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 border="1">
	<tbody>
		<tr>
			<td>Name of the reagent</td>
			<td>Company</td>
			<td>Catalogue number</td>
			<td>Comments (optional)</td>
		</tr>
		<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></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&#39;,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&#176; yellow</td>
			<td>Grale Scientific</td>
			<td>SF41299</td>
			<td></td>
		</tr>
		<tr>
			<td>FLEX IHC Microscope Slides</td>
			<td>DAKO</td>
			<td>K802021-2</td>
			<td></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></td>
			<td></td>
			<td></td>
			<td>
			Non-Standard Solution Preparation for Special Stains
			1% Aqueous Cresyl Violet Stock Solution
			<ul>
				<li>Cresyl violet 1 g</li>
				<li>Distilled water 100 ml</li>
			</ul>
			Cresyl Violet Acetate Working Solution
			<ul>
				<li>1% aqueous cresyl violet stock solution 9 ml</li>
				<li>Distilled water 51 ml</li>
				<li>10% acetic acid 0.5 ml</li>
			</ul>
			Biebrich Scarlet-Acid Fuchsin Solution
			<ul>
				<li>1% Beihrich scarlet 90 ml</li>
				<li>1% acid fuchsin 10 ml</li>
				<li>Glacial acetic acid 1 ml</li>
			</ul>
			Phosphomolybdic-Phosphtungstic Acid Solution
			<ul>
				<li>Phosphomolybdic acid 2.5 g</li>
				<li>Phosphotungstic acid 2.5 g</li>
				<li>Distilled water 100 ml</li>
			</ul>
			Aniline Blue Solution
			<ul>
				<li>Aniline blue 2.5 g</li>
				<li>Glacial acetic acid 2 g</li>
				<li>Distilled water 100 ml</li>
			</ul>
			Solutions for Immunohistochemistry
			Wash Buffer Solution
			<ul>
				<li>0.1% Triton X-100 in Phosphate buffered saline (PBS)</li>
			</ul>
			Serum Block Solution
			<ul>
				<li>10% normal goat serum/0.1% Triton X-100 in PBS</li>
			</ul>
			SUPPLEMENTARY MATERIAL
			Luxol Fast Blue
			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.
			Periodic Acid Schiff
			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.
			Masson&#8217;s Trichrome Blue
			The principle of the Masson&#39;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&#39;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.
			Glial Fibrillary Acidic Protein (GFAP) Antibody
			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&#252;ller cells in the retina), the antigen is made form 428 amino acids and has a molecular weight of 49512 Da.
			Neurofilament-200 antibody
			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.
			Glutamine Synthetase (GS) antibody
			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&#252;ller cells in the retina), the antigen has 373 amino acids and a molecular weight of 42,064 Da.
			</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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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

Performing and Processing FNA of Anterior Fat Pad for Amyloid

Historically, heart, liver, and kidney biopsies were performed to demonstrate amyloid deposits in amyloidosis. Since the clinical presentation of this disease is so variable and non-specific, the associated risks of these biopsies are too great for the diagnostic yield. Other sites that have a lower biopsy risk, such as skin or gingival, are also relatively invasive and expensive. In addition, these biopsies may not always have sufficient amyloid deposits to establish a diagnosis. Fat pad aspiration has demonstrated good clinical correlation with low cost and minimal morbidity. However, there are no standardized protocols for performing this procedure or processing the aspirated specimen, which leads to variable and nonreproducible results. The most frequently utilized modality for detecting amyloid in tissue is an apple-green birefringence on Congo red stained sections using a polarizing microscope. This technique requires cell block preparation of aspirated material. Unfortunately, patients presenting in early stage of amyloidosis have minimal amounts of amyloid which greatly reduces the sensitivity of Congo red stained cell block sections of fat pad aspirates. Therefore, ultrastructural evaluation of fat pad aspirates by electron microscopy should be utilized, given its increased sensitivity for amyloid detection. This article demonstrates a simple and reproducible procedure for performing anterior fat pad aspiration for the detection of amyloid utilizing both Congo red staining of cell block sections and electron microscopy for ultrastructural identification.

