The authors would like to thank Blanka Duvnjak and Nikola Kuhr for their excellent technical assistance.

1. Splitting and seeding of cells
<ol>
	<li>Warm up sterile Dulbecco&#39;s Modified Eagle&#39;s Medium (DMEM) including 10% fetal calf serum (FCS), sterile Phosphate buffered saline (PBS) and sterile trypsin to 37 &#176;C. Activate the clean bench.</li>
	<li>Prepare a 6-well plate by adding one sterile glass cover slip (10 mm) to each well using sterile forceps.</li>
	<li>Put a 10 cm cell culture dish with Cos7 cells under the clean bench. Under the clean bench, aspirate the cells&#39; medium using a vacuum device.</li>
	<li>Hold the angulated cell culture dish in one hand and add 10 mL of sterile PBS to the side of the cell culture dish to avoid rinsing off cells. Put the cell culture dish down so that PBS rinses the complete cell culture dish.</li>
	<li>Remove the PBS and add 1 mL of trypsin to the middle of the cell culture dish and incubate for 5 min at 37 &#176;C.</li>
	<li>Stop the trypsinization reaction by adding 10 mL of DMEM including 10% FCS and pipette the cell solution up and down with a pipettor to manually separate the cells.</li>
	<li>Analyze the cell number per milliliter using a counting chamber.</li>
	<li>Seed 68,000 cells in 2 mL medium per well onto the glass cover slips in the 6-well plate. Distribute the cells within the 6-well plate by cautiously shaking the plate horizontally and vertically. 
	NOTE: Cells should lie scattered.</li>
	<li>Incubate the cells overnight in a 37 &#176;C incubator with 5% CO2.</li>
</ol>
2. Transfection of cells
<ol>
	<li>Prepare two sterile 1.5 mL tubes (one for diluted DNA (A) and one for diluted reagent for cationic lipid transfection (B)). Pipette 0.75 &#181;g of nephrin and 0.75 &#181;g of podocin cDNA expression plasmid per well into one 1.5 mL tube and dilute it into 100 &#181;L of reduced serum medium per well. Add 3 &#181;L of cationic lipid transfection reagent per well to the second 1.5 mL tube and dilute with 100 &#181;L of reduced serum medium. Incubate both reactions for 5 min at room temperature.</li>
	<li>Combine both reactions (A and B) to a DNA-lipid complex and incubate for 20 min at room temperature. Cautiously add 200 &#181;L of the DNA-lipid complex to each well.</li>
	<li>Incubate the transfected cells at 37 &#176;C with 5% CO2 for 48 h without changing the medium.</li>
</ol>
3. Immunolabeling of cellular structures
<ol>
	<li>Prepare a fixing solution (4% (w/v) paraformaldehyde in PBS, 1 mL/well), the permeabilization solution (0.5% (w/v) Triton X-100 in PBS, 1 mL/well), and a blocking solution (5% (v/v) bovine serum albumin in PBS, 1 mL/well).</li>
	<li>Remove the medium with a vacuum device. Add 2 mL of PBS to each well to remove extra medium. Aspirate the PBS completely. 
	NOTE: To avoid washing away cells with PBS, make sure to pipette PBS not onto the glass cover slip directly.</li>
	<li>Fix cells with 4% (w/v) paraformaldehyde (PFA) dissolved in PBS for 10 min at room temperature. 
	NOTE: Alternatively, fix cells in 3% (v/v) glyoxal-ethanol18.</li>
	<li>Discard PFA and wash cells twice in PBS (2 mL each) by avoiding direct pipetting onto the glass cover slips. Permeabilize the fixed cells with Triton X-100 0.5% (w/v) in PBS for 10 min at room temperature.</li>
	<li>Remove the permeabilization solution and wash twice with PBS as indicated in step 3.2.</li>
	<li>Block the cells by adding 1 mL of 5% (v/v) BSA in PBS for 1 h at room temperature. Incubate the cells with 200 &#181;L of the primary antibody (anti-podocin antibody 1:200 in 1% (v/v) BSA in PBS) overnight at 4 &#176;C. 
	NOTE: Alternatively, incubate the primary antibody for 1 h at room temperature.</li>
	<li>Remove the primary antibody and wash three times with PBS as in step 3.2. Add the secondary antibody (goat anti-rabbit Alexa 488 1:1000 in 1% (v/v) BSA in PBS) for 1 h at room temperature. Keep the cells in the dark using a box.</li>
	<li>Discard the secondary antibody and wash with PBS three times.</li>
	<li>Remove PBS and incubate the cells with 200 &#181;L of anti-nephrin antibody 1:100 in 1% (v/v) BSA in PBS for 1 h at room temperature. Wash with PBS three times. Keep the cells in the dark using a box.</li>
	<li>Remove PBS and incubate with the secondary antibody donkey anti-guinea pig 633, 1:200, for 1 h at room temperature. Wash with PBS three times. Keep the cells in the dark using a box.</li>
</ol>
4. Expansion microscopy
<ol>
	<li>Preparation
	<ol>
		<li>To form the spacers for the gelation chamber, cut glass cover slips #1.0 and #1.5 into 5 mm stripes (four each per glass slide) using a diamond knife. Position the #1.5 mm cover slip stripes so that they form a square of 2.5 cm length on a glass slide and place them into a staining chamber (Figure 2A1-C1). Pipette a droplet of ddH2O into the corners of the square to adhere the glass cover slip stripes to each other and to the glass slide (Figure 2A2-C2). 
		NOTE: Avoid complete drying of ddH2O as the adhesion force will get lost. If needed, apply more droplets of ddH2O. Wait for approximately 20 min so that #1.5 mm cover slips are stably attached before starting with step 4.1.2.</li>
		<li>Place four #1.0 glass cover slip stripes per cover slip onto the #1.5 mm cover slips. Adhere the stripes by pipetting a droplet on each #1.5 mm cover slip stripe (Figure 2A3-C3). 
		NOTE: The gel&#39;s thickness should be at least 0.15 mm to facilitate handling in the progress of the protocol. However, to avoid excessive gel on top of the cells, keep the spacers height close to the cell substrate. By using #1.5 and #1.0 cover slip stripes as spacers, the spacers height will be approximately 0.3 mm. The cells adhere to the cover glass (height 0.12 mm). Therefore, the gel will be thick enough to handle but excessive gel on top of the cells is avoided using #1.0 and #1.5 cover glass stripes as spacers.</li>
