Lysophosphatidic acid receptors LPA₄ and LPA₆ differentially promote lymphocyte transmigration across high endothelial venules in lymph nodes

HEV LPA receptors differentially regulate lymphocyte recirculation

Abstract

Naive lymphocytes continuously migrate from the blood into lymph nodes (LNs) via high endothelial venules (HEVs). To extravasate from the HEVs, lymphocytes undergo multiple adhesion steps, including tethering, rolling, firm adhesion and transmigration. We previously showed that autotaxin (ATX), an enzyme that generates lysophosphatidic acid (LPA), is highly expressed in HEVs, and that the ATX/LPA axis plays an important role in the lymphocyte transmigration across HEVs. However, the detailed mechanism underlying this axis’s involvement in lymphocyte transmigration has remained ill-defined. Here, we show that two LPA receptors, LPA₄ and LPA₆, are selectively expressed on HEV endothelial cells (ECs) and that LPA₄ plays a major role in the lymphocyte transmigration across HEVs in mice. In the absence of LPA₄ expression, lymphocytes accumulated heavily within the HEV EC layer, compared to wild-type (WT) mice. This accumulation was also observed in the absence of LPA₆ expression, but it was less pronounced. Adoptive transfer experiments using WT lymphocytes revealed that the LPA₄ deficiency in ECs specifically compromised the lymphocyte transmigration process, whereas the effect of LPA₆ deficiency was not significant. These results indicate that the signals evoked in HEV ECs via the LPA₄ and LPA₆ differentially regulate lymphocyte extravasation from HEVs in the peripheral LNs.

Introduction

Throughout life, naive lymphocytes patrol the body to detect and eliminate invading pathogens as well as aberrant cells that may arise in the host. During this process, which is called lymphocyte recirculation, the naive lymphocytes selectively migrate into lymph nodes (LNs) and Peyer’s patches through a specific type of blood vessel, called the high endothelial venule (HEV). Within the HEVs, naive lymphocytes undergo a multistep adhesion cascade, which is initiated by rolling, followed by firm arrest and then transmigration. These steps enable the large-scale trafficking of blood-borne naive lymphocytes into the LNs. While the molecular requirements for rolling and adhesion are becoming clearer (1, 2), we still know relatively little about the molecular mechanism of lymphocyte extravasation (3).

Lysophosphatidic acid (LPA) is a pleiotropic lipid mediator that regulates a variety of biological responses, including cell adhesion, migration, proliferation, and survival, gap-junction closure and opening, and the production of growth factors and cytokines (4). LPA can be generated by at least two enzymes: autotaxin (ATX or ENPP2 [ectonucleotide pyrophosphatase/phosphodiesterase family member 2]), which hydrolyzes lysophosphatidylcholine (LPC) to LPA, and phospholipase A₁, which hydrolyzes phosphatidic acid to LPA. There are six known receptors for LPA, LPA₁-LPA₆, which are located on the cell surface (5, 6). LPA₁-LPA₃ are members of the endothelial differentiation gene (EDG) family, which also includes sphingosine-1-phosphate receptor 1 (S1P₁), a key regulator of lymphocyte egress from lymphoid tissues (7). LPA₄-LPA₆ are non-EDG family receptors that belong to the purinergic P2Y receptor family (8). These LPA receptors transmit signals through various G proteins, including Gα_i, Gα_12/13, Gα_q and Gα_s. LPA₁ and LPA₄ are known to be involved in blood vessel formation during development (9). LPA₆ shows high sequence homology with LPA₄; it is expressed in vascular endothelial cells (ECs), where it regulates EC contraction in a Gα_12/13-Rho-dependent manner (5).

Previously, others and we have reported that ATX is highly expressed in the HEV ECs and that it promotes lymphocyte transmigration across the HEVs by locally producing LPA, which in turn acts on HEV ECs (10–12). LPA also acts on lymphocytes to induce chemokinesis, cell polarization and transmigration across HEVs, although the responsible receptor(s) has remained unclear (12, 13).

In this study, we examined the mode of action of the ATX/LPA axis in lymphocyte transmigration across HEVs. We first found that HEV ECs express LPA₄ and LPA₆. Using knockout (KO) mice, we found that LPA₄ deficiency caused severe lymphocyte accumulation within the HEV EC layer, which delayed lymphocyte transmigration across this layer in vivo. In contrast, LPA₆ deficiency compromised this cell trafficking process to a much smaller extent. Taken together, these results indicate that LPA₄ and LPA₆ on HEV ECs are differentially involved in the LPA-dependent lymphocyte transmigration across the HEV wall in LNs.

