- Open Access
Assessing diatom-mediated fatty acids in intertidal biofilm: a new conservation concern
Environmental Systems Research volume 10, Article number: 30 (2021)
Biofilm communities on intertidal mudflats are recognized as major producers of nutrients, especially fatty acids. The rising threats posed by both climatic and anthropogenic stressors increase the necessity of understanding and conserving these communities. Shorebirds provide a proxy for studying the complex ecology of biofilm communities because of their heavy reliance on fatty acids from diatomaceous biofilm for successful long-distance migration. Herein, we review biofilm feeding patterns by migratory shorebirds, experimental design considerations for sampling and studying the fatty acid content of biofilm, and the literature describing established and emerging analytical methodology. Techniques for fatty acid analysis include the commonly employed gas chromatography–flame ionization detection (GC/FID) and gas chromatography–mass spectrometry (GC/MS) with derivatization. Liquid chromatography–mass spectrometry (LC/MS) and liquid chromatography–quadrupole time of flight (LC/QTOF) are newly emerging techniques that enable derivatization to be eliminated. In addition, Fourier transform infrared spectroscopy (FT/IR), a common instrument in chemistry laboratories, has applications in fatty acid research, specifically for screening. Using a combination of sampling and analytical methods is necessary for improved understanding of intertidal biofilm, both as a source of essential fatty acids in aquatic systems and a critical food for shorebirds.
Reduced fatty acid production by diatoms in aquatic systems worldwide has emerged as an overarching issue exacerbated by global climate change, with major ramifications to both terrestrial systems and human health (Hixson and Arts 2016; Colombo et al. 2017). Concomitantly, threats to diatomaceous biofilm on intertidal mudflats from climate change and anthropogenic stressors are raising new conservation concerns related to shorebirds (Beninger 2018). Worldwide declines in the abundance of most migratory shorebird species (Clemens et al. 2016; Studds et al. 2017; Rosenberg et al. 2019; Canham et al. 2021) have occurred in tandem with average losses to their intertidal flat habitats at 16% of their total area between 1984 and 2016 (Murray et al. 2019). Mudflat habitats have long been known to provide wide-ranging ecosystem services that support fish and wildlife (Beninger 2018) but have assumed even more importance in recent years with the discovery that many shorebird species consume intertidal biofilm rich in fatty acids produced by diatoms to support their long-distance migration (Schnurr et al. 2019, 2020). Analysis of fatty acids from intertidal biofilm provides critical information on the value of mudflats as habitats for birds, as well as autecological insights into the microbial communities that compose intertidal biofilm (Schnurr et al. 2020; Passarelli et al. 2014). However, techniques for sampling biofilm and assessing the fatty acid content have yet to be fully evaluated or standardized and this will form part of this study’s discussion.
The role of biofilm as a food source for shorebirds appears closely linked to both the physical properties of biofilm itself and the chemistry of diatom-mediated fatty acid production. However, as biofilm is highly labile and logistically challenging to collect, experiments based on biofilm sampling from mudflats need to consider a complex array of spatial–temporal factors, as well as issues including fatty acid sample degradation, and dewatering. For analysis of the samples, an array of techniques exists, including gas chromatography–flame ionization detection (GC/FID) with derivatization (Alfaro et al. 2006; Quinn et al. 2017; Schnurr et al. 2019, 2020), gas chromatography–mass spectrometry (GC/MS) with derivatization (Dahl et al. 2000, 2003; Bergamino et al. 2014), liquid chromatography–mass spectrometry (LC/MS) (Schlotterbeck et al. 2018; Pérez-Navarro et al. 2019), liquid chromatography–quadrupole time of flight (LC/QTOF) (Nguyen et al. 2015; Molina-Calle et al. 2017), and Fourier transform infrared spectroscopy (FT/IR) (Karunathilaka et al. 2017; Aryee et al. 2009; Gieroba et al. 2020). Each technique has a different practicality and suitability for analysing complex mixtures of fatty acids. Furthermore, previously established methods in soil science (Buyer and Sasser 2012) and naphthenic acid analysis (Hao et al. 2005; Shang et al. 2013; Brunswick et al. 2015, 2016, 2017; Bowman et al. 2019) may offer new options with crossover to fatty acid analysis.
The purpose of this review is to evaluate experimental design considerations for sampling and analysing the fatty acid content of biofilm. In order to do so, information on the ecology of intertidal biofilm and the emerging literature on biofilm use by migratory shorebirds was collated to identify the challenges to understanding such an extremely complex medium. Based on the context, we provide initial guidance for assessing diatom-mediated fatty acids in intertidal biofilm and underscore the urgency for further research into this new conservation priority.
What is biofilm?
Microphytobenthos assemblages on intertidal mudflats are primarily composed of microalgae, heterotrophic bacteria, and cyanobacteria, and are frequently dominated by photosynthetic diatoms (Passarelli et al. 2014). These diatoms, together with other microorganisms, secrete a matrix of extracellular polymeric substances (EPS) composed of polysaccharides and proteins, which form biofilms or mats at the surface of the mud. The matrix cover binds the microorganism community onto the sediment, while also adding protection from harsh environmental conditions (Hope et al. 2020). Due to the photosynthetic properties of diatoms and the limited depth of sunlight exposure, biofilm is most abundant in the top 0.5–3 mm of sediment (Macintyre et al. 1996). The biofilm appears either almost colorless or with a green–brown hue, and has a consistency ranging from a low-viscosity slime to a tightly packed gel (Hope et al. 2020; Decho 2000). Microphytobenthic biofilm is a fundamental base to primary production of estuarine ecosystems (Decho 2000). Epibenthic grazers and deposit feeders have a direct dependence on biofilm, which contributes to energy and fatty acid transfer into higher trophic organisms (Herman et al. 2000). Primary production from biofilm is dependent on various environmental conditions, including temperature, salinity, tidal dynamics, light, and nutrients (Underwood and Kromkamp 1999) but, overall, biofilm is estimated to be responsible for approximately 50% of carbon fixation in estuaries (Underwood and Kromkamp 1999). Biofilm also plays a mechanical role in sediment stabilization and protection against scouring by tides and currents (Decho 2000). In the presence of biofilm, the critical erosion threshold of sediments was reported to increase threefold due to the excretion of mucus-like EPS from microorganisms such as diatoms (De Deckere et al. 2001).
