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Chapter 8

Maturation and larval rearing of the Pacific white shrimp, Penaeus vannamei
Lorenzo M. Juarez, Shaun M. Moss


The global shrimp farming industry is largely dependent on domesticated broodstock to provide a predictable and consistent supply of healthy postlarvae to stock into nurseries or growout facilities. This process of captive rearing includes the maturation and spawning of broodstock to produce viable larvae and the rearing of larvae to produce postlarvae for stocking. Maturation refers to the process of conditioning broodstock to stimulate gonad development and to the induction of mating, spawning and hatching of eggs to produce viable larvae. Environmental controls of shrimp maturation include water temperature, photoperiod, diet and water quality. Activities associated with shrimp maturation include the acclimation, stocking and sourcing of gravid broodstock, as well as mating and spawning of fertilized eggs. Eggs hatch into larvae which go through a series of developmental stages, namely nauplii, protozoea, mysis and postlarvae. Protozoea are herbivorous and require phytoplankton, whereas mysis are carnivorous and require zooplankton. Postlarvae are omnivorous and can eat both plant and animal matter. Of paramount importance to both maturation and larval rearing are water quality and animal health. Water filtration and disinfection are important activities for the captive rearing of penaeid shrimp as are biosecurity procedures to minimize the introduction and spread of pathogens. In this chapter, we review commercial maturation and larval-rearing techniques for penaeid shrimp, with a particular emphasis on the Pacific white shrimp, Penaeus vannamei. This species dominates global production largely because of the commercial availability of Specific Pathogen Free (SPF), selectively bred stocks.


Captive rearing of penaeid shrimp includes the conditioning of broodstock to produce viable larvae and the rearing of larvae to produce postlarvae (PL) for stocking into nurseries or growout facilities. Rearing techniques used today in commercial shrimp hatcheries have their origins in earlier work. Larval rearing of penaeid shrimp was first documented in 1935 when Motosaku Fujinaga (Hudinaga) spawned gravid, wild-caught female Kuruma prawns, Penaeus japonicus, and reared the larvae in captivity (Hudinaga, 1935). Methods developed by Fujinaga include the use of large (50 to 500 m3), flat-bottom tanks where female shrimp spawn, and where larvae are reared in water fertilized with nutrients to stimulate the production of planktonic food organisms. Modifications of these methods are still used today and collectively are referred to as the “Japanese method” for larval rearing. During the 1960s, with help from Dr. Fujinaga, Harry Cook and co-workers at the U.S. National Marine Fisheries Service in Galveston, Texas, developed a modified approach to rear larvae of P. setiferus and P. aztecus. This “Galveston method” included the use of smaller tanks (0.5 to 20 m3) with conical bottoms, airlifts for aeration, daily water exchanges and addition of exogenous, live feeds (Mock and Murphy, 1970).

Commercial shrimp farming in the Americas began in the 1970s and was facilitated by favourable geographic, climatic and biological conditions. Of paramount importance to the economic success of these early commercial efforts was the seasonal abundance of wild postlarval Pacific white shrimp, P. vannamei, collected along the Pacific coast of Ecuador and Central America. Although natural supplies of postlarvae were critical to catalyzing a commercial shrimp farming industry, shrimp farmers soon realized that a consistent supply was required for the industry to expand and overcome natural fluctuations in abundance, especially during El Niño years. The predictable and consistent supply of postlarvae from commercial hatcheries, using modifications of the Galveston method, was one of the major factors responsible for the rapid growth of shrimp farming in the region during the 1980s and early 1990s. However, in the 1990s, viral diseases severely impacted the global shrimp farming industry, resulting in significant changes in operations (Lightner et al., 2009). Specifically, wild broodstock were identified as potential disease vectors, and there was a move towards developing domesticated broodstock to help mitigate disease problems. The use of wild broodstock in the Americas decreased from 80% in 1992 (Kawahigashi, 1992) to 3% in 2001 (Crocos and Moss, 2006), based on survey results obtained from commercial hatcheries at those times.

The disease status of domesticated broodstock can be controlled, to a large extent, using specific pathogen free (SPF) shrimp together with a comprehensive biosecurity strategy designed to minimise the introduction and spread of pathogens (Lotz, 1997a; Lightner, 2003; Lightner et al., 2009). SPF shrimp are free of specified pathogens and SPF status depends on the level of biosecurity and disease history of the facility where the shrimp are maintained (Moss et al., 2003; Lightner et al., 2009).

SPF populations of P. vannamei are commercially available on a large scale, and this has led to a spectacular switch from farming P. monodon to P. vannamei in Asia over the past decade. Global production of farmed P. monodon decreased from 630,984 metric tons (MT) in 2000 to 589,888 MT in 2007, a 6.5% reduction. During the same time period, production of farmed P. vannamei increased from 145,386 MT to 2,296,630 MT (FAO, 2009a), a 15-fold increase. Historically, shrimp farmers in the Western Hemisphere have cultured P. vannamei, therefore this change has occurred primarily in Asia where more P. vannamei are now produced than in the Americas (Moss, 2004). The combined value of these two species produced by the global shrimp farming industry in 2007 was $11.7 billion U.S. dollars, of which $2.9 billion corresponded to P. monodon and $8.8 billion to P. vannamei (FAO, 2009b).

In this chapter we review commercial maturation and larval-rearing techniques for penaeid shrimp and emphasize current approaches and emerging technologies. The chapter focuses primarily on P. vannamei as this species now dominates global production. However, references to other penaeid species are made throughout the chapter when relevant.


In the context of shrimp farming, “maturation” commonly refers to the process of conditioning broodstock to stimulate gonad development and to induce mating, spawning and hatching of eggs to produce viable larvae.

Reproductive biology of penaeid shrimp

The male reproductive system consists of testes, vasa deferentia, and terminal ampoules which contain spermatophores (King, 1948). The testes are paired organs located above the heart in the vicinity of the digestive gland and are composed of five to eight lobes on each side. They are connected to the terminal ampoulae by the vasa deferentia, which divided into distal, medial and proximal portions. In mature males, the ampoules harbour the spermatophores which contain the sperm. The male genital ducts end in pores located on the inner surfaces of the coxae of the last pair of pereopods.

The main secondary sexual structure of male shrimp, called the petasma, is a tubular organ resulting from specialization of the endopods of the first pair of pleopods (Dall et al., 1990). In immature males, these endopods appear as two simple structures. As the male matures, the endopods become linked by a series of hook-like setae along the mid line. Another secondary sexual characteristic of males is the appendix masculina. This structure results from specialization of the endopods of the second pair of pleopods. It has been proposed that during copulation, the petasma, aided by the appendix masculina, helps in transferring the spermatophores to the female (Dall et al., 1990). Spermatophores are ejaculated as pairs, one from each side and pressed together to form a compound spermatophore. Sperm cells are stored in a sperm package inside each spermatophore. Spermatozoids are spherical and have a single, non-motile projection (spike) that attaches to the ovum during fertilization. Leung-Trujillo (1990) reported that the number of spermatozoa is directly related to the size of the male, and Wang et al. (1995) found sperm counts of 13.1± 7.2 (SD) million sperm cells per spermatophore in P. vannamei males averaging 41.1 ± 5.2 (SD) g in weight.

The ovaries of female shrimp are much larger than testes of the males and extend from the oesophageal region to the sixth abdominal segment (King, 1948). They consist of two anterior lobes, six to eight lateral lobes and two long posterior lobes (Chamberlain, 1985). Two oviducts lead from the sixth lateral lobe to the genital openings located on the inner surfaces of the coxae of the third pair of pereopods. The seminal receptacle of the female is called the thelycum and consists of modified sternal plates on the seventh and eighth thoracic segments. The structure of the thelycum is unique to each shrimp species and is widely used in taxonomy. In white shrimp, like P. vannamei, thelyca are simple open depressions referred to as “open” thelyca to differentiate them from the “closed” paired and covered pouches characteristic of other penaeid shrimp, such as P. monodon.

Ovarian maturation can be classified into five stages (Fig. 1) according to the external appearance of the ovaries (AQUACOP, 1975; Chamberlain, 1985; Dall et al., 1990; Yano, 1985):

Stage I. Undeveloped. Posterior ovarian lobes are translucent and of smaller diameter than the intestine. Oocites undeveloped.

Stage II. Developing. Posterior ovarian lobes are opaque and diameter equal to that of the intestine. Oocites growing.

Stage III. Nearly ripe. Posterior ovarian lobes are yellowish and larger in diameter than the intestine. Oocites commence vitellin accumulation.

Stage IV. Ripe. Ovarian tubes are deeply pigmented and almost occupy the entire dorsum of the shrimp. Oocites are mature.

Stage V. Spent. Ovaries empty; lobes flaccid and convoluted. Oocites spawned or undergoing reabsorption.


Figure 1. Stages of ovarian maturation in white shrimp (Penaeus vannamei). Shown from left to right are dorsal dissections exposing the ovaries of stages I – V as described in the text.
(Photo courtesy of Traci Holstein).

In P. vannamei, mating occurs after the ovaries have matured and usually towards the end of the moult cycle. Mating behaviour is triggered by photoperiod and occurs around sunset. It has been suggested that females may release pheromones to attract males (Yano et al., 1988), although there is no conclusive evidence to support this suggestion. Courtship starts when a male approaches a female and attempts to get underneath her from behind. The female then swims away and is followed by the male. This chasing behaviour can occur for several minutes, and more than one male may be involved. Males often are observed chasing immature females as well as other males. During insemination, the male briefly turns upside down, while remaining parallel to the female, and grasps her from underneath with his pereopods for one or two seconds at which time spermatophore transfer occurs.

During spawning, eggs are extruded backwards from the genital opening and passed on to a fertilization chamber formed by the coxae and ventral setae of the third and fourth pairs of pereopods. Based on morphology, Dall et al. (1990) proposed that sperm from the spermatophores could be pumped by movements of the coxae and travel forward towards the fertilization chamber through a channel formed by the median plate of the thelycum and the coxae of the fourth pair of pereopods. In P. vannamei, spawning takes place three to five hours after mating. Immediately before spawning, the female rests on the bottom and then moves toward the surface where she begins swimming in circles 0.8 to 1.2 m in diameter. As the eggs are spawned, the movement of the pereopods scatters the eggs behind the female. Spawning takes one to four minutes. Eggs are negatively buoyant and tend to sink slowly. Fertilized eggs can be identified under a microscope by the presence of hatching membranes and developing embryos. Commercial hatcheries commonly report fertilization rates of 65 to 95%. Fecundity in P. vannamei is correlated with female size (Treece, 2000). R Wouters (CENAIM/ESPOL, Ecuador, personal communication) developed an equation for the number of eggs (y) as a function of weight in grams (x), based in 612 wild, female broodstock from Ecuador with weights ranging from 27 to 80 g. The equation is y = 3,665(x) + 22,660 with r2 = 0.1892. Commercial maturation facilities normally obtain between 80,000 and 180,000 nauplii per spawn for natural mating and 40,000 to120,000 nauplii per spawn for artificial insemination (Juarez et al., 2003).

In decapods, gonads are controlled by a system of hormones that promote or inhibit maturation. The gonad-stimulating hormone (GSH), produced by the brain and the thoracic ganglion, promotes sexual maturation. The gonad-inhibitory hormone (GIH), produced in the X-organ-sinus gland complex of the eyestalk is responsible for inhibiting sexual maturation. The actions of GSH and GIH are antagonistic (Adiyodi and Adiyodi, 1970). Reduction in levels of GIH, either naturally or artificially by eyestalk ablation, promote sexual maturation (Dall et al., 1990).

