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>

<ol>
	<li>Hauser, M., Wojcik, M., Kim, D., Mahmoudi, M., Li, W., Xu, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Correlative+Super-Resolution+Microscopy:+New+Dimensions+and+New+Opportunities.">Correlative Super-Resolution Microscopy: New Dimensions and New Opportunities.</a> Chem Rev. , (2017).</li><li>Dempsey, G. T., Vaughan, J. C., Chen, K. H., Bates, M., Zhuang, X. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evaluation+of+fluorophores+for+optimal+performance+in+localization-based+super-resolution+imaging.">Evaluation of fluorophores for optimal performance in localization-based super-resolution imaging.</a> Nat Methods. 8 (12), 1027-1036 (2011).</li><li>F&#246;lling, J., Bossi, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescence+nanoscopy+by+ground-state+depletion+and+single-molecule+return.">Fluorescence nanoscopy by ground-state depletion and single-molecule return.</a> Nat Methods. 5 (11), 943-945 (2008).</li><li>Betzig, E., Patterson, G. H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+intracellular+fluorescent+proteins+at+nanometer+resolution.">Imaging intracellular fluorescent proteins at nanometer resolution.</a> Science (New York, N.Y.). 313 (5793), 1642-1645 (2006).</li><li>Kopek, B. G., Shtengel, G., Grimm, J. B., Clayton, D. A., Hess, H. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Correlative+photoactivated+localization+and+scanning+electron+microscopy.">Correlative photoactivated localization and scanning electron microscopy.</a> PLOS one. 8 (10), e77209 (2013).</li><li>Paez-Segala, M. G., Sun, M. G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fixation-resistant+photoactivatable+fluorescent+proteins+for+CLEM.">Fixation-resistant photoactivatable fluorescent proteins for CLEM.</a> Nat Methods. 12 (3), 215-218 (2015).</li><li>Narayan, K., Subramaniam, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Focused+ion+beams+in+biology.">Focused ion beams in biology.</a> Nat Methods. 12 (11), 1021-1031 (2015).</li><li>Tokuyasu, K. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Immunochemistry+on+ultrathin+frozen+sections.">Immunochemistry on ultrathin frozen sections.</a> Histochem J. 12 (4), 381-403 (1980).</li><li>Suleiman, H., Zhang, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nanoscale+protein+architecture+of+the+kidney+glomerular+basement+membrane.">Nanoscale protein architecture of the kidney glomerular basement membrane.</a> eLife. 2, e01149 (2013).</li><li>Kopek, B. G., Shtengel, G., Xu, C. S., Clayton, D. A., Hess, H. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Correlative+3D+superresolution+fluorescence+and+electron+microscopy+reveal+the+relationship+of+mitochondrial+nucleoids+to+membranes.">Correlative 3D superresolution fluorescence and electron microscopy reveal the relationship of mitochondrial nucleoids to membranes.</a> Proc Nat Acad Sci U S A. 109 (16), 6136-6141 (2012).</li><li>Avanesov, A., Malicki, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Analysis+of+the+retina+in+the+zebrafish+model.">Analysis of the retina in the zebrafish model.</a> Methods Cell Biol. 100, 153-204 (2010).</li><li>Bachmann-Gagescu, R., Phelps, I. G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+ciliopathy+gene+cc2d2a+controls+zebrafish+photoreceptor+outer+segment+development+through+a+role+in+Rab8-dependent+vesicle+trafficking.">The ciliopathy gene cc2d2a controls zebrafish photoreceptor outer segment development through a role in Rab8-dependent vesicle trafficking.</a> Hum Mol Gen. 20 (20), 4041-4055 (2011).</li><li>Bachmann-Gagescu, R., Dona, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Ciliopathy+Protein+CC2D2A+Associates+with+NINL+and+Functions+in+RAB8-MICAL3-Regulated+Vesicle+Trafficking.">The Ciliopathy Protein CC2D2A Associates with NINL and Functions in RAB8-MICAL3-Regulated Vesicle Trafficking.</a> PLOS Gen. 11 (10), e1005575 (2015).</li><li>Mateos, J. M., Guhl, B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Topographic+contrast+of+ultrathin+cryo-sections+for+correlative+super-resolution+light+and+electron+microscopy.">Topographic contrast of ultrathin cryo-sections for correlative super-resolution light and electron microscopy.</a> Sci Rep. 6, 34062 (2016).</li><li>Westerfield, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The zebrafish book. A guide for the laboratory use of zebrafish (Danio rerio). , (2000).</li><li>Thompson, R. E., Larson, D. R., Webb, W. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Precise+nanometer+localization+analysis+for+individual+fluorescent+probes.">Precise nanometer localization analysis for individual fluorescent probes.</a> Biophys J. 82 (5), 2775-2783 (2002).</li><li>Ovesn&#253;, M., K&#345;&#237;&#382;ek, P., Borkovec, J., Svindrych, Z., Hagen, G. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=ThunderSTORM:+a+comprehensive+ImageJ+plug-in+for+PALM+and+STORM+data+analysis+and+super-resolution+imaging.">ThunderSTORM: a comprehensive ImageJ plug-in for PALM and STORM data analysis and super-resolution imaging.</a> Bioinformatics. 30 (16), 2389-2390 (2014).</li><li>Schindelin, J., Arganda-Carreras, I., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fiji:+an+open-source+platform+for+biological-image+analysis.">Fiji: an open-source platform for biological-image analysis.</a> Nat Methods. 9 (7), 676-682 (2012).</li><li>Cardona, A., Saalfeld, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=TrakEM2+software+for+neural+circuit+reconstruction.">TrakEM2 software for neural circuit reconstruction.</a> PLOS one. 7 (6), e38011 (2012).</li><li>Wurm, C. A., Neumann, D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nanoscale+distribution+of+mitochondrial+import+receptor+Tom20+is+adjusted+to+cellular+conditions+and+exhibits+an+inner-cellular+gradient.">Nanoscale distribution of mitochondrial import receptor Tom20 is adjusted to cellular conditions and exhibits an inner-cellular gradient.</a> Proc Nat Acad Sci U S A. 108 (33), 13546-13551 (2011).</li><li>Slot, J. W., Geuze, H. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cryosectioning+and+immunolabeling.">Cryosectioning and immunolabeling.</a> Nat Prot. 2 (10), 2480-2491 (2007).</li></ol>

