Department of Physiology, School of Medicine, University of California at San Francisco, San Francisco, California 94143
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ABSTRACT |
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Olson, Andrew J., Arturo Picones, and Juan I. Korenbrot. Developmental Switch in Excitability, Ca2+ and K+ Currents of Retinal Ganglion Cells and Their Dendritic Structure. J. Neurophysiol. 84: 2063-2077, 2000. In the retina of teleost fish, continuous neuronal development occurs at the margin, in the peripheral growth zone (PGZ). We prepared tissue slices from the retina of rainbow trout that include the PGZ and that comprise a time line of retinal development, in which cells at progressive stages of differentiation are present side by side. We studied the changes in dendritic structure and voltage-dependent Ca2+, Na+, and K+ currents that occur as ganglion cells mature. The youngest ganglion cells form a distinct bulge. Cells in the bulge have spare and short dendritic trees. Only half express Ca2+ currents and then only high-voltage-activated currents with slow inactivation (HVAslow). Bulge cells are rarely electrically excitable. They express a mixture of rapidly inactivating and noninactivating K+ currents (IKA and IKdr). The ganglion cells next organize into a transition zone, consisting of a layered structure two to three nuclei thick, before forming the single layered structure characteristic of the mature retina. In the transition zone, the dendritic arbor is elaborately branched and extends over multiple laminae in the inner plexiform layer, without apparent stratification. The arbor of the mature cells is stratified, and the span of the dendritic arbor is well over five times the cell body's diameter. The electrical properties of cells in the transition and mature zones differ significantly from those in the bulge cells. Correlated with the more elaborate dendritic structures are the expression of both rapidly inactivating HVA (HVAfast) and of low-voltage-activated (LVA) Ca2+ currents and of a high density of Na+ currents that renders the cells electrically excitable. The older ganglion cells also express a slowly activating K+ current (IKsa).
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INTRODUCTION |
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In
the course of development the dendritic structure and functional
connectivity of individual neurons frequently transform from an
initial, immature pattern to a final, mature one through activity-dependent rearrangements (reviews in Fields and Nelson 1992; Goodman and Shatz 1993
; Purves
1994
). Ganglion cells in the vertebrate retina provide
remarkable examples of activity-dependent developmental changes. The
afferent activity of these cells is critical for the development of the
structure and connectivity of the cells' postsynaptic partners (review
in Katz and Shatz 1996
; Shatz 1996
). In
both lower (Olson and Meyer 1991
; Reh and Constantine-Paton 1985
; Schmidt 1985
,
1990
) and higher vertebrates (Casagrande and
Condo 1988
; Galli-Resta et al. 1993
;
Stryker and Harris 1986
), blocking action potentials in
the ganglion cells disrupts the orderly development of their axon
terminals and their connectivity to next order neurons. This block does
not affect the dendritic structure of the ganglion cells themselves
(Campbell et al. 1997
; Wong et al. 1991
).
The input connectivity of ganglion cells also matures with development.
In the rabbit's retina, for example, ganglion cells respond robustly
to light by postnatal day 10 (P10), but only by
P21 do they exhibit the adult pattern of visual field
organization (Bowe-Anders et al. 1975
; Masland 1975
). Developmental maturation of ganglion cell field
organization also occurs in lower vertebrates (Sernagor and
Grzywacz 1995
). The development of the ganglion cells'
dendritic structure depends on incoming electrical activity. Laminar
stratification and receptive fields of ganglion cells fail to mature
when synaptic activity of cells in the pathway between photoreceptors
and ganglion cells is blocked (in higher vertebrates: Bodnarenko
et al. 1995
; in lower vertebrates: Sernagor and Grzywacz
1996
). The mechanism of this phenomenon is unclear
(Tagawa et al. 1999
).
To investigate the mechanisms of activity-dependent dendritic
restructuring in ganglion cells, it must first be established that
developmental maturation occurs both in the electrical properties of
the cells and in their structure. Elegant studies of ganglion cells in
retinas isolated from animals at various developmental stages have
demonstrated that the electrophysiological properties of the cells
change with development. Cells both dissociated and in tissue pieces
have been studied (Rorig and Grantyn 1994; Rothe and Grantyn 1994
; Schmid and Guenther 1996
,
1999
; Skaliora et al. 1995
; Wang
et al. 1997
). In studies of dissociated cells, however, it is
not possible to correlate electrical and structural events. In studies
of retinal pieces, whether slices or whole-mount, developmental changes
can only be established with relatively limited precision, by comparing
tissues isolated from different animals. To overcome some of these
limitations we have developed a new experimental preparation amenable
to electrophysiological and anatomical investigation, a retinal slice
from teleost fish that includes the peripheral growth zone
(Olson et al. 1999
).
Teleost fish are unique among vertebrates because their eyes continue
to grow throughout the life of the animal (Grun 1975; Lyall 1957
; Muller 1952
). Addition and
differentiation of new retinal tissue occurs continually in a narrow
ring at the periphery of the mature retina and in a fissure that
extends along the ventral pole from the center to the edge of the
retina. The ring of developing tissue is known as the peripheral growth
zone (PGZ) (Johns and Easter 1977
; Kock
1982
; Kunz and Callaghan 1989
; Lyall
1957
; Meyer 1978
; Negishi et al.
1985
). In the PGZ there is a continuous gradient of
developmental stages that extends from undifferentiated stem cells at
the very margin to the fully mature, central retina. Thus in the PGZ
time and distance along an equator are equivalent, and every stage of
retinal differentiation is simultaneously present, side by side. Mature
ganglion cells (GC) in the fish retina process visual information with
spatial, spectral, and dynamic complexity similar to that displayed by
GC in mammals (Bilotta and Abramov 1989a
,b
).
We report here on a study of the developmental changes in the intrinsic electrical properties and gross dendritic structure of ganglion cells in the PGZ of the trout retina. We describe some of the salient anatomical features of the developing cells and the changes in electrical excitability and the expression, density, and biophysical properties of the ionic currents that reflect the activity of voltage-gated K+ and Ca2+ ion channels.
