Download Dendritic cells express tight junction proteins and penetrate gut

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Dendritic cells express tight junction
proteins and penetrate gut epithelial
monolayers to sample bacteria
Maria Rescigno1, Matteo Urbano1, Barbara Valzasina1, Maura Francolini2,
Gianluca Rotta1, Roberto Bonasio1, Francesca Granucci1,
Jean-Pierre Kraehenbuhl3 and Paola Ricciardi-Castagnoli1
Penetration of the gut mucosa by pathogens expressing invasion genes is believed to occur mainly
through specialized epithelial cells, called M cells, that are located in Peyer’s patches. However,
Salmonella typhimurium that are deficient in invasion genes encoded by Salmonella pathogenicity
island 1 (SPI1) are still able to reach the spleen after oral administration.This suggests the existence
of an alternative route for bacterial invasion, one that is independent of M cells.We report here a new
mechanism for bacterial uptake in the mucosa tissues that is mediated by dendritic cells (DCs). DCs
open the tight junctions between epithelial cells, send dendrites outside the epithelium and directly
sample bacteria. In addition, because DCs express tight-junction proteins such as occludin, claudin 1
and zonula occludens 1, the integrity of the epithelial barrier is preserved.
that are ideally located for antigen sampling in tissues that interface
with the external environment, such as skin and mucosae, where they
perform a sentinel function for incoming pathogens9–11. DCs with typical features of immature cells12 have been described, in PPs, as forming a dense layer of cells in the subepithelial dome, just beneath the
follicle epithelium13,14. They have also been described, in the lamina
propria of the gut, as inserting into the epithelium15 but they have never
been observed facing the gut lumen. DCs are in close contact with M
cells and colocalize with S. typhimurium 4 h after ligated loop injection16, thus indicating their major role in bacterial handling. However,
it is not clear whether DCs can directly take up bacteria across the
mucosal epithelium or whether they intervene only after bacteria has
been internalized by M cells. If DCs could take up bacteria directly
(because they are distributed along the entire intestinal epithelium),
The intestinal mucosa is covered by a single layer of epithelial cells that
are inaccessible to pathogens due to the presence of a brush border on
the lumenal cell surface and of tight junctions (TJs) between cells1,2.
Thus, the entry of pathogens3–5 occurs mainly through specialized
epithelial cells, called M cells, that are located in Peyer’s patches
(PPs)6. Penetration of M cells by bacteria requires the expression of
invasin proteins. Nevertheless, Salmonella typhimurium that are deficient in invasion genes encoded by Salmonella pathogenicity island 1
(SPI1) are still able to reach the spleen after oral administration7, which
suggests an M cell–independent pathway.
It has been postulated that CD18-expressing phagocytes are
involved in this alternative route for bacterial invasion because noninvasive Salmonellae are unable to invade CD18-deficient mice8. Among
CD18+ cells, dendritic cells (DCs) are migratory and phagocytic cells
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Figure 1. DCs induce transcytosis of fluorescent bacteria across a monolayer of differentiated human Caco-2 cells. A monolayer of Caco-2 cells was grown
on the lower face of 6.5-mm filters. Subsequently, D1 cells were seeded overnight facing the basolateral side of the Caco-2 monolayer and fluorescent (a) S. gordonii or (b)
S. typhimurium incubated from the apical side of the epithelium. Medium facing the basolateral side of the epithelium was collected at indicated times and analyzed by cytofluorimetry for the presence of fluorescent particles (see Methods for more details). (c) Caco-2 monolayers were incubated overnight with supernatant from the D1–Caco2 coculture (D1SN) and tested for transcytosis with S. typhimurium as in b.
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Department of Biotechnology and Bioscience, University of Milano-Bicocca, Milano, Italy. 2CNR, Cellular and Molecular Pharmacology Centre, Milano, Italy. 3Swiss Institute for
Experimental Cancer Research, University of Lausanne, Epalinges, Switzerland. Correspondence should be addressed to P. R. C. ([email protected]).
