Journal of Molecular and Cellular Cardiology 72 (2014) 85–94

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Journal of Molecular and Cellular Cardiology

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Original article

TNF-α promotes early atherosclerosis by increasing transcytosis of LDLacross endothelial cells: Crosstalk between NF-κB and PPAR-γ

Youzhi Zhang 1, Xiaoyan Yang 1, Fang Bian, Pinhui Wu, Shasha Xing, Gao Xu, Wenjing Li, Jiangyang Chi,Changhan Ouyang, Tao Zheng, Dan Wu, Yonghui Zhang, Yongsheng Li, Si Jin ⁎Department of Pharmacology, Tongji Medical College, Huazhong University of Science and Technology, Wuhan, ChinaThe Key Laboratory of Drug Target Research and Pharmacodynamic Evaluation of Hubei Province, Wuhan, China

⁎ Corresponding author at: Department of PharmacHuazhong University of Science, Wuhan 430030, China. T

E-mail address: Jinsi@mail.hust.edu.cn (S. Jin).1 These authors contributed equally to this work.

http://dx.doi.org/10.1016/j.yjmcc.2014.02.0120022-2828/© 2014 Elsevier Ltd. All rights reserved.

a b s t r a c t

a r t i c l e i n f o

Article history:Received 3 August 2013Received in revised form 21 February 2014Accepted 24 February 2014Available online 2 March 2014

Keywords:LDLTNF-αNF-κBPPAR-γAtherosclerosis

Tumor necrosis factor-α (TNF-α) is an established pro-atherosclerotic factor, but the mechanism is notcompletely understood. We explored whether TNF-α could promote atherosclerosis by increasing thetranscytosis of lipoproteins (e.g., LDL) across endothelial cells and how NF-κB and PPAR-γ were involved inthis process. TNF-α significantly increased the transcytosis of LDL across human umbilical vein endothelialcells (HUVECs) and stimulated an increase of subendothelial retention of LDL in vascular walls. These effects ofTNF-αwere substantially blocked not only by transcytosis inhibitors, but also byNF-κB inhibitors and PPAR-γ in-hibitors. In ApoE−/−mice, bothNF-κB and PPAR-γ inhibitors alleviated the early atherosclerotic changes promot-ed by TNF-α. NF-κB and PPAR-γ inhibitors down-regulated the transcriptional activities of NF-κB and PPAR-γinduced by TNF-α. Furthermore, cross-binding activity assay revealed that NF-κB and PPAR-γ could form an ac-tive transcription factor complex containing both the NF-κB P65 subunit and PPAR-γ. The increased expressionsof LDL transcytosis-related proteins (LDL receptor and caveolin-1, -2) stimulated by TNF-αwere also blocked byboth NF-κB inhibitors and PPAR-γ inhibitors. TNF-α promotes atherosclerosis by increasing the LDL transcytosisacross endothelial cells and thereby facilitating LDL retention in vascular walls. In this process, NF-κB and PPAR-γare activated coordinately to up-regulate the expression of transcytosis-related proteins. These observations sug-gest that inhibitors of either NF-κB or PPAR-γ can be used to target atherosclerosis.

© 2014 Elsevier Ltd. All rights reserved.

1. Introduction

Atherosclerosis (AS) is the pathophysiological basis of cardiovascu-lar and cerebral vascular diseases, which are the major causes of deathin elderly people. Due to the complexity in pathogenesis, the mecha-nism of AS is far from fully elucidated.

In the past decade, the “response-to-retention” hypothesis hasattracted much attention, which emphasizes the importance ofsubendothelial retention of apoB-containing lipoprotein particles (e.g.lowdensity lipoprotein, LDL) in the initiation of AS [1]. To be accumulat-ed in the vascular walls, the LDL particles must pass through the endo-thelial barrier. However, since the diameter of the LDL particles(20–30 nm) is much larger than the intercellular junction betweenthe vascular endothelial cells (3–6 nm), the only pathway for the LDLparticles to traffic across the intact endothelial barrier is through atransporting process termed transcytosis [2], whichwas first postulatedto describe the transport of macromolecular cargo from one side to the

ology, Tongji Medical College,el.: +86 27 8369 1182.

other side of cells via membrane-bounded carriers [3]. In terms of LDL,these particleswerefirst ingested by the endothelial cells on the luminalside. After being transported to the basolateral side, these particles werethen excreted to the subendothelial space.

On the other hand, AS is proposed to be a chronic inflammatory dis-ease of the arterial wall [4]. Several pro-inflammatory cytokines, includ-ing tumor necrosis factor-α (TNF-α), IL-1, IL-6 etc. have been reportedto promote AS. Specifically, TNF-α has been suggested to play a keyrole in the development of AS [5]. Previous studies showed that TNF-αwas predominately expressed in early atherosclerotic lesions [6] andthe concentration of TNF-α was higher in the serum of patients withAS [7,8]. However, the mechanism of how TNF-α promotes AS is yetto be fully clarified.

