Hostname: page-component-745bb68f8f-grxwn Total loading time: 0 Render date: 2025-02-06T08:35:30.506Z Has data issue: false hasContentIssue false
  • English
  • Français

Influence of temperature on the larval development of the edible crab, Cancer pagurus

Published online by Cambridge University Press:  20 January 2009

Monika Weiss ,
Sven Thatje ,
Olaf Heilmayer ,
Klaus Anger ,
Thomas Brey  and
Martina Keller
Monika Weiss
Affiliation:
Alfred-Wegener-Institut für Polar-und Meeresforschung, Am Handelshafen 12, 27570 Bremerhaven, Germany
Sven Thatje*
Affiliation:
National Oceanography Centre, Southampton, School of Ocean and Earth Science, University of Southampton, European Way, SO14 3ZH Southampton, United Kingdom
Olaf Heilmayer
Affiliation:
Alfred-Wegener-Institut für Polar-und Meeresforschung, Am Handelshafen 12, 27570 Bremerhaven, Germany National Oceanography Centre, Southampton, School of Ocean and Earth Science, University of Southampton, European Way, SO14 3ZH Southampton, United Kingdom
Klaus Anger
Affiliation:
Biologische Anstalt Helgoland, Stiftung Alfred-Wegener-Institut für Polar- und Meeresforschung, 27498 Helgoland, Germany
Thomas Brey
Affiliation:
Alfred-Wegener-Institut für Polar-und Meeresforschung, Am Handelshafen 12, 27570 Bremerhaven, Germany
Martina Keller
Affiliation:
Alfred-Wegener-Institut für Polar-und Meeresforschung, Am Handelshafen 12, 27570 Bremerhaven, Germany
*
Correspondence should be addressed to: S. Thatje, National Oceanography Centre, Southampton, School of Ocean and Earth Science, University of Southampton, European Way, SO14 3ZH Southampton, United Kingdom email: svth@noc.soton.ac.uk
Rights & Permissions [Opens in a new window]

Abstract

The influence of temperature on larval survival and development was studied in the edible crab, Cancer pagurus, from a population off the island of Helgoland, North Sea. In rearing experiments conducted at six different temperatures (6°, 10°, 14°, 15°, 18° and 24°C), zoeal development was only completed at 14° and 15°C. Instar duration of the Zoea I was negatively correlated with temperature. A model relating larval body mass to temperature and developmental time suggests that successful larval development is possible within a narrow temperature range (14° ± 3°C) only. This temperature optimum coincides with the highest citrate synthase activity found at 14°C. A comparison for intraspecific variability among freshly hatched zoeae from different females (CW 13–17 cm, N = 8) revealed that both body mass and elemental composition varied significantly. Initial larval dry weight ranged from 12.1 to 17.9 μg/individual, the carbon content from 4.6 to 5.8 μg/individual, nitrogen from 1.1 to 1.3 μg/individual, and the C:N ratio from 4.1 to 4.4. A narrow larval temperature tolerance range of C. pagurus as well as the indication of intraspecific variability in female energy allocation into eggs may indicate a potential vulnerability of this species to climate change. Large-scale studies on the ecological and physiological resilience potential of this commercially fished predator are needed.

Keywords

Brachyuraearly ontogenyedible crabelemental composition
Type
Research Article
Information
Journal of the Marine Biological Association of the United Kingdom , Volume 89 , Issue 4 , June 2009 , pp. 753 - 759
Copyright
Copyright © Marine Biological Association of the United Kingdom 2009

INTRODUCTION

The embryonic and larval development of marine invertebrates is affected by extrinsic and intrinsic factors, such as temperature, maternal energy provisioning (Ouellet & Plante, Reference Ouellet and Plante2004) and both pre- and post-hatching environmental conditions (Torres & Escribano, Reference Torres and Escribano2003; Giménez et al., Reference Giménez, Anger and Torres2004; Fischer & Thatje, Reference Fischer and Thatje2008). Variation in environmental key factors such as food availability (Anger & Dawirs, Reference Anger and Dawirs1982), quality (Harms et al., Reference Harms, Anger, Klaus and Seeger1991), salinity (Giménez & Torres, Reference Giménez and Torres2002; Giménez & Anger, Reference Giménez and Anger2003) or temperature (Dawirs, Reference Dawirs1979, Reference Dawirs1985) can have unfavourable effects on growth in the early ontogeny of decapod crustaceans (for review, see Anger, Reference Anger2001).

The early life cycle of the edible crab, Cancer pagurus, consists of five planktotrophic zoeal stages and a megalopa before reaching the first crab stage (Ingle, Reference Ingle1981). This species has a broad geographical distribution from northern Norway to West Africa and is also abundant in the Mediterranean Sea, which implies a wide range of temperature tolerance in adult populations of the species. Studies of abundance and catch rates show that C. pagurus is expanding its biogeographical range further northwards (Woll et al., Reference Woll, van der Meeren and Fossen2006).

