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*
Department of Molecular Pathology and Medicine, Human Immunology Unit, Scientific Institute San Raffaele-DIBIT, Milan, Italy;
Roche Milano Ricerche, Milan, Italy;
Signal Transduction Laboratory, Imperial Cancer Research Fund, London, United Kingdom; and
§
Ateneo Vita-Salute School of Medicine, Milan, Italy
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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i/q
protein-coupled receptor in the process) (11) and that
mice carrying targeted deletions in nonredundant chemokine or
chemokine-receptor genes display leukocytosis and impaired leukocyte
transmigration into sites of inflammation (12, 13, 14, 15)
. A large number of chemokines are capable of up-regulating integrin expression and function in leukocytes belonging to various lineages (5, 6, 7, 8, 9, 10). This functional up-regulation is typically transient and dose-dependent, in some cases showing a bimodal pattern that results in the inhibition of basal integrin function at high nanomolar concentration of the chemokine (6), as expected for a stimulus that is associated with cell movement rather than with static adhesion. Further, chemokines such as monocyte chemotactic protein-1 (MCP-1),5 RANTES, and IL-8 induce preferential activation of selected integrin heterodimers within a given leukocyte lineage subpopulation (8, 10, 16), indicating that integrin subfamilies may have a differential sensitivity and/or may require alternative activation pathways even within the same cell population.
While differences in chemokine-induced adhesion can be accounted for by
variable chemokine receptor expression or lineage-specific behaviors of
integrins, the molecular steps involved in the process downstream of
chemokine receptor occupancy are less well defined. This issue is
particularly relevant for sublineages of T lymphocytes, as they express
a largely overlapping profile of chemokine receptors and have been
shown to possess unique recirculation patterns, in some cases
associated with pathological conditions characterized by acute or
chronic inflammatory responses. Th1 and Th2 cells are functionally
distinct subsets of CD4+ T lymphocytes
orchestrating polarized versions of the immune response.
IFN-
-secreting Th1 cells regulate phagocyte-dependent immune
responses, whereas Th2 cells, producing IL-4 and IL-5, promote IgE
production and eosinophil function (17). Despite their
common lineage origin, Th1 and Th2 cells show clearly distinct homing
behaviors (for a review see Ref. 18). While Th1 cells
migrate and/or reside preferentially into sites of chronic
inflammation, characterized by extensive fibrosis and thickening of the
basement membrane, Th2 cells show preferential migration into sites of
acute inflammation, such as allergic responses, typically associated
with vascular changes such as vasodilation and increased capillary
permeability. Tissue-specific homing of Th1 and Th2 cells to sites of
inflammation is currently thought to be regulated by differential
expression of P- and E-selectin ligands (19) and selected
chemokine receptors in the two lymphocyte subsets
(20, 21, 22, 23, 24, 25, 26). However, given the complexity of the
extravasation process and the multiplicity of the molecular steps
involved, it is likely that additional features confer selected
recirculation patterns to Th1 and Th2 lymphocytes.
In this study, we investigated the expression and function of integrins in the two sublineages of Th lymphocytes. We found that ß1 integrin-mediated adhesion to extracellular matrix (ECM) proteins in response to chemokines is markedly reduced in Th2 cells, suggesting a developmentally regulated uncoupling of chemokine receptor-generated signals leading to ß1 integrin activation. The molecular step(s) responsible for defective Th2 adhesion in response to chemokines appears to lie downstream of phosphoinositide-3 kinase (PI-3 kinase) activation and actin polymerization or in a complementary signaling pathway required for ß1 integrin up-regulation.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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The human recombinant chemokines RANTES and stromal cell-derived
factor-1 (SDF-1) were obtained from R&D Systems (Minneapolis, MN). PMA,
cytochalasin D (CCD), wortmannin (WM), LY294002 (LY), and
bisindolylmaleimide (BIM) were purchased from Calbiochem (La Jolla,
CA). Monoclonal Ab TS1.22 (anti-CD11a) was kindly provided by
T. A. Springer (Harvard Medical School, Boston, MA); mAb
anti-integrin 3 chain (VLA-3),
anti-4 chain (VLA-4), and anti-6
chain (VLA-6) were purchased from AMAC (Westbrook, ME). FITC-conjugated
goat anti-mouse IgG was obtained from Zymed Laboratories (South San
Francisco, CA). Recombinant human ICAM-1, fused to the amino terminal
region of the Ig-binding subunit of Staphylococcus
aureus protein A (zz-ICAM-1) was generously provided by A.
