MEVALONOSOMES: SPECIFIC VACUOLES CONTAINING THE MEVALONATE PATHWAY IN PLOCAMIUM BRASILIENSE CORTICAL CELLS (RHODOPHYTA)1

J. Phycol. 51, 225–235 (2015) © 2015 Phycological Society of America DOI: 10.1111/jpy.12270

MEVALONOSOMES: SPECIFIC VACUOLES CONTAINING THE MEVALONATE PATHWAY IN PLOCAMIUM BRASILIENSE CORTICAL CELLS (RHODOPHYTA)1 Wladimir Costa Paradas Departamento de Biologia Marinha, Universidade Federal Fluminense, Outeiro S~ao Jo~ao Batista, s/no., Niter oi, Rio de Janeiro, Brazil

Thalita Mendes Crespo, Leonardo Tavares Salgado Diretoria de Pesquisas, Instituto de Pesquisas Jardim Bot^anico do Rio de Janeiro, Rua Pacheco Le~ao, 915, Rio de Janeiro, Brazil

Leonardo Rodrigues de Andrade Centro de Ci^encias da Sa ude, Instituto de Ci^ encias Biom edicas, Departamento de Histologia e Embriologia, Universidade Federal do Rio de Janeiro (UFRJ), Av. Carlos Chagas Filho, 373, bloco: B, sala F2-27, Rio de Janeiro, Brazil

Ang
e lica Ribeiro Soares N ucleo de Pesquisas em Ecologia e Desenvolvimento Social de Maca e, Universidade Federal do Rio de Janeiro, Rua Rotary Club, s/no., S~ao Jos e do Barreto, Maca e, Rio de Janeiro, Brazil

Claire Hellio Universite de Bretagne Occidentale, LEMAR UMR 6539, IUEM - Technopole Brest-Iroise, Rue Dumont d’Urville, Plouzan e, France

Ricardo Rogers Paranhos Departamento de Biologia Marinha, Universidade Federal Fluminense, Outeiro S~ao Jo~ao Batista, s/no., Niter oi, Rio de Janeiro, Brazil

Lilian Jorge Hill, Geysa Marinho de Souza Diretoria de Pesquisas, Instituto de Pesquisas Jardim Bot^anico do Rio de Janeiro, Rua Pacheco Le~ao, 915, Rio de Janeiro, Brazil

Alphonse Germaine Albert Charles Kelecom Departamento de Biologia Geral, Universidade Federal Fluminense, Outeiro S~ao Jo~ao Batista, s/no., Niter oi, Rio de Janeiro, Brazil

Bernardo Ant^ o nio Perez Da Gama, Renato Crespo Pereira Departamento de Biologia Marinha, Universidade Federal Fluminense, Outeiro S~ao Jo~ao Batista, s/no., Niter oi, Rio de Janeiro, Brazil

and Gilberto Menezes Amado-Filho2 Diretoria de Pesquisas, Instituto de Pesquisas Jardim Bot^anico do Rio de Janeiro, Rua Pacheco Le~ao, 915, Rio de Janeiro, Brazil

halogenated monoterpenes. P. brasiliense specimens were submitted to a cytochemical analysis of the activity of the 3-hydroxy-3-methylglutaryl-CoA synthase (HMGS). Using transmission electron microscopy (TEM), we confirmed the presence of HMGS activity within the Mev. Because HMGS is necessary for the biosynthesis of halogenated monoterpenes, we isolated a hexanic fraction (HF) rich in halogenated monoterpenes from P. brasiliense that contained a pentachlorinated monoterpene as a major metabolite. Because terpenes are often related to chemical defense, the antifouling (AF) activity of pentachlorinated

This paper has identified, for the first time in a member of the Rhodophyta, a vacuolar organelle containing enzymes that are involved in the mevalonate pathway—an important step in red algal isoprenoid biosynthesis. These organelles were named mevalonosomes (Mev) and were found in the cortical cells (CC) of Plocamium brasiliense, a marine macroalgae that synthesizes several 1

Received 20 April 2014. Accepted 21 January 2015. Author for correspondence: e-mail gilbertoamadofilho@ gmail.com. Editorial Responsibility: J. Raven (Associate Editor) 2

