Exploring Pavlova pinguis chemical diversity: a potentially novel source of high value compounds.

To uncover the potential of Pavlova pinguis J.C. Green as a natural source of value added compounds, its lipophilic extracts were studied before and after alkaline hydrolysis using gas chromatography-mass spectrometry (GC-MS). The GC-MS analysis of the lipophilic extracts showed a wide chemical diversity including 72 compounds distributed by fatty acids (29), sterols (14), fatty alcohols (13) and other lipophilic compounds (16). Fatty acids represented the main class of identified compounds presenting myristic, palmitic, palmitoleic and eicosapentaenoic acids as its main components. Through the ∑ω6/∑ω3 ratio (0.25) and sterol composition it was possible to observe that P. pinguis is a valuable source of ω3 fatty acids and stigmasterol (up to 43% of total sterols). After alkaline hydrolysis, fatty acids and fatty alcohols content increased by 32 and 14% respectively, in contrast to, monoglycerides which decreased by 84%. The long chain alcohols content enables the exploitation of this microalga as a source of these bioactive compounds. Smaller amounts of sugars and other compounds were also detected. The present study is a valuable reference to the metabolite characterization of P. pinguis and shows the potential of this microalga for nutraceutical and pharmaceutical industries.

Introduction

The search for natural products with pharmaceutical and industrial applications has driven the attention of the scientific community towards the marine environment[1,2]. The large spectrum of marine organisms combined with their intrinsic chemical variability make these organisms a huge source from which to isolate new molecules with a broad range of applications[3]. From the marine organisms, microalgae have emerged as versatile cell factories to produce high-value compounds due to their rich biodiversity, growth rate, phenotypic plasticity and simple nutrient requirements[1].

The richness of microalgal biodiversity is often underestimated in the biotechnological field, being restricted to few species of Chlorophytes and Cyanophytes that dominate the market[4]. This fact represents a constraint for the full development of microalgae based industries once it overshadows the diversity of compounds amongst microalgae taxa[5]. Therefore, to exploit the potential of microalgae as versatile cell factories the following challenges are found: microalgal strain selection, cultivation optimization and downstream biomass extraction[4,6]. These can be overcome through a detailed phytochemical characterization and identification of the high-value components of microalgal extracts[6].

Microalgal cell components have been recognized as precious sources of health promoting phytochemicals that can prevent and/or improve cardiovascular diseases, hyper-tension, and arthritis and act as anti-inflammatory, anticarcinogenic and antitumoral agents[7,8]. Included in the health beneficial phytochemicals synthesized by microalgae are terpenes, sterols, phenolics, polyunsaturated fatty acids (PUFA), vitamins, carbohydrates, proteins among other compounds[9,10].

Although much research has focused on the aquaculture potential of several Pavlova species[11-13], only specific algal compounds (e.g. fatty acids and sterols) have been analyzed to determine their biological activity, nutritional value and applicability[14]. This target analysis restricts the detection of compounds that are present in low quantities which, in turn, makes difficult the inclusion of unknowns in microalgal extract analysis[15]. From the classes of widespread natural products, the composition of long-chain aliphatic alcohols (LC-alcohols), steryl glycosides and monoglycerides in microalgae are poorly studied[9].

In the Haptophyta Pavlova pinguis J. C. Green[16] only specific classes of compounds have been analyzed to assess its potential as food for larval hatcheries in aquaculture and as ecological biomarkers[13,17,18]. For instance, Milke, et al.[11] and Parrish, et al.[12] assessed the ability of P. pinguis and other Pavlova species to sustain postlarval sea scallop growth focusing on its proximate, fatty acid and sterol composition. In this microalga the complete characterization of lipid components (simple and complex lipids) is still largely unexplored[6,9]. Thus, in the present study the analysis of the lipophilic fraction of P. pinguis was performed in order to identify its lipophilic features before and after alkaline hydrolysis through gas chromatography–mass spectrometry (GC–MS) and evaluate its prospects for further improvement in bioactive compounds.

Materials and Methods

Growth and culture conditions

The haptophyta Pavlova pinguis (RCC 1539) was obtained from the Roscoff Culture Collection (RCC). The microalgal cultures were made by inoculating starter cultures into 1L of sterile f/2 – Si medium with pH adjusted to 7.0 under 70 μmol m−2 s−1 light intensity with 16:8 h (light: dark cycles) at 25 °C. At the end of the logarithmic phase, the medium was centrifuged for 7 min. at 3720 g and the pellets washed. Microalgae growth was monitored daily with a Neubauer–improved counting chamber (Marienfield–Superior) and a light microscope (Olympus BX41) with a 40x magnification. The specific growth rate was determined as described in Fernandes, et al.[19].

