Accepted Manuscript Title: Cadmium Induces NLRP3 Inflammasome-dependent Pyroptosis in Vascular Endothelial Cells Author: Haiyan Chen Yonghui Lu Zhengwang Cao Qinlong Ma Huifeng Pi Yiliang Fang Zhengping Yu Houxiang Hu Zhou Zhou PII: DOI: Reference:

S0378-4274(16)30015-7 http://dx.doi.org/doi:10.1016/j.toxlet.2016.01.014 TOXLET 9309

To appear in:

Toxicology Letters

Received date: Revised date: Accepted date:

21-11-2015 18-1-2016 19-1-2016

Please cite this article as: Chen, Haiyan, Lu, Yonghui, Cao, Zhengwang, Ma, Qinlong, Pi, Huifeng, Fang, Yiliang, Yu, Zhengping, Hu, Houxiang, Zhou, Zhou, Cadmium Induces NLRP3 Inflammasome-dependent Pyroptosis in Vascular Endothelial Cells.Toxicology Letters http://dx.doi.org/10.1016/j.toxlet.2016.01.014 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

Title: Cadmium Induces NLRP3 Inflammasome-dependent Pyroptosis in Vascular Endothelial Cells

Authors: Haiyan Chena, Yonghui Lub, Zhengwang Caob, Qinlong Mab, Huifeng Pib, Yiliang Fangb, Zhengping Yub, Houxiang Hua*, Zhou Zhoub**

Affiliations: a

Department of Cardiovasology, Affiliated Hospital of North Sichuan Medical College, Nanchong

637000, Sichuan, China b

Department of Occupational Health, Third Military Medical University, Chongqing 400038, China

Corresponding author at: *Department of Cardiovasology, Affiliated Hospital of North Sichuan Medical College, No.63 Wenhua Street, Shunqing District, Nanchong 637000, Sichuan, China. ** Corresponding author at: Department of Occupational Health, Third Military Medical University, No. 30 Gaotanyan Main St., Shapingba District, Chongqing 400038, China. Email address: [email protected] (Houxiang Hu), [email protected] (Zhou Zhou)

Highlights: 1. Cadmium (Cd) exposure induces NLRP3 inflammasome activation in HUVECs. 2. Cd exposure causes pyroptosis in HUVECs. 3. Mitochondrial ROS mediates Cd-induced NLRP3 activation and pyroptosis in HUVECs.

Abstract Cadmium (Cd) is an important and common environmental pollutant that has been linked to cardiovascular diseases, such as atherosclerosis and hypertension. Increasing evidence demonstrates that Cd impairs the cardiovascular system by targeting vascular endothelial cells, but the underlying mechanisms remain obscure. In human umbilical vein endothelial cells (HUVECs), we observed that Cd treatment led to cell death and the generation of inflammatory cytokines. The Cd-induced cell death was identified as pyroptosis, a novel pro-inflammatory form of cell death depending on caspase-1 activation. In addition, exposure of HUVECs to Cd resulted in NLRP3 inflammasome activation as evidenced by cleavage of caspase-1 and downstream interleukin (IL)-1β production. Moreover, knockdown of NLRP3 by small interfering RNA efficiently suppressed Cd-induced caspase-1 cleavage, IL-1β production and pyroptosis in HUVECs. Additional experiments demonstrated that treatment with Cd significantly increased the levels of mitochondrial reactive oxygen species (mtROS) and intracellular ROS in HUVECs. Accordingly, pre-treatment with mtROS scavenger or total ROS scavenger reduced Cd-induced activation of NLRP3 inflammasome and pyroptotic cell death. Taken together, our data suggest that NLRP3 inflammasome, activated by the generation of mtROS, mediates Cd-induced pyroptosis in HUVECs. Our results provide novel insights into Cd-induced cytotoxicity and the underlying mechanism by which Cd induces endothelial injury.

Keywords: Cadmium; Cardiovascular disease; NLRP3 inflammasome; Pyroptosis; ROS; Vascular endothelial cells.

1. Introduction Cd is an occupational and environmental heavy metal pollutant that adversely affects human health. The main sources of Cd exposure in humans include industrial contamination, batteries, food sources, and tobacco smoke (Nawrot et al., 2010). Once absorbed, Cd accumulates in the tissues and leads to a higher incidence of Cd-related diseases. It has been reported that Cd exhibits toxicity in a wide variety of tissues including bone, heart, liver, kidney, brain, lung, endocrine system and immune system, which may cause harmful effects including fragile bone, cardiovascular disease, abnormal liver metabolic function, renal dysfunction, impaired neurodevelopment, pulmonary edema, type-2 diabetes and cancer (Bernhoft, 2013; Satarug and Moore, 2012). Accumulating evidence shows that Cd is an independent risk factor for various cardiovascular diseases in humans, such as hypertension, atherosclerosis, peripheral arterial disease and myocardial infarction (Messner and Bernhard, 2010; Tellez-Plaza et al., 2013a; Tellez-Plaza et al., 2013b), and the vascular endothelium is an important target of Cd toxicity (Dong et al., 2009; Nagarajan et al., 2013; Prozialeck et al., 2006). Experimental and clinical studies demonstrated that endothelial injury is an important process in the development of hypertension induced by Cd (Kim et al., 2012; Prozialeck et al., 2006; Zhang et al., 2015a). Other experimental evidence suggests that Cd exposure can contribute to the initiation and progression of atherosclerosis due to oxidative stress caused by Cd and may increase lipid peroxidation in vascular endothelial cells (VECs) (Angeli et al., 2013). In vitro studies have demonstrated that Cd treatment caused endothelial cell death, including apoptosis and necrosis (Kaji et al., 1995; Kishimoto et al., 1996). Exposure of ApoE knock-out mice, a common mouse model of arteriosclerosis, to 100 mg/L Cd in drinking water for 12 weeks, resulted in more advanced arteriosclerotic lesions, as well as signs of vascular inflammation (Knoflach et al., 2011). Cd has also been shown to cause the release of various pro-inflammatory cytokines from VECs that facilitate the inflammatory components of atherosclerotic process, such as tumor necrosis factor-α (TNF-α) and interleukin (IL)-6 (Dong et al., 2014; Mlynek and Skoczynska, 2005; Szuster-Ciesielska et al., 2000a). However, the underlying mechanisms by which Cd induces endothelial cell death and the inflammatory response have not yet been completely elucidated. Inflammasome is a multi-molecular platform usually comprised of a cytosolic sensor, an adaptor of ASC (apoptotic speck-like protein containing CARD) and an effector of pro-caspase-1 (Schroder and Tschopp, 2010). Inflammasomes usually serve as pattern recognition receptors and play

