Advertisement
JBC

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Originally published In Press as doi:10.1074/jbc.M408303200 on August 15, 2004

J. Biol. Chem., Vol. 279, Issue 44, 45791-45802, October 29, 2004
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
279/44/45791    most recent
M408303200v1
Right arrow Submit a Letter to Editor
Right arrow Alert me when this article is cited
Right arrow Alert me when eLetters are posted
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowRequest Permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Wegele, R.
Right arrow Articles by Frankenberg-Dinkel, N.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Wegele, R.
Right arrow Articles by Frankenberg-Dinkel, N.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg   Add to Reddit   Add to Technorati  
What's this?

The Heme Oxygenase(s)-Phytochrome System of Pseudomonas aeruginosa*

Rosalina Wegele{ddagger}, Ronja Tasler{ddagger}, Yuhong Zeng§, Mario Rivera§, and Nicole Frankenberg-Dinkel{ddagger}

From the {ddagger}Institute for Microbiology, Technical University Braunschweig, Spielmannstrasse 7, 38106 Braunschweig, Germany and the §Department of Chemistry, University of Kansas, Lawrence, Kansas 66045-7582

Received for publication, July 22, 2004 , and in revised form, August 11, 2004.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
For many pathogenic bacteria like Pseudomonas aeruginosa heme is an essential source of iron. After uptake, the heme molecule is degraded by heme oxygenases to yield iron, carbon monoxide, and biliverdin. The heme oxygenase PigA is only induced under iron-limiting conditions and produces the unusual biliverdin isomers IX{beta} and IX{delta}. The gene for a second putative heme oxygenase in P. aeruginosa, bphO, occurs in an operon with the gene bphP encoding a bacterial phytochrome. Here we provide biochemical evidence that bphO encodes for a second heme oxygenase in P. aeruginosa. HPLC, 1H, and 13C NMR studies indicate that BphO is a "classic" heme oxygenase in that it produces biliverdin IX{alpha}. The data also suggest that the overall fold of BphO is likely to be the same as that reported for other {alpha}-hydroxylating heme oxygenases. Recombinant BphO was shown to prefer ferredoxins or ascorbate as a source of reducing equivalents in vitro and the rate-limiting step for the oxidation of heme to biliverdin is the release of product. In eukaryotes, the release of biliverdin is driven by biliverdin reductase, the subsequent enzyme in heme catabolism. Because P. aeruginosa lacks a biliverdin reductase homologue, data are presented indicating an involvement of the bacterial phytochrome BphP in biliverdin release from BphO and possibly from PigA.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
In the presence of a suitable electron donor the enzyme heme oxygenase (HO,1 EC 1.14.99.3 [EC] ) catalyzes the oxidative degradation of heme to yield equimolar amounts of biliverdin (BV), iron, and CO (see Fig. 1). In the well studied eukaryotic HOs, NADPH and NADPH-cytochrome P450 reductase serve as the electron donor (1, 2). Bacterial HOs have only recently been described, and the HO from Corynebacterium diphtheriae (HmuO) was among the first ones to be discovered (3). In C. diphtheriae the HO reaction is utilized to release iron from heme under pathogenic (i.e. free-iron limiting) conditions (3). In fact, the breakdown of heme to mine iron is thought to be the major function of HOs from pathogenic organisms, thus allowing them to overcome the low concentrations of free-iron necessary for successful colonization (infection). Their function in iron utilization has been shown for HOs from several pathogenic bacteria, including Neisseria meningitidis (HemO), Corynebacterium ulcerans (HmuO), and Staphylococcus aureus (IsdG/I) (4–6). One of the most interesting bacterial HOs was recently identified in the Gram-negative opportunistic pathogen Pseudomonas aeruginosa. The pigA gene, originally identified in a screen for iron-regulated genes, was shown to encode a protein with homology to the neisserial HOs (7). The gene, which is part of a five-gene operon (pigA–E), is under the control of the global iron regulator Fur (ferric uptake regulator) and is only expressed under iron-limiting conditions (8). Unlike all other eu- and prokaryotic HOs that regiospecifically cleave the {alpha}-meso carbon of heme, PigA targets the {beta}- and {delta}-meso carbons of the heme macrocycle (7) (Fig. 1). The unusual production of {beta}- and {delta}-BV was shown to be due to an unusual seating of the substrate heme in the active site pocket of PigA (9). The purpose for the production of these BV isomers and their fate in P. aeruginosa, however, is currently unknown. In addition to their function in iron utilization bacterial HOs are also involved in the first step of phycobilin biosynthesis. These linear tetrapyrrole molecules are precursors of the chromophores for the cyanobacterial light-harvesting phycobiliproteins and the photoreceptor phytochrome. Phytochromes are traditionally known as biliprotein photoreceptors in plants but have recently also been discovered in bacteria (10). Unlike plant and cyanobacterial phytochromes, which carry a phytochromobilin or phycocyanobilin chromophore, bacteriophytochromes (BphPs) from non-photosynthetic prokaryotes have been shown to utilize a BV chromophore (11). P. aeruginosa was among the first non-photosynthetic bacteria identified to harbor a gene encoding for a BphP. This bacteriophytochrome is a typical sensor kinase of a prokaryotic two-component signaling system. The bphP gene is located in an operon downstream from a bphO gene, which encodes a putative HO. So far, no gene encoding for the corresponding response regulator has been identified in P. aeruginosa. Herein we present the first biochemical and biophysical evidence that bphO encodes for a functional HO that oxidizes heme to BV IX{alpha}. Because P. aeruginosa is the first known organism to encode two functional HOs, each capable of oxidizing heme to a different BV isomer, their involvement in phytochrome chromophore biosynthesis has been investigated and will also be discussed.



View larger version (18K):
[in this window]
[in a new window]
  		FIG. 1.
Reactions catalyzed by P. aeruginosa heme oxygenases. Two types of bacterial HOs are known so far that show different regiospecificities toward the cleavage of the heme macrocycle. Most HOs cleave the heme at the {alpha}-meso carbon position yielding biliverdin IX{alpha}. The only HO showing a different regiospecificity is PigA from P. aeruginosa, which oxidizes heme to {beta}- and {delta}-biliverdin.

 

   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Reagents
All chemicals, including glutathione-agarose, were purchased from Sigma (Munich, Germany) and were American Chemical Society grade or better. Restriction enzymes were from Invitrogen (Cleveland, OH). MasterTaqTM was purchased from Eppendorf Scientific (Westbury, NY). HPLC-grade acetone and 80% formic acid were purchased from Fisher Scientific (Pittsburgh, PA). The expression vector pGEX-6P-1 and PreScissionTM protease were obtained from Amersham Biosciences. Centricon-10 concentrator devices were purchased from Amicon (Beverly, MA). Cytochrome P450 reductase was from Merck Biosciences (Bad Soden, Germany), all ferredoxins, ferredoxin-NADP+ oxidoreductase, and catalase were from Sigma.

Construction of Expression Vectors
P. aeruginosa PAO I pigA and bphO (PA4116) genes were amplified from chromosomal DNA via polymerase chain reaction with a hot start protocol using the following primers, which contained the indicated and underlined restriction sites: pigABamHIfwd: 5'-CGGGATCCGATACCCTGGCCCCTGAATCC-3', pigAXhoIrev: 5'-CCGCTCGAGTCAGGCGAAGGTACGCTCCAG-3'; bphOBamHIfwd: 5'-GCGGATCCATCTCCCCATCTCCATCGCCAGC-3', bphOSmaIrev: 5'-TCCCCCGGGTCAGGCTTCGTCGAGAACCCATC-3'. The polymerase chain reaction products were then cut with the indicated enzymes and inserted into similarly restricted pGEX-6P-1. The integrity of the plasmid constructs was verified by complete DNA sequence determination of the insert (Davis Sequencing, Davis, CA). Both constructs placed the heme oxygenase gene downstream of and in frame with the glutathione S-transferase (GST) gene of Schistosoma japonicum under the control of a Ptac promoter. A recognition sequence for PreScission protease is located upstream of the cloned gene. Proteolytic cleavage yields the native protein with a small N-terminal extension.

