1Department of Physiology, University of Utah School of Medicine, Salt Lake City 84108; and 2Department of Neurobiology and Anatomy, University of Utah School of Medicine, Salt Lake City, Utah 84132
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ABSTRACT |
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Piper, David R., Tahmina Mujtaba, Mahendra S. Rao, and Mary T. Lucero. Immunocytochemical and Physiological Characterization of a Population of Cultured Human Neural Precursors. J. Neurophysiol. 84: 534-548, 2000. Human neural precursor cells (HNPC) have recently become commercially available. In an effort to determine the usefulness of these cells for in vitro studies, we have grown cultured HNPCs (cHNPCs) according to the supplier specifications. Here we report our characterization of cHNPCs under nondifferentiating and differentiating growth conditions and make a comparison to primary HNPCs (pHNPCs) obtained at the same developmental time point from a different commercial supplier. We found that under nondifferentiating conditions, cHNPCs expressed nestin, divided rapidly, expressed few markers of differentiated cells, and displayed both 4-aminopyridine (4-AP)-sensitive and delayed-rectifier type K+ currents. No inward currents were observed. On changing to differentiating culture conditions, a majority of the cells expressed neuronal markers, did not divide, expressed inward and outward time- and voltage-dependent currents, and responded to the application of the neurotransmitters acetylcholine and glutamate. The outward current densities were indistinguishable from those in undifferentiated cells. The inward currents included TTX-sensitive and -resistant Na+ currents, sustained Ca2+ currents, and an inwardly rectifying K+ current. Comparison of the properties of differentiated cells from cHNPCs with neurons obtained from primary fetal cultures (pHNPCs) revealed two major differences: the differentiated cHNPCs did not express embryonic neural cell adhesion molecule (E-NCAM) immunoreactivity but did co-express GFAP immunoreactivity. The co-expression of neuronal and glial markers was likely due to the growth of cells in serum containing medium as the pHNPCs that were never exposed to serum did express E-NCAM and did not co-express glial fibrillary acidic protein (GFAP). The relevance of these results is discussed and compared with results from other neuronal progenitor populations and cultured human neuronal cells.
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INTRODUCTION |
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Pluripotent stem cells in the CNS can give rise to
glia and neurons either directly or through intermediate precursors
(Kalyani et al. 1999; Kilpatrick and Bartlett
1993
; Price and Thurlow 1988
; Price et
al. 1992
; Reynolds and Weiss 1996
;
Reynolds et al. 1992
; Temple and Davis
1994
; Williams and Price 1995
) and include the neuroepithelial (NEP) cells identified in the developing spinal cord of
day E10.5 rats and E9.0 mice. These NEP cells divide in vivo and in
vitro and have been shown to give rise to lineally restricted,
intermediate precursor cells in both mass and clonal cultures
(Kalyani et al. 1997
; Mujtaba et al.
1998
). Two classes of restricted precursors have been shown to
differentiate directly from multipotent stem cells: a neuron restricted
precursor (NRP) and a glial restricted precursor (GRP or oligosphere).
These intermediate precursor cells can be isolated from E13.5 rat and
E12.0 mouse in vivo or derived from NEP cells in vitro (Kalyani
et al. 1998
, 1999
; Mayer-Proschel et al. 1997
;
Rao et al. 1998
). GRPs may further differentiate into
oligodendrocytes or astrocytes but not neurons. Conversely the NRPs may
generate multiple kinds of neurons but not oligodendrocytes or
astrocytes. NRP cells are characterized by E-NCAM/2F7 immunoreactivity
and the absence of A2B5 immunoreactivity while GRP cells are
characterized by the expression of A2B5 and the absence of 2F7 or
E-NCAM immunoreactivity (Kalyani et al. 1998
, 1999
;
Mayer-Proschel et al. 1997
; Rao and
Mayer-Proschel 1997
; Rao et al. 1998
). A great
deal of evidence has been collected to support this model of lineally
restricted, neural development; however, it represents but one of
several views in the current literature (for a discussion on
alternative models see Kalyani and Rao
1998
).
Several lines of evidence indicate that human neural development may
mirror rodent neural development through a significant period of
embryogenesis (Herschkowitz 1988; Mrzljak et al.
1990
). By week six of embryological development, the human
neural tube has differentiated into an outer mantle layer, and an inner
proliferative zone and some neurons have already differentiated fully.
Neurogenesis and gliogenesis proceed over the next several weeks and
most, though not all, neuronal proliferation is completed by 8-10 wk of gestation. Gliogenesis proceeds for longer periods, and multipotent stem cells, GRPs and NRPs can be isolated from embryos at 10-18 wk of
gestation (Chalmers-Redman et al. 1997
; Kalyani
et al. 1998
; Li et al. 1999
; Quinn et al.
1999
; Svendsen et al. 1997
; Tohyama et
al. 1991
). The antigens expressed by human precursor cells, as
well as their growth factor responses, appear similar to those described in rodent cells (Li et al. 1999
;
Scolding et al. 1999
; Thal et al. 1992
).
Several groups have isolated human stem cells (Carpenter et al.
1999
; Johansson et al. 1999
; Vescovi et
al. 1999
; Yandava et al. 1999
). Li et al.
(1999)
showed that human primary neural tube cultures contained
dividing neuron restricted precursors that expressed E-NCAM and could
differentiate into relatively mature neurons. Precursor cells could be
immortalized and neuronal restricted precursor cell lines generated.
The electrophysiological properties of any of these precursor cells and
the differentiated population were not described, and to date only
morphological and antigenic characteristics have been examined.
To test the properties of human neural precursor cells (HNPCs) we
examined two commercial sources of HNPCs isolated at overlapping gestational ages. Here we report the mitotic, antigenic and
electrophysiological phenotypes of cells from one source (Clonexpress)
in a set of baseline culture conditions. This heterogeneous population,
which has been cultured and passaged multiple times in the presence of
serum (cHNPCs), differs from primary HNPCs (pHNPCs), which have been
passaged only once and in the absence of serum. Comparison of the
electrophysiological properties of neuronal precursor cells described
from other species and spatiotemporal points in development (Feldman et al. 1996; Luskin et al. 1997
)
as well as glia and developing neurons in acute and primary cultures
(Black and Waxman 1996
; Sontheimer et al.
