**Amaranth Meal and Environmental** *Carnobacterium maltaromaticum* **Probiotic Bacteria as Novel Stabilizers of the Microbiological Quality of Compound Fish Feeds for Aquaculture**

### **Iwona Goła´s 1,\*, Jacek Potorski 1, Małgorzata Wo ´zniak 2, Piotr Niewiadomski 3, Ma Guadelupe Aguilera-Arreola 4, Araceli Contreras-Rodríguez <sup>5</sup> and Anna Gotkowska-Płachta <sup>1</sup>**


Received: 24 June 2020; Accepted: 23 July 2020; Published: 25 July 2020

**Abstract:** Fish feed should be characterized by microbiological stability to guarantee the optimal health of farmed fish. The aim of this study was to determine the efficacy of amaranth meal (*Amaranthus cruentus*) and a highly active environmental strain of probiotic bacteria, *Carnobacterium maltaromaticum*, as novel supplements that stabilize the quantitative and qualitative composition of microbiota in compound fish feeds for aquaculture, regardless of storage temperature. The total viable counts of mesophilic bacteria at 28 ◦C (TVC 28 ◦C), hemolytic mesophilic bacteria (Hem 37 ◦C), *Staphylococcus* sp. bacteria, aerobic spore-forming bacteria (ASFB), sulfite-reducing anaerobic spore-forming *Clostridium* sp. bacteria, yeasts, and molds were analyzed in control feed (CF), in feed supplemented with amaranth meal (AF), and in feed supplemented with amaranth meal and *C. maltaromaticum* (ACF), stored at a temperature of 4 ◦C and 20 ◦C for 98 days. *Amaranthus cruentus* and *C. maltaromaticum* significantly reduced bacterial counts in fish feeds, regardless of the temperature and duration of storage. The antibacterial and antifungal effects of the tested additives were statistically significant (*p* ≤ 0.05). The studied novel supplements contribute to the microbiological safety of compound fish feeds. The tested additives could be recognized as the key ingredients of organic, environmentally friendly fish feeds, which guarantee the high quality of fish intended for human consumption.

**Keywords:** aquaculture; compound feed; antimicrobial stabilizers; *Amaranthus cruentus*; *Carnobacterium maltaromaticum*

### **1. Introduction**

Feed is one of the main factors that influence fish welfare and the microbiological status of water in aquaculture and freshwater ecology [1]. The nutritional value and microbiological quality of feed determine fish weight gains, and the sanitary and epidemiological safety of aquatic organisms and the aquatic environment [2–6].

Synbiotics containing probiotics and prebiotics enhance the health benefits of feed. They promote the growth and metabolic activity of beneficial microorganisms in the host's gastrointestinal tract without compromising endogenous gut microbiota [7–9]. Probiotics are natural microbiome bacteria that deliver multidirectional beneficial effects for living organisms (humans and animals) at the local and systemic level [2,10]. The role of probiotic feed microbiota in the maintenance of gut homeostasis is increasingly recognized as a critical success factor in fish breeding [6,11–13].

The group of probiotic bacteria includes members of the genus *Carnobacterium* [14–17]. *Carnobacterium maltaromaticum,* which colonizes natural aquatic habitats, is one of the most metabolically active probiotic bacteria in the digestive tract of animals. This bacterial species easily adapts to changes in habitat conditions such as temperature, salinity, and pH, and it delivers health benefits for the host organism [14,18–20] *Carnobacterium maltaromaticum* effectively inhibits the development of pathogenic bacteria and is regarded as a potent immune stimulator in fish [14,21–23].

The growth and activity of probiotic bacteria are influenced by environmental conditions that can be optimized with the use of prebiotics [24,25]. Prebiotics are non-digestible food ingredients that beneficially affect the host by selectively stimulating the growth and/or activity of bacteria colonizing the gastrointestinal tract [7].

Animal and plant meals are one of the main ingredients of fish feeds [26]. However, plant meals contain anti-nutritional factors, and their applicability in compound fish feeds is limited [27]. One of exceptions is amaranth meal, characterized by a low content of anti-nutritional factors, mainly saponins and phytic acid [28].

Amaranth meal contains lignins and various compounds with antioxidant, antibacterial, antiviral, and fungistatic properties [29–31]. Amaranth seeds are also abundant in other health-promoting substances, such as squalene and fiber [32,33]. In a study by Niewiadomski et al. [34], feed supplemented with 20% of amaranth meal promoted the growth of rainbow trout (*Oncorhynchus myskiss*) and improved the digestibility of dietary nutrients. A microbiological analysis in a pilot study conducted by Potorski and Niewiadomski [35] revealed that amaranth supplementation can prevent excessive growth and proliferation of *Staphylococcus* sp. bacteria, *Clostridium* sp. anaerobic spore-forming bacteria, yeasts, and molds in compound fish feeds. A similar beneficial influence of amaranth meal on selected probiotic strains was also observed by Vieira et al. [36] who demonstrated that amaranth meal stimulated the fermentation ability of ten probiotic strains (*Lactobacillus* spp. and *Bifidobacterium* spp.)

The microbiological composition of fish feeds significantly influences fish health and weight gains. This parameter is particularly important if feeds contain harmful microorganisms that compromise fish health, disrupt digestive metabolism, and compromise the reproduction and survival of farmed fish [4]. Feeds should be characterized by microbiological stability and high quality to guarantee the optimal health status and physiological condition of farmed fish. Nevertheless, not all undesirable microorganisms are eliminated during fish feed production. According to the literature [4,37], the standard extrusion process does not guarantee complete elimination of various microorganisms from fish feeds. Furthermore, the metabolic activity of heterotrophic bacteria that survive in ready-made feeds involves the oxidative degradation of lipids and proteins. As a result, the nutritional value of feeds can be modified by natural feed microbiota or by contamination with exogenous microorganisms. Inadequate storage temperature and prolonged storage can also promote the development and metabolic activity of various groups, genera, and species of heterotrophic microorganisms [38].

Our previous experiment [39], which investigated the effect of *C. maltaromaticum* on heterotrophic microbiota, revealed that probiotic bacteria were the main factor responsible for a decrease in the counts of all analyzed bacterial groups in commercial fish feed. The results of studies conducted by other authors [40–42] demonstrated that amaranth meal increased the survival and growth rates of probiotic bacteria and improved the microbial stability of foods. The combined use of environmental probiotic bacteria and amaranth meal as stabilizers of the microbiological quality of fish feeds remains

insufficiently researched. These facts have prompted the authors to evaluate the effectiveness of a highly active environmental isolate of *C. maltaromaticum* and amaranth meal in stabilizing the microbiological quality of fish feed. The aim of this study was to determine the efficacy of amaranth meal (*Amaranthus cruentus*) and a highly active environmental strain of probiotic bacteria, *C. maltaromaticum*, as novel supplements that stabilize the quantitative and qualitative composition of microbiota in compound fish feeds for aquaculture, regardless of storage temperature.

### **2. Materials and Methods**

### *2.1. Isolation and Identification of C. Maltaromaticum Probiotic Bacteria*

A probiotic strain of *C*. *maltaromaticum* was isolated from water samples collected from the benthic zone of Lake Legi ´nskie (at a depth of 34 m) located in north-eastern Poland (N = 53◦58 51" N and E = 21◦8 4"). The strain had been isolated during a previous study conducted by the Department of Environmental Microbiology of the University of Warmia and Mazury in Olsztyn.

The isolate was identified to species level by matrix-assisted laser desorption/ionization time-of-flight mass spectrometry (MALDI-TOF VITEK® MS) at the Department of Microbiology, Escuela Nacional de Ciencias Biológicas, Instituto Politécnico Nacional, in Mexico City, Mexico. The identification was additionally verified by 16S rDNA (recombinant DNA) sequencing with the BigDye Terminator v3.1 kit in the ABI 3730xl genetic analyzer (Applied Biosystems, Foster City, USA). In addition, 16S rDNA genes were sequenced by PCR with the use of 27F (5 -AGAGTTTGATCATTGGCTCAG-3 ) and 1492R (5 -GGTACC-TTGTTACGACTT-3 ) primers according to the method described by Gillan et al. [43]. The BLAST program available on the website of the National Center of Biotechnology Information [44] was used to identify DNA sequences. The results of 16S rDNA sequencing are presented in Table S1 (Supplementary Materials).

