Fish Physiol Biochem (2008) 34:373–384 DOI 10.1007/s10695-007-9197-7
Development of digestive enzyme activity in larvae of spotted sand bass Paralabrax maculatofasciatus. 1. Biochemical analysis C. A. Alvarez-Gonza´lez Æ F. J. Moyano-Lo´pez Æ R. Civera-Cerecedo Æ V. Carrasco-Cha´vez Æ J. L. Ortiz-Galindo Æ S. Dumas
Received: 14 August 2007 / Accepted: 16 December 2007 / Published online: 7 February 2008 Ó Springer Science+Business Media B.V. 2008
Abstract Spotted sand bass Paralabrax maculatofasciatus is a potential aquaculture species in Northwest Mexico. In the last few years it has been possible to close its life cycle and to develop larviculture technology at on pilot scale using live food, however survival values are low (11%) and improvements in growth and survival requires the study of the morpho-physiological development during the initial ontogeny. In this research digestive activity of several enzymes were evaluated in larvae, from hatching to 30 days after hatching (dah), and in live prey (rotifers and Artemia), by use of biochemical and C. A. Alvarez-Gonza´lez (&) DACBIOL Laboratorio de Acuacultura, Universidad Jua´rez Auto´noma de Tabasco, Carretera Villahermosa Ca´rdenas km 0.5, 86139 Villahermosa, Tabasco, Mexico e-mail:
[email protected] F. J. Moyano-Lo´pez Departamento de Biologı´a Aplicada, Escuela Polite´cnica Superior, La Can˜ada de San Urbano, Universidad de Almerı´a, 04120 Almeria, Spain R. Civera-Cerecedo Laboratorio de Nutricio´n Acuı´cola, Centro de Investigaciones Biolo´gicas del Noroeste (CIBNOR), Mar Bermejo 195, Col. Playa Palo de Santa Rita, 23090 La Paz, BCS, Mexico C. A. Alvarez-Gonza´lez V. Carrasco-Cha´vez J. L. Ortiz-Galindo S. Dumas Unidad Piloto de Maricultivos, CICIMAR-IPN. Av. IPN s/n, Col. Playa Palo de Santa Rita, 23096 La Paz, BCS, Mexico
electrophoretic techniques. This paper, is the first of two parts, and covers only the biochemical analysis. All digestive enzyme activities were detected from mouth opening; however the, maximum activities varied among different digestive enzymes. For alkaline protease and trypsin the maximum activities were detected from 12 to 18 dah. Acid protease activity was observed from day 12 onwards. The other digestive enzymes appear between days 4 and 18 after hatching, with marked fluctuations. These activities indicate the beginning of the juvenile stage and the maturation of the digestive system, in agreement with changes that occur during morpho-physiological development and food changes from rotifers to Artemia. All enzymatic activities were detected in rotifers and Artemia, and their contribution to enhancement the digestion capacity of the larvae appears to be low, but cannot be minimised. We concluded that the enzymatic equipment of P. maculatofasciatus larvae is similar to that of other marine fish species, that it becomes complete between days 12 and 18 after hatching, and that it is totally efficient up to 25 dah. Keywords Amylase Lipase Ontogeny Paralabrax maculatofasciatus Phosphatases Proteases Spotted sand bass Introduction Rearing of marine fish larvae is based primarily on the use of live prey during a variable period lasting
123
374
Fish Physiol Biochem (2008) 34:373–384
Table 1 Feeding schedule during rearing of spotted sand bass larvae Food item
Species
Quantity
Microalgae
Nannochloropsis oculata
300,000 cells ml-1
1–8
Non-enriched rotifers (NER)
Brachionus plicatilis
0.7–2 prey ml-1
1–3
Artemia sp.
2–10 prey ml-1 1–2 prey ml-1
4–15 12–15
2–12 prey ml-1
15–30
0.1–0.2 prey ml-1
20–30
Enriched rotifers (ER)a Non-enriched Artemia nauplii (AN)b Enriched Artemia meta-nauplii (EAMN)a,
b
b, c
Artemia juveniles (AJ) a
The enrichment used in this experiment was Super HUFA enrich (Salt Creek, Salt Lake City, Utah, USA)
b
The Artemia were obtained from Microest Artemia Cyst (Burn Philip Food, Oklahoma, USA)
c
Fed with Chaetoceros calcitrans
from two to several weeks. In recent years, infrastructure, human effort, and energy costs of maintaining live food cultures in hatcheries have increased with the demand for a greater number of juveniles generated by the growing facilities. Moreover, the dependence of hatcheries on a few Artemia suppliers is becoming a bottleneck for the development of fish rearing. In addition, it is well known that traditional live foods (rotifers and Artemia) do not quite cover the nutrient requirements of larvae (Versichelle et al. 1989). The development of artificial diets is considered an important alternative. However, the use of these feeds may be seriously limited by several factors, one of the most important being the ability of the larvae to efficiently digest the principal nutrients. As a result of the difficulty of performing digestibility studies with larvae, the most popular approach to evaluate their potential ability to digest food has been to study their digestive biochemistry. These studies have focused on quantifying the activity of the principal enzymes and assessing their variability throughout the development of the larvae. Several species have been studied, including turbot (Scophthalmus maximus), gilthead seabream (Sparus aurata), winter flounder (Pleuronectes americanus), Atlantic halibut (Hippoglossus hippoglossus), European sea bass (Dicentrarchus labrax), Senegal sole (Solea senegalensis), white seabream (Diplodus sargus), Japanese eel (Anguilla japonica), California halibut (Paralichthys californicus), and kingfish (Seriola lalandi) (Cousin et al. 1987; Moyano et al. 1996; Baglole et al. 1998; Martı´nez et al. 1999; Gawlicka et al. 2000; Zambonino-Infante and Cahu 2001; Ribeiro et al. 2002; Cara et al. 2003; Pedersen
123
Period (dah)
et al. 2003; Alvarez-Gonza´lez et al. 2005; Chen et al. 2006). The spotted sand bass (P. maculatofasciatus) is a serranid that appears to be suitable for aquaculture. It is adaptable to environmental changes, has low territoriality, and market prices are good. Reproduction of this species may be induced almost year-round under controlled laboratory conditions. The possibility of using artificial feed for the larval stages has been tested by Aviles-Quevedo et al. (1995) and Anguas-Ve´lez et al. (2000), who recorded low success with a survival below 2.5%. However, better survival of 11% was obtained by Alvarez-Gonza´lez et al. (2001) in larval culture. The morphology of the digestive system during ontogeny has been described using live prey (Pen˜a-Martı´nez et al. 2003). Considering this information, the purpose of this study was to evaluate the development of the activity of several digestive enzymes and the effect of digestive enzymes from live prey during the early ontogeny of this species by use of biochemical techniques.
