TY - JOUR
AU - Amado-Filho, Gilberto Menezes
AB - Abstract We investigated the organelles involved in the biosynthesis of fatty acid (FA) derivatives in the cortical cells of Laurencia translucida (Rhodophyta) and the effect of these compounds as antifouling (AF) agents. A bluish autofluorescence (with emission at 500 nm) within L. translucida cortical cells was observed above the thallus surface via laser scanning confocal microscopy (LSCM). A hexanic extract (HE) from L. translucida was split into two isolated fractions called hydrocarbon (HC) and lipid (LI), which were subjected to HPLC coupled to a fluorescence detector, and the same autofluorescence pattern as observed by LSCM analyses (emission at 500 nm) was revealed in the LI fraction. These fractions were analyzed by gas chromatography–mass spectrometry (GC-MS), which revealed that docosane is the primary constituent of HC, and hexadecanoic acid and cholesterol trimethylsilyl ether are the primary components of LI. Nile red (NR) labeling (lipid fluorochrome) presented a similar cellular localization to that of the autofluorescent molecules. Transmission and scanning electron microscopy (TEM and SEM) revealed vesicle transport processes involving small electron-lucent vesicles, from vacuoles to the inner cell wall. Both fractions (HC and LI) inhibited micro-fouling [HC, lower minimum inhibitory concentration (MIC) values of 0.1 µg ml–1; LI, lower MIC value of 10 µg ml–1]. The results suggested that L. translucida cortical cells can produce FA derivatives (e.g. HCs and FAs) and secrete them to the thallus surface, providing a unique and novel protective mechanism against microfouling colonization in red algae. Introduction Acyclic hydrocarbons (HCs) that are produced by algae can be categorized into two broadly defined subclasses according to their structure and presumed routes of biosynthesis, namely linear n-alkanes/n-alkenes, which have varying degrees of unsaturation/methylation, and the highly branched isoprenoids (Volkman 2006). These substances serve many functions in algae (Eltgroth et al. 2005, Dahmen and Leblond 2013), but their roles as antifouling (AF) agents in marine macroalgae have not yet been described. Studying the localization, storage and exudation mechanisms of biologically active AF metabolites in macroalgae is necessary to enrich our understanding of the ecological significance of surface-active bioinhibitors (Salgado et al. 2008, Paradas et al. 2015). In addition, numerous studies have shown the activity of macroalgal lipids against fouling organisms (Bazes et al. 2009, Plouguerné et al. 2010), and little is now known about processes related to storage and transport to the macroalgal thallus surface of fatty acid (FA) derivatives (e.g. HCs). With regards to lipids with proposed ecological activity, linear n-alkanes and n-alkenes are under consideration (Volkman 2006). These lipids are derived from the decarboxylation of FAs that are associated with acyl glycerolipids (Dahmen and Leblond 2013). For example, the C21 alkene 3,6,9,12,15,18-heneicosahexaene, which is one of the most common linear alkenes observed in HC-producing microalgae, is thought to be derived from docosahexaenoic acid [22:6(n-3)] (Volkman 2006). The unicellular green alga Pseudochoricystis ellipsoidea (Chlorophyta) (Satoh et al. 2010) and the colonial green alga Botryococcus braunii race A (Chlorophyta) (Yoshida et al. 2012) accumulate fatty HCs. According to Yoshida et al. (2012), B. braunii races B and L produce terpenoid-derived HCs. Haptophytes also produce long-chain ketones, which are known as ‘alkenones’ (Laws et al. 2001, Eltgroth et al. 2005). Eltgroth et al. (2005) used cellular fractionation and microscopy techniques to show that polyunsaturated lipids (long-chain C37–C39 alkenones, alkenoates and alkenes) are produced first in chloroplasts in the cells of the microalgae Isochrysis galbana (Haptophyta), and are then exported to the cytosol, where they are internalized again into cytoplasmic vesicles or lipid bodies (Eltgroth et al. 2005). In another study, Dahmen and Leblond (2013) demonstrated the cellular localization of lipids in the cells of Pyrocystis lunula (Dinophyta), in which the major characterized metabolites were polyunsaturated C27 HCs. However, the function(s) of linear long-chain HCs in algae is currently unclear; however, it has been hypothesized to be linked to buoyancy, energy storage or signaling (Sinninghe Damsté et al. 2000, Eltgroth et al. 2005). In red algae, biochemical data have indicated that FA derivatives are first biosynthesized in chloroplasts and in the endoplasmic reticulum (ER) by prokaryotic and eukaryotic pathways, respectively (Sato and Moriyama 2007). For example, Khozin-Goldberg and Cohen (2011) showed that in Porphyridium cruentum (Rhodophyta), the triacylglycerols contribute to the synthesis, of monogalactosyl diacylglycerol (MGDG) species through the eukaryotic pathway; these species are chloroplast membrane lipids. In addition, the genomic characterization of P. cruentum has revealed genes that are involved in glycerolipid biosynthesis (Bhattacharya et al. 2013), and studies that address FA derivative biosynthesis in red algae are also scarce (Sato and Moriyama 2007, Khozin-Goldberg and Cohen 2011). Some studies have revealed that FA derivatives from seaweeds have shown strong AF activity (Bazes et al. 2009, Plouguerné et al. 2010, Iyapparaj et al. 2014). FAs (C16–C24) from the seagrasses Syringodium isoetifolium (Magnoliophyta) and Cymodocea serrulata (Magnoliophyta), which were assayed for their antimicrobiotic effects, presented minimal inhibitory concentrations (MICs) ranging from 1.0 to 10 µg.ml–1 against fouling microalgae and bacteria (Iyapparaj et al. 2014). With regards to the Laurencia genus (Rhodophyta), many species are prolific producers