This article describes a method for monitoring protein-ligand interactions using nuclear magnetic resonance (NMR) in a cellular environment. The technique involves embedding target protein-expressing cells in agarose and analyzing changes in NMR spectra upon ligand binding.
To monitor the interactions of target proteins with small molecule test ligands in a native cellular environment, begin with an NMR tube filled with a buffer containing a gel plug at the base. Inject the target protein-expressing cells, embedded in agarose into the NMR tube, forming thread-like structures.
Load the tube into a bioreactor's flow unit, supplying a growth medium to maintain cell viability. Insert the flow unit inside a nuclear magnetic resonance or NMR instrument.
Under a strong magnetic field, the atomic nuclei of the proteins' hydrogen atoms get magnetized and align parallel to the field, adapting to a low-energy state.
Apply a radiofrequency pulse, causing nuclei to align anti-parallel to the magnetic field, allowing them to transition to higher-energy states.
After a brief pulse, the nuclei return to their original position, emitting absorbed energy. The detector captures this energy, generating an NMR spectrum representing the hydrogen atom's resonance frequencies. Each spectral peak corresponds to a different hydrogen atom in a specific electronic environment.
Add ligand molecules into the flow unit. These penetrate inside the cells, interact with the target protein, and alter the electronic environment around the hydrogen atoms, causing resonance frequency change.
Compare the NMR spectra. The spectral differences post-ligand binding indicate successful protein-ligand interaction.
With a Pasteur pipette, fill the bottom of the flow unit NMR tube with 60 to 70 microliters of 1.5% agarose gel, and place it on ice to create a 5-millimeter high bottom plug. Heat the cell pellet for 15 to 20 seconds in the block heater, and carefully resuspend the cells in 450 microliters of agarose solution, avoiding the formation of bubbles.
Aspirate the cell-agarose suspension into a 30-centimeter long chromatography PEEK tubing of 0.75-millimeter inner diameter connected to a 1-milliliter syringe. Let the tubing cool at room temperature for 2 minutes. Pre-fill the flow unit NMR tube with 100 microliters of PBS at room temperature. Cast threads of cells embedded in agarose into the flow unit NMR tube by gently pushing the syringe.
Remove the empty NMR tube from the flow unit and increase the flow rate to 2 milliliters per minute for a few minutes to remove residual gas bubbles in the inlet tubing. Set the flow rate to 0.2 milliliters per minute and insert the NMR tube containing the cells by pushing it upwards slowly but steadily. Supply the bioreactor medium at a flow rate of 0.1 milliliters per minute.
Set the temperature in the NMR spectrometer to 310 Kelvin, and insert the flow unit in the spectrometer. Once the bioreactor is inserted in the NMR spectrometer, wait a few minutes to allow the medium exchange. Adjust the matching and tuning of the proton channel, shim the magnet, and calculate the proton 90-degree hard pulse length.
Adjust the proton power levels in each pulse sequence according to the proton hard pulse. Record a first zgesgp proton NMR spectrum to record the sample content and field homogeneity. Copy the zgesgp and the p3919gp, or the sfhmqc experiments to the desired number, and queue them in the acquisitions spooler.
Inject a concentrated solution of the external molecule to the medium reservoir bottle by piercing the silicone tubing with a sterile long-needle syringe. At the end of the NMR experiment, replace the tube containing the cells with an empty tube and rinse the flow unit with water.
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