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

Animal care and use protocols were approved by the Ethics Committee of Wenzhou Medical University and followed the guidelines established by the Association for Research in Vision and Ophthalmology (ARVO). All mice were maintained in a 12-h light and 12-h dark cycle and supplied with a standard chow diet.
1. Retina dissection and fixation
<ol>
	<li>Anesthetize the mice (C57BL/6J, Male, 8-week-old, 20-25 g) by injecting 2,2,2-tribromoethanol (0.25 mg/g of body weight) intraperitoneally. Confirm the depth of anesthetization by no retraction of the back paw after pinching the toe or no blinking reflex. Then, dislocate the cervical vertebra while still under anesthesia to euthanize the animal. 
	NOTE: Tribromoethanol was chosen for its well-documented efficacy in inducing rapid and reliable anesthesia in small animal models, particularly for short-term procedures. While pharmaceutical-grade anesthetics are preferred, tribromoethanol offers specific advantages in certain protocols, making it a suitable alternative when properly prepared and administered. All precautions were taken to ensure its safe and effective use. This study adheres to ethical considerations by ensuring proper preparation, administration, and monitoring to mitigate potential risks associated with its use.</li>
	<li>Remove the eyes with curved scissors and immerse the eyes in a mixed solution containing 2% glutaraldehyde and 2% paraformaldehyde in phosphate buffer (PB, 0.1 M, pH 7.4).</li>
	<li>Remove the anterior segment and vitreous from the eyes with forceps under the dissecting microscope.</li>
	<li>Peel the sclera with the two forceps until the retina is completely isolated from the eyecup.</li>
	<li>Cut the retina rapidly into 100 - 200 &#181;m-thick strips with a razor and handle the retina strips with a Pasteur pipette into a new microcentrifuge (EP) tube containing 2% glutaraldehyde and 2% paraformaldehyde in phosphate buffer (PB, 0.1 M, pH 7.4) for 2 h at room temperature (RT) on the rotator. After completion, place the tubes to fix at 4 &#176;C for 24 h.</li>
</ol>
2. Sample embedding process
<ol>
	<li>After washing in 0.1 M sodium cacodylate buffer (pH 7.4) 5 times&#160;for 15 min each, incubate the retina strips with a solution containing 1.5% potassium ferrocyanide and 1% OsO4(in PB, pH7.4) in 0.1 M sodium cacodylate with 4 mM CaCl2 for 1.5 h in the dark on ice. 
	NOTE: Dark incubation and lack of phosphate can prevent precipitation and artifacts in the sample.</li>
	<li>Wash the retina strips 3 times&#160;in ddH2O for 10 min each, and then treat with 1% thiocarbohydrazide in ddH2O for 30 min in the dark at RT.</li>
	<li>After rinsing in ddH2O 3 times&#160;for 15 min each, incubate the retina strips in 2% OsO4 (in PB, PH 7.4) for 45 min in the dark at RT.</li>
	<li>After washing in ddH2O 3 times&#160;for 15 min each, immerse the retina strips in 1% aqueous uranyl acetate in the dark at 4 &#176;C&#160;overnight.</li>
	<li>Next day, after washing in ddH2O 3 times&#160;for 15 min each, treat the retina strips with 0.66% lead nitrate in L-aspartic acid (0.03 M, pH 5.5) for 40 min at 65 &#176;C.</li>
	<li>After washing in ddH2O 3 times&#160;for 15 min each, dehydrate the retina strips with ascending ethanol concentration of 30%, 50%, 70%, 90%, and 100% for 20 min in each solution.</li>
	<li>Treat retina strips with a mixed solution containing equal ethanol and acetone for 20 min, followed by an acetone wash 2 times&#160;for 20 min each.</li>
	<li>Infiltrate the retina strips in acetone/resin mixed in a ratio of 7:3 in the dark at RT&#160;overnight and then acetone/resin at 3:7 in the dark for 24 h at RT. 
	NOTE: The precise composition of the resin is as follows: Epon812 10.06 mL, DDSA 4 mL, MNA 5.94 mL, and DMP-30 0.2 mL.</li>
	<li>Change the tubes the next day and infiltrate the retina strips with pure resin for 24 h at 45 &#176;C. Change the resin again the next day, and embed the retina strips for 48 h at 60 &#176;C. 
	NOTE: The purpose of increasing the temperature is to improve penetration.</li>
</ol>
3. Selection of the area of interest
<ol>
	<li>Cut the resin blocks into 1 &#181;m thick sections, stain sections with 1% toluidine-blue, and observe the general structure of the retina under light microscopy to select the rough regions of interest.</li>
	<li>Coat the resin blocks with platinum using a sputter coater, followed by milling with 70 nm thick section using a focused-ion-beam scanning electron microscopy under a current of 0.23 nA and acceleration voltage of 30 kV.</li>
	<li>Pre-screen the section using FIB-SEM with 2000x magnification to identify the precise regions of interest.</li>
	<li>Collect the data using FIB-SEM with a beam current of 0.69 nA and acceleration voltage of 2 kV. The other parameters are as follows: dwell time = 2 &#181;s, voxel size = 1.8 nm x 1.8 nm x 15 nm.</li>
</ol>

