Browse Subject Areas

Click through the PLOS taxonomy to find articles in your field.

For more information about PLOS Subject Areas, click here.

  • Loading metrics

Early Mode of Life and Hatchling Size in Cephalopod Molluscs: Influence on the Species Distributional Ranges

Early Mode of Life and Hatchling Size in Cephalopod Molluscs: Influence on the Species Distributional Ranges

  • Roger Villanueva, 
  • Erica A. G. Vidal, 
  • Fernando Á. Fernández-Álvarez, 
  • Jaruwat Nabhitabhata


Cephalopods (nautiluses, cuttlefishes, squids and octopuses) exhibit direct development and display two major developmental modes: planktonic and benthic. Planktonic hatchlings are small and go through some degree of morphological changes during the planktonic phase, which can last from days to months, with ocean currents enhancing their dispersal capacity. Benthic hatchlings are usually large, miniature-like adults and have comparatively reduced dispersal potential. We examined the relationship between early developmental mode, hatchling size and species latitudinal distribution range of 110 species hatched in the laboratory, which represent 13% of the total number of live cephalopod species described to date. Results showed that species with planktonic hatchlings reach broader distributional ranges in comparison with species with benthic hatchlings. In addition, squids and octopods follow an inverse relationship between hatchling size and species latitudinal distribution. In both groups, species with smaller hatchlings have broader latitudinal distribution ranges. Thus, squid and octopod species with larger hatchlings have latitudinal distributions of comparatively minor extension. This pattern also emerges when all species are grouped by genus (n = 41), but was not detected for cuttlefishes, a group composed mainly of species with large and benthic hatchlings. However, when hatchling size was compared to adult size, it was observed that the smaller the hatchlings, the broader the latitudinal distributional range of the species for cuttlefishes, squids and octopuses. This was also valid for all cephalopod species with benthic hatchlings pooled together. Hatchling size and associated developmental mode and dispersal potential seem to be main influential factors in determining the distributional range of cephalopods.


The early life of marine organisms is a decisive phase as it determines survival and recruitment success dictating many aspects of the species population dynamics. The varying synchrony between hatching of larvae and plankton production means that hatching in the right place and time will be decisive for first feeding success and thus, survival and growth. Dispersal of the hatchlings away from their parent source will have an imperative role in this process. In fact, as noted by R. Nathan [1]: “dispersal is a fundamental biological process with important implications at multiple scales of organization: for the survival, growth and reproduction of individuals; for the composition, structure and dynamics of populations and communities; and for the persistence, evolution and geographical distribution of species”. Multiple biotic and abiotic factors regulate dispersal and geographical distribution of marine species. Among them, the hatchling mode of life, benthic or planktonic, can be of notable influence on dispersal [2, 3]. However, few studies have addressed this topic, particularly for marine carnivores. Cephalopods play an important role as predators and prey in marine ecosystems [4] and their relative abundances have increased worldwide in recent years [5]. For adult cephalopods, both latitude and depth range have a significant effect on maximum body size, and temperature seems to be the most important factor in determining the distribution of adult body size along the continental shelves of the Atlantic Ocean [6]. These molluscs have both planktonic and benthic hatchling modes of life depending on the species. For this reason they may be used as models to improve our understanding of the factors that influence marine species distributional ranges. Most of the cephalopod species that have small hatchlings are planktonic during their early life, while those that produce larger hatchlings are usually benthic, with some exceptions.

Depending on the environment in which they live and their early mode of life, cephalopods can be divided into the following groups: holobenthic, when the full life cycle is associated with the benthos (e.g. most cuttlefishes); holopelagic, when the full life cycle is associated with the pelagic environment (e.g., all squids); and merobenthic, when hatchlings are planktonic followed by a benthic life from juvenile to adult (e.g., some octopuses). All three types of life cycle are observed in the octopods, a group with holopelagic, holobenthic and merobenthic representative species [7]. In comparison with other molluscs, cephalopods have direct development, thus hatchlings are essentially miniature of the adults and there are no marked morphological changes during ontogeny [4]. Nevertheless, the term paralarva is used on an ecological context to refer to planktonic hatchlings that have a different mode of life than the adults [8] and, the term juvenile is commonly used to refer individuals after the end of the planktonic phase, as well as for benthic hatchlings

Planktonic hatchlings are transported by currents and an inverse relationship should be expected between hatchling size and dispersal potential. In fact, transport by currents has been shown to be a powerful dispersal mechanism for planktonic squid [914] and octopus paralarvae [15, 16]. In addition to this passive transport, the swimming capacities recorded in the laboratory for planktonic cephalopod hatchlings are within the range or higher than those found for the larval fishes [1722], which could enhance their dispersal potential. In contrast, mark and recapture experiments with hatchlings of large, benthic cuttlefish, showed limited shallow water dispersal, as the individuals remain in the same or adjacent bays to those in which they hatched [23]. Molecular studies revealed some low-scale geographic population structure in cuttlefish species [24], supporting the suspected low dispersal abilities of this group.

As a working hypothesis, we aim to evaluate if hatchling size and early mode of life, planktonic or benthic, are related to the extent of the species distribution. To understand the possible influence of the early mode of life on the geographic distribution of cephalopods, we studied three parameters for each species: the mean hatchling size, their mode of early life (planktonic or benthic) and the latitudinal distribution range of the species. We also selected species with known hatchling size to obtain additional information on the duration of the planktonic phase and on the relative hatchling size in relation to adult size. These parameters are subject to relative variability, both at spatial and temporal scales. However, they can offer interesting insights to understanding early life history strategy, species dispersal potential and biogeography in this group of marine invertebrate predators.

Materials and Methods

Hatchling and adult size

The hatchling size data were obtained through an extensive literature review, selecting only laboratory studies that examined: egg masses spawned in the laboratory from females properly identified at the species level, in vitro fertilization experiments and laboratory-hatched individuals from properly identified egg masses collected from the wild (see Table 1). These criteria were necessary to avoid possible taxonomic misidentifications, size variations and unknown age determination from wild-collected hatchlings. Hatchling size was measured as mantle length (ML; mm) of fresh individuals.

Table 1. Summary of data and literature references of hatchling mantle length (ML) and latitudinal distribution range for the 110 cephalopod species analyzed in the present study.

For each species and publication source, the hatchling ML (mm) was obtained as the mean value provided by the study. When the hatchling ML of a species was obtained from more than one study, the mean of each study was pooled to obtain the mean value representative of that particular species. The hatchling ML of a genus was obtained as the mean ML of all the species from that genus, following the FAO taxonomic criteria [2527]. Hatchling ML from fresh individuals was selected when possible. When only preserved material existed for a species (n = 10 species, see Table 1), a shrinkage correction factor of 25.8% was applied to estimate the fresh ML. This correction factor was obtained as the average shrinkage percentage from five species whose fresh and preserved hatchling ML was known from the same individuals. These species were: Sepietta oweniana [28], Callistoctopus macropus [29], Eledone moschata [30], Graneledone boreopacifica [31] and Scaeurgus unicirrhus [32], with respective shrinkages of 37.5, 25.0, 10.0, 34.6 and 22.0%. Thus, when only measurements from preserved material was available, 25.8% of the preserved ML was added to estimate the fresh ML.

Some species with well-known hatchling size were excluded from the analysis for different reasons. This applied to species currently considered as a species complex, consequently, having uncertain taxonomic and latitudinal distribution such as Sepia pharaonis [33, 34], Sepioteuthis lessoniana [3537], Loligunculla brevis [38] and Sthenoteuthis oualaniensis [39], as well as species with uncertain taxonomic status like Pinnoctopus cordiformis [40]. Ommastrephes bartramii is thought to represent at least three different species [41]. However, the hatchlings previously described in the literature are from one of this species, with a known distribution range in the North Pacific [42]. Hatchlings from species recently described and that have not been recorded outside the type locality were also excluded from the analysis, such as Octopus laqueus [43]. The particular characteristics and anatomy of the nautiluses, which possess very large hatchlings of 26–30 mm [44, 45], are measured through the shell diameter and not by with standard ML, as in other cephalopods. Thus, they were also excluded from the general comparative analysis.

The duration of the planktonic phase (in days, d) in cephalopod species obtained from culture experiments was recorded from the literature (Table 2). For comparisons between species, the maximum duration of the planktonic phase was selected for each species. The hatchling size in relation to the adult size was also explored as an indicator of the relative hatchling size for each species. Here we defined the species hatchling size index (SHSI, %) as: [(mean hatchling ML of the species)/(maximum adult ML of the species)]x100. The maximum adult ML was selected instead of the mean adult ML due to the high intraspecific variability of the latter in the literature. The maximum adult ML (in mm) was recorded in most cases based on recent FAO reviews [2527], except for Octopus vulgaris type II [46] and type IV [47] (Table 1).

Table 2. Size at hatching and at the end of the planktonic phase, rearing temperature and duration of planktonic phase for 15 cephalopod species cultured in the laboratory.

The number of species used in the present study (n = 110) represents 13% of the 845 living cephalopod species described to date [216] and can be considered as a representative sample for this group of molluscs. The number of genus (n = 41) and families (n = 14) analysed here represents 24% and 28% respectively of the 174 genus and 50 families described to date for cephalopods [216]. Nevertheless, some groups may be over-represented in this sample, for example, the number of octopod species analysed here (n = 53) represents nearly half of the total number of species considered in the present study (n = 110), while the total number of octopod species represents nearly one-third of cephalopod species. This fact illustrates that this group is relatively easy to maintain and reproduce in aquaria, making it easier to collect egg masses and hatchlings in comparison with squids, which are more difficult to rear, spawn and consequently, to obtain hatchlings under laboratory conditions [217219].

Latitudinal distribution

Systematics and geographical distribution of the species were determined at first instance according to recent FAO reviews [2527]. Then, for each species, an extensive literature review on its distributional range was conducted. The literature references used to obtain latitudinal distribution assigned to each species are found in Table 1. The maximum and minimum latitudinal distribution ranges obtained for each species was introduced in Google Earth® to determine the latitudinal range of the species in degrees of latitude [220]. This range was transformed into distance (d) in km by applying the haversine formula to calculate the great-circle distance between two points; that is, the shortest distance over the earth’s surface. The haversine formula was used was: d = R·c; where, R = the earth’s radius (mean radius = 6371 km); c = 2·atan2(√a, √(1−a)); a = sin2(Δφ/2) + cos(φ1)·cos(φ2)·sin2(Δλ/2); φ, latitude; λ, longitude.

Data treatment

Values were compared using analysis of variance (ANOVA) and differences were considered significant when P < 0.05. Linear regressions were used for the graphics. Data were assessed using the JMP statistical package.


