Cytoskeletal modulation of sodium current in human jejunal
circular smooth muscle cells
Peter R.
Strege1,2,
Adrian N.
Holm1,2,
Adam
Rich1,2,
Steven M.
Miller1,2,
Yijun
Ou1,2,
Michael G.
Sarr3, and
Gianrico
Farrugia1,2
1 Department of Physiology and Biophysics,
2 Enteric NeuroScience Program and Division of
Gastroenterology and Hepatology, and 3 Department of
Surgery, Mayo Clinic and Mayo Foundation, Rochester, Minnesota
55905
 |
ABSTRACT |
A
Na+ current is present in human jejunal circular smooth
muscle cells. The aim of the present study was to determine the role of
the cytoskeleton in the regulation of the Na+ current.
Whole cell currents were recorded by using standard patch-clamp
techniques with Cs+ in the pipette to block K+
currents. Cytochalasin D and gelsolin were used to disrupt the actin
cytoskeleton and phalloidin to stabilize it. Colchicine was used to
disassemble the microtubule cytoskeleton (and intermediate filaments)
and paclitaxel to stabilize it. Acrylamide was used to disrupt the
intermediate filament cytoskeleton. Perfusion of the recording chamber
at 10 ml/min increased peak Na+ current recorded from
jejunal smooth muscle cells by 27 ± 3%. Cytochalasin D and
gelsolin abolished the perfusion-induced increase in Na+
current, whereas incubation with phalloidin, colchicine, paclitaxel, or
acrylamide had no effect. In conclusion, the Na+ current
expressed in human jejunal circular smooth muscle cells appears to be
regulated by the cytoskeleton. An intact actin cytoskeleton is required
for perfusion-induced activation of the Na+ current.
small intestine; patch clamp; actin; microtubules
 |
INTRODUCTION |
OPENING OF
Na+ channels provides for a large change in membrane
permeability for Na+, resulting in entry of Na+
into the cell with subsequent membrane depolarization (12, 16). The amount of Na+ entering the cell, and hence
the degree of depolarization, is regulated by mechanisms and pathways
that determine channel gating. In epithelial, skeletal, and cardiac
Na+ channels, channel gating appears to be altered by the
cytoskeleton (5, 11, 15, 23), although the exact
mechanisms are still controversial (17). The cytoskeleton
is made up of three major components: microfilaments, of which actin is
the major component (size
60 Å), microtubules (size
150-250 Å), and intermediate filaments, which, as indicated by
their nomenclature, are thicker than microfilaments but thinner than
microtubules (2-4, 22, 24).
Patch-clamp experiments on dissociated human jejunal circular smooth
muscle cells show that a tetrodotoxin (TTX)-resistant Na+
current is present, and the activation-inactivation kinetics suggest
steady-state Na+ entry at physiological voltages
(13). The channel shows close homology with the
TTX-resistant cardiac Na+ channel NaH1/NaSkM2
(13), and electrophysiological and molecular biology
evidence (13) suggests that both tissues express the same
Na+ channel. The molecular sequence of the pore-forming
subunit (SCN5A) of NaH1/NaSkM2 and human jejunal SCN5A contains the
consensus sequence (S/T)XV-COOH, which binds to PDZ domains found on
the cytoskeletal component syntrophin (11, 20). The aim of
this study was to examine the effects of cytoskeletal modulation on the
human jejunal circular smooth muscle Na+ current. We found
that Na+ current is modified by disruption of the actin
cytoskeleton by cytochalasin D and gelsolin, suggesting a link between
the channel and the smooth muscle cytoskeleton.
 |
METHODS |
The use of human jejunum, obtained as surgical waste tissue
during gastric bypass operations performed for morbid obesity, was
approved by the Institutional Review Board. Tissue specimens with warm
ischemia times of ~30 s were harvested directly into chilled
buffer. Single isolated, relaxed circular smooth muscle cells were
obtained from the human jejunal specimens as described previously
(8, 9).
Patch-clamp recordings.
