Autotaxin Produced by Stromal Cells Promotes LFA-1–Independent and Rho-Dependent Interstitial T Cell Motility in the Lymph Node Paracortex

Tomoya Katakai, Naoyuki Kondo, Yoshihiro Ueda and Tatsuo Kinashi
J Immunol July 15, 2014, 193 (2) 617-626; DOI: https://doi.org/10.4049/jimmunol.1400565

Abstract

T cells exhibit high-speed migration within the paracortical T zone of lymph nodes (LNs) as they scan cognate Ags displayed by dendritic cells in the tissue microenvironment supported by the network of stromal cells. Although intranodal T cell migration is controlled in part by chemokines and LFA-1/ICAM-1, the mechanisms underlying their migratory activity independent of these factors remain to be elucidated. In this study, we show that LN stromal cells constitutively express autotaxin (ATX), an ectoenzyme that is important for the generation of lysophosphatidic acid (LPA). Importantly, CCL21+ stromal cells in the T zone produced and immobilized ATX on their cell surface. Two-photon imaging using LN tissue slices revealed that pharmacological inhibition of ATX or LPA receptors significantly reduced T cell migration, and this was further exacerbated by blockage of Gαi signaling or LFA-1. Therefore, T cell motility mediated by the ATX–LPA axis was independent of Gαi and LFA-1. LPA induced slow intermittent movement of T cells in vitro in a LFA-1–independent manner and enhanced CCL21-induced migration. Moreover, LPA and CCL21 cooperatively augmented RhoA activity in T cells, which was necessary for efficient intranodal T cell migration via the downstream ROCK–myosin II pathway. Taken together, T zone stromal cells control optimal migratory behavior of T cells via multiple signaling cues mediated by chemokines and ATX/LPA.

This article is featured in In This Issue, p.465

Introduction

The priming of naive T cells by Ag-bearing dendritic cells (DCs) in the lymph node (LN) paracortex (T zone) is central for the initiation of adaptive immune responses (1). T cells migrate at high speeds within the T zone, reaching an average velocity of >10 μm/min (2, 3). This dynamic migratory behavior of T cells is thought to be important for the efficient detection of rare cognate Ags that are presented by DCs. A stromal cell subset, fibroblastic reticular cells (FRCs), constructs an elaborate network in the T zone and supports the tissue structure (4–7). T zone FRCs also express chemokines, primarily CCL21 (8, 9), which is a potent stimulator of naive T cell motility in vitro and promotes intranodal migration via the receptor CCR7 (10–12). In general, chemokines transmit signals through Gαi-coupled receptors that trigger cytoskeletal remodeling, cell polarization, and integrin activation (13, 14). Indeed, intranodal T cell migration partially depends on Gαi signaling and the integrin LFA-1 (11, 12, 15–17). Recently, ICAM-1 expressed by DCs, but not by stromal cells, was shown to be involved in LFA-1–dependent T cell motility (17). However, the mechanisms that control the motility independent of Gαi-signaling and LFA-1/ICAM-1 remain to be elucidated. Some stromal cell–derived mediators may be involved in such a motility component.

Lysophospholipids such as sphingosine-1-phosphate and lysophosphatidic acid (LPA) regulate a wide range of cellular processes, including cell motility through G protein–coupled receptors (18–20). Although sphingosine-1-phosphate is an established critical mediator for lymphocyte egress from lymphoid organs (21), a clear understanding of LPA in lymphocyte trafficking is still limited. Two groups reported that an LPA-producing ectoenzyme autotaxin (ATX), also referred to as lysophospholipase D or ectonucleotide pyrophosphatase/phosphodiesterase 2 (ENPP2) (22), were highly expressed by high endothelial venules (HEVs) in the LNs and ATX controlled adhesion and transmigration of lymphocytes (23, 24). Kanda et al. (23) suggested the expression of ATX in stromal regions of lymphoid organs and the possible involvement of ATX and LPA in interstitial lymphocyte migration. Moreover, Zhang et al. (25) recently showed that ATX and LPA stimulated naive T cell motility. However, it remains unclear whether the ATX–LPA axis in fact plays a role in T cell migration within the LN paracortex and the contribution of stromal cells in this process.

The small GTPase Rho as well as downstream pathways mediated by the Rho effector kinase ROCK and non-muscle myosin II are involved in the motility of various cells, particularly in densely packed three-dimensional environment in vitro (26–30). Activation of Rho signaling by LPA (31–35) suggests a mechanistic linkage between LPA-mediated extracellular stimuli and cell motility. However, the role of Rho activation by LPA in interstitial T cell migration in the LN remains unclear.

In this study, we report that T zone stromal cells in the LNs produce ATX. Two-photon microscopy using LN tissue slices revealed that intranodal T cell motility was controlled in part by ATX activity and LPA signaling. In vitro studies showed that LPA induced LFA-1–independent chemokinesis of T cells. Moreover, LPA and CCL21 cooperatively augment RhoA activity, which is also necessary for interstitial T cell migration. Taken together, these findings indicate that optimal high-speed T cell migration in the LN is controlled by integration of multiple environmental cues via chemokines and ATX-LPA.

Materials and Methods

Mice

Mice were maintained and bred under specific pathogen-free conditions in the animal facility of Kansai Medical University. C57BL/6 mice were purchased from CLEA Japan. To generate bone marrow (BM) chimera, CAG-EGFP mice were gamma irradiated with a single dose of 10.5 Gy from a cesium source and reconstituted with 1–5 × 106 BM cells. Two months after the BM transfer, the chimeric mice were used for experiments. All animal procedures were approved by the Committee on Animal Research at Kansai Medical University.

Antibodies

The following Abs labeled with or without fluorophores or biotin were used: anti-CD3 (145-2C11), anti-B220 (RA3-6B2), anti-CD11b (M1/70), anti-CD11c (N418), anti-CD45 (30-F11), anti-CD31 (390), anti-podoplanin (8.1.1), rat IgG2a (eBR2a) (eBioscience); ER-TR7 (BMA); anti-CCL21, anti–VCAM-1 (goat polyclonal) (R&D Systems); anti-lysophospholipase D (ATX) (rabbit polyclonal, Cayman Chemical); normal rat IgG (Caltag Laboratories); and goat anti-rat or anti-rabbit Alexa Fluor-488, -594, and -633 (Invitrogen). mAbs against LFA-1 (KBA) (36) and ICAM-1 (YN1/1; obtained from American Type Culture Collection) were purified from hybridoma supernatants.

Isolation and culture of LN stromal cells

LNs were minced using microscissors and digested with 1 mg/ml collagenase D, 0.1 mg/ml liberase-TM, and 0.1 mg/ml DNase I (Roche) in 1% FCS RPMI 1640 medium supplemented with 10 mM HEPES at 37°C for 0.5–1 h. During the enzymatic reaction, undigested tissues fragments were passed sequentially through 21-, 25-, and 27-g needles every 5–10 min. Cell suspensions were washed three times with cold 1% FCS-RPMI 1640 medium by centrifugation at 4°C. The CD45− stromal cell fraction was enriched from the LN cell suspension by magnetic cell sorting using microbeads-conjugated anti-CD45 Ab (Miltenyi Biotec). Cells were suspended in 10% FCS DMEM supplemented with penicillin, streptomycin, and 10 mM HEPES and plated onto 24-well plates coated with fibronectin (10 μg/ml at 37°C for 1 h). After incubation for 1 h, nonadherent cells were removed by gentle washing, and fresh 0.5 ml medium was added to the dishes. Cells were cultured to a confluent monolayer for 4 or 7 d without or with 10 ng/ml TNF-α (PeproTech) and 1 μg/ml anti-LTβR Ab (R&D Systems).