		<li>For the gelation chamber lid, wrap a cover glass (#1.5) with paraffin film. Avoid any folds or dirt on the paraffin film (Figure 2A5-C5).</li>
	</ol>
	</li>
	<li>Anchoring and polymerization (gelation)
	<ol>
		<li>Prepare the anchoring buffer (see Table 1). For anchoring treatment, replace PBS with 250 &#181;L of anchoring buffer per well directly onto the glass cover slip and incubate for 3 h at room temperature. Keep substrate in the dark using a box. 
		&#8203;NOTE: Alternatively, incubate overnight at room temperature. Prepare fresh anchoring buffer for every experiment and wait for 10-15 min until it is dissolved properly. 6-((Acryloyl)amino)hexanoic acid, succinimidyl ester (AcX) should be replaced every 4-5 months to ensure adequate anchoring.</li>
		<li>Remove the anchoring buffer and wash once with 1.5 mL PBS per well.</li>
		<li>To stain actin fibres, thaw an ExM compatible phalloidin-solution and incubate the phalloidin (5 &#181;L of phalloidin diluted in 195 &#181;L of 1% (v/v) BSA in PBS/well) for 45 min at room temperature. Keep samples in the dark using a box.</li>
		<li>In the meantime, dissolve sodium acrylate in ddH2O using a stirring device. On ice, prepare the monomer solution (see Table 1). 
		NOTE: Dissolved sodium acrylate should be a clear and colorless solution. If the solution is yellow, replace with new sodium acrylate.</li>
		<li>Prepare the gelling solution on ice (see Table 1). Pipette ammoniumperoxidsulfate (APS) into the gelling solution right before the gelling solution is applied to the gelation chamber.</li>
		<li>Remove phalloidin from the cells and wash with 1.5 mL of PBS twice at room temperature. Leave 1.5 mL PBS within the well to facilitate removal of the sample on the glass cover slip.</li>
		<li>Place the cells on the cover glass into the gelling chamber using forceps and a cannula to lift the cover glass slip from the 6-well plate. 
		NOTE: The cells should be on the top of the glass cover slip. The glass cover slip should not touch the spacers.</li>
		<li>Add APS to the gelling solution and vortex shortly. Pipette 200 &#181;L of gelling solution on the sample (Figure 2A4-C4). Cautiously close the gelation chamber by avoiding air bubbles within the gel (Figure 2A5-C5).</li>
		<li>Incubate the gelling chamber for at least 1 h at 37 &#176;C to polymerize the gel in the wet staining chamber.</li>
	</ol>
	</li>
	<li>Homogenization (digestion)
	<ol>
		<li>Take the staining chamber out of the incubator. To open the gelation chamber lid, introduce a razor blade between the lid and the spacer. Remove the lid cautiously. Remove the spacers with the razor blade and eliminate all extra gel by cutting it with the razor blade.</li>
		<li>Put the slide with the gel and cover glass into a dish filled with PBS. By shaking gently, remove the detached cover glass from the gel. To remove the gel from the dish easily, put the slide below the gel to attach the gel to the slide.</li>
		<li>With the gel on the slide, divide the gel in small pieces (quarter of the gel is divided into two to three pieces) using the razor blade. Gently push one piece of gel into a well of a 6-well plate with glass bottom and enfold it by a paintbrush. Keep the gel moisturized with a little amount of PBS using the paintbrush to avoid dehydration of the gel. 
		NOTE: The cells face downwards.</li>
		<li>Using an inverted microscope, take overview images with low numerical aperture to determine the expansion factor after expansion.</li>
		<li>Prepare the digestion buffer (see Table 1). Dilute Proteinase K to 4 U/mL in digestion buffer to receive the digestion solution. 
		NOTE: Digestion buffer without Proteinase K can be stored at 4 &#176;C for 1-2 weeks.</li>
		<li>Add 500 &#181;L of the digestion solution to each well and immerse the gel within the solution. Let it digest overnight at room temperature and close the lid keeping the samples in the dark. 
		NOTE: Alternatively, let it digest at 37 &#176;C for 1 h.</li>
	</ol>
	</li>
	<li>Expansion
	<ol>
		<li>Remove the digestion solution with a pipette and discard it. Add 1 mL of ddH2O. Incubate the immersed gel for 10 min at room temperature.</li>
		<li>Remove the water and add 1 mL of fresh ddH2O. Wait for 10 min and continue exchanging water every 10 min until a plateau of expansion is reached. 
		NOTE: Sample expansion up to 4.5-fold is achievable. The gel becomes optically clear.</li>
	</ol>
	</li>
	<li>Imaging
	<ol>
		<li>Remove the water from the gel and directly start microscopy. Using an inverted microscope, primarily use an air objective (low magnification) to find imaged cells in the pre-expansion state (step 4.3.4).</li>
		<li>Switch to a 40x (oil/water) and 63x objective for better resolution. Excite with the wavelength of interest and take the image via the camera.</li>
	</ol>
	</li>
	<li>Validation
	<ol>
		<li>Take an overview image of the sample. Find and match the same structures within the sample that were imaged in step 4.3.4. Use the channel with the best signal-to-noise ratio for validation (Figure 3 A-B). 
		NOTE: Adjust the imaging parameters to achieve similar brightness as in the image acquired in the non-expanded state (step 4.3.4).</li>
		<li>Overlay pre- and post-expansion images by rotating and shifting them with ImageJ. Use the distance measuring tool in ImageJ to measure the distances between clearly identifiable structures. Measure at least 10 different structures. 
		NOTE: Alternatively, use a Python script to measure distances as described in14.</li>
		<li>Calculate the expansion factor by dividing post-/pre-expansion measurements.</li>
		<li>To determine distortions, take images with a higher numerical aperture (Figure 4A-B). Overlay these images and analyze them with ImageJ or as described in15. 
		NOTE: Determination of distortions should be performed routinely but not necessarily on every sample.</li>
	</ol>
	</li>
</ol>

