Inhibitory Action of NoxA1 on Dual Oxidase Activity in Airway Cells^*,S⃞

EXPERIMENTAL PROCEDURES

Antibodies and Plasmids—Polyclonal antibodies directed against c-Myc (A-14) and anti-RhoGDI (A-20) were from Santa Cruz Biotechnology, and actin (A2066) was from Sigma. The expression plasmid for hDuox1 in pcDNA3.1 was kindly provided by F. Miot and subcloned into pGEX-2T. A point mutation, Duox1 (P1497A), was prepared by QuikChange® site-directed mutagenesis kit (Agilent-Stratagene) and verified by sequencing. The full coding sequence of hDuoxA1 was amplified by PCR with Platinum® Pfx (Invitrogen) using total RNA of small airway epithelial cells and specific primers (GenBank™ accession number DQ489735). DuoxA1 was cloned into pcDNA3.1. after adding in-frame a 3′ Myc tag. The expression plasmid for hNoxa1 in pRK5Myc was kindly provided by T. Leto. The Noxa1(V205A) and Noxa1(W436R) mutations were prepared by QuikChange® site-directed mutagenesis kit (Agilent-Stratagene) and verified by sequencing. Duox1 calcium-binding domain (amino acids 775–1026, GST-Duox1-Δ1), Duox1 C-terminal wild type (amino acids 1270–1551, GST-Duox1-Δ2) or mutated GST-Duox1M-Δ2 P1497A), the Nox2 C terminus (amino acids 290–570, GST-Nox2-Δ), or the Nox1 C terminus (amino acids 290–565, GST-Nox1-Δ) and full-length Noxa1 were inserted into pGEX-4T-3 (GE Healthcare). Proteins were expressed and affinity-purified according to standard protocols. Recombinant Noxa1 was purified after thrombin cleavage to remove the GST moiety, assessed for purity and identity by Coomassie Blue staining, and immunoblotting. Purified GST-Duox-Δ1 and Noxa1 proteins were used for polyclonal antibody generation. Antibody specificity controls were performed (supplemental Fig. 2, A and B). The Duox antibody detects Duox1 and Duox2 at 180 kDa.

Cell Culture—Small airway epithelial cells (SAEC) were purchased from Lonza (Walkersville, MD) and used at culture passages 3–4. Cells were grown in serum-free medium (SABM, Lonza) supplemented with SAGM SingleQuots (Lonza) at 37 °C in a humidified 5% CO₂ incubator. For the differentiated lung epithelial model (three-dimensional), cells were detached with trypsin and seeded onto inserts (Costar Transwell-clear culture insert, 0.4 μm pore; Corning Costar) coated with human placenta collagen type VI (15 μg/cm²; Sigma). Airway epithelial cells in three dimensions were grown in serum-free culture medium (50% SABM, 50% Dulbecco's modified Eagle's medium high glucose) supplemented with defined growth factors in the SingleQuots and 20–50 nm retinoic acid, prepared freshly to induce differentiation. After 2–4 days in immersed culture conditions, cell culture was switched to an air-liquid interface system for 3 weeks. Differentiation was assessed by MUC5AC expression, mucin production, resistance, and permeability measurement and electron microscopy. NCI-H292, A549, HEK293, HeLa, and H661 cells were cultured in the appropriate media. Cells were preincubated with DPI (25 μm) or BAPTA/AM (0–0.75 μm; BD Biosciences) for 10 and 5 min, respectively, before exposure to stimuli. Cells were treated with ionomycin (2–3 μm, Sigma) for 30–60 min.

