Name: Whole-mount Retinal Organoid Visualization with Cellular Resolution (Video) | JoVE Journal
Uploaded: 2025-06-20T13:35:00.000+00:00
Duration: 9 min 20 s
Description: Learn whole-mount retinal organoid visualization using optical clearing and immunolabeling with confocal microscopy. This method reveals 3D neuronal layers and cellular connections for studying retinal development and disease mechanisms.

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5 minutes

We are thankful to Angel Sandoval, Eric Calatayud, Gustavo Castro-Olvera, Marta Martin, and Alina Hirschmann (ICFO-Institut de Ci&#232;ncies Fot&#242;niques) for their technical support. The funding entities that supported this work are Fundaci&#243; CELLEX; Fundaci&#243; Mir-Puig; Ministerio de Econom&#237;a y Competitividad - Severo Ochoa program for Centres of Excellence in R&#38;D (CEX2019-000910-S, [MCIN/AEI/10.13039/501100011033]); Ministerio de Ciencia, Innovaci&#243;n y Universidades - Agencia Estatal de Investigaci&#243;n (PID2021-122807OB-C31 and PID21-122807OB-C32); Instituto de Salud Carlos III (PI22/01747); Fundaci&#243;n La ONCE; Generalitat de Catalunya through CERCA program; Laserlab-Europe (EU-H2020 GA no. 871124); and Fondo Social Europeo (PRE2020-095721, M.C.).

