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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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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Acrylamide &#62;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&#228;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>&#34;Menzel&#34; Deckgl&#228;ser (cover glasses)</td>
			<td>Thermo Fischer&#160;</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&#228;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&#160;</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&#39;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>Antibodies</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Actin-ExM 546&#160;</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>Software</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>microscope</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...

Watch this Scientific Journal Video about Imaging of Podocytic Proteins Nephrin, Actin, and Podocin with Expansion Microscopy at JoVE.com

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

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

Method to Visualize and Analyze Membrane Interacting Proteins by Transmission Electron Microscopy

Monotopic proteins exert their function when attached to a membrane surface, and such interactions depend on the specific lipid composition and on the availability of enough area to perform the function. Nanodiscs are used to provide a membrane surface of controlled size and lipid content. In the absence of bound extrinsic proteins, sodium phosphotungstate-stained nanodiscs appear as stacks of coins when viewed from the side by transmission electron microscopy (TEM). This protocol is therefore designed to intentionally promote stacking; consequently, the prevention of stacking can be interpreted as the binding of the membrane-binding protein to the nanodisc. In a further step, the TEM images of the protein-nanodisc complexes can be processed with standard single-particle methods to yield low-resolution structures as a basis for higher resolution cryoEM work. Furthermore, the nanodiscs provide samples suitable for either TEM or non-denaturing gel electrophoresis. To illustrate the method, Ca2+-induced binding of 5-lipoxygenase on nanodiscs is presented.

Many proteins perform their function when attached to membrane surfaces. The binding of extrinsic proteins on nanodisc membranes can be indirectly imaged by transmission electron microscopy. We show that the characteristic stacking (rouleau) of nanodiscs induced by the negative stain sodium phosphotungstate is prevented by the binding of extrinsic protein.

Monotopic proteins exert their function when attached to a membrane surface, and such interactions depend on the specific lipid composition and on the ...

Imaging Amyloid Tissues Stained with Luminescent Conjugated Oligothiophenes by Hyperspectral Confocal Microscopy and Fluorescence Lifetime Imaging

Proteins that deposit as amyloid in tissues throughout the body can be the cause or consequence of a large number of diseases. Among these we find neurodegenerative diseases such as Alzheimer&#39;s and Parkinson&#39;s disease afflicting primarily the central nervous system, and systemic amyloidosis where serum amyloid A, transthyretin and IgG light chains deposit as amyloid in liver, carpal tunnel, spleen, kidney, heart, and other peripheral tissues. Amyloid has been known and studied for more than a century, often using amyloid specific dyes such as Congo red and Thioflavin T (ThT) or Thioflavin (ThS). In this paper, we present heptamer-formyl thiophene acetic acid (hFTAA) as an example of recently developed complements to these dyes called luminescent conjugated oligothiophenes (LCOs). hFTAA is easy to use and is compatible with co-staining in immunofluorescence or with other cellular markers. Extensive research has proven that hFTAA detects a wider range of disease associated protein aggregates than conventional amyloid dyes. In addition, hFTAA can also be applied for optical assignment of distinct aggregated morphotypes to allow studies of amyloid fibril polymorphism. While the imaging methodology applied is optional, we here demonstrate hyperspectral imaging (HIS), laser scanning confocal microscopy and fluorescence lifetime imaging (FLIM). These examples show some of the imaging techniques where LCOs can be used as tools to gain more detailed knowledge of the formation and structural properties of amyloids. An important limitation to the technique is, as for all conventional optical microscopy techniques, the requirement for microscopic size of aggregates to allow detection. Furthermore, the aggregate should comprise a repetitive &#946;-sheet structure to allow for hFTAA binding. Excessive fixation and/or epitope exposure that modify the aggregate structure or conformation can render poor hFTAA binding and hence pose limitations to accurate imaging.

Amyloid deposition is a hallmark of different diseases and afflicts many different organs. This paper describes the application of luminescent conjugated oligothiophene fluorescence staining in combination with fluorescence microscopy techniques. This staining method represents a powerful tool for detection and exploration of protein aggregates in both clinical and scientific setups.

Proteins that deposit as amyloid in tissues throughout the body can be the cause or consequence of a large number of diseases. Among these we find ...

