Unchanged NADPH Oxidase Activity in Nox1-Nox2-Nox4 Triple Knockout Mice: What Do NADPH-Stimulated Chemiluminescence Assays Really Detect?

Abstract

NADPH oxidases of the Nox family are considered important sources of cellular reactive oxygen species (ROS) production. This conclusion is, in part, based on the ability of NADPH to elicit a chemiluminescence signal in tissue/cell homogenates or membrane preparations in the presence of enhancers such as lucigenin, luminol, or L012. However, the ability of these particular assays to specifically detect Nox activity and Nox-derived ROS has not been proven. In this study, we demonstrate that combined knockout of the three main Nox enzymes of the mouse (Nox1-Nox2-Nox4 triple knockout) had no impact on NADPH-stimulated chemiluminescence signals in the aorta, heart, and kidney homogenates. In the NADPH-stimulated membrane assays, no effect of in vivo angiotensin II pretreatment or deletion of Nox enzymes was observed. In in vitro studies in HEK293 cells, the overexpression of Nox5 or Nox4 markedly increased ROS production in intact cells, whereas overexpression of Nox5 or Nox4 had no influence on the signal in membrane assays. In contrast, overexpression of nitric oxide synthase or cytochrome P450 enzymes resulted in an increased chemiluminescence signal in isolated membranes. On the basis of these observations, we propose the hypothesis that NADPH-stimulated chemiluminescence-based membrane assays, as currently used, do not reflect Nox activity. Antioxid. Redox Signal. 24, 392–399.

Innovation

NADPH oxidases of the Nox family are considered important sources of ROS in mammalian tissue, but the assays to determine their activity ex vivo have not been carefully validated. In this study, we demonstrate using newly generated Nox1-Nox2-Nox4 triple knockout mice that the most frequently used technique (enhanced chemiluminescence) in membrane preparations does not detect Nox activity, but rather other enzymes such as NO synthase or cytochrome P450 monooxygenase. Thus, this frequently used technique should no longer be applied to determine Nox NADPH oxidase activity.

NADPH oxidases of the Nox family are important sources of reactive oxygen species (ROS). Much evidence for this came from chemiluminescence assays that made use of the enhancers lucigenin (4), L012, or luminol/horseradish peroxidase (HRP). In these, tissues or cells were homogenized and the chemiluminescence was measured after addition of the substrate, that is, NADH or NADPH to the crude homogenates or membrane preparations. To our knowledge, the true cellular ROS formation by Nox enzymes in living cells has never been systematically compared to the signal obtained in homogenates after the addition of NADPH.

To determine whether the currently used assays are detecting Nox activity, we generated a triple knockout mouse lacking Nox1, Nox2, and Nox4 (3N^−/−) and compared the signals generated in the different assays. Moreover, two separate approaches, that is, in vivo angiotensin II (AngII) treatment and ex vivo transforming growth factor-β (TGF-β) application were used to increase Nox expression.

Triple Nox Knockout Mice Are Viable and Lack Gross Abnormalities

Triple Nox-knockout mice (3N^−/−) showed no obvious phenotype. There was a trend toward lower systolic blood pressure and a significantly higher basal heart rate in 3N^−/− compared to wild-type (WT) mice (Fig. 1A–E). As shown by polymerase chain reaction (PCR), the 3N^−/− animals lacked Nox1, Nox2, and Nox4, whereas Nox3 was expressed at extremely low levels. Vascular expression of endothelial NO synthase (eNOS) was significantly decreased in 3N^−/− (Table 1). Following exposure to AngII, the expression of Nox3 increased in 3N^−/−, but not in WT mice. In the heart, eNOS and Duox2 expression was significantly higher in 3N^−/− mice. In the kidney, Nox3 and Duox2 were increased in the 3N^−/− mice, but no difference in eNOS expression was detected. eNOS expression was decreased in the lungs of 3N^−/− mice, whereas Duox1 was increased (Table 2). These data indicate that some compensation occurs in 3N^−/− mice, which might be relevant for eNOS, Duox1/2, and Nox3.

