Báo cáo khoa học: Thioredoxin reductase from the malaria mosquito Anopheles gambiae Comparisons with the orthologous enzymes of Plasmodium falciparum and the human host pdf

Thioredoxin reductase from the malaria mosquito Anopheles gambiae Comparisons with the orthologous enzymes of Plasmodium falciparum and the human host Holger Bauer 1 , Stephan Gromer 1 , Andrea Urbani 2 , Martina Schno¨ lzer 2 , R. Heiner Schirmer 1 and Hans-Michael Mu¨ ller 3 1 Biochemie Zentrum, Universita ¨ t Heidelberg, Heidelberg, Germany; 2 Deutsches Krebsforschungszentrum, Heidelberg, Germany; 3 European Molecular Biology Laboratory, Heidelberg, Germany The mosquito, Anopheles gambiae, is an important vector of Plasmodium falciparum malaria. Full genome analysis revealed that, as in Drosophila melanogaster,theenzyme glutathione reductase is absent in A. gambiae and func- tionally substituted by the thioredoxin system. The key enzyme of this system is thioredoxin reductase-1, a homo- dimeric FAD-containing protein of 55.3 kDa per subunit, which catalyses the reaction NADPH + H + + thio- redoxin disulfide fi NADP + + thioredoxin dithiol. The A. gambiae trxr gene is located on chromosome X as a single copy; it represents three splice variants coding for two cytosolic and one mitochondrial variant. The predominant isoform, A. gambiae thioredoxin reductase-1, was recomb- inantly expressed in Escherichia coli and functionally com- pared with the wild-type enzyme isolated in a final yield of 1.4 UÆml )1 of packed insect cells. In redox titrations, the substrate A. gambiae thioredoxin-1 (K m ¼ 8.5 l M , k cat ¼ 15.4 s )1 at pH 7.4 and 25 °C) was unable to oxidize NADPH-reduced A. gambiae thioredoxin reductase-1 to the fully oxidized state. This indicates that, in contrast to other disulfide reductases, A. gambiae thioredoxin reduc- tase-1 oscillates during catalysis between the four-electron reduced state and a two-electron reduced state. The thio- redoxin reductases of the malaria system were compared. A. gambiae thioredoxin reductase-1 shares 52% and 45% sequence identity with its orthologues from humans and P. falciparum, respectively. A major difference among the three enzymes is the structure of the C-terminal redox cen- tre, reflected in the varying resistance of catalytic inter- mediates to autoxidation. The relevant sequences of this centre are Thr–Cys–Cys–SerOH in A. gambiae thioredoxin reductase, Gly–Cys–selenocysteine–GlyOH in human thio- redoxin reductase, and Cys–X–X–X–X–Cys–GlyOH in the P. falciparum enzyme. These differences offer an interesting approach to the design of species-specific inhibitors. Notably, A. gambiae thioredoxin reductase-1 is not a selenoenzyme but instead contains a highly unusual redox- active Cys–Cys sequence. Keywords: Anopheles gambiae; Drosophila melanogaster; Diptera; insect redox metabolism; Plasmodium falciparum. The mosquito Anopheles gambiae is of importance as a vector of tropical malaria caused by the protozoan organism Plasmodium falciparum. The genome of A. gambiae is the second insect genome – after the distantly related model organism Drosophila melanogaster [1,2] – that has been completely sequenced [3]. Annotation of the nucleotide sequences allows access to the genetic background of a disease-transmitting dipteran insect and, furthermore, offers the opportunity of comparative sequence analyses from single genes up to genomic organization [4]. Another aspect is highlighted in this report: only on the basis of the full genome sequence does it become possible to exclude the presence of a given protein function in all cells and all developmental stages of an organism. A case in point is the absence of the enzyme glutathione reductase (GR) in Diptera [5]. Our focus is the redox metabolism of insects [6]. Being present at millimolar concentrations, the tripeptide gluta- thione (GSH) is the most abundant antioxidative thiol compound in most cell compartments. Its redox state determines the intracellular redox environment [7]. Thus, GSH is the major redox buffer compound and essential for the detoxification of free radicals and xenobiotics [8,9]. In the majority of pro- and eukaryotic organisms, the oxidized form of glutathione (glutathione disulfide, GSSG) is reduced to the mono-thiol form (GSH) by the Correspondence to R. H. Schirmer, Biochemie Zentrum, Im Neuen- heimer Feld 504, D-69120 Heidelberg, Germany. Fax: + 49 6221 545586, Tel.: + 49 6221 544165, E-mail: heiner.schirmer@gmx.de or H M. Mu ¨ ller, European Molecular Biology Laboratory, Meyerhofstr. 1, D-69117 Heidelberg, Germany. Fax: + 49 6221 387306, Tel.: + 49 6221 387440, E-mail: hmueller@embl-heidelberg.de Abbreviations:EH 2 , enzyme in a two-electron reduced state; EH 4 , enzyme in a four-electron reduced state; E ox ,enzymeinanoxidized state; EST, expressed sequence tag; GR, glutathione reductase; GSH/GSSG, reduced/oxidized glutathione; Sec, selenocysteine; Trx, thioredoxin; TrxR, thioredoxin reductase. (Received 14 July 2003, revised 29 August 2003, accepted 1 September 2003) Eur. J. Biochem. 270, 4272–4281 (2003) Ó FEBS 2003 doi:10.1046/j.1432-1033.2003.03812.x NADPH-dependent flavoenzyme GR, which catalyses the following reaction: NADPH + GSSG + H + fi NADP + + 2GSH [10–12]. D. melanogaster cells exhibit a high 2[GSH]/ [GSSG] ratio but have been shown to lack a typical GR [5]. As reported here, this is also true for A. gambiae.An important candidate able to functionally substitute for GR is the thioredoxin (Trx) system [13], which compri- ses NADPH, thioredoxin reductase(s) (TrxR) and Trxs [14,15]. Trxs are small, ubiquitous thiol proteins with a relative molecularmassof 12 kDa and a redox active cysteine pair represented in a WCGPC sequence motif. Therefore, they cycle between a disulfide (TrxS 2 ) and a dithiol [Trx(SH) 2 ]form. Trxs were first described as electron-donating substrates for ribonucleotide reductase [16], but they cleave disulfide bonds in a number of other proteins equally well. Thus, Trxs take part in the redox control of numerous processes such as protein folding, signalling and transcription [14,17–19]. Trx reduction is catalysed by the flavin-dependent oxido- reductase TrxR, as follows: NADPH + TrxS 2 +H + fi NADP + +Trx(SH) 2 . TrxRs belong to a disulfide reduc- tase superfamily that includes enzymes such as GR, trypanothione reductase, lipoamide dehydrogenase, and mercuric ion reductase. These homodimeric proteins are structurally, as well as mechanistically, closely related [20]. Evolution has produced two classes of TrxRs: small TrxRs (found in bacteria, plants and fungi) and large TrxRs (present in other eukaryotes) [21]. In contrast to GRs and low molecular weight