The seabass (Lates calcarifer Bloch) is one of the species with a high potential for cultivation. Apart from the culinary characteristics that endear it to the consumer, the fish is fastgrowing and euryhaline. The latter fact is seen as a valuable attribute for a species enabling its adoption for pond and cage culture under marine, brackish, and freshwater environments.
The propagation of seabass in captivity has two major objectives. The first is the production of fingerlings that are further reared in captivity to marketable size. The second objective consists of purely biological purposes such as the study of the morphology, ecology and ethology of larval fish, taxonomy, population dynamics analysis, and genetics.
This manual deals mainly with the first objective aiming to describe the technologies that have been developed and successfully tested at the National Seafarming Development Centre, Hanura, Teluk Betung Lampung, from 1987 – 1989.
The breakthrough in induced spawning of the seabass in captivity through hormone manipulation made at the Seafarming Development Centre in April 1987 has a positive impact on the long term development of Indonesian seabass culture industry. It not only solves a constraint in seed supply, but also reduces impact on wild stock juveniles. The processes involving in seabass propagation as practiced at the Centre are presented in Fig. 1.
The terminology used in this manual for the larvae at different stages of development are :
Hatchlings : This stage comprises yolk-sac individuals. It starts from Days 0–4.
Larvae : Refers to individuals after absorption of the yolk sac to the fry stage. It starts from Days 5–21.
Fry : This stage begins when the individual has completed metamorphosis, i.e., when the fish takes on the appearance of sub-adult. It starts from Days 21–40; and
Juveniles : This stage includes larger fingerlings after Day-41.
Fig. 1. General scheme of seabass culture as practiced in Indonesia.
Seabass was first described by Bloch (1790) from specimens received from Dutch merchants returning from Indonesia. His type specimen was named Holocentrus calcarifer for the similarity between the preopercular spines and thorns. The genus Lates (Cuvier and Valenciennes) was erected somewhat later in 1982 to encompass other species including the closely related Nile perch (L. niloticus Linnaeus).
Dunstan (1959) noted that there were differences in body proportions both within and between localities. His late work (Dunstan 1962) also identified morphological differences between the Papuan and Australian fish and those of previous descriptions. Variation in coloration between juvenile and adult fish and between fish in salt and fresh water was also reported by Dunstan (1962) and Reynolds (1978). Notwithstanding the variability reported to that time, Greenwood (1976) proposed only a single species of Lates throughout the IndoPacific. A full taxonomic description of Lates calcarifer (Bloch) can be found in FAO, 1974 (Appendix 1).
Success of fish breeding depends primarily on the availability of mature brooders of high quality which produce high survival and fast growing fish. Normally it takes at least 3–4 years for a hatchery to have enough brood fish for the operation. The brood stock can be obtained either from the stock raising from the juvenile stage in ponds, netcages and/or caught from the wild.
The broodstock can be reared either from hatchery produced fry or from fry collected from the natural ground. The reared brood fish would be ready for spawning activities at the end of their second year when male brood fish reach 50 cm, total length (TL) and 2.5 kg in weight and female weigh about 3.5 kg.
3.1.1. Development of brood fish in netcages (Fig. 2)
The healthy and fast growing fish fry with size 1.5 to 2.0 cm TL (total length) are selected, and kept in netcages measuring 2.0×2. 0×1.0 m with a net mesh size of 2.0 mm at the initial stocking density of 1 000 juveniles/m3. As the fry grows bigger, the stocking density per unit area is reduced and uses larger netcages with bigger mesh size net as shown in Table 1.