1992
) reveal both similarities and differences. We found that
nestin-immunoreactive cells that incorporate 5-bromodeoxyuridine (BRdU)
at high rates are present in these human neural cell cultures. These
cells can differentiate into postmitotic cells that exhibit neuronal
morphologies; express multiple neuronal-specific markers, including
-III tubulin, MAP-2, and neurofilament and a repertoire of voltage-
and neurotransmitter-gated ion channels but do not fire action
potentials. These results show that heterogenous cHNPCs can be directed
to differentiate into a neuronally restricted lineage that appears
arrested at an early stage.
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METHODS |
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Substrate preparation
Poly-L-Lysine (PLL, Sigma, St. Louis, MO) was dissolved in distilled water (13.3 µg/ml) and applied to tissue culture plates for an hour. Excess PLL was withdrawn and the plates were allowed to air dry. Plates were rinsed with medium once before plating the cells.
Human fetal cell cultures
The cHNPCs (Clonexpress, Gaithersberg, MD) are provided as live suspension cells. The cells are obtained from the CNS of 12- to 18-wk embryos. They can be maintained as undifferentiated cultures by plating on uncoated plastic tissue culture dishes (Corning, Corning, NY) in a chemically defined basal medium (provided by Clonexpress) at 37°C in 5% CO2-95% air. The cells can remain undifferentiated for up to five passages.
The pHNPCs (Clonetics, San Diego, CA) were plated in culture dishes (Corning) coated with fibronectin at a dilution of 5,000 cells/dish. Cells were maintained at 37°C in 5% C02-95% air. The basal, chemically defined medium (provided by Clonetics) consisted of Dulbecco's modification of minimal essential medium and Hams F12 medium 1:1 (DMEM/F12) supplemented with additives, basic fibroblast growth factor (bFGF) and epidermal growth factor (EGF).
Growth of cHNPCs cells
Undifferentiated cHNPCs were maintained in medium (Clonexpress) that consisted of DMEM/F12 + 10% FBS (fetal bovine serum) + neuronal cell supplement (NCS, containing bFGF and EGF). NCS was a proprietary supplement provided by Clonexpress and contained insulin, transferrin and an undefined set of additives. Cells will survive for short periods in the absence of serum but could not be passaged or maintained for 7-10 days without serum supplementation as undifferentiated cells.
For neuronal differentiation, cells were plated on PLL-coated dishes at
a density of 1-2 × 105
cells/cm2 as recommended by the supplier. The
differentiation medium consisted of N2 supplement (GIBCO, Grand Island,
NY) to replace serum; NCS (contains bFGF and EGF); and dibutryl cyclic
AMP (DbcAMP, 100 µM/ml, Sigma), nerve growth factor (NGF, 50 ng/ml,
Upstate, Lake Placid, NY), or bone morphogenic protein (BMP-2, 10 ng/ml, Creative Biomolecules, Boston, MA), which was added every 2 day.
The cells were allowed to mature for 4 day in differentiation medium
prior to physiological or immunocytochemical analysis of cells. Under differentiating conditions, the cells could be maintained for 3-4 wk
provided that the medium was changed every 72 h.
Immunocytochemistry
Staining procedures were as described previously (Rao and
Mayer-Proschel 1997). Staining for cell surface markers such as embryonic neural cell adhesion molecule (E-NCAM) was done in cultures of living cells. To stain cells with antibodies against internal antigens, cultures were fixed with 2-4% formaldehyde for 30 min at
room temperature. In general, dishes were incubated with the primary
antibody for one hour, followed by incubation with an appropriate
secondary antibody for an additional hour. Double-labeling experiments
were performed by simultaneously incubating cells in appropriate
combinations of primary antibodies followed by noncrossreactive
secondary antibodies. Negative controls with omission of primary or
secondary antibodies were run simultaneously. The 4',6
diamidino-2-phenylindole (DAPI) histochemistry was
performed as described previously (Kalyani et al. 1997
).
DAPI staining was generally done after all other antibody staining had
been completed. Staining was visualized under phase optics using
special dichroic filters (Omega) that isolate the appropriate
excitation and emission wavelengths associated with each marker and
thereby reduce any marker to marker bleedthrough to low background
levels, though they also reduce overall fluorescence intensity. We also
routinely stain with only one marker to test for bleedthrough across
the filter sets used.
E-NCAM antibody was a hybridoma supernatant obtained from Developmental
Hybridoma Studies Bank (DHSB, University of Iowa, Iowa City, IA).
Neuron-specific -III tubulin and microtubule associated protein 2 (MAP-2) antibodies were obtained from Sigma. An anti-nestin polyclonal
used in some double-labeling experiments was a kind gift of Dr. Keith
Cauley (Signal Pharmaceuticals, San Diego, CA). The A2B5 antibody
(Eisenbarth et al. 1979
) was obtained from American Type
Culture Collection (ATCC, Manassas, VA) and used to label glial
precursor cells as described previously (Rao et al.
1998
). O4 and Gal-C antibodies that recognize specific glycoprotein epitopes expressed by oligodendrocytes were obtained from
DHSB and used as described previously (Rao et al. 1998
). BRdU (Sigma) was used to determine the number of S-phase cells. Mouse
and rat monoclonal anti-BRdU antibodies were obtained from Boehringer
Mannheim (Indianapolis, IN). All secondary antibodies were purchased
from either Jackson ImmunoResearch Laboratories (West Grove, PA) or
Southern Biotechnology Associates (Birmingham, AL).
Electrophysiology
Current- and voltage-clamp recordings were made using the whole
cell patch-clamp technique (Hamill et al. 1981).
Electrodes were pulled from thick-walled borosilicate glass on a
Flaming/Brown P-87 pipette puller (Sutter Instruments, Novato, CA), to
resistances of ~3-4 M
. An Axopatch 200B amplifier was used to
control pipette potentials or inject current, a TL-1 DMA interface to
convert A/D values at 5-50 kHz, filtered at 2-10 kHz, and pClamp 5.5 software to control both the amplifier and the converter (Axon
Instruments, Foster City, CA). Data were analyzed with pClamp 5.5 and
Webfoot (Biodiversity, Park City, UT).
