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Method Article

Three-dimensional Super Resolution Microscopy of F-actin Filaments by Interferometric PhotoActivated Localization Microscopy (iPALM)

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DOI:

10.3791/54774

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December 1st, 2016

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1Department of Biology, South University of Science and Technology of China, Shenzhen, 2Mechanobiology Institute, Singapore, 3Department of Biomedical Engineering, National University of Singapore

In This Article

Summary

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.

Abstract

Fluorescence microscopy enables direct visualization of specific biomolecules within cells. However, for conventional fluorescence microscopy, the spatial resolution is restricted by diffraction to ~ 200 nm within the image plane and > 500 nm along the optical axis. As a result, fluorescence microscopy has long been severely limited in the observation of ultrastructural features within cells. The recent development of super resolution microscopy methods has overcome this limitation. In particular, the advent of photoswitchable fluorophores enables localization-based super resolution microscopy, which provides resolving power approaching the molecular-length scale. Here, we describe the application of a three-dimensional super resolution microscopy method based on single-molecule localization microscopy and multiphase interferometry, called interferometric PhotoActivated Localization Microscopy (iPALM). This method provides nearly isotropic resolution on the order of 20 nm in all three dimensions. Protocols for visualizing the filamentous actin cytoskeleton, including specimen preparation and operation of the iPALM instrument, are described here. These protocols are also readily adaptable and instructive for the study of other ultrastructural features in cells.

Introduction

The visualization of complex cellular structures has long been integral to biological insights and discovery. Although fluorescence microscopy can image cells with high molecular specificity, its resolving power is limited by diffraction to ~ 200 nm in the image plane (x,y, or lateral dimension) and > 500 nm along the optical axis (z, or axial dimension)1,2. Hence, the observation of ultrastructural features has historically been limited to electron microscopy (EM). Fortunately, the recent development of super resolution microscopy has circumvented this limit, enabling spatial resolution in the 10 - 100 nm range1-6. In particular, super resolution approaches based on single molecule localization, known by acronyms such as PALM (PhotoActivated Localization Microscopy)4, FPALM (Fluorescence PhotoActivated Localization Microscopy)5 (d)STORM (direct Stochastic Optical Reconstruction Microscopy)6,7, PAINT (Point Accumulation for Imaging Nanoscale Topography)8, GSDIM (Ground State Depletion Microscopy followed by individual molecular return)9, or SMACM (Single-Molecule Active-Control Microscopy)10, as well as their 3-dimensional (3D) implementations, such as interferometric PALM (iPALM)11 or 3D-STORM12, have been valuable in revealing novel insights into the nanoscale organization of numerous biological structures, including neuronal axons and synapses13, focal adhesions14,15, cell-cell junctions16, nuclear pores17, and centrosomes18-20, to name a few.

Another ultrastructural feature in cells for which super resolution microscopy is potentially useful is the actin cytoskeleton. The complex meshwork of filamentous (f)-actin in the cell cortex plays an essential role in the control of cellular shape and mechanical properties21. The organization of f-actin is actively and dynamically regulated though numerous regulatory proteins that strongly influence polymerization, crosslinking, turnover, stability, and network topology22. However, although the characterization of the f-actin meshwork architecture is important for mechanistic insights into a diverse range of cellular processes, the small size (~ 8 nm) of the f-actin filaments hampers their observation by conventional diffraction-limited light microscopy; thus, the visualization of actin fine structure has hitherto been exclusively performed by EM. Here, we describe protocols for visualizing the f-actin cytoskeleton in adherent mammalian cells, using the iPALM super resolution microscopy technique to take advantage of its very high precision capability in 3D11,23. Although the iPALM instrument is highly specialized, instruction on setting up such an instrument has been described recently23, while access to the iPALM microscope hosted by the Howard Hughes Medical Institute has also been made available to the research community with minimal cost. Additionally, the specimen preparation methods described herein are directly applicable to alternative 3D super resolution approaches, such as those based on astigmatic defocusing of the point spread function (PSF)12 or bi-plane detection24, which are more broadly available.

