TLRs Regulate the Gatekeeping Functions of
the Intestinal Follicle-Associated Epithelium
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J Immunol 2006; 176:4275-4283; ;
doi: 10.4049/jimmunol.176.7.4275
http://www.jimmunol.org/content/176/7/4275
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References
Sophie Chabot, Jessica S. Wagner, Stephanie Farrant and
Marian R. Neutra
The Journal of Immunology
TLRs Regulate the Gatekeeping Functions of the Intestinal
Follicle-Associated Epithelium1
Sophie Chabot, Jessica S. Wagner, Stephanie Farrant, and Marian R. Neutra2
T
he intestinal epithelium is a delicate barrier separating the
foreign macromolecules and microorganisms in the intestinal lumen from the internal milieu. The vast majority of
the small intestinal surface consists of villi that are covered by a
tight monolayer of epithelial cells concerned primarily with absorption of nutrients, maintenance of electrolyte balance, and exclusion of microorganisms. Epithelial cells on intestinal villi act as
sensors of the luminal environment, and intestinal epithelial cell
monolayers in vitro can release inflammatory cytokines in response to pathogen attachment and invasion (1, 2). Intestinal epithelial cells express a variety of potential sensing receptors, including TLRs (3–19), that may play a role in mucosal responses to
microbial components.
TLRs recognize specific patterns of microbial components present
in glycoproteins, lipoproteins, LPS, glycolipids, peptidoglycan
(PGN),3 fatty acids and nucleic acids of bacterial, viral, parasitic, or
fungal origin (20, 21). On most cell types including APCs, activation
of TLRs by ligand binding induces signal transduction cascades that
result in an expression of inflammatory and immunomodulatory genes
Department of Pediatrics, Harvard Medical School, GI Cell Biology Laboratory, Children’s Hospital Boston and Harvard Digestive Diseases Center, Boston, MA 02115
Received for publication October 12, 2005. Accepted for publication January
26, 2006.
The costs of publication of this article were defrayed in part by the payment of page
charges. This article must therefore be hereby marked advertisement in accordance
with 18 U.S.C. Section 1734 solely to indicate this fact.
1
This study was supported by National Institutes of Health (NIH) Research Grant
HD17557, and NIH Center Grant DK34854 (to the Harvard Digestive Diseases Center). S.C. is the recipient of a Canadian Institute of Health Research Post-Doctoral
Fellowship.
2
Address correspondence and reprint requests to Dr. Marian R. Neutra, GI Cell Biology Laboratory, Enders 720, Children’s Hospital Boston, 300 Longwood Avenue,
Boston, MA 02115. E-mail address: [email protected]
3
Abbreviations used in this paper: PGN, peptidoglycan; PP, Peyer’s patch; FAE,
follicle-associated epithelium; DC, dendritic cell; SED, subepithelial dome; MMP,
matrix metalloproteinase; PFA, paraformaldehyde; UEA, Ulex europaeus agglutinin;
WGA, wheat germ agglutinin; DAPI, 4⬘,6⬘-diamidino-2-phenylindole; PGRP, peptidoglycan recognition protein.
Copyright © 2006 by The American Association of Immunologists, Inc.
involved in the regulation of both innate and adaptive immunity
(22–24). Although villus and crypt epithelial cells express TLRs at
their apical poles in vivo (3, 25, 26), the intestinal mucosa nevertheless coexists with the commensal microflora without chronic inflammation (27), and cultured transformed intestinal epithelial cells that
express TLRs in vitro were shown to be relatively unresponsive to
TLR ligands (6, 7, 28). It is thought that sensing of commensal
bacteria through epithelial TLRs in vivo does not initiate inflammatory cascades but rather is required for the maintenance of the
epithelial barrier (29), and contributes to TLR-dependent intestinal
homeostasis observed in vivo (30).
At sites of organized mucosal lymphoid tissues, the intestinal
epithelium is dramatically modified to fulfill a different function.
Mucosal lymphoid follicles, whether solitary or clustered as in
Peyer’s patches (PP), are covered by a specialized follicle-associated epithelium (FAE) containing M cells. M cells take up intact
macromolecules, particles, and pathogens, delivering samples of
the luminal contents by transepithelial transport (31) to underlying
subepithelial dendritic cells (DCs) that go on to initiate adaptive
mucosal immune responses (32, 33). Although M cells constitute
a minority in the FAE, representing only ⬃10% of FAE cells in
humans and mice, they are clearly important in microbial pathogenesis and vaccine targeting because they provide a gateway to
inductive sites of the mucosal immune system. Adherence of Ags and
microorganisms to M cells greatly increases the efficiency of uptake
into PP (31). It was recently reported that M cells in PP of swine
express TLR2 (34) and TLR9 (35), but whether TLRs are expressed
by M cells in other species, and whether they serve as microbial
receptors or have other functions in these cells, is not known.
