Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

The overall goal of this procedure is to examine the confirmational dynamics of membrane proteins utilizing site directed fluorescence labeling on the surface of single cells. This is accomplished by first individually mutating residues at the interface of the extracellular and transmembrane domains to cystine as shown by the red C.It is then necessary to determine whether this residue can be labeled with a cystine reacting fluoro four. The second step of the procedure is to examine whether cystine, mutagenesis, and or fluoro four labeling affects the kinetics and or confirmational state of the protein.

This is performed using the setup shown. The third step of the procedure is to compare and contrast changes in fluorescence intensity to the kinetic parameters of the target protein. A typical fluorescence trace is shown here.

The final step of the procedure is to label mutated cysteine pairs with either donor Fluor fours or donor acceptor Fluor fours to measure photo destruction rates in tandem with voltage clamp measurements to determine distance constraints. This figure shows donor photo destruction rates in the absence of acceptor in black donor photo destruction in the presence of acceptor is in red. Ultimately, results can be obtained that show that changes in fluorescence intensity, which are the result of protein confirmational changes can be correlated to membrane protein function through voltage clamp fluorometry.

This method can help answer key questions in the field of membrane ion transport, such as how the changes in confirmational state of the protein facilitate small molecule and ion transport across the plasma membrane. Begin the procedure by cloning the sodium potassium pump into a vector suitable for xop posts, Lavis oocyte expression. The next step is to make cysteine mutations at residues located at the interface between the transmembrane and extracellular domains as outlined in the accompanying article.

When finished, verify insertion of the mutation by DNA sequencing after transcribing the plasma DNA to mRNA as outlined in the accompanying article, quantitate the yield of mRNA using a spectrophotometer prior to surgery. The frog should be fasted for 12 hours to prevent vomiting during the operation. To begin the surgery the following day, immerse the frog in the anesthetic solution until the frog becomes unresponsive to a toe pinch.

Remove the frog from the anesthetic solution and place it dorsal side down on the absorbent side of a diaper pad. Place a wet paper towel on the frog to keep the frog moist. Also, if the eyes of the frog are open, keep them moist with saline solution.

Make a small smic incision on one side of the frog's midline. Locate the ovary and bring it to the exterior of the frog. Avoid touching the ovary to the skin.

Remove the ovary and place it in a Petri dish containing ringer solution. Check the remaining tissue for bleeding before placing it back inside the smic cavity. If bleeding occurs, apply pressure with a sterile Q-tip until it stops.

Finish the surgery by suturing the incision using an interrupted suture pattern. Return the frog to its housing to recover from anesthesia using scissors. Divide the isolated ovarian lobe into smaller parts.

Transfer the clumps into ringer solution plus calcium with collagenase. Next, gently shake the digest solution for two hours at 18 degrees Celsius. When most of the cytes are separated, wash the oocytes with ringer solution minus calcium.

Then incubate the oocytes for 10 minutes in ringer solution minus calcium at room temperature. At this point, wash the cytes exhaustively in ringers plus calcium solution. Then transfer the oocytes to ringer solution plus calcium for storage prior to injection.

For the next step, retrieve the synthesized mRNA from the minus 20 degrees Celsius freezer. Add 25 nanograms of the mRNA to a final volume of 50 nanoliters. Then inject the mRNA into the cytes after injection.

Place the oocytes in ringer solution and one milligram per milliliter gentamicin and incubate them at 18 degrees Celsius in the dark for three to seven days. This allows the sodium potassium pump to be expressed within the oocyte plasma membrane. On the day of the experiment, remove the oocytes from the incubator and place them in loading buffer for 45 minutes and then in post loading buffer for 45 minutes.

This increases the intracellular sodium concentration to facilitate measuring the sodium potassium pump. For fluorescence measurements, incubate the cytes in a post loading buffer that contains five micromolar of the desired fluorophore, such as TMRM or FM for five to 10 minutes in the dark. After flora four labeling, wash the oocytes exhaustively with dye-free post buffer.

Keep the oocytes in the dark to prevent photo bleaching. To prepare for the two electrode voltage clamp measurements begin by filling the micro electrodes with three molar potassium chloride and testing the resistance. The resistance should be between 0.5 and 1.5 mega ohms.

