, larval length and weight in both groups were measured as 2.36 ± 0.3 mm and 0.64 ± 0.03 mg, respectively. At 40 DAH, weight development was observed in the probiotic-treated groups with measurements of 28.36 ± 3.9 mg compared to the value for the control group was estimated as 23.78 ± 3.6 mg (Table 1). According to measurements at the end of the experiment, larvae in the probiotic-treated group showed >15-fold weight gain between 7 and 40 DAH, while this rate was ∼13-fold for the control group. However, significant differences were observed in weight development between the probiotic and control groups (P < 0.01). Also, final survival rates were 17.3% and 9.6% with SGRs of 7.25% day−1 and 6.82% day−1 for the experimental and control groups, respectively. It’s worth noting that there were no significant differences in survival performance (P > 0.05), but significant differences were found in SGR between the probiotic and control groups (P < 0.01).
Growth parameters of S. aurata larvae at 21 and 40 DAH.
| S1 No | Control | Probiotic | |
|---|---|---|---|
| I | Total length (mm) | ||
| 1 | 21 DAH | 7.49 ± 0.9a | 8.71 ± 0.6b |
| 2 | 40 DAH | 17.81 ± 1.9a | 23.78 ± 3.6b |
| II | Weight (mg) | ||
| 1 | 21 DAH | 14.67 ± 2.6a | 18.13 ± 2.1b |
| 2 | 40 DAH | 24.78 ± 3.4a | 28.36 ± 3.9b |
| III | SGR (% d−1) | ||
| 1 | 21 DAH | 6.42 ± 0.7a | 7.06 ± 0.5b |
| 2 | 40 DAH | 6.83 ± 0.5a | 7.25 ± 0.4b |
| IV | Survival (%) | ||
| 1 | 21 DAH | 38.79 ± 1.8a | 45.28 ± 2.1a |
| 2 | 40 DAH | 9.6 ± 1.2a | 17.3 ± 1.1a |
| S1 No | Control | Probiotic | |
|---|---|---|---|
| I | Total length (mm) | ||
| 1 | 21 DAH | 7.49 ± 0.9a | 8.71 ± 0.6b |
| 2 | 40 DAH | 17.81 ± 1.9a | 23.78 ± 3.6b |
| II | Weight (mg) | ||
| 1 | 21 DAH | 14.67 ± 2.6a | 18.13 ± 2.1b |
| 2 | 40 DAH | 24.78 ± 3.4a | 28.36 ± 3.9b |
| III | SGR (% d−1) | ||
| 1 | 21 DAH | 6.42 ± 0.7a | 7.06 ± 0.5b |
| 2 | 40 DAH | 6.83 ± 0.5a | 7.25 ± 0.4b |
| IV | Survival (%) | ||
| 1 | 21 DAH | 38.79 ± 1.8a | 45.28 ± 2.1a |
| 2 | 40 DAH | 9.6 ± 1.2a | 17.3 ± 1.1a |
Each mean ± SD represents a pool of 30 larvae. Average in the same row with different superscript letters is significantly different (P < 0.01 and P < 0.05).
Growth parameters of S. aurata larvae at 21 and 40 DAH.
