A schematic of the protocols is illustrated in Figure 1.

1. Cell Plating

Plate cells in 96-well plates at different plating densities (Figure 2). For linearity checks on the N2a neuroblastoma cell line, plate 2.5k, 5k, 10k, and 15k cells per well in 3 or 6 wells/group. For linearity checks in rat primary cortical neurons, plate 25k, 50k, 100k, and 200k cells per well in 3 or 6 wells/group. If the cell lines or primary cells of interest look healthy at different plating densities, plate at and around the optimal cell density for that cell type.

Note: In the present study, N2a cells were plated in 100 μl media and primary cortical neurons in 200 μl media on plates that are designed for lower evaporation. For detailed information on cell handling, media, sera, antibiotics, and toxin treatments, please see Unnithan et al. for N2a cells⁸ and Posimo et al. for primary cortex cultures²².

1. 1. Repeat the plating on a second 96-well plate. One plate will be assayed for ATP (Figure 2A) and the other with the infrared assays (Figure 2B). One cannot use the same plate for the ATP and infrared assays because the cells must be lysed open for intracellular ATP measurements.
   2. Be sure to plate at least 3 extra wells for background subtraction in the infrared assays at the optimal plating density (column 2; Figure 2B).
2. Add media without cells or, more inexpensively, sterile water to the outer wells in rows A and H and in columns 1 and 12. Avoid relying on data from wells along the edges, as viability is often lower here from effects of media evaporation and temperature gradients. Such problems with microplates are commonly called edge effects^23,24. Do not keep these wells empty because then rows B and G and columns 2 and 11 suffer from the edge effect.
   Edge effects may be reduced by incubating a freshly seeded plate at room temperature in ambient conditions before transfer into the CO₂ incubator²⁴. Furthermore, a recent study by Carralot describes a mathematical correction method for microtiter plates using a single control column²³. Oliver and colleagues used a specially designed forced air microtitration plate incubator to reduce thermal gradients²⁵. If the edge effect is of significant concern, microplate stability chambers (e.g. BT&C Incorporated) can be purchased to create a homogenous microenvironment with high humidity and even temperature gradients.
3. If more wells than shown in Figure 1 are needed because additional reagent dilutions or more plating densities will be tested, use the edge wells for background subtraction.
4. Wait overnight for attachment of cells and assay the next morning as described below.

