A Neurite Outgrowth Assay and Neurotoxicity Assessment with Human Neural Progenitor Cell-Derived Neurons

Ethics Statement: Fetal specimens were received from the Birth Defects Research Laboratory at the University of Washington in Seattle through a tissue distribution program supported by the National Institute of Health (NIH). The Birth Defects Research Laboratory obtained appropriate written informed consent from the parents and the procurement of tissues was monitored by the Institutional Review Board of the University of Washington. All the work was performed with approval by the Human Subject Research Office at the University of Miami⁸.

1. Isolation and culture of human neural progenitor cells (hNPCs)

1. Place the brain tissue in a 100 mm Petri dish and carefully remove the meninges using forceps.
2. Transfer the brain tissue to a 50 mL conical tube and wash it twice with 20 mL of PBS by gently inverting the tube.
3. Incubate the brain tissue in a new 50 mL conical tube by submerging the tissue in cell dissociation solution (see Table of Materials) and DNase I (10 U/mL) for 10 min at 37 °C.
4. Add 5 mL of neuronal cell culture medium (see Table of Materials) to the brain tissue containing tube and mechanically dissociate the neurospheres by triturating 20 to 30 times through a 1000 µL pipet tip to make a single-cell suspension.
5. Filter the cell suspension through a 70 µm cell strainer to remove cell clusters.
6. Seed the single-cell suspension in a vented T-25 flask provided with 5-10 mL of neuronal cell culture medium supplemented with components detailed in Table 1.
   1. Sterilize the heparin solution by filtration through a 0.2 µm filter. The B-27 supplement without Vitamin A is a serum-free supplement for the cultivation of neural progenitors and stem cells, without inducing differentiation.
      NOTE: Human neural progenitor cells (hNPCs) are isolated from human fetal brain collected from aborted fetus. Following 7–10 days in culture, neural stem cells (NSCs) form free-floating neurosphere colonies, whereas other cell types remain in suspension as single cells or attach to the bottom of the flask. The isolated hNPCs can be cultured as neurospheres in suspension for several months^8,16,17.

Table 1. Components required for making 100 mL of culture media

2. Passaging the hNPCs

1. Collect the media containing the floating spheres, big and small neurospheres, and transfer them to a 50 mL conical tube.
   NOTE: Due to unknown reasons, the timing for splitting the neurospheres is variable from 7 days up to 30 days. However, generally, neurospheres need to be passaged when they reach a diameter greater than 700-900 µm. This is when the center of the neurosphere starts to darken, which is considered as a sign of a high rate of cell death¹⁸.
2. Spin the neurospheres down by centrifugation at 300-400 x g for 3 min.
3. Carefully aspirate the supernatant and then submerge the spheres in 500 µL of defrosted cell dissociation reagent (see Table of Materials).
   1. Make aliquots of cell dissociation reagent by adding 500 µL per 1.5 mL microtubes and store at -20 °C. To avoid losing enzyme activity, thaw the cell dissociation reagent by holding at RT or warm for 5 min in a 37 °C water/bead bath.
4. Depending upon the density and size of the spheres, incubate the submerged spheres at 37 °C for 5-15 min.
5. Add 5-10 mL of pre-warmed culture media to the neurospheres containing 50 mL conical tube and centrifuge at 300-400 x g for 5 min to sediment the neurospheres.
6. Aspirate the supernatant and gently pipette up and down, using a 1000 µL pipette, in 2 mL of culture media until all the neurospheres are in a single cell suspension.
   NOTE: The dissociation will become visible to the naked eye. Before dissociation, the neurospheres are in the form of spheres. After submerging in dissociation reagent and by pipetting up and down, they will become single cells.
   1. Count the cells and plate the single cells, 2 to 3 million cells per T-25 flask in 10 mL of culture media.
7. Feed the cells every 3 days by replacing half the culture media.
   1. Settle neurospheres by leaning the flask so that it is on its bottom corner. Hold the flask in the position for about 1-2 min until the neurospheres sediment. Then aspirate half the media gently by inserting the serological pipette in the media above the settled neurospheres. Dissociated cells can aggregate to form spheres after 2 to 3 days in culture¹⁹.

3. Freezing the hNPCs

1. Prepare the cell freezing medium by adding DMSO to the culture media to a final concentration of 10% (v/v) or use commercially available cryopreservation medium for sensitive cell types (see Table of Materials).
2. Sort out big spheres by transferring the media into a 50 mL conical tube and letting the spheres settle by gravity. Then remove and transfer the big neurospheres into a new 50 mL conical tube for passaging using a 200 or 1000 µL pipet tip.
   NOTE: Neurospheres with a diameter greater than 900 µm are considered big and the ones with a diameter smaller than 500 µm are considered small.
3. Spin the remaining neurospheres down by centrifugation at 300-400 x g for 3 min. Carefully remove the supernatant.
4. Resuspend up to 100 spheres in 1 mL of cryopreservation reagent and transfer it to a cryotube.
5. Store overnight at -80 °C in a cell freezing container (see Table of Materials) and move it to liquid nitrogen the next day for long-term storage.
   NOTE: It is preferable to freeze small to medium-sized neurospheres (lower than 900 µm in diameter) and avoid freezing big-sized neurospheres (greater than 900 µm in diameter) or single cells. In order to reduce cell damage during thawing the sample, keep the cells dense by seeding the thawed neurospheres into a small T25 flask.