After the identification process, *C. maltaromaticum* was considered as a probiotic strain based on the hemolysis assay, and its acid and bile tolerance properties, according to the guidelines developed by a joint FAO/WHO working group [45]. The hemolytic activity of *C. maltaromaticum* was determined on tryptone soya agar (TSA; Oxoid, Basingstoke, UK) with 5% addition of defibrinated sheep blood incubated at 37 ◦C for 48 h [46]. The bile salt tolerance test of the studied strain was performed in MRS broth culture medium (Sigma - Aldrich, Germany) containing 0.5%, 1.0%, or 2.0% bile salts (Oxoid LP0055) according to the procedure proposed by Succi et al. [47]. The *C. maltaromaticum* isolate was tested for acid tolerance based on its growth on medium with varying pH (1.5, 2.5, 3.5, and 4.5), as described by Vijayarama et al. [48].

### *2.2. Determination of the Metabolic Activity of Probiotic Bacteria Based on the Utilization of Di*ff*erent Carbon Sources*

The applicability of the environmental *C. maltaromaticum* isolate for further analysis was determined by analyzing the bacteria's metabolism based on its utilization of various carbon sources. The biochemical activity of the *C. maltaromaticum* probiotic isolate and its potential to compete for nutrients with feed microbiota were estimated using the OmniLog®System (Biolog, USA). A 96-well plate containing various carbon compounds was inoculated with the evaluated bacterial strain. The plate was incubated, and biochemical parameters were read in a microstation reader. The strain's utilization of different carbon compounds as sources of energy was determined based on the intensity of color reactions.

### *2.3. Compound Feed*

The experiment was performed on three types of extruded compound feeds: Control feed (CF) without the addition of amaranth meal, experimental feed containing 20% of amaranth meal (AF), and experimental feed containing 20% of amaranth meal and *C. maltaromaticum* probiotic bacteria (ACF). The composition of each feed is presented in Table 1. All feeds were formulated based on

the recommendations of Hart et al. [49] and NRC [50]. The feeds were extruded with a co-rotating twin screw extruder (Metalchem, Poland) equipped with a Ø 4.5 mm pellet stencil. The following extrusion processing parameters were applied: Screw speed—105–125 rpm, cutter speed—50 rpm, head temperature—120 ◦C, barrel temperature of 130–150◦ C in 30 s, die diameter—2.0 mm. Compound feeds were enhanced with a mixture of fish oil and soybean oil (5% each). The tested strain of *C. maltaromaticum* was added to the oil mixture. Next, the probiotic oil suspension was added to two experimental feed samples. Tthe oil mixture was pumped into the feed at 0.9 Mpa for 5 min with the use of a vacuum pump. The feed contained 40.0% crude protein, 15.0% crude fat, 3.0% crude ash, 37.0 nitrogen-free extract (NFE), and 5% water.


**Table 1.** Feeds composition (g·100 g−<sup>1</sup> dry diet).

1—control feed (CF); 2—feed containing 20% amaranth (AF); 3—feed containing 20% of amaranth and Carnobacterium maltaromaticum (ACF); 4—Composition of the vitamin premix (IU·1 kg−<sup>1</sup> dry diet): Vitamin A—70,000 IU; vitamin D—200,000 IU; vitamin E—17,500 IU; vitamin K—867 IU; vitamin C—28,500 IU; vitamin B1—1067 IU; vitamin B2—2000 IU; vitamin B5—5334 IU; vitamin B6—1334 IU; vitamin B12—400 IU; biotin—200 IU; niacin—12,000 IU; folic acid—800 IU; inositol—20,000 IU; choline chloride—120,000 IU; betaine—75,000 IU; 5—Composition of the mineral premix (g·1 kg−<sup>1</sup> dry diet): FeSO4·H2O—4334 g; KI—0.734 g; CuSO4·5H2O—0.267 g; MnO—0.734 g; ZnSO4·H2O—1250 g; ZnO—0.750 g; Na2SeO4—0.034 g; CFU—colony forming unit.

### *2.4. Experimental Design*

The prepared feeds (CF, AF, and ACF) were used in an experiment that lasted for 98 days. The control feed (CF) was divided into two equal parts, and the feed containing 20% amaranth meal (AF) was divided into four equal parts under sterile conditions. Every CF and AF sample was placed in a separate, sterile, and tightly closed vessel made of dark glass. Two samples (CF 4 ◦C, AF 4 ◦C) were chill-stored at a temperature of 4 ◦C, and two samples (CF 20 ◦C, AF 20 ◦C) were stored at a temperature of 20 ◦C throughout the experiment. Cultures of the environmental *C. maltaromaticum* strain were added to the remaining two samples (AF) at 1.5 <sup>×</sup> <sup>10</sup><sup>9</sup> CFU·g−<sup>1</sup> (Table 1). One of the samples containing probiotic bacteria (ACF 4 ◦C) was chill-stored at 4 ◦C, and the other sample (ACF 20 ◦C) was stored at 20 ◦C for 98 days.

### *2.5. Microbiological Analyses*

All feed samples (CF, AF, and ACF) stored at 4 ◦C and 20 ◦C were subjected to microbiological analyses after 7, 14, 21, 28, 35, 42, 49 56, 63, 70, 77, 84, 91, and 98 days of the experiment. The following parameters were determined: Total counts of *C. maltaromaticum* bacteria on tryptone soya agar (TSA; Oxoid, Basingstoke, UK) with the addition of 3% yeast extract and 1.5% (*w*/*v*) NaCl [14], total counts of mesophilic bacteria on tryptone soya agar (TSA; Oxoid, Basingstoke, UK) incubated at 28 ◦C for 48 h (TVC 28 ◦C), total counts of hemolytic mesophilic bacteria on tryptone soya agar (TSA; Oxoid, Basingstoke, UK) with 5% addition of defibrinated sheep blood incubated at 37 ◦C for 48 h (hemolytic mesophilic bacteria (Hem) 37 ◦C), total counts of aerobic spore-forming bacteria (ASFB) on an agar/broth medium (Biocorp, Warsaw, Poland) with glucose incubated at 28 ◦C for 72 h, counts of *Staphylococcus* sp. bacteria on the Chapman medium (Merck KgaA, Darmstadt, Germany) incubated at 37 ◦C for 48 h, counts of sulfite-reducing anaerobic spore-forming *Clostridium* sp. bacteria on the Wilson-Blair medium (Merck KgaA, Darmstadt, Germany) incubated at 37 ◦C for 18 h, and total yeast and mold counts on the Rose-Bengal-Chloramphenicol Agar (RGBC; Merck KgaA, Darmstadt, Germany) incubated at 28 ◦C for 5 days.

All analyses were performed according to Polish Standard [51]. The potential pathogenicity of Hem 37 ◦C, *Staphylococcus* sp., and *Clostridium* sp. bacteria was determined based on their hemolytic activity on tryptone soya agar (TSA; Oxoid, Basingstoke, UK) with 5% addition of defibrinated sheep blood incubated at 37 ◦C for 48 h. Hemolysis was confirmed when a transparent zone was formed around the inoculated colonies [46]. Mean microbial counts were calculated based on the values determined in three replicates of the same sample of compound fish feed. Finally, the counts of all analyzed microorganisms were expressed in CFU·1 g−<sup>1</sup> of compound feed.

### *2.6. Statistical Analysis*

The mean values, standard deviations, standard errors, and confidence interval (CI = 95%, N = 3) of microbial counts in feeds (CF, AF, ACF) stored at a temperature 4 ◦C and 20 ◦C were calculated. The relationships between *C. maltaromaticum* bacterial counts and microbial (TVC 28 ◦C, Hem 37 ◦C, ASFB, *Staphylococcus* sp., *Clostridium* sp., yeasts, and molds) counts were determined by Spearman's non-parametric rank correlation test (*p* ≤ 0.05). The significance of differences in microbial counts between the analyzed types of fish feed (CF, AF, ACF) stored at different temperatures (4 and 20 ◦C) and for different periods of time (7, 14, 21, 28, 35, 42, 49, 56, 63, 70, 77, 84, 91, and 98 days) was determined by one-way analysis of variance (ANOVA). Leven's test was used to assess the homogeneity of variance. The verified hypothesis was rejected when Leven's test produced statistically significant results. The Kruskal–Wallis test, a non-parametric version of the classical one-way ANOVA, was then applied. Statistical analyses were performed in the Statistica 13.3 software package (TIBCO Software Inc., Palo Alto, USA) [52].