Expression Analysis of Nox/Duox and NADPH Oxidase Components—Total RNA was extracted from NCI-H292 cells, A549 cells, or SAEC cells grown in monolayer or on inserts by using RNAzol B reagent (TelTest, Inc.) according to the manufacturer's protocol. Amplification by reverse transcription (RT)-PCR was performed. Briefly, cDNA was generated from 1 to 2 μg of extracted total RNA using Superscript II RNase H reverse transcriptase and oligo(dT) primers (Invitrogen) according to the supplier's protocol. cDNAs were amplified by standard PCR using TaqDNA polymerase (Invitrogen), except for Noxa1 and Noxo1 PCR, which were performed with the Advantage-GC 2 PCR kit (BD Biosciences). Sense and antisense primers were designed based on human sequences published in GenBank™. The primers used were as follows: 5′-CCATCGACTACACGCAGCTG-3′ (forward primer) and 5′-GTAGGCAGTCGACGTGCAGC-3′ (reverse primer) for Noxa1 cDNA; 5′-GCAGGACATCAACCCTGCACTCTC-3′ (forward primer) and 5′-CTGCCATCTACCACACGGATCTGC-3′ (reverse primer) for Duox1 cDNA; 5′-ATGTTCTTTCCGACGTGGTGAGCGTGGA-3′ (forward primer) and 5′-CGAATCCAGTAGTTGTTGCAGACCCTGAGA-3′ (reverse primer) for Duox2 cDNA; 5′-GCAGTTTAAGACCATTGCAGG-3′ (forward primer) and 5′-GTCAGGTTCTCCATGACTCCG-3′ (reverse primer) for Nox5 long form cDNA; 5′-CAATCCCTGTGCTCCTGC-3′ (forward primer) and 5′-GTCTGACGTTTCCAACACGC-3′ (reverse primer) for Noxo1 cDNA; and 5′-CGGGATTCACAATGGGAAACTGGGTGG-3′ (forward primer) and 5′-CAAGATAGAAGCAAAGGGGGTGAC-3′ (reverse primer) for Nox1 cDNA. RT-PCR products were resolved by electrophoresis on ethidium bromide-stained agarose gels and visualized by UV.

Small Interfering RNA (siRNA) Treatment—Double strand siRNA targeting hDuox was designed with BLOCK-it RNAi Designer (Invitrogen) to target both Duox1 and Duox2 (GenBank™ accession numbers AF213465 and AF267981, respectively). Predesigned hNoxa1 siRNA sets (NOXA1HSS145807, NOXA1HSS145808, and NOXA1HSS145809) were purchased from Invitrogen. The sequences of Duox siRNA 1 were (sense) GGACAAGGAGGAACUGACAUGGGAA and (antisense) UUCCCAUGUCAGUUCCUCCUU GUCC, and the sequences of Duox siRNA 2 were (sense) GCAUAAGUUUGAGGUGUCAGUGUUA and (antisense) UAACACUGACACCUCAAACUUAUGC. Silencer Negative Control siRNA GC medium (Invitrogen) was used as control siRNA. siRNA transfection into cells was carried out by using Lipofectamine 2000. To optimize conditions, different concentrations of siRNA (1–100 nm) and various duration of RNAi treatment were applied. Two separate transfections of the same cell population with siRNA were performed before ROS generation was measured.

Measurement of H₂O₂ Release—H₂O₂ generation was determined by measuring the oxidation of homovanillic acid (HVA) into its fluorescent derivative in the presence of horseradish peroxidase as described previously (27). The activity is expressed in fluorescent units or quantified using a standard curve obtained by incubating increasing amounts of H₂O₂ (0–25 μm) in HVA solution. In parallel, cells were scraped and lysed in RIPA buffer, and the protein concentration was determined using the BCA protein assay kit (28). H₂O₂ generation was then normalized to the protein concentration. The calcium dependence of H₂O₂ production was determined by assaying cells in parallel in HVA solution prepared with calcium-free phosphate-buffered saline or by preincubating cells with BAPTA/AM as described above.

Western Blot Analysis and Co-immunoprecipitation—Protein extracts in Laemmli buffer (2% SDS, 100 mm dithiothreitol) were homogenized by repeatedly passing through a 0.24-mm gauge syringe to obtain total lysate. Protein samples were heated 5 min at 65 °C for Duox detection and subjected to SDS-PAGE followed by transfer to nitrocellulose membrane. Blots were blocked in TBS containing 5% fat-free dry milk and 0.1% Tween before incubation with primary antibodies and secondary horseradish peroxidase-coupled antibodies. Detection was achieved by ECL (Pierce). A nondenaturing lysis buffer (50 mm Tris-HCl, 150 mm NaCl, 2% Triton X-100, 5 mm EGTA, and inhibitors mixture set I and II (Sigma), pH 7.4) was used for immunoprecipitation experiments. After preclearing, the extracts were immunoprecipitated with Noxa1 antibody and then bound to a mix of 50% protein A- and G-Sepharose. The immunoprecipitates were collected by centrifugation and analyzed by SDS-PAGE and Western blotting as described above. The effect of calcium on complex formation between Duox and Noxa1 was analyzed by adding water or calcium chloride (5 and 10 mm) to lysates (prepared with 5 mm EGTA-containing lysis buffer, see above) and split into equal aliquots before addition. Free final calcium concentration in the experiment was estimated to be 0.3–1 μm.

