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. 2018 May 8;13(5):e0196462. doi: 10.1371/journal.pone.0196462

Phytochemical study of the headspace volatile organic compounds of fresh algae and seagrass from the Adriatic Sea (single point collection)

Igor Jerković 1,*, Zvonimir Marijanović 2, Marin Roje 3, Piotr M Kuś 4, Stela Jokić 5, Rozelinda Čož-Rakovac 6
Editor: Vijai Gupta7
PMCID: PMC5940206  PMID: 29738535

Abstract

Performed phytochemical study contributes to the knowledge of volatile organic compounds (VOCs) of Halopteris filicina (Grateloup) Kützing, Dictyota dichotoma (Hudson) J. V. Lamouroux, Posidonia oceanica (L.) Delile and Flabellia petiolata (Turra) Nizamuddin from the Adriatic Sea (single point collection). VOCs were investigated by headspace solid-phase microextraction (HS-SPME) and analysed by gas chromatography and mass spectrometry (GC-MS/FID). H. filicina headspace contained dimethyl sulfide (DMS; 12.8%), C8-compounds (e.g. fucoserratene (I; 9.5%)), benzaldehyde (II; 8.7%), alkane C17, dictyopterene D and C (III, IV), tribromomethane (V), 1-iodopentane, others. F. petiolata headspace was characterized by DMS (22.2%), 6-methylhept-5-en-2-one (9.5%), C17 (9.1%), II (6.5%), compounds I-V. DMS (59.3%), C15 (14.5%), C17 (7.2%) and C19 (6.3%) dominated in P. oceanica headspace. Sesquiterpenes were found in D. dichotoma, predominantly germacrene D (28.3%) followed by other cadinenyl (abundant), muurolenyl and amorphenyl structures. Determined VOCs may be significant for chemosystematics and chemical communications in marine ecosystem.

Introduction

Volatile organic compounds (VOCs) are low-molecular compounds with low to moderate hydrophilicity that can not only be dissolved in water, but also dissipate into the gas phase at air-water interfaces. Prior to 1966, only one volatile substance was identified from wet, undecomposed seaweed and described in the literature as dimethyl sulfide (DMS). Afterwards, the number of identified marine plant VOCs has been constantly growing. In 1976, VOCs from marine algae were reviewed by Moore [1] including non-isoprenoid C11-compounds, acyclic undecapolyenes, less volatile organosulfur compounds, bromine- and iodine-containing haloforms, halogenated compounds, others. The research on the algae VOCs has been continued by different teams [2–4]. The production and role of volatile halogenated compounds from marine algae were reviewed in 2011 [5]. Recently, more research papers on this topic appeared (e.g. [6,7]). VOCs biogeneration by microalgae, their occurrence, behaviour, ecological implications and industrial applications were described in 2016 [8]. The volatile metabolites emission by in vivo microalgae were reviewed in 2017 [9]. Tricyclic sesquiterpenes from marine origin were systematically presented in 2017 [10].

The common VOCs released by terrestrial plants are: ‘‘green leaf volatiles” (GLVs) which consist almost exclusively of C6 aldehydes and alcohols [11], other aliphatic compounds, monoterpenes, sesquiterpenes, phenylpropane derivatives, others [12]. This is in remarkable contrast to marine VOCs that show much higher structural diversity [13]: e.g. only among marine sesquiterpenes 54 different skeletal types of tricyclic sesquiterpenes were found [10]. In terrestrial ecosystems VOCs are very important group of infochemicals (e.g. plant-herbivore interactions, pheromones). Similarly, VOCs of marine organisms can have multiple functions in ecosystem in intraspecific (pheromones) and interspecific (kairomones) communication and in activated defences [14]. In addition, these compounds (usually present in the extracts) may exhibit other biological activities, e.g. antibacterial, cytotoxic and antitumor [10]. Different methods have been used to investigate VOCs from marine plants such as liquid-liquid extraction (LLE), hydrodistillation (HD), simultaneous distillation-extraction (SDE), vacuum-SDE (V-SDE), static and dynamic headspace extraction techniques (SHE and DHE) as well as headspace solid-phase microextraction (HS-SPME).

However, limited number of studies have been carried out on marine algae and seagrass headspace VOCs in general [6,7,15,16]. In addition, to the best of our knowledge, there is only one paper [17] about VOCs (obtained by SDE) of the algae from the Adriatic Sea reporting the chemical composition of Padina pavonia (L.) Gaill. Considering limited data available on the chemical composition of marine plants found in the Adriatic Sea, the focus of this study was on 4 different samples collected from the same location (single point collection): 3 seaweeds (marine macroalgae) - 2 brown algae (Halopteris filicina (Grateloup) Kützing, Dictyota dichotoma (Hudson) J. V. Lamouroux) and 1 green alga (Flabellia petiolata (Turra) Nizamuddin) as well as 1 seagrass (Posidonia oceanica (L.) Delile). F. petiolata has been commonly found in Mediterranean basin [18], often in association with other algae (e.g. Dictyopteris spp., Dictyota spp., Dilophus spp.). It is one of the main components of the phytocoenoses associated with the endemic and endangered seagrass P. oceanica [19,20]. The specific goals of present phytochemical study are: 1) to determine and unlock the headspace VOCs composition of targeted marine plants by headspace solid-phase microextraction (HS-SPME)/gas chromatography and mass spectrometry (GC-MS/FID)—first time report; 2) to compare the chemical biodiversity of found VOCs among the samples with other marine plants with same, similar or related constituents; 3) to discuss possible biosynthetic origin of identified VOCs from the literature data.

Materials and methods

The marine plant samples

We state clearly that no specific permissions were required for the location and collection. There is no need to issue the permission to collect the samples used in this work because the sample were not taken from national park or protected area, but from the public place–sea. The Ministry approved this type of research by approving our research projects: Croatian Science Foundation under the project HRZZ-IP-11-2013-8547 (Research of Natural Products and Flavours: Chemical Fingerprinting and Unlocking the Potential) and by the Croatian Ministry of Science and Education (MZO grant no. KK.01.1.1.01.0002 –The Scientific Center of Excellence for Marine Bioprospecting-BioProCro). We confirm that the field study did not involve endangered or protected species.

The samples of 2 brown algae (Halopteris filicina (Grateloup) Kützing, Dictyota dichotoma (Hudson) J. V. Lamouroux) and 1 green alga (Flabellia petiolata (Turra) Nizamuddin) as well as 1 seagrass (Posidonia oceanica (L.) Delile) were collected in the middle part of the Adriatic Sea, close to Mala Smokvica island, in July 2017 at the same geographic location (43o31'04.4"N, 15o56'32.6"E). Single point sample collection provided representative robust samples for first time chemical profiling. The samples were collected at 11 a.m. under sunny conditions—constantly illuminated, and area receive direct sunlight for >80% of the time from sunrise to sunset. During the samples collection there was no wind and the sea temperature was 22°C. The algae were collected from depth of 0.5–3 m and the sea was collected from the same depth (directly into the plastic bag where collected algae were placed). The samples were separately collected and separately placed in air tight plastic bags containing seawater and were immediately transported to the laboratory. The samples were kept in dark at 4°C until the extractions that were performed within 24 h after the collection. Before HS-SPME, each sample was taken out of the bag, cut into small pieces, and the excess of seawater was removed by placing it between filter paper layers for 2 min (the seawater was not removed completely).

