Tetsuhiro Tanaka

Michael Wiesener

Wanja Bernhardt

Kai-Uwe Eckardt

Dr Christina Warnecke
email: christina.warnecke@uk-erlangen.de

THE HUMAN HYPOXIA-INDUCIBLE FACTOR (HIF)-3α GENE IS A HIF-1 TARGET GENE AND MAY MODULATE HYPOXIC GENE INDUCTION

Keywords: ARNT, arylhydrocarbon receptor nuclear translocator, bHLH, basic helix-loop-helix green fluorescence protein, HIF, hypoxia-inducible factor, HRE, hypoxia-responsive element, IPAS, inhibitory PAS domain protein, NTAD, Nterminal transactivation transcriptional regulation, hypoxic response, promoter, RNA interference, renal carcinoma 2

SYNOPSIS

Hypoxia-inducible factor (HIF)-3α is the third member of the HIF transcription factor family. Whereas HIF-1α and -2α play critical roles in the cellular and systemic adaptation to hypoxia, little is known about the regulation and function of HIF-3α. At least five different splice variants may be expressed from the human HIF-3α locus, which are suggested to exert primarily negative regulatory effects on hypoxic gene induction. Here we report that hypoxia induces the human HIF-3α gene at the transcriptional level in a HIF-1-dependent manner. HIF-3α2 and HIF-3α4 transcripts, the HIF-3α splice variants expressed in Caki-1 renal carcinoma cells, rapidly increased after exposure to hypoxia or chemical hypoxia mimetics. SiRNA-mediated HIF-α knock-down demonstrated that HIF-3α is a specific target gene of HIF-1α, but not affected by HIF-2α knock-down. In contrast to HIF-1α and HIF-2α, HIF-3α is not regulated at the level of protein stability. HIF-3α protein could be detected under normoxia in cytoplasm and nuclei, but increased under hypoxic conditions. Promoter analyses and chromatin immunoprecipitation experiments localized a functional hypoxiaresponsive element 5´ to the transcriptional start of HIF-3α2. SiRNA-mediated knock-down of HIF-3α increased transactivation of a HIF-driven reporter construct and mRNA expression of lysyl oxidase. Immunohistochemistry revealed an overlap of HIF-1α-positive and HIF-3αpositive areas in human renal cell carcinomas. These findings shed light on a novel aspect of HIF-3α as a HIF-1 target gene and point to a possible role as a modulator of hypoxic gene induction.

INTRODUCTION

Of the three hypoxia-inducible transcription factor (HIF) family members identified in mammals so far, HIF-3 is the least characterized and understood. HIF-1 and -2 are heterodimeric transcription factors which mediate the systemic and cellular response to hypoxia and consist of two basic helix-loop-helix (bHLH) PAS (Per Arnt Sim) domain proteins, of which the α-subunit is oxygen-regulated and the β-subunit constitutively expressed (for review see [START_REF] Schofield | Signalling hypoxia by HIF hydroxylases[END_REF]). HIF-β is identical to the arylhydrocarbon receptor nuclear translocator (ARNT) and thus also part of transcription factor complexes involved in the xenobiotic response [START_REF] Hoffman | Cloning of a factor required for activity of the Ah (dioxin) receptor[END_REF]. Two of the alternative α-subunits, HIF-1α and HIF-2α, transactivate many functionally heterogeneous genes involved e.g. in anaerobic glucose metabolism, erythropoiesis and angiogenesis. HIF-1α and HIF-2α show significant structural homology and are primarily regulated at the protein level by oxygen-dependent destabilization, which is preceded by enzymatic hydroxylation of two conserved prolyl residues in their oxygen-dependent degradation domains. At least three prolyl hydroxylase domain proteins (PHD1-3) have been identified which modify the HIF-α subunits using dioxygen and 2-oxoglutarate as co-substrates. This modification enables binding of the von Hippel-Lindau tumor suppressor protein (pVHL), which targets the HIF-α subunits for proteasomal degradation. Under hypoxia, the HIF-α subunits are thus stabilized, dimerize with HIF-β, translocate to the nucleus and transactivate their target genes. In addition to regulation of protein stability the activity of HIF-1α is repressed in an oxygen-dependent manner by the asparagyl hydroxylase factor inhibiting HIF-1 (FIH-1) (for review see [START_REF] Schofield | Signalling hypoxia by HIF hydroxylases[END_REF]). The vast majority of HIF target genes is regulated by HIF-1α, whereas exclusively HIF-2α-dependent genes are scarce and cell type-dependent [START_REF] Warnecke | Differentiating the functional role of hypoxia-inducible factor (HIF)-1α and HIF-2α (EPAS-1) by the use of RNA interference: erythropoietin is a HIF-2α target gene in Hep3B and Kelly cells[END_REF][START_REF] Hu | Differential roles of hypoxia-inducible factor 1α (HIF-1α) and HIF-2α in hypoxic gene regulation[END_REF][START_REF] Raval | Contrasting properties of hypoxia-inducible factor 1 (HIF-1) and HIF-2 in von Hippel-Lindau-associated renal cell carcinoma[END_REF][START_REF] Elvidge | Concordant regulation of gene expression by hypoxia and 2-oxoglutaratedependent dioxygenase inhibition: the role of HIF-1α, HIF-2α, and other pathways[END_REF]. In 1998 a third HIF-α isoform was identified which exhibits about 55% amino acid sequence identity with HIF-1α and HIF-2α in its bHLH PAS domain [START_REF] Gu | Molecular characterization and chromosomal localization of a third α-class hypoxia inducible factor subunit, HIF3α[END_REF]. It also contained an oxygendependent degradation domain (ODD) and a N-terminal transactivation domain (NTAD), similar to HIF-1α and HIF-2α, but no CTAD. In 2001 the murine inhibitory PAS domain protein (IPAS) was shown to be a repressor of HIF-dependent gene induction by virtue of its ability to bind to HIF-1α [START_REF] Makino | Inhibitory PAS domain protein is a negative regulator of hypoxia-inducible gene expression[END_REF]. Subsequently, IPAS was demonstrated to be a splice variant originating from the murine HIF-3α locus. Highly expressed in the mouse brain and eye hypoxic IPAS induction impairs upregulation of vascular endothelial growth factor (VEGF) activity in the avascular cornea during sleep and thereby prevents angiogenesis [START_REF] Makino | Inhibitory PAS domain protein is a negative regulator of hypoxia-inducible gene expression[END_REF]. Interestingly IPAS, but not HIF-3α, is upregulated under hypoxia by differential splicing and transcriptional induction [START_REF] Ferla | Inhibition of erythropoietin gene expression signaling involves the transcription factors GATA-2 and NF-κB[END_REF][START_REF] Makino | Transcriptional up-regulation of inhibitory PAS domain protein gene expression by hypoxia-inducible factor 1 (HIF-1): a negative feedback regulatory circuit in HIF-1-mediated signaling in hypoxic cells[END_REF]. The human HIF-3α locus may give rise to at least five alternatively spliced transcripts, of which three (HIF-3α1-3) contain the ODD and NTAD but differ in their C-terminal and Nterminal ends, whereas the other splice variants lack the complete C-terminal half of the protein including ODD and NTAD [START_REF] Maynard | Multiple splice variants of the human HIF-3α locus are targets of the von Hippel-Lindau E3 ubiquitin ligase complex[END_REF]. Moreover, HIF-3α1-3 differ by alternative first exons resulting in different translational start sites. HIF-3α2 thereby lacks most of the DNA binding bHLH domain. Functional discrimination of the HIF-3α splice variants are impaired by the lack of cell lines expressing HIF-3α at high levels and of isoform-specific antibodies. Interestingly, the human IPAS orthologue HIF-3α4 was detected in the kidney and shown to be reduced in renal cell carcinomas [START_REF] Maynard | Human HIF-3α4 is a dominant-negative regulator of HIF-1 and is down-regulated in renal cell carcinoma[END_REF]. Since its downregulation may enhance hypoxic gene induction in kidney tumors, this effect may contribute to tumorigenesis [START_REF] Maynard | Human HIF-3α4 is a dominant-negative regulator of HIF-1 and is down-regulated in renal cell carcinoma[END_REF]. To gain insight into the function and regulation of HIF-3α, we screened a number of cell types and identified renal carcinoma Caki-1 cells as a cell line with high HIF-3α expression. We found that hypoxia induces different HIF-3α splice variants at the transcriptional level, mediated by HIF-1α and that HIF-3α may exert a modulating effect on specific HIF target genes.

MATERIALS AND METHODS

Cell culture: Caki-1 renal carcinoma cells, Hep3B and HepG2 hepatoma cells, HeLa cervical cancer cells, Kelly neuroblastoma, HT1080 fibrosarcoma, and HEK293 human embryonic kidney cells were purchased from the ATCC or the DSMZ (Braunschweig, Germany). The VHL-deficient and -reconstituted (+VHL) renal carcinoma cell lines 786-0 and RCC4 were provided by P. Ratcliffe (Oxford, U.K.) and P. Maxwell (London, U.K.), and HKC-8 renal tubular cells by L. Racusen (Baltimore, MD, USA). Carcinoma cells were grown in DMEM, 1.0 g glucose/L, 10% fetal bovine serum and antibiotics. Human primary renal tubular epithelial (hPT) cells were prepared from tumor nephrectomies and cultured as described before [START_REF] Kroening | Regulation of connective tissue growth factor (CTGF) by hepatocyte growth factor in human tubular epithelial cells[END_REF]. Hypoxic stimulation was accomplished by exposing cells to 1% oxygen in a Jouan IG750 incubator (Thermo Electron). For chemical HIF induction the iron chelator 2,2´dipyridyl (DP; 100 µM) and the iron-independent hydroxylase inhibitor dimethyloxalylglycine (DMOG; 1 mM) were used.

RNase protection assay (RPA):

Templates for RNA probes were generated by reverse transcriptase PCR using the primers listed in Supplemental table 1. Amplified fragments were cloned into the pSP72 vector (Promega). 32 P-labeled RNA probes were generated by in vitro transcription and RPAs were carried out as described [START_REF] Warnecke | Differentiating the functional role of hypoxia-inducible factor (HIF)-1α and HIF-2α (EPAS-1) by the use of RNA interference: erythropoietin is a HIF-2α target gene in Hep3B and Kelly cells[END_REF][START_REF] Warnecke | The specific contribution of hypoxia-inducible factor-2α to hypoxic gene expression in vitro is[END_REF]. Quantification of signals was performed using a Phosphoimager (FujiBAS 2000, Japan) and the AIDA ® image analysis software (Raytest, Germany).

Reverse transcriptase-polymerase chain reaction (RT-PCR):

RNA was transcribed using random hexamer primers and M-MuLV Reverse Transcriptase (MBI Fermentas). For the differentiation of HIF-3α splice variants, PCR reactions were carried out with isoformspecific primers (Suppl. table 1) and analyzed on 2% agarose gels. For quantification Realtime PCR was performed on an ABI-Prism7000 thermocycler with SYBR Green Master Mix (Applied Biosystems). Data were analyzed using the 2 (-ΔΔCt) method. Relative expression levels were calculated using β-actin mRNA as reference. Primers are listed in Suppl. table 1.

