Helper T cells actively communicate with adjacent cells by secreting soluble mediators, yet crosstalk between helper T cells and endothelial cells remains poorly understood. Here we found that placental growth factor (PlGF), a homolog of the vascular endothelial growth factor that enhances an angiogenic switch in disease, was selectively secreted by the TH17 subset of helper T cells and promoted angiogenesis. Interestingly, the ‘angio-lymphokine’ PlGF, in turn, specifically induced the differentiation of pathogenic TH17 cells by activating the transcription factor STAT3 via binding to its receptors and replaced the activity of interleukin-6 in the production of interleukin-17, whereas it suppressed the generation of regulatory T cells. Moreover, T cell-derived PlGF was required for the progression of autoimmune diseases associated with TH17 differentiation, including experimental autoimmune encephalomyelitis and collagen-induced arthritis, in mice. Collectively, our findings provide insights into the PlGF-dictated links among angiogenesis, TH17 cell development and autoimmunity.
Access optionsAccess options
Subscribe to Journal
Get full journal access for 1 year
only $18.75 per issue
All prices are NET prices.
VAT will be added later in the checkout.
Rent or Buy article
Get time limited or full article access on ReadCube.
All prices are NET prices.
The microarray dataset has been deposited in the GEO with the accession code GSE118281. The data supporting the findings of this study are available from the corresponding author upon reasonable request.
Tran, C. N., Lundy, S. K. & Fox, D. A. Synovial biology and T cells in rheumatoid arthritis. Pathophysiology 12, 183–189 (2005).
Zhou, L., Chong, M. M. & Littman, D. R. Plasticity of CD4+T cell lineage differentiation. Immunity 30, 646–655 (2009).
Dong, C. TH17 cells in development: an updated view of their molecular identity and genetic programming. Nat. Rev. Immunol. 8, 337–348 (2008).
Sakaguchi, S., Yamaguchi, T., Nomura, T. & Ono, M. Regulatory T cells and immune tolerance. Cell 133, 775–787 (2008).
VolinM.V. & Shahrara, S. Role of TH-17 cells in rheumatic and other autoimmune diseases.Rheumatology (Sunnyvale) 1, 2169 (2011).
Peoples, G. E. et al. T lymphocytes that infiltrate tumors and atherosclerotic plaques produce heparin-binding epidermal growth factor-like growth factor and basic fibroblast growth factor: a potential pathologic role. Proc. Natl Acad. Sci. USA 92, 6547–6551 (1995).
Mor, F., Quintana, F. J. & Cohen, I. R. Angiogenesis-inflammation cross-talk: vascular endothelial growth factor is secreted by activated T cells and induces Th1 polarization. J. Immunol. 172, 4618–4623 (2004).
De Falco, S. The discovery of placenta growth factor and its biological activity. Exp. Mol. Med. 44, 1–9 (2012).
Kim, K. J., Cho, C. S. & Kim, W. U. Role of placenta growth factor in cancer and inflammation. Exp. Mol. Med. 44, 10–19 (2012).
Muramatsu, M., Yamamoto, S., Osawa, T. & Shibuya, M. Vascular endothelial growth factor receptor-1 signaling promotes mobilization of macrophage lineage cells from bone marrow and stimulates solid tumor growth. Cancer Res. 70, 8211–8221 (2010).
Dewerchin M. & Carmeliet P. PlGF: a multitasking cytokine with disease-restricted activity. Cold Spring Harb. Perspect. Med 2, a011056 (2012).
Fischer, C. et al. Anti-PlGF inhibits growth of VEGF(R)-inhibitor-resistant tumors without affecting healthy vessels. Cell 131, 463–475 (2007).
Stritesky, G. L., Yeh, N. & Kaplan, M. H. IL-23 promotes maintenance but not commitment to the Th17 lineage. J. Immunol. 181, 5948–5955 (2008).
