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>

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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><tbody><tr><td>&lt;strong&gt;Chemicals&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>low melting point agarose</td><td>VWR</td><td>444152G</td><td></td></tr><tr><td>tricaine</td><td>Sigma</td><td>E10521</td><td></td></tr><tr><td>triton-X-100</td><td>Sigma</td><td>X100</td><td></td></tr><tr><td>glutaraldehyde</td><td>Polysciences</td><td>1909</td><td></td></tr><tr><td>sodium cacodylate</td><td>Sigma</td><td>C0250</td><td></td></tr><tr><td>osmium tetroxide</td><td>Electron Microscopy Sciences</td><td>19114</td><td></td></tr><tr><td>potassium ferrocyanide (K&lt;sub&gt;4&lt;/sub&gt;[Fe(CN)]&lt;sub&gt;6&lt;/sub&gt;)</td><td>Merck</td><td>4984</td><td></td></tr><tr><td>ethanol</td><td>VWR</td><td>20821.365</td><td></td></tr><tr><td>uranyl acetate&amp;nbsp;</td><td>Merck</td><td>8473</td><td></td></tr><tr><td>sodium tetraborate</td><td>Merck</td><td>1063808</td><td></td></tr><tr><td>toluidene blue</td><td>Merck</td><td>1273</td><td></td></tr><tr><td>basic fuchsin</td><td>BDH</td><td>340324</td><td></td></tr><tr><td>lead citrate</td><td>BDH</td><td>discontinued</td><td></td></tr><tr><td>&lt;strong&gt;EPON&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>2-dodecenylsuccinicacid anhydride&amp;nbsp;</td><td>Serva</td><td>20755</td><td></td></tr><tr><td>methylnadic anhydride</td><td>Serva</td><td>29452</td><td></td></tr><tr><td>glycid ether 100</td><td>Serva</td><td>21045</td><td></td></tr><tr><td>DMP-30&amp;nbsp;</td><td>Polysciences</td><td>553</td><td></td></tr><tr><td>standard flat embedding mold</td><td>Electron Microscopy Sciences</td><td>70901</td><td></td></tr><tr><td>diamond knife</td><td>Diatome Inc.</td><td></td><td></td></tr><tr><td>copper grids&amp;nbsp;</td><td>Electron Microscopy Sciences</td><td></td><td></td></tr><tr><td>double sided carbon tape&amp;nbsp;</td><td>Electron Microscopy Sciences</td><td></td><td></td></tr><tr><td>scanning EM Zeiss Supra55</td><td>Zeiss</td><td></td><td></td></tr><tr><td>ultramicrotome Leica EM UC7</td><td>Leica</td><td></td><td></td></tr><tr><td>atlas external scan generator&amp;nbsp;</td><td>Fibics</td><td></td><td></td></tr></tbody></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

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 ...

Focussed Ion Beam Milling and Scanning Electron Microscopy of Brain Tissue

This protocol describes how biological samples, like brain tissue, can be imaged in three dimensions using the focussed ion beam/scanning electron microscope (FIB/SEM). The samples are fixed with aldehydes, heavy metal stained using osmium tetroxide and uranyl acetate. They are then dehydrated with alcohol and infiltrated with resin, which is then hardened. Using a light microscope and ultramicrotome with glass knives, a small block containing the region interest close to the surface is made. The block is then placed inside the FIB/SEM, and the ion beam used to roughly mill a vertical face along one side of the block, close to this region. Using backscattered electrons to image the underlying structures, a smaller face is then milled with a finer ion beam and the surface scrutinised more closely to determine the exact area of the face to be imaged and milled. The parameters of the microscope are then set so that the face is repeatedly milled and imaged so that serial images are collected through a volume of the block. The image stack will typically contain isotropic voxels with dimenions as small a 4 nm in each direction. This image quality in any imaging plane enables the user to analyse cell ultrastructure at any viewing angle within the image stack.

This protocol describes how resin embedded brain tissue can be prepared and imaged in the three dimensions in the focussed ion beam, scanning electron microscope.