Visualizing Retinal Synaptic Structures in 3D with Volume Electron Microscopy

<ol>
	<li>Masland, R. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+fundamental+plan+of+the+retina.">The fundamental plan of the retina.</a> Nat Neurosci. 4 (9), 877-886 (2001).</li><li>Harris, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transmission+electron+microscopy+in+molecular+structural+biology:+A+historical+survey.">Transmission electron microscopy in molecular structural biology: A historical survey.</a> Arch Biochem Biophys. 581, 3-18 (2015).</li><li>Varsano, N., Wolf, S. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electron+microscopy+of+cellular+ultrastructure+in+three+dimensions.">Electron microscopy of cellular ultrastructure in three dimensions.</a> Curr Opin Struct Biol. 76, 102444 (2022).</li><li>Briggman, K. L., Bock, D. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Volume+electron+microscopy+for+neuronal+circuit+reconstruction.">Volume electron microscopy for neuronal circuit reconstruction.</a> Curr Opin Neurobiol. 22 (1), 154-161 (2012).</li><li>Titze, B., Genoud, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Volume+scanning+electron+microscopy+for+imaging+biological+ultrastructure.">Volume scanning electron microscopy for imaging biological ultrastructure.</a> Biol Cell. 108 (11), 307-323 (2016).</li><li>Sterling, P., Matthews, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Structure+and+function+of+ribbon+synapses.">Structure and function of ribbon synapses.</a> Trends Neurosci. 28 (1), 20-29 (2005).</li><li>Heidelberger, R., Thoreson, W. B., Witkovsky, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Synaptic+transmission+at+retinal+ribbon+synapses.">Synaptic transmission at retinal ribbon synapses.</a> Prog Retin Eye Res. 24 (6), 682-720 (2005).</li><li>Thoreson, W. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transmission+at+rod+and+cone+ribbon+synapses+in+the+retina.">Transmission at rod and cone ribbon synapses in the retina.</a> Pflugers Arch. 473 (9), 1469-1491 (2021).</li><li>Moser, T., Grabner, C. P., Schmitz, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sensory+processing+at+ribbon+synapses+in+the+retina+and+the+cochlea.">Sensory processing at ribbon synapses in the retina and the cochlea.</a> Physiol Rev. 100 (1), 103-144 (2020).</li><li>Schmitz, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+making+of+synaptic+ribbons:+how+they+are+built+and+what+they+do.">The making of synaptic ribbons: how they are built and what they do.</a> Neuroscientist. 15 (6), 611-624 (2009).</li><li>Kantardzhieva, A., Peppi, M., Lane, W. S., Sewell, W. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Protein+composition+of+immunoprecipitated+synaptic+ribbons.">Protein composition of immunoprecipitated synaptic ribbons.</a> J Prot Res. 11 (2), 1163-1174 (2011).</li><li>Zampighi, G. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Conical+tomography+of+a+ribbon+synapse:+structural+evidence+for+vesicle+fusion.">Conical tomography of a ribbon synapse: structural evidence for vesicle fusion.</a> PLoS One. 6 (3), e16944 (2011).</li><li>Barnes, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Morphological+diversity+of+the+rod+spherule:+A+study+of+serially+reconstructed+electron+micrographs.">Morphological diversity of the rod spherule: A study of serially reconstructed electron micrographs.