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In This Article

  • Summary
  • Abstract
  • Introduction
  • Protocol
  • Results
  • Discussion
  • Disclosures
  • Acknowledgements
  • Materials
  • References
  • Reprints and Permissions

Summary

This protocol describes the methodology to genetically ablate the retinal pigment epithelium (RPE) using a transgenic zebrafish model. Adapting the protocol to incorporate signaling pathway modulation using pharmacological compounds is extensively detailed. A MATLAB platform for quantifying RPE regeneration based on pigmentation was developed and is presented and discussed.

Abstract

The retinal pigment epithelium (RPE) resides at the back of the eye and performs functions essential for maintaining the health and integrity of adjacent retinal and vascular tissues. At present, the limited reparative capacity of mammalian RPE, which is restricted to small injuries, has hindered progress to understanding in vivo RPE regenerative processes. Here, a detailed methodology is provided to facilitate the study of in vivo RPE repair utilizing the zebrafish, a vertebrate model capable of robust tissue regeneration. This protocol describes a transgenic nitroreductase/metronidazole (NTR/MTZ)-mediated injury paradigm (rpe65a:nfsB-eGFP), which results in ablation of the central two-thirds of the RPE after 24 h treatment with MTZ, with subsequent tissue recovery. Focus is placed on RPE ablations in larval zebrafish and methods for testing the effects of pharmacological compounds on RPE regeneration are also outlined. Generation and validation of RpEGEN, a MATLAB script created to automate quantification of RPE regeneration based on pigmentation, is also discussed. Beyond active RPE repair mechanisms, this protocol can be expanded to studies of RPE degeneration and injury responses as well as the effects of RPE damage on adjacent retinal and vascular tissues, among other cellular and molecular processes. This zebrafish system holds significant promise in identifying genes, networks, and processes that drive RPE regeneration and RPE disease-related mechanisms, with the long-term goal of applying this knowledge to mammalian systems and, ultimately, toward therapeutic development.

Introduction

The methodology described herein details a protocol to genetically ablate the retinal pigment epithelium (RPE) utilizing larval zebrafish. The RPE extends over the back of the eye and resides between the stratified layers of the neural retina and the layer of vasculature constituting the choroid. Trophic support, absorption of phototoxic light, and maintenance of visual cycle proteins are only some of the critical functions the RPE performs that are essential for sustaining the health and integrity of these adjacent tissues1. Damage to mammalian RPE is reparable when lesions are small2; however, damage suffered by larger injuries or progressive degenerative disease is irreversible. In humans, RPE degenerative diseases (e.g., age-related macular degeneration (AMD) and Stargardt disease) lead to permanent vision loss and, with few treatment options available, decreased patient quality of life. The limited ability for mammalian RPE to self-repair has created a knowledge gap in the field of RPE regenerative processes. Given the robust regenerative capacity of the zebrafish across many different tissue types, this protocol was developed to establish an in vivo vertebrate system to facilitate studies on intrinsically regenerating RPE and uncover mechanisms that drive that response. Using the ablation paradigm outlined here, the canonical Wnt signaling pathway3, the mTOR pathway4, and immune-related responses5 have been identified as critical mediators of RPE regeneration, likely with overlapping functions.

In this genetic ablation paradigm, Tg(rpe65a:nfsB-eGFP)3 zebrafish express the bacterial-derived nitroreductase (NTR/nfsB) gene6 fused to eGFP under control of the RPE enhancer element, rpe65a7. Ablation is achieved by adding the prodrug, metronidazole (MTZ), to system water housing zebrafish. Intracellular activation of MTZ by nitroreductase results in DNA crosslinking and apoptosis in NTR/nfsB-expressing cells8,9. This technology has been widely used in zebrafish to ablate cells of the retina10,11,12,13 and other tissues8. Together, these elements enable targeted expression (rpe65a) of an inducible cell ablation methodology (NTR/MTZ)8,9 and a fluorescent marker (eGFP) for visualization.

