You are not always what you eat—Fatty acid bioconversion and lipid homeostasis in the larvae of the sand mason worm Lanice conchilega

Rita M. Franco-Santos; Holger Auel; Maarten Boersma; Marleen De Troch; Martin Graeve; Cedric L. Meunier; Barbara Niehoff

doi:10.1371/journal.pone.0218015

Abstract

The meroplanktonic larvae of benthic organisms are an important seasonal component of the zooplankton in temperate coastal waters. The larvae of the reef-building polychaete Lanice conchilega contribute up to 15% of the summer zooplankton biomass in the North Sea. Despite their importance for reef maintenance (which positively affects the benthic community), little is known about the trophic ecology of this meroplanktonic larva. Qualitative and quantitative estimates of carbon (C) transfer between trophic levels and of fatty acid (FA)—specific assimilation, biosynthesis, and bioconversion can be obtained by compound-specific stable isotope analysis of FA. The present work tested the hypothesis that the concept of fatty acid trophic markers (FATM), widely used for studies on holoplankton with intermediate to high lipid contents, is also applicable to lipid-poor organisms such as meroplanktonic larvae. The incorporation of isotopically-enriched dietary C by L. conchilega larvae was traced, and lipid assimilation did not follow FA-specific relative availabilities in the diet. Furthermore, FAs that were unavailable in the diet, such as 22:5(n-3), were recorded in L. conchilega, suggesting their bioconversion by the larvae. The results indicate that L. conchilega larvae preferentially assimilate certain FAs and regulate their FA composition (lipid homeostasis) independently of that of their diet. Their quasi-homeostatic response to dietary FA availability could imply that the concept of FATM has limited application in lipid-poor organisms such as L. conchilega larvae.

Citation: Franco-Santos RM, Auel H, Boersma M, De Troch M, Graeve M, Meunier CL, et al. (2019) You are not always what you eat—Fatty acid bioconversion and lipid homeostasis in the larvae of the sand mason worm Lanice conchilega. PLoS ONE 14(6): e0218015. https://doi.org/10.1371/journal.pone.0218015

Editor: Vitor Hugo Rodrigues Paiva, MARE – Marine and Environmental Sciences Centre, PORTUGAL

Received: October 30, 2018; Accepted: May 23, 2019; Published: June 6, 2019

Copyright: © 2019 Franco-Santos et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.

Data Availability: The laboratory experimental data and metadata for this study is available through the PANGAEA repository under the link https://doi.pangaea.de/10.1594/PANGAEA.900110.

Funding: RMF-S received a scholarship from the MARES Doctoral Programme on Marine Ecosystem Health and Conservation (www.mares-eu.org) (Framework Agreement Number 2011-0016). MDT was supported by Ghent University (www.ugent.be) (BOF-GOA project 01G02617). CLM was supported by the Bundesministerium für Bildung und Forschung (www.bmbf.de) (BMBF grant no. 01LN1702A). The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

Competing interests: The authors have declared that no competing interests exist.

Introduction

The meroplanktonic larvae of macrobenthic organisms are an important seasonal component of the zooplankton in temperate coastal waters [1]. In the North Sea, the larvae of the sand mason worm Lanice conchilega (Pallas, 1766) contribute up to 15% of the zooplankton biomass between July and September [2], when water temperatures are above 13°C. The reproductive peak of L. conchilega in the southern North Sea occurs in spring and is followed by smaller peaks during the summer and autumn [3], though larval supply can vary between years [4]. The larvae of this polychaete evolve from a short planktonic to a benthic phase and, after a few days, to a second planktonic stage, the aulophore larvae [5]. At this stage larvae already present a (transparent) tube, and display morphological features of a juvenile [6]. They are able to feed in the water column, where they can remain up to 60 days before succeeding to a benthic stage [7]. The benthic, tube-dwelling adults of L. conchilega are non-selective suspension-deposit feeders [8–10], which derive their diet from the organic matter available in the water column and in the sediment. Common food items include diatoms, bacteria and microphytobenthos, and the species is able to switch between food sources [8,10,11].

Lanice conchilega forms reefs capable of structuring the surrounding habitat and creating a complex and heterogeneous environment [12,13]. The reefs provide refuge against predation and physical and chemical stresses, attract a variety of other organisms, and serve as nursery and feeding grounds to several species, including fish and birds [14–19]. Lanice conchilega is an ecosystem engineer whose presence favors species richness and faunal abundance and positively affects the benthic community [12,13,19,20].

In comparison with adult organisms, few studies have been conducted on the ecology of early stages of L. conchilega. These mostly address topics such as juvenile settlement and buoyancy [4,7,21,22], and little information is available on the feeding ecology of the larvae. The survival and development of these meroplanktonic larvae and, thus, the maintenance of L. conchilega reefs, depend, a.o., on their feeding success. To the best of our knowledge there are no studies published on the feeding ecology of the larvae, a gap which needs to be addressed if the larval stage of this species is to be better understood.

Information on the feeding ecology of an organism on short and long time scales, as well as on the pathways of energy flow between prey and predator, can be obtained from the analysis of fatty acid trophic markers (FATM) [23] in combination with the use of stable isotopes. Fatty acid trophic markers are usually assimilated in a conservative manner (i.e., largely unmodified) by consumers [24,25]. The FAs 16:1(n-7), 16:2(n-4), 16:3(n-4), and 20:5(n-3) (eicosapentaenoic acid, EPA) are characteristic of diatoms, whereas 18:4(n-3) and 22:6(n-3) (docosahexaenoic acid, DHA) are considered dinoflagellate markers [8,23,26,27]. Compound-specific stable isotope analysis (CSIA) is a powerful tool for tracing organic matter origin and fate and to investigate biochemical processes and patterns in individuals and ecosystems (e.g., [28–30]). It provides accurate and precise measurements of the isotopic composition of compounds (up to 0.3‰ or 0.0003 atom percent excess, APE) [31]. When applied together with FA determination, CSIA highlights minute changes in FA-specific concentration. When used in feeding experiments with enriched diets, CSIA enables the discrimination of the origin of a FA–internal or dietary–, and also provides quantitative estimates for the transfer of lipid dietary carbon (C) from prey to predator and for FA-specific assimilation and potential biosynthesis and bioconversion [26,27,32,33].

The present work had two objectives. The first was to start filling the gap of knowledge on the feeding ecology of L. conchilega larvae. Experiments investigating FA-related C assimilation have been conducted mostly for holoplanktonic organisms with intermediate to high lipid contents, whose FA profile tends to reflect that of the food items they ingest (e.g., [26,27]). Little is known, however, about how meroplanktonic organisms, which are usually lipid-poor (total lipid content ranging between 5–15% dry mass), incorporate dietary FAs. It is possible that the lipid content and, thus, requirement, of an organism can dictate whether it will assimilate dietary FAs in an unmodified manner or not. We assume that the larvae of L. conchilega are lipid-poor, so our second objective was to test the hypothesis that the concept of FATM can also be applied in a feeding study with lipid-poor meroplanktonic larvae. In order to do so, the polychaete larvae were fed with a diatom culture previously enriched with ¹³C and the incorporation of isotopically enriched FA-specific lipid C into the consumer was followed. This allowed for the recording of C assimilation and turnover in the organism, and also for the investigation of possible bioconversion pathways.

Method

Field sampling

Zooplankton samples for the feeding experiment were collected on June 6^th 2016 with a 500 μm mesh-size CalCOFI net, which was towed horizontally for 15 minutes at 5 m depth off the German island of Helgoland (54^o11’N, 07^o54’E), in the southern North Sea. As zooplankton samples are routinely collected at this location for the Helgoland Roads time series, no specific permissions were required for this location/sampling. Samples were taken to the laboratory and intact and active aulophore larvae of the polychaete L. conchilega were immediately sorted under an Olympus SZX16 stereoscopic microscope.

Algae culture

Batch cultures were created daily for Conticribra weissflogii (Grunow) (Stachura-Suchoples and Williams, 2009) for five consecutive days by diluting a stock solution with fresh f/2 medium (after [34]). Diatoms were enriched with ¹³C by adding ¹³C-enriched sodium bicarbonate (NaH¹³CO₃) to the medium at a concentration of 4 mg L^-1. Cultures were kept in constant light inside a temperature-controlled room at 18°C, and stirred twice a day in order to keep cells suspended. Algae were grown for five days and then used as food suspension (exponential growth phase) for the polychaete larvae during the experiment. Conticribra weissflogii cultures were sampled daily (5 days) during the experiment for determination of cell C, nitrogen (N), and FA content and of FA-specific ¹³C isotopic enrichment, though only 4 of the 5 samples yielded good chromatograms for the FA analysis. This was done by filtering subsamples of known cell concentrations through pre-combusted (500°C for 24 h) Whatman GF/F filters (0.7 μm pore size, 25 mm diameter). Filters for determination of prey C and N content were dried at 60°C for 48 h, folded inside aluminum foil, and stored in a desiccator until analysis. Samples for FA analyses were placed into pre-combusted lipid vials and stored at -80°C. The remaining volume of the cultures was then used to feed polychaete larvae. Cell densities were determined with a BD Accuri C6 Flow Cytometer.

Experimental design

A total of 390 larvae were sorted, 30 for determination of in situ body C and N content, 150 for analyzing the in situ FA content and composition, and 210 for the feeding experiment. The enriched C. weissflogii was used as food source in the feeding experiment with L. conchilega larvae. Feeding experiments with L. conchilega were initiated immediately after sorting of the larvae. Individuals were kept for five days in groups of 70 individuals in triplicate 500 mL glass beakers fitted with a 300 μm meshed-bottom cylinder (140 ind L^-1). The diatom suspension was provided on a daily basis at a concentration of 8000 cells mL^-1. The polychaete larvae were kept in a dark temperature-controlled room at the temperature recorded when they were sampled in situ, i.e., 13.5 ± 0.3°C. The beakers were gently stirred three times a day for food resuspension in the water. A partial water exchange (66%) was performed daily and followed by the addition of new algae culture, with a final diatom concentration > 8000 cells mL^-1. At the end of the experiment the larvae were sampled from each replicate for body C and N content determination and for analysis of FA composition and FA-specific ¹³C isotopic enrichment (n = 3, with 10 and 50 individuals per sample for C/N and FA analyses, respectively). Individuals (inside their tubes) were gently washed in distilled water, placed into pre-weighed tin capsules (5*9 mm, IVA Analysentechnik) for body C and N content determination or into pre-combusted glass vials for FA analyses, and stored at -80°C until further analysis. Tin capsules with larvae were dried at 60°C for 48 h, weighed with an ultra micro-balance (detection limit: 0.1 μg; XP6U Ultra Micro Balance, Mettler Toledo, Germany) and stored in a desiccator until analysis.