2. Luminescent ATP Assay

1. Follow the Cell Titer Glo manufacturer’s recommendations for reconstitution of the substrate with buffer and for incubation times.
   1. Remove 50 or 150 μl media from the 100 or 200 μl of plating media, respectively. Slightly less than 50 μl will remain behind in the well. Add 25 μl of the reagents (substrate plus buffer) to columns 2-6 (Figure 2A) in a 1:2 dilution.
      Note: Varying volumes of media left behind after removal could dilute or concentrate the ATP assay reagents differentially across wells. Take care to ensure that the level of liquid in all the multichannel pipette tips is equivalent. Measure the remaining liquid in select wells across the plate with a pipette immediately before adding the ATP assay reagents to ensure accuracy. If high variability exists from differential rates of media evaporation across the surface of the plate, remove all the old media and add the same volume of fresh media or phosphate buffered saline (PBS) to all wells immediately before addition of ATP assay reagents. If the plates suffer from variable levels of evaporation, try switching to the low evaporation plates from Costar Corning. In the latter plates, the 60 interior wells shown in Figure 2 only suffer from an average of 0.995%±0.41 media evaporation overnight. There is thus negligible variability in media volume at the time of assay.
   2. In columns 7 through 11, rows B through D, remove enough media to leave 50 μl behind, as detailed above, and add 50 μl of reagents in a 1:1 dilution.
   3. In columns 7 through 11, rows E through G, leave cells in 100 μl of media and add 100 μl of reagents, again in a 1:1 dilution. The company recommends that 100 μl of reagents should be diluted 1:1 in 100 μl of media.
      Note: In order to save on cost, it is possible to cut these recommendations both in volume and in dilution. However, before doing this, ensure that it does not reduce the linearity and sensitivity of the assay.
   4. Once one specific volume and dilution factor of the reagents are found to be satisfactory, stick with them in all subsequent experiments. The criteria for satisfactory data are described in the Results section.
   5. Unused reconstituted reagents can be refrozen and used at a later date to save on cost. According to the manufacturer, reconstituted reagents can be stored at -20 °C for 21 weeks with ~3% loss of activity.
2. Place the plate on an orbital shaker for 2 min or a nutating shaker for 10 min. If part of the plate is being assayed for a different measurement and it cannot be shaken, agitate the media with simple up-and-down pipetting only in the wells of interest. However, excessive bubbles generated during this step will interfere with luminescent output.
   1. 10 min after addition of the reagents, transfer 60 μl of the well contents into white 96-well plates. Luminescence values are higher in white plates than in clear or black plates because they reflect light upwards towards the detector.
      Note: In our experience, cells survive better on the low evaporation, clear Costar plates than other plates. These specific plates are not sold with white walls. Second, the opaque-walled and clear-bottomed plates are more expensive than completely clear plates. If one transfers the luminescent liquid to white plates, the white plates can be washed and reused, saving on the cost of using new white plates for every experiment. Transferring liquid is thus the cheaper option in the long run.
   2. Pop any air bubbles prior to reading the plate with a needle or, preferably, with forced air from a plastic transfer pipette bulb. Read the plate on a luminometer with a 1 sec integration time (the company recommends 0.25-1 sec as sufficient integration time). Read the plate 10-12 min after addition of ATP assay reagents. Timing is critical for comparison across different plates, because the luminescent signal is transient with a fast decay rate, as shown by Gilbert and colleagues²⁶.
3. Average the luminescent values from the 3 or 6 wells in each group and plot as a function of cell number in a scatterplot (see Figure 3). First subtract average background luminescence values of the appropriate empty wells (wells 2B – 2G, wells 11B – 11D, and wells 11E – 11G) from each corresponding data point. There will be three different groups for each plating density in this initial experiment (Figure 2A). ATP levels may be linearly correlated with cell densities in one or all of these groups. Proceed with the highest dilution of reagents that still gives satisfactory results to save on cost. Criteria for satisfactory results are described in the Results section.
   Note: With the media used in the present study, background values with this assay are not high and it is possible to skip the blank wells. However, the manufacturer Promega reports differences in luminescence with different culture media. The present study uses Hyclone Fetal Clone III, a synthetic version of fetal bovine serum for N2a cells and bovine calf serum for primary cortical cultures. According to the manufacturer, calf serum decreases luminescence values but does not decrease sensitivity of the assay. Nevertheless, if this is of significant concern, dilute ATP assay reagents in PBS instead of media at the time of assay.
   1. If the cells appear to grow unevenly across columns overnight, try assaying them within 6 hr of plating. However, bear in mind that the density at the usual time of assay (for example, 24 hr after treatment) is the density at which linearity must be achieved.