### **3. Results**

### *3.1. Probiotic Properties of Carnobacterium Maltaromaticum*

The studied *C. maltaromaticum* isolate was not capable of causing hemolysis, which suggested that the strain was not pathogenic.

The strain tolerated the tested pH values. After 3 h acid exposure, the isolate's survival rate was higher at pH 2.5 (76.1%) than at pH 1.5 (65%), and it reached 82.3% at pH 3.5 and 87.8% at pH 4.5. The bile salt tolerance test revealed a small difference in the survival rates of *C. maltaromaticum*. The highest isolate viability (85.2%) was observed at a 2% concentration of bile salts, whereas the lowest viability (79.5%) was noted at a 0.5% concentration of bile salts; 83.2% of *C. maltaromaticum* bacteria survived at a 1.0% concentration of bile salts (data not shown).

The *C. maltaromaticum* isolate tested in our study could be classified as a probiotic strain based on the results of the above analyses and according to the guidelines developed by a joint FAO/WHO working group [45].

### *3.2. Metabolic Activity of C. maltaromaticum Probiotic Bacteria*

The results of the analyses examining the utilization of various carbon sources by the environmental *C. maltaromaticum* isolate are presented in Figure 1. The analyses performed in the Omnilog Gen III system (Biolog, Hayward, CA, USA) revealed that the evaluated strain actively metabolized 70 carbon sources. The studied *C. maltaromaticum* strain was capable of growth at pH 5 and 6, and in the presence of 1%, 4%, and 8% NaCl. The tested isolate did not metabolize the following substrates: L-alanine, L-arginine, L-aspartic acid, L-glutamic acid, histidine, D-gluconic acid, and mucic acid. The analyzed strain did not metabolize vancomycin, tetrazolium blue chloride, L-pyroglutamic acid, α-ketoglutaric acid, α-ketobutyric acid, and acetoacetic acid. These results confirmed the very high biochemical


activity of the studied environmental probiotic isolate, and suggested its potential to compete for nutrients with feed microbiota.

**Figure 1.** The results of a metabolic activity test analyzing the chemical sensitivity of an environmental *Carnobacterium maltaromaticum* probiotic isolate and its ability to utilize different carbon sources (GEN III MicroPlate™). Purple color—metabolic activity of the *C. maltaromaticum* isolate, white color—no metabolic activity of the *C. maltaromaticum* isolate.

### *3.3. The Quantitative and Qualitative Composition of Bacterial Microbiota in Compound Fish Feeds*

The mean (of three replicates) counts of mesophilic bacteria (TVC 28 ◦C), hemolytic mesophilic bacteria (Hem 37 ◦C), *Staphylococcus* sp., *Clostridium* sp., aerobic spore-forming bacteria (ASFB), yeasts and molds in CF, AF, and ACF, and *C. maltaromaticum* bacteria stored at a temperature of 4 ◦C and 20 ◦C during the 98-day experiment are presented in Figure 2. The mean values, standard deviations, standard errors, and confidence interval of three replicates of microbial counts are shown in Table S2 (Supplementary Materials). In CF, microbial counts differed by several orders of magnitude, depending on the analyzed microbial group and the temperature and time of feed storage. In CF 4 ◦C samples, TVC 28 ◦C and *Clostridium* sp. counts increased several-fold after 14 and 28 days of storage, respectively, relative to initial values. The counts of other microbial groups (Hem 37 ◦C, *Staphylococcus* sp., yeasts, and molds) in CF 4 ◦C samples continued to decrease in successive weeks of the experiment. The noted decrease ranged from 10<sup>1</sup> to 10<sup>5</sup> CFU across the analyzed microbial groups, subject to storage time (Figure 2A).

**Figure 2.** The mean (of three replicates) total viable counts (CFU·g<sup>−</sup>1) of mesophilic bacteria (TVC 28 ◦C), hemolytic mesophilic bacteria (Hem 37 ◦C), *Staphylococcus* sp., *Clostridium* sp., aerobic spore-forming bacteria (ASFB), yeasts, and molds in: (**A**) Control feed (CF 4 ◦C) stored at a temperature of 4 ◦C, (**B**) control feed (CF 20 ◦C) stored at a temperature of 20 ◦C, (**C**) feed supplemented with 20% amaranth

meal (AF 4 ◦C) stored at a temperature of 4 ◦C, (**D**) feed supplemented with 20% amaranth meal (AF 20 ◦C) stored at a temperature of 20 ◦C, (**E**) feed supplemented with 20% amaranth meal and *Carnobacterium maltaromaticum* bacteria (ACF 4 ◦C) stored at a temperature of 4 ◦C, and (**F**) feed supplemented with 20% amaranth meal and *Carnobacterium maltaromaticum* bacteria (ACF 20 ◦C) stored at a temperature of 20 ◦C during a 98 day experiment. The mean values, standard deviations, standard errors, and confidence interval of three replicates of microbial counts are shown in Table S2 (Supplementary Materials).

The counts of nearly all microorganisms (excluding ASBF) increased by around 100% in CF 20 ◦C samples after 14, 28, and 42 days. In CF 20 ◦C samples, TVC 28 ◦C and yeast counts peaked on day 28 at 92 <sup>×</sup> 10<sup>7</sup> and 12 <sup>×</sup> 104 CFU·g<sup>−</sup>1, respectively. The highest counts of potentially pathogenic bacteria (Hem 37 ◦C, *Staphylococcus* sp., *Clostridium* sp.) were noted after 42 days of feed storage. The maximum counts of Hem 37 ◦C, *Staphylococcus* sp., *Clostridium* sp., and molds were determined at 2.8 <sup>×</sup> 106, 3.0 <sup>×</sup> 102, 45, and 2.5 <sup>×</sup> 103 CFU·g−1, respectively. A minor decrease in microbial counts was noted in successive weeks of the experiment. However, on day 98, the counts of all evaluated microorganisms in CF 20 ◦C samples were several-fold to several hundred-fold higher than those in CF 4 ◦C samples (Figure 2B).

In feed samples supplemented with 20% amaranth meal stored at a temperature of 4 ◦C (AF 4 ◦C), the counts of all analyzed microbial groups decreased by several orders of magnitude after 14 days of the experiment. On day 28, Hem 37 ◦C (20 CFU·g<sup>−</sup>1) was the only potentially pathogenic microorganism in the studied samples. Toward the end of the experiment, AF 4 ◦C samples were colonized only by TVC 28 ◦C (500 CFU·g<sup>−</sup>1) and ASFB (5 CFU·g<sup>−</sup>1) (Figure 2C).

In AF 20 ◦C samples, the decrease in the counts of potentially pathogenic Hem 37 ◦C bacteria was considerably lower than that in AF 4◦C samples. On day 28, Hem 37 ◦C counts in AF 20 ◦C samples were determined at 1.0 <sup>×</sup> 10<sup>4</sup> CFU·g<sup>−</sup>1, and they were 500-fold higher than those in AF 4 ◦C samples on the same day. The counts of TVC 28 ◦C, ASBF, and yeasts were also several-fold to several dozen-fold higher in AF 20 ◦C samples than in AF 4 ◦C samples on the same days (Figure 2D). Hem 37 ◦C, *Staphylococcus* sp., and *Clostridium* sp. survived for longer periods of time in AF 20 ◦C than in AF 4 ◦C. Hem 37 ◦C, *Staphylococcus* sp., and *Clostridium* sp. were eliminated from AF 20 ◦C samples only after 56 days, and from AF 4 ◦C—already after 14 or 28 days of the experiment (Figure 2C,D).

