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.

1. Imaging Specimen Preparation
<ol>
	<li>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.</li>
	<li>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 (&#62; 15 - 30 min), sample drift is inevitable.</li>
	<li>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.</li>
	<li>In a sterile laminar flow hood, prepare fibronectin solution for coverglass coating by diluting 1 mg/ml fibronectin stock solution in sterile Dulbecco&#39;s phosphate-buffered saline (DPBS) to a final concentration 2 - 10 &#181;g/ml. Rinse each coverglass three times with DPBS and incubate with 2 ml of the fibronectin solution overnight at 4 &#176;C. Subsequently, aspirate out the fibronectin solution and rinse once with DPBS.</li>
	<li>Rinse the cells briefly with DPBS. Incubate with 1 - 2 ml of trypsin for a few min at 37 &#176;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.
	<ol>
		<li>Replate the cells onto the fibronectin-coated coverglass (for sparse density, plate &#60; 50,000 cells per coverglass) and maintain the culture in an incubator set at 95% humidity, 5% CO2, and 37 &#176;C.</li>
	</ol>
	</li>
	<li>For proper preservation of the fine structures of the f-actin cytoskeleton, use buffer solutions based on Good&#39;s buffer reagents27.
	<ol>
		<li>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-&#181;m filter, store it at 4 &#176;C, and dilute it 1:1 with ddH2O before use.</li>
	</ol>
	</li>
	<li>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:
	<ol>
		<li>Aspirate the media from each culture well containing the cell specimen. Gently but quickly dispense 2 ml of warm (37 &#176;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.</li>
		<li>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.</li>
		<li>Aspirate out the fixatives and gently replace them with PHEM buffer. Tilt and swirl gently, and then wash again with PHEM. Repeat twice.</li>
		<li>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.</li>
		<li>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.</li>
		<li>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.</li>
		<li>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 &#181;M. Prepare ~ 60 &#181;l per coverglass using phalloidin-Alexa Fluor 647 in PHEM buffer.</li>
		<li>Pipette 55 - 60 &#181;l 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.</li>
		<li>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.</li>
		<li>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 &#176;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.</li>
	</ol>
	</li>
	<li>Prior to imaging, gently place the coverglass cell side up onto a new 6-well plate containing 2 ml of PHEM buffer per well.</li>
	<li>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 &#181;l of PHEM buffer, mixed well by vortexing)28. Right before imaging, mix 75 &#181;l of 1 M glucose solution, 30 &#181;l of 1 M cysteamine solution, and 3 &#181;l of 100x stock enzyme mixture. Adjust the volume to 300 &#181;l with PHEM buffer and use immediately after mixing to mount the sample.</li>
	<li>For iPALM, prepare the imaging sample using a pre-cleaned #1.5 coverglass (22-mm diameter).
	<ol>
		<li>Alternatively, use a glass slide (3&#34; x 1&#34;) with a double-sided adhesive spacer to aid in the assembly if astigmatism-based 3D-STORM12 is to be used instead.</li>
		<li>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.</li>
		<li>Place the coverglass cell side facing up on a piece of clean lens paper. Rinse the sample by placing 30 - 50 &#181;l of imaging buffer onto the sample, and remove excess buffer by tilting and tapping with folded absorbent paper.</li>
		<li>Repeat the rinsing step a few times, and then place 30 - 50 &#181;l 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.</li>
		<li>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.</li>
		<li>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 (&#60; 15 &#181;m), but not so much as to crush the cells. Gauge the proper thickness by observing the Newton&#39;s rings pattern. Also, make sure to perform this step gently to minimize air bubbles. If needed, practice with empty coverglasses several times beforehand.</li>
	</ol>
	</li>
	<li>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.</li>
</ol>
2. Sample Placement and iPALM Alignment
<ol>
	<li>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.</li>
	<li>Turn on the excitation lasers. Turn on the Electron Multiplying Charge-Coupled Device (EMCCD) camera in frame-transfer mode.
	<ol>
		<li>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.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>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.
	<ol>
		<li>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.</li>
	</ol>
	</li>
</ol>
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:
<ol>
	<li>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.</li>
	<li>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 &#62; 10 can be achieved in optimal cases (Figure 1D).</li>
	<li>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.
	<ol>
		<li>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&#176; (Figure 1B-D).</li>
		<li>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 &#34;Acquire Calibration Scans vs Sample Piezo Position&#34; in the main interface.</li>
	</ol>
	</li>
</ol>
4. Data Acquisition
<ol>
	<li>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).
	<ol>
		<li>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 (&#62; 50,000 frames). When ready, commence the acquisition of raw image sets using the command &#34;Start iPALM Acquisition&#34; in the main interface.</li>
	</ol>
	</li>
	<li>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.</li>
</ol>
5. Data Processing and Analysis
<ol>
	<li>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.
	<ol>
		<li>Import the raw calibration data acquired in step 3.5 using the command &#34;Extract Peaks Multiple Labels&#34; under the &#34;File&#34; 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 &#34;Save Processed as IDL (.sav)&#34; under the &#34;File&#34; menu.</li>
	</ol>
	</li>
	<li>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 &#34;Anchor Fiducial Points&#34; under the &#34;Image Transformations&#34; 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.</li>
	<li>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 &#34;Transform Raw, Save and Save Sum (.dat)&#34; under the &#34;Image Transformations&#34; menu. This intensity ratio contains the z-coordinate information.</li>
	<li>Select a bright fiducial and perform z-calibration fitting using the function &#34;Test Wind Point 3D&#34; in the pop-up dialog by clicking &#34;Z-coordinate Operations&#34; under the &#34;Special Functions&#34; 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.</li>
	<li>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.</li>
	<li>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 &#34;Pick WND File&#34; and &#34;Extract Z Coordinate&#34; in the pop-up dialog by clicking &#34;Z-coordinate Operations&#34; under the &#34;Special Functions&#34; menu. 
	NOTE: Since the acquisition time is &#62; 15 min, mechanical drift is to be expected.</li>
	<li>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 &#34;Test/Write Guide Star&#34; under the &#34;Image Transformations&#34; menu. Z-coordinate registration can be performed similarly by accessing the &#34;Test Guide Star&#34; and &#34;Write Guide Star&#34; functions in the pop-up dialog by clicking &#34;Z-coordinate Operations&#34; under the &#34;Special Functions&#34; menu.</li>
	<li>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 &#34;Remove XYZ Tilt&#34; in the pop-up dialog by clicking &#34;Z-coordinate Operations&#34; under the &#34;Special Functions&#34; menu.</li>
	<li>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 &#34;Render&#34; 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.</li>
</ol>