Hatchling size was recorded for 110 species of cephalopods (30 sepioids, 27 squids, 53 octopods), ranging from 0.6 (Argonauta hians) to 28.0 (Graneledone boreopacifica) mm ML, with an average size of 3.9±3.5 mm ML. Species from 14 families and 41 genera are listed (Table 1). Of these, 38 species have benthic hatchlings and 72 species have planktonic phase of variable duration (Table 2). Sizes and ranges of planktonic and benthic hatchlings are shown in Table 3. The distribution of sizes for species with planktonic hatchlings showed a maximum for Enteroctopus megalocyathus (8.4 mm ML) and a minimum for A. hians. For species with benthic hatchlings, the smallest size was found in Sepiadarium kochii (1.5 mm ML) and the distribution of sizes was highly right-skewed due to the G. boreopacifica hatchling (Fig 1). There was a considerable overlap between the hatchling sizes of benthic and planktonic species for intermediate sizes classes, although the frequency of planktonic hatchlings in these classes was very low (Fig 1). Maximum adult size of the species analysed ranged from 10.5 (Idiosepius biserialis) to 1200 mm (Dosidicus gigas) ML, with an average size of 201±217 mm ML. The species hatchling size index (SHSI) ranged from 0.1 (Thysanoteuthis rhombus) to 21.7% (Rossia molicella), with an average of 4.3±4.2% (Table 1).

Fig 1. Size-frequency distribution from hatchlings of 110 species of cephalopods hatched in the laboratory.

Empty columns represent planktonic hatchlings; black columns, benthic hatchlings.

Table 3. Hatchling size in mantle length (ML) and latitudinal distribution ranges of 110 cephalopod species according to their hatchling mode of life as planktonic or benthic.

In relation to the latitudinal distribution ranges, Hapalochlaena maculosa registered the smallest range (339 km) for a benthic species and the oceanic ommastrephid squid Todaropsis eblanae reached the broadest range for the planktonic species (13070 km) (Table 1). Species with a planktonic phase have significantly smaller hatchling sizes than species that hatched as benthic individuals. In addition, species with planktonic hatchlings display significantly broader latitudinal distribution compared with species that have benthic hatchlings (Table 3).

When all species were pooled together, a significant inverse relationship was observed between hatchling size and latitudinal distributional range, with species with large hatchlings showing smaller latitudinal distribution ranges (ANOVA, F = 7.17, p = 0.009, n = 110) (Fig 2A). The same relationship is observed when grouping all species at the genus level (F = 5.86, p = 0.02, n = 41). This inverse relationship was also significant when analysing all the planktonic species together, where species with smaller planktonic hatchlings reach broader distributions (F = 5.94, p = 0.017, n = 72) (Fig 2C). In contrast, no relationship between hatchling size and latitudinal distribution was found when all the benthic species were analysed together (F = 0.10, p = 0.76, n = 38) (Fig 2E). When the data were analysed between different major cephalopod groups, this relationship was not found for the major benthic cephalopod group, the sepioids (F = 2.61, p = 0.12, n = 30) (Fig 3A). For squids, a group with only planktonic hatchlings, an inverse relationship exists between hatchling size and latitudinal distribution, the smaller the paralarval size the broader the distributional range of the species (F = 4.36, p = 0.047, n = 27) (Fig 3C). For octopods, a cephalopod group with both planktonic and benthic hatchlings, an inverse relationship between hatchling size and species distributional range was also observed (F = 6.90, p = 0.01, n = 53) (Fig 3E).

Fig 2. Relationship between hatchling size in mantle length (ML) and species hatchling size index (SHSI) with the latitudinal distribution range of cephalopod species hatched in the laboratory.

a), c) and e) shows relationship between hatchling size and latitudinal distribution range of the species; b), d) and f), relationship between the SHSI and the latitudinal distribution range of the species. Data from a) and b) are based on all the 110 cephalopod species analysed in this study; data from c) and d) are based on 72 cephalopod species with planktonic hatchlings; data from e) and f) are based on 38 cephalopod species with benthic hatchlings. Logarithmic scale is used for the X-axis. Empty circles represent planktonic species; black circles benthic species.

Fig 3. Relationship between hatchling size in mantle length (ML) and species hatchling size index (SHSI) with the latitudinal distribution range of cephalopod species hatched in the laboratory for the major cephalopod groups.

a), c), e), relationship between hatchling size and latitudinal distribution range of the species; b), d), f), relationship between the SHSI and the latitudinal distribution range of the species. Data from a) and b) are based on 30 species of sepioids, data from c) and d) are based on 27 species of squids, and data from e) and f) are based on 53 species of octopods. Logarithmic scale is used for the X-axis. Empty circles represent planktonic species; black circles benthic species.

When the SHSI is plotted against the latitudinal distribution range of species, the inverse relationship obtained was stronger than when using the mean hatchling size, showing a significant inverse relationship between the relative hatchling size and the latitudinal distribution range for all species (ANOVA, F = 21.45, p = 0.0001, n = 110) (Fig 2B) and for all genera (F = 8.09, p = 0.007, n = 41) pooled together. The same inverse relationship was also observed for all planktonic (F = 8.11, p = 0.006, n = 72) and all benthic (F = 6.51, p = 0.015, n = 38) species (Fig 2D and 2F). In relation to sepioids (F = 6.09, p = 0.02, n = 30), squids (F = 6.16, p = 0.02, n = 27) and octopods (F = 7.88, p = 0.007, n = 53), the same inverse relationship was found (Fig 3B, 3D and 3F).

The duration of the planktonic phase extended from less than a day (8h for Euprymna hyllebergi) to 180 d in Enteroctopus dofleini, with a mean of 53±47 d (Table 2). This duration increased with hatchling size (ANOVA, F = 6.74, p = 0.02, n = 15) (Fig 4A) and no relationship was found between duration of the planktonic phase and latitudinal range (F = 0.08, p = 0.78, n = 14) (Fig 4B). Sepioteuthis lessoniana was excluded from the latter comparison because it is considered a species complex [3537].

Fig 4.

Relationships between (a) hatchling size in mantle length (ML) and duration of the planktonic phase for 15 cephalopod species; and (b) duration of planktonic phase versus latitudinal distribution range for 14 cephalopod species. See Table 2 for details.


Our results revealed that cephalopods with early planktonic developmental modes hatch at smaller sizes and reach broader distributional ranges in comparison with species with large, benthic hatchlings. In addition, for the cephalopod groups with high numbers of species with planktonic hatchlings (squids and octopods), the smaller the hatchling, the broader the latitudinal distributional range of the species. These facts suggest that the developmental mode (planktonic or benthic) along with the hatchling size may influence the distributional range of cephalopods. Furthermore, when the hatchling size index (SHSI) was used for all species with benthic hatchlings, the smaller the hatchlings, the broader the latitudinal distributional range of the species. This also held for the main benthic group, the sepioids. The chances of dispersal by drifting will tend to be greatest in smaller hatchlings. A positive relationship between the presence of a planktonic larval phase and geographic distributional range has been found for prosobranch gastropod species [221], and the present study shows a similar trend for cephalopods.

Lester and Ruttenberg [222] examined the relationship between distributional range and planktonic larval duration of tropical reef fishes and found that this relationship is positively correlated only in the largest ocean basin (the Indo-Pacific). They found that the spatial distribution of habitats and dispersal barriers are of great importance for the dispersal of reef fishes. The authors also noted that the duration of the larval phase is the best quantitative estimate of the dispersal potential of many species. It is conceivable to expect that the smaller the hatchling size, the longer the duration of the planktonic phase. The duration of the planktonic phase in cephalopods has been determined for only 15 species (Table 2), and our results have shown that duration of the planktonic phase increases as hatchling size increases. It is important to stress, however, that this result was strongly influenced by the presence of cold-water octopod species with larger planktonic hatchlings, such as Enteroctopus dofleini and E. megalocyathus (Table 2, Fig 4). Hence, the duration of the planktonic phase in cephalopods seems to be mainly dependent on the species and temperature [7]. Temperature will have a domineering influence, with the potential to increase or decrease the duration of the planktonic phase for a single species. Octopods of the genus Enteroctopus have the largest planktonic cephalopod at hatching and the duration of their planktonic phase extends to 4–6 months, influenced by the relatively cold-water temperatures (11°C) where they develop (Table 2). In addition, E. megalocyathus hatchlings are larger than other octopods at the end of their planktonic phase and settle as juveniles on the bottom, as in Amphioctopus aegina, Octopus joubini, O.vulgaris and Robsonella fontanianus (see Table 2). It would be highly informative to evaluate the factors regulating the duration of the planktonic phase for many species under controlled conditions. Such information could improve our understanding of the influence of the duration of the planktonic phase on the geographical distribution range of cephalopods.

Cephalopods have a relatively short life cycle, around 1–2 years or less in medium-sized coastal species [4], which implies that the time for dispersal is relatively short in comparison with other long-lived marine groups like fishes. Therefore, the planktonic and juvenile phases last relatively longer in relation to the whole life cycle in cephalopod species and must exert a major influence on their dispersal potential, when compared with other marine animals with longer life cycles. The locomotion capacity of the adults is also closely related to species dispersal, although it has only been studied in a few cephalopod species [223]. In this regard, the short life cycle of coastal cephalopods and their relatively brief adult period suggest again the importance of planktonic and juvenile phases for dispersal.

In marine invertebrates, the offspring size variation can arise from different factors such as maternal size and nutrition, habitat quality and stress [224]. The intraspecific variation in hatchling size has the potential to influence the population dynamics. In cephalopods, this intraspecific variability in hatchling size has been reported for several species (see also Table 1) and attributed mainly to egg incubation temperature [91, 104, 225227] and the consequent duration of the embryonic phase [228], seasonality [108] as well as maternal influence [229233]. The effect of temperature on the duration of embryonic development is well known in cephalopods [234]; as temperature increases, the duration of embryonic development decreases, producing smaller hatchlings. On the other hand, at lower temperatures, embryonic development is longer and hatchlings are larger. This temperature effect can occur at seasonal and latitudinal gradients. Moreover, for a single species, larger hatchlings should be expected at higher latitudes [235].

Other factors influencing dispersal that were not considered here include geological history and oceanographic currents. The latter have a domineering effect on dispersal of marine species. For Mediterranean littoral fish species, inshore larvae showed shorter planktonic larval duration than species with offshore larvae. As a result, mean geographic range was smaller for species with inshore larval distribution than for species with offshore larval distribution [236]. These results indicated that planktonic larval duration is certainly not the only factor controlling geographical range, as the main circulation in the inshore-offshore larval habitat as well as the season of planktonic life play important roles in dispersal.

The results of the present study also indicate that hatchling size is related to dispersal potential and display a phenotypic association with the presence of a planktonic phase and developmental mode. Small hatchling size in marine invertebrates is often associated with planktonic developmental mode, high mortality rates, large dispersal potential and likely gene flow. On the other hand, large hatchling size is linked with benthic development and limited dispersal [237]. Genetic data have given general support to the association between developmental mode and intrapopulation variation, with low genetic differentiation being commonly found in planktonic populations, although there are exceptions [238, 239].

