Attenuation of Graft Arterial Disease by Manipulation of the LIGHT Pathway
Objective— The tumor necrosis factor (TNF) superfamily member LIGHT, which binds herpes virus entry mediator (HVEM) and lymphotoxin β receptor (LTβR), plays important roles in regulating the immune response. To clarify the mechanism underlying graft arterial disease (GAD), we investigated the role of the LIGHT pathway in the progression of GAD.
Methods and Results— Hearts from Bm12 mice were transplanted into C57BL/6 (B/6) mice (class II mismatch). Recipients were injected intraperitoneally with HVEMIg (100 μg per treatment) every 7 days for 8 weeks. Treatment with HVEMIg significantly attenuated GAD (luminal occlusion=16.5±7.7% versus control allograft=62.6±12.1%, P<0.05), and significantly decreased intragraft IL-4, IL-6, and interferon-γ (IFN-γ) mRNA expression compared with controls. LTβR was expressed in smooth muscle cells (SMCs) with or without cytokine stimulation, whereas HVEM was detected in SMCs stimulated by IFN-γ. Coculture of SMCs with T cells after transplantation induced SMC proliferation, and addition of HVEMIg resulted in inhibition of SMC proliferation.
Conclusions— These results indicate that the LIGHT pathway plays important roles in the regulation not only of T-cell activation but also of SMC proliferation. Blockade of the LIGHT pathway is a promising avenue for the prevention of GAD.
New immunosuppressive agents have significantly improved early survival after organ transplantation. However, graft arterial disease (GAD) is the major cause of graft failure, threatening the long-term survival of organ transplant recipients.1,2 GAD is diffuse and concentric intimal thickening composed of T cells, macrophages, and smooth muscle cells (SMCs). Although the pathogenesis of GAD remains unclear, multiple factors are thought to be involved.3 In the immune response, 2 distinct signals are necessary for efficient activation of T cells. The first signal is delivered from T-cell receptors, and the second signal is delivered from costimulatory molecules such as CD28. Elimination of signals of the B7-CD28 pathway not only prolongs allograft survival but also attenuates the development of GAD.3,4
A role of CD28-independent pathways such as those of the tumor necrosis factor (TNF) superfamily has also been reported. LIGHT (homologous to lymphotoxins, exhibits inducible expression and competes with herpes simplex virus glycoprotein D for herpes virus entry mediator [HVEM], a receptor expressed by T lymphocytes) was recently described as a member of the TNF superfamily.5 LIGHT is expressed in peripheral blood mononuclear cells, including T and B cells, natural killer cells, monocytes, and granulocytes,5,6 and binds to HVEM, lymphotoxin β receptor (LTβR), and TR6/DcR3.5,7 Although LTβR is not expressed in T or B cells, HVEM is expressed in lymphocytes and endothelial cells.8–10 In vitro studies showed that LIGHT–HVEM interaction is involved in T-cell proliferation, cytokine production, and stimulation of NF-κB activation.11–13
Recent studies also showed that signaling through the LIGHT pathway plays an important role in vivo. Blockade of the LIGHT pathway leads to regression of tumors and amelioration of graft-versus-host disease.11 In a murine cardiac transplantation model, LIGHT-deficient recipients showed prolonged allograft survival.14 However, it has not been reported whether the LIGHT pathway is involved in the development of GAD. We hypothesized that the LIGHT pathway is important in regulating not only acute rejection but also development of GAD in organ transplant recipients. Here, we show that blockade of the LIGHT pathway significantly attenuates the development of GAD and also suppresses of SMC proliferation in cardiac allografts.
C57BL/6 (B/6, H-2b) mice aged 6 to 8 weeks were obtained from Japan Clea (Tokyo, Japan). B6.C-H-2bm12KhEg (Bm12, H-2bm12) mice were obtained from the Jackson Laboratory (Bar Harbor, Me). Animals were maintained in our animal facility. The present study conformed to the Guide for the Care and Use of Laboratory Animals of Tokyo Medical and Dental University.
Preparation of HVEMIg
To prepare a soluble form of HVEM (HVEMIg), an adenovirus vector containing a chimeric gene that encodes a fusion protein consisting of an extracellular domain of murine HVEM and the Fc portion of human IgG1 was constructed. The cDNA of the extracellular domain of HVEM was amplified by reverse-transcription polymerase chain reaction (RT-PCR), and an mRNA isolated from concanavalin A-stimulated splenocytes obtained from a BALB/c mouse was used as the template. The PCR primers used to amplify the cDNA were 5′-TAACTCGAGCTCTTGGCCTGAAGTTTC-3′ and 5′-TTAAGGATCCGAGGAGCAGGTGGTGTCT-3′.
