Morphological analysis of Schlemm’s canal in mice

Abstract

Immunofluorescent confocal microscopy is a powerful tool for analysis of the morphology and development of vascular and lymphatic tissues. Schlemm’s canal (SC) is a large, lymphatic-like vessel in the anterior chamber of the eye, which is essential for aqueous humor drainage required to maintain intraocular pressure and is sensitive to defects in blood and lymphatic vascular signaling pathways. Here, we describe a method to stain and quantify SC area and morphology in enucleated mouse eyes, providing a tool for understanding its development and function in small animal genetic or disease models.

1. Introduction

Schlemm’s canal (SC) is a large, lymphatic-like vessel in the anterior chamber of the eye and is essential for aqueous humor outflow [1–5]. Defects in SC structure and function lead to reduced aqueous humor outflow, elevated intraocular pressure (IOP) and glaucoma [2,5–8]. Imaging methods for evaluation of SC morphology and development are therefore valuable tools in the study of mouse models of ocular hypertension and glaucoma. While mice have a continuous SC morphologically similar to humans, both species exhibit marked variability in diameter along the length of the canal [9,6,3] making analysis using thin sections difficult and labor intensive. Recent studies have overcome this obstacle using whole-mount imaging of the enucleated eye, which allows 3-dimensional analysis of the full SC [2–4,6,7]. In mice this can be readily accomplished using confocal microscopy, as SC lies within 50 μm of the outer surface of the eye and a number of high-quality antibodies and fluorescent reporter mouse lines are available, which robustly label the SC endothelium. Here, we report a method to prepare and stain wholemounts of eye tissue from mice for confocal microscopy and SC analysis. We have successfully used this method to image the SC region in mice of all ages, from embryonic day 18.5 (before SC is present) until adulthood.

2. Materials

Prepare all solutions using ultrapure water and analytical grade reagents. Store all solutions at room temperature except for fixative and blocking buffer, which are stored at 4°C.

2.1: Fixation

1. Fixative: 2% formaldehyde in 0.1M phosphate buffer, pH 7.4. We prepare this solution as two 10X stocks, which can be quickly combined to prepare the fixative working solution. This working solution is prepared by adding 5 ml 1M sodium phosphate buffer and 5 ml 20% formaldehyde to 40 ml water, yielding 2% formaldehyde in 0.1M phosphate buffer. Working solution should be stored at 4°C and used within 2–3 days.

2. 1M sodium phosphate pH 7.4: Dissolve 13.56 g NaH₂PO₄ and 54.94 g Na₂HPO₄ in water to a volume of 400 ml. Adjust pH if needed and set final volume to 500 ml.

3. 20% formaldehyde: Add 20 g powdered paraformaldehyde (see note 1) to a glass beaker and add water to a volume of 90 ml. Place solution on a hot plate in a fume food and heat to 60° with stirring, gradually adding 1N NaOH until paraformaldehyde is dissolved. Do not add more than 1 ml of 1N NaOH or allow solution to heat above 70°. Allow solution to cool to room temperature and set volume to 100 ml with water. Aliquots of 20% formaldehyde should be stored at −20°C and thawed as needed by warming in a 60° water bath (see note 2).

4. Phosphate buffered saline (PBS), pH 7.4: 137 mM NaCl, 2.7 mM KCl, 4.3 mM Na₂HPO₄, 1.47 mM KH₂PO₄.

2.2. Tissue preparation and staining

1. Tris buffered saline (TBS), pH 7.4: 150 mM NaCl, 50 mM Tris-Cl.

2. TBST: TBS containing 0.05% Tween-20.

3. Permeabilization/blocking buffer: 5% donkey serum, 2.5% BSA, 0.5% Triton X100 in TBS. After preparation, this solution should be sterile filtered and stored as aliquots at −20°C. Thawed aliquots can be stored for several weeks at 4°C.

4. Rat anti-CD31/Pecam1 antibody

5. Goat anti-PROX1 antibody

6. Fluorescently labeled donkey anti-rat and anti-goat secondary antibodies.

3. Method

3.1: Tissue preparation

1. Sacrifice mice according to the protocol approved by your local animal care committee (see Note 3). To enucleate the eyes, use fine scissors and make an incision on the lateral side of the socket, cutting through the conjunctiva and surrounding tissue while being careful not to puncture the eye. After making this incision, enucleate the eye by pulling back the tissue surrounding the eye socket and firmly grasping the optic nerve with curved forceps.

2. Fix enucleated eyes by immersion in fixative (described above) and rotating overnight at 4°C.

3. After fixation, place the eye in a 10 cm tissue dish of TBS and, working under a dissecting microscope, use fine scissors (see Note 4) to remove the conjunctiva and any remaining connective tissue from the surface of the eye. Take care not to puncture the eye at this stage.

4. Using scissors, cut the optic nerve where it emerges from the globe.

5. Grasp the eye with forceps by inserting one point into the hole where the optic nerve emerged from the sclera. With the orb thus gripped, use a sharp razor blade (see notes 5 and 6) to make an incision from the optic nerve to the center of the cornea. This incision should be made in one even, gentle stroke, applying little downwards pressure and instead relying on the sharp edge of the blade to make the cut. If the initial cut does not extend the full length of the needed opening, enlarge the incision with scissors (Figure 1 A).

