Epithelial cells form a continuous barrier between the external
environment (i.e. lumen) and interstitium while vectorially
transporting ions, solutes, and macromolecules between these two
compartments. These cells also vectorially secrete their own protein
products; for example, epithelial cells composing the exocrine glands
secrete specific secretory products (e.g. mucins and digestive
enzymes) into the lumen. Nonglandular epithelial cells also secrete a
number of ``housekeeping'' proteins (e.g. cytokines,
protease inhibitors, components of the extracellular matrix, and
antibacterial peptides) that protect and maintain the mucosa and
serosa(1, 2, 3, 4, 5) . Two
well defined classes of protein secretory pathways are utilized by
epithelial as well as nonepithelial cells: (a) the regulated
secretory pathway and (b) the constitutive pathway (for review
see (6) ). The regulated secretory pathway involves the acute,
stimulus-induced release of secretory material from a preformed storage
compartment, such as the mucin storage granules of goblet
cells(7) . Conversely, the constitutive secretory pathway lacks
well defined storage granules and operates at a considerable basal
level of activity. Constitutive membrane traffic pathways are also
subject to regulation; e.g. there is accumulating evidence
that the rate of constitutive membrane traffic from the Golgi apparatus
to the cell surface can be modulated by regulatory factors such as
heterotrimeric G proteins and protein
kinases(8, 9, 10, 11, 12) .
The molecular mechanisms and physiological stimuli underlying the
protein kinase C-dependent regulation of constitutive secretion in rat
basophilic leukemia cells have been partially characterized and involve
the regulated binding of ARF (
)to Golgi
membranes(11) . Less is known about the regulatory role of
cAMP-dependent protein kinase in constitutive membrane traffic. Apical
membrane traffic in Madin-Darby canine kidney epithelial cells is
stimulated by cAMP analogs but only at high concentrations (0.5-5
mM 8-Br-cAMP) (10) or in combination with high
concentrations of phosphodiesterase inhibitors (500 µM 3-isobutyl-1-methylxantine)(8) . Unresolved issues
include: (a) whether or not apical protein secretion can be
stimulated by physiological activators of the cAMP-dependent protein
kinase pathway (e.g. hormone receptors coupled to adenylate
cyclase) in polarized epithelial cells and (b) the identities
of the downstream effectors that mediate the stimulation of
constitutive secretion by cAMP-dependent protein kinase.
In an
earlier work we demonstrated that cAMP inhibits endocytosis and
stimulates the recycling of previously internalized glycoproteins to
the cell surface in pancreatic and intestinal epithelial cells that
express wild type CFTR(13, 14) . Cells that were
homozygous for the most common CFTR mutation (
F508) lacked the
regulation of either endocytosis or exocytosis by cAMP(14) .
Because CFTR itself has been shown to enter the recycling pathway (15) and is present in clathrin-coated vesicles(16) ,
it has been postulated that CFTR modulates the recycling pathway via
its function as a cAMP-regulated Cl
channel within
this pathway(15) . Given that a subset of recycling
glycoproteins recycle through the TGN following their endocytosis from
the cell surface(17, 18, 19) , it is
conceivable that the recycling and biosynthetic pathways share common
compartments and perhaps regulatory elements at the level of the TGN.
On this basis we reasoned that cAMP (perhaps through CFTR) could
regulate not only apical membrane recycling but also protein processing
and polarized secretion along the biosynthetic pathway.
In order to
assess the role of cAMP in regulating the biosynthetic pathway, we
measured polarized protein secretion by colonic epithelial cells
(T
and HT29-CL19A) cultured on permeable supports,
characterized the cargo within the apical and basolateral secretory
pathways, and determined the effects of cAMP both on polarized
secretion and on protein sialylation. We demonstrate that cAMP
regulates apical but not basolateral protein secretion by these cells
in a dose-dependent manner. The apical secretion by colonic epithelial
cells is at least an order of magnitude more sensitive to cAMP
analogues than that described for Madin-Darby canine kidney cells and
is regulated by a physiological stimulator of the cAMP second messenger
system (i.e. VIP). One of the proteins secreted by HT29-CL19A
cells was identified as
1 antitrypsin, which is secreted at both
the apical and basolateral surfaces. Based on several criteria, we
determined that the secretion of AT from HT29-CL19A cells takes place
primarily via the constitutive pathway and that cAMP facilitates the
delivery of AT from the TGN to the apical cell surface. In addition,
when Cl
was replaced by gluconate, a nonpermeant
anion, the regulation was reversed, i.e. cAMP inhibited apical
protein secretion. Finally, we observed that cyclic AMP also stimulates
the rate of AT sialylation within the biosynthetic pathway, indicating
that the TGN is at least one site of action for cAMP. We discuss how
CFTR, a cAMP-regulated Cl
channel that has been
implicated in the acidification of the TGN and in the regulation of
membrane recycling, may be involved in the cAMP-dependent regulation of
protein sialylation and apical protein secretion in colonic epithelial
cells.
