Monitoring On-Target Signaling Responses in Larval Zebrafish – Z-REX Unmasks Precise Mechanisms of Electrophilic Drugs and Metabolites

Zebrafish husbandry and handling procedures at Cornell University (United States) were performed following the guidelines of the National Institutes of Health (NIH) and approved by the Institutional Animal Care and Use Committee (IACUC). Zebrafish husbandry and handling procedures at the zebrafish unit of the Swiss Federal Institute of Technology Lausanne (EPFL, Switzerland) were performed following Animal Welfare Act SR 455 and Animal Welfare Ordinance SR 455.1, with cantonal veterinary authorization VD-H23.

NOTE: In this protocol, Tg(lyz:TagRFP) and Tg(mpeg1:EGFP) fish lines expressing Halo-TeV-Keap1 are used to demonstrate Z-REX. The method can be extended to other proteins of interest, transgenic reporter fish lines, and downstream biological assays. Refer to Supplementary Table 1 for the buffers used in this study. All the reagents, instruments, equipment, antibodies, plasmids, zebrafish strains and equipment are listed in the Table of Materials.

1. mRNA preparation

1. Prepare Halo-TeV-Keap1-2xHA and Halo-2xHA-P2A-Keap1-2xHA mRNA using the mMessage mMachine SP6 in vitro transcription kit.
   NOTE: Follow the manufacturer's instructions and perform 40 µL scale reactions for each mRNA. Redissolve the mRNA pellet in 10 µL of nuclease-free water.
2. Assess the mRNA quality and measure the concentration by microvolume spectrophotometer and agarose gel electrophoresis. An mRNA of good quality should have an A₂₆₀/A₂₈₀ ratio of around or above 2.0.
3. Dilute the mRNA to 1-1.5 mg/mL with nuclease-free water.
4. Aliquot the mRNA solution (1-2 µL per tube), and store the aliquots at -80 °C.

2. Producing fish embryos

1. Option 1: Producing wild-type (WT) fish embryos.
   1. Set up 10 fish crossing pairs in 10 separate tanks, with each tank containing a divider between the male and female parent fish.
      NOTE: A total of 10 crossing pairs typically provides a sufficient number of embryos for the assays. The number of crossing pairs can be adjusted according to the experiment design/need and the fecundity of the parent fish.
   2. The following morning, after setting up the injector, remove the dividers in five of the tanks. Wait for 30 min for the fish to mate.
   3. Move the parent fish to another tank, collect embryos by passing the tank water through a strainer, and then rinse out the embryos from the strainer into a 10 cm Petri dish. These embryos will be used for the first round of injections.
      NOTE: If the egg quality from a certain batch is poor (e.g., eggs are opaque due to aggregation of proteins), do not pool them with the other embryos.
   4. (Optional) Perform similar procedures as described in steps 2.1.2-2.1.3 on the other five tanks for the next round of injection.
      NOTE: It is best only to perform one round of injection, to minimize the age difference between embryos. However, should more embryos than can be injected in one lot be required, performing two rounds of injection is recommended to ensure the embryos remain at a 1-4-cell stage during mRNA injection. The number of injection rounds can be adjusted according to the operator's injection skill and experiment design. However, it is suggested to perform the whole procedure (from the removal of the first divider to the injection of the last embryo) within 2 h. A large age difference across embryos may impair the reliability and reproducibility of the experiment results.
2. Option 2: Producing heterozygous transgenic neutrophil/macrophage reporter fish embryos.
   1. Set up 10 fish crossing pairs in 10 separate tanks, with each tank inserted with a divider between male and female parent fish: WT fish versus Tg(lyz:TagRFP), or WT fish versus Tg(mpeg1:eGFP).
      NOTE: Avoid in-crosses between heterozygous transgenic fish, which may affect downstream fluorescent readouts as homozygous reporter fish show a higher fluorescent signal compared to heterozygous fish. Transgenic reporter lines and WT embryos can be easily distinguished when imaging. Having a mixture of WT and heterozygous embryos in the same pool is not a problem. lyz:TagRFP reports neutrophils, and mpeg:eGFP reports macrophages. This protocol can be applied to other reporter fish lines as well.
   2. Follow steps 2.1.2-2.1.4.