Expression and Purification
Expression and purification were performed according to instructions supplied by the manufacturer (Amersham Biosciences) and as described before (12). Recombinant P. aeruginosa BphP was expressed using a tet promoter-driven Strep-tag system (13) (IBA, Göttingen, Germany). Details of the cloning and purification will be published elsewhere.

Protein Determination
Protein concentration was determined by the Bradford method with bovine serum albumin as a standard or by measuring the absorbance at 280 nm and using the calculated {epsilon}280 nm for each individual protein (14, 15).

Spectrophotometric Analysis of Biliverdin Production
The formation of BV IX{alpha} was detected spectrophotometrically at 450 nm by converting it to bilirubin IX{alpha} (BR) using recombinant rat biliverdin reductase (BVR) (16). An aliquot of ~5 µg of crude recombinant rat BVR (expression clone was a gift of Dr. J. C. Lagarias, University of California at Davis) was added to the purified BphO·BV complex in 25 mM HEPES-KOH, 100 mM KCl, 10% glycerol, pH 7.5, and the reaction was started by adding an NADPH-regenerating system containing 6.5 mM glucose 6-phosphate, 0.82 mM NADP+, and 1.1 unit/ml glucose-6-phosphate dehydrogenase.

Reconstitution of BphO with Heme
The BphO·heme complex was prepared according to protocols previously described for heme·heme oxygenase complexes (3, 17). Heme was added to purified protein at a final ratio 3:1 heme:protein. Aliquots of heme (1.0–30 µM) were added to the cuvette containing 10 µM BphO and to the reference cuvette at room temperature. Spectra were recorded 5 min after heme addition.

Determination of the Extinction Coefficient for the BphO·heme Complex
The millimolar extinction coefficient at 409 nm for the BphO·heme complex was determined using the pyridine hemochrome method (2, 18). The isolated BphO·heme complex was converted to a pyridine hemochrome in a 500-µl sample volume by the addition of 62.5 µl pyridine and 62.5 µl of 0.5 N NaOH solution. The spectrum of the oxidized pyridine hemochrome was recorded. An excess of dithionite was added, and the spectrum of the reduced ferrous pyridine hemochrome was recorded. The spectrum of the reduced form minus the spectrum of the oxidized form was calculated at 557 nm. The concentration of heme was determined from the extinction coefficient using an {epsilon}405 nm value of 34,530 M–1 cm–1.

Spectroscopic Characterization of the Catalytic Turnover of the BphO·heme Complex
The BphO·heme complex (1:3) was prepared as described above and applied to a hydroxylapatite column (0.5 x 1.0 cm) equilibrated with 10 mM potassium phosphate buffer (pH 7.4). The column was washed with the same buffer, and the protein eluted in 200 mM potassium phosphate buffer (pH 7.4). The fractions containing the BphO·heme complex were dialyzed against 25 mM HEPES-KOH containing 100 mM KCl and 10% glycerol (pH 7.5).

The standard assay (0.5-ml final volume) contained 10 µM BphO·heme, 0.15 mg/ml bovine serum albumin, 4.6 µM ferredoxin (Spinacea, Porphyra, and Clostridium), 0.025 unit/ml spinach ferredoxin-NADP+ oxidoreductase in 25 mM HEPES-KOH buffer (pH 7.5). The reaction was started by addition of the NADPH-regenerating system listed above. Spectral changes between 300 and 800 nm were monitored for 30 min at 37 °C. Sodium ascorbate or Trolox (6-hydroxy-2,5,7,8-tetramethylchroman-2-carboxylic acid) was added to a final concentration of 5 mM unless otherwise noted. The reaction in the presence of human cytochrome P450 reductase (hCPR) was carried out as described previously by Wilks and Schmitt (3). When appropriate, catalase from Aspergillus niger was added to reaction cuvettes at a ratio of catalase:BphO·heme 0.5:1 immediately before the addition of reductant.

HPLC Analysis of BphO Reaction Products
BphO/PigA assay mixtures were loaded onto a Waters (Milford, MA) C18 Sep-Pak Light cartridge preconditioned as follows: 3-ml wash with acetonitrile to wet the Sep-Pak, 3-ml wash with MilliQ water, and 3-ml wash with 50 mM 4-methylmorpholine/glacial acetic acid (pH 7.7). After the sample was loaded onto the Sep-Pak, it was washed with 3 ml of 4-methylmorpholine/glacial acetic acid (pH 7.7) followed by 3 ml of 0.1% (v/v) trifluoroacetic acid. The tetrapyrroles were then eluted from the Sep-Pak using 2 ml of 100% acetonitrile. The eluate was dried using a Speed-Vac lyophilizer, and the dried samples were analyzed by HPLC. Samples were first dissolved in 10 µl of Me2SO and then diluted with 200 µl of the HPLC mobile phase. Following brief centrifugation and filtration through a 0.45-µm polytetrafluoroethylene syringe filter, bilins were resolved by reversed phase chromatography using an Agilent Technologies 1100 liquid chromatograph. The HPLC column used for all of the analyses was a 4.6- x 250-mm Phenomenex Ultracarb 5-µm ODS20 analytical column with a 4.6- x 30-mm guard column of the same material. The mobile phase consisted of acetone:20 mM formic acid (50:50 by volume), and the flow rate was 0.6 ml/min. Eluates were monitored at 650 and 380 nm using an Agilent Technologies 1100 series diode array detector.

NMR Spectroscopic Characterization of BphO
Preparation of BphO Reconstituted with 13C-Labeled Heme—13C-Labeled {delta}-aminolevulinic acids (ALAs) were used as biosynthetic precursors for the preparation of protoheme IX (heme). {delta}-[5-13C]Aminolevulinic acid ([5-13C]ALA) and [1,2-13C]ALA were synthesized utilizing methods described previously (19). Heme labeled with 13C was obtained utilizing previously reported methodology (9, 20–22) developed to take advantage of the fact that the first committed precursor in heme biosynthesis is ALA (23). Thus, 13C-labeled heme, which was biosynthesized in Escherichia coli upon addition of suitably labeled ALA, was trapped by simultaneously expressing rat liver outer mitochondrial membrane cytochrome b5. The details of the biosynthetic protocol, which entail the expression and purification of outer mitochondrial membrane cytochrome b5 harboring 13C-labeled heme, have been presented previously (9, 20–22). 13C-Labeled heme was extracted from outer mitochondrial membrane cytochrome b5 as follows: pyridine (15 ml) was added to 2.5 ml of 1 mM outer mitochondrial membrane cytochrome b5 in phosphate buffer (µ = 0.10, pH = 7.0). Slow addition of chloroform, typically 10–12 ml, resulted in the precipitation of the polypeptide, while maintaining the pyridine hemochrome in the supernatant. The latter was subsequently separated from the denatured polypeptide by centrifugation and then dried over anhydrous MgSO4. The desiccant was removed by filtration, and the filtered pyridine-chloroform solution was transferred to a round bottom flask, where it was concentrated to dryness in a rotary evaporator. Finally, the solid was redissolved in 1.5 ml of Me2SO. BphO was reconstituted with a freshly prepared solution of 13C-labeled heme by titrating it into a 25-ml solution of phosphate buffer (µ = 0.10, pH = 7.0) containing ~2 µmol of BphO until the ratio A280/ASoret no longer changed. The resultant solution containing the reconstituted enzyme was incubated at 4 °C overnight, then purified by size exclusion chromatography with the aid of a Sephadex G-50 column (90 x 2.6 cm inner diameter) equilibrated and eluted with 10 mM phosphate buffer, pH 7.0 at 4 °C. Fractions containing pure protein were concentrated in Amicon centrifugal concentrators to ~1 ml and then transferred to smaller Centricon concentrators to exchange the protein into deuterated phosphate buffer, pH 7.0, not corrected for the deuterium isotope effect.

