Algal biomass is a promising resource for the production of renewable fuels. Algae offer the advantages of high aerial productivity, growth in saline and brackish water, use of marginal land for cultivation, and the coproduction of high value products [1,2]. The production of biodiesel from triacylglycerols (TAGs) has been central to the efforts to commercialize algae at a very large scale. Microalgae are known to produce TAGs at up to 50% of their biomass dry weight [3] and thus have been the primary focus of most algal biofuels development and research. Macroalgae have generated much less interest as potential biofuel feedstocks because they have generally been characterized as having low-lipid productivity, often producing less than 5% lipids by dry weight. However, there are multiple examples of macroalgae with lipid production exceeding 10% of the dry biomass. These higher values for observed lipid accumulation, combined with the relative ease of cultivation and harvest of macroalgae as compared to microalgae, make them viable candidates for biofuel production and worthy of additional study in this area [4].
Accurate and repeatable methods for the extraction and quantification of lipids are key to the meaningful evaluation of biomass from diverse algae and their suitability as biodiesel feedstocks. To date, the most commonly employed protocols for lipid extraction from microalgal biomass have been the methods of Folch and of Bligh and Dyer [4][5][6][7]. However, it has been demonstrated that the use of 2-ethoxyethanol (2-EE) provides superior lipid recovery (150-200%) in microalgae when compared to these methods [8]. Here we describe methods for biomass preparation and related metrics, total lipid extraction using 2-EE, and the identification of lipid classes by thin layer chromatography (TLC). The standardized 2-EE protocol is widely used for microalgae and is adapted here for the extraction of lipids from macroalgae.
2.1 Fresh Weight (FW), Dry Weight (DW), and Ash-Free Dry Weight (AFDW) Determination
Prepare all solutions using ultrapure or double-distilled water and analytical grade reagents. Adhere to all waste disposal regulations when disposing of waste materials.
If dried tissue will be used for the lipid extraction, prepare two sets of tissue samples: one set of samples to be reserved for lipid extraction after drying and one set to determine ash-free dry weight (see Fig. 1; see Note 6). The other set will proceed to step 11 for ashing (see Fig. 1).
11. Place the crucible in a muffle furnace set to 550 C.
12. Combust the tissue sample for 2 h then allow the furnace to cool for 1 h.
13. Remove the crucible using crucible tongs and place on an insulated surface-items will be very hot. Allow the crucible and contents to cool to room temperature, place in a desiccator, and weigh once an hour until a constant weight is achieved.
14. Transfer the crucible to the analytical balance and weigh. Record the value. Subtract the weight obtained from step 1 to determine the weight of the ash.
15. Calculate the tissue DW, AFDW, % ash, and % moisture for sample Duplicate 1 (DWD1, AFDWD1, etc.) as follows:
where FWD1 ¼ fresh weight of duplicate #1 in grams (see Subheading 3.1, step 4).
DWD1 ¼ dry weight of duplicate #1 in grams.
AFDW1 ¼ ash-free dry weight of duplicate #1 in grams.
Repeat calculations for each sample Duplicate 2 (DWD2, AFDWD2, etc.).
1. Weigh 25-50 mg of dried tissue reserved in step 8 from Subheading 3.1 and transfer to a 40-mL glass vial. Alternatively, fresh biomass can be used. In which case, an amount of fresh tissue proportional to that of the dried material, based on the ratio of fresh weight to dry weight can be used (see Subheading 3.1, step 13 for dry weight calculation method).
2. Use a serological pipette to transfer 5 mL of MeOH to the vial containing harvested tissue. Cap the vial tightly and extract the tissue by heating in a 60 C water bath for 30 min.
3. Use a serological pipette to transfer 5 mL 2-EE to the vial, cap tightly, and extract lipids by heating in a 60 C water bath for 60 min. Mix by gentle vortexing for a few seconds every 5 min.
4. Transfer the liquid extract to a preweighed 100-mL RBF using a glass Pasteur pipette. To maximize recovery, wash the vial containing the residual solid biomass with acetone and methanol (1:1, v/v), to recover all of the lipids, using chlorophyll as the indicator pigment (see Note 2). Pool the washes for each sample in the 100-mL RBF containing the lipid extract.