of terpenoid and non-terpenoid secondary metabolites (Fujii et al. 2011). This genus has been reported worldwide in both the temperate and tropical oceans of the world, occurring from the intertidal to the subtidal zones up to 65 m in depth (Guiry and Guiry 2010). These metabolites play important ecological roles, but many biotechnological applications have also been described for them, e.g. AF paint (Da Gama et al. 2003). However, of the 12 species that belong to the Laurencia complex in Brazil, only six have been chemically investigated (Fujii et al. 2011). Taxonomically, some Laurencia species are differentiated from other members of the Laurencia complex by the presence of refractive intracellular organelles called corps en cerise (CC) (Feldmann and Feldmann 1950). Concerning cell biology studies, Laurencia dendroidea (Rhodophyta) was shown to store and transport halogenated metabolites from the CC to the thallus surface in response to epiphytic bacteria through vesicular traffic (Salgado et al. 2008, Paradas et al. 2010). Reis et al. (2013) showed that both microfilaments and microtubules have a central role in this L. dendroidea defensive system. In addition, the genomic information for this species has been advanced as a consequence of broad transcriptomic analysis, which unveiled many genes that are responsible for the synthesis of terpenoid compounds (Oliveira et al. 2012, Oliveira et al. 2015). In contrast to L. dendroidea, the endemic Brazilian species Laurencia translucida M.T. Fujii & Cordeiro-Marino (Rhodophyta) does not possess CC, and its outermost cortical cells are small and translucent (Fujii and Cordeiro-Marino 1996). Previous chemical studies on L. translucida hexanic extract (HE) revealed an AF property against the fouling mussel Perna perna (Da Gama et al. 2008) and also high cytotoxic activity against tumor cell lines (Stein et al. 2011). More recently, it was suggested that one possible role for L. translucida translucent cells is the storage of chemical defenses (Fujii et al. 2012). However, neither the metabolites involved in ecological interactions nor microscopy and ultrastructural characterization were performed on this algal species. Thus, the aims of the present work were: (i) to describe the cortical cells of L. translucida using bright field optical microscopy, laser scanning confocal microscopy (LSCM), transmission electron microscopy (TEM) and scanning electron microscopy (SEM); (ii) to localize bioactive (AF) secondary metabolites inside cortical cells through fluorescence spectral scanning; (iii) to characterize the lipid composition of L. translucida by using gas chromatography–mass spectrometry (GC-MS), HPLC and nuclear magnetic resonance (NMR); and (iv) to investigate the antimicrofouling activity of L. translucida in isolated metabolites, specifically against fouling microalgae and bacteria using in vitro bioassays. Results LSCM analyses Multispectral fluorescence analysis by LSCM revealed three different fluorescence emissions (Fig. 1A), with the first one ranging from 450 to 541 nm with an emissions peak at approximately 500 nm, the second one ranging from 550 to 614 nm with its highest emission at appropximately 580 nm and the third one ranging from 623 to 750 nm with its highest emission at approximately 690 nm. Blue autofluorescence was registered along the algal surface and within the cortical cells (Fig. 1B, E, F). The emissions wavelength in the blue range is related to an unidentified L. translucida metabolite. The second and third fluorescence emissions (Fig. 1C, D) correspond to the two primary red photosynthetic pigments in the alga, namely, phycobilins and Chl a, respectively. The co-localization of phycobilins and Chl a fluorescence emissions with the internal blue fluorescence confirms the presence of these metabolites within chloroplasts (Fig. 1E, F). Fig. 1 View largeDownload slide LSCM multispectral fluorescence analysis of L. translucida cortical and subcortical cells. (a) The fluorescence intensity spectrum as a function of the emission wavelength; arrows point to the three primary peaks observed as follows: (1) unidentified metabolite (UM), (2) phycobilins and (3) Chl a. (b–f) LCSM multispectral fluorescence images of L. translucida cortical and subcortical cells. (b) UM, the arrowheads show the unidentified metabolite within cortical cells; (c) phycobilins; (d) Chl a. (e, f) The combined autofluorescence images correspond to the distribution of UM (arrowheads) plus Chl a (arrows) (e) and UM (arrowheads) plus phycobilins (arrows) (f). Scale bars = 18 µm. Fig. 1 View largeDownload slide LSCM multispectral fluorescence analysis of L. translucida cortical and subcortical cells. (a) The fluorescence intensity spectrum as a function of the emission wavelength; arrows point to the three primary peaks observed as follows: (1) unidentified metabolite (UM), (2) phycobilins and (3) Chl a. (b–f) LCSM multispectral fluorescence images of L. translucida cortical and subcortical cells. (b) UM, the arrowheads show the unidentified metabolite within cortical cells; (c) phycobilins; (d) Chl a. (e, f) The combined autofluorescence images correspond to the distribution of UM (arrowheads) plus Chl a (arrows) (e) and UM (arrowheads) plus phycobilins (arrows) (f). Scale bars = 18 µm. During a detailed algal analysis, transversally sectioned, unbroken cortical cells presented the same autofluorescence emissions corresponding to the L. translucida metabolites, thus confirming the presence of these metabolites along the algal surface and associating them with chloroplasts (Fig. 2A–C). By performing deconvolution image processing, the Chl a autofluorescence distribution (Fig. 2) could be spatially distinguished from the bluish fluorescence location in L. translucida cortical cells (Fig. 2D–F). In this case, the