Overview of maturation facilities

Maturation tanks are used to maintain broodstock, either as mixed populations or separated by sex. These tanks provide a controlled environment to promote the development of mature gonads, chasing and mating. Maturation tanks come in a variety of sizes and shapes, usually round or rectangular. In the past, round tanks were the most common shape. However, rectangular raceways are now popular because of their space efficiency. These raceways can be up to 40 m in length, with or without central baffles or rounded ends. In general, maturation tanks are designed to be big enough to accommodate a large number of broodstock yet small enough (in width) so that mated females can be caught from the sides of the tank. P. vannamei broodstock require sufficient space to perform chasing and mating behaviours (Yano et al., 1988), and minimum tank diameter has been estimated to be about 3 m for several penaeid species (Bray and Lawrence, 1992). In a survey conducted by Moss and Crocos (2001), 74% of commercial maturation facilities had between 5 and 50 maturation tanks; 74% of the tanks were round, 54% were lined with plastic, 77% were between 5 and 20 m3 in volume, and 71% were 0.2 to 0.6-m deep.

High-quality and stable water conditions are necessary for the successful maturation and spawning of shrimp (AQUACOP, 1983; Chamberlain, 1985; Wyban and Sweeney, 1991). Historically, water quality was maintained by replacing 100-400% per day of the water in the maturation tanks with new, filtered seawater from an external source (Bray and Lawrence, 1992). However, during the late 1990’s, the industry began using recirculation technologies in response to biosecurity concerns (Courtland, 1999; Ogle and Lotz, 2001; Otoshi et al., 2002, 2003). Although recirculation systems are relatively costly and complicated to operate, they also provide advantages such as greater control over water quality, enhanced biosecurity, better temperature control, higher retention of pheromones and reduced effluent discharge. The shift from flow-through to recirculation systems is reflected in a survey (Moss and Crocos, 2001) where 46% of those facilities used partial or complete water recirculation. The most common types of filtration were fluidized bed filters and pressurized sand filters. For descriptions of water recirculation systems for shrimp maturation see Ogle and Lotz (2001) and Courtland (1999).

Operating protocols: broodstock acclimation, stocking and sourcing

Broodstock arriving at a maturation facility should be acclimated to local water-quality conditions, especially in terms of water temperature, salinity and pH. Practical rates of acclimation are 1°C, 1 ppt salinity and 0.1 pH unit every 15 minutes. After acclimation, it is advisable to release a few broodstock into the receiving water before releasing the entire population. In addition, common reception procedures include segregating new broodstock to modular quarantine units, taking samples for disease status evaluation and applying prophylactic treatments for bacteria and/or external fouling parasites.

Typically, new broodstock are allowed to recover from shipping stress for one to two weeks, at which time they are fed a standard maturation diet. Maturation tanks are stocked with broodstock shrimp at densities of 5-10/m2, with a female:male ratio of 1:1 to 1:1.5. Broodstock should be at least eight months old and weigh a minimum of 32 g for males and 35 g for females. Broodstock can be kept in production for different time periods, according to the needs of the facility, but peak nauplii production and quality usually last up to 120 days after ablation.

Maturation tank bottoms should be siphoned daily to remove uneaten feed, faeces and moults. Broodstock maintained in tanks with excess organic matter are susceptible to bacterial necrosis and melanization of the male reproductive system, a condition characterized by brown or black discoloration of the spermatophore and sometimes the vas deferens (Talbot et. al., 1989). The number of moults and mortalities should be recorded at the time of siphoning in order to track moulting frequency and stock mortality replacements. There appears to be an inverse relationship between moulting and sexual maturation, and there is a decrease in nauplii production during periods of heavy moulting.

Typically, sourcing for mated females is done one hour after sunset. Some operators collect only females with obvious spermatophores, whereas others collect all females with advanced ovarian development. It is advisable to install a pen or net enclosure in the maturation tank to hold unmated females to avoid repeated collecting. Also, it is important that females are handled gently to avoid stress and possible loss of the spermatophore.

Natural mating versus artificial insemination

Surveys by Kawahigashi (1992) and Crocos and Moss (2006) showed that natural mating was the most common method to obtain mated females. Artificial insemination (AI) has been used successfully for routine production in commercial facilities, but it can be a stressful or even lethal procedure, especially for males. On the other hand, AI is an essential technique for breeding programs. For a description of different AI techniques as applied to genetic selection programs see Arce et al. (2000).

Operating protocols: spawning

Mated females can be spawned in individual or communal tanks. Most maturation facilities in the Western Hemisphere use a mass-spawning approach where as many as 25 mated females can be spawned in a 4-5 m diameter spawning tank. It is advisable not to exceed one female, or 40 grams of body weight, per 100 L of water. In a survey by Moss and Crocos (2001), most respondents (67%) placed multiple females in a single spawning tank, 27% placed one female in each spawning tank, and 6% relied on mass spawning within the maturations tanks. Advantages of mass spawning (rather than individual spawning) include reduced labour, better fertilization rates and increased harvest efficiency. The major disadvantage is the inability to collect fecundity and fertilization data for individual females. In breeding programs, individual spawning is an essential tool for pedigree tracking. The diameter of individual spawning tanks should be at least 1 m to allow for unrestricted spawning behaviour. Once a female spawns, she should be returned to her original maturation tank.

Water used in spawning tanks must be filtered for particles larger than 0.5 µm, treated with granular activated carbon (GAC) and disinfected with ultraviolet light. To promote better hatching, spawning water often is adjusted to a salinity of 30 to 32 ppt and treated with 5 to 15 ppm ethylenediaminetetraacetic acid (EDTA), a chelating agent that sequesters metal ions such as Cu2+ and Fe3+. It is important to maintain the same water temperature in the spawning tank as in the maturation tank (normally 27 to 28 °C). Temperature fluctuations or temperatures higher than 30 °C in spawning tanks often result in nauplii deformities. Substandard water quality is a common cause of poor hatching and bacterial infestation of the eggs. Aeration in spawning tanks should be low enough to prevent damage to the eggs but sufficient to prevent settling.

Cooney et al. (2005) suggested that bacteria on the surface of broodstock were the most common source of contamination to which newly hatched larvae were exposed. To minimise the horizontal transmission of pathogens, improve hatching rate and increase the survival of nauplii, shrimp eggs often are harvested from spawning tanks, rinsed, disinfected and transferred to separate hatching tanks. These procedures minimise the horizontal transmission of pathogens. Eggs can be safely harvested from spawning tanks by draining or siphoning 4-6 h after spawning. Eggs can be retained and concentrated in tanks provided with 100-?m plastic screen overflows and rinsed with abundant filtered and ultraviolet irradiated seawater. After washing, eggs can be disinfected with an iodophore compound. Common dip treatments for eggs are 50 ppm Argentyne® for 60 sec., 20 ppm Argentyne® for 5 min., or 1.2 mL of Formalin in 40 L of sea water for 30 sec. Repeated tests at the GMSB shrimp hatchery in Florida showed that rinsing eggs with clean, abundant seawater is more effective than iodine treatments at reducing bacterial load. After rinsing, eggs can be stocked into dedicated hatching tanks at ? 4,000 eggs/L and provided with enough aeration to keep them in suspension until hatching. Nauplii can be harvested by either light-attraction and siphoning, or by draining and further rinsed with abundant, clean seawater. As an additional precaution, stage three nauplii (N-3) can be dip-treated with 25 ppm Argentyne® for 3 min., or stage five nauplii (N-5) with either 50 ppm Argentyne® for 60 sec., 25 ppm Argentyne® for 3 min., or 2 ppm Chloramine-T for 5 min. Treflan TR-10® at 0.01-0.05 ppm is commonly used to prevent fungal infestations of eggs and nauplii in hatching tanks. Late nauplii about to metamorphose to protozoea should not be subjected to chemical treatments or other stressors, including handling and transferring. Spawning tanks and related equipment require scrupulous cleaning, disinfecting, rinsing and drying between batches.

Water temperature and lighting

Water temperature in maturation tanks has a significant impact on gonad development and mating. For consistent reproductive performance, fluctuations in water temperature need to be minimized and maturation tanks should be maintained at 28 ± 1 °C. Deviations below this temperature may reduce mating frequency and inhibit ovarian development. Deviations above this temperature may reduce mating frequency, fecundity, or egg viability and can promote bacterial growth. Kawahigashi (1992) reported changes in nauplii production due to seasonal fluctuations in temperature among 21 commercial maturation facilities surveyed in the Americas. High ocean temperatures (>33 °C) during the summer months were blamed for a 50% decline in nauplii production at commercial facilities in Mexico and Colombia. Several studies have shown a correlation between high water temperatures (? 29 °C) and low sperm counts, resulting in reduced fertility of males (Perez-Velazquez et al. 2001; Perez-Velazquez and Gonzalez-Felix, 2002).

Illumination and photoperiod are important factors in promoting shrimp maturation and mating behaviour. Many commercial and research facilities establish photoperiod to resemble summer conditions, i.e. 13 h light and 11 h darkness. Photoperiod typically is set to accommodate the work schedule of the crew (for example, lights out at 1130 h and on at 2230 h). This photo-shifting allows for the sourcing of mated females shortly after noon and for harvesting of nauplii (spawned the previous day) at about 1500 h, before the end of a normal work day. Dawn and dusk effects can be simulated by placing separate banks of lights on independent timers, or by using more sophisticated dimming timers. Gradual dimming allows for a more natural transition of light intensities and minimizes stress to the shrimp. Other facilities rely on natural photoperiods and adjust the work schedule accordingly. Wurts and Stickney (1984) recommended that light intensity for shrimp maturation should resemble that found in the shrimp’s habitat and depth which, in the case of P. setiferus, was estimated to be no higher than 12 ?W/cm2. Maturation facilities use similar levels of subdued lighting (approximately equivalent to 29 luxes, or 2.7 ft-candles at the water’s surface) to promote maturation.

Broodstock nutrition and diet

Broodstock diet plays a significant role in the maturation process, especially in stimulating ovarian development in females and in determining nauplii quality (Racotta et al., 2001). Protein requirements for maturation diets have not been quantified, but given the accelerated biosynthesis associated with egg development, they are assumed to be high relative to those of non-maturing shrimp. Some of these requirements are likely met by diversion of resources from somatic growth (Harrison 1997; Wouters et al. 2001). The protein level in fresh, raw maturation feed organisms can range from 58-73 % (dry basis), which is higher than the approximately 50% protein content reported in several commercially available broodstock diets (Wouters et al., 2000; Wouters et al., 2001).

In addition to protein, numerous studies on broodstock nutrition indicate the importance of lipids, particularly polyunsaturated fatty acids (PUFAs), on reproductive performance (Bray et al., 1990; Cahu et al., 1995; Wouters et al., 1999a). The most common fatty acids in shrimp ovaries are 16:0, 16:1(n-7), 18:0, 18:1(n-9), 18:1(n-7), 20:4(n-6), 20:5(n-3) and 22:6(n-3). There appears to be few interspecific differences in ovarian fatty acid profiles among penaeid shrimp, as reported by Wouters et al. (1999b) for P. vannamei, Middleditch et al. (1980) for P. setiferus, Teshima and Kanazawa (1983) for P japonicus and Mourente and Rodríguez (1991) for P. kerathurus.

Shrimp have limited abilities to synthesize the n-6 and n-3 families of fatty acids de novo, including the polyunsaturated linoleic (18:2n-6) and linolenic (18:3n-3) acids (Kanazawa et al., 1979a), and they have a limited ability to elongate and de-saturate these PUFAs into highly unsaturated fatty acids (HUFAs), such as arachidonic (20:4n-6), eicosapentanoic (20:5n-3) and decosahexanoic (22:6n-3) acids (Kanazawa et al., 1979a; Kanazawa et al., 1979b; Kayama et al., 1980). Because de novo synthesis of these fatty acids is limited, the high ovarian content of 20:5(n-3) and 22:6(n-3) has been attributed to dietary intake and is of practical importance in maturation diets. Middleditch et al. (1980) attributed the popularity of marine polychaete worms as maturation feed items to their PUFA content and profile. In fact, 66% of the respondents in a recent survey indicated that they include polychaete worms in their maturation diets (Crocos and Moss, 2006).