Carbon, nitrogen and fatty acid content analyses

The C and N contents of all larvae and algae samples were measured with an elemental analyzer (detection limit: 2 μg C / 0.5 μg N; maximum error: ± 3%, Euro EA 3000, EuroVector S.P.A., Milan, Italy) using acetanilide as a standard.

Lipids were extracted and FAs identified as described by Boissonnot et al., 2016 [26]. Samples were homogenized in a dichloromethane:methanol (2:1, v:v) solution, from which total lipids were extracted. A known amount of an internal standard, the tricosanoic acid methyl ester (23:0), was added to each sample. Potassium chloride (KCl 0.88% solution) was added to create a biphasic system and aid in extraction. Lipid extracts were transesterified by heating samples with 3% sulfuric acid (H₂SO₄) in methanol at 80°C under nitrogen atmosphere for 4 hours. Fatty acid methyl esters (FAMEs) were then extracted with cyclohexane, and determined and quantified with a gas chromatograph (HP 6890 N, Agilent Technologies Deutschland GmbH & Co. KG) equipped with a 60m × 0.25mm i.d. wall-coated open tubular capillary column (film thickness: 0.25 μm; liquid phase: DB-FFAP), a split/splitless injector (250°C) and a flame ionization detector (280°C). The peaks in the chromatograms generated were manually identified in relation to known standards analyzed together with samples. Chromatograms were then evaluated with the ChemStation software from Agilent. The A:B(n-X) shorthand notation was used to refer to FAs, where A is the number of carbon atoms, B is the number of double bonds, and (n-X) gives the position of the double bond closest to the terminal methyl group. Total lipid contents of larvae and algae cultures were calculated by adding the mass of all FAs. Fatty acid concentrations are presented in absolute (ng C ind^-1) and relative (% of total FA, TFA) units in order to report on both the quantity of FAs available in the samples and the relative importance of each FA within the total FAs for each sample (respectively). Fatty alcohols were not detected in the samples.

Compound-specific stable isotope analysis (CSIA)

The FA-specific stable isotope composition of carbon in FAMEs extracted was obtained according to Boissonnot et al., 2016 [26] with a Thermo gas chromatography-combustion-isotope ratio mass spectrometry (GC-c-IRMS) system, equipped with a Trace GC Ultra gas chromatograph, a GC Isolink, and a Delta V Plus isotope ratio mass spectrometer connected via a Conflo IV interface (Thermo Scientific Corporation, Bremen, Germany). The chromatograms containing peak areas and C isotope ratios were analyzed with the IRMS software Isodat 3.0. The 14:0 and 18:0 FAME reference standards (Iowa University) with known δ-values used for further calculations.

The equations used by Boissonnot et al., 2016 [26] to calculate carbon assimilation were also applied in the present study, and were:

Isotopic ratios of FAs (1) where R_sample and R_standard are the ratio of ¹³C/¹²C in the sample and in the Vienna Pee De Belemnite reference standard, respectively;

Atom percent (atom%), which are converted δ-values and express isotope data in terms of isotope concentrations to inform on the ¹³C enrichment in each FA (2) where R is the ratio ¹³C/¹²C in the sample;

Atom percent excess (APE), which considers the isotopic enrichment in experimental larvae (in comparison to in situ individuals) due to the assimilation of algae enriched in ¹³C (3) Carbon mass of each FA (C_mass, in μg C ind^-1), which divides the FA mass by the FAME mass (4) where A_12C, A_13C, A_H, and A_O are the atomic masses of ¹²C, ¹³C, H and O, respectively, C_FA is the number of carbon atoms in the FA, H_FAME and O_FAME are the number of hydrogen and oxygen atoms in the FAME, and FA_mass is the mass (in μg ind^-1) of the FA;

Proportion of carbon assimilated (PA) (5) where L is the average enrichment (in atom%) in all algae FAs but 18:0 (L = 3.62%). The averaging is to account for the elongation and/or desaturation of small amounts of dietary FAs when they are assimilated by the larvae [23], and the exclusion of 18:0 is due to its poor enrichment in comparison to other FAs in prey cultures [26,35]. When calculating the PA in the larvae FA 18:0, L was the average of enrichment in algae FA 18:0 (L_{FA 18:0} = 1.87%);

Carbon assimilation (C_assim, in μg C ind^-1), calculated for each FA (6) Total C assimilation was obtained by summing the C_assim values for all FAs. Assimilation rates were obtained by dividing C_assim by the number of experimental days when sample was collected;

Carbon turnover rates (C_T in % day^-1) (7) where _(t) indicates the specific sampling time for which the values of C_assim and C_mass should be used and Δt the number of experimental days when sample was collected.

Statistical analyses

Differences in atom% for the different batches of the diatom culture were investigated with one-way analysis of variance (ANOVA), and the origin of differences was identified by applying the Tukey HSD (Honestly Significant Difference) post-hoc test with a 95% confidence limit. Prior to the ANOVAs, the data were tested for normality and homogeneity of variances with Shapiro-Wilk and Bartlett tests, respectively. The difference between in situ and laboratory-fed polychaete larvae in dry mass (DM), C, N, and total lipid contents, and C:N molar ratio were investigated with a t-test (for each variable). Data used for t-tests was also previously tested for normality and homogeneity of variances. Whenever a p-value was obtained, significance was set at p ≤ 0.05. All univariate analyses were performed using R ver. 3.4.4 [36].

Multivariate analysis of FA composition for L. conchilega individuals was performed with a dendrogram, which clustered individuals with similar FA profiles (based on their group average linkage clustering). The dendrogram was generated from a Bray-Curtis similarity matrix obtained from absolute and percentage (with logit transformation, as suggested by Warton and Hui, 2011 [37]) data on FAs with relative content > 1% TFA. All multivariate analyses were performed with PRIMER 7.0 software [38].

Results

Algal composition and enrichment

The average C and N contents of the C. weissflogii cultures were 577 and 117 μg L^-1, respectively, resulting in an average molar C:N ratio of 5.8 (Table 1). The average lipid content was 116 μg L^-1 (Table 1). The FA profile of the diatom (Table 1, Fig 1) was dominated by polyunsaturated fatty acids (PUFAs), which comprised 61% of total fatty acids (TFA). Saturated (SFA) and monounsaturated (MUFA) fatty acids were present in similar amounts, 22 and 17% TFA, respectively. The most important FAs in the C. weissflogii batches were 20:5(n-3) (EPA), 16:3(n-4), 16:1(n-7), and 16:0 (22, 19, 15 and 13% TFA, respectively). The FA 22:6(n-3) (DHA) was present in a smaller amount, i.e., 5% of TFA.

Download:

Fig 1. Fatty acid relative (% total FA, %TFA) and absolute (ng C ind^-1)concentrations in diatoms (C. weissflogii, n = 4) and in L. conchilega larvae (n = 3 with 50 individuals pooled per sample).

Data for larvae are differentiated between organisms sampled in situ and after 5 days of feeding on the diatom. Standard error bars are shown for the mean values.

https://doi.org/10.1371/journal.pone.0218015.g001

Download:

Table 1. Dry mass (DM, in μg ind^-1); carbon (C), nitrogen (N) and total lipid contents (TLC) (ng larvae^-1 / diatom: μg L^-1 (pg cell^-1)); molar C:N ratio; and FA-specific absolute (ng C larvae^-1) and relative (larvae: ‰ body C (italics, second row) / diatom: ‰ cell C / all: % total FA, in parenthesis) composition of L. conchilega larvae and its diet, the diatom C. weissflogii.

https://doi.org/10.1371/journal.pone.0218015.t001

The ¹³C isotopic enrichment in all diatom FAs (but 18:0) varied from 2.88 ± 0.21 atom% to 4.00 ± 0.19 atom% between the different batches, averaging 3.62 ± 0.50 atom% for all days (Table 2, Fig 2). For the FA 18:0, isotopic enrichment averaged 1.87 ± 0.30 atom% between the different batches (Table 2, Fig 2). Enrichment was significantly different between batches (ANOVA, F = 21.66, df = 3,67, p = 6*10⁻¹⁰), being higher on days 1 and 5 than on day 4 (Tukey, p = 0.03) and on day 2 (Tukey, p < 1*10⁻⁷), and higher on day 4 than on day 2 (Tukey, p = 5*10⁻⁴). The major FAs cited above (or PUFAs in general) showed the highest enrichment values within the FAs (Table 2).

Download:

Fig 2. ¹³C isotopic enrichment (atom%) of C. weissflogii daily batch cultures fed to polychaete larvae during the experiment.