Immunofluorescence—SAEC, NCI-292, and A459 cells were seeded on a glass coverslips coated with human placenta collagen type VI (10 μg/ml), HEK293, and HeLa cells on polylysine or fibronectin. At 60% confluence or after transfection (siRNA or overexpression), the cells were fixed with 4% paraformaldehyde for 10 min. After permeabilization with 0.5% Triton X-100 and blocking with 5% bovine serum albumin in phosphate-buffered saline, coverslips were incubated for 2 h at room temperature with either anti-Duox or anti-Noxa1 antibody in 2% bovine serum albumin/phosphate-buffered saline. Rabbit preimmune sera were used to verify the specificity of the staining. The secondary antibodies were anti-rabbit IgG conjugated with Alexa Fluor 488 or 568 (Invitrogen). Coverslips were mounted and examined with indirect immunofluorescence (Olympus IX70) and confocal microscopy (×6, MRC 2100; Bio-Rad). All images are representative of several independent experiments recording multiple cells in different sections of the coverslip.

Cell Fractionation—With or without stimulation, cells were harvested and resuspended in relaxation buffer (KCl 100 mm, NaCl 3 mm, MgCl₂ 3.5 mm, EGTA 1 mm, Hepes 10 mm, PIPES 0.5 mm, pH 7.4). The cells were then disrupted by two 15-s cycles of sonication at 4 °C using a microprobe sonicator (Microson XL). Unbroken cells and nuclei were pelleted by centrifugation at 600 × g for 10 min at 4 °C. The supernatant (S1) was then centrifuged at 100,000 × g for 30 min at 4 °C in an Optima TLX ultracentrifuge with a TLA 100.3 rotor (Beckman Instruments Inc., Palo Alto, CA). The high speed supernatant (S2) represented the soluble cytosolic fraction. The pellet (P2) was resuspended in relaxation buffer with vigorous mixing, and this sample was again centrifuged for 15 min at 100,000 × g at 4 °C. The final supernatant (S3) represented a wash fraction. The final pellet (P3), representing the membrane fraction, was resuspended in relaxation buffer. An equivalent of 100–200 μg of protein was resolved by gel electrophoresis, and the membranes were probes for Noxa1, Duox, actin, RhoGDI, HSP72, or epidermal growth factor receptor.

Pulldown Assay—Recombinant GST, GST-Duox1-Δ1, GST-Duox1-Δ2, GST-Duox1M-Δ2, GST-Nox1-Δ, and GST-Nox2-Δ bound to Sepharose beads were incubated with purified, recombinant Noxa1 for 3 h at 4 °C. The beads were pelleted and washed three times with phosphate-buffered saline. Proteins were resuspended in Laemmli buffer and boiled 5 min to elute proteins from the beads. The amount of Noxa1 and GST fusion proteins contained in the eluates was evaluated by electrophoresis using anti-Noxa1 and anti-GST antibodies and quantified by densitometry performed with ImageJ software. The quantity of Noxa1 bound to the different GST-coupled peptide fragments was normalized by the ratio Noxa1/GST density versus the background represented by GST beads alone using values from four different experiments. For GST-Nox1-Δ and GST-Duox1M-Δ2, two experiments were performed.

Transfection—HEK293 cells were seeded in 6-well plates for 24 h prior to transfection, which was performed using Lipofectamine 2000 according to the manufacturer's protocol. In studies expressing mutant proteins, equal amounts of mutant plasmid were used in place of wild type plasmid. Twenty four hours after transfection, HEK293 cells were assayed for ROS release (with or without ionomycin stimulation) using the HVA assay, as described above.

Statistical Analysis—Experiments were evaluated with the Student's t test. p < 0.05 was considered statistically significant. The symbol ^* indicates a value of p < 0.05; ^** indicates p < 0.01, and ^*** indicates p < 0.001.