Other interesting in vivo models also exist that can be used to study the regenerative potential of the RPE14. These are broad and include RPE-to-retina transdifferentiation post-retinectomy in amphibians, in which RPE cells lost to retinal regrowth are replaced15,16; RPE restoration post-injury in the "super healing" MRL/MpJ mouse17; and exogenous stimulation of RPE proliferation in a rat model of spontaneous RPE and retinal degeneration18, among others. In vitro models, such as adult human RPE stem cells (RPESCs)19 have also been developed. These models are all valuable tools working to uncover the cellular processes related to RPE regeneration (e.g., proliferation, differentiation, etc.); however, the zebrafish is unique in its capacity for intrinsic RPE repair post-ablation.

While the methodology here is written to focus on understanding the mechanisms driving RPE regeneration, the Tg(rpe65a:nfsB-eGFP) line and this genetic ablation protocol could be utilized to study other cellular processes such as RPE apoptosis, RPE degeneration, and the effect of RPE injury on adjacent retinal and vascular tissues. The ablation protocol can also be modified to include pharmacological manipulation, which is a convenient preliminary strategy to screen signaling pathways of interest. For example, blocking the canonical Wnt pathway using Inhibitor of Wnt Response-1 (IWR-1)20, has been shown to impair RPE regeneration3. This was repeated here to guide users through a pharmacological manipulation experiment and serve as proof-of-concept to validate a MATLAB script (RpEGEN) created to quantify RPE regeneration based on recovery of pigmentation. Like the transgenic line and ablation protocol, the RpEGEN scripts are adaptable and could be used to quantify other markers/cellular processes within the RPE.

Protocol

All methodologies outlined herein are compliant with the Institutional Animal Care and Use Committee (IACUC) of the University of Pittsburgh.

1. Preparation prior to zebrafish embryo collection

  1. Set embryo incubator to 28.5°C.
  2. Prepare a 25x stock solution of the melanogenesis inhibitor, N-phenylthiourea (PTU)21,22. This stock solution is scaled from a common recipe22 and 1x is equal to 0.003% weight per volume (% w/v) (e.g., 0.003 g of PTU powder into 100 mL of liquid solvent).
    1. To make a large 25x PTU stock solution, add 0.75 g of PTU powder to 1 L of purified deionized water (hereafter referred to as dH2O) and mix thoroughly at room temperature (~25 °C) using a stir bar and stir plate. Store at 4°C for up to 3 months protected from light.
      NOTE: It is difficult to get PTU to go into aqueous solution and extended stirring overnight may be necessary.
      CAUTION: PTU is hazardous, and care should be taken to prevent ingestion, inhalation, and/or contact with the skin or eyes. PTU powder and all PTU liquid derivatives described herein may need to be disposed of as chemical waste depending on state and institutional regulations. Confirmation of proper PTU waste disposal methods, if any, is recommended prior to use.
    2. To make a 1.5x PTU working solution (hereafter referred to as 1.5x PTU), add 60 mL of 25x PTU stock solution to 940 mL of zebrafish housing facility water (hereafter referred to as system water). Optimal water quality parameters for zebrafish have been described23 and aquatics facilities should have standard water monitoring procedures in place. Store 1.5x PTU at 28.5°C for 1-2 weeks protected from light.
      NOTE: This protocol is routinely performed using the PTU concentrations, solvents, and storage parameters described in step 1.2. As a precaution, embryos/larvae should be observed every 1-2 days while in PTU to validate efficacy and confirm sustained depigmentation. Dissolution and/or storage conditions should be optimized if decreases in PTU solubility/efficacy are suspected.
  3. Prepare a stock solution of 0.05% w/v methylene blue, a fungal growth inhibitor, by adding 0.05 g of methylene blue powder to 100 mL of dH2O. Mix thoroughly using a stir bar and stir plate. Store at 4°C.
  4. Prepare pipettes for embryo/larva manipulation (e.g., moving embryos/larvae between Petri dishes, separating larvae during fluorescence screening, collecting euthanized larvae into microcentrifuge tubes for fixation, etc.) by cutting back the tapered end of a glass Pasteur pipette using a diamond tip scribing pen. Etch around the circumference of the pipette with the diamond pen and gently pull or snap the end off to make a clean break.
    NOTE: The mouth of the pipette should be smooth and wide enough to easily take up an embryo still inside the chorion without shearing. Use a bulb draw transfer pipette as an alternative. Prepared pipettes can also be used to remove liquid during water changes (rather than pouring) to minimize embryo/larva loss.
  5. Prepare a 4% paraformaldehyde (PFA) solution in 1x phosphate buffered saline (PBS) (e.g., add 10 mL of 16% PFA to 4 mL of 10x PBS and 26 mL of dH2O). Store at 4°C for up to 4 weeks protected from light.
    CAUTION: Paraformaldehyde is a hazardous chemical and should be handled in a chemical fume hood and disposed of properly. Care should be taken, and personal protective equipment (PPE) should be worn, to prevent ingestion, inhalation, and/or contact with the skin or eyes.