bluish emission related to the L. translucida metabolites appeared specifically in vesicles that were closely associated with chloroplasts (Fig. 2E, F). Fig. 2 View largeDownload slide Laurencia translucida cortical cells in LSCM images processed by deconvolution. (a, b). The unidentified metabolite (UM; blue wavelength emission) and chloroplast (red wavelength emission) autofluorescence were detached from one another. The asterisks indicate cortical cells. Arrowheads indicate the UM distribution along the algal surface. (c) The combined autofluorescence images correspond to the distribution of UM and chloroplasts. Scale bar = 17 µm. (d, e) The UMs are distributed at the peripheral region of chloroplasts and in the cytoplasm. (f) The combined autofluorescence images correspond to the distribution of UMs and chloroplasts. The arrows point to chloroplasts. Arrowheads point to the UMs. Scale bar = 5 µm. Fig. 2 View largeDownload slide Laurencia translucida cortical cells in LSCM images processed by deconvolution. (a, b). The unidentified metabolite (UM; blue wavelength emission) and chloroplast (red wavelength emission) autofluorescence were detached from one another. The asterisks indicate cortical cells. Arrowheads indicate the UM distribution along the algal surface. (c) The combined autofluorescence images correspond to the distribution of UM and chloroplasts. Scale bar = 17 µm. (d, e) The UMs are distributed at the peripheral region of chloroplasts and in the cytoplasm. (f) The combined autofluorescence images correspond to the distribution of UMs and chloroplasts. The arrows point to chloroplasts. Arrowheads point to the UMs. Scale bar = 5 µm. NR staining The LSCM images of L. translucida Nile red (NR)-stained specimens revealed the presence of FA derivatives as a yellow emission found in chloroplasts and cytoplasmic vesicles along the thallus surface and inside the cell walls (Fig. 3A–C). In fixed L. translucida individuals (control), only Chl a autofluorescence was observed (which is indicated by the green color in Fig. 3D–F). Fig. 3 View largeDownload slide LCSM fluorescence images of NR staining in L. translucida cortical cells. (a, b) A longitudinal sections (NR) obtained by the analysis of the x- and y-axis where lipids (yellow fluorescence) can be observed inside vesicles, chloroplasts and along the thallus surface. Lipid fluorescence patterns (yellow) were also observed in the upper portion of cortical cells. The arrowhead points to the lipid fluorescence associated with chloroplasts. Arrows show the lipids in cytoplasmic vesicles. Asterisks indicate the presence of lipids above the cell wall. Scale bars = 4 µm. (c) A transversal section (NR) obtained by the analysis of the x- and z-axis where lipids can be observed (yellow) distributed along the surface of the algae, which is associated with the plasmatic membrane and the inside of the cell wall. Arrowheads point to lipids above the thallus surface. Arrows show lipids associated with the plasmatic membrane. Small arrowheads point to lipids inside the cell wall. Scale bar = 7.5 µm. (d, e) The same LSCM optical sections used for NR staining were programmed in samples without NR staining (control). LSCM images only show the Chl a autofluorescence that is artificially colored (green) inside the chloroplasts. Scale bars: a = 8 µm; b and c = 5 µm. Fig. 3 View largeDownload slide LCSM fluorescence images of NR staining in L. translucida cortical cells. (a, b) A longitudinal sections (NR) obtained by the analysis of the x- and y-axis where lipids (yellow fluorescence) can be observed inside vesicles, chloroplasts and along the thallus surface. Lipid fluorescence patterns (yellow) were also observed in the upper portion of cortical cells. The arrowhead points to the lipid fluorescence associated with chloroplasts. Arrows show the lipids in cytoplasmic vesicles. Asterisks indicate the presence of lipids above the cell wall. Scale bars = 4 µm. (c) A transversal section (NR) obtained by the analysis of the x- and z-axis where lipids can be observed (yellow) distributed along the surface of the algae, which is associated with the plasmatic membrane and the inside of the cell wall. Arrowheads point to lipids above the thallus surface. Arrows show lipids associated with the plasmatic membrane. Small arrowheads point to lipids inside the cell wall. Scale bar = 7.5 µm. (d, e) The same LSCM optical sections used for NR staining were programmed in samples without NR staining (control). LSCM images only show the Chl a autofluorescence that is artificially colored (green) inside the chloroplasts. Scale bars: a = 8 µm; b and c = 5 µm. TEM ultrastructural observations To understand how FA derivative biosynthesis occurs in L. translucida cortical cells, we performed conventional ultrastructural observations. TEM images from L. translucida cortical cells showed the presence of many subcellular structures such as nuclei, the Golgi apparatus and chloroplasts (Fig. 4A). Two types of vacuoles were observed inside the cytoplasm, with some possessing granulate electron-dense material (VE) and others displaying smaller translucent vesicles inside (V) (Figs. 4A, 5A, B). Both types of vacuoles (VE and V) were observed in association with regions in which smaller vesicles (translucent nanometric vesicles) were exudated to the cell walls (Fig. 4C, D). A large amount of osmiophilic material was also observed in large vacuoles and in small vesicles in the cytoplasm (Fig. 5A, D). Fig. 4 View largeDownload slide TEM images of a transverse section of L. translucida cortical cells. (a) A general view of a cortical cell showing electron-dense vesicles (VE) and electron-lucent vacuoles (V), chloroplasts (C), Golgi apparatus (GA), osmiophilic material (O), a nucleus (N) and cell walls (CW). Arrowheads indicate the peripheral