Cerebral Cortex Advance Access originally published online on June 15, 2005
Cerebral Cortex 2006 16(3):301-312; doi:10.1093/cercor/bhi120
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Feature Article |
Cell Proliferation in the Adult Hippocampal Formation of Rodents and its Modulation by Entorhinal and Fimbria–Fornix Afferents
1 Development and Regeneration of the CNS, Department of Cell Biology, Barcelona Science Park-IRB, University of Barcelona, E-08028 Barcelona, Spain and 2 Cellular Neurobiology, Cell Biology Department, University of Valencia, Dr. Moliner, 50, Burjassot, E-46100, Spain
Address correspondence to J.A. del Río, Development and Regeneration of the CNS, Department of Cell Biology, Barcelona Science Park, University of Barcelona, Josep Samitier 1–5, E-08028 Barcelona, Spain. Email: jario{at}pcb.ub.es
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Abstract |
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New granule neurons are produced in the dentate gyrus (DG) of rodents throughout adult life. Recent studies have also reported adult neurogenesis in the cerebral cortex in normal animals or after brain injury. However, few of these studies focused on the hippocampal formation (HF), a cortical area involved in learning and memory in which extensive cell death occurs in neurodegenerative diseases. Thus, we studied cell proliferation in the HF of rodents and the intrinsic putative neurogenic potential of entorhinal cortex (EC) progenitors. We show that only the DG generates new neurons in the HF. In addition, neurospheres from the EC differentiate into neurons and glia in vitro and after transplantation in the adult DG. We also analyzed whether the absence of the synaptic input from the main hippocampal afferents induces neuronal generation in the HF outside the DG and/or regulates the proliferation of DG neuroprogenitors. We show that the denervation of the hippocampus does not induce neurogenesis in HF regions other than the DG. However, neuroprogenitor proliferation in the DG is reduced after fimbria–fornix lesions but not after entorhinal deafferentation, which supports the view that neuroprogenitor proliferation and/or differentiation in the DG are controlled from basal forebrain/septal neurons.
Key Words: fimbria–fornix lesion • hippocampal formation • NG2-positive cells • neural progenitor cells • perforant pathway lesion
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Introduction |
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TopAbstractIntroductionMaterial and MethodsResultsDiscussionSupplementary MaterialReferences |
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In the development of the rodent hippocampal formation (HF), pyramidal neurons in the entorhinal cortex (EC) and hippocampus are generated prenatally, whereas granule cells in the dentate gyrus (DG) are mainly generated postnatally (Altman and Das, 1965
, 1966; Schlessinger et al., 1975). Moreover, as well as in the subventricular zone surrounding the anterior lateral ventricles (Lois and Álvarez-Buylla, 1994; Álvarez-Buylla and García-Verdugo, 2002), neurogenesis occurs in the innermost portion of the granule cell layer (GCL) of the dentate gyrus (DG), the subgranular zone (SGZ), throughout adult life (Kuhn et al., 1996; Palmer et al., 1997; Cameron and McKay, 2001; Seri et al.., 2001; van Praag et al., 2002). Adult neuronal production in the DG has been documented in many vertebrates, including rodents (e.g. Cameron and McKay, 2001; Dayer et al., 2003), non-human primates (Gould et al., 1999a, 2001; Kornack and Rakic, 1999) and humans (Eriksson et al., 1998). In recent years, three studies reported adult neuronal generation under non-pathological conditions in the hippocampus proper and the neocortex of rodents (Rietze et al., 2000, Dayer et al., 2005), and in the primate neocortex (Gould et al., 1999b; for further discussion, see also Rakic, 2002). In contrast, several studies have demonstrated that the adult cortex of rodents only generates new neurons after injury (e.g. Magavi et al., 2000; Zhang et al., 2001; for a review of ischemia, stroke or target-directed apoptosis, see Arlotta et al., 2003). As well as these data, proliferating cells identified as glia through several techniques [e.g. tritiated thymidine or bromodeoxyuridine (BrdU) labeling] were identified several years ago in almost all forebrain regions under non-pathological conditions (e.g. Kaplan and Hinds, 1980; Palmer et al., 1999; see also Korr, 1980).
Precise control of adult neurogenesis in the DG appears to play a critical role in the regulation of hippocampal functions (for recent reports, see Gould et al., 1999c; Bruel-Jungerman et al., 2005). Many factors, including endocrine, neural or physiological conditions (e.g. stress; Gould and Tanapat, 1999), greatly influence (negative or positively) the generation and survival of late-generated neurons in the DG (for a review, see Fuchs and Gould, 2000). For example, adrenal steroids inhibit progenitor proliferation in the adult rodent hippocampus through an NMDA-receptor mediated pathway (Cameron et al., 1995, 1998; Gould and Cameron, 1997; Gould et al., 1997). Therefore, glutamatergic input is believed to be a negative regulator of cell proliferation in the DG (Cameron et al., 1995). Indeed, i.p. administration of NMDA competitive antagonists, such as CGP43487, increases cell proliferation in the DG (Gould and Cameron, 1997; Nacher et al., 2001). In addition, brief electrical stimulation of the entorhino-hippocampal pathway (EHP) (Parent et al., 1997), hippocampal kindling stimulation (Bengzon et al., 1997) and several kinds of CNS injuries (e.g. ischemia, Yagita et al., 2001; epileptogenic paradigms, Nakagawa et al., 2000; Parent, 2002) all increase the proliferation of the neuroprogenitors of adult DG (for a review, see Parent and Lowenstein, 2002). Taken together, these data indicate that synaptic activity may also participate in the regulation of the neuroprogenitor proliferation in the DG and/or the survival of adult-generated neurons.