FIG. 1.

Effect of in vivo angiotensin II treatment on cardiovascular parameters. (A) Systolic, (B) diastolic, and (C) mean blood pressure. (D) Heart rate and (E) heart to body weight ratio. *p<0.05 with versus without angiotensin II (AngII, 0.7 mg/kg/day), ^# p<0.05 3N^−/− versus WT. (F, G) Ex vivo vascular function study in mesenteric arteries of animals pretreated with and without AngII for 2 weeks, *p<0.05 with versus without AngII. (F) Acetylcholine-induced relaxation after preconstriction with phenylephrine (80%) *p<0.05 WT+AngII versus other groups, (G) phenylephrine-induced constriction. *p<0.05 WT without AngII versus other groups, n=7–8. WT, wild type.

Table 1.

qRT-PCR for Mouse Aortic Tissue

Target	CT	WT	+AngII	3N^−/−	+AngII
Nox1	35.8	1±0.27	2.28±0.83	n.d.	n.d.
Nox2	29.2	1±0.15	1.17±0.24	n.d.	n.d.
Nox3	35.9	1±0.35	1.54±0.42	0.65±0.14	2.39±0.59^a
Duox1	32.9	1±0.19	1.15±0.33	0.67±0.19	0.66±0.08
Duox2	30.4	1±0.18	0.86±0.17	1.39±0.23	1.02±0.13
Xanthine oxidase	26.2	1±0.09	1.04±0.13	0.89±0.10	0.95±0.07
eNOS	26.5	1±0.18	1.26±0.19	0.72±0.16^b	0.62±0.11^b

Values are expressed relative to the copy numbers determined in control WT mice. CT is provided as a rough estimate of RNA abundance.

p<0.05 relative to 3N^−/− CTR.

p<0.05 relative to WT CTR.

CT, cycle threshold; qRT-PCR, quantitative reverse-transcriptase polymerase chain reaction.

Table 2.

qRT-PCR from Heart, Kidney, and Lung Tissue

	Heart			Kidney			Lung
Target	CT	WT	3N^−/−	CT	WT	3N^−/−	CT	WT	3N^−/−
Duox1	32.7	1±0.14	1.15±0.08	32.4	1±0.08	1.04±0.07	36.1	1±0.21	27.15±5.06^a
Duox2	32.9	1±0.11	1.98±0.34^a	35.5	1±0.30	1.75±0.20^a	30.2	1±0.18	1.16±0.08
Nox3	37.9	1±0.44	1.83±0.20	34.9	1±0.09	1.29±0.09^a	34.6	1±0.38	0.74±0.08
Xanthine oxidase	26.4	1±0.12	1.57±0.30	26.4	1±0.03	1.49±0.40	22.9	1±0.04	1.07±0.07
CytB1	13.8	1±0.04	0.56±0.39	13.8	1±0.09	0.95±0.14	16.3	1±0.03	1.09±0.21
CytC	20.4	1±0.05	1.09±0.34	20.7	1±0.14	1.39±0.06	23.1	1±0.10	1.06±0.06
eNOS	28.3	1±0.13	2.30±0.35^a	29.2	1±0.86	0.87±0.16	26.8	1±0.08	0.67±0.02^a
iNOS	28.6	1±0.17	1.27±0.20	30.3	1±0.13	1.01±0.16	28.0	1±0.18	2.18±0.66
nNOS	33.0	1±0.12	0.88±0.09	30.9	1±0.11	1.15±0.10	33.4	1±0.12	0.66±0.16
SOD1	23.0	1±0.19	0.86±0.34	21.5	1±0.13	2.04±0.69	21.8	1±0.11	0.79±0.03
SOD3	26.8	1±0.07	1.19±0.28	21.6	1±0.09	0.87±0.10	22.0	1±0.09	0.83±0.16
Catalase	25.7	1±0.15	0.75±0.27	22.5	1±0.15	1.5±0.24	23.9	1±0.10	1.16±0.39

Values are expressed relative to the copy numbers determined in WT mice. CT is provided as a rough estimate of RNA abundance.

p<0.05 relative to WT.

