Majority of the work has been performed at the UMCG Microscopy and Imaging Center (UMIC), which is sponsored by NWO 175-010-2009-023 and ZonMW 91111006; STW &#34;Microscopy Valley 12718&#34; to B.N.G.G.. This work was sponsored by a ZonMW VENI grant, a Marie Curie career integration grant (saving dying neurons) and an Alzheimer Nederland fellowship to T.J.v.H.

All experiments with zebrafish larvae were approved by the Animal Experimentation Committee of the University of Groningen according to national and EU guidelines.
1. Sample Preparation
<ol>
	<li>Fixation and Embedding
	<ol>
		<li>Fixation
		<ol>
			<li>Anaesthetize fish larvae in tricaine (0.003%) added to the E3 (5 mM NaCl, 0.17 mM KCl, 0.33 mM CaCl2, 0.33 mM MgSO4, 10 mM HEPES, pH 7.6) zebrafish water and test for depth of anesthesia by poking gently with a 200 &#181;l pipette tip under dissection microscope until motion is no longer detected (Figure 1A).</li>
			<li>When processing samples for correlative EM after acquiring in vivo imaging data using confocal or 2 photon imaging in 1.8% agarose as described,5,15 fix larvae in agarose using 4% PFA in PBS containing Triton-X-100 (0.05%).</li>
			<li>Cut larvae from agarose and process for large-scale EM5.</li>
			<li>Fix larvae in fresh 4% paraformaldehyde in PBS containing Triton-X-100 (0.05%) for 2 hr at room temperature in plastic tubes. 
			Note: Cells grown in culture (in vitro) can be directly fixed in glutaraldehyde, as described in the next step (1.1.1.5).</li>
			<li>Remove solution and add 100 &#181;l 0.5% PFA, 2% glutaraldehyde (GA), and 0.1 M sodium cacodylate (pH 7.4). Store overnight at 4 &#176;C. Rinse samples twice in 0.1 M sodium cacodylate.</li>
			<li>Cut larval heads rostrally to the hindbrain to facilitate penetration of osmium. Use fine syringes/forceps.</li>
			<li>Postfix larvae in 1% osmium tetroxide (OsO4)/1.5% potassium ferrocyanide (K4[Fe(CN)6]) in 0.1 M sodium cacodylate on ice for 2 hr and rinse 3 x 5 min in double distilled H2O.</li>
			<li>Dehydrate in ethanol by 20 min incubations in increasing ethanol concentrations (50%, 70%, 3 times 100%).</li>
		</ol>
		</li>
		<li>Embedding
		<ol>
			<li>Prepare epoxy resin according to standard recipes used in EM labs. 
			Note: We use epoxy resin as follows: mix 19.8 g glycid ether 100, 9.6 g 2-dodecenylsuccinic acid anhydride and 11.4 g methylnadic anhydride using a magnetic stirrer. Add 0.5 g DMP-30 (tris-(dimethylaminomethyl) phenol) and mix again.</li>
			<li>Incubate larvae overnight in 50% epoxy resin in ethanol (v/v) while rotating (RT).</li>
			<li>Replace diluted epoxy-resin with pure resin. Incubate for 30 min, add fresh resin again and incubate for another 30 min. Again refresh the resin and then incubate 3 hr at RT, followed by 15 min at 58 &#176;C and another 1 hr at RT under low pressure (200 mbar).</li>
			<li>Orient the heads in commercially available silicon flat embedding molds under a dissection microscope using a needle or toothpick with the anterior facing the side of the mold or as needed (Figure 1B).</li>
			<li>Polymerize epoxy resin overnight at 58 &#176;C. If epoxy resin is still too soft allow another day to polymerize and harden at 58 &#176;C. Test stiffness by applying pressure using forceps. If this easily leaves an indentation allow another day to polymerize and harden at 58 &#176;C. When specimen is fully hardened proceed to trimming and sectioning.</li>
			<li>Trim away excess resin from the stub using razor blades (Figure 1B).</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Sectioning, Contrasting, and Mounting
	<ol>
		<li>Sectioning
		<ol>
			<li>Section epoxy resin block using an ultramicrotome.</li>
			<li>Use a glass knife or a diamond histoknife to cut semithin sections (500 nm) for toluidine blue/basic fuchsin (Figure 1D) staining to detect the right positioning.</li>
			<li>Transfer semi-thin sections on microscopic slides by picking them up with a glass Pasteur pipet whose tip has been closed by melting in a flame. Dry on a hot plate until no water is left. Stain for 10 sec with 1% toluidine blue in water on a hot plate and after rinsing with water, stain for 10 sec with 0.05% basic fuchsin in 1% sodium tetraborate. Examine with a normal light microscope at 10X to 40X.</li>
			<li>When the proper site/orientation within the brain is reached, continue sectioning the epoxy resin block with a diamond knife to cut ultrathin sections (~ 70 nm; Figure 1E)</li>
			<li>Use easily identifiable anatomic structures in and around the brain, including olfactory pits, eyes, grey-white matter boundaries, to identify the region of interest during ultrathin sectioning and to adjust the sectioning angle when the sample is tilted.</li>
			<li>Mount section on single slot L2 x 1 copper grids (Figure 1F), to allow acquisition uninterrupted by grid bars. Use Formvar coated grids to support the sections.</li>
		</ol>
		</li>
		<li>Contrasting
		<ol>
			<li>Stain ultrathin sections on grids for 2 min in 2% uranyl acetate in methanol followed by Reynolds lead citrate for another 2 min 16.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
2. Image Acquisition
<ol>
	<li>Large Scale STEM 