An increase in lipid levels in maturing ovaries, concurrent with their decrease in the hepatopancreas (HP), has been reported by Teshima and Kanazawa (1983) for P. japonicus, suggesting that lipids are stored in the HP then transported to the ovaries via the hemolymph. Wouters et al. (2000) reported a similar observation in P. vannamei, but highlighted the importance of dietary versus stored lipids. During maturation, most ovarian lipids are transferred to the eggs, and after spawning and hatching, to the yolk sac of the nauplii. Palacios et al. (2000a) showed a correlation between high levels of specific lipids (triglicerol and lysophosphatidylcholine) in eggs and survival of the resulting larvae. Nauplii with a higher lipid content exhibited higher rates of metamorphosis and survival (Lavens et al., 1991). Consequently, lipids levels in eggs and nauplii have been proposed as indicators of the nutritional condition of broodstock and of the quality of larvae (Barclay et al., 1983; Lavens and Sorgeloos, 1991; Lavens et al., 1991). Wouters et al. (1999a) reported that triacylglyceride (TAG) levels increased from 8.3% to 33.8% in P. vannamei ovaries and that this level decreased to 20.6% after spawning. Nauplii from these spawns had 33.5% TAG. The study suggested that TAG are selectively incorporated into the eggs and resulting nauplii and constitute the principal energy source during embryogenesis, hatching and early development.

Carotenoids are a group of pigments that cannot be biosynthesized by broodstock shrimp. During ovarian maturation, carotenoids from the diet are first accumulated in the HP then mobilized via the hemolymph to the ovaries, resulting in their darkening colour (Wouters et al., 2001). Astaxanthin is the principal carotenoid in maturing shrimp ovaries (Dall et al., 1995) and polychaete worms. Artemia and krill are important sources of carotenoids in shrimp maturation diets. Pigment-containing ingredients have been found to be effective at improving reproductive success and nauplii quality. Wyban et al. (1997) reported improved nauplii survival when broodstock were fed squid that had been marinated with paprika, which contains the pigments beta-carotene, beta-cryptoxanthin, capsanthin and capsorubin (D’Abramo et al., 1983), relative to the non-marinated control diets.

Specific vitamins are important in broodstock diets, providing anti-oxidant protection to dietary lipids and cells (Chamberlain, 1988; Alava et al., 1993a; Alava et al., 1993b; Cahu et al., 1995; Sagi et al., 1995). Specifically, vitamins E and C are believed to play important roles in protection from free radicals and in promotion of gonad maturation (Wouters et al., 1999b). Vitamin E has been shown to improve the percentage of normal sperm and the rate of ovarian maturation in P. setiferus (Chamberlain, 1988). Du et al. (2006) provided broodstock of P. vannamei with four different concentrations of vitamin E (tocopherols) and concluded that dietary supplementation with 350 mg/kg a-tocopherol was needed to achieve significantly better ovarian maturation and reproductive performance. Alava et al. (1993a) demonstrated retarded ovarian maturation as an effect of deficiencies in vitamins A, C and E. Maturation diets often are supplemented with ascorbic acid (AA). Deficiencies in AA can result in poor feed conversion, poor growth, incomplete moulting, decreased resistance to stress, impaired wound healing and in extreme cases, death (Sangha et al., 2000). Ascorbic acid has been shown to accumulate in the ovary of Palemon serratus during vitellogenesis (Guary et al., 1975). Cahu et al. (1991) found that supplementation of maturation diets with AA increased hatchability of eggs in P. indicus.

Most studies on the nutritional requirements for broodstock have focused on the female’s requirements for vitellogenesis. Less attention has been given to the male’s requirements because they mature relatively easily in captivity. Wang et al. (1995) found no significant differences in sperm quality in P. vannamei when test groups of males were fed four different diets for periods of 24 d. In contrast, Pérez-Velásquez et al. (2003) reported a significant dietary effect on male reproductive quality in P. vannamei and suggested that the typical dietary combination of fresh-food organisms is not nutritionally optimal for male broodstock. Similarly, Paibulkichakul et al. (2008) reported a dietary effect in adult male P. monodon, as the number of spermatozoa was significantly enhanced by higher concentrations of dietary fish oil and astaxanthin.

Maturation facility managers feed their broodstock a mixed diet consisting of raw, wet feeds such as squid, marine polychaete worms, enriched adult Artemia, mussels, clams and oysters, as well as dry, formulated feeds. Marine polychaete “bloodworms” (Americonuphis reesea) were first used as maturation feed by J. Mountain (Harbour Branch Oceanographic Institution, personal communication) in 1976 at Ralston Purina’s shrimp facility in Panama. Since then, several species of marine worms have been used as the primary source of essential HUFAs in maturation diets. These worms are the most expensive component of maturation rations (Harrison, 1997) and often have to be imported into countries that do not harvest them naturally. Wild North Atlantic bloodworms, Glycera dibranchiata, and sandworms, Nereis viriens, are widely used in maturation facilities in the Western Hemisphere. Commercial companies in Europe have cultured and commercialized N. virens, also known as “ragworms”. These farmed worms have several advantages over their wild counterparts, including a steady supply, consistent quality, a better HUFA profile (Table 1), increased biosecurity and more stable pricing (Pinon, 2000). Laufer et al. (1997) reported that bloodworms increase fecundity and hatch rate in P. vannamei.

Table 1. Comparison of fatty acid profiles between wild and farmed sandworms (Nereis virens).a

Table 1. Comparison of fatty acid profiles between wild and farmed sandworms (Nereis virens).a
Fatty acid Systematic name Wild (mg/g DW) Farmed (mg/g DW) % Fatty acid increment Wild Farmed farmed over wild
C15:0 Pentadecanoic 0.22 1.82 +727%
C16:0 Palmitic 5.41 10.78 +99 %
C16:1n7 Palmitoleic 1.82 3.08 +69 %
C18:0 Estearic 0.86 2.02 +134 %
C18:1n12 Petroselenic 1.10 1.53 +39 %
C18:1n9 Oleic 0.25 12.32 4,828 %
C18:1n6/7 Cis-vaccenic 2.01 3.70 +84 %
C18:2n5 - - - 1.23 9.86 +701 %
C18:2n6 Linoleic 2.15 2.39 +11 %
C18:3n3 Linolenic 0.25 0.48 +92 %
C20:0 Arachidonic 1.18 1.74 +47 %
C20:1n9 Gondoic 0.21 7.10 +3,280 %
C20:2n6 - - - 0.12 3.70 +2983 %
C20:4n6 ARA 0.15 0.58 +286 %
C20:5n3 EPA 5.38 5.54 + 3%
C22:0 Behenic * 5.37 *
C22:1n9 Erucic * 0.54 *
C22:2n6 - - - 0.56 1.81 +223 %
C22:6n6 - - - 1.36 0.68 -50 %
C22:6 n3 DHA 0.47 3.80 +708 %

aData courtesy of Service Aqua, LLC

Enriched Artemia biomass was first used as maturation feed around 1988 in an effort to offset the rising cost of wild marine worms. Enriched Artemia has a good fatty acid profile and a high carotenoid content. Squid is a staple in maturation diets and is considered important in promoting maturation in both sexes. The high nutritional value of squid is attributed not only to its HUFA content but also to its amino acid composition and high sterol levels (Wouters et al., 2001). Mendoza et al. (1997) provided evidence for the presence of a hormonally active compound in squid which promotes ovarian maturation in P. vannamei. Whole squid labelled for human consumption is preferred to that sold as bait, as the former is of higher microbiological quality and free from impurities. Other fresh feeds (such as mussels, clams, krill and oysters) are practical only if locally available and inexpensive. They are used to supplement the primary HUFA components from marine worms and Artemia biomass. Feeding raw crustacean products presents a biosecurity threat, as they can be vectors for shrimp diseases, especially viral pathogens. In an attempt to diminish this risk, some operators cook or irradiate certain fresh feeds.

Although fresh feeds provide good nutrition for broodstock in maturation facilities, they can also be problematic due to variations in quality, availability and cost. They require cold storage and labour-intensive preparation and present a risk for disease transfer. All of these drawbacks make the replacement or partial substitution with dry feeds desirable. The development of a defined diet to meet the nutritional needs of broodstock shrimp has been a subject of research for many years (Harrison 1990, 1997; Wouters et al. 2001), and a number of researchers, farmers, and feed companies have attempted to develop a dry or semi-moist, practical maturation diet for penaeid shrimp. However, attempts to completely replace fresh food with artificial diets have resulted in decreased ovarian maturation, reduced number of spawns, or inferior larval quality (Wouters et al., 2000). Considered primarily as supplements to fresh ingredients, formulated diets are used at most maturation facilities and are available from commercial feed suppliers (Table 2).

Table 2. Commercial shrimp maturation dry-feeds.
Feed and manufacturer Type
Ziegler 3/32” Maturation Pellet, Ziegler Bros, Inc. USA Dry pellet
Ziegler EZ-Mate, Ziegler Bros, Inc. USA Dry premix powder
MadMac-MS®, Aquafauna Biomarine Inc. USA Dry premix powder
Higashimaru, Higashimaru Co. Japan Dry pellet
Royal Oyster®, Bernaqua. Belgium Dry extruded
Rangen Maturation, Rangen, Inc. USA Dry pellet

Quality and freshness of feed items are important for maturation. In order to ensure freshness, frozen inventories should be used within two to three months. Feeds should be defrosted and used immediately. Feeding regimes typically are calculated on a percent body weight basis. In a survey, 59% of respondents fed their broodstock on a percent body weight basis, whereas 38% fed to satiation (Moss and Crocos, 2001). As a general rule, broodstock should be fed 18-30% of the biomass per day (wet weight of feed). The relative contribution of the different feed types can be broken down as follows: 14% bloodworms or sandworms, 48% squid, 8% krill, 28% enriched Artemia biomass and 2% dry feed. Broodstock typically are fed five times per day as follows: 0700 h Artemia; 1000 h squid; 1300 h polychaetes; 1500 h squid; 2300 h krill and dry feed. Excess feed should be siphoned out of the maturation tanks after 2 to 3 h in the tank to avoid deterioration of water quality and tank bottom and feeding quantities should be adjusted daily based on consumption.

Eyestalk ablation

Panouse (1943) reported that ovarian development in the caridean shrimp, Leander serratus, was accelerated by ablating the eyestalk of adult females. This technique was later applied to penaeid shrimp (Caillouet, 1972; AQUACOP, 1975), thus paving the way for controlled reproduction. Broodstock-size female P. vannamei (35–50 g) typically are conditioned to maturation routines and diets 10 – 20 d before ablation. In properly conditioned females, spawning normally occurs four to seven days post-ablation. During this time, weight of the ovaries increases four to nine fold (Mourente and Rodriguez, 1991). Spawning then occurs every 7 to 14 days for a period of 3 to 6 months, before the quantity and quality of the spawn decrease to commercially unacceptable levels. Unilateral eyestalk ablation is practiced in maturation facilities throughout the world with notable exceptions in Colombia and Venezuela (Kawahigashi, 1992). Respondents to a survey of maturation facilities in these two countries reported achieving mating frequencies greater than 4% per night and nauplii production comparable to that of ablated females, without the need for ablation. These results have been attributed to domestication of the broodstock.

From a practical standpoint, ablation should be considered the last push towards sexual maturation. It should be done only after broodstock are the correct age and weight, have fully recovered from shipping and acclimation stress, have been on a maturation photoperiod and diet for one or two weeks, and have started to mature on their own without ablation. Ablating recently moulted or “soft” females should be avoided as it is often lethal to the shrimp. In addition to shrimp reproduction, eyestalk ablation impacts other physiological and metabolic processes. For example, Sainz-Hernández et al. (2008) reported that eyestalk ablation in P. vannamei resulted in a decreased duration of the moult-cycle and affected the concentrations of several hemolymph metabolites important in modulating the immune response, including the phenoloxidase system.

