Quantification of the Abundance and Charging Levels of Transfer RNAs in Escherichia coli

The overall goal of this experiment is to quantify the abundance and charging levels of transfer RNAs in Escherichia coli. This method can help answer key questions in cell biology such as, how tRNA charging levels changes at different growth states. The main advantage of this method is that, RNA levels can be quantitatively compared even between growth states that drastically alter the composition of the RNA in the cells.

Start this experiment by growing experimental and spike-in E.coli cultures according to the text protocol. Grow the spike-in culture at 37 degree celsius, while shaking at 160 RPM. When the spike-in culture has reached an OD436 of about 0.1, induce the expression of tRNA cell C by adding IPTG.

Grow the culture until it reaches an OD436 of about 0.5. Then, move the culture to ice until all samples have been harvested. After that, prewarm one 100 milliliter capped culture flask containing four milliliters of 10%TCA for each sample in a 37 degree celsius water bath.

Measure and record the OD436 of the experimental cultures directly before cell harvest. Then, transfer four milliliters of culture to a prewarmed TCA containing flask. Mix the sample by hand swirling the flask for five seconds, which will immediately disable all enzymatic activities preserving the charging levels of the tRNAs.

When all samples have been collected, add 5%spike-in cells to each sample. Also prepare a control sample consisting only of spike-in cells as described in the text protocol. To start RNA extraction, pour eight milliliters of each sample, including controls into an ice chilled centrifuge tube.

Centrifuge the samples at 9, 500 x G at four degree celsius for 10 minutes. After removing the supernatant completely, re-suspend the pellet in 0.3 milliliters of cold sodium acetate, containing 10 millilimolar EDTA. Transfer each samples into a cold 1.5 milliliter microcentrifuge tube and add 0.3 milliliters of phenol equilibrated with the buffer, used to re-suspend the pellet.

Then, vortex each sample 10 times for 15 seconds each time. With at least one minute at zero degree celsius between each vortex, to keep the samples cold. Next, centrifuge the samples at 20, 000 x G at four degree celsius for 15 minutes.

Transfer the upper or water face to a new 1.5 milliliter microcentrifuge tube. Add 0.3 milliliters of cold phenol to the water face and vortex the samples four times 15 seconds each as previously described. Then, centrifuge the samples at 20, 000 x G at four degrees celsius for 10 minutes.

Transfer the water face to a new cold 1.5 milliliter microcentrifuge tube and add 2.5 volumes of cold 96%ethanol. Then, incubate the tubes at 20 degree celsius for one hour to precipitate the RNA. Centrifuge the samples at 20, 000 x G at four degree celsius for 30 minutes.

After removing the supernatant, carefully add one milliliter of cold 70%ethanol to the RNA pellet, making sure not to disturb it. Invert the tube gently once, to wash it with ethanol. Finally, centrifuge at 20, 000 x G at four degree celsius for 10 minutes.

Remove the supernatant completely, and air dry the pellet until the ethanol is completely removed. At 20 microliters of cold sodium acetate containing 1 millimolar EDTA, vortex vigorously to re-suspend the pellet. To start electrophoresis, mix four microliters of each sample with six microliters of loading buffer.

Load the samples including controls onto a 0.4 millimeter thick 6.5 polyacrylamide gel containing eight molar ureal and 0.1 molar sodium acetate buffer. Then, separate the RNA by running the electrophoresis at 10 volts per centimeter of gel at four degree celsius for approximately 20 hours until the bromophenol blue dye reaches the bottom of the gel. Carefully separate the two glass plates so that the gel remains on one of them.

Use the gel area from the xylene cyanol dye and 20 centimeters down towards the bromophenol dye for blotting. Cut a thin piece of filter paper to the size of the desired blotting area. Place the filter paper on top of the part of the gel that will be used for blotting.

Cut off and discard the parts of the gel not covered with filter paper. Then use the filter paper to carefully lift the gel away from the glass plate. Place a positively charged nylon membrane on top of the gel, and electroblot it in transfer buffer at 20 volts for 90 minutes.

Finish by crosslinking the RNA to the membrane using UV light. Next, place the crosslinked membrane in a hybridization tube and add six milliliters of hybridization solution. Rotate the tube at 42 degree celsius for one hour to pre-hybridize the membrane.

Then, add 30 peak molar radioactive oligo DNA probe, five prime and labeled with P32. Rotate to incubate the membrane with the probe at 42 degree celsius overnight. After incubation, use a disposable 10 milliliter pipette to remove the probe.

Next, quickly wash the membrane twice in the hybridization tube, in 15 milliliters of washing solution at room temperature. Move the membrane to a flat plastic container, cover it with the washing solution and close the lid. Incubate the membrane on a shaker at room temperature for 30 minutes.

Change the solution and continue washing until achieving a satisfying signal to noise ratio. Use a geiger muller tube to obtain the signal to noise ratio, by comparing the radiation from the area where the probe has hybridized to tRNA, to the radiation from an empty area. After that, wrap the membrane tightly in plastic wrap and seal it airtight by welding.

Place this membrane on a phosphorimaging screen. After adequate exposure time, scan the screen using a laser scanner and save the file in gel format. Next, load the gel file into the imaging software.

Draw a vertical line along each of the sample lanes so that it spans most of the width of the lane while excluding the edges of the bands where the signal can be uneven. To visualize counts from each lane, click analysis and then, create graph. Manually define the two peaks that represent the charged and the uncharged tRNA from each lane.

After that, click on analysis and then, area report to obtain counts from each peak. Finally, record the area under each of the two peaks. Calculate the tRNA levels as described in the text protocol.

This method was used to measure the abundance of various tRNAs and E.Coli nf915 before and during our arginine starvation. On the northern plot hybridized with different tRNA probes, there is a clear separation between the bands that represent the charge tRNA from the bands that represent the uncharged tRNA. The radiation profile along each lane, was obtained from the northern blots using imaging software.

The radiation signal from each lane was quantified by isolating each from the blot. A graph that represents the quantification of the signal from a single lane, shows two peaks. The left peak comes from the aminoacylated tRNA and the right from the deacylated tRNA.

The area under each peak is exported for further analysis. Quantification of the bands from the northern blot, shows that the relative abundance of all tested tRNA species rapidly and significantly decreased during amino acid starvation. On the other hand, tRNA charging levels of the arginine accepting tRNA dropped rapidly upon arginine removal and then slightly increased during the starvation period.

However, the charging levels of the other tRNAs increased slightly upon starvation and then decreased to almost pre-starvation levels. After watching this video, you should have a good understanding of how to purify RNA in a way that tRNA charging and abundance can be quantified and how to use spike-in cells for normalization. Throughout this procedure, it’s important to remember to keep the RNA cold and acidic to prevent the hydrolysis of the RNA.