Flow cytometry and cell sorting

Single-cell suspensions were stained with fluorochrome-conjugated Abs by direct or indirect methods. Data were acquired using FACSCalibur or FACSCanto II (BD Biosciences) and analyzed with CellQuest software (BD Biosciences). To isolate the stromal cell fraction for microarray analysis, CD45− cells were pre-enriched from LN cell suspension with magnetic sorting using microbeads-conjugated anti-CD45 Ab (Miltenyi Biotec). The stromal cell–enriched fraction was further stained with anti-podoplanin (rabbit polyclonal IgG; Sigma-Aldrich) Ab and biotin-anti–VCAM-1 (Goat polyclonal IgG; R&D Systems) Ab, followed by PE-Cy7-anti-CD45, PE-anti-CD31, allophycocyanin-streptavidin (eBioscience), and Alexa Fluor-488–anti-rabbit IgG Abs. CD45−CD31−podoplanin+VCAM-1+ cells were then sorted with FACSAria cell sorter (BD Biosciences). Subpopulations for immune cells in LNs were sorted as follows: pan T, CD3+; CD4+ T, CD3+CD4+; CD8+ T, CD3+CD8+; B, B220+; DC, CD11chiCD11blo; and macrophage, CD11cloCD11bhi.

Microarray analysis

Total RNA was extracted from mouse embryonic fibroblasts (MEFs) and sorted stromal cell fractions with an RNeasy Plus Micro Kit (Qiagen). RNA was amplified with the WT cDNA synthesis and amplification kit (Affymetrix), and cDNA fragments were labeled with WT terminal labeling kit (Affymetrix). Samples were then hybridized to the Affymetrix GeneChip Gene 1.0 ST array for mice. Expression values for each probe set were calculated using the GC-RMA method in Affymetrix GeneChip Command Consol and Expression Console software (Affymetrix), and analyzed using GeneSpring GX 7.3.1 software (Agilent Technologies). The microarray data were deposited in the NCBI Gene Expression Omnibus database (http://www.ncbi.nlm.nih.gov/geo/) (accession number GSE55251).

Quantitative RT-PCR

Total RNA was extracted from cells with TRIzol (Invitrogen) reagent, and cDNA was synthesized using the Superscript III first-strand synthesis system and Oligo-dT primer (Invitrogen). Quantitative PCRs were performed using the SYBR Premix Ex Taq (Takara) and the appropriate primer pairs on a Thermal Cycler Dice Real-Time system (Takara). Primers used are as follows, some of them are described elsewhere (37). ATX, forward, 5′-GGAGAATCACACTGGGTAGATGATG-3′; ATX, reverse, 5′-ACGGAGGGCGGACAAAC-3′; LPA receptor (LPAR)1, forward, 5′-GAGGAATCGGGACACCATGAT-3′; LPAR1, reverse, 5′-ACATCCAGCAATAACAAGACCAATC-3′; LPAR2, forward, 5′-GACCACACTCAGCCTAGTCAAGAC-3′; LPAR2, reverse, 5′-CTTACAGTCCAGGCCATCCA-3′; LPAR3, forward, 5′-GCTCCCATGAAGCTAATGAAGACA-3′; LPAR3, reverse, 5′-AGGCCGTCCAGCAGCAGA-3′; LPAR4, forward, 5′-TGTGCGCTGGAGTCTGGATC-3′; LPAR4, reverse, 5′-GCAAGTGGTGGTCGCATTG-3′; LPAR5, forward, 5′-TGTGGGCTCTCATCCTGCTG-3′; LPAR5, reverse, 5′-GCCCTTCCACAGTTCATCGC-3′; LPAR6, forward, 5′-CGTTTGCATTGCTGTGTGGTTC-3′; LPAR6, reverse, 5′-GGCCGCTGGAAAGTTCTCAAAG-3′; GAPDH, forward, 5′-GCCAAGGTCATCCATGACAACT-3′; and GAPDH, reverse, 5′-GAGGGGCCATCCACAGTCTT-3′. To determine the absolute amounts of LPAR transcripts, full-length cDNAs were cloned to pBluscript II and used to obtain standard curve.

Immunohistochemistry

Frozen sections (10 μm) were fixed with cold acetone for 3 min and stained with Abs against ER-TR7 (BMA) and ATX (rabbit polyclonal IgG; Cayman Chemical), followed by Alexa Fluor 488-anti-rat IgG and Alexa Fluor 594-anti-rabbit IgG Abs (Invitrogen). Stained sections were examined using a confocal laser-scanning microscope (LSM510-META; Carl Zeiss). Digital images were prepared using an LSM examiner (Carl Zeiss) and Adobe Photoshop CS5 software (Adobe Systems).

T cell migration in the LN

Imaging of T cell migration in the LN slices was performed as previously described (17). For LN slices, the upper part of LNs in a 4% low gelling temperature agarose gel (type VII-A; Sigma-Aldrich) was cut off using vibration slicer (Linear Slicer Pro 10; Dosaka EM). T cells were freshly isolated from the LNs by magnetic cell sorting using a pan T cell isolation kit (Miltenyi Biotec) and labeled with 5 μM CMTMR (Invitrogen). Fluorescent-labeled T cells (2–5 × 105) were suspended in 10 μl RPMI 1640 medium supplemented with 0.1% fatty acid free (FAF)-BSA, then loaded onto the surface of the LN slice and incubated for 30 min. In some experiments, Abs (20 μg/ml) and/or pharmacological inhibitors were added to the T cell suspension and loaded onto the tissue slice. The inhibitors included 25 μM S32826 (38), 25 μM Ki16425 (39), (Cayman Chemical), 20 μM VPC32183 (40) (Avanti), 50 μM blebbistatin, and 20 μM Y27632 (Sigma-Aldrich). Alternatively, T cells were pretreated with 200 ng/ml pertussis toxin (PTx; Calbiochem) at 37°C for 1 h or 5 μg/ml cell-permeable C3 toxin (C3Tx; Rho inhibitor I; Cytoskeleton) at 37°C for 4 h before being applied to tissue slice. The LN slices were placed in a heated chamber with perfusion and time-lapse images were acquired using a two-photon microscope (FV1000MPE; Olympus). Ti:sapphire laser (MaiTai HP DeepSee-OL; Spectra-Physics) was tuned to 850 or 880 nm. Stacks of 11 to 25 x–y sections with 3-μm z-spacing were acquired every 10 or 20 s using emission wavelengths of 495–540 nm (for EGFP) and 575–630 nm (for CMTMR). Image stacks were transformed into volume-rendered four-dimensional movies and cell motility was examined with semiautomated tracking using Volocity software (Improvision). Cellular motility parameters were calculated based on the x, y, and z coordinates of cell centroids using Volocity or Microsoft Excel.

T cell migration on the stromal cell monolayer

The CD45- LN stromal cell fraction was plated onto ΔT-dishes (Bioptics) coated with fibronectin (10 μg/ml at 37°C for 1 h) and adherent cells were cultured for 7–10 d. The confluent monolayer was stimulated with 10 ng/ml TNF-α (PeproTech) for 4 h. T cells were isolated from mouse LNs by magnetic sorting (Miltenyi Biotec). A total of 2.5 × 105 T cells suspended in Leibovitz’s L-15 medium supplemented with 0.5% FAF-BSA with or without 1-oleoyl LPA (1 μM), Ki16425 (10 μM) (Cayman Chemical), and/or mouse CCL21 (R&D Systems) were loaded onto the stromal cell monolayer. After 1–3 h preincubation, phase-contrast images (512 × 512 pixels) were obtained every 5 s or 30 s at 37°C for 20 min using a LSM510 confocal microscope (Carl Zeiss) equipped with a heated stage adapter for ΔT-dishes (Bioptics) (41). Blocking Abs (20 μg/ml) were added 20 min prior to initiating image collection. To inhibit Gαi activity, cells were pretreated with 200 ng/ml PTx or B-oligomer (Calbiochem) for 1 h before being loading onto the stromal monolayer. For T cell motility under stromal cell–free condition, T cells were applied onto ΔT-dishes that were precoated with 1% FAF-BSA at 37°C for 1 h. The image data were analyzed using the ImagePro Plus software (Media Cybernatics). Alternatively, tracking data from CMTMR-labeled T cells was analyzed with Volocity software (Improvision). The trajectories of T cells that were clearly influenced by the thermal convection of medium, particularly in stromal cell–free setting, were excluded from the analysis.