	Note: We use scanning EM with an external scan generator capable of acquiring multiple large fields of view at high resolution3,5,7,17 using STEM detection. Typically, one image generated this way is equivalent to the fields of view of ~ 100 Transmission Electron Microscope (TEM) images, significantly reducing the amount of stitching.
	<ol>
		<li>Mounting Sample in Microscope
		<ol>
			<li>Place grid with section in the multiple grid sample holder and transfer into the chamber of the SEM.</li>
		</ol>
		</li>
		<li>Alignment of the STEM Detector
		<ol>
			<li>Put the grid with the sample under the beam and move it up so it will be approx. 5 mm from the lens pole. Acquire an image at 20 kV using the in-lens detector.</li>
			<li>Focus on grid edge and adjust working distance to 4.0 mm.</li>
			<li>Move the sample holder to a safe position and bring in the STEM detector.</li>
			<li>Center the hole of the STEM detector in the image-field by turning the adjustments screws while imaging at maximum speed with the in-lens detector.</li>
			<li>Carefully bring back the sample holder which will just fit between the lens pole and the detector.</li>
			<li>Acquire an image with the in-lens detector to be sure to be on the section area.</li>
			<li>Switch to STEM detection (gain medium and all quadrants set on &#39;minus&#39;) and acquire an image and select the area to be scanned.</li>
			<li>Raise acceleration voltage to 21 kV and wait for stabilization of the beam (stable image again) and then go to 29 kV in steps of 2 kV.</li>
		</ol>
		</li>
		<li>Acquiring Images
		<ol>
			<li>Pre-irradiate the sample (to prevent brightness changes caused by the e-beam) by zooming out so the complete area to be scanned fits the image window.</li>
			<li>Change aperture to 120 &#181;m (to keep the proper dynamic range the detector gains have to be reduced one step).</li>
			<li>De-focus so the image is blurred.</li>
			<li>Make the scanned area as tight as possible using the reduced area scan option.</li>
			<li>Set scan speed such that a frame is scanned in approx. 1 - 2 sec. 
			Note: Depending on the size and tissue this typically takes &#189; hr (100 x 100 &#181;m FOV) to up to 3 hr (1,000 x 500 &#181;m).</li>
			<li>Zoom in at least 100X and scan a small area for 10 sec. 
			Note: When this area does not change brightness compared to its surrounding, pre-irradiation is sufficient.</li>
			<li>Bring back the 30 &#181;m aperture (and STEM gain on medium)</li>
			<li>Focus the sample.</li>
			<li>Align aperture using the wobbler (at ~ 40,000X magnification).</li>
			<li>While in focus select the lightest area/feature, set scan speed such that details are visible.</li>
			<li>In the microscope software adjust brightness and contrast by carefully watching the histogram to keep all pixels in the dynamic range. Do the same for the darkest area/features.</li>
			<li>Go back to the bright area and check again. Be on the safe side so there is some space on both sides of the histogram.</li>
			<li>Zoom out so the complete area to be scanned fits the imaging window, and start the large area acquisition program.</li>
			<li>Use the wizard option to set up a mosaic by selecting an area from the screen. Use pixel size 2 - 5 nm depending on what details are necessary. Use dwell time 3 &#181;s for STEM.</li>
			<li>Select option &#34;autofocus on previous tile&#34;, use the large scale acquisition software autofocus with default company settings, check box &#34;verify setting prior to execution&#34; and define where to save the data.</li>
			<li>Press &#34;optimize&#34; to check microscope settings. 
			Note: Now the time needed will be shown. This might be up to 72 hr for 1 x 0.5 mm areas.</li>
			<li>In window &#34;maximum tile resolution&#34; type 20,000 so individual tiles will not be bigger than 20k x 20k, preventing blunt edge effects. Press &#34;execute&#34;.</li>
			<li>When asked for checking focus and Brightness/Contrast (B/C), switch off external scan generator and carefully adjust focus and astigmatism in the microscope software. Do not change B/C. 
			Note: The stage can be moved and magnification changed, but magnification has to be set back to the desired pixel size.</li>
			<li>Switch on external scan generator again and press &#34;continue&#34;.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
3. Data Analysis
<ol style="margin-left: 40px;">
	<li>Open large-scale EM file viewer program and open the file &#34;mosaic info&#34;. This will open all the tiled tif-files. This preferably is performed on another workstation to not obstruct new acquisitions.</li>
	<li>Choose option &#39;auto stitch entire mosaic (parameters overlap mode on half, stitching threshold 0.90, noise reduction automatic. In case stitching criteria cannot be fulfilled, zoom in and manually place tile in position and click &#39;continue as is&#39;).</li>
	<li>Export as an html file, or alternatively as a single TIF. 
	Note that for TIF export a downscaling might be needed. Include the final pixel size in the file name enabling measurements later.</li>
	<li>Perform annotations in the TIF file by opening it in an image editing program. In parallel, open the zoomable html file in a web browser for optimal resolution observation.</li>
</ol>

