Partial characterization and ontogenetic development of pancreatic digestive enzymes in Japanese eel Anguilla japonica larvae
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- Murashita, K., Furuita, H., Matsunari, H. et al. Fish Physiol Biochem (2013) 39: 895. doi:10.1007/s10695-012-9749-3
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The pancreatic digestive enzymes, trypsin, chymotrypsin, lipase and amylase were partially characterized, and changes in their activities were examined during the initial ontogeny of Japanese eel Anguilla japonica larvae from 5 to 34 days post-hatching (dph). The pH optima of the eel larval enzymes were narrower than those other fish species; trypsin activity was highest at pH 9, chymotrypsin and amylase activities were highest at pH 7 and 8, and lipase activity was highest at pH 8 and 9. In an analysis of thermal profiles, the larval pancreatic enzymes had a high optimal temperature and high thermal stability, which are typical of fish from the tropics. At 12 and 13 dph, lipase activity and gene expression levels of trypsin (-a and -b), lipase and amylase decreased markedly, suggesting a marked change in larval metabolism at that time. These data could be useful in the development of artificial larval diets in Japanese eel.
KeywordsJapanese eelLarvaePancreatic digestive enzymesActivityGene expression
Methods for artificially producing Japanese eel Anguilla japonica larvae have not yet been fully established, even though Anguilla species are among the most important aquaculture species around the world. Japanese eel aquaculture is typically capture-based, with wild-caught glass eels collected for aquaculture seed; however, a global shortage of glass eels has become a serious problem in recent years. Tanaka et al. first succeeded in producing leptocephali (Tanaka et al. 2001), which then metamorphosed into glass eels (Tanaka et al. 2003). However, compared to wild eels, the growth rates of these captive-raised larvae were very low, possibly because their larval diet was incomplete (Tanaka et al. 2003).
Considerable research in the area of larval fish nutrition has focused on the development of artificial diets. In these studies, it is widely considered that the development of such diets requires a detailed knowledge of the digestive physiology of the early life stages of the target species (Kolkovski 2001; Moyano et al. 1996; Rønnestad et al. 2007). Consequently, numerous studies have been conducted to understand the digestive physiology during early ontogeny of fish species such as walleye pollock Theragra chalcogramma (Oozeki and Bailey 1995), red drum Sciaenops ocellatus (Lazo et al. 2000), California halibut Paralichthys californicus (Alvarez-González et al. 2006) and Atlantic cod Gadus morhua (Perez-Casanova et al. 2006). In Japanese eel larvae, several important findings have been reported regarding the development of the digestive system; for example, the synthesis of pancreatic enzymes (trypsin, lipase and amylase) is initiated within 8 days post-hatching (dph) (Kurokawa et al. 1995, 2002), and the absorption capacity of the alimentary canal is completed at 7 dph (Ozaki et al. 2006). In addition, the production of pepsinogen and ghrelin in the digestive tract starts during metamorphosis (from late phase of metamorphosis in leptocephali to the early glass eel phase; 269–311 dph), with the gastric endocrine cells (ghrelin cells) developing earlier than the exocrine cells (pepsinogen cells) (Kurokawa et al. 2011). However, no studies on the biochemical characteristics of digestive enzymes in Japanese eel larvae have been undertaken to date, and the early ontogeny of the digestive enzymes remains unclear in this species. Thus, to better understand the digestive physiology of Japanese eel larvae, this study partially characterized the pancreatic digestive enzymes and clarified the changes in digestive enzyme activities during the early stages of eel ontogeny.
Materials and methods
Experimental fish and sampling
Hatched Japanese eel larvae were produced by artificially induced spawning at the Shibushi Laboratory at the National Research Institute of Aquaculture (NRIA; Shibushi, Kagoshima, Japan) using methods developed by the NRIA group (Kagawa et al. 1997, 2012; Ohta et al. 1996, 1997). Larvae at 5 dph were transferred to the Nansei Laboratory of the NRIA (Minamiise, Mie, Japan) where they were maintained in 5 L acrylic resin bowls (water temperature: 23 °C) and reared on a shark egg-based diet developed by Tanaka et al. (2001, 2003) from 6 dph onward. The larvae were fed the diet five times per day (09:00, 11:00, 13:00, 15:00 and 17:00). Samples were taken from the tanks before the morning feeding (09:00, 16 h fasting), when the digestive tracts of the larvae were empty.
For partial characterization of the pancreatic digestive enzymes of Japanese eel larvae, 400 larvae were collected at 27 dph. The larvae were pooled and stored at −80 °C (n = 3, total 1,200 larvae) until analysis.
