Cultivation of Green Microalgae in Bubble Column Photobioreactors and an Assay for Neutral Lipids

1. Setup of Bubble Column Photobioreactors

1. Construct a set of vented lids from the plastic lids that came with the 1 L glass bottles and hybridization tubes (see Figure 1 for schematic and photos). Construct lids for the humidifier, mixing trap, each air lift photobioreactor, and each bottle reactor.
   1. Drill ¼” holes in the lid: 2 holes are needed for bioreactor and humidifier lids; 3 holes are needed for the mixing trap.
   2. Slip a ¼” O-ring over the threads of a 1/8” panel mount Luer fitting, and slide this into the ¼” hole drilled in the lid (Figure 1A).
   3. Slip a second ¼” O-ring over the threads so that the lid is sandwiched between the two O-rings. Slip a locknut onto the threads and tighten to fix the panel mount Luer in place.
   4. Snap lock rings onto the exposed male Luer projecting from the lid. Repeat steps 1.1.2-1.1.4 for each hole in the lid.
   5. For lids that will be used on bubble column and bottle reactors, attach 1/8” female Luer to barb fittings to 1.5” pieces of 1/8” ID PVC tubing. Attach these to each of the exposed male Luer fittings on the lid.
   6. Connect a check valve (pointing away from the lid) to the free end of one of the 1/8” pieces.
      NOTE: This will serve as the exhaust port for the bioreactor.
   7. Connect a male Luer to barb fitting to the second piece of 1/8” tubing projecting from the lid. Click the rotating lock ring into place and fasten a 0.2 μm air filter to this.
      NOTE: This will serve as the inlet port for the reactor.

Figure 1. Schematic and photos for constructing bioreactors. (A) Schematic for construction of the bioreactor lids (B) photo of the assembled bioreactor lid, and (C) photo of the assembled lid used for the humidifier. Note that the humidifier fittings should be coated in water-proof silicone to ensure an airtight seal with the lid. Please click here to view a larger version of this figure.

2. Assemble the air delivery system (see Figure 2A and 2B for a schematic and photo).
   1. Attach 1/8” NPT thread to barb fittings to the inlets and outlets on the back of each rotameter.
      NOTE: 200-2,500 cm³/min rotameters are for air pressure modulation upstream of the humidifier, 100-1,000 cm³/min rotameters are for bottle reactors, 50-500 cm³/min rotameters are for air lift bioreactors, and 5-50 cm³/min rotemeters are for CO₂ flow regulation. It is recommended to mount rotameters to a fixed surface (e.g. plastic sheet) so they do not fall over during operation.
   2. Shut off the compressed air source, then connect ¼” ID flexible PVC tubing to the compressed air source with a hose clamp. Step down the hose diameter to 1/8” ID PVC flexible tubing using a ¼” female to barb fitting and a 1/8” male to barb fitting.
   3. Connect the free end of the 1/8” ID tubing to the inlet of a 200-2,500 cm³/min rotameter.
      NOTE: The outlet of this rotameter will feed the humidifier bottle via 1/8” ID tubing.
   4. Connect the 1/8” tubing to an inlet to a vented lid (use a female Luer to barb fitting to make the connection). Then connect a second piece of 1/8” tubing to the inside of the panel mount fitting.
      NOTE: This piece will hang down into the humidifier and bubble air through the water.
   5. Attach 1/8” female Luer to barb fittings to each end of a piece of 1/8” ID tubing and use this piece to connect the outlet of the humidifier to the inlet of the mixing trap.
   6. In the same manner as 1.2.5, connect the outlet of the CO₂ regulator to a second port on the mixing trap.
   7. Construct a manifold using 1/8” tubing and 1/8” multiport barb (see Figure 2C) to feed air into the rotameter banks.
      NOTE: These rotameters will be used to supply the bioreactors. Avoid constructing more than 6 rotameters in series. Instead, use parallel banks of rotameters to expand the system. Ensure that the total flow demand for all reactors is less than 2,500 cm³/min (or else a larger rotameter will be needed upstream of the humidifier).
   8. Connect the outlet (3^rd port) of the mixing trap to the newly constructed rotameter banks using 1/8” tubing and a 1/8” female to barb Luer.
   9. Connect sufficiently long 1/8” tubing to the outlets of each rotameter in the rotameter bank to supply air to the bioreactors. Label the ends of the tubing as well as the rotameters in the bank.
   10. Apply water-proof silicone around all ports on the humidifier and mixing trap lids to ensure they are air tight.

Figure 2. Schematic and photos for assembling bubble column system. (A) Schematic of the aeration system (B) photo of the humidifier, mixing trap, and rotameter bank, and (C) photo of the manifolds used to connect the rotameter banks together. Please click here to view a larger version of this figure.

