Single Molecule Fluorescence Microscopy on Planar Supported Bilayers

In this video, we will show how to prepare and characterize bio functionalized planar supported lipid bilar. We will also describe in detail the architecture of a total internal reflection microscope with single molecule detection capabilities. We will measure the excitation power density, the fluidity, And the protein density of the lipid blay.

Let's begin with the short introduction To planar glass supported lipid blay. When small uni lanar vesicles encounter a clean glass surface, they burst and spread out to form a contiguous lipid bilayer. On a water cushion, we typically choose palm material.

Ole phosphocholine or POPC has the predominant lipid because it gives rise to fluid lipid bilayers at room temperature and 37 degrees Celsius. For functionalization, we add nickel antiox, a synthetic lipid with a nickel chelating head group, which binds poly histamine tag proteins. For example, the co-stimulatory molecule B seven one, the adhesion molecule, ICAM one, and the fluorescently labeled peptide loaded MHC molecule for T-cell recognition as shown here, however, this bilayer system can in principle be employed to address entirely different processes in a variety of biological systems.

Due to the nature of the planar system, fluorescence events can be monitored in total internal reflection mode, which cuts down cellular background substantially and is hands very well suited for single molecule fluorescence microscopy For lipid preparation, one Needs the lipids, POPC and nickel antiox chloroform, and a clean round bottom flask dissolve the lipids in the desired ratio and chloroform and dry them under vacuum as shown here. After high vacuum treatment overnight, the lipids form a film on the inside of the flask. For lipid rehydration, use Degas PBS at 10 milliliters of the Degas PBS to the dried lipid Mixture.

Do not mix. Now to avoid lipid oxidation, fill the flask with an inert gas such as nitrogen or argon swivel the flask. The lipids form a milky suspension.

Take a small sample as a blank for the spectrophotometer to prevent gas exchange during the next step. It is very important to properly seal the flask containing the lipid suspension. Use a heat stable sticky tape, for example, autoclave tape where air protection during physical formation using an enclosed ultrasonic bath.

After mounting the flask, start the sonication procedure at a maximum power of 170 watts. Sonic Cade for 60 minutes. Due to physical formation, The turbidity has dropped significantly.

This can be verified by spectro photometry at 550 nanometers. Use the aliquot taken prior to sonication as a blank. The optical density at 230 nanometers, which is indicative of lipid Quantity should remain constant.

Here We fill small ultra centrifugation tubes with one milliliter visl suspension and place them into a fixed angle rotor of a tabletop ultracentrifuge. It is very important to use only small uni laina vesicles for bilayer preparation. Larger and more dense multi laina vesicles, which are also formed during sonication, interfere with bilayer fluidity and must therefore be paled And removed by centrifugation.

First, we spin for one hour at 150, 000 G at room temperature. The resulting snat is then centrifuged for additional eight hours at 288, 000 G At four degrees Celsius. Glass lights must Be very clean before lipid vesicles can spread on them to form a contiguous and fluid bilayer.

For this, we place cover slips into a Teflon mound, Which we then put into a glass vessel exercise. Great caution where eye protection, protecting gloves and a lab coat and work in a chemical cabinet. We add one part 30%hydrogen peroxide, and then three parts concentrated sulfuric acid to give rise to very reactive and oxidative perio mono sulfuric acid.

This mixture reacts violently with organic substances, including those of your skin eyes and clothing. It Heats up immediately. Incubate the slides for 30 minutes.

Grab the cover slip with forcet Thoroughly wash off the acid on both sides of the clean cover slip using W distilled or high quality unionized water. Then place the slide on a Teflon stand and let it dry for no longer than 10 minutes to be able to produce several individual lipid bilayers at once. We glue dried cover slips with fast hardening.

Two component epoxy glue to either an eight well or a 16 well lab tech chamber. First, remove the original lept glass slide Carefully from its frame. Mix the two glue components in a one to one ratio.

Spread it evenly into the bottom grooves of the chamber frame. Place the clean cover slip onto the glue covered bottom of the chamber frame. Make sure the glue is evenly spread between the glass slide and the plastic frame.