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<li>Saleem, M. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Co-localization+of+nephrin%2C+podocin%2C+and+the+actin+cytoskeleton+-+Evidence+for+a+role+in+podocyte+foot+process+formation%5BTitle%5D)+AND+%22American+Journal+of+Pathology%22%5BJournal%5D)">Co-localization of nephrin, podocin, and the actin cytoskeleton - Evidence for a role in podocyte foot process formation.</a> American Journal of Pathology. 161 (4), 1459-1466 (2002).</li>
<li>Kestila, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Positionally+cloned+gene+for+a+novel+glomerular+protein+-+nephrin+-+is+mutated+in+congenital+nephrotic+syndrome%5BTitle%5D)+AND+%22Molecular+Cell%22%5BJournal%5D)">Positionally cloned gene for a novel glomerular protein - nephrin - is mutated in congenital nephrotic syndrome.</a> Molecular Cell. 1 (4), 575-582 (1998).</li>
<li>Huber, T. B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Podocin-mediated+recruitment+of+nephrin+into+lipid+rafts+is+required+for+efficient+nephrin+signaling%5BTitle%5D)+AND+%22Journal+of+the+American+Society+of+Nephrology%22%5BJournal%5D)">Podocin-mediated recruitment of nephrin into lipid rafts is required for efficient nephrin signaling.</a> Journal of the American Society of Nephrology. 14, 8(2003).</li>
<li>Boute, N., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((NPHS2%2C+encoding+the+glomerular+protein+podocin%2C+is+mutated+in+autosomal+recessive+steroid-resistant+nephrotic+syndrome%5BTitle%5D)+AND+%22Nature+Genetics%22%5BJournal%5D)">NPHS2, encoding the glomerular protein podocin, is mutated in autosomal recessive steroid-resistant nephrotic syndrome.</a> Nature Genetics. 24 (4), 349-354 (2000).</li>
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<li>Willig, K. I., Rizzoli, S. O., Westphal, V., Jahn, R., Hell, S. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((STED+microscopy+reveals+that+synaptotagmin+remains+clustered+after+synaptic+vesicle+exocytosis%5BTitle%5D)+AND+%22Nature%22%5BJournal%5D)">STED microscopy reveals that synaptotagmin remains clustered after synaptic vesicle exocytosis.</a> Nature. 440 (7086), 935-939 (2006).</li>
<li>van de Linde, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Direct+stochastic+optical+reconstruction+microscopy+with+standard+fluorescent+probes%5BTitle%5D)+AND+%22Nature+Protocols%22%5BJournal%5D)">Direct stochastic optical reconstruction microscopy with standard fluorescent probes.</a> Nature Protocols. 6 (7), 991-1009 (2011).</li>
<li>Rust, M. J., Bates, M., Zhuang, X. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Sub-diffraction-limit+imaging+by+stochastic+optical+reconstruction+microscopy+%28STORM%29%5BTitle%5D)+AND+%22Nature+Methods%22%5BJournal%5D)">Sub-diffraction-limit imaging by stochastic optical reconstruction microscopy (STORM).</a> Nature Methods. 3 (10), 793-795 (2006).</li>
<li>Testa, I., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Multicolor+fluorescence+nanoscopy+in+fixed+and+living+cells+by+exciting+conventional+fluorophores+with+a+single+wavelength%5BTitle%5D)+AND+%22Biophysical+Journal%22%5BJournal%5D)">Multicolor fluorescence nanoscopy in fixed and living cells by exciting conventional fluorophores with a single wavelength.</a> Biophysical Journal. 99 (8), 2686-2694 (2010).</li>
<li>Wassie, A. T., Zhao, Y., Boyden, E. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Expansion+microscopy%3A+principles+and+uses+in+biological+research%5BTitle%5D)+AND+%22Nature+Methods%22%5BJournal%5D)">Expansion microscopy: principles and uses in biological research.</a> Nature Methods. 16 (1), 33-41 (2019).</li>
<li>Tillberg, P. W., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Protein-retention+expansion+microscopy+of+cells+and+tissues+labeled+using+standard+fluorescent+proteins+and+antibodies%5BTitle%5D)+AND+%22Nature+Biotechnology%22%5BJournal%5D)">Protein-retention expansion microscopy of cells and tissues labeled using standard fluorescent proteins and antibodies.</a> Nature Biotechnology. 34 (9), 987-992 (2016).</li>
<li>Truckenbrodt, S., Sommer, C., Rizzoli, S. O., Danzl, J. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+practical+guide+to+optimization+in+X10+expansion+microscopy%5BTitle%5D)+AND+%22Nature+Protocols%22%5BJournal%5D)">A practical guide to optimization in X10 expansion microscopy.</a> Nature Protocols. 14 (3), 832-863 (2019).</li>
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<li>Chang, J. B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Iterative+expansion+microscopy%5BTitle%5D)+AND+%22Nature+Methods%22%5BJournal%5D)">Iterative expansion microscopy.</a> Nature Methods. 14 (6), 593-599 (2017).</li>
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</ol>

The authors have nothing to disclose.