3. Set up fish tanks, stir plates, and lights (Figure 3).
   CAUTION: This system requires a large number of outlets and sufficient circuit capacity to support all components. Avoid stringing together multiple power strips and extension cords in a daisy-chain fashion because this is an electrical hazard. Use of GFI type outlets and power strips is highly encouraged due to the presence of water in the system.
   1. Arrange the low-profile magnetic stirrers on a level surface that is strong enough to hold the weight of water-filled fish tanks.
   2. Place small wooden or plastic blocks (that are slightly taller than the stir plates) around the perimeter of the stir plates to support the weight of the fish tanks.
      CAUTION: Avoid placing the fish tanks directly on the stir plates as the weight will crush them.
   3. Place the fish tanks over the stir plates and support blocks and fill the tanks with water.
   4. Cut a piece of a stiff plastic sheet to fit on top of the fish tank as a lid. Cut holes in this cover to slide the hybridization tubes in and out. Also cut a hole for the fish tank heater.
   5. Arrange the fluorescent light banks next to the fish tank to provide horizontal illumination of the bioreactors. Plug the light bank into a light timer to set a day/night cycle.

Figure 3. System schematic for the bottle bioreactors (left) and the bubble column photobioreactors (right). This figure has been modified from Higgins et al.¹⁷. Please click here to view a larger version of this figure.

2. Preparation of Microalgae Inoculum

1. Obtain microalgae inoculum from a cryo-preserved, plated, or liquid culture.
   NOTE: It is recommended that cryopreserved organisms be plated prior to the use as inoculum to ensure that cells are viable and that the resulting culture is axenic. Agar medium (e.g., ATCC #5 sporulating agar)²¹ is a rich medium that works well for reviving species of Chlorella and Auxenochlorella from cryo-storage.
2. Prepare 2.4 L of mineral medium that is appropriate for the particular microalgae species.
   NOTE: Examples include N8 medium²³ for species of Chlorella, N8-NH₄ medium²¹ for species of Auxenochlorella. Use of a medium appropriate for the algae strain is one of the most important steps toward ensuring robust algae growth.
3. Aliquot 2.4 L of mineral medium equally into three 1 L glass bottles, add stir bars to each bottle and assemble the vented lids (Figure 1) for each bottle. Double check that the aeration tube is on the inlet side and each bottle has a stir bar in it.
4. Autoclave the stock bottles using a liquid sterilization cycle (121 °C) for 30 min. Autoclave 100 mL of deionized water (dH₂O) and some 1.5 mL tubes at the same time, which will later be used for plating. Allow the medium to cool overnight. Alternatively, cool the reactor to room temperature and then aerate for 2 h prior to inoculation.
5. In a biosafety cabinet (BSC), inoculate microalgae from a plate or axenic liquid culture into the stock bottles. Use sterile technique to maintain axenic cultures in the following steps.
   1. Add 20 mL of autoclaved dH₂O to a sterile 50 mL centrifuge tube. Use a sterile 10 µL disposable loop to pick several single colonies from the plate from step 2.1. Dip the loop into the 50 mL tube and wash the algae cells into the 20 mL of autoclaved dH₂O. Shake the 50 mL tube to make a homogenous microalgae solution.
   2. Pipette 6 mL of microalgae solution into each stock bottle with a 10 mL sterile serological pipette. Swirl the bottle to mix the microalgae evenly into the medium.
6. Use a 2 mL sterile serological pipette to draw 1 mL samples from each stock bottle and transfer into sterile 1.5 mL tubes.
   NOTE: Micropipettes are not recommended for this step due to the risk of contamination. Tighten the vented lids on stock bottles.
7. Place the stock bottles on stir plates (~150 rpm) and adjust air flow rate, CO₂, and lighting levels as appropriate for the species. Rotate the stock bottle position every day.
8. Dilute the 1 mL samples obtained during step 2.6 (100-fold dilution in sterile water usually works well) and spread plate onto a rich agar medium.
   NOTE: These plates can be used to check for instances of contamination as well as serve as a source of future algae inoculum for further experiments.
9. Take samples from the bottles (in the BSC) every two days to check the microalgae growth.
   Place samples in a 96-well microplate in triplicate (200 µL) and measure optical density (OD) at 550 nm and 680 nm every two days until OD reaches 0.2-0.3 (which typically requires 5-7 days).
10. Stop the incubation and place the stock bottles on a bench for 24-48 h to allow the algae cells to settle by gravity.
    NOTE: The settled cells will be used next to inoculate the bubble column photobioreactors. If a more rapid cell collection is desired, cells may be centrifuged at no more than 1,000 x g to collect cells.

