Microbiology (2000), 146, 3099–3107
Printed in Great Britain
Prochlorococcus marinus strain PCC 9511, a
picoplanktonic cyanobacterium, synthesizes
the smallest urease
Katarzyna A. Palinska,1† Thomas Jahns,2 Rosmarie Rippka1
and Nicole Tandeau de Marsac1
Author for correspondence : Nicole Tandeau de Marsac. Tel : j33 1 45 68 8415. Fax : j33 1 40 61 3042.
e-mail : ntmarsac!pasteur.fr
1
Unite! de Physiologie
Microbienne, De! partement
de Biochimie et Ge! ne! tique
Mole! culaire, Institut
Pasteur (CNRS, URA 1129),
28 rue du Docteur Roux,
75724 Paris, France
2
Institut fu$ r Mikrobiologie,
Fachrichtung 13.3,
Universita$ t des Saarlandes,
D-66041 Saarbru$ cken,
Germany
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 minN1
(mg protein)N1 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 (γ or UreA ; predicted molecular mass 11 kDa),
13 kDa (β or UreB ; predicted molecular mass 12 kDa) and 63 kDa (α or UreC ;
predicted molecular mass 62 kDa) were detected in the native enzyme,
suggesting a quaternary structure of (αβγ)2. The Km of the purified enzyme was
determined as being 023 mM urea. The urease activity was inhibited by HgCl2,
acetohydroxamic acid and EDTA but neither by boric acid nor by L-methionineDL-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 57 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
INTRODUCTION
Prochlorococcus spp. were discovered about 10 years
ago in the North Atlantic. With cell diameters ranging
from 0n5 to 0n7 µm and a concentration of up to 4i 10&
cells ml−" 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
.................................................................................................................................................
† Present address : Carl von Ossietzky University, ICBM, Geomicrobiology,
PO Box 2503, 26111 Oldenburg, Germany.
The GenBank accession number for the sequence determined in this work
is AF242489.
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
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K. A. P A L I N S K A a n d O T H E RS
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. 0n2–5n0 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 nitrogenlimiting 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
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 mC in liquid medium PCR-Tu
#
(Rippka et al., 2000). Either (NH ) SO , 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−# s−" (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 (0n2 %, w\v, and 0n02 %, w\v,
respectively) and solidified with Difco Bacto agar (1 %, w\v).
Plasmids were maintained in the E. coli strain DH5α Mcr−.
Recombinant E. coli strains were grown at 37 mC in Luria–
Bertani medium supplemented with 100 µg ampicillin ml−".
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−" at 30 mC and pH 8n0.
Specific activities are expressed as U (mg protein)−". 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 OD of about 0n15–0n17 (equivalent to an OD
(&!
'(%
of 0n4–0n5) were harvested by centrifugation at 20 000 g for
30 min at 18 mC and resuspended in 10 ml buffer A (50 mM
Na HPO , pH 7n5 ; 1 mM EDTA ; 3 mM mercaptoethanol) to
#
%
which 30 % (v\v) glycerol was added. After sonication
(Branson B12) for 1 min ml−" with intermittent cooling, cells
were centrifuged at 40 000 g for 60 min at 4 mC ; the supernatant was used as the crude cell-free extract and treated as
follows.
Cell-free extract was heated for 20 min at
55 mC, stored on ice for 30 min and then centrifuged at
40 000 g, for 60 min, at 4 mC ; the pellet was discarded.
(i) Heat treatment.
(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 mC at a flow rate of
approximately 24 ml h−". Fractions of approximately 6 ml
each were collected.
(iii) HIC (hydrophobic interaction chromatography) on PhenylSuperose. Fractions from the SEC column (total volume 24 ml)
containing urease activity were adjusted to 0n5 M (NH ) SO
%# %
by the addition of 3 M (NH ) SO 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 0n6 M (NH ) SO . Urease was eluted at a flow
%# %
rate of 0n4 ml min−" in a linear gradient from 0n6 to 0 M
(NH ) SO .
%# %
(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 0n6 M (NH ) SO . Urease was eluted at a
%# %
flow rate of 0n8 ml min−" in a linear gradient from 0n6 to 0 M
(NH ) SO at a concentration of approximately 5 % (w\v)
%# %
(NH ) SO .
%# %
(v) Ion exchange on Mono-Q HR 5/5. The five fractions with the
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Urease of Prochlorococcus marinus
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 MonoQ column connected to the LKB HPLC system. Urease was
eluted in a linear gradient of 0–0n6 M NaCl in buffer A
(0n3 ml min−"), at a concentration of 320 mM NaCl.
on both strands (Genome Express, Paris, France). The
GenBank accession number of the sequence is AF242489.
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 k20 mC.
P. marinus exhibited specific urease activities of about
0n1 U (mg protein)−" 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
94n6 U (mg protein)−".
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 : 20 000 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 : 0n25 % (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 Na EDTA pH 8n0) for
# for the primary
the blocking reagent, and 1 % (v\v) in NET
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 (OD approx.
'(% resus0n4), washed twice in 10 ml NET. Washed cells were
pended 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), 0n1
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, 0n1 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 mC and 55 mC, 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 : 5h-AAA YTW CAT GAA GAT TGG GG-3h and 5hATG ATG YCA WAC CAT WAY CAT-3h. PCR-based
amplification was performed as described by Collier et al.
(1999). The PCR product was used as a probe after labelling
with [α-$#P]dATP (110 Tbq mmol−") by using a Megaprime
random labelling kit (Amersham).
RESULTS
Purification of the urease complex
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 0n23 mM urea,
measured at pH 7n5 and 30 mC. The Km was only slightly
affected by pH changes between pH 6n8 and 8n0, but the
Vmax (maximal velocity) decreased in the acidic range.
Maximum activity was obtained at 60 mC for both the
purified and non-purified enzyme (Fig. 1 and data not
shown).
Storage of the pure urease at k20 mC 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−",
whereas in the assay with the purified enzyme, protein
concentration was approximately 36 ng ml−". The
enzyme was stable for at least 15 min at temperatures
between 40 and 60 mC, while above 60 mC, a rapid
irreversible inactivation occurred (Fig. 1). Urease was
stable in the range between pH 4n0 and 11n0 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).
Cloning and sequencing of urease genes. A 5n7 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 DH5α 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
Inhibition of urease activity
Acetohydroxamate, HgCl , 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 0n1–0n5 mM acetohydroxamate,
0n02–0n1 mM HgCl and 10 mM EDTA, respectively
#
(Table 2). No significant
inhibitory effect (less than
10 %) was observed with boric acid irrespective of the
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Table 1. Purification of the urease from P. marinus strain PCC 9511
Purification step
Volume
(ml)
Crude extract
Heat treatment
SEC pool
Phenyl-Superose
Phenyl-Sepharose
Mono-Q HR 5\5
11
10
24
16
11
0n9
Protein
(µg ml−1)
3n63
1n18
0n03
0n01
0n01
0n006
Total
protein
(mg)
39n9
11n8
0n72
0n16
0n16
0n0054
Specific
activity [U (mg
protein)−1]
Recovery
(% initial
activity)
Purification
factor
0n105
0n265
4n028