Cells were plated on 12-mm glass coverslips as described in the preceding text. Coverslips were placed in an acrylic chamber on an Olympus IMT-2 microscope and perfused at 1.0-1.8 ml/min. The external bath solution, rat Ringer (RR), consisted of (in mM) 140 NaCl, 3 KCl, 1 MgCl2, 2 CaCl2, 10 HEPES, and 10 glucose. External solutions were set to pH 7.4 with NaOH. Bath ground was established with a 3 M KCl/Agar bridge. External test solutions were made up in RR and included 300 nM tetrodotoxin (TTX), 5 mM 4-AP, and 500 µM acetylcholine (ACh). Test solutions were applied through square glass tubes, which could be rotated into or out of position with respect to the cell using a galvanometer.
The internal pipette solutions varied. To study Na+ currents in isolation, either Cs+ was used to replace internal K+ or internal tetraethylammonium (TEA) was used to block K channels. The CsF internal solution consisted of (in mM) 125 CsF, 15 CsCl, 11 EGTA, and 10 HEPES. The TEA internal solution consisted of (in mM) 125 potassium tetramethylamine-n-oxide (KTMAO), 15 KCl, 5 MgCl2, 11 EGTA, 10 HEPES, and 10 TEA. To measure K+-selective currents, we used a KF internal solution that consisted of (in mM) 125 KF, 15 KCl, 11 EGTA, and 10 HEPES. Some K+-selective currents and action potentials were recorded using a KAsp solution, which consisted of (in mM) 50 KF, 75 KAspartate, 15 NaCl, 11 EGTA, and 10 HEPES. All internal solutions were set to pH 7.2 with CsOH, tetramethylammonium hydroxide or KOH as appropriate. Liquid junction potentials (LJPs) were calculated with Axoscope (Axon Instruments), and data were corrected appropriately. The calculated potentials for all internal solutions with RR as the external solution were: CsF = 9.l mV, KF = 8.4 mV, TEA = 10.0 mV, KAsp = 12.1 mV.
Passive membrane properties such as cell capacitance
(Cm), input resistance
(Ri), and series resistance
(Rs) were calculated from current
records obtained by averaging 24 records stepping from 70 to
80 mV.
Cm was calculated by integrating the
charge under the current transient resulting from the
10 mV step and dividing it by
10 mV. Rs was
calculated by fitting a single exponential to the falling phase of the
capacity transient to obtain the RC time constant,
Rs. Rs
was then calculated as
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Activation parameters were obtained from current-voltage
(I-V) relationships by voltage-clamping the cell membrane to
80 or
100 mV, stepping to test voltages between
80 and +80 mV, in
10-mV increments, for either 10 or 100 ms, and returning to the initial
holding potential. Linear leak currents were subtracted using a P/4
protocol (Armstrong and Bezanilla 1973
). Peak currents elicited by the voltage step were measured in either the inward or
outward direction and plotted against the test voltage, which was
corrected off-line for series resistance and LJP errors. Average activation curves were constructed by averaging the fitted values because the actual test voltages varied with
Rs from cell to cell. Estimates of
activation parameters were obtained by fitting the I-V
relationships to the following form of the Boltzmann equation
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Steady-state inactivation data were acquired by holding the membrane at
120 mV to allow full channel recovery from inactivation, stepping to
a prepulse potential between
120 and
25 mV for 500 ms (3 s for
K+ currents) and then stepping to 0 mV (50 or 60 mV for K+ currents) to measure the available
current. Percent available current was calculated by normalizing the
currents obtained at each potential by the maximum current response,
I/Imax. The distribution I/Imax versus voltage could
then be approximated by a Boltzmann equation
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Intracellular Ca2+ measurements
Calcium-imaging experiments were performed on cHNPCs maintained
under differentiating conditions as described above. Cells were loaded
with 5 µM fura-2 AM and 80 µg/ml Pluronic F127
(Grynkiewicz et al. 1985) in RR for 30 min at
23°C in the dark. The cells were then washed three times with RR, and
the fura-2 AM was allowed to de-esterify for 30 min. Changes in the
ratio of fluorescence emission intensity at 520 nm by excitation at
340/380 nm were measured and correlated to changes in intracellular
calcium, [Ca2+]i, using a
simple two-point calibration scheme
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A Zeiss-Attofluor imaging system (Atto Instruments, Rockville, MD) was
used to acquire and analyze the data, which were sampled at 1 Hz. Cells
were constantly perfused at 1-1.5 ml/min with RR. Neurotransmitters
(500 µM) were made fresh in RR and delivered by bath exchange using a
small volume loop injector (200 µl) located ~250 µl upstream from
the bath. A response to neurotransmitter was defined as a minimum 10%
transient rise over the baseline fluorescence ratio within 60 s
from the time of loop insertion. The dead time from loop to bath inport
was ~10-15 s. Neurotransmitters examined included: -amino-butyric
acid (GABA), glycine (G), dopamine (DA), glutamate (E), and ACh. In
addition, a 50 mM K+ RR solution was used to
depolarize the cells and to test for Ca2+ influx
through voltage-gated channels (45 mM NaCl in RR replaced by 45 mM
KCl). Ascorbic acid (500 µM) was added to dopamine solutions to
prevent oxidation. Control applications of 500 µM ascorbic acid had
no effect (Fig. 9). The pH of all test solutions was adjusted to 7.4 with 1 M NaOH.
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RESULTS |
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Cell Division, morphology, and immunoreactivity
Fetal human CNS cells are now available from two commercial
sources: Clonetics (pHNPCs) and Clonexpress (cHNPCs). Clonetics cells
have been shown to be a mixed population that can generate neurons and
astrocytes (Svendsen et al. 1997). Clonexpress cells are
derived from similar ages but have not been well characterized. To
determine the usefulness of these cells for in vitro studies of human
neuronal development, we obtained cHNPCs, maintained them in culture by
supplier's protocols, and examined their properties under
nondifferentiating and differentiating culture conditions.
Unlike previously described pHNPCs (Li et al. 1999;
Svendsen et al. 1997
), the cHNPCs obtained from
Clonexpress consisted of a relatively undifferentiated population. All
cells expressed nestin and many were dividing in culture as shown by
BRdU incorporation (Fig. 1, 30-40%,
n = 5 independent experiments). Morphologically, cells
appeared flat with few processes, and they formed tight clusters. Cells
could be maintained in an undifferentiated condition for multiple
passages (
5). At both early and late passages, <5% of the cells
expressed GFAP or
-III tubulin immunoreactivity and no cells that
expressed either A2B5 (Eisenbarth et al. 1979
) or other
markers of oligodendrocyte differentiation were detected (data not
shown).
