ATX-LPA1 axis contributes to proliferation of chondrocytes by regulating fibronectin assembly leading to proper cartilage formation

Nishioka, Tatsuji; Arima, Naoaki; Kano, Kuniyuki; Hama, Kotaro; Itai, Eriko; Yukiura, Hiroshi; Kise, Ryoji; Inoue, Asuka; Kim, Seok-Hyung; Solnica-Krezel, Lilianna; Moolenaar, Wouter H.; Chun, Jerold; Aoki, Junken

doi:10.1038/srep23433

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Published: 23 March 2016

ATX-LPA₁ axis contributes to proliferation of chondrocytes by regulating fibronectin assembly leading to proper cartilage formation

Tatsuji Nishioka¹^na1,
Naoaki Arima¹^na1,
Kuniyuki Kano¹^na1,
Kotaro Hama¹^na1,
Eriko Itai¹^na1,
Hiroshi Yukiura¹^na1,
Ryoji Kise¹^na1,
Asuka Inoue^1,2^na1,
Seok-Hyung Kim³^na1,
Lilianna Solnica-Krezel⁴^na1,
Wouter H. Moolenaar⁵^na1,
Jerold Chun⁶^na1 &
…
Junken Aoki^1,7^na1

Scientific Reports volume 6, Article number: 23433 (2016) Cite this article

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Abstract

The lipid mediator lysophosphatidic acid (LPA) signals via six distinct G protein-coupled receptors to mediate both unique and overlapping biological effects, including cell migration, proliferation and survival. LPA is produced extracellularly by autotaxin (ATX), a secreted lysophospholipase D, from lysophosphatidylcholine. ATX-LPA receptor signaling is essential for normal development and implicated in various (patho)physiological processes, but underlying mechanisms remain incompletely understood. Through gene targeting approaches in zebrafish and mice, we show here that loss of ATX-LPA₁ signaling leads to disorganization of chondrocytes, causing severe defects in cartilage formation. Mechanistically, ATX-LPA₁ signaling acts by promoting S-phase entry and cell proliferation of chondrocytes both in vitro and in vivo, at least in part through β1-integrin translocation leading to fibronectin assembly and further extracellular matrix deposition; this in turn promotes chondrocyte-matrix adhesion and cell proliferation. Thus, the ATX-LPA₁ axis is a key regulator of cartilage formation.

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Introduction

Lysophosphatidic acid (LPA) is produced in the extracellular milieu such as in plasma, mainly by autotaxin (ATX) and acts on at least six receptors that are specific to LPA (LPA_1–6) to exert its functions^1,2. Historically, LPA was identified as a growth factor in serum. Indeed, LPA has been shown to stimulate the proliferation of many cell types including fibroblasts and cancer cells^3,4. Especially, LPA was shown to promote cell cycle entry, i.e., promote the G₀/G₁- to S-phase transition⁴, although different cell types can respond in other ways such as with cycle exit for neural progenitor cells⁵. The molecular mechanisms underlying LPA-induced cell cycle progression as well as its in vivo relevance remain unclear. In constitutive LPA receptor knockout (KO) mice, only LPA₁ KO mice showed obvious physical defects, including retardation of physical growth, prominent craniofacial abnormalities (including a shorter snout) and shorter limbs^6,7. ATX is a secreted lysophospholipase D (lysoPLD) which catalyzes a reaction to produce LPA from lysophosphaditylcholine (LPC)⁸. ATX KO mice are embryonic lethal around E9.5–10.5 because of vascular defects⁹. ATX was originally identified as a cell motility-stimulating factor secreted by human melanoma cells^10,11. Because enhanced expression of ATX has been demonstrated in various tumor tissues¹², ATX may promote proliferation and migration of cancer cells through LPA production. ATX is also expressed in various normal tissues and is present at high concentration in various biological fluids. Thus, ATX has an important role in cell proliferation not only in cancer cells but also in various normal cell types. Here, we show that ATX-LPA-LPA₁ signaling promotes the G₀/G₁-phase to S-phase transition in chondrocytes by enhancing integrin-dependent fibronectin assembly. These changes contribute to the normal development of cartilage tissues, based on analyses of both fish and mice.

Results

Loss of ATX-LPA₁ signaling results in dyschondroplasia in zebrafish

LPA-related genes are highly conserved in vertebrates. In zebrafish and mice, the amino acid sequences of ATX and LPA₁ are 67 and 85% identical, respectively. The genes for ATX and the six LPA receptors in vertebrates are completely conserved in zebrafish^13,14. We employed TILLING¹⁵ and identified a zebrafish lpa₁ mutant with a vu374 mutation at G309 that results in a premature stop codon (Fig. S1a) in the first extracellular loop of LPA₁. Homozygous lpa₁ mutants are able to reach adulthood and are fertile but display a craniofacial malformation: a round-shaped cephalic region that is also phenocopied in lpa₁ mutant mice⁶ (Fig. 1a).

A search for other mutant and morphant zebrafish with similar phenotypes identified several cases, e.g. the foxe1 mutant. In most of these cases, cartilage formation was impaired^16,17. In addition, lpa₁ mRNA was highly expressed at 72 and 96 hour post fertilization (hpf) in the zebrafish embryos and the expression pattern overlapped with that of sox9a mRNA (Fig. 1c), a marker of chondrocytes, which shows that chondrocytes express lpa₁ in the cartilage. Staining the cartilage of the wild type and lpa₁ mutant with alcian blue revealed that the mutant embryos had disorganized jaw cartilage at both 96 (Fig. 1a) and 120 hpf (data not shown). Both Meckel’s and ceratohyal cartilages (Fig. S1b), the two main cartilages of the lower jaw, were deformed. The lengths of Meckel’s and ceratohyal cartilages were shorter in the lpa₁ mutants (Fig. 1a and Fig. S1c). Similar abnormalities were observed when LPA₁ was down-regulated either by injection of a morpholino antisense oligonucleotide (MO) against LPA₁¹³ or by treatment with an LPA₁ antagonist, Ki16425 (Fig. 1b and Fig. S1d–f), which is known to be active against zebrafish LPA₁¹⁴.

