ENaC- and CFTR-dependent ion and fluid transport in mammary
epithelia
Sasha
Blaug1,2,
Kevin
Hybiske1,
Jonathan
Cohn3,
Gary L.
Firestone1,
Terry E.
Machen1, and
Sheldon S.
Miller1,2
1 Department of Molecular and Cell Biology and
2 School of Optometry, University of California, Berkeley,
California 94720-3200; and 3 Departments of Medicine and Cell
Biology, Duke University Medical Center, Durham, North Carolina
27710-0001
 |
ABSTRACT |
Mammary epithelial 31EG4 cells (MEC) were grown as monolayers on
filters to analyze the apical membrane mechanisms that help mediate ion
and fluid transport across the epithelium. RT-PCR showed the presence
of cystic fibrosis transmembrane conductance regulator (CFTR) and
epithelial Na+ channel (ENaC) message, and immunomicroscopy
showed apical membrane staining for both proteins. CFTR was also
localized to the apical membrane of native human mammary duct
epithelium. In control conditions, mean values of transepithelial
potential (apical-side negative) and resistance
(RT) are
5.9 mV and 829
· cm2, respectively. The apical membrane
potential (VA) is
40.7 mV, and the mean ratio
of apical to basolateral membrane resistance (RA/RB) is 2.8. Apical
amiloride hyperpolarized VA by 19.7 mV and
tripled RA/RB. A
cAMP-elevating cocktail depolarized VA by 17.6 mV, decreased RA/RB by
60%, increased short-circuit current by 6 µA/cm2,
decreased RT by 155
· cm2, and largely eliminated responses to
amiloride. Whole cell patch-clamp measurements demonstrated
amiloride-inhibited Na+ currents [linear current-voltage
(I-V) relation] and forskolin-stimulated Cl
currents (linear I-V relation). A capacitance probe method
showed that in the control state, MEC monolayers either absorbed or
secreted fluid (2-4
µl · cm
2 · h
1). Fluid
secretion was stimulated either by activating CFTR (cAMP) or blocking
ENaC (amiloride). These data plus equivalent circuit analysis showed
that 1) fluid absorption across MEC is mediated by
Na+ transport via apical membrane ENaC, and fluid secretion
is mediated, in part, by Cl
transport via apical
CFTR; 2) in both cases, appropriate counterions move through
tight junctions to maintain electroneutrality; and 3)
interactions among CFTR, ENaC, and tight junctions allow MEC to either
absorb or secrete fluid and, in situ, may help control luminal
[Na+] and [Cl
].
amiloride; diphenylamine-2-carboxylate; milk secretion; patch
clamp; microelectrodes; electrophysiology; cystic fibrosis; tight
junctions; leaky and tight epithelia; epithelial sodium channel; cystic
fibrosis transmembrane conductance regulator
 |
INTRODUCTION |
THE MAMMARY GLAND is
a branched, convoluted tubular organ with acinar cells that secrete
macromolecules (milk proteins, lactose, fats), salts (including
Na+, K+, Ca2+, Cl
,
and phosphate), and water. Human milk contains ~200 mM lactose, 7 mM
Na+, 13 mM K+, 8 mM Ca2+, and 12 mM
Cl
and is mildly acidic (pH 6.8) (34). The
initial secretion of milk by the acinar cells probably occurs as an
isotonic fluid, largely driven by the production and then secretion of
lactose and osmotically obliged fluid. The duct cells likely modify
this fluid as it moves along the duct and passes out to the nipple. It
is possible that the relatively low ionic content of milk is either
generated or maintained by the reabsorptive properties of acinar and
duct cells, but the specific ion channels and other transporters
involved have not been described.
Abnormal fluid accumulation in the breast is common in premenopausal
women (5, 6, 21, 30). As part of a project to determine
how mammary cysts accumulate fluid, we began a study of the ion
transport properties of the mammary epithelial cell line 31EG4. This
untransformed mouse cell line appears to have properties of both acinar
and ductal mammary epithelia cells (39, 42, 48). Previous
experiments on primary cultures of mouse mammary epithelial cells,
which are a mixture of duct and acinar cells (4), showed
that Na+ is actively absorbed, that this absorption is
stimulated by the lactogenic hormone prolactin, and that absorption is
inhibited by apical amiloride, the well-known blocker of the epithelial Na+ channel (ENaC) (19). 31EG4 cells form
polarized monolayers with tight junctions that are regulated in their
"tightness" by the glucocorticoid dexamethasone, which can stop
growth and induce differentiation (48). This process
includes an increase in transepithelial resistance
(RT) from 100-300
· cm2 to >700-1,000
· cm2 and the appearance of tight junctional
proteins ZO-1 and more "organized" filamentous actin. It is thought
that this regulation of tight junctions recapitulates the
transformation of mammary cells from a leaky to a tight epithelium
characteristic of the lactating gland that must maintain a large
transepithelial (blood to milk) concentration gradient for ions and
macromolecules (29, 35). Coincident with the increase in
RT, 31EG4 cells also express Na+/H+ exchange and
Na+-HCO
cotransport activity in the
basolateral, but not the apical, membrane (41).
The goal of the present work was to determine whether the epithelial
Cl
and Na+ channels [cystic fibrosis
transmembrane conductance regulator (CFTR) and ENaC, respectively]
contribute to the ion and fluid transport properties of the mammary
epithelial cell line 31EG4. We reasoned that 31EG4 cells might also
express CFTR and ENaC, since 1) the mammary gland and sweat
glands are of similar embryological origin and have several
characteristics in common, and 2) sweat duct cells express
high levels of both these ion channels to allow reabsorption of salt
from the fluid that flows down the duct (38). We used
RT-PCR to identify the message and Western blot and immunomicroscopy to
identify the proteins. In experiments utilizing transepithelial, microelectrode, and patch-clamp techniques, amiloride-inhibitable changes in voltage and resistance were used to localize ENaC to the
apical membrane, and stimulation with cAMP and inhibition by
diphenylamine-2-carboxylate (DPC) showed that CFTR is localized to the
apical membrane. The roles of ENaC and CFTR in fluid absorption and
secretion were determined by measuring amiloride- and cAMP-induced changes in fluid transport (capacitance probe method).
 |
MATERIALS AND METHODS |
Cell culture.
31EG4 cells were grown and cultured in DMEM/F-12 medium containing 5%
fetal bovine serum (FBS), 5 µg/ml insulin, and 5 µg/ml gentamicin
sulfate. Upon reaching confluency, these cells, passages 6-10, were plated onto Transwell filters (Costar) at a
density of 105 cells/well. They were grown in DMEM/F-12
medium containing glutamine, 2% FBS, insulin, and gentamicin as
described above. When the cells became confluent on the filters, 1 µM
dexamethasone was added to stop growth and induce differentiation,
including formation of tight junctions (48) and
polarization of ion transport pathways to the apical and basolateral
membranes (41). RT and
transepithelial membrane voltage potential (TEP) were estimated with an
EVOM (Epithelial Voltammeter; World Precision Instruments, New Haven
CT). The experiments were all carried out between 6 and 9 days after
the addition of dexamethasone with no discernible differences in mean
TEP or RT.
