Caenorhabditis elegans and Human Dual Oxidase 1 (DUOX1) “Peroxidase” Domains

Materials and General Methods

Sf9 cells (Invitrogen) were grown in ExCell 420^TM medium (SAFC Biosciences) supplemented with glutamine (2.7 g/liter). High Five^TM cells were grown in Express Five^TM medium (Invitrogen) supplemented with glutamine (2.7 g/liter) and 10% fetal bovine serum. Both cell lines were kept in suspension at 27 °C (100 rpm) and maintained at densities between 0.5 × 10⁶ and 2 × 10⁶ cells/ml. Trifluoroacetic acid, formic acid, H₂O₂ (30% w/w), ABTS, LPO (bovine), CaCl₂, and 5,5-dimethyl-1,3-cyclohexanedione were purchased from Sigma-Aldrich. T4 DNA ligase and restriction endonucleases were obtained from New England Biolabs. DNA sequencing was performed by Elim Biopharmaceuticals. The entire gene insert was completely sequenced for each plasmid construct. For protein identification, mass spectra of all trypsin digests were obtained on a QSTAR-XL hybrid QqTOF mass spectrometer (Applied Biosystems; supplemental Fig. S1). Spectrophotometric measurements were performed on a Cary 50 Bio UV-visible spectrophotometer (Varian). Liquid chromatography-mass spectrometry (LC-MS) was performed on a Waters Micromass ZQ coupled to a Waters Alliance HPLC system (2695 separations module, Waters 2487 dual λ absorbance detector). For experiments utilizing H₂O₂, the concentrations were determined spectrophotometrically at 240 nm by using the molar extinction coefficient ϵ = 43.6 m⁻¹ cm⁻¹ (20). All of the experiments were performed at room temperature unless otherwise stated.

Plasmid Constructs

The original source of the human duox1_1–593 gene was Quick-clone cDNA from human lung (Clontech). The gene encoding the peroxidase domain of hDUOX1 (residues 1–593) was amplified from plasmid pETDUOXnc³ using primers JLM-Bac-ntDUOX1F and JLM-Bac-DUOX1R (supplemental Table S1) with cloned Pfu polymerase (Stratagene); these primers introduced BamHI and EcoRI sites, respectively, onto the 5′ and 3′ ends of the duox1_1–593 open reading frame. After digestion, the PCR product was ligated into the corresponding sites of the baculovirus expression vector pAcGP67-b (BD Biosciences; pJLM04). To introduce a C-terminal His₆ tag and retain the BamHI and EcoRI restriction sites, duox1_1–593 was amplified from pJLM04 with primers JLM-Bac-ntDUOX1F and JLM-BacCDUOX-R (supplemental Table S1) and subcloned into TOPO vector pCR 2.1 (Invitrogen; pJLM05). This plasmid construct was subsequently digested with BamHI and EcoRI, and the gene was inserted by ligation into pAcGP67-b (pJLM08).

The original source of the C. elegans duox1_1–589 gene was a ProQuest C. elegans cDNA Library (Invitrogen). The gene encoding the peroxidase domain of CeDUOX1 (residues 1–589) was amplified from the cDNA library using primers JLM-BacC-Celegans1F and JLM-BacC-Celegans1R that introduced BamHI and PstI sites onto the 5′ and 3′ ends of the duox1_1–589 open reading frame (supplemental Table S1). The PCR product was subcloned into TOPO vector pCR 2.1 (Invitrogen; pJLM022). This plasmid construct was subsequently digested with BamHI and PstI, and the gene was inserted by ligation into pAcGP67-b (pJLM023).

Generation of Recombinant Baculoviruses

Plasmids pJLM08 and pJLM023 were co-transfected with viral baculogold DNA into Sf9 insect cells, according to the manufacturer's instructions (BD Biosciences). The resulting viruses were plaque-assayed to generate high titer recombinant baculovirus stocks amplified (two rounds of amplification in Sf9 cells) from single viral populations.

