Clathrin in gastric acid secretory (parietal)
cells: biochemical characterization and subcellular
localization
Curtis T.
Okamoto1,
Joseph G.
Duman2,
Kamala
Tyagarajan2,
Kent L.
McDonald2,4,
Young Y.
Jeng1,
Jeana
McKinney1,
Trudy M.
Forte3, and
John G.
Forte2
1 Department of Pharmaceutical Sciences, School of Pharmacy,
University of Southern California, Los Angeles 90089-9121; and
2 Department of Molecular and Cell Biology, 3 Lawrence
Berkeley Laboratory, and 4 Electron Microscope Lab, University
of California, Berkeley, California 94720
 |
ABSTRACT |
Clathrin from
H-K-ATPase-rich membranes derived from the tubulovesicular compartment
of rabbit and hog gastric acid secretory (parietal) cells was
characterized biochemically, and the subcellular localization of
membrane-associated clathrin in parietal cells was characterized
by immunofluorescence, electron microscopy, and immunoelectron
microscopy. Clathrin from H-K- ATPase-rich membranes was determined
to be comprised of conventional clathrin heavy chain and a predominance
of clathrin light chain A. Clathrin and adaptors could be induced to
polymerize quantitatively in vitro, forming 120-nm-diameter basketlike
structures. In digitonin-permeabilized resting parietal cells, the
intracellular distribution of immunofluorescently labeled clathrin was
suggestive of labeling of the tubulovesicular compartment. Clathrin was
also unexpectedly localized to canalicular (apical) membranes, as were
-adaptin and dynamin, suggesting that this membrane domain of
resting parietal cells is endocytotically active. At the
ultrastructural level, clathrin was immunolocalized to canalicular
and tubulovesicular membranes. H-K-ATPase was immunolocalized to
the same membrane domains as clathrin but did not appear to be enriched
at the specific subdomains that were enriched in clathrin. Finally, in
immunofluorescently labeled primary cultures of parietal cells, in
contrast to the H-K-ATPase, intracellular clathrin was found not to
translocate to the apical membrane on secretagogue stimulation. Taken
together, these biochemical and morphological data provide a framework
for characterizing the role of clathrin in the regulation of membrane
trafficking from tubulovesicles and at the canalicular membrane in
parietal cells.
hydrogen-potassium-adenosinetriphosphatase; apical membrane
recycling; tubulovesicles; dynamin; gastric microsomes
 |
INTRODUCTION |
THE REGULATION OF THE
TRAFFICKING of membrane transporters is becoming widely
recognized as a basic mechanism by which cells, epithelial cells in
particular, regulate solute transport (8). The gastric
parietal (oxyntic) cell represents a model system in which to study the
means by which solute transport can be regulated by the vesicular
trafficking of a membrane transporter, the H-K-ATPase (13,
21). In the resting parietal cell, the gastric H-K-ATPase is
sequestered in an intracellular system of tubulovesicular membranes. When cells are stimulated to secrete HCl, the tubulovesicular membranes
fuse with the canalicular (apical) membrane, thus delivering the
H-K-ATPase to the apical membrane. When the stimulus is removed, the
H-K-ATPase is retrieved from the canalicular membrane, and the
tubulovesicular compartment is reestablished (22).
The tubulovesicular compartment of the parietal cell has recently been
demonstrated to contain key components of the essential machinery to
regulate the trafficking of the H-K-ATPase, such as the small GTPases
rab11 (10) and rab25 (26), and proteins implicated in vesicular docking/fusion, such as syntaxin 3 and vesicle-associated membrane protein (VAMP) (10, 41). Some of these components, including rab11, show secretagogue-stimulated changes in subcellular membrane localization (11), and
additional functional studies suggest that rab11 is a key
regulator of H-K-ATPase trafficking (20). In addition, a
tyrosine-containing motif in the cytoplasmic domain of the
-subunit of the heterodimeric H-K-ATPase has been implicated to
serve as a sorting signal for the reinternalization of the H-K-ATPase
from the apical membrane on cessation of acid secretion
(17). The motif in the
-subunit is strikingly similar to the internalization signal in the transferrin receptor that allows
the transferrin receptor to interact with clathrin adaptor protein-2
(AP-2) at the plasma membrane. Clathrin and an AP-1 clathrin adaptor
have been recently identified on tubulovesicles (39).
Moreover, the AP-1 adaptor and the H-K-ATPase appear to interact, as
shown by their copurification from tubulovesicles solubilized with
a nondenaturing detergent (39). Thus significant progress in recent years has been made in cataloging components of the
molecular machinery ostensibly involved in the regulation of H-K-ATPase trafficking.
In the present study the role of clathrin in the regulation of
trafficking of the H-K-ATPase in gastric parietal cells was investigated by biochemical and morphological approaches. Clathrin from
H-K-ATPase-rich membranes was further characterized immunologically and
by mass spectrometry. Clathrin and adaptors formed baskets in vitro.
Immunofluorescent labeling and electron-microscopic localization of
clathrin in resting parietal cells suggest a role for clathrin in the
regulation of membrane traffic between tubulovesicular membranes and
the canalicular membrane. Immunofluorescent labeling of clathrin in
secretagogue-stimulated parietal cells suggests that clathrin is not
translocated to the apical membrane with the H-K-ATPase and may
therefore play a direct role in H-K-ATPase trafficking at another stage
of the secretory cycle.
 |
MATERIALS AND METHODS |
Materials.
Anti-clathrin heavy chain monoclonal antibodies (MAbs) X-22
(36) and TD.1 (36); the anti-clathrin light
chain MAbs X-16 (1), LCB.1 (1), and CON.1
(36); and anti-
-adaptin MAb AP.6 (15) were
obtained from two sources: as kind gifts of Dr. Frances Brodsky
(University of California, San Francisco) and from X-22, TD.1, CON.1,
and AP.6 hybridoma cell culture supernatants. The hybridomas were
purchased from the American Type Culture Collection. 4,4-Difluoro-4-bora-3a,4a-diaza-s-indacene (BODIPY-FL)-phallacidin and
rhodamine-phalloidin were purchased from Molecular Probes (Eugene, OR)
and were used according to the vendor's instructions. Anti-dynamin MAb
and anti-clathrin heavy chain MAb 23 were purchased from Transduction
Labs (Lexington, KY). Anti-
-adaptin MAb 100/3 was purchased from
Sigma Chemical (St. Louis, MO). Horseradish peroxidase (HRP)-conjugated
goat anti-mouse antibody was purchased from Bio-Rad (Hercules, CA).
Enhanced chemiluminescence (ECL) reagents were purchased from Pierce
Chemical (Rockford, IL). Cy3-conjugated and FITC goat anti-mouse
antibodies were purchased from Jackson ImmunoResearch Labs (West
Grove, PA). ProLong antifade mounting medium was purchased from
Molecular Probes. Vectashield antifade mounting medium was purchased
from Vector Laboratories (Burlingame, CA). Digitonin was purchased
from Boehringer (Indianapolis, IN). Cimetidine, 8-bromo-cAMP, and
collagenase were purchased from Sigma Chemical. SCH-28080 was a gift
from Dr. J. Kaminsky (Schering, Kenilworth, NJ). All other materials
were reagent grade. Purified gastric microsomes were prepared as
described previously (53). Crude clathrin-coated vesicles
(CCV) from brain were purified according to the protocol of Pearse and
Robinson (40).
SDS-PAGE and related procedures.
Protein determinations were made using the bicinchoninic acid protein
assay (Pierce Chemical). SDS-PAGE was performed according to Laemmli
(32). Because of the sensitivity of the H-K-ATPase to
extended boiling, samples containing H-K-ATPase were boiled for only 2 min in sample buffer before they were loaded on gel. Samples without
H-K-ATPase were boiled for 5 min.
For Western blotting, primary antibodies were diluted in Tris-buffered
saline-0.02% Tween 20 as follows: 1:5,000 for anti-clathrin MAb TD.1,
1:1,000 for anti-light chain A MAb X-16, 1:1,000 for anti-light chain B
MAb LCB.1, and 1:1,000 for anti-light chain MAb CON.1. Hybridoma cell
culture supernatants containing TD.1 and CON.1 were also used neat for
Western blotting. Goat anti-mouse-HRP secondary antibody was used at
1:20,000 dilution. Blocking of nitrocellulose was done in 1-5%
nonfat milk or 0.2% BSA in Tris-buffered saline. HRP was detected by
ECL, and the signal was visualized on Kodak Bio-Max X-ray film.
Hydroxyapatite chromatography of coat proteins from purified
gastric microsomes.
Clathrin coat proteins and other peripheral membrane proteins were
stripped from purified gastric microsomes (starting with 3-30 mg
of microsomal membrane proteins) essentially according to the protocol
of Keen et al. (29). The stripped proteins were dialyzed
overnight against three changes of a solution of 125 mM mannitol, 40 mM
sucrose, 1 mM EDTA, and 5 mM PIPES buffer, pH 6.7 (MSEP). The dialysate
was applied to a 5-ml hydroxyapatite column equilibrated with MSEP, and
proteins were eluted by a stepwise gradient of 12 ml each of 10, 100, 200, and 400 mM sodium phosphate, pH 7.0. NaN3 was added to
the eluted fractions, and these fractions were stored at 4°C. For SDS
gels and Western blots, proteins in 0.3- to 1.2-ml aliquots were
precipitated with 10% TCA and resuspended in SDS gel buffer before electrophoresis.
