Steroid hormone-dependent expression of blockersensitive
ENaCs in apical membranes of A6 epithelia
Lynn M.
Baxendale-Cox1,
Randall
L.
Duncan1,
Xuehong
Liu1,
Kieron
Baldwin2,
Willem J.
Els2, and
Sandy I.
Helman1
1 Department of Molecular and
Integrative Physiology, University of Illinois at Urbana-Champaign,
Urbana, Illinois 61801; and
2 Department of Anatomy and Cell
Biology, University of Cape Town Medical School, Cape Town, South
Africa
 |
ABSTRACT |
Weak channel blocker-induced noise analysis was
used to determine the way in which the steroids aldosterone and
corticosterone stimulated apical membrane
Na+ entry into the cells of
tissue-cultured A6 epithelia. Among groups of tissues grown on a
variety of substrates, in a variety of growth media, and with cells at
passages 73-112, the steroids
stimulated both amiloride-sensitive and amiloride-insensitive
Na+ transport as measured by
short-circuit currents in chambers perfused with either growth medium
or a Ringer solution. From baseline rates of blocker-sensitive
short-circuit current between 2 and 7 µA/cm2, transport was stimulated
about threefold in all groups of experiments. Single channel currents
averaged near 0.3 pA (growth medium) and 0.5 pA (Ringer) and were
decreased 6-20% from controls by steroid due to the expected
decreases of fractional transcellular resistance. Irrespective of
baseline transport rates, the steroids in all groups of tissues
stimulated transport by increase of the density of blocker-sensitive
epithelial Na+ channels (ENaCs).
Channel open probability was the same in control and stimulated
tissues, averaging ~0.3 in all groups of tissues. Accordingly,
steroid-mediated increases of open channel density responsible for
stimulation of Na+ transport are
due to increases of the apical membrane pool of functional channels and
not their open probability.
electrophysiology; sodium channels; tissue culture; cortical
collecting ducts; kidney; noise analysis
 |
INTRODUCTION |
WHEREAS STEROID HORMONES are known to stimulate
Na+ transport in tight epithelia,
the specific steps and mechanisms involved in steroid-mediated
regulation of epithelial amiloride-sensitive apical membrane
Na+ channels (ENaCs) are unknown.
Both mineralocorticoids and glucocorticoids act over periods of hours
to increase Na+ entry into the
cells (19) via open channels (15). Because changes of open channel
density may reflect changes of channel open probability
(Po)
and/or the density of functional channels (NT),
the fundamental question has been whether regulation of transport
occurs by way of
Po
and/or
NT.
Using methods of blocker-induced noise analysis, Baxendale et al. (2)
first reported that aldosterone and corticosterone stimulated transport
by increase of
NT
in tissue-cultured A6 epithelia. In native tissues of rat renal
cortical collecting ducts, patch clamp revealed that stimulation of
transport by diet and aldosterone occurred by increase of
NT
(14). Similarly, A6 epithelia grown on rat tail collagen films respond
to aldosterone by increase of
NT
(12). Because the ways in which cells regulate their channels may
depend on the substrate on which A6 cells are grown (3, 11), we were
led in scope of the experiments reported here to study a variety of
passages of A6 cells (passages
73-112) originating in different laboratories on
substrates other than collagen-coated Nuclepore membranes that were
used in our original experiments (2) of steroid effects on
Po
and
NT.
We report that aldosterone and corticosterone stimulate transport by
increase of
NT
with no change of
Po
of amiloride-sensitive Na+
channels, regardless of substrate, serum, growth medium, or baseline expression of transport.
 |
MATERIALS AND METHODS |
Cell culture.
A6 cells from American Type Culture Collection (ATCC; Rockville, MD) at
passage 69 were subcultured repeatedly
in plastic flasks and used at passages
73-112. Three groups of experiments to be referred
to as groups A, B, and
C were done with differences not only
in passage number but also the growth medium and the permeabilized
substrates on which the tissues were grown. Cells in
group A at passages
73-80 originated in Dr. R. L. Duncan's laboratory
(Renal Division, Jewish Hospital, St. Louis, MO), where confluent
tissues were grown on collagen-coated Nucleopore membranes (0.8-µm
pore size; Worthington, Freehold, NJ), as described previously (9).
