This work was supported in part by grants from the National Key Research and Development Program of China (2022YFA1105503), the State Key Laboratory of Neuroscience (SKLN-202103), and the Zhejiang Natural Science Foundation of China (Y21H120019), the Natural Science Foundation of China (82070981).

1. Obtaining ROs from iPSCs25
NOTE: ROs were derived from iPSCs by modifying the previously reported procedure.
<ol>
	<li>Dissociate iPSCs at ~90% confluence using a bacterial protease (see Table of Materials). Chop up the colonies into pieces and scrape them with a cell lifter.</li>
	<li>After collection, resuspend the clusters of cells in 0.25 mL of ice-cold Matrigel. Following 20 min of incubation at 37 &#176;C, scatter the solidified gel with the DMEM/F12 medium (0.5% N2 supplement, 1% B27 supplement, 0.1 mM &#946;-mercaptoethanol, 0.2 mM L-glutamine substitute, 100 U/mL penicillin and 100 mg/mL streptomycin). Define this time as day 0 of differentiation.</li>
	<li>After 5 days of floating culture, cells group to form cell cysts. Transfer these cell cysts into a 100 mm Petri dish where they will now adhere and grow.</li>
	<li>On day 15, use bacterial protease to dissociate the adhering cells and culture them in DMEM/F12 medium (2% B27 supplement, 0.1 mM non-essential amino acids).</li>
	<li>On day 21, change the medium with 20 mL of serum-containing medium (DMEM/F12 medium, 8% FBS, 2% B27 supplement, 100 mM Taurine, and 2 mM L-glutamine substitute), and replace the medium every week.</li>
</ol>
2. Anterior fixation of ROs
<ol>
	<li>Gently move one of the cultured ROs (~D180, a size of about 1~2 mm in diameter is optimal) from the Petri dish to 1.5 mL microcentrifuge tubes with a pipette and mark the relevant information about the ROs. 
	NOTE: Cut ROs in half or cut off the poorly structured tissue with a razor under the microscope in the dish containing fixative if the ROs are too large.</li>
	<li>Remove the culture medium from 1.5 mL microcentrifuge tubes with a pipette. Add 1.5 mL of 2% paraformaldehyde-2% glutaraldehyde fixative diluted in 0.1 M phosphate buffer (PB), pH 7.4, and fix at room temperature (RT) for 4 h. Store the microcentrifuge tubes at 4 &#176;C overnight. 
	NOTE: This step can be paused for 1 week but restarting as early as possible is recommended. If ROs are left for longer, it may cause autolysis of cells leading to destruction of the ultrastructure of ROs. Due to the fragility of the ROs, avoid rotating, oscillating, flipping, or harsh operations.</li>
</ol>
3. Post fixation of ROs
<ol>
	<li>Remove the 2% paraformaldehyde-2% glutaraldehyde fixative and wash with 1 mL of 0.01 M phosphate buffered saline (PBS) at pH 7.4 for 3 x 10 min. Gently suspend the ROs in the 1.5 mL microcentrifuge tubes during the washing process for fully rinsing. For every additional day of fixation, increase the number of washes by two to remove&#160;the residual fixative as much as possible.</li>
	<li>Remove any remaining PBS using a pipette and replace it with ~150 &#181;L of 1% osmic acid (OsO4) until the tissue blocks are submerged. Place the microcentrifuge tubes in a dark box to protect them from light and infiltrate at RT for 1 h. 
	NOTE: OsO4 is a strong oxidant and must be handled in a fume hood.</li>
</ol>
4. Staining and dehydration
NOTE: The dehumidifier must be turned on to dry the environment from this step.
<ol>
	<li>Remove OsO4 using a pipette, wash with 0.01 M PBS (pH 7.4) 3 x 10 min, and then wash 3 x 10 min with deionized water. 
	NOTE: To ensure complete rinsing of ROs, we recommend suspending the ROs. Aspirate OsO4 into a dedicated disposal bottle.</li>
	<li>Remove the last deionized water wash and replace with ~150 &#181;L per tube (ensure full infiltration of tissue blocks) of uranium acetate and stain for 1-2 h at RT. Protect the microcentrifuge tubes from light.</li>
