Ex Vivo OCT-Based Multimodal Imaging of Human Donor Eyes for Research into Age-Related Macular Degeneration

Podsumowanie

Laboratory assays can leverage prognostic value from the longitudinal optical coherence tomography (OCT)-based multimodal imaging of age-related macular degeneration (AMD). Human donor eyes with and without AMD are imaged using OCT, color, near-infrared reflectance scanning laser ophthalmoscopy, and autofluorescence at two excitation wavelengths prior to tissue sectioning.

Streszczenie

A progression sequence for age-related macular degeneration (AMD) learned from optical coherence tomography (OCT)-based multimodal (MMI) clinical imaging could add prognostic value to laboratory findings. In this work, ex vivo OCT and MMI were applied to human donor eyes prior to retinal tissue sectioning. The eyes were recovered from non-diabetic white donors aged ≥80 years old, with a death-to-preservation time (DtoP) of ≤6 h. The globes were recovered on-site, scored with an 18 mm trephine to facilitate cornea removal, and immersed in buffered 4% paraformaldehyde. Color fundus images were acquired after anterior segment removal with a dissecting scope and an SLR camera using trans-, epi-, and flash illumination at three magnifications. The globes were placed in a buffer within a custom-designed chamber with a 60 diopter lens. They were imaged with spectral domain OCT (30° macula cube, 30 µm spacing, averaging = 25), near-infrared reflectance, 488 nm autofluorescence, and 787 nm autofluorescence. The AMD eyes showed a change in the retinal pigment epithelium (RPE), with drusen or subretinal drusenoid deposits (SDDs), with or without neovascularization, and without evidence of other causes. Between June 2016 and September 2017, 94 right eyes and 90 left eyes were recovered (DtoP: 3.9 ± 1.0 h). Of the 184 eyes, 40.2% had AMD, including early intermediate (22.8%), atrophic (7.6%), and neovascular (9.8%) AMD, and 39.7% had unremarkable maculas. Drusen, SDDs, hyper-reflective foci, atrophy, and fibrovascular scars were identified using OCT. Artifacts included tissue opacification, detachments (bacillary, retinal, RPE, choroidal), foveal cystic change, an undulating RPE, and mechanical damage. To guide the cryo-sectioning, OCT volumes were used to find the fovea and optic nerve head landmarks and specific pathologies. The ex vivo volumes were registered with the in vivo volumes by selecting the reference function for eye tracking. The ex vivo visibility of the pathology seen in vivo depends on the preservation quality. Within 16 months, 75 rapid DtoP donor eyes at all stages of AMD were recovered and staged using clinical MMI methods.

Wprowadzenie

Fifteen years of managing neovascular age-related macular degeneration (AMD) with anti-VEGF therapy under the guidance of optical coherence tomography (OCT) has offered new insights into the progression sequence and microarchitecture of this prevalent cause of vision loss. A key recognition is that AMD is a three-dimensional disease involving the neurosensory retina, retinal pigment epithelium (RPE), and choroid. As a result of the OCT imaging of trial patients and the fellow eyes of treated clinic patients, the features of pathology beyond those seen by color fundus photography, a clinical standard for decades, are now recognized. These include intraretinal neovascularization (type 3 macular neovascularization¹, formerly angiomatous proliferation), subretinal drusenoid deposits (SDDs, also called reticular pseudodrusen)², multiple pathways of RPE fate^3,4, and intensely gliotic Müller cells in atrophy^5,6.

Model systems lacking maculas (cells and animals) recreate some slices of this complex disease^7,8,9. Further success in ameliorating the burden of AMD could come from the discovery and exploration of primary pathology in human eyes, understanding the unique cellular composition of the macula, followed by translation to model systems. This report portrays a three decade collaboration between an academic research laboratory and an eye bank. The goals of the tissue characterization methods described herein are two-fold: 1) to inform evolving diagnostic technology by demonstrating the basis of fundus appearance and imaging signal sources with microscopy, and 2) to classify AMD specimens for targeted (immunohistochemistry) and untargeted molecular discovery techniques (imaging mass spectrometry, IMS, and spatial transcriptomics) that preserve the cone-only fovea and rod-rich para- and perifovea. Such studies could accelerate the translation to clinical OCT, for which a progression sequence and longitudinal follow-up are possible through eye-tracking. This technology, which is designed to monitor treatment effects, registers scans from one clinic visit to the next using retinal vessels. Linking eye-tracked OCT to laboratory results obtained with destructive techniques could provide a new level of prognostic value to molecular findings.

