Prochlorococcus marinus strain PCC 9511, a picoplanktonic cyanobacterium, synthesizes the smallest urease

Katarzyna A. Palinskaa,1, Thomas Jahns2, Rosmarie Rippka1 and Nicole Tandeau de Marsac1

Unité de Physiologie Microbienne, Département de Biochimie et Génétique Moléculaire, Institut Pasteur (CNRS, URA 1129), 28 rue du Docteur Roux, 75724 Paris, France1
Institut für Mikrobiologie, Fachrichtung 13.3, Universität des Saarlandes, D-66041 Saarbrücken, Germany2

Author for correspondence: Nicole Tandeau de Marsac. Tel: +33 1 45 68 8415. Fax: +33 1 40 61 3042. e-mail: ntmarsac{at}pasteur.fr


   ABSTRACT
TOP
ABSTRACT
INTRODUCTION
METHODS
REFERENCES
 
The urease from the picoplanktonic oceanic Prochlorococcus marinus sp. strain PCC 9511 was purified 900-fold to a specific activity of 94.6 µmol urea min-1 (mg protein)-1 by heat treatment and liquid chromatography methods. The enzyme, with a molecular mass of 168 kDa as determined by gel filtration, is the smallest urease known to date. Three different subunits with apparent molecular masses of 11 kDa ({gamma} or UreA; predicted molecular mass 11 kDa), 13 kDa (ß or UreB; predicted molecular mass 12 kDa) and 63 kDa ({alpha} or UreC; predicted molecular mass 62 kDa) were detected in the native enzyme, suggesting a quaternary structure of ({alpha}ß{gamma})2. The Km of the purified enzyme was determined as being 0·23 mM urea. The urease activity was inhibited by HgCl2, acetohydroxamic acid and EDTA but neither by boric acid nor by L-methionine-DL-sulfoximine. Degenerate primers were designed to amplify a conserved region of the ureC gene. The amplification product was then used as a probe to clone a 5·7 kbp fragment of the P. marinus sp. strain PCC 9511 genome. The nucleotide sequence of this DNA fragment revealed two divergently orientated gene clusters, ureDABC and ureEFG, encoding the urease subunits, UreA, UreB and UreC, and the urease accessory molecules UreD, UreE, UreF and UreG. A putative NtcA-binding site was found upstream from ureEFG, indicating that this gene cluster might be under nitrogen control.

Keywords: P. marinus subsp. pastoris, Prochlorales, nitrogen metabolism, biochemical characterization, ure genes

The GenBank accession number for the sequence determined in this work is AF242489.

a Present address: Carl von Ossietzky University, ICBM, Geomicrobiology, PO Box 2503, 26111 Oldenburg, Germany.


   INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
METHODS
REFERENCES
 
Prochlorococcus spp. were discovered about 10 years ago in the North Atlantic. With cell diameters ranging from 0·5 to 0·7  µm and a concentration of up to 4x 105 cells ml-1 within the euphotic zone of the world oceans, they represent the smallest and the most abundant photosynthetic organisms known to date, but their real importance in terms of global production is certainly still underestimated (Partensky et al., 1999 ). Because of their very small cell size and thus presumably low nutrient requirement, they dominate in oligotrophic areas of the oceans, where nutrients such as nitrogen and phosphorus are often limiting. Prochlorococcus spp. exhibit a pigment composition remarkable among oxygen-evolving phototrophs, in lacking phycobilisomes and synthesizing divinyl derivatives of chlorophylls a and b (Partensky et al., 1999 ). A novel type of phycoerythrin has been identified in some members (Hess et al., 1996 ). Prochlorococcus spp. together with Prochloron didemni and Prochlorothrix hollandica, with which they share a similar, though not identical, pigment composition, were considered for several years to represent a division or order, named respectively Prochlorophyta and Prochlorales (Lewin, 1976 ; Florenzano et al., 1986 ), distinct from cyanobacteria. Sequence analyses of the 16S rRNA gene, as well as of some other genes, have demonstrated, however, that members of these three genera are dispersed within the radiation of cyanobacteria and therefore can not be considered as a separate phylum (Turner, 1997 ; Florenzano et al., 1986 ).