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When cells were replated into differentiation conditions (growth in
DMEM/F12 with NCS containing EGF and bFGF, replacement of serum with N2
supplement, and addition of either DbcAMP, NGF, or BMP-2) cells altered
their morphology and a substantial number (>80%, n = 5 independent experiments) appeared phase bright and generated
processes of greater than two cell diameters (Fig. 1). Immunocytochemistry showed that both -III tubulin and GFAP
immunoreactive cells could be detected in culture and occasional A2B5
reactive cells were seen. No Gal-C or O4 immunoreactive cells
(oligodendrocytes and oligodendrocyte precursors) were detected in any
culture condition (data not shown).
-III tubulin immunoreactive
cells expressed other neuronal markers such as neurofilament (NF) and
MAP-2 but did not incorporate BRdU when cells were differentiated.
DbcAMP, NGF, and BMP-2 all arrested mitosis and enhanced neurite
outgrowth (Fig. 1 and data not shown), but in no condition did we
observe fully mature neurons as assessed by cell size, cell
aggregation, and neurite length.
Passive membrane properties and overall ionic currents
We measured the passive membrane properties of 66 cells with
typical neuronal morphologies (Fig. 1B, maturing) and from 6 cells without processes (Fig. 1A, immature). The maturing
cells had Cm = 32.1 ± 2.1 pF,
Rs = 13.6 ± 1.4 M, and
Ri = 810 ± 127 M
; the
immature cells had Cm = 16.6 ± 2.7 pF, Rs = 9.1 ± 1.7 M
, and
Ri = 867 ± 290 M
(± SE
unless noted). Comparison of these values by Student's
t-test revealed a significant difference in Cm for the two cell types
(P < 0.03), presumably due to neurite outgrowth in the
maturing cells, but not for either Ri
or Rs (P > 0.8 and
P > 0.3), suggesting similar levels of resting
conductances and comparable levels of electrical access to the cell
membrane in the two cell types. Figure 2,
A and B, shows representative currents from an
immature and a maturing, multipolar, neuronal cell obtained by holding
the membrane potential at
80 mV and stepping to test potentials
between
60 and +80 mV in 10-mV increments. While both cell types
express significant time- and voltage-dependent outward currents, none
of the immature cells expressed any inward currents. In contrast, 53 of
64 maturing cells (83%) exhibited recognizable time- and
voltage-dependent inward currents. These data suggest that as the
immature cells stop dividing and begin differentiating, they
concurrently begin expressing Na channels without a significant change
in the density of K channel expression. Resting membrane potentials
(Vrest) were measured from 17 maturing, neuronal cells using the KF (n = 7) or KAsp
(n = 10) internal solutions, ranged from
17.1 to
56.5 mV and averaged
34.3 ± 3.0 mV.
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Outward currentspotassium
The outward currents appeared to be a mixture of several outward,
presumably K+-selective, currents
(IK) in both the immature and maturing
neuronal cells. The transient outward current obvious in the records
(Fig. 2, A and B) resembles the A-type,
4-AP-sensitive current, IK(A), described in neurons and many other cell types (Connor and
Stevens 1971; Rudy 1988
). Figure 2, C
and D, displays currents recorded from the same two cells in
the presence of 5 mM 4-AP. 4-AP eliminated the fast outward current and
revealed a slowly activating outward current that resembled a
delayed-rectifier type of K+ current,
IK(DR). Figure 2, E and
F, represents the 4-AP-sensitive currents obtained by
subtracting the current records in the presence of 5 mM 4-AP from those
in the absence of 5 mM 4-AP. Every cell examined displayed some outward
current except for two neuronal cells, (6/6 immature, 34/36 maturing
neuronal). We obtained the outward current densities by dividing the
peak outward current at +50 mV of each cell by its capacitance. In the
immature cells, the outward current density averaged 36.6 ± 13.4 pA/pF (n = 6) and did not differ from the maturing
neuronal cells, 28.3 pA/pF ± 3.5 pA (n = 34, Student's t-test, P > 0.4). These data
demonstrate that as the neuronal cells begin to differentiate, the
density of K channels in the plasma membrane remains relatively
constant. We further characterized the kinetics of the 4-AP-sensitive
current and the steady-state properties of both the 4-AP-sensitive and -insensitive components of the outward current.
4-AP-sensitive K+ channel kinetics
The time to half-peak (t1/2) and
inactivation time constant (a) of
IK(A) were measured or fit by single
exponential distributions, respectively, from 4-AP-subtracted currents
recorded from the immature (n = 5) or maturing
(n = 4) cells. Since there was no significant
difference for either t1/2 or
a between the two cell types, the
distributions were averaged. Figure
3A shows the time to half-peak
versus voltage for both cell morphologies from a sample of nine cells.
Fitting a single exponential function to this distribution (or
distributions obtained from either the immature or maturing neuronal
cell types separately, from
20 to 80 mV) revealed that
t1/2 gets faster with respect to
voltage and follows a
= 14.0 ± 0.8 mV (
). Figure
3B plots
a versus voltage for the
same group of cells. Inactivation kinetics were relatively voltage
independent and the mean time constant across the voltage range shown
(0-80 mV) was
a = 18.1 ± 0.9 ms. The
similarity in kinetics between the two cell populations suggests that
both the immature and maturing neuronal cell types express the same
4-AP-sensitive K channel. Kinetic analysis was not performed on
IK(DR) as we did not isolate it from
other currents including possible Ca2+-activated
K+ currents.
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K+ channel steady-state inactivation and activation
Steady-state activation and inactivation parameters were
obtained as described in METHODS.