Table 1. Netcage specification and stocking density in relation to fish size.
| Size of fish (cm TL) | size of cage (m) | Mesh size (mm) | Stocking density (pcs/m3) |
| 1.5 – 2.0 | 2.0×2.0×1.0 | 2.0 (nylon hapa) | 1000 |
| 2.0 – 10.0 | 2.0×2.0×2.0 | 8.0 (nylon hapa) | 250 |
| 10.0 – 15.0 | 2.0×2.0×2.0 | 10.0 (polyethylene) | 125 |
| 15.0 – 20.0 | 5.0×5.0×2.0 | 12.5 (polyethylene) | 60 |
| >25 | 5.0×5.0×2.0 | 25.0 (polyethylene) | <10 kg/m3 |
Fig. 2. Floating netcages used for broodstock development
One month later, 50% of the healthy and fast growing fry are reselected for further rearing in a bigger netcage of 5.0×5.0×2.0 m. After second year, the fry are reselected again and only half of the healthy and fast growing fry are kept as the potential brood fish.
The holding netcages size as practiced at the Center measuring about 5.0×5.0×2.0 m. The mesh size varies from 4–8 cm. Stocking density is 5–7 kg of fish per ton of water. To ensure good water exchange, the holding netcage is replaced monthly with a clean net.
Trash fish is given once or twice daily. The size of the food depends on the size of fish. Fingerlings smaller than 10 cm TL are fed minced trash fish at rate of 20% body weight daily while those of 10–15 cm TL are fed on chopped pieces of around l cm length at 15% body weight daily. Fish larger than 15 cm TL are fed on larger chopped pieces of around 2.5 cm at 10% body weight daily. At the size of 1 kg, the fish are fed at 5% body weight daily. Mature fish of 3 years or more are fed at 2–3% body weight daily. The feeding rate is further reduced to 1% body weight during the spawning season. After the third year, the fish weighing between 3.5 to 4 kg in weight can be used for the gonadal development
3.1.2. Holding brood fish in broodstock pond.
Concrete tanks or earth ponds built on land are also good for brood fish development. Although these types of cultured facilities provide easier control of environmental factors, but they are much more expensive in comparison with netcages.
The size of the broodstock holding tanks as practiced in Southeast Asia region are ranging from 75–150 t (5.0×10.0×1.5 m and 10.0×10.0×1.5 m). Stocking density is 1 kg of fish/t of water. Water quality in the tank should be maintained as good as natural seawater. Suggested water quality of the concrete holding tanks is given in Table 2.
Brood fish captured from the natural ground during spawning season can obtained by gillnet of 6–10 cm mesh. The net is set in direction perpendicular to the current. The set gill net is checked for captured seabass regularly. This is to ensure that the captured fish are not left struggling for long period. The captured seabass are stocked immediately in the holding tank on board the fishing vessel. Aeration is provided either from an air blower or oxygen from a cylinder.
As all fish caught by this method usually suffer to some extent from body damage, they are treated directly in the holding tank with antibiotics or approximate chemical bath, eg. 5ppm acriflavine for 2–3 hrs. The fish should be transferred to broodstock ponds or netcages immediately after arrival in the hatchery. In general, at least 6 months are needed for the fish to recover from stress and injury and to condition to the confined environment in the broodstock ponds or netcages before they can be used for spawning.
Table 2. Suitable ranges of water quality of the broodstock tank.
| Parameters | Suitable range of values | |
| Temperature | 28 – 32 | °C |
| Salinity | 29 – 32 | ppt |
| Alkalinity (as CaCO3) | 80 – 120 | mg/1 |
| Acidity (pH) | 6.8 – 8.0 | |
| Dissolved oxygen | > 6 | mg/l |
| preferably 100% saturation | ||
| Phosphate | 10 – 100 | mg/l |
| Unionized ammonia (NH3) | < 0.5 | mg/l |
| preferably not detectable | ||
| Ionized ammonie (NH3+) | < 1.5 | mg/l |
| Aluminium sulphate (Alum) | < 80 | mg/l |
| Turbidity : | ||
| suspended solid size > 1 um | 2 – 10 | mg/l |
| suspended solid size < 1 um | 2 – 3 | mg/l |
| BOD (5 day) | maximum 3 | mg/l |
| Nitrite (NO2) | < 1 | mg/l |
| preferably not detectable | ||
| Nitrate (NO3) | < 150 | mg/l |
| Chlorine (CL2) | < 0.8 | mg/l |
Nutrition has profound effect on gonadal growth and fecundity of brood fish. Although precise information on the nutritional requirements for gonadal maturation in seabass broodstock is not available at present, it has been recognized that quality and quantity of the diet significantly affect broodstock reproductive success, spawning, hatching ability of eggs and survival rate of offspring. At the National Seafarming Development Centre, Lampung, it has been observed that broodstock fed with poor nutrition diet resulted in poor hatching ability of eggs and survival rate of larvae and fry. The condition of the brood fish was improved after good quality of fresh marine fish was given and supplemented with vitamin E once a week at 30 mg/kg of fish.