Detection of RhoA and Rap1 activation

T cells were suspended at 2 × 107 cells/ml in RPMI 1640 medium supplemented with 0.1% FAF-BSA and 10 mM HEPES and stimulated with LPA (Cayman Chemical) or mouse CCL21 (R&D Systems) in the presence or absence of Ki16425 (20 μM) at 37°C for 0.5 or 2 min. Cells were collected by centrifugation and immediately resuspended in lysis buffer. Cleared cell lysates were subsequently used for pull-down assays. GTP-bound RhoA was quantitated using the Rho activation assay biochem kit (using GST-Rhotekin-RBD fusion protein; Cytoskeleton) (42). GTP-bound Rap1 was measured using a GST-RalGDS-RBD fusion protein and anti-Rap1 Ab (Transduction Laboratories) as described previously (43).

Statistical analysis

GraphPad Prism was used for statistical analyses. The means of two groups were compared using a Student t test. Mann–Whitney U test was used to compare two nonparametric datasets.

Results

ATX is highly expressed in LN stromal cells

To identify stromal cell–derived factors that regulate immune function, specifically lymphocyte motility, we performed comparative gene expression profiling between LN stromal cells and MEFs as a mesenchymal control. The CD45−CD31−podoplanin+VCAM-1+ stromal cell fraction was sorted from enzymatically digested mouse LNs. Total RNAs extracted from stromal cells and MEFs were used for microarray analysis to assess relative gene expression. We identified genes that were selectively expressed in stromal cells compared with MEFs (Fig. 1A). Importantly, the reliability of the results were demonstrated by the high expression of genes encoding homeostatic chemokines including CXCL13, CCL19, and CCL21, which are the signatures of stromal cells in lymphoid tissues (Fig. 1A, upper panel). High expression of complement components (C1s/r, C2, C3, C4, C7, Cfb, and Cfh) and cytokines such as IL-6, IL-33, and BAFF in stromal cells were also remarkable. Moreover, we found that LN stromal cells expressed Enpp2 encoding ATX more than a hundred fold relative to MEFs (Fig. 1A, upper panel). High expression of Enpp2 in the LN stromal fraction was also confirmed in a public database (ImmGen; http://www.immgen.org) (44). Because ATX and its product LPA have been implicated in cell migration, we focused on this molecule for further in depth analysis. Expression of ATX in the stromal fraction was confirmed by quantitative RT-PCR analysis; T cells, B cells, DCs, and MEFs expressed ATX at negligible levels (Fig. 1B). Of note, cultured LN stromal cells rapidly lost their expression of ATX, even in the presence of TNF-α and lymphotoxin (Fig. 1C), suggesting that ATX expression was optimally maintained by unidentified pressures in the tissue environment in vivo.

FIGURE 1.
FIGURE 1.

The ATX gene was highly expressed in LN stromal cells. (A) Comparative microarray analysis of LN stromal cells and MEFs. A scatter plot representing log2 intensities of relative gene expression. The red-boxed region is magnified in the upper panel, with red arrow indicating ATX (ENPP2). Several known genes that were highly expressed in stromal cells are also indicated. (B) Validation of ATX expression by quantitative RT-PCR analysis. Data shown were normalized to GAPDH. Stromal cells (St), T cells (T), and B cells (B). (C) Quantitative RT-PCR analysis for ATX expression in cultured LN stromal cells. CD45−-enriched LN stromal cells were cultured for 4 or 7 d with or without 10 ng/ml TNF-α and 1 μg/ml anti-LTβR Ab (T+L). ATX expression was lost during the culture.

Stromal cells in the T zone produce and display ATX

Although ATX is a secreted protein (45, 46), we speculated that ATX might be immobilized on the stromal cell surface, similar to chemokines. To test this, single-cell suspensions that were prepared from digested LNs were stained with a polyclonal Ab against ATX and several other markers for flow cytometric analysis. As expected, ATX protein was detected on the surface of a fraction of CD45−CD31− stromal cells (Fig. 2A), suggesting that some distinct stromal subpopulation produces and presents ATX. Similar to previous reports, ATX was also expressed by a fraction of CD45−CD31+ endothelial cells, which would correspond to HEV endothelial cells (23, 24). Consistent with quantitative RT-PCR analysis, surface ATX was undetectable in T cells, B cells, and DCs. To further characterize the subpopulation that displayed ATX, CD45− LN cells were separated based on differential expression of the endothelial marker CD31 and cell surface–bound CCL21, which are characteristic of HEV endothelial cells and FRCs (Fig. 2B). In the CD45− fraction, ATX was detected on CD31+CCL21+ endothelial cells and CD31−CCL21+ FRCs. In contrast, both CD31+CCL21− and CD31−CCL21− cells had low surface expression of ATX (Fig. 2C). Therefore, stromal cells in the T zone produce and display ATX as well as CCL21.

FIGURE 2.
FIGURE 2.

Expression of ATX on the surface of stromal cells in the T zone. (A) Staining of cell surface ATX in LN cell subpopulations. Single-cell suspensions from enzyme-digested LNs were stained for ATX and lineage markers for flow cytometric analysis. Gating was as follows: stromal cells (St), CD45−CD31−gp38+; endothelial cells (End), CD45-CD31+; T cells, CD3+; B cells, B220+; and DCs, CD11chiCD11blow. ATX staining is shown as the shaded histogram, and the white histogram represents control staining with rabbit IgG. Cell surface ATX was detected in a fraction of stromal cells (arrowhead) and endothelial cells (small arrow). (B and C) ATX expression in stromal and endothelial subsets. Single-cell suspensions of LNs were stained for ATX, CD45, CD31, and CCL21 for flow cytometric analysis. (B) Dot plot shows the expression of CD31 and surface-bound CCL21 in CD45−-gated cells. (Ca–Cd) ATX staining of cells in the gated regions (Ba–Bd). Histograms represent ATX (shaded) and control (open) staining. The stromal cell fraction that expressed surface-bound CCL21 also displayed ATX on the surface (Bd and Cd, bold boxes).

We also stained LN tissue sections with Abs against ATX and ER-TR7, a marker for the reticular fiber of the stromal network. ATX staining was relatively strong in HEVs and weak throughout the parenchyma, whereas a fibrous staining pattern was colocalized with the ER-TR7+ stromal network in the T zone (Fig. 3), thus confirming the production of ATX by reticular-forming FRCs in this area.

FIGURE 3.
FIGURE 3.

Detection of ATX in the LN T zone. LN cryosections were stained with control rabbit IgG (upper panels) or anti-ATX Ab (lower panels) plus ER-TR7, and examined by confocal microscopy. The outer cortex of the LN in serial sections is shown. The dotted line indicates the border between the T zone (T) and follicles (B). Some filamentous staining patterns of ATX (arrowheads) were colocalized with FRC network in the T zone. Relatively strong staining of ATX was also detected in the HEV (arrows).