To evaluate the ontogenetic changes in pancreatic digestive enzyme activities, five replicates of 600 larvae (5 dph) were placed in separate tanks (total 3,000 larvae). Thirty larvae were then collected from each tank at 5, 9, 11, 12, 13, 14, 15, 20, 27 and 34 dph. The larvae from each tank were pooled and stored at −80 °C for assays of enzymatic activity (n = 5, total 150 larvae/sample). In addition, three larvae from each tank were pooled and stored in RNAlater (Ambion, Austin, TX) at −80 °C for assays of gene expression (n = 5, total 15 larvae/sample).
Partial characterization of digestive enzymes
Pancreatic enzyme activity assay
Pooled larvae were homogenized in ice-cold distilled water (10 μl/larva) using a 1.5 ml micro tube pestle homogenizer (BioMasher II, nippi, Tokyo, Japan). The homogenate was then centrifuged at 4 °C at 15,000 × g for 15 min. The supernatant was used as the crude enzyme extract in enzymatic activity assays.
The activity of trypsin (E.C. 22.214.171.124) was assayed using N-benzoyl-l-arginine-p-nitroanilide (L-BAPA, Peptide, Osaka, Japan) as a substrate according to a modification of Erlanger et al. (1961). The reaction mixture consisted of 240 μl buffer (containing 20 mM CaCl2, see “pH profile” section), 100 μl 2.4 mM L-BAPA and 50 μl of enzyme extract. Production of p-nitroaniline (pNA) was measured by monitoring the increase in absorbance at 410 nm per minute for 7 min. One unit (U) of activity was defined as the amount of enzyme that caused an increase of 1 absorbance unit at 410 nm in 1 min.
The activity of chymotrypsin (E.C. 126.96.36.199) was assayed using N-succinyl–Ala–Ala–Pro–Phe-p-nitroanilide(SAPFNA, Sigma, St. Louis, MO) as a substrate according to a modification of Geiger (1986). The reaction mixture consisted of 240 μl buffer (containing 20 mM CaCl2, see “pH profile” section), 100 μl 2.4 mM SAPFNA and 50 μl enzyme extract. Production of pNA was measured by monitoring the increase in absorbance at 410 nm per minute for 7 min. One unit (U) of activity was defined as the amount of enzyme that caused an increase of 1 absorbance unit at 410 nm in 1 min.
The activity of lipase (E.C. 3.1.1) was assayed using p-nitrophenyl myristate (PNPM, Sigma) as a substrate according to a modification of Albro et al. (1985). The reaction mixture consisted of 240 μl buffer (see “pH profile” section), 100 μl 3.5 mM PNPM (containing 0.5 % Triton X-100) and 50 μl enzyme extract. Production of p-nitrophenol (pNP) was measured by monitoring the increase in absorbance at 410 nm every minute for 7 min. One unit (U) of activity was defined as the amount of enzyme that caused an increase of 1 absorbance unit at 410 nm in 1 min.
Amylase (E.C. 188.8.131.52) activity was assayed according to a modification of Natalia et al. (2004). A 1 % starch solution was prepared for use as a substrate. Twenty-five microliters of substrate solution and 25 μl of buffer (see “pH profile” section) were added to 50 μl of enzyme extract, and the mixture was incubated for 60 min. This was followed by addition of 50 μl of dinitrosalicylic acid reagent (1 % dinitrosalicylic acid and 30 % sodium potassium tartrate in 0.4 M NaOH) and incubation in a boiling water bath for 5 min. Absorbance of a sixfold diluted reaction mixture was recorded at 540 nm, and the amount of maltose released was determined from standard curve. The activity was calculated in U and was defined as the amount (μmol) of maltose released in 1 min.
Enzyme activities were expressed as specific activity defined as mU per μg protein. Protein concentration was determined using a BCA protein assay kit (Pierce, Rockford, IL) using bovine serum albumin as a standard.
To determine the pH optima for pancreatic digestive enzymes, the activities of the enzymes were assayed over a wide pH range (2, 4, 6, 7, 8, 9, 10, 11 and 12; 100 mM KCl–HCl buffer for pH 2; 100 mM citrate–NaOH buffer for pH 4 and 6; 100 mM tris–HCl buffer for pH 7–9 and 100 mM glycine–NaOH buffer for pH 10–12) at 37 °C.