<ol>
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The authors have nothing to disclose.

EM allows analysis of cellular context with high resolution imaging of macromolecules in biological context. However, this typically limits the field of view. Larger scale 3D EM is particularly suitable for mapping neuronal connectivity by creating nanoscale resolution 3 dimensional reconstructions, requiring complex data processing10. In contrast, 2D large scale EM requires only a single section and stitching of the imaging data, and assessment of the data is possible by anyone with access to an Internet browser. We and others previously used large scale EM to analyze tissues and whole animals. As for most approaches, TEM and SEM-based stitching do have their own advantages. Here, scanning transmission EM (STEM) was used that allows generation of a large field of view at high resolution. Typically, one STEM image is equivalent to the fields of view of approximately 100 TEM images, significantly reducing the amount of stitching when imaging large fields of view at high resolution. If higher resolution is needed, TEM may be advantageous. STEM has the advantage over TEM that non-contrasted samples can be used with good ultrastructural contrast6 .
Additionally, the method described here can be simply adjusted for use with back scattered electron detection (BSD) on sections mounted on silica wafers, broadening the use to multiple microscope systems. HTML-zoomable tissue EM files are very useful for quantification, sharing data that may not have been analyzed to its full content, combining LM and EM data (CLEM)8, presentation purposes in scientific research, to analyze patient data, and for education. Alternatively, in large-scale EM in a SEM, but not in a TEM, silica wafers can be used, which has two main advantages: BSD can be used, which is more generally available on scanning electron microscopes than STEM detection. Second, mounting of large sections (&#62; 1 mm2 areas of interest) is straightforward. Mounting on single slotted grids is labor intensive and technically challenging. Detailed comparison between TEM, SEM and STEM is detailed elsewhere6. A disadvantage of BSD imaging is that, compared to STEM, images have increased noise. This can partly be compensated by increasing the pixel dwell time, resulting in much longer acquisition times.
Although for sample preparation relatively standard EM processing (fixation, embedding and sectioning) is needed5-7 , it is technically challenging to cut large ultrathin sections completely devoid of artifacts. Sections are very fragile, easily break, fold or are destroyed during imaging, which typically takes multiple hours per dataset. However, because of the online sharing of the raw unbiased data, it should become possible to compare more easily to published data, and use published dataset in the open domain as controls. Currently, few EM devices capable of large-scale analysis are in use, and therefore accessibility to this technique is somewhat limited, though most imaging centers welcome collaborative efforts.
For large-scale sections the tissue has to be properly fixed throughout the sample. That is why a mixture of the fast but moderate fixative PFA is used in combination with the slow but strong fixative GA. Cutting and picking up large sections without artifacts is difficult. Working with wafers in combination with BSD is easier compared to collecting sections on single hole grids. Heavy metal post staining is more critical compared to classical EM. Since the whole section is imaged every artifact will be visible. TEM users typically find it difficult to switch to a SEM, because of the difference in microscope operation.
Quantification and data sharing - Quantifying subcellular features is difficult in single EM images. The possibility to zoom in and out of large-scale images readily allows identification of cells of interest, which can be followed by nanoscale measurements inside the cells. These datasets indicate that specific cell types can be rapidly identified and quantified in an unbiased manner in these large tissue sections, based on features detectable at different scales. For example microglia can be identified based on their morphology and dense cytoplasm. Subsequently, upon zooming in on the individual cells, nanoscale subcellular and molecular features of these cells can be measured within the same dataset, as we previously showed for ER width within a large scale EM tissue culture dataset2. An additional advantage of nanotomy is that hosting of large scale datasets online will allow others to inspect the data, maybe for other features, and draw their conclusions on new hypothesis.
