eoe sub articles

EoE Sub Articles

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,&#160;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 electron microscopy (EM) after acquiring&#160;in vivo&#160;imaging data using confocal or 2 photon imaging in 1.8% agarose as described,&#160;fix larvae in agarose using 4% paraformaldehyde (PFA) in phosphate buffered saline (PBS) containing Triton-X-100 (0.05%).</li>
			<li>Cut larvae from agarose and process for large-scale EM.</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 the 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&#160;(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 the 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;&#160;Figure 1E)</li>
			<li>Use easily identifiable anatomic structures in and around the brain, including olfactory pits, eyes, and 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 the 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.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
2. Image Acquisition
<ol>
	<li>Large Scale Scanning Transmission Electron Microscopy (STEM) 
	Note: We use scanning EM with an external scan generator capable of acquiring multiple large fields of view at high resolution&#160;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 ensure it is on the section area.</li>
			<li>Switch to STEM detection (gain medium and all quadrants set on &#39;minus&#39;), acquire an image, and select the area to be scanned.</li>
			<li>Raise the acceleration voltage to 21 kV and wait for stabilization of the beam (stable image again), 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 the aperture to 120 &#181;m (to keep the proper dynamic range, the detector gains have to be reduced by 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 field of view (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 on the sample.</li>
			<li>Align the aperture using the wobbler (at ~ 40,000X magnification).</li>
			<li>While in focus, select the lightest area/feature and set the 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 the option &#34;autofocus on previous tile&#34;, use the large scale acquisition software autofocus with default company settings, check the 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 the 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 the external scan generator again and press &#34;continue.&#34;</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Chemicals</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&#160;</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>osmiumtetroxide</td>
			<td>Electron Microscopy Sciences</td>
			<td>19114</td>
			<td></td>
		</tr>
		<tr>
			<td>potassiumferrocyanide (K4[Fe(CN)]6)</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&#160;</td>
			<td>Merck</td>
			<td>8473</td>
			<td></td>
		</tr>
		<tr>
			<td>sodiumtetraborate</td>
			<td>Merck</td>
			<td>1063808</td>
			<td></td>
		</tr>
		<tr>
			<td>Tolduidene 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>EPON</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>2-dodecenylsuccinicacid anhydride&#160;</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&#160;</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&#160;</td>
			<td>Electron Microscopy Sciences</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>double sided carbon tape&#160;</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&#160;</td>
			<td>Fibics</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

<img alt="Figure 1" src="/files/ftp_upload/22992/22992_Figure1.jpg" /> 
Figure 1: Workflow of Large-scale Tissue Electron Microscopy on Zebrafish Brains. (A)&#160;7 day-old zebrafish larva.&#160;(B)&#160;Zebrafish larval heads embedded side by side in epoxy resin.&#160;(C)&#160;Glass knife for semi -thin sectioning and orienting the specimen.&#160;(D)&#160;Representative semithin toluidine blue stained section showin...

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

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, ...

Take resin-embedded, chemically fixed ultrathin brain sections from healthy and brain-injured zebrafish larvae mounted on a one-hole grid.&#160;
The sections are stained with heavy metals to enhance image contrast during microscopic imaging.
Place the grid in a sample holder and transfer the holder to the microscopy chamber.
Align the grid under the electron gun, which generates an electron beam.
An array of electromagnetic lenses and coils focuses the electron beam onto the sections.
Move the sample holder to scan the sections in a raster pattern.
As the electron beam passes through the tissue sections, the stained electron-dense regions scatter more electrons, while the unstained regions transmit the electrons.
The detector collects these electron signals, creating high-resolution brain images.
Compared to the control, the injured brain shows phagocytic microglia containing numerous intracellular inclusions, a characteristic feature of injury-induced neurodegeneration.

investigating-zebrafish-larval-brain-injury-using-large-scale

Research

JoVE Encyclopedia of Experiments

Neuroscience

Neuropathology

Trauma

93 ビュー. 出典:Kuipers, J. et al., Large-scale Scanning Transmission Electron Microscopy (Nanotomy) of Healthy and Injured Zebrafish Brain.J. Vis. Exp. (2016). このビデオは、走査型電子顕微鏡(SEM)と透過型電子顕微鏡(TEM)の両方の要素を組み合わせて高解像度の画像を生成する技術である大規模な走査型透過型電子顕微鏡(STEM)を使用して、ゼブラフィッシュの幼生の脳損傷を調査する方法を示しています。サンプルの?...

