Preparation and Analysis of Histological Slides of Rat and Mouse Eyeballs to Evaluate the Retina

Summary

This manuscript presents a protocol for the histological preparation of slides from rodent eyeballs. With the proposed method, the inner retina can be easily visualized and assessed under a light microscope. The procedure is presented for the eyeballs of mice and rats.

Abstract

A rodent eyeball is a powerful tool for researching the pathomechanisms of many ophthalmic diseases, such as glaucoma, hypertensive retinopathy, and many more. Preclinical experiments enable researchers to examine the efficacy of novel drugs, develop new methods of treatment, or seek new pathomechanisms involved in the disease's onset or progression. A histological examination provides a lot of information necessary to assess the effects of the conducted experiments and can reveal degeneration, tissue remodeling, infiltration, and many other pathologies. In clinical research, there is rarely any chance of obtaining eye tissue suitable for a histological examination, which is why researchers should take advantage of the opportunity offered by the examination of eyeballs from rodents. This manuscript presents a protocol for the histological preparation of rodent eyeballs' sections. The procedure is presented for the eyeballs of mice and rats and has the following steps: (i) harvesting the eyeball, (ii) preserving the eyeball for further analysis, (iii) processing the tissue in paraffin, (iv) preparing slides, (v) staining with hematoxylin and eosin, (vi) assessing the tissue under a light microscope. With the proposed method, the retina can be easily visualized and assessed in detail.

Introduction

The eyeball is a globe-like structure constituting the organ of sight. The inside of the eyeball can be divided into two segments: the anterior segment, which includes the cornea, iris, ciliary body, lens, anterior chamber, and posterior chamber, and the posterior segment, which includes the vitreous body, retina, choroid, sclera, optic nerve head. The anterior chamber is located between the cornea and the iris^1,2. The iris is a circular structure with an opening in the center called the pupil. At the junction of the cornea and the iris, there is the drainage angle, where a specialized system of trabeculae drains the aqueous humor from the eyeball to the venous system. The posterior chamber is bound by the iris, the ciliary body, and the lens, as well as the suspensory ligaments that hold the lens in place. Behind the lens, the vitreous body is located, which is the largest structure of the eyeball. The vitreous body adheres to the retina and stabilizes its position. The retina is surrounded by the choroid and the sclera¹. The anatomy of the eyeball is summarized in Figure 1.

The eyeball wall constitutes three layers. The outermost layer is the sclera, which protects the eyeball from injury and maintains its shape. At the anterior pole of the eyeball, the sclera turns into a transparent cornea. Under the sclera, there is a vascular structure called the uvea, whose primary function is to nourish the structures of the eye. The uvea forms the choroid, and within the anterior pole, the ciliary body and the iris¹. The innermost layer is the retina, which is a highly specialized structure of the eye, made up of neurons, glial cells, and pigment epithelial cells. The retinal neurons include the photoreceptor cells (rods and cones), horizontal cells, bipolar cells, amacrine cells, and retinal ganglion cells (RGCs). These cells are organized into complex layers, and their role is to receive and process light stimuli^2,3. The glial cells of the retina comprise the Müller cells, astrocytes, and microglia, which are present in all layers of the retina. The many functions of glial cells include the nutrition of neurons, support of the blood-retina barrier, structural support of neurons to maintain the layered structure of the retina, regulation of neurons' metabolism, secretion of biologically active proteins regulating functioning, growth, and survival of neurons, and regulating the immunological processes within the retina⁴. The pigment epithelial cells make up the outermost layer of the retina and act as a barrier between the choroid and photoreceptors. These cells regulate the bidirectional transport of waste products and nutrients and also protect the retina from excessive light-induced damage by absorbing light energy and neutralizing light-generated reactive oxygen species⁵.