3. Microinjector setup

1. Turn on the air source, set the back pressure to 0.2-0.5 psi, and set the injection pressure to 25-30 psi. The specific range of pressure shown is typically recommended.
   NOTE: It is essential to have a stable back pressure to prevent fish media backflow into the needle. When calibrating the injection volume in step 3.8, only the injection time should be changed. Do not change the injection pressure in the following steps; low injection pressure can lead to injection failure due to surface and interfacial tension, whereas high injection pressure can damage embryos.
2. Clean the equipment and injection platform with RNase decontamination solution.
   NOTE: RNase, which degrades mRNA, could come from the operator or the equipment. It is necessary to perform the cleanup before the experiment, and wear gloves.
3. (Optional) If co-injecting mRNA and morpholino, premix the two in a 0.2 mL tube.
   NOTE: Z-REX works well by using Halo-TeV-Keap1-2xHA mRNA solution with a concentration of 250-1500 ng/µL. Several morpholinos have also been used in zebrafish, and the optimal concentrations have been reported⁷. If using a morpholino with an unpublished sequence, the operator should first assess the toxicity and gene knockdown efficiency of the morpholino before using it in Z-REX.
4. Transfer 1-2 µL of mRNA (and/or morpholino where applicable⁷) into an injection needle with a micro-loader pipette tip.
   NOTE: If preparing needles with a Flaming/Brown micropipette puller, the setup is as follows. Heat: 520 units; pull strength: 60 units; velocity: 70 units; delay: 155 units; pressure: 550 units; ramp: 530 units.
5. Install the needle on the microinjection manipulator.
   NOTE: Back pressure from the air source should push the mRNA(/morpholino) solution to the needle tip.
6. Use sharp forceps or a razor blade to break the needle tip, creating a suitable opening for injection.
7. Submerge the needle tip in mineral oil on a stage micrometer.
8. Apply two or three injection pulses to remove air bubbles in the tip.
9. Calibrate the drop size to 2 nL by changing the injection time.
   ​NOTE: This is best performed by injecting into mineral oil (which mimics the viscosity of the yolk sac) laid upon a hemocytometer. Using a microscope, use the gridlines of the hemocytometer to estimate the size of the droplet formed during injection, and adjust the injection time accordingly. Although phenol red dye is sometimes used, the need for it in the mRNA injection procedure described here has not been observed.

4. Microinjection

1. Fill an injection plate with fresh 10% Hank's balanced salt solution (HBSS) medium, and align the embryos in the grooves of the plate with blunt forceps.
   NOTE: The injection plate is prepared with 2% agarose in 10% HBSS medium; the grooves are formed using a plastic mold.
2. Submerge the needle tip in 10% HBSS medium in the injection plate.
3. Apply two or three injection pulses to remove air bubbles in the tip.
4. For each injection, penetrate the chorion and yolk sac in a single move and apply an injection pulse. This injected liquid can be seen right after the injection as a small spheroid within the yolk sac. This small spheroid dissipates relatively rapidly. Repeat this step for other embryos, until a sufficient number of injected embryos has been obtained.
   NOTE: The survival rate of embryos (both injected and non-injected) typically varies from 50%-90%. Aim to inject double the number of embryos needed for each control/experimental group. In the biotin pull-down assay, 100-140 viable embryos are required for each condition. In the qRT-PCR assay, five viable embryos are recommended for each condition. The sample size for live fish imaging and whole-mount immunofluorescence staining assay is user-defined; having at least 20 viable embryos per condition is recommended to yield good statistical power in the analysis.
5. Rinse out the injected embryos to a new 10 cm Petri dish containing fresh 10% HBSS media.
   NOTE: The embryos can be easily rinsed out from the grooves using a squirt bottle.
6. Pool the non-injected embryos into another plate.
   ​NOTE: The non-injected embryos can serve as quality controls for the health of the fish, baseline protein expression, background fluorescence levels, etc., as needed. If the injection procedure worked well, and injected mRNA/morpholino is not lethal to the embryos, injected and non-injected embryos should have similar viability.