Expression and Purification of hDUOX1_1–593

In a typical overexpression, 1 liter of High Five^TM cells (2.0 × 10⁶ cells/ml) supplemented with 250 μm 5-aminolevulinic acid (Fluka) were infected with the hDUOX1_1–593-His₆ viral stock at a multiplicity of infection of 5 for 3 days at 27 °C. After 72 h of infection, cell suspensions were concentrated and purified by nickel affinity chromatography (nickel-nitrilotriacetic acid-agarose; see supplemental data for purification details); the purified protein was stored at −20 °C. The protein stock concentration was determined by Bradford assay in triplicate (21).

Expression and Purification of CeDUOX1_1–589

CeDUOX1_1–589-His₆ was overexpressed and purified as described above for hDUOX1_1–593-His₆; the purified protein was stored at −20 °C, and its concentration was determined by Bradford assay.

Heme Staining Protocol

Detection of heme-protein complexes from SDS-PAGE was achieved by incubation with a solution containing tetramethylbenzidine and the addition of H₂O₂ for color development, as previously described (22). Experimental details are included in the supplemental data.

HPLC Analysis of Covalent Heme Binding

Samples were analyzed by direct injection onto a 150 × 4.6-mm C4 reversed phase column (Vydac 214MS) on an Agilent 1200 series HPLC instrument. The protein was eluted with a linear gradient of 20–50% acetonitrile in water (0.1% trifluoroacetic acid) over 30 min (1 ml/min) with detection at 215 and 400 nm. For covalent heme binding as a function of H₂O₂, CeDUOX1_1–589 (26 μm) was incubated with H₂O₂ (26–520 μm) for 5 min at 25 °C, pH 8.0. Excess H₂O₂ was then consumed by incubation with catalase (5 units) for 5 min. The resulting samples were chromatographed as described above, with the proportion of covalently bound heme determined by the percentage of the total 400-nm integration attributed to the peak(s) that co-eluted with the 215-nm absorbing protein peak (27–30 min).

CeDUOX1_1–589 Dihydroxyheme Isolation

CeDUOX1_1–589 (2 mg/ml, 70 μl) was digested by incubation for 1 h with trypsin (1 mg/ml, 2 μl) at room temperature, followed by the addition of proteinase K (Fermentas; 20 mg/ml, 3 μl); the digest was continued overnight at 37 °C. As a control, hemin was purchased from Sigma-Aldrich; a stock solution was created by dissolving 1–2 mg in 200 μl of 1 n NaOH, followed by the addition of 9.8 ml of 20 mm phosphate buffer, 0.4 m NaCl, pH 8.0); 2.5 μl stock was diluted into 25 μl of buffer for analysis. Digested protein (40 μl) and porphyrin (25 μl) separations were achieved on a Vydac 214MS reversed phase C4 column (150 × 4.6 mm) using water containing 0.1% trifluoroacetic acid (solvent A) and acetonitrile containing 0.09% trifluoroacetic acid (solvent B) on an Agilent 1200 series HPLC instrument. All of the samples were eluted at a flow rate of 1 ml/min, with a 3-min isocratic wash of 15% solvent B, followed by a linear gradient of 15–43% solvent B over 31 min, monitored at 400 nm. The same gradient and column were utilized for analysis of the observed peaks by LC-MS.

Reduction and CO Binding Studies of CeDUOX1_1–589

The ferrous and CO bound absorbance spectra for CeDUOX1_1–589 (6 μm) were collected in 100 mm phosphate buffer, pH 7.0; 800 μl of total sample volume. To achieve reduction, sodium dithionite (8 μl, 100 mm) was added to an argon flushed enzyme sample in a sealed cuvette. After reduction, the enzyme was incubated with CO for 30 s, and the spectra were collected until saturation was demonstrated.

