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1 Department of Neurosurgery and Cellular and Molecular Physiology and, Yale University School of Medicine, New Haven, Connecticut 06520-8082; 2 Interdepartmental Neuroscience Graduate Program, Yale University School of Medicine, New Haven, Connecticut 06520-8082
Submitted 12 December 2002; accepted in final form 10 June 2003
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONDISCLOSURESREFERENCES |
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-tubulin, and recorded cells displayed a morphology typical of the neuronal progenitors known to migrate throughout the SVZ and RMS to the olfactory bulb. Recorded progenitors had depolarized zero-current resting potentials (mean more depolarized than –28 mV), very high input resistances (about 4 G
), and lacked action potentials. Using the reversal potential of K+ currents through a cell-attached patch a mean resting potential of –59 mV was estimated. Recorded progenitors displayed Ca2+-dependent K+ currents and TEA-sensitive-delayed rectifying K+ (KDR) currents, but lacked inward K+ currents and transient outward K+ currents. KDR currents displayed classical kinetics and were also sensitive to 4-aminopyridine and 
-dendrotoxin, a blocker of Kv1 channels. Na+ currents were found in about 60% of the SVZ neuronal progenitors. No developmental changes were observed in the passive membrane properties and current profile of neuronal progenitors. Together these data suggest that SVZ neuronal progenitors display passive membrane properties and an ionic signature distinct from that of cultured SVZ neuronal progenitors and mature neurons. |
INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONDISCLOSURESREFERENCES |
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, 1999; Garcia-Verdugo et al. 1998; Levison and Goldman 1997; Luskin 1998). Of particular interest in this region are the neuronal progenitors that migrate on a rostro-caudal axis to the olfactory bulb by the rostral migratory stream (RMS) (Doetsch et al. 1997; Luskin 1993, 1998). Understanding the biology of the SVZ neuronal progenitors might provide important insights into the development of the CNS and the factors regulating cell proliferation, migration, and differentiation, which constitute the process of neurogenesis. In addition these cells hold promise for future therapeutic intervention in neurodegenerative diseases using transplants and/or enhancement of endogenous neurogenesis. The different steps of neurogenesis can be influenced by a large variety of stimuli (Cameron et al. 1998; Gage et al. 1998; Heller et al. 1996; Morrison 2001; Nakashima and Taga 2002). Although many laboratories have started to reveal the extracellular factors regulating neurogenesis, intrinsic cellular properties such as ion channels, which are known to functionally regulate the different steps of neurogenesis in several cell types, have received very little attention.
K+ channels in particular are known to be involved in the regulation of two critical steps of neurogenesis: cell proliferation and migration (Dubois and Rouzaire-Dubois 1993; Puro et al. 1989; Schlichter et al. 1996; Schwab 2001; Schwab et al. 1999). Moreover K+ channel properties are often influenced by factors known to regulate the distinct steps of neurogenesis, such as cytokines and growth factors (Bowlby et al. 1997; Gamper et al. 2002; Perillan et al. 2002; Timpe and Fantl 1994). Thus a detailed characterization of these channels in neuronal progenitors promises to contribute to an understanding of the mechanisms of action of the various factors stimulating neurogenesis in the SVZ and RMS. Moreover this knowledge could provide additional approaches for selectively regulating cell proliferation and/or migration in these regions. A recent report characterized the K+ channels in neuronal progenitors in primary cultures isolated from the SVZ (Stewart et al. 1999). However, cultured cells may take on a more differentiated phenotype and thus become unrepresentative of cells in their native environment. Therefore it is critical to study the intrinsic properties of progenitors in their near-intact physiological environment.
We thus conducted a comprehensive biophysical study of neuronal progenitors located in the SVZ and RMS. Neuronal progenitors exhibited high-input resistances and depolarized zero-current resting potentials similar to those of cultured SVZ neuronal progenitors (Stewart et al. 1999). However, using the single K+ channel reversal potential the resting potential was estimated at –59 mV. SVZ progenitors were found to express Ca2+-dependent K+ channels and delayed rectifying K+ (KDR) channels but lack transient A-type K+ channels that were reported in cultured SVZ progenitors (Stewart et al. 1999).