ATX–LPA axis promotes interstitial T cell migration in the LN

Given that ATX is produced by the T zone FRCs, we sought to determine whether ATX and its product LPA were involved in interstitial T cell migration. We performed live imaging of LN tissue slices using two-photon microscopy. Freshly isolated T cells were labeled with fluorescent dye and loaded onto the LN slice in the presence or absence of pharmacological inhibitors. In the presence of S32826, an inhibitor of ATX enzymatic activity (38), T cell migration velocity was significantly reduced compared with the control (Fig. 4A, 4B, Supplemental Video 1). To inhibit LPA signals, we used two antagonists of LPARs, Ki16425 (39) or VPC32183 (40). The migration velocity of T cells was inhibited by both LPAR inhibitors to a similar extent as the ATX inhibitor (Fig. 4C, 4D, Supplemental Video 2; data not shown). In addition, simultaneous treatment of S32826 and Ki16425 did not show an additive effect on the inhibition of T cell motility (Fig. 4E), suggesting that ATX and LPA act in the same pathway to stimulate T cell motility. Taken together, these findings suggest that the ATX–LPA axis promotes interstitial migration of T cells in the LN.

FIGURE 4.
FIGURE 4.

Inhibition of the ATX–LPA axis reduces interstitial T cell migration in the LNs. (A and B) Migration velocities of T cells in LN slices treated with DMSO (control) or an ATX inhibitor S32826 (25 μM). (A) The mean velocity of individual cells (circles) and the median (horizontal bars). (B) The median velocity of individual experiments (circles) and the mean (horizontal bars). (C and D) Migration velocities of T cells treated with DMSO (control) or the LPAR inhibitor Ki16425 (25 μM). The mean velocity of individual cells (C) and the median velocity of individual experiments (D). (E) Migration velocities of T cells treated with DMSO (control), S32826 (25 μM), Ki16425 (25 μM), or both inhibitors.

To examine whether ATX/LPA signaling depends on Gαi, T cells pretreated with PTx were applied onto LN slices in the presence or absence of the inhibitors for ATX or LPARs. Interestingly, simultaneous inhibition of ATX or LPAR further exacerbated the PTx-mediated reduction of T cell migration (Fig. 5A, 5B, Supplemental Video 3). To determine whether ATX/LPA-dependent motility required LFA-1 function, we treated LN slices with a combination of anti–LFA-1 Ab and ATX/LPA inhibitors. Similar to PTx, ATX/LPA inhibitors further enhanced the effect of LFA-1 blockage (Fig. 5C, Supplemental Video 4). These findings strongly support the notion that the fraction of T cell motility that is mediated by ATX–LPA axis occurs independently of Gαi-signaling and LFA-1.

FIGURE 5.
FIGURE 5.

ATX–LPA axis is independent of Gi signal and LFA-1 in intranodal T cell migration. (A and B) Migration velocities of T cells in LN slices treated with PTx plus S32826 (A) or Ki16425 (B). The mean velocity of individual cells (circles) and the median (horizontal bars). (C) Migration velocities of T cells in LN slices treated with an anti–LFA-1 Ab and/or ATX/LPAR inhibitors.

LPA induces chemokinesis in T cells

Previous reports have shown that LPA induces T cell motility in vitro (23, 25). We also confirmed the migratory activity of LPA using Transwell chambers. The optimal concentration of LPA was 1 μM in the upper compartment but not in the lower (data not shown), indicating that LPA stimulates chemokinetic rather than chemotactic T cell motility. To determine the role of LPA more specifically, we examined T cell behavior on a monolayer of primary LN stromal cells. Because LN stromal cells have lost the expressions of ATX as well as CCL21 during the culture (Fig. 1C, Supplemental Fig. 1), T cells applied onto the monolayer did not show any active migration. However, the addition of CCL21 induced dose-dependent T cell motility, which was completely dependent upon Gαi and LFA-1/ICAM-1 (Supplemental Fig. 2). Interestingly, the addition of LPA alone significantly augmented T cell motility, which was slower and more random than CCL21 (Figs. 6A, 6B, Supplemental Video 5). LPAR inhibitor Ki16425 abrogated this activity, whereas anti–LFA-1 Ab had no inhibitory effect (Fig. 6B, 6C). LPA was also able to stimulate Ki16425-sensitive T cell motility under a stromal cell–free condition (Fig. 6D, Supplemental Video 6), thus indicating that LPA-induced motility does not require LFA-1–mediated attachment to stromal cells or the indirect effects of LPA through stromal cells. LPA induced an intermittent polarized morphology with migration in a short range before stopping (Fig. 6E, Supplemental Video 7). Moreover, LPA was able to enhance T cell motility that was induced by suboptimal but not higher concentrations of CCL21, and Ki16425 treatment abrogated the effects of LPA (Fig. 6F, 6G). These results show that LPA stimulates LFA-1–independent T cell chemokinesis distinct from that induced by chemokines.

FIGURE 6.
FIGURE 6.

LPA induces LFA-1–independent chemokinesis in T cells in vitro. (A) Trajectories of T cell migration on the monolayer of LN stromal cells in the presence of LPA (1 μM) or CCL21 (100 nM). Cells in the final frame are shown as red silhouettes. (B–D) Migration velocities of T cells in response to LPA plus treatment with Ki16425 (B) or anti–LFA-1 Ab (C), or in the absence of stromal cells (D). The mean velocity of individual cells (circles) and the median (horizontal bars) are shown. (E) Time-lapse images of a T cell migrating in the presence of LPA. The red line represents the trajectory. The cell showed intermittent, short-distance motility with a transient polarized morphology. (F and G) Migration velocities of T cells treated with CCL21 and/or LPA. (F) Enhancing effect of LPA on CCL21-induced T cell migration was observed at a suboptimal (30 nM) but not saturated (100 nM) concentration of CCL21. (G) Ki16425 inhibited the enhancing effect of LPA under suboptimal concentration of CCL21 (30 nM).

Because six LPA-specific receptors (LPAR1-6) have been identified to date (47), quantitative PCR was used to determine the expression of these receptors in LN-derived T cells. Among the six receptors, LPAR5 and LPAR6 were expressed at relatively high levels in both CD4+ and CD8+ T cells (Supplemental Fig. 3A, 3B), whereas LPAR2 was detected at low levels; expression of LPAR1, LPAR3, and LPAR4 were undetectable. Ki16425 is known to inhibit LPAR1 and LPAR3, and it inhibits LPAR2 only slightly at higher concentrations (39). One report demonstrated that Ki16425 did not inhibit LPAR5 (48). In addition, we observed that Ki16425 did not block LPA-induced morphological changes of the Ba/F3 lymphoid cell line reconstituted with LPAR6 (data not shown). Therefore, currently the receptor responsible for LPA-mediated T cell motility sensitive to Ki16425 remains unclear. Uncharacterized LPARs may regulate this process or LPAR2 could be involved, despite its low expression and relative insensitivity to Ki16425. As LPARs are also expressed in stromal cell fraction (Supplemental Fig. 3A, 3C), the inhibitors possibly affect T cell motility indirectly through stromal cells. Although we did not observed this effect in vitro, we could not exclude the autocrine or paracrine effect of LPA on FRCs that possibly affect T cell motility in vivo.

LPA and CCL21 cooperatively augment RhoA activity in T cells

LPA has been reported to activate the small GTPase Rho, which regulates cell morphology and motility (31–35). However, whether LPA can also activate Rho in T cells and the interplay between LPA- and chemokine-signaling remain unknown. Thus we measured RhoA activity after the stimulation of T cells with LPA and/or CCL21. Pull-down detection of the GTP-bound active form of RhoA showed that LPA indeed activated RhoA in T cells (Fig. 7A left). The maximum RhoA activation occurred at 1 μM of LPA, which was in agreement with the migration-stimulating activity of LPA. CCL21 also activated RhoA in a dose-dependent manner (Fig. 7A, right). Notably, LPA and CCL21 showed an additive effect on the activation of RhoA, and Ki16425 selectively inhibited LPA-dependent activity (Fig. 7B, 7C). LPA did not activate the small GTPase Rap1, a critical regulator of chemokine-induced migration and LFA-1 activation in lymphocytes (Fig. 7D) (13, 43). In the pull-down assay, Rap1 activation was only detected within 1 min after CCL21 stimulation, whereas RhoA activation was not detected at this time point. In contrast, LPA activated RhoA but not Rap1 at this early point (1 min). These patterns of small GTPase activation are largely consistent with the observation that LPA does not stimulate robust LFA-1–dependent motility in T cells.