important roles in host defense and sterile inflammatory diseases (Abderrazak et al., 2015). Of these inflammasomes, NLRP3 (NLR pyrin domain containing 3) inflammasome is the most studied, and is composed of NLRP3, ASC, and pro-caspase-1. The NLRP3 inflammasome has been reported to be activated by a wide variety of pathogen-associated molecular patterns (PAMPs) and endogenous danger-associated molecular patterns (DAMPs), such as adenovirus, Candida albicans, extracellular ATP, uric acid and cholesterol crystals (Abderrazak et al., 2015; Tschopp and Schroder, 2010). Upon activation, NLRP3 recruits and cleaves pro-caspase-1 into its active form. The active caspase-1 is composed of a tetramer containing two 20-kD fragments (Casp1 p20) and two 10-kD fragments, which process pro-IL-1β and pro-IL-18 into their mature forms (IL-1β and IL-18) (Schroder and Tschopp, 2010). In addition, caspase-1 activation also results in a novel form of cell death known as pyroptosis. Pyroptosis is a pro-inflammatory cell death which is dependent on caspase-1 activation. Cells undergoing pyroptosis result in DNA damage, pore formation in membrane with positivity for dead cell staining, cell swelling, lysis and the release of pro-inflammatory contents into the extracellular space (Chang et al., 2013; Wree et al., 2014). Accumulating evidence shows that NLRP3 inflammasome activation with or without pyroptosis is critical for the development of cardiovascular disease (Li et al., 2014). NLRP3 inflammasome activation has been demonstrated to be required for atherogenesis and is activated by cholesterol crystals (Duewell et al., 2010). Recent studies indicate that cholesterol crystals cause endothelial dysfunction by NLRP3 inflammasome activation resulting in pyroptosis of VECs (Zhang et al., 2015b). With the exception of PAMPs and DAMPs, the NLRP3 inflammasome has also been shown to be activated by several environmental irritants, including silica, asbestos and alum (Abderrazak et al., 2015; Schroder and Tschopp, 2010). Interestingly, NLRP3 has been reported to be activated by the metal nickel ion (Ni2+) (Li and Zhong, 2014). Importantly, a study revealed that a selective inhibitor of caspase-1 suppressed Cd-induced cell death (Kim et al., 2000). However, to the best of our knowledge, few studies have demonstrated that the cell death induced by Cd, which could be suppressed by a caspase-1 inhibitor, was pyroptosis. Moreover, whether NLRP3 activation and the resulting pyroptosis are involved in Cd-induced endothelial injury and cell death in VECs has not been investigated. The data presented in the current report demonstrate that Cd induced the inflammatory response, NLRP3 inflammasome activation and subsequent pyroptosis in human umbilical vein endothelial

cells (HUVECs). Considering that mitochondrial reactive oxygen species (mtROS) have been proved to play a crucial role in NLRP3 inflammasome activation, we also explored whether Cd induces NLRP3 activation through mtROS production. Our findings also demonstrated that NLRP3 inflammasome activation and pyroptosis were mediated by the increase in mtROS after Cd treatment in HUVECs. These data illustrate a new molecular mechanism of endothelial injury and inflammation caused by Cd, which may be utilized for future clinical treatment of Cd-induced cardiovascular toxicity.

2. Materials and methods 2.1. Reagents and antibodies Dulbecco's modified Eagle's medium (DMEM) and fetal bovine serum (FBS) were obtained from Gibco (Grand Island, NY, USA). Cadmium chloride (CdCl2) and Mito-TEMPO were purchased from Sigma–Aldrich, Co. (St. Louis, MO, USA). Z-YVAD-FMK, Control-siRNA (small interfering RNA), NLRP3-siRNA, anti-cryopyrin antibody and anti-pro-caspase-1 antibody were purchased from Santa Cruz Biotechnology (Santa Cruz, CA, USA). Anti-caspase-1 antibody was purchased from Novus Biologicals (Novus Biologicals, Littleton, CO, USA). Anti-IL-1β antibody, anti-mouse antibody and anti-rabbit antibody were obtained from Abcam (Cambridge, UK). FLICA® 660 in vitro Caspase-1 Detection (FLICA 660-YVAD-FMK) Kit was purchased from Immunochemistry Technologies, LLC (Bloomington, MN, USA). All other chemicals and reagents were purchased from Beyotime Biotechnology (Beyotime, Jiangsu, China) unless otherwise specified.

2.2 Cell culture and treatment HUVECs were obtained from American Type Culture Collection (Manassas, VA, USA). The HUVECs

were

cultured

in

DMEM

supplemented

with

10%

FBS

and

1%

(v/v)

penicillin/streptomycin at 37°C in a humidified atmosphere with 95% air and 5% CO2. HUVECs were primed with 200 ng/mL lipopolysaccharide (LPS) for 3 h prior to CdCl2 exposure. In the experiments involving pharmacological inhibitors, HUVECs were pre-treated by addition of the pyroptosis inhibitor glycine (5 mM), caspase-1 inhibitor Z-YVAD-FMK (20 µM), mtROS scavenger Mito-TEMPO (50 µM) or total ROS scavenger N-acetyl-L-cysteine (NAC, 1 mM) diluted in

medium for 60 min and then treated with CdCl2 in the presence of these inhibitors.

2.3. Cell viability assay Cell viability was determined using the Cell Counting Kit-8 (CCK-8) (Dojindo Kumamoto, Japan) according to the manufacturer’s instructions. Briefly, HUVECs were cultured in 96-well plates to reach the desired confluence. The cells were then incubated with different concentrations of CdCl2 (0, 20, 40, and 80 μM). After 12 and 24 h of CdCl2 treatment, 10 μl CCK-8 was added to each well. The cells were then incubated for 2 h at 37°C. Following incubation, the absorbance was read at 450 nm using a microplate reader (Tecan, Mannedorf, Switzerland).

2.4. Lactate dehydrogenase release assay The release of lactate dehydrogenase (LDH) into the supernatant was regarded as an indicator of cytotoxicity. After the cells had been exposed to the various treatments, the culture supernatants were harvested, and the LDH levels were determined using the LDH cytotoxicity detection kit (Roche, Basel, Switzerland) according to the manufacturer’s instructions. The absorbance of samples was measured at 490 nm using a microplate reader (Tecan).

