Next Article in Journal
New Kunitz-Type HCRG Polypeptides from the Sea Anemone Heteractis crispa
Next Article in Special Issue
Interactions between Carotenoids from Marine Bacteria and Other Micronutrients: Impact on Stability and Antioxidant Activity
Previous Article in Journal
Diatom-Specific Oligosaccharide and Polysaccharide Structures Help to Unravel Biosynthetic Capabilities in Diatoms
Previous Article in Special Issue
A Phospholipid-Protein Complex from Antarctic Krill Reduced Plasma Homocysteine Levels and Increased Plasma Trimethylamine-N-Oxide (TMAO) and Carnitine Levels in Male Wistar Rats
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Marine Drugs
Volume 13
Issue 10
10.3390/md13106019
marinedrugs-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles thumb_up
...
Endorse textsms
...
Comment
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Article

Ocean Warming and CO2-Induced Acidification Impact the Lipid Content of a Marine Predatory Gastropod

by
Roselyn Valles-Regino
1,
Rick Tate
2,
Brendan Kelaher
2,
Dale Savins
3,
Ashley Dowell
3 and
Kirsten Benkendorff
1,*
1
Marine Ecology Research Center, School of Environment, Science and Engineering, Southern Cross University, P.O. Box 157, Lismore, NSW 2480, Australia
2
National Marine Science Centre, Southern Cross University, Coffs Harbour, NSW 2450, Australia
3
Southern Cross Plant Science, Southern Cross University, Lismore, NSW 2480, Australia
*
Author to whom correspondence should be addressed.
Mar. Drugs 2015, 13(10), 6019-6037; https://doi.org/10.3390/md13106019
Submission received: 12 June 2015 / Revised: 8 September 2015 / Accepted: 14 September 2015 / Published: 24 September 2015
(This article belongs to the Special Issue Marine Lipids)
Browse Figures
Versions Notes

Abstract

:
Ocean warming and acidification are current global environmental challenges impacting aquatic organisms. A shift in conditions outside the optimal environmental range for marine species is likely to generate stress that could impact metabolic activity, with consequences for the biosynthesis of marine lipids. The aim of this study was to investigate differences in the lipid content of Dicathais orbita exposed to current and predicted future climate change scenarios. The whelks were exposed to a combination of temperature and CO2-induced acidification treatments in controlled flowthrough seawater mesocosms for 35 days. Under current conditions, D. orbita foot tissue has an average of 6 mg lipid/g tissue, but at predicted future ocean temperatures, the total lipid content dropped significantly, to almost half. The fatty acid composition is dominated by polyunsaturated fatty acids (PUFA 52%) with an n-3:6 fatty acid ratio of almost 2, which remains unchanged under future ocean conditions. However, we detected an interactive effect of temperature and pCO2 on the % PUFAs and n-3 and n-6 fatty acids were significantly reduced by elevated water temperature, while both the saturated and monounsaturated fatty acids were significantly reduced under increased pCO2 acidifying conditions. The present study indicates the potential for relatively small predicted changes in ocean conditions to reduce lipid reserves and alter the fatty acid composition of a predatory marine mollusc. This has potential implications for the growth and survivorship of whelks under future conditions, but only minimal implications for human consumption of D. orbita as nutritional seafood are predicted.

Graphical Abstract

1. Introduction

Climate change is one of the major environmental challenges to humankind and all other life forms on Earth. The consequences of the global climate change have been widely reported [1,2,3] and are likely to worsen over the coming decades. Global sea surface temperatures are projected to increase by 2–4 °C towards the end of the 21st century [1], while the surface ocean pH is predicted to decrease by 0.14–0.35 units, adding to the present decrease of 0.1 units that has already occurred since pre-industrial times [1,2]. The drop in pH is due to the absorption of increasingly greater CO2 concentrations into the oceans as a result of the rising partial pressure of atmospheric CO2 (pCO2) from anthropogenic activities, such as burning of fossil fuels, agriculture and land clearing [1]. These two phenomena, ocean warming and pCO2-induced acidification, are anticipated to have detrimental effects on seawater quality and consequently to marine organisms.
As soft-bodied, slow moving invertebrates, molluscs are highly susceptible to biotic pressures and abiotic changes in their environment. Marine molluscs are considered good models for climate change studies because they are ectothermic animals with limited ability to regulate their internal temperature. They can only survive within a narrow range of tolerable temperatures to which they have adapted [4] and stress outside their optimal temperature range leads to loss of metabolic functions [3,4,5]. Molluscs are also vulnerable to ocean acidification because their calcium carbonate shells are eroded when exposed to low pH [6,7] and an excess of hydrogen ions in the ocean can interfere with shell formation [8]. Furthermore, molluscs tend to have low metabolic rates and cannot easily compensate for pH disturbances [9]. Chronic stress from sub-optimal conditions can result in decreased growth and reproduction, increased susceptibility to disease and reduced survivorship [3,10,11]. Recent meta-analyses have identified molluscs as one of the most vulnerable invertebrate taxa under changing ocean conditions [9,11,12]. However, further studies are required to investigate the biochemical responses that may lead to reduced resilience under future conditions.
Marine molluscs comprise a major invertebrate fishery resource [13,14] and are highly regarded as healthful food for human consumption [15,16,17,18]. Like other shellfish and fish, molluscs are known to contain significant amounts of lipid, that is relatively low in saturated fatty acids (SFA) and high in polyunsaturated fatty acids (PUFA) [19,20]. The large amounts of PUFA in seafood offer nutritional and health benefits, such as the provision of essential fatty acids, carriers of fat-soluble vitamins, and decreasing the risk of cardiovascular disease [21]. Seafood is not only a good source of PUFAs, but also provides n-3 fatty acids with optimal n-3/n-6 ratios [22]. Dietary intake of n-3 fatty acids has a broad range of beneficial health effects in humans, including well-established anti-inflammatory, anti-arrhythmic and prothrombotic properties [23]. They are also reported to reduce the risk of colorectal cancer [24], lower blood pressure associated with hypertension [25], reduce depression and treat type 2 diabetes [26], and Alzheimer’s disease [27]. Omega-6 fatty acids are necessary for good nutrition, but when consumed in large amounts they may become harmful to the human body [26] by increasing the risk of cardiovascular and coronary heart disease [28] and may increase offspring adiposity [29]. To our knowledge, no previous studies have investigated the combined impacts of ocean warming and acidification on fatty acid compositions in marine molluscs, despite possible implications for human health.
Previous studies have shown that fatty acids in marine organisms can be influenced by environmental conditions, such as changes in temperature [30] and CO2 concentrations [31]. Temperature is a key parameter for optimal physiology in organisms, since it modulates the basic rates of all chemical reactions in cells, thus affecting the stability of structural components, particularly lipids and proteins [32]. Studies on plankton [33,34], copepods [35], microalgae [32,36,37], and bacteria [38] have demonstrated that temperature impacts the lipid composition by increasing the unsaturated fatty acid levels as the temperature declines and increasing the saturated fatty acids (SFA) content as the temperature increases. A recent study on the interactive effects of temperature and food quality was demonstrated in Daphnia pulex, where elevated temperature (26 °C) significantly decreased the total body fatty acids and thus negatively affected the PUFA content [39]. Based on comparison of the fatty acid composition of temperate and arctic marine ectotherms, Lewis [40] proposed that fatty acid profiles could actually be used to predict environmental temperatures effects.
Fewer studies have investigated the effects of ocean acidification (OA) on fatty acid composition. However, in a laboratory experiment to test potential OA effects on fatty acid composition in diatoms, elevated CO2 significantly changed the fatty acid concentration and composition of Thalassiosira pseudonana [31]. A significant decline in total fatty acids and the ratio of long-chain polyunsaturated to saturated fatty acids (PUFA:SFA) was found in algae cultured under elevated (pCO2 = 750 ppm; pH = 8.14) compared to present day CO2 concentrations (pCO2 = 380 ppm; pH = 7.94). This impact was directly translated to micro-algal grazing copepods, resulting in an almost tenfold decline in total fatty acids and triple the contribution of SFAs in copepods at high CO2 [31]. This study demonstrates the potential for far-reaching consequences of OA in ocean food webs by changing the nutritional quality of essential macromolecules in primary producers that can then cascade up the food web. However, to date, few studies appear to have investigated the potential for synergistic effects of elevated temperature and acidification on the lipid composition of benthic marine predators.
The Muricidae are a diverse family of predatory marine gastropods that currently comprise over 39% of the world-wide gastropod fisheries harvest [41]. Muricidae fisheries production has experienced a slow increase globally, with only 5000–24,000 tonnes of catch annually from 2002 to 2006, then peaking in 2007, with a catch of about 27,000 tonnes [41]. China is the lead producer [41], although muricids have gained importance in small-scale fisheries industry and aquaculture throughout Asia, Europe, and Central and South America [13,42], both for seafood consumption and as a source of Tyrian purple [43,44,45]. Dicathais orbita (Gmelin) is temperate species of Muricidae native to Australia, and is considered as a useful bioindicator of environmental conditions [45,46,47,48]. D. orbita has also provided a good model for natural product research [45]. However, the fatty acid composition of flesh of this species has not previously been reported.
Thus, the aim of the study was to assess the fatty acid composition in the foot tissue of the D. orbita and to investigate any impacts of ocean warming and acidification on the lipid content and fatty acid profile after a 35-day exposure to future climate change conditions. This study provides valuable information to establish the nutritional value of this Australian whelk and provides insights into the potential effect of future ocean conditions on the fatty acid composition of a predatory gastropod, with potential flow-on effects for human consumption.