Let the glue harden for 30 minutes. Dilute the lipid physical suspension in PBS one to 10 and filter it through a 0.2 micron filter at 120 microliters of this dilution to each well of an eight well chamber or 60 microliters to each well of a 16 well chamber. Make sure the entire glass surface is Covered and wait for 20 minutes.

The bilayer has now formed To get rid of residual les rinse each well with 30 milliliters of PBS. Avoid air exposure of the glass surface at any cost. This would instantly destroy the bilayer to ensure Each well contains the same volume of buffer For the following protein decoration step, fill each well completely and remove the liquid dome.

Take out 330 microliters. This will leave about 350 microliters within each. Well to decorate the bilayer with proteins, prepare a master mix containing poly histamine tech proteins of choice and add 50 microliters of it to each well.

Proteins bind to the nickel anti head group of the dark lipid to ensure reproducible results mixed thoroughly but carefully and always incubate for 60 minutes to remove unbound proteins. Wash again with 30 milliliters of PBS. Let us now focus on the microscopy hardware.

First, we will illustrate the requirements of total internal reflection microscopy for objective based turf microscopy. One needs an oil immersion objective with a high numerical aperture. The incoming laser excitation beam is reflected by the dichroic mirror and focused via a set of two or three lenses onto the back focal plane of the objective only.

Then the laser light leads the objective as a coated beam along the optical axis. This way, the fluorescent specimen is illuminated in its entire depth. When you translocate the beam perpendicularly to the optical axis, the beam remains collated, but leaves the objective in a tilted fashion.

If you continue translocating, you will reach at some point the critical angle at which the beam is totally reflected at the interface between the glass cover slip and the specimen. The intensity of the resulting evanescent wave decays exponentially and only floor fours, which are in close vicinity of the interface up to several hundreds of nanometers are excited. Now, we illustrate how to focus the laser beam manually into the back focal plane of the objective.

Using three or two lenses with the first two lenses, the diameter of the laser beam is increased. The ratio of their focal length determines the size of the illumination spot at the specimen plane. To ensure that the beam stays collated adjust the distance between these lenses so that it equals the sum of both focal length.

The third lens is used to focus the widen beam into the back focal plane of the objective. Because the focal length of this lens needs to accommodate the dimensions of the microscope body and that of an associated periscope, it must be much longer in our case, 75 centimeters varying. The distance between lens two and three has no influence on the beam properties.

Therefore, the infinity space in between can be used to modify the beam in a defined manner without distortions. For example, you can insert an aperture and project precisely onto the specimen plane. In essence, you can achieve the same with the use of two lenses instead of three.

The two lens system is less defined but introduces fewer lens aberrations. Simple features such as an aperture Can still be very well implemented for temporal Modulation. A gas, iron, or solid state laser should be placed in front of a millisecond range shutter, for example, a mechanical shutter or an al optical modulator.

A good alternative is a diode laser, which can be electronically modulated and does not need external shutting. Two mirrors are sufficient to target the beam to any place and in any direction. As discussed before, the two lenses are used to widen the beam and to focus it on the back focal plane of the microscope.Objective.

We can translocate the beam for turf configuration by shifting the lower mirror of the periscope on an optical rail. For image detection, we use a highly sensitive camera. For example, an electron multiplying charge coupled device or em CCDA second laser beam of a different wavelength can be aligned into the beam path.

With the use of mirrors and a dichroic overlay. For simultaneous two color detection, we place a commercially available beam splitting device into the emission path. For experiments, it is very useful to have two independent excitation beam path in place.

For example, one for turf imaging and a second for targeted photobleaching or activation. This can be readily achieved by implementing a set of 2 50 50 beam splitters. As shown in our illustration, two different lens and mirror adjustment systems will give rise to two individual beam profiles and beam angles.

For example, a widened one for imaging in turf and a more focused one for bleaching or activation in non turf. An aperture can be placed into the non turf beam to precisely shape the bleaching or activation spot. Two additional shutters, one for each beam path Enable independent control switch on the laser beam, leaving the objective in a straight non tilted path.

Turn On the power meter and adjust it to the corresponding Laser wavelength. Measure The power of the laser light at the objective and write it down. Now, translocate the mirror behind the microscope to tilt the laser beam at the objective into the turf position.