</a> Plos One. 11 (3), e0150024 (2016).</li><li>Xu, C. S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Enhanced+FIB-SEM+systems+for+large-volume+3D+imaging.">Enhanced FIB-SEM systems for large-volume 3D imaging.</a> Elife. 6, e25916 (2017).</li><li>Crewe, A. V., Lin, P. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+use+of+backscattered+electrons+for+imaging+purposes+in+a+scanning+electron+microscope.">The use of backscattered electrons for imaging purposes in a scanning electron microscope.</a> Ultramicroscopy. 1 (3), 231-238 (1976).</li><li>Richards, R. G., Gwynn, I. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Backscattered+electron+imaging+of+the+undersurface+of+resin-embedded+cells+by+field-emission+scanning+electron+microscopy.">Backscattered electron imaging of the undersurface of resin-embedded cells by field-emission scanning electron microscopy.</a> J Microsc. 177, 43-52 (1995).</li><li>Wergin, W. P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+thin+and+thick+sections+of+biological+tissue+with+the+secondary+electron+detector+in+a+field-emission+scanning+electron+microscope.">Imaging thin and thick sections of biological tissue with the secondary electron detector in a field-emission scanning electron microscope.</a> Scanning. 19 (6), 386-395 (1997).</li><li>Willingham, M. C., Rutherford, A. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+use+of+osmium-thiocarbohydrazide-osmium+(OTO)+and+ferrocyanide-reduced+osmium+methods+to+enhance+membrane+contrast+and+preservation+in+cultured+cells.">The use of osmium-thiocarbohydrazide-osmium (OTO) and ferrocyanide-reduced osmium methods to enhance membrane contrast and preservation in cultured cells.</a> J Histochem Cytochem. 32 (4), 455-460 (1984).</li><li>Buravkov, S. V., Chernikov, V. P., Buravkova, L. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Simple+method+of+specimen+preparation+for+scanning+electron+microscopy.">Simple method of specimen preparation for scanning electron microscopy.</a> Bull Exp Biol Med. 151 (3), 378-382 (2011).</li><li>Masland, R. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neuronal+diversity+in+the+retina.">Neuronal diversity in the retina.</a> Curr Opin Neurobiol. 11 (4), 431-436 (2001).</li><li>Svara, F. N., Kornfeld, J., Denk, W., Bollmann, J. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Volume+EM+reconstruction+of+spinal+cord+reveals+wiring+specificity+in+speed-related+motor+circuits.">Volume EM reconstruction of spinal cord reveals wiring specificity in speed-related motor circuits.</a> Cell Rep. 23 (10), 2942-2954 (2018).</li><li>Kubota, Y., Sohn, J., Kawaguchi, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Large+volume+electron+microscopy+and+neural+microcircuit+analysis.">Large volume electron microscopy and neural microcircuit analysis.</a> Front Neural Circuits. 12, 98 (2018).</li><li>Guo, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+optimized+approach+using+cryofixation+for+high-resolution+3D+analysis+by+FIB-SEM.">An optimized approach using cryofixation for high-resolution 3D analysis by FIB-SEM.</a> J Struct Biol. 212 (1), 107600 (2020).</li><li>Seligman, A. 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The authors have no disclosures.