ATX is an LPA-producing enzyme that was found to be expressed in the cartilage (Fig. 1c). Knockdown of ATX with a high dose of ATX MO1 (2.5 ng) was previously shown to induce severe vascular defects¹⁴. When a low dose of ATX MO1 (0.3 ng) or ATX MO2 (3.2 ng) was injected, most of the embryos survived at 120 hpf but had rounded-shaped heads (Fig. 1b and Fig. S1e), similar to the heads of LPA₁ mutant embryos. ATX morphant embryos also displayed impaired cartilage formation as illustrated by the loss of gill cartilage, i.e. deformation of Meckel’s and ceratohyal cartilages (Fig. 1b). We conclude that the loss of LPA₁ signaling results in dyschondroplasia in zebrafish embryos and that ATX is the main LPA-producing enzyme in cartilage tissues.

To determine how loss of ATX-LPA₁ signaling affects the behavior of chondrocytes in cartilage tissues, we employed col2:EGFP transgenic zebrafish, which expressed EGFP protein specifically in chondrocytes under the control of the col2a1a promoter¹⁸ (Fig. 1d). At 120 hpf, chondrocytes in both Meckel’s and ceratohyal cartilages maintained their intercalated and stacked organization in control embryos (Fig. 1d). In contrast, uneven sized and irregularly aligned chondrocytes were observed in LPA₁ and ATX MO-injected embryos (Fig. 1e) as well as LPA₁ antagonist (Ki16425)-treated embryos (data not shown).

The cartilage elements of the jaw are largely derived from cranial neural crest cells (CNCCs) that arise from dorsal and lateral regions of the neural ectoderm at 12 hpf and migrate into the area of the pharyngeal arches at 24–48 hpf, where the cells differentiate into chondrocytes to form the jaw cartilage at 72 hpf^17,19. The expression patterns of slug and sox10, which are markers of CNCCs, were normal in both LPA₁ and ATX morphant embryos (Fig. S1g) and the expression pattern of sox9a was also normal (data not shown). Thus, the migration of CNCCs and the maturation and differentiation of chondrocytes from CNCCs appeared to be unaffected by the loss of ATX-LPA₁ signaling. It should be noted that bone tissues are not formed in zebrafish embryos until 120 hpf¹⁹, suggesting again that ATX-LPA₁ signaling has a critical role in chondrogenesis.

Loss of ATX-LPA₁ signaling results in dyschondroplasia in mice

We next examined the role of ATX-LPA₁ signaling in cartilage formation in mice. LPA₁ KO mice showed reduced anteroposterior growth of skull bone and shorter femur, tibia and humerus (Fig. 2a–e). We focused on cartilage tissues in the cranial base (Fig. S2a) because the base is important for anteroposterior growth of skull bone. In fact, many mutant mice with defects in the base showed abnormal formation of skull bones like LPA₁ KO mice^20,21,22. We found that intersphenoid synchondrosis (the cartilage that links bones at the cranial base) ossified earlier in LPA₁ KO mice at 3 weeks of age (Fig. S2b). At the cellular level, alignment of chondrocytes in the cartilage tissue was disturbed (Fig. 2f) and the number of cells was also significantly lower in LPA₁ KO mice (Fig. 2g). Similar mislocalization of the chondrocytes was observed in other cartilage tissues such in the costa and femur (Fig. S2c).

Since global ATX KO mice are embryonic lethal because of impaired vascular formation⁹, we set out to produce conditional ATX KO mice. We produced mice with various combinations of ATX wild type, ATX-flox and null alleles, i.e., ATX^+/+, ATX^+/flox, ATX^flox/flox, ATX^+/− and ATX^flox/− mice. We found that one flox allele insertion significantly decreased the serum ATX activity about 15% (Fig. S2d). Interestingly, mice with both ATX-flox and ATX-null alleles (ATX^flox/− mice) showed phenotypes similar to those of LPA₁ KO mice. Because ATX^+/− mice as well as mice with other genotypes did not show obvious abnormality at all, it was speculated that the significant difference between ATX^+/− and ATX^flox/− (~50% minus ~35%, i.e., ~15%) was important for normal cartilage formation. Alternatively, it is possible that the insertion of flox allele impairs the transcription or stability of ATX pre-mRNA in a specific cell type, which affect the normal cartilage formation. From P0 to the adult stage, the ATX^flox/− mice displayed obvious abnormalities in craniofacial morphology and anteroposterior growth of the skull bone and shorter femur, tibia and humerus (Fig. 2a–e). The intersphenoid synchondrosis ossified significantly earlier as well in the ATX^flox/− mice (Fig. S2b). In addition, at the cellular level, the alignment of chondrocytes was significantly disturbed in ATX^flox/− mice (Fig. 2f).

As was observed in zebrafish, loss of ATX-LPA₁ signaling did not affect the differentiation of chondrocytes, since LPA₁ KO did not affect the expressions at P0 of type II collagen (Col II), a marker of resting and proliferating chondrocytes or type X collagen (Col X), a marker of pre-hypertrophic and early hypertrophic chondrocytes (Fig. S2a,e). Taking account of the fact that both LPA₁ and ATX were highly expressed in the cartilage tissues in both zebrafish and mice (Fig. 1c and Fig. S2f), we conclude that ATX-LPA₁ signaling functions after chondrocyte differentiation and that dyschondroplasia is the main defect of LPA₁ KO and ATX^flox/− mice and is the cause of craniofacial abnormalities and retardation of physical growth in these mice.