In 1993, the research laboratory captured color photographs of postmortem fundus on film¹⁰. This effort was inspired by the superb photomicroscopy and histology of the human peripheral retina by Foos and colleagues^11,12,13 and the extensive AMD clinicopathologic correlations by Sarks et al.^14,15. Starting in 2009, ex vivo multimodal imaging (MMI) anchored on spectral domain OCT was adopted. This transition was inspired by the similar efforts of others^16,17 and especially by the realization that so much of the ultrastructure described by the Sarks was available in three dimensions, over time, in the clinic^18,19. The goal was to acquire eyes with attached maculas in a reasonable time frame for well-powered studies of cellular-level phenotypes in the retina, RPE, and choroid. The intent was to move beyond "per eye" statistics to "per lesion type," a standard influenced by the "vulnerable plaque" concepts from cardiovascular disease^20,21.

The protocol in this report reflects experience with nearly 400 pairs of donor eyes accessioned in several streams. In 2011-2014, the Project MACULA website of AMD histopathology was created, which includes layer thicknesses and annotations from 142 archived specimens. These eyes were preserved from 1996-2012 in a glutaraldehyde-paraformaldehyde fixative for high-resolution epoxy-resin histology and electron microscopy. All the fundi had been photographed in color when received and were reimaged by OCT just prior to histology. An eye holder originally designed for optic nerve studies²² was used to accommodate an 8 mm diameter full-thickness tissue punch centered on the fovea. OCT B-scans through the foveal center and a site 2 mm superior, corresponding to histology at the same levels, were uploaded to the website, plus a color fundus photograph. The choice of the OCT planes was dictated by the prominence of AMD pathology under the fovea²³ and the prominence of SDDs in rod-rich areas superior to the fovea^24,25.

Starting in 2013, eyes imaged with OCT-anchored MMI during life were available for direct clinicopathologic correlations. Most (7 of 10 donors) involved patients at a retina referral practice (author: K.B.F.), which offered an advanced directive registry for patients interested in donating their eyes after death for research purposes. The eyes were recovered and preserved by the local eye bank, transferred to the laboratory, and prepared in the same way as the Project MACULA eyes. Pre-mortem clinical OCT volumes were seamlessly read in the laboratory, thus aligning the pathology features seen during life with the features seen under the microscope²⁶.

Starting in 2014, prospective eye collection began by screening for AMD in donor eyes without a clinical history but preserved during a defined time limit (6 h). For this purpose, the eye holder was modified to accommodate a whole globe. This reduced the chance of detachment around the cut edges of the previously used 8 mm punch. The eyes were preserved in 4% buffered paraformaldehyde for immunohistochemistry and transferred to 1% the next day for long-term storage. In 2016-2017 (pre-pandemic), 184 eyes from 90 donors were recovered. The statistics and images in this report are generated from this series. During the pandemic era (2020 lockdowns and aftermath), prospective collections for transcriptomics and IMS collaborations continued at a reduced pace, essentially using the 2014 methods.

Other methods for donor eye assessment are available. The Minnesota Grading System (MGS)^27,28 is based on the AREDS clinical system for color fundus photography²⁹. The limitations of this method include the combining of atrophic and neovascular AMD into one stage of "late AMD". Further, the MGS entails the removal of the neurosensory retina before the photo-documentation of the RPE choroid. This step dislodges SDDs to varying degrees^30,31 and removes the spatial correspondence of the outer retina and its support system. Thus, efforts to link metabolic demand and signaling from the retina to pathology in the RPE-choroid may be impeded. The Utah System implemented MMI using ex vivo color photography and OCT to categorize eyes destined for dissection into regions for RNA and protein extractions³². Although preferable to whole eyecup extractions, the 3 mm diameter area at the highest risk for AMD progression^33,34 represents only 25% of a 6 mm diameter fovea-centered punch. Thus, techniques that can localize findings in reference to the fovea, such as serial sectioning for immunohistochemistry, are advantageous.