In-gel digestion.
A slightly modified in-gel digestion method from Rosenfeld et al.
(44) was performed. Protein bands (~5 µg of protein)
were minced, and the gel slices were destained with three washes of 50% acetonitrile-25 mM NH4HCO3 (~10 min
each). The destained gel pieces were dried in a Speedvac (Savant,
Farmingdale, NY) for 30 min and then rehydrated in 50 µl of 25 mM
NH4HCO3 (pH 8.0) with 0.01 µg/µl trypsin.
The slices were overlaid with 50 µl of 25 mM
NH4HCO3 and incubated for 15 h at 37°C.
Peptides were recovered by three extractions of the digestion mixture
with 50% acetonitrile-5% trifluoroacetic acid. All supernatants were
pooled, concentrated to 5 µl in a Speedvac, and brought back up to 25 µl in 50% acetonitrile-5% trifluoroacetic acid. The peptide mix was
stored at
20°C until further analysis.
Matrix-assisted laser desorption delayed extraction reflection
time of flight mass spectrometry of clathrin peptides.
Aliquots (
) of unseparated tryptic digests were
cocrystallized with
-cyano-4-hydroxycinnamic acid and analyzed using
a matrix-assisted laser desorption delayed extraction reflection
(MALDI) time of flight (TOF) mass spectrometry (MS) instrument
(Perceptive Biosystems, Voyager Elite mass spectrometer, Framingham,
MA) equipped with a nitrogen laser at the University of California, San
Francisco, Mass Spectrometry Facility. Measurements were performed in a
positive ionization mode. All MALDI spectra were externally calibrated
using a standard peptide mixture. For postsource decay (PSD) spectra,
tryptic peptides were fractionated by reverse-phase microbore HPLC. PSD
spectra were acquired on a TofSpec SE MALDI-TOF MS (Micromass,
Manchester, UK) with a nitrogen laser and operated in the reflectron mode.
Database searches for protein identification.
Experimentally determined masses were used for database interrogation
with use of MS-Fit software (16, 43). PSD data
interrogation was performed using MS-Tag. Both software programs were
developed at the University of California, San Francisco, Mass
Spectrometry Facility and are available on the World Wide Web at
http://prospector.ucsf.edu. Protein searches were carried out in the
National Center for Biotechnology Information protein database and the
SwissProt database by using a protein molecular weight of
150-250 kDa, a peptide mass tolerance of 0.5 Da, and a minimum
match of 50% of peptides observed in the total digest.
In vitro polymerization of clathrin and negative-stain electron
microscopy.
For in vitro polymerization of clathrin coats, 100 µg of purified
gastric microsomes were stripped as described above. The stripped
proteins were dialyzed overnight against three changes of 20 mM MES, pH
6.5, and either 1 mM sodium EGTA or 2 mM CaCl2 to analyze
the calcium dependency of clathrin and clathrin adaptor polymerization.
The dialysate was spun at 300,000 g for 20 min in a
miniultracentrifuge (model RC M120EX, Sorvall). The pellet was
resuspended directly into SDS sample buffer, and proteins in the
supernatant were precipitated with 10% TCA. The entire samples were
loaded onto SDS gels for analysis by Coomassie blue staining and
Western blot. Alternatively, 1-4 ml of the hydroxyapatite fraction
enriched in tubulovesicular clathrin and AP-1 adaptors (i.e., proteins
eluted by 200 mM sodium phosphate) were dialyzed overnight against one
change of 20 mM MES, pH 6.5, and two changes of 20 mM MES, pH 6.5, and
either 1 mM sodium EGTA or 2 mM CaCl2. The dialysate was
spun at 300,000 g for 20 min. The supernatant was removed,
and proteins in the supernatant were precipitated in 10% TCA. The
precipitated proteins were resuspended in SDS gel buffer to the same
volume as the 300,000-g pellet. Equal volumes of fractions
were run on gels for Coomassie blue staining or Western blotting. The
efficiency of clathrin and clathrin adaptor polymerization was
quantitated by scanning Coomassie blue-stained gels or films of
immunoblots at 300 dpi with a UMAX Vista 6E color scanner, and the
digitized images were processed using the NIH Image version 1.55 program. For Western blots developed by ECL, films exposed for
different times were analyzed to ensure that the signals from bands
fell within the linear range and were linear relative to each other.
Samples of clathrin baskets for electron-microscopic analyses were
prepared using the same protocol, except the 300,000-g pellet was resuspended in a small volume of MES-CaCl2
buffer. Aliquots of the resuspended pellet were applied to
Formvar-coated grids, negatively stained with 1% uranyl acetate, and
viewed in a JEOL 100CX electron microscope.
High-pressure freezing of isolated gastric glands for
transmission electron microscopy and immunogold electron microscopy.
Gastric glands were isolated from rabbit stomach as previously
described (54). Isolated gastric glands were incubated at 37°C in a balanced salt solution (54) containing
10
4 M cimetidine as the H2 receptor
antagonist to the natural secretagogue histamine. Aliquots of glands
were sedimented by light centrifugation and transferred to the 100-µm
deep well of a type A high-pressure freezing planchette (Ted Pella,
Redding, CA) and then frozen in a high-pressure freezing machine (model
HPM 010, Bal Tec), as described by McDonald (35). Frozen
samples were freeze substituted in 2% osmium tetroxide plus 0.1%
uranyl acetate in acetone for morphological studies or in 0.2%
glutaraldehyde plus 0.1% uranyl acetate in acetone for
immunolocalization studies. Cells were kept in fixative at dry ice
temperature (approximately
78°C) for 3 days and then warmed to room
temperature over 12 h. After three 10-min rinses in pure acetone,
osmium-fixed cells were infiltrated and embedded in Epon-Araldite
resin. Glutaraldehyde-fixed cells were also rinsed three times in
acetone, infiltrated, and embedded in LR White resin. Thin
sections (60-70 nm thick) were cut on a Reichert Ultracut E
(Leica, Deerfield, IL) or an MTX (RMC, Tucson, AZ) ultramicrotome,
poststained with uranyl acetate and lead citrate, and observed in a
JEOL 100CX transmission electron microscope operating at 80 kV.
Immunogold staining protocol.
For immunogold labeling, LR White sections were picked up on 100-mesh
nickel grids coated with Formvar film and carbon, incubated in blocking
buffer (5% BSA, 0.1% fish gelatin, and 0.05% Tween 20 in PBS) for 30 min, and then incubated with primary antibodies diluted in blocking
buffer for 1.5-2 h. MAb X-22 was used as undiluted cell culture
supernatant; MAb 2G11 cell culture supernatant was diluted 1:5.
The commercially procured MAb23 was diluted 1:100. Sections were
rinsed in PBS-Tween, and then PBS and incubated for 1 h in
secondary antibodies conjugated to 10-nm gold particles [goat
anti-mouse IgG F(ab')2 (H + L); Ted Pella] diluted
1:20 in blocking buffer. Sections were washed as described above, fixed in 0.5% glutaraldehyde in PBS for 5 min, and rinsed in PBS and water.
Sections were poststained in 2% uranyl acetate for 5 min and in lead
citrate for 3 min. Identically treated samples were stained with
secondary antibody only as controls and revealed no labeling pattern.
Fractionation of sucrose density gradient-purified gastric
microsomes on discontinuous glycerol velocity gradients.
The additional fractionation of sucrose density gradient-purified
gastric microsomes on discontinuous glycerol velocity gradients was
adapted from Salem et al. (45). Gastric sucrose microsomes (200 µg) sedimenting at the 32% barrier on sucrose density gradients were diluted to 0.4 mg/ml in MSEP. It was layered on top of a discontinuous glycerol gradient comprised of 0.5 ml each of 20, 40, and
80% glycerol, diluted in MSEP. The sample was centrifuged in an
RP55S-485 Sorvall swinging bucket rotor at 55,000 rpm for 30 min in a
Sorvall RC M120EX miniultracentrifuge. The membranes sedimenting at
each interface were collected, diluted to 1.0 ml with MSEP, and
recentrifuged at 150,000 g for 40 min. The pellets were
resuspended in SDS gel sample buffer and analyzed by SDS-PAGE and
Western blot. For the membranes sedimenting at the 20 and 40% glycerol
layers, the entire samples were loaded onto the gels; for membranes
sedimenting at the 80% layer, one-eighth to one-fourth of the sample
was loaded onto the gels.
Immunofluorescent staining of isolated gastric glands and
cultured parietal cells.
Isolated glands were incubated with 10
5 M cimetidine
(resting) for 30 min at 37°C in a total of 10 ml of a suspension of a 1:10 dilution of glands. In the case of staining with anti-dynamin MAb,
glands were fixed in 3.7% formaldehyde in PBS for 20 min and then
permeabilized in 0.5% Triton X-100-PBS for 20 min, both at room
temperature. After fixation and permeabilization, glands were blocked
for 30-60 min at room temperature with a solution of 1% BSA and
0.066% (wt/vol) fish skin gelatin diluted into PBS. Glands were
incubated with anti-dynamin MAb (diluted 1:100 in blocking
solution) for 2 h at room temperature. Glands were then incubated
with Cy3-conjugated secondary antibody diluted 1:500 into PBS-0.05%
Tween 20 for 60 min at room temperature. Glands were counterstained
with BODIPY FL-phallacidin simultaneously with the secondary antibody.