Randi (Glaxo-Wellcome, Stevenage, U.K.). Fura-2 AM, fibronectin,
FITC-conjugated phalloidin, and BSA were obtained from Sigma (St.
Louis, MO). Laminin-1 was purchased from Life Technologies (Grand
Island, NY). The neutralizing anti-cytokine mAbs were purchased
from PharMingen (San Diego, CA). IL-12 was obtained from
Hoffmann-LaRoche (Nutley, NJ). IFN- was kindly provided by
Hoffman-LaRoche AG (Basel, Switzerland). Fluo-3AM was purchased from
Molecular Probes (Eugene, OR). Anti-human Vav and
anti-phosphotyrosine (4G10) mAbs were purchased from Upstate
Biotechnology (Lake Placid, NY). Anti-phosphotyrosine (PY99) mAb was
purchased from Santa Cruz Biotechnology (Santa Cruz, CA).
Generation of polarized human Th lymphocytes
Human neonatal leukocytes were isolated from freshly collected, heparinized, neonatal blood by Ficoll-Hypaque density gradient centrifugation. Polarized Th cell lines were generated by stimulation with 2 µg/ml PHA in the presence of various combinations of cytokines and neutralizing anti-cytokine Abs, as previously described (26). Briefly, Th1 cells were generated by addition of 5 ng/ml IL-12 and 200 ng/ml neutralizing anti-IL-4 Ab. Th2 cells were generated by addition of 10 ng/ml IL-4 and 2 µg/ml neutralizing anti-IL-12 Abs 17F7 and 20C2. Cells were cultured in complete RPMI 1640 medium supplemented with 5% FBS, 2 mM L-glutamine, 1 mM sodium pyruvate, 100 U/ml penicillin-streptomycin. On day 3, the cultures were washed and expanded in complete medium with the addition of 100 U/ml IL-2.
Intracellular staining for cytokines
After 10 days of culture, Th1 and Th2 cells were washed,
collected, and restimulated with 50 ng/ml PMA and 1 µg/ml ionomycin
for 4 h. Then, 10 µg/ml of brefeldin A were added for the last
2 h of culture. Cells were then fixed with 4% paraformaldehyde,
permeabilized with saponin, and stained with FITC-labeled
anti-IFN- and PE-labeled anti-IL-4 Abs (PharMingen). Samples
were analyzed by flow cytometry with a FACScan (Becton Dickinson,
Mountain View, CA).
Evaluation of integrin expression by immunofluorescence
Phenotypic analysis was performed by staining 1 x
106 cells for 30 min on ice with saturating
concentrations of mAbs specific for different surface markers in PBS
containing 1% FBS. Cells were then washed and incubated for 30 min on
ice with 10 µg/ml of FITC-conjugated goat anti-mouse IgG. After
washing, cells were resuspended in PBS and analyzed in a FACScan flow
cytometer (Becton Dickinson, Mountain View, CA).
6 integrin antigenic sites were determined
after calibration of the flow cytometer by using fluorescein-labeled
microspheres (Flow Cytometry Standards, Research Triangle Park, NC; No.
824) as previously described (27).
Analysis of intracellular calcium mobilization
Th1 and Th2 cells were washed twice in Krebs-Ringer-HEPES (KRH) medium, counted, and resuspended at 5 x 106 cells/ml. The fluorescent Ca2+ indicator acetoxymethyl-fura 2 (2 µM) was added to the cells, followed by a 60-min incubation at 37°C on a shaking waterbath. Fluorescence measurements were conducted in a Perkin-Elmer LS50 B spectrofluorometer (Norwalk, CT) under continuous stirring. Calibration of the fluorescent signal was conducted as described previously in detail (27).
Actin polymerization assay
F-actin was quantitated by staining with FITC-conjugated phalloidin. Briefly, 1 x 106 cells were resuspended in RPMI 1640 medium with 10% FBS and pretreated at 37°C for 30 min with or without the following drugs: 10 µM CCD, 200 nM WM, 50 µM LY, or 2 µM BIM. Cells were stimulated with chemokines (100 ng/ml) or PMA (100 nM) at 37°C for the indicated time points. Incubation was stopped by adding three volumes of 3.7% paraformaldehyde at room temperature (RT) for 10 min, followed by washing with PBS and permeabilization on ice for 2 min with 0.1% Triton-HEPES. Thereafter, cells were stained with FITC-phalloidin (2 µg/ml) at RT for 30 min, washed with PBS, and analyzed by flow cytometry with a FACScan.