TrxRs, large TrxRs possess an additional redox centre located in the C-terminal extension which is necessary for the interaction with the substrate Trx. In mammalian enzymes this redox centre is represented by a neighboring cysteine–seleno- cysteine pair [22] and in the TrxR of P. falciparum it is a cysteine pair separated by a spacer sequence of four amino acids [23]. D. melanogaster TrxR was described as the first member of a third type of large TrxRs. It is characterized by two adjacent cysteines preceding the C-terminus [5,24]. High molecular mass TrxRs exhibit a rather broad substrate spectrum that includes a number of natural and also artificial disulfide compounds, such as 5,5¢-dithiobis(2- nitrobenzoate). Glutathione disulfide, however, is not a substrate. In the fruit fly it was shown that GSSG reduction can occur in a dithiol–disulfide exchange reaction with reduced Trx [5]. At physiological concentrations of GSSG and Trx, this system allows GSSG fluxes of > 100 l M Æmin )1 [25]. In this report we introduce the TrxR of the malaria mosquito A. gambiae. The protein could be isolated from whole insects and from cultured insect cells. With the progress of the Anopheles genome project it was possible to identify the complete sequence of the gene and its organi- zation. We cloned, recombinantly expressed and character- ized the enzyme. Our data support the assumption that the substitution of the Trx system for GR, as well as the mechanistic particularities of the TrxR, are a common principle in dipteran insects. In the context of the malarial system, this implies that the TrxRs of insect vector, parasite, and human host differ in their cellular roles as well as their enzyme mechanisms. Experimental procedures D. melanogaster Trx-2, A. gambiae Trx-1 and P. falciparum Trx-1 were prepared and purified as previously described [25,26]. PCR chemicals and restriction enzymes were purchased from MBI Fermentas and Applied Biosystems, precast polyacrylamide gels from Bio-Rad and molecular mass standards from Amersham Pharmacia Biotech. Anti- biotics, substrates for enzyme assays, and other chemicals were from BioMol, Fluka, or Sigma. All compounds were of the highest available purity. Purification of authentic A. gambiae TrxR from insect cells A. gambiae cells (cell line 4a-2s4) were cultured in Schneider’s medium and harvested as described previ- ously [27]. A 0.3-mL volume of lysis buffer (50 m M Tris/ HCl, 3 m M EDTA, 2.5 m M phenylmethanesulfonyl fluoride, 5 l M pepstatin and 5 l M cystatin, pH 7.6) was added per mL of frozen cell pellet. The pellets were thawed at 37 °C in a water bath, fresh phenyl- methanesulfonyl fluoride (ad 500 l M ) was added, and the cells were disintegrated by ultrasound. All subsequent steps were carried out at 4 °C. The suspension was centrifuged for 1 h at 26 000 g. The supernatant was set aside, and the pellet was resuspended in lysis buffer and centrifuged as described above. The combined supernatants were mixed with two volumes of TE buffer (50 m M Tris/HCl, 1m M EDTA, pH 7.6) and slowly loaded onto a cooled 2¢,5¢- ADP–Sepharose column (1.5 mL per 10 mL of cell pellet) equilibrated with TE buffer. The column was washed with two column volumes of TE buffer, 1.5 column volumes of 100 m M KCl in TE buffer, three column volumes of 1 : 3 diluted TE buffer, 1.5 column volumes of 1 m M NADH in 1 : 3 diluted TE buffer, and two column volumes of TE buffer. TrxR activity was then eluted with 2 m M NADP + in TE buffer. Fractions containing significant amounts of activity were pooled, concentrated and washed with TE buffer in a 10-kDa Amicon concentrator. Purity was analyzed by SDS/PAGE and silver staining. GR activity was not detected in the crude extract or in any column fraction, even when the column was washed with 1 M KCl and 1 m M NADPH in TE buffer. The combined and concentrated washing solutions were stored at )80 °Casa source of other NADPH-dependent enzymes of A. gambiae. TrxR assay TrxR assays were conducted at 25 °C with a reaction volume of 1 mL consisting of buffer T (100 m M potassium phos- phate, 2 m M EDTA, pH 7.4) and 100 l M NADPH. For determination of the K m values of Trxs, Trx concentrations were varied from 3 to 50 l M [25]. The assays were started by the addition of 10 milliunits of A. gambiae TrxR-1 (1 unit ¼ 1 lmol of NADPH consumption per minute under substrate saturation), and the Trx-dependent NADPH oxidation was followed spectrophotometrically at 340 nm applying an e-value of 6.22 m M )1 Æcm )1 . K m and V max values were obtained by applying the Michaelis–Menten equation. GR activity was determined using established protocols [28,29]. Ó FEBS 2003 Thioredoxin reductase from Anopheles gambiae (Eur. J. Biochem. 270) 4273 Trx-dependent GSSG-reduction assay The Trx-dependent GSSG-reduction assay was conducted as described previously [25,26]. The mixture contained 100 l M NADPH and A. gambiae Trx-1 in concentrations from 5 to 50 l M . The assay was started by the addition of 1UofA. gambiae TrxR-1, and NADPH oxidation was followed at 340 nm. After reduction of Trx was complete, 1m M GSSG was added, and GSSG reduction was followed by further NADPH consumption. The composition of the assay mixture guarantees that > 98% of Trx is present in the reduced form. L -Dehydroascorbate reduction assay for TrxR L -Dehydroascorbate (dimer; Sigma-Aldrich) was studied as a substrate in the range of 50 l M to 5 m M in TrxR assay mixture containing 200 l M NADPH and 300 n M A. gamb- iae TrxR subunits; NADPH consumption was determined spectroscopically at 340 nm. Purified human TrxR served as a positive control. Protein determination The protein concentration in crude fractions was estimated assuming an absorption of 10 at 280 nm for a 1% solution. For determining the exact concentration of TrxR, flavin absorbance was measured at 450 nm after denaturation of the enzyme sample by 0.1% SDS and heating to 80 °C; the FAD released by this procedure has an e-value of 11.3 m M )1 Æcm )1 . Sequence studies on authentic A. gambiae TrxR A10lg sample of purified authentic A. gambiae TrxR was applied per lane in reducing SDS/PAGE. After electro- phoresis, one lane was Coomassie-stained and the putative TrxR band was subjected to tryptic digestion (see below). Another lane was blotted [in 50 m M sodium borate, 20% (v/v) methanol, pH 9.0, at 150 mA] overnight onto a poly(vinylidene difluoride) membrane and stained with 0.1% amido-black in 2% acetic acid. This yielded a protein band of 58 kDa. An additional minor band of  62 kDa appeared when phenylmethanesulfonyl fluoride was present during all steps of the protein isolation. Both bands were excised and subjected