5. Z-REX

1. Distribute the injected embryos into 10 cm dishes, according to the experiment setup (i.e., numbers of control/experiment groups).
2. In a dark room with red-light illumination, replace the media with 30 mL of 10% HBSS with or without 1 µM Ht-PreHNE(alkyne).
   NOTE: Ht-PreHNE(alkyne) is a light-labile compound. Embryos should be kept in the dark in the following steps.
3. Incubate the embryos at 28.5 °C in the dark.
4. At 30.5 hpf, in a dark room, replace the media with a fresh 30 mL of 10% HBSS.
   NOTE: When replacing the medium, remove as much of the old medium as possible. This is critical for removing the unbound/excess amount of Ht-PreHNE(alkyne) from the embryos.
5. Incubate the embryos at 28.5 °C in the dark for 30 min.
6. Repeat steps 5.4-5.5 two more times.
7. Turn on the UV lamp (365 nm, 3 mW/cm²) for 5 min to prewarm the lamp.
   NOTE: The lamp prewarming step must be performed before step 5.8. The lamp power is lower/unstable in the first few minutes after being turned on. The lamp power should be measured by a UV meter regularly.
8. At 32 hpf, expose the embryos to UV light.
   1. Option 1: For downstream readouts, such as live fish imaging, whole-mount immunofluorescence staining, in-gel fluorescence analysis (click coupling with Cy5 azide), RNA-seq, or qRT-PCR, expose the embryos to UV light for 3 min, swirling the plates every 30 s.
   2. Option 2: For downstream readouts, such as biotin pull-down assay, expose the embryos to UV light for a maximum of 5 min (and minimum of 3 min), swirling the plates every 30 s, and chill the plates on ice for 1 min.
      ​NOTE: If using different probes, the light exposure time needs to be optimized, depending on the t_1/2 of photouncaging for a given photocaged electrophile probe and light source deployed. t_1/2^{photouncaging} can be determined using known procedures⁸. For Ht-PreHNE(alkyne), t_1/2 is < 1 min³; thus, the indicated time above is sufficient.

