The work, including rats, was funded as part of the project TIME 2 MUW - teaching excellence as an opportunity for the development of the Medical University of Warsaw co-financed by the European Social Fund under the Operational Program Knowledge Education Development for 2014-2020, the number of the co-financing agreement: POWR.03.05.00-00-Z040/18-00. The work, including mice, was carried out as part of a project implemented in the years 2020 to 2022, financed by a subsidy for science obtained by the Medical University of Warsaw.

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 2nd 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&#160;preparing sections of mouse eyeballs and was&#160;developed using eyeballs harvested from:&#160;female C57Bl/6 mice aged 11 weeks (average&#160;weight 21 g), female C57Bl/6 mice aged 44&#160;weeks (average weight 25 g), female DBA/2&#160;mice aged 11 weeks (average weight 21.5 g),&#160;and female DBA/2 mice aged 44 weeks with&#160;glaucoma (average weight 25 g).
<ol>
	<li>Enucleation
	<ol>
		<li>Use small forceps to open the eyelids. Press the eyelids for the eyeball to prolapse.</li>
		<li>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.
		<ol>
			<li>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.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Fixation of tissues
	<ol>
		<li>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.</li>
		<li>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 &#176;C. Needle punctures for sucrose infiltration should be avoided to minimize the risk of eyeball damage.</li>
	</ol>
	</li>
	<li>Tissue freezing and storing
	<ol>
		<li>Remove the eyeball from the sucrose solution and dry it by lightly dabbing it with a paper towel.</li>
		<li>Place the eyeball in a clean 1.5 mL microcentrifuge tube and freeze at - 80 &#176;C for future processing. 
		NOTE: If stored at - 80 &#176;C, an eyeball can remain viable for up to 12 months until future processing.</li>
	</ol>
	</li>
	<li>Slide preparation
	<ol>
		<li>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.</li>
		<li>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.</li>
		<li>Clear the eyeball by placing it in xylene for 5 min under a fume hood.</li>
		<li>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 &#176;C and 68 &#176;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.</li>
		<li>Embed the eyeball into a paraffin block as described below.
		<ol>
			<li>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.</li>
			<li>Briefly cool the mold on a cooling plate set to - 15 &#176;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.</li>
		</ol>
		</li>
		<li>Secure a paraffin block in the microtome. Set the microtome to trim the excess paraffin from the block (we set it to cut 15 &#181;m) and trim until the center of the eyeball. Change the microtome settings to cut thinner sections (we set it to cut 3.5 &#181;m) 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 &#181;m or 5 &#181;m 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.</li>
		<li>Place the slide into a laboratory incubator set to 56 &#176;C for 30 min.</li>
		<li>Remove the slide from the incubator and begin staining it with hematoxylin and eosin (H&#38;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).</li>
		<li>Wash in 5 min running water, then 2 min Mayer&#39;s hematoxylin, 2 min Mayer&#39;s hematoxylin (another beaker with a clean solution), 5 min running water, 5 min running water (another, clean beaker).</li>
		<li>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.</li>
		<li>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.</li>
	</ol>
	</li>
</ol>
2. Preparation of rat eyeballs section and slide mounting
NOTE: This protocol presents the method of&#160;preparing sections of rat eyeballs and was&#160;developed using the eyeballs harvested from:&#160;male SHR rats aged 6 weeks (average weight&#160;115.5 g), male SHR rats aged 12 weeks with&#160;arterial hypertension (average weight 262.5 g),&#160;male WKY rats aged 6 weeks (average weight&#160;143.5 g), male WKY rats aged 12 weeks&#160;(average weight 265 g), male Lewis rats aged 12&#160;weeks (average weight 310 g), and male Lewis&#160;rats aged 16 weeks with induced diabetes&#160;mellitus (average weight 259 g).
<ol>
	<li>Enucleation
	<ol>
		<li>Use small forceps to open the eyelids. Press on the eyelids to prolapse the eyeball.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Fixation of tissues
	<ol>
		<li>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.</li>
		<li>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 &#176;C. Needle punctures for sucrose infiltration should be avoided to minimize the risk of eyeball damage.</li>
	</ol>
	</li>
	<li>Tissue freezing and storing
	<ol>
		<li>Remove the eyeball from the sucrose solution and dry it by lightly dabbing it with a paper towel.</li>
		<li>Place the eyeball in a clean 1.5 mL microcentrifuge tube and freeze it at -80 &#176;C for future processing. 
		NOTE: If stored at -80 &#176;C, the eyeball can remain viable for up to 12 months until future processing.</li>
	</ol>
	</li>
	<li>Slide preparation
	<ol>
		<li>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.</li>
		<li>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.</li>
		<li>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.</li>
		<li>Clear the eyeball by placing it in xylene for 7 min under a fume hood.</li>
		<li>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 &#176;C and 68 &#176;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.</li>
		<li>Embed the eyeball into a paraffin block as described below.
		<ol>
			<li>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.</li>
			<li>Briefly cool the mold on a cooling plate set to - 15 &#176;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.</li>
		</ol>
		</li>
		<li>Secure a paraffin block in the microtome. Set the microtome to trim the excess paraffin from the block (we set it to cut 15 &#181;m) 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 &#181;m) 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 &#181;m or 5 &#181;m 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.</li>
		<li>Remove the slide and begin staining with hematoxylin and eosin (H&#38;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).</li>
		<li>Wash in 5 min running water, then 2 min Mayer&#39;s hematoxylin, 2 min Mayer&#39;s hematoxylin (another beaker with a clean solution), 5 min running water, 5 min running water (another, clean beaker).</li>
		<li>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.</li>
		<li>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.</li>
	</ol>
	</li>
</ol>
3. Assessing the eyeball sections
<ol>
	<li>Slide scanning
	<ol>
		<li>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.</li>
	</ol>
	</li>
	<li>Slide analysis
	<ol>
		<li>Open the scanned file with the use of image viewing software. Find cross-pupil cross-sections and analyze three consecutive sections.</li>
		<li>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.</li>
		<li>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 &#181;m. Representative results are shown in Figure 2 and Figure 3.</li>
		<li>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.</li>
		<li>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 &#181;m section, and for the rat retina, use five 100 &#181;m sections.</li>
		<li>Using the ruler, measure the chosen distance along the GCL.Press the right-hand button on the mouse and select Annotate &#62; Saved &#62; 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.</li>
		<li>After analyzing three consecutive sections, draw the average for all the measurements.</li>
	</ol>
	</li>
</ol>