The cDNA was inserted into the XhoI and BamHI sites of a plasmid carrying IgG1-Fc DNA,15 and the sequence was verified. The plasmid was digested with XhoI/XbaI to obtain the HVEMIg DNA fragment. The fragment was blunt-ended and inserted into a SwaI site of the pAxCAwt cosmid vector (Takara, Kyoto, Japan) for preparation of recombinant adenovirus AxHVEMIg. The recombinant adenovirus was prepared with an Adenovirus Expression Vector Kit (Takara) according to the protocol supplied by the manufacturer. HVEMIg protein was purified from the supernatant of AxHVEMIg-infected COS7 cells, as described previously.16
Heterotopic cardiac transplantation was performed as described previously.17 Graft function was evaluated by daily palpation. Recipient mice were injected intraperitoneally with HVEMIg (n=5) or isotype-matched control IgG (human IgG; Jackson ImmunoResearch Laboratories, West Grove, Pa; n=5) every 7 days for 8 weeks after transplantation (100 μg per injection).
Cardiac allografts of combinations of B/6 and Bm12 (class II mismatch) functioned normally for at least 8 weeks without immunosuppression. Allografts were harvested at 8 weeks after transplantation. Harvested allografts were sectioned transversely into 3 parts. The basal section was fixed in 10% formalin and embedded in paraffin for elastic fiber stain. The middle section was embedded immediately in OCT compound and flash-frozen in liquid nitrogen (LN2) for immunohistochemistry. The apical section was used to extract RNA for ribonuclease protection assay (RPA).
Frozen sections (5 μm) were fixed in acetone for 10 minutes at 4°C. To reduce nonspecific binding, sections were incubated with 10% normal rabbit serum at room temperature. To stain for HVEM, LTβR, LIGHT, CD4+, or CD8+ T cells, sections were incubated with goat IgG (Santa Cruz Biotechnology Inc, Santa Cruz, Calif) or antibodies against HVEM, LTβR, LIGHT (Santa Cruz Biotechnology Inc), CD4+, CD8+ T cells (Pharmingen, San Diego, Calif) overnight at 4°C. After a washing in phosphate-buffered saline, sections were incubated with biotinylated secondary antibodies at room temperature for 30 minutes. Antigen-antibody conjugates were detected with avidin-biotin-horseradish peroxidase complex (Nichirei, Tokyo, Japan) according to the manufacturer’s instructions. The complex was visualized with 3-amino-9-ethylcarbazole chromogen. Sections were counterstained with hematoxylin. Cell numbers were quantified by counting stained lymphocytes in 20 fields per graft.
Immunofluorescence double staining was performed to examine HVEM or LTβR expression on SMCs. After incubation with first (anti-HVEM antibody or anti-LTβR antibody) and biotinylated secondary antibody, sections were stained with fluorescein isothiocyanate (FITC)-conjugated anti-αSMA antibody (Pharmingen) and avidin-Texas red (Pharmingen). Sections were observed under confocal microscopy.
Grafts were analyzed by elastica van Gieson stains. All elastin-positive vessels in each section were evaluated.18 The lumen and the internal elastic lamina were carefully traced, and planimetric areas were calculated by an image analysis system (Scion Image beta 4.0.2). Cross-sectional area luminal stenosis was calculated as luminal occlusion=(internal elastic lamina area − luminal area)/internal elastic lamina area ×100 (%).
mRNA isolation was performed with TRIzol (Life Technologies, Rockville, Md) according to the manufacturer’s protocol. In vitro transcription was performed with the template set, T7 polymerase, and [α-32P]UTP. Ten micrograms of total RNA was hybridized with probes at 56°C for 16 hours. All samples were then treated with RNase before treatment with proteinase K. Samples were electrophoresed on denaturing gels containing 5% polyacrylamide. Detection of the mRNA bands was performed with an image analyzer (BAS2000; Fujifilm). mRNA expression levels were quantified and standardized against the expression levels of GAPDH. Standardized mRNA expression levels in the control IgG-treatment groups were expressed as 1.0.19,20
Primary SMCs were obtained from the thoracic aortas of Bm12 mice by an explant technique. Cells were grown in Dulbecco modified Eagle medium (Sigma Chemical Co, St. Louis, Mo) containing 50 μg/mL streptomycin, 50 IU/mL penicillin, and 10% fetal bovine serum at 37°C and 5% CO2.21 Cultured SMCs were identified by the typical hill-and-valley morphology and immunostaining with monoclonal antibody to α-smooth muscle actin. All experiments were performed with cells between passages 3 and 8.
