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Laboratory culture, growth, and the life cycle of the little cuttlefish Sepiola atlantica (Cephalopoda: Sepiolidae).

ABSTRACT Pairs of Sepiola atlantica maintained in aquaria at ~17[degrees]C successfully mated in the "male parallel position" for between 21 min and 77 min. Over a period of several weeks after mating, female S. atlantica laid egg masses containing 8-161 eggs. At 14.4[degrees]C, embryonic development took 33 days and the hatching phase lasted for 23 days (mean hatching success, 32%). Hatchlings emerged from the eggs at a mean dorsal mantle length (DML) of 1.91 mm and entered a pelagic paralarval phase lasting 6 days. Ten to 20 days after hatching, the internal yolk sac became exhausted, whereupon hatchlings were fed ad libitum on wild-caught zooplankton at a density of -90 organisms/L or with enriched adult Artemia (density, 50 organisms/L). Hatchlings maintained on the Artemia diet all died within 38 days, whereas ~38% of those fed on zooplankton survived to this point, and the remaining juveniles subsequently attained adulthood when reared on a diet of Crangon crangon. These laboratory-reared juveniles matured and successfully mated, but the females did not lay any eggs. Females subsequently died 230-250 days after hatching and 10-19 days after mating, at a DML of between 21.7 mm and 23.2 mm, whereas the smaller males died 265-293 days after hatching (DML, 17.4-21.4 mm). Growth (increase in DML) of S. atlantica had 2 phases. Growth during the first 120 days was relatively slow at 0.05 mm/day (0.043 mm/day in males and 0.055 ram/day in females), increasing slightly thereafter to day 210, after which growth leveled off. These data indicate that S. atlantica has a similar life cycle to other sepioids.

KEY WORDS: little cuttlefish, reproduction, paralarvae, growth, Sepiola atlantica, sexual maturity


Cephalopods are primarily maintained in captivity for research into aspects of their biology, physiology, and biochemistry where a supply of healthy, laboratory-maintained animals is essential (Boyle 1991). Members of the family Sepiolidae are characteristically small, have a short life cycle, and generally adapt well to life in captivity, making them excellent model organisms for laboratory studies (Boletzky et al. 1971, Summers 1985). Interest in mutualistic symbiotic relationships between luminescent bacteria (Vibrio fischeri) and their sepiolid cephalopod hosts such as the Hawaiian bobtail squid, Euprymna scolopes, has recently increased (e.g., Nishiguchi et al. 2004, Visick & Ruby 2006, Adin et al. 2009, Mandel et al. 2009, Nyholm et al. 2009, Soto et al. 2009).

The term "paralarva" describes the pelagic posthatching phase in the cephalopod life cycle, which is distinctly different from that of the adults (Young & Harman 1988). Several researchers have noted that high mortalities occur when rearing paralarvae through this phase (Mangold & Boletzky 1973, Hanlon et al. 1997). This is believed to result from either an inadequate rearing environment or the paralarvae starve as a result of an inappropriate food supply (Hanlon 1987, Vidal et al. 2002). Cephalopod paralarvae are carnivorous, feeding predominantly on crustaceans, molluscs, and fish (Nixon 1987) and require prey items that are both high in polyunsaturated fatty acids and docosahexaenoic acid (Navarro & Villanueva 2000).

Like E. scolopes, the little cuttlefish (or Atlantic bobtail squid), Sepiola atlantica Orbigny, (1839-1842) have a photophore (light organ), it is small (reaching up to 21 mm in dorsal mantle length (DML)), and is widely distributed in the northeastern Atlantic from 65[degrees]N to 35[degrees]N from Iceland, the Faroe Islands, and western Norway in the north to Morocco in the south (Yau & Boyle 1996, Collins et al. 2002, Jereb & Roper 2005). Considering the wide distribution and common occurrence of the species from the shallow intertidal zone down to depths of 150 m (Norman 2000, Collins et al. 2001), comparatively little has been reported on the life cycle of this species. In this article we describe the life cycle and experimental laboratory culture of the little cuttlefish, S. atlantica, with the objective of developing a suitable methodology for the laboratory culture of the species through consecutive generations.