Inhibition of ATX-LPA₁ signaling delays S-phase entry in chondrocytes

To examine the effect of LPA signaling on cellular functions of chondrocytes, we established primary cultured chondrocytes from rib cages. We found that proliferation of LPA₁^−/− chondrocytes was significantly slower than that of LPA₁^+/+ and LPA₁^+/− chondrocytes (Fig. 3a). In addition, the cell size of LPA₁^−/− chondrocytes was much smaller (Fig. S3a). We didn’t observe any significant changes in the proliferative activity (Fig. 3a) or cell size (data not shown) between LPA₁^+/+ and LPA₁^+/− chondrocytes. Thus in the following experiments, we used LPA₁^+/− chondrocytes as a control. Time-lapse images of the proliferating chondrocytes revealed that the doubling time for LPA₁^−/− chondrocytes (1660 ± 400 min) was significantly longer than that for LPA₁^+/− chondrocytes (1280 ± 220 min) (Fig. S3b and Videos 1 and 2). Although the duration of the M phase did not differ between the two types of chondrocytes (Fig. S3c and Videos 1 and 2), we found that cell cycle progression from the G₀/G₁ to S-phase as judged by BrdU incorporation was significantly reduced in LPA₁^−/− chondrocytes (Fig. 3c,d). Importantly, the smaller cell size and the reduced proliferative activity were reproduced by adding an LPA₁ antagonist (Ki16425) to cultures of LPA₁^+/− chondrocytes (Fig. 3b–d, Fig. S3a,b and Video 3) or LPA^+/+ chondrocytes (data not shown), indicating that the phenotypes of LPA₁^−/− chondrocytes are not due to congenital changes of the chondrocytes. Addition of an ATX inhibitor (ONO-8430506) also resulted in a smaller cell size (Fig. S3a) and reduced proliferative activity, albeit to a lesser extent than the LPA₁ antagonist (Fig. 3b–d and Video 4). Unlike LPA₁^−/− chondrocytes or chondrocytes treated with LPA₁ antagonist or ATX inhibitor, chondrocytes from ATX^flox/− mice proliferated normally. It should be mentioned that the culture medium contained significant amounts of ATX and its substrate, lysophosphatidylcholine, both of which are present within fetal calf serum (FCS).

In serum-free medium, LPA significantly stimulated S phase entry of LPA₁^+/− chondrocytes whereas the effect was not observed in LPA₁^−/− chondrocytes or Ki16425-treated LPA₁^+/− chondrocytes (Fig. 3e,f). In vivo imaging of LPA₁ KO mice at P0 using 5-ethynyl-2′-deoxyuridine (EdU) showed that the proliferation of chondrocytes in the intersphenoid synchondrosis was significantly decreased (Fig. 3g,h). These in vivo and in vitro findings indicate that in the absence of LPA₁ signaling, the G₀/G₁-to-S phase transition is prolonged, which explains the reduction of cell proliferation and decreased cell number of chondrocytes in the intersphenoid synchondrosis (Fig. 2g and 3g, h).

Integrin-mediated adhesion to fibronectin promotes LPA-induced S-phase entry of chondrocytes

When stimulated by LPA in the presence of EdU, most of the EdU-positive chondrocytes adhered tightly to the culture plates and spread fully (Fig. 4a), suggesting that the adhesive property of the chondrocytes affected their proliferation. Since chondrocytes in cartilage tissues are surrounded by and are in contact with extracellular matrix (ECM), which mainly consists of Col II and fibronectin (FN), we examined the effect of ECM on LPA-induced cell proliferation. Strikingly, FN coating dramatically enhanced the LPA-induced BrdU incorporation of LPA₁^+/− chondrocytes (Fig. 4b). The FN effect was not observed in LPA₁^−/− chondrocytes and was suppressed by treating the cells with LPA₁ antagonist, Ki16425, as well as by treating them with Y27632 and PTX (Fig. 4b,c), indicating that LPA-induced proliferation on FN-coated plates was LPA₁-dependent and mediated via both the G_α12/13 and G_αi–linked pathways. LPA enhanced the spreading of LPA₁^+/− but not LPA₁^−/− chondrocytes on FN-coated plates (Fig. 4d,e). LPA had similar effects on Col II-coated plates and on non-coated plates (data not shown) (Fig. 4b and Fig. S4a,b). On both FN- and Col II-coated plates, cell proliferation was suppressed by GRGDSP, an integrin blocking peptide, but not by a control peptide, GRGESP (Fig. 4f and Fig. S4c), showing an important role for integrins in LPA-induced chondrocyte proliferation.

LPA enhances fibronectin assembly through LPA₁

FN plays an important role in cell adhesion, which, in turn, affects the proliferation and survival of many cell types²³. In LPA₁^+/− chondrocytes at 24 hours after LPA stimulation, FN was distributed in filamentous structures, which overlapped with F-actin and β1-integrin (Fig. 5a). In contrast, in LPA₁^−/− chondrocytes and LPA₁^+/− chondrocytes treated with Ki16425, Y27632 or PTX, such filamentous structures were less developed (Fig. 5a). In the presence of LPA, extracellularly added fluorescently labeled FN (Hilyte-488 FN) was readily incorporated into the filamentous structures and colocalized with F-actin and β1-integrin in LPA₁^+/− but not LPA₁^−/− chondrocytes (Fig. 5b). It thus appears that FN, once secreted from chondrocytes, is incorporated and assembled into filamentous structures in an LPA₁- and integrin-dependent manner. Consistent with this notion, LPA-induced FN assembly was prominently suppressed by Ki16425, Y27632 and PTX and partially inhibited by GRGDSP peptide but not by GRGESP (Fig. 5c). Importantly, most of the EdU-positive cells showed the filamentous FN structures (Fig. 5d,e). On the basis of these observations, we hypothesized that LPA stimulates S-phase entry by enhancing FN assembly downstream of LPA_1, through β1-integrin activation. To further confirm that the interaction of β1 integrin and FN transmits the intracellular signaling, we examined the formation of focal adhesions which are known to be activated by the coordinated action of β1-integrin and FN. We observed that LPA₁ signaling promoted focal adhesion assembly as judged by colocalization of vinculin and actin (Fig. S5). Focal adhesions distributed peripherally in non-stimulated cells even in LPA₁^−/− chondrocytes. But, only in LPA₁^+/− chondrocytes, LPA promoted focal adhesions larger and some of them existed under the cell bodies. and this conformational change was also inhibited by Ki16425, Y27632 and PTX (Fig. S5). These observations confirmed the idea that intracellular signaling via β1-integrin was activated downstream of LPA₁ signaling.