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METHODS |
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Materials
Rainbow trout (Onchorynchus mykiss) were raised from
fertilized eggs received from Mt. Lassen Trout Farm (Red Bluff, CA). The aquaculture facility and conditions of animal care are described elsewhere (Julian et al. 1998). Animals used in these
experiments were juveniles 5-7 cm long, a size that was reached
uniformly 5-10 wk posthatching. The Animal Care Committee at
University of California at San Francisco approved experimental
protocols. All drugs and chemicals were obtained from Sigma Chemicals
(St. Louis, MO).
Retinal slices for electrophysiological studies
The procedure to obtain retinal slices that include the PGZ is
described in detail elsewhere (Olson et al. 1999).
Briefly, after 45-60 min dark adaptation the fish is killed by rapid
decapitation and is double-pithed. A section in the middle of the eye
along the naso-temporal axis that includes the PGZ is removed and
placed, retina side down, on a piece of Millipore filter membrane
(Millipore, Bedford, MA) at the edge of the recording chamber. The
sclera, choroid, and pigment epithelium are lifted away, and the retina is covered with ice-cold normal Ringer solution with 0.1% bovine serum
albumin (BSA; Table 1). Slices (240 µm)
are cut at the equator of the retina with a tissue chopper in which the
blade cuts with a "rolling" motion. Slices are maneuvered into
their proper position in the recording chamber using the protruding ends of the filter membrane, which are secured in slots filled with
silicone vacuum grease.
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Ganglion cell morphology
Dendritic morphology of developing ganglion cells was
investigated both by filling individual cells and by Golgi staining. For single-cell fills, we used either intracellular electrodes of ~40
M resistance filled with 5% neurobiotin/l% Lucifer yellow in 0.1 M
Tris, pH 7.6, or tight-seal electrodes filled with the normal filling
solution containing 5% neurobiotin/l% Lucifer yellow. Trout retina
PGZ slices were prepared as above, and the recording chamber placed on
the fixed stage of an upright microscope equipped with differential
interference contrast (DIC) and epifluorescence optics. Slices were
superfused with Ringer solution. Individual ganglion cells in each of
the three developmental regions (bulge, transition, and monolayer) were
visually identified and then impaled. With intracellular electrodes,
effective cell penetration and cell integrity were assessed by
observing the cell under epifluorescence after filling it with Lucifer
yellow using
2 nA current applied for 5-10 s. If the cell was
intact, it was loaded with biotin using +2 nA current applied for 1-2
min (either constant current, or 50% duty cycle). Five or six fill
attempts could easily be made at intervals along the slice without
danger of overlapping dendritic fields from filled cells. With
tight-seal electrodes, membrane resistance was assessed after achieving
whole cell mode. Cells were maintained under current-clamp at holding
voltages of
60 mV for 5-10 min. Cell filling occurs mostly by
exchange diffusion between the electrode and the cell's interior.
After cell filling, slices were fixed overnight at 4°C with 4% paraformaldehyde in iso-osmotic phosphate-buffered saline (PBS). After 3 × 10-min rinses in PBS, the slices were incubated for 4 h in PBS + 0.1% Triton X100, rinsed again with PBS, then incubated for 1 h in avidin-horse radish peroxidase diluted 1:1,000 in PBS (Vector Labs, Burlingame, CA). After thorough rinsing in PBS, filled cells were labeled with 3,3'-diaminobenzidine (DAB substrate kit, Vector Labs, Burlingame, CA). In some slices, and before immunostaining, we observed the fixed tissue under a confocal microscope (Bio-Rad, Richmond, CA).
Our Golgi staining procedure is based on the method developed for
goldfish retina by Stell (1975). The extent of staining obtained with the Golgi method varied considerably, despite every effort to repeat methodology exactly. In addition to reporting the
concentrations of reagents that typically gave us good staining, we
also give the ranges within which we obtained useful results. A section
of retina containing the PGZ and attached to a piece of Millipore was
held on a glass slide with small dabs of silicone grease. The retinal
piece and filter membrane were then covered with a small square of the
perforated plastic film from a Telfa brand bandage, which was secured
onto the glass slide with cyanoacrylate glue applied at its corners
(Loctite, Hartford, CT). The "sandwiched" retinas were fixed for 1 day in one part 25% glutaraldehyde plus four parts 2.5%
K2Cr2O7
(we found that here, and in the next step, up to 3.5%
K2Cr2O7
can be used). The tissue was treated for 2 days (this time can be
increased to as much as 14 days, if necessary) with 2.5%
K2Cr2O7.
The slides were drained and placed in 1.4% AgNO3
solution and maintained in darkness for 1 day (we found the useful
range of AgNO3 concentration to be 1.0-1.5%).
Following a brief rinse in water, we removed the plastic film and
sliced the retinal piece using the tissue chopper as described above. Individual slices were dehydrated through successive alcohol solutions, cleared in xylene, mounted in histological mounting media (Permount®; Fisher Scientific, Fair Lawn, NJ), and coverslipped.
Two-dimensional profiles of neurobiotin-filled or Golgi-stained cells
were traced with the aid of a computer-controlled upright microscope
using NeuroLucida software (Micro Brightfield, Colchester, VT).
Electrical recordings
For electrophysiological studies, three to four retinal slices cut from the naso-temporal equator were held in the recording chamber on the fixed stage of an upright microscope equipped with DIC optics. We observed them using a ×40, 0.75 N.A. water-immersion objective lens (Zeiss, Oberkochen, Germany) with an overall magnification of ×400. The external solution, normal Ringer (Table 1), was saturated with 100% oxygen and continuously superfused at a flow rate of ~0.5 ml/min.