Between 2006 and 2008, 73 (26 male and 47 female) S. atlantica were collected using a beach seine net (20 x 2.2 m; diameter of mesh at cod end, 5 mm) from Y Foryd Bay (53[degrees]07.328'N, 004[degrees]19.337'W) and Traeth Penrhos (53[degrees]08.736'N, 004[degrees]24.469'W), Anglesey, North Wales, UK. Upon capture, S. atlantica were immediately placed in polythene bags (31 x 39 cm) containing 3 L ambient temperature seawater and a 3 to 4-cm bed of fine sand from the collection site before being placed in insulated boxes (18 x 12 x 17 cm) to reduce temperature fluctuations during transport (up to 2 h) from the collection site to the laboratory. S. atlantica were acclimated (~1 h) by floating the open plastic bags in aquaria, with partial water changes (~20%) every 20 rain under subdued lighting (~55 lux) to reduce stress.

Each of 15 pairs of S. atlantica (1 male and 1 female) were maintained in rectangular black PVC tanks (39 x 28 x 22 cm = ~21 L) with a flow of ambient temperature seawater (~17[degrees]C) through each tank (flow rate, ~0.32 L/min). Each tank was covered with a lid to minimize disturbance, contained a 3-4-cm bed of fine sand, and was aerated and the water circulated with a submersible pump (Micro Jet MC320; Aquarium Systems (Aquarium Systems Inc, Mentor, OH)). Wild-caught S. atlantica were kept in constant darkness, whereas laboratory-cultured animals were maintained on a 12-h light/12-h dark cycle. Plastic pipes, rocks, empty bivalve shells (e.g., Mytilus edulis, Modiolus modiolus, and Arctica islandica) and coral fragments (Lophelia pertusa) were used as spawning substrata. S. atlantica were fed ad libitum on the mysid Neomysis integer and the shrimp Crangon crangon. Salinity, pH, ammonia, nitrite, and nitrate were recorded once a week, with seawater temperature recorded hourly using a Gemini data logger (Gemini Data Loggers, Chichester, West Sussex, UK). A refractometer (for salinity), a handheld pH probe, and standard laboratory test kits were used to monitor water quality (e.g., ammonia, nitrite, and nitrate).

To understand mating behavior in S. atlantica, 13 mating pairs of animals were filmed in darkness using a digital video camera (SONY Handycam DCR-HC96E, SONY, Tokyo, Japan) with an infrared light source. Observations were made in tanks similar to those in which they were maintained. S. atlantica pairs were placed in a tank, separated by a mesh divider for 1 h prior to removal of the divider, and filmed until mating was completed (<90 min). Spawned egg masses were photographed in situ using a Nikon D40 camera (Nikon, Tokyo, Japan). Eggs were counted and their diameter measured (to 0.01 mm) from digital photographs using analySIS image analysis software (Olympus Soft Imaging Solutions GmbH, Munich, Germany). Three eggs were removed at random from the egg masses every third day for measurement, placed in a Petri dish containing seawater, and photographed using a photo microscope (65.560 RZT-SF, Novex-Holland, Arnhem, Holland). The outer chorion was carefully removed from each egg using forceps and rephotographed to document the stages of embryonic development.

The aquaria containing the egg masses were inspected twice daily (~1000 h and ~1700 h) by flashlight, and the number of hatchlings recorded. Hatchlings were removed with a turkey baster pipette (25-mL volume) to avoid net damage and aerial exposure, and were placed randomly at a density of ~1 paralarva/L in 1 of 6 circular black PVC bowls (diameter, 31 cm; depth, 13 cm: volume, ~6L), containing sediment to a depth of 3 mm with a flow-through of ambient temperature seawater (flow rate, 10 ml/min). The seawater in each bowl was vigorously aerated and the bowl with a central stand pipe (1-cm diameter) covered with 150-[micro]m mesh (in 2007) to retain both paralarvae and food items while allowing the overflow of water out of the tank. Each bowl was covered with a thin black polythene sheet to reduce overhead illumination and disturbance from passing workers, and to enhance feeding compared with E. scolopes (Hanlon et al. 1997). During our rearing experiment, the paralarvae reduced in number so that, by day 80, the surviving S. atlantica were assigned to individual bowls and, by day 208, they were transferred to their own broodstock tanks.