Although unilateral eyestalk ablation is now an industry standard, other endocrine manipulations have been investigated as promoters of ovarian maturation. The use of non-ablated females could lead to increased spawn quality, reduced labour and longer reproductive life of broodstock. Administration of compounds related to endocrine functions has been tried with varying degrees of experimental success, but as of yet, no commercial applications appear viable. Among the compounds tested are: methylfarnesoate (MF), serotonin, endocrine gland extracts, steroid hormones (e.g. 17-hydroxy-progesterone), retinoids, eyestalk peptides and antibody-inhibiting hormones (Racotta et al., 2001). MF is a terpenoid hormone produced in the mandibular organ of shrimp; it plays a central role as a regulator of oocyte development (Huberman 2000; Laufer and Biggers 2001). Laufer (1992) reported increased egg production, fertility and survival in broodstock fed diets supplemented with MF, and Tsukimura and Kamemoto (1991) found that the diameter of oocytes from isolated immature shrimp ovaries was significantly increased by MF. However, MF also has been shown to inhibit certain aspects of shrimp reproduction, indicating that its role may be complex and variable (Marsden et al. 2008).

It has been reported that the neurotransmitters serotonin (5 Hydroxy-Tryptamine; 5-HT) and dopamine (DA) affect ovarian maturation and ovulation in the freshwater crayfish Procambrus clarkii (Sarojini et al., 1995), the white shrimp, P. vannamei (Vaca and Alfaro 2000) and the tiger prawn, P. monodon (Wongprasert et al., 2006). Fingerman (1997) reported that 5-HT inhibits MF synthesis, but stimulates release of gonad-stimulating hormone (GSH). DA inhibits gonadal maturation in both sexes by inhibiting the release of GSH. Alfaro et al. (2004) demonstrated that ovarian maturation can be stimulated by a combined injection of 5-HT and the DA antagonist spiperone. The amino acid tryptophan is a precursor of 5-HT and there may be some rationale for adding this amino acid as a dietary supplement to stimulate elevated levels of 5-HT, thereby promoting ovarian maturation.

Metrics of reproductive performance

Reproductive performance of individual shrimp is highly variable (Ibarra et al. 2007). It has been noted that a small proportion of the females spawn multiple times and are responsible for the majority of the offspring, whereas a larger percentage of females spawn infrequently and contribute only a small proportion of the nauplii (Bray et al., 1990; Wyban and Sweeney, 1991, Ibarra et al., 2007). Furthermore, it has been shown that multiple spawns by a single female do not result in decreased egg or nauplii quality (Browdy and Samocha, 1985; Otogalli et al., 1988; Palacios et al., 2000b; Arcos et al., 2003; Arcos et al 2004; Palacios and Racotta, 2003). Reproductive performance can be quantified or monitored using the following metrics:

• Maturation rate. This is the rate of ovarian development of females and is normally expressed as the daily average percentage of the female population reaching stage IV ovarian development during a specified time period.

• Days from ablation to first spawn (latency period).

• Mating rate. The daily average percentage of the female population that gets mated during a specified period of time. This trait is influenced by both females and males. Mating rates of 6-12% are common in commercial facilities.

• Spawning rate. The daily average percentage of the female population that spawns during a specified period of time. This trait is sometimes expressed as the average number of days between spawns.

• Fecundity. The average number of eggs per spawn. This is a female trait.

• Fertility rate. The proportion of eggs that are fertilized. Fertility is influenced by both females and males

• Hatching rate. The proportion of fertilized eggs that successfully hatch. This rate is influenced by both females and males.

For practical reasons, the previous metrics often are grouped into: a) spawning rate and b) nauplii per spawner, or combined in a single index such as Nauplii per Female per Month (NFM), calculated as: NFM = (Spawning rate/100) X Nauplii per Female per Spawn X number of days in a specified month.

Selective breeding to improve reproductive performance

The primary traits of interest for shrimp breeders are growth and resistance to viral pathogens (Clifford and Preston 2006; Moss and Moss, 2009). These traits typically are polygenic and may be controlled by hundreds or thousands of genes (Tave, 1993), although a few genes may have disproportionately large effects on an animal’s phenotype with many genes having smaller effects (Orr, 1999). In addition to genetic influences, an organism’s phenotype is determined by environmental factors and by the interaction between its genes and the environment. Environmental factors relevant to shrimp breeders include stocking density, water temperature, water quality and diet, among others.

Heritability (h2) describes the percentage of phenotypic variance that is inherited in a predictable manner (Tave, 1993), and h2 estimates range from zero to one. Traits with h2 approaching zero are not very heritable and respond better to family selection than to individual selection, whereas traits with h2 approaching one are highly heritable and are amenable to both individual and family selection. Reproductive traits commonly exhibit low heritability (Falconer and Mackay, 1996; Tave 1993; Table 3) and therefore respond better to family selection than to individual selection. Estimates of h2 are important when designing breeding programs, predicting responses to selection, or calculating an individual’s breeding value.

Table 3. Heritability estimates (± standard error) for reproductive traits in penaeid shrimp.
Species Reproductive trait Heritablity ± SE Reference
P. vannamei Fecundity
Days to first spawn after
eyestalk ablation
0.17 ± 0.24
0.54 ± 0.25
Arcos et al. (2004)
P. vannamei Number of spawns
(spawning rate)
0.20a Ibarra et al. (2005)
P. monodon Days between spawns
Nauplii per spawn
Hatching rate
0.47 ± 0.15
0.41 ± 0.18
0.27 ± 0.16
0.18 ± 0.16
Macbeth et al. (2007)

95% confidence interval 0.06-0.43

Published h2 estimates for eggs per spawn (fecundity) range from low to moderate. Arcos et al. (2004) working with P. vannamei reported h2 estimates of 0.09 ± 0.23 (mean ±SE) and 0.17 ± 0.24 when adjusted for the time in the maturation tanks before spawning, whereas Macbeth et al. (2007) reported a higher estimate of 0.41 ± 0.18 for P. monodon (Table 3). It is important to note that h2 estimates for a given trait are not immutable and can change for a number of reasons including variations in gene frequencies within a population, environmental changes, age of the animals, interspecific differences and measurement errors.

Ibarra et al. (2005) reported a h2 estimate of 0.20 (95% CI = 0.06-0.43) for number of spawns in P. vannamei, and Macbeth et al. (2007) reported low to moderate h2 estimates (± SE) for number of viable nauplii/spawn (0.27 ± 0.16) and hatching rate (0.18 ± 0.16) in P. monodon. These values of h2 could be biased because the traits were assumed to be influenced exclusively by the females. See Ibarra et al. (2007) for additional h2 estimates of reproductive performance and for a review of the quantitative and molecular genetics of reproductive performance in penaeid shrimp.

Despite low to moderate h2 estimates for reproductive traits in penaeid shrimp it may be possible to improve these traits by family selection. Improvements may be made because reproductive traits tend to be highly variable. For example, Macbeth et al. (2007) measured days from ablation to first spawn (latency period), number of eggs/spawn (fecundity), number of nauplii per spawn and hatching rate for P. monodon and reported coefficients of variation (CV) of 34, 35, 68 and 71%, respectively. Similarly, Arcos et al. (2004) reported large CVs for latency period (83%) and eggs/spawn (35%) for P. vannamei. The large CVs along with the heritabilities reported by Macbeth et al. (2007) and Arcos et al. (2004) suggest there is sufficient genetic variation to make significant improvements in the reproductive traits of penaeid shrimp.

Broodstock health and disease

All diseases that affect adult shrimp can be present in maturation systems. However, little is known about the sub-lethal effects of adult shrimp diseases on reproduction. Aranguren et al. (2006) demonstrated that necrotizing hepatopancreatitis (NHP) in spawners affects both maturation and larviculture, causing a decrease in the number of eggs/spawn, a reduction in the levels of triglycerides in nauplii and a decrease in growth and resistance to osmotic stress at the 10 day-old postlarvae (PL-10) stage. Table 4 lists common diseases that may occur in maturation facilities and chemical treatments used for their control. Two conditions of special significance to broodstock will be discussed below.

Table 4. Common treatments used to control pathogens in maturation facilities.
Pathogen Treatment Duration
Bacterial infection in eggs 100 ppm formalin dip 30 sec
Bacterial infection in eggs 100 ppm Povidone Iodine dip 30 sec
Bacterial infection in nauplii 100 ppm Povidone Iodine dip 5 min
Fungus in eggs or nauplii 0.05-0.1 ppm Treflan TR-10 Every 6 h
Filamentous bacteria in broodstock gills 14 ppm Cooper Control dip 1 h
Protozoan infestation in broodstock gills 150 ppm formalin dip 1 h

Reproductive tract degeneration syndrome, also known as black spermatophore syndrome, causes necrosis and melanization of the spermatophores and can render affected males infertile (Chamberlain et al., 1983). The condition is characterized by a brown to black discoloration of the terminal ampoule, spermatophore and vasa deferentia, as well as inflamed spermatophores. Several circumstances favour the development of black spermatophores, including organic matter accumulation on tank bottoms and temperatures higher than 30°C. Maintaining males and females in separate tanks tends to reduce ejaculation and regeneration of healthy spermatophores. Although the cause of black spermatophore syndrome is unknown, Carr et al. (1995) reported that broodstock males with black spermatophores were not responsive to oxytetracycline therapy and suggested a non-infectious primary aetiology.

Filamentous bacteria and/or sessile protozoans can colonise the gills and other body surfaces of broodstock, inhibiting respiration and leading to hypoxia and death. This condition is favoured by high levels of organic matter in the water, poor sanitation and factors that inhibit moulting (i.e. sub-optimal rearing temperatures). Fouling disease can be prevented by practices that promote clean water and by adequate sanitation of facilities and equipment. Once present, the condition can be treated with copper compounds or formalin as shown in Table 4.

Larval rearing

After initial work with the Kuruma prawn, P. japonicus, by Hudinaga (1935), the life history of penaeid shrimp was studied in detail by Heldt (1938) who described the larvae of three Mediterranean species. Hudinaga (1942) described larval rearing of P. japonicus in captivity, effectively pioneering shrimp larviculture. Larval development of P. vannamei was described by Kitani (1986) and was found to be similar to that of related brown shrimp, P. californiensis and blue shrimp, P. stylirostris. Developmental stages include nauplii, protozoea, mysis and postlarvae (PL), all of which are described below.


Fertilized eggs of P. vannamei are 260-290 µm in diameter, tend to sink slowly and undergo a complex process of embryonic development which culminates in hatching of a nauplius larvae 14-16 hours after spawning (at 28 ºC). Fig. 2 shows late eggs and recently hatched nauplii of P. vannamei.


Figure 2. Eggs and recently hatched nauplii of white shrimp (Penaeus vannamei). Note the hatched egg membrane in the lower center and the hatching nauplii in the lower right portion of the photo.


Nauplii are the first larval stage of penaeid shrimp and this stage is common among all crustaceans. The name “nauplii refers to crustacean larvae that use appendages originating from the head (antennules and antennae) for swimming. Nauplii have only one compound eye which divides into two at a later stage. Nauplii do not feed but use

their internal yolk reserves for energy. Kitani (1986) described six naupliar sub-stages for P. vannamei, but subsequent workers recognise only five. Nauplii grow from an initial body length (BL), excluding furcal spines, of 330 µm and body width (BW) of 190 µm, to a final BL of 430 µm and BW of 220 µm in 36-40 h at 28-30 ºC. Nauplii swim intermittently and are positively phototactic, a behaviour that is useful for their harvesting. In hatcheries, nauplii often are checked under the microscope for deformities of the antennae and furcal setae, which may result from changes in temperature during development or from metal toxicity. The size, shape and biochemical composition of the yolk sac is influenced by the nutrition of the mother, and these traits may be used as an indication of nauplii quality, with large orange or reddish yolk sacs reflecting better quality. Palacios et al. (1998) reported a positive correlation between levels of triacylglycerol and glucose (but not of cholesterol) and naupliar survival to the protozoeal stage.