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B., Soeller, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Visualization+of+Localization+Microscopy+Data.">Visualization of Localization Microscopy Data.</a> Microsc Microanal. 16, 64-72 (2010).</li><li>El Beheiry, M., Dahan, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=ViSP:+representing+single-particle+localizations+in+three+dimensions.">ViSP: representing single-particle localizations in three dimensions.</a> Nat Methods. 10, 689-690 (2013).</li><li>Schnell, U., Dijk, F., Sjollema, K. A., Giepmans, B. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Immunolabeling+artifacts+and+the+need+for+live-cell+imaging.">Immunolabeling artifacts and the need for live-cell imaging.</a> Nat Methods. 9, 152-158 (2012).</li><li>Shroff, H., Galbraith, C. G., Galbraith, J. A., Betzig, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Live-cell+photoactivated+localization+microscopy+of+nanoscale+adhesion+dynamics.">Live-cell photoactivated localization microscopy of nanoscale adhesion dynamics.</a> Nat Methods. 5, 417-423 (2008).</li><li>Yamashiro, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=New+single-molecule+speckle+microscopy+reveals+modification+of+the+retrograde+actin+flow+by+focal+adhesions+at+nanometer+scales.">New single-molecule speckle microscopy reveals modification of the retrograde actin flow by focal adhesions at nanometer scales.</a> Mol Biol Cell. 25, 1010-1024 (2014).</li><li>Shroff, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dual-color+super+resolution+imaging+of+genetically+expressed+probes+within+individual+adhesion+complexes.">Dual-color super resolution imaging of genetically expressed probes within individual adhesion complexes.</a> P Natl Acad Sci USA. 104, 20308-20313 (2007).</li><li>Chen, B. 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A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Super+resolution+fluorescence+imaging+of+mitochondrial+nucleoids+reveals+their+spatial+range,+limits,+and+membrane+interaction.">Super resolution fluorescence imaging of mitochondrial nucleoids reveals their spatial range, limits, and membrane interaction.</a> Mol Cell Biol. 31, 4994-5010 (2011).</li><li>Huang, F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Video-rate+nanoscopy+using+sCMOS+camera-specific+single-molecule+localization+algorithms.">Video-rate nanoscopy using sCMOS camera-specific single-molecule localization algorithms.</a> Nat Methods. 10, 653-658 (2013).</li><li>Daostorm, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=DAOSTORM:+an+algorithm+for+high-density+super-resolution+microscopy.">DAOSTORM: an algorithm for high-density super-resolution microscopy.</a> Nat methods. 8, 279 (2011).</li><li>Zhu, L., Zhang, W., Elnatan, D., Huang, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Faster+STORM+using+compressed+sensing.">Faster STORM using compressed sensing.</a> Nat Methods. 9, 721-723 (2012).</li><li>Van Engelenburg, S. B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Distribution+of+ESCRT+machinery+at+HIV+assembly+sites+reveals+virus+scaffolding+of+ESCRT+subunits.">Distribution of ESCRT machinery at HIV assembly sites reveals virus scaffolding of ESCRT subunits.</a> Science. 343, 653-656 (2014).</li><li>Vaughan, J. C., Jia, S., Zhuang, X. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ultrabright+photoactivatable+fluorophores+created+by+reductive+caging.">Ultrabright photoactivatable fluorophores created by reductive caging.</a> Nat Methods. 9, 1181-1184 (2012).</li></ol>

The authors have nothing to disclose.