SiRNA transfections: Cells were transfected with HIF-1α and HIF-2α siRNAs at 50 nM as described before [START_REF] Warnecke | Differentiating the functional role of hypoxia-inducible factor (HIF)-1α and HIF-2α (EPAS-1) by the use of RNA interference: erythropoietin is a HIF-2α target gene in Hep3B and Kelly cells[END_REF][START_REF] Warnecke | The specific contribution of hypoxia-inducible factor-2α to hypoxic gene expression in vitro is[END_REF]. SiRNAs directed against luciferase (luc) and green fluorescence protein (GFP) were used as controls. For HIF-3α knock-down normal siRNAs (HIF-3α´ and HIF-3α´´; Qiagen and Eurogentec) and stabilized Stealth ® siRNAs (3α69 and 3α70; Invitrogen) and a GC content-matched Stealth ® control siRNA (H) were used (see Suppl. table 1). All HIF-3α siRNAs were targeting HIF-3α1-4.

Promoter constructs and luciferase reporter assays: Candidate HREs in the HIF-3α genomic locus were localized using the MatInspector ® software [15]. DNA fragments comprising the HREs were PCR-amplified and cloned into the pGL2basic vector (Promega). The functional HRE was inactivated by site-directed mutagenesis. Luciferase reporter assays were carried out as described [START_REF] Warnecke | Differentiating the functional role of hypoxia-inducible factor (HIF)-1α and HIF-2α (EPAS-1) by the use of RNA interference: erythropoietin is a HIF-2α target gene in Hep3B and Kelly cells[END_REF]. The functional role of HIF-3α was investigated by cotransfection of HIF-3α siRNAs with a 6xHRE luciferase reporter (6xHRE-luc). Because of the long half-life of HIF-3α protein, cells were stimulated 32 h after transfection for further 16 h.

Chromatin immunoprecipitation (ChIP):

Binding of HIF-1 to the candidate HREs of the HIF-3α gene was demonstrated by ChIP analyses as described previously [16]. Cells were fixed with 1% formaldehyde, lysed by 1% SDS and sonicated. Samples were then diluted 10fold, divided into two parts, and precipitated with anti-HIF-1α rabbit polyclonal antibody (NB 100-449, Novus Biologicals) or normal rabbit serum and protein A agarose (Roche). Precipitated DNA was used for PCR with the primers listed in Suppl. table 1.

Immunoblotting: Protein expression of HIF-α subunits was analyzed as described [START_REF] Warnecke | Differentiating the functional role of hypoxia-inducible factor (HIF)-1α and HIF-2α (EPAS-1) by the use of RNA interference: erythropoietin is a HIF-2α target gene in Hep3B and Kelly cells[END_REF][START_REF] Warnecke | The specific contribution of hypoxia-inducible factor-2α to hypoxic gene expression in vitro is[END_REF] with polyclonal anti-HIF-3α (ab10134, Abcam), anti-HIF-1α (NB100-449) or anti-HIF-2α (NB100-122, Novus Biologicals) antibodies; β-actin (monoclonal antibody AC-74, Sigma) was used as loading control. X-ray films were scanned and signals quantified densitometrically using the AIDA ® software.

Tumor collection: Tissue specimens were collected from total nephrectomy in patients with renal clear cell carcinoma [17]. All patients gave their informed consent. The study was approved by the local ethics committees. Tissue samples were immediately snap frozen after surgical removal or fixed overnight in 4% paraformaldehyde.

Immunohistochemistry: HIF-3α and HIF-1α protein was immunostained in human kidney tumor sections as described [17]. For immunofluorescence Alexa fluor ® 488 labelled secondary antibodies were used.

Statistical analysis:

Data are expressed as mean±SD or mean±SEM, as appropriate. Statistical analyses were carried out using a Stat-View software (Version 5.0 SAS Institute, USA). Comparisons among groups were made using ANOVA, with the correction of Bonferroni/Dunn´s method, where applicable. P values of <0.05 were considered significant.

RESULTS

Expression and hypoxic induction of HIF-3α splice variants in Caki-1 cells.

We first screened human cell lines for HIF-3α expression by RNase protection assay (RPA) using a probe for the PAS-B domain of HIF-3α, which is contained in all splice variants (Fig. 1 A). Among ten cell lines tested, Caki-1 renal carcinoma cells expressed HIF-3α mRNA by far at the highest level, showing a moderate increase after stimulation with the hypoxia mimetic 2,2´dipyridyl (DP; Fig. 1 B). Hep3B and HeLa cells expressed about 10% of HIF-3α transcript levels found in Caki-1 cells. In VHL-deficient and VHL-reconstituted 786-0 and RCC4 renal carcinoma cells HIF-3α was not detectable by RPA or RT-PCR (Fig. 1 B and data not shown). Therefore, we primarily used Caki-1 cells for subsequent experiments. Hypoxia (1% O 2 ) and the hypoxia mimetics DP and DMOG stimulated HIF-3α mRNA expression approximately 3-fold as determined by RPA (hypoxia 3.0±0.9-fold, DP 3.9±0.7fold and DMOG 3.6±0.3-fold; each p<0.05 vs. control, n=3-4; Fig. 1 C). Five splice variants have been reported and confirmed for the human HIF-3α locus (Fig. 1 A). The ODD containing transcripts for HIF-3α1, 2 and 3 differ structurally in terms of their transcriptional start sites thus allowing further differentiation. RT-PCR analyses identified a specific band for HIF-3α2, while those corresponding to HIF-3α1 and 3 were absent within the amplification range of up to 40 cycles (Fig. 1 D). HIF-3α6 was also absent (not shown). Next we quantified mRNA levels of the HIF-3α splice variants by RPA using two different probes directed against the oxygen-dependent degradation domain (ODD: for HIF-3α1, 2 and 3) and HIF-3α4, respectively (Fig. 1 A andE). After 4 and 16 h hypoxia, HIF-3αODD containing transcripts (here: HIF-3α2) and HIF-3α4 were clearly increased, reaching 2.2±0.2and 2.8±0.4-fold, respectively (both p<0.05 vs. unstimulated controls, n=3; Fig. 1 F). Although total HIF-3α expression was markedly lower, a similar induction was observed in Hep3B cells (not shown). In addition, qRT-PCR analysis identified HIF-3α transcripts in human primary renal tubular epithelial cells (hPT, passage 1-3) and in RNA samples from human normal kidney tissue obtained from tumor nephrectomies. In hPT cells total HIF-3α mRNA expression was induced significantly by hypoxia and DP suggesting that HIF-3α is expressed in tubular cells under physiological conditions and that its hypoxic induction may also be preserved in the kidney (Fig. 1 G).

Hypoxic induction of HIF-3α requires HIF-1α.

A time-course analysis of HIF-3α mRNA induction following DP-stimulation revealed kinetics similar to those of known HIF-1 target genes such as angiopoietin-like 4 (ANGPTL4 [18]; Fig. 2 A). We therefore hypothesized that HIF-3α might be a direct transcriptional target of HIF-1 and examined the effects of siRNAs directed against HIF-1α and HIF-2α on HIF-3α mRNA expression. Knock-down of HIF-1α, but not of HIF-2α, significantly reduced the induction of ODD containing transcripts and HIF-3α4 indicating that HIF-3α requires HIF-1α for transcriptional upregulation (Fig. 2 B). HIF-α immunoblotting confirmed the efficiency of the HIF-α knock-down (Fig. 2 C). To address the possibility that HIF-1 might indirectly increase HIF-3α mRNA levels through the induction of a mediator, we investigated the effect of HIF-α knock-down after short-term DPstimulation. After 4 h DP-stimulation, HIF-1α knock-down already reduced the stimulated mRNA levels of both splice variants by about 30%, and after 8 h by 40% (n=3), suggesting that HIF-3α may be a direct target of HIF-1 (Fig. 2 D). In unstimulated normoxic Caki-1 cells, i.e. in the absence of HIF-1α protein, HIF-1α siRNA transfection did not reduce HIF-3α mRNA expression, as determined by the use of qRT-PCR for reasons of sensitivity (Fig. 2 E).

HIF-1α binds to regulatory elements in the human HIF-3α gene.

Next we searched the HIF-3α genomic locus for consensus (R)CGTG/C HIF binding motifs by the use of the MatInspector ® software (Genomatix, [15]). Screening 20 kbp of the 5´flanking region up to exon 2, we identified six candidate HIF binding sites (Fig. 3 A and Supplemental table 2). We then determined whether HIF-1 binds to these sequences using chromatin immunoprecipitation (ChIP) assays. Of three sets of PCR primers used, primer pair 2 yielded an amplified fragment when hypoxic cell lysates were immunoprecipitated with a HIF-1α antibody, which indicated that HIF-1 binds to the HRE at nt 42496-42508 (termed `c´ in Fig. 3 A) in the HIF-3α gene (Fig. 3 B).

Hypoxic regulation of HIF-3α promoter constructs. The functionality of this HIF binding motif was corroborated in luciferase reporter assays. We cloned three different HIF-3α promoter fragments into the pGL2basic vector to cover all candidate HREs and examined their hypoxic induction in different cell types (Fig. 3 A andC). Following DP-stimulation in Hep3B cells, luciferase activity of the reporter driven by the HIF-3α promoter fragment nt 42365-43656 (seq.2-luc) increased 2.9-fold, whereas all other reporter constructs retained activation levels similar to the pGL2 control vector (1.8-to 1.9-fold). Identical results were obtained by co-transfection of a normoxically stable HIF-1α triple mutant (mHIF-1αTM) in HEK293 cells. Again, the seq.2-luc reporter responded more markedly than the other constructs including pGL2 control (16.9-fold vs. 5.0-to 5.6-fold), suggesting that the seq.2 contains a functional HIF binding motif. Thus, the motif `c´ was considered to play a critical role for the hypoxic induction of HIF-3α mRNA. The region spanning nt 42365-43656 (seq.2) was therefore further characterized. 3´terminal deletion of this promoter fragment from nt 42741 (seq.22) to nt 42484 (seq.23) blunted the hypoxic induction markedly (2.6±0.5 vs. 1.5±0.1-fold, p<0.01; Fig. 3 D). In contrast, deleting the 5´end from nt 42365 (seq.22) to 42484 (seq.25) did not affect the hypoxic response significantly (2.6±0.5 vs. 2.3±0.3-fold, p=n.s.), which suggested that a functional hypoxiaresponsive sequence was located between nt 42484 and 42741. Mutagenesis of the HRE candidate motif `c´ at nt 42496-42508 from ACGTC to AGATC (seq.2mut) markedly reduced the hypoxic activation (3.0±0.5 vs. 1.8±0.3-fold, p<0.01; Fig. 3 D). These results further supported that the HRE at nt 42496-42508 upstream of exon 1b is critical for the hypoxic induction of the HIF-3α promoter.

Expression and subcellular localization of HIF-3α protein.

The HIF-3α antibody used in the present study was raised against a peptide within the region AA 300-400 of HIF-3α and therefore predicted not to detect HIF-3α4. Thus, the band of approximately 60 kDa which was marked by the antibody in Caki-1 cells was considered to represent HIF-3α2 according to the results of the RNA analyses. In some experiments, at longer exposures a second, slower migrating band appeared, which may represent a different splice variant or modified HIF-3α2. In keeping with this notion pre-incubation of the antibody solution with the HIF-3α peptide used for immunization eliminated both bands in immunoblot assays (data not shown). Hypoxic induction of HIF-3α was also observed at the protein level, albeit less strikingly than at the transcript level. In Caki-1 cells, hypoxia induced HIF-3α protein 1.5±0.2-fold and DP 1.2±0.09-fold (n=4; p<0.05, Fig. 4 A). The differences between normoxic and stimulated cells may have been underestimated due to the method of quantification (scanning and densitometric analysis of x-ray films). In concordance with mRNA data, Caki-1 cells expressed HIF-3α protein more abundantly than HeLa cells. HIF-3α protein was also detectable in hPT cells (not shown). When 200 µg protein were loaded, three hypoxiainducible bands were marked by the antiserum. In contrast to HIF-1α, HIF-3α did not accumulate during 2 h DP-stimulation, which indicates that HIF-1α, but not HIF-3α, is subject to oxygen-dependent protein destabilization, which is rapidly inhibited by DP (Fig. 4 B). Translational inhibition by cycloheximide revealed that HIF-3α has a distinctly longer protein half-life (approximately 18 h) in Caki-1 cells than HIF-1α, indicating that these two HIF-α family members are regulated by different molecular mechanisms (Fig. 4

C).

