Introduction

Lipids play an important role in living organisms being essential in aquatic life forms. They are a group of highly energetic molecules used as energy reserves, with regulatory functions in cell membrane physiology, precursors to many animal hormones responding to changes in the environment (Guschina and Harwood 2009). Furthermore, they are fundamental in many cellular and metabolic functions and specific alterations of the lipidome are implicated in the onset and progression of many diseases (Siegel et al. 2019). Lipids, a group of chemically heterogeneous molecules, are fatty acids (FAs) and their derivatives, and substances related biosynthetically or functionally to these compounds (de Carvalho and Caramujo 2018). Their diversity ranges from fatty acids, oxylipins and their ester or amide derivatives, to hydrocarbons, carotenoids, terpenes, sterols and bile acids. Lipids are divided in two main categories: the nonpolar (acylglycerols, sterols, free fatty acids, hydrocarbons, alcohols, wax, and steryl esters) and the polar lipids (phosphoglycerides, glycosylglycerides, and sphingolipids) (Guschina and Harwood 2013).

The neutral lipids, or nonpolar lipids, are not the main constituents of membranes and are essentially stored intracellularly in specialized compartments (lipid particles, lipid droplets or oil bodies). They are a source of sterols, free fatty acids and diacylglycerols as well as energy source. Actually, fatty acids stored in neutral lipids (neutral lipids fatty acids—NLFAs) can either be used for energy production or serve as substrates for phospholipid synthesis (Athenstaedt and Daum 2006).

In general, lipids have in common a ready solubility in organic solvents such as hydrocarbons, chloroform and alcohols but some can also be soluble in water (Guschina and Harwood 2013). For isolation or extraction purposes, several organic solvents or solvent mixtures must be adequately chosen depending on the fraction to be extracted. Polar solvents are used to remove lipids from their association with cell membranes and tissue constituents; and less polar solvents are used to dissolve all triacylglycerols and other nonpolar lipids like free fatty acids (Iverson et al. 2001).

In the last decades, an increasing interest has been placed in a particular lipid fraction, the fatty acid content. FAs are important nonpolar lipid molecules at functional and structural levels found in all organisms (Colombo et al. 2017). They can be constituents of phospholipids, the “building blocks” of cell membranes, storage as part of neutral lipids and play a role in cell signaling (de Carvalho and Caramujo 2018). Some polyunsaturated FAs (PUFAs), such as omega-3 (ω-3) and omega-6 (ω-6) families, are considered essential FAs (EFAs). Few animals have the ability to synthesize EFAs, so they have to acquire these important molecules or the correct precursors through their diet (Parrish 1999; Kainz et al. 2004; Taipale et al. 2013; Twining et al. 2016).

Several methods are available for lipid extraction from biological sources/matrices (Li et al. 2014). Most of them are devoted to enable total lipid extraction, but others only allow the extraction of some fractions like neutral lipids (Anthony and Stuart 2015). The selection of mechanical and/or chemical methods is fundamental to range from total to fraction extraction. The extraction of the total lipid leads to profiles which reflect mainly the major fractions composition, like membrane lipids. Under this circumstance, minor fractions like neutral lipids, are underestimated due to their lower amount in the biological samples. On the other hand, extraction of independent fractions allows to better understand and study minor lipid components and their variation under different situations (e.g., organisms, environments, growth phases).

In aquatic food webs, lipids are a great form of energy (Parrish 2009), and thus can be used to predict the transfer of energy from one trophic level to another (Napolitano 1999). A good biomarker for trophic interactions is achieved with the FAs and sterols transfer up the food chain via classical predator–prey interactions (Ruess and Müller-Navarra 2019). Indeed, ω3PUFAs have already been used as markers for trophic interactions in freshwater plankton (Desvilettes et al. 1997). Among others, essential biochemical resources such as FAs and sterols can directly influence consumers’ growth and reproduction, and consequently the functioning of the ecosystem. These molecules are, generally, incorporated into consumer body tissue without major modification, a fact called “dietary routing” (Stott et al. 1997). It is considered that all PUFAs in food webs thus originate mainly from primary producers (de Carvalho and Caramujo 2018). Well know is the role played by microalgae providing the bulk of de novo PUFA production for all aquatic food webs. Indeed, they are commonly used in the rearing of zooplankton organism in laboratory conditions. Recently, several studies on aquatic food webs pointed to the importance of prokaryotes, and in particular bacteria, as a primary food source for omnivorous, sestonivorous and filtering benthic animals and also in lipid transfer (Nichols 2003; de Carvalho and Caramujo 2012). In fact, the role of prokaryotes is described for nutrient cycling and the degradation of organic matter, and a full function in the microbial loop is not yet completely understood.

