*Article Astragalus membranaceus* **Extract (AME) Enhances Growth, Digestive Enzymes, Antioxidant Capacity, and Immunity of** *Pangasianodon hypophthalmus* **Juveniles**

**Hany M. R. Abdel-Latif 1,\*, Hamada A. Ahmed 2, Mustafa Shukry 3, Md Reaz Chaklader 4, Rasha M. Saleh <sup>5</sup> and Mohamed A. Khallaf <sup>6</sup>**


**Abstract:** The present study evaluated the impacts of powdered *Astragalus membranaceus* extract (AME) on the growth, physiological responses, and serum immunity of *Pangasianodon hypophthalmus* juveniles. Four test diets were formulated to include varying AME levels as 0.0 (control), 1.5 (AME1.5), 3.0 (AME3.0), and 4.5 (AME4.5) g/kg. Fish weighing approximately 11.50 g were stocked into four triplicate groups and hand-fed on the test diets three times daily for two months. At 60 days postfeeding, the growth performance, including weight gain and the specific growth rate, was increased quadratically (R<sup>2</sup> > 0.90) with increasing AME inclusion levels. An improvement in the feed intake and feed conversion ratio were also noticed in groups fed at different AME levels. The whole-body and amino acid composition were unaffected by the test diets. A significant quadratic trend in the digestive enzymes (lipase, α-amylase, and protease) was found along with increasing AME inclusion levels. Liver enzymes associated with liver functions were improved by AME dietary inclusion levels. Meanwhile, the blood urea nitrogen, uric acid, and creatinine values were unaffected by AME dietary inclusion. On the other hand, serum immunity (lysozyme and total Igs) was elevated with a significant quadratic trend along with increasing AME dietary inclusion levels. Liver MDA levels decreased with increasing AME levels. Liver CAT, GPx, and SOD enzyme activities demonstrated a significant increasing trend along with dietary AME inclusion. The aforementioned effects of dietary AME on *P. hypophthalmus* health underpinned the potentiality of AME to be used as a phyto-additive to improve the functionality of aquafeed.

**Keywords:** herbal medicines; growth; digestive enzymes; striped catfish; immunity

#### **1. Introduction**

*Pangasianodon hypophthalmus* is generally known as striped catfish, pangas catfish, or pangasiid catfish. It is a freshwater fish popularly farmed in several Asian countries [1,2]. Its high tolerance to environmental conditions, along with its higher growth rate, lower operational expenditures, and relatively higher economic returns, make it a favorable candidate for finfish aquaculture in several countries [3,4]. This fish is usually cultivated in high stocking density [2], which may increase its susceptibility to various infectious agents, especially bacterial pathogens [5]. However, to achieve the intensification of production, researchers have engaged, in recent years, to find novel immunostimulants to enhance the immunity of cultured species and support antibiotic- and chemotherapeutic-free and

**Citation:** Abdel-Latif, H.M.R.; Ahmed, H.A.; Shukry, M.; Chaklader, M.R.; Saleh, R.M.; Khallaf, M.A. *Astragalus membranaceus* Extract (AME) Enhances Growth, Digestive Enzymes, Antioxidant Capacity, and Immunity of *Pangasianodon hypophthalmus* Juveniles. *Fishes* **2022**, *7*, 319. https://doi.org/10.3390/ fishes7060319

Academic Editors: Marina Paolucci and Shunsuke Koshio

Received: 21 September 2022 Accepted: 30 October 2022 Published: 3 November 2022

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**Copyright:** © 2022 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 (https:// creativecommons.org/licenses/by/ 4.0/).

disease-resistant sustainable aquaculture. Using plant herbal extracts as immunostimulants has been considered a common practice to improve the aquafeed's functionality [6–8]. Several researchers have assessed a variety of plant- or herbal extract-based diets to promote the growth and immunity of *P. hypophthalmus*. For example, *Zingiber officinale*, *Euphorbia hirta*, *Phyllanthus amarus*, *Azadirachta indica*, and *Allium sativum* were proven to be potential immunostimulants for *P. hypophthalmus* [9,10]. Moreover, *Andrographis paniculata* extract also improved the immune responses and disease resistance of juvenile pangas catfish [11]. Our recent study also reported the positive impacts of *Silybum marianum* extract on the growth, immune performance, antioxidant condition, and intestine morphometry of this valuable fish species [8].

*Astragalus membranaceus* (AM) is a frequently used medicinal plant in China. It has well-known immunostimulatory, antioxidant, general tonic, and hepato-protective properties due to its phyto-components of *Astragalus* polysaccharides, flavonoids, phenolics, alkaloids, and saponins [12–14]. There is a plethora of studies on the beneficial roles and applications of AM as a functional feed supplement in several finfish and shellfish aquaculture. For example, feeding *Oreochromis niloticus* on a diet supplemented with 0.1% powdered AME extract increased its resistance to *Aeromonas hydrophila* [15]. Dietary AM plants also promoted the growth performance, antioxidant responses, immunity, and increased stress tolerance of yellow perch (*Perca flavescens*) [16]. Furthermore, dietary application of AM (1.5, 3, and 4.5%) promoted growth and antioxidative status, as well as modulated cold water stress tolerance in bluegill sunfish (*Lepomis macrochirus*) [17]. Dietary supplementation with a dried AM plant (0.25% and 0.5%) also enhanced the growth, immunity, and antioxidant status of *Litopenaeus vannamei* [18]. However, a recently published study demonstrated that dietary supplementation with 0.1–0.8% AM root extract did not improve the growth, hepatic morphology, or immunity of hybrid groupers (*Epinephelus lanceolatus* ♂× *E. fuscoguttatus* ♀) [19].

*Astragalus* polysaccharides (ASP) are dietary supplements that are the subject of extensive research and recent publications in finfish and shrimp species. Diet supplementation with 0.05% and 0.10% ASP promoted growth, digestive enzyme activities, and intestinal morphology of larval large yellow croaker (*Larimichthys crocea*) [20]. Dietary ASP (50 and 150 mg/kg) ameliorated the growth, antioxidant activity, and immunity of turbot (*Scophthalmus maximus*) [21]. In addition, dietary ASP obtained from dried roots of *A. membranaceus* (100 mg/kg) enhanced the growth and immunity of *Carassius auratus* juveniles [22]. Moreover, dietary 0.01% ASP improved the growth, gut health condition, and anti-viral immunity of *Danio rerio* [23]. Furthermore, dietary ASP (1 g/kg) also enhanced the immunity, intestinal microbiota, and disease resistance of *Ctenopharyngodon idella* [24]. It was recently reported that dietary ASP (100, 200, and 300 mg/kg) significantly improved the growth, antioxidant responses, and immunity of *Catla catla* [25]. In addition, dietary supplementation with ASP obtained from dried sliced roots of *A. membranaceus* (30 g/kg) also promoted the growth and improved disease resistance of *L. vannamei* [26]. Lately, it has been demonstrated that dietary 0.15% ASP and 0.30% ASP significantly improved the immunity, disease resistance and attenuated toxicity signs in Nile tilapia [27,28].

Various reports regarding emerging issues have jeopardized the aquaculture industry, such as the elaboration of antibiotic- and therapeutics-resistant bacteria and the deposition of antibiotic and chemotherapeutic compounds in the aquaculture products [29]. These concerns have led to efforts to find suitable, effective, economic, and eco-friendly alternatives such as dietary herbal extracts [6–8]. Herein, though several researchers have conducted several experimental trials on *P. hypophthalmus* after its introduction in Egypt [4,8], there are no known reports on the effects of AME as a feed additive for striped catfish juveniles. Consequently, this study was intended to assess the supplementing impacts of powdered AME on the growth performances, serum immunity, hepato-renal functions, and antioxidant responses of *P. hypophthalmus* juveniles to test its suitability for improving the functionality of striped catfish diets and maintaining sustainable aquaculture.

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

#### *2.1. Herbal Extract and Analysis of Its Bioactive Constituents*

Powdered *Astragalus membranaceus* extract (AME) was procured from Free Trade Egypt Co., Alexandria, EGY. The GC-MS method analyzed the bio-active constituents in the ethanolic extract of AME, as illustrated by Gomathi et al. [30]. Bioactive components were characterized and identified by mass spectral and relative retention time (RT) comparisons with the WILEY 09 and NIST14 Mass Spectral databases and libraries. The flavonoid compounds in AME were determined by HPLC analysis following Mattila et al. [31], whilst the phenolic content of AME ethanolic extract was measured in accordance with the methods outlined by Öztürk et al. [32].

#### *2.2. Fish and Adaptation Conditions*

At a fish hatchery in Borg El Arab, Alexandria, EGY, three hundred juvenile healthy *P. hypophthalmus* were raised in spherical black fiberglass tanks with 1000 L of water capacity for fifteen days before starting the experiments (for adaptation). During this period, fish were fed daily on a well-balanced reference diet (30% crude protein (CP), Aller Aqua Co., October City, Egypt). The feed ingredients of this diet were formulated to contain all the nutritional necessities for rearing the fish in accordance with NRC guidelines [33] (Table 1).



Formerly published in our study [8].1 Danish FM (contains 72.0% CP), procured from TripleNine Fish Protein, DK-6700 (Esbjerg, Denmark).2 Egyptian soybean flour (contains 46.0% CP), procured from Cargill Trading Egypt Co. (Katameya, Cairo, Egypt).<sup>3</sup> AGRI-VET CO. for manufacturing Vitamins and Feed additives (10th of Ramadan City A2, Egypt). The vitamin premix mixture (contains per 1 kg): Vitamin A (17,000 IU); Vitamin D3 (2400 IU); Vitamin E (240 mg); Vitamin K3 (11 mg); Vitamin B1 (24 mg); Vitamin B2 (52 mg); Vitamin B3 (275 mg); Vitamin B6 (25 mg); Vitamin B12 (0.05 mg); Vitamin C (220 mg); Folic acid (15 mg); Calcium d–pantothenate (55 mg); Biotin (1.5 mg); Inositol (125 mg), and Choline chloride (2500 mg). The mineral mixture (contains per 1 kg) is composed of the following: Iron (74.50 mg/kg); Copper (12.5 mg/kg); Manganese (200 mg/kg); Zinc (80 mg/kg); Iodine (2 mg/kg); Selenium (0.330 mg/kg), and Cobalt (1.5 mg/kg).4 NFE = 100 − (CP + EE + CF + Ash). <sup>5</sup> GE was estimated on the basis of 23.60, 39.40, and 17.20 kJ/g of CP, EE, and NFE, respectively following NRC guidelines [33].6 P/E ratio was estimated as mg crude protein/KJ.