Confluent monolayers were brought to Urbana for the experimental part
of the studies that were carried out in 1985-1986. Cells in
group B were purchased from ATCC at
passage 69, subcultured, used at
passage 75 with tissue growth on
Millicell HA substrates (Millipore, Bedford, MA), and studied in 1994 in Urbana. Cells in group C were
obtained as a gift to Dr. W. J. Els from Dr. W. Van Driessche, used at
passages 107-112 with tissue
growth on Millicell HA substrates, and studied in 1995 in Cape Town.
The results of other groups of experiments were the same, with tissues grown on Transwell-Clear (Costar, Cambridge, MA) and Anocell (Whatman, Clifton, NJ) substrates and in a Leibovitz-Ham growth medium (16, 17),
and will not be reported here.
Growth and perfusion media.
The growth medium for group A
experiments was a glutamine-, glucose-, and pyruvate-supplemented
Dulbecco's modified Eagle's medium (D5648, Sigma Chemical, St. Louis,
MO) diluted 15%. NaHCO3 (8 mM),
penicillin (100 U/ml), streptomycin (100 µg/ml), insulin (5 mU/ml),
and 10% fetal bovine serum (FBS) were added to this medium. Cells and
tissues were maintained in a humidified incubator at 28°C with air
containing 1.7% CO2. Ten days
after seeding of the cells on the collagen-coated Nuclepore membranes,
the tissues were fed serum- and insulin-free medium. The tissues were
studied on days 14-26 in their
control, steroid-depleted states of transport.
The growth medium for group B and
C experiments was a Dulbecco's
modified Eagle's medium (84-5022EC, GIBCO, Grand Island, NY) with
N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic
acid (HEPES; 4 mM), penicillin (25 U/ml) and streptomycin (25 µg/ml)
(17-719R, BioWhittaker, Walkersville, MD), and 10% FBS (Hyclone,
Logan, UT). Cells and tissues were grown in the presence of humidified air containing 1% CO2 in an
incubator at 28°C. Removal of the FBS before treatment with
steroids was found to be unnecessary and was not done in these groups
of experiments.
We report the results of experiments in which control and
steroid-treated tissues originated from the same lots of tissues. Before the day of an experiment, tissues were fed either growth medium
(control) or the same medium containing exogenous steroid (0.27 µM
aldosterone or corticosterone). Both control and steroid-treated tissues were studied at intervals between ~4 and 50 h postfeeding and
handled in the same ways during transfer of the tissues from the
inserts to continuous perfusion chambers designed for noise analysis
(1). Group A tissues were perfused
with a Ringer solution consisting of 100 mM NaCl, 2.4 mM
KHCO3, 1.0 mM
CaCl2, and 5 mM glucose.
Group B and
C tissues were perfused with growth
medium minus the FBS and antibiotics, thereby ensuring that the tissues were being studied under essentially the same conditions under which
they were grown.
Electrical measurements.
The methods of study with blocker-induced noise analysis were identical
to those described in detail previously (6, 7, 10). After transfer to
the chambers, the tissues were short-circuited continuously for at
least 1 h to allow the macroscopic short-circuit currents
(Isc) to
stabilize. Thereafter, during periods of ~30 min, the apical
membranes of the cells were exposed in steps to increasing
concentrations (5-50 µM) of the weak
Na+ channel blocker
6-chloro-3,5-diaminopyrazine-2-carboxamide (CDPC; Aldrich Chemical,
Milwaukee, WI). Current noise at each blocker concentration
(B) was amplified, digitized, and Fourier transformed to
yield power density spectra from which the low-frequency plateaus (S0) and corner
frequencies
(fc)
of the induced Lorentzians were determined by nonlinear curve fitting
of the combined Lorentzians, "1/f" noise at the lower
frequencies and amplifier noise at the higher frequencies. Blocker on-
and off-rate coefficients
(kob and
kbo,
respectively) were calculated from the slopes and intercepts of
B
2
fc plots,
respectively, yielding the blocker equilibrium constant
KB = kbo/kob.