For cysteine scanning experiments, equip the fluorescence microscope with a 5 35 DF 50 excitation filter, a 5 65 EFLP emission filter, and a five 70 DRLP diic mirror. Next, place the octe in an RC 10 chamber on the fluorescent microscope stage. Then gently insert both micro electrodes into the octe using the amplifier to hold the membrane potential at a constant value.

Measure the ion flux across the membrane by activating the protein using an appropriate technique such as solution exchange or changing the membrane potential. For concurrent fluorescence measurements, use a 100 watt tungsten light source to excite the donor fluorophore and to pin 0 2 2, A photo DDE to detect changes in fluorescence intensity for fluorescent labeling. Following cysteine scanning.

Measure the change in fluorescent intensity at stationary current using voltage steps that are controlled by P clamp 10 software. A second part of the protocol involves taking anes atropy measurements alongside distance measurements. To calculate the range of kappa squared values, an atropy measures the relative mobility of the fluoro four due to rotation.

See the accompanying article for details of the calculations. To measure distance constraints, use a holo enzyme, which has two accessible extracellular cysteine residues that can be labeled with fluoro fours. Add post loading buffer containing one micromolar FM and four micromolar TMRM.

That is the donor and the acceptor. Fluoro fours to one batch of oocytes add one micromolar FM only. That is just the donor fluoro.

Four to another batch of oocytes incubate both batches of oocytes on ice for 30 minutes in the dark. This allows for measurements to be made of hollo enzymes labeled with and without an acceptor fluoro four. Next, equip the microscope with a 4 75 DF 40 excitation filter, a five 30 DF 30 emission filter, and a 5 0 5 DRLP dichroic mirror.

Then place the oocyte in the chamber on the fluorescence microscope stage. Maintaining a continuous solution flow measure the time dependence of donor bleaching in the presence and absence of the acceptor flora.Four. These results can be used to calculate the distance between the two residues on the sodium potassium pump utilizing forester's.

Equation two electrode voltage clamp measurements should be made concurrently to ensure the protein is functional. Using the clamp X feature in P clamp 10 average the results of the photo destruction of the donor fluoro four for a minimum of four oh site recordings. To measure the relative movement of protein subunits use double cysteine constructs that contain acceptor and donor fluoro fours.

The fluorescence intensity at these residues is insensitive to the confirmational state of the protein and will be a function of the distance between the two fours. Next, change the filter set in the microscope to a 4 75 a F 40 excitation filter, a 5 95 a F 60 emission filter, and a 5 0 5 DRLP dichroic mirror. Then place the OI in the chamber on the fluorescence microscope stage.

Next, the donor is excited with a 100 watt tungsten light source and the fluorescence intensity of the acceptor. Fluoro four is measured using an appropriate method such as voltage ligand or solution exchange, activate the membrane protein and simultaneously measure the change in fluorescence intensity. This figure shows the correlation of ion transport across the cell membrane and changes in fluorescence intensity.

The upper trace shows current clamp measurements in the presence of sodium test solution and potassium test solution in the presence of 10 micromolar and 10 millimolar ing. The lower trace shows changes in fluorescence intensity measured in tandem with the current clamp measurements. This figure shows two oocytes labeled with TMRM, the acceptor fluoro four.

The oocyte on the left expresses wild type sodium potassium atpa, while the oocyte on the right expresses sodium potassium APAs with one accessible cysteine residue. The upper traces of this figure show transient current data upon voltage pulse. The lower traces show real-time recordings of fluorescence intensity changes upon voltage pulse.

This figure shows the time dependence of photobleaching donor photo destruction is measured in the absence and presence of acceptor fluoro four photo bleaching occurs more rapidly without an acceptor fluoro four. Each trace is the average of four oh site recordings. This figure shows relative subunit movement upon confirmational changes.

Fluorescence changes are shown in the red traces and current flow is shown in the black traces. An increase in fluorescence intensity shown on the left indicates the flora floors move closer together. No change in fluorescence intensity shown in the middle indicates the Fluor four distance remains static.

A decrease in fluorescence intensity shown on the right indicates the Fluor fours move further apart While attempting this procedure. It's important to remember to correlate changes in kinetic and steady state fluorescence intensity to the confirmational changes in the protein.