NMR Spectroscopy—1H and 13C NMR spectra were acquired on a Varian Unity Inova spectrometer operating at frequencies of 599.740 (1H) and 150.817 (13C) MHz, respectively. 1H spectra were referenced to the residual water peak at 4.8 ppm, and 13C spectra were referenced to an external solution of dioxane (60% v/v in D2O) at 66.66 ppm. Proton spectra were acquired with pre-saturation of the residual water peak over 10,000 data points, with a spectral width of 24 kHz, a 150-ms acquisition time, a 200-ms relaxation delay, and 1024 scans. 13C spectra were collected over 12,000 data points, with a spectral width of 60 kHz, a 100-ms acquisition time, a 25-ms relaxation delay, and 400,000 scans. HMQC spectra were typically acquired with spectral widths of 24 kHz for 1H and 50 kHz for 13C and a 200-ms relaxation delay. HMQC spectra obtained from samples containing BphO reconstituted with heme labeled at the methyl, propionate-{beta}, and vinyl-{beta} carbons were acquired with refocusing delays based on 1JCH = 140 Hz, whereas spectra obtained from BphO reconstituted with heme labeled at the meso carbons were acquired with 1JCH = 180 Hz. Data were collected as an array of 2,000 x 128 points with 512 scans per t1 increment and processed by zero filling once in t2 and twice in t1 to obtain an 8,000 x 8,000 matrix. This was apodized with a 90°-shifted squared sine bell function and Fourier transformed. Water-eliminated Fourier transform-NOESY (24, 25) spectra were acquired with 24 kHz in both dimensions, 2,000 data points in t2, 256 increments in t1, 352 scans per t1 increment, and (typically) a 40-ms mixing time. The data were processed by zero filling in both dimensions to obtain an 8,000 x 8,000 matrix, apodized with 90°-shifted squared sine bell, and Fourier transformed.

Gel Permeation Chromatography
An Amersham Biosciences Superdex 200 HR10/30 gel permeation chromatography column was equilibrated in 50 mM TES-KOH buffer, pH 7.5, containing 100 mM KCl and 10% (v/v) glycerol (GPC buffer) at a flow rate of 0.4 ml/min. Standards with known Mr (i.e. {beta}-amylase 200,000; alcohol dehydrogenase 150,000; bovine serum albumin 66,000; carbonic anhydrase 29,000; and cytochrome c 12,600) were applied to the column (100 µg), and their elution volumes were determined spectroscopically at 280 nm. 10 µM BphP and 10 µM estimated BV (bound to BphO) were applied to the column and chromatographed under conditions identical to those used for the standard proteins. All samples were also monitored at 650 nm for BV detection.

Phytochrome Difference Spectroscopy
To test chromophore assembly of BphP in vitro, 20 µM recombinant apo-BphP was incubated with 40 µM chromophore or BphO (PigA)·BV complex for 30 min at room temperature in darkness. Absorbance spectra were obtained after 3-min incubation with red light at 630 nm (Pfr spectrum) and far red light at 750 nm (Pr spectrum), respectively, in a volume of 500 µl (50 mM HEPES-KOH, 20 mM KCl, pH 8.0) using an Agilent Technologies 8453 Value Analysis UV-Visible System. Difference spectra (Pr – Pfr) were calculated.

Coupled Oxidation of Heme
The four isomers of BV IX were obtained through coupled oxidation of heme in pyridine as described before (26). The four BV isomers were separated using a semi-preparative reversed phase HPLC of the same material as described above, except that the flow rate was 2 ml/min.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Expression and Purification of Recombinant P. aeruginosa Heme Oxygenases—The P. aeruginosa pigA and bphO genes were expressed using a tac promoter-driven N-terminal GST fusion expression system. All the reported results were obtained for recombinant BphO unless otherwise noted. Recombinant PigA and BphO could be purified to ~99% homogeneity, as shown in Fig. 2A for BphO. One single band was obtained on SDS-PAGE migrating at ~23,000, which corresponds to the predicted relative molecular weight calculated from the amino acid composition (Mr = 21,465). One liter of bacterial culture typically yielded 10 mg of purified BphO. Interestingly, expression of recombinant protein resulted in the development of a bright green color (Fig. 2B as seen by absorption at 650 nm) indicating the production of BV during the recombinant expression of BphO. The produced BV appears to have a high affinity toward BphO, because the BphO·BV complex remained stable throughout the entire purification process; however, the bound BV can be replaced by the substrate heme (see below). The production of BV was further confirmed by its conversion to bilirubin (BR) upon addition of rat biliverdin reductase (BVR). Hence, addition of BVR and NADPH to the purified BphO·BV complex resulted in the conversion of the bound BV to BR, which was accompanied by a color change from green to yellow and by the appearance of a peak at 450 nm (Fig. 2B). For additional confirmation that the production of BV was due to the action of BphO during expression, BphO was expressed under anaerobic culture conditions using nitrate as a terminal electron acceptor. Because any known HO reaction is oxygen-dependent, no formation of BV should be obtained. As expected, no color formation was observed during anaerobic expression and purification, but the spectrum of purified recombinant BphO showed a small peak around 420 nm suggesting the formation of a BphO·heme complex (Fig. 2B, inset). These observations taken together demonstrated that recombinant BphO is already active in E. coli, where it must interact with a reducing agent. The nature of this reductant, however, remains to be elucidated.



View larger version (35K):
[in this window]
[in a new window]
  		FIG. 2.
A, affinity purification of recombinant BphO. SDS-PAGE analysis of whole cell protein extracts before (lane 1) and after (lane 2) induction with IPTG. Lane 3 shows GST-BphO after glutathione agarose affinity chromatography; lane 4, PreScission Protease cleaved GST-BphO; lane 5, BphO after the second glutathione-agarose affinity chromatography; lane M, molecular mass standards. Numbers on the right indicate positions of molecular weight markers. B, absorbance spectroscopy of purified BphO. Electronic absorption spectra of recombinant BphO expressed and purified under aerobic conditions (solid line) and after addition of rat BVR (dashed line). The insert shows the BphO absorbance spectrum after anaerobic expression and purification.

 
Heme Binds to BphO—The formation of a BphO·heme complex was determined by difference spectroscopy. To this end, equal aliquots of hemin were added to the sample cuvette containing the BV complex of BphO in buffer and to the reference cuvette containing only buffer. Addition of hemin to BphO resulted in the replacement of BV, which was evident from a change in color of the solution, from green to brownish red. The electronic absorption spectrum of the BphO·heme complex is different from that of the BphO·BV complex and from that of free heme in solution (Fig. 3). The BphO·heme complex has a Soret band at 409 nm, which is close to that reported for other bacterial HO·heme complexes (27). Attempts to calculate the dissociation constant (Kd) did not result in reliable data, which is most likely a consequence of competition between BV and hemin for the binding site in BphO. Efforts to remove the relatively tightly bound BV from BphO prior to the titration experiments failed. The extinction coefficient of the BphO·heme complex was found to be 97.6 mM–1 cm–1, a value that appears to be the lowest reported for a bacterial HO.



