Lineage-reprogramming of Pericyte-derived Cells of the Adult Human Brain into Induced Neurons

1. Isolation and Culturing of Adult Human Brain Cells

Experiments involving human tissue should be performed in accordance with all relevant governmental and institutional regulations regarding the use of human material for research purposes. The present protocol was developed in accordance with the approval by the ethical committee of the Medical Faculty of the LMU Munich and written informed consent from all patients.

This protocol of preparing cultures of the human adult cerebral cortex has been established using specimen of patients of both sexes suffering from temporal lobe epilepsy or other deep-seated non-traumatic, non-malignant lesions. The tissue obtained from the surgical room comprised exclusively the access channel to the brain lesion and therefore is considered healthy. The age range of the patients was 19-70 years.

1. Prepare growth medium by adding heat-inactivated fetal calf serum (FCS) to DMEM high glucose with GlutaMAX to obtain a final concentration of 20% FCS. Add 5 ml penicillin/streptomycin to a total of 500 ml growth medium. Perform this and all subsequent steps requiring sterile culture conditions in an appropriate laminar flow hood.
2. Keep the adult human brain biopsy obtained from the surgical room in Hanks’ balanced salt solution with CaCl₂ and MgCl₂ (HBSS) medium including HEPES (10 mM final concentration) on ice until processing. Start processing as soon as possible.
3. To start the dissociation into single cells, transfer the tissue into a 65 mm petri dish and mince into small pieces by using two sterile single-use scalpels. For enzymatic digestion use 3-6 ml TrypLE in a 15 ml conical tube and incubate for 15-30 min at 37 °C in a water bath.
4. Add 1 volume of prewarmed growth medium to facilitate dissociation and gently triturate the solution containing tissue pieces up and down by first using a 5 ml disposable pipette, followed by using a glass Pasteur pipette until homogenization of the cell suspension. Typically, some residual tissue pieces, mostly consisting of white matter, will remain in the suspension.
5. Spin down at 157 x g for 5 min and resuspend the pellet in the appropriate amount of growth medium (10 ml per uncoated T75 culture flask). Use one T75 culture flask for a biopsy of 5-10 mm diameter in size and extrapolate from this in case of larger specimens.
6. Place cells into a 37 °C incubator with 5% CO₂ and 5% O₂.
7. Replace half of the medium every 3-4 days until cells have reached confluency. This usually takes up to two weeks (referred to as passage 0 [P0]).

2. In vitro Expansion and Characterization of Adult Human Brain Cells

1. In vitro expansion:
   1. Aspirate the culture medium and wash with phosphate-buffered saline (PBS), kept at room temperature, before adding 3 ml TrypLE.
   2. Incubate at 37 °C for 5-10 min until most of the cells detached. Gently tap the flasks to facilitate lifting of the cells; following addition of 3 volumes of growth medium, transfer the cells into a 15 ml conical tube and spin down at 157 x g for 5 min and resuspend the cells in the appropriate amount of growth medium.
   3. Passage the cells at a ratio of 1:3 for optimal yield into new T75 cell culture flasks.
      Note: We have passaged these cells until P5 without any signs of reduced reprogramming efficiency. While at P0 the culture contains a considerable fraction of endothelial cells (as assessed by CD34 expression), at subsequent passages their number becomes negligible.
2. Characterization of adult human brain cultures:
   1. Immunocytochemistry:
      1. For characterization of the cells, seed ~60,000 cells onto poly-D-lysine-coated glass cover slips in 500 μl fresh growth medium in 24-well tissue culture plates and incubate at 37 °C with 5% CO₂ and 5% O₂.
      2. For immunocytochemical analysis of the cultured human brain cells remove the medium and wash 3x with 1 ml of PBS.
      3. Fix cells by adding 500 μl of 4% paraformaldehyde in PBS for 15 min at room temperature.
      4. Transfer the glass cover slips into an appropriate humidified staining chamber and block with 80 μl PBS containing 3% bovine serum albumin (BSA) and 0.5% Triton X-100 for 1 hr at room temperature.
      5. Add primary antibodies diluted in the same solution (for dilution factors see table of specific reagents and equipment) and incubate overnight at 4 °C or 1 hr at room temperature.
         Note: A substantial but varying fraction of the cells at this stage of culturing are immunoreactive for platelet-derived growth factor receptor β (PDGFRβ), cluster of differentiation 146 (CD146), NG2 chondroitin sulfate proteoglycan, and smooth muscle actin (SMA), i.e. markers characteristic of microvessel-associated pericytes. Only a minority (<1%) express the astroglial markers glial fibrillary acidic protein (GFAP) and S100β. Furthermore, at this stage the culture should be devoid of any cells expressing the neuronal marker βIII-tubulin. Use multiple primary antibody combinations for co-labeling of different markers. Further confirmation of the expression of glial, neuronal, endothelial and pericyte markers can be obtained by reverse transcription and quantitative real time polymerase chain reaction (not described in this protocol).
      6. Following incubation with the primary antibodies wash three times with 1 ml PBS and add the corresponding secondary antibodies (in the appropriate dilutions) to the staining solution and incubate 1 hr at room temperature in the dark. Wash the cells 3x with PBS. If required, prior to mounting include staining with DAPI for 5 min at room temperature.
      7. Use antifading mounting medium and analyze the cover slips using an epifluorescence microscope or a confocal microscope.
   2. Flow cytometry:
      1. For analysis of surface marker expression using flow cytometry, grow cells to confluency in uncoated T25 or T75 cell culture flasks.
      2. Detach cells by using TrypLE as described before and resuspend in staining solution consisting of PBS + 0.5% BSA. Per staining reaction, resuspend 100,000-200,000 cells in 100 μl of staining solution. Prepare single staining reactions per antibody used (CD146-FITC, CD140b-PE, CD34-APC, CD13-FITC, respective isotype control antibodies).
      3. Add fluorochrome-conjugated antibodies to each staining solution (for dilutions see table of specific reagents and equipment) directly into the cell suspension and incubate at 4 °C for 30 min in the dark.
      4. Wash the cells three times with 500 μl PBS and spin down between each washing step at 157 x g for 5 min.
      5. Resuspend the cell pellet in 0.5-1 ml staining solution and transfer cells through a cell strainer (70 μm) to FACS tubes. Protect the sample from exposure to light.
      6. Vortex samples prior to placing them in the FACS instrument.
      7. To analyze the living cells and exclude debris and cell aggregates, gate for FSC-A and SSC-A. Cell duplets are specifically gated out by FSC-A and FSC-W. To determine the right gating conditions for the cell surface markers set the gates with isotype-matched antibody control conjugated to the respective fluorochrome. Then determine the proportion of antibody-bound cells.