Glands were then washed and mounted in ProLong or Vectashield antifade medium.
When glands were probed for clathrin heavy chain (MAb X-22) and
-adaptin (MAb AP.6), they were permeabilized with digitonin (33) before fixation and staining. After incubation with
cimetidine, glands were permeabilized for 5 min with ice-cold 0.004%
digitonin diluted in 12.5 mM HEPES-KOH, 50 mM PIPES-KOH, 1 mM
MgSO4, and 4 mM EGTA, pH 7.0. Subsequent steps for
fixation, blocking, and staining (with MAb X-22 or AP.6) were similar
to those described above.
Cultures enriched in parietal cells were obtained by a modification of
the method of Chew et al. (14). The minced mucosa was
digested in MEM supplemented with HEPES buffer (MEM-HEPES) containing
0.125 mg/ml collagenase (Sigma Chemical) and 0.25 mg/ml BSA at 37°C
for ~30 min. The reaction was stopped by threefold dilution of the
digestion solution with MEM-HEPES. Because of their large size,
relatively intact gastric glands settled out in 10-15 min, leaving
individual cells suspended in the medium. The suspended cells were
strained through nylon mesh and washed three times with MEM-HEPES.
Cells were next incubated for 30 min in medium B (DMEM-F-12;
GIBCO, Rockville, MD) supplemented with 20 mM HEPES, 0.2% BSA, 10 mM
glucose, 8 nM EGF, 1× SITE (selenite, insulin, and transferrin) medium
(Sigma Chemical), 1 mM glutamine, 100 U/ml penicillin-streptomycin, 400 µg/ml gentamicin sulfate, 25 µg/ml amphotericin B, 15 µg/l
geneticin, and 20 µg/ml novobiocin, pH 7.4, to prevent yeast
infection. Cells were then plated onto coverslips coated with Matrigel
(Collaborative Biomedical, Franklin Lakes, NJ) and incubated at 37°C
in culture medium A (medium B without
amphotericin B).
To effect functional alterations in secretory state, parietal cell
cultures were held in a resting state through the addition of
cimetidine to 100 µM or maximally stimulated through addition of
histamine and IBMX (final concentrations 100 and 30 µM,
respectively). In some cases, cells were treated with the proton pump
inhibitor SCH-28080 (5 µM) in addition to the secretagogues. After
addition of drugs, cultures were incubated for 25 min at 37°C. Cells
were then fixed with 3.7% formaldehyde in PBS for 20 min,
permeabilized in 0.3% Triton X-100 in PBS for 15 min, and blocked in
2% BSA in PBS for 15 min. Fixed, permeabilized cells were probed with a variety of antibodies. H-K-ATPase was detected by 1 h of
incubation with a 1:3 dilution of culture supernatant of MAb 2G11, a
mouse MAb against its
-subunit (Affinity Bioreagents, Golden, CO). Clathrin was probed using anti-clathrin heavy chain MAb X-22 and MAb 23 and anti-clathrin light chain MAb CON.1. When the cells were stained
with MAb X-22, 5.0 M urea was included in the permeabilization step.
The clathrin AP-1 adaptor was probed using anti-
-adaptin MAb 100/3.
The primary mouse MAbs were detected by a subsequent 30-min incubation
with FITC-labeled goat anti-mouse IgG. All antibody dilutions were made
in PBS containing 2% BSA. In most cases, cells were also stained for
F-actin by coincident incubation with 80 nM rhodamine-labeled
phalloidin. Coverslips were supported on slides by grease pencil
markings and mounted in Gel/Mount (Biomeda, Foster City, CA).
Stained glands and cells were examined by conventional fluorescence
microscopy or by confocal microscopy. Immunofluorescent images were
captured with a Nikon Microphot FX-2 microscope equipped with a
Photometrics Sensys coupled charge device camera by use of Innovision
software. Confocal images of glands were taken with a Bio-Rad MRC-1024
instrument equipped with a krypton-argon laser with use of a Zeiss
Axioplan microscope and a ×60 plan-Apo 1.4 NA oil immersion objective
or with a Nikon PCM quantitative measuring high-performance confocal
system attached to a Nikon TE300 Quantum upright microscope.
 |
RESULTS |
Immunodetection of clathrin heavy chain and light chains on Western
blots of purified gastric microsomes.
The MAb TD.1 was used to confirm the presence of clathrin in density
gradient fractions of H-K-ATPase-rich gastric microsomes derived from
tubulovesicles of parietal cells. A Coomassie blue-stained gel of the
purified microsomal fractions from rabbit gastric mucosa is shown in
Fig. 1A (lanes 1 and 2), with the position of the
-subunit of the
H-K-ATPase indicated. The Western blots in Fig. 1B show that
clathrin heavy chain, as recognized by MAb TD.1, is present in
microsomes purified from rabbit gastric mucosa, confirming earlier
identification of clathrin heavy chain in these H-K-ATPase-rich
membranes (39).

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Fig. 1.
Immunodetection of clathrin heavy chain on purified
gastric microsomes of parietal cells. A: Coomassie
blue-stained SDS gel of sucrose density gradient-purified gastric
microsomes from rabbit gastric mucosa (24 µg protein/lane). Position
of the H-K-ATPase -subunit (HK ) is indicated. Lane 1,
microsomes purified at the 27% sucrose layer; lane 2,
microsomes purified at the 32% layer. Apparent molecular masses are
indicated, and the identity of the markers is as follows: myosin,
-galactosidase, phosphorylase B, BSA, ovalbumin, and carbonic
anhydrase. B: Western blot of purified gastric microsomes
(15 µg protein/lane) with anti-clathrin heavy chain monoclonal
antibody (MAb) TD.1. Lane 1, 27% sucrose layer; lane
2, 32% sucrose layer. Immunoreactivity was detected by enhanced
chemiluminescence (ECL) with a 2-min exposure to film. Position of
prestained molecular mass markers (myosin and -galactosidase) is
shown. C: comparison of clathrin content of crude
clathrin-coated vesicles (CCVs) from brain and purified gastric
microsomes. Coomassie blue-stained SDS gel of crude CCVs from hog brain
and purified gastric microsomes (10 µg protein/lane) is shown.
Lane 1, crude CCVs from hog brain; lane 2, hog
gastric microsomes purified at the 32% layer. D: Western
blot with MAb TD.1. Lane 1, CCVs from hog brain (5 µg);
lane 2, hog gastric microsomes purified at the 32% layer (5 µg). Signal detected by ECL with a 2-min exposure.
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|
For comparison, the relative amounts of clathrin in crude CCVs from hog
brain and hog gastric microsomes were assessed in Coomassie
blue-stained gels (Fig. 1C). Clathrin heavy chain, migrating as a 160-kDa protein band, is the major protein in the brain CCVs (lane 1). In contrast, H-K-ATPase is the major protein in
gastric microsomes, whereas some ~160-kDa migrating protein bands are also visible (lane 2). The relative amounts of clathrin
associated with purified gastric microsomes and brain CCVs were
estimated from Western blots probed with MAb TD.1 (Fig. 1D).
Brain CCVs (lane 1) contain ~20 times more clathrin than
gastric microsomes (lane 2). One of the reasons that brain
CCVs may possess a higher specific content of clathrin than gastric
microsomes is that the preparation of brain CCVs includes an extraction
step with 1% Triton X-100. This treatment would be expected to
solubilize most membrane proteins and therefore deplete the cargo
proteins for CCVs, but clathrin and associated proteins would remain
and, therefore, be enriched, in pelletable complexes. When crude or
purified gastric microsomes are subjected to the same extraction
procedure, clathrin is recovered but not enriched in the Triton
X-100-insoluble pellet, even though the H-K-ATPase is quantitatively
extracted (data not shown). Thus there appear to be differences between
clathrin on brain CCVs and purified gastric microsomes that confer
greater stability of clathrin on brain CCVs to the Triton X-100
extraction procedure.
Purified hog gastric microsomal membrane fractions were also probed for
clathrin light chains. The immunodetection of clathrin light chain A
(LCa) and light chain B (LCb) in gastric microsomes and brain CCVs was
performed with the MAbs X-16 (LCa specific) and LCB.1 (LCb specific)
(1). LCa (Fig.
2A) is clearly present in
gastric microsomes (lane 2). Although LCb is not detectable in gastric microsomes in the immunoblot shown in Fig. 2B
(lane 2), it becomes detectable when higher amounts of
microsomal protein are assayed. The light chains from brain appear to
be of higher relative molecular weight on SDS gels than their
counterparts in gastric microsomes, possibly because of the presence of
neuron-specific inserts in LCa and LCb (9, 30).

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Fig. 2.
Identification of clathrin light chain isoforms on
purified gastric microsomes by immunoblotting. A: Western
blot of crude CCVs from hog brain (3 µg) and hog gastric microsomes,
32% layer (10 µg), with anti-light chain A MAb X-16. Signal was
detected by ECL with a 2-min exposure. B: Western blot of
crude CCVs from hog brain (3 µg) and hog gastric microsomes, 32%
layer (10 µg), with anti-light chain B MAb LCB.1. Signal was detected
by ECL with a 1-min exposure. C: Western blot of crude CCVs
from hog brain (lanes 1 and 2) and hog gastric
microsomes (lanes 3 and 4) with anti-light chain
(common) MAb CON.1. Lane 1, 3 µg of CCVs; lane
2, 5 µg of CCVs; lane 3, 10 µg of 32% gastric
microsomes; lane 4, 20 µg of 27% gastric microsomes.