225

226

W L A D I M I R CO S T A PA R A D A S E T A L .

monoterpene was tested. We found that the settlement of the mussel Perna perna was reduced by HF treatment (2.25 times less than control; 40% and 90% of fouled surface, respectively; P = 0.001; F9,9 = 1.13). The HF (at 10 lg
mL1) also inhibited three species of fouling microalgae (Chlorarachnion reptans, Cylindrotheca cloisterium, and Exanthemachrysis gayraliae), while at a higher concentration (50 lg
mL1), it inhibited the bacteria Halomonas marina, Polaribacter irgensii, Pseudoalteromonas elyakovii, Shewanella putrefaciens, and Vibrio aestuarianus. The AF activity of P. brasiliense halogenated monoterpenes and the localization of HMGS activity inside Mev suggest that this cellular structure found in CC may play a role in thallus protection against biofouling. Key index words: 3-hydroxy-3-methylglutaryl-CoA synthase; antifouling activity; chemical defense; cortical cell cytochemistry; halogenated monoterpenes; mevalonate pathway; mevalonosome; osmiophilic bodies; Plocamium brasiliense Abbreviations: AF, antifouling; CC, cortical cells; C, chloroplast; CW, cell wall; GC-MS, gas chromatography mass spectrometry; HF, hexanic fraction; HMGS, 3-hydroxy-3-methylglutaryl-CoA synthase; LB, Luria Hinton Broth; MC, medullary cells; Mev, mevalonosome; MIC, minimum inhibition concentration; NMR, nuclear magnetic resonance; OB, osmiophilic bodies; OM, optical microscopy; P, pit connections; SEM, scanning electron microscopy; SER, smooth endoplasmic reticulum; SG, floridean starch grains; TEM, transmission electron microscopy; V, vacuole

Studying the localization, storage, and exudation mechanisms of biologically active secondary metabolites in macroalgae is necessary to enrich our understanding of the ecological significance of surface-active inhibitors (Dworjanyn et al. 1999, Paul et al. 2006, Salgado et al. 2008). Numerous studies have focused on the activity of macroalgal secondary metabolites against fouling organisms (Da Gama et al. 2008, Nylund et al. 2008, Bazes et al. 2009, Bianco et al. 2009, Hellio et al. 2009, Plouguerne et al. 2010, Fusetani 2011, Silkina et al. 2012). However, little is known about the cellular mechanisms involved in secondary metabolite biosynthesis (Dworjanyn et al. 1999, Paul et al. 2006, Salgado et al. 2008). More than half of the reported secondary metabolites from macroalgae are isoprenoid derivatives such as terpenes, steroids, carotenoids, prenylated quinines, and hydroquinones, which are derived from either the classical mevalonic acid pathway (MVA) or from the 2C-methyl-D-erythritol 4-phosphate (MEP) pathway (Maschek and Baker 2008). The key enzymatic precursors in these pathways are 3-hydroxy-3-methylglutaryl-CoA synthase (HMGS)