Urea is a nitrogen source utilized by many cyanobacteria belonging to different taxonomic groups (Kratz & Myers, 1955 ; R. Rippka, unpublished data). Its concentration in natural environments can be similar to those of nitrate and ammonium, e.g. 0·2–5·0 µM, in oceans (DeManche et al., 1973 ). Urease activity is widely distributed in soil and aquatic environments, where it plays an essential role in nitrogen metabolism in plants, algae, some invertebrates, fungi and prokaryotes, including eubacteria and archaea (Mobley & Hausinger, 1989 ). In higher plants, as well as in the prokaryotes examined so far, each molecule of urea is hydrolysed to two molecules of ammonia and one of carbon dioxide by the nickel-requiring metalloenzyme urease (urea amidohydrolase; EC 3 . 5 . 1 . 5), whereas in some fungi and in green algae (Chlorophyceae), urea is first carboxylated to yield allophanate, which is then hydrolysed to two molecules each of ammonia and carbon dioxide (Leftley & Syrett, 1973 ). Ureases are homo- or heteropolymeric enzymes, being composed of one (in jack bean), two (in Helicobacter species) or three (in most bacteria) structural subunits (Mobley et al., 1995 ).

In bacteria, the synthesis of a catalytically active urease of the common three-subunit systems requires a minimum of seven genes (Mobley et al., 1995 ). The ureA, ureB and ureC genes encode two small and one large structural subunits, respectively, and ureD, ureE, ureF and ureG code for the accessory polypeptides required for the assembly of the nickel metallocentre within the urease active site. In some species, urease is constitutively expressed, while in other organisms, urease is induced by urea or derepressed under nitrogen-limiting growth conditions, or controlled by cell growth phase or by pH (Collins & D’Orazio, 1993 ; Mobley et al., 1995 ; De Koning-Ward et al., 1997 ). Seven urease genes have been characterized in the cyanobacteria Synechocystis sp. strain PCC 6803 (Kaneko et al., 1996 ) and Synechococcus sp. strain WH 7805 (Collier et al., 1999 ). In the former strain, the genes are scattered throughout the genome, whereas in the latter they are grouped in two divergently orientated clusters, ureEFG and ureDABC.

In this paper, we report the first biochemical and genetic characterization of the urease complex of the oceanic photosynthetic prokaryote Prochlorococcus marinus strain PCC 9511, an axenic isolate which does not utilize nitrate, urea or ammonium being the preferred sources of nitrogen (Rippka et al., 2000 ).


   METHODS
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ABSTRACT
INTRODUCTION
METHODS
REFERENCES
 
Strain and growth conditions.
The axenic isolate Prochlorococcus marinus Chisholm et al., 1992 , subsp. pastoris subsp. nov. strain PCC 9511 (hereafter designated P. marinus) was grown at 18–20 °C in liquid medium PCR-Tu2 (Rippka et al., 2000 ). Either (NH4)2SO4, urea or alanine at a concentration of 400 µM was used as a nitrogen source. White, true light was supplied by fluorescent tubes (Duro-Lite) providing a photosynthetic photon flux density (PPFD) of 20 µmol photons m-2 s-1 (measured with a LICOR LI-185B quantum/radiometer/photometer equipped with a LI-190SB quantum sensor) with a light/dark cycle of 14 h/10 h. Cultures (6 ml) were maintained in 18 ml culture tubes; 250 ml batch cultures for urease purification were grown in 500 ml Erlenmeyer flasks plugged with cotton stoppers.

The purity of the cultures was checked at each step on plates of medium ASNIII (Rippka et al., 1979 ), supplemented with glucose and Casamino acids (0·2%, w/v, and 0·02%, w/v, respectively) and solidified with Difco Bacto agar (1%, w/v).

