Skip to main page content

• HOME
• Current Issue
• Archive
• Alerts
• About ASM
• Contact us
• Tech Support
• Journals.ASM.org

This Article Abstract Full Text (PDF) Alert me when this article is cited Alert me if a correction is posted Services Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Reprints and Permissions Copyright Information Books from ASM Press MicrobeWorld Citing Articles Citing Articles via Google Scholar Google Scholar Articles by Yeom, J. Articles by Park, W. Search for Related Content PubMed PubMed Citation Articles by Yeom, J. Articles by Park, W.

 Previous Article  |  Next Article 

Journal of Bacteriology, March 2009, p. 1472-1479, Vol. 191, No. 5
0021-9193/09/$08.00+0     doi:10.1128/JB.01473-08
Copyright © 2009, American Society for Microbiology. All Rights Reserved.

Ferredoxin-NADP⁺ Reductase from Pseudomonas putida Functions as a Ferric Reductase

Jinki Yeom,¹ Che Ok Jeon,² Eugene L. Madsen,³ and Woojun Park^1*

Division of Environmental Science and Ecological Engineering, Korea University, Seoul 136-075, Republic of Korea,¹ Department of Life Science, Chung-Ang University, Seoul 156-756, Republic of Korea,² Department of Microbiology, Cornell University, Ithaca, New York 14853-8101³

Received 20 October 2008/ Accepted 19 December 2008

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Pseudomonas putida harbors two ferredoxin-NADP⁺ reductases (Fprs) on its chromosome, and their functions remain largely unknown. Ferric reductase is structurally contained within the Fpr superfamily. Interestingly, ferric reductase is not annotated on the chromosome of P. putida. In an effort to elucidate the function of the Fpr as a ferric reductase, we used a variety of biochemical and physiological methods using the wild-type and mutant strains. In both the ferric reductase and flavin reductase assays, FprA and FprB preferentially used NADPH and NADH as electron donors, respectively. Two Fprs prefer a native ferric chelator to a synthetic ferric chelator and utilize free flavin mononucleotide (FMN) as an electron carrier. FprB has a higher k_cat/K_m value for reducing the ferric complex with free FMN. The growth rate of the fprB mutant was reduced more profoundly than that of the fprA mutant, the growth rate of which is also lower than the wild type in ferric iron-containing minimal media. Flavin reductase activity was diminished completely when the cell extracts of the fprB mutant plus NADH were utilized, but not the fprA mutant with NADPH. This indicates that other NADPH-dependent flavin reductases may exist. Interestingly, the structure of the NAD(P) region of FprB, but not of FprA, resembled the ferric reductase (Fre) of Escherichia coli in the homology modeling. This study demonstrates, for the first time, the functions of Fprs in P. putida as flavin and ferric reductases. Furthermore, our results indicated that FprB may perform a crucial role as a NADH-dependent ferric/flavin reductase under iron stress conditions.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Commonly, Fprs are ubiquitous, monomeric, reversible flavin enzymes. Fprs evidence a profound preference for NADP(H) over NAD(H) (3). They harbor a prosthetic flavin cofactor (FAD) and catalyze the reversible electron exchange between NADPH and either ferredoxin (Fd) or flavodoxin (Fld) (4, 5). In oxygenic photosynthesis, the Fd is reduced by the photosystem and subsequently passes electrons on to NADP⁺ via the Fpr. This reaction provides the cellular NADPH pool required for CO₂ assimilation and other biosynthetic processes (4, 5). In heterotrophic organisms such as bacteria, reduced ferredoxin, owing to the reverse enzymatic activity of the Fpr, can donate an electron to several Fd-dependent enzymes, such as nitrite reductase, sulfite reductase, glutamate synthase, and Fd-thioredoxin reductase, allowing ferredoxin to function in a variety of systems, including oxidative stress (1, 4, 5).

Iron is the fourth most abundant element in the natural environment and exists primarily as an oxidized form, Fe(III), which has very low solubility under neutral pH conditions (9, 34) and thus presents problems in terms of bioavailability. However, ferrous iron, of Fe(II), is soluble and available at neutral pH in bacterial cytosol (34). Most bacteria secrete siderophores, which are natural chelators of ferric iron. After they bind to ferric iron, that complex enters the bacteria and releases ferric iron into the cytosol in ferric or ferrous form (9). In the bacterial cytosol, ferric iron must be reduced to ferrous form, and thus ferric reductase is essential to bacterial iron utilization.

Commonly, prokaryotic ferric reductases are divided into two groups—namely, the bacterial and archaeal types (34). The typical bacterial type ferric reductase is Escherichia coli Fre, which also functions as a flavin reductase. In other words, the ferric reductase can reduce free flavin as flavin reductase, rather than having the flavin cofactor as a prosthetic group in E. coli (38). The archaeal ferric reductase harbors a flavin cofactor in the enzyme and thus does not require a flavin carrier for ferric reduction (26, 34). E. coli Fre includes a Rosmann folding structure at the NAD(P) binding region, whereas the archaeal ferric reductase (FeR) of Archaeoglobus fulgidus does not evidence that folding structure (6, 34). Many bacterial ferric reductases utilize free flavins, such as flavin mononucleotide (FMN), flavin adenine dinucleotide (FAD) and riboflavin, as electron carrier and, NADH (NAD) or NADP as electron donors to ferric reductase (14, 34). However, reduced ferric iron by reduced free flavin gives rise to the Fenton reaction, which generates the hydroxyl radical within the cell (20, 38). The Fenton reaction is known to generate hydroxyl radicals from ferrous iron and hydrogen peroxide (20). The hydroxyl radical is the most reactive radical and can damage DNA, proteins, and membrane lipids (16, 20, 34, 38). Therefore, the fine-tuning of ferric reduction regulation is required for the survival of bacterial cells.