1. Preparation of custom-made fluorescent secondary antibodies
NOTE: This approach serves as an alternative to the use of commercial fluorescent antibodies (see the discussion for the advantages of preparing fluorescent secondary antibodies). Moreover, this protocol can be used for conjugating fluorophores to primary antibodies in case direct immunofluorescence is to be performed and the fluorescent primary antibody is not commercially available. Controlling the fluorophore-to-antibody ratio by adjusting the amount of fluorophore in the labeling reaction produces brighter conjugated antibodies. However, custom-made secondary antibodies are less stable, and the concentration of the resulting conjugate is 10x lower than that of the commercial fluorescent antibodies. This protocol is adapted from the work described by Bálint et al9.
<ol>
	<li>Dissolve 1 mg of the fluorophore in anhydrous DMSO and aliquot into tubes for a final 0.02 mg of fluorophore per tube. Remove all DMSO using a speed vacuum concentrator, running for 1 h and 30 min. Store aliquots at -20 ºC protected from light. For Alexa Fluor 405, ultrapure H2O is the appropriate solvent for making aliquots.</li>
	<li>For performing the labeling reaction, dissolve one aliquot of the fluorophore in 1-10 µL of DMSO (or ultrapure H2O for Alexa Fluor 405). 
	NOTE: The exact amount of DMSO is determined through previous control conjugation tests. These tests are performed when a new batch of a fluorophore is to be used (or when running out of a previous fluorophore batch), or when a new fluorophore is used for the first time in the conjugation. The control conjugation test involves carrying out labelling reactions with varying amounts of fluorophore to determine the optimal fluorophore-to-antibody ratio in the conjugate. In our case, the optimal ratio typically falls between 3 µL and 5 µL.</li>
	<li>Incubate 50 µL of secondary IgG (or primary), 6 µL of 1 M NaHCO3, and 1-5 µL of fluorophore for 40 min at room temperature (RT) on a rocking platform, protected from light.</li>
	<li>While the reaction is progressing, prepare the purification size exclusion columns by removing its lids and letting the buffer pass through. Equilibrate the column (one for each labeling reaction) by running 3 x 2-3 mL of PBS through the column.In case the last equilibration step finishes before the incubation ends, put the lids back to avoid them drying up and wait until the labeling reaction finishes. 
	NOTE: Purification size exclusion columns work by gravity. Make sure that the column does not dry up at any moment, or the purification reaction will not work properly.</li>
	<li>When the reaction finishes (step 1.3), add 140 µL of PBS to the labeling reaction to make the volume approximately 200 µL and vortex it. Add this solution to the center of the column, allow the sample to enter the column, and after the last drop has eluted, push by adding 550 µL of PBS. When the liquid stops falling, elute by adding 300 µL of PBS and collect in a 1.5 mL microcentrifuge tube.</li>
	<li>Calculate the antibody concentration and labeling ratios by measuring the absorbance of the sample at 280 nm and the absorbance of the fluorophores used. Apply the Beer-Lambert Law to determine the concentration (in molarity) as follows: 
	<img alt="IgG concentration formula, diagram with absorbance and correction factor parameters for protein analysis." src="/files/ftp_upload/68384/68384eq1.jpg" style="opacity: 0.9;" data-altupdatedat="1788304762000" data-alt="Equation 1"> 
	Where A stands for the absorbance, ε for the molar extinction coefficient, and CF280 is a measure of the absorbance that the specific fluorophore has at 280 nm that serves as a correction factor. Then, the concentration of the antibody is: 
	<img alt="IgG concentration formula, Afluorophore*dilution/εfluorophore, quantitative analysis equation." src="/files/ftp_upload/68384/68384eq2.jpg" style="opacity: 0.9;" data-altupdatedat="1788304762000" data-alt="Equation 2"> 
	And the labeling ratio is: 
	<img alt="Labeling ratio formula, fluorophore to IgG (M), equation for fluorescence analysis." src="/files/ftp_upload/68384/68384eq3.jpg" data-altupdatedat="1788304762000" data-alt="Equation 3"></li>
	<li>Store the labeled antibodies at 4 ºC, protected from light, for up to 6 months.</li>
</ol>
2. Immunofluorescence labeling
<ol>
	<li>Fix the organoids with 4% paraformaldehyde (PFA) at RT for 45 min. 
	NOTE: OPTIONAL: Incubate the organoids with an antigen-retrieval solution (steps 2.2 and 2.3) to enhance the immunoreactivity of the antigens.</li>
	<li>Prepare 40 mL of antigen-retrieval solution by mixing 0.117 mg of sodium citrate tribasic dihydrate in ultrapure H2O and adjust to pH 9.</li>
	<li>Incubate the organoids with the antigen-retrieval solution with mild shaking (30 rpm) at 60 °C for 1 h.</li>
	<li>Permeabilize the organoids with PBST (PBS with 1% Triton X-100) with mild shaking at RT for 4 h.</li>
	<li>Block the organoids with blocking solution (2% bovine serum albumin (BSA) with 0.1% Triton X-100) at RT for overnight (or over 1 day). 
	NOTE: Primary and secondary antibody concentrations depend on the particular antibody. For immunofluorescence of whole-mount samples, we use a higher concentration in comparison to sectioned samples. We incubate the primary antibody at 10 µg/mL which, in most cases, corresponds to 1:100 from the stock (Table 1). We incubate our custom-made secondary antibodies at 20-50 µg/mL.</li>
