Funding ION and RGB: Swiss National Science Foundation Ambizione-SCORE grant PZ00P3 142404/1 and PZ00P3 163979.

All experiments were performed in accordance with the ARVO Statement for the Use of Animals in Ophthalmic and Vision Research and were approved by the local authorities.
1. Preparation of Ultrathin Sections on Silicon Wafers
<ol>
	<li>Sample fixation
	<ol>
		<li>Prepare the fixative solution containing 0.1% glutaraldehyde, 4% formaldehyde in 0.1 M cacodylate buffer. 
		NOTE: CAUTION! Use caution when working with glutaraldehyde, formaldehyde and cacodylate buffer, wear appropriate personal protective equipment and work in a fume hood.</li>
		<li>Euthanize 5 days post fertilization (5 dpf) zebrafish larvae with tricaine (ethyl 3-aminobenzoate methanesulfonate, 0.4% w/v in PBS (phosphate buffer saline, pH 7)) as previously described15.</li>
		<li>Fix the larvae by immersion in pre-chilled fixative solution on ice. Remove the head and incubate overnight at 4 &#176;C with gentle rocking.</li>
		<li>Dissect the eyes by trimming the tissue around the eye using fine forceps and a microdissection scalpel (under a binocular) in chilled fixative solution on a 1% agarose-bedded plate. Transfer the eyes to a tube with fresh pre-chilled fixative.</li>
		<li>Wash twice in PBS (phosphate buffer saline, pH 7.4) for 5 min each wash at room temperature (RT).</li>
	</ol>
	</li>
	<li>Gelatin infiltration and mounting
	<ol>
		<li>Warm up 15 mL local food brand gelatin 12% w/v in PB&#160;(phosphate buffer, 0.1 M pH 7.4) at 40 &#176;C. Remove the PBS and add gelatin solution to the tubes containing the eyes. Tap the tube gently to ensure the gelatin infiltrates the sample and incubate for 10 - 30 min at 40 &#176;C in a thermoblock with gentle shaking or in a water bath.</li>
		<li>Fill 12 mm x 5 mm x 3 mm silicon or polyethylene flat embedding molds with warm gelatin in a 40 &#176;C water bath. Add two eyes per mold using a pipette, align them properly under a binocular using a dissection needle and let the gelatin cool down at room temperature for 1 min and harden at 4 &#176;C for 20 min.</li>
		<li>Re-trim the gelatin block under the binocular to fit one eye per block using a razor blade.</li>
		<li>Transfer the gelatin embedded eyes to 2.3 M sucrose in PB on ice. Incubate at 4 &#176;C overnight.</li>
		<li>Exchange to new 2.3 M sucrose solution and store at 4 &#176;C or -20 &#176;C; after this step, samples are ready for sectioning or can be stored at -20 &#176;C for several weeks to months.</li>
		<li>Re-trim the gelatin block to almost the size of the eye before transferring to a cryo-pin. Freeze in liquid nitrogen and transfer to a cryo-ultramicrotome.</li>
	</ol>
	</li>
	<li>Cryosectioning
	<ol>
		<li>Cut 110 nm thick-sections at -120 &#176;C with a diamond knife in the cryo-ultramicrotome.</li>
		<li>Pick the sections with a wired loop containing a droplet of 2% methylcellulose (in water) and 2.3 M sucrose solution (1:1). Transfer sections to a 7 mm x 7 mm silicon wafer. Store sections at 4 &#176;C until further processing.</li>
	</ol>
	</li>
</ol>
2. Immunolabelling
<ol>
	<li>Wash wafers with PBS at 0 &#176;C for 20 min on four&#160;drops by placing the wafers upside down on the drops. Wash in PBS 2x 2 min at room temperature.</li>
	<li>Incubate 3 times in 0.15% glycine in PBS, for 1 min each. Wash 3x for 1 min/wash with PBS. Pre-incubate with PBG (PBS with 0.5 % Bovine Serum Albumin BSA and 0.2 % Gelatin type B) for 5 min.</li>
	<li>Incubate with rabbit anti-Tom20 (see the Table of Materials) in PBG (4 &#181;g/mL) for 30 min at room temperature.</li>
	<li>Wash 6x for 1 min/wash in PBG. Pre-incubate with PBG for 5 min at RT. Incubate with anti-rabbit Alexa 647 F(ab&#39;)2 (see the Table of Materials) in PBG (7.5 &#181;g/mL) for 30 min.</li>
	<li>Wash 6x for 1 min/wash in PBG. Wash 3x 2 min in PBS. Incubate with DAPI (4 &#181;g/mL) in PBS for 10 s. Wash 2x 2 min in PBS.</li>
</ol>
3. Super-resolution Microscopy
<ol>
	<li>Incubate the wafer briefly (10 s) by placing it on a droplet of a 1:1 solution of glycerol (80%) and imaging buffer containing an oxygen scavenging system (200 mM Phosphate buffer containing 10% glucose, 0.5 mg/mL glucoseoxidase, 40 &#181;g/mL catalase, 15 mM beta-mercaptoethylamine hydrochloride (MEA HCL), pH 8.0).</li>
	<li>Transfer the wafers (section facing down) to a glass bottom (thickness 170 &#177; 5 &#181;m) Petri dish onto a fresh drop of the 1:1 mixture of glycerol (80%) and imaging buffer (as in step 3.1).</li>