2. Zebrafish embryo collection and maintenance prior to genetic ablation (0-5 days post-fertilization)

  1. Maintain adult zebrafish as described previously3,4,5. The afternoon/evening prior to embryo collection, separate adult zebrafish into breeding tanks for spawning.
  2. The following morning (0 days post-fertilization (dpf)), collect embryos into 10 cm diameter Petri dishes in system water and remove all nonviable or unfertilized eggs, which will appear opaque and/or show irregular cytoplasm and failed cleavage24.
    NOTE: Normal cleavage and developmental staging events will be apparent in healthy embryos as described25. Petri dishes should be kept three-fourths full (~30 mL for a 10 cm diameter dish) throughout the protocol.
    1. Add two drops of 0.05% w/v methylene blue to each Petri dish, mix gently, and store embryos at 28.5°C for the remainder of the protocol.
  3. Around 6 h post-fertilization (hpf)4,5,26, replace system water in embryo Petri dishes with 1.5x PTU (working solution made in step 1.2.2) and replenish methylene blue.
    NOTE: PTU must be added to embryos prior to the onset of pigmentation (i.e., before 24 hpf)25 as already pigmented tissues will not depigment upon addition of PTU21. It should be noted, however, that reduced eye size, ocular and craniofacial deficits, and disruption of some signaling pathways (e.g., thyroid signaling) have been reported in PTU-treated zebrafish27,28,29. The developmental toxicity of PTU appears to be dependent on concentration and timing of PTU addition27,29. As mentioned above for validating PTU efficacy (step 1.2), signs of PTU toxicity should also be carefully monitored and, if suspected, the working concentration and/or time of PTU addition should be optimized.
  4. On 2-3 dpf, dechorionate embryos using freshly made pronase solution.
    1. Dissolve pronase in 1.5x PTU at a concentration of 2 mg/mL by vortexing.
      CAUTION: Pronase is packaged as a very fine powder and is an irritant. Take measures to avoid inhalation and/or contact with skin, eyes, etc.
    2. Separate hatched embryos from unhatched embryos and pronase-treat only unhatched embryos.
    3. Replace 1.5x PTU with 2 mg/mL pronase solution made in step 2.4.1 and leave on unhatched embryos for 4-5 min with gentle agitation (e.g., on a tabletop rotator/shaker or by manual swirling).
    4. Pour off the pronase solution and immediately rinse with fresh 1.5x PTU. Gently triturate 1.5x PTU rinse over embryos with a bulb-draw transfer pipette.
    5. Repeat a second 1.5x PTU rinse to discard all chorion debris, and then replenish 1.5x PTU for maintenance.
      NOTE: Embryos can also be dechorionated manually using fine-tipped forceps. In this case, chorion debris should be removed and 1.5x PTU replenished after manual dechorionation.
  5. Monitor embryo/larval health and replenish 1.5x PTU every 1-2 days. Embryos/larvae are kept in 1.5x PTU until ablation at 5 dpf.
    NOTE: The importance of steps 2.3 and 2.4 are addressed further in the Discussion section.