2. Results and Discussion

2.1. Impacts of Ocean Climate Change on D. orbita Total Lipid Content

The highest lipid content from D. orbita foot tissue was 6.3 mg/g fresh weight in specimens held at current ocean conditions of 23 °C, current pCO2 (control), dropping to an average of 5.0 mg/g in specimens maintained at 23 °C and future pCO2 (Figure 1). The lipid content was less than 4% for all whelks kept at 25 °C, with the lowest mean amount of lipid at 3.6 mg/g from the foot tissue of whelks held in future warming and acidification conditions for 35 days (Figure 1). Permutational univariate analysis (PERMANOVA) revealed that temperature (p < 0.05), but not CO2-induced acidification, significantly affected the total lipid yield and there was no significant interaction between these factors (p > 0.05, Table 1). The reduced lipid content under elevated water temperature conditions in this study suggests that D. orbita is sensitive to the relatively small increases (~2 °C) in temperature predicted from global climate change models [1]. This result substantiates the negative response to elevated temperature in lipid synthesis for the larvae of the hard clam Mercenaria mercenaria and bay scallop Argopecten irradians [49], although these larva also showed a reduction in total lipids at elevated CO2. Metabolic rates generally increase at higher temperatures and under stressful conditions, thus placing more demand on metabolic reserves, such as the stored lipid. Reduced lipid reserves under conditions of elevated temperature could have significant implications for the long-term viability of whelk populations. High water temperature can result in the dysfunction of important biological functions, as lipids are structural components of all cell membranes and are considered cellular fuels [50,51,52]. Reduced lipid storage has been correlated with poor growth condition in predatory fish in response to increasing sea surface temperatures and it has been argued that lipid reserves also provide a good indicator for reproductive performance, as they are crucial for pre-spawning conditioning of eggs [53]. Consequently, the effects of long-term exposure to future ocean conditions on lipid reserves could result in reduced reproduction, development and growth of marine molluscs. This could have serious implications for population viability and sustainable fisheries, unless they are able to adapt to the new conditions. The east coast of Australia has been recognized as an ocean warming hotspot [54] and in this study, D. orbita was collected close to the northern limit of its distribution. Previous studies on invertebrates from thermally stressful intertidal habitats indicate that warm-adapted populations may already be close to their upper thermal tolerance and thus are most vulnerable to ocean warming due to less capacity to acclimatize [55]. Nevertheless, multi-generational studies have indicated the presence of tolerant genotypes in other marine invertebrates [56], suggesting potential for some species to adapt to concurrent ocean warming and acidification.
Marinedrugs 13 06019 g001 1024
Figure 1. Total lipid yield extracted from the foot tissue of D. orbita after 35 days exposure to temperature and pCO2 treatments (n = 6 per group). Error bars show standard error of the mean. * Significantly different to the control at 23 °C and current pCO2 (p < 0.05).
Figure 1. Total lipid yield extracted from the foot tissue of D. orbita after 35 days exposure to temperature and pCO2 treatments (n = 6 per group). Error bars show standard error of the mean. * Significantly different to the control at 23 °C and current pCO2 (p < 0.05).
Marinedrugs 13 06019 g001
Table
Table 1. Summary of the statistical outcomes for all univariate and multivariate analyses. Two factor PERMANOVAs were used to test the effects of temperature and pCO2 induced acidification. Significant effects are in bold.
Table 1. Summary of the statistical outcomes for all univariate and multivariate analyses. Two factor PERMANOVAs were used to test the effects of temperature and pCO2 induced acidification. Significant effects are in bold.
TemperatureAcidificationTemperature × Acidification
Pseudo Fp ValuePseudo Fp ValuePseudo Fp Value
UNIVARIATE
Total lipid yield19.12300.00021.98630.17961.63960.2098
SFA3.61890.07334.29890.04783.10750.0905
MUFA0.41150.54515.71820.01960.00630.9373
PUFA10.9810.00290.02810.87056.72980.0164
n-34.71500.04760.00020.98811.85720.1848
n-65.15510.03401.26600.27420.04520.8261
n-3:n-6 ratio0.336970.56970.202870.66371.08280.3088
MULTIVARIATE
Overall fatty acid composition7.70940.00012.84520.02291.71860.1321