Relatively little is known about the non-M epithelial cells, the
FAE enterocytes, which constitute most of the FAE. FAE enterocytes are morphologically indistinguishable from absorptive enterocytes on villi, but they lack key functions of villus and crypt
cells. They express very low levels of brush border hydrolases and
thus do not participate in nutrient absorption (36), and they lack
polymeric Ig receptors and thus do not secrete IgA (37, 38). FAE
enterocytes produce chemokines, including CCL20 (MIP3␣) and
0022-1767/06/$02.00
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Initiation of adaptive mucosal immunity occurs in organized mucosal lymphoid tissues such as Peyer’s patches of the small
intestine. Mucosal lymphoid follicles are covered by a specialized follicle-associated epithelium (FAE) that contains M cells, which
mediate uptake and transepithelial transport of luminal Ags. FAE cells also produce chemokines that attract Ag-presenting
dendritic cells (DCs). TLRs link innate and adaptive immunity, but their possible role in regulating FAE functions is unknown.
We show that TLR2 is expressed in both FAE and villus epithelium, but TLR2 activation by peptidoglycan or Pam3Cys injected
into the intestinal lumen of mice resulted in receptor redistribution in the FAE only. TLR2 activation enhanced transepithelial
transport of microparticles by M cells in a dose-dependent manner. Furthermore, TLR2 activation induced the matrix metalloproteinase-dependent migration of subepithelial DCs into the FAE, but not into villus epithelium of wild-type and TLR4-deficient
mice. These responses were not observed in TLR2-deficient mice. Thus, the FAE of Peyer’s patches responds to TLR2 ligands in
a manner that is distinct from the villus epithelium. Intraluminal LPS, a TLR4 ligand, also enhanced microparticle uptake by the
FAE and induced DC migration into the FAE, suggesting that other TLRs may modulate FAE functions. We conclude that
TLR-mediated signals regulate the gatekeeping functions of the FAE to promote Ag capture by DCs in organized mucosal
lymphoid tissues. The Journal of Immunology, 2006, 176: 4275– 4283.
4276
Materials and Methods
Mice
BALB/c, C57BL/6, C57BL/10, and C57BL/10ScNCr (TLR4-deficient) female mice 6 – 8 wk of age were obtained from Charles River Laboratories.
C57BL/6-derived TLR2-deficient mice (originally produced by Dr. S.
Akira, Osaka University, Osaka, Japan) were generously provided by Dr.
D. Golenbock (University of Massachusetts, Worcester, MA). Mice were
maintained in laminar flow cages and were free of specific microbial and
viral pathogens as determined by plasma Ab screening. All animal procedures were performed in compliance with the guidelines for animal experimentation of Harvard Medical School, the Children’s Hospital Boston,
and the National Institutes of Health.
Ligated intestinal loops
Mice were anesthetized by i.p. injection of avertin (tribromoethanol in
t-amyl alcohol; 200 mg/kg animal weight), and a 3- to 5-cm segment of
ileum containing a PP was ligated and injected as described previously
(43). Loops were injected with 200 ␮l of PBS containing TLR agonists,
which include Staphylococcus aureus PGN (Fluka; Sigma-Aldrich),
Pam3Cys (EMC), or ultrapure Escherichia coli LPS (InvivoGen) at concentrations of 0.01, 0.1, or 1 mg/ml. Where indicated, TLR ligands were
injected in combination with fluoresbrite plain yellow-green 0.2-␮m microspheres (Polyscience), 2 ␮g/ml anti-TLR2 Abs (Santa Cruz Laboratories), 5 ␮g/ml GM6001 matrix metalloproteinase (MMP) inhibitor (Iomastat; Chemicon) or PBS alone. After 45 or 90 min, animals were sacrificed,
and PP were excised, washed in cold PBS, immediately embedded in Tissue-Tek OCT-embedding medium (Sakura Finetek), frozen in 2-methylbutane (Sigma-Aldrich) cooled with liquid nitrogen, and stored at ⫺80°C
until sectioned. For whole-mount preparations, PP were excised and fixed
in PBS containing 4% paraformaldehyde (PFA) before staining.
Immunohistochemistry
Immunohistochemistry was performed on fresh frozen tissue sections and
on PFA-fixed whole-mount PP preparations. Cryosections (3 ␮m) of frozen
PP were cut on a cryostat model CM3050 (Leica), mounted on S/P Superfrost Plus microscope slides (Allegiance), air-dried overnight, and fixed
with cold acetone for 2 min. Cryosections or PFA-fixed whole-mount PP
preparations were blocked using PBS containing 2% of goat serum, and the
avidin/biotin blocking kit (Vector Laboratories). The following primary
anti-TLR Abs were used: rabbit anti-mouse TLR2 (H-175), TLR4 (H-80),
TLR5 (H-127), TLR9 (H-100) (from Santa Cruz Laboratories), and rat
anti-mouse TLR4/MD2 complex (clone MTS510; eBioscience). Normal
rabbit IgG (Santa Cruz Laboratories), purified rat IgG2a (eBioscience), or
omission of primary Abs served as negative controls. Secondary Abs included biotinylated goat anti-rabbit IgG (SouthernBiotech) for TLR detec-
tion and biotinylated mouse anti-rat IgG (eBioscience) for TLR4/MD2 detection, both amplified using the ABC amplification kit (Vector
Laboratories) followed by incubation with Streptavidin-Alexa555 (Molecular
Probes). DCs were visualized using biotin-conjugated rat anti-mouse-CD11c
(BD Pharmingen) followed by streptavidin-Alexa555. Laminin was stained
with rabbit anti-mouse laminin (ICN) followed by goat anti-rabbit-Alexa488
(Molecular Probes). Counterstaining was performed using FITC-conjugated
lectin Ulex europaeus agglutinin (UEA)-1 (Vector Laboratories) for BALB/c
mice, and FITC- or Cy5-conjugated lectin wheat germ agglutinin (WGA)
(Vector Laboratories). DAPI (4⬘,6⬘-diamidino-2-phenylindole) (Biochemika)
was used for staining of nuclei in tissue sections, and stained sections were
mounted with Mowiol (Calbiochem-Novabiochem).