Simões et al. (2002), using histology and transmission electron microscopy, reported the presence of an open anal pore in nauplii-5 of P. vannamei. This pore, combined with anti-peristaltic movements (“anal drinking”), allows early bacterial colonization of the digestive tract. These findings have implications to larval culture as beneficial or harmful bacteria can be introduced to the digestive tract of the larvae even before the mouth and the digestive tract become fully functional.


The first protozoea has a flattened body which is clearly divided into carapace and abdomen. Compound eyes are present but they do not extend beyond the carapace. The digestive tract is fully formed and functional, exhibiting peristaltic movements that start from the oesophagus region. Protozoea swim continuously and are capable of vigorous feeding immediately after moulting. The second protozoea has a rostrum, prominent stalked compound eyes and forked supra-orbital spines. Average length is 1.3 to 2.15 mm. The third protozoea is characterized by the appearance of biramous uropods and spines on the dorsal and lateral sides of abdominal segments. Average length is 2.1 to 2.7 mm. Fig. 3 shows the three protozoeal sub-stages of P. vannamei.


The change to first mysis is accompanied by drastic changes in appearance and behaviour. The body shape of the larvae now resembles that of an adult shrimp (Fig. 4). Mysis swim head-down with vigorous movements of the tail. Strong mysis tend to swim towards the surface of the water. Feeding habits change from herbivorous in protozea (phytoplankton)


Figure 3. Protozoea of white shrimp (Penaeus vannamei). A 1st – 3rd protozoea; B 1st antenna; C 2nd antenna; D mandible; E 1st maxilla; F 2nd maxilla; G 1st maxilliped; I lateral view, 3rd protozoea. Scale bar denotes 0.5 mm. From Kitani (1986). Reprinted with permission.

to carnivorous (zooplankton) in mysis. In the first mysis, rudimentary pleopod buds appear on the ventral side of the first five abdominal segments. A bifurcated telson appears. Body length is 2.5 to 3.5 mm. In the second mysis, the pleopod buds are elongated but remain un-segmented and slightly curved. Body length is 3 to 4 mm. By the third mysis, the pleopods become segmented and are used to create a current which directs water and food towards the pereopods and mouth. Body length is 3.6 to 4.4 mm.


The first PL are characterized by setae in the swimming legs. They exhibit a radically different swimming behaviour moving forward, and they are able to maintain their position or move at will in the water. They also become increasingly benthic and tend to swim against water currents, with both behaviours getting more pronounced over time. Body length of the first PL can vary from 3.5 to 4.3 mm. It is customary to refer to young PL by the number of days after metamorphosis, i.e., PL-1 for one day old postlarvae, etc. PL age does not refer to a specific moult stage or size, but only to the days after reaching first postlarvae. As PL grow, they become larger, and they also develop more complex gill structures (Fig. 5), which allow for more efficient gas exchange and osmotic regulation.


Figure 4. Mysis and postlarvae of white shrimp (Penaeus vannamei). A lateral view; B 1st antenna; C 2nd antenna; D telson, E 1st maxilla; F 2nd maxilla; G 1st maxilliped; H 2nd maxilliped; I 1st pereiopods; J 4th pereiopod. M1-M3 1st – 3rd mysis. PL postlarvae. Scale bar denotes 0.5 mm. From Kitani (1986).
Reprinted with permission.




Figure 5. Gill development in postlarvae of white shrimp (Penaeus vannamei) in relation to age in days.
A: 1 – 4 days old; B: 5 – 7 days old; C: 8 – 10 days old; D: 11- 14 days old.

Overview of larval culture systems

Larval rearing tanks (LRTs) can be constructed in a variety of materials, shapes and sizes. Tank material greatly affects cost, durability, thermal insulation and surface properties. The most common material used for LRTs is concrete block, often lined with an inert plastic membrane. Fiberglass tanks were common in the past but have been replaced because of their relatively high cost, lack of thermal insulation and high maintenance requirements. Tanks with “V”- or “U”-shaped bottoms favour good water circulation, but larval culture is possible in flat-bottom tanks, provided there is sufficient aeration to move the water effectively and prevent “dead zones” where organic matter can accumulate. Commercial hatcheries typically use tanks ranging in volume from 5 to 30 m3, although larger tanks have also been used (Juarez and Fegan, 2006).

As with broodstock, the health and performance of shrimp larvae depend on high-quality seawater. Important water quality requirements include low concentrations of ammonia and nitrite, high dissolved oxygen and appropriate pH levels (Table 5). The extent to which a hatchery’s intake water is treated depends on the quality of the source. General treatment strategies include:

Table 5. General water quality requirements for shrimp hatcheries.
Variable Acceptable range
Temperature 27-29 ºC
Salinity 30 to 34 ppt
pH 7.6 - 7.8
Dissolved oxygen >5 ppm
NH3-N <0.01 ppm
NO2-N <0.1 ppm

a Some hatcheries use a high temperature regime

(29-34 ºC) for larval rearing

Degasification: This may be needed if the source water contains undesirable gases, such as H2S or CO2. Degasification is achieved using columns packed with plastic media and counter-current air flow from air blowers.

Mechanical filtration: Hatchery water should have low concentrations of total suspended solids (TSS). Particle filtration is used to reduce TSS loads by stripping intake water of progressively smaller-sized particles. In hatcheries with a beach intake, a well point screen (finely slotted well casing) buried in the sand is the first line of filtration, normally followed by some combination of settling reservoirs, sand, cartridge, bag and/or diatomaceous earth filters. In general, maturation water should be filtered to 10 µm, whereas larval-rearing water should be filtered to 0.5-5µm.

Disinfection: Before intake water is used to fill the LRTs, it should be disinfected to minimise the introduction and spread of pathogens or their vectors. Disinfection of intake water is often accomplished by adding 0.5-20 ppm of free chlorine, which can be neutralized with sodium thiosulfate, ascorbic acid, or allowed to volatilize with heavy aeration. Ozone is another disinfectant commonly used in hatcheries, but care must be taken to avoid overdosing which can cause the production of various persistent oxidants, collectively referred to as Total Residual Oxidants (TRO). A general rule of thumb for a safe TRO concentration in invertebrate culture is <0.02 mg/L, which can normally be prevented by limiting ozone injection to that resulting in an oxidation-reduction potential (ORP) of 400 mV or less (Aiken and Smith, 2004). ORP is used only as an indicator, as there is no direct correlation between ORP and TRO. Ozonation also helps eliminate colloidal solids, refractory organic matter and nitrite. Ultraviolet sterilizers also are common means of water disinfection in shrimp hatcheries and often are used as the last step in water treatment.

Absorption: If intake water has unacceptable levels of dissolved organic matter, adsorption-based equipment, such as protein skimmers or granular activated carbon (GAC) filters may be used. GAC filters also help to remove heavy metals and undesirable by-products of ozonation.

Plumbing: If left unchecked, hatchery plumbing can easily become a reservoir for pathogenic bacteria. In well-designed hatcheries, water pipes are installed at a slope and plumbed with threaded tees instead of elbows so they can drain completely and be cleaned with a pipe brush. Most hatcheries implement procedures to disinfect plumbing routinely. Critical plumbing is disinfected with chlorine weekly, and less critical water lines are scheduled for monthly or bimonthly disinfection.

Water management: The health and performance of larvae in LRTs depends on high-quality water, which can be maintained by water exchange through screens of different sizes, based on larval stage. As larvae get older and larger, the mesh size of the screen increases (Table 6). Food must be added to the LRTs in sufficient quantities to replace food consumed by the larvae and food flushed during water exchange. A healthy bacterial community depends on a careful balance among the carrying capacity of the tank, microalgae, other exogenous feed and water replacement. Although frequent and high rates of water exchange promote healthy larvae, they are wasteful with regard to water, feed and heat. High exchanges also have the potential to destabilize the structure of the microbial community and create new niches for pathogenic bacteria (Browdy, 1998). In response to these disadvantages, two alternative approaches to larval rearing have been developed. One approach relies on bio-filtration and water re-use, whereas the other requires the use of probiotics and high-density culture. These approaches will be described below.

The use of bio-filtration to recycle water during larval rearing can reduce water use, but it also presents unique problems (Menasveta et al., 1989; Millamena et al., 1991; Gandy, 2004). For example, the duration of larval rearing is short (~18 days for P. vannamei) relative to the time needed to establish a mature biological filter. Bio-filtration systems present advantages of increased biosecurity, reduced feeding and heating costs and reduced environmental impact of effluents, but their use has been mostly experimental, with few exceptions. In Ecuador, Industrias Bioacuaticas S.A. (INBIOSA), a commercial shrimp hatchery, operated

Table 6. Water-exchange regime and screen sizes used to retain larvae in the flow-through method of raising shrimp larvae.
Larval stage Water exchange (% per day) Screen opening (µm)
Nauplii 0 -
Protozoea-2 50 200
Protozoea-3 50 200
Mysis-1 50 200
Mysis-2 75 300
Mysis-3 75 300
Postlarvae-1 150 500
Postlarvae 2 - 9 200 500
Postlarvae 10 - 12 200 600
Postlarvae 13 - 20 200 700

a recirculating larval-rearing system during 1989-90 under the technical guidance of France Aquaculture (J. Baquerizo and I. Morales, personal communication). Each LRT was connected to a biofilter with crushed coral rock as media. Water from the LTR was recirculated through the bio-filter using an air-lift pump. Coral media was soaked in chlorinated water between larval batches, rinsed and “recharged” in biofilter “booster” tanks supplied with inorganic ammonia. Bio-filters were connected to LRTs when the larvae reached the protozoea-3 stage and were maintained in recirculation for 10 days until the postlarvae were transferred to outdoor raceways. The system performed well and resulting postlarvae were hardy, homogeneous in size and of general good quality. During 2004-2007, Ocean Boy, an intensive shrimp aquaculture project in South Central Florida, operated a commercial hatchery based on similar principles (J. Quintana, personal communication). The system was successful in producing good quality postlarvae for the farm’s intensive heterotrophic ponds.

Recently, a novel, high-density larval-rearing method, using probiotics and no water exchange, has been used by commercial hatcheries in Brazil and Mexico. With this method nauplii are stocked in LRTs at densities of 300 to 500 nauplii/L, which are three to five times higher than the industry standard of 100 nauplii/L. Commercial probiotics and nitrifying bacteria are incubated separately and added daily to the LRTs to maintain detrimental bacteria and nitrogenous compounds under control. The initial water volume is raised to full tank capacity during the first few days but no water is exchanged until the early postlarval stages. Although this method often results in lower than normal survival, production per tank is greatly increased and production costs are reduced due to savings in feed, water pumping and heating.

Larval feeding

In general, protozoea are herbivorous and require microalgae, whereas mysis and PL are carnivorous and require Artemia nauplii. Postlarvae are omnivorous and can eat both plant and animal matter. Dry and/or microencapsulated feeds are used to supplement live feeds. For feeding protozoea, traditional species of microalgae include Tetraselmis (Platymonas) and Chaetoceros spp. Piña et al. (2005) showed that a mono-algal diet based on Chaetoceros muelleri is an acceptable option for early larval feeding of P. vannamei. Barbarito et al. (2006) reported that feeding a combination of microalgae results in better larval nutrition and survival than feeding a single algal species, an observation confirmed by experiences in commercial hatcheries. In the last few years, the use of alternate microalgae such as Thallasiosira and Skeletonema has become common. The influence of algal quality on larval rearing cannot be overemphasized. Only algal cultures in their active growth phase should be used. Two successful routines for feeding shrimp larvae with different algal mixtures are presented in Table 7. Algal-culture methods include batch, semi-continuous and continuous systems. Although not essential for nutrition, some hatchery managers prefer to have microalgae in the culture water during the mysis and early postlarval stages, not only because they provide excellent supplemental nutrition through incidental or indirect ingestion, but also because they enhance water quality by absorbing nitrogenous compounds and carbon dioxide, provide shading and help stabilize the bacterial flora of the water.