Fat pad aspiration is a preferred, minimally invasive, and low cost approach as compared to other methods to detect amyloid for diagnosis of systemic amyloidosis. This video article demonstrates a procedural outline for performing fat pad aspiration with appropriate processing of the specimen for the optimal diagnostic outcome.

Historically, heart, liver, and kidney biopsies were performed to demonstrate amyloid deposits in amyloidosis.  Since the clinical presentation of this ...

How to Measure Cortical Folding from MR Images: a Step-by-Step Tutorial to Compute Local Gyrification Index

Cortical folding (gyrification) is determined during the first months of life, so that adverse events occurring during this period leave traces that will be identifiable at any age. As recently reviewed by Mangin and colleagues2, several methods exist to quantify different characteristics of gyrification. For instance, sulcal morphometry can be used to measure shape descriptors such as the depth, length or indices of inter-hemispheric asymmetry3. These geometrical properties have the advantage of being easy to interpret. However, sulcal morphometry tightly relies on the accurate identification of a given set of sulci and hence provides a fragmented description of gyrification. A more fine-grained quantification of gyrification can be achieved with curvature-based measurements, where smoothed absolute mean curvature is typically computed at thousands of points over the cortical surface4. The curvature is however not straightforward to comprehend, as it remains unclear if there is any direct relationship between the curvedness and a biologically meaningful correlate such as cortical volume or surface. To address the diverse issues raised by the measurement of cortical folding, we previously developed an algorithm to quantify local gyrification with an exquisite spatial resolution and of simple interpretation. Our method is inspired of the Gyrification Index5, a method originally used in comparative neuroanatomy to evaluate the cortical folding differences across species. In our implementation, which we name local Gyrification Index (lGI1), we measure the amount of cortex buried within the sulcal folds as compared with the amount of visible cortex in circular regions of interest. Given that the cortex grows primarily through radial expansion6, our method was specifically designed to identify early defects of cortical development.
In this article, we detail the computation of local Gyrification Index, which is now freely distributed as a part of the FreeSurfer Software (<a href="http://surfer.nmr.mgh.harvard.edu/">http://surfer.nmr.mgh.harvard.edu/</a>, Martinos Center for Biomedical Imaging, Massachusetts General Hospital). FreeSurfer provides a set of automated reconstruction tools of the brain's cortical surface from structural MRI data. The cortical surface extracted in the native space of the images with sub-millimeter accuracy is then further used for the creation of an outer surface, which will serve as a basis for the lGI calculation. A circular region of interest is then delineated on the outer surface, and its corresponding region of interest on the cortical surface is identified using a matching algorithm as described in our validation study1. This process is repeatedly iterated with largely overlapping regions of interest, resulting in cortical maps of gyrification for subsequent statistical comparisons (Fig. 1). Of note, another measurement of local gyrification with a similar inspiration was proposed by Toro and colleagues7, where the folding index at each point is computed as the ratio of the cortical area contained in a sphere divided by the area of a disc with the same radius. The two implementations differ in that the one by Toro et al. is based on Euclidian distances and thus considers discontinuous patches of cortical area, whereas ours uses a strict geodesic algorithm and include only the continuous patch of cortical area opening at the brain surface in a circular region of interest.

Measuring gyrification (cortical folding) at any age represents a window into early brain development. Hence, we previously developed an algorithm to measure local gyrification at thousands of points over the hemisphere1. In this paper, we detail the computation of this local gyrification index.

Cortical folding (gyrification) is determined during the first months of life, so that adverse events occurring during this period leave traces that will ...