We measured whole cell membrane currents under voltage clamp using an
Axopatch 200A patch-clamp amplifier (Axon Instruments, Foster City,
CA). A Ag/AgCl reference electrode was connected to the bath through a
1 M KCl agar bridge. Analog signals were filtered at either 1 or 5 kHz
with an eight-pole low-pass Bessel filter (Frequency Devices,
Haverhill, MA) and digitized on-line with 12-bit accuracy at a sampling
rate of either 2.5 or 12.5 kHz (Indec Systems, Sunnyvale, CA). We used
tight-seal electrodes produced from aluminosilicate glass (Corning
1724, 1.5 × 1.0 mm, OD × ID). Whole cell mode was achieved
either by mechanical disruption or by chemical perforation. For
perforation, the electrode filling solution contained either nystatin
(100 µg/ml) or amphotericin B (112.5 µg/ml) prepared as detailed
elsewhere (Miller and Korenbrot 1994).
Ionic solutions
In whole cell mode, whether attained by disruption or perforation, the tight seal electrode was filled with normal intracellular solution (Normal ICS, Table 1), pH adjusted to 7.3 with KOH, osmotic pressure 303 mosM. The normal extracellular Ringer was modified in various experimental protocols, as detailed in Table 1.
Data analysis
Capacitance was measured under the ionic conditions that
isolated Ca2+ currents (see
Voltage-dependent Ca2+ currents). We
integrated the capacitative current transient generated by either 20-
or +10-mV steps from a holding voltage of
100 mV and divided by the
voltage pulse amplitude. To calculate membrane capacitance, the
electrode capacitance was measured immediately after forming a
giga-seal and this value subtracted from the capacitance measured under
whole cell mode. In the analysis of K+ currents,
we did not determine a normalized K+ current in
each cell (pA/pF), as we did in the analysis of
Ca2+ currents, because the acquisition bandwidth
that we used to record the long epochs necessary to study
K+ currents (up to 4 s) was not sufficiently
fast to accurately determine capacitance in the same cell.
Selected functions were fit to experimental data using nonlinear,
least-square minimization algorithms (Origin, Microcal software, Northampton, MA). Statistical errors throughout are given as means ± SE. To compare means among more than two populations, it is common
to execute ANOVA. However, ANOVA simply tests the null hypothesis and
determines whether any of the means differ from each other, with some
confidence interval. If the means are not the same, other tests of
significance must be executed to determine which two means are not the
same. This paired comparison of means is valid as long as the total
number of comparison is less than or equal to six (Snedecor and
Cochran 1967). Since we only compared three populations at a
time, we report here the significance of difference between means
calculated by pairing any two of the three populations at a time and
applying Student's two-tailed t-test.
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RESULTS |
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Ganglion cell morphology
On the basis of gross histological criteria, we identified three
distinct regions in the ganglion cell (GC) layer of the PGZ in the
retina of trout. The typical features of these regions are illustrated
in Fig. 1 (for a more detailed
description, see Olson et al. 1999). Just beneath the
initial portion of the inner plexiform layer (IPL) is a multi-layered
region of GC somas that stands out as an oval-shaped bulge of cells
(Fig. 1, top left). This region consists of the youngest
GCs, and we refer to it as the bulge. In the morphologically
mature retina, the GC somas form a single layer, so we refer to this
region as the monolayer (Fig. 1, top right). In
between the bulge and monolayer regions, the GC somas form a bilayer;
we call this the transition zone (the initial part of the
transition region can be seen in Fig. 1, top right).
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Individual cells in each of the three regions have different
dendritic morphologies. Figure 2
illustrates tracings of typical neurons in each of the three regions in
the GC layer. The tracings are projections onto a single plane of the
cellular structures. These tracings were created with either of two
methods: 1) hand tracing of biotin-filled single cells using
a camera lucida attachment and observing fixed tissue after development
with an immunomarker linked to avidin and 2)
computer-assisted image reconstructions of Lucifer yellow-filled cells
using a confocal microscope. Results with both methods are
indistinguishable and pooled. In addition, we also traced with the
camera lucida single cells stained with the Golgi method. This method
was, in our hands, of very limited success. The cells we selected for
this illustration reflect the typical features of cells found in each
of the three regions. We studied the anatomical details of 12 cells in
the bulge, 19 cells in the transition, and 15 cells in the mature
region. Our sampling size is sufficient to identify the major gross
features of the dendritic tree in the developing ganglion cells. It is not sufficient to define fine features comparably to previous developmental anatomical studies of others (Hitchcock and Easter 1986). Such detailed studies, moreover, would be unwise in
tissue slices, since dendritic trees are likely truncated.
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Within the restrictions of our anatomical universe, changes in
dendritic structure as ganglion cells matured from bulge to mature
regions are so dramatic as to be readily definable. GC in all three
regions had dendrites, identified as the structures connected to the
cell bodies and located in the IPL. We scored dendritic trees by
1) their length and branching, 2) their extension into the IPL and, most especially, 3) their stratification
within the IPL. A feature common to mature ganglion cells of all
species, and first described in fish retina by Famiglietti et
al. (1977) is the selective distribution and termination
of dendrites in distinct lamina of the IPL. Dendrites are stratified
either in a single lamina or in two laminae (mono- and bistratified
ganglion cells). Sublamina a, nearest the inner nuclear
layer, is the layer in which the dendrites of the hyperpolarizing
(OFF) ganglion cells distribute (Fig. 2, mature GC on the
left). Sublamina b, nearest the GC cell bodies,
is the layer in which the dendrites of the depolarizing
(ON) ganglion cells distribute (Fig. 2, mature GC on the
right). Bulge cells have extremely short and sparse
dendritic trees. On average, the longest single dendrite was 1.35 ± 0.72 times the longest cell body axis. The average long body axis
was 17.2 ± 6.6 µm, and the short body axis was 12.6 ± 5.2 µm. Dendrites in bulge cells did not end in a
specific lamina and, on average, extended 25.5 ± 8.3% of the
width of the IPL. In the first third of the transition zone, the
dendritic trees are long, on average, 3.45 ± 1.3 times the
longest cell body axis. The average long body axis was 12 ± 3.3 µm, and the short body axis was 10.5 ± 1.8 µm. The dendrites
are elaborately branched, and they extend across the full width of the
IPL but are not stratified within the IPL (Fig. 2, transition cells,
left and middle). Stratification of dendritic
trees first appears in the central two-thirds of the transition zone
(Fig. 2, transition cell on the right). We defined a
stratified dendritic tree as one that 1) does not reach
across the full width of the IPL and 2) terminates in a
well-defined laminae of the IPL. Within that lamina, the dendrites
extend parallel with the GC layer. The dendritic trees of all cells in
the monolayer region are stratified (Fig. 2). Along with the growth of
bipolar and amacrine cell synaptic terminals, these changes in
individual GC dendritic morphology are responsible for the rapid
increase in the thickness of the IPL as retinal maturation proceeds
(compare top panels of Fig. 1) (see also Olson et al.