Methods

Mice

C57BL/6 mice were purchased from Japan SLC. GFP transgenic mice (14) were kindly provided by Dr Masaru Okabe (Research Institute for Microbial Diseases, Osaka University). LPA₄ KO mice were generated as described previously (15). The LPA₆/Lpar6 KO mice (Accession No. CDB0977K: http://www.clst.riken.jp/arg/mutant%20mice%20list.html) were generated by three of us (S.I., K.B. and T.A), by crossing LPA₆ ^fl/fl mice and CAG-Cre mice (16), as described in Supplementary Data 1, available at International Immunology Online. Homologous recombinants were isolated, using the HK3 ES cell line established from the C57BL/6N strain (17). The LPA₆ KO mice were genotyped by genomic PCR. The primers were 5′-AAAAATCCGAAATGGCAAAGTAAA-3′ and 5′-GTGACCACATCTGAATAGCAAAGG-3′ for the wild-type (WT) allele, 5′-ACTTCCTGACTAGGGGAGGAGTAGA-3′ and 5′-GTGACCACATCTGAATAGCAAAGG-3′ for the floxed allele and 5′-TTCCGTAAACAACATCTCGGTTC-3′ and 5′-GTGACCACATCTGAATAGCAAAGG-3′ for the mutant allele (see Supplementary Data 1, available at International Immunology Online, for details) and yielded 303-bp, 445-bp and 462-bp products, respectively. All mice were housed at the Institute of Experimental Animal Sciences at Osaka University Medical School, and all animal experiments followed protocols approved by the Ethics Review Committee for Animal Experimentation of Osaka University Graduate School of Medicine.

Reagents and antibodies

Hybridomas for anti-peripheral node addressin (PNAd) mAb, MECA-79, the anti-mucosal vascular addressin cell adhesion molecule-1 (MAdCAM-1) mAbs, MECA-89 and MECA-367, and the ER-TR7 mAb were injected into nude mice i.p., and the antibodies were later purified from the ascites. Purified MECA-79, MECA-89 and ER-TR7 mAbs were labeled with the Alexa Fluor 594 Protein Labeling Kit (Life Technologies, Carlsbad, CA, USA). The MECA-367 and MECA-89 mAbs were biotinylated using the Sulfo-NHS-LC-biotin Reagent (Thermo Fisher Scientific, Waltham, MA, USA). Anti-ATX serum was generated in rabbits after several immunizations with GST-fused recombinant ATX (57S-116A); its specificity is shown in Supplementary Data 2, available at International Immunology Online. Mouse γ-globulins and FITC-anti-α-smooth muscle actin (SMA) mAb were purchased from Sigma-Aldrich (St Louis, MO, USA). Goat IgG, biotinylated anti-CD4 mAb (RM4-5) and allophycocyanin (APC)-anti-CD45 mAb (30-F11) were purchased from Chemicon (Temecula, CA, USA), BD Biosciences (San Jose, CA, USA) and eBioscience (San Diego, CA, USA), respectively. FITC-anti-B220 mAb (RA3-6B2) and purified anti-CD31 mAb (390) were purchased from Biolegend (San Diego, CA, USA). Purified anti-CD31 mAb was labeled with the Alexa Fluor 647 Protein Labeling Kit. Alexa Fluor 647-labeled goat anti-rabbit IgG, Hoechst 33342, lysine fixable FITC-conjugated dextran (MW 70kDa) and CellTracker™ Orange CMTMR (5-[and-6]-[{(4-chloromethyl)benzoyl}amino]tetramethylrhodamine) were all purchased from Life Technologies.

RT–PCR

HEV ECs were isolated as MECA-367⁺CD45⁻ cells from the mesenteric LNs (MLNs) using a FACSVantage cell sorter, and the total RNA was extracted from freshly isolated MECA-367⁺ HEV ECs using the RNAqueous-4PCR Kit (Ambion, Foster, CA, USA). The cDNA was synthesized using the Ovation System (Nugen Technologies, San Carlos, CA, USA). T cells, B cells and dendritic cells (DCs) were isolated from the spleen as, respectively, CD3⁺, B220⁺ and CD11c⁺ cells. Total RNA was extracted with Trizol (Life Technologies), and cDNA was synthesized with Superscript III (Life Technologies). The cDNA fragments of LPA receptors (Lpar1-Lpar6) were amplified by PCR using ExTaq (Takara, Shiga, Japan). The primer pairs are described in Supplementary Data 3, available at International Immunology Online.

