The overall goal of this procedure is to determine the relationship between the formation of intracellular protein aggregates and their impact on cellular oxidative stress using the baker's yeast Saccharomyces cerevisiae. This method can help answer key questions in the field of protein misfolding and aggregation diseases, such as neurodegenerative disorders and non-neuropathic amyloidoses. The main advantage of this technique is that it's simple, fast, and allows quantification of the cellular oxidative stress damage in a large yeast population.
With this method, we can provide insight into the correlation between intracellular soluble or aggregated state of an amyloidogenic protein with oxidative stress levels in yeast. In addition, it can also be applied to other model organisms expressing recombinant proteins. Demonstrating the flow cytometry analysis will be Manuela Costa, a technician from the Flow Cytometry Facility at the UAB.
In this study, the intracellular aggregation state of different A-beta-42 peptide variants is tracked using S.cerevisiae transformed with a plasmid encoding for A-beta-42 fused to GFP under the control of a galactose-inducible promoter. Pick one colony of the transformed yeast cells, and inoculate into 20 milliliters of SC minus Ura medium containing 2%glucose. Grow the culture at 30 degrees Celsius under agitation overnight.
On the following day, inoculate 100 microliters of the overnight culture into five milliliters of fresh SC minus Ura medium, and grow the cells at 30 degrees Celsius for two to three hours. When the culture is at an optical density at 590 nanometers or OD 590 of 0.5, centrifuge the culture at 3, 000 times g for four minutes, discard the supernatant, and resuspend the cells in five milliliters of fresh SC minus Ura medium containing 2%raffinose. Incubate the cells at 30 degrees Celsius under agitation for 30 minutes.
After 30 minutes, centrifuge the cells at 3, 000 times g for four minutes, discard the supernatant, and resuspend the cells in fresh SC minus Ura medium containing 2%galactose to induce recombinant protein expression. Return the cells to the incubator for 16 hours. After 16 hours, harvest the cells by transferring one-milliliter aliquots of the culture to sterile microcentrifuge tubes and centrifuging at 3, 000 times g for four minutes.
Begin this procedure by determining the OD 590 of the 16-hour-induced yeast cells. Then dilute the cells in sterile PBS to an OD 590 of 0.1. Transfer the expressing cell suspensions to appropriately labeled round-bottom polystyrene tubes, and protect them from light.
Prepare non-induced cells as a negative control for the flow cytometry analysis. Add the oxidative stress probe to each sample at a final concentration of five micromolar. Cover the samples with aluminum foil, and incubate at 30 degrees Celsius for 30 minutes.
When the incubation is done, spin down the cells, remove the supernatant, and resuspend the cell pellets in PBS. Wash the cells three times in this manner with PBS. After the third wash, resuspend the cells in the same volume of PBS.
Make sure non-stained cells are included in the flow cytometry analysis. Using the appropriate lasers and filters, perform the flow cytometry analysis to detect GFP and the fluorescent signal of the oxidative stress probe. Start by clicking on the Open New Worksheet panel, and give the experiment a name.
From the toolbar, select the Scatter Plot Tool, and create a plot with the variables Side Scatter Area on the y-axis versus Forward Scatter Area in a linear scale on the x-axis. Click the Scatter Plot Tool, and create a plot with the variables FITC Area on the x-axis versus APC Area in a logarithmic scale on the y-axis. Click the Instrument Setting icon, and set all compensation levels to zero in the Compensation tab.
Then click on the Acquisition tab, and select a total number of 20, 000 events to be recorded with a low flow rate. Because the fluorescent emission signal of one fluorochrome can be detected by another detector, it is important to perform a compensation process to equalize the min signal of each fluorochrome in all detectors by a single-color control. Rename tube 001 as negative control, and click the Acquire tab to start running the non-induced, unstained cells.
Adjust the voltage in the Instrument Settings of the forward and side scatter until the population is distributed in the middle left quadrant. Click on the Polygon icon to set a region R1 around the cell population excluding cell debris, and use this gated population P1 equals R1 for all fluorescent dot plots and histogram representations. To adjust the PMT voltage of the fluorescence signal, run the unstained cells in the FL1 to FL3 dot plot, tuning the gain in the Instrument Setting tab, until the cells are distributed in the lower left quadrant.
Using the Scatter Plot Tool, create a dot plot with the variables FITC-A in the x-axis versus FSC-A and a second dot plot with the variables APC-A in the x-axis versus FSC-A. Next, change the sample to the induced cells to measure GFP fluorescence. In the Instrument Setting tab, set the gain in the FSC-A versus FITC when the population is distributed in the low right quadrant.
Define the GFP-positive cell population with a gate. Change the sample to the non-induced stained cells. Display oxidative stress by the fluorescence on an FSC-A versus APC-A dot plot.
Tune the gain until the cell population is distributed in the left upper quadrant. Gate the positive cell population in P3.With the Histogram icon, make two histogram plots to represent the cell fluorescence, one for FITC and another for APC fluorescence. Create a table displaying the mean fluorescent intensity and median fluorescence with its corresponding standard error and/or the coefficient of variance for GFP fluorescence and oxidative stress levels.
Click the Compensation tab, and set all compensation levels to zero. Click on the Acquisition tab, and select a total number of 20, 000 events to be recorded. Prepare three new tubes of samples, and name them non-induced, induced soluble, and induced aggregated.
Acquire data from the samples with the preset settings. Analyze the data by opening a new sheet and creating a dot plot for the variables FITC-A and APC-A, gating the positive cells for CellROX staining. For each sample, create a statistics table with the mean fluorescence and the standard error for the FITC and APC channels.
It is also possible to sort cells with a FACS sorter following the aggregated protein counting. Yeast cells expressing A-beta-42 variants after an induction period of 16 hours are visualized under a fluorescence microscope to determine the recombinant protein distribution inside the cells. These images are representative of selected A-beta-42 GFP variants.
The formation of protein inclusions, or PI, was confirmed in 10 out of the 20 variants from the analyzed collection. This bar graph indicates the percentage of cells containing different numbers of PI calculated from a total of 500 fluorescent cells for each variant into biological replicates. An excellent agreement between predicted and in vivo aggregation properties was observed.
The protein expression levels in cellular extracts were quantified using an A-beta-specific antibody. As a general trend, PI-forming A-beta-42 variants, colored in green, are present at lower levels than those diffusely distributed in the cytosol, colored in red. Important differences among A-beta-42 variants were observed when their oxidative stress probe fluorescence, protein levels, and GFP fluorescence properties were represented relative to their ability to form PI and their intrinsic aggregation propensities predicted by either the AGGRESCAN or TANGO bioinformatics algorithm.
The PI-forming variants are in green, and the non-PI-forming variants are in red. Following this procedure, other methods such as propidium iodide or annexin V staining can be also performed to answer additional questions like determining the impact of an intracellular expressed protein variant on cell apoptosis or cytotoxicity.
