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. M., Wasserkrug, H. L., Hanker, J. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+new+staining+method+(OTO)+for+enhancing+contrast+of+lipid-containing+membranes+and+droplets+in+osmium+tetroxide-fixed+tissue+with+osmiophilic+thiocarbohydrazide+(TCH).">A new staining method (OTO) for enhancing contrast of lipid-containing membranes and droplets in osmium tetroxide-fixed tissue with osmiophilic thiocarbohydrazide (TCH).</a> J Cell Biol. 30 (2), 424-432 (1966).</li></ol>

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><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>CaCl&lt;sub&gt;2&lt;/sub&gt;</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>Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;.12H&lt;sub&gt;2&lt;/sub&gt;O</td><td>Sigma</td><td>71650</td><td>A component of phosphate buffer</td></tr><tr><td>NaH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;.H&lt;sub&gt;2&lt;/sub&gt;O</td><td>Sigma</td><td>71507</td><td>A component of phosphate buffer</td></tr><tr><td>OsO&lt;sub&gt;4&lt;/sub&gt;</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/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/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/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

473 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

Retinal Pericytes Visualization Using Tissue-Clearing and Fluorescent Labeling

3D Visualization of Retinal Vascular Pericytes in Mice by Immunostaining

Retinal pericytes are essential for vascular development and stability of the retina, playing a key role in maintaining the integrity of the retinal vasculature. To provide a detailed view of the morphological characteristics of pericytes, this study described a new approach combining the retro-orbital injection of a fluorescent agent, immunofluorescent-staining, and tissue-clearing treatment. Firstly, the fluorescent tomato lectin was injected into the retro-orbital sinus of the live mouse to label the retinal vasculature. After 5 min, the mouse was sacrificed, and its intact retina was carefully removed from the retinal cup and immunofluorescently stained with platelet-derived growth factor receptor &#946; to reveal the pericytes. Additionally, the stained retina was treated with tissue-clearing reagent and whole mounted on the microscope slide. Through these approaches, the retinal vasculature and pericytes were clearly observed in the transparent retina. Under a confocal microscope, we obtained more opportunities to take high-resolution images for further reconstructing and analyzing the morphological characteristics of pericytes along the retinal vascular tree in a three-dimensional view. Methodologically, this protocol offers an effective approach for visualizing pericytes within the retinal vasculature, providing valuable insights into their role under both physiological and pathological conditions.

The pericytes in retinal vasculature were examined by immunofluorescent staining with platelet-derived growth factor receptor &#946; after retro-orbital injection of fluorescent tomato lectin. The labeled retina was further treated with the tissue-clearing method and whole mounted for visualizing the three-dimensional views of pericytes surrounding retinal vasculature under a confocal microscope.

Retinal pericytes are essential for vascular development and stability of the retina, playing a key role in maintaining the integrity of the retinal ...

Biochemistry

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 ...

Optimized Protocol for Retinal Wholemount Preparation for Imaging and Immunohistochemistry

Working with delicate tissue can be a complicating factor when performing immunohistochemical assessment. Here, we present a method that utilizes a ring-supported hydrophilized PTFE membrane to provide structural support to both living and fixed tissue during immunohistochemical processing, which allows for the use of a variety of protocols that would otherwise cause damage to the tissue. First, this is demonstrated with bolus loading of fluorescent markers into living retinal tissue. This method allows for quick visualization of targeted structures, while the membrane support maintains tissue integrity during the injection and allows for easy transfer of the preparation for further imaging or processing.
Second, a procedure for antibody staining in tissue fixed with carbodiimide is described. Though paraformaldehyde fixation is more common, carbodiimide fixation provides a superior substrate for the visualization of synaptic proteins. A limitation of carbodiimide is that the resulting fixed tissue is relatively fragile; however, this is overcome with the use of the supporting membrane. Retinal tissue is used to demonstrate these techniques, but they may be applied to any fragile tissue.

The protocol described here is for structural assessment of a wholemount retinal preparation. This includes descriptions of tissue dissection, mounting onto a hydrophilized polytetrafluoroethylene (PTFE) membrane insert, bolus loading with fluorescent markers, and a comparison of fixation with carbodiimide and paraformaldehyde for immunohistochemical analysis of cellular and synaptic components.

Working with delicate tissue can be a complicating factor when performing immunohistochemical assessment. Here, we present a method that utilizes a ...

Biology

Microsurgical Dissection and Tissue Clearing for High Resolution Intact Whole Retina and Vitreous Imaging

Neuronal and vascular structures of the retina in physiologic and pathologic conditions can be better visualized and characterized by using intact whole retina imaging techniques compared to conventional retinal flat mount preparations and sections. However, immunofluorescent imaging of intact whole retina is hindered by the opaque coatings of the eyeball, i.e., sclera, choroid, and retinal pigment epithelium (RPE) and the light scattering properties of retinal layers that prevent full thickness high resolution optical imaging. Chemical bleaching of the pigmented layers and tissue clearing protocols have been described to address these obstacles; however, currently described methods are not suitable for imaging endogenous fluorescent molecules such as green fluorescent protein (GFP) in intact whole retina. Other approaches bypassed this limitation by surgical removal of pigmented layers and the anterior segment of the eyeball allowing intact eye imaging, though the peripheral retina and hyaloid structures were disrupted. Presented here is an intact whole retina and vitreous immunofluorescent imaging protocol that combines surgical dissection of the sclera/choroid/retina pigment epithelium (RPE) layers with a modified tissue clearing method and light sheet fluorescent microscopy (LSFM). The new approach offers an unprecedented view of&#160;unperturbed vascular and neuronal elements of the retina as well as the vitreous and hyaloid vascular system in pathologic conditions.