The major hippocampal afferents are the EHP, the excitatory commissural/associational connections and the basal forebrain/septal projection. They terminate in a laminated fashion on their target neurons: the granule cells in the DG and the pyramidal neurons in the hippocampus proper (Amaral and Whiter, 1995). Extensive cell loss and axonal deficits occur in the HF in neurological disorders linked to memory deficits, such as Alzheimer's disease (Gomez-Isla et al., 1996), as well as in normal aging (Squire, 1992). It is well established that induction of synaptic-specific impairment in the HF (e.g. after perforant pathway lesion) is a good model for examining the synaptic changes that trigger reactive responses in neurons and glial cells in the hippocampus (for a review, see Turner et al., 1998). Indeed, hippocampal denervation induced profound transneuronal reorganization (e.g. reactive synaptogenesis) and a new physiological balance in both projecting neurons and target cells in the hippocampus, which in turn may induce or modulate neuronal production in the region.
The aims of this study were twofold. First, we analyzed in detail cell proliferation in the hippocampus and EC of adult healthy mice by BrdU-labeling methods. In addition, neural progenitors were isolated from the EC and maintained as floating aggregates to characterize their differentiation potential in vitro and after transplantation in vivo. Second, we analyzed whether neurogenesis could take place in the HF outside the DG after axotomy of the main extrahippocampal afferents (entorhinal pathway and basal forebrain/septal afferents) and whether neuroprogenitor cell proliferation in the DG is modulated in the absence of these synaptic inputs. After BrdU-labeling experiments in normal and denervated hippocampus in vivo, we report that the potential to generate new neurons only exists in the SGZ of the DG. However, neural proliferating cells from the EC form neurospheres, which differentiate to give rise to neurons and glial cells in vitro and after transplantation to the adult DG. Finally, we found that axotomy of the entorhinal afferents did not alter the proliferation of neuroprogenitor cells in the DG 1 week later. However, their proliferation dropped greatly after fimbria–fornix (FF) lesion, which implies extrahippocampal control of dentate progenitor cells by factors reaching the hippocampus via FF afferents.
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Material and Methods |
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All procedures were performed in accordance with guidelines approved by the Spanish Ministry of Science and Technology, following European standards. A total of 73 OF-1 adult and 15 postnatal mice (P5–P15 days old) (Iffra Credo, Lyon, France) and 10 Wistar rats (3 months old) were used. We also used Tau-GFP transgenic mice (Pratt et al., 2000
) (n = 10) as an alternative source of neurospheres for transplantation studies. Antibodies
Rabbit polyclonal antibody against glial fibrillary acidic protein (GFAP, 1:500, Dakopatts, Glostrup, Denmark) was used to label astrocytes; monoclonal antibody against class ßIII-tubulin isoform (TUJ1, diluted 1:1000, Babco, Richmond, VA), to detect the cytoskeletal protein expressed by young postmitotic neurons; and rabbit polyclonal antibody against NG2 (diluted 1:200, Chemicon, Temecula, CA) to detect oligodendrocyte precursors. Rat monoclonal antibody anti-BrdU (diluted 1:50, Harlan Sera-Lab, Loughborough, UK) and mouse monoclonal antibody anti-BrdU (diluted 1:50, Dakopatts) were also used. Mouse monoclonal antibody (IgM subtype) anti-PSA-NCAM (kindly provided by Dr G. Rougon, diluted 1:5000) was also used. Mouse monoclonal antibody against NeuN (diluted 1:50; Chemicon) was used to label postmitotic neurons. Rabbit anti-Calbindin (diluted 1:1000; Swant, Bellinzona, Switzerland) was used for the detection of mature granule cells. Rabbit polyclonal antibody against rat collapsing response-mediated protein (rCRMP-4/TUC-4; 1:5000, Chemicon) was used to label neural progenitors.
Neurosphere Isolation and Differentiation In Vitro and In Vivo
For neurosphere preparation, animals (P5–P15) were anesthetized by hypothermia and their brains were aseptically removed from the skull. After rinsing in 0.1 M phosphate-buffered saline (PBS) containing 6.5% glucose, the meninges were removed and the EC and the hippocampus were dissected. Tissue pieces were rinsed in Hank's balanced salt solution (HBSS without Ca2+, Mg2+ and phenol red), containing 0.01 M HEPES and 6.5 mg/ml glucose (HBSS–HEPES–glucose) in 90 mm diam. Petri dishes. After rinsing, slices (300–400 µm thick) of tissue containing both the hippocampus and the EC were taken in the horizontal plane through the caudoventral pole of the cerebral hemisphere using a McIlwain tissue chopper (for details, see Li et al., 1994). Afterwards, slices were selected and transferred to cold HBSS–HEPES–glucose buffer. The main anatomical features of the EC and the hippocampus, including the pyramidal layer, the fimbria and the subiculum, were clearly visible under dark-field optics. The hippocampus and EC were then separated and stored in separate Petri dishes at 4°C. The hippocampus proper was microdissected from the hippocampal slices using fine surgical instruments. For the EC, small tissue explants containing the lateral and the medial entorhinal area were separated from the adjacent isocortex and subicular region using the rhinal fissure as anatomic reference (see Fig. 2). Next, the white matter was removed from these explants by microdissection using fine tungsten knifes. In parallel experiments, the dissected pieces of the EC were divided into two sections containing either upper (from layer I to lamina disecans) or lower (from layer V to white matter) layers (see Fig. 2).