Since 1962 mean annual sea-surface temperature in the North Sea around the island of Helgoland rose 1.1°C, with milder winters and rising summer maxima (Wiltshire & Manly, Reference Wiltshire and Manly2004). Such a shift in ecological conditions may cause changes in the metabolic efficiency or fitness of an organism (Pörtner, Reference Pörtner2001; Heilmayer et al., Reference Heilmayer, Brey and Pörtner2004), which presumably is reflected in its elemental and biochemical composition (Dahlhoff, Reference Dahlhoff2004). Studies on the complete temperature tolerance window of invertebrate larvae are extremely scarce, but may be a clue in future assessments of the potential of species to cope with climate change (e.g. Anger, Reference Anger2001; Pörtner et al., Reference Pörtner, Berdal, Blust, Brix, Colosimo, Wachter, Giuliani, Johansen, Fischer, Knust, Lannig, Naevdal, Nedenes, Nyhammer, Sartoris, Serendero, Sirabella, Thorkildsen and Zakhartsev2001, Reference Pörtner, Storch and Heilmayer2005; Thatje et al., Reference Thatje, Anger, Calcagno, Lovrich, Pörtner and Arntz2005).

In the present study, we provide evidence for temperature-induced changes in the chemical composition and aerobic capacities of C. pagurus larvae. Based on the hypothesis that the early ontogeny is the most vulnerable part of a life cycle (Anger, Reference Anger2001), we discuss the physiological capability of the species to cope with elevated temperatures.

MATERIALS AND METHODS

Sampling and maintenance of adults and larvae

Ovigerous Cancer pagurus (carapace width (CW) 125 to 171 mm) were caught in May 2005 near the island of Helgoland in the North Sea (54° 11′N 7° 53′E) using a bottom trawl. Animals were immediately transported to the laboratory of the Marine Biological Station Helgoland (Biologische Anstalt Helgoland (BAH)) where they were maintained individually in flow-through seawater aquaria (15–20 l) at sea-surface temperature (15.2–17.2°C) and salinity (~32 psu) in a 12:12-hour light/dark cycle. Adults were fed twice a week either with isopods (Idotea sp.) or pieces of mussel (Mytilus edulis) meat. One day after feeding, remains were removed from the aquaria to maintain good water quality.

Freshly hatched larvae were collected in filters receiving water from the overflow of the aquaria. Since most larvae hatched at night, samples were taken every morning. Filters were cleaned every evening to ensure daily larval age did not vary by more than 12 hours (Lovrich et al., Reference Lovrich, Thatje, Calcagno, Anger and Kaffenberger2003). Solely actively moving larvae were used for experiments.

Influence of temperature on larval development

Randomly selected larvae from one randomly selected female (A) were kept in 500 ml glass bowls with a density of 20 to 30 individuals per bowl. In daily intervals, water was changed; larvae checked for moults or mortality and subsequently were fed with freshly hatched Artemia spp. nauplii. One hatch (female A; Table 1) was divided on the day of hatching and subsequently reared at five constant temperatures (6, 10, 14, 18 and 24°C). Larvae reared at 15°C resulted from a female caught in 1985 and were maintained and reared under the same condition as outlined above.

Table 1. Cancer pagurus. Changes in dry weight (W), carbon (C), nitrogen (N) and C:N ratio during time days after hatch (±SD) at five temperatures. Larvae reared at 15°C are from different female.

Minimum time of development for each instar was recorded assuming optimal developmental conditions in larvae (see Figure 1). Samples for determinations of larval dry weight (W) and elemental composition were taken immediately after hatching and later in intervals of one to ten days (see Table 1). Five replicates were collected, or fewer, when too few larvae were available. Each replicate consisted of 20–25 individuals in the Zoea I (Z I), but fewer (see Table 1) in the following (larger) instars.

Fig. 1. Model of body mass increase (C) throughout the larvae development of the edible crab Cancer pagurus. Equation for the model is: C BC =−7.97−10.755 × t + 919.52 × f(T) + 476.634 × t × f(T); N = 463, F = 1599.51, R2 = 0.954; T represents the temperature in Kelvin and t the time in days. Isolines represent carbon content in µg. ZI–ZV are the larval instars. White areas show the realistic range of the model.

Elemental analyses (CHN)

Carbon (C), hydrogen (H) and nitrogen (N) contents were determined following Anger & Dawirs (Reference Anger and Dawirs1982); in brief: larvae were gently rinsed in distilled water, blotted on filter paper, placed into tin cartridges, vacuum-dried for 48 hours at <0.01 mbar in a Lyovac GT 2E (Leybold-Heraeus) apparatus, weighed to the nearest 0.1 µg on a Mettler UM3 microbalance, and stored frozen at –20°C. CHN content was measured with a Fisons (Carlo Erba Science) Model 1108 Elemental Analyzer.