to Edman degradation. Selenium analysis Ten microgram samples of purified authentic A. gambiae TrxR were subjected to atomic absorption spectrometry for selenium determination (Dr Muntean, Labor Seelig, Karls- ruhe, Germany). A negative control (TE buffer) and a positive control (10 lg of human TrxR in TE buffer) were analysed in parallel. Cloning of A. gambiae TrxR-1 The A. gambiae trxr-1 gene was PCR cloned from genomic DNA as well as from the cDNA of adult insects. In the case of the amplification from genomic DNA, the bases coding for the first five amino acids, located on the first exon, were included in the primer. For the cloning of genomic DNA, the following primers were applied: forward, 5¢-CGCAG GATCCGCGCCATTGAATCAGGAAAACTATGAGT ACGATCTGGTG-3¢ (containing a BamHI restriction site); and reverse, 5¢-TCCTAAGCTTCTAGCTGCAG CAGGTCGCCGGCGTCG-3¢ (containing a HindIII restriction site). Dimethylsulfoxide [5% (v/v)] was added to the PCR mixture to improve amplification of the GC-rich gene. PCR conditions were as follows: 94 °C for 60 s; 35 cycles of 30 s at 94 °C, 30 s at 68 °Cand90 sat72°C; then 10 min at 72 °C. The PCR fragment was cloned into the expression vector, pQE-60 (Qiagen), and Escherichia coli NovaBlue cells (Novagen) were transformed with the plasmid. The insert was verified by sequencing. Protein expression Transformed E. coli NovaBlue cells were grown overnight at 34 °Cin2· YT medium containing 50 lgÆmL )1 carbeni- cillin. A. gambiae TrxR-1 expression was then induced with 0.3 m M isopropyl-b- D -thiogalactopyranoside for 4 h at 34 °C. After centrifugation (3000 g,10min,4°C), cells were resuspended in 25 m M TE buffer and treated with lysozyme (0.2 mgÆmL )1 ) and DNase (0.02 mgÆmL )1 )for 20 min at room temperature. Phenylmethanesulfonyl fluo- ride (100 l M ), pepstatin (3 l M )andcystatin(80n M )were added as protease inhibitors and the cells were disintegrated by ultrasound. The homogenate was centrifuged at 38 000 g for 30 min at 4 °C and the supernatant was applied to a 2¢,5¢-ADP–Sepharose column equilibrated with 50 m M TE buffer. The column was washed with five volumes of 25 m M TE buffer and one volume of 50 m M TE buffer. A. gambiae TrxR-1 was eluted by 2 m M NADP + in 50 m M TE, the final yield being  40 mgÆL )1 of protein culture. SDS/ PAGE, using a 10% gel, showed a single band of the expected size, the purity being > 95% as judged by silver staining. MS of tryptic peptides Protein bands were excised from SDS/PAGE, and their cysteine residues were reduced and alkylated with iodoacet- amide. The samples were then digested with porcine trypsin (Promega) in 40 m M ammonium bicarbonate at 37 °Cfor 6–8 h. The reaction was stopped by freezing. Tryptic peptides were extracted by ZipTip C18 reverse phase material (Millipore), chromatographed, and taken up in a saturated solution of a-cyano-4-hydroxycinnamonic acid in 50% (v/v) acetonitrile/water. MALDI mass spectra were recorded in the positive ion mode with delayed extraction on a Reflex IV time-of-flight instrument equipped with an MTP multiprobe inlet and a 337-nm nitrogen laser. Mass spectra were obtained by averaging 50–200 individual laser shots. Calibration of the spectra was internally performed by a two-point linear fit using the autolysis products of trypsin at m/z 842.50 and m/z 2211.10. The peptide masses were screened against the NCBInr database using the peptide search algorithm MASCOT (Matrix Science). Fragments generated by postsource decay experiments were analysed using the database search algorithm MS - TAG (http://prospector.ucsf.edu). 4274 H. Bauer et al. (Eur. J. Biochem. 270) Ó FEBS 2003 Results When this project started, only one Trx (A. gambiae Trx-1) had been described as a part of the Trx-based redox metabolism in the malaria mosquito A. gambiae [25]. Complementary studies on the fruit fly, D. melanogaster [5], suggested investigating, in detail, the redox homeostasis in a disease-transmitting insect. The characterization of A. gambiae TrxR allows the comparison of three different mechanisms of Trx reduction in the P. falciparum malaria system, i.e. that in the parasite, the human host, and the insect vector. Isolation of A. gambiae TrxR from insect cells From 8 mL of pelleted A. gambiae cells, 19 U of A. gamb- iae TrxR activity was extracted. Approximately 60% TrxR was recovered from the 2¢,5¢-ADP–Sepharose affinity column. No GR activity was detected either in the dialyzed crude extract or in any fraction eluted from the column. SDS/PAGE analysis of the TrxR fraction revealed two major bands representing apparent molecular masses of 62 kDa and 58 kDa (inset Fig. 1A). We observed a decrease in intensity of the heavy band when the cell extract was ageing. As this process could be prevented by repeated addition of phenylmethanesulfonyl fluoride, it was conclu- ded that proteolytic cleavage of the 62 kDa protein yielded a product co-migrating with the 58 kDa band. Sequence analysis by Edman degradation and mass spectral analysis Edman degradation of the 58 kDa band resulted in the N-terminal sequence of 17 residues given in Fig. 1B; the 62 kDa protein resisted Edman degradation. For further sequence information on A. gambiae TrxR, we conducted mass spectral analyses of the two bands from the SDS/PAGE gel. To achieve this, the proteins were subjected to tryptic digestion. The patterns of high-yield peptides (Fig. 1B) were indistinguishable, which suggests that the two electrophoretic bands represent splice isoforms of A. gambiae TrxR (see below). Furthermore, sequence comparison of the peptides with D. melanogaster TrxR-1 confirmed that we had indeed isolated a homologue of the Drosophila enzyme [5]. Lack of detectable selenium A particular point of interest was whether TrxR from Anopheles is a selenoprotein (like its relatives from humans and other mammals) [30,31] or whether A. gambiae TrxR is a selenium-free ÔDrosophila-typeÕ enzyme. In mammalian TrxR, the C-terminal redox centre is formed by a Cys– selenocysteine (Sec) pair [22,32]. No significant selenium – less than 0.01 mol per mol of A. gambiae TrxR subunit compared with 0.94 mol per mol of human TrxR subunit – was determined by atomic absorption spectrometry. The absence of a catalytic Sec residue in A. gambiae TrxR was corroborated by the finding that L -dehydroascorbate – which is a substrate of the selenium-dependent TrxRs of mammals [33] – was not a substrate at concentrations up to 5m M . Genomic organization of the A. gambiae TrxR gene The genome sequence analysis of A. gambiae, reported previously [3], enabled us to address the genomic organiza- tion of the gene (Fig. 2). A. gambiae trxr occurs in three different