1999
).
Voltage-dependent ionic currents in ganglion cells
We systematically explored the features of voltage-dependent ionic
currents in the cells in the three developmental regions of the GC
layer. In fish, <1% of the cells in the ganglion cell layer are not
ganglion cells (probably displaced amacrines) (Hitchcock and
Easter 1986). The distinct anatomical regions in the PGZ of rainbow trout described above are correlated with specific changes in
the electrophysiological properties of the cells. We identified the
principal voltage-dependent Ca2+ and
K+ membrane currents in GC using
established biophysical and pharmacological criteria. For each class of
currents, we first describe their defining features and then detail
their developmental changes.
Voltage-dependent Ca2+ currents
We investigated the properties of Ca2+
currents in slices continuously bathed with Ringer containing
tetrodotoxin to block voltage-dependent Na+
currents, tetraethyl ammonium (TEA) and 4-aminopyridine (4-AP) to block
all identified K+ currents (see below) and
diisothiocyanatostilbene-2,2-disulfonic acid (DIDS) to block
Cl currents (Table 1,
Ca2+ Ringer). Whole cell mode was attained either
by membrane disruption or by perforation. In the conventional whole
cell mode, the tight-seal electrode was filled with a solution
containing Cs+ and EGTA (Table 1,
Cs+, EGTA ICS) to further reduce
K+ currents and to attenuate possible
Ca2+-dependent currents, such as
Ca2+-dependent K+ and
Ca2+-dependent Cl
currents. Results with both methods were indistinguishable and are
presented together. Under these experimental conditions, depolarization to between
60 and +10 mV from a conditioning voltage of
100 mV,
activated inward currents that were blocked by external
Co2+ and that therefore we identify as
Ca2+ currents. There was also a small and
variable outward current that is unidentified. We recognized three
distinct Ca2+ currents by their voltage
dependence of activation, and their voltage dependence and time course
of inactivation: 1) high voltage activated and rapidly
inactivating (HVAfast), 2) high voltage activated and slowly
inactivating (HVAslow), and 3) low voltage activated (LVA).
Identification of these current types is an operational definition
based on and consistent with commonly identified
Ca2+ current types (Hille 1992
).
We do not hold that these current types necessarily correspond to a
single molecular ion channel type. The identification of
particular molecular type(s) of Ca2+ channel
requires investigation of the pharmacological action of a collection of
specific toxins (Birnbaumer et al. 1994
), a task now
left for future work.
Voltage dependence of activation and inactivation
Typical features of HVA Ca2+ currents are
illustrated in Fig. 3. Illustrated is a
current with slow inactivation. Holding membrane voltage was 70 mV.
To investigate the activation voltage, membrane voltage was first held
at
100 mV for 500 ms to remove any steady-state inactivation and was
then successively stepped up to +40 mV in 10-mV increments. This
voltage protocol activated inward currents that reached a peak within
20 ms and then inactivated slowly and only partially (Fig. 3, top
left). The current-voltage (I-V) curve of these
currents (Fig. 3, bottom left) showed an activation
threshold between
50 and
40 mV and a maximum amplitude at about
20 mV. For n = 25 cells, on average, threshold
voltage was
40.8 ± 1.6 mV, and peak current occurred at
9.2 ± 1.8 mV.
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HVA currents either inactivated rapidly or very slowly (some not at
all). For rapidly inactivating currents, HVAfast (see below for further
definition), we determined the voltage dependence of the inactivation
by successively holding the membrane voltage for 500 ms at values
between 120 and +10 mV, in 10-mV increments, and then stepping the
voltage to
20 mV (Fig. 3, top right). The inward currents
activated at
20 mV were large only when the conditioning pulse was
more negative than
60 mV. Conditioning pulses more positive than
60
mV reduced the peak amplitude of the current activated at
20 mV,
indicating that the current was inactivated. We analyzed the voltage
dependence of steady-state inactivation by determining the dependence
on conditioning voltage of the normalized current activated at
20 mV.
We normalized the current activated at
20 mV by dividing the peak
amplitude measured after each conditioning pulse by the maximum peak
amplitude, that measured following a
100-mV conditioning voltage. The
normalized current decreased with increasing voltage, and this
dependence is well described by the Boltzmann function (Fig. 3,
bottom right)
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Typical features of LVA currents are illustrated in Fig.
4. Holding membrane voltage was 70 mV.
To investigate the activation voltage of these currents, membrane
voltage was first held at
100 mV for 500 ms to remove any
steady-state inactivation and was then successively stepped in 10-mV
increments up to +40 mV. This voltage protocol activated inward
currents that reached a peak within 10 ms and then inactivated rapidly
and completely (Fig. 4, top left). For four cells the
average time-to-peak of the maximum current was 9.8 ± 3.1 ms. The
I-V curve of these currents (Fig. 4, bottom left)
showed an activation threshold at
70 mV and a maximum amplitude at
about
50 mV. For four cells, on average, threshold voltage was
70 ± 0 mV, and peak current occurred at
50 ± 8.2 mV.
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To investigate the steady-state inactivation of the LVA currents, we
successively held the membrane voltage for 500 ms at voltages between
120 and +10 mV, in 5-mV increments, and then stepped the voltage to
50 mV. The inward current activated at
50 mV was large only when
the conditioning pulse equal to or more negative than
100 mV. Pulses
more positive than
100 mV reduced the current peak amplitude,
indicating that current is inactivated after 500 ms. We measured the
current amplitude activated at
50 mV as a function of the
conditioning voltage and normalized the amplitude as described above
for HVA currents. The dependence of normalized current amplitude on
conditioning voltage is well described by the Boltzmann function
(Eq. 1). For four cells, on average,
V1/2 was
68.4 ± 5.8 mV and
k was 4.0 ± 0.9 mV.