CLEM - In addition to facilitating quantitative EM, large scale EM makes it easier to correlate light microscopic labeling to EM level8. In the present example the presence of phagocytic microglia in a zebrafish ablation model is shown. A major question in neuroscience is what the individual functions and contributions are of microglia and potential other sources of phagocytic and immune cells. Early EM studies have shown distinctive subcellular features of microglia in disease18. Unfortunately, it is difficult to selectively label microglia in particular in a pathological setting, as they exhibit large overlap with other immune cells in gene expression, morphology and function. Therefore, it is unclear whether and when microglia at the ultrastructural level differ from immune cells from other sources including monocyte derived infiltrating macrophages. Understanding whether there are ultrastructural differences between these cells will provide a starting point for analysis of functional differences. Combining selective transgenic or expression markers and CLEM allows detection of ultrastructural features selective to specific populations.
Diagnosis &#38; presentation and education - By visualizing nanoscale to microscale within a single dataset EM data is greatly facilitated to a broad audience. With the increased possibilities and tools for correlative and large-scale EM we anticipate a revival of EM in basic and medical research. Our method represented here is applied to a zebrafish brain injury model5, but has been used on human tissue7, rat brain3, in a rat model for diabetes4 and in cell culture2, and can also be used in combination with a TEM-based approach1 showing the versatility of this method. The microscope operator is no longer recording highly selected, and hence biased images, but all numerous ultrastructural features are recorded and open for worldwide analysis.

Recent technical developments have improved the versatility, applicability and quantitative nature of EM, leading to a revival of ultrastructural analysis. Advances include 3D EM, large scale 2D EM and improved methods and reagents for correlated light microscopy and electron microscopy (CLEM) to compare other modes of microscopic analysis directly to the EM level8-10. Large-scale 2D EM is particularly suitable to quantify or identify (novel) disease features for human pathology, study animal models for disease and tissue culture models. Due to the typically small field of view it is difficult to correlate changes at high magnification to a tissue wide scale, as well as to unbiasedly and quantitatively analyze ultrastructural features.
For pathological analysis of human tissue or the assessment of pathology in animal models, haematoxylin and eosin (H&#38;E) colored sections of formaldehyde fixed paraffin embedded (FFPE) tissue is the standard. Besides simple H&#38;E staining immunolabeling is also performed to identify pathological abnormalities. If such tissues could be analyzed at the EM level, specific cell types, and subcellular changes could be identified. The unbiased nature of large scale EM allows finding unexpected and novel features of disease. By large-scale EM areas up to square millimeters can be visualized. We and others previously applied nanotomy to rat pancreatic islets4, cell culture2, rat brain3, skin and mucosa7 and whole zebrafish larvae1,5 (www.nanotomy.org). Zebrafish are highly suitable for in vivo imaging, in particularly to visualize cell types which are difficult to access in mammalian tissues including brain immune cells11. Here the nanotomy procedure is described in detail, applied to coronal sections of zebrafish heads undergoing conditional neuronal ablation by conversion of metronidazole by neuronal expressed nitroreductase (www.nanotomy.org)5,12-14.&#160; All raw data is presented as zoomable HTML files, visualizing molecular to tissue scale changes. The presentation of the raw data allows unbiased analyses of the datasets from other angles by experts worldwide.