Assessing Retinal Thickness and Ganglion Cell Layer Counts in Rodent Eyeballs

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<li>Polley, E. H., Walsh, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+technique+for+flat+embedding+and+en+face+sectioning+of+the+mammalian+retina+for+autoradiography%5BTitle%5D)+AND+%22J+Neurosci+Method%22%5BJournal%5D)">A technique for flat embedding and en face sectioning of the mammalian retina for autoradiography.</a> J Neurosci Method. 12 (1), 57-64 (1984).</li>
<li>Turner, K. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Hematoxylin+toluidine+blue-phloxinate+staining+of+glycol+methacrylate+sections+of+retina+and+other+tissues%5BTitle%5D)+AND+%22Stain+Technol%22%5BJournal%5D)">Hematoxylin toluidine blue-phloxinate staining of glycol methacrylate sections of retina and other tissues.</a> Stain Technol. 55 (4), 229-233 (1980).</li>
<li>Cheng, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Correlation+of+optical+coherence+tomography+and+retinal+histology+in+normal+and+pro23his+retinal+degeneration+pig%5BTitle%5D)+AND+%22Transl+Vis+Sci+Technol%22%5BJournal%5D)">Correlation of optical coherence tomography and retinal histology in normal and pro23his retinal degeneration pig.</a> Transl Vis Sci Technol. 7 (6), 18(2018).</li>
</ol>

The authors declare no conflict of interest.