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 &amp;lt;111&gt; 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 &quot;Perfect loop&quot;</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&#039;)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>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>&lt;strong&gt;Instruments&lt;/strong&gt;</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. Dit protocol beschrijft het nodige eiwit subcellular localisatie om resultaten te verkrijgen op de zebravis netvlies door correleren super resolutie de lichte microscopie en elektronenmicroscopie afbeeldingen scannen.

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

Imaging Subcellular Structures in the Living Zebrafish Embryo

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such imaging experiments in higher vertebrates such as mammals generally requires surgical access to the system under study. The optical accessibility of embryonic and larval zebrafish allows such invasive procedures to be circumvented and permits imaging in the intact organism. Indeed the zebrafish is now a well-established model to visualize dynamic cellular behaviors using in vivo microscopy in a wide range of developmental contexts from proliferation to migration and differentiation. A more recent development is the increasing use of zebrafish to study subcellular events including mitochondrial trafficking and centrosome dynamics. The relative ease with which these subcellular structures can be genetically labeled by fluorescent proteins and the use of light microscopy techniques to image them is transforming the zebrafish into an in vivo model of cell biology. Here we describe methods to generate genetic constructs that fluorescently label organelles, highlighting mitochondria and centrosomes as specific examples. We use the bipartite Gal4-UAS system in multiple configurations to restrict expression to specific cell-types and provide protocols to generate transiently expressing and stable transgenic fish. Finally, we provide guidelines for choosing light microscopy methods that are most suitable for imaging subcellular dynamics.

Imaging the dynamic behavior of organelles and other subcellular structures in vivo can shed light on their function in physiological and disease conditions. Here, we present methods for genetically tagging two organelles, centrosomes and mitochondria, and imaging their dynamics in living zebrafish embryos using wide-field and confocal microscopy.

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such ...

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 ...

Culture of Adult Transgenic Zebrafish Retinal Explants for Live-cell Imaging by Multiphoton Microscopy

An endogenous regeneration program is initiated by M&#252;ller glia in the adult zebrafish (Danio rerio) retina following neuronal damage and death. The M&#252;ller glia re-enter the cell cycle and produce neuronal progenitor cells that undergo subsequent rounds of cell divisions and differentiate into the lost neuronal cell types. Both M&#252;ller glia and neuronal progenitor cell nuclei replicate their DNA and undergo mitosis in distinct locations of the retina, i.e. they migrate between the basal Inner Nuclear Layer (INL) and the Outer Nuclear Layer (ONL), respectively, in a process described as Interkinetic Nuclear Migration (INM). INM has predominantly been studied in the developing retina. To examine the dynamics of INM in the adult regenerating zebrafish retina in detail, live-cell imaging of fluorescently-labeled M&#252;ller glia/neuronal progenitor cells is required. Here, we provide the conditions to isolate and culture dorsal retinas from Tg[gfap:nGFP]mi2004 zebrafish that were exposed to constant intense light for 35 h. We also show that these retinal cultures are viable to perform live-cell imaging experiments, continuously acquiring z-stack images throughout the thickness of the retinal explant for up to 8 h using multiphoton microscopy to monitor the migratory behavior of gfap:nGFP-positive cells. In addition, we describe the details to perform post-imaging analysis to determine the velocity of apical and basal INM. To summarize, we established conditions to study the dynamics of INM in an adult model of neuronal regeneration. This will advance our understanding of this crucial cellular process and allow us to determine the mechanisms that control INM.