As observed at the mRNA level, siRNA-mediated knock-down of HIF-1α, but not HIF-2α, again notably offset the DP-stimulated increase in HIF-3α protein (Fig. 4 D, for HIF-2α protein knock-down see Fig. 2 C), which confirmed that HIF-3α induction requires HIF-1α activation. HIF-1α knock-down reduced hypoxic HIF-3α protein expression by 32±17% (p<0.05, n=4), whereas HIF-2α knock-down had no significant effect (102±13% of luc siRNA-transfected DP-stimulated cells). Immunofluorescence analysis was performed to determine the subcellular localization of HIF-3α in Caki-1 cells (Fig. 4 E). HIF-3α was found in both cytoplasm and nuclei under normoxia. Overnight exposure to hypoxia or DP enhanced in particular the nuclear signals as determined by co-staining of the nuclei with Hoechst dye.

Effect of siRNA-mediated HIF-3α knock-down on HIF-1α activity. HIF-3α has been

proposed to serve as a competitive inhibitor of other HIFs, notably HIF-1α. Therefore we examined the effect of HIF-3α knock-down on HIF-1-driven reporter gene induction in Caki-1 and Hep3B cells. Probably due to the long half-life of HIF-3α protein, knock-down proved to be difficult. The first siRNA (HIF-3α´) reduced HIF-3α protein levels by 12-16% after 48 hours and the second (HIF-3α´´) by about 50% in both Caki-1 and Hep3B cells, whereas the expression of HIF-1α protein was not affected (Fig. 5 A andB). In Hep3B cells cotransfection of siRNA HIF-3α´´ with a 6xHRE-luc reporter significantly increased DPstimulated reporter activity in comparison with GFP control siRNA-transfected, DPstimulated cells (Fig. 5 C). SiRNA HIF-3α´ had no significant effect. We tried to improve knock-down efficiency with chemically stabilized Stealth ® siRNAs. SiRNAs 3α69 and 3α70 did not significantly alter HIF-1α or HIF-2α protein induction (Fig. 5 D). SiRNA 3α69 reduced DP-stimulated HIF-3αODD transcript levels in Caki-1 cells to 66%±13 and HIF-3α4 transcript levels to 71±3%, siRNA 3α70 HIF-3αODD mRNA to 32±5% and HIF-3α4 mRNA to 50±11% compared with control siRNA H as determined by RPA (n=3, Fig. 5 E). Using the more sensitive qRT-PCR analysis, total HIF-3α knock-down amounted to 18% of DPstimulated control values by the 3α69 and 7% by the 3α70 siRNA, respectively (Fig. 5 E, bar graph). HIF-2α mRNA expression was reduced to 72±5% (p<0.05) by the 3α69 siRNA and HIF-1α mRNA to 76±17% (n.s.) of controls by the 3α70 siRNA. In reporter assays in Caki-1 cells, DP-stimulated 6xHRE-driven luciferase activity significantly increased by 70±16% in the presence of the siRNA 3α69 and by 110±55% in the presence of the siRNA 3α70 (both p<0.05, n=6; Fig. 5 F). These data indicate that HIF-3α splice variants in Hep3B and Caki-1 cells may exert inhibitory effects on HIF-1α-dependent gene activation. The modest reduction of HIF-1α and HIF-2α mRNA may have been due to non-specific effects of siRNA transfection. Searching for endogenous HIF target genes that were regulated by HIF-3α in Caki-1 cells, we analyzed RNA samples of DP-stimulated and siRNA 3α69-or 3α70-transfected cells on Affymetrix microarrays and compared them with DP-stimulated control siRNA-transfected cells (data not shown). However, despite good RNA quality the reverse transcription and labelling process was obviously impaired by the Stealth® siRNAs, resulting in low detection calls. A high proportion of genes not related to the hypoxia response were altered by the two HIF-3α siRNAs, which may suggest a high incidence of off-target effects. However, it cannot be excluded that the expression of genes other than HIF-1α and HIF-2α targets may be regulated by HIF-3α, since HIF-β/ARNT, with which HIF-3α is considered to interact, is also part of other transcription factor complexes. In the microarray the HIF target genes Lox, ETS1, IRS1, TOP2B and IGFBP3 appeared in the group of transcripts slightly upregulated by both HIF-3α siRNAs, whereas ANGPTL4, HIG2 and EGLN2 (=PHD1) were among the downregulated genes. We investigated mRNA expression of (amongst others) Lox, ANGPTL4, HIG2 as well as VEGF-A and PDGF-B, because these two genes were previously associated with IPAS/HIF-3α function, in Caki-1 cells by the use of qRT-PCR. Lox was significantly induced by HIF-3α knock-down and ANGPTL4 reduced (Fig. 5 G). For the other genes consistent effects of the two HIF-3α siRNA could not be confirmed.

Immunohistochemical analyses of human renal cell carcinomas reveal co-localization of HIF-3α and HIF-1α.

To investigate whether HIF-3α expression correlates with HIF-1α protein accumulation in vivo, we analysed human renal cell carcinomas (RCCs), the majority of which are associated with inactivation of the VHL gene and thus characterized by constitutive HIF-α protein expression [17,19]. Staining of semi-serially prepared RCC sections with HIF-1α and HIF-3α antibodies confirmed HIF-3α expression in 14 out of 15 kidney tumors, and comparison of the distribution patterns revealed co-localization of HIF-1α and HIF-3α (Fig. 6 and Suppl. fig. 1). Finally, we analysed RNA from HIF-positive and HIFnegative tumors by RPA. In HIF-positive tumors the expression of HIF-3α, both of ODDcontaining splice variants and HIF-3α4, tended to be higher than in normal kidneys, whereas the expression level in HIF-negative tumors was similar to, or even lower than that in normal kidneys (Fig. 7). Altogether these observations indicate that HIF-3α may also be induced by HIF-1α in vivo. However, the lack of HIF-3α expression observed in some HIF-1α positive tumors and in the RCC cell lines in vitro hints at the contribution of other factors to basal HIF-3α expression.

DISCUSSION

In the present study we analysed the molecular mechanisms governing the hypoxic regulation of the human HIF-3α gene in Caki-1 renal carcinoma cells, a cell line with high endogenous HIF-3α expression. The HIF-3α splice variants identified in Caki-1 cells were simultaneously and rapidly upregulated by hypoxia and DP. Induction kinetics and the effects of HIF-1α knock-down indicated that, for both splice variants, transcriptional activation by HIF-1α was the molecular mechanism underlying mRNA induction. In contrast, the alternatively spliced transcripts of the murine HIF-3α/IPAS gene were shown to be regulated through two complementary mechanisms: transcriptional induction and enhanced alternative splicing [START_REF] Makino | Transcriptional up-regulation of inhibitory PAS domain protein gene expression by hypoxia-inducible factor 1 (HIF-1): a negative feedback regulatory circuit in HIF-1-mediated signaling in hypoxic cells[END_REF]20]. Notably, however, total HIF-3α mRNA has been reported to be induced by systemic hypoxia in rat organs [21] as well as in A549 human lung cancer cells [22]. Three of the human HIF-3α splice variants (HIF-3α1-3) encompass an ODD with a single prolyl hydroxylation motif predicting destabilization by molecular oxygen. However, in the context of the full length protein modulation of protein stability by the HIF-3α ODD was greatly diminished [START_REF] Maynard | Multiple splice variants of the human HIF-3α locus are targets of the von Hippel-Lindau E3 ubiquitin ligase complex[END_REF]. Our results are in keeping with these data and show that protein stabilization is of limited, if any, significance for the hypoxic regulation of HIF-3α. ChIP analyses in conjunction with luciferase reporter assays served to pinpoint a functional hypoxia-responsive element in the human HIF-3α gene. The sequence of this DNA element upstream of exon 1b was ACGTC and thus slightly different from the canonical HIF binding site RCGTG. However, the transcriptional induction of the murine IPAS gene was also shown to be mediated by a non-canonical novel hypoxia-responsive DNA element with no similarity to previously described HIF binding sites [START_REF] Makino | Transcriptional up-regulation of inhibitory PAS domain protein gene expression by hypoxia-inducible factor 1 (HIF-1): a negative feedback regulatory circuit in HIF-1-mediated signaling in hypoxic cells[END_REF]. Inactivation of the hypoxia-responsive element by site-directed mutagenesis led to a reduction of promoter activation confirming its functional significance. However, we cannot exclude the possibility that other cis-active elements may contribute to the hypoxic regulation of HIF-3α. A previous study reported a decrease of HIF-3α4 mRNA in human renal carcinomas irrespective of the presence of functional pVHL [START_REF] Maynard | Human HIF-3α4 is a dominant-negative regulator of HIF-1 and is down-regulated in renal cell carcinoma[END_REF]. We rather found a tendency for a colocalization of HIF-3α and HIF-1α protein in human renal carcinomas, in accordance with the notion of a transcriptional induction of HIF-3α by HIF-1α. However, there were HIF-1α positive tumors lacking immunodetectable HIF-3α. In addition, HIF-3α mRNA was not detectable in the RCC cell lines RCC4 and 786-0. Instead, early passage primary human tubular epithelial cells (hPT) expressed considerable amounts of HIF-3α (comparable to Caki-1 cells), which was hypoxia-inducible in a HIF-1-dependent manner (Fig. 1 G and data not shown). It should be noted that HIF-3α mRNA was downregulated after passage 3 and also reduced in Caki-1 cells after extensive passaging, which suggests that dedifferentiation processes may affect basal HIF-3α expression. To date the current knowledge on human HIF-3α function is largely confined to the variant HIF-3α4, the human ortholog of murine IPAS. In contrast to IPAS, which inhibits HIF-1α function by binding to HIF-1α, human HIF-3α4 was demonstrated to bind to both HIF-1α and ARNT [START_REF] Maynard | Human HIF-3α4 is a dominant-negative regulator of HIF-1 and is down-regulated in renal cell carcinoma[END_REF]23] and may thus also compete with other ARNT-dimerization partners, e.g. transcriptions factors involved in the xenobiotic response [START_REF] Hoffman | Cloning of a factor required for activity of the Ah (dioxin) receptor[END_REF]. Therefore, the modulation of a large number of genes not associated with the hypoxic response, as they were detected in a microarray experiment (data not shown), do not necessarily reflect siRNA off-target effects, but may be related to functions of HIF-3α outside of the HIF system. In our attempt to identify HIF target genes whose induction was further enhanced by HIF-3α knock-down, only Lox (lysyl oxidase) could be confirmed, indicating that the functional significance of HIF-3α as a modulator of the hypoxic response is limited and may be target gene-and cell type-specific. Whether the unexpected reduction of hypoxic ANGPTL4 expression by HIF-3α knock-down was indeed HIF-3α-dependent remains to be clarified. As one of the HIF-1α target genes with the highest induction amplitudes [START_REF] Warnecke | The specific contribution of hypoxia-inducible factor-2α to hypoxic gene expression in vitro is[END_REF], ANGPTL4 is supposed to be highly sensitive towards even subtle alterations in HIF-1α transcription and translation, as they may occur during long-term exposure of cells to siRNAs. Reports on stimulatory effects of other HIF-3α splice variants on hypoxic gene expression are rare [24,25]. The majority of studies on HIF-3α function, which were predominantly based on overexpression experiments, point towards inhibitory effects on HIF-dependent gene regulation. Thus, along with the PHDs and the hypoxia-inducible transcriptional repressor of HIF-dependent gene induction CITED2 [26], HIF-3α may represent a further piece of a multipart system of feedback mechanisms which attenuate the hypoxic response and thereby may prevent detrimental effects of an unopposed, prolonged HIF activation. Interestingly, HIF-3α is most highly expressed in the human placenta and the myocardium [START_REF] Maynard | Multiple splice variants of the human HIF-3α locus are targets of the von Hippel-Lindau E3 ubiquitin ligase complex[END_REF], tissues in which the knock-out of HIF-1α or its modulators such as PHD2 and CITED2 has severe developmental consequences. This implies that for proper development and function of these organs a precisely tuned control of HIF activity may be critical. In keeping with this notion genetic deletion recently revealed the functional significance of HIF-3α and its splice variant NEPAS for the regulation of post-natal remodelling in the heart and the lung [27]. It also became evident in that study that HIF-3α splice variants do not act as global repressors of hypoxic gene induction, but may modulate specific target genes in individual cell types. In conclusion, we identified a cell culture system suitable for the analysis of endogenous HIF-3α and demonstrated that the human HIF-3α gene is a transcriptional target of HIF-1α. The contribution of HIF-3α to gene regulation in vivo remains an important issue for further studies.

limited and modulated by cell type-specifi

Fig. 2. Hypoxic induction of HIF-3α mRNA in Caki-1 cells is HIF-1α-dependent.