Cell adhesion assay
Ninety-six-well ELISA plates (Linbro/Titertek, Huntsville, AL)
were coated overnight at 4°C with 10 ng/ml of laminin 1, fibronectin,
or BSA in PBS, followed by blockade with 2% BSA at 37°C for 1
h. Recombinant zz-ICAM-1 was adsorbed at 2 µg/ml onto plates
precoated for 2 h at RT with 20 mg/ml human IgG (Sandoglobulin,
Sandoz S.A., Basel, Switzerland) in PBS. Because the "zz" tag binds
to Igs, the precoating with human IgGs allows a correct orientation of
bound ICAM-1. Thereafter, plates were washed with PBS blocked with 2%
BSA, and 1 x 105 cells were added to
triplicate wells with or without chemoattractants (100 ng/ml) or PMA
(100 nM). For competitive inhibition experiments, cells were
preincubated with serial dilutions of function-blocking
anti-6 Ab for 20 min on ice and washed
before addition to ligand-coated plates. In some experiments, cells
were pretreated at 37°C for 30 min with or without the following
drugs: WM (200 nM), LY (50 µM), or BIM (2 µM). Plates were
centrifuged for 2 min at 800 rpm to allow rapid cell sedimentation and
were incubated for the indicated time periods at 37°C. Nonadherent
cells were removed by washing four times with medium; adherent cells
were fixed with 3.7% paraformaldehyde at RT for 10 min, washed with
PBS, and stained with 0.5% crystal violet at RT for 10 min. Plates
were washed four times with water, crystal violet was then extracted by
the addition of 1% SDS, and adsorbance at 540 nm was measured in an
ELISA reader. After subtraction of the background binding, as assessed
with BSA-coated control wells, specific binding was calculated as
percentage of total input cells.
Immunoprecipitation and Western blot
Cells (1.5 x 107 cells/sample) were either left untreated or stimulated for 2 min at 37°C with chemokines (RANTES, 200 ng/ml; or SDF-1, 200 ng/ml) or with 50 µg/ml of anti-CD3 (TR66), mAb plus goat anti-mouse IgG Ab. Cells were then lysed in 2x Lysis buffer (100 mM Tris, pH 8, 300 mM NaCl, 2 mM EDTA, 2 mM EGTA, pH 8, 2% Nonidet P-40, 2 nM PMSF, 20 mM NaF, 2 mM Na3VO4, 2 µg/ml aeupeptin, 2 µg/ml apoprotinin) for 10 min on ice. For immunoprecipitation, protein G-Sepharose was preincubated with anti-human Vav Ab, washed twice in PBS, incubated with lysates at 4°C overnight, and eluted in reducing sample buffer. Samples were analyzed by SDS-PAGE and transferred onto nitrocellulose. The membrane was blocked in 5% milk-PBS, incubated with 4 µg/ml of anti-human Vav or with a combination of two anti-phosphotyrosine Abs (Py99, 0.2 µg/ml; and 4G10, 1 µg/ml) followed by an incubation with an appropriate HRP-conjugated Ab. Peroxidase reaction was developed using enhanced chemiluminescence (Amersham, Arlington Heights, IL).
GST-p21-activated kinase (PAK) pull-down assay
The sequence encoding the amino-terminal 252 aa of rat PAKa was amplified using the primers GCT CTA GAA ATG TCA AAT AAC GGC TTA GAC and CTC TAA GCT TGA TCT CCT CAT CAG ACA TTT TAC. The PCR fragment was digested with XbaI and HindIII and cloned into pGEX KG. pXJ40HA wild-type PAKa was a kind gift from L. Lim (UCLA, Los Angeles, CA). To prepare the purified GST-PAK fusion protein, one Escherichia coli colony was inoculated and grown for 16 h at 37°C under agitation in 2x YTA (16 g/L tryptone, 10 g/L yeast extract, 5 g/L NaCl, 100 µg/L ampicillin). The culture was diluted 1:100 in fresh, prewarmed 2x YTA and grown by monitoring OD at 590 nm. Then, 100 mM isopropyl-ß-D-thiogalactoside was added to a final concentration of 0.1 mM, and the culture was grown for additional 2 h. The induced cells were then centrifuged for 10 min at 4°C at 7700 x g and resuspended in 50 µl ice-cold PBS per ml of culture. Cells were disrupted by sonication. Triton X-100 was added to a final concentration of 1%, and samples were incubated for 30 min at 4°C. Cells were centrifuged for 10 min at 4°C and 12,000 x g, and the supernatant was saved and stored in aliquots at -80°C. To purify the expressed protein, to 1 ml of the sonicate 40 µl of in PBS equilibrated glutathione Sepharose (4B; Pharmacia, Upsala, Sweden) was added, and samples were incubated with gentle agitation for 30 min at RT. The suspension was centrifuged at 500 x g for 3 min, and the pellet washed three times with PBS. For pull-down assays, 25 x 106 cells in 0.5 ml PBS were stimulated with 100 nM chemokines or 10 µg/ml anti-CD3 mAb for 30 s at 