4. Differentiation and treatment of hNPCs

NOTE: To induce differentiation, neurospheres are disaggregated into single cells, counted and then seeded on coated plates for 5 days. Then differentiated cells are treated for 24 h with test compounds before immunostaining and fluorescence quantification.

1. Coating
   1. Add 200 µL of poly-L-lysine (PLL) per well of 4-well glass chamber slides (140 µL per well of 8-well chamber slides).
   2. Incubate for 1 h at room temperature (RT).
   3. Wash 3x with PBS.
   4. Let it dry at RT (for about 30 min).
   5. Add 150 µL of laminin (50 µg/mL) per well of 4-well glass chamber slides (120 µL per well of 8-well chamber slides).
   6. Incubate for 2 h at 37 °C.
   7. Wash 3x with PBS.
      NOTE: Coated chamber slides can be stored at 4 °C for 1 month.
2. Plating the cells
   1. Count and plate 80,000 single cell neurospheres (neurospheres in a single cell suspension) per well of 4-well chamber slides (70,000 cells per well of 8-well chamber slide).
   2. Add 500 µL of differentiation media per well of 4-well chamber slide (250 µL media per well of 8-well chamber slides).
      1. To make the differentiation media first, add the following components in Table 2 to a sterile, disposable container to make the neural inducing media (NIM).
      2. Add the following ingredients in Table 3 to 48.5 mL of NIM made in the previous step to make the differentiation media.
   3. Incubate for 5 days at 37 °C.
   4. After 5 days, treat the cells for 24 h by replacing half the media in each well with fresh media mixed with the desired concentration of test compounds, including appropriate controls.

Table 2. Components required for making 50 mL of NIM

Table 3. Components required for making 50 mL of differentiation media

5. Immunocytochemistry (ICC)

NOTE: Cells are fixed with 4% formaldehyde. Permeabilization and blocking is then performed to improve penetration and prevent nonspecific binding of antibodies. Cells are then incubated with primary antibodies overnight. Subsequently, cells are incubated with fluorescently labeled secondary antibodies. Finally, after using DAPI to stain the nucleus, chamber slides are mounted.

1. Fixation
   1. Gently aspirate the media in each well.
   2. Add 500 µL of 4% formaldehyde per well of 4-well chamber slides (250 µL per well of 8-well chamber slides).
   3. Incubate for 15 min at RT.
   4. Gently wash 2x with 500 µL of PBS.
      NOTE: After fixation, by leaving 1 mL of PBS in each well, culture slides can be stored at 4 °C for up to 3 months.
2. Cell permeabilization and blocking
   1. Add the following components in Table 4 to a sterile, disposable container to make the antibody (Ab) buffer.
   2. Mix the following ingredients in Table 5 with the Ab buffer made in the previous step to make the cell permeabilization and blocking solution.
   3. Add 500 µL per well of 4-well chamber slides and incubate for 1 h at RT.
      NOTE: Ab buffer can be stored at 4 °C.

Table 4. Components required for making 200 mL of Antibody buffer

Table 5. Components required for making 3 mL of cell permeabilization and blocking solution

3. Staining
   1. Wash 2x with 500 µL of PBS.
   2. Add the diluted primary antibody, anti-β-tubulin III (1:200), and incubate overnight at 4 °C (200 µL per well of 4-well chamber slides). Dilute the primary antibody in PBS.
   3. Wash 2x with PBS.
   4. Add the diluted, in PBS, fluorescently labeled secondary antibody, Alexa Fluor 488 (1:500), and incubate for 2 h at RT in a place protected from light (250 µL per well of 4-well chamber slides)
   5. Wash 2x with PBS.
   6. Add the diluted, in PBS, DAPI (300 nM concentration) and incubate for 5 min at RT in a place protected from light (300 µL per well of 4-well chamber slides).
   7. Wash 3x with PBS.
   8. Mount the chamber slides using the following instructions.
      1. Take apart the chamber slide by breaking the breakaway tabs and removing the gasket and base.
      2. Add one drop of the mounting solution (see Table of Materials) per well of 4-well chamber slides. Then use a cover slide to cover the whole slide.
      3. Using a tweezer and at an angle, place one side of the cover slip against the slide while making contact with the outer edge of the liquid drop.
      4. Carefully tip the coverslip onto the mounting solution when lowering it into place. Avoid the creation of bubbles. Take a pipette tip and press it down on the cover slip. The bubbles will move to the side.
         NOTE: Bubble formation is inevitable at times. If it occurs, image around them as long as there are a few.
      5. Follow manufacturer’s directions for curing time. Avoid using noncuring mounting solutions due to the difficulty in handling during imaging. Otherwise using an appropriate coverslip sealant on edges is required to prevent the coverslip from sliding during imaging.
      6. To seal the coverslip, use nail polish and make a small line on the edge of the cover slip. Let the nail polish dry for about 2 min.

