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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W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hematoxylin+toluidine+blue-phloxinate+staining+of+glycol+methacrylate+sections+of+retina+and+other+tissues.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Correlation+of+optical+coherence+tomography+and+retinal+histology+in+normal+and+pro23his+retinal+degeneration+pig.">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.
<p class="jove_co...

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.

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			<td>Acetone&#160;</td>
			<td>Chempur&#160;</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>
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			<td>Ethyl alcohol 96 %</td>
			<td>Chempur&#160;</td>
			<td>CH.ET.AL.96%CZDA.CH</td>
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			<td>Ethyl alcohol 99.9 %</td>
			<td>Chempur&#160;</td>
			<td>CH.ETYL.ALK.99,9%BW</td>
			<td>Use to prepare 70% and 80% solutions of EtOH</td>
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			<td>Glacier - 86 &#176;C Ultralow Temperature Freezer NU-9483E</td>
			<td>NuAire</td>
			<td>13027D0034</td>
			<td>Freezing temperature - 80 &#176;C</td>
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		<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>
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			<td>Leica CV5030 Fully Automated Glass Coverslipper</td>
			<td>Leica Biosystems&#160;</td>
			<td>14,04,78,80,101</td>
			<td>Automated slide sealer&#160;</td>
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			<td>Mayer&#39;s hematoxylin</td>
			<td>Chempur&#160;</td>
			<td>124687402</td>
			<td></td>
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			<td>Metal base molds 15 mm&#160; x 15 mm</td>
			<td>Leica Biosystems&#160;</td>
			<td>3803081</td>
			<td></td>
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			<td>Microme HM 340 E</td>
			<td>Thermo Scientific</td>
			<td>90-519-0</td>
			<td>Microtome</td>
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			<td>Microtome razor blades</td>
			<td>Mar-Four</td>
			<td>HI.N.35</td>
			<td>Cutting angle 35&#176;</td>
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		<tr>
			<td>Micro-Twin biopsy cassette</td>
			<td>Mar-Four</td>
			<td>LN.13874550</td>
			<td></td>
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			<td>Microwave Hybrid Tissue Processor</td>
			<td>Milestone</td>
			<td></td>
			<td></td>
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			<td>Multistainer Leica ST5020</td>
			<td>Leica Biosystems&#160;</td>
			<td></td>
			<td>Automated slide stainer&#160;</td>
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			<td>NanoZoomer 2.0-HT slide scanner</td>
			<td>Hamamatsu</td>
			<td>C9600</td>
			<td>Histopathological slide scanner</td>
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			<td>NDP.view2 Image viewing software</td>
			<td>Hamamatsu</td>
			<td>U12388-01</td>
			<td>Image viewing software</td>
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			<td>Paraffin Pathowax Plus</td>
			<td>Mar-Four</td>
			<td>4P.PWP.010</td>
			<td>Melting temperature 56 &#176;C - 58 &#176;C</td>
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			<td>Paraformaldehyde</td>
			<td>Sigma - Aldrich</td>
			<td>441244-1KG</td>
			<td>Prepare a 4% solution in PBS</td>
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			<td>PBS Tablets</td>
			<td>Merck Millipore</td>
			<td>524650-1EA</td>
			<td>Use to prepare a solution of phosphate buffer saline, per manufacturer&#39;s instructions</td>
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			<td>Pierce Microcentrifuge Tubes, 1.5 mL</td>
			<td>Thermo Scientific</td>
			<td>69715</td>
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			<td>Refrigerator-freezer EN14000AW</td>
			<td>Electrolux</td>
			<td>925032562-00</td>
			<td>refrigeration 4 &#176;C</td>
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			<td>Scalpel blade</td>
			<td>Swann-Morton</td>
			<td>T.OST.SKAL00.024</td>
			<td></td>
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			<td>Sucrose</td>
			<td>Chempur&#160;</td>
			<td>427720906</td>
			<td></td>
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			<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>
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			<td>Xylene</td>
			<td>Chempur&#160;</td>
			<td>CH.KSYL.5l.METAL.OP&#160;</td>
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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

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JoVE Journal

Medicine

460 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

Retinal Detachment Model in Rodents by Subretinal Injection of Sodium Hyaluronate

