This research was supported by the KBRI basic research program through Korea Brain Research Institute funded by Ministry of Science and ICT (24-BR-01-03). TMEM192-V5-APEX2 plasmids were kindly provided by Hyun-Woo Rhee (Seoul National University). TEM data were acquired at Brain Research Core Facilities in KBRI.

The step-by-step workflow for sample preparation is shown in Figure 1.
1. Cell culture and transfection
<ol>
	<li>Culture HeLa cells at a density of 1 &#215; 105 cells in 35 mm glass grid-bottomed culture dishes containing DMEM with 10% fetal bovine serum and 100 U/mL of penicillin and streptomycin at 37 &#176;C.</li>
	<li>The day after seeding the cells, co-transfect the cells at 30%-40% confluency with TMEM192-V5-APEX2 and Mito-mEosEM plasmids using the transfection reagent according to the manufacturer&#39;s instructions(1 &#181;g each of TMEM192-V5-APEX2 and Mito-mEosEM plasmid DNA with 3 &#181;L of transfection reagent). After transfection, treat the cells with DMSO or U18666A (2 &#181;g/mL), and Bafilomycin A1 (100 nM) for 18 h.</li>
</ol>
2. Confocal microscopic imaging
<ol>
	<li>After treating the cells with Bafilomycin and U18666A for 18 h, remove the culture media and immediately add 250 &#181;L of warm (30 &#176;C-37 &#176;C) fixative solution (2.5% glutaraldehyde and 2% paraformaldehyde in 0.1 M sodium cacodylate solution [pH 7.0]) by gentle pipetting. Immediately remove the fixative, replace it with 1.5 mL of fresh fixative, and incubate on ice for 30 min. Then, wash the cells for 3 x 10 min in 1 mL of ice-cold 0.1 M sodium cacodylate buffer. 
	NOTE: Aldehyde fumes are toxic. All experiments should be conducted in a ventilated fume hood.</li>
	<li>Add 1 mL of cold 20 mM glycine solution to quench the aldehyde and incubate on ice for 10 min. Wash 3 x 10 min in cold 0.1 M sodium cacodylate buffer.</li>
	<li>Prepare a fresh solution containing 300 &#181;M Amplex-Red by adding 10 mM H2O2 to a 0.1 M cacodylate solution. Add 500 &#181;L of Amplex-Red solution to the sample dish and incubate for 5 min on ice.</li>
	<li>Remove the Amplex-Red solution and rinse 3 x 10 min with 0.1 M sodium cacodylate buffer.</li>
	<li>After Amplex-Red staining, use a confocal microscope to image the area around the cells of interest in 3 x 3 tile scan mode. Capture both differential interference contrast (DIC) and fluorescence signal to identify and record the grid and letters on the grid dish using a 40x (or 60x) objective. 
	NOTE: These overview images serve as a map for identifying specific target cells in the plastic-embedded specimens.</li>
	<li>Image the target cells at high magnification using a 40x (or 60x) objective and acquire a Z-stack image. 
	NOTE: This high-magnification image is superimposed on an electron microscope image to create a correlative image.</li>
</ol>
3. Sample preparation for EM block
<ol>
	<li>After LM imaging, add 1 mL of 1% reduced osmium tetroxide (prepared by containing 3% potassium ferrocyanide in 0.3 M cacodylate buffer with an equal volume of 2% aqueous osmium tetroxide) for postfixation at 4 &#176;C for 1 h, and then rinse for 3 x 10 min with distilled water at 4 &#176;C. 
	NOTE: Osmium tetroxide fumes are toxic. All experiments should be performed in a ventilated fume hood.</li>
	<li>Rinse 3 x 10 min with distilled water at room temperature, then dehydrate in a graded ethanol series for 15 min each time (50%, 60%, 70%, 80%, 90%, 95%, 100%, and 100%) at room temperature.</li>
	<li>Mix ethanol with epoxy resin in volume ratios of 3:1, 1:1, and 1:3, preparing a total of 300 &#181;L of each mixture. Pour each epoxy resin into the dish, allowing it to incubate at room temperature for 1 h. Then, use the 100% epoxy resin for overnight impregnation.</li>
	<li>Prepare the embedding capsule or PCR tube by filling it with fresh epoxy resin, place the tube or capsule upside down on the region of interest, and place it in an oven at 60 &#176;C for 48 h to polymerize.</li>
</ol>
4. Sectioning and TEM imaging
<ol>
	<li>After the epoxy resin has completely polymerized, separate the bottom glass from the polymer block by immersing it in liquid nitrogen.</li>
