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Proc Natl Acad Sci U S A. Oct 28, 2008; 105(43): 16707–16712.
Published online Oct 15, 2008. doi:  10.1073/pnas.0808488105
PMCID: PMC2575484
Medical Sciences

Striatal progenitors derived from human ES cells mature into DARPP32 neurons in vitro and in quinolinic acid-lesioned rats


Substitutive cell therapy using fetal striatal grafts has demonstrated preliminary clinical success in patients with Huntington's disease, but the logistics required for accessing fetal cells preclude its extension to the relevant population of patients. Human embryonic stem (hES) cells theoretically meet this challenge, because they can be expanded indefinitely and differentiated into any cell type. We have designed an in vitro protocol combining substrates, media, and cytokines to push hES cells along the neural lineage, up to postmitotic neurons expressing striatal markers. The therapeutic potential of such hES-derived cells was further substantiated by their in vivo differentiation into striatal neurons following xenotransplantation into adult rats. Our results open the way toward hES cell therapy for Huntington's disease. Long-term proliferation of human neural progenitors leads, however, to xenograft overgrowth in the rat brain, suggesting that the path to the clinic requires a way to switch them off after grafting.

Keywords: cell therapy, Huntington's disease, striatum, cell differentiation, overgrowth

Huntington's disease (HD), a neurodegenerative disease of genetic origin that affects primarily the striatum, has been shown to be partially amenable to treatment by substitutive cell therapy. Fetal neural grafts have provided preliminary clinical success in a small number of HD patients (15). However, this technique is marred by logistic problems, because fetal striatal cells in tissue dissected immediately after elective abortion cannot be adequately expanded in vitro. This restricts considerably the amount of material available for the delivery of an optimally sized transplant to all patients who may benefit from it.

Meeting this challenge requires identifying cells that can be banked and propagated as necessary, and that fully reproduce the phenotype of the fetal neural precursors. Due to their self-renewal and pluripotency properties, human embryonic stem (hES) cells theoretically fulfill both prerequisites. Several authors have already demonstrated the relevance of using hES cells for therapy for Parkinson's disease (PD). They showed that hES cells can be differentiated in vitro into neurons exhibiting major phenotypic characteristics of the nigral dopaminergic neurons (68). Implantation of hES cell-derived neural progenitors committed to that phenotype into the rat brain has revealed appropriate differentiation of a sizeable proportion of those cells (6, 9).

We have undertaken a study aimed at reaching a similar achievement in hES cell therapy for HD: that is, (i) designing the in vitro protocol necessary to obtain the hES cell-derived progeny equivalent to fetal striatal progenitors, and (ii) validating this protocol in vivo by using xenotransplantation in rodents. Via a multistep protocol we have obtained postmitotic neurons exhibiting major phenotypic characteristics of the striatal GABAergic medium-spiny neurons. Xenotransplantation of hES cell-derived striatal progenitors into adult nude rats confirmed the efficiency of the differentiation protocol. It also confirmed the risk of graft overgrowth recently revealed in an experiment with the PD model (9), suggesting the need for antiproliferative safety procedures before proceeding toward the clinic.


Our protocol to differentiate hES cells into striatal neurons was developed based on the simplifying scheme that the normal course of neuronal differentiation may be separated into three successive steps: namely, (i) neural induction, (ii) regional commitment while neural expansion continues, and (iii) neuronal maturation (Fig. 1A). Commitment to a ventral telencephalic identity can be confirmed by detection of specific markers of the lateral ganglionic eminence (LGE), the striatal germinative zone (10), such as BF1 (FOXG1B), GSH2, and DLX2 (11, 12). Terminal striatal differentiation can be established by the detection of MAP-2+/Ki67 postmitotic neurons expressing key striatal markers, such as GABA, GAD67 (GAD1), DARPP32, ARPP21, calbindin, or calretinin. For each step, we sought to optimize our protocol by using distinct combinations of substrates, media, and signaling molecules. Neural induction was carried out as previously described (7) using a coculture with murine stromal cells (MS5). Default commitment toward telencephalon relied on the plating of mechanically isolated rosettes cells on coated dishes in N2 medium supplemented with BDNF (7, 13). Ventralization of the telencephalic neural progenitors was sought by adding SHH and DKK1 (passages 1–4) because these cytokines are involved in the patterning of the forebrain in mouse and chicken (14, 15) and can push neural cells derived from ES cells toward ventral telencephalic progenitors (8, 16, 17). Neuronal terminal differentiation was carried out by replating the committed neural progenitors at a lower density (7) in the absence of patterning molecules but in the presence of dibutyryl-cAMP (dbcAMP) and valproic acid, previously found to stimulate GABA neurogenesis of rat forebrain stem cells (18).

