Abstract

Transcriptional co-activator with PDZ-binding motif (TAZ) is a key downstream effector of the Hippo tumor-suppressor pathway. The functions of TAZ in the kidney, especially in tubular epithelial cells, are not well-known. To elucidate the adaptive expression, protective effects on kidney injury, and signaling pathways of TAZ in response to acute kidney injury (AKI), we used in vitro (hypoxia-treated human renal proximal tubular epithelial cells [RPTECs]) and in vivo (mouse ischemia–reperfusion injury [IRI]) models of ischemic AKI. After ischemic AKI, TAZ was up-regulated in RPTECs and the renal cortex or tubules. Up-regulation of TAZ in RPTECs subjected to hypoxia was controlled by IκB kinase (IKK)/nuclear factor κ-light-chain-enhancer of activated B cell (NF-κB) signaling. TAZ overexpression attenuated hypoxic and oxidative injury, inhibited apoptosis and activation of p38 and c-Jun N-terminal kinase (JNK) proteins, and promoted wound healing in an RPTEC monolayer. However, TAZ knockdown aggravated hypoxic injury, apoptosis, and activation of p38 and JNK signaling, delayed wound closure of an RPTEC monolayer, and promoted G0/G1 phase cell-cycle arrest. Chloroquine and verteporfin treatment produced similar results to TAZ overexpression and knockdown in RPTECs, respectively. Compared with vehicle-treated mice, chloroquine treatment increased TAZ in the renal cortex and tubules, improved renal function, and attenuated tubular injury and tubular apoptosis after renal IRI, whereas TAZ siRNA and verteporfin decreased TAZ in the renal cortex and tubules, deteriorated renal failure and tubular injury, and aggravated tubular apoptosis. Our findings indicate the renoprotective role of tubular TAZ in ischemic AKI. Drugs augmenting (e.g., chloroquine) or suppressing (e.g., verteporfin) TAZ in the kidney might be beneficial or deleterious to patients with AKI.

Introduction

Acute kidney injury (AKI) is one of the most serious conditions worldwide and is associated with mortality [1,2]. The proximal tubule has been considered the primary target of AKI, and tubular injury plays a critical role in subsequent chronic kidney disease (CKD) due to glomerulosclerosis, vascular rarefaction, and fibrosis [3,4]. After the development of CKD, renal function declines gradually and end-stage renal disease (ESRD) is usually inevitable. Moreover, AKI is linked to subsequent serious morbidities: coronary events, severe sepsis, and gastrointestinal hemorrhage [5–7]. Thus, identifying novel preventive or therapeutic targets of AKI is needed to reduce mortality, subsequent CKD and ESRD, and other complications.

Similar to the Wnt signaling pathway, the Hippo signaling pathway delivers signals from the plasma membrane to the nucleus, alters gene expression, and controls cell survival [8]. Transcriptional co-activator with PDZ-binding motif (TAZ) and Yes-associated protein (YAP) are key effectors in the Hippo signaling pathway [9]. TAZ and YAP are closely related and share approximately 50% of sequence identity, a similar topology, and downstream transcription factors [10]. YAP and TAZ can regulate the process of skin wound healing [11]. TAZ was initially recognized as a 14-3-3 binding protein [10]. Previous studies demonstrated that TAZ plays an important role in the growth of organs, for tissue renewal, regeneration, and for cell proliferation [9].

Recently, YAP has been reported to be involved in the pathogenesis of focal segmental glomerulosclerosis and regeneration after ischemic AKI [12–14]. Additionally, YAP and TAZ promote renal tubulointerstitial fibrosis [15,16]. However, little is known about the independent biological role of TAZ in AKI. Therefore, we investigated the expression levels of TAZ during ischemic AKI models and the potential renoprotective role of TAZ.

Materials and methods

In vitro hypoxia models

Human kidney 2 cells, a type of human renal proximal tubular epithelial cells (RPTECs) (Bioresource Collection and Research Center [BCRC], Hsinchu, Taiwan) were cultured in keratinocyte-serum free medium (KSFM, GIBCO-BRL) supplemented with 5 ng/ml of recombinant epidermal growth factor and 40 µg/ml of bovine pituitary extract, and incubated (37°C, 5% CO2). Hypoxia chamber and cobalt chloride (CoCl2; catalog no. C8661, Sigma–Aldrich) were used to create in vitro models of hypoxia. In the hypoxia chamber experiments, human RPTECs were incubated (1% O2, 5% CO2, 37°C) for 24 h. In CoCl2 experiments, RPTECs were treated with CoCl2 (300 or 600 μM) for 6 or 24 h. We examined the expression of hypoxia-inducible factor (HIF)-1α to ensure the success of both the models. For the oxidative stress experiments, RPTECs were treated with H2O2 (100 or 250 μM) for 6 h.

Stable TAZ-overexpressing and knockdown RPTECs

Briefly, recombinant lentiviruses were produced by transiently co-transfecting human embryonic kidney (HEK293T) cells with the indicated plasmids, packaging plasmid Δ8.2, and envelope plasmid VSV-G, using the calcium phosphate precipitation method. Full-length human TAZ (cDNA clone: MGC 19891; purchased from BCRC) was cloned into a cDNA expression lentiviral vector (pLAS2w.Ppuro, National RNAi Core Facility of Academia Sinica, Taipei, Taiwan) to create a TAZ-overexpression stable RPTEC cell line. Moreover, pLKO.1-puro-Luc and pLKO.1-puro-shTAZ (clone ID: TRCN0000019469; National RNAi Core Facility of Academia Sinica) were used to create a TAZ-knockdown stable RPTEC cell line. Plasmid DNA was transfected into HEK293T cells (10 μg of vector per 10-cm cell culture dish) for 48 h. Culture supernatants containing recombinant lentiviruses were harvested and filtered through a 0.45-μm pore size filter. Next, human RPTECs were incubated with recombinant viruses for 24 h. Infected RPTECs were selected using a 48-h puromycin (1 μg/ml) treatment. Finally, the success of the virus infection was examined using Western blotting for TAZ.

IκB kinase siRNA, IκB kinase inhibitor, verteporfin, and chloroquine treatment in vitro

Human RPTECs (2 × 105 cells/ml) were seeded in six-well culture plates for 12 h. RPTECs were treated with CoCl2 (300 μM) and IκB kinase (IKK) siRNA (250 nM, catalog no. M-003503-03-0010, Dharmacon; sequences were 5′-AUGAAUGCCUCUCGACUUA-3′, 5′-GAAGAGGUGGUGAGCUUAA-3′, 5′-GAGCUGUACAGGAGACUAA-3′, and 5′-CCGAUAAGCCUGCCACUCA-3′) and/or IKK inhibitor (5 μM, catalog no. 401486, Merck Millipore) for 6 h. Control siRNA (catalog no. D-001810-10-05, Dharmacon; sequences were 5′-UGGUUUACAUGUCGACUAA-3′, 5′-UGGUUUACAUGUUGUGUGA-3′, 5′-UGGUUUACAUGUUUUCUGA-3′, and 5′-UGGUUUACAUGUUUUCCUA-3′) was used as a comparison group. Additionally, RPTECs were treated with verteporfin (10–250 nM, catalog no. SML0534, Sigma–Aldrich) and CoCl2 for 6 and 24 h, respectively. Moreover, RPTECs were treated with chloroquine (2.5 μM, C6628, Sigma–Aldrich) for 24 h before CoCl2-induced hypoxia. Cell viability and apoptosis were examined using the methylthiazol tetrazolium (MTT) assay and flow cytometry, respectively.