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Dissected tissue pieces from each region were minced into small pieces, rinsed twice in HBSS–HEPES–glucose and trypsinized for 15–20 min at 37°C. Trypsinization was stopped by addition of horse serum (one-quarter volume). After centrifugation, cells were dissociated by trituration 40 times in 0.1 M HBSS–HEPES–glucose containing 0.025% DNase with a fire-polished Pasteur pipette. Dissociated cells were collected by centrifugation at 800 r.p.m. for 5 min, counted and cultivated in 25 ml flasks (1 x 106 cells/flask; Nunc, Roskilde, Denmark) in basic medium containing nutrient mixture F-12 (HAM) and DMEM (Dulbecco's modified Eagle's medium) in the proportion 3:1. Culture medium was supplemented with B27, antibiotics, and 20 ng/ml FGF2 and EGF (Sigma-Aldrich, St Louis, MO). Every week, growing neurospheres were mechanically dissociated and replated in fresh medium. Culture media and supplements were purchased from GIBCO Life Technologies (Merelbeke, Belgium) unless otherwise indicated.
For Western blotting analysis, neurosphere protein samples were obtained in Laemmy sample buffer, electrophoretically separated in 8–10% sodium dodecyl sulfate–polyacrylamide gels and electrotransferred to nitrocellulose membranes. Membranes were blocked with 10% non-fat milk in 0.1 M Tris-buffered saline (pH 7.4) for 2 h and incubated overnight in 0.5% blocking solution containing primary antibodies. After incubation with peroxidase-tagged secondary antibodies, peroxidase activity was developed by an ECL-plus chemiluminescence Western blotting kit (Amersham Biosciences, Buckinghamshire, UK).
For differentiation experiments, neurospheres were left to grow for 10–15 days after mitogen withdrawal on coverslips (12 mm diam.) coated with poly-L-ornitine and laminin (Sigma-Aldrich) in serum-free medium (DMEM containing glutamine, B27 and antibiotics). After differentiation, cultures were fixed with 2% phosphate-buffered paraformaldehyde for 1 h at 4°C. Coverslips were then processed for the immunocytochemical detection of ßIII-tubulin (TUJ1), MAP2, GFAP or NG2, using Alexa-Fluor 488 and 568 tagged secondary antibodies (Molecular Probes, Eugene, OR). After rinsing, cell nuclei were counterstained with bisbenzimide (Hoechst 32444, 1 µM in 0.1 M PBS, 10 min) and the coverslips were mounted in Fluoromount (Vector Labs, Burlingame, CA).
For transplantation experiments, neurospheres from the EC of Tau-GFP transgenic mice were isolated and expanded in vitro, as described above. After several passages (4–5), neurospheres were mechanically dissociated and transplanted into the DG (n = 4 animals, bregma = –1.4 mm, lateral = 1 mm, deep = 2 mm; Slotnick and Leonard, 1975) or the lower layers of the neocortex (n = 5 animals, bregma = –0.7 mm, lateral = 3 mm, deep = 1.5 mm) of wild-type (GFP-negative) mice. After 10–15 days, transplanted mice were perfused with 4% paraformaldehyde and their brains were cryoprotected with a 30% saccharose solution in 0.1 M PBS and processed.
Surgical Procedures
Adult OF1 mice (n = 39, 3 months old) were lesioned unilaterally in the perforant pathway (n = 29) or FF (n =10) with a wire-knife (David Kopf Instruments, Consultants GmbH, Düsseldorf, Germany), following Slotnick and Leonard stereotaxic atlas. Briefly, for perforant pathway lesions, mice were placed in a stereotaxic apparatus after pentobarbital anesthesia (50 mg/kg body wt) and drilled at 0.75 mm caudal and 3 mm lateral from lambda (dissection scheme in Supplementary Fig. 1). A closed wire-knife was then inserted at a 14° lateral angle. Four millimeters ventral to the dura, the wire was unfolded 1.5 mm and the perforant pathway was sectioned by retracting the knife 3 mm. For FF lesions, the knife was inserted 0.5 mm posterior and 1.8 mm lateral from bregma. Four millimeters ventral to dura, the wire was unfolded 1.5 mm and the FF was sectioned by retracting the knife 3 mm upwards. After a range of survival times (4, 7 and 15 days), both lesioned and sham-operated mice were transcardially perfused and processed. Nissl-staining and immunocytochemical methods against GFAP (Supplementary Fig. 1) or PSA-NCAM (Fig. 4) in horizontal sections, 4 and 7 days after axotomy, showed that lesions were mainly confined to the lower portion of the EC and few extended to the subicular region (PaS and PreS). For quantitative analysis, mice with lesions affecting the upper layers of the EC, the DG or the hippocampus proper were discarded. For FF lesions, only unilateral complete section of the fornix columns was considered.