To investigate the growth and survival of paralarvae, 3 replicate groups of animals were fed ad libitum on 1 of 2 diets: a natural zooplankton mix (density, ~90/L) and a diet of enriched adult Artemia sp. (density, ~50/L). Zooplankton was collected using a standard zooplankton tow net (420-[micro]m diameter mesh; net opening diameter, 52 cm; net length, 150 cm) deployed every third day from Menai Bridge Pier in the Menai Strait. The zooplankton comprised the copepods Temora longicornis, Paracalanus parvus, Pseudocalanus elongates, Centropages typicus, Calanus finmarchicus, and Oithona nana; and the malacostracan crustaceans Crangon crangon, Corophium volutator, N. integer, Inachus dorsettensis (megalopa), and Parathemista sp. They were maintained in a 20-L aerated cylinder (35-cm diameter x 15-cm depth) with daily 80% seawater changes and the addition of 500 mL each of the microalgae Pavlova lutheri and Rhinomonas reticulata cultured at densities of ~6,000 cells/[micro]L and 1,000 cells/[micro]L, respectively. Artemia was cultured using the standard methodology of Lavens and Sorgeloos (1996). In 2006, 24-h-old, nonenriched Artemia nauplii (Gold label, Vinh Chfiu, Cantho, Vietnam) were fed in abundance to the S. atlantica paralarvae. The following year (2007), 24 h prior to feeding the S. atlantica, l-wk-old Artemia sp. (size, 2-3 mm total length) were instead enriched with Algamac 2000 (Bio-Marine Brand, Hawthorne, CA) high (>24%) in docosahexaenoic acid and then fed to the S. atlantica. C. crangon (total length, 4-5 mm) and C. volutator (total length, 3-4 mm) were offered and readily consumed by S. atlantica from day 60 onward, when it appeared that the zooplankton species were no longer being eaten.

Mortality of S. atlantica paralarvae was monitored daily. For the first 40 days, 1 randomly selected paralarva from each of the 3 replicate bowls from each diet was removed and euthanized in ice water every fifth day for the measurement of wet body weight and DML. A digital image of the paralarva/juvenile was taken, the DML was measured using the analySIS image analysis software (to 0.01 mm), and the blotted (tissue paper) flesh of each animal was weighed (wet body weight) to the nearest 0.001 g on a top-loading balance (Ohaus Analytical Plus, Pine Brook, NJ) (Summers 1985). As the experiment progressed, mortality increased, thus the time between measurements was extended to every tenth day. After mating had occurred, female S. atlantica were no longer selected for measurement because disturbance following mating affects egg production and delays spawning (Hanlon et al. 1997).


S. atlantica successfully mated and spawned at 17.4 [+ or -] 0.4[degrees]C (SD; range, 16.3-17.9[degrees]C), and paralarvae and juveniles were successfully produced at seawater temperatures ranging between 15.3[degrees]C and 17.7[degrees]C (mean, 17.7 [+ or -] 0.9 [degrees]C). Throughout the experiments, ammonia, nitrite, and nitrate remained within the recommended levels proposed by Hanlon et al. (1997) (i.e., ammonia and nitrite, 0.10 mg/L; nitrate, 20 mg/L), whereas the pH ranged between 8.3 and 8.9 (mean, 8.6 [+ or -] 0.2) and salinity ranged from 31 35[per thousand] (mean, 32[per thousand]).

Male and female S. atlantica were observed in a mating embrace known as the "male parallel position" (Hanlon & Messenger 1996). Analysis of film footage of the 13 mating pairs showed that mating was initiated within 26 [+ or -] 32.1 min (range, 1-83 min) after removal of the tank divider, with the males and females mating on average for 64 [+ or -] 9.8 min (range, 49-77 min). Mating only took place in darkness and no courtship behavior was observed. A male "pounced" on a female from behind, and mating generally occurred on the aquarium floor. At the point of separation after mating, the S. atlantica inked in 8 of the 13 observed matings.

Between 2006 and 2007, 5 females laid eggs at night on the aquaria walls at the air-water interface. Eggs were laid in the corner of the tanks, above aerators where the water was vigorously agitated and well oxygenated. Five batches of eggs were laid between 17 and 83 days postcapture, and contained varying numbers of eggs (8, 12, 123, 132, and 161), and the females survived for 1-8 days after egg laying. Lemon-shaped, initially cream-colored eggs were laid in a single layer; the eggs subsequently turned golden brown during development. The first batch of eggs hatched earlier than expected and all the paralarvae were lost from the bowls, 2 batches of eggs failed to develop, and 1 batch hatched prematurely after physical disturbance of the egg mass and all the hatched paralarvae died by day 14. The fifth S. atlantica laid the most eggs (n = 161) and these developed satisfactorily. Postmortem examination of the 5 females that died after spawning showed that the ovaries were not completely spent. During development, mean egg diameter increased from an initial 2.48 [+ or -] 0.05 mm to a maximum of 3.37 [+ or -] 0.09 mm by day 26, decreasing in size to 2.54 [+ or -] 0.13 mm at first hatching, with a mean diameter of 3.13 [+ or -] 0.18 mm at last hatching. Hatching lasted for 23 days at a mean temperature of 14.4 [+ or -] 0.3[degrees]C, with a hatching success of 32%, whereas a peak in hatching rate (i.e., 12 hatchlings) occurred on day 14 of hatching. On hatching, the paralarvae had a mean DML of 1.91 [+ or -] 0.26 mm (n = 52).