6. Using scissors, remove the lower third of the sclera as shown by the dotted line in Figure 1 A, taking care not to damage the iridocorneal angle / limbal region where SC is located.

7. Using forceps, remove the crystalline lens and retina through the opening created in the previous steps (see Note 7).

Figure 1.

Preparing enucleated eyes for staining and mounting. (A) After removing the conjunctiva and other connective tissue from the surface of the globe, remove the optic nerve by cutting flush with the surface of the eye with scissors. While grasping the eye through the hole left by the optic nerve, a razor blade is used to make an incision from the optic nerve to the center of the cornea. A second cut (dashed line) is then made to remove the lower third of the sclera. (B) After staining, scissors are used to make a series of cuts in the sclera and cornea in preparation for flat mounting—taking care to avoid cutting into the iridocorneal angle/limbal region (orange). (C) After cutting, the eye is opened and laid flat on a glass slide in preparation for mounting.

3.2: Staining

1. Transfer the prepared eyes to 2ml round-bottom eppendorf tubes completely filled with permeabilization/blocking buffer. Rotate on an end-over-end or nutating shaker for 6 hours at room temperature or overnight at 4°C.

2. Prepare primary antibody dilutions in fresh permeabilization/blocking buffer. Mature SC is strongly labeled by antibodies against CD31 (Pecam1) and PROX1 [3,4,7], and we prefer a double-labeling method using both of these antibodies. 100 μl of antibody working solution is sufficient to stain an adult mouse eye in a 2 ml round bottom Eppendorf tube.

3. Aspirate blocking buffer and replace with primary antibody diluted in fresh permeabilization/blocking buffer. Place tubes upright in an Eppendorf tube rack and rotate overnight at 4°C on a bellydancer or nutating shaker.

4. Aspirate antibody solution and wash 6x in TBST. For each wash, completely fill the Eppendorf tube with TBST and rotate at room temperature for 1 hour. Repeat by aspirating and replacing the solution with fresh TBST.

5. Prepare dilutions of appropriate fluorescently labeled secondary antibodies in permeabilization/blocking buffer. Aspirate TBST after the final wash and replace with 100 μl of secondary antibody solution. Place tubes upright in Eppendorf tube rack, wrap in foil to protect from light, and rotate overnight as in step 2.3.

6. Wash as in step 2.4.

7. Mount. Place a clean microscope slide under a dissecting microscope and place a large drop of TBS in the center. Using forceps, transfer the tissue into this drop so that it is submerged and easily visible under the microscope. Use fine scissors to trim the eye as shown in Figure 1 B, making a series of cuts through the sclera towards the limbal region. A similar series of cuts should then be made from the center of the cornea. Still working within the drop of TBS, unroll the eye and flatten it against the slide, with the outer surface facing upwards. Additional cuts in the cornea and sclera can be made as required to ensure that the tissue lies flat as shown in Figure 1 C. When making cuts, take care to avoid cutting through and damaging the limbal region where SC is located, marked in orange in Figure 1. Use tissue paper to slowly aspirate TBS from the surface of the slide, encouraging the tissue to lie flat and stopping as needed to make additional cuts and straighten the tissue with forceps. Once TBS is removed and tissue is fully flattened, gently place a drop of mounting media (see Note 8) onto the slide and cover with an appropriate cover glass.

8. Mounted slides should be dried while protected from light and stored at 4°C until imaging.

3.3: Imaging and analysis

1. Image slides using a confocal microscope equipped with appropriate laser lines and filters for secondary antibody fluorophores selected in step 11 and a 20x objective (see Note 9). When scanning through the iridocorneal angle region in the Z-axis, the CD31-positive superficial vascular plexus [3] and PROX1/CD31-positive limbal lymphatic vasculature [2,5] will be readily apparent (Figure 2). SC resides 15–20 μm below these vessels, and can be identified by its large diameter as well as its PROX1/CD31 double staining (see Note 10). Scanning deeper will reveal the vasculature of the iris (although this may not be visible if using eyes from pigmented mice).

Figure 2.

When imaging the iridocorneal region using a confocal microscope, the superficial vascular (SVP, red) and lymphatic (LP, green) plexuses will be easily visible near the surface of the eye. Schlemm’s canal (SC, blue) resides 10–15 μm below them.

3.4: Measurement of SC area

1. With the confocal pinhole set to 1 Airy unit and a step size of 1.67 μm, capture 3–5 10-image SC Z-stacks at intervals around the circumference of the iridocorneal angle.

2. Using ImageJ Fiji software [10], generate maximum intensity Z-projections of each image and measure the SC area obtained in each field. Average these measurements to obtain a single value for each eye.

3.5. Quantification of focal defects and convolutions

1. To quantify the presence of focal defects and convolutions which do not have a significant impact on total SC area, large area scans encompassing the entire SC may be used. Using a 10 or 20× objective with a fully opened pinhole, focus on the SC and use the large area scan function (see Note 11) of a confocal microscope equipped with a computer controlled stage to capture an image including the entire circumference of the iridocorneal angle and the canal.

2. Use large area scan images to quantify focal defects (i.e. narrowings or gaps) in canal morphology. These can be easily counted manually using Fiji software.