Straight lines represent atom% values for the fatty acid (FA) 18:0, whereas boxplots and open circles represent average atom% values from all the other FAs. The minimum and maximum observation values are represented by the lower and upper whiskers (respectively), the first and third quartiles are represented by the lower and upper hinges (respectively), and the median is shown as the line that divides the box.

https://doi.org/10.1371/journal.pone.0218015.g002

Download:

Table 2. Total fatty acid (TFA) and FA-specific (> 1% TFA) ¹³C isotopic enrichment for C. weissflogii (atom%) and L. conchilega (APE); and carbon assimilation (C_assim, as ng C ind^-1 (% total C assimilated)) and turnover rate (C_turn, as % day^-1) for L. conchilega.

https://doi.org/10.1371/journal.pone.0218015.t002

Larval composition and enrichment

Approximately 85% of the L. conchilega larvae survived at the end of the experiment. The average in situ DM was 55 μg ind^-1, and in situ C and N contents of larvae were 7668 and 1929 ng ind^-1, respectively. The average DM, C, and N values at the end of the experiment were 52 μg ind^-1 and 8511 and 2170 ng ind^-1, respectively (Table 1). Changes in DM (t = 0.59751, df = 2, p = 0.61) and in C (t = -0.42141, df = 2, p = 0.71) and N (t = -0.54494, df = 2, p = 0.64) contents between larvae sampled in situ and after the experiment were not significant, indicating that the organisms did not lose or gain weight during the five-day incubation. The molar C:N ratio of larvae remained the same throughout the experiment at an average of 4.6 (t = -0.02381, df = 2, p = 0.98). The average lipid content was similar between in situ (632 ng ind^-1) and experimental (845 ng ind^-1) individuals (t = -3.4065, df = 2, p = 0.076, Table 1). The lipid C content accounted for approximately 7% of the total C content of larvae sampled in situ and at the end of the experiment. Total lipid content comprised 1.2 ± 0.3% DM for in situ larvae and 1.7 ± 0.4% DM for larvae at the end of the experiment.

The major FAs in L. conchilega larvae were EPA, 16:0 and 22:5(n-3), which corresponded to 22, 20, and 15% TFA at the beginning and to 19, 22, and 15% TFA at the end of the experiment, respectively (Table 1). About a quarter of the FAs identified were only present in small amounts (<1% TFA, Fig 1). The relative FA composition of L. conchilega larvae fed with diatoms remained similar to that of the individuals collected in situ (Table 1, Fig 1), as conveyed by the 95.5% similarity between samples shown in the Bray-Curtis dendrogram (Fig 3A). A similar pattern was observed for the absolute FA composition of the larvae (Table 1, Fig 1), except that one of the in situ replicates was grouped with the experimental ones at 91% similarity, and these were found to be 82% similar to the other in situ replicates (Fig 3B). These results further reinforce the above mentioned statistically non-significant differences in lipid C between in situ larvae and those fed during the experiment. In general, changes (i.e., increase or decrease) in the absolute concentration of a FA matched the change in its relative concentration (Fig 1). The only exception to this pattern were the FAs 18:0, 18:1(n-7) and EPA, whose relative concentrations in the larvae decreased despite an increase in their absolute concentrations (Fig 1).

Download:

Fig 3. Dendrograms generated with Bray-Curtis similarity matrices comparing fatty acid profiles of L. conchilega larvae.

Similarity matrices present data on the FA profile of larvae sampled in situ and after 5 days of feeding on the diatom C. weissflogii. Only FAs with relative content > 1% TFA were used to generate the similarity matrices. a) logit transformed relative values for FA composition (% TFA). b) absolute (ng C ind^-1) values for FA composition.

https://doi.org/10.1371/journal.pone.0218015.g003

The average ¹³C enrichment of L. conchilega FAs was of 1.16 ± 0.48 APE, ranging from 0.49 APE in FA 20:1(n-11) to 2.01 APE for FA 16:3(n-4) (Table 2). The FAs 18:1(n-11), 20:1(n-11), and 22:5(n-3) had the lowest enrichment values (together with 18:0) in L. conchilega, whereas the FAs 16:2(n-4), 16:3(n-4), and 16:1(n-7) (diatom fatty acid trophic markers, FATM) had the highest APE (together with 18:4(n-3)).

Origin of FAs in the larvae

The enrichment of FAs with ¹³C enabled for the discrimination of the source of each FA-specific lipid C content: internal (Table 1), for FAs already present in the in situ larvae, and dietary (Table 2), for FAs assimilated via feeding on the enriched diatoms. The absolute content of internal FAs at the end of the experiment remained the same (~98%) as recorded from in situ larvae for two of the three replicates, and on average represented 92% of the in situ internal FA content (Table 1). Given the similarity of C content of internal FA between the beginning and end of the experiment the FA-specific differences are not considerable. The internal FA that showed the greatest decrease in terms of C content during the experiment was EPA, which fell from 22% TFA in in situ larvae to 19% of the total internal FA in diatom-fed larvae (Table 1). Saturated FAs also showed a slight decrease in C content between the beginning (38% TFA) and end (36% TFA) of the experiment.

Lipid C assimilation and turnover in larvae

Total assimilation of C into lipids in polychaete larvae at the end of the experiment was approximately 200 ng lipid C ind^-1 (Table 2). The relative assimilation of lipid C into FAs (Table 2) followed the relative composition of internal FAs (Table 1). Approximately half of the lipid C was assimilated into PUFAs, a third into SFAs, and only a sixth into MUFAs (Table 2, Fig 4). The majority of the total lipid C was assimilated into 16:0 and EPA (22 and 21%, respectively), followed by 22:5(n-3), DHA, and 18:0 (13, 11 and 10%, respectively). It is worth noting that the incorporation of dietary C into specific FAs did not follow their relative availability in the diet. The absolute concentration of the FAs 17:0, 18:1(n-11), 20:1(n-11), and 22:5(n-3) increased in L. conchilega, even though they were absent from the diatom cultures fed to the larvae (Table 2). The absolute concentration of the FAs 18:0 and DHA also increased in the diatom-fed larvae (Table 2), even though they were available in low amounts in the diet (1 and 5% TFA in C. weissflogii, respectively). Furthermore, the FAs i-15:0, 16:1(n-5), 18:2(n-6), 18:3(n-6), and 20:4(n-3) were only identified in C. weissflogii (Fig 1) and were either not assimilated or bioconverted by the polychaete larvae. Carbon assimilation in FATM, such as 16:1(n-7), 16:2(n-4), and 16:3(n-4), was low in comparison to that into other FAs, but amounted to the highest C turnover rates (Table 2).

Download:

Fig 4. FA-specific carbon assimilation (ng C ind^-1, left panels) and turnover (%, right panels) in L. conchilega larvae after the feeding experiment (n = 3).

Values are shown for the sum of all fatty acids (TFA), saturated (SFA), monounsaturated (MUFA), and polyunsaturated (PUFA) fatty acids (upper panels); and for specific FAs (lower panels).

https://doi.org/10.1371/journal.pone.0218015.g004

The average lipid C turnover rate was of 6% day^-1, but varied between 3 and 11% day^-1 for different FAs (Table 2). Carbon turnover of SFA and PUFA was similar, 7% day^-1, and slightly higher than that of MUFAs, 5% day^-1. Overall, 32% of all lipid C was replaced with enriched dietary lipid C by the end of the experiment (Fig 4).

Discussion

This study is the first to our knowledge to present data on C assimilation and turnover as well as on FA bioconversion in the larvae of the polychaete L. conchilega, and in any other meroplanktonic larvae for that matter. Although these were recorded from a short-term feeding experiment (5 days), they indicate how quickly polychaete larvae can assimilate dietary material. These lipid-poor larvae seem to be able to regulate their lipid composition (homeostasis) by preferentially assimilating certain FAs and/or by controlling their FA metabolism, and through bioconversion of dietary and internal FAs.

Lipid composition, homeostasis, and bioconversion in L. conchilega larvae

The low total lipid content recorded for in situ larvae in the present study (1.2% DM) is similar to that reported for field and starving crab zoeae and bivalve larvae [39–41]. Lanice conchilega larvae thus appear to be lipid-poor organisms.

The relative concentrations of single FAs and of SFAs, MUFAs, and PUFAs were preserved in the larvae during the feeding period. This occurred despite the provision of, and successful feeding upon, a diet with a different FA composition. These results indicate that L. conchilega larvae can selectively accumulate FAs and/or control its FA metabolism and, thus, regulate lipid composition. A quasi-homeostatic response to variation in FA availability in the food source has also been shown for other zooplankton (daphnids) by Müller-Navarra, 2006 [42]. The preferential retention of a FA over another can be easily identified if their relative proportions in the diet are similar, but their assimilation by the larvae differs. In the present study this is the case for the pairs of FAs 16:0 + 16:1(n-7), EPA + 16:3(n-4), and DHA + 18:4(n-3), for which the first FA of each pair was assimilated by L. conchilega in quantities that were 3-20x higher than those of the second FA, indicating their preferential accumulation. The FAs EPA and DHA have an important function for marine consumers as they fuel reproductive processes and neural function, and regulate cell membrane fluidity [43–45]. It has been observed that adult L. conchilega accumulate (n-3) FAs to store energy during gametogenesis [8], but larvae probably use EPA and DHA for growth, as tissue hormones (eicosanoids) are produced from PUFA [46]. The FA 16:0, on the other hand, can be elongated and desaturated into other necessary FAs [23].

There are three different metabolic sources from which the FAs composing the lipid reserves of a consumer can be derived. The FAs can be (1) assimilated unmodified from the food (dietary origin), (2) synthesized de novo (dietary or internal origin), or (3) bioconverted (synthesized via chain elongation and desaturation) from another FA of dietary or internal origin [47]. The bioconversion of FAs has more often been reported for crustaceans [23,32] than for polychaetes [48]. In the present study we report the bioconversion of the FAs 17:0, 18:1(n-11), 20:1(n-11), and 22:5(n-3). It should be noted that the synthesis of odd-chained FAs is uncommon in consumers, and that the FA 17:0 is usually considered a bacterial FATM [23]. The presence of this FA in the lipid profile of L. conchilega larvae indicates that either the organisms are able to synthesize odd-chained FAs or that bacteria were present in small amounts in the experimental units and were ingested by the larvae. Bioconversion occurred for both endogenously-derived FAs, which increased in absolute content during the experiment, and for dietary-derived FAs, which were enriched in ¹³C. The elongation and desaturation of FAs involve reactions catalyzed by enzymes and are subject to isotopic fractionation [49], which could explain why the ¹³C enrichment was lower in the bioconverted FAs. The FA 22:5(n-3) was bioconverted at high quantities and made up 13% of all C assimilated by the larvae. Although marine invertebrates are not able to biosynthesize de novo the FAs 18:2(n-6) and 18:3(n-3), they have a limited ability to convert them into PUFAs via chain elongation and desaturation [23,50]. In the present study neither the diet nor the larvae contained 18:3(n-3), so 22:5(n-3) must have been bioconverted from EPA, as has been suggested for brittle stars [51]. It is noteworthy to mention the high enrichment of the FA 18:4(n-3), which is known to be a FATM for dinoflagellates, in the larvae. Although the connection between the referred FA and dinoflagellates might lead to confusion, in the present study the larvae obtained the FA 18:4(n-3) from their diet–this FA was present in the diatom cultures and attained one of the highest atom% values among diatom FAs.

