The overall goal of this experimental setup is to delineate the main layers of RNA and DNA regulation to uncover principle molecular mechanisms occurring in response to a stimulus or treatment. This method can help answer key questions in many different biological fields, such as immune responses in general, but also molecular responses to drug treatment. The main advantage of this technique is that it enables you to simultaneously display multiple layers of gene regulation over time.

Therefore, it is important to use the same pool of cells for each method. Broadly speaking, these three protocols involve first manipulating cells of interest to different time points. After each manipulation, the cells are pooled and divided into three groups, one for each assay.

This section of the video describes the labeling and purifying of the most recently produced transcripts using 4sU-labeling. Each protocol ultimately produces samples ready for sequencing. First, look up the required 4sU concentration and incubation time needed for the cell line of interest.

For example, 200 micromolar 4sU is used on T cells for one hour. Separate the pool of cells after treatment into one separate dish for each method and time point. Then, thaw out the required number of 4sU aliquots completely, and add the 4sU directly to the cell medium and incubate the cultures as directed.

Next, collect the 4sU-labeled cells in polypropylene tubes. Then, centrifuge the cells. Resuspend the pellet in TRIzol at about three million cells per milliliter, and incubate the cells in TRIzol at room temperature for five minutes.

Now prepare 30 to 80 micrograms of RNA, or more, using the method by Radle and company. Next, mix the thiol-specific biotinylation reaction in this order. Start with nuclease-free water, then 10-fold buffer, then RNA, and last biotin-HPDP.

Then, mix with pipetting and incubate. Proceed with the cleanup step to produce samples of newly transcribed biotinylated RNA mixed with unlabeled, preexisting RNA. Next, heat the samples to 65 degrees Celsius for 10 minutes.

Then, immediately place them on ice. Once each sample cools, add 100 microliters of streptavidin beads, and incubate them at room temperature with rotation for 15 minutes. Now, isolate the beads decorated with newly transcribed RNA using a magnetic column, and then elute the RNA in 11 microliters of nuclease-free water for sequency.

This section describes how to isolate RNA that are actively being translated. Sequencing these transcripts surveys the dynamic translatosome. Coupled with RNA sequencing data, this assay allows for calculation of RNA turnover and translation rates.

Start with adding cycloheximide to the suspension and mixing it using inversions. After one minute at room temperature, centrifuge the cells, remove the medium, and wash cells with at least 10 milliliters of PBS supplemented with cycloheximide. Next, aspirate the medium, and replace it with 100 microliters of mammalian cell lysis buffer per 10 million cells.

Triturate the mixture to lyse the cells using a 20 to 25-gauge needle. Then, transfer the cell lysate to a pre-cooled 1.50-milliliter tube, and incubate the lysate on ice for 10 minutes with periodic inversions. After 10 minutes, centrifuge the lysate and transfer the supernatant into a cool, 1.5-milliliter tube.

Next, prepare a one to 10 dilution of the lysate in nuclease-free water, and measure the absorbance at 260 nanometers. Then, create 200-microliter aliquots of the undiluted lysate, and treat each with 7.5 units of nuclease for each unit of absorbance. Allow the nuclease to react for 45 minutes, and then end the reaction using 15 microliters of RNase inhibitor, or store at negative 80 degrees Celsius.

Then, purify the RPF RNA using a commercial kit, and follow with an rRNA depletion, also using a kit. Next, perform a PAGE purification on 500 nanograms of isolated RPF RNA. After preparing the samples to load into the gel, denature them, the latter and controls, with a five-minute incubation at 95 degrees Celsius.

Then, run the samples at 180 volts over about 75 minutes, and assess the results. For the gel slice collection, poke holes at the bottom of a 0.5-milliliter tube using a sterile 20-gauge needle. Next, excise gel slices corresponding to the 28 to 30 base nucleotide region, and transfer them to the prepared 0.5-milliliter tubes.

Include a sample of the RNA control and a sample of the negative control to check for contamination. Next, load the small, capped sample tubes into larger microcentrifuge tubes. Then, centrifuge the assemblies to shred the gel, and transfer the sample to the 1.5-milliliter tube.

Next, add water, ammonium acetate, and SDS to the sample tubes to elute the RNA in an overnight reaction at four degrees Celsius. Then, use a commercial kit to make a cDNA library from the small RNA, and proceed with sequencing the cDNA library. ChIP-seq is used to map the binding of a transcription factor of interest throughout a genome.

Combined with data from ribosome profiling and the survey of all the new transcripts, ChIP-seq reveals the actual impact of transcription factor binding. To begin, cross-link the cell samples using formaldehyde to a final concentration of 1%After 10 minutes at room temperature with gentle rocking, stop the reaction by raising the glycine concentration to 0.125 molar. Then, collect the cells and wash them three times with ice-cold PBS, and freeze the cell pellet at negative 80 degrees Celsius.

Later, resuspend the cell pellet in one milliliter of ice-cold cell lysis buffer with freshly added protease inhibitors, and incubate the cells on ice for 10 minutes. After 10 minutes, spin down the cells, aspirate the supernatant, and repeat the lysis buffer addition and incubation. Next, use sonication to generate sheared chromatin between 200 and 1, 000 base pairs.

Then, couple the chromatin to antibody-coated magnetic beads, and elute the protein-DNA complexes in 50 microliters of elution buffer. Next, add the five microliters of 40%RNase and elution buffer, and incubate the sample for 30 minutes. Then, add one unit of proteinase K and 20 micrograms of glycogen to the sample, and incubate it for two hours.

Next, move the sample to 65 degrees Celsius overnight to reverse cross-link it. The next day, place the sample against the magnet for at least 30 seconds, and then collect the supernatant, which contains the DNA of interest. Then, purify the DNA, use quantitative PCR for verification, and sequence the DNA.

4sU-labeling can stress cells and lead to apoptosis. Annexin V and 7-AAD staining was used to identify apoptotic and dead cells. After labeling Th1 cells with 4sU using 30 or 60-minute treatments, the cells did not die or undergo apoptosis.

The RNA's integrity was always checked prior to sequencing using a standard assay, as this is of great importance when processing RNA. Amplified libraries had a healthy range of nucleotides. Excessive adapter dimers needed further purification by PAGE.

Too much template or too many cycles results in heavy nucleotides, smeared products, and adapter dimer products Chromatin shearing was found to work best when the main fraction of the sheared chromatin was around 1, 000 base pairs or slightly less. Quantitative PCR was used to test the applicability of the antibody to chromatin immunoprecipitation sequencing. The negative control primers are for genes not expressed in the cells of interest.

While attempting this procedure, it's important to remember to make a detailed plan of the experimental setup, including a time schedule for when to add 4sU and when to harvest the cells for each method. Always verify optimal 4sU-labeling conditions by testing the impact of 4sU on cell viability and stress response in advance. In summary, this approach allow us to visualize all events of gene regulation that might occur during a process of cellular activation.

This method could also be utilized for primary immune cells and their response to a stimuli or a drug.

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