Positions of light chains A and B (LCa and LCb) are indicated. Signal
was detected by ECL with a 15-s exposure.
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By immunoblotting with MAb CON.1, which recognizes both light chains
equally well (36), the ratio of LCa to LCb can be
determined in a single sample. The immunoblot in Fig. 2C
suggests that LCa is the predominant light chain associated with
gastric microsomes (lanes 3 and 4), and these
results are consistent with the results from the immunoblots by use of
the isoform-specific MAbs (Fig. 2, A and B). From
densitometric analyses of these Western blots, the ratio of LCa to LCb
is 3.1:1 ± 1.1 (SD, n = 11 blots from 8 different animals).
Fractionation of gastric microsomal clathrin by hydroxyapatite
chromatography.
Purified gastric microsomes were stripped of their clathrin coats (and
other peripheral membrane proteins) by incubation with 0.5 M
Tris · HCl and 2 mM EDTA. After dialysis in a
low-ionic-strength buffer, the stripped proteins were applied to a
hydroxyapatite column, and proteins were eluted by a stepwise sodium
phosphate gradient. A major protein, eluted by the addition of 0.2 M
sodium phosphate and clearly visible on Coomassie blue-stained gels
(Fig. 3A, lane 4), was
identified as clathrin heavy chain on the basis of its immunoreactivity
with anti-clathrin heavy chain MAb 23 (Fig. 3B) and TD.1
(not shown). Clathrin light chains (with approximately the same ratio
of LCa to LCb as in isolated membranes) were also detected in Western
blots of the 0.2 M sodium phosphate eluate (Fig. 3C). From
densitometric analyses of Coomassie blue-stained gels, clathrin heavy
chain was determined to comprise 14 ± 3% (SD, n = 10 independent preparations) of the total protein eluted in this
fraction. By comparison, in crude CCVs isolated here (Fig. 1C), clathrin comprises ~12% of the total protein. If we
assume that the recovery of clathrin heavy chain by hydroxyapatite
chromatography is 100%, we estimate that gastric microsomes contain
~17 µg of clathrin heavy chain per milligram of microsomal protein.
If one assumes that the H-K-ATPase comprises minimally 50% of the
purified gastric microsomal protein, a ratio of 50 copies of H-K-ATPase per copy of clathrin heavy chain can be calculated.

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Fig. 3.
Hydroxyapatite chromatography of gastric microsomal
peripheral membrane proteins. Gastric microsomal membranes were
stripped of their peripheral membrane proteins in 0.5 M
Tris · HCl buffer and fractionated on hydroxyapatite. Each
fraction was analyzed by SDS-PAGE and immunoblotting. Each lane
contains the proteins from 1/40 of the total volume of each fraction.
A: Coomassie blue-stained gel. Lane 1, nonbinding
fraction; lane 2, proteins eluted with 10 mM sodium
phosphate; lane 3, proteins eluted with 100 mM sodium
phosphate; lane 4, proteins eluted with 200 mM sodium
phosphate; lane 5, proteins eluted with 400 mM sodium
phosphate. Positions of molecular mass markers are shown and are the
same as described in Fig. 1. B: Western blot of the same
fractions in A with anti-clathrin heavy chain MAb 23. C: Western blot of the same fractions in A with
anti-light chain MAb CON.1. D: Western blot of the same
fractions in A with anti- -adaptin MAb 100/3. Signal was
detected by ECL with a 2-min exposure.
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Finally, as reported previously, the majority of
-adaptin (Fig.
3D) and a
-adaptin that is apparently immunologically
distinct from the conventional
1- and
2-adaptins (not shown) also eluted into this
fraction (38). This fraction was used to characterize gastric microsomal clathrin heavy chain by mass spectrometry.
Characterization of gastric microsomal clathrin by MS.
The 160-kDa gel band immunoreactive with anti-clathrin heavy chain MAbs
(Fig. 3, A and B, lane 4) was subjected to in-gel trypsinolysis, and the recovered peptide mixture was analyzed by
MALDI-TOF-MS to yield a peptide mass fingerprint (Fig.
4). When a database search was performed
with 17 input peptide masses, only conventional human, bovine, and rat
clathrin heavy chain (clathrin heavy chain I) matched the input
criteria given in MATERIALS AND METHODS. These
observed peptides are compared with the predicted masses from tryptic
digestion of clathrin heavy chain I in Table 1, all agreeing within a mass accuracy of
0.2 Da.

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Fig. 4.
Matrix-assisted laser desorption delayed extraction reflection
(MALDI) time of flight (TOF) mass spectrometry (MS) analysis of
peptides generated from in-gel tryptic digestion of gastric microsomal
clathrin enriched by hydroxyapatite chromatography. The hydroxyapatite
fraction enriched in immunoreactive clathrin heavy chain was resolved
by SDS-PAGE. The 160-kDa band immunoreactive with anti-clathrin heavy
chain antibodies was excised from the Coomassie blue-stained gel and
subjected to in-gel proteolysis with trypsin. Aliquots (1/25) of
unseparated tryptic digests were cocrystallized with
-cyano-4-hydroxycinnamic acid and analyzed using an MALDI TOF
instrument equipped with a nitrogen laser. Measurements were performed
in a positive ionization mode. The peptide mass fingerprint is shown
with peptides matching clathrin heavy chain I assigned to their
respective masses. m/z, Mass-to-charge ratio.
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Confirmation of the 160-kDa band as clathrin heavy chain I was obtained
from sequence information based on PSD spectra. To obtain good PSD
spectra, the total tryptic peptide mixture was fractionated by
reverse-phase microbore HPLC before MALDI-MS. A representative PSD
spectrum of peptide mass-to-charge ratio (m/z) 1,415.82 (also seen in total digest, Fig. 4) is shown in Fig.
5, which matches a single sequence,
1011IVLDNSVFSEHR1022, from clathrin heavy
chain. Similar PSD sequence confirmation was obtained on peptides with
m/z 1,126.6 (1398Val-Arg1406),
1,304.7 (355Asn-Arg366), 1,433.8 (469Ser-Arg481), 1,943.0 (1482Thr-Arg1498), and 1,971.2 (1227Leu-Arg1245) (data not shown). All
peptides identified from PSD data are indicated by underline in Table
1. The peptide mass and sequence data thus verified gastric microsomal
clathrin to be the conventional clathrin heavy chain (clathrin heavy
chain I) isoform.

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Fig. 5.
Postsource decay (PSD) spectrum of 1,415.82 m/z peptide
purified by reverse-phase microbore HPLC. Sequencing of peptide with
m/z of 1,415.82 confirms that gastric microsomal clathrin
heavy chain is conventional clathrin heavy chain (clathrin heavy chain
I).
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In vitro polymerization of gastric microsomal clathrin and
adaptors.
Proteins stripped from purified gastric microsomes with 0.5 M Tris
buffer were dialyzed against a low-ionic-strength buffer in the absence
or presence of Ca2+. The polymerized proteins were
sedimented and compared with the nonpolymerized proteins. The overall
polypeptide pattern of the sedimented proteins (Fig.
6A, lane 2) differs from that
of the proteins remaining in the supernatant (Fig. 6A, lane
3). The prominent 160-kDa protein in the sedimentable material was
confirmed to be clathrin heavy chain by immunoblot analysis (not
shown). Also, as shown in the Western blot in Fig. 6B,
-adaptin was quantitatively recovered in a sedimentable complex.
However, the quantitative polymerization of clathrin and adaptors did
not appear to depend on the presence of Ca2+ in the
dialysis buffer. Of the total recovered clathrin, 69 ± 21% (SE,
n = 5 independent experiments) was found in the pellet in the absence of Ca2+, whereas 59 ± 30% (SE,
n = 11 independent experiments) was found in the pellet
in the presence of Ca2+.

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Fig. 6.
In vitro polymerization of gastric microsomal clathrin
and adaptors. A: Coomassie blue-stained gel of proteins
assembled into a sedimentable complex on dialysis of proteins stripped
from purified gastric microsomes. Lane 1, starting material
(proteins stripped from purified gastric microsomes by 0.5 M Tris);
lane 2, proteins in a pelletable complex after dialysis in
low-ionic-strength buffer; lane 3, proteins remaining in the
supernatant. For the pellet and the supernatant (lanes 2 and
3), equal proportions of the entire sample were loaded in
each lane. B: Western blot with anti- -adaptin MAb 100/3
of the same samples in A. Signal was detected by ECL with a
2-min exposure. C: Coomassie blue-stained gel of proteins
assembled into a sedimentable complex on dialysis of proteins in the
fraction enriched in clathrin obtained from the hydroxyapatite column
(200 mM sodium phosphate eluate). Lane 1, starting material;
lane 2, proteins in a pelletable complex after dialysis;
lane 3, proteins remaining in the supernatant. For the
pellet and the supernatant (lanes 2 and 3), equal
proportions of the entire sample were loaded in each lane.