SMCs were trypsinized and seeded onto 96-well plates. At confluence, SMCs were arrested in medium with 0.4% fetal bovine serum for 5 days, and then recombinant mouse IL-4, IL-6, or IFN-γ (10 ng/mL each) was added to each well. After 24 hours, cells were incubated with BrdU (Roche, Mannheim, Germany) according to the manufacturer’s instructions. Incorporated BrdU was measured by a microplate-imaging system (BioRad, Hercules, Calif).
Separation of T Cells
Splenocyte suspensions were obtained by disrupting spleens between sterile glass slides. Red blood cells were lysed by ammonium chloride lysis. Cells were washed, and T cells were isolated by magnetic cell sorting according to the manufacturer’s instructions (Miltenyi Biotec, Bergisch Gladbach, Germany). The purity of separated T cells was consistently >95% as assessed by flow cytometry.
Coculture of SMCs and T cells
Trypsinized SMCs were seeded onto 96-well plates. At confluence, SMCs were arrested in medium with 0.4% fetal bovine serum for 5 days. SMCs were then stimulated with IFN-γ (10 ng/mL) for 48 hours. After washing in phosphate-buffered saline, mitomycin-C–inactivated T cells (total 5×105) and HVEMIg (10 μg/mL) were added. SMC proliferation was measured as described.
Total RNA was extracted from SMCs after 24-hour IL-4, IL-6, or IFN-γ stimulation. cDNA was prepared by reverse transcription of 5 μg RNA. cDNA (10 μL) was amplified according to the following parameters : 94°C, 1 minute; 58°C, 1 minute; 72°C, 1 minute; for 30 cycles. Primers for LIGHT, HVEM, LTβR, LTα, and β-actin were as follows: for LIGHT, 5′-GCTTTCTGGGTTTTGAGCTG-3′ and 5′-GGTGCTCAGAAGCTCGTACC-3′; for HVEM, 5′-GTGTCATCCTTTTGCCACT-3′ and 5′-CAGTTGGAGGCTGTCTCCTC-3′; for LTβR, 5′-TTATCGCATAGAAAACCAGACTTGC-3′ and 5′-TCAAAGCCCAGCACAATGTC-3′; for LTα, 5′-CACGAGGTCCAGCTCTTTTC-3′ and 5′-ACCCTTGAAACAACGGTCAG-3′; and for β-actin, 5′-AACTGGGACGACATGGAGAA-3′ and 5′-CATGAGGTAGTCTGTCAGGT-3′.
PCR products were analyzed by ethidium bromide staining of 1.5% agarose gels.
Fluorescence-Activated Cell Sorter Analysis
Graft infiltrating cells were isolated from allografts at 8 weeks after transplantation, as described previously.17 Cells were incubated with anti-LIGHT antibody and stained with biotinylated isotype-matched control IgG or goat IgG at 4°C for 20 minutes. Cells were stained with FITC-conjugated anti-CD4 or anti-CD8 antibody (Pharmingen) and streptavidin-PE at 4°C for 20 minutes.
After arrested SMCs were stimulated with IL-4, IL-6, or IFN-γ (10 ng/mL) for 48 hours, SMCs were incubated with 10 μg/mL anti-HVEM antibody and anti-LTβR antibody, and stained with biotinylated isotype-matched control IgG or goat IgG at 4°C for 20 minutes. Cells were then stained with FITC-conjugated anti-αSMA antibody and streptavidin-PE at 4°C for 20 minutes. Then the cells were analyzed by flow cytometry on a FACSCalibur (Becton Dickinson).
All data are expressed as mean±SEM. Groups of data were compared by 1-way ANOVA. P<0.05 was considered statistically significant.
Expression of HVEM, LTβR, and LIGHT in Cardiac Allografts
To explore whether the LIGHT pathway is involved in the development of GAD, we evaluated the expression of HVEM, LTβR, and LIGHT in cardiac allografts by immunohistochemistry. LIGHT was detected in graft-infiltrating cells in the cardiac allografts at 8 weeks after transplantation; it was not detected in native hearts (Figure IA, available online at http://atvb.ahajournals.org). These cells were positive for LIGHT in CD4+ and CD8+ cells (Figure IB). Interestingly, LTβR was expressed on vascular SMCs in native hearts. However, HVEM and LIGHT were not expressed on SMCs. Furthermore, HVEM and LTβR were detected on SMCs in cardiac allografts at 2 weeks after transplantation (Figures I and II, available online at http://atvb.ahajournals.org).