The presented method enables the investigator to visualize cellular proteins, e.g., podocin, nephrin, and cytoskeletal components, e.g., F-actin. Within this protocol, transfected cos7 cells are used as a model to study interaction of slit diaphragm proteins with F-actin. Unfortunately, immortalized podocyte cell lines do not express sufficient endogenous amounts of slit diaphragm proteins19.
With this method, cellular proteins can be visualized with nanoscale resolutio...

Albuminuria is a surrogate parameter of cardiovascular risk and results from disruption of the glomerular filter1. The glomerular filter is composed of the fenestrated endothelium, the glomerular basement membrane and the slit diaphragm formed by podocytes. Primary and secondary foot processes of podocytes wrap around the capillary wall of the glomerulum2. The delicate structure of foot processes is maintained by cortical actin bundles which also serve as anchors for multiple slit diaphragm proteins and other adaptor proteins2. The slit diaphragm&#39;s backbone protein is called nephrin and interacts in a homophilic manner with nephrin molecules of opposing podocytes. Via diverse adaptor proteins, nephrin is linked to the actin cytoskeleton2,3. Mutations in the nephrin-encoding gene NPHS1 lead to nephrotic syndrome of the Finnish type4.
One of nephrin&#39;s interacting proteins is podocin, a hairpin-like protein of the stomatin family3. Podocin recruits nephrin to lipid rafts and links it to the actin cytoskeleton5. Podocin is encoded by the NPHS2 gene. Mutations in NPHS2 lead to steroid-resistant nephrotic syndrome6.
To visualize and co-localize actin adaptor proteins, immunofluorescence techniques may be used. Unfortunately, the diffraction barrier of the light limits the resolution of conventional fluorescence microscopes to 200-350 nm7. Novel microscopy techniques, e.g., stimulated emission depletion (STED)8, photo-activated localization microscopy (PALM)9, stochastic optical reconstruction microscopy (STORM or dSTORM) or ground state deletion microscopy followed by individual molecule return (GSDIM)9,10,11, enable a resolution up to approximately 10 nm. However, these super resolution techniques require highly expensive microscopes, well-trained personnel and are therefore not available in many laboratories.
Expansion microscopy (ExM) is a novel and simple technique that enables super resolution imaging with conventional microscopes and is potentially available to a large research community12. In protein retention expansion microscopy (proExM), the sample of interest (cells or tissue) is fixed and stained with fluorophores13. Proteins within the sample are then covalently anchored by a small molecule (6-((Acryloyl)amino)hexanoic acid, succinimidyl ester, AcX) into a swellable hydrogel13. Through enzymatic digestion with proteinase K (ProK), proteins and fluorophores maintain their relative position within the gel after expansion13. After swelling of the gel, the sample expands up to 4.5-fold (90-fold volumetric expansion) leading to an effective lateral resolution of approximately 60-70 nm (300 nm/4.5). Modifications of this technique can even allow for a 10-fold expansion (1,000-fold volumetric expansion), rendering a resolution of 20-30 nm on conventional microscopes14,15,16.
Glomerular structures of mouse and human kidneys have been visualized via ExM17. Within this paper, we present a detailed proExM protocol to visualize super resolution images of F-actin and the actin-adapter protein podocin within cells using a conventional fluorescence microscope.

