Disruption-free solid-phase extraction of surface metabolites from macroalgae

The surface chemistry of aquatic organisms is decisive for their biotic interactions. Metabolites in the spatially limited laminar boundary layer mediate processes, such as fouling, allelopathy and chemical defense against herbivores. However, very few methods are available for the investigation of such surface metabolites. Here we give a detailed protocol in which surfaces are extracted by means of C18 solid phase material, elution of the solid phase extraction material with solvent and analysis via liquid chromatography / mass spectrometry (LC/MS) and/or gas chromatography / mass spectrometry (GC/MS). The protocol introduced here is based on a previous publication (Cirri et al. 2016) where validation is described. The method is robust, picks up metabolites of a broad polarity range and is easy to handle. It was developed for the macroalgae Fucus vesiculosus, Caulerpa taxifolia and Gracilaria vermiculophylla, but can be easily transferred to other algae and to other aquatic organisms in general.


Introduction
Surface metabolites play a fundamental role in the mediation of interactions on biotic surfaces of e.g., macroalgae, corals or sponges. Such compounds control settling processes, regulate predator / prey relationships and mediate infection processes (da Gama et al. 2014, Dobretsov et al. 2013, Wahl 2009. A hallmark of such interactions is the locally much focused action of the compounds in question (Dworjanyn et al. 1999, Dworjanyn et al. 2006. The surface concentration of such metabolites often exceeds the concentration in the medium, but it can even exceed concentrations within the tissue since exuded compounds can accumulate in a diffusion limited laminar boundary layer.
Despite their ecological importance, until now, only a few methods allow the estimation of local concentrations of surface metabolites. As a consequence, many investigations on the effect of surface metabolites were based on bioassays with extracts of whole organisms (see, e.g., Hellio et al. 2000). Such experiments do not reflect the real ecological relevance of surface active substances because only metabolites at the surface or near a producer should be considered (Nylund et al. 2007). The determination of metabolites within the laminar boundary layer around an aquatic organism, a thin film of about 100-200 µm that determines the transition between the surface and the surrounding water, is thus crucial for experiment planning and evaluation. So far the laminar boundary layer has been studied to determine uptake rates of nutrients (Wheeler 1980) or to model them (Hadley 2014), or it has been investigated in correlation with climate changes and oceans' acidification (Cornwall 2014), but the study of metabolites identity and concentration at macroalgal surface is crucial both from an ecological and an industrial point of view, especially in the framework of antifouling substances research (Bhadury 2004, Rajan 2016.

State of the Art
Several methods to study the laminar boundary layer have been established during the last 20 years: these include Raman micro spectroscopy (Grosser, et al. 2012), mass spectrometry imaging, like MALDI and DESi (Slaveykova 2009, Andras 2012) and the widely used dipping methods (de Nys et al. 1998, Lachnit et al. 2010. For extraction, algae are immersed in a solvent for a short period, during which the metabolites are partially extracted from the surface. Care has to be taken that the solvent does not damage the cells of the extracted organism. After concentration in vacuum, the extracts can be submitted to analytical methods, such as GC-MS and LC-MS. Dipping methods are really easy to handle and can be used with a lot of different species of macroalgae. By optimizing the solvent (or mixtures of solvents) for extractions, as well as the solvent's volume, these technique allows extracting very different kind of metabolites, proving to be flexible and adaptable to specific questions (see Chapter 19 by Weinberger). Although useful, dipping methods could also be problematic since solvent exposure can cause cell lysis and thereby contamination of the surface extract with intracellular metabolites; these problems depend both on the algal species and on the physiological conditions, as well as on the part of the alga that needs to be extracted. Some algae only tolerate exposure to rather nonpolar solvents such as hexane for few seconds. However, these solvents only cover a very limited range of unpolar metabolites and do not penetrate surface associated water. If solvent mixtures containing methanol are employed, massive damage of the algae could be observed, thereby questioning the validity of results. To overcome these limitations, we developed a new, non-destructive solvent-free and universal method for extracting secondary metabolites from marine macroorganisms (Cirri et al. 2016). The method is based on the adsorption of organic metabolites onto Evans blue staining by red/green ratio analysis at 5, 30, 60, 120, 300, 600 s exposure to C18 material (gray), hexane/methanol dipping (white) and control (black), (n = 5 ± SD).
The technique has been optimized regarding recovery, reproducibility, and ease of use with the brown macro alga Fucus vesiculosus as a model organism. F. vesiculosus is a common, well studied brown alga that can be found on the coasts of the North Sea, the western Baltic Sea, and the Atlantic and Pacific Oceans. However, also the green alga Caulerpa taxifolia and the red alga Gracilaria vermiculophylla were extracted for proof of concept, demonstrating the universality of the method that can be potentially used for all aquatic macroscopic organisms. Here a detailed commented protocol is given, based on our publication Cirri et al. 2016.