After complete washout of CDPC from the apical solution and
restabilization of the
Isc
(~10-15 min), the channels were completely (~99.8%) blocked
with 100 µM amiloride to yield the amiloride-insensitive current
(IAmilsc). When all
Na+ was removed from the apical
solution (1:1 substitution with impermeant cations), the
IAmilsc fell to zero,
indicating that the IAmilsc
was due to amiloride-insensitive Na+ conductive channels at the
apical membranes of the cells.
Defining the blocker-sensitive macroscopic currents
IBNa = IBsc
IAmilsc, the single channel
currents at any B are
|
(1)
|
Extrapolation
of the iBNa to zero blocker
concentration provided the single channel currents in the absence of
blocker (iNa)
before any autoregulatory changes that could have influenced their
values. Blocker-sensitive open channel density in the absence of
blocker is No = INa/iNa
expressed in units of open channels per planar square centimeter or per
100 µm2, where the latter
approximates the area per cell.
Changes of No
arise from changes of
Po and/or
NT, where
No = PoNT.
If the blocker interacts only with the open state of ENaCs (10),
Po
can be estimated from the fractional blocker-dependent decreases of
NBo according to
|
(2)
|
Because
CDPC interacts rapidly with open channels, channels redistribute among
open, blocked, and closed states with a time constant limited by the
very slow spontaneous gating kinetics of the channels (
~1-2
s with mean open and closed times of several seconds). At the
equilibrium redistribution of channels, the fractional inhibition of
open channel density and thus Na+
entry into the cells is dependent on the
Po.
Extrapolation of the apparent values of
Po
at the various B values to zero B circumvents the
autoregulatory increases of
NBo that lead to
underestimates of the
Po at any
B (10; see RESULTS). As
will be evident in
RESULTS, the
Po
values of blocker-sensitive channels estimated by noise analysis as
indicated above are virtually the same as those of the long open time
channels measured by patch clamp of A6 epithelia, underscoring the
validity of the assumption that blocker interacts only with the open
state of the channels in A6 epithelia as in those of frog skin.
The channel densities determined this way are a measure of the pool of
apical membrane channels directly involved in
Na+ transport. Hence,
INa = iNaPoNT,
where the functional
NT = No/Po. Values are means ± SE.
 |
RESULTS |
Blocker-sensitive and blocker-insensitive
Isc.
Compared with previous published reports in studies of frog skin (6, 7,
10), A6 epithelia behaved in identical ways in response to CDPC
inhibition of apical membrane Na+
channels, except the control baseline rates of transport were generally
considerably less than those in native tissues (e.g., frog skin, toad
urinary bladder, cortical collecting duct, and others). As indicated in
Fig. 1, the strip chart
recordings of the changes of
Isc
in response to staircase increases of blocker concentration showed the
typical scalloped appearance, indicative of autoregulatory increases of
channel density (1, 10), and the typical overshoot of the
Isc
after complete washout of CDPC from the apical solution. From peak
values, the currents relaxed toward the original control
Isc
within ~10 min. Addition of 100 µM amiloride to the apical solution
caused marked inhibition of the
Isc,
but not to zero, leaving an
IAmilsc. Removal of all
Na+ from the Ringer solution
perfusing the tissues (group A) caused the
IAmilsc to fall to zero,
indicating that the IAmilsc
was due to highly selective
Na+-conductive but
blocker-insensitive channels.

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Fig. 1.
Unmodified strip chart recording of changes of short-circuit current
(Isc) caused by
step increases of 6-chloro-3,5-diaminopyrazine-2-carboxamide (CDPC)
added to apical solution of a control A6 epithelium. Removal of CDPC
causes an overshoot of
Isc
that relaxes back to control values within ~10 min. Amiloride (100 µM) caused marked inhibition of
Isc, but not to
zero, leaving an amiloride-insensitive current.
|
|
A summary of Isc
and IAmilsc in all groups of
experiments is presented in Table 1. In
general, the IAmilsc represented a relatively small fraction of the
Isc
(1.2-7.4%) in the control and steroid-stimulated states of
transport. Isc were maximally stimulated about threefold in all groups of experiments, regardless of their baseline rates of transport, which ranged from
2.33-7.36 µA/cm2 to as high
as 20.41 µA/cm2 in stimulated
tissues. Subtraction of the
IAmilsc from
Isc
yielded the blocker-sensitive Na+
currents INa in
the absence of blocker and
IBNa at any blocker
concentration.