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METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONDISCLOSURESREFERENCES |
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Slices containing the SVZ or RMS were acutely prepared as previously described for hippocampal and cortical slices (Bordey and Sontheimer 1997; Bordey et al. 2001). Briefly, 15- to 25-day-old CD-1 mice were anesthetized using pentobarbital (50 mg/kg) and decapitated. The brain was quickly removed and chilled (0–4°C) in 95% O2-5% CO2 saturated artificial cerebrospinal fluid (ACSF) containing (in mM): NaCl 125; KCl 2.5; CaCl2 2; MgCl2 2; NaHCO3 25; glucose 10. The tissue of interest was glued to the stage of a Vibratome and 250-µm-thick slices were cut in cold oxygenated ACSF. After a recovery period of 1 h in ACSF, slices were placed in a flow-through chamber, held in position by a nylon mesh glued to a U-shaped platinum wire, and continuously superfused with oxygenated ACSF at room temperature. The chamber was mounted on the stage of an upright microscope (Olympus BX50WI) equipped with a 60x water immersion objective and infrared optics.
Whole cell recordings and drug application
Whole cell patch-clamp recordings were obtained as previously described (Edwards and Konnerth 1992). Patch pipettes were pulled from thin-walled borosilicate glass (OD, 1.55 mm; ID, 1.2 mm; WPI, TW150F-40) on a PP-83 puller (Narishige, Japan). Pipettes had resistances of 6–8 M when filled with the following solutions (in mM): KCl 140; CaCl2 1.0; MgCl2 2.0; ethylene glycol-bis(-aminoethyl ether)-N,N,N',N'-tetraacetic acid (EGTA) 10; Hepes 10; pH adjusted to 7.2 with Tris-base. When noted in the text, EGTA and Ca2+ were removed from the intracellular solution. The osmolarities of the intracellular and extracellular solutions were 295–300 and 305–310 mOsm, respectively. The osmolarity of all solutions was measured with a vapor pressure osmometer 5500 (Wescor) and was adjusted by addition of D-mannitol. For morphological identification, 0.1% Lucifer yellow (LY, dilithium salt) was added to the pipette solution. When a Ca2+-free extracellular solution was used, Ca2+ was replaced with Mg2+ and 1 mM of the Ca2+ chelator EGTA was added. In addition, the control solution contained only 1 mM Ca2+ instead of 2 mM to avoid any complication attributed to surface charge effect. Whole cell and cell-attached recordings were performed using an Axopatch-200B amplifier (Axon Instruments). Recordings were obtained 
7 h after brain removal. Current signals were low-pass filtered at 2–5 kHz and digitized on-line at 5–20 kHz using a Digidata 1320 digitizing board (Axon Instruments) interfaced with an IBM-compatible computer system. Data acquisition, storage, and analysis were performed using PClamp version 8.0.2 (Axon Instruments). Whole cell parameters (capacitance CM and series resistance) were determined by compensating the transients of a small (5 mV) 10-ms hyperpolarizing voltage step. Voltage-clamp data in this study were limited to cells with an input resistance RM >200 M (determined at –80 mV). The capacitance reading of the amplifier was used as the value for the whole cell capacitance. These values and the zero-current resting membrane potential VR were determined in the first 3 min of whole cell recording. Capacitive and leak conductances were not subtracted unless otherwise stated. Peak currents were determined using Clampfit (Axon Instruments), and statistical values were evaluated with a statistical graphing and curve-fitting program (Origin, MicroCal). Statistical comparison of means was performed with the Student's t-test or ANOVA test (Statview, SAS Institute). Data are represented as means ± SD in the text and as means ± SE on the graphs. Ion channel blockers were diluted in ACSF and applied by a rapid bath application system. Blockers were diluted in ACSF in which Hepes replaced NaHCO3 and the pH was adjusted to 7.4 by NaOH or Tris base. No correction of junction potentials was performed.
Using the reversal potential of K+ currents through cell-attached patches to monitor the cell resting potential
The method to measure VR from cell-attached K+ currents is identical to that described by Verheugen et al. (1995, 1999). For these recordings, 150 mM K+ was used in the pipette solution. With a 150 mM K+ pipette solution, which is close to the estimated intracellular [K+] in other cell types (155 mM; Hille 1992), the equilibrium potential for K+ (EK) across the patch is approximately 0 mV, and K+ currents will reverse when the pipette potential (Vpip) cancels VR out. Therefore the cell-holding potential (–Vpip) at which the K+ current reverses direction gives a direct quantitative measure for Vm (at K+ channel reversal, Vpatch = VR – Vpip = EK about 0 mV). Depolarizing voltage ramps (Fig. 3) were applied to activate voltage-gated K+ channels. For analysis of currents evoked by ramp stimulation, a correction was made for a leak component by linear fit and extrapolation of the closed level.
