Cadmium Induces Glomerular Endothelial Cell–Specific Expression of Complement Factor H via the −1635 AP-1 Binding Site

Cadmium (Cd) is a ubiquitous environmental pollutant, and the kidney is one of the primary targets of Cd toxicity (1, 2). With chronic exposure, Cd accumulates in epithelial cells of the proximal tubule, leading to a loss of reabsorptive capacity (3). In addition, Cd disrupts glomerular function (4). In the kidney, the glomerulus and its surrounding Bowman capsule constitute the basic filtration unit (5). The glomerular filtration barrier, composed of glomerular endothelial cells (GECs), glomerular basement membrane, and podocytes, allows the efficient flow of water and small solutes while preventing the passage of plasma proteins (6). The mesangial cells surrounding the glomerulus provide structural support for the capillary loops (6). After entering the human body, Cd circulates in the blood as free ion or bound with plasma proteins such as metallothionein (MT), therefore directly affecting the glomerular endothelium (7, 8). Previous studies have shown that low-dose Cd does not trigger cell death but induces hyperpermeability in GECs (7, 9). However, the mechanisms underlying the effects of Cd on the glomerulus are still unclear.

The complement system is a major component of innate immunity and defends the host against infectious microorganisms (10). It might also be activated inappropriately and function as an inflammatory mediator (11). Complement factor H (CFH) is a soluble 155-kDa protein and a negative regulator of the complement alternative pathway (12). With multiple binding sites for complement component C3b, CFH competes with factor B for C3b binding and prevents the formation of the alternative pathway C3 convertase (13). CFH also accelerates C3 convertase decay (14). In addition, CFH functions in cell adhesion, migration, and immune cell recruitment (15–17).

Defects in CFH result in uncontrolled C3 activation and excessive complement activation, leading to the development of C3 glomerulopathy, which is characterized by low serum C3 levels and C3 deposition in glomeruli (18, 19). CFH gene variants are associated with several diseases, including various renal diseases such as atypical hemolytic uremic syndrome (20), membranoproliferative glomerulonephritis type I, and dense deposit disease (21, 22). In addition, a decrease in CFH activity also leads to tubulointerstitial injury, as it binds to tubular epithelial cells and inhibits complement activation in ischemic renal injury (23). Mice with a deletion of the Cfh gene were hypersensitive to immune-induced renal injury and showed accumulation of C3 along the glomerular basement membrane, which was resolved by administration of recombinant murine or human CFH (19, 24). Thus, CFH is an essential factor in complement-mediated renal diseases.

CFH is mainly produced in the liver and secreted into blood plasma, but it is also expressed by endothelial cells (25). Endothelial cells form the inner cellular lining of blood vessels and display heterogeneous gene expression patterns in each vascular bed to adapt to the underlying tissues (26). The local production of CFH from endothelial cells directly affects the vascular beds and thus contributes to the immune homeostasis of the microenvironment. CFH production from endothelial cells can be regulated by inflammatory cytokines such as TNF-α, IL-1β, and IFN-γ (27). To date, few studies have addressed the regulatory mechanisms of CFH expression.

In this study, we explored the effects of Cd on CFH production from the kidney. Our results showed that Cd distinctively increases Cfh expression in mouse glomeruli. In vitro, Cd does not induce cytotoxicity in human renal GECs (HRGECs) but specifically induces CFH expression in GECs via activation of the JNK pathway. In addition, the −1635 AP-1 motif on the CFH promoter mediates Cd-inducible CFH gene expression.

C57B6/J male mice (10- to 12-wk-old; Vital River Laboratory, Beijing, China) weighing 25 ± 2 g were used for this study. The mice were administered 50 mg/kg CdCl₂ (C3141; Sigma-Aldrich, St. Louis, MO) in distilled water through a gastric tube at 0, 24, or 48 h, and mice from the control group were administered distilled water in the same manner for the same period. At 24, 48, and 72 h (24 h after the daily Cd treatment), the mice were anesthetized with 3% sodium pentobarbital (i.p., 20 mg/kg) (Sigma-Aldrich), and the blood samples were drawn directly from the renal vein or the heart. The mice were killed by exsanguination, and the kidneys of the mice were removed and snap-frozen in liquid nitrogen for further analyses. The urine samples were collected after 72-h Cd treatment. Creatinine levels were detected using an enzymatic method of creatinine plus ver.2 (CREP2) kit according to the manufacturer’s protocol. All the procedures were approved by the Animal Care Committee of Shandong Provincial Qianfoshan Hospital, Shandong University, and performed in accordance with guidelines prescribed by the Animal Care Committee of Shandong University.

