We thank Mickael Lelek, Orestis Faklaris and Nicolas Bourg for fruitful discussion, Andrey Aristov and Elena Rensen for help with the super-resolution technique and Shailaja Seetharaman for careful reading of the manuscript. We gratefully acknowledge the UtechS Photonic BioImaging (Imagopole) Citech of Institut Pasteur (Paris, France) as well as the France-BioImaging infrastructure network supported by the French National Research Agency (ANR-10-INSB-04; Investments for the Future), and the R&#233;gion Ile-de-France (program Domaine d'Int&#233;r&#234;t Majeur-Malinf) for the use of the Elyra microscope. This work was supported by the the Ligue Contre le Cancer and the French National Research Agency (ANR-16-CE13-0019).

1. Coverslips preparation (Day 1, 30 min)
<ol>
	<li>Place the 18-mm n&#186;1.5H (170 &#181;m +/- 5&#181;m) glass coverslips on a plastic rack.</li>
	<li>Put the rack in a 100-mL beaker filled with acetone and soak for 5 min.</li>
	<li>Transfer the rack to a 100-mL beaker filled with absolute ethanol and soak for 5 min.</li>
	<li>Transfer the rack to a new 100 mL beaker filled with absolute ethanol and place the beaker in an ultrasonic cleaner device (see Table of materials) at room temperature. Press &#34;on&#34; and wait for 10 min.</li>
	<li>Put the rack under a laminar flow hood and let the coverslips dry or dry the coverslips with filtered air flow.</li>
	<li>Put the rack with coverslips in a plasma cleaner device. Switch on the vacuum pump, close the door, wait for 3 min to make the vacuum and switch on the plasma for 1 min. Use the coverslips shortly after cleaning. Do not store them.</li>
</ol>
2. Cell plating (Day 1, 20 min)
<ol>
	<li>Grow the glial cell line U373 in Minimum Essential Medium (MEM) with 10% Fetal Bovine Serum, 1% Penicillin-Streptomycin, and 1% non-essential amino acid in 10 cm diameter plastic petri dish.</li>
	<li>Place clean glass coverslips in 12-well plates under the laminar flow hood and add 1 mL of preheated cell medium per well.</li>
	<li>Take a dish containing cells from the cell incubator (37 &#176;C, 5% CO2).</li>
	<li>Remove the medium and add 10 mL PBS.</li>
	<li>Remove the PBS and add 1 mL 0.05% Trypsin-EDTA.</li>
	<li>Place the dish back in the incubator for 2-3 min.</li>
	<li>Add 10 mL of warm medium preheated at 37 &#176;C on the dish and resuspend the cells.</li>
	<li>Seed about 100 000 cells per well (~150 &#181;L of cells) in the 12-well plates that contain the coverslips.</li>
	<li>Wait for at least 6 h (or overnight) to allow cell spreading.</li>
</ol>
3. Cell fixation (Day 2, 2h)
<ol>
	<li>Prepare the cytoskeleton buffer with 80 mM PIPES, 7 mM MgCl2, 1mM EGTA, 150 mM NaCl, and adjust the pH to 6.9.</li>
	<li>Wash the cells once with PBS.</li>
	<li>Remove the PBS and gently add 1 mL per well of extraction/fixation solution (0.25% glutaraldehyde, 0.3% Triton X-100 in the cytoskeleton buffer) preheated at 37 &#176;C.</li>
	<li>Incubate for 60 s at room temperature.</li>
	<li>Remove the solution and gently add 1 mL per well of preheated and freshly prepared 4% PFA (paraformaldehyde) solution diluted in the cytoskeleton buffer. 
	Caution: Use gloves and work under the chemical hood.</li>
	<li>Place the 12-well plate back in the incubator at 37&#176;C for 10 min.</li>
	<li>Remove the PFA and add 3 mL of PBS per well. 
	Note: Work under the chemical hood with gloves and put the recycled PFA in a specific bin.</li>
	<li>Wash twice with PBS.</li>
	<li>Remove the PBS and add freshly prepared 10 mM NaBH4 solution diluted in PBS in order to quench the autofluorescence coming from glutaraldehyde.</li>
	<li>Incubate for 7 min at room temperature.</li>
	<li>Put the recycled NaBH4 in a specific bin.</li>
	<li>Wash thrice 5 min with PBS.</li>
	<li>Remove the PBS and add the blocking solution (5% BSA solution in PBS).</li>
	<li>Incubate for 1 h at room temperature (or overnight at 4 &#176;C). 
	Note: The cytoskeleton buffer can be stored at room temperature for a few months. PFA, NaBH4 and glutaraldehyde should be handled with special care (hood, gloves, and specific recycling bins). Solutions should be added and removed gently in order to preserve the integrity of the cells. It is important to not let the cells dry.</li>
</ol>
4. Immunostaining (Day 2, 5h)
<ol>
	<li>Prepare the solution of primary antibodies using anti-vimentin and anti-tubulin with a 1:500 dilution in the blocking solution.</li>
	<li>Put 100 &#181;L drops of diluted primary antibodies on a paraffin film layer and place the coverslips on top of them with cells facing downward.</li>
	<li>Incubate cells with primary antibodies for 2 h. Put the coverslips back in a 12-well plate filled with PBS. Rinse thrice for 5 min each time in PBS at room temperature on an orbital shaker.</li>
	<li>In the meantime, prepare the secondary antibodies solution using anti-mouse Alexa647 and anti-rat Alexa555 at a dilution of a 1:1, 000 in the blocking solution. Incubate cells with secondary antibodies for 1 h at room temperature. Proceed like 4.2. protecting the cells from light.</li>
	<li>Put the coverslip back in a 12-well plate filled with PBS. Rinse thrice for 5 min each time in PBS at room temperature on an orbital shaker. Remove the PBS and add 0.5 mL of PBS mixed with 1 &#181;L of 0.1 &#181;m tetraspeck beads.</li>
	<li>Put the wells on an orbital shaker for 30 min. Rinse with PBS. Remove the PBS and incubate the cells for 5 min with a 4% PFA solution in PBS. Rinse thrice in PBS. 
	Note: Fixed cells can be stored for a couple of weeks in PBS at 4&#176;C. Immunostaining should be done at the last minute.</li>
</ol>
5. Sample preparation (Day 3, 5 min)
<ol>
	<li>Put aliquots of MEA (cysteamine, 1 M, pH 8.3), glucose oxidase (100x, 50 mg/mL), catalase (500x, 20 mg/mL) in an ice basket.</li>