3. Cultivation of Microalgae in Bubble Column Photobioreactors

1. The day before bioreactor inoculation, prepare appropriate media and transfer 200 mL (or desired volume) to the bubble column photobioreactor tubes (hybridization tubes). Autoclave tubes with media and vented lids in place.
   NOTE: If using wastewater as a growth medium, autoclave the empty bioreactors and add sterile-filtered wastewater (if axenic culture is desired).
2. Concentrate the settled microalgae stock by removing the supernatant using a vacuum pump. Leave less than 100 mL of medium in each bottle but avoid removing settled algae.
   NOTE: Conduct this procedure inside a BSC and follow sterile technique. A simple vacuum apparatus can be constructed using either a vacuum flask or bottle. Fit a sterile serological pipette onto the end of the tubing.
3. Suspend and transfer the algae slurry to sterile 50 mL centrifuge tubes. Centrifuge at 1,000 x g for 5 min to further concentrate algae.
4. In the BSC, remove enough supernatant to achieve a total volume of ~80 mL of algae concentrates for 12 photobioreacters. Avoid vacuuming out the pellet. Transfer the algae concentrate to a sterile container (or the used algae stock bottle).
5. Add 6 mL of algae slurry into each photobioreactor with a sterile 10 mL serological pipette.
6. Sterile filter (0.2 μm syringe or vacuum filter) and add appropriate amounts of any other compounds (e.g. vitamin stocks) which cannot be autoclaved.
7. Swirl bioreactors to mix algae into the medium.
8. Draw a 2 mL sample from each bioreactor using a serological pipette and transfer to a 2 mL tube. Collect a 2 mL sample (in a BSC) every 24 h to monitor culture progress. Check the sample for pH using test strips and adjust the reactor as needed with either 3 M NaOH or 3 M HCl.
9. Tighten the bioreactor lids and place all bioreactors into the fish tank water bath. Adjust the aeration, CO₂, and lighting to the appropriate level for the species. Rotate the bioreactor position each day after sampling (Step 3.8).
10. Apply 200 µL of each culture sample in triplicate to wells of a 96 well microplate. Measure optical density (OD) at 550 nm and 680 nm.
    1. On the last day of the culture period, measure OD under different dilution factors (e.g., a 1x, 2x, 4x, 8x, 16x and 32x) to establish a correlation between the OD and the actual dry weight after harvest (step 4).
11. Centrifuge the 2 mL sample tube at 12,000 x g for 5 min.
12. Filter the supernatant through 0.2 µm non-sterile syringe filter and store the supernatant (and pellet if needed) at no higher than -20 °C for long term storage and later analyses of changes in media composition.

4. Harvest and Freeze Drying of Microalgal Biomass

1. Measure a fixed volume of algae culture from each bioreactor with a graduated cylinder (e.g., 160 mL from a bioreactor that originally contained 200 mL of medium) and transfer into centrifuge bottles. Rinse the graduated cylinder with dH₂O in between each measurement.
2. Centrifuge at 4,696 x g for 5 min. Discard the supernatant by carefully vacuuming it out.
3. Transfer the pellets to labeled 50 mL tubes. Rinse the centrifuge bottles with dH₂O, and transfer contents to the 50 mL tubes. Ensure the total tube volume does not exceed 45 mL.
4. Wash the algae pellets with dH₂O to remove salts.
   1. Centrifuge the 50 mL tubes at 4,696 x g for 5 min and discard the supernatant.
   2. Add 40 mL dH₂O to each 50 mL tube; vortex to mix. Centrifuge again at 4,696 x g for 5 min and discard supernatant.
   3. Repeat step 4.4.2 again.
5. Label and weigh empty 15 mL centrifuge tubes on a 4-decimal balance (label both the lid and tube and weigh them together). Weigh one tube per algae culture. Weigh each 15 mL tube twice to minimize error.
6. After the last wash, discard the supernatant, and add 7.5 mL of dH₂O to each 50 mL tube. Vortex and transfer the algae slurries into the pre-weighed 15 mL tubes. Rinse the 50 mL tubes with additional dH₂O and transfer liquid to the 15 mL tubes. Avoid exceeding 12 mL of total volume in the 15 mL tubes.
7. Centrifuge the 15 mL tubes at 4,696 x g for 5 min and decant the supernatant. Freeze the tubes with pellets at -80 °C for at least 30 min in preparation for freeze-drying.
8. Freeze dry overnight or until dried.
9. Weigh and record the freeze dried 15 mL tubes with algae.