A description of the sequence of embryonic development is based on the eggs laid by the fifth S. atlantica and is shown in Figure 1. Figure 1A (day 9) shows that the developing embryo is not visible when viewed under a dissecting microscope, but by day 14 the embryo is clearly visible (Fig. lB), with orange pigmented eyes and early development of the arms. By day 17 (Fig. 1C), the embryos rotated within their egg capsules and the eye lenses were distinguishable. Internal organs were also visible and, externally, fins started to develop. Figure 1D (day 20) shows the development of suckers on the arms, with the appearance of red eyes; by day 23, orange chromatophores were observed on the dorsal surface of the head. By day 26, chromatophores covered the head and both dorsal and ventral surfaces of the mantle, the eyes were blood red, and the yolk envelope was observed to pulsate. Figure 1E (day 29) shows that the ink sac is clearly visible within the mantle cavity. Brown chromatophores were also visible on both the head and mantle of the embryo. By day 32 (Fig. 1F), orange, brown, and black chromatophores were visible on the head and mantle. The now well-developed eyes were black, and some movement was observed in the arms. By day 34, the fins were seen to move freely and a greater degree of control was seen in the chromatophores as they rhythmically expanded and contracted. Figure lG shows the anterior and posterior lobes of the inner yolk sac along with the lateral lobe of the inner yolk sac visible through the mantle of an S. atlantica hatchling (Fig. 1H).

Upon hatching, S. atlantica paralarvae constantly swam in the water column for 6 days but did not feed. From day 6 onward, however, they were observed hunting and capturing prey, and then began to adopt a more epihenthic lifestyle, settling on the aquarium floor. From these observations, it is assumed that the pelagic paralarval phase (at ~2.4-mm DML) lasts for 6 days, during which time the paralarvae are unable to feed independently. By settling partially buried in sand and adopting a benthic lifestyle similar to that of the adults, S. atlantica are now classified as juveniles (Young & Harman 1988). The internal yolk sac became exhausted 10-20 days after hatching and juveniles showed less interest in the zooplankton diet (day 50); by day 54, they fed and readily captured ~4-5-mm (total length) C. crangon. To achieve this, juveniles would make several approaches toward the shrimp, swimming head down in mid water or at the surface, with their arms forming a cone shape before extending their tentacles to seize the prey in a manner similar to that described for Euprymna berryi (Choe 1966) and S. robusta (Boletzky 1983).

The survival of S. atlantica paralarvae and juveniles reared on a diet of either mixed zooplankton (2007) or enriched Artemia sp. nauplii or adults (2006 and 2007) can be seen in Figure 2. During the first 10 days in 2006, survival was greater than in 2007, with the Artemia nauplii numbers declining rapidly until all animals were dead by day 14. In 2007, survival on Artemia was much better than in 2006; nevertheless, all the paralarvae were dead by day 38. Survival on the zooplankton mixture in 2007 was generally better so that by day 38, ~38% of the S. atlantica were still alive. By day 40, 5 were alive (~25%) and they survived until the following year before finally succumbing after mating (i.e., by day 293). Overall, 9.6% of the hatchlings survived to adulthood. Although these laboratory-reared adults mated successfully in 2008, the 3 females did not spawn and died 230-250 days after hatching at a DML of between 21.7 mm and 23.2 mm, and 10-19 days after mating. The two males outlived the females, dying 265 and 293 days after hatching (DML 17.4 mm and 21.4 mm) respectively.



