Preparation of Mouse Retinal Cryo-sections for Immunohistochemistry

Summary

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

Abstract

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

Introduction

Immunohistochemistry (IHC) is a powerful technique for localizing specific proteins and cellular structures in tissues in situ^1,2,3. Inappropriate fixation methods and sub-optimal sectioning of complex tissues can disrupt tissue structure, generate high background staining or diminish antibody-epitope interactions, resulting in staining artifacts and consequent misinterpretation of IHC data⁴. As the vertebrate retina is a complex and highly organized neural organ composed of strata of interconnected photoreceptors, interneurons and ganglion cells, it is very fragile and can be easily disrupted during dissection and sectioning. A detailed, standardized, and validated protocol from mouse eye dissection and orientation to immunostaining will help significantly reduce IHC artifacts, thereby, increasing the reliability of the results and allowing for more accurate comparative data analysis.

There are many protocols for tissue preparation for IHC, however, not all are suitable for retinal tissue. Strong fixatives like 10% formalin or Bouin's solution preserve retinal structure during dissection and sectioning⁵. Unfortunately, strong fixatives often lead to the enhanced background fluorescence and epitope masking due to the chemical modification of epitopes⁶. On the other hand, milder fixatives, like 4% paraformaldehyde (PFA) can alleviate some of these artifacts but require meticulous dissection and sectioning to preserve the optimal retinal structure. PFA rapidly penetrates tissue, but cross-links proteins very slowly, reducing the risk of epitope masking. Since short time PFA incubation is a relatively mild fixation, tissues often require rapid freezing to preserve antigens. It is important to avoid ice crystal formation during tissue freezing as they distort and damage the integrity of cells and tissues⁷.

Here we describe detailed and standardized protocols for dissection, fixation, and cryo-protection of mouse ocular posterior cups that yield consistent and reliable IHC data.

Protocol

All methods described here were carried out in strict accordance with the recommendations in the National Institutes of Health Guide for the Care and Use of Laboratory Animals provided by the Institute of Laboratory Animal Resources and were approved by the Institutional Animal Care and Use Committee of the University of Pennsylvania.

All tools and equipment for the methods are shown in Figure 1 and listed in the Table of Materials.

1 . Mouse eye enucleation, eye cup dissection, and embedding

1. Euthanize mice with carbon dioxide (CO₂) followed by either decapitation (postnatal mice P0 - P11) or cervical dislocation (adult >P11 mice). For P0-P14 mice, eyes are covered by skin. Ensure to remove the skin prior to subsequent steps.
2. Mark the temporal part of the eye (Figure 2A) by slightly burning the cornea using a tail cauterizer. Avoid burning a hole in the cornea by touching the cornea very lightly and for no more than a split second with the cauterizer.
3. Immediately enucleate mouse eyes using curved Dumont #5/45 forceps. Incubate for 15 min in a 1.5 mL cryotube tube with 1 mL of 4% paraformaldehyde (PFA) in phosphate-buffered saline (PBS) on ice.
4. After 15 min in 4% PFA, transfer the fixed eye using the curved forceps Dumont #5/45 to a modified 35 mm dissection dish (see Table of Materials and Figure 1C) filled with PBS and place under dissecting microscope.
5. Make a small incision at the burn mark, perpendicular to and just above the limbus (border of the cornea) using the micro knife.
6. Insert the curved scissors into the incision and perform a circumferential cut of the cornea following the limbus (Figure 2B).
7. Remove the cornea and extract the lens with a thin Dumont #5 forceps, (Figure 2B) and lift it delicately away from the posterior portion of the eye. The vitreous body, the clear gel that fills the space between the lens and the retina, will come out with the lens (Figure 2B) and the retina will be visible as a white surface covering the inside of the posterior eye cup. Ensure that there is no visible damage to the retina, such as holes or tears (Figure 2C).
8. Wash the eye cups twice in 1 mL of PBS for 10 min at room temperature.
9. Cryoprotect the eye cups by equilibrating them in a solution of 15% sucrose in PBS overnight at 4 °C until the eye cup sinks
10. Transfer the eye cups to 30% sucrose in PBS for 2-3 h until the eye cup sinks.
11. Embed the eye cups in OCT in 10 x 10 mm cryomolds (Figure 3A). Using a dissecting microscope, orient the eye in the cryomold along its dorsal-ventral axis (Figure 2D, Figure 3A) by rotating the eye until the burn mark is on top, the optic nerve is on the left and the cut-out part of the mold faces the right (Figure 2D). Take care to avoid bubbles near the tissue.
    NOTE: To manipulate the eye cup, use a titanium probe made of a titanium wire attached to a pipette tip.
12. To snap-freeze retinal tissue, immerse the cryomold for at least 5 min in a metal beaker containing isopentane and place the beaker in liquid nitrogen or dry ice/100% ethanol bath (up to 1/3 of the height of the metal beaker).
    NOTE: The cryomold should be immersed in isopentane to greater than half of its height but does not have to be completely submerged.
13. Remove the frozen block and wrap it in aluminum foil.
14. Store at -80 °C.
15. Mark the frozen block using a pen or sharpie to record the orientation of the block (Figure 3B).