Feed samples supplemented with 20% amaranth meal and a highly active environmental strain of *C. maltaromaticum* probiotic bacteria (ACF 4 ◦C, ACF 20 ◦C) were characterized by the lowest counts (Figure 2E,F) and the lowest survival rate of all analyzed microbial groups, regardless of storage temperature (Table S3). Potentially pathogenic *Staphylococcus* sp., *Clostridium* sp., and Hem 37 ◦C bacteria were not detected in ACF 4 ◦C and ACF 20 ◦C samples already after 7 days. In the first two weeks of the experiment, TVC 28 ◦C counts decreased around 1000 fold, ASFB counts decreased more than 100-fold, and yeast counts decreased several fold in ACF 4 ◦C and ACF 20 ◦C samples relative to the initial values. On day 98, ACF 4 ◦C samples were colonized only by TVC 28 ◦C and ASBF at 10 and 5 CFU·g<sup>−</sup>1, respectively (Figure 2E). TVC 28 ◦C and ASFB counts were higher in ACF 20 ◦C at 120 and 20 CFU·g<sup>−</sup>1, respectively (Figure 2F). Additionally, Spearman's test revealed significant (*<sup>p</sup>* <sup>≤</sup> 0.05) negative correlations between *C. maltaromaticum* counts and almost all microbial populations (except for *Clostridium* sp. and molds) in ACF, regardless of storage temperature (Table 2).

The differences in the quantitative and qualitative composition of bacterial and fungal microbiota in the analyzed types of fish feeds (CF, AF, and ACF) stored at different temperatures (4 ◦C and 20 ◦C) and for different periods of time were confirmed by the statistical analysis (Table 3). The Kruskal−Wallis test revealed significant (*p* ≤ 0.05) differences in the counts of all analyzed microorganisms between the evaluated feeds (CF, AF, and ACF) and in ASFB and yeast counts in feed samples stored for different periods of time. Significant (*p* ≤ 0.05) differences were also observed in *Staphylococcus* sp., *Clostridium* sp., and mold counts in feed samples stored at different temperatures, and in TVC 28 ◦C, Hem 37 ◦C, *Staphylococcus* sp., and *Clostridium* sp. counts in feed samples stored for different periods of time.

**Table 2.** The values of correlation coefficients between microbial counts in feed supplemented with 20% amaranth meal and *C. maltaromaticum* probiotic bacteria (ACF) stored at 4 and 20 ◦C. The correlations between microbial counts in ACF 4 ◦C (N = 14) and ACF 20 ◦C (N = 14) samples were analyzed with Spearman's test.


1—mesophilic bacteria (TVC 28 ◦C); 2—hemolytic mesophilic bacteria (Hem 37 ◦C); 3—aerobic spore-forming bacteria (ASFB); \*—statistically significant correlations in Spearman's test (*p* ≤ 0.05).

**Table 3.** The quantitative composition of microorganisms in control feed (CF), feed supplemented with 20% amaranth meal (AF), and feed supplemented with 20% amaranth meal and *Carnobacterium maltaromaticum* bacteria (ACF) stored at different temperatures (4 and 20 ◦C) for 7, 14, 21, 28, 35, 42, 49, 56, 63, 70, 77, 84, 91, and 98 days, validated in the Kruskal−Wallis test (N = 84).


1—mesophilic bacteria (TVC 28 ◦C); 2—hemolytic mesophilic bacteria (Hem 37 ◦C); 3—aerobic spore-forming bacteria (ASFB); \*—statistically significant differences; one-way ANOVA, *p* ≤ 0.05.

### **4. Discussion**

The analyses of the quantitative and qualitative composition of microbiota in fish feed samples revealed significant differences (*p* ≤ 0.05) across the examined types of feed (CF, AF, ACF), feed storage temperatures, and feed storage times. Control feed (CF) was characterized by the highest counts, highest survival rates, and longest survival times of all analyzed microbial groups, which indicates that feed ingredients promote the growth of both specific feed microorganisms and potentially pathogenic microorganisms [37,53,54]. Similar results were reported by Petreska [4] and Goła´s et al. [55] who analyzed the counts of heterotrophic mesophilic bacteria and selected potentially pathogenic bacteria, yeasts, and molds in commercial feeds administered to intensively reared *Silurus glanis* L.

In our study, the counts of all specific feed microbiota and potentially pathogenic microorganisms (Hem 37 ◦C, *Staphylococcus* sp., *Clostridium* sp.) in feed supplemented with 20% amaranth meal (AF 4 ◦C, AF 20 ◦C) decreased by 1 to 4 orders of magnitude relative to those determined in CF 4 ◦C and CF 20 ◦C. The survival times of potentially pathogenic bacteria (Hem 37 ◦C, *Staphylococcus* sp., *Clostridium* sp.) were also significantly shorter in AF 4 ◦C and AF 20 ◦C than in CF 4 ◦C and CF 20 ◦C. The obtained results and the presence of significant differences (*p* ≤ 0.05) in the counts of all analyzed microbial groups between CF and AF samples indicate that feed supplementation with 20% amaranth meal inhibits the growth of bacterial and fungal microbiota regardless of storage temperature or duration (Table 3). The antibacterial and antifungal properties of amaranth meal are also confirmed by the decrease in the counts of the remaining microbial groups (TVC 28 ◦C, ASFB, yeasts, molds) in

AF 4 ◦C and AF 20 ◦C samples in successive weeks of the experiment. The above could be attributed to the fact that amaranth meal contains lignins whose antioxidant, antibacterial, antiviral, and fungistatic properties contribute to the maintaining of the adequate microbiological quality of feed [31,56–59]. The addition of amaranth meal stabilizes natural microbiota in animal feeds, enhances the nutritional value of feeds, and improves performance.

Research studies have confirmed the beneficial influence of amaranth-supplemented feeds on the health status and body weight gains of rats [60], intensively farmed pigs [61,62], chickens [63], calves, lambs, sheep, and ruminants [56]. Studies investigating the effect of amaranth-supplemented feeds on fish in different farming systems also demonstrated that amaranth meal stimulated the immune system of fish [64], their growth performance, and the enzymatic activity of their gut microbiota [5,65]. The results of the present study indicate that amaranth meal can be effectively used to improve the quality and microbiological safety of fish feeds.

The counts of all studied microorganisms (TVC 28 ◦C, Hem 37 ◦C, ASFB, *Staphylococcus* sp., *Clostridium* sp., yeasts, and molds) were lowest in ACF 4◦ and ACF 20◦ relative to AF and CF stored at the corresponding temperatures. The counts, percentage viability, and survival times of the evaluated microbial groups were considerably lower in ACF 4◦ and ACF 20◦ than in AF 4◦ and AF 20◦ (Figure 2C–F; Table S3), which indicates that amaranth meal and *C. maltaromaticum* probiotic bacteria exert antibacterial and antifungal effects on natural microbiota and potentially pathogenic microorganisms in compound feed. The synergistic effects of the tested feed additives could be attributed to the symbiotic relationship between amaranth meal and the evaluated probiotic bacteria, and their ability to inhibit the growth and development of various microbial groups and genera. An in vitro study [57,66] revealed that amaranth is a source of bioactive compounds that suppress the proliferation of many microorganisms, including *Staphylococcus aureus*, *Bacillus, Escherichia coli*, *Salmonella typhi*, *Pseudomonas aeruginosa*, *Proteus mirabillis, Klebsiella pneumoniae*, and *Candida albicans*. Amaranth meal also promotes the development of many species of probiotic bacteria, such as *Lactobacillus plantarum,* L. *paralimentarius,* L. *helveticus,* L. *sakei, Pediococcus pentosaceus,* L. *paralimentarius*, *Enterococcus mundtii*, *E. hermanniensis*, *E. durans, Enterococcus* sp., and *Leuconostoc mesenteroides*, whose metabolic activity enhances the nutritional value and health benefits of food products [67–69]. An in vitro study conducted by Gullón et al. [70] demonstrated that amaranth was characterized by a high prebiotic potential and promoted the growth of probiotic microflora isolated from the human digestive tract. By inhibiting the growth and development of naturally occurring microorganisms and pathogenic microbiota in foodstuffs and feedstuffs [5,66] probiotic bacteria and amaranth contribute to improving fish welfare and performance in various aquaculture systems [71–73].