and 1-deoxy-D-xylulose-5-phosphate synthase, respectively (Hemmerlin et al. 2012, see Lohr et al. 2012 for a review). Traditionally, the MVA pathway occurs in the cytoplasm and mitochondria, and the MEP pathway is localized in chloroplasts (C); however, the exact cellular localization of isoprenoid synthesis and storage is poorly understood (Lohr et al. 2012). Recent studies suggest that in some species of red algae, the MVA pathway has been lost (Lohr et al. 2012), but in Galdieria sulphuraria (Rhodophyta), both the MVA and MEP pathways are present (Schwender et al. 1997). In Cyanidioschyzon merolae (Rhodophyta), a close relative of G. sulphuraria, the only terpenoid synthesis gene identified was related to HMGS, whereas all of the genes encoding enzymes for the subsequent reactions in the MVA pathway are absent (Matsuzaki et al. 2004). However, to date, only one study has provided biochemical data supporting the presence of the MVA pathway in Florideophyceae (Barrow and Temple 1985). Similarly, cellular microscopy studies of the cellular localization of secondary metabolites biosynthetic steps have not yet been reported for marine macroalgae. The spatial distribution throughout the cell of components related to secondary metabolism has been detailed for filamentous fungi and plants (Lendenfeld et al. 1993, Hoppert et al. 2001, Kutchan 2005, Lunn 2007, Hong and Linz 2008, 2009, Saikia et al. 2008). For example, in Aspergillus parasiticus, aflatoxisomes constitute a group of specialized trafficking vesicles that participate in the biosynthesis and transport of aflatoxin to the cell exterior (Chanda et al. 2009). In general, sub-cellular localization studies indicate that the enzymes, substrates, intermediates, and end products of secondary metabolism often accumulate at different subcellular locations (Roze et al. 2011, see Ziegler and Facchini 2008 for a review). The intracellular storage sites of secondary metabolites and the locations of their biosynthetic pathways and release mechanisms are highly reflective of their ecological roles (Pereira and Da Gama 2008). The production of antifouling (AF) metabolites is frequently associated with the capability of some species to store some isoprenoids derivatives or secondary metabolites in specialized structures near the surface cell layer (Dworjanyn et al. 1999, Paul et al. 2006, Salgado et al. 2008). In macroalgae, three types of specialized structures have been described so far: gland cells (or vesicle cells), in which a storage vesicle occupies almost the entire cellular space (Young and West 1979, Dworjanyn et al. 1999); the corps en cerise, which are refractive cell inclusions found in some species of Laurencia (Young et al. 1980, Salgado et al. 2008); and the physodes found in Ochrophyta (Schoenwaelder 2002, Ank et al. 2014). With regard to the exudation mechanisms of secondary metabolites, in brown algae, for example,

ME V A L O N A T E P A T H W A Y I N P . B R A S I L I E N S E C O R T I C A L C E L L S

the Golgi apparatus plays a role in the synthesis and transport of phlorotannins to the cell wall (CW) via the exocytic pathway (Schoenwaelder 2002). In Laurencia dendroidea (Rhodophyta), exudation occurs by way of a complex system involving the microfilament- and microtubule-mediated vesicular transport of halogenated metabolites from corps en cerise (storage sites) to the cell periphery (Salgado et al. 2008, Paradas et al. 2010, Reis et al. 2013). The metabolites are then exocytosed at the CW and reach the algal surface, where they act as AF chemicals (Salgado et al. 2008, Paradas et al. 2010). In this context, the study of the genus Plocamium has been restricted mainly to the chemical characterization of the products of secondary metabolism and their activity on other organisms (Kladi et al. 2004). These algae produce cyclic and acyclic halogenated monoterpenes (Diaz-Marrero et al. 2002), which are known to decrease the activities of herbivores, fouling organisms, fungi, and competitors (Kladi et al. 2004). Monoterpenes isolated from Plocamium costatum displayed anti-settling activity against cyprid larvae of the barnacle Balanus amphitrite at 1 lg
cm2, where the unit lg
cm2 indicates the amount of material (lg) used to coat one square centimeter (cm2) of the petri dish internal glass used in B. amphitrite assays (K€ onig et al. 1999a). Thus, although many studies have highlighted the ecological properties of Plocamium compounds (K€ onig et al. 1999a,b, Pereira et al. 2002, Kladi et al. 2004), the cellular structures involved in isoprenoid derivative biosynthesis and storage in Plocamium cortical cells (CC) have not been studied. Moro et al. (2003) described the ultrastructure of Plocamium cartilagineum but did not identify any specific organelles related to isoprenoid biosynthesis or storage. However, published data (Barrow and Temple 1985) guide the present work to investigate the location of the mevalonate pathway and the possible existence of cellular structures involved in the synthesis and storage of the substances involved. Radioactive [3H, 14C] mevalonate precursors have been successfully incorporated into halogenated monoterpenes in Plocamium cartilagineum (Rhodophyta), thus, confirming the occurrence of this metabolic pathway in the Plocamium genus and its role in monoterpene synthesis (Barrow and Temple 1985). The species chosen for the present investigation was Plocamium brasiliense ([Greville] M.A.Howe & W.R.Taylor [Rhodophyta]). Although studies of the chemical ecology of P. brasiliense are scarce, it was demonstrated that P. brasiliense monoterpenes have high anti-herbivore activity (Pereira et al. 2002, Pereira and Vasconcelos 2014). In an applied context, these molecules showed activity as herbicides (Fonseca et al. 2012) and against both herpes virus (Ferreira et al. 2010, Pinto et al. 2014) and snake venom (Claudino et al. 2014). Thus, the aims of the present work were: (i) to describe the ultrastructure of P. brasiliense CC using optical