IK(A) generally activates and
inactivates faster and at more hyperpolarized potentials than
IK(DR) (Connor and Stevens
1971). Figure
4A shows
the raw current traces obtained by the steady-state inactivation
protocols shown in the inset from a maturing neuronal cell expressing
both IK(A) and
IK(DR). The majority of
IK(A) is inactivated by
60 mV,
revealing IK(DR), which begins to
inactivate near
70 mV. Figure 4B plots the average I/Imax versus prepulse
voltage for seven cells that expressed a 4-AP-sensitive, transient
current. Fitting a Boltzmann distribution to the
IK(A) availability data yielded a
Vhalf =
80.3 ± 0.5 mV. For
IK(DR), a fit to the availability data
from three cells yielded Vhalf =
42.1 ± 0.8 mV. Activation parameters were fit to either 4-AP-subtracted currents, IK(A), or
4-AP-resistant currents, IK(DR) as
described in METHODS. The fitted values were averaged and
plotted with the inactivation data in Fig. 4, B and
C: IK(A), Vhalf =
18.9 ± 5.2 mV, and
IK(DR),
Vhalf =10.4 ± 7.6 mV
(n = 7). For both currents, a small "window
current" (centered around
57 mV for
IK(A) and
22 mV for
IK(DR)) can be seen where the activation and availability curves overlap suggesting that around these
voltages, steady-state activation of the particular
K+ channel can occur and partially contribute to
Vrest.
|
Inward currents
POTASSIUM.
In several of the cells studied, a nonsaturating, inward current
artifact was observed, caused by the P/4 subtraction protocol activating nonlinear conductances at hyperpolarized potentials (data
not shown). The currents in Fig.
5A were activated by the hyperpolarizing voltage step protocol shown in the inset. The I-V relationships for the peak currents (, taken from the
1st 20 ms after the test voltage step) and the stationary currents (
, measured at 900 ms after the test voltage step) are shown in Fig.
5B. The kinetics and voltage dependence of these
hyperpolarization-activated currents resemble those of an inwardly
rectifying potassium channel, IK(IR), but we did not characterize
these currents pharmacologically. Other currents that may activate in
these voltage ranges include the hyperpolarization activated
nonselective cation channel,
If/Ih, and the human ether-a-go-go-related gene product, HERG.
Ih, however, typically activates over
an order of magnitude more slowly, and HERG channels would typically be
closed at holding potentials of
80 mV where many of the P/4
subtraction protocols activated this current.
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CALCIUM AND SODIUM.
In our recordings, only the maturing neuronal cells expressed inward
time- and voltage-dependent currents so all subsequent recordings were
made from cells with neuronal morphologies (not round). Some cells
showed slowly activating, sustained inward currents that resembled
Ca2+ currents
(ICa), but we did not isolate or
characterize them pharmacologically. Of 64 cells, 53 displayed fast,
transient inward currents that resembled Na+
currents (INa). From these cells, we
calculated a current density = 9.8 ± 1.1 pA/pF
(n = 53) by dividing the peak
INa by the cell capacitance for each
cell and averaging the values.
Sodium channel pharmacology and activation
We studied 26 of the 53 cells further using a CsF or TEA
containing pipette solution and 300 nM TTX to isolate the current from
the accompanying IK and
ICa. TTX has been used as a specific blocker of Na channels (Henderson et al. 1974), and 19 of 26 cells tested (77%) expressed
INa that was blocked by 300 nM TTX
(TTX-S). Interestingly, 4 of the 26 cells expressed
INa that was only blocked 20
50% by
TTX and 3 of the 26 cells expressed
INa that was completely insensitive or
resistant to 300 nM TTX (TTX-R). Figure
6A shows TTX-subtracted
currents recorded from a cell that expressed TTX-S currents;
application of 300 nM TTX blocked all of the
INa in this cell. Figure 6B
shows currents recorded from a cell that expressed TTX-R currents; 300 nM TTX blocked almost none (13%) of the
INa in this cell. We fit the
I-V relationships (with peak INa > |150 pA|) from the
TTX-S and TTX-R currents to a modified form of the Boltzmann equation
(see METHODS) and found that the half voltage for the TTX-S
and TTX-R INa, differed significantly: Vhalf(TTX-S) =
25.4 ± 3.2 mV
(n = 10), Vhalf(TTX-R) =
34.9 ± 1.9 mV (n = 7, P < 0.0004, Student's t-test). TTX-R sodium currents have been
reported in developing dorsal root ganglion neurons (Caffrey et
al. 1992
; Elliott and Elliott 1993
) but show a
depolarizing shift in the voltage dependence of activation and are
considerably slower to activate and inactivate. Another TTX-R sodium
current was first described in glia (Black et al. 1992
;
Sontheimer and Waxman 1992
; Sontheimer et al.
1992
), and termed INa(G) for
glial sodium current. Two different genes have been identified that may
encode this current, Na-G and NaCh6 (Gautron et al.
1992
; Schaller et al. 1995
), and transcript for
both isoforms has been identified in developing and mature neurons and
in glia (Felts et al. 1997
).
INa(G) differs from
ITTX-R in dorsal root ganglion cells
by displaying a hyperpolarizing shift in both the steady-state activation and inactivation curves compared with the classic neuronal TTX-S INa and by exhibiting little difference in
the kinetics of either activation or inactivation. We examined both the
kinetic and steady-state properties of the TTX-S and TTX-R currents to see how they compared with other Na+ currents.
|
Sodium channel kinetics
The time to half-peak and the decay phase kinetics of the TTX-S
and TTX-R currents were measured. Single exponential fits to the decay
phase of the currents provided the inactivation time constant,
h. No significant difference was found between
ITTX-S or
ITTX-R for either
t1/2 or
h at
any voltage (Student's t-test, 0.5 < P < 1 and 0.08 < P < 0.9, respectively, n = 3). Figure
7A shows that the times to
half-peak for ITTX-S and
ITTX-R decrease with voltage and both
could be fit by a single exponential function between
20 and 80 mV,
yielding a
TTX-S = 28.7 ± 2.3 mV that reached a baseline value of 0.48 ms and a
TTX-R = 35.3 ± 2.1 mV that reached a
baseline value of 0.46 ms. Figure 7B plots
h versus voltage where the mean (
10 to 80 mV) for ITTX-S was
h(TTX-S) = 0.56 ± 0.03 ms and for
ITTX-R was
h(TTX-R) = 0.66 ± 0.008 ms (Student's
t-test, P > 0.16). These data demonstrate
that the activation and inactivation kinetics of both the TTX-S and
TTX-R Na+ currents are similar to each other and
to the analogous currents described by Sontheimer et al.