The lowest counts of all evaluated microbial groups and genera and the shortest microbial survival times were noted in ACF samples regardless of storage temperature and storage time, which indicates that amaranth meal and *C. maltaromaticum* probiotic bacteria exert synergistic effects on the quantitative and qualitative composition of feed microbiota. Feed supplementation with 20% amaranth meal and *C. maltaromaticum* (ACF) bacteria completely inhibited the growth of most analyzed microorganisms (excluding ASFB and TVC 28 ◦C) in feeds stored at 4 ◦C and 20 ◦C for 7 days. The results of our in vitro study were validated statistically, which suggests that the novel tested additives contribute to the microbiological stability of fish feeds regardless of storage conditions and storage time.

### **5. Conclusions**

The results of the present study, which investigated the supplementation of compound fish feeds with innovative additives, amaranth meal, and a highly active environmental strain of probiotic bacteria, *C. maltaromaticum*, indicate that the tested additives exert synergistic effects and contribute to the microbiological stability of fish feeds regardless of the temperature and time of storage. The evaluated components decreased the counts, percentage viability, and survival times of various groups and genera of microorganisms that occur naturally in feeds, which suggests that they can minimize feed losses resulting from the growth and metabolic activity of autochthonous and allochthonous microbiota in feeds that are stored for excessive periods of time and/or at inadequate temperature. Excessive microbial growth lowers the nutritional value of feed, and decreases nutrient digestibility and assimilability, which may negatively affect fish performance in aquaculture. The addition of 20% amaranth meal and a highly active environmental strain of probiotic bacteria, *C. maltaromaticum,* to fish feed inhibited the growth of potentially pathogenic microbiota (Hem 37◦, *Staphylococcus* sp., and *Clostridium* sp.) in vitro, which is important for the growth rate and welfare of fish. Due to their novel synergistic health-promoting properties, amaranth meal and environmental *C. maltaromaticum* bacteria could be recognized as the key ingredients of organic, environmentally friendly fish feeds, which guarantee the high quality of fish intended for human consumption.

**Supplementary Materials:** The following are available online at http://www.mdpi.com/2076-3417/10/15/5114/s1, Table S1: The identification of an environmental strain of *Carnobacterium maltaromaticum* bacteria based on 16S rDNA sequence analysis. Table S2: The mean values (X), confidence interval (CI) (CI = 95%, N = 3), standard deviations (SD), and standard errors (SE) of microbial counts in control feed (CF), in feed supplemented with 20% amaranth meal (AF), and in feed supplemented with 20% amaranth meal and *Carnobacterium maltaromaticum* bacteria (ACF) stored at 4 and 20 ◦C during 98 days of the experiment. Table S3: Survival rates of microorganisms (%) in control feed (CF), feed supplemented with 20% amaranth meal (AF), and feed supplemented with 20% amaranth meal and *Carnobacterium maltaromaticum* bacteria (ACF) stored at 4 and 20 ◦C during 98 days of the experiment.

**Author Contributions:** Conceptualization, I.G. and M.W.; methodology, I.G., J.P., M.W., P.N., M.G.A.-A. and A.C.-R.; software, I.G.; validation, I.G., J.P., M.W. and P.N.; formal analysis, J.P. and P.N.; investigation, I.G., J.P., M.W. and P.N.; resources, I.G., M.W., M.G.A.-A. and A.C.-R.; data curation, I.G. and J.P.; writing—original draft preparation, I.G. and M.W.; writing—review and editing, M.W. and A.G.-P.; visualization, A.G.-P.; supervision, I.G. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research received no external funding.

**Acknowledgments:** This study was supported by research grants No. 18.610-004-300 and No. 18.610.001-300 from the Ministry of Science and Higher Education (Poland). The project was financially co-supported by the Minister of Science and Higher Education under the program entitled "Regional Initiative of Excellence" for the years 2019–2022, Project No. 010/RID/2018/19, amount of funding PLN 12,000,000. The authors would like to thank Z. Filipkowska and E. Korzeniewska, Professors at the University of Warmia and Mazury in Olsztyn for the culturing and sequencing of 16S rDNA of the tested *C. maltaromaticum* strain, and K. Gli ´nska-Lewczuk (University of Warmia and Mazury in Olsztyn) for help in the statistical processing of data.

**Conflicts of Interest:** The authors declare that they have no conflict of interest. The funders had no role in the design of the study; in the collection, analyses, or interpretation of data; in the writing of the manuscript, or in the decision to publish the results.

### **References**


© 2020 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

### *Communication* **E**ff**ects of Colored Light on Growth and Nutritional Composition of Tilapia, and Biofloc as a Food Source**

### **Daniela Lopez-Betancur 1, Ivan Moreno 2, Carlos Guerrero-Mendez 1, Domingo Gómez-Meléndez 1, Manuel de J. Macias P. <sup>3</sup> and Carlos Olvera-Olvera 1,\***


Received: 28 November 2019; Accepted: 30 December 2019; Published: 3 January 2020

**Abstract:** Light stimulation and biofloc technology can be combined to improve the efficiency and sustainability of tilapia production. A 73-day pilot experiment was conducted to investigate the effect of colored light on growth rates and nutritional composition of the Nile tilapia fingerlings (*Oreochromis niloticus*) in biofloc systems. The effect of colored light on the nutritional composition of bioflocs as a food source for fish was measured. Three groups were illuminated in addition to natural sunlight with colored light using RGB light emitting diodes (LEDs) with peak wavelengths (λ) of 627.27 nm for red (R), 513.33 nm for green (G), and 451.67 nm for blue (B) light. LED light intensity was constant (0.832 mW/cm2), and had an 18-h photoperiod of light per day throughout the study. The control group was illuminated only with natural sunlight (natural). Tilapia had an average initial weight of 0.242 g. There was a significant effect of colored light on tilapia growth and composition. The R group showed the best growth rate, highest survival, and highest lipid content. The B group showed homogeneous growth with the lowest growth rate and lipid content, but the highest protein level. On the other hand, the biofloc composition was influenced by the green light in the highest content of lipids, protein, and nitrogen-free extract.

**Keywords:** food science; light; color; LEDs; sustainable aquaculture; fish production; preliminary results

### **1. Introduction**

Aquaculture is one of the fastest-growing food production areas and it is one of the most important sources of food, nutrition, income, and livelihood for hundreds of millions of people worldwide [1]. By 2030, the production of freshwater species such as carp, catfish, and tilapia is expected to represent about 60% of total aquaculture production [2,3]. However, fish farming requires the use of land, freshwater, and environmental resources, which are increasingly scarce and expensive worldwide. By 2030, the world could have a global water deficit of 40% in the usual commercial scenario, and by 2050, the demand for water is expected to increase by 55% in all sectors of production [4]. Therefore, an increase in aquaculture production must be carefully planned, minimizing the environmental impact and optimizing the use of natural resources.

Today, sustainable aquaculture systems produce more fish without affecting the environment, such as using biofloc technology (BFT). In systems with BFT, there are limited exchanges of water and, thus, there is an accumulation of organic matter and nutrients that promote the development of

a microbial community called a bioflocs [5]. Bioflocs are conglomerates of phytoplankton, bacteria, zooplankton, microbial grazers, and particulate organic matter, which are mainly heterotrophic bacteria. When this conglomerate is mixed with an added external carbon source, the growth of heterotrophic bacteria is stimulated and the absorption of nitrogen occurs through the production of microbial proteins, which serve as a food source for fish that is available 24 h per day [6–9]. Biofloc systems are mainly used to cultivate tilapia (*Oreochromis* sp.) and white shrimp (*Litopenaeus vannamei*) because both species can eat biofloc and live in environments with high levels of turbidity [10]. The biofloc community can also be used to improve water quality by adding carbon sources to the pond [11,12].