EXPERIMENTAL PROCEDURES
Materials
Tissue culture media were from Life
Technologies, Inc., and defined fetal bovine serum (FBS) was from
Hyclone (Logan, UT). Acrylamide, bis-acrylamide, ammonium persulfate,
TEMED, urea,
-mercaptoethanol, and ampholytes were from Bio-Rad.
Protein G immobilized on agarose beads was from Boehringer Mannheim.
Dialyzed FBS was prepared by dialyzing FBS against 100 volumes of
phosphate-buffered saline overnight at 4 °C in dialysis bags
(Spectrapor; molecular weight cut-off, 3,500) followed by sterile
filtration. Tran-[
S]-label was purchased from
ICN (Costa Mesa, CA). Forskolin was from Calbiochem (La Jolla, CA), and
Rp-8cpt-cAMPS was from Biolog (La Jolla, CA).
C-SDS-PAGE
molecular weight markers were from Amersham Corp. All other chemicals
were from Sigma.
Tissue Culture and Metabolic Labeling of
Cells
HT29-CL19A cells were isolated as a clonal cell line that
emerged from the parental HT29 cells following the induction of
differentiation by treatment with butyrate(20) . HT29-CL19A
cells represent a tissue culture model of nongoblet colonic crypt cells
based on the fact that less than 1% are mucin positive
cells(20) , the presence of cAMP-induced vectorial
Cl
transport, and high levels of CFTR
expression(21) , i.e. hallmarks of the colonic crypt (22, 23) . T
cells, another colonic cell
line that we used for our studies, also exhibit a very small proportion
of mucin-producing cells (
5%)(24) , display similar ion
transport properties, and also express CFTR(25, 26) .
HT29-CL19A cells were maintained in Dulbeco's modified
Eagle's medium (DMEM) supplemented with 10% FBS. T
cells were cultured in 1:1 DMEM:F12 supplemented with 10% FBS.
For all experiments cells were seeded onto Transwell filters (pore
diameter, 0.4 µm; Costar); either 10
cells for filters
of 6.5-mm diameter or 10
cells for filters of 24.5-mm
diameter. Electrical resistance was monitored using
``chopstick'' electrodes and a high impedance ohmmeter
(Millipore). Cells were typically used for experiments between 10 and
20 days following seeding at which time the electrical resistances were
at 600 ohms
cm
or higher. For metabolic labeling,
cells were incubated in a CO
incubator at 37 °C in
methionine and cysteine-free DMEM supplemented with 5% dialyzed FBS for
30 min followed by treatment with Tran-[
S]-label
in methionine- and cysteine-free DMEM/5% dialyzed FBS for an additional
30 min. Tran-[
S]-label was added to the
basolateral side only; 50 µCi/100 µl volume for the 6.5-mm
filters and 500 µCi/600 µl volume for the 24.5-mm filters.
Macroscopic Assay for the Secretion of Metabolically
Labeled Proteins by Trichloroacetic Acid Precipitation and
Scintillation Counting
Following metabolic labeling, cells were
washed two times with phosphate-buffered saline supplemented with 0.1
mM CaCl
and 1 mM MgCl
. 200
µl of DMEM + 10% FBS was then placed into both the apical and
basolateral compartments with or without secretagogues. Both media
samples were collected following various time periods and placed on
ice. Apical samples were centrifuged at 2,000
g for 2
min to remove any loose cells. When the chase was performed in the
absence of serum, FBS (i.e. carrier) was added to samples to a
final concentration of 10% (v/v) before trichloroacetic acid
precipitation. Trichloroacetic acid was added to the apical and the
basolateral samples to a final concentration of 10%. Following
incubation for 20 min on ice, samples were centrifuged at 3,000
g for 3 min, supernatants were discarded, and pellets were
washed with 10% trichloroacetic acid. Finally, the media samples were
centrifuged at 16,000
g for 3 min, and the pellets
were dissolved in 0.5 N NaOH. The radiolabeled proteins
remaining in the cells were collected by cutting the filters from the
filter cups and placing them into ice-cold 10% (w/v) trichloroacetic
acid. Filter were then washed in 10% trichloroacetic acid followed by
dissolving the cells in 0.5 N NaOH. Samples were counted using
a Packard (Downers Grove, IL) scintillation counter. Secretion was
expressed as (media counts/cell counts)
100 (i.e. the
percentage of radiolabeled proteins released).