| S1 No | Control | Probiotic | |
|---|---|---|---|
| I | Total length (mm) | ||
| 1 | 21 DAH | 7.49 ± 0.9a | 8.71 ± 0.6b |
| 2 | 40 DAH | 17.81 ± 1.9a | 23.78 ± 3.6b |
| II | Weight (mg) | ||
| 1 | 21 DAH | 14.67 ± 2.6a | 18.13 ± 2.1b |
| 2 | 40 DAH | 24.78 ± 3.4a | 28.36 ± 3.9b |
| III | SGR (% d−1) | ||
| 1 | 21 DAH | 6.42 ± 0.7a | 7.06 ± 0.5b |
| 2 | 40 DAH | 6.83 ± 0.5a | 7.25 ± 0.4b |
| IV | Survival (%) | ||
| 1 | 21 DAH | 38.79 ± 1.8a | 45.28 ± 2.1a |
| 2 | 40 DAH | 9.6 ± 1.2a | 17.3 ± 1.1a |
| S1 No | Control | Probiotic | |
|---|---|---|---|
| I | Total length (mm) | ||
| 1 | 21 DAH | 7.49 ± 0.9a | 8.71 ± 0.6b |
| 2 | 40 DAH | 17.81 ± 1.9a | 23.78 ± 3.6b |
| II | Weight (mg) | ||
| 1 | 21 DAH | 14.67 ± 2.6a | 18.13 ± 2.1b |
| 2 | 40 DAH | 24.78 ± 3.4a | 28.36 ± 3.9b |
| III | SGR (% d−1) | ||
| 1 | 21 DAH | 6.42 ± 0.7a | 7.06 ± 0.5b |
| 2 | 40 DAH | 6.83 ± 0.5a | 7.25 ± 0.4b |
| IV | Survival (%) | ||
| 1 | 21 DAH | 38.79 ± 1.8a | 45.28 ± 2.1a |
| 2 | 40 DAH | 9.6 ± 1.2a | 17.3 ± 1.1a |
Each mean ± SD represents a pool of 30 larvae. Average in the same row with different superscript letters is significantly different (P < 0.01 and P < 0.05).
Growth increases (about 10–12 and 13–15-fold increase for the control and probiotic-treated group, respectively) of larvae were observed in both groups, but statistically significant increases were recorded in the probiotic-treated group, clearly indicating that supplementation of live food with probiotics improved the rearing parameters of larvae (P < 0.01). It should be noted herein that not only the enrichment of the live feed with probiotics but also administration to directly rearing water caused about 20-25% better growth performance and contributed to this significant difference. Because of the administration of the probiotic via live food and water, significantly increased the population of Bacillus bacteria in the bacterial microbiota of seabream larvae increased significantly. In such studies, it is noted that probiotics added only to water do not significantly affect both larval development and survival rates. Similarly, in the study where we added Lactobacillus sp. to the culture water of sea bream larvae, we found no significant difference in the control group. However, significant increases in survival and larval development performances were found after the addition of probiotics directly to the live feed and to the tank environment with live feed (Suzer et al. 2008, Avella et al. 2010, Arığ et al. 2013, Kuebutornye et al. 2019). Consequently, findings of these two studies showed similar results in terms of showing the importance of co-application of probiotics to the live feed and tank environment. In another study conducted with seabream larvae, administration of Bacillus sp. strains via Artemia and rotifers, followed by feeding them to larvae (47 DAH) and juveniles (75 DAH), resulted in notable growth enhancements in terms of husbandry parameters when compared to the control group. It is clearly pointed out that especially, at 75 DAH, the group in which the live food was treated with Bacillus showed a significant increase in growth rate (Avella et al. 2010). In addition, B. clausii DE5 and B. pumilus SE5 strains were introduced to copepods as probiotics, and these enriched copepods were then fed to grouper (Epinephelus coioides) larvae for 28 days. It was found from the results at 14 DAH that survival rates were slightly increased with the administration of both Bacillus spp. compared to the control group.
Similarly to these studies, the effects of administration of Virgibacillus proomii and B. mojavensis on both growth and specific activities main digestive enzymes were investigated until 60 DAH during the larviculture of Dicentrarchus labrax. Hamza et al. (2016) reported a significant increase in not only total length and weight gain but also in survival rate and feed conversion ratio (P < 0.05). Furthermore, these results are in line with larvae of seabream (Avella et al. 2010), grouper E. coioides (Sun et al. 2013), beluga, Huso huso (Jafaryan et al. 2010), D. labrax (Md et al. 2015), eel Anguilla japonica (Jang et al. 2023), and C. carpio (Wang and Xu et al. 2006).