Subretinal injection of sodium hyaluronate is a widely accepted method of inducing retinal detachment (RD). However, the height and duration of RD or the occurrence of subretinal hemorrhage can affect photoreceptor cell death in the detached retina. Hence, it is advantageous to create reproducible RDs without subretinal hemorrhage for evaluating photoreceptor cell death. We modified a previously reported method to create bullous and persistent RDs in a reproducible location with rare occurrence of subretinal hemorrhage. The critical step of this modified method is the creation of a self-sealing scleral incision, which can prevent leakage of sodium hyaluronate after injection into the subretinal space. To make the self-sealing scleral incision, a scleral tunnel is created, followed by scleral penetration into the choroid with a 30 G needle. Although choroidal hemorrhage may occur during this step, astriction with a surgical spear reduces the rate of choroidal hemorrhage. This method allows a more reproducible and reliable model of photoreceptor death in diseases that involve RD such as rhegmatogenous RD, retinopathy of prematurity, diabetic retinopathy, central serous chorioretinopathy, and age-related macular degeneration (AMD).

Creating experimental retinal detachments with a reproducible and sustained height of detachment, and without subretinal hemorrhage, is important for studying the pathophysiology of photoreceptor cell loss in retinal disease and evaluating potential therapeutic interventions. Here, we report such a method in detail.

Subretinal injection of sodium hyaluronate is a widely accepted method of inducing retinal detachment (RD). However, the height and duration of RD or the ...

A Mouse Tumor Model of Surgical Stress to Explore the Mechanisms of Postoperative Immunosuppression and Evaluate Novel Perioperative Immunotherapies

Surgical resection is an essential treatment for most cancer patients, but surgery induces dysfunction in the immune system and this has been linked to the development of metastatic disease in animal models and in cancer patients. Preclinical work from our group and others has demonstrated a profound suppression of innate immune function, specifically NK cells in the postoperative period and this plays a major role in the enhanced development of metastases following surgery. Relatively few animal studies and clinical trials have focused on characterizing and reversing the detrimental effects of cancer surgery. Using a rigorous animal model of spontaneously metastasizing tumors and surgical stress, the enhancement of cancer surgery on the development of lung metastases was demonstrated. In this model, 4T1 breast cancer cells are implanted in the mouse mammary fat pad. At day 14 post tumor implantation, a complete resection of the primary mammary tumor is performed in all animals. A subset of animals receives additional surgical stress in the form of an abdominal nephrectomy. At day 28, lung tumor nodules are quantified. When immunotherapy was given immediately preoperatively, a profound activation of immune cells which prevented the development of metastases following surgery was detected. While the 4T1 breast tumor surgery model allows for the simulation of the effects of abdominal surgical stress on tumor metastases, its applicability to other tumor types needs to be tested. The current challenge is to identify safe and promising immunotherapies in preclinical mouse models and to translate them into viable perioperative therapies to be given to cancer surgery patients to prevent the recurrence of metastatic disease.

A mouse tumor model of surgical stress is used to explore how postoperative immune suppression promotes metastatic disease and to evaluate immunostimulatory perioperative therapies.

Surgical resection is an essential treatment for most cancer patients, but surgery induces dysfunction in the immune system and this has been linked to ...

Slow-release Drug Delivery through Elvax 40W to the Rat Retina: Implications for the Treatment of Chronic Conditions

Diseases of the retina are difficult to treat as the retina lies deep within the eye. Invasive methods of drug delivery are often needed to treat these diseases. Chronic retinal diseases such as retinal oedema or neovascularization usually require multiple intraocular injections to effectively treat the condition. However, the risks associated with these injections increase with repeated delivery of the drug. Therefore, alternative delivery methods need to be established in order to minimize the risks of reinjection. Several other investigations have developed methods to deliver drugs over extended time, through materials capable of releasing chemicals slowly into the eye. In this investigation, we outline the use of Elvax 40W, a copolymer resin, to act as a vehicle for drug delivery to the adult rat retina. The resin is made and loaded with the drug. The drug-resin complex is then implanted into the vitreous cavity, where it will slowly release the drug over time. This method was tested using 2-amino-4-phosphonobutyrate (APB), a glutamate analogue that blocks the light response of the retina. It was demonstrated that the APB was slowly released from the resin, and was able to block the retinal response by 7 days after implantation. This indicates that slow-release drug delivery using this copolymer resin is effective for treating the retina, and could be used therapeutically with further testing.