Zebrafish retinal regeneration has mostly been studied using fixed retinas. However, dynamic processes such as interkinetic nuclear migration occur during the regenerative response and require live-cell imaging to investigate the underlying mechanisms. Here, we describe culture and imaging conditions to monitor Interkinetic Nuclear Migration (INM) in real-time using multiphoton microscopy.

An endogenous regeneration program is initiated by M&#252;ller glia in the adult zebrafish (Danio rerio) retina following neuronal damage and death. The ...

Retrograde Labeling of Retinal Ganglion Cells in Adult Zebrafish with Fluorescent Dyes

As retrograde labeling retinal ganglion cells (RGCs) can isolate RGCs somata from dying sites, it has become the gold standard for counting RGCs in RGCs survival and regeneration experiments. Many studies have been performed in mammalian animals to research RGCs survival after optic nerve injury. However, retrograde labeling of RGCs in adult zebrafish has not yet been reported, though some alternative methods can count cell numbers in retinal ganglion cell layers (RGCL). Considering the small size of the adult zebrafish skull and the high risk of death after drilling on the skull, we open the skull with the help of acid-etching and seal the hole with a light curing bond, which could significantly improve the survival rate. After absorbing the dyes for 5 days, almost all the RGCs are labeled. As this method does not need to transect the optic nerve, it is irreplaceable in the research of RGCs survival after optic nerve crush in adult zebrafish. Here, we introduce this method step by step and provide representative results.

We introduce an efficient method to retrograde label retinal ganglion cells (RGCs) in adult zebrafish.

As retrograde labeling retinal ganglion cells (RGCs) can isolate RGCs somata from dying sites, it has become the gold standard for counting RGCs in RGCs ...

Neuroscience

Transplantation of GFP-expressing Blastomeres for Live Imaging of Retinal and Brain Development in Chimeric Zebrafish Embryos

Cells change extensively in their locations and property during embryogenesis. These changes are regulated by the interactions between the cells and their environment. Chimeric embryos, which are composed of cells of different genetic background, are great tools to study the cell-cell interactions mediated by genes of interest. The embryonic transparency of zebrafish at early developmental stages permits direct visualization of the morphogenesis of tissues and organs at the cellular level. Here, we demonstrate a protocol to generate chimeric retinas and brains in zebrafish embryos and to perform live imaging of the donor cells. The protocol covers the preparation of transplantation needles, the transplantation of GFP-expressing donor blastomeres to GFP-negative hosts, and the examination of donor cell behavior under live confocal microscopy. With slight modifications, this protocol can also be used to study the embryonic development of other tissues and organs in zebrafish. The advantages of using GFP to label donor cells are also discussed. 

We demonstrate a protocol to generate chimeric zebrafish embryos for live imaging cellular behavior during embryogenesis.

Cells change extensively in their locations and property during embryogenesis. These changes are regulated by the interactions between the cells and their ...

