The authors wish to thank: Ms. Alexia Saunders and Ms. Michelle McPhedran for experimental assistance; Ms. Helen Feng for electrode fabrication; Dr. Penny Allen and Dr. Jonathan Yeoh for surgical assistance; Staff of the Royal Victorian Eye and Ear Hospital's Biological Research Centre for animal care; Prof. Rob Shepherd for general guidance throughout; and Dr. Bryony Nayagam and Mr. Ronald Leung for critical comments on a draft form of the manuscript.
This work was performed at the Bionics Institute and St Vincent's Hospital, Melbourne. Funding was provided by the Ian Potter Foundation, the John T Reid Charitable Trusts, and the Australian Research Council through its Special Research Initiative in Bionic Vision Science and Technology grant to Bionic Vision Australia (BVA). The Bionics Institute acknowledges the support it receives from the Victorian Government through its Operational Infrastructure Support Program.

1. Enucleation and Post-Fixation
This protocol assumes that the subject has been implanted unilaterally with a suprachoroidal electrode array.
<ol>
	<li>Prepare postfix solutions in glass Schott bottles.
	<ol class="lalpha">
		<li>Davidson's fixative solution, 2x 100 ml
		<ul>
			<li>2 ml formalin (37% formaldehyde)</li>
			<li>10 ml glacial acetic acid (99.7%)</li>
			<li>35 ml ethanol (98%)</li>
			<li>53 ml distilled water</li>
		</ul>
		</li>
		<li>50% ethanol, 2x 100 ml</li>
		<li>70% ethanol, 2x 100 ml</li>
	</ol>
	</li>
	<li>Transcardially perfuse subject with warm (37 &deg;C) heparinized saline followed by cold (4 &deg;C) neutral buffered formalin (10%).</li>
	<li>Tie sutures at the superior limbus (or a location that is remote from the suprachoroidal array) of both eyes - in line with a rectus muscle, to serve as a landmark.</li>
	<li>Enucleate eyes, maintaining any external leads from the implanted eye.</li>
	<li>Place each eye in 100 ml of Davidson's fixative solution and leave to post-fix at room temperature.</li>
	<li>After 18 - 36 hr in Davidson's fixative, transfer to 50% ethanol (room temperature) for 6 - 8 hr, then finally to 70% ethanol (room temperature).</li>
	<li>Refrigerate samples at 4 &deg;C and store until dissection.</li>
</ol>
Intact and pressurized eye globes have been stored in this manner for up to 3 months without adversely affecting the resultant histology.
2. Identification and Tissue Mapping
The above fixation protocol (step 1) has rendered the sclera translucent and the array is now visible - including the individual electrode sites (Figure 3).
<ol>
	<li>Remove the implanted globe from ethanol and carefully trim away excess tissue, including conjunctiva and Tenon's capsule. Trim optic nerve to a 2 - 3 mm length (Figure 3A).</li>
	<li>Using a histological dye marking scheme, label the electrodes with a predefined color code, taking care not to smudge the dye.</li>
</ol>
<ul>
	<li>We chose a commercially available suite of specialized histological dye (refer to appendix). Small amounts of the dye were carefully applied with a fine tipped paint brush (Roymac size 00) to the tissue and allowed to air dry for up to 5 min.</li>
	<li>For our purposes, we have chosen: green for the active electrode(s) that was(were) stimulated maximally, at suprathreshold levels, for the duration of the study; red for other active electrodes; yellow for the larger diameter return electrodes; and blue for additional guides and/or anatomical distance markings (Figures 3B and 3C).</li>
	<li>Some trial and error may be required to find a dye that is stable throughout the subsequent steps. For example, in Figure 4, it can be seen that the green dye was more resilient than the red dye. Diluting the dye in 70% ethanol helps improve lipid penetration and tissue coating.</li>
	<li>It is useful to sketch or photograph the dyed globe for future reference.</li>
</ul>
<ol start="3">
	<li>Return to 70% ethanol to dehydrate the dye.</li>
	<li>Repeat step 2.1 for the unimplanted control globe.</li>
	<li>Using the sutures from step 1.3, align the control eye and the implanted eye as a mirrored pair.</li>
	<li>Place a silicone template (with the same dimensions as the electrode array) in the mirrored location on the control eye as the implant is located on implanted eye (N.B. the translucent sclera and the dye markings from step 2.2 allow ready visualization of the implanted array position).</li>
	<li>Perform steps 2.2 and 2.3 for the control eye, so that each implanted electrode site has a control pair in an anatomically comparable (i.e. mirror matched equivalent) location.</li>
</ol>
3. Dissection and Embedding
<ol>
	<li>Remove the implant array from the eye and remove the front of the eye (including the cornea, iris and lens). Remove the vitreous fluid from the remaining eye cup (posterior chamber).</li>
	<li>Dissect the implanted eye into multiple representative strips (~2 mm thickness) each containing a subset of the dye marked regions (Figure 3D). Note that the orientation of the strips should be selected to assist in assessing various aspects of the tissue response to the implant7.</li>
</ol>
It is useful, for future reference, to make a record of which dye regions are present on each strip and their relative locations (and colors).
<ol start="3">