The optical system of iPALM is based on a 4-&#960; 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 drawings, control schematics, and software, readers are encouraged to inquire with Harald Hess at HHMI for further information. A primary advantage for using iPALM as well as other single-molecule localization microscopy methods for visualizing f-actin is the relative ease of sample preparation compared to EM techniques, which generally require harsh sample processing, laborious preparation, and highly skilled practitioners3,23. Additionally, fluorescence is naturally amenable to multichannel imaging, which remains difficult for EM. Nevertheless, as described further below, there are several current limitations for the approach, both in terms of specimen preparation and imaging process. First, as is the case for EM sample preparation, care must be taken to ensure good ultrastructural preservation of the specimens35, since a protocol adequate for a diffraction-limited level of resolution can cause severe perturbations at the nanoscale level. For actin imaging, proper fixation of the cells is an important factor affecting specimen quality. Glutaraldehyde is a preferred fixative since it preserves the cytoskeleton and membrane very well, though in cases of co-staining with antibody, epitope sites could be affected. In such cases, paraformaldehyde may offer an acceptable compromise, as it tends to better preserve antibody epitope sites. Importantly for single-molecule localization microscopy, these fixatives tend to generate background autofluorescence; thus, post-fixation quenching by borohydride is necessary, particularly in the case of glutaraldehyde.
Additionally, the proper selection of fluorophores and labeling strategies are among the most important factors for successful experiments. Due to the nanoscale dimension of f-actin, high densities of labels are required to capture the underlying filamentous networks, as stipulated by the Nyquist sampling theorem36. For actin, phalloidin allows very high density labeling, with a wide range of organic fluorophores available from many manufacturers. However, since phalloidin is toxic, cell fixation and permeabilization is required. Alternative strategies for live-cell compatible actin labeling include using fusion of fluorescent proteins (FP), either with monomeric actin or with small actin-binding polypeptides. However, we found that these tend to yield a lower labeling density compared to phalloidin (Figure 6). We note that purified actin can also be directly labeled with organic fluorophores and introduced into living cells by electroporation37, but this approach is expected to be costly and labor-intensive. In terms of the fluorophores, Alexa Fluor 647 has generally been found to offer consistently good performance for localization microscopy. This can be described in terms of the contrast ratio-the brightness quotient between the single molecule peak and the background fluorescence38. A higher labeling density requires a higher contrast ratio, since the background could cumulatively degrade the localization precision. In our experience, several other fluorophores, such as ATTO488, ATTO520, and Alexa Fluor 568 (Figure 6B), can be used to visualize the f-actin, although with less consistent results compared to Alexa Fluor 647, which offers robust photoswitching performance across a wide range of imaging buffer conditions.
In terms of the imaging process, limitations in biological imaging methods have often been conceptualized as a trade-off in spatial resolution, temporal resolution, and imaging range (both field-of-view/depth-of-field and duration)39. This equally applies to the advantages and limitations of iPALM. Compared to other 3-dimensional super resolution microscopy techniques, iPALM has remained the highest-resolving optical approach to date, particularly along the z-axis, with the z-resolution nearly 2 times better than the x,y-resolution11. However, the reliance of iPALM on interferometry for a high z-resolution also imposes a restriction on the imaging depth, since the calibration curve is periodic. In other words, z-coordinates repeat every 250 nm or so (for ~ 700 nm emission wavelengths of Alexa Fluor 647). In practice, this can be addressed by using TIRF illumination, which limits the excitation zone to within the evanescent field depth of &#60; 200 nm from the coverglass. Thus, fluorophores deeper into the specimen are not excited and remain invisible. However, this also limits the biological structures that can be imaged to those in close proximity to the coverglass, such as focal adhesions, cortical cytoskeleton, and ventral plasma membrane, while more interior structures are out of reach. For fixed specimen, one remedy is to perform cryosectioning40. However, such an approach is highly laborious and requires specialized expertise that is not commonly accessible.
Alternatively, iPALM can be adapted for an extended z-range by combining interferometry with the astigmatic-defocusing scheme12 used in 3D-STORM. This approach provides dual z-coordinate readouts, the first by interferometry, which is high-precision but periodic, and the second by astigmatic defocusing, which is less precise but non-periodic. The latter can then be used to &#34;unwrap&#34; or break the degeneracy of the interferometric z-coordinates, thus enabling the tripling of the iPALM imaging depth to ~750 nm, effectively approaching the intrinsic focal depth of high NA objective lenses. With phase unwrapping, iPALM has been used to image structures deep within the samples, such as mitochondria, albeit with slightly reduced precision in all 3 dimensions due to the additional defocusing40.
Another intrinsic limitation of iPALM is the imaging speed. Since large numbers of frames are required to obtain a high density of localization coordinates, the camera speed is usually the limit on the acquisition rate. With current EMCCD cameras running at 10 - 20 MHz readout rates, tens of minutes are required for a sufficient number of frames. Such long time scales require that the specimen be fixed. Here, recent advances in camera technology, such as the much faster scientific Complementary Metal Oxide Semiconductor (sCMOS) camera, may enable a rapid increase in imaging speed41, although this has not been implemented for iPALM yet. For some single-molecule localization microscopy approaches, a dense single-molecule field can be analyzed using a modified algorithm42 or compressed sensing43. The adaptation of such strategies may also speed up iPALM by allowing denser single-molecule images and, hence, fewer frames.
With limitations in speed and imaging range and strengths in spatial resolution, a major application for iPALM is as an ultrastructural method that leverages the benefits of fluorescent labeling to unveil nanoscale organization of specific proteins14,15,44. Further improvement, both in terms of optics and fluorophore technologies, should enable further enhancement in iPALM spatial resolution. For example, iPALM image processing currently employs a relatively simple first-order approximation approach, with each single molecule modeled by a 2D-Gaussian function and assumed to be an isotopically averaged dipole. This could be augmented with recent methods, which employ a more accurate model of the point-spread function and dipole orientation, or explicit treatment of optical aberrations across the field of view to significantly improve spatial resolution. Likewise, photocaging has been reported, which enhances fluorophore brightness by orders of magnitude45. Indeed, since key elements of their ultrastructural organization have been well-characterized, the f-actin cytoskeleton could be a highly valuable model system for testing further methodological developments in iPALM. The ability to analyze protein organization by light microscopy with true EM-level resolving power is expected to greatly enhance our understanding of myriad aspects of cellular structure and function.