	<li>Incubate the organoids with the primary antibodies diluted in washing solution (0.1% BSA with 0.1% Triton X-100) at 4 °C for 2 days with mild shaking.</li>
	<li>Wash the organoids for 3 x 15 min in washing solution with mild shaking at RT.</li>
	<li>Incubate the organoids with the secondary antibodies diluted in washing solution at 4 °C for 2 days with mild shaking.</li>
	<li>Wash the organoids for 3 x 15 min in washing solution with mild shaking at RT. 
	NOTE: After immunolabeling, fluorescent dyes of choice can be incubated. We normally use fluorescent nuclei markers, such as DAPI (1:1,000) and DRAQ5 (1:500), or phalloidin (1:100) for visualizing the actin filament network.</li>
	<li>Incubate the organoids with the fluorescent dyes diluted in washing solution at RT for 1 h with mild shaking.</li>
	<li>Wash the organoids for 3 x 15 min in washing solution with mild shaking at RT.</li>
</ol>
3. Optical clearing with FluoClear BABB
NOTE: 1-Propanol might compromise plastic containers, particularly for long-term exposure. To be safe, use glass containers.
<ol>
	<li>Dehydration step
	<ol>
		<li>Prepare 1-propanol solutions in ultrapure H2O at the given percentages (15%, 30%, 45%, 60%, 75%, 90%) and then adjust to pH 9.5 with trimethylamine. 
		CAUTION: Triethylamine is toxic if inhaled. Work under a fume hood. Solutions can be reused if stored properly (well-sealed container). Adjust the pH before usage.</li>
		<li>Incubate the samples with the increasing gradient of 1-propanol solutions (15%, 30%, 45%, 60%, 75%, 90%) for 2 h each at 30 °C with mild shaking.</li>
		<li>Incubate the samples with 100% 1-propanol overnight at 30 °C with mild shaking.</li>
	</ol>
	</li>
	<li>Clearing step 
	CAUTION: BABB is a potent solvent mixture that is known for its ability to dissolve or significantly damage many plastics. To be safe, use glass containers with compatible lids and work inside a fume hood.
	<ol>
		<li>Prepare the BABB by mixing Benzyl alcohol (BA) and Benzyl benzoate (BB) at a 1:2 ratio.</li>
		<li>Immerse the samples in BABB at RT overnight.</li>
		<li>Before imaging, refresh the BABB. 
		NOTE: For ~1 mm diameter organoids, BABB induces a clearing effect that is evident in tens of minutes. Depending on the size and nature of the sample, the degree of clearing could be different and/or require longer exposure time to the clearing solution. We recommend acquiring images before and after BABB immersion to improve the clearing protocol.</li>
		<li>Store BABB cleared samples with BABB solution for the long term at 4 °C. Protect from light to preserve the fluorescence.</li>
	</ol>
	</li>
</ol>
4. Whole-mount image acquisition
<ol>
	<li>Using a glass pipette, position the organoid in a glass-bottom Petri dish with a drop of BABB. Make sure that the sample contacts the surface of the coverglass. 
	CAUTION: Take care that the drop of BABB containing the sample does not come into contact with the plastic edges of the dish, or it will melt the plastic, compromising the sample and the quality of the imaging.</li>
	<li>Using an inverted confocal laser-scanning microscope equipped with low and high magnification objectives, acquire Z-stack images to gain the full 3D insight of the sample with cellular detail. 
	NOTE: For this study, a 10x numerical aperture (NA) 0.4 air objective and a 63x NA 1.4 objective were used, with a step size of 3.5 µm and 1 µm, respectively.</li>
</ol>
5. Image processing
<ol>
	<li>Recalibrate the step size of the Z-stack acquisition considering the refractive index (RI) mismatch between the clearing solution and the immersion media.
	<ol>
		<li>Calculate the recalibrated step size as follows: 
		<img alt="Recalibration formula for step size in microscopy, involving refractive indices." src="/files/ftp_upload/68384/68384eq4.jpg" data-altupdatedat="1788304762000" data-alt="Equation 4"> 
		For a sample embedded in BABB, imaged with an air objective with a 3.5 µm step size, it will be as follows: 
		<img alt="Recalibration formula for step size calculation; equation for optical measurement precision." src="/files/ftp_upload/68384/68384eq5.jpg" style="opacity: 0.9;" data-altupdatedat="1788304762000" data-alt="Equation 5"></li>
		<li>Using ImageJ, change the voxel depth (click on Image | Properties...) for the recalculated step size.</li>
	</ol>
	</li>
	<li>Create image projections of the Z-stack.
	<ol>
		<li>Using ImageJ, inspect the Z-stack in depth (click on Image | Stacks | Orthogonal views). XY, XZ, and YZ views of the retinal organoid will appear.</li>
		<li>Save the desired regions of interest (click on File | Save as...).</li>
	</ol>
	</li>
	<li>Create an animation of the 3D render of the Z-stack using processing software (see the Table of Materials).
	<ol>
		<li>Select the channels to be visualized by enabling their checkboxes.</li>
		<li>Adjust the Z-planes to visualize the interior of the sample (enable Clipping in the checkbox).</li>
		<li>Record the animation using the Movie editor (enable Movie editor in the checkbox). Drag and drop, and zoom in and out directly using the mouse. Click Add to include a different clip for the animation. If adjusting the Z-planes, enable the Clip planes option (click on Settings... | check Clip planes box).</li>
	</ol>
	</li>
</ol>