Microscopy
Stained tissue sections were examined and photographed using a Zeiss
Axiophot microscope (Zeiss) equipped with a Spot camera (Diagnostic
Instruments), or with a Nikon Eclipse TE2000 inverted PerkinElmer ultraview confocal microscope equipped with Yokogawa spinning disk scan
head and Orca ER AG cooled CCD camera (Hamamatsu); Slidebook software (Intelligent Imaging Innovations) was used for image capture and
analysis. Whole-mount PP preparations were viewed with a Nikon Eclipse
E800 confocal microscope equipped with a BioRad MRC-1024 and the
Lasersharp 2000 software.
Quantitation and statistics
For quantitative comparisons of microparticle uptake and DC migration, the
ligated PP from each of four mice in each treatment group was sectioned
through and for each PP, four different FAEs associated with separate lymphoid follicles in sections, which included the apex of the follicle dome, were
selected and photographed. To quantitate microparticle uptake, a standard area
encompassing 0.04 mm2 that included the FAE and underlying tissue was
defined on each photograph. Within this area, all fluorescent microparticles
associated with the FAE and within the tissue were counted, and particle
counts from four FAE areas were totaled to obtain a standard FAE sample for
each mouse. Data were expressed as the mean number of particles per standard
FAE sample (n ⫽ 4 mice) ⫾ the SDs. To quantitate relative numbers of
intraepithelial CD11c⫹ DCs within the FAE, a segment of FAE measuring 223
␮m in length was demarcated, all CD11c⫹ cells located within the segment
were counted, and DC counts in four FAE segments were totaled to obtain a
standard FAE sample for each mouse. To quantitate DCs within the villus
epithelium of the same mice, separate villi sectioned longitudinally and lying
within a 0.04-mm2 area of the PP sections were identified, and the total number
of CD11c⫹ cells within the epithelium of four villi, a standard villus sample,
was recorded for each mouse. Data were expressed as mean DCs per standard
FAE sample or per standard villus sample ⫾ the SDs. Statistical analyses were
performed using Instat (GraphPad). Differences between groups were analyzed
using the Bonferroni multiple comparison one-way ANOVA. Differences with
ⴱ, p ⬍ 0.05; ⴱⴱ, p ⬍ 0.01; and ⴱⴱⴱ, p ⬍ 0.001 were considered statistically
significant. To analyze the difference between LPS-treated samples and respective PBS controls, an unpaired t test was performed.
Results
TLR2 expression is polarized in intestinal FAE cells
Expression of TLR2 in epithelia of unmanipulated animals was
studied using whole-mount preparations of intact mouse PP and thin
(3 ␮m) frozen tissue sections. Surface staining of the epithelium in
formaldehyde-fixed whole mounts of BALB/c mouse PP with WGA
allowed us to visualize the surface of the FAE on an individual
lymphoid follicle surrounded by villi (Fig. 1a). Available anti-TLR2
Abs do not recognize TLR2 in formaldehyde-fixed tissues, and
unfixed PP degenerates rapidly ex vivo. Therefore, to obtain an
overview of TLR2 distribution on the surface of intact, fresh PP
mucosa, rabbit anti-mouse TLR2 Abs or control rabbit IgG was
injected into ligated segments of ileum in vivo, and 1 h later tissue was
excised, fixed, and labeled with fluorophore-conjugated secondary
Abs. The fixed tissue was treated with fluorophore-conjugated UEA-1
to label apical surfaces of M cells and fluorophore-conjugated WGA
to label all epithelial surfaces, and tissue was analyzed by confocal
microscopy. A 1.5-␮m stack of confocal planes containing WGAlabeled epithelial surfaces was selected to visualize the apical poles of
epithelial cells including enterocyte brush borders, M cell surfaces,
and apical endosomes. The WGA signal was subsequently subtracted
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CCL9 (MIP1␥), that are not expressed by villus epithelial cells and
that attract DCs to the subepithelial dome (SED) region under the
FAE (39 – 41). This presumably benefits the host by positioning
DCs for efficient capture of the foreign materials transported by M
cells. Whether TLRs are expressed on FAE enterocytes, and
whether they play a role in the specialized functions of the FAE,
has not been examined.