Many Pseudomonas strains, including Pseudomonas putida, a gram-negative soil model bacteria, and Pseudomonas aeruginosa, a human pathogen bacteria, do not harbor annotated ferric reductase within their genome sequences. Commonly, the pathogens compete with the host for available iron, whichis crucial for their survival within the host. Thus, studies of P. aeruginosa regarding iron utilization, siderophores, and ferric reduction are considered to be essential for a better understanding of human infections (9, 19). Studying the physiology and ecology of P. putida also provides us with a new framework for elucidating the basis of the metabolic versatility and environmental stress response of soil microorganisms. Thus, the study of ferric reductase in strains of Pseudomonas at the molecular level is certainly required. From the structural perspective, ferric reductases are generally considered to be contained within the structurally diverse ferredoxin-NADP⁺ reductase (Fprs; EC 1.18.1.2) superfamily, which is frequently involved in the transfer of electrons between Fd/Fld and NADP(H) (2, 15, 34). Thus, we tested the role of the Fpr as a ferric reductase using free flavin (FMN or FAD), NADH, or NADPH as electron donors, and ferric-citrate or ferric-EDTA as terminal electron acceptors (37). We determined that FprA could efficiently utilize NADPH in ferric reduction. Rather, FprB could use NADH as an electron donor and may perform a crucial role as a NADH-dependent ferric reductase under iron stress conditions.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Bacterial strains, plasmids, and growth conditions. The bacterial strains and plasmids are shown in Table 1. Antibiotics (kanamycin, 100 µg/ml; ampicillin, 50 µg/ml) were added whenever necessary. The open reading frame of the fprA gene was PCR amplified using the FprA-OE1 and FprA-YT2 primer pairs, and the fprB open reading frame was amplified using the FprB-OE1 and FprB-OE2 primer pairs. The amplified fragments, harboring the fprA and fprB genes, were cloned into the NdeI/SalI sites of pET-28a(+), yielding pET-fprA and pET-fprB. pET-fprA and pET-fprB were transformed into E. coli BL21(DE3) cells via electroporation. E. coli BL21(DE3) cells were grown with moderate shaking, at a range of temperatures, in 2-YT medium supplemented with kanamycin (100 µg/ml). The cells were grown to the mid-log phase (optical density at 600 nm [OD₆₀₀] of 0.7) at 37°C, with aeration. The cells were then induced by adding 0.25 mM IPTG (isopropyl-β-D-thiogalactopyranoside) for 5 to 7 h at 30°C. In the case of the growth of wild-type P. putida and fprA and fprB mutant strains in minimal medium (M9), they were cultured with ferric-citrate 0.5 µM at 30°C with aeration.

View this table: [in this window] [in a new window]

TABLE 1. Strains, plasmids and primers

Protein purification. All purification steps were conducted at 4°C, using a fast-performance liquid chromatography (FPLC) system (AKTA FPLC, Unicorn 4.0; Amersham Bioscience). In the case of flavoproteins, or Fprs, E. coli cell pellets were resuspended in buffer A (50 mM Tris-Cl, 1 mM dithiothreitol [pH 7.5]) and disrupted via sonication. After the removal of cell debris by 30 min of centrifugation at 12,000 x g, the soluble fraction was loaded onto an anion-exchange column (1 ml, DEAE-cellulose; Amersham Bioscience) equilibrated with buffer A, and the proteins were eluted with a 20-ml linear gradient of 0 to 1 M NaCl in buffer A (pH 7.5). The fractions (0.5 ml each) were collected and concentrated via ultrafiltration with a Centricon (2 ml of YM-10; Amicon). The concentrates were then applied to a nickel-nitrilotriacetic acid column (1 ml, His-Trap; Amersham Bioscience) equilibrated with binding buffer (20 mM sodium phosphate, 0.5 M NaCl, 40 mM imidazole [pH 7.4]), and the proteins were eluted with 15 ml of elution buffer (20 mM sodium phosphate, 0.5 M NaCl, 250 mM imidazole [pH 7.4]). The fractions were dialyzed via ultrafiltration with a Centricon apparatus and stored at –80°C in 10% glycerol (30, 36). Sodium dodecyl sulfate-10% polyacrylamide gel electrophoresis was used to track the progress of the purification of the two Fprs.

Enzyme kinetics for analysis of catalytic activity. All chemicals and electron acceptors using enzyme kinetics were purchased from Sigma. Enzyme kinetics were monitored by using an Optizen 2120 UV/VIS spectrophotometer (Mecasys, Korea) at 25°C under anaerobic conditions. The ferric reductase assay was recorded at 562 nm in Tris-Cl buffer (100 mM; pH 7.5), and any increase in absorbance at 562 nm indicated the formation of the ferrozine-ferrous complex. Buffer solutions were prepared under anaerobic conditions via 2 h of the application of O₂-free nitrogen with continuous evacuation. The ferric reductase assay utilized 0.15 mM NADPH or NADH as the electron donor and various concentrations of ferric-citrate/ferric-EDTA as the electron acceptors. The steady-state kinetic parameters were dependent on the concentration of ferric-citrate/ferric-EDTA. The ferric reductase assay also contained free flavins, specifically 15 µM FAD or FMN, and 2 µg of FprA or FprB was used for the assay (13, 28). Each kinetic experiment was repeated five times.