HRGECs were obtained from ScienCell Research Laboratories (catalog no. 4000; Carlsbad, CA) and cultured in basal endothelial cell medium supplemented with the EGM-2-MV bullet kit (CC-3156; Lonza, Basel, Switzerland). Human renal mesangial cells (HRMCs) were obtained from ScienCell Research Laboratories (catalog no. 4200) and cultured in DMEM (10-017; Corning, Corning, NY) with 10% FBS (10099-141; Lonza). Human renal podocytes were obtained from Chi Scientific (Maynard, MA) and cultured in RPMI 1640 medium (10–104; Corning) with 10% FBS. All the media contained antibiotics (100 IU/ml penicillin and 100 μg/ml streptomycin). The cell culture was placed in humidified air at 37°C with 5% CO₂. Pyrrolidine dithiocarbamate (PDTC) (sc-253594) and GSK-690693 (sc-363280) were purchased from Santa Cruz Biotechnology (Santa Cruz, CA). PD98059 (9900), SP600125 (8177), and SB203850 (5633) were purchased from Cell Signaling Technology (Danver, MA).

The frozen samples of mouse kidney were sectioned at 10 μm on aminoalylsilane-coated slides (HistoGene; Arcturus Engineering, Mountain View, CA). Laser capture microdissection (LCM) collection was performed by the Arcturus microdissection system (Life Technologies) (28). Specific tissues were collected by transferring them from tissue sections to an adhesive infrared-activated polymer on the CapSure HS LCM Cap (Life Technologies). Total RNA was extracted from the pooled samples with PicoPure RNA Isolation Kit (KIT0204; Life Technologies) according to the manufacturer’s instructions.

HRGEC proliferation was evaluated by an MTT assay kit (Cayman Chemical Company, Ann Arbor, MI) and CCK8 kit (Dojindo, Shanghai, China) following the manufacturer’s protocol. Briefly, the HRGECs were plated with a density of 1 × 10⁴ cells per well in a 96-well plate, and the next day the cells were treated with 4 μM Cd for 24 h. For MTT assay, the staining solution (10 μl) was added to each well and incubated for 4 h. The crystals were solubilized by the addition of 110 μl of formazan dissolving solution. After shaking at room temperature to ensure a homogeneous solution, the OD were determined using a 96-well plate reader (Molecular Devices, Sunnyvale, CA) at a wavelength of 490 nm. For CCK8 assays, 10 μl of staining solution was added into each well. After 1 h of incubation, the OD was detected by a microplate reader at a wavelength of 450 nm (Molecular Devices).

Cell viability was assessed by trypan blue exclusion assay and lactate dehydrogenase (LDH) release assay. After Cd treatment for 24 h, HRGECs were washed and incubated in 0.05% trypsin for 2 min at 37°C. The cells were disaggregated into single-cell suspension and were diluted 9:1 with 0.4% trypan blue (Solarbio Science & Technology, Beijing, China). The percentage of dye-free cells was calculated under a microscope. LDH release assay was performed using a Cytotoxicity LDH Assay Kit-WST (Dojindo) following the manufacturer’s instructions. Briefly, after Cd treatment, 10 μl of lysis buffer was added to each well and incubated at 37°C for 30 min. Then, 100 μl of working solution was added to each well and incubated at the room temperature for 20 min. Finally, 50 μl of stop solution was added to each well. The release of LDH was measured by a microplate reader at a wavelength of 490 nm (Molecular Devices) and presented as the percentage of maximal LDH activity.