In situ hybridization assay

In situ hybridization was performed as previously described (10). The MLNs were embedded in OCT compound (Sakura Finetek, Torrance, CA, USA), and 10-µm-thick serial frozen sections were cut. A 537-bp fragment from nucleotides 182–718 of the LPA₄ cDNA (GenBank accession no. NM_175271) or a 501-bp fragment from nucleotides 23–523 of the LPA₆ cDNA (GenBank accession no. NM_175116) was inserted into the pPCRII vector (Invitrogen). The plasmids were linearized by digestion with XhoI or SpeI, at sites flanked by the T7 and SP6 promoters, respectively. Digoxigenin (DIG)-labeled anti-sense and sense probes were generated by in vitro transcription using the DIG RNA Labeling Mix (Roche Diagnostics, Basel, Switzerland), according to the manufacturer’s instructions.

Measurement of body weight and the total cell number in each organ

The total body weight of male 8-week-old LPA₄-deficient mice, LPA₆-deficient mice and their littermates was measured. The total cell numbers in the spleen, MLNs, inguinal LNs (ILNs) and popliteal LNs (PLNs) were determined by flow cytometry (FACSVerse; BD Biosciences).

Conventional immunohistochemistry

ILNs and MLNs obtained from 8-week-old littermate mice were snap frozen in OCT compound and cut into 10-µm-thick frozen sections. The sections were fixed in methanol, blocked in 10% FCS/PBS containing mouse γ-globulins (20 µg/ml) and stained with FITC-anti-α-SMA, Alexa Fluor 594-anti-PNAd, Alexa Fluor 594-anti-MAdCAM-1, biotinylated-anti-MAdCAM-1, biotinylated-anti-CD4, FITC-anti-B220, APC-anti-CD45, Alexa Flour 594-ER-TR7 or Hoechst 33342 (2 µg/ml). Biotinylated antibody was detected by Alexa Fluor 405-streptavidin (2 µg/ml). ATX expression was detected by rabbit anti-ATX serum (1:2000) and Alexa Fluor 647-goat anti-rabbit antibody (2.5 µg/ml) after blocking with goat IgG (20 µg/ml). Immunofluorescence confocal microscopy was performed with an FV1000-D confocal laser scanning microscope (Olympus, Tokyo, Japan) and an LSM 710 confocal laser microscope (Zeiss, Oberkochen, Germany).

Assessment of leakage from HEVs

Mice were given an injection of Alexa Fluor 594-ER-TR7 (5 µg/foot) into the hindfoot. Six hours later, the mice were given an i.v. injection of Alexa Fluor 647-anti-CD31 (10 µg/mouse). Seven minutes later, an i.v. injection with fixable FITC-dextran (MW 70kDa, 1mg/mouse) was given. Fifteen minutes after the final injection, the mice were killed under isoflurane anesthesia, dissected and the ipsilateral PLNs were collected in 4% paraformaldehyde (PFA). The LNs were washed with PBS and incubated with 30% sucrose/PBS for 30min. The immunofluorescent signals were observed with an FV1000-D confocal laser scanning microscope.

Quantification of lymphocyte accumulation within the HEV EC layer

Quantitative analysis of the lymphocyte accumulation within the HEV EC layer was performed using LN sections as follows. The HEV EC layer was identified by staining with fluoresceinated PNAd or MAdCAM-1 mAbs, and the area of the HEV EC layer was determined using ImageJ. Lymphocytes located within the HEV EC layer were identified by CD45 staining, and lymphocyte accumulation within the HEV EC layer was expressed as the number of CD45⁺ cells per unit HEV EC area (1000 µm²).

Transmission electron microscopy

The ILNs of 8-week-old WT, LPA₄ KO and LPA₆ KO mice were collected and processed as described previously (18).

Trafficking assay using flow cytometry

Mice were i.v. injected with GFP⁺ splenocytes (2×10⁷ cells/mouse). Thirty minutes after adoptive transfer, the mice were anesthetized and transcardially perfused with PBS. The spleens, MLNs and peripheral LNs were collected. The proportion of donor GFP⁺ cells in these tissues was analyzed by flow cytometry.

Trafficking assay using whole-mount analysis

Mice were i.v. injected with GFP⁺ splenocytes (2×10⁷ cells/mouse) and an Alexa Fluor 647-conjugated anti-CD31 mAb (10 µg/mouse). Sixty minutes after the injection, the mice were anesthetized and transcardially perfused with PBS and 4% PFA in phosphate buffer. The mice were dissected, and the LNs were harvested. The LNs were incubated with increasing concentrations (10, 20 and 30%) of sucrose. The immunofluorescent signals were observed with an FV1000-D confocal laser scanning microscope (Olympus), and the acquired images were analyzed with the IMARIS 7.4.2. software (Bitplane, Zurich, Switzerland). Briefly, the images were processed using the surface rendering and contour modes.