2.2. Impacts of Ocean Climate Change on the Major Classes of Fatty Acids

The distribution of fatty acids in D. orbita under current ocean conditions was characterised by a predominance of PUFA (51.6% ± 2.1%) followed by SFA (26.7% ± 1.4%) and a low abundance of MUFA (10.2% ± 0.9%) (Figure 2). This relatively high unsaturated fatty acid content is consist across a wide range of molluscs, including filter feeding bivalves, herbivorous gastropods and predatory whelks [20,57]. D. orbita contains more n-3 (ALA, eicosatrienoic, EPA, DPA, and DHA) than n-6 (LA, and ARA) PUFAs (Table 2), with an n-3:n-6 ratio of approximately 2 (Figure 2). Both n-3 and n-6 fatty acids are required for normal human health, with a recommended ratio between 1 and 4 [58,59]. In Western diets the n-3 to n-6 ratio is approximately 1:20–30, indicating a deficiency of n-3 fatty acids, which has been linked to several diseases, such as heart disease, diabetes and hypertension [60]. Our study reveals that D. orbita provides a good balance of n-3 and n-6 and a high proportion of PUFAs, which could contribute to a healthful human diet.
Marinedrugs 13 06019 g002 1024
Figure 2. Proportions of (a) polysaturated fatty acids (PUFA) (n = 6) showing relative amounts of n-3 and n-6 fatty acids; (b) saturated fatty acids (SFA) (n = 12) and (c) monosaturated fatty acids (MUFA) (n = 12) in D. orbita foot tissue after 35 days exposure to different temperatures and CO2-induced acidification. Error bars show standard error of the mean. * Significant differences (p < 0.05) within the same group of fatty acids, in comparison to the control at 23 °C and current pCO2.
Figure 2. Proportions of (a) polysaturated fatty acids (PUFA) (n = 6) showing relative amounts of n-3 and n-6 fatty acids; (b) saturated fatty acids (SFA) (n = 12) and (c) monosaturated fatty acids (MUFA) (n = 12) in D. orbita foot tissue after 35 days exposure to different temperatures and CO2-induced acidification. Error bars show standard error of the mean. * Significant differences (p < 0.05) within the same group of fatty acids, in comparison to the control at 23 °C and current pCO2.
Marinedrugs 13 06019 g002
Examination of the fatty acids of D. orbita after exposure to future temperature and pCO2-induced acidification treatments for 35 days revealed that the relative proportions of the major fatty acid classes remained similar across all treatments (Figure 2) and there was no significant difference in the ratio of n-3:n-6 (Table 1, p > 0.05). This implies that the nutritional benefit associated with the fatty acid content of these whelks is likely to be retained under future ocean conditions.
Table
Table 2. Fatty acid and lipophilic hydrocarbon profile of D. orbita foot tissue after 35 days exposure to current and future ocean temperatures and acidification conditions. Data are expressed as % of total fatty acid methyl esters (FAMEs) as mean ± SE (n = 12). Others include cyclopropane fatty acids and dimethyl acetal aldehydes from plasmalogen phospholipids.
Table 2. Fatty acid and lipophilic hydrocarbon profile of D. orbita foot tissue after 35 days exposure to current and future ocean temperatures and acidification conditions. Data are expressed as % of total fatty acid methyl esters (FAMEs) as mean ± SE (n = 12). Others include cyclopropane fatty acids and dimethyl acetal aldehydes from plasmalogen phospholipids.
Fatty AcidTrivial NameRetention Time (min)23 °C, Current pCO223 °C, Future pCO225 °C, Current pCO225 °C, Future pCO2
Saturated
C14:0Myristic16.81.46 ± 0.131.56 ± 0.051.74 ± 0.081.28 ± 0.12
C15:0Pentadecanoic18.41.24 ± 0.181.30 ± 0.071.38 ± 0.111.11 ± 0.12
C16:0Palmitic19.89.26 ± 0.498.86 ± 0.179.53 ± 0.398.66 ± 0.23
C17:0Margaric21.31.98 ± 0.121.71 ± 0.101.31 ± 0.121.22 ± 0.13
C18:0Stearic22.68.26 ± 0.498.17 ± 0.157.90 ± 0.248.30 ± 0.12
C24:0Lignoceric29.74.54 ± 0.164.99 ± 0.364.81 ± 0.164.18 ± 0.13
Monounsaturated
C16:1Palmitoleic20.61.87 ± 0.160.84 ± 0.320.89 ± 0.250.86 ± 0.23
C18:1 (n-9)Oleic23.14.54 ± 0.494.96 ± 0.074.89 ± 0.234.42 ± 0.06
C20:1 (n-9)11-Eicosenoic25.73.50 ± 0.113.40 ± 0.193.81 ± 0.163.81 ± 0.19
C22:1 (n-9)Erucic30.20.24 ± 0.040.20 ± 0.010.34 ± 0.110.15 ± 0.07
Polyunsaturated
C18:2 (n-6)Linoleic acid (LA)24.11.54 ± 0.091.51 ± 0.071.66 ± 0.121.69 ± 0.08
C18:3 (n-3)α-Linolenic (ALA)25.10.56 ± 0.040.60 ± 0.030.69 ± 0.060.77 ± 0.04
C20:2Eicosadienoic26.52.50 ± 0.132.89 ± 0.162.60 ± 0.12.45 ± 0.13
C20:3 (n-3)Eicosatrienoic27.10.03 ± 0.030.03 ± 0.0300
C20:4 (n-6)Arachidonic (ARA)27.311.49 ± 0.2211.84 ± 0.2610.88 ± 0.1611.07 ± 0.14
C20:5 (n-3)Eicosapentaenoic (EPA)28.32.61 ± 0.302.76 ± 0.332.00 ± 0.422.05 ± 0.44
C22:2Docosadienoic28.511.15 ± 0.5710.15 ± 0.3911.81 ± 0.5111.40 ± 0.42
C22:5 (n-3)Docosapentaenoic (DPA)30.717.72 ± 0.4916.87 ± 0.5216.13 ± 0.4016.69 ± 0.39
C22:6 (n-3)Docosahexaenoic (DHA)30.94.02 ± 0.273.70 ± 0.133.55 ± 0.263.85 ± 0.35
Others
2-octylcyclo-propanedecanoic26.80.63 ± 0.020.64 ± 0.040.68 ± 0.030.73 ± 0.02
Unknown fatty acid derivative29.20.53 ± 0.040.48 ± 0.060.56 ± 0.050.50 ± 0.05
Dimethyl acetal aldehydes
Hexadecan-1-al18.81.04 ± 0.080.94 ± 0.040.87 ± 0.700.82 ± 0.02
Heptadecan-1-al20.20.21 ± 0.051.03 ± 0.260.86 ± 0.171.78 ± 0.06
Octadecan-1-al21.78.91 ± 0.8310.27 ± 0.2010.80 ± 0.2911.92 ± 0.20
Nonadecan-1-al24.70.17 ± 0.060.30 ± 0.030.30 ± 0.070.29 ± 0.06
Nevertheless, PERMANOVA univariate analysis identified some significant effects on the % PUFA, MUFA and SFAs (Table 1). The relative proportion of PUFAs was affected by a significant interaction between temperature and pCO2 (Table 1, p < 0.05). There was a significant reduction in the % PUFAs at elevated temperatures under current pCO2 conditions, but no effect under future conditions (Figure 2). Temperature also caused a consistent reduction in the amount of n-3 and n-6 fatty acids, as a percent of total lipids (Figure 2, Table 1, p < 0.05). This effect of temperature is consistent with a previous study on bivalves, Ruditapes decussatus and Ruditapes philippinarum, which also showed decreased PUFA content after exposure to warmer waters [61]. Some PUFAs are important in maintaining the fluidity and permeability of biological membranes in response to temperature fluctuations [50]. The decreased proportion of PUFA at higher temperatures could be explained by acclimatization to the prevailing thermal conditions by restructuring cell membranes to maintain optimal fluidity and permeability. This strategy, termed homeoviscous adaptation, can occur via changes in phospholipid head groups, fatty acid composition, and cholesterol content [51]. Nevertheless, it remains unclear whether this biochemical acclimatization response, to offset the direct effects of temperature on membrane lipid fluidity, can be sustained under long-term conditions that also cause an overall reduction in the total lipid content.
The proportion of PUFAs in D. orbita was not significantly affected by future pCO2 conditions at either temperature (Table 1, p > 0.5, Figure 2a). This result agrees with recent research on Pacific oyster Crassostrea gigas that showed no effect on the total fatty acid contents under elevated pCO2 [62]. Nevertheless, exposure to CO2-induced acidification did result in a significant but small reduction (1%–2%) in the relative proportions of SFA and MUFA in D. orbita (Table 1, p < 0.05, Figure 2). Although response to CO2 acidification is species-specific, it has been proposed that elevated CO2 levels promote synthesis and accumulation of SFAs in green algae and can lead to the desaturation of pre-existing fatty acids [63]. SFAs were also found to increase proportionally under elevated pCO2 conditions in diatoms and this effect was magnified in grazing copepods [31]. Our results on a predatory whelk are inconsistent with this, as we found a significant decrease in the relative proportion of SFAs under future CO2 acidification conditions. However, the whelks were fed fresh oysters that were held under current ocean conditions, and thus our study does not account for potential trophic transfer of altered lipid compositions. The integration species interactions into future studies on temperature and pCO2-induced biochemical changes could provide a more realistic assessment of species vulnerabilities, based on functional networks from the molecular to ecosystem scale [64].