The flavin reductase assay was also performed under anaerobic conditions by applying O₂-free nitrogen for 2 h with continuous evacuation. For flavin reductase, glucose-6-phosphate and glucose-6-phosphate dehydrogenase were omitted, and we measured the absorbance at 340 nm to assess the consumption of NAD(P)H (27, 33, 37). In order to calculate the ferric reduction activity by FprA and FprB, the reagent-only control experiments were conducted under identical conditions, except that the free flavin or crude extract were omitted. A single unit of activity was defined as the quantity of enzyme required to oxidize 1 µmol of NAD(P)H per min, and the extinction coefficients of NADPH and NADH were 6.20 and 6.22 M^–1 cm^–1, respectively (27, 37). The flavin reduction assay was repeated three times.

Western blotting of Fprs under various stresses. Hyperimmune rabbit antisera were raised against FprA and FprB. In brief, His-FprA and His-FprB were expressed in E. coli and purified on an anion-exchange column (1 ml, DEAE-cellulose; Amersham Bioscience) and a nickel-nitrilotriacetic acid column (1 ml; His-Trap) using an FPLC system. After collection of preimmune sera, two New Zealand White female rabbits at the age of 2 to 3 months were immunized intramuscularly with the purified Fprs (500 µg) emulsified with an equal volume of complete Freund adjuvant on day 0. Booster injections with the same immunogen (200 µg per rabbit) emulsified in incomplete Freund adjuvant was subcutaneously administered on days 28, 42, and 56. At 2 weeks after the final injection, both animals were sacrificed to obtain a maximal volume of blood through cardiac puncture. After removal of the clotted blood cells by centrifugation, anti-Fpr immune sera were collected and stored at –20°C until use. Immunoglobulin G molecules were precipitated from the hyperimmune antisera with 50% saturated ammonium sulfate solution, resuspended in cold phosphate-buffered saline (PBS), and dialyzed into PBS. Western blotting was conducted by using Western Lighting Chemiluminescence Reagent Plus (Perkin-Elmer) and polyvinylidene difluoride membrane (Bio-Rad) in accordance with the manufacturers’ instructions. The Fpr band was analyzed by using ProXPRESS 2D (Perkin-Elmer) and Total Lab 2.0 Software (Nonlinear Dynamics; BioSystematica, United Kingdom).

Northern blot analysis and green fluorescent protein (GFP) fluorescence measurement. Total RNA was isolated from 6 ml of exponentially growing cells, using an RNeasy minikit (Qiagen) according to the manufacturer's instructions. Northern blot analysis was performed as described previously (24, 25). RNA concentrations were estimated using absorbance at 260 nm. Samples of total RNA (5 to 10 µg) were loaded on denaturing agarose gels containing 0.25 M formaldehyde and electrophoresed, and the gels were stained with ethidium bromide to visualize 23S and 16S rRNA. The fractionated RNA was transferred to nylon membranes (Schleicher & Schuell) by using a Turboblotter (Schleicher & Schuell). The amounts of fprA and fprB mRNA were determined by hybridizing the membrane with a fprA- and fprB-specific, ³²P-labeled probe (Takara) prepared by PCR amplification with the primer pairs fprPp-1/fprPp-2 and FprB-KO1/FprB-KO2, respectively (24).

Bacterial cells, with the P_fprA::gfp and the P_fprB::gfp fusion, at the exponential growth phase (OD₆₀₀ of 0.5) grown in LB medium supplemented with various iron conditions, were collected by using a microcentrifuge (15,800 x g) and washed twice with PBS (137 mM NaCl, 10 mM phosphate, 2.7 mM KCl [pH 7.4]). Then, both the OD₆₀₀ and the GFP fluorescence intensity of the resuspended cultures were quantified by using a microtiter plate reader (Victor³; Bio-Rad). This reporter strains expresses a stable GFP variant that absorbs light at 488 nm (25).

Homology modeling of the protein complex. Homology models of Fprs were generated by using the SWISS-MODEL (http://swissmodel.expasy.org) and the Protein Homology/Analogy Recognition Engine (PHYRE, version 0.2; Imperial College, London, United Kingdom) homology modeling sites (17). The FprA model used the Fpr of A. vinelandii, due to its high degree of similarity with that of P. putida, and the FprB model utilized E. coli Fpr as a template model. The Fre of the E. coli structure (PDB entry 1QFJ) was utilized as a comparison model for the NAD(P) region of Fprs. PDB files were generated by using the DeepView/Swiss PDB-viewer (version 3.7) and PyMOL (version 1.0) (7, 22).