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BrdU Experiments
Control Mice and Rats
Twelve OF-1 mice received a daily BrdU (Sigma-Aldrich) pulse by i.p. injection of 50 mg/kg body wt for 4 days. BrdU-injected mice were assigned to three equivalent experimental groups which were sacrificed 4 h, 7 days or 15 days after the last BrdU injection. To study possible cortical neurogenesis, one group of five mice were injected daily with 50 mg/kg body wt of BrdU for 15 days and sacrificed 15 days later. In parallel experiments, adult rats (n = 10, 3 months old) were injected twice daily with 50 mg/kg i.p. for 2 days and were sacrificed 21 days later. Rodents were perfused with 4% buffered paraformaldehyde and fixed brains were post-fixed in the same fixative solution for an additional 2.5 h. After fixation, brains were cryoprotected and frozen in Isopentane solution for storage at –80°C. Brains were horizontally sectioned using a freezing microtome (30 µm thick) and sections were preserved at –20°C in cryoprotecting solution. Free-floating sections were further processed for immunohistochemistry.
Lesioned Mice
All operated mice received a single i.p. injection of BrdU per day (50 mg/kg body wt) before (pre-lesion labeling group, protocol used for both EHP and FF lesion) or after (post-lesion labeling group, only for EHP lesion) axotomy. A total of 47 adult mice (3 months old) were used.
Thus, 22 mice (12 for EHP axotomy and 10 for FF lesion) received the thymidine analog three days before the lesion and were sacrificed on post-lesion days 4 (six for EHP axotomy and three for FF lesion), 7 (six for EHP axotomy and three for FF lesion) or 15 (four for FF lesion). For the post-lesion labeling group (n = 17 for EHP), 13 mice received a BrdU pulse-injection every 24 h after lesion (for 3 days) and were processed the following day (4 days after lesion; DAL). In addition, four lesioned mice received BrdU injections 4, 5 and 6 DAL and were sacrificed 7 DAL.
Given that EHP in mice is mainly an ipsilateral projection and has an almost negligible contralateral component to the hippocampus (van Groen et al., 2002, 2003), in this study the contralateral hemisphere to the lesion was considered an internal control. In addition, eight mice were prelabeled with BrdU as above, and sham-operated and processed 4 and 7 days later. At the distinct survival times, the BrdU-positive and PSA-NCAM-positive cell counts in the two hemispheres were compared with those obtained in sham-operated and control (unlesioned) mice. We also performed double BrdU/NeuN immunohistochemistry in lesioned mice to analyze whether the injury induces neurogenesis in the HF outside the DG or not.
Immunohistochemical Procedures
Free-floating sections were processed to detect neuronal (PSA-NCAM and NeuN) or glial (GFAP and NG2) antigens previously rinsed in 0.1 M PBS and incubated in a blocking solution containing 10% normal serum for 2 h. For peroxidase staining, endogenous peroxidase activity was inhibited by a solution with 10% methanol and 3% hydrogen peroxide in 0.1 M PBS for 15 min. After blocking, sections were incubated overnight with primary antibodies at 4°C. All primary antibodies were diluted in 0.1M PBS containing 5% normal serum, 0.2% gelatine and 0.5% Triton X-100; tissue-bound primary antibody was detected with the ABC method (Vector Laboratories) after incubation for 2 h with biotinylated goat anti-mouse IgM (Chemicon, 1:500), goat anti-rabbit or goat anti-mouse secondary antibodies (1:150, Vector Laboratories). Peroxidase activity was revealed with 0.03% diaminobenzidine (DAB) and 0.01% hydrogen peroxide. Afterwards, sections were mounted on gelatinized slides, dehydrated and coverslipped with Eukitt (Merck, Darmstadt, Germany).
For BrdU immunohistochemistry, sections were processed essentially as described (del Río and Soriano, 1989). Briefly, sections were pretreated with cold 0.1 N HCl for 15 min and 2 N HCl at 37°C for 20 min to denature DNA. After rinsing in 0.1 M PBS, they were incubated with a Fab goat anti-mouse IgG (1/50 diluted; Jackson Immunocytochemical) for 2 h and incubated with primary anti-BrdU antibodies (see above). Tissue-bound primary antibody was detected using a biotinylated secondary antibody and the ABC method with DAB–Ni as chromogen. Parallel sections were stained with cresyl violet or processed for the double immunofluorescence detection of BrdU combining GFAP, NeuN, PSA-NCAM or NG2, using Alexa-Fluor 488 and Alexa-Fluor 568 tagged secondary antibodies (Molecular Probes, Eugene, USA). Double rCRMP-4/BrdU immunohistochemistry was performed by simultaneous incubation with both primary antibodies, as described by Soriano et al., 1991. Sections were mounted on Fluoromount and analyzed on an Olympus Fluoview SV 500 confocal microscope. All images were obtained in sequential scanning laser mode to avoid fluorochrome cross-excitation and further processed with Silicon Graphics Imaris software to obtain three-dimensional orthogonal projections.