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.
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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 iris1,2. 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 stimuli2,3. The glial cells of the retina comprise the M&#252;ller 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&#39; 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 nerve7,8,9.
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&#39;s quality of life. To further the understanding of many diseases&#39; 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 diseases12,13.
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.

<table><tbody><tr><td>Acetone&amp;nbsp;</td><td>Chempur&amp;nbsp;</td><td>111024800</td><td></td></tr><tr><td>Dumont Tweezers #5, 11cm, 0.05 x 0.01mm Tips</td><td>World Precision Instruments</td><td>14095</td><td>small, microsurgical forceps</td></tr><tr><td>Eosin</td><td>Mar-Four</td><td>4P.05.2001/L</td><td>Aqueous solution</td></tr><tr><td>Ethyl alcohol 96 %</td><td>Chempur&amp;nbsp;</td><td>CH.ET.AL.96%CZDA.CH</td><td></td></tr><tr><td>Ethyl alcohol 99.9 %</td><td>Chempur&amp;nbsp;</td><td>CH.ETYL.ALK.99,9%BW</td><td>Use to prepare 70% and 80% solutions of EtOH</td></tr><tr><td>Glacier - 86 &amp;deg;C Ultralow Temperature Freezer NU-9483E</td><td>NuAire</td><td>13027D0034</td><td>Freezing temperature - 80 &amp;deg;C</td></tr><tr><td>Glass slides</td><td>Mar-Four</td><td>MI.15.01673</td><td>Ground glass slides with a double-sided matte field for description</td></tr><tr><td>Leica CV5030 Fully Automated Glass Coverslipper</td><td>Leica Biosystems&amp;nbsp;</td><td>14,04,78,80,101</td><td>Automated slide sealer&amp;nbsp;</td></tr><tr><td>Mayer&#039;s hematoxylin</td><td>Chempur&amp;nbsp;</td><td>124687402</td><td></td></tr><tr><td>Metal base molds 15 mm&amp;nbsp; x 15 mm</td><td>Leica Biosystems&amp;nbsp;</td><td>3803081</td><td></td></tr><tr><td>Microme HM 340 E</td><td>Thermo Scientific</td><td>90-519-0</td><td>Microtome</td></tr><tr><td>Microtome razor blades</td><td>Mar-Four</td><td>HI.N.35</td><td>Cutting angle 35&amp;deg;</td></tr><tr><td>Micro-Twin biopsy cassette</td><td>Mar-Four</td><td>LN.13874550</td><td></td></tr><tr><td>Microwave Hybrid Tissue Processor</td><td>Milestone</td><td></td><td></td></tr><tr><td>Multistainer Leica ST5020</td><td>Leica Biosystems&amp;nbsp;</td><td></td><td>Automated slide stainer&amp;nbsp;</td></tr><tr><td>NanoZoomer 2.0-HT slide scanner</td><td>Hamamatsu</td><td>C9600</td><td>Histopathological slide scanner</td></tr><tr><td>NDP.view2 Image viewing software</td><td>Hamamatsu</td><td>U12388-01</td><td>Image viewing software</td></tr><tr><td>Paraffin Pathowax Plus</td><td>Mar-Four</td><td>4P.PWP.010</td><td>Melting temperature 56 &amp;deg;C - 58 &amp;deg;C</td></tr><tr><td>Paraformaldehyde</td><td>Sigma - Aldrich</td><td>441244-1KG</td><td>Prepare a 4% solution in PBS</td></tr><tr><td>PBS Tablets</td><td>Merck Millipore</td><td>524650-1EA</td><td>Use to prepare a solution of phosphate buffer saline, per manufacturer&#039;s instructions</td></tr><tr><td>Pierce Microcentrifuge Tubes, 1.5 mL</td><td>Thermo Scientific</td><td>69715</td><td></td></tr><tr><td>Refrigerator-freezer EN14000AW</td><td>Electrolux</td><td>925032562-00</td><td>refrigeration 4 &amp;deg;C</td></tr><tr><td>Scalpel blade</td><td>Swann-Morton</td><td>T.OST.SKAL00.024</td><td></td></tr><tr><td>Sucrose</td><td>Chempur&amp;nbsp;</td><td>427720906</td><td></td></tr><tr><td>SuperCut Spring Scissors, 12.5cm, Curved</td><td>World Precision Instruments</td><td>501925</td><td>curved, small microsurgical scissors</td></tr><tr><td>Tape for automatic sealers</td><td>Mar-Four</td><td>KP-3020/SMRH001</td><td></td></tr><tr><td>Xylene</td><td>Chempur&amp;nbsp;</td><td>CH.KSYL.5l.METAL.OP&amp;nbsp;</td><td></td></tr></tbody></table>