The development of new technologies and scientific studies in aquaculture is essential to improve intensive fish production. A promising improvement in aquaculture comes from light emitting diode (LED) lighting. It has been shown that lighting in aquaculture can influence embryonic development, releasing reproductive hormones that increase fish growth [13,14]. However, fish are visual feeders that need a minimum light intensity to eat and, thus, grow and develop [15]. In addition to the influence on embryonic development, the intensity and spectrum light in certain photoperiods (intensity, duration, and periodicity) can be used to alter and control the growth of fish [16–20]. Photoperiods also influence the release of reproductive hormones, which play an important role in fish reproduction and growth [13]. In addition, under short wavelengths such as blue light, melatonin (which is the hormone that is responsible for sleep) decreases in the bass fish, and the lower the melatonin, the longer the fish are awake, and the more they feed [21]. However, banana shrimp (*Penaeus merguiensis*) have a faster growth related to the intensity of the lighting, and the higher the light intensity, the less the shrimp feed, but they grow faster, possibly because of better efficiency of food assimilation [22]. For shrimp with BFT, it was observed that when low light intensity was used, shrimp production decreased by 48%, and the density of microalgae, zooplankton, and rotifers also decreased by 60%, 60%, and 90%, respectively [23].

Tilapias are generally diurnal feeders that feed at different time periods during the day [24]. Using fluorescent tube lamps with a photoperiod of 18L:6D and illuminance of 2500 lux, it is possible to produce more Nile tilapia seeds (percentage of spawning synchrony and percentage of the sac and swim-up fry stages) compared to a shorter photoperiod with less illumination (2500 lux/15 h, 2500 lux/12 h, 500 lux/18 h, 500 lux/15 h, and 500 lux/12 h) [14]. The promising new LED light technology, which has not been widely explored in aquaculture, especially in BFT systems, can be a useful light source tool if the light parameters (intensity, color, and periodicity) are analyzed and applied to obtain benefits in the production of Nile tilapia in BTF. The rapid development of LED technology in recent years has exceeded the characteristics of incandescent lamps in luminous efficiency, low heat emission, robustness, environment resistance, non-toxicity, durability, and adjustable light intensity and wavelength, which allows precise control of the light spectrum [25,26]. The aim of this study was to investigate the effect of using colored LED light on tilapia growth, the nutritional composition of the Nile tilapia fingerlings, and the composition of the bioflocs that are used as a food source for fish.

### **2. Materials and Methods**

### *2.1. Ethics Statement*

All work with animals in this research was done in accordance with the "Guidelines for the Use of Fishes in Research" published by the American Society of Ichthyologists and Herpetologists (https://www.asih.org/sites/default/files/2018-05/asf-guidelines-use-of-fishes-inresearch-2013.pdf) and complied with the Mexican law on experimental animals according to the protocols: NOM-062-ZOO-1999 and NOM-033-SAG/ZOO-2014.

The experimental design and the fish-use protocol were approved within the project "LED lighting to improve the production of tilapia in biofloc systems" by the ethics committee for animal research at the "Autonomous University of Zacatecas" (authorization number: ACS/UAZ/036/2018).

The light intensities used in all the experiments did not exceed values that were observed in natural waters. This study did not include endangered or protected species. All fish were acclimatized for greenhouse conditions for 2 weeks, with the lamps off (the fish only received natural light), before the start of the experiment. The duration of our experiment was 73 days and 489 Nile tilapia fingerlings (*Oreochromis niloticus*) were used. During the planning stage of the experiment, water quality parameters (temperature, dissolved oxygen, and pH) were monitored twice a day to confirm optimal living conditions for tilapia. The average water quality parameters for fishponds were: 28.50 ◦C for temperature, 6.68 mg/L for dissolved oxygen, 7.9 for pH, 0.08 mg/L for ammonia, and 0.83 mg/L for total ammonia nitrogen. The average ammonia and total ammonia nitrogen values were monitored every 8 days. The fishponds were supplied with a 300 W thermostat heater (Grupo acuario LOMAS, Ciudad de Mexico, CDMX, Mexico) to avoid temperature variations. Aeration was provided by a Sino-Aqua blower (1/2 Hp of power, and 9" membrane diffuser discs). Additionally, 10% of the tank water was exchanged each week. During the progress of this experiment, 105 fish died from natural causes such as acclimatization, and no specific pathologies were observed to determine if the fish should be euthanized. At the end of the experiment 384 fish were sacrificed. We carefully used standardized procedures for fish euthanasia. Since Nile tilapia do not tolerate cold water, they were sacrificed by rapid chilling (hypothermic shock) by keeping the tilapia in ice water at 2 ◦C for 10 min after the opercular movement stopped. The fish were stored in a freezer until chemical analysis of their nutritional composition (approximately 72 h).

### *2.2. Laboratory Facilities and Fish Stocking*

A 73-day experiment was performed in a greenhouse in the prototypes laboratory at the Autonomous University of Zacatecas, Mexico. Before the experiment started, 489 Nile tilapia (*O. niloticus*) fingerlings were purchased from a commercial hatchery (AQUAMOL S.C. DE R.L., Jamay, Jal., Mexico), and moved to the greenhouse, where they underwent a period of acclimatization. The Nile tilapia fingerlings had an initial average weight of 0.242 ± 0.01 g. They were randomly distributed, with a same density of 0.2 kg/m<sup>3</sup> (approximately 123 fish per tank, and exactly 30 g of biomass per tank), in circular tanks (150-L capacity, 39 cm high × 70 cm in diameter) under natural light. This is a pilot study because in this experimentation there were no replicated tanks (*n* = 1). All results presented are preliminary results, but significant because each tank contained a large number of fish.

### *2.3. Experimental Design and Setup*

The experiment was designed to study the effect of colored light under real light conditions. The experiment could also be transferred to real production because real conditions were present in the experiments, such as all four treatments received natural light, which is consistent with aquaculture farms.

This investigation implemented a special change, in addition to receiving natural light, three treatments were illuminated with red, green, and blue LED lamps for 18 h per day. The illumination was provided by RGB (red, green, and blue monochromatic light) LED lamps with peak wavelengths (λ) of λ = 627.27 nm for the red light treatment (R), λ = 513.33 nm for the green light treatment (G), and λ = 451.67 nm for the blue light treatment (B). LED lamps were positioned 25 cm above the water surface, with the purpose that the lamp opening angle (110◦) should focus completely on the water surface of the entire tank diameter. LED light intensity of all the lamps was constant at 0.832 mW/cm2, and the natural light provided by the sunlight inside the greenhouse had a maximum of irradiance of 0.95 mW/cm2 at 14:00 h. The fourth treatment, which we called "Natural" light treatment, only received natural light, and served as a control treatment. The lighting conditions for the four treatments are shown in Figure 1.

**Figure 1.** Light periods received by each tank during this research. All treatments received natural light, but three of the treatments were also illuminated with red, green, and blue colored light, which was provided by an LED lamp, with a photoperiod of 18 h per day.

### *2.4. Characterization of LED Lamps*

The LED lamps used in this experiment were optically characterized using a spectrophotometer (USB2000, Ocean Optics, Largo, FL, USA) and a NIST calibrated radiometer (ILT-1400, International Light Technologies Inc., Peabody, MA, USA). Wavelength (λ) and spectral width (Δλ) parameters were measured using the spectrophotometer device. The irradiance or radiant power density on the water surface of each tank was measured using the radiometer. The measured LED lamp parameters are presented in Table 1, where the average irradiance was 0.832 mW/cm2. Figure 2 also shows the spectral power distribution of light emitted by each colored LED lamp.

**Table 1.** Optical properties of light colors in light emitting diode (LED) lamps: red (R), green (G), and blue (B).

**Figure 2.** The spectral power distribution of the RGB LED lamps used in the study: (**a**) R LED lamp; (**b**) G LED lamp; and (**c**) B LED lamp.

### *2.5. Nile Tilapia Fingerlings and the Biofloc System*

All treatment groups began the study with tap water, which was previously aerated for 1 week to allow water dechlorination. No water exchange was made, and the formation of the biofloc started from zero when the tilapia arrived. Each tank was fixed at 29 ◦C using a 300 W thermostat heater to avoid high temperature variations, and to provide a comfortable temperature for the fish. Pure cane sugar was added to each tank as a carbon source, which was calculated using the method described by De Schryver et al. [27]. The use of a carbon-to-nitrogen (C/N) ratio of 20:1 in the tanks ensures the water quality during the biofloc formation because we guarantee a low level of total ammonia nitrogen (TAN) in the water [28,29]. The fish were fed by hand twice a day using the commercial diet I during the first month, and then using the commercial diet II, according to the requirements of the growth stage. The nutritional composition of the commercial diet I, commercial diet II, and cane sugar used are listed in Table 2. The daily amount of food was adjusted every week (the feeding rate started with 10% of the fish's weight per day and it decreased to 5% per day) according to the growth of the fish, fish survival, and the biofloc that was expected in each tank. The fish feed composition was provided by the manufacturer and was also corroborated based on a bromatology test to guarantee the consumption and quality of the feed. During the experimental period, the tanks were checked by hand every day, and any dead fish were removed and recorded.