2.3. Impacts of Ocean Climate Change on Fatty Acid Composition

In the present study, we have identified 21 fatty acids in D. orbita lipid extracts (Table 2, Figure S1), with carbon atoms ranging from 14 to 22, including saturated, monoenoic, polyenoic and cyclopropane-containing fatty acids (CFA) (e.g., Figure S2). The biophysical properties of these cyclic fatty acids are similar to unsaturated fatty acids, hence they are usually considered as part of the unsaturated fatty acid component [65]. Several studies have confirmed the presence of CFAs in marine organisms, such as the Okinawan ascidian Diplosoma sp. [66] and the Caribbean sponge P. suberitoides [67]. CFAs have also been previously reported in freshwater molluscs, specifically the prosobranch gastropods Viviparus (Bellayma) bengalensis and Pila globosa [68]. CFAs are common in bacteria and function to increase cellular membrane stability when bacteria are exposed to low pH [69]. Consistent with this we observed a slight increase in 2-octylcyclo-propanedecanoic methyl ester in whelks held under future CO2 conditions (Table 2). The relative proportion of CFAs in D. orbita also increased at elevated water temperatures, which may be due to the post-synthesis modification of unsaturated fatty acids [70].
A series of dimethyl acetals of aliphatic aldehydes were also identified in the lipid composition of D. orbita (Table 2, Figure S3), based on the common fragment ion m/z 75, representing the McLafferty rearrangement ion (CH3O)2CH− [71]. The detection of these dimethyl acetals of aldehydes in D. orbita tissue is consistent with the presence of animal cell plasmalogens, which are glycerolphospholipids with a vinyl ether linkage at the sn-1 position and an ester linkage at the sn-2 position [72]. The aldehydes are generated when the vinyl ether bond is broken, then immediately converted to dimethyl acetals during acidic transesterification in BF3 methanol [71]. Plasmalogens play many important physiological roles in animals and have been proposed to provide a sink for polyunsaturated fatty acids in some tissues [72]. In a study on the mitochondrial fraction of the marine bivalve Arctica islandica, the abundance of plasmalogens (evaluated on the basis of dimethyl acetal content) was found to slightly increase as a result of warmer temperature [73]. This is in agreement with our results. In D. orbita, the proportional increase of dimethyl acetals at elevated temperature coincides with the decrease of PUFA (Table 1, Figure 3). This may be because PUFAs are degraded by autoxidation chain reactions within the cell membranes, thereby releasing reactive fatty aldehydes [61,73]. Multivariate PERMANOVA revealed that there is a significant effect of both temperature and acidification on the overall fatty acid composition of D. orbita (Table 1, p < 0.05). However, there was no synergistic interaction between temperature and acidification (Table 1, p > 0.05). Principal coordinate ordination (PCO) with trajectory overlay was used to explore the differences in the lipid composition between temperature and acidification treatments (Figure 3). The whelks held at 23 °C and current pCO2 treatment are spread across the right hand side of the ordination plot, whereas the points representing whelks held at 25 °C and future pCO2 treatment are clustered mostly towards the left hand side (Figure 3). Vector overlay using Spearman rank correlation suggests that the whelks held at 25 °C in future pCO2 conditions are characterized by higher dimethyl acetal of octadecan-1-al, which could represent changes to the phospholipid cell membranes in whelks held under future stressful conditions. In comparison, the majority of whelks maintained at 23 °C in current pCO2 conditions are characterized by higher margaric acid and DPA (Figure 3, Table 2). Variation in the concentrations of docosadienoic acid and EPA along the Y axis explains much of the variation between replicate whelks held under the same treatment conditions (Figure 3).
Marinedrugs 13 06019 g003 1024
Figure 3. Principal coordinate ordination (PCO) of the lipophilic compound profile from D. orbita based on a Euclidian distance similarity matrix of the percent composition data with vector overlay from Spearman rank correlation of 0.6.
Figure 3. Principal coordinate ordination (PCO) of the lipophilic compound profile from D. orbita based on a Euclidian distance similarity matrix of the percent composition data with vector overlay from Spearman rank correlation of 0.6.
Marinedrugs 13 06019 g003
Despite significant changes in the fatty acid composition of whelks held under future temperature and pCO2 conditions, the degree of impact on specific PUFAs was perhaps not as apparent as expected (Table 2). By comparison, a study on the gill of an eastern oyster Crassostrea virginica maintained at 25 °C showed a 14% increase in the amount of ARA compared to gills from the oysters kept at 12 °C. However, the magnitude of temperature change in this previous study on oysters was much greater (52% increase), in comparison to just 2 °C (8%) increase in our study, as this is more relevant to future ocean warming predictions. The detection of even relatively minor statistically significant changes to the fatty acid composition and glycerophospholipids of D. orbita, after just 35 days exposure to small predicted increases in water temperature, is of concern. These small changes in relative composition are magnified by the large decreases in total lipid content (Figure 1), with an average of 43% reduction in all lipids under future warming and pCO2 conditions, relative to current conditions. Long-term exposure and multi-generational studies are required to assess the potential for acclimatization and adaptation of the metabolic and biosynthetic capabilities of predatory marine whelks under future ocean conditions.
Furthermore, whilst this study has examined the impacts of ocean climate change stressors on whelk lipid storage and fatty acid composition, we have not accounted for bioaccumulated effects on dietary derived fatty acids likely to result from predator-prey interactions in natural ecosystems experiencing long term climate change. The combined effects of ocean warming and acidification have demonstrated effects on the biosynthesis of PUFAs in marine algae, with flow-on effects for herbivores [31] and ultimately this could further magnify the effects on higher order consumers. Therefore whilst the statistically significant changes in fatty acid proportions in D. orbita, along with the overall reduction in total lipid yield under future ocean conditions, can be considered a negative outcome from ocean warming and acidification, the effects may actually be under-estimated, with unknown consequences for the nutritional requirements and long-term survival of the species. Future mesocosm studies, involving a range of marine predator and prey interactions, could investigate the potential accumulation of biochemical changes in key primary metabolites under prolonged stressful conditions. This would help establish whether marine species at higher trophic levels are generally at greater risk of metabolic dysfunction under future ocean conditions and any consequent implications for the production of sustainable healthful seafood.

3. Experimental Section

3.1. Study Site and Experimental Design

To test the hypothesis that ocean warming and acidification will impact the fatty acid composition of Dicathais orbita, 144 whelks were subjected to 35 days under experimental conditions at the National Marine Science Centre (NMSC), Coffs Harbour, Australia (30°16′3.70″ S, 153°8′15.31″ E). The experiment utilized D. orbita (51–79 mm shell length) from rock platforms around Coffs Harbour and a two-factor factorial design with four treatments (mean ± SD): (1) current conditions, 22.9 ± 0.6 °C and 378.6 ± 35.6 ppm; (2) elevated temperatures, 25.2 ± 0.6 °C and 382.2 ± 35.5 ppm; (3) elevated pCO2 = 22.9 ± 0.5 °C and 749.9 ± 80.6 ppm; and (4) increased temperature 25.3 ± 0.6 °C and elevated pCO2 = 763.0 ± 104.6 ppm. Ambient water temperatures were based on data collated by Navy Meteorology and Oceanography indicating an average (±SD) sea surface temperature off the Coffs Harbour coast between September and November of approximately 21.3 ± 1.0 °C [74]. Ambient pH during the experimental period was estimated to be ~8.2 [75]. Future ocean conditions were based on the IPCC [76] trajectory for a drop in 0.3 pH (i.e., increased pCO2 to 750–800 ppm) and a 3 °C rise in sea surface temperature by the year 2100 under climate change model RCP8.5.
To maintain experimental conditions, twelve 1100 L header tanks were filled with seawater pumped from the open ocean adjacent to the NMSC. This seawater was passed through a sand filter and 50 µm filters before being allowed to flow into the header tanks. The temperature of seawater in the header tanks was controlled by heater chiller units (Aquahort Ltd., Auckland, New Zealand). Seawater pH (pCO2) was manipulated by bubbling CO2-enriched air through experimental treatments after pre-mixing the gases using a gas mixer (PEGAS 4000MF, Columbus Instruments, Columbus, OH, USA). The water temperature, pH, conductivity and salinity were measured daily and total alkalinity was measured weekly through potentiometric titration using an automated titrator (888 Titrando, Metrohom, Riverview, FL, USA). The pH, alkalinity, temperature and salinity readings were used to calculate the partial pressure of CO2 using the CO2SYS program [77] with constants from Mehrbach et al. [78] as adjusted by [79].
Each header tank supplied temperature and pCO2 controlled water at 3 L/min−1 to a tray (860 mm × 650 mm × 96 mm) that housed four cages (305 mm × 360 mm × 90 mm) with three D. orbita in each cage. D. orbita were acclimated in experimental conditions for one week before feeding trials commenced. Whelks were fed Sydney rock oysters (Saccostrea glomerata), a common prey item in field conditions. The whelks in each cage were initially fed 4 small oysters ranging 30–50 mm, with new oysters of similar size added daily. After 35 days exposure to the experimental conditions, three male and three female whelks were sampled from each tank.