RT-PCR.
Total RNA was extracted from 31EG4 cells by using the RNAzol B method
(Teltest) per manufacturer's instructions. First-strand cDNA synthesis
was carried out by using 0.5 pg of total RNA, 20 pM oligo(dT) primers,
0.5 mM dNTP mix, and 200 units of Moloney murine leukemia virus reverse
transcriptase in a final volume of 20 ml of 50 mM Tris · HCl
(pH 8.3), 75 mM KCl, and 3 mM MgCl2. The RNA and oligo(dT)
were annealed by first mixing the two items, heating to 70°C for 2 min, and then cooling on ice, followed by the addition of the remaining
reaction components. The mixture was incubated at 42°C for 1 h
for first-strand synthesis and then heated to 94°C for 4 min to stop
the reaction. The mixture was then diluted 1:5 with sterile distilled
water. A control reaction containing no reverse transcriptase was
included for each tissue-reverse transcriptase reaction to assure that
no genomic DNA was being amplified (data not shown).
PCR was carried out by using oligonucleotide primers (sense
gtgattggagctatagcagttgtcg; antisense cccacatctggagcccacagc) designed to
cover a 463-bp region of CFTR and a 596-bp region of ENaC (sense tgcaactaccggaacttcacg; antisense gtactgggtgtcattgcagg). The primers (0.5 mM final concentration) were added to a 1-ml aliquot of the first-strand synthesis mixture with the following: 0.2 mM dNTPs (each),
1.25 units of Thermus aquaticus polymerase, PCR buffer with
Mg2+ (Boehringer Mannheim), and water to bring the mixture
to a total volume of 50 µl. The reaction was then overlaid with two
drops of mineral oil (Sigma, St. Louis, MO). The mixture was then
incubated in a thermal cycler (Stratagene) with the following
amplification profile: 1 cycle at 94°C for 4 min; 37 cycles at 94°C
for 1 min, 55°C for 1 min, and 72°C for 1 min; and 1 cycle of
72°C for 10 min. The PCR product was run on a 1.5% agarose gel. This
product was excised from the gel with a razor and then purified and
sequenced by the University of California at Berkeley Sequencing Center.
Western blots.
31EG4 cells were cultured to confluence on filters as described in
Cell culture. Proteins were isolated by placing the
cells into 100 µl of protein isolation buffer containing (in mM) 65 NaCl, 2 MgCl2, 1 EDTA, and 5 Tris-acetate, pH 7.4, as well
as the following protease inhibitors (in µg/ml): 2 aprotinin, 2 leupeptin, 1 pepstatin A, 2 antipan, 100 phenylmethylsulfonyl fluoride,
50 N
-p-tosyl-l-lysine
chloromethyl ketone, and 100 N-tosyl-l-phenylalanine chloromethyl
ketone. The cells were homogenized by sonicating (Branson,
Danbury, CT) on ice for 60 s. Protein concentration was determined
using BCA (bicinchoninic acid) protein assay kit (Pierce, Rockford,
IL). To separate protein from membranes, we incubated the sample for
4 h at room temperature (RT) in 4% digitonin (wt/vol) in 0.2 M
sodium phosphate buffer, pH 8.6. Electrophoresis of the samples was
carried out by using Bio-Rad Mini-Protein ready gels (7.5%
Tris · HCl) and the Bio-Rad Mini-Protean cell (Hercules, CA).
The running buffer contained 25 mM Tris, 200 mM glycine, and 0.1% SDS.
Electroblotting of the proteins onto polyvinylidene difluoride (PVDF)
membranes was carried out in a Bio-Rad Mini-Trans-Blot cell at 4°C.
The transfer buffer contained 25 mM Tris and 200 mM glycine, pH
8.2-8.5. The blotted PVDF membranes were then blocked in 2%
casein [in 0.1% (vol/vol) Tween 20 in PBS] for 1 h at RT and
then incubated with a 1:1,000 dilution of primary anti-CFTR antibody
overnight at 4°C (pAbECL1-19, peptides corresponding to amino acids
103-109, 109-114, 114-119 of CFTR; a generous gift from
Genzyme). Similar procedures were used with a primary anti-ENaC antibody (
-subunit, a generous gift from Pascal Barbry). The membranes were rinsed three times (5 min each time) with Blotto (5%
nonfat dried milk and 0.1% Tween 20 in PBS). The membrane was then
incubated in a 1:5,000 dilution of horseradish peroxidase-conjugated anti-rabbit secondary antibody (Sigma) for 1 h at RT, followed by
three rinses (5 min each time) with Blotto and then 5 min with PBS. The
labeled membranes were developed by using the enhanced chemiluminescence method (NEL-10 Renaissance; NEN).
Immunohistochemistry.
31EG4 cells were grown to confluency on Transwell filters, and the
monolayer and filter membranes were punched out, rinsed in PBS for 1 min at RT, and then fixed in 4% formaldehyde-PBS for 30 min. The
sample was then rinsed three times, over the course of 10 min, with PBS
at RT and placed in 30% sucrose-PBS solution to equilibrate overnight
at 4°C. The samples were next embedded in OCT (optimum cutting
temperature) compound (Tissue-Tek, Torrance, CA) and incubated at
20°C for at least 1 h. The samples were then sectioned in 10- to 20-µm slices with a cryostat (Leica), placed on Superfrost glass
slides (Fisher Scientific), and left overnight to dry at RT. Sections
were rinsed for 5 min in PBS and incubated in 10% BSA-PBS solution for
2 h at RT. The slides were then incubated overnight at 4°C in
1:100 (PBS/BSA) dilutions of either polyclonal anti-CFTR or anti-ENaC
(
-subunit). The sections were rinsed three times (10 min each time)
in PBS at RT and then incubated in a 1:2,000 dilution of FITC-labeled
secondary antibodies for 1 h at RT. The sections were finally
rinsed three times (10 min each time) in PBS and mounted on coverslips
with Prolong mounting solution (Molecular Probes, Eugene, OR). Images
were obtained with the use of a Zeiss Axiophot microscope with a ×63 objective.
Immunomicroscopy to identify CFTR in intact mammary gland was also
performed on cryosections (4 µm) of promptly frozen normal human
surgical specimens by using an affinity-purified rabbit antibody [a1468, a synthetic peptide corresponding to amino acids 1468-1480 of CFTR (11, 31)] at a 1:200 dilution. The
specificity of a1468 as a reagent for detecting endogenously expressed
CFTR in human colonocytes has been verified in a variety of tissues by
immunoblot, immunoprecipitation, and immunostaining (11-13, 18, 31). Sections were incubated first with a1468 and then with
a monoclonal antibody to cytokeratin 19 (ICN Biomedical, Costa Mesa,
CA). Sections were then washed and incubated with TRITC- and
FITC-conjugated goat F(ab')2 fragments (Tago, Burlingame, CA). Controls for immunostaining included sections stained without primary antibody or with normal rabbit serum.
Fluid transport.