Statistics

Data are presented as the mean ± SD. Raw data were analyzed using the GraphPad Prism 6 software (GraphPad Software, San Diego, CA, USA). The Mann–Whitney U-test was used for comparisons of body weight, the total number of accumulated lymphocytes and lymphocyte extravasation. The Kruskal–Wallis test was used for comparison of the lymphocyte accumulation in three groups before the Mann–Whitney U-test. The Student’s t-test was used for the comparison of lymphocyte trafficking by flow cytometry.

Results

HEV ECs express two LPA receptors, LPA₄ and LPA₆

While we previously showed that the ATX/LPA axis regulates lymphocyte transmigration across the basal lamina of HEVs, primarily by acting on HEV ECs (11), the LPA receptor(s) involved in this event remained unknown. Therefore, we first examined the LPA receptor expression in HEV ECs and leukocytes by RT–PCR. As shown in Fig. 1, HEV ECs, which were rigorously purified by cell sorting, expressed LPA₄ and LPA₆, whereas lymphocytes expressed LPA₂, LPA₅ and LPA₆, and DCs expressed all of the LPA receptors, at the mRNA level. The LPA₄ and LPA₆ expression on HEV ECs was further confirmed by in situ hybridization of LN sections (Fig. 1B). Although we tried to examine the protein expression as well, the commercially available anti-LPA₄ and anti-LPA₆ antibodies we used gave substantial signals not only in WT mice but also in LPA₄ KO and LPA₆ KO mice, indicating that they were not specific enough for our purpose. However, as described later, mice deficient in the LPA ₄ or LPA ₆ gene showed distinct HEV phenotypes compared with WT mice, indicating that the LPA₄ and LPA₆ proteins are indeed functionally expressed in HEV ECs. Although we previously reported that LPA₁ is expressed in HEV ECs (10), that observation was not confirmed in the present study, possibly because of the more rigorous isolation methods used here. These results demonstrated that HEV ECs preferentially express two LPA receptors: LPA₄ and LPA₆.

Fig. 1.

LPA₄ and LPA₆ are expressed in HEV ECs. (A) HEV ECs were sorted as MECA-367⁺CD45⁻ cells from the MLNs. The total RNA was extracted from T cells, B cells, DCs and HEV ECs, and RT–PCR was performed. (B) LPA₄ and LPA₆ expression in MLNs was examined by in situ hybridization. Scale bars indicate 20 µm. HEVs are indicated by yellow lines. Data shown are representative of two independent experiments.

Neither LPA₄ deficiency nor LPA₆ deficiency grossly affects the LN architecture

To examine the biological significance of the LPA₄ and LPA₆ receptors in the immune system, we next examined the LNs and spleen of WT, LPA₄ KO and LPA₆ KO mice. As shown in Fig. 2(A), neither the LPA₄ nor the LPA₆ deficiency caused a significant difference in the total cell number in the spleen or MLNs. Of the peripheral LNs, while the ILNs and PLNs tended to be larger in the LPA₄ KO mice than in their WT littermates, there was considerable heterogeneity, and no statistically significant difference in the total cell numbers was found. The peripheral LNs of LPA₆ KO mice were comparable in size to those of the WT littermates. In addition, no difference in the total body weight was observed among these groups (Fig. 2A). As shown in Fig. 2(B), immunohistological analysis indicated that neither the LPA₄ nor the LPA₆ deficiency compromised the formation of HEVs, B-cell follicles or T-cell areas in the LNs. FACS analysis of cells obtained from the LNs showed no abnormalities in CD4, CD8 or B220 expression (data not shown).

Fig. 2.

Neither LPA₄ deficiency nor LPA₆ deficiency grossly affects the LN architecture or ATX production. (A) Total cell numbers of the spleen, MLN, ILN and PLN and total body weight of WT, LPA₄ KO and LPA₆ KO littermates. The cell numbers are shown for the spleen (×10⁷ cells), MLN (×10⁶ cells), ILN (×10⁶ cells) and PLN (×10⁵ cells). Points indicate data from individual mice, and bars show the arithmetic means ± SD for three to four mice from each group. Mann–Whitney U-test, N.S.: not significant. (B) Sections of peripheral LNs were stained for CD4 (blue), B220 (green), ER-TR7 (red), α-SMA (green) and PNAd (red). ATX expression was identified by rabbit anti-ATX serum and secondary antibodies (white). Scale bars indicate 50 µm. (C) Blood vessel distribution pattern in the PLN visualized by i.v. injection of the Alexa Fluor 647-anti-CD31 antibody (10 µg/mouse, white). Scale bars indicate 100 µm. (D) HEVs of the PLN visualized by s.c. injection of Alexa Fluor 594-ER-TR7 (5 µg, red) and i.v. injection of anti-CD31 antibody (10 µg/mouse, blue) and FITC-dextran (70kDa, 1mg/mouse, green). Scale bars indicate 100 µm. Similar experiments were performed at least three times (LPA₄ KO mice) and once (LPA₆ KO mice) using 2-photon intravital microscopy.