2.5. Real-time RT-PCR HUVECs were incubated with different concentrations of CdCl2 (0, 20, 40, and 80 μM) for the indicated time periods. Total RNA was extracted using RNAiso Plus (Takara, Dalian, China) as described by the manufacturer. The concentrations of RNA were measured by a NanoDrop 2000 spectrophotometer (Thermo Scientific, Waltham, MA, USA). 1 μg of total RNA was reversed transcribed using Prime Script Reverse Transcriptase (Takara) to synthesis cDNA. The RT-PCR reactions were performed using the CFX96TM Real-time System (Bio-Rad, Hercules, CA, USA) with the SYBR Premix Ex Taq II (Tli RNaseH Plus) Kit (Takara). The primer sense and antisense sequences were as follows: IL-1β: forward 5´-CAC GAT GCA CCT GTA CGA TCA-3´, reverse 5´-GTT GCT CCA TAT CCT GTC CCT-3´; TNF-α: forward 5´-GTG ATC GGC CCC CAG AGG GA-3´, reverse 5´-CAC GCC ATT GGC CAG GAG GG-3´; monocyte chemoattractant protein 1 (MCP-1): forward 5´-GGA CCA CCT GGA CAA GCA AA-3´, reverse 5´-GGT GTC TGG GGA AAG CTA GG-3´; IL-6: forward 5´-CCA CTC ACC TCT TCA GAA C-3´, reverse 5´-CTT TGC

TGC TTT CAC ACA T-3´; IL-8: forward 5´-GCA GAG GGT TGT GG A GA A-3´, reverse 5´-ACT GGC ATC TTC ACT GAT TC-3´; GAPDH: forward 5´-TTT GGC TAC AGC AAC AGG GT-3´, reverse 5´-GGG AGA TTC AGT GTG GTG GG-3´.

2.6. Western blot analysis After the indicated treatments, HUVECs were lysed using RIPA Lysis Buffer (Beyotime) and the proteins levels were determined using a BCA protein assay kit (Beyotime). The extracted protein was separated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis and then transferred to polyvinylidene fluoride membranes (Bio-Rad). Membranes were blocked with 5% non-fat milk for 1 h at room temperature and incubated overnight at 4◦C with the following primary antibodies: rabbit anti-cryopyrin (1:400), rabbit anti-pro-caspase-1 (1:400); rabbit anti-caspase-1 (1:400); rabbit anti-IL-1β (1:1000) and mouse anti-β-actin (1:2000, Sigma). The membranes were then incubated with either anti-mouse or anti-rabbit IgG antibody (1:5000, Abcam) for 2 h at room temperature. The bands were visualized using a chemiluminescent substrate (Millipore, Billerica, MA, USA) and detected using Chemi DocTM XRS+ imaging system (Bio-Rad).

2.7. Measurement of intracellular and mitochondrial ROS levels The production of intracellular ROS and mtROS was measured using a Reactive Oxygen Species Assay Kit (Beyotime) and MitoSOXTM Red (Molecular Probes, Eugene, OR, USA) according to the manufacturer’s protocols. After the appropriate treatments, the HUVECs were loaded with DCFH-DA (10 µM) or MitoSOXTM Red (5 µM) in serum-free medium at 37°C for 20 min and protected from light. The cells were then washed three times with phosphate-buffered saline (PBS) and the fluorescence was measured with a microplate reader (Tecan).

2.8. Flow cytometry To assess pyroptosis in HUVECs, active caspase-1 was determined using the FLICA® 660 in vitro Caspase-1 Detection (FLICA 660-YVAD-FMK) Kit as described previously (Alfonso-Loeches et al., 2014; Wree et al., 2014). SYTOX® Green stain (Molecular Probes), a fluorescent nucleic acid dye which only penetrates ruptured cell membranes, was used to mark cells with membrane pore formation. Briefly, after treatment, the cells were harvested and incubated with red caspase-1

detection probe (FLICA 660-YVAD-FMK) for 60 min at 37◦C in the dark. At the end of incubation, the unbound FLICA reagent was washed away using cellular wash buffer. Cells were then stained with 1 μM SYTOX® Green for a further 10 min at 37◦C protecting from light. The cells were then analyzed using a flow cytometer (BD Biosciences, San Jose, CA, USA). Pyroptotic cells were defined as double positive for FLICA 660-YVAD-FMK and SYTOX® Green.

2.9. RNA interference of NLRP3 HUVECs were transfected with either 10 μM NLRP3-targeting siRNA (Santa Cruz) or a control nonspecific siRNA (Santa Cruz) using Lipofectamine 2000 reagent (Invitrogen, Carlsbad, CA, USA) according to the manufacturer’s protocol. The cells were then primed with LPS and exposed to Cd for 24 h. Successful depletion of NLRP3 protein expression was confirmed by western blot analysis.

2.10. Propidium iodide (PI) staining To confirm the pore formation in cell membrane, PI staining were performed after the treatments. The cells were seeded in a 24-well plate and allowed to adhere for 24 h. Following treatment as mentioned above, the cells were washed with PBS and then stained with Hoechst 33342 (5 μl) and PI (5 μl) for 20 min at 37◦C in darkness. Cells with blue and red fluorescence were observed using a fluorescence microscope (Leica, Wetzlar, Germany). PI positive cells were counted in approximately 200 cells from three random microscopic fields for each sample and are expressed as percentages of the total cells.

2.11. Statistical analysis All statistics were calculated using GraphPad Prism 5.0 software (GraphPad Software Inc., La Jolla, CA, USA) and presented as means ± SEM. Each experiment was performed at least three times. Data were compared using one-way analysis of variance or Student’s t-test. Differences were considered statistically significant when p < 0.05.

3. Results 3.1. Cadmium cytotoxicity in HUVECs Previous studies showed that VECs did not appear to be particularly sensitive to Cd-induced

cytotoxicity. Cd-induced death of VECs was observed only when Cd concentrations up to 1–5 μM for 24–72 h in serum free condition, and higher dose of Cd (10 μM–1 mM) was required for serum supplement or shorter exposure period (Prozialeck et al., 2006).Thus, based on previous studies, we exposed HUVECs to 0, 20, 40, and 80 μM CdCl2 for 12 and 24 h after LPS priming, and then cell viability and LDH release were determined. As shown in Figure 1, in the 12 h-incubation protocol, only the highest concentration of CdCl2 (80 μM) resulted in marked cytotoxicity in HUVECs as indicated by decreased cell viability and elevated LDH release. Following treatment with CdCl2 for 24 h, cell viability was decreased to 65% and 25% of the control at 40 μM and 80 μM, respectively. In line with the observed cell viability, LDH release increased 2-fold and 2.5-fold following treatment with 40 μM and 80 μM CdCl2, suggesting that CdCl2 caused cell death. These results indicate that Cd induced significant cytotoxicity in HUVECs in a dose- and time-dependent manner. Accordingly, the moderate cytotoxicity of Cd in HUVECs was observed under the exposure model of 40 μM for 24 h, which was then employed by further study.

3.2. Cadmium induces a pro-inflammatory response in vascular endothelial cells As a rapidly early event, the gene expression of pro-inflammatory cytokines in HUVECs might be affected by Cd exposure. Thus, the mRNA levels of the cytokines were determined after 12 h of treatment with CdCl2. As shown in Figure 2A, the mRNA level of IL-8 was increased by more than 100-fold after exposure to 40 μΜ CdCl2 and more than 200-fold after exposure to 80 μΜ CdCl2 for 12 h. CdCl2 also upregulated the gene expression of TNF-α and IL-6 in a dose-dependent manner (Fig. 2B and C). The real-time PCR results also showed that treatment with 40 and 80 μΜ CdCl2 resulted in a dose-dependent increase in the mRNA levels of IL-1β in HUVECs (Fig. 2D). With regard to the chemokine MCP-1, increased mRNA expression was observed after treatment with the high concentration of CdCl2 (80 μΜ) (Fig. 2E).