<table border="0" cellpadding="0" cellspacing="0" width="738">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:248px;">Chemicals</td>
			<td style="width:204px;"></td>
			<td style="width:119px;"></td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:248px;">Low melting point agarose</td>
			<td style="width:204px;">VWR</td>
			<td style="width:119px;">444152G</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">tricaine</td>
			<td style="width:204px;">Sigma</td>
			<td style="width:119px;">E10521</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Triton- X-100&#160;</td>
			<td style="width:204px;">Sigma</td>
			<td style="width:119px;">X100</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">glutaraldehyde</td>
			<td style="width:204px;">Polysciences</td>
			<td style="width:119px;">1909</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Sodium cacodylate</td>
			<td style="width:204px;">Sigma</td>
			<td style="width:119px;">C0250</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">osmiumtetroxide</td>
			<td style="width:204px;">Electron Microscopy Sciences</td>
			<td style="width:119px;">19114</td>
			<td></td>
		</tr>
		<tr height="25">
			<td height="25" style="height:25px;">potassiumferrocyanide (K4[Fe(CN)]6)</td>
			<td style="width:204px;">Merck</td>
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			<td height="21" style="height:21px;">glycid ether 100</td>
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</table>

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 EM dataset of coronal sections of the head of 1-week-old zebrafish larvae shows many tissue and cellular features in a single large scale image (www.nanotomy.org).
Nanotomy of control brain sections reveals typical ultrastructural features of neural tissue of the rostral forebrain including olfactory fiber bundles, neuronal nuclei and neuronal sub-compartments including synapses (Figure 2A, www.nanotomy.org). Cell types in the head that can be distinguished include chondrocytes with large vacuoles in the lower jaw, osmiophilic myelin of the olfactory nerve cells, endothelial cells of a small vessel, structures include the meninx (the zebrafish counterpart of the meninges, the membranous layer encasing the mammalian brain). Subcellular structures include the glycocalyx of the epidermis, different nuclear morphologies and foci in different cell types and organelles including Golgi apparatus, endoplasmic reticulum (ER), mitochondria and synaptic vesicles and post-synaptic densities.
Unexpectedly the nuclei of cells lining the ventricle are very electron dense compared to other cells dispersed more laterally in the brain. Additionally, these ventricular nuclei show dark foci that are absent in other cells in the brain and appear different in size and structure from the heterochromatin, which is found in nuclei of phagocytic microglia (Figure 2). Cells lining the ventricle in developing zebrafish include mostly radial glial cells and neuronal progenitors, which may reflect an altered chromatin state than more differentiated cells. As stem cells in tissue are generally quite rare, nanotomy to correlate tissue labeling for stem cell markers with tissue scale EM could therefore be used to identify ultrastructural features of stem cells.
Large-scale EM (www.nanotomy.org) of a coronal section in a zebrafish larva undergoing neuronal ablation reveals many specific tissues, and cellular and molecular features not found in control animals. Cellular features include phagocytic microglia and vacuoles the size of cells, likely representing dying cells (Figure 2B, www.nanotomy.org).&#160; Previous EM studies have identified microglia in vertebrates18,19. Characteristic features of microglia include elongated nuclei, clumps of patchy chromatin next to the nuclear envelope, prominent Golgi apparatus, free polyribosomes, granular endoplasmic reticulum (ER) with long narrow cisternae, relatively dark/dense cytoplasm, and numerous inclusions, such as phagosomes, lipid droplets and lysosomes. All these hallmarks, attributed to microglia in mammalian tissues, were also found in these cells in zebrafish brains (Figure 2B, www.nanotomy.org).
<img alt="Figure 1" src="/files/ftp_upload/53635/53635fig1.jpg" /> 
Figure 1: Workflow of Large-scale Tissue Electron Microscopy on Zebrafish Brains. (A) 7 day-old zebrafish larva. (B) Zebrafish larval heads embedded side by side in epoxy resin. (C) Glass knife for semi -thin sectioning and orienting the specimen. (D) Representative semithin toluidine blue stained section showing two side-by-side embedded zebrafish larvae. (E) Diamond knife to cut ultrathin (70 nm) sections. (F) Edited image in (D) of whole zebrafish larval brain section on one-hole grid to indicate positioning. (G) Microscopy system used for large scale 2D EM in this study. (H) Flow of image processing. <a href="https://www.jove.com/files/ftp_upload/53635/53635fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/53635/53635fig2.jpg" /> 
Figure 2: Nanotomy Results: From Macromolecule to Tissue. (A) Nanotomy of brains of 7 days post fertilization control and (B) treated (neuronal ablation; Figure 1), showing features specific to the neuronal ablated degenerative brain. (B) These include phagocytic microglia, dark cells undergoing cell death and spongy appearance of neural tissue ((A) and (B)). Upper panels (adapted from5 ): 10X magnified view of region indicated in middle panels. Middle panels: 10X magnified view of region indicated in lower panels. (A, middle panel) High magnification view of neuropil showing synapse in (A, lower panel), synaptic vesicles (SVs) and postsynaptic density (PD). (B, Middle panel) High magnification view of amoeboid microglia or possibly a macrophage (middle panel, M) showing typical amoeboid microglial features (lower panel) including prominent Golgi apparatus (G), inclusions including lysosomal vacuoles (L). Scale bars: 50 &#181;m (top), 5 &#181;m (middle) and 0.5 &#181;m (bottom). This figure has been modified from5. <a href="https://www.jove.com/files/ftp_upload/53635/53635fig2large.jpg" target="_blank">Please click here to view a larger version of this figure</a>, or visit the full data online (nanotomy.org).