splicing variants (AJ459821, AJ549084, AJ549085) as a single-copy gene on chromosome X. In contrast to D. melanogaster TrxR-1 – where the coding sequence is interrupted by three introns – the A. gambiae TrxR coding sequences were found to be separated by a single intron corresponding to the proximal intron in D. melanogaster TrxR-1 (Fig. 2). The identification of three types of expressed sequence tags (ESTs), varying in the sequence of the first exon only, suggests that three alternative transcription start sites are operative (exons 1–3 of A. gambiae trxr in Fig. 2). Exon 1 is Fig. 1. Characterization of authentic Anopheles gambiae thioredoxin reductase-1 (TrxR-1) by physicochemical analyses. (A) The absorption spectra of 6.6 l M TrxR-1 in the oxidized form E ox (dashed curve) and in the four-electron reduced form (EH 4 ) (solid curve), which was obtained by adding 33 l M NADPH to the E ox sample. In the EH 4 sample, the absorption at wavelengths below 400 nm is largely a result of excess NADPH. The inset shows A. gambiae TrxR species in a silver-stained gel after SDS/PAGE. In lane 1, the two variants of A. gambiae TrxR isolated from cultured insect cells can be distin- guished; lane 2 shows recombinant A. gambiae TrxR-1, and the outer lane marker proteins. (B) The results of peptide analyses. The DNA- deduced sequence of A. gambiae TrxR-1 is shown in standard script. Tryptic peptides of authentic enzyme that were identified by MS are underlined and marked in bold. These peptides were found in both protein bands shown on lane 1 in the inset. The N-terminal sequence (17 residues in bold italics) was identified by Edman degradation of the protein isolated from the major band of the SDS/PAGE gel. The minor band of 62 kDa resisted Edman degradation. Ó FEBS 2003 Thioredoxin reductase from Anopheles gambiae (Eur. J. Biochem. 270) 4275 represented in the NCBI database by 14 ESTs that overlap with exon 4. Joining of exon 1 (encoding the five amino acids MAPLN) with exon 4 (encoding QENYEY and further 491 amino acids), leads to a protein of 502 residues which is the orthologue of D. melanogaster TrxR-1 (Figs 1B and 3). This protein is introduced here as A. gambiae TrxR-1 (CAD30858). Two alternative 5¢ exons–exon2andexon3–were identified, each represented by a single EST. The N-terminal segment contributed by exon 2 (MATAVLARPARS LINVVQCVRL IRTQATVMFA) shows the properties of a mitochondrial targeting sequence containing the predicted cleavage site between IRT and QAT. The last four amino acids (VMFA) are not encoded by EST BM603316, but the correct overlap with exon 4 was proven by sequencing a PCR product amplified with an oligonucleotide pair specific for exon 1 and exon 4 (data not shown). The deduced N-terminal sequence of the putative mitochondrial enzyme is thus QATVMFA|KENEY, the change from Q to K in position 5 resulting from splicing. Exon 3 occurs in EST BM583435, which extends into exon 4. The resulting N-terminal sequence – MAAATAAE| QENYEY – probably represents a second cytosolic species of TrxR. As judged by the number of EST sequences and by Edman degradation of enzyme isolated from insect cells, we can state that A. gambiae TrxR-1 is the major isoform in vivo. Similarly to the fruit fly, no GR-like sequence could be identified in the mosquito genome [34]. This is consistent with the absence of detectable genuine GR activity in Anopheles cell extracts. Cloning and characterization of A. gambiae TrxR-1 The A. gambiae trxr-1 gene was cloned and recombinantly expressed in E. coli. In SDS/PAGE, this protein co-migrates with wild-type A. gambiae TrxR-1 – isolated from cultured Anopheles cells or whole insects – at a position representing a molecular mass of 58 kDa (Fig. 1A). The discrepancy between this value and the molecular mass of 54.5 kDa deduced from the amino acid sequence has also been observed in other disulfide reductases; they all show a subunit molecular mass (deduced by SDS/PAGE) that is overestimated by 7–10%. The molecular basis for this electrophoretic behaviour is unknown [35]. The identity between authentic and recombinant A. gambiae Trx-1 is supported by the concurrence of deduced and experimentally determined amino acid sequences (Fig. 1B). Thus, TrxR-1 from A. gambiae con- tains 502 amino acids per subunit; the calculated molecular mass is 54.5 kDa per subunit for the apoprotein and 2 · 55.3 kDa for the FAD-containing homodimeric holo- enzyme. The protein shares 69% sequence identity with its orthologue from D. melanogaster (Fig. 3). Both insect enzymes have sequence elements that are typical for large TrxRs, including the flavin-near redox-active Cys–Val– Asn–Val–Gly–Cys motif, as well as a C-terminally located redox centre (Fig. 3). In the sequence of A. gambiae TrxR-1 and D. melanogaster TrxR-1 a sequentially adjacent cys- teine pair (Cys500¢ and Cys501¢) is present. Thus, the two insect Trxs known, to date, are typical members of the large TrxR enzyme class characterized by an additional redox centre. However, in contrast to mammalian TrxRs or TrxR from P. falciparum, this part of the active site is structurally distinct in the insect enzymes. Indeed a redox centre formed by two sequential Cys residues is highly unusual [24]. A. gambiae TrxR-1 as a Trx-reducing enzyme TrxR-1 was tested with Trxs from A. gambiae (A. gambiae Trx-1), D. melanogaster (D. melanogaster Trx-2), and P. falciparum (P. falciparum Trx-1) as oxidizing substrates (Table 1). The catalytic efficiency of A. gambiae TrxR-1 is Fig. 2. Genomic organization of the Anopheles gambiae thioredoxin reductase-1 (TrxR-1)geneincomparisonwiththeDrosophila melano- gaster TrxR gene. In both insects the trxr locusislocatedonchro- mosome X. Numbered boxes represent exons within the gene. Coding regions are shown in black, untranslated sequences are shaded in grey. Scale bar divisions are in kilobases. A. gambiae trxr occurs in three possible splice isoforms (AJ459821, AJ549084, AJ549085) that differ in the first exon (exon 1, 2 or 3) joined to exon 4. The orthologous Drosophila locus, shown below, is similarly organized, except that the sequence corresponding to exon 4 of A. gambiae trxr is interrupted by two short introns, which results in exons 4–6. Exons 1, 2 or 3 joined to exons 4–6 yield transcripts CG2151-RA, CG2151-RB and CG2151- RC, respectively. In both insect species, exon 1 encodes the N-terminal sequence of an abundant cytosolic, exon 2 of a mitochondrial, and exon 3 of a minor cytosolic TrxR form. Table 1. Kinetic parameters of Anopheles gambiae thioredoxin reductase-1 (TrxR-1) with different thioredoxins and 5,5¢-dithiobis(2-nitrobenzoic acid) (DTNB). All values were determined in 100 m M potassium phosphate buffer, 2 m M EDTA, pH 7.4. As expected, A. gambiae Trx-1 was the best substrate of A. gambiae TrxR-1, but the k cat /K m value was only marginally better than for thioredoxin-2 (Trx-2) from Drosophila melanogaster. Plasmodium falciparum thioredoxin-1 (Trx-1) showed the highest k cat value, but the K m was significantly lower than for the insect thioredoxins. DTNB was included as an artificial disulfide substrate which is reduced by most high molecular mass thioredoxin reductases. Substrate K m (l M ) V max (UÆmg )1 ) k cat (s )1 ) k cat /K m ( M )1 Æs )1 ) A. gambiae Trx-1 8.5 ± 1.5 16.9 ± 1.6 15.4 ± 1.5 1.81 · 10 6 D. melanogaster Trx-2 9.0 ± 1.0 15.7 ± 1.3 14.3 ± 1.2 1.58 · 10 6 P. falciparum Trx-1 33 ± 5 17.2 ± 1.2 15.7 ± 1.1 0.48 · 10 6 DTNB 700 ± 200 6.0 ± 0.7 5.5 ± 0.6 7.9 · 10 3 4276 H. Bauer et al. (Eur. J. Biochem. 