Whole cell patch-clamp recordings were made with standard and
amphotericin-perforated patch-clamp whole cell techniques. Whole cell
recordings were obtained with Kimble KG-12 glass pulled on a P-97
puller (Sutter Instruments, Novato, CA). Electrodes were coated with
R6101 (Dow Corning, Midland, MI) and fire polished to a final
resistance of 3-5 M
. Currents were amplified, digitized, and
processed with an Axopatch 200A amplifier, a Digidata 1200, and pCLAMP8
software (Axon Instruments, Foster City, CA). Whole cell records were
sampled at 10 kHz and filtered at 4 kHz with an eight-pole Bessel
filter with the pulse protocols shown in Figs. 1, and 3-8.
Seventy to seventy-five percent series resistance compensation (lag of
10 µs) was applied during each recording. The cell capacitance
(Cm) was 40-70 pF, and the access
resistance (Ra) was 5-10 M
. All records
were obtained at room temperature (22°C).

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Fig. 1.
Perfusion increases peak Na+ current. A:
representative Na+ current recordings obtained from a human
jejunal circular smooth muscle cell at a holding voltage of 100 mV
with the pulse protocol shown in the inset. Recordings were
taken in a still bath with Ringer solutions and then during perfusion
with the Ringer solution at 10 ml/min. B: normalized
current-voltage relationships before and after perfusion. C:
% increase in maximal peak Na+ current evoked by perfusion
(*P < 0.01).
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Drugs and solutions.
The pipette solution contained (in mM) 145 Cs+, 20 Cl
, 2 EGTA, 5 HEPES, and 130 methanesulfonate. The bath
solution contained normal Ringer solution (in mM: 149.2 Na+, 4.74 K+, 156.5 Cl
, and 2.54 Ca2+) buffered with 5 mM HEPES and nifedipine (1 µM)
unless indicated otherwise. Drugs were purchased from Sigma (St. Louis,
MO). Nifedipine stock solution was made up in 100% ethanol. The final
dilution of alcohol applied was <1:1,000. At this concentration,
preliminary experiments showed that ethanol had no effect on currents
recorded (data not shown).
Cytochalasin D and gelsolin were used to depolymerize the actin
cytoskeleton and phalloidin to stabilize it (2, 14, 21). Colchicine was used to disassemble the microtubule cytoskeleton (4) and paclitaxel to stabilize it (4).
Collapse of the microtubule cytoskeleton is associated with concomitant
collapse of the intermediate filament cytoskeleton. Acrylamide was used to disrupt the intermediate filament cytoskeleton (3). The optimal dose and specificity of each agent were determined by staining
for each component of the cytoskeleton in the presence and absence of
each drug. Mouse anti-
-smooth muscle actin (Sigma A2547) primary
antibodies were used to stain the actin (microfilament) cytoskeleton,
and rabbit anti-desmin (Sigma D8281) antibodies were used to stain the
intermediate filament cytoskeleton. The primary antibodies were
visualized with affinity-purified secondary antibodies conjugated to
rhodamine or CY3 (Chemicon, Temecula, CA). Primary antibodies were
diluted 1:100 in 0.1 M PBS containing 5% normal donkey serum and 0.3%
Triton X-100. Secondary antibodies were diluted 1:100 in 0.1 M PBS
containing 2.5% normal donkey serum and 0.3% Triton X-100. Cells were
examined with a laser scanning confocal microscope (Zeiss LSM 510). On
the basis of these preliminary experiments, cells were incubated with
the respective drugs for 10-15 min, except for acrylamide, which
was incubated for 30 min. The following concentrations were used:
cytochalasin D, 5 µg/ml; gelsolin, 1 µM; phalloidin, 25 µM;
colchicine, 10 µM; paclitaxel, 25 µM; and acrylamide, 5 mM.
Drugs were applied by complete bath changes with the solution
containing the drug, except for gelsolin, which is not membrane permeant. The extent of membrane permeability of phalloidin is uncertain. Preliminary experiments were performed with cells incubated with tetramethylrhodamine isothiocyanate (TRITC)-labeled phalloidin. These experiments showed that phalloidin can cross the intact cell
membrane. Experiments using phalloidin were carried out with both
intracellular (delivered via pipette) and extracellular (in separate
experiments) phalloidin. No difference was noted between the two sets
of experiments; therefore, the data were combined. Gelsolin was also
placed in the recording pipette, and the data obtained with gelsolin
and phalloidin were compared with separate controls. For drugs applied
to the bath, comparisons were made with the peak control
Na+ current before application of the drug, in which every
cell acted as its own control. Records were obtained in the presence of
nifedipine (1 µM) to block Ca2+ currents, except for the
acrylamide experiments because the incubation time was considerably
longer for this drug and we did not want to expose the cells to
nifedipine for prolonged periods.