Growth in a mixed-sex population of laboratory-reared S. atlantica had 2 phases (Fig. 3). Growth in DML during the first 120 days was relatively slow, increasing slightly thereafter to day 210, after which growth leveled off. During the first 120 days, S. atlantica grew at a rate of 0.05 mm/day (0.043 mm/day in males and 0.055 mm/day in females). The mean population daily growth rate over 210 days was 0.08 mm/day (males, 0.07 mm/ day; females, 0.09 mm/day). The growth of S. atlantica slowed markedly after mating and, although the females died off after this point, the growth rate of the remaining live males continued at a slow rate of 0.04 mm/day from day 210 until the completion of the experiment. Large variability in the size of individuals was evident between day 200 and day 260. The increase in DML was independent of seawater temperature, and S. atlantica grew at a similar rate at 10[degrees]C and 15[degrees]C; growth rate was not significantly different (F = 1.34, P = 0.274; data normally distributed and homogeneity of variance) between the 5 S. atlantica (2 males and 3 females) between day 90 and day 180, when they became sexually mature. It was not possible to test for statistical differences in growth rate among the 5 animals after sexual maturation until death (nonnormally distributed data and unequal variance). However, the data suggest that the females grew faster than males after they became sexually mature.



Hanlon et al. (1997) have proposed that optimal conditions for broodstock management should be maintained in order for "normal" reproductive behavior and fecundity to occur. Only 5 female S. atlantica laid eggs despite the provision of excellent water quality, abundant and enriched food sources, suitable spawning substrata, and the manipulation of lighting regimes. It is therefore likely that spawning is triggered by one or more environmental factors that were not always present in the relevant quantities in our experiments. The data indicate that S. atlantica spawned during periods of increasing seawater temperatures, as Ambrose (1988) has similarly noted for other cephalopod species. Batches of 8-161 lemon-shaped eggs were laid in a single layer and in similar numbers to those laid in egg masses of wild and laboratory-maintained sepiolids. For example, E. scolopes lays between 12 eggs and 310 eggs (Singley 1983, Claes & Dunlap 2000), whereas Steer et al. (2004) noted that between 3 eggs and 107 eggs were laid by Euprymna tasmanica, 25-50 eggs were laid by Rossia pacifica (Summers 1985, Summers & Colvin 1989, Anderson 1991), and Sepietta oweniana laid between 2 eggs and 176 eggs (Bergstrom & Summers 1983, Bello & Deickert 2003, Deickert & Bello 2005). Female S. atlantica died 1-8 days after spawning, similar to laboratory-held R. pacifica, S. obscura, S. oweniana, E. scolopes, S. robusta, and E. hyllebergi (Bergstrom & Summers 1983, Boletzky 1983, Singley 1983, Summers 1985, Anderson 1991, Nabhitabhata et al. 2005).

A hatching success of 32% achieved in this study is considerably less than the 70-90% recorded for E. scolopes by Claes and Dunlap (2000) and the more than 80% recorded by Choe (1966) in E. berryi and 82-100% in E. hyllebergi (Nabhitabhata et al. 2005). Seawater quality and lighting levels remained relatively constant in our study, so the low hatching successes could be a result of physical disturbance to the eggs during their removal for measurement and observation. Differences in embryo mortality and reproductive output have been attributed to maternal ration (Steer et al. 2004). For example, reduction in lipid intake by the female subsequently increased embryo mortality because they had insufficient lipid reserves to complete embryogenesis. In our study, the S. atlantica broodstock were fed to excess on shrimp that were themselves fed daily on mussel flesh, so it seems unlikely that later embryo mortality would have been affected by female ration. S. atlantica hatchlings had a mean DML of 1.9 mm, similar to the 2.2 mm in S. robusta (Boletzky 1983), 2.5 mm for S. oweniana (Bergstrom & Summers 1983), 1.5-1.9 mm in E. scolopes (Hanlon et al. 1997, Singley 1983), and 2.2 mm in E. hyllebergi (Nabhitabhata et al. 2005).

Survival of juvenile S. atlantica fed on a diet of enriched adult Artemia was poor, but improved when they were reared on a mixed zooplankton diet. E. scolopes fed entirely on Artemia suffered high mortality rates and exhibited poor growth (Singley 1983). Sepioids have been fed various diets in abundance. For example Hanlon et al. (1997) recorded 100% mortality in E. seolopes by day 8 fed on a zooplankton mix as well as in a control group that were not fed. Survival rates of 10% were achieved by feeding E. scolopes on postlarval mysids and ~16% survival on a diet of various postlarval shrimp, whereas ~13% of those reared on a combination of postlarval mysids, fish, zooplankton, and adult mysids survived to settlement. In the current study, S. atlantica mortality leveled off after day 40, suggesting that the first 40 days in the life cycle are critical for survival.