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Cell Proliferation in the Adult Hippocampal Formation of the Mouse
Cell proliferation in the adult HF in situ was monitored by BrdU incorporation into mitotically active cells 4 h, 7 days or 15 days after pulse injections of BrdU over 4 days (Fig. 1). In all cases, BrdU-positive cells were observed in the SGZ, the GCL, the hilus and molecular layer (ML) of the DG and the fimbria, and were scattered throughout the cornus ammonis regions 1–3 of the hippocampus proper (Fig. 1A). They were also detected in the periventricular region of the hippocampus with increasing numbers of positive cells at dorsal hippocampal levels (not shown). In addition, small BrdU-positive cells (10 µm diam. nuclei main axis) were abundant in the EC (Fig. 1B), frequently contiguous to large NeuN-positive cells in double immunoreacted sections (Fig. 1C). Double-labeling with BrdU and glial antigens (GFAP or NG2) demonstrated that the majority of BrdU-labeled cells in the EC were NG2-positive (68% after 15 days of the last BrdU injection; Fig. 1D) and a small percentage of the remaining BrdU-labeled cells were GFAP-immunoreactive (7% after 15 days; Fig. 1E), whereas none of the BrdU-positive cells in the EC were immunopositive for NeuN at any stage studied (800 BrdU-positive cells analyzed). In parallel experiments performed in rats, sections were processed to rCRMP-4 (Fig. 1F,G) 21 days after BrdU labeling (see Material and Methods for details). Double rCRMP-4/BrdU positive cells were found only in the DG (Fig. 1G), while in the EC multipolar rCRMP-4-positive/BrdU-negative cells were observed (Fig. 1F). To avoid underestimation of BrdU-labeled proliferating cells and to determine whether BrdU-positive cells give rise to neurons in the HF outside the DG as suggested for other regions (Kaplan, 1981; Rietze et al., 2000; Dayer et al., 2005), we also performed long-term BrdU-labeling experiments over 15 days (see Material and Methods for details). Again, we were unable to observe any double-labeled NeuN–BrdU cell in the EC or the hippocampus proper 15 days after the last BrdU injection (700 BrdU-positive cells analyzed) (Fig. 1H). Taken together, these results demonstrate and confirm that the potential to generate new neurons in the normal rodent HF is restricted to the SGZ of the DG.
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Finally, we examined the evolution of the number of BrdU-labeled cells in the EC, hippocampus proper and DG, 4 h, 7 days and 15 days after a 4-day BrdU-labeling protocol (Fig. 1I). The highest BrdU-positive cell numbers in the EC and hippocampus proper were detected 4 h (17.4 ± 2.4 for the EC and 23.5 ± 1.6 for the hippocampus proper) post-injection. In contrast, a significant increase (1.53 fold increase) in the number of BrdU-positive cells was observed between 4 h (10.3 ± 1.62) and 7 days (15.8 ± 2.5) post-injection in the DG. However, all three regions decreased the number of BrdU-positive cells 15 days after BrdU-labeling: by 50% in the EC and hippocampus proper and by 25% in the DG which may reflect cell death or nucleotide dilution.
Isolation of Entorhinal and Hippocampal Progenitors and Differentiation In Vitro and after Transplantation In Vivo
As the white matter of the corpus callosum contains cells with neurogenic potential (e.g. Palmer et al., 1999; Marshall et al., 2003), we first compared the number of neurospheres obtained from the upper (layer I to lamina disecans) or lower (V to white matter) layers of the EC one week after the isolation and expansion of the progenitor cells (Fig. 2A–C). Dissociated entorhinal cells from lower cortical layers generated eightfold more neurospheres in the presence of FGF2 and EGF than upper dissected cells (Fig. 2B), which supports the view that dissected regions containing the remnant of the developmental neurogenic zone (e.g. white matter) hold large numbers of multipotent precursors (see Palmer et al., 1999; Marshall et al., 2003). Thus, to avoid interference with these precursors, we removed the white matter of the EC and only neurospheres obtained from the parenchyma (Fig. 2C) were used in the following experiments.
EC-derived (entNPC) and hippocampus-derived (hipNPC) neurospheres were maintained for 
1 year (>60 passages, one passage a week). Nine days after the last passage, both types of neurospheres were allowed to differentiate in vitro for 10–15 days in serum-free media without growth factors (Fig. 2E,F) and the relative proportions of TUJ1-, MAP2-, GFAP- and NG2-positive cells were counted (Fig. 2G). Relative percentages of TUJ1- or MAP2-positive neurons and GFAP-positive cells were similar in entNPC and hipNPC (TUJ1: 7.9 and 6.1%, MAP2: 8.4 and 6.5%, GFAP: 50 and 55%, respectively). However, NG2-positive cells were significantly more abundant in entNPC- than in hipNPC-derived cultures (2.52-fold increase). To ascertain whether cells forming neurospheres were from endogenous proliferating glial cells (mainly oligodendroglial precursors), as observed in vivo (see above), we measured NG2 in protein extracts taken from EC neurospheres cultured 3 and 7 days after tissue dissociation by Western blot. The 200–220 kDa NG2 band was detected 3 days post-dissection, which suggests that NG2-positive cells are present in the early formed neurosphere (Fig. 2D). In addition, both neuronal and glial differentiation occurred in the floating entNPC early after passage, since GFAP and TUJ1 antigens were also detected by Western blot 3 days post-passage (Fig. 2D). These results show that EC contains endogenous neural progenitors, which can be cultured long-term as floating neurospheres and which differentiate into astroglia, oligodendroglia and neurons in vitro (Fig. 2E,F).
Finally, we analyzed the lineage potential in vivo of progenitors from the EC by transplanting neurospheres obtained from Tau-GFP transgenic mice into the DG and the neocortex of wild-type (GFP-negative) adult mice (Fig. 2H–J). Fifteen days post-transplantation, double-labeled GFP-positive and NeuN- or Calb-positive neurons were observed in the SGZ of the hippocampus. In contrast, GFP-positive progenitors differentiated exclusively in GFAP-positive cells when transplanted into the neocortex (not shown). Thus, these data indicate that neural progenitors of the EC are able to differentiate into neurons when transplanted into the adult DG, which may well contain specific factors responsible for this differentiation.