We note that a necessary ingredient for single-molecule localization-based super resolution microscopy in general is the photoswitchable fluorophore25, which allows the three critical requirements for single-molecule localization-based super resolution microscopy to be fulfilled: i) high single-molecule brightness and contrast relative to background signals; ii) sparse distribution of single molecules in a given image frame; and iii) high spatial density of labeling sufficient to capture the profile of the underlying structure (also known as Nyquist-Shannon sampling criterion)26. Thus, for satisfactory results, emphasis should be placed equally on both the proper preparation of specimens to optimize fluorophore photoswitching and to preserve the underlying ultrastructure, as well as on the instrumentation and acquisition aspects of the experiments.

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Protocol

1. Imaging Specimen Preparation

  1. Since background fluorescence signals interfere with fluorescence from fluorophore labels, clean the coverglasses by first rinsing them in de-ionized water (ddH2O) and then air-drying them using compressed air. Subsequently, perform plasma etching in a plasma cleaner for 15 sec, or longer if necessary.
  2. To enable drift correction and iPALM calibration, use #1.5 round (22-mm diameter) pre-cleaned coverglasses embedded with fluorescent nanoparticles as fiducial marks, which serve as highly photostable fiducials for reliable calibration and drift correction. Due to the long acquisition time required to accumulate sufficient fluorophore density (> 15 - 30 min), sample drift is inevitable.
  3. Place each fiducialed coverglass into a 6-well tissue culture plate. Sterilize them by ultraviolet (UV) radiation in a laminar flow hood for 15 min.
  4. In a sterile laminar flow hood, prepare fibronectin solution for coverglass coating by diluting 1 mg/ml fibronectin stock solution in sterile Dulbecco's phosphate-buffered saline (DPBS) to a final concentration 2 - 10 µg/ml. Rinse each coverglass three times with DPBS and incubate with 2 ml of the fibronectin solution overnight at 4 °C. Subsequently, aspirate out the fibronectin solution and rinse once with DPBS.
  5. Rinse the cells briefly with DPBS. Incubate with 1 - 2 ml of trypsin for a few min at 37 °C until the cells detach and quench with ~ 10 ml of fresh serum-containing cell culture medium. For example, for human umbilical vein endothelial cells (HUVEC), use large vessel endothelial culture medium supplemented with large vessel endothelial factors and penicillin or streptomycin.
    1. Replate the cells onto the fibronectin-coated coverglass (for sparse density, plate < 50,000 cells per coverglass) and maintain the culture in an incubator set at 95% humidity, 5% CO2, and 37 °C.
  6. For proper preservation of the fine structures of the f-actin cytoskeleton, use buffer solutions based on Good's buffer reagents27.
    1. For example, prepare PHEM buffer as a 2x stock solution (120 mM PIPES, 50 mM HEPES, 20 mM EGTA, 4 mM MgCl2, pH 7.0 with KOH) by dissolving 6.5 g of HEPES, 3.8 g of EGTA, and 190 mg of MgCl2 in ~ 300 ml of ddH2O, with the pH adjusted to 7.0 by dropwise addition of concentrated KOH solution; then, add ddH2O to bring the volume to 500 ml. Sterilize the buffer using a 0.22-µm filter, store it at 4 °C, and dilute it 1:1 with ddH2O before use.
  7. For best results with high density labeling of f-actin, use phalloidin conjugated with organic fluorophores, such as Alexa Fluor 647. At the desired time point after cell replating, fix the cells as follows:
    1. Aspirate the media from each culture well containing the cell specimen. Gently but quickly dispense 2 ml of warm (37 °C) extraction fixatives containing 0.25% glutaraldehyde in PHEM buffer with 0.25% Triton X-100. Incubate it at room temperature for 1 - 2 min. For subsequent steps, use 2 ml of fixative or quenching buffer per coverglass, unless indicated otherwise.
    2. Replace the extraction fixative with a 2.5% glutaraldehyde fixative in PHEM buffer and let the samples incubate for 10 - 12 min. This and following steps are all carried out at room temperature.
    3. Aspirate out the fixatives and gently replace them with PHEM buffer. Tilt and swirl gently, and then wash again with PHEM. Repeat twice.