Transepithelial fluid flows (JV) were measured
by using the capacitance probe technique (22, 24). A
monolayer on a filter (0.5-cm2 exposed area) was mounted
between two water-impermeable Kel F half-chambers.
JV was determined with a very sensitive
oscillator circuit (Acumeasure 1000; Mechanical Technology, Albany, NY)
connected to two probes, one on either side of the tissue, which
measure the capacitance between the probe tips and the fluid meniscuses connected to each half-chamber. Ion-linked fluid is driven across the
tissue from its apical to basolateral surfaces, or vice versa. Fluid
movement across the epithelium is recorded by the changes in probe
output voltage. Ports in the bottom of the half-chambers allow for
solution and chemical composition changes on either side of the tissue.
The capacitance probe in each half-chamber was calibrated by injecting
1-µl volumes of fluid and measuring the probe output in millivolts
(737 mV/µl). This technique has a resolution of ~1 nl/min,
corresponding to a fluid transport rate approximately one-tenth the
average baseline fluid transport rate seen in the present experiments.
Voltage-sensing and current-passing bridges built into each
half-chamber permit continuous monitoring of TEP and
RT, the latter being calculated from the voltage
deflections in response to transepithelial current pulses of known
magnitude. The control Ringer solution for measurements of
JV, TEP, and RT contained
(in mM) 113.5 NaCl, 5 KCl, 26 NaHC03, 1.8 CaCl2, 0.8 MgS04, 1.0 NaH2P04, and 5.5 glucose, pH 7.4. In some
experiments, a cAMP cocktail was added to the control Ringer solution:
500 M 3-isobutyl-1-methylxanthine (IBMX), 100 µM
8-(4-chlorophenylthio)adenosine 3',5'-cyclic monophosphate (CPT-cAMP),
and 12 µM forskolin. In other experiments, the following inhibitors
were tested: 10-50 µM amiloride (to block ENaC), 0.5-1.0 mM
DPC (to block CFTR), and 50 µM 5-nitro-2-(3-phenylpropylamino)benzoic
acid (NPPB; to block CFTR).
Patch-clamp electrophysiology.
Cells were plated on glass coverslips at low density. Patch pipettes
were pulled from glass (WPI, Sarasota, FL) on a microelectrode puller
(Narishige, Tokyo, Japan) and fire-polished on a microforge (Narishige)
to a resistance of 2-10 M
when filled with intracellular solution (see below). Seal resistances were typically 10 G
. Data were recorded in the voltage-clamp mode with an amplifier (Axopatch-1D; Axon Instruments, Foster City, CA) connected to a personal computer with an analog-to-digital board. Voltage sweeps and currents were recorded with the use of Clampex acquisition software (Axon
Instruments). Leak subtraction was not performed on any of the data.
Solution changes were made via a seven-node perfusion chamber. The bath chamber was maintained at 37°C with a battery-operated resistive element mounted on the bottom of the chamber and controlled by a
battery-operated feedback circuit.
To identify CFTR, whole cell patch pipettes were filled with
intracellular solution containing (in mM) 121.5 N-methyl-D-glucamine (NMDG)-gluconate, 13.5 NMDG-Cl, 1.8 ATP, 0.09 GTP, 9 HEPES, and 9 glucose. Extracellular
solution contained (in mM) 141 NaCl, 4 KCl, 1 KH2PO4, 1 MgSO4, 1 CaCl2, 10 HEPES, and 10 glucose, with either 10 µM
forskolin or 2 mM DPC added. Voltage sweeps were run from
80 mV to
100 mV, with data sampled at a frequency of 10 kHz.
Whole cell experiments were also performed in an attempt to identify
ENaC in single 31EG4 cells. Cells were plated at low density and then
treated with 1 µM dexamethasone for roughly 24 h before the
experiment to increase cellular differentiation. Pipette solution for
these experiments contained (in mM) 121.5 NMDG-gluconate, 2.7 NMDG-Cl,
0.9 CaCl2, 0.9 MgCl2, 9 HEPES, 4.5 glucose,
2.25 ATP, 0.09 GTP, and 9.9 EGTA. Two different extracellular solutions
were employed for these experiments. Extracellular Na-gluconate solution contained (in mM) 145 Na-gluconate, 7 NaCl, 1 CaCl2, 1 MgCl2, 10 HEPES, and 5 glucose.
Extracellular NaCl solution contained (in mM) 150 NaCl, 1 CaCl2, 1 MgCl2, 10 HEPES, and 5 glucose.
Voltage sweeps were run from
80 mV to 100 mV, with data sampled at a
frequency of 10 kHz. Conductances values were obtained by measuring the
slopes of the current-voltage (I-V) curves at around 20 mV.
Transepithelial and microelectrode electrophysiology.
The recording setup and perfusion system has been described previously
(36). Transwells with confluent 31EG4 monolayers and
RT > 300
· cm2
were used for the electrophysiology measurements. The monolayers on
filters were mounted on a nylon mesh support and clamped into a
modified Ussing chamber. RT and the ratio of the
apical to basolateral membrane resistance (a) were obtained
by passing 4-µA current pulses across the tissue and measuring the
resultant changes in TEP and voltages across the apical
(VA) and basolateral membranes (VB). Current pulses were bipolar, with a period
of 3 s applied at various time intervals.
RT is the resulting change in TEP divided by 2 µA, and a is the ratio of voltage change in
VA divided by the change in
VB (a =
VA/
VB). The
current-induced voltage deflections were digitally subtracted from the
records for clarity. Short-circuit current (Isc)
measurements were performed with the use of similar chambers and
electrophysiological apparatus, with TEP clamped to zero and using
solution and resistance compensation. Pulses of 5 mV were often
utilized to measure RT in these experiments. The
control Ringer solutions (in mM) used for measurements of TEP,
RT, and Isc are identical
to those used for the measurements described in Fluid
transport.
Calculating membrane and shunt resistances and equivalent
electromotive forces.
The intracellular measurements were analyzed with an equivalent circuit
model (see Fig. 1) that has been
previously described (26, 32). We used the microelectrode
data to calculate resistances (R, in
· cm2) and equivalent electromotive forces
(E, in mV) for the apical and basolateral membranes and the
shunt for 31EG4 cells. The general approach used (26) was
to measure TEP as well as VA and
VB and determine a values
(a =
VA/
VB = RA/RB) in control conditions and then during
treatment with amiloride. This allowed us to calculate shunt resistance
(RS). It was assumed that amiloride only altered EA and RA. Calculation of
equivalent circuit parameters was based on the following steady-state
considerations
|
(1)
|
where i is the loop current generated by the
difference in VA and VB,
and EA and EB are the
apical and basolateral membrane potentials, respectively, that would be
recorded if RS were infinite. By definition
|
(2)
|
RT is given by
|
(3)
|
Combining Eqs. 1 and 2 gives
|
(4)
|
Similarly,
|
(5)
|
Using Eq. 3 before and after (denoted with
asterisk) amiloride, we obtain
|
(6)
|
Using Eqs. 4-6 and the data collected before and
shortly after amiloride treatment, we calculated
RS and then RA,
RB, EA, and EB for control and amiloride-treated conditions.