Apoptosis of HRGECs was determined by annexin V–FITC and propidium iodide (PI) staining using an assay kit (Neobiosciences, Shenzhen, China) according to the manufacturer’s protocol. Briefly, The HRGECs were cultured in six-well plates. After 24-h Cd treatment, cells were harvested, washed twice with cold PBS, and resuspended into single-cell suspension. Then, 1 × 10⁶ cells were stained with annexin V–FITC (0.025%) for 3 min and PI (20 μg/ml) for 10 min. Positive staining of the HRGECs was detected using a FACSAria II flow cytometer (BD Biosciences, San Jose, CA), and the data were analyzed using the FACSDIVA acquisition and analysis software (BD Biosciences).

Quantitative real-time PCR was performed as previously described (7). Total RNA of HRGECs, human renal podocytes, and HRMCs were extracted with the E.Z.N.A. Total RNA Kit II (R6934-02; OMEGA Bio-tek, Norcross, GA) following the manufacturer’s protocol. The cDNA synthesis was performed using the RevertAid First Strand cDNA Synthesis Kit (K1622; Thermo Fisher Scientific, Grand Island, NY). Quantitative real-time PCR was performed using a ViiA7 Real-Time PCR System (Applied Biosystems, Waltham, MA). Reaction conditions were 95°C for 15 min, 40 cycles of 95°C for 10 s, and 60°C for 32 s. Relative expression was calculated using GAPDH as an endogenous internal control. The following primers were used: 5′-GTGAAGTGTTTACCAGTGACAGC-3′ and 5′-AACCGTACTGCTTGTCCAAA-3′ for human CFH; 5′-GCCCGGTAGAGTGTATGGGCCAG-3′ and 5′-GCTCTGCCCCACTCCTGCCTT-3′ for human F3; 5′-TGATGACATCAAGAAGGTGGTGAAG-3′ and 5′-TCCTTGGAGGCCATGTGGGCCAT-3′ for human GAPDH; 5′-AGAGATTCCATTGAGTCC-3′ and 5′-ATGTCACTTGTTCTCCTGTCC-3′ for mouse Cfh; and 5′-GGATGCAGAAGGAGATTACTGC-3′ and 5′-CCACCGATCCACACAGAGTA-3′ for mouse β-actin.

At each time point of Cd treatment, HRGECs were lysed using RIPA buffer (20 mM Tris pH 7.5, 150 mM NaCl, 50 mM NaF, 1% NP40, 0.1% deoxycholate, 0.1% NaDodSO₄, and 1 mM EDTA) supplemented with 1 μg/ml aprotinin, 10 μg/ml leupeptin, and 1 mM PMSF. Equal amounts of protein extracts were electrophoresed through an 8% NaDodSO₄–polyacrylamide gel and then transferred to a PVDF membrane (IPVH00010; Sigma-Aldrich). The membranes were blocked with 5% nonfat milk in TBST (20 mM Tris, 150 mM NaCl, 0.1% Tween 20, pH 7.5) at room temperature for 3 h and incubated overnight at 4°C with primary Abs. After washing three times with TBST, the membranes were incubated with the secondary Ab (1:6000) at room temperature for 2 h. The primary Abs were rabbit anti-SAPK/JNK (9258), rabbit anti–phospho-SAPK/JNK (4668), rabbit anti–c-Fos (2250), rabbit anti–c-Jun (9165), rabbit anti-GAPDH (2118) (Cell Signaling Technology), and a mouse mAb against human CFH (ab118820, OX24; Abcam, Cambridge, MA). The anti-CFH Ab recognizes complement control protein domain 5 (CCP5) of the CFH protein (29), and the same domain of factor H–like protein 1 (FHL-1), which is also derived from the CFH gene through alternative splicing (30). The secondary Abs were human renal podocyte–conjugated goat anti-rabbit IgG (SA00001-2) and goat anti-mouse IgG (SA00001-1) (Proteintech, Chicago, IL). The immunoreactive signals were visualized with ECL reagents (WBKLS0100; Millipore, Billerica, MA), and the densitometry analysis was performed with ImageJ software (National Institutes of Health, Bethesda, MD).