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 &#62; 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.

<table><tbody><tr><td>optical table</td><td>Newport, CA</td><td>RS4000</td><td>iPALM, installed on 4 Newport Stabilizer vibration isolators</td></tr><tr><td>vibration isolator for optical table</td><td>Newport, CA</td><td>S-2000</td><td></td></tr><tr><td>laser-642</td><td>Newport, CA</td><td>1185055</td><td>output power=100 mw</td></tr><tr><td>laser-561</td><td>Newport, CA</td><td>1168931</td><td>output power=200 mw</td></tr><tr><td>laser-488</td><td>Newport, CA</td><td>1137970</td><td>output power=200 mw</td></tr><tr><td>laser-405</td><td>Newport, CA</td><td>1142279</td><td>output power=100 mw</td></tr><tr><td>broadband dielectric mirrors</td><td>Thorlabs, NJ</td><td>BB1-E02</td><td>laser combiner</td></tr><tr><td>dichroic beamsplitter</td><td>Semrock, NY</td><td>LM01-427-25</td><td></td></tr><tr><td>acousto-optic tunable filter</td><td>AA Opto-Electronic, France</td><td>AOTFnC-VIS-TN</td><td></td></tr><tr><td>Linear polarizer</td><td>Newport, CA</td><td>05LP-VIS-B</td><td></td></tr><tr><td>baseplate</td><td>local workshop</td><td>customized</td><td></td></tr><tr><td>turning mirror (22.5&amp;deg;)</td><td>Reynard Corpporation, CA</td><td>customized</td><td>22.5&amp;deg; mirror</td></tr><tr><td>motorized optic mounts</td><td>New Focus, CA</td><td>8816</td><td></td></tr><tr><td>motorized XYZ translation stage</td><td>Thorlabs, NJ</td><td>MT3/M-Z6</td><td>sample holder</td></tr><tr><td>T-Cube DC servo motor controller</td><td>Thorlabs, NJ</td><td>TDC001</td><td></td></tr><tr><td>Piezo Phase Shifter</td><td>Physik Instrumente, Germany</td><td>S-303.CD</td><td></td></tr><tr><td>objective lens</td><td>Nikon, Japan</td><td>MRD01691</td><td>objective. Apo TIRF 60X/1.49 oil</td></tr><tr><td>translation stage</td><td>New Focus, CA</td><td>9062-COM-M</td><td></td></tr><tr><td>Pico Motor Actuator</td><td>New Focus, CA</td><td>8301</td><td></td></tr><tr><td>rotary Solenoid/Shutter</td><td>DACO Instruments, CT</td><td>5423-458</td><td></td></tr><tr><td>3-way beam splitter</td><td>Rocky Mountain Instruments, CO</td><td>customized</td><td>beamsplitter</td></tr><tr><td>Piezo Z/Tip/Tilt scanner</td><td>Physik Instrumente, Germany</td><td>S-316.10</td><td></td></tr><tr><td>motorized five-axis tilt aligner</td><td>New Focus, CA</td><td>8081</td><td></td></tr><tr><td>Picmotor ethernet controller</td><td>New Focus, CA</td><td>8752</td><td></td></tr><tr><td>Piezo controllers/amplifier/digital operation module</td><td>Physik Instrumente, Germany</td><td>E-509/E-503/E-517</td><td></td></tr><tr><td>band-pass filter</td><td>Semrock, NY</td><td>FF01-523/20</td><td>filters</td></tr><tr><td>band-pass filter</td><td>Semrock, NY</td><td>FF01-588/21</td><td></td></tr><tr><td>band-pass filter</td><td>Semrock, NY</td><td>FF01-607/30</td><td></td></tr><tr><td>band-pass filter</td><td>Semrock, NY</td><td>FF01-676/37</td><td></td></tr><tr><td>notch filter</td><td>Semrock, NY</td><td>NF01-405/488/561/635</td><td></td></tr><tr><td>motorized filter wheel with controllter</td><td>Thorlabs, NJ</td><td>FW103H</td><td></td></tr><tr><td>EMCCD</td><td>Andor, UK</td><td>DU-897U-CSO-#BV</td><td>3 sets</td></tr><tr><td>Desktop computers for controlling cameras and synchronization</td><td>Dell</td><td>Precision T3500</td><td>PC, 4 sets</td></tr><tr><td>coverslips with fiducial</td><td>Hestzig, VA</td><td>600-100AuF</td><td>sample preparation. fiducial marks with various density and spectra available</td></tr><tr><td>fibronectin</td><td>Millipore, MT</td><td>FC010</td><td></td></tr><tr><td>paraformaldehyde</td><td>Electron Microscopy Sciences, PA</td><td>15710</td><td>fixation. 16%</td></tr><tr><td>glutaraldehyde</td><td>Electron Microscopy Sciences, PA</td><td>16220</td><td>25%</td></tr><tr><td>triton X-100</td><td>Sigma aldrich, MO</td><td>T8787</td><td></td></tr><tr><td>HUVEC cells</td><td>Life Technologies, CA</td><td>C-015-10C</td><td></td></tr><tr><td>Medium 200</td><td>Life Technologies, CA</td><td>M-200-500</td><td></td></tr><tr><td>Large Vessel Endothelial Factors</td><td>Life Technologies, CA</td><td>A14608-01</td><td></td></tr><tr><td>Dulbecco&#039;s Phosphate Buffered Saline</td><td></td><td>14190367</td><td></td></tr><tr><td>Pennicillin/Streptomycin</td><td></td><td>15140122</td><td></td></tr><tr><td>Trypsin/EDTA</td><td>Life Technologies, CA</td><td>25200056</td><td></td></tr><tr><td>PIPES</td><td>Sigma aldrich, MO</td><td>P1851</td><td>PHEM</td></tr><tr><td>HEPES</td><td>1st base, Malaysia</td><td>BIO-1825</td><td></td></tr><tr><td>EGTA</td><td>Sigma aldrich, MO</td><td>E3889</td><td></td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Millipore, MT</td><td>5985</td><td></td></tr><tr><td>Alexa Fluor 647 Phalloidin</td><td>Invitrogen, CA</td><td>A22287</td><td>staining</td></tr><tr><td>sodium borohydride (NaBH&lt;sub&gt;4&lt;/sub&gt;)</td><td>Sigma aldrich, MO</td><td>480886</td><td>quenching</td></tr><tr><td>glucose</td><td>1st base, Malaysia</td><td>BIO-1101</td><td>imaging buffer</td></tr><tr><td>glucose oxidase</td><td>Sigma aldrich, MO</td><td>G2133</td><td></td></tr><tr><td>catalase</td><td>Sigma aldrich, MO</td><td>C9322</td><td></td></tr><tr><td>cysteamine</td><td>Sigma aldrich, MO</td><td>30070</td><td></td></tr><tr><td>Epoxy</td><td>Thorlabs, NJ</td><td>G14250</td><td></td></tr><tr><td>vaseline</td><td>Sigma aldrich, MO</td><td>16415</td><td>sample sealing</td></tr><tr><td>lanolin</td><td>Sigma aldrich, MO</td><td>L7387</td><td></td></tr><tr><td>parafin wax</td><td>Sigma aldrich, MO</td><td>327204</td><td></td></tr><tr><td>Immersion oil</td><td>Electron Microscopy Sciences, PA</td><td>16915-04</td><td>imaging. Cargille Type HF</td></tr></tbody></table>

