Thermodynamique du repliement des protéines membranaires mesuré par fluorescence Spectroscopie

The overall goal of the following experiment is to measure the thermodynamics associated with membrane protein folding. This is achieved by first assembling small LAR vesicles to provide a folding environment for the membrane protein. Once constructed, vesicles are used to prepare samples containing protein and systematically varied concentrations of urea.

Next, the tryptophan fluorescent spectrum of each sample is measured In order to gather data required to generate a membrane protein unfolding curve results are obtained that show how the Gibbs free energy of membrane protein unfolding can be obtained based on unfolding curves generated from tryptophan fluorescent spectra. Hi, my name is Judy Kim. I'm an assistant professor at the University of California at San Diego.

The method that you're about to watch reveals an important property of membrane proteins. Specifically, we'll show you how to measure the stability of a protein that is folded and inserted into a lipid bilayer. My name is Diana Ottinger, and I'm a graduate student in the Department of Chemistry at uc, San Diego.

Today, we'll demonstrate this method on a representative membrane protein. However, this technique is general and may be applied to other biological systems such as membrane associated peptides and soluble proteins. So let's get started.

Prior to the start of this protocol, purchase a solution of one two DME oil, SN glycerol, three phosphocholine or DMPC, lipid in chloroform and Eloqua into clean glass vials. At 20 milligram per vial quantities, add a layer of nitrogen gas to each vial to prevent lipid oxidation and seal the vials. A single vial containing 20 milligrams of lipid in chloroform is used for each experiment.

To prepare the stock vesicle solution begin by drying the contents of the vial under a stream of nitrogen gas for one hour, using a Teflon septum and needles until no solvent remains. Add one milliliter of 20 millimolar potassium phosphate buffer, pH 7.3 to the dried lipids and resus suspend in a water bath ator for approximately 30 seconds. Transfer the resulting cloudy suspension to a plastic 15 milliliter tube with a conical bottom pipette.

An additional one milliliter of phosphate buffer to the empty glass vial that originally contained the lipid, and repeat the sonication. Add the resulting suspension to the same 15 milliliter tube. Repeat this process a total of four times to obtain a final volume of four milliliters and a lipid concentration of five milligrams per milliliter.

Place the lipid vesicle suspension into a warm water bath and sonicate using an ultrasonic ator micro tip for one hour at 50%duty cycle. The water bath serves to prevent the lipid solution from becoming too hot and keeps the temperature of the aqueous lipid solution above the bi-layer phase. Transition temperature during sonication.

Filter the SONICATED solution through a 0.22 micron syringe filter to remove debris from the ator tip and allow the filtered solution to equilibrate overnight at 37 degrees Celsius. To prepare the sample for initial fluorescence unfolding, make a stock solution of 10 molar urea in 20 millimolar phosphate buffer and a stock buffer solution of 20 millimolar phosphate. According to the written protocol, the urea concentration should be determined experimentally by measuring refractive index with a refractometer and comparing to known values.

Next, prepare a stock solution of unfolded protein at a concentration of approximately 200 micromolar protein in eight molar urea, 20 millimolar phosphate buffer. In this video, the membrane protein OPA A is used to demonstrate the protocol samples for fluorescent studies are prepared by combining appropriate volumes of stock protein stock lipid solution stock 10 molar urea solution and stock phosphate buffer to make samples containing approximately four micromolar protein, one milligram per milliliter lipid and zero to eight molar urea in one molar increments resulting in a total volume of 200 microliters. Blank samples are made in the same manner.

However, four microliters of eight molar urea solution is substituted for protein. These blanks are made in order to subtract scattering and background signals that appear in the fluorescent spectra of protein. Incubate the protein samples in blanks at 37 degrees Celsius for at least two hours before measurement of fluorescent spectra.

To ensure complete folding turn on the fluorimeter one hour prior to measurements. Membrane proteins generally fold into synthetic lipids only when the temperature is above the bilayer phase transition temperature. Therefore, in the case of DMPC, the temperature of the sample holder in the fluorimeter is set to 30 degrees Celsius.