2 . Sectioning using a cryostat

1. After adjusting the cryostat temperature (between -20 and -25 °C), allow the molds containing the embedded eye cups to equilibrate to the cryostat temperature for 1 h.
2. Install the block with dorso-ventral orientation (Figure 3C) and cut 10 µm thick serial sections. Carefully place the retinal sections on annotated and numbered microscope slides.
3. Store slides in slide boxes at -20 °C or -80 °C until IHC procedures.

3 . Fluorescence immunostaining using the Slide Rack

NOTE: The Slide Rack system (e.g., Sequenza) holds the glass microscope slides with a cover plate, creating a ~100 µL capillary gap between the slide and the plate.

1. Thaw the cryo-sections at room temperature (RT) for 2 h to 4 h to allow them to dry and attach to microscope slides.
   NOTE: It is also possible to dry them at 30-37 °C for 30 min to 1 h.
2. Place slides in Slide Rack and wash the retinal sections twice in 100-200 µL PBS for 2 min.
3. Permeabilize the retinal sections by incubating them in 0.25% Triton-X / 0.05% NaN₃in PBS for 5 min at RT.
4. Add 100 µL of blocking solution to the slides.
5. Add 100 µL of primary antibody diluted in blocking solution to the slides.
6. Incubate retinal sections overnight at 4 ˚C.
   NOTE: During this time, cover the Slide Rack to avoid desiccation of the sections.
7. Wash the retinal sections 3 times, each for 15 min, using 200 µL PBS.
   NOTE: The addition of 0.001- 0.002% Triton-X100 to PBS (PBS-T) allow a more stringent wash but can disrupt some membranes and cellular structures.
8. Incubate retinal sections in fluorescent-labeled secondary antibodies for 1 h at RT. From now on protect from light.
9. Wash the retinal sections 3 times, each for 15 min.
10. Incubate the retinal sections at RT for 5 min with a nuclear marker such as Hoechst 33342 or DAPI solution.
11. Wash the retinal sections 1 time for 15 min.
12. Place a coverslip on top of a paper towel on the lab bench.
13. Add 2 drops of anti-fading mounting media to the center of the coverslip.
14. Turn the slide over to have the retinal cryo-section facing down.
15. Carefully mount the coverslip slowly, at a ~45˚ angle, tip the slide onto the mounting media.
    NOTE: Avoiding forming bubbles as you lower the slide in place.
16. Apply even pressure, using a heavy book or catalog (see below), to the slide-coverslip combination to eliminate excess mounting media.
    NOTE: To ensure consistency in sample preparation, always use the same object, (i.e., the same book or catalog) to apply even pressure to the coverslip during slide mounting.
17. Keep flat and allow to harden overnight at RT in the dark. Store slides flat at 4 °C indefinitely.
18. Image via wide field or confocal fluorescence microscopy (Figure 4).