#### *2.3. Formulation of AME-Based Diets*

The powdered AME was fully blended with the reference diet at varying inclusion levels of 0.0, 1.5, 3.0 and 4.5 g/kg diet to make 4 experimental diets termed AME0.0 (control), AME1.5, AME3.0, and AME4.5, respectively. For 15 min, we thoroughly combined and

pulverized all of the feed ingredients. A suitable amount of sunflower oil and warm distilled water (20.0 ◦C) were added to the feed ingredients to make a dough. The formulated feed pellets of 2.0 mm in diameter and 2.0 mm in length were produced by passing the dough through a meat mincer. The prepared experimental diets were subjected to the sun to dry, then preserved in Ziplock bags, labeled with the group names, and stored in the refrigerator (−18 ◦C) until used for feeding the fish groups during the feeding trial.

#### *2.4. Fish Rearing and Experimental Design*

Juvenile striped catfish with an initial body weight of (11.50 ± 0.5 g) were allocated into four triplicate groups and were cultivated into twelve rectangular glass aquaria. Each glass aquarium was sized 1.0 m × 0.90 m × 0.75 m with a 100 L water capacity. There were 30 individuals per group (10 fish per aquarium). Throughout the duration of the experiment, fish were raised in these aquariums for two months. The lighting schedule was configured to a 12 h cycle of light and dark, whereas fish reared in the day sunlight for 12 h and reared in the night for 12 h using Medium Bi-Pin Fluorescent Lamps with a Power of 36.0 Watt. Two air stones attached to the air pumps supported each aquarium to maintain enough aeration. On the previously formulated test diets, fish were hand-fed three times a day (6:00 a.m., 1:00 p.m., and 8:00 p.m.) with 3% of their wet body weight. Equal meals were given to each group during each feeding time. Every two days, the water column in each replica was exchanged, and well-aerated water was used (for around 33% of the water column). Fecal matter and remaining food particles were siphoned off during water exchange to avoid deterioration of the water quality parameters.

#### *2.5. Water Quality Measurements*

Using the HI9829 multiparameter equipment (HANNA instruments, EGY), the daily evaluation of dissolved oxygen (DO) and pH values was performed. A water thermometer measured the water's temperature (◦C). A pH meter was used to check the pH levels (HANNA 8424, Hungary). A DREL portable spectrophotometer 2000 was used to measure the amounts of nitrite (NO2; mg/L) and unionized ammonia (NH3; mg/L) (HACH Co., Loveland, CO, USA). DO, water temperature, pH, NO2, and NH3 values were kept at 6.6 ± 0.4 mg/L, 29.0 ± 1 ◦C, 8.0 ± 0.05, 0.03 ± 0.01, and 0.02 ± 0.01 mg/L, respectively.

#### *2.6. Determination of Growth, Feed Utilization, and Survival Rates*

The final fish weight (FW) was determined by dividing the total fish weight in each aquarium by their number. The following equations were used to estimate the growth, feed utilization and fish survival:

Weight gain (WG; g) = Final weight (FW) − Initial weight (IW);

Weight gain percentage (WG%) = 100 × (FW − IW)/IW;

Specific growth rate (SGR; %/day) = [Ln FW − Ln IW] × 100/60 (experiment period in days);

Feed intake (FI; g feed/fish/day) = The total amount of diets used by fish all over the whole feeding period;

Feed conversion ratio (FCR) = FI (g)/WG (g);

Fish survival rates (SR; %) = [Fish number per group after the feeding trial/their initial number] × 100.

#### *2.7. Proximate Composition of the Whole-Body and Amino Acid Retention*

After the end of the feeding trial, three fish per replicate (9 fish per group) were frozen at −20 ◦C then transferred to the laboratory to analyze the whole-body proximate chemical analysis and amino acid retention. According to AOAC procedures, the proximate chemical composition of the entire fish body, including crude protein (CP), moisture (%), ether extract (EE), and ash (%), was assessed [34]. The composition analysis of the essential amino acids (EAAs) and non-essential amino acids (% of total amino acid) retention in the

whole-body of the treated fish was conducted using Amino Acid Analyzer (SupNIR-2700 series) following the guidelines provided by the manufacturer.

#### *2.8. Sampling Procedures*

All fish groups fasted for one day after the feeding trial so that blood and liver samples could be taken at a uniform time. Clove oil (50 μL/L; Algomhuria company, Alexandria, Egypt) was used to induce anesthesia. The serum was separated from blood samples. Liver samples and homogenates were taken from fish in a sterile environment.

#### Serum Collection and Preparation of Tissue Homogenates

Nine fish were sampled for each group. Blood was sampled from their caudal vessels into sterile Eppendorf tubes without using anticoagulant. Blood was centrifuged at 3000 rpm for 10 min at 4 ◦C, and then the serum samples were kept at −20 ◦C until used. The liver samples (9 per group) were sampled aseptically and kept on ice. Liver samples were homogenized, centrifuged for 15 min at 5000 rpm at 4 ◦C, and then supernatants were collected in sterile test tubes and stored at −20 ◦C until they were used to measure liver antioxidants, while sediments were discarded.

#### *2.9. Serum Biochemical Assays*

Specific diagnostic kits (Biodiagnostic Co., Cairo, Egypt) were used to measure enzymatic tests at a wavelength of 540 nm for enzymes such as aspartate transaminase (AST), alanine transaminase (ALT), and alkaline phosphatase (ALP) according to the protocols outlined in [35,36]. The digestive enzymes were assessed in serum samples by diagnostic kits (Cusabio Biotech Co. Ltd., Wuhan, China) following the procedures provided by the supplier. Amylase, lipase, and protease enzyme activities were determined by the methods represented in [37–39]. Following the protocols outlined, blood urea nitrogen, creatinine, and uric acid levels were measured using diagnostic kits (Biodiagnostic Co., Cairo, Egypt) [40–42].

#### *2.10. Serum Immunity Parameters*

Serum lysozyme (LZ) activity was measured using a turbidimetric technique with a *Micrococcus lysodeikticus* suspension (Sigma-Aldrich, St. Louis, MI, USA) [43]. LZ activities in serum were determined using a standard curve established from LZ extracted from chicken egg white (Sigma-Aldrich, USA). The serum samples' total immunoglobulin content was measured following the manufacturer's instructions of the diagnostic kits (Cusabio Biotech Co. Ltd., China) [44,45].

#### *2.11. Hepatic Antioxidant Biomarkers*

Diagnostic kits were used to check the levels of the hepatic antioxidant enzymes as catalase (CAT), superoxide dismutase (SOD), and glutathione peroxidase (GPx) (MyBioSource Inc., San Diego, CA, USA) following the manufacturer's guidelines [46–48]. Malondialdehyde (MDA; the lipid peroxidation marker) was measured in liver homogenate using the thiobarbituric acid (TBA) technique at OD 532 nm [49,50].

#### *2.12. Statistical Analysis*

Results were analysed as a function of dietary supplementation of AME using complex regression models. Best-fitted models were applied with checking inbuilt options for the normality od residuals using the D'Agostino–Pearson Omnibus Test. Adjusted R squares < 0.2 were rejected in favour of simpler models.

#### **3. Results**

#### *3.1. The Phyto-Components, Flavonoids, and Phenolics Present in AME Supplement*

GC-MS spectra of AME showed the peaks that exhibited the main constituents of AME as identified by GC-MS analysis (Supplementary Materials). GC-MS chromatogram signifies the separated bioactive constituents of AME. The compounds have been found and cross-referenced with their counterparts in the WILEY 09 and NIST14 mass spectrum databases. The compound names, retention time (RT), Area %, molecular formula, and molecular weight are described in detail (Supplementary Materials). Table 2 shows the HPLC analysis of the concentration (μg/mL), and RT of phenolics and flavonoids detected in the AME feed supplement. An ample number of flavonoids (such as rutin, catechin, quercetin, kaempferol, luteolin, chrysoeriol, naringin, and apigenin) and phenolic compounds (such as syringic acid, caffeic acid, ferulic acid, protocatechuic acid, gallic acid, ellagic acid, p-coumaric acid, resveratrol, vanillic acid, and gentisic acid) have been found in the AME that has been used in the present experiment (Table 2).


**Table 2.** HPLC results of the flavonoids and phenolics present in *Astragalus membranaceus* extract (AME) used in the present study.

#### *3.2. Growth Performance, Feed Utilization, and Survival Rates*

The growth performance, including WG (Figure 1A) and SGR (Figure 1B) of juvenile *P. hypophthalmus*, were increased linearly (R<sup>2</sup> > 0.90) with increasing AME inclusion levels, as determined by second-order polynomial regression analysis. A significant linear increase in FI (Figure 1C) and decrease in FCR (Figure 1D) following quadratic regression were found in fish fed with all AME inclusion levels compared to the AME0.0 diet. Interestingly, after the end of the feeding experiment, the fish survival rates (SR%) were not considerably altered between the experimental and the CONT groups. It was found that the SR% was 100% in all groups including the CONT group.

#### *3.3. Whole-Body Proximate Analysis and Amino acid Composition*

At the end of the trial, AME dietary supplementation did not influence the body composition, including moisture, CP, EE, and ash of juvenile *P. hypophthalmus*, manifested by no statistically significant variation and linear trend (Table 3). The effects of AMEbased diets on the essential and non-essential amino acid retention in the whole-body of juvenile *P. hypophthalmus* are exemplified in Table 4. Results showed that none of the tested diets influenced the essential and non-essential amino acid composition, manifested by no statistically significant variations (Table 4).

**Figure 1.** Growth performance including weight gain (WG) (**A**) and specific growth rate (SGR) (**B**) and feed utilization including feed intake (FI) (**C**) and feed conversion ratio (FCR) (**D**) of *P. hypophthalmus* fed diets with different AME inclusion levels. Data lines denote best fit models for the data. Equations and R square in the figures demonstrates the relationship between the measured parameters and AME inclusion levels. Individual marker indicates the number of technical replicates (n = 3).

**Table 3.** Whole body composition analysis of striped catfish juveniles fed diets supplemented with AME inclusion of levels after 2 months feeding trial.



**Table 4.** Effects of dietary AME levels on the amino acid composition (% of total amino acids) retention in the whole-body of striped catfish juveniles.

Values are depicted as means ± standard error of means of three technical replicates.