	<li>Prepare 50 mL of Tris-NaCl buffer (50 mM Tris, 10 mM NaCl, pH = 8) supplemented with 10% of glucose (100 mg/mL).</li>
	<li>Prepare 1.2 mL of imaging buffer just before imaging with 10 mM MEA, 0.5 mg/mL (75 U/mL) glucose oxidase, 40 &#181;g/mL catalase (80 - 200 U/mL) in the Tris-NaCl buffer with 10% glucose.</li>
	<li>Mount the coverslip containing the labeled cells on the magnetic sample holder (e.g. chamlide chamber) and add the imaging buffer to fill entirely the chamber. Put the lid on and make sure that there is no air bubble between the imaging buffer and the lid.</li>
	<li>Clean the bottom of the sample with absolute ethanol and verify that there is not leak. Use grease to seal the sample holder properly if necessary. 
	Note: Prepare 1 M stock of MEA solution (pH 8.3 adjusted using HCl), stock of glucose oxidase at 50 mg/mL and stock of catalase at 20 mg/mL in water, make aliquots and store them at -20&#176;C for up to one month for MEA and a year for glucose oxidase and catalase. Do not refreeze aliquots. Prepare the Tris-NaCl buffer in advance and store it at room temperature for several months. Add glucose on the day of the experiment (step 5.3).</li>
</ol>
6. Image acquisition (Day 3, ~1h per cell)
<ol>
	<li>Switch on the microscope. Switch on the acquisition software and press Start system.</li>
	<li>Select the Acquisition Tab. Expand the Imaging Setup tool in the Setup Manager tool group. Select the laser WF (wide field) mode of acquisition. Select the 100X NA 1.46 objective. Select the TIRF u-HP mode which uses a collimator to reduce the field of view being illuminated and concentrate the laser energy into a smaller area. Use the optovar lens 1.6x that gives a pixel size of 100nm.</li>
	<li>Select the Time series tool and set up the time lapse with 50 000 frames and 0.0 ms between frames. Expand the Channels tool in the Acquisition parameter tool group and select &#34;show all&#34;.</li>
	<li>Set the Alexa647 track: check the 642 and 405 laser lines for excitation and accept to turn them on. Select the &#34;655 LP&#34; emission filter in the Imaging Setup. Rename &#34;647&#34;.</li>
	<li>Click on &#39;+&#39; in the Channels tool to add another light pass and select the mode &#34;switch track every: frame&#34; in the Imaging Setup tool. Check 561, 488 and 405 laser lines and accept to turn them on. Select the &#34;BP 570-650+LP750&#34; emission filter and use optovar lens 1.6x. Check that the TIRF-uHP mode is selected. Rename the track &#34;555&#34;.</li>
	<li>Select the Acquisition mode tool and set the frame size to 200 x 200 pixels. Select the Focus devices and strategy tool and set the definite focus to 1 000 frames (if there is a focus-lock system).</li>
	<li>Put oil on the objective and put the sample on. Select the Locate tab, and open the shutter of the fluorescence lamp. Find the focus and look for the cell to image using the eyepiece. Put it in the center of the field of view using the joystick.</li>
	<li>Select the Acquisition tab to go back to the acquisition mode. Select the &#34;647&#34; track, set the 642-laser power to 0.2%, set the 405-laser power to 0, set the exposure time to 50 ms and the gain of 50. Choose the Continuous acquisition mode to observe the cell on the screen. Adjust the focus. Make sure there is at least one tetraspeck bead in the field of view, increase the contrast to see them if necessary. Use an auto-scale mode for the display. Take a wide-field epifluorescence image of the vimentin network by clicking on snap and save it. To check the TIRF illumination, click on TIRF, adjust the angle of illumination and/or collimator if necessary to have a homogeneous field of illumination and save it. 
	NOTE: It is important that the TIRF and epifluorescent images are of high quality before starting the raw data acquisition. Discontinuous, non-uniformly labeled filaments will give rise to poor quality dSTORM images.</li>
	<li>Uncheck the &#34;647&#34; track and select and check the &#34;555&#34; track. Take an epifluorescence image of the microtubule network and save it. Take a TIRF image and save it too.</li>
	<li>Uncheck the &#34;555&#34; track, select and check the &#34;647&#34; track, then press Continuous. Put the gain to 0 and exposure time to 10 ms. Set the 642-nm laser power to 100% in order to pump most of the Alexa647 dyes to the dark state (~2 kW/cm2). Once the fluorophores start blinking, put the exposure to 15 ms and gain to 300. Select the TIRF mode of illumination. Use the Range indicator mode without auto-scale to verify that single molecule signals do not saturate the camera (indicated by the red color). In that case, decrease the gain until the saturation disappears. Tetraspeck beads will remain saturated at the beginning of the acquisition. Press Stop to stop the continuous observation of the cell.</li>
	<li>Expand the Online processing tool and check Online processing PALM to visualize the STORM image during acquisition. Use the following parameters: 9 for the Peak mask size (9 pixels diameter), and 6 for the Peak intensity to noise. Theses parameters will define the molecules that are taken into account in the processing (bright enough and not overlapping). In the PAL-Statistics tool, select Plot type: histogram, and Histogram Source: Precision. Press Start acquisition.</li>
	<li>During acquisition, refocus if necessary (if there is no focus-lock device). Adjust the parameters (exposure, laser powers) and eventually switch on the 405-nm laser line (starting at 0.001%) to keep a medium density of fluorophores with a minimum distance of 1 &#181;m between single molecules (&#62; 9 pixels of 100 nm, the Peak mask size) (Figure 1A-B). If the molecule density is a little bit too high, use the HILO (Highly Inclined and Laminated Optical sheet) mode of illumination instead of TIRF. This will increase the laser power by 20%. Make sure that the peak of localization precision on the histogram below the reconstructed image is centered around or below 10 nm. 