Perfusion was used to mechanically perturb the cell membrane. Perfusion
and positive pressure in the whole cell mode and negative pressure in
the on-cell mode have been used to mechanoactivate ion channels
(7). Of the three methods, perfusion was chosen in this
study because it may most closely mimic the effects of movement of the
extracellular matrix and adjacent smooth muscle on ion channels present
on the cell surface and because of the marked repeatability of its
effects (7).
Data analysis.
Electrophysiological data were analyzed with pCLAMP8 software or custom
macros in Excel (Microsoft, Redmond, WA). Voltages were adjusted for
the junction potential. Paired Student's t-test or ANOVA
with Tukey correction was used to evaluate statistical significance. A
P value of <0.05 was considered significant. Values in the
text are presented as mean ± SE maximal peak inward currents. The
mean percent changes in current reported reflect the mean of the
percent increase in current for each experiment. Data in Figs. 1 and
3-8 are presented as normalized values with the preperfusion current normalized to 100% for the bar graphs and the current-voltage relationships normalized to the maximal inward current of the control
current set at 1.
 |
RESULTS |
Activation of Na+ current by
perfusion.
Initial experiments showed that the Na+ current increased
in size when the bath was perfused with Ringer solution, suggesting that the jejunal circular smooth muscle channel was shear stress sensitive. Therefore, the recording chamber was perfused for 30 s
at 10 ml/min to mechanically perturb the cellular surface. Perfusion increased peak Na+ current in human jejunal circular smooth
muscle cells by 27 ± 3% (147 ± 21 to 175 ± 23 pA,
n = 41, P < 0.01; Fig.
1). Thirty-nine of the forty-one cells
studied showed an increase in current with perfusion. Time to peak
inward Na+ current (activation) was faster at voltages
ranging from
50 to
2 mV (n = 13, P < 0.05). Inactivation, measured as a single tau, of the currents after
perfusion was not different from before perfusion (n = 6; Fig. 2). Activation kinetics of the
difference current, that is, the current selectively activated by
perfusion, were faster at all voltages measured, as was inactivation of
the difference current, again measured as a single tau (Fig. 2). The increase in Na+ current with perfusion was used to evaluate
the effect of the cytoskeleton-altering drugs. In cells perfused with
N-methyl-D-glucamine (NMDG) in the bath to
replace Na+, no effect of perfusion was seen at all
voltages tested (
90 to +30 mV, n = 3), suggesting
that the effects of perfusion were on a Na+ current.

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Fig. 2.
Effect of perfusion on activation and inactivation. Activation
(time to peak; A) and inactivation (single tau;
B) for the Na+ current before and after
perfusion and for the difference current are shown. Time to peak was
faster after perfusion at voltages ranging from 50 to 2 mV.
Difference current kinetics were faster at all voltages tested
(*P < 0.05).
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Actin cytoskeleton-altering drugs.
Cytochalasin D was used to disassemble the actin cytoskeleton. Cells
were patch clamped, and nifedipine (1 µM) was added to the bath to
block Ca2+ current. After a 5-min equilibrium period, cells
were perfused for 30 s with NaCl Ringer solution alone.
Cytochalasin D (10 µM) and nifedipine (1 µM) were then added to the
bath, and after a 15-min incubation period the bath perfused for
30 s. The mean maximal peak inward Na+ current was
158 ± 45 pA before and 163 ± 44 pA (n = 6)
after incubation with cytochalasin D (P > 0.05). The
perfusion-induced increase of Na+ current was markedly
reduced from 21 ± 3% (P < 0.01, n = 6) before cytochalasin D to 6 ± 2%
(P > 0.05, n = 6) after incubation
with cytochalasin D (Fig. 3). Control
experiments with two perfusions of Ringer solution 15 min apart did not
show any decrease in the perfusion-induced increase in Na+
current between the first and second perfusion (data not shown).