SDS-PAGE Analysis of the Secreted Proteins
For
SDS-PAGE analysis, filters were labeled as above and washed twice with
phosphate-buffered saline supplemented with 0.1 mM CaCl
and 1 mM MgCl
. 30 µl of DMEM without
serum was placed into each compartment either with or without
secretagogues. In some experiments, the chase was performed in a
modified Earle's buffered salt solution (MEBSS) (116 mM NaCl, 5.4 mM potassium acetate, 0.4 mM MgSO
, 1.8 mM CaSO
, 0.9 mM NaH
PO
, 5.5 mM glucose, 4.9 mM sodium pyruvate, 26 mM NaHCO
in 5% CO
atmosphere) or in the same buffer but containing sodium gluconate
instead of NaCl (MEBSS gluconate). Following the chase period, the
media samples were collected and loose cells were removed from the
apical samples as described above. 5
SDS sample buffer (60
mM Tris (pH 6.8), 2%SDS, 5%
-mercaptoethanol, 0.1%
bromphenol blue, 50% glycerol) was then added to each sample followed
by heating to 95 °C for 10 min. Samples and
C-methylated molecular weight markers were resolved on 8%
T, 3% C SDS-PAGE gels, stained with Coomassie Blue, dried, and analyzed
using a Molecular Dynamics PhosphorImager (Sunnyvale, CA). Phosphor
image analysis was performed using IPlab Spectrum software (Signal
Analytics, Vienna, VA) on a Macintosh IIci computer.
Immunoprecipitation of
1-Antitrypsin
Cell
pellets were lysed at 4 °C for 60 min in RIPA buffer (50 mM Tris (pH 7.5), 24 mM sodium deoxycholate, 150 mM NaCl, 1% Triton X-100 (v/v), 0.1% SDS) supplemented with protease
inhibitors (0.1 mg/ml aprotinin, 0.1 mg/ml leupeptin, 1 mM phenylmethylsulfonyl fluoride). Lysates were clarified by
centrifugation at 16,000
g for 4 min. Supernatants
were precleared by treatment with protein G immobilized on agarose at 4
°C for 1 h. Precleared supernatants were then incubated with AT
antibody or with an identical amount of nonimmune IgG for 90 min
followed by incubation with immobilized protein G overnight.
Immunoprecipitates were washed twice with RIPA buffer and solubilized
in sample buffer either for SDS-PAGE or for one-dimensional IEF.
One-dimensional Isoelectric Focusing and Two-dimensional
Polyacrylamide Gel Electrophoresis
For one-dimensional IEF, gels
with a composition of 5% T, 3.3% C, 9 M urea, 1% (v/v) Biolyte
3/10, 4% Biolyte 5/7, 2% Triton X-100 were cast in minigel slabs
(Bio-Rad). Immunoprecipitated AT was dissolved in a sample buffer
containing 9 M urea, 1% (v/v) Biolyte 3/10, 4% Biolyte 5/7, 2%
Triton X-100, 5% 2-mercaptoethanol and loaded onto the slab gel.
Electrophoresis was performed at 5 watts constant power for 4 h and 8
watts constant power for 2 h using 20 mM NaOH as catholyth and
10 mM H
PO
as anolyth. Gels were fixed
in 10% trichloroacetic acid, washed in 1% trichloroacetic acid, and
dried prior to phosphor imaging. The first dimension of two-dimensional
PAGE was performed using the aforementioned gel composition and the
Mini-PROTEAN II tube system (Bio-Rad). Isoelectric focusing was
performed at 300 V for 90 min, 500 V for 120 min, and 1000 V for 60
min. For the second dimension gel tubes were adapted briefly in 100
mM Tris (pH 6.8), 2% SDS, 5% 2-mercaptoethanol, and 8%
glycerol and then subjected to SDS-PAGE using 8% T, 3% C gel slabs.
Gels were stained, dried, and subjected to phosphor imaging or
autoradiography.
RESULTS
General Considerations Regarding the Approach
In
order to assess the regulation of polarized protein secretion by
filter-grown cultures of colonic epithelial cells, we analyzed the
secretion of metabolically pulse-labeled proteins by three different
methods. First, labeled proteins that were secreted into either the
apical or basolateral compartment, as well as labeled proteins retained
within the cells, were trichloroacetic acid precipitated and quantified
by scintillation counting. The rates of secretion were then calculated
as the percentage of radiolabeled proteins released into either
compartment as a function of time. This technique provided a
macroscopic assay of the kinetics and regulation of apical and
basolateral secretion. Second, the apical and basolateral media samples
were directly analyzed by SDS-PAGE followed by phosphor imaging. This
``medium resolution'' technique provided an initial
qualitative assessment of the cargo within the apical and basolateral
secretory pathways and quantitative information regarding the secretion
of individual protein bands. Third, we used a combination of
two-dimensional gel electrophoresis, immunoprecipitation, and
one-dimensional isoelectric focusing to identify specific proteins
within the secretory pathway and to monitor protein processing in the
biosynthetic pathway.