94n575
100
75
69


12
1
2n5
57n5


900
, Not determined.
350
1
Specific activity
300
2
kDa
250
232
200
150
168
158
100
50
35 40 45 50 55 60 65 70 75
65
Temperature (°C)
.................................................................................................................................................
Fig. 1. Temperature dependence of the activity (#) and
stability (
) 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 mC.
Table 2. Inhibition studies on the purified urease of
P. marinus strain PCC 9511
Inhibitor
Concn
(mM)
Remaining activity
(%)
HgCl
0n020
0n050
0n1
0n1
0n5
1n0
2n0
5n0
10
10
1n0
5n0
41
8
0
41
6
100
100
91
95
70
91
90
#
Acetohydroxamic acid
Boric acid
EDTA
MSX*
* MSX, -methionine--sulfoximine.
43
.................................................................................................................................................
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).
concentration tested (1 and 5 mM) (Table 2). Similarly,
-methionine--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).
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
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Urease of Prochlorococcus marinus
PCC
7120
PCC
9511
132
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.
78
Characterization of the urease genes
kDa
216
45·7
32
18·4
7·5
.................................................................................................................................................
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/Nostoc PCC 7120.
.................................................................................................................................................
Fig. 4. Physical organization of the urease genes of P. marinus
strain PCC 9511. Arrows indicate the orientation of the genes.
A 5n7 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-N TAC motif upstream from ureE. In cyanobacteria, )a
similar motif GTA-N -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.
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
Table 3. Urease genes of P. marinus strain PCC 9511
Gene
designation
Gene
product
(aa)
Mol. mass
(Da)*
pI*
Percentage identity†
WH 7805
PCC 6803
Other bacteria
Subunit
UreA (γ)
UreB (β)
UreC (α)
ureA
ureB
ureC
100
106
569
11 173
11 678
61 653
5n76
4n69
5n76
81
64
79
67
60
69
49–66
48–65
55–66
Accessory
proteins
UreD
UreE
UreF
UreG
ureD
ureE
ureF
ureG
297
154
228
201
33 494
18 027
26 682
21 780
7n80
9n58
6n78
5n41
37
43
37
68
28
32
27
66
21–30
24–33
15–25
49–64
* Molecular mass of apoproteins and theoretical pI calculated by using the GCG program version 8.1 (Wisconsin Sequence Analysis
Package).
† Calculated by using the NCBI  program version 2.0.11. WH 7805, Synechococcus WH 7805 ; PCC 6803, Synechocystis PCC 6803 ;
other bacteria, all bacteria whose urease gene sequences are available in GenBank. The ureC gene product from Synechococcus PCC 7002
(GenBank accession number AF035751) is 75 % identical to UreC of P. marinus.
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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. marinus sequence), 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.
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.
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
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Urease of Prochlorococcus marinus
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 0n23 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 94n6 U (mg
protein)−". 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)−" have been measured
(Jahns et al., 1995). In other bacteria, specific activities
up to 180 000 U (mg protein)−" 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
Hg#+ 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
11 173 Da (γ or UreA), 11 678 Da (β or UreB), and
61 653 Da (α or UreC) (Table 3). The predicted mol-
ecular 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 60n3 and 61n0 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 (αβγ) . 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 (αβγ) structure, and displays the smallest molecular
#
mass among
all the ureases purified to date, may have
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K. A. P A L I N S K A a n d O T H E RS
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).
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.................................................................................................................................................
Received 10 March 2000 ; revised 24 August 2000 ; accepted
20 September 2000.
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