227

(OM), transmission (TEM), and scanning electron microscopy (SEM); (ii) to localize mevalonic acid enzyme activity inside CC by using a specific cytochemical assay; and (iii) to investigate the AF activity of P. brasiliense halogenated monoterpenes using in vitro bioassays. MATERIALS AND METHODS

Algal sampling. Specimens of P. brasiliense were collected by self-contained underwater breathing apparatus in the subtidal zone at Forno beach in 2011 from June to October (Rio de Janeiro State; Brazil; 22°450 42.17″ S; 41°520 29.25″ W). After collection, living algal samples were stored in filtered seawater inside a dark isothermic chamber and transported to the laboratory. Optical microscopy. Morphological characterization of living CC of P. brasiliense was performed using the optical microscope Olympus BX 50 (Olympus, Tokyo, Japan) equipped with a 1009 N.A. 1.3 objective lens using oblique illumination and connected to a CoolSnap-Pro Color RS Photometrics camera (Photometrics, Tucson, AZ, USA). Digital images were analyzed using ImageJ software (Abramoff et al. 2004) to describe the morphological aspects of the organelles, such as area, circularity, and diameter (n = 60). Transmission electron microscopy. Specimens were fixed in a solution of 4% formaldehyde and 5% glutaraldehyde (SigmaAldrich Company, Saint Louis, MO, USA) in 0.1 M cacodylate buffer pH 7.3 diluted in sterile seawater. Post-fixation was performed in OsO4 1% (Sigma-Aldrich Company) for 2 h at room temperature. Algal samples were then dehydrated in a crescent acetone series (to 100%) and embedded in Spurr resin (Sigma-Aldrich Company) at room temperature. The polymerization process was performed at 70°C. Ultrathin sections (50 nm) were obtained by using an ultra-microtome Leica EM UC7 (Leica Microsystems Company, Wetzlar, Hessen, Germany), collected on copper grids (300 mesh; Electron Microscopy Sciences Company, Hatfield, PA, USA) and observed on a JEOL 1010 EX (Jeol Company, Tokyo, Japan) TEM microscope operated at 80 kV. Localization of mevalonic acid enzyme activity. Specimens of P. brasiliense were exposed to enzymatic bioassays in which algal fragments were treated with prospected substrates for the enzymes acetyl-CoA and acetoacetyl-CoA (from SigmaAldrich Company). This treatment is based on prior knowledge of a mevalonate-dependent pathway, in which the HMGS transforms acetyl-CoA and acetoacetyl-CoA into 3-hydroxy-3-methylglutaryl-CoA. Another enzymatic product of this reaction is free Coenzyme A-SH (CoA-SH). The first step of the cytochemical reaction occurs when potassium ferricyanide (Sigma-Aldrich Company) is combined with the incubation solution and reacts with CoA-SH, reducing it to ferrocyanide. The ferrocyanide then reacts with added uranyl acetate (Sigma-Aldrich Company) to form uranyl ferrocyanide, which precipitates and appears as a highly particulate electron-dense material in TEM. According to Croteau et al. (2000), the enzymology of isopentenyl pyrophosphate (IPP) biosynthesis—the precursor of all terpenoids compounds—by the acetate/mevalonate pathway is widely accepted. This cytosolic IPP pathway involves the two-step condensation of three acetyl-CoA molecules catalyzed by thiolase (TH) and HGMS (Croteau et al. 2000). The resulting product, 3-hydroxy-3-methylglutaryl-CoA, is subsequently reduced by HMGR in two coupled reactions to form mevalonic acid (Croteau et al. 2000). The TH, HMGS, and HMGR reactions are some of the many chemical reactions in which free CoA-SH is released (Croteau et al. 2000). For