A, HIF-3αODD and HIF-3α4 mRNA induction after DP-stimulation paralleled the induction of the HIF-1 target gene ANGPTL4. Quantitative data are means of n=2. B, effect of HIF-1α (1α) siRNA, but not HIF-2α (2α) or luciferase (luc) control siRNA, on HIF-3α mRNA induction after 16 h DP stimulation; for quantification of knock-down effects, mRNA expression of luc siRNA-transfected, DP stimulated cells were set to 1. Data in bar graphs are means±SD of n=3. C, HIF-1α and HIF-2α immunoblots demonstrating the efficiency of the HIF-α knock-down in Caki-1 cells. D, effect of HIF-1α knock-down at earlier time points (4 h and 8 h DP); line graph: quantification of HIF-3α transcript levels in HIF-1α siRNAtransfected cells in comparison with luc siRNA-transfected cells (=100%), representative of n=3. E, in unstimulated cells, HIF-1α knock-down did not reduce HIF-3α transcripts. Upper three panels: immunoblots for HIF-1α, HIF-2α and β-actin; lower four panels: RPA of different HIF-3α transcripts; bar graph: qRT-PCR analysis of total HIF-3α mRNA expression (means of n=3).

Fig. 3. The human HIF-3α gene is a transcriptional target of HIF-1.

A, sequence analysis identified six putative HIF binding sites (a-e) in the 5´flanking region and the first intron of the HIF-3α gene (upper). PCR primer design for ChIP analysis (middle) and design of promoter luciferase reporter constructs (lower) are shown in parallel; numbering of nucleotide positions according to acc. no. AC007193. B, ChIP analysis revealed binding of HIF-1α protein to the DNA region comprising HRE `c´ amplified by primer pair 2 (nt 42369-42613). Representative of four independent experiments. C, activation of HIF-3α promoter constructs (as depicted in Fig. 3A) in Hep3B cells after DP-stimulation (upper panel), and in HEK293 cells after co-transfection of a normoxically stable mHIF-1α triple mutant (HIF-1αTM, lower panel). Luciferase activities were normalized to the respective βgalactosidase activities. Data are representative of at least four independent experiments with three replicates per sample (means±SD). D, characterization of the HIF-3α promoter fragment nt 42365-43656 (acc. no. AC007193) in Hep3B cells. Inactivation of the candidate HRE significantly reduced hypoxic activation. Data are means±SEM, # p<0.01, n≥3.

. Effect of HIF-3α knock-down on HIF-1-dependent reporter gene induction

A, immunoblot analysis demonstrating different knock-down efficiencies of two HIF-3α siRNAs in Caki-1 cells: HIF-3α protein levels were reduced by siRNA HIF-3α´´ by 50%; the HIF-3α´ siRNA was virtually inactive. HIF-1α protein expression was not affected. B, immunoblot analysis of Hep3B cells showed that HIF-3α splice variants are more effectively reduced by siRNA HIF-3α´´ than by siRNA HIF-3α´. C, DP-stimulated induction of a 6xHRE-driven luciferase reporter was increased by the HIF-3α´´ siRNA (#p<0.05 vs. GFP-siRNA transfected, DP-stimulated cells; mean±SD of n=4). HIF-1α siRNA served as positive control. D-F, two Stealth ® siRNAs (3α69 and 3α70) directed against all HIF-3α variants also reduced HIF-3α protein, but not HIF-1α or HIF-2α (D). HIF-3α transcripts after 3α69 and 3α70 siRNA transfection (E, bar graph=quantification by qRT-PCR, n=5). HIF-3α knockdown increased 6xHRE-driven luciferase activity in Caki-1 cells (F) in comparison with the control siRNA H (#p<0.05, means±SD of n=6). G, HIF-3α knock-down increased DPinduced Lox mRNA expression and decreased ANGPTL4 induction as determined by qRT-PCR. Mean of n=3-4 (#p<0.05, ##p<0.01 vs. DP-stimulated controls.)

Fig. 1 .

 1 Fig. 1. mRNA expression and hypoxic regulation of HIF-3α splice variants.A, design of RPA probes generated to differentiate HIF-3α splice variants (schematic representation, based on acc. no.: HIF-3α1 NM_152794; HIF-3α2 NM_022462; HIF-3α3 NM_152795; HIF-3α4 AB118749; HIF-3α6 AK024095). B, expression of HIF-3α mRNA in various cell lines as determined by RPA. U6 small nuclear RNA (U6sn) served as loading control; where indicated cells were stimulated with DP (100 µM) for 16 h. C, induction of total HIF-3α mRNA after 16 h exposure to 1% O 2 (hyp), DP or DMOG (DG, 1 mM). D, RT-PCR analysis for the three PAS-B domain-containing HIF-3α splice variants (HIF-3α1, 2 and 3) revealed that HIF-3α2 is the predominant HIF-3α transcript in Caki-1 cells. Genomic DNA served as positive control for HIF-3α1 and the cDNA clone acc. no. BC026308 as control for HIF-3α3. E, mRNA induction of ODD containing HIF-3α splice variants (HIF-3α2) and HIF-3α4 in Caki-1 cells. F, quantification of RPA results; data given are means±SD, #p<0.05 vs. normoxia, n=4. G, in hPT cells total HIF-3α mRNA induction after 16 h hypoxia or DP was quantified by qRT-PCR. Means of three independent experiments with three different hPT preparations.

Fig. 4 .Fig. 5

 45 Fig. 4. Expression, hypoxic induction and subcellular localization of HIF-3α protein.A, immunoblot analysis of Caki-1 and HeLa extracts demonstrating the hypoxic induction of HIF-3α protein; β-actin served as loading control. B, in comparison with HIF-1α, HIF-3α protein induction by DP was distinctly delayed. C, translation inhibition by cycloheximide (CHX, 450 µM) after DP pre-stimulation revealed that HIF-3α has a much longer protein half-life than HIF-1α. D, siRNA-mediated knock-down of HIF-1α, but not HIF-2α, abrogates the induction of HIF-3α protein in Caki-1 cells (see Fig.2C for HIF-2α knockdown), for HIF-3α two independent experiments are depicted (panel HIF-3α (exp. 2)), representative of n=4. E, immunofluorescence staining of HIF-3α in Caki-1 cells after 16 h hypoxia (hyp) or DP. Hoechst staining was used to visualize the nuclei. Overlay (merge) of HIF-3α signals with Hoechst-staining demonstrated that HIF-3α is present in cytoplasm and nuclei already under normoxia (no). Hypoxia and DP enhanced HIF-3α immunoreactivity,

Fig. 6 .

 6 Fig. 6. HIF-3α and HIF-1α expression patterns in human renal cell carcinomas support the notion of a HIF-1α-inducible HIF-3α expression.Immunostaining of semi-serially prepared human renal carcinoma sections revealed colocalization for HIF-1α (Α and C) and HIF-3α (B and D); magnification 200x.

Fig. 7 .

 7 Fig. 7. HIF-3α mRNA expression in renal cell carcinomas. RPA showing mRNA expression of ODD containing HIF-3α transcripts and HIF-3α4 in a HIF-1α/-2α positive and a HIF-1α/-2α negative tumor in comparison with normal kidney tissue. Positions of specific bands are indicated by arrows.

Licenced copy. Copying is not permitted, except with prior permission and as allowed by law. © 2009 The Authors Journal compilation © 2009 Portland Press Limited

Acknowledgements:

The authors thank P. Ratcliffe, C. Pugh (Oxford, U.K.) and P. Maxwell (London, U.K.) for the U6snRNA probe, the 6xHRE luc plasmid and the renal carcinoma cell lines, and S. Kroening (Erlangen, Germany) for primary hPT. T.T. received a grant of the Alexander von Humboldt-Foundation. C.W. was supported by the Roche Foundation for Anemia Research and the ELAN-Fonds of the University of Erlangen-Nuremberg, Germany. The authors thank B. Teschemacher, B. Rogge and H. Fees for excellent technical assistance.


Susana A L Lobo

Amanda Brindley

Martin J Warren
email: m.j.warren@kent.ac.uk

Lígia M Saraiva

Functional characterization of the early steps of tetrapyrrole biosynthesis and modification in Desulfovibrio vulgaris Hildenborough

Keywords: sulphate reducing bacteria, tetrapyrrole biosynthesis, precorrin-2, sirohydrochlorin ALA, 5-aminolevulinic acid, PBG, porphobilinogen, HMB, hydroxymethylbilane, SAM, S-adenosyl-L-methionine, NAD + , nicotinamide adenine dinucleotide, SRB, sulphate reducing bacteria

The biosynthesis of the tetrapyrrole framework has been investigated in the sulphate reducing bacterium Desulfovibrio vulgaris Hildenborough by characterization of the enzymes required for the transformation of 5-aminolevulinic acid into sirohydrochlorin. Porphobilinogen synthase (HemB) was found to be a zinc-dependent enzyme that exists in its native state as a homohexamer. Porphobilinogen deaminase (HemC) was shown to contain the dipyrromethane cofactor. Uroporphyrinogen III synthase is found fused with a uroporphyrinogen III methyltransferase (HemD-CobA). Both activities could be demonstrated in this amalgamated protein and the individual enzyme activities were separated by dissecting the relevant gene to allow the production of two distinct proteins. A gene annotated on the genome as a bifunctional precorrin-2 dehydrogenase/sirohydrochlorin ferrochelatase was in fact shown to act only as a dehydrogenase and is simply capable of synthesizing sirohydrochlorin rather than sirohaem. Genome analysis also reveals a lack of any uroporphyrinogen III decarboxylase, an enzyme necessary for the classical route to haem synthesis. However, the genome does encode some predicted haem d 1 biosynthetic enzymes even though the bacterium does not contain the cd 1 nitrite reductase. We suggest that sirohydrochlorin acts as a substrate for haem synthesis using a novel pathway that involves homologues of the d 1 biogenesis system. This explains why the uroporphyrinogen III synthase is found fused with the methyltransferase, bypassing the need for uroporphyrinogen III decarboxylase activity.