Triple Knockout of Nox Results in an Attenuated Response to AngII

In response to AngII, the heart rate was more markedly decreased in the 3N^−/− mice, so that in the presence of AngII, the heart rate in both strains was similar (Fig. 1D). The increase in blood pressure in response to AngII was similar in both strains, although there was a trend toward an attenuated response in 3N^−/− mice (Fig. 1A–D). As judged by the heart to body weight ratio, only WT animals developed cardiac hypertrophy to AngII (Fig. 1E). Similarly, AngII induced a rightward shift in the endothelium-dependent relaxation of mesenteric arteries from WT mice, but not 3N^−/− mice (Fig. 1F).

Interestingly, the constrictor response in mesenteric arteries from WT mice was less compared with that of all the other groups, potentially indicating that these animals had the highest basal NO availability (Fig. 1G). The compensations occurring in a triple knockout mouse are obviously complex, and the genetic situation is hard to control. Nevertheless, it is fascinating that the 3N^−/− mice comprise a mixed phenotype of the individual Nox knockout; similar to the Nox1 and Nox2 knockout mice (7), they do not develop endothelial dysfunction in response to AngII, but due to the lack of Nox4 (9) their endothelium-dependent relaxation under basal conditions is not enhanced. Similar to Nox4^−/− mice, expression of eNOS is attenuated in tissue rich of endothelial cells, whereas the failure to induce cardiac hypertrophy in response to AngII was previously noted in Nox2^−/− mice (3).

Aortic Rings from WT and Triple Knockout Give Similar Signals in Chemiluminescence Assays

To correlate these functional data with Nox activity assays, measurements with lucigenin and L012 were performed. There was no detectable difference in the basal lucigenin signal of the rings in the absence of NADPH, but its addition resulted in a massive increase in the signal. The latter was, however, similar in tissues from WT and 3N^−/− mice (Fig. 2A), indicating that the lucigenin signal generated by isolated mouse aortic rings is not derived from Nox1, Nox2, or Nox4. In the L012 assay, aortic rings from 3N^−/− mice yielded a slightly (but nonsignificantly) higher signal than WT mice. In vivo treatment with AngII had no effect on the chemiluminescence signal, but the acute application of PMA, to stimulate Nox2, induced a significant increase in the signal from WT aortic rings, but not rings from 3N^−/− mice (Fig. 2B). The decrease in the signal in response to PMA in the 3N^−/− group most probably reflects the fact that the solvent DMSO is a strong quencher of the L012 signal. Thus, L012 chemiluminescence can detect the PMA-stimulated activation of Nox2, but is unable to detect the increase in Nox1 expression/activity associated with AngII treatment with the current amount of tissue and number of samples included.

FIG. 2.

Chemiluminescence signal of intact mouse aortic rings. Three aortic rings were pooled from the strains as indicated, with or without in vivo treatment with AngII, and subsequently analyzed by chemiluminescence in response to lucigenin (A, 10 μmol/L) or L012 (B, 200 μmol/L). ⁺Indicates values obtained after addition of compounds. NADPH (100 μmol/L), PEG-SOD (50 U/ml), or +PMA (100 nmol/L in 0.1% DMSO). N=6–8, *p<0.05 with versus without compound.