View larger version (24K):
[in this window]
[in a new window]
  		FIG. 3.
Absorption difference spectra of heme binding to BphO. Increasing amounts of heme (1–30 µM) were added both to the reference and sample cuvettes.

 
Spectroscopic Characterization of the Catalytic Turnover of the BphO·heme Complex—The catalytic turnover of the ferric BphO·heme complex in the presence of spinach ferredoxin and a ferredoxin reductase system (i.e. ferredoxin-NADP+-oxidoreductase and a NADPH-regenerating system) was measured spectrophotometrically. In the presence of reduced spinach ferredoxin the BphO·heme complex was immediately converted to the oxyferrous complex, which was accompanied by a shift in the Soret band from 409 to 412 nm and by the appearance of {beta}/{alpha}-bands at 545 and 579 nm, respectively (Fig. 4A). Within 30 min the oxyferrous complex was fully degraded to ferric-BV (Fig. 4A). Interestingly, when the reaction mixture contains the hydrogen peroxide scavenger catalase in addition to Fd, to rule out non-enzymatic coupled oxidation of the BphO·heme complex, the reaction proceeded much slower and was arrested at the oxyferrous stage; the latter remained stable over at least 30 min (Fig. 4B). Because it has been reported that for some bacterial HOs the Fd-dependent conversion of heme to BV requires a second reductant such as ascorbate (28), ascorbate or Trolox was added to the arrested reaction. This resulted in the full conversion of the oxyferrous complex to iron-free BV, as was evident from the development of green color and from the appearance of a peak at 680 nm (Fig. 4C). In comparison, ascorbate by itself supported the oxidation of the BphO·heme complex to ferric (Fe3+-) BV (Fig. 4D). Addition of catalase to the ascorbate-mediated heme degradation did not alter the reaction.



View larger version (25K):
[in this window]
[in a new window]
  		FIG. 4.
Spectroscopic characterization of the catalytic turnover of the BphO·heme complex in the presence of spinach ferredoxin. A, catalytic turnover of BphO·heme in the presence of spinach Fd. The oxyferrous complex is formed immediately and degraded to ferric-BV. B, same reaction as A but in the presence of catalase. The reaction is arrested at the oxyferrous complex stage. C, same reaction as B. After addition of ascorbate the oxyferrous complex is further degraded to yield iron-free BV. D, catalytic turnover of BphO·heme in the presence of 5 mM sodium ascorbate. Initial spectra are shown in dashes, final spectra in bold. Spectra during this conversion are light gray. The direction of spectral changes are indicated by arrows.

 
To distinguish the observed spectral changes from a coupled oxidation reaction, the BphO·heme complex was incubated with 30 or 300 µM H2O2, respectively. Only the high concentration of H2O2 promoted the coupled oxidation reaction, which resulted in the formation of Fe3+-verdoheme (Table I). If the same reaction also contained catalase, no heme degradation was observed, which confirmed that in the presence of peroxides heme in the BphO·heme complex was degraded via coupled oxidation.


View this table:
[in this window]
[in a new window]
  		TABLE I
Tested reductants in the BphO assay

 
Reductant Specificity of BphO—The reductant specificity of BphO was tested using several commercially available reductants in our assay system. We tested the Fds from Clostridium sp. [4Fe-4S], Porphyra sp. [2Fe-2S], together with spinach ferredoxin-NADP+ oxidoreductase as well as human cytochrome P450 reductase (hCPR). All Fds tested were capable of supporting the degradation of the BphO·heme complex, whereas hCPR failed to mediate the oxidation of the BphO·heme complex. Furthermore, sodium ascorbate was a potent reducing partner (as described above). Experiments using a single reductant always resulted in the production of ferric-BV. Only if a second reductant was present iron-free BV was the final product. The results are summarized in Table I.

HPLC Analysis of the BphO and PigA Reaction Products— Upon completion of the Fd-mediated degradation of the BphO·heme complex, the reaction products were extracted using C18 SepPak cartridges and subsequently subjected to reverse phase HPLC. The four BV IX isomers were used as a standard as well as the reaction products of the PigA reaction. Fig. 5 shows a chromatogram confirming that BV IX{alpha} is the only product of the BphO reaction, whereas PigA produces the {beta}- and {delta}-isomers of BV IX.



View larger version (12K):
[in this window]
[in a new window]
  		FIG. 5.
HPLC analysis of the reaction products of the two P. aeruginosa heme oxygenases. Purified recombinant BphO and PigA were assayed for HO activity. Bilins were extracted from the incubation mixture using a SepPak C reversed phase cartridge and analyzed by reversed phase HPLC as 18described under "Experimental Procedures." The HPLC solvent was acetone:20 mM formic acid (50:50, v/v), and the eluate was monitored at 650 nm. The top trace shows the metabolites obtained by the BphO reaction; the bottom trace shows the products of the PigA reaction. Retention times for the four BV IX isomers are indicated by arrows.

 
NMR Spectroscopic Characterization of BphO—Chemical shifts for 1H and 13C nuclei originating from heme methyl and meso positions of cyanide-inhibited BphO (BphO-CN) were identified with a combination of one-dimensional non-de-coupled 13C and HMQC spectra, according to the assignment strategies outlined in previous work (9, 21, 22). The 1H resonance assignments corresponding to heme substituents were subsequently obtained from the NOESY map shown in Fig. 6. The significance of these assignments (summarized in Table II) is presented below.



View larger version (47K):
[in this window]
[in a new window]
  		FIG. 6.
Water-eliminated Fourier transform-NOESY (30 °C) spectrum of BphO-CN. Resonance assignments for heme substituents are obtained by following the dipolar connectivities starting from meso-H{delta} at 7.19 ppm and walking through the macrocycle in two directions: 1) meso-H{delta} -> 1Me -> 2V{beta} -> 2V{alpha} -> meso-H{alpha} -> 3Me -> 4V{beta} -> meso-H{beta} -> 5Me -> 6P{alpha}. The 4V{alpha} resonance can also be identified by its cross-peak with 4V{beta}; 2) meso-H{delta} -> 8Me -> 7P{alpha}.

 


View this table:
[in this window]
[in a new window]
  		TABLE II
1H and 13C chemical shifts (ppm) from heme substituents in cyanide-inhibited BphO at 30 °C

 
It has been proposed that the pattern of paramagnetically affected heme methyl resonances in the 1H NMR spectra of HO enzymes is diagnostic of the heme in-plane orientation, and hence regioselectivity of meso carbon oxidation (9, 21). For instance, the cyanide-inhibited forms of {alpha}-biliverdin-producing HO enzymes (HO-CN) exhibit 1H NMR spectra with only 1 heme methyl resonance above 12 ppm (3Me). Fig. 7 (A and B) illustrates the downfield portion of the 1H NMR spectra of the {alpha}-biliverdin producing HmuO-CN and HemO-CN, respectively, with only the 3Me resonance from its major (3Me) and minor (3me) heme orientational isomer resolved from the diamagnetic envelope of resonances (21, 29). The spectra of the {alpha}-biliverdin-producing human and rat HO-1-CN enzymes are essentially identical to those shown in Fig. 7 (A and B) (30, 31). In comparison, the 1H NMR spectrum of PigA-CN, where the heme is rotated in-plane ~100° relative to the heme of the {alpha}-biliverdin-producing enzymes, exhibits three heme methyl resonances from the more abundant heme orientational isomer (5Me, 1Me, and 8Me) and three from the less abundant isomer (me) (9) (see Fig. 7C). It is therefore evident from the resolved portion of the 1H NMR spectrum of BphO·CN (Fig. 7D), which only displays the methyl resonance corresponding to 3Me, that this enzyme should oxidize heme to produce {alpha}-biliverdin. Indeed, this prediction based on the diagnostic feature of the downfield resolved 1H NMR spectrum is in good agreement with the fact that assays of heme oxidation regio-selectivity yield only {alpha}-biliverdin.