Cyanide Binding to CeDUOX1_1–589

The cyanide (CN⁻) bound absorbance spectrum for CeDUOX1_1–589 (6 μm) was collected via titration with KCN in 100 mm phosphate buffer, pH 7.0. A stock solution of KCN (10 mm) was utilized to titrate from 12.5 to 225 μm until enzyme saturation was demonstrated. A titration curve was established by plotting the change in absorbance (ΔA), from subtraction of difference spectra maxima (423 nm) and minima (402 nm), against CN⁻ concentration; a dissociation constant was determined from this plot at half the maximum ΔA observed.

ABTS Activity Assay Comparison of the DUOX1 Homologs

Peroxidase activity was measured using 500 nm ABTS incubated with 500 nm LPO, hDUOX1_1–593, or CeDUOX1_1–589 in 50 mm phosphate buffer (pH 7.0, treated with Chelex-100 resin (Bio-Rad)). The reactions were initiated by the addition of H₂O₂ (50 μm). Each reaction was monitored by recording the absorbance intensity at 414 nm as a function of time. The molar extinction coefficient for ABTS at 414 nm is ϵ = 36,800 m⁻¹ cm⁻¹ (23).

Peroxidase Activity Assay(s) with Tyrosine Ethyl Ester

Enzyme peroxidase activity was measured using tyrosine ethyl ester as a substrate (1 mm) in phosphate-buffered saline (pH 7.4, treated with Chelex-100 resin (Bio-Rad)). LPO, CeDUOX1_1–589, and hDUOX1_1–593 (1 μm) were assayed; each reaction was initiated with the addition of H₂O₂ (100 μm). Initial reaction kinetics of tyrosine cross-linking were monitored using a Fluorolog 3 Spectrofluorometer (Horiba Jobin Yvon), with excitation at 295 nm and emission monitored at 414 nm using 1-nm excitation and emission slits at room temperature. After 1 h of reaction time, LPO and CeDUOX1_1–589 assays were further analyzed by direct injection onto a 50 × 2.1-mm C18 reversed phase column (Waters XTerraMS) on an Agilent 1200 series HPLC instrument. The assays were characterized utilizing an isocratic gradient of 9% acetonitrile in water (0.1% formic acid), followed by a linear gradient of 9–10% acetonitrile in water (0.1% formic acid) over 10 min (0.2 ml/min) with detection at 280 nm and fluorescence detection by excitation at 295 nm and emission at 414 nm. The same gradient and column were utilized for analysis of the observed peaks by LC-MS.

Superoxide Dismutase Activity Assay(s)

SOD activity was measured by the percentage reaction inhibition rate of enzyme (SOD, hDUOX1_1–593, or CeDUOX1_1–589) with WST-1 substrate (a water soluble tetrazolium dye) and xanthine oxidase using a SOD assay kit (Fluka; 19160) according to the manufacturer's instructions. Each end point assay was monitored by absorbance at 450 nm (the absorbance wavelength for the colored product of WST-1 reaction with superoxide) after 20 min of reaction time at 37 °C. All of the enzyme stocks were made in 50 mm HEPES buffer, pH 7.0, 5% glycerol (to allow for CaCl₂ addition); each assay was performed in triplicate.

Bromide Oxidation Activity Assay(s)

5,5-Dimethyl-1,3-cyclohexanedione (1 mm, 50 μl), NaBr (2 m, 50 μl), and enzyme (50 nm LPO, 50 μl; 5 μm CeDUOX1_1–589 or hDUOX1_1–593, 50 μl) were mixed in a quartz cuvette. To this solution was added 850 μl of H₂O₂ (0.52 mm in 50 mm citrate buffer, pH 4.4). The resulting solution was mixed, and the absorbance at 292 nm was recorded as a function of time. The initial velocity was calculated from the change in absorbance between 10 and 30 s; the molar extinction coefficient of 5,5-dimethyl-1,3-cyclohexanedione at 292 nm is ϵ = 20,000 m⁻¹ cm⁻¹ (24).