	<li>Remove from all sides, with a pipette, most of the liquid underneath the wafer. Use silicone stripes to fix the wafer to the bottom of the Petri dish. 
	NOTE: Silicone stripes are made out of two-component silicone-glue (3 mm x 12 mm).</li>
	<li>Image sections on an inverted microscope using a high numerical aperture (e.g. 160X&#160;/ NA 1.43; see the Table of Materials) oil immersion super resolution dedicated objective. Prior to imaging, let the sample equilibrate to microscope temperature in order to minimize/reduce lateral and axial drift.</li>
	<li>Center the area of interest and acquire first widefield epifluorescence reference images. Change to the super resolution operation mode. Adjust the exposure time of the camera to 15 ms and set the electron multiplying (EM) gain to the maximum of 300.</li>
	<li>Illuminate sample with the 642 nm continuous wave laser at maximum laser power (corresponding to ~2.8 kW/cm2) in epifluorescence mode. As soon as the single molecule blinks are well separated in each frame, so that the probability is low that individual signals overlap, set the laser power to ~0.7 kW/cm2. Record the raw image in epifluorescence mode by acquiring a minimum of 30,000 frames. 
	NOTE: All these parameter can vary depending on different specimens and labeling densities.</li>
	<li>From the raw data, generate a reconstruction event list (localizations of each single molecule blink in the raw image) using a detection threshold of 30 photons (needs to be adjusted according to the sample) by clicking &#34;Evaluate&#34; in the &#39;t-series analysis&#39; under &#39;Tools&#39;.</li>
	<li>Visualize the super resolution image by Gaussian fitting16 applying a rendering pixel size of 4 nm by clicking &#34;Create Image&#34; in the &#39;event list processing panel&#39; under &#39;Tools.&#39; 
	NOTE: For generating the super resolved image the integrated software tools of the system used in this study were employed. However, the described super resolution imaging could be performed on any other single molecule localization based system, and the data could be processed with open source software tools as thunderSTORM17.</li>
</ol>
4. Platinum Shadowing
<ol>
	<li>Remove silicon stripes and add a drop of PBS close to the edges of the wafer to lift it up from the Petri dish. Wash the wafer 2x 2 min in PBS, then post fix it with 0.1% glutaraldehyde in PBS for 5 min. Wash 2x for 2 min/wash in PBS.</li>
	<li>Incubate the wafer twice for 5 min/incubation in 1 drop of methylcellulose 2 % in water on ice. Insert the wafer in a centrifuge tube and centrifuge at 14,100 x g for 90 s. Mount it on a SEM aluminum stub with conducting carbon cement.</li>
	<li>Add a layer of 2 to 10 nm of platinum/carbon on the sample by rotary shadowing at 8&#176; using an electron beam evaporation device settings: 1.55 kV, 55 mA, at 0.3 nm/s, and an angle of 8&#176;, rotation level 4.</li>
</ol>
5. Scanning Electron Microscopy
<ol>
	<li>Image sections with a scanning electron microscope at 1.5 kV, 2 mm working distance and with an In lens secondary electron detector.</li>
</ol>
6. Alignment of Light and Electron Microscopy Images
<ol>
	<li>Open both types of images with Fiji18 by clicking &#34;File|Open&#34;. Adjust canvas size by clicking &#34;Image| Adjust| Canvas size&#34; and bring both images to a stack by clicking &#34;Image| Stacks| Images to stack&#34;. Save the stack as tiff file type.</li>
	<li>In Fiji, open a new TrackEM219 interface by clicking &#34;File|new|TrackEM2 (new)&#34;. Import the stack with both images by right clicking on the black window and selecting &#34;Import stack&#34;.</li>
	<li>Align light microscopy image to electron microscopy image manually with landmarks by using a right mouse click on the image and selecting &#34;Align|Align layer manually with landmarks&#34;.
	<ol>
		<li>Pick the select tool (the black arrow) to add landmarks. Use the shape of nuclei as reference to select the same edges in both images (Supplementary Figure 1). Add several points (minimum three points need to be selected). Apply alignment with an affine model by right mouse clicking, and selecting &#34;Apply transform| Affine model&#34;.</li>
	</ol>
	</li>
	<li>Change layer transparency (see Supplementary Figure 1) to assess the quality of the alignment.</li>
</ol>