Fig. 1.
Phenotypic characterization of striatal progenitors and neurons generated in vitro from hES cells. (A) Outline of the multistep protocol for the differentiation of striatal progenitors and neurons. DIV, days in vitro. (B and C) Proliferative neural rosette ...

In Vitro Assessment of a Striatal Differentiation Protocol.

Neural, neuronal, and striatal differentiation was monitored from day in vitro 0 (DIV 0) to DIV 63 by using immunocytochemistry (Fig. 1) and quantitative PCR (Q-PCR) (Fig. 2). Neural rosettes were obtained after 3 weeks of coculture with MS5 and were amenable to passaging, expansion, and patterning [supporting information (SI) Fig. S1]. Relative gene expression analysis showed a time-dependent loss of the ES marker NANOG upon rosette formation (at DIV 21–23, stage A), consistent with a peak of expression of the early neural marker SIX3 and a delayed peak of expression of the neural marker SOX1 (Fig. 2A). After 4–5 weeks of culture of these rosettes in the presence of SHH and DKK1, hES cell progeny formed a monolayer of neural progenitors. Most cells were immunopositive for the neural markers nestin (92 ± 3%, n = 5) and Pax6 (77 ± 3%, n = 3) and were Ki67+ proliferating progenitors (Fig. 1 B, C, and H). At this stage, few cells expressed neuronal markers, such as Tuj1 (18 ± 1%, n = 3) or MAP2 (4 ± 1%, n = 5) (Fig. 1 C and H). As previously described with primate ES cell progeny (19), BF1 expression was already detected at the rosette stage (stage A) and reached a plateau at DIV 29–30 (stage B) (Fig. 2B). Striatal enrichment of the culture was detected by the peak of expression of GSH2 and DLX2 markers at stage B (Fig. 2B). We observed a significant increase in LGE markers on exposure of stage A culture for 12 days to BDNF, SHH, and DKK1 compared with BNDF alone (3.6 ± 0.8-fold, P < 0.01 for GSH2, 4.1 ± 0.7-fold, P < 0.01 for DLX2; n = 5) (Fig. 2C). Conversely, BF1 expression did not change significantly in the same cultures (1.1 ± 0.1 fold; n = 5). These findings indicate that generation of striatal progenitors from hES cell-derived neural rosettes can be regulated by extrinsic signals. Terminal maturation of the expanded and partially patterned neural cells (DIV 46–59, stage D) over two additional weeks resulted in a cell population (DIV 62–72, stage E) that contained up to 22 ± 2% (n = 3) MAP2+ postmitotic neurons (Fig. 1H). Accordingly, Q-PCR showed a continuous increase in expression of the neuronal marker MAP2 coinciding with the decreased expression of markers SIX3 and SOX1 (Fig. 2A). Among the MAP2 immunopositive neurons, 36 ± 2% were GABA+ (n = 4), and 53 ± 6% expressed DARPP32 (n = 4) (Fig. 1 E, G, and I). In addition, 10 ± 2% of MAP2+ neurons were immunoreactive for calbindin (n = 4), and 55 ± 7% were immunoreactive for calretinin (n = 3) (Fig. 1 D, F, and I). Less than 2% of the neurons expressed tyrosine hydroxylase (TH) (Fig. 1I and Fig. S1F). Even fewer neurons were serotonergic (5HT+). Human GFAP-immunoreactive astrocytes were very scarce (Fig. 1H and Fig. S1H). Numerous neural progenitors (nestin+/Pax6+) persisted in these long-term cultures in the absence of any exogenous mitogens, patterning factors, or stromal feeders. Striatal commitment of the terminally differentiated culture was also confirmed by Q-PCR, which showed increased expression of DARPP32, GAD67, and calbindin (Fig. 2B). We next tested in vitro function of neurons in our striatal culture by patch clamp studies. Single-cell recordings demonstrated the induction of single-action potential (5 of 9) or rhythmic activity (4 of 9) in response to depolarizing currents (Fig. S2).