Solvent extraction

The extraction of non-polar phases was made as described by Ma et al.[20], with some modifications. To 0.10 g of microalgal freeze dried biomass an aqueous solution (methanol:water in a 1:1 ratio) and chloroform in 1:1 ratio were added. After homogenization, the mixture was left stirring for 15 min. and centrifuged at 4430 g for 10 min. The organic layer was carefully removed and transferred into pre-weighted tubes. The insoluble residue was washed three times with chloroform and dried in Na2SO4 filters. The extracts were evaporated in a nitrogen atmosphere and the amount of extractable substances was gravimetrically quantified and expressed as a percentage by weight of the freeze dried biomass (dry weight, dw). The extractable substances are presented as an average of at least three replicates.

Fourier transform infrared (FTIR) spectroscopy

FTIR with attenuated total reflectance (ATR) was used to identify the major functional groups in the raw microalga and chloroform extracts. FTIR-ATR spectra were collected on a Perkin–Elmer Spectrum Two instrument coupled with a Diamond ATR accessory (DurasamplIR II, Smiths Detection, UK) scanning over the wavenumber range of 4000–650 cm−1 at a resolution of 4 cm−1 and 36 scans.

Alkaline hydrolysis

The alkaline hydrolysis was performed in two aliquots of the chloroform extracts to detect molecules in their esterified forms according to Santos, et al.[9]. To each extract, 10 mL of 0.5 M NaOH in aqueous methanol was added and the mixtures were heated at 100 °C for 1 h in a nitrogen atmosphere. Then, the samples were allowed to cool, prior to the acidification of the mixtures to pH 2 with 1 M HCl. Following this step the hydrolyzed samples were extracted with dichloromethane. The solvent was evaporated to dryness under nitrogen.

Gas chromatography–mass spectrometry (GC-MS) analysis

Prior to GC–MS analysis, the extracts were silylated accordingly to Santos, et al.[9]. Two aliquots of each dried extract (before and after alkaline hydrolysis) and an accurate amount of internal standard (tetracosane, 0.30 and 0.40 mg) were dissolved in 250 μL of pyridine, 250 μL of N,O-bis(trimethylsilyl)trifluoroacetamide and 50 μL of trimethylclorosilane. Then, the mixture was kept at 70 °C for 30 min to proceed to the conversion of the hydroxyl and/or carboxyl groups into trimethylsilyl (TMS) ethers and/or esters, respectively. TMS were analyzed in a gas chromatographer (Agilent HP 6890) equipped with a mass selective detector (Agilent 5973) and a ValcoBon 17704 capillary column VB1 (30 m × 0.25 mm inner diameter, 0.25 µm film thickness). The chromatographic conditions were as follows: oven initial temperature was 80 °C for 5 min.; increasing 4 °C min−1 until reach the 208 °C; followed by 2 °C min−1 to 260 °C; and 5 °C min−1 until reaching the final temperature of 300 °C for 4 min. The temperature of the injector was 250 °C; the transfer line, 290 °C; and the split ratio was 33:1. Helium was used as the carrier gas at a constant flow of 1.0 mL min−1. The identification of the extracted compounds as TMS derivatives was made by comparison of the mass spectra fragmentation to those in the GC-MS spectral library (Wiley-NIST Mass Spectral Library 1999), literature data[21-24] or by injection of standards. For semi-quantitative analysis, GC–MS was calibrated with pure reference compounds (mannose, trans-ferulic acid, nonadecan-1-ol, eicosan-1-ol, 5α–Chlolestane, cholesterol, stigmasterol, hexadecanoic, and nonadecanoic acids) relative to tetracosane.

Antioxidant activity

The 2,2-diphenyl-1-picrylhydrazyl (DPPH) radical scavenging activity was determined according to Maadane, et al.[25] with some modifications. Stock solutions of butylated hydroxytoluene (BHT, 1 mg mL−1) and extract (3 mg mL−1) were prepared in methanol and dimethyl sulfoxide, respectively. The stock solutions were added to 1300 µL of DPPH radical solution (83 µM). Then the absorbance of samples was measured at 520 nm with a UV/Vis spectrometer Lambda 25 (Perkin Elmer), after 30 min in the dark at room temperature. The DPPH scavenging effect was calculated by the Equation (1):where Asample is the absorbance of DPPH solution with the sample or standard, Asample blank is the absorbance of sample without DPPH and Acontrol is the absorbance of DPPH solution without sample.

Statistical analysis

Statistical analysis of the data was carried out using the software IBM SPSS Statistics 24. Differences between treatments were assessed with Student’s t-test, p-values < 0.05 were considered to be statistically significant.

Results and Discussion

Growth and extraction yield

The overall yield of a specific desired product is dependent on the microalgal growth rate and on the product content[26]. Thus, when exploring the potential of a microalgal strain for further improvement in high valued compounds, the microalgal growth should be considered. The growth rate determined for P. pinguis was 0.8 day−1 and the maximum cell concentration, reached by this microalga, was 8.46 × 106 cells mL−1, Fig. 1. According to Steinrucken, et al.[27] this microalgal strain can be considered as a high growth rate strain since its growth rate is ≥0.7 day−1. The average dry biomass production observed at the end of the batch cultivation was 368 mg L−1 achieved in 7 days, this value being close to the dry weight (dw) estimated by Mansour, et al.[18] of 390 mg L−1.