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The authors have nothing to disclose.

This method combines super-resolved protein localization with context information to determine the precise position of proteins in an organelle. We demonstrate here the completion of the experiment to visualize the expression of Tom20 in the outer membrane of mitochondria, and its relation to other organelles like nuclei or outer segments of the photoreceptor in the larval zebrafish retina.
Tokuyasu cryo-sectioning requires some training to acquire well-preserved sections. However, this is a method employed in many laboratories with demonstrated success21. Transfer of the sections to the silicon wafer is very simple and no special considerations are needed. The use of glycerol in the imaging buffer is a very critical step to avoid drying of the sections. For super-resolution imaging the best results are obtained when the wafer is very close to the glass bottom of the Petri dish. The silicone stripes help maintaining the wafer in this position. Special care has to be taken while removing the stripes in order to avoid damaging the sections.
The thickness of the cryo-sections, around 100 nm, thinner than the optical resolution in the Z-dimension, additionally facilitates the accuracy of this correlative method as the super-resolution microscopy signal is coming only from this thin layer and the scanning electron microscope signal is displaying the topography of the sample. This method could also be combined with multicolor imaging. However, special care must be taken that the sample can be imaged under same conditions (e.g. imaging buffer) and cross talk should be prevented.
One limitation of the method is its two dimensional approach, since only a limited number of serial sections (about 3 to 7 per wafer) can be collected. Thus, projects dealing with volume analysis would not be ideal. However, it is a method that can be applied to detect protein expression in any type of tissue by simple sectioning of the sample. We provide a method without use of classical contrast agents like uranyl acetate or lead citrate. The contrast by platinum shadowing provides very informative topographical contrast, but in some cases membranes are difficult to resolve. This may pose problems for projects that need, for example, to determine the expression of proteins in small vesicles.
Our protocol, based on Tokuyasu cryo-sections, uses standard equipment to obtain correlative results from super-resolution and scanning electron microscopes. The collection of sections on silicon wafers and the use of platinum for contrast are simple steps to provide stability and reproducibility to the sample preparation.

Methods to determine the subcellular localization of proteins and their relationship to different compartments of the cell are essential tools to understand their functions and possible interactions. Super-resolution microscopy in combination with electron microscopy provides such information1. Ground state depletion microscopy followed by individual molecule return (GSDIM) is a super-resolution microscopy technology compatible with a wide range of organic and genetically encoded fluorophores2 and achieves a lateral resolution up to 20 nm3. The incorporation of methods with higher resolution than standard diffraction-limited microscopy improves the accuracy of the correlation4,5,6. In order to achieve the best correlation of protein expression with a specific subcellular compartment and to reduce the volume of uncertainty7 the use of the same ultrathin section for light and electron microscopy is recommended. Among the different sectioning methods, Tokuyasu cryo-section protocol does not require dehydration or resin embedding and, in addition, preserves the antigenicity of many epitopes and provides good tissue ultrastructure8. Several methods have demonstrated the applicability of these sections in correlative light and electron microscopy (CLEM)4,5,9,10.
The zebrafish retina is a valuable model to study visual development and human disease mechanisms given its highly conserved structure and function across vertebrates. In particular, retinal photoreceptors display the same architecture as mammalian photoreceptors, with a basal synapse, an apico-basally elongated nucleus, clustering of mitochondria in the more apical inner segment and an outer segment composed of membrane disks in the most apical position11. Protein localization to the diverse cellular compartments is conserved between zebrafish and human, allowing investigation of the biological function of human disease-relevant proteins12,13.
Here we present a protocol to prepare larval zebrafish retina samples to resolve the localization of the mitochondrial outer membrane protein Tom20 by correlative super-resolution light and electron microscopy. The method is based on collecting cryo-sections on silicon wafers and obtaining contrast by topographical information produced after application of a thin layer of platinum. These steps are clear technical improvements in terms of ease of use, reproducibility, and time to complete experiments. We have recently demonstrated the applicability of the method to detect nuclear pores and mitochondrial proteins in mouse tissue14.

