Ethics statement

This study conformed to the Australian National Health and Medical Research Council's published Code of Practice for the Use of Animals in Research, and experiments were approved by the Florey Neuroscience Institutes animal ethics committee. All animals were housed on a 12 h light/dark cycle with ad libitum access to food and water.

All experiments for iPS and human embryonic stem cell lines were performed in accordance with approvals obtained from the University of Melbourne Human Ethics Committee (#0605017, #0830010 and #0829937).

Cell culture and differentiation

IPS cells derived from FRDA patient fibroblasts (FA3 iPS and FA4 iPS) [14], control iPS cells (iPS1, WiCell USA), and the human embryonic stem cell line H9 (WA-09, WiCell USA) cell lines were cultured as previously described [15]. Briefly, mitomycin-C treated human foreskin fibroblasts in KSR media consisting of DMEM/nutrient mixture F-12, supplemented with 0.1 mM β-mercaptoethanol, 1% nonessential amino acids, 2 mM glutamine, 25 U/ml penicillin, 25 µg/ml streptomycin and 20% knockout serum replacement (all from Invitrogen). All cells were cultured at 37°C, 5% CO₂. Colonies were mechanically dissected every 7 days and transferred to freshly prepared human embryonic fibroblasts. The FA3-GFP cells were generated by transfecting FA3 iPS cells with the piggyBac transposon vector (Wellcome Trust Sanger Institute) modified to contain a GFP expression cassette, driven by the human elongation factor 1 alpha promoter. Positive GFP-expressing cells were enriched by mechanical dissection of transfected FA3 iPS cells, using a Leica MZFIII Fluorescence Stereomicroscope. For neural induction, colonies were treated with 500 ng/ml human recombinant noggin (PeproTech) and 4 ng/ml bFGF in neural basal media (NBM) [16]. After 14 days, colony pieces were mechanically harvested and cultured in suspension in NBM supplemented with 20 ng/ml bFGF and 20 ng/ml EGF to promote neurosphere formation. Neuronal differentiation was performed by plating neurospheres on poly-D-lysine/laminin substrates in NBM for at least 1 week, as previously described [16].

Electrophysiology

Neurons cultured on individual coverslips were transferred to the recording chamber mounted on the stage of an upright microscope (AxioExaminer D1, Zeiss) fitted with a 40X water-immersion objective lens. Cells were visualised with phase contrast or Dodt optics, and a monochrome CCD camera (Spot RT SE18, Diagnostic Instruments). Electrophysiological recordings in whole cell patch-clamp were performed at room temperature (22–25°C). Micropipette electrodes were fabricated from borosilicate tubing (1.0 mm O.D., 0.58 mm I.D.) using a Sutter P-2000 puller (Sutter Instrument Company) and had a tip resistance of 2–6 MW. During recordings, cells were superfused at 1–3 ml/min with extracellular buffer (137 mM NaCl, 5 mM KCl, 10 mM HEPES, 1 mM MgCl₂, 2 mM CaCl₂, 10 mM glucose, pH 7.35, 300–305 mosmol/kg), and micropipettes contained internal solution (115 mM K-gluconate, 10 mM HEPES, 7 mM KCl, 0.05 mM EGTA, 2 mM Na₂ATP, 2 mM MgATP, 0.5 mM Na₂GTP, pH 7.3, 294 mosmol/kg). Tetrodotoxin (TTX), tetraethylammonium chloride (TEA) and 4-aminopyridine (4-AP) were diluted daily to final concentrations in the bath perfusate, and administered by superfusion via a gravity-fed system. Signals were recorded with a MultiClamp 700B amplifier (Molecular Devices), and data acquisition system (Digidata 1440A, Molecular Devices), and AxoGraph X analysis software (AxoGraph Scientific). Records were digitized at 50 kHz and filtered at 10 kHz. Series resistance (R_S) was monitored in response to a 10 mV voltage step (mean 18.0±0.8 MW; cells with R_S greater than 30 MW were excluded). In current clamp, pipette capacitance neutralization was applied and bridge balance utilized to compensate errors due to R_S. All recordings were made from a holding potential of −73 mV, and corrections for liquid junction potential (12.8 mV; JPCalcW, courtesy of Prof. P H Barry, Sydney) were made offline. Results are presented as mean ± SEM.