3. Enrichment for Pericyte-derived Cells by Fluorescence Activated Cell Sorting

1. Purify the pericyte cell population (resuspended in growth medium) prior to transduction via FACS sorting using a 70 μm nozzle. Use CD34-APC in combination with CD140b-PE and CD146-FITC to select for pericytes. Sort for CD34-negative, CD140b-, and CD146-positive cells.
2. Collect the sorted cells in growth medium and plate onto poly-D-lysine-coated glass cover slips in 24-well tissue culture plates and incubate at 37 °C with 5% CO₂ and 5% O₂.
3. 48 h after sorting replace the medium with fresh growth medium and perform transduction as described in  Protocol 4.

4. Retroviral Transduction of Pericyte-derived Cells and Reprogramming into Induced Neurons

Note: Refer to local biosafety guidelines when handling retroviral particles. In Germany the use of the retroviruses employed here require biosafety level 2. For production of retroviral particles refer to protocol⁶. For retroviral constructs used for reprogramming, refer to Karow et al. (2012). Retroviruses were pseudotyped with VSV-G (vesicular stomatitis virus-glycoprotein)⁵. For production please refer to Gavrilescu et al⁶.

1. 24 hr prior to retroviral transduction seed ~60,000 cells onto poly-D-lysine-coated glass cover slips in 500 μl fresh growth medium in 24-well tissue culture plates and incubate at 37 °C with 5% CO₂ and 5% O₂.
2. The next day, replace the growth medium with 500 μl fresh, prewarmed growth medium and add 1 μl of the respective concentrated supernatants containing retroviral particles (pCAG-IRES-dsred for control, especially during establishing the protocol in the lab), pCAG-ascl1-IRES-dsred, pCAG-sox2-IRES-gfp.
3. One day later, remove the medium including the viral particles from the cells and add 1 ml fresh, prewarmed B27-differentiation medium, composed of 49 ml DMEM high glucose with GlutaMAX and 1 ml B27 serum-free supplement. Maintain the cells in culture at 37 °C in 5% CO₂ and 5% O₂ for 4-8 weeks without changing the medium as repetitive medium changes become harmful as iNs mature.

5. Characterization of Induced Neurons by Immunocytochemistry

1. To determine the cellular identity of the transduced cells, in particular to demonstrate the acquisition of a neuronal phenotype, perform immunocytochemistry against neuronal antigens such as bIII-tubulin, MAP2, and NeuN (for antibodies and their respective dilutions, see table of specific reagents and equipment; follow the same procedure as indicated in 2.2.1).
2. For improving the maturation of PdiNs, co-culture these cells with neurons derived from E14 mouse cerebral cortices. To this end, dissect the cerebral cortex and dissociate the tissue mechanically with a fire-polished glass Pasteur pipette. Add 10,000-50,000 murine neurons to cultures of pericyte-derived cells 2-3 weeks following transduction with sox2- and ascl1-encoding retroviruses and continue culturing. Transduced cells can be distinguished from murine neurons by their reporter (GFP and dsRed) expression, either by live epifluorescence or following immunocytochemistry for GFP and dsRed.
3. To assess the formation of synapses onto PdiNs by murine neurons, perform immunocytochemistry against vesicular neurotransmitter receptors such as vesicular glutamate transporter 1 (vGluT1).
4. Probe transduced cells for their electrical properties (i.e. ability of generating action potentials) and the establishment of functional synapses by patch-clamp electrophysiology. For details of the procedure see Heinrich et al. (2011).