270) Ó FEBS 2003 comparable with the value previously determined for D. melanogaster TrxR-1 [5,25]. Expectedly, with a K m value of 8.5 l M and a k cat of 15.4 s )1 , A. gambiae Trx-1 is the best substrate of the enzyme but D. melanogaster Trx-2, i.e. the orthologue of A. gambiae Trx-1 in Drosophila,isanalmost equally good substrate. The K m value for the reducing substrate NADPH was found to be 5.0 l M . By comparison with other TrxRs [21,24,36], we can delineate the pathways of electrons in A. gambiae TrxR-1 during catalysis (Figs 3 and 4). The reducing equivalents flow from the nicotinamide of NADPH via the flavin and the pair Cys57/Cys62 to the redox centre Cys500’/Cys501’ of the other subunit, and hence to the disulfide of the substrate Trx. The thiolate of Cys62 in A. gambiae TrxR-1 forms a charge transfer complex with the reoxidized flavin during catalysis [20,24,37]. This charge transfer gives rise to the absorption band at  530 nm (Fig. 1A) and thus to the orange/red colour of stable catalytic intermediates that contain both oxidized flavin and Cys62 as a thiolate. Unexpectedly, freshly prepared A. gambiae TrxR was found to be orange/red, which indicated the presence of reduced Cys62. Auto-oxidation of the A. gambiae TrxR-1 preparations was very slow and proceeded over a range of hours to days, finally producing the typical yellow colour of oxidized enzyme E ox (Fig. 1A). The freshly isolated enzyme also resisted oxidation by its native substrate, A. gambiae Trx. In contrast, most disulfide reductases, when present in reduced forms, can be easily oxidized by their native substrates [20]. Consequently, we conducted redox titration experiments on A. gambiae TrxR-1, starting out with the oxidized form, E ox , and monitoring the absorbance Fig. 3. Multiple sequence alignment ( CLUSTAL W ) of high molecular mass thioredoxin reductases (TrxR). The search was conducted (NCBI accession numbers in parentheses) with the enzymes from Anopheles gambiae (AgTrxR-1, CAD30858), Drosophila melanogaster (DmTrxR-1, AAG25639), humans (hTrxR, AAB35418), and from the malaria parasite Plasmodium falciparum (PfTrxR-1, CAA60574). The enzyme of Anopheles shares 69%, 52%, and 45% sequence identity with the TrxRs of D. melanogaster, humans, and P. falciparum, respectively. The sequences of the redox- active centres are shaded in grey; U (residue 498) in the human enzyme represents selenocysteine (Sec). Ó FEBS 2003 Thioredoxin reductase from Anopheles gambiae (Eur. J. Biochem. 270) 4277 at 530 nm. As shown in Fig. 1A, the absorption coefficient at 530 nm was 0.4 m M )1 Æcm )1 for the oxidized enzyme, E ox , 1.6 m M )1 Æcm )1 after addition of one equivalent NADPH, leading to the two-electron reduced enzyme species EH 2 , and 3.0 m M )1 Æcm )1 for the enzyme reduced with two or more equivalents NADPH, giving rise to the four-electron reduced enzyme species EH 4 . After reoxidation with 100 l M A. gambiae Trx-1, the e-value fell to 1.6 m M )1 Æcm )1 , indi- cative of a two-electron reduced enzyme species (EH 2 ). In contrast, reoxidation of EH 4 with 125 l M potassium ferricyanide, as described previously [16,24], led to the E ox species with an e-value of 0.4 m M )1 Æcm )1 at 530 nm. These data confirm that the native substrate does not reoxidize the enzyme to E ox but only to the EH 2 state where the redox-active Cys residues 57, 62, 500’, and 501’ are present partly as thiols so that the thiolate of Cys62 can still form a charge transfer complex with flavin. For catalysis, this implies that the very first catalytic cycle is primed by two NADPH molecules, which results in the four-electron reduced state. Oxidation with TrxS 2 then leads to the two- electron reduced state, where the two disulfide bridges are partially reduced, i.e.: priming reaction, E ox + 2NADPH + 2H + fi EH 4 + 2NADP + ; catalytic cycle, EH 4 +TrxS 2 fi EH 2 +Trx(SH) 2 ;EH 2 +NADPH+ H + fi EH 4 +NADP + .TrxS 2 +NADPH+H + fi Trx(SH) 2 +NADP + . This balance reaction of the cata- lytic cycle, of course, represents the net reaction catalyzed by TrxR. Discussion The isolation and characterization of A. gambiae TrxR contributes to the understanding of the redox metabolism in Diptera. The principles of redox homeostasis that were tentatively postulated for the fruit fly can be extended to a disease-transmitting insect. In short, a GR is absent, although GSH is a key compound of the redox networks also in insects [38]. The nonenzymatic reduction of GSSG by reduced Trx is probably a major pathway for GSH reduction in these organisms [5]. Thus, TrxR indirectly substitutes for the function of GR. As described previously, A. gambiae Trx-1 is a highly expressed protein in vivo [25]. The efficiency of GSSG reduction by A. gambiae Trx-1 is similar to its orthologue (Trx-2) in D. melanogaster and probably sufficient to maintain physiological needs. In the Anopheles mosquito, one TrxR gene is present which occurs in three splice variants. Alternative use of first exons was previously reported for mammalian and Droso- phila TrxR genes [39]. In D. melanogaster, three alternative transcripts have been identified: CG2151-RA is the ortho- logue of A. gambiae TrxR-1; CG2151-RB corresponds to a mitochondrial TrxR form; and CG2151-RC encodes a second cytosolic TrxR. Transcripts coding for the latter form are rare, as only two ESTs corresponding to CG2151- RC have been identified (compared with more than 80 CG2151-RA ESTs). Thus, the trxr loci in Anopheles and in Drosophila are structurally organized in a similar way: there are three alternative first exons, coding probably for a major cytosolic, a mitochondrial, and a minor cytosolic form (Fig. 2). Unlike in A. gambiae, a second TrxR gene, trxr-2,was identified in the genome of D. melanogaster.However,a D. melanogaster TrxR-1 null mutant leads to death, at