This protocol describes how biological samples, like brain tissue, can be imaged in three dimensions using the focussed ion beam/scanning electron ...

Neuroscience

Live Imaging of the Zebrafish Embryonic Brain by Confocal Microscopy

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the developing zebrafish brain (Gutzman et al, 2008). Such analysis affords a new understanding of the underlying cell biology required for development of the 3D structure of the vertebrate brain, and significantly increases our ability to study neural tube morphogenesis. The embryonic zebrafish brain is shaped beginning at 18 hours post fertilization (hpf) as the ventricles within the neuroepithelium inflate. By 24 hpf, the initial steps of neural tube morphogenesis are complete. Using the method described here, embryos at the one cell stage are injected with mRNA encoding membrane-targeted green fluorescent protein (memGFP). After injection and incubation, the embryo, now between 18 and 24 hpf, is mounted, inverted, in agarose and imaged by confocal microscopy. Notably, the zebrafish embryo is transparent making it an ideal system for fluorescent imaging. While our analyses have focused on the midbrain-hindbrain boundary and the hindbrain, this method could be extended for analysis of any region in the zebrafish to a depth of 
80-100 &mu;m.

In this video, we demonstrate a method by which to analyze the developing vertebrate brain in live zebrafish embryos at single cell resolution by confocal microscopy. This includes the method by which we inject the single-cell zebrafish embryo and subsequently mount and image the developing brain.

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the ...

Biology

Visualizing the Developing Brain in Living Zebrafish using Brainbow and Time-lapse Confocal Imaging

Development of the vertebrate nervous system requires a precise coordination of complex cellular behaviors and interactions. The use of high resolution in vivo imaging techniques can provide a clear window into these processes in the living organism. For example, dividing cells and their progeny can be followed in real time as the nervous system forms. In recent years, technical advances in multicolor techniques have expanded the types of questions that can be investigated. The multicolor Brainbow approach can be used to not only distinguish among like cells, but also to color-code multiple different clones of related cells that each derive from one progenitor cell. This allows for a multiplex lineage analysis of many different clones and their behaviors simultaneously during development. Here we describe a technique for using time-lapse confocal microscopy to visualize large numbers of multicolor Brainbow-labeled cells over real time within the developing zebrafish nervous system. This is particularly useful for following cellular interactions among like cells, which are difficult to label differentially using traditional promoter-driven colors. Our approach can be used for tracking lineage relationships among multiple different clones simultaneously. The large datasets generated using this technique provide rich information that can be compared quantitatively across genetic or pharmacological manipulations. Ultimately the results generated can help to answer systematic questions about how the nervous system develops.

In vivo imaging is a powerful tool that can be used to investigate the cellular mechanisms underlying nervous system development. Here we describe a technique for using time-lapse confocal microscopy to visualize large numbers of multicolor Brainbow-labeled cells in real time within the developing zebrafish nervous system.

Development of the vertebrate nervous system requires a precise coordination of complex cellular behaviors and interactions. The use of high resolution in ...

Imaging and 3D Reconstruction of Cerebrovascular Structures in Embryonic Zebrafish

Zebrafish are a powerful tool to study developmental biology and pathology in vivo. The small size and relative transparency of zebrafish embryos make them particularly useful for the visual examination of processes such as heart and vascular development. In several recent studies transgenic zebrafish that express EGFP in vascular endothelial cells were used to image and analyze complex vascular networks in the brain and retina, using confocal microscopy. Descriptions are provided to prepare, treat and image zebrafish embryos that express enhanced green fluorescent protein (EGFP), and then generate comprehensive 3D renderings of the cerebrovascular system. Protocols include the treatment of embryos, confocal imaging, and fixation protocols that preserve EGFP fluorescence. Further, useful tips on obtaining high-quality images of cerebrovascular structures, such as removal the eye without damaging nearby neural tissue are provided. Potential pitfalls with confocal imaging are discussed, along with the steps necessary to generate 3D reconstructions from confocal image stacks using freely available open source software.