Headspace solid-phase microextraction (HS-SPME)

The headspace extraction was performed with a manual SPME holder using polydimethylsiloxane/divinylbenzene (PDMS/DVB) fibre obtained from Supelco Co. (Bellefonte, PA, USA). The fibre was conditioned prior to the extraction according to the instructions by Supelco Co. For HS-SPME, previously prepared samples (1 g) were placed separately in 5 mL glass vials and hermetically sealed with PTFE/silicone septa. The vials were maintained in a water bath at 60 oC during equilibration (15 min) and HS-SPME (45 min). After sampling, the SPME fibre was withdrawn into the needle, removed from the vial, and inserted into the injector (250 oC) of the GC-FID and GC-MS for 6 min where the extracted volatiles were thermally desorbed directly to the GC column. HS-SPME was done in triplicate for each sample.

Gas chromatography and mass spectrometry (GC-FID and GC-MS) analyses

The GC-FID analyses were carried out with an Agilent Technologies (Palo Alto, CA, USA) gas chromatograph model 7890A equipped with a flame ionization detector (FID) and a HP-5MS capillary column (5% phenyl-methylpolysiloxane, Agilent J and W). The GC conditions were similar to those described previously [21]. In brief, the oven temp. was programmed isothermal at 70 oC for 2 min, increasing from 70–200 oC at 3 oC min-1, and held isothermally at 200 oC for 15 min; carrier gas was helium (He 1.0 mL min-1). The GC-MS analyses were performed using an Agilent Technologies (Palo Alto, CA, USA) gas chromatograph model 7820A equipped with a mass selective detector (MSD) model 5977E (Agilent Technologies) and a HP-5MS capillary column, under the same conditions as for the GC-FID analysis. The MSD (EI mode) was operated at 70 eV, and the mass range was 30–300 amu.

The identification of VOCs was based on the comparison of their retention indices (RI), determined relative to the retention times of n-alkanes (C9-C25), with those reported in the literature [22] and their mass spectra with the spectra listed in Wiley 9 (Wiley, New York, NY, USA) and NIST 14 (D-Gaithersburg) mass spectral libraries. The percentage composition of the samples was computed from the GC peak areas using the normalization method (without correction factors). The average component percentages in Table 1 were calculated from GC-FID and GC-MS analyses (triplicates).

Table 1. VOCs composition determined by headspace solid-phase microextraction (HS-SPME).