Injecting a fluoresceinated anti-CD31 mAb intravenously, which illuminated all of the vascular trees in the LNs, revealed no obvious differences in the overall vascular distribution pattern in the LPA₄ KO or LPA₆ KO mice compared to WT mice (Fig. 2C). Intravenously injected FITC-dextran (70kDa) showed some leakage from HEVs, but most of it was retained within the vasculature at comparable levels in the LPA₄ KO, LPA₆ KO and WT mice (Fig. 2C). The expression of the LPA-producing enzyme, ATX, appeared unaltered in the HEVs and fibroblastic reticular cells of LPA₄ KO and LPA₆ KO mice (Fig. 2B), indicating that the local LPA production was unimpaired. These results indicated that neither the LPA₄ deficiency nor the LPA₆ deficiency grossly affected the tissue architecture or ATX production in LNs.

LPA₄ deficiency and LPA₆ deficiency differentially induce lymphocyte accumulation within the HEV EC layer

Next, to analyze the HEVs of LPA₄- and LPA₆-deficient mice, we examined cross-sections of the PNAd⁺ HEVs in the ILNs and MLNs of WT, LPA₄ KO and LPA₆ KO mice (Fig. 3A). The HEV ECs, pericytes and lymphocytes were observed by staining for PNAd, α-SMA and CD45, respectively. The PNAd staining appeared sparser and less intense in substantial proportions of the HEVs of the LPA₄ KO and LPA₆ KO mice compared to WT mice (upper panels), which was apparently due to a greater abundance of lymphocytes within the HEV EC layer (lower panels).

Fig. 3.

LPA₄ deficiency and LPA₆ deficiency differentially induce lymphocyte accumulation within the HEV EC layer. (A) Sections of ILNs obtained from WT, LPA₄ KO or LPA₆ KO mice were stained for α-SMA (green), PNAd (red), CD45 (white) and nuclei (blue) and analyzed by confocal microscopy (top panel). Yellow dotted lines and asterisks indicate the HEV EC layer and CD45⁺ lymphocytes located within the EC layer, respectively (bottom panel). Scale bars indicate 20 µm. (B) Lymphocytes located within the HEV EC layer were enumerated in the ILN obtained from female LPA₄ ^+/+, LPA₄ ^+/− and LPA₄ ^−/− mice and from male LPA₄ ^+/y, LPA₄ ^−/y, LPA₆ ^+/+ and LPA₆ ^−/− mice. More than 10 photographs of the ILN section from each mouse were obtained, and three to five mice from each group were analyzed. Kruskal–Wallis test and Mann–Whitney U-test, ***P < 0.0001. N.S.: not significant.

To quantify the lymphocyte accumulation, we enumerated the number of lymphocytes within the HEV EC layer per constant HEV area (1000 µm²) in female mice; the gene dosage effect can be examined in female littermates, because the LPA ₄ gene is located on the X chromosome. As shown in Fig. 3(B), the targeted disruption of LPA₄ increased the lymphocyte accumulation within the HEV EC layer of the ILN in a manner dependent on the extent of gene deletion (LPA₄ ^−/− > LPA₄ ^+/− > LPA₄ ^+/+) (left panel). In male mice, the LPA₄ deficiency (LPA₄ ^−/y) also significantly increased the lymphocyte accumulation within the HEV EC layer compared with littermate LPA₄ ^+/y mice (middle panel). In contrast, LPA₆ deletion caused only a minimal (statistically non-significant) increase in lymphocyte accumulation within the HEV EC layer (right panel). These results suggested that the LPA₄- and LPA₆-mediated signals are differentially involved in the regulation of lymphocyte transmigration across the HEV wall in peripheral LNs.