	<li>Place the specimen block in the sample holder of the ultramicrotome and set it in the trimming block. Use a razor blade to trim the resin into a trapezoidal or rectangular shape around the object of interest.</li>
	<li>Place the trimmed specimen block in the sample holder and adjust the diamond blade. Ensure that the leading edge of the specimen block is perfectly parallel to the edge of the diamond blade.</li>
	<li>Using an ultramicrotome, cut ultrathin sections approximately 60-70 nm from the flat-embedded cells. Collect the serial sections on a formvar-coated slot grid and dry them on a hot plate.</li>
	<li>Stain the section with the contrast stain for 2 min and lead citrate for 1 min.</li>
	<li>Place the grid in the TEM sample holder for observation. Identify the target cell of interest based on the DIC image from the light microscope. Image the entire cell area using tiled, overlapping images at 1,700x magnification.</li>
</ol>
5. Image stitching using imageJ Fiji.
<ol>
	<li>To begin image stitching, start imageJ Fiji25 and click on File | New | TrakEM2 (blank) | Select storage folder to create a new TrakEM226 project (Figure 2A1).</li>
	<li>Open the tile image dataset by dragging and dropping the raw tile image data into the workspace (Figure 2A2).</li>
	<li>Click the right mouse button and select Align | Montage all images in this layer | elastic (non-linear block correspondences), and then click OK to accept the default values remaining parameters (Figure 2A3).</li>
	<li>Click the right mouse button and select Export | Make flat image to create a stitched image, then click File | Save As to save the image.</li>
</ol>
6. Resizing the fluorescence image
<ol>
	<li>Open the photo enlargement software (see Table of Materials). Click on File | Open and select the fluorescence image.</li>
	<li>Enter the width and height to be the same size as the electron microscope image, and select the algorithm to be applied in the resize method (Figure 2B1).</li>
	<li>Click on File | Save AS to save the resized image.</li>
</ol>
7. Landmark-based image registration with the Big Warp plugin
<ol>
	<li>Open ImageJ Fiji and drag and drop the resized fluorescence micrograph (FM image) and the stitched electron micrograph (EM image).</li>
	<li>Click on Plugins | BigData Viewer | Big Wrap to start the registration , then click on the dropdown menu to select the FM image as the moving image and the EM image as the target image (Figure 2C1). 
	NOTE: Each FM and EM image will have three new popup windows (BigWrap moving Image, BigWrap fixed image, and Landmarks) to display landmarks.</li>
	<li>Compare the FM and EM images using functions such as zoom (mouse wheel), image rotation (moving the mouse while holding down the left button), and move (moving the mouse while holding down the right button), and then press the space bar on the keyboard to switch Landmark mode on, and press the left mouse button to mark at least three landmarks identified in the FM and EM images (Figure 2C2, pink dots and red arrow).</li>
	<li>After marking all the identified landmarks, go to File | Export Moving Image | OK and a new popup window will appear with the transformed FM image (Figure 2C3). 
	NOTE: The newly created popup window is an FM image transformed based on the basis of the landmark. Review the alignment and make adjustments, if necessary, by adding or modifying landmarks and recalculating the transformation.</li>
	<li>Click on File | Save As to save the transformed FM image.</li>
</ol>
8. Overlay of CLEM images
<ol>
	<li>In ImageJ, drag and drop the transformed FM image and the stitched EM image.</li>
	<li>Click on Image | Type | 8-Bit Color to set the same image type for both the LM and EM images (Figure 2D1). 
	NOTE: The EM image must be converted twice in the following order: 8-bit -&#62; RGB Color -&#62; 8-bit Color.</li>
	<li>Click on Image | Color &#62;| Merge Channels to combine the images into a composite image by selecting the FM and EM images from the dropdown menu, respectively (Figure 2D2).</li>
	<li>Click on File | Save As to save the CLEM image.</li>
</ol>

Correlative Microscopy Reveals Stress-Induced Mitochondria-Lysosome Interactions

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The authors have no conflicts of interest to disclose.