Materials and methods Rearing and sampling of larvae Larvae were obtained from a spontaneous spawning of P. maculatofasciatus broodstock in the Laboratorio de Biologı´a Experimental, CICIMAR-IPN. One-dayold larvae were stocked at a density of 50 larvae l-1 in 12 glass fiber tanks (140 l) connected to a recirculating system, as described by Alvarez-Gonza´lez et al. (2001). The feeding schedule applied to the rearing of larvae is detailed in Table 1. The water parameters in
Fish Physiol Biochem (2008) 34:373–384
the experiment included salinity: 35.8 ± 1.1 ppt, dissolved oxygen: 6.4 ± 0.3 mg l-1, and temperature: 23.1 ± 1.1°C. Individual dry weight of larvae (lg individual-1) was calculated by counting and from the weight of each pooled sample of freeze-dried larvae taken on each day, by use of an analytical balance (Sartorious, Gottingen, Germany; precision of 1 9 10-4 g). Thirteen samples of feed larvae were taken from different tanks, in triplicate (the numbers in parentheses are the numbers of larvae sampled per replicate) on day 0 (embryos, 1,000), 1 (860), 2 (440), 3 (420), 4 (380), 5 (290), 7 (230), 9 (230), 12 (90), 15 (60), 18 (50), 25 (40), and 30 (20) after hatching. Additionally, five samples of starved larvae were obtained on day 0 (embryos, 950), 1 (830), 2 (370), 3 (440), and 4 (460) after hatching. All feed larvae were sampled at 7:00 am before feeding to avoid food contamination. Finally, four samples of live prey were taken—non-enriched rotifers (NER), enriched rotifers (ER), enriched Artemia meta-nauplii (EAMN), and Artemia juveniles (AJ). Larvae were anesthetized with tricaine methanesulfonate (MS 222), and all samples were rinsed with distilled water, frozen at -50°C, freeze-dried, and stored at 50°C for later analysis.
Biochemical analyses Three pooled samples of fed and starved larvae taken on each day, and live prey (rotifers and Artemia) were homogenized (30 mg ml-1) in cold 50 mmol l-1 Tris–HCl 20 mmol l-1 CaCl2 buffer, pH 7.5. Supernatant obtained after centrifugation (16,000g for 15 min at 5°C) was stored at -20°C to be used for enzyme analysis. The concentration of soluble protein in pooled samples was determined by the Bradford (1976) method using bovine serum albumin as a standard. Digestive enzyme activity was expressed as U mg protein-1; U larvae-1 or U prey-1 was estimated using the number of individuals in each pooled sample. Total alkaline protease activity was measured using casein (0.5%) in 50 mmol l-1 Tris–HCl buffer, pH 9.0, following the method of Kunitz (1947), modified by Walter (1984). Acid protease activity was evaluated according to Anson (1938) using 0.5% hemoglobin in 0.1 mmol l-1 glycine–
375
HCl, pH 2.0. One unit of enzyme activity was defined as 1 lg tyrosine released per minute. Chymotrypsin activity in extracts was determined using BTEE (N-benzoyl-L-tyrosine ethyl ester) according ´ sgeirsson and Bjarnason (1991). One unit of to A enzyme activity was defined as 1 lmol BTEE hydrolyzed per minute. The amidase activity of alkaline proteases (trypsin amidase activity) was assayed using BAPNA (N-a-benzoyl-DL-arginine 4nitroanilide hydrochloride) as substrate according to Erlanger et al. (1961). One unit of enzyme activity was defined as 1 lmol p-nitroaniline released per minute. Leucine aminopeptidase was determined using leucine p-nitroanilide (0.1 mmol l-1 in DMSO) as substrate, according to Maraux (1973). One unit of enzyme activity was defined as 1 lg nitroanilide released per minute. Determination of a-amylase activity was carried out following the Somoyi-Nelson procedure described by Robyt and Whelan (1968). One unit of activity was defined as the amount of enzyme able to produce 1 lg maltose per minute. Lipase activity was quantified using b-naphthyl caprylate according to Versaw et al. (1989). One unit of activity was defined as 1 lg naphthol released per minute. Acid and alkaline phosphatases were assayed using 4-nitrophenyl phosphate in acid citrate buffer (pH 5.5) or glycine–NaOH buffer (pH 10.1) according to Bergmeyer (1974). One unit was defined as 1 lg nitrophenyl released per minute. All assays were performed by triplicate at 37°C.
Statistical analysis Growth of larvae was determined with an exponential model Y = aebX, with previous data transformation using logarithm to the base 10, and the parameters of the model were calculated by use of the least-squares technique. A Mann–Whitney U-test was used to compare digestive enzyme activity between starved larvae, fed larvae at 2, 3, and 4 dah, and live foods. A Kruskal–Wallis test was used to compare the enzyme activity between fed larvae and live foods. A nonparametric Nemenyi test was used when significant differences were detected. All tests were carried out with the software Statistica v7.0. (StatSoft, Tulsa, OK, USA).