Given the importance of LDL transcytosis in the initiation and devel-opment of AS, we hypothesized that TNF-α promotes AS by increasingthe transcytosis of LDL across endothelial cells. To test this hypothesis,we first quantified the amount of LDL transcytosis in an in-vitro modelof an endothelial cell monolayer and found that TNF-α indeed signifi-cantly increased the transcytosis of LDL across endothelial cells. Tofurther clarify the molecular mechanisms involved, the roles of twoTNF-α-related transcription factors, nuclear factor kappa B (NF-κB)and peroxisome proliferator-activated receptor gamma (PPAR-γ)

86 Y. Zhang et al. / Journal of Molecular and Cellular Cardiology 72 (2014) 85–94

were also investigated. The present study demonstrates that TNF-αpromotes atherosclerosis by increasing the LDL transcytosis across en-dothelial cells and thereby facilitating LDL retention in vascular walls.In this process, NF-κB and PPAR-γ are activated coordinately to up-regulate the expression of transcytosis-related proteins.

2. Material and methods

2.1. Primary culture of human umbilical vein endothelial cells (HUVECs)

The collection of human umbilical cords was approved by the EthicsCommittee of Tongji Medical College, Huazhong University of Scienceand Technology (Wuhan, China). All subjects provided written in-formed consent prior to the initiation of the study. HUVECs were isolat-ed as previously reported [9] and cultured in endothelial cell medium

Fig. 1. Effect of TNF-α on LDL transcytosis in HUVECs. A. HUVECs were cultured in a monolawell. a, The medium in the apical chamber contained FITC-LDL (50 μg/mL), and the amtransported (control insert). b, The medium in the apical chamber contained FITC-LDL (50in the basal chamber as the amount of LDL transported paracellularly (naive insert). The amthe control insert by the naive insert. B. HUVECs were pretreated with Bay, PDTC, GW9662The amount of LDL transcytosis was measured. The amount of LDL transcytosis was normab 0.01 vs. TNF-α; n = 4. C. Upper panel: the representative Western blots showing the exppanel: the transfected HUVECs were exposed to TNF-α for 3 h. The amount of LDL transcexpressions of NF-κB p65 and PPAR-γ for 48 h after specific SiRNA transfection. Lowertranscytosis was measured. The amount of LDL transcytosis was normalized to that obtaineout TNF-α group; †, P b 0.05, ††, P b 0.01 vs. scrambled siRNA with TNF-α group; n = 3.

(ECM) (ScienCell). Cells were passaged when 70%–80% confluent andused between passages 2 and 6.

2.2. LDL labeling

As described previously [10], FITC (120 μg; Sigma) and LDL (2 mg;Yiyuan biotechnology, China) were incubated at 37 °C for 2 h. UnboundFITC was removed by dialysis against PBS overnight at 4 °C.

2.3. Measurement of LDL transcytosis

As described previously [11], the amount of LDL transcytosis wasmeasured by a non-radioactive method in vitro. As shown in Fig. 1A,HUVECs were seeded (~2 × 105 cells per insert) on a polyester mem-brane (Costar transwell, 6.5 mm diameter, 0.4 μm pore size). The

yer on a polyester membrane (0.4 μm) placed in the apical chamber of a dual-chamberount of FITC-LDL was measured in the basal chamber as the amount of total LDL

μg/mL) and 6-fold excess of unlabeled LDL, and the amount of FITC-LDL was measuredount of LDL transcytosis is the difference of FITC fluorescent intensity subtracted from, T007, NEM, MCD and 50 μg/mL FITC-LDL for 24 h, and then exposed to TNF-α for 3 h.lized to that obtained in the vehicle control group. **, P b 0.01 vs. vehicle group; ††, Pressions of caveolin-1 and caveolin-2 for 48 h after specific SiRNA transfection. Lowerytosis was measured. D. Upper panel: the representative Western blots showing thepanel: the transfected HUVECs were exposed to TNF-α for 3 h. The amount of LDLd in the scrambled siRNA without TNF-α group. **, P b 0.01 vs. scrambled siRNA with-