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The authors have no conflicts of interest to declare.

The protocol described here facilitates the study of the spatiotemporal distribution of the cells forming the retinal organoid at different stages of maturation, preserving its natural 3D architecture, a critical aspect for understanding these models in the context of future personalized medicine. It provides a detailed view of the retinal architecture at the level of distinct layers and individual cell types, enabling visualization of subcellular structures while preserving the global 3D context.
The alternative to whole-mount imaging is postprocessing the sample for performing imaging on conventional sections (paraffined or cryopreserved). While sectioning provides valuable information about the localization of proteins and structures, this data is restricted to a single plane, limiting the understanding of spatial cellular connections. As a result, features such as the proportion of cones and rods or the formation of axonal bundles in specific regions may not be accurately represented, complicating the interpretation in three dimensions. Retinal organoids are highly heterogeneous, with different regions potentially containing different cell types and/or exhibiting varying degrees of complexity and differentiation. Whole-mount imaging solves this problem as it allows analyzing heterogeneity gradients globally without relying on the selection of a subset of sections that might not represent the entirety of the sample.
Light-sheet fluorescence microscopy (LSFM)21,22,23 could be used as an alternative to laser-scanning confocal microscopy. In LSFM, the specimen is illuminated with a thin sheet of light of a few microns thick, and all the fluorescence generated from the illuminated optical section is detected using a camera. LSFM is a widefield-based detection technique that represents a highly efficient strategy for fast visualization of large volumes (up to centimeter-scale samples) with minimal photobleaching and phototoxicity effects. However, organoids may exhibit significant spherical aberration because light passing through the curved surfaces is refracted differently depending on the angle of incidence. In these experiments, this distortion produced a loss of focus, leading to blurring and compromising the resolution of deeper structures. For imaging our retinal organoids (approximately 1 mm in diameter), laser-scanning confocal microscopy proved to be the most effective approach.
This protocol is tailored to our experimental requirements. It can be used as a starting guide for researchers venturing into whole-mount imaging of optically cleared retinal organoids and should be adapted if those differ. In this section, the critical steps, modifications, and limitations of the methodology are described in detail.
A critical step is the decision on the clearing protocol. Many criteria need to be considered. First, consider the typology of the sample. Clearing bones is not the same as clearing brain tissue, just as clearing retinal organoids differs from clearing gastric organoids, as each requires a distinct approach. To visualize cleared organoids of a different type, it is essential to review the literature to identify protocols that have been optimized specifically for the sample of interest. Fructose/glycerol is a fast and non-toxic clearing method that has been proven to be useful for clearing organoids derived from different tissues (human airway, colon, kidney, liver, and breast tumor organoids, and mouse mammary gland organoids)12 but was not suitable for making human retinal organoids transparent. To the best of our knowledge, no study reports fructose/glycerol application to neural tissue/organoid. Therefore, we conclude that this clearing method is not appropriate for clarifying neural tissues. Further investigation is warranted to elucidate the underlying reasons for this incompatibility.
Another critical factor to consider is the FoV required, as it directly impacts the imaging strategy and the choice of optical components. Large FoVs are typically achieved with low magnification objectives, which are often designed for use without immersion media (e.g., air, RI = 1.00). These objectives experience a larger RI mismatch when imaging BABB-cleared samples (RI = 1.56), leading to shifts in depth focusing that must be corrected during post-processing. In contrast, oil immersion objectives (RI = 1.51) present less RI mismatch with clearing solutions like BABB (RI = 1.56) and, therefore, do not require recalibration.