Primary renal tubular epithelial cells

Primary renal tubular epithelial cells (PRTECs) were obtained from C57BL/6 mice (National Laboratory Animal Center, Taipei, Taiwan), as previously described [17]. Briefly, mice were killed and both kidneys were removed under general anesthesia. The kidney cortices were minced and digested with 1 mg/ml collagenase type 1 at 37°C for 30 min. The undigested kidney tissues were removed using a 100-μm and a 70-μm sieve. Then, the filtered cell suspension was centrifuged at 50×g for 5 min to collect the first pellet. The supernatant was centrifuged at 50×g for 5 min to collect the second pellet. Then, the first pellet was resuspended in 20 ml of culture medium and was centrifuged at 50×g for 5 min to collect the third pellet. Next, we resuspended the second and third pellets in 1 ml of culture media for subculture. PRTECs were cultured in DMEM/F12 (Gibco) supplemented with recombinant EGF (50 ng/ml; Sigma–Aldrich), fetal bovine serum (2%; Sigma–Aldrich), transferrin (5 µg/ml; Sigma–Aldrich), insulin (5 µg/ml; Sigma–Aldrich), and hydrocortisone (36 ng/ml; Sigma–Aldrich). To suppress the expression of TAZ, PRTECs were treated with two different doses (20 or 200 nM) of TAZ siRNA (5′-AUACUUCCUUAAUCACAUtt-3′, catalog no. 4457308, Thermo Fisher Scientific) for 48 h.

Western blotting

Human RPTECs and homogenized mouse renal cortex tissues were lysed in RIPA buffer at 4°C for 30 min. The protein concentration of the whole lysate was quantified using the bicinchoninic acid assay kit (Thermo Scientific). We used a nuclear protein extraction kit (catalog no. 2900, Merck Millipore), according to the manufacturer’s instructions. Immunoblotting was performed using primary antibodies against TAZ (catalog no. 4883, Cell Signaling), HIF-1α (catalog no. 610958, BD Biosciences), cleaved caspase-8 (catalog no. 9496, Cell Signaling), cleaved caspase-9 (catalog no. 20750, Cell Signaling), nuclear factor κ-light-chain-enhancer of activated B cells (NF-κB, p65) (catalog no. 8242, Cell Signaling), phospho-NF-κB (p-p65) (catalog no. 3033, Cell Signaling), p38 (catalog no. 9212, Cell Signaling), phospho-p38 (catalog no. 9211, Cell Signaling), c-Jun N-terminal kinase (JNK; catalog no. 9258, Cell Signaling), phospho-JNK (catalog No. 07-175, Millipore), B-cell lymphoma 2 (Bcl-2; catalog no. 2872, Cell Signaling), Bcl-2-associated X protein (Bax; catalog no. 5023, Cell Signaling), Lamin A/C (catalog no. Ab108922, Abcam), β-actin (catalog no. NB600-501, Novus), and GAPDH (catalog no. Ab181602, Abcam). HRP-conjugated anti-rabbit (catalog no. GTX213110-01, GeneTex) and anti-mouse (catalog no. GTX213111-01, GeneTex) secondary antibodies were used. β-actin or GAPDH was used as an internal control and reference in the Western blotting.

Immunofluorescence staining

Human RPTECs were treated with CoCl2 (300 μM) for 6 h. Following fixation (4% formaldehyde) and permeabilization (0.1% Triton X-100), cells were incubated with anti-TAZ antibody (catalog no. 4883; Cell Signaling) and fluorescent-dye conjugated secondary antibodies (Alexa Fluor® 594, catalog no. 111-585-003 or Fluorescein, catalog no. 111-095-003, Jakson ImmunoResearch). Nuclear staining was performed using DAPI (catalog no. 17510, AAT Bioquest). Images were captured using an Olympus Confocal Laser Scanning Microscope FV3000.

Immunohistochemistry staining

Formalin-fixed, paraffin-embedded renal tissue sections (4 μm) were processed through deparaffinization, rehydration, and antigen retrieval (citrate buffer, 20 min), then incubated with anti-TAZ antibodies (1:200; catalog no. 4883, Cell Signaling) (8 h, 4°C). After incubation with goat anti-rabbit IgG, 3,3′-diaminobenzidine tetrahydrochloride (DAB) substrate and Hematoxylin were used for color development and counterstaining. Computer-assisted quantitative analysis was performed to determine the expression level of TAZ, as previously described [18,19]. Briefly, ten randomly selected non-overlapping high-power field images (×200) for renal cortical sections were captured using an Olympus BX61 microscope and an Olympus DP81 digital color camera (Japan). The images were analyzed using Image Pro-Plus software (v6.0, Media Cybernetics). The quantitative immunohistochemical staining value (QISV) was calculated as the integrated optical density divided by the total area occupied by the DAB- and Hematoxylin-stained cells of each slide.

Histopathology

Renal tissue sections were stained using Hematoxylin and Eosin. The severity of the tubulointerstitial damage was examined under ten random non-overlapping high-power fields (×200). Tubular injury was graded on a 0–4 scale (0, 1, 2, 3, and 4 for no lesions, lesions involving <25%, 25–50%, 50–75%, and >75% of the cortical parenchyma, respectively) [18,19].

Tumor necrosis factor-α-induced apoptosis

Human RPTECs (6.73 × 103 or 2 × 105 cells/well) were seeded in 96-well or 6-well culture plates for 12 h and treated with recombinant human tumor necrosis factor-α (TNF-α; 10–50 ng/ml; catalog no. 300-01A, PeproTech) for 72 or 96 h. Cell viability and the activation of cleaved caspase-8 were measured using the MTT assay and Western blotting, respectively.

MTT assay

Human RPTECs (6.73 × 103 cells/well) were seeded in 96-well cell culture plates for 12 h. RPTECs were incubated under hypoxia conditions (1% O2 for 24 h or CoCl2 for 6 and 24 h). These cells were incubated with MTT (5 mg/ml) for 1 h and the formazan crystals were dissolved by dimethyl sulfoxide (DMSO). The absorbance (570 nm) in MTT-treated cells is an index of cell viability.

Analysis of apoptosis

Transfected human RPTECs (2 × 105 cells/well) were seeded in six-well cell culture plates for 12 h. Following CoCl2 treatment for 6 and 24 h, RPTECs were stained using propidium iodide (PI) and FITC-conjugated annexin-V (BD Biosciences) for 30 min. The percentages of cell death (PI+ Annexin-V+ cells/total cells) were measured using flow cytometry (FC-500, Beckman). In other experiments, CoCl2-treated RPTECs were incubated with FITC-conjugated anti-active caspase-3 antibodies (BD Biosciences) for 30 min. Caspase-3-positive cells (caspase-3+ cells/total cells) were analyzed using flow cytometry.

Scratch assay

Artificial wounds were created using Culture-inserts (Ibidi). Briefly, Culture-inserts were placed on RPTEC monolayers until confluent monolayers of RPTECs were established on dishes. After removing the Culture-insert, a 500-μm gap was created. Two pictures per wound were taken using a phase contrast microscope at different time points (0, 8, and 24 h). The gap was assessed using ImageJ.

Animal models and in vivo renal ischemia–reperfusion injury

C57BL/6 mice (National Laboratory Animal Center, Taipei, Taiwan) were housed in groups of four in filter-top cages with unlimited access to food and water. Cages, food, and water were autoclaved before use. During the experimental procedure, mice were kept well hydrated with phosphate-buffered saline (PBS), at 38°C. Body temperature was maintained using a 45°C heating pad and a heating lamp during surgery and recovery, respectively. All procedures on animals were performed at the Changhua Christian Hospital and experiments were conducted according to the Taiwan animal protection laws which were approved by the Animal Center of Changhua Christian Hospital. Mice were anesthetized with 5% isoflurane and 2.5% isoflurane for induction throughout surgery, respectively. For induction of renal ischemia–reperfusion injury (IRI), bilateral renal arteries and veins were clamped for 20 (chloroquine groups) or 10 min (TAZ siRNA and verteporfin groups) with small vascular clips. Renal ischemia time was reduced (10 min) in order to minimize mortality after bilateral renal IRI in TAZ siRNA and verteporfin groups. After 24 h, the mice were killed and the ischemic kidneys were snap-frozen in liquid nitrogen and fixed in 10% buffered formalin.