preparation and analysis of histological slides of rat and mouse eyeballs to evaluate the retina

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.

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 -<...

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

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.

A rodent eyeball is a powerful tool for researching the pathomechanisms of many ophthalmic diseases, such as glaucoma, hypertensive retinopathy, and many ...

preparation-analysis-histological-slides-rat-mouse-eyeballs-to

Research

JoVE Journal

Medicine

824 Views.  Medical University of Warsaw. 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.

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

A Protocol to Evaluate and Quantify Retinal Pigmented Epithelium Pathologies in Mouse Models of Age-Related Macular Degeneration

Age-related macular degeneration (AMD) is a debilitating retinal disorder in aging populations. It is widely believed that dysfunction of the retinal pigmented epithelium (RPE) is a key pathobiological event in AMD. To understand the mechanisms that lead to RPE dysfunction, mouse models can be utilized by researchers. It has been established by previous studies that mice can develop RPE pathologies, some of which are observed in the eyes of individuals diagnosed with AMD. Here, we describe a phenotyping protocol to assess RPE pathologies in mice. This protocol includes the preparation and evaluation of retinal cross-sections using light microscopy and transmission electron microscopy, as well as that of RPE flat mounts by confocal microscopy. We detail the common types of murine RPE pathologies observed by these techniques and ways to quantify them through unbiased methods for statistical testing. As proof of concept, we use this RPE phenotyping protocol to quantify the RPE pathologies observed in mice overexpressing transmembrane protein 135 (Tmem135) and aged wild-type C57BL/6J mice. The main goal of this protocol is to present standard RPE phenotyping methods with unbiased quantitative assessments for scientists using mouse models of AMD.

Mouse models can be useful tools for investigating the biology of the retinal pigmented epithelium (RPE). It has been established that mice can develop an array of RPE pathologies. Here, we describe a phenotyping protocol to elucidate and quantify RPE pathologies in mice using light, transmission electron, and confocal microscopy.

Age-related macular degeneration (AMD) is a debilitating retinal disorder in aging populations. It is widely believed that dysfunction of the retinal ...

Genetics

Assessment of Vascular Regeneration in the CNS Using the Mouse Retina

The rodent retina is perhaps the most accessible mammalian system in which to investigate neurovascular interplay within the central nervous system (CNS). It is increasingly being recognized that several neurodegenerative diseases such as Alzheimer&#8217;s, multiple sclerosis, and amyotrophic lateral sclerosis present elements of vascular compromise. In addition, the most prominent causes of blindness in pediatric and working age populations (retinopathy of prematurity and diabetic retinopathy, respectively) are characterized by vascular degeneration and failure of physiological vascular regrowth. The aim of this technical paper is to provide a detailed protocol to study CNS vascular regeneration in the retina. The method can be employed to elucidate molecular mechanisms that lead to failure of vascular growth after ischemic injury. In addition, potential therapeutic modalities to accelerate and restore healthy vascular plexuses can be explored. Findings obtained using the described approach may provide therapeutic avenues for ischemic retinopathies such as that of diabetes or prematurity and possibly benefit other vascular disorders of the CNS.

The rodent retina has long been recognized as an accessible window to the brain. In this technical paper we provide a protocol that employs the mouse model of oxygen-induced retinopathy to study the mechanisms that lead to failure of vascular regeneration within the central nervous system after ischemic injury. The described system can also be harnessed to explore strategies to promote regrowth of functional blood vessels within the retina and CNS.