Fig. 2.
Expression of key regulatory genes during differentiation. (A and B) Q-PCR analyses of undifferentiated hES cells, stages A, B, D, E, and human FB. (A) Time-dependent gene expression suggesting the sequential differentiation of hES cells (NANOG+) into ...

Intracerebral Transplantation of hES Cell-Derived Striatal Progenitors.

Fetal striatal neural grafts survive and integrate best when tissue is retrieved at a stage at which progenitors are present in the ganglionic eminence—i.e., committed to a ventral telencephalic fate—but still proliferating and not terminally differentiated (20). Accordingly, the most likely candidates for in vivo transplantation should be found between the rosette stage and the neuronal stage. Our first series of experiments aimed at defining a more precise optimal time window for transplantation. For this purpose, we transplanted cells treated using our protocol for 23–72 days and qualitatively assessed their survival, the formation of teratoma-like regions in the grafts, and both neural progenitor and striatal neuron differentiation. Cells were transplanted into the quinolinate-lesioned right striatum of immunocompetent rats that were killed 4–6 weeks after surgery (n = 48). For sake of comparison, in vitro differentiation was divided up arbitrarily into the five stages (A to E) as defined above. All animals undergoing transplantation had surviving grafts with cells positive for the human nuclear antigen (HNA). Together, all studied parameters demonstrated that the cell content of the grafts depended on the duration of their in vitro differentiation. Human grafts transplanted at early stages (stages A and B) included regions that were not expressing neuroectodermal markers (“teratoma-like regions”) in most cases (Fig. 3 A–C). They seldom maintained a homogenous neural specification, and then only exhibited structures resembling neural rosettes (Fig. 3 A, D, and E). In contrast, more mature cells (stages C, D, and E) produced grafts that were fully committed to the neural lineage and did not include teratoma-like regions (Fig. 3 A and I). Regarding striatal neuron maturation, stage D was better than the others. Only a few of the DARPP32+ neurons were detected with cells transplanted at stages C or E (Fig. 3 A, F, and G). In contrast, areas containing clustered DARPP32+ and calbindin+ striatal neurons were observed in all stage D transplants (Fig. 3 H, J, K, N, and O). These areas were negative for acetylcholinesterase (Fig. 3 L and M).

Fig. 3.
Comparative in vivo assessment of different cell preparations 4–6 weeks after xenografting in rat. (A) Key properties of the grafted cells (stages A to E) after transplantation. (B and C) Formation of nonneuroectodermal teratoma-like regions after ...

Using cells at stage D (DIV 45), we then explored the fate of transplanted cells over a longer period in a search for their full differentiation potential. A total of 50,000 cells were transplanted into the quinolinate-lesioned right striatum of immunotolerant nude rats that were killed after 13 to 21 weeks (n = 24). All animals undergoing transplantation had large spherical surviving grafts that compressed the host brain. Immunocytochemistry revealed large striatal donor-derived DARPP32+/HNA+ regions spread over the entire grafts (Fig. 4 A and B and Fig. S3A). DARPP32+ cells displayed a bipolar neuronal morphology with extensive neuritic outgrowth and with medium-sized cell bodies (10–16 μm in diameter) (Fig. 4C). They coexpressed the neuronal markers MAP2 (Fig. 4 D and E and Fig. S3A) and NeuN (Fig. S3B). Neurons and DARPP32+ cell-rich regions coincided, with DARPP32+ cells comprising an average of 21 ± 7% of the NeuN+ population. Although the volume of the grafts increased over time, the proportion of DARPP32+ neurons was not significantly different in brains analyzed at 13 (n = 5), 15–16 (n = 4), and beyond 17 weeks (n = 4) after transplantation (Tukey–Kramer test P = 0.05). In contrast to what we observed in short-term transplants, DARPP32+ regions were often calbindin-negative (Fig. S3 C and D). There were very few TH-positive neurons (Fig. S3E), and there was neither AChE staining nor cholinergic, serotonergic, or cortical neurons (MAP2+/Pax6+). Human GFAP+ astrocytes also were very scarce. Scattered neurites positive for human-specific nestin or human-specific NCAM were detected in host major fiber tracts, such as the corpus callosum or the pyramidal tract down to the brainstem (Fig. S3F). In contrast, there was no migration of transplanted HNA+ human cells into the host parenchyma beyond the edge of the grafts. Grafts contained numerous cells of host rat origin, in particular HNA/IB4+ microglia and endothelial cells forming blood vessels, and HNA/GFAP+ astrocytes at the graft–host interface (Fig. S4).