Intertidal biofilm is an important food source for migratory shorebirds. Elner et al. (2005) established that western sandpipers (Calidris mauri) and dunlin (Calidris alpina) graze biofilm at a major stopover site on the Fraser River estuary, British Columbia, Canada, on their northward journey from Central and South America to Alaska for breeding. Their findings used scanning electron microscopy to show that the tongues of such birds had bristle-like structures capable of scraping biofilm from the sediment surface. The initial discovery fostered new research on the dynamics and ecology of shorebird feeding on biofilm (Beninger and Elner 2020). Kuwae et al. (2008) investigated western sandpiper grazing using a combination of stable isotope, video recording, and stomach content analysis techniques, and estimated that 45–59% of their diet consisted of biofilm. Subsequently, many other species of shorebirds in tidal flats around the world have been shown to graze biofilm, including the semipalmated sandpiper (Calidris pusilla) and red-necked stint (Calidris ruficollis) (Beninger and Elner 2020; Quinn and Hamilton 2012). The underlying rationale for the birds’ novel food choice may be partly explained by the fact that diatoms in biofilm produce large quantities of lipids and fatty acids (Scholz and Liebezeit 2013; Stonik and Stonik 2015; Mathot et al. 2018), essential nutrients that are not as readily available in their invertebrate prey. Moreover, the consumption of biofilm has been shown to physiologically enhance muscle performance for long distance flight, as well as provide other benefits for successful migration and reproduction (Jenni and Jenni-Eiermann 1998; Price 2010; Guglielmo 2010).
The abundance of diatoms in biofilm is responsible for its high fatty acid content (Passarelli et al. 2014; Scholz and Liebezeit 2013; Stonik and Stonik 2015; Mathot et al. 2018). Fatty acids are organic molecules with a structural formula of CH3(CH2)nCOOH, characterized by a carboxylic acid group and an aliphatic chain of varying length. The aliphatic chain may be saturated, termed saturated fatty acids (SFA), or unsaturated with one or more double bonds, designated monounsaturated fatty acids (MUFA) and polyunsaturated fatty acids (PUFA), respectively (Fig. 1). The content profiles of biofilm lipid and fatty acids are not consistent across a mudflat. Changing environmental conditions affect biofilm composition and productivity, consequently altering fatty acid production in diatoms (Schnurr et al. 2019, 2020; Passarelli et al. 2014; Scholz and Liebezeit 2013). Factors such as temperature, light, season, pH, salinity, and tidal cycles all affect diatom growth, and, thus, the concentration of fatty acids in biofilm (Schnurr et al. 2019, 2020; Hu et al. 2008; Li et al. 2014; Hu and Gao 2006; Venkata Mohan and Devi 2014; Sharma et al. 2012). When conditions for growth are favorable, diatoms produce membrane glycerolipids, which primarily consist of PUFAs that play structural roles (Hu et al. 2008). In contrast, under environmental stress when diatom growth is unfavorable, there is a higher production of SFAs and MUFAs which contribute to storage lipids such as triacylglycerol (TAG) (Li et al. 2014).
What can the study of fatty acids reveal?
Temporally changing mudflat conditions result in fluctuating lipid and fatty acid profiles (Schnurr et al. 2019, 2020; Quinn and Hamilton 2012). This affects the organisms that thrive and forage in estuaries, such as migratory shorebirds. As noted previously, different fatty acids accumulate in biofilm depending on environmental conditions (Schnurr et al. 2019, 2020; Hu et al. 2008; Li et al. 2014; Hu and Gao 2006; Venkata Mohan and Devi 2014; Sharma et al. 2012). As the type of fatty acids present in biofilm help determine its nutritional quality, not all intertidal biofilm is equal. The benefits that specific fatty acids have for shorebirds are extensive, with SFAs, MUFAs, and PUFAs all contributing in different ways (Jenni and Jenni-Eiermann 1998; Price 2010; Guglielmo 2010). Both SFAs and MUFAs comprise a major portion of the energy requirements of migratory birds during long-distance flight. PUFAs, particularly omega-3 (n-3), and omega-6 (n-6) fatty acids, are critical as birds are not able to synthesize these de novo (Stevens 1996) and they have physiological benefits for migration, including increased flight muscle performance and recovery (Price 2010; Guglielmo 2010; Arts et al. 2001; Maillet and Weber 2006). The n-3 fatty acids, eicosapentaenoic acid (EPA), and docosahexaenoic acid (DHA) are all essential nutrients for both birds and other organisms (Maillet and Weber 2007).
Recent studies have linked seasonal changes in fatty acid content of intertidal biofilm to shorebird presence at their migratory stopover site in the Fraser River estuary, such that the period of highest fatty acid abundance aligns with western sandpiper arrival en masse during their northward spring migration (Schnurr et al. 2019, 2020). Similarly, during their southward migration via the upper Bay of Fundy (New Brunswick, Canada), staging semipalmated sandpipers were shown to ingest more biofilm in areas of the mudflats where PUFAs were most abundant (Quinn et al. 2017). While the fundamental importance of fatty acids to the health of aquatic systems is well established (Maillet and Weber 2007), targeted multidisciplinary research is needed to test the various hypotheses on the nutritional properties of biofilm and the effect that its different fatty acid compositions have on avian migratory performance (Schnurr et al. 2019; Price 2010).
The use of fatty acids as biomarkers has broad applications in benthic ecosystems. Although not a perfect trophic marker, they can provide useful qualitative information (Schnurr et al. 2020; Dalsgaard et al. 2003; Kelly and Scheibling 2012). The fatty acid profile of mudflat biofilm can provide insight into its microorganism community composition. However, fluctuations in diatom growth conditions that result in changes in fatty acid content and the fact that multiple estuarine invertebrates may contain the same fatty acids are considered complications to using fatty acids as biomarkers (Dalsgaard et al. 2003; Kelly and Scheibling 2012). Schnurr et al. (2020) studied seasonal changes in the ratios of specific fatty acids compared to the total fatty acid concentrations for particular microorganism groups as chemotaxonomic biomarkers in the Fraser River estuary. They observed community compositional changes in fatty acid biomarkers for bacteria (odd-chain length fatty acids), cyanobacteria (18:2n-6), dinoflagellates (22:6n-3 and 18:1n-9), and diatoms (16:1n-7 and 20:5n-3). Further, from spring to summer, diatom abundance in the intertidal biofilm decreased, while the abundance of cyanobacteria, bacteria, and dinoflagellates increased. Thus, the availability of diatoms appears correlated with the seasonal abundance of shorebirds and may help explain why their southward summer migration through the Fraser River estuary is protracted and dissipated relative to the concentrated spring migration (Schnurr et al. 2020).
Fatty acid biomarkers have been used extensively for studying a range of consumer and food web relationships (Dalsgaard et al. 2003; Kelly and Scheibling 2012; Dahl et al. 2003; Alfaro et al. 2006; Van Der Heijden et al. 2019). Dahl et al. (2003) discerned food chain linkages by studying the fatty acid profiles in muscle tissues of three bird species: common eider (Somateria mollissima), black-legged kittiwake (Rissa tridactyla), and northern fulmar (Fulmarus glacialis). From the fatty acid composition, they found that, of the three species, the common eider had the lowest trophic level and the strongest relationship with the benthic food chain. The northern fulmar was found to have the highest trophic level, followed by the black-legged kittiwake, with both of these species associated with the pelagic food chain. Alfaro et al. (2006) investigated the use of fatty acid biomarkers and stable isotope analysis to study estuarial food webs in northern New Zealand. Information on the preferred food sources of various organisms including fish, filter feeding invertebrates, and shrimp was found using fatty acid biomarkers. Using this method, they also found that higher consumers have a mixed diet consisting of a wide variety of prey from different trophic linkages (Alfaro et al. 2006). When used in tandem with other methods, such as stable isotope analysis and compound specific isotope analysis, fatty acid biomarkers become even more powerful tools for studying consumer-resource systems and community composition (Dalsgaard et al. 2003; Kelly and Scheibling 2012; Alfaro et al. 2006; Van Der Heijden et al. 2019).