Introduction

Sulphate reducing bacteria (SRB) are environmentally important microorganisms that are found in a wide range of habitats but are reliant on being able to reduce sulphate in a dissimilatory manner to produce sulphide. To assist them in their metabolic challenge SRB employ a rich repertoire of redox groups including porphyrins and related modified tetrapyrroles. Genomic analysis suggests that SRB are able to make at least three types of modified tetrapyrroles, including haem, sirohaem and adenosylcobalamin, the coenzyme form of vitamin B 12 .

Modified tetrapyrroles have a similar structural architecture reflecting the fact that they are all derived from a common macromolecular intermediate, uroporphyrinogen III. Thus all modified tetrapyrroles are made along a branched biosynthetic pathway, as outlined in Figure 1. The pathway starts with the generation of 5-aminolevulinic acid 1 (ALA), the first common intermediate of tetrapyrrole biosynthesis, which is synthesized either from succinylcoenzyme A and glycine (in the case of some proteobacteria, fungi, yeast and animals) or from a tRNA-bound glutamate (in plants, algae, archaea and eubacteria) [START_REF] Gibson | Initial stages in the biosynthesis of porphyrins. 2. The formation of delta-aminolevulinic acid from glycine and succinyl-coenzyme A by particles from chicken erythrocytes[END_REF][START_REF] Jahn | Glutamyl-transfer RNA: a precursor of heme and chlorophyll biosynthesis[END_REF]. The next step involves the asymmetric condensation of two ALA molecules to form the monopyrrole porphobilinogen (PBG), in a Knorr-type condensation reaction catalysed by the enzyme porphobilinogen synthase (also called 5-aminolevulinic acid dehydratase), which is encoded by the hemB gene [START_REF] Nandi | Delta-aminolevulinic acid dehydratase of Rhodopseudomonas spheroides. 3. Mechanism of porphobilinogen synthesis[END_REF]. Porphobilinogen synthases are distinguished according to their requirement in the active site for zinc or magnesium atoms and the need for magnesium in an allosteric binding site. The only known porphobilinogen synthases where these metal binding sites are absent are from the bacterial genus Rhodobacter [START_REF] Jaffe | An unusual phylogenetic variation in the metal ion binding sites of porphobilinogen synthase[END_REF]. R. capsulatus porphobilinogen synthase was shown not to require any metal ion for catalysis, having the added peculiarity of forming a hexameric structure [START_REF] Bollivar | Rhodobacter capsulatus porphobilinogen synthase, a high activity metal ion independent hexamer[END_REF], unlike most other studied porphobilinogen synthases that form octamers (eg. [START_REF] Senior | Comparative studies on the 5aminolaevulinic acid dehydratases from Pisum sativum, Escherichia coli and Saccharomyces cerevisiae[END_REF]).

Four molecules of the pyrrole PBG are subsequently polymerised into the linear tetrapyrrole hydroxymethylbilane (HMB) by the action of porphobilinogen deaminase, which is coded by hemC. The enzyme forms stable enzyme-intermediate complexes with one, two and three PBG-derived pyrrole molecules. A feature of this enzyme is the presence of a covalently bound dipyrromethane cofactor where the growing poly-pyrrole chain is assembled [START_REF] Jordan | Evidence for a dipyrromethane cofactor at the catalytic site of E. coli porphobilinogen deaminase[END_REF][START_REF] Jordan | Mechanism of action of porphobilinogen deaminase. The participation of stable enzyme substrate covalent intermediates between porphobilinogen and the porphobilinogen deaminase from Rhodopseudomonas spheroides[END_REF]. The HMB intermediate is further modified and cyclised by uroporphyrinogen III synthase (encoded by the hemD gene) to yield uroporphyrinogen III [START_REF] Burton | 13C-NMR evidence for a new intermediate preuroporphyrinogen in the enzymic transformation of porphobilinogen into uroporphyrinogens[END_REF]. In the absence of uroporphyrinogen III synthase, HMB cyclises spontaneously to yield the uroporphyrinogen I isomer [START_REF] Burton | 13C-NMR evidence for a new intermediate preuroporphyrinogen in the enzymic transformation of porphobilinogen into uroporphyrinogens[END_REF]. The synthase is able to invert the terminal pyrrole unit of HMB, ring D, and cyclise the macromolecule to give the asymmetric and only biologically relevant type III isomer. In many organisms uroporphyrinogen III represents the first branch point in the pathway, where the action of enzymes such as HemE, F (N) and G results in the formation of protoporphyrin IX, a precursor of modified tetrapyrroles such as haem and chlorophyll. Alternatively, uroporphyrinogen III can undergo an S-adenosyl-L-methionine (SAM) dependent-transmethylation at positions 2 and 7 (by CysG, Met1p or CobA depending on the organism) to generate precorrin-2, a highly unstable yellow dipyrrocorphin [10,[START_REF] Sattler | Cloning, sequencing, and expression of the uroporphyrinogen III methyltransferase cobA gene of Propionibacterium freudenreichii (shermanii)[END_REF]. Precorrin-2 is further modified by a dehydrogenase (CysG, Met8p or SirC enzymes) to yield sirohydrochlorin. This isobacteriochlorin can be chelated with either ferrous iron to form sirohaem by the action of sirohydrochlorin ferrochelatase (CysG, Met8p or SirB enzymes) or with cobalt to generate cobalt-sirohydrochlorin, an intermediate that is directed along the cobalamin biosynthetic pathway. Thus, as with uroporphyrinogen, sirohydrochlorin also represents an important branch point in the biosynthesis of modified tetrapyrroles.

In E. coli and Salmonella enterica the transformation of uroporphyrinogen III into sirohaem is performed by a single multifunctional enzyme, sirohaem synthase (CysG). The SAM-dependent methyltransferase activity is associated with the C-terminal domain of GysG (CysG A ) and the NAD + -dependent dehydrogenation and ferrochelation activities are linked to the N-terminal domain (CysG B ) [START_REF] Stroupe | CysG structure reveals tetrapyrrole-binding features and novel regulation of siroheme biosynthesis[END_REF][START_REF] Warren | Gene dissection demonstrates that the Escherichia coli cysG gene encodes a multifunctional protein[END_REF]. In yeast, two independent enzymes, Met1p

(SAM-dependent methyltransferase) and Met8p (NAD + -dependent dehydrogenase/ferrochelatase) are required for the sirohaem synthesis [START_REF] Hansen | Siroheme biosynthesis in Saccharomyces cerevisiae requires the products of both the MET1 and MET8 genes[END_REF]. A third alternative is found in Bacillus megaterium, where the three reactions are catalyzed independently, by SirA (SAM-dependent uroporphyrinogen III methyltransferase), SirC (NAD + -dependent precorrin-2 dehydrogenase) and SirB (sirohydrochlorin ferrochelatase) [15].

Fused enzyme systems are not just restricted to the sirohaem branch of the pathway. In some bacterial species, such as Selenomonas ruminantium [START_REF] Anderson | A gene, cobA + hemD, from Selenomonas ruminantium encodes a bifunctional enzyme involved in the synthesis of vitamin B12[END_REF], Lactobacillus reuteri [17] and Clostridium josui [START_REF] Fujino | Cloning and sequencing of some genes responsible for porphyrin biosynthesis from the anaerobic bacterium Clostridium josui[END_REF], the uroporphyrinogen III methyltransferase and uroporphyrinogen III synthase activity is performed by one single protein, since the cobA and hemD genes are linked together. The biochemical logic and metabolic importance of such fusions is not fully understood.

The sulphate reducers of the Desulfovibrio genus have many different haemproteins [START_REF] Timkovich | Iron Uroporphyrin I and a Heme c-Derivative are Prosthetic Groups in Desulfovibrio gigas Rubredoxin Oxidase[END_REF][START_REF] Romao | Iron-coproporphyrin III is a natural cofactor in bacterioferritin from the anaerobic bacterium Desulfovibrio desulfuricans[END_REF][START_REF] Murphy | Siroheme and sirohydrochlorin. The basis for a new type of porphyrin-related prosthetic group common to both assimilatory and dissimilatory sulfite reductases[END_REF] that play essential roles in the metabolism of the organism. Although tetrapyrrole biosynthesis has not been studied in any molecular detail in the SRB, earlier research on haem synthesis in D. vulgaris had uncovered one of the most interesting discoveries in tetrapyrrole biosynthesis of recent times as it demonstrated that haem is made by a novel route utilizing precorrin-2 or sirohydrochlorin as an intermediate [22]. This new pathway is likely also to be the major route for haem synthesis in the archaea. A follow-up study in D. vulgaris identified 12,18-didecarboxysirohydrochlorin (or its reduced form) as a possible intermediate but no specific enzymes/genes were linked with the metabolic activity. The authors also suggested that this novel pathway proceeded via coproporphyrinogen III, but this was not rigorously shown [START_REF] Ishida | A primitive pathway of porphyrin biosynthesis and enzymology in Desulfovibrio vulgaris[END_REF]. Despite the novelty of this intriguing pathway, no genomic or biochemical investigation into tetrapyrrole synthesis in D. vulgaris has been undertaken even though the genome has been fully sequenced. In this paper we searched the D. vulgaris Hildenborough genome [24] and noted it contains genes encoding enzymes for the transformation of ALA into sirohydrochlorin via uroporphyrinogen III. Consistent with the presence of a novel haem biosynthetic pathway, the organism appears to be missing the majority of genes that encode the classical enzymes required for the transformation of uroporphyrinogen III into haem. In this paper we report the characterization of the early enzymes of tetrapyrrole assembly and modification in D. vulgaris and suggest enzymes that may be involved in the novel haem pathway.

Materials and Methods

Cloning and expression of recombinant proteins D. vulgaris Hildenborough genes encoding putative porphobilinogen synthase (hemB), porphobilinogen deaminase (hemC), uroporphyrinogen III methyltransferase/synthase (cobA/hemD), sirohaem synthase (cysG B ) were PCR amplified from genomic DNA and specific oligonucleotides (Table S1, supplementary material). Two additional oligonucleotides were designed for amplification of truncated forms of cobA/hemD to generate hemD or cobA encoding regions. The individual DNA fragments were cloned into pET vectors (Novagen) (Table S1), which produced proteins with a 6x-His- Tag tail in the N-terminal or C-terminal, depending on the vector utilized. The genes cobA/hemD, cobA and cysG B were also subcloned into pETac [START_REF] Raux | A role for Salmonella typhimurium cbiK in cobalamin (vitamin B12) and siroheme biosynthesis[END_REF] for use in complementation studies. The integrity of the gene sequence for all cloned fragments was confirmed by DNA sequencing.

Plasmids pET-28a(+)cysG B and pET-22b(+)cobA/hemD were transformed into E. coli BL21Gold(DE3) cells (Stratagene) and plasmids pET-14b(+)hemB, pET-14b(+)hemC, pET-14b(+)cobA and pET-14b(+)hemD into E. coli BL21Star(DE3)(pLysS) cells (Invitrogen). In order to over express the proteins, the E. coli cells were grown at 37 ºC, in Luria-Bertani (LB) medium with appropriate antibiotics, until an OD600 between 0.5 and 0.8 was reached. Induction of the genes was achieved by addition of 200-400 µM of IPTG and by growing the cells for 4h at 37 ºC, for cobA/hemD and cysG B genes, or overnight at 19 ºC for hemB, hemC, cobA and hemD genes. The truncated forms of cobA/hemD are designated cobA d and hemD d (where "d" stands for domain).