Never look directly into the laser light. In the lower right corner, you see the intensity profile of the excitation laser as visualized through the emission of a bilayer decorated with fluorescent proteins. Here we use a false color representation to highlight regions of different intensities.

Measure the average intensity of the background within a large enough region of interest. This value depends on specific camera settings such as EM gain and readout speed and will have to be subtracted from all measured intensities. Determine the average intensity within the entire illuminated area.

Choose the diameter large enough so that intensity values at the periphery are similar to that of the camera background measure the average intensity of the central area. This area defines the region of interest in which microscopy experiments will be performed later. Now, subtract the average background intensity value from the intensities of both the eliminated and central area.

Next, multiply the background corrected intensities with the corresponding pixel number of the region size to arrive at integrated intensities. Set the integrated intensity of the illuminated area as one and calculate the fraction for the central area. The power measured with the power meter.

In our case five milliwatt is directly proportional to the integrated intensity of the whole illuminated area. Multiply this value with the power fraction of the central area to determine the power within the central area. In our example, 1.15 milliwatt, the factor to convert the number of pixels to square microns depends on the actual pixel size of the camera chip and the magnification of the emission pathway.

It can be easily determined with a micrometer slide. In our example, the central area corresponds to 170.8 square microns. The power density refers to the measured power divided by the area and thus equals 1.15 milliwatt divided by 170.8 square microns or 0.007 milliwatt per square micron.

That is 0.7 Kilowatt per square centimeter. To determine the fluidity of The bilayer, we perform a fluorescence recovery after photobleaching of wrap experiment. As shown on the lower left movie, we image the fluorescent bilayer at low power density.

On the lower right movie, you can see the corresponding lab tech chamber on the microscope. We then bleach a circular area with the short high power density pulses and monitor the recovery at low power density. Again, the montage provides an overview of the experiment for different time points with zero time being the bleach pulses.

We quantitate the average background corrected fluorescence intensity within the mark circle and normalize all measured intensities to the intensity at the beginning of the experiment. Prior to the bleach pulses, we plot the normalized intensity values against time. The saturation value represents the mobile Fraction in our case over 90%First, take an image of The bilayer at low excitation power density.

For example, by placing an optical density filter with a logarithmic attenuation value of two, this reduces the power by a factor of 100. The exposure time should be constant throughout the entire procedure and sufficient to image single fluid force at the non attenuated power density in our case, 10 milliseconds at 0.7 kilowatt per square centimeter. Use false color representation to highlight areas of different intensities.

Choose one region of interest in which single molecule measurements will be later performed and one region of the same size. For background subtraction, calculate the background subtracted integrated intensity. Now remove the optical density filter from the excitation beam path, bleach the area and take an image.

You will see individual flu force diffusing in the bleached area. Single flu force are diffraction limited with a typical diameter of two to three pixels, take one region of seven by seven pixels around each single molecule signal and the second region close by. For background subtraction, it is important to quantitate the signal of single Fluor force only.

Therefore, it is best to image proteins on the BI layer with a to protein ratio of less than one or for SAT specifically labeled proteins. Even more diet to protein ratio of one as shown in our example, measure the integrated intensities for both regions and subtract the background from the signal to not take the last observed event shown in our example as bleaching has probably occurred during exposure average. The corrected signals here, we do this for one flu of four, which we observe over nine frames.

However, this procedure should be performed for at least several hundred single molecule events. Let's go back to our original bilayer image observed at attenuated excitation power density. The molecular density is determined by correcting the integrated intensity of the region of interest for low excitation power density through multiplication with a hundred and dividing it by the average single molecule intensity and the area.

We are dealing with a density of 421.4 Fluor force per square micron. In our example, with a diet to protein ratio of one, this equals to the same number of molecules per square Micron. After watching this Video, you should be able to prepare and characterize a protein functionalized lipid bilayer using single molecule fluorescence microscopy.

Furthermore, you should be acquainted with the basic principles to modify and upgrade a standard microscope for this purpose. We will later use this methodology to determine the binding kinetics of bilayer bound antigen to cell bound T-cell receptors, but many other applications based on this are certainly Feasible.