Kinetics of inactivation
The typical inactivation features of HVA and LVA
Ca2+ currents are illustrated in Fig.
5. Based on kinetics, we identified HVA currents as HVAfast or HVAslow. For HVAfast, 90% of the inactivating component exhibited a time course well described by a single
exponential
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All LVA current inactivated rapidly at a rate that was voltage
dependent (Fig. 5, middle). At any given voltage, the
inactivation is well described by a single exponential function
(Eq. 2; Fig. 5, bottom). For four cells, the
average value of at the voltage that generated the maximum peak
amplitude was 8.0 ± 2.9 ms.
Developmental changes in voltage-dependent Ca2+ currents
Over all our sample of cells that expressed Ca2+ currents (n = 29), in all but two we observed only one type of Ca2+ current, as defined above. Of these, both were mature cells that coexpressed LVA and HVAslow.
In the bulge zone, Ca2+ currents of any type were expressed in only 46% of the cells (6 of 13), but about 90% of the cells in the transition and monolayer zones expressed Ca2+ currents (23 of 26; Fig. 6, left). There was no statistically significant difference in the fraction of cells expressing these currents between transition and monolayer zones.
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HVASLOW CURRENTS. About half of the bulge cells studied (6 of 13) expressed any voltage-dependent Ca2+ currents, and these were exclusively HVAslow currents (Fig. 6).
To compare current amplitudes in the various cell stages, we normalized the Ca2+ current amplitude measured in each cell by the capacitance of the same cell. We measured cell capacitance through an analysis of the capacitative transient in the voltage-clamped current, as detailed under METHODS. Assuming that specific membrane capacitance (1 µF/cm2) does not change with development, dividing current amplitude by the cell's capacitance is a reliable method to assess membrane current density. The mean capacitance was as follows: bulge cells, 11.1 ± 3.7 pF; transition cells, 6.7 ± 1.2 pF; and monolayer cells, 7.6 ± 1.6 pF. The values are not significantly different. In the bulge cells that expressed HVAslow, the current density tended to be smaller in amplitude than in transition or monolayer cells (Fig. 7, top). This difference in amplitude must now be stated as a tendency, rather than a statistical difference, because the universe of cells is small and the dispersion in the data significant (Fig. 7). There are no statistically significant differences between the HVA currents in transition and monolayer cells with respect to their amplitude and voltage dependence of activation (Fig. 7).
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HVAFAST CURRENTS. Bulge cells did not express HVAfast currents (0 of 13). Of the cells that expressed Ca2+ currents in transition and monolayer stages, about 35% of the cells expressed HVAfast currents (8 of 23, Fig. 6, right). There is no statistically significant difference in the fraction of cells expressing this current in transition and monolayer stages. There were no statistically significant differences between the HVAfast currents in transition and monolayer cells with respect to their amplitude, voltage dependence of activation (Fig. 7, middle), voltage dependence of inactivation, or inactivation rates.
LVA CURRENTS. Bulge cells did not express LVA currents (0 of 13). These currents were observed only in transition and monolayer cells, in 17% of those cells with Ca2+ currents (4 of 23, Fig. 6, right). The biophysical features of LVA were essentially the same in all cells that expressed them. Cells expressing LVA currents did not simultaneously express any other type of Ca2+ current. Figure 7 (bottom) illustrates average I-V curves of normalized LVA currents measured in transition and monolayer GC. There are no statistically significant differences between the currents in their amplitude, voltage dependence of activation, voltage dependence of inactivation, or inactivation rates. Thus only the more mature transition and monolayer cells express HVAfast and LVA currents, the youngest (bulge) cells exclusively express HVAslow. Once the currents are expressed, however, their biophysical features are essentially invariant, all that changes are the levels of expression.
Development of electrical excitability
Mature ganglion cells are electrically excitable and sustain
Na+-dependent action potentials (Kaneda
and Kaneko 1991a; Lipton and Tauck 1987
). In the
course of development, spiking activity in GC is first spontaneous,
then correlated among neighboring cells (Meister et al.
1991
; Wong et al. 1993
), and, finally, light driven (Bowe-Anders et al. 1975
; Masland
1975
). We wondered whether at their earliest developmental
stage, GC that express voltage-dependent Ca2+
currents might also sustain Ca2+-dependent action
potentials. In other excitable cells, a developmental switch from
Ca2+ to Na+-dependent
spikes has been documented (Moody 1995
; Spitzer
and Ribera 1998
). We investigated electrical excitability by
measuring voltage under current clamp using the whole cell mode and a
patch-clamp amplifier. While this method is a technical compromise that
underrepresents the true time course and amplitude of the action
potentials (Magistretti et al. 1998
), it provides,
nonetheless, an effective means to test cell excitability.
In normal Ringer, we recorded from each GC under voltage clamp. If the
cell exhibited inward current between 50 and
10 mV, we then
switched to current clamp. The mean zero current membrane voltage was
not statistically different among the various developmental stages:
bulge,
34 ± 20 mV (n = 6); transition,
43 ± 9 mV (n = 7); and monolayer,
52 ± 9 mV (n = 3). We applied the current necessary to hold
the membrane voltage at
75 mV and then applied successive current
steps to both hyper- and depolarize the cell (Fig.
8). Membrane depolarization above
55 mV
readily generated action potentials in nearly all transition and
monolayer cells (5 of 6 cells), but action potentials could not be
elicited in bulge cells even at voltages near zero mV (5 of 6 cells;
Fig. 8). The action potentials were reversibly blocked with 1 µM TTX (Table 1, TTX Ringer, data not shown). Thus in agreement with all
previous findings (review in Robinson and Wang 1998
),
mature GC sustain Na+-dependent action
potentials, and the youngest GC are simply not excitable, even if they
express voltage-dependent Ca2+ currents.