Table 7. Algal-feeding routines used in the larval culture of white shrimp (Penaeus vannamei).


Routine 1 (cells/mL) Routine 2 (cells/mL)

4041.png 4043.png

Stage/Algae Chaetoceros Tetraselmis Thalassiosira Tetraselmis

gracilis suecica weisflogii suecica

4048.png Nauplii 60,000 40,000

Protozoea-1 80,000 40,000

Protozoea-2 100,000 20,000 80,000

Protozoea-3 100,000 20,000 80,000 20,000

Mysis-1 80,000 20,000 80,000 20,000

Mysis-2 60,000 20,000 40,000 20,000

Mysis-3 60,000 20,000 40,000 20,000

Postlarvae 1-10 40,000 20,000 40,000 20,000


Freshly hatched Artemia nauplii are an excellent food for mysis and one of the main sources of lipids for larval shrimp (see Table 8 for an effective feeding regime). Artemia usually are fed live, but some hatchery operators prefer to feed frozen Artemia, or to kill them by submersing in 80°C water for 30 sec. This approach has the added advantage of reducing bacterial load. Killed Artemia can be fed to larvae as early as protozoea-2. Puello-Cruz et al. (2002) demonstrated a variable larval enzyme response starting at protozoea-2 and suggested that P. vannamei are physiologically adapted to a more carnivorous diet after this stage. Larvae fed with Artemia starting at protozoea-2 reach mysis earlier than those fed exclusively with microalgae. Common usage rates vary from 0.05 kg of Artemia cysts per million larvae per day at protozoea-2 to 0.25 kg at mysis-3 and from 0.3 kg in PL-1 to 0.5 kg in PL-5. Fluctuations in supply and variations in cyst prices and quality continue to stimulate interest in inert and alternative live feeds which could serve as complete or partial replacements for Artemia cysts. Zelaya et al. (2007) reported no significant difference in survival between postlarval shrimp fed Artemia and those fed good-quality dry diets. Alternative live feed replacements for Artemia include nematodes, Panagrellus redivivus (Biedenbach et al., 1989), rotifers Brachionus plicatilis (Samocha et al., 1989) and copepods, Apocyclops dengizicus (Farhadian et al.,2009). Despite efforts to develop alternative diets, Artemia nauplii remain the staple feed for mysis and early postlarval shrimp.

Table 8. Artemia-feeding regime used in the larval rearing of white shrimp (Penaeus vannamei).


Larval stage Average number of Artemia nauplii consumed per shrimp larvae or postlarvae per daya


Protozoea-2 1.5 b

Protozoea-3 8 b

Mysis-1 15

Mysis-2 21

Mysis-3 25

Postlarvae-1 30

Postlarvae-2 40

Postlarvae-3 50

Postlarvae-4 57

Postlarvae-5 64

Postlarvae-6 70

Postlarvae 7-10 76


a Daily amount is normally divided in two to four feedings per day. Some of the daily feedings can consist of heat-killed or frozen Artemia nauplii.

b Protozoea-2 and -3 usually are fed with killed Artemia nauplii.

Formulated feeds are used in larval rearing as a supplement to live feeds, as means of introducing specific nutrients and to reduce the quantity and expense of Artemia cysts. Commercial feeds are available in a variety of brands, particle sizes and presentations (Table 9). Most hatchery managers prefer to mix different feeds for the same stage rather than depend on a single product. Table 10 presents a generalized dry-feed schedule for shrimp larvae, but specific amounts should be determined based on manufacturer’s recommendations, careful observation of feeding activity, water quality and experience. It is important to avoid overfeeding as this may lead to deterioration of water quality and to larval disease and mortality. Most hatcheries divide the daily rations into at least four feedings per day and some feed every one or two hours. Frequent feeding promotes ingestion of fresh feed and helps avoid deterioration of water quality.

Table 9. Common commercial feeds used in the larval rearing of shrimp.


Feed and manufacturer Type


EZ Larva®, Ziegler Bros, Inc. USA Microencapsulated liquid

Larval AP100®, Ziegler Bros, Inc. USA Dry powder

Z Plus®, Ziegler Bros, Inc. USA Dry powder

Algamac®, Aquafauna Biomarine, Inc. USA Dry powder

Artemac®, Aquafauna Biomarine, Inc. USA Dry powder

Micromac®, Aquafauna Biomarine, Inc. USA Dry powder

Frippak®, INVE Aquaculture. Belgium Microencapsulated powder

Lansy-shrimp®, INVE Aquaculture. Belgium Dry powder

Microfeast®, Salt Creek, Inc. Salt Lake City. USA Microencapsulated powder

Progression®, Salt Creek, Inc. Salt Lake City. USA Microencapsulated powder

Epifeed®, Epicore, Eastampton. USA Microencapsulated liquid

Liqualife®, Cargill, Minneapolis. USA Microencapsulated liquid

Biospheres®, Bernaqua. Belgium Microencapsulated liquid

Royal Caviar®, Bernaqua. Belgium Dry powder

Royal Seafood®, Bernaqua. Belgium Dry powder

Royal Pepper®, Bernaqua. Belgium Dry powder

Golden Pearls®, Bernaqua. Belgium Microencapsulated powder

P. vannamei hatchery feed, Higashimaru Co. Japan Dry feed


Table 10. Typical dry-feed regime used to supplement microalgae and Artemia in the larval rearing of white shrimp (Penaeus vannamei).


Larval stage Dry feed Particle sizes

(g/m3/d)a (µm)

4502.png Protozoea-1 5 5-30

Protozoea-2 7 5-30

Protozoea-3 9 5-30

Mysis-1 16 30-50

Mysis-2 22 50-150

Mysis-3 24 50-150

Postlarvae-1 26 150-250

Postlarvae-2 28 150-250

Postlarvae-3 29 250-300

Postlarvae-4 29 250-300

Postlarvae-5 31 250-300

Postlarvae-6 32 250-300

Postlarvae-7 34 250-300

Postlarvae-8 35 250-300

Postlarvae-9 36 250-300

Postlarvae 10-15 37 250-500


aDaily amount is normally divided in four to twelve feedings per day.

Larval feeds have improved significantly in recent years. Commercial feeds have benefited from improved manufacturing techniques and novel ingredients. Digestibility, binding, water stability, attraction, palatability, size distribution and colour have improved significantly. In addition to traditional micro-particulate feeds, advanced micro-encapsulated and liquid-based feeds are now available for shrimp larvae. They not only have the advantage of being highly stable in water but also allow for the introduction of specific ingredients, like probiotics or immune boosters.

The potential to stimulate a shrimp’s immune system via feeds has generated much interest in recent years. The immune system of invertebrates is not well understood (Loker et al., 2004). It is generally assumed to lack a true specific response (Soderhall and Thornqvist, 1997; Thornqvist and Soderhall, 1997). However, some indications of specific immunity in penaeids have been suggested (Namikoshi et al., 2004; Witteveldt et al., 2004; Bright-Singh et al., 2005; Rajeshkumar et al., 2009). The most important innate immune defence of crustaceans is the phenoloxidase (PO) system (Sritunyalucksana and Soderhall, 2000; Lee and Soderhall, 2002). Oral administration of the immune booster ß-1, 3 glucan has been shown to enhance PO activity and increase survival to viral exposure in P. monodon (Chang et al., 2003). Larval feeds supplemented with immune booster formulations are intended to enhance resistance to diseases and to increase larval survival. Several are commercially available and have been used with varying degrees of success.

Production routines

Most hatcheries obtain favourable results when modular larval-rearing rooms are stocked with discrete batches of larvae in an all-in, all-out sequence, followed by disinfection and dry-out procedures. Large hatcheries are thus designed with separate modules to allow for continuous production. Modules of LRTs are stocked with nauplii in a short period of time, normally two to four days.

Between-batch cleaning and disinfection should include tanks, plumbing, floors, walls and all relevant equipment. Table 11 outlines methods commonly used for hatchery sanitation. After drying for a period of three to ten days, LRTs can be filled to initial capacity (normally 50 to 75% of the final working volume) for stocking. It is important to ensure the initial fill water is properly filtered and disinfected (no bacterial growth on TCBS agar). Algae are introduced to the tanks once nauplii reach the 4th sub-stage.

Table 11. Methods used for sanitation of different components of hatchery systems.


Components Sanitation methodsa


Larval-rearing tanks After each cycle brush with detergent, rinse.

Sponge-in a solution of muriatic acid of pH 2, rinse well. Brush-in with 200 ppm chlorinated water

Rinse and dry (Never mix bleach and acid).

Building floors and walls Brush with detergent, spray with 100 ppm chlorinated water.

Water lines Disinfect weekly by running 100 ppm chlorinated water

Air lines Disinfect weekly by running 100 ppm chlorinated water

Miscellaneous equipment Submerge in 100 ppm PVP-Iodine or 100 ppm chlorinated water


aUse of these chemicals requires specialized training, careful observation of safety precautions and use of protective equipment such as respirators, goggles, boots and special clothing.

Assessment of larval stage and quality is the basis for determining appropriate management with regard to feeding, water exchange and other treatments. The simplest assessment is based on gross observations of the culture system and larval characteristics, and consists of a routine morning evaluation of tank and larval conditions. Walk-through evaluations are highly dependant on the skill of the observer and include:

• Verification of water volume, colour, turbidity and smell.

• Checking for presence of foam or organic debris.

• Confirmation that correct screens and water exchanges are in place.

• Verification and adjustment of aeration as needed.

• Collection of samples of larvae from tanks and observation with the naked eye for the following:

- Larval stage. An experienced operator can approximate the stage with the naked eye, but precise determination requires observation through the microscope.

- Larval activity and swimming behaviour.

- Feeding activity, larval behaviour and the presence of faecal strands.

- Presence and grade of faeces, Artemia nauplii, dry feeds and/or larval moults in the water.

- Larval mortality.

- Water temperature.

More detailed observations require a trained operator and use of a microscope. These include:

• Checking quality and quantity of microalgae.

• Staging the larvae.

• Examination of the larvae for colour and muscle opacity (expanded chromatophores and/or opaque muscle tissue are signs of stress or disease).

• Assessment of the condition of the hepatopancreas. This is an indication of larval feeding and digestion. It can be observed in wet mounts on a microscope at 40X magnification. In healthy and well fed larvae, the hepatopancreas should be full of small bubbles (digestive or “lipid” vacuoles).

• Examination of the gut and intestinal content and checking for peristaltic activity of the gut.

• Checking larvae for clinical signs of disease and conditions described in the following section.

Larval diseases


Larval shrimp are affected by diseases of viral, bacterial, fungal, protozoan and toxic aetiologies. Bacterial diseases, in particular, have a major impact on hatchery production. Many diseases can be prevented with good management practices.  In general, viral diseases are best prevented through exclusion of pathogens by using Specific Pathogen Free (SPF) stocks and practicing biosecurity. Non-excludable diseases, like those caused by ubiquitous bacteria and fungi, are best prevented by sanitation and maintenance of good rearing conditions. Principal diseases of concern to hatcheries are described below.