ビデオ: 大規模走査型透過電子顕微鏡によるゼブラフィッシュ幼生脳損傷の検討 - 概念

Visualization and Quantitative Analysis of Embryonic Angiogenesis in Xenopus tropicalis

Blood vessels supply oxygen and nutrients throughout the body, and the formation of the vascular network is under tight developmental control. The efficient in vivo visualization of blood vessels and the reliable quantification of their complexity are key to understanding the biology and disease of the vascular network. Here, we provide a detailed method to visualize blood vessels with a commercially available fluorescent dye, human plasma acetylated low density lipoprotein DiI complex (DiI-AcLDL), and to quantify their complexity in Xenopus tropicalis. Blood vessels can be labeled by a simple injection of DiI-AcLDL into the beating heart of an embryo, and blood vessels in the entire embryo can be imaged in live or fixed embryos. Combined with gene perturbation by the targeted microinjection of nucleic acids and/or the bath application of pharmacological reagents, the roles of a gene or of a signaling pathway on vascular development can be investigated within one week without resorting to sophisticated genetically engineered animals. Because of the well-defined venous system of Xenopus and its stereotypic angiogenesis, the sprouting of pre-existing vessels, vessel complexity can be quantified efficiently after perturbation experiments. This relatively simple protocol should serve as an easily accessible tool in diverse fields of cardiovascular research.

Visualization and Quantitative Analysis of Embryonic Angiogenesis in Xenopus tropicalis

This protocol demonstrates a fluorescence-based method to visualize the vasculature and to quantify its complexity in Xenopus tropicalis. Blood vessels can be imaged minutes after the injection of a fluorescent dye into the beating heart of an embryo after genetic and/or pharmacological manipulations to study cardiovascular development in vivo.

の胚性血管新生の可視化と定量分析<em&gt;ツメガエル</em

Blood vessels supply oxygen and nutrients throughout the body, and the formation of the vascular network is under tight developmental control. The ...

JoVE Journal

発生生物学

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.

ゼブラフィッシュの網膜におけるタンパク質局在を解決する相関超解像および電子顕微鏡観察

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

Analysis of Brain Mitochondria Using Serial Block-Face Scanning Electron Microscopy

Human brain is a high energy consuming organ that mainly relies on glucose as a fuel source. Glucose is catabolized by brain mitochondria via glycolysis, tri-carboxylic acid (TCA) cycle and oxidative phosphorylation (OXPHOS) pathways to produce cellular energy in the form of adenosine triphosphate (ATP). Impairment of mitochondrial ATP production causes mitochondrial disorders, which present clinically with prominent neurological and myopathic symptoms. Mitochondrial defects are also present in neurodevelopmental disorders (e.g. autism spectrum disorder) and neurodegenerative disorders (e.g. amyotrophic lateral sclerosis, Alzheimer&#39;s and Parkinson&#39;s diseases). Thus, there is an increased interest in the field for performing 3D analysis of mitochondrial morphology, structure and distribution under both healthy and disease states. The brain mitochondrial morphology is extremely diverse, with some mitochondria especially those in the synaptic region being in the range of &#60;200 nm diameter, which is below the resolution limit of traditional light microscopy. Expressing a mitochondrially-targeted green fluorescent protein (GFP) in the brain significantly enhances the organellar detection by confocal microscopy. However, it does not overcome the constraints on the sensitivity of detection of relatively small sized mitochondria without oversaturating the images of large sized mitochondria. While serial transmission electron microscopy has been successfully used to characterize mitochondria at the neuronal synapse, this technique is extremely time-consuming especially when comparing multiple samples. The serial block-face scanning electron microscopy (SBFSEM) technique involves an automated process of sectioning, imaging blocks of tissue and data acquisition. Here, we provide a protocol to perform SBFSEM of a defined region from rodent brain to rapidly reconstruct and visualize mitochondrial morphology. This technique could also be used to provide accurate information on mitochondrial number, volume, size and distribution in a defined brain region. Since the obtained image resolution is high (typically under 10 nm) any gross mitochondrial morphological defects may also be detected.

Mitochondrial visualization and analysis from mammalian brain tissue is a challenging task. Here, we describe how three dimensional (3D) reconstruction analysis from the serial block-face scanning electron microscopy (SBFSEM) can be used to gain insights on the morphological and volumetric analysis of this critical energy generating organelle.

シリアルブロック・フェイス走査型電子顕微鏡を用いて脳ミトコンドリアの解析

Human brain is a high energy consuming organ that mainly relies on glucose as a fuel source. Glucose is catabolized by brain mitochondria via glycolysis, ...

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

記録

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