Ectocarpus siliculosus (Dilwyn) Lyngbye (Ectocarpales, Phaeophyceae) unialgal strain 32 (accession CCAP 1310/4, origin San Juan de Marcona, Peru, 2002) was cultivated in 10 L plastic flasks in a culture room at 14 °C, using filtered and autoclaved seawater (SW) with a salinity of 33 psu; SW was enriched in Provasoli nutrients (Starr & Zeikus 1993), that is 0.8 mm nitrate, 50 µm phosphate, 0.45 mm boric acid, 6 µm iron, 18 µm manganese, 1.9 µm zinc, 0.48 µm cobalt, 1.5 nm vitamin B12, 0.3 nm thiamine, 4 nm biotin, 0.8 mm Tris, 75 µm EDTA (all concentrations are the final concentrations in the medium), and adjusted to a pH of 8. These nutrients are adequate to grow E. siliculosus even in otherwise nutrient-free artificial SW. PAR (400–700 nm) was provided by Philips daylight fluorescence tubes at a photon flux density (PFD) of 40 µmol m−2 s−1 for 14 h per day, a PFD that was sufficient for cultures to grow well, but low enough not to produce high light stress. Cultures were aerated with filtered (0.22 µm) compressed air. One week before the experiment, several thalli per condition, time point, and replicate (three replicates per condition, about 0.5 g fresh weight each) were transferred to individual Petri dishes containing 100 mL of fresh culture medium and maintained as described earlier, but without aeration.
Stress experiments were started about 30 min after the beginning of the light phase by replacing the culture media with pre-prepared stress media. To be able to compare this study with a previous transcriptomic study (Dittami et al. 2009), identical stress conditions were chosen. For hyposaline stress, 33 psu SW was diluted to 12.5% of its original concentration with distilled water (i.e. final salinity: 4 psu). For hypersaline stress, 60 g NaCl were added per liter of 33 psu SW (i.e. final salinity: 93 psu). For oxidative stress, H2O2 (Sigma-Aldrich, St. Louis, MO, USA) was added immediately before beginning the stress experiment at a final concentration of 1 mm. For the control condition, 33 psu SW was used. Identical final quantities of Provasoli nutrients were added to each of these media. All three tested stress conditions were previously shown to be sub-lethal for the examined strain of E. siliculosus (Dittami et al. 2009), as it was able to survive indefinitely in stress media and even resumed growth (data not shown). However, the treatments reduced its photosynthetic quantum yield (measured by pulse amplitude modulation fluorometry) by 50–70% after 6 and 24 h stress exposure. In addition, after restoring control conditions for algae submitted to saline stresses, it took cultures between 3 and 6 d to completely recover their original photosynthetic quantum yield (Dittami et al. 2009). This was also true for the H2O2 treatments, although H2O2 concentrations in the medium probably decreased significantly throughout the experiment.
Transcriptomic changes were monitored after 6 h in our previous study (Dittami et al. 2009), as this corresponds to the time interval between high and low tide. For metabolite profiling, two time points (6 h and 24 h) were examined as transcriptomic changes might also translate into variations of metabolites later. Cultures were harvested by concentrating the culture on 40 µm nylon mesh filters (Cell Strainer, BD, NJ, USA), quickly drying the algae with a paper towel, and immediately freezing them in liquid nitrogen. This whole process did not take more than 1 min.
For the analysis of amino acids, carbohydrates, polyols, organic acids and urea, samples were ground in liquid nitrogen and freeze-dried. Amino acids, non-structural carbohydrates and organic acids were extracted and quantified as follows: between 10.0 and 12.0 mg of freeze-dried sample, corresponding to approximately 50 mg fresh weight, were weighed and ground using a ball mill. The powder was suspended in 400 µL of a methanolic solution containing 100 µm of DL-3-aminobutyric acid (i.e. β-aminobutyric acid) and 200 µm ribitol, followed by 15 min of agitation at room temperature. Then, 200 µL of chloroform were added, followed by a 5 min agitation step. Finally, 400 µL of water were added, and samples were vortexed vigorously before centrifugation at 13 000 g for 5 min. Fifty µL and 200 µL aliquots of the upper phase, which contained polar metabolites including amino acids, polyols and carbohydrates, were transferred to clean vials and vacuum-dried for subsequent chromatographic analysis.
For amino acid profiling, the vacuum-dried polar phase aliquots were resuspended in 50 µL of ultra-pure water, and 10 µL was used for the derivatization using the AccQ-Tag Ultra derivatization kit (Waters, Milford, MA, USA). Derivatized amino acids were analysed using an Acquity UPLC system (Waters) according to Jubault et al. (2008), using DL-3-aminobutyric acid as internal standard. Peaks were identified according to their retention time by comparison with commercial standards. When changing the elution gradient, all of the identified peaks in the E. siliculosus samples showed identical shifts in retention time compared with the standard (data not shown), further confirming their identity.