Pediococcus acidilactici is also a lactic acid bacterium and is frequently used in aquaculture as a supplementary probiotic. Lamari et al. (2013) conducted a study with seabass larvae, P. acidilactici, administered at 5 DAH in two different doses, alongside L. casei. It was noted that both probiotics significantly impacted growth parameters, especially at 22 DAH; however, the osteocalcin gene was overexpressed in the group fed with L. casei, suggesting that it may play a role in early bone development. In another study with turbot larvae (Psetta maxima), P. acidilactici were supplemented through water and rotifers at different times. The results showed that probiotics given with rotifers were more prevalent in the gastrointestinal tract of larvae.
Specific activities of pancreatic enzymes
The specific activity of trypsin showed a consistent exponential increase of ~3-fold across all groups up to 35 DAH. Subsequently, this activity gradually declined till the end of the trials. The probiotic-treated group exhibited the highest specific activity of trypsin, measured at 116.5 ± 14.3 mU/mg protein−1 (Fig. 2). It was significantly higher compared to the control group (P < 0.05).
Specific activities of trypsin, chymotrypsin, amylase, and lipase of S. aurata larvae up to 40 DAH. In the case of all digestive enzymes except amylase, there is a steady increase and a gradual decrease in specific activities of both groups depending on larval age and size.
Similar to trypsin, chymotrypsin activity showed a slow increasing profile until day 35 in both experimental groups and then gradually decreased until 40 DAH. At the end of the experiment, chymotrypsin activity in the probiotic-treated group presented highest activity (211.3 ± 24.8 mU/mg protein−1) and significant differences were recorded compared to the non-treated group (P < 0.05).
Amylase specific activity presented a sudden increase as above 5-fold in the probiotic-treated group (568.9 ± 35.9 mU/mg protein−1), while this increase was relatively slower at ~3-fold in the control group (328.9 ± 56.7 mU/mg protein−1) until 14 DAH. After this date, the specific activity exhibited a slowly decreasing profile in both groups until the end of the experiment. The peak of specific activity was found at 14 DAH in the probiotic-supplemented group at 568.9 ± 35.2 mU/mg protein−1 (Fig. 2) and it was significantly different than from that in the control group (P > 0.05).
Lipase specific activity exhibited a slow increase (as above 2.5-fold) in the probiotic-treated group up to 21 DAH (324.8 ± 19.4 mU/mg protein−1), while this increasing profile continued until 28 DAH in the control group (239.3 ± 22.1 mU/mg protein−1). After that, it showed a decreasing trend with some fluctuations in both groups. The maximum lipase activity was measured in probiotic-treated group on day 21 at 324.8 ± 19.4 mU/mg protein−1 (Fig. 2), which is significantly higher than that noted in the control group (P < 0.05).
According to a heatmap, there was a positive strong correlation among specific activities of all assayed enzymes between experimental and control groups, indicating that the supplementation treatment responded positively to these activities except amylase. Moreover, amylase activity presented neither a negative nor a positive correlation and exhibited a neutral profile compared to the other digestive enzymes (Fig. 3).
Heatmap illustration of specific activities of measured digestive enzymes. Positive strong correlations were found among specific activities of all assayed enzymes between experimental and control groups, indicating that the supplementation treatment responded positively to these activities except amylase and LAP. However, strong negative correlations were observed among all other enzyme activities.
Specific activities of intestinal enzymes
The specific activities of the intestinal enzymes AP and AN showed a similar profile between the experimental groups. Both enzymes showed a sudden increasing trend until 28 DAH (221.5 ± 36.7 and 345.7 ± 24.9 mU/mg protein−1 for the control and probiotic-treated group, respectively), after which they showed a slower upward trend until the end of the experiment. The highest specific activities of AP and AN were determined on day 40 in the probiotic-treated groups as 732.4 ± 112.3 and 477.2 ± 55.8 mU/mg protein−1, respectively (Fig. 4). Additionally, activities of these intestinal enzymes in probiotic-supplemented groups were found significantly different compared to the control groups (P < 0.05) (Fig. 4).