Sex of seabass can only be determined accurately in mature fish, even though the the fish has some dimorphic characters to enable sex determination. For seabass of the same age, males are generally smaller and with a more slender body and narrower body depth than the female. During the spawning season, the milt can be observed at the genital opening if slight pressure is applied on the abdomen of the mature male. The female can be recognized from the big soft round belly (Fig. 3) with the red-pink papilla extruding out at the urogenital aperture. If the female has a fully ripe egg, the egg will be visible when the abdomen is pressed by hand (Fig.4).
Fig. 3. Mature seabass brooder
In order to avoid using immature spawners, the potential female can be selected on the basis of egg size. The stages of maturity for male and female seabass are given in Table 3. Spawning takes place between stage V (fully ripe), and stage VI (spent).
Fig. 4 Checking the readiness of broodfish by pressing on abdomen of the fish
Table 3. Stage of gonodal development in seabass, Lates calcarifer.
| Stage | Female | Gonadal condition male |
| I Virgin | Glassy, rounded and ¼ the body cavity in length | Colorless thin strap lying along the blood vessel. One half body cavity in length. |
| II Maturing virgin and recovering spent. | Definite gonadal appearance. The same length as stage I | Whitish and has assumed a definite gonadal appearance. The same length as stage I. |
| III Developed gonad | Yellowish and easily detectable as female. Ovary about ⅔ of body cavity. | Whitish with gonadal appearance. |
| IV Developing | Fills half the body Eggs can be distinguished separately. | Fills half the body cavity. Whitish. |
| V Fully ripe | Eggs are separate and fill the entire body cavity | Milt fills the body cavity and can be expelled without difficulty White and sticky. |
| VI Spent | Ovary flaccid. May have some eggs remaining. | Testes thin although not as flaccid as female. Some spawner may have the testes remain and fill to one half of body cavity. |
| VII Resting | Ovaries reddish and small. Easily confused with stage II. Identification under microscope may be necessary. | Testes are small and thin. They are sharp viewed from the edge. |
(Modified from Broadhead, 1953)
The assessment of the maturity of the female fish can be made from eggs sampled by catherization from the mid-portion of the ovary, using a polyethylene cannula. The sampling process is made by inserting a polyethylene cannula 1.2 mm in diameter into the oviduct of the fish for a distance of 6–7 cm through the genital opening (Fig. 5). The eggs are placed in the test tube containing 1% formalin in 0.6% saline (sodium chloride) solution for subsequent checking of egg size and appearance. Egg diameters are measured on a glass slide using an ocular micrometer under a microscope. The diameter of egg is measured by everaging the long and short diameters. Females bearing uniform, spherical and nonadhesive eggs with a mean diameter 0.4 – 0.5 mm or more are selected for the inducement of gonadal maturation. Males are selected when white and creamy milt oozes out from the genital opening upon gentle stripping of the belly. Males yielding watery and curdled milt are not suitable for use in spawning.
Fig. 5 Checking maturity of eggs using a polyethylene cannula
In Lampung area, induced spawning by hormone manipulation can be started from November-July. Synthetic hormones, Human Chorionic Gonadotropin (HCG), Puberogen and Pregnyl and pituitary gland (PG) of common carp are commonly used. Normally 50 IU of HCG plus 0.5–1 dose of PG is the appropriate dose for the first injection to induce ovulation. The second dose 100–200 IU of HCG plus 1.5–2.0 doses of PG is applied after 12 hours. Overdoses of HCG should be avoided, since it may result in fattening of the fish, causing regression of the female's ovaries. Male (stage IV-V), milt formation can also be stimulated by injection of 0.5 dose of PG plus 25–50 IU HCG.