3.3. Cadmium treatment results in NLRP3 inflammasome activation in HUVECs Considering the important roles of NLRP3 in endothelial inflammation and dysfunction as mentioned previously, we determined NLRP3 inflammasome activation after Cd treatment in HUVECs. The cleavage of caspase-1 and maturation of IL-1β are the hallmarks of NLRP3 inflammasome activation (Takahashi, 2014), thus we measured the levels of cleaved caspase-1

(Casp1 p20) and 17-kDa mature IL-1β to examine the activation of NLRP3. HUVECs were primed with 200 ng/mL LPS for 3 h followed by incubation with different concentrations of CdCl2 for an additional 24 h. Immunoblotting results showed that, in LPS-primed HUVECs, Cd treatment resulted in a significant dose-dependent increase in the protein expression of Casp1 p20 and 17-kDa mature IL-1β (Fig. 3), which suggested the NLRP3 inflammasome was activated, although the total protein level of NLRP3, pro-caspase-1 (Pro-Casp1), and pro-IL-1β were not elevated (Fig. 3).

3.4. Cadmium induces vascular endothelial cell pyroptosis In view of Cd-induced NLRP3 activation, we also determined whether Cd treatment caused pyroptosis in HUVECs. As previously mentioned, pyroptosis depends on caspase-1 activation, leads to membrane pore formation and cell lysis, resulting in positive dead cell staining and leakage of cellular contents. Therefore, we assessed pyroptosis by determining caspase-1 activation, dead cell staining and LDH release as performed in previous studies (Fink et al., 2008; Wree et al., 2014). Active caspase-1 and pore formation were detected by flow cytometry using FLICA 660-YVAD-FMK and SYTOX Green as described in the methods section. Moreover, PI staining was also performed to confirm membrane rupture. Pyroptosis was determined in LPS-primed HUVECs after exposure to 40 μM CdCl2 for 24 h. To confirm that the cell death induced by Cd was pyroptosis, the caspase-1 inhibitor, Z-YVAD-FMK (YVAD), was used. YVAD effectively abolished caspase-1 cleavage and subsequent IL-1β maturation induced by Cd exposure (Fig. 4A). Flow cytometry results revealed that Cd treatment markedly increased the number of double positive cells for caspase-1 and SYTOX Green, which was reduced by YVAD pre-treatment (Fig. 4C). PI staining also showed that Cd exposure increased PI uptake by HUVECs, and this increase was reduced by YVAD and the specific pyroptosis inhibitor glycine (Fig. 4D). Cd-induced pyroptosis was further confirmed by the results of LDH release which showed the inhibitory effect of YVAD and glycine on Cd-induced LDH release (Fig. 4B).

3.5. NLRP3 inflammasome activation mediates Cd-induced pyroptosis in vascular endothelial cells Despite the fact that Cd induces NLRP3 inflammasome activation which results in active caspase-1 in HUVECs, it was demonstrated that pyroptosis can also be triggered by the activation of

other inflammasomes (Cerqueira et al., 2015). Therefore, we knocked down NLRP3 with a specific siRNA to investigate whether Cd-induced HUVEC pyroptosis is mediated by the NLRP3 inflammasome. Transfection of NLRP3 with siRNA in HUVECs was highly efficient and diminished NLRP3 protein levels (Fig. 5A). Transfected cells were then cultured for 24 h with or without CdCl2 (40 μM) after LPS priming. We found that knockdown of NLRP3 with siRNA reduced the effect of Cd on activation of capase-1 and production of mature IL-1β in LPS-primed HUVECs (Fig. 5B). Moreover, silencing of NLRP3 expression reduced the ability of Cd to induce HUVEC pyroptosis as proved by decreased caspase-1 and SYTOX Green double-positive cells (Fig. 5C), indicating that the NLRP3 inflammasome plays a vital role in caspase-1 activation and pyroptosis induction in our model. Furthermore, we demonstrated that siRNA against NLRP3 significantly decreased membrane pore formation and cell death of HUVECs as shown by the reduction in PI positive cells and LDH release (Fig. 5D, 5E), confirming the role of NLRP3 in pyroptotic cell death induced by Cd.

3.6. Mitochondrial ROS mediate Cd-induced NLRP3 activation and pyroptosis in HUVECs ROS production has been demonstrated to be one of the mechanisms involved in NLRP3 inflammasome activation. Recently, mtROS generation was demonstrated to be particularly critical for NLRP3 activation (Zhou et al., 2011). To determine whether mtROS are required for Cd-induced NLRP3 inflammasome activation in HUVECs, the levels of mtROS and intracellular ROS were determined, and their specific inhibitors were used. Our results showed that Cd treatment promoted intracellular ROS and mtROS generation in a dose-dependent manner (Fig. 6A, 6B). Then, the mtROS was inhibited with a specific mtROS scavenger (Mito-TEMPO) and a total ROS scavenger (NAC) considering that mtROS is one source of intracellular ROS. HUVECs were pre-treated with Mito-TEMPO or NAC, and then exposed to CdCl2 (40 μM) for 24 h in the presence of the inhibitors. Pre-treatment of cells with Mito-TEMPO or NAC inhibited Cd-induced caspase-1 activity and mature IL-1β production (Fig. 6C). And the inhibitors per se did not affect caspase-1 activation and IL-1β expression (Fig. 6D). To investigate the role of mtROS in Cd-induced pyroptosis, we measured caspase-1 and SYTOX Green double-positive cells in the presence or absence of Mito-TEMPO or NAC. As shown, the presence of Mito-TEMPO and NAC markedly decreased pyroptosis induced by Cd (Fig. 6E). In addition, Mito-TEMPO and NAC significantly reduced LDH release and PI uptake induced by Cd exposure (Fig. 6F and G), confirming that Cd causes HUVEC pyroptosis via mtROS

production.