Specific activities of aminopeptidase N, AP, and LAP of S. aurata larvae up to 40 DAH. In all measured digestive enzymes except LAP, there is a steady increase in specific activities of both groups depending on larval age and size.
Unlike AP and AN activities, LAP exhibited a comparatively higher activity at the beginning of the experiment, followed by a gradual decline over time in both experimental groups. Sudden declines were analyzed after 21 DAH (277.2 ± 32.7 and 312.3 ± 34.8 mU/mg protein−1 for the control and probiotic-treated group, respectively) in synchronization with increase in AP activity. At 40 DAH, peak of LAP activity was measured in the probiotic-supplemented group on day 7 as 411.8 ± 42.5 mU/mg protein−1, but no significant differences were recorded in both groups (P > 0.05) (Fig. 4).
In heatmap, while the specific activities of other intestinal enzymes increased synchronously with the age/size of larvae, LAP activity exhibited a decreasing profile in all groups from the initiation until the end of the experiment. LAP enzyme showed a strong negative correlation with all other enzyme activities (Fig. 3).
For this mechanism to work effectively, a crucial step is for probiotics to accumulate and then colonize the intestinal tissue during the larval development of cultured fish. To corroborate this phenomenon, we supplemented Bacillus with live food and water in seabream larvae. Significant differences were found in alkaline protease enzyme activity in the intestine (P < 0.05), while there were insignificant differences in acid protease activity in the stomach (P > 0.05) (Arığ et al. 2013). Additionally, probiotics aid in preventing intestinal disorders and facilitating the pre-digestion of anti-nutritional factors present in feed ingredients (Verschuere et al. 2000). After passing through the stomach, probiotics undergo germination in the intestine, where they utilize significant quantities of carbohydrates for their growth. This process involves the synthesis of essential digestive enzymes such as amylase, protease, and lipase, which play vital roles in breaking down and absorbing nutrients. In aquaculture, probiotics can be introduced either through dietary supplements via live food and/or extruded diets or by adding them directly to the aquatic environment (Moriarty 1998).
Digestive enzyme activity is commonly recognized as a reliable biomarker of larval nutritional conditions and, to a certain extent, reflects digestive capacity relative to the type of feed provided. Additionally, evaluating the expressions and specific activities of digestive enzyme serves as a comparative measure for assessing the rate of larval growth parameters, nutritional capabilities, digestive performance, and subsequent survival rates (Ueberschär 1995, Suzer et al. 2008, 2013). In this sense, probiotics secrete digestive enzymes to facilitate the digestion of cultured organisms and take part in the synthesis of vitamins to improve the nutrition of these organisms and increase the feed efficiency. Among the probiotic bacteria commonly used in aquaculture, Bacillus is a genus of Gram-positive, rod-shaped bacteria that can survive in extreme environments such as low pH and high temperature, form spores that are pathogenic and non-toxic to the host, and produce various antibacterial substances and thus they considered as preferred candidates compared to other probiotic strains (Kuebutornye et al. 2019). The obtained results distinctly illustrate that supplementing Bacillus via rotifer to S. aurata larvae led to a notable enhancement in the specific activities of all measured both pancreatic and intestinal enzymes. This increase might be related to improved capacity of digestion caused by probiotics and the enhanced absorption of food resulting from this process contributed to the improved survival and growth observed in S. aurata.The relatively improved growth performance obtained from larvae in the experimental group compared to the control group is thought to be due to the probiotic-induced increase in pancreatic and intestinal digestive enzyme activity. In addition to these, another point to be considered as the reason for increased digestive capacity and enzymatic activity in seabream larvae is that Bacillus sp. secrete a wide range of exoenzymes as a working mechanism of probiotics (Ziaei-Nejad et al. 2006, Arığ et al. 2013, Suzer et al. 