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Figure 2. DCs cross the filter and infiltrate the epithelial monolayer. (a–c)
D1 cells were added to the compartment facing the basolateral side of the Caco-2
monolayers. After 5 h of coculture, monolayers were incubated with bacteria added
on the apical side; the TER was unchanged throughout the whole experiment. Filters
were then fixed and processed for transmission electron microscopy. (a) D1 cells
crossed the filters and infiltrated between epithelial cells (EC) opening their TJs without changing microvilli organization (magnification: ×15,000). (b) and (c) The boxed
region in a was magnified further to reveal TJ-like structures, indicated by arrowheads, between DCs and Caco-2 in two serial sections (magnification: ×47,000).
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then they could enormously increase the mucosal surface exposed to
microorganisms. Thus, we have investigated the interaction of DCs
with epithelial cells both in vitro in a transwell coculture system, and
in vivo in the mouse, to dissect out the involvement of DCs in bacterial uptake. We have found that DCs express TJ proteins, open the TJs
between epithelial cells and can take up microorganisms directly, preserving the integrity of the epithelial barrier.
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Results
DCs allow the transcytosis of bacteria
To study the interaction of DCs with epithelial cells, we established a
transwell coculture system in vitro. We cultured a monolayer of the differentiated human enterocyte cell line Caco-2 on the lower face of 6.5mm Corning filters until a transepithelial electric resistance (TER) of
∼330 Ω/cm2 was achieved. Subsequently, growth factor–dependent D1
cells, which represented mouse immature DCs10,17,18, were seeded for 4
h on the other side of the filter, facing the basolateral membrane of the
Caco-2 cells. After incubation the transwells were extensively washed
to eliminate DCs that had not attached to the filter and bacteria were
seeded from the apical side to analyze whether the presence of DCs
would influence bacterial transport across the monolayer.
Trancytosis of fluorescent bacteria from the apical to the basolateral
side of the epithelium was analyzed by cytofluorimetry. We used the
auxotrophic S. typhimurium AroA SL7207 strain, which retains the
capacity to colonize the gut mucosa despite impairment of its replicating potential19, and Streptococcus gordonii, a nonpathogenic commensal bacterium20. We generated S. typhimurium that expresses green fluorescent protein (GFP), whereas S. gordonii was labeled with carboxyfluorescein diacetate succinimidyl ester (CFSE). We were able to discriminate between active transport through the epithelium or paracellular leakage of macromolecules by monitoring the passage of the fluorescent particles for 60 min at 4 °C and then at 37 °C after a rapid temperature shift.
Bacteria were detected at the basolateral side of the Caco-2 cells
only in the presence of DCs, which suggested that bacteria cannot be
transcytosed by the Caco-2 cells (Fig. 1a,b). Only a small proportion
of the bacteria seeded from the apical lumen was transported across the
epithelium: only 1 per 10000 seeded fluorescent bacteria were recovered from the DC side. Almost 50% of these were free in solution, with
the remaining 50% found associated with DCs that had detached from
the filter. Transport of bacteria across the Caco-2 monolayers is mediated by the direct interaction of DCs with epithelial cells because conditional medium from Caco-2–DC cocultures could not induce the
transcytosis of GFP-conjugated Salmonella (Fig. 1c).
DCs open TJs between epithelial cells
We next examined whether DCs induced the formation of M cells in the
cocultures and whether they disorganized the monolayers. DCs crossed
the filters and homed into the Caco-2 monolayers only when bacteria
were added apically, as revealed by transmission electron microscopy
(Fig. 2). In contrast to lymphocytes21, DCs did not induce the formation
of M cell–like cells. However, the DCs that had crept between epithelial cells were able to open the epithelial TJs without disorganizing the
brush border (Fig. 2 a,c); this could not be observed with B cells (data
Figure 3. DCs can uptake bacteria directly through a
monolayer of epithelial cells. D1 cells were labeled with
CMTMR and cells (4×105) were seeded on the upper face of
the filter (facing the basolateral side of Caco-2 monolayer) for
4 h. Subsequently, GFP-conjugated SL7207 (108 CFUs) were
incubated in the lower chamber (facing the apical side of the
epithelium) for 45 min.The TER did not vary throughout incubation. Filters were fixed in 1% PFA and analyzed by laser fluorescent confocal microscopy. GFP and CMTMR fluorescence
were recorded independently to exclude any interference in
their respective channels. Merged images show either (a) contact (c) colocalization of DCs with bacteria (yellow) (b) both
colocalization and contact or (d) the internalization. The
scheme depicts the filter organization: CMTMR-conjugated
D1 cells, red; GFP-conjugated bacteria, green; filter, black bar.)