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Steady-state parameters for activation and inactivation
To establish steady-state activation or inactivation curves, the peak current I was measured at each potential, and the corresponding conductance G was calculated using the following equation: G = I/(V – Vi), where V is the membrane command potential and Vi is the equilibrium (Nernst) potential for the ion under consideration (for potassium, VK = –101 mV). The measured peak amplitudes and the calculated peak conductances were then normalized with respect to the maximum values and plotted as a function of the membrane potential during the test pulse. The resulting activation and inactivation curves were fitted to the Boltzmann equation: Gi/Gi(max) = 1/(1 + exp[(V1/2 – V)k]), where Gi(max) is the maximum ionic conductance at peak current; V1/2 is the voltage where Gi is one half of Gi(max), and k is the slope factor that determines the voltage-dependent relation of Gi.
Cell identification
Images of cells visually chosen for recordings were archived using LG3 frame grabber (Scion Corporation, Frederick, MD) for later (off-line) comparison to LY fills. After recordings, slices were fixed in 4% paraformaldehyde in phosphate-buffered saline (PBS) overnight. Slices were washed three times in 2x Tris-buffered saline (TBS) over 1 h, permeabilized for 15 min with 1.25% Triton X-100 and 10% bovine serum albumin (BSA, Sigma) in 2x TBS, and then blocked for 1 h in blocking solution containing 0.2% Triton and 10% BSA in TBS. Slices were subsequently incubated with primary antibodies specific for nestin (Rat-401, supernatant, Developmental Studies Hybridoma Bank developed under the auspices of the NICHD and maintained by the University of Iowa) or neuron-specific class III -tubulin (TuJ1, monoclonal anti--tubulin Cy3 conjugate; Sigma, St. Louis, MO). For TuJ1 immunostaining, slices were incubated with Cy3-conjugated antibody against TuJ1 (1:100) for 2 h at room temperature in blocking solution. For nestin immunostaining, slices were incubated overnight at 4°C with primary antibody in blocking solution. Slices were subsequently washed three times with TBS for 5 min and incubated with secondary antibody (goat anti-mouse IgG) conjugated to Texas red (dilution 1:200; Vector) for 2 h at room temperature. For both stainings, slices were then washed in TBS and mounted on glass coverslips with fluorescent mounting medium (Vectashield; Vector) and were finally viewed on an epifluorescence microscope (Olympus microscope BX51) using standard procedures. Images were captured with a DVC color camera and printed on an Epson color printer.
Chemicals were purchased from Sigma except the recombinant peptide toxins, agitoxin-2 and -dendrotoxin, which were purchased from Alomone Labs (Jerusalem, Israel). Toxins were stored lyophilized at –20°C and made up in extracellular solution containing 0.1% BSA.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONDISCLOSURESREFERENCES |
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The location and morphology of recorded cells identify them as neuronal progenitors
LOCATION. Coronal slices containing either the RMS or the SVZ were located between sections 9 and 16 (both approximately) based on the brain map described by Swanson (2002), as shown in a sagittal section of the brain with the section number on a rostro-caudal axis (Fig. 1A, left panel). Images of slices shown in Fig. 1 were taken before or during the recordings. At section 11 (approximately) on the brain map, the RMS took on an elongated shape, distinguishable from the surrounding tissue by its darker appearance, as indicated by the location of the two patch pipettes in Fig. 1B. In more rostral slices, the RMS had a round appearance (data not shown). In section >12, the lateral ventricle could be observed together with the SVZ [Fig. 1C, section 15 (approximately)]. At this level, the SVZ is located along the lateral wall of the ventricle and striatum and extends between the striatum and the corpus callosum (see diagram in Fig. 1A, right panel).
CELLS RECORDED IN THE SVZ AND RMS DISPLAY SIMILAR MORPHOLOGIES, TYPICAL OF IMMATURE CELLS. Cells in the RMS (Fig. 1D) and the SVZ (Fig. 1E) were densely packed, with a morphological appearance distinct from that of mature cells. They were characterized by a small round cell body with a main diameter of 6–8 µm, as well as occasional short processes that could be observed particularly when the patch pipette was penetrating inside the tissue. In the RMS they formed a channel of cells that often appeared to protrude from the surrounding mature tissue (Fig. 1D). In contrast, the SVZ consisted of a 30- to 50-µm-wide layer of cells, separated from the ventricle by ependymal cells displaying characteristic cilia. This layer of cells narrowed in more caudal sections. To permit further visualization of the cell morphology, cells were routinely filled with Lucifer yellow (LY) during the recording, and a photograph of each recorded cell was taken and stored. Most of the recorded cells in the SVZ and RMS had a similar morphology characterized by a round to fusiform cell body with one or occasionally two processes (Fig. 4A, top panel). These recorded cells closely resemble neuronal progenitors described in vivo (Doetsch et al. 1999) and in vitro (Stewart et al. 1999; Wichterle et al. 1997). A few recorded cells had a larger cell body with one long process and were found only in the SVZ; other recorded cells were found both in the SVZ or RMS but had a more elaborated morphology with 2 or 3 long processes. Data from these morphologically distinct cells were not included in the present study.