Genetic studies in populations of holobenthic octopods showed that individuals are unable to disperse between sites separated by tracts of deep ocean, which apparently present a major physical barrier to dispersal, such as depths > 1000 m for Pareledone turqueti [240]. This isolation, mediated by the limited movement of benthic adults, seems to promote the population differentiation pattern in continuous habitats as in the holobenthic Octopus pallidus [241]. In contrast, merobenthic octopuses showed less consistent patterns, with interactions of multiple factors, such as oceanic currents, duration of the planktonic phase and fitness with settlement areas influencing species dispersal and connectivity [241]. For Octopus vulgaris type II, microsatellite data have revealed significant genetic differentiation in four populations from the SW Atlantic, however, no relationship between geographic distance and genetic differentiation was found [242]. The combination of morphological and microsatellite data, has provide evidences of phylogeographic boundaries for Loligo reynaudii in southern Africa [243], despite its narrow distribution range with no obvious physical boundaries. Many topics remain to be investigated, such as the interactions between paralarval swimming behaviour and wind-driven circulation, which may strongly affect dispersal and retention patterns, leading to many possible explanations for genetic and morphologic diversity.

According to Strahmann and Strahmann [244], the recruitment variability inherent to species with small-sized hatchlings and planktonic development seems to be incompatible with short life cycles and low fecundity of small-sized species. Planktonic developmental mode plays both the role in feeding and dispersal. Thus, a greater incidence of planktonic development should be expected in large animals with longer life spans. Interestingly, though, a study that correlated adult size with oocyte size and inferred developmental mode in shallow water benthic octopuses suggested that oocyte size is negatively related to body size and thus, species with larger body sizes tend to have smaller oocytes (and likely planktonic hatchlings) compared to smaller body sized species [171]. It should be noted that octopus species with considerably large body sizes such as Enteroctopus dofleini and E. megalocyathus, have planktonic hatchlings, in agreement with the suggestions of both Guzik [171] and Strahmann and Strahmann [244]. On the other hand, polar regions seem to have selected for the production of large benthic hatchlings as there is abundant food during the productive summers months minimizing post-settlement competition [245]. This appears to apply particularly to polar and deep-sea octopus species, which have very large hatchlings that are produced over long to exceptionally long incubation periods, as in Graneledone boreopacifica, the species with the longest egg brooding period (53 months) ever registered for an animal [246].

The deep-sea and polar octopuses show a clear tendency for large hatchlings. Phylogenetic evidence suggests that polar and deep-sea octopuses, all with benthic hatchlings, have shallow water ancestors with planktonic paralarvae [247, 248]. In the deep-sea, the main incirrate octopods such as Bathypolypus, Benthoctopus and Graneledone, have very large eggs, suggesting benthic hatchlings [249251]. In this bathyal and abyssal environment, the suborder of the cirrate octopods probably represents an exception to the rule. These typical deep-sea cephalopods with a gelatinous consistency and well-developed fins, spawn very large eggs suggesting direct developing juveniles [252]. However, the cirrate octopod family Cirroteuthidae (Cirrothauma, Cirroteuthis, Stauroteuthis) are essentially pelagic, but live generally close to the sea floor, and are characterized by very large fins and swimming behaviour [253255]. The Cirroteuthidae morphology suggests a possible planktonic or benthopelagic mode of development for large hatchlings in the deep-sea, because very large fins were observed in advanced cirrate embryos of 9 mm ML [256]. This exception could also be extended to the old cephalopod lineage of the nautiluses, with very large and pelagic hatchlings [44, 45, 257]. Increased dispersal capacity has the potential to impact recruitment variability and thus, may have several consequences for the whole life cycle. It is important to emphasize, however, that dispersal is not only achievable during the larval phase of the life cycle, but that other factors such as adult body size and locomotion capacities also play a role. There may be interesting insights to be gained from exploring the importance of developmental mode and dispersal with gene flow in cephalopod populations.


Cephalopod species with smaller planktonic hatchlings seem to reach larger distributional extensions in comparison with species with large, benthic hatchlings. This seems evident for squids and octopods, where species with larger hatchlings have geographical distributions of comparatively minor extension. This general tendency was not detected for sepioids, a more homogeneous group composed mainly of species with large and benthic hatchlings. However, when observing the relative size of the hatchlings in comparison with the adults, the smaller the hatchlings, the broader the latitudinal distribution range of sepioids. This was also valid for all species with benthic hatchlings pooled together, thus confirming the influence of hatchling size on dispersal potential. The duration of the planktonic phase is also believed to be an important factor influencing the species geographical distribution. However, this has been determined only for a few cephalopod species to date and future research is needed on this topic.


Ms. Marta Campos helped in obtaining species latitudinal distributions using Google Earth®. We wish to thank Enrique Macpherson for his comments on an early version of the manuscript.

Author Contributions

  1. Conceptualization: RV EAGV FAFA JN.
  2. Formal analysis: RV EAGV FAFA JN.
  3. Investigation: RV EAGV FAFA JN.
  4. Methodology: RV EAGV FAFA JN.
  5. Writing – original draft: RV EAGV FAFA JN.
  6. Writing – review & editing: RV EAGV FAFA JN.