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An aliquot of the hydroxyapatite fraction enriched in gastric
microsomal clathrin and AP-1 adaptors (200 mM sodium phosphate fraction; Fig. 3, lane 4) was dialyzed in low-ionic-strength
buffer to induce the polymerization of clathrin and adaptors. As
observed above, the polypeptide pattern of the sedimenting proteins
differed significantly from those proteins remaining in the supernatant (Fig. 6C, lanes 2 and 3), suggesting specificity
in the proteins incorporated into the polymerized complex.
Negative-stain electron microscopy of polymerized clathrin and AP-1
adaptors from gastric microsomes revealed basketlike structures very
similar to those polymerized in vitro from clathrin and adaptors from
brain (Fig. 7). One difference between
the clathrin cages assembled in vitro from brain and gastric microsomal
clathrin is that gastric microsomal clathrin baskets appear to be of
slightly larger diameter: 120 nm compared with 80 nm for reassembled
clathrin-AP-1 adaptor baskets from brain (40). Another
difference is that gastric microsomal clathrin baskets appeared to be
less regular in shape than those from brain. Although other proteins
present in the sedimentable material may play a role in determining the size and shape of clathrin baskets formed in vitro from gastric microsomal clathrin and adaptors, the polymerized baskets appear to be
more similar, rather than dissimilar, to baskets polymerized in vitro
from clathrin isolated from other sources.

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Fig. 7.
Electron micrographs of gastric microsomal clathrin and
AP-1 clathrin adaptors assembled in vitro into baskets. Baskets were
visualized by negative staining with 1% uranyl acetate. Clathrin and
AP-1 clathrin adaptors were assembled by dialysis of the fraction from
hydroxyapatite chromatography that is enriched in clathrin. The
pelletable material was resuspended and processed for negative-stain
electron microscopy. Scale bar, 100 nm.
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Immunofluorescent labeling of clathrin in digitonin-permeabilized
resting parietal cells.
With most of our previously used immunostaining protocols, the
distribution of clathrin immunostaining within parietal cells was
suggestive of labeling of the tubulovesicular compartment (39); however, we could not clearly distinguish between
labeling of clathrin on tubulovesicles or other intracellular membranes and labeling of a cytoplasmic pool of clathrin. Thus a prefixation step
of permeabilization by digitonin was employed to visualize membrane-associated clathrin (33). Figure
8, A and A', shows confocal scanning laser micrographs of digitonin-permeabilized gastric
glands doubly stained for F-actin and clathrin, respectively. Canalicular (apical) membranes of resting parietal cells are delineated by intense staining of F-actin with BODIPY FL-phallacidin (Fig. 8A). The canaliculi within parietal cells are clearly
identifiable as a network of tubular structures projecting from the
gland lumen (Fig. 8, A-C). The lumen of the gland,
including the apical membranes of parietal cells as well as all other
cell types in the gastric gland, is also prominently stained with
BODIPY FL-phallacidin. Abundant intracellular staining for clathrin is
evident in parietal cells, with a significant amount of staining
associated with the regions peripheral to canalicular membranes; this
staining pattern is consistent with clathrin immunoreactivity on
tubulovesicular membranes. Interestingly, clathrin immunoreactivity is
also readily visible at the apical canalicular membranes of parietal
cells (Fig. 8A', arrowheads), as indicated by its
colocalization with F-actin staining of canaliculi (Fig.
8A).

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Fig. 8.
Confocal immunofluorescence microscopy of nonstimulated rabbit
gastric glands doubly stained for F-actin (A-C) and
clathrin (A'), dynamin (B'), and -adaptin
(C'). BODIPY FL-phallacidin was used for F-actin staining.
F-actin staining was prominent along the gland lumen, which is made up
of the apical membrane of nonparietal and parietal cells. Within
parietal cells, F-actin staining is also obvious on the basolateral
membrane and the intracellular canaliculi (arrowheads), which are
invaginated and branching apical membranes of these cells. Gastric
glands were permeabilized with digitonin before fixation and costained
for F-actin (A) and anti-clathrin heavy chain MAb X-22
(A'). Immunostaining for clathrin was observed on
canalicular membranes (arrowheads) and abundantly throughout the
cytoplasm in parietal cells. This latter staining may represent
staining of tubulovesicles. Gastric glands were fixed, permeabilized,
and costained for F-actin (B) and dynamin by use of
anti-dynamin MAb (B'). The most prominent staining for
dynamin is on canaliculi of parietal cells (arrowheads).
Digitonin-permeabilized gastric glands costained for F-actin
(C) and the -adaptin subunit (C') of the AP-2
clathrin adaptor. Canalicular staining for -adaptin is indicated
(arrowheads).
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The presence of clathrin at the canalicular membrane suggests that this
membrane domain is endocytotically active, even in resting parietal
cells. Thus other major proteins that regulate endocytosis at the cell
membrane should also be present. Subsequent staining demonstrated that
canalicular membranes are also immunoreactive for dynamin (Fig.
8B') and the AP-2 clathrin adaptor subunit
-adaptin (MAb
AP.6; Fig. 8C'). This report is the first to identify
clathrin and
-adaptin at the canalicular membrane of parietal cells
and confirms the enrichment of dynamin at the canalicular membrane in
parietal cells (11). Moreover, the expression of this
immunoreactive form of dynamin appears to be significantly higher in
parietal cells than in the nonparietal glandular cells. These results
suggest that the budding of CCVs at the canalicular membrane is
mediated by the AP-2 clathrin adaptor and a member of the dynamin
family of GTPases.
Identification and localization of clathrin-coated pits and
vesicles in parietal cells by thin-section electron microscopy of
isolated rabbit gastric glands prepared by high-pressure rapid freezing
and freeze substitution.
Previous ultrastructural studies have not been able to demonstrate
convincingly the presence of clathrin-coated membranes in parietal
cells (7, 23, 28, 47). Yet, the biochemical and
immunofluorescence data suggest that clathrin is a significant component of tubulovesicles and apical membranes. Thus we sought to reexamine the identification and localization of clathrin in parietal cells at the ultrastructural level. Isolated rabbit gastric glands were prepared for thin-section electron microscopy by
high-pressure rapid freezing and freeze substitution. A
low-magnification (×4,000) image of a resting, nonsecreting parietal
cell is shown in Fig. 9A; this
micrograph demonstrates the excellent morphological preservation of
parietal cells by the high-pressure rapid-freezing technique. Prominent
features observed previously in electron micrographs of resting cells
that are also observed here are the numerous mitochondria,
intracellular canaliculi (which are invaginations of the apical
membrane), and, barely visible, the elaborate system of tubulovesicles
in proximity to intracellular canaliculi. At higher magnification,
coated membranes are clearly visible along the intracellular
canaliculus, which also includes nicely preserved microvilli and their
microfilaments (Fig. 9B). Coated pits are prominent at the
canalicular membranes (Fig. 9C, arrows), and most of the
coated pits appear at the base of canalicular microvilli. On average,
these pits are 60-90 nm diameter. Coated membranous structures in
the subapical cytoplasm (Fig. 9C, arrowhead) are occasionally visible. The ability to identify a significant number of
coated pits at the canalicular membrane reinforces the conclusion that
this membrane in resting parietal cells is an endocytotically active
zone. Moreover, these coats possess a morphology that is highly
reminiscent of conventional clathrin-coated membranes, suggesting that
clathrin is involved in endocytotic processes at the canalicular
membrane.

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Fig. 9.
Transmission electron micrographs of isolated
rabbit gastric glands fixed by high-pressure rapid freezing.
A: low-power magnification of a parietal cell in a gastric
gland. Numerous electron-dense mitochondria (m) and intracellular
canaliculi (c) are visible. A glancing section through a nucleus (n) is
also seen. Tubulovesicles are barely visible at this magnification in
the regions around the intracellular canaliculi. B:
high-power magnification of a region surrounding an intracellular
canaliculus with short microvilli projecting into the lumen of the
canaliculus. Microfilaments are evident in the microvilli lining the
lumen of the intracellular canaliculus. Several endocytic structures,
some of which appear to possess a coat at the cytosolic face of the
membranes, are visible. Mitochondria and numerous tubulovesicular
structures are seen in the surrounding cytoplasm. C:
higher-power view of a region surrounding an intracellular canaliculus.
Several coated endocytic structures are seen invaginating from the
canalicular surface (arrows) or as coated vesicles in the subapical
cytoplasm (arrowhead). Cytoplasmic space also includes numerous
tubulovesicles, several of which appear as indented saccules or
C-shaped structures. Scale bars, 1 µm.
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Despite the excellent preservation of coated pits at the canalicular
membrane, a distinctive clathrin-like coat is not observed on
tubulovesicles, although proteinaceous material is occasionally visible
on the cytoplasmic faces of tubulovesicular membranes. Thus the
ultrastructural identification of clathrin on tubulovesicles could not
be made by this protocol. On the other hand, an interesting morphological feature of these resting parietal cells is the presence of numerous subapical tubular and cup-shaped membranous structures (Fig. 9, B and C), ranging from 45 to 60 nm
diameter and from 300 to 380 nm long. With respect to shape, these
structures are reminiscent of the subapical endocytotic/transcytotic
vesicles observed in Madin-Darby canine kidney (MDCK) cells
(25), but the tubules in parietal cells appear to be
somewhat larger.
Immunogold electron-microscopic localization of clathrin and
H-K-ATPase in resting parietal cells.
Isolated rabbit gastric glands were processed for immunogold
localization of clathrin and H-K-ATPase within resting parietal cells.