To evaluate the effects of TAZ knockdown on AKI, TAZ siRNA (5′-AUACUUCCUUAAUCACAUtt-3′, catalog no. 4457308, Thermo Fisher Scientific) was injected into bilateral renal pelvis 30 min before IRI (50 μmol administered to each kidney). TAZ siRNA was mixed with a 100 μl transfection reagent (TransIT-QR Delivery Solution, catalog no. MIR 5240, Mirus Bio) before injection. The distribution of siRNAs administered intrapelvically was examined using fluorescence molecular tomography (FMT; Supplementary Figure S1) and confocal microscopy (Supplementary Figure S2). To investigate the effects of chloroquine on renal IRI, mice were divided into groups 1, 2, and 3, which were treated with PBS, chloroquine at 10 and 50 mg/kg intraperitoneally per day, 5 days before induction of IRI, respectively. To reveal the effects of verteporfin on renal IRI, mice were divided into groups 1 and 2, which were treated with DMSO and verteporfin at 100 mg/kg intraperitoneally every other day, 5 days before induction of IRI, respectively.

FMT

The distribution of siRNA injected intrapelvically was assessed using the FMT 2500 In Vivo Imaging System (PerkinElmer Inc.). First, the mice treated with Alexa Fluor® 633-conjugated siRNA were placed in the supine position after anesthesia, centrally in the imaging cassette, to capture the whole body, except the head. After mice were imaged in vivo (6 h after intrapelvic siRNA injection), they were killed and the organs (kidneys, ureters, and urinary bladder) were taken out for ex vivo FMT imaging. Both in vivo and ex vivo FMT imaging were performed at 635 nm.

Terminal-deoxynucleotidyl transferase-mediated digoxigenin-deoxyuridine nick-end labeling assay

Terminal-deoxynucleotidyl transferase-mediated digoxigenin-deoxyuridine nick-end labeling (TUNEL) assay was performed using In Situ Cell Death Detection Kit (Roche Diagnostics) according to the instructions of the manufacturer. Briefly, renal tissue sections (4 μm) were treated with proteinase K (10 μg/ml, 37°C, Amresco Inc.) for 10 min. The sections were then blocked with H2O2 (3%, 10 min) and normal goat serum (Jackson ImmunoResearch), and then were reacted with the TUNEL reaction mixture (37°C, 40 min) and converter-POD solution (37°C, 20 min). For the color development and counterstaining, DAB and Hematoxylin were used, respectively. Apoptosis in the renal cortical tissue was determined as the percentage of TUNEL-positive cells.

Statistical analysis

Data were expressed as mean ± SEM (shown in Figures 19). Comparisons between two groups and more were performed using the unpaired t test and one-way ANOVA with Tukey’s post hoc, respectively. All statistical analyses were performed using SPSS 24 (IBM SPSS Inc.) or GraphPad 7.0.4 (GraphPad Software). P-values <0.05 were considered statistically significant.

Hypoxic injury up-regulates TAZ in vivo and in cultured RPTECs

Figure 1
Hypoxic injury up-regulates TAZ in vivo and in cultured RPTECs

(A) Representative Western blots (left upper panel) with densitometric quantitative results (right upper panel) of TAZ and β-actin in renal cortical tissues of sham-operated and renal IRI mice. Representative immunohistochemical staining images (left lower panel) and QISVs (right lower panel) show the differential expression of TAZ in renal cortical tissues of sham-operated and IRI mice. TAZ was expressed mainly in the renal tubules (left lower panel). n=4 per group, t test. *P<0.05. Scale bar, 50 μm. (B) Immunofluorescence revealed that there was no nuclear localization of TAZ in proximal (AQP1-positive) renal tubules of sham-operated mice. However, some nuclei of proximal renal tubules were filled with TAZ staining in IRI mice. (C) Representative Western blots (top panel) with densitometric quantitative results (bottom panel) of HIF-1α, TAZ, and GAPDH in control and CoCl2-induced hypoxia groups. Each experiment was repeated in triplicate. After a 6-h CoCl2 treatment, HIF-1α and TAZ were significantly increased in human RPTECs. (D) Representative Western blots and densitometric quantitative results of TAZ and β-actin in normoxia and low oxygen (hypoxia) groups (n=3 per group) (E) and in the control and H2O2-treated groups (n=3 per group). RPTECs were treated with H2O2 for 6 h. (F) Cytosolic distribution of TAZ decreased after hypoxia (6-h CoCl2 treatment, assessed using confocal microscopy). Scale bar, 50 μm. (G) Western blots of TAZ after extracting cytosol and nuclear proteins further confirms that TAZ was up-regulated and translocated into nucleus after hypoxia. *P<0.05; **P<0.01; ***P<0.001. Abbreviations: AQP1, aquaporin 1; DAPI, 4′,6 -diamidino-2-phenylindole; kDa, kilodalton.

Figure 1
Hypoxic injury up-regulates TAZ in vivo and in cultured RPTECs

(A) Representative Western blots (left upper panel) with densitometric quantitative results (right upper panel) of TAZ and β-actin in renal cortical tissues of sham-operated and renal IRI mice. Representative immunohistochemical staining images (left lower panel) and QISVs (right lower panel) show the differential expression of TAZ in renal cortical tissues of sham-operated and IRI mice. TAZ was expressed mainly in the renal tubules (left lower panel). n=4 per group, t test. *P<0.05. Scale bar, 50 μm. (B) Immunofluorescence revealed that there was no nuclear localization of TAZ in proximal (AQP1-positive) renal tubules of sham-operated mice. However, some nuclei of proximal renal tubules were filled with TAZ staining in IRI mice. (C) Representative Western blots (top panel) with densitometric quantitative results (bottom panel) of HIF-1α, TAZ, and GAPDH in control and CoCl2-induced hypoxia groups. Each experiment was repeated in triplicate. After a 6-h CoCl2 treatment, HIF-1α and TAZ were significantly increased in human RPTECs. (D) Representative Western blots and densitometric quantitative results of TAZ and β-actin in normoxia and low oxygen (hypoxia) groups (n=3 per group) (E) and in the control and H2O2-treated groups (n=3 per group). RPTECs were treated with H2O2 for 6 h. (F) Cytosolic distribution of TAZ decreased after hypoxia (6-h CoCl2 treatment, assessed using confocal microscopy). Scale bar, 50 μm. (G) Western blots of TAZ after extracting cytosol and nuclear proteins further confirms that TAZ was up-regulated and translocated into nucleus after hypoxia. *P<0.05; **P<0.01; ***P<0.001. Abbreviations: AQP1, aquaporin 1; DAPI, 4′,6 -diamidino-2-phenylindole; kDa, kilodalton.

Results

Hypoxic injury up-regulates TAZ protein expression and nuclear localization

A recent research examined temporal-specific gene expression patterns after murine renal IRI using RNA-sequencing analysis. According to their uploaded RNA-sequencing database (NCBI GEO accession: GSE98622), an increasing trend of TAZ mRNA levels in the kidneys was observed from 4 h to 3 days after renal IRI [20]. To confirm this, we investigated the in vivo effects of TAZ on a murine model of renal IRI and found that TAZ expression was significantly increased in injured renal cortical tissues 24 h after renal IRI. Moreover, immunohistochemical staining revealed that the differential expression of TAZ was mainly in renal tubular epithelial cells (Figure 1A). Immunofluorescence showed that TAZ was absent from the nucleus of aquaporin-1-positive RPTECs in sham-operated mice. In contrast, TAZ was increased in both the cytoplasm and nucleus of RPTECs after IRI (Figure 1B). To establish a hypoxic culture condition, human RPTECs were treated with CoCl2 or low (1%) oxygen, which triggered TAZ expression in human RPTECs (Figure 1C,D). Additionally, oxidative stress increased tubular TAZ expression (Figure 1E). Immunofluorescence staining revealed that intracellular and cytosolic TAZ increased and decreased during hypoxia, respectively (Supplementary Figure S3; Figure 1F). We further extracted nuclear and cytoplasmic proteins from RPTECs and confirmed that TAZ was largely localized in the nucleus after hypoxia (Figure 1G).