6. Downstream assays

1. Option 1: Phenotypic assay. Live imaging of transgenic neutrophil/macrophage reporter
   ​fish lines, Tg(lyz:TagRFP) and Tg(mpeg1:eGFP) (Figure 2)
   1. Anesthetize the embryos by incubation at 4 °C for 10 min.
      NOTE: It is recommended to have at least 20 viable embryos per condition.
   2. Remove the unfertilized/dead embryos from the plate.
      NOTE: The unfertilized/dead embryos are cloudy/non-transparent, and can be visually identified. If a high death rate is seen, check back to the injection procedure or try to reduce the concentration of mRNA or morpholino.
   3. Dechorionate the embryos with sharp forceps. Hold the chorion with one pair of forceps, without touching the larval fish, and use the other pair of forceps to rip off the chorion.The embryo is fragile. Only touch the chorion when performing dechorionation.
      NOTE: It is common, especially in beginners, to damage some embryos during dechorionation. Therefore, always have more embryos than the minimum needed.
   4. Mount the embryos on a 2% agarose plate (prepared with 10% HBSS medium) and image the embryos with a stereomicroscope (bright field and respective fluorescent channels) (Figure 2A, C, G).
      NOTE: After Z-REX [combination of Halo-TeV-Keap1-2xHA mRNA injection and Ht-PreHNE(alkyne) treatment], the depletion of neutrophils (lyz:TagRFP) was found to be most significant at 36 hpf (4 h post Z-REX), whereas the reduction of macrophages (mpeg1:eGFP) was most significant at 34 hpf (2 h post Z-REX) (Figure 2E,F). Other time points can be used if using different reporter lines, mRNA/morpholino, or probes. The exposure time and/or gain must be optimized for visualizing single cells or the specific (ultra)structures desired.
   5. Count the neutrophil/macrophage number of each fish by ImageJ (NIH) (Figure 2B, D-F, H). Use the Freehand Selection tool in ImageJ to circle the whole fish, and use the option Find Maxima to count the fluorescent cells.
2. Option 2: Target-labeling assessment. Biotin azide-click coupling and biotin pull-down assay (Figure 3)
   1. Anesthetize the embryos by incubation at 4 °C. This usually takes 10 min.
      NOTE: To obtain enough fish lysate, 100-140 viable embryos are required for each condition.
   2. Remove the unfertilized/dead embryos from the plate.
   3. Perform dechorionation and deyolking. Perform the two manipulations by holding the chorion with a pair of sharp forceps, using the other pair of forceps to penetrate the yolk sac, and then separating the yolk sac while removing the chorion to allow the yolk proteins to come out.
      NOTE: It is critical to remove the yolk proteins in this step. The abundant yolk proteins in the sample interfere with later analysis.
   4. Transfer the deyolked embryos into a 1.5 mL tube.
      NOTE: Swirl the plate to center the deyolked embryos, which facilitates collection. The lighter chorion debris winnows away during this process.
   5. After the embryos settle down to the tube bottom, remove the supernatant, and add 1 mL of chilled HEPES-buffered saline (pH 7.6).
   6. Repeat step 6.2.5 two more times.
   7. (Optional) If not intending to proceed with the next step immediately, remove the buffer, flash freeze the samples in liquid nitrogen, and store them at -80 °C.
      NOTE: Deyolked fish pellets can be flash-frozen in liquid nitrogen and stored at -80 °C. Flash-frozen samples can be stored at -80 °C for up to 2 weeks.
   8. Resuspend the fish pellets in lysis buffer.
      NOTE: Lysis buffer (pH 7.6) is composed of 50 mM HEPES, 100 mM NaCl, 1% Triton X-100, 0.3 mM TCEP, 2x Roche cOmplete Mini EDTA-free protease inhibitors, and 0.1 mg/mL trypsin inhibitor from soybean. One embryo gives around 2 µg of lysate. Use 100 µL of lysis buffer for every 120 embryos. Two Roche cOmplete Mini EDTA-free protease inhibitors and trypsin inhibitors from soybean should be added to the lysis buffer just before use.
   9. Add 20% v/v zirconia beads to the tube.
   10. Vortex for 20 s, flash freeze in liquid nitrogen, and thaw in a 37 °C water bath.
   11. Repeat step 6.2.10 another two times.
   12. Centrifuge the solution at 21,000 x g at 4 °C for 10 min.
   13. Transfer the supernatant to a new, prechilled 1.5 mL tube.
   14. Measure the protein concentration by Bradford assay.
   15. Dilute the lysate to 1 mg/mL.
   16. For each condition, transfer 170 µg of lysate into a 2 mL tube.
   17. Mix the lysate from step 6.2.16 with 0.2 mg/mL TeV protease(S219V), and incubate the solution at 37 °C for 30 min.
       NOTE: For non-TeV protease-treated groups, simply mix the lysate with an equal voloume of lysis buffer to the TeV protease solution used in other groups.
   18. Prepare a 10x master mix for the biotin-azide click reaction: 10% w/v SDS, 10 mM CuSO₄, 1 mM Cu(TBTA), 1 mM biotin-azide, and 20 mM TCEP.
       NOTE: Add TCEP to the mix just before step 6.2.19.