This protocol describes dual-color CLEM to observe the interaction of mitochondria and lysosomes in fixed cells using genetically encoded mEosEM and APEX2 tags. Over the past few decades, advances in microscopy have greatly improved the ability to observe changes in organelle networks32,33, and the study of organelle interactions has become increasingly interesting. Super-resolution microscopy has been used to observe changes in highly dynamic contacts between mi...

Mitochondria and lysosomes are the principal membrane-bound organelles that are essential for the maintenance of cellular homeostasis. Mitochondria are highly dynamic cellular organelles modulated by fission and fusion events1. In general, mitochondria are referred to as the powerhouse of the cell and are known for their important role in oxidative phosphorylation, ATP production, and metabolite storage such as&#160;calcium2. In addition to their role in energy metabolism, mitochondria are involved in a number of other cellular signaling processes, including apoptosis3,4, calcium signaling5,6, and lipid metabolism7,8. Similarly, lysosomes are highly dynamic cellular organelles that play multiple roles as cellular recycling centers to maintain cellular cleanliness through protein degradation, autophagy, endocytosis, and phagocytosis9. Apart from their role in degradation, lysosomes also play roles in cellular physiology such as nutrient sensing, homeostasis10, lipid metabolism11, and cellular dedifferentiation12.
Despite the existence of numerous studies on the functions of individual cellular organelles, our recent research has concentrated on the less-understood crosstalk between mitochondria and lysosomes, which has been demonstrated to influence both cellular physiology and pathology. Contacts between mitochondria and lysosomes have been observed in various cell types, including cancer cells, neurons, and induced pluripotent stem cell (iPSC)-derived cells13, and they are mainly observed under cellular stress or in neurodegenerative diseases such as Parkinson&#39;s disease13,14 and Charcot-Marie-Tooth disease15. The contact between mitochondria and lysosomes plays an important role in maintaining metabolic homeostasis through the exchange of ions (e.g., Ca2+)16, cholesterol17, and iron18 between the two organelles. Furthermore, the contact between mitochondria and lysosomes plays a crucial role in the quality control of cellular organelles. For example, damaged mitochondria can be targeted for degradation through the process of mitophagy, in which they are engulfed by autophagosomes and delivered to lysosomes for degradation. Consequently, the removal of dysfunctional mitochondria can be facilitated before they cause further damage to the cell or to normal mitochondria. This interaction between mitochondria and lysosomes is crucial for maintaining cellular homeostasis and preventing the accumulation of dysfunctional organelles. Thus, only damaged mitochondria are selectively removed, allowing healthy mitochondria to function optimally within the cell.
Correlative light and electron microscopy (CLEM) is an imaging technique that has recently received attention as a research method by combining the advantages of light microscopy (LM) and electron microscopy (EM) to provide detailed structural information at different scales. In CLEM, a sample is first imaged using LM to identify specific regions of interest, followed by EM sample preparation steps such as fixation, dehydration, and embedding. The same sample is then observed under an electron microscope. Although live-cell CLEM methods have been reported recently19,20,21,22, it remains challenging to accurately combine optical images of highly dynamic biological samples with electron microscopy images of fixed cells into a single plane. For highly dynamic organelles like mitochondria and lysosomes, their precise localization can change as they are observed live under the light microscope, and they continue to move even during the short delay time required for fixation.