37°C. Then, 0.5 ml ice-cold 2x lysis buffer (100 mM Tris, pH 7.5, 150 mM NaCl, 2 mM sodium orthovanadate, 100 mM NaF, 2 mM PMSF, 20 mM MgCl2, 2% Triton X-100, 1 mM DTT, 20 µg/ml leupeptin and aprotinin) was added, and lysates were vortexed and were immediately cleared for 1 min at 13,000 rpm. The cleared lysates were added to 40 µl GST-PAK beads and incubated for 5 min at 4°C under agitation. The GST-PAK beads were then quickly washed twice with wash buffer (50 mM Tris, pH 7.5, 500 mM NaCl, 10 mM MgCl2, 0.1% Triton X-100, 2 mM DTT), and bound proteins were immediately eluted with sample buffer (2% SDS, 0.005% bromophenol blue, 10% glycerol, 50 mM Tris, pH 6.8). A standard SDS-PAGE was performed, and proteins were transferred to a nitrocellulose membrane (Amersham). GST-PAK-bound Rac1 was identified by immunoblotting with an anti-Rac1 mAb (Transduction Laboratories, Lexington, KY).
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Following their expansion in the presence of polarizing
chemokines, Th1 and Th2 cells were defined phenotypically by their
reciprocal expression of IFN- and IL-4, respectively (Fig. 1
). A cytofluorometric analysis of
surface-expressed integrins in polarized Th cell subsets revealed that
the overall levels of ß2 integrins are higher
in Th1 cells, while surface ß1 integrins appear
to be comparatively similar in the two subsets, as judged by staining
with Abs recognizing the common ß subunits of the various integrin
subfamilies. Conversely, the composition of ß1
integrin heterodimers expressed at the cell surface is qualitatively
heterogeneous in the two subsets: the
4ß1 integrin, a
receptor for fibronectin and VCAM-1, is more highly expressed on Th2
cells, while the 6ß1
integrin, the major laminin receptor, displays consistently higher
expression levels in Th1 cells (Fig. 2).
The surface density of the 6 subunit in
particular was found to range between 20,000 and 25,000 sites/cell and
between 45,000 and 55,000 sites/cells in Th2 and Th1 lymphocytes,
respectively. The observed differences in surface expression of
6ß1 in the two subsets
are consistent with the finding that the steady-state levels of the
mRNA coding for the 6 subunit are markedly
up-regulated by IL-12 treatment in Th1 cells (28). The
3ß1 and
5ß1 receptors were
found to be equally expressed in the two subsets, while the
ß4 integrin subunit, which in selected cell
types dimerizes with the 6 subunit to
yield a receptor for laminin V, was not expressed in either cell
subpopulation.
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To assess the functional properties of integrins expressed by Th1
and Th2 cells, we performed adhesion assays on immobilized integrin
ligands under static conditions. Fig. 3A shows that Th1, but not
Th2, lymphocytes display chemokine-inducible adhesion to fibronectin
and laminin I in response to both RANTES and SDF-1. Such inducible
adhesion reaches a plateau at 15 min poststimulation and is typically
transient, returning to prestimulation levels within 45–60 min (Fig. 3B). The defective response observed in Th2 cells can be
bypassed using phorbol ester stimulation, which induces a long-lasting
up-regulation of integrin avidity, or by performing the adhesion assay
in the presence of 1 mM Mn2+, a condition known
to promote an allosteric transition in integrins’ extracellular
domains resulting in a persistent increase in ligand affinity
(29). Competitive inhibition experiments using
function-blocking Abs specific for the various integrin subunits
confirmed that the adhesion of both cell subsets to fibronectin is
cooperatively mediated by the
4ß1 and
5ß1 heterodimers (not
shown), while adhesion to laminin is entirely dependent on the
6ß1 integrin (see
below). Experiments performed to assess ß2
integrin function in the two subsets, using immobilized ICAM-1 as a
ligand, showed a high baseline adhesion by both subsets, which could be
only marginally up-regulated by chemokine or phorbol ester treatment
(not shown). These findings demonstrate that intrinsic structural or
functional alterations of integrins are unlikely to account for the
reduced adhesion observed in Th2 cells in response to chemokines and
suggest that a molecular step involved in the signaling pathway linking
chemokine receptor stimulation to functional up-regulation of
ß1 integrins is defective in the latter cell
population.