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Fig. 3.
Cytochalasin D inhibits the perfusion-induced increase in peak
Na+ current. A and B: representative
Na+ current recordings obtained with the pulse protocol in
the inset. A: recordings were taken of a cell
incubated in a solution of NaCl Ringer with 1 µM nifedipine, and then
the cell was perfused at 10 ml/min with NaCl Ringer solution.
B: recordings from the same cell after a 15-min incubation
in NaCl Ringer solution containing 10 µM cytochalasin D and 1 µM
nifedipine and then perfusion with Ringer solution at 10 ml/min.
C: normalized current voltage-relationships. D:
mean % increase in maximal peak Na+ current evoked by
perfusion (*P < 0.01). Cytochalasin inhibited the
perfusion-induced increase in Na+ current.
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The actin-severing protein gelsolin was used to further investigate the
effects of altering the actin cytoskeleton on the perfusion-induced
increase in Na+ current. Intracellular application of
gelsolin (1 µM) markedly decreased the perfusion-induced increase in
Na+ current from 27 ± 3% in control cells to 5 ± 2% (P > 0.05, n = 7) in the
presence of gelsolin (Fig. 4).

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Fig. 4.
Intracellular gelsolin inhibits perfusion-induced increase in
Na+ current. A and B: representative
Na+ current recordings with the pulse protocol in the
inset. A: currents obtained from a cell
patch-clamped with 1 µM gelsolin in the intracellular solution and
NaCl Ringer solution in the bath. B: recording from the same
cell perfused at 10 ml/min with NaCl Ringer solution 15 min after
break-in. C: normalized current-voltage relationships.
D: mean % increase in maximal peak Na+ current
evoked by perfusion. Perfusion did not induce an increase in maximal
peak Na+ currents in the presence of gelsolin.
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Phalloidin was used to stabilize the actin cytoskeleton. Phalloidin had
no effect on the maximal peak Na+ current (160 ± 46 pA before and 161 ± 46 pA after incubation with phalloidin,
n = 7, P > 0.05). In contrast to
cytochalasin D, phalloidin did not affect the perfusion-induced
increase in Na+ current. The perfusion-induced increase in
Na+ current in phalloidin-exposed cells was 25 ± 3%
(P < 0.01, n = 7, Fig.
5) before and 21 ± 3%
(P < 0.01, n = 7) after incubation with phalloidin in same-cell controls (P > 0.05 between the 2 perfusions).

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Fig. 5.
Phalloidin does not alter perfusion-induced increase in maximal
peak Na+ current. A and B:
representative Na+ current recordings with the pulse
protocol in the inset. A: control recordings of a
cell in NaCl Ringer solution subsequently perfused with the same
solution at 10 ml/min. B: recordings of the same cell
incubated for 15 min in Ringer solution with 25 µM phalloidin and
then perfused at 10 ml/min with NaCl Ringer solution alone.
C: normalized current-voltage relationships. D:
mean % increase in maximal peak Na+ current evoked by
perfusion (*P < 0.01). Phalloidin incubation did not
alter the perfusion-induced increase in Na+ current.
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Microtubular cytoskeleton-altering drugs.
Colchicine was used to disassemble the microtubule (and intermediate
filament) cytoskeleton and paclitaxel to stabilize it. Both colchicine
and paclitaxel had no effect on either peak maximal Na+
current or perfusion-induced increase in Na+ current. After
a 15-min incubation with colchicine (10 or 100 µM), maximal peak
current was 207 ± 72 pA compared with 191 ± 67 pA
(n = 10, P > 0.05) before addition of
the drug. The perfusion-induced increase in Na+ current was
33 ± 8% (P < 0.05, n = 10)
before colchicine and 18 ± 2% (P < 0.05, n = 10) after incubation with colchicine (Fig. 6; P > 0.05 between the
2 perfusions). Similarly, after a 15-min incubation with paclitaxel (25 µM) maximal peak current was 142 ± 25 pA compared with 150 ± 26 pA (n = 6, P > 0.05) before
addition of the drug. The perfusion-induced increase in Na+
current was 21 ± 3% (P < 0.01, n = 6) before paclitaxel and 17 ± 4%
(P < 0.05, n = 6) after incubation
with paclitaxel (Fig. 7;
P > 0.05 between the 2 perfusions).