Lipid homeostasis in lipid-poor and lipid-rich marine planktonic organisms

The ability to regulate FA composition does not come without an energetic cost. Lipid homeostasis can be easy to maintain when an organism feeds upon a prey of similar lipid composition, but will require that energy be allocated to bioconversion if prey have a different biochemical make-up. Furthermore, the way in which an individual utilizes its assimilated FAs is life-stage specific [23]. Based on our results and on the available literature on lipid content and assimilation of mero- and holoplanktonic marine species ([26,27,52,53], other references shown in Table 3, Franco-Santos et al., unpublished data), we put forward the hypothesis that 3 different patterns in lipid homeostasis can be found among lipid-poor and lipid-rich planktonic organisms.

Download:

Table 3. Total lipid content (TLC, in % dry mass) and wax ester content (WE, in % total lipid content) of several meroplanktonic and holoplanktonic species.

https://doi.org/10.1371/journal.pone.0218015.t003

Lipid-poor meroplanktonic larvae from decapods, bryozoans, and vertebrates, a.o., generally display total lipid contents in the order of 5–15% DM (Table 3). According to the available literature, total lipid content is on the lower side of this range for bivalve larvae (2–8% DM) and on the higher part for barnacle larvae (13–15% DM) and octopus paralarvae (< 15% DM) (Table 3). Both lecithotrophic (which possess and feed on yolk reserves) and planktotrophic (which feed on plankton) larvae rely on obtaining enough energy to sustain metabolic functions and growth into juvenile stages, and cannot dispense energy for storage purposes. Lipid classes which are destined for energy storage, such as wax esters (WE) and triacylglycerols, represent a small proportion of total lipid content in lipid-poor meroplankton (Table 3). These organisms have specific requirements of FAs to sustain their body functions, such as formation of biomembranes, and will thus display lipid homeostasis, even though this may require the allocation of further energy for FA bioconversion. This is the case of the meroplanktonic larvae of L. conchilega, as discussed in the previous section. It appears, however, that the larvae of hermatypic and soft corals are lipid-rich (total lipid content reported to range between 41–68% DM; [71, 72]). These would be an exception within the lipid-poor meroplankton, and more information is necessary before we can infer on their ability to regulate lipid composition during the larval stage.

Holoplanktonic organisms with intermediate total lipid content (15–25% DM) include small copepods, tunicates, chaetognaths, and ostracods (Table 3). Generally speaking, the lower end of this range is observed in organisms found in tropical regions, whereas the higher end is recorded for individuals from temperate and polar regions (Table 3). The WE content of organisms varies between different groups, and ranges from values as low as 1% total lipid content to as high as 52% total lipid content (Table 3). These holoplanktonic organisms generally have a small to modest ability for energy storage as lipid reserves, a strategy whose focus is likely the maximization of reproductive output. Such species will mostly assimilate dietary FAs in an unmodified manner (non-homeostatic).

Lipid-rich holoplanktonic species, on the other hand, will display total lipid contents > 30% DM (Table 3). The species for which data were available were mostly from polar regions, and represent amphipods (55% DM), large copepods (31–74% DM), krill (7–60% DM), decapods (35–39% DM), and pteropods (19–31% DM) (Table 3). These organisms need to store energy in the most dense and effective way to survive periods of food shortage during winter and for buoyancy purposes [73], and will thus bioconvert dietary FAs into long chain MUFAs / WE (semi-homeostatic). This can be exemplified by the large proportion of WE within the total lipid content in the mentioned groups, which is approximately 50% in decapods and krill and 34–90% in copepods (Table 3). Only pteropods and amphipods have a lower proportion of WE, which is approximately 15–20% of the total lipid content (Table 3).

The applicability of the FATM concept

In theory, FATM are assimilated by consumers in a conservative manner [24,25]. The applicability of the FATM concept has been broadly acknowledged to depend, a.o., on the qualitative and quantitative transfer of FAs between trophic levels and on their metabolic stability and non-selective incorporation into consumer tissues [23]. In the present study the diatom FATMs were transferred between the producer and consumer trophic levels in a qualitative manner. In quantitative terms, however, the dietary FAs 16:0, EPA, and DHA were preferentially assimilated by L. conchilega larvae. Furthermore, we have postulated that EPA, which is a diatom FATM, was likely bioconverted (via chain elongation, as indicated in Fig 1 of [74]) into 22:5(n-3) by the polychaete larvae. Unlike the trends shown by lipid-rich or lipid-accumulating organisms, the results of this study indicate that the applicability of the concept of FATM is limited in the lipid-poor larvae of L. conchilega. We also suggest that this will be the case for other lipid-poor meroplanktonic larvae, as they probably also display preferential assimilation and bioconversion of FAs in order to sustain body functions.

Experimental conditions

The active uptake of dietary C during the experiment was sufficient to compensate for the larval metabolic costs of living during the 5 days. This indicates that diatoms are a nutritious source of food for L. conchilega larvae, which readily accept these algae and utilize it as metabolically necessary. The lack of literature on feeding studies with meroplanktonic larvae prompted us to look for guidelines on food concentrations from laboratory experiments with small planktonic copepods (e.g., [75]), so it is possible that individuals would have shown higher growth if they had received more food.

A common value of 30–40% DM is usually attributed for C content in marine zooplankton (e.g., [76]). The values obtained in the present study for L. conchilega larvae (~15% DM) were only a third to half of that, but within the 16–44% DM range recorded for polychaete by Parsons et al. [76]. It should be noted, however, that sample C and N contents and weight might have been affected by sampling larvae while they were still inside their tubes. The larval tube is composed of a thin organic layer, to which sand and shell fragments are attached at later stages [21]. An attempt was made to remove them, but we observed that it was not possible to do so without damaging the individuals or without the application of a sedative.

Conclusions

The present study documented preferential assimilation of FAs by L. conchilega larvae, which seem to be able to regulate their FA composition regardless of the FA profile of their food source. Bioconversion was also recorded in these individuals, which synthesized the PUFA 22:5(n-3) from dietary FA precursors. Our results show that dietary FAs are not transferred in a conservative (quantitative) manner, indicating that the concept of FATM is of limited applicability in the trophic study of lipid-poor L. conchilega larvae. This is probably also the case for other lipid-poor meroplanktonic larvae. Based on the results of this study and on data available from the literature, we propose that lipid homeostasis depends upon the lipid content of zooplanktonic organisms. Lipid-poor meroplanktonic larvae should display lipid homeostasis in order to maintain vital lipid-based body functions; holoplanktonic organisms with intermediate lipid levels will usually assimilate dietary lipids in a conservative manner and invest assimilated energy into reproduction; whereas lipid-rich holoplanktonic organisms will be semi-homeostatic, converting stored energy (FAs) into WE in order to survive starvation conditions during winter and to increase buoyancy.

Acknowledgments

We kindly thank the crew of the Aade research vessel (AWI-BAH) for providing us with zooplankton samples and the technicians from the Ecological Chemistry department at the AWI in Bremerhaven for the help with lipid analysis.