The retina can be divided into ten layers: (i) the retinal pigment epithelium, (ii) the photoreceptor segment layer (rods and cones), (iii) external limiting membrane (ELM), (iv) outer nuclear layer (ONL), (v) outer plexiform layer (OPL), (vi) inner nuclear layer (INL), (vii) inner plexiform layer (IPL), (viii) ganglion cell layer (GCL), (ix) retinal nerve fiber layer (RNFL), and (x) internal limiting membrane (ILM)². The nucleated layers of the retina include the ONL, INL, and GCL, while the layers with synaptic connections between cells include the OPL and IPL⁶. The nuclei of rods and cones form the ONL, and their axons extend into the OPL, where they connect to the dendrites of bipolar and horizontal cells. Within the INL, the horizontal, bipolar, and amacrine cell nuclei are located. Within the IPL, axon terminals of bipolar and amacrine cells synapse with dendrites of RGCs. Within the GCL, the RGCs make up most of the cell bodies, but also some dislocated amacrine cells can be found there. The axons of RGCs form the RNFL and, further - the optic nerve^7,8,9.

Many ophthalmic diseases, such as neurodegenerative disorders of the retina, lead to irreversible and incurable blindness¹⁰. Vision is considered one of the most important senses¹¹, and permanent blindness significantly reduces the patient's quality of life. To further the understanding of many diseases' pathomechanisms, preclinical research using cell cultures or animal models is broadly performed. Preclinical studies using experimental animals provide an opportunity to assess the pathomechanisms underlying eye diseases and to develop new therapeutic strategies. Eye diseases can be modeled in both large animals (monkeys, cows, dogs, and cats) and small animals (rabbits, rats, mice, and zebrafish). The use of larger animals allows better access to the eye due to its size. Experiments performed with rodents, on the other hand, due to their relatively rapid reproduction, allow the breeding of inbred strains characterized by susceptibility to certain diseases^12,13.

In animal models, enucleation is possible, which enables performing a detailed histopathological examination of the eyeball, with the assessment of minor pathologies that appear in particular structures of the eye during the development of the disease - an examination that can be very rarely performed among humans. Histopathological assessment is an extremely valuable tool while researching the pathomechanisms of ocular disorders. In the published literature, the descriptions of the methodology for histological examination of eyeballs are very limited, and step-by-step guides are lacking, which makes it difficult for beginners to recreate. This study aims to provide a simple and concise method of preparing histological slides and assessing the rat and mouse retina.

Protocol

The research was performed in compliance with institutional guidelines. Eyeballs used in this study were obtained from animals included in an experiment conducted based on the study approval number WAW2/122/2020 issued by the 2^nd Local Ethics Committee at the Warsaw University of Life Sciences in Warsaw, Poland. Our protocol was developed by examining mice aged 11 and 44 weeks and rats aged 6, 12, and 16 weeks.

1. Preparation of mouse eyeballs

NOTE: This protocol presents the method of preparing sections of mouse eyeballs and was developed using eyeballs harvested from: female C57Bl/6 mice aged 11 weeks (average weight 21 g), female C57Bl/6 mice aged 44 weeks (average weight 25 g), female DBA/2 mice aged 11 weeks (average weight 21.5 g), and female DBA/2 mice aged 44 weeks with glaucoma (average weight 25 g).