Protein purification

Cells expressing HemB, HemC, CobA/HemD, CobA d (Met1-His246), HemD d (Lys247-Lys503) and CysG B proteins were harvested and resuspended in 20 mM Tris-HCl buffer, pH 8 containing 500 mM NaCl and 10 mM imidazole (buffer A); for the purification of CysG B all buffers contained 5 % (v/v) glycerol. All cells were lysed by sonication and centrifuged for 20 minutes at ~30000 x g. The soluble fraction was loaded on to a NiCl 2 charged chelating Sepharose resin (4 ml) (GE, Healthcare) equilibrated with buffer A. After washing with 5-10 volumes of buffer A containing 50 and 100 mM imidazole, all proteins were eluted with 2-5 volumes of buffer A with 500 mM imidazole, except CysG B which was eluted with 100 mM imidazole. Protein fractions were pooled together after detection with Bio-Rad protein assay (BioRad), concentrated with Millipore Ultra Centrifugal Filters with a 10-kDa membrane and buffer exchanged, by means of a PD10 column (GE Healthcare), into buffer B (50 mM Tris-HCl pH 8), with the exception of CobA d protein, which was buffer exchanged into buffer B containing 500 mM NaCl and 200 mM imidazole. The purity of the proteins was assessed by SDS-PAGE [START_REF] Laemmli | Cleavage of structural proteins during the assembly of the head of bacteriophage T4[END_REF] and protein concentration was determined by the bicinchoninic acid method [START_REF] Smith | Measurement of protein using bicinchoninic acid[END_REF] using protein standards from Sigma. The native molecular mass of the various proteins was determined by gel filtration chromatography on either a Superdex 200 or 300 HR 10/30 column (GE, Healthcare), previously equilibrated with 20 mM Tris-HCl, pH 7.5.

The detection of the dipyrromethane cofactor of D. vulgaris porphobilinogen deaminase was determined by incubating 0.5 ml of the enzyme (0.2 mg) with an equivalent volume of Ehrlich's reagent [28] and by following the UV-visible spectral changes over 20 min.

Enzymatic assays

The various activity measurements were performed in a Hewlett Packard 8452A photodiode array spectrophotometer, a Shimadzu UV-1700 spectrophotometer or a BMG Labtech Flurostar Optima plate reader. All activities were assayed in buffer B, with the exception of porphobilinogen synthase which was assayed in 100 mM Tris-HCl pH 8.

Porphobilinogen synthase. HemB (0.5-5 µg) was pre-incubated for 5 min at 37 ºC, and the reaction started by addition of ALA (5 mM), in a final volume of 0.5 ml. The reaction mixture was incubated for a further 5 min at 37 ºC. The reaction was stopped by the addition of 500 µl of 10% (w/v) TCA in 100 mM HgCl A c c e p t e d M a n u s c r i p t min at 9700 x g and an equal volume of a modified Ehrlich's reagent [28] was added to the supernatant. After a 15 min incubation, at room temperature, the absorbance at 555 nm was measured (ε 555 = 6.02×10 4 M -1 cm -1 ) [28].

Porphobilinogen deaminase. The HemC activity was determined essentially as described in [29], using 10-30 µg of enzyme and 200 nmol of PBG, and a ε 405 = 5.48×10 5 M - 1 cm -1 . One unit of enzyme is the amount of enzyme necessary to catalyze the utilization of 1 µmol of PBG in one hour.

Uroporphyrinogen III methyltransferase. Uroporphyrinogen III methyltransferase activity was followed by coupling it to precorrin-2 dehydrogenase (SirC) activity, allowing the transformation of uroporphyrinogen III into sirohydrochlorin (ε 376 = 2.4 ×10 5 M -1 cm -1 ) to be monitored. The substrate, uroporphyrinogen III, was generated anaerobically by incubating HemC and HemD from B. megaterium with 2 mg of PBG. The reaction mixture was prepared in an anaerobic chamber (Belle Technology, <2ppm O 2 ), in a final volume of 250 µl with NAD (100 µM), uroporphyrinogen III (2 µM), B. megaterium SirC (20 µg) and with different amounts of D. vulgaris bi-functional CobA/HemD (0.5-10 µg); the reaction was started by addition of SAM (100 µM).

Sirohaem synthase (CysG B ). Precorrin-2 dehydrogenase and sirohydrochlorin chelatase activities were assayed anaerobically following the formation of sirohydrochlorin or cobalt-sirohydrochlorin and sirohaem, respectively (ε 376 = 2.4 ×10 5 M -1 cm -1 ) [START_REF] Schubert | The structure of Saccharomyces cerevisiae Met8p, a bifunctional dehydrogenase and ferrochelatase[END_REF]. The substrates, precorrin-2 and sirohydrochlorin, were generated anaerobically as described previously [START_REF] Schubert | The structure of Saccharomyces cerevisiae Met8p, a bifunctional dehydrogenase and ferrochelatase[END_REF]. The dehydrogenase assay was performed in a 1 ml reaction, at 30 ºC, with precorrin-2 (2 -3 µM), NAD + (800 µM) and different amounts of D. vulgaris CysG B (0.5-10 µg). The chelatase activity was measured with 4.2 µM of sirohydrochlorin, 20 µM of Co 2+ or Fe 2+ and varying the amount of enzyme (10-50 µg).

In vitro generation of precorrin-2 with D. vulgaris enzymes

HemB, HemC and CobA/HemD enzymes of D. vulgaris were tested simultaneously and individually in a linked assay for the in vitro generation of precorrin-2 [START_REF] Raux | The role of Saccharomyces cerevisiae Met1p and Met8p in sirohaem and cobalamin biosynthesis[END_REF]. The combination of enzymes and substrates used for each assay is described in Table 1. All the reaction mixtures were performed in 50 mM Tris-HCl pH 8, in a final volume of 3 ml and contained between 0.05-5 mg of each enzyme, SAM (1 mg) and ALA (0.5 mg) or PBG (0.2 mg). The reaction was incubated for 2 h at room temperature, and after colour development UV-visible spectra were recorded. B. megaterium SirC (0.1 mg) was added to the assays to generate sirohydrochlorin, as a further confirmation of precorrin-2 production (Table 1).

Complementation of E. coli cysG and E. coli hemD mutant strains

The pETac plasmids containing the genes cobA/hemD, cobA and cysG B were transformed into E. coli cysG mutant strain 302∆a (cysteine auxotrophic) (Table 2). E. coli cysG transformants were selected on LB plates, with 100 µg/ml ampicillin and 35 µg/ml chloramphenicol, and the mutant phenotype rescue tested on minimal medium plates with the appropriate antibiotics, in the absence and presence of cysteine. The plasmids pET14b(+)cobA/hemD and pET14b(+)hemD were introduced in E. coli SASZ31 (a hemD gene mutant strain) and selected on LB plates, with 100 µg/ml ampicillin by incubation at 37 ºC, for approximately 20 h. The mutant strains were re-streaked on LB plates and incubated at 37 ºC for a shorter time period (~12 h) to evaluate cell growth.

Results and Discussion

The genome of D. vulgaris [START_REF] Raux | A role for Salmonella typhimurium cbiK in cobalamin (vitamin B12) and siroheme biosynthesis[END_REF] was searched using the basic local alignment search tool (BLAST) of NCBI (http://blast.ncbi.nlm.nih.gov/Blast.cgi) and DNA sequences encoding known bacterial enzymes involved in the early steps of tetrapyrrole biosynthesis and modification, namely HemB, HemC, CobA/HemD and CysG. Four gene-derived amino acid sequences that have a high degree of sequence identity with these enzymes from different organisms, such as E. coli and B. subtilis, were identified (Table 3). To study the D. vulgaris proteins the corresponding genes were cloned, the recombinant proteins produced and biochemically characterized.

D. vulgaris porphobilinogen synthase is active as a hexamer

A comparison of the gene deduced amino acid sequence of D. vulgaris porphobilinogen synthase with the porphobilinogen synthases of other organisms showed that D. vulgaris HemB contains the conserved cysteine rich sequence for Zn 2+ binding (C121, C123 and C131, D. vulgaris HemB residue numbering), the determinant residues for allosteric magnesium binding (R11 and E238, D. vulgaris HemB residue numbering) [START_REF] Jaffe | An unusual phylogenetic variation in the metal ion binding sites of porphobilinogen synthase[END_REF] and the amino acid residues known to be involved in the binding of the substrate and reaction mechanism [START_REF] Kervinen | Mechanistic basis for suicide inactivation of porphobilinogen synthase by 4,7dioxosebacic acid, an inhibitor that shows dramatic species selectivity[END_REF][START_REF] Erskine | Xray structure of 5-aminolevulinic acid dehydratase from Escherichia coli complexed with the inhibitor levulinic acid at 2.0 A resolution[END_REF][START_REF] Frankenberg | High resolution crystal structure of a Mg2+-dependent porphobilinogen synthase[END_REF] (Figure 2). The recombinant D. vulgaris HemB migrates by SDS-PAGE analysis with an apparent molecular mass of ~36 kDa whereas gel filtration studies indicated the enzyme has a native mass of 226 kDa. This suggests that D. vulgaris exists as a homo hexamer.

The specific activity of D. vulgaris HemB was determined in the presence of zinc to be 45 µmol of porphobilinogen h -1 mg -1 with a K M value for ALA of 0.05 mM (Table 4). This activity is within the values measured for other bacterial porphobilinogen synthases such as E. coli [START_REF] Senior | Comparative studies on the 5aminolaevulinic acid dehydratases from Pisum sativum, Escherichia coli and Saccharomyces cerevisiae[END_REF], also assayed in the presence of zinc, and has a similar K M .

Porphobilinogen deaminase of D. vulgaris contains the dipyrromethane cofactor in its active site

The purified recombinant D. vulgaris HemC migrates by SDS-PAGE with an apparent molecular mass of about 38 kDa, which is in close agreement with the predicted mass of 34 kDa. The UV-visible spectrum of the oxidized protein, at pH 8, contains two weak bands around 410 nm and 500 nm (data not shown), similar to the spectrum reported for the E. coli HemC enzyme [START_REF] Jordan | Evidence for a dipyrromethane cofactor at the catalytic site of E. coli porphobilinogen deaminase[END_REF]. To investigate the presence of the dipyrromethane cofactor, the enzyme was mixed with Ehrlich's reagent, and the reaction followed by UVvisible spectroscopy over a 20 min period. A change in absorbance from 566 nm to 495 nm (Figure 3) was observed, consistent with the presence of the dipyrromethane cofactor at the catalytic site of D. vulgaris HemC [START_REF] Jordan | Evidence for a dipyrromethane cofactor at the catalytic site of E. coli porphobilinogen deaminase[END_REF]. The specific activity of enzyme was measured as being 20 µmolh -1 mg -1 , which is similar to the activity of E. coli HemC enzyme (43 µmol h -1 mg -1 ) [29]. D. vulgaris HemC has a K M of 214 µM for porphobilinogen, which is higher than the range of values measured for HemC of E. coli, Rhodopseudomonas sphaeroides, Clostridium josui and Chlorella regularis (19-89 µM) [START_REF] Fujino | Cloning and sequencing of some genes responsible for porphyrin biosynthesis from the anaerobic bacterium Clostridium josui[END_REF]29,[START_REF] Jordan | Purification and properties of uroporphyrinogen I synthetase from Rhodopseudomonas spheroides[END_REF][START_REF] Shioi | Purification by affinity chromatography and properties of uroporphyrinogen I synthetase from Chlorella regularis[END_REF].