123
376
Results Growth during larviculture was observed to be exponential. The weights of the fed and the starved larvae were statistically different (P \ 0.05) at days 4 and 5 after hatching, with fed larvae larger than starved larvae (Fig. 1). Changes in the activity of the different proteases recorded throughout larval development are detailed in Figs. 2a–j. Alkaline proteases and trypsin specific activity (Figs. 2a, e) showed similar patterns where two main peaks were detected, a small one on day 4 after hatching and the maximum peak between days 12 and 18 after hatching. Chymotrypsin specific activity increased rapidly from day 4 and continued to increase, with fluctuation, until day 18 after hatching, when maximum activity was detected, then decreased gradually on days 25 and 30 after hatching (Fig. 2g). Acid protease specific activity (Fig. 2c) was detected at 12 dah, remained constant until 25 dah, and reached a maximum at 30 dah. Leucine aminopeptidase specific activity (Fig. 2i) had its maximum activity on day 12, and decreased rapidly from day 15 onwards. On the other hand, activity of all the individual alkaline proteases (including trypsin, chymotrypsin, and leucine aminopeptidase) increased exponentially from days 3 and 4 after hatching to a maximum value at 30 dah. Finally, individual acid protease activity showed a progressive increase, also reaching a maximum value at 30 dah (Fig. 2d). Lipase specific activity (Fig. 3a) reached a maximum (P \ 0.05) 7 dah, followed by a gradual decrease and then an increase to reach a second peak
Fig. 1 Mean dry weight (lg larva-1 ± std. dev., n = 3 pooled larvae) of spotted sand bass larvae
123
Fish Physiol Biochem (2008) 34:373–384
on day 30 after hatching. The specific amylase activity profile (Fig. 3c) showed two peaks, the first on day 4 after hatching and the second peak (maximum) on 18 dah. The specific activity of the acid phosphatases was greater than that of the alkaline phosphatases during the first day. The activity decreased for both phosphatases from 4 to 7 dah, whereas a sharp increase was detected at 12 dah when maximum activity was recorded for both digestive enzymes (Fig. 3e). Finally, the individual activity of lipase, amylase, and phosphatase were exponential (Figs. 3b, d, f), with statistically significant maximum activity recorded 30 dah. Comparison of all enzymatic activity between starved and the fed larvae on days 2, 3, and 4 after hatching (Table 2) showed significant differences (P \ 0.05). From day 2 onwards, specific lipase, amylase, trypsin, chymotrypsin, leucine aminopeptidase, and acid phosphatase activity was higher in the fed larvae. However, significant differences were only detected on day 3 after hatching for the specific alkaline protease and alkaline phosphatase activity; again, activity was maximum in fed larvae. On the other hand, individual trypsin, chymotrypsin, and lipase showed significant differences at day 2 after hatching, when fed larvae had the maximum values. For individual alkaline protease, leucine aminopeptidase, amylase, and alkaline phosphatase showed significant differences at day 3 after hatching, when activity was highest for fed larvae. Finally, individual acid phosphatase activity did not show significant differences. Digestive enzyme activity in live prey was the same as for the spotted sand bass larvae. Significant differences were detected (Table 3) for alkaline protease, amylase, trypsin, and leucine aminopeptidase between different organisms as follows: Artemia juveniles (AJ) [ enriched rotifers (ER) [ nonenriched rotifers (NER) [ enriched Artemia metanauplii (EAMN). Acid protease and acid phosphatase activity were higher in ER and NER than in EAMN and AJ. A higher lipase activity was detected for AJ, followed by NER, ER, and, finally, EAMN, while chymotrypsin activity was higher in NER followed by AJ and then EAMN. Lower activity was detected for ER. Calculating the values of digestive enzyme activity per prey, the maximum statistically significant value was obtained for Artemia juveniles (AJ) follow by enriched Artemia meta-nauplii (EAMN),
Fish Physiol Biochem (2008) 34:373–384
377
Fig. 2 Digestive enzyme activity during spotted sand bass larviculture (mean ± SD, n = 3 pooled larvae). (a) Specific alkaline protease activity, (b) individual alkaline protease activity, (c) specific acid protease activity, (d) individual acid protease activity, (e) specific trypsin activity, (f) individual trypsin activity, (g) specific chymotrypsin activity, (h) individual chymotrypsin activity, (i) specific leucine aminopeptidase activity, (j) individual leucine aminopeptidase activity. NER: non-enriched rotifers, ER: enriched rotifers, AN: Artemia nauplii, EAMN: enriched Artemia metanauplii, AJ: Artemia juvenile
non-enriched rotifers (NER), and enriched rotifers (ER). Using the data of rotifer and Artemia individual weight (Theilacker and Kimball 1984; Garı´a-Ortega et al. 1998) and consumption during the larviculture, we calculated the percentage of exogenous alkaline and acid proteases (4.3 and 2.3% from rotifers, and 1.7 and 3.2% from Artemia respectively), trypsin (1.3% from rotifers and 0.2% from Artemia), chymotrypsin (0.5% from rotifers and 0.2% from Artemia), leucine aminopeptidase (1.8% from rotifers and 1.2% from Artemia), acid and alkaline phosphatases (9.9 and 6.0% from rotifers, and 1.8 and 3.5%
from Artemia, respectively), lipase (5.3% from rotifers and 2.1% from Artemia), and amylase (9.7% from rotifers and 2.2% from Artemia).