We implemented the OTO&#39;s Volume EM sample preparation protocol to analyze the photoreceptors&#39; terminal structure in retinal tissue. The focus was on detailing the entire procedure, starting from the detachment and fixation of the retina to showcasing the results of 3D reconstruction of photoreceptor axon terminals.
The distinctive feature of retinal tissue, unlike brain tissue, lies in its lack of regional differences. Comprising three layers of neuronal cell bodies and two layers of synaptic connections, the retina exhibits a regular and hierarchical structure despite the complexity of its neuronal and synaptic components20. In dissecting the retina, it becomes imperative to ensure that the thin strips are minimized to guarantee uniform contrast in subsequent sample preparation stages. The photoreceptor axon terminals, critical structures of interest, are situated in the outer plexiform layer (OPL) of the retina. The hierarchical and organized nature of the retina allows for efficient localization of the target structures, facilitating precise sampling during EM and ensuring the accuracy of subsequent 3D reconstruction results.
Volume EM facilitates three-dimensional visualization of local neural circuits and synaptic connections within the retina at the nanoscale, utilizing serial sections21,22. This technique enables the automatic generation of high-resolution 3D images, and FIB-SEM holds the potential to unveil intricate details of both physiological and pathological activities. Therefore, maintaining stable image contrast throughout the entire block is crucial during the sample preparation process. To achieve uniform conductivity, we employ the pre-embedding OTO method, a classical approach widely acknowledged for its effectiveness in enhancing contrast in scanning electron microscopy (SEM)18,23,24. This method involves a double osmium coating combined with thiocarbohydrazide, ensuring optimal bulk conductivity for large samples. Additionally, the double osmium treatment contributes to the superior preservation of lipid membrane structures, adding a layer of precision to our investigations. The retinal samples are hierarchical, the tissue blocks are small and easy to penetrate, and the OTO method can maintain better cell membrane contour and uniform image contrast.
Several critical factors contribute to the success of this protocol. Firstly, swift retinal harvesting and precise fixation times are paramount for maintaining optimal subcellular structure. Secondly, ensuring appropriate concentrations of osmium and thiocarbohydrazide is essential to preserve contrast in the membrane structure of the series images. Osmium, in particular, plays a crucial role by binding with lipid molecules, enhancing the BSE signal.
Our retinal sample preparation protocol is user-friendly, requires no additional equipment, and minimizes processing time. This characteristic highlights its potential applicability to other Volume EM techniques, suggesting versatility and ease of adoption in diverse experimental setups. The OTO&#39;s method notably enhances the contrast of FIB-SEM images. However, the details about the cell membrane, especially the bilayer structure, may still appear somewhat blurred. Our current sample preparation scheme involves some sacrifice of small details in cell structure to a certain extent. We aim to develop optimized methods in the future to address and improve this limitation.