    4. To quench autofluorescence from glutaraldehyde, which can overwhelm signals from the desired fluorophores, incubate the specimens with a freshly prepared quenching buffer containing NaBH4 at a mass concentration of 0.1% in PHEM. Profuse bubbles will be observed. Occasionally tap the sample dish gently to dislodge the bubbles. Let it incubate for 5 - 10 min.
    5. Aspirate out the quenching buffer and gently replace it with PHEM buffer. Rinse a few times to clear away the bubbles. Pipette in 2 ml of PHEM buffer and let it incubate in the dark for 5 min. Repeat twice and let the specimen rest in PHEM buffer when done.
    6. Prepare a humidity chamber for phalloidin incubation using a large plastic Petri dish padded with a piece of paper towel generously moistened with 5 - 10 ml of ddH2O. Place a large sheet of clean Parafilm on top of the wet paper towel.
    7. Due to the relatively high cost of labeled phalloidin, use a small volume for each labeling. For high density labeling, start with a concentration of 0.3 µM. Prepare ~ 60 µl per coverglass using phalloidin-Alexa Fluor 647 in PHEM buffer.
    8. Pipette 55 - 60 µl of the phalloidin solution onto the Parafilm sheet in the humidity chamber. Using fine forceps, gently remove the specimen coverglass. Be careful to note the correct cell-containing face.
    9. Quickly and gently tap away excess buffer by touching the edge of the coverglass with a folded piece of delicate absorbent paper, and then place the coverglass cell side facing down onto the drop of phalloidin solution on the Parafilm. Make sure that there are no bubbles trapped by the coverglass.
    10. Place the cover on the humidity chamber. Wrap the chamber in aluminum foil to protect it from ambient light, and let the sample incubate overnight at 4 °C. The sample can be kept in this condition for several days. Make sure that the humidity chamber remains moist if long storage is planned.
  8. Prior to imaging, gently place the coverglass cell side up onto a new 6-well plate containing 2 ml of PHEM buffer per well.
  9. Prepare oxygen-scavenged thiol-based imaging buffers using the following stock solutions: 1 M glucose, 1 M cysteamine, and 100x glucose oxidase/catalase enzyme mixture (4 mg of catalase and 10 mg of glucose oxidase in 100 µl of PHEM buffer, mixed well by vortexing)28. Right before imaging, mix 75 µl of 1 M glucose solution, 30 µl of 1 M cysteamine solution, and 3 µl of 100x stock enzyme mixture. Adjust the volume to 300 µl with PHEM buffer and use immediately after mixing to mount the sample.
  10. For iPALM, prepare the imaging sample using a pre-cleaned #1.5 coverglass (22-mm diameter).
    1. Alternatively, use a glass slide (3" x 1") with a double-sided adhesive spacer to aid in the assembly if astigmatism-based 3D-STORM12 is to be used instead.
    2. To assemble the imaging sample, use fine forceps to gently remove the specimen coverglass from the buffer well. Then, quickly and gently tap away excess buffer by touching the edge of the coverglass with folded absorbent paper.
    3. Place the coverglass cell side facing up on a piece of clean lens paper. Rinse the sample by placing 30 - 50 µl of imaging buffer onto the sample, and remove excess buffer by tilting and tapping with folded absorbent paper.
    4. Repeat the rinsing step a few times, and then place 30 - 50 µl of imaging buffer onto the sample. Blot dry the edge of the coverglass and place multiple very small dots of fast-curing epoxy onto the dried area.
    5. Slowly lower another pre-cleaned #1.5 coverglass (plain, round coverglass, 18-mm diameter) onto the center of the cell-containing 22-mm coverglass. Let the imaging buffer wet both coverglasses by capillary action. The small dots of fast-curing epoxy should adhere to both coverglasses.
    6. 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 (< 15 µm), but not so much as to crush the cells. Gauge the proper thickness by observing the Newton's rings pattern. Also, make sure to perform this step gently to minimize air bubbles. If needed, practice with empty coverglasses several times beforehand.
  11. Seal the sample with melted vaseline-lanolin-paraffin (VALAP, stock prepared from 100 g each of petroleum jelly, lanolin, and paraffin, melted together)29, rinse the sealed sample with ddH2O, and blow dry with compressed air. The sample is now ready for mounting onto the microscope for imaging.