3.2. Extraction and Preparation of Fatty Acid Methyl Esters

To prepare the samples, soft tissue of the whelks was extracted by crushing the shell using a bench-top vice. Tissues from three replicate male whelks were pooled from each tank and similarly three replicate females were pooled to represent one replicate sample per tank (n = 3 tanks per treatment combination for each gender). The samples were prepared from the foot tissue (~1.0 gram) and were soaked in methanol:chloroform (2:1) for 1.5 h. The solvent extract was then filtered using Sigma-Aldrich Whatman filter paper 90 mm into a clean test tube. The tissue was soaked further in fresh solvent, decanted and replaced until a colourless solution was obtained. At least three washes were made to maximize lipid recovery. All the chloroform fractions from each sample were combined and dried using a rotary evaporator (Buchi Vacuum System) at 40 °C maintained at 337 mbar Hg. The samples were then transfered to a clean pre-weighed vial using sequential resuspension in minimum volumes of methanol:chloroform solution and were further concentrated under a stream of high purity (100%) nitrogen gas and then weighed and stored in a −20 °C freezer until utilized.
The above lipid extracts were derivatised to generate fatty acid methyl esters according to Kanthilatha et al. [80]. The dried lipid extracts were dissolved in 1.5 mL of 0.5 M saturated sodium hydroxide in methanol and then heated in a dri-block at 100 °C for 10 min. To completely methylate the extracts, 2 mL of boron trifluoride in methanol was added and then heated again for another 30 min. After cooling at room temperature, 1 mL of hexane was added to extract the fatty acid methyl esters (FAMEs). The tube was shaken vigorously for 30 s. Five milliliters of sodium chloride solution was added to aid in phase separation and shaken again for 5 s to separate the hydrophilic layer from the lipophilic layer. The FAMEs that were formed were recovered from the lower phase and collected in an autosampler vial ready for gas chromatography (GC) injection.

3.3. FAMEs and GC Analysis

The FAMEs samples were analysed using a GC (Agilent 6890N, Santa Clara, CA, USA) coupled with a flame ionisation detector (FID) with Agilent 6890 split/splitless injection and a fitted with BPX 70 capillary column (70% cyanopropyl polysilphenylene-siloxane, 50 m length, 0.22 mm internal diameter and 0.25 µm thickness). The FID was operated at 260 °C while the split injector was kept at 230 °C. The carrier gas was high-purity helium maintained with a linear flux of 1 mL/min. The GC oven was initially held at 100 °C for 5 min and then raised to 240 °C at a rate of 5 °C/min. One microliter of the sample extract was injected with a split ratio of 200:1 and a column flow of 1 mL/min.
FAMEs were identified by the peak, retention time and elution order and compared against the reference FAMEs standard test mix (SUPELCO 37-Component FAME Mix CRM47885, Bellefonte, PA, USA). Some samples were further analyzed using an Agilent gas chromatography-mass spectrometer (GC-MS) with an Agilent 5973 Mass Selective Detector to confirm the identity of the fatty acids. The mass spectra were recorded at 70 eV ionisation voltage over the mass range of 35 to 550 amu. Individual peaks were identified by comparison to library mass spectra (WILEY 275 and NIST98). To facilitate the identification of DPA, which was not in the test mix, a soft ionisation MS technique at 40 eV ionisation voltage was employed to ionise the lipid molecules in the D. orbita samples without causing extensive fragmentation (Figure S4). Spectrum was compared on the MS database, retention time and elution order from extensive literature search such as the American Oil Chemists’ Society. The relative composition of each identified fatty acid was done by peak integration from the GC and expressed for individual FAMEs as percentages of the total in each run.

3.4. Statistical Analyses

The data are expressed as means ± standard error. All statistical analyses were undertaken using PRIMER v 6 + PERMANOVA add-on. Initially three-factor PERMANOVAs (temperature, pCO2 and gender) were used to investigate the effects of temperature, pCO2-induced pH and snail gender. However, in all cases gender was not statistically significant and did not interact with the other factors (p > 0.05). Consequently, two-factor PERMANOVAs (temperature and pCO2) were used to investigate the effects of temperature and CO2-induced acidification. Univariate PERMANOVAs were used to compare the total yield of lipid extract, the percent SFA, MUFA and PUFA, as well as the percent of n-3 and n-6 fatty acids between each of the experimental conditions. Multivariate analyses were used to assess the total fatty acid composition (relative abundance standardized by the total). In all cases, Euclidean distance similarity matrices were created and PERMANOVAs were run using the full model and 9999 permutations of the data to determine overall differences between treatments followed by pairwise analyses on the interaction term when this was significant. Principal component ordination (PCO) was also undertaken to visually represent the patterns in the multivariate data (overall fatty acid composition standardized by total), with vector overlay using Spearman correlation >0.6. In all analyses, a statistically significant result was accepted for p < 0.05.

4. Conclusions

D. orbita has been found to contain a complex fatty acid composition with high PUFA levels and a good ratio of n-3 and n-6 fatty acids, which is typical of healthful seafood. Small temperature and pCO2 increases predicted with future ocean warming and acidification can, however, negatively impact the total yield of lipids and the overall fatty acid composition. Overall, our study indicated that ocean warming significantly decreases the proportion of PUFA and increases plasmalogen derived dimethyl acetals of aliphatic aldehydes, whilst elevated pCO2 decreases SFA and MUFA and alters the overall fatty acid composition. However, with the exception of total lipids, the percent changes in composition are relatively small and the n-3:n-6 ratio remains the same, suggesting minimal implications for human consumption of D. orbita as nutritional seafood. Nevertheless, there are possible implications for the long term viability of the species resulting from reduced lipid reserves and the potential for reduced bioaccumulation of lipids through the food web in predatory species.

Supplementary Files

Supplementary File 1

Acknowledgments

The first author greatly acknowledges the support from Davao Oriental State College of Science and Technology (DOSCST), Mati City, Davao Oriental, Philippines and the Australia Awards Scholarships. We thank L. Yee for the meaningful encouragement and guidance for this project. Southern Cross University (SCU), School of Environment, Science and Engineering and Marine Ecology Research Centre provided post-graduate research funding. The National Marine Science Centre (NMSC-SCU) and Southern Cross Plant Science (SCPS) contributed the use of their laboratory facilities. P. Mouatt and M. Russell helped identify fatty acid peaks.