Dipstick assays for complex I activity, complex IV activity and frataxin protein

Dipstick assay for frataxin, mitochondrial complexes I (CI) and IV (CIV) were obtained from Mitosciences, and performed as per the manufactures instructions in dissociated neurospheres (day 17–19 post neuronal induction).

TUNEL assay

Neurosphere frozen sections for TUNEL analysis were permeabilised in 0.1% Triton X-100 for 2 min on ice, followed by TUNEL labeling according to the manufacturer's protocol (Roche, In situ Cell Death Detection Kit, Fluorescein). Sections were photographed using an Olympus BX51 microscope and ULH100HG reflected fluorescence system with DP70 camera.

Post-image acquisition, images were rescaled from 1376 by 1038 pixels to 5504 by 4152 pixels (ImageJ, plugin TransformJ using quintic b-splines interpolation) [17]. Illumination correction was performed (CellProfiler software) [18], and contrast enhanced for each image uniformly (ImageJ, plugin CLAHE) [19]. A pixel classifier for use with CellProfiler was generated using the open source software Ilastik version 0.5 (www.ilastik.org) [20]. A CellProfiler pipeline was designed to collect intensity data from the TUNEL channel, using the areas of nuclei identified in the DAPI channel as a mask (Figure S2). For each nuclei identified in the DAPI channel, nuclei area, nuclei centroid location in Cartesian coordinates, integrated intensity of TUNEL, median intensity of TUNEL, and an image of the mask was collected. Data was then transformed with an arcsine transformation to approximate a normal distribution, and an ANOVA test performed.

ATP synthesis assay

ATP synthesis rates were measured in dissociated neurospheres (day 17–19 post neuronal induction) as previously described [21], with cells permeabilised in buffer containing 0.005% digitonin with substrate/inhibitors as described (mitochondrial complex II (CII)-dependent substrate succinate (10 mM), CII inhibitor malonate (1 mM), CI-dependent substrates glutamate (10 mM), pyruvate (10 mM) and malate (10 mM), and CI inhibitor rotenone (2.5 µM)).

Enzyme activity assays by spectrophotometry

Citrate synthase activity of dissociated neurospheres (day 19–24 post neuronal induction) was determined relative to total protein at 30°C as previously described [22].

The activity of CI was also detected spectrophotometrically in whole cell homogenates, as previously described [22]. However, due to the very large amount of error, the measurements were deemed unreliable (data not shown). This was likely due to the small amount of biological material available for analysis, and the high rotenone-insensitive rate of NADH oxidation in these samples (61%±6.3 (SEM)).

Mitochondrial membrane potential (ΔΨ_m)

Dissociated adherent neurospheres (day 25–33 post neuronal induction) grown in a black 96 well plate were stained in media containing 75 nM tetramethyl rhodamine methyl ester (TMRM) (Ex 544 nm, Em 590 nm) and 2 µg/ml Hoechst-33258 (Ex 346 nm, Em 460 nm) for 40 min at 37°C, 5% CO₂. Cells were washed in warm phosphate-buffered saline (PBS), then the mitochondrial membrane potential (ΔΨ_m) was determined as the ratio of TMRM to Hoechst fluorescence (FLUOstar OPTIMA). Control wells were treated with 20 µM carbonylcyanide-ρ-trifluoromethoxyphenyl hydrazone (FCCP) to dissipate the ΔΨ_m.

The ΔΨ_m was also determined by microscopy in dissociated adherent neurospheres (day 21 post neuronal induction) grown in 35 mm glass bottom dishes (World Precision instruments). Cells were stained in Hanks Buffered Salt Solution with magnesium and calcium (Gibco) containing 75 nM TMRM for 30 min at 37°C, 5% CO₂. Cells were visualised at 37°C, 5% CO₂ on a Delta Vision OMX V3 Imaging System (Applied Precision) using the TRITC filter set with oil-immersion objective (Olympus, IX71), fitted with a CoolSNAP HQ² camera (Photometrics). After initial imaging, 20 µM FCCP was applied to dishes to dissipate the ΔΨ_m, and regions of interest were visualized again. Images were de-convoluted, cropped and projected with maximum intensity (Applied Precision, SoftWoRx v5.5). Mitochondrial rich perinuclear regions were selected in greater than 80 cells per cell line and average TMRM fluorescence determined pre- and post-FCCP application.

Western blot analysis

Dissociated neurospheres (day 17–19 post neuronal induction) were analysed by western blot analysis as previously described [23], using the human OXHOS complex antibody cocktail (Abcam, MS601).