Figure 1

Growth curve of P. pinguis in f/2 growth medium.

The yields of the chloroform extractable substances, in P. pinguis, accounted for 11.92% dw. This yield was higher than that previously reported by Mansour, et al.[18] for this microalga cultivated in the fE and GSe growth media and extracted with a modified Bligh and Dyer, in which, the chloroform extracts accounted 7.6% and 3.5% dw, respectively. The total chloroform extractable substances obtained in this study (5.17 pg cell−1) were also higher than that found for other P. pinguis strains (2 pg cell−1) grown in f/2 medium and extracted with chloroform through modified Bligh and Dyer[22].

FTIR analysis

The FTIR-ATR was employed to P. pinguis biomass and chloroform extracts as a first approach to perform a qualitative analysis of the extractable substances of P. pinguis (Fig. 2). Through the FTIR spectrum of the raw microalga, it is possible to visualize three main regions, that relate to the main macromolecular pools: (i) the carbohydrate region between 1200–900 cm−1 (νC–O–C of carbohydrates); (ii) the protein bands at 1655 cm−1 and 1545 cm−1 (νC-O of amide II and δN–H of amide I, respectively); (iii) and the lipid associated peaks at 1740 (νC=O of the ester functional groups) and 3050–2800 cm−1 [28]. This observation indicates the co-presence of lipids, proteins and polysaccharides in the microalgal biomass. In the FTIR spectrum of the chloroform extract it is possible to visualize that the signals often attributed to the characteristic functional groups of lipids increased their intensity, namely those in the 3050–2800 cm−1 region (C-H stretch), which indicates that lipids are the major component of the lipophilic extract. Moreover, the presence of peaks at around 720, 1745 and 3010 cm−1, that are related to the CH2 rocking, C=O and C-H stretch, respectively, indicates the presence of unsaturated hydrocarbons in the lipophilic extract[29].

Figure 2

FTIR–ATR spectra of the lipophilic extractives and the raw marine microalga P. pinguis, the characteristic bands of the lipophilic extracts are highlighted.

GC-MS analysis

To screen microalgae for commercial purposes, different aspects have to be covered: the first is to explore the chemical diversity of microalgae; the second is to evaluate the quality of these extracts by searching for known bioactive compounds[30].

The microalgal strain under study presented a wide chemical diversity in the extract: 29 fatty acids, 14 sterols, 13 fatty alcohols, and 16 other compounds. Through Table 1 it is possible to observe qualitative and quantitative differences, in the P. pinguis chloroform extracts, before and after hydrolysis. Before hydrolysis 71% of the extractable substances were quantified while after hydrolysis this percentage increased to 88%.

Table 1

Compounds detected in the lipophilic extracts of P. pinguis before (BH) and after (AH) alkaline hydrolysis.

