Immunostaining of Whole-Mount Retinas with the CLARITY Tissue Clearing Method

Summary

Here we present a protocol to adapt the CLARITY method of the brain tissues for whole-mount retinas to improve the quality of standard immunohistochemical staining and high-resolution imaging of retinal neurons and their subcellular structures.

Abstract

The tissue hydrogel delipidation method (CLARITY), originally developed by the Deisseroth laboratory, has been modified and widely used for immunostaining and imaging of thick brain samples. However, this advanced technology has not yet been used for whole-mount retinas. Although the retina is partially transparent, its thickness of approximately 200 µm (in mice) still limits the penetration of antibodies into the deep tissue as well as reducing light penetration for high-resolution imaging. Here, we adapted the CLARITY method for whole-mount mouse retinas by polymerizing them with an acrylamide monomer to form a nanoporous hydrogel and then clearing them in sodium dodecyl sulfate to minimize protein loss and avoid tissue damage. CLARITY-processed retinas were immunostained with antibodies for retinal neurons, glial cells, and synaptic proteins, mounted in a refractive index matching solution, and imaged. Our data demonstrate that CLARITY can improve the quality of standard immunohistochemical staining and imaging for retinal neurons and glial cells in whole-mount preparation. For instance, 3D resolution of fine axon-like and dendritic structures of dopaminergic amacrine cells were much improved by CLARITY. Compared to non-processed whole-mount retinas, CLARITY can reveal immunostaining for synaptic proteins such as postsynaptic density protein 95. Our results show that CLARITY renders the retina more optically transparent after the removal of lipids and preserves fine structures of retinal neurons and their proteins, which can be routinely used for obtaining high-resolution imaging of retinal neurons and their subcellular structures in whole-mount preparation.

Introduction

The vertebrate retina is perhaps the most accessible part of the central nervous system (CNS), and it serves as an excellent model for studying the development, structure, and function of the brain. Five classes of neurons in the retina are distributed in three nuclear layers separated by two plexiform layers. The outer nuclear layer (ONL) consists of classical photoreceptors (rods and cones) that convert light into electrical signals. Electrical signals are processed by neurons in the inner nuclear layer (INL), including bipolar, horizontal, and amacrine cells, and then transmitted to retinal ganglion cells (RGCs) in the ganglion cell layer (GCL). RGCs are the output neurons of the retina, with the axons projecting to the brain to contribute to image-forming and non-image-forming visual function. In addition, three types of glial cells (Muller cells, astroglia, and microglia) provide nutrients to neurons and protect neurons from harmful changes in their extracellular environment.

One specialized subpopulation of amacrine cells produces and releases dopamine, an important neuromodulator in the CNS, reconfiguring retinal neural circuits during light adaptation^1,2. Dopaminergic amacrine cells (DACs) have a unique feature of morphological profiles. Their somata are located in the proximal INL with dendrites ramifying in the most distal part of the inner plexiform layer (IPL). Axon-like processes of DACs are unmyelinated, thin and long, sparsely branched, and bear varicosities (the sites of dopamine release). They form a dense plexus with dendrites in the IPL, including ring-like structures around the somata of AII amacrine cells. The axons also run through the INL toward the OPL, forming a centrifugal pathway across the retina³. We have demonstrated that DAC processes express receptors in response to glutamate release from presynaptic neurons, including bipolar cells and intrinsically photosensitive retinal ganglion cells (ipRGCs)^4,5,6. However, it is unclear whether glutamate receptors express on the axons, dendrites, or both since they are cut off in vertical retinal sections and cannot be distinguished from each other^5,6. Immunostaining needs to be carried out in whole-mount retinas to reveal three-dimensional branching of DACs and the presence of glutamate receptors on subcellular compartments. Although the retina is relatively transparent, the thickness of a mouse whole-mount retina is approximately 200 µm, which limits the penetration of antibodies into the deep tissue as well as reduces light penetration for high-resolution imaging due to tissue light-scattering. To overcome these limitations, we adapted the immunostaining compatible tissue hydrogel delipidation method (CLARITY) developed recently for thick brain sections to whole-mount mouse retinas⁷.