What experimental conditions and designs should be considered?
When designing experiments for studying the relationship between fatty acids, biofilm, and ecosystems, researchers must take spatial and temporal variability into account. On intertidal mudflats, the composition of biofilm varies spatially between different areas and temporally throughout the year (Bergamino et al. 2014). Frankenbach et al. (2020) studied the primary productivity of microphytobenthos in the Ria de Aveiro Estuary, Portugal, and placed an emphasis on characterizing the local variability. To study spatial variability, samples were collected in areas with differing sediment grain size, salinity, hydrodynamics, and distances to the mouth of the estuary. To study temporal variability, samples were taken each hour, fortnight, and season. Including these spatial and temporal variables in the study allowed them to acquire detailed data on photosynthetic productivity, enabling accurate estuary carbon-fixation estimates. Similarly, Bergamino et al. (2014) examined spatial and temporal changes in organic material and sediments in the Kowie Estuary, South Africa. Sampling took place in each of the four seasons and were collected in three areas of the estuary with different marine influences. Using both fatty acid and stable isotope analyses, the organic compositions were found to vary spatially and temporally. While adopting a spatial and temporal approach to estuarial research is time-consuming and complex, it should be explicitly considered as it provides deeper insight into ecosystem dynamics (Bergamino et al. 2014; Frankenbach et al. 2020).
Biofilm contamination and degradation are unavoidable factors that must also be carefully considered. Quinn et al. (2017) noted that sediment contamination is inescapable in biofilm samples collected in the field and that the presence of sediment can alter the measured fatty acid content. In Schnurr et al. (2019), while invertebrates larger than 0.5 mm were removed from biofilm samples before analyses, meiofaunal invertebrates smaller than 0.5 mm remained. As a result, a small portion of the total fatty acids and biomass measured could be attributed to meiofauna. The degradation of fatty acid samples can also corrupt targeted organic compounds unless samples are frozen promptly after collection (Rudy et al. 2016). Degradation can occur through hydrolysis of samples by lipases present in aqueous conditions and through other chemical reactions such as oxidation. Both processes result in changes to the total fatty acid content, potentially compromising the integrity of a study. To avoid this, if immediate analysis is not available, samples should ideally be stored at − 80 °C following collection (Rudy et al. 2016; Metherel et al. 2013; Sasaki and Capuzzo 1984). The low temperature reduces deterioration of the fatty acids present in the samples.
Other sediment effects, such as compaction and dewatering, can change how algal biomass data is processed and interpreted. A study in the Eden Estuary, Scotland (Perkins et al. 2003) investigated the effects of sediment compaction and dewatering on algal biomass data. The results differed based on whether the algal biomass data (expressed as chlorophyll a and EPS) was presented as a concentration (µg Chl a per mg dry weight sediment) or as content (µg Chl a per cm2 sediment). A theoretical model was used to predict the effects that these sediment transformations had on Chl a and EPS values over the course of a 6-h tidal emersion period. The predicted algal biomass values from the theoretical model were then compared to experimentally obtained Chl a and EPS values for diel and seasonal emersion periods. The predicted biomass Chl a values expressed in units of concentration and content showed opposing trends in the face of dewatering effects (Table 1). These findings highlight the importance of correcting for dewatering effects in sediment data and understanding the implications of expressing algal biomass as either concentration or content.
What analytical techniques are available for fatty acid analysis?
Gas chromatography (GC) based methods are cost effective, robust, and commonly employed to identify and quantify fatty acids (Fisk et al. 2014). While these aspects make GC a widely accepted technique, due to the low volatility of targeted fatty acids, extensive sample preparation and derivatization is often required (Fisk et al. 2014; Drozd 1975). Sample preparation for fatty acids frequently involves a lipid extraction method, such as the Folch Method (Folch et al. 1957). Another well-known lipid extraction method is the Bligh and Dyer method (Bligh and Dyer 1959), which, along with the Folch method, have many published modifications involving different solvents and solvent ratios (Kumar et al. 2015; Ren et al. 2017). The optimal choice of extraction method depends on the lipid chemical behavior and the nature of the sample matrix, as well as the researcher’s specific recovery goals and time frame (Kumar et al. 2015; Work and Work 1972; Ulmer et al. 2018). Extraction preferentially also includes an internal standard that displays similar behaviour to the analyte, in order to give an indication of fatty acid recovery. Deuterated fatty acids and odd-numbered long chain fatty acids such as tridecanoic, heptadecanoic, and nonadecanoic acids have previously been used as internal standards (Quehenberger et al. 2011; Reich et al. 2012; Politz et al. 2013).
Extracted fatty acids are frequently analysed by both GC/FID and GC/MS (Tang and Row 2013). GC/FID does not generally require prior derivatization of the fatty acids before column separation and FID detection. FID detection applies a destructive air–hydrogen flame with the electric current from this combustion being measured by a detector. The drawbacks of FID detection are that all peaks are determined by current response rather than accurate mass, as is the case by a mass spectrometer-based method. For fatty acids analysis, GC/MS in positive electron ionization mode (EI+) is more sensitive than GC/FID, has better specificity, and provides more compound structural information. By GC/MS, compounds are separated on a GC capillary column, with eluting analytes ionized by electron bombardment, resulting in fragmentation ions that are detected by their mass to charge ratios. However, to be amenable to GC/MS EI+ detection, fatty acids require derivatization (Fisk et al. 2014). Derivatization involves the replacement of a fatty acid functional group in order to increase the volatility of the analyte (Fisk et al. 2014; Drozd 1975). A commonly used derivatization method for fatty acids involves methylation with reagents such as boron trifluoride (BF3) and methanol to create fatty acid methyl esters (FAME), which possess volatilities sufficient for GC analysis (Topolewska et al. 2015). Other less common derivatization methods used for analysis of fatty acids include silylation to generate trimethylsilyl (TMSi) esters, as well as the production of tert-butyldimethylsilyl (TBDMSi) derivatives (Woo and Kim 1999; Moon et al. 2018). The choice of derivatization method generally depends upon the demonstrated stability of the product analyte and can be dependent upon the matrix. Each derivatization method has benefits as well as drawbacks; for example, the synthesis of FAME derivatives using BF3 and methanol have been found to have a lower reaction yield compared to the production of TBDMSi derivatives by silylation using N-tert-butyldimethylsilyl-N-methyltrifluoroacetamide (MTBSTFA) as a reagent (Topolewska et al. 2015). Additionally, TBDMSi derivatives have higher hydrolytic stability compared to TMSi derivatives, which can easily hydrolyse when exposed to water (Woo and Kim 1999). Several previously mentioned papers employed GC, as well as the FAME method of derivatization, including Schnurr et al. (2019, 2020), Quinn et al. (2017), Dahl et al. (2003), Alfaro et al. (2006), and Bergamino et al. (2014). A summary of the advantages and disadvantages of GC/FID and GC/MS, as well as other analytical techniques discussed in following sections, is provided in Table 2. In addition to the previously mentioned techniques, the use of comprehensive two-dimensional gas chromatography (GC × GC) has been reported for the analysis of algal fatty acids (Akoto et al. 2008; Gu et al. 2011). This technique has been found to have lower detection limits and improved separation, as compared to traditional GC analysis (Akoto et al. 2008; De Geus et al. 2001).