No. Compound Molecular formula Compound class RI Area percentage (%)
1 2 3 4
1. Ethanol C2H5O Alcohol < 900 1.2 0.6 5.8 1.2
2. Methylthiomethane (Dimethyl sulfide) C2H6S Organosulfur compound < 900 12.8 22.2 59.3 1.0
3. 3-Methylbutanal C5H10O Aldehyde < 900 0.9 1.0 0.0 0.0
4. 2-Methylbutanal C5H10O Aldehyde < 900 0.5 0.0 0.0 0.0
5. Benzene C6H6 Aromatic compound < 900 0.0 0.0 0.0 0.1
6. Pent-1-en-3-one C5H8O Unsaturated ketone < 900 2.6 2.9 0.0 0.2
7. Pentanal C5H10O Aldehyde < 900 0.0 1.6 0.0 0.1
8. (2E)-Pent-2-enal C5H8O Aldehyde < 900 0.0 0.0 0.0 0.1
9. Dimethyl disulfide C2H6S2 Organosulfur compound < 900 0.0 0.0 0.0 0.1
10. 3-Methylbut-2-enal C5H8O Aldehyde < 900 1.5 1.6 0.0 0.1
11. (2Z)-Pent-2-en-1-ol C5H10O Unsaturated alcohol < 900 0.9 0.6 0.0 0.1
12. Methylbenzene C7H8 Aromatic compound < 900 0.0 0.0 0.0 0.1
13. Hexanal C6H12O Aldehyde < 900 4.7 1.5 0.2 0.3
14. (3E)-Octa-1,3-diene C8H14 Alkene < 900 2.3 0.7 0.0 0.7
15. (2E)-Hex-2-enal C6H10O Unsaturated aldehyde < 900 0.5 0.2 0.0 0.0
16. Hexan-1-ol C6H14O Alcohol < 900 2.4 0.0 0.0 0.0
17. (3Z,5E)-Octat-1,3,5-triene (Fucoserraten) C8H12 Alkene < 900 9.5 3.2 0.0 1.6
18. Tribromomethane CHBr3 Polyhalo-methane < 900 2.1 1.8 0.0 0.0
19. Heptanal C7H14O Aldehyde 903 0.3 0.5 0.0 0.1
20. 1-Iodopentane* C5H11I Organoiodine 929 0.8 0.0 0.0 0.0
21. (2E,4E,6E)-Octa-2,4,6-triene* C8H12 Alkene 941 0.2 0.0 0.0 0.1
22. α-Pinene C10H16 Monoterpene 942 0.0 0.0 0.1 0.0
23. (2Z)-Hept-2-enal C7H12O Unsaturated aldehyde 961 0.3 0.6 0.0 0.1
24. Benzaldehyde C7H6O Aldehyde 968 8.7 6.5 0.2 0.5
25. Dimethyl trisulfide C2H6S3 Organosulfur compound 978 0.0 0.0 0.0 0.1
26. (5Z)-Octa-1,5-dien- 3-ol C8H14O Unsaturated alcohol 978 0.7 0.0 0.0 0.0
27. Oct-1-en-3-one C8H14O Unsaturated ketone 981 0.5 1.1 0.0 0.1
28. Oct-1-en-3-ol C8H16O Unsaturated alcohol 983 1.2 2.5 0.0 0.2
29. Octan-3-one C8H16O Ketone 987 4.1 0.0 0.0 0.0
30. 6-Methylhept- 5-en-2-one C8H14O Unsaturated ketone 989 0.0 9.5 0.0 0.7
31. Octan-2-one C8H16O Ketone 993 0.0 1.6 0.0 0.0
32. 2-Pentylfuran C9H14O Furan derivative 994 0.7 0.0 0.2 0.0
33. (2E,4Z)-Hepta-2,4-dienal C7H10O Unsaturated aldehyde 999 2.4 0.7 0.0 0.2
34. Octanal C8H16O Aldehyde 1004 4.7 0.5 0.0 0.0
35. (2E,4E)-Hepta-2,4-dienal C7H10O Unsaturated aldehyde 1014 0.7 0.8 0.0 0.0
36. 1,8-Cineole C10H18O Monoterpene 1037 0.0 0.6 0.0 0.1
37. Phenylacetladehyde C8H8O Aromatic aldehyde 1050 0.9 0.7 0.1 0.0
38. (2E)-Oct-2-enal C8H14O Unsaturated aldehyde 1062 0.7 0.5 0.0 0.1
39. Octan-1-ol C8H18O Alcohol 1077 5.1 0.0 0.0 0.3
40. Nonanal C9H18O Aldehyde 1105 0.4 0.3 0.0 0.1
41. 6-[(1Z)-1-Butenyl]-1,4-cycloheptadiene (Dictyopterene D) C11H16 Alkene 1156 1.9 7.4 0.0 0.0
42. 6-Butyl-1,4-cycloheptadiene (Dictyopterene C) C11H18 Alkene 0.7 0.7 0.0 0.0
43. Decanal C10H20O Aldehyde 1207 0.2 0.0 0.0 0.0
44. β-Cyclocitral C10H16O C13-Norisoprenoid 1223 0.5 0.8 0.0 0.0
45. Decan-1-ol C10H22O Alcohol 1279 0.0 0.0 0.0 0.1
46. (2E,4Z)-Deca-2,4-dienal C10H16O Unsaturated aldehyde 1295 0.5 0.0 0.0 0.0
47. (2E,4E)-Deca-2,4-dienal C10H16O Unsaturated aldehyde 1318 0.4 0.0 0.0 0.0
48. α-Cubebene C15H24 Sesquiterpene 1352 0.0 0.0 0.0 1.5
49. Cycloisosativene C15H24 Sesquiterpene 1367 0.0 0.0 0.0 0.5
50. α-Ylangene C15H24 Sesquiterpene 1372 0.0 0.0 0.0 0.3
51. α-Copaene C15H24 Sesquiterpene 1377 0.0 0.0 0.0 0.7
52. β-Bourbonene C15H24 Sesquiterpene 1386 0.0 0.0 0.0 5.1
53. β-Cubebene C15H24 Sesquiterpene 1390 0.0 0.0 0.0 0.8
54. Tetradec-1-ene C14H28 Alkene 1393 0.0 1.4 0.0 0.0
55. Tetradecane C14H30 Alkane 1400 0.0 0.0 0.6 0.0
56. Dodecanal C12H24O Aldehyde 1409 0.3 0.0 0.0 0.0
57. α-Ionone C13H20O C13-Norisoprenoid 1429 0.4 0.9 0.0 0.0
58. Aromadendrene C15H24 Sesquiterpene 1440 0.0 0.0 0.0 0.3
59. (E)-β-farnesene C15H24 Sesquiterpene 1459 0.0 0.0 0.0 1.0
60. γ-Muurolene C15H24 Sesquiterpene 1465 0.0 0.0 0.0 2.1
61. β-Cadinene C15H24 Sesquiterpene 1474 0.0 0.0 0.0 2.8
62. α-Amorphene C15H24 Sesquiterpene 1478 0.0 0.0 0.0 3.5
63. Dodecan-1-ol C12H26O Alcohol 1479 0.4 0.0 0.0 0.0
64. Germacrene D C15H24 Sesquiterpene 1481 0.0 0.0 0.0 28.3
65. Ar-curcumene C15H22 Sesquiterpene 1484 0.7 0.0 0.0 0.9
66. β-Ionone C13H20O C13-Norisoprenoid 1487 1.2 0.7 0.0 0.0
67. Pentadec-1-ene C15H30 Alkene 1493 3.3 3.2 0.0 0.0
68. Bicyclogermacrene C15H24 Sesquiterpene 1496 0.0 0.0 0.0 4.7
69. Epizonarene C15H24 Sesquiterpene 1498 0.0 0.0 0.0 4.3
70. α-Muurolene C15H24 Sesquiterpene 1499 0.0 0.0 0.0 2.2
71. Pentadecane C15H32 Alkane 1500 2.6 1.4 14.5 0.0
72. δ-Selinene C15H24 Sesquiterpene 1507 0.0 0.0 0.0 2.7
73. Tridecanal C13H26O Aldehyde 1511 0.5 0.0 0.2 0.0
74. γ-Cadinene C15H24 Sesquiterpene 1515 0.0 0.0 0.0 3.4
75. δ-Cadinene C15H24 Sesquiterpene 1525 0.0 0.0 0.0 8.3
76. trans-γ-Bisabolene C15H24 Sesquiterpene 1533 0.3 0.0 0.0 0.0
77. trans-Cadina-1,4-diene C15H24 Sesquiterpene 1534 0.0 0.0 0.0 1.2
78. α-Cadinene C15H24 Sesquiterpene 1539 0.0 0.0 0.0 0.9
79. α-Calacorene C15H24 Sesquiterpene 1545 0.0 0.0 0.0 1.3
80. Germacrene B C15H24 Sesquiterpene 1555 0.3 0.0 0.0 3.6
81. Dendrolasin C15H22O Furan derivative 1575 0.3 1.2 0.0 0.0
82. Hexadecane C16H34 Alkane 1600 0.0 0.0 0.4 0.0
83. α-Cadinol C15H26O Sesquiterpene 1645 0.0 0.0 0.0 0.2
84. τ-Muurolol C15H26O Sesquiterpene 1657 0.0 0.0 0.0 0.1
85. (8E)-Heptadec-8-ene C17H34 Alkene 1679 0.7 1.6 0.1 0.0
86. Heptadec-1-ene C17H34 Alkene 1693 1.1 0.6 0.0 0.0
87. Heptadecane C17H36 Alkane 1700 4.0 9.1 7.2 0.1
88. Octadecanal C18H36O Aldehyde 1715 0.3 0.0 1.5 0.0
89. Octadecane C18H38 Alkane 1800 0.0 0.0 0.6 0.0
90. Nonadecane C19H40 Alkane 1900 0.0 0.0 6.3 0.1
91. Heneicosane C21H44 Alkane 2100 0.0 0.0 0.6 0.0
Total identified (%) 95.0 92.8 97.9 89.9
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The symbols in the table are: 1 –H. filicina; 2 –F. petiolata; 3 –P. oceanica; 4 –D. dichotoma; RI–retention indices relative to C9-C25 alkanes

*—tentatively identified

Results and discussion

Headspace solid-phase microextraction (HS-SPME) with the fibre polydimethylsiloxane/ divinylbenzene (PDMS/DVB) was successfully applied first time to investigate VOCs from four fresh marine plants (Halopteris filicina (Grateloup) Kützing, Dictyota dichotoma (Hudson) J. V. Lamouroux, Posidonia oceanica (L.) Delile and Flabellia petiolata (Turra) Nizamuddin) that were collected from the Adriatic Sea from the same location. Single point sample collection provided representative robust samples for first time chemical profiling. Representative chemical profiles of their headspace composition were obtained without the application of heat that can lead to the artefacts (as in other methods for VOCs isolation e.g. hydrodistillation). The first insight into the results (Table 1) reveals striking differences among the obtained chemical profiles. As was expected, significant contrast with common VOCs released by terrestrial plants can also be immediately noticed. The results are further discussed in separate subtitles for each seaweed or seagrass and for possible biosynthetic origin of identified VOCs.