Cell Proliferation in the Hippocampal Formation of the Mouse after Axotomy of the EHP
Stereotaxic lesions of the EHP in adult mice resulted in an anterograde degeneration of perforant pathway fibers in the hippocampus and DG (e.g. Fagan and Gage, 1994; Jensen et al., 1994). Entorhinal lesions are accompanied by considerable glial reactivity and proliferation shortly after lesion (between 24 and 48 h), which declines in the following days. Thus, after perforant path knife-lesions, glial cells proliferate actively in the outer two-thirds of the dentate ML and in the stratum lacunosum-moleculare of the hippocampus proper, as described (Supplementary Fig. 1).
Studies to analyze the proliferation of neuroprogenitor cells have used cumulative BrdU-pulses after CNS lesions. With this approach the BrdU content of the DNA of neuroprogenitor cells may be underestimated by the partial dilution of the nucleotide analogue in reactive proliferating glia. In addition, the identification of neuroprogenitors may be hindered by abundant glial reactivity. Thus, to label endogenous neuroprogenitor cells and follow their fate, we compared findings between our pre-lesion BrdU-pulse-labeling procedure and conventional post-lesion BrdU-labeling procedures (see Materials and Methods for details) after EHP lesion. The thymidine analogue was injected for three consecutive days before the lesion, which render a high percentage of both BrdU-labeled neuroprogenitors and postmitotic cells at the moment of lesion (Dayer et al., 2003). This labeling enabled us to follow short-time effects on the proliferation rate of neuroprogenitors and to ascertain the survival of postmitotic cells generated during the days of BrdU labeling (e.g. Dayer et al., 2003).
Four and seven days after EHP axotomy, with the BrdU-prelesion method numerous double-labeled NG2–BrdU cells were seen around the lesion site (Fig. 3). This indicates that NG2-positive endogenous oligodendrocyte progenitors proliferate locally near the lesioned area (Fig. 3A,B,D). However, no BrdU–NeuN double-labeled cells were observed in the HF outside the DG after EHP axotomy (not shown). Similar negative results were obtained after FF lesions (see below).
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In the DG, constant numbers of BrdU-positive (pre-labeling method) (Fig. 4A,B,H) or PSA-NCAM-positive (Fig. 4D–F,H) and double-labeled BrdU–PSA-NCAM cells (not shown) were observed in the SGZ and in the GCL 4 and 7 DAL. In addition, cell counts after unilateral EHP lesion showed that the number of BrdU-positive cells (pre-lesion labeling method) in the SGZ and in the GCL were similar in the DG ipsilateral and contralateral to the lesion as also observed in sham animals (I/C
1; Fig. 4A,B,G). In contrast, at 4 DAL the I/C index increased (4.5-fold increase; Fig. 4G) when the BrdU post-labeling method was used, but it decreased at 7 DAL to the levels observed in sham animals, mimicking the results obtained at 4 and 7 DAL using the BrdU pre-labeling method. Double-immunocytochemistry against BrdU and glial markers revealed that most of these BrdU-positive cells observed at 4 DAL in the hippocampus and DG were reactive astroglia and microglia (see Supplementary Fig. 1). Moreover, BrdU-labeled pyknotic nuclei were rare in the SGZ and the GCL with both types of labeling both 4 and 7 days after EHP lesion. Taken together, these results indicate that EHP axotomy does not induce adult neurogenesis in the EC and hippocampus, but does cause NG2-positive cells near the lesion to proliferate. In addition, EHP axotomy does not increase the number of BrdU-positive cells (pre-labeling method) in the SGZ of the DG, which implies that neuroprogenitors do not increase their proliferation rate and that survival of postmitotic neurons is not compromised at least 1 week after lesion.
Cell Proliferation in the Adult DG of the Mouse after Unilateral FF Axotomy
It is believed that fornix–fimbrial fibers modulate electrical activity in normal hippocampal physiology (e.g. Olton, 1977; Cassel et al., 1997; Kirk, 1998). Thus, we next performed experiments after unilateral lesions of the fornix–fimbria axons that were similar to those after EHP lesion and also used the BrdU pre-labeling method. As after EHP axotomy, unilateral FF transection did not induce neurogenesis in the hippocampus proper (not shown). However, BrdU cell counts revealed a 40% decrease (7.5 ± 1.7 in FF-lesioned animals versus 12.4 ± 1.3 in sham-operated animals) in the number of BrdU-positive cells in the SGZ and GCL in the ipsilateral deafferented DG 7 days after unilateral FF lesion (Fig. 5). This decrease further correlates in our experiments with a 33% reduction in the number of PSA-NCAM-positive cells in the GCL 15 DAL (Fig. 5C). Since no significant BrdU-positive or Nissl-stained pyknotic nuclei numbers were observed at any survival time (3 of 400 BrdU-counted cells), these data suggest that axotomy of the fornix–fimbria afferents decreases the proliferation of the neuroprogenitor cells in the DG of the adult mouse.