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IMMUNOSTAININGS FOR NESTIN AND TUJ1. To verify that the described regions contained immature cells, immunostaining was performed for nestin, a class VI intermediate filament protein that distinguishes progenitors from more differentiated cells (Hockfield and McKay 1985; Lendahl et al. 1990). In our slices, nestin staining was selectively observed in the SVZ and RMS [Fig. 2A, left panel, sections 12–13 (approximately)]. At high magnification, nestin staining in the dorsolateral SVZ had a honeycomb appearance, probably as a result of the round cell somata and the lack of staining in the nucleus (Fig. 2A, right panel). Virtually every cell in the SVZ was nestin-positive, identifying them as progenitors. In addition, to verify that the described regions contained neuronal progenitors, immunostaining was performed for TuJ1, a marker of neuronal progenitors. Slices containing the RMS and the rostral (or anterior) SVZ displayed strong immunostaining for TuJ1 (Fig. 2B), identifying a large proportion of the cells in these regions as neuronal progenitors, as previously reported (Brock et al. 1998; Doetsch et al. 1997; Luskin 1998). As expected, the corpus callosum, the striatum, and the olfactory limb of the anterior commissure stained positive for TuJ1.
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Thus the location, morphology, and immunostaining data strongly suggest that the cells recorded in the SVZ and RMS correspond to the neuronal progenitors known to migrate to the olfactory bulb in postnatal animals (Gritti et al. 2002; Lois et al. 1996; Luskin 1993, 1998). In addition, because glial progenitors have not been reported in the RMS (Doetsch et al. 1997), recorded cells are unlikely to be glial progenitors. In the following sections, we set out to determine and compare the electrophysiological properties of RMS and SVZ neuronal progenitors.
Neuronal progenitors in the SVZ and RMS display similar passive membrane properties and current profile
PASSIVE MEMBRANE PROPERTIES. The zero-current VR, CM, and RM were determined for 49 and 112 neuronal progenitors recorded in the RMS and the SVZ, respectively (Table 1). Both SVZ and RMS progenitors exhibited similar and very high input resistances (mean of about 4,000 M) and depolarized zero-current VR (Table 1). The mean zero-current VR values were more depolarized than –28 mV. Both SVZ and RMS progenitors also displayed similar and small CM (mean <8 pF, Table 1), indicating their small cell size. Because developmental changes in the architecture of the SVZ occur during the third postnatal week in rats (Peretto et al. 1999), we questioned whether the passive membrane properties of the neuronal progenitors may also show developmental changes between postnatal day 15 and day 25. To address this question, we plotted CM, zero-current VR, and RM as functions of the animal age (data not shown). There was no correlation between these parameters and the age of the animals as measured by plotting these respective parameters as a function of the postnatal day (linear coefficient of correlation r = –0.06 and 0.02 for CM, –0.07 and 0.07 for zero-current VR, and 0.07 and 0.03 for RM in the RMS and SVZ, respectively). In a few cells, we estimated the membrane time constant from a monoexponential fit to the voltage response after a –100-pA current injection at –70 mV. The membrane time constant of neuronal progenitors was 46.1 ± 8.6 ms (n = 7), which is between that of dentate granule neurons and CA3 pyramidal neurons [40 and 66 ms at their resting potentials (Spruston and Johnston 1992)].
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It is important to point out that the zero-current VR of neuronal progenitors likely does not represent the true resting potential of these cells because neuronal progenitors have a very high input resistance (about 4 G) and the seal resistances were between 3 and 10 G. In these circumstances, the seal conductance value is close to that of the membrane conductance (estimated as the input conductance minus the seal conductance), which results in an artifactual depolarization of recorded cells (Pongracz et al. 1991). In fact, there was a weak correlation of the input resistance plotted as a function of the zero-current VR (r = 0.4, n = 121, Fig. 3). To estimate the VR of neuronal progenitors with another method more appropriate in this circumstance, we performed cell-attached recordings and used the reversal potential of K+ channels as a monitor of VR as previously described for lymphocytes (Verheugen et al. 1995) and hippocampal interneurons (Verheugen et al. 1999) (Fig. 3B, schematic of the recording conditions). To activate K+ channels, voltage ramps from –140 to +200 mV (–Vpip, Fig. 3, C and D) were repeated at 0.5 Hz without any significant inactivation of K+ channels. Sufficiently large outward currents were activated in about 20% of the patches. Only patches displaying inward K+ currents in addition to outward K+ currents were analyzed (Fig. 3C). In addition, in 2 patches single-channel currents likely the result of Ca2+-dependent K+ channel activity were observed (Fig. 3D). At the K+ channel reversal, the cell membrane potential, which represents the cell resting potential at rest was –59.1 ± 2.4 mV (n = 7).