Design and Expression of Soluble DUOX1 Peroxidase Domain Constructs

hDUOX1 and CeDUOX1 are large membrane-bound proteins (∼180 kDa) predicted to contain an extracellular N-terminal domain tethered by a transmembrane helix to a C-terminal region with six TM domains, two cytosolic EF hand motifs, and an NADPH oxidase domain (Fig. 1A). This N-terminal extracellular domain is homologous to classical peroxidases such as MPO and TPO (43% sequence similarity) (17); however, neither the hDUOX1 nor the CeDUOX1 peroxidase domain contains the full complement of heme-binding and catalytic residues, as shown in Fig. 1B. This partial sequence alignment with the classical peroxidases demonstrates that both DUOX homologs lack the catalytic distal histidine residue (Ser¹⁰⁸, hDUOX1; and Tyr¹⁰⁵, CeDUOX1) and the aspartate residue that in MPO and LPO covalently binds to the heme co-factor through an ester linkage (Leu¹⁰⁷, hDUOX1; and Ala¹⁰⁴, CeDUOX1) (25, 26). hDUOX1 lacks additional critical peroxidase residues: the glutamate that forms the second covalent bond to the heme (Arg²⁴¹) and the proximal histidine residue (Ser³³¹). Both of these amino acids are conserved in the CeDUOX1 homolog. Despite sequence similarity, the only critical residue of the heme-binding region conserved among mammalian peroxidases and the DUOX proteins is the catalytic arginine residue (Arg⁴⁰⁵, hMPO; Arg²³⁸, hDUOX1; and Arg²³⁵ CeDUOX1). Soluble constructs are required to elucidate the function of this region for both DUOX homologs.

Initially, the full-length human and C. elegans DUOX1 proteins were truncated using the TMHMM Server version 2.0 transmembrane helix algorithm to identify the primary sequences likely to compose their entire extracellular peroxidase domains (27). Based on this analysis, we designed constructs of hDUOX1 (residues 1–593) and CeDUOX1 (residues 1–589) for expression in Escherichia coli. N- and C-terminal His₆-tagged and glutathione S-transferase-tagged constructs were created but attempts at expression under various conditions did not yield soluble protein (data not shown). As a result, our expression efforts shifted to a baculovirus system. This technique often yields recombinant proteins with the proper fold, oligomerization, and post-translational modifications, thus generating a protein structurally and functionally similar to its native counterpart. Both truncated DUOX1 homologs were inserted into baculovirus expression vector pAcGP67-b, which allows for secretion of the recombinant protein (resulting in the N-terminal addition of five amino acids to the expressed protein as part of the secretion signal sequence; supplemental Fig. S1). N- and C-terminal His₆-tagged constructs of the DUOX1 proteins were co-transfected with baculovirus DNA into Sf9 cells to obtain high titer viral stocks for infection (see “Experimental Procedures” for details).

hDUOX11_-593 and CeDUOX1_1–589 were stably overexpressed (via infection of High Five^TM cells) as C-terminally His₆-tagged proteins (no protein was obtained from the N-terminally tagged constructs) and purified by nickel-nitrilotriacetic acid-agarose affinity chromatography (Fig. 2A); each DUOX1 protein was eluted with ∼100 mm imidazole. Yields were reproducible at levels of ∼0.7 mg/liter for both hDUOX1_1–593 and CeDUOX1_1–589.

Visual inspection of each protein stock solution, supported by absorbance spectra (Fig. 2B), demonstrated that heme was associated with CeDUOX1_1–589 and not with the isolated hDUOX1_1–593. Further investigation by heme titration failed to demonstrate specific heme binding by hDUOX1_1–593. Absorbance spectroscopy of CeDUOX1_1–589 identified a Soret maximum at 408 nm with a 408/280-nm ratio of 0.55. This Soret is significantly blue-shifted from that seen for LPO, EPO, or TPO (412–413 nm), which have two heme-covalent bonds, or MPO (428 nm), which has three such bonds (28). This absorbance shift suggests that the number of covalent bonds or other structural aspects of the heme pocket are responsible for the Soret difference for CeDUOX1_1–589. The nature of the heme co-factor interaction with CeDUOX1_1–589 was initially evaluated by tetramethylbenzidine staining of denaturing polyacrylamide gels. Under these separating conditions, the presence of heme was confirmed by its coincident migration with the CeDUOX1_1–589 peroxidase domain (Fig. 2A), establishing that at least a portion of the heme co-factor is indeed covalently bound to the recombinant protein.