Halopteris filicina (Grateloup) Kützing headspace VOCs

Halopteris filicina (Grateloup) Kützing is a greenish-brown, fern-like seaweed that belongs to the family Stypocaulaceae and it has been usually found in the Mediterranean and warm seas. Preliminary phytochemical screening of its crude extracts revealed the presence of alkaloids, saponins, flavonoids and terpenes [23]. Free amino acids, amino sulfonic acids, sugars and sterols have been examined and quantitatively determined [24], but there is no report on its volatile constituents. Its methanolic extract showed inhibition against Staphylococcus aureus, Staphylococcus epidermidis, Bacillus subtilis, Bacillus spp., Salmonella typhi, and Escherichia coli [23].

The headspace VOCs composition of H. filicina is reported in Table 1. Dimethyl sulfide (DMS) was the most abundant (12.8%). DMS serves as a chemoattractant for phytoplankton, bacteria, zooplankton, fish, and sea birds [25,26]. Tribromomethane (2.1%) was found and it can be classified as polyhalomethane. The VOCs of Ascophyllum nodosum contained tribrommethane [27]. One organoiodine was identified: 1-iodopentane (0.8%). Asparagopsis taxiformis, Asparagopsis armata and Falkenbergia rufolanosa synthesized more than 100 different organoiodines including 1-iodopentane [28]. It has also been demonstrated that the volatile halogenated compounds are involved in the defence mechanism (allelopathy) of the algae [29] as one of several types of allelochemicals. They are probably important in the global cycling of gaseous organohalogen species [30]. The most widespread were aliphatic compounds, particularly C8-compounds (total 9 compounds, Table 1) such as: octan-1-ol (5.1%), octanal (4.7%), octan-3-one (4.1%), (3E)-octa-1,3-diene (2.3%), (3Z,5E)-octat-1,3,5-riene (9.5%), or oct-1-en-3-ol (1.2%). Two typical marine algae C11-hydrocarbons were also present: dictyopterene D (1.9%) and dictyopterene C (0.7%). Most of the chemical signals (specific volatile pheromones) in various genera of macroalgae are unsaturated, acyclic and/or alicyclic C11-hydrocarbons of different ring size and different degrees of unsaturation which are active at picomolar concentrations [14,31]. C8-Hydrocarbon fucoserratene ((3Z,5E)-octa-1,3,5-triene) is also used for intraspecific chemical communication (pheromone) in Fucales [14,32]. Another group of abundant aliphatic compounds were higher acyclic hydrocarbons such as heptadecane (4.0%), pentadecane (2.6%) or pentadec-1-ene (3.3%). Pentadecane was most typical in Ulva spp. and two species of Enterornorpha. Unbranched alkanes were identified in Ulva rigida with heptadecane predominance [33]. Three C13-norisoprenoids were detected: α-ionone (0.4%), β-ionone (1.2%) and β-cyclocitral (0.5%). C13-norisoprenoids, such as the potent flavour compound β-ionone, are produced by a diversity of algae taxa, for example Ulothrix fimbriata and Porphyra tenera [14]. Two phenypropane derivatives were found (Table 1): benzaldehyde (8.7%) and phenylacetladehyde (0.9%). Benzaldehyde was predominant in seaweed Monostroma nitidum [6].

Previous research applying GC-MS on H. filicina reported the amounts of fatty acids (evaluated as the sum of all fatty acids (originally present and those resulting from the alkaline hydrolysis) by GC-MS after derivatization with BF3 and purification) that ranged from 74 to 1897 mg kg-1, dry weight [34]. H. filicina significantly differs from the other species as it shows a much smaller fatty acids total content, although it contains important acids (e.g. hexadecanoic acid (C16:0), octadecanoic acid (C18:0), or eicosapentaenoic acid (EPA, C20:5 ω-3)). In addition, valuable compounds in macroalgae extracts were determined [35] by GC-MS after derivatisation with trimethylsilyltrifluoroacetamide (MSTFA). H filicina extract contained proline, mannitol, hexadecanoic acid, octadecanoic acid, cholestanol and fucosterol. Sterols were identified by comparison of GC retention data of their TMSi ethers with those of authentic compounds and by GC-MS of their acetates [24]. The sterol fraction contained fucosterol, cholesterol and 24-methylenecholesterol; minute amounts of 22-dehydrocholesterol and 24-methyl-cholesta-5,22-dien-3β-ol.

Flabellia petiolata (Turra) Nizamuddin headspace VOCs

Flabellia petiolata (Turra) Nizamuddin is a green alga that belongs to Udoteaceae family (Chlorophyta, Bryopsidales) commonly found in the Mediterranean basin [36]. Compared to many other green algae, F. petiolata appears to be a particularly interesting species, since antibacterial, antiviral, antimitotic, antifungal and cytotoxic activities of its raw extract have been observed [37]. Selected constituents from F. petiolata from Turkey were determined [38] such as: the average cellulose content (18.86±0.69%), the average crude protein content (22.45±0.62%) and the crude fat content (1.08±0.24%). There were no available data on its VOCs composition.

DMS, a highly-volatile organosulfur compound was quite abundant in the headspace of F. petiolata (22.2%; Table 1) and in general serves to the algae as a chemoattractant [25,26]. Since DMS distribution in the water column does not correlate well with phytoplankton biomass [39], it led to suggestions that only certain groups of marine algae (e.g. Emiliania huxleyi) produce significant amounts of DMS. Polyhalomethane tribromomethane was present with low percentage (1.8%). Variety of aliphatic compounds were found in the headspace (Table 1). Low-molecular aliphatic compounds up to 7 carbons, particularly ketones and aldehydes, were quite widespread, e.g. 3-methylbutanal (1.0%), pentanal (1.6%), 3-methylbut-2-enal (1.6%), hexanal (1.5%), or pent-1-en-3-one (2.9%). In distinction from lipid-derived low-molecular aliphatic compounds, 3-methylbutanal is derived from leucine [16] and it is generally associated with heated products; however fresh P. petiolata was used for this research. It is frequently found in many sea products, such as crayfish or lobster as well as in red algae Palmaria palmate [14,38]. 6-Methylhept-5-en-2-one (9.5%) was the most abundant ketone and previously it was found in relatively high amount (along with 6-methylheptan-2-one) in V-SDE extract of the green alga Capsosiphon fulvescens [40]. Seven C8-compounds were identified with major representatives (3Z,5E)-octat-1,3,5-riene (fucoserratene; 3.2%), oct-1-en-3-ol (2.5%) and oct-1-en-3-one (1.1%). Identified dictyopterene D (7.4%) and dictyopterene C (0.7%) belong to C11-hydrocarbons (volatile pheromones). Heptadecane (9.1%) was the most abundant among alkanes and it was already observed to be one of the most common hydrocarbons in marine algae, detected in relevant amount (26.38%) in Hypnea cornuta [33]. C13-norisoprenoids were represented by α-ionone (0.9%), β-ionone (0.7%) and β-cyclocitral (0.8%). β-Ionone and β-cyclocitral were found previously among the major volatile compounds of the essential oil isolated from Capsosiphon fulvescens [40] by V-SDE. Benzaldehyde was the most abundant (6.5%) among phenylpropane derivatives. Only one monoterpene was found in low percentage (1,8-cineole; 0.6%).