RESULTS

Expression of Oxidase Components in Human Airway Epithelial Cells—Several NADPH oxidases have been implicated in ROS production in the airways. The calcium-activated oxidases Duox1 and Duox2 were linked to H₂O₂ generation in lung epithelial cells (16, 17). Other reports connected the Nox1/Noxo1 system as well as Nox3 to tumor necrosis factor receptor or Toll-like receptor4 signaling in ROS-mediated lung alterations (29–31). RT-PCR and immunoblotting were performed to assess expression of NADPH oxidases and oxidase-regulatory components in primary human small airway cells (SAEC) and in the lung cancer cell lines NCI-H292 and A549. Duox1 and Duox2 message and protein were expressed in SAEC and NCI-H292 cells, whereas A549 cells showed faint transcripts for Duox2 only, and expressed no detectable Duox protein by immunoblot analysis. SAEC cell lysates showed a 180-kDa band representing mature Duox (Fig. 1A, see also Fig. 1C) together with a sporadic appearance of an unidentified, faster migrating band that disappeared upon airway cell differentiation and upon Duox RNAi. Comparison of SAEC cells cultured in monolayer versus air-liquid interface (ALI) conditions on inserts (three-dimensional SAEC) indicated a 5–10-fold increase in Duox expression after differentiation (Fig. 1A). This result correlates well with the initial suggestion by Geiszt and co-workers (16) that Duox expression increases with cell differentiation. Other oxidases or certain oxidase regulatory components were absent from the cell types used in this study (supplemental Fig. 1A and data not shown).

Duox-mediated ROS Generation—As a physiological stimulus for Duox activation remains unidentified, the Ca²⁺ ionophore ionomycin was added to SAEC or NCI-H292 cells for 30–60 min to stimulate calcium influx. In resting conditions H₂O₂ release was not detected, whereas exposure to ionomycin led to substantial H₂O₂ production. Duox expression levels correlated with the extent of H₂O₂ production, with ∼5-fold higher ROS production in differentiated SAEC (three-dimensional SAEC) than in SAEC cultured in monolayer (Fig. 1B). ATP and thapsigargin, two other stimuli known to increase calcium availability in cells, also led to a substantial increase in H₂O₂ generation in differentiated SAEC (data not shown). Similar results were obtained with Duox-expressing primary human bronchial epithelial cells and an immortalized SAEC cell line (data not shown).

By using ALI-cultured SAEC, the location of Duox-mediated H₂O₂ production was probed by treatment of the upper and lower chamber of the inserts with ionomycin. H₂O₂ production was only detected on the apical side of ionomycin-stimulated cells. This ROS production was abolished after pretreatment with diphenylene iodonium (DPI), an irreversible flavoenzyme inhibitor (Fig. 1B). The contribution of Duox proteins to ROS generation by airway cells was probed with RNAi. Two different siRNAs were designed to regions identical in Duox1 and Duox2, thus targeting both Duox homologs at the same time. Time courses for both Duox1 and Duox2 mRNA and protein expression were performed in SAEC and NCI-H292 cells (Fig. 1C and supplemental Fig. 1B). Immunoblotting confirmed depletion of Duox1 and Duox2 after 96 h of RNAi treatment, which did not affect cell viability. After 96 h of treatment with Duox siRNA ROS production was inhibited by 90–100% (Fig. 1D), whereas control siRNA had no effect. These data indicate that Duox proteins are the main source for calcium-stimulated H₂O₂ generation in primary human lung epithelial cells and H292 cells.

Noxa1 Translocates from the Membrane to the Cytosol upon Calcium Influx—The oxidase regulatory component Noxa1 was detected in cell lysates of lung epithelial cells or lung cancer cells (Fig. 2A). In resting cells immunostaining of endogenous Noxa1 showed localization at the plasma membrane on intracellular membrane structures and in the nucleus (Fig. 2B (-)). After ionomycin stimulation for 5–15 min, the majority of the Noxa1 staining at the membrane edges disappeared in SAEC and NCI-H292 cells (Fig. 2B (+)). Translocation of Noxa1 from intracellular membranes to cytosolic fractions was difficult to visualize in immunofluorescence, but it was clearly detected when cellular fractionation was used. These experiments demonstrated that a significant fraction of the membrane-bound Noxa1 protein translocated to the cytosol after ionomycin stimulation of SAEC or H292 cells (Fig. 2C). Noxa1 depletion from the plasma membrane already occurred at the earliest stages of ionomycin stimulation (5 min) and seemed to coincide with the beginning of H₂O₂ production. Duox-mediated ROS generation was an event following Noxa1 translocation rather than preceding it, because pretreatment of cells with DPI did not affect stimulation-dependent Noxa1 movement to the cytosol (data not shown).