#### *3.4. Digestive Enzymes*

The effect of different supplemental AME levels on the different serum digestive enzymes of juvenile *P. hypophthalmus*, including lipase E, α-amylase E, and protease E, is presented in Figure 2. Lipase E (Figure 2A), α-amylase E (Figure 2B), and protease E (Figure 2C) activities have been increased with a significant quadratic trend in fish fed diets supplied with different AME inclusion levels when compared with the AME0.0 group.

#### *3.5. Serum Biochemical Variables*

A panel of serum biochemistry in juvenile *P. hypophthalmus* fed diets with different AME inclusion levels is presented in Figure 3. Liver enzymes (AST, ALT, and ALP) (Figure 3A–C) were improved by the dietary AME inclusion levels, manifested by a significant quadratic decreasing trend in the levels of those enzymes in fish fed with different AME levels. Meanwhile, there was no relationship between the blood urea nitrogen (Figure 3D), uric acid (Figure 3E), and creatinine (Figure 3F) and the AME inclusion levels.

**Figure 2.** Serum lipase E (**A**), α-amylase E (**B**), and protease E (**C**) of juvenile *P. hypophthalmus* fed diets with different AME inclusion levels over the feeding trial period of 56 days. Data lines denote best fit models for the data. Equations and R square in the figures demonstrate the relationship between the measured parameters and AME inclusion levels. Individual marker indicates the number of technical replicates (n = 3). Individual marker is the mean of two biological replicates.

**Figure 3.** Serum biochemistry including AST (**A**), ALT (**B**), ALP (**C**), blood urea nitrogen (**D**), uric acid (**E**), and creatinine (**F**) of juvenile *P. hypophthalmus* fed diets with different AME inclusion levels over the feeding trial period of 56 days. Data lines denote best fit models for the data. Equations and R square in the figures demonstrate the relationship between the measured parameters and AME inclusion levels. Individual marker indicates the number of technical replicates (n = 3). Individual marker is the mean of two biological replicates.

#### *3.6. Serum Immunity and Hepatic Antioxidant Activity*

The impacts of dietary AME on serum immune response and antioxidant activity of juvenile *P. hypophthalmus* is demonstrated in Figure 4. Serum immunity, including lysozyme (Figure 4A) and total Igs (Figure 4B), has been elevated considerably with a strong quadratic trend in fish fed with different AME levels when compared with the control. Hepatic MDA levels (Figure 4C) were decreased with increasing AME levels. Hepatic CAT (Figure 4D) and SOD (Figure 4E) have been increased with a quadratic trend with the inclusion of AME in fish diets. Hepatic GPx (Figure 4F) activity showed a similar result.

**Figure 4.** Serum immune responses (lysozyme and total Ig) (**A**,**B**) and hepatic antioxidant activity (**C**–**F**) of juvenile *P. hypophthalmus* fed diets with different AME inclusion levels over the feeding trial period of 56 days. Data lines denote best fit models for the data. Equations and R square in the figures demonstrate the relationship between the measured parameters and AME inclusion levels. Individual marker indicates the number of technical replicates (n = 3). Individual marker is the mean of two biological replicates.

#### **4. Discussion**

The observed enhancement in the growth performance, in terms of WG and SGR, in juvenile *P. hypophthalmus* in response to AME-based diets was the result of the improvements recorded in the FI and FCR values. The growth and feed utilization results were concomitant with digestive enzymes observations, suggesting the ability of AME to improve the functionality of diets. The growth-stimulating effects of dietary AM were also confirmed in Nile tilapia [51] and bluegill sunfish [17]. Dietary supplementation with *Astragalus* polysaccharides (ASP) also supported the growth performance of several finfish species such as *Schizothorax prenanti* [52], Nile tilapia [28,53], turbot [21], large yellow croaker [20], largemouth bass juveniles [54], crucian carp juveniles [22], and Zebrafish [23]. Differently, in a recently published paper by Sun et al. [19], AME in the diet was shown to have no discernible effect on hybrid grouper's weight increase or feed efficiency. Variations in fish species, diet, experiment design, time spent feeding, or other factors may all play a role in these disparities. The growth-promoting effects of dietary AME may be attributed to the presence of functional bioactive constituents in the AME, such as phenolic acids and flavonoids, as presented in Table 2. These important phytochemicals have already proven to increase the voluntary feeding intake and feed efficiency and improve protein retention [55]. These functional bioactive compounds could also positively enhance nutrient digestibility, which may, in turn, help improve feed utilization [56]. As reported in our current study, dietary ASP could also improve digestive enzyme activities, which subsequently helped to

increase nutrient digestibility [53]. Dietary ASP has been described to be improved the gut health of fish by boosting the intestinal mucosal barrier functions [20] and the abundance of several gut-beneficial microbial communities [23,24], which were not considered in the present study, thus deserving further studies. One hundred percent survival rates across groups at the end of the feeding trial suggested that the treated fish suffered no hazardous effects from receiving AME in their diets. A possible explanation for these results involves the functional bioactive phytochemicals included in the AME, which have potent growthpromoting, hepatoprotective, antioxidant, and immunostimulant properties [55,57,58].

The whole-body composition, including moisture, CP, EE, and ash and amino acids composition, was unchanged by the test diets, suggesting that AME supplementation did not affect the assimilation of whole-body and amino acid composition. Similarly, it was found that diets supplemented with *Yucca schidigera* or *Quillaja saponaria* did not significantly affect the whole-body composition of *P. hypophthalmus* [59,60]. Moreover, dietary AME did not considerably alter CP, moisture, EE, and ash in the hybrid grouper's whole body and muscles [19]. Our findings were also in harmony with Sun et al. [21], who found that ASP-supplemented diets did not substantially change the whole-body composition of turbot. Farag et al. [28] also found that ASP-based diets did not significantly influence the whole-body proximate composition of Nile tilapia. However, our findings did not correspond to those reported by Liu et al. [20], who found that dietary supplementation with ASP at a dose rate of 0.10 or 0.15% significantly increased the whole-body CP content in large yellow croaker larvae. Several factors may be responsible for the obvious differences in the findings, as mentioned earlier, including different supplementation doses, fish species, experimental setup, feeding duration, and others. Conversely, our results also showed that none of the tested diets had influenced the essential and non-essential amino acid composition. This result is linked to non-significant changes in CP content among experimental groups. The reasons for the insignificant differences in body proximate composition and amino acid composition of groups fed AME-supplemented diets and those fed the control diet are ill-defined and require additional investigation.

It is well-documented that enhancement in the fishes' digestive enzyme activities is associated with improving the digestibility and availability of nutrients for fish [61]. A significant improvement in lipase, α-amylase, and protease enzyme production in juvenile *P. hypophthalmus* fed with different AME inclusion levels indicated that AME-based diets might have positively enhanced the nutrient digestibility and promoted nutrient absorption capacity, underpinning the growth-promoting effects of AME. This was proven by a study in which 0.1% of AST supplementation improved intestinal metabolisms by stimulating intestinal mucosal barrier function and beneficial bacteria [62]. Similarly, supplementation with *Eleutherine bulbosa*, a medicinal herb, improved the intestinal digestive enzymes in *P. hypophthalmus* [63]. Several previously published studies reported the functional ability of ASP to enhance the digestive enzymes in several other finfish species. For instance, the intestinal protease, amylase, and lipase enzyme activities are elevated in crucian carps fed ASP-based diets [22]. A similar reflection was observed in the intestinal trypsin enzymatic activity in large yellow croaker larvae [20] and intestinal protease and amylase activities in Catla when fed ASP-based diets [25]. Furthermore, dietary ASP (0.10–0.2%) increased the midgut digestive enzymes, for instance, protease and lipase activities in Asian seabass (*Lates calcarifer*) compared with those fed the reference diet [64]. However, a recent study showed no variations in α-amylase and protease enzyme activities in Nile tilapia-fed ASPenriched diets associated with the controls [28]. These inconsistencies may be attributable to variables such as dietary supplementation dose and experiment design.

Fish liver enzymes such as AST and ALT are used as indications of liver health [65], and the increases in these enzymes in sera are considered a potential indicator of liver injury, causing leakage of these enzymes into the blood circulation from the hepatocytes [66]. Liver health was improved in *P. hypophthalmus* juveniles fed AME-based diets, manifested by a considerable decrease in liver enzymes (AST, ALT, and ALP) in fish fed diets with different AME levels compared to those provided with the reference diet. These data may suggest the

hepatoprotective impacts of dietary AME, suggesting the potential application of AME as a functional additive to prevent negative impact on the liver caused by intrinsic or extrinsic factors. This could be further strengthened by the study of Jia et al. [67], who observed potential hepatoprotective effects of ASP against CCl4-induced hepatic injury in common carp by inhibiting the elevation of AST, ALT, and lactate dehydrogenase (LDH) enzymes in the hepatocytes. Another study found a similar dietary effect of ASP on the serum ALT and AST enzyme levels in crucian carp juveniles [22]. The functional flavonoids in AME might have potential effects on the improvement in liver functions of *P. hypophthalmus* juveniles. These flavonoids conferred effective hepatoprotective functions [68]. Moreover, an earlier study also reported that *A. membranaceus* root had potent hepato-protective effects and protected hepatic cells from pathological injuries [69]. Several other reports have demonstrated the potential hepatoprotective effects of polysaccharides in *A. membranaceus* [13,67,70]. Another theory showed that AM has potent antioxidant constituents such as astragalosides, flavonoids, and polysaccharides, effectively preventing tissue injury via their antioxidant mechanisms [71]. Jin et al. have reviewed that ASP could increase the enzymatic antioxidant activities, which helps to limit and eliminate oxidative stress caused by free radicals and oxygen radicals [13].

Similar to liver enzymes, blood urea nitrogen, an indicator of the occurrence of renal damage and gill dysfunction [72], was improved by dietary AME, indicating the optimistic impacts of dietary AST supplementation on the kidney functions of the treated fish with no renal injuries. As far as we know, there were no previous reports on the effects of AME on the fish's kidney functions. *A. membranaceus* root has been described as used in treating kidney diseases in Chinese medicine [73]. It also has renal protective effects against nephropathy via the modulation of kidney function biomarkers in the blood [74]. In animal models, the positive renal protective effect of *A. membranaceus* root has been closely associated with the presence of astragalosides (astragalus saponins); they are most wellknown for their ability to protect renal tubules from damage caused by free radicals [75]. However, our previously published study showed that kidney function markers such as creatinine, uric acid, and blood urea nitrogen were not significantly altered in common carp fingerlings fed diets supplemented with *Origanum vulgare* essential oil [65]. Apart from the beneficial effect of AME on enzymes associated with liver and kidney function, further research should consider the histological microstructure of the liver and kidney in relation to stress-relevant gene expression to better understand the actual modes of action of phytochemicals present in AME.