 Baculovirus penaei (BP) is an occluded, double-stranded DNA virus that affects shrimp mainly in the protozoea, mysis and early postlarval stages (Couch, 1974). Epizootics often result in high mortality. The disease can be diagnosed by observing the presence of prominent occlusion bodies, readily observable by direct microscopy in squash preparations of the hepatopancreas, midgut or faeces (Fig. 6). Occlusion bodies are tetrahedral or pyramidal in shape and range in size from less than 0.1 µm to nearly 20 µm from base to peak, with a modal vertical length of 8-10 µm (Lightner, 1996). In hatcheries, transmission of BP can occur in LRTs by ingestion of faeces, contaminated water, or by cannibalism of diseased shrimp and in spawning tanks via faecal contamination of the spawn by infected females. Prevention can be accomplished by techniques designed to interfere with transmission from spawning females to eggs. These procedures involve harvesting fertilized eggs from the spawning tanks, rinsing with abundant clean, ultraviolet-treated water and disinfecting with organic iodine compounds. The importance of BP as a major pathogen in shrimp hatcheries has declined significantly since the introduction of these techniques (OIE, 2000). Today, the presence of BP in larvae indicates that the sanitary management of the hatchery facility is inadequate.



Figure 6. Wet mount of feces from a larval white shrimp (Penaeus vannamei) infected with Baculovirus penaei. The tetrahedral occlusion bodies (arrows) are diagnostic for infection of the shrimp’s hepatopancreas or midgut epithelial cells. Phase contrast, no stain. Magnification 700X.
Photo Courtesy of DV Lightner, University of Arizona, Aquaculture Pathology Laboratory.

Bacterial diseases are the most common cause of mortality in shrimp hatcheries. Bacteria most often associated with disease belong to the genus Vibrio. However, other Gram-negative bacteria, belonging to the genera Aeromonas, Pseudomonas and Flavobacterium, are occasionally diagnosed as disease-causing agents. Adverse environmental conditions, handling stress, poor nutrition or mechanical injuries are important predisposing factors for bacterial disease.

 In the past, hatchery managers and growout farmers relied on antibiotics to prevent or treat bacterial outbreaks (Reed et al., 2004), but there is growing concern about the use of antimicrobial drugs in aquaculture (Verschuere et al., 2000). Widespread use of antibiotics in shrimp aquaculture has led to the detection of antibiotic residues in farmed shrimp (Nogueira-Lima et al. 2006) and to the establishment of antibiotic-resistant strains of bacteria (Nakayama et al. 2006). In addition to potential human health problems, antibiotics can kill both beneficial and pathogenic bacteria in the LRTs, which can destabilize microbial community structure (Brown, 1989). A stable microbial environment is important for the survival of larval shrimp. Otta et al. (2001) showed that Vibrio spp were less dominant in natural seawater (30% prevalence) than in LRTs (up to 73% prevalence). Because of concerns associated with antibiotics, their use is now discouraged, and efforts have shifted towards maintaining healthy conditions by the use of probiotics and by improving biosecurity, sanitation and frequency of dry-outs (Browdy, 1998).

Another strategy employed to mitigate bacterial problems in hatcheries is the use of probiotics, live microbial supplements designed to out-compete opportunistic pathogens and maintain a healthy environment. The use of probiotics in shrimp hatcheries started in Ecuador with the observation that LRTs experiencing disease showed a decrease in populations of Vibrio alginolyticus concurrent with an increase in V. parahemolyticus (Griffith, 1995). Experiments with locally isolated Vibrio strains confirmed the pathogenicity of V. parahemolyticus and the relative benign influence of V. alginolyticus. A system was developed for the routine isolation, mass culture and larval tank inoculation of benign bacterial strains (Garriques and Arevalo, 1995; Krauss et al., 1996). This approach has worked in some situations, but care must be exercised as different strains of V. alginolyticus can show marked differences in their effects on the larvae, from benign to pathogenic. Furthermore, the transfer of virulence factors among bacterial isolates has been reported, either by genetic processes (Pizzutto and Hirst, 1995) or by bacteriophages (Alday et al., 2000). Vandenberghe et al. (1999) reported high heterogeneity among V. alginolyticus strains, and suggested that putative probiotic and pathogenic strains have specific genotypes. For these reasons, most commercial probiotics consist of non-pathogenic bacteria such as Bacillus spp.   Probiotics have helped minimise bacterial problems, reduce hatchery down-time and increase survival. Probiotic additions can begin before the nauplii are stocked into LRTs and can continue throughout the rearing period, according to the manufacturer’s specifications. Successful use of probiotics depends upon careful evaluation, implementation and cost-benefit analysis.

Luminescent bacterial septicemia of shrimp larvae is one of the major causes of mortality in hatcheries (Lavilla-Pitogo et al., 1990). Larvae affected with luminous vibriosis glow blue-green when observed in complete darkness; therefore, nightly observation of larval rearing tanks by a trained observer is an excellent surveillance method for this disease. Microscopically, moribund larvae show swarms of motile bacteria in the body cavity. This disease often results in heavy mortalities. Studies at shrimp hatcheries in India have demonstrated that shrimp broodstock are the main reservoir of luminescent bacteria (Abraham and Palaniappan, 2004; Chrisolite et al., 2008). General sanitation and good management practices, such as egg washing and chemical disinfection, help prevent luminous vibriosis in LRTs. Presence of luminescent V. harveyi and/or related species during the nauplii and protozoea stages is an indicator of a possible outbreak of luminescent bacterial disease in subsequent larval stages. Control of luminescent bacteria using antibiotics has been effective in laboratory trials (Baticados et al., 1990) but may not be cost-effective under commercial conditions. Alternative approaches to combat vibriosis include sanitary practices and the use of disinfectants, immunostimulants and/or probiotics (Alday et al. 2006). Lio-Po et al. (2005) found anti-luminous Vibrio factors in bacteria, fungus, microalgae and tilapia skin mucus associated with the “green water” method of farming P. monodon in ponds in the Philippines. Notably, the diatoms, Chaetoceros calcitrans and Nitzchia sp., demonstrated inhibition of luminous Vibrio. Bacteriophages, viruses that infect and kill specific bacterial hosts, also have been suggested as potential alternatives to antibiotics in controlling bacterial infections (Sulakvelidze et al., 2001; Karunasagar et al., 2007; Okano et al., 2007). Vinod et al. (2006) reported microcosm studies with P. monodon larvae infected with luminescent V. harveyi, and showed that larval survival in the presence of bacteriophages was enhanced (80%) compared to that of controls (25%). Treatment with bacteriophages also improved larval survival and reduced luminescent V. harveyi counts in hatchery tanks.

 Non-luminous bacterial infections of larvae also can occur as localized or generalized infections. Affected larvae may show necrosis of the appendages, expanded chromatophores and reduced feeding (Fig. 7 and Fig. 8). Mortalities may be very high within a few days of infection. Common bacteria involved are: V. harveyi, V. splendidus, V. parahaemolyticus, V. alginolyticus, V. vulnificus, V. penaeicida, V. campbellii and V. anguillarum (Lavilla-Pitogo et al., 1990; Chen et al., 1992; Lightner et al., 1992; Limsuwan, 1993; Ruangpan et al., 1995; Hameed et al., 1996; Regunathan and Wesley, 2004). Microscopic demonstration of motile bacteria in the body cavity of moribund larvae and isolation and identification of pathogenic bacteria help in the diagnosis of the disease. Prevention consists of good sanitary practices, maintenance of good rearing conditions and reduction of the organic load in the rearing water. The addition of 10-15 ppm EDTA to the water reportedly has therapeutic effects against vibriosis in hatcheries (Pathak et al., 1996).

Another larval disease that results in hatchery mortalities is known as “bolitas” (Spanish for “little balls”). This syndrome is characterized by the detachment of epithelial cells from the intestine and hepatopancreas which slough off and appear as distinct spheres in the lumen of hepatopancreatic tubules or digestive tract   (Fig. 9). The syndrome also is characterized by reduced feeding and activity, retarded


Figure 7. Wet mount of bacterial necrosis from appendage tips of a postlarval blue shrimp (Penaeus stylirostris). No stain. Magnification 300X. Photo Courtesy of DV Lightner, University of Arizona, Aquaculture Pathology Laboratory.


Figure 8. Postlarval blue shrimp (Penaeus stylirostris) with vibriosis. Necrosis of several appendages (pleopods and pereiopods) is indicated by melanized foci or tips. A dark oral region is indicative of bacterial colonization of the cuticle of the oesophagus and mouth appendages. Wet-mount; no stain. Magnification 50X. Photo courtesy of DV Lightner, University of Arizona, Aquaculture Pathology Laboratory.



Figure 9. Protozoea-2 of P. vannamei displaying “bolitas” (= “little balls”). These ball-like structures are hepatopancreas or midgut epithelial cells that have rounded-up and sloughed into the gut lumen in response to toxins released from Vibrio spp. Photo Courtesy of Marisol Morales-Covarrubias, CIAD-Mazatlan, Mexico.

development and mass larval mortality. “Bolitas” are thought to be a reaction to bacterial toxins and/or possibly to heavy metals. Robertson et al. (1998) experimentally induced “bolitas” by exposing nauplii and zoea of P. vannamei to 105 cells/ml of Vibrio harveyi in the culture water and confirmed Koch’s postulates for the syndrome by re-isolation and identification of the bacterium. Esteve and Herrera (2000) also induced sloughing of the cellular lining of the hepatopancreas in response to experimental infections with V. alginolyticus. Vandenberghe et al. (1999) isolated eleven dominant strains of Vibrio associated with the “bolitas” syndrome from Ecuadorian hatcheries, including different strains of V. harveyi and V. alginolyticus. Prevention of “bolitas” in commercial hatcheries involves good sanitation and feeding practices, the use of probiotics and all-in, all-out stocking procedures, followed by dry-out and disinfection of facilities.

Zoea-II syndrome (Z-IIs) is the name given to a condition causing larval mortalities in hatcheries throughout the Americas since 1993. The syndrome is characterized by cessation of feeding, lethargy, inflammation of the digestive tract, atrophy of the hepatopancreas and rapid evacuation of intestinal contents (Juarez, 1997). Clinical signs are first noticed at the late zoea-I or early zoea-II stages and result in mortalities of up to 95% of the larval population before metamorphosis to zoea-III or mysis-I. Larvae that survive typically continue their larval development unaffected. The cause of Z-IIs is not fully established, but Vandenberghe et al. (1999) found several strains of V. alginolyticus associated with larvae affected with Z-IIs. No treatments are consistently effective against Z-IIs, and commercial hatcheries rely on preventive measures, such as dry-out and disinfection of facilities between larval runs and the use of probiotics in larval rearing. The use of multi-phase systems in hatcheries is largely a strategy designed to counteract the effects of bacterial diseases and Z-IIs. Larval rearing is divided in two or three phases, with each phase taking place in special tanks located in separate areas. These systems reduce disease risk by increasing control and sanitation between the phases.

 Exuvial entrapment is characterized by incomplete moulting which results in the moults sticking to the heads of the larvae, deformities, impediment to feeding and ultimately starvation and death. This syndrome is most often observed during the mysis stage but can also occur in PL. The cause of the syndrome is not clear, but poor feed quality and/or bacterial disease have been implicated. Thus, increased water exchange and revision of feeding protocols are suggested to combat this problem (FAO, 2003). Vandenberghe et al. (1999) isolated eight dominant strains of bacteria from moribund larvae with mysis moult syndrome and six were identified as V. alginolyticus.

 Filamentous bacteria can foul the gills, setae, appendages and body surface of shrimp in all life stages, including larvae and PL. These infestations lead to impaired moulting, hypoxia and death. Bacteria implicated in this type of infection include Leucothrix mucor (Fig. 10), Thiothrix sp., Flexibacter sp. and Cytophaga spp. (Aguirre and Ascencio, 2000). Recently Mouriño et al. (2008) reported mortalities up to 92% in mysis and postlarval P. vannamei in Brazilian hatcheries, caused by the Gram-negative, filamentous bacillus Flexibacter maritimus. Filamentous bacterial disease during larval rearing occurs when there is abundant organic matter in the water. Increased water exchange and other measures designed to reduce organic loading generally help in preventing or reducing problems associated with filamentous bacteria. Other control measures for filamentous bacteria in LRTs include treatment with organic copper compounds, such as Copper Control® or Cutrine-Plus® at 0.5 ppm in a 6-hour bath (Delves-Broughton and Poupard, 1978).