In order to study intraspecific variability in the maternal energy provisioning of offspring, we sampled freshly hatched larvae from eight ovigerous females (always after the first night of the hatching period; female CW = 13–17 cm; Table 2) and compared their initial body mass (measured as dry weight, W) and elemental composition (carbon, hydrogen and nitrogen; collectively CHN).

Table 2. Cancer pagurus. Size of eight different females and elemental composition (dry weight (W), carbon (C), nitrogen (N), hydrogen (H), C:N and C:H ratio of freshly hatched larvae (hatch 1)).

Enzyme assay

Citrate synthase (CS) (E.C. 4.1.3.7) is a key regulatory enzyme in the tricarbonic acid (TCA) cycle and was chosen as an indicator of aerobic capacity. Investigations on metabolic enzymes in larval stages are scarce and studies on shrimp larvae show that CS activity is dependent on growth during ontogeny (Lemos et al., Reference Lemos, Salomon, Gomes, Phan and Buchholz2003). Samples for determinations of enzyme activity of CS were taken in the premoult period of the Zoea I instar of larvae reared at three temperatures (10, 14 and 18°C) and analysed following a modified method of Sidell et al. (Reference Sidell, Driedzic, Stowe and Johnston1987). Three replicates were collected, each consisting of 10 larvae. No samples were taken for 6° and 24°C as an insufficient amount of material was available.

Frozen samples were homogenized in ~0.3 µl extraction buffer (75 mM Tris–HCl, 1 mM EDTA; pH 7.6) per 1µg larval W (dry weight) to get a 1:10 (w/v) ratio with a Branson Sonifier 450 (0°C, output control 8, duty cycle 50%, 15 minutes). Homogenates were centrifuged for 5 minutes at 7400 g and 0°C with an Eppendorf Centrifuge 5810R. The concentration of soluble protein in the extracts was measured after Bradford (Reference Bradford1976). The samples were first diluted 1:5 with 0.9% NaCl and were then applied in duplicate (5 µl) on microplates. Subsequently 250 µl dye reagent (Biorad protein assay 500 0006, diluted 1:5 with aqua dest) were added and the optical density was measured at 620 nm in a microplate reader (FLUOstar Galaxy). Bovine serum albumin (BSA, 0–3.5 µg per well) was run parallel as standard.

For enzyme assays the absorption of the supernatant was measured at four temperatures (10, 14, 18 and 24°C) in three aliquots with a microplate reader at 405 nm. Homogenates (2 µl/well) were assayed in 150 µl of 100 mM Tris–HCl buffer (pH 8.0), 10 µl 5 mM DTNB (5.5′-Dithio-bis-(2-nitrobenzoic acid)) and 4 µl Acetyl-CoA (20 mM). 4 µl Oxalacetat (20 mM) was added to start the reaction (omitted for the blanks). Standards of 0.5 mM Dithiothreitol DTT (5–40 µl per well) were run in parallel. The activity was expressed as the change of absorption per time and protein weight units (ΔA mingprt−1−1).

Statistical analyses

All data were tested with the Nalimov test to exclude outliers from analysis (Kaiser & Gottschalk, Reference Kaiser and Gottschalk1972). A general additive model (Hastie & Tibshirani, Reference Hastie and Tibshirani1990) was used to describe larval mass (µg C) as a function of time (t, days) and temperature (T, Kelvin):

C_{BC} = a+b_1 \times t + b_2 \times f\lpar T\rpar + b_3 \times t \times f\lpar T\rpar \quad\lsqb \mu g\comma \; d\comma \; K \rsqb

where CBC is the Box–Cox transformed larval mass (Sokal & Rholf, Reference Sokal and Rohlf1981) and f(T) a function that models the temperature effect according to a normal distribution with mean MT, standard deviation SDT and skewing factor SKT. The latter was introduced to allow for asymmetric effects of temperatures above and below the optimum, as observed in many temperature tolerance studies (see e.g. Pörtner et al., Reference Pörtner, Berdal, Blust, Brix, Colosimo, Wachter, Giuliani, Johansen, Fischer, Knust, Lannig, Naevdal, Nedenes, Nyhammer, Sartoris, Serendero, Sirabella, Thorkildsen and Zakhartsev2001, Reference Pörtner, Storch and Heilmayer2005) and implied by the original data:

f\lpar t\rpar = \left(1/\left(SD_T \times \sqrt{2\pi}\right)\right)\times e^{-0.5 \times \lpar \lpar \lpar T - M_T\rpar + SK_T \times \lpar T - M_T\rpar \rpar /SD_T\rpar ^2} \quad \hbox{for T} \gt = \hbox{M}_{\rm T}
{f\lpar t\rpar = \left(1/\left(SD_T \times \sqrt{2\pi}\right)\right)\times e^{-0.5 \times \lpar \lpar \lpar T - M_T\rpar - SK_T \times \lpar T - M_T\rpar \rpar /SD_T\rpar ^2} \quad \hbox{for T} \lt \hbox{M}_{\rm T}}

For the comparison of elemental composition of larvae of different females and temperature dependence of CS activity a one-way ANOVA was used. Post-hoc tests were conducted with the Student–Newman–Keuls method. C/N ratio data were transformed logarithmically prior to analysis in order to achieve a normally distributed data set.