Lysozyme, a mucolytic enzyme excreted by leukocytes, can activate leukocytes and macrophages to lyse the bacterial cell walls [76,77]. Total Igs are a major part of the fish's humoral immunity playing a significant role in the fish's immune system defense and are deemed a biomarker of the fish's adaptive immune responses [44]. Improved serum immunity, including lysozyme activity and total Igs in *P. hypophthalmus* groups fed diets with different AME levels compared with the CONT, suggested the immunostimulatory roles of dietary AME. It is well documented that plant herbal extracts can stimulate the immune response of *P. hypophthalmus*. For instance, it was found that dietary supplementation with *Euphorbia hirta* extract for one month considerably increased serum lysozyme and total Igs content in *P. hypophthalmus* [9]. Correspondingly, supplementing diets with *Psidium guajava* and *Phyllanthus amarus* extracts also improved the serum lysozyme and total Igs in *P. hypophthalmus* [10]. The immunomodulatory effects of dietary AME or ASP have been described in several studies in other finfish species. Ardó et al. [15] found that dietary *A. membranaceus*-based diets significantly increased serum lysozyme and total Igs in Nile tilapia. Serum lysozyme pursuits were also increased in Nile tilapia-fed ASP-enriched diets [53]. Furthermore, ASP liposome-based diets significantly increased nitric oxide production and boosted the phagocytic activities of head kidney macrophages as well as serum lysozyme activity in large yellow croaker [78]. It was reported that polysaccharides, saponins, and water decoction of AME significantly increased phagocytic activity and serum lysozyme activities of spotted maigre (*Nibea albiflora*) [79]. It was recently reported

that dietary ASP significantly increased serum lysozyme and total Igs in Nile tilapia [27]. The immunostimulatory functions of AME might be associated with the functional phytochemicals such as polyphenols, flavonoids, and phenolic acids, as mentioned above [55,80]. In addition, astragalosides present in *A. membranaceus* possess immune-boosting, antiinflammatory, and immune-regulatory effects [81]. Further challenge trials considering the immune response at a different post-challenge time should be conducted to understand the immunostimulatory effects of AME.

The enzymatic endogenous antioxidant enzymes such as CAT, SOD, and GPx can safeguard the host against oxidative stress [82,83]. MDA is a biomarker of the lipid peroxidation process [84]. An elevation in the hepatic production of CAT, GPx, and SOD enzymes coupled with the lower production of MDA in *P. hypophthalmus* fed AME-based diets with regard to those reared in the AME0.0 group suggested the antioxidant effects of dietary AME. A study by Wu et al. [51] observed similar responses in SOD, GPx, CAT, and MDA enzyme activities to low-temperature stress in the liver of Nile tilapia when fed AME powder. In a similar pattern, dietary AME increased the enzymatic antioxidant capacity (via increased SOD, GPx, and CAT enzyme activities) of bluegill sunfish exposed to cold-water stress [17]. *Astragalus* polysaccharides also modulated the antioxidant activity of several finfish species, such as Nile tilapia [53], large yellow croaker [20], and turbot [21]. Furthermore, it was reported that ASP significantly modulated CCl4-induced oxidative stress through increasing SOD enzyme activity and total antioxidant capacity (T-AOC) and decreasing MDA concentrations in the liver of common carp [67]. The antioxidant effects of dietary AME may be associated with ASP, which has potential efficacy in increasing antioxidant enzyme activities and counteracting the negative impacts of free radicals and reactive oxygen species [85]. Notably, the HPLC analysis of AME used in the present study found many flavonoids and phenolics, compounds with strong antioxidant properties [55,86–88]. We hypothesize that these phytochemicals can positively enhance the antioxidative capacity of juvenile *P. hypophthalmus*.

#### **5. Conclusions and Prospects**

In summary, feeding AME-supplemented diets for two months improved the growth performance of juvenile *P. hypophthalmus*. This was supported by improvement in feed utilization with concurrent elevated levels of digestive enzymes. In addition, inclusion of AME in feed improved immunity and antioxidant activity without impacting liver and kidney functions. The overall improvement in fish health might be attributed to the phytochemicals (flavonoids and phenolic acids) found in the AME used in the present study. In addition, dietary supplementation of 1.0–4.5 g AME/kg diet could be regarded as a promising phyto-additive to promote the health condition and welfare of farmed *P. hypophthalmus* in addition to potentially being used as a functional additive to maintain hepatorenal functions. It may also prevent liver and kidney damage caused by intrinsic and extrinsic factors. Nevertheless, more in-depth experiments and investigations are still necessary to decipher the existent roles of the phytochemicals in AME in influencing the gut health, intestinal histomorphology, and gene expression analysis of the examined fish species.

**Supplementary Materials:** The following Supplementary Materials are available for download at: https://www.mdpi.com/article/10.3390/fishes7060319/s1.

**Author Contributions:** Conceptualization, R.M.S., H.M.R.A.-L. and M.S.; methodology, M.S. and H.A.A.; software, M.A.K. and R.M.S.; validation, H.A.A., M.S. and M.R.C.; formal analysis, M.R.C.; investigation, H.M.R.A.-L. and M.S.; resources, M.A.K. and M.S.; data curation, M.R.C.; writing original draft preparation, H.M.R.A.-L. and M.R.C.; writing—review and editing, H.M.R.A.-L. and M.R.C.; visualization, H.M.R.A.-L.; supervision, R.M.S. and H.A.A.; project administration, H.M.R.A.- L. and R.M.S.; funding acquisition, M.A.K. and R.M.S. Each author has contributed equally to this work. All authors have read and agreed to the published version of the manuscript.

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

**Institutional Review Board Statement:** The experimental methods, procedure, and fish rearing were approved by the Faculty of Veterinary Medicine, Kafrelsheikh University, Egypt. This study was conducted according to the guidelines of the Local Experimental Animal Care Committee, Faculty of Veterinary Medicine, Kafrelsheikh University, Egypt.

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** Data are available upon request.

**Acknowledgments:** All authors thank Basem Elkhayat, The Fish and Shrimp Consultant at Fish hatchery, Borg El Arab, Alexandria, Egypt, for his aid and support throughout fish rearing and experimental work.

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

#### **References**


## *Article* **Comparison of Three Artificial Diets for the Larviculture of Giant Kokopu ( ¯** *Galaxias argenteus***)**

**William McKay 1,\* and Andrew Jeffs 1,2**


**\*** Correspondence: wjgmckay@gmail.com

**Abstract:** The selection of artificial feed is critical to the success of larviculture of fin fish and requires knowledge of the varied species-specific dietary and nutritional requirements. With the emergence of commercial aquaculture of giant kokopu, ¯ *Galaxias argenteus*, there is a need to understand the species-specific needs for artificial feeds in larviculture. Consequently, this study compares three commercial artificial dry feeds; Otohime, Artemac and O.range on the growth of recently weaned giant kokopu. Larvae fed with Otohime outperformed both Artemac and O.range treatments by ¯ achieving the highest wet weight after 67 days, greater by at least 47% on average than both Artemac and O.range. These differences in larval performance are likely to be due to the higher protein:energy ratio and EPA content of Otohime. High DHA and ARA in the diets in absolute terms or in relation to EPA did not result in added benefit for growth performance. This study provides an important first step in identifying the nutritional needs of larval giant kokopu which can assist in improving ¯ their commercial aquaculture production.

**Keywords:** aquaculture; whitebait; larval diet; formulated feed; PUFA; HUFA

**Citation:** McKay, W.; Jeffs, A.

Comparison of Three Artificial Diets for the Larviculture of Giant Kokopu ¯ (*Galaxias argenteus*). *Fishes* **2022**, *7*, 310. https://doi.org/10.3390/ fishes7060310

Academic Editor: Terje van der Meeren

Received: 16 September 2022 Accepted: 24 October 2022 Published: 28 October 2022

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2022 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 (https:// creativecommons.org/licenses/by/ 4.0/).

#### **1. Introduction**

Reliable larviculture to produce high quality juvenile fin fish in large quantities is one of the greatest challenges facing the aquaculture industry currently [1,2]. Low survival rates and poor growth of fin fish larvae are frequently major contributing factors to this production bottleneck [3,4]. Poor outcomes from larviculture is often the result of a combination of factors, such as a lack of knowledge of larval nutritional requirements, difficulties in supplying feeds that can be processed by rudimentary larval digestive systems, providing husbandry of small and fragile larval fish, as well as the appropriate management of the rapid and complex changes that fish undergo during this early phase of development [1–3,5–7]. The preparation and provision of appropriate feed is particularly critical for successful fish larviculture and typically accounts for 30–70% of total production costs [8], so that the optimization of larval nutrition and feeding regimes are priority targets for research in early stage aquaculture businesses [9].

Generally, feeding regimes for the larviculture of fin fish begin with several weeks of live food provision, followed by a period of co-feeding with live and artificial diets before larvae can be fully weaned to artificial diets [3,6,10]. Rotifers and *Artemia* are the most commonly used live feeds, but while these organisms have unlocked the potential for aquaculture of a multitude of fin fish species in the last few decades, they still impose limitations to successful commercial larviculture [7]. These limitations include their high costs, from both their purchase and the infrastructure and resources required to culture or prepare them [10]. Furthermore, the nutritional condition of live feeds can vary widely, with poor nutritional condition resulting in inferior survival and growth performance in cultured larvae [10,11]. As a result, significant effort is focus on identifying optimal artificial diets to replace live feeds.

Artificial diets provide several key advantages over live prey for larviculture. Most importantly, they can be customized to the species and age-specific nutritional needs of the animal in culture and are reliably uniform in composition [6,10]. This allows the reliability that is required for consistently delivering optimal survival and growth in commercial larviculture. Artificial diets are also far more practical, in that they are easily stored, require far fewer resources for preparation than live feeds, and are immediately dispensable when required [6].

Issues remain in preparing artificial larval feeds which provide for nutrient stability, as well as perceptibility, palatability, digestibility and nutritional value [5,6,12]. Of critical importance is identifying the specific needs of the species in culture, such that the optimal feed can be provided and economical production be achieved [13]. The provision of essential fatty acids are especially critical for larval marine fin fish [14–16]. The absolute and relative requirements of essential fatty acids, particularly EPA (eicosapentaenoic acid), ARA (arachidonic acid) and DHA (docosahexaenoic acid), are highly species-specific due to differences in their metabolic capabilities and nutritional requirements for development [13,15,17–21].

