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Vol. 14, Issue 7, 2689-2705, July 2003
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Department of Cell Biology, Johns Hopkins University School of Medicine, Baltimore, Maryland 21205
Submitted December 12, 2002;
Revised March 3, 2003;
Accepted March 13, 2003
Monitoring Editor: Jennifer Lippincott-Schwartz
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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;
Gu et al., 2001;
Tuma and Hubbard, 2001). Most
of the signals contain either a tyrosine in the context of a short degenerate
sequence or a dileucine motif. Some signals overlap with those used at the PM
in receptor-mediated endocytosis, whereas other basolateral signals, such as
that in the polymeric IgA receptor (pIgA-R), are unique.
The identification of cytoplasmic signals that mediate apical PM targeting
has been much more elusive. Many apical residents (e.g., single transmembrane
domain [TMD] ectoenzymes) have very short cytoplasmic tails (6–8 amino
acids) or lack them entirely (e.g., glycophoshatidylinositol [GPI]-anchored
proteins). However, two apical sorting signals have been proposed:
extracellular glycans on the apical residents or the incorporation of apical
residents into specialized membrane domains
(Scheiffele et al.,
1995; Harder and Simons,
1997). The latter sorting mechanism is the focus of this
study.
The "raft hypothesis" for apical protein sorting emerged from
two fundamental observations. First, glycosphingolipids and cholesterol are
enriched in the apical surfaces of epithelial cells. The intrinsic properties
of these lipid species are thought to promote their assembly into specialized
membrane domains called "rafts" (for review, see
Harder and Simons, 1997;
Brown and London, 1998).
Second, selected proteins are recruited to these domains based on their
biophysical properties. In particular, GPI-anchored proteins, which are
predominantly expressed at the apical PM, are found in rafts. Thus, according
to the raft hypothesis for protein sorting, rafts form in the biosynthetic
pathway where they recruit apically destined proteins (especially GPI-anchored
proteins), and then they with their recruited cargo are transported in
vesicles directly to the apical domain.
Do lipid rafts sort apical residents in polarized hepatocytes? Unlike most simple polarized epithelial cells, the predominant route to the hepatic apical PM is indirect. Newly synthesized apical proteins are delivered from the trans-Golgi network (TGN) first to the basolateral PM where they are selectively internalized and transcytosed to the apical surface. Are apical proteins sorted into rafts at the TGN for delivery to the basolateral PM? Once delivered to the basolateral PM, are apical proteins recruited into rafts that are necessary for apical targeting along the transcytotic route? If so, are rafts present at the basolateral domain or in other transcytotic intermediates?
We determined that only a subset of apical residents in polarized WIF-B cells was detergent insoluble. Examination of the acquisition of insolubility of newly synthesized proteins indicated that apical residents become insoluble with different kinetics and at different places along the biosynthetic pathway. In cholesterol or glycosphingolipid-depleted cells, the basolateral-to-apical transcytosis of all apical residents examined and pIgA-R was inhibited. Apical proteins were basolaterally internalized, but blocked at early endosomes, indicating that rafts are required for endosomal transcytotic efflux. Biochemical and morphological examination of apical proteins in Fao cells further revealed that raftdependent sorting is conserved in nonpolarized cells.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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-cyclodextrin (mCD), cholesterol-loaded mCD,
fumonisin B1 (FB1), horseradish peroxidase (HRP) (type VI), and cytochalasin D
were purchased from Sigma-Aldrich (St. Louis, MO). Latrunculin B was purchased
from BIOMOL Research Laboratories (Plymouth Meeting, PA). Lovastatin was from
A.G. Scientific (San Diego, CA). HRP-conjugated secondary antibodies and Super
Signal West Pico chemiluminescence substrate were from Amersham Biosciences
(Piscataway, NJ) and Pierce Chemical (Rockford, IL), respectively.
Alexa-conjugated secondary antibodies were from Molecular Probes (Eugene, OR).
Anti-transferrin receptor (Tf-R) antibodies were purchased from Accurate
Chemical & Scientific (Westbury, NY) and anti-V5 epitope tag antibodies
were from Invitrogen (Carlsbad, CA). Anti-5'nucleotidase (5'NT)
(monoclonal and affinity purified polyclonal), -CD59 (affinity-purified
monoclonals), and -Tf-R (polyclonal) were kindly provided by J.P. Luzio
(Cambridge University, Cambridge, United Kingdom), P. Morgan (University of
Wales College of Medicine, Cardiff, United Kingdom), and M. Farquhar
(University of California, San Diego, San Diego, CA), respectively. Antibodies
against aminopeptidase N (APN), asialoglycoprotein receptor (ASGP-R), CE9,
pIgA-R, dipeptidylpeptidase IV (DPP IV), and HA321 were prepared in the
Hubbard laboratory and have been described previously
(Bartles et al., 1985;
Hubbard et al., 1985;
Barr and Hubbard, 1993;
Ihrke et al., 1993;
Shanks et al.,
1994).
Cell Culture
WIF-B and Fao cells were grown in a humidified 7% CO2 incubator
at 37°C as described previously (Ihrke
et al., 1993; Shanks
et al., 1994). Briefly, cells were grown in F-12 medium
(Coon's modification), pH 7.0, supplemented with 5% fetal bovine serum. WIF-B
medium was also supplemented with 10 µM hypoxanthine, 40 nM aminoterpin,
and 1.6 µM thymidine. In general, cells were seeded onto glass coverslips
at 1.3 x 104 cells/cm2. Fao cells were cultured
for 3–5 d and WIF-B cells for 8–12 d until they reached maximum
density and polarity.
Exogenous Expression of DPP IV and pIgA-R
WIF-B or Fao cells were infected with recombinant adenovirus particles
(0.7–1.4 x 1010 virus particles/ml) encoding V5/His6
epitope-tagged full-length DPP IV or pIgA-R for 30 min at 37°C as
described previously (see Bastaki et
al., 2002 for detailed methods and description of
constructs). The cells were washed with complete medium and incubated an
additional 18 h to allow expression.