3. Screening zebrafish larvae for rpe65a:nfsB-eGFP and genetic ablation of the retinal pigment epithelium (5-6 days post-fertilization)

  1. Make a fresh 10 mM metronidazole (MTZ) solution on 5 dpf (day of ablation). This process takes 2 h to complete.
    1. Add MTZ powder to system water without PTU and mix thoroughly by vigorous shaking (e.g., 250 rotations per min) for 1 h at 37 °C.
    2. Cool 10 mM MTZ solution for an additional 1 h at room temperature on a tabletop rotator/shaker and ensure complete dissolution prior to adding to Petri dishes with larvae.
      NOTE: Fluorescence screening and separation of eGFP+ larvae (step 3.2) can be performed during the 37 °C and room temperature incubations.
      CAUTION: MTZ is hazardous, and care should be taken to prevent ingestion, inhalation, and/or contact with the skin or eyes. MTZ powder and all liquid derivatives described herein may need to be disposed of as chemical waste depending on state and institutional regulations. Confirmation of proper MTZ waste disposal methods, if any, is recommended prior to use.
  2. Screen zebrafish larvae for the rpe65a:nfsB-eGFP transgene.
    1. Anesthetize larvae with 0.168 g/L of tricaine (MS-222) and separate transgenic (eGFP+) larvae (Figure 1) from non-transgenic (eGFP-) larvae using a fluorescence stereo microscope with a 488 nm excitation laser/filter.
      NOTE: Tricaine should be added to 1.5x PTU and/or pharmacological compound solutions as larvae should remain immersed in the treatment while being screened for eGFP. Incubate larvae in tricaine only for the duration of screening (e.g., ≤10 min for a single 10 cm Petri dish containing 50 larvae).
    2. Wake screened larvae up immediately by pipetting directly into a Petri dish with fresh 1.5x PTU without tricaine.
    3. Upon completion of screening, further separate the eGFP+ larvae into two groups of Petri dishes: one group to receive MTZ treatment (ablated/MTZ+) and one group to be the unablated (MTZ-) control.
  3. Ablate the retinal pigment epithelium.
    1. Remove 1.5x PTU from the unablated (MTZ-) control dishes and add fresh system water without PTU.
    2. Remove 1.5x PTU from the ablated (MTZ+) treatment dishes and add the freshly made 10 mM MTZ solution (step 3.1.).
    3. Remove the 10 mM MTZ solution after exactly 24 h (designated 1 day post-injury (dpi)) and add fresh system water without PTU. Change out the fresh system water without PTU on the MTZ- dish(es). Larvae will not be exposed to PTU again for the remainder of the protocol.
      NOTE: It may be difficult to pipette or pour off all 1.5x PTU (steps 3.3.1 and 3.3.2) or 10 mM MTZ (step 3.3.3) between solution exchanges without larval loss as animals are actively swimming around. In this event, wash(es) of system water without PTU can be added to ensure successful solution exchange.

4. Larval maintenance post-genetic ablation (6+ days post-fertilization)

  1. Check larvae and replenish system water without PTU daily until euthanasia (step 5.6) or return to zebrafish housing facility.
  2. Monitor the success and extent of ablation in vivo on 2 dpi (7 dpf) using transmitted light illumination on a stereo microscope (Figure 2).

5. Incorporating pharmacological treatment into zebrafish retinal pigment epithelium ablation protocol

NOTE: As performed previously3, treatment with 15 µM IWR-1 or volume-matched dimethyl sulfoxide (DMSO) vehicle control starting at 4 dpf is outlined here as an example experiment to test RpEGEN. Concentrations and timelines may vary with different pharmacological compounds and recommendations for dose-response validation, treatment duration, screening, and other aspects of experimental design for pharmacological manipulation studies are addressed in the Discussion section. Follow steps 6 and 7 if image analysis is required.