The rodent retina is perhaps the most accessible mammalian system in which to investigate neurovascular interplay within the central nervous system (CNS). ...

Neuroscience

Glaucoma-inducing Procedure in an In Vivo Rat Model and Whole-mount Retina Preparation

Glaucoma is a disease of the central nervous system affecting retinal ganglion cells (RGCs). RGC axons making up the optic nerve carry visual input to the brain for visual perception. Damage to RGCs and their axons leads to vision loss and/or blindness. Although the specific cause of glaucoma is unknown, the primary risk factor for the disease is an elevated intraocular pressure. Glaucoma-inducing procedures in animal models are a valuable tool to researchers studying the mechanism of RGC death. Such information can lead to the development of effective neuroprotective treatments that could aid in the prevention of vision loss. The protocol in this paper describes a method of inducing glaucoma - like conditions in an in vivo rat model where 50 &#181;l of 2 M hypertonic saline is injected into the episcleral venous plexus. Blanching of the vessels indicates successful injection. This procedure causes loss of RGCs to simulate glaucoma. One month following injection, animals are sacrificed and eyes are removed. Next, the cornea, lens, and vitreous are removed to make an eyecup. The retina is then peeled from the back of the eye and pinned onto sylgard dishes using cactus needles. At this point, neurons in the retina can be stained for analysis. Results from this lab show that approximately 25% of RGCs are lost within one month of the procedure when compared to internal controls. This procedure allows for quantitative analysis of retinal ganglion cell death in an in vivo rat glaucoma model.

Glaucoma-inducing Procedure in an In Vivo Rat Model and Whole-mount Retina Preparation

Glaucoma is characterized by damage to retinal ganglion cells. Inducing glaucoma in animal models can provide insight into the study of this disease. Here, we outline a procedure that induces loss of RGCs in an in vivo rat model and demonstrates the preparation of whole-mount retinas for analysis.

Glaucoma is a disease of the central nervous system affecting retinal ganglion cells (RGCs). RGC axons making up the optic nerve carry visual input to the ...

Preparation of Mouse Retinal Cryo-sections for Immunohistochemistry

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.

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.

Preparation of high-quality mouse eye sections for immunohistochemistry (IHC) is critical for assessing the retinal structure and function and for ...

Full-Circle Cauterization of Limbal Vascular Plexus for Surgically Induced Glaucoma in Rodents

Glaucoma, the second leading cause of blindness worldwide, is a heterogeneous group of ocular disorders characterized by structural damage to the optic nerve and retinal ganglion cell (RGC) degeneration, resulting in visual dysfunction by interrupting the transmission of visual information from the eye to the brain. Elevated intraocular pressure is the most important risk factor; thus, several models of ocular hypertension have been developed in rodents by either genetic or experimental approaches to investigate the causes and effects of the disease. Among those, some limitations have been reported such as surgical invasiveness, inadequate functional assessment, requirement of extensive training, and highly variable extension of retinal damage. The present work characterizes a simple, low-cost, and efficient method to induce ocular hypertension in rodents, based on low-temperature, full-circle cauterization of the limbal vascular plexus, a major component of aqueous humor drainage. The new model provides a technically easy, noninvasive, and reproducible subacute ocular hypertension, associated with progressive RGC and optic nerve degeneration, and a unique post-operative clinical recovery rate that allows in vivo functional studies by both electrophysiological and behavioral methods.

The goal of this protocol is to characterize a novel model of glaucomatous neurodegeneration based on 360&#176; thermic cauterization of limbal vascular plexus, inducing subacute ocular hypertension.

Glaucoma, the second leading cause of blindness worldwide, is a heterogeneous group of ocular disorders characterized by structural damage to the optic ...