Immunoblot Analysis
Cells were rinsed in cold phosphate-buffered saline (PBS) and extracted for
30 min on ice in 0.15 ml of lysis buffer [1% (vol/vol) Triton X-100, 150 mM
NaCl, 25 mM HEPES, pH 7.4] containing 1 µg/ml each of aprotinin, pepstatin,
antipain, leupeptin, benzamidine, and phenylmethylsulfonyl fluoride. The
samples were sheared using a 26-gauge needle and centrifuged at 120,000
x g for 30 min at 4°C. The resultant pellet was resuspended
to volume with SDS-PAGE sample buffer and boiled for 3 min. Reducing sample
buffer was used for analysis of all proteins except HA321 and CD59 both of
which required nonreducing conditions (lacking -mercaptoethanol) for
optimal immunodetection. For immunoblotting, anti-5'NT (affinity
purified polyclonal) was diluted to 1:200, anti-CE9, pIgAR and -Tf-R (rabbit
polyclonal sera) were diluted 1:5000, whereas ASGP-R and APN polyclonal sera
were diluted to 1:1000 and 1:2000, respectively. Anti-CD59 (affinity purified
monoclonal) was diluted to 1 µg/ml. HRP-conjugated secondary antibodies
were used at 5 ng/ml, and immunoreactivity was detected with enhanced
chemiluminescence. The relative levels of immunoreactive species in the
soluble and insoluble fractions were determined by densitometric comparison of
immunoreactive bands.
In Figure 3, cells were
treated with 5 mM mCD for the indicated times in medium prepared with 5%
LPDM and extracted as described above. In
Figure 5, cells were treated
for 24 or 48 h in complete medium with 25 µM FB1 diluted in methanol. For
the 48-h samples, cells were renewed with fresh medium and drug after the
first 24 h. The control cells were treated with the same methanol
concentration for 48 h, renewing after 24 h.
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Metabolic Labeling
WIF-B cells were incubated in cysteine- and methionine-free medium for 1 h
at 37°C. Cells were then labeled for 10 min at 37°C in the same medium
containing 100–200 µCi of trans-[35S]methionine.
Cells were rinsed with Hanks' buffered saline solution, placed in prewarmed,
complete medium containing excess unlabeled cysteine (64 µg/ml) and
methionine (20 µg/ml) and chased at 37°C for the indicated times. For
the 20°C block, cells were either first chased for 10 min at 37°C
after labeling or transferred directly to precooled chase medium at 20°C
for 2 h. To release the 20°C block, prewarmed medium was added and the
cells returned to 37°C. At the indicated times, the cells were rinsed with
Hanks' buffered saline solution and once with cold PBS before extracting and
immunoprecipitation.
Immunoprecipitations
APN and HA321. The detergent-soluble and -insoluble samples
were prepared as described above for immunoblotting with a few modifications.
The coverslips were instead solubilized in 0.9 ml of lysis buffer,
centrifuged, and the resultant pellet solubilized in 0.2 ml of solubilization
buffer (1% SDS, 50 mM Tris, 5 mM EDTA, pH 8.8), sheared with a 26-gauge needle
until fully resuspended, and diluted to 1.0 ml with lysis buffer. The
supernatants were corrected to contain the same concentration of
solubilization buffer components and diluted to 1.0 ml with lysis buffer. The
detergent-soluble and -insoluble samples were serially immunoprecipitated,
first with anti-APN polyclonal antibodies (1:1000) at 4°C for 16 h.
Protein A-Sepharose was added for 4 h and samples processed as described
previously (Bartles et al.,
1987). The samples were then incubated with affinity-purified
polyclonal anti-HA321 (1:1000) and processed as for anti-APN
immunoprecipitations. Immunoprecipitates were separated by SDS-PAGE and
transferred to nitrocellulose. The membranes were exposed from 18 h to 4 d to
phosphorimaging plates that were scanned using a PhosphorImager (Fuji, Tokyo,
Japan).
5'NT. Cells were rinsed in cold PBS
and lysed in parallel at 4°C for 30 min or at 37°C for 15 min in 0.9
ml of lysis buffer. The 4°C lysates were centrifuged at 120,000 x
g for 30 min at 4°C and the 37°C lysates at 25°C. The
supernatants were supplemented to contain 20 mM octylglucoside, 10 mM Tris, 1
mM EDTA and diluted to 1.0 ml with lysis buffer. Directly conjugated
monoclonal antibody-Sepharose was used to immunoprecipitate 5'NT as
described previously (Schell et
al., 1992).
Thin Layer Chromatography (TLC)
Cholesterol was measured in control or treated cells grown on coverslips.
At the indicated times, total lipids were extracted as described previously
(Bligh and Dyer, 1959) and
separated on 10 x 10 cm high-performance TLC plates with a mobile phase
of hexane/ethyl acetate (80:20). To measure sphingomyelin (SM), control or
FB1-treated cells were extracted as described previously
(Bligh and Dyer, 1959) and the
lipids separated with a mobile phase of chloroform/methanol/concentrated
ammonia (20:5:0.5). Lipids were visualized by charring plates that had been
sprayed with 3% cupric acetate (wt/vol) in 8% phosphoric acid
(Macala et al.,
1983).
Immunofluorescence Microscopy
Control or treated cells were fixed on ice with chilled PBS containing 4%
PFA for 1 min and permeabilized with ice-cold methanol for 10 min. Cells were
processed for indirect immunofluorescence as described previously
(Ihrke et al., 1993).
Alexa 488- or 568-conjugated secondary antibodies were used at 3–5
µg/ml.
Internalization Assays
Cholesterol-depleted Cells. WIF-B or Fao cells were
pretreated for 5 min in LPDM in the absence or presence of 5 mM mCD.