The cellular infiltrate was examined immunohistochemically in allografts at 2 weeks after transplantation. The number of infiltrating CD4+ and CD8+ T cells in the HVEMIg-treated group was significantly reduced in comparison to that of the control IgG-treated group (Figure IC and ID).
Prevention of GAD by Treatment With HVEMIg
We attempted to determine whether blockade of the HVEM pathway attenuates GAD. Elastic fiber staining of tissue sections showed that severe GAD developed in allografts from control IgG-treated mice at 8 weeks (Figure 2A). However, allografts treated with HVEMIg at 8 weeks after transplantation showed significantly less GAD (Figure 2B). Elastin-positive vessels were used to evaluate the severity of luminal stenosis. The mean luminal occlusion in mice treated with HVEMIg (16.5±7.73%) was significantly less than that in untreated mice (62.6±12.1%, P<0.02) (Figure 2C).
Suppression of Cytokine mRNA in Cardiac Allografts Treated With HVEMIg
The expression of multiple cytokines is altered in the development of GAD. To examine cytokine expression in cardiac allografts, we performed RPA on cardiac allografts at 2 and 8 weeks after transplantation. The expression of IL-4 and IFN-γ mRNAs was suppressed in HVEMIg-treated allografts at 2 weeks after transplantation (data not shown). mRNAs for IL-4, IL-10, IL-15, IL-6, and IFN-γ were detected in control IgG-treated allografts at 8 weeks after transplantation (Figure 3, left). However, treatment with HVEMIg significantly suppressed the expression of IL-4, IL-6, and IFN-γ mRNAs at 8 weeks after transplantation (Figure 3, right; Figure III, available online at http://atvb.ahajournals.org; P<0.05).
Proliferation of SMCs in Response to IL-4, IL-6, and IFN-γ
SMCs were isolated from the thoracic aorta and treated with various cytokines. SMCs treated with IL-4 showed significant proliferation in comparison to untreated cells (Figure 4A). Similarly, IL-6 and IFN-γ enhanced the proliferation of SMCs (Figure 4B and 4C).
Expression of HVEM and LTβR by SMCs
Because SMCs express HVEM and LTβR (Figure 1), we examined the expression of these molecules in vitro using RT-PCR. Consistent with immunohistological findings, LTβR was expressed in untreated cells and SMCs stimulated by IL-4, IL-6, and IFN-γ (Figure 5A and 5C). HVEM expression was induced in SMCs treated with IFN-γ (Figure 5B and 5C); it was not detected in untreated cells (Figure 5). IL-4 and IL-6 treatment did not induce expression of HVEM (data not shown). LIGHT and LTα were not detected in untreated SMCs (Figure 5A) or in response to IL-4, IL-6, or IFN-γ (data not shown).
Proliferation of SMCs by Interaction With T Cells
On the basis of expression of HVEM and LTβR by SMCs (Figure 1), we examined whether SMC proliferation is involved in the LIGHT pathway. Naïve T cells did not stimulate proliferation of unstimulated SMCs and SMCs in response to IFN-γ (Figure 6A), IL-4, or IL-6 (data not shown). Chemokine-unstimulated SMCs also did not proliferate in response to the addition of T cells from mice than underwent transplantation (Figure 6B). However, coculture of SMCs stimulated by IFN-γ and activated T cells after transplantation induced significant proliferation of SMCs, and addition of HVEMIg to the coculture suppressed SMC proliferation (Figure 6B).
GAD decreases the survival time in patients after organ transplantation. Therefore, it is important to prevent the development of GAD. In this study, we showed conclusively that the LIGHT pathway plays a pivotal role in the proliferation of SMCs and intimal thickening in allografts and that the blockade of this pathway prevents GAD development by suppressing cytokine expression and attenuating SMC proliferation.
We confirmed that LIGHT is expressed by CD4+ and CD8+ T cells in allografts and that blockade of the LIGHT pathway with HVEMIg significantly reduced the development of GAD. It has been reported that costimulatory molecules are involved in the development of GAD. Treatment with CTLA4Ig attenuated GAD development in a rat cardiac transplantation model.3 Blockade of the CD40/CD154 pathway by the generation of CD154-deficient mice did not prevent the development of GAD;19 however, long-term administration of anti-CD154 monoclonal antibody suppressed GAD development.22 Although the mechanism of interaction between GAD and costimulatory molecules remains to be determined, there are 2 possible means by which blockade of the HVEM pathway prevents GAD development. One involves the prevention of interaction of SMCs and T cells, and the other involves the suppression of cytokine expression.