distribution of VE. A white asterisk indicates a small osmiophilic vesicle in the cytoplasm. Scale bar = 2 µm. (b–d) A detail of the small vesicles being released to the cell wall. Arrowheads indicate the vesicles. Scale bars b = 0.3 µm; c = 0.5 µm and d = 0.2 µm. (e, f) Endoplasmatic reticulum (ER) in the periphery of the cell, near the cell wall. Black asterisks indicate vesicles budding from the ER. Scale bar = 0.2 µm. Fig. 4 View largeDownload slide TEM images of a transverse section of L. translucida cortical cells. (a) A general view of a cortical cell showing electron-dense vesicles (VE) and electron-lucent vacuoles (V), chloroplasts (C), Golgi apparatus (GA), osmiophilic material (O), a nucleus (N) and cell walls (CW). Arrowheads indicate the peripheral distribution of VE. A white asterisk indicates a small osmiophilic vesicle in the cytoplasm. Scale bar = 2 µm. (b–d) A detail of the small vesicles being released to the cell wall. Arrowheads indicate the vesicles. Scale bars b = 0.3 µm; c = 0.5 µm and d = 0.2 µm. (e, f) Endoplasmatic reticulum (ER) in the periphery of the cell, near the cell wall. Black asterisks indicate vesicles budding from the ER. Scale bar = 0.2 µm. Fig. 5 View largeDownload slide TEM images of a transverse section of L. translucida cortical cells. (a) Golgi apparatus (GA), electron-dense vacuoles (VE), electron-lucent vacuoles (V), mitochondria (M), osmiophilic material (O) and the nucleus (N); white asterisks indicate small osmiophilic vesicles in the cytoplasm. Scale bar = 10 µm. (b) Chloroplasts (C), O, V; a black asterisk indicates vesicle–vacuole fusion. Scale bar = 7 µm. (c) Arrowheads indicate vesicle exportation from the Golgi apparatus (GA) and dyctiosomes (D) and the asterisk shows the tubular continuous structure of the endoplasmatic reticulum (ER). Scale bar = 4 µm. (d) Asterisks indicates small osmiophilic vesicles that were observed in detail in the cytoplasm. Scale bar = 200 nm. Fig. 5 View largeDownload slide TEM images of a transverse section of L. translucida cortical cells. (a) Golgi apparatus (GA), electron-dense vacuoles (VE), electron-lucent vacuoles (V), mitochondria (M), osmiophilic material (O) and the nucleus (N); white asterisks indicate small osmiophilic vesicles in the cytoplasm. Scale bar = 10 µm. (b) Chloroplasts (C), O, V; a black asterisk indicates vesicle–vacuole fusion. Scale bar = 7 µm. (c) Arrowheads indicate vesicle exportation from the Golgi apparatus (GA) and dyctiosomes (D) and the asterisk shows the tubular continuous structure of the endoplasmatic reticulum (ER). Scale bar = 4 µm. (d) Asterisks indicates small osmiophilic vesicles that were observed in detail in the cytoplasm. Scale bar = 200 nm. Many Golgi apparatus were observed in L. translucida cortical cells. Each Golgi apparatus consisted of 7–8 tightly stacked cisternae (Fig. 5C). The polarity of the Golgi dictyosomes was determined by the presence of the vesicles coming out from the trans-Golgi network (Fig. 5C). There were numerous vesicles with increasing sizes budding from the trans-Golgi network (Fig. 5C); additionally, a vesicle–vacuole fusion was observed (Fig. 5B). ER was frequently observed in L. translucida cortical cells close to the nucleous, chloroplasts, cell wall and vacuoles (Figs. 4E, F, 5C). ER with budding vesicles was also observed, and they (vesicles) appear to be being transported to the cell wall at a later time (Fig. 4E, F). SEM ultrastructure observation SEM images showed cortical cell images with the cell walls removed and the plasmatic membrane exhibiting small vesicles over its surface (Fig. 6A, B). In addition, when the cell membrane was fractured, vacuoles with small vesicles could be observed inside (Fig. 6C, D). Fig. 6 View largeDownload slide SEM images of intracellular constituents from fractured L. translucida cortical cells. (a, b) The cell wall was removed and the plasmatic membrane with vesicles above can be observed. Arrowheads indicate vesicles. Scale bars: a = 4 µm; b = 2 µm. (c, d) The plasmatic membrane and the vacuole membrane were removed and many vacuoles are observed; notably, a huge amount of small vesicles can be observed inside the vacuoles, and arrowheads indicate vacuoles and vesicles. Scale bars: c = 2.5 µm; d = 2 µm. Fig. 6 View largeDownload slide SEM images of intracellular constituents from fractured L. translucida cortical cells. (a, b) The cell wall was removed and the plasmatic membrane with vesicles above can be observed. Arrowheads indicate vesicles. Scale bars: a = 4 µm; b = 2 µm. (c, d) The plasmatic membrane and the vacuole membrane were removed and many vacuoles are observed; notably, a huge amount of small vesicles can be observed inside the vacuoles, and arrowheads indicate vacuoles and vesicles. Scale bars: c = 2.5 µm; d = 2 µm. Chemical analyses Thin-layer chromatography (TLC) analyses of the L. translucida HE showed the major presence of HCs at 74 ± 3% and sterols (STs) at 6 ± 0.9% (Supplementary Fig. S1) in these extracts, and the minor compounds identified here were FAs at 0.5 ± 0.3%, triglycerides (TGs) at 0.5 ± 0.5%, monoglycerides (MGs) at 2 ± 0.2% and diglycerides (DGs) at 2 ± 0.3% (Supplementary Fig. S1). The 1H-NMR data thereby corroborated the TLC analyses of L. translucida HEs, once they revealed the presence of a complex mixture of FA derivatives in L. translucida HE, in which the aliphatic region of these substances was present at δ 0.86 p.p.m. and δ 1.70 p.p.m. in the 1H-NMR experiment because of strong overlapping resonance signals from hydrogens. The α-CH2 acyl group was observed at δ 2.32 (m). To investigate the autofluorescence properties of FA derivatives from L. translucida, the two isolated fractions, namely HC and LI (ST, FA, TG, MG and DG), were subjected to HPLC autofluorescence scanning analyses (excitation, 405 nm; emission, 500 