Triple Nox Knockout Has No Effect on the Chemiluminescence Signal of Cardiac Membranes

Membrane preparations from WT and 3N^−/− mouse hearts were compared using all three enhancers. Whereas the membranes produced no basal chemiluminescence with lucigenin alone (data not shown), a signal was readily detectable after the addition of NADPH. This, however, was identical in samples from WT and 3N^−/− mice and not affected by in vivo AngII treatment (Fig. 3A). Likewise, with luminol/HRP, there was no difference in the NADPH-induced signal between samples from WT and 3N^−/− mice (Fig. 3B). Interestingly, the L012 assay yielded a signal by membranes even in the absence of NADPH. This signal might reflect an alteration of the autoxidation of L012 mediated by peroxidases or small molecules (F.R., unpublished observation). In response to NADPH, the signal increased by almost 10-fold and extracts from 3N^−/− hearts generated more signals than those from the WT hearts. Nevertheless, there was no difference in the signal to tissues from AngII-treated mice (Fig. 3C). On this basis, it has to be concluded that all three assays failed to detect the activity of Nox1, Nox2, and Nox4 in cardiac membranes.

FIG. 3.

Assay comparison in cardiac membranes. Preparations were obtained from hearts of WT or 3N^−/− mice treated with and without angiotensin II (AngII, 0.7 mg/kg/day, 2 weeks). (A) Lucigenin assay in the presence of NADPH (100 μmol/L) with and without PEG-SOD (50 U/ml). (B) Luminol/HRP (100 μmol/L, 1 U/ml HRP, NADPH 10 μmol/L) and (C) LO12 (200 μmol/L) with and without NADPH (10 μmol/L). N=6–7, *p<0.05 WT versus 3N^−/−. HRP, horseradish peroxidase.

Kidney Membranes Generate a Complex Chemiluminescence Signal

The kidney is the organ with the highest Nox4 expression. Amplex red/HRP assays in freshly isolated tissue revealed lower levels of H₂O₂ in samples from 3N^−/− mice (Fig. 4A). Interestingly, this effect was not observed when the samples were further processed and the assay repeated using renal membrane preparations stimulated with NADPH in the presence of luminol/HRP, which is generally assumed to detect H₂O₂ (Fig. 4B). A somewhat but not significantly lower NADPH-amplified lucigenin chemiluminescence signal was detected in membranes from 3N^−/− mice versus WT counterparts, despite the fact that Nox4 produces H₂O₂ rather than superoxide (Fig. 4C). As with the cardiac membrane preparations, L012 chemiluminescence yielded an autoxidation signal in membrane samples without NADPH, which was higher in 3N^−/− than WT preparations. Upon addition of NADPH, the L012 chemiluminescence increased and was similar between the WT and 3N^−/− preparations (Fig. 4D).

FIG. 4.

Assay comparison in kidney preparations. (A) Exemplary photo and quantification of Amplex red/HRP assay from minced living kidney. (B–D) Chemiluminescence signal of renal membranes obtained from mice pretreated with or without angiotensin II (AngII, 0.7 mg/kg/day, 2 weeks) enhanced by luminol/HRP (100 μmol/L, 1 U/ml, B), lucigenin (10 μmol/L, C), and L012 (200 μmol/L, D), n=7–8, *p<0.05 WT versus 3N^−/−, ^# p<0.05 with versus without NADPH (10 μmol/L).

In vivo AngII treatment slightly increased the NADPH-stimulated signal in the luminol/HRP assay and L012 assay of WT mouse kidney membranes, and this effect was not observed in membranes prepared from 3N^−/− mice. In contrast, the lucigenin signal of renal membranes was similar between fractions obtained from mice with and without in vivo AngII treatment (Fig. 4B–D). Collectively, these data suggest that renal membranes of 3N^−/− may have a reduced NADPH-stimulated lucigenin signal, which cannot, however, be linked directly to Nox activity.