<table>
	<tbody>
		<tr>
			<td>Paraformaldehyde</td>
			<td>Sigma-Aldrich</td>
			<td>#158127</td>
			<td></td>
		</tr>
		<tr>
			<td>Glutaraldehyde EM Grade</td>
			<td>EMS, USA</td>
			<td>#16220</td>
			<td></td>
		</tr>
		<tr>
			<td>Cacodylate</td>
			<td>Merck</td>
			<td>#8.20670</td>
			<td></td>
		</tr>
		<tr>
			<td>Tricaine</td>
			<td>Sigma-Aldrich</td>
			<td>#886-86-2</td>
			<td></td>
		</tr>
		<tr>
			<td>Agarose, peqGOLD Universal</td>
			<td>VWR International GmbH</td>
			<td>35-1020</td>
			<td></td>
		</tr>
		<tr>
			<td>Flat embedding molds</td>
			<td>BEEM Flat</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Local food brand gelatin</td>
			<td>Dr.Oetker</td>
			<td>Extra Gold</td>
			<td></td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>Merck</td>
			<td>#1.07687</td>
			<td></td>
		</tr>
		<tr>
			<td>Methylcellulose</td>
			<td>Sigma</td>
			<td>#M-6385</td>
			<td></td>
		</tr>
		<tr>
			<td>Glycine</td>
			<td>Sigma</td>
			<td>#G-7126</td>
			<td></td>
		</tr>
		<tr>
			<td>Gelatin type B</td>
			<td>Sigma</td>
			<td>#G-6650</td>
			<td></td>
		</tr>
		<tr>
			<td>BSA</td>
			<td>Applichem</td>
			<td>#A6588.0050</td>
			<td></td>
		</tr>
		<tr>
			<td>Silicon wafer</td>
			<td>Si-Mat Silicon Materials</td>
			<td></td>
			<td>Type: P/Boron; Orientation &#60;111&#62; ON; Growth method: CZ; Resistivity:1-30 ohm/cm; Surface: polished; Laser cut at 7 x7 mm</td>
		</tr>
		<tr>
			<td>Cryo-pin</td>
			<td>Baltic Preparation</td>
			<td>#16701950</td>
			<td></td>
		</tr>
		<tr>
			<td>Wired loop &#34;Perfect loop&#34;</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Silicone stripes</td>
			<td>Picodent</td>
			<td>Twinsil 22</td>
			<td></td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td>Sigma-Aldrich</td>
			<td>#G8270</td>
			<td></td>
		</tr>
		<tr>
			<td>glucoseoxidase</td>
			<td>Sigma-Aldrich</td>
			<td>#G7141</td>
			<td></td>
		</tr>
		<tr>
			<td>catalase</td>
			<td>Sigma-Aldrich</td>
			<td>#C40</td>
			<td></td>
		</tr>
		<tr>
			<td>beta-Mercaptoethylamine hydrochloride</td>
			<td>Sigma-Aldrich</td>
			<td>#M6500</td>
			<td></td>
		</tr>
		<tr>
			<td>anti-Tom20</td>
			<td>Santa Cruz Biotechnology</td>
			<td>#sc - 11415</td>
			<td></td>
		</tr>
		<tr>
			<td>AlexaFluor 647 AffiniPure F(ab&#39;)2 fragment donkey anti rabbit IgG</td>
			<td>Jackson Immuno-Research</td>
			<td>#711-606-152</td>
			<td></td>
		</tr>
		<tr>
			<td>DAPI</td>
			<td>ROCHE, Switzerland</td>
			<td>#10236276001</td>
			<td></td>
		</tr>
		<tr>
			<td>Glycerol solution</td>
			<td>Sigma-Aldrich</td>
			<td>#49782</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass bottom petri dish</td>
			<td>Ibidi, Germany</td>
			<td>u-Dish 35mm, high glass bottom, #81158</td>
			<td></td>
		</tr>
		<tr>
			<td>SEM aluminium stub</td>
			<td>Agar Scientific</td>
			<td>#G301F</td>
			<td></td>
		</tr>
		<tr>
			<td>Conducting carbon cement Leit-C</td>
			<td>Plano, Germany</td>
			<td>AG3300</td>
			<td></td>
		</tr>
		<tr>
			<td>Name</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>Instruments</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Diamond Knife (Cryo Immuno)</td>
			<td>Diatome</td>
			<td>DCIMM3520</td>
			<td></td>
		</tr>
		<tr>
			<td>Cryo-ultramicrotome</td>
			<td>Leica Microsystems</td>
			<td>Leica EM FCS</td>
			<td></td>
		</tr>
		<tr>
			<td>Widefield-TIRF microscope GSD Leica SR GSD 3D</td>
			<td>Leica Microsystems</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>160x 1.43 TIRF objective</td>
			<td>Leica Microsystems</td>
			<td>11523048</td>
			<td></td>
		</tr>
		<tr>
			<td>SEM - Zeiss Supra 50 VP</td>
			<td>Zeiss</td>
			<td>Supra 50 VP</td>
			<td></td>
		</tr>
		<tr>
			<td>FIB-SEM - Zeiss Auriga 40</td>
			<td>Zeiss</td>
			<td>Auriga 40</td>
			<td></td>
		</tr>
	</tbody>
</table>

correlative super-resolution and electron microscopy to resolve protein localization in zebrafish retina

We present a method to investigate the subcellular protein localization in the larval zebrafish retina by combining super-resolution light microscopy and scanning electron microscopy. The sub-diffraction limit resolution capabilities of super-resolution light microscopes allow improving the accuracy of the correlated data. Briefly, 110 nanometer thick cryo-sections are transferred to a silicon wafer and, after immunofluorescence staining, are imaged by super-resolution light microscopy. Subsequently, the sections are preserved in methylcellulose and platinum shadowed prior to imaging in a scanning electron microscope (SEM). The images from these two microscopy modalities are easily merged using tissue landmarks with open source software. Here we describe the adapted method for the larval zebrafish retina. However, this method is also applicable to other types of tissues and organisms. We demonstrate that the complementary information obtained by this correlation is able to resolve the expression of mitochondrial proteins in relation with the membranes and cristae of mitochondria as well as to other compartments of the cell.