The retina is densely packed with intertwining neuronal axons and dendrites that form synapses between them1. Microscopy is an indispensable tool for studying retinal anatomy as it has fine, intricate, and small structures. Although electron microscopy (EM) provides unparalleled power to investigate the ultrastructure of subcellular organelles and the accurate localization of specific proteins at the nanometer level2, it produces images limited to the two-dimensional (2D) plane, leading to potential loss of key information.
The development of emerging high-resolution volume electron microscopy (Volume EM) techniques supports the provision of more comprehensive and larger-scale three-dimensional (3D) structural information. Some 3D EM methods have been recently reviewed by others3,4,5. 3D EM allows for the reconstruction of neuronal shape and connectivity details, enabling precise quantitative analysis of structures of interest. This demonstrates that the data obtained by volume EM are more systematic, complete, and accurate.
Retinal photoreceptors, constituting the initial neurons in visual signaling6,7, establish synapses with dendrites of second-order bipolar and horizontal cells in the photoreceptor&#39;s terminal to facilitate excitatory signals8,9. These terminals, referred to as cone pedicles and rod spherules, encompass three crucial components: mitochondria, synaptic ribbons, and synaptic vesicles. While previous studies have predominantly concentrated on the general structure of ribbon synapses, there has been a notable absence of investigation into the fine structure of major components, including mitochondria, ribbon, vesicle pools, and their spatial organization in terminals10,11,12,13. A precise and systematic analysis of each component, along with an understanding of their inter-association within photoreceptor terminals, is vital for unraveling spatial organization and comprehensively grasping visual processing functions. In photoreceptors, mitochondria are mainly present in the inner segment, cell body, and terminal. We focused here on the mitochondria in the terminal photoreceptors. Focused ion beam scanning electron microscopy (FIB-SEM), a type of volume EM boasting high resolution (x, y, and z resolution &#60; 5 nm) and a relatively large volume flux4,14, stands as a potent tool for accurately visualizing the 3D structure of photoreceptor terminals.
Both FIB-SEM and Serial Block Face Scanning Electron Microscope (SBF-SEM) are Volume EM based on SEM for obtaining the image of tissues by scanning the surface of the sample. The ultrastructural features of a specimen&#39;s surface are revealed through the contrast created by the intensity of secondary or backscattered electrons (BSE) when the electron beam scans the sample15. Essentially, detecting BSE or secondary electrons from the cross-section surface of a resin-embedded tissue sample in SEM allows for obtaining images of the embedded sample16,17. When BSE or secondary electrons are less generated, information on the sample surface can only be obtained. Achieving consistent contrast and high-quality serial images necessitates sufficient deposition of heavy metals in the sample. Therefore, specific sample preparation protocols are crucial for subsequent segmentation, 3D reconstruction, and analysis when utilizing SEM for serial imaging. The osmium-thiocarbohydrazide-osmium (OTO) method is a typical sample preparation scheme for 3D electron microscopy of biological samples, preserving the structure of lipid-containing membranes and maintaining good contrast18,19.
Here, we developed the OTO method for the preparation of retinal samples for the use of Volume EM. This process particularly focuses on dissecting the retina, determining the optimal fixation time for retinal tissue, and detailing the specific procedures and precautions in 3D sample preparation. Additionally, segmentation and 3D reconstruction of the target structure are integral steps in this extended application. The retina, being a small and challenging structure to obtain materials from, requires swift and precise operations for EM, with fixed times and fresh reagents prepared for immediate use.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>2,2,2-Tribromoethanol</td>
			<td>Sigma-Aldrich</td>
			<td>T48402</td>
			<td></td>
		</tr>
		<tr>
			<td>Acetone</td>
			<td>Electron Microscopy Science</td>
			<td>10000</td>
			<td></td>
		</tr>
		<tr>
			<td>Amira 6.8</td>
			<td>Thermo Fisher Scientific</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>CaCl2</td>
			<td>Sigma</td>
			<td>C-2661</td>
			<td></td>
		</tr>
		<tr>
			<td>Embedding mold</td>
			<td>Beijing Zhongjingkeyi Technology</td>
			<td>GP10590</td>
			<td></td>
		</tr>
		<tr>
			<td>Epon resin</td>
			<td>Electron Microscopy Science</td>
			<td>14900</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethanol</td>
			<td>Sigma</td>
			<td>64-17-5</td>
			<td></td>
		</tr>
		<tr>
			<td>Glutaraldehyde</td>
			<td>Electron Microscopy Science</td>
			<td>16020</td>
			<td></td>
		</tr>
		<tr>
			<td>Helios NanoLab 600i dual-beam SEM</td>
			<td>FEI</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>L-aspartic acid</td>
			<td>Sigma</td>
			<td>56-84-8</td>
			<td></td>
		</tr>
		<tr>
			<td>Lead nitrate</td>
			<td>Sigma</td>
			<td>10099-74-8</td>
			<td></td>
		</tr>
		<tr>
			<td>Na2HPO4.12H2O</td>
			<td>Sigma</td>
			<td>71650</td>
			<td>A component of phosphate buffer</td>
		</tr>
		<tr>
			<td>NaH2PO4.H2O</td>
			<td>Sigma</td>
			<td>71507</td>
			<td>A component of phosphate buffer</td>
		</tr>
		<tr>
			<td>OsO4</td>
			<td>TED PELLA</td>
			<td>4008-160501</td>
			<td></td>
		</tr>
		<tr>
			<td>Paraformaldehyde</td>
			<td>Electron Microscopy Science</td>
			<td>157-8</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium ferrocyanide</td>
			<td>Sigma</td>
			<td>14459-95-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium cacodylate</td>
			<td>Sigma</td>
			<td>6131-99-3</td>
			<td></td>
		</tr>
		<tr>
			<td>Sputter coater</td>
			<td>Leica</td>
			<td>ACE200</td>
			<td></td>
		</tr>
		<tr>
			<td>Thiocarbohydrazide</td>
			<td>Sigma</td>
			<td>2231-57-4</td>
			<td></td>
		</tr>
		<tr>
			<td>Uranyl acetate</td>
			<td>TED PELLA</td>
			<td>CA96049</td>
			<td></td>
		</tr>
	</tbody>
</table>

