두 개의 상보적인 세포 동기화 프로토콜에 의한 세포주기 조절 유전자 발굴 연구

The overall goal of this protocol is to provide a context for the analysis of gene expression in a cell-cycle specific manner. This method can help answer key questions in the cancer field related to cell-cycle dependent transcriptional events that underlie malignant transformation as well as anti-cancer treatment. The main advantage of this protocol is that it establishes with accuracy cell-cycle phase specific gene expression patterns both in number two conditions and after exposure to chemotherapy.

U2OS cells are used for this experiment, and should be seeded in 100 millimeter dishes in the evening around seven p.m. so that subsequent steps can be carried out during working hours. Let cells attach by incubating the dishes at 37 degrees celsius in a humidified atmosphere with 5%CO2 for 24 hours.

On the following day examine the cells to confirm that they are at 50%confluency. For the Thymidine block, add 100 microliters of a freshly prepared 200 millimolar thymidine stock to each 100 millimeter culture dish. Incubate the cells with Thymidine at 37 degrees celsius in a humidified atmosphere with 5%CO2 for 20 hours.

On the following day, at about three p.m. release cells from the Thymidine block by removing Thymidine containing growth medium. Wash cells twice with pre-warmed 1X PBS.

And then add 10 milliliters of complete medium to each 100 millimeter dish. Incubate cells at 37 degrees celsius for five hours. For mitotic cell arrest add nocodazole to a final concentration of 50 nanograms per milliliter.

Incubate cells with nocodazole for no longer than 10 to 11 hours. On the following morning, between six and seven a.m. check the cells under the microscope to confirm that they are indeed blocked in G2-M, indicated by their rounded appearance.

Detach mitotic cells by shaking each plate, and gently pipetting nocodazole containing growth medium with detached cells from each 100 millimeter plate. Combine mitotic cells from all dishes into a 50 millileter sterile tube. Centrifuge 300 times g for five minutes at four degrees celsius.

Wash cells twice by adding cold 1X PBS, followed by centrifugation. After removing the PBS from the second wash, re-suspend mitotic cells in complete culture medium. Save two milliliters for RNA extraction, and two milliliters for FACS analysis for the zero hour time point.

Re-plate remaining mitotic cells for subsequent time points in six well plates. To begin this procedure, seed 0.25 times 10 to the sixth U2OS cells per well in six well plates. On the cover of each six well plate write the experimental condition for each well.

For example, untreated or treated with different concentration of drug and time points. Let cells attach by incubating the six well plates overnight. On the following morning examine the cells to confirm that they are at 50%confluency.

Remove complete medium from the wells, and add two milliliters of pre-warmed, FBS free DMEM-Glutamine medium per well. Incubate the cells for an additional 24 hours. To arrest cells with hydroxyurea, remove medium from the wells, and replace with freshly prepared four micromolar hydroxyurea containing complete medium.

Incubate cells in hydroxyurea containing medium for 24 hours. Prior to releasing cells from hydroxyurea mediated arrest, check the cells to confirm that they are arrested. Remove hydroxyurea containing medium from wells, and rinse wells twice with pre-warmed 1X PBS.

After removing the PBS from the second wash, add two milliliters of complete medium per well. Keep the plates in the incubator until sample collection. Samples from both synchronization protocols are collected in the same way for FACS analysis and for RNA extraction.

For FACS analysis, rinse each well with two milliliters of pre-warmed 1X PBS, and then add 0.3 milliliters of pre-warmed Trypsin-EDTA solution to detach the cells. After five minutes, inactivate Trypsin-EDTA by adding one milliliter of complete medium. Collect each sample in a separate 15 milliliter tube.

Centrifuge cells at 300 times G at room temperature for five minutes. Discard the supernatant, wash with 1X PBS and spin again. Remove the PBS and save the cellular pellet.

In order to fix cells, re-suspend each cellular pellet in one milliliter of chilled 70%ethanol by gently vortexing. Place the cells on ice for approximately 15 minutes prior to propidium iodide staining and FACS analysis. For RNA extraction remove the medium and rinse each well with two milliliters of pre-warmed 1X PBS.

Take the plate to a safety cabinet, and add one milliliter of suitable RNA isolation reagent per well. Pipet up and down to pipet and lyse the cells and then transfer each cell lysate to a 1.5 milliliter microcentrifuge tube. Incubate at room temperature for five minutes.

Start the RNA extraction procedure by retrieving from the freezer the microcentrifuge tubes with samples in RNA isolation reagent, and thawing them at room temperature inside a safety cabinet for chemicals. Add 400 microliters of chloroform to each sample and shake vigorously without vortexing until completely mixed. Incubate the samples at room temperature for five minutes.

Centrifuge the tubes for 15 minutes in a bench top microcentrifuge. Transfer the aqueous phase of each sample to a new 1.5 milliter microcentrifuge tube. Add one volume of 100%ethanol slowly, drop by drop, to the aqueous phase while mixing.

Do not centrifuge. Transfer up to 700 microliters of each sample including any precipitate that may have formed into a spin column in a two milliliter collection tube provided by a commercial RNA mini-prep kit, and close the lid. Centrifuge for 15 seconds.

Discard the flow-through. If starting from more than 700 microliters per sample, transfer the remaining sample into the spin column, and centrifuge again. After washing the RNA per the manufacturer's instructions, elute each sample by pipetting 30 to 40 microliters of nuclease free water into the spin column, and centrifuging at room temperature for one minute.

Determine RNA concentration and purity of samples by absorbance measurements. Store the RNA samples at minus 80 degrees celsius until use for gene expression analysis. Treatment by the Thymidine-Nocodazole protocol arrests U2OS cells in m-phase entry, and provides a cell population that progresses synchronously through G1 and into S phase as shown by flow cytometry analysis.

Treatment by the Hydroxyurea protocol arrests cells at the G1/S boundary. Upon release, cells transition synchronously through S and G2 phases. The Thymidine-Nocodazole protocol is most suitable for analysing mRNA profiles of genes with peak expression during G1 phase.

As shown for E2F1 expression. By contrast, the Hydroxyurea protocol works better for the analysis of genes with preferential expression in S or G2, as illustrated for E2F7 expression. The cell-cycle is typically perturbed by anti-tumor drugs, shown is a mitomycin C imposed permanent G2 phase arrest in hydroxyurea synchronized cells.

Cell sychronization helps discriminate genes that are responsive to the anti-tumor agent from those that are solely affected by cell-cycle perturbations imposed by the agent, for example, a reduction in E2F1 mRNA levels after mitomycin C treatment is likely an indirect affect of the drug on cell-cycle dynamics. By contrast, the peak of p21-Cip1 mRNA expression after mitomycin C treatment is directly the result of a transcriptional program triggered by this drug. Following this procedure genome wide transcriptomic as well as proteomic analysis can be performed, in order to unravel global gene expression changes affecting a specific cell-cycle phases.

Either in number two conditions or upon anti-tumor treatments. After watching this video, you should have a good understanding of the procedures that are required to perform gene expression analysis in synchronized cell cultures. Don't forget that working with propidium iodide or RNA isolation reagents can be extremely hazardous and precautions such as gloves and safety cabinet for chemicals should always be taken while performing this procedure.