	<li>Place the strip on its side into a shallow pool of liquefied agar (4%; 80 - 90 &deg;C) with the side to be cut first facing downwards (Figure 3E).</li>
	<li>Once the agar has set, cut around the sample and place into a tissue cassette supported by foam inserts (Figure 3F).</li>
	<li>Repeat steps 3.1-3.3 for the control eye - taking care to dissect mirror-matched strips.</li>
	<li>Process all cassettes via standard (automated overnight) paraffin processing technique.</li>
	<li>Embed processed tissue in paraffin with the side to be cut first facing downwards.</li>
</ol>
4. Cutting and Staining
<ol>
	<li>Cut paraffin blocks into 5 &mu;m sections referring regularly to the notes from Steps 2 and 3.</li>
	<li>Collect sections from each of the dyed regions and mount on slides.</li>
</ol>
The region immediately adjacent to each dyed spot of sclera can now be assessed with confidence that it was in closest proximity to the corresponding electrode in the array (Figure 4).
<ol start="3">
	<li>Stain or perform immunofluorescence as desired. Example stains and immunohistochemistry shown in Figure 5 are: H&amp;E; Luxol fast blue (LFB); cresyl violet; Masson's trichrome blue; periodic acid schiff (PAS); Perls' Prussian blue stain; anti-glutamine synthetase (GS); anti-neurofilament protein (NF); anti-glial fibrillary acidic protein (GFAP) and 4',6-diamidino-2-phenylindole (DAPI).</li>
	<li>Standard and special stains were performed as detailed:
	<ol>
		<li>H&amp;E staining was performed according to the protocol provided by Leica multistainer bath array ST5020 (Leica Biosystems).</li>
		<li>LFB and cresyl violet (modified from 11):
		<ol>
			<li>Dewax in 3 changes of xylene (3x 2 min) and 2 changes of ethanol (100%, 100%) (2x 1 min) and rinse in tap water (45 sec).</li>
			<li>Hydrate sections to 95% ethanol.</li>
			<li>Place in LFB solution 11 at 37 - 42 &deg;C (overnight).</li>
			<li>Rinse off excess stain with 70% ethanol.</li>
			<li>Rinse in distilled water.</li>
			<li>Place in dilute lithium carbonate (8%) (few seconds).</li>
			<li>Differentiate in 70% ethanol (few seconds).</li>
			<li>Rinse in distilled water.</li>
			<li>Repeat steps 4.4.2.6 to 4.4.2.8, if necessary, until background is colorless.</li>
			<li>Place in cresyl violet acetate working solution at 37 &deg;C (10 min).</li>
			<li>Do not wash in water.</li>
			<li>Dehydrate in 3 changes of absolute alcohol (1x 30 sec, 2x 20 sec) and 3 changes of xylene (1x 1 min 30 sec, 2x 1 min).</li>
			<li>Mount and coverslip with DPX.</li>
		</ol>
		</li>
		<li>Masson's trichrome blue (modified from 11):
		<ol>
			<li>Dewax as per 4.4.2.1.</li>
			<li>Bring sections to water (2 min).</li>
			<li>Stain the nuclei using Weigert's iron haematoxylin (2 min).</li>
			<li>Wash in tap water, rinse in distilled water.</li>
			<li>Stain in Biebrich scarlet-acid fuchsin solution (5 min).</li>
			<li>Rinse in distilled water.</li>
			<li>Differentiate in phosphomolybdic-phosphotungstic acid until collagen is pale pink (6 min).</li>
			<li>Rinse in distilled water.</li>
			<li>Counterstain in aniline blue (1 min).</li>
			<li>Wash well in 1% acetic acid (1 min).</li>
			<li>Dehydrate, mount and coverslip as per 4.4.2.12 and 4.4.2.13.</li>
		</ol>
		</li>
		<li>PAS (modified from 11):
		<ol>
			<li>Dewax as per 4.4.2.1.</li>
			<li>Oxidize in 1% periodic acid (15 min).</li>
			<li>Wash in running water then distilled water.</li>
			<li>Stain in Schiff's reagent (15 min).</li>
			<li>Wash in water (5 min).</li>
			<li>Counterstain with Harris haematoxylin (1 min).</li>
			<li>Wash briefly in tap water.</li>
			<li>Differentiate in 0.5% acid alcohol (1 dip).</li>
			<li>Wash well in tap water.</li>
			<li>Blue in Scott's tap water (1 min).</li>
			<li>Wash well in water.</li>
			<li>Dehydrate, mount and coverslip as per 4.4.2.12 and 4.4.2.13.</li>
		</ol>
		</li>
		<li>Perls' Prussian blue stain as described in 11.</li>
	</ol>
	</li>
	<li>Immunofluorescence staining was performed as detailed:
	<ol>
		<li>Dewax as per 4.4.2.1.</li>
		<li>Rinse in distilled water (2x 5 min).</li>
		<li>Perform antigen retrieval using 0.01% citrate buffer, pH 6 at 75 - 80 &deg;C (20 min) and allow to cool in 0.01% citrate buffer solution (20 min).</li>
		<li>Wash sections in phosphate buffered saline (PBS) (2x 5 min).</li>
		<li>Permeabilize the cell membrane in wash buffer solution (10 min).</li>
		<li>Incubate in serum block solution (2 hr).</li>
		<li>Incubate sections in a single primary antibody diluted in antibody diluent (10% normal goat serum/0.1% Triton X-100/PBS) (overnight in fridge). The titre of each primary antibody used is shown in Table 1.</li>
		<li>After overnight incubation, wash sections in wash buffer solution (5x 3 min). From this point on, keep sections in the dark.</li>
		<li>Incubate sections in the corresponding secondary antibody at a titre of 1:500 for a specific duration (see Table 1 for details).</li>
		<li>Wash sections in wash buffer solution (3x 5 min).</li>
		<li>Incubate sections in DAPI (1:25,000 / PBS) (30 min).</li>
		<li>Wash sections in wash buffer solution (3x 5 min).</li>
		<li>Mount and coverslip using fluorescent mounting media.</li>
	</ol>
	</li>
</ol>
Test samples and control samples are ready for pairwise comparison.