Plasmids were maintained in the E. coli strain DH5{alpha} Mcr-. Recombinant E. coli strains were grown at 37 °C in Luria–Bertani medium supplemented with 100 µg ampicillin ml-1.

Assay for urease activity.
Urease activity was determined by measuring the amount of ammonium released from urea, using the indophenol method (Chaney & Marbach, 1962 ). The activity in fractions eluting from the Phenyl-Superose and Phenyl-Sepharose column containing high amounts of ammonium were measured as described by Moore & Kauffman (1970) . One unit of enzyme activity (U) is defined as the decomposition of one µmol of urea min-1 at 30 °C and pH 8·0. Specific activities are expressed as U (mg protein)-1. Protein content was estimated by using the Bio-Rad protein assay with bovine serum albumin (Sigma A-9647) as standard.

Purification of the urease complex.
Pellets from 16 l culture grown to an OD750 of about 0·15–0·17 (equivalent to an OD674 of 0·4–0·5) were harvested by centrifugation at 20000 g for 30 min at 18 °C and resuspended in 10 ml buffer A (50 mM Na2HPO4, pH 7·5; 1 mM EDTA; 3 mM mercaptoethanol) to which 30% (v/v) glycerol was added. After sonication (Branson B12) for 1 min ml-1 with intermittent cooling, cells were centrifuged at 40000 g for 60  min at 4 °C; the supernatant was used as the crude cell-free extract and treated as follows.

(i) Heat treatment.
Cell-free extract was heated for 20 min at 55 °C, stored on ice for 30 min and then centrifuged at 40000 g, for 60 min, at 4 °C; the pellet was discarded.

(ii) SEC column (Sephacryl S300HR).
The supernatant resulting from the heat treatment (10 ml) was loaded onto a SEC column and eluted overnight at 4 °C at a flow rate of approximately 24 ml h-1. Fractions of approximately 6 ml each were collected.

(iii) HIC (hydrophobic interaction chromatography) on Phenyl-Superose.
Fractions from the SEC column (total volume 24 ml) containing urease activity were adjusted to 0·5 M (NH4)2SO4 by the addition of 3 M (NH4)2SO4 and loaded onto a Phenyl-Superose column (bed volume 5 ml) connected to the HPLC system (LKB) (Jahns et al., 1995 ) and equilibrated with buffer A containing 0·6 M (NH4)2SO4. Urease was eluted at a flow rate of 0·4 ml min-1 in a linear gradient from 0·6 to 0 M (NH4)2SO4.

(iv) HIC on Phenyl-Sepharose FF.
The fractions with the highest urease activity were pooled and applied to a Phenyl-Sepharose FF column (bed volume 30 ml) previously equilibrated with buffer A containing 0·6 M (NH4)2SO4. Urease was eluted at a flow rate of 0·8 ml min-1 in a linear gradient from 0·6 to 0 M (NH4)2SO4 at a concentration of approximately 5% (w/v) (NH4)2SO4.

(v) Ion exchange on Mono-Q HR 5/5.
The five fractions with the highest urease activity were pooled (total volume 11 ml). The pooled fractions were diluted with buffer A to obtain a final ammonium concentration of 50 mM and applied to a Mono-Q column connected to the LKB HPLC system. Urease was eluted in a linear gradient of 0–0·6 M NaCl in buffer A (0·3 ml min-1), at a concentration of 320 mM NaCl.

Fractions containing the pure enzyme were mixed with an equal volume of buffer A and glycerol at a final concentration of 30% (v/v) and then kept on ice or stored at -20 °C.