Growing Protein Crystals with Distinct Dimensions Using Automated Crystallization Coupled with In Situ Dynamic Light Scattering

The automated crystallization device is a patented technique1 especially developed for monitoring protein crystallization experiments with the aim to precisely maneuver the nucleation and crystal growth towards desired sizes of protein crystals. The controlled crystallization is based on sample investigation with in situ Dynamic Light Scattering (DLS) while all visual changes in the droplet are monitored online with the help of a microscope coupled to a CCD camera, thus enabling a full investigation of the protein droplet during all stages of crystallization. The use of in situ DLS measurements throughout the entire experiment allows a precise identification of the highly supersaturated protein solution transitioning to a new phase &#8211; the formation of crystal nuclei. By identifying the protein nucleation stage, the crystallization can be optimized from large protein crystals to the production of protein microcrystals. The experimental protocol shows an interactive crystallization approach based on precise automated steps such as precipitant addition, water evaporation for inducing high supersaturation, and sample dilution for slowing induced homogeneous nucleation or reversing phase transitions.

Growing Protein Crystals with Distinct Dimensions Using Automated Crystallization Coupled with In Situ Dynamic Light Scattering

Here we present a protocol for controlled production of protein microcrystals. The process uses an automated device allowing controlled manipulation of several crystallization parameters. The protein crystallization is carried-out by controlled and automated addition of crystallization solutions while monitoring and investigating the radius distribution of particles in the crystallization droplet.

The automated crystallization device is a patented technique1 especially developed for monitoring protein crystallization experiments with the aim to ...

Probing The Structure And Dynamics Of Nucleosomes Using Atomic Force Microscopy Imaging

Chromatin, which is a long chain of nucleosome subunits, is a dynamic system that allows for such critical processes as DNA replication and transcription to take place in eukaryotic cells. The dynamics of nucleosomes provides access to the DNA by replication and transcription machineries, and critically contributes to the molecular mechanisms underlying chromatin functions. Single-molecule studies such as atomic force microscopy (AFM) imaging have contributed significantly to our current understanding of the role of nucleosome structure and dynamics. The current protocol describes the steps enabling high-resolution AFM imaging techniques to study the structural and dynamic properties of nucleosomes. The protocol is illustrated by AFM data obtained for the centromere nucleosomes in which H3 histone is replaced with its counterpart centromere protein A (CENP-A). The protocol starts with the assembly of mono-nucleosomes using a continuous dilution method. The preparation of the mica substrate functionalized with aminopropyl silatrane (APS-mica) that is used for the nucleosome imaging is critical for the AFM visualization of nucleosomes described and the procedure to prepare the substrate is provided. Nucleosomes deposited on the APS-mica surface are first imaged using static AFM, which captures a snapshot of the nucleosome population. From analyses of these images, such parameters as the size of DNA wrapped around the nucleosomes can be measured and this process is also detailed. The time-lapse AFM imaging procedure in the liquid is described for the high-speed time-lapse AFM that can capture several frames of nucleosome dynamics per second. Finally, the analysis of nucleosome dynamics enabling the quantitative characterization of the dynamic processes is described and illustrated.

Here, we present a protocol to characterize nucleosome particles at the single-molecule level using static and time-lapse atomic force microscopy (AFM) imaging techniques. The surface functionalization method described allows for the capture of the structure and dynamics of nucleosomes in high-resolution at the nanoscale.

Chromatin, which is a long chain of nucleosome subunits, is a dynamic system that allows for such critical processes as DNA replication and transcription ...