View larger version (17K):
[in this window]
[in a new window]
  		FIG. 7.
Downfield portion of the 1H NMR spectra of cyanide inhibited HemO (A), HmuO (B), PigA (C), and BphO (D). The presence of only one heme methyl resonance in this region, 3Me, is diagnostic of all known {alpha}-hydroxylating HOs, where the proximal histidine-imidazole plane lies approximately parallel to the {beta}-{delta}-meso axis (see Fig. 8). The same region of the 1H NMR spectrum of PigA, where the heme is rotated in-plane ~100° relative to the {alpha}-biliverdin-producing HOs, shows three heme methyl resonances. These resonances indicate that the proximal histidine-imidazole plane in PigA lies approximately parallel to the {alpha}-{gamma}-meso axis (see Fig. 8).

 
The plot shown in Fig. 8 permits a straightforward correlation of the observed shifts for the four heme methyl groups as a function of the angle {phi} formed between the axial histidine-imidazole plane and the molecular x-axis (32). Interpretation of the heme methyl resonance assignments in the context of this plot made it possible to correctly predict an angle {phi} of 125° for the proximal imidazole of human heme oxygenase (32) before the x-ray crystal structure was solved (33). Moreover, interpretation of the heme methyl resonances from PigA in the context of the plot of Fig. 8 also made it possible to predict (9) that the heme in this enzyme is rotated in-plane ~110° relative to the heme in {alpha}-biliverdin HO enzymes before the x-ray crystal structure of PigA (34) confirmed this prediction. Hence, the order of the heme methyl resonances, 3Me > 8Me > 5Me > 1Me (see Table II) suggests that the proximal histidine-imidazole plane in BphO forms an angle of ~133° with respect to the molecular x-axis, as defined in Fig. 8. This indicates that the proximal histidine-imidazole plane in BphO lies almost parallel to the {beta}-{delta}-meso axis (shown schematically in Fig. 8) as is the case for all {alpha}-biliverdin producing HOs whose structure is known (human (33) and rat (35) HO-1, HemO (36), and HmuO (37)).



View larger version (32K):
[in this window]
[in a new window]
  		FIG. 8.
Plot illustrating the dependence of the heme methyl chemical shifts on the angle {phi} made by the projection of the proximal histidine-imidazole plane and the molecular x-axis (32, 49). In all {alpha}-biliverdin-producing HOs, including BphO, the proximal histidine-imidazole plane lies nearly parallel to the {delta}-{beta}-meso axis (bottom right). The order and position of the heme methyl chemical shifts in BphO indicate that the angle {phi} in this enzyme is ~133° (see plot), a value very similar to that seen in the crystal structures of HmuO (37), HemO (36), and HO-1 (33). The heme methyl chemical shifts of PigA have been used to predict (9) that its heme is rotated in-plane nearly 100° (top right) so that the proximal histidine-imidazole plane lies nearly parallel to the {alpha}-{gamma}-meso axis and forms an angle {phi} of ~35°. This prediction was corroborated in the recent crystal structure of PigA (34).

 
Interaction between HO·BV Complexes and BphP—It has been mentioned above that recombinant BphO purifies as a stable BphO·BV complex. The bound BV cannot be removed by gel filtration, ion exchange chromatography, or changes in solution pH, indicating that the release of the BV product is probably the rate-limiting step in this reaction and that BV has a high affinity toward BphO. The release of BV is known to be the rate-limiting step in other HOs. In the case of mammalian HOs the rate limitation is overcome by the presence of BVR, the subsequent enzyme in heme catabolism; the presence of BVR accelerates the release of BV (38). Because P. aeruginosa lacks a BVR homologue, another enzyme might be involved in the release of BV from BphO. Because the bphO gene is located in an operon together with the gene bphP, which encodes a bacterial phytochrome, we tested the interaction between recombinant BphO and BphP (a) to probe if BphO is able to produce the chromophore for BphP and (b) to investigate if the release of BV is facilitated through the interaction with BphP. Hence, purified BphO·BV was incubated with purified apo-BphP while monitoring changes in absorbance. The mixture was subsequently subjected to red- and far-red light-induced difference spectroscopy (39) to test for the formation of holo-BphP. As shown in Fig. 9A the spectrum of BphO·BV changes upon addition of BphP leading to the formation of a photoactive holo-BphP (Fig. 9B). To further confirm that BphO and BphP interact to transfer the BV produced by BphO to BphP, we subjected both proteins to gel permeation chromatography (GPC) to see whether we obtain any physical interaction between both proteins. As seen in Fig. 9C both proteins by themselves elute as single peaks on a Superdex 200 GPC column (upper two elution profiles). Monitoring the absorbance values at both 280 nm (protein) and 650 nm (BV) confirmed that BphO forms a very stable complex with BV. After incubation of both proteins in a 1:1 ratio and subsequent chromatography, we were still able to separate both proteins from each other. However, a major portion of the absorbance at 650 nm (i.e. BV) was then associated with BphP, indicating that BV was removed from BphO and was attached to BphP. This attachment was identified to be covalent by zinc blotting (40) (data not shown) and is in agreement with other known bacterial phytochromes that form a covalent complex with BV (11, 41). Experiments to identify real physical interaction between BphO and BphP using the GPC method after Hummel and Dreyer (42) with BphO·BV in the mobile phase did not reveal a stable complex of both proteins (data not shown).



View larger version (24K):
[in this window]
[in a new window]
  		FIG. 9.
Characterization of the interaction between HO·BV complexes with BphP. A, absorbance spectra of purified BphO·BV (dashed line), apo-BphP (dotted line), and a 1:1 mixture of both (solid line). The inset shows an enlargement of the region between 500 and 800 nm. B, red- and far-red light-induced difference spectroscopy of the 1:1 mixture of BphO·BV and apo-BphP. C, gel permeation chromatography of BphO and BphP using a Superdex 200 column. Absorbance values were monitored at 280 nm (protein; solid lines) and 650 nm (BV; dashed lines). Upper chromatogram, BphP; second chromatogram, BphO·BV; lower chromatogram, 1:1 mixture of BphP and BphO·BV. D, red- and far-red light-induced difference spectroscopy of the 1:1 mixture of PigA·BV and apo-BphP.

 
Because PigA, the second HO in P. aeruginosa, also purifies as a stable PigA·BV complex, similar experiments were performed using the PigA·BV complex. Incubation of PigA·BV with BphP also resulted in an absorbance change yielding a functional holo-BphP (Fig. 9D). In contrast to BphO, which only produced the {alpha}-isomer of BV IX, PigA produced two different isomers of BV IX. Because the purified PigA·BV complex most likely consisted of a mixture of PigA·BV IX{beta} and PigA·BV IX{delta}, it was necessary to establish which BV isomer is responsible for the obtained photoactive phytochrome. Therefore, we obtained each BV isomer through coupled oxidation of heme and tested whether they are able to bind to apo-BphP. As shown in Fig. 10 only the BV isomers IX{alpha} and IX{delta} are able to form a photoactive holo-phytochrome. The {Delta}{Delta}A of the obtained difference spectrum with BV IX{delta} is ~one-fifth of that with BV IX{alpha} implying a lower affinity of BV IX{delta} toward BphP. Furthermore, it was shown that both isomers ({alpha} and {delta}) form a covalent bond with BphP (data not shown). These results indicate that the obtained difference spectrum after incubation of PigA·BV with BphP is due to the covalent attachment of BV IX{delta} to BphP.