At the Seafarming Development Centre at Hanura, Teluk Betung Lampung, brood fish, both male and female weighing about 2–5 kg injecting first dose of 50 RU Puberogen + 250 IU HCG at 08.00 hr and following a second dose of 100 RU Puberogen + 500 IU HCG at 08 00 hr of the following day, spawned between 23 00 – 04 00 hr after the second injection.
To immobilize the brood fish during hormones injection process, a tranguilizer such as Tricane (MS - 222) or Ether at doses ranging between 1 : 12 500–1:25 000 is recommended Other fish anesthetics available in the market and suitable for general handling of fish are quinaldine, tertiary amulalcohol, choralhydrate and Phenoxethol. At the Centre, Ethylene glycol was used at dose between 1:8 000–1 :10 000. The brood fish will be immobilized within 3–5 mins.
Injection of hormones is given intramuscular on area between the lateral line, above the base of the pectoral, and dorsal fin (Fig.6) using a 5 ml disposal syringe and a gauge No.21 –22 needle. All instruments used for the operation should be clean to avoid bacterial infection
Seabass can be induced to spawn in captivity either by hormone-inducing or by natural spawning. Four weeks before spawning season, the brood fish is transferred into the spawning tank at a rate of 1 kg of fish/t of water with a sex ratio of 1:1 Due to handling during the transfer, the fish may refrain from eating one to two days.
To maintain good water quality in the spawning tank, periodic draining and refilling of water are necessary. Normally 80–100 percent of the total volume of water is changed every day. Salinity of water is maintained at 30 ppt. However, to ensure that the spawning tanks have the desirable water quality, the installation of a jet spray head and aeration system in the tank are preferred.
Fig. 6. Intramuscular injection of hormone
If the water quality and environment in the spawning tank is suitable with proper feeding, the female fish gradually appears with swollen abdomen, swimming awkwardly. Approximately 1–2 weeks before spawning, the female fish separates from the school and reduces feeding activity while the male fish continues their normal activity and active.
Natural spawning in controlled tank takes place at the same time as natural spawning ground. It starts from the beginning of November to the end of July. Spawning activity occurs between 19.00 and 23.00 hr on the first to the 8th day of the full moon or new moon. As the female approaches full maturity, there is an increase in spawning play activity. The ripe male and female swim together often turning laterally when swimming and then spawn.
If the female fish is not in a fully ripe stage, a certain dose of hormones are required for inducing the spawning. The first injection of 250 IU HCG/kg (body weight of female fish) and 50 RU Puberogen/kg (body weight of male fish) is practiced at the Seafarming Development Centre. The second injection of 500 IU HCG/kg female fish and 100 RU Puberogen/kg male fish is followed after 24 hrs. The fish showed a good response with satisfactory result.
For broodstock newly caught from the natural ground during the spawning season, the eggs and milt can be stripped and fertilized artificially. In this case, the readiness of spawner has to be checked before stripping. The healthy ripe running eggs have a diameter of 0.8 mm, contained oil globule of 0.2 mm in diameter, round with smooth surface, transparent, have light yellow color, loosely separate, float in water at 28–30 ppt salinity, and there is no yolk vesicle. The stage of ripe running eggs can be checked also by dropping a sample of eggs into a clear glass of water. The ripe running eggs show individually scattering whereas the unripe running eggs tend to group together and sink down to the bottom of the glass.
Strip eggs and milt into a dry clean tray. Stir it thoroughly with a feather. Add filtered fresh seawater salinity of about 28–30 ppt to cover all the eggs. Stir continuously for 1–3 mins, then wash it with seawater 3–4 times through a strain to remove mucus and foreign substances which could cause subsequent bacterial infection. The eggs are placed in incubation tank for further hatching.
One to two months before spawning season, the brood fish everage size of about 4 kg are transferred to spawning tanks. The stocking density is about one fish per 4–5 m3. The initial salinity in the spawning tank should be the same as the salinity in the netcages or ponds where the brood fish are kept before transferring.