nm). These analyses showed one prominent autofluorescent peak in the LI chromatogram with a retention time of 3.75 min (Fig. 7A) and a small peak in the HC chromatogram with a retention time of 3.25 min (Fig. 7B). No peak was observed in chromatograms of methanol alone (data not shown). The GC-MS analyses of the HC fraction indicated that docosane was the major compound in L. translucida HE (Supplementary Fig. S2; Supplementary Table S1). The L. translucida HC fraction composition order was as follows: docosane (61.66%) > octadecane (18.63%) > nonadecane (8.64%) > hexadecane (7.70%) > icosane (3.37%) (Supplementary Table S1). In relation to the LI fraction, the GC-MS analyses identified 20 FAs (Supplementary Table S2), and the three major FAs were ordered hexadecanoic acid, methyl ester (57.86%; saturated) > 9-octadecenoic acid (Z)-, methyl ester (12.14%; monounsaturated) > octadecanoic acid, methyl ester (10.08%; saturated) (Supplementary Table S2). The GC-MS analyses identified only one ST in the L. translucida LI fraction and that was the cholesterol known as trimethylsilyl ether (100%) (Supplementary Table S2). Fig. 7 View largeDownload slide HPLC autofluorescence scanning analyses (excitation 405 nm; emission 500 nm) chromatograms of isolated fractions of L. translucida (HC, hydrocarbons; LI, lipids). (a) LI, a characteristic peak with a retention time of 3.75 min. (b) HC, a small peak with a retention time of 3.25 min. Fig. 7 View largeDownload slide HPLC autofluorescence scanning analyses (excitation 405 nm; emission 500 nm) chromatograms of isolated fractions of L. translucida (HC, hydrocarbons; LI, lipids). (a) LI, a characteristic peak with a retention time of 3.75 min. (b) HC, a small peak with a retention time of 3.25 min. AF tests The L. translucida HE inhibited all the bacterial strains (Halomonas marina, Polaribacter irgensii, Pseudoalteromonas elyakovii, Shewanella putrefaciens and Vibrio aestuarianus) and microalgae species (Chlorarachnion reptans, Cylindotheca cloisterium, Exanthemachrysis gayraliae, Navicula jeffreyae and Chlorarachnion globosum) with a MIC of 50 µg ml–1 (Table 1). The L. translucida HC fraction inhibited P. elyakovii, V. aestuarianus, C. reptans and N. jeffreyae (Table 1) at a MIC value of 0.1 µg ml–1, and it inhibited H. marina, P. irgensii, S. putrefaciens, C. cloisterium, E. gayraliae and C. globosum at 1 µg ml–1 (Table 1). The L. translucida LI fraction was able to inhibit H. marina, P. irgensii, C. cloisterium and E. gayraliae at a MIC value of 10 µg ml–1 (Table 1). Table 1 Effects of L. translucida HE, HC and LI fractions that were tested at the MIC (µg ml–1) on sensitive strains of marine fouling bacteria and microalgae   Antibacterial activity (MIC)     Halomonas marina  Polaribacter irgensii  Pseudoalteromonas elyakovii  Shewanella putrefaciens  Vibrio aestuarianus  HE  50  50  50  50  50  HC  1  1  0.1  1  0.1  LI  10  10  50  >100  >100    Antibacterial activity (MIC)     Halomonas marina  Polaribacter irgensii  Pseudoalteromonas elyakovii  Shewanella putrefaciens  Vibrio aestuarianus  HE  50  50  50  50  50  HC  1  1  0.1  1  0.1  LI  10  10  50  >100  >100    Antimicroalgal activity (MIC)     Chlorarachnion reptans  Cylindrotheca closterium  Exanthemachrysis gayraliae  Navicula jeffreyae  Chlorarachnion globosum  HE  50  50  50  50  50  HC  0.1  1  1  0.1  1  LI  50  10  10  50  100    Antimicroalgal activity (MIC)     Chlorarachnion reptans  Cylindrotheca closterium  Exanthemachrysis gayraliae  Navicula jeffreyae  Chlorarachnion globosum  HE  50  50  50  50  50  HC  0.1  1  1  0.1  1  LI  50  10  10  50  100  View Large Discussion In the present work, the chemical analyses suggested that the major L. translucida HE metabolites are primarily made up of FA derivatives (HC, ST, FFA, TG, MG and DG), and HCs account for more than half of these compounds. In addition, the presence of a fluorescent LI fraction (ST, FA, TG, MG and DG) was also determined by HPLC, with an emission wavelength at 500 nm. The same pattern of fluorescence emission (500 nm) found in the LI fraction was also found in L. translucida cortical cells by using LSCM multispectral fluorescence analysis, thus revealing the distribution of these substances specifically along the thallus surface and inside chloroplasts and adjacent vesicles (which is more evident in the deconvoluted images). Complementarily, the intracellular distribution of autofluorescent substances (bluish autofluorescence) coincided with some regions that were labeled with NR fluorochrome (which specifically binds neutral lipids), confirming the lipidic nature of these autofluorescent regions. NR staining has commonly been used to localize lipids in algal cells, e.g. for the localization of neutral long-chain lipids in I. galbana (Eltgroth et al. 2005) and triacylglycerides and FAs in Chlamydomonas reinhardtii (Chlorophyta) (Wang et al. 2009). As we show with NR labeling, one of the primary lipid sites within the L. translucida cortical cells is the chloroplast. FA biosynthesis in Plantae is well documented to take place within plastids, where it is initiated by the ATP-dependent carboxylation of acetyl-CoA (Somerville et al. 2000). Acetyl-CoA carboxylase catalyzes the formation of malonyl-CoA, and FA synthase catalyzes the extension of the growing acyl chain, usually for the final products 16:0, 18:0 and C18:1 (FA species) (Ohlrogge and Jaworski 1997). Because hexadecanoic acid (C16:0), octadecanoic acid (C18:0) and 9-octadecenoic acid (Z) (C18:1) were the most commonly found FA compounds in the L. translucida LI fraction, we propose that L. translucida chloroplasts could be involved in the first step of FA biosynthesis in this alga. In this study, conventional TEM images of L. translucida cortical cells showed osmiophilic material precipitation inside the vesicles, which