This paper details how Elvax 40W can be used as a slow-release method for drug delivery to the adult rat retina. The protocol for preparing, loading, and delivering the drug-resin complex to the eye is described.

Diseases of the retina are difficult to treat as the retina lies deep within the eye. Invasive methods of drug delivery are often needed to treat these ...

Do's and Don'ts in the Preparation of Muscle Cryosections for Histological Analysis

Histological evaluation of muscle biopsies has served as an indispensable tool in the understanding of the development and progression of pathology of neuromuscular disorders. However, in order to do so, proper care needs to be taken when excising and preserving tissues to achieve optimal staining. One method of tissue preservation involves fixing tissues in formaldehyde and then embedding them with paraffin wax. This method preserves morphology well and allows for long-term storage at RT but is cumbersome and requires handling of toxic chemicals. Further, formaldehyde fixation results in antigen cross-linking, which necessitates antigen retrieval protocols for effective immunostaining. On the contrary, frozen sectioning does not require fixation and thus retains biological antigen conformation. This method also provides a distinct advantage in quick turn around time, making it especially useful in situations needing quick histological evaluation like intraoperative surgical biopsies. Here we describe the most effective method of preparing muscle biopsies for visualization with different histological and immunological stains.

Do&#039;s and Don&#039;ts in the Preparation of Muscle Cryosections for Histological Analysis

Here we demonstrate the most efficient methods for freezing, embedding, cryosectioning, and staining of muscle biopsies to avoid freezing artifacts.

Histological evaluation of muscle biopsies has served as an indispensable tool in the understanding of the development and progression of pathology of ...

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 ...

Dissection of the Mouse Pancreas for Histological Analysis and Metabolic Profiling

We have been investigating the pancreas specific transcription factor, 1a cre-recombinase; lox-stop-lox- Kristen rat sarcoma, glycine to aspartic acid at the 12 codon (Ptf1acre/+;LSL-KrasG12D/+) mouse strain as a model of human pancreatic cancer. The goal of our current studies is to identify novel metabolic biomarkers of pancreatic cancer progression. We have performed metabolic profiling of urine, feces, blood, and pancreas tissue extracts, as well as histological analyses of the pancreas to stage the cancer progression. The mouse pancreas is not a well-defined solid organ like in humans, but rather is a diffusely distributed soft tissue that is not easily identified by individuals unfamiliar with mouse internal anatomy or by individuals that have little or no experience performing mouse organ dissections. The purpose of this article is to provide a detailed step-wise visual demonstration to guide novices in the removal of the mouse pancreas by dissection. This article should be especially valuable to students and investigators new to research that requires harvesting of the mouse pancreas by dissection for metabolic profiling or histological analyses.

This video article provides a detailed demonstration of the procedures required to successfully remove the pancreas from a mouse by dissection for histological analysis and metabolic profiling.

We have been investigating the pancreas specific transcription factor, 1a cre-recombinase; lox-stop-lox- Kristen rat sarcoma, glycine to aspartic acid at ...

Echocardiographic and Histological Examination of Cardiac Morphology in the Mouse

An increasing number of genetically modified mouse models has become available in recent years. Moreover, the number of pharmacological studies performed in mice is high. Phenotypic characterization of these mouse models also requires the examination of cardiac function and morphology. Echocardiography and magnetic resonance imaging (MRI) are commonly used approaches to characterize cardiac function and morphology in mice. Echocardiographic and MRI equipment specialized for use in small rodents is&#160;expensive and requires a dedicated space. This protocol describes cardiac measurements in mice using a clinical echocardiographic system with a 15 MHz human vascular probe. Measurements are performed on anesthetized adult mice. At least three image sequences are recorded and analyzed for each animal in M-mode in the parasternal short-axis view. Afterwards, cardiac histological examination is performed, and cardiomyocyte diameters are determined on hematoxylin-eosin- or wheat germ agglutinin (WGA)-stained paraffin sections. Vessel density is determined morphometrically after Pecam-1 immunostaining. The protocol has been applied successfully to pharmacological studies and different genetic animal models under baseline conditions, as well as after experimental myocardial infarction by the permanent ligation of the left anterior descending coronary artery (LAD). In our experience, echocardiographic investigation is limited to anesthetized animals and is feasible in adult mice weighing at least 25 g.

