Microbiology (2004), 150, 3527–3546
Review
DOI 10.1099/mic.0.27303-0
Nitrate reduction and the nitrogen cycle in archaea
Purificación Cabello,1 M. Dolores Roldán2 and Conrado Moreno-Vivián2
Correspondence
Conrado Moreno-Vivián
[email protected]
1
Departamento de Biologı́a Vegetal, Área de Fisiologı́a Vegetal, Universidad de Córdoba, Spain
Departamento de Bioquı́mica y Biologı́a Molecular, Edificio Severo Ochoa, 1a planta, Campus
Universitario de Rabanales, Universidad de Córdoba, 14071-Córdoba, Spain
2
The nitrogen cycle (N-cycle) in the biosphere, mainly driven by prokaryotes, involves different
reductive or oxidative reactions used either for assimilatory purposes or in respiratory processes
for energy conservation. As the N-cycle has important agricultural and environmental implications,
bacterial nitrogen metabolism has become a major research topic in recent years. Archaea
are able to perform different reductive pathways of the N-cycle, including both assimilatory
processes, such as nitrate assimilation and N2 fixation, and dissimilatory reactions, such as nitrate
respiration and denitrification. However, nitrogen metabolism is much less known in archaea
than in bacteria. The availability of the complete genome sequences of several members of
the eury- and crenarchaeota has enabled new approaches to the understanding of archaeal
physiology and biochemistry, including metabolic reactions involving nitrogen compounds.
Comparative studies reveal that significant differences exist in the structure and regulation of some
enzymes involved in nitrogen metabolism in archaea, giving rise to important conclusions and
new perspectives regarding the evolution, function and physiological relevance of the different
N-cycle processes. This review discusses the advances that have been made in understanding
nitrate reduction and other aspects of the inorganic nitrogen metabolism in archaea.
Introduction
As nitrogen is an essential element for living organisms, the
availability of a suitable nitrogen source often limits primary
productivity in both natural environments and agriculture.
Nitrogen can be found in several oxidation states, from +5
in the most oxidized compound (nitrate) to 23 in the most
reduced form (ammonium), but in biological compounds
it is almost exclusively present in the fully reduced state.
The global biogeochemical nitrogen cycle (N-cycle) allows
the interconversions of nitrogen compounds to maintain a
relatively small amount of fixed or combined nitrogen in a
continuous exchange with atmospheric dinitrogen (N2),
a huge nitrogen reservoir. The N-cycle includes both
reductive and oxidative processes, in which prokaryotes
play a predominant role (Fig. 1). Assimilatory pathways,
such as N2 fixation and nitrate assimilation, generate
ammonium that is further incorporated into cell material,
mainly by the glutamine synthetase–glutamate synthase
(GS–GOGAT) route. However, whereas N2 fixation is only
driven by free-living or symbiotic diazotrophic prokaryotes,
assimilatory nitrate reduction is carried out by bacteria,
fungi, algae and higher plants, and almost all living organisms are able to incorporate ammonium into carbon skeletons.
On the other hand, some prokaryotes can obtain metabolic
energy by redox processes involving nitrogen compounds,
such as nitrification and denitrification. Autotrophic nitrification is a two-step process consisting of the oxidative conversion of ammonia to nitrite via hydroxylamine,
0002-7303 G 2004 SGM
carried out by ammonia-oxidizing bacteria, and the further
oxidation of nitrite to nitrate, performed by nitriteoxidizing chemolithoautotrophic bacteria. Denitrification
is a respiratory process whereby nitrate is successively
reduced to nitrite, NO, N2O and N2, which takes place
predominantly under anaerobic conditions in many facultative bacteria (Berks et al., 1995; Zumft, 1997; MorenoVivián & Ferguson, 1998; Moreno-Vivián et al., 1999;
Richardson & Watmough, 1999). Some heterotrophic
bacteria are able to combine nitrification and aerobic
denitrification by oxidizing ammonia to nitrite, which
is later reduced to N2 (de Boer & Kowalchuk, 2001).
Ammonification is the dissimilatory reduction of nitrate
to ammonia that does not serve the purpose of nitrogen
autotrophy, although this term normally describes the
release of ammonia from organic nitrogen compounds
(Zumft, 1997). Finally, anaerobic ammonium oxidation
(anammox) is a reaction that produces N2 by reducing
nitrite and oxidizing ammonium. This process, performed
by bacterial communities of planctomycetes, seems to be of
ecological importance in marine environments (Dalsgaard
et al., 2003; Kuypers et al., 2003).
Bacteria are involved in all N-cycle pathways, and the
different aspects of their inorganic nitrogen metabolism
have been extensively studied both biochemical and genetically (for specific reviews see Berks et al., 1995; Zumft, 1997;
Ferguson, 1998; Moreno-Vivián et al., 1999; Richardson &
Watmough, 1999; Eisenberg et al., 2000; Halbleib & Ludden,
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3527
P. Cabello, M. D. Roldán and C. Moreno-Vivián
Fig. 1. The biological N-cycle. The different
nitrogen compounds are arranged according
to their oxidation states and the main reductive or oxidative pathways are indicated by
arrows. Conversion of hydroxylamine to
nitrous oxide by heterotrophic nitrification,
and production of N2 from ammonium and
nitrite by anaerobic ammonium oxidation
(anammox), are not indicated. The enzymes
involved in the different pathways are: 1–2,
glutamine synthetase–glutamate synthase
(GS–GOGAT); 3, glutamate dehydrogenase
(GDH); 4, nitrogenase; 5, assimilatory nitrate
reductase (Nas); 6, assimilatory nitrite reductase (sirohaem-Nir); 7, dissimilatory and
respiratory nitrate reductases (Nap and Nar);
8, respiratory nitrite reductases (Cu-Nir and
cd1-Nir); 9, nitric oxide reductase (Nor); 10,
nitrous oxide reductase (Nos); 11, ammonia
monooxygenase; 12, hydroxylamine oxidase;
13, nitrite oxidase.
2000; Arcondéguy et al., 2001; Kowalchuk & Stephen, 2001;
Richardson et al., 2001). Archaea form a monophyletic
domain of organisms distinct from bacteria and eukarya,
with two major subdivisions: crenarchaeota, which are
predominantly hyperthermophiles and anaerobic respirers,
and euryarchaeota, which include methanogens and extreme
halophiles (Woese et al., 1990; Brown & Doolittle, 1997). In
general, archaeal replication, transcription and translation
are more related to eukaryal than to bacterial processes
(Bell & Jackson, 2001; Kelman & Kelman, 2003; Reeve,
2003), but archaeal central metabolism components are
more closely related to bacterial proteins. In addition to
ammonium assimilation, members of archaea can drive all
the reductive pathways of the N-cycle, either dissimilatory
reactions, such as nitrate respiration and denitrification,
or assimilatory pathways like N2 fixation and nitrate assimilation. Because archaea are the predominant microbial
populations in extreme environments, such as highly saline
water and hot springs, they sustain the N-cycle in these
special and severe ecosystems. In spite of this, to date, there
is little information on inorganic nitrogen metabolism
in archaea, especially on nitrate reduction. Although the
archaeal genome sequencing projects are bringing out new
data, the understanding of inorganic nitrogen metabolism
in archaea is still an open research field. This review covers
the current knowledge on the nitrate reduction processes
and other pathways of the N-cycle in archaea.
Assimilatory and respiratory nitrate reduction in
archaea
Many archaea are able to reduce nitrate by assimilatory or
respiratory pathways, dissimilatory nitrate reduction being
much more frequent than nitrate assimilation (Martı́nezEspinosa et al., 2001b). In addition, genes encoding
putative nitrate transporters, nitrate reductases and nitrite
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reductases have been found in the sequenced genomes
of members of both crenarchaeota and euryarchaeota
(Table 1). However, in most cases, either functional or
sequence data, but not both, are available. Therefore, future
research should be directed to characterizing at the physiological and biochemical levels the products of the genes
identified by these genome sequencing projects and their
regulation. This will be of special evolutionary interest
because significant differences between the bacterial and
archaeal nitrate-reducing systems are emerging.