  1. 1. Nathan R. The challenges of studying dispersal. Trends in Ecology & Evolution. 2001;16(9):481–3.
  2. 2. Boero F, Bouillon J. Zoogeography and life cycle patterns of Mediterranean hydromedusae (Cnidaria). Biological journal of the Linnean Society. 1993;48(3):239–66.
  3. 3. Cowen RK, Sponaugle S. Larval dispersal and marine population connectivity. Annual Review of Marine Science. 2009;1(1):443–66.
  4. 4. Boyle PR, Rodhouse PG. Cephalopods: ecology and fisheries: Blackwell Publishers; 2005. 452 p.
  5. 5. Doubleday ZA, Prowse TAA, Arkhipkin A, Pierce GJ, Semmens J, Steer M, et al. Global proliferation of cephalopods. Current Biology. 2016;26(10):R406–R7. pmid:27218844
  6. 6. Rosa R, Gonzalez L, Dierssen H, Seibel B. Environmental determinants of latitudinal size-trends in cephalopods. Marine Ecology Progress Series. 2012;464:153–65.
  7. 7. Villanueva R, Norman MD. Biology of the planktonic stages of benthic octopuses. Oceanography and Marine Biology: An Annual Review. 2008;46:105–202. pmid:ISI:000256878700004.
  8. 8. Young RE, Harman RF. "Larva," "paralarva" and "subadult" in cephalopod terminology. Malacologia. 1988;29(1):201–7.
  9. 9. Rowell TW, Trites RW, Dawe EG. Distribution of short-finned squid (Illex illecebrosus) larvae and juveniles in relation to the Gulf stream frontal zone between Florida and Cape Hatteras. NAFO Scientific Council Studies. 1985;9:77–92.
  10. 10. Saito H, Kubodera T. Distribution of Ommastrephid rhynchoteuthion paralarvae (Mollusca, Cephalopoda) in the Kuroshio region. In: Okutani T, O'Dor RK, Kubodera T, editors. Recent advances in Fisheries Biology. Tokyo: Tokai University Press; 1993. p. 457–66.
  11. 11. Bower JR, Seki MP, Young RE, Bigelow KA, Hirota J, Flament P. Cephalopod paralarvae assemblages in Hawaiian Islands waters. Marine Ecology Progress Series. 1999;185:203–12.
  12. 12. Roberts MJ, van den Berg M. Currents along the Tsitsikamma coast, South Africa, and potential transport of squid paralarvae and ichthyoplankton. African Journal of Marine Science. 2005;27:375–88.
  13. 13. Martins RS, Roberts MJ, Lett C, Chang N, Moloney CL, Camargo MG, et al. Modelling transport of chokka squid (Loligo reynaudii) paralarvae off South Africa: reviewing, testing and extending the ‘Westward Transport Hypothesis’. Fisheries Oceanography. 2014;23(2):116–31.
  14. 14. Downey-Breedt NJ, Roberts MJ, Sauer WHH, Chang N. Modelling transport of inshore and deep-spawned chokka squid (Loligo reynaudi) paralarvae off South Africa: the potential contribution of deep spawning to recruitment. Fisheries Oceanography. 2016;25(1):28–43.
  15. 15. Kubodera T. Distribution and abundance of the early life stages of octopus, Octopus dofleini Wülker, 1910 in the North Pacific. Bull Mar Sci. 1991;49(1–2):235–43.
  16. 16. Otero J, Alvarez-Salgado XA, Gonzalez AF, Gilcoto M, Guerra A. High-frequency coastal upwelling events influence Octopus vulgaris larval dynamics on the NW Iberian shelf. Marine Ecology-Progress Series. 2009;386:123–32. pmid:CCC:000268552500011.
  17. 17. Packard A. Jet propulsion and the giant fibre response of Loligo. Nature. 1969;221:875–7. pmid:5765066
  18. 18. Chen DS, Dykhuizen , Hodge J, Gilly WF. Ontogeny of copepod predation in juvenile squid (Loligo opalescens). Biol Bull. 1996;190:69–81. pmid:8852631
  19. 19. Villanueva R, Nozais C, Boletzky SV. Swimming behaviour and food searching in planktonic Octopus vulgaris Cuvier from hatching to settlement. Journal of Experimental Marine Biology and Ecology. 1997;208(1–2):169–84. Ipmid:SI:A1997WC65300011.
  20. 20. Bartol IK, Krueger PS, Stewart WJ, Thompson JT. Pulsed jet dynamics of squid hatchlings at intermediate Reynolds numbers. J Exp Biol. 2009;212(10):1506–18. pmid:CCC:000265680000016.
  21. 21. Bartol IK, Krueger PS, Thompson JT, Stewart WJ. Swimming dynamics and propulsive efficiency of squids throughout ontogeny. Integrative and Comparative Biology. 2008;48(6):720–33. pmid:CCC:000262760700004.
  22. 22. Yoo H-K, Yamamoto J, Saito T, Sakurai Y. Laboratory observations on the vertical swimming behavior of Japanese common squid Todarodes pacificus paralarvae as they ascend into warm surface waters. Fish Sci. 2014;80(5):925–32.
  23. 23. Dan S, Hamasaki K, Yamashita T, Oka M, Kitada S. Age-based life cycle traits of the broadclub cuttlefish Sepia latimanus confirmed through release and recapture experiments. Aquatic Biology. 2012;17(2):181–95.
  24. 24. Perez-Losada M, Guerra A, Carvalho GR, Sanjuan A, Shaw PW. Extensive population subdivision of the cuttlefish Sepia officinalis (Mollusca: Cephalopoda) around the Iberian Peninsula indicated by microsatellite DNA variation. Heredity. 2002;89(6):417–24. pmid:12466983
  25. 25. Jereb P, Roper CFE. Cephalopods of the world. An annotated and illustrated catalogue of cephalopod species known to date. Rome: FAO Species catalogue for Fishery Purposes No. 4, Vol. 1 Chambered nautiluses and sepioids (Nautilidae, Sepiidae, Sepiolidae, Sepiadariidae, Idiosepiidae and Spirulidae); 2005. 262 p.
  26. 26. Jereb P, Roper CFE. Cephalopods of the world. An annotated and illustrated catalogue of cephalopod species known to date. Rome: FAO Species catalogue for Fishery Purposes No. 4, Vol. 2. Myopsid and Oegopsid Squids.; 2010. 605 p.
  27. 27. Jereb P, Roper CFE, Norman MD, Finn JK. Cephalopods of the world. An annotated and illustrated catalogue of cephalopod species known to date. Volume 3. Octopods and Vampire Squids. FAO Species Catalogue for Fishery Purposes. Rome, FAO. 2013. 370 p.
  28. 28. Bergstrom B, Summers WC. Sepietta oweniana. In: Boyle PR, editor. Cephalopod life cycles Volume 1 Species accounts: Academic Press London; 1983. p. 75–91.
  29. 29. Boletzky SV, Fuentès M, Offner N. First record of spawning and embryonic development in Octopus macropus (Mollusca: Cephalopoda). Journal of the Marine Biological Association of the United Kingdom. 2001;81(4):703–4.
  30. 30. Boletzky SV. Le développment d'Eledone moschata (Mollusca, Cephalopoda) élevée au laboratoire. Bulletin de la Societé Zoologique de France. 1975;100(3):361–7.
  31. 31. Voight JR, Drazen JC. Hatchlings of the deep-sea octopus Graneledone boreopacifica are the largest and most advanced known. Journal of Molluscan Studies. 2004;70:406–8.
  32. 32. Boletzky SV. The embryonic development of the octopus Scaeurgus unicirrhus (Mollusca, Cephalopoda). Additional data and discussion. Vie et Milieu. 1984;334:87–93.
  33. 33. Anderson FE, Engelke R, Jarrett K, Valinassab T, Mohamed KS, Asokan PK, et al. Phylogeny of the Sepia pharaonis species complex (Cephalopoda: Sepiida) based on analyses of mitochondrial and nuclear DNA sequence data. Journal of Molluscan Studies 2011 77 (1): 65–75
  34. 34. Nabhitabhata J, Tuanapaya S. Allopatric variation of Sepia pharaonis Ehrenberg, 1831 complex (Cephalopoda: Sepiida) in two oceans of Thai waters based on morphology and mitochondrial DNA sequences. Cephalopod International Advisory Council; Hakodate, Japan2015. p. 97.
  35. 35. Jivaluk J, Nabhitabhata J, Nateewathana A, Watprasit P. Description of the Thai type of Bigfin Reef squid, Sepioteuthis lessoniana, hatchling with note on comparison to Japanese types. Phuket mar biol Cent Res Bull. 2005;66:117–26.
  36. 36. Triantafillos L, Adams M. Genetic evidence that the northern calamary, Sepioteuthis lessoniana, is a species complex in Australian waters. ICES Journal of Marine Science. 2005;62:1665–70.
  37. 37. Cheng SH, Anderson FE, Bergman A, Mahardika GN, Muchlisin ZA, Dang BT, et al. Molecular evidence for co-occurring cryptic lineages within the Sepioteuthis cf. lessoniana species complex in the Indian and Indo-West Pacific Oceans. Hydrobiologia. 2014;725(1):165–88.
  38. 38. Sales JBL, Markaida U, Shaw PW, Haimovici M, Ready JS, Figueredo-Ready WMB, et al. Molecular phylogeny of the genus Lolliguncula Steenstrup, 1881 based on nuclear and mitochondrial DNA sequences indicates genetic isolation of populations from north and south Atlantic, and the possible presence of further cryptic species. PLOS ONE. 2014;9(2):e88693. pmid:24586371
  39. 39. Staaf D, Ruiz-Cooley R, Elliger C, Lebaric Z, Campos B, Markaida U, et al. Ommastrephid squids Sthenoteuthis oualaniensis and Dosidicus gigas in the eastern Pacific show convergent biogeographic breaks but contrasting population structures. Marine Ecology Progres Series. 2010;418:168–78.
  40. 40. Carrasco SA. The early life history of two sympatric New Zealand octopuses: eggs and paralarvae of Octopus huttoni and Pinnoctopus cordiformis. New Zealand journal of zoology. 2014;41(1):32–45.
  41. 41. Fernández-Álvarez F, Sánchez P, Cuesta-Torralvo E, Escánez A, Martins C, Vidal E, et al. The genus Ommastrephes d'Orbigny, 1834: a single species or more than one hidden behind a single name?. Cephalopod International Advisory Council; Hakodate, Japan: Book of Abstracts; 2015. p. 103.
  42. 42. Roper CFE, Nigmatullin CM, Jereb P. Family Ommastrephidae. In Jereb P. & Roper C.F.E., eds. Cephalopods of the world. An annotated and illustrated catalogue of species known to date. Volume 2. Myopsid and Oegopsid Squids. FAO Species Catalogue for Fishery Purposes. No. 4, Vol. 2. Rome, FAO. pp. 269–347. 2010.
  43. 43. Kaneko N, Oshima Y, Ikeda Y. Egg brooding behavior and embryonic development of Octopus laqueus (Cephalopoda: Octopodidae). Molluscan Research. 2006;26(3):113–7. pmid:CCC:000243238600001.
  44. 44. Arnold JM, Awai M, Carlson B. Hatching of Nautilus embryos in the Waikiki Aquarium. Journal of Cephalopod Biology. 1990;1:117.
  45. 45. Wood JB, O'Dor RK. Do larger cephalopods live longer? Effects of temperature and phylogeny on interspecific comparisons of age and size at maturity. Mar Biol. 2000;136(1):91–9. pmid:ISI:000085966300012.
  46. 46. Teixeira PB. Biologia reprodutiva do polvo, Octopus vulgaris Cuvier 1797 no sul do Brasil: Universidade Federal de Santa Catarina; 2011.
  47. 47. Gleadall IG. Octopus sinensis (Cephalopoda: Octopodidae): valid species name for the commercially valuable East Asian common octopus. Species Diversity. 2016; 21:31–42.
  48. 48. Choe S. On the eggs, rearing, habits of the fry and growth of some cephalopods. Bulletin of Marine Science. 1966;16:330–48.
  49. 49. Reid A, Jereb P. Family Sepiolidae. In: Jereb P, Roper CFE, editors. Cephalopods of the world An annotated and illustrated catalogue of species known to date Vol 1: Chambered nautiluses and sepioids (Nautilidae, Sepiidae, Sepiolidae, Sepiadariidae, Idiosepiidae and Spirulidae). FAO Species Catalogue for Fishery Purposes. Rome: FAO; 2005. p. 153–203.