Therefore, in this protocol, two different molecular tags that remain active even within fixative solutions were employed to visualize the interaction between mitochondria and lysosomes, which play an important role in maintaining metabolic homeostasis through the exchange of ions (e.g., Ca2+)16, cholesterol17, and iron18 between these organelles. The constructs TMEM192-V5-APEX2 and Mito-mEosEM were used to observe the interactions of mitochondria and lysosomes in cultured cells at both the EM and LM levels. For mitochondria, the mEosEM tag was selected for its ability to preserve the fluorescence signal during EM sample preparation in a fixative solution and OsO423. For lysosomes, the APEX2 tag, a protein of approximately 28 kDa, was used to study the localization of a specific protein at both EM and LM levels24. In the presence of hydrogen peroxide, the APEX2&#39;s substrate, Amplex-Red, forms an insoluble polymer that can be observed by confocal microscopy at the tag-targeted location. In this protocol, dual-color CLEM was used to visualize the structure of the interactions between lysosomes and mitochondria in fixed cells.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Chemicals and reagents</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>25% Glutaraldehyde</td>
			<td>Electron Microscopy Sciences</td>
			<td>16200</td>
			<td>Aldehyde fumes are extremely toxic. Use only in fume hood.</td>
		</tr>
		<tr>
			<td>30% Hydrogen peroxide solution</td>
			<td>Merck</td>
			<td>107210</td>
			<td></td>
		</tr>
		<tr>
			<td>4% Paraformaldehyde solution</td>
			<td>Biosesang</td>
			<td>PC2031-100-00</td>
			<td>ldehyde fumes are extremely toxic. Use only in fume hood.</td>
		</tr>
		<tr>
			<td>Amplex Red</td>
			<td>ThermoFisher</td>
			<td>A12222</td>
			<td></td>
		</tr>
		<tr>
			<td>Epon 812</td>
			<td>Electron Microscopy Sciences</td>
			<td>14120</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethanol</td>
			<td>Merck</td>
			<td>100983</td>
			<td></td>
		</tr>
		<tr>
			<td>Fugene HD</td>
			<td>Promega</td>
			<td>E2311</td>
			<td>Plasmid transfection</td>
		</tr>
		<tr>
			<td>Glycine</td>
			<td>SIGMA</td>
			<td>G8898</td>
			<td></td>
		</tr>
		<tr>
			<td>Lead Citrate 3%</td>
			<td>Electron Microscopy Sciences</td>
			<td>22410</td>
			<td></td>
		</tr>
		<tr>
			<td>Osmium tetroxide 4 % aqueous solution</td>
			<td>Electron Microscopy Sciences</td>
			<td>16320</td>
			<td>Very toxic. Use only in fume hood.</td>
		</tr>
		<tr>
			<td>Potassium hexacyanoferrate(II) trihydrate&#160;</td>
			<td>SIGMA</td>
			<td>P3289</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium cacodylate trihydrate</td>
			<td>Sigma-Aldrich</td>
			<td>C0250-500G</td>
			<td></td>
		</tr>
		<tr>
			<td>Uranyl acetate</td>
			<td>Electron Microscopy Sciences</td>
			<td>22400</td>
			<td></td>
		</tr>
		<tr>
			<td>UranyLess EM stain</td>
			<td>Electron Microscopy Sciences</td>
			<td>22409</td>
			<td></td>
		</tr>
		<tr>
			<td>Plasmid construction</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Mito-mEosEM</td>
			<td>addgene</td>
			<td>132706</td>
			<td>DOI:&#160;10.1016/j.chembiol.2024.02.007</td>
		</tr>
		<tr>
			<td>TMEM192-V5-APEX2</td>
			<td>Sharma N.&#160; et al.</td>
			<td>N/A</td>
			<td>DOI:&#160;10.1016/j.chembiol.2024.02.007</td>
		</tr>
		<tr>
			<td>Tools</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>0.22 um syringe filter</td>
			<td>Sartorius</td>
			<td>16534</td>
			<td></td>
		</tr>
		<tr>
			<td>35 mm Gridded coverslip dish</td>
			<td>Mattek</td>
			<td>P35G-1.5-14-CGRD</td>
			<td></td>
		</tr>
		<tr>
			<td>BEEM Embedding Capsules</td>
			<td>Ted Pella</td>
			<td>130</td>
			<td></td>
		</tr>
		<tr>
			<td>Formvar Supported Single slot copper grid</td>
			<td>Ted Pella</td>
			<td>01705</td>
			<td></td>
		</tr>
		<tr>
			<td>Diamond knife</td>
			<td>DiATOME</td>
			<td>DU4530</td>
			<td></td>
		</tr>
		<tr>
			<td>Ultra-microtome</td>
			<td>Leica</td>
			<td>ARTOS 3D</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscopes</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Inverted confocal microscopy</td>
			<td>Nikon</td>
			<td>A1 Rsi/Ti-E</td>
			<td></td>
		</tr>
		<tr>
			<td>Transmission electron microscopy</td>
			<td>FEI</td>
			<td>Tecnai G2</td>
			<td></td>
		</tr>
		<tr>
			<td>Software and Algorithms</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Fiji/Image J</td>
			<td>NIH / open source</td>
			<td></td>
			<td>https://imagej.nih.gov/ij/</td>
		</tr>
		<tr>
			<td>ImageJ BigWarp software package</td>
			<td>NIH / open source</td>
			<td></td>
			<td>https://imagej.net/plugins/bigwarp</td>
		</tr>
		<tr>
			<td>Photozoom Pro 8&#160;</td>
			<td>BenVista</td>
			<td></td>
			<td>https://www.benvista.com/photozoompro 
			 