budded from a cisterna-shaped organelle (possibly ER) and close to chloroplasts and the Golgi apparatus. The TEM images also showed the ER releasing electron-dense vesicles to the cytoplasmic regions near the cell wall, which could indicate the involvement of the ER in later steps in the synthetic pathway of unsaturated lipids, and, perhaps, their release to the cell wall. In B. braunii, many enzymes involved in HC biosynthesis are localized to the ER, which is also responsible for the transport of vesicles to the outer cell wall region (Weiss et al. 2012). The TEM images also showed large electron-dense bodies in Emiliania huxleyi (Prymnesiophyta), a species that produces polyunsaturated long-chain lipids (C37–C39). It is important to mention that the osmium tetroxide (OsO4) used to post-fix the material during TEM preparation reacts with unsaturated lipids, producing osmium precipitation over the sectioned material (Korn 1967, Bozzola and Russell 1998). These bodies in E. huxleyi were confirmed as lipid bodies through NR staining (Eltgroth et al. 2005). Thus, we propose that electron-dense and autofluorescent vesicles, some of the structures of which were labeled with NR, presumably represent the location of one of the algal lipid fractions, specifically the unsaturated FAs identified by GC-MS (9-hexadecanoic acid; 9,12-octadecadoenoic acid Z-Z; 9-octadecenoic acid E; 5,8,11,14,17-eicosapentaenoic acid; 8,11,14-eicosatrienoic acid Z-Z-Z; and 15-tetracosenoic acid). With regards to the location of saturated FA derivatives, the TEM images also displayed vacuoles with granular electron-dense material (VE) and vacuoles that enveloped small electron-lucent vesicles (V). Interestingly, these small vesicles were exudated to the cell walls according to the TEM. Similar nanometric vesicles were identified in SEM images of fractured cortical cells, above the plasmatic membrane and inside fractured vacuoles of cortical cells. Because the TEM images of L. translucida cortical cells primarily showed electron-lucent vesicles being transported to the inner region of the cell wall, along with the fact that these vacuoles only have electron-lucent vesicles (saturated FA does not react with OsO4), we suggest that saturated FA derivatives are being transported by vesicle exudation to the algal thallus surface in L. translucida cortical cells. These assertions were corroborated by the LSCM image localization of lipid derivatives labeled with NR probe above the algal thallus surface and inside the cell walls. GC-MS analyses allowed for the identification of five linear saturated HCs in L. translucida HE. NR staining has also been used to localize HCs in algal cells, e.g. the localization of linear C27 HCs in P. lunula (Dahmen and Leblond 2013) and HCs in B. braunii (Weiss et al. 2012). A large amount of vesicle budding was observed in association with an active trans-Golgi network apparatus in L. translucida cortical cells. These vesicles formed structures resembling the vacuoles with vesicular contents, the latter of which are similar to the exudated material. Taken together, these findings could indicate a role for the Golgi apparatus in the formation of these vacuoles and for sorting subsequent exudations. In plants, the trans-Golgi network-derived vesicles were already known to form an intermediate compartment, between the late trans-Golgi sorting site and the vacuole (Marty 1999). These vesicles have been collectively referred to as pre-vacuoles because they act ontogenetically as the immediate progenitors of the vacuole (Marty 1999). They also mediate transport between the ER/the Golgi complex to the vacuole and thus take functional precedence along the path to the vacuole (Marty 1978). In many systems, the ultrastructure of the Golgi is an expression of its function in the secretion process (Griffing 1991). In plants, the Golgi apparatus is involved in polysaccharide, FA, ST and phytosterol synthesis (Andreeva et al. 1998, Hawes 2005). The Golgi apparatus in L. translucida cortical cells seems to be involved in vacuole ontogeny, and, based on GC-MS analyses and LCSM images that localized FA derivatives inside smaller vacuoles, the present data suggest that the Golgi apparatus can also be involved in the compartmentalization of FA derivatives in L. translucida cortical cells. With respect to the chemical profile approach, the GC-MS analyses revealed that the L. translucida HE is made of a complex mixture of FA derivatives containing the following major compounds: docosane (HC, 61.66%), hexadecanoic acid, methyl ester (FA, 57.86%) and cholesterol trimethylsilyl ether (ST, 100%). FAs and HCs were already determined in some algal species such as Pseudochoricystis ellipsoidea (Chlorophyta) (Satoh et al. 2010) and Botryococcus braunii (Chlorophyta) (Yoshida et al. 2012). These lipids are usually found in the Rhodophyta species, but not as the major secondary metabolite. In the present study, the large amount of FA/HC and the identification of only one ST, along with the absence of typical Rhodophyta isoprenoids (terpenes and phytosterols), suggests that the most active secondary metabolite biosynthetic route in L. translucida is the FA pathway (Ohlrogge and Jaworski 1997, Brown 1998). We tested L. translucida HE, HC and LI against bacteria and microalgae species which are typical components of the marine fouling community (Aguila-Ramírez et al. 2014, Trepos et al. 2015). We determined that both HCs and LI fractions inhibited marine fouling bacteria and microalgae with MIC values ranging from 0.1 to 1.0 µg ml–1 and from 10 to 100 µg ml–1, respectively. It was demonstrated that the FA derivatives from L. translucida had lower MIC values than those found for S. isoetifolium and C. serrulata (1.0–10 µg ml–1; Iyapparaj et al. 2014) and for P. brasiliense (10–50 µg