Discussion Protease activity The presence of alkaline protease activity was detected from 2 dah onwards in P. maculatofasciatus, which is considered early compared with other
123
378
Fish Physiol Biochem (2008) 34:373–384
Fig. 3 Digestive enzyme activity during spotted sand bass larviculture (mean ± SD., n = 3 pooled larvae). (a) Specific lipase activity, (b) individual lipase activity, (c) specific amylase activity, (d) individual amylase activity, (e) specific acid and alkaline phosphatase activity, (f) individual acid and alkaline phosphatase activity. NER: non-enriched rotifers, ER: enriched rotifers, AN: Artemia nauplii, EAMN: enriched Artemia meta-nauplii, AJ: Artemia juvenile
species such as S. auratus and D. labrax (Zambonino-Infante and Cahu 1994; Moyano et al. 1996). Since the profile obtained for total alkaline protease activity coincides with that obtained for trypsin, it is possible that this enzyme is responsible for most of the total activity. Both trypsin and trypsinogen have been identified during the early stages of larval development in other species (Gawlicka et al. 2000; Cuvier-Pe´res and Kestemont 2002; Alvarez-Gonza´lez et al. 2005; Kva˚le et al. 2007). In contrast with the sharp variations in the specific activity of trypsin observed during the first 15 days of life, chymotrypsin activity was more constant from 3 dah onwards. In fact, this activity may represent the main alkaline protease secreted during the first week after hatching, a result already reported for the Dover sole (Solea solea) (Clark et al. 1986). The variations observed in the specific activity of the alkaline proteases of P. maculatofasciatus during larval development have been reported for many other species including S. aurata, D. sargus, and S. lalandi (Cara et al. 2003; Chen et al. 2006). These
123
fluctuations may be explained by the combination of three different factors: 1.
Variable induction as a response to changes in the amount and composition of the live food provided. An increase in activity was detected in 12day-old larvae when Artemia nauplii were added. This increase may be explained by the different nutritional composition of rotifers and Artemia (Moyano et al. 1996; Morais et al. 2004).
2.
Changes in the amount of soluble protein in the larvae as a result of growth that may affect the expression of activity in relation to this fraction. As reported for D. sargus (Cara et al. 2003), the pronounced decrease in the activity observed from day 20 may be partly explained by this factor. Changing expression of different enzymes as a response to variations in larval metabolism, for example yolk resorption and stomach development. It has been suggested that alkaline proteases are responsible for protein digestion
3.
larvae
-1
larvae
-1
0.2 ± 0.0
22.7 ± 1.1
0.0 ± 0.0
0.2 ± 0.0
16.2 ± 1.2
0.1 ± 0.0
5.9 ± 0.4
0.5 ± 0.1
45.0 ± 11.4
25.9 ± 0.4
0.1 ± 0.1
2.5 ± 0.4a
0.7 ± 0.1
a
123.0 ± 18.7a
0.1 ± 0.0
a
0.6 ± 0.1a
12.7 ± 1.6
2.0 ± 0.4
0.0 ± 0.0b
3.6 ± 0.7b
0.6 ± 0.1
b
100.8 ± 15.0b
0.0 ± 0.0
b
0.4 ± 0.1b
14.9 ± 1.9
b
2.4 ± 0.3b
Starved larvae 3
0.1 ± 0.0a
10.0 ± 0.8a
2.3 ± 0.1
a
324.9 ± 12.0a
0.1 ± 0.0
a
1.3 ± 0.1a
93.0 ± 2.8
a
14.1 ± 0.4a
Fed larvae
0.0 ± 0.0b
2.7 ± 0.7b
0.4 ± 0.1
b
76.5 ± 10.6b
0.0 ± 0.0
b
0.5 ± 0.2b
12.6 ± 1.7
b
2.9 ± 0.4b
Starved larvae 4
0.1 ± 0.0a
8.8 ± 1.6a
2.7 ± 0.2a
251.7 ± 16.2a
0.1 ± 0.0a
1.2 ± 0.1a
129.5 ± 2.9a
16.4 ± 0.4a
Fed larvae
0.1 ± 0.0
14.7 ± 1.2b
0.1 ± 0.0
7.6 ± 1.0
1.3 ± 0.1
155.1 ± 6.5b
34.5 ± 4.4b
0.1 ± 0.0
22.7 ± 0.6a
0.1 ± 0.0
10.9 ± 1.0
1.3 ± 0.2
195.5 ± 36.6a
51.1 ± 4.7a
0.1 ± 0.0
11.0 ± 1.2b
0.0 ± 0.0
b
8.0 ± 0.3b
1.9 ± 0.2b
316.7 ± 32.7b
48.7 ± 0.5b
0.1 ± 0.0
13.9 ± 0.6a
0.1 ± 0.0
a
19.1 ± 1.8a
6.4 ± 0.2a
970.7 ± 35.8a
78.5 ± 0.4a
0.1 ± 0.0
12.7 ± 0.7
0.0 ± 0.0
b
8.5 ± 1.7b
1.0 ± 0.2b
236.8 ± 46.8b
20.0 ± 2.8b
0.1 ± 0.0
12.6 ± 0.6
0.1 ± 0.0a
17.6 ± 1.7a
7.0 ± 1.1a
882.0 ± 82.6a
122.0 ± 11.3a
4106.1 ± 529.2b 7986.7 ± 734.5a 7835.1 ± 82.5b 11911.7 ± 53.2a 4621.7 ± 644.4b 15478.1 ± 1437.8a
0.0 ± 0.0
1.9 ± 0.4b
0.6 ± 0.1
b
73.2 ± 8.0b
0.0 ± 0.0
b
0.4 ± 0.1b
17.8 ± 2.2
2.1 ± 0.3
Fed larvae
Different superscripts in columns show significant differences (P \ 0.05) between starved and fed larvae on days 2, 3, and 4 after hatching
mU larvae
-1
U mg prot-1
Acid phosphatase
mU larvae
-1
U mg prot-1
3.5 ± 0.2
0.6 ± 0.1
mU larvae-1
Alkaline phosphatase
59.0 ± 11.8
9.9 ± 0.8
U mg prot-1
Amylase
U larvae
-1
0.0 ± 0.0
1.3 ± 0.5
1.1 ± 0.1
143.2 ± 10.6
0.0 ± 0.0
0.8 ± 0.5
12.7 ± 3.7
1.1 ± 0.3
Yolk-sac larvae Starved larvae 1 2
1040.9 ± 86.7 2182.7 ± 33.7
0.0 ± 0.0
mU 9 10-3 larvae-1
Lipase U mg prot-1
1.1 ± 0.3
1.3 ± 0.1
133.5 ± 11.7
0.0 ± 0.0
0.5 ± 0.4
15.2 ± 2.1
1.6 ± 0.2
mU 9 10-3 mg prot-1
Leucine aminopeptidase
mU 9 10
-3
mU 9 10-3 mg prot-1
Chymotrypsin
mU 9 10
-3
mU 9 10-3 mg prot-1
Trypsin
mU larvae
-1
U mg prot-1
Alkaline protease
Days after hatching (dah) Embryo 0
Table 2 Comparison of digestive enzyme activity between starved and fed larvae of spotted sand bass (from embryos until day 4 after hatching) (mean ± std. dev.)