To evaluate the thermal profile of the pancreatic digestive enzymes, the optimal temperature and thermal stability were analyzed. To determine the optimal temperature, enzyme activity was assayed at different temperatures (10, 20, 30, 40, 45, 50, 60 and 70 °C). For thermal stability, enzyme extracts were initially incubated at different temperatures (10, 20, 30, 40, 45, 50, 60, 70 and 80 °C) for 10 min on a temperature-controlled aluminum block before being cooled immediately in ice water. The residual activity was then assayed at 37 °C. The buffer pH that was used for these assays were based on the results obtained in the pH assay described in “pH profile” section above; pH 8 was used for the chymotrypsin and amylase assays, and pH 9 was used for the trypsin and lipase assays.
Ontogenetic development of pancreatic digestive enzymes
Ontogenetic changes in pancreatic digestive enzyme activities were analyzed based on the results described in Sect. “Partial characterization of digestive enzymes”. Trypsin and lipase activity was measured at 40 °C and pH 9, and chymotrypsin and amylase activities were measured at 40 °C and pH 8. Enzyme activities were expressed as total activity (mU per larva) or as specific activity (mU per μg protein).
Primers designed for qPCR analysis in the present study
The results were analyzed by one-way analysis of variance (ANOVA) followed by Tukey’s multiple comparison tests using InStat 3.0 (GraphPad Software, San Diego, CA).
Ontogenetic development of pancreatic digestive enzymes
Enzyme activity of trypsin, chymotrypsin, lipase and amylase
Gene expression of trypsin (-a and -b), lipase and amylase
In this study, the pancreatic digestive enzymes in Japanese eel larvae were partially characterized and the physiological characteristics of the enzymes during early ontogeny were clarified. In general, the pancreatic digestive enzymes in fish exhibit some flexibility with respect to their pH optima (Heu et al. 1995; Lazo et al. 2007; Raae and Walther 1989). This flexibility has also been observed in adult Japanese eel, in which optimal trypsin and chymotrypsin activities have been reported to range from pH 6–9 using crude enzyme extract (Chiu and Pan 2002). Yoshinaka et al. (1985) purified two trypsins (named trypsin 1 and 2) from adult eel pancreas, and both of the purified trypsins showed highest activities only at around pH 8, suggesting that crude enzyme extract of adult eel includes some other trypsin isozymes exhibiting different pH optima. However, the pancreatic digestive enzymes of the eel larvae examined in this study exhibited very narrow pH specificities (pH 9 for trypsin, pH 7–8 for chymotrypsin and amylase and pH 8–9 for lipase). Eel larvae might have fewer digestive isozymes compared to the adult stage. These differences in the optimal pH range of digestive enzymes in adults and larvae imply that the pH specificities of these enzymes change over the course of Japanese eel development. Narrow pH ranges in digestive enzymes (protease) have also been observed in gilthead seabream Sparus aurata during early ontogeny (Moyano et al. 1996). The stomach of the Japanese eel develops during metamorphosis (269–311 dph) (Kurokawa et al. 2011), implying that ingested food flows directly into the intestine during the early larval stage. Consequently, changes in the pH specificity of digestive enzymes may be an important factor affecting larval diet and digestive physiology.
High optimum temperatures for trypsin have been reported in tuna Thunnus orientalis (de la Parra et al. 2007), T. thynnus (Stevens and McLeese 1984), T. albacores (Klomklao et al. 2006b), T. tonggol (Klomklao et al. 2006a) and yellowtail Seriola quinqueradiata (Kishimura et al. 2006b); these species show maximal activities at temperatures greater than 60 °C. In the present study, maximum trypsin activity in the eel larvae was also observed at a high temperature (70 °C). However, the optimal temperatures for most fish species are lower and range between 40 and 50 °C (Cao et al. 2000; Kishimura et al. 2006a, b, 2007, 2008; Liu et al. 2007; Lu et al. 2008; Zhou et al. 2012). Kanno et al. (2010) have reported a positive correlation between habitat temperature and the thermal profile of trypsin in some of fish species such as masu salmon Oncorhynchus masou, walleye pollock, greenling Pleurogrammus azonus, sardine Sardinops melanostictus, spotted mackerel Scomber australasicus, skipjack tuna Katsuwonus pelamis and T. albacores, which indicates that the temperature optima for trypsin are higher in tropical-zone fish than in temperate-zone and rigid-zone fish. Tsukamoto et al. (2011) recently collected eggs, newly hatched larvae and spawning-condition adults of Japanese eel near the West Mariana Ridge and determined that the temperature of the water where spawning occurred was 25–27 °C. As with trypsin, the chymotrypsin and lipase activities in the eel larvae of this study had relatively higher optimal temperatures (60 °C) than other fish species (chymotrypsin, 35–55 °C; lipase, 40–50 °C) (Borlongan 1990; Castillo-Yáñez et al. 2009; de la Parra et al. 2007; Heu et al. 1995; Kristjánsson and Nielsen 1992; Lazo et al. 2007; Nolasco et al. 2011; Yang et al. 2009). Furthermore, like T. orientalis, which inhabits the tropical-zone, eel larval amylase exhibited the highest activity at 45 °C (de la Parra et al. 2007). The activities of all of the enzymes assayed for thermal stability in this study decreased at high temperatures (trypsin, 70 °C; chymotrypsin, 70 °C; lipase, 60 °C; amylase, 60 °C), which seems higher than most other fish species (Castillo-Yáñez et al. 2009; Gjellesvik et al. 1992; Heu et al. 1995; Kanno et al. 2010; Klomklao et al. 2007; Kristjánsson and Nielsen 1992; Liu et al. 2007; Nolasco et al. 2011; Smichi et al. 2010). In our assay, thermal stability was determined after incubation for 10 min; however, in the previous studies mentioned above, an incubation period of 15–30 min was used. Since the thermal stability of enzymes does not only vary between species, but also between assay conditions, it is possible that the differences in the methods used to estimate enzyme activities may have accounted for the differences observed between studies. Using the methods described above to characterize pH and thermal profiles, the ontogenetic development of larval enzyme activity was clarified.