Proteins present at the basolateral PM in WIF-B cells or the PM in Fao cells
were continuously labeled with specific antibodies diluted in LPDM in the
continued absence or presence of mCD. Rabbit polyclonals against ASGP-R,
APN, and DPP IV were diluted 1:100, 1:200, and 1:500, respectively. Mouse
anti-5'NT, anti-V5, and anti-CD59 were diluted 1:1000, 1:2000, and
1:500, respectively, and hybridoma supernatant containing anti-TF-R antibodies
was diluted 1:5. After labeling, cells were fixed as described above, and the
trafficked antibodies were labeled with Alexa-488 or -568–conjugated
secondary antibodies (3–5 µg/ml) as described previously
(Ihrke et al.,
1998).
For recovery assays, cells were pretreated in LPDM containing 5 mM
mCD for 60 min. Cells were rinsed twice in prewarmed LPDM and incubated
an additional 60 min in LPDM in the absence or presence of 365 µg/ml
cholesterol-loaded mCD (equivalent to the addition of 20 µg/ml free
cholesterol). Antigens at the basolateral PM (in WIF-B cells) or PM (in Fao
cells) were continuously labeled for 60 min in the continued presence of the
drug, fixed, permeabilized, and stained as described above.
Sphingolipid-depleted Cells. WIF-B cells were pretreated for 48 h in complete medium in the absence or presence of 25 µM FB1 (medium and drug renewed daily). For experiments where DPP IV or pIgA-R expression was required, cells were infected with recombinant adenovirus after the first day of FB1 treatment. Cells were antibody labeled, fixed, and stained as described above.
Kinetic Assays
Total IgG from serum (APN or DPP IV) or ascites (5'NT) were purified
(EZ-Sep; Pharmacia AB, Uppsala, Sweden) and biotinylated (EZ-Link
sulfo-NHS-biotin; Pierce Chemical) according to the manufacturers'
instructions. To measure internalization, WIF-B cells were continuously
labeled with biotinylated antibodies for the indicated times at 37°C. The
remaining surface-associated antibodies were eluted with isoglycine (200 mM
glycine, 150 mM NaCl, pH 2.5) for 5 min at room temperature and the cells
lysed in isoglycine containing 20 mM octylglucoside and 0.5% Triton X-100 for
30 min on ice. Aliquots of the eluate and lysate were incubated in
streptavidin-coated 96-well plates (Pierce Chemical). Bound antibodies were
detected with HRP-conjugated secondary antibodies followed by colorimetric
detection with an HRP substrate detection kit (BioRad, Hercules, CA).
Recycling Assays
Fao PM proteins were continuously labeled as described for the
internalization assays. To strip surface-associated antibodies, cells were
incubated in isoglycine as described above. The cells were rinsed with PBS,
placed in prewarmed LPDM in the absence or presence of 5 mM mCD, and
incubated at 37°C for 60 min. Cells were fixed and labeled as described
for the internalization assays.
Imaging
Labeled cells were visualized by epifluorescence (Axioplan Universal
microscope; Carl Zeiss, Jena, Germany). Images were acquired with a Princeton
MicroMax cooled charge-coupled device camera (Roper Scientific, Trenton, NJ)
and IP Labs software (Scanalytics, Fairfax, VA). Further image processing and
figure compilation was performed using Photoshop (Adobe Systems, Mountain
View, CA) and Microsoft PowerPoint software (Microsoft, Redmond, WA).
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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Incorporation into Rafts at the TGN Is Not Required for Apical PM
Sorting of 5'NT
Because 5'NT, CD59, and APN were partially detergent insoluble, we
determined where in their life cycles insolubility was acquired. The approach
we chose was metabolic pulse labeling followed by extraction into soluble and
insoluble pools and immunoprecipitation with specific antibodies. However,
5'NT recovered in the insoluble fraction could not be resolubilized for
subsequent immunoprecipitation. Although APN was quantitatively recovered in
immunoprecipitates from the pelleted fraction resuspended in solubilization
buffer and boiled for 3 min (see MATERIALS AND METHODS and
Figure 2), none of the >85%
of insoluble 5'NT was recovered
(Figure 1B). Changing the
solubilization temperature or time also did not allow for 5'NT
immunoprecipitation from this fraction
(Figure 1B).
To overcome this experimental barrier, we adopted an alternative method
(Cerneus et al.,
1993) for determining the insolubility of newly synthesized
5'NT. Unlike at 4°C, all of 5'NT was soluble in Triton X-100
at 37°C (Figure 1C). From
the densitometric analysis of the Western blot shown, the steady-state amount
of 5'NT recovered in the supernatant and pelleted fractions at 4°C
was the same as that recovered in the supernatant at 37°C (160 versus 163
arbitrary density units). Furthermore, 5'NT in the 37°C extracts was
quantitatively immunoprecipitated (Figure
1C). Thus, the proportion of insoluble 5'NT was determined
by subtracting the amount of 5'NT immunoprecipitated from the 4°C
soluble pool (the soluble fraction) from the amount recovered in the 37°C
extracts (the total population) (Figure
1C). Using this method, we determined that 87.0 ± 6.6% of
5'NT is Triton X-100 insoluble, nearly identical to the value determined
from steady state Western blotting (Figure
1A). Thus, this method was suitable for the subsequent metabolic
labeling experiments.