(1992)
.
|
Sodium channel availability
We measured the steady-state availability of sodium channels using
a typical prepulse protocol to various test potentials between 120
and
30 mV to inactivate the channels before stepping to a depolarized
potential (0 mV) to measure the fraction of current left available.
Figure 8A shows the
TTX-subtracted currents evoked in this manner for a cell expressing
TTX-S currents while Fig. 8B shows the currents evoked in
the presence of TTX for a cell expressing TTX-R currents. Figure
8C plots I/Imax
versus prepulse voltage for cells with only TTX-S currents (circles)
and for cells with only TTX-R currents (triangles). The availability
curve for each current was fit by a Boltzmann equation (see
METHODS) to estimate the half-point of inactivation and
Vhalf(TTX-S) =
60.9 ± 1.0 mV
(n = 4) while
Vhalf(TTX-R) =
89.1 ± 1.0 mV
(n = 3). Similar to the activation curves, the
ITTX-R availability curves are
significantly shifted to hyperpolarized potentials when compared with
ITTX-S. Since most of the cells we
studied showed lower INa densities
than typical mature neurons (Feldman et al. 1996
) and 7/26 (27%) expressed TTX-R currents, we expect that this neuronal cell
population represents intermediate progenitors or precursors to fully
mature neurons.
|
Intracellular calcium changes in response to neurotransmitters
To ascertain whether or not the neuronal cells could respond to neurotransmitters like mature neurons, we used fura-2 imaging techniques to monitor [Ca2+]i in response to application of 500 µM GABA, G, DA, ascorbic acid, E, ACh, and elevated extracellular potassium (50 mM) (Fig. 9). Of the 68 cells tested, 34 responded to E (50%), 68 responded to ACh (100%), and 3 responded to 50 mM extracellular K+ (~4%). None of the 68 responded to GABA, G, DA, or ascorbic acid.
|
Electrophsyiological response to ACh
We also measured the currents elicited by application of 500 µM
ACh from two cells in voltage-clamp mode while simultaneously monitoring a group of cells under visual control using fura-red dye.
Like the fura-2-loaded cells, most of the cells under visual control
exhibited an increase in
[Ca2+]i in response to
ACh. Of the two cells we recorded from electrically, both exhibited
inward currents of about 130 pA in response to 300-ms applications of
ACh. Figure 9B shows the responses of one of the cells to a
300- and a 900-ms application of 500 µM ACh. In cells with
Ri ~ 800 M
, these currents could
provide ample depolarization to evoke action potentials, providing the
Na channel density is high enough to support regenerative firing
behavior (V = IR, 80 mV =100 pA*800 M
).
Action potentials
While the neuronal cells expressed sodium currents, the densities
of these currents do not compare to those found in mature neurons
(Feldman et al. 1996). To test whether or not the sodium current could support action potential generation, we used whole-cell current-clamp to inject small, depolarizing currents for 1-200 ms from
various resting potentials (set by constant current injections) and
measured the resulting changes in membrane potential. Of the nine cells
tested, none fired even weak action potentials (data not shown). These
results suggest that the failure of 65/68 cells to increase
[Ca2+]i in response to
elevated extracellular potassium arises, at least in part, from
inadequate densities of available sodium current, and the inability to
generate action potentials in response to membrane depolarization.
Co-expression of GFAP
While the cHNPCs grown in differentiating conditions expressed
multiple neuronal markers and displayed electrophysiological properties
characteristic of neurons, they nevertheless did not appear to mature
completely in culture. A possible reason for the failure to mature may
be exposure to serum. The cHNPCs are grown in serum containing medium
and appear to require serum for their survival. Switching cHNPCs to
serum-free medium prior to initiating differentiation is not possible,
as cells did not survive the change in medium (data not shown). Serum,
however, has been shown to inhibit neuronal differentiation and alter
growth characteristics (Kilpatrick and Bartlett 1993).
Likewise, transformed glial precursor cells will undergo a mesenchymal
transformation and no longer differentiate into oligodendrocytes
(Noble and Mayer-Proschel 1997
). To test if exposure to
serum may have altered the properties of cHNPCs, we examined the
expression of E-NCAM and GFAP by
-III tubulin immunoreactive
cHNPC-derived cells. Cells were grown in Clonexpress-supplied medium,
were allowed to differentiate as previously described, and were
immunostained for co-expression of
-III tubulin with E-NCAM and
GFAP. As can be seen in Fig. 10,
-III tubulin cells derived from cHNPCs did not exhibit E-NCAM immunoreactivity. The failure to detect E-NCAM immunoreactivity was
likely due to culture conditions, as first-passage human fetal cultures
(pHNPCs) derived at the same developmental age, cultured in the absence
of serum, co-expressed
-III tubulin and E-NCAM (Fig. 10).
|
Exposure to serum also appeared to alter GFAP immunoreactivity. We
noted that -III tubulin immunoreactive cells that were derived from
cHNPCs grown in serum co-expressed GFAP. This co-expression was not
seen if primary human cells (pHNPCs) were grown in nonserum containing
medium (compare B and B' with D and
D'). Undifferentiated cHNPCs could be kept in culture
without serum for
7 days but still expressed GFAP when forced to
differentiate by addition of DbcAMP, NGF, or BMP-2. Longer time periods
were not tested due to loss of cell viability in the absence of serum.
Thus culture conditions can modify the properties of precursor cells
and lead to atypical phenotypes and antigen expression.
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DISCUSSION |
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Morphology, mitotic activity, and immunoreactivity
Developing human CNS cells obtained from Clonexpress (12-18 wk
gestation) could be maintained in a mitotically active state when grown
in medium supplemented with serum, bFGF, and EGF. These cells were
without processes and displayed a flat, round morphology with
nestin immunoreactivity. By replacing serum with N2 supplement and
adding DbcAMP, NGF, or BMP-2, we found that the cells could be forced
to stop dividing and differentiate. The differentiating cells send out
long processes, over two cell diameters, in one, two, or multiple
directions and begin to resemble neurons. These neurite outgrowths were
correlated with an increase in whole-cell capacitance. In addition, the
differentiating cells lost nestin immunoreactivity and many displayed
-III tubulin, NF, MAP-2, or GFAP immunoreactivity. Only a small
amount of staining was found for A2B5 and no staining was found for
nestin, GalC, or O4, suggesting the relative absence of oligodendrocyte
precursors, oligodendrocytes, astrocyte precursors, and type 2 astrocytes. Thus the exchange of growth factors induces the
proliferating human CNS cells to become a postmitotic population of
cells that resemble neurons in morphology and antigenicity
(
-III tubulin+, NF+,
MAP-2+), but also display the glial antigenic
marker GFAP. We went on to describe the functional phenotype of
these maturing, neuronal cells by studying their functional properties
using whole-cell patch-clamp and fura-2 imaging techniques.