Growth in cephalopods is influenced by biotic and abiotic factors such as seawater temperature, diet, age, gender, and maturity (Forsythe 1984, Forsythe & Van Heukelem 1987, Forsythe & Hanlon 1988, Forsythe 1993). Variations in the growth of individuals can influence a population in a number of ways, such as its population size and age structure, reproductive dynamics, and hatchling survival, which in turn affect the abundance of the species (Leporati et al. 2007). Growth studies of captive cephalopods have shown a 2-phase growth pattern, generally comprising a short exponential stage of high growth, with a subsequent decline in growth rate at a point when energy is diverted from growth to reproduction (Mangold 1983). S. atlantica conforms similarly to this pattern of growth. Boletzky (1983) reported a mean growth rate of 0.1 mm/day (DML) in S. robusta reared at 20[degrees]C, followed by a general decrease in growth accompanied by increased variability in the growth of individuals. Boletzky et al. (1971) reported similar growth rates in Sepiola and Sepietta of 2.5 mm/mo (DML) during the first 5 mo in Sepiola and 3 mo in Sepietta, whereas the growth rate later increased to 5 mm/mo in Sepietta, and concluded that growth rate was independent of seawater temperature because S. obscura grew fastest during periods with the lowest temperatures. Summers (1985) and Summers and Colvin (1989) observed that growth rates did not remain constant throughout the life cycle of R. pacifica, with growth rates of 0.034 mm/day during the first 6 mo, increasing to 0.065 mm/day and 0.25 mm/ day for males and, females respectively, thereafter. Bergstrom and Summers (1983) recorded a mean growth rate of 4.8 mm/ mo in S. oweniana, with females growing significantly faster than males (5.3 mm/mo in females, 4.2 mm/mo in males) during the first 6 mo of life. Sexual maturation in members of the Sepiolinae is generally independent of environmental cues such as light and seawater temperature (Boletzky 1983). MangoldWirz (1963) and Mangold et al. (1975) found that no single factor governed the onset of sexual maturation in S. oweniana, whereas sexual maturation of captive S. affinis was unaffected by constant illumination but delayed by low food ration (Boletzky 1975). In our laboratory study, S. atlantica became sexually mature ~7 mo after hatching (~15 mm DML in males, ~17 mm in females). Yau (1994) demonstrated that male S. atlantica collected from a wild population in Scotland attained sexual maturity earlier, and at a slightly smaller size (12.8-mm DML) than females (15.0-mm DML). Although none of the laboratory-reared animals spawned, it can be estimated that the life cycle of S. atlantica is complete within 8 mo, although the longest lived individual survived for almost 10 mo. These observations are in agreement with a longevity of less than 12 mo proposed by Yau (1994), and similar to the 6-9 mo in the Sepiolinae proposed by Boletzky (1974). Female S. robusta become sexually mature by 17 mm DML, with a complete life cycle of 5.5 mo, whereas males were smaller in length on maturation (Boletzky 1983). Singley (1983) found that E. scolopes reached sexual maturity when 4-6 mo old, with a life span of 7-10 too. Male and female S. robusta usually die within a few months of reaching sexual maturity, with females dying days after spawning, followed closely by males, who are generally 6-8 mo old at death (Boletzky 1983). Boletzky et al. (1971) found that male S. neglecta, S. obscura, and Sepiola sp. reached adult size between 110 days and 190 days, with S. rondeleti spawning ~140 days after hatching and ~170 days in S. robusta. In conclusion, S. atlantica appears to have a similar life cycle to a number of other Sepiolids that have previously been cultured in the laboratory through consecutive generations. Although it was not achieved here, it seems entirely possible that a second captive generation of S. atlantica could be reared in the future. The environmental cues that trigger spawning in this species remain unclear, although a greater understanding of the nutritional requirements of both paralarvae and juveniles would increase survival of laboratoryreared animals through the critical initial phase of the life cycle.


N. J. thanks the European Social Fund for supporting his PhD studies. We thank Berwyn Roberts and Andy Marriott for assistance in the field, and Gwyn Hughes for technical support in the laboratory.


Adin, D. M., J. T. Engle, W. E. Goldman, M. J. McFall-Ngai & E. V. Stabb. 2009. Mutations in ampG and lytic transglycosylase genes affect the net release of peptidoglycan monomers from Vibrio fischeri. J. Bacteriol. 191(7):201-2022.

Ambrose, R. F. 1988. Population dynamics of Octopus bimaculatus: influence of life history patterns, synchronous reproduction and recruitment. Malacologia 29:23-39.

Anderson, R. C. 1991. Aquarium husbandry of the sepiolid squid Rossia pacifica. AAZPA Ann. Conf. Proc. 206-211.