In general, the total activities of digestive enzymes increased with age during early larval development (Lazo et al. 2007; Caruso et al. 2001); however, the total lipase activity of eel larvae decreased temporarily at 12 and 13 dph. A similar temporal fluctuation was also observed in specific lipase activity. A marked decrease in the specific activity of digestive enzymes at specific periods during early ontogeny has also been observed in yellowtail kingfish Seriola lalandi (Chen et al. 2006), red drum (Lazo et al. 2007), white seabass Atractoscion nobilis (Galaviz et al. 2011), rock bream Oplegnathus fasciatus (He et al. 2012) and common snook Centropomus undecimalis (Jimenez-Martinez et al. 2012). Furthermore, gene expression levels of trypsin (-a and -b), lipase and amylase all decreased at 12 and 13 dph in this study, indicating that a reduction in enzyme synthesis occurred at that time. Pedersen et al. (2003) reported that the content of water-soluble proteins decreased temporarily in 11 dph eel larvae. Such temporal changes may reflect metabolic changes occurring during larval development.
In many fish species, the expression of genes for digestive enzymes has been detected at early stages of ontological development, even before mouth opening or during the embryonic stage (Darias et al. 2006; García-Gasca et al. 2006; Kortner et al. 2011; Srivastava et al. 2002; Sæle et al. 2010). In an analysis of mRNA expression of pancreatic enzymes (trypsin-a, lipase and amylase) in Japanese eel larvae, Kurokawa et al. (2002) detected the first RT-PCR signal (gel band) at 6 dph (trypsin-a and amylase) or 8 dph (lipase). Our qPCR results indicate that these genes are expressed before first feeding at 5 dph. This discrepancy in the timing of digestive enzyme expression in eel larvae could be attributed to differences in sensitivity of the assays used.
Miura et al. (2009) reported that the trypsin plays essential role in spermatogenesis in Japanese eel. Specifically, they found that when the trypsin-b gene was cloned from eel testis, immunohistochemical trypsin signals were detected in the sperm. Although the expression of trypsin-a and trypsin-b genes exhibited similar changes in the early ontogeny of eel larvae, trypsin-b gene expression levels were higher than those of trypsin-a gene in this study. In addition, since pooled samples of whole fish larvae were used for the assays in this study, it is possible that the trypsin-b gene might be expressed in non-pancreatic tissue, while the trypsin-a gene is only expressed in pancreatic cells (Kurokawa et al. 2002). Compared to expression levels at 5 dph, enzyme levels were markedly higher at 9 dph, probably because feeding was initiated at 6 dph in this study.
In this study, thermal and pH optima of digestive enzymes were determined in Japanese eel larvae, and ontogenetic development of the enzymes was analyzed. The high thermal profiles of larval eel enzymes resembled those of tropical-zone fish. The digestive enzymes of eel larvae have a very narrow pH profile. Moreover, the decrease in enzyme activity and gene expression observed during early ontogeny suggests that marked changes in larval physiology may occur at that time.
We are grateful to Dr. Y. Masuda, Mr. H. Imaizumi, Mr. T. Jinbo (Sibushi Laboratory, NRIA) and Mr. H. Hashimoto (Yaeyama Laboratory, Seikai National Fisheries Research Institute) for providing the Japanese eel larvae. This study was supported by a grant-in-aid for “Development of Sustainable Aquaculture Technology Independent of Wild Fishery Resources” from the Ministry of Agriculture, Forestry and Fisheries, Government of Japan.