Imaging of Extracellular Vesicles by Atomic Force Microscopy

Exosomes and other extracellular vesicles (EVs) are molecular complexes consisting of a lipid membrane vesicle, its surface decoration by membrane proteins and other molecules, and diverse luminal content inherited from a parent cell, which includes RNAs, proteins, and DNAs. The characterization of the hydrodynamic sizes of EVs, which depends on the size of the vesicle and its coronal layer formed by surface decorations, has become routine. For exosomes, the smallest of EVs, the relative difference between the hydrodynamic and vesicles sizes is significant. The characterization of vesicles sizes by the cryogenic transmission electron microscopy (cryo-TEM) imaging, a gold standard technique, remains a challenge due to the cost of the instrument, the expertise required to perform the sample preparation, imaging and data analysis, and a small number of particles often observed in images. A widely available and accessible alternative is the atomic force microscopy (AFM), which can produce versatile data on three-dimensional geometry, size, and other biophysical properties of extracellular vesicles. The developed protocol guides the users in utilizing this analytical tool and outlines the workflow for the analysis of EVs by the AFM, which includes the sample preparation for imaging EVs in hydrated or desiccated form, the electrostatic immobilization of vesicles on a substrate, data acquisition, its analysis, and interpretation. The representative results demonstrate that the fixation of EVs on the modified mica surface is predictable, customizable, and allows the user to obtain sizing results for a large number of vesicles. The vesicle sizing based on the AFM data was found to be consistent with the cryo-TEM imaging.&#160;

A step-by-step procedure is described for label-free immobilization of exosomes and extracellular vesicles from liquid samples and their imaging by atomic force microscopy (AFM). The AFM images are used to estimate the size of the vesicles in the solution and characterize other biophysical properties.&#160;

Exosomes and other extracellular vesicles (EVs) are molecular complexes consisting of a lipid membrane vesicle, its surface decoration by membrane ...

Mitochondria and Endoplasmic Reticulum Imaging by Correlative Light and Volume Electron Microscopy

Cellular organelles, such as mitochondria and endoplasmic reticulum (ER), create a network to perform a variety of functions. These highly curved structures are folded into various shapes to form a dynamic network depending on the cellular conditions. Visualization of this network between mitochondria and ER has been attempted using super-resolution fluorescence imaging and light microscopy; however, the limited resolution is insufficient to observe the membranes between the mitochondria and ER in detail. Transmission electron microscopy provides good membrane contrast and nanometer-scale resolution for the observation of cellular organelles; however, it is exceptionally time-consuming when assessing the three-dimensional (3D) structure of highly curved organelles. Therefore, we observed the morphology of mitochondria and ER via correlative light-electron microscopy (CLEM) and volume electron microscopy techniques using enhanced ascorbate peroxidase 2 and horseradish peroxidase staining. An en bloc staining method, ultrathin serial sectioning (array tomography), and volume electron microscopy were applied to observe the 3D structure. In this protocol, we suggest a combination of CLEM and 3D electron microscopy to perform detailed structural studies of mitochondria and ER.

We present a protocol to study the distribution of mitochondria and endoplasmic reticulum in whole cells after genetic modification using correlative light and volume electron microscopy including ascorbate peroxidase 2 and horseradish peroxidase staining, serial sectioning of cells with and without the target gene in the same section, and serial imaging via electron microscopy.

Cellular organelles, such as mitochondria and endoplasmic reticulum (ER), create a network to perform a variety of functions. These highly curved ...

Production of Dynein and Kinesin Motor Ensembles on DNA Origami Nanostructures for Single Molecule Observation

Cytoskeletal motors are responsible for a wide variety of functions in eukaryotic cells, including mitosis, cargo transport, cellular motility, and others. Many of these functions require motors to operate in ensembles. Despite a wealth of knowledge about the mechanisms of individual cytoskeletal motors, comparatively less is known about the mechanisms and emergent behaviors of motor ensembles, examples of which include changes to ensemble processivity and velocity with changing motor number, location, and configuration. Structural DNA nanotechnology, and the specific technique of DNA origami, enables the molecular construction of well-defined architectures of motor ensembles. The shape of cargo structures as well as the type, number and placement of motors on the structure can all be controlled. Here, we provide detailed protocols for producing these ensembles and observing them using total internal reflection fluorescence microscopy. Although these techniques have been specifically applied for cytoskeletal motors, the methods are generalizable to other proteins that assemble in complexes to accomplish their tasks. Overall, the DNA origami method for creating well-defined ensembles of motor proteins provides a powerful tool for dissecting the mechanisms that lead to emergent motile behavior.

The goal of this protocol is to form ensembles of molecular motors on DNA origami nanostructures and observe the ensemble motility using total internal reflection fluorescence microscopy.

Cytoskeletal motors are responsible for a wide variety of functions in eukaryotic cells, including mitosis, cargo transport, cellular motility, and ...