preparation of retinal samples for volume electron microscopy

Volume electron microscopy (Volume EM) has emerged as a powerful tool for visualizing the 3D structure of cells and tissues with nanometer-level precision. Within the retina, various types of neurons establish synaptic connections in the inner and outer plexiform layers. While conventional EM techniques have yielded valuable insights into retinal subcellular organelles, their limitation lies in providing 2D image data, which can hinder accurate measurements. For instance, quantifying the size of three distinct synaptic vesicle pools, crucial for synaptic transmission, is challenging in 2D. Volume EM offers a solution by providing large-scale, high-resolution 3D data. It is worth noting that sample preparation is a critical step in Volume EM, significantly impacting image clarity and contrast. In this context, we outline a sample preparation protocol for the 3D reconstruction of photoreceptor axon terminals in the retina. This protocol includes three key steps: retina dissection and fixation, sample embedding processes, and selection of the area of interest.

Figure 1A shows the image of retinal photoreceptor terminals prepared using the traditional chemical double fixation method, and Figure 1B shows the image of retinal photoreceptor terminals prepared using the OTO method. Both were sampled by FIB-SEM. It can be clearly seen that the cell membrane structure can be retained as much as possible by using the OTO method, and even the outline of vesicles can be clearly seen. In addition, the contrast of images obtained by using the OTO method is also clearer.
Figure 2 illustrates the experimental workflow employed for reconstructing the ultrastructural characteristics of photoreceptor terminals. The reconstruction focused on four types of photoreceptor terminals, including M-cone pedicles, S-cone pedicles, rod1 spherules, and rod2 spherules. The main emphasis was on essential components, such as mitochondria, synaptic ribbons, and synaptic vesicle pools within the photoreceptor terminals.
Figure 3 displays the reconstructed mitochondria and synaptic ribbons in the rod photoreceptor terminals. Notably, the synaptic ribbons of the rod2 spherules exhibit a distinctive horseshoe shape, typically featuring a single ribbon per spherule. Moreover, Figure 2 highlights the presence of two mitochondria in rod spherules, with a consistent position within the central space of the terminals, located above the synaptic ribbons. All the structures were traced manually, page by page.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/66589/66589fig01.jpg" /> 
Figure 1: Examples of images sampled using FIB-SEM for traditional chemical double fixation method and OTO method. (A) Traditional chemical double fixation method. (B) OTO method. Scale bar: 1000 nm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66589/66589fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/66589/66589fig02.jpg" /> 
Figure 2: 3D reconstruction workflow for photoreceptor terminals using FIB-SEM. Following alignment using 3D data visualization software, the serial images were manually segmented and rendered for further analysis. (A) Original serial images. (B) Aligned images. (C) Segmented images. (D) Reconstructed images. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66589/66589fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/66589/66589fig03.jpg" /> 
Figure 3: 3D reconstruction. The representative 3D reconstruction images depict photoreceptor terminals in white, mitochondria in orange, and synaptic ribbons in blue. Scale bar: 500 nm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66589/66589fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Author Spotlight: Retinal Neuroscience Studies with Volume Electron Microscopy

This protocol describes the detailed steps for preparing retinal samples for volume electron microscopy, focusing on the structural features of retinal photoreceptor terminals.