	Note: Usually the number of molecules decreases with time, and the 405-nm laser power is increased slowly accordingly. The goal is to decrease the localization precision (histogram displayed below the super-resolution image) as much as possible with a peak below 10 nm (Figure 1C). Increase the contrast of the PALM image if necessary, if too many bright beads are present in the field.</li>
	<li>Press Stop to stop the movie when more than 106 localizations have been reached (typically after 40 000 frames). Put the lasers back to 0.2% (642 nm) and 0 (405 nm). Save the raw data of the STORM acquisition with the .czi format. Uncheck track &#34;647&#34; in the Channels tool, and select and check track &#34;555&#34;.</li>
	<li>Set the exposure time to 25 ms and gain to 0. Press the Continuous button and set both 488 and 561 lasers at 100% to pump Alexa555 dyes to the dark state (~2 kW/cm2 each). Once the fluorophores start blinking, put the gain to 250, use the HILO illumination mode and check that single molecules do not saturate the camera with the range indicator mode. If that is the case, decrease the gain. Press Stop. Decrease the 488-laser power to 50%.</li>
	<li>Press Start acquisition. During acquisition, increase progressively the gain to always be at the limit of saturation. Increase step by step the 405-nm laser power to keep a medium density of single molecule in the field of view (See step 6.18). Decrease the 488 laser power to 20-30% when the 405 laser is higher than 15%. Then decrease gradually the 488 while increasing the 405 laser line in order to keep the blinking of the molecules as fast as possible. The 488 laser line coupled with the 561 excitation laser at maximal intensity increases both the on and off rates of the fluorophores, whereas the 405 laser line only increases the on rate.</li>
	<li>Stop the acquisition when more than 500 000 localizations have been reached (after about 20 000 frames) or more when no more molecules are blinking. Save the raw data of the STORM acquisition with the .czi format. 
	Note: Always start with the higher wavelength to avoid bleaching (or activation) of the other colors. Since the chamlide chamber is not sealed, it is possible to change the imaging buffer regularly, e.g. to test different buffer conditions. Using the 488 laser line to deplete the Alexa 555 dyes is necessary only when the 561 laser power is limited (with 100 mW lasers).</li>
</ol>
7. Image reconstruction (5 min)
<ol>
	<li>In the PAL-Drift tool, use the Model-based correction type with automatic segments with a maximum size of 8. Press Apply multiple times in a row until the drift is corrected. This model-based correction applies an algorithm based on cross-correlation analysis which corrects the drift.</li>
	<li>Using the PAL-Filter tools, improve the appearance of the STORM image by selecting only the molecules which were best localized. For example, limit the Localization precision to 30 nm and the PSF to 120-180 nm.</li>
	<li>Use a pixel resolution of 5 or 10 nm in the PAL-render tool, as well as a Gaussian display mode, with an expansion factor of 1 PSF. Select Render auto dynamic range HR with a 95% value and Render auto dynamic range SWF with a 90% value. Select Render best quality.</li>
	<li>Select PALM convert, in the PALM menu of the Processing tab (post-processing mode). Press Select and then Apply. Save the created image with a format. LSM (and not .czi, very important). 
	Note: The image reconstruction can be performed immediately after the acquisition or later using the image processing mode of the acquisition software. In the off line post-processing mode, it is possible to reanalyze the stacks with different values of parameters (peak mask size, peak intensity to noise). Always choose to &#34;discard overlapping molecules&#34; and not &#34;average before localization&#34;.</li>
</ol>
8. Estimation of the localization precision of a STORM image (10 min)
<ol>
	<li>Open your STORM raw data and use the Image processing tab. Select the PALM method and then PALM again. Press Select.</li>
	<li>Press Apply to start the reconstruction using a peak mask size of 9 and a peak intensity to noise of 6 (or the parameters you choose for the image).</li>
	<li>Select the rectangle Graphics tool and draw a rectangle on the raw image around a single molecule present on the glass surface. There are usually a few single molecules there.</li>
	<li>Press Apply again, and only the molecules present inside the rectangle region will be analyzed.</li>
	<li>In the PAL-Statistics tool, select Plot type: histogram, and Histogram Source: X position or Y position, to visualize the position histograms.</li>
	<li>Save the table with the list of localizations on the left below the images.</li>
	<li>Using a software of data analysis, create histograms of X and Y positions (Figure 1D).</li>
	<li>Fit the distributions by a Gaussian and save the &#963;X and &#963;Y values. The localization precision &#963;SMLM is estimated by (&#963;X*&#963;Y)^0.5.</li>
	<li>Repeat step 8.3-8.8 on as many molecules as possible and average the &#963;SMLM</li>
</ol>
9. Channel registration (5 min)
<ol>
	<li>Open the STORM image of each color using the Fiji (Image J) software.</li>
	<li>Select the wand tool to measure the positions of each tetraspeck beads.</li>
	<li>Calculate the X and Y shifts for each bead and average them.</li>
	<li>Use the &#34;Translate&#34; command to translate the second image with the calculated X and Y shifts.</li>
	<li>Merge the two images using the Merge Channels command.</li>
</ol>