Xu, L. et al. Placenta growth factor overexpression inhibits tumor growth, angiogenesis, and metastasis by depleting vascular endothelial growth factor homodimers in orthotopic mouse models. Cancer Res. 66, 3971–3977 (2006).
Eriksson, A. et al. Placenta growth factor-1 antagonizes VEGF-induced angiogenesis and tumor growth by the formation of functionally inactive PlGF-1/VEGF heterodimers. Cancer Cell 1, 99–108 (2002).
El-Behi, M. et al. The encephalitogenicity of T(H)17 cells is dependent on IL-1- and IL-23-induced production of the cytokine GM-CSF. Nat. Immunol. 12, 568–575 (2011).
Korn, T. et al. IL-21 initiates an alternative pathway to induce proinflammatory T(H)17 cells. Nature 448, 484–487 (2007).
Ichiyama, K. et al. Transcription factor Smad-independent T helper 17 cell induction by transforming-growth factor-beta is mediated by suppression of eomesodermin. Immunity 34, 741–754 (2011).
Basu, A. et al. Cutting edge: Vascular endothelial growth factor-mediated signaling in human CD45RO + CD4 + T cells promotes Akt and ERK activation and costimulates IFN-gamma production. J. Immunol. 184, 545–549 (2010).
Delgoffe, G. M. et al. Stability and function of regulatory T cells is maintained by a neuropilin-1-semaphorin-4a axis. Nature 501, 252–256 (2013).
Errico, M. et al. Identification of placenta growth factor determinants for binding and activation of Flt-1 receptor. J. Biol. Chem. 279, 43929–43939 (2004).
Mamluk, R. et al. Neuropilin-1 binds vascular endothelial growth factor 165, placenta growth factor-2, and heparin via its b1b2 domain. J. Biol. Chem. 277, 24818–24825 (2002).
Yang, X. O. et al. STAT3 regulates cytokine-mediated generation of inflammatory helper T cells. J. Biol. Chem. 282, 9358–9363 (2007).
Bellik, L., Vinci, M. C., Filippi, S., Ledda, F. & Parenti, A. Intracellular pathways triggered by the selective FLT-1-agonist placental growth factor in vascular smooth muscle cells exposed to hypoxia. Br. J. Pharmacol. 146, 568–575 (2005).
Schust, J., Sperl, B., Hollis, A., Mayer, T. U. & Berg, T. Stattic: a small-molecule inhibitor of STAT3 activation and dimerization. Chem. Biol. 13, 1235–1242 (2006).
Fujino, M. & Li, X. K. Role of STAT3 in regulatory T lymphocyte plasticity during acute graft-vs.-host-disease. JAKSTAT 2, e24529 (2013).
Allen, I. C. Delayed-type hypersensitivity models in mice. Methods Mol. Biol. 1031, 101–107 (2013).
Dong, C. Targeting Th17 cells in immune diseases. Cell Res. 24, 901–903 (2014).
Parsonage, G. et al. Prolonged, granulocyte-macrophage colony-stimulating factor-dependent, neutrophil survival following rheumatoid synovial fibroblast activation by IL-17 and TNFalpha. Arthritis Res. Ther. 10, R47 (2008).
Freeman, M. R. et al. Peripheral blood T lymphocytes and lymphocytes infiltrating human cancers express vascular endothelial growth factor: a potential role for T cells in angiogenesis. Cancer Res. 55, 4140–4145 (1995).
Chung, A. S. et al. An interleukin-17-mediated paracrine network promotes tumor resistance to anti-angiogenic therapy. Nat. Med. 19, 1114–1123 (2013).
Yoo, S. A. et al. Placental growth factor-1 and −2 induce hyperplasia and invasiveness of primary rheumatoid synoviocytes. J. Immunol. 194, 2513–2521 (2015).
Yoo, S. A. et al. Role of placenta growth factor and its receptor flt-1 in rheumatoid inflammation: a link between angiogenesis and inflammation. Arthritis Rheum. 60, 345–354 (2009).