Galaxiid fishes are a family of freshwater fishes, found in cool-temperate southern hemisphere [22]. Most galaxiids maintain an entirely freshwater life cycle, however, amphidromy is common, such as in the case of the giant kokopu ( ¯ *Galaxias argenteus*) which is endemic to New Zealand [23,24]. The eggs of this species are deposited on river-banks, incubating terrestrially, until flooding rain events stimulate hatching, washing the larvae down rivers and out to sea [25–28]. After 3–6 months, giant kokopu larvae, along with ¯ larvae from four other galaxiid species, undertake a mass migration back into the freshwater habitats occupied by adults [27,29]. These mass migrations occurring in confined fresh waterways provide the opportunity for targeting their harvesting to provide e highly prized "whitebait" [30,31]. Increasingly, studies show that the abundance of the species which make up this fishery are in decline due to habitat loss and predation by introduced species, with the giant kokopu now considered threatened [ ¯ 31–33].

Protection of these galaxiid species largely involves increasing regulatory controls on wild fisheries activities, however, larviculture of giant kokopu is in the early stages of ¯ commercialisation and is proposed to reduce the reliance on harvesting threatened wild populations [28,34,35]. Optimum diet selection for larvae is a significant issue impeding further commercialisation of giant kokopu aquaculture. Recent work on this species ¯ has developed knowledge on their morphometry and energetic demands, indicating the benefits of earlier provision of larger feed particles [36,37]. Still, a significant portion of the larval production cycle (approximately 30 of 77 days) expensive live feeds are required, in large part due to the unknown nutritional of requirements of the larvae. Some early research that has been undertaken on the closely related inanga, *Galaxias maculatus*, identified that its larval diet requires high levels of alpha-linolenic fatty acid under certain salinity culture conditions [38]. However, these studies were undertaken at much lower salinities (0 and 15 ppt) than those in which giant kokopu are reared. ¯

This study aims to improve knowledge of the nutritional requirements of larval giant kokopu through the comparison of the growth performance of larvae fed on three different ¯ commercially available artificial dry feeds. The results have the potential to be useful for improving the efficiency of larviculture, which is important for securing the future commercial success of the giant kokopu aquaculture industry. ¯

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

#### *2.1. Experimental Animals*

Gametes from 80 female and 20 male giant kokopu were stripped and fertilized before ¯ being subjected to a 4 week incubation period in UV treated freshwater filtered to 1 μm at 4 ◦C. On 4 July 2016, approximately 1.2 million giant kokopu were hatched directly into ¯ one 2500 L conical commercial larval rearing tank containing UV treated water, 35 ppt and ambient temperature < 18 ◦C. At 2 DAH (days after hatching) around 9000 fish were randomly selected from the commercial tank and split evenly among nine 20 L experimental tanks. This was achieved by estimating the total number of fish per liter in the transfer vessel by careful mixing and taking random 200 mL samples and then counting the number of fish in each sample to produce a mean estimate of the total number of fish.

#### *2.2. Tank Design and Recirculation System*

Experimental tanks were made from 20 L plastic (HDPE), round, blue pails, 270 mm in diameter and 380 mm in height. The water outflow pipe was set 80 mm below the rim of the pail so that each tank held 18 L. For the first 4 weeks the outflow pipe was fitted with a banjo filter using 600 μm filter mesh to prevent the escape of giant kokopu larvae while ¯ allowing the passage of suspended particles. For the remainder of the experiment, banjo filters with 1 mm mesh were used. Surfboard wax was applied in a thick, 40 mm width strip around the inside of the tank at the water level to inhibit the climbing ability of the fish larvae.

The experimental tanks were connected to a recirculation system with the outflow from each tank being directed to a filter basket for removal of insoluble particles by a 5 μm filter mat. After passing through the filter mat, the water entered a 300 L sump containing 40 L of plastic Kaldnes-K3 media (Krüger Kaldnes AS, Norway) for biological filtration which had been preconditioned in a commercial giant kokopu RAS system for at least ¯ six months immediately prior to experimental use. Protein was skimmed from the sump manually, as required. Each day the filter mat was changed, and 100 L of seawater was removed from the sump and replaced with natural seawater, 35 ppt, filtered to 5 μm and UV sterilized.

From the sump seawater was pumped through a UV filter and then distributed into each experimental tank using 4 mm tubing connected at the surface and bottom of the tank. Water flow to each tank was 0.28 L min−<sup>1</sup> over the first 14 days of the experiment. However, inflow was suspended for 30 min during feeding events for the first 7 days. Flow rate was increased to 0.37 L min−<sup>1</sup> from 15–28 days before increasing to 0.49 L min−<sup>1</sup> for the following 14 days, and finally to 0.62 L min−<sup>1</sup> for the remainder of the experiment.

Tanks were aerated by an air-stone at the bottom of the tank producing two medium sized (0.5 mm diameter) bubbles per second for the first 28 days of the experiment. The air-stone was then changed to provide a high number of very fine bubbles for the remaining experimental period.

Illumination of experimental tanks was provided by three 58 W fluorescent tubes, suspended 100 cm above the top edge of the tanks. Light reaching the tanks was dimmed by hanging shade cloth over the tanks for 30 min either side of the lights coming on at 0745 h and off at 1800 h.

Seawater temperature was not controlled, but was measured every 6 h with a glass thermometer (Aqua One) during the experimental period and found to vary between 14 and 18 ◦C, and was consistent among all tanks. Nitrate (<5 mg/L), nitrite (<0.25 mg/L), ammonia (NH3/NH4 +) (<0.25 mg/L), carbonate hardness and pH (7–8) were measured every second day using API® test kits to ensure water treatment was maintaining suitable conditions for the larvae and was within the acceptable ranges reported for the rearing of larvae [39–42]. Water quality was never found to be outside these acceptable ranges for rearing larvae from set up through to the conclusion of the experiment and was consistent among experimental tanks due to the recirculation system.

#### *2.3. Experimental Design*

Three commercially available artificial dry feeds for larval giant kokopu were tested. ¯ Larvae were provided solely live food for the first 14 days before a prolonged weaning period after which (from 45 DAH) only artificial food was provided (Table 1).

**Table 1.** Experimental feeding regime for giant kokopu larvae showing the feed provision of each ¯ feeding event and number of feeding events per day by larvae age. Instar-I *Artemia* (in–I), instar–II *Artemia* (in–II), small particle (SP), medium particle (MP), large particle (LP). \* Indicates that for the period 36–44 DAH a ration of 2 g of instar–II *Artemia* were also administered with the fourth feed of the day only.


The first feeding treatment "OTO" used the Otohime products A (75–250 μm), B1 (250–360 μm) and B2 (360–650 μm) (Marubeni Nisshin Feed Co., Ltd., Tokyo, Japan) and are referred to as Small Particle, Medium Particle and Large Particle or "SP", "MP", and "LP", respectively (Table 2).

**Table 2.** Percent dry matter feed composition, energy content, and fatty acid profiles of the three larval diets tested in the present study.


(i) Data on percent dry matter was obtained from product manufacturer's specifications except for the moisture content of O.range which came from [43]. (ii) Digestible energy calculated with the equation: crude protein × 5.64 kcal/g + crude lipid × 9.44 kcal/g [44,45]. (iii) Gross energy calculated with the equation: crude protein × 5.64 kcal/g, crude lipid × 9.44 kcal/g [46] and carbohydrate (nitrogen free extract) × 4.11 kcal/g [44,45,47]. (iv) Protein energy ratio calculated with the equation: crude protein × 5.64 kcal/g × 0.877, crude lipid × 9.44 kcal/g × 0.982 [46], and carbohydrates (nitrogen free extract) × 4.11 kcal/g × 0.90 [44,45,47]. (v) EPA, DHA data obtained from manufacturers for Artemac and O.Range, and for Otohime as well as ARA for each product from [13,43,48].

The second treatment "ART" made use of the Artemac products 2 (100–200 μm), 3 (200–300 μm) and 4 (300–500 μm) (Aquafauna Bio-Marine, Inc., Hawthorne, CA, USA), again referred to as "SP", "MP", and "LP", respectively (Table 2).

The final feed treatment "ORA" used O.range products START-S (100–200 μm), WEAN-S (200–400 μm) and WEAN–L (300–500 μm) (INVE Aquaculture Inc., Salt Lake City, UT, USA) also referred to as "SP", "MP", and "LP", respectively (Table 2).

#### *2.4. Live Food Production*

*Artemia* cysts used to produce live feed throughout this experiment were GSL Sep-Art (INVE Aquaculture Inc., Salt Lake City, UT, USA) from the same batch.

Live feeds were administered by total wet weight, with *Artemia* being harvested and poured through a 100 μm sieve that was allowed to drip dry on a towel for 1 min and measured with electronic scales to the nearest 0.1 g.

Instar-I *Artemia* were produced by incubating cysts in natural seawater 35 ppt for 17 h at 29 ◦C with constant, vigorous aeration while exposed to light in 250 l *Artemia* cones. Instar-I *Artemia* were separated from unhatched cysts and husks with a magnet before rinsing in a 100 μm sieve with clean 35 ppt water. Instar-I *Artemia* were fed out immediately after harvesting from cysts.

Instar-II *Artemia* were produced and prepared for feeding under the same conditions, however, received a 27 h incubation with enrichment. After separation live animals were enriched in a 400 L tank for between 23–31 h using a proprietary enrichment formula that combines the commercially available instant algae products—Rotigrow Plus, Nanno 3600 and Tetraselmis 3600 (Reed Mariculture Inc., Salt Lake City, UT, USA). At the beginning of enrichment and 23 h later an aliquot of 60 mL of enrichment formula was added to the enrichment tank.

#### *2.5. Sampling of Larvae*

Two sampling events of larval giant kokopu took place in this experiment. The initial ¯ sampling took place on 27 July 2016 when larvae were 23 DAH, once larvae had been exposed to SP weaning diets for the first week to establish if initial weaning performance may set a foundation for subsequent outcomes. The second sampling was at the conclusion of the experiment on 9 September 2016 when larvae were 67 DAH to determine the overall outcome of the comparative weaning treatments.

At both sampling events, randomly sampled larvae were euthanized by placing in ice water (0 ◦C) for 20 min, measured for total length (i.e., snout to tip of tail) and body depth (i.e., center of body at the anus across to the dorsal surface) from each tank. Measurements were conducted by placing fish on 46 μm grid plastic sheets and photographing fish under a microscope using an Olympus TG-4 camera. Images were later processed using ImageJ (ver. 1.53, National Institutes of Health) to derive measurements.