Fish Physiol Biochem (2008) 34:373–384 379
123
380
Fish Physiol Biochem (2008) 34:373–384
Table 3 Digestive enzyme activity from live food used in spotted sand bass larviculture (mean ± std. dev.) Digestive activity
NER
ER
EAMN
AJ
Alkaline protease (U mg protein-1)
3.1 ± 0.3b
-1
(mU prey ) Acid protease
0.09 ± 0.01
(U mg protein-1)
4.3 ± 0.3b c
4.2 ± 0.4a
-1
(mU prey )
0.12 ± 0.01
0.07 ± 0.01
c
0.08 ± 0.01
23.4 ± 0.7a
b
137.2 ± 4.2a
1.9 ± 0.2
4.7 ± 0.8a d
2.3 ± 0.3b
2.3 ± 0.1bc c
3.0 ± 0.4b
b
1.9 ± 0.0
17.5 ± 2.1a
Trypsin (mU 9 10-3 mg protein-1) (mU 9 10-6 prey-1)
1.2 ± 0.2b
1.5 ± 0.2b
0.5 ± 0.1c
8.0 ± 0.6a
0.02 ± 0.00c
0.02 ± 0.00c
0.3 ± 0.0b
77.6 ± 5.8a
265.1 ± 33.1a
114.0 ± 10.6d
172.3 ± 17.5c
201.2 ± 7.0b
c
c
b
0.09 ± 0.01
2.0 ± 0.1a 44.7 ± 3.5a
Chymotrypsin (mU 9 10-3 mg protein-1) (mU 9 10
-3
-1
prey )
0.01 ± 0.00
0.01 ± 0.00
Leucine aminopeptidase (mU 9 10-3 mg protein-1) (mU 9 10
-6
-1
prey )
34.6 ± 2.0b
35.7 ± 2.1b
7.1 ± 2.1c
c
c
b
0.6 ± 0.0
0.6 ± 0.0
3.9 ± 1.1
436.5 ± 34.3a
420.3 ± 33.1c
13397.2 ± 675.6a
Lipase (U mg protein-1)
6736.7 ± 669.6b
-1
(U prey )
0.20 ± 0.02
Amylase (U mg protein-1) -1
(U prey )
d
5666.2 ± 404.7b 0.10 ± 0.01
c
645.1 ± 34.9b
774.1 ± 47.0b
c
c
0.02 ± 0.00
0.01 ± 0.00
b
0.4 ± 0.0
78.5 ± 9.8a
49.4 ± 11.9c
1442.4 ± 94.2a
b
0.4 ± 0.1
8.5 ± 0.6a
1.7 ± 0.6d
4.9 ± 1.2c
b
1.4 ± 0.1
28.5 ± 6.7a
4.8 ± 0.6b
12.6 ± 2.3a
Acid phosphatase (mU mg protein-1) (mU 9 10
-3
-1
prey )
13.0 ± 0.3b 0.40 ± 0.01
21.5 ± 0.7a c
0.40 ± 0.01
c
Alkaline phosphatase (mU mg protein-1) (mU 9 10
-3
-1
prey )
10.2 ± 1.0a 0.30 ± 0.03
11.3 ± 1.3a c
0.20 ± 0.02
c
b
3.9 ± 0.5
73.6 ± 13.3a
NER: non-enriched rotifers, ER: enriched rotifers, EAMN: Enriched Artemia meta-nauplii, AJ: Artemia juvenile. Different superscripts in columns show significant differences (P \ 0.05)
until the stomach is fully developed (Ribeiro et al. 2002). A decrease in the activity of the alkaline proteases may be related to an increase in the functionality of this organ (Cahu and Zambonino-Infante 1994; Baglole et al. 1998; Kva˚le et al. 2007). Histological studies have shown that the stomach of P. maculatofasciatus larvae is fully functional earlier than in other species (Pen˜a-Martı´nez et al. 2003). Pepsin activity was detected in this study as early as 10 dah. Activity increased and was very high by 25 dah. In the case of the enzymes involved in parietal digestion, a progressive increase should be expected together with the progressive maturation of enterocytes. Leucine aminopeptidase activity was initially
123
low but increased as a response to the presence of food. Similar results have been reported for other species such as S. maximus (Cousin et al. 1987) and D. labrax (Cahu and Zambonino-Infante 1997). The pattern of activity coincided with that of other brushborder enzymes which should indicate a maturation of enterocytes (Zambonino-Infante and Cahu 2001; Hakim et al. 2007), while the clear drop in all activity from 12 dah onwards may be related to the onset of stomach activity. True lipase activity Several studies have evaluated the activity of lipases during the larval stage of marine fish (Martı´nez et al.