cAMP Stimulates Apical but Not Basolateral Protein
Secretion
Fig. 1summarizes the results of our initial
experiments performed using the macroscopic assay of polarized protein
secretion. Both colonic cell lines behaved very similarly regarding the
polarized secretion of metabolically labeled proteins analyzed with
this method. Secretion exhibited linear kinetics throughout the 120-min
chase period and the rate of basolateral secretion by both cell lines
was severalfold greater than the apical secretion (Fig. 1, a and b). Secretion in both directions was abolished at 4
°C, as expected for a vesicle-mediated secretory process (Fig. 1b). When the chase was performed in the presence
of cpt-cAMP (Fig. 1, a and b), the rate of
apical secretion was stimulated significantly over baseline in both
colonic cell lines. Forskolin, a direct activator of adenylate cyclase,
also stimulated apical secretion by T
cells (Fig. 1b). Basolateral secretion was unaffected by
cyclic AMP in either of these two cell lines. Responsiveness to
secretagogues was more consistent for HT29-CL19A cells (Fig. 1, error bars); accordingly, these cells were utilized for the
more detailed analysis of the apical secretory pathway (see Fig. 2Fig. 3Fig. 4Fig. 5Fig. 6Fig. 7).
Figure 1:
Regulation of polarized protein
secretion as determined by trichloroacetic acid precipitation and
scintillation counting. Macroscopic assay was performed as described
under ``Experimental Procedures.'' The data shown are the
percentages of incorporated label (i.e. cell-associated label)
secreted as a function of time.
, control;
, 800 µM cpt-cAMP;
, 10 µM forskolin;
, 4
°C. Effect of forskolin on secretion by HT29-CL19A cells was not
evaluated by this technique. a, HT29-CL19A; b,
T
cells.
Figure 2:
a, electrophoretic profile of secreted
proteins under control and stimulated conditions. Assay was performed
as described under ``Experimental Procedures.'' Shown are
duplicate samples for each condition. Molecular mass markers are shown
in the middle (200, 97.4, 69, 46, or 30 kDa). Arrowheads,
major protein bands (95 and 60 kDa) observed in both apical and
basolateral samples; arrows, protein bands (105 and 120 kDa)
limited to the basolateral samples; hollow arrowheads, protein
bands (135 and >200 kDa) found only in apical samples. b,
dose-response relationship between cpt-cAMP concentration and apical
secretion. The percentage of stimulation was determined by measuring
the change in intensity of the 60-kDa band on phosphor images of apical
media samples. Shown is the mean ± S.E. of six determinations
from three separate experiments (each performed in duplicate). c, the subarea indicated in b depicted at a higher
resolution. Phosphor image analysis of the 95-kDa band revealed similar
quantitative results; however, for simplicity and because the 60-kDa
band was later identified as
1-antitrypsin, we show only the data
regarding the 60-kDa band.
Figure 3:
cAMP is a specific regulator of apical
protein secretion by HT29-CL19A cells. a, phosphor image of
the apically secreted 60-kDa band from a selected experiment shown
along with corresponding densitometry data averaged from three separate
experiments performed in duplicate (mean ± S.E., n = 6). The percentage of stimulation of the intensity of the
60-kDa band in the apical and basolateral media was assessed as
described for Fig. 2. Individual data points were normalized to
the mean intensity of the apical 60-kDa band determined for duplicate
samples under control conditions. b, corresponding phosphor
images and densitometry data of basolateral samples (mean ±
S.E., n = 6). Basolateral data are also normalized to
the apical control value to emphasize the differences in the baseline
levels of apical and basolateral secretion. Asterisks indicate
statistically significant differences from control (p <
0.0005). Lanes 1, control; lane 2, 200 µM cpt-cAMP added both apically and basolaterally; lane 3,
10 µM forskolin added both apically and basolaterally; lane 4, 100 nM VIP added basolaterally; lane
5, 100 nM VIP added apically; lane 6, 100 nM VIP added basolaterally along with 500 µM Rp-8cpt-cAMPS (i.e. a membrane-permeable, stereospecific
inhibitory analog of cAMP) added to both
sides.
Figure 4:
Two-dimensional PAGE analysis of apically
secreted proteins;
1-antitrypsin as the predominant secretory
protein. The direction of the first dimension is indicated. Molecular
markers mass were run in parallel (left side of panel, 200, 97.4, 69, 46, and 30 kDa). The arrow indicates the protein that corresponds to the 60-kDa protein
observed on one-dimensional gels. Isoelectric point of this protein
corresponds to 4.9-5.3 as determined by comparison with
two-dimensional protein markers (Bio-Rad). The position and appearance
of this protein is identical to the position and appearance of
1-antitrypsin secreted by HepG2 cells (Swiss 2D databank). The
protein corresponding to the 95-kDa band (asterisk) has not
been identified.