RESULTS

Influence of temperature on larval development

Complete zoeal development occurred only at 14 and 15°C (Table 1; Figure 1). At 6°C, the Zoea I survived for up to 25 days without moulting to the Zoea II stage. At 10°C, first larvae moulted after 12 days to the Zoea II and survived only for another three days. Larvae reared at 14°C reached the Zoea V 33 days after hatching. At 18°C, larvae died already after 6 days in the Zoea II stage, and at 24°C they reached the Zoea III, dying ten days later.

The minimum duration of development through the ZI stage decreased with increasing temperature. This pattern can be described as a linear relationship between ln temperature and ln instar duration with the equation:

\ln \hbox{D} = \ln 6.5482 - 1.8096 \times \ln T\semicolon \; \hbox{R}^{2} = 0.991\comma \; P \lt 0.001

where D = time of development (days) and T = temperature (°C). Changes in W and CHN during the course of larval development are shown in Table 1.

Larval body mass (C in µg/ind) is predicted from day (t) and temperature (T in K) by the model

C_{BC} = - 7.97 - 10.755 \times t + 919.52 \times f \lpar T\rpar + 476.634 \times t \times f\lpar T\rpar

N = 463, F = 1599.51, R2 = 0.954, P < 0.001 for the whole model and each term with

{\eqalign{f\lpar T\rpar =& \ 0.02452 \times e^{-0.5 \times \lpar \lpar \lpar \lpar T - 295.243\rpar + \lpar T - 295.243\rpar + 0.620 \times \lpar T - 295.243\rpar \rpar \div 16.273\rpar ^2\rpar } \hbox{for T} \gt = 295.243}}
{\eqalign{f\lpar T\rpar =&\, 0.02452 \times e^{-0.5 \times \lpar \lpar \lpar \lpar T - 295.243\rpar + \lpar T - 295.243\rpar - 0.620 \times \lpar T - 295.243\rpar \rpar \div 16.273\rpar ^2\rpar } \hbox{for T} \lt 295.243}}

using

C_{BC} = \lpar C^{0.2} - 1\rpar /0.0283318325559

this solves to

C = \lpar 0.779 - 0.305 \times t + 26.0517 \times f\lpar T\rpar + 13.504 \times t \times f\lpar T\rpar \rpar ^5

Note that this model predicts a larval mass for just any combination of time and temperature whereas our experiments indicate that the time × temperature space where larvae do exist is limited (see discussion). Figure 1 provides a plot of larval mass in the time × temperature space, where the ‘unrealistic’ range has been shadowed. The residual plot (Figure 2) indicates that the model fit the data quite well up to about 40 µg C, but severely underestimates larval mass >50 µg C. At the upper threshold temperatures in higher instars, changes in carbon values were generally lower than predicted by the model (see Figure 2). Further increase or decrease in temperature cause death of the larvae. To give an example: at 6°C larvae did not moult and die after 25 days; at 24°C, carbon values in the Zoea III are much lower than they were predicted by the model (also compare Table 1), e.g. 20°C, and the larvae died before reaching the Zoea IV. According to the model, the final Zoea V body mass of 60 µg C observed at 16°C would be obtained after 36 days at 18°C, which already lies outside the realistic assumption, and would be unattainable at temperatures <= 10°C.

Fig. 2. Residual by predicted plot from the model predicting larval growth: plot of regression residuals versus predicted values. The plot does not indicate a problem with the model.

Citrate synthase activity (CSA)

Activities of citrate synthase (CS) in C. pagurus larvae measured at four different temperatures are shown in Figure 3. Larvae of all acclimation temperatures show lowest activity levels at 10°C assay temperature. 14°C acclimated larvae show the highest activity at all assay temperatures. 18°C acclimated larvae show no significant differences over the whole measured temperature range. Significantly higher CSA rates compared to larvae at hatching are only observed at 14°C assay temperature.

Fig. 3. Citrate synthase activity (U * g Prot−1) of Cancer pagurus larvae acclimated to three different temperatures (light grey bars—10°C acclimated; white bars—14°C acclimated; dark grey bars—18°C acclimated) compared with larvae immediately after hatching (black bars). Bars within an assay temperature not connected with the same letter are significantly different (P > 0.05).