Cells were 35S-labeled for 10 min and then chased for
0–120 min at 37°C. At each time point, cells were extracted with 1%
Triton X-100, the detergent-insoluble and -soluble pools prepared and the
specific molecules immunoprecipitated. Mature APN reached steady-state
insolubility levels (
45%) quickly, after only 30 min of chase
(Figure 2a). Insoluble
populations of mature 5'NT were also detected after 30-min chase, but
accounted for only 20% of the total mature protein pool
(Figure 2b). After 60 min of
chase, when 5'NT molecules were fully mature, only 50% of the
5'NT population was insoluble. Steady-state insolubility levels were not
observed until 3–5 h of chase (our unpublished data), suggesting that
apical delivery was required for complete insolubility to be achieved, a
process that takes >4 h in intact hepatocytes
(Schell et al., 1992)
or >3 h in WIF-B cells (Ihrke et
al., 1998). We also noticed that after the 10-min label
(0-min chase), 25% of 5'NT was already mature, yet none of this
population was insoluble. This suggested that insolubility might not be
achieved until basolateral delivery (see below). For comparison, we examined
the solubility properties of the newly synthesized basolateral resident, HA321
(Figure 2c). Like APN and
5'NT, mature forms of HA321 were detected after short chase periods, and
by 60 min, the molecules were fully mature. HA321 was not detected in the
insoluble fractions consistent with its steady-state solubility
properties.
To determine whether the insolubility of APN and 5'NT was acquired in
the TGN, the presumed site of raft formation
(Simons and Ikonen, 1997),
metabolically pulse-labeled cells were subjected to a 2-h 20°C temperature
block before an additional chase at 37°C. The 20°C incubation produces
a block in post-Golgi transport such that newly synthesized proteins
accumulate in the Golgi (Matlin and
Simons, 1983; Griffiths et
al., 1985; Saraste et
al., 1986; our unpublished data). Although only 30% of APN
was detected in its mature form after the 2 h block (0 min after release),
25% of that population was detected in an insoluble pool
(Figure 2d). As more newly
synthesized APN matured during the subsequent chase at 37°C, the
proportion of insoluble APN remained constant, suggesting that APN acquired
its insolubility in the TGN and maintained it thereafter. In contrast, 75% of
35S-labeled 5'NT was mature after the temperature block
(Figure 2e), but none was
detected in an insoluble pool. To make sure that the 20°C block was not
accumulating 5'NT in compartments preceding the TGN, cells were chased
for 10 min at 37°C before imposing the 20°C block to allow further
transit along the biosynthetic pathway. As before, nearly all of newly
synthesized 5'NT was mature under these conditions, but no insoluble
populations were detected (our unpublished data). HA321 examined under the
same conditions was never detected in an insoluble pool
(Figure 2f). These results
indicated that different hepatic apical residents were incorporated into
detergent-insoluble domains with differing kinetics and at different places
along the biosynthetic pathway. A percentage of the single TMD protein, APN,
was likely incorporated into rafts at the TGN, whereas the GPI-anchored
protein, 5'NT, became raft-associated much later. These observations
together with the finding that many apical PM proteins are completely
detergent-soluble indicate that raft association is not a universal
requirement for sorting apical residents from the TGN to the basolateral PM in
polarized hepatic cells.
Transcytosis Is Cholesterol Dependent
To determine whether raft association was important in regulating
transcytosis in hepatic cells, we depleted polarized WIF-B cells of
cholesterol, a condition reported to dissociate raft components
(Ohtani et al., 1989;
Kilsdonk et al.,
1995; Scheiffele et
al., 1997). As measured by TLC, cholesterol was rapidly
depleted by the addition of 5 mM mCD
(Figure 3A). After only 15 min
of treatment, cholesterol levels fell to 52.5 ± 6.4% of control, and by
60 min, were 23.0 ± 10.7% of control. To ensure that mCD was not
toxic when cholesterol was depleted to such low levels, we measured cell
viability by trypan blue exclusion. Cells incubated for 60 min in LPDM alone
(control) or in the presence of 5 mM mCD had similar viability levels
(91.4 ± 1.9% of control and 89.7 ± 2.6% of treated). Moreover,
cells remained polarized (our unpublished data). We also examined the effects
of adding lovastatin, another commonly used cholesterol-depleting drug.
However, even after 3 d of treatment in LPDM, cholesterol levels fell by only
20% (our unpublished data).
We next determined the solubility properties of apical proteins in
cholesterol depleted WIF-B cells. In the presence of mCD, only the
solubility of APN was significantly altered. After 15 min of treatment, nearly
all of this single TMD protein was detected in the soluble fraction
(Figure 3, B and C). In
contrast, only small percentages of 5'NT and CD59 (12 and 14%,
respectively) were soluble after 60 min, a time point at which 80% of the
cholesterol was extracted (Figure 3, B and
C).
We then determined whether cholesterol depletion impaired transcytosis of
basolaterally located apical proteins. Cells were pretreated for 5 min with
LPDM in the absence or presence of 5 mM mCD. Transcytosis was monitored
by continuously labeling the basolateral pool of selected apical proteins or
recycling receptors in the absence or presence of mCD for 60 min. The
cells were fixed, permeabilized, and the trafficked antibody–antigen
complexes visualized with secondary antibodies. Although cholesterol depletion
did not change the steady-state apical protein distributions (our unpublished
data), it severely impaired transcytosis of all apical molecules tested. As
reported previously, control cells showed intense fluorescence labeling of all
apical markers tested at the apical PM after 60 min (see
Figure 4a for an example). This
result indicated successful transcytotic delivery. In contrast, the apical
domains in treated cells had no apparent fluorescence labeling for any apical
resident tested, whereas a reciprocal increase in basolateral staining was
often observed. CD59, APN, DPP IV, and pIgA-R trafficking are shown in
Figure 4, b, c, e, and g, respectively. We also examined the trafficking of Tf-R and ASGP-R, recycling
receptors that, like pIgA-R, are internalized by clathrin-mediated mechanisms.
Surprisingly, both receptors were internalized in the same cells where a block
in transcytosis of the apical residents was observed. Cotrafficking of APN and
Tf-R is shown in Figure 4, c and
d, whereas DPP IV or pIgA-R cotrafficking with ASGP-R is shown in
Figure 4, e and f, and g and h, respectively. The robust internalization of the fluid phase marker HRP was
also observed in control and mCD-treated cells (our unpublished data).