CURRENT PROFILE. Whole cell currents were activated in voltage-clamped progenitors by applying hyperpolarizing and depolarizing voltage steps from –100 mV to +100 mV from a holding potential of –80 mV. In both RMS and SVZ cells, nonleak subtracted traces were similar and characterized by voltage-dependent outward currents, resulting in a strictly outwardly rectifying current–voltage (I–V) curve (Fig. 4, A and B for a SVZ progenitor). The conductance density (conductance divided by CM) of outward currents was similar in SVZ and RMS progenitors (mean of 270 and 340 pS/pF, respectively, Table 1). In addition, the density of outward current amplitude did not change with the age of the animal (r = –0.07 and –0.01 for RMS and SVZ cells, respectively). A tail current analysis of the outward currents gave a reversal potential of about –80 mV, close to EK in our recording conditions (Fig. 4, C and D), and the sensitivity of these outward currents to TEA identified them as voltage-dependent K+ currents (see next section and Figs. 5 and 6). The K+ currents recorded in the RMS and SVZ were similar in their extent of voltage-dependent inactivation. To quantify this observation, the ratio of the peak current amplitude to the current at the end of a 300-ms pulse to +20 mV was calculated. This ratio was similar in both groups of neuronal progenitors (ratio of 1.3 in RMS cells and 1.2 in SVZ cells, Table 1). A substantial cumulative inactivation of the K+ currents for voltage steps applied every 4 s was also observed in both RMS and SVZ neuronal progenitors (data not shown).
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In summary, neuronal progenitors in the SVZ and RMS display similar morphology, passive membrane properties, and current profile, confirming that they belong to the same progenitor type (i.e., neuronal progenitors).
Biophysical and pharmacological properties of K+ currents in neuronal progenitors
To activate outward currents, cells were held at –80 mV and depolarizing voltage steps were applied after a prepulse to –50 mV (Figs. 5 and 6) or to –110 mV (Fig. 7) to allow the activation of transient A-type K+ currents in addition to delayed-rectifying K+ (KDR) currents. In our recordings, neuronal progenitors displayed outward currents that showed delayed activation and slow inactivation (Figs. 5 and 6). Virtually no transient A-type K+ currents were observed in SVZ and RMS neuronal progenitors.
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PRESENCE OF TEA AND CHARYBDOTOXIN-SENSITIVE K+ CURRENTS. Stewart et al. (1999) showed that a step to +0 mV resulted in opening of Ca2+-dependent K+ channels (KCa) as a result of Ca2+ entry through voltage-dependent Ca2+ channels. Thus we tested the effects of a Ca2+-free extracellular solution (see METHODS), 100 µM TEA, and 100 nM charybdotoxin (ChTx) on the outward K+ currents. TEA (100 µM) has been shown to block KCa channels without affecting KDR channels (Iwatsuki and Petersen 1985). ChTx is known to block KCa channels. Using a Ca2+-free extracellular solution or bath applying TEA or ChTx did not significantly reduce the outward current amplitude measured at +20 mV, suggesting that in our recording conditions KCa currents did not significantly contribute to the current amplitude measured at +20 mV. However, when cells were recorded with an intracellular solution containing no EGTA/Ca2+, a Ca2+-free extracellular solution and 100 µM TEA reversibly blocked outward K+ currents by 11.6 ± 2.7% (n = 3, Fig. 5) and 14.7 ± 2.3% (n = 3, data not shown), respectively. ChTx irreversibly reduced outward currents when cells were recorded without internal EGTA. The Ca2+-sensitive K+ currents were voltage dependent and activated at about –10 mV (Fig. 5A). These data suggest the presence of Ca2+ and voltage-dependent K+ channels in neuronal progenitors, as previously shown by Stewart et al. (1999). All the following experiments were performed with an intracellular solution containing EGTA to limit KCa channel activation.