Watch this Scientific Journal Video about Large-scale Scanning Transmission Electron Microscopy Nanotomy of Healthy and Injured Zebrafish Brain at JoVE.com

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 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 ...

large-scale-scanning-transmission-electron-microscopy-nanotomy

Research

JoVE Journal

Developmental Biology

11.0K Views.  UMC Groningen. 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. 

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

Large-scale Zebrafish Embryonic Heart Dissection for Transcriptional Analysis

The zebrafish embryonic heart is composed of only a few hundred cells, representing only a small fraction of the entire embryo. Therefore, to prevent the cardiac transcriptome from being masked by the global embryonic transcriptome, it is necessary to collect sufficient numbers of hearts for further analyses. Furthermore, as zebrafish cardiac development proceeds rapidly, heart collection and RNA extraction methods need to be quick in order to ensure homogeneity of the samples. Here, we present a rapid manual dissection protocol for collecting functional/beating hearts from zebrafish embryos. This is an essential prerequisite for subsequent cardiac-specific RNA extraction to determine cardiac-specific gene expression levels by transcriptome analyses, such as quantitative real-time polymerase chain reaction (RT-qPCR). The method is based on differential adhesive properties of the zebrafish embryonic heart compared with other tissues; this allows for the rapid physical separation of cardiac from extracardiac tissue by a combination of fluidic shear force disruption, stepwise filtration and manual collection of transgenic fluorescently labeled hearts.

To analyse cardiac gene expression profiles during zebrafish heart development, total RNA has to be extracted from isolated hearts. Here, we present a protocol for collecting functional/beating hearts by rapid manual dissection from zebrafish embryos to obtain cardiac-specific mRNA.

The zebrafish embryonic heart is composed of only a few hundred cells, representing only a small fraction of the entire embryo. Therefore, to prevent the ...

Isolation and Culture of Adult Zebrafish Brain-derived Neurospheres

The zebrafish is a highly relevant model organism for understanding the cellular and molecular mechanisms involved in neurogenesis and brain regeneration in vertebrates. However, an in-depth analysis of the molecular mechanisms underlying zebrafish adult neurogenesis has been limited due to the lack of a reliable protocol for isolating and culturing neural adult stem/progenitor cells. Here we provide a reproducible method to examine adult neurogenesis using a neurosphere assay derived from zebrafish whole brain or from the telencephalon, tectum and cerebellum regions of the adult zebrafish brain. The protocol involves, first the microdissection of zebrafish adult brain, then single cell dissociation and isolation of self-renewing multipotent neural stem/progenitor cells. The entire procedure takes eight days. Additionally, we describe how to manipulate gene expression in zebrafish neurospheres, which will be particularly useful to test the role of specific signaling pathways during adult neural stem/progenitor cell proliferation and differentiation in zebrafish.