Animals and transplantation surgery

Dissociated neurospheres derived from the FA3 GFP iPS cells were grafted into 4 adult (250 g), female athymic (nude; CBH-rnu) rats. The animals were deeply anaesthetised using 2% isoflurane as an inhalant and placed in a stereotaxic frame (Kopf) in a flat skull position. Donor neural progenitors were injected into the right cerebellar hemisphere using a 5 µl microsyringe (SGE Analytical Science) fitted with a pulled glass capillary as previously described [24]. A total of 1 µl of the cell suspension (0.5×10⁵ cells/µl) was injected over 2 min; 10.8 mm caudal to bregma; 3.6 mm lateral to the midline; and 4.0 mm below the dural surface. The cannula was left in place a further 5 min after the contents had been delivered to prevent backflow up the needle track. The animals were killed (lethal dose of pentobarbitone; intravenous perfusion) 12 weeks later for histological analysis. Animals were transcardially perfused with saline (0.9%) followed by paraformaldehyde (4% in 0.1 M PBS). The brains were removed and post-fixed for a further 2 h in 4% paraformaldehyde and cryoprotected overnight in sucrose (25% in 0.1 M PBS) before being sectioned on a freezing microtome (Leica). Parasagittal sections were collected in 12 series at a thickness of 30 µm.

Immunohistochemistry and imaging

Immunohistochemical procedures for transplantation studies were performed as previously described in [25]. Detection of the antibody conjugates was achieved using either a peroxidase-based reaction followed by precipitation of diaminobenzidine (DAB; Sigma) or conjugation of a fluorophore. The primary antibodies and dilutions used were as follows: mouse anti-APC (1∶500; Calbiochem); rabbit anti-GABA (1∶5000; Sigma); rabbit anti-GFAP (1∶200; DAKO); chicken anti-GFP (1∶1000; AbCam); mouse anti-NeuN (1∶200; Millipore); rabbit anti-Olig2 (1∶500; Millipore); mouse anti-PSA-NCAM (1∶100; Santa Cruz); rabbit anti-Tbr1 (1∶1000; Millipore). For fluorescent labeling, secondary antibodies raised in donkey were conjugated with Alexa-488; -549 or -633 (Molecular Probes). For DAB labeling, biotin-conjugated goat anti-rabbit was used (Jackson ImmunoResearch) followed by incubation with peroxidase-conjugated streptavidin (Vectastain ABC kit, Vector Laboratories). No significant non-specific signal was generated from the secondary antibodies, as determined by control experiments where the primary antibody was omitted from the immunohistochemical procedure.

The overview of DAB-labeled GFP images were generated as a montage of single dark-field images captured using a 10X objective on a Leica DM6000 upright light microscope with the Leica X-Y tiling software module (v3.8). Fluorescent images of the grafted cells were captured using a Zeiss upright laser-scanning confocal microscope and ZEN software.

Statistics

Unless otherwise described, error bars represent SEM, and p values are generated from student t-tests (GraphPad, PRISM V6.0b) and reported when ≤0.05.

1. FA iPS cells differentiate in vitro to functionally active neurons

We have previously described the generation and characterization of two FRDA iPS cell lines, known as ‘FA3’ and ‘FA4’, derived from adult FRDA human skin biopsies, and demonstrated their capacity to differentiate to neurons in vitro [14]. Here we extend these studies to determine the functionality of FA-derived neurons using whole cell patch clamp recordings. We find then that FA-derived neurons are electrically active and display core electrophysiological characteristics typical of neurons derived from human stem cells, namely I_Na and I_K [26]–[28].

Electrophysiological recordings were obtained for 62 FA3 iPS-derived neurons displaying an ovoid soma (∼10 mm diameter) and bipolar processes (Figure 1A–B). The mean resting membrane potential was −49.5±1.3 mV, and membrane capacitance 7.7±0.3 pF. In current clamp, intracellular depolarizing current steps (20 to 190 pA; 300 ms duration) elicited action potentials in 58 of 62 (94%) of FA iPS-derived neurons (Figure 1C). Firing threshold was −31.8±0.7 mV, with an average action potential amplitude of 96.4±1.9 mV, and mean action potential latency of 6.8±0.3 ms and half-width of 4.8±0.4 ms during supra-threshold input (+190 pA). Of the neurons that generated action potentials, 53 displayed a single action potential fired at stimulus onset. The remaining four cells exhibited between two and four action potentials at threshold, decreasing to one or two spikes with increasing stimulation (Figure 1C). Spontaneous firing in the absence of stimulation was never observed. Exposure to 1 mM tetrodotoxin (TTX), the selective blocker of voltage-gated sodium channels, inhibited firing of action potentials (Figure 1E). Injection of hyperpolarizing current (−5 to −30 pA) resulted in membrane hyperpolarization (Figure 1D), but failed to evoke a voltage-dependent depolarizing sag indicative of a hyperpolarization-activated mixed cation current, and rebound action potentials at stimulus offset were also absent.

