Multi-Color-Lokalisierung Mikroskopie des einzigen Membranproteine in Organellen der lebenden Säugetier-Zellen

This technique can answer key questions in regulatory processes such as which role protein dynamics and organization play in functional adaptation. The main advantage of this technique is that it can be done in living cells, thereby revealing protein dynamics on the single-molecule level. Though this method was originally implemented to provide insights into the spatiotemporal organization of mitochondrial membrane proteins, it can also be applied for other organelles and other membrane proteins.

Demonstrating the technique will be Timo Appelhans, a grad student from my lab. After transfecting and preparing cell samples and the microscope according to the text protocol, mount the prepared sample with fluorescent beads in the sample holder between the PTFE ring and the red rubber ring. Start the microscope, all hardware components, and all software needed for microscopy.

Use a fresh, lint-free tissue sprayed with isopropanol to clean the objective lens and the bottom of the cover slip. Then place a droplet of immersion oil on the pupil of the objective lens. Next, place the sample holder with the bead sample onto the microscope stage so that the bottom of the cover slip contacts the oil.

Then focus on the beads by using transmission light or a laser line. Adjust the power of the two excitation lasers to achieve similar signal density in the two fluorescence channels. Then search for an area with many distinct fluorescent signals.

Generate a merged view of the fluorescent channels by using the camera control software. Then use the screws at the image splitter to manually tilt the internal optical image splitter to achieve the best overlay of the signals from the two fluorescent channels. Start the TIRF microscope-controlling software and choose to display individual channels in live stream mode.

Take a snapshot image, exciting fluorescents in all channels. Then use this image to produce the transformation matrix. Start the software analysis plugin and load the previously-recorded dual-color images of fluorescent beads into the software.

Choose the used orientation of the fluorescent channels. Click Yes when asked for calibrate images, and select the previously-taken snapshot. Open the unit manager to define unit conversion factors, then open the localization manager.

To determine the point spread function, or PSF, press the button PSF Radius. In the PSF Estimator window that opens, define the numerical aperture and the emission maximum. Then click Estimate PSF Radius.

Next, accept the obtained experimental PSF. Define the evaluation box, number of deflation loops, and how many cores of the computer are used for calculation. Then press Localize to start fitting the intensity distribution of single particles, using a 2D symmetric Gaussian function.

Open the calibration manager. In the rendered, merged image of the two channels, the original signal and the localized centers will be shown. Choose Affine mode.

Then manually connect the corresponding pairs of localized centers in the two channels, that have originated from the same fluorescent bead, by drawing a connection line. Connect the corresponding signals distributed over the field of view. Then press Accept, and save the transformation matrix.

To carry out single molecular imaging of mitochondrial membrane proteins, mount the specimen cover slip between the rubber and PTFE rings. Then fill the chamber with 0.5 to 0.8 milliliters of imaging medium. Clean the objective lens and bottom of the cover slip.

Then use transmission light or a laser line to focus on the cells, after adding a drop of oil to the lens. Then adjust the power of the two excitation lasers to achieve a similar intensity, and search for an area with many fluorescent signals. Adjust the illumination angle in the TIRF microscope-controlling software to create an incident angle that is smaller than the critical angle, or the TIR-mode, to excite the specific region of interest via a HILO sheet.

Adapting the illumination of the TIRF microscope to excite cell organelles using the HILO illumination requires quick judgment of the signal-to-background ratio and experience in TIRF microscopy, as well as a knowledge of cell morphology. Set the EM gain and choose an exposure time suitable for the experiment that collects sufficient photons per frame. Then adjust the laser power in order to achieve a high signal-to-noise ratio, as this ratio directly corresponds to the localization precision.

Find an area in the cell periphery with non-overlapping, elongated mitochondria and single-molecule signals. If no single signals are visible in one or both channels, wait until bleaching results in the appearance of single-molecule signals. In the case only one channel needs to be bleached, switch the laser of the other channel off, to avoid bleaching, before recording this channel.

Record until the number of signals is too low for reasonable continuation. Start the imaging-processing software and check for mitochondrial structures by generating a cumulative rendered-sum image of at least 1, 000 recorded frames. Start the software analysis plugin.

To carry out data processing for localization and tracking, load the raw data. Check multicolor. Choose the correct orientation of both channels.

When asked for Calibrate images, load the transformation matrix. The channels will be separately displayed. Open the raw data from the image container by clicking Show.

Then open unit manager in the top tab, and define unit conversion factors in each channel. Open the localization manager in the top tab, and define the evaluation box and number of deflation loops. Click on PSF Radius to open the PSF estimator.

Set numerical aperture and emission maximum to get the theoretical PSF value. Press Accept. Repeat this for all channels.

Then press Localize to start the single-particle localization for the recorded signals in both channels. Wait for the software to finish. Now, a super-resolved image for each channel is obtained in the image container of the software analysis plugin.

PINK1 is an important factor guaranteeing mitochondrial functionality. To determine the localization of PINK1 relative to Tom20 in functional and dysfunctional mitochondria, a time series of 1, 000 frames was recorded. The cumulative image of all signals over time from one channel showed that PINK1 was localized in the cytosol, at the outer membrane, and inside mitochondria under normal conditions, but preferentially in the outer membrane of depolarized mitochondria.

As the summed images of localized particles show, PINK1 is distributed mainly inside polarized mitochondria, while Tom20 is localized in the outer mitochondrial membrane. This becomes even clearer in the merged image of localized Tom20 and PINK1 particles, which shows that the distribution of the outer membrane protein Tom20 is much broader. To verify the localization of PINK1 inside mitochondria, the 30 kilodalton subunit of respiratory complex one was fused to photoactivable GFP and recorded by fluorescence photoactivation localization microscopy.

The cumulative triple-color image and the cross-section distribution show the overlay of complex one and PINK1, confirming the import of full-length PINK1. These data demonstrate that subpopulations of PINK1 occupy several mitochondrial micro-compartments. Once mastered, this technique can be done in 20 hours, including recording and post-processing of data, if it’s done properly.

While attempting this procedure, it is important to remember to test for organelle movement. Following this procedure, the relative localization of proteins can be determined. After its development, this technique paved the way for researchers in the field of single-particle tracking of mitochondrial proteins to observe their behavior in Z2, and different metabolic conditions, and under stress.

After watching this video, you should have a good understanding of how to perform dual-column microscopy of mitochondrial proteins to reveal their sub-organelle position and trafficking.