Nox4 Is Not the Source of the Signal of the NADPH-Stimulated Luminol/HRP Assay

To further dissect the previous data, experiments were performed in murine dermal fibroblasts in the absence and presence of TGF-β (16 h, 10 ng/ml), which markedly increased Nox4 protein expression (Fig. 5A). Intact WT cells generated a higher basal signal than 3N^−/− cells in the Amplex red/HRP and luminol/HRP assays (Fig. 5B, C). Paralleling the induction of Nox4, TGF-β increased the chemiluminescence signals from WT, but not 3N^−/− cells (Fig. 5B, C). To determine whether measurements could also be made in disrupted cells, NADPH-enhanced luminol/HRP chemiluminescence was assessed in homogenates from the same cells. The signals generated by homogenates from WT and 3N^−/− fibroblasts, however, were similar and unaffected by TGF-β (Fig. 5D). Thus, although HRP-based assays are capable of detecting Nox4-dependent H₂O₂ production in living cells, the signal obtained in homogenates is unrelated to Nox4.

FIG. 5.

Assay comparison in murine dermal fibroblasts. (A) Representative Western blot for the proteins indicated in cells pretreated with transforming growth factor-β (TGF-β) (10 ng/ml, 16 h). Numbers left of the lane indicate the bands of the molecular weight marker. (B) Amplex red assay/HRP assay, n=8, (C, D) Luminol/HRP (100 μmol/L, 1 U/ml) chemiluminescence assay in intact cells (C, with original recording), or cell homogenates (D) of WT or 3N^−/− fibroblasts. N=3–4. *p<0.05 with versus without TGF-β preincubation. ^# p<0.05 WT versus 3N^−/−.

Chemiluminescence Assays in Membrane Fractions Detect CYP and eNOS Activity Rather Than Nox-Derived ROS Production

Among the membrane-anchored enzymes, NOS and cytochrome P450 enzymes produce ROS. To assess whether assay signals can be attributed to these enzymes, HEK293 cells overexpressing eNOS, CYP2C8, Nox4 or Nox5 were studied. In intact HEK293 cells, Nox4 and Nox5 produced a strong signal in the presence of luminol/HRP, whereas in the presence of L012 or lucigenin, which both are thought to detect primarily O₂ ⁻; the signal was strongest in the Nox5-expressing cells (Fig. 6A–C). Neither CYP2C8 nor eNOS overexpression resulted in chemiluminescence under these conditions. This changed after addition of NADPH, so that there was a pronounced increase in the signal in the eNOS-transfected cells as well as in the CYP2C8-transfected cells. In contrast, the chemiluminescence of WT or Nox4-transfected cells only doubled, and the WT and Nox4- and Nox5-overexpressing cells all generated similar signal intensities. Thus, NADPH obscures the signal generated by Nox enzymes (Fig. 6D). A similar picture was obtained using membrane fractions whereby the Nox4- and Nox5-overexpressing cells were indistinguishable from WT cells in NADPH-enhanced luminol/HRP or L012 chemiluminescence, while a fivefold higher signal was detected in eNOS- or CYP2C8-overexpressing cells (Fig. 6E, F). In the lucigenin assay using isolated membranes, there was no clear signal generated by the Nox- or CYP2C8-expressing cells, while eNOS resulted in the highest signal (Fig. 6G).

FIG. 6.

Comparison of chemiluminescence assay in HEK293 cells transiently transfected with the plasmids indicated. N4: Nox4, N5: Nox5, eNOS: eNOS, Cyp: Cytochrome P450-2C8. (A–C): Measurements in intact cells, (D) in intact cells in the presence of NADPH (100 μmol/L), and (E–F) in membrane preparations in the presence of NADPH (E, F 10 μmol/L; G 100 μmol/L). N≥3, *p<0.05 versus WT.

Thus, NADPH-stimulated membrane assays fail to detect Nox activity, but are reasonable indicators of eNOS activity. The latter effect is a potential explanation as to why so many studies detected differences in the lucigenin assay, which they attributed to Nox: the expression of eNOS is redox sensitive and H₂O₂, depending on the concentration applied, can induce or impair eNOS expression. Similarly, stimuli, which can activate Nox2, like TNFα, also activate eNOS and lead to an increase of the inducible NOS. That NOS is a special oxidoreductase is well established due to its diaphorase activity—the ability to oxidize reduced NADH or NADPH. Even in the absence of NADPH, NOS can reduce electron acceptors like formazan (5), and thus, it is conceivable that compounds like lucigenin, which also act as electron acceptors, are directly reduced by NOS in the presence of NADPH. Similarly, cytochrome P450 monooxygenases are oxidoreductases and also, these have been shown to directly reduce lucigenin (2).