View larger version (10K):
[in this window]
[in a new window]
  		FIG. 10.
Difference spectroscopy of BphP incubated with the BV IX isomers produced by P. aeruginosa. A, BphP incubated with BV IX{alpha}; B, BphP incubated with BV IX{delta}; C, BphP incubated with BV IXb.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
bphO Encodes a Second Heme Oxygenase in P. aeruginosa— The Gram-negative opportunistic pathogen P. aeruginosa is the first example of an organism that harbors two heme oxygenases with different regiospecificity of heme cleavage. The PigA enzyme has already been well characterized and is known to be involved in the mining of heme-iron under the free-iron limiting conditions likely to occur during infection of a host cell (7). On the basis of 1H NMR and resonance Raman studies, it was recently determined that the unusual regiospecificity of PigA is due to a rotated seating of the heme substrate in the active site pocket (9). This observation was further confirmed by the recently published crystal structure of PigA (34). The biological reason for the production of BV IX{beta} and BV IX{delta}, however, remains unknown. In comparison, to date, there is only indirect evidence that bphO genes encode for functional HOs (11). In this report we present biochemical and biophysical evidence demonstrating that recombinant P. aeruginosa BphO is indeed a heme oxygenase that catalyzes the conversion of heme to BV IX{alpha}.

BphO Prefers Ferredoxins and Ascorbate as Reducing Partners—Most HOs are capable of utilizing an endogenous E. coli electron donor, which is manifested as a green color in E. coli cells expressing HOs (3, 27, 43). In vitro, however, the activity of most bacterial HOs is usually tested using either mammalian CPR or ascorbate as an electron donor. E. coli possesses several reductants that could possibly serve as electron donors; among them are several Fds and flavodoxin (44). We therefore tested commercially available Fds, as well as ascorbate, for their ability to serve as electron donors for the P. aeruginosa HOs. Indeed, we were able to show that reduced Fds can serve as the electron donor in the BphO reaction, yielding ferric-BV as the final product. In the presence of catalase the conversion from heme to ferric-BV was arrested at the stage of the oxyferrous complex, suggesting that the conversion of the oxyferrous complex to ferric-BV is partly due to coupled oxidation. Fd has a sufficiently negative reduction potential to reduce O2 to superoxide (45); hydrogen peroxide is then spontaneously formed through dismutation of the superoxide radical. The presence of H2O2, which is known to react with FeII-heme to hydroxylate the heme (46), is scavenged by the presence of catalase. Only when a second reductant such as ascorbate is added to the reaction mixture, is the oxyferrous complex oxidized to BV, thus indicating that the reaction is catalyzed by BphO. The requirement for a second reductant in addition to Fd is in agreement with observations described for HO from algae, plants, and cyanobacteria (28, 47). Interestingly, it has been shown that the cyanobacterial HO-1 reaction is also terminated at the oxyferrous complex stage with cytochrome P450 reductase (27) indicating that this is a stable intermediate and that the following steps in the catalytic cycle might require a second reductant. Alternatively, it is also possible that the true physiological reductant in P. aeruginosa is capable of supporting the complete catalytic cycle. In contrast to other described bacterial HOs, hCPR was unable to mediate the conversion of heme to BV by BphO in our assay system, indicating that with regard to the redox partner BphO is quite different from other bacterial HOs and more similar to those described for cyanobacteria and plants. In agreement with many other described HOs, ascorbate is also capable of serving as an electron donor in the BphO reaction. Because catalase has no effect in the ascorbate-supported reaction, no coupled oxidation is involved during catalysis. Therefore it appears that ascorbate is the most efficient reductant tested for the in vitro activity of BphO. The origins of the endogenous reductant in E. coli and the one supporting the activity in P. aeruginosa remain to be elucidated.

The Second HO in P. aeruginosa, BphO, is an {alpha}-Meso-hydoxylating Enzyme—PigA was initially reported to oxidize heme to produce mostly {beta}-biliverdin (7). Subsequent studies established that PigA oxidizes heme to a mixture of {beta}- (30%) and {delta}- (70%) biliverdin (9). Further, NMR spectroscopic studies revealed that the heme in PigA is rotated in-plane ~100° relative to the heme in all {alpha}-hydroxylating heme oxygenase enzymes (9) (see Fig. 11A). This in-plane rotation locates the {delta}-meso carbon within the fold of PigA in the same place where {alpha}-biliverdin forming HOs place the {alpha}-meso carbon (9) (see Fig. 11B). Hence, attack of the {delta}-meso carbon in PigA leads to the formation of 70% {delta}-biliverdin. Inspection of Fig. 11A also reveals that a 180° rotation of the heme about the {alpha}-{gamma}-meso axis, a phenomenon common in heme proteins where the heme is not covalently attached to the polypeptide (48), places the {beta}-meso carbon where it is attacked, thus explaining the formation of 30% {beta}-biliverdin (9). More recently it has been shown that the heme, in one of the heme orientational isomers of the R177E mutant of C. diphtheriae HmuO, rotates ~85° and places the {beta}-meso carbon in the place typically occupied by the {alpha}-meso carbon, which results in the formation of ~50% {beta}-biliverdin (21). Results obtained from the NMR spectroscopic analysis of BphO strongly suggest that the fold of BphO is likely very similar to that of the other heme oxygenase enzymes. Moreover, these observations also imply that one of the most important structural differences between the two heme oxygenase enzymes in P. aeruginosa is the in-plane conformation of the heme, with the heme in PigA being rotated ~110° relative to that of the heme in BphO. This different in-plane conformation of the heme allows BphO to locate the {alpha}-meso carbon where it can be hydroxylated by the FeIII-OOH intermediate.



View larger version (46K):
[in this window]
[in a new window]
  		FIG. 11.
A, the in-plane heme orientation in PigA places the {delta}-meso carbon within the fold where it is susceptible to oxidation. A 180° rotation of the heme about the {alpha}-{gamma}-meso carbon places the {beta}-meso carbon where it is susceptible to oxidation (9, 34). B, the heme in {alpha}-biliverdin producing HOs (HemO is used as an example (36)) places the {alpha}-meso carbon in a location equivalent to that of the {delta}-meso carbon in PigA, thus resulting in {alpha}-meso carbon oxidation.

 
Interaction between BphO and BphP Facilitates the Release of BV from BphO—Single turnover experiments of human HO-1 have revealed that the release of BV from HO-1 is the rate-limiting step in the conversion from heme to BV (38). In the presence of BVR, this step is accelerated and one of the earlier electron transfer steps becomes rate-limiting. The data presented here indicate that the bacterial phytochrome BphP from P. aeruginosa whose gene is encoded in the same operon as bphO is able to interact with the BphO·BV complex and facilitates the release of BV. The BV is "pulled" out of the HO and forms a stable covalent adduct with BphP, which was confirmed by difference spectroscopy and zinc blotting. BphP is not only able to use iron-free BV as a substrate (as shown with purified BphO·BV complex) but also the Fe3+-BV, which is the product of the Fd-mediated reaction (data not shown). These results indicate that BphO produces the chromophore for the bacterial phytochrome BphP and that the activity of BphO might be regulated through the interaction between BphO and BphP. The biological role of BphP still remains to be elucidated.