2. Sample Placement and iPALM Alignment

  1. Move the spring-loaded top objective lens upward to allow for the removal of the sample holder. Place the sealed specimen prepared in step 1.10 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.
  2. Turn on the excitation lasers. Turn on the Electron Multiplying Charge-Coupled Device (EMCCD) camera in frame-transfer mode.
    1. Rotate in the proper emission filters. Activate a mechanical shutter to block the top beam path (Figure 1A) and open the bottom beam path. Bring the bottom objective lens into focus by translation in small increments using the piezo actuator.
    2. 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.
  3. 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 micro-fine set screws, until the fiducial images are overlapped as closely as possible, ideally within one pixel.
    1. Subsequently, perform fine adjustments so that the fiducial images in step 2.3 overlap within one tenth of an EMCCD pixel. Adjust the top 2-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 the bottom objective views via the computer display to guide the process.

3. Calibration of the iPALM Setup

NOTE: Since fluorescence emission is incoherent, for interference to be observed in iPALM, the path lengths through the top and bottom objectives must be close to each other, within a few microns. This can be achieved as follows:

  1. 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 nm.
  2. Taking advantage of the fact that, when out of optimal alignment, the fiducial intensity varies little with the oscillation, 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 > 10 can be achieved in optimal cases (Figure 1D).
  3. 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 8 nm z-steps over 800 nm.
    1. Monitor the fiducial intensity among cameras #1 - 3. 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 #1 relative to camera #2 is maximized, ideally at 120° (Figure 1B-D).
    2. 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, carry out the procedure in step 3.4 again, and record the calibration curve for use in subsequent z-coordinate extractions using the command "Acquire Calibration Scans vs Sample Piezo Position" in the main interface.

4. Data Acquisition

  1. 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. Increase the excitation power of the 642-nm laser to maximum. For Alexa Fluor 647, an initial period of fluorophore switching off may be needed (Figure 2A).
    1. Expose with a constant 642-nm excitation for 5 min, or longer if necessary, until single molecule blinking is observed (Figure 2B). The software allows an automatic increase of 405-nm photo activation during the course of the acquisition. Large numbers of acquisition frames are usually required for filamentous features to be clearly visible (> 50,000 frames). When ready, commence the acquisition of raw image sets using the command "Start iPALM Acquisition" in the main interface.
  2. During the acquisition, tune the photoactivation level by adjusting the intensity of the 405-nm laser to maintain the proper blinking density as needed (see examples in Figure 2B).
    NOTE: Once the acquisition is completed, the software will automatically convert the image files into the proper binary format. The data repository in the computation server is mounted as a networked drive, allowing data to be directly copied there for further processing.