Protokół

The institutional review board at the University of Alabama at Birmingham approved the laboratory studies, which adhered to Good Laboratory Practices and Biosafety Level 2/2+. All US eye banks conform to the 2006 Uniform Anatomical Gifts Act and US Food and Drug Administration. Most US eye banks, including Advancing Sight Network, conform to the medical standards of the Eye Bank Association of America.

The Table of Materials lists the supplies and equipment. Supplementary Material 1 provides an overview of the dissection, color fundus photography, and OCT-based MMI. Supplementary Material 2 provides details of the OCT-based MMI.

1. Criteria for tissue collection

1. To maximize the yield of AMD eyes in a screen of undocumented donors, set the following criteria for acceptable donors: age ≥80 years, white, non-diabetic, and ≤6 h death-to-preservation (DtoP).
   NOTE: DtoP is defined as the time between death and when the eye is placed in a provided preservative, either in the hospital or in the laboratory.

2. Preservation medium and other preparation (laboratory)

1. Make 4% phosphate-buffered paraformaldehyde from 20% stock (purchased). Prepare 1 L by adding 200 mL of 20% paraformaldehyde (dilution factor of 5) to 800 mL of 0.1 M Sorenson's phosphate buffer. Test and adjust to ensure the pH is 7.2, if necessary. Store at 4 ˚C.
2. Dispense 30 mL of 4% phosphate-buffered paraformaldehyde into 40 mL jars.
3. Stock labeled 40 mL containers of preservative at the eye bank so that the tissue can be recovered at any time and on any day.
4. For the storage of preserved eyes, make 1% paraformaldehyde from the 4% solution. Prepare 1 L by adding 250 mL of 4% paraformaldehyde (dilution factor of 4) to 750 mL of 0.1 M Sorenson's phosphate buffer. Test and adjust the pH if necessary. Store at 4 ˚C.
   1. Prepare 1 L of a 0.1 M solution from the 0.2 M solution of Sorenson's phosphate buffer, pH 7.2, by combining 500 mL of distilled water and 500 mL of Sorenson's buffer.
   2. Adjust the pH drop by drop using 1 N hydrochloric acid or 1 N sodium hydroxide. Store at 4 ˚C.
5. Create a holder to stabilize the eyes during dissection. Fill a Petri dish with dental wax heated until liquid. When it is slightly cool, make a hemispheric impression in it with a large ball bearing, and then freeze the dish to facilitate the removal of the ball bearing.

3. Eye bank methods

1. To ensure the rapid recovery of research tissues, recover all the tissues rapidly, including those intended for transplantation.
2. Receive death referrals, as required by law, within 1 h of death, and track each donor with a referral sequence number that follows the tissue.
3. To find cases with potential clinical documentation, ask AMD- and eye disease-specific questions in the research Donor Risk Assessment Interview.
4. To minimize the travel time, recover whole globes (as distinct from only corneas for transplantation) within a compact area (i.e., city and adjacent suburban county).
5. Bring two jars of preservative (buffered 4% paraformaldehyde) to the decedent's hospital room for tissue recovery (as opposed to waiting for the body to be moved to a morgue).
6. Before and during recovery, communicate with the investigators to confirm the delivery and timing.
7. Recover the globes on-site in the hospital, open them by consistent handling methods, and immerse the opened eye in preservative (Figure 1).
   1. Hold the excised donor eye in place using a sheath of gauze stabilized by a hemostat.
   2. To facilitate the removal of the cornea with a 2 mm wide rim of sclera, use an 18 mm diameter trephine to score the globe.
   3. To free the cornea with an accompanying rim of sclera, cut along the scored circle using spring-loaded scissors with curved tips while stabilizing the globe with the hemostat-clipped gauze.
   4. Lift the cornea off the sclera, exposing the iris and ciliary body.
   5. To facilitate the penetration of the preservative into the vitreous chamber, make a 2 mm long slit in the iris perpendicular to the pupillary margin. Place the eyes into specimen jars with 30 mL of preservative at 4 °C, and transfer to the laboratory on wet ice.
8. Transmit de-identified donor information electronically from the eye bank to the research laboratory database.
   NOTE: The database maintains the referral number, the eye bank tissue number, and the laboratory ID number for tracking, plus other relevant information.