The expression of the protein Tom20, a subunit of the translocase mitochondrial outer membrane complex20, was determined, in thin sections of larval zebrafish retina, by super-resolution light microscopy (Figure 1) and this information was complemented with the topographical signal obtained by scanning electron microscopy after platinum shadowing of the same sections. These correlative data confirm the localization of a protein in association with a particular compartment, the outer mitochondrial membrane and, in addition provide information about the relation of the protein with other organelles of the cell.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56113/56113fig1.jpg" /> 
Figure 1: CLEM on zebrafish retina. A. Low magnification widefield image of a 5 dpf zebrafish retinal section, nuclei stained with DAPI (cyan). B. Scanning electron microscopy of the same area. C. Higher magnification widefield image of frame in B. Nuclei stained with DAPI (cyan) and Tom20 mitochondrial staining appears in red. D. Widefield image of same section at higher magnification. The pattern of Tom20 expression is at the clusters of mitochondria. E. Expression of Tom20 (red dots) detected by GSDIM microscopy. Nuclei stained with DAPI (cyan). F. Same section as E combining correlative super-resolution and scanning electron microscopy. Tom20 staining (red dots) appears at the mitochondrial cluster (M) at the outer membranes of mitochondria. Fluorescence DAPI signal in the nuclei (N) corresponds with the topography of the SEM image. G. High magnification image of frame in F. The scanning electron microscopy image provides context to the GSDIM image (red dots). Mitochondrial cristae are clearly visible and the Tom20 staining is localized to the outer membranes of mitochondria. The membranes of the outer segment the photoreceptors (OS) are clearly resolved. Image pixel size 5 nm. Scale bars: A, B, and C: 10 &#181;m; D: 2 &#181;m; E and F: 1 &#181;m and G: 0.2 &#181;m. <a href="http://cloudflare.jove.com/files/ftp_upload/56113/56113fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplementary Figure 1: Alignment of light and electron microscopy images. A. Screenshot from TrackEM2 interface with SEM image and numbered landmarks (yellow) along different nuclei. B. Screenshot from TrackEM2 interface with fluorescence image and numbered landmarks (yellow) along different DAPI stained nuclei. To change layer transparency the sliders on the left upper part of the menu can be used. Scale bar: 1 &#181;m. <a href="http://cloudflare.jove.com/files/ftp_upload/56113/Supplementary_Figure_1.tif">Please click here to download this file.</a>

Watch this Scientific Journal Video about Correlative Super-resolution and Electron Microscopy to Resolve Protein Localization in Zebrafish Retina at JoVE.com

Correlative Super-resolution and Electron Microscopy to Resolve Protein Localization in Zebrafish Retina

This protocol describes the necessary steps to obtain subcellular protein localization results on zebrafish retina by correlating super-resolution light microscopy and scanning electron microscopy images.

We present a method to investigate the subcellular protein localization in the larval zebrafish retina by combining super-resolution light microscopy and ...

correlative-super-resolution-electron-microscopy-to-resolve-protein

Research

JoVE Journal

Developmental Biology

9.4K Views.  University of Zurich. This protocol describes the necessary steps to obtain subcellular protein localization results on zebrafish retina by correlating super-resolution light microscopy and scanning electron microscopy images.

Video: Correlative Super-resolution and Electron Microscopy to Resolve Protein Localization in Zebrafish Retina

Measuring Protein Stability in Living Zebrafish Embryos Using Fluorescence Decay After Photoconversion (FDAP)

Protein stability influences many aspects of biology, and measuring the clearance kinetics of proteins can provide important insights into biological systems. In FDAP experiments, the clearance of proteins within living organisms can be measured. A protein of interest is tagged with a photoconvertible fluorescent protein, expressed in vivo and photoconverted, and the decrease in the photoconverted signal over time is monitored. The data is then fitted with an appropriate clearance model to determine the protein half-life. Importantly, the clearance kinetics of protein populations in different compartments of the organism can be examined separately by applying compartmental masks. This approach has been used to determine the intra- and extracellular half-lives of secreted signaling proteins during zebrafish development. Here, we describe a protocol for FDAP experiments in zebrafish embryos. It should be possible to use FDAP to determine the clearance kinetics of any taggable protein in any optically accessible organism.

Measuring Protein Stability in Living Zebrafish Embryos Using Fluorescence Decay After Photoconversion FDAP

Protein levels in cells and tissues are often tightly regulated by the balance of protein production and clearance. Using Fluorescence Decay After Photoconversion (FDAP), the clearance kinetics of proteins can be experimentally measured in vivo.

Protein stability influences many aspects of biology, and measuring the clearance kinetics of proteins can provide important insights into biological ...

Analysis of Zebrafish Larvae Skeletal Muscle Integrity with Evans Blue Dye

The zebrafish model is an emerging system for the study of neuromuscular disorders. In the study of neuromuscular diseases, the integrity of the muscle membrane is a critical disease determinant. To date, numerous neuromuscular conditions display degenerating muscle fibers with abnormal membrane integrity; this is most commonly observed in muscular dystrophies. Evans Blue Dye (EBD) is a vital, cell permeable dye that is rapidly taken into degenerating, damaged, or apoptotic cells; in contrast, it is not taken up by cells with an intact membrane. EBD injection is commonly employed to ascertain muscle integrity in mouse models of neuromuscular diseases. However, such EBD experiments require muscle dissection and/or sectioning prior to analysis. In contrast, EBD uptake in zebrafish is visualized in live, intact preparations. Here, we demonstrate a simple and straightforward methodology for performing EBD injections and analysis in live zebrafish. In addition, we demonstrate a co-injection strategy to increase efficacy of EBD analysis. Overall, this video article provides an outline to perform EBD injection and characterization in zebrafish models of neuromuscular disease.

In this study, we describe a straightforward method to perform Evans Blue Dye (EBD) analysis on zebrafish larvae. This technique is a powerful tool for the characterization of skeletal muscle integrity and delineation of zebrafish models of muscular dystrophy, and is a valuable method for the development of novel therapeutics.