Simple immersion methods usually do not achieve high transparency and are not suitable for acquiring high-quality images of whole-mount samples. Tissue clearing protocols influence effective resolution via swelling or shrinkage of the samples. Depending on the goal of the experiment, either swelling or shrinkage can be beneficial. Protocols that expand the sample can enhance the resolution of fine details in the same optical system incapable of showing the same details before. Shrinkage can be desired to visualize large samples that otherwise would not fit into the sample chamber or adjust to the objective working distances. However, because shrinkage compresses the sample and may distort internal distances, this technique is not suitable for dimensional assessment or precise measurement of intercellular distances.
Another important factor to be considered is compatibility with fluorescent proteins. Mostly organic solvent-based protocols quench fluorescent proteins as they require high temperatures or degrade the fluorescent proteins while embedding with RI-matching solutions. Fortunately, most of the methods are compatible with immunolabeling techniques. However, the compatibility of each epitope should be confirmed experimentally.
Moreover, penetration of large IgG molecules through tissue can be obstructed, decreasing staining efficiency and limiting visualization of the targeted structure to the sample surface. Staining whole-mount specimens with antibodies might take several days (or even weeks), depending on the size and the nature of the sample, substantially increasing the duration of the protocol.
Lastly, compatibility with the optical instrument is mandatory. Most organic solvent-based methods (except ECi) are very corrosive and not chemically compatible with commercial sample chambers and immersion objectives. Furthermore, some clearing solutions have an RI that does not match the range required by the available objectives (usually water).
There are some steps in the protocol that need special attention. FluoClear BABB, as a hydrophobic clearing method, requires a previous dehydration step. The pH plays an important role in preserving the fluorescence of endogenous proteins such as GFP. Then, it is crucial to ensure that the alcohol solutions have the pH properly adjusted. Dehydration is key for complete clearing. Optimal results are achieved by incubating the samples with an increasing gradient of alcohol solutions.
The size of the sample plays an important role in allowing antibodies to reach their specific targets. Antibodies might fail to penetrate homogeneously and be retained in the outside of your sample. If this occurs, consider switching to another type of detergent (Tween-20 or Saponin), increasing the concentration and incubation time of the detergent and the antibodies. If these adjustments are insufficient, alternative strategies can improve antibody permeabilization. For whole-mount immunolabelling of big specimens (centimeter-scale), researchers have found that the SDS can facilitate deeper antibody penetration24. SDS induces a partial loss of the tertiary structure of antibodies, effectively linearizing the protein and allowing it to penetrate deeper into the tissue. Then, washing out the SDS restores the antibodies&#39; antigen-binding properties, enabling more uniform staining of thicker tissues and reducing the commonly observed gradient of stronger labeling on the surface of the sample.
Preparing your own fluorescent secondary antibodies offers several advantages, as it provides greater flexibility in immunolabeling design, allowing for the selection of the specific fluorophore-secondary antibody combination, increasing the number of colors that can be used in the same sample. Additionally, the fluorophore-to-antibody ratio can be tuned, so they are generally brighter. However, some aspects must be considered. Custom-made secondary antibodies are generally less stable than the commercially available ones and should be used fresh. Most retain their ligand affinity for 3-6 months at 4 &#176;C, but stability needs to be experimentally determined, as some combinations could degrade more quickly. Moreover, some antibody-fluorophore pairings can alter the affinity of the antibody for its epitope, resulting in partial or complete nonspecific labeling. To ensure reliability, it is advisable to first test custom-made secondary antibodies using controls, such as cells in culture, before applying them to experimental specimens.
This optimized protocol integrates optical clearing with immunolabeling to facilitate confocal imaging of whole-mount retinal organoids. It preserves structural integrity and enables the identification of key retinal cell types within the organoid. Thus, the protocol provides researchers with a powerful tool to investigate retinal organoid maturation and spatial organization.