This study was initiated to investigate the expression and localization of TLRs in epithelial cells of the FAE in mouse PP, including TLR2, -4, -5, and -9 that recognize pathogen-associated
molecular patterns commonly present in enteric microorganisms
and that are targets of mucosal vaccine adjuvants (42). Because the
specialized phenotypes of epithelial cells and their interaction with
DCs in PP have not been reproduced in vitro, we focused this study
on PP in vivo. The distribution of epithelial TLRs was polarized in
epithelial cells of the FAE. Injection of TLR2 ligands into the
intestinal lumen in vivo had dramatic effects on TLR2 localization
in FAE cells, but not in epithelia of villi or crypts. TLR2 ligands
enhanced M cell transport of microparticles and induced DC migration into the FAE. Similar effects were observed using the
TLR4 ligand LPS. Thus, the FAE has a unique response to TLR
ligands, and TLR-mediated signals promote mucosal Ag sampling
in organized mucosal lymphoid tissues. These findings are potentially important for understanding microbial pathogenesis in mucosal tissues and for the design of mucosal vaccines.
TLRs AFFECT FUNCTIONS OF THE FAE
The Journal of Immunology
4277
from the confocal images shown. Anti-TLR2 Abs were present at
apical poles of both FAE and villus epithelium (Fig. 1b). In the FAE,
highest levels of anti-TLR2 staining occurred on FAE enterocytes,
and lower levels were observed on some (but not all) UEA-1⫹ M cells
(Fig. 1b). Tissues of normal BALB/c mice exposed to nonspecific
rabbit IgG showed no significant label (Fig. 1b), and no staining was
observed when anti-TLR2 was injected into ligated loops of TLR2deficient mice (data not shown). Because labeling of the intact
mucosal preparations in vivo presumably allowed uptake of TLR2-Ab
complexes, the distribution of TLR2 was confirmed by application of
anti-TLR2 Abs to frozen sections of unfixed PP tissue. This revealed
TLR2 on or near the apical surfaces of FAE, villus, and crypt
epithelial cells, including a subset of M cells (Fig. 1, c and f). TLR2
was also present at the basal poles of FAE cells (Fig. 1c) but not at
basal poles of villus or crypt epithelial cells (Fig. 1f). Double-staining
with WGA, which labels the brush border glycocalyx located above
the tips of epithelial microvilli, showed that TLR2 was localized
below the glycocalyx, consistent with an apical membrane and/or
apical endosome distribution (Fig. 1d). A similar distribution of TLR2
was seen in PP tissues of C57BL/6 mice (data not shown). Control
tissue sections labeled with nonspecific rabbit IgG (Fig. 1e), and
anti-TLR2 labeling of sections from TLR2-deficient mice (Fig. 1g)
showed no staining, confirming the specificity of the anti-TLR2
Abs used.
Luminal TLR2 ligands induce the redistribution of TLR2 in cells
of the FAE but not in villus or crypt epithelium
To test the effect of receptor ligation on the distribution of TLR2
in epithelial cells, PGN or Pam3Cys, ligands for TLR2, were injected into ligated loops containing a single PP in the mid-region
of the small intestine, and PP tissues were harvested and frozen 90
min later. For most experiments, a high dose (1 mg/ml) of TLR2
ligands was used in an attempt to override possible effects of endogenous TLR2 ligands from commensal bacteria, but 0.01– 0.1
mg/ml PGN produced similar effects on TLR2 distribution. AntiTLR2 labeling of frozen tissue sections showed a reduction, but
not loss, of TLR2 on villus epithelial surfaces (Fig. 2a). In the FAE
exposed to PGN or Pam3Cys, UEA⫹ M cells appeared to contain
higher levels of TLR2 intracellularly than in controls, whereas
TLR2 was no longer detected in FAE enterocytes (Fig. 2b).
TLR2⫹ cells were detected in the subepithelial region underneath
the FAE only after exposure to PGN or Pam3Cys (Fig. 2b), and
double-labeling with anti-CD11c Abs indicated that most of these
cells were DCs (data not shown).
It has been proposed that interaction of TLR4 and TLR2 may be
required for TLR2 signaling (44) because PGN-mediated responses in
cultured cells were dependent on expression of both TLR2 and TLR4
(45). To determine whether PGN-mediated redistribution of TLR2 in
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FIGURE 1. TLR2 is expressed in
mouse PP epithelial cells. a and b, Intact,
whole-mount preparation of mouse PP.
a, Staining of the intact tissue with the
lectin WGA and analysis by confocal
microscopy shows the FAE of an individual follicle surrounded by villi. b, AntiTLR2 Abs were applied in vivo and
visualized on intact tissue at lower (top
panels; scale bar, 30 ␮m) and higher
magnification (bottom panels; scale bar,
10 ␮m). Anti-TLR2 (green) is detected
in apical poles of cells in the FAE (left
panels) but not in control FAE exposed
to nonspecific rabbit IgG (right panels).