Analytical Techniques for Assaying Nitric Oxide Bioactivity

Nitric oxide (NO) is a diatomic free radical that is extremely short lived in biological systems (less than 1 second in circulating blood)1. NO may be considered one of the most important signaling molecules produced in our body, regulating essential functions including but not limited to regulation of blood pressure, immune response and neural communication. Therefore its accurate detection and quantification in biological matrices is critical to understanding the role of NO in health and disease. With such a short physiological half life of NO, alternative strategies for the detection of reaction products of NO biochemistry have been developed. The quantification of relevant NO metabolites in multiple biological compartments provides valuable information with regards to in vivo NO production, bioavailability and metabolism. Simply sampling a single compartment such as blood or plasma may not always provide an accurate assessment of whole body NO status, particularly in tissues. The ability to compare blood with select tissues in experimental animals will help bridge the gap between basic science and clinical medicine as far as diagnostic and prognostic utility of NO biomarkers in health and disease. Therefore, extrapolation of plasma or blood NO status to specific tissues of interest is no longer a valid approach. As a result, methods continue to be developed and validated which allow the detection and quantification of NO and NO-related products/metabolites in multiple compartments of experimental animals in vivo. The established paradigm of NO biochemistry from production by NO synthases to activation of soluble guanylyl cyclase (sGC) to eventual oxidation to nitrite (NO2-) and nitrate (NO3-) may only represent part of NO's effects in vivo. The interaction of NO and NO-derived metabolites with protein thiols, secondary amines, and metals to form S-nitrosothiols (RSNOs), N-nitrosamines (RNNOs), and nitrosyl-heme respectively represent cGMP-independent effects of NO and are likely just as important physiologically as activation of sGC by NO. A true understanding of NO in physiology is derived from in vivo experiments sampling multiple compartments simultaneously. Nitric oxide (NO) methodology is a complex and often confusing science and the focus of many debates and discussion concerning NO biochemistry. The elucidation of new mechanisms and signaling pathways involving NO hinges on our ability to specifically, selectively and sensitively detect and quantify NO and all relevant NO products and metabolites in complex biological matrices. Here, we present a method for the rapid and sensitive analysis of nitrite and nitrate by HPLC as well as detection of free NO in biological samples using in vitro ozone based chemiluminescence with chemical derivitazation to determine molecular source of NO as well as ex vivo with organ bath myography.

The endogenous production of nitric oxide (NO) regulates a wide variety of biological functions. It is becoming increasingly clear that disruption or dysregulation of NO based signaling is involved in many human diseases. Methods to quantify relevant NO metabolites may provide novel diagnostic or prognostic biomarkers for human disease.

Nitric oxide (NO) is a diatomic free radical that is extremely short lived in biological systems (less than 1 second in circulating blood)1. NO may be ...

Processing of Human Reduction Mammoplasty and Mastectomy Tissues for Cell Culture

Experimental examination of normal human mammary epithelial cell (HMEC) behavior, and how normal cells acquire abnormal properties, can be facilitated by in vitro culture systems that more accurately model in vivo biology. The use of human derived material for studying cellular differentiation, aging, senescence, and immortalization is particularly advantageous given the many significant molecular differences in these properties between human and commonly utilized rodent cells1-2. Mammary cells present a convenient model system because large quantities of normal and abnormal tissues are available due to the frequency of reduction mammoplasty and mastectomy surgeries.
The mammary gland consists of a complex admixture of many distinct cell types, e.g., epithelial, adipose, mesenchymal, endothelial. The epithelial cells are responsible for the differentiated mammary function of lactation, and are also the origin of the vast majority of human breast cancers. We have developed methods to process mammary gland surgical discard tissues into pure epithelial components as well as mesenchymal cells3. The processed material can be stored frozen indefinitely, or initiated into primary culture. Surgical discard material is transported to the laboratory and manually dissected to enrich for epithelial containing tissue. Subsequent digestion of the dissected tissue using collagenase and hyaluronidase strips stromal material from the epithelia at the basement membrane. The resulting small pieces of the epithelial tree (organoids) can be separated from the digested stroma by sequential filtration on membranes of fixed pore size. Depending upon pore size, fractions can be obtained consisting of larger ductal/alveolar pieces, smaller alveolar clusters, or stromal cells. We have observed superior growth when cultures are initiated as organoids rather than as dissociated single cells. Placement of organoids in culture using low-stress inducing media supports long-term growth of normal HMEC with markers of multiple lineage types (myoepithelial, luminal, progenitor)4-5. Sufficient numbers of cells can be obtained from one individual's tissue to allow extensive experimental examination using standardized cell batches, as well as interrogation using high throughput modalities.
Cultured HMEC have been employed in a wide variety of studies examining the normal processes governing growth, differentiation, aging, and senescence, and how these normal processes are altered during immortal and malignant transformation4-15,16. The effects of growth in the presence of extracellular matrix material, other cell types, and/or 3D culture can be compared with growth on plastic5,15. Cultured HMEC, starting with normal cells, provide an experimentally tractable system to examine factors that may propel or prevent human aging and carcinogenesis.