N°	Identified Compounds	MW	Content (mg g−1 of microalgal dw)		Content (mg g−1 of extract)
			BH	AH	BH	AH
	Fatty acids		57.34 ± 1.33a	75.60 ± 4.49b	482.95 ± 26.06a	631.77 ± 23.20b
	Saturated		24.99 ± 0.67a	35.55 ± 2.60b	210.60 ± 12.07a	297.02 ± 14.12b
2	Octanoic acid	216	n.d.	1.03 ± 0.00	n.d.	8.65 ± 0.27
4	Nonanoic acid	230	0.77 ± 0.00a	1.05 ± 0.01b	6.49 ± 0.23a	8.78 ± 0.20b
6	Decanoic acid	244	n.d.	1.03 ± 0.01	n.d.	8.62 ± 0.34
13	Dodecanoic acid	272	0.78 ± 0.01a	1.05 ± 0.01b	6.54 ± 0.28a	8.79 ± 0.27b
17	Myristic acid	300	7.43 ± 0.41a	9.43 ± 0.54b	62.62 ± 5.37a	78.86 ± 4.03b
20	Pentadecanoic acid	314	1.06 ± 0.02a	1.38 ± 0.03b	8.91 ± 0.45a	11.53 ± 0.25b
28	Palmitic acid (PA)	328	7.32 ± 0.22a	9.97 ± 1.66b	61.71 ± 3.62a	83.13 ± 11.56b
18	Hexadecanoic methyl ester	270	0.85 ± 0.01a	1.05 ± 0.01b	7.14 ± 0.34a	8.79 ± 0.36b
30	Heptadecanoic acida	342	2.54 ± 0.01a	3.26 ± 0.02b	21.37 ± 0.58a	27.29 ± 0.71b
39	2-Octyl-Cyclopropaneheptanoic acid	354	0.82 ± 0.00a	1.09 ± 0.01b	6.89 ± 0.23a	9.10 ± 0.31b
38	Octadecanoic acid	356	1.81 ± 0.03a	3.07 ± 0.50b	15.24 ± 0.67a	25.61 ± 3.40b
46	Eicosanoic acid	384	0.79 ± 0.00a	1.07 ± 0.01b	6.69 ± 0.24a	8.93 ± 0.20b
53	Docosanoic acid	412	0.83 ± 0.02a	1.07 ± 0.01b	7.00 ± 0.24a	8.94 ± 0.23b
	Monounsaturated		15.20 ± 0.54a	17.83 ± 1.00b	128.12 ± 8.53a	149.07 ± 5.78b
24	Palmitoleic acida	326	10.97 ± 0.39a	13.06 ± 0.78b	92.46 ± 6.15a	109.19 ± 5.00b
25	7-Hexadecenoic acid	326	0.92 ± 0.02a	1.17 ± 0.01b	7.75 ± 0.37a	9.79 ± 0.31b
37	Vaccenic acid	354	1.27 ± 0.08a	1.72 ± 0.16b	10.68 ± 0.98a	14.33 ± 0.91b
36	Oleic acid	354	1.25 ± 0.08a	1.89 ± 0.08b	10.55 ± 0.93a	15.76 ± 0.28b
44	Gondoic acid	382	0.79 ± 0.00	n.d.	6.68 ± 0.22a	n.d.
	Polyunsaturated		16.34 ± 0.29a	21.09 ± 0.92b	137.38 ± 5.69a	176.31 ± 4.50b
22	4,7,10,13-Hexadecatetraenoic acid	322	0.82 ± 0.01a	1.08 ± 0.01b	6.91 ± 0.27a	9.01 ± 0.29b
26	Methyl-4,7,10,13-hexadecatetraenoate	262	0.85 ± 0.01a	1.05 ± 0.00b	6.86 ± 0.27a	8.80 ± 0.28b
34	Linoleic acid (LA)a	352	1.86 ± 0.03a	2.58 ± 0.03b	15.65 ± 0.71a	21.59 ± 0.81b
35	α-Linolenic acid (ALA)	350	0.89 ± 0.01a	1.18 ± 0.02b	7.49 ± 0.30a	9.90 ± 0.29b
33	Stearidonic acid (SA)	348	2.82 ± 0.09a	3.04 ± 0.41a	23.80 ± 1.47a	25.39 ± 2.67a
32	3,6,9,12,15-Octadecapentaenoic acid	346	0.90 ± 0.01a	1.18 ± 0.02b	7.60 ± 0.30a	9.84 ± 0.33b
42	Eicosapentaenoic acid (EPA)	374	3.49 ± 0.09a	4.73 ± 0.26b	29.43 ± 1.44a	39.54 ± 1.51b
48	7,10,13,16,19-Docosapentaenoic acid	402	1.59 ± 0.08a	2.05 ± 0.08b	13.36 ± 0.68a	17.14 ± 0.51b
49	4,7,10,13,16-Docosapentaenoic acid	402	1.11 ± 0.02a	1.53 ± 0.06b	9.37 ± 0.32a	12.80 ± 0.32b
47	Docosahexaenoic acid (DHA)	400	2.01 ± 0.10a	2.67 ± 0.15b	16.91 ± 0.76a	22.30 ± 0.95b
	Diacids		0.81 ± 0.01a	1.12 ± 0.03b	6.86 ± 0.27a	9.37 ± 0.33b
50	Octadecenedioic acid	456	0.81 ± 0.01a	1.12 ± 0.03b	6.86 ± 0.27a	9.37 ± 0.33b
	Fatty Alcohols		9.69 ± 0.26a	10.90 ± 0.39b	81.54 ± 0.32a	91.14 ± 0.90b
1	1-Octanol	202	n.d.	1.20 ± 0.08	n.d.	10.05 ± 0.35
7	1-Undecanol	244	0.63 ± 0.01a	0.83 ± 0.00b	5.31 ± 0.24a	6.93 ± 0.23b
12	1-Dodecanol	258	0.63 ± 0.00a	0.83 ± 0.01b	5.28 ± 0.18a	6.99 ± 0.25b
14	1-Tridecanol	272	0.64 ± 0.01a	0.83 ± 0.01b	5.36 ± 0.26a	6.98 ± 0.25b