IKK/NF-κB signaling regulates TAZ protein expression during hypoxia in RPTECs

CoCl2 increased the phosphorylation of NF-κB (p65) in human RPTECs (Figure 2A,B). Next, IKK siRNA and inhibitor were used to evaluate the regulatory role of the IKK/NF-κB axis in TAZ expression in human RPTECs with hypoxia. IKK siRNA and inhibitor significantly suppressed the phosphorylation of NF-κB (p65) and reduced TAZ expression in human RPTECs (Figure 2A–C and Supplementary Figure S4).

TAZ expression was regulated by IKK/NF-κB signaling in CoCl2-induced hypoxia in cultured RPTECs

Figure 2
TAZ expression was regulated by IKK/NF-κB signaling in CoCl2-induced hypoxia in cultured RPTECs

Representative Western blots (A) with densitometric quantitative results (B,C) of HIF-1α, NF-κB, TAZ, and β-actin in RPTECs in response to hypoxia and IKK inhibition. (B) Hypoxia activated the phosphorylation of NF-κB (p65) in human RPTECs. IKK siRNA suppressed the hypoxia-induced NF-κB activation (n=3 per group, ANOVA). (C) Hypoxia up-regulated TAZ expression in human RPTECs. Additionally, IKK siRNA suppressed TAZ expression after hypoxia (n=3 per group, ANOVA). RPTECs were treated with control or IKK siRNAs (1 μM, 48 h and CoCl2 (300 μM), 6 h). **P<0.01; ***P<0.001.

Figure 2
TAZ expression was regulated by IKK/NF-κB signaling in CoCl2-induced hypoxia in cultured RPTECs

Representative Western blots (A) with densitometric quantitative results (B,C) of HIF-1α, NF-κB, TAZ, and β-actin in RPTECs in response to hypoxia and IKK inhibition. (B) Hypoxia activated the phosphorylation of NF-κB (p65) in human RPTECs. IKK siRNA suppressed the hypoxia-induced NF-κB activation (n=3 per group, ANOVA). (C) Hypoxia up-regulated TAZ expression in human RPTECs. Additionally, IKK siRNA suppressed TAZ expression after hypoxia (n=3 per group, ANOVA). RPTECs were treated with control or IKK siRNAs (1 μM, 48 h and CoCl2 (300 μM), 6 h). **P<0.01; ***P<0.001.

TAZ overexpression ameliorates hypoxia-induced damage, apoptosis, activation of p38 and JNK, and accelerates epithelial wound healing in RPTECs

To examine the independent renoprotective effect of TAZ on hypoxia-induced damage, we established a stable high TAZ expression RPTEC cell line (Figure 3A), in which TAZ was substantially increased both in the nucleus and cytosol after CoCl2 treatment (Figure 3B). This overexpression significantly enhanced cell survival in human RPTECs incubated in hypoxic conditions (CoCl2, 1% oxygen) and under oxidative stress (Figure 3C,D; Supplementary Figure S5). Additionally, it reduced apoptotic cells and decreased the activation of caspase-3 and caspase-8 in CoCl2-treated RPTECs (Figure 3E,F). We investigated whether the protective role of TAZ during hypoxic conditions was associated with inhibition of the caspase-8-dependent pathway. The levels of TNF-α-induced cytotoxicity and activation of caspase-8 were lower in TAZ-overexpressing RPTECs compared with the transfection control (transfected with an empty vector) (Figure 4A). Furthermore, we tested the effects of TAZ overexpression on epithelial monolayer injury by mechanical scratching of the RPTEC monolayers and TAZ overexpression significantly accelerated epithelial wound closure (Figure 4B). Moreover, hypoxia substantially activated mitogen-activated protein kinase (MAPK) intracellular signals, including phosphorylation of JNK and p38; however, TAZ overexpression reduced the activation of JNK and p38-MAPK signaling (Figure 4C).

TAZ overexpression suppressed hypoxic injury and apoptosis in cultured RPTECs

Figure 3
TAZ overexpression suppressed hypoxic injury and apoptosis in cultured RPTECs

(A) TAZ was highly expressed in the TAZ-overexpressing human RPTECs compared with empty vector-transfected and wild-type human RPTECs (WT). (B) Hypoxia further increased TAZ in the cytoplasm and nucleus in TAZ-overexpressing human RPTECs compared with empty vector-transfected human RPTECs. (C) Cell viability (via MTT assay) was higher in TAZ-overexpressing compared with empty vector-transfected human RPTECs after 6 or 24-h CoCl2 treatments (n=5 per group, t test). (D) Cell viability (MTT assay) was higher in TAZ-overexpressing human RPTECs compared with empty vector-transfected human RPTECs in low oxygen (1% O2) for 24 h (n=4 per group, t test). (E) TAZ overexpression inhibited CoCl2-induced apoptosis (600 μM, 6 h). Apoptotic or caspase-3-positive cells were lower in TAZ-overexpressing human RPTECs compared with empty vector-transfected human RPTECs (n=3 per group, t test). (F) Representative Western blots (left panel) and densitometric results (right panel) of cleaved caspase-8 and caspase-9 in TAZ-overexpressing and empty vector-transfected human RPTECs after CoCl2-induced hypoxia (n=4 per group, t test). *P<0.05; **P<0.01; ***P<0.001.

Figure 3
TAZ overexpression suppressed hypoxic injury and apoptosis in cultured RPTECs

(A) TAZ was highly expressed in the TAZ-overexpressing human RPTECs compared with empty vector-transfected and wild-type human RPTECs (WT). (B) Hypoxia further increased TAZ in the cytoplasm and nucleus in TAZ-overexpressing human RPTECs compared with empty vector-transfected human RPTECs. (C) Cell viability (via MTT assay) was higher in TAZ-overexpressing compared with empty vector-transfected human RPTECs after 6 or 24-h CoCl2 treatments (n=5 per group, t test). (D) Cell viability (MTT assay) was higher in TAZ-overexpressing human RPTECs compared with empty vector-transfected human RPTECs in low oxygen (1% O2) for 24 h (n=4 per group, t test). (E) TAZ overexpression inhibited CoCl2-induced apoptosis (600 μM, 6 h). Apoptotic or caspase-3-positive cells were lower in TAZ-overexpressing human RPTECs compared with empty vector-transfected human RPTECs (n=3 per group, t test). (F) Representative Western blots (left panel) and densitometric results (right panel) of cleaved caspase-8 and caspase-9 in TAZ-overexpressing and empty vector-transfected human RPTECs after CoCl2-induced hypoxia (n=4 per group, t test). *P<0.05; **P<0.01; ***P<0.001.

TAZ overexpression suppressed TNF-α-induced apoptosis, promoted wound healing of RPTECs, and inhibited activation of p38 and JNK

Figure 4
TAZ overexpression suppressed TNF-α-induced apoptosis, promoted wound healing of RPTECs, and inhibited activation of p38 and JNK

(A) Cell survival was higher in TAZ overexpressing human RPTECs compared with empty vector-transfected human RPTECs after treatment with various concentrations of TNF-α (10–50 ng/ml) for 72 h (n=5 per group, t test). In contrast, less activated caspase-8 was observed in TAZ-overexpressing human RPTECs compared with transfection with an empty vector. (B) A pro-regenerative effect was observed using an in vitro scratch assay on monolayers of TAZ-overexpressing human RPTECs (n=3 per group, t test). Scale bar, 200 μm. (C) Representative Western blots (left panel) and densitometric quantitative results (right panel) of phospho-p38 (p-p38), p38, phospho-JNK (p-JNK), JNK, and β-actin proteins in human RPTECs with or without TAZ overexpression after CoCl2-induced hypoxia (n=3–4 per group, t test). *P<0.05; **P<0.01; ***P<0.001.

Figure 4
TAZ overexpression suppressed TNF-α-induced apoptosis, promoted wound healing of RPTECs, and inhibited activation of p38 and JNK

(A) Cell survival was higher in TAZ overexpressing human RPTECs compared with empty vector-transfected human RPTECs after treatment with various concentrations of TNF-α (10–50 ng/ml) for 72 h (n=5 per group, t test). In contrast, less activated caspase-8 was observed in TAZ-overexpressing human RPTECs compared with transfection with an empty vector. (B) A pro-regenerative effect was observed using an in vitro scratch assay on monolayers of TAZ-overexpressing human RPTECs (n=3 per group, t test). Scale bar, 200 μm. (C) Representative Western blots (left panel) and densitometric quantitative results (right panel) of phospho-p38 (p-p38), p38, phospho-JNK (p-JNK), JNK, and β-actin proteins in human RPTECs with or without TAZ overexpression after CoCl2-induced hypoxia (n=3–4 per group, t test). *P<0.05; **P<0.01; ***P<0.001.