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6ß1 receptor
function in Th1 and Th2 cells
To assess whether the observed difference in
6ß1 surface expression
levels is responsible for the variable adhesion efficiency of the two
lymphocyte subsets, we performed competitive inhibition experiments
using serial dilutions of a function-blocking
anti-6 mAb. Fig. 4A shows a titration curve
generated to determine the saturating concentration of the
anti-6 mAb in the two subsets. At 0.1
µg/ml of Ab, virtually all surface-expressed
6ß1 integrin is
saturated in Th2 cells, while 
50% of the
6ß1 receptors
expressed on Th1 cells are still unsaturated at this mAb concentration.
Based on the estimated receptor density reported above for the two cell
subsets, the number of
6ß1 sites available
for laminin binding in Th1 cells, in the presence of 0.1 µg/ml
anti-6 Ab, should approximate the total
number of receptors expressed by untreated Th2 cells. The experiment
shown in Fig. 4B demonstrates that in the presence of 0.1
µg/ml anti-6 mAb, the adhesion
efficiency of SDF-1-stimulated Th1 cells is still significantly higher
compared with Th2 cells in the absence of Ab (24 ± 2% vs 7
± 3%, respectively). Thus, the defective adhesion to laminin
displayed by chemokine-stimulated Th2 cells cannot be explained by a
lower surface expression of the related
6ß1 integrin receptor.
Of note, chemokine-induced adhesion to laminin of Th1 cells is
completely abolished by as little as 1 µg/ml
anti-6 mAb, further indicating that this
is the only functional laminin receptor expressed by this cell
subset.
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A simple explanation for the observed differences in the adhesive
response of Th1 and Th2 cells following chemokine stimulation is that
Th2 cells are less responsive to the chemokines under study at the
receptor level due to lower receptor density and/or sensitivity. As one
of the earliest functional responses to chemokines is a rise in free
[Ca2+]i levels resulting
from phospholipase C activation and breakdown of membrane
phosphoinositides, we analyzed such response in the two lymphocyte
subsets using RANTES and SDF-1 at nanomolar concentrations. Fig. 5 shows that indeed Th2 cells display a
less pronounced elevation of free
[Ca2+]i in response to
RANTES when compared with Th1 cells. This is consistent with previous
reports showing that Th1 cells do express higher levels of selected
receptors for RANTES, such as CCR1 (28) and CCR5, and
respond preferentially to RANTES in chemotactic migration assays. In
contrast, the elevation of free [Ca2+]i induced by
SDF-1 is virtually identical in the two subsets, despite their
differential response to this chemokine in the static adhesion assays
reported in Fig. 3
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To investigate the mechanisms underlying the observed variability
in chemokine-induced ß1 integrin functional
up-regulation, we performed chemokine- or phorbol ester-driven adhesion
assays in Th1 cells in the presence of selected enzymatic inhibitors.
Interestingly, chemokine-induced adhesion to ß1
integrin ligands is markedly inhibited in Th1 cells by both PI-3 kinase
and protein kinase C (PKC) inhibitors (Fig. 6), suggesting that parallel signaling
pathways, involving PI-3 kinase and PKC activation, are triggered by
the chemokines under study and are both required for the regulated
adhesion of Th1 cells to ß1 integrin ligands.
Persistent activation of PKC by the irreversible agonist PMA is
sufficient per se, independently of PI-3 kinase activation, in driving
ß1 integrin functional up-regulation, as judged
by its resistance to PI-3 kinase inhibitors and by its complete
inhibition with the selective PKC inhibitor BIM.