Retinal Pathophysiological Evaluation in a Rat Model

A posterior segment eye disease like diabetic retinopathy alters the physiology of the retina. Diabetic retinopathy is characterized by a retinal detachment, breakdown of the blood-retinal barrier (BRB), and retinal angiogenesis. An in vivo rat model is a valuable experimental tool to examine the changes in the structure and function of the retina. We propose three different experimental techniques in the rat model to identify morphological changes of retinal cells, retinal vasculature, and compromised BRB. Retinal histology is used to study the morphology of various retinal cells. Also, quantitative measurement is performed by retinal cell count and thickness measurement of different retinal layers. A BRB breakdown assay is used to determine the leakage of extraocular proteins from the plasma to vitreous tissue due to the breakdown of BRB. Fluorescence angiography is used to study angiogenesis and leakage of blood vessels by visualizing retinal vasculature using FITC-dextran dye.

Diabetic retinopathy is one of the leading causes of blindness. Histology, blood-retinal barrier breakdown assay, and fluorescence angiography are valuable techniques to understand the pathophysiology of the retina, which could further enhance the efficient drug screening against diabetic retinopathy.

A posterior segment eye disease like diabetic retinopathy alters the physiology of the retina. Diabetic retinopathy is characterized by a retinal ...

Microdissection of the Rodent Eye

The ocular micro-dissection of the rodent eye involves the segmentation of the enucleated eyeball with the attached nictitating membrane, or third eyelid, to obtain the anterior and posterior eyecups. With this technique, the sub-parts of the eye, including the corneal tissue, neural tissue, retinal pigment epithelial (RPE) tissue, and lens, can be obtained for wholemounts, cryo-sectioning, and/or single-cell suspensions of a specific ocular tissue. The presence of the third eyelid presents unique and significant advantages, as it benefits the maintenance of the orientation of the eye, which is important for understanding eye physiology following any localized intervention or in studies involving ocular analysis relating to the eye&#39;s spatial topography.
In this method, we enucleated the eyeball at the socket along with the third eyelid by carefully and slowly cutting through the extraocular muscles and severing the optic nerve. The eyeball was pierced through the corneal limbus using a microblade.&#160;The incision was used as the point of entry, allowing for cutting along the corneal-scleral junction by inserting micro-scissors through the incision point. Small and continuous cuts along the circumference were made until the cups separated. These could be further dissected by gently peeling the translucent layer of the neural retina using Colibri suturing forceps to obtain the neural retina and RPE layers. Further, three/four equidistant cuts were made from the periphery perpendicularly to the optic center until the optic nerve was reached. This opened the hemispherical cups into a floret shape so that they fell flat and could be easily mounted. This technique has been used in our lab for corneal wholemounts and retinal sections. The presence of the third eyelid delineates the nasal-temporal orientation, which allows for the study of various cell therapy interventions post-transplantation and, thus, the targeted physiological validation vital for visualization and accurate representation in such studies.

This paper presents a protocol for ocular micro-dissection in rodents. The process involves the enucleation of the eyeball along with the nictitating membrane (i.e., the third eyelid). This is then followed by the separation of the posterior and anterior eye cups.

The ocular micro-dissection of the rodent eye involves the segmentation of the enucleated eyeball with the attached nictitating membrane, or third eyelid, ...

Synaptic Circuits Analysis and Protein Localization in the Mouse Retina

Investigating Retinal Circuits and Molecular Localization by Pre&#45;Embedding Immunoelectron Microscopy

The retina comprises numerous cells forming diverse neuronal circuits, which constitute the first stage of the visual pathway. Each circuit is characterized by unique features and distinct neurotransmitters, determining its role and functional significance. Given the intricate cell types within its structure, the complexity of neuronal circuits in the retina poses challenges for exploration. To better investigate retinal circuits and cross-talk, such as the link between cone and rod pathways, and precise molecular localization (neurotransmitters or neuropeptides), such as the presence of substance P-like immunoreactivity in the mouse retina, we employed a pre-embedding immunoelectron microscopy (immuno-EM) method to explore synaptic connections and organization. This approach enables us to pinpoint specific intercellular synaptic connections and precise molecular localization and could play a guiding role in exploring its function. This article describes the protocol, reagents used, and detailed steps, including (1) retina fixation preparation, (2) pre-embedding immunostaining, and (3) post-fixation and embedding.