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Ferric reductase activity of Fpr in vitro. Commonly, the bacterial ferric reductase uses NAD(P)H as an electron donor and free flavin as an electron carrier (34). Thus, we tested NADH/NADPH as an electron donor and free FMN/FAD as an electron carrier. In the present study, ferric-citrate and ferric-EDTA were used as the terminal electron acceptor, which represent a native ferric chelator and a synthetic ferric iron chelator, respectively. When NADPH was utilized as an electron donor, FprA evidenced profound catalytic activity (k_cat = 37.6 ± 1.3 s^–1 for ferric-citrate, 11.2 ± 0.2 s^–1 for ferric-EDTA) in the presence of free FMN (Table 2). The K_m value of FprA (6.25 µM) was extremely low compared to those of other ferric reductases (Tables 2 and 3). Experiments utilizing either free FAD or Fpr alone resulted in a low level of catalytic efficiency (k_cat/K_m). In the case of the NADPH-dependent ferric reductase activity of FprB, the K_m value was significantly higher than that observed with FprA using free FMN as an electron carrier (Table 2). As such, FprA may function as an NADPH-dependent ferric reductase using free FMN (Table 2). Surprisingly, the moderate k_cat and very low K_m (2.35 ± 0.04 µM for ferric-citrate, 6.50 ± 0.03 µM for ferric-EDTA) of FprB was observed when paired with NADH. The data implied that FprB preferentially uses NADH as an electron donor in the context of ferric reduction and uses free FMN as an electron carrier (Table 2). The K_m value of FprB was also quite low (2.35 µM) compared to those of other ferric reductases (Tables 2 and 3). Commonly, bacterial Fprs are known to utilize NADPH as an electron donor (3-5). However, in the present study, FprB evidently used NADH as an electron donor in the ferric reductase assay. The NADH preference of FprB was also noted in the diaphorase assay (J. Yeom et al., unpublished data). Thus, FprB may play the role of an efficient ferric reductase, using NADH as an electron donor and free FMN as an electron carrier. Since the overall catalytic efficiency (k_cat/K_m) of both Fprs with ferric-citrate is higher than with ferric-EDTA, the Fprs of P. putida prefer the natural ferric-citrate to the synthetic ferric-EDTA in ferric iron reduction (Table 2). Although ferric-citrate is reduced by the ferric-citrate transport gene (Fec) cluster in the periplasm of proteobacteria, ferric-citrate may function as a soluble native ferric chelator (9). Therefore, the reduction of ferric-citrate is considered to be crucial to our understanding of the role of ferric reductase.

View this table: [in this window] [in a new window]

TABLE 2. Kinetic parameters of electron transfer by FprA and FprB, determined using the ferric reductase assaya

View this table: [in this window] [in a new window]

TABLE 3. Kinetic data comparison between P. putida and other organisms as determined by ferric reductase assay

Growth phenotypes of Fpr-deficient mutants under iron-limited conditions. In order to verify the functions of the Fprs as ferric reductases in vivo, the growth rates of two Fpr-deficient mutants (i.e., the fprA and fprB mutants) were monitored in M9 minimal medium containing Na₂HPO₄·7H₂O (6.8 g), KH₂PO₄ (3 g), NaCl (0.5 g), NH₄Cl (1 g), MgSO₄ (2 mM), and CaCl₂ (0.1 mM) with glucose (2 g/liter) as a carbon source and ferric-citrate (0.5 µM), which provided a sufficient iron concentration for supporting the growth of soil bacteria and their ferric reductase activity (27). In the control experiments, in which no extra ferric iron was added, the growth rate of the wild-type strain was not different from that of the fprB mutant (wild type, 1.09 ± 0.03 h^–1; fprB mutant, 1.13 ± 0.03 h^–1). However, the growth rate of the fprA mutant was slightly higher than those of the wild type and fprB mutant (fprA mutant, 1.30 ± 0.07 h^–1). In the M9 medium with ferric-citrate, the Fpr-deficient mutants evidenced significantly lower growth rates (fprA mutant, 2.02 ± 0.09 h^–1; fprB mutant, 1.84 ± 0.04 h^–1) than were observed with the wild-type strain (3.11 ± 0.11 h^–1). Under iron-amended conditions, the growth rate of the fprA mutant was also found to be slightly higher than that of the fprB mutant. These data indicated that both fpr gene products were involved in the utilization of ferric iron and may function as ferric reductases. However, the explanation for the faster growth of the fprA mutant than the fprB mutant under both conditions is not straightforward. However, we had observed that the expression of the fprB gene was increased in the fprA mutant (Fig. 1A). We also speculated that FprB may be involved in the iron-sulfur center assembly, since its gene is located near the isc-hsc gene cluster region, which may be linked to this growth pattern (12, 21).

View larger version (54K): [in this window] [in a new window]

FIG. 1. Expression analysis for Fprs under various conditions. (A) Northern blot analysis of the fprB gene expression in wild-type and the fprA mutant strains. Growth-phase-dependent expression of fprB was evaluated. Lane 0, overnight culture (16 h); the incubation times (in hours) are indicated above each lane. (B to D) Western blot analysis of Fprs under various stresses and iron conditions. Shown in each panel are the protein induction levels of Fprs under various stress conditions (B), in minimal medium (M9) with 0.5 µM ferric-citrate (C), and in minimal medium (M9) with ferrous sulfate (D). Each lane was loaded with 10 µg of crude extract, which was quantified via the Bradford method. In panel B, PQ (paraquat, 0.5 mM), MD (menadione, 1.0 mM), H2O2 (1.0 mM), and t-BooH (tert-butyl hydroperoxide, 0.5 mM) were applied to the cells for 30 min. Treatments within panel C: C, no treatment; Fe-C, ferric-citrate; Fe-E, ferric-EDTA and Di, 2',2'-dipyridal. Treatments within panel D: C (0.5 µM ferric-citrate), Fe-C1 (ferric-citrate, 1 µM), Fe-C5 (ferric-citrate, 5 µM), Fe-E2 (ferric-EDTA, 2 µM), Fe-E7 (ferric-EDTA, 7 µM), Di1 (dipyridal, 1.0 µM), and Di2 (dipyridal, 2.0 µM). Treatments within panel E: C (0.5 µM ferric-citrate), Fe(II)2 (iron sulfate, 2 µM), and Fe(II)7 (iron sulfate, 7 µM) treated for 30 min. The number below each lane shows the intensity of the band relative to the nontreated sample (lanes C), and bands were analyzed by using ProXPRESS 2D and Total Lab 2.0 software. (E) Northern blot analysis of the fpr genes under various conditions. Shown in each panel are the mRNA induction level of fpr genes in minimal medium (M9) with 0.5 µM ferric-citrate. All chemicals treated with 10 µM concentration, and "C" means no treatment. The ethidium bromide-stained gel prior to blotting demonstrated consistent loading in all lanes. The number below each lane shows the intensity of the band relative to the nontreated sample (sample C), and bands were analyzed by using ProXPRESS 2D and Total Lab 2.0 software. (F) Quantification of GFP expression in fprA and fprB genes reporter strains grown in the presence of various iron conditions. GFP was measured as described in Materials and Methods. All chemicals were treated with 2 mM final concentrations. The GFP intensity was determined after 3 h.