Phylogenetic relationships of the nitrate reductases
Three different nitrate-reducing systems (Nas, Nar and
Nap) have been described in bacteria (Berks et al., 1995;
Moreno-Vivián et al., 1999; Richardson, 2000; Richardson
et al., 2001). Assimilatory nitrate reductases (Nas) are
usually cytoplasmic enzymes which are repressed by
ammonium and use either NADH or ferredoxin as
physiological electron donor. The NADH-Nas enzymes of
Klebsiella and Rhodobacter capsulatus are dimers composed of a small FAD-containing diaphorase and a large
catalytic subunit with one Fe–S centre and a molybdo-bismolybdopterin guanine dinucleotide (Mo-bisMGD) cofactor,
whereas cyanobacterial ferredoxin-dependent enzymes are
monomers (Moreno-Vivián et al., 1999; Richardson et al.,
2001). Membrane-bound nitrate reductases (Nar) are
involved in anaerobic nitrate respiration and denitrification, being negatively regulated by O2 and unaffected by
ammonium. Nar enzymes are composed of a catalytic
subunit with MGD cofactor (NarG) and an electrontransfer subunit with four iron–sulfur centres (NarH). This
complex is bound to a membrane bihaem b quinoloxidizing component (NarI). Recently, the crystal structure
of Nar has been determined at high resolution, revealing
that NarG protein has an aspartate residue acting as the
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Microbiology 150
Nitrogen cycle in archaea
Table 1. Presence of putative nitrate transporters and enzymes involved in nitrate reduction and denitrification in archaea
Evidence for the presence of the different proteins is derived from gene bank sequences and genome sequence analysis (genetic) conducted
on websites http://gib.genes.nig.ac.jp and http://www.genome.ad.jp, from physiological and biochemical studies (functional) or both (genetic
and functional). The accession numbers of the corresponding genes are indicated. ND indicates that the protein or enzyme has not been
detected by genetic or functional analysis. ? indicates that no clear assignation is deduced only on the basis of sequence similarities. For
details and references, see the text.
Archaea
Nitrate
transporter
Crenarchaeota
Aeropyrum pernix
ND
Sulfolobus solfataricus
ND
Sulfolobus tokadaii
Pyrobaculum aerophilum
Euryarchaeota
Archaeoglobus fulgidus
Nitrate
reductase
Nitrite
reductase
Nitric oxide
reductase
Nitrous oxide
reductase
Nar (genetic)
APE1288 (NarG),
APE1294 (NarH),
APE1300 (NarJ)
Nar? (genetic)
SSO1579 (NarH?),
SSO1580 (NarG?)
Sirohaem-Nir
(genetic)
APE1326 (NirA)
ND
ND
ND
qNor? (genetic)
SSO1573 (N-term.),
SS01571 (C-term.)
ND
ND
ND
ND
ND
ABC (genetic)
PAE3619 (NrtC)
Nar (genetic and
functional)
PAE3610 (NarJ),
PAE3611 (NarG),
PAE3612 (NarH),
PAE3613, PAE2614
(NarI)
Sirohaem-Nir
(genetic)
PAE2577 (NirA)
cd1-Nir (genetic
and functional)
PAE3600, PAE3598
(NirS)
qNor (genetic and
functional)
PAE3603 (NorZ)
Nos? (genetic)
ST2395 (Cu quinol
oxidase), ST0105
(Cu cyt c oxidase)
Nos (functional)
ABC (genetic)
AF0638 (NrtC),
AF0639 (NrtB),
AF0640 (NrtA),
AF0086 (NrtB),
AF0087 (NrtC),
AF0088 (NrtA)
Nar (genetic)
AF0173 (NarJ),
AF0174, AF0175
(NarH), AF0176
(NarG), AF0545
(NarV), AF546
(NarI), AF0501
(NarI)
Nar (genetic and
functional)
CAD22069 (NarG),
CAD22070 (NarH),
CAD22073 (NarJ)
Sirohaem-Nir
(genetic)
AF0164 (NirA)
ND
ND
Cu-Nir (genetic
and functional)
CAB93142 (NirK)
Nor (functional)
ND
Haloarcula marismortui
ND
Halobacterium sp.
ND
ND
ND
ND
Haloferax denitrificans
ND
ND
Cu-Nir (genetic
and functional)
CAD89521 (NirK)
Sirohaem-NirA
(functional)
Cu-Nir or cd1Nir (functional)
Nor (functional)
Haloferax mediterranei
Nar (genetic and
functional)
CAD82168 (NarH)
Ferredoxin-Nas
(functional)
Nar (functional)
Nos? (genetic)
VNG2377G (NosY)
Nos (functional)
ND
ND
Haloferax volcanii
ABC (genetic and
functional)
AJ238877, AJ238878,
AJ238879
Nar (functional)
ND
ND
ND
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P. Cabello, M. D. Roldán and C. Moreno-Vivián
Table 1. cont.
Archaea
Nitrate
transporter
Nitrate
reductase
Nitrite
reductase
Nitric oxide
reductase
Nitrous oxide
reductase
Methanothermobacter
thermautotrophicus
ND
Nas (genetic)
MTH1567 (NasA),
MTH1547
ND
ND
ND
Methanocaldococcus jannaschii
ABC (genetic)
MJ0412 (NrtC),
MJ0413 (NrtB)
ABC (genetic)
MK0601(NrtA),
MK0602 (NrtB),
MK0603 (NrtC)
ND
Sirohaem-Nir
(genetic)
MJ0551 (NirA)
Sirohaem-Nir
(genetic)
MK0801 (NirA)
ND
ND
ND
ND
ND
ND
ND
ND
ND
ND
ND
ND
Nor (genetic)
MM2988
ND
Methanopyrus kandleri
Methanosarcina acetivorans
Methanosarcina mazei
Pyrococcus abyssi
Pyrococcus furiosus
ND
ND
ND
ND
ND
ND
ABC (genetic)
AA113496 (NrtC)
ND
ND
ND
Pyrococcus horikoshii
ND
ND
Sirohaem-Nir?
(genetic)
PF1197 (NirA?)
Sirohaem-Nir
(genetic)
PH0827 (NirA)
ND
ND
Thermoplasma acidophilum
ABC (genetic)
Ta1177 (NrtB),
Ta1178 (NrtC)
ABC (genetic)
TVN0387 (NrtB)
Nas? (genetic)
Ta0136 (Nia)
ND
ND
ND
Nas? (genetic)
TVN0215 (Nia)
ND
ND
ND
Thermoplasma volcanium
Mo-ligand for the Mo-bisMGD cofactor, and also includes
an unusual iron–sulfur cluster, previously undetected
(Bertero et al., 2003; Jormakka et al., 2004). Finally, the
periplasmic Nap systems of many Gram-negative bacteria
participate in different processes, like redox balancing,
scavenging nitrate in nitrate-limited environments, and
aerobic or anaerobic denitrification (Potter et al., 2001;
Gavira et al., 2002; Ellington et al., 2003). Usually, Nap
enzymes are heterodimers with a catalytic subunit (NapA)
and a cytochrome c (NapB), which receives electrons from
NapC, a membrane cytochrome c (Berks et al., 1995; Reyes et
al., 1996, 1998). However, the enzyme of Desulfovibrio
desulfuricans is a single NapA subunit, for which the crystal
structure has been solved (Dias et al., 1999).
Preliminary sequence comparisons of Nar, Nas and Nap
indicated that these enzymes represent different phylogenetic groups (Reyes et al., 1996). A recent study of the
sequences of eukaryotic and prokaryotic nitrate reductases
revealed the existence of three different clades, corresponding to the eukaryotic nitrate reductase, the prokaryotic
respiratory Nar, and the prokaryotic Nas/Nap enzymes,
with the Nap from Desulfovibrio desulfuricans providing
the evolutionary link between Nas and Nap subclades
(Stolz & Basu, 2002). It is worth noting from this study
that the deepest branch of the Nar clade is the euryarchaeote Haloarcula marismortui, and that the Nar enzymes
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of the thermophilic bacterium Thermus thermophilus and
the crenarchaeote Aeropyrum pernix are closely related.
Phylogenetic trees based on NarH sequences also correlate
well with the 16S rDNA tree, making the archaea A. pernix
and Pyrobaculum aerophylum the outgroup and closely
joining these archaeal sequences to the T. thermophilus
bacterial subtree (Petri & Imhoff, 2000; Philippot, 2002).
However, the NarG-based phylogeny is not entirely
consistent with the 16S rDNA-based taxonomy (Chèneby
et al., 2003; Gregory et al., 2003). As the nar genes were
present before the phylogenetic divergence of bacteria and
archaea, it can be assumed that respiratory nitrate reductase
played a key role in energy metabolism during pre-oxic
times (Petri & Imhoff, 2000). In addition, there is evidence
that Nap and Nar have probably been acquired in some
bacteria by horizontal gene transfer (Stolz & Basu, 2002).