Blood samples were centrifuged for 15 min at 1000 × g at 4°C, and the plasma was collected for detection of mouse Cfh protein. Cells were treated with serum-free basal endothelial cell media containing 4 μM Cd, and the media were collected after 0-, 1-, 2-, 6-, 12-, and 24-h exposure. The ELISAs of mouse plasma and culture media were performed using CFH Quantikine kit (LS-F14942; LifeSpan BioSciences, Seattle, WA) or CFH Quantikine ELISA kit (ab137975; Abcam) according to the manufacturer’s protocol (31). Briefly, an aliquot of 100 μl of standard, media, or plasma samples was added to each well and incubated for 2 h covered with a plate sealer at 37°C. After aspiration, Detection Reagent A was added to each well for 1 h. Then, the wells were washed three times, and Detection Reagent B was added to each well and incubated for 1 h. After washing, 90 μl of 3, 3′, 5, 5′-tetramethybenzidine substrate solution was added to each well and incubated for 15–30 min at 37°C. Finally, 50 μl of stop solution was added to each well. The OD (OD value) was immediately determined using a microplate reader (Molecular Devices) at a wavelength of 450 nm.

HRGEC monolayers were grown on fibronectin-coated glass chamber slides and were treated with 4 μM Cd for 24 h. The medium was aspirated, and the monolayers were washed with PBS, fixed with 4% paraformaldehyde for 10 min, and washed three times with PBS. Then, the cells were permeabilized with 0.1% Triton X-100 for 10 min and washed three times with PBS. Immunofluorescence was performed by staining with a mouse mAb against human CFH (1:50; ab118820; Abcam) overnight at 4°C and a tetramethylrhodamine-conjugated rabbit anti-mouse IgG (1:250; CW0222; Boster, Wuhan, China) for 1 h at room temperature. The cells were photographed using an Olympus FSX100 Imaging System (Olympus Corporation, Tokyo, Japan) with an excitation wavelength of 546 nm.

HRGECs were fixed with 1% paraformaldehyde for 10 min and then treated with 0.2 M glycine for 5 min. The cells were then washed twice with PBS containing an EDTA-free protease inhibitor mixture (Roche, Basel, Switzerland) and collected by a cell scraper. After centrifugation, the cell pellets were sonicated with a Scientz Sonifier to shear the chromatin into 300–600-bp fragments. Immunoprecipitation was performed using a chromatin immunoprecipitation (ChIP) assay kit (catalog no. 17-371; Upstate Biotechnology, Lake Placid, NY) according to the manufacturer’s instructions (32). The chromatin fragments were immunoprecipitated with the Ab against c-Jun and c-Fos (Cell Signaling Technology). The DNA fragments were detected by semiquantitative PCR. The sequences of the primers are listed in Supplemental Table I.

Nuclear protein extracts were prepared from the HRGECs using Pierce NE-PER Nuclear and Cytoplasmic Extraction Reagents (Thermo Fisher Scientific). Briefly, HRGECs were transferred to a 1.5-ml microcentrifuge tube and pelleted by centrifugation at 500 × g for 2–3 min. After discarding the supernatant, ice-cold Cytoplasmic Extraction Reagent I was added to the cell pellet. Then, the tube was vortexed vigorously at 12,000 × g for 15 s to fully suspend the cell pellet and incubated on ice for 10 min before addition of ice-cold Cytoplasmic Extraction Reagent II. After centrifugation and transfer of the supernatant (cytoplasmic extract), the insoluble (pellet) fraction was suspended in ice-cold Nuclear Extraction Reagent. Then, the tube was vortexed and centrifuged at 12,000 × g in a microcentrifuge for 10 min. Finally the supernatant (nuclear extract) fraction was transferred to a clean prechilled tube. EMSA was performed using a LightShift Chemiluminescent EMSA Kit (Thermo Fisher Scientific) (33). The probe sequence was 5′-TGGGAGGCCTGACACTGGGCGAAG-3′. Equal amounts of nuclear extract were incubated with the biotin-labeled double-stranded probes and poly(dI:dC) for 20 min in binding reaction buffer. The DNA–protein complexes were electrophoresed through a 6% polyacrylamide gel and transferred onto a positively charged nylon membrane (Thermo Fisher Scientific). The membrane was then cross-linked with UV radiation and visualized by chemiluminescence reagents (Thermo Fisher Scientific).