Regarding the involvement of LPA₄ signaling, very similar results were observed in the PNAd⁺ HEV ECs of MLNs (Supplementary Data 4, available at International Immunology Online), in which the HEV ECs express either PNAd or MAdCAM-1 as a tissue-specific EC adhesion molecule (19). In the MLNs of both male and female LPA₄ KO mice, elevated lymphocyte accumulation was observed within the EC layer of the PNAd⁺ HEV ECs, albeit less prominently than in the PNAd⁺ HEV ECs of the peripheral LNs (Supplementary Data 4, available at International Immunology Online). These results together indicated that lymphocyte transmigration across HEVs depends at least partly on LPA₄ signaling in both peripheral LNs and MLNs.

Accumulated lymphocytes in the HEV EC layer are located between the ECs and the underlying basal lamina

We next compared the HEVs of LPA₄ KO, LPA₆ KO and WT mice by transmission electron microscopy. As shown in Fig. 4(A), compared with WT mice, a substantial proportion of the HEVs in both the LPA₄ KO and LPA₆ KO mice contained more lymphocytes within the EC layer, which occasionally caused luminal narrowing. Regarding the precise location of the lymphocytes within the EC layer, it could clearly be seen in the LPA₄-deficient mice that, although some lymphocytes were present between the ECs, most of them were nested between the ECs and the underlying basal lamina (Fig. 4B).

Fig. 4.

Lymphocytes are more abundant in the HEV pockets of LPA₄- and LPA₆-deficient mice than in WT mice. (A) Transmission electron micrographs of HEVs in the ILNs of WT, LPA₄ KO and LPA₆ KO mice. Color was added (lower panel of each image) to show that the lymphocytes (yellow) were located in the HEV EC layer including the perivenular channels. The luminal surface of HEV ECs is indicated by red lines. Scale bars, 5 µm. (B) Transmission electron micrographs of HEVs in the ILNs of WT, LPA₄ KO and LPA₆ KO mice. Color was added (lower panels of each image) to show the nuclei of the HEV ECs (green), the discontinuous basal lamina (light green), lymphocytes in the HEV EC layer (blue) and lymphocytes in the vicinity of the basal lamina (red). Scale bars, 2 µm. (C) HEV pockets containing >3 lymphocytes were selected, and the number of lymphocytes per pocket was enumerated in LPA₄ KO, LPA₆ KO and WT littermates. Mann–Whitney U-test, **P < 0.01, ***P < 0.0001. More than 10 HEVs in each group were statistically analyzed.

This finding was consistent with an observation reported by Mionnet et al. (20), that the HEV ECs create pockets where lymphocytes reside for several minutes before they extravasate. These pockets were readily observed in all of the mice we examined, with different degrees of lymphocyte accumulation. Whereas most of the pockets housed up to four to five lymphocytes in WT mice, those in LPA₄ KO and LPA₆ KO mice held statistically greater numbers of lymphocytes (Fig. 4C), with >10 lymphocytes being occasionally present in a single pocket in the LPA₄ KO and LPA₆ KO mice. When this occurred, the ECs overlying such a pocket bulged out into the vascular lumen, resulting in luminal narrowing (Fig. 4A). Given that the pockets in the HEV EC layer are the areas where lymphocytes are retained before they enter the LN parenchyma (20), these data raise the possibility that LPA₄ and LPA₆ are both involved in driving lymphocytes from the pockets into the LN parenchyma.

The absence of LPA₄ signaling delays lymphocyte extravasation across HEVs

To further evaluate the importance of LPA₄- and LPA₆-mediated signaling in the lymphocyte transmigration across HEVs, we adoptively transferred WT donor cells from GFP mice into WT, LPA₄ KO or LPA₆ KO mice and quantified the lymphocyte migration from the blood into the LNs and spleen, by flow cytometry. As shown in Fig. 5(A), LPA₄ deficiency mildly reduced the lymphocyte entry into the peripheral LNs but not into the spleen, whereas LPA₆ deficiency did not affect the lymphocyte entry into any of the lymphoid tissues examined at appreciable levels. Phenotypic analysis of migrated cells indicated that T-cell migration is more strongly affected than B-cell migration in the absence of LPA₄ signaling (Supplementary Data 5, available at International Immunology Online). When LPA₆-deficient cells were transferred to LPA₆-deficient mice, they showed uncompromised migration to the LNs and spleen compared with WT cells (Supplementary Data 6, available at International Immunology Online), indicating that LPA₆ in lymphocytes is dispensable for lymphocyte migration in vivo.

Fig. 5.