GC methods are well established in soil science, in which the use of phospholipid fatty acids (PLFAs) as biomarkers has been gaining popularity since the 1990s (Frostegård et al. 2011). Mudflats and terrestrial soils have many similarities and involve the study of microorganisms living in complex substrates. Likely, PLFAs could be adapted for studying intertidal biofilm. Buyer and Sasser (2012) used a high throughput GC sample preparation method for soil analysis that could potentially be applied to biofilm. Their method described the preparation of ninety-six samples, as well as blanks, in only a day and a half. The previously mentioned Bligh–Dyer lipid extraction (Bligh and Dyer 1959) was used, as well as a solid phase extraction. Such an efficient method could reduce the extensive preparation time when preparing biofilm samples for GC analysis. Due to similarities between fatty acids and naphthenic acids, it is likely that the two will behave in comparable ways when analyzed in the laboratory. Therefore, additional potential procedures based on this association need to be reviewed. Fatty acids and naphthenic acids are very similar chemical species, both containing a carboxylic acid with an aliphatic chain (Table 3) (Shang et al. 2013). The main difference between the two is the presence of saturated ring structures, such as cyclohexyl and cyclopentyl, on naphthenic acids. These descriptions encompass the classic definition of naphthenic acids, which can be described with the formula CnH2n+zO2 (Shang et al. 2013). The “z” in this formula is either zero, or an even, negative integer, which represents the hydrogen deficiency. It is expected that established methods from naphthenic acid studies could be applied to fatty acids in the future, reducing the time required for planning and designing an experiment. Comprehensive two-dimensional gas chromatography time-of-flight mass spectrometry (GC × GC/TOFMS) has been used for naphthenic acid mixture characterization along with the use of mass deconvolution software and spectral database searches (Hao et al. 2005). Recently, this GC × GC/TOFMS technique has been applied to the naphthenic acid monitoring of a wetland reclamation site in the Athabasca oils sands region in northern Alberta, Canada (Bowman et al. 2019). Both of these studies involved the derivatization of the samples into methyl esters. The internal standard used in the second method was methyl 2-hexyl-decanoate dissolved in dichloromethane (Bowman et al. 2019). Techniques for liquid chromatography analysis of naphthenic acids are discussed in the following section.
Liquid chromatography is a separation technique that can be applied to fatty acid analysis (Pérez-Navarro et al. 2019; Molina-Calle et al. 2017; Nguyen et al. 2015; Brondz 2002). While sample clean-up and derivatization prior to LC analysis can improve the selectivity and sensitivity of the instrument, it is not absolutely required (Brondz 2002; Snyder and Kirkland 1979; Kataoka 2017). When performing trace analysis in ppb and ppt, contamination is a major problem, especially when studying fatty acids that are commonly found in the laboratory environment (Mizuike and Pinta 1978). Application of direct LC analysis without derivatization can avoid the time-consuming preparative steps that risk contamination from ubiquitous fatty acids in the lab. Palmitic acid and stearic acid are fatty acids that have been found to be common laboratory contaminants in disposable plasticware used to avoid cross contamination between samples (Cheng and Yu 2020). Using glass, rather than plastic, helped to reduce the contamination and decrease the limit of detection when studying these fatty acids. Ubiquitous fatty acid contaminants have also been found in commercially available solid phase extraction columns, which can be particularly troubling when using these products during sample preparation (Cavonius and Carlsson 2015). It is important to be aware of these pervasive fatty acids and plan experiments to minimize the effect of contamination. As with other chromatographic techniques, compound separation occurs by selection of a column phase that shows affinity for the analytes of interest, in combination with complementary mobile phases. There are numerous publications listing LC parameter choices including the LC column, mobile phases, and eluent gradients that have achieved separation of fatty acids. Chu et al. (2009), Samburova et al. (2013), and Schlotterbeck et al. (2018) are examples of literature that describe the use of ultra and ultra-high performance liquid chromatography to study fatty acids, with Schlotterbeck et al. (2018) describing methods for both targeted and untargeted analysis. The LC parameters used in these papers are summarized in Table 4 for the benefit of future researchers.
LC/MS and LC/QTOF are widely used LC instruments that are highly selective and sensitive, with their ability to provide analyte structural information showing promise for applications to fatty acid analysis. Advantages and disadvantages for LC/MS and LC/QTOF are provided in Table 2 to contrast the previously mentioned GC methods. Pérez-Navarro et al. (2019) used liquid chromatography tandem mass spectrometry (LC/MS/MS) to determine the free fatty acid profile in Vitis vinifera grape cultivars. Molina-Calle et al. (2017) tentatively identified fatty acids using LC/QTOF to research Stevia leaves. Nguyen et al. (2015) used a combination of GC/QTOF and LC/QTOF to analyze both fatty acids and triacylglycerols from fruits in the Apiaceae family. Additionally, several established LC techniques for naphthenic acid analysis have potential applications in fatty acid method development. Shang et al. (2013) described a specific and sensitive LC/MS method for quantification of naphthenic acids that included the use of large volume injection. Sample preparation was simple, with samples only being centrifuged after the addition of the internal standard. Brunswick et al. (2015, 2016, 2017) reported a high resolution, high throughput, LC/QTOF method that also involved limited sample preparation by pH adjustment to an alkaline pH > 10. Applicability of the naphthenic acid methodology to fatty acids is supported by their use of deuterated fatty acids as internal standards and reference standards.
Fourier transform infrared (FT/IR) spectroscopy
Fourier transform infrared (FT/IR) spectroscopy is a technique used to analyse the characteristic molecular absorption of infrared light to determine structure, typically in the region of 4000–400 cm−1 (Griffiths 1983; Faix 1992). It is a rapid and non-destructive analytical technique with a broad range of applications and is readily available in many laboratories for routine work (Gieroba et al. 2020). FT/IR was used by Gieroba et al. (2020) to analyze bacterial biofilm on human teeth. They were able to identify lipids and proteins in biological samples but noted caution because the presence of water can distort results. The use of FT/IR to study free fatty acids in fish oils was described by Aryee et al. (2009). FT/IR was demonstrated to be an adequate method for determination of free fatty acids due to its simplicity and reproducibility. FT/IR could have potential applications for studying biofilm but is most likely reserved as a rapid and less sensitive screening tool to initially assess the composition of the sample.