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R., Bell, T. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Image+artifacts+in+single+molecule+localization+microscopy:+why+optimization+of+sample+preparation+protocols+matters.">Image artifacts in single molecule localization microscopy: why optimization of sample preparation protocols matters.</a> Sci Rep. 5, 7924 (2015).</li><li>Ovesny, M., Krizek, P., Borkovec, J., Svindrych, Z., Hagen, G. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=ThunderSTORM:+a+comprehensive+ImageJ+plug-in+for+PALM+and+STORM+data+analysis+and+super-resolution+imaging.">ThunderSTORM: a comprehensive ImageJ plug-in for PALM and STORM data analysis and super-resolution imaging.</a> Bioinformatics. 30 (16), 2389-2390 (2014).</li><li>Olivier, N., Keller, D., Gonczy, P., Manley, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Resolution+doubling+in+3D-STORM+imaging+through+improved+buffers.">Resolution doubling in 3D-STORM imaging through improved buffers.</a> PLoS One. 8 (7), e69004 (2013).</li><li>Nahidiazar, L., Agronskaia, A. V., Broertjes, J., van den Broek, B., Jalink, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optimizing+Imaging+Conditions+for+Demanding+Multi-Color+Super+Resolution+Localization+Microscopy.">Optimizing Imaging Conditions for Demanding Multi-Color Super Resolution Localization Microscopy.</a> PLoS One. 11 (7), e0158884 (2016).</li><li>Huang, B., Wang, W., Bates, M., 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=Three-dimensional+super-resolution+imaging+by+stochastic+optical+reconstruction+microscopy.">Three-dimensional super-resolution imaging by stochastic optical reconstruction microscopy.</a> Science. 319 (5864), 810-813 (2008).</li><li>Mikhaylova, M., 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=Resolving+bundled+microtubules+using+anti-tubulin+nanobodies.">Resolving bundled microtubules using anti-tubulin nanobodies.</a> Nat Commun. 6, 7933 (2015).</li><li>Nieuwenhuizen, R. P., 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=Measuring+image+resolution+in+optical+nanoscopy.">Measuring image resolution in optical nanoscopy.</a> Nat Methods. 10 (6), 557-562 (2013).</li><li>Schnitzbauer, J., Strauss, M. T., Schlichthaerle, T., Schueder, F., Jungmann, R. <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+microscopy+with+DNA-PAINT.">Super-resolution microscopy with DNA-PAINT.</a> Nat Protoc. 12 (6), 1198-1228 (2017).</li></ol>