5. Data Processing and Analysis

  1. Perform localization analysis using a custom-developed software to extract best-fit parameters for all single molecules as well as for the fiducials11,15,23. This yields not only the x,y-coordinate, but also the intensity that is used for calibration curve analysis.
    1. Import the raw calibration data acquired in step 3.5 using the command "Extract Peaks Multiple Labels" under the "File" menu to perform the single-molecule localization. Following the initial localization analysis, coordinates obtained from cameras #1 - 3 are present in red, green, and blue channels, respectively, and can be saved for further analysis using the command "Save Processed as IDL (.sav)" under the "File" menu.
  2. To bring data from cameras #1 - 3 into registration using the fluorescent fiducials embedded on the coverslips, select several bright fiducials to provide coverage of the central image (e.g., Figure 1B) using the command "Anchor Fiducial Points" under the "Image Transformations" menu. Use the triple sets of localization coordinates from cameras #1 - 3 obtained from the bright fiducials in step 5.1 to calculate the rotation and scaling matrix that will bring cameras #1 and #3 into register with camera #2. With a sufficiently large number of fiducials, higher-order polynomial warping can be performed for better fits.
  3. Once the transformation matrix is calculated, transform the raw data of cameras #1 - 3 together to obtain the summed raw data. Perform another round of localization analysis to yield a more precise x,y-coordinate and to determine the relative contribution of each camera channel to the summed raw data; use the command "Transform Raw, Save and Save Sum (.dat)" under the "Image Transformations" menu. This intensity ratio contains the z-coordinate information.
  4. Select a bright fiducial and perform z-calibration fitting using the function "Test Wind Point 3D" in the pop-up dialog by clicking "Z-coordinate Operations" under the "Special Functions" menu. This will fit 3-sinusoidal functions to the intensities of the 3 camera channels to determine the calibration curve. The calibration file is then saved for further use on the main datasets.
  5. To test the quality of the calibration curve, perform z-coordinate extraction, as described previously23. For well-behaved fiducials in a well-calibrated system, the z-coordinate should scale linearly, since the calibration datasets are taken with a linear sweep in z-position. Furthermore, the z-positions of all fiducials should scale with a similar slope.
  6. Once a satisfactory calibration is obtained, perform localization and transformation analysis for the raw image datasets acquired in step 4 following same procedure outlined in steps 5.1-5.3. When done, load the calibration file from step 5.4 and perform z-coordinate extraction using the functions "Pick WND File" and "Extract Z Coordinate" in the pop-up dialog by clicking "Z-coordinate Operations" under the "Special Functions" menu.
    NOTE: Since the acquisition time is > 15 min, mechanical drift is to be expected.
  7. To perform drift correction in x,y, select a bright fiducial from the localization coordinates. This fiducial should be present in all frames. Then, use the x,y-coordinate drift of the fiducial to align all other coordinates within the same frame back into registration (Figure 3A-B) using the function "Test/Write Guide Star" under the "Image Transformations" menu. Z-coordinate registration can be performed similarly by accessing the "Test Guide Star" and "Write Guide Star" functions in the pop-up dialog by clicking "Z-coordinate Operations" under the "Special Functions" menu.
  8. As the sample may exhibit slight tilt, perform tilt correction by providing the x,y,z-coordinates of 3 reference points defining the plane that should be set level using the function "Remove XYZ Tilt" in the pop-up dialog by clicking "Z-coordinate Operations" under the "Special Functions" menu.
  9. Subsequent to the drift and tilt corrections, save the localization coordinates. Reconstruct a super resolution image for further analysis (Figure 4A-D) by using the "Render" command on the main interface. Color can be used to indicate the z-coordinate. Alternatively, a side view of the selected area can also be rendered. The localization coordinates can be exported as text files for further quantification, or the reconstructed image can be saved as a .tif file.

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Results

Critical requirements for iPALM are the alignment, registration, and calibration of the optical systems. These are necessary to ensure proper interference within the 3-way beam splitter requisite for z-coordinate extraction. To enable continuous monitoring, constant point sources of fluorescence are necessary. This can be achieved using fluorescent Au or bi-metallic nanoparticles23 whose photoluminescence arise from localized surface plasmon resonance (LSPR). They act as a stab...

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Discussion

The optical system of iPALM is based on a 4-π dual-opposed objective design, as shown in Figure 1A. The setup is constructed using both custom-machined and commercial opto-mechanical parts, as described earlier23 and listed in Table 1. In addition to our setup, the Howard Hughes Medical Institute (HHMI) hosts a system that is accessible to the scientific community at the Advanced Imaging Center at the Janelia Research Campus. For the full mechanical drawi...

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Disclosures

The authors have nothing to disclose.

Acknowledgements

YW and PK gratefully acknowledge funding support from the Singapore National Research Foundation, awarded to PK (NRF-NRFF-2011-04 and NRF2012NRF-CRP001-084). We also thank the MBI open lab and microscopy core facilities for infrastructure support.