A method to process human mammary surgical discard material is described. Processed tissue, in the form of organoids, can be stored frozen indefinitely or placed in culture for long-term growth. This method enables experimental examination of normal human epithelial cell biology, and the effects of exogenous perturbations.

Experimental  examination of normal human mammary epithelial cell (HMEC) behavior, and how  normal cells acquire abnormal properties, can be facilitated ...

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 ...

Quantitative Fundus Autofluorescence for the Evaluation of Retinal Diseases

The retinal pigment epithelium (RPE) is juxtaposed to the overlying sensory retina, and supports the function of the visual system. Among the tasks performed by the RPE are phagocytosis and processing of outer photoreceptor segments through lysosome-derived organelles. These degradation products, stored and referred to as lipofuscin granules, are composed partially of bisretinoids, which have broad fluorescence absorption and emission spectra that can be detected clinically as fundus autofluorescence with confocal scanning laser ophthalmoscopy (cSLO). Lipofuscin accumulation is associated with increasing age, but is also found in various patterns in both acquired and inherited degenerative diseases of the retina. Thus, studying its pattern of accumulation and correlating such patterns with changes in the overlying sensory retina are essential to understanding the pathophysiology and progression of retinal disease. Here, we describe a technique employed by our lab and others that uses cSLO in order to quantify the level of RPE lipofuscin in both healthy and diseased eyes.

The retinal pigment epithelium (RPE) supports the sensory retina through recycling visual cycle byproducts, which accumulate as lipofuscin. These products are autofluorescent and can be qualitatively imaged in vivo. Here, we describe a method to quantitatively image RPE lipofuscin using confocal scanning laser ophthalmoscopy.

The retinal pigment epithelium (RPE) is juxtaposed to the overlying sensory retina, and supports the function of the visual system. Among the tasks ...

A Step by Step Protocol for Subretinal Surgery in Rabbits

Age related macular degeneration (AMD), retinitis pigmentosa, and other RPE related diseases are the most common causes for irreversible loss of vision in adults in industrially developed countries. RPE transplantation appears to be a promising therapy, as it may replace dysfunctional RPE, restore its function, and thereby vision.
Here we describe a method for transplanting a cultured RPE monolayer on a scaffold into the subretinal space (SRS) of rabbits. After vitrectomy xenotransplants were delivered into the SRS using a custom made shooter consisting of a 20-gauge metallic nozzle with a polytetrafluoroethylene (PTFE) coated plunger. The current technique evolved in over 150 rabbit surgeries over 6 years. Post-operative follow-up can be obtained using non-invasive and repetitive in vivo imaging such as spectral domain optical coherence tomography (SD-OCT) followed by perfusion-fixed histology.
The method has well-defined steps for easy learning and high success rate. Rabbits are considered a large eye animal model useful in preclinical studies for clinical translation. In this context rabbits are a cost-efficient and perhaps convenient alternative to other large eye animal models.

Retinal pigment epithelium (RPE) replacement strategies and gene-based therapy are considered for several retinal degenerative conditions. For clinical translation, large eye animal models are required to study surgical techniques applicable in patients. Here we present a rabbit model for subretinal surgery geared towards RPE transplantation, which is versatile and cost-efficient.

Age related macular degeneration (AMD), retinitis pigmentosa, and other RPE related diseases are the most common causes for irreversible loss of vision in ...

An Acute Retinal Model for Evaluating Blood Retinal Barrier Breach and Potential Drugs for Treatment

A low-cost, easy-to-use and powerful model system is established to evaluate potential treatments that could ameliorate blood retinal barrier breach. An inflammatory factor, histamine, is demonstrated to compromise vessel integrity in the cultured retina through positive staining of IgG outside of the blood vessels. The effects of histamine itself and those of candidate drugs for potential treatments, such as lipoxin A4, are assessed using three parameters: blood vessel leakage via IgG immunostaining, activation of M&#252;ller cells via GFAP staining and change in neuronal dendrites through staining for MAP2. Furthermore, the layered organization of the retina allows a detailed analysis of the processes of M&#252;ller and ganglion cells, such as changes in width and continuity. While the data presented is with swine retinal culture, the system is applicable to multiple species. Thus, the model provides a reliable tool to investigate the early effects of compromised retinal vessel integrity on different cell types and also to evaluate potential drug candidates for treatment.