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Fig. 6.
Colchicine does not inhibit the perfusion-induced increase in
Na+ current. A and B: representative
Na+ current recordings obtained with the pulse protocol in
the inset. A: control recordings of a cell in
NaCl Ringer solution that was then perfused with the same solution at
10 ml/min. B: recordings of the same cell incubated for 15 min in NaCl Ringer solution with 10 µM colchicine and then perfused
at 10 ml/min. C: normalized current-voltage relationships.
D: mean % increase in maximal peak Na+ current
evoked by perfusion (*P < 0.05).
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Fig. 7.
Paclitaxel does not inhibit the perfusion-induced increase in
Na+ current. A and B: representative
Na+ current recordings with the pulse protocol in the
inset. A: control recordings from a cell in NaCl
Ringer solution subsequently perfused with the same solution at 10 ml/min. B: recordings of the same cell incubated for 15 min
in NaCl Ringer solution with 25 µM paclitaxel and then perfused at 10 ml/min with NaCl Ringer solution alone. C: normalized
current-voltage relationships. D: mean % increase in
maximal peak Na+ current evoked by perfusion
(*P < 0.05).
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Intermediate filament cytoskeleton-altering drugs.
Acrylamide was used to disrupt the intermediate filament cytoskeleton.
Human jejunal circular smooth muscle cells were incubated with
acrylamide for 30 min. Because of the length of incubation, nifedipine
was not added to the bath solution and same-cell control perfusions
were not obtained. The perfusion-induced increase in Na+
current was not affected by incubation with acrylamide. After incubation with acrylamide, perfusion induced a 38 ± 9%
(P < 0.01, n = 8; Fig.
8) increase in maximal current compared
with 27 ± 3% in control cells (P > 0.05 between
acrylamide and controls).

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Fig. 8.
Acrylamide does not alter the perfusion-induced increase in maximal
peak Na+ current. A and B:
representative Na+ current recordings obtained with the
pulse protocol in the inset. Note the slower
Ca2+ current present in these cells because nifedipine was
not added to the bath. A: recordings from a cell incubated
in NaCl Ringer solution with 5 mM acrylamide for 30 min. B:
recording from the same cell perfused at 10 ml/min with NaCl Ringer
solution. C: normalized current-voltage relationships.
D: mean % increase in maximal peak Na+ current
evoked by perfusion (*P < 0.01).
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The L-type Ca2+ channel current present in human jejunal
circular smooth muscle cells is also activated by perfusion.
Experiments similar to those described above were carried out on the
L-type Ca2+ current. None of the cytoskeleton-modifying
agents used had any effect on the perfusion-induced increase in
Ca2+ current (data not shown).
 |
DISCUSSION |
The main finding of the present study was that the actin
cytoskeleton appears to modulate the increase in Na+
current in human jejunal circular smooth muscle cells evoked by
perturbation of the cell membrane. Baseline unstimulated
Na+ current was not altered by agents altering the
cytoskeletal structure, suggesting that gating of unstimulated
Na+ channels mediating the inward Na+ current
is not dependent on the cytoskeleton but activation of jejunal circular
smooth muscle Na+ channels by membrane perturbation is. The
luminal diameter of the gastrointestinal tract is constantly changing
as a result of digestive and interdigestive contractile activity and
secondary to the passage of food boluses. The gastrointestinal smooth
muscle cell membrane is subsequently under a constantly varying amount of shear stress as it is sandwiched between the extracellular matrix
and the rigid cytoskeleton. Regulation of Na+ entry by
shear stress may alter the contractile activity of intestinal smooth
muscle. Block of the intestinal smooth muscle Na+ channel
results in membrane hyperpolarization (13), and an increase in Na+ entry in response to increased shear stress
would be expected to depolarize intestinal smooth muscle, bringing the
membrane potential closer to the contractile threshold. The actin
cytoskeleton may serve as a mechanism to transmit force to the
Na+ channel by constraining the lipid bilayer or by
directly interacting with the Na+ channel.