Single channel currents and open channel densities.
Despite the low baseline values of transport (compared with native
tissues like frog skin and toad urinary bladder), noise analysis was
possible at
Isc
somewhat less than 1 µA/cm2 at
B
5 µM CDPC. As indicated in Fig.
2, Lorentzians were easily resolved at
5-50 µM CDPC. For all groups of experiments and as indicated for
group A experiments summarized in Fig.
3, the
fc varied linearly with B in both control and steroid-treated
tissues. S0
and IBNa of control and
steroid-treated tissues changed in accordance with a kinetic scheme of
closed
open
blocked states, where the blocker interacts only
with the open state of the channel. As indicated in Fig. 3 for
group A tissues and summarized in
Table 1 for all groups, the
iNa
were less in steroid-treated tissues than in controls due at least in
part to depolarization of apical membrane voltage in their stimulated
states of transport (3, 7).

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Fig. 2.
Current-noise power density spectra at 5 and 50 µM CDPC at low
(A) and intermediate
(B) rates of
Na+ transport. Data were fit by
nonlinear regression to 3 components, including a Lorentzian
{S0/[1 + (f/fc)2],
where S0 is
low-frequency plateau, f is frequency,
and fc is corner
frequency}, noise at low frequencies
(S1/f ),
and noise at higher frequencies
(S2f ),
originating at input stage of voltage amplifier. Spectra at intermediate concentrations are not shown.
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Fig. 3.
Blocker concentration dependence of blocker-sensitive macroscopic
currents (IBNa),
S0 and
fc of
Lorentzians, and single channel currents
(iBNa) in control and
steroid-treated tissues from group A.
Single channel currents in absence of blocker were determined by
extrapolation of iBNa to zero
blocker concentration.
|
|
Because single channel currents changed little compared with
INa, open channel
density, No,
calculated from the quotient
INa/iNa must increase with steroid stimulation of transport. Plotted in Fig.
4 are the individual values of
No against
INa for all
groups of experiments, with means of control and steroid-treated
tissues summarized in Table 1. In maximally stimulated tissues,
No is ~50
channels/100 µm2 or ~50
channels/cell. At 1 µA/cm2,
there can only be about two to three open channels per cell on average,
which explains the difficulty in finding channels by patch clamp in
steroid-depleted A6 epithelia. In this regard, noise analysis permitted
study of the blocker-sensitive channels without difficulty at this
extreme lower bound of transport.

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Fig. 4.
Relationship between blocker-sensitive open channel densities
(No) and
Isc in control
and steroid-treated tissues.
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Blocker on- and off-rates.
The kob and
kbo measured in
A6 epithelia, summarized in Table 2, were
virtually the same as those reported for
Na+ channels in frog skin.
Accordingly, the blocker site appears to be conserved between these
tissues. Both the access time to the blocker site and the residency
time at the site were essentially the same among all passages of cells
and conditions of growth, yielding
KB
that averaged ~30 µM. It was apparent that the channels recruited
by steroid possessed virtually the same kinetic interactions with CDPC.
Po and NT.
It is evident from the data summarized in Fig.
5 that steroid stimulation of transport
does not occur by increase of
Po.
Po averaged between ~0.25 and 0.35 in all groups of tissues and was not
changed by steroid treatment of the tissues. The mean values of
Po
measured here by noise analysis are virtually the same as those of the
long open time channel described by Kemendy et al. (12) in their
patch-clamp studies of A6 epithelia.

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Fig. 5.
Apparent open probabilities of blocker-sensitive channels
(PBo) are plotted against
blocker concentration and appear to decrease owing to autoregulatory
increases of channel density (10). Autoregulatory effect is manifest in
strip chart recordings as a secondary slow increase of
Isc, with
transient overshoot of
Isc
after removal of CDPC from apical solution. Extrapolation of apparent
values of open probability to zero blocker concentration provided
values of
Po
of blocker-sensitive channels before any effect of autoregulation on
channel density and
Po.
Po
was not changed by steroid. Similarity in values of apparent
PBo at various blocker
concentrations indicates similarity in autoregulatory increases of
channel density expressed as a fraction of
NT.
|
|
The density of functional channels undergoing spontaneous fluctuations
between closed and open states, NT, was
calculated from the quotient
No/Po and
summarized in Fig. 6. In all groups, steroid treatment resulted in increase of
NT
and no change of Po.