Figure 5:
Linear kinetics and Ca
independence of AT secretion in the absence or the presence of cAMP. a and b, representative data from one experiment are
shown. Individual data points represent the intensity of the 60-kDa
protein band determined by phosphor image analysis of apical (a) and basolateral (b) samples collected following
various chase periods in the presence or the absence of cpt-cAMP (200
µM). Correlation coefficients from linear curve fitting:
apical control R = 0.976, apical cAMP R = 0.979,
basolateral control R = 0.965, basolateral cAMP R =
0.945. c, extracellular [Ca
] and
ionomycin (1 µM) have no effect on AT secretion. Shown are
the normalized mean intensities of the 60-kDa apical band following 2 h
of chase as determined by phosphor image analysis of duplicate samples
from two experiments (mean ± S.E., n = 4).
Individual data points were normalized to the mean intensity of the
60-kDa apical band determined for duplicate samples under control
conditions. The presence (+) or the absence(-) of
extracellular Ca
(1 mM) during the chase is
indicated.
Figure 6:
The regulation of apical protein secretion
by cAMP is dependent on the presence of Cl
. Pulse
labeling of the cells was followed by chase in DMEM (white
bars), in MEBSS (black bars), or in MEBSS gluconate (hatched bars) in the presence or the absence of 200
µM cpt-cAMP as indicated. Phosphor image of the 60-kDa
band from a selected experiment is shown with corresponding mean
intensities of duplicate samples (mean ±
S.D.).
Figure 7:
cAMP
stimulates the sialylation of AT. Pulse labeling and chase were
performed at 20 °C to prevent traffic from the TGN to the cell
surface. Immunoprecipitation and one-dimensional IEF of AT was
performed as described under ``Experimental Procedures.'' a, IEF profiles of AT following the indicated chase periods
(in hours) in the presence or the absence of 200 µM cpt-cAMP. b, corresponding densitograms. Top to bottom on
the gels corresponds to left to right on the densitograms (i.e. alkaline, top/left; acidic, bottom/right). Only
the sialylated forms of AT are shown on this figure, excluding the
large signal associated with more alkaline forms (i.e. earlier
glycosylation intermediates) of the molecule. The total
immunoprecipitated signal (i.e. the signal shown on this
figure plus the signal associated with less mature forms) was equal in
all samples.
Our macroscopic secretion data suggested that protein secretion by
colonic epithelial cells is polarized, because the rate of basolateral
secretion was severalfold greater than apical secretion. However, this
quantitative difference may simply reflect the greater basolateral
surface area of these cells as observed by transmission electron
microscopy (data not shown) rather than a qualitative difference in the
nature of these pathways. In order to assess the profiles of apically
and basolaterally secreted proteins and to obtain qualitative
information regarding the polarity of secretion, the secretory products
were analyzed by SDS-PAGE followed by phosphor imaging and
densitometry. This analysis (Fig. 2a) revealed that
protein secretion by HT29-CL19A cells is indeed qualitatively
polarized. For example, major bands of 105 and 120 kDa were observed
exclusively in the basolateral secretion (Fig. 2a, right, arrows), whereas several lower intensity bands
were characteristic of the apical pathway (Fig. 2a, hollow arrowheads). Interestingly, in spite of the
qualitatively distinct protein profiles of the apical and basolateral
pathways, many bands were present in both, including two major bands
corresponding to 60 and 95 kDa (Fig. 2a, solid
arrowheads). Consistent with our macroscopic data, the apical but
not basolateral secretion of the 60 and 95 kDa proteins increased in
response to cpt-cAMP, as evidenced by the increased density of the
corresponding bands on the phosphor images (Fig. 2a, left). The density of the 60-kDa band that appeared in the
apical medium during a 2-h chase period increased by 2.5-fold (i.e. a 150% increase over the density of the apical control band) in
the presence of 200 µM cpt-cAMP, as determined by phosphor
image analysis (see Fig. 2b and Fig. 3a, second column). The stimulation by
cAMP was not restricted to the 60-kDa protein but was generally
observed for the majority of apically secreted proteins (Fig. 2a; see 95-kDa band and bands indicated by hollow arrowheads).
Fig. 2b illustrates
that the rate of apical secretion was stimulated at cpt-cAMP
concentrations as low as 12.5 µM. Shown is the
dose-response relationship between the concentration of cpt-cAMP and
the relative stimulation of secretion of the 60-kDa band (Fig. 2b). We observed a characteristic two-phase
regulation of apical secretion by cAMP. In the first phase,
concentrations of cpt-cAMP between 0-100 µM caused a
2-fold stimulation of secretion, plateauing at 50 µM (Fig. 2c). In the second phase, concentrations of
cpt-cAMP greater than 100 µM resulted in additional
stimulation of secretion that plateaued at approximately 500 µM cpt-cAMP and that corresponded to a nearly 4-fold stimulation over
control values.