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Neural Eng. 2, S121-S128 (2005).</li><li>Morimoto, T., Kamei, M., 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=Chronic+implantation+of+newly+developed+suprachoroidal-transretinal+stimulation+prosthesis+in+dogs.">Chronic implantation of newly developed suprachoroidal-transretinal stimulation prosthesis in dogs.</a> Invest. Ophthalmol. Vis. Sci. 52, 6785-6792 (2011).</li><li>Bron, A. J., Tripathi, R. C., Tripathi, B. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Wolff's anatomy of the eye and orbit. , (2001).</li><li>Stein, J. J., Johnson, S. A., Berson, D. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Distribution+and+coverage+of+beta+cells+in+the+cat+retina.">Distribution and coverage of beta cells in the cat retina.</a> J. Comp. Neurol. 372, 597-617 (1996).</li><li>Dagnelie, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Psychophysical+evaluation+for+visual+prosthesis.">Psychophysical evaluation for visual prosthesis.</a> Annu. Rev. Biomed Eng. 10, 339-368 (2008).</li></ol>

Authors have nothing to disclose.

Assessing the safety of the implant is a primary consideration for preclinical studies. Before a retinal prosthesis can be implanted in humans it is essential to verify that it will not cause any harm. This requires an assessment of both the implant biocompatibility 17-23 as well as any potential damage that could be caused by chronic electrical stimulation 24-26. Experience with similar neural prostheses, such as the cochlear implant, provides insight into the broad range of stimulation, and physical, parameters that do not cause tissue damage 27. However, the retina has different characteristics to other implantation sites and therefore precise safety limits (such as maximum charge density) cannot be assumed from previous studies in other systems. Charge densities are highest at the active electrode sites and this corresponds with the greatest potential for damage. Therefore a method for localizing the tissue directly adjacent to the individual active electrodes would greatly increase the usefulness of these studies. The tissue adjacent to specific regions of the implant may also be highlighted for closer inspection. This targeted approach permits fewer sections to be analyzed, whilst retaining confidence that the "worst case scenario" was examined.
The scleral dye localization method described here has been extensively tested with hand-shaped and molded silicone/Pt electrode arrays 28-30. It has been used with thin-film polyamide arrays 7,31 and also thin-film silicone/Pt arrays 32. Some retinal prosthesis studies employed "bullet shaped" electrodes with intrascleral implantations 33. The present technique should be effective for assessing this type of electrode arrays. Feline eyes with thin sclera (thickness at posterior pole 90 - 200 &mu;m) allowed visualization of individual electrodes in an array. However, even if individual electrodes were not visible, their positions could be readily inferred using a silicone template when the outline of the array can be seen (similar to that used in step 2.6).
The dye mapping limits the need to obtain non-relevant tissue sections as only sections with the dye present are obtained. The ability to accurately infer electrode position not only allows relevant histopathological analysis and greater statistical power, but has a major impact on time and cost management by reducing the quantity of slides and blades used as well as the cost of labor. The use of dye that is adherent and can withstand tissue processing whilst also being visible in unstained tissue sections is an important initial consideration. Knowledge of the location of the dye and orientation of the tissue at dissection and during embedding assists the cutting process. This includes correctly orientating the block to achieve a section that is not obliquely cut and gives a full representation of the tissue. It is therefore important that the person performing the cutting is present at the time of dye application, dissection and embedding.
The overall protocol is robust to minor alteration, particularly with fixation timings. However, the eye dissection and dye marking steps are delicate procedures that benefit from good manual dexterity to avoid damaging the specimen. Whilst Davidson's fixative has proved effective for reducing artifactual detachment of the retina near an implant, it does not prevent direct mechanical damage during dissection. Davidson's fixative was not readily compatible with some special histological and immunohistochemical procedures. Therefore there was a need to modify/optimize these steps as detailed above. Incremental changes in staining times and concentrations were made until the optimum result was achieved - for more details, refer to the supplementary materials.
Quantification of retinal histology remains problematic. The retina is non-homogeneous and both the thickness and the cell density change with increasing distance from area centralis 34,35. Therefore, neighboring tissue within the implanted eye cannot be used for comparisons and a control eye is employed instead. Pairwise matching of each section with the control eye gives us a more powerful statistical comparison of pathological changes; efficacy and safety outcomes with retinal prosthetics are better assessed when comparing the fellow eyes in a subject 36. Special care must be taken to minimize oblique cutting as this would affect quantification.
In summary, the methods described above have significantly improved our histopathological analysis, which in turn has led to a more efficient preclinical workflow. The results of preclinical studies using these methods directly led to a clinical trial in which 3 RP patients have been safely implanted with suprachoroidal retinal prostheses. These methods are relevant both to retinal stimulators and other ocular implants where the pathological response to long term implantation needs to be evaluated. This method provided worthwhile advantages for maintaining the cytoarchitecture and registration of the pathology to regions of interest on the implant. Although not specifically tested, a modification of these procedures may be used in other species and other anatomical locations, particularly where an array is implanted superficially.

Retinal prostheses may soon become useful clinical interventions for treating various forms of severe vision loss. Certain conditions, such as retinitis pigmentosa (RP), result in the widespread degeneration of the photoreceptors in the eye; these are the cells responsible for transducing light into neural signals. However, some cells in other layers of the retina remain and are potential targets for electrical stimulation with a retinal prosthesis 1. Early results from human clinical trials have been promising for electrode arrays implanted in the epiretinal 2,3, subretinal 4,5, and intrascleral 6 locations. These devices are made from different materials with different shapes, but they all use electrical pulses to activate the remaining viable neurons of the retina in order to create visual percepts.
Our wider group (Bionic Vision Australia) has developed a retinal implant that has progressed to a clinical pilot study with 3 RP patients implanted with suprachoroidal electrode arrays (Clinical Trial Number: NCT01603576). Figure 1A shows this suprachoroidal prototype retinal prosthesis.
Patient safety is paramount in clinical studies and thus, before commencing human trials, we performed extensive acute and chronic testing in preclinical models (Figure 1B). Histopathological assessments were essential to refine the surgical procedures, iterate through electrode design, and ultimately establish the safety of the implant. To maintain an efficient workflow dovetailed with standard clinical pathology practices, a paraffin embedding technique was employed. It was also desirable to utilize staining procedures comparable with clinical standards, such as haematoxylin and eosin (H&amp;E), which are readily interpreted by pathologists. We also wanted a technique that could be adapted for immunohistochemical analyses, in order to facilitate further cellular evaluation of the tissues.
The biocompatibility of the implant materials, and the safety of chronic electrical stimulation, need to be carefully assessed. It is important to be able to examine the histopathology of the ocular tissue adjacent to the individual stimulating electrodes within the array. However, the arrays needed to be removed prior to processing the samples so they can be tested, and because their metallic components could not be readily sectioned. Consequently, our established tissue preparation did not permit localization and mapping of tissue with respect to the implanted electrodes 7. In addition to the lack of a tissue localization method, the standard formaldehyde-based fixation and processing techniques resulted in widespread artifacts and detachment of the retinal due to the differential shrinkage of the various layers of the eye (Figure 2). Due to the non-homogeneity of the retina, it was difficult to make valid comparisons with control tissue, particularly for quantitative measurements. These combined limitations complicated accurate assessments of the potential damage caused by our implanted arrays.
Here we present novel techniques for overcoming these limitations. We have modified specialized eye fixation and processing techniques 8-10 to maintain compatibility with paraffin-based histology. We have modified standard histological and immunohistochemical stains to give comparable results, based on feedback from clinical pathologists. After rendering the scleral tissue translucent, a dye-based color-code was employed to mark the ocular tissue adjacent to each individual electrode, before removing the array from the suprachoroidal space. By marking the electrode array and the individual electrode sites, samples could be accurately collected from electrode adjacent regions. Matched samples could be gathered from the control eye in order to facilitate pairwise comparisons.