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The results described above indicate that the molecular step(s)
responsible for the reduced adhesive response to chemokines of Th2
cells lies downstream of receptor occupancy, phospholipase C
activation, and second messenger generation, at least if one considers
the dissociated functional response of Th2 lymphocytes to SDF-1 by
comparing the induction of free
[Ca2+]i elevations vs
ß1 integrin-mediated adhesion. Moreover, our
findings show that two major enzymatic pathways triggered by
chemokines, namely PI-3 kinase and PKC, are required to up-regulate
ß1 integrin function in Th1 cells. Among the
requirements for inducible integrin-mediated adhesion are an increased
rate of actin polymerization and an extensive reorganization of the
actin-based cytoskeleton, both of which are under the control of small
GTPases of the Rho family and are largely abrogated by CCD pretreatment
(30, 31, 32). Therefore, we set out to investigate whether
chemokines indeed promote net actin polymerization in Th1 and Th2
cells, and the potential mechanisms involved in such response. Fig. 7A shows that following
chemokine stimulation both cell subsets undergo a rapid burst of actin
polymerization, which peaks at 30 s to 1 min and subsides to
baseline levels within 5–10 min poststimulation. Such response is
completely abrogated by pretreatment of the cells with CCD, which binds
irreversibly to free actin monomers and selectively blocks the
ATP-dependent actin polymerization process. Similar to the
[Ca2+]i response reported
above, Th2 cells displayed a delayed and quantitatively reduced actin
polymerization response upon RANTES stimulation, whereas the two cell
subsets showed an identical response when stimulated with the
chemokine SDF-1. Both subsets responded to PMA treatment with a
moderate but persistent increase of actin polymerization. To analyze
the enzymatic pathways involved in such response, we pretreated the two
subsets with selected inhibitors before chemokine or phorbol ester
stimulation. Fig. 7B shows that the two chemokines and PMA
display a reciprocal pattern of inhibition: the burst of actin
polymerization induced by both chemokines is substantially inhibited by
the PI-3 kinase inhibitors WM and LY, but is unaffected by
the selective PCK inhibitor BIM. Conversely, PMA-induced actin
polymerization is resistant to both PI-3 kinase inhibitors but is
completely abrogated by BIM. These findings demonstrate that the PI-3
kinase-dependent burst of actin polymerization induced by chemokines
appears to be equally rapid and efficient in both Th1 and Th2
lymphocytes and parallels the chemokines’ ability to generate second
messengers following receptor occupancy.
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To clarify the signaling mechanisms responsible for the observed
phenomena, we investigated the expression and functional activation of
a number of molecular intermediates reportedly involved in promoting
net actin polymerization and increased integrin-mediated adhesion upon
exposure to chemokines in the two T lymphocyte subsets. As a positive
control in the assay, we used solid phase-bound anti-CD3 Abs, as
most of the signal trasduction pathways analyzed are known to be
triggered by such treatment in T lymphocytes. To indirectly assess the
functional activation of PI-3 kinase, we analyzed the phosphorylation
levels of protein kinase B (PKB)/Akt, a known downstream effector of
PI-3 kinase (33). Fig. 8
shows that a fraction of PKB/Akt undergoes rapid phosphorylation in
both subsets following chemokine stimulation, indicating that PI-3
kinase becomes activated by both chemokines. This is further suggested
by the observation that chemokine-induced PKB/Akt phosphorylation was
completely abrogated by PI-3 kinase inhibitors (not shown). Comparable
levels of PKB/Akt phosphorylation were induced in both subsets by
SDF-1, whereas RANTES consistently induced higher PKB/Akt
phosphorylation levels in Th1 as opposed to Th2 lymphocytes, again
confirming the existence of a higher sensitivity of the former T cell
subset to RANTES. Activation of the Rho-like GTPase Rac1 has been
proposed to link PI-3 kinase activation to actin polymerization,
adhesion, and chemotaxis in several cell types. To assess whether Rac1
undergoes GDP to GTP exchange following chemokine stimulation, we used
an agarose matrix-immobilized GST-PAK fusion protein in pull-down
assays, which specifically identify the GTP-bound fraction of Rac1
(34). Such assays demonstrate that a small but significant
fraction of endogenous Rac1 binds to the effector protein PAK rapidly
after chemokine stimulation. Such binding was observed at comparable
levels in the two subsets and was completely abrogated by pretreatment
of the cells with PI-3 kinase inhibitors (not shown). Among the
exchange factors known to promote the conversion of GDP for GTP in
Rho-like GTPases, Tiam-1 and Vav have been reported to be
preferentially active on Rac1 (35, 36) and to be expressed
in lymphoid cells. Fig. 8 shows that indeed both proteins are expressed
at comparable levels in Th1 and Th2 cells. However, unlike immobilized
anti-CD3 Abs, neither RANTES nor SDF-1 induce detectable
phosphorylation of Vav on tyrosine residues, an event which appears to
be required for the acquisition of full catalytic activity by this
exchange factor (37). These findings indicate that a major
signaling pathway involved in receptor-mediated actin polymerization
and cell motility is activated at comparable levels in both Th1 and Th2
lymphocytes following chemokine stimulation, despite the differential
ability of the two subsets to up-regulate ß1
integrin adhesion in response to chemokines.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Despite their demonstrated role in supporting lymphocyte
transendothelial migration, integrins have not been extensively
analyzed phenotypically or functionally in polarized subsets of Th
cells. As expected for T lymphocytes cultured for extended periods in
the presence of mitogens, we found that both Th1 and Th2 cells express
high surface levels of integrins. However, ß2
integrins appear to be expressed at higher levels in Th1 cells, while
ß1 integrins are expressed at comparable levels
in the two subsets. On a more qualitative basis, Th1 cells express
higher surface levels of the 6 integrin
subunit, which, as previously reported by our group, appears to be a
transcriptional target of the Th1-polarizing cytokine IL-12
(28). In contrast, Th2 cells express higher levels of the
4 integrin subunit. Competitive inhibition
experiments demonstrated that in our experimental model adhesion to
laminin appeared to be exclusively mediated by the
6ß1 receptor, while
adhesion to fibronectin was cooperatively mediated by
4ß1and
5ß1 integrins.