16	1-Tetradecanol	286	0.67 ± 0.01a	0.88 ± 0.01b	5.61 ± 0.26a	7.36 ± 0.17b
21	1-Hexadecanol	314	1.50 ± 0.09a	1.21 ± 0.08b	12.63 ± 0.39a	10.07 ± 0.42b
31	1-Octadecanol	342	1.18 ± 0.06a	1.17 ± 0.07a	9.94 ± 0.16a	9.81 ± 0.31a
29	Octadece-9-nol	340	1.46 ± 0.12a	1.21 ± 0.12b	12.29 ± 0.62a	10.10 ± 0.72b
43	1-Eicosanol	368	0.69 ± 0.01	n.d.	5.77 ± 0.15a	n.d.
51	1-Docosanol	396	0.65 ± 0.01a	0.85 ± 0.00b	5.48 ± 0.13a	7.10 ± 0.19b
58	1-Octacosanol	482	0.97 ± 0.02a	1.02 ± 0.06a	8.13 ± 0.18a	8.54 ± 0.25b
71	1-Dotriacontanol	538	0.68 ± 0.00a	0.86 ± 0.01b	5.73 ± 0.15a	7.21 ± 0.21b
	Sterols		14.26 ± 1.04a	12.79 ± 1.01a	120.26 ± 11.78a	106.86 ± 6.16a
57	22-Stigmasten-3-one	412	0.26 ± 0.02	n.d.	2.15 ± 0.20a	n.d.
59	Stigmastane-3,6-dione	428	0.34 ± 0.06a	0.19 ± 0.01b	2.84 ± 0.58a	1.56 ± 0.09b
60	Campesterol	472	0.57 ± 0.02a	0.56 ± 0.03a	4.81 ± 0.29a	4.71 ± 0.26a
61	Stigmasterol	484	6.14 ± 0.52a	5.07 ± 0.29b	51.75 ± 5.69a	42.38 ± 1.99b
62	24-Ethyl-δ(22)-coprostenol	486	0.75 ± 0.10a	0.63 ± 0.04a	6.31 ± 0.92a	5.25 ± 0.31a
63	β-Sitosterol	486	0.78 ± 0.03a	0.90 ± 0.21a	6.53 ± 0.31a	7.46 ± 1.55a
64	4α,24-Dimethyl-5α-cholestan-3β-ol	488	1.09 ± 0.04a	0.95 ± 0.11a	9.22 ± 0.57a	7.97 ± 0.72b
66	4α-methyl,24-ethyl-5α-cholest-22E-en-3β-ol	500	0.77 ± 0.04a	0.70 ± 0.10b	6.52 ± 0.54a	5.82 ± 0.71a
68	4α-methyl-24-ethyl-5α-cholestan-3-ol	502	n.d.	0.34 ± 0.03	n.d.	2.86 ± 0.23
69	4α,24β-dimethyl-5α-cholestan-3β,4β-diol	504*	0.37 ± 0.04a	0.32 ± 0.02a	3.10 ± 0.44a	2.65 ± 0.2a
72	4α-methyl-24β-ethyl-5α-cholestan-3β,4β-diol	518*	0.71 ± 0.09a	0.58 ± 0.05b	6.04 ± 0.93a	4.88 ± 0.32a
67	Unidentified C30 Sterol		1.96 ± 0.12a	1.64 ± 0.18b	16.48 ± 1.35a	13.66 ± 1.12b
70	Unidentified C30 Sterol		0.17 ± 0.01a	0.21 ± 0.01b	1.45 ± 0.08a	1.76 ± 0.12b
65	Unidentified C30 Sterol		0.36 ± 0.02a	0.71 ± 0.07b	3.06 ± 0.26a	5.90 ± 0.46b
	Monoglycerides		0.27 ± 0.01a	0.04 ± 0.01b	2.31 ± 0.12a	0.37 ± 0.05b
40	1-Monotridecanoin	432	0.06 ± 0.00a	0.04 ± 0.01b	0.52 ± 0.05a	0.37 ± 0.05b
45	Monomyristin	446	0.09 ± 0.01	n.d.	0.72 ± 0.05a	n.d.
52	Monopalmitin	474	0.07 ± 0.01	n.d.	0.59 ± 0.07a	n.d.
55	Monostearin	502	0.06 ± 0.01	n.d.	0.48 ± 0.07a	n.d.
	Sugars		0.17 ± 0.01a	0.41 ± 0.02b	1.42 ± 0.10a	3.64 ± 0.02b
19	Rhamnose	452	n.d.	0.21 ± 0.01	n.d.	1.85 ± 0.02
27	Deoxyglucose	452	n.d.	0.20 ± 0.01	n.d.	1.79 ± 0.00
56	Glucosamine	612	0.17 ± 0.01	n.d.	1.42 ± 0.10a	n.d.
	Others		2.02 ± 0.02a	5.60 ± 0.07b	17.05 ± 0.83a	46.84 ± 1.29b
3	2,4,6,8-Tetramethyl-1-undecene	210	0.03 ± 0.00	n.d.	0.21 ± 0.04a	n.d.
5	2-Methyltetradecane	212	0.05 ± 0.01	n.d.	0.46 ± 0.07a	n.d.
8	2-Methyl-4-nonadecene	280	0.03 ± 0.00	n.d.	0.24 ± 0.04a	n.d.
10	3-Methyl-4-nonadecene	280	0.04 ± 0.01a	0.05 ± 0.01a	0.32 ± 0.07a	0.41 ± 0.10a
9	2,6-bis(1,1-Dimethylethyl)phenol	278	0.99 ± 0.02a	1.56 ± 0.08b	8.43 ± 0.43a	13.02 ± 0.28b
15	Methylsuccinic acid	276	0.05 ± 0.01	n.d.	0.45 ± 0.04a	n.d.
11	3-Methoxycinnamic acid	250	n.d.	1.18 ± 0.01	n.d.	9.88 ± 0.38
41	Dehydroabietic acid	372	0.18 ± 0.02a	0.66 ± 0.05b	1.49 ± 0.13a	5.55 ± 0.49b
54	Pinoresinol	502	n.d.	1.30 ± 0.03	n.d.	10.85 ± 0.55
23	Mannitol	614	0.65 ± 0.00a	0.85 ± 0.01b	5.46 ± 0.21a	7.13 ± 0.29b
	Total Identified		83.75 ± 2.00a	105.34 ± 5.90b	705.52 ± 37.96a	880.41 ± 29.68b