Preparation of Retinal Samples for Volume Electron Microscopy

Volume electron microscopy (Volume EM) has emerged as a powerful tool for visualizing the 3D structure of cells and tissues with nanometer-level ...

preparation-of-retinal-samples-for-volume-electron-microscopy

Nanyang Technological University

Research

JoVE Journal

Neuroscience

471 Views.  Wenzhou Medical University. This protocol describes the detailed steps for preparing retinal samples for volume electron microscopy, focusing on the structural features of retinal photoreceptor terminals.

Video: Author Spotlight: Retinal Neuroscience Studies with Volume Electron Microscopy

Preparation of Mouse Brain Tissue for Immunoelectron Microscopy

Transmission electron microscopy (TEM) is extremely useful for visualizing microglial, oligodendrocytic, astrocytic, and neuronal subcellular compartments (dendrite, dendritic spine, axon, axon terminal, perikaryon), as well as their intracellular organelles and cytoskeleton, in the central nervous system at high spatial resolution. Combined with TEM, pre-embedding immunocytochemistry allows the discrimination of cellular elements with few distinctive features and identification criteria (e.g., microglial perikarya and processes, when using an antibody against the microglia-specific marker Iba1 (ionized calcium binding adaptor molecule 1; as presented here)), identifying the neurotransmitter contents of cellular elements (e.g., serotonergic) and their ultrastructural localization of soluble or membrane-bound proteins (e.g., 5 HT1A and EphA4 receptors). Here, we describe a protocol for transcardiac perfusion of mice with acrolein fixative, removal and sectioning of the brain, as well as immunoperoxidase-diaminobenzidine (DAB) staining, resin embedding, and ultrathin sectioning of the brain sections. Upon completion of these procedures, the immunostained material is ready for examination with TEM. When rigorously performed, this technique provides an excellent compromise between optimal ultrastructural preservation and immunocytochemical detection.

We describe a protocol for transcardiac perfusion of mice, removal and sectioning of the brain, as well as immunoperoxidase staining, resin embedding, and ultrathin sectioning of the brain sections. Upon completion of these procedures, the immunostained material is ready for examination with transmission electron microscopy.

Transmission electron microscopy (TEM) is extremely useful for visualizing microglial, oligodendrocytic, astrocytic, and neuronal subcellular compartments ...

An Isolated Retinal Preparation to Record Light Response from Genetically Labeled Retinal Ganglion Cells

The first steps in vertebrate vision take place when light stimulates the rod and cone photoreceptors of the retina 1. This information is then segregated into what are known as the ON and OFF pathways. The photoreceptors signal light information to the bipolar cells (BCs), which depolarize in response to increases (On BCs) or decreases (Off BCs) in light intensity. This segregation of light information is maintained at the level of the retinal ganglion cells (RGCs), which have dendrites stratifying in either the Off sublamina of the inner plexiform layer (IPL), where they receive direct excitatory input from Off BCs, or stratifying in the On sublamina of the IPL, where they receive direct excitatory input from On BCs. This segregation of information regarding increases or decreases in illumination (the On and Off pathways) is conserved and signaled to the brain in parallel.
The RGCs are the output cells of the retina, and are thus an important cell to study in order to understand how light information is signaled to visual nuclei in the brain. Advances in mouse genetics over recent decades have resulted in a variety of fluorescent reporter mouse lines where specific RGC populations are labeled with a fluorescent protein to allow for identification of RGC subtypes 2 3 4 and specific targeting for electrophysiological recording. Here, we present a method for recording light responses from fluorescently labeled ganglion cells in an intact, isolated retinal preparation. This isolated retinal preparation allows for recordings from RGCs where the dendritic arbor is intact and the inputs across the entire RGC dendritic arbor are preserved. This method is applicable across a variety of ganglion cell subtypes and is amenable to a wide variety of single-cell physiological techniques.