A low-cost, easy-to-use and powerful system is established to evaluate potential treatments that could ameliorate blood retinal barrier breach induced by histamine. Blood vessel leakage, M&#252;ller cell activation and the continuity of neuronal processes are utilized to assess the damage response and its reversal with a potential drug, lipoxin A4.

A low-cost, easy-to-use and powerful model system is established to evaluate potential treatments that could ameliorate blood retinal barrier breach. An ...

Methodology for Sputum Induction and Laboratory Processing

The technique of sputum induction and processing is a recognized non-invasive method allowing the collection and analysis of cells from the airways, which is interesting in various respiratory diseases like asthma, chronic obstructive pulmonary disease (COPD), chronic cough, or idiopathic pulmonary fibrosis. This technique is well tolerated, safe and non-invasive, but is currently limited to research services and specialized centers in clinical practice because it is technically demanding, time-consuming, and requires trained staff. The success rate of sputum induction and analysis is about 80%.
Here, we describe the induction and laboratory processing of sputum samples. Sputum is induced by inhalation of hypertonic or isotonic saline with salbutamol. For the processing, we use the whole sputum technique. Dithiothreitol (DTT) is used to allow mucolysis of sputum samples. The primary aim of sputum processing is to obtain a differential cell count to study the cell types present in the airway lumen. Additional analyses may also be performed on sputum supernatant and sputum cells, which may allow further investigation into inflammatory processes and immune mechanisms. Examples include studying mediators in sputum supernatant and performing a large spectrum of analysis on sputum cells such as flow cytometry, genomics, or proteomics.
Finally, representative results of sputum analysis in healthy controls, asthmatics, and COPD patients are presented.

Here we describe the technique of sputum induction. This protocol also explains the sputum processing to perform a differential cell count and to collect sputum supernatant and cells for further analysis.

The technique of sputum induction and processing is a recognized non-invasive method allowing the collection and analysis of cells from the airways, which ...

Studying Diabetes Through the Eyes of a Fish: Microdissection, Visualization, and Analysis of the Adult tg(fli:EGFP) Zebrafish Retinal Vasculature

Diabetic retinopathy is the leading cause of blindness among middle-aged adults. The rising prevalence of diabetes worldwide will make the prevention of diabetic microvascular complications one of the key research fields of the next decades. Specialized, targeted therapy and novel therapeutic drugs are needed to manage the increasing number of patients at risk of vision-loss. The zebrafish is an established animal model for developmental research questions with increasing relevance for modeling metabolic multifactorial disease processes. The advantages of the species allow for optimal visualization and high throughput drug screening approaches, combined with the strong ability to knock out genes of interest. Here, we describe a protocol which will allow easy analysis of the adult tg(fli:EGFP) zebrafish retinal vasculature as a fast read-out in settings of long-term vascular pathologies linked to neoangiogenesis or vessel damage. This is achieved via dissection of the zebrafish retina and whole-mounting of the tissue. Visualization of the exposed vessels is then achieved via confocal microscopy of the green EGFP reporter expressed in the adult retinal vasculature. Correct handling of the tissue will lead to better outcomes and less internal vessel breakage to assure the visualization of the unaltered vascular structure. The method can be utilized in zebrafish models of retinal vasculopathy linked to changes in the vessel architecture as well as neoangiogenesis.

Studying Diabetes Through the Eyes of a Fish: Microdissection, Visualization, and Analysis of the Adult tgfli:EGFP Zebrafish Retinal Vasculature

Here, we discuss a method protocol which will allow an easy analysis of the adult tg(fli:EGFP) zebrafish retinal vasculature as a fast read-out in settings of long-term vascular pathologies linked to neoangiogenesis and structural changes.

Diabetic retinopathy is the leading cause of blindness among middle-aged adults. The rising prevalence of diabetes worldwide will make the prevention of ...