<table><tbody><tr><td>The Davidson marking system</td><td>Bradley Products</td><td>1163-4 - red 1163-5 - blue 1163-2 - yellow 1163-1- green</td><td>&amp;nbsp;</td></tr><tr><td>Rabbit Polyclonal Anti-Glial Fibrillary Acidic Protein Antibody</td><td>Millipore</td><td>AB5804</td><td>1:1500, overnight incubation</td></tr><tr><td>Mouse Monoclonal Anti-Glutamine Synthetase Antibody</td><td>Millipore</td><td>MAB302</td><td>1:100 overnight incubation</td></tr><tr><td>Monoclonal Mouse Anti-Neurofilament 200 Antibody</td><td>Sigma-Aldrich</td><td>N0142</td><td>1:100 overnight incubation</td></tr><tr><td>4&#039;,6-diamindino-2-phenylindole, dilactate (dilactate)</td><td>Life Technologies</td><td>D3571</td><td>1:25,000 30 min incubation</td></tr><tr><td>Alexa Fluor 488 goat anti-mouse IgG</td><td>Life Technologies</td><td>A11029</td><td>1:500</td></tr><tr><td>Superfrost 90&amp;deg; yellow</td><td>Grale Scientific</td><td>SF41299</td><td>&amp;nbsp;</td></tr><tr><td>FLEX IHC Microscope Slides</td><td>DAKO</td><td>K802021-2</td><td>&amp;nbsp;</td></tr><tr><td>Alexa Fluor 594 goat anti-rabbit IgG</td><td>Life Technologies</td><td>A11037</td><td>1:500</td></tr><tr><td>Normal goat serum</td><td>Life Technologies</td><td>PCN5000</td><td>10% serum/0.1% Triton X-100/PBS</td></tr><tr><td colspan="4"> &lt;p class=&quot;jove_content&quot;&gt;&lt;strong&gt;Non-Standard Solution Preparation for Special Stains&lt;/strong&gt;&lt;/p&gt; &lt;p class=&quot;jove_content&quot;&gt;&lt;em&gt;1% Aqueous Cresyl Violet Stock Solution&lt;/em&gt;&lt;/p&gt; &lt;ul&gt; &lt;li&gt;Cresyl violet 1 g&lt;/li&gt; &lt;li&gt;Distilled water 100 ml&lt;/li&gt; &lt;/ul&gt; &lt;p class=&quot;jove_content&quot;&gt;&lt;em&gt;Cresyl Violet Acetate Working Solution&lt;/em&gt;&lt;/p&gt; &lt;ul&gt; &lt;li&gt;1% aqueous cresyl violet stock solution 9 ml&lt;/li&gt; &lt;li&gt;Distilled water 51 ml&lt;/li&gt; &lt;li&gt;10% acetic acid 0.5 ml&lt;/li&gt; &lt;/ul&gt; &lt;p class=&quot;jove_content&quot;&gt;&lt;em&gt;Biebrich Scarlet-Acid Fuchsin Solution&lt;/em&gt;&lt;/p&gt; &lt;ul&gt; &lt;li&gt;1% Beihrich scarlet 90 ml&lt;/li&gt; &lt;li&gt;1% acid fuchsin 10 ml&lt;/li&gt; &lt;li&gt;Glacial acetic acid 1 ml&lt;/li&gt; &lt;/ul&gt; &lt;p class=&quot;jove_content&quot;&gt;&lt;em&gt;Phosphomolybdic-Phosphtungstic Acid Solution&lt;/em&gt;&lt;/p&gt; &lt;ul&gt; &lt;li&gt;Phosphomolybdic acid 2.5 g&lt;/li&gt; &lt;li&gt;Phosphotungstic acid 2.5 g&lt;/li&gt; &lt;li&gt;Distilled water 100 ml&lt;/li&gt; &lt;/ul&gt; &lt;p class=&quot;jove_content&quot;&gt;&lt;em&gt;Aniline Blue Solution&lt;/em&gt;&lt;/p&gt; &lt;ul&gt; &lt;li&gt;Aniline blue 2.5 g&lt;/li&gt; &lt;li&gt;Glacial acetic acid 2 g&lt;/li&gt; &lt;li&gt;Distilled water 100 ml&lt;/li&gt; &lt;/ul&gt; &lt;p class=&quot;jove_content&quot;&gt;&lt;strong&gt;Solutions for Immunohistochemistry&lt;/strong&gt;&lt;/p&gt; &lt;p class=&quot;jove_content&quot;&gt;&lt;em&gt;Wash Buffer Solution&lt;/em&gt;&lt;/p&gt; &lt;ul&gt; &lt;li&gt;0.1% Triton X-100 in Phosphate buffered saline (PBS)&lt;/li&gt; &lt;/ul&gt; &lt;p class=&quot;jove_content&quot;&gt;&lt;em&gt;Serum Block Solution&lt;/em&gt;&lt;/p&gt; &lt;ul&gt; &lt;li&gt;10% normal goat serum/0.1% Triton X-100 in PBS&lt;/li&gt; &lt;/ul&gt; &lt;p class=&quot;jove_content&quot;&gt;&lt;strong&gt;SUPPLEMENTARY MATERIAL&lt;/strong&gt;&lt;/p&gt; &lt;p class=&quot;jove_content&quot;&gt;&lt;em&gt;Luxol Fast Blue&lt;/em&gt;&lt;/p&gt; &lt;p class=&quot;jove_content&quot;&gt;The purpose of performing a Luxol Fast Blue (LFB) stain was to identify the ganglion cells in the ganglion cell layer through the counterstain of cresyl violet. While LFB stains myelin, the cresyl violet stain binds to the Nissl substances in neurons, composed of rough endoplasmic reticulum and ribosomes. Numerous cells in the retina contain Nissl substance including amacrine cells. These cells usually are located in the inner boundary layer of the inner nuclear layer, but displaced amacrine cells can be found in the ganglion cell layer. The staining of Nissl substance is helpful in differentiating large, heavily stained ganglion cells from smaller displaced amacrine cells. To aid visualisation of these cells, we modified the cresyl violet working solution protocol by increasing the volume of the cresyl violet stock solution in the working solution. We increased the volume in 1.0 ml increments from 6.0 ml to 9.0 ml, in turn reducing the volume of distilled water. Assessing the ease of visualising and identifying the ganglion cells, optimal staining was achieved with 9 ml of additional cresyl violet stock solution and 51 ml of distilled water. Cresyl violet is a basic dye, though to dye the Nissl substance, it is required to be used in an acidic solution, and therefore the volume of acetic acid remained unchanged at 0.5 ml.