Flavin reductase activity of Fpr using crude extract. We subsequently assessed flavin reductase activity using the crude extracts from the wild-type strain and the two Fpr mutant strains (37). Since both Fprs utilize free flavin, particularly free FMN, as an electron carrier for ferric reduction, the flavin reductase assay using crude extract is physiologically relevant to ferric reduction within the cell. There is no FAD-dependent flavin reductase activity in both wild-type and mutant cells (Table 4). Commonly, ferric reductase, which does not harbor flavin cofactor in the enzyme, has the characteristics of a flavin reductase (38). Since the flavin reductase can give rise to the Fenton reaction, this flavin reductase feature is essential for bacteria (14). Interestingly, the crude extract from the fprB mutant evidenced no NADH-dependent flavin reductase activity using free FMN (Table 4). The crude extracts of the fprA and fprB mutants using NADPH as an electron donor and free FMN as an electron acceptor also evidenced low flavin reduction activity relative to that of the wild-type strain; but the fprA mutant using NADH as an electron donor retained some activity. Collectively, the results of the ferric and flavin reductase assays (Tables 2 and 4) implied that (i) FprA may have low-level ferric reductase activity relative to that of FprB, (ii) FprB is essential for NADH-dependent ferric reductase using free FMN as an electron acceptor in vivo, and (iii) FprB can use both NADH and NADPH as an electron donor for ferric reduction and flavin reduction.

View this table: [in this window] [in a new window]

TABLE 4. Flavin reductase activity in extracts from P. putida wild type and FprA and FprB deletion mutants grown on ferric-citratea

Expression analysis of Fprs under various conditions. In an effort to gain further insight into the functions of Fprs in P. putida, Fpr expression at the protein level was assessed via Western blot analysis. Previously, Northern blot analysis has demonstrated that the fprA gene is induced by oxidative stress, whereas the fprB gene is slightly induced by salt stress but is not induced by oxidative stress (24, 25). Consistently, Western blot analysis showed that FprA was induced by a variety of oxidative stress reagents, and FprB was not (Fig. 1B). In order to verify our results regarding ferric reductase, Western blot analysis was conducted under ferric iron excess and ferrous iron depletion conditions in M9 minimal medium. The ferric iron excess condition was achieved via the addition of 0.5 µM ferric-citrate and ferric-EDTA in M9 medium and the ferrous iron depletion condition was achieved via the addition of 2'2'-dipyridyl, a ferrous iron chelator, in minimal medium. Interestingly, both Fprs of P. putida were induced by both conditions. The induction level of FprB was stronger than that of FprA under both ferric iron excess and ferrous iron depletion conditions; however, both Fprs were shown not to be induced by ferrous iron (Fig. 1C and D). Northern blot and GFP-reporter assays were consistent with our Western blot data. (Fig. 1E and F). The data presented here indicated that both Fprs perform crucial functions in ferric iron metabolism.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Commonly, the bacterial type Fprs are divided into two subclasses: subclass I and subclass II. The bacterial Fpr subclass I is represented by Azotobacter vinelandii Fpr, and their functions are modulated by phenylalanine residues. Bacterial subclass II is exemplified by E. coli Fpr, and the tryptophan residue of the C terminus of E. coil Fpr is crucial to their catalytic functions (4, 5, 8, 35). The majority of proteobacteria harbor only one Fpr, but A. vinelandii and Pseudomonas species contain two Fprs in their chromosomes. FprA (NP_743795.1) and FprB (NP_746755) of P. putida have 83.7 and 36.7% amino acid similarity, respectively, with the Fprs of A. vinelandii (ZP_00417949) and E. coli (NP_418359.1). However, the functions of their Fprs remain poorly understood. Also, ferric reductase was not annotated in the pseudomonad chromosome, and thus we assessed the function of P. putida Fprs as ferric reductases. The ferric reductase (Fre) of E. coli is included in the Fpr family with regard to its structure; however, ferric reductase differs from the Fpr enzymes in regard to its function (34). The Fpr of proteobacteria is not currently known to function as a ferric reductase. Thus, we assessed the Fpr of P. putida using a variety of methods, including kinetics, regulation, and structural studies.

Previously, we have demonstrated that the expression of the fprA gene is induced by oxidative stress and is regulated by the finR gene product (24, 31). However, the fprB gene is not induced by oxidative stress but rather is upregulated by salt stress (25). Consistent with this previous observation, the expression of the FprA protein under oxidative stress conditions is also upregulated (Fig. 1B). By way of contrast, the induction of E. coli Fpr occurs under superoxide stress conditions as the result of the soxRS regulatory system (23). However, the Fpr of P. putida is not influenced by the soxRS system (31). Recently, Newman's group reported that the soxR gene has different functions among the proteobacteria (11). Moreover, the soxS gene does not exist on the chromosome of P. putida, and therefore the regulation and function of Fpr in P. putida may differ from those of E. coli (31). Interestingly, the Fpr of E. coli is representative of bacterial subclass II, but the oxidative stress-induced FprA of P. putida, which belongs to bacterial subclass I by virtue of its structure, resembles the Fpr of E. coli (Fig. 2). We have shown here for the first time that the levels of FprA and FprB protein expression are increased under ferric iron stress conditions (Fig. 1C). Much stronger expression was noted with the FprB under such conditions. We speculated that FprA may be more relevant to the defense of oxidative stress in order to control the NADPH/NADP⁺ pool, as is the case with the E. coli Fpr. However, it is apparent that FprB may perform a significant function as a ferric reductase under ferric iron stress in P. putida. Many redox enzymes, for example, sulfite reductase and thiol reductase, can adventitiously reduce free flavins, albeit at a lower rate than does Fre in E. coli (29, 38). Thus, we cannot dismiss the possibility that many other redox enzymes will also function as flavin reductases in P. putida. However, the contribution of FprB's function as a ferric reductase is significant, as demonstrated by the in vivo and in vitro data presented herein.