the latest during the second larval instar [40], and both cytosolic and mitochondrial TrxR-1 forms have been shown to be necessary for survival [41]. Thus, the putative activity of TrxR-2 is not sufficient to compensate for the lack of either the cytosolic or the mitochondrial TrxR-1. The A. gambiae TrxR preparation from insect cells results in two enzyme species that can be distinguished by SDS/PAGE (Fig. 1). The predominant band represents the cytosolic variant A. gambiae TrxR-1, and the 62 kDa band is possibly the mitochondrial precursor variant. This assumption is supported by the size of the protein and by the observation that it is stabilized by protease inhibitors. A. gambiae TrxR-1 shares 69% sequence identity with D. melanogaster TrxR-1, including the important redox- active Cys–Cys motif on the C-terminal extension (Figs 3 and 4). For the Drosophila enzyme it was shown that both cysteines are essential for the interaction with the natural substrate Trx [5,24]. In the case of rat TrxR, which is a selenoprotein with a Cys–Sec–sequence instead, the Sec fi Cys exchange results in a mutant with less than 1% catalytic activity when compared with the wild-type enzyme [30,42]. The main difference between the insect enzymes and the TrxR from rat concerns the amino acid residues adjacent to the cysteines. In mammalian TrxRs, including the human orthologue, we find a Gly– Cys–Cys–Gly sequence, whereas in A. gambiae TrxR-1, it is Thr–Cys–Cys–Ser and in D. melanogaster TrxR-1 it is Ser– Cys–Cys–Ser. There is evidence that the hydroxyl functions Fig. 4. Sketch of homodimeric Anopheles gambiae in the four-electron reduced (EH 4 )form.The dimer interface is shown as a diagonal line with a black circle at the centre. This circle represents the molecular two-fold symmetry axis. The sketch shows the EH 4 form where all four redox-active cysteines are present in the reduced form. The thiolate 62 forms a charge transfer complex with oxidized flavin, and Cys501¢ is ready to attack the disulfide bond of the substrate thioredoxin disulfide. By analogy with other disulfide reductases, the most probable scenario leading from oxidized enzyme (E ox )to EH 4 is as follows. When NADPH binds to one subunit (upper right), its reducing equivalents flow via the flavin to the disulfide Cys62/Cys57. The resulting dithiol is reoxidized by exchange with the disulfide bridge between residues 500¢ and 501¢ of the other subunit. Subsequently, binding and oxidation of a second NADPH molecule leads to re-reduction of the disulfide between Cys57 and Cys62. 4278 H. Bauer et al. (Eur. J. Biochem. 270) Ó FEBS 2003 of the flanking amino acid residues are crucial for catalysis (H. S. Gromer et al., unpublished data). It is interesting to note that despite the evolutionary distance of 250 million years [34] between the fruit fly and the mosquito, not only the basic principles of redox metabolism and the genomic organization of the TrxR gene, but also the mechanistic peculiarities of these orthologous enzymes, have remained highly conserved. With a k cat of 15–22 s )1 for Trx [5,24], the insect TrxRs exhibit a somewhat lower turnover number than their mammalian relatives ( 35–45 s )1 ) [30,43], but they have the advantage of being independent of the rare trace element selenium. This evolutionary adaptation is plausible because TrxR is apparently the mainstay enzyme of the antioxidant metabolism in insects, whereas mammals have a second, GR-based system. Redox processes also represent an interesting aspect of parasite–host interaction. For human malaria it is well known that disturbance of the antioxidative metabolism results in an inhibition of parasitic growth in erythrocytes. The most prominent example is glucose-6-phosphate-dehy- drogenase (Glc6PDH) deficiency, an inherited disease also known as favism [44,45]. A major effect of Glc6PDH deficiency is an impaired NADPH production, which affects the ability of the erythrocyte to resist oxidative stress. The effects of Glc6PDH deficiency can be imitated by GR-inhibitors such as carmustine [46] or isoalloxazines [47]. The current interpretation of Glc6PDH deficiency, as a condition protecting from severe malaria, is based on the observation that the anion channel protein of the erythro- cyte membrane undergoes oxidative changes when ring- stage parasites are present in the red blood cell. This oxidized band-3 protein is recognized by a specific antibody that initiates immunologic processes to eliminate the parasitized cell [48,49]. With respect to the insect vector A. gambiae,ithasalso been shown that nitrosative and oxidative stress imposed by NO and peroxynitrite play a dominating role in the host’s defence against the parasite [50,51]. The insect cells, in turn, have to protect themselves against these reactive agents. When Anopheles cells are challenged by oxidative stress, transcription of numerous genes that are associated with the Trx system are induced, prominently among them the TrxR gene [52]. Interestingly, a similar response occurs after exposing the cells to bacterial peptidoglycan. Trx system-related genes are also transcribed in the salivary glands of A. gambiae. It is assumed that the corresponding proteins are especially important for pro- tecting the glands from heme-driven free radical attack [53]. Thus, redox processes play a major role in host– parasite interactions, not only in human blood but also inside the insect vector. The differences between the enzyme systems involved in antioxidative metabolism offer an interesting novel target for the development of insect-specific TrxR inhibitors. The potential of TrxR as a target for novel insecticides is supported by the fact that TrxR-1 knockouts in Drosophila are lethal in early embryonic stages [40]. Based on the genomic data and comparative genome analyses, it can be reasonably assumed that this is also true for Anopheles.The development of novel insecticides is an important approach in the fight against malaria which is becoming more and more complicated, not least as a result of the occurrence of insecticide resistance [54]. In this context it should be noted that inhibitors of the Trx system have toxic effects themselves but, in addition, they sensitize organisms for other toxic agents [13,55,56]. Consequently, inhibitors of A. gambiae TrxR-1 are expected to protect other insecti- cides from the development of resistance. Acknowledgements We are indebted to Dr Stefan M. Kanzok for the early studies on the Anopheles thioredoxin system and for his help in database searches. Our work was supported by the Deutsche Forschungsgemeinschaft (Grants B2 and C1 of SFB 544 ÔControl of Tropical Infectious DiseasesÕ to R.H.S. and H.M.M., respectively, as well as grant GR2028/1-1 to S.G.) and by the Fonds der Chemischen Industrie (Grant 161576) to R.H.S. References 1. Adams, M.D., Celniker, S.E., Holt, R.A., Evans, C.A., Gocayne, J.D., Amanatides, P.G., Scherer, S.E., Li, P.W., Hoskins, R.A., Galle, R.F., George, R.A., Lewis, S.E., Richards, S., Ashburner, M., Henderson, S.N., Sutton, G.G., Wortman, J.R., Yandell, M.D., Zhang, Q., Chen, L.X. et al. (2000)Thegenomesequenceof Drosophila melanogaster. Science 287, 2185–2195. 