In addition to the BphO-BphP interaction, we were able to show that BphP also facilitates the release of BV from PigA. From reports by other groups it is known that an A-ring endo vinyl-group of the BV is necessary for binding to BphP (41). Therefore, only the BV IX{delta} isomer should yield a photoactive holo-BphP. Indeed, using the BV isomers obtained from a coupled oxidation reaction, we were able to show that only BV IX{delta} binds to BphP. The attachment was shown to be covalent. This indicates that the release of the bound BV from the HOs is determined by the type of bound BV rather than by the nature of the protein. Therefore, the holo-BphP obtained upon incubation of PigA·BV with BphP arises from the release of BV IX{delta} from PigA. Because PigA in vitro produces the BV isomers in a ratio of 30:70 (IX{beta}:IX{delta}) (9), BV IX{delta} is a possible chromophore for BphP, although the amount of resultant holo-BphP is only one-fifth that of the BV IX{alpha}-BphP adduct. This could be due to a lower affinity of BV IX{delta} toward BphP. Therefore, the function of this photoactive phytochrome might be different from that with bound BV IX{alpha}. Although we were able to show in this study that BphP is able to bind BV IX{delta}, its physiological significance remains to be proven, because PigA is only expressed under iron-limiting conditions, and it is presently unknown if BphP is expressed under these conditions as well. The role of the interaction with BphP in controlling enzymatic activity of the two HOs and the physiological consequences of the formed holo-BphP are subjects of continuing efforts in our laboratories.


   FOOTNOTES
 
* This work was supported by the Emmy-Noether-Program of the Deutsche Forschungsgemeinschaft and funds from the Fonds der Chemischen Industrie (to N. F.-D.) and by Grant GM 50503 from the National Institutes of Health (to M. R.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

To whom correspondence should be addressed: Tel.: 49-531-391-5815; Fax: 49-531-391-5854; E-mail: n.frankenberg{at}tu-bs.de.

1 The abbreviations used are: HO, heme oxygenase; BR, bilirubin; BV, biliverdin; BVR, biliverdin reductase; Fd(s), ferredoxin(s); Fur, ferric uptake regulator; GPC, gel permeation chromatography; GST, glutathione S-transferase; (h)CPR, (human) cytochrome P450 reductase; HPLC, high performance liquid chromatography; ALA, {delta}-aminolevulinic acid; HMQC, heteronuclear multiple quantum coherence; TES, 2-{[2-hydroxy-1,1-bis(hydroxymethyl)ethyl]amino}ethanesulfonic acid; NOESY, nuclear Overhauser effect spectroscopy. Back


   ACKNOWLEDGMENTS
 
The cloning and initial experiments of this project were started in the laboratory of J. Clark Lagarias (University of California at Davis). His encouragement and continuous support as well as the gift of a rat BVR expression clone is greatly acknowledged. We thank Paul Ortiz de Montellano for the gift of an hCPR expression clone. Thanks are also due to Angela Wilks for helpful discussions.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Maines, D. M. (1988) FASEB J. 2, 2557–2568[Abstract]
  2. Wilks, A. (2002) Antiox. Redox Signal. 4, 603–614[CrossRef][Medline] [Order article via Infotrieve]
  3. Wilks, A., and Schmitt, M. P. (1998) J. Biol. Chem. 273, 837–841[Abstract/Free Full Text]
  4. Skaar, E. P., Gaspar, A. H., and Schneewind, O. (2004) J. Biol. Chem. 279, 436–443[Abstract/Free Full Text]
  5. Zhu, W., Hunt, D. J., Richardson, A. R., and Stojiljkovic, I. (2000) J. Bacteriol. 182, 439–447[Abstract/Free Full Text]
  6. Schmitt, M. (1997) J. Bacteriol. 179, 838–845[Abstract/Free Full Text]
  7. Ratliff, M., Zhu, W., Deshmukh, R., Wilks, A., and Stojiljkovic, I. (2001) J. Bacteriol. 183, 6394–6403[Abstract/Free Full Text]
  8. Vasil, M. L., and Ochsner, U. A. (1999) Mol. Microbiol. 34, 399–413[CrossRef][Medline] [Order article via Infotrieve]
  9. Caignan, G. A., Deshmukh, R., Wilks, A., Zeng, Y., Huang, H. W., Moenne-Loccoz, P., Bunce, R. A., Eastman, M. A., and Rivera, M. (2002) J. Am. Chem. Soc. 124, 14879–14892[CrossRef][Medline] [Order article via Infotrieve]
  10. Davis, S. J., Vener, A. V., and Vierstra, R. D. (1999) Science 286, 2517–2520[Abstract/Free Full Text]
  11. Bhoo, S.-H., Davis, S. J., Walker, J., Karniol, B., and Viestra, R. D. (2001) Nature 414, 776–779[CrossRef][Medline] [Order article via Infotrieve]
  12. Frankenberg, N., Mukougawa, K., Kohchi, T., and Lagarias, J. C. (2001) Plant Cell 13, 965–978[Abstract/Free Full Text]
  13. Skerra, A. (1994) Gene (Amst.) 151, 131–135[CrossRef][Medline] [Order article via Infotrieve]
  14. Bradford, M. M. (1976) Anal. Biochem. 72, 248–254[CrossRef][Medline] [Order article via Infotrieve]
  15. Gill, S. C., and von Hippel, P. H. (1989) Anal. Biochem. 182, 319–326[CrossRef][Medline] [Order article via Infotrieve]
  16. Muramoto, T., Kohchi, T., Yokota, A., Hwang, I., and Goodman, H. M. (1999) Plant Cell 11, 335–348[Abstract/Free Full Text]
  17. Yoshida, T., and Kikuchi, G. (1978) J. Biol. Chem. 253, 4224–4229[Free Full Text]
  18. Berry, E. A., and Trumpower, B. L. (1987) Anal. Biochem. 161, 1–15[CrossRef][Medline] [Order article via Infotrieve]
  19. Bunce, R. A., Shilling, C. L., III, and Rivera, M. (1997) J. Labelled Compd. Radiopharm. 39, 669–675[CrossRef]
  20. Rivera, M., and Walker, F. A. (1995) Anal. Biochem. 230, 295–302[Medline] [Order article via Infotrieve]
  21. Zeng, Y., Deshmukh, R., Caignan, G. A., Bunce, R. A., Rivera, M., and Wilks, A. (2004) Biochemistry 43, 5222–5238[CrossRef][Medline] [Order article via Infotrieve]
  22. Rivera, M., and Caignan, G. A. (2004) Anal. Bioanal. Chem. 378, 1464–1483[Medline] [Order article via Infotrieve]
  23. Warren, M. J., and Scott, A. I. (1990) Trends Biochem. Sci. 15, 426
  24. Patt, S. L., and Sykes, B. D. (1972) J. Chem. Phys. 56, 3182–3184[CrossRef]
  25. Lankhorst, P. P., Wille, G., van Boom, J. H., Altona, C., and Haasnoot, C. A. G. (1983) Nucleic Acids Res. 11, 2839–2856[Abstract/Free Full Text]
  26. Bonnett, R., and McDonagh, A. F. (1973) J. Chem. Soc. 9, 881–888
  27. Migita, C. T., Zhang, X., and Yoshida, T. (2003) Eur. J. Biochem. 270, 687–698[Medline] [Order article via Infotrieve]
  28. Cornejo, J., Willows, R. D., and Beale, S. I. (1998) Plant J. 15, 99–107[CrossRef][Medline] [Order article via Infotrieve]
  29. Li, Y., Syvitski, R. T., Auclair, K., Wilks, A., Ortiz de Montellano, P. R., and La Mar, G. N. (2002) J. Biol. Chem. 277, 33018–33031[Abstract/Free Full Text]
  30. Hernandez, G., Wilks, A., Paolesse, R., Smith, K. M., Ortiz de Montellano, P. R., and La Mar, G. N. (1994) Biochemistry 33, 6631–6641[CrossRef][Medline] [Order article via Infotrieve]
  31. Gorst, C. M., Wilks, A., Yeh, D. C., Ortiz de Montellano, P. R., and La Mar, G. N. (1998) J. Am. Chem. Soc. 120, 8875–8884[CrossRef]
  32. Shokhirev, N. V., and Walker, F. A. (1998) J. Biol. Inorg. Chem. 3, 581–594[CrossRef]
  33. Schuller, D. J., Wilks, A., Ortiz de Montellano, P. R., and Poulos, T. L. (1999) Nat. Struct. Biol. 6, 860–867[CrossRef][Medline] [Order article via Infotrieve]
  34. Friedman, J., Lad, L., Li, H., Wilks, A., and Poulos, T. L. (2004) Biochemistry 43, 5239–5245[CrossRef][Medline] [Order article via Infotrieve]
  35. Sugishima, M., Omata, Y., Kakuta, Y., Sakamoto, H., Noguchi, M., and Fukuyama, K. (2000) FEBS Lett. 471, 61–66[CrossRef][Medline] [Order article via Infotrieve]
  36. Schuller, D. J., Zhu, W. M., Stojiljkovic, I., Wilks, A., and Poulos, T. L. (2001) Biochemistry 40, 11552–11558[CrossRef][Medline] [Order article via Infotrieve]
  37. Hirotsu, S., Chu, G. C., Unno, M., Lee, D.-S., Yoshida, T., Park, S.-Y., Shiro, Y., and Ikeda-Saito, M. (2004) J. Biol. Chem. 279, 11937–11947[Abstract/Free Full Text]
  38. Liu, Y., and Ortiz de Montellano, P. R. (2000) J. Biol. Chem. 275, 5297–5307[Abstract/Free Full Text]
  39. Terry, M. J., and Lagarias, J. C. (1991) J. Biol. Chem. 266, 22215–22221[Abstract/Free Full Text]
  40. Berkelman, T. R., and Lagarias, J. C. (1986) Anal. Biochem. 156, 194–201[CrossRef][Medline] [Order article via Infotrieve]
  41. Lamparter, T., Michael, N., Caspani, O., Miyata, T., Shirai, K., and Inomata, K. (2003) J. Biol. Chem. 278, 33786–33792[Abstract/Free Full Text]
  42. Hummel, J. P., and Dreyer, W. J. (1962) Biochim. Biophys. Acta 63, 530–532[Medline] [Order article via Infotrieve]
  43. Zhang, X., Sato, M., Sasahara, M., Migita, C. T., and Yoshida, T. (2004) Eur. J. Biochem.
  44. Blattner, F. R., Plunkett, G., Bloch, C. A., Perna, N. T., Burland, V., Riley, M., ColladoVides, J., Glasner, J. D., Rode, C. K., Mayhew, G. F., Gregor, J., Davis, N. W., Kirkpatrick, H. A., Goeden, M. A., Rose, D. J., Mau, B., and Shao, Y. (1997) Science 277, 1453–1474[Abstract/Free Full Text]
  45. Arora, A., Sairam, R. K., and Srivastava, G. C. (2002) Curr. Sci. 82, 1227–1238
  46. Avila, L., Huang, H. W., Damaso, C. O., Lu, S., Moenne-Loccoz, P., and Rivera, M. (2003) J. Am. Chem. Soc. 125, 4103–4110[Medline] [Order article via Infotrieve]
  47. Muramoto, T., Tsurui, N., Terry, M. J., Yokota, A., and Kohchi, T. (2002) Plant Physiol. 130, 1958–1966[Abstract/Free Full Text]
  48. La Mar, G., N., Burns, P. D., Jackson, J. T., Smith, K. M., Langry, K. C., and Strittmatter, P. (1981) J. Biol. Chem. 256, 6075–6079[Free Full Text]
  49. Bertini, I., Luchinat, C., Parigi, G., and Walker, F. A. (1999) J. Biol. Inorg. Chem. 4, 515–519[Medline] [Order article via Infotrieve]