The authors declare no competing financial interests.

Critical steps in the protocol
We present here a protocol which minimizes artifacts in the dSTORM images of microtubules and IFs in glial cells. Artifacts can be created at every step of the sample preparation and imaging: fixation, blocking, immunolabeling, drift during acquisition, non-optimal blinking conditions23. We list below the most critical steps.
Cleaning the coverslips is an important step to limit the non-specific...

Cytoplasmic intermediate filaments (IFs) are 10 nm-diameter homo- or heteropolymers of a cell type specific subset of IF proteins. IFs participate in a large range of cellular functions such as cell motility, proliferation and stress responses. Their key role is highlighted by the fact that more than 90 human diseases are directly caused by mutations in IF proteins; for instance, changes in IF composition accompanies tumour growth and spreading1,2,3. There is growing evidence that the three cytoskeleton systems work in collaboration to control cellular functions such as cell polarization, division and migration2,3. Since there is a tight coupling between spatial architecture and functions, it is crucial to get insights into the structural organization of the IF with the other filaments and better understand the cytoskeletal cross-talk. this article provides a protocol to perform the super-resolution technique dual-color dSTORM (direct STochastic Optical Reconstruction Microscopy)4 and how it is used to investigate the interaction between IF and microtubules in fixed, cultured cells in 2 dimensions (2D).
Several super-resolution techniques have been developed over the past decade, and there discoveries were at the origin of the Nobel Prize in Chemistry in 20145. Among all these techniques lies the single molecule localization-based methods like PALM6 (Photo-Activated Localization Microscopy) which uses photoswitchable fluorescent proteins, STORM7 with pairs of fluorophores or dSTORM4 with conventional fluorophores. All these methods are based on the same principle which consists of (i) the switch of most of the fluorescent reporters into an &#34;off&#34; state&#34; (non-fluorescent), (ii) the stochastic activation of a subset of them into a fluorescent state in order to localize their spatial position with nanometer accuracy, and (iii) the repetition of this process in order to activate as many subsets of fluorophores as possible. A final image is reconstructed using all the localizations of the activated molecules, providing a lateral resolution down to ~20-40 nm. Several commercialized optical systems allowing PALM/STORM are now available for biologists for routine experiments. Here, one such system was used to study the structural association of microtubules and IFs. Among all the single-molecule localization-based methods, the dual-color dSTORM technique was selected, because it is well suited to observe very thin lamellar regions (&#60;1 &#181;m) of adherent cells and can provide significant improvement of image resolution with minimal time and money investment. Indeed, dSTORM is a very convenient and versatile technique that is compatible with the standard organic dyes routinely used for cellular staining and immunofluorescence.
While one color dSTORM images are relatively simple to obtain using the fluorophore Alexa6478, dual color dSTORM requires to optimize the experimental conditions so that two dyes can blink properly in the chosen buffer, especially when there is limited laser power. Excellent papers are already available on the methods to set up multi-color imaging with dSTORM9,10,11,12,13, and these papers explain in detail the possible sources of artifacts and degraded image resolution as well as how to overcome them. In this article, the optimal experimental conditions in terms of cells fixation, immunostaining, sample preparation, imaging acquisition and image reconstruction are described in order to acquire images of dense cytoplasmic IF networks and microtubules in glial cells with dual-color dSTORM. Briefly, the extraction/fixation method described in Chazeau et al12 was adapted to glial cells and used with a post-fixation step, optimized antibodies concentration, a STORM buffer with 10 mM MEA described in Dempsey9 et al. which was found to be optimal for the experimental set up and sample type.
Glial cells mainly express three types of IF proteins: vimentin, nestin and GFAP (glial acidic fibrillary protein). These three proteins were shown to co-polymerize in astrocytes14. We previously showed using super-resolution structured illumination microscopy (SR SIM) that these three IF proteins can be found in the same single IF filament and that they display similar distribution and dynamics in glial cells 15. Due to the similarities between the three IF proteins, vimentin staining was used as a reporter for the whole IF network. Using dSTORM, we managed to resolve how IF form bundles along microtubules, which was not possible with diffraction limited microscopy techniques15. These observations may help to understand how vimentin IFs can template microtubules and regulate their growth trajectory, promoting the maintenance of a polarity axis during cell migration16. Super-resolution images provided key information on the mutual interaction of the two cytoskeletal subsystems, and brought insight into the link between spatial association and local function which could be cell-type specific17. In general, dual-color dSTORM can be used to study the crosstalk between cytoskeletal elements or other types of organelles, provided that good immunostaining conditions are reached in terms of density and specificity. This technique will be useful to better characterize the cytoskeleton changes observed during astrogliosis and in glioblastoma, the most common and most malignant tumor of the central nervous system, where the expression of IF proteins is altered 18,19,20,21.