The present review synthesizes literature on the roles and properties of intertidal biofilm, with special reference to migratory shorebirds, and underscores the value of diatoms as a major source of fatty acids. In support of future studies, experimental design factors such as sample storage, spatial–temporal sampling, and accounting for dewatering effects on sediment are emphasized. The established methodologies of biofilm fatty acid analysis including GC/FID and GC/MS remain effective, but more forthcoming techniques, including LC/MS and LC/QTOF should be considered, alongside the possible incorporation of FT/IR as a screening tool. In addition, potential is noted for applying soil science and naphthenic acid techniques to fatty acid research.
Worldwide, a growing appreciation for the benefits of intertidal mudflat habitats has given alarm to the potential negative effects of climate change and coastal development to both mudflat quality and quantity. In particular, as the quality of intertidal biofilm can be gauged by the production of diatom-mediated fatty acids (Schnurr et al. 2020; Canham et al. 2021), the predicted reduced production of polyunsaturated fatty acids in algae due to global warming (Hixson and Arts 2016; Colombo et al. 2020) compounded by losses in mudflat quantity (Murray et al. 2019) are likely to result in irreversible and fundamentally adverse changes to these coastal systems. Establishing robust and routine analytical techniques to temporally monitor the quality and condition of mudflats, in tandem with research programs to achieve a deeper understanding of mudflat ecology, are core to effective conservation. In particular, determining the quality of biofilm as a food source for migratory shorebirds is of prime importance. In this regard, the determination of specific SFAs, MUFAs, and PUFAs ratios are a key goal. Perhaps the overarching challenge is to discover the specific triggers to the lipid accumulation in diatoms and related responses in intertidal biofilm, given the fundamental role of mudflats in supporting not only migratory shorebirds but also the whole ecosystem of surrounding wildlife and fisheries (Mathot et al. 2018).
Availability of data and materials
Gas chromatography–flame ionization detection
Gas chromatography–mass spectrometry
Liquid chromatography–mass spectrometry
Liquid chromatography–quadrupole time-of-flight
Fourier transform infrared spectroscopy
Extracellular polymeric substances
Saturated fatty acid
Monounsaturated fatty acids
Polyunsaturated fatty acids
- Chl a :
Positive electron ionization mode
- BF3 :
Fatty acid methyl esters
- GC × GC:
Two-dimensional gas chromatography
Phospholipid fatty acids
- GC × GC/TOFMS:
Two-dimensional gas chromatography time-of-flight mass spectrometry
Liquid chromatography tandem mass spectrometry
Akoto L, Stellaard F, Irth H, Vreuls RJJ, Pel R (2008) Improved fatty acid detection in micro-algae and aquatic meiofauna species using a direct thermal desorption interface combined with comprehensive gas chromatography–time-of-flight mass spectrometry. J Chromatogr A 1186:254–261. https://doi.org/10.1016/j.chroma.2008.02.005
Alfaro AC, Thomas F, Sergent L, Duxbury M (2006) Identification of trophic interactions within an estuarine food web (northern New Zealand) using fatty acid biomarkers and stable isotopes. Estuar Coast Shelf Sci 70:271–286. https://doi.org/10.1016/j.ecss.2006.06.017
Arts MT, Ackman RG, Holub BJ (2001) “Essential fatty acids” in aquatic ecosystems: a crucial link between diet and human health and evolution. Can J Fish Aquat Sci 58:122–137. https://doi.org/10.1139/cjfas-58-1-122
Aryee ANA, van de Voort FR, Simpson BK (2009) FTIR determination of free fatty acids in fish oils intended for biodiesel production. Process Biochem 44:401–405. https://doi.org/10.1016/j.procbio.2008.12.004
Beninger PG (2018) Mudflat ecology. Springer International Publishing, Cham
Beninger PG, Elner RW (2020) On the tip of the tongue: natural history observations that transformed shorebird ecology. Ecosphere. https://doi.org/10.1002/ecs2.3133
Bergamino L, Dalu T, Richoux NB (2014) Evidence of spatial and temporal changes in sources of organic matter in estuarine sediments: stable isotope and fatty acid analyses. Hydrobiologia 732:133–145. https://doi.org/10.1007/s10750-014-1853-1
Bligh EG, Dyer WJ (1959) A rapid method of total lipid extraction and purification. Can J Biochem Physiol 37:911–917. https://doi.org/10.1139/o59-099
Bowman DT, Warren LA, McCarry BE, Slater GF (2019) Profiling of individual naphthenic acids at a composite tailings reclamation fen by comprehensive two-dimensional gas chromatography–mass spectrometry. Sci Total Environ 649:1522–1531. https://doi.org/10.1016/j.scitotenv.2018.08.317
Brondz I (2002) Development of fatty acid analysis by high-performance liquid chromatography, gas chromatography, and related techniques. Anal Chim Acta 465:1–37
Brunswick P, Shang D, van Aggelen G, Hindle R, Hewitt LM, Frank RA, Haberl M, Kim M (2015) Trace analysis of total naphthenic acids in aqueous environmental matrices by liquid chromatography/mass spectrometry–quadrupole time of flight mass spectrometry direct injection. J Chromatogr A 1405:49–71. https://doi.org/10.1016/j.chroma.2015.05.048
Brunswick P, Hewitt LM, Frank RA, Van Aggelen G, Kim M, Shang D (2016) Specificity of high resolution analysis of naphthenic acids in aqueous environmental matrices. Anal Methods 8:6764–6773. https://doi.org/10.1039/c6ay01912a
Brunswick P, Hewitt LM, Frank RA, Kim M, van Aggelen G, Shang D (2017) A traceable reference for direct comparative assessment of total naphthenic acid concentrations in commercial and acid extractable organic mixtures derived from oil sands process water. J Environ Sci Health A 52:274–280. https://doi.org/10.1080/10934529.2016.1253399
Buyer JS, Sasser M (2012) High throughput phospholipid fatty acid analysis of soils. Appl Soil Ecol 61:127–130. https://doi.org/10.1016/j.apsoil.2012.06.005
Canham R, Flemming SA, Hope DD, Drever MC (2021) Sandpipers go with the flow: correlations between estuarine conditions and shorebird abundance at an important stopover on the Pacific Flyway. Ecol Evol 11:2828–2841. https://doi.org/10.1002/ece3.7240