Author Spotlight: Understanding the Ultrastructural Basis of Retinal Synaptic Connectivity and Neurotransmitter Localization in Mice 

This protocol outlines the detailed steps of pre-embedding immunoelectron microscopy, with a focus on exploring synaptic circuits and protein localization in the retina.

The retina comprises numerous cells forming diverse neuronal circuits, which constitute the first stage of the visual pathway. Each circuit is ...

Retinal Cryo-sections, Whole-Mounts, and Hypotonic Isolated Vasculature Preparations for Immunohistochemical Visualization of Microvascular Pericytes

Retinal pericytes play an important role in many diseases of the eye. Immunohistochemical staining techniques of retinal vessels and microvascular pericytes are central to ophthalmological research. It is vital to choose an appropriate method of visualizing the microvascular pericytes. We describe retinal microvascular pericyte immunohistochemical staining in cryo-sections, whole-mounts, and hypotonic isolated vasculature using antibodies for platelet-derived growth factor receptor &#946; (PDGFR&#946;) and nerve/glial antigen 2 (NG2). This allows us to highlight advantages and shortcomings of each of the three tissue preparations for the visualization of the retinal microvascular pericytes. Cryo-sections provide transsectional visualization of all retinal layers but contain only a few occasional transverse cuts of the microvasculature. Whole-mount provides an overview of the entire retinal vasculature, but visualization of the microvasculature can be troublesome. Hypotonic isolation provides a method to visualize the entire retinal vasculature by the removal of neuronal cells, but this makes the tissue very fragile.

We demonstrate three different tissue preparation techniques for immunohistochemical visualization of rat retinal microvascular pericytes, i.e., cryo-sections, whole-mounts, and hypotonic isolation of the vascular network.

Retinal pericytes play an important role in many diseases of the eye. Immunohistochemical staining techniques of retinal vessels and microvascular ...

Biology

Cryosectioning and Immunostaining of Mouse Retina

Tissue sectioning and immunohistochemistry are essential techniques in histological and pathological studies of retinal diseases using animal models. These methods enable detailed examinations of tissue morphologies and the localization of specific proteins within the tissue, which provide valuable insights into disease processes and mechanisms. Mice are the most widely used model for this purpose. However, because mouse eyeballs are small and mouse retinas are extremely delicate tissues, obtaining high-quality retinal sections and immunostaining images from mouse eyeballs is typically challenging. This study describes an improved protocol for cryosectioning mouse retinas and performing immunohistochemistry. An essential point of this protocol involves coating the eyeball with a layer of super glue, which prevents deformation of the eyeballs during the processes of cornea removal, lens extraction, and embedding. This step ensures the integrity of retinal morphologies is well preserved. This protocol highlights critical technical considerations and optimization strategies for consistently producing high-quality retinal sections and achieving excellent immunostaining results.

A protocol for preparing mouse retinal cryosections and performing immunostaining on photoreceptors is described. This article enables researchers to consistently produce mouse retinal frozen sections with well-preserved morphology and high-quality immunostaining results.

Tissue sectioning and immunohistochemistry are essential techniques in histological and pathological studies of retinal diseases using animal models. ...

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

Summary

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A Protocol to Evaluate and Quantify Retinal Pigmented Epithelium Pathologies in Mouse Models of Age-Related Macular Degeneration

Assessment of Vascular Regeneration in the CNS Using the Mouse Retina

Glaucoma-inducing Procedure in an In Vivo Rat Model and Whole-mount Retina Preparation

Preparation of Mouse Retinal Cryo-sections for Immunohistochemistry

Full-Circle Cauterization of Limbal Vascular Plexus for Surgically Induced Glaucoma in Rodents

Retinal Pathophysiological Evaluation in a Rat Model

Microdissection of the Rodent Eye

Investigating Retinal Circuits and Molecular Localization by Pre-Embedding Immunoelectron Microscopy

Retinal Cryo-sections, Whole-Mounts, and Hypotonic Isolated Vasculature Preparations for Immunohistochemical Visualization of Microvascular Pericytes

Cryosectioning and Immunostaining of Mouse Retina