1. Enucleation
   1. Use small forceps to open the eyelids. Press the eyelids for the eyeball to prolapse.
   2. Dissect the eyeball from the socket by cutting off the tissue behind the eyeball with small, sharp scissors. Once the eyeball is removed from the orbit, cut off the protruding connective tissue and periocular muscles.
      1. Mark the eyeballs for better orientation, for example, by tying a small suture on the stump of the tendon of the lateral rectus muscle of the eyeball to mark the temporal side of the eyeball.
2. Fixation of tissues
   1. Place the enucleated eyeball in 1.5 mL microcentrifuge tubes and let it rest in 4 % solution of paraformaldehyde (PFA) dissolved in phosphate buffer saline (PBS) for 24 h.
   2. Following this, treat the eyeball for 24 h in a 10% solution of sucrose, then remove and place it for 24 h in a 20% solution of sucrose, and finally, remove and place for 24 h in a 30% solution of sucrose.
      NOTE: Throughout the process, keep the eyeball at a low temperature of 4 °C. Needle punctures for sucrose infiltration should be avoided to minimize the risk of eyeball damage.
3. Tissue freezing and storing
   1. Remove the eyeball from the sucrose solution and dry it by lightly dabbing it with a paper towel.
   2. Place the eyeball in a clean 1.5 mL microcentrifuge tube and freeze at - 80 °C for future processing.
      NOTE: If stored at - 80 °C, an eyeball can remain viable for up to 12 months until future processing.
4. Slide preparation
   1. Thaw the eyeball at room temperature. Move it from the microcentrifuge tube into a cassette and rinse under running water for 20 min to wash out the sucrose.
   2. Dehydrate the eyeball by placing it into, and then removing it from the following solutions under a fume hood: 70% solution of ethyl alcohol (EtOH) for 10 min, 80 % solution of EtOH for 10 min, 96 % solution of EtOH for 10 min, 99.9 % solution of EtOH for 10 min, and acetone for 5 min.
   3. Clear the eyeball by placing it in xylene for 5 min under a fume hood.
   4. Melt the paraffin by placing some solid paraffin into a beaker and putting it into a laboratory incubator set to a temperature higher than the melting temperature specified by the paraffin manufacturer (the melting temperature of paraffin is between 46 °C and 68 °C). After the paraffin is fully melted (the time depends on the amount of used paraffin), move the eyeball into the beaker with paraffin and keep warm in the incubator for 1 h. During 1 h of paraffin infiltration into the tissue, stir the paraffin every 10 - 15 min.
      NOTE: A tissue processor that enables paraffin infiltration while stirring the solution may be used for this step.
   5. Embed the eyeball into a paraffin block as described below.
      1. Open the cassette and remove the top cover. Choose a metal mold and pour a small amount of paraffin to cover the bottom of the mold. Remove the eyeball from the cassette with forceps and place it onto the paraffin in the mold. Place the eyeball on its side to ensure cutting in the sagittal plane.
         NOTE: Placing the eyeball on its side (with the anterior pole to one side and the posterior pole to the other) will enable cutting in the sagittal plane. This way, sections through the cornea, iris, ciliary body, pupil, peripheral, and central retina will be obtained.
      2. Briefly cool the mold on a cooling plate set to - 15 °C for the paraffin to slightly solidify to anchor the tissue in place. Carefully cover the tissue with paraffin and the cassette bottom to form a block. Put the block on a cooling plate to fully solidify. Then, remove the block from the metal mold and place the paraffin blocks back onto the cooling plate for at least 15 min to cool before cutting.
   6. Secure a paraffin block in the microtome. Set the microtome to trim the excess paraffin from the block (we set it to cut 15 µm) and trim until the center of the eyeball. Change the microtome settings to cut thinner sections (we set it to cut 3.5 µm) and place the section onto a slide. If using a microtome with an adjacent water bath, remove the excess water from the slide with a moist tissue.
      NOTE: The goal is to obtain at least 3 cross-sections through the pupil. The less experienced the researcher, the more sections should be obtained to ensure sectioning at the level of the pupil. Place a couple of sections onto one slide. Also, for less experienced researchers, cut the tissue thicker, for example, into sections 4 µm or 5 µm thick. The use of a microtome with an adjacent water bath makes the process of transferring tissue sections onto slides much easier when compared to using microtomes without water baths. Another point worth noting is that the lens may be difficult to cut through. To minimize the risk of tissue damage by the brittle lens, only use sharp microtome blades designated to cut through hard tissues. Consider using one blade for trimming and another for cutting sections to minimize the blunting of the blade.
   7. Place the slide into a laboratory incubator set to 56 °C for 30 min.
   8. Remove the slide from the incubator and begin staining it with hematoxylin and eosin (H&E) manually by sequentially putting and removing the slide to and from the following solutions. Start with 7 min xylene, 5 min xylene (another beaker with a clean solution), and 5 min xylene (another beaker with a clean solution). Follow this with 2 min 96% EtOH, 1 min 96% EtOH (another beaker with a clean solution), 1 min 96% EtOH (another beaker with a clean solution).
   9. Wash in 5 min running water, then 2 min Mayer's hematoxylin, 2 min Mayer's hematoxylin (another beaker with a clean solution), 5 min running water, 5 min running water (another, clean beaker).
   10. Add 1% Eosin for 2 min, followed by 1 min 96% EtOH, 30 s 96% EtOH (another beaker with a clean solution), 30 s 96% EtOH (another beaker with a clean solution), 2 min xylene, 1 min xylene (another beaker with a clean solution), 2 min xylene (another beaker with a clean solution).
       NOTE: Alternatively, use an automated slide stainer.
   11. Seal the slide using an automated slide sealer. The prepared slide is ready for storage and assessment under a light microscope.
       NOTE: Alternatively, perform slide sealing manually by placing a thin layer of polystyrene mountant on the stained tissue and covering it with a glass coverslip.
       CAUTION: Steps that include PFA, EtOH, acetone, and xylene should be performed under a laboratory fume hood.