Kang, M. C. et al. Gestational loss and growth restriction by angiogenic defects in placental growth factor transgenic mice. Arterioscler. Thromb. Vasc. Biol. 34, 2276–2282 (2014).
Tarallo, V., Tudisco, L. & De Falco, S. A placenta growth factor 2 variant acts as dominant negative of vascular endothelial growth factor A by heterodimerization mechanism. Am. J. Cancer Res. 1, 265–274 (2011).
Yoo, S. A. et al. A novel pathogenic role of the ER chaperone GRP78/BiP in rheumatoid arthritis. J. Exp. Med. 209, 871–886 (2012).
Bolstad, B. M., Irizarry, R. A., Astrand, M. & Speed, T. P. A comparison of normalization methods for high density oligonucleotide array data based on variance and bias. Bioinformatics 19, 185–193 (2003).
Lee, H. J. et al. Direct transfer of alpha-synuclein from neuron to astroglia causes inflammatory responses in synucleinopathies. J. Biol. Chem. 285, 9262–9272 (2010).
Hwang, D. et al. A data integration methodology for systems biology. Proc. Natl Acad. Sci. USA 102, 17296–17301 (2005).
Yu, G. C., Wang, L. G., Han, Y. Y. & He, Q. Y. ClusterProfiler: an R package for comparing biological themes among gene clusters. Omics 16, 284–287 (2012).
Hwang, S. H. et al. Leukocyte-specific protein 1 regulates T-cell migration in rheumatoid arthritis. Proc. Natl Acad. Sci. USA 112, E6535–E6543 (2015).
Arnett, F. C. et al. The American Rheumatism Association 1987 revised criteria for the classification of rheumatoid arthritis. Arthritis Rheum. 31, 315–324 (1988).
We thank the members of the Center for Integrative Rheumatoid Transcriptomics and Dynamics at the Catholic University of Korea for their assistance. This work was supported by grants the National Research Foundation of Korea (NRF) funded by the Ministry of Science and ICT (grant no. 2015R1A3A2032927) to W.U.K. and (grant nos. 2014R1A6A3A04054066 and 2019R1A2C2010897) to S.A.Y., respectively, and by Project SATIN-POR Campania FESR 2014/2020 to S.D.F.
The authors declare no competing interests.
Peer review information: Zoltan Fehervari was the primary editor on this article and managed its editorial process and peer review in collaboration with the rest of the editorial team.
Publisher’s note: Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Integrated supplementary information
(a) VEGF production by sorted CD4+ and CD8+ T cells (n = 6). After mouse T cells were stimulated with anti-CD3 and anti-CD28 (α-CD3/28), PMA (50 ng/ml) plus ionomycin (IONO, 5 μg/ml), LPS (1 μg/ml), or IL-1β (10 ng/ml) for 48 hours, the levels of VEGF (VEGF164 plus VEGF120) in the culture supernatants were measured using an ELISA. (b and c) VEGF and PlGF production by TH1, TH2 and TH17 cells. CD4+ T cells from WT mice (n = 3) were activated with anti-CD3 and anti-CD28 (1 μg/ml) under TH1 cell, TH2 cell, and TH17 cell-polarizing conditions (with IL-23) for 5 days using a protocol modified from the standard T cell polarization protocol described in the Methods section of the present study. VEGF, PlGF, IL-17, and IFN-γ concentrations in the culture supernatants were determined using ELISAs (b). Flow cytometry analysis was performed under the same conditions to confirm the number of IFN-γ+TBET+ cells produced under TH1-polarizing conditions, IL-4+GATA3+ cells produced under TH2-polarizing conditions, and IL-17+RORC+ cells produced under TH17-polarizing conditions (c). Data are presented as the mean and SEM of three independent experiments.