Set up the fluorescence experiment on a steady state fluorimeter and measure the tryptophan fluorescence of each sample in blank. The excitation wavelength should be set to 290 nanometers to avoid excitation of tyrosine residues and the emission collected from 305 to 500 nanometers. Use an entrance and exit band pass of three nanometers.

The wavelength increment and integration time can be optimized for signal to noise ratio. Typical values for wavelength increment and integration time are one nanometer per step and 0.5 seconds per step respectively. Pipette 200 microliters of the sample into a micro volume fuse silica Q Vet with 160 microliter capacity and place the qve into the qve holder of the machine.

Finally, fluorescence measurements can begin load an XY data sheet of the fluorescence spectra into software such as Igor Pro, MATLAB origin, or Excel. For this demonstration, Igor Pro will be used to remove the urea and lipid vesicle background, multiply the fluorescent spectrum of the blank sample by the S scaler, and subtract it from the raw fluorescent spectrum of the protein lipid solution. At the corresponding urea concentration tabulate the wavelength of maximum fluorescence or lambda max from the corrected spectrum.

For each urea concentration. A typical value for a fully unfolded membrane protein is 350 nanometers. While that of a folded protein is approximately 330 nanometers, translate the tabulated wavelengths to fraction unfolded by converting the range of lambda max values to a range between zero on one correlating the lambda max value to fraction of unfolded population.

Then plot the fraction of unfolded protein against urea concentration. The plot of fraction unfolded F versus urea concentration C can be fit to an equation that assumes the protein can exist in either the folded or unfolded state. The coefficient M corresponds to the rate of change of the free energy with respect to the denature and concentration, and CM corresponds to the midpoint urea concentration at which the folded population equals the unfolded population.

R is the gas law constant, and T is the temperature. In Kelvin. The fitting procedure yields values for the coefficients m and cm.

These coefficients are multiplied to determine delta G, the Gibbs free energy of unfolding in the absence of urea. The unfolding curve described reveals the range of urea concentrations that causes the greatest change in the lambda max. This range is typically small and the majority of unfolding occurs within this small range in order to precisely determine the free energy of unfolding.

Additional samples should be analyzed in this region to obtain a plot that contains the many data points necessary for optimized lease squares fitting. For example, if the protein unfolds between two and four molar urea samples can be made between these urea concentrations in 0.2 molar increments to more precisely reveal the shape of this important region in the unfolding curve. A blank should be included at least every 0.5 molar step and every 0.2 moer step for best results.

The free energy of unfolding obtained from this plot may differ slightly from that obtained in the original unfolding curve, but reflects a more precise value shown. Here are tryptophan fluorescent spectra of the representative membrane protein om a that contains a single tryptophan residue. Raw fluorescent spectrum of the blank are subtracted from the raw fluorescent spectra of the protein as described in the protocol resulting in the corrected spectrum.

The corrected tryptophan fluorescent spectra of membrane protein for numerous urea concentrations is shown here from the methods described. In this demonstration, an unfolding curve was generated under the assumption that the protein is either folded or unfolded and that there is no stable state. The parameter M reflects the amount of polypeptide that is exposed to solvent upon unfolding, whereas CM is an important value that describes the denature and concentration required to achieve an equal population of folded and unfolded populations.

Large values of CM and M indicate a stable protein. The Gibbs free energy of unfolding was calculated to be 6.2 kilo calories per mole for OPA a Following this procedure. Other methods of unfolding such as heat, pH and other chemical Dena may be used to provide analogous information about the thermodynamics of protein folding.

After watching this video, you should have a good understanding of how to measure the stability of a protein based on the different fluorescence properties of the folded and unfolded states. Alternatively, you could also generate an unfolding curve using other methods such as circular, dim, as long as the method reports on protein confirmation, regardless of the analytical tool that you choose. The method of unfolding curves is versatile and can be applied to a wide range of membrane and soluble proteins.