Fish Physiol Biochem (2008) 34:373–384
1999; Zambonino-Infante and Cahu 1999; Gawlicka et al. 2000; Hoehne-Reitan et al. 2001; Sidell and Hazel 2002). Lipase activity was detected in P. maculatofasciatus larvae from 0 to 7 dah, after which it decreased markedly. Early activity of lipases in embryos and yolk-sac larvae of some marine fish species has been reported by several authors (Ozkizilcik et al. 1996; Ribeiro et al. 2002). Initial high lipase activity followed by a marked decrease has also been reported. Martı´nez et al. (1999) recorded a maximum lipase activity 10 dah in S. senegalensis larvae and related it to the development of the pancreas, while Cousin et al. (1987) observed lipase activity only at 15 dah in S. maximus larvae. These authors explained that lipid catabolism during the first day (before stomach formation) can only be carried out by esterases. When the digestive system is fully developed, lipid digestion is supported by true lipases. It is now widely accepted there are two types of bile salt-dependent lipase; the first type should be activated during the embryonic period and helps in the digestion of yolk lipids (Diaz et al. 2002) whereas the second type should be directly secreted by the pancreas to digest lipids from food (Buchet et al. 2000). The use of only one type of substrate (a derivative of a short-chain fatty acid) in this study may have provided a limited picture of the evolution of lipid digestion, because the progressive appearance of pancreatic lipases may have been better evidenced by the use of more suitable substrates.
381
Phosphatase activity As previously mentioned, the importance of the parietal digestive enzymes (leucine aminopeptidase, and phosphatases) indicates maturation of the enterocytes. In this sense, phosphatases have two major functions. The first is in the hydrolysis of inorganic phosphates to be used for energy production; the second is in the transportation of nutrients through membranes into cells (absorption process). When enterocytes reach their maximum capacity, phosphatase activity increases and is related to the decay of exopeptidase activity. Other enzymes such as phosphatase are also indirectly involved in digestion, facilitating the activity of other enzymes by modifying phosphate side-chains of amino acids, thereby promoting nutrient absorption by Ca2+ transport from lumen into cells (Copeland 1996). When the brushborder membrane enzymes, for example phosphatase, abruptly increase in activity during the course of the larval stage, this indicates the time the intestine matures and attains a more adult character with improved luminal digestive capacity (Ribeiro et al. 2002; Zambonino Infante and Cahu 2007). This increment was detected at day 12 after hatching for P. maculatofasciatus, which coincides with a decrease of aminopeptidase activity and the beginning of acid protease activity.
Digestive enzymes in starved larvae and live prey Amylase activity The specific activity of amylase in P. maculatofasciatus larvae showed an increase in yolk absorption from 2 dah until 18 dah, when a sharp decrease was noted. A high activity followed by a pronounced reduction has also been reported for other species (Zambonino-Infante and Cahu 1994; Moyano et al. 1996; Martı´nez et al. 1999; Buchet et al. 2000 Cuvier-Pe´res and Kestemont 2002; German et al. 2004). It has been suggested that early expression of amylase may be genetically determined (Pe´res et al. 1996) but is maintained if the composition of the diet makes it necessary (Munilla-Moran and SaboridoRey 1996). In the case of P. maculatofasciatus larvae, the absence of carbohydrates in the diet should result in a decrease in secretion.
All enzymes were present in the starved larvae of P. maculatofasciatus, although activity was low compared with in fed larvae. The early appearance of digestive enzyme activity before the addition of food has also been detected in D. labrax larvae (Zambonino-Infante and Cahu 2001). Some studies have reported the participation of exogenous enzymes from live prey. Pedersen and Andersen (1992) studied C. harengus larvae fed different foods (copepods) and microspheres, and demonstrated that the pancreatic trypsinogen content, food size, and prey dimension are the most important factors in trypsinogen secretion, especially in stomach-less larvae. In this sense, the contribution of exogenous enzymes from the live prey could be significant in improving the digestion of the larvae. For P. maculatofasciatus, the contribution in general is low for all digestive
123
382
enzymes (from 0.2% for Artemia trypsin to 9.7% for rotifer amylase). This is not in total agreement with results obtained by Gawlicka et al. (2000) who found that the exogenous contribution of amylase to H. hippoglossus larvae was of 50%, although participation in the activity of trypsin, lipase, and alkaline phosphatase from Artemia was only 10%. Additionally, Perez-Casanova et al. (2006) reported a contribution from live prey to haddock (Melanogrammus aeglefinus) and Atlantic cod (Gadus morhua) of 13 and 16.4%, respectively, for proteases and 100% for amylase activity provided by rotifers. On the other hand, it has been reported that although the activity of enzymes increases significantly after addition of live food, participation of the exogenous enzymes might be worthless (Moyano et al. 1996; Morais et al. 2004) such as observed in this research. In fact, larvae have their own genetically programmed enzymatic equipment, which is expressed in the first day of life. Kurokawa et al. (1998) worked with Sardinops melanoticus larvae and reported that the contribution of proteases from rotifers to digesting capacity was approximately 0.6% of total protease activity. Thus, the contribution of exogenous enzymes is negligible for some species whereas for others it is vital. However, it seems to be necessary to increase the innate pancreatic enzyme secretion of the larvae in both cases. In addition, rotifers and Artemia present some auto-hydrolysis after being swallowed by larvae, resulting in their nutrients becoming available. This has been demonstrated for S. aurata and D. dentex larvae (Dı´az et al. 1997). It is concluded that P. maculatofasciatus larvae are precocious organisms with some digestive activity from the time of hatching and yolk absorption. Their digestive enzymatic equipment is complete on day 12 after hatching when the stomach becomes functional and all morpho-physiological changes are concluded (Pen˜a-Martı´nez et al. 2003). By this time, the larvae have a monogastric digestive system characterized by the digestion of food in the stomach and nutrient absorption in the intestine. On the basis of these results, early weaning at day 12 after hatching could be possible. Acknowledgements This research was supported by the institutional projects: CGPI 998022, 20010825 and 20020357, by the Consejo Nacional de Ciencia y Tecnologı´a, Mexico
123
Fish Physiol Biochem (2008) 34:373–384 (CONACyT) Project 31666B, by the Universidad de Almerı´a, and by the Centro de Investigaciones Biolo´gicas del Noroeste (CIBNOR). We also want to thank the Comisio´n de Operacio´n y Fomento de Actividades Acade´micas (COFAA-IPN) and the Programa Institucional para la Formacio´n de Investigadores (PIFI-IPN).