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Figure 4. DCs express proteins involved in the formation of TJs. (a) D1 cells were either untreated (0 h) or incubated with S. typhimurium AroA at a ratio of 1:10
for 60 min in the absence of antibiotics. Cells were then washed and incubated for an additional hour (2 h) or for 17 more hours (18 h) in medium containing gentamycin
(50 µg/ml) and tetracyclin (10 µg/ml). ZO-1, E-cadherin, β-catenin, occludin and claudin-1 mRNAs were amplified by RT-PCR. (b) Occludin mRNA was quantified by TaqMan
PCR on mRNA derived from cells treated with LPS (10 µg/ml). (c,d) Protein expression by D1 cells or bone marrow–derived DCs that were untreated or treated with bacteria for the indicated times was analyzed with the use of immunofluorescence. (c) In agreement with the PCR data, ZO-1 staining was undetectable in nontreated cells. (d)
In contrast, occludin was already expressed at 0 h, up-regulated 2 h after bacterial encounter and mostly down-regulated at the later time-points. (e) Morphology of bone
marrow–derived DCs was viewed with Normarski optics. (f) β-catenin protein expression was quantified by immunoblot analysis of extracts from cells treated with LPS for
indicated times.
not shown). We also found that when low numbers of DCs (<4×105
cells) were seeded at the basolateral side, the TER was not affected
(data not shown). In addition, we found that the integrity of the epithelial barrier function was preserved by the presence of TJ-like structures
between DCs and epithelial cells (Fig. 2b,c, see arrowheads). DCs
added to the apical compartment did not migrate into the epithelial
monolayer (not shown), which suggests that interaction with epithelial
cells and/or the release of epithelial chemotactic signals is polarized.
bacteria, including attenuated Salmonella, is well established16,22,23,
whereas uptake from the apical membrane of differentiated Caco-2 is
minimal. Therefore, translocation of Salmonella (Fig. 1) can only
occur via DCs that are directly exposed on the apical side of the
monolayers or via dendrites sent through the TJs.
Thus, we incubated 108 colony-forming units (CFUs) of GFP-conjugated salmonellae from the apical side of the cocultured transwell and
analyzed the position of the bacteria in the filter throughout their transport across the monolayer (Fig. 3). We examined more than 100 sections per six different transwells, in three different experiments, and
observed that bacteria were found associated only with DCs stained
with CellTracker™ Orange CMTMR and not with Caco-2 cells (Fig.
3a,d), which indicated that uptake is restricted to DCs. Indeed GFPconjugated bacteria were found in contact with (Fig. 3a), inside (Fig.
DCs access the lumen and take up bacteria
The ability of DCs to gain access to the lumenal side of the epithelium and their role in mediating bacterial transport across the monolayer was analyzed by confocal fluorescent microscopy with the use
of GFP-conjugated Salmonella. The capacity of DCs to internalize
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Figure 5. DCs are recruited at the site of infection in vivo. Mice were anesthetized and stretches of
the small intestine corresponding to ∼3 cm were ligated
at their extremities. PBS or S. typhimurium (109) were
injected in the loop and at the indicated times, intestines
were isolated and snap frozen. (a) Cryosections (7 µm)
were fixed with 1% PFA and immunostained for CD11c
(green) and occludin (red). (Bar, 10 µm.) (b) The average±s.d. number of CD11c+ cells per section counted on
five sections per experiment.