Cavonius LR, Carlsson N-G (2015) Fatty acid contaminations originating from commercially available solid phase-extraction columns. Chem Sci Rev Lett 16:1206–1209
Cheng YY, Yu JZ (2020) Minimizing contamination from plastic labware in the quantification of C16 and C18 fatty acids in filter samples of atmospheric particulate matter and their utility in apportioning cooking source contribution to urban PM2.5. Atmosphere 11:1120. https://doi.org/10.3390/atmos11101120
Chu X, Zhao T, Zhang Y, Zhao A, Zhou M, Zheng X, Dan M, Jia W (2009) Determination of 13 free fatty acids in pheretima using ultra-performance LC–ESI–MS. Chromatographia 69:645–652. https://doi.org/10.1365/s10337-009-0962-x
Clemens RS, Rogers DI, Hansen BD, Gosbell K, Minton CDT, Straw P, Bamford M, Woehler EJ, Milton DA, Weston MA, Venables B, Weller D, Hassell C, Rutherford B, Onton K, Herrod A, Studds CE, Choi CY, Dhanjal-Adams KL, Murray NJ, Skilleter GA, Fuller RA (2016) Continental-scale decreases in shorebird populations in Australia. Emu 116:119–135. https://doi.org/10.1071/MU15056
Colombo SM, Wacker A, Parrish CC, Kainz MJ, Arts MT (2017) A fundamental dichotomy in long-chain polyunsaturated fatty acid abundance between and within marine and terrestrial ecosystems. Environ Rev 25:163–174
Colombo SM, Rodgers TFM, Diamond ML, Bazinet RP, Arts MT (2020) Projected declines in global DHA availability for human consumption as a result of global warming. Ambio 49:865–880. https://doi.org/10.1007/s13280-019-01234-6
Dahl TM, Lydersen C, Kovacs KM, Falk-Petersen S, Sargent J, Gjertz I, Gulliksen B (2000) Fatty acid composition of the blubber in white whales (Delphinapterus leucas). Polar Biol 23:401–409. https://doi.org/10.1007/s003000050461
Dahl T, Falk-Petersen S, Gabrielsen G, Sargent J, Hop H, Millar R (2003) Lipids and stable isotopes in common eider, black-legged kittiwake and northern fulmar: a trophic study from an Arctic fjord. Mar Ecol Prog Ser 256:257–269. https://doi.org/10.3354/meps256257
Dalsgaard J, St. John M, Kattner G, Müller-Navarra D, Hagen W (2003) Fatty acid trophic markers in the pelagic marine environment. Adv Mar Biol 46:225–340
De Deckere EMGT, Tolhurst TJ, De Brouwer JFC (2001) Destabilization of cohesive intertidal sediments by infauna. Estuar Coast Shelf Sci 53:665–669. https://doi.org/10.1006/ecss.2001.0811
De Geus HJ, Aidos I, De Boer J, Luten JB, Brinkman UAT (2001) Characterisation of fatty acids in biological oil samples using comprehensive multidimensional gas chromatography. J Chromatogr A 910:95–103. https://doi.org/10.1016/S0021-9673(00)01183-3
Decho AW (2000) Microbial biofilms in intertidal systems: an overview. Cont Shelf Res 20:1257–1273. https://doi.org/10.1016/S0278-4343(00)00022-4
Drozd J (1975) Chemical derivatization in gas chromatography. J Chromatogr A 113:303–356. https://doi.org/10.1016/S0021-9673(00)95303-2
Elner RW, Beninger PG, Jackson DL, Potter TM (2005) Evidence of a new feeding mode in western sandpiper (Calidris mauri) and dunlin (Calidris alpina) based on bill and tongue morphology and ultrastructure. Mar Biol 146:1223–1234. https://doi.org/10.1007/s00227-004-1521-5
Faix O (1992) Fourier transform infrared spectroscopy. Springer, Berlin, pp 83–109
Fisk HL, West AL, Childs CE, Burdge GC, Calder PC (2014) The use of gas chromatography to analyze compositional changes of fatty acids in rat liver tissue during pregnancy. J vis Exp 85:51445. https://doi.org/10.3791/51445
Folch J, Lees M, Sloane GH (1957) A simple method for the isolation and purification of total lipides from animal tissues. J Biol Chem 226:497–509
Frankenbach S, Ezequiel J, Plecha S, Goessling JW, Vaz L, Kühl M, Dias JM, Vaz N, Serôdio J (2020) Synoptic spatio-temporal variability of the photosynthetic productivity of microphytobenthos and phytoplankton in a tidal Estuary. Front Mar Sci 7:170. https://doi.org/10.3389/fmars.2020.00170
Frostegård Å, Tunlid A, Bååth E (2011) Use and misuse of PLFA measurements in soils. Soil Biol Biochem 43:1621–1625. https://doi.org/10.1016/j.soilbio.2010.11.021
Gieroba B, Krysa M, Wojtowicz K, Wiater A, Pleszczyńska M, Tomczyk M, Sroka-Bartnicka A (2020) The FT-IR and raman spectroscopies as tools for biofilm characterization created by cariogenic streptococci. Int J Mol Sci. https://doi.org/10.3390/ijms21113811
Griffiths PR (1983) Fourier transform infrared spectrometry. Science 222:297–302. https://doi.org/10.1126/science.6623077
Gu Q, David F, Lynen F, Vanormelingen P, Vyverman W, Rumpel K, Xu G, Sandra P (2011) Evaluation of ionic liquid stationary phases for one dimensional gas chromatography–mass spectrometry and comprehensive two dimensional gas chromatographic analyses of fatty acids in marine biota. J Chromatogr A 1218:3056–3063. https://doi.org/10.1016/j.chroma.2011.03.011
Guglielmo CG (2010) Move that fatty acid: fuel selection and transport in migratory birds and bats. Integrative and comparative biology. Oxford Academic, Oxford, pp 336–345
Hao C, Headley JV, Peru KM, Frank R, Yang P, Solomon KR (2005) Characterization and pattern recognition of oil-sand naphthenic acids using comprehensive two-dimensional gas chromatography/time-of-flight mass spectrometry. J Chromatogr A 1067:277–284. https://doi.org/10.1016/j.chroma.2005.01.041
Herman P, Middelburg J, Widdows J, Lucas C, Heip C (2000) Stable isotopes as trophic tracers: combining field sampling and manipulative labelling of food resources for macrobenthos. Mar Ecol Prog Ser 204:79–92. https://doi.org/10.3354/meps204079
Hixson SM, Arts MT (2016) Climate warming is predicted to reduce omega-3, long-chain, polyunsaturated fatty acid production in phytoplankton. Glob Change Biol 22:2744–2755. https://doi.org/10.1111/gcb.13295
Hope JA, Paterson DM, Thrush SF (2020) The role of microphytobenthos in soft-sediment ecological networks and their contribution to the delivery of multiple ecosystem services. J Ecol 108:815–830. https://doi.org/10.1111/1365-2745.13322
Hu H, Gao K (2006) Response of growth and fatty acid compositions of Nannochloropsis sp. to environmental factors under elevated CO2 concentration. Biotechnol Lett 28:987–992. https://doi.org/10.1007/s10529-006-9026-6