K+ currents
The presence of outward K+ currents in both
the round, immature, mitotic, nestin+ cells and
the neuronal, postmitotic, -III tubulin+ cells
was not surprising since most cell types have been shown to express K
channels (Rudy 1988
). Both cell types had at least two
kinds of K+ currents, a delayed-rectifier type
K+ current and a transient, A-type,
4-AP-sensitive K+ current. These currents had
similar steady-state and kinetic properties in both cell types and
resembled those described in neurons, glia, and the precursor cells
that give rise to either of these neural cell types (Barres et
al. 1990
; Feldman et al. 1996
; Ritchie
1991
; Sontheimer 1994
; Stewart et al.
1999
; Swanson et al. 1990
; Verma Kurvari
et al. 1997
). The presence of K+
currents in the dividing population of cells contrasts with the observations of Feldman et al. (1996)
. They reported
that dividing, EGF-dependent cells in neurospheres, derived from the
subventricular zone (SVZ) of postnatal rat (P0-P3), do not express K
channels until they begin differentiating following attachment to a
suitable substrate. Similar to our observations, Luskin et al.
(1999)
, recorded several K+ currents from
dividing,
-III tubulin+, neuronal progenitor
cells, obtained directly from the SVZ of postnatal rat (P0-P1). Our
data indicate that the human, EGF/bFGF-dependent, mitotic, neural
progenitors express at least two K+ currents, as
do their progeny, similar to the SVZ cells. Although we expected to
find K+ currents expressed in the
nestin+ cells, we imagined that the postmitotic,
neuronal cells would express a higher density of current, which was not
the case. Instead, the main difference in the repertoire of ion channel
expression between the two cell types was the absence of inward current
in the mitotic, nestin+ cells. These inward
currents include an inwardly rectifying K+
current, a slowly activating and inactivating
Ca2+ current, and a fast activating and
inactivating Na+ current. We next
concentrated on describing the INa expressed by
the postmitotic,
-III tubulin+, maturing
neuronal cells.
Na+ currents
We used external TTX, a common and specific Na channel blocker, to
study the TTX-subtracted currents, which should represent the
Na+ current specifically. Surprisingly, we noted
that seven cells expressed some INa
that was not blocked by 300 nM TTX. The TTX-S and -R currents were
compared and found to differ in steady-state activation and
inactivation properties but not kinetics. The TTX-S INa resembled the classical neuronal
INa in that it was half-maximally activated at 25 mV and half inactivated at
52 mV. Like the neuronal INa, the TTX-S current turned on and
off quickly (
h) and was sensitive to nanomolar
concentrations of TTX. The TTX-R INa
steady-state parameters were shifted in the hyperpolarized direction
compared with the TTX-S so that TTX-R was half-maximally activated at
35 mV and half-maximally inactivated at
80 mV; similar to the TTX-R current, INa(G), described in
astrocytes, Schwann cells, and some neurons (Barres et al.
1989
; Howe and Ritchie 1990
; Sontheimer and Waxman 1992
). Like the currents described by
Sontheimer et al. (1992)
, but not the other groups, the
kinetics of the TTX-R current recorded from these neuronal precursor
cells were fast, essentially identical to those of the TTX-S current.
While the expression of Na channels suggests a neuronal phenotype, the
densities of the currents in these cells (~9 pA/pF at peak) were
lower than those found in mature neurons and more comparable with those
found in cultured glial and neural progenitor cells (Feldman et
al. 1996
). Although Luskin et al. (1999)
did not
report Na+ current densities, they did note that
28/32 cells tested had INa, but 11/12
tested could not fire action potentials, suggesting a low density of
Na+ current. The
INa recorded by Feldman et al.
(1996)
from EGF-dependent neurosphere-derived cells, which
differentiated in the presence of EGF approached mature, neuronal
densities (~50 ± 35 pA/pF); while
INa recorded from cells that
differentiated in the presence of bFGF (9.4 ± 3.7 pA/pF) remained
closer to the values that we observed. They also noted a decrease in K
channel density when bFGF was included in the culture medium,
suggesting that bFGF may generally inhibit ion channel expression in
these culture systems. Several reports have described the expression of
Na channels in developing neurons and have shown that several Na
channel isoforms are up-regulated as neurons mature, including both the
"glial," INa(G) (TTX-R) and the
"neuronal," rat brain IIA (TTX-S), isoforms (Felts et al.
1997
). The cHNPC derived, maturing, neuronal cells may resemble
their in vivo cousins in that they may first express a low density of
Na channels in early differentiated states, followed by a sharp rise in
the density of Na channels in terminally differentiated states. For
reasons unknown, we may have arrested the differentiation process at an
early step, where early neuronal markers and fundamental channel
expression have been turned on but not amplified or fine-tuned. In cell
cultures, transcriptional regulation can be influenced dramatically by
serum, growth factors, or co-culture with other cell types. In
particular, the expression pattern of ion channels in both neurons and
glia appears highly dependent on whether or not these two cell types
are cultured together (MacDonald et al. 1996
;
Maxwell et al. 1996
; Thio et al. 1993
),
and only additional studies will allow us to understand the factors
necessary to promote both proliferation of progenitor cells and
complete differentiation and maturation of progeny. This work provides
a structural and functional baseline to compare with neurons induced to
differentiate under different culture conditions: various growth
factors, co-culture with glia, glial or neuronal conditioned medium,
explants, or transplants.