Author Contributions

R.V.-R., K.B. and B.K. conceived and designed the experiments. R.T. maintained and monitored the climate change experiment and assisted R.V.-R. with the preparation of samples. D.S. ran the FAMEs and GC-MS with advice from A.D., R.V.-R., D.S., A.D. and K.B. all contributed to the identification of fatty acids. R.V.-R. and K.B. undertook the statistical analyses and R.V.-R. wrote the first draft of the paper, with further contributions from all authors. This manuscript was substantially revised by K.B. and edited by R.V.-R. in response to constructive feedback from reviewers.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. IPCC. Climate Change 2007. The Physical Science Basis. Contribution of Working Group 1 to the Fourth Assessment, Report of the Intergovernmental Panel on Climate Change; Solomon, S., Qin, D., Manning, M., Chen, Z., Marquis, M., Averyt, K.B., Tignor, M., Miller, H.L., Eds.; Cambridge University Press: Cambridge, UK, 2007. [Google Scholar]
  2. Caldeira, K.; Wickett, M.E. Oceanography: Anthropogenic carbon and ocean pH. Nature 2003, 425, 365–365. [Google Scholar] [CrossRef] [PubMed]
  3. Byrne, M. Impact of ocean warming and ocean acidification on marine invertebrate life history stages: Vulnerabilities and potential for persistence in a changing ocean. Oceanogr. Mar. Biol. 2011, 49, 1–42. [Google Scholar]
  4. Harley, C.D.G.; Denny, M.W.; Mach, K.J.; Miller, L.P. Thermal stress and morphological adaptations in limpets. Funct. Ecol. 2009, 23, 293–301. [Google Scholar] [CrossRef]
  5. Nguyen, K.D.T.; Morley, S.A.; Lai, C.H.; Clark, M.S.; Tan, K.S.; Bates, A.E.; Peck, L.S. Upper temperature limits of tropical marine ectotherms: Global warming implications. PLoS ONE 2012, 6. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  6. Parker, L.M.; Ross, P.M.; Pörtner, H.O.; Scanes, E.; Wright, J.M. Predicting the response of molluscs to the impact of ocean acidification. Biology 2013, 2, 651–692. [Google Scholar] [CrossRef]
  7. Fabry, V.J.; Seibel, B.A.; Feely, R.A.; Orr, J.C. Impacts of ocean acidification on marine fauna and ecosystem processes. ICES J. Mar. Sci. 2008, 65, 414–432. [Google Scholar] [CrossRef]
  8. Orr, J.C.; Fabry, V.J.; Aumont, O.; Bopp, L.; Doney, S.C.; Feely, R.A.; Gnanadesikan, A.; Gruber, N.; Ishida, A.; Joos, F.; et al. Anthropogenic ocean acidification over the twenty-first century and its impact on calcifying organisms. Nature 2005, 437, 681–686. [Google Scholar] [PubMed]
  9. Wittmann, A.C.; Pörtner, H. Sensitivities of extant animal taxa to ocean acidification. Nat. Clim. Chang. 2013, 3, 995–1001. [Google Scholar]
  10. Hooper, C.; Day, R.; Slocombe, R.; Benkendorff, K.; Handlinger, J.; Goulias, J. Effects of severe heat stress on immune function, biochemistry and histopathology in farmed Australian abalone (hybrid Haliotis laevigata × Haliotis rubra). Aquaculture 2014, 432, 26–37. [Google Scholar] [CrossRef]
  11. Kroeker, K.J.; Kordas, R.L.; Crim, R.; Hendriks, I.E.; Ramajo, L.; Singh, G.S.; Duarte, C.M.; Gattuso, J.P. Impacts of ocean acidification on marine organisms: Quantifying sensitivities and interaction with warming. Glob. Chang. Biol. 2013, 19, 1884–1896. [Google Scholar] [CrossRef] [PubMed]
  12. Byrne, M.; Przeslawski, R. Multistressor impacts of warming and acidification of the ocean on marine invertebrates’ life histories. Integr. Comp. Biol. 2013, 53, 582–596. [Google Scholar] [CrossRef] [PubMed]
  13. Leiva, G.E.; Castilla, J.C. A review of the world marine gastropod fishery: Evolution of catches, management and the Chilean experience. Rev. Fish Biol. Fish. 2002, 11, 283–300. [Google Scholar] [CrossRef]
  14. Food and Agriculture Organization. The State of World Fisheries and Aquaculture; FAO: Rome, Italy, 2012. [Google Scholar]
  15. Ponder, W.; Hutchings, P.; Chapman, R. Overview of the Conservation of Australian Marine Invertebrates; A Report for Environment Australia; Australian Museum: Sydney, Australia, 2002. [Google Scholar]
  16. Szabó, K.; Amesbury, J.R. Molluscs in a world of islands: The use of shellfish as a food resource in the tropical island Asia-Pacific region. Quat. Int. 2011, 239, 8–18. [Google Scholar] [CrossRef]
  17. Koutsoubas, D.; Galinou-Mitsoudi, S.; Katsanevakis, S.; Leontarakis, P.; Metaxatos, A.; Zenetos, A. Bivalve and gastropod molluscs of commercial interest for human consumption in the Hellenic seas. In State of the Hellenic Fisheries; Papaconstantinou, C., Zenetos, A., Tserpes, G., Vassilopoulou, V., Eds.; HCMR Publications: Athens, Greece, 2007; pp. 70–84. [Google Scholar]
  18. Flores-Garza, R.; García-Ibáñez, S.; Flores-Rodríguez, P.; Torreblanca-Ramírez, C.; Galeana-Rebolledo, L.; Valdés-González, A.; Suástegui-Zárate, A.; Violante-González, J. Commercially important marine mollusks for human consumption in Acapulco, México. Nat. Res. 2012, 3, 11–17. [Google Scholar]
  19. Saito, H.; Aono, H. Characteristics of lipid and fatty acid of marine gastropod Turbo cornutus: High levels of arachidonic and n-3 docosapentaenoic acid. Food Chem. 2014, 145, 135–144. [Google Scholar] [CrossRef] [PubMed]
  20. Nichols, P.D.; Virtue, P.; Mooney, B.D.; Elliott, N.G.; Yearsley, G.K. Seafood the Good Food. The Oil (Fat) Content and Composition of Australian Commercial Fishes, Shellfishes and Crustaceans; CSIRO Division of Marine Research; Fisheries Research and Development Corporation: Deakin, Australia, 1998. [Google Scholar]
  21. Ruxton, C.H.S.; Reed, S.C.; Simpson, M.J.A.; Millington, K.J. The health benefits of omega-3 polyunsaturated fatty acids: A review of the evidence. J. Hum. Nutr. Diet. 2007, 20, 275–285. [Google Scholar] [CrossRef]
  22. Meyer, B.J.; Mann, N.J.; Lewis, J.L.; Milligan, G.C.; Sinclair, A.J.; Howe, P.R.C. Dietary intakes and food sources of omega-6 and omega-3 polyunsaturated fatty acids. Lipids 2003, 38, 391–398. [Google Scholar] [CrossRef]
  23. Covington, M.B. Omega-3 fatty acids. Am. Fam. Physician 2004, 70, 133–140. [Google Scholar] [PubMed]
  24. Kimura, Y.; Kono, S.; Toyomura, K.; Nagano, N.; Mizoue, T.; Moore, M.A.; Mibu, R.; Tanaka, M.; Kakeji, Y.; Maehara, Y.; et al. Meat, fish and fat intake in relation to subsite-specific risk of colorectal cancer: The Fukuoka colorectal cancer study. Cancer Sci. 2007, 98, 590–597. [Google Scholar] [PubMed]
  25. De Wilde, M.; Hogyes, E.; Kiliaan, A.J.; Farkas, T.; Luiten, P.G.M.; Farkas, E. Dietary fatty acids alter blood pressure, behavior and brain membrane composition of hypertensive rats. Brain Res. 2003, 988, 9–19. [Google Scholar] [CrossRef]
  26. Kendall-Tackett, K. Long-chain omega-3 fatty acids and women’s mental health in the perinatal period and beyond. J. Midwifery Womens Health 2010, 55, 561–567. [Google Scholar] [CrossRef] [PubMed]
