Measuring Cell Cycle Progression Kinetics with Metabolic Labeling and Flow Cytometry

The overall goal of the following procedure is to measure the kinetics and progression of the cell cycle. First cells are pulse chase labeled with romo, deoxy, uridine or BRDU, which is incorporated into the cell's DNA in place of thymidine. The cells are then immuno stained with antibodies against BRDU and analyzed by flow cytometry immediately after staining.

Only cells in S-phase are labeled as the labeled cells progress through the cell cycle to G two. They are detected as having four NDNA content following mitosis. The DNA content is reduced to two N.Thus, analysis of the fluorescence profiles generated by flow cytometry can be used to determine the kinetics of cell cycle progression.

The main advantage of this technique over other protocols that use chemicals for cell synchronization is that there are no physiological perturbations that might inadvertently affect the cell division machinery. We first used this method when our attempts at studying cell cycle progression rate using chemical synchronization gave us unexpected results that interfered with our analysis. This method can help answer key questions in the field of cell biology.

For example, what is the basis for differential growth rates between different cell types? Begin this protocol by pulse Chase labeling the cells with BRDU to do this. Begin by labeling one 10 centimeter plate for each time point plus one for each of the following controls.

BRDU only PI only BRDU negative PI negative seed each plate with five times 10 to the fifth. MCF seven cells in supplemented DMEM incubate the plates for 24 to 48 hours to allow the cells to recover and attach. Once the cells reach 60%conf fluency, replace the medium with fresh DMEM containing 10 micromolar.

BRDU incubate the cells for one hour at 37 degrees Celsius 5%CO2 to allow for BRDU incorporation into the DNA. Be sure to leave one plate untreated to act as the negative control or zero time point after one hour. Remove the pulse labeling medium and rinse the cells briefly with PBS to the two to eight hour time points.

Add fresh growth medium and place them back in the incubator. Then immediately harvest the zero and one hour time point plates as described in the next section. To harvest the cells, aspirate the medium and rinse the plate.

Once with PBS, then trypsin eyes the cells to detach them from the plate and collect them in DMEM centrifuge at three 90 times G for five minutes. Discard the supernatant. Then rinse the cells in one to five milliliters ice cold PBS centrifuge at three 90 times G for five minutes following the spin, discard the supernatant and resuspend the cells in a hundred microliters of ice.

Cold PBS containing 1%FBS. The FBS helps prevent cell clumping. One of the most important aspects of this procedure is to ensure that you have a single cell suspension from your harvested cells.

For facts. To achieve this, you can add your cells directly to the ethanol drop by drop. This is critical Next to fix the cells, use the pipette to add the cells dropwise to five milliliters of ice code.

70%ethanol in a 15 milliliter conical tube. Incubate the cells on ice for 30 minutes or store them at four degrees Celsius overnight. This is an ideal step at which to stop the collection of samples from the various time points, the samples can be left in ethanol for several days, allowing them to be processed simultaneously through the remainder of the protocol to perform BRDU and propidium iodide staining.

Remove the cells from four degrees Celsius and centrifuge them at three 90 times G for five minutes. Following the spin aspirate most of the fixative leaving about 50 microliters. Then vortex.

To loosen the pellet to denature the DNA while vortexing. Slowly add one milliliter of two normal HCL Triton X 100 dropwise then incubate samples at room temperature for 30 minutes. Pellet cells by centrifuging samples for five minutes at three 90 times G next aspirate and discard the supernatant.

Then resuspend the cell pellet in one milliliter of 0.1 molar sodium tetra borate. After centrifuging, the cells again aspirate and discard the supernatant. Then resuspend the cell pellet in 75 microliters of BRDU staining.

Mix incubate at room temperature for 45 minutes, protected from light following the incubation. Pellet the cells by centrifugation as before and resuspend the cell pellet in one milliliter of PBS containing five micrograms per milliliter per perdium iodide. When running the PI positive sample through the flow cytometer create a forward scatter versus side scatter plot to ensure the proper size distribution of cells.

Both of these parameters should be plotted on a linear scale. Simultaneously view a plot of FL three H versus FL two W to view the PI staining versus fluorescence pulse width. This plot is used to create a gait to isolate the fraction of cells that are thought to be within the normal distribution pattern of the cell cycle.

In general, this distribution pattern is characterized by two clusters of cells from the two N and four NDNA content of PI staining representing the G one and G two phases of the cell cycle respectively. A string of cells located between these two clusters is representative of ongoing DNA replication that occurs in the S phase of the cell cycle. Create a plot displaying the gated cells with FL one H or FITC on a log scale on the Y axis and PI on a linear scale on the X axis.

This plot will be used to set the remaining parameters using the various controls as well as to collect data. For analysis, place the PI only stain sample in the loading port of the flow cytometer adjust the gain to place G one at about 200 on the x axis. This will be easier to visualize in a histogram plot of the PI stain.

Next place the BRDU only sample in the loading port and run. Adjust the gain so that the two populations of cells, BRDU positive and BRDU negative appear on the plot. Ideally, the BRDU negative cells are positioned just at or below 10 to the one.

The final control is the BDU slash PI negative sample. Run this negative control to ensure that the cells do not appear in any of the upper or right hand quadrants. Once the parameters of the various plots have been set, the flow cytometer is calibrated and ready for processing of BRDU and PI.Stained cells collect data from a minimum of 10, 000 PI gated cells for each sample.

Then using software such as FlowJo or fax diva, create a plot of FITC versus PI with cells gated as PI positive from the PI versus FL two W plot. To distinguish the cycling population. Next, generate a second gate to isolate the BRDU positive population.

Then to track the time course of cell cycle progression, plot the number of BRDU positive cells with G one or G two content as a function of time to compare cell cycle of progression. Kinetics between two colorectal cell lines HT 29 and LS 180. Cells were collected every hour for eight hours following the removal of A-B-R-D-U pulse.

Comparing the kinetics of the two profiles, the emergence of a G one peak is evident at t equals four hours post BRDU in the LS 180 cells compared with the corresponding time point for the HT 29 cell line, which lacks this peak. This indicates an accelerated cell cycle progression through the G two phase of the cell cycle. In the colorectal cell line LS 180 to generate a cycling profile of the breast cancer cell line.

MC seven cells were collected every hour for eight hours. Following the removal of the BRDU pulse. The immuno labeled and DNA counterstain cells were then sorted based on their fluorescent signals.

As can be seen here, CF seven cells cycled at a significantly slower rate than either one of the two colorectal cell lines tested. After watching this video, you should have a good understanding of how metabolic labeling of DNA in conjunction with time point harvest allows you to track cell cycle kinetics.