In kidney homogenates, loss of Nox enzymes resulted in a non-significantly attenuated NADPH-stimulated lucigenin chemiluminescence without affecting, however, any other assay. Moreover, overexpression of Nox4 in HEK293 cells had no significant effect on NADPH-stimulated lucigenin chemiluminescence. Probably, additional methodologically independent NADPH oxidase assays have to be applied to address this aspect further. However, NADPH interferes with most ROS detectors, even cytochrome C, although chemiluminescence assays are particularly sensitive to this artifact. Thus, ROS-independent measures of Nox activity have to be applied. NADPH consumption in homogenates is high and basically Nox independent (R.P.B., unpublished observation). In leukocyte homogenates, in the classic Nox assay, oxygen consumption is recorded. In other cell types, the impact of Nox enzymes on oxygen consumption was too low to yield a reliable measurement (R.P.B., unpublished observation).

A potential explanation for our failure to detect Nox activity in membrane preparations is that these may lack a constituent of Nox enzymes. For example, it was suggested to add flavin adenine dinucleotide (FAD) to membrane assays (8) and also the traditional cell-free assays of leukocytes, in which oxygen consumption is measured and requires addition of FAD, flavin mononucleotide, and the cytosolic cofactors. Addition of FAD or using crude homogenates rather than isolated membranes, however, also did not yield a Nox-attributable chemiluminescence signal in our hands (data not shown). It also appears possible that Nox enzymes degrade more easily than eNOS or CYP450 enzymes due to their dependency on p22phox or through irreversible oxidation. It has been shown that diamide and thiol oxidation lowers Nox activity (6). In fact, we did not include thiol protection by DTT in our assays, as this does not belong to the standard protocols for Nox measurement in membranes.

In conclusion, we generated and characterized Nox1-Nox2-Nox4 triple knockout mice, which develop normally and have a phenotype with aspects of knockout of the individual Nox homologues. Importantly, NADPH-stimulated chemiluminescence is grossly unaffected by the triple knockout. Thus, the present study casts serious doubt on the validity of Nox membrane assays. Scientists are advised to measure ROS formation in intact tissue rather than boosting their signal with NADPH or using homogenates when addressing ROS production to Noxes.

Notes

Nox^−/− animals and animal procedure

Triple knockout mice (3N^−/−) for Nox1, Nox2, and Nox4 were generated by crossing Nox1 and Nox2 mice, as both genes are on the X chromosome. The Nox1/2 double knockout animals obtained were further crossed with Nox4^−/− animals. All mice were previously backcrossed at least for 10 generations into the C57/Bl6J background. Nox2^−/− mice were obtained from Jackson laboratories, and Nox1^−/− mice were kindly provided by K.H. Krause (University of Geneva). Nox4^−/− mice were generated by the authors. All experiments performed with animals were in accordance with the German animal protection law and were carried out after approval by the local authorities. Animals were housed in groups with free access to chow and water in a specified pathogen-free facility with a 12-h day/12-night cycle. Given the impact of gender on ROS production, only male animals aged 8–16 weeks were used in this study.

Quantitative reverse-transcriptase polymerase chain reaction

Total mRNA from frozen homogenized tissue was isolated with the RNA-Mini-Kit (Bio&Sell) according to the manufacturer's protocol. Random hexamer primers (Promega) and SuperScript III Reverse Transcriptase (Invitrogen) were used for cDNA synthesis. Semiquantitative real-time PCR was performed with the Mx3000P qPCR cycler (Agilent Technologies) using the PCR Eva Green qPCR Mix with ROX (Bio&Sell) with appropriate primers. Relative expression of target genes was normalized to eukaryotic translation elongation factor 2 (EF2), analyzed by the delta–delta-Ct method, and represented as percent of control samples. Primer sequences are listed in Table 3.