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We do not report here on the features of voltage-dependent
Na+ currents of developing GC, although we
observed them under voltage clamp, as is obviously expected. Inadequate
clamp of Na+ currents was apparent by the absence
of a graded, voltage-dependent increase in time-to-peak of these
currents (data not shown). The issue of quality of space clamp in
ganglion cells studied in retinal slices is thorny. In general,
experimental and theoretical arguments suggest that space clamp is
adequate for only relatively slow events (slower than a few ms)
(Taylor et al. 1996; Velte and Miller 1996
). Note that Ca2+ currents (see
above) are sufficiently slow to be well clamped, as evidenced by the
fact that the activation threshold, I-V curves and voltage
dependence of inactivation of these currents are similar to those of
dissociated neurons (Hille 1992
).
Voltage-dependent K+ currents
We investigated the properties of K+
currents of GC recorded in conventional whole cell mode using
electrodes filled with normal intracellular solution. Under these
experimental conditions, depolarization to +50 mV, from a holding
voltage of 100 mV, activated outward currents that were attenuated by
5 mM external TEA (data not shown), and therefore we identify them as
K+ currents. We recognized three distinct
K+ currents by the kinetics and voltage
dependence of activation, the kinetics of inactivation, and the
sensitivity to specific K+ channel blockers:
1) a noninactivating delayed rectifier (IKdr), 2)
a slowly activating delayed rectifier (IKsa), and 3) a
rapidly inactivating, "A" type current (IKA).
Kinetics of inactivation
Typical features of K+ currents that
demonstrate an inactivating component are illustrated in Fig.
9. Three different cells are illustrated
that represent the range of kinetic behavior we observed. To
investigate the inactivation kinetics, cells were bathed in Ringer
containing TTX and Co2+ to block
Na+ and Ca2+ currents
(Table 1, Co2+, TTX Ringer). The membrane
voltage, held at 70 mV, was first stepped to
100 mV for 750 ms to
remove any steady-state inactivation and was then successively stepped
for 3 s in 10-mV increments up to +50 mV. Pulses were repeated at
4-s intervals. At depolarizing voltages (
0 mV), the current rapidly
reached peak (
30 ms) and then inactivated. In every instance tested
(6 of 6), the inactivating component reversibly disappeared in the
presence of 5 mM 4-AP, a specific blocker of IKA type of inactivating
current (Table 1, Co2+, TTX, 4-AP Ringer). The
residual currents (those measured with the same voltage protocols in
the continuous presence of 4-AP) are illustrated for each cell in the
middle panels of Fig. 9. These currents are voltage
dependent and essentially of constant amplitude for the duration of the
voltage pulse. The bottom panels illustrate the
4-AP-sensitive current, computed from the difference between the
initial and the residual currents. In Fig.
10 we illustrate the I-V
curves measured at the maximum amplitude of the residual (top
left) and the peak of the 4-AP-sensitive (top right)
currents in each of the three cells shown. Residual and 4-AP-sensitive currents have nearly identical I-V curves. Since these
currents 1) were measured under conditions that block
Na+ and Ca2+ currents,
2) are both attenuated by external TEA, and 3)
have the same voltage dependence of activation, we identify them as K+ currents of two types: inactivating (IKA) and
noninactivating (IKdr).
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The kinetics of IKA inactivation indicate that about 90% of the
inactivating component declined with a single exponential time course
with a time constant that is voltage dependent (Fig. 10,
bottom). At +50 mV, this time constant ranged in value
between 80 and 700 ms over all our cell population. We did not average these values because they did not distribute uniformly nor did they
cluster at particular values. We did not discern a pattern to the
distribution of time constant values. The large range of values likely
reflects a mixture of the expression level of the many molecular forms
of the members of the Shaker family of inactivating K+ channels (Panyi and Deutsch
1996).
Activation kinetics
IKdr and IKA activated rapidly; for all cells measured time to
maximum or peak currents was 20 ms at +50 mV. We distinguished a
third type of K+ current by its kinetics of
activation. Typical features of these currents are illustrated in Fig.
10. Membrane voltage was held at
100 mV for 1 s and was then
successively stepped in 10-mV increments up to +50 mV for 900 ms.
Pulses were repeated at 4-s intervals. The outward currents activated
at large depolarizing voltages activated instantaneously and then
continued to grow in amplitude. This behavior was observed for all
voltages greater than +10 mV. The extent and speed of activation of the
slow component was voltage dependent, but the value of the rate of
activation varied from cell to cell. The cells illustrated in Fig.
11 present the range of kinetic
behavior we observed for IKsa. At +50 mV the time to reach midway
between the starting and final values ranged from 38 to 258 ms. We did
not average data because these values were widely dispersed. The
I-V curves of the currents measured for the cells shown
either instantaneously or at the end of the voltage pulse are shown in
Fig. 11. The curves have the same shape, they simply differ in
amplitude. Since the instantaneous and slowly activating currents were
attenuated by external TEA and since they have the same voltage
dependence of activation, we identify them as K+
currents of two types: slowly activating (IKsa) and rapidly activating, noninactivating (IKdr, delayed rectifier type). IKsa currents are
reminiscent of slow K+ currents described in
other tissues (Hille 1992
).
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Developmental changes in voltage-dependent K+ currents
Every cell that we investigated expressed K+ currents (68 of 68). Some expressed only one type of current, invariably IKdr (27 of 68, 39%) while others expressed a mixture of only two currents (41 of 68, 61%), one of which was always IKdr. Developmental changes were reflected in two parameters: the level of expression of IKA and IKsa and the amplitude of the total peak K+ current.
The expression of IKA and IKsa currents changed with development. Every bulge cell we studied expressed both IKA and IKdr (9 of 9), but none expressed IKsa (0 of 9). In the transition zone all cells expressed IKdr (25 of 25, 100%), and of these about half (13 of 25, 52%) expressed IKdr alone. The combination IKdr + IKA was observed in 40% of the cells (10 of 25) and the combination of IKdr + IKsa in only 8% (2 of 25). In the monolayer, all cells expressed IKdr (34 of 34), and of these, again, nearly half (14 of 34, 41%) expressed IKdr alone. The combination IKdr + IKA was observed in 50% of the cells (17 of 34) and the combination of IKdr + IKsa in only 9% (3 of 34). At all stages of development, in cells with mixed current we could not discern any pattern in the ratio of the two currents present. Thus the expression of IKA and IKsa appear to be developmentally regulated: IKA is found in all bulge cells, but only in about 40-50% of the transition and mature cells. Expression of IKsa is relatively infrequent, about 8 to 9%, and only occurs in transition and mature cells.