The CLARITY method was originally developed by the Deisseroth laboratory for immunostaining and imaging of thick brain samples⁷. It uses a strong detergent, sodium dodecyl sulfate (SDS) and electrophoresis to remove the lipid components (that cause tissue light-scattering), leaving the proteins and nucleic acids in place. The removed lipids are replaced with a transparent scaffold made up of hydrogel monomers such as acrylamide to support the remaining protein structure. The cleared tissue can be labeled via immunohistochemistry and imaged with substantially increased light penetration depth through the tissue (up to several millimeters below the tissue surface). Since then, the CLARITY method has been optimized and simplified by several research groups^8,9,10. A modified CLARITY protocol uses a passive clearing technique to avoid the possible tissue damage produced by electrophoresis for clearing the whole-brain and other intact organs¹¹. However, this method has not yet been applied to whole-mount retinas. Here, we adapted the passive CLARITY technique for whole-mount retinas to make them more transparent for immunohistochemistry and imaging. We found that a majority of the retinal proteins tested were preserved during this process for immunohistochemistry. Using the refractive index matching solution, we were able to image retinal neurons across the approximately 200 µm thickness from the ONL to the GCL in whole-mount retinas.

Protocol

Mouse care and all experimental procedures were conducted according to the National Institutes of Health guidelines for laboratory animals and were approved by the Institutional Animal Care and Use Committees at Oakland University (protocol no. 18071).

NOTE: Names of the solutions and their compositions are listed in Table 1.

1. Tissue preparation

1. Euthanize the mouse with an overdose of CO₂, followed by cervical dislocation.
2. Enucleate the eyes with curved forceps and transfer them to a small petri dish with 0.1 M PBS (Table 1). Under a dissection microscope, poke a small hole along the cornea-sclera junction with a needle. Transfer to 4% paraformaldehyde (PFA) for 1 hour.
3. Transfer the eye back to a dish with PBS. Under a dissection microscope, use dissection scissors to cut all the way around the cornea-sclera junction. Remove the cornea and lens. Cut at the base of the optic nerve and carefully peel the sclera off with forceps to isolate the retina.
4. Make four small cuts evenly around the retina and use a fine tip brush dipped in PBS to lay it flat (GCL side down) in a clover-like shape on a small square cut from nitrocellulose filter paper to stabilize the retina.
5. Transfer the retina using forceps to hold the corner of the nitrocellulose paper (without touching the mounted retina) and place it in a 48-well plate with 4% PFA for 1 hour.
6. Transfer the filter paper and retina to a well with PBS and wash (3x for 5 min each).
7. Transfer to A4P0 (Table 1) and incubate overnight at 4 °C with gentle agitation.
8. Pipette vegetable oil into the well to completely cover the A4P0 solution. Incubate in a water bath at 40 °C for 3 hours with no shaking.
9. Wash (3x for 5 min each) in PBS, making sure all the oil has been rinsed off. If necessary, use a pipet to carefully remove remaining oil from the top of the well before the last rinse.
10. Incubate in 10% SDS at 40 °C for two days with gentle shaking. Replace SDS with fresh solution on the second day.
11. Transfer the filter paper and retina to PBS with Triton-X-100 (PBST, Table 1) and wash (5x for 1.5 h each).
12. Store at 4 °C in PBST with 0.01% sodium azide (NaN₃) or move directly to immunostaining.

2. Immunostaining and refractive index matching

1. Remove the retina from the filter paper by gently peeling it off with a fine tip brush in PBST.
2. Incubate the retina in primary antibody (Table 2) diluted in blocking solution (Table 1) for 2 days at 40 °C with gentle shaking.
3. Wash (5x, 1.5 h each) in PBST.
4. Incubate with the appropriate secondary antibodies (Table 3) diluted in blocking solution for 2 days at 40 °C with gentle shaking and protect from light through the remainder of the procedure.
5. Wash (5x, 1.5 h each) in 0.02 M phosphate buffer (see Table 1).
6. Incubate in sorbitol-based Refractive Index Matching Solution (sRIMS, see Table 1) at 40 °C overnight with gentle shaking.