This calculation assumes that amiloride affects only the apical
membrane, while RS and RB
remain constant.

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Fig. 1.
Equivalent circuit for 31EG4 mammary epithelia. Apical and
basolateral membranes are represented by equivalent resistance
(RA and RB, respectively)
in series with an electromotive force (EA and
EB, respectively) that would be measured in the
absence of a shunt. Shunt resistance (RS)
represents the parallel combination of the junctional complex
resistance and the mechanical seal resistance around the tissue. A loop
current (is) flows through the circuit due to
the difference between the measured apical and basolateral membrane
voltages (VA and VB,
respectively). The transepithelial potential (TEP) represents the
difference between VB and
VA.
|
|
Data are presented as means ± SE, unless otherwise specified.
Student's unpaired t-test was used to compare groups, and
P < 0.05 is considered statistically significant.
 |
RESULTS |
PCR, Western blot analysis, and immunocytochemistry.
PCR and Western blot analysis were performed on 31EG4 cells grown to
confluence in the presence of dexamethasone to increase differentiation
of the cells. The PCR experiments summarized in Fig.
2 showed that mRNA for both ENaC and CFTR
was present in 31EG4 cells. In Fig. 3,
Western blots with bands at 170 (CFTR) and 85 kDa (ENaC
subunit)
indicate that both proteins were being made.

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Fig. 2.
PCR products for epithelial Na+ channel
(ENaC; 596 bp) and cystic fibrosis transmembrane conductance regulator
(CFTR; 463 bp). The amplified products were run on a 1.5% agarose gel
and stained with ethidium bromide. The 100-bp ladder is shown at
left (ladder).
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Fig. 3.
Western blot analysis of CFTR and -ENaC protein in
31EG4 cells. Cell homogenates were loaded for SDS-PAGE and analyzed
subsequently by Western blotting. The migration of molecular mass
markers (in kDa) is indicated. See text for details.
|
|
Immunomicroscopy was performed on 31EG4 cells grown to confluence on
filters. As shown in Fig. 4, A
and B, both ENaC and CFTR appeared to be expressed in the
apical portions of the 31EG4 cells. CFTR immunoreactivity was also
detected in intact human mammary glands by indirect immunofluorescence
(Fig. 5). The predominant site of
staining was the apical domain of the epithelial cells lining the ducts
of the mammary gland. Cytokeratin staining identified duct cells and
demonstrated tissue architecture for orientation.

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Fig. 4.
Immunofluorescence localization of ENaC (A) and CFTR
(C) and their respective bright-field images (B
and D) in a 31EG4 monolayer. Cryostat sections were stained
with primary antibodies to the -subunit of ENaC and to CFTR and were
then FITC-conjugated with secondary antibodies. The Transwell mesh is
visible below the monolayer (B and D). Staining
with primary antibodies showed that both ENaC (A) and CFTR
(C) were located in the apical regions of the cells.
Original magnifications, ×630.
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Fig. 5.
Detection of CFTR and cytokeratin 19 in intact human breast by
double-label indirect immunofluorescence with a1468 plus a monoclonal
antibody to cytokeratin 19. A: a1468 staining of the apical
domain of ductal epithelial cells as detected from the FITC (green)
fluorescence. B: combined fluorescence with both antibodies
in a double exposure showing fluorescence. TRITC fluorescence
(orange-red) identifies duct cells on the basis of their content of
cytokeratin 19. FITC fluorescence (green) occurs at the apical domain
of these duct cells. Original magnification, ×400.
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Transepithelial and patch-clamp electrophysiology.
31EG4 monolayers were mounted in Ussing chambers and then treated with
amiloride (to block ENaC), followed by forskolin (to stimulate cAMP
production and active PKA) and DPC (to block CFTR). As shown in Fig.
6, 10 µM amiloride in the apical
solution decreased Isc, although not to zero,
indicating that there was some residual anion secretion or cation
absorption. The identity of this ionic current has not been identified.
Instead, we concentrated on determining the potential contribution of
ENaC and CFTR to the ion transport properties of 31EG4 cells. In the
presence of apical amiloride, forskolin (10 µM) increased
Isc, and subsequent addition of apical DPC (500 µM) decreased Isc. In five experiments,
amiloride decreased Isc by 2.5 ± 0.7 µA/cm2 (mean ± SE); in the presence of amiloride,
forskolin increased Isc by 7.8 ± 1.4 µA/cm2, and subsequent addition of DPC decreased
Isc by 5.5 ± 0.8 µA/cm2.

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Fig. 6.
Effects of amiloride, forskolin, and
diphenylamine-2-carboxylate (DPC) on short-circuit current
(Isc) of 31EG4 monolayers. Cells grown to
confluency on filters were mounted in the chamber, and
Isc was measured. Spikes in the
Isc trace represent changes induced by briefly
clamping epithelium to 5 mV. Apical addition of amiloride caused
Isc to decrease from ~13.5
µA/cm2 to 11.5 µA/cm2. Subsequent treatment
with 10 µM forskolin (to increase cAMP) caused
Isc to increase from 13.5 µA/cm2
to 15 µA/cm2, and DPC reduced Isc
to 9 µA/cm2. Experiment is typical of 4 others.
|
|
Whole cell patch-clamp measurements were also performed on 31EG4 cells.
As shown in Fig. 7A, forskolin
increased cellular conductance from 1.7 nS to 34 nS and shifted the
reversal potential from about
20 mV to
25 mV. DPC reversed the
forskolin-induced changes by bringing the reversal potential and the
conductance back to their resting, baseline levels. Figure
7B summarizes the effects of forskolin and DPC on 31EG4 cell
conductance.

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Fig. 7.
Effects of forskolin and DPC on whole cell patch-clamp
current, current-voltage (I-V) relationship, and
conductance. A: whole cell I-V recording of a
resting cell (control) with a basal Cl conductance of 1.7 nS and a reversal potential of 20 mV. Upon addition of forskolin
(+fsk), conductance increased to 34 nS, and the reversal potential
shifted to 25 mV. Addition of 2 mM DPC [+DPC(+fsk)] decreased the
conductance to 2.7 nS and shifted reversal potential back to 20 mV.
B: summary of 7 similar experiments. Average whole cell
conductance of resting cells was 3.4 ± 1.2 nS, stimulated
conductance was 27.4 ± 4.1 nS, and post-DPC conductance was
2.7 ± 0.6 nS.
|
|
Amiloride-sensitive currents were also identified in whole cell
patch-clamp experiments. These currents were quite small in the resting
state (Fig. 8A), so the
inhibitory effects of amiloride were difficult to identify in every
cell. Figure 8A shows an experiment in which the cells were
incubated in a NaCl-containing Ringer solution; the I-V
curve showed small, linear currents (mean whole cell conductance = 0.6 nS, reversal potential =
15 mV). Conductance was inhibited
by amiloride to 0.3 nS, and the reversal potential shifted in the
negative direction to
22 mV, consistent with inhibition of ENaC. Two
other experiments were also performed on cells in which the bathing
solution was Na-gluconate Ringer (to eliminate contribution of
Cl
currents), and amiloride had similar effects on cell
currents and conductances, although the absolute conductances were, for some unexplained reason, significantly higher. Figure 8B
summarizes the results from two experiments in which the cells were
incubated in a NaCl-containing Ringer and two other experiments in
which the cells were bathed in Na-gluconate Ringer.

