Gentle Isolation of Nuclei from the Brain Tissue of Adult African Turquoise Killifish, a Naturally Short-Lived Model for Aging Research

Animal care and animal experimentation were performed in accordance with the University of Southern California IACUC under approved protocols #21215. For any work using this protocol, it is necessary to obtain approval from the institution's IACUC prior to starting any research work on vertebrate animals.

NOTE: A complete run through the protocol starting from flash-frozen brain tissue (starting from step 3) should take ~2.5 h for six samples (Figure 1).

1. Dissect killifish brains

1. Humanely euthanize killifish using a solution of 1.5 g/L of methanosulfonate/tricaine (MS-222) in the system water.
   1. Use sharp scissors to rapidly decapitate killifish after all body and gill movement has ceased.
2. Dissect the decapitated head on a clean Petri dish as previously described¹⁹.
   1. Place the dish under a dissecting microscope (using an 8x-35x magnification range). Turn the head such that the branchiostegal rays are in focus
   2. Using scissors, cut through the dentary such that the branchiostegal rays and the cranium are revealed.
   3. Using forceps, hold down the branchiostegal rays on either side of the cranium. Use another pair of forceps to remove any remaining tissue fragments from the cranium. The V-shaped optical chiasm that connects the brain and the eyes should become visible.
   4. Cut the optical chiasm using scissors to detach both eyes from the brain. Use forceps to gently free the cranium.
   5. Invert the cranium over and use forceps to scrape the muscle from the dorsal side of the cranium.
   6. Use one pair of forceps to hold the cranium in place and another pair of forceps to gently remove the bones of the cranium, revealing the brain.
   7. The brain appears white and soft. Once visible, it is not required to remove all cranial bones. Remove the brain with clean forceps by gently lifting and scraping.
3. Flash freeze the brain by placing it into a microcentrifuge tube stored on dry ice.
   ​NOTE: The frozen brain tissue can be stored at -80 °C until all samples to be processed together have been collected to limit batch effects when processing for library generation. While samples stored at -80 °C cannot be stored indefinitely, this protocol has been successfully performed on killifish brains stored for up to 12 months with no notable drop in quality.

2. Prepare fresh buffers

1. Prepare nuclei wash buffer (2% bovine serum albumin [BSA] in 1x PBS, phosphate-buffered saline pH 7.4, and 0.2 U/µL RNase inhibitor) and store on ice. Prepare 250 mL of nuclei wash buffer for up to eight samples. For 250 mL, use 50 mL of 10% BSA stock solution, 25 mL of a 10x PBS stock solution, 1.25 mL of the 40 U/µL RNase inhibitor stock, and then bring volume to 250 mL with RNAse-free ddH₂O.
2. Prepare nuclei lysis buffer (10 mM Tris-HCl pH 7.4, 10 mM NaCl, 3 mM MgCl₂, 0.01875% v/v NP-40, and 0.2 U/µL RNase inhibitor) and store on ice. Prepare 10 mL of nuclei lysis buffer for up to eight samples. For 10 mL, use 100 µL of 1 M Tris-HCl pH 7.4 stock solution, 20 µL of 5 M NaCl stock solution, 30 µL of 1 M MgCl₂ stock solution, 18.75 µL of a 10% NP-40 stock solution, 50 µL of the 40 U/µL RNase inhibitor stock, and then bring the volume to 10 mL with RNAse-free ddH₂O.