In order to document that cpt-cAMP stimulated apical
protein secretion by a cAMP-specific mechanism, we utilized a panel of
pharmacological mediators that act on the cAMP signaling pathway in
different ways. Fig. 3summarizes our data regarding the
pharmacological profile of regulation, as assessed by phosphor imaging
and densitometry of the 60-kDa protein. 10 µM forskolin
added to both sides or 100 nM VIP added to the basolateral
side (i.e. to the side where its receptors are present) evoked
increases in apical secretion similar in magnitude to that induced by
200 µM cpt-cAMP (see Fig. 3a). VIP added
to the apical side had no effect, and the administration of 500
µM Rp-8-cpt-cAMPS, a cAMP-antagonist, abolished the effect
of basolaterally administered VIP. The same treatments had no
significant effects on the basolateral secretion of the 60-kDa protein (Fig. 3b). Given that the only common feature of these
three agents is their ability to activate the cAMP signaling pathway,
we conclude that cAMP is a specific regulator of apical protein
secretion by HT29-CL19A cells.
Identification of the 60-kDa Secretory Protein as
1-Antitrypsin
In order to begin identifying some of the
proteins secreted by HT29-CL19A cells, we analyzed the secreted
proteins by two-dimensional PAGE and compared the results with the
SWISS two-dimensional PAGE database(27) . The 60-kDa secreted
protein was identified as AT, based on the characteristic appearance of
its multiple sialylated forms, its isoelectric point, and its molecular
weight (Fig. 4, arrow). This finding was verified by
immunoprecipitation using specific antisera and relevant controls (data
not shown). Two-dimensional PAGE analysis that was performed on both
apically and basolaterally secreted proteins confirmed that AT as well
as the 95-kDa protein (Fig. 4, asterisk) is secreted
bidirectionally (only apically secreted proteins are shown). We also
compared the apical secretions with or without stimulation by 200
µM cpt-cAMP by two-dimensional PAGE and failed to observe
any novel protein species that were released by the cells as a
consequence of cAMP stimulation (data not shown).
Apical Protein Secretion Exhibits Linear Kinetics and Is
Ca
-insensitive
In order to further
characterize the apical secretory pathway that is regulated by cAMP in
HT29-CL19A cells, we evaluated the time-course of secretion over an
extended period of time and the Ca
dependence of
polarized protein secretion. Fig. 5(a and b)
illustrates that AT secretion by HT29-CL19A cells follows a linear time
course over 6 h in both the apical (Fig. 5a) and the
basolateral (Fig. 5b) directions. In the presence of
200 µM cpt-cAMP, the basolateral rate of secretion was
unchanged, whereas the same cpt-cAMP concentration elicited a sustained
increase in the rate of apical AT secretion. Fig. 5c demonstrates that the Ca
ionophore, ionomycin,
had no effect on the apical secretion of AT either in the absence or in
the presence of extracellular Ca
. Ionomycin used at
the same concentration evokes a robust potentiation of cAMP-induced
Cl
secretion by these cells (data not shown); thus,
the inability of ionomycin to stimulate protein secretion in these
cells is not due to a lack of effect on intracellular Ca
activity. Neither ionomycin (data not shown) nor the removal of
extracellular Ca
(Fig. 5c) affected
the extent of stimulation by cAMP. Furthermore, incubation of the
monolayers with 20 µM BAPTA-AM for 60 min prior to
metabolic labeling and during the subsequent labeling and chase periods
did not affect either the polarity or the cAMP-dependent regulation of
secretion (data not shown). Thus, apical protein secretion by
HT29-CL19A cells and its regulation by cAMP are largely insensitive to
intra- and extracellular [Ca
].
Constitutive Apical Secretion in HT29-CL19A Cells Is
Regulated by cAMP in a Cl
-dependent
Fashion
Because CFTR (i.e. a cAMP-regulated
Cl
channel) has been implicated in the regulation of
apical membrane traffic (see the introduction), we also examined the
Cl
dependence of the effect of cAMP on apical protein
secretion. Fig. 6demonstrates the results of a representative
experiment in which the secretion assay was performed either in tissue
culture medium (Fig. 6, white bars), a buffered salt
solution containing Cl
(Fig. 6, black
bars), or the latter solution in which Cl
was
replaced by gluconate (Fig. 6, hatched bars), i.e. an anion that is poorly conducted by most Cl
channels(28, 29, 30, 31) . A
stimulation of apical secretion by cAMP was also observed when the
assay was performed in a physiological buffer solution instead of
tissue culture medium (Fig. 6, black bars). However,
when the Cl
in this buffer was replaced by gluconate,
cAMP failed to stimulate secretion. Instead, cAMP inhibited apical
protein secretion in Cl
-depleted cells (Fig. 6, hatched bars).