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Fig. 8.
Effect amiloride on whole cell patch-clamp current,
I-V relationship, and conductance in NaCl and Na-gluconate
Ringer solutions. A: whole cell patch recording of a resting
cell (control) with a basal conductance in NaCl solution of 0.6 nS and
a reversal potential of 15 mV. Addition of 10 µM amiloride (+amil)
caused conductance to decrease to 0.3 nS and shifted the reversal
potential to 22 mV. B: summary of 2 experiments performed
with NaCl bathing solutions ( ) and 2 experiments
performed with Na-gluconate solutions ( ). Average
resting conductance was 1.2 nS in NaCl solution and 0.7 nS after
amiloride. Average resting conductance in Na-gluconate solution was
16.5 nS, decreasing to 8.8 nS after amiloride.
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Intracellular recordings, calculated electromotive forces, and cell
membrane resistances.
Figure 9 shows a typical intracellular
microelectrode recording with resting, unstimulated levels of
VA, VB, TEP,
RT, and RA/RB. When a cocktail of
forskolin, CPT-cAMP, and IBMX was added to the apical bath, there was a
rapid and reversible increase in TEP, since VA
depolarized more than VB; concomitantly,
RT dropped and
RA/RB decreased from 3.5 to 2. All these responses are consistent with a conductance increase of
an apical membrane channel whose equilibrium potential is depolarized
with respect to VA.

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Fig. 9.
Effects of cAMP cocktail on membrane voltage and
resistance in 31EG4 cells. A: continuous traces represent
VA and VB, and open
circles represent the ratio of apical to basolateral membrane
resistance (RA/RB). The
cAMP-elevating cocktail (see MATERIALS AND METHODS) caused,
after a brief delay, depolarizations of both VA
and VB by >10 mV, whereas
RA/RB decreased.
B: TEP (continuous trace) increased by 2 mV,
RT ( ) decreased by 200 · cm2, and
RA/RB decreased by
approximately a factor of 2, from 3.4 to 1.7. These results are
consistent with cAMP opening Cl channels in the apical
membrane.
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|
As summarized in Table 1, the mean
resting RT and TEP are 829
· cm2 and
5.9 mV, respectively. The
TEP is apical-side negative, since VB is more
hyperpolarized than VA. In addition, the mean
RA/RB of ~3 indicates
that the apical membrane has a threefold greater resting resistance
than the basolateral membrane. Table 1 also summarizes the cAMP-induced
changes in membrane voltages and resistance, which are all consistent
with an increase in apical membrane conductance. From the summary data
in Table 1, the calculated (Ohm's law) equivalent
Isc of the resting monolayer increased from 7.1 µA/cm2 to 12.1 µA/cm2 after treatment with
the cAMP-raising cocktail, similar to the forskolin-induced increase in
Isc measured in the experiment in Fig. 6. The
results shown in Figs. 6 and 9 and summarized in Table 1 are consistent
with the hypothesis that cAMP activates apical membrane CFTR and
increases Cl
transport from the basolateral to the apical
surface at a rate of ~5 µA/cm2.
Because the CFTR-blocker DPC appeared to block the forskolin-induced
increases in Isc (Fig. 6), it was expected that
the cAMP-induced changes in cellular electrophysiology would also be
blocked by NPPB, another fairly specific CFTR blocker (25,
40). In the presence of apical NPPB (see Fig.
10), the cAMP cocktail had relatively little or even opposite effects on RT,
RA/RB, TEP,
VA, and VB (compare with
the following control or Fig. 10). For example, the slow decrease in
TEP shows that VB depolarized faster than
VA in the presence of NPPB, opposite to what is
seen for the cAMP-induced changes in TEP and membrane potential.
Practically identical NPPB-induced responses were observed in four
experiments. In summary, elevating cell cAMP depolarized
VA and VB, decreased
RA/RB and
RT, and increased TEP (Table 1). These changes
and their blockade by apical NPPB (or DPC, in Fig. 6) are all
consistent with the presence of CFTR at the apical membrane.

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Fig. 10.
cAMP-induced changes in membrane voltage and resistance
are blocked by 5-nitro-2-(3-phenylpropylamino)benzoic acid (NPPB) in
31EG4 cells. A: continuous traces represent
VA and VB, and open
circles represent RA/RB.
B: continuous trace represents TEP, and open squares
represent RT. In the presence of NPPB, the cAMP
cocktail elicited only small electrophysiological responses. Removal of
NPPB caused VA and VB to
depolarize by >10 mV, whereas TEP increased and both
RT and
RA/RB decreased.
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Intracellular recordings were also used to measure the effects of
amiloride on membrane voltages and resistances. Results from a typical
microelectrode experiment are shown in Fig.
11 where apical amiloride (20 µM)
decreased TEP and increased RT.
RA/RB increased by more
than a factor of six, and VA hyperpolarized by
24 mV. All the electrophysiological changes were reversed when amiloride was removed from the apical bath. Similar amiloride-induced changes were obtained in 16 experiments and are summarized in Table 1.
These results indicate that amiloride blocked Na+ entry
through apical ENaC.

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Fig. 11.
Amiloride-induced changes in membrane voltage and
resistance. A: continuous traces represent
VA and VB, and open
squares represent RA/RB.
In control Ringer, VA = 46 mV. Apical
amiloride (20 µM) hyperpolarized VA by 24 mV;
RA/RB increased by
approximately a factor of 6. B: TEP (continuous trace)
decreased by 2 mV, and RT ( ) increased by
100 · cm2. Effects of amiloride were readily
reversed upon removal from the apical bath.
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Addition of 20 µM amiloride to the basolateral solution caused small
and equal depolarizations (<5 mV) in VA and
VB with no change in TEP, a 30% drop in
RA/RB, and a 5% increase
in RT (n = 3; not shown). These
small alterations in membrane voltage and resistance may have been due
to the effect of amiloride on the basolateral
Na+/H+ exchanger (41) and
secondary changes in cellular pH.
The data summarized in Table 1 were used as described in
MATERIALS AND METHODS to calculate resistances and
electromotive forces for the apical and basolateral membranes and shunt
for 31EG4 cells in control and during amiloride. The results of these calculations are summarized in Table 2.
The assumption (MATERIALS AND METHODS) that amiloride had
no affect on RS or RB was
tested by allowing either RS or
RB to change up to ±20% after amiloride treatment. In both cases, all of the calculated parameters were within
0.5 SD of the mean values shown in Table 1, indicating that within the
error of these experiments, the effects of amiloride were restricted to
the apical membrane.
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Table 2.
Equivalent circuit parameters calculated from control data and from
electrical changes induced by addition of amiloride
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|
Comparison of amiloride effects under control and cAMP- or
ATP-treated conditions.