Download:

• PPT
  PowerPoint slide
• PNG
  larger image
• TIFF
  original image

Figure 1. Firing properties of neurons derived from FA iPS cells.

(A–B). Patch-clamp recordings were made from iPS cells displaying an ovoid soma and bipolar processes. (C). Action potentials were activated in response to membrane depolarization. In the example shown here, multiple action potentials are observed at threshold (gray), whilst a single spike occurs in response to supra threshold stimulation (black). (D). Negative current injection resulted in membrane hyperpolarisation, without the addition of a depolarizing sag or rebound action potentials at stimulus offset. (E). Action potentials were blocked by addition of the sodium channel antagonist TTX (gray). (F). In voltage-clamp recordings, the transient inward current observed during membrane depolarization (black) was abolished in the presence of TTX (gray). (G). I–V curve demonstrating the voltage-dependence of the TTX-sensitive sodium current.

https://doi.org/10.1371/journal.pone.0101718.g001

In voltage clamp, responses measured during 10 mV increments in voltage (−63 to +27 mV; 250 ms duration) revealed a small rapidly activating and inactivating inward current (Figure 1) followed by a sustained outward current (Figure 2). The transient inward current was completely and reversibly blocked by application of 1 mM TTX (Figure 1F), indicative of a sodium current (I_Na). The peak TTX-sensitive I_Na was −550±73 pA (−83±7 pA/pF) at −13 mV (n = 3), measured in the presence of 10 mM TEA and 1 mM 4-aminopyridine (4-AP) (Figure 1G). Consistent with a potassium-mediated current (I_K), the sustained outward current that followed I_Na (Figure 2A) was reduced by the potassium channel antagonists TEA and 4-AP (Figure 2B). The mean total steady-state current (I_K ss) was 535.4±31.8 pA (Figure 2D), with a mean maximum peak current (I_K peak) of 565.0±33.1 pA at 27 mV (Figure 2C; n = 44). The steady-state portion of the slowly inactivating current was blocked by 80.2% by 10 mM TEA (n = 8), or 82.7% by 10 mM TEA and 1 mM 4-AP (Figure 2D; n = 5), whilst the peak current was blocked by 72.5% by 10 mM TEA (n = 8), or 82.4% by 10 mM TEA and 1 mM 4-AP (Figure 2C; n = 5). Hyperpolarization of the membrane did not evoke activation of a slowly activating inward current (I_h; data not shown).

Download:

• PPT
  PowerPoint slide
• PNG
  larger image
• TIFF
  original image

Figure 2. Outward currents recorded from FA iPS-derived neurons.

(A) Membrane depolarization activates a transient sodium current followed by a sustained outward current that is attenuated by potassium channel blockers TEA (10 mM) and 4-AP (1 mM) (B). C–D. I–V curves depicting peak outward potassium current (C; I_K peak) and steady-state current (D; I_K ss, measured at bar in A) as a function of membrane potential. Currents recorded in the absence of all channel blockers (closed) were reduced in the presence of TEA (open), or TEA and 4-AP (gray).

https://doi.org/10.1371/journal.pone.0101718.g002

2. Frataxin expression is reduced in FA iPS-derived neurospheres, but level of cell death is normal

Although FA iPS cells are able to differentiate into functional neurons, the residual level of frataxin protein expression has not been quantified. Consistent though with the transcriptional expression data previously described [14], frataxin protein levels were significantly lower in both FA3 iPS-derived neurospheres (39%±5.3 SEM) and FA4 iPS-derived neurospheres (28%±4.7 SEM) relative to control iPS1-derived neurospheres (Figure 3A; p<0.001). These levels correspond to the upper quartile range of frataxin protein levels found in FRDA patients, as determined in patient buccal cells [29], [30].

Download:

• PPT
  PowerPoint slide
• PNG
  larger image
• TIFF
  original image

Figure 3. Frataxin expression and level of cell death in neurospheres derived from FA iPS cells.