**Table 2.** Feed composition for tilapia and cane sugar used as a carbon source.

### *2.6. Water Quality Parameters*

Temperature, dissolved oxygen (measured by the YSI model 550A dissolved oxygen meter device (Yellow Springs Instrument Co., Yellow Springs, OH, USA)), and pH (measured by the Hanna model HI 98127 device (Hanna Instruments, Woonsocket, RI, USA)) were measured twice a day in each tank at 10:00 and 20:00 h. Ammonia–nitrogen (*NH*<sup>3</sup> − *N*), which is one form of ammonia in TAN that is toxic for fish, was regulated by adding cane sugar. *NH*<sup>3</sup> − *N* levels were measured once per week in the tanks using the ammonia checker device (Hanna model HI715 (Hanna Instruments, Woonsocket, RI, USA)). In addition, the TAN value was calculated using a modified mathematical expression described by Boyd and Tucker [30], which is written as follows:

$$TAN = \left(NH\_3 - N\right) \times \left\{1 + antilog \left[0.09018 + \left(2729.92 / (273.15 + T)\right)\right] - pH\right\},\tag{1}$$

where *TAN* is the total ammonia nitrogen, (*NH*3-N) is the ammonia–nitrogen, *T* is the water temperature (◦C), and pH is the water pH.

### *2.7. Growth Rates*

Throughout the experiment, biometric measurements were made every week in a group of randomly selected of 15 fish in each tank. At the end of the experiment, all the fish were weighed and measured, and the growth rates were reported as the mean ± standard deviation. According to the final number of fish in each treatment, there were 127 fish that underwent R treatment, 79 fish that underwent G treatment, 94 fish that underwent B treatment, and 84 fish that underwent the Natural treatment. The mean ± standard deviation is presented for the fish weight and length. Additional measurements to evaluate fish growth were used, as follows: initial weight (g/fish), final weight (g/fish), final body length (cm/fish), survival (%), specific growth rate (SGR), daily weight gain (DWG), and feed conversion rate (FCR). The mathematical expressions of the evaluation metrics were extracted from published studies [31,32]. These evaluation metrics are written as follows:

$$\text{Initial weight (g)} = \text{(initial biomass (g))} \text{(initial number of fish)}, \tag{2}$$

$$\text{Final weight (g)} = \text{(final biomass (g))} \text{(final number of fish)}, \tag{3}$$

$$\text{Final body length (cm)} = \frac{\sum\_{i=0}^{\text{Final number of fish}} \text{fish body length (i)}}{\text{final number of fish}},\tag{4}$$

$$\text{Survival} \left( \% \right) = \text{(final number of fish)} \text{(initial number of fish)} \times 100\% \tag{5}$$

$$\text{SGR} \left( \% / \text{day} \right) = \frac{\ln(\text{final weight} \left( \text{g} \right)) - \ln(\text{initial weight} \left( \text{g} \right))}{\text{number of days}} \times 100\% \tag{6}$$

$$\text{DWG } (\text{g/day}) = (\text{final weight} \, (\text{g}) - \text{initial weight} \, (\text{g})) / (\text{number of days}), \tag{7}$$

$$\text{FCR} = \left( \text{total feed intake (g)} \right) / \left( \text{total wet weight gain (g)} \right), \tag{8}$$

where the initial biomass is the weight of all the fish in each tank at the beginning of the experiment, and the final biomass is the weight of all the fish at the end of the experiment. The total feed intake is the amount of food supplied to the fish, and the total wet weight gain is the difference between the final weight and the initial weight of the entire biomass.

### *2.8. Nutritional Composition of the Fish Body and Bioflocs*

Composition of the fish body is valuable information about the nutritional contribution that the fish meat will have for a consumer. A biofloc nutritional composition study also provided information on whether the diet of the fish could be completed correctly using the biofloc. At the end of the experiment, all the fish were caught for a biomass analysis. The water in the tanks was sieved using a 200-mesh screen to maintain the greatest amount of biofloc. The composition of the fish body and the bioflocs were determined by the composition of lipids, moisture, crude protein, crude fiber, ash, and nitrogen-free extract, according to standard methods [33,34]. The values were determined using Soxhlet extraction apparatus while the other parameters were determined with bromatology using gravimetric techniques. The nutritional composition analysis was performed by the Chemical Laboratory of Special Studies at the Autonomous University of Zacatecas.

### *2.9. Statistical Analysis*

All the measured data and the metrics that were calculated in this research were analyzed to achieve a performance evaluation and a broad comparison between the four light treatments. Since the parameters obtained from the tanks are mean values, a one-way analysis of variance (ANOVA), which compares the "variation" of a group of mean values, was used.

Water quality parameters cannot be analyzed using ANOVA because there are no replicate tanks (*n* = 1). However, growth rates can be analyzed because each tank had many final fish as follows: *n* = 127 for R treatment, *n* = 79 for G treatment, *n* = 94 for B treatment, and *n* = 84 for Natural treatment. Thus, the growth rates were analyzed using the one-way ANOVA, followed by Tukey's test, with a significance level of 5%. Before performing the one-way ANOVA evaluation, the hypothesis of equality (homogeneity) of variances was verified using the Levene's test. Data analysis of the biofloc and fish nutritional composition was performed using the mean values of three sample replicates.

### **3. Results**

### *3.1. Water Quality Parameters*

The average values of the water parameters remained approximately equal between all groups during the experimental period. Thus, a constant temperature was maintained, with a variation of less than 1 ◦C, the pH value showed a difference below 1.25%, the variation of ammonia–nitrogen (*NH*3-N) was less than 0.03 mg/L, and the difference in dissolved oxygen and TAN were less than 0.07 and 0.17 mg/L, respectively. The mean values of water quality parameters were calculated, and they are shown in Table 3.


**Table 3.** Water quality parameters.

Data are presented as the mean ± standard deviation.

### *3.2. Growth Rates*

Mean weight, mean body length, survival, specific growth rate, and feed conversion ratio were used as metrics to evaluate the effect of colored light. According to the measured data, the R group showed the highest final weight among the fish, and it also had the lowest initial weight. Therefore, the R group had the highest performance, with the best growth rates and DWG (see Table 4). In addition, the data obtained show that the R and G groups presented the best gains in weight and length. However, the G group had the worst survival. The feed conversion ratio was better in the R and Natural groups compared with the other groups. The SGR value was significantly higher in the groups R and G compared with the other groups. The quantitative distribution of the final weight of the fish for each group is shown in Figure 3.



SGR = specific growth rate; DWR = daily weight gain; FCR = feed conversion rate. Data are presented as the mean ± standard deviation. Values in the same row with different superscript letters indicate significant differences based on the one-way ANOVA and Tukey's test (*p* < 0.05).

**Figure 3.** The mean final weight of the fish in each treatment. Each bar represents the mean with the standard deviation. Different letters (such as a, b, and ab) represent significant differences (*p* < 0.05).

### *3.3. Nutritional Composition of the Fish Body and Bioflocs*

The mean values for nutritional composition of the fish and the nutritional composition of the bioflocs as a food source for fish are presented in Figures 4 and 5, respectively. The moisture value was higher in the body of the fish of group B. In addition, the body of the fish in groups B and Natural presented a higher concentration of crude protein compared to the other treatments. The mineral content (ash) in the fish in groups R, G, and Natural was less than that measured in the fish in group B. Lipids in the body composition of the fish were most influenced by the use of colored lights. Fish in group R showed a higher lipid content and nitrogen-free extract.

**Figure 4.** Box-and-whisker plots showing the nutritional composition of the fish (%).

Measurements of the mean nutritional composition of the bioflocs in the G and Natural groups showed a higher concentration of crude protein compared to the other treatments. The ash composition in the bioflocs from the R and G groups was lower than the content measured in the bioflocs from the B and Natural groups. In addition, the mean nutritional composition of the biofloc in group G showed a higher content of lipids and nitrogen-free extract.