Add to CiteULike CiteULike   Add to Complore Complore   Add to Connotea Connotea   Add to Del.icio.us Del.icio.us   Add to Digg Digg   Add to Reddit Reddit   Add to Technorati Technorati    What's this?


This article has been cited by other articles:


Home page
 J. Biol. Chem.Home page
A. P. Kaur, I. B. Lansky, and A. Wilks
The Role of the Cytoplasmic Heme-binding Protein (PhuS) of Pseudomonas aeruginosa in Intracellular Heme Trafficking and Iron Homeostasis
J. Biol. Chem., January 2, 2009; 284(1): 56 - 66.
[Abstract] [Full Text] [PDF]

Home page
 J. Biol. Chem.Home page
W. C. Lee, M. L. Reniere, E. P. Skaar, and M. E. P. Murphy
Ruffling of Metalloporphyrins Bound to IsdG and IsdI, Two Heme-degrading Enzymes in Staphylococcus aureus
J. Biol. Chem., November 7, 2008; 283(45): 30957 - 30963.
[Abstract] [Full Text] [PDF]

Home page
 J. Biol. Chem.Home page
M. D. L. Suits, N. Jaffer, and Z. Jia
Structure of the Escherichia coli O157:H7 Heme Oxygenase ChuS in Complex with Heme and Enzymatic Inactivation by Mutation of the Heme Coordinating Residue His-193
J. Biol. Chem., December 1, 2006; 281(48): 36776 - 36782.
[Abstract] [Full Text] [PDF]

Home page
 J. Bacteriol.Home page
S. Puri and M. R. O'Brian
The hmuQ and hmuD Genes from Bradyrhizobium japonicum Encode Heme-Degrading Enzymes.
J. Bacteriol., September 1, 2006; 188(18): 6476 - 6482.
[Abstract] [Full Text] [PDF]

Home page
 J. Biol. Chem.Home page
I. B. Lansky, G. S. Lukat-Rodgers, D. Block, K. R. Rodgers, M. Ratliff, and A. Wilks
The Cytoplasmic Heme-binding Protein (PhuS) from the Heme Uptake System of Pseudomonas aeruginosa Is an Intracellular Heme-trafficking Protein to the {delta}-Regioselective Heme Oxygenase
J. Biol. Chem., May 12, 2006; 281(19): 13652 - 13662.
[Abstract] [Full Text] [PDF]

Home page
 Plant Physiol.Home page
T. J. Emborg, J. M. Walker, B. Noh, and R. D. Vierstra
Multiple Heme Oxygenase Family Members Contribute to the Biosynthesis of the Phytochrome Chromophore in Arabidopsis
Plant Physiology, March 1, 2006; 140(3): 856 - 868.
[Abstract] [Full Text] [PDF]

Home page
 J. Bacteriol.Home page
E. P. Skaar, A. H. Gaspar, and O. Schneewind
Bacillus anthracis IsdG, a Heme-Degrading Monooxygenase
J. Bacteriol., February 1, 2006; 188(3): 1071 - 1080.
[Abstract] [Full Text] [PDF]

Home page
 Proc. Natl. Acad. Sci. USAHome page
M. D. L. Suits, G. P. Pal, K. Nakatsu, A. Matte, M. Cygler, and Z. Jia
Identification of an Escherichia coli O157:H7 heme oxygenase with tandem functional repeats
PNAS, November 22, 2005; 102(47): 16955 - 16960.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
279/44/45791    most recent
M408303200v1
Right arrow Submit a Letter to Editor
Right arrow Alert me when this article is cited
Right arrow Alert me when eLetters are posted
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowRequest Permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Wegele, R.
Right arrow Articles by Frankenberg-Dinkel, N.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Wegele, R.
Right arrow Articles by Frankenberg-Dinkel, N.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg   Add to Reddit   Add to Technorati  
What's this?

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
 All ASBMB Journals   Molecular and Cellular Proteomics 
 Journal of Lipid Research   ASBMB Today 
Copyright © 2004 by the American Society for Biochemistry and Molecular Biology.
Advertisement
spacer
Advertisement
Advertisement