The -2200 to +298 fragment of the CFH promoter was generated by DNA synthesis. The −1635 AP-1 mutation (5′-TGACACT-3′ to 5′-GTCACAG-3′) on the CFH promoter fragment was generated by PCR and cloned into pGL3-basic vector (Promega, Madison, WI). The resulting pGL3-CFH (wild-type) and pGL3-CFH (mutant) constructs were transfected into HRGECs using Lipofectamine 3000 (Thermo Fisher Scientific) following the manufacturer’s protocol. A plasmid containing the Renilla luciferase reporter gene under the control of a CMV enhancer/promoter (Promega) was used as an internal control. At 48 h, 4 μM Cd was applied to the culture, and the cells were collected by Passive Lysis Buffer after 24-h treatment. The cell lysates were examined using Dual-Luciferase Reporter (DLR) Assay System (Promega) and measured by a GloMax-Multi Jr Single Tube Multimode Reader (Promega) at a wavelength of 350–650 nm.

C3b binding assay was performed as described previously (34). Briefly, HRGECs were cultured in six-well plates and treated with Cd for 24-h. Then, HRGECs were incubated in 0.05% trypsin for 2 min at 37°C, resuspended as single-cell solution in PBS, and incubated with anti-Endoglin mAb for 30 min at 4°C. Following washing with PBS/1% BSA, HRGECs were incubated with 20% normal human serum in M199 for 3 h at 37°C. After washing with PBS/1% BSA, the anti-C3b mAb (catalog no. MA1-70053; Thermo Fisher Scientific) at 1:50 dilution was added to the suspension for 30 min at 4°C. These HRGECs were then stained with FITC-conjugated donkey anti-mouse IgG (ab150105; Abcam), and the presence of C3b was detected using a FACSAria II flow cytometer (BD Biosciences). Negative control was the HRGECs treated with heat-inactivated human serum.

Statistical significance was assessed using unpaired sample t tests for comparison between two groups and one-way ANOVA for comparison among multiple groups. p < 0.05 was considered significant. Statistical analyses were performed by using SPSS 17.0 software (SPSS, Chicago, IL).

To examine the effects of Cd on CFH production, mice were treated with Cd for 3 d. The peripheral blood and renal blood were collected at 24, 48, and 72 h, and the plasma CFH was examined by ELISA. As shown in Fig. 1A, Cd significantly increases the levels of CFH in renal venous blood at 24 h, which remain at a high level at 48 and 72 h, whereas the CFH levels in peripheral blood were not significantly changed at all time points. Thus, Cd treatment specifically increases CFH secretion from kidney, which is diluted in the peripheral blood. The tissues in different locations of the mouse kidney were precisely collected by LCM technology and examined for Cfh gene expression. We found that 24-h Cd treatment specifically increases Cfh mRNA levels in the cortex region (Fig. 1B). Within the cortex, Cfh gene expression was upregulated by Cd in the glomeruli (Fig. 1C) but not in Bowman capsule, afferent/efferent arterioles, proximal/distal tubules, or intertubular capillaries. Immunohistochemistry revealed that Cd-induced CFH expression is restricted to the GECs of mouse glomeruli (Fig. 1D). In addition, we examined the effects of Cd treatment on renal functions of the mice after day 3. Compared with the untreated mice, the morphology of glomerulus, serum, and urine creatinine levels do not show a significant change in Cd-treated mice (Fig. 1E–G), suggesting that Cd treatment might not induce renal injuries at this time point.