References Alvarez-Gonza´lez CA, Ortı´z-Galindo JL, Dumas S, Martı´nezDı´az SF, Herna´ndez-Ceballos DE, Grayeb del Alamo T, Moreno-Legorreta M, Pen˜a-Martı´nez R, Civera-Cerecedo R (2001) Effect of stocking density on the growth and survival of spotted sand bass Paralabrax maculatofasciatus larvae in a closed recirculating system. J World Aquacult Soc 32:130–137 Alvarez-Gonza´lez CA, Cervantes-Trujano M, Tovar-Ramı´rez D, Conklin DE, Nolasco H, Gisbert E, Piedrahita R (2005) Development of digestive enzymes in California halibut Paralichthys californicus larvae. Fish Physiol Biochem 31:83–93 Anguas-Ve´lez BH, Civera-Cerecedo R, Contreras-Olguı´n M, Rueda-Jasso RA, Guillaume J (2000) Preliminary study on the timing of weaning of spotted sand bass (Paralabrax maculatofasciatus) larvae with a prepared diet: effects on growth and survival. J Appl Aquacult 10:1–15 Anson ML (1938) The estimation of pepsin, trypsin, papain and cathepsin with hemoglobin. J Gen Physiol 22:79–89 ´ sgeirsson B, Bjarnasson JB (1991) Structural and kinetic A properties of chymotrypsin from Atlantic cod (Gadus morhua). Comparison with bovine chymotrypsin. Comp Biochem Physiol 99B:327–335 Avile´s-Quevedo A, McGregor-Pardo U, Rodrı´guez-Ramos R, Hirales-Cosio O, Huerta-Bello M, Izawa M (1995) Biologı´a y cultivo de la cabrilla arenera Paralabrax maculatofasciatus (Steindachner, 1869). SEPESCA, Me´xico Baglole CJ, Goff GP, Wright GM (1998) Distribution and ontogeny of digestive enzymes in larval yellowtail and winter flounder. J Fish Biol 53:767–784 Bergmeyer HV (1974) Phosphatases. Methods of enzymatic analysis, vol 2. Academic Press Bradford MM (1976) A rapid and sensitive method for the quantization of microgram quantities of protein utilizing the principle of protein dye binding. Anal Biochem 72:248–254 Buchet V, Zambonino-Infante JL, Cahu CL (2000) Effect of lipid level in a compound diet on the development of red drum (Sciaenops ocellatus) larvae. Aquaculture 184:339– 347 Cahu CL, Zambonino-Infante JL (1994) Early weaning of sea bass (Dicentrarchus labrax) larvae with a compound diet: effect on digestive enzymes. Comp Biochem Physiol 109A:213–222 Cahu CL, Zambonino-Infante JL (1997) Is the digestive capacity of marine fish larvae sufficient for compound diet feeding? Aquacult Int 5:151–160 Cara JB, Moyano FJ, Ca´rdenas S, Ferna´ndez-Dı´az C, Yu´fera M (2003) Assessment of digestive enzyme activities during larval development of white bream. J Fish Biol 63:48–58
Fish Physiol Biochem (2008) 34:373–384 Chen BN, Qin JG, Kumar MS, Hutchinson W, Clarke S (2006) Ontogenetic development of the digestive system in yellowtail kingfish Seriola lalandi larvae. Aquaculture 256:489–501 Clark J, Murray KR, Stark JR (1986) Protease development in dover sole (Solea solea (L.)). Aquaculture 53:253–262 Copeland RA (1996) Structural components of enzymes. In: Enzymes, a practical introduction to structure, mechanism and data analysis. Wiley, New York, pp 35–65 Cousin JCB, Baudin-Laurencin F, Gabaudan J (1987) Ontogeny of enzymatic activities in fed and fasting turbot, Scophthalmus maximus L. J Fish Biol 30:15–33 Cuvier-Pe´res A, Kestemont P (2002) Development of some digestive enzymes in Eurasian perch larvae Perca fluviatilis. Fish Physiol Biochem 24:279–285 Dı´az M, Moyano FJ, Garcı´a-Carren˜o LF, Alarco´n FJ, Sarasquete MC (1997) Substrate-SDS-PAGE determination of protease activity through larval development in sea bream. Aquacult Inter 5:461–471 Diaz JP, Mani-Ponset L, Blasco C, Connes R (2002) Cytological detection of the main phases of lipid metabolism during early post-embryonic development in three teleost species Dicentrarchus labrax, Sparus aurata and Stizostedion lucioperca. Aquat Living Resour 15:196–178 Erlanger B, Kokowsky N, Cohen W (1961) The preparation and properties of two new chromogenic substrates of trypsin. Arch Biochem Biophys 95:271–278 Garı´a-Ortega A, Verreth JAJ, Coutteau P, Segner H, Huisman EA, Sorgeloos P (1998) Biochemical and enzymatic characterization of decapsulated cysts and nauplii of the brine shrimp Artemia at different developmental stages. Aquaculture 161:501–514 Gawlicka A, Parent B, Horn MH, Ross N, Opstad I, Torrinsen OJ (2000) Activity of digestive enzymes in yolk-sac larvae of Atlantic halibut (Hippoglossus hippoglossus): indication of readiness for first feeding. Aquaculture 184:303–314 German DP, Horn MH, Gawlicka A (2004) Digestive enzyme activities in herbivorous and carnivorous prickleback fishes (Teleostei: Stichaeidae): ontogenetic, dietary, and phylogenetic effects. Physiolog Biochem Zool 77:789–804 Hakim Y, Rowland SJ, Guy JA, Mifsud C, Uni Z, Harpaz S (2007) Effects of genetic strain and holding facility on the characteristics of alkaline phosphatase