(a) T cells from Plgf-Tg mice (n = 3) have higher levels of PlGF expression than WT cells (n = 3). Splenic CD4+ T cells from WT and Plgf-Tg mice were stimulated with anti-CD3 and anti-CD28 (α-CD3/28, 1 μg/ml). After 16 hours, levels of PlGF mRNA were measured by quantitative real-time PCR; level of Gapdh mRNA was used as an internal control (left panel). Concentrations of PlGF in the culture supernatants also were determined using an ELISA 24 or 72 hours after stimulation of CD4+ T cells (1 × 106) of wild-type (WT) and Plgf-Tg‘ mice with α-CD3/28 (right panel). PlGF production by TH17 cells is shown for comparison (See Fig. 1d). * P < 0.05. (b) T-cell–derived PlGF increases tube formation by endothelial cells. CD4+ T cells (5 × 105) from WT (n = 5) and PlGF transgenic (Tg) mice (n = 5) were cultured without or with α-CD3/28 (1 μg/ml) for 48 hours; culture supernatants from the T cells were then added to mouse endothelial cells (H5V cells, 8 × 104) seeded on Matrigel-coated plates for 12 hours. The total length of the tube network was calculated using ImageJ software. * P < 0.05. (c) PlGFCM specifically increases capillary tube formation. H5V cells were incubated with conditioned media containing PlGF-2 (PlGFCM, 100 ng/ml) in the absence or presence of neutralizing antibody against PlGF (α-PlGF Ab) for 12 hours. Isotype control Ab (Con Ab) was used as a control. * P < 0.05 compared with isotype control Ab. (d) CD4+ T cells (5 × 105) from WT (n = 4), Plgf-KO (n = 4), and Plgf-Tg mice (n = 4) were stimulated with anti-CD3 and anti-CD28 (1 μg/ml) for 48 hours. The extent of capillary tube formation was assessed 12 hours after the addition of culture supernatants of TCR-stimulated CD4+ T cells to endothelial cells. (e) Increase in migration of endothelial cells induced by T cell-secreted PlGF in the wound migration assay. Confluent H5V cells were incubated with culture supernatants from CD4+ T cells (activated with α-CD3/28) from WT (n = 4) or Plgf-Tg mice (n = 4), or PlGFCM (100 ng/ml) vs ConCM, for 24 hours. Cells migrating beyond the reference line were photographed and counted. * P < 0.05 compared with WT CD4+ T cells or ConCM. Scale bars = 100 μm. (f) PlGFCM specifically increases endothelial chemotaxis. H5V cells (5 × 104) were loaded into the upper wells of a Boyden chamber. The lower wells were supplied with conditioned media containing PlGF-2 (PlGFCM, 100 ng/ml) in the absence or presence of α-PlGF Ab. After 12 hours, the migrated cells were stained violet using a Diff-Quik kit. * P < 0.05 compared with control Ab. Scale bars = 100 μm. (g) PlGFCM increases angiogenesis in vivo. Matrigels (500 μl) containing ConCM, PlGFCM (200 ng/ml) or recombinant VEGF (100 ng/ml) were subcutaneously injected into C57BL/6N mice (n = 12). After 7 days, the gels were excised and the hemoglobin content of the gels was determined using Drabkin’s reagent (bottom panel). Representative gels are shown in the top panel. * P < 0.05 compared with ConCM. The data shown in (a) through (f) are representative or mean±SEM of more than two independent experiments.
Supplementary Fig. 3 PlGF does not affect IFN-γ and IL-10 production, T cell proliferation, or cellular composition of splenocytes.