Biology

Microdissection of Zebrafish Embryonic Eye Tissues

Zebrafish is a popular animal model for research on eye development because of its rapid ex utero development and good fecundity. By 3 days post fertilization (dpf), the larvae will show the first visual response. Many genes have been identified to control a proper eye development, but we are far from a complete understanding of the underlying genetic architecture. Whole genome gene expression profiling is a useful tool to elucidate genetic regulatory network for eye development. However, the small size of the embryonic eye in zebrafish makes it challenging to obtain intact and pure eye tissues for expression analysis. For example, the anterior-posterior length of the eye between day 2 and 3 is only approximately 200-300 &mu;m, while the diameter of the lens is less 100 &mu;m. Also, the retinal pigment epithelium (RPE) underlying the retina is just a single-layer epithelium. While gene expression profiles can be obtained from the whole embryo, they do not accurately represent the expression of these tissues. Therefore pure tissue must be obtained for a successful gene expression profiling of eye development. To address this issue, we have developed an approach to microdissect intact retina and retina with RPE attached from 1-3 dpf, which cover major stages of eye morphogenesis. All procedures can be done with fine forceps and general laboratory supplies under standard stereomicroscopes. For retinal dissection, the single-layer RPE is removed and peeled off by brushing action and the preferential adherence of the RPE remnants to the surface of the culture plate for dissection. For RPE-attached retinal dissection, the adherence of RPE to the dissection plate is removed before the dissection so that the RPE can be completely preserved with the retina. A careful lifting action of this tissue can efficiently separate the presumptive choroid and sclera. The lens can be removed in both cases by a chemically etched tungsten needle. In short, our approach can obtain intact eye tissues and has been successfully utilized to study tissue-specific expression profiles of zebrafish retina1, 2 and retinal pigment epithelium3.

This article describes an approach to microdissect zebrafish retinas with and without retinal pigment epithelium attached, from one to three days postfertilization embryos.

Zebrafish is a popular animal model for research on eye development because of its rapid ex utero development and good fecundity. By 3 days post ...

In vivo Electroporation of Morpholinos into the Adult Zebrafish Retina

Many devastating inherited eye diseases result in progressive and irreversible blindness because humans cannot regenerate dying or diseased retinal neurons. In contrast, the adult zebrafish retina possesses the robust ability to spontaneously regenerate any neuronal class that is lost in a variety of different retinal damage models, including retinal puncture, chemical ablation, concentrated high temperature, and intense light treatment 1-8. Our lab extensively characterized regeneration of photoreceptors following constant intense light treatment and inner retinal neurons after intravitreal ouabain injection 2, 5, 9. In all cases, resident M&uuml;ller glia re-enter the cell cycle to produce neuronal progenitors, which continue to proliferate and migrate to the proper retinal layer, where they differentiate into the deficient neurons. 
	We characterized five different stages during regeneration of the light-damaged retina that were highlighted by specific cellular responses. We identified several differentially expressed genes at each stage of retinal regeneration by mRNA microarray analysis 10. Many of these genes are also critical for ocular development. To test the role of each candidate gene/protein during retinal regeneration, we needed to develop a method to conditionally limit the expression of a candidate protein only at times during regeneration of the adult retina. 
	Morpholino oligos are widely used to study loss of function of specific proteins during the development of zebrafish, Xenopus, chick, mouse, and tumors in human xenografts 11-14. These modified oligos basepair with complementary RNA sequence to either block the splicing or translation of the target RNA. Morpholinos are stable in the cell and can eliminate or "knockdown" protein expression for three to five days 12. 
	Here, we describe a method to efficiently knockdown target protein expression in the adult zebrafish retina. This method employs lissamine-tagged antisense morpholinos that are injected into the vitreous of the adult zebrafish eye. Using electrode forceps, the morpholino is then electroporated into all the cell types of the dorsal and central retina. Lissamine provides the charge on the morpholino for electroporation and can be visualized to assess the presence of the morpholino in the retinal cells. 
	Conditional knockdown in the retina can be used to examine the role of specific proteins at different times during regeneration. Additionally, this approach can be used to study the role of specific proteins in the undamaged retina, in such processes as visual transduction and visual processing in second order neurons.

In vivo Electroporation of Morpholinos into the Adult Zebrafish Retina

A method to conditionally knockdown a target protein&#x2019;s expression in the adult zebrafish retina is described, which involves intravitreally injecting antisense morpholinos and electroporating them into the retina. The resulting protein is knocked down for several days, which allows testing the protein&#x2019;s role in the regenerating or intact retina.

Many devastating inherited eye diseases result in progressive and irreversible blindness because humans cannot regenerate dying or diseased retinal ...

Preparing Fresh Retinal Slices from Adult Zebrafish for Ex Vivo Imaging Experiments