and brush border enzymes in silver perch (Bidyanus bidyanus). Aquacul Res 38:361–372 Hoehne-Reitan K, Kjorsvik E, Reitan KI (2001) Bile saltdependent lipase in larval turbot, as influenced by density and lipid content of fed prey. J Fish Biol 58:746–754 Kunitz M (1947) Crystalline soybean trypsin inhibitor II. General properties. J Gen Physiol 30:291–310 Kurokawa T, Shiraishi M, Suzuki T (1998) Quantification of exogenous proteases derived from zooplankton in the intestine of Japanese sardine (Sardinops melanoticus) larvae. Aquaculture 161:491–499 Kva˚le A, Mangor-Jensen A, Moren M, Espe M, Hamre K (2007) Development and characterisation of some intestinal enzymes in Atlantic cod (Gadus morhua L.) and Atlantic halibut (Hippoglossus hippoglossus L.) larvae. Aquaculture 264:457–468
383 Maraux S, Louvard D, Baratti J (1973) The aminopeptidase from hog-intestinal brush border. Biochem Biophys Acta 321:282–295 Martı´nez MI, Moyano FJ, Ferna´ndez-Dı´az C, Yu´fera M (1999) Digestive enzyme activity during larval development of the Senegal sole (Solea senegalensis). Fish Physiol Biochem 21:317–323 Morais S, Lacuisse M, Conceic¸a˜o LEC, Dinis MT, Rønnestad I (2004) Ontogeny of the digestive capacity of Senegalese sole (Solea senegalensis), with respect to digestion, absorption and metabolism of amino acids from Artemia. Mar Biol 145:243–250 Moyano FJ, Diaz M, Alarco´n FJ, Sarasquete MC (1996) Characterization of digestive enzyme activity during larval development of gilthead seabream (Sparus aurata). Fish Physiol Biochem 15:121–130 Munilla-Mora´n R, Saborido-Rey F (1996) Digestive enzymes in marine species. II. Amylase activities in gut from seabream (Sparus aurata), turbot (Scophthalmus maximus) and red fish (Sebastes mentella). Comp Biochem Physiol 113B:827–834 Ozkizilcik S, Chu F-LE, Place AR (1996) Ontogenetic changes of lipolytic enzymes in striped bass (Morone saxatilis). Comp Biochem Physiol 113B:631–637 Pedersen BH, Andersen KP (1992) Induction of trypsinogen in herring larvae (Clupea harengus). Mar Biol 112:559–565 Pedersen BH, Ubercha¨r B, Kurokawa T (2003) Digestive response and rates of growth in pre-leptocephalus larvae of the Japanese eel Anguilla japonica reared on artificial diets. Aquaculture 215:321–338 Pen˜a-Martı´nez R, Dumas S, Villalejo-Fuerte M, Ortı´z-Galindo JL (2003) Ontogenetic development of the digestive tract in reared spotted sand bass Paralabrax maculatofasciatus larvae. Aquaculture 219:633–644 Pe´res A, Cahu CL, Zambonino-Infante JL, Legall MM, Quazuguel P (1996) Amylase and trypsin responses to intake of dietary carbohydrate and protein depend on the developmental stage in sea bass (Dicentrarchus labrax) larvae. Fish Physiol Biochem 15:237–242 Perez-Casanova JC, Murray HM, Gallant JW, Ross NW, Douglas SE, Johnson SC (2006) Development of the digestive capacity in larvae of haddock (Melanogrammus aeglefinus) and Atlantic cod (Gadus morhua). Aquaculture 251:377–401 Ribeiro L, Zambonino-Infante JL, Cahu CL, Dinos MT (2002) Digestive enzymes profile of Solea senegalensis post larvae fed Artemia and a compound diet. Fish Physiol Biochem 27:61–69 Robyt JF, Whelan WJ (1968) The b-amylases. In: Radley JA (ed) Starch and its derivatives. Academic Press, London, pp 477–497 Sidell BD, Hazel JR (2002) Triacylglycerol lipase activities in tissues of Antarctic fishes. Polar Biol 1:517–522 Theilacker GH, Kimball AS (1984) Rotifers and copepods as larval fish foods. In: Hunter JR (ed) Synopsis of culture methods for marine fish larvae, CalCOFI Rep., vol XXV. pp 80–86 Versaw W, Cuppett SL, Winters DD, Williams LE (1989) An improved colorimetric assay for bacterial lipase in nonfat dry milk. J Food Sci 54:232–254
123
384 Versichelle D, Le´ger P, Lavens P, Sorgeloos P (1989) L’utilisation d’arte´mia. In: Barnabe´ G (ed) Aquaculture, Technique et Documentation, Lavoisier, Paris Walter HE (1984) Proteinases: methods with hemoglobin, casein and azocoll as substrates. In: Bergmeyer HJ (ed) Methods of enzymatic analysis, vol V. Verlag Chemie, Weinham Zambonino-Infante JL, Cahu CL (1994) Development and response to a diet change of some digestive enzymes in sea bass (Dicentrarchus labrax) larvae. Fish Physiol Biochem 12:399–408
123
Fish Physiol Biochem (2008) 34:373–384 Zambonino-Infante JL, Cahu CL (1999) High dietary lipid levels enhance digestive tract maturation and improve Dicentrarchus labrax larval development. J Nutr 129:1195–1200 Zambonino-Infante JL, Cahu CL (2001) Ontogeny of the gastrointestinal tract of marine fish larvae. Comp Biochem Physiol 130C:477–487 Zambonino-Infante JL, Cahu CL (2007) Dietary modulation of some digestive enzymes and metabolic processes in developing marine fish: applications to diet formulation. Aquaculture 268:98–105