Neurotransmitter responses
Having established that the neuronal cells express ion channels
typical of neurons, we next examined their response profile to a set of
neurotransmitters. Fura-2 Ca2+ imaging
demonstrated that all of the maturing neuronal cells responded to ACh,
a majority to E, a few to elevated extracellular potassium, and that
none responded to GABA, G, DA, or ascorbic acid. The lack of response
to ascorbic acid is expected as a negative control. Because GABA and
glycine receptor ion channels are selective for chloride ions,
intracellular Ca2+ responses to GABA or glycine
are not expected even in the presence of these receptors unless the
reversal potential for chloride depolarizes the cell in response to
receptor activation, leading to an opening of
Ca2+ channels. Likewise, if dopamine receptors
are present, they may act on signaling mechanisms that are uncoupled
from both membrane depolarization and Ca2+ fluxes
and so the lack of response to dopamine does not rule out surface
expression of certain dopamine receptor subtypes. The responses to
glutamate were invariably smaller, ~10% of baseline, and slower, 30- to 40-s latency, than the responses to ACh, ~200% of baseline,
~10-20 s latency (see Fig. 9A, latencies uncorrected for
dead time). These results differ from those obtained when similar cells
were examined from rat and mouse (Kalyani et al. 1998;
Mujtaba et al. 1999
). In both the rat and mouse, NRP
cells gave rise to a postmitotic population of
E-NCAM+ cells with neuronal morphologies and
antigenicities. The differentiated rat cells responded to both ACh and
E with similar high-frequency and magnitude, some cells responding to
either or both. A substantial fraction responded to DA and elevated
extracellular potassium while a few responded to GABA and G
(Kalyani et al. 1998
). The differentiated mouse cells
also displayed a high-frequency and magnitude of response to both ACh
and E, with only a small fraction responding to DA, and few responding
to elevated extracellular potassium (Mujtaba et al.
1999
). Together, these data suggest that the population of
human, differentiated neuronal cells is less heterogeneous than those
of the rat or mouse, and perhaps less mature. In particular, the
failure of the human (and mouse) cells to respond to elevated
extracellular potassium suggests any or all of the following
mechanisms: cells may not set Vrest via K channels, cells may lack voltage-dependent Ca channels, or cells
may lack a sufficient density of available Na channels to fire an
action potential.
Action potentials
To test whether Na channel density was sufficient to generate
action potentials, we examined several of the maturing neuronal cells
using current-clamp to inject depolarizing currents and monitor the
membrane potential response. In nine cells tested by injecting currents
from 10 to 200 pA for 1 to 200 ms, we were unable to elicit an action
potential. In some cases, the decay of the membrane depolarization
appeared slower than the onset and suggested that a
Na+ or Ca2+
conductance may be sustaining the response slightly, but the density of
such depolarizing influences was simply not enough to fire a classic,
regenerative, neuronal action potential. These results are similar to
those obtained by Luskin et al. (1999), who showed that
despite a recognizable sodium current in the dividing SVZ cells, the
SVZ cells were also unable to fire action potentials in response to
depolarizing current injections. So despite a neuronal complement of
ion channels and receptors, these differentiating cells have not
developed the proper balance of channels to support action potential
generation. Future work examining perturbations in the culture
conditions will be necessary to determine what factors are necessary to
allow or promote full maturation of human neurons in culture.
Significance of staining patterns
Although bFGF has been reported to enhance neuronal proliferation
and differentiation in some culture systems the opposite has been
reported in others (Feldman et al. 1996; Kuhn et
al. 1997
; Murphy et al. 1990
; Vescovi et
al. 1993
). In particular, the work of Feldman et al.
(1996)
should be noted in which the EGF-dependent neuronal
precursors appeared "stunted" in the differentiation process if
cultured in the presence of bFGF instead of EGF. In addition, these
authors reported that many of the
-III
tubulin+ cells in the presence of bFGF expressed
GFAP. Several other groups have observed either permissive or
inhibitory effects on differentiation that were related to the presence
of serum in the culture medium (Kilpatrick and Bartlett
1993
). Double labeling of the differentiating Clonexpress cells
for
-III tubulin+ and GFAP confirmed
co-expression of these markers on single cells and revealed that
-III tubulin+ cells did not stain for ENCAM. A
different population of human precursor cells, isolated from
overlapping gestational ages (Li et al. 1999
) was shown
to express ENCAM and not to co-express GFAP but was never exposed to
serum. These data underscore the direct effect culture conditions have
on the developing phenotypes of differentiating neuronal cells.
Collectively, these data show that a self-renewing population of cells
can be isolated from human developing CNS that can give rise to both
neurons and glia. External factors play a critical role in shaping the
fates of these cells as they divide and differentiate. The replacement
of serum with N2 supplement and addition of NGF, DbcAMP, or BMP-2
arrests mitosis and initiates a process of neural differentiation that
restricts some of the cells to what may be an astrocytic fate while
most of them take on neuronal phenotypes. The maturing neuronal
phenotype is characterized by a lack or slow rate of cell division;
-III tubulin, NF, and MAP-2 immunoreactivity; expression of K, Ca,
and Na channels typical of neuronal tissue; functional responses to ACh
and glutamate; an inability to fire action potentials; and
co-expression of GFAP. When primary human precursor cells were grown
and differentiated in the absence of serum the cells did not co-express
-III tubulin and GFAP. We suggest that the presence or absence of an
undetermined factor arrests the full maturation of these developing
neurons in culture. Future studies will be aimed at understanding the
nature of these factors, their interactions with these developing cells
and the expression patterns they dictate.
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ACKNOWLEDGMENTS |
---|
The authors thank Dr. W. C. Michel, Dr. T. Parks, and M. Zabel for critical reviews of this manuscript. The authors thank the Developmental Studies Hybridoma Bank (DSHB) for the antibodies used. DSHB is maintained by the University of Iowa under contract NO1-HD-7-3263 from the National Institute of Child Health and Human Development). M. S. Rao gratefully acknowledges the constant support of Dr. S. Rao through all phases of this project.
This work was supported by the Muscular Dystrophy Association and a National Institutes of Health first award to M. S. Rao; D. R. Piper and M. T. Lucero were supported by National Institute on Deafness and Other Communication Disorders Grant DC-82994.
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Address for reprint requests: M. T. Lucero, Dept. of Physiology, University of Utah School of Medicine, 410 Chipeta Way, Salt Lake City, UT 84108.
Received 12 November 1999; accepted in final form 23 February 2000.
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REFERENCES |
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