&lt;/p&gt; &lt;p class=&quot;jove_content&quot;&gt;&lt;em&gt;Periodic Acid Schiff&lt;/em&gt;&lt;/p&gt; &lt;p class=&quot;jove_content&quot;&gt;The Periodic Acid Schiff (PAS) stain was performed to highlight polysaccharides especially glycogen, found in basement membranes and fungal cell walls, indicating a fungal infection. The process of the PAS stain is to oxidise the hydroxyl groups in polysaccharides using periodic acid, producing aldehyde groups. The application of the Schiff reagent, binds to the aldehyde groups producing a magenta color with haematoxylin counterstaining producing purple cell nuclei. Our slides showed weak staining at the inner limiting membrane. This may be due to the polysaccharides present, as not all polysaccharides can be oxidised with a standard time frame, especially those containing anionic polysaccharides. We therefore increased the duration of the oxidation step from 10 min to 12, 15, and 20 min. We found that a 15 min oxidation step was most effective in PAS staining.&lt;/p&gt; &lt;p class=&quot;jove_content&quot;&gt;&lt;em&gt;Masson&amp;rsquo;s Trichrome Blue&lt;/em&gt;&lt;/p&gt; &lt;p class=&quot;jove_content&quot;&gt;The principle of the Masson&#039;s trichrome blue stain is to stain collagen and muscle, which in our retinal slides can be used to indicate an increase in collagen tissue which may indicate fibrosis due to injury. This stain is sensitive to fixation techniques, so alterations were needed to obtain optimal staining. The process of this stain is to initially stain cytoplasm and muscle fibres red using the acidic Biebrich scarlet-acid fuchsin red dye. Differentiation with phosphomolybdic/phosphotungstic acid competes with the red dye in the collagen fibres. In the sclera, the large phosphomolybdic/phosphotungstic acid molecules are able to penetrate the loose connective tissue, as opposed to the dense muscle fibres which are penetrable to small molecules only. Aniline blue dye is then applied to replace the acid in the collagen fibres producing blue dyed sclera. To optimise this stain in retinal sections, the duration of staining with Biebrich scarlet-acid fuchsin dye was reduced to create a balance between the blue and the red dye. The use of Davidson&#039;s fixative also required an extended differentiation time to remove the red dye from the collagen. We trialled differentiation at 5, 6, 8, and 10 min, with optimal results occurring at 6 min. Differentiation also varies on the thickness and cutting of smooth sections, with greater than 6 min differentiation possibly being required.&lt;/p&gt; &lt;p class=&quot;jove_content&quot;&gt;&lt;em&gt;Glial Fibrillary Acidic Protein (GFAP) Antibody&lt;/em&gt;&lt;/p&gt; &lt;p class=&quot;jove_content&quot;&gt;Polyclonal, host species is rabbit, has a molecular weight of 50 kDa. GFAP antigen is an intermediate filament found in the cytoplasm (the cytoplasm of astrocytes and M&amp;uuml;ller cells in the retina), the antigen is made form 428 amino acids and has a molecular weight of 49512 Da.&lt;/p&gt; &lt;p class=&quot;jove_content&quot;&gt;&lt;em&gt;Neurofilament-200 antibody&lt;/em&gt;&lt;/p&gt; &lt;p class=&quot;jove_content&quot;&gt;Monoclonal, host species is mouse, clone N52, isotype IgG1, it recognises phosphorylated and non-phosphorylated forms of the heavy neurofilaments which have a molecular weight of 200 kDa. The antibody will bind to an epitope in the tail domain of the neurofilament. The antibody will stain neurofilaments in the neuronal perikarya, dendrites and axons.&lt;/p&gt; &lt;p class=&quot;jove_content&quot;&gt;&lt;em&gt;Glutamine Synthetase (GS) antibody&lt;/em&gt;&lt;/p&gt; &lt;p class=&quot;jove_content&quot;&gt;Monoclonal, clone GS-6, host species is mouse, has a molecular weight of 45 kDa and antibody isotype IgG2a. The GS antigen is found in the cell cytoplasm (cytoplasm of M&amp;uuml;ller cells in the retina), the antigen has 373 amino acids and a molecular weight of 42,064 Da.&lt;/p&gt; </td></tr></tbody></table>