Duox Is Localized on the Plasma Membrane and on Intracellular Membranes—Duox localization was confirmed by immunofluorescence and cell fractionation in human lung epithelial cells (Fig. 2, D and E). Confocal images showed Duox protein at the plasma membrane and on intracellular membranes in SAEC and NCI-H292 cells. The intracellular Duox pool may correspond to Duox on intracellular vesicles or to Duox retained in the endoplasmic reticulum and/or Golgi compartments (see lower exposure of SAEC cells, Fig. 2D, upper right). Because the Duox antibody recognizes Duox1 and Duox2, it is also conceivable that Duox isoforms are located at distinct membranes. Immunofluorescence of A549 cells showed minimal Duox protein in the endoplasmic reticulum (Fig. 2D). This correlates well with the immunoblot results and the lack of calcium-stimulated H₂O₂ production in this cell type (Fig. 1, A and B). Membrane localization of Duox was confirmed by immunoblotting of H292 membrane and cytosol fractions (Fig. 2E).

Association of Duox and Noxa1—The shared plasma membrane localization of Duox and Noxa1 in resting cells prompted us to determine complex formation of these proteins. Overlapping migration of Noxa1 protein and the upper IgG band did not permit immunodetection of Noxa1 in Noxa1 immunoprecipitates. Thus, we performed a control experiment to validate the Noxa1 antibody for immunoprecipitation. Noxa1 antibody immunoprecipitated Noxa1 from HEK293 cell lysates expressing Myc-Noxa1 as visualized with anti-Myc immunoblotting (Fig. 3A). To test association of endogenous proteins, Noxa1 was immunoprecipitated with increasing amounts of anti-Noxa1 antibody followed by immunoblotting for Duox. We detected increasing amounts of Duox in Noxa1 immunoprecipitates (180 kDa; Fig. 3B). Several controls were carried out to validate these data. Immunoprecipitation with anti-Noxa1 in lysates derived from Duox-deficient A549 cells versus Duox-containing NCI-H292 cells demonstrated the absence of the 180-kDa band in A549 cells (Fig. 3B). Similarly, Duox siRNA in NCI-H292 cells resulted in a significant decrease in recovered Duox after Noxa1 immunoprecipitation, when compared with control siRNA-treated cells (Fig. 3B). The presence of Noxa1 on the plasma membrane of airway cells was dependent on plasma membrane-bound Duox as demonstrated by immunofluorescence staining of SAEC cells treated with control or Duox siRNA (Fig. 3C). Some Noxa1 staining on intracellular membranes remained, indicating that either Duox knockdown was not complete or that Noxa1 may have a secondary membrane target.

Disruption of the Noxa1-Duox Association by Calcium Elevation— Duox requires the presence of micromolar concentrations of calcium to generate ROS (8). Indeed, when calcium was omitted from buffers used for ROS detection, H₂O₂ was not produced upon ionomycin stimulation (data not shown). Similarly, when cells were pretreated with the calcium chelator BAPTA/AM, H₂O₂ production was inhibited (Fig. 3D). In these conditions, translocation of Noxa1 from the membrane to the cytosol did not occur (see immunoblot Fig. 3D). This suggests that calcium elevation is, at least in part, responsible for the translocation of Noxa1. To substantiate this hypothesis, we investigated if the interaction between Duox and Noxa1 was calcium-dependent. Resting NCI-H292 cells were lysed in the presence of 5 mm EGTA with or without the addition of 5–10 mm CaCl₂ before immunoprecipitation of Noxa1 was performed. As shown in Fig. 3E, an increase in free calcium to ∼0.3–1 μm free Ca²⁺ substantially decreased the association of Duox and Noxa1 in vitro.