2013, Kuebutornye et al. 2019). The results of these studies and our study with Lactobacillus showed that probiotic supplementation has a significant effect on enzymatic activity in larvae of S. aurata; especially, specific activities of both pancreatic and intestinal enzymes of probiotic administered in live feed were found to be significantly different among experimental groups (Suzer et al. 2008). It has been confirmed that the administration of probiotics in live food directly affects bacterial colonization in larval intestine, increases the bacterial population, and shows more effective results on digestive (both pancreatic and intestinal) enzymes, thanks to its increased exoenzymes. In addition, it is clearly emphasized that not only this but also by increasing microbial enzyme activity, it directly affects the digestion process in larvae and improves both microbial balance and nutrition and feed utilization in seabream larval culture as in other culture species. Moreover, it is expected that increased feed utilization and improved digestion may positively increase larval culture and pancreatic and intestinal enzyme activity as well as growth performance. For instance, in D. labrax larvae, a mixture of V. proomii and B. mojavensis was administered by live food (rotifer and Artemia) and microdiet for 60 days and significant differences were noted on both pancreatic enzyme, trypsin, and intestinal enzymes, AN, AP, and LAP specific activities (P < 0.05) (Hamza et al. 2016). Similarly, effects of probiotic–prebiotic symbiosis on main digestive enzymes were investigated, and it was reported that the supplementation of fructooligosaccharides, mannan oligosaccharides, and B. clausii in different concentrations led to significant and positive effects on specific activities of intestinal protease and amylase in the Paralichthys olivaceus juveniles (Ye et al. 2011). The results obtained in different studies on larvae and probiotic treatment compared are very close to the results obtained in our study on seabream larvae and it was clearly demonstrated that the supplementation of probiotics had positive effects on the specific activities of main digestive enzymes.
Conclusions
In conclusion, this study could be considered as the first trial to analyze the effects of the addition of three different Bacillus spores solely via rotifer with rearing water on not only pancreatic but also intestinal enzymes in seabream larvae during early life development. Therefore, this study confirmed that supplementing live feed with probiotics proved to be the most effective approach, yielding significant and beneficial outcomes for both growth parameters and pancreatic and intestinal digestive enzymes. The findings presented here offer an innovative perspective on the burgeoning on biotechnology, potentially paving the way for advancements aimed at bolstering the productivity and competitiveness of the aquaculture sector. Further studies should be focused on exploring the effects of supplementation of candidate probiotic-bacteria species on larval rearing of common cultured species.
Acknowledgments
The authors wish to thank the staff of the Kılıç Sea Products Inc., Oren, Milas, Muğla, Türkiye, where the experiments were conducted, for their technical assistance. Additionally, the authors express their gratitude to Nektar Yem Corporation for kindly providing the probiotics used in this study.
Author contributions
Şükrü Yıldırım (Conceptualization [equal], Writing – original draft [equal], Writing – review & editing [equal]), Cüneyt Suzer (Conceptualization [equal], Formal analysis [equal], Methodology [equal], Supervision [equal], Writing – original draft [equal], Writing – review & editing [equal]), Kürşat Fırat (Formal analysis [equal], Investigation [equal]), Şahin Saka (Formal analysis [equal], Investigation [equal]), Müge Hekimoğlu (Formal analysis [equal], Investigation [equal]), Deniz Çoban (Investigation [equal], Methodology [equal], Writing – review & editing [equal]), Ali Yıldırım Korkut (Investigation [equal], Methodology [equal], Writing – review & editing [equal]), İbrahim Köse (Investigation [equal]), Onurkan Antepli (Investigation [equal]), Alize Gökvardar (Investigation [equal]), and Fatih Perçin (Investigation [equal])
Conflict of interest
None declared.
Data Availability
Data will be made available upon a reasonable request to the corresponding author.