Values (means ± SD of four replicates) in the same row, not sharing a common superscript are significantly different (p < 0.05). Compounds are numbered by their elution order (see Fig. 2). All the compounds containing hydroxyl and/or carboxyl groups are identified as the correspondent TMS derivatives. MW – Molecular weight of compounds after silylation. aContains the iso- and anteiso-isomers; bContains cis and trans isomers; *Identified as the mono-TMS ether; n.d. – non detected; dw – dry weight.

Alkaline hydrolysis is often used for the analysis of compounds in their esterified forms[31]. In the chloroform extracts, submitted to alkaline hydrolysis, total fatty acids increased by 32% being the major increase verified for the unsaturated fatty acids (46%). This slight increase might be explained from poor extraction, such that only the smaller lipids will be available for derivatization and analysis, including volatile compounds. Moreover, these observations reveal that 32% of total fatty acids were in complex forms. Additionally, the absence of significant differences (p < 0.05) in sterols amounts, in contrast to, sugars and monoglycerides levels, after alkaline hydrolysis, indicates that the major complex forms present in the chloroform extracts of P. pinguis where mainly glycolipids and fatty acids esterified with glycerol (mono, di and triglycerides). The chromatogram obtained for the derivatized chloroform extract of P. pinguis after alkaline hydrolysis is displayed in Fig. 3. According to Milke et al.[11] the levels of free fatty acids in Pavlova spp., Chaetoceros muelleri and Placopecten magellanicus samples ranged 0.30 and 3.10%. In the P. pinguis studied, fatty acids before hydrolysis accounted 5.71% of microalgal dry weight. These differences might be explained by several factors such as: (i) the strain-to-strain variation; (ii) the cultivation conditions; (iii) the methodology applied.

Figure 3

Chromatogram of the derivatized P. pinguis lipophilic extract after alkaline hydrolysis. Peak identification as in Table 1. IS – Internal Standard (Tetracosane, 0.40 mg).

Fatty acids

In Fig. 4 it are represented the main families identified in the P. pinguis chloroform extractable substances before (Fig. 4a) and after (Fig. 4b) alkaline hydrolysis. Through this figure it is possible to visualize that fatty acids were the main family present in the chloroform extracts accounting up to 72% of the total compounds identified (Fig. 4b). The major fatty acids found in P. pinguis were palmitoleic (C16:1ω7 – PAA), myristic (C14:0 – MA), palmitic (C16:0 – PA) and eicosapentaenoic (C20:5ω3 – EPA) acids which together accounted over 30% of the total identified compounds before and after hydrolysis.

Figure 4

Main families identified in P. pinguis lipophilic extracts (a) before and (b) after alkaline hydrolysis, in percentage of the total identified compounds.

Microalgae are the primary producers of essential fatty acids that cannot be synthesized by humans, which, in turn, must obtain them through their diet[32]. These are linoleic (C18:2ω6 - LA) and α-linolenic (C18:3ω3 - ALA) acids which represent the omega-6 LC-PUFA and the omega-3 LC-PUFA, respectively[32]. P. pinguis presented a high content of LA (1.86 mg g−1 dw) and minor amounts of ALA (0.89 mg g−1 dw). As main precursor of the ω3 fatty acid synthesis, the minor amounts of ALA can be explained by the high levels of stearidonic (SA; 2.82 mg g−1 dw), EPA (3.49 mg g−1 dw) and docosahexaenoic (2.01 mg g−1 dw) acids[8,33]. These results are consistent with those in literature that point P. pinguis as a high value omega-3 LC-PUFA producing strain[14].

Docosahexaenoic acid (C22:6ω3 – DHA) is one of the main components of the structural lipids of the brain, whereas EPA display an important role in cardiovascular and immunological health[32]. The government health agencies worldwide recommend a dietary intake of DHA and EPA ranging from 200 and 670 mg day−1 [34]. In P. pinguis, DHA and EPA accounted a total of 5.50 mg g−1 dw, which means that 122 g of dry microalga represents the highest dietary reference value.