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integrity of the cell monolayer was determined by a method describedpreviously [2,12]. Two inserts of cell monolayers with equal integritywere assigned into the same group: the control insert and the naive in-sert. The control insert was incubated with FITC-LDL for 3 h to deter-mine the total amount of transendothelial LDL. Paracellular transportwas determined by incubating the cells with FITC-LDL (50 μg/mL) and6-fold excess of unlabeled LDL in the naive insert. The choice of 6-foldwas due to the fact that LDL with a concentration of more than350 μg/mL is significantly toxic to endothelial cells (data not shown),and a saturating concentration of LDL is 100 μg/mL in peripheral aorticendothelial cell monolayers as reported previously [13,14]. Sampleswere then collected from the outer chambers and further dialyzedagainst PBS to remove the free FITC due to possible degradation or me-tabolism in the cell. The FITC fluorescent intensity was measured via afluorescence spectrophotometer (TECAN, INFINITE F200PRO)with exci-tation and emission wavelengths of 490 nm and 520 nm, respectively.Background fluorescence was determined by measuring the serum-free ECM. We then subtracted it from the FITC fluorescent intensity ofeach sample. As a matter of fact, the amount of LDL transcytosis is thedifference in FITC fluorescent intensity between the control insert andthe naive insert. HUVECs were assigned into eight groups. They werepretreated with transcytosis inhibitor Filipin (1 mg/L; Sigma), N-ethylmaleimide (NEM, 10 μmol/L; Sigma) or Methyl-beta-cyclodextrin(MCD, 3 mmol/L; Sigma), NF-κB inhibitor Bay-11-7082(Bay, 1 μmol/L;Cayman Chemical) or ammonium pyrrolidinedithiocarbamate (PDTC,3 μmol/L; Sigma), or PPAR-γ inhibitor GW9662 (3 μmol/L; CaymanChemical) or T0070907 (T007, 10 μmol/L; Cayman Chemical) for 24 h,and then exposed to 30 ng/mL TNF-α for 3 h.

2.4. Confocal imaging and analysis of FITC-LDL in HUVECs and vascularwalls

HUVECs or human umbilical venous rings were incubated with var-ious inhibitors for 24 h at 37 °C, followed by FITC-LDL (50 μg/mL) andTNF-α for 3 h. For HUVECs, the individual microscopic field was ran-domly selected to include at least 15 cells and the number of cells wascounted. Fluorescence intensity was normalized to the number of cells[15]. Images were obtained with a confocal laser scan microscopy(Olympus FV500) using a 40× objective. For fluorescence imaging,wavelengths of 490-nmand 520-nmwere used for excitation and emis-sion respectively. For human umbilical venous rings, the 10 μm-thickfrozen sections were cut horizontally, and labeled with DAPI. Imagesof each area were captured at the same laser intensity, gain, and offsetto ensure consistency between sections from different groups. Foreach optical section, the region above the basilar membrane was de-fined as the region of interest (ROI). The integral optical density (IOD)and area were quantified using Image Pro Plus and the average opticaldensity of each ROI was IOD / area. All fluorescence images were quan-tified using Image J software.

2.5. Animal procedures

Animals were treated in accordance with the guide for the Care andUse of Laboratory Animals published by the US National Institutes ofHealth and approved by the local animal care committee. Seven-week-old male ApoE−/− mice weighing 20–30 g were purchased fromVital River Laboratory Animal Technology Co. Ltd in Beijing and housedin an air-conditioned room at 25 °Cwith a 12-h light/dark cycle. Allmicewere given a western chow diet (21% fat, 0.15% cholesterol; Vital RiverLaboratory) and water ad libitum. The procedure of the animal experi-ment is shown in Fig. 4A. Mice were randomly assigned into 6 groups(n = 6 per group) and intraperitoneally (i.p.) injected with 1 μg/kg re-combinantMouse TNF-α once everyweek [16,17] except Control group.After a 7-week administration, the hearts and aortas were collected andfixed in 4% paraformaldehyde.

2.6. Assessment of the atherosclerotic lesion size

Hearts were cut paralleling the leaflet and the upper portions wereembedded and cut at 8-μmin thickness though the aortic sinus. Sectionsof the aortic sinus were stained with Oil-red O. Morphometric analysisof microscopic images was performed by using Image-J software todetermine the ratio of lipid-stained plaque area to the vessel diameteras previously reported [18].

2.7. Immunohistochemistry analysis for CD154

Sections (8-μm thickness) of the aortic sinus were stained with pri-mary CD154 antibody (1:100, Proteintech Ltd) as described previously[19]. Images were obtained at a magnification of ×40. For the red quan-tification in the plaque area, the sectionswere examined at amagnifica-tion with a semi-quantitative scoring system as previously reported[19]. All analyses were performed in a blinded manner.

2.8. NF-κB and PPAR-γ activity assay

HUVECs were pretreated with Bay, PDTC, GW9662 and T007 for30 min or 24 h, followed by TNF-α (30 ng/mL) stimulation for 30 min.Then the NF-κB and PPAR-γ activity assay was carried out accordingto an ELISA-based method described previously [20,21]. To excludethe false positive, we used the anti-CREB antibody as the negative con-trol, which does not bind to KBRE or PPRE probe. The blank controlswere also set up. The blanks are realized following the same procedureas the tests, except that lysis buffer is incubated in the microwellsinstead of cell lysate. The results are expressed after subtraction of theblank values.