Retinal organoids
Retinal organoids are complex 3D structures that mimic the human retina, offering a valuable platform for studying retinal development, disease mechanisms, and potential therapeutic strategies1. They maintain the natural 3D architecture of the retina, demonstrate transcriptomic fidelity and functional competence, reproducing phototransduction and synaptic connectivity2. Additionally, they closely mimic in vivo development3,4, allowing for a more realistic representation as compared to flat cell cultures. In previous studies, Isla-Magran&#233; et al. established a two-step protocol to obtain multiocular organoids derived from human induced pluripotent stem cells (hiPSC) to model cellular features of the human eye in development5,6. Due to the scarcity of fetal tissue, human eye development research has been restricted to anatomical studies.
The retinal organoids obtained with this protocol exhibited proper layering and contained all major retinal cell subtypes, such as photoreceptors, bipolar cells, ganglion cells, amacrine and horizontal cells, and macroglia, along with key morphological features and correct spatial organization and connections. Organoid models, although a potential solution, exhibit significant variability, making it difficult to achieve consistent results. Therefore, there is a pressing need to employ a variety of advanced technologies in more reliable human models. To date, these organoids have only been studied using histological sections, as their complex cellular composition, density, and size hinder light penetration, resulting in poor optical imaging performance in deeper layers, thus limiting comprehensive analysis.
Whole-mount imaging
Traditional histological techniques, which involve sectioning and staining of fixed tissues, can provide detailed information about specific regions but fail to capture the overall architecture and spatial relationships within the entire organoid. To overcome this limitation, whole-mount imaging emerges as a valuable tool for studying retinal organoids. Whole-mount imaging allows visualization of the entire organoid in its intact 3D form, preserving the spatial distribution of cell types and extracellular matrix components. This approach is particularly beneficial for investigating the cellular composition within and between organoids, providing a more complete understanding of their developmental processes and functional organization. However, due to their cellular density and compaction, retinal organoids are opaque (Figure 1, uncleared). The inherent opaqueness of samples causes light scattering, which hinders detailed visualization of internal structures using microscopy. Here, optical clearing techniques emerge as a powerful tool.
Optical clearing techniques
Optical clearing refers to physicochemical treatments that make thick biological samples transparent by homogenizing the refractive index (RI) gradients of the tissue to match those of the clearing agent and removing light-absorbing substances7. These methods dramatically reduce light scattering and increase transparency. This transformation allows us to see through the entire retinal organoid, enabling comprehensive analysis of its intricate cellular organization and morphology through whole-mount imaging techniques. In practice, this is achieved by impregnating the sample with a high RI substance and removing lipids and pigments.
Different clearing methods are available, differing in various aspects (e.g., degree of transparency, compatibility with microscopic techniques, specimens, and fluorescent molecules, feasibility, duration), but none stands out as the best common protocol8. They are classified into four major groups: (1) simple immersion, (2) hydrophobic of organic solvent-based, (3) hyperhydration-based, and (4) tissue transforming- or hydrogel embedding-based, as nicely summarized by Avilov et al7. All methods include a step for the RI matching, usually by dipping the sample in a solution that matches the average RI of the tissue. In hydrophobic methods, lipids and water are removed by dipping the sample in an increasing organic solvent concentration (such as methanol or propanol).
The presented protocol is an optimized method that integrates optical clearing with immunolabeling to facilitate whole-mount 3D visualization of retinal organoids with confocal microscopy, enabling full-volume imaging while preserving structural integrity. This approach allows for the identification of key retinal cell types within the organoid, providing researchers with a powerful tool to investigate retinal organoid maturation and spatial organization in a physiologically relevant context.