The ω6 fatty acids (main precursors of pro-inflammatory mediators) and the ω3 fatty acids (major precursors of anti-inflammatory molecules) compete for the same enzyme sets when metabolized[35]. Therefore, a balanced intake of ω6:ω3 fatty acids close to 1:1 is recommended. Western diets are characterized by high levels of ω6 Fatty acids and an unbalanced Σω6/Σω3 fatty acids ratio of 20:1 promoting the pathogenesis of various diseases[32]. This trend might be inverted by decreasing the intake of ω6 rich sources and increasing the intake of ω3 rich sources[32]. P. pinguis presented high amounts of ω3 fatty acids (13.36 mg g−1 dw) and a low Σω6/Σω3 fatty acids ratio (1:4) which, in turn, makes it suitable for dietary supply of ω3 fatty acids. This ratio was close to that obtained by Slocombe, et al.[14] (1:3) for P. pinguis.

Fatty alcohols

Fatty alcohols have been studied for their antibacterial activity and cholesterol-lowering ability[36,37]. The biological properties of these biomolecules are linked with the carbon chain length that is thought to determine their antibacterial activity and mode of action in biological systems[36]. Fatty alcohols accounted up to 12% of the total compounds identified (Fig. 4a). The major fatty alcohols found in P. pinguis were hexadecanol (C16-OH), octadece-9-nol (C18:1-OH) and octadecanol (C18-OH) which together accounted up to 40% of the total fatty alcohols. The mass fragmentation of C18:1-OH is displayed in Fig. 5a. Through this figure it is possible to observe three key fragment ions from alcohols: the base peak at m/z 75 [(CH3)2SiOH]+, the m/z 325 [M − 15]+ and the molecular ion [M]+ at m/z 340.

Figure 5

Mass spectra of some assigned peaks as trimethylsilyl (TMS) ethers and/or esters: (a) Octadece-9-nol (peak 29), (b) Monomyristin (peak 45), (c) Pinoresinol (peak 54) and (d) 4α-methyl-24β-ethyl-5α-cholestan-3β,4β-diol as mono TMS ether (peak 72).

In P. pinguis, it was identified two very long chain alcohols: octacosanol (C28-OH) and dotriacontanol (C32-OH). The consumption of 5–20 mg day−1 of very long aliphatic chain alcohols is known to decrease the low-density lipoprotein (LDL) cholesterol[9,37]. Therefore, 3.04–12.15 g of microalgal biomass and 0.36–1.44 g of microalgal extract are the quantities needed to fulfill these requirements (Table 1). Moreover, alcohols such as undecanol (C11-OH), dodecanol (C12-OH) and tridecanol (C13-OH), have been pointed in previous studies by their bactericidal activity[36]. After alkaline hydrolysis the fatty alcohols increased by 12%, with the highest increase verified for the C12-OH (32%).

The detected docosanol (C22-OH) is known by its antiproliferative effect of chinese hamster ovary cells K1 (CHO-K1) and human melanoma (CRL-1974TM) cell lines[38]. Investigations concerning the effects of long chain alcohols are often performed with long chain alcohols isolated from sugarcane where these compounds make up 0.10–0.30% of its mass[39]. In the present study this class of compounds comprised 0.16 and 0.19% of P. pinguis dried biomass, before and after alkaline hydrolysis respectively.

Sterols

Microalgae are recognized for their wide diversity of sterols that are often used for chemotaxonomic and phylogenetic comparisons[7]. This diversity along with the high sterol content make microalgae promising sources of novel sterols with potential novel activities[40].

Sterols accounted up to 17% of the total identified compounds (Fig. 4a). The major sterols were stigmasterol (40–43%), 4α, 24-Dimethyl-5α.cholestan-3β-ol (7–8%) and an unidentified sterol (13–14%), which together accounted over 50% of total sterols (Table 1). The dominance of stigmasterol across Pavlova species namely P. pinguis have been reported in studies targeting aquaculture and chemotaxonomy[11,22]. β-sitosterol, campesterol and stigmasterol - in their non-esterified forms, have been subject to the Food and Drug Admnistration (FDA) health claim for reduced risk of coronary heart disease[40]. P. pinguis presented high levels of stigmasterol which alone represented up to 7% of the total identified compounds in the lipophilic profile. Campesterol and β-sitosterol were also identified in the P. pinguis chloroform extracts accounting 0.57 and 0.78 mg g−1 of microalgal biomass and 4.81 and 6.53 mg g−1 of extract, respectively (Table 1). In contrast to Milke, et al.[11] stigmastanol and cholesterol were not detected in P. pinguis lipophilic extracts.

Pavlova species are often recognized by their unusual dihydroxylated sterols called Pavlovols[17]. In P. pinguis two dihydroxylated sterols were identified, 4α-methyl-24β-ethyl-5α-cholestan-3β, 4β-diol (ethylpavlovol) and 4α, 24β-dimethyl-5α-cholestan-3β, 4β-diol (methylpavlovol), and a structurally isomeric form of dinosterol, 4α-methyl-24-ethyl-5α-cholest-22E-en-3β-ol. As with fatty acids, the sterol profiles of microalgae are species-specific and often used as chemotaxonomic markers[17].