4. Discussion The principal findings of this study are the presence of pyroptotic cell death due to Cd cytotoxicity with hyper-activation of the NLRP3 inflammasome. The identification of pyroptosis in cells exposed to Cd was defined by the presence of both active caspase-1 and SYTOX Green positivity and was confirmed by the inhibiting effects of both a caspase-1 inhibitor (Z-YVAD-FMK) and a pyroptosis inhibitor (glycine) on Cd-induced cell death. To our knowledge, this is the first demonstration of this form of cell death in Cd-induced endothelial toxicity, which may provide a new perspective in understanding cell death due to Cd cytotoxicity. Healthy cells respond to death-inducing stimuli by initiating a wide variety of molecular pathways leading to cell death (Fink and Cookson, 2005). Besides the extensive use of the apoptosis and necrosis paradigm, many other forms of cell death exist, including autophagy, pyroptosis, NETosis, oncosis, pyronecrosis, and necroptosis (LaRock and Cookson, 2013). To date, the mechanism of Cd-induced endothelial injury has mainly concentrated on pro-inflammatory effects, oxidative stress and apoptosis. Previous studies have demonstrated that Cd treatment can result in apoptosis in multiple cells in vitro, such as osteoblasts and hepatocytes (Liu et al., 2014b; Wang et al., 2014a). In VECs, apoptosis has also been reported to be induced by Cd treatment (Kishimoto et al., 1996; Szuster-Ciesielska et al., 2000b). Our results also showed that Cd treatment induced an increase in LDH release in HUVECs. Elevated LDH indicated that, in HUVECs, cell death may be a component of the cytotoxicity induced by Cd. Furthermore, we recognized a novel form of cell death known as pytoptosis after Cd treatment in HUVECs. Pyroptosis is a recently proposed form of cell death which is different from other types of cell death and is characterized by active caspase-1, membrane pore formation and leakage of intracellular contents. Thus, according to our results, Cd cytotoxicity is partially caused by the induction of pyroptosis, at least in VECs. These findings have expanded the current understanding of Cd-induced endothelial toxicity and provide a new therapeutic target for the cardiovascular diseases caused by Cd. In various diseases, especially atherosclerosis, inflammation plays an important role in the initiation and development of disease. Many studies have shown that Cd can cause the production of pro-inflammatory cytokines in various cells and tissues, including vascular cells and tissues (Deng et

al., 2015; Mlynek and Skoczynska, 2005; Prozialeck et al., 2006). The mechanism underlying Cd-induced inflammation is not fully understood. Based on the current study, Cd-induced pyroptosis may be an important mechanism through which Cd causes pro-inflammatory responses in HUVECs. Unlike apoptosis, which is caspase-3 dependent, pyroptosis relies on the activation of caspase-1, a primary pro-inflammatory serine protease. Once activated, caspase-1 cleaves the pro-inflammatory cytokine precursors, pro-IL-1β and pro-IL-18, into their mature forms, allowing their release into the extracellular space (Bergsbaken et al., 2009; Kepp et al., 2010). In addition, membrane pore formation during pyroptosis allows leakage of cytosolic contents (DAMPs) accompanied by abundant pro-inflammatory cytokine production (Lin et al., 2013). This pathway is therefore inherently pro-inflammatory. The present study demonstrated that Cd exposure induced the gene expression of multiple pro-inflammatory cytokines, including IL-6, IL-8, IL-1β, TNF-α and MCP-1 in HUVECs. Therefore, it can be assumed that pyroptosis may be one of the mechanisms through which Cd induces pro-inflammatory responses in cells and tissues, then initiates or promotes pathophysiologic progression, including atherosclerosis. As mentioned above, pyroptosis is dependent on caspase-1 activation, which is regulated by the inflammasomes. There are a number of inflammasomes composed of different subunits, and the best-characterized is the NLRP3 inflammasome. Activation of the NLRP3 inflammasome can be initiated by numerous stimuli, which have been traditionally PAMPs and DAMPs, as well as environmental crystalline substances including silica and asbestos (Tschopp and Schroder, 2010). Recently, Ni2+ was shown to activate the NLRP3 inflammasome in antigen-presenting cells (Li and Zhong, 2014). This finding was consistent with a previous study, in which the authors found that NLRP3 activation induced by multi-walled carbon nanotubes was associated most strongly with nickel contamination on the particles (Hamilton et al., 2012). Recently, Toomey et al. suggested that HgCl2 exposure in B10.S mice led to significant inflammation together with increased expression of the NLRP3 inflammasome and pro-inflammatory cytokines such as IL-1β, TNF-a, and interferon-γ (Toomey et al., 2014). These results suggest that some metal ions may be activators of the NLRP3 inflammasome. In support of this conclusion, our results demonstrated that exposure of HUVECs to Cd led to NLRP3 inflammasome activation, as characterized by caspase-1 activation and mature IL-1β production. Thus, these findings also support Cd-induced pyroptosis in HUVECs. Although the exact mechanism by which caspase-1 causes pyroptosis is unknown, it is clear that active

caspase-1 induces DNA damage and leads to membrane pore formation, cell swelling and eventual cell lysis. Thus, the measurement of caspase-1 activation, LDH release and dead cell staining has been used to indicate pyroptosis (Wree et al., 2014; Yang et al., 2014b). Our data support pyroptosis after Cd exposure with evidence of double positivity for active caspase-1 and SYTOX Green staining (a DNA fluorescent dye which cannot penetrate the membrane), PI uptake and LDH release. Cd-induced pyroptosis was also confirmed by the caspase-1 inhibitor Z-YVAD-FMK which reduced these effects. NLRP3 inflammasome activation has been observed in VECs and implicated in atherosclerosis. Vascular endothelial cells are key sentinel cells which sense pathogens and xenobiotics. The NLRP3 inflammasome has been reported to be activated by ATP and xenobiotic pregnane X receptor (PXR) agonists in HUVECs (Wang et al., 2014b; Yang et al., 2014a). Moreover, NLRP3 activation was proved to promote the formation of foam cells induced by oxidized low density lipoprotein and was required for atherosclerosis progression and activated by cholesterol crystals (Duewell et al., 2010; Liu et al., 2014a). Therefore, Cd-induced NLRP3 activation and subsequent pyroptosis may be a novel link in Cd associated cardiovascular diseases and atherosclerosis. However, caspase-1 is also a subunit of other inflammasomes, such as NLRP1, NLRC4 (NLR family CARD domain-containing protein 4) and AIM2 (absent in melanoma 2) inflammasome (Lamkanfi and Dixit, 2014). The activation of NLRP1 and NLRC4 inflammasomes has been reported to mediate pyroptosis through active caspase-1 (Cerqueira et al., 2015; Masters et al., 2012). In the present study, although Cd exposure induced NLRP3 inflammasome activation and resulted in caspase-1 activation in HUVECs, whether pyroptosis was mediated by NLRP3 activation is unknown and should be confirmed. We have provided evidence to show that knockdown of NLRP3 with a specific siRNA diminished the activation of caspase-1, membrane pore formation and cell death induced by Cd, confirming the role of NLRP3 inflammasome in Cd-induced pyroptosis in HUVECs after LPS priming. Further research to determine whether the activation of other inflammasomes is also involved in Cd-induced pyroptosis is essential. Several common upstream mechanisms implicated in NLRP3 inflammasome activation have been suggested, including potassium efflux, generation of ROS, lysosomal destabilization and release of lysosomal cathepsins (Lamkanfi and Dixit, 2014). In particular, ROS generation has been identified as an important mechanism by which NLRP3 inflammasome is activated and mtROS production

