Molecular Probe Optimization to Determine Cell Mortality in a Photosynthetic Organism (Microcystis aeruginosa) Using Flow Cytometry

The overall goal of this procedure is to demonstrate how to create a protocol that can analyze the cyanobacterial physiological state effectively, using flow cytometry and molecular probes. This protocol can help answer key questions about the cell physiology of microbial communities. The main advantage with this procedure is that an optimized fluorescent probe protocol can distinguish between the physiological states of cyanobacteria at the individual cell level in real-time.

Demonstrating this procedure will be David Hartnell, a PhD student working the cytometry lab at Bournemouth University. Before beginning the experiment, place an empty hemolysis tube onto the sample injection probe. Click unclog.

And then back flush to start the flow cytometry cleaning process. At the end of the back flush, load a new hemolysis tube containing two milliliters of ultra pure filtered water onto the sample injection probe, or SIP, and set the time limit to 10 to 15 minutes, and the fluidic speed to fast. Next, select a new data cell, and set the relevant fluorescence and light scatter thresholds to reduce the background noise.

Then click run. If the total events per second are not less than the manufacturer's recommendation, run two milliliters of decontamination solution for two minutes on fast. And repeat the back flush and ultra pure water flushing steps.

To prepare a primary M.aeruginosa monoculture, autoclave 98 milliliters of ultra pure filtered water, mixed with two milliliters of algal medium, in a 250-milliliter beaker for 20 minutes at 120 degrees Celsius. When a culture is in a high steady state density, disaggregate two milliliters of cells by vortexing. And then transfer the cells under the sample injection probe.

Confirm that the cells are evenly dispersed by light microscopy. Then select a histogram plot on the flow cytometer software to record the forward light scatter data. And click log to view the data in a log scale on the x-axis.

In a separate output, set another log axis histogram to record the natural fluorescence of the M.aeruginosa cells. Next, select a light source that can excite phycocyanin, and a detector that can filter the emissions from the resulting fluorescence. For recording at the highest resolution, select the settings closest to the core size of the target organism.

And set a relatively slow flow rate. Before acquiring the data, set a threshold to gate out any light scatter and fluorescence signals caused by electronic background noise or cell sample debris. Then select a new data cell.

Create a density plot with forward and side scatter parameters on a log scale, and click run. Now apply the forward light scatter and natural fluorescence gate to the forward by side scatter data to exclude any low-level scatter signals, and to include only the higher relative fluorescence phycocyanin signals. Then collect events until the sample is finished, and use the data to determine the initial cell count.

To optimize the molecular probe cell uptake, expose half of the previously prepared monoculture to the appropriate conditions for generating a dead control. Check the variations in the sample micro environment to confirm the death of the culture. And then disaggregate the colony formation as just demonstrated.

Next, select a 488-nanometer laser alongside a detector that can record fluorescence from the green and orange nucleic acid probes. And the 640-nanometer laser to record the phycocyanin signals through their respective detectors. In a new data sheet, set up a density plot with forward and side scatter parameters.

Then create one histogram using the respective molecular probe optical detector channel. One histogram to detect the phycocyanin emissions, and one histogram for forward scatter events, all on a log scale. Run the cyanobacterial samples using different concentrations and incubation times.

Create a gate in the forward scatter histogram to include events only with the target organism's cell size. And apply it to the corresponding fluorescence probe channel. Next, in the fluorescence probe channel, create another inclusive software gate against the highest peak in the histogram.

And subsequently gate the corresponding positive probe fluorescence onto the density plot. Compare the number of fluorescence signals to the number of dead control cells, where the highest percentage of cell nucleic probe uptake that does not produce non-specific staining. For testing the fluorescence interference overlap from intrinsic or non-specific cell staining, select the 50 percent live and 50 percent dead mixed-culture data.

And remove the fluorescent gates. Apply gates to include only the forward scatter histogram for the target organism's cell size on the phycocyanin channel histogram. Gating the highest and lowest phycocyanin peaks, and labeling them live'and dead, respectively.

Then separately apply the gates to the live and dead phycocyanin signals. And record both mean wavelengths. To determine the protocol sensitivity, ratio the mean wavelengths of the positive dead molecular probe fluorescence, and the intrinsic non-specific live signal.

Finally, select the optimized protocol where the highest amount of dead cells has been stained without the occurrence of non-specific staining. In these graphs, the representative forward and side light scatter outputs, for cellular size and internal complexity of an M.aeruginosa batch culture in the exponential phase are shown. Gating can be performed by refining the data between certain points of the forward light scatter output.

Phycocyanin produces a strong signal when interrogated by a red light source, which can be used to further gate the populations of interest. From the forward light scatter and fluorescent signals, data can then be gated from the original output, to specific data on the M.aeruginosa sample for final cell counts. When live higher-pigmented, and dead lower-pigmented controls are mixed, the decreasing shift in auto fluorescence becomes clear.

In membrane-compromised cells, the nucleic acid probes produce an additional signal that can be observed by flow cytometry and further confirmed by epifluorescence microscopy. Fluorescence discrimination between live and dead cells increases with time between the 0.05 and 0.5 micromolar concentrations, but decreases between the one and 100-micromolar concentrations for both probes. Indeed, the optimal concentration for the green nucleic acid probe appears to be 0.5 micromolar with an incubation time of 30 minutes.

For the orange nucleic acid probe, the optimal concentration is one micromolar for a 10-minute incubation. Once mastered, for each molecular probe, optimization can be completed in a day, if it is performed properly. While attempting this procedure, it is important to remember to keep the probes in a stable environment, under the appropriate temperature, pH, and light conditions.

After its development, this technique paved the way for researchers in the field of microbiology to explore community heterogeneity in phytoplankton. Having watched the video, you should now have a good understanding of how to develop an optimal protocol for assessing cellular physiology using flow cytometry and molecular probes. Don't forget that working with molecular probes and toxic organisms can be extremely hazardous, and that precautions such as wearing the appropriate P.P.E, and having the full understanding of COSHH materials, should always be taken while performing this procedure.