Our research focuses on the dynamics of catalytic active RNAs and how these dynamics change under different environmental conditions. We use single-molecule FRET to study the influence of factors such as monovalent and divalent midlines, chaperone proteins, and crowding agents on RNA dynamics. Precisely incorporating donor and acceptor fluorophores without disrupting RNA structure and function is challenging.
Additionally, tracking the folding of a large dynamic RNA is difficult. However, the established labeling and encapsulation protocol allows us to monitor the folding dynamics of the functional RNA over time. Our protocol addresses the challenge of observing large, catalytically active RNAs in real time and under neophysiological conditions.
By covalently labeling and encapsulating the RNA and vesicles, which are then surface immobilized, we can monitor RNA function with single-Molecule TIRF. We focus on covalent FRET labeling of long catalytic RNAs and particularly group II introns. This is a challenge due to their structural complexity.
When encapsulated in vesicles, we get to keep the RNA within the evanescent field for TIRF microscopy, and yet the RNA can freely float. In this way, smFRET can reveal the dynamic behavior of RNA without hindrance. We will continue trying to understand how nature really works, focusing on the structure-function relationship and dynamics of ribosomes, riboswitches, and other regulatory RNAs using a multi-technique approach.
So moving towards investigations under physiological and cellular conditions will be a major emphasis of our work in the future. To begin, aliquot the RNA of interest, having 50 to 75 micrograms of RNA per tube in 55 microliters of double-distilled water, then add 45 microliters of the EDC/NHS sodium acetate pH 6 mixture to the RNA. Incubate the mixture for four hours at 25 degrees Celsius while shaking at 500 RPM.
After incubation, resuspend the activated RNA pellet in 95 microliters of 100-millimolar MOPS buffer pH 7.5. Then add five microliters of two-millimolar sulfonated cyanine-3 amine to the activated RNA, and incubate at 25 degrees Celsius for 16 hours at 500 RPM to couple the fluorophore to the activated 5-prime phosphate. The next day, add 200 microliters of double-distilled water to improve the separation.
Then purify the 5-prime phosphate labeled RNA by ethanol precipitation. After purification, resuspend the labeled RNA in 100 microliters of 100-millimolar tris HCl, and incubate for two hours at 25 degrees Celsius. Then to remove the free dyes, wash the labeled RNA in double-distilled water with centrifugal filtration.
For dual-end-labeling, aliquot the 5-prime end-labeled RNA, having approximately 50 to 75 micrograms of RNA per tube in 90 microliters of solution. Add five microliters of one-molar sodium acetate buffer pH 5.5. Then add four microliters of freshly prepared 500-millimolar sodium metaperiodate stock solution.
For 3-prime end single-labeling, add eight microliters of metaperiodate stock solution. After incubating the solutions in the dark for two hours, pipette 30 microliters of 50%glycerol to stop the reaction. After 30 minutes of incubation, add 400 microliters of ice cold ethanol sodium acetate precipitation mixture.
Resuspend the oxidized RNA pellet in 95 microliters of 50-millimolar sodium acetate buffer pH six. Purify the dual-end-labeled RNA by ethanol precipitation and centrifugal filtration, as demonstrated previously. Elute the purified RNA in double-distilled water.
Visually compare the supernatant and pellet colors obtained after each step. Calculate the RNA and conjugate dye concentrations on a UV-visible spectrophotometer. Fluorescent gel electrophoresis confirmed the successful single and dual labeling of RNA, as shown by the comigration of the RNA with the fluorophores.
FRET efficiency increased in the presence of metal ions, indicating RNA folding, which led to a decrease in donor emission and an increase in acceptor emission. To prepare a microfluidic chamber, use a diamond driller to drill four holes in the quartz slides to form two channels. Place the drilled quartz slides and approximately twice as many coverslips into a glass Coplin staining jar with a cover.
Sonicate the slides and coverslips in double-distilled water for five minutes. Next, sonicate the jar in a 10%laboratory glassware cleaning solution for 30 minutes at 50 degrees Celsius. After rinsing the jar with double-distilled water, sonicate in one-molar potassium hydroxide solution for 30 minutes and leave it in the solution overnight.
After repeated rinsing and sonication with distilled water, dry the slides and coverslips using nitrogen gas flow. Treat the dried slides and coverslips with an oxygen plasma cleaner for 30 minutes. Place the clean slides and coverslips in a Coplin staining jar containing 3%AP-test ethanol solution Sonicate for one minute and then incubate for 30 minutes.
Rinse the slides and coverslips three times each with absolute ethanol and double-distilled water, then dry them under nitrogen gas flow. Next, prepare a humid box by filling an empty pipet tip box halfway with double-distilled water. Place the slides inside the box with the side to be treated facing up.
Add 30-microliter droplets of freshly prepared bPEG/mPEG mixture to the center of the slide to cover both channels. Cover the droplet with a clean coverslip and close the humid box. Incubate the box overnight in the dark for PEGylation.
The next day, rinse the PEG-passivated and biotinylated slides and coverslips with double-distilled water and observe the change in hydrophobicity of the treated surfaces. Attach the double-sided sticker to the slide, ensuring the sticker covers the area of interest. Then carefully place the coverslip on top of the slide, aligning the PEGylated surfaces to face each other.
Transfer the assembled chamber to a 50-milliliter centrifuge tube, and fill the tube with nitrogen gas. To begin, use a sterile needle to poke a hole from the inside out in the lid of a sterile clean two-milliliter tube. Add 109 microliters of bPE-DMPC mixture into the tube.
Place a cardboard cell partition insert from a microtube storage box into a 500-milliliter Schlenk flask to serve as a tube holder. Using long tweezers, carefully place the tubes containing the lipid mixture into the tube holder. Evaporate the chloroform under a low flow of nitrogen gas overnight.
After obtaining the dried lipid layer, seal the tubes with parafilm to cover the holes. Assemble the extruder using a 100-nanometer polycarbonate membrane and a 10-millimeter polyester drain disc. Equilibrate the syringe and membranes with anti-blinking buffer.
After mixing dual-end-labeled RNA with anti-blinking buffer, hydrate the lipid cake with RNA anti-blinking buffer mixture at 30 degrees Celsius. Shake for five minutes at 1, 400 RPM, followed by 15 minutes at 700 RPM. Centrifuge the mixture for two minutes at 13, 000 G, and dilute the supernatant with 250 microliters of anti-blinking buffer.
Fill the syringe with the RNA lipid suspension, and extrude 35 times at 30 degrees Celsius on a heating block to encapsulate the dual-end-labeled RNA in phospholipid vesicles of 100-nanometer diameter. Flush the chamber two times with 200 microliters of 5X standard buffer at room temperature. Fill the chamber with 50 microliters of 20 microliters per milliliter streptavidin solution, and incubate for five minutes.
Then flush the chamber with 200 microliters of standard buffer and 100 microliters of anti-blinking buffer. Add 75 microliters of vesicle suspension and incubate for 10 minutes for immobilization of the encapsulated RNA on the surface. After immobilization, flush the chamber with 200 microliters of freshly prepared imaging buffer.
For data acquisition, mount the chamber on the TIRF microscope and acquire smFRET data with alternating laser excitation. Single-molecule FRET imaging using TIRF microscopy tracked individual RNA molecules and detected real-time changes in FRET efficiency. Dynamic Single-Molecule traces showed anti-correlated fluctuations between donor and acceptor signals, indicating confirmational changes in RNA.