3. Infrared Assays

1. The morning after plating, fix the cells in the second plate at room temperature in a fume hood. Be sure to wear gloves for this step and dispose of the fixative as chemical waste because formaldehyde is a carcinogen. Add 4% formaldehyde and 4% sucrose in 0.1 M phosphate buffer to the existing media in a 1:1 dilution. The media can also be washed off with PBS before fixing in full-strength fixative. Either technique works well.
   1. Incubate in fixative for 20 min and then remove the fixative and wash 3x in 200 μl PBS.
2. Store the plate in PBS with 0.2% sodium azide at 4 °C if the assay will not take place on the same day. Otherwise, proceed with the assay and place the cells in blocking solution to minimize nonspecific binding of antibodies.
   1. Dilute the fish serum Odyssey block 1:1 in PBS and add 0.3 % Triton-X 100 as a cell permeabilizer. Make enough blocking solution as well as primary and secondary antibody solutions so that 35 μl can be pipetted into each well. However, if a lot of bubbles are observed during pipetting, make >50 μl for each well, otherwise the bubbles reach down into the cells and block antibodies from binding. Excessive soapy bubbles will appear as an unstained circular spot in the middle of the well. Try to adjust the pipetting if this occurs.
   2. Incubate in this blocking solution for 30-60 min at room temperature.
      Note: 5% bovine serum albumin or normal sera from the same species as the secondary antibody can also be used as blocking solutions to save on cost of the fish serum block. Blocking solutions can affect performance of the antibody. Thus, the optimal block for each antibody is best determined empirically.
      Note: Some investigators place In-Cell Western plates on shakers during incubations. However, this is not necessary.
3. Make the primary antibodies: 1:10,000 dilution for anti-α-tubulin (mouse monoclonal, Table 1) and 1:2,000 dilution for anti-MAP2 (mouse monoclonal, Table 1). These antibodies are specific to the proteins of interest in these cellular models; specificity is essential for any immunocytochemical stain.
   Note: MAP2 is a marker for neurons and is appropriate for mixed neuronal/glial cultures. The postnatal cultures shown here also contain some glia (~25%) because astrocytes appear first on embryonic day 18, with their numbers peaking in the early neonatal period^27,28. One cannot distinguish between astrocytes and neurons in the ATP and DRAQ5 + Sapphire assays. In order to measure neurons and astrocytes separately, use simultaneous MAP2 and GFAP staining in the 800 and 700 nm channels, as described by Mullett and colleagues^4,29, leaving out DRAQ5 + Sapphire. However, if all cells in the culture wish to be assessed, use a pan-cellular marker such as α-tubulin, β-actin, or GAPDH.
   Note: These dilutions have been optimized for the N2a and primary cortical cells and may not generalize to every model. Therefore, try at least two primary antibody dilutions, one on the left half of the plate and one on the right half (see Figure 2B). Alternatively, try 3-4 dilutions of primary antibodies on 3 wells each. For example, the recommended dilution on the antibody insert sheet can be flanked with two-fold changes. In other words, if the recommended dilution for immunocytochemistry is 1:500, also try 1:250 and 1:1,000 dilution factors.
   1. Dilute antibodies in 1:1 Odyssey block:PBS and add 0.3% Triton-X. To save on cost, try making the antibodies in the blocking solution that was applied to the cells in step 3.2.1 by removing this solution at the end of the incubation. Keep the cells in PBS while doing this; they must not dry out.
   2. Incubate in primary antibodies either 1-2 hr at room temperature or overnight at 4 °C. For weakly binding antibodies and proteins that are not abundant, overnight incubations can help increase signal.
      Note: Leave at least 3 wells in blocking solution for background subtraction at each secondary antibody concentration (wells 2B-2D and wells 2E-2G in Figure 2B). Do not expose these wells to any primary antibody whatsoever. They reveal the extent of nonspecific binding by the secondary antibody and are useful for calculations of signal-to-noise ratios. If there is a concern that the secondary antibodies will lead to high levels of nonspecific binding, also include control wells that are not exposed to either primary or secondary antibody but receive the same number of washes. The difference between these wells and the “secondary only” wells will reveal the extent of nonspecific binding caused by the secondary antibody alone. In our test on mouse N2a cells, signal intensity with anti-mouse secondary antibody alone was 0.557±0.032, signal with anti-rabbit secondary antibody alone was 0.533±0.041, signal with no secondary antibody was 0.357±0.003, and signal with primary and secondary antibodies was 11.867±0.911. We have thus not observed high levels of nonspecific binding even when using anti-mouse secondary antibodies on mouse cells. There are several manufacturer’s for infrared secondary antibodies; we recommend only buying the highly cross-adsorbed ones. Note that the concentration of secondary IgG antibodies may vary depending upon the source.
4. Wash off primary antibodies with 3 washes of 200 μl PBS per well, 10 min each. Primary antibodies can be saved at 4 °C for a few weeks in 0.2% sodium azide and reused until tiny specks of debris become apparent when the solutions are held up to light. This is only done to save on cost. If cost is not an issue, make fresh antibodies for each use.
5. Dilute the secondary antibody by 1:1,000 or 1:2,000 in 1:1 Odyssey block:PBS with 0.3% Triton-X (Figure 2B). Add the 1:1,000 dilution to the top half of the plate and 1:2,000 to the bottom half of the plate. These concentrations could be further reduced to save on cost, but check for linearity before committing to this.
   Note: Be sure to add the appropriate secondary antibody solution to the background subtraction wells (column 2 in Figure 2B).