M cell surfaces are stained with the lectin
UEA-1 (red). The whole-mount preparations shown are representative of three
mice. c, Tissue section of BALB/c
mouse PP stained with anti-TLR2
(green) and UEA-1 (red). TLR2 is
present at apical and basal poles of FAE
cells. Nuclei are stained blue with DAPI
(scale bar, 5 ␮m). d, FAE stained with
anti-TLR2 (green) and WGA (red)
shows that TLR2 is present below the
brush border glycocalyx (scale bar, 5
␮m). e, Negative control section stained
with nonspecific rabbit IgG (green) and
UEA-1 (red) (scale bar, 8 ␮m). f, Tissue
section stained with anti-TLR2 (green)
shows that TLR2 is confined to apical
poles of villus and crypt epithelial cells
(scale bar, 8 ␮m). Pictures shown (c–f)
are representative of staining observed in
BALB/c (n ⫽ 15) and C57BL/6 (n ⫽ 9)
mice. g, TLR2 is not detected in the FAE
of TLR2-deficient mice (scale bar, 8
␮m). The dotted line indicates the basal
side of the FAE. Staining shown is representative of all TLR2-deficient mice
(n ⫽ 11).
4278
TLRs AFFECT FUNCTIONS OF THE FAE
FIGURE 2. Luminal TLR2 ligands induce rapid redistribution of TLR2 in the FAE, but not in villus epithelium. Ligated intestinal loops in BALB/c mice were injected with 1 mg/ml PGN or PBS (control) and harvested
after 90 min. Tissue sections were stained with anti-TLR2
(green), UEA-1 (red), and DAPI (blue). a, In villi, TLR2
on epithelial cells is diminished, but not lost, following
exposure to PGN (scale bar, 10 ␮m). b, Sections of PP
mucosa exposed to PGN or PAM3Cys show loss of TLR2
(green) from FAE enterocytes, but some UEA-1⫹ M cells
(red) remain positive for TLR2. Similar results were observed after lower doses of TLR2 ligands (10 and 100
␮g/ml). TLR2-positive DCs (arrowheads) are present in
tissue exposed to PGN or Pam3Cys. The dotted line indicates the basal side of the FAE. Pictures shown are
representative of staining observed in PBS control (n ⫽
15), PGN-injected (n ⫽ 11), and Pam3Cys-injected (n ⫽
5) mice. Scale bar, 15 ␮m.
shown), but only at the apical poles of villus epithelial cells (Fig.
3). The distribution of TLR2, -5, and -9 in TLR4-deficient mice
was indistinguishable from that of control mice. Furthermore, the
distribution of TLR4, -5, and -9 in FAE or villus epithelium was
not affected by treatment with luminal PGN in either wild-type or
TLR4-deficient mice (data not shown), suggesting that the observed redistribution of TLR2 in response to TLR2 ligands was
due to specific TLR2-ligand interaction.
TLR2 ligands enhance FAE transport of microparticles
We then sought to determine whether TLR2 ligands might have an
effect on FAE functions such as M cell transport. M cells in rabbits
(47) and mice (48) are known to endocytose and transport fluorescent latex microspheres across the FAE. Ligated intestinal loops
of control BALB/c, C57BL/6, and TLR2-deficient mice were
therefore injected with 200-nm fluorescent latex microspheres, and
tissue was harvested, rinsed, and fixed 90 min later. Microspheres
on the surface of the FAE, within the epithelium and in the underlying tissue, were counted in standard areas on sections of PP
follicle domes as described in Materials and Methods. In untreated
or PBS-injected wild-type and TLR2-deficient mice, the mean
numbers of microspheres per standard area were similar (Fig. 4, a
and b). When 1 mg/ml PGN was injected along with microspheres,
however, the number of microspheres within and under the FAE
FIGURE 3. Expression of TLR4, -5, and -9 in the FAE
of mouse PP. Fresh frozen sections of PP from control
BALB/c mice were stained with Abs against TLR4, TLR4/
MD2 complex, TLR5, or TLR9 (red), and counterstained
with anti-CD11c (green) and DAPI (blue). TLR4 is expressed at very low levels at apical poles of both villus and
FAE cells. In contrast, TLR4/MD2 complex is not detected
in FAE cells, but is present in CD11c⫹ DC located in the
SED region. TLR5 is detected at apical surfaces of FAE
cells and in supranuclear structures where it colocalizes
with H. pomatia agglutinin (HPA), a Golgi marker (inset).
TLR9 is expressed at apical and basolateral poles of FAE
cells, and only at apical poles of villus epithelial cells. V,
villi. The dotted line indicates the basal side of the FAE.
Pictures shown are representative of staining observed in
BALB/c mice (n ⫽ 9). (Scale bar, 20 ␮m.)
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the FAE in vivo was dependent on the presence of TLR4, PGN was
injected into ligated intestinal loops in C57BL/10ScNCr TLR4deficient and C57BL/10 control mice. PGN induced the same redistribution of TLR2 in FAE of both control C57BL/10 (TLR4⫹) mice
and TLR4-deficient mice (data not shown), as observed in BALB/c
mice (Fig. 2b), indicating that the response was not TLR4 dependent.