228

W L A D I M I R CO S T A PA R A D A S E T A L .

example, carnitine acetyltransferase (ACT) also releases CoASH. The discussed reactions are described below: TH: Acetyl-CoA + Acetyl-CoA ? Acetoacetyl-CoA (+ CoASH) HMGS: Acetyl-CoA + Acetoacetyl-CoA ? 3-hydroxy-3-methylglutaryl-CoA (+ CoA-SH) HMGR: 3-hydroxy-3-methylglutaryl-CoA ? Mevalonic acid (+ CoA-SH) ACT: Carnitine + Acetyl-CoA ? Acetylcarnitine (+ CoASH) In this way, the localization of acetyl-CoA, acetyl-CoA Ctransferase (TH), and 3-hydroxy-3-methylglutaryl-CoA reductase (HGMR) are indistinguishable from that of HGMS because both reactions produce CoA-SH, which precipitates as uranyl ferrocyanide (Curry 1987). The HGMS reaction necessarily labels HGMR. The highest level of precipitation is expected to be associated with HGMS and not TH because a high quantity of enzymatic substrate was presented to HGMS (Curry 1987). In these experiments, specimens of P. brasiliense were fixed for 30 min in 4% formaldehyde and 1% glutaraldehyde in 0.05 M sodium cacodylate buffer (pH 7.0) diluted in sterile seawater, followed by a buffer rinse (0.05 M cacodylate, pH 7.0). Subsequently, the algal tissue was pre-incubated at ambient temperature for 20 min in 3 mM potassium ferricyanide in 0.05 M cacodylate buffer (pH 7.0) followed by a buffer rinse (Curry 1987). Then, algae were incubated for 45 min at ambient temperature to allow enzyme catalysis and precipitation to occur (Curry 1987). Complete media including acetoacetyl-CoA or carnitine were used along with several controls (Table 1). Specimens of P. brasiliense were then post-fixed for 1 h at ambient temperature in 2% (w/v) osmium tetroxide in 0.05 M sodium cacodylate buffer (pH 7.0) followed by a buffer rinse. Then, the algal tissue was dehydrated in a crescent acetone series (to 100%) and embedded in Spurr resin at room temperature. The polymerization process was performed at 70°C. Thin sections (200 nm) were obtained on an ultra-microtome Leica EM UC7 and collected on single slot copper grids (Electron Microscopy Sciences Company) coated with a Formvar film (~20 nm). The grids were observed in a FEI TECNAI G20 (FEI Company, Hillsboro, OR, USA) TEM with an accelerated voltage of 200 kV. Scanning electron microscopy. To provide three-dimensional information on organelle distribution, SEM was performed using a Zeiss EVO 40 (Zeiss Company, Oberkochen, BW, Germany) with an accelerated voltage of 15 kV. Fragments of P. brasiliense were fixed in 4% glutaraldehyde and 5% formaldehyde in 0.1 M pH 7.4 sodium cacodylate buffer diluted in seawater. Thereafter, samples were washed with buffered sterile seawater (0.05 M sodium cacodylate, pH 7.4) and postfixed in 1% OsO4 for 1 h at 20°C. After washing in buffered sterile seawater, the samples were dehydrated in a series of