Cyclic AMP Also Stimulates Protein Sialylation:
Regulation at the TGN
Inhibition of TGN acidification results in
reduced protein sialylation and in a reduced rate of constitutive
protein secretion in HepG2 cells(32) . Because CFTR, a
cAMP-regulated Cl
channel, has been implicated in TGN
acidification in those cells that express
it(33, 34, 35, 36) , we reasoned
that the regulation of constitutive secretion by cAMP in colonic
epithelial cells could be related to a cAMP-dependent regulation of TGN
acidification. AT sialylation can be used as an indirect indicator of
acidification in the TGN, due to the close correlation between TGN
acidification and the efficiency of sialylation(32) . In order
to examine the effect of cAMP on the rate of sialylation in HT29-CL19A
cells, we analyzed the extent of AT sialylation at 20 °C when newly
synthesized AT is entrapped within the TGN(8, 37) ,
thus allowing the analysis of AT sialylation within a single
intracellular pool. Fig. 7a demonstrates the results of
a representative experiment in which AT was immunoprecipitated from
cell lysates and analyzed by one-dimensional IEF. One-dimensional IEF
allows the estimation of the extent of sialylation based on the
pronounced acid shift in the isoelectric point of AT as a consequence
of the addition of successive sialic acid residues. When the chase was
performed in the presence of 200 µM cpt-cAMP, a shift
toward the most acidic forms of AT was observed (compare second and fourth lanes of Fig. 7a). Fig. 7b shows densitograms corresponding to the second and fourth lanes of Fig. 7a (top to bottom on gels corresponds to left to right on
densitogram). The pronounced increase in the density of bands
representing the most acidic (i.e. sialylated) forms of AT in
the cAMP-treated sample verifies that cAMP stimulates sialylation
within the TGN of HT29-CL19A cells.
DISCUSSION
cAMP Regulates Constitutive Membrane Traffic from the
TGN to the Apical Cell Surface
Our results indicate that cAMP
regulates apical but not basolateral protein secretion in colonic
epithelial cells. The specificity of this regulation by cAMP was
documented using a panel of secretagogues including a physiological
activator of the cAMP pathway (i.e. VIP), the stimulatory
effect of which was blocked by a stereo-specific inhibitor of cAMP. The
apical secretory pathway in colonic epithelial cells is at least an
order of magnitude more sensitive to cAMP analogs than that reported
for Madin-Darby canine kidney cells(8, 10) . It
remains to be determined if this differential sensitivity of the
biosynthetic pathway to cAMP is due to differences in cAMP metabolism
or signaling between these cell types (e.g., differences in
the expression levels of phosphodiesterase or cAMP-dependent protein
kinase isoforms) or due to different downstream effector mechanisms. On the basis of the following considerations we conclude that cyclic
AMP regulates protein secretion by stimulating constitutive membrane
traffic to the apical cell surface: (a) the linear kinetics of
secretion under both control and stimulated conditions, (b)
the insensitivity of secretion to Ca
, a classical
stimulator of regulated secretory pathways, and (c) the lack
of accumulation of fully processed secretory material in well defined
storage granules in these cells (data not shown). The observed
stimulation of apical protein secretion by cAMP is not due to an
elevated rate of protein synthesis, because that would have affected
the basolateral rate of secretion. In addition, we determined that
there is no further incorporation of radiolabeled amino acids into
trichloroacetic acid precipitable material following the initial pulse
period (i.e. during the time period when secretagogues are
present; data not shown). The polarity of regulation indicates that TGN
to apical cell surface traffic is at least one site of regulation by
cAMP (i.e. there are no known polarized compartments within
the biosynthetic pathway proximal to the TGN). This notion is also in
agreement with the finding that the regulatory subunit (RII) of
cAMP-dependent protein kinase associates with the TGN in epithelial
cells(38) . A feasible mechanism by which cAMP could regulate
constitutive membrane traffic is the stimulation of secretory vesicle
formation at the TGN, similar to the regulation of constitutive
secretion in rat basophilic leukemia cells by protein kinase
C(11) . In rat basophilic leukemia cells the regulation of
constitutive secretion by protein kinase C involves a stimulation of
ARF binding to Golgi membranes that drives coat formation and vesicle
budding from the Golgi(39) . AT secretion in HT29-CL19A cells
is likely ARF-dependent, because we observed that brefeldin A (i.e. a drug that inhibits the exchange of guanine nucleotides bound to
ARF(40) ) completely blocks protein secretion by HT29-CL19A
cells (data not shown). Therefore, it is conceivable that mechanisms
that regulate ARF binding to the TGN in a cAMP-dependent manner could
modulate constitutive secretion in HT29-CL19A cells (see also below).