			Alternative software 
			Image Resizer : https://imageresizer.com/ 
			Gigapixel AI : https://www.topazlabs.com/gigapixel 
			ON1 resize : https://www.on1.com/products/resize-ai/</td>
		</tr>
	</tbody>
</table>

dual-color correlative light and electron microscopy for the visualization of interactions between mitochondria and lysosomes

Cellular organelles, such as mitochondria and lysosomes, display dynamic structures. Despite the higher resolution of transmission electron microscopy for structural analysis, light microscopy is essential for the visualization of dynamic organelles by target-specific labeling. The following protocol describes a method that combines dual-color correlative light and electron microscopy (CLEM) to observe the interactions between mitochondria and lysosomes. In this study, mitochondria were labeled with mEosEM (Mito-mEosEM) and lysosomes with TMEM192-V5-APEX2. The results obtained from CLEM images enable us to observe the changes in the interactions between mitochondria and lysosomes under external stress conditions. Treatment with bafilomycin (BFA), which inhibits lysosomal function, resulted in an increase in contact between mitochondria and lysosomes, leading to the formation of fragmented mitochondria trapped inside lysosomes. Conversely, treatment with U18666A, which inhibits cholesterol export from lysosomes, caused lysosomes to be surrounded by mitochondria, indicating a distinct form of interaction. This study presents an effective method for observing the interactions between mitochondria and lysosomes in fixed cells. Furthermore, CLEM imaging with dual-color probes offers a powerful tool for future investigations of organelle dynamics and their implications for cell function and pathology.

To visualize the interactions between mitochondria and lysosomes using this dual-color CLEM protocol, we employed Mito-mEosEM and TMEM192-V5-APEX2. Fixed cells were first imaged using a light microscope, followed by EM sampling and imaging. The results of the correlated images are shown in Figure 3A,B and Figure 4A. Following the protocol described above, we checked whether we could observe the changes in the interaction between mitochondria and...

Dual-color Correlative Light and Electron Microscopy for the Visualization of Interactions between Mitochondria and Lysosomes

This protocol describes a mechanism for using correlative light and electron microscopy to visualize the interaction of mitochondria and lysosomes labeled with mEosEM and APEX2, respectively.

Cellular organelles, such as mitochondria and lysosomes, display dynamic structures. Despite the higher resolution of transmission electron microscopy for ...

dual-color-correlative-light-electron-microscopy-for-visualization

Research

JoVE Journal

Neuroscience

473 Views.  Korea Brain Research Institute. This protocol describes a mechanism for using correlative light and electron microscopy to visualize the interaction of mitochondria and lysosomes labeled with mEosEM and APEX2, respectively.

Video: Dual-color Correlative Light and Electron Microscopy for the Visualization of Interactions between Mitochondria and Lysosomes

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 ...