ECM formed through LPA₁ signaling supports the proliferation of chondrocytes

To examine the biological significance of LPA₁-mediated FN assembly, we prepared ECM by culturing LPA₁^+/− and LPA₁^−/− chondrocytes. After 10 days of culture, filamentous FN structures were well developed in LPA₁^+/− chondrocytes but less developed in LPA₁^−/− chondrocytes (Fig. 6a). Furthermore, LPA₁^−/− chondrocytes had significantly less deoxycholate-insoluble FN protein (Fig. 6b) and significantly less total ECM proteins (Fig. 6c). Under these conditions, both types of chondrocytes showed equal cell numbers per well (Fig. S6a) and FN and Col II mRNA levels (Fig. 6d), suggesting that deposition of ECM components was attenuated in LPA₁^−/− chondrocytes. Transmission electron micrographs of LPA₁^+/− and LPA₁^−/− chondrocytes confirmed that thick filamentous bundles were better developed in LPA₁^+/− chondrocytes than in LPA₁^−/− chondrocytes (Fig. S6b). We next compared the abilities of decellularized ECMs (Fig. S6c) formed by LPA₁^+/− and LPA₁^−/− chondrocytes to support cell proliferation of LPA₁^+/− chondrocytes. After 5 days of culture, ECMs formed by LPA₁^+/− chondrocytes had significantly more cells than ECMs formed by LPA₁^−/− chondrocytes (Fig. 6e). Thus LPA₁ signaling regulates the nature of the ECM and provides the proper external milieu for cell proliferation.

Discussion

LPA₁ KO mice show obvious craniofacial abnormalities, retardation of physical growth and a low bone mass^6,7. Previous reports indicated that loss of LPA₁ signaling resulted in impaired differentiation of osteoclasts and osteoblasts in vitro^7,24,25. However, the fundamental causes of the defects of LPA₁ KO mice has remained unclear. In this study we showed that LPA₁ is highly expressed in chondrocytes in both fish (Fig. 1c) and mice (Fig. S2f) and has a critical role in stimulating the proliferation and positioning of chondrocytes (Fig. 1e and 2f), thereby promoting proper cartilage formation. Bones, especially long limb bones, are principally formed by a process called endochondral ossification, in which cartilage is replaced by bone and chondrocytes serve as a center of bone growth²⁶. Because the cartilage phenotype was observed before bone tissues had formed, we conclude dyschondroplasia is the primary cause of impaired bone development and retarded physical growth of LPA₁ KO mice. We also showed that ATX is highly expressed in chondrocytes of both zebrafish and mice and that inactivation of ATX in both species phenocopied dyschondroplasia as observed when LPA₁ was attenuated or deleted. We therefore conclude that ATX is the major LPA-producing enzyme in cartilage tissues.

The dyschondroplasia of LPA₁ KO and ATX^flox/− mice can be attributed to dysfunction of chondrocytes. In vitro experiments revealed that the ATX-LPA₁ axis has a pivotal role in chondrocyte proliferation. We also examined the effect of an ATX inhibitor and LPA₁ antagonist on cell proliferation of various cell types and found that chondrocytes are more sensitive to LPA₁ signaling than other cell types (data not shown). The ATX-LPA₁ axis promoted cell cycle progression of chondrocytes, in agreement with LPA having growth factor-like activities^3,4,27,28 and anti-apoptoc effects as observed in the nervous system^5,29,30. Pathophysiological effects of LPA as a growth factor remain less clear, however it can disrupt normal processes relevant to neurodevelopmental disorders like hydrocephalus and schizophrenia^31,32. To the best of our knowledge, the present study is the first to demonstrate a physiological meaning for LPA-induced cell proliferation through cell cycle progression.

Our present results raise a new question: how does LPA support the proliferation of chondrocytes downstream of LPA₁? Kingsbury et al. reported that LPA-induced cell growth of neural progenitor cells is not due to increased proliferation but rather to reduced cell death via LPA_1/2⁵. However, this is not the case, because we did not observe any sign of cell death when LPA₁ signaling was attenuated (Videos 1,2,3,4). These observations also exclude the possibility that loss of LPA₁ signal induced anoikis in chondrocytes. Clues to answering this question are that LPA₁^−/− chondrocytes were less adhesive to ECM than LPA₁^+/− chondrocytes (Fig. 4d,e) and that LPA-induced chondrocyte proliferation was dramatically enhanced by the presence of FN and suppressed by an integrin-blocking RGD peptide (Fig. 4b,f). These findings suggest that LPA supports chondrocyte proliferation by upregulating β1-integrin-mediated cell adhesion to FN. Indeed, mice in which cartilage-specific β1-integrin is conditionally knocked out exhibited dyschondroplasia similar to that of LPA₁ KO mice³³. Furthermore, topical administration of anti-α5β1-integrin antibody or RGD-containing peptide to the upper limbs of mouse fetuses suppressed chondrocyte proliferation and shortened the upper limbs³⁴. Together, these findings indicate that β1-integrin and LPA₁ have closely related roles in the formation of cartilage.

Another important observation for understanding the mechanism of LPA-induced proliferation is that FN was assembled to form filamentous structures (Fig. 5a–c and Fig. S5). We presumed that the filamentous FN structure is a multimeric FN, i.e., FN fibril. FN fibrils within the ECM play central roles in both physiological and pathological processes during development and tissue regeneration by coordinating cell adhesion, growth, migration and differentiation. FN is assembled to form FN fibrils in an integrin- and actin stress fiber-dependent manner^35,36,37,38. In vitro studies using fibroblasts and platelets have suggested that integrins and some agonists for G protein–coupled receptors including LPA are involved in the assembly of FN fibrils^39,40,41. The present results clearly demonstrate that LPA₁ signaling induced assembly of the filamentous FN structures undersurface of chondrocytes; the FN structures then modified the proliferative and adhesive property of the cells. The structures were co-localized with actin stress fiber and β1-integrin (Fig. 5a,b) and their formation was canceled by Y27632 and PTX (Fig. 5a,c). The coordinated changes in integrins and the actin cytoskeleton observed here parallel cadherin and focal adhesion assembly associated with actin changes observed in Schwann cells of the nervous system⁴². All together, these findings support the idea that the filamentous FN structure is an FN fibril and that the LPA-LPA₁ axis regulates actin stress fiber formation and then β1-integrin translocation. These changes stimulate the FN assembly to form FN fibrils, which enhances the proliferation of chondrocytes (Fig. 7). This model is further supported by the observation that the ECM formed by chondrocytes in the presence of LPA₁ signaling supports cell proliferation more efficiently than ECM formed in the absence of LPA₁ signaling (Fig. 6b). The observation also implicates that integrin activity itself can be regulated in both an inside-out and outside-in manner^43,44. The present results indicate that the ATX-LPA₁ axis regulates integrins in an inside-out manner (from step (i) to (v) in Fig. 7), which results in the formation of FN fibrils and the subsequent organization of other ECM components such as Col II. The ECM thus formed then regulates integrins in an outside-in manner (from step (v) to (vi) in Fig. 7), which contributes to proper cell adhesion and proliferation of chondrocytes, leading to normal cartilage development. Thus, in addition to the previously indicated LPA₁-G_αi-Akt pathway⁴⁵, we propose a new model of ATX-LPA-LPA₁ axis-induced cell proliferation that is mediated by integrin-dependent fibronectin assembly, occurring in an “inside-outside-in” manner (Fig. 7).