References

1. Umezawa Y, Tamaki A, Suzuki T, Takeuchi S, Yoshimizu C, Tayasu I. Phytoplankton as a principal diet for callianassid shrimp larvae in coastal waters, estimated from laboratory rearing and stable isotope analysis. Mar Ecol Prog Ser. 2018;592: 141–158.
- View Article
- Google Scholar
2. Hickel W. The mesozooplankton in the Wadden Sea of Sylt (North Sea). Helgoland wiss Meer. 1975;27: 254–262.
- View Article
- Google Scholar
3. Van Hoey G. Spatio-temporal variability within the macrobenthic Abra alba community, with emphasis on the structuring role of Lanice conchilega. 2006. Thesis, Ghent University, Belgium.
4. Strasser M, Pieloth U. Recolonization pattern of the polychaete Lanice conchilega on an intertidal sand flat following the severe winter of 1995/96. Helgol Mar Res. 2001;55: 176–181.
- View Article
- Google Scholar
5. Bhaud M. The two planktonic larval periods of Lanice conchilega (Pallas, 1766) annelida polychaeta, a peculiar example of the irreversibility of evolution. Ophelia. 1988;29: 141–152.
- View Article
- Google Scholar
6. Rabaut M. Lanice conchilega, fisheries and marine conservation–Towards an ecosystem approach to marine management. 2009. Thesis, Ghent University, Belgium.
7. Bhaud M, Cazaux CP. Buoyancy characteristics of Lanice conchilega (Pallas) larvae (Terebellidae). Implications for settlement. J Exp Mar Biol Ecol. 1990;141: 31–45.
- View Article
- Google Scholar
8. Braeckman U, Provoost P, Sabbe K, Soetaert K, Middelburg JJ, Vincx M, et al. Temporal dynamics in the diet of two marine polychaetes as inferred from fatty acid biomarkers. J Sea Res. 2012;68: 6–19.
- View Article
- Google Scholar
9. Bühr KJ. Suspension-feeding and assimilation efficiency in Lanice conchilega (Polychaeta). Mar Biol. 1976;38: 373–383.
- View Article
- Google Scholar
10. Bühr KJ, Winter JE. Distribution and maintenance of a Lanice conchilega association in the Weser Estuary (FRG), with special reference to the suspension-feeding behaviour of Lanice conchilega. In: Keegan BF, Ceidigh PO, Boaden PJS, editors. Biology of benthic organisms. Pergamon; 1977. pp. 101–113.
11. Dubois S, Orvain F, Marin-Leal JC, Ropert M, Lefebvre S. Small-scale spatial variability of food partitioning between cultivated oysters and associated suspension-feeding species, as revealed by stable isotopes. Mar Ecol Prog Ser. 2007;336: 151–160.
- View Article
- Google Scholar
12. Zühlke R, Blome D, Heinz van Bernem K, Dittmann S. Effects of the tube-building polychaete Lanice conchilega (Pallas) on benthic macrofauna and nematodes in an intertidal sandflat. Senck Marit. 1998;29: 131–138.
- View Article
- Google Scholar
13. Zühlke R. Polychaete tubes create ephemeral community patterns: Lanice conchilega (Pallas, 1766) associations studied over six years. J Sea Res. 2001;46: 261–272.
- View Article
- Google Scholar
14. Alves RMS, van Colen C, Vincx M, Vanaverbeke J, de Smet B, Guarini JM, et al. A case study on the growth of Lanice conchilega (Pallas, 1766) aggregations and their ecosystem engineering impact on sedimentary processes. J Exp Mar Biol Ecol. 2017;489: 15–23.
- View Article
- Google Scholar
15. Braber L, De Groot SJ. The food of five flatfish species (Pleuronectiformes) in the southern North Sea. Neth. J Sea Res. 1973;6: 163–172.
- View Article
- Google Scholar
16. De Smet B, Godet L, Fournier J, Desroy N, Jaffré M, Vincx M, et al. Feeding grounds for waders in the Bay of the Mont Saint-Michel (France): the Lanice conchilega reef serves as an oasis in the tidal flats. Mar Biol. 2013;160: 751–761.
- View Article
- Google Scholar
17. Godet L, Toupoint N, Olivier F, Fournier J, Retière C. Considering the functional value of common marine species as a conservation stake: The case of Sandmason Worm Lanice conchilega (Pallas 1766) (Annelida, Polychaeta) beds. Ambio. 2008;37: 347–355. pmid:18828280
- View Article
- PubMed/NCBI
- Google Scholar
18. Petersen B, Exo KM. Predation of waders and gulls on Lanice conchilega tidal flats in the Wadden Sea. Mar Ecol Prog Ser. 1999;178: 229–240.
- View Article
- Google Scholar
19. Rabaut M, Guilini K, Van Hoey G, Vincx M, Degraer S. A bio-engineered soft-bottom environment: The impact of Lanice conchilega on the benthic species-specific densities and community structure. Estuar Coast Shelf S. 2007;75: 525–536.
- View Article
- Google Scholar
20. Callaway R. Tube worms promote community change. Mar Ecol Prog Ser. 2006;308: 49–60.
- View Article
- Google Scholar
21. Callaway R. Juveniles stick to adults: recruitment of the tube-dwelling polychaete Lanice conchilega (Pallas, 1766). Hydrobiologia. 2003;503: 121–130.
- View Article
- Google Scholar
22. Ropert M, Dauvin JC. Renewal and accumulation of a Lanice conchilega (Pallas) population in the Baie des Veys, western Bay of Seine. Oceanol Acta. 2000;23: 529–546.
- View Article
- Google Scholar
23. Dalsgaard J, John MS, Kattner G, Müller-Navarra D, Hagen W. Fatty acid trophic markers in the pelagic marine environment. Adv Mar Biol. 2003;46: 225–340. pmid:14601414
- View Article
- PubMed/NCBI
- Google Scholar
24. Graeve M, Kattner G, Hagen W. Diet-induced changes in the fatty acid composition of Arctic herbivorous copepods: Experimental evidence of trophic markers. J Exp Mar Biol Ecol. 1994;182: 97–110.
- View Article
- Google Scholar
25. Lee RF, Nevenzel JC., Paffenhöfer GA. Importance of wax esters and other lipids in the marine food chain: phytoplankton and copepods. Mar Biol. 1971;9: 99–108.
- View Article
- Google Scholar
26. Boissonnot L, Niehoff B, Hagen W, Søreide JE, Graeve M. Lipid turnover reflects life-cycle strategies of small-sized Arctic copepods. J Plankton Res. 2016;38: 1420–1432.
- View Article
- Google Scholar
27. Graeve M, Albers C, Kattner G. Assimilation and biosynthesis of lipids in Arctic Calanus species based on feeding experiments with a ¹³C labelled diatom. J Exp Mar Biol Ecol. 2005;317: 109–125.
- View Article
- Google Scholar
28. Guilini K, van Oevelen D, Soetaert K, Middelburg JJ, Vanreusel A. Nutritional importance of benthic bacteria for deep-sea nematodes from the Arctic ice margin: Results of an isotope tracer experiment. Limnol Oceanogr. 2010;55: 1977–1989.
- View Article
- Google Scholar
29. Rix L, de Goeij JM, van Oevelen D, Struck U, Al-Horani FA, Wild C, et al. Reef sponges facilitate the transfer of coral-derived organic matter to their associated fauna via the sponge loop. Mar Ecol Prog Ser. 2018;589: 85–96.
- View Article
- Google Scholar
30. Wegener G, Bausch M, Holler T, Thang NM, Mollar XP, Kellermann MY, et al. Assessing sub-seafloor microbial activity by combined stable isotope probing with deuterated water and 13C-bicarbonate. Env Microbiol. 2012;14: 1517–1527.
- View Article
- Google Scholar
31. Meier-Augenstein W. Stable isotope analysis of fatty acids by gas chromatography-isotope ratio mass spectrometry. Anal Chim Acta. 2002;465: 63–79.
- View Article
- Google Scholar
32. De Troch M, Boeckx P, Cnudde C, Van Gansbeke D, Vanreusel A, Vincx M, et al. Bioconversion of fatty acids at the basis of marine food webs: insights from a compound-specific stable isotope analysis. Mar Ecol Prog Ser. 2012;465: 53–67.
- View Article
- Google Scholar
33. Middelburg JJ, Barranguet C, Boschker HTS, Herman PMJ, Moens T, Heip CHR. The fate of intertidal microphytobenthos carbon: An in situ ¹³C-labeling study. Limnol Oceanogr. 2000;45: 1224–1234.
- View Article
- Google Scholar
34. Guillard RRL. Culture of phytoplankton for feeding marine invertebrates. In: Smith WL, Chanley MH, editors. Culture of marine invertebrate animals. US: Springer; 1975. pp. 29–60.
35. Li HY, Lu Y, Zheng JW, Yang WD, Liu JS. Biochemical and genetic engineering of diatoms for polyunsaturated fatty acid biosynthesis. Mar Drugs. 2014;12: 153–166. pmid:24402175
- View Article
- PubMed/NCBI
- Google Scholar
36. R Core Team. 2018. R: A language and environment for statistical computing. R Foundation for Statistical Computing, Vienna, Austria. Available from https://www.R-project.org/.
37. Warton DI, Hui FKC. The arcsine is asinine: the analysis of proportions in ecology. Ecology. 2011;92: 3–10. pmid:21560670
- View Article
- PubMed/NCBI
- Google Scholar
38. Clarke KR, Gorley RN. PRIMER v7: User Manual/Tutorial. PRIMER-E, Plymouth; 2015.
39. Harms J, Anger K, Klaus S, Seeger B. Nutritional effects on ingestion rate, digestive enzyme activity, growth, and biochemical composition of Hyas araneus L. (Decapoda: Majidae) larvae. J Exp Mar Biol Ecol. 1991;145: 233–265.
- View Article
- Google Scholar
40. Harms J, Meyer-Harms B, Dawirs RR, Anger K. Growth and physiology of Carcinus maenas (Decapoda, Portunidae) larvae in the field and in laboratory experiments. Mar Ecol Prog Ser. 1994;108: 107–118.
- View Article
- Google Scholar
41. His E, Maurer D. Shell growth and gross biochemical bomposition of oyster larvae (Crassostrea gigas) in the field. Aquaculture. 1988;69: 185–194.
- View Article
- Google Scholar
42. Müller-Navarra DC. The nutritional importance of polyunsaturated fatty acids and their use as trophic markers for herbivorous zooplankton: Does it contradict? Arch Hydrobiol. 2006;167: 501–513.
- View Article
- Google Scholar
43. Brett MT, Müller-Navarra DC. The role of highly unsaturated fatty acids in aquatic foodweb processes. Freshwater Biol. 1997;38: 483–499.
- View Article
- Google Scholar
44. Jónasdóttir SH, Visser AW, Jespersen C. Assessing the role of food quality in the production and hatching of Temora longicornis eggs. Mar Ecol Prog Ser. 2009;382: 139–150.
- View Article
- Google Scholar
45. Thor P, Koski M, Tang KW, Jónasdóttir SH. Supplemental effects of diet mixing on absorption of ingested organic carbon in the marine copepod Acartia tonsa. Mar Ecol Prog Ser. 2007;331: 131–138.
- View Article
- Google Scholar
46. Johnston PV. Dietary fat, eicosanoids, and immunity. Adv Lipid Res. 1985;21:103–141. pmid:3895844
- View Article
- PubMed/NCBI
- Google Scholar
47. Budge SM, Iverson SJ, Koopman HN. Studying trophic ecology in marine ecosystems using fatty acids: a primer on analysis and interpretation. J Exp Biol. 2006;214: 3790–3800.
- View Article
- Google Scholar
48. Jeffreys RM, Wolff GA, Murty SJ. The trophic ecology of key megafaunal species at the Pakistan Margin: Evidence from stable isotopes and lipid biomarkers. Deep-Sea Res. 2009;56: 1816–1833.
- View Article
- Google Scholar
49. Budge SM, Wang SW, Hollmén TE, Wooller MJ. Carbon isotopic fractionation in eider adipose tissue varies with fatty acid structure: implications for trophic studies. Mar Mammal Sci. 2011;22: 759–801.
- View Article
- Google Scholar
50. Pond DW, Allen CE, Bell MV, Van Dover CL, Fallick AE, Dixon DR, et al. Origins of long-chain polyunsaturated fatty acids in the hydrothermal vent worms Ridgea piscesae and Protis hydrothermica. Mar Ecol Prog Ser. 2002;225: 219–226.
- View Article
- Google Scholar
51. Mansour MP, Holdsworth DG, Forbes SE, Macleod CK, Volkman JK. High contents of 24:6(n-3) and 20:1(n-13) fatty acids in the brittle star Amphiura elandiformis from Tasmanian coastal sediments. Biochem Syst Ecol. 2005;33: 659–674.
- View Article
- Google Scholar
52. Kreibich T, Saborowski R, Hagen W, Niehoff B. Influence of short-term nutritional variations on digestive enzyme and fatty acid patterns of the calanoid copepod Temora longicornis. J Exp Mar Biol Ecol. 2011;407: 182–189.
- View Article
- Google Scholar
53. Mann R, Gallager SM. Physiological and biochemical energetics of larvae of Teredo nevalis L. and Bankia gouldi. (Bartsch) (Bivalvia: Teredinidae). J Exp Mar Biol Ecol. 1985;85: 211–228.
- View Article
- Google Scholar
54. Whyte JNC, Bourne N, Ginther NG, Hodgson CA. Compositional changes in the larva to juvenile development of the scallop Crassadoma gigantea (Gray). J Exp Mar Biol Ecol. 1992;163: 13–29.
- View Article
- Google Scholar
55. Orellana MC, Cancino JM, Hughes RN. Is settlement in lecithotrophic bryozoan larvae constrained by energy reserves? In: Gordon DP, Smith AM, Grant-Mackie JA, editors. Bryozoans in space and time. Proceedings of the 10th International Bryozoology Conference; Wellington, New Zealand. National Institute of Water and Atmospheric Research Ltd., Wellington; 1996. 442p.
56. Holland DL, Walker G. The biochemical composition of the cypris larva of the barnacle Balanus balanoides L. J Const Int Explor Mer. 1975;36: 162–165.
- View Article
- Google Scholar
57. Lourenço S, Roura A, Fernández-Reiriz MJ, Narciso L, González AF. Feeding relationship between Octopus vulgaris (Cuvier, 1797) early life-cycle stages and their prey in the Western Iberian upwelling system: Correlation of reciprocal lipid and fatty acid contents. Front Physiol. 2017;8: 1–11. pmid:28154536
- View Article
- PubMed/NCBI
- Google Scholar
58. Thatje S, Lovrich GA, Torres G, Hagen W, Anger K. Changes in biomass, lipid, fatty acid and elemental composition during the abbreviated larval development of the subantarctic shrimp Campylonotus vagans. J Exp Mar Biol Ecol. 2004;301: 159–174.
- View Article
- Google Scholar
59. Rotllant G, Anger K, Durfort M, Sardà F. Elemental and biochemical composition of Nephrops norvegicus (Linnaeus 1758) larvae from the Mediterranean and Irish Seas. Helgol Mar Res. 2004;58: 206–210.
- View Article
- Google Scholar
60. Wang M, O’Rorke R, Waite AM, Beckley LE, Thompson P, Jeffs AG. Fatty acid profiles of phyllosoma larvae of western rock lobster (Panulirus cygnus) in cyclonic and anticyclonic eddies of the Leeuwin Current off Western Australia. Progr Oceanogr. 2014;122: 153–162.
- View Article
- Google Scholar
61. Reinhardt SB, Van Vleet ES. Lipid composition of twenty-two species of Antarctic midwater zooplankton and fish. Mar Biol. 1986;91: 149–159.
- View Article
- Google Scholar
62. Vázquez R, González S, Rodríguez A, Mourente G. Biochemical composition and fatty acid content of fertilized eggs, yolk sac stage larvae and first-feeding larvae of the Senegal sole (Solea senegalensis Kaup). Aquaculture. 1994;119: 273–286.
- View Article
- Google Scholar
63. Lee RF. Lipids of zooplankton from Bute Inlet, British Columbia. J Fish Res Board Can. 1974b;31: 1577–1582.
- View Article
- Google Scholar
64. Goswami SC, Rao TSS, Matondkar SGP. Biochemical composition of zooplankton from the Andaman Sea. Indian J Mar Sci. 1981;10: 296–300.
- View Article
- Google Scholar
65. Lee RF. Lipids of Arctic zooplankton. Comp Biochem Physiol. 1975;51: 263–266.
- View Article
- Google Scholar
66. Antony G, Antony A. Biochemical composition of Lucifer hanseni from the Cochin estuary. Indian J Fish. 2001;48: 41–47.
- View Article
- Google Scholar
67. Lee RF. Lipid composition of the copepod Calanus hyperboreas from the Arctic Ocean. Changes with depth and season. Mar Biol. 1974a;26: 313–318.
- View Article
- Google Scholar
68. Diel S, Tande K. Does the spawning of Calanus finmarchicus in high latitudes follow a reproducible pattern? Mar Biol. 1992;113: 21–31.
- View Article
- Google Scholar
69. Scott CL, Kwasniewski S, Falk-Petersen S, Sargent JR. Lipids and life strategies of Calanus finmarchicus, Calanus glacialis and Calanus hyperboreus in late autumn, Kongsfjorden, Svalbard. Polar Biol. 2000;23: 510–516.
- View Article
- Google Scholar
70. Hagen W, Kattner G, Terbrüggen A, Van Vleet ES. Lipid metabolism of the Antarctic krill Euphausia superba and its ecological implication. Mar Biol. 2001;139: 95–104.
- View Article
- Google Scholar
71. Ben-David-Zaslow R, Benayahu Y. Biochemical composition, metabolism, and amino acid transport in planula-larvae of the soft coral Heteroxenia fuscescens. J Exp Zool. 2000;287: 401–412. pmid:11074452
- View Article
- PubMed/NCBI
- Google Scholar
72. Harii S, Nadaoka K, Yamamoto M, Iwao K. Temporal changes in settlement, lipid content and lipid composition of larvae of the spawning hermatypic coral Acropora tenuis. Mar Ecol Prog Ser. 2007;346: 89–96.
- View Article
- Google Scholar
73. Sargent JR, Falk-Petersen S. The lipid biochemistry of calanoid copepods. Hydrobiologia. 1988;167/168: 101–114.
- View Article
- Google Scholar
74. Leonard AE, Pereira SL, Sprecher H, Huang YS. Elongation of long-chain fatty acids. Prog Lipid Res. 2004;43: 36–54. pmid:14636670
- View Article
- PubMed/NCBI
- Google Scholar
75. Franco-Santos RM, Auel H, Boersma M, De Troch M, Meunier CL, Niehoff B. Bioenergetics of the copepod Temora longicornis under different nutrient regimes. J Plankton Res. 2018;40: 420–435.
- View Article
- Google Scholar
76. Parsons TR, Takahashi M, Hargrave B. Chemical composition. In: Parsons TR, Takahashi M, Hargrave B, editors. Biological Oceanographic Processes. 2nd ed. Oxford: Pergamon Press; 1977. pp. 40–64.