The distribution of TLR4, -5, and -9 in PP of normal BALB/c
and C57BL/6 mice was then examined by immunostaining of frozen tissue sections. In both mouse strains, Abs recognizing the N
terminus of TLR4 revealed low or undetectable levels of TLR4 at
apical surfaces of FAE and villus epithelial cells (Fig. 3), and
higher levels at apical surfaces of crypt cells (data not shown) as
reported previously (26). Labeling was TLR4 specific, because the
Abs failed to label tissues from TLR4-deficient mice (data not
shown). Abs specific for the TLR4/MD2 signaling complex labeled subepithelial CD11c⫹ DCs, but not the FAE (Fig. 3). Although mice deficient in TLR5 and TLR9 were not available to
verify the specificity of the Abs used, anti-TLR5 labeled apical
poles and supranuclear structures in FAE as well as villus (Fig. 3)
and crypt epithelial cells (data not shown). The supranuclear structures were also labeled with the lectin Helix pomatia agglutinin
(Fig. 3, inset), a marker for nascent complex carbohydrates in the
Golgi complex (46). TLR9 was detected at both apical and basal
poles of the FAE (Fig. 3) and crypt epithelial cells (data not
The Journal of Immunology
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FIGURE 4. PGN enhances uptake of fluorescent latex microspheres by the FAE in a TLR2-dependent manner. a, Ligated intestinal loops were injected
with 200-nm fluorescent latex microspheres (beads) and either 1 mg/ml PGN or PBS (control) in wild-type mice (above) and TLR2-deficient mice (below).
PGN increased uptake of fluorescent latex microspheres across the FAE in wild-type but not in TLR2-deficient mice. b and c, Relative number of fluorescent
latex microspheres associated with the FAE and underlying tissue. Bars represent the mean number of microspheres per standard FAE sample ⫾ SD. A
standard FAE sample was obtained from each of four mice in all groups (n ⫽ 4), except control C57BL/10 mice (n ⫽ 2). Each standard FAE sample
represents the total number of microspheres counted in standard areas of FAE sections from four separate follicles in a single PP as described in Materials
and Methods. b, PGN- (1 mg/ml) enhanced microsphere uptake in wild-type C57BL/6 (TLR2 wild-type; wt), wild-type C57BL/10 (TLR4 wt), and
TLR4-deficient C57BL/10ScNCr (TLR4⫺/⫺) mice, but not in TLR2-deficient C57BL/6 (TLR2⫺/⫺) mice. Significance between PGN and respective PBS
control are indicated (ⴱⴱⴱ, p ⬍ 0.001; ⴱⴱ, p ⬍ 0.01). c, Increased uptake of microspheres induced by PGN was significant at 45 and 90 min and was dose
dependent. d, A section of FAE from a BALB/c mouse exposed to PGN (100 ␮g/ml) and fluorescent latex microspheres (green) stained with UEA-1 (red),
a marker for M cells, confirms that uptake of microspheres was mediated by M cells. The picture shown is representative of staining observed in all
PGN-exposed tissues of BALB/c mice (n ⫽ 15).
was significantly enhanced in wild-type BALB/c mice (data not
shown), wild-type C57BL/6 mice, wild-type C57BL/10 mice, and
TLR4-deficient mice, but not in TLR2-deficient mice (Fig. 4b).
The effect of PGN on microsphere uptake could be detected after
45 min, and the effect was dose dependent (Fig. 4c). UEA-1 la-
beling of tissue sections from BALB/c mice given injections with
100 ␮g/ml PGN showed colocalization of microspheres with
UEA-1⫹ M cells, as expected (Fig. 4d). These results suggest that
luminal PGN modulates the transport functions of M cells in the
FAE through a TLR2-mediated mechanism.
4280
TLR2 ligands induce migration of subepithelial DCs into the
FAE
Another important function of the FAE is to attract DCs to the
SED region, presumably to capture transported Ags (39 – 41, 49).
