Precise Cellular Ablation Approach for Modeling Acute Kidney Injury in Developing Zebrafish

This study was carried out in accordance with the recommendations in the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health. The protocol was approved by the NYIT College of Osteopathic Medicine Institutional Animal Care and Use Committee (NYITCOM IACUC). All surgery and in vivo experimentation was performed under tricaine anesthesia, and all efforts were made to minimize suffering.

1. Obtaining and Maintaining Embryos

1. Obtain embryos by crossing kidney GFP fluorescent zebrafish.
   NOTE: Examples of fluorescently labeled zebrafish transgenic lines are listed in Table 1.
2. Keep the embryos at 28.5 °C in E3 solution during the 24 h post-fertilization.
3. After 24 h, replace the medium with an E3 solution that contains 0.003% 1-phenyl-2-thiourea (PTU).
   NOTE: PTU is used because it blocks the tyrosinase-dependent steps in melanogenesis and hence prevents pigmentation. Here, this solution is referred to as E3-PTU.
4. Raise the fertilized embryos at 28.5 °C until they are >6 dpf, at which point the kidneys have matured.
   NOTE: It is best to use larvae in the 7-9 dpf window.
5. Using a fluorescent dissecting microscope at 40X magnification and green fluorescence emission filters, select the larvae with bright kidney GFP fluorescence.

2. Mounting the Zebrafish for Live-imaging

1. If photoablating the embryos prior to 3 dpf, remove the chorion from any unhatched zebrafish before mounting the zebrafish for imaging.
   1. Manually dechorinate with forceps (preferred) or use a chemical treatment with a protease mixture.
      NOTE: Dechorination allows for the best imaging results.
2. Rinse out the chorion debris in the petri dish using E3-PTU solution and refill the dish with E3-PTU.
3. Prepare 1-2% Low-Melting Point (LMP) agarose with 0.2 mg/mL tricaine solution to mount the zebrafish for kidney photoablation and live imaging.
   1. If LMP agarose was already prepared beforehand, reheat it in the microwave to melt it.
4. Once the LMP agarose is prepared, place a 35 mm plastic petri dish on a dissecting microscope. Place a pulled glass probe nearby to properly orient the anesthetized larvae.
   NOTE: It is important to do this step in a timely fashion and to have everything in place, because the embryos must be oriented before the agarose solidifies in about 1-3 min.
5. Before transferring the embryo to the agarose, make sure that the agarose is cool (i.e. at about body temperature). Use a plastic or glass transfer pipette to place the embryo into the cooled-down melted agarose-tricaine solution.
   NOTE: This is done because agarose that is too hot can be detrimental to the embryo, leading to cardiac arrest or death
6. Using a transfer pipette, redraw the embryo in the agarose solution and position the embryo/larva in the middle of a 35 mm dish.
7. Quickly spread the agarose containing the embryo/larva to evenly cover the bottom of the 35 mm dish; the best volume of agarose to use is ~1-1.1 mL.
8. Use the glass probe to orient the embryo in the best possible position for photoablation and imaging.
   NOTE: Movie 1 shows the best orientation of the embryo/larva for unilateral photoablation. This step is the most time-sensitive and should be performed quickly before the agarose starts to gel, as any attempt to reorient the fish after gelling begins is detrimental.
   1. Orient the larva so that the kidney segment of interest is perpendicular to the beam path while the contralateral segment is away from the laser beam (see Movie 1).
      NOTE: This can be achieved by turning the larva as shown in Movie 1.
9. Allow about 15 min for the agarose to solidify. Cover the petri dish to minimize evaporation. Once the agarose has solidified, the embryo or larva is ready for photoablation.