Variation of transport between ~2 and 20 µA/cm2 among all control and
steroid-treated tissues could be ascribed simply to increase of
NT,
as illustrated in Fig. 7.

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Fig. 6.
Summary of changes of blocker-sensitive
NT
and
Po
caused by steroid treatment of A6 epithelia in groups
A, B, C1, and C2.
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Fig. 7.
Relationship between blocker-sensitive
Na+ transport and
NT. Values are
means ± SE from all groups of experiments (A, B,
C1, and C2) of
control and steroid-treated tissues.
|
|
 |
DISCUSSION |
Regardless of baseline rates of
Na+ transport, age and origin of
the cells, substrates on which the cells were grown, growth media, and
serum, our results demonstrate that the steroid hormones aldosterone
and corticosterone increase Na+
transport by increase of the density of functional channels at the
apical membranes of the cells. The channels are the same, as judged by
the similarity in the kinetics of blocker interaction of the channels
and by the magnitudes of the single channel currents that are in the
same range as those measured by patch clamp and those expected for
single channel conductance near 5 pS (3). In comparison with frog skin,
the channels appear to be identical, possessing extremely high
selectivity for Na+ where the
predominant population of channels is amiloride inhibitable. Because
the Po is not
changed by aldosterone or corticosterone, stimulation of transport must
be due to increase in the apical membrane pool size of functional
channels involved in Na+
transport. It is widely appreciated that channels may be recruited from
pools of nonfunctional channels present in the apical membranes and/or trafficked to the apical membrane from the cytosol. Our experiments do not address this issue or the mechanism whereby steroids
increase
NT.
Using identical methods of blocker-induced noise analysis, Granitzer et
al. (8) reported that the glucocorticoid dexamethasone stimulates
NT
in A6 epithelia with no change of Po. Pácha
et al. (14) reported that
Po
was not influenced by mineralocorticoid status in the rat cortical
collecting duct; in contrast, Kemendy et al. (12) reported that
aldosterone shifted channels from a short to a long open time state in
A6 cells. Accordingly, regardless of the origin of the channels,
Na+ transport must be carried out
principally by blocker-sensitive channels with long open and closed
times and by channels in which Po
is the same at various states of mineralocorticoid and glucocorticoid status.
Blocker-sensitive and blocker-insensitive
Isc.
Since discovery of amiloride as a potent inhibitor of epithelial
Na+ transport, it has been widely
acknowledged that this diuretic inhibits most, but not all, of the
Isc in target
tissues like frog skins and toad urinary bladders. The same appears to
be true for A6 epithelia in steroid-depleted and steroid-stimulated A6 epithelia. Regardless of the permeabilized substrate on which the cells
are grown (11) and as indicated in Table 1, the amiloride- or
blocker-insensitive currents 1)
represent a rather small fraction of the macroscopic rates of transport
into the cells as measured in absolute and relative terms and
2) are stimulated by steroid hormone. Because the IAmilsc
decreases to zero after removal of all
Na+ from the apical solution, the
apical membrane must contain channels that are conductive to
Na+ but that are not blocked by
amiloride at concentrations
100 µM that far exceed by several
orders of magnitude the KAmilB of blocker-sensitive channels. Accordingly, apical membranes must possess blocker-sensitive and blocker-insensitive pools of channels and
corresponding blocker-sensitive and blocker-insensitive
Isc. Because
blocker-induced noise analysis measures only those channels that are
blocker sensitive and because blocker-insensitive
Na+ currents represent a very
small fraction of Na+ transport,
increases of transport must be attributable to increases of
blocker-sensitive
NT.
Nevertheless, blocker-insensitive currents are also increased by
steroids but represent a comparatively small fraction of
Na+ transport in the presence and
absence of exogenous steroids. The gating kinetics and single channel
conductance of blocker-insensitive Na+ channels are unknown.