Zooplankton, a group of non-specific filtering organisms, has an intermediate position between the lower and higher trophic levels and thereby it serves as driver of lipids from primary producers to predators (Arts 1999). A well-studied and standard organism in aquatic ecotoxicology is the microcrustacean D. magna which has its diet conditioned by a single food source (a microalga) restraining it from nutritional diversity [e.g., carbon and energy (lipids) sources] (Bychek et al. 2005). In the last years, our research group has studied the potential use of bacteria as the only food source or as a nutritional supplement to the traditional single algal source (Antunes et al. 2016; Lage and Antunes 2018; Marinho et al. 2018, 2019). The focus has been put in a particular phylum of bacteria, the Planctomycetes and in particular in the species R. rubra. This bacterium proved to be, especially in addition to R. subcapitata, a good nutritional complement to D. magna food regimen by increasing D. magna size and fertility with a significant increase of offsprings (Marinho et al. 2019).

The present study aims to compare the total fatty acids (TFAs) reported in the literature (here reported as conventional method—CM), with neutral lipid fatty acid (NLFA) profiles assessed by the here reported method (RM), in the bacterium R. rubra and the alga Raphidocelis subcapitata, which are both food sources for D. magna. Furthermore, D. magna NLFAs were assessed when this crustacean was fed with the bacterium and the alga, individually or in combination. These analyses allowed a deeper insight into the NLFA profiles under different D. magna feeding regimens.

Materials and methods

Cultures of organisms

R. rubra was cultured in modified medium M13 (Lage and Bondoso 2011). This bacterium was grown on solid medium at 26 °C and then transferred into a small volume of liquid medium and cultivated under continuous stirring at 200 rpm. The culture was then upscaled to a volume of 600 mL. After 3 days of growth, corresponding to exponential phase, cells were collected by centrifugation at 4000 rpm for 10 min. Cell pellets were resuspended in distilled water to achieve an optical density adjusted to 0.2 absorbance units (AU) at λ = 600 nm. Aliquots with 50 mL of cell suspension were stored at − 20 °C for later use in the D. magna feeding assays (see below Feeding assays). Due to the formation of cell clusters, before each use, the bacterial cell suspension was sonicated for 30 s with a Misonix Microson Ultrasonic Cell Disruptor XL set to intensity of 10 W. After this procedure, the bacterial cell suspension was again adjusted to 0.2 AU at λ = 600 nm before being provided as food for D. magna in the feeding assays.

The microalga Raphidocelis subcapitata Korshikov (Hindak) (formerly known as Pseudokirchneriella subcapitata) was kept in cultures with Woods Hole MBL medium (Stein 1973), under controlled conditions of temperature (20 ± 2 °C) and continuous light (~ 6000 lx), were harvested in the exponential growth phase (5–7 days old) and inoculated in fresh medium (Environment Canada 1992; OECD 2006). Algal cell concentration was calculated based on the correlation of absorbance measured at 440 nm and cell density, previously determined.

Monoclonal cultures of D. magna (clone A, sensu Baird et al. 1989a) were kept over several generations of pure parthenogenetic cultures under controlled conditions of photoperiod (16hL:8hD) and temperature (20 ± 2 °C). Cultures were maintained in ASTM synthetic hard water medium according to standard procedures (ASTM 1980) and supplemented with standard organic additive (suspension extracted from brown algae Ascophyllum nodosum) (Baird et al. 1989b) to provide essential microelements to Daphnia. The culture medium was renewed three times a week, and D. magna was fed with the microalga R. subcapitata at a ratio of 3.0 × 105 cells mL−1 day−1. Neonates born between the third and the fifth brood with less than 24 h, were used to initiate assays or to renew the cultures.

Feeding assays

Feeding assays were conducted for 21 days (adapted by OECD 2012 guidelines) under the same conditions as described for rearing cultures. In the assay, D. magna was fed with three different diets: (1) R. subcapitata as the only food source added, in a ratio of 3.0 × 105 cells mL−1 day−1; (2) R. rubra as the only food source added, − 2500 µL of cell suspension (0.2 AU at λ = 600 nm), and (3) the same cell density of bacterium described above plus a standard food source (R. subcapitata). Ten individual D. magna replicates were kept in glass cups filled with 50 mL of ASTM medium. The organisms were fed with alga or/and bacteria every day and the medium was renewed every 2 days.