TEA-SENSITIVE DELAYED RECTIFYING K+ CURRENTS CONTRIBUTE TO OUTWARD K+ CURRENTS. In the presence of EGTA in the intracellular solution to limit KCa channel activation, a bath application of 40 mM TEA completely blocked voltage-dependent outward currents in all neuronal progenitors tested (n = 7, Fig. 6A). TEA-sensitive currents and their respective I–V curves are displayed in Fig. 6B. TEA-sensitive currents activate at –40 mV and display voltage-dependent inactivation as observed on their mean I–V curves (Fig. 6C). The mean steady-state activation curves (see METHODS) of TEA-sensitive currents gave a mean half activation voltage V1/2 of –4.5 mV and a mean slope k of 8.6 mV–1 (Table 2, Fig. 6D). In neuronal progenitors, the mean TEA-sensitivity curve could be fitted with a sigmoidal curve that gave an IC50 value of 3.2 mM (Fig. 6, E and F). This IC50 is close to that reported for KDR currents in cultured neuronal progenitors (4.1 mM) (Stewart et al. 1999).
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OUTWARD K+ CURRENTS ARE SENSITIVE TO 4-AMINOPYRIDINE (4-AP) AND A KV1 CHANNEL BLOCKER, A-DENDROTOXIN. 4-AP (2 mM) completely blocked KDR currents in neuronal progenitors (Fig. 7A), and 4-AP–sensitive currents (Fig. 7B) displayed kinetic characteristics similar to those of TEA-sensitive currents. They activated at –40 mM, displayed voltage-dependent inactivation (Fig. 7C), and their steady-state activation curve parameters were similar to those obtained for TEA-sensitive currents (Table 2). Voltage-gated K+ channels are assembled from subunits of 4 major subfamilies, designated as Kv1, Kv2, Kv3, and Kv4 (Coetzee et al. 1999; Pongs 1992). Because of the similarity between the properties (kinetics, and TEA and 4-AP sensitivity) of KDR currents found in neuronal progenitors and those of Kv1 currents (Coetzee et al. 1999), we tested two selective blockers of Kv1 channels. -Dendrotoxin (80 and 200 nM), which potently blocks Kv1.1, Kv1.2, and Kv1.6 (Coetzee et al. 1999), reduced outward K+ currents in 6/8 neuronal progenitors tested. -Dendrotoxin (80 nM) reduced outward currents by 62 ± 21% (data not shown). Agitoxin-2 (10 nM), an extremely potent Kv1.3 blocker and less potent Kv.1.1 blocker (Garcia et al. 1994, 1997), reduced outward K+ currents in only 2/8 cells tested (70 ± 4%, n = 2, data not shown).
SVZ (60%) and RMS (80%) neuronal progenitors express Na+ channels
Transient inward currents were activated with voltage steps more positive than –40 mV, peaked within <3 ms, and inactivated rapidly (Fig. 8A). Current amplitudes were voltage-dependent, peaked between –10 and 0 mV, and decreased with more depolarized voltage steps (Fig. 8B). The mean I–V curve was generated from 3 cells in normal solution without TEA and from 4 cells in the presence of 40 mM TEA to minimize contributions from K+ currents. Data obtained with and without TEA were combined to obtain a mean I–V curve. Extrapolation of the linear fit of the mean I–V curve between –10 and +40 mV gave a current reversal potential of +60 mV (n = 7). The mean steady-state activation curve of Na+ currents gave a mean V1/2 of –20 mV and a mean slope of –6 mV–1 (Fig. 8D, filled circles; Table 2). The steady-state inactivation properties of Na+ currents were determined by measuring peak current amplitudes at –10 mV after prepulse potentials ranging from –104 to 0 mV (Fig. 8C). Fitting each inactivation curve of Na+ currents (Fig. 8D, open circles) to a Boltzmann equation yielded average values of –61 mV for V1/2 and 7.4 mV–1 for the slope (n = 4). In 6 cells in which it was tested, 100 nM TTX blocked the inward currents, confirming that these currents are Na+ currents (n = 3 cells in the SVZ and 3 cells in the RMS, data not shown). TTX block was partially reversible.