Visualization of Proprioceptors in Drosophila Larvae and Pupae

Proprioception is the ability to sense the motion, or position, of body parts by responding to stimuli arising within the body. In fruitflies and other insects proprioception is provided by specialized sensory organs termed chordotonal organs (ChOs) 2. Like many other organs in Drosophila, ChOs develop twice during the life cycle of the fly. First, the larval ChOs develop during embryogenesis. Then, the adult ChOs start to develop in the larval imaginal discs and continue to differentiate during metamorphosis.
The development of larval ChOs during embryogenesis has been studied extensively 10,11,13,15,16. The centerpiece of each ChO is a sensory unit composed of a neuron and a scolopale cell. The sensory unit is stretched between two types of accessory cells that attach to the cuticle via specialized epidermal attachment cells 1,9,14. When a fly larva moves, the relative displacement of the epidermal attachment cells leads to stretching of the sensory unit and consequent opening of specific transient receptor potential vanilloid (TRPV) channels at the outer segment of the dendrite 8,12. The elicited signal is then transferred to the locomotor central pattern generator circuit in the central nervous system.
Multiple ChOs have been described in the adult fly 7. These are located near the joints of the adult fly appendages (legs, wings and halters) and in the thorax and abdomen. In addition, several hundreds of ChOs collectively form the Johnston's organ in the adult antenna that transduce acoustic to mechanical energy 3,5,17,4.
In contrast to the extensive knowledge about the development of ChOs in embryonic stages, very little is known about the morphology of these organs during larval stages. Moreover, with the exception of femoral ChOs 18 and Johnston's organ, our knowledge about the development and structure of ChOs in the adult fly is very fragmentary.
Here we describe a method for staining and visualizing ChOs in third instar larvae and pupae. This method can be applied together with genetic tools to better characterize the morphology and understand the development of the various ChOs in the fly.

Visualization of Proprioceptors in Drosophila Larvae and Pupae

A method to immunostain and visualize chordotonal organs in larvae and pupae of Drosophila melanogaster is described.

Proprioception is the ability to sense the motion, or position, of body parts by responding to stimuli arising within the body. In fruitflies and other ...

Genetic Manipulation of Cerebellar Granule Neurons In Vitro and In Vivo to Study Neuronal Morphology and Migration

Developmental events in the brain including neuronal morphogenesis and migration are highly orchestrated processes. In vitro and in vivo analyses allow for an in-depth characterization to identify pathways involved in these events. Cerebellar granule neurons (CGNs) that are derived from the developing cerebellum are an ideal model system that allows for morphological analyses. Here, we describe a method of how to genetically manipulate CGNs and how to study axono- and dendritogenesis of individual neurons. With this method the effects of RNA interference, overexpression or small molecules can be compared to control neurons. In addition, the rodent cerebellar cortex is an easily accessible in vivo system owing to its predominant postnatal development. We also present an in vivo electroporation technique to genetically manipulate the developing cerebella and describe subsequent cerebellar analyses to assess neuronal morphology and migration.

Genetic Manipulation of Cerebellar Granule Neurons In Vitro and In Vivo to Study Neuronal Morphology and Migration

Neuronal morphogenesis and migration are crucial events underlying proper brain development. Here, we describe methods to genetically manipulate cultured cerebellar granule neurons and the developing cerebellum for the assessment of morphology and migratory characteristics of neurons.

Developmental events in the brain including neuronal morphogenesis and migration are highly orchestrated processes. In vitro and in vivo analyses allow ...

Correlative Light and Electron Microscopy to Study Microglial Interactions with &#946;-Amyloid Plaques

A detailed protocol is provided here to identify amyloid A&#946; plaques in brain sections from Alzheimer&#39;s disease mouse models before pre-embedding immunostaining (specifically for ionized calcium-binding adapter molecule 1 (IBA1), a calcium binding protein expressed by microglia) and tissue processing for electron microscopy (EM). Methoxy-X04 is a fluorescent dye that crosses the blood-brain barrier and selectively binds to &#946;-pleated sheets found in dense core A&#946; plaques. Injection of the animals with methoxy-X04 prior to sacrifice and brain fixation allows pre-screening and selection of the plaque-containing brain sections for further processing with time-consuming manipulations. This is particularly helpful when studying early AD pathology within specific brain regions or layers that may contain very few plaques, present in only a small fraction of the sections. Post-mortem processing of tissue sections with Congo Red, Thioflavin S, and Thioflavin T (or even with methoxy-X04) can label &#946;-pleated sheets, but requires extensive clearing with ethanol to remove excess dye and these procedures are incompatible with ultrastructural preservation. It would also be inefficient to perform labeling for A&#946; (and other cellular markers such as IBA1) on all brain sections from the regions of interest, only to yield a small fraction containing A&#946; plaques at the right location. Importantly, A&#946; plaques are still visible after tissue processing for EM, allowing for a precise identification of the areas (generally down to a few square millimeters) to examine with the electron microscope.