This seems to be the case for the nar gene cluster of
T. thermophilus (Ramı́rez-Arcos et al., 1998b) and for the
plasmid-encoded nap genes of Rhodobacter sphaeroides
(Castillo et al., 1996). Results of NarG-based trees are also
consistent with the possibility of lateral gene transfer
(Gregory et al., 2003). Evidence of recent lateral gene
transfer among hyperthermophilic archaea has been published (DiRuggiero et al., 2000), but the relative importance
of this mechanism of genetic exchange among archaea, and
also from and to bacteria, should be determined in future
studies.
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Nitrogen cycle in archaea
Nitrate reductase cofactors
Genome sequences of several archaea contain MGD cofactor
biosynthesis genes, and archaeal nitrate reductases seem to
be molybdoenzymes. However, tungsten and molybdenum
have similar chemistry and it is known that hyperthermophilic archaea possess W-containing enzymes, such as
aldehyde ferredoxin oxidoreductase, with the basic structure
of the pterin cofactor shared in the Mo- and W-enzymes
(Hille, 2002). Recently, Mo-free nitrate reductases with
vanadium and haem c as cofactors have been isolated from
vanadate-reducing, iron-reducing and sulfur-oxidizing bacteria (Antipov et al., 1998, 2003; Murillo et al., 1999). In
addition, active W-substituted molybdoenzymes have been
described in mesophilic bacteria (Buc et al., 1999; Gates
et al., 2003). Thus, tungstate can function at the active
site of the Paracoccus pantotrophus NapA, although the
enzyme shows lower affinity for nitrate (Gates et al., 2003).
Although nitrate reductases and other Mo-enzymes are
usually inactivated by tungstate, this inhibition is not
relevant in vivo since bacteria exhibit high-affinity Mouptake systems for extracting traces (micromolar or
nanomolar) of Mo from the media, and because Mo is
usually present in natural environments at higher concentrations than W. However, Mo(IV) forms a highly
insoluble sulfide (MoS2), while V (as VO2+ or V3+) and W
or WS2{
(as WO2{
4
4 ) form relatively soluble salts, and
therefore there are chemical reasons to believe that both
W and V were more available than Mo in the primitive
strongly reducing sulfide-rich environments (Williams &
Fraústo da Silva, 2002). In addition, some extreme environments such as hot springs and deep-sea hydrothermal vents
usually contain high levels of tungsten, and in fact an
obligate W requirement for the anaerobic growth of the
hyperthermophile Pyrobaculum aerophilum has been reported
(Afshar et al., 1998). The respiratory nitrate reductase of
this archaeon is a Mo-enzyme, but its activity is not
abolished at high W concentrations, in contrast to bacterial
Nar systems. Moreover, the presence of tungstate in the
culture medium stimulates the anaerobic growth with
nitrate (Völkl et al., 1993). The ability to respire nitrate in
these conditions has been interpreted as an adaptation of
the archaeon to the W-rich hyperthermophilic environment (Afshar et al., 1998, 2001). Clearly, further biochemical studies will be necessary to determine the possible
involvement of alternative metal cofactors in archaeal
nitrate reduction, but the possibility that V or W, rather
than Mo, may be included in some archaeal nitrate reductase cofactors cannot be excluded.
Nitrate transport systems
Two types of nitrate transporters are involved in bacterial
assimilatory nitrate reduction: the ATP-dependent ABC
transporters composed of an integral membrane subunit,
a cytoplasmic ATP-binding component and a periplasmic
substrate-binding protein; and the monomeric NarK-type
transporters belonging to the major facilitator superfamily,
which depend on proton-motive force. However, only
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transport systems of the NarK type are involved in bacterial respiratory nitrate reduction (Moir & Wood, 2001).
Within the bacterial NarK-like transporters there are two
distinct subgroups. Recent studies in the bacterium Paracoccus pantotrophus suggest that NarK1 is a proton : nitrate
symporter that allows initiation of nitrate respiration,
whereas NarK2 is a nitrate : nitrite antiporter required for
maintenance of a steady-state rate (Wood et al., 2002).
Genome sequencing projects reveal that ABC-type transporters are widespread in archaea (Table 1). In particular,
putative ABC-type nitrate transporters are present in the
crenarchaeote Pyrobaculum aerophilum (Fitz-Gibbon et al.,
2002) and the euryarchaeota Archaeoglobus fulgidus (Klenk
et al., 1997), Methanocaldococcus jannaschii (Bult et al.,
1996), Methanopyrus kandleri (Slesarev et al., 2002), Thermoplasma acidophilum (Ruepp et al., 2000) and Thermoplasma
volcanium (Kawashima et al., 2000). However, substrate
specificity for these putative transporters has been assigned
solely on the basis of sequence similarities. A functional
characterization of archaeal nitrate transporter genes has
been only performed in Haloferax volcanii, where three
ABC-type transporters are essential for nitrate respiration
(Wanner & Soppa, 1999). The type of transport system used
for nitrate uptake has a serious impact on the bioenergetics of the process because an ATP-driven nitrate uptake
system is equivalent to consuming about three protons and
this energetic cost makes its involvement in dissimilatory
processes unlikely (Moir & Wood, 2001). For this reason,
as mentioned above, bacterial nitrate assimilation usually
requires ATP-dependent ABC nitrate transporters whereas
nitrate respiration is associated with proton-motive-forcedriven NarK transporters (Moreno-Vivián et al., 1999; Moir
& Wood, 2001), even in the extreme thermophilic bacterium
Thermus thermophilus (Ramı́rez et al., 2000). It is worth
noting that most archaea with putative ABC nitrate transporters seem to contain respiratory nitrate reductases rather
than assimilatory Nas enzymes. However, as discussed
below, nitrate uptake may not be required for respiratory
nitrate reduction in archaea, in contrast to bacteria, and the
putative ATP-dependent nitrate transporters may indeed
be involved in nitrate assimilation.
Assimilatory nitrate reduction in archaea
Most nitrate-reducing archaea use nitrate as alternative
electron acceptor in anaerobic respiration, but recently,
assimilatory nitrate and nitrite reductases have been purified
from Haloferax mediterranei (Martı́nez-Espinosa et al.,
2001a, b). This extreme halophilic archaeon grows aerobically with nitrate as the sole nitrogen source, but it can
also denitrify because it contains both Nas and Nar
enzymes. Assimilatory nitrate reductase is a dimer of 105
and 50 kDa subunits and uses ferredoxin as reductant, but
not NAD(P)H (Martı́nez-Espinosa et al., 2001b). This could
indicate a different structure for archaeal Nas because
bacterial ferredoxin-Nas enzymes are usually monomeric,
whereas NADH-dependent Nas enzymes are heterodimers
(Moreno-Vivián et al., 1999). The assimilatory enzyme of
H. mediterranei has a Km for nitrate of 0?95 mM and an
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P. Cabello, M. D. Roldán and C. Moreno-Vivián
optimum temperature of 80 uC at 3?1 M NaCl, but 60 uC
at 1?3 M NaCl. Activity is induced by nitrate and repressed
by ammonium, as described for the bacterial Nas (Martı́nezEspinosa et al., 2001b). The assimilatory nitrite reductase
of H. mediterranei has also been purified. It is a 66 kDa
monomer which uses ferredoxin as electron donor, has a
Km for nitrite of 8?6 mM, and shows maximal activity at
60 uC with 3?3 M NaCl. Like the bacterial assimilatory
nitrite reductases, the H. mediterranei enzyme contains sirohaem and Fe–S centres (Martı́nez-Espinosa et al., 2001a).
The 21 kDa ferredoxin from this halophilic archaeon has
been purified and characterized, and its physiological role as
the electron donor for both assimilatory nitrate and nitrite
reductases has been confirmed (Martı́nez-Espinosa et al., 2003).
Analysis of the published archaeal genome sequences
suggests the presence of putative assimilatory nitrate/nitrite
reductases (Table 1). Thus Methanothermobacter thermautotrophicus has a putative nitrate reductase (MTH1567
gene product) homologous to bacterial Nas (Smith et al.,
1997). Thermoplasma strains contain nitrate reductase
related proteins (T. acidophilum Ta0136 and T. volcanium
VN0215 gene products), which are similar to each other
and to the eukaryotic assimilatory NADPH-nitrate reductases (Kawashima et al., 2000; Ruepp et al., 2000), although
these putative proteins are too small (about 200 residues)
compared to the nia-encoded eukaryotic nitrate reductases.
