Biology
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Three-dimensional Super Resolution Microscopy of F-actin Filaments by Interferometric PhotoActivated Localization Microscopy (iPALM)
Chapters
Summary December 1st, 2016
We present a protocol for the application of interferometric PhotoActivated Localization Microscopy (iPALM), a 3-dimensional single-molecule localization super resolution microscopy method, to the imaging of the actin cytoskeleton in adherent mammalian cells. This approach allows light-based visualization of nanoscale structural features that would otherwise remain unresolved by conventional diffraction-limited optical microscopy.
Transcript
The overall goal of this procedure is to visualize the actin cytoskeleton in adherent mammalian cells at the ultrastructural level, using a three-dimensional super resolution microscopy method called iPALM. iPALM imaging provides sub-trending nanometer spatial resolution in all three dimensions. This enables ultrastructures to be visualized using florescence probes, allowing high labeling specificity.
The procedure presented in this video should be rapidly adaptable and instructive for visualizing other ultrastructural features in cells by single molecule localization based in super resolution microscopy. Begin this procedure by preparing the cells on cover glasses in a six well plate. To assemble the imaging sample, use fine forceps to gently remove the specimen cover glass from the buffer well.
Then quickly and gently tap away excess buffer by touching the edge of the cover glass with folded, absorbent paper. Place the cover glass cell side facing up on a piece of clean lens paper. Rinse the sample by placing 30 to 50 microliters of the imaging buffer onto the sample.
Remove excess buffer by tilting and tapping with folded absorbent paper. After repeating the rinsing step a few times, place 30 to 50 microliters of imaging buffer onto the sample. Blot dry the edge of the cover glass and place multiple very small dots of fast curing epoxy onto the dried area.
Slowly lower another pre-cleaned number 1.5 cover glass onto the center of the cell containing 22 millimeter cover glass. Let the imaging buffer wet both cover glasses by capillary action. The small dots of fast curing epoxy should adhere to both cover glasses.
Gently press upon the assembled sample using folded absorbent paper to spread the pressure evenly. Use sufficient pressure to make the sample cell thin and even, but not so much as to crush the cells. Gauge the proper thickness by observing the Newton's rings pattern.
Make sure to perform this step gently to minimize air bubbles. If needed, practice with empty cover glasses several times beforehand. Next, seal the sample with melted Vaseline lanolin paraffin.
Then rinse the sealed sample with deionized water. Blow dry the sample with compressed air to make ready for mounting onto the microscope. Move the spring-loaded top objective lens upward to allow for the removal of the sample holder.
Place the sealed specimen onto the sample holder and secure with several small rare-earth magnets. Apply immersion oil on both sides of the imaging sample. Place the sample holder back into the optical path and gently lower the top objective lens.
Turn on the excitation lasers. Also turn on the EMCCD cameras in frame transfer mode. Next, rotate in the proper emission filters.
Activate a mechanical shutter to block the top beam path and open the bottom beam path. Bring the bottom objective lens into focus by translation in small increments using the piezo actuator. Once the fiducial is in focus, open the top beam path while blocking the bottom beam path and bring the top objective lens into focus in a similar manner.
Monitor the width of the fiducial on the computer display for optimal focus. For proper centration, open both the top and bottom beam paths. Manually adjust the top objective lens while the bottom objective is held constant using a pair of microfine set screws until the fiducial images are overlapped as closely as possible, ideally within one pixel.
Subsequently, perform fine adjustments so that the fiducial images overlap within one tenth of an EMCCD pixel. Adjust the top two axis piezo mounted mirrors via the control software while holding both objective lenses and the bottom reflection mirrors constant. Compare the centers of the fiducials of the top and bottom objective views via the computer display to guide the process.
With the lasers on, the cameras continuously streaming, and both the top and bottom beam paths open, oscillate the sample holder Z piezo using a sinusoidal voltage waveform generated by the control software for a continuous Z-axis oscillation over a magnitude of 400 nanometers. Manually translate the motorized beam splitter assembly up or down until the intensity of the fiducial oscillates. Due to the desired single photon interference effect, this signifies the close matching of the optical path lengths.
A peak to valley ratio of greater than 10 can be achieved in optimal cases. To ensure that both the amplitude and phase at each surface are as uniform as possible across the field, adjust the bottom mirror of the beam splitter assembly in small steps to fine tune the gap and the tilt angles. Perform beam splitter fine alignment by translating the sample in eight nanometer Z steps over 800 nanometers.
Monitor the fiducial intensity among cameras one through three. Adjust the height, position, and tilt of the bottom mirror within the beam splitter assembly in small steps such that the oscillation phase of camera one relative to camera two is maximized, ideally at 120 degrees. Once the initial alignment is complete, translate the sample to scan for a suitable field of view that contains both cells to image and multiple fiducials nearby.
Place an enclosure around the system to block stray light and ambient perturbations. Once the imaging area is found, translate the sample again and record the calibration curve for use in subsequent Z coordinate extractions. Using the command acquire calibration scans versus sample piezo position in the main interface.
Once a desired area is found and the calibration curve is obtained, enter the appropriate file names into the software. Open both the top and bottom beam paths, then increase the excitation power of the 642 nanometer laser to the maximum. For Alexa Fluor 647, an initial period of Fluor-4 switching off may be needed.
Expose with a constant 642 nanometer excitation for five minutes, or longer if necessary, until single molecule blinking is observed. The software allows an automatic increase of 405 nanometer photoactivation during the course of the acquisition. Large numbers of acquisition frames are usually required for filamentous features to be clearly visible.
When ready, commence the acquisition of raw image sets using the command start iPALM acquisition in the main interface. During the acquisition, tune the photo activation level by adjusting the intensity of the 405 nanometer laser to maintain the proper blinking density as needed. When the sample cover slip is assembled properly, Newton's rings can be observed by naked eyes under ambient light or standard fluorescent lighting.
Extraction of the axial positions of fluorescence emitters requires simultaneous multi-phase interference in iPALM imaging, which is calibrated by the intensity changes of a given fiducial in three EMCCDs. The intensities are then normalized and fit to determine the phase differences. Here is the reconstruction iPALM image with each localization coordinate rendered by a 2D Gaussian with the width corresponding to localization uncertainty.
The Z coordinate is colored according to the color bar. Areas of dense and sparse filamentous features can be seen. Shown here are reconstruction iPALM images of actin cytoskeleton of HUVEC cells labeled by Alexa Fluor 647 phalloidin.
The image color indicates the Z coordinates according to the color bar. The transverse cross section histogram is generated from the XY localization coordinates along the long axis of the boxed area. Gaussian fitting shows full-width at half maximum of 43.88 nanometers associated with the transverse cross section.
This histogram is of the Z position of localization coordinates in the boxed area. Gaussian fitting shows full-width at half maximum of 17.50 nanometers. Once mastered, this technique can be done in 90 minutes if it is performed properly.
After watching this vide, you should have a understanding of how to visualize the actin cytoskeleton at the ultrastructural using iPALM through simple preparation and mounting, optical alignment, raw data acquisition, and the subsequent analysis for reconstructing super resolution images. After its development, this technique paved the way for researches in the field of cell biology to explore the nanoscale architecture of the actin cytoskeleton, as well as cell adhesion structures such as the focal adhesions.
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