View larger version (64K): [in this window] [in a new window]

FIG. 2. Modeling the structure of the C-terminal of FprB in P. putida and of Fre in E. coli. The green color indicates the FprB of P. putida, and the blue color indicates the Fre of E. coli. The pink-colored β-sheet structure in the FprB of P. putida indicates the structurally distinctive region between FprB and Fre.

Subsequently, the C-terminal region of FprB, which is known to be the NAD(P) interacting region, is structurally quite similar to the C-terminal region of fre, a gene that encodes for ferric reductase in E. coli (37.5% amino acid similarity) (Fig. 2). The NAD(P) domain of FprB, but not that of FprA, overlaps with the NAD-binding region of Fre, as determined via homology modeling (Fig. 2). Fre in E. coli utilizes NADH rather than NADPH as an electron donor (34). FprB evidenced the potential to bind to NADH in all of the assays reported above (ferric and flavin reductase assay; Tables 2 and 4). However, one of the β-sheet structures of the C-terminal region of FprB in P. putida differs from that of the Fre region of E. coli (Fig. 2, pink color). That distinctive β-sheet of the FprB C-terminal region features a group of residues near the key terminal tryptophan residue (which interacts with the FAD cofactor [5]). We speculate that this difference in β-sheet residues allows FprB to function with an FAD cofactor, whereas Fre in E. coli does not (34). The catalytic activities (k_cat) of FprB with ferric-citrate and ferric-EDTA are also quite a bit higher than those of FprA, and this suggests that FprB may be capable of catalysis with a broad range of ferric complexes (Table 2). Thus, the ferric reduction of FprB is determined by K_m values, and this indicates that the interaction affinity of the electron donor is crucial to the ferric reduction of FprB. Commonly, the flavin reduction activity and degree of the Fenton reaction are determined by a concentration of NADH in E. coli, and that is confirmed by the existence of cyanide, which disrupts the oxidative phosphorylation cycle in the bacterial membrane (38). Thus, the high catalytic activity (k_cat) and low K_m value of FprB using NADH as an electron donor are similar kinetic characteristics, especially compared to the Fre of E. coli (Tables 2 and 3). Interestingly, FprB prefers NADH to NADPH as an electron donor in the ferric reductase reaction.

Ferric reductase in E. coli has no flavin cofactor and rather uses free flavin as an electron carrier, and the ferric reductase of the archaeon A. fulgidus harbors a flavin cofactor within the enzyme (34). It has been known that bacterial ferric iron reductase appears to be a flavin reductase, which uses free flavin as an electron carrier. Then, reduced flavin can reduce various ferric iron complexes. Interestingly, we found that Fprs have significant ferric reductase activities without flavin cofactors, as shown in Table 2. This finding is probably due to the fact that Fprs have FAD cofactor in their N-terminal domains (5). It has been known that the ferric reductase of the archaeon A. fulgidus harbors a flavin cofactor within the enzyme and can either reduce free flavin or complexed ferric iron (37). Therefore, we proposed that Fprs can function similar with A. fulgidus ferric reductase.

The Fprs of P. putida harbor a flavin cofactor (FAD) and utilize free flavin (FMN) as an electron carrier. In eukaryotic organisms, the cytochrome P450 reductase utilizes both FAD and FMN cofactors, and electron transfer occurs between FAD and FMN (10, 32). These observations demonstrate that there may be an evolutionary trend across taxa and domains toward increased versatility in Fprs and their ability to transfer electrons to both FAD and free FMN (see below). One other study has evaluated a putative ferric reductase from Pseudomonas species (18, 19, 34). The results of that study demonstrated that the role of the 27.5-kDa ferric reductase is broadly related to ferric reduction using NADH and free FMN (34), a finding consistent with the data presented here.

In the present study, the function of two Fprs as ferric reductase was determined by kinetics, growth tests with targeted mutants, expression analysis, and structural studies. The Fpr has been shown to control the NADP⁺/NADPH pool in proteobacteria. Interestingly, two Fprs of P. putida can affect the reduction of ferric iron with free FMN, and its function appears to be quite important to P. putida from the standpoint of iron acquisition. All of the results of the present study are illustrated in Fig. 3. The data presented here expand the traditional view of Fprs that function as electron carriers between the electron donor and Fd/Fld to ferric/flavin reductase in proteobacteria.

View larger version (33K): [in this window] [in a new window]

FIG. 3. Proposed roles of ferric/flavin reductase by two Fprs in P. putida KT2440. The thickness of an arrow represents the catalytic efficiency (kcat/Km) of the redox partner. The dashed lines indicate that Fprs can either reduce free flavin or complexed ferric iron.

 
This study was supported by a National Core Research Center grant (R15-2003-002-01002-0) and a Korea Science and Engineering Foundation grant (R01-2008-000-10697-0) programs to W.P.