  27. Solfrizzia, V.; D’Intronoa, A.; Colaciccoa, A.M.; Capursob, C.; del Parigic, A.; Capursoa, S.; Gadaletaa, A.; Capursoa, A.; Panzaa, F. Dietary fatty acids intake: Possible role in cognitive decline and dementia. Exp. Gerontol. 2005, 40, 257–270. [Google Scholar] [CrossRef] [PubMed]
  28. Ramsden, C.E.; Zamora, D.; Leelarthaepin, B.; Majchrzak-Hong, S.F.; Faurot, K.R.; Suchindran, C.M.; Ringel, A.; Davis, J.M.; Hibbeln, J.R. Use of dietary linoleic acid for secondary prevention of coronary heart disease and death: Evaluation of recovered data from the Sydney diet heart study and updated meta-analysis. BMJ 2013, 346, 1–18. [Google Scholar] [CrossRef] [PubMed]
  29. Moon, R.J.; Harvey, N.C.; Robinson, S.M.; Ntani, G.; Davies, J.H.; Inskip, H.M.; Godfrey, K.M.; Dennison, E.M.; Calder, P.C.; Cooper, C.; et al. Maternal plasma polyunsaturated fatty acid status in late pregnancy is associated with offspring body composition in childhood. J. Clin. Endocrinol. Metab. 2013, 98, 299–307. [Google Scholar] [PubMed]
  30. Noyola, J.; Mascaró, M.; Caamal-Monsreal, C.; Noreña-Barroso, E.; Díaz, F.; Re, D.; Sánchez, A.; Rosas, C. Effect of temperature on energetic balance and fatty acid composition of early juveniles of Octopus maya. J. Exp. Mar. Biol. Ecol. 2013, 445, 156–165. [Google Scholar] [CrossRef]
  31. Rossoll, D.; Bermudez, R.; Hauss, H.; Schulz, K.G.; Riebesell, U.; Sommer, U.; Winder, M. Ocean acidification-induced food quality deterioration constrains trophic transfer. PLoS ONE 2012, 7. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  32. Wagenen, J.V.; Miller, T.W.; Hobbs, S.; Hook, P.; Crowe, B.; Huesemann, M. Effects of light and temperature on fatty acid production in Nannochloropsis salina. Energies 2012, 5, 731–740. [Google Scholar] [CrossRef]
  33. Farkas, T.; Herodek, S. The effect of environmental temperature on the fatty acid composition of crustacean plankton. J. Lipid Res. 1964, 5, 369–373. [Google Scholar] [PubMed]
  34. Thompson, P.A.; Guo, M.; Harrison, P.J.; Whyte, J.N.C. Effects of variation in temperature. II. On the fatty acid composition of eight species of marine phytoplankton. J. Phycol. 1992, 28, 488–497. [Google Scholar]
  35. Nanton, D.A.; Castell, J.D. The effects of temperature and dietary fatty acids on the fatty acid composition of harpacticoid copepods, for use as a live food for marine fish larvae. Aquaculture 1999, 175, 167–181. [Google Scholar] [CrossRef]
  36. Renaud, S.M.; Zhou, H.C.; Parry, D.L.; Thinh, L.V.; Woo, K.C. Effect of temperature on the growth, total lipid content and fatty acid composition of recently isolated tropical microalgae Isochrysis sp., Nitzschia closterium, Nitzschia paleacea, and commercial species Isochrysis sp. (clone T.ISO). J. Appl. Phycol. 1995, 7, 595–602. [Google Scholar]
  37. Sushchik, N.N.; Kalacheva, G.S.; Zhila, N.O.; Gladyshev, M.I.; Volova, T.G. A temperature dependence of the intra- and extracellular fatty-acid composition of green algae and cyanobacterium. Russ. J. Plant Physiol. 2003, 50, 374–380. [Google Scholar] [CrossRef]
  38. Allen, E.E.; Facciotti, D.; Bartlett, D.H. Monounsaturated but not polyunsaturated fatty acids are required for growth of the deep-sea bacterium Photobacterium profundum ss9 at high pressure and low temperature. Appl. Environ. Microbiol. 1999, 65, 1710–1720. [Google Scholar] [PubMed]
  39. Przytulska, A.; Bartosiewicz, M.; Rautio, M.; Dufresne, F.; Vincent, W.F. Climate effects on high latitude Daphnia via food quality and thresholds. PLoS ONE 2015, 10. [Google Scholar] [CrossRef] [PubMed]
  40. Lewis, R.W. Temperature and pressure effects on the fatty acids of some marine ectotherms. Comp. Biochem. Physiol. 1962, 6, 75–89. [Google Scholar] [CrossRef]
  41. Fishery and Aquaculture Statistics. FishStatJ—Software for Fishery Statistical Time Series. Available online: http://www.Fao.Org/fishery/statistics/software/fishstatj/en (accessed on 4 April 2015).
  42. Woodcock, S.H.; Benkendorff, K. The impact of diet on the growth and proximate composition of juvenile whelks, Dicathais orbita (Gastropoda:Mollusca). Aquaculture 2008, 276, 162–170. [Google Scholar] [CrossRef]
  43. Naegel, L.C.A.; Cooksey, C. Tyrian purple from marine muricids, especially from Plicopurpura pansa (Gould, 1853). J. Shellfish Res. 2002, 21, 193–200. [Google Scholar]
  44. Benkendorff, K. Aquaculture and the production of pharmaceuticals and nutraceuticals. In New Technologies in Aquaculture; Burnell, G., Allan, G., Eds.; Woodhead Publishing Limited: Cambridge, UK, 2009; pp. 866–891. [Google Scholar]
  45. Benkendorff, K. Natural products in the Australian marine invertebrate Dicathais orbita. Mar. Drugs 2013, 11, 1370–1398. [Google Scholar] [CrossRef] [PubMed]
  46. Bech, M. A survey of imposex in muricids from 1996 to 2000 and identification of optimal indicators of tributyltin contamination along the east coast of Phuket Island, Thailand. Mar. Pollut. Bull. 2002, 44, 887–896. [Google Scholar] [CrossRef]
  47. Gibson, C.P.; Wilson, S.P. Imposex still evident in eastern Australia 10 years after tributyltin restrictions. Mar. Environ. Res. 2003, 55, 101–112. [Google Scholar] [CrossRef]
  48. Bech, M.; Strand, J.; Jacobsen, J.A. Development of imposex and accumulation of butyltin in the tropical muricid Thais distinguenda transplanted to a TBT contaminated site. Environ. Pollut. 2002, 119, 253–260. [Google Scholar] [CrossRef]
  49. Talmage, S.S.; Gobler, C.J. Effects of elevated temperature and carbon dioxide on the growth and survival of larvae and juveniles of three species of Northwest Atlantic bivalves. PLoS ONE 2011, 6. [Google Scholar] [CrossRef] [PubMed]
  50. Rais, A.; Miller, N.; Stillman, J.H. No evidence for homeoviscous adaptation in intertidal snails: Analysis of membrane fluidity during thermal acclimation, thermal acclimatization, and across thermal microhabitats. Mar. Biol. 2010, 157, 2407–2414. [Google Scholar] [CrossRef] [PubMed]
  51. Hazel, J.R. Thermal adaptation in biological membranes: Is homeoviscous adaptation the explanation? Annu. Rev. Physiol. 1995, 57, 19–42. [Google Scholar] [CrossRef] [PubMed]
  52. Zhukova, N.V. Lipid classes and fatty acid composition of the tropical nudibranch mollusks Chromodoris sp. and Phyllidia coelestis. Lipids 2007, 42, 1169–1175. [Google Scholar] [CrossRef] [PubMed]
  53. Todd, C.D.; Hughes, S.L.; Marshall, C.T.; MaClean, J.C.; Lonergan, M.E.; Biuw, E.M. Detrimental effects of recent ocean surface warming on growth condition of Atlantic salmon. Glob. Chang. Biol. 2008, 14, 1–13. [Google Scholar]
  54. Hobday, A.J.; Pecl, G.T. Identification of global marine hotspots: Sentinels for change and vanguards for adaptation action. Rev. Fish Biol Fish. 2014, 24, 415–425. [Google Scholar] [CrossRef]