ml–1; Paradas et al. 2015) when tested against the same microbial strains used in this study. Among the bacteria tested, P. elyakovii is a common component of macroalgal microflora and can also be involved in spreading of disease; moreover it was isolated from wounded tissue of Laminaria japonica (Sawabe et al. 2000). Thus, based on the MIC values obtained, we can hypothesize a possible role for FA derivatives in L. translucida for protection against fouling. Nevertheless, the FA fractions should be tested against isolated sympatric bacteria from L. translucida to confirm the AF role of these compounds in natural conditions. When comparisons were made between MIC values from L. translucida HCs and LI isolated fractions, the HCs presented the lowest MIC values against microfouling species (bacteria and microalgae). However, the autofluorescence emission wavelength (500 nm) observed above the surface of the algae (LSCM) was not determined in the isolated HC fraction (HPLC). Otherwise, NR labeling was observed in regions inside cell walls, which could indicate HC transport to the algal surface. Some authors have attributed the NR staining to cellular HC localization (Kimura et al. 2004, Eltgroth et al. 2005, Leblond et al. 2010, Dahmen and Leblond 2013). In P. lunula, NR staining localized C27 HCs inside lipid bodies and above outer sheath cells, where these HCs are transported to the outer region of the cell wall (Dahmen and Leblond 2013). In B. braunii cells, NR staining coupled with confocal Raman microspectroscopy has shown the presence of botryococcenes (HCs) in the extracellular matrix (ECM) and in multiple cytoplasmic lipid bodies (Weiss et al. 2010), supporting the hypothesis that the lipid body botryococcenes are secreted out of the cells (Metzger and Largeou 2005, Metzger et al. 2008). In the present work, NR staining in L. translucida strongly labeled the thalli surface. Perhaps this fluorescence could be related to both HC- and LI-specific localization, being transferred by vesicle exudation to the outer region of the cell wall where it provides protection against biofouling. The current knowledge about linear n-alkane biosynthesis in eukaryotic algae indicates that these compounds are the result of FA decarboxylation (Dahmen and Leblond 2013). Interestingly, the GC-MS data identified 20 species of FA in the L. translucida LI fraction, where some of them, namely the tricosanoic acid (23:0), tetracosanoic acid (24:0) and 15-tetracosenoic acid (24:1ω9), have higher carbon numbers than all the linear n-alkanes identified here (hexadecane-C16, octadecane-C18, nonadecane-C19, icosane-C20 and docosane-C22). In this way, the present data strongly support the hypothesis that the HCs in L. translucida cortical cells are being produced by the decarboxylation of saturated and unsaturated FAs. However, the confirmation of HC distribution over the thallus surface requires a complementary chemical approach, perhaps by DESI-MS and MALDI-TOF imaging. In conclusion, this study was the first to identify the cellular structures responsible for FA derivative (e.g. HCs) biosynthesis in L. translucida cortical cells. The biosynthetic pathway involves chloroplasts, the Golgi apparatus, the ER and vacuoles. These compounds are stored inside vacuoles and vesicles for later transport by vesicle exudation to the thallus surface. The autofluorescence results primarily confirmed the secretion of FAs to cell surfaces, and NR staining also revealed the location of whole lipids in L. translucida cortical cells. These results, together with the electron microscopy results and chemical analysis, also indicate the secretion of HCs to the algal thallus surface, where a lipidic layer may play a role in AF protection. Materials and Methods Algal sampling Laurencia translucida specimens were collected from the intertidal zone at Rasa beach from 2010 to 2013, from June to October (Rio de Janeiro State; Brazil; 22°43′58′′S, 41°57′25′′W). After collection, living algal samples were stored in filtered seawater inside a dark isothermic chamber and transported to the laboratory. LSCM analyses Longitudinal sections of living algae that had been collected on a glass slide with a glass cover slip were transferred to an LSCM Leica TCS SPE AOBS (Leica Microsystems Company), and a spectral fluorescence scanning analysis was performed to determine the autofluorescence emission pattern. The microscope was previously set to discover the autofluorescence emission peaks under the same operational conditions as the fluorimetric analysis that was performed with L. translucida HE, as follows: (i) excitation wavelength = 405 nm; (ii) beginning of emissions spectra at 450 nm; (iii) end of emissions spectra at 750 nm; (iv) emissions wavelengths analyzed at intervals of 10 nm (e.g. 450, 460, 470 nm,…, 740, 750 nm); (v) acquisition time = 200 s; and (vi) Z-thickness = 1 µm. For the final acquisition of the autofluorescent images, the resulting resolution was 2,048 × 2,048, and, to improve the image quality, some of them were processed by deconvolution with LAS AF software (Leica Microsystems Company). Nile red (NR) staining Laurencia translucida fragments (algae sliced into longitudinal sections) were fixed with 4% formaldehyde (diluted in seawater) and stained in 1 ml aliquots of NR (Sigma-Aldrich Company) solution prepared in seawater (10 µg ml–1). Afterwards, the samples were transferred to a glass slide with a glass cover slip for observation under the LSCM Leica TCS SPE AOBS (Leica Microsystems Company). The emission patterns of FA derivatives that were related to NR were determined according to Eltgroth et al. (2005). For the control, images from L. translucida fixed cells were obtained using the same acquisition parameters used for NR-stained samples, but they only excluded the NR incubation. Both