The eukaryotic nitrate reductases are quite distinct from
any of the bacterial Nas, Nar or Nap enzymes. They share
no sequence similarity and do not bind the Mo-bisMGD
cofactor, but instead have the so-called molybdopterincofactor (Mo-co), in addition to FAD and cytochrome b557
(Moreno-Vivián et al., 1999; Campbell, 2001). Therefore,
the assignation of a role in nitrate reduction to the
Thermoplasma small nitrate-reductase-related proteins will
be of special evolutionary interest and should be explored
in future studies.
The Pyrobaculum aerophilum genome contains a gene (nirA)
encoding a putative assimilatory ferredoxin-dependent
sirohaem nitrite reductase (PAE2577), in addition to putative genes encoding a respiratory cytochrome cd1-nitrite
reductase (Fitz-Gibbon et al., 2002). Putative nirA homologous genes are also present in the genomes of Pyrococcus
furiosus, Pyrococcus horikoshii (PF1197 and PH0827, respectively; Maeder et al., 1999), Aeropyrum pernix (APE1326;
Kawarabayasi et al., 1999), Archaeoglobus fulgidus (AF0164;
Klenk et al., 1997), Methanocaldococcus jannaschii (MJ0551;
Bult et al., 1996), and Methanopyrus kandleri (MK0801;
Slesarev et al., 2002). However, biochemical and functional
studies are necessary to confirm the assimilatory nature of
these putative nitrate and nitrite reductases and to establish
if these archaeal enzymes are structurally different from the
bacterial assimilatory enzymes.
Respiratory nitrate reduction in archaea
The ability to use nitrate as a terminal electron acceptor in
energy metabolism is found in a variety of halophilic and
3532
hyperthermophilic archaea; many of them can perform the
entire denitrification process (see below). Respiratory Nar
enzymes have been purified from several denitrifying halophilic euryarchaeota, including three Haloferax species and
Haloarcula marismortui (Table 1). A salt-requiring Nar has
been purified from Haloferax mediterranei, with an activity
maximum at 89 uC in 3?2 M NaCl and Km for nitrate values
in the range from 2?5 to 6?7 mM depending on salt concentration (Álvarez-Ossorio et al., 1992). The membranebound Nar of Haloferax denitrificans is composed of two
subunits of 116 and 60 kDa, with a Km for nitrate of
0?2 mM. Curiously, the enzyme is stable in the absence of
salt, and activity decreases with increasing NaCl concentrations (Hochstein & Lang, 1991). Haloferax volcanii contains
a membrane-bound trimeric Nar, with 100, 61 and 31 kDa
subunits. Activity is salt-independent, with a temperature
optimum of 80 uC and a Km for nitrate of 0?36 mM (BickelSandkötter & Ufer, 1995). The Haloarcula marismortui
Nar has a Km for nitrate of 80 mM with 2?0 M NaCl, and
activity is enhanced by salt. The enzyme was first described
as a homotetramer of a 63 kDa polypeptide (Yoshimatsu
et al., 2000), but further characterization revealed that it
is a heterodimer with 117 and 47 kDa subunits, the 63 kDa
band being an incomplete denaturation state of the enzyme
(Yoshimatsu et al., 2002). It has been proposed that the
H. marismortui Nar is a new archaeal type of membranebound nitrate reductase (Yoshimatsu et al., 2002). First, the
enzyme is composed of two subunits, which are homologous to the NarGH bacterial subunits, but lacks the NarI
membrane-associated protein. Second, the H. marismortui
NarGH complex has significant sequence and structure
similarity to the dissimilatory selenate reductase from
Thauera selenatis, although it does not reduce selenate.
Recently, the similarity of Nar to selenate reductases has
also been indicated on the basis that both enzymes have
Asp ligands to the Mo atom (Jormakka et al., 2004). In
addition, the N-terminal region of NarG includes a typical
twin-arginine signal peptide for protein translocation across
the membrane by the Tat export pathway (Berks et al., 2000)
and the enzyme has activity in situ with both membranepermeable benzyl viologen and membrane-impermeable
methyl viologen, suggesting that the catalytic site is located
in the outside of the membrane, as described for the T.
selenatis selenate reductase (Yoshimatsu et al., 2002). In
contrast, bacterial NarG faces the cytoplasmic side of the
membrane and nitrate reduction takes places in the
cytoplasm (Berks et al., 1995; Moreno-Vivián et al., 1999).
Finally, although the H. marismortui NarGH complex does
not contain any quinol-oxidizing cytochrome b subunit, a
gene encoding a homologue to SerC, the cytochrome b
subunit of selenate reductase, is found downstream of
the narGH genes and together with a narJ-homologous
gene, which probably encodes a nitrate-reductase-specific
chaperone for MGD cofactor assembly.
A gene cluster encoding a putative respiratory Nar has also
been identified in Archaeoglobus fulgidus (Klenk et al., 1997;
Table 1). The AF0176 gene encodes a protein with 42 %
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Nitrogen cycle in archaea
similarity to Escherichia coli NarG, although this A. fulgidus
NarG protein is smaller (80 kDa) and contains a twinarginine signal peptide, suggesting that it is translocated
across the membrane like the H. marismortui and probably
other archaeal NarG proteins. The AF0175 product is
predicted to bind four Fe–S clusters, whereas the AF0174
gene encodes an integral membrane protein, although no
haem groups are bound to this protein. Finally, a NarJ
homologue is also found in the gene cluster (Klenk et al.,
1997; Richardson et al., 2001). In addition, genes encoding
an ABC-type nitrate transporter (AF0638–AF0640), a putative nirA gene encoding the cytoplasmic sirohaem nitrite
reductase (AF0164) and other putative iron–sulfur proteins
clustered to narI-like genes (AF0499–AF0501 and AF0543–
AF0547) are also present in the A. fulgidus genome. Some
hyperthermophilic archaea are also able to respire nitrate.
Thus, Aeropyrum pernix has a putative nar cluster
(Kawarabayasi et al., 1999) and Pyrobaculum aerophilum
uses nitrate as respiratory electron acceptor (Völkl et al.,
1993). The purified Nar of this archaeon shows very high
specific activity, with a Km for nitrate of 58 mM and an
optimal temperature of 95 uC, and is active in the presence
of high tungstate concentrations. Interestingly, the P.
aerophilum Nar is a heterotrimer, with subunits of 130, 52
and 32 kDa, and contains a b-type cytochrome similar to
the Thauera selenatis SerC, in addition to molybdenum
cofactor and iron–sulfur cluster (Afshar et al., 2001).
Bacterial and archaeal Nar systems also differ in the organization of the nar genes (Philippot, 2002). The main narGHJI
operon organization is conserved in all bacteria, including
Thermus thermophilus (Ramı́rez-Arcos et al., 1998a), although
this organism has an extra narC gene in the operon, and
Paracoccus pantotrophus includes the narK gene in the
narKGHJI cluster (Wood et al., 2001). However, archaeal
nar genes do not conserve this organization. Thus, in
Pyrobaculum aerophilum a putative narJ gene is located
upstream from the narGH genes, in the opposite direction.
A gene encoding a cytochrome b is present downstream
from narH, although its role as Nar subunit is conjectural,
and another putative narI homologue is located elsewhere
in the genome (Fitz-Gibbon et al., 2002). In Haloarcula
marismortui, two small ORFs and a narJ homologue are
located downstream from the narGH genes, and putative
genes encoding an iron–sulfur protein and a cytochrome b
are located upstream from narGH (Yoshimatsu et al.,
2002). In Aeropyrum pernix, no narI-like gene has been
found and narGH genes are followed by a putative cytochrome b gene and narJ, which is located about 2 kb downstream from narH (Kawarabayasi et al., 1999). Therefore,
based on subunit composition, subcellular location of the
active site and nar gene organization, it can be concluded
that archaeal Nars are a new type of enzyme with the active
site facing the outside and anchored to the membrane by a
SerC-like cytochrome b, or stabilized by the lipid environment in the membrane as described for the P. aerophilum
Nar (Afshar et al., 2001). This system could be an ancient
respiratory nitrate reductase, although the nitrite formed
could also be assimilated (Richardson et al., 2001). Figs 2
Fig. 2. Schematic representation of the denitrification pathway in bacteria. The periplasmic nitrate reductase (NAP), the
membrane-bound nitrate reductase (NAR), the periplasmic nitrite reductases (Cu-NIR or cd1-NIR), the membrane-bound nitric
oxide reductase (NOR) and the periplasmic nitrous oxide reductase (NOS) are represented without indicating their subunit
composition and cofactors. Note that not all represented components are present in a single organism (i.e. NAP and NAR,
Cu-NIR and cd1-NIR). cyt bc1, proton-pumping cytochrome bc1 complex; cyt c, NapC membrane-bound tetrahaem
cytochrome c; cyt c550, cytochrome c550; MQH2/MQ, menaquinol/menaquinone pool.