3. Isolate nuclei

1. If using frozen samples, thaw them on ice for 10 min. If using freshly dissected brains, proceed directly to step 3.2
   NOTE: Although this protocol can work on single brains, smaller samples from young fish (5-6 weeks old) will have a poorer yield. Sufficient yields can always be obtained by processing two brains as one sample, regardless of the fish's age and sex (Table 1). This protocol has successfully isolated high-quality nuclei from fish aged 5-26 weeks, including animals from both GRZ and ZMZ1001 strains. There are no issues expected in applying this protocol to other strains.
2. Dounce the brain in 1 mL of ice-cold nuclei lysis buffer in a 2 mL Dounce homogenizer. Keep the homogenizer on ice and avoid generating bubbles while lysing the sample. Dounce at an approximate rate of 0.5 s/downstroke and 0.5 s/upstroke with a 180° twist per cycle. Dounce using the loose pestle for 15 strokes, or until the tissue/lysis buffer mixture appears homogeneous if additional strokes are needed.
   NOTE: Dounce homogenizers should be washed, rinsed with distilled water, and sterilized with standard dry autoclave sterilization cycles between uses.
   1. Dounce using the tight pestle for 30 strokes with small breaks, 30 s to 1 min, every 10 strokes. Dounce at an approximate rate of 0.5 s/downstroke and 0.5 s/upstroke with a 180° twist per cycle. Let the sample rest on ice in the Dounce homogenizer for 2 min.
      NOTE: This incubation time ensures adequate lysis but should be shortened if nuclear overlysis is observed.
3. Strain the sample through a 70 µm cell filter into a 15 mL conical tube precoated in 5% BSA.
4. Add 4 mL of nuclei wash buffer to the sample by pipetting the wash buffer over the 70 µm cell filter from step 3.3 and gently mix by inversion five times.
   NOTE: This allows for the recovery of nuclei that may be trapped on the filter and improves yield.
5. Centrifuge using a swinging bucket rotor at 500 x g, 4 °C, for 10 min. Discard the supernatant by gently pouring into a liquid waste receptacle. Return the sample to an upright position and remove the remaining wash buffer using a P1000 pipette.
   NOTE: The nuclei pellet is likely to be visible when starting from two brains in one sample. However, when processing single brains from young animals (5-6 weeks old), a lower yield is expected, and pellets may not always be visible. In this case, the supernatant should be carefully removed while assuming the position of the pellet at the bottom of the tube.
6. Immediately resuspend the nuclei in 1 mL of nuclei wash buffer with a wide bore P1000 pipette tip.
7. Bring the sample to 5 mL by adding 4 mL of nuclei wash buffer and gently mix by inversion five times. Centrifuge using a swinging bucket rotor at 500 x g, 4 °C, for 10 min, and discard the supernatant.
8. Resuspend the nuclei in 1 mL of nuclei wash buffer with a wide bore pipette tip and transfer to a flow cytometry (FACS) tube.
9. Add 300 µL of debris removal solution (Table of Materials) and mix thoroughly by pipetting with a wide bore tip until the mixture is homogeneous.
10. Gently overlay 1 mL of nuclei wash buffer on top of the nuclei solution prepared in step 3.9 using a P1000 pipette and holding the tube at a slight angle (Supplementary Figure 2). The debris removal solution fraction and the nuclei wash buffer fraction should form a clear interface if layered correctly.
11. Centrifuge in a swinging bucket rotor at 3000 x g, 4 °C, for 10 min. Completely discard the top two phases using a P1000 pipette.
    NOTE: The clear FACS tube will permit the visualization of the three distinct phases following centrifugation (top, interphase, bottom). To facilitate maximal debris removal, it is advised to be more aggressive than conservative in removing the top two phases (i.e., removing a small amount of the bottom phase is preferable to carrying over any of the top phases; Supplementary Figure 2). Additionally, the interphase layer may be too thin to visualize. In this case, remove the top (apparent) layer, which contains the interphase layer. The bottom layer that is retained should contain ~1 mL total volume.
12. Transfer the bottom layer to a 15 mL conical tube precoated with 5% BSA.
13. In the conical tube, bring the volume to 15 mL with nuclei wash buffer and invert three times to mix.
14. Centrifuge in a swinging bucket rotor at 1000 x g, 4 °C, for 10 min. Remove the supernatant carefully.
15. Immediately resuspend in 150 µL of nuclei wash buffer using a wide bore pipette tip.
16. Switch to a standard bore pipette tip set to ~300 µL. Uptake the entire sample and then attach a 40 µm on-tip filter to the end of the pipette tip, taking care not to expel the sample.
    NOTE: On-tip filters are necessary to obtain smaller volumes, which are needed to maintain sufficient nuclei concentration for downstream applications. For instance, for snRNA-seq using microfluidics, a minimum concentration of >300 nuclei/µL is recommended, although the optimal range is 700-1200 nuclei/µL¹⁸. Higher concentrations of nuclei can also be easily diluted if necessary.
17. Filter the sample by forcefully expelling the sample through the filter into a low DNA binding 1.5 mL microcentrifuge tube. The final elution volume should be 120-150 µL.
    ​NOTE: Some frothing may result from filtration. This does not appear to affect the nuclei quality.