Although virtually no apical labeling was observed for DPP IV, intracellular
labeling was observed that colocalized with ASGP-R
(Figure 4, e and f, insets; see
below).
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The exogenous addition of cholesterol reversed the mCD-induced defect
in transcytosis. Cells were first treated for 60 min with mCD, which
impaired transcytosis (Figure
4i), and then the drug was removed and the cells incubated for 60
min in LPDM containing 365 µg/ml cholesterol-loaded mCD. Cells were
subsequently labeled with anti-APN antibodies for an additional hour in the
continued presence of the agent. As shown in
Figure 4j, APN staining was
detected at the apical PM with a concomitant decrease in basolateral PM
staining, indicating reversal of the mCD-induced impairment. No such
recovery was observed in cells that did not receive exogenous cholesterol (our
unpublished data). Thus, the defect in transcytosis was likely due to
cholesterol depletion directly.
We quantitated the morphological observations shown in
Figure 4 by measuring the
relative fluorescence intensity present at the apical and basolateral surfaces
of 100–300 individual cells (Table
1). In control cells, the ratio of apical-to-basolateral
fluorescence for all markers tested was >1, indicating that higher levels
of fluorescence labeling was detected at the apical PM. Notably, the ratios
for 5'NT and pIgA-R were higher than the others, likely reflecting their
faster transcytosis kinetics (Ihrke et
al., 1998). In contrast, the ratios of apical-to-basolateral
PM staining in treated cells were all <1 and represented decreases from 70
to 90% relative to control cells (values in parentheses). This reciprocal
staining is evident in Figure
4. These results indicate that cholesterol is required for
transcytosis of single TMD and GPI-anchored apical residents, even though it
seems not to be relevant to their detergent solubility properties.
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Transcytosis Is Glycosphingolipid Dependent
We next determined whether depletion of glycosphingolipids, other raft
components, also impaired transcytosis. We treated cells with 25 µM FB1 and
monitored glycosphingolipid depletion by measuring SM levels by TLC. Although
SM levels do not specifically reflect cellular gylcosphingolipid levels, FB1
inhibits the formation of a common biosynthetic precursor. Also, SM is far
more abundant than specific glycosphingolipids in WIF-B cells, allowing for
more accurate TLC measurement. After 24 h in FB1, 40% of the SM pool was
depleted, and by 48 h, >60% was depleted
(Figure 5A). These TLC
conditions also resolved cholesterol and as indicated by the arrow in
Figure 5A, its levels were
unchanged by FB1 treatment. The detergent solubility of 5'NT, CD59, and
APN were not altered by FB1 treatment alone
(Figure 5B), although addition
of both FB1 and mCD solubilized APN and CD59, but only
50% of 5'NT (Figure 5C).
We also examined the effect of two actin-depolymerizing agents, cytochalasin D
and latrunculin B, alone and in combination with lipid-depleting agents on
apical protein insolubility. None of the three apical residents were released
into a high-speed detergent supernatant after the agents were used alone. When
both mCD and cytochalasin D were added, APN and CD59 were mostly
solubilized, but 5'NT remained in the pelleted fraction. Together, these
results indicate that other factors impart detergent insolubility to
5'NT.
Although no changes in the solubility properties of the apical residents
were observed after 48 h of FB1 treatment, transcytosis of all apical proteins
tested was significantly impaired. The bright apical labeling of 5'NT in
control cells (Figure 6a) was
clearly absent from cells treated with FB1
(Figure 6b). Apical delivery of
APN, DPP IV, and pIgA-R was also significantly decreased
(Figure 6, c, e, and g). Our
quantitative analysis (Table 1)
confirmed these observations and further revealed that the FB1-induced block
on transcytosis was less severe than that caused by cholesterol depletion. In
all cases, the ratio of apical-to-basolateral fluorescence was greater in
FB1-treated cells than in mCD-treated cells. The overall decreased
severity likely reflects the incomplete depletion of glycosphingolipids by
FB1. In contrast, internalization of ASGP-R and Tf-R seemed to be more similar
to that in control cells. Cotrafficking of APN and Tf-R is shown in
Figure 6, c and d, and DPP IV
or pIgA-R with ASGP-R in Figure 6, e and f,
and g and h, respectively. Although little apical labeling by DPP
IV or pIgA-R was observed, intracellular labeling was detected that
colocalized with ASGP-R (Figure 6,
e–h, insets; see below).
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Apical Proteins Are Basolaterally Internalized in Cholesterol- and
Glycosphingolipid-depleted Cells
To further characterize the block imposed by raft disruption in WIF-B
cells, we determined the step in the transcytotic pathway that was impaired by
mCD or FB1 treatment. First, we examined whether the block was occurring
at the basolateral PM by measuring internalization of basolaterally labeled
apical proteins in control and treated cells. As shown in
Figure 7A, the internalization
of 5'NT was only minimally impaired (<20%) by mCD, and only
after 60 min of treatment. This minor impairment does not account for the
>80% decrease in apical delivery observed
(Table 1). Likewise, the
impairment of APN internalization by mCD (35%), albeit greater than
that seen for 5'NT, could not account for the >90% decrease in APN
apical delivery (Table 1).
Furthermore, DPP IV basolateral internalization was virtually unaffected by
mCD; after 60 min of treatment, internalization levels were 97% of
control (our unpublished data). Thus, cholesterol depletion does not
significantly block internalization from the basolateral PM in WIF-B
cells.
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Consistent with the biochemical results, we detected intracellular labeling
of transcytosing apical proteins in both control and lipid-depleted cells.