 
* Corresponding author. Mailing address: Division of Environmental Science and Ecological Engineering, Korea University, Seoul 136-075. Phone: 82-2-3290-3067. Fax: 82-2-953-0737. E-mail: wpark{at}korea.ac.kr

Published ahead of print on 29 December 2008.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 

1
1. Aliverti, A., R. Faber, C. M. Finnerty, C. Ferioli, V. Pandini, A. Negri, P. A. Karplus, and G. Zanetti. 2001. Biochemical and crystallographic characterization of ferredoxin-NADP⁺ reductase from nonphotosynthetic tissues. Biochemistry 40:14501-14508.[CrossRef][Medline]
2
2. Aliverti, A., V. Pandini, A. Pennati, M. de Rosa, and G. Zanetti. 2008. Structural and functional diversity of ferredoxin-NADP⁺ reductases. Arch. Biochem. Biophys. doi:10.1016/j.abb.2008.02.014.
3
3. Batie, C. J., and H. Kamin. 1986. Association of ferredoxin-NADP⁺ reductase with NADP(H) specificity and oxidation-reduction properties. J. Biol. Chem. 261:11214-11223.[Abstract/Free Full Text]
4
4. Carrillo, N., and E. A. Ceccarelli. 2003. Open questions in ferredoxin-NADP⁺ reductase catalytic mechanism. Eur. J. Biochem. 270:1900-1915.[Medline]
5
5. Ceccarelli, E. A., A. K. Arakaki, N. Cortez, and N. Carrillo. 2004. Functional plasticity and catalytic efficiency in plant and bacterial ferredoxin-NADP(H) reductases. Biochim. Biophys. Acta 1698:155-165.[Medline]
6
6. Chiu, H., E. Johnson, I. Schröder, and D. C. Rees. 2001. Crystal structure of a novel ferric reductase from the hyperthermophilic archaeon Archaeoglobus fulgidus and its complex with NADP⁺. Structure 9:311-319.[Medline]
7
7. Comeau, S. R., D. W. Gatchell, S. Vajda, and C. J. Camacho. 2004. ClusPro: a fully automated algorithm for protein-protein docking. Nucleic Acids Res. 32:W96-W99.[Abstract/Free Full Text]
8
8. Coves, J., and M. Foncave. 1993. Reduction and mobilization of iron by a NAD(P)H:flavin oxidoreductase from Escherichia coli. Eur. J. Biochem. 211:635-641.[Medline]
9
9. Crosa, J. H., A. R. Mey, and S. M. Payne. 2004. Iron transport in bacteria. ASM Press, Washington, DC.
10
10. Degtyarenko, K. N., and T. A. Kulikova. 2001. Evolution of bioinorganic motifs in P450-containing systems. Biochem. Soc. Trans. 29:139-147.[CrossRef][Medline]
11
11. Dietrich, L. E., T. K. Teal, A. Price-Whelan, and D. K. Newman. 2008. Redox-active antibiotics control gene expression and community behavior in divergent bacteria. Science 321:1203-1206.[Abstract/Free Full Text]
12
12. Dos-Santos, P. C., D. C. Johnson, B. E. Ragle, M. C. Unciuleac, and D. R. Dean. 2007. Controlled expression of nif and isc iron-sulfur protein maturation components reveals target specificity and limited functional replacement between the two systems. J. Bacteriol. 189:2854-2862.[Abstract/Free Full Text]
13
13. Fischer, F., D. Raimondi, A. Aliverti, and G. Zanetti. 2002. Mycobacterium tuberculosis FprA, a novel bacterial NADPH-ferredoxin reductase. Eur. J. Biochem. 269:3005-3013.[Medline]
14
14. Fontecave, M., J. Coves, and J. Pierre. 1994. Ferric reductases or flavin reductase? Biometals 7:3-8.[Medline]
15
15. Giró, M., N. Carrillo, and A. R. Krapp. 2006. Glucose-6-phosphate dehydrogenase and ferredoxin-NADP(H) reductase contribute to damage repair during the soxRS response of Escherichia coli. Microbiology 152:1119-1128.[Abstract/Free Full Text]
16
16. Green, J., and M. S. Paget. 2004. Bacterial redox sensors. Nat. Rev. Microbiol. 2:954-966.[CrossRef][Medline]
17
17. Guex, N., and M. C. Peitsch. 1997. SWISS-MODEL and the Swiss-PdbViewer: an environment for comparative protein modeling. Electrophoresis 18:2714-2723.[CrossRef][Medline]
18
18. Halle, F., and J. M. Meyer. 1989. Ferripyoverdie-reductase activity in Pseudomonas fluorescens. Biol. Metals 2:18-24.[CrossRef][Medline]
19
19. Halle, F., and J. M. Meyer. 1992. Ferrisiderophore reductase of Pseudomonas: purification, properties, and cellular location of the Pseudomonas aeruginosa ferriptoverdine reductase. Eur. J. Biochem. 209:613-620.[Medline]
20
20. Imlay, J. A. 2006. Iron-sulphur clusters and the problem with oxygen. Mol. Microbiol. 59:1073-1082.[CrossRef][Medline]
21
21. Jung, Y. S., H. S. Gao-Sheridan, J. Christiansen, D. R. Dean, and B. K. Burgess. 1999. Purification and biophysical characterization of a new [2Fe-2S] ferredoxin from Azotobacter vinelandii, a putative [Fe-S] cluster assembly/repair protein. J. Biol. Chem. 274:32402-32410.[Abstract/Free Full Text]
22
22. Kozakov, D., R. Brenke, S. R. Comeau, and S. Vajda. 2006. PIPER: an FFT-based protein docking program with pairwise potentials. Proteins 65:392-406.[CrossRef][Medline]