2. Preparation of rat eyeballs section and slide mounting

NOTE: This protocol presents the method of preparing sections of rat eyeballs and was developed using the eyeballs harvested from: male SHR rats aged 6 weeks (average weight 115.5 g), male SHR rats aged 12 weeks with arterial hypertension (average weight 262.5 g), male WKY rats aged 6 weeks (average weight 143.5 g), male WKY rats aged 12 weeks (average weight 265 g), male Lewis rats aged 12 weeks (average weight 310 g), and male Lewis rats aged 16 weeks with induced diabetes mellitus (average weight 259 g).

1. Enucleation
   1. Use small forceps to open the eyelids. Press on the eyelids to prolapse the eyeball.
   2. Dissect the eyeball from the socket by cutting off the tissue behind the eyeball with small, sharp scissors.
      NOTE: Mark the eyeballs for better orientation, for example, by tying a small suture on the stump of the tendon of the lateral rectus muscle of the eyeball to mark the temporal side of the eyeball.
2. Fixation of tissues
   1. Place the enucleated eyeball in a 1.5 mL microcentrifuge tube and let it rest in a 4% solution of paraformaldehyde (PFA) dissolved in phosphate buffer saline (PBS) for 24 h.
   2. Following this, treat the eyeball for 24 h in a 10% solution of sucrose, then remove and place it for 24 h in a 20% solution of sucrose, and finally remove and place it for 24 h in a 30% solution of sucrose.
      NOTE: Throughout the process, keep the eyeball at a low temperature of 4 °C. Needle punctures for sucrose infiltration should be avoided to minimize the risk of eyeball damage.
3. Tissue freezing and storing
   1. Remove the eyeball from the sucrose solution and dry it by lightly dabbing it with a paper towel.
   2. Place the eyeball in a clean 1.5 mL microcentrifuge tube and freeze it at -80 °C for future processing.
      NOTE: If stored at -80 °C, the eyeball can remain viable for up to 12 months until future processing.
4. Slide preparation
   1. Remove the eyeball from the freezer and from the microcentrifuge tube.Cut the eyeball into half along the sagittal plane with a sharp scalpel.Remove and discard the lens.
      NOTE: This step is easiest to perform on a frozen eyeball, freshly taken from the freezer. During eyeball cutting, the lens may damage the surrounding tissue. To minimize the risk of damage, use a very sharp scalpel and remain cautious.
   2. Thaw the eyeball at room temperature. Move the two halves into a cassette and rinse under running water for 20 min to wash out the sucrose.
   3. Dehydrate the eyeball by placing it into and then removing it from the following solutions under a fume hood: 70% solution of ethyl alcohol (EtOH) for 15 min, 80% solution of EtOH for 15 min, 96% solution of EtOH for 15 min, 99.9% solution of EtOH for 15 min, and acetone for 7 min.
   4. Clear the eyeball by placing it in xylene for 7 min under a fume hood.
   5. Melt the paraffin by placing some solid paraffin into a beaker and putting it into a laboratory incubator set to a temperature higher than the melting temperature specified by the paraffin manufacturer (the melting temperature of paraffin is between 46 °C and 68 °C). After the paraffin is fully melted (the time depends on the amount of paraffin used), move the eyeball into the beaker with paraffin and keep warm in the incubator for 1 h. During 1 h of paraffin infiltration into the tissue, stir the paraffin every 10 - 15 min.
      NOTE: A tissue processor that enables paraffin infiltration while stirring the solution may be used for this step.
   6. Embed the eyeball into a paraffin block as described below.
      1. Open the cassette and remove the top cover. Choose a metal mold and pour a small amount of paraffin to cover the bottom of the mold. Remove the eyeball from the cassette with forceps and place it onto the paraffin in the mold. Place one eyeball half with the bottom of the cup up, and the other - down.
         NOTE: Placing the eyeball halves with one half with the bottom of the cup up, and the other - down will enable tissue cutting in the sagittal plane. This way, sections through the cornea, iris, ciliary body, pupil, peripheral, and central retina will be obtained.