	<li>Remove the uranium acetate and replace it with 1 mL of 50% acetone in the fume hood, dehydrating for 10 min. Then, use 70%, 80%, 90%, 100%, and 100% acetone for gradient dehydration of the ROs successively. Each dehydration lasts 10 min. 
	NOTE: Leave some liquid to soak the ROs for each exchange to prevent exposure to air and moisture. Additionally, operate quickly as soon as possible because acetone is volatile. Aspirate uranium acetate into a dedicated disposal bottle.</li>
</ol>
5. Infiltration
<ol>
	<li>Discard the acetone and add ~150 &#181;L of the mixture consisting of acetone and Epon-812 resin at a ratio of 1:1. Place it in a 37 &#176;C oven for 1 h.</li>
	<li>Remove the above mixture and replace it with a mixture consisting of acetone and Epon-812 resin at a ratio of 1:4. Place it in a 37 &#176;C oven overnight.</li>
	<li>Transfer the ROs into the new tubes containing ~500 &#181;L of pure epoxy resin carefully and place them in a 45 &#176;C oven for 1 h.</li>
</ol>
6. Embedding
<ol>
	<li>Prepare the 30-50 mL of pure epoxy resin in a 50 mL centrifuge tube by mixing the reagent kit according to the manufacturer&#39;s instructions in advance. Rotate the pure epoxy resin upside down slowly for 30 min to reduce air bubbles introduced by the mixer. Then, leave it in a 37 &#176;C oven for 20 min. Add 2/3 volume of pure epoxy resin to the groove of the embedding mold.</li>
	<li>Use a toothpick to pick ROs into the embedding mold equipped with pure epoxy resin.</li>
	<li>Fill the groove with pure epoxy resin until it slightly protrudes or bulges.</li>
	<li>Adjust the position of ROs at both ends of the embedding mold for directional embedding. Place the embedding mold in a 45 &#176;C oven for 1-2 h and then put it in a 65 &#176;C oven for 48-72 h.</li>
</ol>
7. Semi-thin positioning
<ol>
	<li>Sand the excess epoxy resin around the tissue from the ends of the longer four sides of the embedding block to form a trapezoidal face. Install the embedding block into the sample mounting stand and screw down the bolt with an L screwdriver. Then, install glass knives on the knife stand and tighten the nut manually.</li>
	<li>Trim away the excess resin until the desired area is exposed.</li>
	<li>Fill the trough of the glass knife with water. Cut semi-thin slices to a thickness of 1 &#181;m using a semi-thin microtome and lay them on a microscope slide with a needle.</li>
	<li>Add 50 &#181;L of 1% toluidine blue to dye semi-thin slices and incubate for 1 min at 95 &#176;C on the heating plate. Rinse with distilled water for 2 min. Use a 20x ordinary optical microscope to observe the structure of the sections.</li>
</ol>
8.&#160;Ultra-thin sectioning
<ol>
	<li>Install the embedding blocks and glass knives as described in step 7.1. Adjust the distance between the knife and the tissue block, the liquid level of the sink, and the slicing speed. Cut the ultrathin sections with a thickness of ~70-100 nm with an ultrathin microtome and scoop the slices onto the copper mesh (200 mesh) with the supporting film.</li>
	<li>Stain ultrathin sections with 3% lead citrate for 15 min to enhance contrast in a dual staining procedure.</li>
</ol>
9. Imaging ROs by TEM
<ol>
	<li>Launch the software (see the Table of Materials).</li>
	<li>Click on the Memory mode button and mark each ultrathin section roughly.</li>
	<li>Search for synaptic structures.
	<ol>
		<li>Distinguish the laminated structure of ROs at low magnification.</li>
		<li>Locate the outer nuclear layer (ONL), which shows a nuclear structure with a deeper electron density. Subsequently, navigate to the terminal of the photoreceptor where numerous vesicles accumulate. Search for the rod terminals with a single synaptic ribbon and the cone terminals with multiple synaptic ribbons.</li>
		<li>Move to the inner plexiform layer (IPL) and seek out a similar structure with a small ribbon.</li>
	</ol>
	</li>
	<li>Capture the microscopic image.
	<ol>
		<li>Click Start and input the corresponding magnifying power.</li>
		<li>Save the images.</li>
	</ol>
	</li>
</ol>