ethanol/water solutions (30%, 50%, 70%, and 100%). Algal samples were dried using the critical point dryer LEICA CPD (Leica Microsystems Company), and the dried pieces were mounted on aluminum sample holders for SEM using double-sided carbon tape (SPI Supplies Company, West Chester, PA, USA). The thalli of P. brasiliense were fractured, and double-sided adhesive was used to remove tissue fragments (in the following order: stub/carbon tape/alga/adhesive tape). The tissue fracture resulted in cellular fracture, thus exposing the intracellular contents of the fractured cells. The samples were then coated with a thin gold layer (~20 nm) with a Sputter coater BalTec SCD 050 (Baltec Company, Manchester, NH, USA). Extraction, fractionation, and chemical analyses. Crude extract from P. brasiliense (0.6% dry weight, DW) was obtained by extraction in dichloromethane (Merck & Co. Inc., Readington Township, NJ, USA) for 15 d (the solvent was exchanged three times) following previously described procedures for natural product extraction (Da Gama et al. 2002). To obtain a chemical profile of the compounds in the extract, it was initially analyzed with 1H nuclear magnetic resonance (NMR; 300 MHz, CDCl3) and gas chromatography mass spectrometry (GC-MS). After that, the crude extract (97.3 g) was subjected to a preparative chromatography fractionation to isolate the most abundant terpenes. Then, 10 g of silica gel (Merck 60 F254 0.5 mm; Merck & Co. Inc.) was placed inside a glass filter holder (Merck & Co. Inc.) attached to a Kitasato glass (1,000 mL). Afterward, the crude extract was carefully inserted into the silica gel and then filtered under a vacuum with 25 mL of hexane (Merck & Co. Inc.) four times. The filtered and hexanic fractions (HF) were collected separately for solvent elimination under reduced pressure in a rotary evaporator at room temperature. The HF was analyzed by 1D and 2D NMR techniques (at 300 MHz to 1H and 75 MHz to 13 C in CDCl3) and by GC-MS. All the GC-MS analyses were performed using a QP2010 plus series instrument (Shimadzu Corporation, Kyoto, Japan). The GC-MS was equipped with a polysiloxane capillary column (RTXâ-1MS; 30 m 9 0.25 mm i.d. 9 film thickness 0.25 lm; Restek Corporation, Bellefonte, PA, USA). The temperature was programmed to hold at 100°C for 2 min and then increase from 8°C
min1 to 310°C, where it was maintained for 1 min. Helium was used as a carrier gas at a flow rate of 1.2 mL
min1. The injector and interface temperatures were 260°C and 320°C, respectively. Electron impact spectra were recorded at 70 eV with scan time of 1 s. The compounds were identified by comparison of their mass spectra with those provided in a mass spectral database (NIST 2008) and in the literature (Mynderse and Faulkner 1975, Afolayan et al. 2009). The GC-MS solution version 2.53 software (Shimadzu Corporation) was used for data processing. Antifouling assays. Mussel assay: The HF from P. brasiliense was used in assays of the response of the fouling mussel Perna perna. AF activity was measured by following the method described by Da Gama et al. (2003) with some modification.

TABLE 1. Enzymatic substrate mediums used to treat to Plocamium brasiliense tissue. (A) Complete acetoacetyl-CoA, (B) complete carnitine, (C) substrate control for acetoacetyl-CoA, (D) substrate control for carnitine, (E) primary reaction product control, and (F) final reaction product control (Curry 1987). Chemical substrates and reagents

A

B

C

D

E

F

2.0 mg
mL1 potassium ferricyanide 1.0 mg
mL1 uranyl acetate 0.8 mg
mL1 acetyl-CoA sodium salt 1.6 mg
mL1 acetoacetyl-CoA sodium salt 1.6 mg
mL1 DL carnitine 0.05 M sodium cacodylate buffer (pH 7.0)