Electrophoresis and immunoblotting analysis.
Native- and SDS-PAGE, and immunoblotting, were carried out as described previously (Forchhammer & Tandeau de Marsac, 1994 ) in a Bio-Rad Mini-Protean system. The urease was visualized using rabbit polyclonal antisera (1:1000 dilution) raised against the purified Bacillus pasteurii urease as the primary antibody. Peroxidase-conjugated anti-rabbit antibodies obtained from Sigma (A-6154) were used as the secondary antibody (1:20000 dilution). Urease was visualized by silver staining as described by Wray et al. (1981) or immunoblotted. The ECL system (Amersham) was used according to the manufacturer’s instructions, with the following modifications: 0·25% (v/v) Tween (Sigma P-4675) was added and dried skim milk was replaced by bovine serum albumin (Sigma A-9647) at concentrations of 3% in NET (6 mM Tris; 1 M NaCl and 100 mM Na2EDTA pH 8·0) for the blocking reagent, and 1% (v/v) in NET for the primary and secondary antibody solutions.

DNA isolation, preparation of the DNA hybridization probe, Southern blots.
DNA of P. marinus was extracted from pooled pellets corresponding to 1 l culture (OD674 approx. 0·4), washed twice in 10 ml NET. Washed cells were resuspended in 10 ml lysis buffer (10 mM Tris, 20 mM EDTA pH 8). After two extractions with phenol/chloroform (1:1) followed by two with chloroform/isoamyl alcohol (24:1), 0·1 vol. 3 M sodium acetate at pH 5 was added and the DNA was precipitated with chilled ethanol (Merck, 100%). The pellet was washed once in 70% (v/v) ethanol, air-dried and resuspended in 10 mM Tris, 0·1 mM EDTA, pH 8.

DNA gel electrophoresis, blotting and hybridizations were carried out as described (Damerval et al., 1989 ). Prehybridization (4  h) and hybridization (16 h) experiments were performed at 65 °C and 55 °C, respectively. Degenerate oligonucleotides were designed to match the highly conserved regions found at residues 217–223 (KLHEDWG) and 315–322 (MLMVCHHL) of UreC in the alignment presented by Mobley et al. (1995) . The sequences of two degenerate primers were: 5'-AAA YTW CAT GAA GAT TGG GG-3' and 5'-ATG ATG YCA WAC CAT WAY CAT-3'. PCR-based amplification was performed as described by Collier et al. (1999) . The PCR product was used as a probe after labelling with [{alpha}-32P]dATP (110 Tbq  mmol-1) by using a Megaprime random labelling kit (Amersham).

Cloning and sequencing of urease genes.
A 5·7 kbp fragment of genomic DNA, digested with EcoRI, gave a strong hybridization signal with the PCR-mediated ureC probe. A partial library was constructed by ligating DNA fragments of approximately 6 kbp into the dephosphorylated pBluescript SK- vector as described by Sambrook et al. (1989) . Ligated DNA was transformed by electroporation (Bio Rad, Gene Pulser) into E. coli DH5{alpha} Mcr- (Dower et al., 1988 ). The clone carrying the correct insert was selected by colony hybridization with the ureC probe. The recombinant plasmid DNA was purified with the QIA filters Qiagen kit (ref. 12262) according to the manufacturer’s instructions and sequenced on both strands (Genome Express, Paris, France). The GenBank accession number of the sequence is AF242489.

RESULTS
Purification of the urease complex
P. marinus exhibited specific urease activities of about 0·1 U (mg protein)-1 in media containing 400 µM ammonia, urea or alanine. After purification of urease from urea-grown cells, an enrichment of about 900-fold and recovery of 12% was obtained (Table 1). Significant amounts of urease were lost at each purification step, since in order to obtain a homogeneous enzyme preparation, only fractions exhibiting the highest urease activities were used for further enrichment. The highest specific activity obtained with the purified enzyme was 94·6 U (mg protein)-1.


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Table 1. Purification of the urease from P. marinus strain PCC 9511

 
Biochemical characterization
Purified enzyme was used for all subsequent studies. The urease from P. marinus exhibited Michaelis–Menten kinetics with a Km value of about 0·23 mM urea, measured at pH 7·5 and 30 °C. The Km was only slightly affected by pH changes between pH 6·8 and 8·0, but the Vmax (maximal velocity) decreased in the acidic range. Maximum activity was obtained at 60 °C for both the purified and non-purified enzyme (Fig. 1 and data not shown).