  1. Collect and maintain embryos as described in step 2. Dechorionate embryos on 2 dpf.
  2. Screen eGFP+ larvae on 4 dpf as described in step 3.2. Rather than Petri dishes, place eGFP+ larvae into 6-well plates at a density of n ≤ 10 larvae per 6-well for pharmacological treatment. Designate separate 6-well plates for larvae that will be unablated (MTZ-) and larvae that will be ablated (MTZ+).
    NOTE: The eGFP signal is visible on 4 dpf but appears dimmer than signal intensity on 5 dpf.
  3. Pretreat 4 dpf eGFP+ larvae with 15 µM IWR-1 or volume-matched DMSO vehicle control for exactly 24 h prior to genetic ablation with 10 mM MTZ solution.
    NOTE: Often, very little compound is needed for pharmacological treatment experiments. To avoid weighing out small quantities of IWR-1 powder, this pharmacological compound is purchased already in DMSO solution, at a concentration of 25 mM, and aliquoted into smaller volumes upon arrival to avoid repeated freeze-thaw cycles.
    1. Determine the volume of pharmacological and vehicle control treatments needed and aliquot 1.5x PTU into conical tubes accordingly. A volume of 5 mL/well of a 6-well plate is recommended.
    2. Add IWR-1 stock to 1.5x PTU for a final concentration of 15 µM IWR-1 (e.g., 3 µL of 25 mM IWR-1 per 5 mL of 1.5x PTU). Add a matched volume of DMSO stock to 1.5x PTU (e.g., 3 µL ≥ 99.7% DMSO per 5 mL of 1.5x PTU). Here, this will end up yielding a final concentration of 0.06% volume/volume (% v/v) DMSO. Mix well by vortexing and visually confirm dissolution of compounds.
      CAUTION: DMSO, IWR-1, and other pharmacological compounds and solvents may need to be disposed of as chemical waste depending on state and institutional regulations. Confirmation of hazard level and proper waste disposal methods for these compounds, if any, is recommended prior to use.
    3. Remove 1.5x PTU from the eGFP+ larvae in 6-well plates and add 5 mL/well of freshly made 0.06% v/v DMSO or 15 µM IWR-1 treatments from step 5.3.2.
  4. On 5 dpf, ablate the RPE. In addition to the 24 h pretreatment (step 5.3), larvae will remain immersed in pharmacological and vehicle control treatments during 24 h of genetic ablation with 10 mM MTZ and during post-ablation recovery, until fixation (e.g., from 4-9 dpf).
    1. Make 10 mM MTZ solution 2 h prior to performing genetic ablation (step 3.1).
    2. Determine the volume of pharmacological and vehicle control treatments needed for both unablated (MTZ-) and ablated (MTZ+) 6-well plates and aliquot appropriate volumes of either fresh system water without PTU (MTZ-) or 10 mM MTZ solution (MTZ+) into conical tubes. This will yield four treatment conditions: 1) 0.06% v/v DMSO, MTZ-; 2) 15 µM IWR-1, MTZ-; 3) 0.06% v/v DMSO, MTZ+; and 4) 15 µM IWR-1, MTZ+.
    3. Add IWR-1 and DMSO stock solutions to respective conical tubes as performed in step 5.3.2. Mix well by vortexing and visually confirm dissolution of compounds.
    4. Remove 0.06% v/v DMSO and 15 µM IWR-1 treatments in 1.5x PTU (step 5.3.2) from designated unablated (MTZ-) and ablated (MTZ+) 6-well plates and replenish with the appropriate treatments made in step 5.4.3.
    5. Remove 0.06% v/v DMSO and 15 µM IWR-1 treatments in 10 mM MTZ solution after exactly 24 h and replenish with treatments in fresh system water without PTU. Replenish 0.06% v/v DMSO and 15 µM IWR-1 treatments in fresh system water without PTU on the unablated (MTZ-) 6-well plate(s).
  5. Follow larval maintenance post-ablation as outlined in step 4 and replenish 0.06% v/v DMSO or 15 µM IWR-1 treatments daily in system water without PTU.
  6. Euthanize larvae on 9 dpf (4 dpi for age-matched MTZ-treated siblings) by immersing animals in 0.3 g/L tricaine solution (lethal overdose) coupled with rapid chilling (e.g., place Petri dishes on ice) for at least 20 min30. Check to make sure that larvae are unresponsive to touch and fix in 4% PFA (step 1.5) for 3 h at room temperature or at 4 °C overnight.
  7. Process post-fixation larval tissue for z-stack image acquisition on a confocal microscope as described previously5,31 and here in the Representative Results section. Analysis in steps 6 and 7 will require, at minimum, acquisition of nuclear marker (e.g., DAPI) and brightfield z-stack images.