Variability in initial larval body mass among broods

Female carapace width (CW), larval body weight (W) and elemental composition (CHN) are compared in Table 2. Initial body mass and elemental composition of freshly hatched Zoea I larvae varied significantly among the eight females (Table 3) without showing a clear pattern between larval energy provisioning and female size. For example, females B and D had the same CW, but larval W and C, H, and N contents differed significantly.

Table 3. Cancer pagurus. One-way ANOVA to evaluate the maternal influence on body mass (as dry weight (W), carbon (C), and nitrogen (N)]) and C:N ratio of freshly hatched larvae (first day of the hatching period 1). *, ANOVA on ranks because equal variance test failed.

DISCUSSION

Larval development of C. pagurus of the Helgoland population up to Zoea V is achieved within a narrow temperature window (15–16°C) only. Below and above this window we see initial development that completes stages Zoea I to Zoea III, depending on temperature. Zoea I development times indicate a positive temperature effect on larval growth up to 24°C, too. Our predictive model captures this feature quite well, indicating that optimum temperature should be around 22°C. Hence, what causes the failure of larval development outside the 15–16°C window? Obviously, this is not a straightforward temperature effect on metabolism, e.g. through cellular oxygen deficiency beyond Pejus temperatures as postulated by Pörtner (Reference Pörtner2001). Additionally, reduced feeding activity at cold temperatures can result in insufficient ingestion rates, while at high temperatures higher maintenance costs cannot be compensated (Dawirs & Dietrich, Reference Dawirs and Dietrich1986; Anger et al., Reference Anger, Lovrich, Thatje and Calcagno2004; Heilmayer et al., Reference Heilmayer, Thatje, McClelland, Conlan and Brey2008). We hypothesize that the transition from one stage to the next is the phase of failure. Either, energy investment and thus oxygen demand is distinctly enhanced during transition from one stage to the next, and/or the complex metamorphotic process by itself is more temperature sensitive, as it has long been known that ecdysis is the critical point in the development of decapod larvae (Anger, Reference Anger2001).

The residual plot (Figure 2) shows a general good fit of the model (i.e. random distribution of residuals) and slightly lower carbon values in higher instars (Z V) than predicted by the model (Figure 2), which might be due to problems during metamorphosis to the megalopa stage. It is known that larvae are able to postpone their metamorphosis to the megalopa stage if the cue for the suitable habitat cannot be detected (Krimsky & Epifanio, Reference Krimsky and Epifanio2008), but the enduring lack of those cues cause stress and the depletion of resources.

Changes in the kinetic characteristics of enzymes reflect differences in metabolic regulation and are inevitably involved in adaptation and acclimation to ambient temperature (Wells et al., Reference Wells, Lu, Hickey and Jeffs2001; Somero, Reference Somero2005). A decrease in CS activity is a strong indication for metabolic reduction. In C. pagurus CSA decreases with increasing enzyme assay temperature, indicating an optimum temperature at 14°C. The generally lower citrate synthase activity, i.e. lower aerobic capacity, of 18°C larvae over the whole measured temperature range indicate that acclimation to above-optimum temperatures cannot be compensated. The compensation over certain temperature ranges has been reported for several fish and crustaceans (e.g. Salomon & Buchholz, Reference Salomon and Buchholz2000; Lannig et al., Reference Lannig, Eckerle, Serendero, Sartoris, Fischer, Knust, Johansen and Pörtner2003; Lemos et al., Reference Lemos, Salomon, Gomes, Phan and Buchholz2003).

No complete larval development was observed in our laboratory experiments at 18°C, although this is a temperature which larvae may encounter in Helgoland waters during summer. Only little information is available about the southern distribution boundaries, where larvae should encounter much warmer water temperatures. There is evidence for a northern expansion of this species in Norway (Woll et al., Reference Woll, van der Meeren and Fossen2006), probably indicating that warming of the ocean drives an expansion if not shift of the geographical range of C. pagurus towards the northern North Sea (see also Wiltshire & Manly, Reference Wiltshire and Manly2004). It is thus necessary to determine the significance of the temperature tolerance window of this species for its distribution boundaries more accurately.