<table><tbody><tr><td>Acrylamide &gt;99%</td><td>Sigma-Aldrich</td><td>A3553-100G</td><td></td></tr><tr><td>6-((Acryloyl)amino)hexanoic acid, succinimidyl ester, Acryloyl-X, SE</td><td>invitrogen</td><td>A-20770</td><td>store up to 4 months</td></tr><tr><td>APS</td><td>Sigma-Aldrich</td><td>A3678-25G</td><td></td></tr><tr><td>Deckgl&amp;auml;ser (cover glasses)</td><td>Engelbrecht</td><td>K12432</td><td>24x32mm #1.0</td></tr><tr><td>Diamont cutter</td><td>VWR</td><td>201-0392</td><td>for cutting the cover slips</td></tr><tr><td>Guanidine HCl</td><td>Sigma-Aldrich</td><td>G3272-100G</td><td>8M Stock can be kept at RT</td></tr><tr><td>Marten hair paintbrush</td><td>Leon Hardy</td><td>3 (770)</td><td></td></tr><tr><td>&quot;Menzel&quot; Deckgl&amp;auml;ser (cover glasses)</td><td>Thermo Fischer&amp;nbsp;</td><td>15654786</td><td>24x24mm #1.5</td></tr><tr><td>N,N`-Methylenbisacrylamide</td><td>Sigma-Aldrich</td><td>M7256-25G</td><td></td></tr><tr><td>Objekttr&amp;auml;ger UniMark</td><td>Marienfeld</td><td>703010</td><td></td></tr><tr><td>Proteinase K</td><td>New England Biolabs</td><td>P8107S</td><td></td></tr><tr><td>Sodium Acrylate</td><td>Sigma-Aldrich</td><td>408220</td><td>check purity&amp;nbsp;</td></tr><tr><td>Sodium Bicarbonate</td><td>Sigma-Aldrich</td><td>S5761</td><td></td></tr><tr><td>Staining chamber</td><td></td><td></td><td>produced at the university&#039;s workshop</td></tr><tr><td>TEMED</td><td>ROTH</td><td>2367.1</td><td></td></tr><tr><td>6-Well glass bottom plates</td><td>Cellvis</td><td>P06-1.5H-N</td><td></td></tr><tr><td>&lt;strong&gt;Antibodies&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Actin-ExM 546&amp;nbsp;</td><td>chrometra</td><td>non-available</td><td>1:40</td></tr><tr><td>Anti Podocin produced in rabbit</td><td>Sigma</td><td>P-0372-200UL</td><td>1:200</td></tr><tr><td>Donkey anti guinea-pig CF633</td><td>Sigma</td><td>SAB4600129-50UL</td><td>1:200</td></tr><tr><td>Goat anti rabbit 488</td><td>Life Technologies</td><td>A11034</td><td>1:1000</td></tr><tr><td>Guinea pig anti nephrin</td><td>Origene</td><td>BP5030</td><td>1:100</td></tr><tr><td>&lt;strong&gt;Software&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>FIJI</td><td></td><td></td><td></td></tr><tr><td>Visiview</td><td></td><td></td><td></td></tr><tr><td>&lt;strong&gt;microscope&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>AXIO Observer Z1</td><td>Zeiss</td><td>non-available</td><td></td></tr></tbody></table>

imaging of podocytic proteins nephrin, actin, and podocin with expansion microscopy

Disruption of the glomerular filter composed of the glomerular endothelium, glomerular basement membrane and podocytes, results in albuminuria. Podocyte foot processes contain actin bundles that bind to cytoskeletal adaptor proteins such as podocin. Those adaptor proteins, such as podocin, link the backbone of the glomerular slit diaphragm, such as nephrin, to the actin cytoskeleton. Studying the localization and function of these and other podocytic proteins is essential for the understanding of the glomerular filter&#39;s role in health and disease. The presented protocol enables the user to visualize actin, podocin, and nephrin in cells with super resolution imaging on a conventional microscope. First, cells are stained with a conventional immunofluorescence technique. All proteins within the sample are then covalently anchored to a swellable hydrogel. Through digestion with proteinase K, structural proteins are cleaved allowing isotropical swelling of the gel in the last step. Dialysis of the sample in water results in a 4-4.5-fold expansion of the sample and the sample can be imaged via a conventional fluorescence microscope, rendering a potential resolution of 70 nm.

The concept and timing of this proExM protocol is depicted in Figure 1. On day 5, transfected cells are fixed and stained with fluorescent antibodies targeting the protein of interest (Figure 1A,B). On day 6, treatment with AcX leads to formation of amine groups on all proteins (including fluorophores) (Figure 1A,B)12. Upon polymerization of t...

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Imaging of Podocytic Proteins Nephrin, Actin, and Podocin with Expansion Microscopy

The presented method enables visualization of fluorescently labeled cellular proteins with expansion microscopy leading to a resolution of 70 nm on a conventional microscope.

Disruption of the glomerular filter composed of the glomerular endothelium, glomerular basement membrane and podocytes, results in albuminuria. Podocyte ...

imaging-podocytic-proteins-nephrin-actin-podocin-with-expansion

Research

JoVE Journal

Biochemistry

6.8K Views.  Heinrich-Heine-University. Dit microscopische expansieprotocol stelt de onderzoeker in staat om glomerulaire eiwitten met nanoschaalresolutie te visualiseren op een conventionele microscoop. Het belangrijkste voordeel van expansiemicroscopie is dat het toegankelijk is voor een grote onderzoeksgemeenschap. Begin met het maken van afstandhouders voor de gelatiekamer.Gesneden nummer één en nummer 1.5 glazen deksel glijdt in vijf millimeter strepen, met behulp van een diamantmes. Plaats een glazen schuif in een kl...

Video: Imaging of Podocytic Proteins Nephrin, Actin, and Podocin with Expansion Microscopy

Isolation of Glomeruli and In Vivo Labeling of Glomerular Cell Surface Proteins

Proteinuria results from the disruption of the glomerular filter that is composed of the fenestrated endothelium, glomerular basement membrane, and podocytes with their slit diaphragms. The delicate structure of the glomerular filter, especially the slit diaphragm, relies on the interplay of diverse cell surface proteins. Studying these cell surface proteins has so far been limited to in vitro studies or histologic analysis. Here, we present a murine in vivo biotinylation labeling method, which enables the study of glomerular cell surface proteins under physiologic and pathophysiologic conditions. This protocol contains information on how to perfuse mouse kidneys, isolate glomeruli, and perform endogenous immunoprecipitation of a protein of interest. Semi-quantitation of glomerular cell surface abundance is readily available with this novel method, and all proteins accessible to biotin perfusion and immunoprecipitation can be studied. In addition, isolation of glomeruli with or without biotinylation enables further analysis of glomerular RNA and protein as well as primary glomerular cell culture (i.e., primary podocyte cell culture).

Isolation of Glomeruli and In Vivo Labeling of Glomerular Cell Surface Proteins

Here we present a protocol for murine in vivo labeling of glomerular cell surface proteins with biotin. This protocol contains information on how to perfuse mouse kidneys, isolate glomeruli, and perform endogenous immunoprecipitation of the protein of interest.