4. Tissue preparation for ex vivo color fundus photography

1. Use two stereo microscopes, one for dissection and one for color fundus photography.
   NOTE: For transillumination to visualize pigmentary changes, use a base for dark-field microscopy.
2. Remove the anterior segment remnants (iris and lens). To stabilize the eye during dissection, place it in the Petri dish filled with wax (Supplementary Material 1, slides 7-8). To prevent the ciliary body and attached retina from collapsing into the vitreous cavity, minimize the perturbation of the ring of thick vitreous attached to the ciliary body (vitreous base).
3. Mark the superior pole for orientation. Place the globe anterior side down in the dish. Find the insertion tendons of the superior rectus and superior oblique muscles. Using a wooden applicator, sparingly apply a marking ink in a 10 mm line in an anterior to posterior direction (i.e., perpendicular to the tendinous insertion of the superior rectus muscle).
4. Before photography, fill the fundus with cold Sorensen's phosphate buffer.
5. Insert a 1 mm ruby bead into the fundus as an internal scale bar to appear in each image²⁷.
6. Acquire color images with a single-lens reflex camera mounted to a stereo microscope fitted with a ring flash. Use trans-, epi-, and flash illumination at each of three standard magnifications to capture images intended to highlight specific areas (Supplementary Material 1, slide 11): 1) the fundus to the equator, 2) the posterior pole (vascular arcades, optic nerve head, fovea), and 3) the fovea within the macula lutea (yellow spot).

5. Preparation for ex vivo color fundus photography

1. Turn the camera and monitor on. Plug in the remote shutter, and release the actuator in the high-definition multimedia interface (HDMI) camera/television (TV) monitor and display cable.
2. Set the camera settings to manual ISO function and the mirror lock-up position (locked in place to reduce vibration).
   NOTE: Refer to the manufacturer's user manual for the settings on the camera used. Learn from the camera display the over/underexposure settings relative to the histogram and exposure readouts.
3. Arrange two light sources, each with two flexible light guides positioned perpendicular to each other, for illumination in the four cardinal directions on the microscope stage (Supplementary Material 1, page 10).
4. Turn on the epi-illumination light sources to full power.
   NOTE: It is helpful but not necessary to have a black felt shroud around the stage to limit light/flash scatter for the photographer.
5. At the dissecting microscope, use forceps to insert the posterior pole into a 30 mL quartz crucible filled with buffer. Allow the tissue to sink to the bottom. Insert a bracing, such as a tissue sponge, between the eye and the wall of the crucible to prevent movement. Insert the 1 mm ruby bead into the posterior pole.
   NOTE: The bead may fall into the optic nerve cup.
6. Carefully place the globe in the crucible onto the stereo microscope stage, and observe the interior ocular fundus through the microscope eyepieces. Using the lowest magnification, orient the eye by identifying the tissue mark at the 12 o'clock position, the optic nerve head (ONH), and the fovea 5° below the ONH. Rotate the eye so that the fovea falls below a line through the ONH by 5°.
   NOTE: If it is a right eye, the ONH is to the right of the fovea, as seen through the oculars of the microscope. If it is a left eye, the ONH is to the left of the fovea.
7. Turn on the remote monitor viewing from the camera. Ensure that the microscope beam-splitter slider is set to observe through the photo port and not through the port for the oculars. Depending on the light and mirror path, be prepared to rotate the tissue 180˚ on the stage for proper orientation.

