A Simple and Rapid Protocol for Measuring Neutral Lipids in Algal Cells Using Fluorescence

The overall goal of the following experiment is to measure the neutral lipid content of algal cells using Nile red fluorescent dye. This is achieved by first suspending the cells in an ethanol solution to permeable the cell wall and cell membrane. As a second step Nile red fluorescent dye is added, which enters the cells and accumulates in the non-polar regions of the cell, including the neutral lipid bodies.

Next, the samples are exposed to an excitation light source of 530 nanometers, causing the Nile red absorbed in the neutral lipid bodies to fluoresce at 604 nanometers. Results are obtained on a mass scale by conducting the individual tests in a micro well multipl and using a spectrophotometer to produce rapid fluorescence readings. Readings show a linear relationship between the neutral lipid content of the lgal cells and the intensity of the fluorescence measurement.

Conversion of fluorescence measurements to oil content can be achieved with a calibration curve prepared with cells of known oil composition. The main advantage of this technique over existing protocols, such as the Bly dire gravimetric method, is it allows for rapid determination of neutral lipid content using materials available to most labs. This method provides immediate feedback on bioprocess performance during algae cultivation.

Also, the high throughput allowed by the 96 Well plate reader is ideal for screening studies and factorial experiments. Though this technique has been optimized for chlorella and s Desus, it can be easily modified for analysis of other lgal species.Okay. To begin, remove a sample volume from the growing algal culture that will provide at least 200 milligrams of dry biomass.

400 to 600 milligrams is preferable. Centrifuge the sample at four degrees Celsius for 10 minutes at 10, 000 times. G, discard the supernatant and wash the pellet with an equal volume of phosphate buffer formulated to the same pH as the growth media.

Repeat this process for a total of three washing steps. Resuspend the pellet in deionized water and transfer to a pre weighed weigh dish. Let the sample dry at 50 degrees Celsius for 48 hours.

The dried algal cultures can be stored at room temperature for future use. Measure approximately 50 milligrams of dry algae biomass in a whey dish. Transfer the biomass to a mortar pre-washed with hexane if necessary, wash the whey dish with a small amount of hexane using a past pipette in order to completely transfer the biomass to the mortar order.

Next, grind the algal biomass for five minutes using a pestle. Begin with gentle grinding and gradually increase the intensity. Grind the biomass into a fine and smooth paste.

During the five minute period, add a few milliliters of hexane to the mortar and mix the resulting slurry with the pestle until it is homogenized. Ensure that all the cell debris adhered to the walls of the mortar are knocked free and suspended in the liquid. Next, transfer the hexane cell mass mixture to a compatible centrifuge tube.

Repeat three to five times until all the biomass has been transferred to the centrifuge tube. Centrifuge the sample at four degrees Celsius for 20 minutes at 10, 000 times. G.Following centrifugation, carefully pipette the supernatant into a pre weighed metal way dish, which can be stored in the fume hood.

Perform a second hexane extraction by adding three milliliters of hexane to the pellet and vortexing vigorously. For one minute, repeat the centrifugation and transfer of the supernatant. Determine the mass of oil extracted gravimetric after the hexane has completely evaporated.

For fluoro metric quantification, prepare all algal samples at the same biomass concentration and in the same manner as the standards used in the measurement. Do this by suspending pre dried samples in the appropriate amount of phosphate buffer. If necessary, use a homogenizer to facilitate resus suspension of dried lgal cells.

For each sample mix 80 microliters of a prepared 30%ethanol solution, 10 microliters of a prepared Nile red solution and 10 microliters of algae suspension in a single well of a 96 well plate. In order to properly account for the variability of the fluorescence measurement, perform five replicates of each sample. Next, run a two point calibration curve with standards prepared previously.

In order to account for day-to-day variations in the instrument and preparation to perform the fluorescence measurements in a multi-well plate reader spectrophotometer, set the parameters to shake at 1, 200 RPM and orbit three millimeters for 30 seconds, followed by incubation at 40 degrees Celsius for 10 minutes. Another shake at 1, 200 RPM and orbit three millimeters for 30 seconds, and to record fluorescence with excitation at 530 nanometers and emission at 604 nanometers. Then start the run as a final step.

Convert the fluorescence measurements to oil content using the results from the internal standards to perform fluorescence microscopy. Use the sample prepared with the Nile red stain to prepare a microscope slide of the process sample according to standard laboratory procedures. Starting with the microscope in transmission mode, load the prepared slide into the microscope and locate the cells at the desired magnification.

Once focused, switch the microscope from transmission to epi fluorescence illumination mode. The light source should now be coming directly from the objective lens, which can be confirmed visually by observing the space between the slide and the objective lens. Insert a green excitation filter into the light source and a red emission filter into the observation light path.

The fluorescence of the stained cells should now be directly visible through the eyepiece. Switch the microscope observation mode from the eyepiece to the mounted camera and use viewing software to capture an image of the fluorescing cells. Specific settings will vary with instruments, equipment, and cell types.

Representative algal cells stained with Nile red dye are shown images of a prothe grown in excess nitrogen lead to very low intracellular lipid accumulation, which are shown in contrast to samples of a OIDs grown under nitrogen limitation. Under transmission illumination, the lipid bodies of the cells can be visualized with careful inspection where they appear as shiny circular structures and constitute the majority of the cell volume. The cells shown here are only 5%oil by dry weight and do not contain significant levels of lipid bodies when shown the appropriate light conditions.

The differences in these samples are magnified. The oil lean cells appear as fluorescent rings with dark bodies, while the oil rich cells display a bright orange red glow where they have accumulated lipid bodies. Nile red will fluoresce at different wavelengths depending on the presence of neutral lipids, phospholipids located on the cell wall and nonpolar cellular proteins, which is why the oil lean cells appear as fluorescent rings with dark bodies.

This non specificity is also what causes the background level of fluorescence during the spectrophotometer measurements of the neutral lipids under the optimized conditions of the assay, calibration curves with R square values greater than 0.980 are readily achievable. The relationship between cell oil content and fluorescence becomes non-linear if the cells are stained in a solution, lacking a carrier solvent such as ethanol. The data presented here have an R square of 0.395, and were obtained by carrying out the protocol in a 0%ethanol solution.

Once mastered, this technique can be done in 15 to 20 minutes if performed properly. While performing this procedure, it is critical to ensure that all samples are prepared at the same biomass concentration. After watching this video, you should have a good understanding of how to measure the neutral lipid content of lgal cells using both traditional hexane extraction protocols and the simpler NAL red fluorescent staining procedure.