Fig. 4.
Grafted HNA+ cells (stage D, DIV 45) matured into DARPP32 neurons 3–5 months after transplantation. (A–C) DARPP32+ (green)/HNA+ (red) cells are (A and B) spread over the graft and display a neuronal morphology, as shown in C in a confocal ...

Xenograft Overgrowth and Persistent Proliferation of Human Neural Progenitors.

Two months and beyond after transplantation, many animals started to exhibit lethargy, rapid weight loss, and sometimes hemiparesis. At killing, these animals displayed particularly large grafts that expanded beyond the limits of the striatum. At 13 weeks, the graft volume ranged between 200 mm3 and 800 mm3. Analyses of these overgrown grafts revealed neither persistency of undifferentiated hES cells, as all grafts were Oct4-negative, nor teratoma formation, necrosis, or evidence of overt anaplasia. All grafts comprised a mosaic of two types of regions with different cellular characteristic (Figs. 4 A and D and and55D). The first region type consisted almost exclusively of nestin+ neural progenitors, many of which displayed the pan-neural marker Pax6 colocalizing with the proliferation marker Ki67 (Fig. 5 B–D). The second region type showed no Ki67+ proliferating cells and, in contrast, contained more mature neural cells, including numerous differentiated MAP2+/DARPP32+ striatal neurons (Fig. S5). There was no specific arrangement of the two types of regions and, in particular, the more differentiated ones were not preferentially localized within the periphery.

Fig. 5.
Massive proliferation of neural progenitors 3–5 months after transplantation. (A) Cresyl violet-stained coronal section (right) and outlines of serial coronal sections spanning over ≈10 mm (left) illustrating the size of the graft (red) ...

Flow cytometry analyses of the grafted cells retrieved from four animals confirmed that they were committed to the neural lineage, with 96 ± 1% CD56/NCAM+ and 86 ± 7% nestin+ cells (Fig. S6A). Q-PCR analyses were conducted to compare 3-month-old dissected grafts (n = 4) to pregrafting cells at stages D (DIV 45) and E (DIV 63). There was no apparent dedifferentiation of grafted cells, because NANOG did not increase in the grafts compared with in vitro culture (stages D and E). Early neural (SOX1) and LGE (GSH2 and DLX2) markers decreased, whereas forebrain and striatal neuronal markers (BF1, MAP2, ARPP21, and DARPP32) were significantly increased in grafts (Fig. S6B).

Four rats were killed at 13 or 15 weeks after transplantation, and their oversized graft was dissected out to perform additional in vitro analyses. Small pieces of the dissected grafts were dissociated, and cells were seeded back onto coated culture dishes containing medium supplemented with serum and N2 (n = 4) to evaluate their proliferative potential and commitment to the neural lineage. These cultures appeared fully committed to the neural lineage (nestin+ or Tuj1+) and amenable to both passaging and neuronal or glial differentiation (Fig. 5 E–H). However, the neuronal differentiation of those progenitors did not lead to a significant number of DARPP32+ neurons.

A second series of transplantations (n = 12) using similar conditions was conducted using DIV 56 cells—i.e., the upper limit of the stage D maturation time window. Results were similar to those described above for cells transplanted at DIV 45, the lower limit of stage D, in terms of both striatal fate and overgrowth.


The main result of this study is the development and validation of a protocol, both in vitro and in vivo following xenografting, to direct the differentiation of hES cells into neurons that exhibit phenotypic features of the medium-spiny GABAergic neurons (MSNs) of the striatum. Such hES-derived striatal progeny may provide a potential cell therapy product to substitute for fetal neural precursors that are currently used in clinical trials to treat patients with HD. On the path to the clinic, our results suggest that specific measures should be taken to control neural cell proliferation to ensure postgrafting safety. Substitutive cell therapy has shown some long-term success in patients when ganglionic eminences (GEs) of human fetuses were grafted (2, 3). This cell product is, unfortunately, in short supply, because GE cells are not easily propagated. This limitation has led to a yet-unfruitful search for alternative cell sources (21). Our results demonstrate that this goal could be achieved by using hES cells as a source.