6. Image acquisition using ex vivo color fundus photography

1. With the epi-illumination turned on, set the magnification so that the entire 18 mm trephine incision occupies the entire field of view. Increase the magnification so that it is possible to focus on the bottom of the foveal pit. Reduce the magnification to the previous setting.
2. Adjust the ISO settings in the range of 1,600-3,000 so that exposure times fall in the center range of the meter on the camera.
3. Press the remote shutter trigger. Listen for the mirror to lock in the up position. Press the trigger again for exposure.
4. Observe the image on the monitor, with the preset metadata showing red, green, blue (RGB), and a color histogram for the correct exposure. If the exposure is acceptable, proceed; if not, then delete, re-evaluate the parameters, and re-image.
5. Turn off the gooseneck lamps for epi-illumination to highlight drusen, and turn on the flash, with a 1/4 s exposure, a camera shutter speed of 1/250 s, and an ISO of 100-320. Set the flash speed to the default, or modify it during the initial setup of the camera. Acquire an image, and check the histogram for proper exposure.
6. Turn off the flash, and turn on the trans-illumination light source. Reset the ISO to above 5,000, and ensure the exposure time does not go below 1/30 s due to potential vibration in the photographic system. Acquire an image, and check the histogram for proper exposure.
7. Increase the magnification to view both the ONH and fovea in the field of view. Turn on the epi-illumination lamps. Set the ISO range to 1,600-3,000. Ensure that the exposure times fall in the center range of the camera.
8. Turn off the epi-illumination lamps, and turn on the flash, with a 1/4 s exposure, the preset camera shutter speed set to 1/250 s, and the ISO set to 400-800. Acquire an image, and check the histogram for proper exposure.
9. Turn off the flash, and turn on the trans-illumination using the dark-field base. Reset the ISO to above 5,000. Ensure that the exposure time is not slower than 1/30 s due to potential vibration in the photographic system. Acquire an image, and check for a proper histogram.
10. Turn off the transillumination, and turn on the epi-illumination lamps again.
11. Increase the magnification to that used in step 6.7. Refocus if needed.
12. Increase the ISO within the range of 3,000-6,000, and adjust the exposure time to fall within the proper exposure range of the camera. Acquire an image.
    NOTE: The ruby bead may no longer appear in the field of view. If so, capture separate images of the bead at this magnification for reference.
13. Turn off the epi-illumination lamps. Turn on the flash with a 1/4 s exposure and with the camera shutter speed to 1/250 s. Set the flash speed to the default, or change it during the initial setup of the camera, and set the ISO to 500-1,000. Acquire the image. Check the histogram for proper exposure.
14. Turn off the flash, and turn on the trans-illumination lamps. Reset the ISO to above 8,000 to allow a faster exposure time with a more sensitive image sensor. Ensure that the exposure time does not go below 1/30 s due to potential vibration in the photographic system. Acquire an image.
15. Export the images from the camera to a computer. Review the images before removing the specimen from the microscope stage in case some need to be captured again.

7. Image acquisition overview for ex vivo OCT and scanning laser ophthalmoscopy (SLO)

1. For spectral domain OCT, place the globes in phosphate buffer in a custom eye holder with a chamber with a 60 diopter lens³⁵. Mount the eye holder to a bracket on a clinical OCT imaging device, and attach it to where a patient would rest their forehead. The OCT device automatically inserts a scale bar into each image.
2. At the same time, using the same device and with the eye still in the holder, image the globes with a scanning laser ophthalmoscope (SLO) using near-infrared reflectance (NIR, used as a locator image by the indwelling software), 488 nm excitation fundus autofluorescence (FAF), and red-free (RF) reflectance.
   NOTE: The 787 nm excitation FAF in this device is only occasionally suitable because a beam-splitter reduces the light transmittance to the SLO. For this reason, a second device for 787 nm FAF images (see next point) is used.
3. Separately, but with the eye still in the eye holder, image the globes with 787 nm FAF for detecting RPE disturbance³⁶ using a separate device that displays this modality well.