The zebrafish model is an emerging system for the study of neuromuscular disorders.  In the study of neuromuscular diseases, the integrity of the muscle ...

Large-scale Scanning Transmission Electron Microscopy (Nanotomy) of Healthy and Injured Zebrafish Brain

Large-scale 2D electron microscopy (EM), or nanotomy, is the tissue-wide application of nanoscale resolution electron microscopy. Others and we previously applied large scale EM to human skin pancreatic islets, tissue culture and whole zebrafish larvae1-7. Here we describe a universally applicable method for tissue-scale scanning EM for unbiased detection of sub-cellular and molecular features. Nanotomy was applied to investigate the healthy and a neurodegenerative zebrafish brain. Our method is based on standardized EM sample preparation protocols: Fixation with glutaraldehyde and osmium, followed by epoxy-resin embedding, ultrathin sectioning and mounting of ultrathin-sections on one-hole grids, followed by post staining with uranyl and lead. Large-scale 2D EM mosaic images are acquired using a scanning EM connected to an external large area scan generator using scanning transmission EM (STEM). Large scale EM images are typically ~ 5 - 50 G pixels in size, and best viewed using zoomable HTML files, which can be opened in any web browser, similar to online geographical HTML maps. This method can be applied to (human) tissue, cross sections of whole animals as well as tissue culture1-5. Here, zebrafish brains were analyzed in a non-invasive neuronal ablation model. We visualize within a single dataset tissue, cellular and subcellular changes which can be quantified in various cell types including neurons and microglia, the brain&#39;s macrophages. In addition, nanotomy facilitates the correlation of EM with light microscopy (CLEM)8 on the same tissue, as large surface areas previously imaged using fluorescent microscopy, can subsequently be subjected to large area EM, resulting in the nano-anatomy (nanotomy) of tissues. In all, nanotomy allows unbiased detection of features at EM level in a tissue-wide quantifiable manner.

Large-scale Scanning Transmission Electron Microscopy Nanotomy of Healthy and Injured Zebrafish Brain

Large-scale 2D electron microscopy (EM), or nanotomy, is the tissue-wide application of nanoscale resolution EM. Here we describe a universal method for nanotomy applied to investigate the zebrafish larval brain in health and upon non-invasive brain injury.

Large-scale 2D electron microscopy (EM), or nanotomy, is the tissue-wide application of nanoscale resolution electron microscopy. Others and we previously ...

Analysis of Zebrafish Kidney Development with Time-lapse Imaging Using a Dissecting Microscope Equipped for Optical Sectioning

In order to understand organogenesis, the spatial and temporal alterations that occur during development of tissues need to be recorded. The method described here allows time-lapse analysis of normal and impaired kidney development in zebrafish embryos by using a fluorescence dissecting microscope equipped for structured illumination and z-stack acquisition. To visualize nephrogenesis, transgenic zebrafish (Tg(wt1b:GFP)) with fluorescently labeled kidney structures were used. Renal defects were triggered by injection of an antisense morpholino oligonucleotide against the Wilms tumor gene wt1a, a factor known to be crucial for kidney development.
The advantage of the experimental setup is the combination of a zoom microscope with simple strategies for re-adjusting movements in x, y or z direction without additional equipment. To circumvent focal drift that is induced by temperature variations and mechanical vibrations, an autofocus strategy was applied instead of utilizing a usually required environmental chamber. In order to re-adjust the positional changes due to a xy-drift, imaging chambers with imprinted relocation grids were employed.
In comparison to more complex setups for time-lapse recording with optical sectioning such as confocal laser scanning&#160;or light sheet microscopes, a zoom microscope is easy to handle. Besides, it offers dissecting microscope-specific benefits such as high depth of field and an extended working distance.
The method to study organogenesis presented here can also be used with fluorescence stereo microscopes not capable of optical sectioning. Although limited for high-throughput, this technique offers an alternative to more complex equipment that is normally used for time-lapse recording of developing tissues and organ dynamics.

The method described here allows time-lapse analysis of organ development in zebrafish embryos by using a fluorescence dissecting microscope capable of performing optical sectioning and simple strategies of readjustment to correct focal and planar drift.

In order to understand organogenesis, the spatial and temporal alterations that occur during development of tissues need to be recorded. The method ...

Using Light Sheet Fluorescence Microscopy to Image Zebrafish Eye Development

Light sheet fluorescence microscopy (LSFM) is gaining more and more popularity as a method to image embryonic development. The main advantages of LSFM compared to confocal systems are its low phototoxicity, gentle mounting strategies, fast acquisition with high signal to noise ratio and the possibility of imaging samples from various angles (views) for long periods of time. Imaging from multiple views unleashes the full potential of LSFM, but at the same time it can create terabyte-sized datasets. Processing such datasets is the biggest challenge of using LSFM. In this protocol we outline some solutions to this problem. Until recently, LSFM was mostly performed in laboratories that had the expertise to build and operate their own light sheet microscopes. However, in the last three years several commercial implementations of LSFM became available, which are multipurpose and easy to use for any developmental biologist. This article is primarily directed to those researchers, who are not LSFM technology developers, but want to employ LSFM as a tool to answer specific developmental biology questions. 
Here, we use imaging of zebrafish eye development as an example to introduce the reader to LSFM technology and we demonstrate applications of LSFM across multiple spatial and temporal scales. This article describes a complete experimental protocol starting with the mounting of zebrafish embryos for LSFM. We then outline the options for imaging using the commercially available light sheet microscope. Importantly, we also explain a pipeline for subsequent registration and fusion of multiview datasets using an open source solution implemented as a Fiji plugin. While this protocol focuses on imaging the developing zebrafish eye and processing data from a particular imaging setup, most of the insights and troubleshooting suggestions presented here are of general use and the protocol can be adapted to a variety of light sheet microscopy experiments.