  50. 50. Nabhitabhata J, Nilaphat P, Promboon P, Jaroongpattananon C. Life cycle of cultured bobtail squid, Euprmna hyllerbergi Nateewathana, 1997. Phuket Mar Biol Cent Res Bull. 2005;66:351–65.
  51. 51. Aungtonya C, Nateewathana A, SecherTendal O, Nabhitabhata J. New records of the bobtail squid Euprymna hyllebergi Nateewathana, 1997 with designation of a neotype. Phuket Mar Biol Cent Res Bull 2011;70:15–27.
  52. 52. Hanlon RT, Claes MF, Ashcraft SE, Dunlap PV. Laboratory culture of the sepiolid squid Euprymna scolopes: a model system for bacteria—animal symbiosis. Biological Bulletin. 1997;192:364–74.
  53. 53. Singley CT. Euprymna scolopes. In: Boyle PR, editor. Cephalopod life cycles Volume 1 Species accounts: Academic Press London; 1983. p. 69–74.
  54. 54. Moltschaniwskyj NA. pers comm.
  55. 55. Nabhitabhata J. Distinctive behaviour of Thai pygmy squid, Idiosepius thailandicius Chotiyaputta, Okutani & ChaitiaMVong, 1991. Phuket Marine Biological Center Special Publication. 1998;18(1):25–40.
  56. 56. Jv Byern, Söller R, Steiner G. Phylogenetic characterisation of the genus Idiosepius (Cephalopoda; Idiosepiidae). Aquatic Biology. 2012;17(1):19–27.
  57. 57. Natsukari Y. Egg-laying behavior, embryonic development and hatched larva of the pygmy cuttlefish Idiosepius pygmaeus paradoxus Ortmann. Bulletin of the Faculty of Fisheries, Nagasaki University. 1970;30:15–29.
  58. 58. Sato N, Awata S, Munehara H. Seasonal occurrence and sexual maturation of Japanese pygmy squid (Idiosepius paradoxus) at the northern limits of their distribution. Ices Journal of Marine Science. 2009;66(5):811–5. pmid:CCC:000266350100003.
  59. 59. Nabhitabhata J, Nilaphat P, Reunreng A, Promboon P. Culture, growth and behaviour of sharp-tail Pygmy squid, Idiosepius pygmaeus Steenstrup, 1881. Rayong Coastal Fisheries Research and Development Center. 2004; (Contribution No. 27).
  60. 60. Norman MD, Reid A. A guide to squid, cuttlefish and octopuses of Australasia. Victoria, Australia: CSIRO Publishing; 2000. 96 p.
  61. 61. Voss GL. Cephalopods of the Phillipine Islands. United States National Museum Bulletin. 1963;234:1–180.
  62. 62. Grasse B. The biological characteristics, life cycle, and system design for the flamboyant and paintpot cuttlefish, Metasepia sp., cultured through multiple generations. Drum and Croaker. 2014;45:58–71.
  63. 63. Reid A, Jereb P, Roper CFE. Family Sepiidae. In P. Jereb & C.F.E. Roper, eds. Cephalopods of the world. An annotated and illustrated catalogue of species known to date. Volume 1. Chambered nautiluses and sepioids (Nautilidae, Sepiidae, Sepiolidae, Sepiadariidae, Idiosepiidae and Spirulidae). FAO Species Catalogue for Fishery Purposes. No. 4, Vol. 1. Rome, FAO. 2005:pp. 57–152.
  64. 64. Umezu Y, Tsuchiya K, Segawa S. Growth, maturation and metabolism of reared Metasepia tullbergi. Cephalopod International Advisory Council; Florianopolis, Brazil: Official Program; 2012. p. 28.
  65. 65. Boletzky SV, Boletzky MV. Observations on the embryonic and early post-embryonic development of Rossia macrosoma (Mollusca: Cephalopoda). Helgol Mar Res. 1973;20:135–61.
  66. 66. Okutani T, Sasaki T. Eggs of Rossia mollicella (Cephalopoda: Sepiolidae) deposited in a deep-sea sponge. J Mollus Stud. 2007;73(3):287–9.
  67. 67. Nesis KN. Cephalopods of the World. Burguess LA, editor. Neptune City: T.F.H. Publications; 1987. 351 p.
  68. 68. Summers WC, Colvin L. On the cultivation of Rossia pacifica (Berry, 1911). Journal of Cephalopod Biology. 1989;1:21–32.
  69. 69. Cronin ER, Seymour RS. Respiration of the eggs of the giant cuttlefish Sepia apama. Marine Biology. 2000;136(5):863–70.
  70. 70. Warnke KM, Kaiser R, Hasselmann M. First observations of a snail-like body pattern in juvenile Sepia bandensis (Cephalopoda: Sepiidae). A note. Neues Jahrbuch Fur Geologie Und Palaontologie-Abhandlungen. 2012;266(1):51–7. pmid:WOS:000309508800007.
  71. 71. Natsukari Y, Tashiro M. Neritic squid resources and cuttlefish resources in Japan. Marine Behaviour and Physiology. 1991;18(3):149–226. pmid:ISI:A1991FW46200001.
  72. 72. Okutani T. Studies on early life cycle of Decapodan Mollusca—VII. Eggs and newly hatched larvae of Sepia latimanus Quoy & Gaimard. Venus. 1978;37:245–8.
  73. 73. Boletzky SV, Hanlon RT. A review of the laboratory maintenance, rearing and culture of cephalopod molluscs. Memoirs of the National Museum of Victoria. 1983;44:147–87.
  74. 74. Watanabe K, Sakurai Y, Segawa S, Okutani T. Development of the ommastrephid squid Todarodes pacificus, from fertilized egg to rhynchoteuthion paralarva. Am Malacol Bull. 1996;13:73–88.
  75. 75. Boletzky SV. Sepia officinalis. In: Boyle PR, editor. Cephalopod life cycles Volume 1 Species accounts: Academic Press London; 1983. p. 31–52.
  76. 76. Forsythe JW, Derusha RH, Hanlon RT. Growth, reproduction and life span of Sepia officinalis (Cephalopoda, Mollusca) cultured through seven consecutive generations. Journal of Zoology. 1994;233:175–92. pmid:ISI:A1994NV72000001.
  77. 77. Boletzky SV. Preliminary observations on laboratory-reared Sepia orbignyana (Mollusca, Cephalopoda). Rapports et Procès-verbaux des Réunions Commission International pour l'Exploration Scientifique de la Mer Méditerranée. 1988;31:256.
  78. 78. Nabhitabhata J, Nilaphat P, Promboon P, Jaroongpattananon C, Nilaphat G, Reunreng A. Performance of simple large-scale cephalopod culture system in Thailand. Phuket marine biological Center Research Bulletin. 2005;66:337–50.
  79. 79. Reid A. 2005. Family Sepiadariidae. In P. Jereb & C.F.E. Roper, eds. Cephalopods of the world. An annotated and illustrated catalogue of species known to date. Volume 1. Chambered nautiluses and sepioids (Nautilidae, Sepiidae, Sepiolidae, Sepiadariidae, Idiosepiidae and Spirulidae). FAO Species Catalogue for Fishery Purposes. No. 4, Vol. 1. Rome, FAO. pp. 204–207. 2005.
  80. 80. Nabhitabhata J. Life cycle of three cultured generations of spineless cuttlefish, Sepiella inermis (Férussac and D´Orbigny, 1848). Phuket Marine Biological Center Special Publication. 1997;17(1):289–98.
  81. 81. Zheng XD, Xiangzhi L, Zhaokai W, Ruihai Y, Chuanyuan T, Qi L. Studies on the life cycle of cultured spineless cuttlefish Sepiella japonica. Transactions of Oceanology and Limnology. 2010;3:24–8.
  82. 82. Boletzky SV, Boletzky MV, Frösch D, Gätzi V. Laboratory rearing of Sepiolinae (Mollusca, Cephalopoda). Mar Biol. 1971;8:82–7.
  83. 83. Mauris E. Colour patterns and body postures related to prey capture in Sepiola affinis (Mollusca: Cephalopoda). Marine Behaviour and Physiology. 1989;14:189–200.
  84. 84. Jones NJE, Richardson CA. Laboratory culture, growth, and the life cycle of the little cuttlefish Sepiola atlantica (Cephalopoda: Sepiolidae). Journal of Shellfish Research. 2010;29(1):241–6. pmid:CCC:000276267900031.
  85. 85. Rodrigues M, Garci ME, Troncoso JS, Guerra A. Spawning strategy in Atlantic bobtail squid Sepiola atlantica (Cephalopoda: Sepiolidae). Helgol Mar Res. 2011;65(1):43–9. pmid:ISI:000287329300005.
  86. 86. Boletzky SV. Sepiola robusta. In: Boyle PR, editor. Cephalopod life cycles Volume 1 Species accounts: Academic Press London; 1983. p. 53–67.
  87. 87. Guerra A, Rocha F, Gonzalez AF, Buckle LF. Embryonic stages of the Patagonian squid Loligo gahi (Mollusca: Cephalopoda). Veliger. 2001;44(2):109–15. pmid:ISI:000167997400001.
  88. 88. Arkhipkin AI, Laptihovsky VV, Middleton DAJ. Adaptations for cold water spawning in loliginid squid: Loligo gahi in Falkland waters. Journal of Molluscan Studies. 2000;65:551–64.
  89. 89. McConathy DA, Hanlon RT, Hixon RF. Chromatophore arrangements of hatchling loliginid squids (Cephalopoda, Myopsida). Malacologia. 1980;19(2):279–88. pmid:ISI:A1980JP39900005.
  90. 90. Yang WT, Hixon RF, Turk PE, Krejci ME, Hulet WH, Hanlon RT. Growth, behavior, and sexual maturation of the market squid, Loligo opalescens, cultured through the life cycle. Fishery Bulletin. 1986;84(4):771–98. pmid:ZOOR:ZOOR12400026273.
  91. 91. Vidal EAG, DiMarco FP, Wormuth JH, Lee PG. Influence of temperature and food availability on survival, growth and yolk utilization in hatchling squid. Bull Mar Sci. 2002;71(2):915–31. pmid:ISI:000182787600025.
  92. 92. Hanlon RT, Turk PE, Lee PG, Yang WT. Laboratory rearing of the squid Loligo pealei to the juvenile stage: growth comparisons with fishery data. Fish Bull. 1987;85:163–7.
  93. 93. Baeg GH, Sakurai Y, Shimazaki K. Embryonic stages of Loligo bleekeri Keferstein (Mollusca: Cephalopoda). Veliger. 1992;35(3):234–41.
  94. 94. Ikeda YS I; Ito K; Sakurai Y; Matsumoto G. Rearing of squid hatchlings, Heterololigo bleekeri (Keferstein 1866) up to 2 months in a closed seawater system. Aquac Res. 2005;36:409–12.
  95. 95. Segawa S, Yang WT, Marthy HJ, Hanlon RT. Illustrated embryonic stages of the Eastern Atlantic squid Loligo forbesi. Veliger. 1988;30(3):230–43. pmid:ISI:A1988Q265400003.
  96. 96. Hanlon RT, Yang WT, Turk PE, Lee PG, Hixon RF. Laboratory culture and estimated life span of the Eastern Atlantic squid, Loligo forbesi Steenstrup, 1856 (Mollusca: Cephalopoda). Aquaculture and Fisheries Management. 1989;20:15–34.
  97. 97. Vecchione M, Lipinski MR. Descriptions of the paralarvae of two loliginid squids in southern African waters. South African Journal of Marine Science. 1995;15:1–7.
  98. 98. Blackburn S, Sauer WH, Lipinski MR. The embryonic development of the chokka squid Loligo vulgaris reynaudii d'Orbigny, 1845. The Veliger. 1998;4(3):249–58.
  99. 99. Vidal EAG, Roberts MJ, Martins RS. Yolk utilization, metabolism and growth in reared Loligo vulgaris reynaudii paralarvae. Aquatic and Living Resources. 2005;18:386–93.
  100. 100. Martins RS, Roberts MJ, Vidal ÉAG, Moloney CL. Effects of temperature on yolk utilization by chokka squid (Loligo reynaudii d'Orbigny, 1839) paralarvae. Journal of Experimental Marine Biology and Ecology. 2010;386(1–2):19–26.
  101. 101. Villanueva R, Sanchez P. Cephalopods of the Benguela Current off Namibia—new additions and considerations on the genus Lycoteuthis. Journal of Natural History. 1993;27(1):15–46. pmid:ISI:A1993KQ57500002.
  102. 102. Turk PE, Hanlon RT, Bradford LA, Yang WT. Aspects of feeding, growth and survival of the European squid Loligo vulgaris Lamarck, 1799, reared through the early growth stages. Vie et Milieu. 1986;36:9–13.