4. Assess nuclei quality by microscopy

1. In a separate microcentrifuge tube, mix 10 µL of the filtered sample with 10 µL of 0.4% trypan blue solution by gentle pipetting. Samples require no extended incubation time and are ready for visualization immediately after mixing with trypan blue solution.
2. Deposit 10 µL of the stained nuclei into a chamber of a counting chamber slide.
   NOTE: Alternatively, deposit 10 µL of the stained nuclei sample onto a hemocytometer or microscopy slide and cover with a cover slip.
3. Assess the nuclei quality by light microscopy.
   NOTE: Lower magnifications should only be used to zoom in on nuclei, since nuclei defects will only be visually apparent at higher magnifications (i.e., 60x or higher). This step can be done quickly to assess the success of nuclei preparation and does not require secondary analysis beyond visual inspection. High quality, intact nuclei will have a sharply defined border²⁰ (Figure 2A). Poor quality nuclei will have disrupted nuclear membranes, patchy trypan blue staining near the nuclear membrane indicative of potential nucleic acid leakage, and/or evidence of blebbing²⁰ (Figure 2B). Healthy nuclei are expected to be 10-15 µm in diameter.

5. Quantify singlet nuclei, debris, and multiplet proportion by flow cytometry

NOTE: The specific terminology and interface of the flow cytometer software may differ based on the brand of the machine, but these steps should be easily adapted to other systems if required.