Chloroquine triggers tubular TAZ, protects RPTECs from hypoxia injury, and accelerates epithelial wound healing

Chloroquine has been shown to activate NF-κB in several cell lines [21,22]. Recently, we found that the Akt-NF-κB pathway up-regulates YAP, a TAZ structurally related protein, in 3T3-L1 cells [23]. Therefore, we examined whether chloroquine could regulate TAZ in human RPTECs, and indeed it can substantially increase TAZ expression though YAP expression in human RPTECs (Figure 5A and Supplementary Figure S6). Additionally, chloroquine significantly improved cell survival and reduced apoptosis after hypoxia injury (Figure 5B,C). Moreover, chloroquine accelerated RPTEC monolayer wound healing, compared with the vehicle group (Figure 5D).

Chloroquine triggered TAZ expression, protected RPTECs from hypoxic injury, and promoted epithelial wound healing

Figure 5
Chloroquine triggered TAZ expression, protected RPTECs from hypoxic injury, and promoted epithelial wound healing

(A) Human RPTECs were treated with chloroquine (2.5 μM) for 24 h, and representative Western blots (left panel) and densitometric quantitative results (n=3 per group, t test) for TAZ are shown. (B) Cell survival (MTT assay) after CoCl2-induced hypoxia in human RPTECs treated with or without chloroquine (n=3 per group, ANOVA). (C) CoCl2 (600 μM) significantly increased apoptotic RPTECs but the apoptosis was inhibited by chloroquine treatment (n=3 per group, t test). (D) A pro-regenerative effect was observed on wound healing in human RPTECs after 8- or 24-h chloroquine treatments (n=3 per group, t test). Scale bar, 200 μm. *P<0.05; **P<0.01; ***P<0.001.

Figure 5
Chloroquine triggered TAZ expression, protected RPTECs from hypoxic injury, and promoted epithelial wound healing

(A) Human RPTECs were treated with chloroquine (2.5 μM) for 24 h, and representative Western blots (left panel) and densitometric quantitative results (n=3 per group, t test) for TAZ are shown. (B) Cell survival (MTT assay) after CoCl2-induced hypoxia in human RPTECs treated with or without chloroquine (n=3 per group, ANOVA). (C) CoCl2 (600 μM) significantly increased apoptotic RPTECs but the apoptosis was inhibited by chloroquine treatment (n=3 per group, t test). (D) A pro-regenerative effect was observed on wound healing in human RPTECs after 8- or 24-h chloroquine treatments (n=3 per group, t test). Scale bar, 200 μm. *P<0.05; **P<0.01; ***P<0.001.

TAZ knockdown increased susceptibility to hypoxia-induced injury, apoptosis, and cell arrest in RPTECs

To confirm the biological role of TAZ, we established a stable TAZ-knockdown RPTEC cell line (Figure 6A). TAZ knockdown decreased cell survival and increased apoptotic cells in RPTECs incubated with 600 μM CoCl2 (Figure 6B,C). Moreover, TAZ knockdown dampened and delayed RPTEC wound healing compared with the empty vector group (Figure 6D,E). Regarding the activation of p38 and JNK, TAZ knockdown increased the phosphorylation of p38 and JNK in CoCl2-treated RPTECs (Figure 6F; Supplementary Figure S7). Furthermore, TAZ knockdown in RPTECs enhanced cell cycle arrest at the G0/G1 phase (Supplementary Figure S8). PRTECs were obtained from C57BL/6 mice to confirm the results from human RPTECs. The effects of TAZ knockdown in human RPTECs were similar to those in primary renal tubular cells (Supplementary Figure S9).

TAZ knockdown exaggerated hypoxic damage and apoptosis, impeded epithelial wound healing, and activated p38 and JNK in RPTECs

Figure 6
TAZ knockdown exaggerated hypoxic damage and apoptosis, impeded epithelial wound healing, and activated p38 and JNK in RPTECs

(A) Representative Western blots (left) and densitometric quantitative results (right panel) of TAZ and β-actin in human RPTECs transfected with empty vector or TAZ shRNA (TAZKD) (n=5 per group, t test). (B) Cell viability measured (MTT assay) in human RPTECs with or without TAZ knockdown (n=5 per group, t test). (C) Flow cytometry analysis showed that apoptotic cells (annexin V+PI+ cells) were significantly increased in TAZ knockdown RPTECs (n=5 per group, t test). (D,E) Representative images (D) and quantitative results (E) of the in vitro scratch assay in human RPTECs with or without TAZ knockdown (n=3 per group, t test). Scale bar, 200 μm. (F) Representative Western blots of p-p38, p38, p-JNK, JNK, and β-actin proteins in human RPTECs with or without TAZ knockdown after CoCl2-induced hypoxia. *P<0.05; **P<0.01; ***P<0.001.

Figure 6
TAZ knockdown exaggerated hypoxic damage and apoptosis, impeded epithelial wound healing, and activated p38 and JNK in RPTECs

(A) Representative Western blots (left) and densitometric quantitative results (right panel) of TAZ and β-actin in human RPTECs transfected with empty vector or TAZ shRNA (TAZKD) (n=5 per group, t test). (B) Cell viability measured (MTT assay) in human RPTECs with or without TAZ knockdown (n=5 per group, t test). (C) Flow cytometry analysis showed that apoptotic cells (annexin V+PI+ cells) were significantly increased in TAZ knockdown RPTECs (n=5 per group, t test). (D,E) Representative images (D) and quantitative results (E) of the in vitro scratch assay in human RPTECs with or without TAZ knockdown (n=3 per group, t test). Scale bar, 200 μm. (F) Representative Western blots of p-p38, p38, p-JNK, JNK, and β-actin proteins in human RPTECs with or without TAZ knockdown after CoCl2-induced hypoxia. *P<0.05; **P<0.01; ***P<0.001.

Verteporfin suppresses TAZ, exaggerates hypoxic and oxidative injury, and impedes epithelial wound healing in RPTECs

Verteporfin reduces TAZ levels in fibroblasts and the kidney [15,24]. We found that verteporfin can substantially suppress the expression of TAZ (Figure 7A), decrease cell viability (Figure 7B), increase cell apoptosis and the activation of caspase-3 in CoCl2-treated human RPTECs (Figure 7D), decrease cell viability in H2O2-treated human RPTECs (Figure 7C), and inhibit epithelial wound healing of human RPTECs (Figure 7E).

Verteporfin suppressed TAZ expression, exaggerated hypoxic injury, oxidative stress and apoptosis, and impeded epithelial wound healing in RPTECs

Figure 7
Verteporfin suppressed TAZ expression, exaggerated hypoxic injury, oxidative stress and apoptosis, and impeded epithelial wound healing in RPTECs

(A) Representative Western blots (left) and densitometric quantitative results (right panel) of TAZ and β-actin in human RPTECs treated with CoCl2 (300 μM, 6 h) and different concentrations (62.5–250 nM) of VP (n=3 per group, ANOVA). *P<0.01 versus control; #P<0.01 versus 300 μM CoCl2; ##P<0.001 versus 300 μM CoCl2. (B) Cell viability (MTT assay) in human RPTECs treated with CoCl2 (300 μM) and different concentrations (62.5–250 nM) of Verteporfin (VP; n=3–6 per group, ANOVA). *P<0.05 versus control; #P<0.05 versus CoCl2 alone. (C) Cell viability measured by MTT in human RPTECs treated with H2O2 (250 μM) and different concentrations (62.5–250 nM) of VP (n=5 per group, ANOVA). *P<0.05 versus control; #P<0.05 versus H2O2 alone. (D) Apoptotic cells (annexin V+PI+ cells, left panel) and cleaved caspase-3 (right panel) by flow cytometry in human RPTECs treated with CoCl2 and different concentrations (62.5–250 nM) of VP (n=4–8 per group, ANOVA). **P<0.01 versus control, ***P<0.001 versus control; ###P<0.001 versus 300 μM CoCl2; §P<0.05 versus 300 μM CoCl2 plus 62.5 nM VP; §§§P<0.001 versus 300 μM CoCl2 plus 62.5 nM VP; &&&P<0.001 versus 300 μM CoCl2 plus 125 nM VP. (E) VP treatment (10 nM) inhibited wound healing in an epithelial monolayer composed of human RPTECs (n=3 per group, t test). Scale bar, 200 μm. *P<0.05 versus vehicle; **P<0.01 versus vehicle.