MSN differentiation occurs in a large proportion of neurons when hES cells are treated in vitro with our protocol. Progression along the striatal lineage is characterized by the timed expression of specific developmental genes, such as GSH2 and DLX2, and the detection of DARPP32 protein in terminally differentiated neurons. During early development, immature neural progenitors bear, by default, a rostral fate (22, 23). Consistently, we found that the neuroectodermal cells produced up to the rosette stage express genes associated with a forebrain fate, such as BF1. The antagonizing actions of SHH and Wnt signaling play a major role in the fine tuning of subregional cell fate in the developing forebrain, as demonstrated in many mammals, and most recently by using ES cells (8, 14, 16, 17). We made use of and confirmed these data to obtain a subregionalization to ventral telencephalic structures—in particular the GE—by treating rostral neural cells with SHH and DKK1, a Wnt pathway inhibitor. We obtained a large number of cells displaying a MSN phenotype after grafting using progenitors retrieved at our stage D, which is equivalent to 7.5–9.5 weeks after conception. Three months after grafting using stage D cells, the yield of DARPP32+ neurons was high, in the range of several hundred thousand striatal neurons following implantation of 50,000 neural progenitors—i.e., probably relevant to a clinical application. It is interesting to note that this staging matches the time window used for human fetal cell retrieval for grafts in HD patients (3, 24), suggesting that the striatal and neuronal differentiation in vitro bears some similarity to the actual chronobiology of the ontogenetic process. Consequently, longer-term studies allowing to follow up grafted cells up to establishment of a full set of connections—which may occur as late as more than 1 year after grafting, taking into account data from fetal neural transplants (25)—would be of prime interest to determine the final differentiation stage of ES-derived neurons.

However, despite these promising data, results of the long-term transplant experiments suggest the existence of a potential roadblock on the path to the clinic, because the human xenografts systematically overgrew in the rat brain over time. This overgrowth was massive, with up to several thousand-fold increase in cell content, and rapid, because it was visible already at 4 to 6 weeks and provoked clinical symptoms within 3–5 months. Xenograft overgrowth following intracerebral transplantation of neural progenitors derived from two hES cell lines (H1 and H9) different from the one we used has been reported recently (9), and compressive grafts also were mentioned following implantation in the rat brain of nonhuman primate ES cell-derived neural progenitors (19, 26). This adverse effect is not related, therefore, to one particular hES cell line or to the human species.

One main question raised by these data deals with the “physiological” or otherwise “pathological” nature of these large grafts—i.e., whether human neural progenitors are simply proliferating as they are supposed to, or else have lost their responsiveness to signals that normally switch off their proliferation. Our analysis of the phenotype of the proliferating cells (in vivo) underlined that they were typical neural progenitors with no apparent phenotypic abnormality. They were capable of differentiating into neurons and astrocytes after replating in vitro. These results were consistent with those recently presented by Roy et al. (9) on dopaminergic neural xenografts. Still, these authors showed that their dopamniergic transplants were intermingled with the host tissue, whereas proliferating progenitors occupied the graft core. This observation suggested that host cells might deliver differentiation cues inaccessible to the proliferating cells of the inner core and led these authors to support the hypothesis of a pathologic state of the proliferating cells. In the present study, this suggestive topographic arrangement was not observed, and regions containing either proliferative or differentiated cells were interspersed. Despite the decreased expression in early neural markers (e.g., SOX1) in the 3- to 5-month-old graft compared with stage A–D cultures, we cannot exclude that graft overgrowth is at least in part due to the presence of a small proportion of early-stage “rosette-like” precursors not yet committed to a striatal fate that would have remained undetected. Besides, in our case, the chronobiology of the ontogenesis in the GE supports the physiological hypothesis. Indeed, progenitors in these germinative zones not only participate in striatal neurogenesis up to the 15th week of gestation but also give rise to many cortical interneurons over several months and to gliogenesis well beyond (2729). The fact that cells replated in vitro from overgrown grafts differentiated normally into neurons and glia but failed to give rise to MSN, the neurogenesis time schedule of which had passed, is in agreement with this suggestion. It is interesting to mention that human-to-rat xenotransplantation of fetal GE tissue led to overgrowth (30) and that areas containing potentially proliferating cells were observed in grafts analyzed in a patient autopsied 18 months after grafting (25). Apparent overgrowth of human xenograft in rat brain may, therefore, only be a normal consequence of the physiological capacity of proliferation of human neural precursors, essentially revealed because of the size difference between rat and human brain. Nevertheless, as one may not exclude some impact of this phenomenon in human allografts, this calls for specific precautionary action on the path to the clinic, allowing for blocking precursor proliferation if need be. Potential action may involve isolation of pure striatal population with limited or no proliferative capacity, preemptive in vitro treatment of the grafted cells with antimitotic agents, or genetic engineering of the transplant to provide a means to eliminate unwanted dividing cells in vivo, such as by using a suicide gene paradigm.