Light sheet fluorescence microscopy is an excellent tool for imaging embryonic development. It allows recording of long time-lapse movies of live embryos in near physiological conditions. We demonstrate its application for imaging zebrafish eye development across wide spatio-temporal scales and present a pipeline for fusion and deconvolution of multiview datasets.

Light sheet fluorescence microscopy (LSFM) is gaining more and more popularity as a method to image embryonic development. The main advantages of LSFM ...

Analysis of Protein-protein Interactions and Co-localization Between Components of Gap, Tight, and Adherens Junctions in Murine Mammary Glands

Cell-cell interactions play a pivotal role in preserving tissue integrity and the barrier between the different compartments of the mammary gland. These interactions are provided by junctional proteins that form nexuses between adjacent cells. Junctional protein mislocalization and reduced physical associations with their binding partners can result in the loss of function and, consequently, to organ dysfunction. Thus, identifying protein localization and protein-protein interactions (PPIs) in normal and disease-related tissues is essential to finding new evidences and mechanisms leading to the progression of diseases or alterations in developmental status. This manuscript presents a two-step method to evaluate PPIs in murine mammary glands. In protocol section 1, a method to perform co-immunofluorescence (co-IF) using antibodies raised against the proteins of interest, followed by secondary antibodies labeled with fluorochromes, is described. Although co-IF allows for the demonstration of the proximity of the proteins, it does make it possible to study their physical interactions. Therefore, a detailed protocol for co-immunoprecipitation (co-IP) is provided in protocol section 2. This method is used to determine the physical interactions between proteins, without confirming whether these interactions are direct or indirect. In the last few years, co-IF and co-IP techniques have demonstrated that certain components of intercellular junctions co-localize and interact together, creating stage-dependent junctional nexuses that vary during mammary gland development.

Intercellular junctions are requisites for mammary gland stage-specific functions and development. This manuscript provides a detailed protocol for the study of protein-protein interactions (PPIs) and co-localization using murine mammary glands. These techniques allow for the investigation of the dynamics of the physical association between intercellular junctions at different developmental stages.

Cell-cell interactions play a pivotal role in preserving tissue integrity and the barrier between the different compartments of the mammary gland. These ...

Multimodal Hierarchical Imaging of Serial Sections for Finding Specific Cellular Targets within Large Volumes

Targeting specific cells at ultrastructural resolution within a mixed cell population or a tissue can be achieved by hierarchical imaging using a combination of light and electron microscopy. Samples embedded in resin are sectioned into arrays consisting of ribbons of hundreds of ultrathin sections and deposited on pieces of silicon wafer or conductively coated coverslips. Arrays are imaged at low resolution using a digital consumer like smartphone camera or light microscope (LM) for a rapid large area overview, or a wide field fluorescence microscope (fluorescence light microscopy (FLM)) after labeling with fluorophores. After post-staining with heavy metals, arrays are imaged in a scanning electron microscope (SEM). Selection of targets is possible from 3D reconstructions generated by FLM or from 3D reconstructions made from the SEM image stacks at intermediate resolution if no fluorescent markers are available. For ultrastructural analysis, selected targets are finally recorded in the SEM at high-resolution (a few nanometer image pixels). A ribbon-handling tool that can be retrofitted to any ultramicrotome is demonstrated. It helps with array production and substrate removal from the sectioning knife boat. A software platform that allows automated imaging of arrays in the SEM is discussed. Compared to other methods generating large volume EM data, such as serial block-face SEM (SBF-SEM) or focused ion beam SEM (FIB-SEM), this approach has two major advantages: (1) The resin-embedded sample is conserved, albeit in a sliced-up version. It can be stained in different ways and imaged with different resolutions. (2) As the sections can be post-stained, it is not necessary to use samples strongly block-stained with heavy metals to introduce contrast for SEM imaging or render the tissue blocks conductive. This makes the method applicable to a wide variety of materials and biological questions. Particularly prefixed materials e.g., from biopsy banks and pathology labs, can directly be embedded and reconstructed in 3D.

This protocol targets specific cells in tissue for imaging at nanoscale resolution using a scanning electron microscope (SEM). Large numbers of serial sections from resin-embedded biological material are first imaged in a light microscope to identify the target and then in a hierarchical manner in the SEM.

Targeting specific cells at ultrastructural resolution within a mixed cell population or a tissue can be achieved by hierarchical imaging using a ...