Noxa1 Binds to the Duox1 C Terminus in Vitro—To determine whether the association between Noxa1 and Duox is direct or mediated by another protein, protein fragments encompassing intracellular domains of Duox1 were generated. GST fusion proteins, including a fragment containing the calcium-binding EF hands (GST-Duox1-Δ1) or encompassing the C terminus (GST-Duox1-Δ2), were generated. GST protein and a C-terminal fragment of Nox1 served as positive control, whereas a C-terminal fragment of Nox2 was prepared as a negative control (supplemental Fig. 3A). These fragments were tested for their ability to interact with purified, recombinant Noxa1 in in vitro pulldown assays. Although we noticed some unspecific binding of Noxa1 to the beads, including the GST control beads, Noxa1 binding to the C terminus of Duox1 (GST-Duox-Δ2) was 8–10 times higher than the GST background values (Fig. 3F). By determining the ratio of GST fusion proteins bound to the beads versus Noxa1 binding to these proteins, the Nox2 C terminus and the EF hand-containing fragment of Duox1 exhibited clearly less binding of Noxa1 than the Duox1 C terminus. As anticipated, the C terminus of Nox1 bound to Noxa1 in the pulldown assay (supplemental Fig. 3). These results indicate that Noxa1 binds directly to the Duox1 C terminus in vitro.

Noxa1 Inhibits Duox-mediated ROS Generation in Resting Cells— The maturation factor DuoxA1 is required for Duox expression in heterologous cell systems (32). Co-transfection of Duox1 and Myc-tagged DuoxA1 in HEK293, HeLa, and H661 cells generated calcium-inducible ROS (Fig. 4A). In fact, co-expression of Duox1 and Myc-DuoxA1 generated also basal, unstimulated H₂O₂ in HEK293 and HeLa cells. This was not observed in H661 cells, which are deficient in Duox and DuoxA proteins, but express Noxa1 (see Fig. 5C). We assessed if Noxa1 expression alters the basal and stimulated H₂O₂ production by Duox1. Duox1 and DuoxA1 were co-transfected without or with Noxa1 into HEK293 cells, followed by HVA assay and immunoblotting (Fig. 4B, C). As expression levels did not change significantly between the different conditions, ROS production by Duox1/DuoxA1 was set to 100%. An inhibition of ∼40–50% of the basal H₂O₂ production was observed in the presence of Noxa1. Only a slight inhibitory effect of Noxa1 on ionomycin-stimulated H₂O₂ production was detected (Fig. 4B). This suggests that the association of Noxa1 with Duox1 acts as a negative Duox1 regulator, keeping the oxidase inactive in resting cells via complex formation. Release of Noxa1 from Duox1 was observed in transfected HEK293 cells, showing loss of Noxa1 from the membrane after ionomycin stimulation. Although, in contrast to endogenous Noxa1, overexpressed Noxa1 was partially bound to the plasma membrane in the absence of Duox1, this fraction of Noxa1 did not translocate to the cytosol after ionomycin treatment (data not shown).

The importance of the Noxa1 activation domain on Duox-mediated H₂O₂ generation was tested. Transfection of Noxa1 or Noxa1(V205A) inhibited equally well basal ROS generation by Duox1 (Fig. 4, B and C). Thus, the activation domain of Noxa1 is not required for Duox function. We then set out to evaluate the effect of Noxa1 knockdown on basal ROS production by primary airway cells. siRNA treatment decreased Noxa1 protein ∼50–60%. Noxa1 RNAi for more than 36–48 h or at higher concentration was not well tolerated by SAEC. Knockdown of Noxa1 augmented basal H₂O₂ production, but it did not alter significantly ionomycin-stimulated ROS generation (Fig. 4, D and E, and data not shown).

The Duox C-terminal PXXP Motif Is Involved in Noxa1 Binding—The Duox1 C terminus, the site of Duox1-Noxa1 association in vitro (Fig. 3F), contains a proline-rich motif, which represents a putative Noxa1 SH3 domain-binding site. We generated a Duox1 mutant in which proline 1497 was replaced by alanine and compared Duox wild type and Duox mutant for their ability to support basal and stimulated H₂O₂. Co-transfection of Noxa1 did not alter basal and stimulated H₂O₂ production by mutant Duox1(P1497A) (Duox1M) (Fig. 5, A and B). Unexpectedly, the Duox1(P1497A) mutant displayed increased ROS production, suggesting that this mutation leads to a conformational change in the Duox1 C terminus and/or altered FAD/NADPH binding. This substantially higher H₂O₂ production by Duox1M was observed in three different cell types (Fig. 5C and data not shown).