<table>
	<tbody>
		<tr>
			<td>Minimum Essential Media (MEM)</td>
			<td>Gibco</td>
			<td>41090-028</td>
			<td>cell culture</td>
		</tr>
		<tr>
			<td>non essential amino acid</td>
			<td>Gibco</td>
			<td>1140-050</td>
			<td>cell culture 1/100e</td>
		</tr>
		<tr>
			<td>Penicillin-Streptomycin</td>
			<td>Gibco</td>
			<td>15140-122</td>
			<td>cell culture 1/100e</td>
		</tr>
		<tr>
			<td>F&#339;tal Bovine Serum</td>
			<td>Gibco</td>
			<td>10270-106</td>
			<td>cell culture 1/10e</td>
		</tr>
		<tr>
			<td>0,05 % Trypsin-EDTA (1X)</td>
			<td>Gibco</td>
			<td>25300-054</td>
			<td>cell culture</td>
		</tr>
		<tr>
			<td>PBS 10x</td>
			<td>fischer scientific</td>
			<td>11540486</td>
			<td>cell culture</td>
		</tr>
		<tr>
			<td>coverslip 18 mm n&#176;1,5H</td>
			<td>Marienfield/Dominic dutsher</td>
			<td>900556</td>
			<td>coverslips</td>
		</tr>
		<tr>
			<td>paraformaldehyde (PFA) solution</td>
			<td>Sigma</td>
			<td>F8775</td>
			<td>cell fixation</td>
		</tr>
		<tr>
			<td>glutaraldehyde</td>
			<td>Sigma</td>
			<td>G5882</td>
			<td>cell fixation</td>
		</tr>
		<tr>
			<td>triton X100</td>
			<td>Sigma</td>
			<td>T9284</td>
			<td>non ionic surfactant. Used for cell fixation</td>
		</tr>
		<tr>
			<td>sodium borohydride (NaBH4)</td>
			<td>Sigma</td>
			<td>452904-25ML</td>
			<td>cell fixation</td>
		</tr>
		<tr>
			<td>BSA</td>
			<td>Sigma</td>
			<td>A7906</td>
			<td>sample passivation 3% in PBS</td>
		</tr>
		<tr>
			<td>Monoclonal Anti-Vimentin (V9) antibody produced in mouse</td>
			<td>Sigma</td>
			<td>V6630</td>
			<td>primary antibodies</td>
		</tr>
		<tr>
			<td>Rat anti alpha tubulin</td>
			<td>Biorad</td>
			<td>MCA77G</td>
			<td>primary antibodies</td>
		</tr>
		<tr>
			<td>Goat anti-Rat IgG (H+L) Secondary Antibody, Alexa Fluor 555</td>
			<td>Fisher scientific</td>
			<td>10635923</td>
			<td>secondary antibodies</td>
		</tr>
		<tr>
			<td>Donkey anti-Mouse IgG (H+L) Secondary Antibody, Alexa Fluor 647</td>
			<td>Fisher scientific</td>
			<td>10226162</td>
			<td>secondary antibodies</td>
		</tr>
		<tr>
			<td>TetraSpeck Microspheres, 0.1 &#181;m, fluorescent blue/green/orange/dark red</td>
			<td>Fisher scientific</td>
			<td>T7279</td>
			<td>fluorescent beads</td>
		</tr>
		<tr>
			<td>Chamlide magnetic chamber</td>
			<td>Live Cell Instrument</td>
			<td>CM-B18-1</td>
			<td>magnetic sample holder, maximum volume 1,2 mL</td>
		</tr>
		<tr>
			<td>Cysteamine (MEA)</td>
			<td>Sigma</td>
			<td>30070</td>
			<td>for the blinking buffer</td>
		</tr>
		<tr>
			<td>glucose oxidase from Aspergillus niger</td>
			<td>Sigma</td>
			<td>G0543</td>
			<td>for the blinking buffer</td>
		</tr>
		<tr>
			<td>glucose</td>
			<td>sigma</td>
			<td></td>
			<td>for the blinking buffer</td>
		</tr>
		<tr>
			<td>catalase from bovine liver</td>
			<td>Sigma</td>
			<td>C9322</td>
			<td>for the blinking buffer</td>
		</tr>
		<tr>
			<td>Tris (trizma)</td>
			<td>Sigma</td>
			<td>T1503</td>
			<td>for the blinking buffer</td>
		</tr>
		<tr>
			<td>Wash-N-Dry&#160;coverslip rack</td>
			<td>Sigma</td>
			<td>Z688568</td>
			<td></td>
		</tr>
		<tr>
			<td>Parafilm</td>
			<td>Sigma</td>
			<td>P7793-1EA</td>
			<td>paraffin film</td>
		</tr>
		<tr>
			<td>ultrasonic cleaner</td>
			<td>Fisher scientific</td>
			<td>15-335-6</td>
			<td></td>
		</tr>
		<tr>
			<td>plasma cleaner</td>
			<td>Harrick plasma</td>
			<td>PDC-32G-2</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

A microscope equipped with 50 mW 405 nm and 100 mW 488, 561 and 642 nm solid-state lasers, an EMCCD 512x512 camera, an alpha Plan Apo 100X/1.46 objective and Band Pass 570-650 / Long Pass 655 emission filters was used for the representative results presented below.
Figure 1A gives an example of molecule density and signal to noise ratio that should be used during raw image acquisition. Good quality ...

Watch this Scientific Journal Video about Imaging Intermediate Filaments and Microtubules with 2-dimensional Direct Stochastic Optical Reconstruction Microscopy at JoVE.com

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

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

The cytoskeleton, composed of actin microfilaments, microtubules, and intermediate filaments (IF), plays a key role in the control of cell shape, ...

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10.8K Views.  Institut Pasteur. 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 

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

Labeling Stem Cells with Fluorescent Dyes for non-invasive Detection with Optical Imaging

Optical imaging (OI) is an easy, fast and inexpensive tool for in vivo monitoring of new stem cell based therapies. The technique is based on ex vivo labeling of stem cells with a fluorescent dye, subsequent intravenous injection of the labeled cells and visualization of their accumulation in specific target organs or pathologies. The presented technique demonstrates how we label human mesenchymal stem cells (hMSC) by simple incubation with the lipophilic fluorescent dye DiD (C67H103CIN2O3S) and how we label human embryonic stem cells (hESC) with the FDA approved fluorescent dye Indocyanine Green (ICG). The uptake mechanism is via adherence and diffusion of the lypophilic dye across the phospholipid cell membrane bilayer. The labeling efficiency is usually improved if the cells are incubated with the dye in serum-free media as opposed to incubation in serum-containing media. Furthermore, the addition of the transfection agent Protamine Sulfate significantly improves contrast agent uptake.

This video shows techniques for labeling of human embryonic stem cells and mesenchymal stem cells with fluorescent dyes. This technique can be used for an in vivo tracking of transplanted stem cells with optical imaging and for histopathological correlations with fluorescence microscopy.

Optical imaging (OI) is an easy, fast and inexpensive tool for in vivo monitoring of new stem cell based therapies. The technique is based on ex vivo ...

Proper Care and Cleaning of the Microscope

Keeping the microscope optics clean is important for high-quality imaging. Dust, fingerprints, excess immersion oil, or mounting medium on or in a microscope causes reduction in contrast and resolution. DIC is especially sensitive to contamination and scratches on the lens surfaces. This protocol details the procedure for keeping the microscope clean.