seems to play a critical role in NLRP3 inflammasome activation (Tschopp and Schroder, 2010; Zhou et al., 2011). Indeed, previous studies have demonstrated that Cd-induced ROS generation participates in Cd toxicity in both BEAS-2B cells and BAECs (Jing et al., 2012; Szuster-Ciesielska et al., 2000b). Consistent with these findings, our results also showed that stimulation with Cd resulted in the accumulation of both intracellular ROS and mtROS in HUVECs. The role of mtROS in NLRP3 inflammasome activation was confirmed by data which demonstrated that the inhibition of mtROS, at least partially, diminished both caspase-1 activation and IL-1β maturation induced by Cd, and subsequently reduced Cd-induced pyroptosis in HUVECs. However, previous data showed that potassium efflux and phagosomal destabilization were also involved in inflammasome activation (Cassel and Sutterwala, 2010; Chen and Nunez, 2010; Hornung et al., 2008; Segovia et al., 2012). Whether these well-recognized mechanisms of inflammasome activation also participate in Cd-induced NLRP3 inflammasome activation is unclear. Further investigation into the upstream mechanisms involved in Cd-triggered NLRP3 inflammasome activation in VECs is warranted. In conclusion, the present study showed the Cd-induced caspase-1-dependent pyroptosis and inflammatory response in vascular endothelial cells. We demonstrated, for the first time, that activation of the NLRP3 inflammasome contributes to Cd-induced pyroptosis in HUVECs. Moreover, the generation of ROS is involved in Cd-induced activation of the NLRP3 inflammasome and pyroptosis. Our data could be the basis for further extensive study of these mechanisms in animal models and may provide novel methods to improve occupational health protection, and the prevention and chemotherapy of cardiovascular disease caused by Cd.

Acknowledgments This work was financially supported by the National Natural Science Foundation of China (Grant Numbers: 81070101 and 31200627).

References Abderrazak, A., Syrovets, T., Couchie, D., El Hadri, K., Friguet, B., Simmet, T., Rouis, M., 2015. NLRP3 inflammasome: from a danger signal sensor to a regulatory node of oxidative stress and inflammatory diseases. Redox biology 4, 296-307. Alfonso-Loeches, S., Urena-Peralta, J.R., Morillo-Bargues, M.J., Oliver-De La Cruz, J., Guerri, C., 2014. Role of mitochondria ROS generation in ethanol-induced NLRP3 inflammasome activation and cell death in astroglial cells. Frontiers in cellular neuroscience 8, 216. Angeli, J.K., Cruz Pereira, C.A., de Oliveira Faria, T., Stefanon, I., Padilha, A.S., Vassallo, D.V., 2013. Cadmium exposure induces vascular injury due to endothelial oxidative stress: the role of local angiotensin II and COX-2. Free radical biology & medicine 65, 838-848. Bergsbaken, T., Fink, S.L., Cookson, B.T., 2009. Pyroptosis: host cell death and inflammation. Nature reviews Microbiology 7, 99-109. Bernhoft, R.A., 2013. Cadmium toxicity and treatment. TheScientificWorldJournal 2013, 394652. Cassel, S.L., Sutterwala, F.S., 2010. Sterile inflammatory responses mediated by the NLRP3 inflammasome. European journal of immunology 40, 607-611. Cerqueira, D.M., Pereira, M.S., Silva, A.L., Cunha, L.D., Zamboni, D.S., 2015. Caspase-1 but Not Caspase-11 Is Required for NLRC4-Mediated Pyroptosis and Restriction of Infection by Flagellated Legionella Species in Mouse Macrophages and In Vivo. Journal of immunology 195, 2303-2311. Chang, W., Lin, J., Dong, J., Li, D., 2013. Pyroptosis: an inflammatory cell death implicates in atherosclerosis. Medical hypotheses 81, 484-486. Chen, G.Y., Nunez, G., 2010. Sterile inflammation: sensing and reacting to damage. Nature reviews Immunology 10, 826-837. Deng, Z.Y., Hu, M.M., Xin, Y.F., Gang, C., 2015. Resveratrol alleviates vascular inflammatory injury by inhibiting inflammasome activation in rats with hypercholesterolemia and vitamin D2 treatment. Inflammation research : official journal of the European Histamine Research Society [et al] 64, 321-332. Dong, F., Guo, F., Li, L., Guo, L., Hou, Y., Hao, E., Yan, S., Allen, T.D., Liu, J., 2014. Cadmium induces vascular permeability via activation of the p38 MAPK pathway. Biochemical and biophysical research communications 450, 447-452. Dong, Z., Wang, L., Xu, J., Li, Y., Zhang, Y., Zhang, S., Miao, J., 2009. Promotion of autophagy and inhibition of apoptosis by low concentrations of cadmium in vascular endothelial cells. Toxicology in vitro : an international journal published in association with BIBRA 23, 105-110.

Duewell, P., Kono, H., Rayner, K.J., Sirois, C.M., Vladimer, G., Bauernfeind, F.G., Abela, G.S., Franchi, L., Nunez, G., Schnurr, M., Espevik, T., Lien, E., Fitzgerald, K.A., Rock, K.L., Moore, K.J., Wright, S.D., Hornung, V., Latz, E., 2010. NLRP3 inflammasomes are required for atherogenesis and activated by cholesterol crystals. Nature 464, 1357-1361. Fink, S.L., Bergsbaken, T., Cookson, B.T., 2008. Anthrax lethal toxin and Salmonella elicit the common cell death pathway of caspase-1-dependent pyroptosis via distinct mechanisms. Proceedings of the National Academy of Sciences of the United States of America 105, 4312-4317. Fink, S.L., Cookson, B.T., 2005. Apoptosis, pyroptosis, and necrosis: mechanistic description of dead and dying eukaryotic cells. Infection and immunity 73, 1907-1916. Hamilton, R.F., Jr., Buford, M., Xiang, C., Wu, N., Holian, A., 2012. NLRP3 inflammasome activation in murine alveolar macrophages and related lung pathology is associated with MWCNT nickel contamination. Inhalation toxicology 24, 995-1008. Hornung, V., Bauernfeind, F., Halle, A., Samstad, E.O., Kono, H., Rock, K.L., Fitzgerald, K.A., Latz, E., 2008. Silica crystals and aluminum salts activate the NALP3 inflammasome through phagosomal destabilization. Nature immunology 9, 847-856. Jing, Y., Liu, L.Z., Jiang, Y., Zhu, Y., Guo, N.L., Barnett, J., Rojanasakul, Y., Agani, F., Jiang, B.H., 2012. Cadmium increases HIF-1 and VEGF expression through ROS, ERK, and AKT signaling pathways and induces malignant transformation of human bronchial epithelial cells. Toxicological sciences : an official journal of the Society of Toxicology 125, 10-19. Kaji, T., Mishima, A., Machida, M., Yabusaki, K., Suzuki, M., Yamamoto, C., Fujiwara, Y., Sakamoto, M., Kozuka, H., 1995. Comparative cytotoxicity of exogenous cadmium-metallothionein and cadmium ion in cultured vascular endothelial cells. Bulletin of environmental contamination and toxicology 54, 501-506. Kepp, O., Galluzzi, L., Zitvogel, L., Kroemer, G., 2010. Pyroptosis - a cell death modality of its kind? European journal of immunology 40, 627-630. Kim, J., Lim, W., Ko, Y., Kwon, H., Kim, S., Kim, O., Park, G., Choi, H., Kim, O., 2012. The effects of cadmium on VEGF-mediated angiogenesis in HUVECs. Journal of applied toxicology : JAT 32, 342-349. Kim, M.S., Kim, B.J., Woo, H.N., Kim, K.W., Kim, K.B., Kim, I.K., Jung, Y.K., 2000. Cadmium induces caspase-mediated cell death: suppression by Bcl-2. Toxicology 145, 27-37. Kishimoto, T., Oguri, T., Yamabe, S., Tada, M., 1996. Effect of cadmium injury on growth and migration of cultured human vascular endothelial cells. Human cell 9, 43-48. Knoflach, M., Messner, B., Shen, Y.H., Frotschnig, S., Liu, G., Pfaller, K., Wang, X., Matosevic, B., Willeit, J., Kiechl, S., Laufer, G., Bernhard, D., 2011. Non-toxic cadmium concentrations induce vascular inflammation and promote