In D. vulgaris the uroporphyrinogen III synthase and methyltransferase activities are performed by a single enzyme

A BLAST search with the locus DVU0734 revealed that the N-terminal domain of the protein (amino acids 1-246) exhibits significant amino acid sequence similarity with the uroporphyrinogen III methyltransferase enzymes (CobA), whereas the C-terminal region of 7 The initial characterization of DVU0734 was done by complementation. In E. coli the multifunctional enzyme CysG synthesizes sirohaem from uroporphyrinogen III. A deletion of this gene produces a mutant strain that is not able to produce sirohaem and thus cannot synthesize cysteine. The mutant phenotype can be rescued by complementation with the expression of genes from other organisms, which encode enzymes that catalyze the reactions performed by CysG (SAM-dependent methyltransferase of uroporphyrinogen III, NAD +dependent dehydrogenation of precorrin-2 and ferrochelation of sirohydrochlorin) [START_REF] Raux | A role for Salmonella typhimurium cbiK in cobalamin (vitamin B12) and siroheme biosynthesis[END_REF][START_REF] Raux | The role of Saccharomyces cerevisiae Met1p and Met8p in sirohaem and cobalamin biosynthesis[END_REF][START_REF] Lobo | Two distinct roles for two functional cobaltochelatases (CbiK) in Desulfovibrio vulgaris Hildenborough[END_REF]. To test if the bi-functional CobA/HemD and CobA d of D. vulgaris have uroporphyrinogen III methyltransferase activity in vivo, they were used to rescue the E. coli cysG mutant strain by co-expression with the bi-functional MET8 of yeast, which encodes a protein with precorrin-2 NAD + -dependent dehydrogenation and sirohydrochlorin ferrochelation activities (Table 2). The E. coli cysG strain expressing MET8 of Saccharomyces cerevisiae and cobA/hemD or cobA d of D. vulgaris is able to grow in the absence of cysteine, demonstrating that the D. vulgaris enzyme variants are able to perform the in vivo transmethylation of uroporphyrinogen III to yield precorrin-2.

The in vivo uroporphyrinogen III synthase activity of the D. vulgaris CobA/HemD and HemD d variants was investigated by complementation of an E. coli hemD strain (SASZ31) (see [START_REF] Chartrand | Uroporphyrin-and coproporphyrin I-accumulating mutant of Escherichia coli K12[END_REF]). This strain grows very poorly in LB medium, forming mini colonies, and thus complementation is observed only when the bacterium is transformed with a gene that allows the organism to form normal-sized colonies. Transformation of the mutant strain with plasmids harbouring D. vulgaris cobA/hemD or hemD d resulted in fast growing strains, demonstrating that the D. vulgaris cobA/hemD eliminates the cell growth deficiency and indicating that the encoded protein is functioning in vivo. Furthermore, E. coli cells producing the D. vulgaris CobA/HemD exhibited a reddish pigmentation, which is due to the accumulation of sirohydrochlorin and trimethylpyrrocorphin and is consistent with increased uroporphyrinogen III methyltransferase activity [START_REF] Warren | Gene dissection demonstrates that the Escherichia coli cysG gene encodes a multifunctional protein[END_REF]39].

The in vitro activity of the recombinantly produced and purified enzyme variants was also studied. After purification, the CobA/HemD, which has a molecular mass of ~55 kDa, was tested for uroporphyrinogen III synthase and methyltransferase activity. A specific activity of 3 nmol.min -1 mg -1 was determined for the uroporphyrinogen III methyltransferase activity of the fused enzyme, with a K M for uroporphyrinogen III of 0.4 µM (Table 4). This is within the range of values observed previously, e.g. the Pseudomonas denitrificans CobA A c c e p t e d M a n u s c r i p t of HMB. If uroporphyrinogen III synthase is added, the reaction proceeds to the synthesis of precorrin-2 and the development of a characteristic yellow colour. When the CobA/HemD fusion was supplied to this assay (Table 1), the generation of precorrin-2 was confirmed with the appearance of a characteristic UV-visible spectrum and the development of the typical yellow chromophore of the compound. The further addition of the B. megaterium SirC enzyme, which oxidizes precorrin-2, led to the formation of sirohydrochlorin providing further evidence for the formation of precorrin-2 in the reaction mixture (data not shown).

The activities of the two truncated proteins were investigated separately in an in vitro linked assay. This revealed that the CobA d had uroporphyrinogen III methyltransferase activity but not uroporphyrinogen III synthase activity, while HemD d was found to possess uroporphyrinogen III synthase activity (Table 1). During the course of these studies a mutant variant, F446A, was serendipitously obtained in HemD d and this variant showed no measurable uroporphyrinogen III synthase activity. The function of F446 in HemD enzymes is unknown, and although is not strictly conserved this residue seems to play an essential role. However, more data will be required to confirm the catalytic role of F446.

The D. vulgaris CysG B has precorrin-2 dehydrogenase activity

The D. vulgaris CysG B has 17, 21 and 26 % sequence identity with Saccharomyces cerevisiae MET8, B. megaterium SirC and to the N-terminal part of the E. coli CysG, respectively (Table 3). Although this is a relatively low value of amino acid identity, it is in the same range of identity observed between the first 223 amino acids of E. coli CysG and MET8 (16 %). A common feature of all of these proteins is the presence of a consensus NAD + binding motif at the start of the N-terminal region [START_REF] Bellamacina | The nicotinamide dinucleotide binding motif: a comparison of nucleotide binding proteins[END_REF]. This motif on the D. vulgaris CysG B (GxGxxGx 10 G) is identical to the one observed in MET8 [START_REF] Hansen | Siroheme biosynthesis in Saccharomyces cerevisiae requires the products of both the MET1 and MET8 genes[END_REF] and similar to the one found in the N-terminal domain of the B. megaterium SirC (GxGxxAx 10 G) and E. coli CysG (GxGxxAx 3 Ax 6 G).

CysG B of D. vulgaris was assayed for NAD + -dependent precorrin-2 dehydrogenase activity by following the appearance of sirohydrochlorin, and was found to have a specific activity of ~700 nmol.min -1 .mg -1 and a K M for NAD + of 70 µM (Table 4). The specific activity of D. vulgaris CysG B is significantly higher than the NAD + -dependent precorrin-2 dehydrogenase activity reported for the B. megaterium SirC (60 nmol.min -1 .mg -1 ) [15]. However, the D. vulgaris enzyme is not able to perform the insertion of iron or cobalt into sirohydrochlorin, a result that is consistent with complementation studies, where the corresponding gene was unable to rescue the E. coli cysG mutant strain phenotype (Table 2). Cumulatively, these results show that the putative CysG B of D. vulgaris, although annotated in the D. vulgaris genome as an orthologue of the yeast bi-functional enzyme Met8p [START_REF] Raux | The role of Saccharomyces cerevisiae Met1p and Met8p in sirohaem and cobalamin biosynthesis[END_REF], is in fact a functional SirC enzyme, a single NAD + -dependent precorrin-2 dehydrogenase that does not possess any chelatase activity [15].

Conclusion D. vulgaris makes sirohydrochlorin

In the work reported herein, the biochemical and functional characterization of several enzymes that catalyze the early steps of the modified tetrapyrrole biosynthetic pathway in D. vulgaris was undertaken. Thus, the D. vulgaris porphobilinogen synthase (HemB) was produced recombinantly and shown to be a zinc-dependent enzyme that appears to have a native hexameric structure, in contrast to the majority of bacterial HemB enzymes that are octamers. For porphobilinogen deaminase (HemC), the presence of the dipyrromethane cofactor, which covalently binds the growing poly-pyrrole chain, further supports the universal nature of this essential prosthetic group.

The recombinant D. vulgaris CobA/HemD was shown to be active both in vivo and in vitro, and represents the first such characterisation of a bi-functional uroporphyrinogen III synthase/methyltransferase. This suggests that, in this organism, uroporphyrinogen III is not released as a free intermediate. Rather, uroporphyrinogen III seems to be directed towards the formation of precorrin-2. This highly unstable intermediate is acted upon by a SirC-like protein, which, in the presence of NAD + , transforms it into sirohydrochlorin. It is known that sirohydrochlorin acts as an intermediate for B 12 biosynthesis, since it is chelated with cobalt to generate cobalt-sirohydrochlorin by CbiK [START_REF] Lobo | Two distinct roles for two functional cobaltochelatases (CbiK) in Desulfovibrio vulgaris Hildenborough[END_REF]. It also plays a role in sirohaem synthesis where ferrous chelation generates this prosthetic group, which is required by sulfite reductase and which is also found in D. vulgaris. However, it is also likely that sirohydrochlorin acts as an intermediate in the biosynthesis of haem.

D. vulgaris does not contain HemE, N, G or H

One of the interesting features of the D. vulgaris genome is the absence of the genes that encode enzymes for the classical synthesis of haem from uroporphyrinogen III. Thus, there are no homologues to hemE, hemN, hemG or hemH, which encode the enzymes uroporphyrinogen III decarboxylase, coproporphyrinogen III oxidase, protoporphyrinogen IX oxidase and protoporphyrin IX ferrochelatase, respectively. The absence of these genes is consistent with the presence of a novel pathway for the transformation of precorrin-2 or sirohydrochlorin into haem. Previous work by Sano and co-workers had shown that haem must be made via precorrin-2, since the two methyl groups found on rings A and B of the final porphyrin product are derived from methionine and not from the C2 position of ALA [22]. However, in their follow-up work, it was reported that an incubation of crude cell extracts of D. vulgaris with uroporphyrinogen III resulted in the appearance of sirohydrochlorin, 12,18-didecarboxysirohydrochlorin, coproporphyrin III and protoporphyrin IX. However, in this study only the structure of the novel 12,18-didecarboxysirohydrochlorin was fully characterised [START_REF] Ishida | A primitive pathway of porphyrin biosynthesis and enzymology in Desulfovibrio vulgaris[END_REF].

If coproporphyrinogen III is an intermediate, then it could be acted upon by an anaerobic coproporphyrinogen oxidase such as HemN. Interestingly, the D. vulgaris genome does encode an orthologue of the radical SAM-reliant coproporphyrinogen III oxidase. The HemN proteins are characterised by the presence of an Fe-S redox group, which is constituted by 4 cysteines within the conserved motif (CxxxCxxCxC). However, in the D. vulgaris homologue, one of the cysteines is missing, suggesting that this D. vulgaris protein is not a HemN. Proteins missing this conserved motif have been shown not to be involved in haem biosynthesis [START_REF] Sofia | Radical SAM, a novel protein superfamily linking unresolved steps in familiar biosynthetic pathways with radical mechanisms: functional characterization using[END_REF]. Thus, if coproporphyrinogen III is an intermediate, it is not clear how it is converted into protoporphyrin IX as there are no recognized enzymes present to undertake the required reactions.

Possible proteins involved in the novel haem synthesis pathway

A scan of the genome reveals a number of genes encoding enzymes that are designated for the synthesis of haem d 1 (nirD and nirJ), the cofactor of the nitrite reductase cd 1 . However, D. vulgaris does not seem to have cd 1 . These haem d 1 synthetic homologues are also found in some genomes of methanogens and other bacteria that do not have cd 1 . In the methanogens, it has also been shown that haem is also made via a dimethylated uroporphyrinogen III derivative such as precorrin-2 or sirohydrochlorin [44]. Significantly, there are some mechanistic similarities between the pathways for haem d 1 synthesis and haem construction via precorrin-2. In haem d 1 synthesis the two propionic acid side chains attached at C3 and C8 of the macrocycle are lost and are replaced by oxygen. To achieve haem synthesis from precorrin-2/sirohydrochlorin, the acetic acid side chains at C2 and C7 have to be removed. Both processes may involved radical chemistry and thus could be mediated via a radical SAM enzyme system. It is therefore of interest to note that NirJ is proposed to be a radical SAM protein. We thus propose that the D. vulgaris nirJ and nirD genes are involved in the transformation of either sirohydrochlorin or sirohaem into haem.