three-dimensional super resolution microscopy of f-actin filaments by interferometric photoactivated localization microscopy (ipalm)

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 &#62; 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.

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

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.

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

Fluorescence microscopy enables direct visualization of specific biomolecules within cells. However, for conventional fluorescence microscopy, the spatial ...

three-dimensional-super-resolution-microscopy-f-actin-filaments

Nanyang Technological University

Research

JoVE Journal

Biology

10.7K Views.  South University of Science and Technology of China, Shenzhen.  We presenteren een protocol voor de toepassing van interferometrische Photoactivated Localization Microscopy (iPALM), een 3-dimensionale single-molecule lokalisatie super resolutie microscopie methode, aan de beeldvorming van het actine cytoskelet in hechtende zoogdiercellen. Deze aanpak maakt het mogelijk op basis van licht visualisatie van nanoschaal structurele kenmerken die anders onopgelost zouden blijven door middel van conventionele diffractie beperkt optische mic...

Video: Three-dimensional Super Resolution Microscopy of F-actin Filaments by Interferometric PhotoActivated Localization Microscopy iPALM

Live Cell Imaging of F-actin Dynamics via Fluorescent Speckle Microscopy (FSM)

In this protocol we describe the use of Fluorescent Speckle Microscopy (FSM) to capture high-resolution images of actin dynamics in PtK1 cells. A unique advantage of FSM is its ability to capture the movement and turnover kinetics (assembly/disassembly) of the F-actin network within living cells. This technique is particularly useful in deriving quantitative measurements of F-actin dynamics when paired with computer vision software (qFSM).  We describe  the selection, microinjection and visualization of fluorescent actin probes in living cells. Importantly, similar procedures are applicable to visualizing other macomolecular assemblies. FSM has been demonstrated for microtubules, intermediate filaments, and adhesion complexes.  

Live Cell Imaging of F-actin Dynamics via Fluorescent Speckle Microscopy FSM

Selection, microinjection, and imaging of fluorescently-labeled F-actin via fluorescent speckle microscopy (FSM).

