Voltage-clamp Fluorometry in Xenopus Oocytes Using Fluorescent Unnatural Amino Acids

This article describes an enhancement of conventional Voltage-Clamp Fluorometry (VCF) where Fluorescent Unnatural Amino Acids (fUAA) are used instead of maleimide dyes, to probe structural rearrangements in ion channels. The procedure includes Xenopus oocyte DNA injection, RNA/fUAA coinjection, and simultaneous current and fluorescence measurements.

The overall goal of this procedure is to introduce a fluorescent unnatural amino acid into a membrane protein expressed in Xenopus oocytes and to determine the protein's function and structural rearrangement simultaneously, using voltage-clamp fluorometry. This technique will help answer key questions in the field of structure function relations of membrane proteins. Using voltage-clamp fluorometry, we can directly correlate structural rearrangements with protein function.

Adding fluorescent unnatural amino acid labeling to it help to overcome previous limitations on the labeling. This was once the accessibility of the labeling sites as well as background labeling due to endogenous binding sites. The implication of this technique extends towards a better biophysical understanding of membrane proteins because you can now probe domains, which previously were inaccessible with traditional voltage-clamp fluorometry.

After surgical removal of the Xenopus oocytes, wash the oocytes three times with SOS solution. Select large and healthy oocytes individually, and incubate them at 18 degrees Celsius for at least four hours in Barth's solution supplemented with antibiotics and 5%horse serum before injection. For nuclear injection of DNA, prepare a long and thin injection tip to reach the nucleus without damaging the oocytes.

Then, fill the injection tip with oil, and mount the injection tip on the nanoinjector device. Next, install the nanoinjector under a stereo microscope and break the end of the tip with forceps. After that, eject the oil until there is no air bubble trapped inside the end of the tip.

Following that, place one microliter of 0.1 microgram per microliter pAnap in nuclease-free water on a piece of Parafilm under the stereoscope, and fill the injection tip with DNA. Then, transfer the oocytes to a mesh-containing injection dish containing Barth's solution supplemented with antibiotics. As the oocyte nucleus is located in the animal pole, aim the injection tip at the center of the animal pole and impale such that the tip reaches near the center of the animal hemisphere.

Next, injection 9.2 nanoliters of the pAnap solution into the nucleus of each oocyte. Subsequently, incubate the oocytes in two milliliters of Barth's solution supplemented with antibiotics and 5%horse serum at 18 degrees Celsius for six to 24 hours to allow robust expression of Anap-specific tRNAs and tRNA synthetases. Now, prepare the nanoinjector again, but this time, the injection tip does not need to be as thin as the one for DNA injection.

Work in only red light from this point to prevent photobleaching of the Anap. Mix one microliter of one-millimolar Anap with one microliter of one to two micrograms per microliter mRNA directly on a piece of Parafilm, and fill the injection tip with the mix solution. Impale the tip in the vegetal pole just below the membrane, and inject 46 nanoliters of the mix solution in each pAnap-injected oocyte.

Then, protect the oocytes from light by placing them in a light-tight box or by wrapping the flask in aluminum foil. Incubate them in Barth's solution supplemented with antibiotics and 5%horse serum at 18 degrees Celsius for two to three days. Replace it with fresh Barth's solution every day, and remove the dead oocytes to avoid contamination.

In this setup, the cut-open voltage-clamp equipment is integrated into a fluorescence microscopy setup. The recording chamber is installed on a slider that allows it to move between the standard stereoscope for placing the oocyte and the microscope to perform fluorescence measurements. A photodiode detection system is connected to the C-mount exit port of the fluorescence microscope, and the photocurrent readout is connected to a second input channel in the digital signal processor.

A 100-watt, 12-volt halogen lamp is used as a light source for the fluorescence excitation. The excitation is controlled by a mechanical shutter between the excitation light source and the microscope. The activation is triggered via a TTL pulse coming from the digital signal processor that is timed in the recording software such that the shutter opens about 100 milliseconds before the beginning of the recording, and an appropriate filter cube is required in the filter cube turret.

For two-color VCF, incubate the prepared oocytes in five-micromolar TMR maleimide in labeling solution for 15 minutes immediately before recording. Then, wash the oocytes with the labeling solution three times to remove excess dye before recording. At this point, the protein is specifically labeled with two different fluorophores.

After oocyte mounting, with the animal pole facing upwards and permeabilized, slide the chamber to the microscope and focus on the oocyte using a 4X objective. Next, impale the oocyte with a voltage-sensing V1 electrode. Then, switch to the 40X water immersion objective.

Focus on the animal pole, which is facing upwards. After that, turn off the red light. Select the Anap filter cube by turning the filter cube turret and the optical exit port connected to the photodiode.

Following that, turn on the halogen lamp at the highest intensity, and open the shutter for two to five seconds to read the background fluorescence intensity originating from the oocyte. Afterward, turn on the clamp, flip the bath/guard switch to active, and adjust the membrane potential to the command potential by turning the knob on the headstage. Subsequently, select the holding potential, step protocol, number, and length of pulses in the recording software.

Then, record the voltage-dependent currents and Anap fluorescence intensities. For two-color VCF, select the TMR filter cube by turning the cube turret. Read the background fluorescence for TMR, as described for Anap, and record voltage-dependent currents and TMR fluorescence intensity simultaneously.

This figure displays Anap and TMR fluorescence signals using step protocols obtained from the same oocyte expressing a Shaker mutant. By adding a cysteine, accessible from the external solution, into the L382 stop W434F background and labeling it with TMR maleimide, it is possible to probe real-time movements in different regions of the same protein simultaneously. Fluorescence voltage relationship is obtained by plotting the steady-state fluorescence intensity against voltage.

After watching this video, you should have a good understanding of how to express fluorescent unnatural amino acids into Xenopus oocytes and how to perform one-color or two-color voltage-clamp fluorometry. Once you master the technique, it should roughly take about the same amount of time as traditional electrophysiology experiments, you just have the extra step of injecting DNA into the nucleus of the oocyte. When attempting this procedure, it's important to remember to check for leak expression in the absence of the unnatural amino acid.

This has to be done for every mutation, as the amount of leak expression depends on the insertion site of the stop codon within the protein. This technique now enables researchers to study conformational changes on the cytosolic site of the protein, where many of the regulatory mechanisms occur.