Optimized Protocol for Retinal Organoid Sample Preparation for TEM

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H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ribbon+synapses+of+the+retina.">Ribbon synapses of the retina.</a> Cell Tissue Res. 326 (2), 339-346 (2006).</li><li>Kolb, H., Fernandez, E., Nelson, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Webvision: The organization of the retina and visual. , (1995).</li><li>Soto, F., Zhao, L., Kerschensteiner, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Synapse+maintenance+and+restoration+in+the+retina+by+ngl2.">Synapse maintenance and restoration in the retina by ngl2.</a> Elife. 7, e30388 (2018).</li><li>Kruczek, K., Swaroop, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pluripotent+stem+cell-derived+retinal+organoids+for+disease+modeling+and+development+of+therapies.">Pluripotent stem cell-derived retinal organoids for disease modeling and development of therapies.</a> Stem Cells. 38 (10), 1206-1215 (2020).</li><li>O'Hara-Wright, M., Gonzalez-Cordero, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Retinal+organoids:+A+window+into+human+retinal+development.">Retinal organoids: A window into human retinal development.</a> Development. 147 (24), (2020).</li><li>Norrie, J. 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The authors have no conflicts of interest to disclose.

In this article, we presented a detailed protocol for observing conventional and ribbon synaptic ultrastructure in ROs by TEM. This protocol is based on the previously described retinal preparation methods with some modifications20. To improve the success rate of sample treatment and the quality of TEM micrographs, consider the following key points. First, it is important to acknowledge that ROs develop from iPSCs, forming a cell mass lacking vasculature6,<sup cl...