After the fish get used to the tank environment, normally it takes about 2–3 days, the salinity of water in spawning tank is reduced to 20–25 ppt. Keep the broodfish in the 20– 25 ppt salinity for 7 days then gradually increase it to 30–32 ppt through daily water changing About 60–70% of water are changed daily until the salinity reached 30–32 ppt. Changing in salinity is a natural stimulation during its migration from feeding ground in the brackishwater to spawning ground in the sea environments.
On the day during full moon or new moon period, the water is reduced to about 30 cm depth at noon and exposed to the sun light for 3–4 hrs to increase the water temperature to 30–32 degree Celsius. New seawater is added to manipulate rising tide conditions and decrease the water temperature to 27–28 degree Celsius as found in normal spawning environment.
If the fish is in the right stage and good condition, it will spawn in the same night or the next after at 19 00–22 00 hr. If the fish does not respond or responds but with abnormal eggs, the injection of hormones is required.
Besides spawning in tanks on shore, the seabass can also be spawned at sea in floating netcages. For spawning in the floating netcages, the small mesh size of net is installed the night of the second dose of the inducement of the spawning to prevent the spawned eggs from being washed away by the tidal current. The brooders are not fed so long as they are in the spawning cage to prevent deterioration of water quality inside the netcages.
Major factors affecting gonadal maturation and spawning of seabass in captivity are food, water quality, salinity, stress, size of broodstock, age of brood fish and lunar cycle.
Quality and quantity of food given to brood fish affect the gonadal maturation.
Normally fresh marine fishes such as sardine, anchovy and yellow strip caranx are used as a main food for brood fish at the rate of 5% of body weight. One to two months before spawning season, the feed is reduced to 1% of body weight and fed once a day. At this period vitamin E should be given at a dose of 30–50 mg/kg of fish as a supplementary feed to increase tocopherol in the female brood fish.
Keeping the brood fish in floating netcages have advantage of having natural good seawater. For broodstock tanks on shore, the water quality have to be maintained as good as in nature. Normally 50–80% of water are changed daily to keep the water quality to the acceptable level (Table 2.) Poor water quality will result in a negative effect on gonadal maturation and spawning.
Salinity of the water in broodstock tanks prior to spawning season can be maintained within the range of 20–25 ppt, but it should be increased to the range of 28–32 ppt during spawning season.
Disturbances to brood fish during spawning season should be avoided. Excessive noise and vibration due to vehicles near-by are also stressing the brood fish having a negative effect on spawning.
Male and female brood fish should have a uniform size. Sex ratio between male and female is usually 1:1. If the male are much smaller than the female, the number of male brood fish should be increased.
Mature females of 3–7 years (body weight 3.5–12kg) and males of 3–5 years (body weight 3.0–7.5 kg) are good for inducement of the spawning.
Spawning activity of seabass in captivity is found closely related to the lunar cycle. In Indonesia water natural spawning activity of seabass occurs between 19 00 and 23 00 hr during high tide at 1st–8th day after new moon or full moon. Usually it spawns at night both during the new moon and full moon phases. However, the fish will yield more eggs and better quality at full moon than at new moon phase.
Two methods may be used for collecting seabass eggs from a spawning tank.
A small seine net mesh size about 200 u can be used to collect the egg from the spawning tank in the morning after spawning (Fig.7).
Collecting the eggs from spawning tanks by seine net is necessary to stop the aeration to allow the eggs afloat in upper layer of the water column. This period might reduce the dissolved oxygen in water to a level that could stress the spawners especially in high stocking density broodstock tanks. More over turbulence caused by fish movement in the tank tends to redistribute the eggs in the water column resulting in uncompleted collection of eggs. To avoid the problems, the eggs can be transferred into the egg collector through the continuous flow of water. Seawater should be flowed after the fish spawned. The overflowing water carries the eggs into the egg collector made of a fine netting (200 u) of 10–20 1 capacity (Fig. 8).
Fig. 7 Collecting seabass eggs using a small seine net.