X X X X

X X X

X X

X X

X X X

X X

X

X

X

X X X

X

X X

229

ME V A L O N A T E P A T H W A Y I N P . B R A S I L I E N S E C O R T I C A L C E L L S

Juvenile specimens of P. perna collected during low tide from the rocky coastal area of Itaipu beach (Niter oi City, Rio de Janeiro State, Brazil; 23°000 34″ S; 44°260 10″ W) were acclimated in a 2,301 recirculating laboratory aquarium (equipped with biological filtering, protein skimming and activated carbon) at a constant temperature (20°C), salinity (35) and aeration for 12 h. Treatment filters were soaked in the P. brasiliense HF fraction at physiological concentration (0.24%, DW). Next, all filter paper circles were allowed to air dry. Filter circles were then placed in the bottom of sterile polystyrene petri dishes (9 cm diameter 9 1.0 cm height) and completely filled with sterile seawater (~80 mL), and three mussel specimens (1.5–2.5 cm length) were added to each petri dish (n = 10). Antibacterial assay: The P. brasiliense HF was tested for inhibitory activity against growth of five strains of biofilmforming marine bacteria (Bressy et al. 2014) obtained from the collection of the University of Portsmouth (School of Biological Sciences): Halomonas marina (ATCC 25374), Polaribacter irgensii (ATCC 700398), Pseudoalteromonas elyakovii (ATCC 700519), Shewanella putrefaciens (ATCC 8071), and Vibrio aestuarianus (ATCC 35048). Each treatment condition and control (culture media) was replicated six times. The bacteria (2 9 108 cells
mL1) were incubated in 96 well-plates with the HFs at concentrations of 0.1, 10, 50, 125, 250, and 500 lg
mL1 in Luria Hinton Broth (LB) medium (SigmaAldrich Company) supplemented with NaCl (35 g
L1) at 30°C for 72 h (Mar echal et al. 2004). Minimum inhibitory concentrations (MICs) compared with the control were determined using the microtiter broth dilution method (Amsterdam 1996, Plouguern e et al. 2008, 2010). Antimicroalgal assay: The P. brasiliense HF was tested for inhibitory activity against five strains of marine microalgae obtained from Algobank-Caen (Universit e de Caen Basse-Normandie, France): Chlorarachnion reptans (AC132), Cylindrotheca cloisterium (AC170), Exanthemachrysis gayraliae (AC15), Navicula jeffreyi (AC181), and Chlorarachnion globosum (ATCC 8071). All microalgal cultures and assays were performed under controlled conditions (23°C  2°C under 54 lmol photons
m2
s1 from a cool-white fluorescent lamp). F/2 medium (Guillard and Ryther 1962) was used for cultivation. Microalgae for AF assays were cultivated as outlined by Tsoukatou et al. (2002). All experiments were carried out in six replicates. Briefly, 100 lL of a culture at 0.4 lg
mL1 of chl a was introduced into 96-well plates containing the fractions at concentrations of 0.1, 1, 10, 50, and 100 lg
mL1 (Plouguerne et al. 2010). After 48 h, MICs were determined by comparison of the cell growth between treatments and controls (Tsoukatou et al. 2002). Statistical analyses. A t-test for independent samples was performed to compare mussel byssal thread attachment data between filters in HF treatment and control conditions using the Statistica 8.0 software (Statsoft Inc., Tulsa, OK, USA). When normality or variance homogeneity assumptions were not met, data were transformed using the square root of X + 1 prior to t-test. Differences were considered significant whenever P < 0.05 (a = 5%). RESULTS

Optical microscopy. A large spherical and refractive organelle not previously described, here termed the mevalonosome (Mev), was observed in the CC (1 per CC, n = 60) of P. brasiliense (Fig. 1, a and b). These organelles had a diameter, circularity, and area of 10  2, 0.92  0.027, and 24.12  1.30 lm, respectively. Several contour measures revealed that

FIG. 1. Image of a longitudinal section of the living thallus from Plocamium brasiliense obtained by optical microscopy (a, b). The large spherical organelles (arrowheads) were observed in all cortical cells; bar = 10 lm.

TABLE 2. Morphometry of the cortical and medullary cells and spherical organelles of Plocamium brasiliense. Mean and SD in lm (n = 60). Cell diameter

Mean SD

Spherical organelles

Cortical (lm)

Medullary (lm)

Area (lm2)

Diameter (lm)

Circularity

22.03 4.7

99.25 14.25

24.12 1.30

10.00 2.00

0.92 0.02

these refractive organelles were circular (Fig. 1, a and b; Table 2). Transmission electron microscopy. Images obtained by TEM show the presence of a thin biofilm mainly composed of bacteria and microalgae on the P. brasiliense surface (Fig. 2a). In P. brasiliense CCs, the following cellular structures were observed: the large spherical organelles (Mev), C with electron-dense inclusions (350 nm  50, n = 60), inclusions of osmiophilic bodies (OB), pit connections (P), floridean starch grains (SG), smooth endoplasmic reticulum (SER), and small vacuoles (V; Fig. 2b). Osmiophilic vesicles were observed in association with Mev (Fig. 2c), and a large number of OB were observed forming clusters in many regions of the cytoplasm and near the CW (Fig. 2, b and e). Localization of mevalonic acid enzyme activity. The TEM images of P. brasiliense CC treated with complete acetoacetyl-CoA medium (A) indicated the presence of granular, electron-dense material within Mev (Fig. 3a), CCs in control mediums (C and E) did not present granular electron-dense material within Mev (Fig. 3b). In algae submitted to the complete carnitine medium (B), the TEM images showed the presence of electron-dense vesicles surrounding the Mev (Fig. 3, c and d) and electrondense clusters near C (Fig. 3e). In carnitine substrate control medium (D), no electron-dense material was observed (Fig. 3f). No electron-dense material was observed in TEM images of algae treated with the control mediums C and F (data not showed). Scanning electron microscopy. The SEM images of P. brasiliense cells revealed the thallus surface and the intracellular constituents of fractured CC and medullar cells (MC; Fig. 4a). The Mev was observed