This article provides a description of how to dissect and record from the isolated retinal preparation in mouse. In particular, we describe how to record light responses from a fluorescently labeled ganglion cell population and subsequently identify and analyze its morphology.

The first steps in vertebrate vision take place when light stimulates the rod and cone photoreceptors of the retina 1.  This information is then ...

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 ...

Preparation of Primary Neurons for Visualizing Neurites in a Frozen-hydrated State Using Cryo-Electron Tomography

Neurites, both dendrites and axons, are neuronal cellular processes that enable the conduction of electrical impulses between neurons. Defining the structure of neurites is critical to understanding how these processes move materials and signals that support synaptic communication. Electron microscopy (EM) has been traditionally used to assess the ultrastructural features within neurites; however, the exposure to organic solvent during dehydration and resin embedding can distort structures. An important unmet goal is the formulation of procedures that allow for structural evaluations not impacted by such artifacts.
Here, we have established a detailed and reproducible protocol for growing and flash-freezing whole neurites of different primary neurons on electron microscopy grids followed by their examination with cryo-electron tomography (cryo-ET). This technique allows for 3-D visualization of frozen, hydrated neurites at nanometer resolution, facilitating assessment of their morphological differences. Our protocol yields an unprecedented view of dorsal root ganglion (DRG) neurites, and a visualization of hippocampal neurites in their near-native state. As such, these methods create a foundation for future studies on neurites of both normal neurons and those impacted by neurological disorders.

To preserve neuronal processes for ultrastructural analysis, we describe a protocol for plating of primary neurons on electron microscopy grids followed by flash freezing, yielding samples suspended in a layer of vitreous ice. These samples can be examined with a cryo-electron microscope to visualize structures at the nanometer scale.

Neurites, both dendrites and axons, are neuronal cellular processes that enable the conduction of electrical impulses between neurons. Defining the ...

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 ...

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. ...

Microdissection and Whole Mount Scanning Electron Microscopy Visualization of Mouse Choroid Plexus

The choroid plexus (CP), a highly vascularized structure protruding into the ventricles of the brain, is one of the most understudied tissues in neuroscience. As it is becoming increasingly clear that this tiny structure plays a crucial role in health and disease of the central nervous system (CNS), it is of utmost importance to properly dissect the CP out of the brain ventricles in a way that allows downstream processing, ranging from functional to structural analysis. Here, isolation of the lateral and fourth brain ventricle mouse CP without the need for specialized tools or equipment is described. This isolation technique preserves the viability, function, and structure of cells within the CP. On account of its high vascularization, the CP can be visualized floating inside the ventricular cavities of the brain using a binocular microscope. However, transcardial perfusion required for downstream analysis can complicate the identification of the CP tissue. Depending on the further processing steps (e.g., RNA and protein analysis), this can be solved by visualizing the CP via transcardial perfusion with bromophenol blue. After isolation, the CP can be processed using several techniques, including RNA, protein, or single cell analysis, to gain further understanding on the function of this special brain structure. Here, scanning electron microscopy (SEM) on whole mount CP is used to get an overall view of the structure.

The choroid plexus (CP), an understudied tissue in neuroscience, plays a key role in health and disease of the central nervous system. This protocol describes a microdissection technique for isolating the CP and the use of scanning electron microscopy to obtain an overall view of its cellular structure.

Preparation of Retinal Samples for Volume Electron Microscopy

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Preparation of Mouse Brain Tissue for Immunoelectron Microscopy

An Isolated Retinal Preparation to Record Light Response from Genetically Labeled Retinal Ganglion Cells

Focussed Ion Beam Milling and Scanning Electron Microscopy of Brain Tissue

Preparation of Primary Neurons for Visualizing Neurites in a Frozen-hydrated State Using Cryo-Electron Tomography

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

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

Preparation of Mouse Retinal Cryo-sections for Immunohistochemistry

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

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

Microdissection and Whole Mount Scanning Electron Microscopy Visualization of Mouse Choroid Plexus