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assessed by TaqMan polymerase chain reaction (PCR) and showed a
fourfold increase after 6 h of stimulation with LPS; only a slight
decrease was observed at late time-points (18 h after LPS activation)
(Fig. 4b). Consistent with the reverse transcription (RT)-PCR data, ZO1 was detected by immunofluorescence only after bacterial activation
of both D1 cells and bone marrow–derived DCs (Fig. 4c). In contrast,
after an initial increase, the labeling density of surface occludin
decreased over time, which was more pronounced in the D1 cells than
in bone marrow–derived DCs (Fig. 4d).
The reduction of occludin probably reflected protein instability
rather than a decrease in rate of synthesis because the corresponding
mRNA was only slightly down-regulated, as already described in
epithelial cells26. In addition, in bone marrow–derived DCs, occludin
labeling was first associated both with the soma and dendrites of DCs
(time 0 and 2 h after bacterial encounter, compare the fluorescent staining with the cell morphology in Fig. 4e). Later, occludin could be
observed only in the Golgi, which suggested new protein synthesis. The
pattern of expression of β-catenin was similar to that of occludin, with
a parallel down-regulation 6 h after bacterial activation (Fig. 4f).
Therefore, occludin and claudin 1 can mediate the interaction between
DCs and epithelial cells by opening preexisting TJs and forming new
TJ-like structures.
3c) and both inside and outside (Fig. 3b) DCs and during the DC internalization process (Fig. 3d). At this time-point (45 min into coincubation) we could count, per filter, an average of 1550±160 bacteria that
were associated with DCs, of which one-third were internalized. We
also found that there was no drop in the TER (not shown), an observation which was consistent with the capacity of DCs to establish tight
contacts with epithelial cells that were similar to TJs (Fig. 2b,c).
However, it is likely that some bacteria might remain attached in clusters to DC dendrites (as in Fig. 3c) and are released on the other side of
the epithelium without being internalized, thus explaining why 50% of
the bacteria were found free in solution after transcytosis.
DCs express TJ proteins
TJs consist of a continuous, circumferential belt that seals the apex of
epithelial and endothelial cells and prevents the paracellular diffusion
of solutes, macromolecules and microorganisms. The major constituents of the TJ strands include two integral membrane proteins—
occludin and claudin24—that are associated with the cytoplasmic proteins zonula occludens 1 (ZO-1), E-cadherin and catenins, which play
an important role in the formation and localization of TJs25. To understand how DCs open TJs up and form TJ-like structures with neighboring epithelial cells to maintain the integrity of the epithelial barrier
while sampling lumenal microorganisms, we tested whether D1 cells or
primary bone marrow–derived DCs express the machinery necessary to
assemble TJs.
DCs express all the TJ proteins, as was reflected by protein and
mRNA analysis (Fig. 4). We also found that
expression is regulated during the maturation
process, which is induced by bacteria or bacterial
a
lypopolysaccharide (LPS). Occludin, claudin-1,
E-cadherin and β-catenin mRNA were detected at
all time-points, whereas ZO-1 mRNA was
observed only 2 h after treatment with bacteria
(Fig. 4a). The amount of occludin mRNA was
Figure 6. In the presence of bacteria, DCs creep
between epithelial cells and send dendrites out to
the intestinal lumen in vivo. Ligated loops were injected
with GFP-conjugated S. typhimurium (109) and 30 or 60 min
later, intestines were isolated and snap frozen. Cryosections
(7 µm) were fixed with 1% PFA and immunostained for
CD11c. (a,b) DC dendrites were sent outside, to the
epithelium, by 30 min after infection (see arrow). The inset
in b is magnified ×1.7. (c,d) DCs are surrounded by epithelial cells (see arrow) or (e) are found in the lumen, although
they are still attached to the epithelium (see arrow). (f,g)
Precise DC processes creeping out of the epithelial barrier
and contacting GFP-conjugated S. typhimurium (see arrows).