techniques for processing eyes implanted with a retinal prosthesis for localized histopathological analysis

With the recent development of retinal prostheses, it is important to develop reliable techniques for assessing the safety of these devices in preclinical studies. However, the standard fixation, preparation, and automated histology procedures are not ideal. Here we describe new procedures for evaluating the health of the retina directly adjacent to an implant. Retinal prostheses feature electrode arrays in contact with eye tissue. Previous methods have not been able to spatially localize the ocular tissue adjacent to individual electrodes within the array. In addition, standard histological processing often results in gross artifactual detachment of the retinal layers when assessing implanted eyes. Consequently, it has been difficult to assess localized damage, if present, caused by implantation and stimulation of an implanted electrode array. Therefore, we developed a method for identifying and localizing the ocular tissue adjacent to implanted electrodes using a (color-coded) dye marking scheme, and we modified an eye fixation technique to minimize artifactual retinal detachment. This method also rendered the sclera translucent, enabling localization of individual electrodes and specific parts of an implant. Finally, we used a matched control to increase the power of the histopathological assessments. In summary, this method enables reliable and efficient discrimination and assessment of the retinal cytoarchitecture in an implanted eye.

Figure 4 shows a representative section obtained from cutting the sample depicted in Figure 3. The section was cut at 5 &mu;m thickness and stained with H&amp;E according to the protocols described above. The gross shape of the sample strip is preserved with minimal differential tissue artifacts (Figure 4A). Both dye markings were visible on the sclera, although the green dye was more resilient than the red (Figure 4B). The retinal layers were not artifactually detached and the retinal morphology is preserved (Figure 4C). The retinal tissue adjacent to the dye markings, which corresponds to the location of the electrodes in the array, can be readily identified.
Figure 5 shows representative special staining and immunohistochemistry of tissue sections prepared according to the current protocol. Refer to Figure 5 caption and Supplementary Materials for more detail on the specific stains and the modifications made to their usage. Post-modification, these stains and immunohistochemistry were equivalent to established standards - as verified by pathologists.
<table border="1">
	<tbody>
		<tr>
			<td>Type of primary antibody and titre (in parenthesis)</td>
			<td>GFAP (1:1500)</td>
			<td>NF200 (1:100)</td>
			<td>GS (1:100)</td>
		</tr>
		<tr>
			<td>Type of secondary antibody and titre (in parenthesis)</td>
			<td>Alexa Fluor 594</td>
			<td>Alexa Fluor 488</td>
			<td>Alexa Fluor 488</td>
		</tr>
		<tr>
			<td>Duration of incubation of secondary antibody</td>
			<td>1 hr</td>
			<td>2 hr</td>
			<td>45 min</td>
		</tr>
	</tbody>
</table>
Table 1. Antibody Type, Titre and Duration of Labeling.
<img alt="Figure 1" src="/files/ftp_upload/50411/50411fig1.jpg" /> 
Figure 1. Suprachoroidal Prototype Electrode Array. A) Macro photograph of a clinical grade molded array. B) Photomicrograph of a handmade preclinical array.
<img alt="Figure 2" src="/files/ftp_upload/50411/50411fig2.jpg" /> 
Figure 2. Former Retinal Histology. Standard histological methods were used to prepare these, H&amp;E stained, sections of an eye implanted with a suprachoroidal electrode array. A) An orthogonal section through the electrode array cavity (depicted by a star). The retina is detached from the outer eye tissue. This is particularly evident beneath the implanted region (arrow). It is impossible to determine which portion of retina was adjacent to each individual electrode of the array. Scalebar = 1 mm. B) A higher magnification of the boxed region in Panel A. There are several, regularly spaced, artifacts in the retinal layers (arrows), as well as major detachment from the tapetal layer (the reflective layer in feline eyes). Scalebar = 100 &mu;m. C) A higher magnification of the boxed region in panel B. Arrow indicated the outer segments of the photoreceptors, as well as the pigmental epithelium, which remain intact, suggesting that the detachment was an artifactual side-effect of the processing, as opposed to being caused by an in vivo trauma or pathology. Scalebar = 20 &mu;m.
<img alt="Figure 3" src="/files/ftp_upload/50411/50411fig3.jpg" /> 
Figure 3. Localization and Dissection of Electrode-Adjacent Tissue. A-D) High dynamic range macro photographs of an enucleated eye, with a suprachoroidal electrode array in situ. A) Post fixed with Davidson's fixative, and prior to array removal, the translucent sclera enables visualization of the individual electrodes in the array (single example indicated with arrow). B) The electrodes are marked with a predefined dye color. C) Dye markings at regular spacing from anatomical landmarks are used to maintain consistency across experiments. D) Upon removal of the array, the eye is dissected by hand into sample strips. Colored arrows indicate two of the electrode adjacent sites on the sclera. Yellow dashed line indicates plane of sectioning. E) Macro-photograph of sample strip embedded within an agar block. F) Macro photograph of the agar-embed sample strip from Panel E, cut out and placed in an embedding cassette supported by foam biopsy pads to minimize movement of the section during processing.
<img alt="Figure 4" src="/files/ftp_upload/50411/50411fig4.jpg" /> 
Figure 4. New Retinal Histology. The above sample strip (from Figure 3D), containing the electrode pocket, was paraffin embedded, sectioned at 5 &mu;m and stained with H&amp;E. A) Green and red dyed regions (indicated by arrows, same as Figure 3D) are visible in a section taken at the level of the yellow dashed line in Figure 3D. Scale bar = 1,000 &mu;m. The retina is not detached, neither beneath the array pocket nor remotely. B) The boxed region from Panel A with visible red and green dye indicated by arrows, and intact retina throughout. Scale bar = 500 &mu;m. C) The boxed region from Panel B, showing the intact, attached, retina adjacent to the green dye. Scale bar = 50 &mu;m. Knowing that the location of the dye indicates the position of an electrode, we can confidently assess the adjacent retinal tissue for damage, if any, caused by electrical stimulation.
<img alt="Figure 5" src="/files/ftp_upload/50411/50411fig5.jpg" /> 
Figure 5. Examples of modified retinal cytohistochemistry using special stains and immunofluorescence. The use of Davidson's fixative, as opposed to standard formalin fixation techniques, required alterations to the usual staining protocols, as outlined in the main protocol text. Pathologists have confirmed that these modifications resulted in equivalent staining. A) H&amp;E; haematoxylin stains cell nuclei purple while eosin stains cell cytoplasm, collagen and supportive tissue various shades of pink. B) LFB stains myelin blue while the cresyl violet counterstain can be used to identify ganglion cells by staining Nissl bodies within the perikaryon blue and highlighting the satellite cells surrounding the cells. C) Masson's trichrome blue; the Biebrich scarlet-acid fuchsin solution stains muscle fibers red, while the aniline blue stains the collagen, in this case sclera, blue. D) PAS; glycoprotein components of basement membrane and connective tissue components are stained purple, while the haematoxylin counterstains cell nuclei purple. E) Perls' Prussian blue; at the site of haemorrhage, the formation of haemosiderin from degraded red blood cells and release of iron complexes produces a purple color while the addition of neutral red stain colors lysosomes red. F) GS; this neurotransmitter degrading enzyme found in M&uuml;ller cells 13 (green), can be seen extending in both directions with the M&uuml;ller cell bodies in the inner nuclear layer and M&uuml;ller cell endfeet forming the inner limiting membrane. G) NF-200; this heavy chain cytoskeletal protein (green) found in the cell soma and processes, crosslinks with other neurofilaments to maintain the structure of neurons 14,15. Ganglion cells and their axons can be seen in the ganglion cell layer and nerve fiber layer, while axons of horizontal cells located at the outer boarder of the inner nuclear layer in the feline retina, are also highlighted. Counterstaining with DAPI (blue) allows visualization of retinal layers. H) GFAP; this cytoskeletal protein (red) proliferated in M&uuml;ller cells and astrocytes during gliosis 16, can be seen lining the nerve fiber layer with M&uuml;ller cells forming thin extensions through the inner to outer retinal layers. Counterstaining with DAPI (blue) allows for visualization of the retinal layers. Scale bar = 50 &mu;m in all panels. The inner retina is shown at the top of each image; the outer retina is shown at the bottom of each image, excluding Panel E, which shows subscleral reaction.

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Techniques for Processing Eyes Implanted With a Retinal Prosthesis for Localized Histopathological Analysis

Techniques for visualizing retinal cytoarchitecture directly adjacent to individual electrodes within a retinal stimulator.

With  the recent development of retinal prostheses, it is important to develop  reliable techniques for assessing the safety of these devices in ...

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Nanyang Technological University

Research

JoVE Journal

Medicine

19.7K Views.  Bionics Institute. Techniques for visualizing retinal cytoarchitecture directly adjacent to individual electrodes within a retinal stimulator.