For gas chromatography/mass spectrometry (GC-MS) profiling of non-structural carbohydrates and organic acids, the 200 µL vacuum-dried polar phase aliquots were resuspended in 40 µL of 20 g L−1 methoxyamine-hydrochloride (Sigma-Aldrich, St. Louis, MO, USA) in pyridine before incubation under orbital shaking at 30 °C for 90 min. After addition of one volume of N,O-bis(trimethylsisyl)trifluoroacetamide (Sigma-Aldrich), samples were incubated 37 min at 30 °C, transferred to glass vials and incubated at room temperature overnight before injection. GC-MS analysis was performed according to Roessner et al. (2001). The GC-MS system consisted of a TriPlus autosampler, a Trace GC Ultra chromatograph and a Trace DSQII quadrupole mass spectrometer (Thermo Fischer Scientific Inc, Waltham, MA, USA). Chromatograms were deconvoluted using the AMDIS software v2.65 (http://chemdata.nist.gov/mass-spc/amdis/) associated with NIST libraries. Metabolite levels were quantified using ribitol as internal standard and by comparison with individual external standards.
In addition, mannitol concentrations were confirmed by an alternative method, based on an extraction in 1 mL of 70% aqueous ethanol (v/v) at 70 °C for 4 h according to Karsten et al. (1991). After centrifugation for 5 min at 5000 g, 700 µL of the supernatant were evaporated to dryness under vacuum, re-dissolved in the same volume of distilled water and analysed with an isocratic Agilent HPLC system equipped with a differential refractive index detector, a Bio-Rad resin-based column (Aminex Fast Carbohydrate Analysis, Bio-Rad, Hercules, CA, USA; 100 × 7.8 mm) and a Phenomenex Carbo-Pb2+ (4 × 3 mm) guard cartridge. Mannitol was eluted with water at a flow rate of 1 mL min−1 at 70 °C, identified by comparison of the retention time with that of a commercial standard prepared as 1 mm aqueous solution and quantified by peak area.
Urea was determined following the protocol outlined by Beale & Croft (1961). Thirty milligrams [dry weight (DW)] of sample were resuspended in 500 µL of zinc/sodium sulphate solution (9 mm ZnSO4, 84 mm Na2SO4). Then 7.5 µL of 1 m NaOH were added, and the mixture was thoroughly vortexed for 2 min before centrifugation for 10 min at 20 000 g. Two 120 µL-samples of the supernatant (technical replicates) were each mixed with 1 volume of DAM-PAA reagent (1 volume of 1% w/v diacetylmonoxime in 0.02% acetic acid, 1 volume of phenylanthranilic acid in 20% v/v ethanol with 120 mm NaCO3). One mL of activated acid phosphate (1.3 m NaH2PO4, 10 mm MnCl2, 0.4 mm NaNO3, 0.2 m HCl, in 31% v/v H2SO4) was added before incubation in boiling water for 12 min. The tubes were left to cool at room temperature and centrifuged for 1 min at 20 000 g before measuring absorption at 535 nm using a spectrophotometer. A standard curve was created using 0, 50, 100, 250 nmol urea dissolved in 10 µL of distilled water instead of the freeze-dried samples. Some Ectocarpus samples spiked with 50, 100 and 250 nmol urea were also included as positive controls to test for possible urease activity in the extract.
For the measurement of total fatty acids, approximately 400 mg (fresh weight) of sample were ground in liquid nitrogen and extracted with 2 mL of ethyl acetate as previously described (Küpper et al. 2006). As an internal standard, 250 ng of 12-OH-lauric acid were added. Extracts were evaporated under a stream of nitrogen, resuspended in 1 mL of BF3 10% in methanol for esterification and resuspended in 100 µL hexane before GC-MS analysis as described in Le Quéréet al. (2004). Fatty acids were measured only for the 6 h data point.
All metabolite concentrations were calculated per g DW, except fatty acids, which were determined as percentage of total fatty acids. As E. siliculosus is known to accumulate high concentrations of NaCl in response to salt stress, dry weights were corrected for the quantity of NaCl in each sample. This quantity was calculated based on the ratio dry weight to fresh weight (on average 20.1%) and the intracellular Na+ concentrations determined by flame photometry as described in our previous study (Dittami et al. 2009). We assumed that changes in Na+ also correspond to changes in Cl-, as this is the predominant anion in E. siliculosus, about five times more abundant than nitrate (Dittami et al. unpublished data). All analyses were performed both with and without this correction. Although the observed differences between the two analyses were only quantitative and not qualitative, we chose to show the corrected data, as it is biologically more relevant.