The eggs are placed in plastic buckets. The unfertilized eggs sinking to the bottom are discarded. The fertilized eggs are washed through 1.5–2.0 mm mesh screen in order to remove debris or foreign materials that may attach to the eggs. The number of fertilized eggs is estimated and treated with 5 ppm of acriflavine solution or in ovadine at dilution of 1 ml/ 100 ml water for 1 min for disinfection purposes. Since survival of eggs and disinfectant are most effective at neutrality, an adjustment of disinfectant solution to pH 7.0 is necessary. Rinse the eggs in seawater two to three times before placing in the incubation tanks.
| 1= water inlet, | S= screen, | N= egg - collecting net, and |
| 0= water outlet |
Fig. 8. Diagram of a spawning tank with egg-collecting system.
The eggs are incubated in a circular tank. The stocking density recommended is around 60–100 eggs/1. Hatching rates of seabass eggs vary with salinity (Table 4). The salinity ranging between 28–30 ppt is recommended for hatching of the egg. During the hatching process, aeration should be given gently to allow the water to ciruculate and prevent the eggs from settle on the bottom. Unfertilized eggs are siphoned out by stopping the aeration so that fertilized eggs will float to the upper layer of the water column. After the 10th hour, 50% of the water in the incubator tank are changed by siphoning out the lower layer and replaced it with a new seawater of the same salinity (28–30 ppt).
Table 4. Effects of salinity on hatching rate of seabass eggs.
| Salinity (ppt) | Rate of hatching (%) | |
| 0 | 00.0 | |
| 5 | 02.9 | |
| 10 | 58.5 | |
| 15 | 75.0 | |
| 20 | 82.4 | |
| 25 | 83.4 | |
| 30 | 80.8 | |
| 35 | 46.9 |
Source : Tattanon and Maneewongsa, 1982a.
The fertilized eggs hatched out within 17–18 hrs at a water temperature of 26–28 degree Celsius. The stages of development of the seabass egg recorded by Maneewongsa and Tattanon (1982) is given in Table 5, Fig. 9. The newly hatched larvae (size about 1.5 mm) contain a big yolk sac with an oil drop at the anterior end. When it does not swim the fish has its head in an upward position. While moving, the body of the fish will be at a 45–90 degree angle with the horizontal. The body is slender and compressed. The pigment is scattered in spots. The eye, digestive system and intestines can be seen clearly. The mouth parts appear when the fish is three days old. The yolk sac is completely absorbed at Day 4 (Table 6, Fig. 10).
Table. 5 Embryonic development of seabass eggs at 27°C
| Stage of development | Hour Minutes | |
| Fertilized egg | 00 | |
| 1 - cell | 00 35 | |
| 2 - cell | 00 40 | |
| 4 - cell | 00 45 | |
| 8 - cell | 01 00 | |
| 32 - cell | 02 15 | |
| 64 - cell | 02 45 | |
| 128 - cell | 02 55 | |
| Multi - cell | 03 15 | |
| Blastula | 05 30 | |
| Gastrula | 06 30 | |
| Morula | 08 30 | |
| Early embryo with eye vesicle | 11 20 | |
| Heart function, free movement of tail | 15 30 | |
| Hatching | 17 30 |
Source : Maneewongsa and Tattanon 1982.
Table 6. Rate of absorption of the yolk
| Diameter of Yolk (mm) | Days | |
| 0.88 | 0 | Source : Maneewongsa and Tattanon, 1982 |
| 0.35 | 1 | |
| 0.28 | 2 | |
| 0.15 | 3 | |
| 0.01 | 4 | |
| 0.00 | 5 |
Fig. 9 Embryonic development of seabass
(Source : Maneewongsa and Tattanon, 1982)
Newly hatched larvae float in the upper layer of the water column. Chromatophores are observed behind the eyes, in the body, and on the oil globule. The larvae completed its larval stage within 3 weeks (Fig. 10).