Proteinuria results from the disruption of the glomerular filter that is composed of the fenestrated endothelium, glomerular basement membrane, and ...

Long-term Intravital Immunofluorescence Imaging of Tissue Matrix Components with Epifluorescence and Two-photon Microscopy

Besides being a physical scaffold to maintain tissue morphology, the extracellular matrix (ECM) is actively involved in regulating cell and tissue function during development and organ homeostasis. It does so by acting via biochemical, biomechanical, and biophysical signaling pathways, such as through the release of bioactive ECM protein fragments, regulating tissue tension, and providing pathways for cell migration. The extracellular matrix of the tumor microenvironment undergoes substantial remodeling, characterized by the degradation, deposition and organization of fibrillar and non-fibrillar matrix proteins. Stromal stiffening of the tumor microenvironment can promote tumor growth and invasion, and cause remodeling of blood and lymphatic vessels. Live imaging of matrix proteins, however, to this point is limited to fibrillar collagens that can be detected by second harmonic generation using multi-photon microscopy, leaving the majority of matrix components largely invisible. Here we describe procedures for tumor inoculation in the thin dorsal ear skin, immunolabeling of extracellular matrix proteins and intravital imaging of the exposed tissue in live mice using epifluorescence and two-photon microscopy. Our intravital imaging method allows for the direct detection of both fibrillar and non-fibrillar matrix proteins in the context of a growing dermal tumor. We show examples of vessel remodeling caused by local matrix contraction. We also found that fibrillar matrix of the tumor detected with the second harmonic generation is spatially distinct from newly deposited matrix components such as tenascin C. We also showed long-term (12 hours) imaging of T-cell interaction with tumor cells and tumor cells migration along the collagen IV of basement membrane. Taken together, this method uniquely allows for the simultaneous detection of tumor cells, their physical microenvironment and the endogenous tissue immune response over time, which may provide important insights into the mechanisms underlying tumor progression and ultimate success or resistance to therapy.

The extracellular matrix undergoes substantial remodeling during wound healing, inflammation and tumorigenesis. We present a novel intravital immunofluorescence microscopy approach to visualize the dynamics of fibrillar as well as mesh-like matrix components with high spatial and temporal resolution using epifluorescence or two-photon microscopy.

Besides being a physical scaffold to maintain tissue morphology, the extracellular matrix (ECM) is actively involved in regulating cell and tissue ...

Bioengineering

Using Expansion Microscopy to Physically Enlarge Whole-Mount Drosophila Embryos for Super-Resolution Imaging

The workhorse of developmental biology is the confocal microscope, which allows researchers to determine the three-dimensional localization of tagged molecules within complex biological samples. While traditional confocal microscopes allow one to resolve two adjacent fluorescent point sources located a few hundred nanometers apart, observing the finer details of subcellular biology requires the ability to resolve signals in the order of tens of nanometers. Numerous hardware-based methods for super-resolution microscopy have been developed to allow researchers to sidestep such resolution limits, although these methods require specialized microscopes that are not available to all researchers. An alternative method for increasing resolving power is to isotropically enlarge the sample itself through a process known as expansion microscopy (ExM), which was first described by the Boyden group in 2015. ExM is not a type of microscopy per se but is rather a method for swelling a sample while preserving the relative spatial organization of its constituent molecules. The expanded sample can then be observed at an effectively increased resolution using a traditional confocal microscope. Here, we describe a protocol for implementing ExM in whole-mount Drosophila embryos, which is used to examine the localization of Par-3, myosin II, and mitochondria within the surface epithelial cells. This protocol yields an approximately four-fold increase in sample size, allowing for the detection of subcellular details that are not visible with conventional confocal microscopy. As proof of principle, an anti-GFP antibody is used to distinguish distinct pools of myosin-GFP between adjacent cell cortices, and fluorescently labeled streptavidin is used to detect endogenous biotinylated molecules to reveal the fine details of the mitochondrial network architecture. This protocol utilizes common antibodies and reagents for fluorescence labeling, and it should be compatible with many existing immunofluorescence protocols.

Using Expansion Microscopy to Physically Enlarge Whole-Mount Drosophila Embryos for Super-Resolution Imaging

Here, a protocol for the implementation of expansion microscopy in early Drosophila embryos to achieve super-resolution imaging using a conventional laser-scanning confocal microscope is presented.

The workhorse of developmental biology is the confocal microscope, which allows researchers to determine the three-dimensional localization of tagged ...

Developmental Biology

Optical Clearing and Imaging of Immunolabeled Kidney Tissue

Optical clearing techniques render tissue transparent by equilibrating the refractive index throughout a sample for subsequent three-dimensional (3-D) imaging. They have received great attention in all research areas for the potential to analyze microscopic multicellular structures that extend over macroscopic distances. Given that kidney tubules, vasculature, nerves, and glomeruli extend in many directions, which have been only partially captured by traditional two-dimensional techniques so far, tissue clearing also opened up many new areas of kidney research. The list of optical clearing methods is rapidly growing, but it remains difficult for beginners in this field to choose the best method for a given research question. Provided here is a simple method that combines antibody labeling of thick mouse kidney slices; optical clearing with cheap, non-toxic and ready-to-use chemical ethyl cinnamate; and confocal imaging. This protocol describes how to perfuse kidneys and use an antigen-retrieval step to increase antibody- binding without requiring specialized equipment. Its application is presented in imaging different multicellular structures within the kidney, and how to troubleshoot poor antibody penetration into tissue is addressed. We also discuss the potential difficulties of imaging endogenous fluorophores and acquiring very large samples and how to overcome them. This simple protocol provides an easy-to-setup and comprehensive tool to study tissue in three dimensions.