6. Confocal microscope z-stack image preprocessing in FIJI (ImageJ)

  1. Import and format confocal microscope z-stack images using FIJI32.
    1. Open microscope images using the Bio-Formats Import Options set to View stack with: Hyperstack and Color mode: Grayscale.
    2. Generate a maximum intensity projection of the imported microscope image by choosing Image | Stacks | Z Project. In the ZProjection window, set Start Slice and Stop Slice to include all slices for that image. For example, set Start Slice: 1 and Stop Slice: 18 for a z-stack that has a total of 18 slices. Choose Projection Type: Max Intensity and click on OK.
    3. Convert the maximum intensity projection file to an 8-bit image (if not already) by choosing Image | Type | 8-bit.
    4. Reorient the image so that the dorsal side is up and distal (i.e., the lens) is left by choosing Image | Transform | Flip Horizontally (for the last command, choose the option that best suits the directionality needed for that image). For a multi-channel image, click on Yes in the Process Stack? window to reorient all channels.
      NOTE: This step is critical, but not needed if the image is already in the dorsal up, distal left orientation. See Figure 3A-D and Figure 4 for images with eyes in the correct directionality for processing.
    5. Save the 8-bit maximum intensity projection as a tagged image file format (TIF) file by choosing File | Save As | Tiff....
  2. Generate RPE region of interest (ROI) using FIJI.
    1. Open an 8-bit TIF image generated as described in step 6.1. Start the ROI Manager by choosing Analyze | Tools | ROI Manager.
    2. Use Image | Zoom and toggle between the DAPI and brightfield channels to identify the point(s) at which the apical side of the RPE is adjacent to the tip of the outer limiting membrane (OLM) (Figure 3B', B", D', D"; blue arrowheads). Use this anatomical landmark as the ROI starting point.
    3. Create the RPE ROI with the Polygon Selections tool (in the FIJI toolbar) using both the DAPI and brightfield image channels and zoom function (Image | Zoom) to identify apical and basal RPE boundaries. Bring the dorsal and ventral ends of the ROI (i.e., where the ROI transitions from the apical to basal side of the RPE) to a sharp point rather than blunting or rounding off (Figure 3B', B", D', D"; magenta lines).
      NOTE: Creating a pointed ROI end is a critical step for optimizing endpoint detection using RpEGEN and is addressed in the Discussion section.
    4. Add the ROI by clicking on Add in the ROI Manager. Adjust the ROI as needed and click on Update in the ROI Manager.
    5. Save the ROI file by choosing More>> | Save... within the ROI Manager. Use identical names for matched ROI and TIF image files (e.g., [file name].tif and [file name].roi).
  3. Combine 8-bit TIF and ROI files for each condition into a single folder. For example, the folder, DMSO_9dpf, will contain all the matched TIF and ROI files for the 9 dpf unablated (MTZ-) 0.06% v/v DMSO-treated larval group.