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Na+ currents were observed in 59% of SVZ (54/92) and 79% of RMS (35/44) neuronal progenitors, which was significantly different (P < 0.02, chi-square and Fisher exact test). The mean density of Na+ conductance was about 100 pS/pF in both SVZ and RMS neuronal progenitors (Table 2). Although overall 65% (89/136) of both SVZ and RMS neuronal progenitors express Na+ channels, Na+ conductances were too low to allow these progenitors to generate a fast action potential from –70 mV in response to current injections (Fig. 8F). Figure 8E displays a plot of GK/CM as a function of GNA/CM that shows the range of both K+ and Na+ conductances. In addition, this plot illustrates that GK > GNA. As for the passive membrane properties and the K+ current amplitude, there was no change in the amplitude of the Na+ currents over the age period studied (r = 0.05 for the density of Na+ current amplitude in cells from both regions plotted as a function of the postnatal age). Similarly, there was no significant difference between the numbers of cells expressing Na+ channels between postnatal days 15 and 19 (61% SVZ cells, n = 41/67; 82% RMS cells, n = 14/17) and postnatal days 21 and 25 (56% SVZ cells, n = 13/25; 78% RMS cells, n = 21/27).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONDISCLOSURESREFERENCES |
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Identification of neuronal progenitors in the SVZ
Cells were recorded in both the anterior SVZ and the RMS. These two regions are essentially composed of cells positive for nestin, a marker of progenitors cells (Hockfield and McKay 1985; Lendahl et al. 1990). Indeed LY-filled cells recorded in these regions displayed a morphology typical of immature cells and were thus considered to be progenitors. The SVZ is known to contain distinct progenitor types, including stem cells, bipotential progenitors, and committed progenitors, that can be distinguished morphologically (Doetsch et al. 1997, 1999; Garcia-Verdugo et al. 1998; Luskin 1998). The presumed SVZ stem cells have a fairly complex astrocyte-like morphology, whereas bipotential progenitors have no processes and are relatively rare (Doetsch et al. 1997). Cells recorded in our study resembled neither of these two types, but were characterized by a relatively small soma (6–8 µm in diameter) and one or occasionally two processes, identifying them as committed progenitors, which could be either neuronal or glial progenitors. Staining for class III -tubulin, a marker of neurons and neuronal progenitors, showed that a large proportion of the SVZ cells were neuronal progenitors. In addition, the RMS has been shown to be composed of about 80% neuronal but not glial progenitors and about 20% astrocyte-like stem cells (Doetsch et al. 1997; Luskin 1993), identifying the cells recorded in the RMS as neuronal progenitors. The similar morphology of the cells recorded in the RMS and SVZ suggest that the recorded SVZ cells are the neuronal progenitors known to originate and migrate through the SVZ and reach the olfactory bulb by the RMS (Doetsch et al. 1997; Luskin 1993, 1998).
RMS and SVZ neuronal progenitors display similar passive membrane properties and current profile
PASSIVE PROPERTIES. Neuronal progenitors have depolarized zero-current resting membrane potentials (mean > –28 mV), a very high input resistance (mean >4 G) and a low mean cell capacitance consistent with their small size. The mean zero-current VR is close to that reported in cultured SVZ neuronal progenitors (–32 mV; Stewart et al. 1999). However, because the VR measured as the zero-current potential is affected by the seal conductance in high-resistance cells, we estimated the resting potential with another method. Using the reversal of K+ channels in patches VR was estimated at –59 mV. It is worth mentioning that this method gives us an estimate and not the true resting potential because we do not know the intracellular K+ levels. However, as mentioned by Verheugen et al. (1999), with a intracellular [K+] of 155 mM a difference of 15 mM would result in an error of RT/F x ln (155/140) < 3 mV. In addition, some neuronal progenitors had a relatively hyperpolarized resting potential (< –50 mV, Fig. 3A). Postnatal neuronal progenitors thus have a resting potential similar to that of embryonic neuronal progenitors (–65/–60 mV; Owens et al. 1996).
When recorded cells have a high-input resistance, it is usually thought that they lack electrical coupling. However, postnatal SVZ cells have been shown to express connexin 43 (Miragall et al. 1997). In addition we occasionally observed spontaneous 20-mV–amplitude stepwise changes in the membrane potential similar to those reported in island of Calleja granule cells (Halliwell and Horne 1998). Halliwell and Horne suggested that these stepwise changes observed in granule cells that have a 3-G input resistance are attributed to electrical coupling. Thus to conclude on the lack or presence of electrical coupling would require further investigation.
EXPRESSION OF CA2+-DEPENDENT K+ CHANNELS AND DELAYED RECTIFYING K+ CHANNELS. Voltage-dependent outward K+ currents in neuronal progenitors displayed slow activation and inactivation. These outward K+ currents were composed of charybdotoxin-sensitive Ca2+-dependent K+ currents and TEA-sensitive delayed rectifying K+ currents. KCa and KDR currents activated at about –20 and –40 mV, respectively. KDR currents were totally blocked by 40 mM TEA and were significantly reduced by 2 mM 4-AP. The IC50 for TEA blockade of KDR was 3.2 mV, which is similar to that reported for cultured SVZ neuronal progenitors (4.2 mM; Stewart et al. 1999). Voltage-gated outward K+ channels are assembled from distinct subunits of 4 major subfamilies (Kv1, Kv2, Kv3, and Kv4), which can form heterotetramers, and each different subunit composition can confer unique kinetic and pharmacological properties on the resulting channel (Coetzee et al. 1999). A reduction of KDR currents by -dendrotoxin, a blocker of Kv1.1, Kv1.2, and Kv1.6 (Coetzee et al. 1999), suggests the presence of Kv1 subunits in these cells.