FIGURE 7.
FIGURE 7.

LPA induces Rho activation in T cells. (A) Activation of RhoA in T cells stimulated with LPA or CCL21. T cells were treated with the indicated concentrations of LPA or CCL21 for 2 min at 37°C, and cell lysates were harvested. Activated GTP-RhoA in cell lysates was detected by pull-down and Western blotting analysis. Total RhoA in cell lysates was detected by direct western blotting. (B and C) RhoA activation in T cells stimulated with LPA (1 μM) and/or CCL21 (100 nM). (C) Ki16425 inhibited LPA-mediated but not CCL21-mediated RhoA activation. Quantitative measurements of the bands were normalized to total RhoA (lower graph in C) and the fold-increase of band intensities shown was relative to untreated control (none). (D) Activation of Rap1 and RhoA in T cells in response to LPA and/or CCL21. T cells were treated with LPA and/or CCL21 for 1 min at 37°C, and activated GTP-Rap1 or GTP-RhoA in cell lysates were detected by pull-down assays. Note that CCL21 does not activate RhoA at this time point.

Rho–ROCK–myosin II axis is required for T cell migration in the LN

The role of Rho activation in intranodal T cell migration remains to be elucidated. To this end, we took advantage of a cell permeable form of botulinum C3Tx that prevents the activation of all three Rho subtypes (49). T cells pretreated with C3Tx were loaded onto LN slices and examined with two-photon microscopy. Although a fraction of Rho-inhibited T cells could enter the tissue, motility of the cells was markedly impaired (Fig. 8A, Supplemental Video 8). An inhibitor of ROCK, Y27632, and a myosin II inhibitor, blebbistatin, also reduced T cell migration in LNs (Fig. 8B, 8C), indicating that the Rho–ROCK–myosin II pathway plays a critical role in interstitial T cell migration.

FIGURE 8.
FIGURE 8.

The Rho–ROCK–myosin II pathway controls intranodal T cell migration and LFA-1–independent motility. (A) Migration velocities of C3Tx-pretreated T cells on LN slices. (B and C) Migration velocities of T cells in LN slices in the presence of Y27632 (B) or blebbistatin (C). (D) Representative migratory morphology of T cells treated with C3Tx or blebbistatin. Z-projected time-lapse images obtained using LN slices from GFP BM chimera mice. The migration of T cells (red) in the network of FRCs (green) is shown. C3Tx (middle) or blebbistatin (bottom) induced a typical T cell morphology with a swollen rear part and no uropod-like squeezed shape (asterisks) but extensive protrusions in the front (arrows). Note that the time spans in the image sequences are 50 s for control, 2 min 48 s for C3Tx, and 1 min 50 s for blebbistatin. (E and F) Migration velocities of T cells in LN slices treated with combinations of anti–LFA-1 Ab and Y27632 (E) or blebbistatin (Blebb.) (F).

Migrating T cells show a dynamic polarized morphology with a cycle of elongation and contraction. C3Tx treatment influenced the morphology of migrating T cells. Many cells had a rounded shape, but some were still motile as characterized by an extensive protrusion at the front of the cell and a swollen rear part without an uropod-like squeezed shape (Fig. 8D, middle, Supplemental Videos 9, 10). These migrating cells exhibited a sliding-like slow movement. Blebbistatin-treated T cells showed similar morphological and migratory features (Fig. 8D, bottom, Supplemental Video 9), suggesting that the appropriate cycles of morphological changes controlled by Rho and myosin II are required for the efficient forward movement of T cells. Importantly, the residual motility in the presence of anti–LFA-1 Ab was abrogated by the addition of Y27632 or blebbistatin (Fig. 8E, 8F, Supplemental Video 11). Therefore, LFA-1–independent T cell motility in the LNs is largely regulated by the Rho–ROCK–myosin II axis.

Discussion

This study shows that ATX and LPA produced by stromal cells in the LN T zone serve as migratory cues that drive high-speed interstitial T cell migration in this region. Importantly, the fraction of T cell motility mediated by the ATX–LPA axis was independent of Gαi-signaling and LFA-1, but likely depended on the Rho–ROCK–myosin II pathway. Our current findings taken together with our previous observations (17) suggest that multiple environmental cues provided by the network of FRCs and DCs coordinately regulate intranodal T cell migration in the LN T zone (Fig. 9).

FIGURE 9.
FIGURE 9.

FRCs and DCs stimulate high-speed T cell migration in the LN T zone. Schematic representation of environmental migration cues provided by FRCs and DCs in the LN T zone that support high-speed T cell migration via stimulation of signaling pathways.

We previously demonstrated that LFA-1–dependent T cell motility requires ICAM-1 expressed by DCs but not by stromal cells (17). In the current study, we showed that FRCs play an indispensable role in T cell migration via the production of motility factors, including chemokines and ATX/LPA. CCL21 secreted from FRCs is immobilized on the cell surface but a fraction of CCL21 also can bind to the surface of DCs through interactions with heparan sulfate proteoglycans (50), where it is able to stimulate T cell motility and facilitate LFA-1–dependent adhesion to ICAM-1. ATX is also able to bind to heparan sulfate through an internal motif rich in basic amino acid residues (51) and it is detected on the surface of stromal cells, but not DCs. Although the mechanism underlying the delivery of ATX-derived LPA to T cells remains obscure, recent structural information has suggested a possibility of direct LPA transport from ATX to the receptors (52, 53). This suggests that immobilized ATX/LPA works locally on the stromal cell surface, which stimulates LFA-1–independent motility. Thus, relative amounts of CCL21 and ATX/LPA in the microenvironment could determine in situ dependency of T cell migration on LFA-1. These two pathways may act complementary or compensatory to obtain the optimal total migratory activity of T cells. This idea is consistent with the fact that LPA enhances T cell motility induced by a suboptimal amount of CCL21. It would be intriguing to examine in the future whether this balance changes under inflammatory conditions.

To date, there are at least six LPARs (LPAR1–6), and recent reports have identified putative LPARs (47). We determined that T cells express relatively high levels of LPAR6 and LPAR5, and low level of LPAR2; expression of LPAR1 and LPAR3, which are the known targets of Ki16425, are undetectable. Interestingly in our in vitro experimental settings including a motility analysis under stromal cell–free condition or Rho-activation assay, Ki16425 directly inhibited the responsiveness of T cells to LPA, suggesting that the presence of Ki16425-sensitive unknown receptors or even the low expression of LPAR2 were sufficient to transmit motility signals, which could be inhibited by relatively high concentrations of Ki16425. Nevertheless, we could not completely exclude the possibility that LPA facilitated T cell motility indirectly through FRCs or DCs in vivo. The patterns of Gα types coupled to LPARs are diverse depending on receptors and tissues (19). At least in intranodal T cell migration, LPA signals are independent of Gαi signaling, which is consistent with the fact that LPA-mediated phenomena are independent of the Gαi–Rap1–LFA-1 axis.