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3533
P. Cabello, M. D. Roldán and C. Moreno-Vivián
Fig. 3. Schematic representation of the denitrification pathway in archaea. Membrane location and dependence on
menaquinol as the electron donor are assumed for the four reductases (nitrate reductase, NAR; nitrite reductase, NIR; nitric
oxide reductase, NOR; and nitrous oxide reductase, NOS), as suggested for Pyrobaculum aerophilum. These reductases are
represented without indicating their subunit composition and cofactors. An outside location for the nitrate reduction site in the
NAR complex is also assumed (see text for details). The identity of the proton-translocating dehydrogenase (DH) that
regenerates menaquinol from menaquinone is presently unknown. The putative ABC-type nitrate/nitrite transporter present in
some archaea is assumed to serve for assimilatory purposes rather than for the respiratory nitrate reduction, since the active
site of NAR is assumed to be located on the outside of the membrane. MQH2/MQ, menaquinol/menaquinone pool.
and 3 schematize the different organization of the bacterial
and archaeal respiratory nitrate-reducing systems, and the
other enzymes involved in the whole denitrification pathway
(see below).
archaeal species would be probably involved in nitrate
assimilation rather than in nitrate respiration.
The possible outside location of the catalytic site of NarG
in the archaeal respiratory nitrate reductase has important
bioenergetic implications. First, bacterial nitrate respiration involves the generation of a proton-motive force by a
redox-loop mechanism, with two protons consumed in the
nitrate reduction reaction in the cytoplasm and two protons
released from quinol oxidation on the periplasmic side of
the membrane (Berks et al., 1995; Jormakka et al., 2003),
although periplasmic nitrate reduction by the Nap enzyme
and other reductases involved in the denitrification pathway are coupled to proton-pumping protein complexes
(Fig. 2). However, the possible outside location of the
nitrate reduction site in archaeal Nar would imply the
coupling of this process to a proton-translocating complex,
although the identity of this complex is presently unknown
(Fig. 3). The genome sequence of Pyrobaculum aerophilum
reveals that this organism apparently lacks a cytochrome
bc1 complex (Fitz-Gibbon et al., 2002), but the occurrence
of membrane-bound bc complexes and different protonpumping NADH dehydrogenases and hydrogenases has
been described in archaea (Castresana & Moreira, 1999;
Deppenmeier et al., 1999; Schütz et al., 2000; Sapra et al.,
2003). In addition, an active nitrate-uptake system would
not be required for respiratory nitrate reduction in
archaea, thus increasing the energetic yield of the nitrate
reduction process. This could also explain the apparent
absence of NirK-type nitrate transporters in archaea and
suggests that, as indicated above, the putative ATPdependent ABC-type nitrate transporters found in several
Denitrification can be considered as the modular assemblage of four partly independent respiratory processes:
nitrate, nitrite, nitric oxide and nitrous oxide reduction
(Zumft, 1997; Figs 2 and 3). All denitrifiers isolated so far
are also able to respire oxygen, indicating a close evolutionary relationship between the two processes, and denitrification has been considered the ancestor of aerobic
respiration (Saraste & Castresana, 1994). Therefore, knowledge of denitrification in archaea is of special interest from
an evolutionary point of view. Denitrification has been
described for several halophilic archaea, such as Haloferax
and Haloarcula strains (Tomlinson et al., 1986; Inatomi &
Hochstein, 1996; Ichiki et al., 2001), and extreme thermophilic archaea, such as Ferroglobus placidus (Vorholt et al.,
1997) and Pyrobaculum aerophilum (Völkl et al., 1993; de
Vries & Schröder, 2002). As the archaeal respiratory nitrate
reductases that catalyse the first step of denitrification have
been described above, we will focus now on the nitrite and
N-oxide respiration processes in archaea.
3534
Denitrification in archaea
Nitrite respiration
In bacteria, there are two different and unrelated NOproducing respiratory nitrite reductases: the homotrimeric
copper-containing enzyme encoded by nirK, and the
homodimeric cytochrome cd1-nitrite reductase encoded
by nirS. The crystal structures of both enzymes, which are
located in the periplasm, are known and have been
reviewed in detail elsewhere (Zumft, 1997; Ferguson,
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Nitrogen cycle in archaea
1998; Richardson & Watmough, 1999). Azurin and pseudoazurin are the electron donors to Cu-Nir, whereas cytochrome c551 is the reductant for cd1-Nir (Zumft, 1997).
Denitrifying archaea also contain either Cu-Nir or cd1-Nir.
Haloarcula marismortui has a nirK gene encoding a CuNir. EPR analysis reveals the presence of type 1 and type 2
Cu centres in the purified enzyme, which has optimal
activity at 2 M NaCl. A 46 kDa precursor of this protein
with an N-terminal signal peptide for translocation across
the plasma membrane has been detected by SDS-PAGE
(Ichiki et al., 2001). Although the physiological electron
donor is unknown, the archaeal halocyanin, a blue Cuprotein present in some archaea (Scharf & Engelhard,
1993), could be the putative reductant. Protein sequence
analysis of the H. marismortui nirK gene product reveals a
close relationship with the Neisseria gonorrhoeae Cu-Nir,
thus suggesting lateral gene transfer of the nirK gene
(Ichiki et al., 2001; Philippot, 2002). Interestingly, the
Haloferax denitrificans nirK gene sequence is also available
(CAD89521, Table 1), and is highly similar to both H.
marismortui and N. gonorrhoeae Cu-Nir. However, antiserum against bacterial Cu-Nir does not recognize the H.
denitrificans enzyme, suggesting possible structural differences (Inatomi & Hochstein, 1996).
et al., 1993). In most denitrifying bacteria, Nor is a
membrane complex of a 17 kDa cytochrome c (encoded
by norC) and a 38 kDa cytochrome b with 12 transmembrane regions (encoded by norB). This enzyme,
known as cNor, receives electrons from cytochrome c.
The denitrifying bacterium Ralstonia eutropha and several
non-denitrifying pathogenic bacteria such as Neisseria
species possess a monomeric Nor with 14 transmembrane
regions, which is called qNor because it is a quinoloxidizing enzyme. This protein, encoded by norZ, is
similar to the norB gene product but contains an Nterminal extension absent in NorB, with a quinone-binding
site. In Gram-positive bacteria, a menaquinol Nor with
an additional small CuA-containing subunit has been
described as qCuANor (Hendriks et al., 2000; de Vries &
Schröder, 2002; Philippot, 2002).
N-oxide respiration
Genetic and biochemical evidence indicates that the
archaeon Pyrobaculum aerophilum has a qNor-type NO
reductase (Table 1). Putative nir (nitrite reductase) and
nor (nitric oxide reductase) genes are located 3?3 kb
upstream from the nar (nitrate reductase) genes (Völkl
et al., 1993; Fitz-Gibbon et al., 2002). The purified Nor
is a single membrane subunit with two modified haem
groups and one non-haem iron, and uses menaquinol as
electron donor (de Vries & Schröder, 2002). The genome
of Methanosarcina mazei also contains a NO reductase
gene (MM2988; Deppenmeier et al., 2002) and Sulfolobus
solfataricus includes a putative gene encoding a haem NO
reductase, but it is truncated by a transposase separating
the N-terminal (SSO1573) and C-terminal (SSO1571)
domains (She et al., 2001). Sequenced genomes of other
archaea do not include nor homologous genes (Table 1),
although the presence of NO reductase has been reported
in some denitrifying halophilic archaea that have not yet
been sequenced. The presence of qNor in some archaea is of
evolutionary interest because qNor could be the ancestor
not only of the cNor, by acquiring haem c-binding residues
in the N-terminal region and separating this domain as a
small cytochrome c subunit, but also of the whole superfamily of haem-copper oxidases and aerobic respiratory
enzymes. The capacity for N-oxide respiration appeared
early in evolution and qNor may have arisen to detoxify
NO; this could still be the role of the enzyme in nondenitrifying bacteria. After the appearance in the atmosphere of oxygen generated photosynthetically, these
enzymes could have replaced non-haem iron by copper
and evolved to generate the proton-pumping cytochrome
oxidases (Saraste, 1994; Saraste & Castresana, 1994;
Richardson, 2000; de Vries & Schröder, 2002).
