Simultaneous Multicolor Imaging of Biological Structures with Fluorescence Photoactivation Localization Microscopy

The overall goal of this procedure is to image multiple protein species simultaneously with nanometer precision in fixed or living cells. This is accomplished by first adjusting the position of a camera and optics until a focused image from the microscope is projected onto the camera chip. The second step is to arrange laser beams so they're directly aligned onto a sample on the microscope stage.

Next, the prepared cell sample is illuminated and a cell expressing the desired proteins is chosen. The final step is to image the cell with fluorescence photo activation localization microscopy by illuminating the sample with lasers, and then directing the cell fluorescence to the camera chip to acquire a dataset set. Ultimately, fluorescence photo activation localization microscopy is used to show localization of multiple protein species at the nanometer spatial scale in fixed or living cells and in either wide field or when using total internal reflection fluorescence to isolate a thin region of the sample.

The main advantage of this technique over existing methods such as confocal or electron microscopy, is that one can image multiple proteins simultaneously with nanometer resolution in fixed or living cells. The following steps refer to a numbering system as shown here, which is a schematic for the multicolor F Om setup provided as figure one in the accompanying text protocol to align the microscope begin by placing a calibration scale or radical onto the microscope stage with a 10 x objective in place and the lamp set for transmitted light. Center the vertical in the center of the field of view.

Next, adjust the microscope for coer illumination by closing the field aperture and looking through the oculars at the vertical. If the edges of the field aperture are out of focus, adjust the height of the condenser until both the field aperture and redle are in focus. Then adjust the lateral position of the field aperture until it is centered with respect to the field of view and close the field aperture until only the center grid on the redle is illuminated.

The first time that these components are assembled, adjust the path lengths of each channel to be equal. To accomplish this project, the redle onto the camera chip adjust mirrors seven and nine and close the detection aperture to prevent spatial overlap between the two channels. Then focus the image of the radical in the reflected light channel and check to see if it is also in focus in the transmitted light channel.

If the image in the transmitted light channel is not in focus, translate mirror nine until the radical image is in focus simultaneously in both channels. Start aligning the lasers by removing lens one from the laser path and blocking the activation and readout beams. Then place a white card flush against mirror four, open the microscope shutter and focus until the radical image projects onto the card in front of mirror four.

Next, unblock the readout beam and adjust mirror one to center the readout laser onto the cross hairs of the radical image on mirror four. Once centered, project the radical image onto mirror five and adjust mirror four until the beam is centered on the image cross hairs of mirror five. The readout beam should now be centered at both M four and M five.

Then block the readout beam by closing shutter one. Next, project the radical image onto mirror three. Remove the beam expander from the laser path and unblock the activation laser.

Now adjust mirror two to center the activation beam onto the cross hairs of the radical image on mirror three. Once centered, replace the beam expander between mirrors two and three and adjust the position of the beam expander until the beam is centered on the cross hairs of the radical image. On mirror three, project the radical image onto mirror five and focus.

Using the microscope's focus knob, adjust the angle of the dichroic mirror number one until the activation beam is centered on the redle image. Then block activation laser by closing shutter two with no objective in place and the microscope shutter open. Open the readout shutter shutter one and project the laser through the back aperture of the microscope.

Adjust mirror five until the beam emerges from the microscope, so the beam exits the microscope vertically with lens one and the objective back in place. Place a sample containing 100 micromolar bromine B on the stage with the activation laser blocked. Project the readout laser through a 60 x objective and into the dye.

With the electron multiplying gain disabled. Send this image to the camera. Next, focus the objective into the sample.

Open the aperture wide enough to allow imaging of the full beam profile. Then translate the aperture laterally so that the center of the beam profile and aperture are concentric. Using the camera software, choose the region of interest to allow the smallest camera readout region encapsulating both channels and record these coordinates.

At this point, record a single snapshot to represent the readout beam profile. Next, block the readout laser by closing shutter one and open shutter two. In order to begin to measure the activation beam profile project the activation laser to the sample, if necessary, use an electron multiplying gain of less than 100 and adjust the first dichroic mirror until the beam is centered in each field of view.