2.9. NF-κB and PPAR-γ cross-binding activity assay

This test is similar to the above ELISA-based method for NF-κB andPPAR-γ activity assay as shown in Figs. 6C &G. HUVECswere stimulatedwith TNF-α for 30 min. Cell extracts were incubated in a 96-well plate,which had been coated with the oligonucleotide containing the NF-κBresponse elements (KBRE; sequence: 5′-GGGACTTTCC-3′); or PPAR-γresponse elements (PPRE; sequence: 5′-AGGTCAAAGGTCA-3′).The for-mation of complexes between KBRE and activated transcription factorproteins was detected by NF-κB p65 antibody as in regular assay andby PPAR-γ antibody. Similarly, the formation of complexes betweenPPRE and activated transcription factor proteins was detected by thePPAR-γ antibody as in the regular assay and by the NF-κB p65 antibody.The activities were quantified from absorbance values measured at awavelength of 450 nm.

2.10. Western blot analysis

HUVECs were pretreated with various inhibitors for 24 h, followedby TNF-α stimulation for 3 h. Cells were lysed with the RIPA lysis buffer(Beyotime Institute of Biotechnology) containing protease inhibitors.The proteins were separated by SDS-PAGE gel and transferred to thePVDF membrane. The membranes were probed with primary antibod-ies against LDLR (1:700; Proteintech Group), caveolin-1 (1:8000, CellSignaling), caveolin-2 (1:100; Santa Cruz) and β-actin (1:4000;Proteintech Group).

2.11. RNA interference

HUVECs were transfected with the SiRNA targeted for caveolin-1(Santa Cruz, sc-29241), caveolin-2 (Santa Cruz, sc-40388), NF-κB p65(Santa Cruz, sc-29410) or PPAR-γ (Santa Cruz, sc-29455) usingHiperfect Transfection Reagent (Qiagen) according to manufacturer'sinstruction. A SiRNA consisting of a scrambled sequence of similarlength was transfected as control. Before transfection, HUVECs at

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passage 3 were seeded (~6 × 104 cells per insert) on a polyester mem-brane (Costar transwell, 6.5 mm diameter, 0.4 μm pore size) and theywere 30–50% confluent at the time of transfection. The cells were incu-bated with the transfection complexes under their normal growthcondition for 48 h after transfection. Then the protein expressions ofcaveolin-1, caveolin-2, NF-κB p65 or PPAR-γ were evaluated usingwestern blots and the LDL transcytosis was measured. As shown inthe upper panel of Fig. 1C, the expressions of caveolin-1 and caveolin-2 were both reduced by (50.8 ± 15.4) % and (40.6 ± 10.9) % at 48 hafter SiRNA transfection, respectively. And the expressions of NF-κB p65or PPAR-γ were both reduced by (46.6 ± 10.6) % and (51.7 ± 15.0) %,respectively (upper panel of Fig. 1D).

2.12. Statistical analyses

Data are presented as the mean ± SE. Multiple group comparisonswere evaluated by one-way ANOVA with Duncan's multiple-rangetesting. P b 0.05 was considered statistically significant.

Fig. 2. Confocal analysis of FITC-LDL uptake in HUVECs. HUVECs were first incubated withFITC-LDL and various inhibitors for 24 h, followed by exposure to TNF-α for 3 h. A. Confo-cal microscopic images of FITC-LDL uptake in HUVECs incubated with LDL (a), F-LDL(b), TNF-α (c) alone or with pretreatment of Bay (d), PDTC (e), GW9662 (f), T007(g) or NEM (h) MCD (i). B. Quantitative summary of FITC-LDL uptake in HUVECs.** P b 0.01 vs. Ctr; ††, P b 0.01 vs. TNF-α; n = 3. Scale bar = 100 μm.

3. Results

3.1. TNF-α increased LDL transcytosis in HUVECs

TNF-α is a promoting factor in the development of AS. In thisstudy, we first tested whether TNF-α could increase LDL transcytosisusing the established non-radioactive in-vitro method. The amountof LDL transcytosis was normalized to that obtained in the controlgroup. At 10, 30, 60, 120 or 180 min after TNF-α (30 ng/mL) incuba-tion, the transportation of LDL across the HUVEC monolayer in-creased in a time dependent manner, with a peak at 120–180 mintime point (Fig. S1A). Increasing the concentration of TNF-α (1, 3,10, 30 and 100 ng/mL) resulted in increase in the amount of LDLtranscytosis, with a plateau at 30 ng/mL (Fig. S1B). As shown inFig. 1B, exposure to TNF-α (30 ng/mL) for 3 h significantly increasedLDL transcytosis. Pretreatment with NF-κB inhibitors (Bay, PDTC),PPAR-γ inhibitors (GW9662, T007) or transcytosis inhibitors (Filipin,NEM, MCD) largely attenuated TNF-α-induced LDL transcytosis.Specific knockdown of caveolin-1, caveolin-2, NF-κB p65 or PPAR-γ

Fig. 3. Confocal analysis of FITC-LDL retention in human umbilical venous walls. Thehuman umbilical venous rings were incubated with FITC-LDL and various inhibitors for24 h, followed by exposure to TNF-α for 3 h. A. Confocal microscopic images of FITC-LDLretention in human umbilical venous rings incubated with LDL (a), F-LDL (b), TNF-α(c) alone or with Bay (d), PDTC (e), GW9662 (f), T007 (g) or NEM (h), MCD (i). B. Quan-titative summary of FITC-LDL retention in vessels. **, P b 0.01, vs. Ctr; ††, P b 0.01 vs. TNF-α;n = 3. Scale bar = 100 μm.