The immunogold localization of clathrin by use of anti-clathrin heavy
chain MAb X-22 is shown in Fig.
10A in a region including and surrounding a canaliculus. Gold particles clearly decorate invaginated membranes or pits at the canalicular surface, usually at
the bases of microvilli (arrowheads). These data thus confirm that the
coated pits observed in Fig. 9, B and C, are
comprised of clathrin. In Fig. 10A and in the more extensive
subapical cytoplasmic view in Fig. 10B, densely staining,
80- to 100-nm-diameter vesicular profiles are also decorated with gold
particles, as are the ends of the tubulovesicles (arrows). Thus,
although a distinctive coat was not observed on these membranes by
standard electron-microscopic staining protocols, clathrin can be
localized to these sites by immunogold labeling. Similar results were
obtained with another anti-clathrin heavy chain antibody, MAb 23 (not
shown). This identification of clathrin at the ultrastructural level on
tubulovesicles of parietal cells very likely corresponds to the
clathrin identified biochemically on purified gastric microsomes.

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Fig. 10.
Immunogold labeling of clathrin with anti-clathrin heavy chain MAb
X-22 in parietal cells. Nonstimulated gastric glands were prepared for
immunoelectron microscopy by high-pressure rapid freezing.
A: section showing canaliculus and subcanalicular space.
B: section deeper within the cytoplasm of parietal cell.
Small clusters of gold particles indicated that clathrin at the apical
canalicular membrane was predominantly associated with the membrane at
the base of microvilli (arrowheads), usually concentrated in apical
pits or invaginations, or subapical vesicles. Within the cytoplasm,
accumulations of clathrin-associated gold particles were frequently
found on the ends of tubulovesicles and on small dense structures that
appeared to be near, or sometimes budding from, tubulovesicles
(arrows). Scale bars, 0.5 µm.
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To assess the degree to which the distributions of clathrin and
H-K-ATPase overlap, the anti-H-K-ATPase
-subunit antibody MAb 2G11
was used for immunogold staining of H-K-ATPase (Fig. 11). As expected, anti-H-K-ATPase
staining was observed predominantly in two places in resting parietal
cells: along the central regions of intracellular tubular membrane
profiles and along the microvillar membranes of the canaliculus. The
microvillar staining was more obvious along their lengths than at their
bases. Despite the abundance of anti-H-K-ATPase staining, virtually no
gold particles were associated with mitochondria or intracellular
organelles other than tubulovesicles. However, anti-H-K-ATPase
staining, although it appeared in the same general areas of clathrin
staining (at the canalicular membrane and on tubulovesicles), did not
appear to be concentrated at the same sites in which anti-clathrin
immunolabeling was observed (in invaginations of the canalicular
membrane and at the ends of tubulovesicles), although labeling for the
H-K-ATPase could occasionally be found in pits at the canalicular
membrane and in densely staining vesicles in the cytoplasm. Thus,
overall, clathrin and the H-K-ATPase appear to be segregated to
different regions or subsets of the apical canalicular membrane
and tubulovesicles.

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Fig. 11.
Immunogold labeling of H-K-ATPase in parietal cells.
Immunoelectron microscopy was carried out on the same preparations used
for clathrin staining in Fig. 10, but with MAb 2G11 used for
H-K-ATPase. Gold particles were abundant in the cytoplasm in close
association with tubulovesicles. The apical canalicular membrane was
also decorated with gold particles, usually more obvious along the
lengths of the microvilli and less concentrated at the microvillar
bases. Virtually no gold particles were associated with mitochondria or
other intracellular organelles. Scale bar, 0.5 µm.
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Subfractionation of gastric microsomes by glycerol gradient
centrifugation.
The immunogold labeling of clathrin in parietal cells suggests that at
least two different types of intracellular membranes may possess a
clathrin coat: the ends of tubulovesicles and the densely staining
vesicles. Such membranes may cofractionate with "conventionally"
purified H-K-ATPase-rich gastric microsomes. Thus we subjected purified
gastric microsomes to additional fractionation on a discontinuous
glycerol gradient. Figure 12 shows
Coomassie blue-stained gels and Western blots of purified gastric
microsomes subfractionated on a discontinuous glycerol gradient. By
this approach, a population of clathrin-rich and H-K-ATPase-poor
membranes was identified at the 40% glycerol boundary, whereas the
majority of H-K-ATPase was found in membranes sedimenting at the 80%
glycerol boundary (Fig. 12A). It is clear from the
immunoblots that clathrin is also present in the 80% glycerol fraction
(Fig. 12B), and after correction for the total amount of
protein, this fraction still contained the majority of the total
microsomal clathrin. Interestingly, the 80% fraction is also enriched
in the
-adaptin subunit of the AP-1 clathrin adaptor (Fig.
12C). Thus the glycerol gradient can effect the separation
of two types of clathrin-coated membranes, those poor in H-K-ATPase and
those rich in H-K-ATPase, and may correspond to two (or more)
intracellular populations of clathrin-coated membranes identified by
immunogold electron microscopy.

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Fig. 12.
Fractionation of purified gastric microsomes on
discontinuous glycerol velocity gradients. Gastric microsomes purified
on a 32% sucrose density gradient were further fractionated on
discontinuous glycerol velocity gradients. A: Coomassie
blue-stained gel of glycerol gradient fractions. The position of the
migration of the H-K-ATPase -subunit is shown. B: Western
blot with anti-clathrin heavy chain MAb TD.1. C: Western
blot with anti- -adaptin MAb 100/3. Signals on Western blots were
detected by ECL with films exposed for 2 min. Lane 1, 20 µg of 32% sucrose density gradient-purified gastric microsomes;
lane 2, membranes fractionating at the 20% glycerol layer;
lane 3, membranes fractionating at the 40% glycerol layer;
lane 4, membranes fractionating at the 80% glycerol layer,
representing of total protein fractionating at this layer.
The migration of molecular markers is shown.
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Immunofluorescent staining of clathrin, AP-1 clathrin adaptors, and
H-K-ATPase in resting and stimulated primary cultures of rabbit
parietal cells.
With the characterization of the steady-state localization of clathrin
in resting cells, we sought to characterize the role of clathrin in the
dynamic membrane trafficking processes occurring during functional
resting-to-stimulated transition of parietal cells. Primary cultures of
parietal cells have proven to be a good system with which to evaluate
membrane recruitment and structural rearrangement associated with
stimulation (2).
Within a few hours of being isolated and placed in culture, the
apical canalicular membrane becomes sequestered into the parietal cell
and now appears as a collection of vacuoles that are clearly identifiable by differential interference contrast microscopy or by
labeling the membrane with probes for F-actin. Figure
13 shows cultured parietal cells in the
resting state stained variously for H-K-ATPase, clathrin (with MAb
X-22), and F-actin. F-actin staining clearly demarcates the apical
membrane vacuoles and the basolateral membrane surrounding the cell
(Fig. 13, B and C). In resting cells, H-K-ATPase
can be seen in a punctate distribution throughout the cytoplasm and, to
some extent, within the apical membrane vacuoles (Fig. 13, A
and B'). Staining with MAb X-22 indicates that distribution
of clathrin is similar to that of H-K-ATPase, i.e., throughout the
cytoplasm and within the apical membrane vacuoles (Fig. 13,
A' and C').

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Fig. 13.
Immunofluorescent labeling of clathrin heavy chain and
F-actin or H-K-ATPase in cultured parietal cells under various
secretory conditions. Primary parietal cell cultures were held in a
resting state through the addition of 100 µM cimetidine as an
H2 receptor antagonist (resting) or maximally stimulated
through addition of 100 µM histamine and 30 µM IBMX (stimulated).
In some cases, cells were treated with the proton pump inhibitor
SCH-28080 (5 µM) in addition to histamine plus IBMX (stimulated + SCH-28080). After 25 min of incubation with drugs at 37°C, cells
were fixed and processed for immunofluorescence. Some cells were doubly
stained for F-actin (Phall) and clathrin (X-22) or for F-actin and
H-K-ATPase (HK). In those cases where cells were doubly stained for
clathrin and H-K-ATPase, the MAb 2G11 antibody for H-K-ATPase was
previously biotinylated, and rhodamine-conjugated avidin was used as
the fluorescent probe. In the resting state, F-actin can be seen
predominantly at the apical membrane vacuoles within the parietal cells
and somewhat less intensely at the surrounding plasma (basolateral)
membrane. In these resting cells, clathrin heavy chain and H-K-ATPase
are distributed in a similar, although not identical, pattern, i.e.,
some staining at the apical membrane vacuoles and punctate staining
throughout most of the cytoplasmic space. In stimulated parietal cells,
the apical membrane vacuoles have expanded to enormous size because of
accumulation of HCl and water. There appears to be a great deal of
overlap in the staining for F-actin, clathrin, and H-K-ATPase, possibly
because of the diminution of cytoplasmic space as the vacuoles have
expanded. For cells stimulated in the presence of the proton pump
inhibitor (stimulated + SCH-28080), the apical membrane vacuoles
are not expanded or are only slightly enlarged, and in most cases there
have been some peripheral extensions in the form of lamellipodia. Even
though HCl secretion does not occur, H-K-ATPase has been recruited to
the apical membrane vacuoles and has been virtually cleared from the
cytoplasm (tubulovesicles), essentially colocalizing with F-actin at
the apical membrane vacuoles. On the other hand, much of the clathrin
is still distributed throughout the cytoplasm. Scale bar, 10 µm.