2. Myers, E.W., Sutton, G.G., Delcher, A.L., Dew, I.M., Fasulo, D.P., Flanigan, M.J., Kravitz, S.A., Mobarry, C.M., Reinert, K.H., Remington, K.A., Anson, E.L., Bolanos, R.A., Chou, H.H.,Jordan,C.M.,Halpern,A.L.,Lonardi,S.,Beasley,E.M., Brandon, R.C., Chen, L., Dunn, P.J. et al. (2000) A whole-genome assembly of Drosophila. Science 287, 2196–2204. 3. Holt, R.A., Subramanian, G.M., Halpern, A., Sutton, G.G., Charlab, R., Nusskern, D.R., Wincker, P., Clark, A.G., Ribeiro, J.M., Wides, R., Salzberg, S.L., Loftus, B., Yandell, M., Majoros, W.H., Rusch, D.B., Lai, Z., Kraft, C.L., Abril, J.F., Anthouard, V., Arensburger, P. et al. (2002) The genome sequence of the malaria mosquito Anopheles gambiae. Science 298, 129–149. 4. Kanzok, S.M. & Zheng, L. (2003) The mosquito genome – a turning point? Trends Parasitol. 19, 329–331. 5. Kanzok, S.M., Fechner, A., Bauer, H., Ulschmid, J.K., Mu ¨ ller, H.M., Botella-Munoz, J., Schneuwly, S., Schirmer, R.H. & Becker, K. (2001) Substitution of the thioredoxin system for glutathione reductase in Drosophila melanogaster. Science 291, 643–646. 6. Hemingway, J. & Ranson, H. (2000) Insecticide resistance in insect vectors of human disease. Annu. Rev. Entomol. 45, 371–391. 7. Schafer, F.Q. & Buettner, G.R. (2001) Redox environment of the cell as viewed through the redox state of the glutathione disulfide/ glutathione couple. Free Radic. Biol. Med. 30, 1191–1212. 8. Flohe ´ , L. (2003) Lest I forget thee, glutathione. Biol. Chem. 384, 503–687. 9. Dolphin, D., Poulson, R. & Avramovic, O., (eds). (1989) Coen- zymes and Cofactors – Glutathione,Vol.III.PartAandB.John Wiley & Sons, New York. 10. Schirmer, R.H., Krauth-Siegel, R.L. & Schulz, G.E. (1989) Glu- tathione reductase. In Coenzymes and Cofactors – Glutathione (Dolphin,D.,Poulson,R.&Avramovic,O.,eds),Vol.III,pp. 553–596. John Wiley & Sons, New York. 11. Schirmer, R.H., Bauer, H. & Becker, K. (2002) Glutathione reductase. In Wiley Encyclopedia of Molecular Medicine (Creighton T.E., ed.), Vol. 5, pp. 1471–1475. John Wiley & Sons, New York. 12. Sarma, G.N., Savvides, S.N., Becker, K., Schirmer, M., Schirmer, R.H. & Karplus, P.A. (2003) Glutathione reductase of the malarial parasite Plasmodium falciparum: crystal structure and inhibitor development. J. Mol. Biol. 328, 893–907. Ó FEBS 2003 Thioredoxin reductase from Anopheles gambiae (Eur. J. Biochem. 270) 4279 13. Gromer,S.,Urig,S.&Becker,K.(2004)Thethioredoxinsystem– from science to clinic. Med. Res. Rev. in press. 14. Arne ´ r, E.S. & Holmgren, A. (2000) Physiological functions of thioredoxin and thioredoxin reductase. Eur. J. Biochem. 267, 6102–6109. 15. Holmgren, A. (1985) Thioredoxin. Annu. Rev. Biochem. 54, 237–271. 16. Holmgren, A. (1989) Thioredoxin and glutaredoxin systems. J. Biol. Chem. 264, 13963–13966. 17. Holmgren, A. (1984) Enzymatic reduction-oxidation of protein disulfides by thioredoxin. Methods Enzymol. 107, 295–300. 18. Schenk, H., Klein, M., Erdbrugger, W., Dro ¨ ge,W.&Schulze- Osthoff, K. (1994) Distinct effects of thioredoxin and antioxidants on the activation of transcription factors NF-kappa B and AP-1. Proc.NatlAcad.Sci.USA91, 1672–1676. 19. Follmann,H.&Ha ¨ berlein, I. (1995) Thioredoxins: universal, yet specific thiol-disulfide redox cofactors. Biofactors 5, 147–156. 20. Williams, C.H. Jr (1992) Lipoamide dehydrogenase, glutathione reductase, thioredoxin reductase, and mercuric ion reductase – a family of flavoenzyme transhydrogenases. In Chemistry and Biochemistry of Flavoenzymes (Mu ¨ ller F., ed.), Vol. 3, pp. 121–211. CRC Press, Inc., Boca Raton, FL. 21. Williams, C.H., Arscott, L.D., Mu ¨ ller, S., Lennon, B.W., Ludwig, M.L.,Wang,P.F.,Veine,D.M.,Becker,K.&Schirmer,R.H. (2000) Thioredoxin reductase: two modes of catalysis have evolved. Eur. J. Biochem. 267, 6110–6117. 22. Zhong, L., Arne ´ r, E.S. & Holmgren, A. (2000) Structure and mechanism of mammalian thioredoxin reductase: the active site is a redox-active selenolthiol/selenenylsulfide formed from the con- served cysteine-selenocysteine sequence. Proc. Natl Acad. Sci. USA 97, 5854–5859. 23. Gilberger, T.W., Bergmann, B., Walter, R.D. & Mu ¨ ller, S. (1998) The role of the C-terminus for catalysis of the large thioredoxin reductase from Plasmodium falciparum. FEBS Lett. 425, 407–410. 24. Bauer, H., Massey, V., Arscott, L.D., Schirmer, R.H., Ballou, D.A. & Williams, C.H. Jr (2003) The mechanism of high M r thioredoxin reductase from Drosophila melanogaster. J. Biol. Chem. 278, 33020–33028. 25. Bauer, H., Kanzok, S.M. & Schirmer, R.H. (2002) Thioredoxin-2 but not thioredoxin-1 is a substrate of thioredoxin peroxidase-1 from Drosophila melanogaster: isolation and characterization of a second thioredoxin in D. melanogaster and evidence for distinct biological functions of Trx-1 and Trx-2. J. Biol. Chem. 277, 17457– 17463. 26. Kanzok, S.M., Schirmer, R.H., Tu ¨ rbachova, I., Iozef, R. & Becker, K. (2000) The thioredoxin system of the malaria parasite Plasmodium falciparum: glutathione reduction revisited. J. Biol. Chem. 275, 40180–40186. 27. Mu ¨ ller, H.M., Dimopoulos, G., Blass, C. & Kafatos, F.C. (1999) A hemocyte-like cell line established from the malaria vector Anopheles gambiae expresses six prophenoloxidase genes. J. Biol. Chem. 274, 11727–11735. 28. Fa ¨ rber, P.M., Arscott, L.D., Williams, C.H. Jr, Becker, K. & Schirmer, R.H. (1998) Recombinant Plasmodium falciparum glutathione reductase is inhibited by the antimalarial dye methy- lene blue. FEBS Lett. 422, 311–314. 29. Worthington, D.J. & Rosemeyer, M.A. (1976) Glutathione reductase from human erythrocytes: catalytic properties and aggregation. Eur. J. Biochem. 67, 231–238. 30. Zhong, L. & Holmgren, A. (2000) Essential role of selenium in the catalytic activities of mammalian thioredoxin reductase revealed by characterization of recombinant enzymes with selenocysteine mutations. J. Biol. Chem. 275, 18121–18128. 31. Gladyshev, V.N. & Hatfield, D.L. (1999) Selenocysteine-contain- ing proteins in mammals. J. Biomed. Sci. 6, 151–160. 32. Gladyshev, V.N., Jeang, K.T. & Stadtman, T.C. (1996) Seleno- cysteine, identified as the penultimate C-terminal residue in human T-cell thioredoxin reductase, corresponds to TGA in the human placental gene. Proc. Natl Acad. Sci. USA 93, 6146–6151. 