Figure 7
Verteporfin suppressed TAZ expression, exaggerated hypoxic injury, oxidative stress and apoptosis, and impeded epithelial wound healing in RPTECs

(A) Representative Western blots (left) and densitometric quantitative results (right panel) of TAZ and β-actin in human RPTECs treated with CoCl2 (300 μM, 6 h) and different concentrations (62.5–250 nM) of VP (n=3 per group, ANOVA). *P<0.01 versus control; #P<0.01 versus 300 μM CoCl2; ##P<0.001 versus 300 μM CoCl2. (B) Cell viability (MTT assay) in human RPTECs treated with CoCl2 (300 μM) and different concentrations (62.5–250 nM) of Verteporfin (VP; n=3–6 per group, ANOVA). *P<0.05 versus control; #P<0.05 versus CoCl2 alone. (C) Cell viability measured by MTT in human RPTECs treated with H2O2 (250 μM) and different concentrations (62.5–250 nM) of VP (n=5 per group, ANOVA). *P<0.05 versus control; #P<0.05 versus H2O2 alone. (D) Apoptotic cells (annexin V+PI+ cells, left panel) and cleaved caspase-3 (right panel) by flow cytometry in human RPTECs treated with CoCl2 and different concentrations (62.5–250 nM) of VP (n=4–8 per group, ANOVA). **P<0.01 versus control, ***P<0.001 versus control; ###P<0.001 versus 300 μM CoCl2; §P<0.05 versus 300 μM CoCl2 plus 62.5 nM VP; §§§P<0.001 versus 300 μM CoCl2 plus 62.5 nM VP; &&&P<0.001 versus 300 μM CoCl2 plus 125 nM VP. (E) VP treatment (10 nM) inhibited wound healing in an epithelial monolayer composed of human RPTECs (n=3 per group, t test). Scale bar, 200 μm. *P<0.05 versus vehicle; **P<0.01 versus vehicle.

Manipulation of renal TAZ expression in vivo affects severity of renal damage after ischemic AKI in mice

To evaluate the role of tubular TAZ in renal IRI, TAZ or control siRNAs were administered into the bilateral renal pelvis of each mouse before IRI. The expression and nuclear localization of TAZ in renal tubules decreased after administration of TAZ siRNA (Figure 8A,B). In vivo TAZ knockdown exaggerated renal dysfunction, tubular injury (Figure 8C), and apoptosis (Figure 8D) in the renal cortex after renal IRI.

TAZ knockdown in renal tubular epithelial cells by bilateral intrapelvic administration of TAZ siRNA exaggerated renal dysfunction, tubular injury, and apoptosis after ischemic AKI

Figure 8
TAZ knockdown in renal tubular epithelial cells by bilateral intrapelvic administration of TAZ siRNA exaggerated renal dysfunction, tubular injury, and apoptosis after ischemic AKI

(A) Immunofluorescence staining showed that TAZ siRNA significantly reduced tubular TAZ expression after renal IRI. (B) Representative Western blots and densitometric quantitative results of TAZ and β-actin between control (IRI+TR) and TAZ knockdown (IRI+TAZ siRNA) mice after IRI. (C) TAZ knockdown exaggerated tubular damage (representative images of the left panel and TIS of the right panel) and renal dysfunction (right panel). (D) TAZ knockdown increased apoptotic (TUNEL+) renal tubular cells (n=6 per group, t test). *P<0.05, **P<0.01, and ***P<0.001 versus IRI+TR group. Scale bar, 50 μm. Abbreviations: AQP1, aquaporin 1; DAPI, 4′,6-diamidino-2-phenylindole; TIS, tubular injury score; TR, transfection reagent.

Figure 8
TAZ knockdown in renal tubular epithelial cells by bilateral intrapelvic administration of TAZ siRNA exaggerated renal dysfunction, tubular injury, and apoptosis after ischemic AKI

(A) Immunofluorescence staining showed that TAZ siRNA significantly reduced tubular TAZ expression after renal IRI. (B) Representative Western blots and densitometric quantitative results of TAZ and β-actin between control (IRI+TR) and TAZ knockdown (IRI+TAZ siRNA) mice after IRI. (C) TAZ knockdown exaggerated tubular damage (representative images of the left panel and TIS of the right panel) and renal dysfunction (right panel). (D) TAZ knockdown increased apoptotic (TUNEL+) renal tubular cells (n=6 per group, t test). *P<0.05, **P<0.01, and ***P<0.001 versus IRI+TR group. Scale bar, 50 μm. Abbreviations: AQP1, aquaporin 1; DAPI, 4′,6-diamidino-2-phenylindole; TIS, tubular injury score; TR, transfection reagent.

Moreover, to evaluate the effects of pharmacologically elevating or suppressing renal TAZ expression on ischemia–reperfusion-induced renal damage, we administered chloroquine or verteporfin intraperitoneally to alter TAZ expression in the kidneys. Mice treated with chloroquine (10 or 50 mg/kg per day intraperitoneally for 6 consecutive days) significantly increased TAZ expression in renal cortical tissues after IRI compared with those treated with PBS (Figure 9A). Chloroquine treatment enhanced nuclear translocation of TAZ in mouse proximal renal tubules (Supplementary Figure S10), preserved renal function, ameliorated tubular injury, and reduced apoptosis in the renal cortex after IRI in a dose-dependent manner (Figure 9B,C; Supplementary Figure S11). Next, we tested the nephrotoxicity of verteporfin and found that it (100 mg/kg intraperitoneally every other day for three times) did not cause AKI or tubulotoxicity in mice (Supplementary Figure S12). Moreover, verteporfin treatment reduced cytosolic and nuclear TAZ expression, aggravated renal tubular injury and apoptosis in the renal cortex while elevating serum creatinine and blood urea nitrogen levels in mice with IRI compared with vehicle-treated mice (Figure 9D,E; Supplementary Figures S13 and S14).

Chloroquine (CQ) up-regulated TAZ expression in mouse renal cortical tissues and ameliorated renal failure, tubular injury, and apoptosis after ischemic AKI

Figure 9
Chloroquine (CQ) up-regulated TAZ expression in mouse renal cortical tissues and ameliorated renal failure, tubular injury, and apoptosis after ischemic AKI

In contrast, verteporfin (VP) down-regulated TAZ expression, and exaggerated renal failure, tubular injury, and apoptosis following ischemic AKI. (A) Representative images (left upper panel, Western blots; left lower panels, immunohistochemical staining) and quantitative results (left upper panel, densitometry; right panel, immunohistochemical staining values [QISV]) of TAZ in renal cortical tissues of renal IRI mice with and without chloroquine treatment (n=6 per group). Low- (10 mg/kg/day intraperitoneally for 6 consecutive days) and high-dose (50 mg/kg/day intraperitoneally for 6 consecutive days) chloroquine treatments (LDCQ and HDCQ, respectively) significantly increased TAZ expression in renal tubules. *P<0.05 versus IRI plus PBS, **P<0.01 versus IRI+PBS. Scale bar, 50 μm. (B) Representative images of Hematoxylin and Eosin staining of sham-operated mice and IRI mice treated with PBS, LDCQ, and HDCQ. (C) Quantitative results of blood urea nitrogen (BUN), creatinine (Cr), and renal tubular injury score (TIS) after IRI (n=5–6 per group, ANOVA). *P<0.05 versus IRI plus PBS group; #P<0.05 versus IRI plus LDCQ group. Scale bar, 50 μm. (D) Verteporfin treatment decreased TAZ expression in renal cortical tissue after IRI. Left upper panel shows Western blots of TAZ in renal cortex of IRI mice. Left lower and right panels show the representative immunohistochemical staining images and QISV of TAZ in renal cortex of IRI mice. (n=6 per group, t test) *P<0.05 versus IRI+DMSO group. Scale bar, 50 μm. (E) Treatment of verteporfin increased TISs, serum levels of Cr, and BUN in mice with renal IRI. (n=6 per group, t test) *P<0.05 versus IRI+DMSO group. Scale bar, 50 μm. Abbrevaitions: CQ, chloroquine; HD, high dose; LD, low dose.