A key question is whether cells other than chondrocytes utilize this system for their proliferative activity. Chondrocytes and fibroblasts are both derived from mesenchymal stem cells. LPA₁ KO mice were resistant to bleomycin-induced lung fibrosis⁴⁶ and unilateral ureteral obstruction-induced renal fibrosis⁴⁷. Thus it is possible that LPA-LPA₁ signaling upregulates proliferation of fibroblasts in a manner similar to what we propose in chondrocytes. In addition, because LPA₁ and ATX are overexpressed in several cancers^8,48,49, the enhanced proliferation of certain LPA₁-positive cancer cells, such as glioblastoma cells⁵⁰, may reflect a similar mechanism, which is currently being examined.

Materials and Methods

Reagents

Lysophospholipids including 1-oleoyl (18:1)-LPA and 1-myristoyl (14:0)-LPC were purchased from Avanti Polar Lipids. Lysophospholipids were dried under nitrogen gas and dissolved in 0.1% fatty acid-free bovine serum albumin (Sigma-Aldrich)-PBS using water bath sonication and stocked in −20 °C. Y27632, ROCK inhibitor, was purchased from Merck. Pertussis toxin was purchased from Wako Pure Chemicals, GRGDSP and GRGESP were purchased from AnaSpec. Ki16425 was chemically synthesized as described previously⁵¹. ATX inhibitor, ONO-8430506 was a generous gift from Ono pharmaceutical Co., Ltd⁵².

Zebrafish lines

Zebrafish were maintained according to the Guidelines for Animal Experimentation of Tohoku University and the protocol was approved by the Institutional Animal Care and Use Committee at Tohoku University. Wild-type AB zebrafish were obtained from Zebrafish International Resource Center (University of Oregon, Eugene, OR). The LPA₁ mutant was generated by target-selected mutational inactivation of genes (TILLING) according to standard methods⁵³. Fish were maintained at 27–28 °C under a controlled 13.5 h light/10.5 h dark cycle. Embryos were obtained from natural spawning and staged according to morphology. The standard staging of zebrafish embryos is used and determined in hpf (hour post fertilization) or dpf (day post fertilization) at 28 °C⁵⁴.

Generation of col2a1:egfp transgenic lines

To generate a col2a1:egfp transgenic line, the col2a1 promoter¹⁸ was introduced upstream of egfp using the Tol2kit system⁵⁵. Briefly, the col2a1 promoter was introduced into p5E-MCS vector and then p5E-col2a1, pME-EGFP, p3E-polyA and pDestTol2pA2 were combined with LR clonase II plus (Life Technologies). Twenty-five ng of the DNA plasmid was injected into the embryos at the 1-cell stage to establish a col2a1:egfp transgenic line that selectively expresses EGFP in cartilage.

Microscopic Analysis

Zebrafish larvae were anesthetized with 0.016% tricaine methanesulfonate solution (Sigma-Aldrich) and positioned in 3% methylcellulose (Sigma-Aldrich) on a slide glass. Images were captured with a Leica M80 stereomicroscope equipped with a Leica DFC425 digital camera (Leica Microsystems). Col2a1:egfp transgenic fish larvae were positioned in 1.5% agarose (Sigma-Aldrich) on a glass-bottom well (MatTek, MA) and imaged with a LSM 700 confocal laser-scanning microscope (Carl Zeiss).

Alcian blue staining

Zebrafish cartilage and bone were double stained with alcian blue and alizarin red as described previously⁵⁶. At 4 or 5 dpf, larvae were fixed with 4% PFA in PBS, rocked at room temperature for 2 h, washed and dehydrated with 50% EtOH at room temperature for 10 min. After removing the EtOH, the larvae were double stained with acid-free stain solution (0.02% alcian blue, 0.005% alizarin red, 150 mM MgCl₂ in 70% EtOH), rocked overnight at room temperature, washed with water, depigmented with bleaching solution (1.5% H₂O₂ in 1% KOH), rocked at room temperature for 20 min and cleared with successive changes of a solution of glycerol and KOH. The lengths of Meckel’s and ceratohyal cartilages were measured with a Zeiss Axio Imager (Carl Zeiss MicroImaging). Larvae with smaller or abnormally bent cartilage compared with control larvae were classified as ‘malformed’ larvae.

Antisense morpholino injection

The morpholino (MO) sequences were:

LPA₁ MO, 5′-TGGAGCACTTACCCAATACAATCAC-3′¹³;

ATX MO1 (MO1-enpp2), 5′-GGAGAATACCTGGGTCGAGACACCG-3′¹⁴;

ATX MO2 (MO2-enpp2), 5′-TTCTTTCAAAGTCCCTACCTTTGCA-3′¹⁴.

LPA1 (2.5 ng), ATX MO1 (0.3 ng) and ATX MO2 (3.2 ng) of each MOs were injected into the yolks of one- to two-cell stage embryos.

Ki16425 treatment for zebrafish embryos

Embryos were treated with 120 μM of Ki16425 in the embryo medium with 1% DMSO⁵⁷ from 48 hpf. Embryo medium containing Ki16425 was replaced approximately every 24 hours.

Genotyping of lpa₁ mutant

The lpa₁ mutation was genotyped by restriction fragment length polymorphism (RFLP) analysis. All the primers were designed by a web-based program, dCAPS Finder 2.0⁵⁸. The primer sequences were:

LPA₁-180, 5′-ACAAGAAGGCTAACGGTTAGCACGTG-3′;

LPA₁-181, 5′-ACAAGAAGGCTAACGGTTAGCAGGTG-3′;

LPA₁-182, 5′-CATGACGATAGACATGGTCCAGATGATG-3′.