LPA stimulates RhoA activation in T cells, which triggers a downstream cascade toward the reorganization of the actin cytoskeleton and myosin II–mediated contractility. Efficient intranodal T cell migration requires signaling via the Rho–ROCK–myosin II pathway. Inhibition of the ATX–LPA axis is not as effective as the inhibition of the Rho pathway, which is likely because CCL21 also activates RhoA. Of note, the Rho–myosin II pathway appears to be a driving force for LFA-1–independent interstitial T cell motility facilitated by the “confinement effect” in a three-dimensional densely packed tissue microenvironment (28, 29). Although the ATX–LPA axis is a potential mediator of this type of migration, the traction force generated by the transient adhesion of T cells on the tissue scaffold via LFA-1/ICAM-1 is also indispensable for extremely high-speed migration (17). Simultaneous inhibition of these pathways, which was achieved using the LN slice technique, dramatically suppressed T cell migration. Therefore, these findings highlight the requirement of multiple molecular axes that regulate both adhesion-dependent and confinement-dependent migration modalities.

Taken together, our findings suggest that the migratory cues produced by T zone microenvironment are essential for the optimal high-speed interstitial T cell motility that leads to efficient Ag scanning within this immunologically specialized LN tissue compartment.

Disclosures

The authors have no financial conflicts of interest.

Acknowledgments

We thank M. Okabe for providing the CAG-EGFP mice, J. Aoki and S. Okudaira for ATX Abs, and R. Hamaguchi, H. Takano, and A. Kawasaki for technical assistance.

Footnotes

  • This work was supported in part by Grants-in-Aid for Young Scientists (20689005 and 22790194; to T. Katakai), a Grant-in-Aid for Scientific Research (22370072; to T. Kinashi), Grants-in-Aid for Scientific Research on Innovative Areas “Fluorescence Live Imaging” (23113517; to T. Katakai), “Analysis and Synthesis of Multidimensional Immune Organ Network” (24111005; to T. Katakai), and “Cross-Talk between Moving Cells and Microenvironment” (22111003; to T. Kinashi) from the Ministry of Education, Culture, Sports, Science, and Technology of Japan; a Core Research for Evolutional Science and Technology grant from the Japan Science and Technology Agency (to T. Kinashi); and a research grant from the Takeda Science Foundation (to T. Kinashi). The funding agencies had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

  • The microarray data presented in this article have been submitted to the Gene Expression Omnibus (http://www.ncbi.nlm.nih.gov/geo/) under accession number GSE55251.

  • The online version of this article contains supplemental material.

  • Abbreviations used in this article:

    ATX
    autotaxin
    BM
    bone marrow
    C3Tx
    C3 toxin
    DC
    dendritic cell
    ENPP2
    ectonucleotide pyrophosphatase/phosphodiesterase 2
    FAF
    fatty acid free
    FRC
    fibroblastic reticular cell
    HEV
    high endothelial venule
    LN
    lymph node
    LPA
    lysophosphatidic acid
    LPAR
    LPA receptor
    MEF
    mouse embryonic fibroblast
    PTx
    pertussis toxin.

  • Received March 4, 2014.
  • Accepted May 15, 2014.