In this context it might be important to recognize that initial larval biomass at hatching varied significantly within our small sample size (N = 8) but without statistically significant correlation between female size and the body mass of freshly hatched Zoea I. However, it must be taken into consideration that our data (12.5–17.1 cm CW) do not cover the whole size-range of mature females and thus future study investigating the full range of female maturity (11.5 to 19 cm; Neal & Wilson, Reference Neal and Wilson2004) may come to a different conclusion. Energy allocation of females into offspring may be controlled by genetic disposition (Reznick, Reference Reznick1981), size (DeMartini et al., Reference DeMartini, DiNardo and Williams2003), temperature (Fischer & Thatje, Reference Fischer and Thatje2008), or the nutritional status of the female (Bernardo, Reference Bernardo1996). A positive relationship between maternal size and offspring size has been observed in both invertebrates (Marshall & Keough, Reference Marshall and Keough2004) and vertebrates (Birkeland & Dayton, Reference Birkeland and Dayton2005), but not universally. Among crustaceans, there are species that show such a relationship, e.g. the xanthoid crab Pseudocarcinus gigas (Gardner, Reference Gardner1997), but also other species not following this pattern, e.g. the American lobster (Homarus americanus) (Ouellet & Plante, Reference Ouellet and Plante2004).

Future studies covering the full size-range of mature C. pagurus need to reveal whether intraspecific variability in energy offspring is a matter of individual variability in female fitness and/or a female size related trade off. Intraspecific variability may also affect larval fitness and survival and thus a future large-scale study is needed to reveal whether the herein reported temperature tolerance pattern for larvae from a single female can be easily translated into populations and consequently management approaches of this species.

ACKNOWLEDGEMENTS

We would like to thank U. Nettelmann for help with the larval culture maintenance and K. Bickmeyer for CHN-analyses. This study was partially conducted in the frame of the EU-project CENSOR (Climate variability and El Niño Southern Oscillation: Impacts for natural coastal resources and management) (Contract No. 511071) and is CENSOR publication No. 0309 with additional support by the Marine Biodiversity and Ecosystem Functioning Network of Excellence MarBEF (Contract No. GOCE-CT-2003-505446) of the FP6.