Table 3.

Primers

m Elongation factor EEF
Forward	5′-GACATCACCAAGGGTGTGCAG-3′
Reverse	5′-GCGGTCAGCACACTGGCATA-3′
m Nox2
Forward	5′-AGCTATGAGGTGGTGATGTTAGTGG-3′
Reverse	5′-GCGGTCAGCACACTGGCATA-3′
m Nox1
Forward	5′-CCTCCTGACTGTGCCAAAGG-3′
Reverse	5′-ATTTGAACAACAGCACTCACCAA-3′
m Nox3
Forward	5′-CTCACCATAGCTGTGCTGGCTTAC-3′
Reverse	5′-GATGCAGGGCAAGTTAGGATCAGG-3′
m Nox4
Forward	5′-TGTTGGGCCTAGGATTGTGTT-3′
Reverse	5′-AGGGACCTTCTGTGATCCTCG-3′
m Duox1
Forward	5′-ATTCCTCGACCAAAGTGCGT-3′
Reverse	5′-TTCTCCTCTGGGGAGCCTTT-3′
m Duox2
Forward	5′-TCTTCACCATGATGCGGTCC-3′
Reverse	5′-GGAGTCCGGTTGATGAACGA-3′
m Xanthine oxidase
Forward	5′-CCTCCTGACTGTGCCAAAGG-3′
Reverse	5′-ATTTGAACAACAGCACTCACCAA-3′
m Nos1/nNos
Forward	5′-CTCCCGCCTCGGGCAAACAG-3′
Reverse	5′-GTGCACCCCGTTTCCAGCGT-3′
m Nos2/iNos
Forward	5′-GCTCGCTTTGCCACGGACGA-3′
Reverse	5′-AAGGCAGCGGGCACATGCAA-3′
m Nos3/eNos
Forward	5′-CTCACCATAGCTGTGCTGGCTTAC-3′
Reverse	5′-GATGCAGGGCAAGTTAGGATCAGG-3′
m SOD1
Forward	5′-TCGAGCAGAAGGCAAGCGGTGA-3′
Reverse	5′-TCCGCCGGGCCACCATGTTT-3′
m SOD3
Forward	5′-CTTGGGAGAGCCTGACAGGTGCA-3′
Reverse	5′-ACTTTGGCATGCGTGTCGCC-3′
m Catalase
Forward	5′-GCATCGAGCCCAGCCCTGAC-3′
Reverse	5′-GCACATGGGGCCATCACGCT-3′
m CytB
Forward	5′-CAATCGTTCACCTCCTCTTC-3′
Reverse	5′-TCTGGGTCTCCTAGTATGTC-3′
m Cycs
Forward	5′-GACCAAATCTCCACGGTCTG-3′
Reverse	5′-CTGCCCTTTCTCCCTTCTTC-3′

Angiotensin treatment and blood pressure measurements

Animals (n≥7 per group) were treated for 2 weeks with AngII (0.7 mg/kg/day) delivered by a mini-osmotic pump (Alzet). Tail cuff measurements were performed with a six-channel setup (Vistech BP2000) and measurements of 5 days were averaged per mouse.

Organ chamber experiments

Organ chamber experiments were performed using mesenteric vessels in a carbogen-aerated Krebs-Henseleit buffer containing diclofenac (10 μmol/L). The phenylephrine concentration was cumulatively increased (0.03–0.3 μmol/L) to obtain a preconstriction level of ∼80% of the initial KCl constriction. Subsequently, acetylcholine (ACh) was cumulatively added and dose–response curves for the relative constrictor response to phenylephrine and the dilator response to acetylcholine were constructed.