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Materials

List of materials used in this article
NameCompanyCatalog NumberComments
optical tableNewport, CARS4000iPALM, installed on 4 Newport Stabilizer vibration isolators
vibration isolator for optical tableNewport, CAS-2000
laser-642Newport, CA1185055output power=100 mw
laser-561Newport, CA1168931output power=200 mw
laser-488Newport, CA1137970output power=200 mw
laser-405Newport, CA1142279output power=100 mw
broadband dielectric mirrorsThorlabs, NJBB1-E02laser combiner
dichroic beamsplitterSemrock, NYLM01-427-25
acousto-optic tunable filterAA Opto-Electronic, FranceAOTFnC-VIS-TN
Linear polarizerNewport, CA05LP-VIS-B
baseplatelocal workshopcustomized
turning mirror (22.5°)Reynard Corpporation, CAcustomized22.5° mirror
motorized optic mountsNew Focus, CA8816
motorized XYZ translation stageThorlabs, NJMT3/M-Z6sample holder
T-Cube DC servo motor controllerThorlabs, NJTDC001
Piezo Phase ShifterPhysik Instrumente, GermanyS-303.CD
objective lensNikon, JapanMRD01691objective. Apo TIRF 60X/1.49 oil
translation stageNew Focus, CA9062-COM-M
Pico Motor ActuatorNew Focus, CA8301
rotary Solenoid/ShutterDACO Instruments, CT5423-458
3-way beam splitterRocky Mountain Instruments, COcustomizedbeamsplitter
Piezo Z/Tip/Tilt scannerPhysik Instrumente, GermanyS-316.10
motorized five-axis tilt alignerNew Focus, CA8081
Picmotor ethernet controllerNew Focus, CA8752
Piezo controllers/amplifier/digital operation modulePhysik Instrumente, GermanyE-509/E-503/E-517
band-pass filterSemrock, NYFF01-523/20filters
band-pass filterSemrock, NYFF01-588/21
band-pass filterSemrock, NYFF01-607/30
band-pass filterSemrock, NYFF01-676/37
notch filterSemrock, NYNF01-405/488/561/635
motorized filter wheel with controllterThorlabs, NJFW103H
EMCCDAndor, UKDU-897U-CSO-#BV3 sets
Desktop computers for controlling cameras and synchronizationDellPrecision T3500PC, 4 sets
coverslips with fiducialHestzig, VA600-100AuFsample preparation. fiducial marks with various density and spectra available
fibronectinMillipore, MTFC010
paraformaldehydeElectron Microscopy Sciences, PA15710fixation. 16%
glutaraldehydeElectron Microscopy Sciences, PA1622025%
triton X-100Sigma aldrich, MOT8787
HUVEC cellsLife Technologies, CAC-015-10C
Medium 200Life Technologies, CAM-200-500
Large Vessel Endothelial FactorsLife Technologies, CAA14608-01
Dulbecco's Phosphate Buffered Saline14190367
Pennicillin/Streptomycin15140122
Trypsin/EDTALife Technologies, CA25200056
PIPESSigma aldrich, MOP1851PHEM
HEPES1st base, MalaysiaBIO-1825
EGTASigma aldrich, MOE3889
MgCl2Millipore, MT5985
Alexa Fluor 647 PhalloidinInvitrogen, CAA22287staining
sodium borohydride (NaBH4)Sigma aldrich, MO480886quenching
glucose1st base, MalaysiaBIO-1101imaging buffer
glucose oxidaseSigma aldrich, MOG2133
catalaseSigma aldrich, MOC9322
cysteamineSigma aldrich, MO30070
EpoxyThorlabs, NJG14250
vaselineSigma aldrich, MO16415sample sealing
lanolinSigma aldrich, MOL7387
parafin waxSigma aldrich, MO327204
Immersion oilElectron Microscopy Sciences, PA16915-04imaging. Cargille Type HF

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iPALM MicroscopyActin CytoskeletonSingle Molecule LocalizationMultiphase InterferometryFluorescence MicroscopyEMCCD CamerasLaser ExcitationSample PreparationImage Reconstruction
Three-dimensional Super Resolution Microscopy of F-actin Filaments by Interferometric PhotoActivated Localization Microscopy (iPALM)
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