Hu Q, Sommerfeld M, Jarvis E, Ghirardi M, Posewitz M, Seibert M, Darzins A (2008) Microalgal triacylglycerols as feedstocks for biofuel production: perspectives and advances. Plant J 54:621–639
Jenni L, Jenni-Eiermann S (1998) Fuel supply and metabolic constraints in migrating birds. J Avian Biol 29:521–528. https://doi.org/10.2307/3677171
Karunathilaka SR, Mossoba MM, Chung JK, Haile EA, Srigley CT (2017) Rapid prediction of fatty acid content in marine oil omega-3 dietary supplements using a portable Fourier transform infrared (FTIR) device & partial least-squares regression (PLSR) analysis. J Agric Food Chem 65:224–233. https://doi.org/10.1021/acs.jafc.6b04463
Kataoka H (2017) Sample preparation for liquid chromatography. Liquid chromatography. Elsevier, Amsterdan, pp 1–37
Kelly J, Scheibling R (2012) Fatty acids as dietary tracers in benthic food webs. Mar Ecol Prog Ser 446:1–22. https://doi.org/10.3354/meps09559
Kumar RR, Rao PH, Arumugam M (2015) Lipid extraction methods from microalgae: a comprehensive review. Front Energy Res. https://doi.org/10.3389/fenrg.2014.00061
Kuwae T, Beninger PG, Decottignies P, Mathot KJ, Lund DR, Elner RW (2008) Biofilm grazing in a higher vertebrate: the western sandpiper, Calidris mauri. Ecology 89:599–606. https://doi.org/10.1890/07-1442.1
Li HY, Lu Y, Zheng JW, Yang WD, Liu JS (2014) Biochemical and genetic engineering of diatoms for polyunsaturated fatty acid biosynthesis. Mar Drugs 12:153–166. https://doi.org/10.3390/md12010153
Macintyre HL, Geider RJ, Miller DC (1996) Microphytobenthos: the ecological role of the “secret garden” of unvegetated, shallow-water marine habitats. I. Distribution, abundance and primary production. Estuaries Coast 19:186–201. https://doi.org/10.2307/1352224
Maillet D, Weber JM (2006) Performance-enhancing role of dietary fatty acids in a long-distance migrant shorebird: the semipalmated sandpiper. J Exp Biol 209:2686–2695. https://doi.org/10.1242/jeb.02299
Maillet D, Weber JM (2007) Relationship between n-3 PUFA content and energy metabolism in the flight muscles of a migrating shorebird: evidence for natural doping. J Exp Biol 210:413–420. https://doi.org/10.1242/jeb.02660
Mathot KJ, Piersma T, Elner RW (2018) Shorebirds as integrators and indicators of mudflat ecology. Mudflat ecology. Springer International Publishing, Cham, pp 309–338
Metherel AH, Aristizabal Henao JJ, Stark KD (2013) EPA and DHA levels in whole blood decrease more rapidly when stored at −20 °C as compared with room temperature, 4 and −75 °C. Lipids 48:1079–1091. https://doi.org/10.1007/s11745-013-3827-x
Mizuike A, Pinta M (1978) Contamination in trace analysis. Pure Appl Chem 40:1519–1529
Molina-Calle M, Priego-Capote F, Luque de Castro MD (2017) Characterization of Stevia leaves by LC–QTOF MS/MS analysis of polar and non-polar extracts. Food Chem 219:329–338. https://doi.org/10.1016/j.foodchem.2016.09.148
Moon JY, Kong TY, Jang HJ, Kang HC, Cho YY, Lee JY, Lee HS (2018) Simultaneous quantification of 18 saturated and unsaturated fatty acids and 7 sterols as their tert-butyldimethylsilyl derivatives in human saliva using gas chromatography–tandem mass spectrometry. J Chromatogr B Anal Technol Biomed Life Sci 1092:114–121. https://doi.org/10.1016/j.jchromb.2018.06.003
Murray NJ, Phinn SR, DeWitt M, Ferrari R, Johnston R, Lyons MB, Clinton N, Thau D, Fuller RA (2019) The global distribution and trajectory of tidal flats. Nature 565:222–225. https://doi.org/10.1038/s41586-018-0805-8
Nguyen T, Aparicio M, Saleh MA, McPhee DJ (2015) Accurate mass GC/LC-quadrupole time of flight mass spectrometry analysis of fatty acids and triacylglycerols of spicy fruits from the apiaceae family. Molecules 20:21421–21432. https://doi.org/10.3390/molecules201219779
Passarelli C, Meziane T, Thiney N, Boeuf D, Jesus B, Ruivo M, Jeanthon C, Hubas E (2014) Seasonal variations of the composition of microbial biofilms in sandy tidal flats: focus of fatty acids, pigments and exopolymers. Estuar Coast Shelf Sci 153:29–37. https://doi.org/10.1016/j.ecss.2014.11.013
Pérez-Navarro J, Da Ros A, Masuero D, Izquierdo-Cañas PM, Hermosín-Gutiérrez I, Gómez-Alonso S, Mattivi F, Vrhovsek U (2019) LC–MS/MS analysis of free fatty acid composition and other lipids in skins and seeds of Vitis vinifera grape cultivars. Food Res Int 125:108556. https://doi.org/10.1016/j.foodres.2019.108556
Perkins RG, Honeywill C, Consalvey M, Austin HA, Tolhurst TJ, Paterson DM (2003) Changes in microphytobenthic chlorophyll a and EPS resulting from sediment compaction due to de-watering: opposing patterns in concentration and content. Cont Shelf Res 23:575–586. https://doi.org/10.1016/S0278-4343(03)00006-2
Politz M, Lennen R, Pfleger B (2013) Quantification of bacterial fatty acids by extraction and methylation. Bio-Protoc. https://doi.org/10.21769/bioprotoc.950
Price ER (2010) Dietary lipid composition and avian migratory flight performance: development of a theoretical framework for avian fat storage. Comp Biochem Physiol A Mol Integr Physiol 157:297–309. https://doi.org/10.1016/j.cbpa.2010.05.019
Quehenberger O, Armando AM, Dennis EA (2011) High sensitivity quantitative lipidomics analysis of fatty acids in biological samples by gas chromatography–mass spectrometry. Biochim Biophys Acta Mol Cell Biol Lipids 1811:648–656
Quinn JT, Hamilton DJ (2012) Variation in diet of semipalmated sandpipers (Calidris pusilla) during stopover in the upper Bay of Fundy, Canada. Can J Zool 90:1181–1190. https://doi.org/10.1139/Z2012-086
Quinn JT, Hamilton DJ, Hebert CE (2017) Fatty acid composition and concentration of alternative food of semipalmated sandpipers (Calidris pusilla) in the upper bay of Fundy, Canada. Can J Zool 95:565–573. https://doi.org/10.1139/cjz-2016-0246
Reich M, Hannig C, Al-Ahmad A, Bolek R, Kümmerer K (2012) A comprehensive method for determination of fatty acids in the initial oral biofilm (pellicle). J Lipid Res 53:2226–2230. https://doi.org/10.1194/jlr.D026260