8. Image acquisition protocol for ex vivo OCT/ SLO (see slides in Supplementary Material 2)

1. Indicate the superior rectus muscle with tissue marking dye. Turn on the laser (blue arrow, Supplementary Material 2, page 1).
2. Referring to page 2 of Supplementary Material 2, position the OCT head by moving the entire unit in two axes with respect to the base (green arrow) and then raising the height (y) by rotating the joystick clockwise/counterclockwise (cw/ccw, blue arrows). Focus the image by rotating the knob (orange arrow), with the black lever in position R (*). Lock down the unit by securing the thumb screw (purple arrow).
3. Insert the eye into the holder, and stabilize from the posterior aspect with spacers (Supplementary Material 1, page 13). Exert as little pressure as possible to avoid denting the sclera. Orient with the tissue mark for the superior rectus muscle facing up.
   NOTE: The approximate distance from the front of the eye holder to the OCT device is 25 mm.
4. Open the proprietary visualization and analysis software for the OCT device. A patient list will appear in the left column. The eye donors indexed by an internal code number are the "patients." Refer to the user manual from the manufacturer.
5. Select the New Patient Icon. Complete the patient data information as needed. Select OK. Use a logically ordered numbering system for individual eyes, such as YYYYNNNL,R_agesex. For example, this could be 2017016R_97F.
6. After continuing the data entry on the following window, press OK. Select the operator and study from the drop-down menu.
   NOTE: The entered information will appear in the exportable meta-data.
7. After viewing a blank screen, touch the yellow button to start the image acquisition.
8. Press IR + OCT (near infrared reflectance + OCT). Allow the laser to acquire a live SLO image of the fundus and OCT B-scan.
   1. Finalize the correct anatomical position based on the ONH and fovea, and use a wooden applicator to adjust the eye position in the holder. On the control panel, rotate the black round button to adjust the intensity.
   2. Press the same button to average 9-100 frames (red arrows; 9 is sufficient, 100 looks creamy). If the unit is oriented correctly, the fundus should be in focus, and the OCT B-scan should appear in the top third of the display (Supplementary Material 2, page 9, double red arrow).
   3. On the fundus image, use the cursor to move the blue line to center the fovea (Supplementary Material 2, page 9, white arrow). Press Acquire.
      NOTE: Other default buttons are Retina, EDI (off), and Line Scan.
9. Press RF + OCT for the next acquisition. Recheck the position to ensure that the image has not moved or degraded. Press the black button for averaging. Press Acquire.
10. Switch the internal cube for autofluorescence imaging to the 488 nm and 787 nm excitation wavelengths (Supplementary Material 2, page 10).
    NOTE: The cube position after the switch is shown.
11. Select Autofluorescence mode. Recheck the alignment. Press Acquire.
12. For suspected cases of RPE disruption and atrophy, select ICGA (indocyanine green angiography) for 787 nm autofluorescence. Recheck the eye position, and then press Acquire.
    NOTE: A fundus image often does not appear in this modality because the internal cube attenuates the beam.
13. Switch the internal cube back to R for IR and red free imaging for volume acquisition.
    NOTE: The cube position after the switch is shown on page 13 of Supplementary Material 2.
14. To acquire the OCT volumes, select IR and the volume setting. Match all the settings on page 14 of Supplementary Material 2 by toggling the appropriate buttons on the control module (30° macula cube, 30 µm spacing, averaging depending on the experimental requirements).
15. Notice that the near-infrared reflectance fundus view is covered in blue B-scan lines. Recheck the OCT position in the upper third of the right window. Press Acquire on the Control Module, and wait 5 min for the volume scan to complete. When the imaging acquisition is completed, select EXIT. The images will be saved (red arrow).
    NOTE: The blue lines delimit the distances shown in the previous view in micrometers. The scans start numbering from the bottom (inferior) and proceed upward. Note the red line progression.
16. Allow the computer to process the acquired images, which will appear on the screen (Supplementary Material 2, page 16).
17. When imaging the fellow eyes of one donor, do not change the position of the mounting bracket between the eyes. If the left eye is imaged first, the results will show in the OD (right eye) column. Right-click to select all the images, and then select exchange OS/OD. The images will shift to the OS column.
18. Press Select > Window > Database. The screen defaults to panel 6 with the addition of the donor eye imaged in the right column (Supplementary Material 2, page 18). Right-click on the patient in the dropdown menu, choose Batch, and export the E2E file.
19. Browse to the pre-determined folder created on the desktop for file transfers. Select OK. The folder contains the E2E files to be copied onto an external hard drive and archived.
20. Bring the eye in its holder to the scanning laser ophthalmoscope, which is principally used for 787 nm autofluorescence.
    NOTE: refer to the manufacturer's user guide for the acquisition and archiving of the images.
21. Turn on the computer and the laser.
22. Select the New Patient icon. Complete the patient data as needed.
23. To keep the eye datasheet the same, press OK. As the C-curve stays the same, press Continue.
24. Select Study mode, and enter the password if required. Keep the C-curve at 7.7 mm. Press Ok.
25. Select Continue to verify the C-curve. Select the yellow indicator to start the camera.
26. Select the R position. Align and orient the globe. Select the IR mode to focus and orient as above.
27. Orient the camera head by moving the entire unit in two axes with respect to the base (Supplementary Material 2, page 28, green arrow) and then raising the height (y) of the unit (blue arrows). Focus the image by rotating the knob (orange arrow). The black lever is in position R (*). After this head is in position, lock down the unit by turning the thumb screw (purple arrow).
28. Note that the screen appears as shown on page 29 of Supplementary Material 2.
29. Move the selector knob from R to A. Select ICGA (to achieve 787 nm autofluorescence) in blue, 100% intensity, a 30° field of view, and single-phase imaging.
    NOTE: Like the dye indocyanine green, melanin is excited by 787 nm wavelength light.
30. Note that the screen appears as shown on page 31 of Supplementary Material 2. Press the black disc for averaging, and then select Acquire.
31. Choose Window > Database. Select Import E2E files from the SLO device; these files are stored on an external hard drive and uploaded to the desktop.
32. Select Open. Check the marks are the default, and select OK.
33. Note that the patient now has two tabs: one showing the images acquired from the SLO (blue arrow), and the other tab showing the images acquired from the OCT device (red arrow) (Supplementary Material 2, page 34).
34. Right-click on the 786 nm (ICGA) image, and export the picture to a file labeled SLO 786 on the desktop.
35. To save the 786 nm autofluorescence SLO image, select the patient, and right-click to export images as RAW files (for .vol files) to a folder on the desktop.
36. To export images from the OCT device for transfer to the archive computer, copy/paste from RAWEXPORT to a folder labeled RAW.