Lymphocytes accumulate more heavily in the HEV EC layer in LPA₄-deficient mice than in LPA₆-deficient mice. (A) GFP⁺ lymphocytes (2×10⁷ cells/mouse) were injected into the tail vein of WT, LPA₄ KO and LPA₆ KO mice. Thirty minutes after cell transfer, spleens and peripheral LNs were harvested. The cells that had migrated into these tissues were then quantified by flow cytometry. The number of donor cells found in the spleens (×10⁵ cells) and peripheral LNs (×10³ cells) is shown. Results are shown as the mean ± SD of three mice from each group. Student’s t-test, N.S.: not significant. The data in the figure are representative of two separate experiments. (B) GFP⁺ lymphocytes (2×10⁷ cells/mouse) and Alexa Fluor 647-conjugated anti-CD31 mAb (10 µg/mouse) were co-injected into the tail vein of WT, LPA₄ KO and LPA₆ KO mice. Sixty minutes after cell transfer, the mice were perfused with PBS and 4% PFA. The LNs were collected and subjected to confocal microscopic analysis (upper panel; donor cells appear green and CD31⁺ blood vessels appear red). Subsequently, donor cells within the HEV EC layer and those located within 20 µm of the outside of HEVs were marked in green and blue, respectively, using the IMARIS software (middle panel). CD31⁺ blood vessels and donor cells in the xy, xz and yz planes (lowest panel). Scale bars indicate 30 µm. The data shown are representative of two separate experiments. (C) Density of transferred cells around HEVs, and the surface area and volume of HEVs in LPA₄ KO, LPA₆ KO and WT mice. Lymphocyte density = the number of donor cells within the HEV EC layer/the number of recently extravasated cells (those located within 20 µm from the outside of an HEV). The HEV surface area and volume per 100 µm of HEV were calculated by the IMARIS software. Mann–Whitney U-test, *P < 0.05. N.S.: not significant. The data in the figure are representative of two separate experiments, with two to three mice per experiment.

We next examined whether LPA₄ or LPA₆ deficiency compromised the extravasation of adoptively transferred lymphocytes from HEVs, by the whole-mount analysis of LNs. To this end, we focused on the particular segment of HEVs, termed orders IV-V, where the lymphocyte rolling and subsequent steps preferentially occur (21). We first identified the adoptively transferred lymphocytes and the order IV-V venules in whole-mount LN preparations (Fig. 5B, top panel) and differentially marked the extravasated donor cells located outside but within 20 µm of the HEV basal lamina and those still located within the HEV EC layer, using the IMARIS software (Fig. 5B, middle and lower panels). We then determined the ratio of the cells resident within the HEV EC layer to those recently extravasated from HEVs. This analysis revealed that the LPA₄ deficiency significantly increased the proportion of lymphocytes located within the HEV EC layer (Fig. 5C), in agreement with the results obtained by conventional immunohistological and electron microscopic analyses (Figs 3 and 4), whereas the samples with LPA₆ deficiency did not show a statistical difference from WT.

In addition, reflecting the lymphocyte accumulation within the EC layer, the HEVs of the LPA₄ KO mice had a wider surface area and larger volume compared to those of WT mice, whereas no statistical difference was observed between these data for LPA₆ KO and WT mice (Fig. 5C), although HEVs with a narrowed lumen, apparently due to lymphocyte accumulation within the HEV cell layer, were occasionally found in these mice. In conjunction with the electron microscopy data presented in the previous section, these results indicated that the LPA₄ and LPA₆ receptors play differential roles in the regulation of lymphocyte transmigration across the HEV EC layer, with LPA₄ signaling playing a more influential role than LPA₆ signaling in the HEV ECs.

Discussion

In the present study, we showed that expression of the LPA receptors LPA₄ and LPA₆ on HEV ECs is required for the effective transport of lymphocytes from the HEV EC layer to the LN parenchymal compartment. Without the signal from either of these receptors, lymphocytes were more frequently trapped in the EC layer, where they accumulated in EC pockets and often narrowed the HEV lumen. These results provide new details about the process of lymphocyte transmigration from HEVs by identifying the LPA₄ and LPA₆ receptors as key receptors in this process.