LIGHT is expressed in activated lymphocytes, natural killer cells, and immature dendritic cells,5,6,12 and HVEM is expressed in monocytes, B cells, resting and activated T cells, and endothelial cells.8–10 LTβR is found in monocytes and stromal cells but is absent in T and B lymphocytes.8 Although CD154 and CD40 are shown to be expressed constitutively by SMCs in vitro and CD154 is induced by stimulation with proinflammatory cytokines,23 expression of LIGHT, HVEM, and LTβR by SMCs has not been shown. We, for the first time to our knowledge, found that SMCs in cardiac allografts after transplantation expressed HVEM and LTβR as shown by immunohistochemical staining, although SMCs in native hearts express only LTβR. LIGHT was not expressed by SMCs in native hearts or rejected cardiac allografts. Consistent with the immunohistochemical staining, RT-PCR showed that LTβR mRNA alone was expressed in unstimulated and stimulated SMCs, and HVEM mRNA was increased in response to treatment with IFN-γ. LIGHT mRNA and LTα mRNA were not detected in untreated and stimulated SMCs.
To examine the effect of the interaction between SMCs and T cells, we studied SMC proliferation after coculture with T cells. Naïve T cells do not express LIGHT, whereas activated T cells do.6 Naïve T cells did not stimulate proliferation of untreated or IFN-γ–treated SMCs. However, activated T cells did induce proliferation of untreated SMCs. Moreover, they significantly induce proliferation of IFN-γ–treated SMCs. Treatment with HVEMIg suppressed this proliferation. These results indicate that SMCs induce HVEM expression in response to cytokine stimulation and interaction with activated T cells and that signaling through the LIGHT pathway induces the proliferation of SMCs. Immunohistochemical studies have shown that the number of infiltrating T cells was reduced in allografts treated with HVEMIg. This result shows that treatment with HVEMIg may have a significant effect on T-cell activation and recruitment. However, it is very important to suppress interaction of T cell with SMCs to prevent SMC proliferation, because there are infiltrating T cells in allografts treated with control IgG and HVEMIg.
To examine the second potential mechanism, we examined cytokine expression in cardiac allografts by RPA. T cells induce production of Th cell types 1 and 2 (Th1 and Th2) cytokines, and these cytokines regulate the immune response, inflammation, and SMC proliferation. IFN-γ–deficient recipients or treatment with anti-IFN-γ monoclonal antibody prevented GAD,24 indicating that IFN-γ, which is a typical Th1 cytokine, contributes to the development of GAD. IL-4 neutralization with anti-IL-4 monoclonal antibody also resulted in the prevention of GAD.25 However, cardiac allografts from IL-4 knockout recipients showed concentric vascular thickening.26 In addition, exogenous IL-10 exacerbated GAD.27 Although Th2 cytokines such as IL-4 and IL-10 downregulate the production of Th1 cytokines and may be involved in graft survival and tolerance, it is not clear whether Th2 cytokines play a role in the prevention of GAD. A previous study showed that cardiac allografts from LIGHT-deficient recipients that had been treated with cyclosporin A suppressed the upregulation of cytokine mRNAs such as those for IFN-γ, IL-2, and IL-10.14 Scheu et al also showed that LIGHT-deficient splenocytes had reduced levels of IFN-γ, IL-2, IL-4, and IL-10 in mixed lymphocyte reaction.28 In this study, we found that IFN-γ, IL-4, and IL-6 mRNAs were suppressed in cardiac allografts from mice treated with HVEMIg at 8 weeks after transplantation compared with those in allografts from mice treated with control IgG. This suggests that both Th1 and Th2 cytokines may be involved in GAD development. Because the development of intimal hyperplasia after transplantation involves SMC proliferation, we examined whether the proliferation of SMCs by cytokines is reduced after treatment. IFN-γ, IL-4, and IL-6 stimulated proliferation of SMCs in vitro, as described previously.29,30 This result indicates that blockade of the LIGHT pathway suppresses the expression of IFN-γ, IL-4, and IL-6, and consequently prevents the proliferation of SMCs.
In conclusion, the present findings indicate that the LIGHT pathway participates in SMC proliferation and cytokine expression. Therefore, blockade of the LIGHT pathway attenuates the development of GAD through suppression of cytokine expression and SMC proliferation. Manipulation of the LIGHT pathway may have clinical potential for the prevention of GAD.
This study was supported by grafts from the Japan Cardiovascular Research Foundation and the Vehicle Racing Commemorative Foundation, grants-in-aid from the Japanese Ministry of Education, Culture, Sports, Science, and Technology, and the Ministry of Health, Labor and Welfare. We thank Noriko Tamura for excellent technical assistance.
- Received February 10, 2004.
- Accepted April 30, 2004.
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