9. Imaging review

1. Assemble the images (color, .vol file, SLO images) in folders for each donor with subfolders for each eye, and index the folders consecutively by laboratory ID.
2. Record the tissue impressions (quality, pathology) in a database for a standardized report.
3. Review the exported OCT volumes with an in-house ImageJ plugin.
   1. Find the fovea center, which can be recognized by the center of the foveal dip, the inward rise of the outer retinal bands, or the thinnest point. In most cases, these will coincide, but not always, as this depends on the foveal preservation, individual variability, and the presence of pathology.
   2. Find a standard scan in the superior perifovea (2 mm/67 B-scans away in the superior direction; i.e., increasing scan numbers).
   3. Save a .tif stack of the entire volume for quick reference in the future.
   4. Scroll through the OCT volume in its entirety, starting from scan 1 (inferior), while recording the scan numbers in which features are recognized.
   5. Check the peripapillary area in addition to the macula.
      NOTE: Peripapillary chorioretinal atrophy in older eyes includes a distinctive basal laminar deposit and neovascularization. This is a common area of pigmentary change in myopic eyes and glaucoma, as well as in aging and AMD³⁷.
4. Inspect the color photographs, and link any findings to those seen by OCT if possible.
   NOTE: In general, it is easier to see most findings by OCT first. Color photographs do provide a large view of the area outside the area on the SLO, choroidal pigmentation including nevi, the presence of heme and hard exudates, and black pigment in neovascular AMD. Dark pigment in the fovea may represent loose melanosomes from the anterior segment and should be washed away gently with buffer using a pipette.
5. Inspect the SLO images, and link any findings to those seen by OCT if possible.
6. Save standard B-scans at the fovea and perifovea, extra B-scans with notable pathology or other features, and SLO images in a pathology report.
7. Categorize the eyes as follows: Unremarkable, Questionable, Early-Intermediate AMD, Atrophic AMD, Neovascular AMD, Other, Unknown, Not Gradable, Not Recorded. "Questionable" is used if it is not clear if changes are severe enough for another categorization. "Not Gradable" is reserved for eyes lacking useful OCT scans, such as those with severely detached retinas. "Not Recorded" is reserved for eyes that are processed immediately upon receipt without photography.
8. Use these criteria for AMD: severe RPE change with either drusen or subretinal drusenoid deposits, with or without signs of neovascularization, such as fluid or fibrosis, and without evidence of other causes (updated from Curcio et al.¹⁰ to accommodate SDDs).
   NOTE: The final diagnosis is confirmed by histologic analysis.

