Biochemistry
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Sample Preparation for Mass Spectrometry-based Identification of RNA-binding Regions
Chapters
Summary September 28th, 2017
We describe a protocol to identify RNA-binding proteins and map their RNA-binding regions in live cells using UV-mediated photocrosslinking and mass spectrometry.
Transcript
The overall goal of this procedure is to identify RNA binding proteins in live cells and map their RNA binding regions using UV mediated photo cross-linking followed by mass spectrometry. This method can help answer key questions in the field of epigenetics, such as which epigenetic regulators are bound to RNA and what protein regions are required for the interactions. The main advantage of this technique is that it does not involve purification of the protein RNA complexes, and therefore requires little material and identifies proteins bound to any type of RNA.
As presented, this method focuses on nuclear proteins because that is our scientific interest, but with minor modifications in the fractionation steps, it could be used to study RNA and binding proteins in other stu-nu-lar compartments. After culturing and expanding mouse ESCs according to the text protocol, expand the cells to the required number of 10 centimeter plates. Before the plates reach confluency, add four SU directly into the medium to a final concentration of 500 micro molar.
Leave 4sU control dishes untreated. Gently shake the plates to homogeneously distribute the 4sU throughout the medium, then return the plates to the same tissue culture incubator for two hours. The efficiency of 4sU incorporation varies across cell types, and therefore it might be necessary to optimize it's concentration and incubation time to ensure efficient cross-linking and minimize toxicity.
Next, transfer the plates to an ice bucket and discard the medium. Then, use 5 milliliters of ice-cold PBS to gently rinse the plates. Add 2 milliliters of ice-cold PBS and place the plates without lids into a cross-linker equipped with UV-B emitting bulbs.
Irradiate the plates at an energy setting of 1 Joule per square centimeter. Efficient UV cross-linking is critical. We recommend using UV-B light, but UV-A can also be used.
In any case, the amount of UV energy used should be optimized and can be monitored by PAR-CLIP or ChIRP. Use cell lifters to collect the cells and transfer them to ice-cold PBS in conical tubes. Then, centrifuge the cells at 2000 times g and four degrees celsius for five minutes.
Remove the supernatant and re-suspend the cell pellet in five milliliters of ice-cold PBS with 2 millimolar EDTA and 0.2 millimolar PMSF. With a hemocytometer, count the cells. Expect five to ten, and ten to twenty million cells from 10 centimeter and 15 centimeter plates, respectively.
Then, centrifuge the cells at 2000 times g and 4 degrees celsius for five minutes. To isolate nuclei from the cells, prepare ice-cold buffer A as outlined in the text protocol. Prior to use, add 0.5 millimolar DTT and 0.2 millimolar PMSF to the desired amount of buffer A.Add 2 milliliters of the cold buffer to the cells to wash them, then spin the cells at 2500 times g and four degrees celsius for five minutes.
Discard the supernatant, and add buffer A supplemented with 0.2 percent of a non-ionic, non-denaturing detergent, then rotate the sample at 4 degrees celsius for five minutes. After spinning the cells at 2500 times g for five minutes, remove the supernatant. The pellet comprises intact nuclei devoid of cytoplasm.
To lyse the nuclei, add 200 microliters of lysis buffer to the tube. Use a 1/8th inch probe to sonicate the sample to shear the chromatin at a 50 percent amplitude setting for five to ten seconds. Spin the sample at 12000 times g for five minutes and collect only 175 microliters of supernatant to avoid the pellet.
Next, measure the protein concentration via the Bradford assay or similar technique. Then, use lysis buffer to dilute the protein in the different samples to 1 milligram per milliliter or the lowest sample concentration. Add 5 millimolar DTT solution to the nuclear lysate and incubate the sample at room temperature for one hour.
Then, add 14 millimolar iodoacetamide from fresh stock and incubate the lysate at room temperature in the dark for 30 minutes. Dilute the lysate with five times the volume of 50 millimolar ammonium bicarbonate, then add trypsin. Incubate the sample at 37 degrees celsius overnight.
To prepare one custom-made stage tip per sample, place a disk of solid phase extraction C18 material into the bottom of a 200 microliter pipette tip. Add 50 microliters of Oligo R3 reverse-phased resin on top of the C18 disk, then place the stage tip inside a centrifugation adapter and place the adapter inside a 1.5 milliliter microfuge tube. Add 100 microliters of 100 percent acetonitrile to the tips to wash them.
Then centrifuge the tips at 1000 times g for one minute to remove the solvent. Equilibrate the tips with 50 microliters of 0.1 percent trifluoroacetic acid or TFA. Then, spin them again.
Add 100 percent formic acid to the digested protein sample to decrease the PH to 2 to 3. Then, load the sample onto a custom-made stage tip and centrifuge the sample at 1000 times g for one minute until all of the sample goes through the tip. Wash the bound peptides with 50 microliters of 0.1 percent TFA.
Then, after spinning the sample at 1000 times g for one minute, elute the peptides by adding 50 microliters of elution buffer to the stage tip and centrifuge at 1000 times g for one minute to force the elution buffer out of the tip. Use a vacuum centrifuge set at 150 times g and 1 to 3 kilopascals for one hour to dry the sample. To remove cross-linked RNA from the sample, re-suspend the pellet in 50 microliters of 2x nuclease buffer.
Measure the peptide concentration at 280 nanometers assuming 1 milligram per milliliter of peptides for and A280 of 1. Use 2x nuclease buffer to adjust all samples to one milligram per milliliter or to the lowest concentration, then prepare a master mix of 50 microliters of water plus 1 microliter of high purity nuclease per sample. Combine 50 microliters of peptide sample with 50 microliters of water plus nuclease master mix.
Then, incubate the sample at 37 degrees celsius for one hour. Finally, carry out nanoliquid chromatography, mass spectrometry, and data processing according to the text protocol. This figure shows a volcano plot of an RBR id result from mouse ESCs.
Peptides that overlapped with an RNA recognition motif domain are in blue and show highly consistent depletion levels. An RNA binding peptide from HNRNPC is highlighted in red. Shown here is a heat map of RBR id scores that are calculated as an estimate of a protein region's RNA binding likelihood.
The score is projected on the surface of the spliceosomal subunit U1-70k. The bright red color in the vicinity of the RNA contact, as determined by the crystal structure, indicates correct identification of the protein RNA interaction. In this figure, TET2 is presented as a novel RBP and the RNA binding activity is mapped to a C terminal region by plotting the RBR id score along the primary sequence of the protein.
Finally, the requirement for this novel RBR could be verified by performing PAR-CLIP using the wild type sequence and comparing the signal to that of a mutant lacking the predicted RBR. Once mastered, this technique can be done in seven to eight hours over two days if it is performed properly. Prior to cross-linking, cells can be stimulated with various treatments in order to answer additional questions like how does differentiation start or how does the cell cycle regulate RNA protein interactions.
After watching this video, you should have a good understanding of how to generate extras containing cross-linked protein RNA complexes to identify sites of protein RNA interactions with mass spectrometry.
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