Echocardiographic examination is frequently used in mice. Expensive high-resolution ultrasound devices have been developed for this purpose. This protocol describes an affordable echocardiographic procedure combined with histological morphometric analyses to determine cardiac morphology.

An increasing number of genetically modified mouse models has become available in recent years. Moreover, the number of pharmacological studies performed ...

Using a Knee Arthrometer to Evaluate Tissue-specific Contributions to Knee Flexion Contracture in the Rat

Normal knee range of motion (ROM) is critical to well-being and allows one to perform basic activities such as walking, climbing stairs and sitting. Lost ROM is called a joint contracture and results in increased morbidity. Due to the difficulty of reversing established knee contractures, early detection is important, and hence, knowing risk factors for their development is essential. The rat represents a good model with which the effect of an intervention can be studied due to the similarity of rat knee anatomy to that of humans, the rat's ability to tolerate long durations of knee immobilization in flexion, and because mechanical data can be correlated with histologic and biochemical analysis of knee tissue.
Using an automated arthrometer, we demonstrate a validated, precise, reproducible, user-independent method of measuring the extension ROM of the rat knee joint at specific torques. This arthrometer can be used to determine the effects of interventions on knee joint ROM in the rat.

The goal of the protocol is to measure the extension range of motion of the rat knee. The effects of various diseases that increase the stiffness of the knee joint and the effectiveness of treatments can be quantified.

Normal knee range of motion (ROM) is critical to well-being and allows one to perform basic activities such as walking, climbing stairs and sitting. Lost ...

A Non-Invasive Method for Generating the Cyclic Loading-Induced Intra-Articular Cartilage Lesion Model of the Rat Knee

The pathophysiology of primary osteoarthritis (OA) remains unclear. However, a specific subclassification of OA in relatively younger age groups is likely correlated with a history of articular cartilage damage and ligament avulsion. Surgical animal models of OA of the knee play an important role in understanding the onset and progression of post-traumatic OA and aid in the development of novel therapies for this disease. However, non-surgical models have been recently considered to avoid traumatic inflammation that could affect the evaluation of the intervention. 
In this study, an intra-articular cartilage lesion rat model induced by in vivo cyclic compressive loading was developed, which allowed researchers to (1) determine the optimal magnitude, speed, and duration of load that could cause focal cartilage damage; (2) assess post-traumatic spatiotemporal pathological changes in chondrocyte vitality; and (3) evaluate the histological expression of destructive or protective molecules that are involved in the adaptation and repair mechanisms against joint compressive loads. This report describes the experimental protocol for this novel cartilage lesion in a rat model.

Here, we present the cyclic loading-induced intra-articular cartilage lesion model of the rat knee, generated by 60 cyclic compressions over 20 N, resulting in damage to the femoral condylar cartilage in rats.

The pathophysiology of primary osteoarthritis (OA) remains unclear. However, a specific subclassification of OA in relatively younger age groups is likely ...

Evaluation of Capillary and Other Vessel Contribution to Macular Perfusion Density Measured with Optical Coherence Tomography Angiography

Parafoveal circulation of the superficial retinal capillary plexus is usually measured with vessel density, which determines the length of capillaries with circulation, and perfusion density, which calculates the percentage of the evaluated area that has circulation. Perfusion density also considers the circulation of vessels larger than capillaries, although the contribution of these vessels to the first is not usually evaluated. As both measurements are automatically generated by optical coherence tomography angiography devices, this paper proposes a method for estimating the contribution of vessels larger than capillaries by using a coefficient of determination between vessel and perfusion densities. This method can reveal a change in the proportion of perfusion density from vessels larger than capillaries, even when mean values do not differ. This change could reflect compensatory arterial vasodilatation as a response to capillary dropout in the initial stages of retinal vascular diseases before clinical retinopathy appears. The proposed method would allow the estimation of the changes in the composition of perfusion density without the need for other devices.

We describe the evaluation of a coefficient of determination between vessel and perfusion density of the parafoveal superficial capillary plexus to identify the contribution of vessels larger than capillaries to perfusion density.

Parafoveal circulation of the superficial retinal capillary plexus is usually measured with vessel density, which determines the length of capillaries ...