In the 198-bp PCR products derived from the wild-type allele with LPA₁-180 and LPA₁-182, an ScrFI restriction site was introduced and therefore the ScrFI treatment degraded the 198-bp PCR product from the wild-type allele into a 174-bp fragment. The PCR products were cleaved with ScrFI and resolved on a 4% agarose gel. Similarly, the 198-bp PCR products from the mutant alleles with LPA₁-181 and LPA₁-182, HphI restriction site were introduced. The HphI restriction enzyme degrades the PCR product from mutant allele into a 163-bp DNA fragment.

Whole mount in situ hybridization

Antisense RNA probes labeled with digoxigenin (DIG) for lpa1, atx, slug, sox10 and sox9a were prepared with an RNA labeling kit (Roche Applied Science). Whole-mount in situ hybridization was performed as previously described⁵⁹.

Mice

Mice were maintained according to the Guidelines for Animal Experimentation of Tohoku University and the protocol was approved by the Institutional Animal Care and Use Committee at Tohoku University. The LPA₁ KO mice generated by Contos et al.⁶ were transferred and maintained on a mixed 129SvJ/C57BL/6J background. Experiments comparing LPA₁ KO and HT mice used offspring of mice heterozygous for the LPA₁ mutant allele. The ATX conditional KO mice with two floxP alleles (ATX^flox/flox) were generated by van Meeteren et al.⁶⁰. ATX heterozygous KO mice (ATX^+/−) were generated by Tanaka et al.⁹. Mice with a floxP allele and ATX null allele (ATX^flox/−) were generated by crossing ATX^+/− and ATX^flox/flox mice.

Skeletal analysis

Plain radiographs were taken using a soft X-ray apparatus (LaTheta LCT-200, Hitachi-Aloka). The size of the head and the lengths of the femur, tibia and humerus were determined with ORS Visual (Nihon Binary) software. Whole-mount skeletal staining was conducted as described previously, with slight modifications⁶¹. Briefly, mice were skinned, eviscerated and dehydrated in 95% (v/v) ethanol overnight. Skeletons were stained with 0.3% (w/v) alcian blue (Sigma-Aldrich) for 24 hours, stained with 1.5% (w/v) alizarin red (Wako) for 2 hours, treated with 1% (w/v) potassium hydrate and stored in glycerol/EtOH.

Isolation of primary mouse rib chondrocytes

Chondrocytes were isolated as previously described⁶². In short, chondrocytes from rib were isolated from newborn wt, LPA₁ HT or KO mice. Rib cages were dissected, rinsed in PBS, incubated at 37 °C for 45 min in 3 mg/ml collagenase type II (Worthington), cleaned of adherent tissues and digested with 0.5 mg/ml collagenase type II at 37 °C overnight. The cell suspension was filtered through a 40 μm cell strainer. The cells were washed, counted and plated. For all experiments, chondrocytes were plated 48 hr before any treatment.

Evaluation of cell proliferation

Cell proliferation was determined with a Cell Counting Kit-8 (Dojindo). Cells were plated in 96-well plates at 1 × 10⁴ cells per well and cultured in the growth medium. Sixty min before the indicated time points, CCK-8 solution was added to each well and the cell numbers were measured by reading the absorbance (450 nm). To evaluate DNA synthesis, cells were seeded on cell culture-treated or Col II- or FN-coated (Sigma Aldrich, 5 μg/mL each) wells in 96-well plates. After 24 hr starvation, the cells were treated with 10 μM 5′-BrdU (Sigma-Aldrich) and 10% FCS or 10 μM LPA for the indicated times. When an inhibitor was used, cells were treated with Ki16425 (5 μM), ONO-8430506 (10 μM), Y27632 (10 μM), GRGDSP (500 μM) or GRGESP (500 μM) for 30 min before the addition of FCS or LPA. PTX (200 ng/mL) was treated at the same time as starvation. 5′-BrdU incorporation was quantified by counting 5′-BrdU-positive cells. 5′-BrdU was detected by anti-BrdU antibody conjugated to fluorescein (Roche, 1:1000). For EdU (Life Technologies) incorporation in vitro, 10 μM EdU was used instead of 5′-BrdU. To examine the incorporation of EdU in vivo, pups (P0-1) were subcutaneously injected with 50 mg/kg EdU and were sacrificed 2 hours after the injection. Heads of the embryos were dissected and fixed in 4% paraformaldehyde overnight at 4 °C. Tissues were embedded in Optimal Cutting Temperature (OCT) compound and stained according to the manufacturer’s instructions. EdU-positive cells were counted in a resting zone within a unit area. All images were captured with the LSM700 confocal microscope equipped with 10×/0.45 M27 Plan-Apochromat.

Tissue and cell staining

For histological analysis, newborn mice were fixed in 4% fresh paraformaldehyde in PBS, pH 7.2, overnight, dehydrated in a graded alcohol series (50, 70, 90, 95 and 99.5%) and embedded in paraffin. Sections were cut at 10 μm and stained with hematoxylin and eosin (Mutoh Industries).

For immunohistochemistry, paraformaldehyde-fixed tissues were embedded in OCT compound or in paraffin. For paraffin sections, antigens were retrieved with 10 mM citrate buffer (pH 7.0) at 120 °C for 10 min. Tissue sections were cut with a cryostat (Leica Microsystems) or microtome (Leica Microsystems). Primary antibodies were used with predetermined optimal concentrations. The primary antibody was anti-Col II (Abcam, ab21291, 1:250). The secondary antibody to detect Col II was conjugated with Alexa Fluor 488 or 568 (Life Technologies, 1:200). Nuclei were counterstained with 4′, 6-diamidino-2-phenylindole (DAPI).

For Immunofluorescence, cells were plated on glass coverslips (Thermo Fisher Scientific) that had been treated with cell culture or coated with Col II or FN and were fixed with 4% paraformaldehyde in PBS for 30 min, permeabilized with 0.5% TritonX-100 in PBS for 15 min and blocked with 3% BSA/PBS for 30 min. The primary antibodies were goat anti-FN (Santa Cruz Biotechnology, sc-6952, 1:50), rabbit anti-vinculin (Abcam, ab73412, 1:150) and rat anti-β1-integrin (Millipore, MAB1997, 1:1000). The secondary antibodies for detecting goat anti-FN, rabbit anti-vinculin and rat anti-β1-integrin were conjugated with Alexa Fluor 488 or 647 (Life Technologies. 1:1000). Actin filaments were stained with Alexa594-conjugated phalloidin (Life Technologies, A12381, 1:250). Fluorescent images were captured with the LSM700 confocal microscope equipped with 63×/1.40 Oil DIC M27 Plan-Apochromat.