(A) Frataxin protein was measured in cell lysates using immuno-capture dipsticks and quantified relative to the control iPS line. N = 3 experiments, with at least 12 neurospheres per sample for each replicate experiment (B) Levels of cell death in neurospheres cultured for 7–26 days were determined by percentage of TUNEL expression. (C) Representative TUNEL staining images from day 26. N = 3 separate experiments per time point, with at least 5 neurospheres per time point for each experiment. *p<0.001. Error bars where shown  = SEM.

https://doi.org/10.1371/journal.pone.0101718.g003

It is unknown however if there is cell death occurring in specific subpopulations of neurons/neural progenitors during neural differentiation that may alter findings. In order to address this, a temporal analysis of apoptotic cells was performed in neurospheres cultured for up to 26 days without passage (Figure 3B, C). Following neurosphere formation both FA3 and FA4 iPS-derived neurospheres, like controls, show increasing levels of cell death from two weeks onwards (Figure 3B). However, at no stage is there a statistically significant increase in cell death in the FA3 and FA4 iPS-derived neurospheres compared to control iPS1 and hES-derived neurospheres (Figure 3B).

3. Mitochondrial functional analyses in FA iPS-derived neurospheres and neurons

Since FRDA is considered a mitochondrial disease, mitochondrial function was also analysed in FA iPS-derived neurospheres and from control iPS1 and hES-derived neurospheres. No significant changes were observed in citrate synthase activity in either FA3 and FA4 iPS-derived neurospheres relative to controls, indicative of no difference in mitochondrial volume per cell [31] (Figure 4A). The mitochondrial oxidative phosphorylation (OXPHOS) system, in which complexes I, II and III (CI, CII and CIII) contain Fe-S clusters [32], was also monitored. The activities of CI and CIV in FA3 and FA4 iPS-derived neurospheres were equivalent to control iPS1 and hES-derived neurospheres (Figure 4B and 4C). Expression of representative OXPHOS subunits were also unchanged in FA3 and FA4 iPS-derived neurospheres relative to controls (Figure 4D), suggesting no marked disruption to OXPHOS complex assembly or stability. The ΔΨ_m was measured by the accumulation of the cationic dye TMRM in the mitochondria [21], and was found to be unchanged in both FA3 and FA4 iPS-derived neurons (dissociated neurospheres) relative to control iPS1 and hESC-derived neurons (Figure 4E; Figure S1). Finally, CI-dependent (glutamate + malate, pyruvate + malate) and CII-dependent (succinate) ATP synthesis capacities were consistent in FA3 and FA4 iPS-derived neurospheres (Figure 4F) with controls. Overall, these data demonstrate that mitochondrial volume and function are not altered in FA iPS neural derivatives relative to controls.

Download:

• PPT
  PowerPoint slide
• PNG
  larger image
• TIFF
  original image

Figure 4. Mitochondrial functional analyses of FA iPS neural progenitors and neurons.

(A) Citrate synthase activity was measured in neurosphere lysates spectrophotometrically and expressed relative to total protein. (B) CI, and (C) CIV activities were measured in neurosphere lysates by immuno-capture dipsticks, and expressed relative to control iPS1. (D) Expression of subunits representative of the OXPHOS CI-V was determined in neurosphere lysates by western blot analysis. (E) The ΔΨ_m was assessed in dissociated neurospheres plated as a monolayer by measuring the accumulation of the cationic dye TMRM relative to cell number (as determined by Hoechst staining). (F) The rate of ATP synthesis was measured in digitonin permeabilised neurospheres, with indicated substrate inhibitor combinations. Error bars are SEM. N = 3–5 experiments, with at least 12 neurospheres per condition for each replicate experiment.

https://doi.org/10.1371/journal.pone.0101718.g004

4. In vivo differentiation potential of FA iPS-derived neural progenitors in the adult cerebellum

FA iPS derived neural progenitors and neurons do not show an overt phenotype in mitochondrial function or cell death in vitro. We therefore proceeded to examine in vivo differentiation potential of FA iPS-derived neural progenitors to determine their capacity to differentiate and integrate into the adult nervous system. These transplantation studies provide a basis to examine the in vivo use of FA iPS neural derivatives.