Despite in Milke, et al.[11] the pavlovols have not been found for Pavlova species, the authors recognize that they can constitute the sterol composition of this microalga specie. Moreover, Volkman, et al.[22] found the existence of these unusual sterols in the composition of two other P. pinguis strains and pointed pavlovols as chemotaxonomic markers of Pavlovales.

Figure 5b shows the mass spectrum of the compound identified as ethylpavlovol. Through comparison of the obtained mass spectrum with the one obtained previously by Volkman, et al.[22], it was possible to identify the ethylpavlovol as its mono TMS ether. The assignment was done by the presence of the following mass fragments: m/z 43, 487, 503 and 518, as well as, the base peak at m/z 371 [M-147(C3H6O2TMS)]+.

Phytosterols are playing a key role in nutraceutic and pharmaceutical industries, as precursors of some bioactive molecules[7,40]. Moreover, it is estimated that the dietary intake of phytosterols is in the range of 150 to 400 mg day−1. P. pinguis can contribute to the intake of around 143 mg of free sterols per 100 g of microalgal dry weight. After alkaline hydrolysis it were not verified significant differences (p < 0.05) in the amounts of sterols (Table 1). This observation indicates that sterols were non-esterified and were as free sterols.

Other compounds

In the classes of monoglycerides, sugars and other components, compositional differences before and after hydrolysis were verified. In Table 1 it is possible to observe that after alkaline hydrolysis the monoglycerides: monomyristin, monopalmitin and monostearin, were not detected. Sugars and the other components classes were those who presented the highest increase after alkaline hydrolysis, 3 and 5 times higher respectively. The increase of the sugar content suggests the presence of polar lipids, namely glycolipids incorporating rhamnose and deoxyglucose, in the chloroform extracts of P. pinguis. The sulfoquinovosyl diacylglycerols (SQDGs) are one of the most abundant glycolipids found in microalgal cells[41]. SQDGs are constituted by a 6-sulfoquinovose unit, which, in turn, is constituted by a sulphur group attached to the quinovose (6-deoxyglucose)[42]. Moreover, it has been reported that microalgal glycolipids may contain other sugar moieties than galactose such as mannose and rhamnose[43].

The mass fragmentation of monomyristin is presented in Fig. 5c. Although in the mass spectrum it is not possible to visualize the molecular ion, the presence of the following ions: m/z 73, 103, 147, 205 347 indicates that this compound is monomyristin. The fragment m/z 347 corresponds to the mass fragment [M-103 (CH2OTMS)]+ and the m/z 103, 147 and 205 are associated to the silylated glycerol backbone.

The detection of smaller amounts of 3-methocycinnamic acid and pinoresinol 1.12 and 1.23% of total identified compounds, respectively, was observed after alkaline hydrolysis. According to Klejdusa, et al.[44] the cinnamic acid derivatives are precursors in the phenyl-propanoid pathway for the synthesis of polyphenols, which indicates that it is possible that this microalgal strain have other phenols that could be extracted by polar solvents. In Fig. 5d it is possible to visualize the fragmentation pattern of obtained for pinoresinol, namely the base peak m/z 73 [(CH3)3Si]+, the m/z 487 resultant from the loss of a methyl group [M-15]+ and the molecular ion [M]+ m/z 502. Phenols are natural products that are recognized by their antioxidant, antimicrobial and antiviral activities[44].

P. pinguis chloroform extracts presented a strong concentration-dependent DPPH radical scavenging activity with a determination coefficient close to 1 (R2 = 0.99), Fig. 6. The presence of phenols in the lipophilic fraction after alkaline hydrolysis suggest that this ability might be resultant from polyphenol-associated lipids. The estimated EC50 for P. pinguis chloroform extracts was of 1 057 µg mL−1.

Figure 6

2,2-diphenyl-1-picrylhydrazyl (DPPH) radical scavenging activity (%) of P. pinguis chloroform extracts (µg mL−1). Butylated hydroxytoluene (BHT) was used as a reference compound and its EC50 value is 9.79 𝜇g mL−1.

Besides nutraceutics and pharmaceutics the rich composition verified for P. pinguis as for other species of the genus Pavlova[45] can also be used in aquaculture for animal consumption as dietetic supply and in food industry as additive and/or nutritional supplements attributing to consumer a higher level of bioactive compounds.

Conclusions

The need for naturally derived health promoting phytochemicals instead of the chemically derived drugs have prompted the search for new sources of natural products. Pavlova pinguis presented a manifold range of metabolites which demonstrated its versatility and potential as a source of high value compounds. The high content of unsaturated fatty acids, long chain aliphatic alcohols and stigmasterol, demonstrate the potential of this microalga not only for aquaculture but also for nutraceutics and pharmaceutics uses. To fully exploit the phytochemical features of microalgae for commercial purposes, a non-targeted approach should be taken in order to uncover whole extract chemical diversity.