   1. If a second protein will be assayed in place of DRAQ5 + Sapphire, label cells for α-tubulin or MAP2 with 700 nm goat anti-mouse IgG and the second protein in the 800 nm channel with primary antibodies from a species other than mouse. The 800 nm channel has less background than the 700 nm channel and should be reserved for the most critical protein of interest.
   2. Incubate in secondary antibodies for 1 hr at room temperature in a drawer away from light.
6. Wash off secondary antibodies with 3 washes of 200 μl PBS per well, 10 min each.
7. Make the DRAQ5 + Sapphire solutions. For the left half of the plate, dilute DRAQ5 1:10,000 (0.5 μM final concentration) and Sapphire 1:1000 in PBS with 0.3% Triton-X (columns 3 – 6; Figure 2B). For the right half of the plate, dilute DRAQ5 1:20,000 (0.25 μM) and Sapphire 1:2000 (columns 7 – 10; Figure 2B).
   Note: DRAQ5 used to be sold at a 1 mM stock instead of a 5 mM stock. Some previously published reports therefore used 1:4,000 or 1:2,000 dilutions of DRAQ5 ⁸.
   1. To save on cost, if two plates are being assayed simultaneously, the same DRAQ5 + Sapphire solutions can be used on two plates in sequence, pipetting it off the first plate and adding it to the second. If the same solutions will be used twice in this manner, use them up within one day. Diluted DRAQ5 + Sapphire solutions cannot be saved at 4 °C or frozen for later use.
   2. Incubate in these solutions for 30 min at room temperature away from light. If time is short and the DRAQ5 + Sapphire solutions will not be reused on other plates with different secondaries, add these stains to the secondary antibody solutions in step 3.5 and incubate for 1 hr.
      Note: Do not add DRAQ5 + Sapphire solution to the background subtraction wells (column 2 in Figure 2B) as these wells should not be stained in the 700 nm channel.
8. Wash the plate 3x with 200 μl PBS per well for 10 min each. Put 0.2% sodium azide in the final PBS wash if the plate is going to be stained with other visible-range secondary antibodies after the infrared imaging is complete.
9. Scan the plate on an Odyssey Imager. Begin by scanning the plates at intensity 5 and 169 mm resolution. Either “medium quality” or “low quality” settings are sufficient. The company does not release detailed excitation/emission filter information. However, the emission filters are approximately 20 nm wide and are centered around 720 and 820 nm, according to the manufacturer.
   Note: It is possible to scan plates while still wet as investigators may wish to continue to stain them with other markers. However, the company suggests in its online protocol that dry plates result in less well-to-well signal spread. If you scan them both wet and then dry to compare the data, be sure to remove all the PBS from the well because remaining salts after evaporation can lead to high background fluorescence along the edges.
   1. The Odyssey Imager scans up and through the bottom of the plate. Plates from different manufacturers may demand different focus offsets because the bottom of the wells can vary in their thickness and depth in the plate. Try scanning at different focus offsets and see where the highest signal-to-noise ratio and crispest (most in focus) signal is achieved: 2.5 mm, 3.0 mm, 3.5 mm, 4.0 mm. Try also to measure the depth of the well from the bottom of a plate with a ruler to confirm the findings on the Odyssey. Remember that the plastic in the bottom of the well can add additional height to the ruler measurements. The Costar plates used in the present study demand a focus offset of 4.0 mm.
   2. Use the In-Cell Western function in the software to place the right-sized grid onto the image of the plate and export the data into Microsoft Excel. Examine the “integrated intensity” values of the same wells across different focus offsets to find the highest fluorescence values. The focus offset that yields the highest signal-to-noise ratio (signal in immunostained wells versus “no primary negative control” wells) is the one to choose hereafter.
   3. Once one focus offset is decided upon, scan again in smaller increments. For example, if the brightest signal was at 3.5 and 4.0 mm, try scanning at 3.6, 3.7, 3.8, and 3.9 mm and see if there is any further improvement in signal strength. If the focus offset is wrong, signal intensities across the 12 columns on an empty plate (when all columns should have equal signal) will appear as a U-shaped curve instead of a flat line. If this continues to be a problem with plastic plates, try glass-bottomed plates.
10. Subtract the average of the integrated intensities in the background subtraction wells (Figure 2B; wells 2B – 2D or wells 2E – 2G) from every corresponding data point in the 700 and 800 nm channels. Then average the integrated signal intensities for each group of wells. Plot the data as a scatterplot against cell density (see Figure 3).
    1. Compare the results of the two different dilutions of primary and secondary antibodies and the results of the two different dilutions of DRAQ5 + Sapphire to settle on one dilution each for subsequent experiments. If linearity is not achieved, try scanning at a different intensity. Rescan with intensities 3 and 7 instead of 5 and reanalyze the data. If the data improve at one of those intensities but are still not satisfactory, rescan in increments around that intensity.
    2. Be careful not to analyze images where the software shows white spots in the wells; those spots signify saturated signal out of the range of the imager. Scan at a lower intensity if this occurs.
    3. If linearity is not achieved, try also to fix the cells within 6 hr of plating because they might grow or die unevenly in different columns overnight. Also try different plating densities, different scanning intensities, further optimizations of reagent dilution factors, glass-bottomed plates, DRAQ5 by itself as a nucleus-only stain, or different antibodies other than anti-α-tubulin or anti-MAP2.