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also inhibited TNF-α-stimulated LDL transcytosis (lower panel ofFigs. 1C and D).

3.2. TNF-α exposure increased the uptake of FITC-LDL in HUVECs

The uptake of LDL by endothelial cells is an intermediate step ofLDL transcytosis. After incubation with FITC-LDL, the HUVECs were

Fig. 4. Effect of TNF-α exposure on atherosclerotic lesion formation in ApoE−/− mice. A. The pQuantitative summary of the percentage of the area of atherosclerotic lesion in the aortic rootroot sections. Bar = 200 μm. (E) Quantitative summary of the staining of CD154. **, P b 0.01 vs

filled with small, individual, discrete vesicles throughout the cells(Figs. 2A-b). The fluorescent intensity in individual cells reflectedthe extent of LDL uptake. As shown in Fig. 2, TNF-α exposure signif-icantly increased the LDL uptake. In contrast, NF-κB inhibitors (Bay,PDTC), PPAR-γ inhibitors (GW9662, T007) or transcytosis inhibitors(NEM, MCD) significantly diminished TNF-α-stimulated FTIC-LDLinternalization.

rocedure of animal experiment. B. Oil Red O-stained aortic root sections. Bar = 600 μm C.. **, P b 0.01, vs. Ctr; ††, P b 0.01 vs. TNF-α; n = 6. D. Immunostaining for CD154 in aortic. Ctr; †, P b 0.05; ††, P b 0.01 vs. TNF-α; n = 6.

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3.3. TNF-α exposure increased the retention of LDL in human umbilicalvenous walls

Subendothelial retention of apoB-containing lipoprotein particles(e.g., LDL) is important for the initiation of AS. The fluorescence-labeled LDL tracer (FITC-LDL) was used to examine LDL retention inhuman umbilical venous walls. As shown in Figs. 3A-b, only a smallamount of LDL was detained in human umbilical venous walls after in-cubated with FITC-LDL (50 μg/mL) for 3 h. TNF-α exposure enhancedLDL transportation from the endothelial cells to the internal elastic lam-ina. In the presence of Bay, PDTC, GW9662 and T007, the transportedLDL was primarily located in the endothelial cells, which indicatedthat these four inhibitors decreased LDL transport in human umbilicalvenous walls. When NEM and MCD were used, transported LDL wasalmost completely diminished (Fig. 3).

3.4. TNF-α accelerated atherosclerotic lesion formation in ApoE−/− mice

To address whether TNF-α-induced LDL transcytosis is relevant toatherosclerosis, ApoE−/− mice were injected with TNF-α alone orin combination with various inhibitors. Treatment with TNF-α for7 weeks (1 μg/kg, i.p. weekly) resulted in a 5-fold-higher mean aorticplaque area compared with the saline controls. NEM and MCD almostcompletely blocked the effect of TNF-α on promoting AS. PDTCand GW9662 also reduced the increase of the lesion size induced byTNF-α (Figs. 4B–C).

CD154 is amarker of early atherosclerosis and is distributed in aorticplaques in ApoE−/− mice (Figs. 4D–E). TNF-α-challenged ApoE−/−

Fig. 5. Effects of NF-κB and PPAR-γ inhibitors on TNF-α-stimulated NF-κB and PPAR-γ activitiesstimulation for 30min, andNF-κB activities (A) or PPAR-γ activities (B)were assayed. C,D.HUVE30 min, and NF-κB activities (C) or PPAR-γ activities (D) were assayed.* P b 0.05, ** P b 0.01, v

mice had stronger CD154 signals, whereas the PDTC- or GW9662-pretreated groups had relatively weaker signals. The NEM- or MCD-pretreated groups also showed reduced CD154 signals.

3.5. TNF-α increased transcriptional activity of both NF-κB and PPAR-γ inHUVECs

In our study, we found that NF-κB and PPAR-γwere both involved inTNF-α-induced LDL transcytosis and the acceleration of atherosclerosis.We further examined the effect of TNF-α on the transcriptional activi-ties of NF-κB and PPAR-γ. The transcriptional activities of NF-κB(Figs. 5A & C) and PPAR-γ (Figs. 5B & D) were significantly increasedby TNF-α stimulation. Pretreatment with NF-κB inhibitors for 30 minor 24 h not only blocked TNF-α-induced NF-κB activation, but alsoprevented TNF-α-stimulated PPAR-γ activation. Pretreatment withPPAR-γ inhibitors also had similar effects, whether it was for 30 minor 24 h. The parallel changes in the transcriptional activities of NF-κBand PPAR-γ suggested that TNF-α may stimulate an interactionbetween NF-κB and PPAR-γ.