The retina, a vital visual sensory organ in humans and mammals, exhibits a distinct laminated structure characterized by three nuclear layers housing neuron somas and two plexiform layers formed by synaptic connections1, including conventional synapses and the specialized ribbon synapse2,3. The ribbon synapse plays a crucial role in transmitting vesicle impulses in a graded manner2,3. The vision process involves electro-optical signal transmission across various levels of neurons and synapses, ultimately reaching the visual cortex4,5.
Retinal organoids (ROs) represent a three-dimensional (3D) culture system derived from induced pluripotent stem cells (iPSCs), mimicking the physiological states of retinal tissue in vitro1,6,7. This approach holds promise for studying retinal diseases8, drug screening9, and serving as a potential therapy for irreversible retinal degenerative conditions such as retinitis pigmentosa10&#160;and glaucoma11. As a powerful in vitro optical conduction system, the synapse within ROs is a crucial structure facilitating effective signal transformation and transfer5.
RO development can be roughly divided into three stages according to their morphological traits and molecular expression profiles6,12. ROs in stage 1 (around D21-D60) comprise neural progenitor cells of the retina, many retinal ganglion cells (RGCs), and a few starburst amacrine cells (SACs), corresponding to the first epoch of human fetal development. In stage 2 (around D50-D150), ROs express some photoreceptor precursors, interneurons, and synaptogenesis-related genes, which represents a phase of transition. Photoreceptors develop maturity in stage 3 ROs (around after D100-D150), corresponding&#160;to the third stage of human fetal development6,12,13. Notably, compared with ROs in stage 1 and stage 2, ROs in stage 3 have a distinct lamellar structure whose synapses have matured12, including the presence of ribbon synapses14. Moreover, a recent study has confirmed that the mature synapses exist the transmission of light signals, indicating they are functional13. Thus, ROs in stage 3 are often selected to investigate synaptic structure.
Immunohistochemistry is widely applied to the study of the expression of various molecular proteins. However, the limitation of optical microscopy lies in its ability to observe only a restricted number of specific cells and molecules at a time, resulting in a lack of comprehensive analysis of the relationships between cells and their surrounding environment. Transmission Electron Microscopy (TEM) is characterized by high resolution, with a limited resolution of 0.1-0.2 nm, surpassing the light microscope by ~10-20 times15. It makes up for the defects of optical microscopy and is used to elucidate retinal development and synapse maturation in humans16,17 and various species18,19,20,21. TEM enables the direct distinction of presynaptic and postsynaptic components18,20, and even allows for comprehensive observation of subcellular structures such as ribbons2,3, vesicles22, and mitochondria23. Therefore, TEM is an essential tool for identifying types of synapses and exploring the ultrastructure of synaptic contacts in ROs at the nanoscale.
It is crucial to note that sample preparation is of great importance for acquiring high-quality electron micrographs. Although some studies have performed EM on ROs12,13,24, the detailed procedures are unclear. Since the quality of the electron microscopy image depends on the effect of RO fixation and reagent permeation to a large extent, various important factors need to be considered during preparation. Consequently, to better investigate synaptic contacts in ROs, we present a method with good reproducibility that shows the operation points of RO fixing, embedding, and the identification of observation sites.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>100 mm Petri dish</td>
			<td>Corning</td>
			<td>430167</td>
			<td></td>
		</tr>
		<tr>
			<td>Acetone</td>
			<td>Electron Microscopy Science</td>
			<td>10000</td>
			<td></td>
		</tr>
		<tr>
			<td>B27 supplement</td>
			<td>Gibco</td>
			<td>A3582801</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell lifter</td>
			<td>Santa Cruz</td>
			<td>sc-395251</td>
			<td></td>
		</tr>
		<tr>
			<td>Copper grids</td>
			<td>Beijing Zhongjingkeyi Technology Co., Ltd.</td>
			<td>AZH400HH</td>
			<td></td>
		</tr>
		<tr>
			<td>DigitalMicrograph Software</td>
			<td>Gatan, Inc.</td>
			<td></td>
			<td>Software</td>
		</tr>
		<tr>
			<td>Dispase</td>
			<td>StemCell Technologies</td>
			<td>#07913</td>
			<td>Bacterial protease</td>
		</tr>
		<tr>
			<td>DMEM/F12 medium</td>
			<td>Gibco</td>
			<td>#11320033</td>
			<td></td>
		</tr>
		<tr>
			<td>Embedding mold</td>
			<td>Beijing Zhongjingkeyi Technology Co., Ltd.</td>
			<td>GZ10592</td>
			<td></td>
		</tr>
		<tr>
			<td>Epon-812 resin</td>
			<td>Electron Microscopy Science</td>
			<td>#14900</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum (FBS)</td>
			<td>Biological Industries</td>
			<td>#04-0021A</td>
			<td></td>
		</tr>
		<tr>
			<td>Glutaraldehyde</td>
			<td>Electron Microscopy Science</td>
			<td>16020</td>
			<td></td>
		</tr>
		<tr>
			<td>hiPSC</td>
			<td>Shownin Biotechnology Co. Ltd.</td>
			<td>RC01001-A</td>
			<td></td>
		</tr>
		<tr>
			<td>Lead citrate</td>
			<td>Beijing Zhongjingkeyi Technology Co., Ltd.</td>
			<td>GZ02618</td>
			<td></td>
		</tr>
		<tr>
			<td>L-GlutaMax</td>
			<td>Life Technologies</td>
			<td>#35050061</td>
			<td>L-glutamine substitute</td>
		</tr>
		<tr>
			<td>Matrigel</td>
			<td>Corning</td>
			<td>356234</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope slide</td>
			<td>CITOTEST</td>
			<td>80312-3161</td>
			<td></td>
		</tr>
		<tr>
			<td>N2 supplement</td>
			<td>Gibco</td>
			<td>17502048</td>
			<td></td>
		</tr>
		<tr>
			<td>Na2HPO4&#183; 12H2O</td>
			<td>Sigma</td>
			<td>71650</td>
			<td>A component of PB/PBS</td>
		</tr>
		<tr>
			<td>NaH2PO4&#183; H2O</td>
			<td>Sigma</td>
			<td>71507</td>
			<td>A component of PB/PBS</td>
		</tr>
		<tr>
			<td>Non-essential amino acids</td>
			<td>Sigma</td>
			<td>#M7145</td>
			<td></td>
		</tr>
		<tr>
			<td>Optical microscope</td>
			<td>Lab Binocular Biological Microscope</td>
			<td>Xsz-107bnii</td>
			<td></td>
		</tr>
		<tr>
			<td>OsO4</td>
			<td>TED PELLA</td>
			<td>4008-160501</td>
			<td></td>
		</tr>
		<tr>
			<td>Oven</td>
			<td>Bluepard</td>
			<td>BPG9040A</td>
			<td></td>
		</tr>
		<tr>
			<td>Paraformaldehyde</td>
			<td>Electron Microscopy Science</td>
			<td>157-8</td>
			<td></td>
		</tr>
		<tr>
			<td>Penicillin-Streptomycin</td>
			<td>Gibco</td>
			<td>#15140-122</td>
			<td></td>
		</tr>
		<tr>
			<td>Semi/ultrathin microtome</td>
			<td>Reichert-Jung</td>
			<td>396649</td>
			<td></td>
		</tr>
		<tr>
			<td>Taurine</td>
			<td>Sigma</td>
			<td>#T0625</td>
			<td></td>
		</tr>
		<tr>
			<td>Toluidine blue</td>
			<td>Sangon Biotech</td>
			<td>E670105-0100</td>
			<td></td>
		</tr>
		<tr>
			<td>Transmission Electron Microscopes</td>
			<td>HITACHI</td>
			<td>H-7500</td>
			<td></td>
		</tr>
		<tr>
			<td>Uranyl acetate</td>
			<td>TED PELLA</td>
			<td>CA96049</td>
			<td></td>
		</tr>
		<tr>
			<td>&#946;-mercaptoethanol</td>
			<td>Sigma</td>
			<td>444203</td>
			<td></td>
		</tr>
	</tbody>
</table>