However, the extent that the intracellular apical proteins colocalized with
recycling receptors in the two conditions was very different. In control
cells, the transcytosing proteins colocalized in juxta-apical structures. As
shown in Figure 7B, a and b,
nearly perfect colocalization was observed for the single TMD protein, APN,
and the GPI-anchored protein, 5'NT. These structures represent the
supapical compartment (SAC), the previously identified transcytotic
intermediate, because they largely excluded cotrafficked ASGP-R, which
recycles between early endosomes and the PM
(Figure 7B, e and f)
(Wall and Hubbard, 1985;
Ihrke et al., 1998;
Tuma and Hubbard, 2001).
Although apical proteins traverse the early endosome in control cells, the
absence of significant colocalization of ASGP-R and the apical proteins
suggests that the latter quickly traverse this compartment. Similarly, the
transcytosing apical residents and Tf-R that traverses the early endosome en
route to the recycling endosome were not observed in the same intracellular
puncta (Figure 7B, i and j). In
contrast, ASGP-R and Tf-R were partially colocalized in control cells
(Figure 7B, m and n). These
puncta likely represent the early endosome, a common intermediate in the
cellular itineraries of both receptors.
Similar to control cells, substantial overlap of apical proteins in
intracellular structures was observed in mCD-treated cells
(Figure 7B, c and d), but these
puncta were larger and contained ASGP-R
(Figure 7B, g and h, and see
Figure 4, e and f). The
ASGP-R–positive puncta also partially overlapped with Tf-R
(Figure 7B, k and l), and
unlike for control cells, the Tf-R-positive puncta also contained
transcytosing pIgA-R (Figure 7B, o and
p). Similar staining patterns were observed in FB1-treated cells
(our unpublished data; Figure 6, e and
f). Thus, proteins that normally have different final destinations
were found in the same structures after lipid depletion. The simplest
interpretation is that mCD and FB1 block transcytotic efflux from
basolateral early endosomes, common trafficking intermediates of apical
residents and recycling receptors.
Cholesterol Depletion Impairs Apical Protein Trafficking in
Nonpolarized Hepatic Fao Cells
We have recently shown that apical proteins are selectively internalized
from the nonpolarized PM of Fao cells, delivered to a novel compartment
containing only other apical proteins and rapidly recycled back to the PM
(Tuma et al., 2002).
Are the solubility properties of apical proteins and the cholesterol
dependence of apical protein trafficking shared by nonpolarized cells or do
they depend on the polarized cell context? To answer this question, we
examined apical proteins in nonpolarized hepatic Fao cells biochemically and
morphologically.
The solubility properties of the different classes of apical proteins in
Fao cells were strikingly similar to those observed in polarized WIF-B cells.
Both of the GPI-anchored proteins were virtually insoluble in Triton X-100,
whereas APN was 50% insoluble (Figure
8A). As in polarized cells, CE9, Tf-R and ASGP-R were entirely
soluble (our unpublished data). Interestingly, 25% of DPP IV was
insoluble in Fao cells, which may reflect its much higher endogenous
concentration than in WIF-B cells (Shanks
et al., 1994; our unpublished data). In fact, when
overexpressed in WIF-B cells, 25% of DPP IV was insoluble (our
unpublished data; see DISCUSSION). These results suggest the presence of an
apical domain is not required for the steady-state insolubility properties of
apical proteins. Also striking is the finding that the solubility properties
of these proteins were altered similarly by mCD
(Figure 8A). CD59 and
5'NT were solubilized 8–11% in Fao cells compared with
12–14% in WIF-B cells, whereas APN was completely solubilized in both
cell types.
|
Unlike for WIF-B cells, mCD treatment changed the steady-state
distributions of the apical proteins in Fao cells. In control cells, all
apical proteins we have examined reside at the PM and in a novel,
intracellular compartment (Tuma et
al., 2002). An example of this staining pattern is shown in
Figure 8B, a. After mCD
treatment, no intracellular labeling of 5'NT was present and a
reciprocal increase in PM staining was observed
(Figure 8B, b). To determine
whether the loss of intracellular staining reflected impaired apical protein
internalization or recycling, we monitored the trafficking of surface-labeled
apical proteins in control and treated cells. In control cells, transport to
the intracellular compartment was readily detectable after 60 min of labeling
(Figure 8C, a). However, in the
presence of mCD, no intracellular labeling of any apical protein tested
was observed. 5'NT, APN, and DPP IV trafficking are shown in
Figures 8C, b, c, and e,
respectively. As in polarized WIF-B cells, we found that both ASGP-R and Tf-R
were internalized in the same cells in which apical protein trafficking was
blocked. Cotrafficking of APN and Tf-R and of DPP IV and ASGP-R is shown in
Figures 8C, c and d, and e and
f, respectively. Quantitation of these results is shown in
Table 1. In this case, the
intracellular fluorescence intensity was measured relative to the intensity
present at the PM of the same cell. The apical compartment to PM ratios were
all >1 in control cells, whereas in treated cells, the ratios were
drastically reduced (70–90%). Thus, cholesterol depletion impaired
trafficking to the apical compartment in nonpolarized cells. Addition of
exogenous cholesterol reversed the trafficking defect; 5'NT trafficking
to the intracellular compartment returned in the presence of 365 µg/ml
cholesterol-loaded mCD (Figure 9, a
and b).
|
To determine whether recycling from the apical compartment was also
impaired, we examined the trafficking of staged apical proteins.