Our whole-mount LN analysis, electron microscopy studies and lymphocyte transfer studies all showed that LPA₄ signaling plays a larger role than LPA₆ signaling in lymphocyte migration through the HEV EC layer. Although we hoped to determine whether this observation could be explained by a difference in their protein expression levels, immunohistochemical analysis failed to produce meaningful results, due to the non-specific reactivity of the commercially available anti-LPA₄ and anti-LPA₆ antibodies we used. Another possible reason for the different effects of LPA₄ versus LPA₆ is a difference in their ligand-binding ability. Yanagida et al. (5) showed that LPA₆ has a much lower affinity for LPA compared with other LPA receptors including LPA₄. One can thus envision a situation in which the amount of LPA available in the HEV EC layer determines whether LPA₄ is preferentially stimulated or both receptors are simultaneously stimulated. Not mutually exclusive with these possibilities, it is also possible that LPA₄ and LPA₆ invoke qualitatively different signals in HEV ECs. Previous reports show that LPA₄ activates Gα_12/13- and Rho-mediated signaling in neuronal cells (22, 23). LPA₄ signaling also induces calcium ion mobilization by activating Gα_q and Gα_i, and intracellular cyclic AMP accumulation by activating Gα_s, which makes this receptor unique among LPA receptors (22). On the other hand, LPA₆ is coupled with Gα_12/13 proteins (5), and its activation leads to increases in intracellular calcium ions and ERK1/2 phosphorylation coupled with Gα_i and Gα_12/13 in intestinal epithelial cells (24). While it remains unclear whether LPA₄ and LPA₆ induce different signals in HEV ECs, the development of specific antagonists for LPA₄ and LPA₆ may help resolve this issue.

The LPA₄/LPA₆ receptor requirement for lymphocyte transmigration across HEVs was not absolute, because the HEV ECs deficient in LPA₄ or LPA₆ also allowed lymphocytes to leave the EC compartment and enter the LN parenchyma, albeit to lesser extents compared to those of WT mice. To examine whether LPA₄ and LPA₆ compensate for each other, or if both LPA₄ and LPA₆ are simultaneously required for efficient lymphocyte transmigration across HEVs, we are in the process of generating conventional and conditional LPA₄/LPA₆ double-KO mice.

We previously reported that LPA₁ is expressed in HEV ECs (10), but we did not reproduce this observation in the present study. This discrepancy could be at least partly explained by the fact that in this study, we carefully sorted the HEV ECs to exclude pericytes and smooth muscle cells, which are known to express LPA₁, from the EC preparation.

Upon leaving the HEV luminal surface, naive lymphocytes pass through an endothelial barrier and form pockets within the EC layer (20), where they are transiently retained before being released into the LN parenchyma. In LPA₄- or LPA₆-deficient mice, these pockets contained greater numbers of lymphocytes in substantial proportions of the HEVs compared to WT, although the increase was less obvious in the LPA₆-deficient animals. On the basis of the results of our whole-mount analysis of LNs, we propose that this accumulation was due to a defect in lymphocyte exit from the pockets to the LN parenchyma, and not to an increase in lymphocyte ingress from the HEV luminal surface. One possibility is that LPA₄/LPA₆ signaling promotes the exit process by enhancing the motility of HEV ECs and/or by impeding the strong adhesive interactions between HEV ECs and lymphocytes, allowing lymphocytes to be released from the EC layer more easily. We had hoped to study the in vitro abilities of HEV ECs to support lymphocyte ingress into and egress from the HEV EC layer in the presence or absence of LPA₄ or LPA₆ signaling. However, we have been unable to obtain HEV ECs at sufficient purity for this experiment from mice deficient in LPA₄ or LPA₆, although we could obtain them from WT animals successfully. It could well be that disruption of the LPA₄ or LPA₆ gene has affected the physiology of HEV ECs such that they could not be isolated successfully with the conventional method that we have been using. Future studies focusing on the details of LPA₄/LPA₆ signaling in HEV ECs are still needed.

Despite the slow lymphocyte egress from the EC compartment into the parenchymal compartment, only a minor reduction was observed in lymphocyte entry into the peripheral LNs in the LPA₄-deficient mice, using a conventional lymphocyte trafficking assay. Although the reason for this observation remains unclear, one possibility is the existence of a homeostatic mechanism that regulates overall lymphocyte trafficking efficiency by sensing the trapping of egress-incompetent lymphocytes in the HEV endothelial pockets.

Collectively, these results indicate that LPA₄ and LPA₆ are differentially involved in the LPA-dependent lymphocyte transmigration across the HEV wall in LNs, by signaling through the HEV ECs. As shown previously by our group (11), HEV ECs secrete ATX, which locally converts LPC to LPA; the LPA then acts on HEV ECs in LPA₄- and LPA₆-dependent manners, with the LPA/LPA₄ axis playing a major role. LPA₄ signaling in HEV ECs is thus an interesting target for immunomodulation.

Supplementary data

Supplementary data are available at International Immunology Online.

Funding

JSPS Kakenhi (Grant Numbers 21390151, 22021027, 24111005); Grant-in-Aid for JSPS Fellows (Number 25 9108).