preparing retinal organoid samples for transmission electron microscopy

Retinal organoids (ROs) are a three-dimensional culture system mimicking human retinal features that have differentiated from induced pluripotent stem cells (iPSCs) under specific conditions. Synapse development and maturation in ROs have been studied immunocytochemically and functionally. However, the direct evidence of the synaptic contact ultrastructure is limited, containing both special ribbon synapses and conventional chemical synapses. Transmission electron microscopy (TEM) is characterized by high resolution and a respectable history elucidating retinal development and synapse maturation in humans and various species. It is a powerful tool to explore synaptic structure in ROs and is widely used in the research field of ROs. Therefore, to better explore the structure of RO synaptic contacts at the nanoscale and obtain high-quality microscopic evidence, we developed a simple and repeatable method of RO TEM sample preparation. This paper describes the protocol, reagents used, and detailed steps, including RO fixation preparation, post fixation, embedding, and visualization.

The establishment of 3D ROs through iPSC differentiation provides a powerful tool for studying retinal disease mechanisms and stem cell replacement therapy. Although others have demonstrated the synaptic connections in ROs functionally and immunocytochemically, direct evidence of conventional and ribbon synapses is very limited. Here we present a method for investigating the ultrastructure of two types of synapses in ROs by TEM. After 180 days of culture, ROs were fixed, stained, embedded, and ultrathin sliced. TEM obser...

Author Spotlight: Analyzing the Synaptic Ultrastructure in Mature Retinal Organoids Using TEM

This protocol provides an optimized and elaborate preparation procedure for retinal organoid samples for transmission electron microscopy. It is suitable for applications that involve the analysis of synapses in mature retinal organoids.

Preparing Retinal Organoid Samples for Transmission Electron Microscopy

Retinal organoids (ROs) are a three-dimensional culture system mimicking human retinal features that have differentiated from induced pluripotent stem ...

preparing-retinal-organoid-samples-for-transmission-electron

Research

JoVE Journal

Neuroscience

766 Views.  Wenzhou Medical University. This protocol provides an optimized and elaborate preparation procedure for retinal organoid samples for transmission electron microscopy. It is suitable for applications that involve the analysis of synapses in mature retinal organoids.

Video: Author Spotlight: Analyzing the Synaptic Ultrastructure in Mature Retinal Organoids Using TEM

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 ...

The Olfactory System as a Model to Study Axonal Growth Patterns and Morphology In Vivo

The olfactory system has the unusual capacity to generate new neurons throughout the lifetime of an organism. Olfactory stem cells in the basal portion of the olfactory epithelium continuously give rise to new sensory neurons that extend their axons into the olfactory bulb, where they face the challenge to integrate into existing circuitry. Because of this particular feature, the olfactory system represents a unique opportunity to monitor axonal wiring and guidance, and to investigate synapse formation. Here we describe a procedure for in vivo labeling of sensory neurons and subsequent visualization of axons in the olfactory system of larvae of the amphibian Xenopus laevis. To stain sensory neurons in the olfactory organ we adopt the electroporation technique. In vivo electroporation is an established technique for delivering fluorophore-coupled dextrans or other macromolecules into living cells. Stained sensory neurons and their axonal processes can then be monitored in the living animal either using confocal laser-scanning or multiphoton microscopy. By reducing the number of labeled cells to few or single cells per animal, single axons can be tracked into the olfactory bulb and their morphological changes can be monitored over weeks by conducting series of in vivo time lapse imaging experiments. While the described protocol exemplifies the labeling and monitoring of olfactory sensory neurons, it can also be adopted to other cell types within the olfactory and other systems.

The Olfactory System as a Model to Study Axonal Growth Patterns and Morphology In Vivo

We describe a protocol for in vivo labeling of olfactory sensory neurons by electroporation and subsequent confocal laser-scanning or multiphoton microscopy to visualize neuronal morphology and its development over time.

The olfactory system has the unusual capacity to generate new neurons throughout the lifetime of an organism. Olfactory stem cells in the basal portion of ...

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, ...