In this regard, Kemendy et al. (12) observed by patch clamp of A6
epithelia two populations of Na+
channels distinguished by large differences in their mean open times
but otherwise exhibiting the same single channel currents and, hence,
single channel conductance. Short mean open time channels (
o ~ 40 ms) coexisted with
mean long open time channels (
o ~ 1,600 ms) in the same patches. It is unknown whether the short open
time channels observed by patch clamp are blocker sensitive or
insensitive. Thus the role and function of short open time channels
remain unknown but could perhaps be related to blocker-insensitive Na+ channels. If the densities of
short and long open time channels are in the same range, then the
contribution of short open time channels to the macroscopic rates of
Na+ transport would be in the
range of a few percent of the
Isc. With
identical channel densities and closed times and with
o of 40 and 1,600 ms, the
contribution of short open time channels to the
Isc is 2.5%.
Origin of channels.
It is of particular interest to know the origin of the long open time
channels, the density of which is increased by steroids. The hypothesis
by Kemendy et al. (12) that steroids change open probability of
preexisting short open time channels into long open time channels by an
all-or-none mechanism rests on critical observations. First, short open
time channels must exist before exposure of the tissues to steroids.
Second, assuming that the density of short open time channels is
constant and that steroids stimulate transport by an all-or-none
increase of the mean open time of these channels, decreases of short
open time channels must be accompanied by identical increases of long
open time channels if this is the only source of channels.
It is appreciated that interpretation of patch-clamp data is
exacerbated, especially for the particular case where channel open
times are small compared with their closed times (4), as is the case
for the short open time channels. With 40-ms mean open time and
3,000-ms mean closed time of the short open time channels, there would,
at a 95% level of confidence, have to be at least 13 channels in a
patch to observe a double opening of this channel within 20 min of
continuous recording (4, 12). The probability of observing multiple
openings of these channels is even more remote, so estimates of short
open time channel number in a patch are practically impossible.
Accordingly, it would be impossible practically to know whether
steroids cause changes of short open time channel densities and, hence,
whether long open time channels originated from short open time
channels as has been suggested (12).
The source of the apical pool of functional channels is unknown and
remains a topic of particular interest. Our own experiments reported
here shed no light on this issue. Our experiments do not rule out
sources of nonfunctional channels within the apical membrane or sources
originating from channel-containing intracellular vesicles. Aldosterone
is known to induce a variety of proteins, among which may be channel
subunits and other proteins involved in sorting and trafficking the
channels to the apical membranes. Subunits are expressed in
steroid-depleted tissues, and aldosterone does not change the levels of
subunit RNA in the same way in all tissues, if at all (13).
cDNA-injected oocytes express long open time channels (18) with no
reports of short open time channels like those observed in A6 (12).
Because apical membrane capacitance is increased by aldosterone in
tissues treated overnight (13a) and in a time-dependent way during
exposure to aldosterone for 6 h (11a), it is possible that steroids
stimulate transport by trafficking of channels to the plasma membrane.
Accordingly, steroids stimulate
Na+ transport by increase of
apical membrane
NT with no change
of Po. The
question of origin of the channels remains open.
 |
ACKNOWLEDGEMENTS |
We thank A. L. Helman for excellence in maintaining our tissue
culture facility (Urbana, IL), the care and feeding of the cells, and
assistance in the preparation of the manuscript.
 |
FOOTNOTES |
This study was supported by National Institutes of Health Grants
DK-16663 and DK-30824 (to S. I. Helman) and RR-05491 (to R. L. Duncan)
and grants from the South African Medical Research Council and the
National Kidney Foundation of South Africa (to W. J. Els).
L. M. Baxendale-Cox was an American Heart Association (Illinois
Affiliate) Fellow. X. Liu is a doctoral student in the Dept. of
Molecular and Integrative Physiology (Urbana). K. Baldwin is a
master's degree student in the Dept. of Anatomy and Cell Biology (Cape
Town, South Africa).
Present addresses: L. M. Baxendale-Cox, School of Nursing, Johns
Hopkins University, Baltimore, MD 21205; R. L. Duncan, Dept. of
Orthopaedic Surgery, Indiana University School of Medicine, Indianapolis, IN 46202.
Address for reprint requests: S. I. Helman, Dept. of Molecular and
Integrative Physiology, 524 Burrill Hall, 407 South Goodwin Ave.,
University of Illinois at Urbana-Champaign, Urbana, IL 61801.
Received 21 February 1997; accepted in final form 8 July 1997.
 |
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