Imaging of cerebrovascular development in larval zebrafish is described. Techniques to facilitate 3D imaging and modify cerebrovascular development using chemical treatments are also provided.

Zebrafish are a powerful tool to study developmental biology and pathology in vivo.  The small size and relative transparency of zebrafish embryos make ...

In vivo Imaging of Fully Active Brain Tissue in Awake Zebrafish Larvae and Juveniles by Skull and Skin Removal

Understanding the ephemeral changes that occur during brain development and maturation requires detailed high-resolution imaging in space and time at cellular and subcellular resolution. Advances in molecular and imaging technologies have allowed us to gain numerous detailed insights into cellular and molecular mechanisms of brain development in the transparent zebrafish embryo. Recently, processes of refinement of neuronal connectivity that occur at later larval stages several weeks after fertilization, which are for example control of social behavior, decision making or motivation-driven behavior, have moved into focus of research. At these stages, pigmentation of the zebrafish skin interferes with light penetration into brain tissue, and solutions for embryonic stages, e.g., pharmacological inhibition of pigmentation, are not feasible anymore.
Therefore, a minimally invasive surgical solution for microscopy access to the brain of awake zebrafish is provided that is derived from electrophysiological&#160;approaches. In teleosts, skin and soft skull cartilage can be carefully removed by micro-peeling these layers, exposing underlying neurons and axonal tracts without damage. This allows for recording neuronal morphology, including synaptic structures and their molecular contents, and the observation of physiological changes such as Ca2+ transients or intracellular transport events. In addition, interrogation of these processes by means of pharmacological inhibition or optogenetic manipulation is feasible. This brain exposure approach provides information about structural and physiological changes in neurons as well as the correlation and interdependence of these events in live brain tissue in the range of minutes or hours. The technique is suitable for in vivo brain imaging of zebrafish larvae up to 30 days post fertilization, the latest developmental stage tested so far. It, thus, provides access to such important questions as synaptic refinement and scaling, axonal and dendritic transport, synaptic targeting of cytoskeletal cargo or local activity-dependent expression. Therefore, a broad use for this mounting and imaging approach can be anticipated.

Here we present a method to image the zebrafish embryonic brain in vivo upto larval and juvenile stages. This microinvasive procedure, adapted from electrophysiological approaches, provides access to cellular and subcellular details of mature neuron and can be combined with optogenetics and neuropharmacological studies for characterizing brain function and drug intervention.

Understanding the ephemeral changes that occur during brain development and maturation requires detailed high-resolution imaging in space and time at ...

Deep-Tissue Three-Photon Fluorescence Microscopy in Intact Mouse and Zebrafish Brain

Multiphoton microscopy techniques, such as two-photon microscopy (2PM) and three-photon microscopy (3PM), are powerful tools for deep-tissue in vivo imaging with subcellular resolution. 3PM has two major advantages for deep-tissue imaging over 2PM that has been widely used in biology laboratories: (i) longer attenuation length in scattering tissues by employing ~1,300 nm or ~1,700 nm excitation laser; (ii) less background fluorescence generation due to higher-order nonlinear excitation. As a result, 3PM allows high-contrast structural and functional imaging deep within scattering tissues such as intact mouse brain from the cortical layers to the hippocampus and the entire forebrain of adult zebrafish.
Today, laser sources suitable for 3PM are commercially available, enabling the conversion of an existing two-photon (2P) imaging system to a three-photon (3P) system. Additionally, multiple commercial 3P microscopes are available, which makes this technique readily available to biology research laboratories. This paper shows the optimization of a typical 3PM setup, particularly targeting biology groups that already have a 2P setup, and demonstrates intravital 3D imaging in intact mouse and adult zebrafish brains. This protocol covers the full experimental procedure of 3P imaging, including microscope alignment, prechirping of ~1,300 and ~1,700 nm laser pulses, animal preparation, and intravital 3P fluorescence imaging deep in adult zebrafish and mouse brains.