treatments were subjected to 488 nm fluorescence excitation. TEM analyses For the ultrastructural characterization of L. translucida cortical cells, the samples were fixed in a buffered seawater solution of 5% formaldehyde and 5% glutaraldehyde (diluted in 0.1 M Na cacodylate buffer, pH 7.4; from Sigma-Aldrich Company; for details see Supplementary Methods S1). SEM analyses To observe the interface among the cell wall surfaces and the plasmatic membrane from L. translucida cortical cells, SEM images of fractured cells were obtained as already described elsewhere (Paradas et al. 2010). Analyses were performed with a Zeiss EVO 40 scanning electron microscope (Zeiss Company) using an acceleration voltage of 15 kV; for details, see Supplementary Methods S2. Chemical analyses Crude extract from L. translucida (0.26% DW) was obtained by extraction in n-hexane (Merck & Co. Inc.) by following previously described procedures for FA derivative (e.g. HCs) extraction (Frenz et al. 1989). The extract was then filtered and the solvent was eliminated under reduced pressure in a rotatory evaporator at room temperature. The lipids from L. translucida HE were characterized and isolated by preparative one-dimensional TLC (silica gel 60 F254S; Merck & Co. Inc.) with the following mobile phase: hexane, diethyl ether and acetic acid at 90 : 7.5 : 1 (v/v/v) (Manilla-Perez et al. 2010); for details see Supplementary Methods S3. Two isolated fractions were obtained from the L. translucida HE, LT1 and LT2, which were named here as HC and LI, respectively. The L. tranlucida HE was also analyzed by 1H-NMR (300 MHz, CDCl3), in which primarily FA derivative signals were observed in the spectra. To understand the fluorescence properties of isolated FA derivatives and to look for similar patterns in the autofluorescence microscopy images, both HC and LI fractions were re-suspended in methanol and injected into a Shimadzu HPLC (Shimadzu Corporation) for autofluorescence emission analyses. The HPLC was equipped with a pump (LC-20AT), a degasser (DGU-20AS), a communicator module (CBM-20 A), a fluorescence detector (RF-20 A), a shim pack VP-ODS column (150 mm × 4.6 µm i. d.) and LCsolution software (Shimadzu Corporation). The RF-20 A detector was set to excitation at 405 nm and emission at 500 nm, and the peak autofluorescence emission observed by LSCM did not correspond to any photosynthetic pigments. Based on the TLC lipid profile, specific GC-MS analysis methods were chosen for compound identification: HCs (Borik 2014), FAs (Christie 1989) and STs (Fridberg et al. 2008) (Supplementary Methods S4). Antibacterial tests The L. translucida fractions of HE, HC and LI were tested for their inhibitory activity against the growth of five strains of biofilm-forming marine bacteria (Paradas et al. 2015) that were obtained from a collection from the University of Portsmouth (ATCC), namely H. marina (25374), P. irgensii (700398), P. elyakovii (700519), S. putrefaciens (8071) and V. aestuarianus (35048). Each treatment and control (culture media) was replicated six times. Antimicroalgal tests The L. translucida fractions of HE, HC and LI were tested for inhibitory activity against the benthic phase of four strains of marine microalgae (Paradas et al. 2015) that were obtained from the Université de Caen Basse-Normandie (AC) and the ATCC, namely C. reptans (AC 132), C. cloisterium (AC 170), E. gayraliae (AC 15), N. jeffreyae (AC 181) and C. globosum (ATCC 8071). Funding This work was supported by the Coordenação de Aperfeiçoamento de Pessoal de Nível Superior [No. 06/2011 to G.M.A.F.]; Conselho Nacional de Desenvolvimento Científico, Tecnológico [grant No. 06/2011 to G.M.A.F., No. 09/2011]; Fundação de Amparo à Pesquisa do Estado do Rio de Janeiro [No. 09/2011 to G.M.A.F.]; Ministério de Ciência e Tecnologia [No. 06/2011 to G.M.A.F.]; the Ministério da Educação [No. 06/2011 to G.M.A.F.]. Abbreviations Abbreviations AF antifouling DG diglyceride ER endoplasmic reticulum FA fatty acid FFA free fatty acid GC-MS gas chromatography–mass spectrometry HC hydrocarbon HE hexanic extract LI lipid fraction LSCM laser scanning confocal microscopy MIC minimum inhibitory concentration MG monoglyceride NMR nuclear magnetic resonance NR Nile red PAL palmityl palmitate SEM scanning electron microscopy ST sterol TEM transmission electron microscopy TG triglyceride TLC thin-layer chromatography Disclosures The authors have no conflicts of interest to declare. Acknowledgements The authors are grateful to CAPES and CNPq for their financial support. R.C.P., G.M.A.F., L.T.A. and A.R.S. would like to thank CNPq for research fellowships. This study is part of the PhD Thesis of W.C.P. The authors thank Dr. Marcos Farina (Laboratório de Biomineralização, ICB, UFRJ) and Dr. Wanderley de Souza (Laboratório de Ultraestrutura Celular Hertha Meyer, IBCCF, UFRJ) for the use of their microscopy facilities. The authors also thank Dr. Luzineide Tinoco (LAMAR-NPPN/UFRJ) for performing the NMR analyses, BSc. Mileane Busch (Laboratório de Bioquímica de Lipídeos e Lipoproteínas IbqM-UFRJ) for the GC-MS analyses, Dr. Miria Gomes Pereira (Laboratório de Ultraestrutura Celular Hertha Meyer, IBCCF, UFRJ) for helpful suggestions on development of the TLC method, and Dr. Benoit Veron (Algobank, Caen, France) for providing the microalgae strains. References Andreeva A.V. Kutuzov M.A. Evans D.E. Hawes C.R. 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TI - A Novel Antifouling Defense Strategy from Red Seaweed: Exocytosis and Deposition of Fatty Acid Derivatives at the Cell Wall Surface
JF - Plant and Cell Physiology
DO - 10.1093/pcp/pcw039
DA - 2016-03-02
UR - https://www.deepdyve.com/lp/oxford-university-press/a-novel-antifouling-defense-strategy-from-red-seaweed-exocytosis-and-042xGSCi62
SP - 1008
EP - 1019
VL - 57
IS - 5
DP - DeepDyve
ER -