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Fig. 1. Temperature dependence of the activity ({circ}) and stability ({blacksquare}) of the urease of P. marinus strain PCC 9511. The urease activity was determined at the temperatures indicated after a preincubation of the purified enzyme for 3 min prior to the determination of urease activity. The temperature stability was determined after a preincubation of the purified enzyme for 15 min at the temperature indicated and rapid cooling at 30 °C.

 
Storage of the pure urease at -20 °C for 24 h resulted in an approximately 40% loss of activity, and a remaining activity of 20% was observed after storage on ice for 10 d. The enzyme in the cell-free extracts lost only about 10% of its activity within 15 d on ice (data not shown). Similarly, a higher temperature stability was observed in cell-free extract as compared to the purified enzyme. The stabilization of the enzyme may be favoured at high protein concentration, since the protein concentration in the cell-free extract was approximately 40 µg ml-1, whereas in the assay with the purified enzyme, protein concentration was approximately 36 ng ml-1. The enzyme was stable for at least 15 min at temperatures between 40 and 60 °C, while above 60 °C, a rapid irreversible inactivation occurred (Fig. 1). Urease was stable in the range between pH 4·0 and 11·0 in three different buffer systems (50 mM citrate, 50 mM phosphate and 50 mM diethylbarbiturate). Below or above these pH values, an irreversible inactivation was observed (data not shown).

Inhibition of urease activity
Acetohydroxamate, HgCl2, EDTA and boric acid are known to be common urease inhibitors (Mobley & Hausinger, 1989 ; Mobley et al., 1995 ). Activity of purified urease of P. marinus was 59–94%, 59–92% and 30% inhibited by using 0·1–0·5 mM acetohydroxamate, 0·02–0·1 mM HgCl2 and 10 mM EDTA, respectively (Table 2). No significant inhibitory effect (less than 10%) was observed with boric acid irrespective of the concentration tested (1 and 5 mM) (Table 2). Similarly, L-methionine-DL-sulfoximine (MSX), an inhibitor of glutamine synthetase, the first enzyme involved in the assimilation of ammonium in cyanobacteria, which has been suggested to block ammonium uptake (Singh et al., 1983 ), did not decrease the activity of the purified urease of P. marinus (Table 2).


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Table 2. Inhibition studies on the purified urease of P. marinus strain PCC 9511

 
Molecular mass and subunit composition
The molecular mass of the native urease was determined by gel filtration on a Superdex 200 HR 10/30 as being approximately 168 kDa, using ferritin (450 kDa), catalase (240 kDa), aldolase (158 kDa) and BSA (68 kDa) as standards. Silver-stained native PAGE confirmed the result obtained by gel filtration: a single band corresponding to approximately 168 kDa was observed (Fig. 2). Silver-stained SDS-PAGE gels of the purified urease revealed three bands, corresponding to three subunits of 11 kDa, 13 kDa and 63 kDa. In Fig. 3, the SDS-PAGE immunoblot of the urease purified from cells of P. marinus is compared with that of Anabaena/Nostoc PCC 7120.



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Fig. 2. Silver-stained nondenaturing-PAGE (7%, w/v) of the purified urease. Lane 1, purified urease; lane 2, molecular mass standards (catalase, 232 kDa; aldolase, 158 kDa; bovine serum albumin, 65  kDa; egg albumin 43 kDa).

 


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Fig. 3. SDS-PAGE (12%, w/v) immunoblot of the purified urease of P. marinus strain PCC 9511 and crude cell-free extract of Anabaena/NostocPCC 7120.