TGN Acidification as a Possible Mechanism Underlying the
Regulation of Constitutive Secretion by Cyclic AMP
Both the
rates of constitutive secretion and of AT sialylation have been shown
to be dependent on TGN acidification in HepG2 cells. Namely,
concanamycin B, an inhibitor of vacuolar ATPases, decreased the rate of
AT secretion and sialylation in these cells(32) . The
dependence of sialylation on TGN acidification is probably due in part
to the acidic pH optimum of sialic acid transferase(41) ;
however, the molecular mechanisms that link acidification to secretion
are unknown. Interestingly, ARF binding to microsomal membranes in
vitro has been shown to be dependent on vesicular
acidification(42) . The acidification dependence of ARF binding
to membranes could account for the inhibitory effect of concanamycin B
on the rate of constitutive secretion in HepG2 cells and provides a
feasible mechanism by which cAMP could regulate secretion (i.e. by regulating TGN acidification) in HT29-CL19A cells. Our
observation of a stimulatory effect of cAMP on the rate of sialylation
of AT that was entrapped in the TGN using a 20 °C temperature block
is consistent with the notion that cAMP enhances TGN acidification in
HT29-CL19A cells (see the connection between sialylation and
acidification above). Vesicular acidification requires a mechanism to
shunt the membrane potential generated by the vacuolar
H
-ATPase that otherwise limits the accumulation of
protons. A Cl
conductance of vesicular membranes can
function as such a shunt mechanism, and a cAMP-regulated Cl
conductance has been shown to regulate endosome acidification in
kidney epithelial cells(43) . CFTR, a cAMP-regulated
Cl
channel, has been implicated in TGN acidification
and in protein sialylation in epithelial
cells(33, 35, 36) . Therefore, CFTR is a
reasonable candidate for regulating TGN acidification in a
cAMP-dependent manner in HT29-CL19A cells. The Cl
dependence of the regulation of apical constitutive secretion by
cAMP in HT29-CL19A cells that we observed is consistent with the role
of a cAMP-regulated Cl
channel in this regulation.
Possible Origins for the Polarity of
Regulation
The TGN of normal rat kidney cells is composed of
multiple tubules, as indicated by three-dimensional reconstruction of
high voltage electron microscopy images. Each tubule corresponds to an
already polarized entity, and all vesicles budding from a single tubule
are coated with only one of two morphologically distinct coat
structures observed in the TGN of these cells(44) . It cannot
be determined from this morphological analysis whether membrane
proteins destined for a single plasma membrane domain (i.e. apical or basolateral) of polarized epithelial cells are
restricted to distinct tubules within the TGN. Nevertheless, if such a
polarity exists within the TGN, then cAMP could selectively stimulate
secretory vesicle budding from a TGN subcompartment that is dedicated
to apical delivery. Such a mechanism would be consistent with a role
for CFTR in this process, i.e. CFTR is targeted to the apical
domains of HT29-CL19A cells(21) . Alternatively, if the budding
of basolateral and apical secretory vesicles takes place from a common
compartment within the TGN, the cAMP-dependent recruitment of a
direction-specific cytosolic component to apically destined vesicles
that are budding from the TGN could determine the polarity of
regulation. The cytoskeletal motor protein, dynein, which has been
shown to participate selectively in the apical delivery of transport
vesicles, could be such a direction-specific cytosolic factor (45) . It remains to be determined if budding from an already
polarized TGN domain, the recruitment of direction-specific cytosolic
factors or both are responsible for the polarity of regulation of the
constitutive secretory pathway by cyclic AMP.
Concluding Remarks
Our data demonstrate the
existence of a high sensitivity, cAMP-dependent regulation of apical
secretion by colonic epithelial cells. Thus, apical protein secretion
and electrolyte and fluid secretion are regulated coordinately by cAMP
in colonic crypt cells. We identified the predominant secretory product
of HT29-CL19A cells as
1-antitrypsin and provided compelling
evidence that the regulation of protein secretion by cAMP in these
cells represents the stimulation of constitutive membrane traffic from
the TGN to the apical cell surface. Our data are consistent with the
notion that protein secretion via the constitutive secretory pathway
takes place via the stimulation of constitutive vesicle generation (i.e. budding) from the TGN, in contrast to the regulation of
secretory granule consumption (i.e. regulated exocytosis) by
glandular cells. We propose that the regulation of protein sialylation
and apical protein secretion by cAMP may have a common origin, i.e. regulated TGN acidification. This hypothesis is consistent with
the genetic evidence that cyclic AMP-regulated CFTR Cl
channels regulate protein sialylation and TGN acidification in
epithelial tissues and is supported by the Cl
dependence of the regulation of protein secretion. Defining the
molecular basis for the regulation of protein processing and secretion
by cyclic AMP, including the possible role of CFTR in this process,
should contribute to our understanding of how polarized epithelial
cells modulate mucosal homeostasis.