This article describes a protocol for visualizing amyloid A&#946; plaques in Alzheimer&#39;s disease mouse models using methoxy-X04, which crosses the blood-brain barrier and selectively binds to &#946;-pleated sheets found in dense core A&#946; plaques. It allows pre-screening of plaque-containing tissue sections prior to immunostaining and processing for electron microscopy.

A detailed protocol is provided here to identify amyloid A&#946; plaques in brain sections from Alzheimer&#39;s disease mouse models before pre-embedding ...

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.

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, ...

Visualization of Motor Axon Navigation and Quantification of Axon Arborization In Mouse Embryos Using Light Sheet Fluorescence Microscopy

Spinal motor neurons (MNs) extend their axons to communicate with their innervating targets, thereby controlling movement and complex tasks in vertebrates. Thus, it is critical to uncover the molecular mechanisms of how motor axons navigate to, arborize, and innervate their peripheral muscle targets during development and degeneration. Although transgenic Hb9::GFP mouse lines have long served to visualize motor axon trajectories during embryonic development, detailed descriptions of the full spectrum of axon terminal arborization remain incomplete due to the pattern complexity and limitations of current optical microscopy. Here, we describe an improved protocol that combines light sheet fluorescence microscopy (LSFM) and robust image analysis to qualitatively and quantitatively visualize developing motor axons. This system can be easily adopted to cross genetic mutants or MN disease models with Hb9::GFP lines, revealing novel molecular mechanisms that lead to defects in motor axon navigation and arborization.

Here, we describe a protocol for visualizing motor neuron projection and axon arborization in transgenic Hb9::GFP mouse embryos. After immunostaining for motor neurons, we used light sheet fluorescence microscopy to image embryos for subsequent quantitative analysis. This protocol is applicable to other neuron navigation processes in the central nervous system.

Spinal motor neurons (MNs) extend their axons to communicate with their innervating targets, thereby controlling movement and complex tasks in ...

Serial Block-Face Scanning Electron Microscopy (SBEM) for the Study of Dendritic Spines

Three-dimensional electron microscopy (3D EM) gives a possibility to analyze morphological parameters of dendritic spines with nanoscale resolution. In addition, some features of dendritic spines, such as volume of the spine and post-synaptic density (PSD) (representing post-synaptic part of the synapse), presence of presynaptic terminal, and smooth endoplasmic reticulum or atypical form of PSD (e.g., multi-innervated spines), can be observed only with 3D EM. By employing serial block-face scanning electron microscopy (SBEM) it is possible to obtain 3D EM data easier and in a more reproducible manner than when performing traditional serial sectioning. Here we show how to prepare mouse hippocampal samples for SBEM analysis and how this protocol can be combined with immunofluorescence study of dendritic spines. Mild fixation perfusion allows us to perform immunofluorescence studies with light microscopy on one half of the brain, while the other half was prepared for SBEM. This approach reduces the number of animals to be used for the study.

Serial Block-Face Scanning Electron Microscopy SBEM for the Study of Dendritic Spines

Serial Block-Face Scanning Electron Microscopy (SBEM) is applied to image and analyze dendritic spines in the murine hippocampus.

Three-dimensional electron microscopy (3D EM) gives a possibility to analyze morphological parameters of dendritic spines with nanoscale resolution. In ...

Primary Cultures of Rat Astrocytes and Microglia and Their Use in the Study of Amyotrophic Lateral Sclerosis