 
Characterization of the urease genes
A 5·7 kbp DNA fragment which hybridized to a PCR product corresponding to part of the ureC gene used as a probe was cloned and sequenced (see Methods for details). The genes encoding the three structural subunits, UreA, UreB and UreC, and four accessory proteins, UreD, UreE, UreF and UreG, of the urease of P. marinus were identified by homology with other bacterial urease genes. The physical organization of the P. marinus urease genes is shown in Fig. 4. The two gene clusters ureDABC and ureEFG are divergent and separated by a 47 nt sequence that contains a GTT-N8-TAC motif upstream from ureE. In cyanobacteria, a similar motif GTA-N8-TAC is recognized by NtcA, a DNA-binding protein and transcriptional effector involved in global nitrogen regulation (Flores & Herrero, 1994 ). Putative Shine–Dalgarno sequences were found upstream of ureD, ureA, ureC and ureG (AGA), and of ureF (GGAG), but not upstream of ureB and ureE.



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Fig. 4. Physical organization of the urease genes of P. marinus strain PCC 9511. Arrows indicate the orientation of the genes.

 
The predicted amino acid sequences of the ureA, ureB, ureC and ureG genes of P. marinus share high similarities with their counterparts in the euryhaline cyanobacterium Synechocystis PCC 6803 and the marine Synechococcus WH 7805, but those of the ureD, ureE and ureF genes are less conserved (Table 3). As shown in the alignment of the predicted amino acid sequences of the structural ureC genes of P. marinus, Synechococcus PCC 7002 and WH 7805, Synechocystis PCC 6803 and of selected bacteria (Fig. 5), several conserved residues can be recognized: residues H136, H138, K219, H248, H274 and D362 (numbering according to the P. marinus sequence), which might be ligands for the nickel metallocentre, and A169, G279, C321, A365 and M366, forming part of the putative binding pocket of the enzyme and including H322 and H221, which are probably implicated in catalysis and in substrate binding.


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Table 3. Urease genes of P. marinus strain PCC 9511

 


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Fig. 5. Alignment of a portion (amino acids 136–366) of the predicted amino acid sequences of the structural gene ureC of P. marinus strain PCC 9511, and of selected cyanobacteria (Synechococcus WH 7805 and PCC 7002; Synechocystis PCC 6803) and bacteria (R. meliloti, Rhizobium meliloti; K. aerogenes, Klebsiella aerogenes [K. pneumoniae]; P. mirabilis, Proteus mirabilis; B. pasteurii, Bacillus pasteurii; A. pleuropneumoniae, Actinobacillus pleuropneumoniae).Percentage similarity and identity relative to the derived amino acid sequence of strain PCC 9511 are presented for both the partial and the complete sequences (values in parentheses). Consensus residues were identified by the program a Multalin, INRA, France (Corpet, 1988 ), where each capital letter corresponds to a maximum number of common bases in the corresponding amino acid codon. The conserved residues implicated in the coordination of the two nickel ions at the active site of urease (H136, H138, K219, H248, H274 and D362, numbering according to the P. marinussequence), as well as those responsible for substrate binding (H221) and catalysis (H322) in the putative binding pocket (A169, G279, C321, A365 and M366), are in bold.

 
DISCUSSION
Urease has been purified and characterized from a number of bacteria and the corresponding genes were characterized (Mobley et al., 1995 ). However, only two ureases from cyanobacteria have been purified to homogeneity (Jahns et al., 1995 ), and the urease genes have been sequenced and analysed from only one cyanobacterial species, Synechococcus sp. strain WH 7805 (Collier et al., 1999 ), though they have also been identified on the basis of the genome sequence of Synechocystis sp. strain PCC 6803. In this study, we purified the urease enzyme of P. marinus and characterized the corresponding ure genes.