  103. 103. Hanlon RT, Boletzky SV, Okutani T, Perez-Gandaras G, Sanchez P, Sousa-Reis C, et al. Suborder Myopsida Orbigny, 1845. In: Sweeney MJ, Roper CFE, Mangold KM, Clarke MR, Boletzky SV, editors. "Larval" and juvenile cephalopods: A manual for their identification. 513. Washington, D.C.: Smithsonian Institution Press; 1992. p. 37–53.
  104. 104. Villanueva R. Effect of temperature on statolith growth of the European squid Loligo vulgaris during early life. Marine Biology. 2000;136(3):449–60. pmid:ISI:000087050800007.
  105. 105. Mladineo I, Valic D, Jozic M. Spawning and early development of Loligo vulgaris Lamarck,1798, under experimental conditions. Acta Adriatica. 2003;44:77–83.
  106. 106. Fernández-Álvarez F. personal observation.
  107. 107. Cardoso F, Hochberg FG. Revision of the genus Lolliguncula Steenstrup, 1881 (Cephalopoda: Loliginidae) off the Pacific Coast of South America. Revista Peruana de Biología. 2013;20:129–36.
  108. 108. Steer MA, Pecl GT, Moltschaniwskyj NA. Are bigger calamary Sepioteuthis australis hatchlings more likely to survive? A study based on statolith dimensions. Marine Ecology-Progress Series. 2003;261:175–82. pmid:ISI:000186704400014.
  109. 109. Steer MA, Moltschaniwskyj NA, Jordan AR. Embryonic development of southern calamary (Sepioteuthis australis) within the constraints of an aggregated egg mass. Marine and Freshwater Research. 2003;54:217–26.
  110. 110. LaRoe ET. The culture and maintenance of the loliginid squids Sepioteuthis sepioidea and Doryteuthis plei. Mar Biol. 1971;9(1):9–25.
  111. 111. Yatsu A, Tafur R, Maravi C. Embryos and rhynchoteuthion paralarvae of the jumbo flying squid Dosidicus gigas (Cephalopoda) obtained through artificial fertilization from Peruvian waters. Fish Sci. 1999;65:904–8.
  112. 112. Staaf DJ, Camarillo-Coop S, Haddock SHD, Nyack AC, Payne J, Salinas-Zavala CA, et al. Natural egg mass deposition by the Humboldt squid (Dosidicus gigas) in the Gulf of California and characteristics of hatchlings and paralarvae. Journal of the Marine Biological Association of the United Kingdom. 2008;88(4):759–70. pmid:ISI:000257947000015.
  113. 113. Bower JR, Seki K, Kubodera T, Yamamoto J, Nobetsu T. Brooding in a Gonatid squid off Northern Japan. Biological Bulletin. 2012;223(3):259–62. pmid:WOS:000313389600003.
  114. 114. Roper CFE, Jorgensen EM, Katugin ON, Jereb P. Family Gonatidae. In P. Jereb & C.F.E. Roper, eds. Cephalopods of the world. An annotated and illustrated catalogue of species known to date. Volume 2. Myopsid and Oegopsid Squids. FAO Species Catalogue for Fishery Purposes. No. 4, Vol. 2. Rome, FAO. pp. 200–222. 2010.
  115. 115. Arkhipkin AI, Seibel BA. Statolith microstructure from hatchlings of the oceanic squid, Gonatus onyx (Cephalopoda, Gonatidae) from the Northeast Pacific. Journal of Plankton Research. 1999;21(2):401–4. pmid:ISI:000078695000013.
  116. 116. Okutani T, Kubodera T, Jefferts K. Diversity, distribution and ecology of gonatid squids in the subarctic Pacific: a review. Bulletin of the Ocean Research Institute University of Tokyo. 1988;26(1). pmid:ZOOR:ZOOR12500048282.
  117. 117. Sakai M, Brunetti NE, Elena B, Sakurai Y. Embryonic development and hatchlings of Illex argentinus derived from artificial fertilization. South African Journal of Marine Science-Suid-Afrikaanse Tydskrif Vir Seewetenskap. 1998;20:255–65. pmid:ISI:000165531100028.
  118. 118. Villanueva R, Quintana D, Petroni G, Bozzano A. Factors influencing the embryonic development and hatchling size of the oceanic squid Illex coindetii following in vitro fertilization. Journal of Experimental Marine Biology and Ecology. 2011;407(1):54–62.
  119. 119. Durward RD, Vessey E, O'Dor RK, Amaratunga T. Reproduction in the squid, Illex illecebrosus: first observation in captivity and implications for the life cycle. International Commision for the Nortwest Atlantic Fisheries Selected Papers. 1980;6:7–13.
  120. 120. O'Dor RK, Foy EA, Helm PL, Balch N. The locomotion and energetics of hatching squid, Illex illecebrosus. Am Malacol Bull. 1986;4(1):55–60.
  121. 121. Sakurai Y, Young RE, Hirota J, Mangold K, Vecchione M, Clarke MR, et al. Artificial fertilization and development through hatching in the oceanic squids Ommastrephes bartramii and Sthenoteuthis oualaniensis (Cephalopoda: Ommastrephidae). Veliger. 1995;38:185–91.
  122. 122. Miyahara K, Katsuya F, Ota T, Minami T. Laboratory observations on the early life stages of the Diamond squid Thysanoteuthis rhombus. Journal of Molluscan Studies. 2006;72:199–205.
  123. 123. Roper CFE, Jereb P. Family Thysanoteuthidae. In P. Jereb & C.F.E. Roper, eds. Cephalopods of the world. An annotated and illustrated catalogue of species known to date. Volume 2. Myopsid and Oegopsid Squids. FAO Species Catalogue for Fishery Purposes. No. 4, Vol. 2. Rome, FAO. pp. 384–387. 2010.
  124. 124. Puneeta P, Vijai D, Yoo H-K, Matsui H, Sakurai Y. Observations on the spawning behavior, egg masses and paralarval development of the ommastrephid squid Todarodes pacificus in a laboratory mesocosm. J Exp Biol. 2015;218(Pt 23):3825–35. pmid:26632456
  125. 125. Fernández-Álvarez FA, Martins CPP, Vidal EAG, Villanueva R. Towards the identification of the ommastrephid squid paralarvae (Mollusca: Cephalopoda): morphological description of three species and a key to the Northeast Atlantic species. Zoological Journal of the Linnean Society. 2016;in press
  126. 126. Hayashi S. Fishery biological studies of the firefly squid, Watasenia scintillans, in Toyama Bay. Bulletin of Toyama Prefectural Fisheries Research Institute. 1995;7:1–128.
  127. 127. Roper CFE, Jereb P. Family Enoploteuthidae. In P. Jereb & C.F.E. Roper, eds. Cephalopods of the world. An annotated and illustrated catalogue of species known to date. Volume 2. Myopsid and Oegopsid Squids. FAO Species Catalogue for Fishery Purposes. No. 4, Vol. 2. Rome, FAO. pp. 183–200. 2010.
  128. 128. Young RE. Brooding in a bathypelagic octopus. Pacific Science. 1972;26:400–4.
  129. 129. Thore S. Investigations on the "Dana" Octopoda. I. Bolitaenidae, Amphitrelidae, Vitreledonellidae, and Allopsidae. Dana Report. 1949;33:1–85.
  130. 130. Hochberg FG, Nixon M, Toll RB. Order Octopoda Leach, 1818. In: Sweeney MJ, Roper CFE, Mangold KM, Clarke MR, Boletzky SV, editors. "Larval" and juvenile cephalopods: A manual for their identification. 513. Washington, D.C.: Smithsonian Institution Press; 1992. p. 213–80.
  131. 131. Thomas RF. Systematics, distribution, and biology of cephalopods of the genus Tremoctopus (Octopoda: Tremoctopodidae). Bulletin of Marine Science. 1977;27(3):353–92.
  132. 132. O'Shea S. The marine fauna of New Zealand: Octopoda (Mollusca: Cephalopoda). NIWA Biodiversity Memoir. 1999;112:280 p. pmid:ZOOR:ZOOR13600052945.
  133. 133. Finn J. Family Argonautidae. In P. Jereb, C.F.E. Roper, M.D. Norman & J. K Finn eds. Cephalopods of the world. An annotated and illustrated catalogue of cephalopod species known to date. Volume 3. Octopods and Vampire Squids. FAO Species Catalogue for Fishery Purposes. No. 4, Vol. 3. Rome, FAO. pp. 228–237. 2013.
  134. 134. Sukhsangchan C, Nabhitabhata J. Embryonic development of muddy paper nautilus, Argonauta hians Lightfoot, 1786, from Andaman Sea, Thailand. Kasetsart J (Nat Sci). 2007;41:531–8.
  135. 135. Ignatius B, Srinivasan M. Embryonic development in Octopus aegina Gray, 1849. Current Science. 2006;91(8):1089–92. pmid:CCC:000241849000025.
  136. 136. Promboon P, Nabhitabhata J, Duengdee T. Life cycle of the marbled octopus, Amphioctopus aegina (Gray) (Cephalopoda: Octopodidae) reared in the laboratory. Sci Mar. 2011;75(4):811–21. pmid:WOS:000299491700017.
  137. 137. Norman MD, Finn J, Hochberg FG. Family Octopodidae. In P. Jereb, C.F.E. Roper, M.D. Norman & J.K. Finn eds. Cephalopods of the world. An annotated and illustrated catalogue of cephalopod species known to date. Volume 3. Octopods and Vampire Squids. FAO Species Catalogue for Fishery Purposes. No. 4, Vol. 3. Rome, FAO. pp. 36–215. 2013.
  138. 138. Forsythe JW, Hanlon RT. Aspects of egg development, post-hatching behaviour, growth and reproductive biology of Octopus burryi Voss, 1950 (Mollusca: Cephalopoda). Vie Milieu. 1985;35(3/4):273–82.
  139. 139. Adam W. Céphalopodes de l'archipel du Cap-Vert, de l'Angola et du Mozambique. Trabalhos do centro de biologia piscatoria: n° 32 a 35 Memorias da Junta do Investigaçoes do Ultramar. Lisboa1962. p. 7–64.
  140. 140. Hanlon RT, Hixon RF, Forshythe JW, Hendrix JP. Cephalopods attracted to experimental night lights during a saturation dive at St. Croix, U.S., Virgin Islands. The Bulletin of the American Malacological Union. 1980;1979:53–8.
  141. 141. Segawa S, Nomoto A. Laboratory growth, feeding, oxygen consumption and ammonia excretion of Octopus ocellatus. Bulletin of Marine Science. 2002;71(2):801–13. pmid:ISI:000182787600017.
  142. 142. Kim BG, Chung EY, Jun JC, Kim CH. Spawning, hatching, survival and cannibalism of Octopus ocellatus. Korean Journal of Malacology. 2001;17:85–94.
  143. 143. Yamazaki A, Yoshida M, Uematsu K. Post-hatching development of the brain in Octopus ocellatus. Zoological Science. 2002;19:763–71. pmid:12149577
  144. 144. Nabhitabhata J. personal observation.
  145. 145. Nabhitabhata J, Nilphat P, Jaroongpattanon C, Promboon P. Culture, growth and behaviour of King Octopus, Octopus rex Nateewathana & Norman, 1999. Rayong Coastal Fisheries Research and Development Center, Bureau of Coastal Fisheries Research and Development, Department of Fisheries. 2003;Contribution No. 26:17.
  146. 146. Sreeja V, Bijukumar A, Norman MD. First report of Amphioctopus neglectus (Nateewathana & Norman, 1999) and A. rex (Nateewathana & Norman, 1999) (Mollusca: Cephalopoda) from the Indian coast. Molluscan Research. 2012;32(1):43–9. pmid:WOS:000305081200006.
  147. 147. Wood JB, Kenchington E, O'Dor RK. Reproduction and embryonic development time of Bathypolypus arcticus, a deep-sea octopod (Cephalopoda: Octopoda). Malacologia. 1998;39(1–2):11–9. pmid:ISI:000073796800002.
  148. 148. Muus B. The Bathypolypus-Benthoctopus problem of the North Atlantic (Octopodidae, Cephalopoda). Malacologia. 2002;44(2):175–222. pmid:WOS:000177725100001.
  149. 149. Mangold K. The Octopodinae from the eastern Atlantic Ocean and the Mediterranean Sea. Smithsonian Contributions to Zoology. 1998;586:521–8. pmid:ZOOR:ZOOR13500001895.