3. Mounting

1. Outline a 18 mm x 18 mm x 1.5 mm glass coverslip with a fine-tip permanent marker to mark a square boundary on the back of a glass microscope slide.
2. Flip the slide over and use a syringe to trace the boundary with a thin line of silicone grease on the front of the slide, leaving a small gap in one corner for excess mounting solution to escape.
3. Transfer the retina to the center of the bounded area and arrange with a fine-tip brush so that it lies flat with the photoreceptor side against the glass slide.
4. Pipette approximately 60 µL of sRIMS so that it covers the flattened retina and extends to one corner of the enclosure, taking care that the retina stays flat and in place.
5. Apply the coverslip starting from the corner with the sRIMS and slowly lower it until it touches the grease on all sides, avoiding the formation of air bubbles.
6. Place a stack of 3 coverslips on each side of the mounted retina as a spacer. Use the long edge of another slide to press down the coverslip so that the mount is flat and even.
7. Store slides flat at 4 °C until imaging.

4. Imaging

1. Image samples on either a conventional fluorescence microscope or a confocal microscope (Table of Materials). Begin by placing the slide on the microscope stage and locating the sample.
   NOTE: If an inverted objective microscope is being used, place the slide upside down on the stage, first ensuring that the exposed areas of the slide are clear of all silicone grease and mounting solution.
2. To obtain z-stacked images of co-labeled samples, first focus on the signal in each channel individually and set the exposure time or scanning speed, for fluorescence or confocal microscopes, respectively.
3. Set the range for the z-stack either by manually setting the focal plane at the top and bottom of the desired range, or by setting the midpoint and then specifying a range around the midpoint.
4. Adjust the step size or number of slices as desired.
5. Capture the image and save the original file as well as exporting it as a TIFF file or other desired format.

5. Image analysis

1. Use the image analysis software of choice (Table of Materials) to adjust the brightness and contrast in each channel until optimum clarity is achieved in both the single images and the 3-dimensional rendering of the z-stack.
   NOTE: If the selected step size is sufficiently small, 3D deconvolution can also be performed to enhance the signal.

Results

Modified CLARITY-processed retinas are optically transparent tissue.
To formulate a tissue clearing method that is compatible with immunohistochemical applications in the retina while providing adequate delipidation and retaining the structural integrity of the cellular proteins, we adapted the CLARITY tissue clearing method to whole-mount mouse retinas. We were able to simplify the protocol and modify it for whole-mount retinas (see Protocol). After completing tissue hybridization, clearing, and r...

Discussion

Modification of the CLARITY protocol for whole-mount retinas.
We have simplified the CLARITY protocol to achieve adequate polymerization without the need for a vacuum evacuation or desiccation chamber, as is used in most previous studies^7,9,11. The polymerization process is inhibited by oxygen, requiring that the sample be isolated from air during the polymerization step of the protocol. However, rather th...

Materials

Name	Company	Catalog Number	Comments
16% Paraformaldehyde	Electron Microscopy Sciences	15710	Fixative
Acrylamide	Fisher Biotech	BP170	Hydrogel monomer
Axio Imager.Z2	Zeiss		Fluorscence microscope
BSA	Fisher Scientific	BP1600	Blocking agent
Eclipse Ti	Nikon Instruments		Scanning confocal microscope
KCl	VWR	BDH0258	Buffer component
KH2PO4	Sigma	P5655	Buffer component
Na2HPO4	Sigma Aldrich	S9763	Buffer component
NaCl	Sigma Aldrich	S7653	Buffer component
NaH2PO4	Sigma Aldrich	S0751	Buffer component
NaN3	Sigma Aldrich	S2002	Bacteriostatic preservative
NDS	Aurion	900.122	Blocking agent
NIS Elements AR	Nikon		Image analysis software
SDS	BioRad	1610301	Delipidation agent
Sorbitol	Sigma Aldrich	51876	Buffer component
Triton-X-100	Sigma	T8787	Surfactant
Tween-20	Fisher Scientific	BP337	Surfactant
VA-044	Wako Chemicals	011-19365	Thermal initiator