The assembly of an active urease is a complex process that involves at least seven genes. These genes are clustered in most bacterial genomes (Mobley et al., 1995 ). In both P. marinus and Synechococcus sp. strain WH 7805 (Collier et al., 1999 ), the three genes encoding the urease structural subunits, UreA, UreB and UreC, and the four genes encoding the accessory molecules, UreD, UreE, UreF and UreG, are partly overlapping and organized in two clusters, ureDABC and ureEFG, in opposite orientation. This physical organization differs from that observed for Synechocystis sp. strain PCC 6803 (Kaneko et al., 1996 ), and most probably from that of Synechococcus sp. strain PCC 7002 (Sakamoto et al., 1998 ), in which the ure genes are scattered on the genome. Since a putative binding site for NtcA is found in front of ureEFG in P. marinus, the expression of this gene cluster might be under the global nitrogen control common among cyanobacteria (Flores & Herrero, 1994 ; Flores et al., 1999 ). In Anabaena doliolum (Singh, 1990 ), as well as in Anacystis nidulans and Nostoc muscorum (Singh, 1992 ), ammonium represses the biosynthesis of urease, provided that this nitrogen source is metabolized via the glutamine synthetase/glutamate synthase pathway. In Synechococcus WH 7805, a lower urease activity is observed in ammonium-grown cells than in the presence of urea or nitrate and the control might be NtcA-dependent (Collier et al., 1999 ). In contrast to these cyanobacteria, but similarly to Anabaena variabilis (Ge et al., 1990 ), P. marinus cells grown on different utilizable nitrogen sources (ammonium, urea or alanine) exhibit a very similar urease activity. Therefore, for the latter two cyanobacteria it remains to be established if NtcA is indeed implicated in the control of urease biosynthesis.

Residues responsible for coordinating the nickel ions at the active site, identified by X-ray crystallography of a bacterial urease (Jabri et al., 1995 ), as well as the residues interacting in substrate binding and catalysis (Fig. 5), are conserved in the sequence of P. marinus, suggesting a similar structure and function of this enzyme. The pH and Km values estimated during this study are very similar to those determined for Synechococcus sp. strain WH 7805, and within the range reported for most other cyanobacterial ureases (Rai, 1989 ; Rai & Singh, 1987 ; Singh, 1990 ; Carvajal et al., 1982 ; Jahns et al., 1995 ; Collier et al., 1999 ). The Km value of 0·23 mM is lower, however, than that of the ureases of many other bacteria (Mobley & Hausinger, 1989 ), indicating that the cyanobacterial enzyme has a high affinity for its substrate. The highest specific activity determined for the urease of P. marinus was 94·6 U (mg protein)-1. This specific activity is higher than that observed for most cyanobacterial ureases purified until now (Carvajal et al., 1982 ; Rai, 1989 ; Argall et al., 1992 ; Collier et al., 1999 ), with the exception of the enzymes of Leptolyngbya boryana PCC 73110 and Anabaena/Nostoc PCC 7120, for which specific activities up to 350 U (mg protein)-1 have been measured (Jahns et al., 1995 ). In other bacteria, specific activities up to 180000 U (mg protein)-1 were observed (Mobley & Hausinger, 1989 ). Inhibition of urease by acetohydroxamate was similar for the enzyme of P. marinus (this study) and B. pasteurii (T. Jahns, unpublished results) and in the range of the inhibition observed for the enzymes of Anacystis nidulans and Anabaena doliolum (Rai & Singh, 1987 ). In contrast, the P. marinus enzyme appeared to be less sensitive to inhibition by Hg2+ than the urease of the latter two strains (Rai & Singh, 1987 ).

Further biochemical characterization using the P. marinus enzyme purified to homogeneity confirmed the results of other groups (Jahns et al., 1995 ; Collier et al., 1999 ), who reported a typical UreA-UreB-UreC subunit composition similar to other bacterial ureases for the cyanobacteria Leptolyngbya boryana PCC 73110, Anabaena/Nostoc PCC 7120 and Synechococcus WH 7805. The ureases of the cyanobacteria Spirulina maxima (Carvajal et al., 1982 ), Anabaena doliolum (Rai, 1989 ) and Anabaena cylindrica (Argall et al., 1992 ), being homopolymeric, appear to differ significantly from those of other cyanobacteria and bacteria. However, none of these enzymes were purified to homogeneity. The apparent native molecular mass of the P. marinus urease was determined to be 168 kDa, and the enzyme is composed of three subunits of 11173 Da ({gamma} or UreA), 11678 Da (ß or UreB), and 61653 Da ({alpha} or UreC) (Table 3). The predicted molecular masses of the structural subunits UreC and UreB are only slightly different from those determined from the silver-stained SDS-PAGE gels (63 and 13 kDa, Fig. 3); much greater variations of up to 20% between the calculated and apparent molecular masses have been observed for other bacterial ureases, e.g. Proteus mirabilis (Breitenbach & Hausinger, 1988 ; Jones & Mobley, 1988 ) or Bacillus sp. strain TB-90 (Maeda et al., 1994 ). Apparent molecular masses of 72 and 73 kDa have been reported for UreC from Klebsiella and Proteus mirabilis, respectively (Breitenbach & Hausinger, 1988 ; Todd & Hausinger, 1987 ), while calculated molecular masses of 60·3 and 61·0 kDa have been described (Mulrooney & Hausinger, 1990 ; Jones & Mobley, 1988 ). The observed differences between the molecular mass of UreC of Prochlorococcus and Anabaena/Nostoc (Fig. 3) therefore fall between the possible discrepancies reported for other bacteria.