Mutation of the Duox1 PXXP site may abolish binding of Noxa1 to Duox1. When overexpression of DuoxA1, Duox1, or Duox1M and Noxa1 was carried out in HEK293 cells, expression levels of Duox1 and Duox1M were comparable (Fig. 5D). Immunoprecipitation with anti-Noxa1 antibody revealed that the Duox1 mutant can still bind to Noxa1, albeit with less efficiency (Fig. 5D). Similar results were obtained in in vitro pulldown experiments using the C termini of Duox1 or Duox1M (supplemental Fig. 3B). Additional mutation of the Noxa1 SH3 domain was carried out for co-expression experiments of Noxa1(W436R) and Noxa1wt with Duox1wt or Duox1(P1497A). Noxa1 immunoprecipitation revealed that the association between wild type and mutant Noxa1 and the Duox mutant (Duox1M) was perturbed. On the other hand, we did not observe a significant change in Noxa1 or Noxa1 mutant association with wild type Duox1. This suggests that the interaction between the Duox C-terminal PXXP motif and the Noxa1 SH3 domain alone is not sufficient to convey high affinity Noxa1 binding in vitro. A second binding site or another modification on Duox or Noxa1 seems to be required for the Duox1-Noxa1 interaction at the cell membrane and for release of Noxa1 into the cytosol. This scenario would be reminiscent of our recent observations regarding Noxa1 dissociation from the membrane-bound Nox1 complex, which was facilitated by protein kinase A and 14-3-3 protein (33).

DISCUSSION

Inflammatory cells such as alveolar macrophages or activated neutrophils are considered the major source for ROS in the lung. Lately, however, cells of the pulmonary endothelium and epithelium are also recognized as important ROS generators. The discovery of the NADPH oxidases Duox, which reside in the airway epithelium and thyroid, stimulated new studies elucidating how pulmonary ROS are produced and how these enzymes might be regulated. To analyze and understand the complex and varied functions of human airway epithelia in a controlled setting and in the absence of other cell types, an in vitro differentiation model was used in this study. This three-dimensional model system, which uses primary human bronchial or small airway cells, creates a mucociliated, pseudostratified airway epithelium after 3–4 weeks of continuous culture (34–36). Several markers for differentiation, biophysical measurements, and electron microscopy were used to ensure the presence of the phenotypic features described for a well differentiated airway epithelium (data not shown). When we examined the expression of the two Duox isoforms, Duox1 and Duox2, it became clear that the differentiation process leads to highly increased expression of Duox protein. It is presently not feasible to distinguish between Duox1- or Duox2-dependent functions, because available antibodies, including ours, cannot differentiate between the Duox isoforms. Elevated Duox expression caused, as expected, 5-fold higher output of H₂O₂ after stimulation with Ca²⁺ ionophore. Confirming earlier suggestions (16, 17), H₂O₂ generation was confined to the apical surface of the epithelial model. Knockdown studies in two different cell types determined Duox as the source of calcium-stimulated H₂O₂ generation in airway cells. It is important to note that we could not detect functional Duox in the lung cancer cell line A549, which is widely used as airway type II cell model.

Screening of human primary lung epithelial cells revealed the presence of the NADPH oxidase component Noxa1, whereas many of the other previously described oxidase proteins were missing. Noxa1 exhibits only ∼30% identity with the NADPH oxidase component p67^phox but presents very similar domain architecture (19–21). In contrast to the role of p67^phox in the Nox2-based phagocyte oxidase, Noxa1 seems to fulfill different functions in epithelial cells. The Noxa1 SH3 domain was not required for phorbol ester-induced Nox1-dependent ROS generation (37) and decreased Noxo1-stimulated Nox3 activity by ∼50% (23, 38). Here we show that endogenous Noxa1 is membrane-bound in primary lung epithelial cells and associates with Duox via its SH3 domain. Upon calcium influx Noxa1 dissociates from the membrane compartment and translocates to the cytosol. Calcium entry, but not ROS production, was required for Noxa1 translocation, indicating that dissociation was an early step in the Duox activation process. This suggests that Noxa1 may act as an inhibitory adaptor rather than an activator of Duox activity.