For the initial sampling event from each tank three samples of 50 fish were taken at random by gently swirling the tank, collecting fish with a small jar and pouring through 300 μm mesh. Mean wet weight (WW) of fish was determined by weighing followed by mean dry weight (DW) after freeze drying, re-weighing and dividing by the total number of fish. These lyophilized samples were then used to determine total lipid and total protein content. A further 20 fish were randomly sampled from each tank to measure total length and body depth.

For the final sampling event 20 fish were randomly sampled to total length and body depth measurements. Due to reduced numbers of fish from mortality amongst all treatments early in the experiment resulting from stress of transfer of larval into the experimental tanks, three samples of 20 fish per sample were taken from each tank to undertake WW and DW measurements. These same sampled fish were then also lyophilized and used for protein and lipid analyses. The ORA treatment was an exception where one tank had sufficient numbers for only 20, 20 and 19 fish per replicate sample and another tank with only 15 fish per replicate sample. Unfortunately, it was not possible to accurately recover and record respective mortalities of larvae throughout the experiment.

#### *2.6. Specific Growth Rate*

The mean specific growth rate (SGR) for WW was determined for each treatment tank across the duration of the 44 day experimental period between initial and final sampling events using Equation (1) [49].

$$SGR = 100(\mathbf{e}^{\mathbf{g}} - 1)\tag{1}$$

where: g = (lnfinal mass − lninitial mass)/(number of days between sampling events).

#### *2.7. Lipid and Protein Composition*

Lipid was extracted from larval fish samples using a modified Bligh and Dyer [50] solvent extraction method [51]. A 1.9 mL aliquot of chloroform, methanol and deionized water mixture (ratios 2:1:0.4) was added to the lyophilized samples before being vortexed for 30 s and then left to stand for 16 h. An aliquot of 0.5 mL of 0.7% sodium chloride and 0.5 mL of chloroform were added, followed by 30 s of vortexing, then centrifuging for 10 min at 1000 rpm. The chloroform-lipid layer was removed and placed in a preweighed glass vial. The residual layer was washed with 1 mL of chloroform, followed by 30 s vortexing, centrifuging for 10 min at 1000 rpm and the chloroform-lipid removed

and added to pre-weighed glass vial. This step was repeated again using only 0.5 mL of chloroform. The glass vials were then placed in a thermal evaporator held at 39 ◦C under flowing nitrogen gas to remove the chloroform. The glass vials were then re-weighed to determine lipid mass which was then divided by the total larval sample dry mass and multiplied by 100 to provide lipid content as a percentage of dry weight (%DW). The total lipid (per larva) was determined by multiplying the lipid proportion (%DW) by the mean DW of individual larvae in the respective sample. For both the three replicate lipid proportion measures and the three replicate total lipid measurements per replicate tank were then used to determine the tank average, with the results from the three tanks per treatment averaged to give the treatment mean.

The protein content of larvae was measured using a bicinchoninic acid (BCA) assay (Micro BCA™ Protein Assay Kit, ThermoFisher Scientific, Auckland, New Zealand). After removing the lipid content the residual larval tissues were freeze dried and ground before the addition of sodium hydroxide and incubation in a water bath at 50 ◦C for 16 h. Samples were diluted and then centrifuged at 4000 rpm for 10 min at 4 ◦C. The resulting samples and a set of bovine serum albumin standards were placed into a 96 well-plate and reagents added to each well followed by reading absorbance at 562 nm. The protein content of the larvae was calculated using the standard curve as a percentage of dry weight (%DW). The mean total protein (per larva) was determined by multiplying the protein proportion (%DW) by the mean DW of larvae in the respective sample.

#### *2.8. Fatty Acid Profiles*

Fatty acid analyses were conducted on an aliquot of the total lipid previously extracted gravimetrically. The derivatization process was based on Lepage & Roy [52]. Laboratory controls were included during the derivatization process. This comprised a positive control containing 52 reference standards of FAs all with different concentrations and a negative control containing C19 and C23 FAs in the same concentration range as the samples. An extraction solution of 2 mL of methanol:toluene (4:1 *v*/*v*, Analytical Grade, Merck) containing internal standards (C19: nonadecanoic acid 0.083 mg mL−<sup>1</sup> and C23: tridecanoic acid 0.082 mg mL−1, Nu-Chek Prep., Elysian, MN, USA) was added to each sample and transferred to borosilicate tubes with Teflon-lined screw caps. Magnetic stirring bars were added to each tube. Acetyl chloride (200 mL, ECP) was added slowly, dropwise to each sample over a period of 1 min. The tubes were placed in a heating and stirring dry block at 100 ◦C for 1 h. After 1 h, the tubes were cooled in water and 5 mL of an aqueous solution of 6% potassium carbonate were added to each tube. The tubes were vortexed, then centrifuged at 3500× *g* (5 min at room temperature). The upper toluene phase was recovered and transferred to a gas chromatography (GC) vial with an insert, and a further 25% dilution was done using toluene, for analysis by GC-mass spectrometry (GC–MS) at the Auckland Science Analytical Services, at the University of Auckland. GC–MS instrument parameters were based on Kramer et al. [53]. The instrument used was an Agilent 7890B gas chromatograph coupled to a 5977C mass spectrometer with a split/splitless inlet [54]. A sample of 1 μL was injected using a CTC PAL autosampler into a glass 4 mm ID straight inlet liner packed with deactivated glass wool (Restek Sky®). The inlet temperature was 250 ◦C, in splitless mode, and the column flow was set at 1 mL min<sup>−</sup>1, with a column head pressure of 62 kPa, giving an average linear velocity of 19 cm s<sup>−</sup>1. Purge flow was set to 50 mL min−<sup>1</sup> at 1 min after injection. Column selection was based on the recommendations from the official methods for the determination of trans fat (American Oil Chemists Society—[55]). The column was a fused silica Rtx-2330, which was 100 m long, 0.25 mm internal diameter, 0.2 μm highly polar stationary phase (90% biscyanopropyl and 10% cyanopropylphenyl polysiloxane, Shimadzu). Carrier gas was instrument grade helium (99.99%, BOC). The GC oven temperature programming started isothermally at 45 ◦C for 2 min, increased by 10 ◦C min−<sup>1</sup> to 215 ◦C, held for 35 min and then increased by 40 ◦C min−<sup>1</sup> to 250 ◦C and held for 10 min. The transfer line to the mass spectrometric detector (MSD) was maintained at 250 ◦C, the MSD source at 230 ◦C and the MSD quadropole at 150 ◦C. The detector was

turned on 14.5 min into the run. The detector was run in positive-ion, electron-impact ionization mode, at 70 eV electron energy, with electron multiplier set with no additional voltage relative to the autotune value. Data were acquired at 1463 amu s−<sup>1</sup> in scan mode from 41 to 420 amu, with a detection threshold of 100 ion counts. Resulting GC-MS peaks were identified on fatty acid methyl ester mass spectral library and each FA peak was quantified using an inhouse R package (RStudio, ver. 1.2.1335). The data were screened for chromatographic retention time drift, and manual correction/integration was carried out where necessary. The data set was normalized by the response of the internal standard (nonadecanoic acid), and a blank treatment was applied to correct the baseline response. The resulting normalized peak area values were used to quantify the total of each FA using linear calibration information obtained from seven calibration curve standards. The total amount of FA measured in the lipid aliquot was then adjusted for the total lipid extract to calculate the proportional contribution of each FA to the total FAMEs.

#### *2.9. Statistical Analyses*

The initial and final mean DW, WW, total length, body depth, total lipid and protein, percent lipid and protein, fatty acids, and SGR were compared among treatments using ANOVA where parametric data assumptions were satisfied. Initial and final lipid and protein concentrations as well as fatty acid profiles and SGR were arc-sine transformed prior to analysis to correct for any data distribution bias associated with percentage data [56]. Normality and equality of variance of data were tested and confirmed using the Shapiro–Wilk's and Levene's tests prior to analyses. Where data conformed to parametric assumptions a linear mixed model ANOVA was fitted to control the random effects of the tanks in each analysis. When ANOVA identified overall experimental treatment effects the differences between pairs of individual means were identified with a Tukey's test with adjustment for false discovery. Estimated mean difference and 95% confidence intervals were calculated and are presented.

Data requirements for performing parametric tests were not met for the mean final total length and depth variables for giant kokopu larvae. Consequently, Kruskal–Wallis ¯ tests were used to compare these data and where significant differences were found, then Mann–Whitney-Wilcox post hoc comparison tests were used to compare means between treatment groups.

All statistical analyses were performed using R (RStudio, ver. 1.2.1335). All measures of variability of sampled means is reported as standard error of the mean.

#### **3. Results**

#### *3.1. Weight*

There was no difference among treatments for mean initial WW of larval fish (F(2,6) = 0.38, *p* = 0.70); i.e., OTO 7.45 ± 0.13 mg, ART 7.52 ± 0.24 mg and ORA 7.21 ± 0.10 mg (Figure 1a). However, at the final sampling the mean WW of the fish was different among the treatments (F(2,6) = 15.99, *p* < 0.01). Fish in the OTO treatment had a greater mean final WW (73.23 ± 3.14 mg), between 12.70 and 34.60 mg greater than ART (49.58 ± 2.15 mg, *p* < 0.01), and between 19.10 and 41.00 mg greater than ORA (43.16 ± 1.74 mg, *p* < 0.01) (Figure 1b). The mean final WW of the ART and ORA treatment groups were not different (*p* = 0.30).

The mean initial DW of the fish larvae was not different among treatments (F(2,6) = 1.03, *p* = 0.41), OTO 1.39 ± 0.03 mg, ART 1.32 ± 0.03 mg and ORA 1.34 ± 0.02 mg (Figure 1a). However, at the final sampling there were significant differences among treatments for DW (F(2,6) = 20.84, *p* < 0.01). OTO mean final DW (15.05 ± 0.62 mg), was between 0.26 and 0.59 times greater than ART (9.83 ± 0.42 mg, *p* < 0.01), and between 0.35 and 0.69 times greater than ORA (8.95 ± 0.37 mg, *p* < 0.01). There was no significant difference in the mean final DW between ART and ORA (*p* = 0.32) (Figure 1b).