Visualization of Estrogen Receptors in Colons of Mice with TNBS-Induced Crohn's Disease using Immunofluorescence

Crohn&#39;s disease is the most diagnosed type of inflammatory bowel disease. Chronic inflammation developing in the intestine leads to peristalsis disorder and damage of intestinal mucosa and seems to be associated with an increased risk of colon neoplastic transformation. Accumulating evidence indicates that estrogens and estrogen receptors affect not only hormone-sensitive tissues, but also other tissues not directly related to estrogens, such as the lungs or colon. Here, we describe the protocol for the successful immunofluorescence staining of estrogen receptors in colon obtained from a murine model of TNBS-induced Crohn&#39;s disease. A detailed protocol for the induction of Crohn&#39;s disease in mice and intestine preparation is provided as well as a step-by-step immunohistochemical procedure using formalin-fixed paraffin-embedded intestine sections. The described methods are not only useful for estrogen receptor detection and estrogen signaling investigation in vivo but can also be applied to for other proteins which may be involved in the development of colitis.

The protocol presents a complete validated TNBS-induced murine model of Crohn&#39;s disease and methods for visualization of estrogen receptors by immunohistochemistry using immunofluorescence of formalin-fixed colon sections embedded in paraffin.

Crohn&#39;s disease is the most diagnosed type of inflammatory bowel disease. Chronic inflammation developing in the intestine leads to peristalsis ...

Characterizing Cellular Proteins with In-cell Fast Photochemical Oxidation of Proteins

Fast photochemical oxidation of proteins (FPOP) is a hydroxyl radical protein footprinting method used to characterize protein structure and interactions. FPOP uses a 248 nm excimer laser to photolyze hydrogen peroxide producing hydroxyl radicals. These radicals oxidatively modify solvent exposed side chains of 19 of the 20 amino acids. Recently, this method has been used in live cells (IC-FPOP) to study protein interactions in their native environment. The study of proteins in cells accounts for intermolecular crowding and various protein interactions that are disrupted for in vitro studies. A custom single cell flow system was designed to reduce cell aggregation and clogging during IC-FPOP. This flow system focuses the cells past the excimer laser individually, thus ensuring consistent irradiation. By comparing the extent of oxidation produced from FPOP to the protein&#39;s solvent accessibility calculated from a crystal structure, IC-FPOP can accurately probe the solvent accessible side chains of proteins.

Here, we characterize protein structure and interaction sites in living cells using a protein footprinting technique termed in-cell fast photochemical oxidation of proteins (IC-FPOP).

Fast photochemical oxidation of proteins (FPOP) is a hydroxyl radical protein footprinting method used to characterize protein structure and interactions. ...

Transverse Sectioning of Mature Rice (Oryza sativa L.) Kernels for Scanning Electron Microscopy Imaging Using Pipette Tips as Immobilization Support

Starch granules (SGs) exhibit different morphologies depending on the plant species, especially in the endosperm of the Poaceae family. Endosperm phenotyping can be used to classify genotypes based on SG morphotype using scanning electron microscopic (SEM) analysis. SGs can be visualized using SEM by slicing through the kernel (pericarp, aleurone layers, and endosperm) and exposing the organellar contents. Current methods require the rice kernel to be embedded in plastic resin and sectioned using a microtome or embedded in a truncated pipette tip and sectioned by hand using a razor blade. The former method requires specialized equipment and is time-consuming, while the latter introduces a new host of problems depending on rice genotype. Chalky rice varieties, particularly, pose a problem for this type of sectioning due to the friable nature of their endosperm tissue. Presented here is a technique for preparing translucent and chalky rice kernel sections for microscopy, requiring only pipette tips and a scalpel blade. Preparing the sections within the confines of a pipette tip prevents rice kernel endosperm from shattering (for translucent or &#8216;vitreous&#8217; phenotypes) and crumbling (for chalky phenotypes). Using this technique, endosperm cell patterning and the structure of intact SGs can be observed.

Transverse Sectioning of Mature Rice Oryza sativa L. Kernels for Scanning Electron Microscopy Imaging Using Pipette Tips as Immobilization Support

This protocol allows for the preparation of transverse sections of cereal seeds (e.g., rice) for the analysis of endosperm and starch granule morphology using scanning electron microscopy.