In vitro, the three types of glomerular cells, including HRGECs, human renal podocytes, and HRMCs, were cultured and treated with 4 μM Cd. In HRGECs, dose-responsive examination showed that CFH gene expression was significantly increased at Cd concentrations higher than 4 μM, whereas CFH secretion and mRNA expression (together with F3 expression) were significantly increased in a time-dependent manner (Fig. 2A, 2B, Supplemental Fig. 1A, 1B). CFH secretion and mRNA expression in human renal podocytes and HRMCs were not affected by Cd treatment at all time points (Fig. 2C–F). In addition, Western blotting indicated that extracellular CFH from the culture media and intracellular CFH from cell lysate of HRGECs were markedly increased by the Cd treatment (Fig. 2G, 2H). Immunoflouresence staining with a CFH Ab also demonstrated a significant increase of CFH protein in Cd-treated HRGECs (Fig. 2I). To validate the function of increased CFH, the effect of Cd on C3b deposition on HRGECs was examined in the presence of normal human serum. As shown in Supplemental Fig. 1C and 1D, C3b deposition is significantly decreased on Cd-treated HRGECs, suggesting the presence of Cd-induced CFH. Together, Cd specifically increases CFH expression and secretion in GECs. Dose-responsive examinations showed that Cd induce cytotoxicity and proliferation inhibition at 10 μM but not at 4 μM (Supplemental Fig. 1E, 1F). In addition, growth and viability of HRGECs were examined after exposure to 4 μM Cd. Fig. 3A and 3B showed that the relative proliferation by MTT and CCK8 assays was similar in HRGECs with or without Cd treatment. In the trypan blue exclusion assay and LDH release assay, Cd exposure did not reduce cell viability (Fig. 3C, 3D). Flow cytometry analyses exhibited that the percentage of annexin V/PI–positive cells and viable cells remained unchanged in HRGECs with or without Cd treatment (Fig. 3E, 3F). Thus, Cd exposure increases CFH expression without induction of cytotoxicity in HRGECs.

Cd is known to activate multiple signaling pathways (7, 35). HRGECs were pretreated with inhibitors of ERK, JNK, p38, NF-κB, and protein kinase B (Akt) pathways for 1 h before treatment with Cd for 24 h. We found that SP600125, an inhibitor of the JNK pathway, eliminated Cd-induced CFH expression and secretion, whereas other inhibitors did not (Fig. 4A, 4B). Although Cd increases CFH gene expression in HUVECs, pretreatment of JNK inhibitor SB203850 does not eliminate Cd-induced CFH transcription, suggesting that Cd induces CFH expression in HUVECs independent of the JNK pathway (Supplemental Fig. 1G). Thus, JNK mediation of Cd-inducible CFH gene expression is restricted to HRGECs. In addition, Western blotting indicated that Cd increases phosphorylation of JNK in HRGECs, whereas the levels of total JNK were unchanged (Fig. 4C). c-Jun and c-Fos, the downstream effectors of the JNK pathway, were also increased in HRGECs by Cd treatment (Fig. 4D). In a time course study, pretreatment with SP600125 abolished the Cd-induced increase of phosphorylation in JNK, cJun/c-Fos, and CFH expression in HRGECs at all time points (Supplemental Fig. 2A–D), suggesting that Cd increases CFH expression via activation of the JNK pathway.

Heterodimers of the transcription factors c-Jun and c-Fos form the AP-1 complex, which binds an AP-1 motif at the promoter and enhancer regions of target genes (36). We screened the region of -2200 to the first exon (+220) of the CFH promoter region by ChIP assay with nuclear extract from HRGECs (Supplemental Fig. 3, Supplemental Table 1). As shown in Fig. 5A and 5B, the DNA fragments including the −1635 AP1 motif were pulled down by the c-Jun or c-Fos Abs, and Cd treatment significantly increased binding activity of c-Jun and c-Fos on the fragment. On the EMSA gel, the probes with the −1635 AP1 motif were shifted by a nuclear extract of HRGECs and supershifted by nuclear extract together with c-Jun or c-Fos Abs (Fig. 5C). In addition, luciferase constructs of the CFH promoter with or without a mutation of the −1635 AP1 binding site were generated and transfected into HRGECs. After Cd treatment, CFH promoter activity was significantly upregulated as shown by the luciferase assay (Fig. 5D). However, the activity of the CFH promoter with the mutation of the −1635 AP1 binding site was not affected by Cd treatment (Fig. 5D). Collectively, these results indicate that the AP-1 complex binds with the −1635 AP-1 motif and controls the CFH promoter activity in HRGECs (Fig. 6).

Cd is an environmental toxicant that is known to target the glomerulus (37). In this study, we found that Cd increases glomerular CFH production by specifically upregulating CFH expression in GECs without induction of cell death. In HRGECs, Cd activates the JNK pathway and increases c-Jun and c-Fos. A JNK inhibitor, SP600125, specifically abolishes Cd-induced CFH expression. Using EMSA and ChIP, the −1635 AP-1 motif on the CFH promoter was identified as the binding element for c-Jun and c-Fos. Mutation of this motif eliminates the Cd-induced increase in CFH promoter activity that is observed in a luciferase activity assay. Taken together, Cd specifically induces CFH expression in GECs via activation of the JNK pathway and increase of c-Jun and c-Fos, which bind the −1635 AP-1 motif on the CFH promoter to activate Cd-inducible CFH gene expression (Fig. 6).