[ref1] 1. Umezawa Y, Tamaki A, Suzuki T, Takeuchi S, Yoshimizu C, Tayasu I. Phytoplankton as a principal diet for callianassid shrimp larvae in coastal waters, estimated from laboratory rearing and stable isotope analysis. Mar Ecol Prog Ser. 2018;592: 141–158.
View Article
Google Scholar

[2] View Article

[3] Google Scholar

[ref2] 2. Hickel W. The mesozooplankton in the Wadden Sea of Sylt (North Sea). Helgoland wiss Meer. 1975;27: 254–262.
View Article
Google Scholar

[5] View Article

[6] Google Scholar

[ref3] 3. Van Hoey G. Spatio-temporal variability within the macrobenthic Abra alba community, with emphasis on the structuring role of Lanice conchilega. 2006. Thesis, Ghent University, Belgium.

[ref4] 4. Strasser M, Pieloth U. Recolonization pattern of the polychaete Lanice conchilega on an intertidal sand flat following the severe winter of 1995/96. Helgol Mar Res. 2001;55: 176–181.
View Article
Google Scholar

[9] View Article

[10] Google Scholar

[ref5] 5. Bhaud M. The two planktonic larval periods of Lanice conchilega (Pallas, 1766) annelida polychaeta, a peculiar example of the irreversibility of evolution. Ophelia. 1988;29: 141–152.
View Article
Google Scholar

[12] View Article

[13] Google Scholar

[ref6] 6. Rabaut M. Lanice conchilega, fisheries and marine conservation–Towards an ecosystem approach to marine management. 2009. Thesis, Ghent University, Belgium.

[ref7] 7. Bhaud M, Cazaux CP. Buoyancy characteristics of Lanice conchilega (Pallas) larvae (Terebellidae). Implications for settlement. J Exp Mar Biol Ecol. 1990;141: 31–45.
View Article
Google Scholar

[16] View Article

[17] Google Scholar

[ref8] 8. Braeckman U, Provoost P, Sabbe K, Soetaert K, Middelburg JJ, Vincx M, et al. Temporal dynamics in the diet of two marine polychaetes as inferred from fatty acid biomarkers. J Sea Res. 2012;68: 6–19.
View Article
Google Scholar

[19] View Article

[20] Google Scholar

[ref9] 9. Bühr KJ. Suspension-feeding and assimilation efficiency in Lanice conchilega (Polychaeta). Mar Biol. 1976;38: 373–383.
View Article
Google Scholar

[22] View Article

[23] Google Scholar

[ref10] 10. Bühr KJ, Winter JE. Distribution and maintenance of a Lanice conchilega association in the Weser Estuary (FRG), with special reference to the suspension-feeding behaviour of Lanice conchilega. In: Keegan BF, Ceidigh PO, Boaden PJS, editors. Biology of benthic organisms. Pergamon; 1977. pp. 101–113.