To examine the extent of heme attachment to CeDUOX1_1–589, an HPLC system was utilized under acidic conditions that favor dissociation of noncovalently bound heme from the apoprotein (29). As shown in Fig. 2C, only 60% of the prosthetic heme co-elutes with the 215-nm absorbing protein peak; this is consistent with results for LPO expressed in baculovirus (29). Investigation of this recombinant LPO covalent heme binding has shown the process to be autocatalytic, as opposed to requiring an enzymatic system such as that required for covalent heme attachment in cytochrome c (30–32). To determine whether a similar autocatalytic process is responsible for heme oxidation and covalent incorporation in CeDUOX1_1–589, the protein was preincubated with varying concentrations of H₂O₂, and the extent of covalent heme binding was evaluated by HPLC. Fig. 2D highlights the results of H₂O₂ treatment, clearly demonstrating that the covalent binding reaction is autocatalytic, because the fraction of protein-bound heme increases as a function of H₂O₂ concentration. A maximum for the amount of heme incorporation (81%) was achieved with 5 equivalents of H₂O₂; the Soret maximum shifted from 408 to 410 nm at this percentage of incorporation after H₂O₂ treatment.

With covalent heme binding established as a major interaction between co-factor and enzyme, the nature of this linkage in CeDUOX1_1–589 was examined. Mammalian peroxidases form covalent linkages to the 1- and 5-methyl groups of the heme porphyrin through conserved aspartate and glutamate residues (Fig. 1B). The result of enzymatic digestion and LC-MS analysis of the classic peroxidases with two covalent bonds (LPO, TPO, and EPO) is a 1,5-dihydroxyheme derivative. Mutation studies of heme proteins (horseradish peroxidase F41E, horseradish peroxidase S73E, LPO D225E, and LPO E375D) to investigate the autocatalytic nature and sequence of covalent bond formation have also led to non-native linkages at the 3- and 8-methyl positions (33, 34). Because of single covalent bond formation in each mutant enzyme, once digested (trypsin/Pronase or proteinase K), each liberated heme co-factor was found as a monohydroxyheme (1-OH, 3-OH, 5-OH, or 8-OH) derivative. For further investigation of the nature of covalent heme binding, CeDUOX1_1–589 was digested and analyzed by HPLC and LC-MS to determine whether a monohydroxy- or dihydroxyheme derivative would result. A heme derivative was isolated (m/z 650, t = 13.2 min), and determined, both by retention time and mass, to be dihydroxyheme, suggestive of two covalent protein-heme linkages (Fig. 3). This covalent bond character, although consistent with that of the known mammalian peroxidases, was highly unexpected in the case of CeDUOX1_1–589 because the aspartate residue required for covalent heme binding is not conserved in this enzyme. It is clear that a different carboxyl residue or a different amino acid altogether is involved in formation of the second covalent bond to the heme.

Structural Stability of the DUOX1 Proteins

Circular dichroism measurements were performed on hDUOX1_1–593 and CeDUOX1_1–589 to confirm that the protein constructs are properly folded (supplemental Fig. S2). The far-UV spectra (195–250 nm) for both homologs support an α-helical structure, with prominent troughs at 208 and ∼220 nm, characteristic of LPO and other mammalian peroxidases (35–37). For further insight into the stability of hDUOX1_1–593 and CeDUOX1_1–589, Trp fluorescence emission spectra were collected for each upon excitation at 292 nm, exhibiting broad emission bands with λ_max at 341 and 348 nm, respectively (supplemental Fig. S2). Both maxima suggest a folded protein structure resulting in protection of Trp residues from solvent exposure. The DUOX1 homologs contain a large number of Trp residues (hDUOX1_1–593, 19 Trp; CeDUOX1_1–589, 11 Trp); both the disparity in the number of Trp residues and quenching by the heme chromophore likely contribute to the difference in emission intensity. The observed wavelength maxima shift to ∼351 nm upon thermal unfolding (80 °C; supplemental Fig. S2) and decrease in fluorescence intensity, consistent with quenching caused by solvent exposure upon unfolding. Chemical denaturation of both proteins using 7 m guanidine hydrochloride resulted in a further red shift of the λ_max to ∼360 nm; this result suggests that the DUOX peroxidase domain(s) retain some tertiary structure upon thermal denaturation, consistent with previous studies on LPO (37).