Video: Techniques for Processing Eyes Implanted With a Retinal Prosthesis for Localized Histopathological Analysis

Experimental Models for Study of Retinal Pigment Epithelial Physiology and Pathophysiology

We have developed a cell culture procedure that can produce large quantities of confluent monolayers of primary human fetal retinal pigment epithelium (hfRPE) cultures with morphological, physiological and genetic characteristics of native human RPE. These hfRPE cell cultures exhibit heavy pigmentation, and electron microscopy show extensive apical membrane microvilli. The junctional complexes were identified with immunofluorescence labeling of various tight junction proteins. Epithelial polarity and function of these easily reproducible primary cultures closely resemble previously studied mammalian models of native RPE, including human. These results were extended by the development of therapeutic interventions in several animal models of human eye disease. We have focused on strategies for the removal of abnormal fluid accumulation in the retina or subretinal space. The extracellular subretinal space separates the photoreceptor outer segments and the apical membrane of the RPE and is critical for maintenance of retinal attachments and a whole host of RPE/retina interactions.

We provide a reproducible method for culturing confluent monolayers of human fetal retinal pigment epithelial cells (hfRPE) cells that exhibit morphology, physiology, polarity, and protein and gene expression patterns of adult native tissue. This work has been extended to an animal model of several eye diseases.

We have developed a cell culture procedure that can produce large quantities of confluent monolayers of primary human fetal retinal pigment epithelium ...

Neuroscience

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). ...

Techniques for Processing Eyes Implanted with a Retinal Prosthesis for Localized Histopathological Analysis: Part 2 Epiretinal Implants with Retinal Tacks

Retinal prostheses for the treatment of certain forms of blindness are gaining traction in clinical trials around the world with commercial devices currently entering the market. In order to evaluate the safety of these devices, in preclinical studies, reliable techniques are needed. However, the hard metal components utilised in some retinal implants are not compatible with traditional histological processes, particularly in consideration for the delicate nature of the surrounding tissue. Here we describe techniques for assessing the health of the eye directly adjacent to a retinal implant secured epiretinally with a metal tack.
Retinal prostheses feature electrode arrays in contact with eye tissue. The most commonly used location for implantation is the epiretinal location (posterior chamber of the eye), where the implant is secured to the retina with a metal tack that penetrates all the layers of the eye. Previous methods have not been able to assess the proximal ocular tissue with the tack in situ, due to the inability of traditional histological techniques to cut metal objects. Consequently, it has been difficult to assess localized damage, if present, caused by tack insertion.
Therefore, we developed a technique for visualizing the tissue around a retinal tack and implant. We have modified an established technique, used for processing and visualizing hard bony tissue around a cochlear implant, for the soft delicate tissues of the eye. We orientated and embedded the fixed eye tissue, including the implant and retinal tack, in epoxy resin, to stabilise and protect the structure of the sample. Embedded samples were then ground, polished, stained, and imaged under various magnifications at incremental depths through the sample. This technique allowed the reliable assessment of eye tissue integrity and cytoarchitecture adjacent to the metal tack.

Here we describe histological techniques for visualising ocular tissue directly adjacent to a metal epiretinal tack and retinal prosthesis.

Retinal prostheses for the treatment of certain forms of blindness are gaining traction in clinical trials around the world with commercial devices ...

Optimized Protocol for Retinal Wholemount Preparation for Imaging and Immunohistochemistry

Working with delicate tissue can be a complicating factor when performing immunohistochemical assessment. Here, we present a method that utilizes a ring-supported hydrophilized PTFE membrane to provide structural support to both living and fixed tissue during immunohistochemical processing, which allows for the use of a variety of protocols that would otherwise cause damage to the tissue. First, this is demonstrated with bolus loading of fluorescent markers into living retinal tissue. This method allows for quick visualization of targeted structures, while the membrane support maintains tissue integrity during the injection and allows for easy transfer of the preparation for further imaging or processing.
Second, a procedure for antibody staining in tissue fixed with carbodiimide is described. Though paraformaldehyde fixation is more common, carbodiimide fixation provides a superior substrate for the visualization of synaptic proteins. A limitation of carbodiimide is that the resulting fixed tissue is relatively fragile; however, this is overcome with the use of the supporting membrane. Retinal tissue is used to demonstrate these techniques, but they may be applied to any fragile tissue.

The protocol described here is for structural assessment of a wholemount retinal preparation. This includes descriptions of tissue dissection, mounting onto a hydrophilized polytetrafluoroethylene (PTFE) membrane insert, bolus loading with fluorescent markers, and a comparison of fixation with carbodiimide and paraformaldehyde for immunohistochemical analysis of cellular and synaptic components.

Working with delicate tissue can be a complicating factor when performing immunohistochemical assessment. Here, we present a method that utilizes a ...

Biology

Subretinal Transplantation of Human Embryonic Stem Cell-Derived Retinal Tissue in a Feline Large Animal Model

Retinal degenerative (RD) conditions associated with photoreceptor loss such as age-related macular degeneration (AMD), retinitis pigmentosa (RP) and Leber Congenital Amaurosis (LCA) cause progressive and debilitating vision loss. There is an unmet need for therapies that can restore vision once photoreceptors have been lost. Transplantation of human pluripotent stem cell (hPSC)-derived retinal tissue (organoids) into the subretinal space of an eye with advanced RD brings retinal tissue sheets with thousands of healthy mutation-free photoreceptors and has a potential to treat most/all blinding diseases associated with photoreceptor degeneration with one approved protocol. Transplantation of fetal retinal tissue into the subretinal space of animal models and people with advanced RD has been developed successfully but cannot be used as a routine therapy due to ethical concerns and limited tissue supply. Large eye inherited retinal degeneration (IRD) animal models are valuable for developing vision restoration therapies utilizing advanced surgical approaches to transplant retinal cells/tissue into the subretinal space. The similarities in globe size, and photoreceptor distribution (e.g., presence of macula-like region area centralis) and availability of IRD models closely recapitulating human IRD would facilitate rapid translation of a promising therapy to the clinic. Presented here is a surgical technique of transplanting hPSC-derived retinal tissue into the subretinal space of a large animal model allowing assessment of this promising approach in animal models.

Presented here is a surgical technique for transplanting human pluripotent stem cell (hPSC)-derived retinal tissue into the subretinal space of a large animal model.

Retinal degenerative (RD) conditions associated with photoreceptor loss such as age-related macular degeneration (AMD), retinitis pigmentosa (RP) and ...