(a–e) CD11c stains green, occluding stains red (bar, 5 µm in
a–c; 3 µm in d; 4 µm in e). (f–g) CD11c stains red, GFP-conjugated S. typhimurium stains green (bar, 3 µm).
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DCs also take up bacteria in vivo
To assess whether, in vivo, DCs could also take up bacteria across
epithelia, we experimented on the ligated loop of the small intestine in
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Figure 7. DCs are recruited in the intestine
and can also uptake nonpathogenic E. coli.
Ligated loops were infected with (a,b,e) E. coli
DH5α (109) expressing the red fluorescence protein
(D5) or (c,d,f) a combination of 5×108 D5 + 5×108
S. typhimurium (SL) for 30 or 60 min. Intestines were
then isolated and snap frozen. (a–d) Cryosections (7
µm) were fixed with 1% PFA and immunostained for
CD11c (green) and occludin (red). CD11c+ cells
were found (a) creeping between epithelial cells (see
arrows) in search of E. coli D5 (red, circled) and (b)
internalizing bacteria (see arrow and inset, which is
magnified ×2), which stained yellow due to the colocalization. (c–d) Coinfection of E. coli (D5) with S.
typhimurium (SL) increases the number of recruited
DCs and accelerates the process and magnitude of
bacterial internalization. (c) Cell full of bacteria (see
arrow indicate and inset, which is magnified ×2). (d)
DC dendrite outside the epithelium during the
process of internalization. Some red bacteria are still
visible (inset magnification: ×2). (e–f) Average±s.d.
number of CD11c+ cells (open bars) or CD11c+D5+
cells (filled bars) in each section after counting five
sections per experiment. (Bars, 7 µm in a,c; 8 µm in
b,d.)
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CD11c+ cells internalizing bacteria was very similar both in the presence and absence of pathogens (13–15%). In addition, 1 h after infection with a mixture of Salmonella + E. coli (but not with E. coli only)
CD11c+ cells carrying bacteria were found deeper in the lamina propria
at the base of the villi, which suggested they had migrated outside the
intestine (not shown). This indicates that DCs play a major role in bacterial sampling across the mucosae in vivo and that they can be responsible for the internalization of pathogenic and nonpathogenic bacteria,
that is, bacteria that are unable to induce their own phagocytosis
through the M cells.
mice. We infected the loops with GFP-conjugated Salmonella or PBS,
as a control. Thirty minutes, 1 or 2 h later the ligated loops were snapfrozen and processed for immunohistochemistry with anti-CD11c to
detect DCs (Fig. 5a). First, we observed that DCs were quickly
recruited into the infected loops, peaking at 30 min (Fig. 5b). The
increase in DC numbers was rapid and transient because the quantity
of DCs was comparable to that observed in noninfected mice a few
hours before infection (numbers of CD11c+ cells increased sevenfold
after 30 min, 2.6-fold after 1 h and 1.6-fold after 2 h). Just 30 min after
bacterial infection, DC dendrites were sent into the lumen (Fig. 6) and
some of them were found in close contact with GFP-conjugated bacteria (Fig. 6f,g, see arrows). We also found that when the villi were
sectioned very close to their edges, DCs were found either surrounded
by epithelial cells (Fig. 6c) or outside of, but still in contact with, the
epithelium (Fig. 6e).
Discussion
The internalization of dietary antigens appears to be widespread
throughout the intestinal epithelium, and is carried out by epithelial
cells27, whereas the uptake of bacteria mainly occurs in the PPs, via
the M cells6. Although only invasive bacteria can induce their own
phagocytosis through M cells, noninvasive bacteria have been shown
to enter the epithelium at a lower efficiency7. It has been postulated
that CD18+ cell types, which include DCs, are involved in this alternative route because noninvasive Salmonellae could not invade
CD18-deficient mice8.