Preparation of Non-human Primate Brain Tissue for Pre-embedding Immunohistochemistry and Electron Microscopy

Despite all the technological advances at the light microscopy&#160;level, electron microscopy remains the only tool in neuroscience to examine and characterize ultrastructural and morphological details of neurons, such as synaptic contacts. Good preservation of brain tissue for electron microscopy can be obtained by rigorous cryo-fixation methods, but these techniques are rather costly and limit the use of immunolabeling, which is crucial to understand the connectivity of identified neuronal systems. Freeze-substitution methods have been developed to allow the combination of cryo-fixation with immunolabeling. However, the reproducibility of these methodological approaches usually relies on costly freezing devices. Moreover, achieving reliable results with this technique is very time-consuming and skill-challenging. Hence, the traditional chemically fixed brain, particularly with acrolein fixative, remains a time-efficient and low-cost method to combine electron microscopy with immunohistochemistry. Here, we provide a reliable experimental protocol using chemical acrolein fixation that leads to the preservation of primate brain tissue and is compatible with pre-embedding immunohistochemistry and transmission electron microscopic examination.

Here, we provide an easy, low-cost, and time-efficient protocol to chemically fix primate brain tissue with acrolein fixative, allowing for long-term preservation that is compatible with pre-embedding immunohistochemistry for transmission electron microscopy.

Despite all the technological advances at the light microscopy&#160;level, electron microscopy remains the only tool in neuroscience to examine and ...

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

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

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

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

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

Preparation of Mouse Retinal Cryo-sections for Immunohistochemistry

Preparation of high-quality mouse eye sections for immunohistochemistry (IHC) is critical for assessing the retinal structure and function and for determining the mechanisms underlying retinal diseases. Maintaining structural integrity throughout the tissue preparation is vital for obtaining reproducible retinal IHC data but can be challenging due to the fragility and complexity of retinal cytoarchitecture. Strong fixatives like 10% formalin or Bouin's solution optimally preserve the retinal structure, they often impede IHC analysis by enhancing the background fluorescence and/or diminishing antibody-epitope interactions, a process known as epitope masking. Milder fixatives, on the other hand, like 4% paraformaldehyde, reduces background fluorescence and epitope-masking, meticulous dissection techniques must be utilized to preserve the retinal structure. In this article, we present a comprehensive method to prepare mouse ocular posterior cups for IHC that is sufficient to preserve most antibody-epitope interactions without loss of retinal structural integrity. We include representative IHC with antibodies to various retinal cell type markers to illustrate tissue preservation and orientation under optimal and sub-optimal conditions. Our goal is to optimize IHC studies of the retina by providing a complete protocol from ocular posterior cup dissection to IHC.

This report describes comprehensive methods for preparing frozen mouse retina sections for immunohistochemistry (IHC). Methods described include dissection of the ocular posterior cup, paraformaldehyde fixation, embedding in Optimal Cutting Temperature (OCT) media and tissue orientation, sectioning and immunostaining.

Preparation of high-quality mouse eye sections for immunohistochemistry (IHC) is critical for assessing the retinal structure and function and for ...

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 ...

Determination of Mitochondrial Respiration and Glycolysis in Ex Vivo Retinal Tissue Samples

Mitochondrial respiration is a critical energy-generating pathway in all cells, especially retinal photoreceptors that possess a highly active metabolism. In addition, photoreceptors also exhibit high aerobic glycolysis like cancer cells. Precise measurements of these metabolic activities can provide valuable insights into cellular homeostasis under physiological conditions and in disease states. High throughput microplate-based assays have been developed to measure mitochondrial respiration and various metabolic activities in live cells. However, a vast majority of these are developed for cultured cells and have not been optimized for intact tissue samples and for application ex vivo. Described here is a detailed step-by-step protocol, using microplate-based fluorescence technology, to directly measure oxygen consumption rate (OCR) as an indicator of mitochondrial respiration, as well as extracellular acidification rate (ECAR) as an indicator of glycolysis, in intact ex vivo retinal tissue. This method has been used to successfully assess metabolic activities in adult mouse retina and demonstrate its application in investigating cellular mechanisms of aging and disease.

Determination of Mitochondrial Respiration and Glycolysis in Ex Vivo Retinal Tissue Samples

Described here is a detailed protocol for performing mitochondrial stress assay and glycolytic rate assay in ex vivo retinal tissue samples using a commercial bioanalyzer.

Mitochondrial respiration is a critical energy-generating pathway in all cells, especially retinal photoreceptors that possess a highly active metabolism. ...

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 ...