   19. Add 8.5 µL of t-BuOH and 17 µL of 10x click reaction master mix to the (TeV protease-digested) lysate from step 6.2.17. Vortex, centrifuge, and incubate the solution at 37 °C for 15 min.
   20. Add another 1 mM TCEP to the solution, and then vortex, centrifuge, and incubate the solution at 37 °C for another 15 min. The incubation time for steps 6.2.19-6.2.20 is 30 min in total.
       NOTE: This supplement of TCEP, a reducing reagent for generating Cu(I), improves the click reaction efficiency.
   21. Add 600 µL of -20 °C ethanol to each tube, vortex the solution, and incubate it at -80 °C overnight.
       NOTE: The samples can be stored at -80 °C for 1 week; if not, proceed with the next step immediately.
   22. Centrifuge the solution at 21,000 x g at 4 °C for 1 h.
       NOTE: A pellet should form in the bottom of the tube after centrifuging, which is the desired fraction.
   23. Remove the supernatant, add 1 mL of -20 °C ethanol, vortex, and centrifuge the solution at 21,000 x g at 4 °C for 10 min.
   24. Repeat step 6.2.23.
   25. Remove the supernatant, add 1 mL of -20 °C acetone, vortex, and centrifuge the solution at 21,000 x g at 4 °C for 10 min.
   26. Remove the supernatant. Allow excess residual acetone to evaporate, though not completely to dryness.
   27. Resuspend the pellet in 100 µL of resuspension buffer (8% w/v lithium dodecyl sulfate [LDS], 1 mM EDTA in 50 mM HEPES saline, pH 7.6), vortex for 15 s, and sonicate until the pellet is dissolved.
   28. Centrifuge the solution at 21,000 x g at room temperature (RT) for 5 min.
   29. Transfer the supernatant to a new 2 mL tube, and add 1.5 mL of 50 mM HEPES saline, pH 7.6.
       NOTE: The final concentration of LDS in this step is 0.5%. Higher LDS concentrations may undermine the pull-down efficiency. Thus, although increasing the LDS concentration can aid the reduction of non-specific binding, it may also reduce the efficiency of the pulldown. Accordingly, it is recommended not to change the LDS concentration at this step.
   30. Collect the "input" sample (Figure 3): transfer 30 µL of 1 mg/mL lysate to a new 1.5 mL tube, and add 10 µL of 4x Laemmli sample buffer containing 6% β-Mercaptoethanol (BME). Flash freeze the solution, and store it at -80 °C.
   31. Transfer 100 µL of bed-volume streptavidin high-capacity resin to a new 2 mL tube. Add 1 mL of 0.5% LDS in 50 mM HEPES saline (pH 7.6), centrifuge at 1,500 x g at RT for 2 min, and remove the supernatant. Repeat the wash with another 1 mL of 0.5% LDS in 50 mM HEPES saline (pH 7.6).
   32. Transfer the solution from step 6.2.29 to the tube containing prewashed streptavidin high-capacity resin from step 6.2.31, and incubate the solution on an end-over-end mixer at RT for 4-6 h.
   33. Centrifuge the mixture at 1,500 x g at RT for 2 min, take 30 µL of the supernatant, and mix it with 10 µL of 4x Laemmli sample buffer containing 6% BME for the "flowthrough" sample. Then, remove the remaining supernatant.
       NOTE: "Flowthrough" samples can be analyzed by western blotting to check the streptavidin pull-down efficiency. It is essential to remove as much of the supernatant as possible to wash out the unbound proteins. First, remove most of the supernatant using a P-1000 pipette, and then remove the remaining supernatant using a P-20 pipette with a gel loading tip.
   34. Add 1 mL of 0.5% LDS in 50 mM HEPES saline (pH 7.6) to the resin, and incubate the mixture for 30 min at RT with end-over-end rotation.
   35. Centrifuge the mixture at 1,500 x g at RT for 2 min, and remove the supernatant.
       NOTE: Typically, 0.5% LDS is enough for removing most non-specific binding proteins. If non-specific binding signals are still seen in the later analysis, the LDS concentration in the wash buffer can be augmented.
   36. Repeat step 6.2.34-6.2.35 two more times.
   37. Add 40 µL of 2x Laemmli sample buffer containing 6% BME to the resin.
   38. Elute the bound proteins by incubating the mixture at 98 °C for 5 min.
   39. Centrifuge the mixture at 21,000 x g at RT for 5 min, and transfer the supernatant to a new 1.5 mL tube. This is the "elute" sample.
       NOTE: Directly loading solution containing resin from step 6.2.38 into the gel may affect SDS-PAGE analysis.
   40. Load 20 µL into each well of 10-lane 10% polyacrylamide gel, and run gel electrophoresis.
       NOTE: Run the gel at a lower voltage (120 V) until the dye front reaches the resolving gel, and change the voltage to 170 V. Stop the program after the dye front comes out.
   41. Perform western blotting with anti-HA, anti-Halo, or other antibodies detecting housekeeping proteins (Figure 3).
3. Option 3: Transcriptomic analysis. RNA-seq and qRT-PCR (Figure 4)
   NOTE: It is strongly recommended to use embryos laid within 15 min of one another for this assay. The age difference of the embryos significantly affects the assay results.