Recent identification of Duox maturation factors (32) such as DuoxA1 permits the processing of Duox1 and the recovery of a mature, active form of Duox1 on the plasma membrane of epithelial cell lines. We noticed that co-transfection of Duox1 with DuoxA1 triggered H₂O₂ production without prior stimulation in HEK293 cells. This ROS generation was substantially enhanced after stimulation with ionomycin. Basal ROS generation was not observed in primary human lung epithelial cells or Duox-deficient lung cancer cells (H661) where Duox was reintroduced via transfection. These cell types express Noxa1, whereas HEK293 or HeLa cells lack expression of this protein. This prompted us to investigate the role of Noxa1 in regulating ROS production by Duox in more detail. Co-transfection of Noxa1 together with Duox1 and DuoxA1 into HEK293 cells inhibited basal production of H₂O₂, but it did not affect calcium-stimulated ROS generation when a strong calcium flux inducer like ionomycin was used. Additionally, we observed a substantial increase of basal ROS generation when endogenous Noxa1 protein was knocked down by RNAi. These data support the idea that Noxa1 functions as a negative regulator of basal Duox stimulation, which could occur in the course of spontaneous calcium oscillations. In accord, mutational analysis revealed that the activation domain of Noxa1 was dispensable for Duox regulation.

Because mutation of a proline-rich motif in the C terminus of Duox1 diminished Noxa1-dependent inhibition of basal ROS generation, we propose that the PXXP domain of Duox is involved in the Duox regulation. The C terminus of both Duox isoforms is highly homologous, suggesting that Noxa1 may regulate Duox1 and Duox2. This scenario is reminiscent of studies suggesting calcium-mediated regulation of Nox5 via intramolecular interaction of the Nox5 N terminus with the Nox5 C terminus (39). In the case of Nox5, an increase of intracellular calcium leads to binding of Ca²⁺ to the EF hands in the N terminus. This may cause a conformational change, which then provides an interface for the interaction between the N terminus and the C terminus, leading to electron transport and oxidase activation. Although detailed in vitro studies with recombinant Duox might be necessary to deduce the Duox activation process, our data suggest an inhibitory mechanism via the Duox C terminus. This inhibition is relieved when Ca²⁺ binds to the two EF hands, causing Noxa1 to drop off. Most of the endogenous Noxa1 is membrane-bound in airway cells, whereas Noxa1 when expressed in model cell lines localizes to the membrane and the cytosol. Thus, the overexpression system does not exactly reflect the Duox-Noxa1 regulation in lung epithelial cells. This may account for the only partial effect of Duox1 P1497A or Noxa1 W436R mutations in abolishing Duox1-Noxa1 interaction. It is also likely that additional modifications on Duox or Noxa1 are required to keep Noxa1 in the cytosol during Duox1 activation and that other yet unknown proteins are involved in Noxa1 shuttling from the membrane to the cytosol.

Regions in close proximity to the Duox PXXP domain seem to be critical for regulation of oxidant production by Duox, because a point mutation in this region increased basal and calcium-stimulated H₂O₂ generation significantly. This might reflect the importance of the Duox C terminus as control mechanism for Ca²⁺ binding or for termination of the electron flow. Limited proteolysis of thyroid NADPH oxidase by α-chymotrypsin results in irreversible, calcium-independent H₂O₂ production (40). These data together with our observation support the idea that Duox activity might be regulated via the displacement or conformational change of an inhibitory domain. Although Nox5 and Duox regulation have some regulatory features in common, such as Ca²⁺-mediated conformational change and activation by relieving inhibition, there seem to be also very distinctive differences. A role for Noxa1 or any other regulatory protein has not been reported for Nox5. Duox and Nox5 are emerging as NADPH oxidases that use an inhibitory mechanism in the basal state, which represents a novel way of oxidase regulation. Because binding of calcium is also used as activation mechanisms for plant oxidases (41, 42), this model of activation may have broader relevance for Ca²⁺-activated oxidases in general.

Supplementary Material