Cytoskeletal modifiers have been shown to alter Na+
channels in cardiac, skeletal, and epithelial tissues (5, 11, 15, 23). Cytochalasin D, a disrupter of the actin cytoskeleton, is
most commonly linked to modulation of Na+ current, but
microtubular cytoskeletal modifiers such as colchicine and paclitaxel
have also been shown to modulate Na+ current
(19).
In skeletal muscle, voltage-gated Na+ channels are
concentrated at postsynaptic membrane sites (6). A family
of intracellular membrane-associated proteins called syntrophins
(1, 2, 10, 18) are partly responsible for the aggregation
of Na+ channels. Syntrophins associate with dystrophin,
which in turn is linked to the cell membrane and extracellular matrix
through actin and dystroglycans (2, 20). Thus syntrophin
links signaling proteins to the actin cytoskeleton and the
extracellular matrix. Syntrophins have a PDZ domain that binds to the
consensus sequence (S/T)XV-COOH (11, 20). Previous work
showed that the molecular sequence of the
-subunit of the
TTX-resistant Na+ channel (SCN5A) in human jejunal circular
smooth muscle (13) and cardiac muscle is similar (GenBank
accession numbers AY038064 and NM000335). The
electrophysiological and pharmacological properties of native
TTX-resistant Na+ channels in human jejunal circular smooth
muscle and cardiac muscle are also similar (13). Both the
human jejunal circular smooth muscle and the cardiac TTX-resistant
Na+ channel
-subunit contain the sequence DRESIV, which
binds strongly to syntrophins (11, 20), suggesting
syntrophins as a possible link between the Na+ channel and
the actin cytoskeleton. Human jejunal circular smooth muscle cells also
express a mechanosensitive L-type Ca2+ channel that is
activated by internal changes in pressure and by perturbation of the
cell membrane by shear forces (7). All cloned
Ca2+ channel subunits to date do not have a PDZ binding
domain. In this context, the lack of effect of agents altering the
cytoskeletal structure on the perfusion-induced increase in
Ca2+ current again suggests that the effects of agents that
modify the actin cytoskeleton on the Na+ current are
mediated through a specific disruption of the actin-syntrophin cytoskeleton link to Na+ channels rather than a nonspecific
effect on the cytoskeleton.
The results obtained in the present study suggest that the cytoskeleton
may not only serve to anchor Na+ channels to the membrane
but may also affect function. Recent work reported the effects of
cytoskeletal modulators on the
-subunit rSkM1 expressed in Cho cells
(17). The investigators reported minimal effects on peak
current, current-voltage relationships, and kinetic properties and
suggested that the cytoskeleton did not directly interfere with
Na+ channel function (17). Our data suggest
that such an interaction may not be apparent under unstimulated
conditions but may become apparent when the membrane and attached
cytoskeleton are deformed, as presumably occurs with perfusion.
In summary, human jejunal circular smooth muscle cells express a
Na+ channel that is activated when the cell membrane is
mechanically perturbed. The increase in Na+ channel current
evoked by perfusion was abolished by agents that disrupt the actin
cytoskeleton but not by other cytoskeleton-altering agents, suggesting
that the actin-related cytoskeleton complex may specifically alter the
regulation of Na+ channel current and subsequent
Na+ entry into the cell in response to membrane perturbation.
 |
ACKNOWLEDGEMENTS |
We thank Dr. Simon Gibbons for helpful discussions, Gary Stoltz for
technical assistance, and Kristy Zodrow for secretarial assistance.
 |
FOOTNOTES |
National Institute of Diabetes and Digestive and Kidney Diseases Grants
DK-52766, DK-57061, DK-17238, and DK-39337 supported this work.
Address for reprint requests and other correspondence:
G. Farrugia, Guggenheim 8, Enteric NeuroScience Program, Div. of
Gastroenterology and Hepatology, Mayo Clinic and Mayo Foundation, 200 First St., SW, Rochester, MN 55905 (E-mail:
farrugia.gianrico{at}mayo.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.
10.1152/ajpcell.00532.2001
Received 7 November 2001; accepted in final form 3 September 2002.
 |
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