ROS measurements

Radical production was assessed by chemiluminescence with luminol (100 μmol/L)/horseradish peroxidase (HRP at 1 U/ml), L012 (200 μmol/L), or lucigenin (10 μmol/L) in a Berthold six-channel luminometer (LB9505; Berthold). For measurements with cell homogenates or membrane fractions, 20 μg of protein was used for each sample and measurements were initiated by addition of NADPH (10 μmol/L for luminol and L012; 100 μmol/L for lucigenin). Chemiluminescence is expressed as arbitrary units. All measurements were performed in the HEPES-Tyrode (HT) buffer containing in mmol/L the following: 137 NaCl, 2.7 KCl, 0.5 MgCl₂, 1.8 CaCl₂, 5 glucose, 0.36 NaH₂PO₄, and 10 HEPES.

Isolation of skin fibroblasts

Murine dermal fibroblasts were isolated by collagenase type II digestion (1000 U/ml). Cells were cultured in the DMEM/F12 medium containing 10% fetal calf serum and antibiotics.

Amplex red measurements from cells and renal tissue

Cells were cultured in 12-well plates and treated overnight with or without TGF-β (10 ng/ml). H₂O₂-dependent Amplex Red (50 μmol/L; Invitrogen) conversion to resorufin catalyzed by horseradish peroxidase (2 U/ml) was carried out in the HT buffer. Fluorescence was determined in the supernatant at 540/580 nm excitation/emission.

Kidneys were freshly isolated and the HT buffer was added to 1 ml/mg tissue. Organs were then minced with a scalpel, and H₂O₂-derived fluorescence was measured in the supernatant of 100 mg tissue.

Immunoblotting analysis for Nox4

Western blot for Nox4 was performed according to the standard technique. Cells were lysed in a 1% triton Tris-HCl pH7.5 buffer containing protease inhibitors. Samples were not heat denatured and TCEP [tris(2-carboxyethyl)phosphine] was used as a reducing agent. Nox4 was detected by an antibody reported by Anilkumar et al. (1).

Overexpression system using HEK293 cells

HEK293 cells were either stably transfected with a plasmid coding for Nox4 or transiently transfected with 1 μg plasmid DNA encoding for Nox5, eNOS, and CYP4502C8 using Lipofectamine according to the manufacturer's instructions.

Membrane fraction preparation from cells and tissue homogenates

Cells and tissues were homogenized by pottering in the HT buffer containing a protease inhibitor mix (antipain, aprotinin, chymostatin, leupeptin, pepstatin, trypsin inhibitor; AppliChem), okadaic acid, and calyculin A. Cell homogenates were cleared by centrifugation (3000 g, 10 min, 4°C) and membrane fractions were obtained by centrifugation at 100,000 g (1 h, 4°C). The membrane pellet was resuspended in the HT buffer, and the protein concentration was determined by the Bradford assay. Twenty micrograms were used for chemiluminescence measurements.

Statistics

Unless otherwise indicated, data are given as mean±standard error of mean (SEM). n refers to the number of animals, where tissue/animal data were shown, and to the number of independent experiments where cell culture data were reported. Calculations were performed with Prism 5.0 or BiAS.10.12. The latter was also used to test for normal distribution and similarity of variance. In case of multiple testing, Bonferroni correction was applied. For multiple group comparisons, ANOVA followed by post hoc testing was performed. Individual statistics of unpaired samples was performed by the T-test and, if not, normally distributed by the Mann–Whitney test. p-Value of<0.05 was considered significant. Unless otherwise indicated, N indicates the number of individual experiments.

Footnotes

Acknowledgments

We are grateful for the excellent technical assistance of Susanne Schütz. The study was supported by the DFG Excellence Cluster Cardiopulmonary System–ECCPS, SFB 815 (TP A1 to K.S. and TP A16 to I.F.), and SFB 834 (TPA2 to R.P.B. and TP A9N to I.F.), the Faculty of Medicine, Goethe-Universität, Frankfurt am Main, Germany, and the Heinrich und Fritz-Riese-Stiftung to F.R.

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