  150. 150. Young RE, Harman RF, Hochberg FG. Octopodid paralarvae from Hawaiian waters. Veliger. 1989;32(2):152–65.
  151. 151. Norman MD. Octopus ornatus Gould, 1852 (Cephalopoda, Octopodidae) in Australian waters—morphology, distribution, and life-history. Proceedings of the Biological Society of Washington. 1993;106(4):645–60. pmid:ISI:A1993MN65500003.
  152. 152. Mangold K, Boletzky SV, Frösch D. Reproductive biology and embryonic development of Eledone cirrhosa (Cephalopoda, Octopoda). Mar Biol. 1971;8:109–17.
  153. 153. Boyle PR. Eledone cirrhosa. In: Boyle PR, editor. Cephalopod life cycles Volume 1 Species accounts: Academic Press London; 1983. p. 365–86.
  154. 154. Mangold K. Eledone moschata. In: Boyle PR, editor. Cephalopod life cycles Volume 1 Species accounts: Academic Press London; 1983. p. 387–400.
  155. 155. Okubo S. Spawning and rearing of hatchlings of the Giant Pacific Octopus. Animals and Nature. 1979;9(3):2–6.
  156. 156. Lang MA, Hochberg FG, editors. Proccedings of the workshop on the fishery and market potential of octopus in California. Washington: Smithsonian Institution; 1997.
  157. 157. Hartwick B. Octopus dofleini. In: Boyle PR, editor. Cephalopod life cycles Volume 1 Species accounts: Academic Press London; 1983. p. 277–91.
  158. 158. Ortiz N, Ré MA, Márquez F. First description of eggs, hatchlings and hatchling behaviour of Enteroctopus megalocyathus (Cephalopoda: Octopodidae). Journal of Plankton Research. 2006;28:881–90.
  159. 159. Voss GL, Pearcy WG. Deep-water octopods (Mollusca; Cephalopoda) of the northeastern Pacific. Proceedings of the California Academy of Sciences. 1990;47(3):47–94. pmid:ZOOR:ZOOR13100020378.
  160. 160. Overath H, Boletzky SV. Laboratory observations on spawning and embryonic development of a blue-ringed octopus. Marine Biology. 1974;27:333–7.
  161. 161. Tranter DJ, Augustine O. Observations on the life history of the blue-ringed octopus Hapalochlaena maculosa. Mar Biol. 1973;18:115–28.
  162. 162. Batham EJ. Care of eggs by Octopus maorum. Transactions of the Royal Society of New Zealand. 1957;84:629–38.
  163. 163. Stranks TN. Biogeography of Octopus species (Cephalopoda: Octopodidae) from southeastern Australia. American Malacological Bulletin. 1996;12(1–2):145–51. pmid:ISI:A1996WD95300014.
  164. 164. Hanlon RT, Forsythe JW. Advances in the laboratory culture of octopuses for biomedical research. Laboratory Animal Science. 1985;35(1):33–40. pmid:ISI:A1985ABW3900001.
  165. 165. Ambrose RF. Observations on the embryonic development and early post-embryonic behaviour of Octopus bimaculatus (Mollusca, Cephalopoda). Veliger. 1981;24:139–46.
  166. 166. Hanlon RT. Octopus briareus. In: Boyle PR, editor. Cephalopod life cycles Volume 1 Species accounts: Academic Press London; 1983. p. 251–66.
  167. 167. Voss GL, Toll RB. The systematics and nomenclatural status of the Octopodinae described from the western Atlantic Ocean. Smithsonian Contributions to Zoology. 1998;586:457–74. pmid:ZOOR:ZOOR13500001892.
  168. 168. Rodaniche AF. Iteroparity in the Lesser Pacific Striped Octopus Octopus chierchiae (Jatta, 1889). Bulletin of Marine Science. 1984;35(1):99–104. pmid:ISI:A1984TJ10000011.
  169. 169. Caldwell RL. personal communication.
  170. 170. Van Heukelem WF. Octopus cyanea. In: Boyle PR, editor. Cephalopod life cycles Volume 1 Species accounts: Academic Press London; 1983. p. 267–76.
  171. 171. Guzik MT. Molecular phylogenetics and evolutionary history of reproductive strategies in benthic shallow-water octopuses [PhD]. TownSVille: James Cook University; 2004.
  172. 172. Alejo-Plata M, Herrera-Alejo S. First description of eggs and paralarvae of green octopus Octopus hubbsorum (Cephalopoda: Octopodidae) under laboratory conditions. Am Malacol Bull. 2014;32:132–9.
  173. 173. Domínguez-Contreras JF, Ceballos-Vázquez BP, Hochberg FG, Arellano-Martínez M. A new record in a well-established population of Octopus hubbsorum (Cephalopoda: Octopodidae) expands its known geographic distribution range and maximum size. American Malacological Bulletin. 2013;31(1):95–9.
  174. 174. Brough EJ. Egg-care, eggs and larvae in the midget octopus, Robsonella australis (Hoyle). Transactions of the Royal Society of New Zealand. 1965;6:7–19.
  175. 175. Higgins FA, Bates AE, Lamare MD. Heat tolerance, behavioural temperature selection and temperature-dependent respiration in larval Octopus huttoni. J Therm Biol. 2012;37(1):83–8. pmid:WOS:000300264600012.
  176. 176. Lenz TM, Elias NH, Leite TS, Vidal EAG. First description of the eggs and paralarvae of the tropical octopus, Octopus insularis, under culture conditions. Am Malacol Bull. 2015;33(1):101–9.
  177. 177. Vidal EAG. personal observation.
  178. 178. Leite TS. personal communication.
  179. 179. Forsythe JW, Toll RB. Clarification of the Western Atlantic ocean pygmy octopus complex: the identity and life history of Octopus joubini (Cephalopoda: Octopodinae). Bulletin of Marine Science. 1991;49(1–2):88–97. pmid:ISI:A1991HC56800008.
  180. 180. Hanlon RT. Octopus joubini. In: Boyle PR, editor. Cephalopod life cycles Volume 1 Species accounts: Academic Press London; 1983. p. 293–310.
  181. 181. Warnke K. Observations on the embryonic development of Octopus mimus (Mollusca: Cephalopoda) from Northern Chile. Veliger. 1999;42(3):211–7. pmid:ISI:000081353100002.
  182. 182. Zheng X-D, Qian Y-S, Liu C, Li Q. Octopus minor. In: Iglesias J, Fuentes L, Villanueva R, editors. Cephalopod Culture. New York: Springer; 2014. p. 415–26.
  183. 183. Cheng RB, Zheng XD, Lin XZ, Yang JM, Li Q. Determination of the complete mitochondrial DNA sequence of Octopus minor. Molecular Biology Reports. 2012;39(4):3461–70. pmid:WOS:000301108500011.
  184. 184. Ylitalo HA, Watling L, Toonen RJ. First description of hatchlings and eggs of Octopus oliveri (Berry, 1914) (Cephalopoda: Octopodidae). Molluscan Research. 2014;34(2):79–83.
  185. 185. Kaneko N, Kubodera T, Iguchi A. Taxonomic study of shallow-water octopuses (Cephalopoda: Octopodidae) in Japan and adjacent waters using mitochondrial genes with perspectives on Octopus DNA barcoding. Malacologia. 2011;54(1–2):97–108. pmid:WOS:000296211300004.
  186. 186. Bozzano A, Pankhurst PM, Moltschaniwskyj NA, Villanueva R. Eye development in southern calamary, Sepioteuthis australis, embryos and hatchlings. Marine Biology. 2009;156(7):1359–73. pmid:ISI:000266010300001.
  187. 187. Mangold-Wirz K, Boletzky SV, Mesnil B. Biologie de reproduction et distribution d'Octopus salutti Verany (Cephalopoda, Octopoda). Rapport de la Commission Internationale pour l'Exploration Scientifique de la Mer Méditerranée. 1976;23(8):83–97.
  188. 188. Collins MA, Yau C, Allcock L, Thurston MH. Distribution of deep-water benthic and bentho-pelagic cephalopods from the north-east Atlantic. Journal of the Marine Biological Association of the United Kingdom. 2001;81(1):105–17. pmid:ISI:000168203900012.
  189. 189. Iribarne OO. Life history and distribution of the small south-western Atlantic octopus, Octopus tehuelchus. Journal of Zoology. 1991;223:549–65. pmid:ISI:A1991FL02700002.
  190. 190. Haimovici M, Perez JAA. Coastal cephalopod fauna of Southern Brazil. Bulletin of Marine Science. 1991;49(1–2):221–30. pmid:ISI:A1991HC56800018.
  191. 191. Joll LM. Mating, egg-laying and hatching of Octopus tetricus (Mollusca: Cephalopoda) in the laboratory. Mar Biol. 1976;36:327–33.
  192. 192. Ramos JE, Pecl GT, Moltschaniwskyj NA, Strugnell JM, León RI, Semmens JM. Body size, growth and life span: implications for the polewards range shift of Octopus tetricus in south-eastern Australia. PLOS ONE. 2014;9(8):e103480. pmid:25090250
  193. 193. Villanueva R. Experimental rearing and growth of planktonic Octopus vulgaris from hatching to settlement. Canadian Journal of Fisheries and Aquatic Sciences. 1995;52(12):2639–50. pmid:ISI:A1995UA09600011.
  194. 194. Carrasco JF, Arronte JC, Rodríguez C. Paralarval rearing of the common octopus, Octopus vulgaris (Cuvier). Aquac Res. 2006;37:1601–5.
  195. 195. Vidal ÉAG, Fuentes L, da Silva LB. Defining Octopus vulgaris populations: a comparative study of the morphology and chromatophore pattern of paralarvae from Northeastern and Southwestern Atlantic. Fisheries Research. 2010;106(2):199–208.
  196. 196. Itami K, Izawa Y, Maeda S, Nakai K. Notes on the laboratory culture of octopus larvae. Bulletin of the Japanese Society of Scientific Fisheries. 1963; 29:514–20.
  197. 197. Ito H. Temperature and salinity tolerances of larvae of Paroctopus conispadiceus (Mollusca: Cephalopoda). Bulletin of the Hokkaido Regional Fisheries Research Laboratory. 1985;50:99–115.
  198. 198. Okutani T, Tagawa M, Horikawa H. Cephalopods from continental shelf and slope around Japan. The intensive research of unexploited fishery resources of continental slopes. Tokyo: Japan Fisheries Resources Conservation Association; 1987.
  199. 199. DeRusha RH, Forsythe JW, Hanlon RT. Laboratory growth, reproduction and life span of the Pacific pygmy octopus, Octopus digueti. Pacific Science. 1987;41(1–4):104–21. pmid:ZOOR:ZOOR12600013438.
  200. 200. Norman MD. Cephalopods: a World Guide. Hackenheim: ConchBooks; 2000. 320 p.
  201. 201. González ML, Arriagada SE, López DA, Pérez MC. Reproductive aspects, eggs and paralarvae of Robsonella fontanianus (d'Orbigny, 1834). Aquac Res. 2008;39:1569–73.
  202. 202. Uriarte I, Hernandez J, Dorner J, Paschke K, Farias A, Crovetto E, et al. Rearing and growth of the octopus Robsonella fontaniana (Cephalopoda: Octopodidae) from planktonic hatchlings to benthic juveniles. Biol Bull. 2010;218(2):200–10. pmid:20413796
  203. 203. Ortiz N, Ré ME. The eggs and hatchlings of the octopus Robsonella fontaniana (Cephalopoda: Octopodidae). Journal of the Marine Biological Association of the United Kingdom. 2011;91(03):705–13.
  204. 204. Ortiz N, Ré ME, Márquez F, Glembocki NG. The reproductive cycle of the red octopus Enteroctopus megalocyathus in fishing areas of Northern Patagonian coast. Fisheries Research. 2011;110(1):217–23.
  205. 205. Sánchez P, Alvarez JA. Scaeurgus unicirrhus (Orbigny, 1840) (Cephalopoda, Octopodidae)—first record from the Southeast Atlantic. South African Journal of Marine Science-Suid-Afrikaanse Tydskrif Vir Seewetenskap. 1988;7:69–74. pmid:ISI:A1988T203700008.
  206. 206. Miske V, Kirchhauser J. First record of brooding and early life cycle stages in Wunderpus photogenicus Hochberg, Norman and Finn, 2006 (Cephalopoda: Octopodidae). Molluscan Research. 2006;26(3):169–71. pmid:CCC:000243238600007.