Fig. 10. Larval development of seabass (Source : Konsutarak and Watanabe, 1984)
Day 1 (TL 2.20 ± 0.08 mm)
Large part of the yolk sac absorbed. Mouth still closed. Anus can be observed. Eyes unpigmented. Pectoral fin appeared as bud. The larvae distributed uniformly in the rearing tank.
Day 2 (TL 1.52 ± 0.06 mm)
Yolk sac almost disappeared. Mouth opened. The larvae gather near aeration or in the direction of the light. Oil globule is still apparent (Konsutarak and Watanabe, 1984).
Day 3 (TL 2.61 ± 0.008 mm)
Air bladder appeared. Yolk sac absorbed, but the oil globule is still observed.
Day 4 (TL 2.78 ± 0.15 mm)
Mouth opened with developed upper and lower jaws. Nostrils appeared on snout. Pectoral fin developed in round shape as finfold. Digestive tract extended relatively in thickness. Melanopores presented on dorsal and ventral profiles, body mid-line and abdomen. Melanophores scattered on mid-brain and lower jaw. Oil globule disappeared.
Day 5 (TL 3.08 ± 0.09 mm)
Teeth appeared on the upper jaw
Day 6 (TL 3.10 ± 0.13 mm)
Under part of caudal end whitish
Day 7 (TL 3.44 ± 0.09 mm)
Rudiments of dorsal and anal fins appeared. Serrated spine appeared at the pre-operculum. Melanophores from snout to tail pronounce, making the larvae black in color (Konsutarak and Watanabe, 1984).
Day 8 (TL 3.58 ± 0.13 mm)
Teeth appeared in the lower jaw
Day 9 (TL 3.49 ± 0.26 mm)
Caudal end of the notochord bent. Soft ray part of the caudal fin developed.
Day 10 (TL 3.81 ± 0.27 mm).
Three serrated spines developed on the posterior margin of the pre-operculum. Head slightly rounded, and the body depth extended with developing dorsal and anal fin base. Tip of notochord flexed to develop the caudal fin. Segmented soft rays appeared. Melanophores pronounced from snout to tail and to the abdomen.
Day 11 (TL 3.87 ± 0.24 mm)
Posterior margins of the dorsal and anal fins were deeply cut. Larval membrance in front of the anal fin small. The number of serrated spines at the posterior margin of the pre-operculum increased from three to four.
Day 12 (TL 4.41 ± 0.09 mm)
Segmented soft rays appeared in the dorsal fin.
Day 13 (TL 4.58 ± 0.17 mm)
The number of serrated spines on the posterior margin of the preoperculum increased to four. Larval membrane in front of anal fin disappeared.
Day 14 (TL 4.75 ± 0.32 mm)
Dorsal and anal fins separated from the caudal fin, and the rudiment of pelvic fin appeared. The chordal well developed and the vertebrae could be counted (11+14). Melanophore spreaded to the whole abdomen, dorsal and anal fins. A white band distinguished from the center of the dorsal fin to the anal fin with the naked eye.
Day 15 (TL 5.41 ± 0.50 mm)
Spines and soft rays of dorsal and anal fins well developed. One-two serrated spines on the upper part of the posterior margin of the operculum developed.
Day 16 (TL 6.56 ± 0.56 mm)
Each fin completely separated. Number of spines and soft rays of the dorsal and anal fins constant, 19 and 11 respectively. Serrated spines of the anterior margin of the pre-operculum disappeared.
Day 18 (5.5 ± 0.40 mm)
Nostrils well developed. Maxillary reached to the center of eye. Single opercular spine presented on the upper part of operculum. Body depth increased relatively. The spines and soft rays of the dorsal, anal, and caudal fins well developed. Pectoral fin partially developed while the pelvic fin appeared as bud. Melanophores scattered on head and most of body and the posterior part. The body has two vertical bands and divided by mid-point of body. There is no melanophore on caudal peduncle.
Day 21 (TL 8.91 ± 1.19 mm)
Numbers of spines and soft rays of the dorsal and anal fins constant. Scales appeared in the mid-lateral surface above the anal fin. The body color changes from black to brown (Konsutarak and Watanbe, 1984).