[ref11] 11. Dubois S, Orvain F, Marin-Leal JC, Ropert M, Lefebvre S. Small-scale spatial variability of food partitioning between cultivated oysters and associated suspension-feeding species, as revealed by stable isotopes. Mar Ecol Prog Ser. 2007;336: 151–160.
View Article
Google Scholar

[26] View Article

[27] Google Scholar

[ref12] 12. Zühlke R, Blome D, Heinz van Bernem K, Dittmann S. Effects of the tube-building polychaete Lanice conchilega (Pallas) on benthic macrofauna and nematodes in an intertidal sandflat. Senck Marit. 1998;29: 131–138.
View Article
Google Scholar

[29] View Article

[30] Google Scholar

[ref13] 13. Zühlke R. Polychaete tubes create ephemeral community patterns: Lanice conchilega (Pallas, 1766) associations studied over six years. J Sea Res. 2001;46: 261–272.
View Article
Google Scholar

[32] View Article

[33] Google Scholar

[ref14] 14. Alves RMS, van Colen C, Vincx M, Vanaverbeke J, de Smet B, Guarini JM, et al. A case study on the growth of Lanice conchilega (Pallas, 1766) aggregations and their ecosystem engineering impact on sedimentary processes. J Exp Mar Biol Ecol. 2017;489: 15–23.
View Article
Google Scholar

[35] View Article

[36] Google Scholar

[ref15] 15. Braber L, De Groot SJ. The food of five flatfish species (Pleuronectiformes) in the southern North Sea. Neth. J Sea Res. 1973;6: 163–172.
View Article
Google Scholar

[38] View Article

[39] Google Scholar

[ref16] 16. De Smet B, Godet L, Fournier J, Desroy N, Jaffré M, Vincx M, et al. Feeding grounds for waders in the Bay of the Mont Saint-Michel (France): the Lanice conchilega reef serves as an oasis in the tidal flats. Mar Biol. 2013;160: 751–761.
View Article
Google Scholar

[41] View Article

[42] Google Scholar

[ref17] 17. Godet L, Toupoint N, Olivier F, Fournier J, Retière C. Considering the functional value of common marine species as a conservation stake: The case of Sandmason Worm Lanice conchilega (Pallas 1766) (Annelida, Polychaeta) beds. Ambio. 2008;37: 347–355. pmid:18828280
View Article
PubMed/NCBI
Google Scholar

[44] View Article

[45] PubMed/NCBI

[46] Google Scholar

[ref18] 18. Petersen B, Exo KM. Predation of waders and gulls on Lanice conchilega tidal flats in the Wadden Sea. Mar Ecol Prog Ser. 1999;178: 229–240.
View Article
Google Scholar

[48] View Article

[49] Google Scholar

[ref19] 19. Rabaut M, Guilini K, Van Hoey G, Vincx M, Degraer S. A bio-engineered soft-bottom environment: The impact of Lanice conchilega on the benthic species-specific densities and community structure. Estuar Coast Shelf S. 2007;75: 525–536.
View Article
Google Scholar

[51] View Article

[52] Google Scholar

[ref20] 20. Callaway R. Tube worms promote community change. Mar Ecol Prog Ser. 2006;308: 49–60.
View Article
Google Scholar

[54] View Article

[55] Google Scholar

[ref21] 21. Callaway R. Juveniles stick to adults: recruitment of the tube-dwelling polychaete Lanice conchilega (Pallas, 1766). Hydrobiologia. 2003;503: 121–130.
View Article
Google Scholar

[57] View Article

[58] Google Scholar

[ref22] 22. Ropert M, Dauvin JC. Renewal and accumulation of a Lanice conchilega (Pallas) population in the Baie des Veys, western Bay of Seine. Oceanol Acta. 2000;23: 529–546.
View Article
Google Scholar

[60] View Article

[61] Google Scholar

[ref23] 23. Dalsgaard J, John MS, Kattner G, Müller-Navarra D, Hagen W. Fatty acid trophic markers in the pelagic marine environment. Adv Mar Biol. 2003;46: 225–340. pmid:14601414
View Article
PubMed/NCBI
Google Scholar

[63] View Article

[64] PubMed/NCBI

[65] Google Scholar

[ref24] 24. Graeve M, Kattner G, Hagen W. Diet-induced changes in the fatty acid composition of Arctic herbivorous copepods: Experimental evidence of trophic markers. J Exp Mar Biol Ecol. 1994;182: 97–110.
View Article
Google Scholar

[67] View Article

[68] Google Scholar

[ref25] 25. Lee RF, Nevenzel JC., Paffenhöfer GA. Importance of wax esters and other lipids in the marine food chain: phytoplankton and copepods. Mar Biol. 1971;9: 99–108.
View Article
Google Scholar

[70] View Article

[71] Google Scholar

[ref26] 26. Boissonnot L, Niehoff B, Hagen W, Søreide JE, Graeve M. Lipid turnover reflects life-cycle strategies of small-sized Arctic copepods. J Plankton Res. 2016;38: 1420–1432.
View Article
Google Scholar

[73] View Article

[74] Google Scholar

[ref27] 27. Graeve M, Albers C, Kattner G. Assimilation and biosynthesis of lipids in Arctic Calanus species based on feeding experiments with a ¹³C labelled diatom. J Exp Mar Biol Ecol. 2005;317: 109–125.
View Article
Google Scholar

[76] View Article

[77] Google Scholar

[ref28] 28. Guilini K, van Oevelen D, Soetaert K, Middelburg JJ, Vanreusel A. Nutritional importance of benthic bacteria for deep-sea nematodes from the Arctic ice margin: Results of an isotope tracer experiment. Limnol Oceanogr. 2010;55: 1977–1989.
View Article
Google Scholar

[79] View Article

[80] Google Scholar

[ref29] 29. Rix L, de Goeij JM, van Oevelen D, Struck U, Al-Horani FA, Wild C, et al. Reef sponges facilitate the transfer of coral-derived organic matter to their associated fauna via the sponge loop. Mar Ecol Prog Ser. 2018;589: 85–96.
View Article
Google Scholar

[82] View Article

[83] Google Scholar

[ref30] 30. Wegener G, Bausch M, Holler T, Thang NM, Mollar XP, Kellermann MY, et al. Assessing sub-seafloor microbial activity by combined stable isotope probing with deuterated water and 13C-bicarbonate. Env Microbiol. 2012;14: 1517–1527.
View Article
Google Scholar

[85] View Article

[86] Google Scholar

[ref31] 31. Meier-Augenstein W. Stable isotope analysis of fatty acids by gas chromatography-isotope ratio mass spectrometry. Anal Chim Acta. 2002;465: 63–79.
View Article
Google Scholar

[88] View Article

[89] Google Scholar

[ref32] 32. De Troch M, Boeckx P, Cnudde C, Van Gansbeke D, Vanreusel A, Vincx M, et al. Bioconversion of fatty acids at the basis of marine food webs: insights from a compound-specific stable isotope analysis. Mar Ecol Prog Ser. 2012;465: 53–67.
View Article
Google Scholar

[91] View Article

[92] Google Scholar

[ref33] 33. Middelburg JJ, Barranguet C, Boschker HTS, Herman PMJ, Moens T, Heip CHR. The fate of intertidal microphytobenthos carbon: An in situ ¹³C-labeling study. Limnol Oceanogr. 2000;45: 1224–1234.
View Article
Google Scholar

[94] View Article

[95] Google Scholar

[ref34] 34. Guillard RRL. Culture of phytoplankton for feeding marine invertebrates. In: Smith WL, Chanley MH, editors. Culture of marine invertebrate animals. US: Springer; 1975. pp. 29–60.

[ref35] 35. Li HY, Lu Y, Zheng JW, Yang WD, Liu JS. Biochemical and genetic engineering of diatoms for polyunsaturated fatty acid biosynthesis. Mar Drugs. 2014;12: 153–166. pmid:24402175
View Article
PubMed/NCBI
Google Scholar

[98] View Article

[99] PubMed/NCBI

[100] Google Scholar

[ref36] 36. R Core Team. 2018. R: A language and environment for statistical computing. R Foundation for Statistical Computing, Vienna, Austria. Available from https://www.R-project.org/.

[ref37] 37. Warton DI, Hui FKC. The arcsine is asinine: the analysis of proportions in ecology. Ecology. 2011;92: 3–10. pmid:21560670
View Article
PubMed/NCBI
Google Scholar

[103] View Article

[104] PubMed/NCBI

[105] Google Scholar

[ref38] 38. Clarke KR, Gorley RN. PRIMER v7: User Manual/Tutorial. PRIMER-E, Plymouth; 2015.

[ref39] 39. Harms J, Anger K, Klaus S, Seeger B. Nutritional effects on ingestion rate, digestive enzyme activity, growth, and biochemical composition of Hyas araneus L. (Decapoda: Majidae) larvae. J Exp Mar Biol Ecol. 1991;145: 233–265.
View Article
Google Scholar

[108] View Article

[109] Google Scholar

[ref40] 40. Harms J, Meyer-Harms B, Dawirs RR, Anger K. Growth and physiology of Carcinus maenas (Decapoda, Portunidae) larvae in the field and in laboratory experiments. Mar Ecol Prog Ser. 1994;108: 107–118.
View Article
Google Scholar

[111] View Article

[112] Google Scholar

[ref41] 41. His E, Maurer D. Shell growth and gross biochemical bomposition of oyster larvae (Crassostrea gigas) in the field. Aquaculture. 1988;69: 185–194.
View Article
Google Scholar

[114] View Article

[115] Google Scholar

[ref42] 42. Müller-Navarra DC. The nutritional importance of polyunsaturated fatty acids and their use as trophic markers for herbivorous zooplankton: Does it contradict? Arch Hydrobiol. 2006;167: 501–513.
View Article
Google Scholar

[117] View Article

[118] Google Scholar

[ref43] 43. Brett MT, Müller-Navarra DC. The role of highly unsaturated fatty acids in aquatic foodweb processes. Freshwater Biol. 1997;38: 483–499.
View Article
Google Scholar

[120] View Article

[121] Google Scholar

[ref44] 44. Jónasdóttir SH, Visser AW, Jespersen C. Assessing the role of food quality in the production and hatching of Temora longicornis eggs. Mar Ecol Prog Ser. 2009;382: 139–150.
View Article
Google Scholar

[123] View Article

[124] Google Scholar

[ref45] 45. Thor P, Koski M, Tang KW, Jónasdóttir SH. Supplemental effects of diet mixing on absorption of ingested organic carbon in the marine copepod Acartia tonsa. Mar Ecol Prog Ser. 2007;331: 131–138.
View Article
Google Scholar

[126] View Article

[127] Google Scholar

[ref46] 46. Johnston PV. Dietary fat, eicosanoids, and immunity. Adv Lipid Res. 1985;21:103–141. pmid:3895844
View Article
PubMed/NCBI
Google Scholar