  207. 207. Hochberg FG, Norman MD, Finn J. Wunderpus photogenicus n. gen. and sp., a new octopus from the shallow waters of the Indo-Malayan Archipelago (Cephalopoda: Octopodidae). Molluscan Research. 2006;26(3):128–40. pmid:CCC:000243238600005.
  208. 208. Vidal EAG, DiMarco FP, Wormuth JH, Lee PG. Optimizing rearing conditions of hatchling loliginid squid. Marine Biology. 2002;140:117–27.
  209. 209. Sugimoto C, Ikeda Y. Ontogeny of schooling behavior in the oval squid Sepioteuthis lessoniana. Fisheries Science. 2012;78(2):287–94. pmid:WOS:000303878600009.
  210. 210. Nabhitabhata J. Life cycle of cultured big fin squid, Sepioteuthis lessoniana Lesson. Phuket Marine Biological Center Special Publication. 1996;16:83–95.
  211. 211. Okubo S. Culture of the larvae and juveniles of Octopus dofleini in an aquarium for one year and two months. Shima Marineland Quarterly. 1980;25:4–5.
  212. 212. Snyder S. Laboratory culture of Octopus dofleini from hatching to settlement. Am Malacol Bull. 1986;4(2):241.
  213. 213. Snyder S. Successful rearing of Octopus dofleini from hatchling to settlement. American Association of Zoological Parks and Aquariums 1986 Annual Conference Proceedings. 1986:pp. 781–3.
  214. 214. Uriarte I, Farías A. Enteroctopus megalocyathus. In: Iglesias J, Fuentes L, Villanueva R, editors. Cephalopod Culture. New York: Springer; 2014. p. 365–82.
  215. 215. Iglesias J, Otero JJ, Moxica C, Fuentes L, Sánchez FJ. The completed life cycle of the octopus (Octopus vulgaris, Cuvier) under culture conditions: paralarval rearing using Artemia and zoeae, and first data on juvenile growth up to 8 months of age. Aquacult Int. 2004;12:481–7.
  216. 216. Hoving H- JT, Perez JAA, Bolstad KSR, Braid HE, Evans AB, Fuchs D, et al. The study of deep-sea cephalopods. In: Erica AGV, editor. Advances in Marine Biology. Volume 67: Academic Press; 2014. p. 235–359. pmid:24880796
  217. 217. Iglesias J, Fuentes L, Villanueva R, editors. Cephalopod Culture. New York Heidelberg Dordrecht London: Springer; 2014.
  218. 218. Vidal EAG, Boletzky SV. Loligo vulgaris and Doryteuthis opalescens. In: Iglesias J, Fuentes L, Villanueva R, editors. Cephalopod Culture. New York: Springer; 2014. p. 271–313.
  219. 219. Villanueva R, Staaf DJ, Argüelles J, Bozzano A, Camarillo-Coop S, Nigmatullin CM, et al. A laboratory guide to in vitro fertilization of oceanic squids. Aquaculture. 2012;342–343(0):125–33.
  220. 220. Veness C. Calculate distance, bearing and more between Latitude/Longitude points: Movable Type Scripts; 2002–2016. Available: Accessed 2016.
  221. 221. Scheltema RS. Planktonic and non-planktonic development among prosobranch gastropods and its relationship to the geographic range of species. In: Ryland JS, Tyler PA, editors. Reproduction, genetics and distributions of marine organisms. Fredensborg, Denmark,: Olsen and Olsen; 1989. p. 183–8.
  222. 222. Lester SE, Ruttenberg BI. The relationship between pelagic larval duration and range size in tropical reef fishes: a synthetic analysis. Proceedings of the Royal Society of London B: Biological Sciences. 2005;272:585–91.
  223. 223. Markaida UR, C J.J.; Gilly W.F. Tagging studies on the jumbo squid (Dosidicus gigas) in the Gulf of California, Mexico. Fishery Bulletin. 2005;103:219–26.
  224. 224. Marshall DJ, Keough MJ. The evolutionary ecology of offspring size in marine invertebrates. Advances in Marine Biology. 2007;53:1–60. pmid:17936135
  225. 225. Bouchaud O. Energy consumption of the cuttlefish Sepia officinalis L (Mollusca, Cephalopoda) during embryonic development, preliminary results. Bulletin of Marine Science. 1991;49(1–2):333–40. pmid:ISI:A1991HC56800030.
  226. 226. Gowland FC, Boyle PR, Noble LR. Morphological variation provides a method of estimating thermal niche in hatchlings of the squid Loligo forbesi (Mollusca: Cephalopoda). Journal of Zoology. 2002;258:505–13. pmid:ISI:000180485900011.
  227. 227. Ceriola L, Jackson GD. Growth, hatch size and maturation in a southern population of the loliginid squid Loliolus noctiluca. Journal of the Marine Biological Association of the UK. 2010;90(04):755–67.
  228. 228. Ikeda Y, Wada Y, Arai N, Sakamoto W. Note on size variation of body and statoliths in the oval squid Sepioteuthis lessoniana hatchlings. Journal of Marine Biological Association of United Kingdom. 1999;79:757–9.
  229. 229. Sakaguchi H, Hamano T, Nakazono A. Growth of Octopus vulgaris in the Northeastern Iyo-Nada of the Seto Inland Sea, Japan. Bull Jap Soc Fish Oceanogr. 2002;66(1):11–5.
  230. 230. Farias A, Navarro JC, Cerna V, Pino S, Uriarte I. Effect of broodstock diet on the fecundity and biochemical composition of eggs of the Patagonian red octopus (Enteroctopus megalocyathus Gould 1852). Ciencias Marinas. 2011;37(1):11–21. pmid:ISI:000288639500002.
  231. 231. Márquez L, Quintana D, Lorenzo A, Almansa E. Biometrical relationships in developing eggs and neonates of Octopus vulgaris in relation to parental diet. Helgol Mar Res. 2013;67(3):461–70.
  232. 232. Caamal-Monsreal C, Mascaró M, Gallardo P, Rodríguez S, Noreña-Barroso E, Domingues P, et al. Effects of maternal diet on reproductive performance of O. maya and its consequences on biochemical characteristics of the yolk, morphology of embryos and hatchling quality. Aquaculture. 2015;441(0):84–94.
  233. 233. Quintana D, Márquez L, Arévalo JR, Lorenzo A, Almansa E. Relationships between spawn quality and biochemical composition of eggs and hatchlings of Octopus vulgaris under different parental diets. Aquaculture. 2015;446(0):206–16.
  234. 234. Boletzky SV. Embryonic phase. In: Boyle PR, editor. Cephalopod life cycles Volume 2 Comparative reviews: Academic Press London; 1987. p. 5–31.
  235. 235. Pecl GT, Steer MA, Hodgson KE. The role of hatchling size in generating the intrinsic size-at-age variability of cephalopods: extending the Forsythe Hypothesis. Marine and Freswater Research. 2004;55:387–94.
  236. 236. Macpherson E, Raventós N. Relationship between pelagic larval duration and geographic distribution of Mediterranean littoral fishes. Marine Ecology Progress Series. 2006;327:257–65.
  237. 237. Levin LA, Bridges TS. Pattern and diversity in reproduction and development. In: Mc Edward L, editor. Ecology of marine invertebrate larvae. Boca Raton, FL: CRC Press; 1995. p. 1–48.
  238. 238. Palumbi SR. Marine speciation on a small planet. Trends in Ecology & Evolution. 1992;7(4):114–8.
  239. 239. Palumbi SR. Using genetics as an indirect estimator of larval dispersal. In: Mc Edward L, editor. Ecology of marine invertebrate larvae. Boca Raton, FL: CRC Press; 1995. p. 369–87.
  240. 240. Allcock AL, Brierley AS, Thorpe JP, Rodhouse P. Restricted gene flow and evolutionary divergence between geographically separated populations of the Antarctic octopus Pareledone turqueti. Mar Biol. 1997;129(1):97–102.
  241. 241. Higgins KL, Semmens JM, Doubleday ZA, CP B. Comparison of population structuring in sympatric octopus species with and without a pelagic larval stage. Marine Ecology Progress Series. 2013;486:203–12.
  242. 242. Moreira AA, Tomás ARG, Hilsdorf AWS. Evidence for genetic differentiation of Octopus vulgaris (Mollusca, Cephalopoda) fishery populations from the southern coast of Brazil as revealed by microsatellites. Journal of Experimental Marine Biology and Ecology. 2011;407(1):34–40.
  243. 243. Van Der Vyver JSF, Sauer WHH, McKeown NJ, Yemane D, Shaw PW, Lipinski MR. Phenotypic divergence despite high gene flow in chokka squid Loligo reynaudii (Cephalopoda: Loliginidae): implications for fishery management. Journal of the Marine Biological Association of the United Kingdom. 2015;FirstView:1–19.
  244. 244. Strathmann RR, Strathmann MF. The relationship between adult size and brooding in marine invertebrates. American Naturalist. 1982;119:91–101.
  245. 245. Bridges TS. Reproductive investment in four developmental morphs of Streblospio (Polychaeta: Spionidae). Biological Bulletin. 1993;184:144–52.
  246. 246. Robison B, Seibel B, Drazen J. Deep-sea octopus (Graneledone boreopacifica) conducts the longest-known egg-brooding period of any animal. PLOS ONE. 2014;9(7):e103437. pmid:25075745
  247. 247. Strugnell JM, Cherel Y, Cooke IR, Gleadall IG, Hochberg FG, Ibáñez CM, et al. The Southern Ocean: source and sink? Deep Sea Research Part II: Topical Studies in Oceanography. 2011;58(1–2):196–204.
  248. 248. Ibáñez CM, Peña F, Pardo-Gandarillas MC, Méndez MA, Hernández CE, Poulin E. Evolution of development type in benthic octopuses: holobenthic or pelago-benthic ancestor? Hydrobiologia. 2014;725(1):205–14.
  249. 249. Voss GL. Evolution and phylogenetic relationships of deep-sea octopods (Cirrata and Incirrata). In: Clark MR, Trueman ER, editors. The Mollusca. 12. San Diego: Academic Press 1988. p. 253–76.
  250. 250. Barratt IM, Johnson MP, Allcock AL. Fecundity and reproductive strategies in deep-sea incirrate octopuses (Cephalopoda: Octopoda). Mar Biol. 2007;150(3):387–98. pmid:CCC:000242830200007.
  251. 251. Laptikhovsky V. Reproductive strategy of deep-sea and Antarctic octopods of the genera Graneledone, Adelieledone and Muusoctopus (Mollusca: Cephalopoda). Aquatic Biology. 2013;18(1):21–9.
  252. 252. Collins MA, Villanueva R. Taxonomy, ecology and behaviour of the cirrate octopods. Oceanography and Marine Biology: Ann Annual Review. 2006;44:277–322. pmid:ISI:000243635700006.
  253. 253. Vecchione M, Young RE. Aspects of the functional morphology of cirrate octopods: locomotion and feeding. Vie et Milieu. 1997;47(2). pmid:ZOOR:ZOOR13400029012.
  254. 254. Villanueva R, Segonzac M, Guerra A. Locomotion modes of deep-sea cirrate octopods (Cephalopoda) based on observations from video recordings on the Mid-Atlantic Ridge. Mar Biol. 1997;129(1):113–22. pmid:ISI:A1997XQ30000014.
  255. 255. Johnsen S, Balser EJ, Fisher EC, Widder EA. Bioluminescence in the deep-sea cirrate octopod Stauroteuthis syrtensis Verrill (Mollusca: Cephalopoda). Biological Bulletin. 1999;197(1):26–39. pmid:ISI:000082403400005.
  256. 256. Boletzky SV. On eggs and embryos of cirromorph octopods. Malacologia. 1982;22(1–2):197–204.
  257. 257. Okubo S, Tsujii T, Watabe N, Williams DF. Hatching of Nautilus belauensis Saunders, 1981, in captivity—culture, growth and stable-isotope compositions of shells, and histology and immunohistochemistry of the mantle epithelium of the juveniles. Veliger. 1995;38:192–202.