7. Quantification and visualization of RPE regeneration using RpEGEN scripts

  1. Install and prepare RpEGEN scripts.
    1. Download the latest RpEGEN scripts from the GitHub repository (https://github.com/burchfisher/RpEGEN) by clicking on Code | Download ZIP.
    2. Unzip the folder and place it in the desired workspace location (e.g., Desktop).
    3. Open MATLAB.
    4. Navigate to the RpEGEN folder in the Current Folder pane (usually on the left side).
    5. Right click on the RpEGEN folder and choose Add to Path | Selected Folders and Subfolders. This adds the folder to the MATLAB path so it can automatically find and run any scripts in the folder.
    6. Double click on the RpEGEN folder in the Current Folder pane to show all the subfolders and M files.
    7. Double click on the RpEGEN.m file to open in the Editor pane.
    8. Under the USER-DEFINED VARIABLES section of the RpEGEN.m file, enter the directory locations for the folders containing the ROI files (.roi), image files (.tif), and where output files should be saved. Enter the group name for the .mat file to be exported (e.g., DMSO_9dpf, DMSO_4dpi, etc.) and the location of the brightfield channel in the TIF image stack (e.g., 3, if brightfield is the third channel in an image stack). If the image file only contains the brightfield image, then this should be equal to 1.
  2. Run RpEGEN.m script and validate the results.
    NOTE: RpEGEN requires the Image Processing Toolbox, the Curve Fitting Toolbox, and the Statistics and Machine Learning Toolbox to be activated on the user MATLAB license in order to run. In addition, the freely available ReadImageJROI toolbox33 is required to import FIJI ROIs into MATLAB; however, it is provided in the RpEGEN folder along with other function M files that do not require any activation.
    1. Run the script by clicking on the Run button in the Editor menu at the top of MATLAB.
      NOTE: Once initiated, the Command window will provide verbose output indicating the progression of the script. After saving the MAT file containing the extracted data, a three-panel figure will appear and be saved as a PDF to the output directory for each image run. These are quality control (QC) figures to make sure that everything has run properly and include: 1) the brightfield image overlaid by the ROI (Figure 4A); 2) the ROI with centerline and associated angular distance (degrees) (Figure 4G); and 3) the ROI with the centerline median intensity values (0-255, 8-bit color scale) (Figure 4H). Wait until the QC PDFs have been saved to the output folder and the last figure has disappeared before proceeding to the next step.
    2. Open the individual PDFs exported to the output folder in any PDF viewer and verify that all ROIs match the brightfield images, that centerline values are reasonable approximations of the center of the ROIs, and that median intensity values are appropriately populated with data (i.e., not all the same value across the entire centerline).
      NOTE: A detailed description and structure of each variable saved in the MAT file by RpEGEN.m can be found in Table 1.
  3. Run the RpEGEN_PermPlot.m script.
    NOTE: The RpEGEN_PermPlot.m script uses the output of RpEGEN.m to run statistical comparisons using a permutation simulation of the medians of two groups and also provides the code for reproducing the plots in this paper using the freely available GRAMM toolbox34, which has also been included in the RpEGEN folder.
    1. Double click on the RpEGEN_PermPlot.m file to open in a new Editor tab.
    2. Under the SECTION 1 - USER-DEFINED VARIABLES in the RpEGEN_PermPlot.m file, enter the directory location for the output folder containing the MAT files from running RpEGEN.m and enter each MAT file name to be loaded (e.g., DMSO_4dpi.mat, IWR1_4dpi.mat).
    3. Run this section of the script by clicking on the Run Section button in the Editor menu at the top of MATLAB.
    4. In Section 2, enter the names of the two groups for statistical comparison in the data_A and data_B variables (these are the groups from which medians will be derived using the permutation simulation). In the bin_sz variable, enter the number of degrees over which to integrate the median intensity values for the datasets (default is 1-degree bins).
      NOTE: The reps variable indicates the number of permutations to use to build a probability distribution and can be set to any number (default value is 20,000). In general, a higher number of repetitions will be more statistically robust but will increase processing time.
    5. Run this section of the script by clicking on the Run Section button in the Editor menu at the top of MATLAB. This section may take some time to complete depending on the number of repetitions specified but does provide a continuous status update in the command window.
    6. Run HEATMAP FIGURE and GROUP RESULTS AND P-VALUES sections independently using the Run Section button. Edit data variables in any sections commented with "ENTER DATA HERE". PDFs of these figures are automatically saved for each and can be easily modified in any vector software post processing.
      NOTE: The plots generated in the RpEGEN_PermPlot.m file are ad hoc and will likely require modifications based on each user's specific data and visualization needs. However, the figures do provide a solid foundation that can be easily individualized using both MATLAB and GRAMM websites.

Results

Inhibiting the canonical Wnt signaling pathway is known to significantly impair zebrafish RPE regeneration using the genetic ablation paradigm (rpe65a:nfsB-eGFP) and pharmacological manipulation methodology (IWR-1) described in the protocol3. This experiment was repeated here to validate an automated method for quantifying zebrafish RPE regeneration based on pigmentation. The results summarized below encompassed all steps of the protocol, from the day of fertilization (0 dpf) to quantific...

Discussion

This protocol describes methodology to genetically ablate the RPE and study mechanisms of degeneration and regeneration in larval-aged zebrafish. This protocol has also been successfully performed in adult zebrafish3 but with less extensive characterization, which is why larvae are the focus here. Critical aspects of this part of the protocol (steps 1-4) include: 1) adding 1.5x PTU to embryos prior to the onset of melanogenesis, 2) dechorionating PTU-treated embryos on 2-3 dpf, 3) careful screenin...