At the end of the assay (21 days), fatty acid profile characterization was conducted in the organisms of each feeding replicate. A pooled sample with three organisms was collected and stored at − 20 °C until fatty acid content analysis was performed.

Neutral lipid fatty acid (NLFA) extraction and profile characterization

NLFAs were analyzed following ISO 12966-2 (2017)—Rapid Method, with slight modifications, which consists of an initial procedure of fat extraction followed by Fatty Acid Methyl Ester (FAME) formation by a trans-esterification reaction. Initially, a mechanical extraction (cell disruption) procedure was performed. For D. magna, manual maceration of organisms (pooled sample in 1.5 mL microtube) was performed with a Teflon pestle in 0.3 mL of distilled water and 100 mg of quartz sand. After obtaining a homogeneous paste, 0.4 mL of distilled water and 0.6 mL of cyclohexane were added. For R. rubra and R. subcapitata, a High Pressure Homogenizer (HPH) (Emulsiflex C3; Avestin®) was used for the same purpose. Concentrated alga and bacteria cultures were centrifuged, and pellets were resuspended in distilled water. These cell suspensions were subjected to several cycles at 1500 bar until cell disruption was achieved (accessed by optical microscopy). To an aliquot of 0.7 mL, for all samples (D. magna, R. rubra and R. subcapitata), 0.6 mL of cyclohexane was added and mixed in a 1.5 mL microtube. All the suspensions (obtained either from maceration or HPH) were mixed in a vortex for 30 s and centrifuged at 8000 rpm for 30 s; the supernatant was then transferred to a clean microtube. A second extraction step was performed by adding 0.6 mL of cyclohexane to the bottom liquid phase, mixing and centrifuging as described above; the resulting supernatant was added to the one initially collected. Then, 70 µL of trans-esterification reagent (2 M KOH in methanol) was added to the extract, followed by repeated (5×) mixing in the vortex for 30 s, along a 15 min resting period. After this period, samples were centrifuged at 8000 rpm for 45 s and the supernatant was collected and transferred to a gas chromatograph vials and stored at − 4 °C until further analysis.

FAMEs were analyzed in gas chromatograph (GC; YL 6500) equipped with a Split/Splitless injector, a flame ionization detector and a capillary column (DN-Wax, 30 m × 0.25 mm id × 0.25 µm film thickness; Dani Instruments®). The injector was kept at 240 °C, with a split ratio of 1:20 using He as carrier gas (1 mL min−1). The oven started at 50 °C for 2 min, ramped up until 150 at 4 °C min−1 and then up to 230 °C at 3 °C min−1 and remained at final temperature for 5 min. Total analysis time was 60 min. All samples were analyzed in triplicate and the respective standard deviations were calculated. For identification purposes, a standard Supelco® 37 Component FAME Mix was analyzed under the same conditions, and an identification calibration was built based on each compound retention time and equivalent chain length (ECL). ECLs were calculated for this chromatographic system according to Bannon et al. (1988). The same standard was also analyzed before each set of samples as a system suitability test. Identified peaks were integrated for area and the % area was considered to characterize the FAME profile (RM—results of the here-reported method). Since not all peaks could be clearly identified since the standard had only 37 compounds, the most relevant and not identified peaks were assigned to the respective ECL.

For the establishment of the total fatty acid (TFA) profile (CM—results from literature) of R. subcapitata, values considered were reported by Nascimento et al. (2013); and for R. rubra values considered were reported by Bondoso et al. (2014).

Statistical analysis

Statistical analysis was carried out with two-tailed Student’s t test for fatty acid profiles comparison between the different D. magna diet regimes (R. subcapitata, R. rubra, and R. subcapitata + R. rubra), and the fatty acid profiles of the respective food sources provided.

Only p values < 0.05 were considered significant. Error bars in graphs represent standard deviation of the mean for three independent measurements.

Results

Table 1 summarizes the 30 identified chromatographic peaks obtained by the RM (here-reported method) and the respective identification according to standard Supelco® 37 and ECL calculation. Values in bold were described in the literature but not detected in our samples and/or method. Due to their relevance (high value reported), they were empirically placed in the table based on the carbon chain length. For normalization purposes, compound name will be primarily used, or when absent, ECL nomenclature (see Table 1 for equivalent designations).