  55. Somero, G.N. The physiology of climate change: How potentials for acclimatization and genetic adaptation will determine “winners” and “losers”. J. Exp. Biol. 2009, 213, 912–920. [Google Scholar] [CrossRef] [PubMed]
  56. Foo, S.A.; Dworjanyn, S.A.; Poore, A.G.B.; Byrne, M. Adaptive capacity of the habitat modifying sea urchin Centrostephanus rodgersii to ocean warming and ocean acidification: Performance of early embryos. PLoS ONE 2012, 7. [Google Scholar] [CrossRef] [PubMed]
  57. Zarai, Z.; Frikha, F.; Balti, R.; Miled, N.; Gargouri, Y.; Mejdoub, H. Nutrient composition of the marine snail (Hexaplex trunculus) from the Tunisian Mediterranean coasts. J. Sci. Food Agric. 2011, 91, 1265–1270. [Google Scholar] [CrossRef] [PubMed]
  58. Yannakopoulos, A.; Tserveni-Gousi, A.; Christaki, E. Enhanced egg production in practice: The case of bio-omega-3 egg. Int. J. Poult. Sci. 2005, 4, 531–535. [Google Scholar]
  59. Milinsk, M.C.; Padre, R.G.; Hayashi, C.; Souza, N.E.; Matsushita, M. Influence of diets enriched with different vegetable oils on the fatty acid profiles of Helix aspersa maxima. Food Chem. 2003, 82, 553–558. [Google Scholar] [CrossRef]
  60. Simopoulus, A.P. Essential fatty acids in health and chronic disease. Am. J. Clin. Nutr. 1999, 70, 560S–569S. [Google Scholar]
  61. Anacleto, P.; Maulvault, A.L.; Bandarra, N.M.; Repolho, T.; Nunes, M.L.; Rosa, R.; Marques, A. Effect of warming on protein, glycogen and fatty acid content of native and invasive clams. Food Res. Int. 2014, 64, 439–445. [Google Scholar] [CrossRef]
  62. Schiffman, E.T.; Coffey, W.D.; Hua, W.; Nunn, B.L.; Dickinson, G.H.; Roberts, S.B. Shotgun proteomics reveals physiological response to ocean acidification in Crassostrea gigas. BMC Genom. 2014, 15. [Google Scholar] [CrossRef]
  63. Sato, N.; Tsuzuki, M.; Kawaguchi, A. Glycerolipid synthesis in Chlorella kessleri 11 h II. Effect of the CO2 concentration during growth. Biochim. Biophys. Acta 2003, 1633, 35–42. [Google Scholar] [PubMed]
  64. Pörtner, H. Ecosystem effects of ocean acidification in times of ocean warming: A physiologist’s view. Mar. Ecol. Prog. Ser. 2008, 373, 203–217. [Google Scholar] [CrossRef]
  65. Zhang, Y.M.; Rock, C.O. Membrane lipid homeostasis in bacteria. Nat. Rev. Microbiol. 2008, 6, 222–233. [Google Scholar] [CrossRef] [PubMed]
  66. Rob, T.; Ogi, T.; Maarisit, W.; Taira, J.; Ueda, K. Isolation of C11 compounds and a cyclopropane fatty acid from an Okinawan ascidian, Diplosoma sp. Molecules 2011, 16, 9972–9982. [Google Scholar] [CrossRef] [PubMed]
  67. Carballeira, N.M.; Montano, N.; Vicente, J.; Rodriguez, A.D. Novel cyclopropane fatty acids from the phospholipids of the Caribbean sponge Pseudospongosorites suberitoides. Lipids 2007, 42, 519–524. [Google Scholar] [CrossRef] [PubMed]
  68. Misra, K.K.; Shkrob, I.; Rakshit, S.; Dembitsky, V.M. Variability in fatty acids and fatty aldehydes in different ograns of two prosobranch gastropod mollusks. Biochem. Syst. Ecol. 2002, 30, 749–761. [Google Scholar] [CrossRef]
  69. Brown, J.L.; Ross, T.; McMeekin, T.A.; Nichols, P.D. Acid habituation of Escherichia coli and the potential role of cyclopropane fatty acids in low pH tolerance. Int. J. Food Microbiol. 1997, 37, 163–173. [Google Scholar] [CrossRef]
  70. Denich, T.J.; Beaudette, L.; Lee, H.; Trevors, J.T. Effect of selected environmental and physico-chemical factors on bacterial cytoplasmic membranes. J. Microbiol. Methods 2003, 52, 149–182. [Google Scholar] [CrossRef]
  71. Christie, W. Mass Spectra of Some Miscellaneous Lipophylic Components. Available online: http://lipidlibrary.aocs.org/Analysis/content.cfm?ItemNumber=39508 (accessed on 18 September 2015).
  72. Nagan, N.; Zoeller, R.A. Plasmalogens: Biosynthesis and functions. Prog. Lipid Res. 2001, 40, 199–229. [Google Scholar] [CrossRef]
  73. Munro, D.; Blier, P.U. Age, diet, and season do not affect longevity-related differences in peroxidation index between Spisula solidissima and Arctica islandica. J. Gerontol. Biol. Sci. 2015, 70, 434–443. [Google Scholar] [CrossRef] [PubMed]
  74. Navy Metoc. Coastal Sea Surface Temperatures. Available online: http://www.metoc.gov.au/products/data/aussst.php (accessed on 25 April 2015).
  75. Poore, A.G.; Graba-Landry, A.; Favret, M.; Sheppard Brennand, H.; Byrne, M.; Dworjanyn, S.A. Direct and indirect effects of ocean acidification and warming on a marine plant-herbivore interaction. Oecologia 2013, 173, 1113–1124. [Google Scholar] [CrossRef]
  76. IPCC. IPCC Summary for Policy Makers. In Climate Change 2013: The Physical Science Basis; Contribution of Working Group 1 to the Fifth Assessment Report of the Intergovernmental Panel on Climate Change; Stocker, T.F., Qin, D., Plattner, G.-K., Tignor, M., Allen, S.K., Boschung, J., Nauels, A., Xia, Y., Bex, V., Midgley, P.M., Eds.; Cambridge University Press: Cambridge, UK, 2013. [Google Scholar]
  77. Pierrot, D.; Lewis, E.; Wallace, D.W.R. MS Excel Program Developed for CO2 System Calculations, Ornl/Cdiac-105; Carbon Dioxide Information Analysis Center, Oak Ridge National Laboratory U.S. Department of Energy: Oak Ridge, TN, USA, 2006. [Google Scholar]
  78. Mehrbach, C.; Culberson, C.H.; Hawley, J.E.; Pytkowicz, R.M. Measurement of the apparent dissociation constants of carbonic acid in seawater at atmospheric pressure. Limnol. Oceanogr. 1973, 18, 897–907. [Google Scholar] [CrossRef]
  79. Dickson, A.G.; Millero, F.J. A comparison of the equilibrium-constants for the dissociation of carbonic-acid in seawater media. Deep Sea Res. Part A Oceanogr. Res. Pap. 1987, 34, 1733–1743. [Google Scholar] [CrossRef]
  80. Kanthilatha, N.; Boyd, W.; Dowell, A.; Mann, A.; Chang, N.; Wolhlmuth, H.; Parr, J. Identification of preserved fatty acids in archeological floor sediments from prehistoric sites at Ban Non Wat and Nong Hua Raet in Northeast Thailand using gas chromatography. J. Archaeol. Sci. 2014, 46, 353–362. [Google Scholar] [CrossRef]

Share and Cite

MDPI and ACS Style

Valles-Regino, R.; Tate, R.; Kelaher, B.; Savins, D.; Dowell, A.; Benkendorff, K. Ocean Warming and CO2-Induced Acidification Impact the Lipid Content of a Marine Predatory Gastropod. Mar. Drugs 2015, 13, 6019-6037. https://doi.org/10.3390/md13106019

AMA Style

Valles-Regino R, Tate R, Kelaher B, Savins D, Dowell A, Benkendorff K. Ocean Warming and CO2-Induced Acidification Impact the Lipid Content of a Marine Predatory Gastropod. Marine Drugs. 2015; 13(10):6019-6037. https://doi.org/10.3390/md13106019

Chicago/Turabian Style

Valles-Regino, Roselyn, Rick Tate, Brendan Kelaher, Dale Savins, Ashley Dowell, and Kirsten Benkendorff. 2015. "Ocean Warming and CO2-Induced Acidification Impact the Lipid Content of a Marine Predatory Gastropod" Marine Drugs 13, no. 10: 6019-6037. https://doi.org/10.3390/md13106019

Article Metrics

Back to TopTop