(a) IFN-γ and IL-10 production by splenocytes from Plgf-KO (n = 6) and WT mice (n = 10). Mouse splenocytes (5 × 105) were stimulated with anti-CD3 and anti-CD28 (α-CD3/28, 1 μg/ml) for 48 hours. The levels of IFN-γ and IL-10 in the culture supernatants were measured using ELISAs. (b) CD4+ T cells (1 × 105) of WT and Plgf-KO mice were differentiated under TH1, TH2 or TH17 polarizing conditions for 5 days (see Methods). The concentrations of IL-17 and IFN-γ in the culture supernatants were determined by an ELISA. * P < 0.05 versus TH17 cells of Plgf-KO mice. (c) Splenocyte proliferation in response to TCR stimulation. Splenocytes from Plgf-KO (n = 8) and WT mice (n = 8) were stimulated with α-CD3/28 for 48 hours. Cell proliferation was measured using the [3H] thymidine incorporation assay. (d) Immune cell compositions of Plgf-KO (n = 2) and WT splenocytes (n = 2). Splenocytes from these mice were stained with APC-labeled anti-CD4, PE-labeled anti-CD8, FITC-labeled anti-CD19, and FITC-labeled anti-CD11. Cells were analyzed using flow cytometry. (e) No effect of exogenous PlGFCM on production of IFN-γ by T cells. Sorted CD4+ T cells (1 × 105, n = 3) were differentiated under TH1, TH2, and TH17-polarizing conditions with exogenous PlGFCM (100 ng/ml) for 5 days. Concentrations of IFN-γ in the supernatants were analyzed by ELISA. Data are presented as the mean and SEM of three independent experiments performed in triplicate.
Supplementary Fig. 4 PlGF-induced increase in IL-17 production and p-STAT3 activation via FLT1 and NRP1 receptor.
(a) Downregulation of PlGF expression in CD4+ T cells by siRNA. CD4+ T cells were transfected with siRNAs for Flt1 or Nrp1 using a T-cell nucleofector kit. After 24 hours, expressions of Flt1 and Nrp1 mRNA were determined by real-time PCR. * P < 0.05 compared to control siRNA (Con). (b) Sorted CD4+ T cells (n = 16) were cultured in RPMI 1640 with 10% FBS and stimulated with PlGFCM (100 ng/ml) for 72 hours in the absence or presence of soluble FLT1 (sFLT1, 10 μg/ml) and soluble NRP1 (sNRP1, 10 μg/ml). Levels of IL-17 in culture supernatants were measured by ELISA. * P < 0.05 compared to cells incubated with PlGFCM but not sFLT1 or sNRP1. Data presented in (a), and (b) are the mean and SEM of more than three independent experiments. (c) CD4+ T cells were incubated with ConCM or PlGFCM (100 ng/ml) for the indicated times. Phosphorylated STAT3 expression was measured using flow cytometry. Presented data are from two independent experiments with similar results.
(a) Recombinant PlGF-induced increase in IL-17 secretion from T cells. Splenic CD4+ T cells (1 × 106) from WT mice maintained in conventional facilities were stimulated with α-CD3/28 (1 μg/ml) plus recombinant PlGF (rPlGF; Sino Biological Inc., Wayne, PA, USA) in the absence or presence of the anti-PlGF antibody (α-PlGF Ab, 50 μ/ml) for 48 hours. IL-17 levels in the culture supernatants were measured using an ELISA. *P < 0.05, †P < 0.05 compared with PlGF 200 ng/ml alone. (b) Increase in p-STAT3 levels by recombinant PlGF. Splenic CD4+ T cells were incubated with recombinant PlGF (200 ng/ml, Sino Biological Inc.) in the absence or presence of α-PlGF Ab (50 μ/ml) for 1 hour. The levels of p-STAT3 were assessed by immunoblotting. (c) Upregulation of Rorc expression by recombinant PlGF. Sorted CD4+ T cells were stimulated with α-CD3/28 plus recombinant PlGF (Sino Biological Inc.) for 12 hours. Levels of the Rorc mRNA were determined by quantitative real-time PCR. Gapdh mRNA was used as an internal control. Fold inductions were calculated using the 2-ΔΔCt method. * P < 0.05. (d) Production of PlGF-2-Fc in HEK-293T cells. The Fc-fused PlGF-2-overexpressing vectors for the production of PlGF-2-Fc in HEK-293T cells were constructed. Mouse or human Fc-fused PlGF-2 was produced using transient gene expression with serum-free suspended HEK-293T cells (see the Methods section). Purified Fc-tagged PlGF-2 protein was analyzed on BoltTM 4–12% Bis-Tris Plus Gels under nonreduced and reduced conditions. Lane 1: non-reduced homodimeric form of human Fc-fused PlGF-2 (arrow), lane 2: reduced monomeric form of human Fc-fused PlGF-2 (arrowhead), lane 3: non-reduced homodimeric form of mouse Fc-fused PlGF-2 (arrow), and lane 4: reduced monomeric form of mouse Fc-fused PlGF-2 (arrowhead). (e and f) Increase in IL-17 production and Rorc expression in CD4+ T cells by the PlGF-2 that we isolated and purified from -overexperssed HEK-293T cells, determined by ELISA and quantitative real-time PCR, respectively. * P < 0.05.