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Parietal cells respond vigorously to secretagogues, recruiting
H-K-ATPase from the compartment of cytoplasmic tubulovesicles to the
apical vacuoles and pumping HCl and water into those structures. As
shown in Fig. 13, D-F, the apical vacuoles greatly
expand as a consequence of 1) the fusion of tubulovesicles
and 2) the osmotic force created by the pump. In this
maximally stimulated configuration where the vacuolar space occupies
most of the viewable cell, staining for F-actin (Fig. 13, E
and F), H-K-ATPase (Fig. 13, D and
F'), and clathrin (Figs. 13, D'and E')
appear to overlap largely at the periphery of the ballooned vacuolar
structures. However, the shrinking of the residual cytoplasmic space
makes it difficult to conclude anything regarding specific localization.
Treatment of parietal cells with proton pump inhibitors has been shown
to produce a definitive pattern of stimulation-dependent structural
changes. When stimulated parietal cells were treated with the pump
inhibitor SCH-28080, H-K-ATPase was cleared from the cytoplasm and
translocated to the apical membrane vacuoles, where it is colocalized
with F-actin. The vacuoles become slightly enlarged, but there is ample
remaining cytoplasmic space that has been cleared of H-K-ATPase
(2, 20) (Fig. 13, H and H'). Thus
membrane recruitment and trafficking of H-K-ATPase remained intact,
even though ion transport was inhibited. On the other hand, staining
with MAb X-22 reveals that although some clathrin is associated with
the apical membrane vacuoles, much parietal cell clathrin remains
distributed throughout the cytoplasm (Fig. 13, G and
G'), sometimes even extending into the lamellipodia, thus
demonstrating that a component of cytoplasmic clathrin can be
segregated from H-K- ATPase on secretory activation.
Probing of parietal cell cultures with other anti-clathrin heavy chain
(MAb 23; Fig. 14,
A'-C') or light chain (MAb CON.1; Fig. 14,
D'-F') antibodies or for the AP-1 clathrin adaptor (MAb 100/3; Fig. 14, G' and H') produced results
identical to those for MAb X-22. In resting cells, clathrin was seen
throughout the cytoplasm, as well as at the apical membrane vacuoles.
When cells were stimulated in the presence of SCH-28080, the pattern of
clathrin and AP-1 clathrin adaptor staining was not significantly
different from that of resting cells, i.e., much of the signal for MAbs 23, CON.1, and 100/3 remained distributed throughout the cytoplasm (Fig. 14, C', F', and H'). These data
together suggest that clathrin and the AP-1 adaptor are not
translocated en masse to the apical membrane on stimulation of the
parietal cell and, therefore, do not appear to accompany the H-K-ATPase
to the apical membrane in stimulated cells.

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|
Fig. 14.
Immunofluorescent double labeling of F-actin and
clathrin heavy chain, clathrin light chain, or -adaptin in cultured
parietal cells under various secretory conditions. Primary parietal
cell cultures were held in a resting state through the addition of 100 µM cimetidine (resting), maximally stimulated by the addition of 100 µM histamine and 30 µM IBMX (stimulated), or treated with the
proton pump inhibitor SCH-28080 (5 µM) in addition to histamine plus
IBMX (Stim + SCH-28080). After 25 min of incubation with the
drugs, cells were fixed, permeabilized, and probed for F-actin
(A, B, and C) and clathrin heavy chain by use of
MAb 23 (A', B', and C'), for F-actin (D,
E, and F) and clathrin light chain by use of CON.1 MAb
(D', E', and F') or for F-actin (G and
H) and -adaptin by use of MAb 100/3 (G' and
H'). Morphological responses to stimulants plus inhibitor
were the same as for Fig. 13; i.e., vacuoles expanded when stimulated
and the enlarged swelling was prevented by the pump inhibitor. A large
portion of clathrin, when probed for heavy chain (C') or
light chain (F'), remained distributed throughout the
cytoplasm in parietal cells stimulated in the presence of SCH-28080.
Similarly, much of the signal for -adaptin (H') remained
distributed throughout the cytoplasm in parietal cells stimulated in
the presence of SCH-28080. In parallel tests, H-K-ATPase was found to
be cleared from the cytoplasm and recruited to the apical membrane
vacuoles when the cells were stimulated in the presence of SCH-28080
(not shown). Scale bars, 10 µm.
|
|
 |
DISCUSSION |
Biochemical characterization of gastric microsomal clathrin.
Previously, clathrin and an AP-1 clathrin adaptor were identified on
gastric microsomes from parietal cells and preliminarily characterized
(39). In this study, gastric microsomal clathrin was
characterized biochemically as a first step in the elucidation of its
function in the regulation of membrane trafficking in the gastric
parietal cell. Clathrin appears to constitute a significant fraction of
the total peripheral membrane proteins of purified gastric microsomes.
The enrichment of clathrin from gastric microsomes on hydroxyapatite
columns reported here should serve as a convenient preliminary step in
the purification of clathrin from gastric microsomes. Gastric mucosal
tissue may represent an easily obtainable source of clathrin from
epithelial cells and should therefore facilitate the biochemical
analysis of clathrin and associated proteins from a secretory
epithelial cell.
Clathrin on gastric microsomes is apparently comprised of a
conventional heavy chain and a light chain, with a predominance of LCa.
The predominance of LCa on gastric microsomes, which are highly
enriched in tubulovesicles, is an intriguing finding, given that
tubulovesicles are a regulated secretory compartment. Previously, a
predominance of LCb was demonstrated in cells that possess a regulated
secretory pathway, such as those from brain and adrenal gland and in
certain cultured cells, such as rat pheochromocytoma (PC-12) cells
(1). On the other hand, LCa was found to predominate over
LCb in cells that do not possess a regulated secretory pathway (cultured cells such as fibroblasts and MDCK cells) and cells in
kidney. However, the present data suggest that perhaps the specific
type of regulated secretory pathway may be important in dictating the
ratio of LCa to LCb, rather than the presence of a regulated secretory
pathway per se. Alternatively, the type of light chain may influence
the size of CCVs or clathrin-coated tubules (9). For
example, CCVs from brain are generally smaller than those from other
tissues, which may be a reflection of not only a predominance of LCb
over LCa, but also a result of both light chains containing neuronal
tissue-specific inserts (9, 30). Consistent with this
hypothesis, we observed that baskets polymerized from tubulovesicular
clathrin and AP-1 adaptors are larger than those polymerized from brain
clathrin and AP-1 adaptors (40). It is also known that
clathrin light chains negatively affect polymerization of clathrin
triskelions (55) and influence the stability of
triskelions (27). The ratio of LCa to LCb may thus impart
physiologically relevant properties to clathrin with respect to
membrane trafficking in the parietal cell secretory cycle.
Despite the relative biochemical abundance of clathrin on purified
gastric microsomes, shown here and in previous work (39), a morphologically distinct clathrin coat has not been reported on
tubulovesicles in any of the previous electron-microscopic analyses
(7, 23, 28, 47). On the other hand, the clathrin baskets
polymerized in vitro may differ markedly from those assembled onto
membranes in vivo. For example, atypical clathrin-coated membranes have
been reported in association with adhesion plaques (37)
and at postsynaptic membranes (5). There is also evidence from yeast for a novel clathrin assembly complex (42).
Moreover, the inability to detect clathrin on some membranes by
standard electron-microscopic techniques may not be unique; only
recently have clathrin-coated buds on endosomes been identified by
immunoelectron microscopy in several different studies (18, 24,
34, 48). Thus the past inability to detect clathrin at the
ultrastructural level on tubulovesicles may be a reflection of some
fundamental properties of clathrin, such as its polymerized structure,
that obfuscates its detection by standard electron-microscopic techniques.
Ultrastructural localization of clathrin on tubulovesicles.
With the high-pressure rapid-freezing technique for preservation of
gastric glands for immunoelectron microscopy, we have finally
successfully identified clathrin on intracellular membranes resembling
tubulovesicles in parietal cells. The amount of anti-clathrin labeling
we observe on tubulovesicles at the ultrastructural level relative to
the amount of H-K-ATPase immunoreactivity appears to be consistent with
our biochemical estimation of the amount of clathrin relative to
H-K-ATPase. An intriguing feature of the distribution of clathrin is
its localization to specific sites on tubulovesicular membranes, i.e.,
at their ends. The implication of these findings is that clathrin may
be involved in the formation of vesicles budding from tubulovesicles in
resting parietal cells, with the canalicular membrane as a potential
target membrane domain (see below). For example, clathrin on
tubulovesicles may be involved in recycling of apical membrane
components from tubulovesicles to the apical membrane, such as
receptors for soluble N-ethylmaleimide-sensitive factor
attachment protein (t-SNARES) like syntaxin 3 (41).
The other clathrin-coated intracellular membranes identified by
immunogold labeling are the densely staining, 80- to 100-nm vesicles.
The origin of these vesicles is unknown. They may represent endocytotic
vesicles, vesicles budded from tubulovesicles, or vesicles budding from
another distinct subcellular membrane compartment, such as early
endosomes. Alternatively, they may represent nascent buds from
tubulovesicles viewed in cross section. They are not enriched in
H-K-ATPase and may be the vesicles that can be subfractionated from
purified gastric microsomes by glycerol gradients. Scaling up the
glycerol gradient might allow for the further characterization of this
subpopulation of clathrin-coated membranes.