Antibody-labeled APN was continuously internalized for 60 min to load the
intracellular compartment (Figure
9c). Residual antibodies were surface stripped using isoglycine
(Figure 9d). Recycling to the
PM was monitored in the absence (Figure
9e) or presence of mCD
(Figure 9f). In both cases, PM
fluorescence was regained indicating that recycling does not require
cholesterol.
|
DISCUSSION |
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|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
|---|
; Bartles and Hubbard,
1988; Schell et al.,
1992; Ihrke et al.,
1998). Only two intermediates in the hepatic basolateral-to-apical
transcytotic pathway have been identified: the basolateral early endosome and
SAC (Barr et al.,
1995; Ihrke et al.,
1998). The early endosome also serves as a trafficking
intermediate in at least four separate pathways. One is recycling, where
internalized proteins are returned directly to the basolateral PM (e.g.,
ASGP-R; Wall and Hubbard,
1985). Another is endocytic targeting to lysosomes represented by
HRP. A third pathway is transport to recycling endosomes depicted by Tf-R, and
fourth is the transcytotic pathway that is taken by apical residents. At
present, it is not known whether internalized apical proteins normally recycle
between the early endosome and basolateral PM. Also, except for pIgA-R, the
mechanisms that mediate basolateral internalization of apical residents are
not known.
|
In mCD- or FB1-treated cells, the apical delivery of apical residents
was significantly impaired. Because both the rate and extent of their
basolateral internalization were not significantly decreased, we propose that
TGN to basolateral PM delivery was not impaired, i.e., the same amount of
protein was synthesized and available for transcytosis. Similarly, VSV-G
delivery from the TGN to the basolateral PM was not impaired in
cholesterol-depleted Madin-Darby canine kidney (MDCK) cells
(Keller and Simons, 1998). The
internalization results also indicate that raft depletion specifically
inhibits transcytotic efflux from basolateral early endosomes. This conclusion
is consistent with the detection of intracellular populations of the
transcytosing proteins in treated cells that colocalized with trafficked
ASGP-R and partially overlapped with Tf-R. We suggest these structures are
early endosomes, a common early transport intermediate of each of these
molecules. A block at the early endosome also accounts for impaired pIgA-R
transcytosis; it occurs downstream of clathrin-mediated uptake.
In contrast, endosomal trafficking of recycling receptors or HRP was not
significantly affected by lipid depletion. Internalization of these markers
was readily observed and their staining patterns were largely unchanged. The
finding that Tf-R and ASGP-R remained partially colocalized in treated cells
further implies Tf-R delivery to recycling endosomes via early endosomes was
maintained. Based on studies performed in cholesterol-depleted Chinese hamster
ovary cells where Tf-R recycling to the PM was unchanged
(Subtil et al.,
1999), we further propose Tf-R recycled normally to the
basolateral PM in lipid-depleted WIF-B cells.
Why was basolateral staining of most apical residents increased in treated cells when internalization was not impaired? Because there was no dramatic intracellular accumulation of internalized apical residents in treated cells, we propose that the apical residents rapidly recycled from the early endosome to the basolateral PM. Thus, normal TGN to basolateral delivery in combination with increased endosomal recycling would account for increased basolateral labeling. This is further substantiated by the finding that recycling, when assayed directly in Fao cells, was not significantly altered.
This model suggests that early endosomes contain raft domains. Although not
tested directly, other intracellular populations of cholesterol have been
identified. Recycling endosomes were found to be major storage compartments
for cholesterol when the dynamics of fluorescently labeled cholesterol analogs
were monitored (Mukherjee et al.,
1998). Immunoisolated recycling endosomes from polarized MDCK
cells were also found to be enriched in SM and the raft-associated proteins
caveolin and flotillin (Gagescu et
al., 2000). Late endosomes and lysosomes also have been shown
to contain cholesterol, but to much lesser extents
(Maxfield and Wustner,
2002).
The finding that basolateral internalization of apical proteins was not
impaired in treated cells further suggests that rafts are not present at the
basolateral PM or they do not mediate internalization of transcytosing apical
proteins. The former possibility is consistent with results from experiments
measuring fluorescence resonance energy between PM-associated raft markers
where rafts were found to be at best a minor component of the cell surface
(Kenworthy et al.,
2000). However, this contradicts reports from other studies
indicating that 70–80% of the PM contains raft-like domains based on
their solubility properties (Maxfield,
2002). Because other factors may also contribute to detergent
insolubility (see below), this may be an overestimation. The possibility that
raft depletion does not impair internalization has been suggested from
experiments examining PM-to-Golgi delivery of ricin, lactosylceramide, and
cholera toxin (Puri et al.,
1999; Rodal et al.,
1999; Shogomori and Futerman,
2001). In all cases, no decreases in internalization were
observed. However, clathrin-mediated internalization of epidermal growth
factor, transferrin, and Tf-R was severely impaired in cholesterol-depleted
Chinese hamster ovary and Hep-2 cells where invagination of coated pits was
perturbed (Rodal et al.,
1999; Shogomori and Futerman,
2001). Although we observed increased ASGP-R and Tf-R basolateral
staining in treated WIF-B cells, the impairment of internalization was
apparently mild given the intense intracellular labeling observed. This
discrepancy may be explained by the different experimental conditions used. We
had to use lower concentrations of mCD (5 versus 10 mM in other studies)
because WIF-B cells did not tolerate the higher doses (our unpublished data).
Similarly, in hippocampal neurons treated with lower concentrations of
mCD, Tf-R internalization was not impaired
(Shogomori and Futerman,
2001).
A General Component of the Transcytotic Trafficking Machinery
Requires Raft Association
Why does depletion of raft components inhibit transcytosis from early
endosomes? Findings that only a subset of the transcytosing proteins are
detergent insoluble and that lipid depletion does not significantly alter
their insolubility indicates that apical residents themselves do not require
raft association for transcytotic sorting. Thus, we suggest that
raft-dependent sorting is conferred by a general regulator of transcytosis
whose activity requires raft association. One good candidate for this
regulator is MAL2. The MAL family of raft-associated proteins are 20-kDa
tetra-spanning TMD proteins. Using an antisense approach, direct apical
delivery was found to be decreased in MDCK cells lacking MAL; the ectopic
expression of human MAL rescued the defect
(Puertollano et al.,
1999; Martin-Belmonte et
al., 2000). Thus, MAL has been implicated as an important
player in direct apical sorting. Interestingly, liver does not express this
MAL isoform, a finding consistent with the absence of direct apical delivery
of single-TMD and GPI-anchored apical residents in hepatocytes. Recently,
another MAL isoform was identified, MAL2, that is enriched in hepatic cells
(Wilson et al., 2001;
De Marco et al.,
2002). In HepG2 cells treated with antisense oligonucleotides,
transcytosis of pIgA was blocked from early endosomes to what appeared to be
the SAC (De Marco et al.,
2002). Thus, raft depletion may prevent MAL2 from sorting
transcytosing proteins from early endosomes.