This protocol demonstrates how to prepare primary cultures of glial cells, astrocytes, and microglia from the cortices of Sprague Dawley rats and how to use these cells for the purpose of studying the pathophysiology of amyotrophic lateral sclerosis (ALS) in the rat hSOD1G93A model. First, the protocol shows how to isolate and culture astrocytes and microglia from postnatal rat cortices, and then how to characterize and test these cultures for purity by immunocytochemistry using the glial fibrillary acidic protein (GFAP) marker of astrocytes and the ionized calcium-binding adaptor molecule 1 (Iba1) microglial marker. In the next stage, methods are described for dye-loading (calcium-sensitive Fluo 4-AM) of cultured cells and the recordings of Ca2+ changes in video imaging experiments on live cells.
The examples of video recordings consist of: (1) cases of Ca2+ imaging of cultured astrocytes acutely exposed to immunoglobulin G (IgG) isolated from ALS patients, showing a characteristic and specific response compared to the response to ATP as demonstrated in the same experiment. Examples also show a more pronounced transient rise in intracellular calcium concentration evoked by ALS IgG in hSOD1G93A astrocytes compared to non-transgenic controls; (2) Ca2+ imaging of cultured astrocytes during a depletion of calcium stores by thapsigargin (Thg), a non-competitive inhibitor of the endoplasmic reticulum Ca2+ ATPase, followed by store-operated calcium entry elicited by the addition of calcium in the recording solution, which demonstrates the difference between Ca2+ store operation in hSOD1G93A and in non-transgenic astrocytes; (3) Ca2+ imaging of the cultured microglia showing predominantly a lack of response to ALS IgG, whereas ATP application elicited a Ca2+ change. This paper also emphasizes possible caveats and cautions regarding critical cell density and purity of cultures, choosing the correct concentration of the Ca2+ dye and dye-loading techniques.

We present here a protocol on how to prepare primary cultures of glial cells, astrocytes, and microglia from rat cortices for time-lapse video imaging of intracellular Ca2+ for research on pathophysiology of amyotrophic lateral sclerosis in the hSOD1G93A rat model.

This protocol demonstrates how to prepare primary cultures of glial cells, astrocytes, and microglia from the cortices of Sprague Dawley rats and how to ...

Investigation of Spatial Interaction Between Astrocytes and Neurons in Cleared Brains

Combining viral vector transduction and tissue clearing using the CLARITY method makes it possible to simultaneously investigate several types of brain cells and their interactions. Viral vector transduction enables the marking of diverse cell types in different fluorescence colors within the same tissue. Cells can be identified genetically by activity or projection. Using a modified CLARITY protocol, the potential sample size of astrocytes and neurons has grown by 2-3 orders of magnitude. The use of CLARITY allows the imaging of complete astrocytes, which are too large to fit in their entirety in slices, and the examination of the somata with all their processes. In addition, it provides the opportunity to investigate the spatial interaction between astrocytes and different neuronal cell types, namely, the number of pyramidal neurons in each astrocytic domain or the proximity between astrocytes and specific inhibitory neuron populations. This paper describes, in detail, how these methods are to be applied.

Combining viral vector transduction and brain clearing using the CLARITY method allows the investigation of a large number of neurons and astrocytes simultaneously.

Combining viral vector transduction and tissue clearing using the CLARITY method makes it possible to simultaneously investigate several types of brain ...

Coherent Anti-Stokes Raman Spectroscopy (CARS) Application for Imaging Myelination in Brain Slices

Coherent anti-Stokes Raman spectroscopy (CARS) is a technique classically employed by chemists and physicists to produce a coherent signal of signature vibrations of molecules. However, these vibrational signatures are also characteristic of molecules within anatomical tissue such as the brain, making it increasingly useful and applicable for Neuroscience applications. For example, CARS can measure lipids by specifically exciting chemical bonds within these molecules, allowing for quantification of different aspects of tissue, such as myelin involved in neurotransmission. In addition, compared to other techniques typically used to quantify myelin, CARS can also be set up to be compatible with immunofluorescent techniques, allowing for co-labeling with other markers such as sodium channels or other components of synaptic transmission. Myelination changes are an inherently important mechanism in demyelinating diseases such as multiple sclerosis or other neurological conditions such as Fragile X Syndrome or autism spectrum disorders is an emerging area of research. In conclusion, CARS can be utilized in innovative ways to answer pressing questions in Neuroscience and provide evidence for underlying mechanisms related to many different neurological conditions.

Coherent Anti-Stokes Raman Spectroscopy CARS Application for Imaging Myelination in Brain Slices

Visualizing myelination is an important goal for many researchers studying the nervous system. CARS is a technique that is compatible with immunofluorescence that can natively image lipids within tissue such as the brain illuminating specialized structures such as myelin.

Coherent anti-Stokes Raman spectroscopy (CARS) is a technique classically employed by chemists and physicists to produce a coherent signal of signature ...