We measured the total K+ current amplitude as the
peak of the outward current activated by a voltage step to +50 mV from
a 70-mV holding voltage. Figure 12
illustrates the changes in total current amplitude with cell
maturation. The average peak currents were as follows: bulge cells,
628 ± 180 pA (n = 15); transition cells,
1,494 ± 227 pA (n = 14); and monolayer cells,
1,296 ± 219 (n = 21). The mean of the bulge cells
differs significantly (P < 0.05) from those of either
transition or monolayer cells, which do not differ from each other.
Since the average cell capacitance does not change with maturation (see
above), the data indicate that the surface density of
K+ channels increases with development and the
changes in current amplitude do not simply reflect changes in cell
dimension.
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DISCUSSION |
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Developmental changes in dendritic structure
In the ganglion cell layer of the PGZ, three different regions can be discerned by gross histological criteria: bulge, transition, and monolayer zones. Cells in the bulge have spare and short dendritic trees that extend, at most, about 25% of the width of the IPL. As the cells mature, their dendritic structure changes. In the cells in the most peripheral third of the transition zone, the dendrites are elaborately branched and extend over the full width of the IPL, but do not stratify or terminate in specific sublamina of the IPL. Over the central two-thirds of the transition zone and in the monolayer, dendrites of the GC stratify in the IPL into distinct sublaminae a and b. Expression and biophysical features of voltage-dependent currents in the young GC of the bulge are different from those in transition and monolayer GC, which do not differ from each other. That is, voltage-dependent ionic currents change dramatically between cells that have a sparse and minimal dendritic structure and those with an elaborate dendritic arbor, but do not change with the stratification of the dendrites.
Anatomical studies in 250-µm-thick retinal slices likely
underrepresent the magnitude of changes in dendritic arbor, yet
facilitate the identification of changes in stratification. In elegant
studies, Hitchcock and Easter (1986) comprehensively
documented the developmental changes in arbor size of GC labeled by
retrograde transport of horseradish peroxidase (HRP) added to the optic
nerve. Our results, while in general agreement with their findings,
extend information in two respects: 1) we have mapped the
extent of dendritic stratification, a feature that can be readily
analyzed in slices, but not in the whole-mounts studied by
Hitchcock and Easter (1986)
; and 2) we explored the dendritic structure of bulge cells, cells that cannot be
filled by retrograde transport of HRP, because their axons are not
sufficiently long to emerge from the eyeball in the optic nerve.
As in other vertebrates, mature ganglion cells in fish have been
recognized to be of distinct anatomical subtypes. Specific classification schemes have been proposed by Naka and Carraway (1975), Murakami and Shimoda (1977)
, Kock
and Reuter (1978)
, Dunn-Meynell and Sharma
(1986)
, Hitchcock and Easter (1986)
, Cook
et al. (1992)
, and Cook and Sharma (1995)
. In
our studies we did not attempt to classify the developing GC into
specific categories because we could not discern systematic patterns of
change in the population we sampled.
Developmental changes in voltage-dependent ionic currents
The ionic currents in mature ganglion cells of trout are similar
to those previously described in other ganglion cells, both in higher
and lower vertebrates (rodent: Guenther et al. 1994; Lipton and Tauck 1987
; Rothe and Grantyn
1994
; Schmid and Guenther 1996
; Wang et
al. 1997
; cat: Huang and Robinson 1998
;
Kaneda and Kaneko 1991a
,b
; Skaliora et al.
1993
; ferret: Wang et al. 1998
; goldfish:
Bindokas and Ishida 1996
; turtle: Liu and Lasater
1994
; review in Robinson and Wang 1998
).
In brief summary, we have found that about half of the youngest identified GC (bulge cells) do not express Ca2+ currents, and those that do only express HVAslow. Bulge cells are not electrically excitable. All bulge cells express K+ currents, invariably a mixture of IKA and IKdr.
Expression and biophysical features of all currents in transition and monolayer cells are essentially the same, but they differ significantly from those in the bulge cells. Ca2+ currents are expressed in the vast majority of the cells; LVA currents and rapidly inactivating HVA currents are first switched on. Transition and monolayer cells are electrically excitable, and their action potentials are blocked by TTX. This indicates that they express voltage-gated Na+ currents. In the bulge, we found that 67% of the cells (10 of 15) expressed voltage-gated Na+ currents; their density, however, is evidently not sufficient to sustain action potentials. All transition and monolayer GC express K+ currents, but IKA is switched off in about half the cells. About half of the cells express only IKdr, while the other half express a mixture of two currents, one of which is always IKdr and the other is, commonly, IKA and, infrequently, IKsa.
Developmental changes in voltage-dependent currents, then, are of two principal types: 1) switching currents on or off and 2) changing the current density. On the other hand, we did not observe large changes in the biophysical features, e.g., voltage dependence of activation and inactivation, of the currents expressed.
Developmental changes in currents have previously been investigated in
GC dissociated from fetal cat retinas. LVA, HVAslow, and HVAfast
Ca2+ currents have been identified. All are
expressed at every developmental stage investigated, with a continuous
increase in the current density and in the fraction of cells expressing
LVA and HVAfast (Huang and Robinson 1998). The fact that
a significant fraction of the youngest GC we sampled lack
Ca2+ currents suggests that our sample included
GC at an earlier developmental stage than those in the cat.
Na+ currents are expressed in the youngest cells
examined and increase in density as time progresses (Skaliora et
al. 1993
). K+ currents of both the IKA
and IKdr type are expressed and young cells generally express both, but
as they mature it is more frequent to find cells expressing IKdr alone
(Skaliora et al. 1995
). These important reports
demonstrate that changes in the electrical properties occur, but their
correlation with specific changes in structure or network function
cannot be assessed in dissociated cells.