To test the possibility that PGN affected the movements of PP
DCs, the distribution of DCs in PP exposed to PGN for 90 min was
compared with that of PBS-injected controls by labeling frozen
tissue sections with anti-CD11c Abs. To visualize the boundary
between the epithelium and connective tissue, basement membranes were stained with anti-laminin Abs. In normal BALB/c
mice and PBS-injected controls, many CD11c⫹ DCs were present
in the SED region under the FAE, and a few DCs were detected
within the FAE (Fig. 5a) as reported by others (39, 49). In PP
TLRs AFFECT FUNCTIONS OF THE FAE
exposed to luminal PGN at concentrations of 0.01, 0.1, or 1 mg/ml,
a significant increase in the numbers of CD11c⫹ DCs in the FAE
was observed in BALB/c (Fig. 5a), wild-type C57BL/6, and
TLR4-deficient mice, but not in TLR2-deficient mice (Fig. 5c). In
contrast, PGN had no effect on the number of DCs in the villus
epithelium of any mouse strain (Fig. 5, b and c). Similar results
were observed after injection of Pam3Cys (data not shown). Finally, intraluminal injection of GM6001 (5 ␮g/ml), a broad-range
inhibitor of MMPs, significantly inhibited PGN-induced migration
of DC into the FAE (Fig. 5d). However, GM6001 treatment did not
prevent the redistribution of TLR2 in the FAE or the enhanced
uptake of microspheres mediated by PGN (data not shown). Taken
together, these results suggest that DCs in the SED region of PP
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FIGURE 5. PGN induces rapid migration of DCs into the FAE, but not into villus epithelium. Tissue sections of BALB/c mouse PP from ligated
intestinal loops injected with 1 mg/ml PGN or PBS (control) were stained for CD11c (green), laminin (red), and nuclei (blue). a and b, Arrows indicate
DCs in the epithelium after 90 min of PGN exposure. Similar results were obtained using lower doses of PGN (10 –100 ␮g/ml). PGN induced migration
of DCs into the FAE (a) but not into villus epithelium (b). Pictures shown (a and b) are representative of staining observed in BALB/c mice (PBS control,
n ⫽ 15; PGN-treated, n ⫽ 11). c and d, Quantitation of CD11c⫹ DCs in FAE or villus epithelium, as described in Materials and Methods. Values shown
are expressed as the mean number of DCs per standard FAE sample ⫾ SD (n ⫽ 4 mice). Each standard FAE sample represents the total number of DCs
counted in four standard lengths of FAE from four separate follicles in a single PP from one mouse. c, PGN (1 mg/ml) significantly increased DCs in the
FAE of both wild-type strains (wt) and TLR4-deficient (⫺/⫺) mice, but not in TLR2-deficient (⫺/⫺) mice. ⴱ, p ⬍ 0.05; ⴱⴱ, p ⬍ 0.01; and ⴱⴱⴱ, p ⬍ 0.001,
PGN treated compared with respective PBS controls. d, A broad-range MMP inhibitor, GM6001, prevented the PGN-induced migration of DCs into the
FAE of BALB/c mice. ⴱⴱⴱ, p ⬍ 0.001, MMP inhibitor/PGN treated compared with PGN alone.
The Journal of Immunology
respond to TLR2-mediated signals, and that TLR2-mediated DC
migration into the FAE is dependent on the activation of MMPs.
LPS, a TLR4 ligand, also modulates gatekeeping functions of
the FAE
To determine whether other TLR ligands can affect M cell transport
and/or DC migration in the FAE, LPS (100 ␮g/ml) was injected into
ligated loops of BALB/c mice, and the effects on microparticle uptake
and DC migration were evaluated after 90 min. LPS significantly
enhanced the uptake of microparticles by the FAE (Fig. 6a) and also
induced migration of DCs into the FAE (Fig. 6b). These results
suggest that TLRs other than TLR2 can modulate FAE functions.
Discussion
FIGURE 6. LPS modulates gatekeeping functions of the FAE. a, Ultrapure E. coli LPS (100 ␮g/ml) injected into ligated intestinal loops for 90
min enhanced the rate of transepithelial transport of microparticles by the
FAE. The numbers of fluorescent latex microspheres associated with the
FAE and underlying tissue in BALB/c mice (n ⫽ 4) were counted as
described in Materials and Methods. b, LPS also induced DC migration
into the FAE of BALB/c mice (n ⫽ 4). For both graphs, values are expressed as the mean of number of beads or DCs per standard FAE sample ⫾ SD. Each standard FAE sample included the total number of beads
or DCs counted in standard areas of FAE in sections from four separate
follicles in a single PP. ⴱ, p ⬍ 0.05 compared with PBS control.
that TLR2, -4, -5 and -9 are also present in the FAE of mouse PP,
and that TLR4 and -5 show the same distribution in FAE cells as
in villus epithelial cells. In contrast, TLR2 and TLR9 were observed at both apical and basal poles of cells in the FAE, but at
apical poles only in villus and crypt epithelial cells. Apical TLR2,
-5, and -9 were located just below the lectin-labeled glycocalyx,
suggesting that these receptors may be present in microvillus
membranes, apical endosomes, or both. Apical membrane components are known to recycle between microvillus membrane and
apical endosomes (50), which is consistent with the availability of
TLRs at intestinal cell surfaces as previously reported for TLR2
and TLR4 (8). The presence of basolateral TLR2 and TLR9 on
FAE but not on villus epithelium may reflect the contrasting functions of these distinct epithelia. The villus epithelium is designed
to prevent the entry of Ags and microorganisms, and apical TLRs
on villus enterocytes may participate in the “sensing” of luminal
microorganisms (1) that is essential for maintaining epithelial homeostasis (28). In contrast, the apical and basal distribution of
TLR2 and TLR9 on the FAE may reflect the presence of microorganisms and their components on both sides of the epithelial
barrier as a result of constitutive transepithelial transport by M
cells. For the recognition of PGN in particular, other receptors
such as peptidoglycan recognition protein (PGRP)-L and PGRP-S,
which are exclusively expressed on the FAE cells, may also be
involved (51).