33. May, J.M., Mendiratta, S., Hill, K.E. & Burk, R.F. (1997) Reduction of dehydroascorbate to ascorbate by the selenoenzyme thioredoxin reductase. J. Biol. Chem. 272, 22607–22610. 34. Zdobnov, E.M., von Mering, C., Letunic, I., Torrents, D., Suyama, M., Copley, R.R., Christophides, G.K., Thomasova, D., Holt, R.A., Subramanian, G.M., Mueller, H.M., Dimopoulos, G., Law, J.H., Wells, M.A., Birney, E., Charlab, R., Halpern, A.L.,Kokoza,E.,Kraft,C.L.,Lai,Z.et al. (2002) Comparative genome and proteome analysis of Anopheles gambiae and Drosophila melanogaster. Science 298, 149–159. 35. Krauth-Siegel, R.L., Mu ¨ ller, J.G., Lottspeich, F. & Schirmer, R.H. (1996) Glutathione reductase and glutamate dehydrogenase of Plasmodium falciparum, the causative agent of tropical malaria. Eur. J. Biochem. 235, 345–350. 36. Sandalova, T., Zhong, L., Lindqvist, Y., Holmgren, A. & Schneider, G. (2001) Three-dimensional structure of a mammalian thioredoxin reductase: implications for mechanism and evolution of a selenocysteine-dependent enzyme. Proc. Natl Acad. Sci. USA 98, 9533–9538. 37. Rietveld, P., Arscott, L.D., Berry, A., Scrutton, N.S., Deonarain, M.P., Perham, R.N. & Williams, C.H. Jr (1994) Reductive and oxidative half-reactions of glutathione reductase from Escherichia coli. Biochemistry 33, 13888–13895. 38. Sohal, R.S., Arnold, L. & Orr, W.C. (1990) Effect of age on superoxide dismutase, catalase, glutathione reductase, inorganic peroxides, TBA-reactive material, GSH/GSSG, NADPH/ NADP + and NADH/NAD + in Drosophila melanogaster. Mech. Ageing Dev. 56, 223–235. 39. Sun, Q.A., Zappacosta, F., Factor, V.M., Wirth, P.J., Hatfield, D.L. & Gladyshev, V.N. (2001) Heterogeneity within animal thioredoxin reductases: evidence for alternative first exon splicing. J. Biol. Chem. 276, 3106–3114. 40. Missirlis, F., Phillips, J.P. & Ja ¨ ckle, H. (2001) Cooperative action of antioxidant defense systems in Drosophila. Curr. Biol. 11, 1272–1277. 41. Missirlis, F., Ulschmid, J.K., Hirosawa-Takamori, M., Gronke, S., Scha ¨ fer, U., Becker, K., Phillips, J.P. & Ja ¨ ckle, H. (2002) Mitochondrial and cytoplasmic thioredoxin reductase variants encoded by a single Drosophila gene are both essential for viability. J. Biol. Chem. 277, 11521–11526. 42. Lee, S.R., Bar-Noy, S., Kwon, J., Levine, R.L., Stadtman, T.C. & Rhee, S.G. (2000) Mammalian thioredoxin reductase: oxidation of the C-terminal cysteine/selenocysteine active site forms a thiosele- nide, and replacement of selenium with sulfur markedly reduces catalytic activity. Proc.NatlAcad.Sci.USA97, 2521–2526. 43. Gromer, S., Arscott, L.D., Williams, C.H. Jr, Schirmer, R.H. & Becker, K. (1998) Human placenta thioredoxin reductase: iso- lation of the selenoenzyme, steady state kinetics, and inhibition by therapeutic gold compounds. J. Biol. Chem. 273, 20096–20101. 44. Golenser, J., Miller, J., Spira, D.T., Kosower, N.S., Vande Waa, J.A. & Jensen, J.B. (1988) Inhibition of the intraerythrocytic development of Plasmodium falciparum in glucose-6-phosphate dehydrogenase deficient erythrocytes is enhanced by oxidants and by crisis form factor. Trop. Med. Parasitol. 39, 273–276. 45. Ginsburg, H., Atamna, H., Shalmiev, G., Kanaani, J. & Krugliak, M. (1996) Resistance of glucose-6-phosphate dehydrogenase deficiency to malaria: effects of fava bean hydroxypyrimidine glucosides on Plasmodium falciparum growth in culture and on the phagocytosis of infected cells. Parasitology 113, 7–18. 46. Becker, K. & Schirmer, R.H. (1995) 1,3-Bis (2-chloroethyl)-1- nitrosourea as thiol-carbamoylating agent in biological systems. Methods Enzymol. 251, 173–188. 4280 H. Bauer et al. (Eur. J. Biochem. 270) Ó FEBS 2003 47. Scho ¨ nleben-Janas, A., Kirsch, P., Mittl, P.R., Schirmer, R.H. & Krauth-Siegel, R.L. (1996) Inhibition of human glutathione reductase by 10-arylisoalloxazines: crystalline, kinetic, and elec- trochemical studies. J. Med. Chem. 39, 1549–1554. 48. Giribaldi, G., Ulliers, D., Mannu, F., Arese, P. & Turrini, F. (2001) Growth of Plasmodium falciparum induces stage-dependent haemichrome formation, oxidative aggregation of band 3, mem- brane deposition of complement and antibodies, and phagocytosis of parasitized erythrocytes. Br. J. Haematol. 113, 492–499. 49. Cappadoro, M., Giribaldi, G., O’Brien, E., Turrini, F., Mannu, F., Ulliers, D., Simula, G., Luzzatto, L. & Arese, P. (1998) Early phagocytosis of glucose-6-phosphate dehydrogenase (G6PD) - deficient erythrocytes parasitized by Plasmodium falciparum may explain malaria protection in G6PD deficiency. Blood 92, 2527– 2534. 50. Tahar, R., Boudin, C., Thiery, I. & Bourgouin, C. (2002) Immune response of Anopheles gambiae to the early sporogonic stages of the human malaria parasite Plasmodium falciparum. EMBO J. 21, 6673–6680. 51. Dimopoulos, G., Seeley, D., Wolf, A. & Kafatos, F.C. (1998) Malaria infection of the mosquito Anopheles gambiae activates immune-responsive genes during critical transition stages of the parasite life cycle. EMBO J. 17, 6115–6123. 52. Dimopoulos, G., Christophides, G.K., Meister, S., Schultz, J., White, K.P., Barillas-Mury, C. & Kafatos, F.C. (2002) Genome expression analysis of Anopheles gambiae: responses to injury, bacterial challenge, and malaria infection. Proc. Natl Acad. Sci. USA 99, 8814–8819. 53. Francischetti, I.M., Valenzuela, J.G., Pham, V.M., Garfield, M.K. & Ribeiro, J.M. (2002) Toward a catalog for the transcripts and proteins (sialome) from the salivary gland of the malaria vector Anopheles gambiae. J. Exp. Biol. 205, 2429–2451. 54. Feyereisen, R. (1995) Molecular biology of insecticide resistance. Toxicol. Lett. 82–83, 83–90. 55. Powis, G., Mustacich, D. & Coon, A. (2000) The role of the redox protein thioredoxin in cell growth and cancer. Free Radic. Biol. Med. 29, 312–322. 56. Rahlfs, S., Schirmer, R.H. & Becker, K. (2002) The thioredoxin system of Plasmodium falciparum and other parasites. Cell. Mol. Life Sci. 59, 1024–1041. Ó FEBS 2003 Thioredoxin reductase from Anopheles gambiae (Eur. J. Biochem. 270) 4281 . Thioredoxin reductase from the malaria mosquito Anopheles gambiae Comparisons with the orthologous enzymes of Plasmodium falciparum and the human host Holger Bauer 1 , Stephan Gromer 1 , Andrea. AAB35418), and from the malaria parasite Plasmodium falciparum (PfTrxR-1, CAA60574). The enzyme of Anopheles shares 69%, 52%, and 45% sequence identity with the TrxRs of D. melanogaster, humans, and. (Figs 3 and 4). The reducing equivalents flow from the nicotinamide of NADPH via the flavin and the pair Cys57/Cys62 to the redox centre Cys500’/Cys501’ of the other subunit, and hence to the disulfide