Nitric oxide, the product of the respiratory nitrite reductases, is a toxic compound that is reduced to N2O by nitric
oxide reductases (Nor) immediately after it has been
generated. Several enzymes with Nor activity have been
described. In fungal denitrification, NO reduction to N2O
is catalysed by a soluble monomeric enzyme of the cytochrome P-450 family (Shoun & Tanimoto, 1991; Nakahara
On the other hand, the E. coli flavorubredoxin has been
recently described as a novel type of prokaryotic NO
reductase. The purified protein has a turnover number
similar to that of canonical bacterial Nor and also has
oxygen reductase activity, although it shows lower affinity
for O2 than for NO (Gomes et al., 2002). Flavorubredoxins
are A-type flavoproteins which are present in bacteria and
The existence of a nirS homologous gene in the genome
of the hyperthermophile Pyrobaculum aerophilum suggests
the presence of a cd1-nitrite reductase (Table 1). Recently, it
has been reported that this archaeon has a membranebound respiratory nitrite reductase which uses menaquinol
as the physiological electron donor, thus having different
subcellular location and reductant than bacterial cd1-Nir
(Figs 2 and 3). It is worth noting that, in contrast to
denitrifying bacteria, all four reductases involved in denitrification in P. aerophilum are membrane-bound enzymes
which use menaquinol as electron donor (de Vries &
Schröder, 2002). The dependence of the menaquinol pool
and the membrane confinement of all these enzymes in
archaeal denitrification could be related to the relatively
small volume between the S-layer and the cytoplasmic
membrane, in comparison with the larger periplasmic
space in bacteria, or could reflect the ancient origin of this
pathway. The membrane location of all denitrification
enzymes in P. aerophilum could also explain the absence in
this archaeon of soluble components of the denitrification
pathway, such as c-type cytochromes or cupredoxins
(Lubben & Morand, 1994; de Vries & Schröder, 2002; see
Figs 2 and 3).
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3535
P. Cabello, M. D. Roldán and C. Moreno-Vivián
some archaea, such as Pyrococcus furiosus, Archaeoglobus
fulgidus, Methanocaldococcus jannaschii and Methanothermobacter thermautotrophicus. Therefore, it has been
proposed that this protein is a widespread class of NO
reductase, which could detoxify NO or oxygen, allowing
versatility in response to environmental conditions (Gomes
et al., 2002). However, the physiological relevance of this
protein should be determined in future studies.
The last step of denitrification, the reduction of N2O to N2,
is of great environmental importance because it closes the
N-cycle. Although N2O is less toxic than NO or nitrite, and
bacteria could manage without converting N2O into N2,
most denitrifying bacteria contain nitrous oxide reductases (Nos) encoded by the nosZ gene. These enzymes are
periplasmic multicopper homodimers that receive electrons
from cytochrome c or pseudoazurin (Zumft, 1997). In each
65 kDa monomer, the copper ions are organized in two
centres, a di-copper cluster CuA resembling that of cytochrome oxidase, and a CuZ cluster in which four Cu
atoms are ligated by seven His residues. A bridging inorganic
sulfur in the CuZ centre has also been identified (Brown
et al., 2000; Rasmussen et al., 2000). In some archaea, such
as Haloarcula marismortui and Ferroglobus placidus, the
predominant gas species produced by denitrification is
N2O (Werber & Mevarech, 1978; Vorholt et al., 1997),
although other archaea such as Haloferax denitrificans and
Pyrobaculum aerophilum are able to reduce N2O to N2,
having a complete denitrification pathway (Tomlinson
et al., 1986; Völkl et al., 1993; de Vries & Schröder, 2002).
However, in P. aerophilum, N2O formed by qNor accumulates because its reduction to N2 by the nitrous oxide
reductase is very slow. In contrast to the periplasmic
bacterial Nos, the P. aerophilum enzyme is located in the
membrane and uses menaquinol as electron donor, like
the other three reductases of the denitrification pathway
in this hyperthermophile (de Vries & Schröder, 2002;
Fig. 3). N2O reduction has not been investigated in other
species of archaea and putative nos genes are not present
in the sequenced archaeal genomes (Table 1), although two
long hypothetical quinol oxidases (ST2395 and ST0105)
with the motif of the Nos binuclear copper centre are found
in Sulfolobus tokodaii (Kawarabayasi et al., 2001).
N2 fixation in archaea
N2 is the most abundant gas in Earth’s atmosphere, but
it is highly stable and unreactive and is only used as nitrogen source by some prokaryotic organisms. Biological N2
fixation is catalysed by nitrogenase, an O2-sensitive enzyme
complex which generates two molecules of NH3 by reduction of N2. Nitrogenase can also reduce protons to H2 and
several substrates with double or triple bonds, such as
acetylene and cyanide (Halbleib & Ludden, 2000). Nitrogenase consists of a molybdenum–iron protein (dinitrogenase), which receives electrons from an iron protein
known as dinitrogenase reductase. MoFe protein is an a2b2
tetramer encoded by the nifD and nifK genes, and Fe protein
is a homodimer of the nifH gene product (Howard & Rees,
3536
1996). Several nif genes are also involved in cofactor
biosynthesis, electron transfer and regulation. Alternative
nitrogenases have been found in several bacteria, with V or
Fe substituting for Mo. These enzymes have lower catalytic
efficiency than Mo-nitrogenases and are encoded by nif
homologous genes, termed vnf (V-nitrogenase) or anf
(Fe-nitrogenase). Alternative nitrogenases include an additional small subunit d, encoded by vnfG or anfG (Eady,
1996; Rees, 2002). N2 fixation is highly regulated since is
an energy-expensive process requiring large amounts of
reducing power and ATP. Ammonium causes a reversible
short-term inhibition of enzyme activity, called switch-off/
on (Zumft & Castillo, 1978), and a long-term effect at
the transcriptional level. Ammonium switch-off occurs
by ADP-ribosylation of NifH by DraT/DraG, an ADPribosyltransferase/glycohydrolase system. Expression of nif
genes requires NifA, a transcriptional activator under the
control of Ntr, a two-component regulatory system of
bacterial nitrogen metabolism (Ludden, 1994; Merrick &
Edwards, 1995; Halbleib & Ludden, 2000).
N2 fixation in archaea, first described in Methanosarcina
barkeri (Murray & Zinder, 1984) and Methanococcus
thermolithotrophicus (Belay et al., 1984), is exclusive of,
but widespread within, methanogenic euryarchaeota
(Table 2). However, neither Methanococcus voltae nor
Methanocaldococcus jannaschii grows diazotrophically,
despite the presence of nifH-like genes in their genomes
(Leigh, 2000). Mo-nitrogenase predominates in diazotrophic methanogens, which contain at least the nifHDKENX
genes (Lobo & Zinder, 1990; Chien & Zinder, 1996; Kessler
et al., 1998; Leigh, 2000). nifHDK are the structural genes
for nitrogenase, whereas nifENX code for FeMo-cofactor
biosynthesis proteins (Moreno-Vivián et al., 1989). The
diazotrophic growth of Methanococcus maripaludis shows
Mo dependence, indicating that N2 fixation is catalysed by
a Mo-nitrogenase (Kessler et al., 1997), although Methanosarcina species contain alternative nitrogenases (Table 2).
M. barkeri has two clusters of nitrogenase genes, the nif2
gene cluster encoding the Mo-nitrogenase and the nif1 (vnf )
cluster encoding an alternative V-nitrogenase, which also
includes a vnfG gene (Chien et al., 2000). Like the nif regions
of other methanogens, both gene clusters contain two
glnB genes encoding nitrogen signal transduction proteins
(Chien et al., 2000). The M. acetivorans genome includes nif,
vnf and anf genes, being unique among archaea in having
all three nitrogenase systems (Galagan et al., 2002).
There are significant differences in gene organization and
regulation between bacterial and archaeal nitrogenase
systems. First, all nif genes and the glnB genes are cotranscribed in a single unit (nifHglnBIglnBIInifDKENX), this
being a gene organization unique to archaea (Kessler et al.,
1998; Leigh, 2000). Transcription of nif genes occurs from
typical archaeal promoters, probably reflecting the differences between archaeal and bacterial transcription apparatus, and is regulated by repression, whereas an activation
mechanism involving NifA operates in bacteria. As stated
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Table 2. Presence of genes encoding ammonium transporters and proteins involved in N2 fixation and ammonium assimilation in the completely sequenced genomes of
archaea
Genome sequence analysis was conducted on websites http://gib.genes.nig.ac.jp and http://www.genome.ad.jp and the accession numbers of the corresponding genes in the different
genomes are indicated. ND indicates that putative genes encoding these proteins or enzymes have not been detected. For details and references, see the text.