We also examined the effect of NF-κB inhibitors or PPAR-γ inhibitorson the basal activities of NF-κB and PPAR-γ. Incubation with NF-κBinhibitors or PPAR-γ inhibitors for 30 min inhibited the transcriptionalactivities of NF-κB (Fig. 5A) and PPAR-γ (Fig. 5B). However, incubationwith these inhibitors for 24hmildly up-regulated the transcriptional ac-tivities of NF-κB (Fig. 5C) and PPAR-γ (Fig. 5D). The mild up-regulationis probably associated with the hypersensitivity of the receptor regula-tion, which is usually observed in the long-term treatment with inhibi-tors of receptors, just as β-blockers do.

. A, B. HUVECs were treated with various inhibitors for 30min alone or followed by TNF-αCswere treatedwith various inhibitors for 24 h alone or followedbyTNF-α stimulation fors. vehicle control group; † P b 0.05, ††, P b 0.01 vs. TNF-α; n = 3.

Fig. 6. Effect of TNF-α on the cross-binding activity of NF-κB and PPAR-γ. HUVECswere treatedwith TNF-α for 30min. A, C, E, and G represent the schematic diagrams of ELISA-based crossbinding activity assay. Quantitative resultswere summarized inB, D, F, andH respectively.*, P b 0.05; ** P b 0.01, vs. Ctr;n=3.KBRE=NF-κB response elements; PPRE=PPAR-γ responseelements; Ab = antibody.

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3.6. TNF-α induced NF-κB activity binding to PPAR-γ responsive elements(PPRE) and PPAR-γ activity binding to NF-κB responsive elements (KBRE)in HUVECs

In order to verify the above speculation, we performed a cross-binding test. Cell extracts were incubated in a 96-well plate, whichhad been coated with the NF-κB binding probe, namely KBRE.Transcriptional complex binding to KBRE was detected not only by theNF-κB p65 antibody but also by the PPAR-γ antibody (Figs. 6A & C).TNF-α stimulation increased the transcriptional activity of NF-κB(Fig. 6B), and simultaneously increased the activity of PPAR-γ bindingto KBRE (Fig. 6D). Simultaneously, cell extracts were coated with thePPAR-γ binding probe, namely PPRE. Transcriptional complex bindingto PPRE was detected not only by the PPAR-γ antibody but also by theNF-κB p65 antibody (Figs. 6B & D). We also found that TNF-α stimula-tion increased the transcriptional activity of PPAR-γ (Fig. 6F), and simul-taneously increased the activity of NF-κB binding to PPRE (Fig. 6H).These results provided evidences that TNF-α indeed induced the forma-tionof an active transcription factor complex containing bothNF-κBp65and PPAR-γ. Co-immunoprecipitation assay further demonstrated theformation of an active transcription factor complex containing bothNF-κB p65 and PPAR-γ in the presence of TNF-α. NF-κB inhibitor orPPAR-γ inhibitor disrupted the interaction between NF-κB p65 andPPAR-γ (Fig. S2).

3.7. TNF-α stimulated the expression of molecules involved in LDLtranscytosis across endothelial cells

As shown in Fig. 7, the expression of proteins involved in LDLtranscytosis (LDLR, caveolin-1 and caveolin-2) was up-regulated in

Fig. 7. Effect of TNF-α on the expression of LDLR, caveolin-1 and -2 in HUVECs. A. Repre-sentative Western blots showing the expression of LDLR, caveolin-1 and caveolin-2 withvarious inhibitors for 24 h and followed by TNF-α stimulation for 3 h. B. Quantitative anal-ysis of the expression of proteins. **, P b 0.01, vs. Ctr; ‡, P b 0.01 vs. TNF-α; n = 3.

HUEVCs by TNF-α stimulation for 3 h. Thus, TNF-α-induced LDLtranscytosis may partly be due to the elevated expression of these pro-teins mediating the LDL transcytosis. Both NF-κB inhibitors (Bay orPDTC) and PPAR-γ inhibitors (GW9662 or T007) significantly blockedthe up-regulation of LDLR, caveolin-1 and caveolin-2 stimulated byTNF-α.

4. Discussion

Although TNF-α has long been recognized as a putative inflammato-ry mediator to provoke atherosclerosis, the molecular mechanisms arenot completely understood. Here, we demonstrate that TNF-α signifi-cantly upregulates the transcytosis of LDL across endothelial cells andpromotes the retention of LDL particles in vessel walls, thereby acceler-ating the development of atherosclerosis. In this process, the two ubiq-uitous transcriptional factors, NF-κB and PPAR-γ crosstalk with eachother and orchestrate the whole event.