Visualizing the Node and Notochordal Plate In Gastrulating Mouse Embryos Using Scanning Electron Microscopy and Whole Mount Immunofluorescence

The post-implantation mouse embryo undergoes major shape changes after the initiation of gastrulation and morphogenesis. A hallmark of morphogenesis is the formation of the transient organizers, the node and notochordal plate, from cells that have passed through the primitive streak. The proper formation of these signaling centers is essential for the development of the body plan and techniques to visualize them are of high interest to mouse developmental biologists. The node and notochordal plate lie on the ventral surface of gastrulating mouse embryos around embryonic day (E) 7.5 of development. The node is a cup-shaped structure whose cells possess a single slender cilium each. The proper subcellular localization and rotation of the cilia in the node pit determines left-right asymmetry. The notochordal plate cells also possess single cilia albeit shorter than those of the node cells. The notochordal plate forms the notochord which acts as an important signaling organizer for somitogenesis and neural patterning. Because the cells of the node and notochordal plate are transiently present on the surface and possess cilia, they can be visualized using scanning electron microscopy (SEM). Among other techniques used to visualize these structures at the cellular level is whole mount immunofluorescence (WMIF) using the antibodies against the proteins that are highly expressed in the node and notochordal plate. In this report, we describe our optimized protocols to perform SEM and WMIF of the node and notochordal plate in developing mouse embryos to help in the assessment of tissue shape and cellular organization in wild-type and gastrulation mutant embryos.

The node and notochordal plate are transient signaling organizers in developing mouse embryos that can be visualized using several techniques. Here, we describe in detail how to perform two of the techniques to study their structure and morphogenesis: 1) scanning electron microscopy (SEM); and 2) whole mount immunofluorescence (WMIF).

The post-implantation mouse embryo undergoes major shape changes after the initiation of gastrulation and morphogenesis. A hallmark of morphogenesis is ...

A Layered Mounting Method for Extended Time-Lapse Confocal Microscopy of Whole Zebrafish Embryos

Dynamics of development can be followed by confocal time-lapse microscopy of live transgenic zebrafish embryos expressing fluorescence in specific tissues or cells. A difficulty with imaging whole embryo development is that zebrafish embryos grow substantially in length. When mounted as regularly done in 0.3-1% low melt agarose, the agarose imposes growth restriction, leading to distortions in the soft embryo body. Yet, to perform confocal time-lapse microscopy, the embryo must be immobilized. This article describes a layered mounting method for zebrafish embryos that restrict the motility of the embryos while allowing for the unrestricted growth. The mounting is performed in layers of agarose at different concentrations. To demonstrate the usability of this method, whole embryo vascular, neuronal and muscle development was imaged in transgenic fish for 55 consecutive hours. This mounting method can be used for easy, low-cost imaging of whole zebrafish embryos using inverted microscopes without requirements of molds or special equipment.

This article describes a method to mount fragile zebrafish embryos for extended time-lapse confocal microscopy. This low-cost method is easy to perform using regular glass-bottom microscopy dishes for imaging on any inverted microscope. The mounting is performed in layers of agarose at different concentrations.

Dynamics of development can be followed by confocal time-lapse microscopy of live transgenic zebrafish embryos expressing fluorescence in specific tissues ...

Whole-Mount In Situ Hybridization in Zebrafish Embryos and Tube Formation Assay in iPSC-ECs to Study the Role of Endoglin in Vascular Development

Vascular development is determined by the sequential expression of specific genes, which can be studied by performing in situ hybridization assays in zebrafish during different developmental stages. To investigate the role of endoglin(eng) in vessel formation during the development of hereditary hemorrhagic telangiectasia&#160;(HHT), morpholino-mediated targeted knockdown of eng in zebrafish are used to study its temporal expression and associated functions. Here, whole-mount in situ RNA hybridization (WISH) is employed for the analysis of eng and its downstream genes in zebrafish embryos. Also, tube formation assays are performed in HHT patient-derived induced pluripotent stem cell-differentiated&#160;endothelial cells (iPSC-ECs; with eng mutations). A specific signal amplifying system using the whole amount In Situ Hybridization &#8211; WISH provides higher resolution and lower background results compared to traditional methods. To obtain a better signal, the post-fixation time is adjusted to 30 min after probe hybridization. Because fluorescence staining is not sensitive in zebrafish embryos, it is replaced with diaminobezidine (DAB) staining here. In this protocol, HHT&#160;patient-derived iPSC lines containing an eng mutation are differentiated into endothelial cells. After coating a plate with basement membrane matrix for 30 min at 37 &#176;C, iPSC-ECs are seeded as a monolayer into wells and kept at 37 &#176;C for 3 h. Then, the tube length and number of branches are calculated using microscopic images. Thus, with this improved WISH protocol, it is shown that reduced eng expression affects endothelial progenitor formation in zebrafish embryos. This is further confirmed by tube formation assays using iPSC-ECs derived from a patient with HHT. These assays confirm the role for eng in early vascular development.

Presented here is a protocol for whole-mount in situ RNA hybridization analysis in zebrafish and tube formation assay in patient-derived induced pluripotent stem cell-derived endothelial cells to study the role of endoglin in vascular formation.

Vascular development is determined by the sequential expression of specific genes, which can be studied by performing in situ hybridization assays in ...