Table 1 Measured chromatographic peaks. Name and formula for the FAMES included in the standard Supelco® 37
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Regarding Fig. 1, both microorganisms present, overall, a similar fatty acid profile as assessed by the RM, except for oleic/elaidic acid (ECL 18.219) and ECL 19.833 which amounts are much higher in R. subcapitata than in R. rubra. However, fatty acid profile as assessed by the CM (literature values) is different for both microorganisms. The main fatty acid present in R. subcapitata is palmitic acid (ECL16.000), while in R. rubra is oleic/elaidic acid (ECL18.219). Moreover, small relative amounts (< 5% of relative amount) of lauric acid (ECL12.000) were only quantified in R. subcapitata, while ECL 18.558, C17:1n8c and pentadecanoic acid (ECL 15:000) were quantified only in R. rubra.

Fig. 1
figure 1

Comparison between the total fatty acid (TFA) profile reported in the literature (♦, CM) and the herein reported neutral fatty acids (bars, RM), for R. subcapitata and R. rubra. Error bars indicate standard deviations for the herein reported neutral fatty acids

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Concerning method comparison, for R. subcapitata and R. rubra, high values of ECLs 19.550, 19.673 and 19.833 were recorded by the RM method, while these compounds were not reported in the CM. On the other hand, some compounds were only detected by the CM method, namely the palmitoleic acid (ECL16.260) for both species and pentadecanoic acid (ECL 15.000), C17:1n8c and ECL 18.558 for R. rubra. For the two species analyzed, higher values (more than half) of palmitic acid (ECL16.000) were reported in the CM comparatively to RM. Another relevant difference observed between the two methods was recorded for oleic/elaidic acid (ECL 18.219) in R. rubra which the value is the highest in the CM (≈ 44% in CM against < 5% in RM).

Figure 2 shows the results of the fatty acid profiles determined in the D. magna after being fed with different diet regimes (R. subcapitata, R. rubra, and R. subcapitata + R. rubra), and the fatty acid profiles of the respective food sources provided. A similar pattern of D. magna fatty acid profiles was observed for the two diet regimens where R. subcapitata was provided (Fig. 2a, c). On the other hand, such similarity was not observed when D. magna is fed with R. rubra (Fig. 2b, d). Overall, the oleic/elaidic acid (ECL 18.219) and palmitic acid (ECL 16.000) were the most abundant fatty acids quantified in D. magna, independently of the diet provided (where lower values were detected in both species). Although R. rubra presents low levels of oleic/elaidic acid (ECL 18.219) in the RM fraction, it is quite high in the CM fraction as reported by the literature (see Fig. 1) what may justify its levels in D. magna fed with this bacterium. The pentadecanoic acid (ECL 15.000) is present in D. magna profiles but not in R. rubra or R. subcapitata. ECL16.192 which is present in relative high amount in D. magna (≈ 20%) is practically absent in R. rubra and R. subcapitata. These results evidence the relevance of these fatty acids to D. magna. For palmitoleic acid (ECL16.260), low levels were observed in D. magna fed with R. subcapitata and with the mix diet (< 2%). Indeed, this fatty acid was only present in high amount in the RM fraction corresponding to D. magna fed only with bacterium (≈ 12%) and in the CM for bacterium (> 15%). This high amount disappears with the introduction of the alga in the diet. Stearic acid (ECL18.000) although present in high amount in the bacteria and the alga (≈ 10%), is comparatively lower (< 5%) in D. magna. ECLs 19.550 and 19.673 were quantified in high levels (> 20%) in the RM in both R. rubra and R. subcapitata, but not in D. magna (< 10%). When Daphnia was fed with only R. rubra, the levels of ECLs 19.550 and 19.673 were higher than when R. subcapitata was included in the diet. These results suggest that D. magna is nutritionally selective regarding these fatty acids. This selectivity is evidenced in Fig. 3.