Disclosures

L.L.L. is the co-inventor on US Patent #9,458,428, which describes an expedited method to derive retinal pigment epithelium from human pluripotent stem cells; this is unrelated to the content herein. J.M.G. and G.B.F. have nothing to disclose.

Acknowledgements

Work described herein was supported by the National Institutes of Health (RO1-EY29410 to J.M.G, and NIH CORE Grant P30-EY08098 to the Department of Ophthalmology); the UPMC Immune Transplant & Therapy Center (to L.L.L. and J.M.G.); and the E. Ronald Salvitti Chair in Ophthalmology Research (to J.M.G.). Additional support was received from the Wiegand Fellowship in Ophthalmology (to L.L.L), the Eye & Ear Foundation of Pittsburgh, and an unrestricted grant from Research to Prevent Blindness, New York, NY. Authors also wish to thank Amanda Platt for technical assistance and Dr. Hugh Hammer and the aquatics staff for excellent animal care support.

Materials

NameCompanyCatalog NumberComments
Lab Material/Equipment
2-(4-Amidinophenyl)-6-indolecarbamidine dihydrochloride (DAPI)Millipore SigmaD9542
6-well platesFisher Scientific07-200-83
Conical Polypropylene Centrifuge TubesFisher Scientific05-539-13Catalog number is for 50 mL tubes
Diamond tip scribing penFisher Scientific50-254-51Manufactured by Electron Microscopy Sciences, items similar to this part number are adequate
Dimethyl sulfoxide (DMSO) ≥99.7 %Fisher ScientificBP231Check instiutional chemical waste disposal requirements
Embryo incubator (large)Fisher Scientific3720A
Embryo incubator (mini/tabletop)LabnetI5110A
Fluorescence stereo microscopeZeissAxio Zoom.V16Or similar, with 488 nm excitation laser/filter
Glass Pasteur pipetteFisher Scientific13-678-4Manufactured by Corning, non-sterile
InSolution Wnt Antagonist I, IWR-1-endoMillipore Sigma5.04462Manufactured by Calbiochem; 25 mM in DMSO; check instiutional chemical waste disposal requirements
Methylene blue (powder)Fisher ScientificBP117-100Also available as a premade aqeuous solution
Metronidazole (MTZ)Millipore SigmaM3761Check instiutional chemical waste disposal requirements
N-phenylthiourea (PTU)Millipore SigmaP7629Check instiutional chemical waste disposal requirements
Paraformaldehyde (16 % w/v) methanol freeFisher ScientificAA433689MChemical waste, proper disposal required
Petri dishesFisher ScientificFB087571210 cm diameter
Phosphate buffered saline (powder packets)Millipore SigmaP3813Used to make 10 X PBS stock
PronaseMillipore SigmaPRON-RO
Shaking incubatorBenchmarkH2010Used for incubating MTZ for 1 hour at 37 degrees Celcius
Stereo microscopeLeicaS9iOr similar, with transmitted light illumination
Student Dumont #5 forcepsFine Science Tools91150-20Fine-tipped forceps for manual dechorionation
Tabletop rotator/shakerScilogexSK-D1807-E
Transfer pipetteMillipore SigmaZ1350033.2 mL bulb draw, non-sterile
Tricaine methanesulfonate (MS-222)PentairTRS1, TRS2, TRS5Also available from Fisher Scientific (NC0342409)
VECTASHIELD Antifade Mounting Medium with DAPIVector LaboratoriesH-1200
Software Material
FIJI (Fiji is Just ImageJ)FIJI (Fiji is Just ImageJ)https://imagej.net/software/fiji/Version: 2.0.0-rc-69/1.52p; Build: 269a0ad53f; Plugin needed: Bio-Formats
GRAMM examples and how-tosMathWorkshttps://www.mathworks.com/matlabcentral/fileexchange/54465-gramm-complete-data-visualization-toolbox-ggplot2-r-like.
MATLABMathWorkshttps://www.mathworks.com/products/get-matlab.htmlToolboxes needed to run RpEGEN: Image Processing Toolbox, Curve Fitting Toolbox, Statistics and Machine Learning Toolbox
MATLAB supportMathWorkshttps://www.mathworks.com/support.html

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