Three-photon microscopy enables high-contrast fluorescence imaging deep in living biological tissues, such as mouse and zebrafish brains, with high spatiotemporal resolution.

Multiphoton microscopy techniques, such as two-photon microscopy (2PM) and three-photon microscopy (3PM), are powerful tools for deep-tissue in vivo ...

In Vivo Whole-Brain Imaging of Zebrafish Larvae

In Vivo Whole-Brain Imaging of Zebrafish Larvae Using Three-Dimensional Fluorescence Microscopy

As a vertebrate model animal, larval zebrafish are widely used in neuroscience and provide a unique opportunity to monitor whole-brain activity at the cellular resolution. Here, we provide an optimized protocol for performing whole-brain imaging of larval zebrafish using three-dimensional fluorescence microscopy, including sample preparation and immobilization, sample embedding, image acquisition, and visualization after imaging. The current protocol enables in vivo imaging of the structure and neuronal activity of a larval zebrafish brain at a cellular resolution for over 1 h using confocal microscopy and custom-designed fluorescence microscopy. The critical steps in the protocol are also discussed, including sample mounting and positioning, preventing bubble formation and dust in the agarose gel, and avoiding motion in images caused by incomplete solidification of the agarose gel and paralyzation of the fish. The protocol has been validated and confirmed in multiple settings. This protocol can be easily adapted for imaging other organs of a larval zebrafish.

Author Spotlight: In Vivo Whole-Brain Imaging of Zebrafish Larvae Using Three-Dimensional Fluorescence Microscopy  

Presented here is a protocol for in vivo whole-brain imaging of larval zebrafish using three-dimensional fluorescence microscopy. The experimental procedure includes sample preparation, image acquisition, and visualization.

As a vertebrate model animal, larval zebrafish are widely used in neuroscience and provide a unique opportunity to monitor whole-brain activity at the ...

Behavior Tracking and Neural Activity Pattern in the Zebrafish Forebrain

Two-Photon Calcium Imaging of Forebrain Activity in Behaving Adult Zebrafish

Adult zebrafish (Danio rerio) exhibit a rich repertoire of behaviors for studying cognitive functions. They also have a miniature brain that can be used for measuring activities across brain regions through optical imaging methods. However, reports on the recording of brain activity in behaving adult zebrafish have been scarce. The present study describes procedures to perform two-photon calcium imaging in the dorsal forebrain of adult zebrafish. We focus on steps to restrain adult zebrafish from moving their heads, which provides stability that enables laser scanning imaging of the brain activity. The head-restrained animals can freely move their body parts and breathe without aids. The procedure aims to shorten the time of head restraint surgery, minimize brain motion, and maximize the number of neurons recorded. A setup for presenting an immersive visual environment during calcium imaging is also described here, which can be used to study neural correlates underlying visually triggered behaviors.

Author Spotlight: Advancements in Adult Zebrafish Brain Research 

Here, we present a protocol to perform two-photon calcium imaging in the dorsal forebrain of adult zebrafish.

Adult zebrafish (Danio rerio) exhibit a rich repertoire of behaviors for studying cognitive functions. They also have a miniature brain that can be used ...

Investigating a Zebrafish Larval Brain Injury Using Large-Scale Scanning Transmission Electron Microscopy

Source: Kuipers, J. et al., <a href="https://app.jove.com/v/53635">Large-scale Scanning Transmission Electron Microscopy (Nanotomy) of Healthy and Injured Zebrafish Brain.</a>&#160;J. Vis. Exp. (2016).
This video demonstrates a method to investigate zebrafish larval brain injury using large-scale scanning transmission electron microscopy (STEM), a technique that combines elements of both scanning electron microscopy (SEM) and transmission electron microscopy (TEM) to produce high-resolution images. It outlines the steps involved in preparing samples and the acquisition of high-resolution STEM images to identify brain injury.

1. Sample Preparation

	Fixation and Embedding
	
		Fixation
		
			Anaesthetize fish larvae in tricaine (0.003%) added to the E3 (5 mM NaCl, 0.17 mM KCl, ...