TLR2 ligands injected into the intestinal lumen induced a dramatic loss of immunoreactive apical and basolateral TLR2 in FAE
enterocytes, whereas apical TLR2 in the villus epithelium was decreased but not lost. Others have reported that low levels of TLRs
are correlated with unresponsiveness to TLR ligands, including
PGN, and with the relatively high levels of inhibitory Tollip
mRNA recovered by laser capture microdissection from human
villus enterocytes (6). The rapid loss of TLR2 from FAE enterocytes could reflect endocytosis that may be required for TLR-mediated signaling (8, 52, 53) and for degradation of receptors resulting in the termination of TLR-mediated responses (54, 55).
Further studies following the fate of TLR2 in PP early after exposure to TLR2 ligands will be required to elucidate the fate of
TLR2-ligand complexes. Nevertheless, the observed loss of TLR2
suggests that FAE enterocytes play a role in TLR2-mediated responses of the FAE.
In contrast to FAE enterocytes, M cells in PP remained strongly
TLR2-positive after exposure to high doses of PGN. However, the
distribution of TLR2 in these cells appeared to shift from predominantly apical, to apical, intracellular, and basolateral, consistent with
possible entry of TLR2-ligand complexes into the well-documented
transepithelial vesicular transport pathway of these cells (31). The
persistence of TLR2 in M cells is consistent with the fact that
materials taken up by these cells are generally not directed to lysosomes and are not degraded; rather, M cell endocytic vesicles deliver
their content directly to the basolateral cell surface (31). The fact that
immunoreactive TLR2 was readily detected intracellularly in PGNexposed M cells suggests that PGN did not interfere with Ab binding
sites on TLR2, and that the loss of TLR2 observed in FAE enterocytes
was not due simply to blocking of Ab binding sites by PGN. The
uptake of TLR2 by M cells suggests that this receptor may carry
endocytosed ligand and mediate the delivery of microbial components
such as PGN, or intact Gram-positive bacteria displaying TLR2
ligands, across the epithelium into the PP. Such a transport function
has been suggested for the alternate PGN receptor PGRP-S expressed
on UEA⫹ M cells in mice (51).
TLR2 ligation also had a rapid effect on M cell endocytic and
transcytotic activity, significantly increasing the uptake and transport of latex microspheres. It is important to note that enhanced
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The FAE associated with organized mucosal lymphoid tissues represents only a small fraction of the intestinal surface area, but it
plays a crucial role in the initiation of mucosal immune responses
against foreign Ags and pathogens. M cells in the FAE transport
samples of the luminal content across the epithelial barrier (31)
and FAE enterocytes release chemokines that attract Ag-presenting DCs to the subepithelial region (39 – 41), where these samples
can be efficiently captured. We have now shown that the distribution of TLR2 in epithelial cells of the FAE, including M cells,
differs from that of villus and crypt epithelial cells. Injection of
TLR2 ligands into the intestinal lumen induced responses in the
FAE that were not observed in adjacent villi. These included TLR2
redistribution, enhanced uptake of microparticles by M cells, and
induced migration of DCs into the epithelium.
In the intestinal villi and crypts of the normal BALB/c and
C57BL/6 mice used in this study, the TLRs detected by immunocytochemistry (TLR2, -4, -5, and -9) were located at the apical
poles of epithelial cells. This is consistent with previous reports of
the polarized apical distribution of TLR3 (10), TLR4/MD2 (26),
and TLR5 (3) in normal mouse intestinal epithelium. We observed
4281
4282
Epithelial-DC interactions at mucosal surfaces are thought to
provide a crucial link between innate and adaptive mucosal immunity (64 – 66). The TLR2-mediated effects on M cell transport
and DC migration that we observed in vivo would be expected to
result in enhanced Ag capture in organized mucosal lymphoid tissues, but not in the general mucosa. The TLR2-induced migration
of DCs into the FAE but not into villus epithelium that we observed is consistent with evidence that DCs associated with the
FAE have a distinct phenotype from those of the lamina propria
(39), and with other evidence that subepithelial DC functions are
influenced by epithelial signals (39 – 41, 48). Crosstalk between
villus epithelial cells and DCs may promote intestinal immune
homeostasis (65), whereas crosstalk between FAE cells and DCs
may influence the nature of immune responses initiated in mucosal-organized lymphoid tissues (67). Our results support the hypothesis that a unique crosstalk between epithelial cells of the FAE
and DCs occurs in mucosal-organized lymphoid tissues, and that
TLRs participate in this important interaction.
Acknowledgments
We thank Dr. Douglas Golenbock (University of Massachusetts, Worcester, MA) for providing TLR2-deficient mice generated by Dr. Shizuo Akira
(Osaka University, Osaka, Japan). Thanks also to Drs. Wee Yong (University of Calgary, Alberta, Canada) and Egil Lien (University of Massachusetts, Worcester, MA) for helpful advice and discussion.
Disclosures
The authors have no financial conflict of interest.
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large congregation of DCs that normally occupies the SED region
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