In this protocol we describe the use of Fluorescent Speckle Microscopy (FSM) to capture high-resolution images of actin dynamics in PtK1 cells. A unique ...

Cortical Actin Flow in T Cells Quantified by Spatio-temporal Image Correlation Spectroscopy of Structured Illumination Microscopy Data

Filamentous-actin plays a crucial role in a majority of cell processes including motility and, in immune cells, the formation of a key cell-cell interaction known as the immunological synapse. F-actin is also speculated to play a role in regulating molecular distributions at the membrane of cells including sub-membranous vesicle dynamics and protein clustering. While standard light microscope techniques allow generalized and diffraction-limited observations to be made, many cellular and molecular events including clustering and molecular flow occur in populations at length-scales far below the resolving power of standard light microscopy. By combining total internal reflection fluorescence with the super resolution imaging method structured illumination microscopy, the two-dimensional molecular flow of F-actin at the immune synapse of T cells was recorded. Spatio-temporal image correlation spectroscopy (STICS) was then applied, which generates quantifiable results in the form of velocity histograms and vector maps representing flow directionality and magnitude. This protocol describes the combination of super-resolution imaging and STICS techniques to generate flow vectors at sub-diffraction levels of detail. This technique was used to confirm an actin flow that is symmetrically retrograde and centripetal throughout the periphery of T cells upon synapse formation.

To investigate flow velocities and directionality of filamentous-actin at the T cell immunological synapse, live-cell super-resolution imaging is combined with total internal reflection fluorescence and quantified with spatio-temporal image correlation spectroscopy.

Filamentous-actin plays a crucial role in a majority of cell processes including motility and, in immune cells, the formation of a key cell-cell ...

Immunology and Infection

Simultaneous Interference Reflection and Total Internal Reflection Fluorescence Microscopy for Imaging Dynamic Microtubules and Associated Proteins

Several techniques have been employed for the direct visualization of cytoskeletal filaments and their associated proteins. Total-internal-reflection-fluorescence (TIRF) microscopy has a high signal-to-background ratio, but it suffers from photobleaching and photodamage of the fluorescent proteins. Label-free techniques such as interference reflection microscopy (IRM) and interferometric scattering microscopy (iSCAT) circumvent the problem of photobleaching but cannot readily visualize single molecules. This paper presents a protocol for combining IRM with a commercial TIRF microscope for the simultaneous imaging of microtubule-associated proteins (MAPs) and dynamic microtubules in vitro. This protocol allows for high-speed observation of MAPs interacting with dynamic microtubules. This improves on existing two-color TIRF setups by eliminating both the need for microtubule labeling and the need for several additional optical components, such as a second excitation laser. Both channels are imaged on the same camera chip to avoid image registration and frame synchronization problems. This setup is demonstrated by visualizing single kinesin molecules walking on dynamic microtubules.

We present a protocol for implementing interference-reflection microscopy and total-internal-reflection-fluorescence microscopy for the simultaneous imaging of dynamic microtubules and fluorescently labeled microtubule-associated proteins.

Several techniques have been employed for the direct visualization of cytoskeletal filaments and their associated proteins. ...

Biochemistry

Using Microfluidics and Fluorescence Microscopy to Study the Assembly Dynamics of Single Actin Filaments and Bundles

In order to decipher the complex molecular mechanisms that regulate the assembly and disassembly of actin filaments, it is a great asset to monitor individual reactions live in well-controlled conditions. To do so, live single-filament experiments have emerged over the past 20 years, mostly using total internal reflection fluorescence (TIRF) microscopy, and have provided a trove of key results. In 2011, in order to further expand the possibilities of these experiments and to avoid recurring problematic artifacts, we introduced simple microfluidics in these assays. This study details our basic protocol, where individual actin filaments are anchored by one end to the passivated coverslip surface, align with the flow, and can be successively exposed to different protein solutions. We also present the protocols for specific applications and explain how controlled mechanical forces can be applied, thanks to the viscous drag of the flowing solution. We highlight the technical caveats of these experiments and briefly present possible developments based on this technique. These protocols and explanations, along with today&#39;s availability of easy-to-use microfluidics equipment, should allow non-specialists to implement this assay in their labs.

We present protocols for simple actin filament microfluidic assays, in combination with fluorescence microscopy, that allow one to accurately monitor individual actin filaments in real-time while sequentially exposing them to different protein solutions.

In order to decipher the complex molecular mechanisms that regulate the assembly and disassembly of actin filaments, it is a great asset to monitor ...