Wyniki

Table 1 shows that during 2016-2017, 184 eyes from 94 white non-diabetic donors >80 years of age were recovered. The mean death-to-preservation time was 3.9 h (range: 2.0-6.4 h). Of the 184 eyes reviewed, 75 (40.2%) had certain AMD. The following categories were identified: Unremarkable (39.7%), Questionable (11.4%), Early-Intermediate AMD (22.8%), Atrophic (7.6%), Neovascular (9.8%), Other (8.7%), and Unknown/Not Recorded/Not Gradable (<1%). Figure 2,

Dyskusje

Using a population-based screening approach during a 16 month period in the pre-COVID era, it was possible to procure 75 donor eyes with AMD. All were recovered with a short DtoP and staged using OCT-anchored MMI. The age criterion (>80 years) is outside the typical age range for tissue recoveries intended for transplantable corneas. Despite the advanced age, our criteria resulted in eyes at all stages of AMD. Many RPE phenotypes are common to all AMD stages, and some are exclusive to neovascular AMD

Materiały

Name	Company	Catalog Number	Comments
Beakers, 250 mL	Fisher	# 02-540K
Bottles, 1 L, Pyrex	Fisher	# 10-462-719	storage for preservative
Bunsen burner or heat source	Eisco	# 17-12-818	To melt wax
Camera, digital	Nikon D7200	D7200
Computer and storage	Apple	iMac Pro; 14 TB external hard drive	Image storage
Container, insulated	Fisher	# 02-591-45	For wet ice
Containers, 2 per donor, 40 mL	Fisher	Sameco Bio-Tite  40 mL # 13-711-86	For preservative
Crucible, quartz 30 mL	Fisher	# 08-072D	Hold globe for photography
Cylinder, graduate, 250 mL	Fisher	# 08-549G
Disinfectant cleaning supplies			https://www.cardinalhealth.com/en/product-solutions/medical/infection-control/antiseptics.html
Eye holder with lens and mounting bracket	contact J. Messinger	jeffreymessinger@uabmc.edu	custom modification of Heidelberg Engineering original design
Face Protection Masks	Fisher	# 19-910-667
Forceps, Harmon Fix	Roboz	# RS-8247
Forceps, Micro Adson	Roboz	# RS-5232
Forceps, Tissue	Roboz	# RS-5172
Glass petri dish, Kimax	Fisher	# 23064
Gloves Diamond Grip	Fisher	# MF-300
Gowns GenPro	Fisher	# 19-166-116
Image editing software	Adobe	Photoshop 2021, Creative Suite
KimWipes	Fisher	# 06-666
Lamps, 3 goosenecks	Schott Imaging	# A20800
Microscope, stereo	Nikon	SMZ 1000	for dissection
Microscope, stereo	Olympus	SZX9	color fundus photography
Paraformaldehyde, 20%	EMS	# 15713-S	for preservative; dilute for storage
pH meter	Fisher	# 01-913-806
Phosphate buffer, Sorenson’s, 0.2 M pH 7.2	EMS	# 11600-10
Ring flash	B & H Photo Video	Sigma EM-140 DG
Ruby bead, 1 mm diameter	Meller Optics	# MRB10MD
Safety Glasses 3M	Fisher	# 19-070-940
Scanning laser ophthalmoscope	Heidelberg Engineering	HRA2
Scissors, curved spring	Roboz	# RS-5681
Sharps container	Fisher	# 1482763
Shutter cord, remote	Nikon	MC-DC2
Spectral Domain OCT device	Heidelberg Engineering	Spectralis HRA&OCT	https://www.heidelbergengineering.com/media/e-learning/Totara-US/files/pdf-tutorials/2238-003_Spectralis-Training-Guide.pdf
Stainless steel ball bearing, 25.4 mm diameter	McMaster-Carr	# 9529K31
Tissue marking dye, black	Cancer Diagnostics Inc	# 0727-1
Tissue slicer blades	Thomas Scientific	# 6767C18
Trephine, 18-mm diameter	Stratis Healthcare	# 6718L
TV monitor (HDMI) and cord for digital camera	B&H Photo Video	BH # COHD18G6PROB	for live viewing and remote camera display features
Wax, pink dental	EMS	# 72670
Wooden applicators	Puritan	# 807-12