5. Remove the solvents using the rotary evaporator at 60 C at a moderate speed. Weigh the flask periodically. The flask with residue will ultimately achieve a constant dry weight with no appreciable reduction due to loss of solvent(s). Record the weight of the RBF with lipid residue. 6. Using a micropipettor, resuspend the lipid extract in 2 mL 2:2:1:1 (hexane-toluene-acetone-methanol, v/v/v/v) with gentle swirling and transfer to a 5-mL glass vial; rinse the RBF with another 2 mL of 2:2:1:1 and pool with the previous (see Note 3).
7. Dry the RBF using the rotary evaporator at 60 C until a constant weight is achieved. Record the value.
8. Subtract the weight of the RBF minus extract from the weight of the RBF with extract to obtain the grams of TLE. Divide the grams of total lipids extracted by the resuspension volume (4 mL) to obtain the concentration of lipids in grams per liter for subsequent TLC analyses.
If beginning with dried material, calculate the % biomass extracted as total lipid as follows for sample Duplicate 1 (D1): %TLED1 of dry weight ¼ TLED1=DWD1 100% %TLED1 of ash-free dry weight ¼ TLED1=AFDWD1 100% where TLED1 ¼ total lipid extract of duplicate #1 in grams.
DWD1 ¼ dry weight of duplicate #1 in grams.
AFDW1 ¼ ash-free dry weight of duplicate #1 in grams.
Perform statistical averages, %RSD, and SD of replicate samples, D1 and D2. 9. If beginning with fresh material, calculate the moisture correction (mc) for duplicate samples D1 and D2 as follows to estimate the sample DW and AFDW: %Moisture of D1 ¼ ðFWD1 DWD1Þ=FWD1 100% %Moisture of D2 ¼ ðFWD2 DWD2Þ=FWD2 100% Average%moisture of D1&D2 ¼ ð%moisture D1 þ %moisture D2Þ=2 DWD1mc ¼ FWD1 ð100 average%moisture of D1&D2Þ=100 AFDWD1mc ¼ DWD1mc average ash weight for D1&D2 Perform statistical averages, %RSD, and SD of replicate samples, D1 and D2. FWD1 ¼ fresh weight of duplicate #1 in grams. DWD1mc ¼ moisture-corrected dry weight of duplicate #1 in grams.
DWD2mc ¼ moisture-corrected dry weight of duplicate #2 in grams.
AFDWD1mc ¼ moisture-corrected ash-free dry weight of duplicate #1 in grams.
1. Prepare a 10 cm 10 cm TLC plate; using a pencil, label the Biomass Lipids by origin, solvent front, mid-way point, and sample spots as
2. Spot 5 and 10 μL of each extract for TLC with capillary micropipettors along with 5 μL of lipid reference standards (see Fig. 2; see Note 4).
3. Add polar solvent system 65:10:20:10:3 chloroform-methanol-acetone-acetic acid-water (v/v/v/v/v) to the chromatography tank and cover (see Notes 5 and 8).
4. Carefully insert the TLC plate straight down and lean behind the ridge with the face of the plate toward the front, and then cover the tank. Develop the plate to the mid-way point, then remove the plate with flat-tipped forceps, lean against the front of the tank, and allow to dry completely (front and back should be dry).
5. Next, develop the plate with the nonpolar solvent system 80:20:1 hexane-diethyl ether-acetic acid (v/v/v) up to the solvent front, then remove the plate from tank (see Notes 5 and 8).
6. Permit to dry, then expose the TLC plate to UV light using the handwand at each wavelength to examine absorption and fluorescence properties. Take images of the plate in visible light and with UV.
7. Place the TLC plate in the chamber containing iodine-silica 1:1 (w/w) and seal. Expose for 2-3 min; lipids will stain darker with increased exposure time. 7. Use a dull #2 pencil to lightly draw a line at the origin, a line at the solvent front, a dash at the mid-way point, small dots where spots are to be applied, and sample labels. Take care not to break the surface of the silica gel. A 1-cm rule should be applied-the origin and solvent front are 1-cm from the bottom and top of the plate, respectively, and sample spots should be 1 cm from the edges and 1 cm between samples.
8. Do not exceed 0.8 cm in depth (usually just below the ridge in standard tanks) to avoid loss of sample. Equilibrate the chromatography tank with developing solvent for at least 15 min prior to developing the plate.