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Immunocytochemistry (ICC) staining is a powerful tool that enables specific proteins or other molecules to be qualitatively and quantitatively visualised in a cellular context. As such, ICC can provide insights into cellular phenotypes, protein expression, and sub-cellular localisation.

In both  ALS/FTD research and drug discovery, ICC is widely used to study TDP-43, a protein that is key to both conditions. A hallmark of these diseases is the mis-localisation, accumulation, and aggregation of TDP-43, of which ICC is a great tool to visualise all of these phenotypes in in vitro cell models. In motor neurons with TDP-43 pathology, ICC has been used to visualise the abnormal accumulation of the protein in the cytoplasm.

High-content analysis of ICC data allows for the precise quantification of TDP-43 levels and its localisation, aiding in the identification of potential therapeutic targets and monitoring the effects of candidate drugs. This approach has been essential in developing phenotypic screening assays, which help pinpoint compounds capable of modulating TDP-43 pathology, offering crucial insights into potential therapies for ALS and FTD.

This versatile and widely used technique provides researchers with valuable information about cell structure, function, and behaviour. While ICC staining of iPSC-derived cells is a relatively straightforward process, it does require careful consideration of antibody pairings, delicate cell treatment, and optimisation to reduce non-specific signals. Here, we provide an optimised protocol for the ICC staining of ioMotor Neurons.

• ioMotor Neurons (io1027) cultured in a 24 well plate following the user manual to day 11 at a seeding density of 30,000 cells/cm²

• Biological safety cabinet with a carbon filter (MSC-CF)

• Normoxic cell culture incubator (37°C, 5% CO²)

• -80°C freezer

• 1000 μL, 200 μL, 20 μL and 10 μL pipettes

• Standard light microscope

• Epifluorescent microscope

This protocol is split into 4 sections:

1: Cell fixation

2: Blocking & permeabilization

3: Primary antibody labelling

4: Secondary antibody labelling

1. Cell fixation

1.1. Prepare the following reagents, according to the instructions in Appendix 1:

1.1.1. 4% paraformaldehyde/PBS

1.1.2. 0.1% Triton-X-100/DPBS

1.1.3. Blocking Solution

1.2. Carefully remove spent culture medium, without disturbing the cells.

1.3. Add 200 μL of DPBS to each well.

1.4. Remove the DPBS without disturbing the cells. Carefully add 200 μL of cold 4% paraformaldehyde/PBS to each well.

1.5. Incubate at room temperature for 10 minutes.

1.6. Remove the 4% paraformaldehyde/PBS solution from each well without disturbing the cells.

1.7. Carefully add 200 μL of DPBS to each well.

1.8. Remove the DPBS without disturbing the cells.

1.9. Repeat steps 1.6 to 1.8 a further 2 times.

1.10. Carefully add 200 μL of DPBS to each well on the centre of the well.

1.11. If the staining will not proceed immediately, wrap the plate with parafilm and store at 4°C overnight. Otherwise, continue to the next section.

2. Blocking and permeabilisation

2.1. Carefully remove the DPBS.

2.2. Gently add 200 μL of blocking solution down the side of each well.

2.3. Incubate at room temperature for 1 hour.

3. Primary antibody labelling

3.1. Prepare the primary antibody solutions described in Appendix 1.

3.2. Following the incubation described in step 2.3, aspirate blocking solution.

3.3. Carefully add 250 μL of the primary antibody mixture to the appropriate wells.

3.4. Add 250 μL of 0.25% TritonX-100/PBS to the negative control wells.

3.5. Seal plates with parafilm and incubate overnight at 4°C.

3.6. Remove the liquid from each well without disturbing the cells.

3.7. Carefully add 250 μL of DPBS to each well.

3.8. Incubate cells for 5 minutes at room temperature.

3.9. Repeat steps 3.6 to 3.8 a further 2 times, leaving the cells in 250 μL of DPBS before moving on to secondary antibody labelling.

4. Secondary antibody labelling:

4.1. Prepare the secondary antibody solutions described in Appendix 1.

4.2. Aspirate the DPBS from the wells.

4.3. Carefully add 250 μL of secondary antibody (with DAPI) to the appropriate wells.

4.4. Incubate cells with the secondary antibody mixture for 1 hour at room temperature.

4.5. After incubation, remove the liquid from each well without disturbing the cells.

4.6. Carefully add 250 μL of DPBS to each well.

4.7. Incubate cells for 5 minutes at room temperature - wrap the plates in foil or place them in a cupboard to prevent fluorophore bleaching.

4.8. Repeat steps 4.5 to 4.7 a further 3 times, leaving the cells in 500 μL of DPBS.

4.9. Image each well using a fluorescent microscope with the fluorescent channel most appropriate for each antibody.

Preparation of 4% paraformaldehyde/PBS:

1. Add 30 mL of DPBS to a 50 mL centrifuge tube.

2. Add 10 mL of 16% paraformaldehyde.

3. Mix gently and store in the fridge until use. Keep for a maximum of 10 days.

Preparation of 0.25% Triton-X-100/DPBS:

1. Add 1250 μL of Triton-X-100 stock solution to 500 mL of DPBS.

2. Mix thoroughly and store at room temperature until use.

Preparation of blocking solution:

1. Add 19 mL of 0.25% Triton-X-100/DPBS to a 50 mL centrifuge tube.

2. Add 1000 μL of Fetal Bovine Serum.

3. Mix gently and store in the fridge until use.

Preparation of primary and secondary antibody solutions:

1. Prepare the base antibody solution by diluting 1 mL of blocking solution in 9 mL of DPBS.

2. Refer to the table below for antibody dilution recommendations.

3. Dilute each antibody in a relevant volume of the base antibody solution. Keep for a maximum of 10 days and observe whether the liquid is transparent or if there are particles in suspension. 

Secondary antibodies should be diluted in DPBS + DAPI (1:500)

Table 1: Validated antibody information for the general characterisation of ioMotor Neurons.

If you have any questions or need assistance, please reach out to technical@ÎÞÓǶÌÊÓƵ and we will do our best to support you.

Published February, version 1