Optimizing the Setup and Conditions for Ex Vivo Electroretinogram to Study Retina Function in Small and Large Eyes

Measurements of retinal neuronal light responses are critical to investigating the physiology of the healthy retina, determining pathological changes in retinal diseases, and testing therapeutic interventions. The ex vivo electroretinogram (ERG) allows the quantification of contributions from individual cell types in the isolated retina by addition of specific pharmacological agents and evaluation of tissue-intrinsic changes independently of systemic influences. Retinal light responses can be measured using a specialized ex vivo ERG specimen holder and recording setup, modified from existing patch clamp or microelectrode array equipment. Particularly, the study of ON-bipolar cells, but also of photoreceptors, has been hampered by the slow but progressive decline of light responses in the ex vivo ERG over time. Increased perfusion speed and adjustment of the perfusate temperature improve ex vivo retinal function and maximize response amplitude and stability. The ex vivo ERG uniquely allows the study of individual retinal neuronal cell types. In addition, improvements to maximize response amplitudes and stability allow the investigation of light responses in retina samples from large animals, as well as human donor eyes, making the ex vivo ERG a valuable addition to the repertoire of techniques used to investigate retinal function.

Optimizing the Setup and Conditions for Ex Vivo Electroretinogram to Study Retina Function in Small and Large Eyes

Modification of existing multielectrode array or patch clamp equipment makes the ex vivo electroretinogram more widely accessible. Improved methods to record and maintain ex vivo light responses facilitate the study of photoreceptor and ON-bipolar cell function in the healthy retina, animal models of eye diseases, and human donor retinas.

Measurements of retinal neuronal light responses are critical to investigating the physiology of the healthy retina, determining pathological changes in ...

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 ...

Characterization of a Novel Human Organotypic Retinal Culture Technique

Previous human organotypic retinal culture (HORC) models have utilized detached retinas; however, without the structural support conferred by retinal pigment epithelium-choroid (RPE-choroid) and sclera, the integrity of the fragile retina can easily be compromised. The aim of this study was to develop a novel HORC model that contains the retina, RPE-choroid and sclera to maintain retinal integrity when culturing retinal explants.
After cutting circumferentially along the limbus to remove iris and lens, four deep incisions were made to flatten the eyecup. In contrast to previous HORC protocols, a trephine was used to cut through not only the retina but also the RPE-choroid and sclera. The resultant triple-layered explants were cultured for 72 h. Hematoxylin and Eosin staining (H&#38;E) was used to assess anatomical structures and retinal explants were further characterized by immunohistochemistry (IHC) for apoptosis, M&#252;ller cell integrity and retinal inflammation. To confirm the possibility of disease induction, explants were exposed to high glucose (HG) and pro-inflammatory cytokines (Cyt), to mimic diabetic retinopathy (DR). The Luminex magnetic bead assay was used to measure DR-related cytokines released into the culture medium.
H&#38;E staining revealed distinct retinal lamellae and compact nuclei in retinal explants with the underlying RPE-choroid and sclera, while retinas without the underlying structures exhibited reduced thickness and severe nuclei loss. IHC results indicated absence of apoptosis and retinal inflammation as well as preserved M&#252;ller cell integrity. The Luminex&#160;assays showed significantly increased secretion of DR-associated pro-inflammatory cytokines in retinal explants exposed to HG + Cyt relative to baseline levels at 24 h.
We successfully developed and characterized a novel HORC protocol in which retinal integrity was preserved without apoptosis or retinal inflammation. Moreover, the induced secretion of DR-associated pro-inflammatory biomarkers when exposing retinal explants to HG + Cyt suggests that this model could be used for clinically translatable retinal disease studies.

This study aims to develop a novel human organotypic retinal culture (HORC) model that prevents compromising retinal integrity during explant handling. This is achieved by culturing the retina with the overlying vitreous and the underlying retinal pigment&#160;epithelium-choroid (RPE-choroid) and sclera.

Previous human organotypic retinal culture (HORC) models have utilized detached retinas; however, without the structural support conferred by retinal ...

Subretinal Transplantation of Human Embryonic Stem Cell Derived-retinal Pigment Epithelial Cells into a Large-eyed Model of Geographic Atrophy

Geographic atrophy (GA), the late stage of dry age-related macular degeneration is characterized by loss of the retinal pigment epithelial (RPE) layer, which leads to subsequent degeneration of vital retinal structures (e.g., photoreceptors) causing severe vision impairment. Similarly, RPE-loss and decrease in visual acuity is seen in long-term follow up of patients with advanced wet age-related macular degeneration (AMD) receiving intravitreal anti-vascular endothelial growth factor (VEGF) treatment. Therefore, on the one hand, it is fundamental to efficiently derive RPE cells from an unlimited source that could serve as replacement therapy. On the other hand, it is important to assess the behavior and integration of the derived cells in a model of the disease entailing surgical and imaging methods as close as possible to those applied in humans. Here, we provide a detailed protocol based on our previous publications that describes the generation of a preclinical model of GA using the albino rabbit eye, for evaluation of the human embryonic stem cell derived retinal pigment epithelial cells (hESC-RPE) in a clinically relevant setting. Differentiated hESC-RPE are transplanted into naive eyes or eyes with NaIO3-induced GA-like retinal degeneration using a 25 G transvitreal pars plana technique. Evaluation of degenerated and transplanted areas is performed by multimodal high-resolution non-invasive real-time imaging.

Retinal pigment epithelial cells could serve as a cell-replacement therapy for the advanced form of dry age-related macular degeneration. This protocol describes the generation of a large-eyed model of geographic atrophy and the subretinal transplantation of human embryonic stem cell-derived retinal pigment epithelial cells into this model of disease.

Geographic atrophy (GA), the late stage of dry age-related macular degeneration is characterized by loss of the retinal pigment epithelial (RPE) layer, ...

Techniques for Processing Eyes Implanted With a Retinal Prosthesis for Localized Histopathological Analysis

Summary

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Chapters in this video

Experimental Models for Study of Retinal Pigment Epithelial Physiology and Pathophysiology

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

Techniques for Processing Eyes Implanted with a Retinal Prosthesis for Localized Histopathological Analysis: Part 2 Epiretinal Implants with Retinal Tacks

Optimized Protocol for Retinal Wholemount Preparation for Imaging and Immunohistochemistry

Subretinal Transplantation of Human Embryonic Stem Cell-Derived Retinal Tissue in a Feline Large Animal Model

Optimizing the Setup and Conditions for Ex Vivo Electroretinogram to Study Retina Function in Small and Large Eyes

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

Characterization of a Novel Human Organotypic Retinal Culture Technique

Subretinal Transplantation of Human Embryonic Stem Cell Derived-retinal Pigment Epithelial Cells into a Large-eyed Model of Geographic Atrophy