Archaea
Crenarchaeota
Aeropyrum pernix
Ammonium
transporter
Nitrogenases
N2-fixation-related
proteins
ND
ND
Sulfolobus solfataricus
AmtB
SSO1054
ND
NifS
SSO2215
Sulfolobus tokadaii
AmtB
ST0661
ND
NifW
STS185
NifB-like ST1383, ST2412
Pyrobaculum aerophilum
ND
ND
ND
AmtB
AF0977, AF1746,
AF1749
ND
Halobacterium sp.
ND
ND
DraG
AF1724
NifU AF0185, AF0565, AF0632
NifS AF0186, AF0564
NifU
VNG2472G
NifS VNG2471G
Methanothermobacter
thermautotrophicus
AmtB
MTH661, MTH663
Methanocaldococcus
jannaschii
AmtB
MJ0058, MJ1343
Methanopyrus kandleri
AmtB
MK0053
Nif
MTH571, MTH643, MTH1711,
MTH1560 (NifH), MTH1522,
MTH1563 (NifD), MTH1564
(NifK), MTH1482, MTH1565
(NifE), MTH1566 (NifN)
Nif
MJ0879 (NifH), MJ0685 (NifH)
Vnf
MJ1423 (VnfN)
Nif
MK1385 (NifD), MK1416 (NifH)
Euryarchaeota
Archaeoglobus fulgidus
GlnA
APE2142
GlnA
SSO0366, SSO2440,
SSO2554
GlnA
ST0156, ST0568,
ST1387
GlnA
PAE2556, PAE3442
Glutamate
synthase
Small subunit
APE0935
Large subunit
SSO0684 (GltB)
Large subunit
ST2198
Glutamate
dehydrogenase
GdhA
APE1386
GdhA
SSO1457, SSO1907,
SSO1930, SSO2044
GdhA1
ST0100, ST224
Small subunit
PAE3227 (GltD),
PAE3443,
PAE3444
GdhA
PAE1996, PAE3438
GlnA
AF0949
Large subunit
AF0953 (GltB)
ND
GlnA
VNG2093G
ND
GdhAB
VNG0161G (GdhB),
VNG0628G (GdhA),
VNG1204G (GdhA)
GlnB-like
MTH1561, MTH1562
NifB-like MTH1871
NifS MTH1389
GlnA
MT1570
Large subunit
MTH105, MTH194,
MTH1666
ND
DraG
MJ1187
NifB-like
MJ1093
NifB-like
MK0243, MK0650, MK1488
GlnA
MJ1346
Large subunit
MJ1351 (GltB)
ND
GlnA
MK0338
Large subunit
MK0550, MK1063,
MK1064 (GltB)
ND
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Nitrogen cycle in archaea
ND
Glutamine
synthetase
Archaea
Microbiology 150
Ammonium
transporter
Nitrogenases
N2-fixation-related
proteins
Glutamine
synthetase
Methanosarcina
acetivorans
AmtB
MA3917, MA3918,
MA4207
Nif
MA1633, MA2032, MA2033, MA3627,
MA3628, MA3895 (NifH), MA3898
(NifD), MA3899 (NifK), MA3900
(NifE), MA3901 (NifN)
Vnf
MA1213 (BNF), MA1216 (VnfD),
MA1217 (VnfG), MA1218 (VnfK),
MA1219 (VnfE), MA1220 (VnfN),
MA1631 (VnfE), MA1632 (VnfN)
Anf
MA1205 (AnfH), MA1208 (AnfK),
MA1209 (AnfG), MA1210 (AnfD)
GlnA
MA3382, MA4216
Small subunit
MA3787
GdhA
MA2892, MA3169
Methanosarcina mazei
AmtB
MM0733, MM0957
Nif
MM0514, MM0687, MM0719,
MM2086 (NifH), MM0722 (NifD),
MM0723 (NifK), MM0724 (NifE),
MM0725 (NifN)
GlnB-like
MA1211, MA1212, MA1214,
MA1215, MA3916, MA4208
DraG
MA4144
NifU
MA0807, MA1225, MA1226,
MA2217, MA3265
NifS
MA0236
NifB-like
MA0175, MA1114, MA1486,
MA2936, MA3185, MA3448,
MA3482, MA4195
GlnB-like
MM0720, MM0721
NifS
MM1517, MM1955
NifU
MM0110, MM1954
NifB-like
MM0309, MM0757, MM0759,
MM0788, MM1105, MM1474,
MM2468
GlnA
MM0964, MM3188
Large subunit
MM0966, MM0967,
MM0968
Small subunit
MM0664
GdhA
MM0357, MM3297,
MM3298
Pyrococcus abyssi
ND
ND
ND
Pyrococcus furiosus
ND
ND
NifH-like
PF2028 (N-term.), PF2029
(C-term.)
NifS
PF0164
GlnA
PAB1292
GlnA
PF0450
GdhA
PAB0391
GdhA
PF1602
Pyrococcus horikoshii
ND
ND
ND
Thermoplasma acidophilum
AmtB
Ta0180, Ta0181
AmtB
TVG1095610
ND
ND
ND
ND
Small subunit
PAB1214, PAB1738
Small subunit
PF1327, PF1852,
PF1910
Large subunit
PF0205
Small subunit
PH0876, PH1873
Small subunit
Ta0414
Small subunit
TVG1211894
Thermoplasma volcanium
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GlnA
PH0359
GlnA
Ta1498
GlnA
TVG1571797
Glutamate
synthase
Glutamate
dehydrogenase
GdhA
PH1593
GdhA
Ta0635, Ta0776
GdhA
TVG0773051,
TVG0774569
P. Cabello, M. D. Roldán and C. Moreno-Vivián
3538
Table 2. cont.
Nitrogen cycle in archaea
below, a repressor NrpR found only in euryarchaeota
regulates nif and glnA (glutamine synthetase) gene expression (Lie & Leigh, 2003). Finally, post-transcriptional
regulation by ammonium (switch-off) also occurs in
archaea (Lobo & Zinder, 1990; Kessler & Leigh, 1999;
Kessler et al., 2001), but does not involve ADP-ribosylation
or covalent modification of NifH and requires two
members of the GlnB family, termed NifI1 and NifI2
(Kessler et al., 2001).
Phylogenetic analysis suggests that N2 fixation may have
originated in a common ancestor of bacteria and archaea,
before branching of these domains and prior to the appearance of oxygen in the atmosphere (Fani et al., 2000; Leigh
2000; Berman-Frank et al., 2003). Comparative analysis of
the amino acid sequences of the NifDK and NifEN proteins
suggests a two-step in-tandem duplication of an ancestor
gene, giving rise first to a bicistronic operon, which later
underwent a paralogous duplication leading to the nifDK
and nifEN genes before the divergence of archaea and
bacteria (Fani et al., 2000). The function of this ancient
enzyme might have depended on the composition of the
early atmosphere, being either a nitrogenase for N2 fixation in a neutral anoxic atmosphere containing N2 but not
NH3, or alternatively, a cyanide-detoxifying enzyme in a
strongly reducing atmosphere, which could change its
specificity to N2, another triple-bond substrate, after progressive exhaustion of combined nitrogen, thus enabling
survival of ancient micro-organisms in N-deficient environments before the appearance of photosynthetic O2 (Fani
et al., 2000; Berman-Frank et al., 2003). After the divergence
of archaea and bacteria, many organisms lost nif genes
due to the weakening of selection pressure and the high
energetic cost of N2 fixation, although lateral gene transfer
of nif genes might have also occurred (Fani et al., 2000).
However, there are no apparent reasons to explain the lost
of diazotrophy in all archaea except methanogens. Earth’s
earliest organisms may have been similar to the thermophiles found in deep-sea hydrothermal vents, but currently
the most thermophilic micro-organism known, Methanococcus thermolithotrophicus, is able to fix N2 at 64 uC (Mehta
et al., 2003). This makes especially interesting a recent paper
describing the first evidence of potential nitrogen fixers in
diffuse hydrothermal vent fluids and deep-sea water, and
reporting that all of the nifH genes from these samples are
most closely related to M. thermolithotrophicus (Mehta et al.,
2003).
Transport and assimilation of ammonium in
archaea
Glutamine synthetase–glutamate synthase (GS–GOGAT)
and glutamate dehydrogenase (GDH) are the major pathways for ammonium assimilation. The GS–GOGAT route
requires ATP but has a high affinity for ammonium, whereas
GDH does not consume ATP but is less effective in cells
growing in N-limited conditions. As these enzymes are
essential in ammonia assimilation and amino acid metabolism, they are present in all three domains of life. Movement
http://mic.sgmjournals.org
of ammonium across biological membranes is also an
important process, and ammonium transporters (Amt) are
found in bacteria, archaea and eukarya.