Supplementary Fig. 6 Effects of exogenous PlGFCM on expression of Foxp3, Tbx21, and Gata3 under helper T cell-polarizing conditions.
Resting CD4+ T cells (2 × 105, n = 3) were differentiated into TH1, TH2, TH17, or Treg cells in the absence or presence of PlGFCM (100 ng/ml) for 5 days. Levels of the Foxp3 (a), Tbx21 (b), and Gata3 (c) mRNAs were measured by quantitative real-time PCR. The level of Gapdh mRNA was used as an internal control. Fold inductions were calculated using the 2-ΔΔCt method. Data are presented as the mean and SEM of three independent experiments. * P < 0.05 compared with ConCM.
(a) Mean clinical scores and incidence of EAE in Rag1-KO mice. Prior to EAE induction, CD4+ T cells were adoptively transferred from WT mice to Rag1-KO mice (WT → Rag1-KO, n = 7) or from Plgf-KO mice to Rag1-KO mice (Plgf-KO → Rag1-KO, n = 8). * P < 0.05. (b) IL-17 and IFN-γ measurements. Cells were collected from the spleens and lymph nodes of WT → Rag1-KO mice (n = 5) and Plgf-KO → Rag1-KO mice (n = 8) on day 35 after EAE induction and stimulated with anti-CD3 and anti-CD28 for 72 hours. Levels of IL-17 and IFN-γ in culture supernatants were measured using ELISAs. * P < 0.05. (c) The frequency of IL-17 and IFN-γ-producing CD4+CD44+ T cells in the central nervous systems (CNS), lymph nodes (LN), and spleens of WT → Rag1-KO mice (n = 6) and Plgf-KO → Rag1-KO mice (n = 6) was determined by flow cytometry at 3 weeks after the primary immunization. *P < 0.05. Representative data are shown on the left. The data in a through c are presented as the mean and SEM.
Supplementary Fig. 8 Model of the PlGF-induced self-perpetuating cycle of chronic inflammation and TH17 cell generation.
Following the formation of new vessels, T cells migrate to inflamed sites and are activated by pre-existing endogenous or exogenous antigens. TCR-activated T cells then secrete PlGF, which in turn may further increase angiogenesis and thereby recruit more T cells and other inflammatory cells into the sites, establishing a vicious cycle of chronic inflammation and the amplification of the initial angiogenic signal. ′Angio-lymphokine′ PlGF also promotes TH17 cell differentiation and conversion to TH17 cells from other T cell subtypes. Resultant TH17 cells produce more PlGF, creating a positive feedback loop between PlGF and IL-17 productions. In these self-perpetuating cycles, TCR activation by antigenic challenge appear to be a prerequisite since PlGF alone was ineffective in IL-17 protein production (Fig. 1).
About this article
Nature Reviews Immunology (2019)