Three-dimensional Imaging of Bacterial Cells for Accurate Cellular Representations and Precise Protein Localization

The shape of a bacterium is important for its physiology. Many aspects of cell physiology such as cell motility, predation, and biofilm production can be affected by cell shape. Bacterial cells are three-dimensional (3D) objects, although they are rarely treated as such. Most microscopy techniques result in two-dimensional (2D) images leading to the loss of data pertaining to the actual 3D cell shape and localization of proteins. Certain shape parameters, such as Gaussian curvature (the product of the two principal curvatures), can only be measured in 3D because 2D images do not measure both principal curvatures. Additionally, not all cells lie flat when mounting and 2D imaging of curved cells may not accurately represent the shapes of these cells. Accurately measuring protein localization in 3D can help determine the spatial regulation and function of proteins. A forward convolution technique has been developed that uses the blurring function of the microscope to reconstruct 3D cell shapes and to accurately localize proteins. Here, a protocol for preparing and mounting samples for live cell imaging of bacteria in 3D both to reconstruct an accurate cell shape and to localize proteins is described. The method is based on simple sample preparation, fluorescent image acquisition, and MATLAB-based image processing. Many high-quality fluorescent microscopes can be simply modified to take these measurements. These cell reconstructions are computationally intensive and access to high-throughput computational resources is recommended, although not necessary. This method has been successfully applied to multiple bacterial species and mutants, fluorescent imaging modalities, and microscope manufacturers.

This protocol explains how to prepare and mount bacterial samples for live three-dimensional imaging and how to reconstruct the three-dimensional shape of E. coli from those images.

The shape of a bacterium is important for its physiology. Many aspects of cell physiology such as cell motility, predation, and biofilm production can be ...

Super-resolution Imaging of the Bacterial Division Machinery

Bacterial cell division requires the coordinated assembly of more than ten essential proteins at midcell1,2. Central to this process is the formation of a ring-like suprastructure (Z-ring) by the FtsZ protein at the division plan3,4. The Z-ring consists of multiple single-stranded FtsZ protofilaments, and understanding the arrangement of the protofilaments inside the Z-ring will provide insight into the mechanism of Z-ring assembly and its function as a force generator5,6. This information has remained elusive due to current limitations in conventional fluorescence microscopy and electron microscopy. Conventional fluorescence microscopy is unable to provide a high-resolution image of the Z-ring due to the diffraction limit of light (~200 nm). Electron cryotomographic imaging has detected scattered FtsZ protofilaments in small C. crescentus cells7, but is difficult to apply to larger cells such as E. coli or B. subtilis. Here we describe the application of a super-resolution fluorescence microscopy method, Photoactivated Localization Microscopy (PALM), to quantitatively characterize the structural organization of the E. coli Z-ring8.
	PALM imaging offers both high spatial resolution (~35 nm) and specific labeling to enable unambiguous identification of target proteins. We labeled FtsZ with the photoactivatable fluorescent protein mEos2, which switches from green fluorescence (excitation = 488 nm) to red fluorescence (excitation = 561 nm) upon activation at 405 nm9. During a PALM experiment, single FtsZ-mEos2 molecules are stochastically activated and the corresponding centroid positions of the single molecules are determined with &lt;20 nm precision. A super-resolution image of the Z-ring is then reconstructed by superimposing the centroid positions of all detected FtsZ-mEos2 molecules.
	Using this method, we found that the Z-ring has a fixed width of ~100 nm and is composed of a loose bundle of FtsZ protofilaments that overlap with each other in three dimensions. These data provide a springboard for further investigations of the cell cycle dependent changes of the Z-ring10 and can be applied to other proteins of interest.

We describe a super-resolution imaging method to probe the structural organization of the bacterial FtsZ-ring, an essential apparatus for cell division. This method is based on quantitative analyses of photoactivated localization microscopy (PALM) images and can be applied to other bacterial cytoskeletal proteins.

Bacterial cell division requires the coordinated  assembly of more than ten essential proteins at midcell1,2. Central  to this process is the formation of ...

Simultaneous Multicolor Imaging of Biological Structures with Fluorescence Photoactivation Localization Microscopy

Localization-based super resolution microscopy can be applied to obtain a spatial map (image) of the distribution of individual fluorescently labeled single molecules within a sample with a spatial resolution of tens of nanometers. Using either photoactivatable (PAFP) or&#160;photoswitchable (PSFP) fluorescent proteins fused to proteins of interest, or organic dyes conjugated to antibodies or other molecules of interest, fluorescence photoactivation localization microscopy (FPALM) can simultaneously image multiple species of molecules within single cells. By using the following approach, populations of large numbers (thousands to hundreds of thousands) of individual molecules are imaged in single cells and localized with a precision of ~10-30 nm. Data obtained can be applied to understanding the nanoscale spatial distributions of multiple protein types within a cell. One primary advantage of this technique is the dramatic increase in spatial resolution: while diffraction limits resolution to ~200-250 nm in conventional light microscopy, FPALM can image length scales more than an order of magnitude smaller. As many biological hypotheses concern the spatial relationships among different biomolecules, the improved resolution of FPALM can provide insight into questions of cellular organization which have previously been inaccessible to conventional fluorescence microscopy. In addition to detailing the methods for sample preparation and data acquisition, we here describe the optical setup for FPALM. One additional consideration for researchers wishing to do super-resolution microscopy is cost: in-house setups are significantly cheaper than most commercially available imaging machines. Limitations of this technique include the need for optimizing the labeling of molecules of interest within cell samples, and the need for post-processing software to visualize results. We here describe the use of PAFP and PSFP expression to image two protein species in fixed cells. Extension of the technique to living cells is also described.

We demonstrate the use of fluorescence photo activation localization microscopy (FPALM) to simultaneously image multiple types of fluorescently labeled molecules&#160;within cells. The techniques described yield the localization of thousands to hundreds of thousands of individual fluorescent labeled proteins, with a precision of tens of nanometers&#160;within single cells.