Presented here is a protocol for intact whole retina imaging in which the outer opaque/pigmented layers of the eyeball are surgically removed, and optical clearing is applied to render retina transparent enabling the visualization of the peripheral retina and hyaloid vasculature in intact retina using light sheet fluorescent microscopy.

Neuronal and vascular structures of the retina in physiologic and pathologic conditions can be better visualized and characterized by using intact whole ...

Two Peeling Methods for the Isolation of Photoreceptor Cell Compartments in the Mouse Retina for Protein Analysis

Rod photoreceptors are highly polarized sensory neurons with distinct compartments. Mouse rods are long (~80 &#181;m) and thin (~2 &#181;m) and are laterally packed in the outermost layer of the retina, the photoreceptor layer, resulting in alignment of analogous subcellular compartments. Traditionally, tangential sectioning of the frozen flat-mounted retina has been used to study the movement and localization of proteins within different rod compartments. However, the high curvature of the rod-dominant mouse retina makes tangential sectioning challenging. Motivated by the study of protein transport between compartments, we developed two peeling methods that reliably isolate the rod outer segment (ROS) and other subcellular compartments for western blots. Our relatively quick and simple techniques deliver enriched and subcellular-specific fractions to quantitatively measure the distribution and redistribution of important photoreceptor proteins in normal rods. Moreover, these isolation techniques can also be easily adapted to isolate and quantitatively investigate the protein composition of other cellular layers within both healthy and degenerating retinae.

This protocol presents two techniques to isolate the subcellular compartments of murine rod photoreceptors for protein analysis. The first method utilizes live retinae and cellulose filter paper to separate rod outer segments, while the second employs lyophilized retinae and adhesive tape to peel away rod inner and outer segment layers.

Rod photoreceptors are highly polarized sensory neurons with distinct compartments. Mouse rods are long (~80 &#181;m) and thin (~2 &#181;m) and are ...

Quantification of Vascular Parameters in Whole Mount Retinas of Mice with Non-Proliferative and Proliferative Retinopathies

Retinopathies are a heterogeneous group of diseases that affect the neurosensory tissue of the eye. They are characterized by neurodegeneration, gliosis&#160;and a progressive change in vascular function and structure. Although the onset of the retinopathies is characterized by subtle disturbances in visual perception, the modifications in the vascular plexus are the first signs detected by clinicians. The absence or presence of neovascularization determines whether the retinopathy is classified as either non-proliferative (NPDR) or proliferative (PDR). In this sense, several animal models tried to mimic specific vascular features of each stage to determine the underlying mechanisms involved in endothelium alterations, neuronal death and other events taking place in the retina. In this article, we will provide a complete description of the procedures required for the measurement of retinal vascular parameters in adults and early birth mice at postnatal day (P)17. We will detail the protocols to carry out retinal vascular staining with Isolectin GSA-IB4 in whole mounts for later microscopic visualization. Key steps for image processing with Image J Fiji software are also provided, therefore, the readers will be able to measure vessel density, diameter and tortuosity, vascular branching, as well as avascular and neovascular areas. These tools are highly helpful to evaluate and quantify vascular alterations in both non-proliferative and proliferative retinopathies.

This article describes a well-established and reproducible lectin stain assay for the whole mount retinal preparations and the protocols required for the quantitative measurement of vascular parameters frequently altered in proliferative and non-proliferative retinopathies.

Retinopathies are a heterogeneous group of diseases that affect the neurosensory tissue of the eye. They are characterized by neurodegeneration, ...

Studying Bipolar and Horizontal Neuronal Cells in the Murine Split Retina

Split Retina as an Improved Flatmount Preparation for Studying Inner Nuclear Layer Neurons in Vertebrate Retina

Bipolar cells and horizontal cells of the vertebrate retina are the first neurons to process visual information after photons are detected by photoreceptors. They perform fundamental operations such as light adaptation, contrast sensitivity, and spatial and color opponency. A complete understanding of the precise circuitry and biochemical mechanisms that govern their behavior will advance visual neuroscience research and ophthalmological medicine. However, current preparations for examining bipolar and horizontal cells (retinal whole mounts and vertical slices) are limited in their capacity to capture the anatomy and physiology of these cells. In this work, we present a method for removing photoreceptor cell bodies from live, flatmount mouse retinas, providing enhanced access to bipolar and horizontal cells for efficient patch clamping and rapid immunolabeling. Split retinas are prepared by sandwiching an isolated mouse retina between two pieces of nitrocellulose, then gently peeling them apart. The separation splits the retina just above the outer plexiform layer to yield two pieces of nitrocellulose, one containing the photoreceptor cell bodies and another containing the remaining inner retina. Unlike vertical retina slices, the split retina preparation does not sever the dendritic processes of inner retinal neurons, allowing for recordings from bipolar and horizontal cells that integrate the contributions of gap junction-coupled networks and wide-field amacrine cells. This work demonstrates the versatility of this preparation for the study of horizontal and bipolar cells in electrophysiology, immunohistochemistry, and in situ hybridization experiments.