**Figure 1.** (**a**) Mean initial wet weight (WW) and dry weight (DW) for larval giant kokopu from three ¯ different feed treatments; OTO (Otohime), ART (Artemac) and ORA (O.range) (mean ± SE). Means with different superscripts are significantly different among treatments for WW and among treatment for DW (*p* < 0.05). (**b**) Mean final wet weight (WW) and dry weight (DW) for larval giant kokopu from ¯ three different feed treatments; OTO (Otohime), Artemac (ART) and ORA (O.range) (mean ± SE). Means with different superscripts are significantly different among treatments for WW and among treatment for DW (*p* < 0.05).

#### *3.2. Length and Depth*

At the initial sampling the total length of the giant kokopu larvae was not signifi- ¯ cantly different among treatments (F(2,6) = 0.11, *p* = 0.90); i.e., OTO 15.05 ± 0.21 mm, ART 15.02 ± 0.21 mm and ORA 14.87 ± 0.18 mg (Figure 2). However, at final sampling the mean total length of larvae was different among treatments (X2 = 8.52, *p* = 0.01); i.e., OTO 27.97 ± 0.55 mm, ART 24.85 ± 0.44 mm and ORA 25.74 ± 0.58 mm (Figure 2). Mean final total length was greater in the OTO treatment than ART (*p* < 0.01), while there was no difference between OTO and ORA (*p* = 0.10), and between ART and ORA (*p* = 0.13).

The mean initial depth of the body of the larval fish was the same among treatments (i.e., 23 DAH) (F(2,6) = 0.02, *p* = 0.98); i.e., OTO 1.09 ± 0.02 mm, ART 1.09 ± 0.02 mm and ORA 1.10 ± 0.02 mm (Figure 3). At the final sampling there was a significant difference in mean body depth among treatments (X2 = 22.08, *<sup>p</sup>* < 0.01); i.e., OTO 2.55 ± 0.06 mm, ART 2.11 ± 0.06 mm and ORA 2.23 ± 0.07 mm (Figure 3). Mean final total body depth was greater in the OTO treatment than both ART (*p* < 0.01) and ORA (*p* < 0.01). There was no difference in final total body depth between ART and ORA (*p* = 0.13).

**Figure 2.** Mean initial and final total length of larval giant kokopu from three different feed treatments; ¯ OTO (Otohime), ART (Artemac) and ORA (O.range) (mean ± SE). Means with different superscripts are significantly different among treatments for mean initial total length and among treatments for mean final total length (*p* < 0.05).

**Figure 3.** Mean initial and final body depth for larval giant kokopu in three different feed treatments; ¯ OTO (Otohime), ART (Artemac) and ORA (O.range) (mean ± SE). Means with different superscripts are significantly different among treatments for mean initial body depth and among treatments for mean final body depth (*p* < 0.05).

#### *3.3. Specific Growth Rate*

There was a significant difference in the SGR among the three feed treatments (F(2,6) = 29.88, *p* < 0.01). The OTO treatment achieved a greater SGR, 5.59 ± 0.10%, than both ART,

4.42 ± 0.06% (*p* < 0.01) and ORA, 4.14 ± 0.09 % (*p* < 0.01) (Figure 4). No difference was found in the SGR between ORA and ART treatments (*p* = 0.20).

**Figure 4.** Mean specific growth rate (SGR) for larval giant kokopu for three different feed treatments; ¯ OTO (Otohime), ART (Artemac) and ORA (O.range) (mean ± SE). Means with different superscripts are significantly different (*p* < 0.05).

#### *3.4. Lipid and Protein*

At the initial sampling there was no difference in proportional lipid content of the larvae among the three feed treatments (F(2,6) = 1.30, *p* = 0.34); i.e., OTO 16.1 ± 0.2% DW, ART 15.8 ± 0.2% DW and ORA 16.0 ± 0.4% DW (Figure 5). There was no significant difference in proportional lipid content of the larvae among treatments at the final sampling (F(2,6) = 0.21, *p* = 0.82); i.e., OTO 17.0 ± 0.6% DW, ART 15.5 ± 0.8% DW and ORA 16.7 ± 0.6% DW (Figure 5).

**Figure 5.** Mean proportional lipid content of larval giant kokopu as a percentage of dry weight (DW) ¯ for at the initial and final sampling events for three different feed treatments; i.e., OTO (Otohime), ART (Artemac) and ORA (O.range) (mean ± SE). Means with different superscripts are significantly different within each of the set of three treatment means for each sampling event (*p* < 0.05).

There was also no difference in total lipid among treatments at the initial sampling (F(2,6) = 3.08, *p* = 0.12); i.e., OTO 0.24 ± 0.00 mg, ART 0.20 ± 0.00 mg and ORA 0.22 ± 0.00 mg (Figure 6). However, there was a significant difference in mean total lipid among the three feed treatments at the final sampling event (F(2,6) = 17.72, *p* < 0.01); i.e., OTO 2.43 ± 0.10 mg, ART 1.56 ± 0.05 mg and ORA 1.43 ± 0.01 mg (Figure 6). OTO accumulated between 0.51 and 1.23 mg more lipid than ART (*p* < 0.01) and between 0.64 and 1.35 mg more lipid than ORA (*p* < 0.01). There was no difference in the total lipid content of larvae between the ART and ORA treatments (*p* > 0.1).

At the initial sampling there was no difference in the proportional protein content of the larval giant kokopu among the three feed treatments (F ¯ (2,6) = 2.29, *p* = 0.18), i.e., OTO 76.8 ± 2.3% DW, ART 77.7 ± 1.5% DW and ORA 72.0 ± 1.6% DW (Figure 7). Likewise, there was no difference in the proportional protein content of larvae among the three feed treatment groups at the final sampling (F(2,6) = 0.22, *p* = 0.81); i.e., OTO 73.5 ± 2.5% DW, ART 77.6 ± 4.7% DW and ORA 73.8 ± 1.9% DW (Figure 7).

There were no differences among treatments in the mean initial total protein of the larvae (F(2,6) = 1.17, *p* = 0.37); i.e., OTO 1.07 ± 0.04 mg, ART 1.03 ± 0.03 mg and ORA 0.96 ± 0.03 mg (Figure 8). However, there was a significant difference in the total protein of larval fish at the final sampling (F(2,6) = 13.82, *p* < 0.01) (Figure 8). The mean final total protein for OTO was 10.65 ± 0.67 mg, between 1.92 and 5.18 mg more than for ART (7.42 ± 0.11 mg, *p* = 0.01), and between 2.45 and 5.98 mg more than for ORA (6.77 ± 0.17 mg, *p* = 0.01) (Figure 8). There was no difference in mean final total protein between ART and ORA (*p* = 0.50) (Figure 8).

**Figure 7.** Mean proportional protein content measured as a percentage of dry weight (DW) for larval giant kokopu at initial and final sampling for three feed treatments; i.e., OTO (Otohime), ¯ ART (Artemac) and ORA (O.range) (mean ± SE). Means with different superscripts are significantly different within each of the set of three treatment means for each sampling event (*p* < 0.05).

**Figure 8.** Mean initial and final total protein content of larval giant kokopu for three feed treatments; ¯ i.e., OTO (Otohime), ART (Artemac) and ORA (O.range) (±SE). Means with different superscripts are significantly different within each of the set of three treatment means for each sampling event (*p* < 0.05).

#### *3.5. Fatty Acid Profiles*

Thirty one fatty acids were identified across the three treatment groups with 18 fatty acids being present as >1% of total fatty acids (Table 3).

**Table 3.** Mean initial and final percent fatty acid composition (±SE) of larval giant kokopu for three ¯ feed treatments. Only fatty acids that are present at >1% are included, "Other" is the sum of all other fatty acids present at <1% (12:0, 15:0, 16:0, 16:1n-7t, 17:1-7c, 18:1n-9t, 18:3n-6c, 20:0, 20:2n-6c, 20:3n-6c, 21:0, 22:0, 24:0, 24:1n-9c). Different superscripts indicate significant differences along the row. There were no differences among initial fatty acids. ARA: arachidonic acid; DHA: docosahexaenoic acid; EPA: eicosapentaenoic acid; PUFA: polyunsaturated FA; HUFA: highly unsaturated FA.


For the initial sampling of larvae at the outset of the experiment, there were no differences for the proportion of any individual fatty acid detected among treatments (Table 3).

For the final sampling the proportions of some fatty acids differed among treatments (Table 3). ARA (F(2,6) = 6.15, *p* = 0.04) was higher in ORA (1.33% ± 0.03) than OTO (1.11% ± 0.03, *p* = 0.04) but not greater than in ART (1.27% ± 0.05, *p* = 0.41), while ART did not have a higher proportion of ARA than OTO (*p* = 0.07). EPA (F(2,6) = 56.21, *p* < 0.01) was higher in OTO (10.08% ± 0.08) than ART (7.73% ± 0.05, *p* < 0.01) and ORA (7.55 ± 0.06%, *p* < 0.01). DHA (F(2,6) = 21.37, *p* < 0.01) was proportionately higher in larvae from the ORA treatment (14.86% ± 0.11) compared to ART (11.00 ± 0.15, *p* < 0.01) and OTO (11.07% ± 0.08, *p* < 0.01).

Larvae in the ORA treatment had a higher proportion of PUFA (F(2,6) = 58.55, *p* < 0.01) (48.19% ± 0.12) than OTO (40.09% ± 0.08, *p* < 0.01) and ART (37.8% ± 0.08, *p* < 0.01). Likewise, HUFA (F(2,6) = 19.13, *p* < 0.01) were proportionately more abundant in ORA (26.66% ± 0.12), followed by OTO (24.76% ± 0.08, *p* < 0.05) and ART (22.11% ± 0.11, *p* < 0.01).

Ratios of EPA, DHA and ARA were different among treatments EPA:ARA (F(2,6) = 66.21, *p* < 0.01) and DHA:APA (F(2,6) = 62.97, *p* < 0.01), but not differ for DHA:ARA EPA (F(2,6) = 4.14, *p* = 0.07). OTO had the highest EPA:ARA ratio (9.15% ± 0.11), greater than that of ART (6.13% ± 0.07) (*p* < 0.01) and ORA (5.69% ± 0.03). ORA DHA:EPA (1.97% ± 0.03) was greater than OTO (1.10% ± 0.04) (*p* < 0.01) and ART (1.43 ± 0.06) (*p* < 0.01).