References

REFERENCES

Anger, K. (2001) The biology of decapod crustacean larvae. Lisse: A.A. Balkema Publishers, Crustacean Issues 14, 420 pp.Google Scholar
Anger, K. and Dawirs, R.R. (1982) Elemental composition (C, N, H) and energy in growing and starving larvae of Hyas araneus (Decapoda, Majidae). Fisheries Bulletin 80, 419–433.Google Scholar
Anger, K., Lovrich, G.A., Thatje, S. and Calcagno, J.A. (2004) Larval and early juvenile development of Lithodes santolla (Molina, 1782) (Decapoda: Anomura: Lithodidae) reared at different temperatures in the laboratory. Journal of Experimental Marine Biology and Ecology 306, 217–230.CrossRefGoogle Scholar
Bernardo, J. (1996) The particular maternal effect of propagule size, especially egg size: patterns, models, quality of evidence and interpretations. American Zoologist 36, 216–236.CrossRefGoogle Scholar
Birkeland, C. and Dayton, P.K. (2005) The importance in fishery management of leaving the big ones. Trends in Ecology and Evolution 20, 356–358.CrossRefGoogle ScholarPubMed
Bradford, M.M. (1976) A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein–dye binding. Analytical Biochemistry 72, 248–254.CrossRefGoogle ScholarPubMed
Dahlhoff, E.P. (2004) Biochemical indicators of stress and metabolism: applications for marine ecological studies. Annual Review of Physiology 66, 183–207.CrossRefGoogle ScholarPubMed
Dawirs, R.R. (1979) Effects of temperature and salinity on larval development of Pagurus bernhardus (Decapoda, Paguridae). Marine Ecology Progress Series 1, 323–329.CrossRefGoogle Scholar
Dawirs, R.R. (1985) Temperature and larval development of Carcinus maenas (Decapoda) in the laboratory: prediction of larval dynamics in the sea. Marine Ecology Progress Series 24, 297–302.CrossRefGoogle Scholar
Dawirs, R.R. and Dietrich, A. (1986) Temperature and laboratory feeding rates in Carcinus maenas L. (Decapoda: Portunidae) larvae from hatching through metamorphosis. Journal of Experimental Marine Biology and Ecology 99, 133–147.CrossRefGoogle Scholar
DeMartini, E.E., DiNardo, G.T. and Williams, H.A. (2003) Temporal changes in population density, fecundity, and egg size of the Hawaiian spiny lobster (Panulirus marginatus) at Necker Bank, Northwestern Hawaiian Islands. Fisheries Bulletin 101, 22–31.Google Scholar
Fischer, S. and Thatje, S. (2008) Temperature-induced oviposition in the brachyuran crab Cancer setosus along a latitudinal cline: aquaria experiments and analysis of field data. Journal of Experimental Marine Biology and Ecology 357, 157–164.CrossRefGoogle Scholar
Gardner, C. (1997) Effect of size on reproductive output of giant crabs Pseudocarcinus gigas (Lamarck): Oziidae. Marine and Freshwater Research 48, 581–587.CrossRefGoogle Scholar
Giménez, L. and Anger, K. (2003) Larval performance in an estuarine crab, Chasmagnathus granulata, is a consequence of both larval and embryonic experience. Marine Ecology Progress Series 249, 251–264.CrossRefGoogle Scholar
Giménez, L. and Torres, G. (2002) Larval growth in the estuarine crab Chasmagnathus granulata: the importance of salinity experienced during embryonic development, and the initial larval biomass. Marine Biology 141, 877–885.CrossRefGoogle Scholar
Giménez, L., Anger, K. and Torres, G. (2004) Linking life history traits in successive phases of a complex life cycle: effects of larval biomass on early juvenile development in an estuarine crab, Chasmagnathus granulata. Oikos 104, 570–580.CrossRefGoogle Scholar
Harms, J., Anger, K., Klaus, S. and Seeger, B. (1991) Nutritional effects on ingestion rate, digestive enzyme activity, growth, and biochemical composition of Hyas araneus L. (Decapoda: Majidae) larvae. Journal of Experimental Marine Biology and Ecology 145, 233–265.CrossRefGoogle Scholar
Hastie, T.J. and Tibshirani, R.J. (1990) Generalized Additive Models. London: Chapman & Hall/CRC Press, 335 pp.Google Scholar
Heilmayer, O., Brey, T. and Pörtner, H.O. (2004) Growth efficiency and temperature in scallops: a comparative analysis of species adapted to different temperatures. Functional Ecology 18, 641–647.CrossRefGoogle Scholar
Heilmayer, O., Thatje, S., McClelland, C., Conlan, K. and Brey, T. (2008) Changes in biomass and elemental composition during early ontogeny of the Antarctic isopod crustacean Ceratoserolis trilobitoides. Polar Biology 31, 1325–1331.CrossRefGoogle Scholar
Ingle, R.W. (1981) The larval and post-larval development of the edible crab, Cancer pagurus Linnaeus (Decapoda: Brachyura). Bulletin of the British Musuem of Natural History (Zoology) 40, 211–236.Google Scholar
Kaiser, R. and Gottschalk, G. (1972) Ausreissertest nach Nalimov. Elementare Tests zur Beurteilung von Messadaten. Mannheim, Wien, Zürich: Bibliographisches Institut, pp. 18–21.Google Scholar
Krimsky, L.S. and Epifanio, C.E. (2008) Multiple cues from multiple habitats: effect on metamorphosis of the Florida stone crab, Menippe mercenaria. Journal of Experimental Marine Biology and Ecology 358, 178–184.CrossRefGoogle Scholar
Lannig, G., Eckerle, L.G., Serendero, I., Sartoris, F.J., Fischer, T., Knust, R., Johansen, T. and Pörtner, H.O. (2003) Temperature adaptation in eurythermal cod (Gadus morhua): a comparison of mitochondrial enzyme capacities in boreal and Arctic populations. Marine Biology 142, 589–599.CrossRefGoogle Scholar
Lemos, D., Salomon, M., Gomes, V., Phan, V.N. and Buchholz, F. (2003) Citrate synthase and pyruvate kinase activity during early life stages of the shrimp Farfantepenaeus paulensis (Crustacea, Decapoda, Penaeidae): effects of development and temperature. Comparative Biochemistry and Physiology B 135, 707–719.CrossRefGoogle ScholarPubMed
Lovrich, G.A., Thatje, S., Calcagno, J.A., Anger, K. and Kaffenberger, A. (2003) Changes in biomass and chemical composition during lecithotrophic larval development of the southern king crab, Lithodes santolla (Molina). Journal of Experimental Marine Biology and Ecology 288, 65–79.CrossRefGoogle Scholar
Marshall, D.J. and Keough, M.J. (2004) When the going gets rough: effect of maternal size manipulation on larval quality. Marine Ecology Progress Series 272, 301–305.CrossRefGoogle Scholar
Neal, K.J. and Wilson, E. (2004) Cancer-pagurus. Edible crab. Marine Life Information. Network: Biology and Sensitivity Key Information Sub-programme (online). Plymouth: Marine Biological Association of the United Kingdom.Google Scholar
Pörtner, H.O. (2001) Climate change and temperature-dependent biogeography: oxygen limitation of thermal tolerance in animals. Naturwissenschaften 88, 137–146.Google ScholarPubMed
Pörtner, H.O., Berdal, B., Blust, R., Brix, O., Colosimo, A., Wachter, B., Giuliani, A., Johansen, T., Fischer, T., Knust, R., Lannig, G., Naevdal, G., Nedenes, A., Nyhammer, G., Sartoris, F.J., Serendero, I., Sirabella, P., Thorkildsen, S. and Zakhartsev, M. (2001) Climate induced temperature effects on growth performance, fecundity and recruitment in marine fish: developing a hypothesis for cause and effect relationships in Atlantic cod (Gadus morhua) and common eelpout (Zoarces viviparus). Continental Shelf Research 21, 1975–1997.CrossRefGoogle Scholar
Pörtner, H.O., Storch, D. and Heilmayer, O. (2005) Constraints and trade-offs in climate dependent adaptation: energy budgets and growth in a latitudinal cline. Scientia Marina 69, 39–55.CrossRefGoogle Scholar
Ouellet, P. and Plante, F. (2004) An investigation of the sources of variability in American lobster (Homarus americanus) eggs and larvae: female size and reproductive status, and interannual and interpopulation comparisons. Journal of Crustacean Biology 24, 481–495.CrossRefGoogle Scholar
Reznick, D. (1981) ‘Grandfather effects’: the genetics of interpopulation differences in offspring size in the Mosquito Fish. Evolution 35, 941–953.Google ScholarPubMed
Salomon, M. and Buchholz, F. (2000) Effects of temperature on the respiration rates and the kinetics of citrate synthase in two species of Idotea (Isopoda, Crustacea). Comparative Biochemistry and Physiology 125, 71–81.CrossRefGoogle ScholarPubMed
Sidell, B.D., Driedzic, W.R., Stowe, D.B. and Johnston, I.A. (1987) Biochemical correlations of power development and metabolic fuel preferenda in fish hearts. Physiological Zoology 60, 221–232.CrossRefGoogle Scholar
Sokal, R.R. and Rohlf, F.J. (1981) Biometry—the principles and practice of statistics in biological research. San Francisco: W.H. Freeman, 859 pp.Google Scholar
Somero, G.N. (2005) Linking biogeography to physiology: evolutionary and acclimatory adjustments of thermal limits. Frontiers in Zoology 2.CrossRefGoogle ScholarPubMed
Thatje, S., Anger, K., Calcagno, J.A., Lovrich, G.A., Pörtner, H.O. and Arntz, W.E. (2005) Challenging the cold: crabs reconquer the Antarctic. Ecology 86, 619–625.CrossRefGoogle Scholar
Torres, C.G. and Escribano, R. (2003) Growth and development of Calanus chilensis nauplii reared under laboratory conditions: testing the effects of temperature and food resources. Journal of Experimental Marine Biology and Ecology 294, 81–99.CrossRefGoogle Scholar
Wells, R.M.G., Lu, J., Hickey, A.J.R. and Jeffs, A.G. (2001) Ontogenetic changes in enzyme activities associated with energy expenditure during development in the spiny lobster, Jasus edwardsii. Comparative Biochemistry and Physiology 130, 339–347.CrossRefGoogle ScholarPubMed
Wiltshire, K.H. and Manly, B.F.J. (2004) The warming trend at Helgoland Roads, North Sea: phytoplankton response. Helgoland Marine Research 58, 269–273.CrossRefGoogle Scholar
Woll, A.K., van der Meeren, G.I. and Fossen, I. (2006) Spatial variation in abundance and catch composition of Cancer pagurus in Norwegian waters: biological reasoning and implications for assessment. ICES Journal of Marine Science 63, 421–433.CrossRefGoogle Scholar
Figure 0