Posidonia oceanica (L.) Delile headspace VOCs

Posidonia oceanica (L.) Delile represents one of the most widespread species endemic to the Mediterranean Sea. Previous phytochemical studies showed that the mature leaves of P. oceanica are rich in amino acids, chicoric acid, p-coumaric acid, vanillin, ferulic acid, gentisic acid, caffeic and cinnamic acids, sterols and phenolic derivatives [41]. In chloroformic extract from P. oceanica a novel methylated sesquiterpene was identified (2E)-3,7,12-trimethyltridec-2-en-1-ol named posidozinol along with β-sitosterol and four fatty acids (palmitic, palmitoleic, oleic and linoleic) [42]. The occurrence of dimethylsulfoniopropionate (DMSP) and its variability during the seasonal cycle in the leaves of P. oceanica was reported [43]. However, there is no report on its headspace VOCs.

The main compound of P. oceanica headspace was DMS (59.3%; Table 1) that was expected since its precursor DMSP was previously identified in the leaves of this seagrass [43]. DMSP has been proposed to exhibit physiological roles (e.g. as an intracellular osmolyte and antioxidant), and also serves as a chemoattractant [26,44]. DMS (mediated by both bacterial and algal DMSP lyases) has been generated in oceans at remarkably high amounts (>107 tons per year) and is a key component of the ocean sulfur cycle and has a global role in atmosphere-ocean feedback processes [45,46]. Chlorophyceae, especially Ulva, Enteromorpha, and Codium, and red alga, Polysiphonia, are capable of producing large amounts of DMS in contrast to Phaeophyceae (brown algae) which produce little [47,48] that is in agreement with present research (Table 1). Namely, DMS percentage was the highest in P. oceanica, than in green alga (F. petiolata) and the lowest in brown algae (Cystoseira sp. and D. linearis). Besides higher alkanes pentadecane (14.5%) and heptadecane (7.2%) that were abundant in F. petiolata and Cystoseira spp. (Table 1), nonadecane was also found (6.3%).

Dictyota dichotoma (Hudson) J. V. Lamouroux headspace VOCs

The genus Dictyota is represented by more than 40 species, thus being the richest genus of the family Dictyotaceae. In the Mediterranean in general, only the species Dictyota dichotoma and Dictyota linearis are found [49]. Brown algae of the family Dictyotaceae produce a significant number of secondary metabolites, especially diterpenes with three types of carbon skeletons: xenicanes; dolabellanes and ‘‘extended sesquiterpenes”. Many members of the family produce cyclic diterpenes, unique in the structural variety of marine natural products [49]. Terpenoid metabolites were identified in crude extract of D. dichotoma [49], e.g.: bicyclosesquiphellandrene, germacrene D, dictyoxide, pachydictyol A, isopachydictyol A, axenol, acetyldictyolal, dictyol B acetate, isopachydictyolal, 10-acetoxy-18-hydroxy-2,7-dolabelladiene, dictyol E, fucosterol, acetoxycrenulide, hydroxyacetyldictyolal, isodictyohemiacetal, dictyol A, dictyol B, dictyol C, and hydroxycrenulide. In addition, D. linearis contains different natural compounds [49] such as: dictyoxide, isopachydictyol A, pachydictyol A, dictyodial, 18-hydroxy-2,7-dolabelladiene, neodictyolactone, acetylsanadaol, acetyldictyolal, 10-acetoxy-18-hydroxy-2,7-dolabelladiene, dictyol E, 10,18-dihydroxy-2,7-dolabelladiene and dictyol C. There is no report on D. dichotoma headspace VOCs.

In distinction form other investigated samples, the major identified headspace VOCs of D. dichotoma were sesquiterpenes. Predominant sesquiterpene was germacrene D (28.3%; Table 1) that was previously found in D. dichotoma [49]. Bicyclogermacrene (4.7%) was also present. Sesquiterpenoids bicyclosesquiphellandrene, germacrene D and axenol were previously isolated from Taonia atomaria [50]. In this research, an array of sesquiterpenes biosynthetically connected with germacrene D were found (Table 1). They probably derived from protonation of germacrene D resulting in cadinenyl, muurolenyl, and amorphenyl cations that further react to form final products. Cadinenyl type sesquiterpenes were abundant: δ-cadinene (8.3%), γ-cadinene (3.4%), β-cadinene (2.8%), and trans-cadina-1,4-diene (1.2%). Epizonarene was found with 4.3% and it can be formed directly from germacrene D or from other intermediate cadinenes. It has been proposed that cyclization of germacrene D or of its endocyclic double bond isomer could result in the formation of bourbonene and copaene skeletons. β-Bourbonene was identified with (5.1%) and α-copaene with lower percentage. As a consequence of the thermodynamically unfavourable process to cis-decalin systems, the muurolenes and amorphenes are usually formed in smaller quantities that is in agreement with current research since they were found with lower percentages (α-muurolene (2.2%), γ-muurolene (2.1%), and α-amorphene (3.5%)).

Possible biosynthetic origin of identified VOCs

The formation of DMS [1] results from an enzymatic decomposition of dimethyl-β-propiothetin, a metabolite of methionine that is fairly widespread in marine plants. Formed dimethylsulfoniopropionate (DMSP), a tertiary sulfonium compound involved in osmoregulation in algae, is the precursor of DMS. Recently, the algal enzyme responsible for formation of DMS from DMSP has been identified and characterized in algae Emiliania huxleyi [26].

Marine macroalgae exhibit a high ability to fix halide ions and form a variety of halogenated secondary metabolites [1]. In present research only tribromomethane and 1-iodopentane were found. A halogenating enzyme, haloperoxidase [29] is considered to participate in their synthesis in the presence of halides and hydrogen peroxide. Among them bromoperoxidases were detected in seaweeds (e.g. from Corallina pilulifera or Ascophyllum nodosum). The enzyme produces CHBr3 by its reaction with ketoacids, halide ions and hydrogen peroxide [51].

Both in terrestrial and aquatic ecosystems, VOCs of primary producers are usually dominated by lipid degradation products, and the overall mechanism for their enzymatic release is identical as in terrestrial plants [11] and algae [13]. The enzyme cascade is initiated by activated phospholipase, followed by lipoxygenase and hydroperoxide lyase, which leads to VOCs liberation. However, the particular enzymes are highly species- and sometimes even strain-specific [13] that can explain large biodiversity of volatile lipid degradation products (e.g. carbonyl compounds, alcohols, hydrocarbons). Marine algae contain C20, C22 and C18 unsaturated fatty acids, and they can produce both plant (C18) and animal type (C20 and C22) fatty acid hydroperoxides. Short-chain aldehydes (e.g. C6, C9) and middle-chain aldehydes (e.g. C10) that were particularly present in H. filicina and F. petiolata are mainly formed from fatty acids (C20) in marine algae (via hydroperoxides), whereas they are formed from C18 fatty acids in higher plants [52,53]. For example, the formation of hexanal is proposed via linoleic acid cascade and arachidonic acid cascade through their hydroperoxides as intermediates by the lipoxygenase/fatty acid hydroperoxide lyase pathway. It could also be provided by oxidation from other polyunsaturated fatty acids, as well as heptanal [54]. Lower aliphatic alcohols may be formed by decomposition of secondary hydroperoxides of fatty acids by the reduction of the corresponding aldehydes [55]. Octanal and nonanal could originate from ω9 mono-unsaturated fatty acids (MUFAs) and also from ω6 PUFAs such as linoleic acid [55]. Following this general concept of lipid peroxidation, and subsequent oxidative cleavage of the carbon skeleton, the biosynthesis of C11- and C8-hydrocarbons could start from a single precursor (e.g. eicosapentaenoic acid). The polyunsaturated fatty acid substrate could be activated [56] either by 9-lipoxygenase or by 12-lipoxygenase, and resulting 9- or 12-hydroperoxides that cleave oxidatively to produce characteristic C11- and C8-hydrocarbons (e.g. fucoserratene). However, the origin of 3-methylbutanal is well known. It is obtained from amino acids, and more particularly from leucine, during Maillard reactions by Strecker degradation [16].