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Proliferating Cells in Adult Hippocampal Formation of Rodents
Two reports indicate adult neurogenesis in the healthy cerebral cortex of primates (Gould et al., 1999b) and the hippocampus proper (Rietze et al., 2000). In addition, a recent study describes adult neurogenesis in the neocortex and in the striatum (Dayer et al., 2005). Although we did not examine brain regions other than HF, our BrdU labeling study indicates that adult cortical neurogenesis in the HF only occurs in the SGZ of the DG. Pioneer autoradiographic studies in rodents by Smart (1961) and in primates by Lewis (1968) described the presence of mitotically active subependymal cells lining the lateral ventricles. Parallel to these studies, proliferating glial cells were identified in almost all forebrain regions in adult animals (for pioneer studies, see e.g. Walker and Leblond, 1958; Messier et al., 1958; Smart and Leblond, 1961; Dalton et al., 1968; for further reviews, see also Korr, 1980; Kaplan and Hinds, 1980; Gensert and Goldman, 1996; for more recent descriptions, see Palmer et al., 1995, 1999). As commented above, we observed numerous double-labeled NG2–BrdU cells in our short- and long-term BrdU-labeling experiments in the HF, but observed no double-labeled NeuN–BrdU or rCRMP–4/BrdU cells in the EC and the hippocampus proper. This observation seems to contradict the finding that BrdU-positive cells of the adult EC parenchyma (most of then NG2-positive) differentiate into neurons in vivo, as indicated in the hilus of the DG or in the neocortex (Belachew et al., 2003; Dayer et al., 2005), and suggests that a subset of NG2-positive cells, if not all, is a proliferating pool of glial precursors which may further differentiate into oligodendroglia and astroglia in the EC (for a review, see Marshall et al., 2003). Postnatally, neuronal and glial progenitors migrate from the proliferative SVZ within a coronal plane into the striatum and the dorsal, medial and lateral regions of the cerebral cortex and very few of them enter the ventral archicortex (Marshall et al., 2003; Suzuki and Goldman, 2003). These migratory patterns could explain the differences between Dayer's findings in the neocortex and our study, focused in the HF. However, several populations of NG2-positive cells with different physiological properties have been described in the postnatal cerebral cortex (Mallon et al., 2002; Chittajallu et al., 2004). All these findings pose numerous challenging questions on the physiological role of these subsets of NG2-positive cells in the cerebral cortex and their neurogenic potential in different cortical regions.
Isolated Neural Progenitors of the EC Can Form Neurospheres
Our study shows that neural progenitors can be isolated from the EC as floating neurospheres as well as from the neocortex (Palmer et al., 1999), and that they display similar characteristics (i.e. neuronal lineage potential) to other well-characterized neural progenitors (e.g. hippocampal progenitors; see Palmer et al., 1997). It is known that expansion of neural progenitors in floating neurosphere culture represses cell-type differentiation by interfering with the normal cascade of cell specification (Hack et al., 2004). In our experiments, EC-derived neural progenitors growing as floating aggregates differentiate inside the neurosphere shortly after passage, as do hippocampal neurospheres (Vicario-Abejon et al., 2000). This finding supports the view that growth factors in culture media play a dual effect on both cell proliferation and differentiation, alongside other effects such as cell density (Vicario-Abejon et al., 2000). In this regard, FGF2 is involved in cell proliferation and differentiation of hippocampal neuroprogenitor cells in healthy and lesioned brains (Yoshimura et al., 2001; Cheng et al., 2002; Nakatomi et al., 2002) and also regulate gene expression changes in neurospheres (Hack et al., 2004) or dissociated cultures (Kessaris et al., 2004).
Several studies have reported that native precursor cells in the neocortex, striatum and hypothalamus generate neurons and glia in culture (Palmer et al., 1995, 1999; Markakis et al., 2004). These studies appear contrast with those of Seaberg and co-workers using neurospheres (Seaberg and van der Kooy, 2002, 2003; Seaberg et al., 2005). Although we did not analyze neurospheres from the isocortex in our study, the data from van der Kooy (Seaberg and van der Kooy, 2002) appear to contrast with our observations in the EC. Although technical considerations (neurosphere versus dissociated cultures, cell densities or different dissociation procedures) cannot be ruled out, our data, which show that neurospheres cultured for long periods can differentiate into neurons, astrocytes and NG2-positive cells, support the notion that the EC contains cells with neurogenic potential in vitro, as also reported in brain regions other than the SVZ or the hippocampus (Kondo and Raff, 2000; Gabay et al., 2003; Dayer et al., 2005).
The white matter of the corpus callosum and the antero-posterior residuum of the SVZ contains cells with neurogenic potential (e.g. Palmer et al., 1999; Marshall et al., 2003; Dayer et al., 2005). In our experiments, we found more BrdU-positive cells in the dorsal hippocampal white matter than in ventral regions. In addition, we obtained, though at lower efficiency, neurospheres from upper layers of the EC. Pioneer autoradiographic studies determined that 0.2% of glial cells in the adult brain proliferate in the adult cortex (for a discussion, see Korr, 1980) and that glial cells in cortical parenchyma deriving from postnatal SVZ proliferate throughout the lifetime of the mouse, though proliferation decreases with age (see also Marshall et al., 2003; present results). These results raise the question of whether the cells that we isolate as neurospheres derived from neural progenitors different than remnant proliferating glial cells (e.g. NG2-positive) in the cortical parenchyma. Despite the need for further study, we observed that the first sign of NG2-immunoreactivity in the growing neurosphere appeared rapidly in culture, indicating that NG2-positive cells are present during the early stages of neurosphere formation. This seems to corroborate recent data from Belachew et al. (2003), who showed that young postnatal FACS-isolated NG2-positive/CNP-GFP cells can form neurospheres. On the basis of our data, we cannot rule out either a participation of NG2-expressing cells in the formation of the neurosphere or an early differentiation of some undifferentiated EC neural progenitors to NG2-positive cells, because FGF2 may also induce oligodendrocyte differentiation in vitro (Kessaris et al., 2004).