Small Ligand Binding to CeDUOX1_1–589

CO and CN⁻ molecules bind tightly to mammalian peroxidases (38, 39). These interactions result in steady-state spectral changes from association with the heme iron in the distal binding pocket. To determine whether the binding pocket of CeDUOX1_1–589 will allow such interactions, steady-state absorbance spectra were collected and are shown in Fig. 4. Ferric CeDUOX1_1–589 exhibits a Soret band maximum at 408 nm with bands at 533 and 623 nm in the visible region (Fig. 4, A and B, and supplemental Table S2). Upon reduction with sodium dithionite, CeDUOX1_1–589 undergoes a red shift to 425 nm, with Q band maxima at 533 and 557 nm. Exposure of the reduced protein to CO leads to the formation of a complex with the ferrous heme iron, demonstrated by a blue shift of the Soret band to 419 nm, with Q bands at 539 and 563 nm.

Cyanide adduct formation, in contrast to CO, results in a stable complex with the ferric heme iron. Consistent with other mammalian peroxidases, upon formation of the Fe⁺³-CN⁻ adduct, the Soret band of CeDUOX1_1–589 exhibited a red shift (Fig. 4C). In the case of LPO (consistent with the other classical mammalian peroxidases; supplemental Table S2), upon complexation with CN⁻, the Soret maxima experiences a large shift of 20 nm, from 412 to 432 nm; CeDUOX1_1–589 exhibits a smaller shift of 7 nm to a Soret maxima of 415 nm at saturation (40). The reduced magnitude of the shift may be related to a difference in the properties of the heme binding pocket and/or the presence of a tyrosyl residue in place of the distal histidine. The hydrogen bonding interactions of the ligated cyanide are likely to differ significantly with a distal tyrosine rather than an imidazole residue. To further characterize the cyanide binding event, a titration was used to monitor the UV-visible spectral shift upon the addition of potassium cyanide (12.5–225 μm) to 6 μm CeDUOX1_1–589. This study established a dissociation constant for cyanide binding of 25 μm; this value is consistent with dissociation constants found for mammalian peroxidases LPO (42 μm) and TPO (10 μm) at neutral pH (39, 41).

Activity Studies for the DUOX Peroxidase Domain(s)

Because of the absence of several residues that are conserved in peroxidases, including the distal histidine residue, the actual catalytic function of the hDUOX1 and CeDUOX1 peroxidase domains remains uncertain. It has been shown through site-specific mutagenesis that the distal catalytic histidine in peroxidases is crucial to their activity. When mutated in such enzymes as horseradish peroxidase, cytochrome c peroxidase, and dehaloperoxidase, a significant loss of activity is observed (42–46). This fact has led researchers to postulate that the N-terminal region of DUOX1 may not act as a classical peroxidase but rather as a SOD (8, 12). In an effort to clarify the function of these domains, now available as purified and structurally sound proteins, we examined each homolog with regard to its peroxidase and SOD activity.

Catalytic activity studies were first conducted with ABTS, a substrate commonly used to evaluate peroxidase activity. In contrast to a standard peroxidase, LPO ((3.3 ± 1.0) × 10³ mol oxidized per min/mol of enzyme) hDUOX1_1–593 exhibited no peroxidase activity, whereas CeDUOX1_1–589 had a modest activity of 14.1 ± 0.8 mol oxidized per min/mol of enzyme.