      2. Briefly cool the mold on a cooling plate set to - 15 °C for the paraffin to slightly solidify and anchor the eyeball in place. Carefully cover the eyeball with paraffin and the cassette bottom to form a block. Put the block on a cooling plate to fully solidify. Then, remove the block from the metal mold and place the paraffin blocks back onto the cooling plate for at least 15 min to cool before cutting.
   7. Secure a paraffin block in the microtome. Set the microtome to trim the excess paraffin from the block (we set it to cut 15 µm) and trim until you reach the center of the eyeball. Change the microtome settings to cut thinner sections (we set it to cut 3.5 µm) and place the section onto a slide. If you are using a microtome with an adjacent water bath, remove the excess water from the slide with a moist tissue.
      NOTE: The goal is to obtain at least 3 cross-sections through the pupil. The less experienced the researcher, the more sections should be obtained to ensure sectioning at the level of the pupil. Place a couple of sections onto one slide. Also, for less experienced researchers, cut the tissue thicker, for example, into sections 4 µm or 5 µm thick. The use of a microtome with an adjacent water bath makes the process of transferring tissue sections onto slides much easier when compared to using microtomes without water baths.
   8. Remove the slide and begin staining with hematoxylin and eosin (H&E) manually by sequentially putting and removing the slide to and from the following solutions. Start with 7 min xylene, 5 min xylene (another beaker with a clean solution), and 5 min xylene (another beaker with a clean solution). Follow this with 2 min 96% EtOH, 1 min 96% EtOH (another beaker with a clean solution), 1 min 96% EtOH (another beaker with a clean solution).
   9. Wash in 5 min running water, then 2 min Mayer's hematoxylin, 2 min Mayer's hematoxylin (another beaker with a clean solution), 5 min running water, 5 min running water (another, clean beaker).
   10. Add 1% Eosin for 2 min, followed by 1 min 96% EtOH, 30 s 96% EtOH (another beaker with a clean solution), 30 s 96% EtOH (another beaker with a clean solution), 2 min xylene, 1 min xylene (another beaker with a clean solution), 2 min xylene (another beaker with a clean solution).
       NOTE: Alternatively, use an automated slide stainer.
   11. Seal the slide using an automated slide sealer. The prepared slides are ready for storage and assessment under a light microscope.
       NOTE: Alternatively, perform slide sealing manually, by placing a thin layer of polystyrene mountant on the stained tissue and cover by a glass coverslip.
       CAUTION: Steps that include PFA, EtOH, acetone, and xylene should be performed under a laboratory fume hood.

3. Assessing the eyeball sections

1. Slide scanning
   1. Put the slide into a histopathological slide scanner and scan the sections using the scanning mode: 40x. Save the images as high-resolution digital files.
2. Slide analysis
   1. Open the scanned file with the use of image viewing software. Find cross-pupil cross-sections and analyze three consecutive sections.
   2. Find the thickest part of the retina and magnify the image to 20x by pressing on the magnification box in the top right-hand corner and choosing 20x. Move the mouse to the level of ELM in the thickest part of the retina and press the right-hand button on the mouse, choose Annotate, and Ruler.
   3. Move the mouse towards the ILM and click the left-hand button. Title the measurement RT and check the boxes show title and show length. This way, the retinal thickness (RT) will be measured, and the measurement will be shown in µm. Representative results are shown in Figure 2 and Figure 3.
   4. Use the ruler, measure the thickness of each retinal layer, including the ONL, OPL, INL, and IPL, and label them. Representative results are shown in Figure 4 and Figure 5.
   5. To count the number of cells within the GCL, establish a predetermined length of the retina. For example, for the mouse retina, use one 500 µm section, and for the rat retina, use five 100 µm sections.
   6. Using the ruler, measure the chosen distance along the GCL.Press the right-hand button on the mouse and select Annotate > Saved > 1x RBC. Click on each cell identified as a round, hematoxylin-stained body along the GCL over the predetermined length of the retina. Count the number of encircled cells. Representative results are shown in Figure 6 and Figure 7.
   7. After analyzing three consecutive sections, draw the average for all the measurements.