Terpenoids are enzymatically synthesized from acetyl-CoA (mevalonate (MVA) pathway) and/or pyruvate (deoxyxylulose-5-phosphate (DXP) pathway) via precursor 2-isopentenyl pyrophosphate (2-IPP) and its isomer 3-isopentenyl pyrophosphate (3-IPP). Regular monoterpenes derived from geranyl pyrophosphate (GPP), but are not widespread among marine algae. More often are sesquiterpenes derived metabolically from 300 distinct C15 -hydrocarbon skeletons, which in turn are produced from farnesyl diphosphate (FPP) by the action of sesquiterpene synthases [57]. The structure elucidation, biosynthesis, and biological activity of marine carbotricyclic sesquiterpene compounds were reviewed [10].

Carotenoid cleavage dioxygenases (CCDs) catalyze oxidative cleavage of carotenoids, resulting in the production of norisoprenoids—apocarotenoids [58] via glycosylation and breakdown of stored glycosides by glycosidase enzymes. CCDs often exhibit substrate promiscuity, which probably contributes to their natural diversity. Non-enzymatic reactions involving one or several steps of carotenoid degradation, stimulated by light, oxygen, temperature and acid hydrolysis can also occur [59]. The breakdown products of carotenoids found in this research belong to C13-norisoprenoids with megastigmane structure. In addition, C11-non-isoprenoids were identified. Dictyopterene A ((1R,2S)-cis-1-vinyl-2-(trans-1’-hexenyl)cyclopropane) and dictyoterpene B ((1R,2S)-cis-1-vinyl-2-(trans-1,cis-3-hexadienyl)cyclopropane) are proposed precursors [1] of dictyopterene C (6-butylcyclohepta-1,4-diene) and dictyopterene D (6-[(1Z)-but-1-enyl]cyclohepta-1,4-diene). Proposed biogenesis of dictyopterenes starts from (3S)-1,cis-5-undecadien-3-ol and (3S)-1,cis-5,cis-8-undecatrien-3-ol by dehydration and cyclization.

Benzenoid and phenylpropanoid volatile compounds, primarily derived from phenylalanine require shortening of the carbon skeleton side chain by a C2-unit, which can potentially occur via either the β-oxidative pathway or non-oxidatively [60].

Conclusions

Considering limited data available on the chemical composition of marine plants from the Adriatic Sea, the present research is contribution toward their better chemical characterization. Significant differences were found among the headspace VOCs from 3 seaweeds and 1 seagrass. High abundance of DMS was found in P. oceanica followed by F. petiolata and H. filicina indicating those plants as source of sulfur compounds in marine ecosystem. Their headspace contained individually variety of C8-compounds (e.g. fucoserratene), benzaldehyde, alkanes C15, C17 and C19, dictyopterene D and C, others. Sesquiterpenes were found in D. dichotoma, predominantly germacrene D indicating similarity to terrestrial aromatic plants. Identified VOCs contain different types of organic compounds that may be significant for chemosystematics and ecology (chemical communications in marine ecosystems).

Data Availability

All relevant data are within the paper. Data set necessary to replicate our research is already provided in the experimental part. There are no ethical or legal restrictions on sharing data sets.

Funding Statement

This research was supported by the Croatian Science Foundation under the project HRZZ-IP-11-2013-8547 (Research of Natural Products and Flavours: Chemical Fingerprinting and Unlocking the Potential) to Igor Jerković and by the Croatian Government and the European Union through the European Regional Development Fund - the Competitiveness and Cohesion Operational Programme (KK.01.1.1.01) – The Scientific Centre of Excellence for Marine Bioprospecting -BioProCro to Rozelinda Čož Rakovac.