Fig. 2
figure 2

Results of the here-reported method (RM) for neutral lipid fatty acids—NLFA profile: a values of D. magna fed with R. subcapitata (grey bars) and R. subcapitata (◦); b values of D. magna fed with R. rubra (grey bars) and R. rubra (◦); c values of D. magna fed with both food source (grey bars) with R. subcapitata (◦) and d values of D. magna fed with both food source (grey bars) with R. rubra (◦). *p < 0.05; ←FAME values only found in one of the situations

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Fig. 3
figure 3

Results of the here-reported method (RM) for neutral lipid fatty acids—NLFA profile: a values of D. magna fed with R. subcapitata (white bars) and values of D. magna fed with R. subcapitata and R. rubra (grey bars); b values of D. magna fed with R. rubra (black bars) and values of D. magna fed with R. subcapitata and R. rubra (grey bars). *p < 0.05; ↓FAMEs values only found in one of the situations

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Discussion

In organisms, the total lipid content is an important parameter for the assessment of biochemical, physiological, and nutritional characterization (Iverson et al. 2001). However, also the partial fractions may be relevant to highlight differences in different biological processes. Depending on the extraction method, different fractions may be available for characterization, namely the neutral fraction which includes the neutral lipid fatty acids. Our results clearly show that NLFA fraction differs from total fraction (here reported by CM). The abundance of acids palmitic (ECL 16.000) and oleic/elaidic (ECL18.219) in the R. subcapitata and R. rubra total fraction indicates a major contribution from membrane phospholipids, which are polar compounds (Guschina and Harwood 2013). On the other hand, when analyzing the neutral fraction (RM), the most abundant FAs are ECLs 19.550 and 19.673, indicating that the membrane phospholipids play a shadow effect on the FA composition. This must be due to the relative negligible amount of these two FAs in the analysis of the total FAs as reported by CM, which are only evident when only neutral lipids are quantified. However, for oleic/elaidic acid (ECL18.219), both fractions (NLFAs—RM and total fatty acids—CM) present the same relative amount in R. subcapitata but not in R. rubra. This different profile shows that different types of organisms (in this case bacterium vs alga) present different fatty acid compositions in different cellular structures, revealing different biosynthetic pathways. Actually, FAs stored in neutral lipids can have different functions such as energy production, substrate for phospholipid synthesis, second messenger in signal transduction and hormone precursors (Athenstaedt and Daum 2006) while polar lipids play fundamental roles in membrane structure (Siegel et al. 2019).

NLFA profiles for R. rubra and R. subcapitata are quite similar as evidenced in our results. However, regarding total FA profile, literature evidences differences between these two organisms. When D. magna is fed with one of these organisms independently (Fig. 2a, b), its NLFA profile is different. Differences found in D. magna NLFAs between feeding regimens can be explained by the differences in total FAs and the small differences in NLFAs of the two feeding sources. This is clear in the case of ECL 19.673 and ECL 19.550, which are in higher relative amount in the R. rubra and R. subcapitata neutral fractions (Fig. 1), but different in the neutral fraction of D. magna fed with each one of them separately (Fig. 2a, b). One exception is the content of palmitic acid (ECL 16.000) which is the core molecule in the fatty acid metabolism (Kanehisa et al. 2010), since its amount is always around 20% of the NLFA fraction in D. magna. Although neither bacterium nor alga have such a high amount in their NLFA fraction, D. magna always manage to accumulate the needed amount which may be supplied by the total FA, mainly the membrane phospholipids, or de novo synthesized (Figs. 1 and 2).

When D. magna is fed with both R. rubra and R. subcapitata, its NLFA profile resembles the one presented when the sole food source is the alga (Fig. 3). This result evidences that the alga should be the preferred food source for D. magna. However, even though R. rubra is not the preferred food source, it allows a standard performance of D. magna in terms of viability and reproductive output. In fact, this study (Marinho et al. 2019) showed that life history parameters (reproductive output and the somatic growth) and the size of offspring of D. magna improved when its nutrition was supplemented with the planctomycete R. rubra.

The here obtained results show that D. magna has the ability to use and change the FAs provided in the diet according to its needs. As daphnids have a complex enzymatic machinery with several fatty acid metabolism pathways, they can use fatty acids as they are provided, or degrade them to acetylCoA and proceed with a de novo synthesis. In fact, robust lipidomic remodeling to preserve membrane physical properties occurred when membrane homeostasis in mammalian cells was challenged by dietary lipids (Levental et al. 2018). Furthermore, D. magna can also synthesize de novo from other organic sources like carbohydrates. Actually, daphnids adapt to available resources in their environments by adjusting their biochemistry and physiology. This capacity associated with D. magna filtering feeding capacity allows D. magna to survive and reproduce in diverse conditions of food availability of food.