6. Image acquisition, neurite outgrowth and fluorescence intensity quantification

NOTE: Following staining, use a confocal microscope with a 20x objective and an image size of 1024 x 1024 pixels to acquire the images of the treated cells. Take image at least from two fields per biological replicate per condition. Then use Fiji image analysis software (ImageJ 1.51u) for quantification of the neurite length. Briefly, measure the length of the longest neurite for each neuron and after averaging the values per treatment, use student’s t test for independent groups to compare the means between experimental groups and control group.

NOTE: Several commercial (Imaris, Volocity, Amira) and open source (ImajeJ, CellProfiler, Vaa3D, BioImageXD, Icy, KNIME) image processing programs are available. Among these programs, ImageJ has become the tool of choice for biological image analysis^20,21. The ImageJ portal at https://imagej.net/Introduction is a useful source of information providing a thorough description of ImageJ’s basic, and built-in functions including image processing, colocalization, deconvolution, registration, segmentation, tracking, and visualization.

1. Measuring neuronal outgrowth with Fiji image analysis software
   1. As illustrated in Figure 1, open the image either by dragging and dropping it onto Fiji software or by selecting File | Open.
   2. Select Analyze | Tools | ROI manager, and then right-click on the 5^th icon in the toolbar Straight and switch to freehand line. Optionally, double-click on the same icon to change the line width to 10, and then trace the longest neurite, beginning near the cell body and extending to the tip of the neurite.
   3. Press Ctrl + T & then F to add the measurement to the ROI Manager and highlight the measured neuron. Select all numbers in the ROI, click on Measure, select all calculated lengths, and copy/paste into a spreadsheet.
2. Measuring fluorescence intensity of labeled cells with Fiji image analysis software
   1. As illustrated in Figure 2, after opening the image, click on the 4^th icon in the toolbar Freehand selections. Draw the shape of the cell.
   2. Select Analyze | Set Measurements and select for the following values: Area, Integrated density, Mean grey value | Analyze | Measure (a pop-up box with a stack of values opens).
   3. Select a region next to the cell as background (size is not important) and then select Analyze | Measure. Select all measured data and copy/paste into a spreadsheet.
      NOTE: Integrated Density (IntDen) is the item used for determining the fluorescent intensities. In this study fluorescent intensity of the target molecule is measured to initially analyze its distribution in the cell and subsequently quantifying the abundance of the target molecule under various treatments. Consequently, the effectiveness of treatments will be measured and compared with control.

7. Neurotoxicity assessment

NOTE: Cytotoxicity of test compounds are evaluated in 384-well plates (see Table of Materials) using a luminescent cell viability assay (see Table of Materials). The hNPCs are prepared following the same method, except slight modifications, described in the “Differentiation and Treatment of hNPCs” section. Subsequently, a luminescent signal generated by luminescent cell viability assay is measured utilizing a microplate reader. The luminescent signal is proportional to the cellular ATP concentration which itself is directly proportional to the number of viable cells present in each well.

1. Coating
   1. Add 30 µL of poly-L-lysine (PLL) per well of 384-well plate.
   2. Incubate for 1 h at RT.
   3. Wash 2x with PBS.
   4. Let it dry at RT (for about 30 min).
   5. Add 30 µL of laminin (50 µg/mL) per well of 384-well plate.
   6. Incubate for 2 h at 37 °C.
   7. Wash 2x with PBS.
      NOTE: Coated 384-well plates can be stored at 4 °C for 1 month.
2. Plating the cells
   1. Count and plate 20,000 single cell neurospheres per well of 384-well plate in 25 µL of differentiation media.
   2. Incubate for 5 days at 37 °C.
   3. After 5 days, treat the cells for 24 h by test compounds prepared at 6x the final desired concentration in 5 µL volume (to make the final volume of 30 µL per well).
3. Cell viability assay
   1. Add 30 µL of luminescent cell viability assay reagent per well of 384-well plate.
      NOTE: Add a volume of luminescent cell viability assay reagent equal to the volume present in each well. Thaw the luminescent cell viability assay reagent and equilibrate to RT prior to use.
   2. Shake on a plate shaker for 2 minutes (to mix and induce cell lysis).
   3. Spin the mixture down by centrifugation at 300-400 x g for 30 s.
   4. Incubate the 384-well plate for 10 min at RT in a place protected from light (to stabilize the luminescent signal).
   5. Record luminescence with a microplate reader.
      NOTE: Use appropriate controls for the viability assay including Velcade (at a final concentration of 10 µM) as positive and HBSS containing DMSO (with final concentration of 0.1% or 0.2%) as negative control.