Then record a snapshot of the activation beam profile in order to begin acquiring multicolor F palm images. Eliminate all room lighting. Then project the mercury lamp via the flip mount onto transfected cells and change the filter cube to one containing the appropriate excitation wavelength of the pre photo switch.State.

Once a cell is chosen, move the flip mount down in order to allow the lasers to pass into the microscope, change the filter turret to that containing the appropriate dichroic mirror for imaging as described in the accompanying text protocol. Then project this image to the camera to distinguish transfected cells from background fluorescence and to confirm that molecules are photoable. Briefly illuminate the sample with a low power of less than 10 micro watts from the activation laser.

Next, prepare the camera software for kinetic series acquisition by setting the electron multiplying game to 200 and the desired number of frames to between five and 10, 000. Also set the exposure to between 10 and 30 milliseconds. Then block the activation beam, unblock the readout beam, and project the image of the illuminated cell to the camera.

Choose a focal plane near the bottom cellular membrane by shifting the focus down until the individual molecules are no longer visible. Then gradually move the focus upward until molecules first become visible. Now unblock the activation beam and illuminate the sample with less than one watt per squared centimeter of intensity.

Begin to acquire these data through the camera software while maintaining a density of 0.1 to one visible photoable molecules per square micron by dynamically adjusting the neutral density filter in front of the activation laser. For total internal reflection fluorescence imaging mount mirror five and lens one onto a single translation stage to be moved laterally just behind the entrance to the microscope. As mirror five and lens one are translated, the lasers exiting the objective upward through the sample will gradually tip to one side, continue to translate the stage until the angle of the lasers reaches 90 degrees from the vertical.

At this point, the emerging laser itself will vanish and the incoming laser will be back reflected into the aperture traveling anti-parallel to the incoming beam and displaced to the side. You should see a reduction in background when the sample enters total internal reflection fluorescence upon completion of the image acquisitions, close the microscope shutter immediately and block both beams. Disable the electron multiplying gain.

Set the camera to record one frame and set the camera readout region to its maximum size. Finally, block one channel by placing a card over F three or F four with a long pass filter mounted on the microscope lamp. Illuminate the sample and project this image to the camera.

Record a snapshot of the cell to represent the whole cell in transmitted light. Shown here is an example of a two color F palm acquisition of an NIH three T three cell expressing both DENDRA two Hemagglutinin and PAM Cherry actin. The transmitted channel on the left contains longer wavelengths than the reflected channel on the right.

Shown here are snapshots from the same two color F OM acquisition following background, subtraction, and transformation to overlay the left and right channels. Individual molecules can be identified and localized. Some molecules appear brighter in the transmitted channel and some have a more even distribution of emission between the two channels.

This indicates the difference in emission spectra between PAM cherry and DENDRA two respectively and is used in analysis to identify these two species. The histogram shown here indicates a ratio of red transmitted channel intensity divided by total intensity for all localized molecules after tolerances had been applied. The pixels appear white in the lower left image and appear red in the final merged image shown on the lower right where those in the green region are shown as white in the upper right image and appear green in the final merged image shown on the lower right.

The threshold levels that are chosen when rendering greatly affects the degree of noise and appearance of colocalization.Shown. Here are three different options for rendering which result in varying degrees of bleed through the most conservative threshold levels are shown on the bottom two. Color F palm alpha histograms are best to interpret when there are two discernible peaks in the image.

While the histogram on the left is a good candidate for further analysis, the images resulting from the other two histograms will be much more difficult to interpret Following this procedure. Methods like total internal reflection, microscopy and live cell imaging can be for performed in order to answer additional questions like how proteins are organized at the nanoscale in living cell membranes. After watching this video, you should have a good understanding of how to set up your own FAR microscope and use it to acquire data.

Don't forget that working with lasers can be extremely hazardous, and safety training should be completed before attempting this procedure.