Results

To illustrate how these protocols, ensure optimal retinal preservation for IHC, we probed retinal sections from P28 WT mice (C57BL/6N) with antibodies to rhodopsin (a photoreceptor marker)⁸, glutamic acid decarboxylase 65 (Gad65, an amacrine cell marker)⁹, glutamine synthetase (GS, a Müller cell marker)¹⁰, and calbindin (a horizontal cell marker)¹¹ (Figure 4

Discussion

Mouse retina dissection is a delicate process due to the small size and shape of mouse eyes and the fragility of retinal tissue. Even though performing high-quality dissection is a matter of practice, having a detailed protocol, providing efficient methods and tips is a necessity to obtain retinal sections and IHC. In addition to the protocols described here, there are several tips that allow for consistent high-quality retinal sections that are suitable for reproducible IHC.

Prior to commenci...

Materials

Name	Company	Catalog Number	Comments
6qt. Stainless Steel Beaker (185 x 218mm)	Gilson	#MA-48	For liquid nitrogen (Figure 1E)
Aluminum foil
Cauterizer	Bovie	#AA01	To mark eye orientation (Figure 1A)
Curved forceps, Dumont #5/45 tweezers, 45-degrees bent	Electron Microscopy Sciences	#72703-D	For mouse eye enucleation and maintenance (Figure 1A)
Curved scissors, Vannas Spring Scissors - 2.5mm Cutting Edge	Fine Science Tools	#15002-08	To circumferentially cut the cornea (Figure 1B)
Dental wax-coated 35mm dissection dish			For eye cup dissection (Figure 1C)
Dissecting microscope	Leica, Houston, USA	MZ12.5 High-performance stereomicroscope
Micro Knives - Plastic Handle	Fine Science Tools	#10315-12	To make a small incision at the burn mark (Figure 1A)
Pink dental wax	Electron Microscopy Sciences	#72660	For coating dissection dish
Slide Rack and Coverplate	Ted Pella	#36107	For retinal IHC (Figure 1G)
Stainless Steel Beaker (89 x 114mm)	Gilson	#MA-40	For isopentane bath (Figure 1E)
Styrofoam box			For insulated cooler
Thin forceps Dumont #5	Fine Science Tools	#11254-20	To remove the cornea and extract the lens (Figure 1B)
Tissue-Tek cryomold  (10mm x 10mm x 5mm)	Electron Miscroscopy Sciences	#62534-10	Mold for OCT embedding
Buffers and Reagents	Company	Catalog Number	Comments
Blocking solution			2% Normal Horse Serum, 1.5% Cold Fish skin Gelatin (at 40-50% in H2O, cat #G7765), 5% bovine serum albumin (cat #AK1391-0100) in permeabilization buffer.
Gelvatol (anti-fading mounting media)			Under the fume hood, dissolve 0.337g DABCO (Sigma cat #D2522-25G) in 10mL Fluoromount G (Fisher cat #OB100-01).  Adjust to pH 8-8.5 with 12N HCl (~ 5 drops).  Store in amber drop bottle at 4°C  (8).
Hoechst 33342 1000x Stock	Thermo Scientific	#62249	1 mg/ml in PBS (1000x). Aliquot and store in dark, at 4°C for up to 1 year or at -20°C for long term storage. Prepare fresh  at 1 ug/mL (1x)
Isopentane, 99%	GFS Chemicals	#2961
Liquid Nitrogen
Paraformaldehyde (PFA) in PBS	Electron Microscopy Sciences	#15710	Prepared fresh 4% PFA from 16% PFA stock. Store the remaining 16% PFA at 4°C in the dark.
Permeabilization solution			PBS + 0.25% Triton-X (AMRESCO cat #0694-1L) + 0.05% NaN3.  Prepare fresh.
Phosphate-buffered saline (PBS)	Bioworld	#41620015-20	0.02 g/L KCl, 0.02 g/L KH2PO4, 0.8 g/L NaCl, 0.216 g/L Na2HPO4 , pH 7.4.  Prepared from 10x stock
Sucrose solutions	Fisher Scientific	#S-5-500	Dissolve the appropriate amount of sucrose in  37°C PBS (takes ~15 min to dissolve). May be stored for a few days at 4°C.
Tissue-Tek OCT-compound	Sakura	#4583
Antibodies	Company	Catalog Number	Comments
anti-calbindin	Sigma-Aldrich	C8666	To label horinzotal cells
anti-GAD65	Chemicon	AB5082	To label GABAergic amacrine cells
anti-GS	BD Transduction	610517	To label Müller cells
anti-rhodopsin	Millipore	MAB5316	To label rods