References

  • 1.Moore RE. Volatile compounds from marine algae. Acc Chem Res. 1997; 10: 40–47. 10.1021/ar50110a002 [DOI] [Google Scholar]
  • 2.Sugisawa H, Nakamura K, Tamura H. The aroma profile of the volatiles in marine green algae Ulva pertusa. Food Rev Int. 1990; 6: 573–589. 10.1080/87559129009540893 [DOI] [Google Scholar]
  • 3.Milkova T, Talev G, Christov R, Dimitrova-Konaklieva S, Popov S. Sterols and volatiles in Cystoseira barbata and Cystoseira crinita from the Black sea. Phytochem. 1997; 45: 93–95. 10.1016/S0031-9422(96)00588-2 [DOI] [Google Scholar]
  • 4.Kajiwara T, Hatanaka A, Kawai T, Ishihara M, Tsuneya T. Study of flavor compounds of essential oil extracts from edible Japanese kelps. J Food Sci. 1988; 53: 960–962. 10.1111/j.1365-2621.1988.tb08995.x [DOI] [Google Scholar]
  • 5.Paul C, Pohnert G. Production and role of volatile halogenated compounds from marine algae. Nat Prod Rep. 2011; 28: 186–195. 10.1039/c0np00043d [DOI] [PubMed] [Google Scholar]
  • 6.Yamamoto M, Baldermann S, Yoshikawa K, Fujita A, Mase N, Watanabe N. Determination of volatile compounds in four commercial samples of Japanese green algae using solid phase microextraction gas chromatography mass spectrometry. Sci World J. 2014; 2014: Article ID 289780. 10.1155/2014/289780 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 7.Zhou L, Chen J, Xu J, Li Y, Zhou C, Yan X. Change of volatile components in six microalgae with different growth phases. J Sci Food Agric. 2017; 97: 761–769. 10.1002/jsfa.7794 [DOI] [PubMed] [Google Scholar]
  • 8.Santos A B, Vieira K R, Nogara G P, Wagner R, Jacob-Lopes E, Zepka L Q. Biogeneration of volatile organic compounds by microalgae: occurrence, behavior, ecological implications and industrial applications In book: Moore J P. Volatile organic compounds: occurrence, behavior and ecological implications, New York: Nova Science Publishers; 2016; 1–24. [Google Scholar]
  • 9.Achyuthan K E, Harper J C, Manginell R P, Moorman M W. Volatile metabolites emission by in vivo microalgae—an overlooked opportunity? Metabolites 2017; 7: 39; 10.3390/metabo7030039 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 10.Le Bideau F, Kousara M, Chen L, Wei L, Dumas F. Tricyclic sesquiterpenes from marine origin. Chem Rev. 2017; 117: 6110–6159. 10.1021/acs.chemrev.6b00502 [DOI] [PubMed] [Google Scholar]
  • 11.Hatanaka A. The biogeneration of green odor by green leaves. Phytochem. 1993; 34: 1201–1218. 10.1016/0031-9422(91)80003-J [DOI] [Google Scholar]
  • 12.Schwab W, Davidovich-Rikanati R, Lewinsohn E, Biosynthesis of plant-derived flavor compounds, Plant J. 2008; 54: 712–732. 10.1111/j.1365-313X.2008.03446.x [DOI] [PubMed] [Google Scholar]
  • 13.Pohnert G, Boland W. The oxylipin chemistry of attraction and defense in brown algae and diatoms. Nat Prod Rep. 2002; 19:108–122. 10.1039/A806888G [DOI] [PubMed] [Google Scholar]
  • 14.Fink P, Ecological functions of volatile organic compounds in aquatic systems, Mar Freshw Behav Physiol. 2007; 40: 155–168. 10.1080/10236240701602218 [DOI] [Google Scholar]
  • 15.Gally A, Yiannovits N, Poulos C, The aroma volatiles from Cystoseira stricta var. amentacea. J Essent Oil Res. 1993; 5: 27–32. 10.1080/10412905.1993.9698166 [DOI] [Google Scholar]
  • 16.Le Pape M-A, Grua-Priol J, Prost C, Demaimay M. Optimization of dynamic headspace extraction of the edible red algae Palmaria palmata and identification of the volatile components. J Agric Food Chem. 2004; 52: 550–556. 10.1021/jf030478x [DOI] [PubMed] [Google Scholar]
  • 17.Kamenarska Z, Gasic MJ, Zlatovic M, Rasovic A, Sladic D, Kljajic Z, et al. Chemical composition of the brown alga Padina pavonia (L.) Gaill. from the Adriatic Sea, Bot Mar. 2002; 45: 339–345. 10.1515/BOT.2002.034 [DOI] [Google Scholar]
  • 18.Nizamuddin M. Observations on the genus Flabellia (Caulerpales, Chlorophyta). Nova Hedwigia 1987; 44:175–188. [Google Scholar]
  • 19.Gallardo T, Gőmez Garreta A, Ribera MA, Cormaci M, Furnari G, Giaccone G et al. Check-list of Mediterranean seaweeds. II. Chlorophyceae Wille s. l. Bot Mar 1993; 36: 399–422. 10.1515/botm.1993.36.5.399 [DOI] [Google Scholar]
  • 20.Personnic S, Boudouresque CF, Astruch P, Ballesteros E, Blouet S, Bellan-Santini D et al. An ecosystem-based approach to assess the status of a Mediterranean ecosystem, the Posidonia oceanica seagrass meadow. PloS One 2014; 9:e98994 10.1371/journal.pone.0098994 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 21.Jerković I, Kranjac M, Marijanović Z, Zekić M, Radonić A, Tuberoso CIG. Screening of Satureja subspicata Vis. honey by HPLC-DAD, GC-FID/MS and UV/VIS: prephenate derivatives as biomarkers. Molecules 2016; 21: 377 10.3390/molecules21030377 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 22.National Institute of Standards and Technology (NIST) Chemistry WebBook, NIST Standard Reference Database Number 69. http://webbook.nist.gov/chemistry/
  • 23.Alghazeer R, Whida F, Abduelrhman E, Gammoudi F, Azwai S. Screening of antibacterial activity in marine green, red and brown macroalgae from the western coast of Libya. Natural Science 2013; 5: 7–14. 10.4236/ns.2013.51002 [DOI] [Google Scholar]
  • 24.Amico V, Oriente G, Piattelli M, Tringali C, Fattorusso E, Magno S, et al. Amino acids, sugars and sterols of some Mediterranean brown algae. Biochem Syst Ecol. 1976; 4: 143–146. 10.1016/0305-1978(76)90028-4 [DOI] [Google Scholar]
  • 25.Seymour JR, Simó R, Ahmed T, Stocker R. Chemoattraction to dimethylsulfoniopropionate throughout the marine microbial food web. Science 2010; 329: 342–345. 10.1126/science.1188418 [DOI] [PubMed] [Google Scholar]
  • 26.Alcolombri U, Ben-Dor S, Feldmesser E, Levin Y, Tawfik DS, Vardi A. Identification of the algal dimethyl sulfide–releasing enzyme: A missing link in the marine sulfur cycle, Science 2015; 348: 1466–1469. 10.1126/science.aab1586 [DOI] [PubMed] [Google Scholar]
  • 27.Newman KA, Gschwend PM. A method for quantitative determination of volatile organic compounds in marine macroalgae. Limnol Oceanogr. 1987; 32: 702–708. 10.4319/lo.1987.32.3.0702 [DOI] [Google Scholar]
  • 28.Houriet R Gaumann T. The chemical ionization of organic compounds. 2nd communication. Linear alkyl halides. Helv Chim Acta 1976; 59: 119–124. 10.1002/hlca.19760590111 [DOI] [Google Scholar]
  • 29.Franssen MCR. Halogenation and oxidation reactions with haloperoxidases. Biocatalysis 1994; 10: 87–111. 10.1016/0920-5861(94)80117-7 [DOI] [Google Scholar]
  • 30.Itoh N, Tsujita M, Ando T, Hisatomi G, Higashi T. Formation and emission of monohalomethanes from marine algae. Phytochem. 1997; 45: 67–73. 10.1016/S0031-9422(96)00786-8 [DOI] [Google Scholar]