230

W L A D I M I R CO S T A PA R A D A S E T A L .

FIG. 2. Transmission electron microscopy (TEM) images of Plocamium brasiliense cortical cells. (a) Bacteria and microalgae present in a microfouling community on the alga surface. Asterisks: bacteria; Arrowheads: microalgae; CW: cell walls; bar = 1 lm. (b) Common cellular structures found in cortical cells: chloroplasts (C), large spherical organelle (black asterisk), starch grains (GA), osmiophilic bodies (OB), pit connection (P), cell wall (CW), smooth endoplasmic reticulum (SER), and vacuoles (V). White asterisks show electron-dense inclusions; bar = 1 lm. (c) Osmiophilic bodies are transported from spherical organelles to the cytoplasm, where they form clusters among the organelles. Arrowheads indicate vesicle transport from large spherical organelles to the cytoplasm; bar = 1 lm. (d) SER and associated vesicles being released to the cytoplasm. Arrowheads are pointing to vesicles coming from the SER; bar = 50 nm. (e) Osmiophilic bodies accumulating in distinct regions of the cytoplasm, including near the cell wall. Arrowheads are pointing to OB clusters in the cytoplasm; bar = 1 lm.

inside CC (Fig. 4b) and was always surrounded by vesicles (Fig. 4b). More cellular contents were observed in CCs than in MCs (Fig. 4,b and c). Chemical analyses. The analysis of the 1H NMR spectra from P. brasiliense crude extract shows groups of signals characteristic of halogenated terpenes between d 7.00–5.20 ppm, d 4.60–4.50 ppm, and d 1.90–1.70 (ranges related to the olefin region, to methine groups bonded to halogen atoms and to methyl groups bonded to quaternary carbons, respectively). The crude extracts were then analyzed by GC-MS. The clear fragmentation pat-

tern for halogenated compounds observed at mass spectra of major component compounds corroborated the presence of halogenated terpenes in P. brasiliense crude extract. In light of the GC-MS data, the crude extract was fractionated and the chemical structure of the major compound comprising HF was suggested by NMR (1H, APT experiments, HSQC and HMBC) and MS analysis. Based on the comparison of our data with the literature (Mynderse and Faulkner 1975, Antunes et al. 2011) the compound was identified as pentachlorinated monoterpene (Fig. 5), previously reported in different species of the Plocamium genus (Mynderse and Faulkner 1975, Afolayan et al. 2009, Antunes et al. 2011). Characterization of the pentachlorinated monoterpene: 1H NMR (400 MHz, CDCl3) d 6.97 (3H, s, H9), 6.38 (1H, dd, 12.6, 3.6, H-5), 6.33 (1H, d, 12.6, H-6), 6.29 (1H, s, H-8), 6.08 (1H, dd, 12.0, 9.0, H2), 5.45 (1H, d, 12.0, H-1a), 5.30 (1H, d, 9.0, H-1b), 4.55 (1H, d, 3.0, H-4), 1.77 (3H, s, H-10). 13C (100 MHz, CDCl3) d 139.4(CH, C-2), 137.9 (C, C-7), 129.3 (CH, C-6), 126.4 (CH, C-5), 118.6 (CH, C-8), 115.5 (CH2, C-1), 70.8 (C, C-3), 67.9 (CH, C-4), 64.6 (CH, C-9), 24.1 (CH3, C-10). EIMS (70 eV) m/ z (rel. int.): 306/308/310/312/314M+ (