1. In a FACS tube, resuspend 10 µL of the filtered nuclei sample in 90 µL of the buffer recommended for flow cytometry for a 1:10 dilution. For this purpose, 0.22 µm filtered sterilized PBS, pH 7.2, with 2 mM EDTA and 0.5% BSA is used in this study.
   NOTE: Although this dilution will work for most nuclei preps, a lower dilution (e.g., 1:5) may be needed to sample enough nuclei for quality assessment in the case of a lower yield sample (i.e., <30 nuclei/µL in the diluted sample).
2. Stain the samples with propidium iodide (PI) at a final concentration of 1 µg/mL. No incubation time is required, and nuclei can be analyzed immediately.
   NOTE: Alternatively, stain the samples with a final concentration of 0.1 µg/mL DAPI. Note that DAPI should be read on a different fluorescence channel than PI (Vioblue-A V1-A channel; excitation at 405 nm, emission at 450/50 nm).
3. Analyze the samples on a flow cytometer as follows:
   1. Set up a workspace:
      1. Have the flow cytometer in Acquisition mode. Click on File, then select New Workspace from the drop-down menu. Under the Experiment tab in the left panel, enter the Name of Project and Sample ID into their respective fields.
      2. In the upper left toolbar, click on the New Analysis Window button (scatter plot icon). Click the box with three plots (Plot 4t) from the pop-up menu. Three plots will appear on the screen. Click on the Analysis Mode button in the upper left toolbar (A icon) so that it is gray and not orange.
   2. Set up analysis plots:
      1. Change the axes of the three plots as follows, ordered as X-axis and Y-axis, respectively:
         Plot 1: X-axis: FSC-A, Y-axis: SSC-A
         Plot 2: X-axis: HDR-T, Y-axis: SSC-A
         Plot 3: X-axis: PerCP-Vio700-A B3-A, Y-axis: SSC-A
      2. Change the axes by clicking the axis label and selecting the desired channel from the drop-down menu.
      3. Set the plots to be density color-coded by clicking the square gray "i" icon that appears next to the upper right corner of each plot. A Properties window opens with a panel of buttons on the left. In the Properties window, click on the Scatterplot icon (second from the top on the left of the panel). Click on OK.
         NOTE: The PerCP-Vio700-A B3-A fluorescence channel corresponds to an excitation at 488 nm, and emission in the 650-730 nm range.
   3. Set up instrument settings:
      1. Click on the Channels tab in the left panel. A list of all channels will appear with three boxes to the right. Set B3, FSC-A, and SSC-A to Hlog by clicking the down arrow on their respective first boxes on the right and selecting Hlog from the drop-down menu. Set the voltage of SSC to 640 V, FSC to 300 V, and B3 to 480 V by typing in these numbers in their respective middle boxes to the right and pressing the Enter key. Note the third box to the right is a toggle that can be used alternately for typing.
      2. Under the Trigger header, set the FSC trigger to 4 and the SSC and B3 trigger to 0 by clicking the down arrow, selecting them from the drop-down menu on the left, typing in the number in the box on the right, and then pressing the Enter key. Note that there is a toggle box on the right that can be used alternatively to typing.
         NOTE: PMT voltages need to be optimized for each particular flow cytometry machine, but the trigger for nuclei must be lower than what is typically used for cells. Furthermore, the axis scale should be set to h-log.
   4. Type 100 µL in the text box Sample Volume and 50 µL in the text box Uptake Volume on the left side of the flow software window. Verify that the Mix Sample parameter is set to Mix Gentle to avoid lysing nuclei at this step. Press the triangle within a circle (Play icon) on the bottom right corner to start flow.
   5. Verify flow quality:
      1. Use side scatter area (SSC-A) versus HDR-T to check for air bubbles or clumps in the sample. Ensure that this plot is a smooth continuum of points. Air bubbles or clumps will result in blank regions in the SSC-A versus HDR-T plot.
      2. Check the side scatter (SSC-A) versus forward scatter (FSC-A) plot to verify the voltages used are correct for the sample.
         NOTE: Unlike cells, nuclei will not fully resolve from debris in a side scatter vs. forward scatter plot. However, there should be a high-density cluster of events toward the middle of the plot corresponding to nuclei (warmer colored) with a lower density of events (cooler colored) in the background corresponding to debris.
   6. Analyze nuclei count and quality:
      1. Double click the SSC-A vs. PerCp-Vio-700A plot for an enlarged view. Click the button on the upper left toolbar with perpendicular lines (Quadrant) to select a quadrant gate. Then, click anywhere in the SSC-A vs. PerCp-Vio-700A plot and quadrants will appear inside the plot.
      2. Click anywhere on the quadrant dividing lines and slide the cursor to modify the quadrant placement. Place the quadrants such that the upper left quadrant contains the far left population (debris), and the upper right quadrant contains several teardrop-shaped clusters (nuclei) (see Figure 3 for gate setup and examples of Flow results for typical results with preps including or not including a debris removal step).
   7. Debris versus nuclei counts:
      1. Click on the square gray "i" icon in the upper right corner of the plot to open the Properties window. Click on the Region Functions tab; lists of plot regions and functions will appear.
      2. Under the Regions list, ensure a green check mark next to the top item, indicating the entire plot is selected. Under the Functions list, click on Count/µL to produce a green check mark. Click on OK, and a counts/µL metric will appear in each quadrant.
      3. To view raw counts and percentages, select Count and %-T from the Region Functions tab if desired. The counts/µL in the upper left and right quadrants correspond with the debris and nuclei yield, respectively.
   8. Singlet counts:
      1. The left-most teardrop-shaped cluster in the lower portion of the upper right quadrant represents singlets. Draw a polygonal gate around this population by clicking the polygon button (Polygon) on the upper left toolbar.
      2. Click a single time, slide the mouse to start drawing, and click again to finish a linear segment of the gate and begin the next one. Double click to finish the gate. Click the border of the gate and slide the mouse to reposition the gate.
      3. Click the vertices of the gate to modify the shape. Counts/µL of the gated population (singlets) will appear. This is the concentration of the nuclei prep with the initial dilution.
         NOTE: The forward scatter area versus height 1:1 linear relationship, commonly used to determine singlet rate when flowing cells, does not apply to nuclei and can be disregarded.
   9. Using the concentration of the diluted nuclei prep and the dilution factor, back-calculate the concentration of the initial prep (e.g., for a 1:10 dilution, multiply the observed concentration by 10). Concentrations from >300 nuclei/µL are suitable for snRNA-seq.