Results

With the use of the presented protocol, the anatomy and morphology of specific structures of the eyeball can be assessed in detail. Our team focuses on researching the pathomechanisms of neurodegeneration in retinal disorders, such as glaucoma, diabetic retinopathy, and hypertensive retinopathy. To assess the features of neurodegeneration in the retina, we evaluated the RT (mouse eyeball -Figure 2, rat eyeball -Figure 3), ONL, OPL, INL, and IPL (mouse eyeball -<...

Discussion

A histological examination of the mouse and rat eyeball is a powerful tool in the field of experimental ophthalmology. It can provide new information related to changes in the ocular structures in the course of ophthalmic diseases, which would otherwise not be observed in clinical settings. The described protocols present a simple method of conducting a histological analysis of the retina. The subtleties of the procedure needed to process anterior tissues, such as the cornea, are not described here.

Materials

Name	Company	Catalog Number	Comments
Acetone	Chempur	111024800
Dumont Tweezers #5, 11cm, 0.05 x 0.01mm Tips	World Precision Instruments	14095	small, microsurgical forceps
Eosin	Mar-Four	4P.05.2001/L	Aqueous solution
Ethyl alcohol 96 %	Chempur	CH.ET.AL.96%CZDA.CH
Ethyl alcohol 99.9 %	Chempur	CH.ETYL.ALK.99,9%BW	Use to prepare 70% and 80% solutions of EtOH
Glacier - 86 °C Ultralow Temperature Freezer NU-9483E	NuAire	13027D0034	Freezing temperature - 80 °C
Glass slides	Mar-Four	MI.15.01673	Ground glass slides with a double-sided matte field for description
Leica CV5030 Fully Automated Glass Coverslipper	Leica Biosystems	14,04,78,80,101	Automated slide sealer
Mayer's hematoxylin	Chempur	124687402
Metal base molds 15 mm  x 15 mm	Leica Biosystems	3803081
Microme HM 340 E	Thermo Scientific	90-519-0	Microtome
Microtome razor blades	Mar-Four	HI.N.35	Cutting angle 35°
Micro-Twin biopsy cassette	Mar-Four	LN.13874550
Microwave Hybrid Tissue Processor	Milestone
Multistainer Leica ST5020	Leica Biosystems		Automated slide stainer
NanoZoomer 2.0-HT slide scanner	Hamamatsu	C9600	Histopathological slide scanner
NDP.view2 Image viewing software	Hamamatsu	U12388-01	Image viewing software
Paraffin Pathowax Plus	Mar-Four	4P.PWP.010	Melting temperature 56 °C - 58 °C
Paraformaldehyde	Sigma - Aldrich	441244-1KG	Prepare a 4% solution in PBS
PBS Tablets	Merck Millipore	524650-1EA	Use to prepare a solution of phosphate buffer saline, per manufacturer's instructions
Pierce Microcentrifuge Tubes, 1.5 mL	Thermo Scientific	69715
Refrigerator-freezer EN14000AW	Electrolux	925032562-00	refrigeration 4 °C
Scalpel blade	Swann-Morton	T.OST.SKAL00.024
Sucrose	Chempur	427720906
SuperCut Spring Scissors, 12.5cm, Curved	World Precision Instruments	501925	curved, small microsurgical scissors
Tape for automatic sealers	Mar-Four	KP-3020/SMRH001
Xylene	Chempur	CH.KSYL.5l.METAL.OP