5. Lipid Extraction using a Modified Folch method²⁴

1. Weigh out 20 mg of freeze-dried algal biomass into a 2 mL screw cap polypropylene tube (check manufacturer label to ensure product is suitable for bead extractions).
2. Add 1.5 mL of Folch solvent (2:1 chloroform/methanol) to each 2 mL tube (which contains 20 mg of freeze-dried algae). Pour ~0.5 mL zirconia/silica beads (0.5 mm) into each tube until liquid level in tube reaches 2 mL.
   CAUTION: Handle chloroform and methanol in a fume hood and avoid breathing fumes or skin contact.
3. Homogenize the algae samples in a bead mill for 20 s at a speed of 6.5 m/s. Transfer tubes to ice for 30 s to chill samples. Repeat five more times to fully extract lipids.
4. Filter the homogenate through a 5 mL syringe containing a stainless-steel wire mesh disk (#60 mesh) to strain out the beads, collecting filtrate in a 15 mL tube.
5. Wash the beads with 1.5 mL of Folch solvent, pushing liquid through with the syringe as necessary. Repeat this wash two more times and collect all filtrate in the 15 mL tube, yielding a final volume of approximately 6 mL.
6. Add 1.2 mL of 0.9% (w/v) NaCl solution to the Folch extract in the 15 mL tube and vortex to mix well.
   NOTE: If necessary, more Folch solvent can be used to wash beads (use 0.2x the total wash volume of 0.9% NaCl solution to induce phase separation).
7. Centrifuge the 15 mL tubes at 6,000 x g for 5 min. Record the bottom chloroform (green) phase volume to the nearest 0.1 mL using lines on the side of the 15 mL tube. Transfer the bottom phase to a glass vial (with the lid) using a glass Pasteur pipette.
8. Store the lipid at -20 °C or (-80 °C if there are plans to use this extract for fatty acid analysis).

6. Neutral Lipid Assay using a Microplate Method (adapted from Higgins et al. 2014²²)

1. Prepare stock solutions. Prepare 10 mL 1 mg/mL vegetable oil standard in chloroform and store at -20 °C.
   NOTE: Any vegetable oil may be used in this assay because it is not sensitive to the types of fatty acids. Prepare 10 mL of 200 µg/mL Nile Red solution in dimethyl sulfoxide (DMSO) and store in the dark at room temperature.
2. Pre-heat dry microplate block to 55 °C in a fume hood. While this is heating, dilute the lipid extracts and vegetable oil standard 3-fold with methanol.
   NOTE: This dilution can be altered based on the lipid content of the algae, but this level works well for most Chlorella.
3. For each diluted sample, add 80 µL to a 96 well polypropylene microplate in quadruplicate.
   CAUTION: Use of polystyrene plasticware is not recommended for use with organic solvents.
4. For the solvent blank, apply 80 µL of 2:1 methanol/chloroform in quadruplicate. For standards, add 10, 30, 60, 90, and 120 µL of the diluted vegetable oil standard in quadruplicate.
5. Place the microplate in a dry block heater at 55 °C for 20-30 min until all solvent has evaporated. While the solvent evaporates, prepare the working Nile Red solution (need 200 µL of 1 µg/ml solution per plate well). As an example, twelve samples and a full set of standards requires 16 mL of 1.0 µg/mL solution; prepare by dissolving 80 µL of the 200 µg/mL stock (in DMSO) into 16 mL dH₂O.
6. Remove the microplate from the heating block and let cool to room temperature. Add 30 µL of isopropyl alcohol to each well and mix by pipetting up and down. Ensure all pipette channels are mixing the solution and resuspending the lipids, yielding a homogeneous green liquid.
7. Add 200 µL of Nile Red solution (1 µg/mL) to each well, pipette up/down 10 times to mix. Incubate the plate for 5 min at room temperature. While waiting, prepare a 50% bleach solution by mixing bleach (6% hypochlorite) with dH₂O. 20 µL per well are needed. Preparation of 3 mL of 50% bleach is sufficient for 12 samples and a full set of standards.
8. Add 20 µL of bleach solution to each microplate well and pipette up and down 5 times to mix well. Incubate 30 min at room temperature.
9. After 30 min, read fluorescence in the samples every 5-10 min at 530 nm excitation/575 nm emission with auto cutoff set to 570 nm until the signal from algae samples stabilizes. Typically, 60 min of total incubation is sufficient.
10. Create a calibration curve for the vegetable oil standards (in the range of 0-40 ng oil/well).
    NOTE: A linear fit works well for low (<30 ng/well) oil concentrations and a polynomial fit may be used if the standard exceeds 30 ng/well. Use this correlation to quantify the neutral lipid in the sample wells.