References

  1. ↵
    1. Steinman R. M.,
    2. M. Pack,
    3. K. Inaba
    . 1997. Dendritic cells in the T-cell areas of lymphoid organs. Immunol. Rev. 156: 25–37.
    OpenUrlCrossRefPubMed
  2. ↵
    1. Miller M. J.,
    2. S. H. Wei,
    3. I. Parker,
    4. M. D. Cahalan
    . 2002. Two-photon imaging of lymphocyte motility and antigen response in intact lymph node. Science 296: 1869–1873.
    OpenUrlAbstract/FREE Full Text
  3. ↵
    1. Mempel T. R.,
    2. S. E. Henrickson,
    3. U. H. Von Andrian
    . 2004. T-cell priming by dendritic cells in lymph nodes occurs in three distinct phases. Nature 427: 154–159.
    OpenUrlCrossRefPubMed
  4. ↵
    1. Gretz J. E.,
    2. A. O. Anderson,
    3. S. Shaw
    . 1997. Cords, channels, corridors and conduits: critical architectural elements facilitating cell interactions in the lymph node cortex. Immunol. Rev. 156: 11–24.
    OpenUrlCrossRefPubMed
    1. Katakai T.,
    2. T. Hara,
    3. J. H. Lee,
    4. H. Gonda,
    5. M. Sugai,
    6. A. Shimizu
    . 2004. A novel reticular stromal structure in lymph node cortex: an immuno-platform for interactions among dendritic cells, T cells and B cells. Int. Immunol. 16: 1133–1142.
    OpenUrlAbstract/FREE Full Text
    1. Katakai T.,
    2. T. Hara,
    3. M. Sugai,
    4. H. Gonda,
    5. A. Shimizu
    . 2004. Lymph node fibroblastic reticular cells construct the stromal reticulum via contact with lymphocytes. J. Exp. Med. 200: 783–795.
    OpenUrlAbstract/FREE Full Text
  7. ↵
    1. Bajénoff M.,
    2. J. G. Egen,
    3. L. Y. Koo,
    4. J. P. Laugier,
    5. F. Brau,
    6. N. Glaichenhaus,
    7. R. N. Germain
    . 2006. Stromal cell networks regulate lymphocyte entry, migration, and territoriality in lymph nodes. Immunity 25: 989–1001.
    OpenUrlCrossRefPubMed
  8. ↵
    1. Cyster J. G.
    1999. Chemokines and cell migration in secondary lymphoid organs. Science 286: 2098–2102.
    OpenUrlAbstract/FREE Full Text
  9. ↵
    1. Luther S. A.,
    2. H. L. Tang,
    3. P. L. Hyman,
    4. A. G. Farr,
    5. J. G. Cyster
    . 2000. Coexpression of the chemokines ELC and SLC by T zone stromal cells and deletion of the ELC gene in the plt/plt mouse. Proc. Natl. Acad. Sci. USA 97: 12694–12699.
    OpenUrlAbstract/FREE Full Text
  10. ↵
    1. Worbs T.,
    2. T. R. Mempel,
    3. J. Bölter,
    4. U. H. von Andrian,
    5. R. Förster
    . 2007. CCR7 ligands stimulate the intranodal motility of T lymphocytes in vivo. J. Exp. Med. 204: 489–495.
    OpenUrlAbstract/FREE Full Text
  11. ↵
    1. Okada T.,
    2. J. G. Cyster
    . 2007. CC chemokine receptor 7 contributes to Gi-dependent T cell motility in the lymph node. J. Immunol. 178: 2973–2978.
    OpenUrlAbstract/FREE Full Text
  12. ↵
    1. Huang J. H.,
    2. L. I. Cárdenas-Navia,
    3. C. C. Caldwell,
    4. T. J. Plumb,
    5. C. G. Radu,
    6. P. N. Rocha,
    7. T. Wilder,
    8. J. S. Bromberg,
    9. B. N. Cronstein,
    10. M. Sitkovsky,
    11. et al
    . 2007. Requirements for T lymphocyte migration in explanted lymph nodes. J. Immunol. 178: 7747–7755.
    OpenUrlAbstract/FREE Full Text
  13. ↵
    1. Kinashi T.
    2005. Intracellular signalling controlling integrin activation in lymphocytes. Nat. Rev. Immunol. 5: 546–559.
    OpenUrlCrossRefPubMed
  14. ↵
    1. Thelen M.,
    2. J. V. Stein
    . 2008. How chemokines invite leukocytes to dance. Nat. Immunol. 9: 953–959.
    OpenUrlCrossRefPubMed
  15. ↵
    1. Boscacci R. T.,
    2. F. Pfeiffer,
    3. K. Gollmer,
    4. A. I. C. Sevilla,
    5. A. M. Martin,
    6. S. F. Soriano,
    7. D. Natale,
    8. S. Henrickson,
    9. U. H. von Andrian,
    10. Y. Fukui,
    11. et al
    . 2010. Comprehensive analysis of lymph node stroma-expressed Ig superfamily members reveals redundant and nonredundant roles for ICAM-1, ICAM-2, and VCAM-1 in lymphocyte homing. Blood 116: 915–925.
    OpenUrlAbstract/FREE Full Text
    1. Raab M.,
    2. H. Wang,
    3. Y. Lu,
    4. X. Smith,
    5. Z. Wu,
    6. K. Strebhardt,
    7. J. E. Ladbury,
    8. C. E. Rudd
    . 2010. T cell receptor “inside-out” pathway via signaling module SKAP1-RapL regulates T cell motility and interactions in lymph nodes. Immunity 32: 541–556.
    OpenUrlCrossRefPubMed
  17. ↵
    1. Katakai T.,
    2. K. Habiro,
    3. T. Kinashi
    . 2013. Dendritic cells regulate high-speed interstitial T cell migration in the lymph node via LFA-1/ICAM-1. J. Immunol. 191: 1188–1199.
    OpenUrlAbstract/FREE Full Text
  18. ↵
    1. Ishii I.,
    2. N. Fukushima,
    3. X. Ye,
    4. J. Chun
    . 2004. Lysophospholipid receptors: signaling and biology. Annu. Rev. Biochem. 73: 321–354.
    OpenUrlCrossRefPubMed
  19. ↵
    1. Choi J. W.,
    2. D. R. Herr,
    3. K. Noguchi,
    4. Y. C. Yung,
    5. C. W. Lee,
    6. T. Mutoh,
    7. M. E. Lin,
    8. S. T. Teo,
    9. K. E. Park,
    10. A. N. Mosley,
    11. J. Chun
    . 2010. LPA receptors: subtypes and biological actions. Annu. Rev. Pharmacol. Toxicol. 50: 157–186.
    OpenUrlCrossRefPubMed
  20. ↵
    1. Lin D. A.,
    2. J. A. Boyce
    . 2006. Lysophospholipids as mediators of immunity. Adv. Immunol. 89: 141–167.
    OpenUrlCrossRefPubMed
  21. ↵
    1. Cyster J. G.,
    2. S. R. Schwab
    . 2012. Sphingosine-1-phosphate and lymphocyte egress from lymphoid organs. Annu. Rev. Immunol. 30: 69–94.
    OpenUrlCrossRefPubMed
  22. ↵
    1. Nakanaga K.,
    2. K. Hama,
    3. J. Aoki
    . 2010. Autotaxin—an LPA producing enzyme with diverse functions. J. Biochem. 148: 13–24.
    OpenUrlAbstract/FREE Full Text
  23. ↵
    1. Kanda H.,
    2. R. Newton,
    3. R. Klein,
    4. Y. Morita,
    5. M. D. Gunn,
    6. S. D. Rosen
    . 2008. Autotaxin, an ectoenzyme that produces lysophosphatidic acid, promotes the entry of lymphocytes into secondary lymphoid organs. Nat. Immunol. 9: 415–423.
    OpenUrlCrossRefPubMed
  24. ↵
    1. Nakasaki T.,
    2. T. Tanaka,
    3. S. Okudaira,
    4. M. Hirosawa,
    5. E. Umemoto,
    6. K. Otani,
    7. S. Jin,
    8. Z. Bai,
    9. H. Hayasaka,
    10. Y. Fukui,
    11. et al
    . 2008. Involvement of the lysophosphatidic acid-generating enzyme autotaxin in lymphocyte-endothelial cell interactions. Am. J. Pathol. 173: 1566–1576.
    OpenUrlCrossRefPubMed
  25. ↵
    1. Zhang Y.,
    2. Y. C. Chen,
    3. M. F. Krummel,
    4. S. D. Rosen
    . 2012. Autotaxin through lysophosphatidic acid stimulates polarization, motility, and transendothelial migration of naive T cells. J. Immunol. 189: 3914–3924.
    OpenUrlAbstract/FREE Full Text
  26. ↵
    1. Fukata Y.,
    2. M. Amano,
    3. K. Kaibuchi
    . 2001. Rho-Rho-kinase pathway in smooth muscle contraction and cytoskeletal reorganization of non-muscle cells. Trends Pharmacol. Sci. 22: 32–39.
    OpenUrlCrossRefPubMed
    1. Riento K.,
    2. A. J. Ridley
    . 2003. Rocks: multifunctional kinases in cell behaviour. Nat. Rev. Mol. Cell Biol. 4: 446–456.
    OpenUrlCrossRefPubMed
  28. ↵
    1. Petrie R. J.,
    2. K. M. Yamada
    . 2012. At the leading edge of three-dimensional cell migration. J. Cell Sci. 125: 5917–5926.
    OpenUrlAbstract/FREE Full Text
  29. ↵
    1. Jacobelli J.,
    2. F. C. Bennett,
    3. P. Pandurangi,
    4. A. J. Tooley,
    5. M. F. Krummel
    . 2009. Myosin-IIA and ICAM-1 regulate the interchange between two distinct modes of T cell migration. J. Immunol. 182: 2041–2050.
    OpenUrlAbstract/FREE Full Text
  30. ↵
    1. Jacobelli J.,
    2. R. S. Friedman,
    3. M. A. Conti,
    4. A. M. Lennon-Dumenil,
    5. M. Piel,
    6. C. M. Sorensen,