Previous experiments have shown increased Na+ and fluid
absorption in cystic fibrosis (CF) tissues (7, 20, 24),
and heterologous expression of CFTR is known to decrease the activity of ENaC in both frog oocytes (27) and Madin-Darby canine
kidney cells (43, 44). However, it also has been shown
that ENaC activity is much reduced in freshly isolated CF sweat ducts
that lack functional CFTR (38). Because 31EG4 cells
express both CFTR and ENaC, we tested whether cAMP treatment could
alter ENaC activity. A typical experiment is shown in Fig.
12A. In the first part of
the experiment, amiloride (20 µM) hyperpolarized
VA by 20 mV and increased baseline
RA/RB by a factor of 10, from 0.25 to 2.5; in addition, RT increased and
TEP decreased. Subsequent addition of the cAMP cocktail, in the
continued presence of amiloride, depolarized VA
by ~24 mV and decreased
RA/RB to its baseline
level in control Ringer. cAMP addition also decreased
RT and increased TEP. Similar results were
obtained in two other experiments (not shown). The voltage changes were
somewhat larger than those exhibited in the absence of amiloride,
probably because amiloride increased the driving force for
Cl
exit across the apical membrane. These results show
that the blockade of ENaC activity with amiloride does not appreciably alter the cAMP-induced changes in membrane voltage and resistance.

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Fig. 12.
Amiloride-induced changes in membrane voltage and
resistance in the absence and presence of cAMP-elevating cocktail.
A: amiloride caused characteristic changes in
VA, VB,
RA/RB ( ),
TEP, and RT ( ), and subsequent
addition of cAMP cocktail in the continued presence of amiloride
produced cAMP responses similar to those measured in control Ringer
(Table 1). B: 31EG4 cells were first treated with cAMP
cocktail, which caused characteristic depolarization of
VA, decrease of
RA/RB ( ),
and increase in TEP. Subsequent addition of amiloride in the continued
presence of cAMP produced no significant changes in
RT ( ),
RA/RB, TEP, and
VA. Summary data are shown in Table 1.
C: amiloride-induced changes in TEP and
RT ( ) in the presence and
absence of cAMP or ATP were measured in the same monolayer.
Top: control TEP and RT were 5.5 mV
and 950 · cm2, respectively. Apical amiloride
reversibly decreased TEP (1.5 mV) and increased
RT (125 · cm2). The
tissue was returned to control Ringer, and then cAMP cocktail was added
to the apical bath; TEP increased by 1.7 mV and
RT decreased by 110 · cm2. In the presence of cAMP, apical
amiloride produced no changes in TEP or RT. The
monolayer was then returned to control Ringer (30 min); TEP and
RT returned to their control levels.
Bottom: apical ATP increased TEP (2 mV) and decreased
RT (200 · cm2). In the
presence of ATP, apical amiloride reversibly decreased TEP by 1 mV and
increased RT by 75 · cm2. Subsequent ATP removal decreased TEP and
increased RT.
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|
In contrast, the data in Fig. 12B show that apical amiloride
caused significantly smaller electrophysiological changes in the presence of cAMP stimulation: VA and
VB hyperpolarized by only 5 mV, and
RA/RB,
RT, and TEP hardly changed in the presence of cAMP (Table 3). Equivalent circuit
calculations of membrane resistances and voltages were also performed
on the subset of microelectrode experiments in which amiloride was
added to monolayers that had been treated with cAMP. Results from these
calculations have been summarized in Table
4. Compared with control conditions
(Table 2), cAMP caused RA to decrease from 1,958
· cm2 to 157
· cm2; in
the presence of cAMP, amiloride had essentially no effect on
RA (Table 4).
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Table 3.
Steady-state levels of membrane voltage and resistance in the presence
of cAMP alone and amiloride responses in the presence of cAMP
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Table 4.
Equivalent circuit parameters calculated from changes induced by cAMP
in the absence and presence of amiloride
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|
The apparent reduction in ENaC activity might be secondary to
cAMP-induced changes in membrane voltage or resistance. This possibility was tested by activating another set of apical
Cl
channels to see if they also reduced the effects of
amiloride. We have found that the apical membrane of 31EG4 cells
contains P2Y receptors that, when activated by ATP, elevate cell
Ca2+ (Blaug and Miller, unpublished observations) and
produce membrane voltage and resistance changes very similar to those
produced by elevating cell cAMP: in seven experiments, apical ATP (50 µM) depolarized VA by ~17 mV, from
43.9 ± 1.7 mV to
26.9 ± 1.1 mV; RA/RB decreased from
3.5 ± 0.2 to 1.8 ± 0.3, and RT
decreased by 109
· cm2, from 518 ± 39
· cm2 to 409 ± 20
· cm2 (means ± SE), changes not
significantly different from those produced by cAMP (Table 1). If the
reduction in ENaC activity were entirely dependent on cAMP-induced
changes in membrane voltage or resistance, then activation of
Ca2+-activated Cl
channels by ATP should also
inhibit the amiloride-induced changes in TEP and
RT.
The experiment summarized in Fig. 12C tested this notion by
comparing the amiloride responses in the same monolayer, first in the
presence of cAMP (top) and then in the presence of ATP (bottom). Identical results were obtained independent of
order. Apical amiloride characteristically decreased TEP and increased RT, and these responses were reversible. In the
presence of cAMP (which increased TEP and decreased
RT), amiloride had no significant effect on TEP
or RT. The cAMP cocktail and amiloride were
removed, and the monolayer was returned to control Ringer for 30 min.
Figure 12C, bottom (same monolayer), shows that the addition
of ATP to the apical bath had effects on TEP and
RT nearly identical to those produced by cAMP.
In the presence of ATP, amiloride decreased TEP and increased
RT by amounts that were very similar to those in
control, untreated monolayers. These results are summarized in Table
5. The amiloride-induced changes in TEP
and RT are significantly reduced in the presence
of cAMP but not in the presence of ATP.
Fluid transport.
Rates of transepithelial fluid movement along with TEP and
RT were measured under baseline, control
conditions in 31EG4 monolayers grown on filters and then after the
addition of either a cAMP-stimulating cocktail or amiloride to the
apical bath. Figure 13 summarizes the
data from a cAMP experiment. The baseline JV was
~3.5 µl · cm
2 · h
1, in
the absorption direction. Addition of cAMP cocktail reversibly altered
TEP and RT, as in the electrophysiology
experiments. Consistent with increased Cl
secretion, cAMP
reversed the direction of steady-state fluid flow from absorption to
secretion (approximately
2.5
µl · cm
2 · h
1); this
JV response was partially reversible.

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Fig. 13.
cAMP-induced stimulation of steady-state fluid secretion. Apical
addition of the cAMP cocktail reversed the direction of net
steady-state fluid transport from absorption (3.5 µl · cm 2 · h 1) to
secretion ( 3.0
µl · cm 2 · h 1) and
simultaneously increased TEP and decreased RT
consistent with the electrophysiological responses summarized in Table
1. The obligatory control-to-control solution change (probes out; see
MATERIALS AND METHODS) during the first 40 min produced
very little change in JV, TEP, and
RT ( ). Removal of cAMP from the
apical bath brought JV back to absorption at
~1.0 µl · cm 2 · h 1. The
electrical changes were also reversible.