Ren X, Zhao X, Turcotte F, Deschênes JS, Tremblay R, Jolicoeur M (2017) Current lipid extraction methods are significantly enhanced adding a water treatment step in Chlorella protothecoides. Microb Cell Fact 16:26. https://doi.org/10.1186/s12934-017-0633-9
Rosenberg KV, Dokter AM, Blancher PJ, Sauer JR, Smith AC, Smith PA, Stanton JC, Panjabi A, Helft L, Parr M, Marra PP (2019) Decline of the North American avifauna. Science (80-) 366:120–124. https://doi.org/10.1126/science.aaw1313
Rudy MD, Kainz MJ, Graeve M, Colombo SM, Arts MT (2016) Handling and storage procedures have variable effects on fatty acid content in fishes with different lipid quantities. PLoS ONE. https://doi.org/10.1371/journal.pone.0160497
Samburova V, Lemos MS, Hiibel S, Kent Hoekman S, Cushman JC, Zielinska B (2013) Analysis of triacylglycerols and free fatty acids in algae using ultra-performance liquid chromatography mass spectrometry. J Am Oil Chem Soc 90:53–64. https://doi.org/10.1007/s11746-012-2138-3
Sasaki GC, Capuzzo JM (1984) Degradation of Artemia lipids under storage. Comp Biochem Physiol B 78:523–531
Schlotterbeck J, Kolb A, Lämmerhofer M (2018) Free fatty acid profiling in marine algae extract by LC–MS/MS and isolation as well as quantification of the ω-3 fatty acid hexadeca-4,7,10,13-tetraenoic acid. J Sep Sci 41:4286–4295. https://doi.org/10.1002/jssc.201800780
Schnurr PJ, Drever MC, Kling HJ, Elner RW, Arts MT (2019) Seasonal changes in fatty acid composition of estuarine intertidal biofilm: implications for western sandpiper migration. Estuar Coast Shelf Sci 224:94–107. https://doi.org/10.1016/j.ecss.2019.04.047
Schnurr PJ, Drever MC, Elner RW, Harper J, Arts MT (2020) Peak abundance of fatty acids from intertidal biofilm in relation to the breeding migration of shorebirds. Front Mar Sci. https://doi.org/10.3389/fmars.2020.00063
Scholz B, Liebezeit G (2013) Biochemical characterisation and fatty acid profiles of 25 benthic marine diatoms isolated from the Solthörn tidal flat (southern North Sea). J Appl Phycol 25:453–465. https://doi.org/10.1007/s10811-012-9879-0
Shang D, Kim M, Haberl M, Legzdins A (2013) Development of a rapid liquid chromatography tandem mass spectrometry method for screening of trace naphthenic acids in aqueous environments. J Chromatogr A 1278:98–107. https://doi.org/10.1016/j.chroma.2012.12.078
Sharma KK, Schuhmann H, Schenk PM (2012) High lipid induction in microalgae for biodiesel production. Energies 5:1532–1553. https://doi.org/10.3390/en5051532
Snyder LR, Kirkland JJ (1979) Introduction to modern liquid chromatography, 2nd edn. Wiley, Hoboken
Stevens L (1996) Avian biochemistry and molecular biology. Cambridge University Press
Stonik VS, Stonik I (2015) Low-molecular-weight metabolites from diatoms: structures, biological roles and biosynthesis. Mar Drugs 13:3672–3709. https://doi.org/10.3390/md13063672
Studds CE, Kendall BE, Murray NJ, Wilson HB, Rogers DI, Clemens RS, Gosbell K, Hassell CJ, Jessop R, Melville DS, Milton DA, Minton CDT, Possingham HP, Riegen AC, Straw P, Woehler EJ, Fuller RA (2017) Rapid population decline in migratory shorebirds relying on Yellow Sea tidal mudflats as stopover sites. Nat Commun 8:1–7. https://doi.org/10.1038/ncomms14895
Tang B, Row KH (2013) Development of gas chromatography analysis of fatty acids in marine organisms. J Chromatogr Sci 51:599–607. https://doi.org/10.1093/chromsci/bmt005
Topolewska A, Czarnowska K, Haliński ŁP, Stepnowski P (2015) Evaluation of four derivatization methods for the analysis of fatty acids from green leafy vegetables by gas chromatography. J Chromatogr B Anal Technol Biomed Life Sci 990:150–157. https://doi.org/10.1016/j.jchromb.2015.03.020
Ulmer CZ, Jones CM, Yost RA, Garrett TJ, Bowden JA (2018) Optimization of Folch, Bligh–Dyer, and Matyash sample-to-extraction solvent ratios for human plasma-based lipidomics studies. Anal Chim Acta 1037:351–357. https://doi.org/10.1016/j.aca.2018.08.004
Underwood GJC, Kromkamp J (1999) Primary production by phytoplankton and microphytobenthos in Estuaries. Adv Ecol Res 29:93–153. https://doi.org/10.1016/S0065-2504(08)60192-0
Van Der Heijden LH, Graeve M, Asmus R, Rzeznik-Orignac J, Niquil N, Bernier Q, Guillou G, Asmus H, Lebreton B (2019) Trophic importance of microphytobenthos and bacteria to meiofauna in soft-bottom intertidal habitats: a combined trophic marker approach. Mar Environ Res 149:50–66. https://doi.org/10.1016/j.marenvres.2019.05.014
Venkata Mohan S, Devi MP (2014) Salinity stress induced lipid synthesis to harness biodiesel during dual mode cultivation of mixotrophic microalgae. Bioresour Technol 165:288–294. https://doi.org/10.1016/j.biortech.2014.02.103
Woo KL, Kim JI (1999) New hydrolysis method for extremely small amount of lipids and capillary gas chromatographic analysis as N(O)-tert-butyldimethylsilyl fatty acid derivatives compared with methyl ester derivatives. J Chromatogr A 862:199–208. https://doi.org/10.1016/S0021-9673(99)00934-6
Work TS, Work E (1972) Lipid extraction procedures. Laboratory techniques in biochemistry and molecular biology, vol 3. Elsevier, Amsterdam, pp 347–353. https://doi.org/10.1016/S0075-7535(08)70549-7
The authors acknowledge the support and input of their colleagues, notably Liane Chow, Honoria Kwok, and Jeffrey Yan of the Pacific Environmental Science Centre, Environment and Climate Change Canada. Thanks is extended to the Science and Technology Branch of Environment and Climate Change Canada for advice. Appreciation is directed to the UBC Science Co-op program for facilitating student work opportunities.
This work was supported by the Science and Technology Branch of Environment and Climate Change Canada, Government of Canada.
Ethics approval and consent to participate
Consent for publication
The authors declare that they have no competing interests.
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
About this article
Cite this article
Ollinik, J.E., Chua, C.C., Brunswick, P. et al. Assessing diatom-mediated fatty acids in intertidal biofilm: a new conservation concern. Environ Syst Res 10, 30 (2021). https://doi.org/10.1186/s40068-021-00236-2
- Fatty acids
- Climate change
- Migratory shorebirds
- Gas chromatography
- Liquid chromatography