In this study, we describe a newly identified mechanism that allows
DCs to sample environmental microorganisms without compromising
the epithelial barrier function and to deliver them into lymphoid tissues
where an efficient immune response can be mounted. Indeed, we have
shown that, after bacterial infection, DCs are recruited to the inflamed
site, probably attracted by inflammatory chemokines such as
macrophage inflammatory protein 3α (MIP-3α) that are secreted by
epithelial cells14,28. Next, DCs are induced to up-regulate TJ proteins
and establish TJ-like structures with epithelial cells to take up antigen
without changing the TER. This type of junction is not species-restricted because it occurs between human epithelial cells and mouse DCs,
which is probably due to the high degree of conservation of these proteins in the two species (∼90% identity).
The mechanism that allows the DCs to destabilize preexisting epithelial TJs is not known. A peptide that corresponds to one of the two
extracellular loops of occludin (OCC2 peptide) has been shown to perturb, in a dose-dependent and reversible manner, the tightness of the
Nonpathogenic bacteria can induce DC recruitment
The capacity of DCs to phagocytose both pathogenic and nonpathogenic bacteria is well established9,22,23. Thus, we studied the ability of
DCs to take up nonpathogenic bacteria across intestinal epithelia in
vivo. Two types of experiments were carried out: ligated loops were
infected either with 109 CFUs of nonpathogenic Escherichia coli
(DH5α) expressing the red fluorescent protein (DsRed, D5) or with a
combination of 5×108 CFUs of unlabeled pathogenic Salmonella +
5×108 CFUs of DsRed-conjugated nonpathogenic E. coli. In both cases
it was found that DCs home to the epithelium and take up bacteria
directly (Fig. 7). In the presence of pathogens, the number of CD11c+
cells recruited to the lumen was doubled when compared to E. coli
alone (647±96 versus 275±26 after 30 min; compare Fig. 7e,f).
However, although the number of CD11c+ cells decreased over time,
when the loops were injected with Salmonella it remained constant
after infection with E. coli alone (Fig. 7e).
DC dendrites directly capturing bacteria (Fig. 7d) as well as colocalization of bacteria with DCs could be observed (Fig. 7) in both conditions, although with different kinetics and magnitude. Indeed, the
presence of pathogenic bacteria accelerated the process and increased
the number of DCs internalizing bacteria threefold. This is probably
due to the higher number of recruited DCs because the percentage of
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In the presence of noninvading bacteria, an increase in the number of
DCs in the intestine and, in particular, in the epithelium, as well as an
increase in the uptake of microorganisms, could be observed. However,
unlike the effect of other pathogens, the number of DCs present in the
apex of villi remained constant over time and cells carrying bacteria
could not be observed deeper in the villi, which suggests that the DCs
stayed in the lamina propria. Immunoglobulin A (IgA) responses to
commensal bacteria can be induced by a primitive T cell–independent
mechanism that does not require organized follicular lymphoid tissue33.
This suggests the involvement of DCs in direct IgA induction against
commensal bacteria in the lamina propria. Finally, this mechanism
could also be involved in the transport of apoptotic intestinal epithelial
cells to T cell areas of mesenteric lymph nodes, as has been suggested11. Indeed, DCs could engulf exhausted epithelial cells without perturbing the integrity of the epithelial barrier.
Thus, our studies provide in vivo evidence of bacterial uptake at
mucosal surfaces. This mechanism could be exploited by pathogenic
microorganisms in order to spread throughout the body and by the
immune system for induction of a primitive IgA response to microorganisms.
Figure 8. Scheme of the events occurring during a bacterial infection.
Under resting conditions, infiltrating DCs establish loose contacts with preexisting
epithelial TJs. Upon bacterial infection, DCs are recruited from the blood and activated, probably via epithelial cell signals.They up-regulate the expression of occludin,
which in turn allows DCs to compete for epithelial occludin and open up the TJs, like
a zip. Infiltrating DCs then face the gut lumen and can directly sample bacteria.
Bacterial components such as LPS trigger the reorganization of TJ proteins via upregulation of ZO-1 and the disappearance of occludin, thus allowing the DCs to
detach from junctions with epithelial cells and to migrate into the draining lymph
nodes.