Localization-based super resolution microscopy can be applied to obtain a spatial map (image) of the distribution of individual fluorescently labeled ...

Super-resolution Imaging of the Cytokinetic Z Ring in Live Bacteria Using Fast 3D-Structured Illumination Microscopy (f3D-SIM)

Imaging of biological samples using fluorescence microscopy has advanced substantially with new technologies to overcome the resolution barrier of the diffraction of light allowing super-resolution of live samples. There are currently three main types of super-resolution techniques &#8211; stimulated emission depletion (STED), single-molecule localization microscopy (including techniques such as PALM, STORM, and GDSIM), and structured illumination microscopy (SIM). While STED and single-molecule localization techniques show the largest increases in resolution, they have been slower to offer increased speeds of image acquisition. Three-dimensional SIM (3D-SIM) is a wide-field fluorescence microscopy technique that offers a number of advantages over both single-molecule localization and STED. Resolution is improved, with typical lateral and axial resolutions of 110 and 280 nm, respectively and depth of sampling of up to 30 &#181;m from the coverslip, allowing for imaging of whole cells. Recent advancements (fast 3D-SIM) in the technology increasing the capture rate of raw images allows for fast capture of biological processes occurring in seconds, while significantly reducing photo-toxicity and photobleaching. Here we describe the use of one such method to image bacterial cells harboring the fluorescently-labelled cytokinetic FtsZ protein to show how cells are analyzed and the type of unique information that this technique can provide.

Super-resolution Imaging of the Cytokinetic Z Ring in Live Bacteria Using Fast 3D-Structured Illumination Microscopy f3D-SIM

Spatiotemporal information about dynamic proteins inside live cells is crucial for understanding biology. A type of super-resolution microscopy called fast 3D-structured illumination microscopy (f3D-SIM) reveals unique information about the cytokinetic Z ring in bacteria: both its bead-like appearance and the rapid dynamics of FtsZ within the ring.

Imaging of biological samples using fluorescence microscopy has advanced substantially with new technologies to overcome the resolution barrier of the ...

Fluorescence Imaging with One-nanometer Accuracy (FIONA)

Fluorescence imaging with one-nanometer accuracy (FIONA) is a simple but useful technique for localizing single fluorophores with nanometer precision in the x-y plane. Here a summary of the FIONA technique is reported and examples of research that have been performed using FIONA are briefly described. First, how to set up the required equipment for FIONA experiments, i.e., a total internal reflection fluorescence microscopy (TIRFM), with details on aligning the optics, is described. Then how to carry out a simple FIONA experiment on localizing immobilized Cy3-DNA single molecules using appropriate protocols, followed by the use of FIONA to measure the 36 nm step size of a single truncated myosin Va motor labeled with a quantum dot, is illustrated. Lastly, recent effort to extend the application of FIONA to thick samples is reported. It is shown that, using a water immersion objective and quantum dots soaked deep in sol-gels and rabbit eye corneas (&#62;200 &#181;m), localization precision of 2-3 nm can be achieved.

Fluorescence Imaging with One-nanometer Accuracy FIONA

Single fluorophores can be localized with nanometer precision using FIONA. Here a summary of the FIONA technique is reported, and how to carry out FIONA experiments is described.

Fluorescence imaging with one-nanometer accuracy (FIONA) is a simple but useful technique for localizing single fluorophores with nanometer precision in ...

Imaging Intermediate Filaments and Microtubules with 2-dimensional Direct Stochastic Optical Reconstruction Microscopy

The cytoskeleton, composed of actin microfilaments, microtubules, and intermediate filaments (IF), plays a key role in the control of cell shape, polarity, and motility. The organization of the actin and microtubule networks has been extensively studied but that of IFs is not yet fully characterized. IFs have an average diameter of 10 nm and form a network extending throughout the cell cytoplasm. They are physically associated with actin and microtubules through molecular motors and cytoskeletal linkers. This tight association is at the heart of the regulatory mechanisms that ensure the coordinated regulation of the three cytoskeletal networks required for most cell functions. It is therefore crucial to visualize IFs alone and also together with each of the other cytoskeletal networks. However, IF networks are extremely dense in most cell types, especially in glial cells, which makes its resolution very difficult to achieve with standard fluorescence microscopy (lateral resolution of ~250 nm). Direct STochastic Optical Reconstruction Microscopy (dSTORM) is a technique allowing a gain in lateral resolution of one order of magnitude. Here, we show that lateral dSTORM resolution is sufficient to resolve the dense organization of the IF networks and, in particular, of IF bundles surrounding microtubules. Such tight association is likely to participate in the coordinated regulation of these two networks and may, explain how vimentin IFs template and stabilize microtubule organization as well as could influence microtubule dependent vesicular trafficking. More generally, we show how the observation of two cytoskeletal components with dual-color dSTORM technique brings new insight into their mutual interaction.

The overall goal of this methodology is to give the optimal experimental conditions from sample preparation to image acquisition and reconstruction in order to perform 2D dual color dSTORM images of microtubules and intermediate filaments in fixed cells

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

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