  • 31.Boland W. The chemistry of gamete attraction: chemical structure, biosynthesis, and (a)biotic degradation of algal pheromones. Proc Natl Acad Sci USA 1995; 92: 37–43. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 32.Mtiller DG, Gassmann G. Sexual reproduction and the role of sperm attractants in monoecious species of the brown algae order fucales (fucus, hesperophycus, pelvetia, and pelvetiopsis). J Plant Physiol 1985; 118: 401–408. 10.1016/S0176-1617(85)80200-5 [DOI] [PubMed] [Google Scholar]
  • 33.Mironov OG, Shchekaturina TL, Tsimbal IM. Saturated hydrocarbons in marine organisms. Mar Ecol Prog Ser. 1981; 5: 303–309. [Google Scholar]
  • 34.Silva G, Pereira RB, Valentão P, Andrade PB, Sousa C. Distinct fatty acid profile of ten brown macroalgae. Braz J Pharmacog. 2013; 23: 608–613. 10.1590/S0102-695X2013005000048 [DOI] [Google Scholar]
  • 35.Andrade PB, Barbosa M, Matos RP, Lopes G, Vinholes J, Mouga T, et al. Valuable compounds in macroalgae extracts. Food Chem. 2013; 138: 1819–1828. 10.1016/j.foodchem.2012.11.081 [DOI] [PubMed] [Google Scholar]
  • 36.Gnavi G, Garzoli L, Poli A, Prigione V, Burgaud G, Varese GC. The culturable mycobiota of Flabellia petiolata: First survey of marine fungi associated to a Mediterranean green alga, PLOS ONE 2017; 12(4): e0175941 10.1371/journal.pone.0175941 April 20, 2017 1–20 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 37.Ballesteros E, Martin D, Uriz MJ. Biological-activity of extracts from some Mediterranean macrophytes. Bot Mar. 1992; 35: 481–485. 10.1515/botm.1992.35.6.481 [DOI] [Google Scholar]
  • 38.Bílgín F, Ertan ÖO. Selected chemical constituents and their seasonal variations in Flabellia petiolata (Turra) Nizam. and Halimeda tuna (Ellis & Sol.) J. V. Lamour. in the gulf of Antalya (North-eastern Mediterranean). Turk J Bot. 2002; 26: 87–90. [Google Scholar]
  • 39.Bates TS, Cline JD. The role of the ocean in a regional sulfur cycle. J Geophys Res. 1985; 90: 9168–9172. 10.1029/JC090iC05p09168 [DOI] [Google Scholar]
  • 40.Sun S-M, Chung G-H, Shin T-S. Volatile compounds of the green alga, Capsosiphon fulvescens, J Appl Phycol. 2012; 24: 1003–1013. 10.1007/s10811-011-9724-x [DOI] [Google Scholar]
  • 41.Haznedaroglu MZ, Zeybec U. HPLC determination of chicoric acid in leaves of Posidonia oceanica, Pharm Biol. 2007; 45: 745–748. 10.1080/13880200701585717 [DOI] [Google Scholar]
  • 42.Hammami S, Salem AB, Ashour ML, Cheriaa L, Graziano G, Mighri Z. A novel methylated sesquiterpene from seagrass Posidonia oceanica (L.) Delile, Nat Prod Res. 2012; 14: 1265–1270. 10.1080/14786419.2012.725401 [DOI] [PubMed] [Google Scholar]
  • 43.Borges AV, Champenois W. Seasonal and spatial variability of dimethylsulfoniopropionate (DMSP) in the Mediterranean seagrass Posidonia oceanica. Aquat Bot. 2015; 125: 72–79. 10.1016/j.aquabot.2015.05.008 [DOI] [Google Scholar]
  • 44.Sunda W, Kieber DJ, Kiene RP, Huntsman S. An antioxidant function for DMSP and DMS in marine algae. Nature 2002; 418: 317–320. 10.1038/nature00851 [DOI] [PubMed] [Google Scholar]
  • 45.Charlson RJ, Lovelock JE, Andreae MO, Warren SG. Oceanic phytoplankton, atmospheric sulphur, cloud albedo and climate. Nature 1987; 326: 655–661. 10.1038/326655a0 [DOI] [Google Scholar]
  • 46.Quinn PK, Bates TS. The case against climate regulation via oceanic phytoplankton sulphur emissions. Nature 2011; 480: 51–56. 10.1038/nature10580 [DOI] [PubMed] [Google Scholar]
  • 47.White RHJ. Analysis of dimethyl sulfonium compounds in marine algae. Mar Res. 1982; 40: 529–36. [Google Scholar]
  • 48.Reed RH. Measurement and osmotic significance of ß–dimethyl–sulphoniumpropionate in marine macroalgae. Mar Biol Letts. 1983; 4: 173–81. [Google Scholar]
  • 49.Siamopoulou P, Bimplakis A, Iliopoulou D, Vagias C, Cos P, Berghe DV et al. Diterpenes from the brown algae Dictyota dichotoma and Dictyota linearis. Phytochem. 2004; 65: 2025–2030. 10.1016/j.phytochem.2004.06.018 [DOI] [PubMed] [Google Scholar]
  • 50.De Rosa S, De Giulio A, Iodice C, Zavodink N. Sesquiterpenes from the brown alga Taonia atomaria. Phytochem. 1994; 37: 1327–1330. 10.1016/S0031-9422(00)90407-2 [DOI] [Google Scholar]
  • 51.Ohshiro T, Nakano S, Takahashi Y, Suzuki M, Izumi Y. Occurrence of bromoperoxidase in the marine green macro-alga, bulvella lens, and emission of volatile brominated methane by the enzyme. Phytochem. 1999; 52: 1211–1215. 10.1016/S0031-9422(99)00404-5 [DOI] [Google Scholar]
  • 52.Boonprab K, Matsui K, Akakabe Y, Yoshida M, Yotsukura N, Chirapart A et al. Formation of aldehyde flavour (n-hexanal, 3Z-nonenal and 2E-nonenal) in the brown alga, Laminaria angustata. J Appl Phycol 2006; 18: 1573–1576. 10.1007/s10811-006-9038-6 [DOI] [Google Scholar]
  • 53.Akakabe Y, Matsui K, Kajiwara T. 2,4-Decadienals are produced via (R)-11-HPITE from arachidonic acid in marine green alga Ulva conglobata. Bioorg Med Chem. 2003; 11: 3607–3609. 10.1016/S0968-0896(03)00364-X [DOI] [PubMed] [Google Scholar]
  • 54.Enoiu M, Wellman M, Leroy P, Ziegler J-M, Mistrea N, Siest G. Gas and liquid chromatography—mass spectrometry of aldehydic products from lipid peroxidation. Analysis 2000; 28: 285–290. 10.1051/analusis:2000280285 [DOI] [Google Scholar]
  • 55.Tanchotikul U, Hsieh TC-Y. Volatile flavor components in crayfish waste. J. Food Sci. 1989; 54: 1515–1520. 10.1111/j.1365-2621.1989.tb05149.x [DOI] [Google Scholar]
  • 56.Hombeck M, Boland W. Biosynthesis of the algal pheromone fucoserratene by the fresh- water diatom Asterionella formosa (Bacillariophyceae). Tetrahedron 1998; 54: 11033–11042. 10.1016/S0040-4020(98)00660-7 [DOI] [Google Scholar]
  • 57.Miller DJ, Allemann RK. Sesquiterpene synthases: passive catalysts or active players? Nat Prod Rep. 2012; 29: 60–71. 10.1039/c1np00060h [DOI] [PubMed] [Google Scholar]
  • 58.Schmidt H, Kurtzer R, Eisenreich W, Schwab W. The carotenase AtCCD1 from Arabidopsis thaliana is a dioxygenase. J Biol Chem. 2006; 281: 9845–9851. 10.1074/jbc.M511668200 [DOI] [PubMed] [Google Scholar]
  • 59.Winterhalter P, Rouseff R. in Carotenoid-derived aroma compounds, eds. Winterhalter P. and Rouse ff R., ACS Symposium Series, American Chemical Society, Washington, DC, 2001: 1–17. [Google Scholar]
  • 60.Boatright J, Negre F, Chen X, Kish CM, Wood B, Peel G et al. N. Understanding in vivo benzenoid metabolism in petunia petal tissue. Plant Physiol. 2004; 135: 1993–2011. 10.1104/pp.104.045468 [DOI] [PMC free article] [PubMed] [Google Scholar]

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Data Availability Statement

All relevant data are within the paper. Data set necessary to replicate our research is already provided in the experimental part. There are no ethical or legal restrictions on sharing data sets.

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