TJs of confluent epithelial monolayers26. By analogy, the presence of
occludin in DCs may be sufficient to loosen the epithelial TJ, a destabilization that is followed by the rapid formation of new junctions
between the epithelium and the infiltrating DCs. Occludin is constitutively expressed in immature DCs, and is distributed uniformly in the
cells. We propose that, under resting conditions, infiltrating DCs establish loose contacts with preexisting epithelial TJs (Fig. 8). Upon bacterial infection, DCs are recruited from the blood and are activated, probably via epithelial cell signals. They then up-regulate the expression of
occludin and distribute it to the cell surface and dendrites. This allows
DCs to compete for epithelial occludin and open up the TJs like a zip.
Infiltrating DCs then face the gut lumen and can directly sample bacteria. Bacterial components such as LPS trigger the reorganization of TJ
proteins with up-regulation of ZO-1 and the disappearance of occludin
from the cell surface. This is presumably as a result of proteolytic
degradation because the mRNA expression is only slightly down-regulated. This allows the DCs to detach from the junctions with epithelial
cells and to migrate into the draining lymph nodes, after a change in the
chemokine receptor program triggered by bacterial components such as
LPS29. Probably other “danger” signals delivered to DCs in situ accelerate the reorganization of TJ proteins because the number of CD11c+
cells is drastically reduced 1 h after infection with pathogens, although
not with nonpathogenic bacteria.
It is also known that neutrophils are able to migrate across epithelia
without affecting the TER and that occludin is involved in the modulation of this process30. Thus, neutrophils could employ the same mechanism to get through epithelial cells without perturbing the epithelial
barrier. In addition, a similar mechanism for the opening of TJs
between endothelial cells could be employed by leukocytes for their
transmigration in the cerebrospinal fluid during experimental meningitis, as blockage of the interaction with junctional adhesion molecule
(JAM) by a specific antibody inhibits the recruitment of both monocytes and neutrophils in the brain parenchyma31.
Because DCs are migratory cells, they can transport pathogens to the
mesenteric lymph node and the spleen for the induction of systemic
responses8, which suggests that this alternative route of bacterial internalization has an important physiological relevance. Indeed the number
of CD11c+ cells is only transiently increased in the intestinal villi when
the loops are infected with pathogens. Accordingly, it has been reported that, in caspase 1–deficient mice, Salmonella do not colonize the
PPs but that they are contained within the spleen32. We propose that the
bacteria are carried to the spleen by DCs directly after their uptake.
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Methods
Cells, bacterial strains and reagents. The D1 cells were maintained in vitro as growth
factor–dependent immature DCs in complete Iscove’s modified Dulbecco’s medium supplemented with 30% R1 conditioned medium containing granulocyte-macrophage colonystimulating factor (30 ng/ml), as described17. Bone marrow–derived DCs were generated
from a single cell suspension of marrow from the femurs of C57BL/6 mice. Bone marrow
cells were split every 5 days with PBS to detach only loosely adherent cells. After 15–20
days of culture in DC medium, the homogeneity of DCs was evaluated by cytofluorimetry.
Only cultures with >90% cells expressing intermediate amounts and <10% cells expressing high amounts of major histocompatibility class II (I-Ab) and B7.2 were used for the
experiments. Caco-2 cells were grown in Dulbecco’s modified Eagle’s medium supplemented with 10% fetal calf serum and nonessential amino acids. The auxotrophic S.
typhimurium aroA strain SL7207—S. typhimurium 233765 derivative hisG46, DEL407
(aroA:Tn10[Tc-s])—was provided by B. A. D. Stocker (Stanford, CA). The wild-type S.
gordonii strain GP204 was provided by G. Pozzi (Siena, Italy)20. GFP-conjugated SL7207
was generated by transformation with a plasmid (pkk223.3) containing the gene encoding
GFP. S. gordonii was labeled with CFSE according to manufacturer’s instructions. E. coli
DH5α expressing the red fluorescent protein (DsRed, D5) was generated by transformation with the plasmid pDS-Red (Clontech, Palo Alto, CA). Bacteria were grown in brainheart infusion at density of A600=0.6. GFP-conjugated D1 cells were generated as
described34. Labeling of cells with CellTracker™ Orange CMTMR (5-(and-6)–4chloromethyl-benzoyl-amino-tetramethylrhodamine, Molecular Probes, Eugene, OR) was
according to manufacturer’s instructions.