We also showed that EC-derived neurospheres give rise to neuronal and glial phenotypes after in vivo transplantation in neurogenic areas (DG), and to glial phenotypes in non-neurogenic regions. This limited potential is also displayed by other neural progenitors (e.g. Herrera et al., 1999; Shetty, 2004). These data indicate that the in vivo differentiation pattern of neurosphere-derived progenitors does not reflect the complete lineage potential of the resident progenitor cells. In addition, these results reinforce the notion that the absence of local environmental factors regulating cell proliferation and neuronal differentiation in ‘neuronal niches’ of the CNS (e.g. noggin/BMP4 signalling: Lim et al., 2000; neurogenesin-1: Ueki et al., 2003; synaptic activity: present results) appears to limit the in situ acquisition of a neural lineage in non-neurogenic areas (see below). In this respect, blocking of noggin function with antisense oligonucleotides decreases cell proliferation in the adult DG, which may reinforce this hypothesis (Fan et al., 2004)
Control of Neuroprogenitor Proliferation in the Rodent SGZ by Hippocampal Afferents
Several studies have identified endocrine, neural and experimental factors that regulate the generation and survival of new neurons in the DG. For example, estrogens (Banasr et al., 2001) and enriched environment (Kempermann et al., 1997) are considered positive regulators of adult-generated neurons. In contrast, glutamatergic excitatory input (Cameron et al., 1995), adrenal steroids (e.g. Cameron et al., 1998) and stress (Gould and Tanapat, 1999) are described as negative regulators. In addition, several studies have shown that various insults stimulate the generation of neurons in neurogenic (Magavi et al., 2000) and non-neurogenic areas (Yamamoto et al., 2001; Jiang et al., 2001; Nakatomi et al., 2002). Our results indicate that axotomy of the entorhinal and fornix–fimbrial afferents does not induce neurogenesis in the HF outside the DG. In addition, the entorhino-hippocampal lesion does not increase the proliferation of neuroprogenitor cells in the DG 1 week after lesion, which contrasts with the findings of other authors (Cameron et al., 1995) but corroborates other recent results (Gama Sosa et al., 2004). Although differences between animal species cannot be ruled out, the different conclusions of our study compared with those of Cameron and co-workers may be due to distinct methodological approaches. Our axotomy procedure is highly reproducible and induces less dramatic effects on the neuronal parenchyma than other lesion procedures do (e.g. death of entorhinal projection neurons; see Jensen et al., 2000; Poulsen et al., 2000; Mingorance et al., 2004). Furthermore, we used combined pre-lesion and post-lesion BrdU-labeling methods that enabled us to discriminate between neuroprogenitor cells and reactive glia, since our pre-lesion labeling procedure well suited to avoiding nucleotide dilution in reactive glia, which could lead to underestimation of the number of proliferating cells (for technical details, see Gould and Gross, 2002). However, this procedure has certain limitations, such as the long survival time analysis of the newly generated cells. Finally, we corroborated BrdU-labeling data by counting PSA-NCAM-positive cells, a marker of neuroprogenitor and newly generated postmitotic granule cells (Seki and Arai, 1993, 1999), which paralleled increased BrdU proliferation in the DG in control mice and after lesion (e.g. after amygdaloid kindling, Saegusa et al., 2004; or NMDA receptor antagonist treatment, Nacher et al., 2001).
Our present data corroborate recent findings that the unilateral axotomy of the FF leads to a rapid decrease in the number of BrdU-positive cells in the DG (Lai et al., 2003). These authors argue that the anterograde transport of Shh from basal forebrain is important in regulating the mitotic activity of neural progenitor cells in the DG, as demonstrated for other neural precursor cells in CNS populations (for recent reports, see Rowitch et al., 1999; Palma et al., 2005). Functions of Shh corroborate the abnormalities in the cytoarchitecture of the DG and the decrease in the number of neural progenitors in the subventricular zone and the hippocampus in conditionally null mice lacking Shh signalling (Machold et al., 2003). Although Shh is relevant to the modulation of neuroprogenitor proliferation in the DG, this phenomenon may be additionally affected by other factors acting over a long distance and transported to the hippocampus via FF, such as the neurotransmitters serotonin (Brezun and Daszuta, 2000a,b) and acetylcholine (Cooper-Kuhn et al., 2004; Mohapel et al., 2005).
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TopAbstractIntroductionMaterial and MethodsResultsDiscussionSupplementary MaterialReferences |
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Supplementary materials can be found at http://www.cercor.oxfordjournals.org.
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Acknowledgments |
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The authors are grateful to Dr Alberto Martínez Serrano (CBM-UAM, Madrid, Spain) for providing the HNSC line, to John Mason (Edinburgh, UK) for the Tau-GFP mice and to Dr Genevieve Rougon for the anti-PSA-NCAM antibody. We also thank Susana Maqueda and Maica López for their technical assistance and Robin Rycroft for editorial help. This study was supported by grants from the Spanish Ministry of Science and Technology (MCYT, EET2002-05149 and BFI2003-03459), from ‘La Caixa Foundation’ and the Fundación Mútua Madrileña Automobilística to J.A.D.R., the MCYT (BFI2003-01254) to J.N., and from the MCYT (SAF2004-07929 and ‘Pfizer Foundation’) to E.S. X.F. is a fellow of the Spanish Ministry of Education and Science.
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