   1. Anesthetize the embryos by incubating them at 4 °C for 10 min, 2 h after Z-REX.
   2. Perform dechorionation with sharp forceps (step 6.1.3).
   3. (Optional) Perform segmentation with forceps (e.g., separate the head from the tail) if different segments are to be analyzed separately.
   4. If extracting RNA from a whole embryo, transfer three to five embryos into a 1.5 mL tube. If extracting RNA from the head or tail, transfer 10-12 dissected segments into a 1.5 mL tube.
      NOTE: It is recommended to perform the experiment with three to five biological replicates.
   5. Add 1 mL of TRIzol reagent and glass beads to the tube.
      NOTE: It was found that glass beads work better than zirconia beads for RNA extraction.
   6. Vortex the mixture for 30 s.
      NOTE: If not proceeding with the next step immediately, the solution can be stored at -80 °C for 1−3 weeks.
   7. Extract the RNA according to the manufacturer's instructions.
   8. Assess the RNA quality and concentration by microvolume spectrophotometer and agarose gel electrophoresis.
      NOTE: RNA of good quality should have an A₂₆₀/A₂₈₀ ratio of around or above 2.0.
   9. Either submit the RNA for sequencing, or treat 1 µg of RNA with amplification-grade DNase I, and reverse transcribe using superscript III reverse transcriptase and oligo-(dT)₂₀. Perform this step according to the manufacturer's instructions.
   10. Perform qRT-PCR and analyze the data by the ΔΔCT method⁹ (Figure 4B-D).
4. Option 4: POI expression and colocalization analysis. Whole-mount immunofluorescence staining assay (Figure 5)
   NOTE: Formaldehyde-fixed embryos are fragile. Avoid vigorous shaking, and handle with care.
   1. Dechorionate the embryos by following steps 6.1.1-6.1.3.
   2. Transfer the embryos to a 1.5 mL tube.
      NOTE: Each tube should have an equal number of embryos, and no more than 40 embryos.
   3. After the embryos settle to the bottom of the tube, remove the supernatant, and add 1 mL of phosphate buffered saline (PBS) (pH 7.6).
   4. Repeat step 6.4.3 one more time.
   5. Remove the supernatant, add 1 mL of 4% formaldehyde in PBS (pH 7.6), and incubate the tube at 4 °C overnight with gentle rocking.
      NOTE: The samples in formaldehyde solution can be stored at 4 °C for 1 week.
   6. Remove the supernatant, add 1 mL of -20 °C methanol, and incubate the tube on its side at -20 °C for at least 18 h.
      NOTE: The samples can be stored at -20 °C for 1 month or more.
   7. Remove the supernatant, and add 1 mL of PDT buffer (0.3% v/v Triton X-100, 0.1% v/v Tween-20, and 1% v/v dimethyl sulfoxide [DMSO] in PBS buffer).
   8. Repeat step 6.4.7, and incubate the tube at RT for 30 min with gentle rocking.
   9. Remove the supernatant, add 1 mL of blocking buffer (10% v/v heat-inactivated fetal bovine serum [FBS], 2% w/v bovine serum albumin [BSA], and 0.1% v/v Tween-20 in PBS buffer), and incubate the tube at RT for 1 h with gentle rocking.
   10. Remove the supernatant, and add 200 µL of primary antibody solution (diluted in blocking buffer).
   11. Remove the supernatant, add 500 µL of primary antibody solution (diluted in blocking buffer), and incubate the tube at 4 °C overnight with gentle rocking.
       NOTE: If using a new primary antibody, it is worth including some samples without primary antibody staining to serve as negative controls. However, ideally, morpholino-knockdown or engineered gene-knockout embryos, or embryos in which expression of the target protein has been stimulated, are more reliable means to validate the antibody.
   12. Remove the supernatant, add 1 mL of PDT buffer, and incubate the tube at RT for 30 min with gentle rocking.
   13. Repeat step 6.4.12.
   14. Remove the supernatant, add 1 mL of blocking buffer, and incubate the tube at RT for 1 h with gentle rocking.
       NOTE: The samples should be protected from light after this step, to prevent photobleaching of the fluorophore conjugated on the secondary antibody.
   15. Remove the supernatant, and add 200 µL of secondary antibody solution (diluted in blocking buffer).
   16. Remove the supernatant, add 500 µL of secondary antibody solution (diluted in blocking buffer), and incubate the tube at RT for 1.5 h with gentle rocking.
   17. Remove the supernatant, add 1 mL of PDT buffer, and incubate the tube at RT for 30 min with gentle rocking.
   18. Repeat step 6.4.17.
   19. Mount the embryos on a 2% agarose plate (made with PBS, pH 7.6) and image the embryos with a stereomicroscope (bright field and respective fluorescent channels) (Figure 5A,B,D,F).
       NOTE: If using the Leica M165 FC fluorescence stereomicroscope, use a magnification of 25x to obtain images with good resolution.
   20. Quantify/analyze the fluorescent signal intensity by ImageJ (NIH). Use the Freehand Selection tool in ImageJ to quantify the signal in the region of interest.