Materials and Methods

Human ES Cell Culture.

Human ES cells (SA-01, XY, passages 20–80; Cellartis; and H9, XX, passages 40–60; WiCell Research Institute) were maintained on a layer of mitotically inactivated STO (ATCC no. CRL-1503) feeders. The hES cells were cultured in DMEM/F12 Glutamax supplemented with 20% knockout serum replacement, 1 mM nonessential amino acids, 0.55 mM 2-mercaptoethanol, and 10 ng/ml recombinant human FGF2 (all from Invitrogen). Cultures were fed daily and manually passaged every 5–7 days.

Induction and Differentiation of Striatal Progenitors.

Induction of striatal progenitors and GABAergic neurons from hES cells was performed by using a modification of a previously described protocol (7). Briefly, hES cells were plated on mitotically inactivated murine bone marrow-derived stromal feeder cells MS5 in serum replacement medium containing DMEM/F12 supplemented with 15% knockout serum replacement, 1 mM nonessential amino acids, and 0.55 mM 2-mercaptoethanol (all from Invitrogen). After 12 days in these conditions, serum replacement medium was replaced with an N2 medium that consisted of DMEM/F12 supplemented with insulin (25 μg/ml), transferrin (50 μg/ml), putrescine (100 μM), selenium chloride (30 nM), and progesterone (20 nM) (all from Sigma–Aldrich). Medium was changed every 2–3 days, and growth factors were added as described: 200 ng/ml SHH, 100 ng/ml DKK1, 20 ng/ml BDNF (R & D Systems), 0.5 mM dbcAMP (Sigma–Aldrich), and 0.5 mM valpromide (Lancaster Synthesis). After ≈3 weeks of differentiation, rosette structures were collected mechanically and transferred to 15 μg/ml polyornithine/1 μg/ml laminin-coated culture dishes in N2 medium supplemented with SHH, DKK1, and BDNF (passage 1). After 8–12 days, cells were exposed to Ca2/Mg2-free Hanks balanced salt solution (HBSS) for 3 h at 37°C, spun at 150 × g for 5 min, resuspended in N2 medium, and then plated onto coated culture dishes in the presence of SHH, DKK1, and BDNF (passage 2). After another two or three passages in the same conditions, cells were differentiated by replating them onto coated culture dishes (25–50 × 103 cells per cm2) in the presence of BDNF, dbcAMP, and valproic acid. Approximately 104 hES cells were plated (three colonies 2–3 mm in diameter) per 6-cm plate of MS5 feeder. At stage A (3 weeks in vitro), between 500,000 and 1 million neural progenitors could be harvested. These neural cells were expanded (first passage plating ratio 1:1, following passage plating ratio 1:3: 105 cells per cm2) up to stage D. After 45–60 days of culture, the initial 104 hES cells may generate up to 3 × 106 cells ready to be grafted, corresponding to an average doubling time of 4–5 days.

Real-Time Q-PCR.

Total RNA was isolated by using the RNeasy Mini Kit according to the manufacturer's instructions (Qiagen). A total of 1 μg of RNA was reverse transcribed into cDNA with SuperScript II (Invitrogen) using random primers. Real-time Q-PCRs were performed with Power SYBR Green PCR Mix (Applied Biosystems) and a Chromo 4TM Real-Time system (Bio-Rad). The amplification efficiency of each pair of primers was determined by comparison with a standard curve generated with serially diluted cDNA of fetal brain (FB) (Clontech). Quantification was performed at a threshold detection line (Ct value). The Ct of each target gene was normalized against that of the cyclophilin housekeeping gene. The 2−ΔΔCt method was used to determine the relative level of expression of each gene, and FB cDNA was used as calibrator. Data were expressed as mean ± SEM. Primer sequences are listed in Table S1.