Keeping the microscope optics clean is important for high-quality imaging. Dust, fingerprints, excess immersion oil, or mounting medium on or in a microscope causes reduction in contrast and resolution. DIC is especially sensitive to contamination and scratches on the lens surfaces. This protocol details the procedure for keeping the microscope clean.

Keeping the microscope optics clean is important for high-quality imaging. Dust, fingerprints, excess immersion oil, or mounting medium on or in a ...

Major Components of the Light Microscope

The light microscope is a basic tool for the cell biologist, who should have a thorough understanding of how it works, how it should be aligned for different applications, and how it should be maintained as required to obtain maximum image-forming capacity and resolution. The components of the microscope are described in detail here.

The light microscope is a basic tool for the cell biologist, who should have a thorough understanding of how it works, how it should be aligned for different applications, and how it should be maintained as required to obtain maximum image-forming capacity and resolution. The components of the microscope are described in detail here.

The light microscope is a basic tool for the cell biologist, who should have a thorough understanding of how it works, how it should be aligned for ...

Phase Contrast and Differential Interference Contrast (DIC) Microscopy

Phase-contrast microscopy is often used to produce contrast for transparent, non light-absorbing, biological specimens. The technique was discovered by Zernike, in 1942, who received the Nobel prize for his achievement. DIC microscopy, introduced in the late 1960s, has been popular in biomedical research because it highlights edges of specimen structural detail, provides high-resolution optical sections of thick specimens including tissue cells, eggs, and embryos and does not suffer from the  phase halos  typical of phase-contrast images. This protocol highlights the principles and practical applications of these microscopy techniques.

Phase Contrast and Differential Interference Contrast DIC Microscopy

This protocol highlights the principles and practical applications of Phase and Differential Interference Contrast (DIC) Microscopy

Phase-contrast microscopy is often used to produce contrast for transparent, non light-absorbing, biological specimens. The technique was discovered by ...

Imaging Exocytosis in Retinal Bipolar Cells with TIRF Microscopy

Total internal reflectance fluorescence (TIRF) microscopy is a technique that allows the study of events happening at the cell membrane, by selective imaging of fluorescent molecules that are closest to a high refractive index substance such as glass1. In this article, we apply this technique to image exocytosis of synaptic vesicles in retinal bipolar cells isolated from the goldfish retina. These neurons are very suitable for this kind of study due to their large axon terminals. By simultaneously patch clamping the bipolar cells, it is possible to investigate the relationship between pre-synaptic voltage and synaptic release2,3. Synaptic vesicles inside the bipolar cell terminals are loaded with a fluorescent dye (FM 1-43&reg;) by co-puffing the dye and a ringer solution containing a high K+ concentration onto the synaptic terminals. This depolarizes the cells and stimulates endocytosis and consequent dye uptake into the glutamatergic vesicles. After washing the excess dye away for around 30 minutes, cells are ready for being patch clamped and imaged simultaneously with a 488 nm laser. The patch pipette solution contains a rhodamine-based peptide that binds selectively to the synaptic ribbon protein RIBEYE4, thereby labeling ribbons specifically when terminals are imaged with a 561 nm laser. This allows the precise localization of active zones and the separation of synaptic from extra-synaptic events.

In this video, we demonstrate how to label and visualize single synaptic vesicle exocytosis and trafficking in goldfish retinal bipolar cells using total internal reflectance fluorescence (TIRF) microscopy.

Total internal reflectance fluorescence (TIRF) microscopy is a technique that allows the study of events happening at the cell membrane, by selective ...

Direct Imaging of ER Calcium with Targeted-Esterase Induced Dye Loading (TED)

Visualization of calcium dynamics is important to understand the role of calcium in cell physiology. To examine calcium dynamics, synthetic fluorescent Ca2+ indictors have become popular. Here we demonstrate TED (= targeted-esterase induced dye loading), a method to improve the release of Ca2+ indicator dyes in the ER lumen of different cell types. To date, TED was used in cell lines, glial cells, and neurons in vitro. TED bases on efficient, recombinant targeting of a high carboxylesterase activity to the ER lumen using vector-constructs that express Carboxylesterases (CES). The latest TED vectors contain a core element of CES2 fused to a red fluorescent protein, thus enabling simultaneous two-color imaging. The dynamics of free calcium in the ER are imaged in one color, while the corresponding ER structure appears in red. At the beginning of the procedure, cells are transduced with a lentivirus. Subsequently, the infected cells are seeded on coverslips to finally enable live cell imaging. Then, living cells are incubated with the acetoxymethyl ester (AM-ester) form of low-affinity Ca2+ indicators, for instance Fluo5N-AM, Mag-Fluo4-AM, or Mag-Fura2-AM. The esterase activity in the ER cleaves off hydrophobic side chains from the AM form of the Ca2+ indicator and a hydrophilic fluorescent dye/Ca2+ complex is formed and trapped in the ER lumen. After dye loading, the cells are analyzed at an inverted confocal laser scanning microscope. Cells are continuously perfused with Ringer-like solutions and the ER calcium dynamics are directly visualized by time-lapse imaging. Calcium release from the ER is identified by a decrease in fluorescence intensity in regions of interest, whereas the refilling of the ER calcium store produces an increase in fluorescence intensity. Finally, the change in fluorescent intensity over time is determined by calculation of &Delta;F/F0.