#### **4. Discussion**

The results from this experiment demonstrate the effectiveness of the artificial dry food (Otohime) for the rearing of larval giant kokopu compared to two other commercially ¯ available artificial dry foods, ART and ORA. The OTO treatment produced giant kokopu ¯ larvae with the greatest WW, DW, SGR, total body length and depth when compared to the ART and ORA treatments. The artificial dry food treatments did not have any influence on the proportions of either lipid or protein of the larvae at the end of the experiment. However, both the total lipid and protein content of the larvae were greater for the OTO treatment as a result of the larger overall size of the fish, than for fish provided with the ART and ORA feed treatments. At the end of the experiment there were no differences in any of the morphometric or biochemical parameters between the larvae provided with the ART and ORA treatments.

The intake of food particles is a crucial determining factor of the suitability of a feed products for larval fish and is affected by several characteristics. Perceptibility, capture/handling and palatability can impact the intake and the effectiveness of a feed item on the growth performance of larval fin fish [6]. Despite similar size and color of the different feed particles, perceptibility may have impacted on growth performances among the different dietary treatments in this experiment. The visual attractiveness of the feed, the speed at which feed particles sink through the water column of the tank and any chemical attractant can influence differences in the feeding response by the larvae [6]. Larval eyes are pigmented at hatch in giant kokopu and relative eye size throughout the first 77 DAH of ¯ development indicate have good visual acuity [36]. However, weaning experiments show artificial feed intake is very low in the first 21 DAH indicating that these larvae are not able to recognize these feed particles [37]. Given that the current experiment included a weaning period, the non-nutritional characteristic of the artificial feed products used will have had an impact on the growth performance of larvae, potentially to the benefit of the darker colored and wider size range of Otohime. Future experiments should seek to separate out the weaning period and the artificial only period so as to determine both the best feeds by non-nutritional and nutritional characteristics and include larger feed particle items which giant kokopu larvae have proven well adept at capturing [ ¯ 36,37]. The incremental increases in feed particle sizes and rates of water flow during the experiment, although kept consistent among all treatments, may have influenced food availability through affecting the period that food particles remained in suspension and available for consumption. The density and sinking rate of feed particles and their suspension via water turbulence are collectively important factors that influence feeding intake but are difficult to measure in practice [57,58].

Nutritional value is a key consideration in the selection of feed in fin fish aquaculture and is likely to have influenced the variation in growth performance amongst the feed treatments in this experiment. Formation of musculature accounts for the majority of mass increase in larval fish with dietary protein providing the amino acids required for muscle construction [59]. Dietary lipid is thought to be the primary source of energy, which is in significant demand in rapidly developing larval fish [59]. In order to realize maximal growth rates the optimum balance between the two macronutrients must be achieved [60]. The protein to energy ratio (P:E) is species-specific, with diets providing P:E ratios either side of the optimum will typically result in reduced growth. A low P:E can incur inadequate protein intake because consumption is also regulated by energetic requirements [8,44], while excessive dietary P:E lacks the energy required for catabolic and anabolic activity [8,44,61]. Despite the OTO feed treatment containing the lowest total protein (i.e., 51–51% versus 57% for ORA and 56% for ART) it is possible that the 123 mg kcal−<sup>1</sup> P:E is most suitable or more readily available to meet the requirements of larval giant kokopu as this treatment group achieved the greatest final mass and total ¯ protein. The ART diet has a lower P:E of 113 mg kcal−<sup>1</sup> and corresponding with lower growth in the larvae. However, the ORA diet has a similar P:E (i.e., 124 mg kcal<sup>−</sup>1) to that of OTO, but resulted in significantly lower growth performance. This outcome exemplifies

the difficulty of larval fin fish diet selection due to the complex and interacting factors characteristics of aquaculture feeds where proximate analyses and ratios alone cannot be used in isolation to determine the optimum larval diet.

Generally, the fatty acid composition of a fish is a reflection of its diet [62–65]. However, digestive capability in larval fin fish is generally poor, limited by the lack of development of organs and physiological activities required to break down feed into usable nutrients [66–68]. Digestibility is fundamental to the transformation of food into utilizable nutrients and subsequently biomass and it is likely that this has had a material impact on the growth performance of larvae in this trial.

The delivery of sufficient essential PUFAs through the diet is critical during larviculture because they are utilized in tissue construction, especially for vital nervous and optic tissues [14,69]. The levels of EPA, DHA and ARA in each of the treatment diets did not always correlate directly with levels of accumulation in larval tissues. Otohime and O.Range have equally high levels of DHA in the feed particles but larvae in the ORA treatment accumulated more DHA as a proportion of total fatty acids. Although Otohime has almost twice as much DHA as Artemac, larvae from the OTO and ART treatments accumulated similar relative levels of this fatty acid. Subsequently, DHA was likely not the limiting factor to growth in these feed treatment. However, for each treatment Final DHA levels were two orders of magnitude greater than respective Initials indicating the demand and possible lack of DHA in the early diet which largely consisted with *Artemia* nauplii. This is consistent with earlier studies where DHA appears to be absent from these live feeds and a limiting factor to growth [37]. Furthermore, it has been highlighted that, current weaning protocols in the commercial hatchery have little effect on growth performance, and that weaning could be undertaken earlier as a result may in fact also reflect the improved nutritional provision of the artificial over the live feeds [36,37].

ARA content in the three larval diets is an order of magnitude lower in the diet than EPA and DHA, and was accumulated in larval tissues to a similarly low degree. ARA is highest in O.Range, of the three diets, with larvae in the ORA treatment accumulating a higher relative proportion of ARA in tissue when compared against the other treatments. Despite the importance of ARA to early development of larvae the differences in accumulated ARA within the tissues of the larvae of each treatment indicates that higher levels of ARA are not advantageous and that the levels found in the OTO diet appear to be sufficient [15,70]. This is contrary to previous findings for other species where higher levels of ARA and lower ratios of EPA:ARA were advantageous, further indicating the likely critical importance of EPA to larval giant kokopu development [ ¯ 13,20,71]. Otohime has the highest EPA proportion of the three diets tested and larvae in the OTO treatment accumulated the highest levels of EPA relative to total fatty acid content. Both Artemac and O.Range have similar, low levels of EPA (roughly half that of Otohime) and EPA accumulated in larvae was equal at the end of the experiment for the latter two treatments. Furthermore, the larvae fed the O.Range diet which has an equal proportion of DHA to Otohime and slightly more ARA, performed as poorly as larvae fed Artemac which has much lower levels of each of these fatty acids. These observations are critical as they highlight the importance of high levels of EPA to giant kokopu growth performance, as has ¯ been noted in many other species [14,21,69,72–74].

Although not measured in this study, the leaching of nutrients from feed particles (i.e., the loss of water soluble compounds out of feeds) can have negative implications for the performance of the feed for provisioning for fish growth. Upon introduction of feed particles to rearing tanks, any rapid leaching of water soluble compounds represents a subsequent loss of the nutritional value of subsequently consumed feed particles to the larvae. Due to high surface area to volume ratio of larval fish microdiets as much as 50–95% of free amino acids and protein hydrolysates can be lost through leaching within minutes of being fed into water [5,68,75–79]. Any differences in nutrient leaching of the feeds among the three treatments may have resulted in differing biomass growth rates between feed treatments with the OTO treatment potentially being more stable and therefore better able to provide nutrients to larvae after their consumption. A further study should determine the differences in the nutrient leaching among these three weaning diets.

The experimental set up may have had an impact on the performance on the artificial feeds due to stress caused due to initially transferring larvae from the commercial tanks in which they hatched. The majority of mortality occurred in the first 48 h after larvae were transferred into the experimental tanks as has been previously observed in prior experiments with this species [80]. Furthermore, recent studies have shown that the predominance of instar-I *Artemia* is not optimal as a first feed for giant kokopu larvae, ¯ potentially compounding the post-transfer stress [80]. As such, the larvae that survived and became the subjects of this study may not best represent the performance of larvae in the commercial scale setting where the transfer in particular does not occur.

Given the scale of the experimental systems water quality was easily managed, maintaining very low ammonia levels which may not be possible at commercial scale. Closely related species *Galaxias maculatus* and *Galaxias fasciatus* have demonstrated tolerance to ammonia toxicity 1.47 and 0.80 mg/L, respectively; however, future experiments on this species should confirm the species-specific optimum ranges for giant kokopu [ ¯ 42]. Furthermore, to ensure the most commercially relevant information is attained, this should focus on the early larval rearing stage [81,82]. A critical learning from this experiment is that the use of full commercial scale larviculture systems would be advantageous for future experimentation to avoid any unnecessary impact on larvae performance. As previously mentioned, larval performance may have been impacted in this study through initial handling for tank transfers, low quality live feeds as well as the small experimental scale for the tank experiment. The latter of which can impact water flow dynamics, affecting larvae behavior, as well as water quality as a result of RAS performance, all of which is likely to impact the performance of the larvae themselves [83–86]. These variations need to be removed in order to attain the most commercially relevant information possible.

This study has confirmed marked differences in performance of larval giant kokopu ¯ fed with different commercially available feeds and that examining the differences among the nutritional status of the resulting fish and that of their diet can provide insights into basis of the observed differences. EPA appears to be more critical than DHA and ARA while a high P:E ratio is also advantageous. There are still several key aspects for future research which would lead to better understanding of the variation among these artificial diets and consequently their suitability as larval giant kokopu diets. Nutrient leaching ¯ rate on rehydration of diets may impact whether key nutrients are still available to larvae when consumed. Furthermore, understanding differences in settling rates among diets may help to identify the characteristics of artificial diets which initiates a feeding response from larval giant kokopu. Analyses should also be undertaken to better understand the ¯ digestive capabilities of these larvae. Furthermore, additional variables should be examined, particularly mortality rates, given the impact this has on total productivity of a larviculture system. However, the present study enables a baseline for a suitable diet from which future studies can build upon to further optimize growth performance and survival from dietary provisions. Given the paucity of information on galaxiid species in aquaculture more generally, these data should also act to inform the development of techniques for closely related species globally.

**Author Contributions:** Conceptualisation, W.M. and A.J.; methodology, W.M.; data analyses, W.M.; writing and editing, W.M. and A.J. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was funded by Callaghan Innovation under project NZPWH1501/PROP-47490-FELLOW. Logistic support was provided New Zealand Premium Whitebait Ltd.

**Institutional Review Board Statement:** Ethical review and approval was not required for this study under New Zealand's Animal Welfare Act 1999 because it explicitly excludes larval stages.

**Data Availability Statement:** The data presented in this study are available on request from the corresponding author. The data are not publicly available due to stakeholder privacy.

**Acknowledgments:** We would like to thank New Zealand Premium Whitebait Ltd. for the larvae and, in particular, Paul Decker and Tagried Kurwie for sharing their experiences and expertise on the larval rearing of galaxiids.

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

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