Ureases studied to date have been reported to be homotrimeric for Brevibacterium ammoniagenes (Nakano et al., 1984 ), homohexameric for Spirulina maxima (Carvajal et al., 1982 ) and Anabaena cylindrica (Argall et al., 1992 ), heterodimeric for Helicobacter spp. (Clayton et al., 1990 ; Ferrero & Labigne, 1993 ; Solnick et al., 1994 ) and heterotrimeric for most other bacterial ureases (Mobley et al., 1995 ). A dimeric structure has been reported for the Ureaplasma urealyticum urease (Saada & Kahane, 1988 ). These results, however, were in contradiction with subsequent studies (Blanchard, 1990 ; Thirkell et al., 1989 ), which demonstrated a heterotrimeric structure. Similarly, the reported homotetrameric structure of the Bacillus pasteurii urease (Christians & Kaltwasser, 1986 ) was later corrected, the enzyme being found to be a heterotrimer (GenBank accession number X78411). Since it is now generally assumed that all ureases possess equal numbers of each of their distinct subunit polypeptides (Mobley et al., 1995 ), the observed and the calculated molecular masses of the subunits of the P. marinus urease described here would correspond to a quaternary structure made up of two heterotrimers ({alpha}ß{gamma})2. This structure yields a predicted native molecular mass of 169 kDa, and the observed apparent native molecular mass of 168 kDa of the P. marinus urease makes it the smallest urease purified so far. The apparent molecular masses for bacterial ureases vary from 200 to 360 kDa, the highest being 800 kDa (Mobley & Hausinger, 1989 ). Within the lower range are the molecular masses of the cyanobacterial ureases of Anabaena/Nostoc PCC 7120 and Leptolyngbya boryana PCC 73110 (220 kDa) (Jahns et al., 1995 ), Anabaena doliolum (228 kDa) (Rai, 1989 ), Anabaena cylindrica (197 kDa) (Argall et al., 1992 ), Spirulina maxima (232 kDa) (Carvajal et al., 1982 ) and Synechococcus WH 7805 (420 kDa) (Collier et al., 1999 ). An anomalously low value (125 kDa) was reported for an urease isolated from a mixed population but never for a purified enzyme (Mobley & Hausinger, 1989 ).

The fact that the urease of P. marinus possibly exhibits an ({alpha}ß{gamma})2 structure, and displays the smallest molecular mass among all the ureases purified to date, may have consequences for its X-ray crystallographic configuration. The relevance of these biochemical results needs to be established.


   ACKNOWLEDGEMENTS
 
We wish to thank Ms R. Schepp for excellent technical support and A. M. Castets for help in the assembling of the nucleotide sequence. This work was supported by the Institut Pasteur, the Centre National de la Recherche Scientique (CNRS, URA 1129) and the European Union MAST III program PROMOLEC (MAS3-CT97-0128).


   REFERENCES
TOP
ABSTRACT
INTRODUCTION
METHODS
REFERENCES
 
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Received 10 March 2000; revised 24 August 2000; accepted 20 September 2000.