Caco-2-D1 Transwell coculture system. Caco-2 cells were seeded either on the lower or
upper face of 6.5-mm filters (3-µm pore Transwell filters, Costar, Cambridge, MA) for
10–15 days in a 24-well plate until a TER of ∼330 Ω/cm2 was achieved. D1, GFP-conjugated D1 or CMTMR-conjugated D1 cells (4×105) were cultured either on the basolateral
or the apical side of Caco-2 monolayer for 2–18 h. Fluorescent bacteria (108 CFUs) were
always incubated from the apical side of the epithelium first at 4 °C for 60 min and then at
37 °C. At different time-points after addition of D1 cells or after subsequent addition of bacteria, the TER was measured and the composition of medium facing the basolateral side of
Caco-2 cells analyzed by FACScan (Becton Dickinson, San Jose, CA). Filters were either
fixed with 1% paraformaldehyde (PFA) for 60 min at 4 °C and processed for fluorescent
confocal microscopy or with glutarahldehyde and processed for transmission electron
microscopy.
Immunofluorescence staining and immunoblotting. DCs were plated on poly(L)
lysine–treated coverslips and incubated either with LPS (E. coli serotype 026:B6, Sigma, St.
Louis, MO) or with bacteria (multiplicity of infection of 10) for varying times between 2–18
h. Cells were fixed in 1% paraformaldehyde for 60 min at 4 °C and processed for immunofluorescence. Fluorescein isothiocyanate– or phycoerythrin-conjugated antibodies to
CD11c were from PharMingen (San Diego, CA). Rabbit anti-occludin was from Zymed
Laboratories (San Diego, CA), rabbit anti–β-catenin and anti–E-cadherin were from Sigma
and mouse anti–ZO-1 clone T8-754 was a gift of M. Furuse (Kyoto, Japan). For
immunoblot analysis, 10–20 µg of total cell extracts were loaded on 10% SDS-polyacrylamide gels. Electrophoretically separated proteins were blotted on nitrocellulose filters.
Filters were blocked and incubated with primary antibodies as above.
RT-PCR. Total RNA was prepared from 5×106 SL7207-treated and untreated DCs by TRIzol
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A RTICLES
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reagent (Gibco-BRL, Gaithersburg, MD), according to manufacturer’s instructions, and
mRNA was reverse-transcribed to cDNA. The following primers were used for PCR amplification: Occludin: 5′–CGGCTATGGAGGCTATGGCTATG–3′ and 5′–ATGAACCCCAGGACAATGGC–3′; ZO-1: 5′–ATCCCAAATAAGAACAGAGC–3′ and 5′–GGCGTTACATCTA
ATAAAGC–3′; E-cadherin: 5′–GCACATATGTAGCTCTCATC–3′ and 5′–CCTTCACAGTC
ACACACATG–3′; β-catenin: 5′–CTGTTCTACGCCATCACGAC–3′ and 5′–TGAAAGGTT
TCTGAGAGTCC–3′; Claudin 1: 5′–TTGTTTGCAGAGACCCCATCAC–3′ and 5′–GGAGT
AAATCTTCCACTGGGGC–3′.
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Acknowledgments
We thank K. Giese (Atugen, Berlin) for the TaqMan analysis on occludin mRNA and R.
Steinman and D. Grdic for helpful discussions. Supported by grants from the Italian
Association against Cancer (AIRC); the National Research Council (CNR Project in
Biotechnology); the EC contract MUCIMM; and the Swiss National Science Foundation.
Received 8 January 2001, accepted 26 February 2001.
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