[129] View Article

[130] PubMed/NCBI

[131] Google Scholar

[ref47] 47. Budge SM, Iverson SJ, Koopman HN. Studying trophic ecology in marine ecosystems using fatty acids: a primer on analysis and interpretation. J Exp Biol. 2006;214: 3790–3800.
View Article
Google Scholar

[133] View Article

[134] Google Scholar

[ref48] 48. Jeffreys RM, Wolff GA, Murty SJ. The trophic ecology of key megafaunal species at the Pakistan Margin: Evidence from stable isotopes and lipid biomarkers. Deep-Sea Res. 2009;56: 1816–1833.
View Article
Google Scholar

[136] View Article

[137] Google Scholar

[ref49] 49. Budge SM, Wang SW, Hollmén TE, Wooller MJ. Carbon isotopic fractionation in eider adipose tissue varies with fatty acid structure: implications for trophic studies. Mar Mammal Sci. 2011;22: 759–801.
View Article
Google Scholar

[139] View Article

[140] Google Scholar

[ref50] 50. Pond DW, Allen CE, Bell MV, Van Dover CL, Fallick AE, Dixon DR, et al. Origins of long-chain polyunsaturated fatty acids in the hydrothermal vent worms Ridgea piscesae and Protis hydrothermica. Mar Ecol Prog Ser. 2002;225: 219–226.
View Article
Google Scholar

[142] View Article

[143] Google Scholar

[ref51] 51. Mansour MP, Holdsworth DG, Forbes SE, Macleod CK, Volkman JK. High contents of 24:6(n-3) and 20:1(n-13) fatty acids in the brittle star Amphiura elandiformis from Tasmanian coastal sediments. Biochem Syst Ecol. 2005;33: 659–674.
View Article
Google Scholar

[145] View Article

[146] Google Scholar

[ref52] 52. Kreibich T, Saborowski R, Hagen W, Niehoff B. Influence of short-term nutritional variations on digestive enzyme and fatty acid patterns of the calanoid copepod Temora longicornis. J Exp Mar Biol Ecol. 2011;407: 182–189.
View Article
Google Scholar

[148] View Article

[149] Google Scholar

[ref53] 53. Mann R, Gallager SM. Physiological and biochemical energetics of larvae of Teredo nevalis L. and Bankia gouldi. (Bartsch) (Bivalvia: Teredinidae). J Exp Mar Biol Ecol. 1985;85: 211–228.
View Article
Google Scholar

[151] View Article

[152] Google Scholar

[ref54] 54. Whyte JNC, Bourne N, Ginther NG, Hodgson CA. Compositional changes in the larva to juvenile development of the scallop Crassadoma gigantea (Gray). J Exp Mar Biol Ecol. 1992;163: 13–29.
View Article
Google Scholar

[154] View Article

[155] Google Scholar

[ref55] 55. Orellana MC, Cancino JM, Hughes RN. Is settlement in lecithotrophic bryozoan larvae constrained by energy reserves? In: Gordon DP, Smith AM, Grant-Mackie JA, editors. Bryozoans in space and time. Proceedings of the 10th International Bryozoology Conference; Wellington, New Zealand. National Institute of Water and Atmospheric Research Ltd., Wellington; 1996. 442p.

[ref56] 56. Holland DL, Walker G. The biochemical composition of the cypris larva of the barnacle Balanus balanoides L. J Const Int Explor Mer. 1975;36: 162–165.
View Article
Google Scholar

[158] View Article

[159] Google Scholar

[ref57] 57. Lourenço S, Roura A, Fernández-Reiriz MJ, Narciso L, González AF. Feeding relationship between Octopus vulgaris (Cuvier, 1797) early life-cycle stages and their prey in the Western Iberian upwelling system: Correlation of reciprocal lipid and fatty acid contents. Front Physiol. 2017;8: 1–11. pmid:28154536
View Article
PubMed/NCBI
Google Scholar

[161] View Article

[162] PubMed/NCBI

[163] Google Scholar

[ref58] 58. Thatje S, Lovrich GA, Torres G, Hagen W, Anger K. Changes in biomass, lipid, fatty acid and elemental composition during the abbreviated larval development of the subantarctic shrimp Campylonotus vagans. J Exp Mar Biol Ecol. 2004;301: 159–174.
View Article
Google Scholar

[165] View Article

[166] Google Scholar

[ref59] 59. Rotllant G, Anger K, Durfort M, Sardà F. Elemental and biochemical composition of Nephrops norvegicus (Linnaeus 1758) larvae from the Mediterranean and Irish Seas. Helgol Mar Res. 2004;58: 206–210.
View Article
Google Scholar

[168] View Article

[169] Google Scholar

[ref60] 60. Wang M, O’Rorke R, Waite AM, Beckley LE, Thompson P, Jeffs AG. Fatty acid profiles of phyllosoma larvae of western rock lobster (Panulirus cygnus) in cyclonic and anticyclonic eddies of the Leeuwin Current off Western Australia. Progr Oceanogr. 2014;122: 153–162.
View Article
Google Scholar

[171] View Article

[172] Google Scholar

[ref61] 61. Reinhardt SB, Van Vleet ES. Lipid composition of twenty-two species of Antarctic midwater zooplankton and fish. Mar Biol. 1986;91: 149–159.
View Article
Google Scholar

[174] View Article

[175] Google Scholar

[ref62] 62. Vázquez R, González S, Rodríguez A, Mourente G. Biochemical composition and fatty acid content of fertilized eggs, yolk sac stage larvae and first-feeding larvae of the Senegal sole (Solea senegalensis Kaup). Aquaculture. 1994;119: 273–286.
View Article
Google Scholar

[177] View Article

[178] Google Scholar

[ref63] 63. Lee RF. Lipids of zooplankton from Bute Inlet, British Columbia. J Fish Res Board Can. 1974b;31: 1577–1582.
View Article
Google Scholar

[180] View Article

[181] Google Scholar

[ref64] 64. Goswami SC, Rao TSS, Matondkar SGP. Biochemical composition of zooplankton from the Andaman Sea. Indian J Mar Sci. 1981;10: 296–300.
View Article
Google Scholar

[183] View Article

[184] Google Scholar

[ref65] 65. Lee RF. Lipids of Arctic zooplankton. Comp Biochem Physiol. 1975;51: 263–266.
View Article
Google Scholar

[186] View Article

[187] Google Scholar

[ref66] 66. Antony G, Antony A. Biochemical composition of Lucifer hanseni from the Cochin estuary. Indian J Fish. 2001;48: 41–47.
View Article
Google Scholar

[189] View Article

[190] Google Scholar

[ref67] 67. Lee RF. Lipid composition of the copepod Calanus hyperboreas from the Arctic Ocean. Changes with depth and season. Mar Biol. 1974a;26: 313–318.
View Article
Google Scholar

[192] View Article

[193] Google Scholar

[ref68] 68. Diel S, Tande K. Does the spawning of Calanus finmarchicus in high latitudes follow a reproducible pattern? Mar Biol. 1992;113: 21–31.
View Article
Google Scholar

[195] View Article

[196] Google Scholar

[ref69] 69. Scott CL, Kwasniewski S, Falk-Petersen S, Sargent JR. Lipids and life strategies of Calanus finmarchicus, Calanus glacialis and Calanus hyperboreus in late autumn, Kongsfjorden, Svalbard. Polar Biol. 2000;23: 510–516.
View Article
Google Scholar

[198] View Article

[199] Google Scholar

[ref70] 70. Hagen W, Kattner G, Terbrüggen A, Van Vleet ES. Lipid metabolism of the Antarctic krill Euphausia superba and its ecological implication. Mar Biol. 2001;139: 95–104.
View Article
Google Scholar

[201] View Article

[202] Google Scholar

[ref71] 71. Ben-David-Zaslow R, Benayahu Y. Biochemical composition, metabolism, and amino acid transport in planula-larvae of the soft coral Heteroxenia fuscescens. J Exp Zool. 2000;287: 401–412. pmid:11074452
View Article
PubMed/NCBI
Google Scholar

[204] View Article

[205] PubMed/NCBI

[206] Google Scholar

[ref72] 72. Harii S, Nadaoka K, Yamamoto M, Iwao K. Temporal changes in settlement, lipid content and lipid composition of larvae of the spawning hermatypic coral Acropora tenuis. Mar Ecol Prog Ser. 2007;346: 89–96.
View Article
Google Scholar

[208] View Article

[209] Google Scholar

[ref73] 73. Sargent JR, Falk-Petersen S. The lipid biochemistry of calanoid copepods. Hydrobiologia. 1988;167/168: 101–114.
View Article
Google Scholar

[211] View Article

[212] Google Scholar

[ref74] 74. Leonard AE, Pereira SL, Sprecher H, Huang YS. Elongation of long-chain fatty acids. Prog Lipid Res. 2004;43: 36–54. pmid:14636670
View Article
PubMed/NCBI
Google Scholar

[214] View Article

[215] PubMed/NCBI

[216] Google Scholar

[ref75] 75. Franco-Santos RM, Auel H, Boersma M, De Troch M, Meunier CL, Niehoff B. Bioenergetics of the copepod Temora longicornis under different nutrient regimes. J Plankton Res. 2018;40: 420–435.
View Article
Google Scholar

[218] View Article

[219] Google Scholar

[ref76] 76. Parsons TR, Takahashi M, Hargrave B. Chemical composition. In: Parsons TR, Takahashi M, Hargrave B, editors. Biological Oceanographic Processes. 2nd ed. Oxford: Pergamon Press; 1977. pp. 40–64.

Figures

Abstract

Introduction

Method

Field sampling

Algae culture

Experimental design

Carbon, nitrogen and fatty acid content analyses

Compound-specific stable isotope analysis (CSIA)

Statistical analyses

Results

Algal composition and enrichment

Larval composition and enrichment

Origin of FAs in the larvae

Lipid C assimilation and turnover in larvae

Discussion

Lipid composition, homeostasis, and bioconversion in L. conchilega larvae

Lipid homeostasis in lipid-poor and lipid-rich marine planktonic organisms

The applicability of the FATM concept

Experimental conditions

Conclusions

Acknowledgments

References