Fungal infections represent another significant cause of mortality of young larvae in hatcheries. The most common pathogen is the phycomycetous fungus Lagenidium callinectes, along with fungi of the genera Sirolpidium, Phythium, Leptolegnia and Haliphthoros (Lightner, 1996). Fungal infection is easily diagnosed by microscopic observation of wet mounts of dead or moribund larvae showing non-septate and highly branched fungal mycelia throughout the body and/or appendages (Fig. 11), eventually replacing most of the host’s tissue and leading to death in one to three days. Lagenidium infects all life stages of shrimp, including adults, but nauplii and zoea larvae are most sensitive. In hatcheries the disease spreads from spawners to eggs and nauplii via bi-flagellated, motile zoospores released from specialized sporangia protruding from the body of the shrimp. The first line of prevention against larval mycoses is good sanitation practice, especially thorough cleaning and disinfection of spawning tanks and related equipment.


Figure 10. Wet mount of gills from a juvenile blue shrimp (Penaeus stylirostris) with a mixed infestation by filamentous bacteria. No stain. Magnification 900X. Photo Courtesy of DV Lightner, University of Arizona, Aquaculture Pathology Laboratory.


Figure 11. Shrimp larvae (Penaeus setiferus) with a severe infestation by the fungus Lagenidium sp.
No stain. Magnification 70X. Photo Courtesy of DV Lightner, University of Arizona,
Aquaculture Pathology Laboratory.

The second line of prevention is chemoprophylaxis of the adult shrimp, eggs and nauplii. Antifungal compounds, when used properly, are effective against the motile zoospores, and thus are useful in preventing transmission, but not at treating shrimp that are already infected. In adults, Treflan TR-100® (10% Trifluralin) has been used at 5 ppm in a 1 h bath treatment. More commonly, eggs, nauplii and zoea are treated with 0.05–0.2 ppm. Once in contact with seawater, Treflan is unstable and must be “dripped-in” or re-applied every 5-8 hrs. Benzalkonium chloride at 0.01 to 0.05 ppm also has been reported to destroy the spores (FAO, 2007).

Several ciliated protozoan epibionts cause infestations in larval shrimp, most commonly Zoothamnium spp. (Fig. 12) or Vorticella spp. These protozoans are indicative of excessive organic matter in the culture water or of conditions that retard or inhibit the normal moulting of the larvae. Protozoan infestations can be prevented by good sanitation of tanks and equipment and by management techniques designed to keep good water quality and to promote healthy moulting of the larvae. Larvae infested with protozoans can be treated with Cholamine-T in 2-hour static bath at dosages from 1 ppm for mysis to 5 ppm for PL. Older PL can be treated with varying concentrations of formalin for one hour, followed by a water exchange, but this chemical is harsh and should be used with caution and tested on a small sample of PL. Treatment levels of 10 to 12 ppm can be used for strong PL-1 to PL-2 and up to 35 ppm can be used for PL-8 and older. Formalin creates an oxygen demand in the water, therefore efficient aeration is recommended during treatment.


Figure 12. Wet mount of a scraping from the cuticle of a shrimp with a highly branched colony of stalked protozoans (Zoothamnium sp). No stain. Magnification 180X. Photo Courtesy of DV Lightner,
University of Arizona, Aquaculture Pathology Laboratory.

Postlarval quality

There are several methods to evaluate PL quality. They include various observations of shrimp behaviour, stress tests, microscopic examinations and disease diagnostic methods. Some of the methods are similar to those described previously for larval shrimp.

Swimming activity is one of the easiest behaviours to evaluate. PL are transferred to a container and the water is stirred to create a circular current. Strong PL align along the sides of the container and vigorously swim against the current. Weak PL are carried by the current and end up in the centre of the container. By determining the percentage of strong versus weak PL, hatchery operators can make assessments about PL quality.

Total length and weight at a certain PL age and homogeneity of sizes are important quality indicators. Fig. 13 shows length-weight relationships for 1- to 13-day PL sampled from numerous LRTs over several production seasons. Comparison with these data can provide an idea about the condition of a specific PL batch. Total length is measured from the tip of the rostrum to the end of the telson. To weigh PL, a sample of approximately 100 individuals is wrapped in a disk of Whatman® #2 filter paper. The wrap is weighed on a balance capable of reading 0.1 mg. Once the weight is recorded, PL are removed from the disk and the weight of the wet disk is subtracted from the previous reading. The PL are then hand-counted and the mean weight is calculated.


Figure 13. Length-weight relationship for postlarval P. vannamei (PL-1 to PL-13). Weight data represent means of 100 samples (100 PLs per sample) from different larval-rearing tanks and conditions, weighed throughout several production seasons. Total length data represent means of 100 individual PLs from the same tanks. Error bars = SD. Previously unpublished data from the authors.

The development of gills influences the ability of PL to cope with low levels of dissolved oxygen and osmotic stress. Hatchery and pond operators often use gill development as a criterion to decide when to transport or stock PL in ponds. Operators tend to avoid oxygen or osmotic stress to the PL before they develop tertiary gill lamellae (as in Fig. 5c). A related indicator of fitness can be estimated by performing salinity stress tests. A common test is to subject a small sample of PL to a sudden exposure to fresh water (keeping temperature constant), maintain them in the fresh water for 30 minutes and then return them to the original salinity. PL are allowed to recover from the salinity shock for another 30 minutes, after which time the percentage of live versus dead PL is estimated. Only PL standing on their legs or swimming are counted as live (those on their sides are counted as dead, even if technically still alive). Survival greater than 80% is characteristic of strong PL. Racotta et al. (2004) found a positive correlation between results of a similar salinity stress test performed at PL-1 and survival from PL-1 to PL-15 and concluded that salinity test criteria are useful for predicting larval and postlarval survival to further stages.

Another measure of postlarval fitness and nutritional status is the muscle to gut ratio (MGR) described by Bauman and Scura (1990) for P. monodon. It is estimated by looking at the 6th abdominal segment of postlarvae under a microscope and measuring the thickness of the ventral abdominal muscle in relation to the diameter of the gut. MGR between 1:1 and 3:1 are considered fair and ratios larger than 3:1 are considered good.

Quantification of PL

PL populations can be quantified by volumetric and gravimetric methods. In general, volumetric methods are more accurate for smaller PL. The volumetric method involves concentrating PL in a known volume of water, homogenizing by vigorous mixing and taking several small aliquots for hand-counting and extrapolation to the larger volume. Hardin et al. (1985) reported that water temperature and PL size had the greatest effect on precision of volumetric estimates, with results ranging from 100% of the hand-counted verified population for 4 mm PL at 18 ºC to as low as 57% for 10.5 mm PL at º30 C. Juarez et al. (1996) evaluated volumetric methods to estimate larval and PL populations in shrimp hatcheries. PL population had a major effect on the accuracy of estimates. The coefficient of variation increased from 4% at a 100% density (one million PL in a 70-L harvest bucket) to 13% at 12.5% of the original density. No significant difference was observed between samples evaluated at 20 and 27 ºC. These results suggest that population and proper mixing are more important than temperature in obtaining accurate counts.

The gravimetric method involves weighing a sub-population of PL after concentrating on a mesh screen and draining as much water as possible. A small sub-sample of PL is then weighed and hand counted to estimate mean PL weight. The total population number can be extrapolated from this value. This method is more accurate for larger PL and juveniles and requires practice to reduce handling stress to the shrimp.

Transportation of PL

After harvesting and counting, PL can be prepared for shipping to farms in either live haulers or bags and coolers. Both methods are appropriate for long distances and transit times of up to 48 hours. Preparation for shipment begins with a gradual acclimation to the shipping temperature, which depends on the time the PL are expected to be in transit. Lower temperatures decrease their metabolic rate, thereby reducing swimming activity, feeding, excretion and oxygen consumption. Common shipping temperatures range from 25 ºC for short transit times of 1-3 hours to 18 ºC for shipments that take ten hours or more. The rate of temperature decrease should not exceed 10C every 15 min. If salinity needs to be adjusted, it is better to do it before shipping to minimize stress of packing and transportation. Acclimation for salinity can be accomplished at rates of 3 ppt per hour from 32 to 20 ppt, at 1 ppt per hour from 20 to10 ppt and at 0.5 ppt per hour for salinities lower than 10 ppt. Water used for acclimation and transportation should be filtered and sterilized by ultraviolet radiation.

Postlarvae can be transported by truck in live haulers. Hauling tanks are supplied with pure oxygen and aeration for mixing. Transport densities vary according to the age/size of the PL and the duration of the trip. Shipping by air is normally done in bags/coolers, which allows shipping to practically any destination in the world. PL are placed in plastic bags with 1/3 water and 2/3 space for pure oxygen. Bags are then placed inside insulated Styrofoam coolers and the coolers are placed inside cardboard boxes. Packing densities shown in Table 12 can be used as a guide for shipping in live haulers or in bags/coolers.

Granular activated carbon, usually in pellets, is often added to the packing water at a rate of 1g/L to absorb organic matter and other toxic substances. Buffers, such as Triss®, can be added to the water to keep the pH stable during long trips. Magallon-Barajas et al. (2006) found that pH can be a greater source of stress during transportation and acclimation than ammonia or temperature. For long transit times, PL can be fed sparingly with Artemia nauplii, but care should be taken not to overfeed, which could result in poor water quality and subsequent PL mortality.

Table 12. Packing densities (postlarvae per liter) for shipping shrimp postlarvae of different ages over various transit times.


Postlarval Hours in transit

age 4 8 12 24 48


PL-6 4,000 3,700 3,500 1,800 1,000

PL-8 3,200 2,800 2,500 1,200 800

PL-10 2,500 2,000 1,800 900 600

PL-12 2,000 1,600 1,300 700 500

PL-16 1,800 1,400 1,000 600 400

PL-18 1,200 900 800 500 300

PL-20 1,000 800 600 400 250


Research priorities

The following areas of interest to the shrimp hatchery industry represent priorities for research with foreseeable benefits to shrimp aquaculture:

Studies with purified or semi-purified diets could elucidate the effects of specific nutrients on ovarian maturation, reproduction and PL quality. This could lead to the development of complete maturation feeds and to independence from fresh feeds, which are expensive and inconsistent, require substantial labour for preparation and often present a biosecurity risk. The replacement of marine polychaetes, often cited as the number one operating expense in maturation facilities, is one of the primary goals in this field. The nutritional and environmental requirements of male broodstock also require more attention as most research to date has focused on females.

Advances in knowledge of the nutrition and endocrinology of shrimp may one day result in artificial diets that enhance reproduction without the need for eyestalk ablation. Stress from ablation and the ensuing demand for reproductive energy often result in broodstock mortality (0.5 to 1% per day is common in maturation facilities) and reduced reproductive life (3 to 4 months). Reproduction without the need for eyestalk ablation could lead to higher fecundity and viability.

The genetic components of reproduction require further research as current programs for shrimp selection focus on the immediate goals of increased growth and disease resistance, which could have undesirable effects on the reproductive ability of selected stocks. Related areas that require better understanding include the genetic basis of sex determination, induction of polyploidy and cryopreservation of shrimp gametes.

The development of larval diets to substitute live feeds, such as Artemia cysts and microalgae, could lead to more consistent larval production and to reduced bacterial problems in hatcheries. Continued research into advanced larval feeds will lead to increased use of feeds as carriers of specific nutrients, probiotics, immune stimulants and medications.


The authors wish to express their appreciation to Victoria Alday and Carlos Pantoja for critically reviewing the disease section. Don Lightner and Marisol Morales kindly provided photos of larval diseases. Margaret Barlow, Iliana Morales, Edward Scura, Richard Towner and Guillermo Baquerizo provided useful data and comments. Appreciation is also expressed to Traci Holstein for proofreading the manuscript.


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