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|
In control Ringer, the direction and magnitude of fluid transport was
variable, independent of time in culture (6-9
days).1 Of 17 cultures
(summarized in Fig. 14), 9 exhibited a
baseline absorption (range: 0.5 to 7.6 µl · cm
2 · h
1) and 8 exhibited a baseline secretion (range: 0.75 to
5
µl · cm
2 · h
1). cAMP
always caused fluid
secretion,2 which ranged from
1 to
8 µl · cm
2 · h
1.
The cAMP-induced changes in TEP and RT were very
similar for both fluid-absorbing and -secreting tissues and were very
similar to the electrophysiological results summarized in Table 1. In six other monolayers, two absorbing and four secreting (Fig. 14), apical amiloride increased the mean secretory rate to
6.7 ± 4.8 µl · cm
2 · h
1 (mean ± SD).

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Fig. 14.
Summary of fluid transport experiments with cAMP
(n = 11) or amiloride (n = 6) addition.
A subset of monolayers (n = 9) absorbed fluid at a mean
JV of 3.6 ± 1.1 µl · cm 2 · h 1. Another
subset of cultures (n = 8) secreted fluid at a mean
rate of 4.6 ± 0.8 µl · cm 2 · h 1. Both cAMP
and amiloride induced fluid secretion.
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|
In Fig. 15 we plotted the magnitude
(|
JV|) of the cAMP-induced
fluid-secretory response as a function of steady-state
JV. |
JV| was largest for
those monolayers that absorbed fluid in the baseline condition (before
addition of cAMP) and lowest for those that secreted fluid. Figure 15
also shows that for monolayers in the baseline condition (no amiloride
or cAMP), RT was highest in monolayers that
absorbed fluid and lowest in monolayers that secreted fluid in the
baseline condition.

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Fig. 15.
Magnitude of cAMP-induced fluid secretory response
(| JV|) and RT are plotted as
a function of baseline, steady-state JV in 31EG4
cells. cAMP-induced fluid secretion rate and RT
increase linearly (as net fluid absorption rate increases). The
correlation coefficients, 0.6 and 0.8, respectively, are statistically
significant (P < 0.01).
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 |
DISCUSSION |
CFTR, ENaC, ion transport, tight junctions, and mammary gland
physiology.
The normal (i.e., nontransformed) mammary epithelial cell line 31EG4
expresses CFTR and ENaC in the apical membranes. Messages for both CFTR
and ENaC were identified, and Western blots showed that proteins with
the expected molecular sizes were present. Immunomicroscopy of the
cultures and intact human mammary tissue (Figs. 4 and 5) indicated that
both channels are located either in or very close to the apical
membrane. Although the immunomicroscopy could not determine the precise
membrane locations of CFTR and ENaC, electrophysiology indicated that
these channels were active in the apical membranes of the cells.
In the short-circuited condition, or when open-circuit TEP and
RT data were used to calculate equivalent
Isc, 31EG4 cells exhibited currents of 2-7
µA/cm2 that were blocked by apical, but not basolateral,
amiloride, consistent with the presence of functional apical ENaC.
Whole cell patch-clamp measurements also demonstrated
amiloride-inhibited currents and conductance as well as linear
I-V curves expected for ENaC. Microelectrode experiments
showed that amiloride hyperpolarized EA and
caused large increases in RA, consistent with
blocking apical ENaC. The comparatively smaller amiloride responses in the patch-clamp vs. the microelectrode experiments is probably due to
poor seals in the patch-clamp experiments and the relatively low
resistance of the shunt compared with the cellular pathway.
cAMP depolarized EA and reduced
RA by more than a factor of 10 (compare Tables 2
and 3) and also increased basolateral-to-apical anion current (likely
Cl
or HCO
) by 5 µA/cm2;
these changes were inhibited by apical DPC or NPPB. Patch-clamp experiments also showed forskolin-stimulated, DPC-inhibitable currents
and conductance as well as linear I-V curves. cAMP treatment also may have altered the voltage or conductance properties of other
Cl
channels at the basolateral membrane
(46), but clearly a major effect of cAMP stimulation was
to activate apical CFTR. 31EG4 cells also express apical membrane
ATP-activated Cl
channels and apical and basolateral
K+ channels (unpublished observations). These channels may
contribute to the negative values of EA and
EB and could also be important for ion and fluid
secretion (and absorption) that occur in both baseline (control) and
cAMP-stimulated conditions.
Our calculations that RS (1,000-1,400
· cm2) is smaller than total cellular
resistance (Rcell = RA + RB = 2,100-2,500
· cm2) indicate that although
RS (Table 1) is large compared with many tissues
with "leaky" tight junctions (RS < 100-200
· cm2; see Ref. 14),
it is apparently still a major pathway for current flow. The low value
of RS compared with Rcell
also helps keep the TEP in 31EG4 cells relatively small,
6 mV,
similar to the situation in leaky epithelia (32). As
discussed below, this tight junctional permeability also contributes to
the ability of 31EG4 cells, a clonally derived cell line (39, 42,
48) with roughly equal distributions of ENaC and CFTR in all the
cells of the monolayer (Figs. 4 and 5), to both secrete and absorb
fluid (Fig. 14 and footnote 1). It will be interesting to determine
whether acinar and ductal cells in situ retain this capability or if
the present results represent a culture-to-culture variation in the ratio of secretory (acinar?) to absorptive (ductal?) epithelial cells.
The present demonstration of functional ENaC in the apical membrane of
31EG4 cells is consistent with previous experiments showing
amiloride-sensitive Na+ absorption across short-circuited
primary cultures of both midpregnant and lactating mammary gland cells
(3). Similarly, localization of CFTR to the luminal
membrane of 31EG4 cells is consistent with the presence of apical CFTR
in native human mammary duct cells (Fig. 5). Our results therefore
indicate that ENaC and CFTR of mammary duct cells may be involved in
generating or maintaining the characteristic Na+ and
Cl
concentrations of milk (34).
ENaC and CFTR also may be involved in the abnormal generation and
accumulation of breast fluid (so-called cystic disease of the breast)
that occurs in 7-10% of women in Western countries, mainly in the
premenopausal decade. There is now strong evidence to indicate that
these women are at a two- to fourfold higher risk of later developing
breast cancer (5, 8, 10, 16, 30). Although the chemical
compositions of these fluid-filled cysts are quite complicated
(1, 9), there appears to be a distinct pattern of
Na+ and Cl
concentrations (17,
33) that could reflect alterations in luminal CFTR and ENaC
activity. Clinically, it has been shown that tamoxifen reduces the
number and size of such cysts (28). Because tamoxifen can
block Cl
channels (15, 47), it may be
exerting its therapeutic effect by decreasing fluid secretion.