We first demonstrated that TNF-α could significantly increase thetranscytosis of LDL across endothelial cells by a non-radioactive in-vitro method. To further determine the roles of NF-κB and PPAR-γ inTNF-α-stimulated LDL transcytosis, two structurally different, but bothselective inhibitors, were employed as tool drugs for each of these twotranscription factors. For NF-κB, we used Bay-11-7082 (Bay) and PDTCas inhibitors [22]. For PPAR-γ, GW9662 and T0070907 (T007) wereused [23,24]. It was found that TNF-α-stimulated LDL transcytosiswas largely blocked not only by NF-κB inhibitors, Bay and PDTC,but also by PPAR-γ inhibitors, GW9662 and T007, thereby stronglysuggesting critical roles of these two transcription factors in mediatingthe TNF-α-stimulated LDL transcytosis.

An intermediate event of LDL transcytosis across endothelial cells isthat LDL particles are endocytosed into the cells, but still have not beenexcreted out through the other side of the cells. When the LDL particlesare further excreted outside of the cells, these particles are retained inthe sub-endothelial space. Thus, the determination of the intermediateform of LDL transcytosis and the subendothelial retention of LDL invascular wall reflects the activity of transcytosis. Indeed we found thatobservations in these two sets of experiments were consistentwith those observed in the in-vitro transcytosis model mentionedabove. Both NF-κB inhibitors and PPAR-γ inhibitors prevented the up-regulation of LDL transcytosis by TNF-α stimulation. These data provid-ed compelling evidences proving that NF-κB and PPAR-γwere involvedin the TNF-α-stimulated LDL transcytosis.

In ApoE−/− mice, we found that TNF-α injections accelerated theformation of atherosclerotic plaques in the arteries, further supportingthe long-standing view of TNF-α as a pro-atherosclerotic factor. Block-ade of the LDL transcytosis by NEM and MCD substantially preventedthe early atherosclerotic changes in artery walls, suggesting a criticalrole of LDL transcytosis in the initiation or development of AS. Mean-while, consistent with the above in-vitro findings, early atheroscleroticchanges in artery walls promoted by TNF-α were not only reversed byNF-κB inhibitors, but also by PPAR-γ inhibitors. In addition, we studiedthe expression of the CD40 ligand, CD154, which is involved in the earlyatherogenesis and contributes to the initial recruitment of inflammato-ry cells to damaged endothelium [25]. It has been described previouslythat the inhibition of the CD40-CD154 signal retarded the initiation ofarterial plaque formation [26]. Our results showed that both NF-κB in-hibitors and PPAR-γ inhibitors lowered the CD154 expression inplaques which were substantially elevated by TNF-α in ApoE−/− mice.These observations further support the in-vivo significance of theinhibition of NF-κB or PPAR-γ, as strategies for the prevention or treat-ment of AS.

To further explore how NF-κB and PPAR-γ were involved in thetranscytosis of LDL in endothelial cells, we further studied the alter-ations of these two transcription factors and their downstream genes.By using an ELISA-based transcriptional factor-DNA binding activityassay, we found that TNF-α could significantly activate NF-κB, which

93Y. Zhang et al. / Journal of Molecular and Cellular Cardiology 72 (2014) 85–94

was consistent with many previous reports [27–29]. However, wefound that the activity of PPAR-γwas also up-regulated by TNF-α stim-ulation, which was different from some previous reports showing thatTNF-α inhibited PPAR-γ [30–32]. NF-κB inhibitors (Bay and PDTC)primarily prevented the TNF-α-stimulatedNF-κB activation, but also re-versed the TNF-α-stimulated PPAR-γ activity to some extent. Likewise,PPAR-γ inhibitors, GW9662 and T007 almost completely abolished theTNF-α-stimulated PPAR-γ activation, and also blocked the TNF-α-stimulated NF-κB activity. This was consistent with a previous reportdemonstrating that NF-κB inhibitor, Bay, also attenuated the PPAR-γ ac-tivity in HepG2 cells [33]. We found that NF-κB activation could also beaugmented by PPAR-γ, because the direct inhibition of PPAR-γ couldalso down-regulate the NF-κB activity. These findings strongly supporta positive crosstalk between NF-κB and PPAR-γ, which means NF-κBand PPAR-γmaywell interact with each other and enhance each other'sactivation. To further verify this speculation, we designed and conduct-ed a cross-binding activity assay. We were surprised to find that, uponthe stimulation of TNF-α, not only could the active transcription factorprotein complex binding with KBRE bind with the NF-κB P65 antibody,but it could also do sowith the PPAR-γ antibody. Meanwhile, the activetranscription factor complex binding with PPRE was found to not onlybe bound with the PPAR-γ antibody, but also bound with the NF-κBP65 antibody. These results strongly suggest that the activated tran-scription factor complex stimulated by TNF-α contains both the NF-κBP65 subunit and PPAR-γ. This active transcription factor complex in-deed has binding affinities to both KBRE and PPRE, thereby controllingthe transcription and expression of target genes of both NF-κB and