Recorded progenitors lacked fast activating and inactivating A-type outward K+ currents and inward K+ currents. The lack of A-type K+ currents contrasts with a previous study reporting large transient A-type currents in cultured SVZ neuronal progenitors (Stewart et al. 1999). Potential explanations for this difference might involve a change in the subunit composition of K+ channels or a different regulation of the K+ channel properties in the two systems, culture and slice preparation. It is also possible that after 24 h in culture, SVZ neuronal progenitors acquire a more differentiated phenotype. Indeed, several studies reported the development of transient A-type outward K+ currents during neuronal differentiation (Barish 1986; Wakazono et al. 1997).
NEURONAL PROGENITORS EXPRESS NA+ CHANNELS. Interestingly, the percentage of neuronal progenitors expressing Na+ channels is significantly higher in the RMS (about 80%) than that in the SVZ (about 60%), which is consistent with the hypothesis that Na+ channels are expressed as an early event of differentiation as cells begin to migrate in the RMS. Consistent with their immature phenotype and the smaller amplitude of Na+ currents than that of K+ currents (GK/GNa >1), neuronal progenitors were not able to generate an action potential in response to current injections. This was also reported for cultured SVZ neuronal progenitors (Liu et al. 1999; Stewart et al. 1999). Although Na+ channels contribute to the generation of action potentials in mature neurons, their function in neuronal progenitors is unclear. They have been shown to be important in fueling the Na+/K+ pump in astrocytes, which may be one of their functions in neuronal progenitors as well (Sontheimer et al. 1994).
It is interesting to point out that neuronal progenitors display differences in the pharmacological profiles of KDR channels and in the amplitude of K+ currents. In addition only a subset of cells expresses Na+ channels. These differences may reflect the fact that recorded cells are at different stages of differentiation or proliferation. Differences in the pharmacological profile of K+ currents could also be attributable to the presence of distinct neuronal progenitor populations. This is in fact expected because SVZ neuronal progenitors give rise to two types of olfactory bulb interneurons, granule cells and periglomerular cells (Luskin 1998). Our data do not allow us to distinguish between the distinct possibilities explaining differences in neuronal progenitor properties and further studies would be required to address this important issue.
Functional implications of the properties of neuronal progenitors
PASSIVE PROPERTIES. Changes in the cell resting potential have been implicated in the control of protein synthesis, proliferation/migration, and voltage-dependent channel and transport system activity. Regulating the resting membrane potential might provide a direct control of the behavior of the neuronal progenitors, which are known to migrate, proliferate, and progressively differentiate along the RMS (Baker et al. 2001; Coskun and Luskin 2002). Importantly, because of the very high input resistance of the neuronal progenitors, induction of a small current flow can induce a significant change in the cell membrane potentials and could thus strongly influence the behavior of the cells. Such a current flow might result from the opening and closing of K+ channels or the activation of GABAA receptors, which are present in neuronal progenitors in vitro (Stewart et al. 2002). Thus the passive membrane properties of neuronal progenitors support the idea that progenitor behavior is likely to be tightly controlled by small local changes in extracellular signals, such as the neurotransmitter GABA.
ION CHANNEL EXPRESSION. Although ion channels are commonly involved in the generation of action potentials in mature neurons, their function in neuronal progenitors remains unclear. However, TEA-sensitive K+ channels are known to be functionally involved in the regulation of cell proliferation of immature cells and many cell types other than neurons (Attali et al. 1997; Gallo et al. 1996; Kotecha and Schlichter 1999; Pappas and Ritchie 1998; Puro et al. 1989). It is thought that blockade of K+ channels influences intracellular Ca2+ homeostasis, which in turn regulates cell proliferation (Platoshyn et al. 2000; Wohlrab et al. 2002). Thus the potential regulation of KDR currents by growth factors or cytokines might influence the proliferation of neuronal progenitors.
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONDISCLOSURESREFERENCES |
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Address for reprint requests and other correspondence: A. Bordey, Department of Neurosurgery, Yale University, 333 Cedar Street, LSOG 228, New Haven, CT 06520-8082 (E-mail: angelique.bordey{at}yale.edu).
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