The aim of this work was to evaluate the ability of chemokines to functionally up-regulate ß1 integrins in polarized T helper cells. The most significant finding reported here is that in Th2 but not Th1 lymphocytes, the early response induced by chemokines, in terms of phospholipase C activation and Ca2+ influx, is functionally uncoupled from the activation of integrin-mediated adhesion. The experiments performed using known activators of integrin-dependent adhesion, such as phorbol esters or buffers containing Mn2+ ions, suggest that integrins are indeed competent to engage their specific ligands in Th2 lymphocytes and that one or more molecular intermediates linking chemokine responses to integrin functional up-regulation are defective in these cells when compared with Th1 cells. In search of such intermediates, we explored selected intracellular pathways that are known to be involved in transducing membrane receptor-generated signals required for cytoskeletal reorganization, adhesion, and motility in most lymphoid and nonlymphoid cells. One such pathway involves the activation of PI-3 kinase by G protein-coupled receptors, including chemokine receptors, and the subsequent activation of subfamilies of Rho-like GTPases. Two lines of evidence in this work support the conclusion that PI-3 kinase is efficiently activated by chemokines at comparable levels in Th1 and Th2 cells: first, we observed comparable levels of phosphorylation of PKB/Akt, a known downstream effector of PI-3 kinase, in both Th1 and Th2 cells upon exposure to SDF-1, a chemokine that promotes comparable chemotattractant responses in the two subsets; second, selective inhibitors of PI-3 kinase abolish the burst of actin polymerization induced by SDF-1 in both lymphocyte subsets. Furthermore, using a sensitive biochemical assay, we provide for the first time evidence for the rapid activation of Rac1 by chemokines, which is compatible with the role played by this Rho-like GTPase in supporting cell adhesion and motility. The activation of Rac1 appears to lie downstream of PI-3 kinase activation and is likely to be primarily responsible for the burst of actin polymerization observed in both T cell subsets. Indeed, both the exchange of GDP for GTP in Rac1 and the observed burst in actin polymerization are efficiently blocked by selective PI-3 kinase inhibitors, and dominant negative Rac1 mutants have been shown by others to prevent actin polymerization induced by various agonists (38, 39, 40). Other members of the Rho family, such as Cdc42, have been reported to be involved in chemokine-induced morphologic changes in myeloid cells, suggesting that Rho GTPases are key regulators of chemoattractant responses in multiple cell types (41, 42). Several recent reports have highlighted a potential mechanism linking PI-3 kinase activation to the accelerated excange of GTP for GDP in Rho-like GTPases by showing that Vav, a specific exchange factor for Rac1, binds to and is directly controlled by substrates and products of PI 3-kinase (37). Interestingly, these reports show that the product phosphatidylinositol- 3,4,5-trisphosphate enhances phosphorylation and full catalytic activation of Vav by srk-family kinases such as Lck (37). Our findings demonstrate that Vav and Tiam-1, two major exchange factors for Rac1, are expressed at comparable levels in Th1 and Th2 cells. However, we could not detect any tyrosine-phosphorylated Vav in either T cell subset upon short-term exposure to chemokines, suggesting that Vav is unlikely to be fully activated by chemokine stimulation in T lymphocytes. Collectively, our findings suggest that the reduced activation of ß1 integrin-dependent adhesion observed in Th2 cells exposed to chemokines cannot be explained by a defective activation of PI-3 kinase, Rac-1, and the related processes leading to a burst of actin polymerization, which in several models is required to support integrin-mediated adhesion, spreading, and motility.