**Figure 5.** Box-and-whisker plots showing the nutritional composition of the biofloc (%).

### **4. Discussion**

To investigate the effect of colored light in the Nile tilapia fingerlings, the parameters that were not related to the light color were kept constant, but there were slight experimental variations in these parameters that are expected not to significantly change this condition of the research. The mean water temperature was maintained at less than 1 ◦C of variation, and all the tank water temperatures were in accordance with the optimum range for the growth of the Nile tilapia fingerlings (26–30 ◦C), as indicated in [35]. The mean water temperature in the four treatments was 28.50 ◦C, which is very close to the ideal temperature of 28 ◦C, as reported in [36]. The dissolved oxygen value did not have a very significant variation between all the groups and it was above the level that was acceptable for Nile tilapia (4–5 mg/L), in accordance with [37]. The pH fluctuated between 7 and 8 during the experimental stage, which is the most appropriate range in tilapia aquaculture to obtain the optimum growth and survival rate, as recommended by [38]. The pH value was not different between the R, G, and B groups, which could be because of a greater total amount of bacterial activity that is influenced by the use of artificial light compared to the Natural group. Using a C/N ratio of 20, the TAN concentration has a

similar behavior value to pH and it was maintained at levels that are suitable for Nile tilapia, according to [39]. The *NH*<sup>3</sup> − *N* values did not exceed the levels that were considered lethal for Nile tilapia during the experiment, and this value remained below the recommended limit of 0.1 mg/L [40].

Light color was the only intentionally different parameter among all treatments. Therefore, each treatment was only influenced by a certain color or wavelength of the light. There are not many studies reported of the influence of colored light on the growth of Nile tilapia and fish of different species in general [18], and as far as we know, this is the first study on the fingerling stage of development of tilapia. Even more, all previous studies on light effect use other experimental conditions, which include significant differences in: The lack of biofloc, the type or duration of photoperiod, the temperature, the light intensity, peak wavelengths, light spectrum shapes, etc.

In this research, we studied the effect of colored light in Nile tilapia fingerlings and bioflocs, the R light treatment was the aquaculture system that showed superior performance in many aspects. All the groups received the amount of food according to the biomass that was present in each tank.

It is possible to ensure that the R treatment presented the best growth performance because it obtained the best values of total biomass gained, SGR, and DWG. Additionally, there was a great demand for feed intake; when food was provided it was consumed immediately, which was confirmed because the R treatment had the best FCR of 0.8. This is consistent with [41], where feeding conduct was analyzed, but in juvenile Nile tilapia under other experimental conditions, showing (in terms of time (seconds), and chemical cues) that feeding motivation was higher in their red light environment. As well, a similar behavior was found in adult rainbow trout [42], where the growth of fish was increased at red illumination under other experimental conditions, showing low cortisol levels in their blood (less stressful environments).

Nevertheless, the G light environment showed the second-best weight gain, but also had the lowest survival. It is possible to rule out that the low survival was caused by a peak of ammonia and TAN, because during the period in which these fish deaths occurred, measurements of ammonia and TAN were in the optimal range for the life of tilapia. A similar low survival phenomenon using green light was also observed and reported in [43], but in small fish groups (four fish per tank) of adult tilapia under fluorescent lights and other experimental conditions. One possible reason to this phenomenon could be that an aggressive behavior of the fish was observed in a green light environment. Other related study determined (by blood test) the basal plasma cortisol levels (stress response) in adult Nile tilapia, and it was found that cortisol levels increased with green light [44]. This could be a cause of fish mortality because the green light may create stressful environment and increase cortisol levels. A high level of cortisol also affects the nutritional composition of tilapia, causing high moisture levels and low protein levels [45], which agree with our results. The low level of protein can be attributed to an intensive use of muscle protein to get higher energy than that obtained from food, which could be caused by the stress due to green light [46]. Green light may be an important factor of mortality, but research with replicate tanks should be carried out to see all possible mortality factors.

Aly et al. [45] reported that hybrid red tilapia in saline water had a lower growth performance under blue light and other experimental conditions. These results were attributed to a reduced vision of the fish under green light, which reduced feeding (also see [47]). In addition, they reported that hybrid red tilapia that were illuminated under red light had the best growth performance [45]. These observations are in agreement with our results, where the Nile tilapia fingerlings in freshwater under blue light had the lowest growth performance, and those under red light had the best growth performance. In addition, in [43], yellow light was shown to have the best growth performance, which also agrees with our results, because the yellow light has a long wavelength that is close to the wavelength of the red light.

In our research, the effect of colored light on the nutritional composition of Nile tilapia fingerlings, and on the biofloc composition as a food source for tilapia was also measured, and this has not been reported before. The effect of light on body composition of tilapia showed that the R group had the highest lipid content and also the lowest moisture level (this indicates that moisture and lipid content

are inversely related, which is in accordance with [48]). However, under blue light, the fish had the highest percentage of protein and moisture and the lowest level of lipids, while under green light, the fish maintained a low content of lipids and proteins. Under natural light, the fish obtained a high crude protein content and the highest content of ash.

The effect of light color on bioflocs showed that the G group had the highest lipid content. In addition, the bioflocs in the G and Natural groups had the highest protein level. For the ash content, B and Natural groups had the highest percentage. In addition, although the biofloc of B group had a high level of lipids, the fish nutritional composition under B light had a low level of lipids. This opposite relationship could be explained by the stress reduction under blue light [44], and also a stress reduction under the lipid-rich diet [49]. Such a possible stress reduction, would reduce the formation of lipid deposits in fish, indicating that such good health condition of tilapia under B light could be because the lack of stress.

The results discussed above may have an important application in tilapia aquaculture with biofloc systems because they improve our understanding of the significant effect that light color has on the growth of fish and their nutritional composition of both the fish body and the bioflocs.

### **5. Conclusions**

We investigated the influence of RGB colored light on the growth and nutritional composition in tilapia in biofloc systems. We also measured the light effect on the biofloc nutritional composition. The results showed that the peak wavelength or color of light had a significant effect on growth performance and nutritional composition of Nile tilapia, which can be used to improve the sustainability and efficiency of tilapia production based on the needs of the consumer. Fish under blue light had the lowest growth performance, but blue light significantly improved the protein levels, had a low lipid level, and obtained the most homogeneous growth among the treatments. The effect of the red light on fish growth was significant. Tilapia under red light showed the best weight gain, body length, and survival rate, and also showed the highest lipid content. The fish also had a lower ash content under colored light compared to natural light. In addition, colored light influenced the lipid, protein, and ash content in bioflocs.

These results provide support for further studies in aquaculture to improve efficient, sustainable, and intensive fish production. For example, our results suggest that growth rates, feed utilization efficiency, survival, and nutritional body composition may be controlled using colored light, which can significantly modify the nutritional quality provided by Nile tilapia and bioflocs. In addition, future research may include the effect of different intensity levels, photoperiods, and light spectra. For example a broad light spectrum with a designed special shape [50], resulting from the combination of LEDs of different colors and intensities, to obtain the best effects of each light wavelength and maximize the nutritional quality of tilapia and bioflocs. This can be adjusted based on the needs of the consumer and, in the future, the scarcity of food resources.

**Author Contributions:** Conceptualization, D.L.-B., C.O.-O. and I.M.; Methodology, D.L.-B. and C.G.-M.; Validation, C.O.-O. and I.M.; Formal analysis, D.L.-B., and C.G.-M.; Investigation, D.L.-B. and C.G.-M.; Resources, C.O.-O., D.G.-M. and M.d.J.M.P.; Data curation, D.L.-B. and M.d.J.M.P.; Writing-original draft preparation, D.L.-B.; Writing—review and editing, I.M., and D.L.-B.; Visualization, C.G.-M. and D.G.-M.; Supervision, C.O.-O. and I.M.; Project administration, I.M., C.O.-O. and M.d.J.M.P. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research received no external funding.

**Acknowledgments:** This research was supported by CONACYT (Mexican Council for Science and Technology) through financing scholarship granted to Daniela Lopez-Betancur.

**Conflicts of Interest:** The authors declare no conflict of interest.

### **References**


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