Bello, G. & A. Deickert. 2003. Multiple spawning and spawning batch size in Sepietta oweniana (Cephalopoda: Sepiolidae). Cah. Biol. Mar. 44:307-314.

Bergstrom, B. & W. C. Summers. 1983. Sepietta oweniana. In: P. R. Boyle, editor. Cephalopod life cycles. Vol. I. Species accounts. London: Academic Press. p. 73-91.

Boletzky, S. V. 1974. Elevage des cephalopods en aquarium. Vie Milieu 24:309-340.

Boletzky, S. V. 1975. The reproductive cycle of Sepiolidae (Mollusca, Cephalopoda). Pubbl. Staz. Zool. Napoli 39, Suppl. 84-95.

Boletzky, S. V. 1983. Sepiola robusta. In: P. R. Boyle, editor. Cephalopod life cycles Vol. I. Species accounts. London: Academic Press. p. 53-67.

Boletzky, S. V., M. V. Boletzky, D. Frosch & V. Gatzi. 1971. Laboratory rearing of Sepiolinae (Mollusca: Cephalopoda). Mar. Biol. 8:82-87.

Boyle, P. R. 1991. The UFAW handbook on the care and management of cephalopods in the laboratory. Aberdeen: Universities Federation for Animal Welfare. p. 63.

Choe, S. 1966. On the eggs, rearing, habits of the fry, and growth of some cephalopoda. Bull. Mar. Sci. 16(2):330-348.

Claes, M. F. & P. V. Dunlap. 2000. Aposymbiotic culture of the sepiolid squid Euprymna scolopes: role of the symbiotic bacterium Vibrio fischeri in host animal growth, development, and light organ morphogenesis. J. Exp. Zool. 286:280-296.

Collins, M. A., C. Yau, L. Allcock & M. H. Thurston. 2001. Distribution of deep water benthic and bentho-pelagic cephalopods from the north-east Atlantic. J. Mar. Biol. Assoc. UK 81:105-117.

Collins, M. A., C. Yau, P. R. Boyle, D. Friese & U. Piatkowski. 2002. Distribution of cephalopods from plankton surveys around the British Isles. Bull. Mar. Sci. 71(1):239-254.

Deickert, A. & G. Bello. 2005. Egg masses of Sepietta oweniana (Cephalopoda: Sepiolidae) collected in the Catalan Sea. Sci. Mar. (Barc.) 69(2):205-209.

Forsythe, J. W. 1984. Octopusjoubini (Mollusca: Cephalopoda): a detailed study of growth through the full life cycle in a closed seawater system. J. Zool. 202:393-417.

Forsythe, J. W. 1993. A working hypothesis of how seasonal temperature change may impact the field growth of young cephalopods. In: T. Okntani, R. K. O'Dor & T. Kubodera, editors. Recent advances in cephalopod fisheries biology. Tokyo: Tokai University Press. p. 133-143.

Forsythe, J. W. & R. T. Hanlon. 1988. Effect of temperature on laboratory growth, reproduction and life span of Octopus bimaculoides. Mar. Biol. 98:369-379.

Forsythe, J. W. & W. F. Van Heukelem. 1987. Growth. In: P. R. Boyle, editor. Cephalopod life cycles, vol. II. London: Academic Press. p. 135-155.

Hanlon, R. T. 1987. Mariculture. In: P. R. Boyle, editor. Cephalopod life cycles, vol. II. London: Academic Press. p. 291-305.

Hanlon, R. T., M. F. Claes, S. E. Ashcraft & P. V. Dunlap. 1997. Laboratory culture of the sepiolid squid Euprymna scolopes: a model system for bacteria-animal symbiosis. Biol. Bull., Mar. Biol. Lab. Woods Hole 192:364-374.

Hanlon, R. T. & J. B. Messenger. 1996. Cephalopod behaviour. Cambridge: Cambridge University Press. p. 232.

Jereb, P. & C. F. E. Roper, editors. 2005. Cephalopods of the world: an annotated and illustrated catalogue of cephalopod species known to date. Chambered Nautiluses and Sepioids (Nautilidae, Sepiidae, Sepiolidae, Sepiadariidae, Idiosepiidae and Spirulidae). FAO species catalogue for fishery purposes. No. 4, vol. 1. Rome: FAO. p. 262.

Lavens, P. & P. Sorgeloos, editors. 1996. Manual on the production and use of live food for aquaculture. FAO fisheries technical paper no. 361. Rome: FAO. p. 295.