To detect transplanted FA cells in vivo, FA3 iPS cells were transduced to constitutively express GFP and then differentiated to neural progenitors. GFP-expressing FA iPS neural progenitors were transplanted into the cerebellar regions of adult rats. Immunohistochemistry for GFP at 12 weeks after transplantation revealed surviving grafts in all four animals. The grafts were positioned in the posterior lobe of the cerebellum, predominately in the grey matter, but also entering the adjacent white matter of the arbor vitae (Figure 5A). The gross morphology of the grafts presented as discrete or contained deposits of cell mass with uniform density and no sign of overt overgrowth or tumourigenesis (Figure 5B). Clusters of sparsely distributed GFP+ cells could be found in the host parenchyma, adjacent to the main graft deposit, that allowed for detailed analysis of cell morphology. Various morphological profiles consistent with both immature and terminally differentiated neural cell types, including neurons and glia, could be identified (Figure 5C and D). Cells with the morphology of migrating neuroblasts (Figure 5F) were distributed extensively throughout the white matter of the grafted hemisphere (Figure 5E). The mitochondrial location of GFP and consequent widespread distribution throughout cells allowed also for detailed resolution of graft derived fiber patterns. Many single GFP+ fibers appeared as spiny dendrites (Figure 5G), consistent with integration at the synaptic level. Large fasciculated fiber bundles were observed originating from the graft and traversing on an anterior pathway through the white matter and deep cerebellar nuclei (Figure 5H), however, no GFP+ fibers were observed outside of the cerebellum. Dense networks of GFP+ fibers were also seen throughout the cerebellar grey matter (Figure 5I).

Download:

• PPT
  PowerPoint slide
• PNG
  larger image
• TIFF
  original image

Figure 5. Structure and morphology of FA-iPS donor grafts revealed by immunohistochemistry for GFP.

(A) Darkfield imaging of a representative section illustrates the size and placement of the grafts within the cerebellum. (B) The GFP+ cells were distributed uniformly throughout the grafts. (C, D) less densely packed cells adjacent to graft had morphological features consistent with differentiated neurons and glia. (E, F) cells with the morphology of migrating neuroblasts could be found throughout the cerebellar white matter. GFP+ fibres appeared in various forms including (G) spiny dendrites, (H) fasciculated bundles and (I) dense networks within the cerebellar grey matter. Scale bars: A, E - 1 mm; B, H, I – 200 µm; C, F – 100 µm; D - µm; G – 25 µm.

https://doi.org/10.1371/journal.pone.0101718.g005

Double labeling for antigenic markers of cell phenotype revealed the presence of GFP+ cells with neuronal or glial identity as well as migrating neuroblasts. The grafts appeared rich in neurons based not only on morphology but also on the presence of large numbers of cells expressing the neuronal nuclear marker NeuN (Figure 6A). The vast majority of GFP+ neurons in the grafts also expressed the inhibitory neurotransmitter γ-aminobutyric acid (GABA; Figure 6B and C). Cells with antigenic features of glia were more sparsely distributed in the grafts and included GFAP+ astrocytes as well as cells expressing markers consistent with immature (Olig2+/APC-) and mature (Olig2+/APC+) oligodendrocytes (Figure 6D and E). Another sparsely distributed population of cells expressed the transcription factor Tbr1 (Figure 6F). The cells with morphology of migrating neuroblasts, distributed widely throughout the white matter, were found to express the cell adhesion molecule PSA-NCAM (Figure 6G). Taken together, FA iPS-derived neural progenitors show a robust capacity to integrate and differentiate into neuronal and glial lineages within the adult nervous system.

Download:

• PPT
  PowerPoint slide
• PNG
  larger image
• TIFF
  original image

Figure 6. Neuronal phenotypes of grafted FA-iPS neural progenitors.

(A) The grafts were rich in differentiated neurons based on immunohistochemistry for GFP and NeuN. (B, C) Many of the neurons also expressed the neurotransmitter GABA (arrowhead in B shows host GABA+ Purkinje neurons). GFP+ cells with glial phenotypes were also present based on immunohistochemistry for (D) the astrocyte marker GFAP and (E) the oligodendrocyte markers Olig2 and APC. (F) A small population of GFP+/TBR1+ cells were scattered throughout the graft core. (G) The majority of the GFP+ cells with the morphology of migrating neuroblasts also expressed the cell adhesion molecule PSA-NCAM. Scale bars: A, C, D, E – 50 µm; B – 200 µm; F, G – 100 µm.

https://doi.org/10.1371/journal.pone.0101718.g006