The combination of antibody labeling, optical clearing, and advanced light microscopy allows three-dimensional analysis of complete structures or organs. Described here is a simple method to combine immunolabeling of thick kidney slices, optical clearing with ethyl cinnamate, and confocal imaging that enables visualization and quantification of three-dimensional kidney structures.

Optical clearing techniques render tissue transparent by equilibrating the refractive index throughout a sample for subsequent three-dimensional (3-D) ...

Medicine

Nanoscopic Imaging of Human Tissue Sections via Physical and Isotropic Expansion

In modern pathology, optical microscopy plays an important role in disease diagnosis by revealing microscopic structures of clinical specimens. However, the fundamental physical diffraction limit prevents interrogation of nanoscale anatomy and subtle pathological changes when using conventional optical imaging approaches. Here, we describe a simple and inexpensive protocol, called expansion pathology (ExPath), for nanoscale optical imaging of common types of clinical primary tissue specimens, including both fixed-frozen or formalin-fixed paraffin embedded (FFPE) tissue sections. This method circumvents the optical diffraction limit by chemically transforming the tissue samples into tissue-hydrogel hybrid and physically expanding them isotropically across multiple scales in pure water. Due to expansion, previously unresolvable molecules are separated and thus can be observed using a conventional optical microscope.

Nanoscale imaging of clinical tissue samples can improve understanding of disease pathogenesis. Expansion pathology (ExPath) is a version of expansion microscopy (ExM), modified for compatibility with standard clinical tissue samples, to explore the nanoscale configuration of biomolecules using conventional diffraction limited microscopes.

In modern pathology, optical microscopy plays an important role in disease diagnosis by revealing microscopic structures of clinical specimens. However, ...

Single-channel Analysis and Calcium Imaging in the Podocytes of the Freshly Isolated Glomeruli

Podocytes (renal glomerular epithelial cells) are known to regulate glomerular permeability and maintain glomerular structure; a key role for these cells in the pathogenesis of various renal diseases has been established since podocyte injury leads to proteinuria and foot process effacement. It was previously reported that various endogenous agents may cause a dramatic overload in intracellular Ca2+ concentration in podocytes, presumably leading to albuminuria, and this likely occurs via calcium-conducting ion channels. Therefore, it appeared important to study calcium handling in the podocytes both under normal conditions and in various pathological states. However, available experimental approaches have remained somewhat limited to cultured and transfected cells. Although they represent a good basic model for such studies, they are essentially extracted from the native environment of the glomerulus. Here we describe the methodology of studying podocytes as a part of the freshly isolated whole glomerulus. This preparation retains the functional potential of the podocytes, which are still attached to the capillaries; therefore, podocytes remain in the environment that conserves the major parts of the glomeruli filtration apparatus. The present manuscript elaborates on two experimental approaches that allow 1) real-time detection of calcium concentration changes with the help of ratiometric confocal fluorescence microscopy, and 2) the recording of the single ion channels activity in the podocytes of the freshly isolated glomeruli. These methodologies utilize the advantages of the native environment of the glomerulus that enable researchers to resolve acute changes in the intracellular calcium handling in response to applications of various agents, measure basal concentration of calcium within the cells (for instance, to evaluate disease progression), and assess and manipulate calcium conductance at the level of single ion channels.

Changes in the intracellular calcium levels in the podocytes are one of the most important means to control the filtration function of glomeruli. Here we explain a high-throughput approach that allows detection of real-time calcium handling and single ion channels activity in the podocytes of the freshly isolated glomeruli.

Podocytes (renal glomerular epithelial cells) are known to regulate glomerular permeability and maintain glomerular structure; a key role for these cells ...

Biology

Ultrastructural Expansion Microscopy in In Vitro Life Cycle of Trypanosoma cruzi

Ultrastructural Expansion Microscopy in Three In Vitro Life Cycle Stages of Trypanosoma cruzi

We describe here the application of ultrastructure expansion microscopy (U-ExM) in Trypanosoma cruzi, a technique that allows increasing the spatial resolution of a cell or tissue for microscopic imaging. This is performed by physically expanding a sample with off-the-shelf chemicals and common lab equipment.
Chagas disease is a widespread and pressing public health concern caused by T. cruzi. The disease is prevalent in Latin America and has become a significant problem in non-endemic regions due to increased migration. The transmission of T. cruzi occurs through hematophagous insect vectors belonging to the Reduviidae and Hemiptera families. Following infection, T. cruzi amastigotes multiply within the mammalian host and differentiate into trypomastigotes, the non-replicative bloodstream form. In the insect vector, trypomastigotes transform into epimastigotes and proliferate through binary fission.The differentiation between the life cycle stages requires an extensive rearrangement of the cytoskeleton and can be recreated in the lab completely using different cell culture techniques. 
We describe here a detailed protocol for the application of U-ExM in three in vitro life cycle stages of Trypanosoma cruzi, focusing on optimization of the immunolocalization of cytoskeletal proteins. We also optimized the use of N-Hydroxysuccinimide ester (NHS), a pan-proteome label that has enabled us to mark different parasite structures.

This study shows a detailed protocol to perform ultrastructure expansion microscopy in three in vitro life cycle stages of Trypanosoma cruzi, the pathogen responsible for Chagas disease. We include the optimized technique for cytoskeletal proteins and pan-proteome labeling.