Direct Imaging of ER Calcium with Targeted-Esterase Induced Dye Loading TED

Targeted-esterase induced dye loading (TED) supports the analysis of intracellular calcium store dynamics by fluorescence imaging. The method bases on targeting of a recombinant Carboxylesterase to the endoplasmic reticulum (ER), where it improves the local unmasking of synthetic low-affinity Ca2+ indicator dyes in the ER lumen.

Visualization of calcium dynamics is important to understand the role of calcium in cell physiology. To examine calcium dynamics, synthetic fluorescent ...

Imaging and 3D Reconstruction of Cerebrovascular Structures in Embryonic Zebrafish

Zebrafish are a powerful tool to study developmental biology and pathology in vivo. The small size and relative transparency of zebrafish embryos make them particularly useful for the visual examination of processes such as heart and vascular development. In several recent studies transgenic zebrafish that express EGFP in vascular endothelial cells were used to image and analyze complex vascular networks in the brain and retina, using confocal microscopy. Descriptions are provided to prepare, treat and image zebrafish embryos that express enhanced green fluorescent protein (EGFP), and then generate comprehensive 3D renderings of the cerebrovascular system. Protocols include the treatment of embryos, confocal imaging, and fixation protocols that preserve EGFP fluorescence. Further, useful tips on obtaining high-quality images of cerebrovascular structures, such as removal the eye without damaging nearby neural tissue are provided. Potential pitfalls with confocal imaging are discussed, along with the steps necessary to generate 3D reconstructions from confocal image stacks using freely available open source software.

Imaging of cerebrovascular development in larval zebrafish is described. Techniques to facilitate 3D imaging and modify cerebrovascular development using chemical treatments are also provided.

Zebrafish are a powerful tool to study developmental biology and pathology in vivo.  The small size and relative transparency of zebrafish embryos make ...

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) maintain the turnover of all mature blood cell lineages. The coordination of the complex signals leading to specific HSC fates relies upon the interaction between HSCs and the intricate bone marrow microenvironment, which is still poorly understood[1-2].
We describe how by combining a newly developed specimen holder for stable animal positioning with multi-step confocal and two-photon in vivo imaging techniques, it is possible to obtain high-resolution 3D stacks containing HSPCs and their surrounding niches and to monitor them over time through multi-point time-lapse imaging. High definition imaging allows detecting ex vivo labeled hematopoietic stem and progenitor cells (HSPCs) residing within the bone marrow. Moreover, multi-point time-lapse 3D imaging, obtained with faster acquisition settings, provides accurate information about HSPC movement and the reciprocal interactions between HSPCs and stroma cells.
Tracking of HSPCs in relation to GFP positive osteoblastic cells is shown as an exemplary application of this method. This technique can be utilized to track any appropriately labeled hematopoietic or stromal cell of interest within the mouse calvarium bone marrow space.

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

The nature of the interactions between hematopoietic stem and progenitor cells (HSPCs) and bone marrow niches is poorly understood. Custom hardware modifications and a multi-step acquisition protocol allow the use of two-photon and confocal microscopy to image ex vivo labeled HSPCs homed within bone marrow areas, tracking interactions and movement.

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) ...

Tandem High-pressure Freezing and Quick Freeze Substitution of Plant Tissues for Transmission Electron Microscopy

Since the 1940s transmission electron microscopy (TEM) has been providing biologists with ultra-high resolution images of biological materials. Yet, because of laborious and time-consuming protocols that also demand experience in preparation of artifact-free samples, TEM is not considered a user-friendly technique. Traditional sample preparation for TEM used chemical fixatives to preserve cellular structures. High-pressure freezing is the cryofixation of biological samples under high pressures to produce very fast cooling rates, thereby restricting ice formation, which is detrimental to the integrity of cellular ultrastructure. High-pressure freezing and freeze substitution are currently the methods of choice for producing the highest quality morphology in resin sections for TEM. These methods minimize the artifacts normally associated with conventional processing for TEM of thin sections. After cryofixation the frozen water in the sample is replaced with liquid organic solvent at low temperatures, a process called freeze substitution. Freeze substitution is typically carried out over several days in dedicated, costly equipment. A recent innovation allows the process to be completed in three hours, instead of the usual two days. This is typically followed by several more days of sample preparation that includes infiltration and embedding in epoxy resins before sectioning. Here we present a protocol combining high-pressure freezing and quick freeze substitution that enables plant sample fixation to be accomplished within hours. The protocol can readily be adapted for working with other tissues or organisms. Plant tissues are of special concern because of the presence of aerated spaces and water-filled vacuoles that impede ice-free freezing of water. In addition, the process of chemical fixation is especially long in plants due to cell walls impeding the penetration of the chemicals to deep within the tissues. Plant tissues are therefore particularly challenging, but this protocol is reliable and produces samples of the highest quality.

Obtaining high-quality transmission electron microscopy images is challenging, especially in the case of plant cells, which have abundant large water-filled vacuoles and aerated spaces. Tandem high-pressure freezing and quick freeze substitution greatly reduce preparation time of plant samples for TEM while producing samples with excellent ultrastructural preservation.

Since the 1940s transmission electron microscopy (TEM) has been providing biologists with ultra-high resolution images of biological materials. Yet, ...

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.

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.

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