<table><tbody><tr><td>10x NA0.40 DRY objective</td><td>Leica</td><td>HC PL APO CS2</td><td>-</td></tr><tr><td>63x NA1.40 OIL objective</td><td>Leica</td><td>HC PL APO CS2</td><td>-</td></tr><tr><td>anti-GFP antibody</td><td>Abcam</td><td>ab13970</td><td>-</td></tr><tr><td>Benzyl alcohol (BA)</td><td>Sigma Aldrich</td><td>13162</td><td>-</td></tr><tr><td>Benzyl benzoate (BB)</td><td>Sigma Aldrich</td><td>W213802</td><td>-</td></tr><tr><td>bottom-glass petri dish</td><td>MatTek</td><td>P35G-1.5-14-C</td><td>-</td></tr><tr><td>Bovine Serum Albumine (BSA)</td><td>Sigma Aldrich</td><td>A7030</td><td>-</td></tr><tr><td>Chx10 antibody</td><td>Sta Cruz Biotech</td><td>sc-365519</td><td>-</td></tr><tr><td>Cytiva Illustra Nap-5 gel filtration columns, Sephadex G-25</td><td>Sigma Aldrich</td><td>17-0853-02</td><td>-</td></tr><tr><td>DAPI</td><td>Sigma Aldrich</td><td>D9542</td><td>-</td></tr><tr><td>DMSO</td><td>Sigma Aldrich</td><td>D8418</td><td>-</td></tr><tr><td>DRAQ5</td><td>Thermofisher</td><td>62251</td><td>-</td></tr><tr><td>Dye (carboxylic acid, succinimidyl ester)</td><td>Invitrogen or Abberior</td><td>-</td><td>-</td></tr><tr><td>ImageJ/Fiji&amp;nbsp;</td><td>NIH</td><td>&amp;nbsp;v1.50i</td><td>-</td></tr><tr><td>Inverted confocal laser-scanning microscope</td><td>Leica microsystems</td><td>TCS SP8 STED 3X-FALCON</td><td>-</td></tr><tr><td>LAS X 3D processing software</td><td>Leica microsystems</td><td>-</td><td>-</td></tr><tr><td>mQ H&lt;sub&gt;2&lt;/sub&gt;O</td><td>-</td><td>-</td><td>Ultrapure water from the Synergy water purification system</td></tr><tr><td>NaHCO&lt;sub&gt;3&lt;/sub&gt;</td><td>Acros Organics</td><td>424270010</td><td>-</td></tr><tr><td>Non-labelled secondary IgG</td><td>Jackson Immunoresearch</td><td>-</td><td>-</td></tr><tr><td>Opsin blue antibody</td><td>Millipore&amp;nbsp;&amp;nbsp;</td><td>AB5407&amp;nbsp;</td><td>-</td></tr><tr><td>Opsin RG antibody</td><td>Millipore</td><td>AB5745&amp;nbsp;</td><td>-</td></tr><tr><td>Paraformaldehyde (PFA) 4%</td><td>Thermo scientific&amp;nbsp;</td><td>J199943-K2</td><td>-</td></tr><tr><td>PBS Solution</td><td>Gibco</td><td>18912-014</td><td>-</td></tr><tr><td>Phalloidin labelling probes</td><td>Thermofisher</td><td>-</td><td>-</td></tr><tr><td>Phosphate-Buffered Saline (PBS)</td><td>Gibco&amp;nbsp;</td><td>10010023</td><td>-</td></tr><tr><td>RECOV antibody</td><td>Millipore</td><td>AB5585-I&amp;nbsp;</td><td>-</td></tr><tr><td>RHO (RET-P1) antibody</td><td>Sigma Aldrich</td><td>O4886&amp;nbsp;</td><td>-</td></tr><tr><td>Sodium citrate tribasic dihydrate&amp;nbsp;</td><td>Sigma Aldrich</td><td>C8532</td><td>-</td></tr><tr><td>Triton X-100</td><td>Sigma Aldrich</td><td>T8787</td><td>-</td></tr><tr><td>TUJ1 antibody</td><td>Covance&amp;nbsp;</td><td>MMS-435P&amp;nbsp;</td><td>-</td></tr></tbody></table>