Time-lapse

In live-cell imaging of chondrocytes, phase-contrast images were taken (LD plan-NEOFLUAR, 20x/0.4, Ph2, 6frames/hr) with a Zeiss inverted microscope (Axio Observer.Z1) equipped with a heated chamber (37 °C) and CO₂ controller (5.0%) over a period of 48 hr. Primary chondrocytes were seeded onto 12-well plates (Greiner), incubated for 2 days at 37 °C and treated with an LPA₁ antagonist (Ki16425, 5 μM) or an ATX inhibitor (ONO-8430506, 10 μM). Live-cell images were taken every 5 or 10 min for 48 hr. We chased cells that divided twice. The doubling time was taken as the time between the two divisions. The duration of the M phase was evaluated by cell morphology. Supplementary Videos 1,2,3,4 were simplified as 1.5 frames/hr for downsizing.

Evaluation of cell spreading

Cells were plated on glass coverslips coated with Col II or FN and were stained with phalloidin. Cell images were captured with the LSM700 confocal microscope equipped with 63×/1.40 Oil DIC M27 Plan-Apochromat. and the cell spreading areas were calculated using the Zeiss Efficient Navigation system (Carl Zeiss).

Quantitative RT-PCR

To prepare total RNA from tissues, tissues were first embedded in OCT compound and frozen sections were cut at 25 μm thickness and mounted on poly-l-lysin–coated LCM transfer film (LEICA-BEST) on glass slides. The chondrocytes in the tissue sections were dissected with a Leica LMD7000 Laser Microdissection System. RNA of the harvested chondrocytes was extracted with an RNAqueous Micro Kit (Ambion). Total RNA from cultured cells was isolated using a GenElute Mammalian Total RNA Miniprep Kit (Sigma-Aldrich). Total RNA samples were reverse-transcribed using High-Capacity cDNA RT Kits (Applied Biosystems) according to the manufacturer’s instructions. PCR reactions were performed with SYBR Premix Ex Taq (Takara Bio) on an ABI Prism 7300 thermocycler (Applied Biosystems). Standard plasmids ranging from 10² to 10⁸ copies per well were used to quantify the absolute number of transcripts of cDNA samples. The numbers of transcripts were normalized to the number of transcripts of a house-keeping gene (GAPDH) in the same sample.

The primers used to determine mouse gene expressions were:

mGAPDH 5′-AGGAGCGAGACCCCACTAAC-3′ and 5′-CGGAGATGATGACCCTTTTG-3′;

mLPA₁ 5′-AGGAGGAATCGGGACACCA-3′ and 5′-AGCACACATCCAGCAATAACAA -3′;

mLPA₂ 5′-TGCCGCTTGACTGGATGT-3′ and 5′-GCTCCTTGCCGCTGTTATT-3′; mLPA₃ 5′-ACCAACGTCTTATCTCCACACAC-3′ and 5′-CAGCAGCAGAACCACCAGAC-3′;

mATX 5′-GGAGAATCACACTGGGTAGATGATG-3′ and 5′-ACGGAGGGCGGACAAAC-3′;

mFibronectin 5′-CAGAACCAGAGGAGGCACAAG-3′ and 5′-CAATGGCGTAATGGGAAACC-3′;

mCollagen type II 5′-AGAGGGGACTGAAGGGACAC-3′ and 5′-GCACCCTGATCTCCAGAAGG-3′.

Evaluation of Fibronectin assembly

Cells were plated on Col II-coated glass coverslips and stimulated with LPA after 24 hr starvation for 24 hr. To measure FN incorporation, 1 μg/ml Hilyte-488 FN (Cytoskeleton) was added with LPA. After 12 hours, cells were fixed and stained. Cells were treated with Ki16425 (5 μM), Y27632 (10 μM), GRGDSP (500 μM) or GRGESP (500 μM) for 30 min before LPA stimulation. PTX (200 ng/mL) treatment was applied at the same time as starvation. Fluorescent images were captured with the LSM700 confocal microscope equipped with 63×/1.40 Oil DIC M27 Plan-Apochromat.

Decellularization

Cells were plated in 24-well plates with glass coverslips or 96-well plates at 1 × 10⁵ cells per well and cultured in the growth medium. After 10 days, cells were washed with PBS and treated with 0.5% Triton X-100 and 20 mM ammonium hydroxide in PBS for 5 min. Decellularized ECMs were treated with DNase (Sigma-Aldrich) at 50 unit/mL for 60 min at 37 °C, then gently washed with PBS. 24-well plates were stained with anti-FN antibody and images were captured with a Leica TCS SP-8 confocal microscope. 96-well plates were stored at −80 °C until use. To assay cell proliferation, cells were plated onto decellularized ECM plates at 0.5 × 10⁴ cells per well.

Western blot analysis of deoxycholate (DOC)-insoluble fibronectin

After 10 days culture, cells were washed with PBS and solubilized with deoxycholate lysis buffer (DOC-buffer) containing 2% sodium deoxycholate, protease inhibitors (Complete Protease Inhibitor Cocktail, Roche), 20 mM Tris-HCl pH 8.8, 2 mM EDTA, 2 mM iodoacetamide and 2 mM N-ethylmaleimide. Homogenates were passed ten times through a 23-gauge needle and centrifuged at 15,000 × g for 15 min at 4 °C. DOC-insoluble fractions were washed with DOC-buffer 3 times and then solubilized with SDS lysis buffer (1% SDS, 25 mM Tris-HCl pH 8.0, 2 mM EDTA, protease inhibitors, 2 mM iodoacetamide and 2 mM N-ethylmaleimide). Equal volumes of DOC-insoluble samples were analyzed by SDS-PAGE using 5% polyacrylamide gels. The primary antibody was goat anti-FN (Santa Cruz Biotechnology, sc-6952, 1:200). A secondary antibody conjugated with horse radish peroxidase (Dako, P0449, 1:1000) against goat IgG was used. Images were captured with a digital gel imaging system (ChemiDoc XRS⁺, BIO-RAD).