atherosclerosis. Circulation journal : official journal of the Japanese Circulation Society 75, 2491-2495. Lamkanfi, M., Dixit, V.M., 2014. Mechanisms and functions of inflammasomes. Cell 157, 1013-1022. LaRock, C.N., Cookson, B.T., 2013. Burning down the house: cellular actions during pyroptosis. PLoS pathogens 9, e1003793. Li, X., Deroide, N., Mallat, Z., 2014. The role of the inflammasome in cardiovascular diseases. Journal of molecular medicine 92, 307-319. Li, X., Zhong, F., 2014. Nickel induces interleukin-1beta secretion via the NLRP3-ASC-caspase-1 pathway. Inflammation 37, 457-466. Lin, J., Shou, X., Mao, X., Dong, J., Mohabeer, N., Kushwaha, K.K., Wang, L., Su, Y., Fang, H., Li, D., 2013. Oxidized low density lipoprotein induced caspase-1 mediated pyroptotic cell death in macrophages: implication in lesion instability? PloS one 8, e62148. Liu, W., Yin, Y., Zhou, Z., He, M., Dai, Y., 2014a. OxLDL-induced IL-1 beta secretion promoting foam cells formation was mainly via CD36 mediated ROS production leading to NLRP3 inflammasome activation. Inflammation research : official journal of the European Histamine Research Society [et al] 63, 33-43. Liu, W., Zhao, H., Wang, Y., Jiang, C., Xia, P., Gu, J., Liu, X., Bian, J., Yuan, Y., Liu, Z., 2014b. Calcium-calmodulin signaling elicits mitochondrial dysfunction and the release of cytochrome c during cadmium-induced apoptosis in primary osteoblasts. Toxicology letters 224, 1-6. Masters, S.L., Gerlic, M., Metcalf, D., Preston, S., Pellegrini, M., O'Donnell, J.A., McArthur, K., Baldwin, T.M., Chevrier, S., Nowell, C.J., Cengia, L.H., Henley, K.J., Collinge, J.E., Kastner, D.L., Feigenbaum, L., Hilton, D.J., Alexander, W.S., Kile, B.T., Croker, B.A., 2012. NLRP1 inflammasome activation induces pyroptosis of hematopoietic progenitor cells. Immunity 37, 1009-1023. Messner, B., Bernhard, D., 2010. Cadmium and cardiovascular diseases: cell biology, pathophysiology, and epidemiological relevance. Biometals : an international journal on the role of metal ions in biology, biochemistry, and medicine 23, 811-822. Mlynek, V., Skoczynska, A., 2005. The proinflammatory activity of cadmium. Postepy higieny i medycyny doswiadczalnej 59, 1-8. Nagarajan, S., Rajendran, S., Saran, U., Priya, M.K., Swaminathan, A., Siamwala, J.H., Sinha, S., Veeriah, V., Sonar, P., Jadhav, V., Jaffar Ali, B.M., Chatterjee, S., 2013. Nitric oxide protects endothelium from cadmium mediated leakiness. Cell biology international 37, 495-506. Nawrot, T.S., Staessen, J.A., Roels, H.A., Munters, E., Cuypers, A., Richart, T., Ruttens, A., Smeets, K., Clijsters, H.,

Vangronsveld, J., 2010. Cadmium exposure in the population: from health risks to strategies of prevention. Biometals : an international journal on the role of metal ions in biology, biochemistry, and medicine 23, 769-782. Prozialeck, W.C., Edwards, J.R., Woods, J.M., 2006. The vascular endothelium as a target of cadmium toxicity. Life Sci 79, 1493-1506. Satarug, S., Moore, M.R., 2012. Emerging roles of cadmium and heme oxygenase in type-2 diabetes and cancer susceptibility. The Tohoku journal of experimental medicine 228, 267-288. Schroder, K., Tschopp, J., 2010. The inflammasomes. Cell 140, 821-832. Segovia, J., Sabbah, A., Mgbemena, V., Tsai, S.Y., Chang, T.H., Berton, M.T., Morris, I.R., Allen, I.C., Ting, J.P., Bose, S., 2012. TLR2/MyD88/NF-kappaB pathway, reactive oxygen species, potassium efflux activates NLRP3/ASC inflammasome during respiratory syncytial virus infection. PloS one 7, e29695. Szuster-Ciesielska, A., Lokaj, I., Kandefer-Szerszen, M., 2000a. The influence of cadmium and zinc ions on the interferon and tumor necrosis factor production in bovine aorta endothelial cells. Toxicology 145, 135-145. Szuster-Ciesielska, A., Stachura, A., Slotwinska, M., Kaminska, T., Sniezko, R., Paduch, R., Abramczyk, D., Filar, J., Kandefer-Szerszen, M., 2000b. The inhibitory effect of zinc on cadmium-induced cell apoptosis and reactive oxygen species (ROS) production in cell cultures. Toxicology 145, 159-171. Takahashi, M., 2014. NLRP3 inflammasome as a novel player in myocardial infarction. International heart journal 55, 101-105. Tellez-Plaza, M., Guallar, E., Howard, B.V., Umans, J.G., Francesconi, K.A., Goessler, W., Silbergeld, E.K., Devereux, R.B., Navas-Acien, A., 2013a. Cadmium exposure and incident cardiovascular disease. Epidemiology 24, 421-429. Tellez-Plaza, M., Jones, M.R., Dominguez-Lucas, A., Guallar, E., Navas-Acien, A., 2013b. Cadmium exposure and clinical cardiovascular disease: a systematic review. Current atherosclerosis reports 15, 356. Toomey, C.B., Cauvi, D.M., Hamel, J.C., Ramirez, A.E., Pollard, K.M., 2014. Cathepsin B regulates the appearance and severity of mercury-induced inflammation and autoimmunity. Toxicological sciences : an official journal of the Society of Toxicology 142, 339-349. Tschopp, J., Schroder, K., 2010. NLRP3 inflammasome activation: The convergence of multiple signalling pathways on ROS production? Nature reviews Immunology 10, 210-215. Wang, J., Zhu, H., Liu, X., Liu, Z., 2014a. Oxidative stress and Ca(2+) signals involved on cadmium-induced apoptosis in rat hepatocyte. Biological trace element research 161, 180-189. Wang, S., Lei, T., Zhang, K., Zhao, W., Fang, L., Lai, B., Han, J., Xiao, L., Wang, N., 2014b. Xenobiotic pregnane X receptor (PXR) regulates innate immunity via activation of NLRP3 inflammasome in vascular endothelial cells. The Journal of