Figure 9
Chloroquine (CQ) up-regulated TAZ expression in mouse renal cortical tissues and ameliorated renal failure, tubular injury, and apoptosis after ischemic AKI

In contrast, verteporfin (VP) down-regulated TAZ expression, and exaggerated renal failure, tubular injury, and apoptosis following ischemic AKI. (A) Representative images (left upper panel, Western blots; left lower panels, immunohistochemical staining) and quantitative results (left upper panel, densitometry; right panel, immunohistochemical staining values [QISV]) of TAZ in renal cortical tissues of renal IRI mice with and without chloroquine treatment (n=6 per group). Low- (10 mg/kg/day intraperitoneally for 6 consecutive days) and high-dose (50 mg/kg/day intraperitoneally for 6 consecutive days) chloroquine treatments (LDCQ and HDCQ, respectively) significantly increased TAZ expression in renal tubules. *P<0.05 versus IRI plus PBS, **P<0.01 versus IRI+PBS. Scale bar, 50 μm. (B) Representative images of Hematoxylin and Eosin staining of sham-operated mice and IRI mice treated with PBS, LDCQ, and HDCQ. (C) Quantitative results of blood urea nitrogen (BUN), creatinine (Cr), and renal tubular injury score (TIS) after IRI (n=5–6 per group, ANOVA). *P<0.05 versus IRI plus PBS group; #P<0.05 versus IRI plus LDCQ group. Scale bar, 50 μm. (D) Verteporfin treatment decreased TAZ expression in renal cortical tissue after IRI. Left upper panel shows Western blots of TAZ in renal cortex of IRI mice. Left lower and right panels show the representative immunohistochemical staining images and QISV of TAZ in renal cortex of IRI mice. (n=6 per group, t test) *P<0.05 versus IRI+DMSO group. Scale bar, 50 μm. (E) Treatment of verteporfin increased TISs, serum levels of Cr, and BUN in mice with renal IRI. (n=6 per group, t test) *P<0.05 versus IRI+DMSO group. Scale bar, 50 μm. Abbrevaitions: CQ, chloroquine; HD, high dose; LD, low dose.

Discussion

For the first time, we demonstrated the renoprotective role of TAZ in ischemic AKI (Figure 10). TAZ adaptively increases in RPTECs subjected to hypoxia or oxidative stress. During hypoxia, TAZ increases by the regulation of IKK/NF-κB signaling and translocates into the nucleus. TAZ overexpression protects RPTECs against hypoxia and oxidative stress, inhibits caspase-8-dependent apoptosis, accelerates wound healing, and inhibits the activation of p38 and JNK in RPTECs. Contrarily, TAZ knockdown produced the opposite effects. Moreover, pharmacologically enhancing or suppressing TAZ by chloroquine or verteporfin in vivo significantly ameliorated or exaggerated renal failure, tubular injury, and apoptosis in tubular cells after renal IRI, respectively.

A simple schematic drawing illustrating the protective effects of tubular TAZ on ischemic AKI

Figure 10
A simple schematic drawing illustrating the protective effects of tubular TAZ on ischemic AKI

Hypoxia augments TAZ expression and nuclear translocation through IKK/NF-κB signaling in the proximal tubular cells, followed by down-regulation of JNK, p38, caspase-8 and caspase-3, inhibition of apoptosis, and promotion of cell proliferation, thereby ameliorating ischemic AKI.

Figure 10
A simple schematic drawing illustrating the protective effects of tubular TAZ on ischemic AKI

Hypoxia augments TAZ expression and nuclear translocation through IKK/NF-κB signaling in the proximal tubular cells, followed by down-regulation of JNK, p38, caspase-8 and caspase-3, inhibition of apoptosis, and promotion of cell proliferation, thereby ameliorating ischemic AKI.

TAZ and YAP are important downstream effectors of the highly conservative Hippo tumor suppressor pathway, which is strongly related to cell growth, organ size control, and tumorigenesis [9,25,26]. Recent studies have shown new functions for the Hippo pathway, where TAZ/YAP and their upstream Hippo kinases Lats 1/2 can control myofibroblast formation and renal fibrogenesis [15,24,27]. Moreover, there the role of the Hippo pathway in AKI has been investigated. YAP promoted the repair of injured renal tubules after renal IRI in rats and was important for cell proliferation and cell cycle progression in human RPTECs [12]. Moreover, YAP activation plays an essential role during recovery from AKI [14]. However, there is a differential regulation of TAZ and YAP in response to oxidative stress, since reactive oxygen species augmented TAZ, but not YAP, in HEK293 cells [28]. We demonstrated the beneficial effects of TAZ on AKI and provided insight on the Hippo pathway as a novel target for preventing AKI in the future. Further studies are needed to elucidate whether TAZ and YAP can collaboratively protect kidneys against AKI.

Traditionally, the expression and biological function of TAZ are regulated by Hippo tumor suppressor pathway. However, other TAZ regulations have been reported [29,30]. HIF-1α has been reported to bind to the WWTR1 gene to promote the transcription and nuclear localization of TAZ in breast cell lines [29]. Moreover, HIF-1α reciprocally interacted with TAZ and synergistically drove the expression of downstream genes in hypoxic breast cancer cells [31]. Moreover, TAZ expression is regulated by PI3K/Akt signaling pathway in HeLa, fibroblast, and breast cancer cells [30]. We found that TAZ was up-regulated and NF-κB was activated in RPTECs during hypoxia. Moreover, inhibition of IKK and NF-κB suppressed the expression of TAZ induced by hypoxia in RPTECs. Therefore, pharmacological targeting of the IKK/NF-κB pathway may regulate TAZ expression and its biological effects in renal tubular cells subjected to hypoxia injury. Our findings could facilitate future novel drug screens looking to modify TAZ expression for clinical applications in AKI.

Additionally, TAZ inhibits apoptosis in lung cancer and breast cell lines [32,33], which was also observed here, with RPTECs. TAZ overexpression increased anti-apoptotic connective tissue growth factor (CTGF) and Bcl-2, decreased Bax, and activated the PI3K/Akt pathway in breast cell lines [33]. Here, the caspase-9-dependent mitochondrial pathway was not inhibited by TAZ overexpression (Supplementary Figure S15). Instead, TAZ overexpression inhibited caspase-8-dependent and TNF-α-induced apoptosis in RPTECs. Moreover, we showed that TAZ overexpression inhibited p38 and JNK MAPK signaling pathways in RPTECs. These findings may underlie major anti-apoptotic/pro-survival effects of TAZ in RPTECs.

Several kinases, including p38 or JNK of MAPK pathway phosphorylate TAZ or YAP [34]. We found a link between TAZ and MAPK pathway that may underly the beneficial effects of TAZ in renal tubular cells on hypoxic injury. To the best of our knowledge, the regulation of MAPK, such as JNK or p38, by TAZ has not been studied. Further studies clarifying the interaction between TAZ and JNK or p38 MAPK pathways are needed.