Another possible class of regulators is the soluble
N-ethylmaleimide-sensitive factor attachment protein receptors
(SNAREs). In both PC12 and MDCK cells, PM-associated t-SNAREs were detected in
detergent-insoluble complexes (Lafont
et al., 1999;
Chamberlain et al.,
2001). Confocal microscopic inspection of isolated PMs further
revealed that t-SNAREs were present in clusters
(Lang et al., 2001).
Cholesterol depletion dispersed the clusters and correlated with decreased
exocytic activity (Chamberlain et
al., 2001; Lang et
al., 2001). Similarly, the apical SNAREs, syntaxin 3 and
Ti-VAMP, were solubilized by mCD in MDCK cells
(Lafont et al.,
1999). These results suggest that the cholesterol-dependent
organization of SNAREs is required for proper vesicle docking and fusion. If
this organization is shared by SNAREs present on intracellular organelles,
then the defect in transcytosis we observed might be explained by perturbed
vesicle docking and fusion with the SAC or apical PM.
Why Do the Same Molecules Have Different Solubility Properties and
What Imparts Detergent Insolubility?
The solubility properties of the apical residents presented in this study
do not completely agree with other published determinations. For example,
pIgA-R in intact enterocytes was 50% insoluble, whereas it was completely
soluble in WIF-B cells or FRT cells
(Hansen et al., 1999;
Sarnataro et al.,
2000). Also, endogenous DPP IV is soluble in WIF-B, but partially
insoluble in Fao cells, Caco-2 cells, and intact enterocytes
(Danielsen, 1995;
Alfalah et al., 2002).
There are several possible explanations for these disparate results. First,
not all soluble and insoluble fractions are created equally. Many methods are
cited for the preparation of detergent-insoluble domains. Perhaps the most
prevalent one used includes a short, low-speed centrifugation (11,800 x
g for 2 min). At the other extreme is a 30-min spin at 120,000
x g. When we tested these methods directly we found
insolubility varied two- to threefold for APN and 5'NT (our unpublished
data). Thus, it is important to note the methods used when making
comparisons.
Perhaps a more interesting explanation for the differential solubility properties may reflect important differences in the cellular context. In particular, variations in membrane lipid composition may alter detergent insolubility. Similarly, these differences may also point to specialized mechanisms for protein sorting present in specific epithelial cell types. Are different MAL isoforms or SNAREs regulating specific protein sorting steps? Also important to consider are the different protein concentrations in different cell types. The more concentrated population of DPP IV at the Fao PM was more insoluble than in WIF-B cells. The reasons for this are not clear, but this finding is important to consider when comparing results from different cell types.
What factors contribute to detergent insolubility? Although FB1
significantly decreased SM levels, all apical proteins tested maintained
control insolubility levels. FB1 also failed to solubilize GPI-anchored
proteins in FRT cells (Lipardi et
al., 2000). Likewise, addition of mCD did not alter the
solubility of GPI-anchored proteins in Fao or WIF-B cells, of cholera toxin in
T84 cells, or of galectin-4 in the intact enterocyte brush border
(Hansen et al., 2001;
Wolf et al., 2002).
Simultaneous depletion of both lipid species still rendered 5'NT 50%
insoluble. Although Triton X-100 insolubility was originally a diagnostic test
for cytoskeletal association, the addition of cytochalasin D alone or in
combination with mCD failed to solubilize all apical residents
(Figure 6;
Brown and Rose, 1992). Thus,
other unidentified factors are contributing to detergent insolubility.
Alternatively, this result may reflect the uncertain relationship between
rafts observed in vivo and detected in vitro.
Another Example of the Presence of Polarized Transport Pathways in
Nonpolarized Cells
The results presented in this study add to the growing list of similarities
between trafficking patterns in polarized and nonpolarized cells. Recently, we
reported that like polarized cells, newly synthesized apical proteins in
nonpolarized cells are delivered to the PM where they are selectively
internalized and delivered to structures containing only other apical proteins
(Tuma et al., 2002).
Likewise, in both polarized and nonpolarized cells, raft depletion impairs
trafficking of apical residents to these specific cellular destinations.
However, in nonpolarized cells, these apical proteins normally recycle back to
the PM, unlike their counterparts in fully polarized WIF-B cells where
actin-dependent mechanisms normally prevent return. This recycling is not
affected by raft depletion, leading to the disappearance of the apical
compartment despite continuous internalization. Thus, the machinery required
for specific intracellular sorting of apical residents is present in
nonpolarized cells. Also striking was the observation that apical proteins in
Fao cells had nearly identical solubility properties as in WIF-B cells.
Together, these results indicate that raft-dependent sorting does not depend
on the polarized state of a cell.
|
ACKNOWLEDGMENTS |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
|---|
|
Footnotes |
|---|
Abbreviations used: APN, aminopeptidase N; ASGP-R, asialoglycoprotein
receptor; DPP IV, dipeptidyl peptidase IV; FB1, fumonisin B1; GPI,
glycophoshatidylinositol; LPDM, lipoprotein deficient medium; mCD,
methyl--cyclodextrin; 5'NT, 5'nucleotidase; pIgA-R,
polymeric IgA receptor; PM, plasma membrane; SAC, subapical compartment; SM,
sphingomyelin; TLC; thin layer chromatography; Tf-R, transferrin receptor;
TMD; transmembrane domain.
Present address: Department of Biology, The Catholic University of America,
Washington, DC. 
* Corresponding author. E-mail address: alh{at}jhmi.edu.
|
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