Tyrosine cross-linking has been established as the biological function of the DUOX enzymes in thyroid hormone biosynthesis for humans and in cuticle formation for C. elegans. Each domain was therefore assayed via fluorescence for dityrosine formation with tyrosine ethyl ester as a substrate (14). Similar results were obtained with this more natural substrate, i.e. no activity with hDUOX1_1–593 and modest activity with CeDUOX1_1–589 (Fig. 5A). To confirm that any increase in fluorescence observed was due directly to the formation of dityrosine, HPLC analysis of the reactions was performed to isolate the dityrosine product (Fig. 5B); separation isolated the fluorescent product formed (t = 8 min) from unreacted tyrosine ethyl ester (t = 2.8 min). LC-MS identified the fluorescent product as dityrosine (m/z 417).

As an alternative to peroxidase activity, the DUOX1 domains may function as SODs, converting two molecules of superoxide to O₂ and H₂O₂. The hDUOX1_1–593 peroxidase domain, which appears unable to bind heme, could conceivably bind a metal ion necessary for SOD activity. To determine whether any metal ions have co-purified with hDUOX1_1–593, the enzyme was submitted to metal analysis by inductively coupled argon plasma optical emission spectroscopy (Garratt Callahan Analytical Laboratory). This analysis showed that the only protein-associated metal ion was calcium, consistent with conservation of a calcium-binding site among the classical peroxidases and the DUOX family (47). Calcium does not belong to the known set of metals utilized by SODs to support their function (the mammalian enzymes utilize copper, zinc, and/or manganese (48–51)); therefore, this result suggests that hDUOX1_1–593 may also not function as a SOD. To further investigate, in the unlikely event calcium may be utilized for enzyme function, hDUOX1_1–593 and CeDUOX1_1–589 were evaluated for their ability to react with superoxide by a colorimetric inhibition assay utilizing WST-1 as a substrate (52, 53). Authentic superoxide dismutase was assayed as a control over a profile of concentrations (Fig. 5C). hDUOX1_1–593 and CeDUOX1_1–589 show less than 10% inhibition over concentrations from 10–120 ng/ml, over which SOD increases in inhibition strength to 70%. Superoxide dismutase demonstrates a high turnover rate even at low concentrations, which may not be shared by the DUOX enzymes. To ascertain whether any activity may be obtained at higher concentrations, each enzyme was assayed under the same conditions at 1 μm concentration, both with and without the addition of 100 μm CaCl₂. As shown in Fig. 5D, even at this higher concentration, no significant dismutase activity is exhibited by the DUOX1 homologs, in either the presence or the absence of additional calcium.

In sum, this collective activity data demonstrates hDUOX1_1–593 to be inactive as a peroxidase or superoxide dismutase, and CeDUOX1_1–589 to be inactive as a SOD but exhibiting low level peroxidase activity. This low activity level is realistic for CeDUOX1_1–589 in vivo and the DUOX family as a whole to prevent cellular oxidative stress and to function in a controlled manner toward polymerization of tyrosines and/or generation of signaling molecules. In contrast, mammalian peroxidases require high peroxidase activity levels, herein demonstrated by the high level of ABTS turnover and tyrosine ethyl ester cross-linking observed with LPO. These levels are necessary for in vivo generation of antimicrobial agents, i.e. hypohalides, from bromide, chloride, and thiocyanate oxidation. Specifically, LPO has the characterized ability to rapidly oxidize bromide at acidic pH (24, 54). Therefore, to support the theory that DUOX enzymes support controlled reactivity and should not oxidize halides, LPO was utilized as a control with both DUOX1 homologs to assay bromide oxidation. This study confirmed that neither DUOX peroxidase domain could oxidize bromide (LPO, (3.5 ± 0.7) × 10³ mol oxidized per min/mol enzyme; CeDUOX1_1–589 and hDUOX1_1–593, no significant oxidation). Although a low activity level appears native to CeDUOX1_1–589, the lack of an observed hDUOX1_1–593 function suggests that specific protein interactions may play a role in heme binding or in enhancing the activity of the protein.