The retina is a complex tissue that initiates and integrates the first steps of vision. Dysfunction of retinal cells is a hallmark of many blinding diseases, and future therapies hinge on fundamental understandings about how different retinal cells function normally. Gaining such information with biochemical methods has proven difficult because contributions of particular cell types are diminished in the retinal cell milieu. Live retinal imaging can provide a view of numerous biological processes on a subcellular level, thanks to a growing number of genetically encoded fluorescent biosensors. However, this technique has thus far been limited to tadpoles and zebrafish larvae, the outermost retinal layers of isolated retinas, or lower resolution imaging of retinas in live animals. Here we present a method for generating live ex vivo retinal slices from adult zebrafish for live imaging via confocal microscopy. This preparation yields transverse slices with all retinal layers and most cell types visible for performing confocal imaging experiments using perfusion. Transgenic zebrafish expressing fluorescent proteins or biosensors in specific retinal cell types or organelles are used to extract single-cell information from an intact retina. Additionally, retinal slices can be loaded with fluorescent indicator dyes, adding to the method&#39;s versatility. This protocol was developed for imaging Ca2+ within zebrafish cone photoreceptors, but with proper markers it could be adapted to measure Ca2+ or metabolites in M&#252;ller cells, bipolar and horizontal cells, microglia, amacrine cells, or retinal ganglion cells. The retinal pigment epithelium is removed from slices so this method is not suitable for studying that cell type. With practice, it is possible to generate serial slices from one animal for multiple experiments. This adaptable technique provides a powerful tool for answering many questions about retinal cell biology, Ca2+, and energy homeostasis.

Preparing Fresh Retinal Slices from Adult Zebrafish for Ex Vivo Imaging Experiments

Imaging retinal tissue can provide single-cell information that cannot be gathered from traditional biochemical methods. This protocol describes preparation of retinal slices from zebrafish for confocal imaging. Fluorescent genetically encoded sensors or indicator dyes allow visualization of numerous biological processes in distinct retinal cell types.

The retina is a complex tissue that initiates and integrates the first steps of vision. Dysfunction of retinal cells is a hallmark of many blinding ...

4-Dimensional Imaging of Zebrafish Optic Cup Morphogenesis

Visual system function requires the establishment of precise tissue and organ structures. In the vertebrate eye, structural defects are a common cause of visual impairment, yet mechanisms of eye morphogenesis are still poorly understood. The basic organization of the embryonic eye is conserved throughout vertebrates, thus live imaging of zebrafish embryos has become a powerful approach to directly observe eye development at real time under normal and pathological conditions. Dynamic cell processes including movements, morphologies, interactions, division, and death can be visualized in the embryo. We have developed methods for uniform labeling of subcellular structures and timelapse confocal microscopy of early eye development in zebrafish. This protocol outlines the method of generating capped mRNA for injection into the 1-cell zebrafish embryo, mounting embryos at optic vesicle stage (~12 hours post fertilization, hpf), and performing multi-dimensional timelapse imaging of optic cup morphogenesis on a laser scanning confocal microscope, such that multiple datasets are acquired sequentially in the same imaging session. Such an approach yields data that can be used for a variety of purposes, including cell tracking, volume measurements, three-dimensional (3D) rendering, and visualization. Our approaches allow us to pinpoint the cellular and molecular mechanisms driving optic cup development, in both wild type and genetic mutant conditions. These methods can be employed directly by other groups or adapted to visualize many additional aspects of zebrafish eye development.

This protocol describes an approach for in toto labeling and multidimensional imaging of zebrafish early eye development. We describe labeling, embedding, and four dimensional (4D) imaging using laser scanning confocal microscopy, and considerations for optimizing acquisition of datasets for dissecting mechanisms of optic cup morphogenesis.

Visual system function requires the establishment of precise tissue and organ structures. In the vertebrate eye, structural defects are a common cause of ...

Synaptic Circuits Analysis and Protein Localization in the Mouse Retina

Investigating Retinal Circuits and Molecular Localization by Pre&#45;Embedding Immunoelectron Microscopy

The retina comprises numerous cells forming diverse neuronal circuits, which constitute the first stage of the visual pathway. Each circuit is characterized by unique features and distinct neurotransmitters, determining its role and functional significance. Given the intricate cell types within its structure, the complexity of neuronal circuits in the retina poses challenges for exploration. To better investigate retinal circuits and cross-talk, such as the link between cone and rod pathways, and precise molecular localization (neurotransmitters or neuropeptides), such as the presence of substance P-like immunoreactivity in the mouse retina, we employed a pre-embedding immunoelectron microscopy (immuno-EM) method to explore synaptic connections and organization. This approach enables us to pinpoint specific intercellular synaptic connections and precise molecular localization and could play a guiding role in exploring its function. This article describes the protocol, reagents used, and detailed steps, including (1) retina fixation preparation, (2) pre-embedding immunostaining, and (3) post-fixation and embedding.

Author Spotlight: Understanding the Ultrastructural Basis of Retinal Synaptic Connectivity and Neurotransmitter Localization in Mice 

This protocol outlines the detailed steps of pre-embedding immunoelectron microscopy, with a focus on exploring synaptic circuits and protein localization in the retina.

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

Samenvatting

Meer video's verkennen