Leporati, S. C., G. T. Pecl & J. M. Semmens. 2007. Cephalopod hatchling growth: the effects of initial size and seasonal temperatures. Mar. Biol. 151:1375-1383.

Mandel, M. J., M. S. Wollenberg, E. V. Stabb, K. L. Visick & E. G. Ruby. 2009. A single regulatory gene is sufficient to alter bacterial host range. Nature 458(7235):215 218.

Mangold, K. 1983. Food, feeding and growth in cephalopods. Mem. Nat. Mus. Victoria 44:81-93.

Marigold, K. & S. v. Boletzky. 1973. New data on the reproductive biology and growth of Octopus vulgaris. Mar. Biol. 19:7-12.

Mangold, K., D. Froesch, R. Boucher-Rodini & V. L. Rowe. 1975. Factors affecting sexual maturation in cephalopods. Pubbl. Stn. Zool. Napoli 39:259-266.

Mangold-Wirz, K. 1963. Biologic des cephalopods benthiques et nectonique de la Mer Catalane. Vie Milieu 13:1-285.

Nabhitabhata, J., P. Nilaphat, P. Promboon & C. Jaroongpattananon. 2005. Life cycle of cultured bobtail squid, Euprymna hyllebergi Nateewathana, 1997. Phuket Mar. Biol. Cent. Res. Bull. 66:351-365.

Navarro, J. C. & R. Villanueva. 2000. Lipid and fatty acid composition of early stages of cephalopods: an approach to their lipid requirements. Aquaculture 183:161-177.

Nishiguchi, M. K., J. E. Lopez & S. V. Boletzky. 2004. Enlightenment of old ideas from new investigations: more questions regarding the evolution of bacteriogenic light organs in squids. Evol. Dev. 6(1):41-49.

Nixon, M. 1987. Cephalopod diets. In: P. R. Boyle, editor. Cephalopod life cycles, vol II. London: Academic Press. pp. 201-209.

Norman, M. 2000. Cephalopods: A world guide. Hachenheim, Germany: Conch Books. p. 318.

Nyholm, S. V., J. J. Stewart, E. G. Ruby & M. J. McFall-Ngai. 2009. Recognition between symbiotic Vibriofischeri and the haemocytes of Euprymna scolopes. Environ. Microbiol. 11(2):483-493.

Singley, C. T. 1983. Euprymna scolopes. In: P. R. Boyle, editor. Cephalopod life cycles. Vol. I. Species accounts. London: Academic Press. pp. 69-74.

Soto, W., J. Gutierrez, M. D. Remmenga & M. K. Nishiguchi. 2009. Salinity and temperature effects on physiological responses of Vibrio fischeri from diverse ecological niches. Microb. Ecol. 57:140-150.

Steer, M. A., N. A. Moltschaniwskyj, D. S. Nichols & M. Miller. 2004. The role of temperature and maternal ration in embryo survival: using the dumpling squid Euprymna tasmanica as a model. J. Exp. Mar. Biol. Ecol. 307:73-89.

Summers, W. C. 1985. Ecological implications of life stage timing determined from the cultivation of Rossia pacifica (Mollusca: Cephalopoda). Vie Milieu 35(3/4):249254.

Summers, W. C. & L. J. Colvin. 1989. On the cultivation of Rossia pacifica (Berry, 1911). J. Cephalopod Biol. 1(1):21-31.

Vidal, E. A. G., N. Koueta, J. Riba & E. Boucaud-Camou. 2002. Influence of temperature and food availability on survival, growth and yolk utilization in hatchling squid. Bull. Mar. Sci. 7(2):915-932.

Visick, K. L. & E. G. Ruby. 2006. Vibriofischeri and its host: it takes two to tango. Curr. Opin. Microbiol. 9:632-638.

Yau, C. 1994. The ecology and ontogeny of cephalopod juveniles in Scottish waters. PhD diss. The University of Aberdeen.

Yau, C. & P. R. Boyle. 1996. Ecology of Sepiola atlantica (Mollusca: Cephalopoda) in the shallow sublittoral zone. J. Mar. Biol. Assoc. UK 76:733-748.

Young, R. E. & R. F. Harman. 1988. "Larva," "paralarva" and "subadult" in cephalopod terminology. Malacologia 29(1):201-202.


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Author:Jones, Nicholas J.E.; Richardson, Christopher A.
Publication:Journal of Shellfish Research
Article Type:Report
Geographic Code:4EUUK
Date:Apr 1, 2010
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