Author Spotlight: Using the Split Retina Technique for Enhanced Access and Accelerated Experiments 

This work presents an alternative flatmount retina preparation in which the removal of photoreceptor cell bodies enables faster antibody diffusion and improved patch pipette access to inner retinal neurons for immunohistochemistry, in situ hybridization, and electrophysiology experiments.

Bipolar cells and horizontal cells of the vertebrate retina are the first neurons to process visual information after photons are detected by ...

Optimized Protocol for Retinal Organoid Sample Preparation for TEM

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.

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.

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

Synaptic Circuits Analysis and Protein Localization in the Mouse Retina

Investigating Retinal Circuits and Molecular Localization by Pre&#45;Embedding Immunoelectron Microscopy

The retina comprises numerous cells forming diverse neuronal circuits, which constitute the first stage of the visual pathway. Each circuit is characterized by unique features and distinct neurotransmitters, determining its role and functional significance. Given the intricate cell types within its structure, the complexity of neuronal circuits in the retina poses challenges for exploration. To better investigate retinal circuits and cross-talk, such as the link between cone and rod pathways, and precise molecular localization (neurotransmitters or neuropeptides), such as the presence of substance P-like immunoreactivity in the mouse retina, we employed a pre-embedding immunoelectron microscopy (immuno-EM) method to explore synaptic connections and organization. This approach enables us to pinpoint specific intercellular synaptic connections and precise molecular localization and could play a guiding role in exploring its function. This article describes the protocol, reagents used, and detailed steps, including (1) retina fixation preparation, (2) pre-embedding immunostaining, and (3) post-fixation and embedding.

Author Spotlight: Understanding the Ultrastructural Basis of Retinal Synaptic Connectivity and Neurotransmitter Localization in Mice 

This protocol outlines the detailed steps of pre-embedding immunoelectron microscopy, with a focus on exploring synaptic circuits and protein localization in the retina.

The retina comprises numerous cells forming diverse neuronal circuits, which constitute the first stage of the visual pathway. Each circuit is ...

Retinal Cryo-sections, Whole-Mounts, and Hypotonic Isolated Vasculature Preparations for Immunohistochemical Visualization of Microvascular Pericytes

Retinal pericytes play an important role in many diseases of the eye. Immunohistochemical staining techniques of retinal vessels and microvascular pericytes are central to ophthalmological research. It is vital to choose an appropriate method of visualizing the microvascular pericytes. We describe retinal microvascular pericyte immunohistochemical staining in cryo-sections, whole-mounts, and hypotonic isolated vasculature using antibodies for platelet-derived growth factor receptor &#946; (PDGFR&#946;) and nerve/glial antigen 2 (NG2). This allows us to highlight advantages and shortcomings of each of the three tissue preparations for the visualization of the retinal microvascular pericytes. Cryo-sections provide transsectional visualization of all retinal layers but contain only a few occasional transverse cuts of the microvasculature. Whole-mount provides an overview of the entire retinal vasculature, but visualization of the microvasculature can be troublesome. Hypotonic isolation provides a method to visualize the entire retinal vasculature by the removal of neuronal cells, but this makes the tissue very fragile.

We demonstrate three different tissue preparation techniques for immunohistochemical visualization of rat retinal microvascular pericytes, i.e., cryo-sections, whole-mounts, and hypotonic isolation of the vascular network.

Preparation of Retinal Samples for Volume Electron Microscopy

Podsumowanie

Przeglądaj więcej filmów

Rozdziały w tym wideo

3D Visualization of Retinal Vascular Pericytes in Mice by Immunostaining

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

Optimized Protocol for Retinal Wholemount Preparation for Imaging and Immunohistochemistry

Microsurgical Dissection and Tissue Clearing for High Resolution Intact Whole Retina and Vitreous Imaging

Two Peeling Methods for the Isolation of Photoreceptor Cell Compartments in the Mouse Retina for Protein Analysis

Quantification of Vascular Parameters in Whole Mount Retinas of Mice with Non-Proliferative and Proliferative Retinopathies

Split Retina as an Improved Flatmount Preparation for Studying Inner Nuclear Layer Neurons in Vertebrate Retina

Preparing Retinal Organoid Samples for Transmission Electron Microscopy

Investigating Retinal Circuits and Molecular Localization by Pre-Embedding Immunoelectron Microscopy

Retinal Cryo-sections, Whole-Mounts, and Hypotonic Isolated Vasculature Preparations for Immunohistochemical Visualization of Microvascular Pericytes