Fucus vesiculosus
1. Fucus vesiculosus samples can be collected independently of the season. The alga can be extracted directly after collection in the field or can be kept in aquaria as described below.
2. Artificial sea water (ASW): 33 g of Instant Ocean™ (Aquarium Systems, France) per liter of deionized water is stirred at least for 12 h to accomplish a complete dissolution. In our experiment, ASW was diluted with deionized water to half of the initial concentration to reproduce the salinity of the Baltic See.
6. Air pump for constant ventilation of aquaria.

4 ml glass vials with a black cap.
14. 1.5 ml glass vials with a blue cap.

Experimental procedures
The following operations (Figure 20

Extraction procedure
1. Spread C18 SPE material (Notes 1, 2) on the small thecae of the petri dish and weight it. For a Petri dish with a diameter of 9.2cm (92 mm × 10 mm), an amount of 0.5 g of SPE material is sufficient to reach uniform covering of the plate (Note 3).

Weight the empty polypropylene cartridge (Note 2).
3. Take the piece of alga that you desire to extract out of the aquarium (in this case, the fronds of F. vesiculosus). Pieces of around 40 cm 2 of this alga were 7 small enough to fit in a 9.2 cm diameter Petri dish and sufficient for generating surface extracts that can be investigated in GC/MS and LC/MS.

Hold the alga for 2 minutes so that excess of water can drop off (Note 4).
5. Place the alga inside the dish, close it and shake it manually for ca 10 s, taking care to obtain a homogenous cover of the surface with SPE material.
6. Leave the alga in the Petri dish for 60 s (Note 5).
7. Take the alga out of the Petri dish using forceps, shaking it gently to remove excess SPE material.  10. Wash the funnel and the cartridge with the SPE material with excess (ca. 20 ml) MilliQ water to remove salt from the seawater/medium, taking care that all SPE material is going into the cartridge. While washing, you can apply a gentle vacuum (≈ 550 mbar) to make the powder settle in the cartridge (0.5-1 mL bed volume) (Note 7).

11.
Add an appropriate internal standard for recovery calculations 12.
Elute the compounds with methanol (3 times 0.5 mL) into a four mL glass vial under ambient pressure. Optimal flow rate should be maximum one drop/second (Note 8).

13.
Remove the solvent under a stream of nitrogen and re-dissolve the sample in 100 µL of methanol (Notes 9, 10). Transfer the sample into a 250 µL glass insert placed in a 1.5 mL glass vial (or other vials suitable for the autosampler of your own GC or HPLC). where TP is the thallus projection.

16.
After the C18 material is completely dry, weight the cartridge to calculate the amount of extraction powder used for the experiment.

Data evaluation
17. After LC-MS or GC-MS measurements, integrate both the chromatographic peak area of the internal standard and the substance(s) of interested with the appropriate software associated with your analytical instrument (e.g., Waters ® Masslynx or Thermo ® Xcalibur).
18. Calculate the ratio between the chromatographic peak area of the internal standard and the substance(s) of interested for a relative quantification. For absolute quantification, the ratio should be compared to an external calibration curve.
19. Normalize the quantity (relative or absolute) to the surface area previously calculated with ImageJ. 9 20. For recovery calculation, normalize the chromatographic peak area of the internal standard (in our experiment, canthaxanthin) to the weight of the C18 material effectively extracted. Note 4. It is essential that the surface is not getting completely dry: the thin layer of water that remains on the surface (laminar boundary layer) is the environment from which metabolites shall be determined.

Notes
Note 5. Different extraction times were tested, and 60s was chosen as the best compromise between a good interaction and absorption of metabolites with C18 material and an easy recovery of the powder from the alga.
Note 6. The amount of water used for the washing steps is not a crucial parameter, but for more precise absolute quantification of metabolites, this step can also be easily standardized, always using the same amount of water.
Note 7. Different vacuum setups could be used, as a vacuum manifold (Visiprep™, Supelco, USA) for multiple, simultaneous extractions. Anyway, the use of a vacuum pump is not necessary. In case the vacuum is not applicable, the settlement of the SPE material will require longer time and a larger amount of water to make sure that the absorbing material is properly packed.
Note 8. Our method was optimized for 1.5 mL of methanol as an extraction solvent, but as in all SPE methods, the amount and the polarity of the solvent (or mixture solvents) can be adapted. Other solvents can be applied depending on the used solid phase and the nature of the metabolites which should be extracted. C18 material does not need to be dry before elution; however, each material has its specific treatment to be fully effective. For all these information, also check the recommendations of the manufacturer.
Note 9. If needed or desired, the sample can be stored at this point at -20 °C, and the drying step could be done afterward Note 10. The volume of re-dissolution should be adapted to some surface metabolites present and the sensitivity of the analytical instrument to obtain a response in the linear range. For GC-MS analysis of polar compounds, a derivatization protocol can be used starting from this step. For more information about derivatization, see chapter 18 by Kuhlisch et al.) and Vidoudez and Pohnert (2011).