Fig. 8. Schematicmodel of the crosstalk betweenNF-κB and PPAR-γ. A. NF-κB and PPAR-γ remaof IκB and NF-κB P65 translocates into the nuclear. In the nucleus, NF-κB P65 binds with PPARexpression of caveolin-1,2 and LDLR, respectively. C. In the presence of NF-κB inhibitors, Bay aform an active complex with PPAR-γ. Thus, PPAR-γ can not bind to PPRE and also can not incT007, these antagonists bind to PPAR-γ, preventing the binding of PPAR-γ to NF-κB P65 and thnot be activated and their target genes’ expression is inhibited.

PPAR-γ, respectively. When one of these two transcription factors wasinhibited by its selective inhibitors, these inhibitors also blocked theinteraction between these two transcription factors, preventing the for-mation of active transcription factor complex, thus blocking the activa-tion of both transcription factors. These results are similar to a recentpaper demonstrating that NF-κB's activation is dependent on the forma-tion of a complex between P65 and PPAR-γ, which suggests an active,but not inactive complex of NF-κB and PPAR-γ [34]. Here, we furthercame up with the notion that this complex was also functional toPPRE. Thus, PPAR's activation is also dependent on NF-κB, or so atleast in part. These models of TNF-α-stimulated crosstalk betweenNF-κB and PPAR-γ are summarized in Fig. 8.

To answer the questionwhy the inhibition of NF-κB or PPAR-γ couldreduce LDL transcytosis across endothelial cells and thereby prevent theformation of AS, we further conducted some studies. The endothelialcells are abundant with caveolae structures and LDL receptors are alsomainly located in these specialized membrane raft domains. The struc-tural proteins of endothelial caveolae, caveolin-1, and -2, play importantroles in the transcytosis of LDL across endothelial cells [35,36]. Caveolin-1 is expressed in the majority of cell types involved in the developmentof an atheroma [12,37,38]. Caveolin-1 is present on both the luminaland basolateral plasma membranes, whereas caveolin-2 is enriched onthe basolateral surface where caveolae are present [39]. In ECs, thedown-regulation of caveolin-1 prevents the transcytosis of LDL acrossendothelial cells, indicating a critical role of caveolae in mediating thetranscytosis of LDL particles. By determining the expression of these es-sential proteins involved in LDL transcytosis, including the LDL receptor

in inactive in the absence of TNF-α. B. TNF-α triggers the phosphorylation and degradation-γ. They form an active complex and bind to both KBRE and PPRE, and then promote thend PDTC, TNF-α can not drive NF-κB P65 to translocate into nuclear and thereby can notrease the target genes' expression. D. In the presence of PPAR-γ inhibitors, GW9662 ande active complex between NF-κB and PPAR-γ can not form. Thus the KBRE and PPRE can

94 Y. Zhang et al. / Journal of Molecular and Cellular Cardiology 72 (2014) 85–94

(LDLR), caveolin-1 and -2 [40,41], we found that TNF-α could signifi-cantly up-regulate the expression of these three proteins, but these ef-fects could also be reduced by inhibitors of either NF-κB or PPAR-γ.These results are consistent with previous studies, which report NF-κBand PPAR-γ dependent expression of caveolin-1 and -2. However, ithas also been reported that there are no NF-κB binding sites in the pro-moter region of LDLR gene [42], and therefore the down-regulation ofLDLR by NF-κB inhibitors may not directly be due to the inhibition ofNF-κB, but rather indirectly be due to the inhibition of PPAR-γ, whichhas been shown to regulate the expression of LDLR directly [43,44].These results at least in part explain the anti-atherosclerotic effectsbrought on by the inhibition of NF-κB or PPAR-γ.

In conclusion, the present study for the first time demonstrates thatthe putative inflammatory factor, TNF-α, exerts its pro-atheroscleroticaction by directly increasing the transcytosis of LDL particles acrossthe endothelial cells. More specifically, NF-κB and PPAR-γ are both acti-vated upon TNF-α stimulation and potentiate each other's activation inan interdependent manner. These findings strongly support the utiliza-tion of inhibitors of NF-κB or PPAR-γ to prevent or treat AS-baseddisorders.

Disclosures

None declared.

Acknowledgment

This study was supported by grants from the National NaturalScience Foundation of China (81072634, 81000080, 30870997,81373413), grants from the Ministry of Education of China (NCET-10-0409, 2011TS069, 20101561, 2013QN182, 2013YGYL008) aswell as from the National Science and Technology Major Projects(2013zx09103-001-020, 2011zx09102-004-001).

Appendix A. Supplementary data

Supplementary data to this article can be found online at http://dx.doi.org/10.1016/j.yjmcc.2014.02.012.

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