Another unique morphological feature in parietal cells revealed by the
high-pressure freezing technique is the appearance of tubulovesicles as
cup-shaped tubules and flattened saccular membranes in the subapical
cytoplasm. The cup-shaped tubules are morphologically similar to those
observed in MDCK cells (25), although parietal cell
tubules appear somewhat larger. In MDCK cells, these tubules were shown
to be derived from an endocytic compartment and are thought to
correspond to the "apical recycling compartment" in these cells
(4, 6, 25, 51). Although these membranes in parietal cells
are not precisely characterized, a large fraction of them appear to
stain positively for H-K-ATPase and are likely to represent bona fide
tubulovesicles. Also, the cup-shaped and saccular structures in
parietal cells may be the equivalent of the apical recycling
compartment or apical early endosomes of other epithelial cells.
Localization of clathrin and associated proteins at the canalicular
membrane.
In addition to clathrin,
-adaptin and a member of the dynamin family
of large GTPases were also immunolocalized to canalicular membranes.
Using the high-pressure rapid-freezing protocol and standard staining
techniques or immunogold labeling, we have also been able to visualize
coated pits and membranes at the canalicular surface that are
morphologically very similar to conventional clathrin-coated pits.
Taken together, these results provide the first evidence that the
canalicular membrane of the resting parietal cell is endocytotically
active in a process involving clathrin, the AP-2 clathrin adaptor, and
a dynamin. It would be of interest to identify the endocytic cargo in
resting parietal cells to characterize further the role of clathrin in
the physiology of resting cells; in this pathway, a possible candidate
for endocytotic cargo might be a v-SNARE, e.g., VAMP (10,
41). Interaction of v-SNARE with endocytotic machinery has been
shown in the case of synaptotagmin-AP-2 adaptor interactions at
neuronal synapses (46, 56).
The detection of dynamin at the canalicular membrane confirms the
findings of Calhoun et al. (11), who, using the same
anti-dynamin MAb used in this study, reported the presence of dynamin
on canalicular membranes in cultured parietal cells. The
immunoreactivity of the canalicular membrane with the anti-dynamin MAb
is intriguing, given the data presented suggesting that the expression
of this immunoreactive form of dynamin in gastric glands appears to be almost exclusive to the parietal cell and enriched at the canalicular membrane. The anti-dynamin MAb was raised against a region in dynamin I
(a COOH-terminal fragment spanning amino acids 698-851) that is a
relatively conserved region among all the dynamins. Because
dynamin I is thought to be neuron specific (49), it is
likely that the anti-dynamin MAb is recognizing another member of the
dynamin family (dynamin II or III). Inasmuch as many splice variants of
dynamin I, II, and III have been identified (12), it will
be of interest to identify at the molecular level the dynamin isoform
expressed in parietal cells, particularly if this isoform is one that
may be preferentially targeted to the apical membrane of all epithelial
cells. Relative to this speculation, dynamin I heterologously expressed
in MDCK cells is targeted to the apical membrane (3),
whereas in pancreatic acinar cells, an endogenous dynamin II has been
immunolocalized to the apical membrane (50). Also, an
endogenous dynamin in Caenorhabditis elegans is targeted to
the apical membrane of its intestinal epithelial cells
(31).
Clathrin and membrane trafficking in the parietal cell secretory
cycle.
Morphological data presented here suggest that the canalicular membrane
of resting parietal cells is endocytotically active, and this process
is mediated by clathrin, the AP-2 clathrin adaptor, and dynamin. The
endocytotic cargo is likely to be destined for tubulovesicles, early
endosomes, or some yet uncharacterized intracellular membrane
compartment. To maintain the steady-state tubulovesicular or
intracellular membrane surface area, membranes need to be recycled from
these intracellular compartments to the apical membrane. This process
also appears to be mediated by clathrin. Three models may account for
the steady-state localization of clathrin on canalicular and
intracellular membranes in resting parietal cells. The first model is
one in which continuous endocytosis and recycling is occurring in
resting cells, as occurs in most cells. Thus one would predict that the
population of intracellular membranes would be comprised of a set of
clathrin-coated early endosomal membranes and a distinct set of
regulated secretory H-K-ATPase-rich tubulovesicular membranes that
might not participate in the constitutive endocytotic-recycling pathway
and might not be clathrin coated. However, the clathrin-coated early
endosomes may copurify with the H-K-ATPase-rich membranes on sucrose
density gradients, thereby giving the impression at the biochemical
level that clathrin resides on H-K-ATPase-rich membranes.
The second model is that constitutive membrane trafficking in resting
cells may represent an extension of the "recovery" phase of the
parietal cell after stimulation, in which the retrieval of membrane and
H-K-ATPase from the apical membrane on the cessation of HCl secretion
may be effected by a two-step process: a relatively rapid, wholesale
uptake of apical membrane followed by an extended phase of recovery
involving more specific sorting of membrane proteins, such as SNARES or
unretrieved H-K-ATPase, to their appropriate steady-state locations.
These processes would be analogous to bulk flow membrane traffic and
signal-mediated sorting, respectively (52). Both of these
steps could be mediated by clathrin, with the second step requiring the
action of clathrin and clathrin adaptors.
A third model for clathrin in resting cells is that it performs two
functions in two distinct populations of tubulovesicles. In one
population of tubulovesicles, poor in H-K-ATPase, it could mediate the
exchange of membrane and membrane protein with the apical membrane;
this exchange may be required for some yet uncharacterized "housekeeping" function, analogous to trafficking through an early endosomal compartment, as described above. In another population of
tubulovesicles, rich in H-K-ATPase, it could sequester fusogenic (i.e.,
v-SNARE-rich) domains to prevent premature fusion of tubulovesicles. In
support of this dual hypothesis, fusogenic domains of Golgi membranes
have been shown to be mechanically separable from other domains
(19), and the AP-3 adaptor has been shown to interact with
a synaptic vesicle v-SNARE to mediate the budding of synaptic vesicles
from endosomes (18, 45). The identity of endocytotic and
recycling cargo will be important to elucidate with respect to
validation of these models, and these models are not necessarily mutually exclusive. Some approaches to evaluate the profiles of intracellular membranes of the parietal cell would be a more thorough fractionation and characterization of the membranes constituting a
conventional preparation of purified gastric microsomes, the development of an in vitro budding assay, and colocalization of clathrin, H-K-ATPase, or other membrane markers at the
immunoelectron-microscopic level.
Because the resting parietal cell appears to be endocytotically active,
on stimulation of the parietal cell, the volume of exocytosis must be
stimulated such that it greatly exceeds that of endocytosis. In
stimulated cells, clathrin is predominantly localized to the cytoplasm;
thus clathrin does not appear to accompany the H-K-ATPase to the apical
membrane on stimulation. Such an outcome might have been predicted if
one considers that if tubulovesicles are the clathrin-coated membrane
compartment, they must be uncoated before their fusion with the
canalicular membrane. Thus, in this scenario, it would appear that
clathrin's main role in the parietal cell secretory cycle may be the
retrieval of membrane and H-K-ATPase when HCl secretion ceases.
Alternatively, clathrin may remain intracellular because of its
association with membranes, such as early endosomes described above,
that do not fuse with the apical membrane on stimulation. One challenge
will be to develop a recycling model in which these hypotheses might be tested.
In summary, the morphological and biochemical data reported here
suggest that the pattern of membrane trafficking and the proteins
regulating this traffic in parietal cells may be more complex than
previously thought. However, these data have given us the ability to
establish a framework for developing testable hypotheses to elucidate
the function of clathrin in parietal cells and, by extension, in apical
membrane trafficking in secretory epithelial cells. The tools are now
available to launch a multidimensional approach to address this
fundamental issue in epithelial cell biology with the parietal cell as
a model system.
 |
ACKNOWLEDGEMENTS |
The authors thank the laboratory of Dr. Vincent Lee for rabbit
stomachs and Drs. Frances Brodsky, Andy Wilde, and Shu-Hui Liu for
generous gifts of antibodies and advice.
 |
FOOTNOTES |
Mass spectra were obtained at the University of California, San
Francisco, Mass Spectrometry Facility, which was supported by the
Biomedical Research Technology Program of the National Center for
Research Resources Grants RR-01614 and RR-08282. This work was
supported by grants from the University of Southern California Gastrointestinal and Liver Diseases Center, the National American Heart
Association, the Burroughs Wellcome Fund, the American Foundation for
Pharmaceutical Education (C. T. Okamoto), and National Institute of Diabetes and Digestive and Kidney Diseases Grants DK-51588 (C. T. Okamoto) and DK-10141 and DK-38972 (J. G. Forte).
Address for reprint requests and other correspondence: C. T. Okamoto, Dept. of Pharmaceutical Sciences, School of Pharmacy, University of Southern California, 1985 Zonal Ave., Los Angeles, CA
90089-9121 (E-mail: cokamoto{at}hsc.usc.edu).
The costs of publication of this
article were defrayed in part by the
payment of page charges. The article
must therefore be hereby marked
"advertisement"
in accordance with 18 U.S.C. §1734 solely to indicate this fact.
Received 22 January 2000; accepted in final form 3 April
2000.
 |
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