Table 1. Cancer pagurus. Changes in dry weight (W), carbon (C), nitrogen (N) and C:N ratio during time days after hatch (±SD) at five temperatures. Larvae reared at 15°C are from different female.

Figure 1

Fig. 1. Model of body mass increase (C) throughout the larvae development of the edible crab Cancer pagurus. Equation for the model is: CBC =−7.97−10.755 × t + 919.52 × f(T) + 476.634 × t × f(T); N = 463, F = 1599.51, R2 = 0.954; T represents the temperature in Kelvin and t the time in days. Isolines represent carbon content in µg. ZI–ZV are the larval instars. White areas show the realistic range of the model.

Figure 2

Table 2. Cancer pagurus. Size of eight different females and elemental composition (dry weight (W), carbon (C), nitrogen (N), hydrogen (H), C:N and C:H ratio of freshly hatched larvae (hatch 1)).

Figure 3

Fig. 2. Residual by predicted plot from the model predicting larval growth: plot of regression residuals versus predicted values. The plot does not indicate a problem with the model.

Figure 4

Fig. 3. Citrate synthase activity (U * g Prot−1) of Cancer pagurus larvae acclimated to three different temperatures (light grey bars—10°C acclimated; white bars—14°C acclimated; dark grey bars—18°C acclimated) compared with larvae immediately after hatching (black bars). Bars within an assay temperature not connected with the same letter are significantly different (P > 0.05).

Figure 5

Table 3. Cancer pagurus. One-way ANOVA to evaluate the maternal influence on body mass (as dry weight (W), carbon (C), and nitrogen (N)]) and C:N ratio of freshly hatched larvae (first day of the hatching period 1). *, ANOVA on ranks because equal variance test failed.