We describe here the application of ultrastructure expansion microscopy (U-ExM) in Trypanosoma cruzi, a technique that allows increasing the spatial ...

Expansion Microscopy for Robust 11-Fold Expansion

Universal Molecular Retention with 11-Fold Expansion Microscopy

The nanoscale imaging of biological specimens can improve the understanding of disease pathogenesis. In recent years, expansion microscopy (ExM) has been demonstrated to be an effective and low-cost alternative to optical super-resolution microscopy. However, it has been limited by the need for specific and often custom anchoring agents to retain different biomolecule classes within the gel and by difficulties with expanding standard clinical sample formats, such as formalin-fixed paraffin-embedded tissue, especially if larger expansion factors or preserved protein epitopes are desired. Here, we describe Magnify, a new ExM method for robust expansion up to 11-fold in a wide array of tissue types. By using methacrolein as the chemical anchor between the tissue and gel, Magnify retains multiple biomolecules, such as proteins, lipids, and nucleic acids, within the gel, thus allowing the broad nanoscale imaging of tissues on conventional optical microscopes. This protocol describes best practices to ensure robust and crack-free tissue expansion, as well as tips for handling and imaging highly expanded gels.

Author Spotlight: Universal Molecular Retention with 11-Fold Expansion Microscopy  

Presented here is a new version of expansion microscopy (ExM), Magnify, that is modified for up to 11-fold expansion, conserving a comprehensive array of biomolecule classes, and is compatible with a broad range of tissue types. It enables the interrogation of the nanoscale configuration of biomolecules using conventional diffraction-limited microscopes.

The nanoscale imaging of biological specimens can improve the understanding of disease pathogenesis. In recent years, expansion microscopy (ExM) has been ...

Preparation and Imaging of Murine Ocular Lens Whole Mounts

Whole Mount Imaging to Visualize and Quantify Peripheral Lens Structure, Cell Morphology, and Organization

The ocular lens is a transparent flexible tissue that alters its shape to focus light from different distances onto the retina. Aside from a basement membrane surrounding the organ, called the capsule, the lens is entirely cellular consisting of a monolayer of epithelial cells on the anterior hemisphere and a bulk mass of lens fiber cells. Throughout life, epithelial cells proliferate in the germinative zone at the lens equator, and equatorial epithelial cells migrate, elongate, and differentiate into newly formed fiber cells. Equatorial epithelial cells substantially alter morphology from randomly packed cobble-stone-shaped cells into aligned hexagon-shaped cells forming meridional rows. Newly formed lens fiber cells retain the hexagonal cell shape and elongate toward the anterior and posterior poles, forming a new shell of cells that are overlaid onto previous generations of fibers. Little is known about the mechanisms that drive the remarkable morphogenesis of lens epithelial cells to fiber cells. To better understand lens structure, development, and function, new imaging protocols have been developed to image peripheral structures using whole mounts of ocular lenses. Here, methods to quantify capsule thickness, epithelial cell area, cell nuclear area and shape, meridional row cell order and packing, and fiber cell widths are shown. These measurements are essential for elucidating the cellular changes that occur during lifelong lens growth and understanding the changes that occur with age or pathology.

Author Spotlight: In-Depth Morphometric Examination and Quantification of Native Lens Structure Using Whole Mount Imaging 

The present protocols describe novel whole mount imaging for the visualization of peripheral structures in the ocular lens with methods for image quantification. These protocols can be used in studies to better understand the relationship between lens microscale structures and lens development/function.

The ocular lens is a transparent flexible tissue that alters its shape to focus light from different distances onto the retina. Aside from a basement ...

Expansion Microscopy for Visualizing Intracellular Sialylated Glycans via MOE

Visualizing Intracellular Sialylation with Click Chemistry and Expansion Microscopy

Metabolic labeling techniques allow the incorporation of bioorthogonal reporters into glycans, enabling the targeted bioconjugation of molecular dyes within cells through click and bioorthogonal chemistry. Metabolic oligosaccharide engineering (MOE) has attracted considerable interest due to the essential role of glycosylation in numerous biological processes that involve molecular recognition and its impact on pathologies ranging from cancer to genetic disorders to viral and bacterial infections.
Although MOE is better known for the detection of cell surface glycoconjugates, it is also a very important methodology for the study of intracellular glycans in physiological and pathological contexts. Such studies greatly benefit from high spatial resolution. However, super-resolution microscopy is not readily available in most laboratories and poses challenges for daily implementation. Expansion microscopy is a recent alternative that enhances the resolution of microscopy by physically enlarging biological specimens labeled with fluorescent markers. By embedding the sample in a swellable gel and causing it to expand isotropically through chemical treatment, subcellular structures can be visualized with enhanced precision and resolution without the need for super-resolution techniques.
In this work, we illustrate the capacity of expansion microscopy to visualize intracellular sialylated glycans through the combined use of MOE and click chemistry. Specifically, we propose a procedure for bioorthogonal labeling and expansion microscopy that employs a reporter targeting sialylation, which may be associated with immunofluorescence for co-localization studies. This protocol enables localization studies of sialoconjugate biosynthesis, intracellular trafficking, and recycling.

Here, we propose a simple protocol combining metabolic oligosaccharide engineering, click chemistry, and expansion microscopy that allows bioimaging of intracellular sialylated N-glycoproteins with improved resolution using routine microscopy equipment.

Metabolic labeling techniques allow the incorporation of bioorthogonal reporters into glycans, enabling the targeted bioconjugation of molecular dyes ...

Imaging of Podocytic Proteins Nephrin, Actin, and Podocin with Expansion Microscopy

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