In summary, the research described in this paper outlines how the basic tetrapyrrole framework is synthesized up to the first branch point, which we believe to be sirohydrochlorin. Genomic analysis thus suggests that cobalamin and sirohaem are made by classical previously described routes but that haem is synthesized along a novel route using homologues of the haem d 1 biosynthetic apparatus. The presence of a bifunctional CobA/HemD is consistent with uroporphyrinogen III not being a branch point in this bifurcated pathway and may represent a useful marker for an alternative haem biosynthetic route. More research is required to elucidate this surrogate synthesis. It is not clear why some of the Desulfovibrio or archaea have an alternative haem biosynthetic pathway. Possibly, the pathway evolved as a method of producing the more oxidized porphyrin ring system under anaerobic conditions at a time when the major tetrapyrroles were sirohaem and cobalamin. Then as molecular oxygen became more abundant so the haem biosynthesis pathway involving several oxidative steps in the synthesis of protoporphyrin evolved and became the more common route. A c c e p t e d M a n u s c r i p t ) japonicum (P45622), Pseudomonas (P.) aeruginosa (AAG08628) and Pisum (Pi.) sativum (AAA33640) (adapted from [START_REF] Kervinen | Mechanistic basis for suicide inactivation of porphobilinogen synthase by 4,7dioxosebacic acid, an inhibitor that shows dramatic species selectivity[END_REF]) (NCBI accession numbers in parenthesis). Strictly conserved residues are shaded. The symbols represent functionally relevant residues in the E. coli enzyme [START_REF] Kervinen | Mechanistic basis for suicide inactivation of porphobilinogen synthase by 4,7dioxosebacic acid, an inhibitor that shows dramatic species selectivity[END_REF][START_REF] Erskine | Xray structure of 5-aminolevulinic acid dehydratase from Escherichia coli complexed with the inhibitor levulinic acid at 2.0 A resolution[END_REF] which are also conserved in D. vulgaris porphobilinogen synthase: (#) cysteine residues that bind zinc; (*) residues involved in allosteric magnesium binding; (") arginine residues involved in binding A-side ALA; (-) residues of the lid region that covers the active site; (&") lysine residue required for correct function of the enzyme; (&) lysine residue that forms the Schiff base with P-side ALA; ($) residues involved in binding of P-side ALA.

4

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2 .5

 2 The mixture was centrifuged for 5 Biochemical Journal Immediate Publication. Published on 06 Mar 2009 as manuscript BJ20090151 THIS IS NOT THE VERSION OF RECORD -see doi:10.1042/BJ20090151

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Biochemical

  Journal Immediate Publication. Published on 06 Mar 2009 as manuscript BJ20090151 THIS IS NOT THE VERSION OF RECORD -see doi:10.1042/BJ20090151 A c c e p t e d M a n u s c r i p t DVU0734 (amino acids 247-503) has similarity with uroporphyrinogen III synthases (HemD). This suggests that in D. vulgaris the two activities are fused into a single multifunctional enzyme, which was named CobA/HemD. An amino acid sequence alignment of D. vulgaris DVU0734 with other separate bacterial CobA and HemD enzymes, allowed us to predict accurately the point of fusion between the two functional domains. Consequently, three proteins were recombinantly produced: the whole CobA/HemD protein, the truncated form corresponding to the N-terminal region (named CobA d ), and a second shorter protein corresponding to the C-terminal domain, (named HemD d ).

  (1 µM) [40]. In contrast to the B. megaterium and P. denitrificans CobAs [10, 40], the D. vulgaris CobA/HemD did not display substrate inhibition with uroporphyrinogen III, a characteristic that had previously been observed with an archaeal uroporphyrinogen III methyltransferase [41]. The uroporphyrinogen III synthase activity of the CobA/HemD fusion protein was evaluated in a linked assay. Here, all the enzymes required to transform ALA into precorrin-2, except for the uroporphyrinogen III synthase, are mixed together. When ALA and SAM are added to this incubation the reaction stalls with the accumulation 8 Biochemical Journal Immediate Publication. Published on 06 Mar 2009 as manuscript BJ20090151 THIS IS NOT THE VERSION OF RECORD -see doi:10.1042/BJ20090151

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Biochemical

  Journal Immediate Publication. Published on 06 Mar 2009 as manuscript BJ20090151 THIS IS NOT THE VERSION OF RECORD -see doi:10.1042/BJ20090151 A c c e p t e d M a n u s c r i p t A c c e p t e d M a n u s c r i p t A c c e p t e d M a n u s c r i p t new analysis and information visualization methods. Nucleic Acids Res 29, 1097-1106 44 Buchenau, B., Kahnt, J., Heinemann, I. U., Jahn, D. and Thauer, R. K. (2006) Heme biosynthesis in Methanosarcina barkeri via a pathway involving two methylation reactions. J. Bacteriol. 188, 8666-8668 45 Larkin, M. A., Blackshields, G., Brown, N. P., Chenna, R., McGettigan, P. A., McWilliam, H., Valentin, F., Wallace, I. M., Wilm, A., Lopez, R., Thompson, J. D., Gibson, T. J. and Higgins, D. G. (2007) Clustal W and Clustal X version 2.0. Bioinformatics 23, 2947-2948 46 Raux, E., Schubert, H. L. and Warren, M. J. (2000) Biosynthesis of cobalamin (vitamin B12): a bacterial conundrum. Cell. Mol. Life Sci. 57, 1880-1893 15 Biochemical Journal Immediate Publication. Published on 06 Mar 2009 as manuscript BJ20090151 THIS IS NOT THE VERSION OF RECORD -see doi:10.1042/BJ20090151 A c c e p t e d M a n u s c r i p t

Figure 1

 1 Figure 1 Tetrapyrrole metabolic pathway. Adapted from [46].

Figure 2

 2 Figure 2Amino acid sequence alignment of porphobilinogen synthases from D. vulgaris (AAS95336), E. coli (NP_414903), Bradyrhizobium (Br.) japonicum (P45622), Pseudomonas (P.) aeruginosa (AAG08628) and Pisum (Pi.) sativum (AAA33640) (adapted from[START_REF] Kervinen | Mechanistic basis for suicide inactivation of porphobilinogen synthase by 4,7dioxosebacic acid, an inhibitor that shows dramatic species selectivity[END_REF]) (NCBI accession numbers in parenthesis). Strictly conserved residues are shaded. The symbols represent functionally relevant residues in the E. coli enzyme[START_REF] Kervinen | Mechanistic basis for suicide inactivation of porphobilinogen synthase by 4,7dioxosebacic acid, an inhibitor that shows dramatic species selectivity[END_REF][START_REF] Erskine | Xray structure of 5-aminolevulinic acid dehydratase from Escherichia coli complexed with the inhibitor levulinic acid at 2.0 A resolution[END_REF] which are also conserved in D. vulgaris porphobilinogen synthase: (#) cysteine residues that bind zinc; (*) residues involved in allosteric magnesium binding; (") arginine residues involved in binding A-side ALA; (-) residues of the lid region that covers the active site; (&") lysine residue required for correct function of the enzyme; (&) lysine residue that forms the Schiff base with P-side ALA; ($) residues involved in binding of P-side ALA.

Figure 3 D

 3 Figure 3 D. vulgaris porphobilinogen deaminase spectral features. Reaction with Ehrlics's reagent assessed by UV-visible. The spectra were recorded at the indicated times.

Figure 4

 4 Figure4UV-visible spectra of precorrin-2 (A) and sirohydrochlorin (B) generated in the linked assay performed with D. vulgaris recombinant enzymes.

Figure 1 21

 1 Figure 1

Figure 2 22

 2 Figure 2

Figure 3 23

 3 Figure 3

Figure 4 24

 4 Figure 4

Table 2 Strains and plasmids used in complementation studies together with D. vulgaris enzymes

 2 HemD activity tested in the presence of CobA/HemD protein and HemD d . ¶ CysG B protein was tested for precorrin-2 dehydrogenase and sirohydrochlorin ferrochelatase activity. ∆ CysG B protein was tested for sirohydrochlorin ferrochelatase activity. Pseudomonas (P.); Saccharomyces (Sa); Methanothermobacter (M.).

		Biochemical Journal Immediate Publication. Published on 06 Mar 2009 as manuscript BJ20090151
		D. vulgaris	Strain	Plasmids	Plasmid description	References
		enzyme
	THIS IS NOT THE VERSION OF RECORD -see doi:10.1042/BJ20090151	pETac-cobA/hemD pCIQ-met8p pETac-cobA pCIQ-met8p pETac-PdcobA pCIQ-met8p pET14b(+)-cobA/hemD pET14b(+)hemD pET14b(+)EchemD pETac-cysG B pCIQ-PdcobA pETac-met8p pCIQ-PdcobA pETac-cysG B pCIQ-sirCcobA A c c e p t e d CobA/HemD* E. coli 302∆a CobA d * E. coli 302∆a E. coli 302∆a (Positive control) CobA/HemD § E. coli SASZ31 HemD d § E. coli SASZ31 E. coli SASZ31 (Positive control) CysG B ¶ E. coli 302∆a E. coli 302∆a (Positive control) CysG B ∆ E. coli 302∆a M a n u s c r i p t D. vulgaris cobA/hemD cloned in pETac This study Sa. met8p cloned in pCIQ This study D. vulgaris cobA cloned in pETac This study Sa. cerevisiae met8p cloned in pCIQ This study P. denitrificans cobA cloned in pETac Sa. cerevisiae met8p cloned in pCIQ This study This study D. vulgaris cobA/hemD cloned in pET14b(+) This study D. vulgaris hemD cloned in pET14b(+) This study E. coli hemD cloned in pET14b(+) This study D. vulgaris cysG B cloned in pETac This study P. denitrificans cobA cloned in pCIQ [37] Sa. cerevisiae met8p cloned in pETac P. denitrificans cobA cloned in pCIQ [30] [37] D. vulgaris cysG B cloned in pETac This study M. thermoautotrophicus sirC and [37]
					P. denitrificans cobA cloned in
					pETac
						17

§

Table 3 Amino acid sequence identities and similarities of D. vulgaris enzymes with orthologs

 3 Biochemical Journal Immediate Publication. Published on 06 Mar 2009 as manuscript BJ20090151 THIS IS NOT THE VERSION OF RECORD -see doi:10.1042/BJ20090151

		Identity (%) Similarity (%)
		D. vulgaris HemB
	E. coli	47	66
	Br. japonicum	42	60
	P. aeruginosa	44	62
	Pi. sativum	41	54
		D. vulgaris HemC
	E. coli	52	63
	S. enterica	51	63
	R. capsulatus	40	54
	C. josui	33	50
		D. vulgaris CobA/HemD
	Se. ruminantium	52	63
	L. reuteri	29	47
	C. josui	36	56
	Li. innocua	35	53
		D. vulgaris CobA d
	P. denitrificans	40	56
	Br. megaterium	48	67
	Pa. denitrificans	34	47
	M. thermautotrophicus	42	62
		D. vulgaris HemD d
	E. coli	23	33
	S. enterica	22	33
	B. megaterium	27	47
	Pa. denitrificans	16	27
		D. vulgaris SirC
	E. coli CysG	12	20
			18

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© 2009 The Authors Journal compilation © 2009 Portland Press Limited

Biochemical Journal Immediate Publication. Published on 06 Mar 2009 as manuscript BJ20090151 THIS IS NOT THE VERSION OF RECORD -see doi:10.1042/BJ20090151

Acknowledgements: This work was financed by the FCT project PTDC/BIA-PRO/61107/2006, and Susana Lobo is recipient of the SFRH/BD/19813/2004 grant. Financial support from the Biotechnology and Biological Sciences Research Council (BBSRC) is also acknowledged.