TAZ i also important for wound healing in RPTECs. TAZ has an oncogenic role in promoting cell proliferation, migration, and epithelial–mesenchymal transition [35–37]. TAZ and YAP knockdowns delay the rate of skin wound closure [11]. TAZ acts as a redox sensor in response to oxidative stress [28]. However, the biological function of TAZ in RPTECs or animal models of AKI have not been studied. We showed that TAZ knockdown induced G0/G1 cell-cycle arrest. Contrarily, TAZ overexpression enhanced cell proliferation and accelerated RPTEC wound closure. Thus, TAZ could be a potential target for preventing AKI and tubular regeneration after AKI.

To establish the independent role of TAZ in hypoxia-induced damage in RPTECs, we used two TAZ stable (overexpression and knockdown) cell lines to confirm the results. However, we did not use TAZ knockout mice because they would develop polycystic kidney disease at the age of 8 weeks and, thus, may not be appropriate for AKI studies [38]. Verteporfin, a member of the porphyrin family, is widely used for photodynamic therapy for patients with macular degeneration [39]. Verteporfin has been demonstrated to have anti-tumor effects through the inhibition of TAZ/YAP [25]. Verteporfin suppresses TAZ/YAP expression in fibroblasts, and in the kidney, and inhibits fibrogenesis after unilateral ureteral obstruction [15,24]. Here, verteporfin treatment or TAZ knockdown elicited similar results in RPTECs: down-regulated TAZ expression, exaggerated hypoxic injury and oxidative stress, increased apoptosis, and impeded wound healing. In vivo, verteporfin treatment also exaggerated renal failure and tubular injury after renal IRI. These findings suggest that verteporfin should be used carefully in patients with or at a high risk of AKI.

Chloroquine induces the activation of NF-κB in several cell lines [21,22] and it has a high tissue-to-plasma concentration ratio (∼670) in the murine kidney [40]. Since TAZ is regulated by IKK/NF-κB signaling in RPTECs, we hypothesized that chloroquine could enhance TAZ expression in RPTECs and kidneys. We found that chloroquine up-regulated TAZ in RPTECs and in vivo. Moreover, our previous study showed the beneficial effect of hydroxychloroquine, a similar drug to chloroquine, in the development of CKD in patients with rheumatoid arthritis [41]. Chloroquine has been shown to both attenuate septic or ischemic AKI [42,43] and to inhibit autophagy and enhance AKI when administered right before the renal injury and daily afterward [44]. Here, chloroquine treatment was started 5 days before the induction of renal IRI to increase the expression of TAZ in the kidney, instead of inhibiting autophagy during AKI. Although the inhibition of toll-like receptor 9 or autophagy, stimulation of nitric oxide synthase, or inhibition of proinflammatory cytokines have been observed, the mechanism for the protective role of chloroquine in AKI remains unclear [43]. We also found that chloroquine can inhibit the autophagic flux in RPTECs (Supplementary Figure S6), which is consistent with previous studies. Chloroquine treatment or TAZ overexpression also elicited similar results, indicating that TAZ may be a key player in renal protection. However, the renoprotective effect of chloroquine on ischemic AKI seems to be dose-dependent, despite the fact that no further increase in TAZ expression in the high-dose chloroquine group indicates the involvement of multiple pathways. Further clinical studies are needed to determine the impact of chloroquine or verteporfin on AKI.

We showed the potential beneficial role of tubular TAZ in ischemic AKI, which was inconsistent with a previous study [14], in which inducible deletion of TAZ in RPTECs did not exaggerate renal failure after renal IRI. We propose three possible reasons. First, the protective effect of TAZ might not be exclusively in RPTECs, but also in other tubular segments in the kidney. Second, mice with different genetic backgrounds may have variable susceptibility to IRI. Their representative figures show a trend of worsened tubular damage in mice with TAZ gene deletion in RPTECs, though it was not statistically significant. Third, the possible off-target effects of TAZ shRNA cannot be excluded. Therefore, more studies are needed to clarify the role of TAZ in other renal tubular segments or constructing RNAi-resistant TAZ in renal tubular cells to eliminate the off-target effect.

Overall, TAZ was adaptively up-regulated in renal tubular cells in response to hypoxia and oxidative stress via the IKK/NF-κB signaling pathway. Hypoxia promotes the translocation of TAZ from the cytosol to the nucleus. In RPTECs or in vivo, TAZ overexpression promoted tubular cell survival and regeneration, reduced apoptosis, and ameliorated renal failure after hypoxic injury. Suppressing TAZ resulted in the opposite effects. We demonstrate for the first time that TAZ plays a protective role in AKI and suggest TAZ as a potential preventive target against AKI.

Clinical perspectives

  • The Hippo signaling pathway regulates organ size, cell proliferation, and apoptosis, and it has been shown to be involved in the pathogenesis of kidney diseases.

  • We identified a novel preventive target of AKI, TAZ. We demonstrate that TAZ adaptively increases in response to in vitro hypoxia or in vivo IRI. TAZ overexpression and knockdown ameliorates and aggravates hypoxic injury in renal tubular cells (RTECs), respectively. Moreover, manipulating TAZ expression with siRNA, chloroquine, or verteporfin in the kidneys produces similar results.

  • We reveal a previously unclear role of TAZ in RTECs and suggest that elevating tubular TAZ expression may be beneficial against AKI.

Competing Interests

The authors declare that there are no competing interests associated with the manuscript.

Funding

This work was supported by the Changhua Christian Hospital Research Foundation [grant numbers 106-CCH-IRP-076, 106-CCH-NFP-001, 107-CCH-NFP-001]; the Ministry of Science and Technology [grant numbers MOST 106-2314-B-010-039-MY3, 108-2314-B-371-003, 109-2314-B-010-056-MY3]; the ‘Center for Intelligent Drug Systems and Smart Bio-devices (IDS2B)’ from The Featured Areas Research Center Program within the framework of the Higher Education Sprout Project by the Ministry of Education (MOE) in Taiwan; and the Foundation for Poison Control.

Author Contribution

C.-L.W. wrote the manuscript. C.-L.W., D.-C.T., and C.-H.C. conceived and designed the experiments. C.-L.W., T.-H.Y., and A.C.-D.T. performed the analyses. C.-L.W., T.-H.Y., A.C.-D.T., and J.-L.W. performed the experiments and collected the data. C.-L.W., T.-H.Y., C.-C.C., A.C.-D.T., J.-L.W., C.-H.C., and D.-C.T. contributed to the discussion and manuscript revision. C.-H.C. and D.-C.T. conceived the study and are the guarantors of this publication. All authors approved the final version of the manuscript.

Acknowledgements

The authors thank Drs. Ting-Huan Chen and Tzu-Cheng Su for their excellent technical assistance.

Abbreviations

     
  • AKI

    acute kidney injury

  •  
  • Bax

    Bcl-2-associated X protein

  •  
  • Bcl-2

    B-cell lymphoma 2

  •  
  • BCRC

    Bioresource Collection and Research Center

  •  
  • CKD

    chronic kidney disease

  •  
  • CoCl2

    cobalt chloride

  •  
  • DAB

    3,3′-diaminobenzidine tetrahydrochloride

  •  
  • DMSO

    dimethyl sulfoxide

  •  
  • ESRD

    end-stage renal disease

  •  
  • FMT

    fluorescence molecular tomography

  •  
  • HIF

    hypoxia-inducible factor

  •  
  • IKK

    IκB kinase

  •  
  • IRI

    ischemia–reperfusion injury

  •  
  • JNK

    c-Jun N-terminal kinase

  •  
  • MAPK

    mitogen-activated protein kinase

  •  
  • MTT

    methylthiazol tetrazolium

  •  
  • NF-κB

    nuclear factor κ-light-chain-enhancer of activated B cell

  •  
  • PBS

    phosphate-buffered saline

  •  
  • PI

    propidium iodide

  •  
  • PRTEC

    primary renal tubular epithelial cell

  •  
  • RPTEC

    renal proximal tubular epithelial cell

  •  
  • TAZ

    transcriptional co-activator with PDZ-binding motif

  •  
  • TNF

    tumor necrosis factor

  •  
  • TUNEL

    terminal-deoxynucleotidyl transferase-mediated digoxigenin-deoxyuridine nick-end labeling

  •  
  • YAP

    Yes-associated protein

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Supplementary data