The limitations of cell dissociation have been addressed in studies of
the electrical properties of GC in isolated retinas or retinal slices
prepared from rats or mice at various developmental stages
(Rorig and Grantyn 1994; Schmid and Guenther
1996
, 1999
). In both rats and mice, there is an
early onset of both Na+ and
Ca2+ currents, with a significant fraction of the
cells failing to express either current. There is a progressive
increase in density of both Ca2+ and
Na+ currents, reaching mature values slightly
earlier for Na+ than Ca2+
currents (in rats at around P6 for Na+
and P18 for Ca2+). Among
Ca2+ currents, LVA are expressed in the youngest
cells and are entirely absent in mature ones. In the cells expressing
HVA currents, the fraction of cells expressing inactivating versus
noninactivating HVA was, essentially, constant (Schmid and
Guenther 1996
). The absence of LVA currents in mature GC of
retinal slices is surprising because dissociated, mature rat GC express
a complex mixture Ca2+ current types, including a
prominent LVA component (Guenther et al. 1994
). This
discrepancy is unexplained; it could be a sampling problem or a true
effect of dissociation.
GC in all species evidence a developmental regulation of
Ca2+ currents, a phenomenon observed in other
neurons as well (Desmadryl et al. 1998; Hilaire
et al. 1996
; Tarasenko et al. 1998
), but there
are striking species-specific differences. In rodents, only LVA
currents are expressed in the youngest GC cells, and the fraction of
cells expressing these currents decreases as development progresses, just as the expression level of HVA currents increases. In fish, in
contrast, the youngest cells do not express LVA currents or inactivating HVA currents, and the fraction of cells expressing these
currents increases with maturation. GC in cats have not been explored
at nearly as early an age as in fish and rats, nonetheless their
developmental pattern is more like that of fish than of rats. Younger
cells express LVA and inactivating HVA currents at low levels, and the
fraction of cells expressing these currents increases with development.
The differences in developmental changes between GC of rats and those
of fish and cats may reflect the need to build functionally distinct
neuronal networks in the retina. Rodent retinas are nearly free of cone
photoreceptors and the animals are essentially scotopic; their visual
system is driven almost exclusively by rod inputs. Fish and cats, in
contrast, are animals in which rod- and cone-driven pathways coexist
and the intensity of the background light determines which of the two
pathways dominates the ganglion cell signal.
Functional significance
Elevated extracellular K+ promotes the
expression of neuronal phenotypes in dissociated cultures of developing
rat retinal cells, specifically favoring the differentiation and
maturation of ganglion cells (Araki et al. 1995).
Elevated K+ is also a necessary condition for
neurotrophic factors to promote survival of dissociated rat GC
(Meyer-Franke et al. 1995
). In both of these reports, it
is argued that the effect of K+ is not direct,
but depends on its ability to depolarize the cells. Changes in the
density and identity of K+ currents, relative to
other currents in the cell, must unavoidably affect the value of
resting membrane potential. Indeed, we observed a systematic difference
in the density and identity of K+ currents and of
resting potential of GC in the bulge versus the more mature cells. If
the effects of membrane voltage observed in vitro indeed occur in vivo,
then the changes we observe in K+ currents and
resting potential may be important factors in the normal progression of
GC development.
It is plausible that changes in Ca2+ current
correlated with developmental processes are causally related (e.g.,
Desmadryl et al. 1998; Hilaire et al.
1996
; Schmid and Guenther 1999
), but proving a
causative role is difficult. We must await direct experimental tests of
this hypothesis, for which the fish model system is well suited. As a
matter of speculation, however, it is interesting to consider that the
most dramatic effect we have observed are switches in the expression of
Ca2+ current types. A different set of
Ca2+ channels and a different resting potential
as cells develop inevitably implies that the resting cytoplasmic
Ca2+ concentration changes with development.
Since young GC are not excitable, there is no reason to suspect large
changes in membrane voltage, although changing synaptic input may be of
consequence. Thus in the youngest fish GC, the higher resting membrane
potential and the expression of HVAslow current suggests that a steady
inward flux of Ca2+ exists at all times. In the
youngest rat GC, in contrast, at comparable resting potentials and
given the exclusive expression of LVA currents, a continuous inward
flux of Ca2+ is unlikely. Most
Ca2+-dependent regulatory processes in neurons,
whether cytoplasmic or nuclear, respond only slowly to
Ca2+ concentration changes and their functional
performance, therefore depends on the standing
Ca2+ concentration. Significant differences may
exist between younger and older GC in the standing cytoplasmic
Ca2+ concentration that may be an important
biochemical regulator of cellular maturation.
By identifying changes in expression of voltage-dependent currents of retinal ganglion cells and correlating them with the developmental state of the cells, our studies provide a basic framework for further investigation of the role of electrical activity in GC maturation. Our results suggest several potentially fruitful lines of inquiry. Simultaneous anatomical and electrophysiological studies should reveal whether specific patterns of current switching correlate with anatomical classes. Such a study would also provide a developmental map in the PGZ with finer spatial resolution, which might reveal tighter correlations between expression of particular currents and dendritic stratification. Finally, for future studies of the development of neurotransmitter-gated currents, these results have provided us with an understanding of the maturational changes in intrinsic membrane properties on which the developing synaptic currents are overlaid.
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ACKNOWLEDGMENTS |
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We thank C. Chung, M. P. Faillace, D. Hackos, T. Ohyama, and T. Rebrik for helpful comments and discussion. We are indebted to L. Bocksai for creative and unfaltering technical support and to Dr. W. K. Stell for insightful advice on the Golgi staining method.
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FOOTNOTES |
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* A. J. Olson and A. Picones contributed equally to the research effort.
Address for reprint requests: J. I. Korenbrot, Dept. of Physiology, School of Medicine, Box 0444, University of California at San Francisco, San Francisco, CA 94143 (E-mail: juan{at}itsa.ucsf.edu).
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.
Received 7 July 1999; accepted in final form 14 June 2000.
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REFERENCES |
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