23
23. Krapp, A. R., R. E. Rodriguez, H. O. Poli, D. H. Paladini, and J. F. Palatnik. 2002. The flavoenzyme ferredoxin (flavodoxin)-NADP(H) reductase modulates NADP(H) homeostasis during the soxRS response of Escherichia coli. J. Bacteriol. 184:1474-1480.[Abstract/Free Full Text]
24
24. Lee, Y., S. Peña-Llopis, Y. S. Kang, H. D. Shin, B. Demple, E. L. Madsen, C. O. Jeon, and W. Park. 2006. Expression analysis of the fpr (ferredoxin-NADP⁺ reductase) gene in Pseudomonas putida KT2440. Biochem. Biophys. Res. Commun. 339:1246-1254.[CrossRef][Medline]
25
25. Lee, Y., J. Yeom, Y. S. Kang, J. Kim, J. S. Sung, C. O. Jeon, and W. Park. 2007. Molecular characterization of fprB (ferredoxin-NADP⁺ reductase) in Pseudomonas putida KT2440. J. Microbiol. Biotechnol. 17:1504-1512.[Medline]
26
26. Lesuisse, E., M. Casteras-Simon, and P. Labbe. 1997. Cytochrome P-450 reductase is responsible for the ferrireductase activity associated with isolated plasma membranes of Saccharomyces cerevisiae. FEMS Microbiol. Lett. 156:147-152.[Medline]
27
27. Mazoch, J., R. Tesarík, V. Sedlácek, I. Kucera, and J. Turánek. 2004. Isolation and biochemical characterization of two soluble iron(III) reductases from Paracoccus denitrificans. Eur. J. Biochem. 271:553-562.[Medline]
28
28. Nascimento, A. S., D. L. Catalano-Dupuy, A. Bernardes, O. Neto-Mde, and M. A. Santos. 2007. Crystal structures of Leptospira interrogans FAD-containing ferredoxin-NADP⁺ reductase and its complex with NADP⁺. BMC Struct. Biol. 7:69.[CrossRef][Medline]
29
29. Nijvipakul, S., J. Wongratana, C. Suadee, B. Entsch, D. P. Ballou, and P. Chaiyen. 2008. LuxG is a functioning flavin reductase for bacterial luminescence. J. Bacteriol. 190:1531-1538.[Abstract/Free Full Text]
30
30. Pandini, V., G. Caprini, N. Thomsen, A. Aliverti, F. Seeber, and G. Zanetti. 2002. Ferredoxin-NADP⁺ reductase and ferredoxin of the protozoan parasite Toxoplasma gondii interact productively in vitro and in vivo. J. Biol. Chem. 277:48463-48471.[Abstract/Free Full Text]
31
31. Park, W., S. Pena-Llopis, Y. Lee, and B. Demple. 2006. Regulation of superoxide stress in Pseudomonas putida KT2440 is different from the SoxR paradigm in Escherichia coli. Biochem. Biophys. Res. Commun. 341:51-56.[CrossRef][Medline]
32
32. Porter, T. D., and C. B. Kasper. 1986. NADPH-cytochrome P-450 oxidoreductase: flavin mononucleotide and flavin adenine dinucleotide domains evolved from different flavoproteins. Biochemistry 25:1682-1687.[CrossRef][Medline]
33
33. Puzon, G. J., J. N. Petersen, A. G. Roberts, D. M. Kramer, and L. Xun. 2002. A bacterial flavin reductase system reduces chromate to a soluble chromium(III)-NAD⁺ complex. Biochem. Biophys. Res. Commun. 294:76-81.[CrossRef][Medline]
34
34. Schröder, I., E. Johnson, and S. de Vries. 2003. Microbial ferric iron reductases. FEMS Microbiol. Rev. 27:427-447.[CrossRef][Medline]
35
35. Seo, D., A. Tomioka, N. Kusumoto, M. Kamo, and I. Enami. 2001. Purification of ferredoxins and their reaction with purified reaction center complex from the green sulfur bacterium Chlorobium tepidum. Biochim. Biophys. Acta 1503:377-384.[Medline]
36
36. Sridhar-Prasad, G., N. Kresge, A. B. Muhlberg, A. Shaw, Y. S. Jung, and B. K. Burgess. 1998. The crystal structure of NADPH:ferredoxin reductase from Azotobacter vinelandii. Protein Sci. 7:2541-2549.[Medline]
37
37. Takeda, K., M. Iizuka, T. Watanabe, J. Nakagawa, S. Kawasaki, and Y. Niimura. 2007. Synechocystis DrgA protein functioning as nitroreductase and ferric reductase is capable of catalyzing the Fenton reaction. FEBS J. 274:1318-1327.[CrossRef][Medline]
38
38. Woodmansee, A. N., and J. A. Imlay. 2002. Reduced flavins promote oxidative DNA damage in non-respiring Escherichia coli by delivering electrons to intracellular free iron. J. Biol. Chem. 277:34055-34066.[Abstract/Free Full Text]

---

Journal of Bacteriology, March 2009, p. 1472-1479, Vol. 191, No. 5
0021-9193/09/$08.00+0     doi:10.1128/JB.01473-08
Copyright © 2009, American Society for Microbiology. All Rights Reserved.

This Article Abstract Full Text (PDF) Alert me when this article is cited Alert me if a correction is posted Services Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Reprints and Permissions Copyright Information Books from ASM Press MicrobeWorld Citing Articles Citing Articles via Google Scholar Google Scholar Articles by Yeom, J. Articles by Park, W. Search for Related Content PubMed PubMed Citation Articles by Yeom, J. Articles by Park, W.