Evaluation of ECM amount

After decellularization, ECMs were solubilized with 5.0 M Urea, 2.0 M Thiourea, 50 mM DTT and 0.1% SDS in PBS and scraped with a rubber policeman⁶³. The collected lysates were placed in 95 °C water for 5 min and centrifuged at 12000 × g for 10 min at 4 °C. Protein concentration was evaluated by the Bradford method.

Transmission electron microscopy (TEM)

Samples were fixed with 2% paraformaldehyde and 2% glutaraldehyde in 0.1M cacodylate buffer pH 7.4 at 4 °C overnight, washed 3 times with 0.1 M cacodylate buffer for 30 min each, postfixed with 2% osmium tetroxide in 0.1M cacodylate buffer at 4 °C for 3hr, dehydrated in graded ethanol solutions (50%, 70%, 90% and 100%), infiltrated with propylene oxide (PO) 2 times for 30 min each, transferred to a 70:30 mixture of PO and resin (Quetol-812, Nisshin EM Co.) for 1h, allowed to stand overnight to volatize the PO, transferred to fresh 100% resin and heated at 60 °C for 48 hr to polymerize the resin. Seventy-nm sections were cut from the polymerized resins (Ultracut UCT, Leica), mounted on copper grids, stained with 2% uranyl acetate at room temperature for 15 min, washed with distilled water, secondary-stained with lead stain solution (Sigma-Aldrich) at room temperature for 3 min and observed with a transmission electron microscope (JEM-1400Plus, JEOL) at an acceleration voltage of 80 kV.

In situ hybridization

Heads from newborn mice were embedded in OCT compound. Seven-μm-thick sections were cut, placed on MAS-coated glass slides (Matsunami Glass), fixed with 4% PFA-PBS, acetylated with 0.5% (v/v) acetic anhydride/0.1M triethanolamine pH 8.0, permeabilized with 0.3% (v/v) TritonX-100/PBS, hybridized with digoxigenin labeled-RNA probes corresponding to 4551-4988 nt of Col2a1 cDNA (NM031163) or 1302-1816 nt of Col10a1 cDNA (NM009925), incubated with anti-digoxigenin antibody conjugated with alkaline phosphatase (Roche), stained with NBT/BCIP (BM purple AP, Roche Diagnostics) and photographed with a Zeiss Axio Imager (Carl Zeiss MicroImaging).

Determination of lysophospholipase D activity

Lysophospholipase D activity of mice plasma was determined as described using 14:0 lysophosphatidylcholine as substrate⁸.

Statistics

All statistical analyses were carried out using EXSAS. Differences were considered significant at P < 0.05. All data are expressed as means ± s.d.

Additional Information

How to cite this article: Nishioka, T. et al. ATX-LPA₁ axis contributes to proliferation of chondrocytes by regulating fibronectin assembly leading to proper cartilage formation. Sci. Rep. 6, 23433; doi: 10.1038/srep23433 (2016).

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Acknowledgements

We thank Drs. Rodney M. Dale and Jacek Topczewski for generous gift of col2a1a promoter. We also thank Ono pharmaceutical Co., Ltd for the gift of the ATX inhibitor, ONO-8430506. The present work was supported partly by AMED-CREST (Japan Agency for Medical Research and Development, Core Research for Evolutional Science and Technology) for J.A., PRESTO (Japan Science and Technology Agency, Precursory Research for Embryonic Science and Technology) for A.I. and Ministry of Education, Culture, Sports, Science and Technology (MEXT) Grant-in-Aid for Scientific Research for K.H. and J.A. Work in LSK lab was supported by R01HD076585 from NIH.

Author information

Nishioka Tatsuji and Arima Naoaki contributed equally to this work.

Authors and Affiliations

Graduate School of Pharmaceutical Sciences, Tohoku University, 6-3, Aoba, Aramaki-aza, Aoba-ku, 980-8578, Sendai, Japan
Tatsuji Nishioka, Naoaki Arima, Kuniyuki Kano, Kotaro Hama, Eriko Itai, Hiroshi Yukiura, Ryoji Kise, Asuka Inoue & Junken Aoki
Japan Science and Technology Agency, Precursory Research for Embryonic Science and Technology (PRESTO), Kawaguchi City, 332-0012, Saitama, Japan
Asuka Inoue
Department of Medicine, Medical University of South Carolina, Charleston, 29425, SC, USA
Seok-Hyung Kim
Department of Developmental Biology, Washington University School of Medicine, St. Louis, 63110, MO, USA
Lilianna Solnica-Krezel
Division of Cell Biology, The Netherlands Cancer Institute, Plesmanlaan 121, 1066CX, Amsterdam, The Netherlands
Wouter H. Moolenaar
Department of Molecular and Cellular Neuroscience, Dorris Neuroscience Center, The Scripps Research Institute, La Jolla, CA, 92037, USA
Jerold Chun
Japan Agency for Medical Research and Development, Core Research for Evolutional Science and Technology (AMED-CREST), Chiyoda-ku, 100-0004, Tokyo, Japan
Junken Aoki

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Tatsuji Nishioka
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T.N. and N.A. designed the study on chondrocytes and zebrafish, respectively and carried out most of the experiments and wrote the draft of manuscript. K.H. designed the study on zebrafish with N.A., E.I. performed part of experiments using chondrocytes. K.K. and A.I. performed and analyzed some experiments using mice and gave many useful comments. H.Y. and R.K. performed and analyzed some experiments using zebrafish and gave many useful comments. S.H.K. and L.S.K. isolated zebrafish LPA₁ mutant. W.M. and J.C. provided the ATXflox and LPA₁ KO mice, respectively, modified the manuscript and gave many useful comments. J.A. supervised all aspects of the study, including experimental design, discussion, data interpretation and modified the manuscript.

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Nishioka, T., Arima, N., Kano, K. et al. ATX-LPA₁ axis contributes to proliferation of chondrocytes by regulating fibronectin assembly leading to proper cartilage formation. Sci Rep 6, 23433 (2016). https://doi.org/10.1038/srep23433

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Received: 12 November 2015
Accepted: 07 March 2016
Published: 23 March 2016
DOI: https://doi.org/10.1038/srep23433

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