whole-mount retinal organoid visualization with cellular resolution

Whole-mount Retinal Organoid Visualization with Cellular Resolution

Retinal organoids derived from human induced pluripotent stem cells are intricate 3D structures that mimic the human retina, offering a powerful platform for studying retinal development, disease mechanisms, and potential therapeutic strategies. Moreover, as they are derived from patients, they are becoming increasingly popular as they hold great promise as a tool for personalized medicine. Unlike conventional 2D cell cultures, retinal organoids preserve the native 3D architecture of the retina, allowing for a more realistic representation and enabling more physiologically relevant studies. However, their structural complexity, high cellular density, and diverse composition present significant challenges for characterization. 
To address these challenges and enhance our understanding of retinal organoid maturation while preserving the 3D context, we combined optical clearing methods with immunolabeling to visualize the entire structure of whole-mount organoids with confocal microscopy. For this, we employed a clearing method compatible with low- and high-numerical-aperture objectives, facilitating full-volume imaging and capturing certain regions of interest with cellular resolution. Using this approach, we identified the morphology and distribution in 3D of the three main neuron paths responsible for the visual information transmission: cone and rod photoreceptors, bipolar and ganglion cells. Our findings shed more light on the visualization techniques to address the spatial organization of retinal cells within the organoid.

This protocol combines optical clearing and immunolabeling for full-volume confocal imaging of whole-mount retinal organoids. It preserves 3D structure, enabling detailed visualization of key neuronal pathways, improving the study of retinal maturation, spatial organization, and its potential applications in disease modeling and personalized medicine.

Retinal organoids derived from human induced pluripotent stem cells are intricate 3D structures that mimic the human retina, offering a powerful platform ...

whole-mount-retinal-organoid-visualization-with-cellular-resolution

Research

JoVE Journal

Neuroscience

The entire 3D structure and cellular content of organoids, as well as their phenotypic resemblance to the original tissue can be captured using the single-cell resolution 3D imaging protocol described here. This protocol can be applied to a wide range of organoids varying in origin, size and shape.

Single-Cell Resolution Three-Dimensional Imaging of Intact Organoids

Presented here is a surgical technique for transplanting human pluripotent stem cell (hPSC)-derived retinal tissue into the subretinal space of a large animal model.

Subretinal Transplantation of Human Embryonic Stem Cell-Derived Retinal Tissue in a Feline Large Animal Model

This study aims to develop a novel human organotypic retinal culture (HORC) model that prevents compromising retinal integrity during explant handling. This is achieved by culturing the retina with the overlying vitreous and the underlying retinal pigment epithelium-choroid (RPE-choroid) and sclera.

Characterization of a Novel Human Organotypic Retinal Culture Technique

Using a self-organizing method, we develop a protocol with the addition of COCO that could significantly increase the generation of photoreceptors.

Directed Induction of Retinal Organoids from Human Pluripotent Stem Cells

Here we describe an optimized retinal organoid induction system, which is suitable for various human pluripotent stem cell lines to generate retinal tissues with high reproducibility and efficiency.

Retinal Organoid Induction System for Derivation of 3D Retinal Tissues from Human Pluripotent Stem Cells

This protocol describes an efficient method of differentiating hiPSCs into eye field clusters and generating neuro-retinal organoids using simplified culture conditions involving both adherent and suspension culture systems. Other ocular cell types, such as the RPE and corneal epithelium, can also be isolated from mature eye fields in retinal cultures.

Generation of Retinal Organoids from Healthy and Retinal Disease-Specific Human-Induced Pluripotent Stem Cells

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

Preparing Retinal Organoid Samples for Transmission Electron Microscopy

Microglia are unique resident immune cells in the retina, playing crucial roles in various retinal degenerative diseases. Generating a co-culture model of retinal organoids with microglia can facilitate a better understanding of the pathogenesis and development progress of retinal diseases.

Assembling Retinal Organoids with Microglia

The pericytes in retinal vasculature were examined by immunofluorescent staining with platelet-derived growth factor receptor β 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.

3D Visualization of Retinal Vascular Pericytes in Mice by Immunostaining

The retina exhibits complex spatial organization across its laminar structure. Modern spatial transcriptomics and sequencing techniques require tissue sections &lt; 20 µm thick, limiting their retinal research applicability. This fresh-frozen cryosectioning method produces thin en face sections of mouse retina while preserving planar spatial relationships, enabling comprehensive molecular mapping across laminae.

En face Cryosectioning of Mouse Retina for High-dimensional Spatial Molecular Analysis

Learn whole-mount retinal organoid visualization using optical clearing and immunolabeling with confocal microscopy. This method reveals 3D neuronal layers and cellular connections for studying retinal development and disease mechanisms.

Whole-mount Retinal Organoid Visualization with Cellular Resolution

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