In animal models, Cd triggers excessive complement activation by induction of complement component C3, the key element of the complement system (38, 39). Activated C3 is cleaved to create C3a and C3b, the latter of which binds to the surface of pathogens or other target cells (11). CFH acts through its capacity to recognize polyanionic structures on host tissues and thereby inactivates C3b (11). As a negative regulator of C3 activation, CFH ameliorates C3 glomerulopathy in which abnormal complement activation results in predominant C3 fragment deposition within the glomerulus and glomerular damage (40, 41). The current study is the first, to our knowledge, to provide evidence for the effect of Cd on CFH expression and production in glomeruli, suggesting Cd as a crucial regulator of the innate immune system. Cd affects multiple organs, including liver and kidney, and the vascular system is also one of the primary targets of Cd-induced toxicity (42, 43). At lower concentrations, Cd does not induce cytotoxicity in endothelial cells (7, 9, 44) but increases expression of vascular hemostastic factors, such as tissue factor and von Willebrand factor (45). In response to Cd, GECs produce additional CFH, which might help protect the glomerular filtration barrier from dysregulation of the complement system.

The JNK pathway is activated by Cd in various cell types (7, 46). Activation of JNK leads to the increase of c-Jun and c-Fos, which form the AP-1 complex that modulates target gene expression (47). In this study, we have demonstrated that Cd causes both an increase in activated JNK in HRGECs as well as increases in levels of c-Fos and c-Jun. Our study also found that although Cd activates multiple pathways in HRGECs, only the JNK pathway inhibitor eliminated the Cd-induced increase of CFH gene expression and CFH secretion, suggesting that the JNK/AP-1 pathway plays a key role in regulating the transcription of the CFH gene. In murine astrocytes, Fraczek et al. (48) identified a 241-bp region at −416 bp to −175 bp on the mouse Cfh gene promoter that showed transcriptional activity with possible binding interactions with c-Jun and c-Fos. In the current study, we did not find AP-1 binding in the corresponding region on human CFH promoter in HRGECs but identified an AP-1 binding site at −1635 of the human CFH promoter, which mediates c-Jun– and c-Fos–regulated CFH gene expression. Our study suggests distinct regulatory mechanisms of human and murine CFH transcription, although both of them might be induced by activation of the JNK pathway. As polymorphic variations in the CFH promoter have been shown to change CFH gene expression (49), it is possible that polymorphisms existing in the −1635 AP1 motif influence gene expression in humans.

Our findings also provide mechanistic insights into cell type–specific regulatory mechanisms of CFH expression. In glomeruli, Cd activates the JNK pathway in podocytes and mesangial cells (46) without induction of CFH expression. Oxidative stress, which also activates the JNK pathway, downregulates CFH expression in retinal pigment epithelial cells (50). Thus, JNK/AP1 mediation of Cd-inducible CFH gene expression is restricted to HRGECs. Endothelial cells are highly responsive to changes of the microenvironment, and their phenotypes are differentially regulated in time and space (51). In addition, endothelial cells of specific vascular beds display significant heterogeneity in structure and function (26). The GECs are a unique cell type with a large fenestrated area constituting between 20 and 50% of the cellular surface, which is indispensable for their function in the glomerular filtration barrier (52, 53). In response to extracellular stimuli, GECs exhibit distinct gene expression profiles from other types of endothelial cells (54). Further studies are needed to determine additional molecular mechanisms that might specifically facilitate JNK/AP1–regulated CFH expression in HRGECs.

In summary, we show that Cd induces CFH expression specifically in GECs in vivo and in vitro, possibly a compensatory mechanism to Cd-induced complement activation. In HRGECs, Cd activates the JNK pathway and increases c-Jun and c-Fos, which upregulate the CFH promoter activity by interaction with the −1635 AP-1 motif. These results provide novel mechanistic insights for understanding the role of Cd in glomerular dysfunction.

The authors have no financial conflicts of interest.