Here we provide a reproducible method to examine adult neurogenesis using a neurosphere assay derived from the whole brain or from either the telencephalic, tectal or cerebellar regions of the adult zebrafish brain. Additionally, we describe the procedure to manipulate gene expression in zebrafish neurospheres.

The zebrafish is a highly relevant model organism for understanding the cellular and molecular mechanisms involved in neurogenesis and brain regeneration ...

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 ...

Large-Scale Production of Cardiomyocytes from Human Pluripotent Stem Cells Using a Highly Reproducible Small Molecule-Based Differentiation Protocol

Maximizing the benefit of human pluripotent stem cells (hPSCs) for research, disease modeling, pharmaceutical and clinical applications requires robust methods for the large-scale production of functional cell types, including cardiomyocytes. Here we demonstrate that the temporal manipulation of WNT, TGF-&#946;, and SHH signaling pathways leads to highly efficient cardiomyocyte differentiation of single-cell passaged hPSC lines in both static suspension and stirred suspension bioreactor systems. Employing this strategy resulted in ~ 100% beating spheroids, consistently containing &#62; 80% cardiac troponin T-positive cells after 15 days of culture, validated in multiple hPSC lines. We also report on a variation of this protocol for use with cell lines not currently adapted to single-cell passaging, the success of which has been verified in 42 hPSC lines. Cardiomyocytes generated using these protocols express lineage-specific markers and show expected electrophysiological functionalities. Our protocol presents a simple, efficient and robust platform for the large-scale production of human cardiomyocytes.

Here, we present a robust, fast and scalable cardiomyocyte differentiation protocol for human pluripotent stem cells (hPSCs). Cardiomyocytes derived using this large-scale method can provide sufficient cell numbers for their effective use in human cardiovascular disease modeling, high-throughput drug screening, and potentially clinical applications.

Maximizing the benefit of human pluripotent stem cells (hPSCs) for research, disease modeling, pharmaceutical and clinical applications requires robust ...

Dissection of Larval Zebrafish Gonadal Tissue

Although wild zebrafish possess a ZZ/ZW sex-determination system, domesticated zebrafish have lost the sex chromosome. They utilize a polygenic sex determination system, where several genes distributed throughout the genome collectively determine the sex identities of individual fish. Currently, the genes involved in regulating gonad development and how they work remain elusive. Normally, isolating gonadal tissue is the first step to examine the sex developmental processes. Here, we present a procedure to isolate gonadal tissue from 17 dpf (days post fertilization) and 25 dpf zebrafish larvae. The isolated gonadal tissue may be subsequently examined by morphology and gene expression profiling.

Here, we present a protocol for isolating gonadal tissue of larval zebrafish, which will facilitate investigations of zebrafish sex differentiation and maintenance.

Although wild zebrafish possess a ZZ/ZW sex-determination system, domesticated zebrafish have lost the sex chromosome. They utilize a polygenic sex ...

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.

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 ...

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 ...

Adult Zebrafish Injury Models to Study the Effects of Prednisolone in Regenerating Bone Tissue

Zebrafish are able to regenerate various organs, including appendages (fins) after amputation. This involves the regeneration of bone, which regrows within roughly two weeks after injury. Furthermore, zebrafish are able to heal bone rapidly after trepanation of the skull, and repair fractures that can be easily introduced into zebrafish bony fin rays. These injury assays represent feasible experimental paradigms to test the effect of administered drugs on rapidly forming bone. Here, we describe the use of these 3 injury models and their combined use with systemic glucocorticoid treatment, which exerts bone inhibitory and immunosuppressive effects. We provide a workflow on how to prepare for immunosuppressive treatment in adult zebrafish, illustrate how to perform fin amputation, trepanation of calvarial bones, and fin fractures, and describe how the use of glucocorticoids affects both bone forming osteoblasts and cells of the monocyte/macrophage lineage as part of innate immunity in bone tissue.

Here, we describe 3 adult zebrafish injury models and their combined use with immunosuppressive drug treatment. We provide guidance on imaging of regenerating tissues and on detecting bone mineralization therein.

Zebrafish are able to regenerate various organs, including appendages (fins) after amputation. This involves the regeneration of bone, which regrows ...