3. Laser Ablation

1. Operate the confocal imaging system.
   NOTE: When using the referenced imaging platform (see the table of materials), the recommended settings include 40X magnification, a 0.8 NA water dipping lens, and the first dichroic mirror set to allow both 488 & 405 nm illumination of the sample. The detection should be performed on the green channel.
2. Position the dish containing the immobilized zebrafish on a confocal stage.
   NOTE: This procedure is optimized for the upright configuration using a 40-60X water dipping lens. However, it should be possible to modify the procedure for an inverted microscope.
3. In the upright configuration, position the petri dish containing the embryo on top of a microscope slide and keep it in place using modeling clay (e.g., plasticine). Position the glass slide with the petri dish onto the stage of the confocal microscope.
4. Add 3 mL of the imaging solution E3-PTU (see step 1.3) with 0.2 mg/mL tricaine.
   NOTE: The imaging solution can also be added immediately before placing the slide on the stage. The addition of imaging solution should be done very carefully to prevent the agarose from floating off the bottom of the dish.
5. Switch to the water dipping lens (40 or 60X). Slowly raise the stage, making sure that the lens is slightly angled to prevent air from becoming trapped in the beam path. Once the lens touches the solution at an angle, center it so that it is in the field of view of the embryos.
6. Find the segment of interest using the fluorescence light source. Superficially position the branch targeted for injury to minimize light scattering (see Movie 1).
   NOTE: The subsequent steps make use of referenced software (see the Table of Materials), but the procedure can be easily adapted to other platforms.
7. Open the software. Navigate to "View"| "Acquisition Control"| "C2plus Compact GUI" and set "pixel dwell time" to "1.9 µs," "frame size" to "512 x 512 pixels," and "pinhole size" to "90 µm." Set "405 nm (or 408 nm in some systems) laser intensity" to zero and "488 laser intensity" to "low" (preferably <1%). Adjust the "gain" to obtain a good dynamic range but not to saturate the signal.
   NOTE: These values are not critical and are used for consistency. When a 405 nm laser is used, it will result in increases in GFP fluorescence. To avoid saturation of the signal during photobleaching (step 3.11), the signal gain value should be scaled down on the green channel. The blue channel gain can be left at zero, as it is not used for sampling.
8. Click "Scan" in the software interface to scan the kidney using a 488 nm laser. Find the segment of interest that is perpendicular to the beam path and that is well-visualized, with good GFP fluorescence. Once that region has been identified, draw a region of interest using an elliptical Region-of-Interest (ROI).
   1. Navigate to "ROI" | "Draw Elliptic ROI."
      NOTE: This will be used to continuously measure average GFP intensity while the photobleaching takes place, permitting the administration of a precise dose of laser treatment.
9. Navigate to "View" | "Acquisition controls"| "C2plus Scan Area" and choose "Band Scan Area." The rectangular mask will appear within that interface. Manually define a rectangular scan window using the cursor (i.e. drag, resize, and turn the mask) to cover the segment targeted for laser ablation. Right-click within the mask area to accept the selection.
   NOTE: Only the selected region will be scanned.
10. Begin the time measurement while monitoring the green channel using just the 488 nm laser to activate GFP. Ensure that the intensity of the 488 laser is relatively low (~1%). Measure the average intensity of the GFP in the region of interest by selecting "Measure" | "Time Measurement." Use the "Graph" or "Data" tabs to monitor the average intensity values within the ROI.
    NOTE: The rate of acquisition is defined by pixel dwell time and the size of the rectangular window set in step 3.9, but in general, it allows for the fast monitoring of the signal (~2-10 frames/s).
11. Induce damage to the kidney cells with the violet laser (405 nm) by increasing the intensity to 100% by sliding the "laser power" control all the way to the right. The 488-nm laser can remain active if the intensity is low.
    1. Observe a line graph using the "Graph" tab or the numerical values of GFP intensity using the "Data" tab and wait until it drops to a desired level, for example 50% of baseline (as shown in Figure 1).
       NOTE: A 20 mW laser has been used as part of the referenced laser unit (see the Table of Materials); maximal intensity may vary between systems. Lower intensity can be compensated for by longer exposure (i.e. using percent GFP bleaching as a measure of total exposure).
12. Once the GFP intensity within the elliptical ROI (step 3.8) has dropped to a desired level, immediately decrease the intensity of the violet (405 nm) laser to "0" by moving the "laser power" slider all the way to the left in the software control panel.
    1. Obtain a new baseline of GFP fluorescence. Confirm the ablation by obtaining a ratio of X/Y.
       NOTE: See Figure 1B; Y = average intensity before the ablation and X = average intensity after the ablation, using an average of 4 consecutive samples within the ROI. The exact target ratio (50%, 60%, etc.) depends on the amount of injury that is desired for a given experiment.

4. Time-lapse Microscopy with Propidium Iodide Staining

1. Apply general time lapse considerations (outlined in a previous study¹⁵) to visualize propidium iodide staining as a correlate of kidney cell injury after photoablation.
2. Pre-incubate the larvae in 30 µM propidium iodide solution (1% DMSO in E3-PTU) for 3 h prior to embedding and photoablation, as described in steps 2 and 3.7, respectively.
   NOTE: No additional propidium iodide is required in the agarose or the imaging solution.
3. Induce segmental kidney injury, as described in steps 3.1-3.12.
4. Obtain time lapse image stacks, as described in a previous study¹⁴, using a 488 nm laser (GFP) and a 461-nm laser (Propidium Iodide, PI).
   NOTE: The two colors are superimposed in confocal stacks to correlate the disappearance of GFP and the appearance of propidium iodide staining.
5. Set the parameters for obtaining the time-lapse image stacks as follows: virtual slice thickness = 4 µm, z-stack interval = 2 µm, pixel dwell time = 1.9 µs, image dimensions = 512 x 512, and averaging = 2.
   NOTE: These parameters allow for continuous, 10  to 15 min interval recording of up to 4 larvae and ensure that there is minimal photobleaching of the sample.
6. Use imaging software compatible with the recording file format to visualize and analyze the data.