A complementary signaling pathway that impacts on chemokine-induced
adhesion involves the activation of the small GTPase RhoA and the
functionally interdependent enzymatic activities of phospholipase D and
PKC (43). Several reports have indicated that RhoA
activity is required for agonist-induced ß1 and
ß2 integrin activation in lymphoid cells
(7, 30). Likewise, a large body of experimental work has
shown that PKC activation plays a central role in integrin function and
integrin-cytoskeletal interactions (44, 45). Our findings
indicate that the selective inhibition of PKC in Th1 cells abolishes
ß1 integrin-dependent adhesion in response to
chemokines and, as expected, to phorbol esters. Interestingly, the
activity of PKC does not seem to be required for chemokine-induced
actin polymerization. This suggests that two independent signaling
pathways, one involving PKC and the other requiring PI-3 kinase
activity, are both required for the observed adhesion to ECM ligand
upon chemokine stimulation and may be differentially regulated in Th1
vs Th2 cells. Although our findings do not provide a mechanistic
explanation for the markedly reduced fucntional activation of
ß1 integrins in Th2 cells exposed to
chemokines, they demonstrate that the observed differences are not due
to a defective activation of a signaling pathway involving PI-3 kinase,
the small GTPase Rac1, and other intracellular regulators of actin
polymerization. Conversely, the possibility exists that the variable
efficiency of chemokine-induced ß1 integrin
function in the two subsets might reside in the differential expression
and functional regulation of selected isoforms of PKC expressed by
lymphoid cells and/or by regulators and effectors of the RhoA GTPase.
In support of this conclusion, recent reports (46) have
demonstrated the involvement of selected PKC isoforms, such as the
atypical PKC-
, in chemokine-induced adhesion to integrin ligands. In
apparent contrast with this hypothesis, we show here that Th2 cells
respond efficiently to PMA in terms of integrin-dependent adhesion.
However, being phorbol esters metabolically stable and devoid of
selectivity as PKC activators, the possibility exists that subtle
differences in the expression and regulation of selected PKC isoforms
or their specific substrates may be responsible for the observed
behavior of ß1 integrins in
chemokine-stimulated Th2 lymphocytes.
In conclusion, our data indicate that in addition to the demonstrated differential expression of chemokine receptors and selectin ligands, Th1 and Th2 cells show different requirements for ß1 integrin functional activation upon exposure to chemokines. If confirmed by appropriate experimental models in vivo, these findings may provide an additional mechanistic clue to the understanding of the markedly different homing behavior of Th1 vs Th2 cells. Reduced ß1 integrin-mediated adhesion in Th2 cells could restrain their ability to invade and/or reside in sites of chronic inflammation, which are characterized by thickening of basement membranes and extensive fibrosis, requiring efficient interaction with organized ECMs. In contrast, such limitations may not affect margination of Th2 cells into areas of acute inflammation, in which a host of soluble mediators, by increasing microvascular permeability, reduce the selectivity of lymphocyte transmigration.
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Acknowledgments |
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Footnotes |
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2 B.C. and D.D. contributed equally to this work.
3 Current address: The Basel Institute for Immunology, CH-4005 Basel, Switzerland.
4 Address correspondence and reprint requests to Dr. Ruggero Pardi, Human Immunology Unit, DIBIT-Scientific Institute San Raffaele, via Olgettina, 58 I-20132 Milano, Italy. E-mail address:
5 Abbreviations used in this paper: MCP-1, monocyte chemotactic protein-1; SDF-1, stromal cell-derived factor-1; PI-3 kinase, phosphoinositide-3 kinase; BIM, bisindolylmaleimide; WM, wortmannin; LY, LY294002; CCD, cytochalasin D; ECM, extracellular matrix; PKC, protein kinase C; PKB, protein kinase B; [Ca2+]i, intracellular Ca2+ concentration; RT, room temperature; KRH, Krebs-Ringer-HEPES; PAK, p21-activated kinase.
Received for publication September 29, 1999. Accepted for publication January 3, 2000.
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) and Th2 (
) cells were
plated onto BSA-, laminin-, or fibronectin-coated wells in the presence
of RANTES, SDF-1 (100 ng/ml), or Mn2+ (1 mM) and incubated
for 30 min at 37°C, followed by removal of nonadherent cells and
quantitation of adhesion as described in Materials and
Methods. Data are expressed as percentage adhesion over the
total input cells and are the mean ± SD of 10 separate
experiments. B, Adhesion was performed as described in
A, but cells were left in culture for the indicated
time-points before removal of nonadherent cells and quantitation of
specific adhesion. Results are the mean ± SD of three independent
experiments.