Quinolinic Acid Lesion and Cell Transplantation.

All animal experiments were conducted in accordance with the Direction des Services Vétérinaires, Ministère de l'Agriculture of France, and with the European Communities Council Directive (86/609/EEC). Adult OFA and Nude rats (weight 220–260 g at the time of grafting; Charles River Laboratories) were used. All surgical procedures were achieved under full anesthesia using a mixture of ketamine (15 mg/kg) and xylazine (3 mg/kg; Bayer Health Care) and using a stereotaxic frame. Unilateral lesions were made by injecting 1 μl of 80 nmol/μl quinolinic acid dissolved in 0.1 M PBS into the right striatum, according to the following coordinates (in mm): anteroposterior (A) = +0.3; lateral (L) = −2.7; ventral (V) = −4.7; and tooth bar = −3. One week after the lesion, rats received transplants of cells (50,000-200,000 in 2 μl of HBSS supplemented with 0.05% DNase I; Invitrogen) (A = +0.3; L = −3; and V = −5.5 and −4.5).

Tissue Processing.

At different times after transplantation, rats were terminally anesthetized with 1 g/kg i.p. sodium pentobarbital (Ceva Santé Animale), and their brains were fixed by transcardial perfusion with 100–150 ml of 0.1 M PBS (pH 7.4), followed by 250 of ml buffered 4% paraformaldehyde (PFA). Brains were removed, postfixed overnight at 4°C in 4% PFA, and then cryoprotected in 30% sucrose solution at 4°C. Coronal brain sections (50 μm) were cut on a freezing Microtome, collected serially (interspace, 600 μm), and stored at −20°C in a cryoprotectant solution until analysis.

For in vitro culture, FACS (see SI Methods), and gene expression analyses of 3-month-old grafts, rats were transcardially perfused with PBS, brains were removed, and the grafted cells were dissociated in HBSS. Cells were cultured on coated culture dishes in N2 medium containing 2% FBS and were passaged every week with trypsin (Invitrogen). Terminal differentiation was induced by replating the cells at lower density in N2 medium supplemented with 20 ng/ml BDNF.

Immunocytochemistry and Immunohistochemistry.

Cells were fixed with 4% PFA for 10 min at room temperature and incubated with primary antibody overnight at 4°C. Secondary antibodies and DAPI counterstain were applied for 1 h at room temperature. Brain sections were incubated with primary antibodies for 36 h at 4°C. Secondary antibodies and DAPI counterstain were applied for 3 h at room temperature.

Both cells and tissue were stained with the primary antibodies listed in SI Methods. Confocal analysis was performed on a Leica TCS SP2 microscope (Leica Microsystems).


Quantitative immunocytochemical analysis was performed on randomly selected visual fields from at least two independent differentiation experiments. In each field, images of separate channels were acquired at 20× magnification on a fluorescence microscope (ImagerZ1; Carl Zeiss) using the Axiovision image capture equipment and software and exported to an imaging software (Adobe Photoshop; Adobe Systems), where separate channel images as well as the corresponding overlaid images were counted. On average, 10 visual fields were acquired per 16-mm coverslip, and a total of 100–500 cells were counted per field. The total number of cells expressing the different markers was plotted as a percentage for either all cells (DAPI) or neurons (MAP2+ cells).

Graft volumes were estimated by using Histolab software (Microvision Instruments). In the grafts, cell number was quantified per uniform randomly selected section within the region of interest. Data are presented as mean ± SEM.


See SI Methods.

Statistical Analysis.

The data were processed using JMP7 software (SAS). Values are reported as mean and SEM. Comparisons among values for all groups were performed by one-way ANOVA, and Tukey–Kramer multiple-comparison test was used to determine the level of significance.

Supplementary Material

Supporting Information:


We thank Stéphane Supplisson, Marc Lechuga, Michael Melkus, and Daniel Stockholm for their assistance. This work was supported in part by additional grants from the European Commission (STEM-HD, FP6), Genopole, and the cluster MediCen Paris Region (IngeCELL). L.A. is the recipient of fellowships from the Fondation pour la Recherche Médicale (Prix Pomaret Delalande 2005), Association Huntington France, and Fédération Huntington Espoir.


The authors declare no conflict of interest.

This article contains supporting information online at www.pnas.org/cgi/content/full/0808488105/DCSupplemental.


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