Effects of cAMP on CFTR and ENaC: evidence for cross talk?
cAMP caused, in addition to the expected increase in CFTR activity and
consequent drop in RT and
RA, an almost complete inhibition of the
amiloride-induced changes in TEP, RT,
RA/RB, and
RA (Fig. 12B; Tables 1-4). This
apparent inhibitory effect of activated CFTR on ENaC may be understood
by a direct interaction between the channels or indirectly by other
mechanisms. 1) The amiloride-induced voltage responses may
have been decreased because cAMP depolarized the apical membrane and
reduced the driving force for Na+ entry. However, similar
inhibitory effects were seen in other experiments (not shown) in which
the apical membrane was current-clamped close to its resting level in
the absence of cAMP. 2) The cAMP-induced increase in apical
membrane conductance may have been sufficiently large to mask the
amiloride-induced changes in membrane resistance and voltage. Against
that possibility are the data summarized in Fig. 12 and Table 5, which
show that cAMP and ATP had practically identical effects on TEP and
RT (and VA and
RA/RB; not shown) but
that the amiloride responses were totally obliterated in cAMP and
relatively unaffected by ATP. However, the conclusion that activation
of CFTR inhibits ENaC is not certain (27, 43) because the
cAMP-induced, but not the ATP-induced, increase in apical membrane
conductance may have been sufficient to mask the amiloride-induced changes in membrane resistance and voltage. A more definitive experiment would be to measure the amiloride-induced changes in RA/RB in ATP- vs.
cAMP-treated cells. However, we have been unable to hold microelectrode
penetrations long enough to make this comparison.
Critical role of tight junctions in ion and fluid transport.
The present data based on 31EG4 monolayers and evidence from other
epithelial systems (25, 37) indicate that net vectorial transport depends critically on ENaC and CFTR operating in concert with
the tight junctions. Fluid absorption is mainly controlled by the
transport of Na+ down its electrochemical gradient through
apical ENaC, while fluid secretion is regulated by cell-to-lumen
movement of Cl
and/or HCO
through
cAMP-stimulated CFTR. In both cases, the obligatory movement of
counterions (cations to accompany active Cl
secretion and
anions to accompany active Na+ absorption) likely takes
place predominantly through leaky tight junctions driven by the basal
side-positive TEP. Thus "leaky" tight junctions are important for
the ability of an epithelium to exhibit both absorption and secretion.
The net balance between secretion and absorption in 31EG4 cells may be
determined in part by the activity of CFTR in the apical membrane. This
idea is consistent with the observation that fluid absorption increases
monotonically with RT, and the cAMP-induced alteration in JV is smallest for control tissues
that secreted fluid and largest for control tissues that absorbed fluid
(Fig. 15). Thus, when CFTR conductance is lowest,
RT is highest, and the monolayers absorb
Na+, Cl
, and fluid. Subsequent elevation of
cell cAMP could then activate the maximum number of CFTR channels and
produce the maximum alteration in JV (Fig. 15).
Conversely, cultures that have a relatively low RT may have a relatively large complement of
active CFTR in the apical membrane and therefore secrete fluid in the
basal state (Fig. 15); secretion is increased by amiloride block of
ENaC (Fig. 14).
The tight junctions will also play a role in determining secretion vs.
absorption because the intraepithelial current (i = TEP/RS) (32), determined in part by
the low resistance of the junctions (RS < RA + RB),
hyperpolarizes VA, thereby increasing Na+ entry (ENaC) into and Cl
(CFTR) exit from
the cells. During cAMP stimulation VA
depolarizes by 18 mV (Table 1), and the increase in fluid secretion
indicates that the net electrochemical gradient for Cl
is
outward across the apical membrane.
These considerations emphasize the importance of the integrated
activities of ion channels, tight junctions, and ion transporters in
determining the direction of net Cl
movement through
CFTR. They also suggest a functional distinction among epithelia with
apical ENaC and CFTR. Epithelia with tight junctions that are
essentially impermeable to ions can only absorb salts and fluid via the
cellular pathway. In these tissues, the shunt current is zero, and
VA is sufficiently depolarized such that the
electrochemical driving forces for both Cl
and
Na+ are inward across the apical membrane. Therefore,
activation of CFTR with cAMP elicits absorption of Cl
,
Na+, and fluid through the cells (see footnote 2 in
Fluid transport). Examples include human sweat ducts
and bovine trachea (38, 46) and also may be true in the
lactating mammary gland, which has large lumen-negative TEP (
35 mV),
characteristic of all tight epithelia, and low concentrations of
Na+ and Cl
in the milk (35).
In contrast, epithelia like 31EG4 cells that have relatively leaky
junctions generate a shunt current that hyperpolarizes VA, increasing Cl
exit (CFTR) and
Na+ entry (ENaC) across the apical membrane. The
conductance of the shunt to anions and cations and the existence of a
TEP permit these cells to either secrete or absorb salts and fluid,
depending on the ratio of ENaC to CFTR conductance and on the direction and magnitude of the electrochemical driving forces for these ions
across the apical membrane. This also appears to be the case in airway
submucosal glands. When CFTR is activated, these cells secrete fluid
driven by the transport of HCO
and Cl
through CFTR (2, 23, 45) with Na+ following
through ion-permeable tight junctions to maintain electroneutrality. In
CF, when CFTR is inactive, submucosal gland cells absorb
Na+ (through ENaC), Cl
(through the
paracellular pathway), and fluid (25). This capability to
both secrete and absorb fluids also may be characteristic of the
nonlactating mammary gland (with its leaky tight junctions) (35).
 |
ACKNOWLEDGEMENTS |
We thank Connie Yu for help with experiments summarized in Fig. 6
and Van Nguyen, Steve Jalickee, and Natalia Zhuravel for experiments
summarized in Fig. 14. We also thank Beate Illek, Horst Fischer, and
Jonathan Widdicombe for helpful comments on an earlier version of the manuscript.
 |
FOOTNOTES |
This work was supported by grants from the Department of the Army
Breast Cancer Research Program DAMD17-96-1-6315 (S. S. Miller), DAMD17-97-5231 (T. E. Machen), California Breast Cancer Research Program 5JB-0077 (S. S. Miller), and National Institute of
Diabetes and Digestive and Kidney Diseases Grant DK-51799 (T. E. Machen).
Address for reprint requests and other correspondence: S. S. Miller, 360 Minor Hall, Univ. of California at Berkeley, Berkeley, CA 94720-2020 (E-mail:
smiller{at}socrates.berkeley.edu).
1
In a comparison of 6-, 8-, and 14-day-old
cultures from the same set of cells, the mean (±SE)
JV was
0.9 ± 1.6 (n = 8;
4 absorbing),
2.6 ± 0.8 (n = 9; 3 absorbing),
and
3.3 ± 2.9 µl · cm
2 · h
1
(n = 11; 3 absorbing), respectively. These values are
not statistically different from each other (P > 0.3).
2
In two experiments (not shown), net fluid
absorption was increased by cAMP. As expected, cAMP decreased
RT in these two experiments, but TEP also
decreased, which is opposite to the results obtained in all the other
experiments. This finding suggests that the electrochemical gradient
for Cl
was inward in these two cases, as was previously
shown in bovine trachea (46).
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 1 March 2000; accepted in final form 26 March 2001.
 |
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