    7. R. S. Adelstein,
    8. M. F. Krummel
    . 2010. Confinement-optimized three-dimensional T cell amoeboid motility is modulated via myosin IIA-regulated adhesions. Nat. Immunol. 11: 953–961.
    OpenUrlCrossRefPubMed
  31. ↵
    1. Moolenaar W. H.,
    2. L. A. van Meeteren,
    3. B. N. G. Giepmans
    . 2004. The ins and outs of lysophosphatidic acid signaling. BioEssays 26: 870–881.
    OpenUrlCrossRefPubMed
    1. Kranenburg O.,
    2. M. Poland,
    3. F. P. G. van Horck,
    4. D. Drechsel,
    5. A. Hall,
    6. W. H. Moolenaar
    . 1999. Activation of RhoA by lysophosphatidic acid and Gα12/13 subunits in neuronal cells: induction of neurite retraction. Mol. Biol. Cell 10: 1851–1857.
    OpenUrlAbstract/FREE Full Text
    1. Bian D.,
    2. C. Mahanivong,
    3. J. Yu,
    4. S. M. Frisch,
    5. Z. K. Pan,
    6. R. D. Ye,
    7. S. Huang
    . 2006. The G12/13-RhoA signaling pathway contributes to efficient lysophosphatidic acid-stimulated cell migration. Oncogene 25: 2234–2244.
    OpenUrlCrossRefPubMed
    1. Herroeder S.,
    2. P. Reichardt,
    3. A. Sassmann,
    4. B. Zimmermann,
    5. D. Jaeneke,
    6. J. Hoeckner,
    7. M. W. Hollmann,
    8. K. D. Fischer,
    9. S. Vogt,
    10. R. Grosse,
    11. et al
    . 2009. Guanine nucleotide-binding proteins of the G12 family shape immune functions by controlling CD4+ T cell adhesiveness and motility. Immunity 30: 708–720.
    OpenUrlCrossRefPubMed
  35. ↵
    1. Kim J. H.,
    2. R. S. Adelstein
    . 2011. LPA(1)-induced migration requires nonmuscle myosin II light chain phosphorylation in breast cancer cells. J. Cell. Physiol. 226: 2881–2893.
    OpenUrlCrossRefPubMed
  36. ↵
    1. Kato K.,
    2. N. Sato,
    3. T. Tanabe,
    4. H. Yagita,
    5. T. Agatsuma,
    6. Y. Hashimoto
    . 1991. Establishment of mouse lymphokine-activated killer cell clones and their properties. Jpn. J. Cancer Res. 82: 456–463.
    OpenUrlCrossRef
  37. ↵
    1. Kishi Y.,
    2. S. Okudaira,
    3. M. Tanaka,
    4. K. Hama,
    5. D. Shida,
    6. J. Kitayama,
    7. T. Yamori,
    8. J. Aoki,
    9. T. Fujimaki,
    10. H. Arai
    . 2006. Autotaxin is overexpressed in glioblastoma multiforme and contributes to cell motility of glioblastoma by converting lysophosphatidylcholine to lysophosphatidic acid. J. Biol. Chem. 281: 17492–17500.
    OpenUrlAbstract/FREE Full Text
  38. ↵
    1. Ferry G.,
    2. N. Moulharat,
    3. J. P. Pradère,
    4. P. Desos,
    5. A. Try,
    6. A. Genton,
    7. A. Giganti,
    8. M. Beucher-Gaudin,
    9. M. Lonchampt,
    10. M. Bertrand,
    11. et al
    . 2008. S32826, a nanomolar inhibitor of autotaxin: discovery, synthesis and applications as a pharmacological tool. J. Pharmacol. Exp. Ther. 327: 809–819.
    OpenUrlAbstract/FREE Full Text
  39. ↵
    1. Ohta H.,
    2. K. Sato,
    3. N. Murata,
    4. A. Damirin,
    5. E. Malchinkhuu,
    6. J. Kon,
    7. T. Kimura,
    8. M. Tobo,
    9. Y. Yamazaki,
    10. T. Watanabe,
    11. et al
    . 2003. Ki16425, a subtype-selective antagonist for EDG-family lysophosphatidic acid receptors. Mol. Pharmacol. 64: 994–1005.
    OpenUrlAbstract/FREE Full Text
  40. ↵
    1. Heasley B. H.,
    2. R. Jarosz,
    3. K. R. Lynch,
    4. T. L. Macdonald
    . 2004. Initial structure-activity relationships of lysophosphatidic acid receptor antagonists: discovery of a high-affinity LPA1/LPA3 receptor antagonist. Bioorg. Med. Chem. Lett. 14: 2735–2740.
    OpenUrlCrossRefPubMed
  41. ↵
    1. Katakai T.,
    2. H. Suto,
    3. M. Sugai,
    4. H. Gonda,
    5. A. Togawa,
    6. S. Suematsu,
    7. Y. Ebisuno,
    8. K. Katagiri,
    9. T. Kinashi,
    10. A. Shimizu
    . 2008. Organizer-like reticular stromal cell layer common to adult secondary lymphoid organs. J. Immunol. 181: 6189–6200.
    OpenUrlAbstract/FREE Full Text
  42. ↵
    1. Ren X. D.,
    2. W. B. Kiosses,
    3. M. A. Schwartz
    . 1999. Regulation of the small GTP-binding protein Rho by cell adhesion and the cytoskeleton. EMBO J. 18: 578–585.
    OpenUrlAbstract/FREE Full Text
  43. ↵
    1. Shimonaka M.,
    2. K. Katagiri,
    3. T. Nakayama,
    4. N. Fujita,
    5. T. Tsuruo,
    6. O. Yoshie,
    7. T. Kinashi
    . 2003. Rap1 translates chemokine signals to integrin activation, cell polarization, and motility across vascular endothelium under flow. J. Cell Biol. 161: 417–427.
    OpenUrlAbstract/FREE Full Text
  44. ↵
    1. Malhotra D.,
    2. A. L. Fletcher,
    3. J. Astarita,
    4. V. Lukacs-Kornek,
    5. P. Tayalia,
    6. S. F. Gonzalez,
    7. K. G. Elpek,
    8. S. K. Chang,
    9. K. Knoblich,
    10. M. E. Hemler,
    11. et al
    . 2012. Transcriptional profiling of stroma from inflamed and resting lymph nodes defines immunological hallmarks. Nat. Immunol. 13: 499–510.
    OpenUrlCrossRefPubMed
  45. ↵
    1. Jansen S.,
    2. C. Stefan,
    3. J. W. Creemers,
    4. E. Waelkens,
    5. A. Van Eynde,
    6. W. Stalmans,
    7. M. Bollen
    . 2005. Proteolytic maturation and activation of autotaxin (NPP2), a secreted metastasis-enhancing lysophospholipase D. J. Cell Sci. 118: 3081–3089.
    OpenUrlAbstract/FREE Full Text
  46. ↵
    1. Koike S.,
    2. K. Keino-Masu,
    3. T. Ohto,
    4. M. Masu
    . 2006. The N-terminal hydrophobic sequence of autotaxin (ENPP2) functions as a signal peptide. Genes Cells 11: 133–142.
    OpenUrlCrossRefPubMed
  47. ↵
    1. Chun J.,
    2. T. Hla,
    3. K. R. Lynch,
    4. S. Spiegel,
    5. W. H. Moolenaar
    . 2010. International union of basic and clinical pharmacology. LXXVIII. Lysophospholipid receptor nomenclature. Pharmacol. Rev. 62: 579–587.
    OpenUrlAbstract/FREE Full Text
  48. ↵
    1. Lee C.-W.,
    2. R. Rivera,
    3. S. Gardell,
    4. A. E. Dubin,
    5. J. Chun
    . 2006. GPR92 as a new G12/13- and Gq-coupled lysophosphatidic acid receptor that increases cAMP, LPA5. J. Biol. Chem. 281: 23589–23597.
    OpenUrlAbstract/FREE Full Text
  49. ↵
    1. Aktories K.,
    2. C. Wilde,
    3. M. Vogelsgesang
    . 2004. Rho-modifying C3-like ADP-ribosyltransferases. Rev. Physiol. Biochem. Pharmacol. 152: 1–22.
    OpenUrlCrossRefPubMed
  50. ↵
    1. Wendland M.,
    2. S. Willenzon,
    3. J. Kocks,
    4. A. C. Davalos-Misslitz,
    5. S. I. Hammerschmidt,
    6. K. Schumann,
    7. E. Kremmer,
    8. M. Sixt,
    9. A. Hoffmeyer,
    10. O. Pabst,
    11. R. Förster
    . 2011. Lymph node T cell homeostasis relies on steady state homing of dendritic cells. Immunity 35: 945–957.
    OpenUrlCrossRefPubMed
  51. ↵
    1. Houben A. J. S.,
    2. X. M. R. van Wijk,
    3. L. A. van Meeteren,
    4. L. van Zeijl,
    5. E. M. A. van de Westerlo,
    6. J. Hausmann,
    7. A. Fish,
    8. A. Perrakis,
    9. T. H. van Kuppevelt,
    10. W. H. Moolenaar
    . 2013. The polybasic insertion in autotaxin α confers specific binding to heparin and cell surface heparan sulfate proteoglycans. J. Biol. Chem. 288: 510–519.
    OpenUrlAbstract/FREE Full Text
  52. ↵
    1. Nishimasu H.,
    2. S. Okudaira,
    3. K. Hama,
    4. E. Mihara,
    5. N. Dohmae,
    6. A. Inoue,
    7. R. Ishitani,
    8. J. Takagi,
    9. J. Aoki,
    10. O. Nureki
    . 2011. Crystal structure of autotaxin and insight into GPCR activation by lipid mediators. Nat. Struct. Mol. Biol. 18: 205–212.
    OpenUrlCrossRefPubMed
  53. ↵
    1. Moolenaar W. H.,
    2. A. Perrakis
    . 2011. Insights into autotaxin: how to produce and present a lipid mediator. Nat. Rev. Mol. Cell Biol. 12: 674–679.
    OpenUrlCrossRefPubMed
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