biological chemistry 289, 30075-30081. Wree, A., Eguchi, A., McGeough, M.D., Pena, C.A., Johnson, C.D., Canbay, A., Hoffman, H.M., Feldstein, A.E., 2014. NLRP3 inflammasome activation results in hepatocyte pyroptosis, liver inflammation, and fibrosis in mice. Hepatology 59, 898-910. Yang, H., Xiao, L., Yuan, Y., Luo, X., Jiang, M., Ni, J., Wang, N., 2014a. Procyanidin B2 inhibits NLRP3 inflammasome activation in human vascular endothelial cells. Biochemical pharmacology 92, 599-606. Yang, H.H., Jun, H.K., Jung, Y.J., Choi, B.K., 2014b. Enterococcus faecalis activates caspase-1 leading to increased interleukin-1 beta secretion in macrophages. Journal of endodontics 40, 1587-1592. Zhang, R., Witkowska, K., Ng, F., Caulfield, M.J., Ye, S., 2015a. Lb03.08: Hypertension Related Variant of Solute Carrier Family 39 Member 8 Gene Influences Cadmium Uptake and Cell Toxicity. Journal of hypertension 33 Suppl 1, e128. Zhang, Y., Li, X., Pitzer, A.L., Chen, Y., Wang, L., Li, P.L., 2015b. Coronary endothelial dysfunction induced by nucleotide oligomerization domain-like receptor protein with pyrin domain containing 3 inflammasome activation during hypercholesterolemia: beyond inflammation. Antioxidants & redox signaling 22, 1084-1096. Zhou, R., Yazdi, A.S., Menu, P., Tschopp, J., 2011. A role for mitochondria in NLRP3 inflammasome activation. Nature 469, 221-225.

Fig. 1. Cytotoxicity of Cd in vascular endothelial cells. HUVECs were exposed to different concentrations of CdCl2 for 12 and 24 h. Cell viability was determined using the CCK-8 Test Kit and LDH release was determined using the LDH Cytotoxicity Detection Kit. Values are mean ± SEM for three independent experiments. *, #P< 0.05 and **, ##P< 0.01 vs. the corresponding control group (con).

Fig. 2. mRNA expression of inflammatory cytokines induced by Cd treatment. (A) – (E) HUVECs were incubated with different concentrations of CdCl2 (0, 20, 40 and 80 μM). The expression of (A) IL-8, (B) TNF-α, (C) IL-6, (D) IL-1β and (E) MCP-1 was measured using real-time RT-PCR at 12 h after CdCl2 exposure. Values are mean ± SEM for three independent experiments. *p < 0.05 and **P < 0.01 vs. control groups (con).

Fig. 3. Cd induces caspase-1 activation and IL-1β production. HUVECs were primed with 200 ng/mL LPS for 3 h followed by stimulation with different concentrations of CdCl2 (0, 20, 40 and 80 μM) for an additional 24 h. Upper panel: representative western blotting bands for NLRP3, pro-caspase-1 (Pro-Casp1), cleaved caspase-1 (Casp1 p20), pro-IL-1β and IL-1β. Lower panel: photodensitometry of western blotting images. Values are mean ± SEM for three independent experiments. *p < 0.05 and **p < 0.01 vs. the control group (0 μM).

Fig. 4. Cd induces vascular endothelial cell pyroptosis. HUVECs were primed with 200 ng/mL LPS for 3 h, then incubated with Z-YVAD-FMK (YVAD) (20 μM) or glycine (5 mM) before and during CdCl2 (40 μM, 24 h) treatment. (A) Immunoblotting analysis of cleaved caspase-1 (Casp1 p20) and IL-1β. (B) Effect of YVAD and glycine on Cd-induced LDH release. (C) Membrane pore formation (SYTOX Green staining, horizontal) and caspase-1 activation (FLICA 660-YVAD-FMK staining, vertical) measured by flow cytometry. (D) Photomicrographs of double-fluorescent staining with PI (red) and Hoechst 33342 (blue). Values are mean ± SEM for three independent experiments. Scale bar indicates 100 μm. *p < 0.05 and **p < 0.01 vs. the control group (con), #p < 0.05 and ##p < 0.01 vs. the Cd group.

Fig. 5. Role of the NLRP3 inflammasome in Cd-induced pyroptosis in vascular endothelial cells. (A) Efficiency of siRNA against NLRP3 tested by western blot. (B) Effect of NLRP3 silencing on Cd-induced cleaved caspase-1 (Casp1 p20) and IL-1β measured by western blot. (C) Effect of siRNA against NLRP3 on Cd-induced caspase-1 (vertical) and SYTOX Green (horizontal) double positivity. (D) PI staining after treatments with Cd in HUVECs transfected with control (con) and NLRP3 targeting siRNA. (E) Effect of NLRP3 silencing on Cd-induced LDH release in HUVECs. Values are mean ± SEM for three independent experiments. Scale bar indicates 100 μm. *p < 0.05 and **p < 0.01 vs. the si-control group (si-con), #p < 0.05 and ##p < 0.01 vs. the si-control + Cd group (si-con + Cd).

Fig. 6. Role of mtROS in Cd-induced NLRP3 activation and pyroptosis in HUVECs. HUVECs were incubated with different doses of CdCl2 for 24 h, and intracellular ROS (A) and mtROS (B) were then analyzed. (C) Effects of Mito-TEMPO (TEMPO) and NAC on Cd-induced caspase-1 activation (Casp1 p20) and IL-1β expression. (D) Immunoblotting analysis of Casp1 p20 and IL-1β after YVAD, TEMPO and NAC alone treatments. (E) Effects of TEMPO and NAC on Cd-induced double positivity for caspase-1 (vertical) and SYTOX Green (horizontal) in HUVECs. (F) LDH release induced by Cd treatment was alleviated by TEMPO and NAC. (G) TEMPO and NAC attenuated Cd-induced PI positivity in HUVECs. Values are mean ± SEM for three independent experiments. Scale bar indicates 100 μm. *p < 0.05 and **p < 0.01 vs. the control group (con), #p < 0.05 and ##p < 0.01 vs. the Cd group.