Ammonium transport
N2 fixation and nitrate assimilation generate ammonium
for incorporation into carbon skeletons inside the cells.
However, when ammonium is available in the environment, organisms usually take up this ion by a specific
transport system, inhibiting and/or repressing the reductive assimilatory pathways of the N-cycle. Although NH3
diffuses through biological membranes, and this could
represent a significant process in organisms growing in
alkaline environments, ammonium transporters are found
in both prokaryotic and eukaryotic organisms. Amt proteins are a family of ammonium and methylammonium
transporters, which are integral membrane proteins of 45–
50 kDa with 10–12 transmembrane helices (Thomas et al.,
2000). It is widely accepted that Amt proteins mediate an
active NHz
4 uniport which depends on proton-motive
force, although it has been also proposed that Amt proteins
facilitate diffusion of NH3 across the plasma membrane
bidirectionally (Soupene et al., 2002). The E. coli AmtB
protein is essential for transport of ammonium and is able
to complement a Saccharomyces cerevisiae mutant (Blakey
et al., 2002). Putative ammonium transporters (AmtB) are
also present in most archaeal genomes (Table 2). In bacteria
and archaea, the amtB gene is invariably linked to the
regulatory glnK gene, which encodes a GlnB-like protein
(PII protein). Both GlnK and AmtB proteins interact for
regulation of the transport activity in response to nitrogen
availability (Thomas et al., 2000; Coutts et al., 2002). In
E. coli, AmtB is inactivated by formation of a membranebound complex with GlnK, a process regulated by uridylylation/deuridylylation of GlnK in response to the nitrogen
status of the cells, and the AmtB activity is also required
for GlnK deuridylylation. Therefore, AmtB could be considered as an ammonium sensor, and transport is also an
integral part of the signal transduction cascade (Javelle et al.,
2004).
The glutamine synthetase–glutamate synthase cycle
Glutamine synthetase (GS) is a ubiquitous enzyme with a
central role in nitrogen metabolism. Three types of GS
catalyse the ATP-dependent synthesis of glutamine from
glutamate and ammonia. GSI, found in most prokaryotes,
is a dodecameric protein composed of identical 50–56 kDa
subunits arranged in two hexagonal rings. GSII is mainly
present in eukaryotes and has an octameric structure, with
identical subunits of 45–50 kDa. GSIII is only found in a
few bacterial species, such as Bacteroides fragilis, and has
a hexameric structure with identical subunits of about
83 kDa (Pesole et al., 1995; DiRuggiero & Robb, 1996;
Eisenberg et al., 2000).
Putative glnA genes encoding GSI are present in archaeal
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3539
P. Cabello, M. D. Roldán and C. Moreno-Vivián
genomes, sometimes in several copies (Table 2). Sequence
analysis of the glnA gene reveals two main subdivisions:
GSI-a, including euryarchaeota, Thermotoga and low-G+C
Gram-positive bacteria; and GSI-b, with all other bacteria. Biochemical properties also indicate that GSs from
archaea and Gram-positive bacteria are functionally similar
(Bhatnagar et al., 1986). Lateral gene transfer from archaea
to Thermotoga and Gram-positive bacteria could explain the
close evolutionary relation between these GSs (Tiboni et al.,
1993; Pesole et al., 1995). However, the crenarchaeote
Sulfolobus solfataricus GS is not placed in either the GSI-a
or the GSI-b division (Brown et al., 1994), although the
Sulfolobus acidocaldarius glnA is more related to GSI-b,
despite its enzyme properties being more similar to GSI-a
(Yin et al., 1998). Most GSI-b enzymes are regulated by
adenylylation/deadenylylation, but GSI-a enzymes are not
controlled by this reversible mechanism. Purified GSs from
Pyrococcus sp., S. acidocaldarius and Methanobacterium
ivanovi are dodecamers of identical subunits that are not
regulated by adenylylation (Bhatnagar et al., 1986; Rahman
et al., 1997; Yin et al., 1998). Therefore, branching of GSI-a
and GSI-b could precede the separation of archaea and
bacteria.
The relevance of GS in ammonium assimilation varies
depending on the archaeal species. Most hyperthermophilic
archaea show very high glutamate dehydrogenase activity,
suggesting that this enzyme is the major pathway for
ammonium assimilation. In fact, Thermococcus profundus
has no detectable GS activity (Kabayashi et al., 1995) and
the GS of Pyrococcus sp. has low biosynthetic activity,
suggesting that the reaction is biased towards glutamate
production. Therefore, hyperthermophilic archaea seem
not to use GS predominantly for ammonium assimilation.
In contrast, the inability to obtain a glnA null mutant in
Methanococcus maripaludis suggests that this archaeon
cannot transport glutamine and that GS is an essential
enzyme, even in the presence of alternative nitrogen sources
(Cohen-Kupiec et al., 1999).
Glutamate synthase (glutamine : 2-oxoglutarate amidotransferase, GOGAT) is an iron–sulfur flavoprotein that
catalyses the transfer of the amide group of glutamine to
2-oxoglutarate, yielding two molecules of glutamate. In
bacteria, GOGAT is usually an NADPH-dependent enzyme
arranged in an a4b4 structure, with large a subunits of
about 135 kDa and small b subunits of approximately
53 kDa (Trotta et al., 1974). The genomes of most archaea
contain putative genes encoding either the large or the
small GOGAT subunit (Table 2). The gltA gene encoding
a putative small GOGAT subunit in the hyperthermophile
Pyrococcus sp. has been cloned. The purified GltA protein
is a tetramer with identical subunits of 53 kDa. This
GOGAT uses NADPH and has both glutamine- and
ammonium-dependent activities, showing different temperature optima, 80 uC for the glutamine reaction and 90 uC
for the ammonium-dependent reaction (Jongsareejit et al.,
1997). The amino acid sequence of this GOGAT includes
3540
two Cys clusters, and adenylate- and FAD-binding domains,
which are conserved in the small subunits of bacterial
GOGAT. It is worth noting that Pyrococcus sp. lacks the
large subunit gene and GOGAT is encoded by the gltA gene
alone. In contrast, genomes of other archaea include only
the gltB gene encoding the large subunit (Table 2). Methanocaldococcus jannaschii has only a gltB gene encoding a
55 kDa protein, which includes features of the bacterial
large subunit and probably has GOGAT activity (Bult
et al., 1996). It has been proposed that both M. jannaschii
GOGAT (GltB) and Pyrococcus sp. GOGAT (GltA) represent
ancestral forms of the large and small subunits, respectively,
which are functional without their subunit counterpart
(Jongsareejit et al., 1997). On the other hand, in eukaryotes,
GOGAT is a monomeric enzyme homologous to either the
large bacterial subunit, as the maize ferredoxin-GOGAT,
or to both large and small bacterial subunits, as the alfalfa
NADH-GOGAT (Jongsareejit et al., 1997).
Glutamate dehydrogenase
Glutamate dehydrogenase (GDH) catalyses both the oxidative deamination of glutamate and the reductive incorporation of ammonium into 2-oxoglutarate. There are three
types of GDH based on coenzyme specificity. Dual NAD(P)GDHs are found in animals and higher plants, but NADPGDHs are mostly involved in ammonia assimilation in
bacteria, fungi and algae. Both NADP-dependent and dual
NAD(P)-GDH are hexamers composed of 48–55 kDa
identical subunits. On the other hand, NAD-GDHs consist
of either four or six identical subunits and usually have a
catabolic role, being involved in the oxidative deamination
of glutamate (Smith et al., 1975).
Most hyperthermophilic archaea studied to date express
high levels of GDH, and it is assumed that GDH plays a
major role in ammonium assimilation in these extremophiles (DiRuggiero & Robb, 1996). The enzymes from
Pyrococcus furiosus, Thermococcus litoralis, Thermococcus
profundus and Sulfolobus solfataricus are hexameric, as in
mesophilic bacteria, and show a high temperature optimum for activity. However, the coenzyme specificity of
GDH from these hyperthermophiles resembles that of the
eukaryotes rather than bacteria, since the enzyme uses
both NADH and NADPH (DiRuggiero & Robb, 1996). The
three-dimensional structure of the P. furiosus NAD(P)GDH has been determined at 2?2 Å resolution (Yip et al.,
1995). The gdhA genes encoding GDH have been cloned or
identified in the genome of several archaea (Table 2). The
P. furiosus gdhA gene is transcribed from a typical archaeal
promoter, but it has been expressed in E. coli, yielding a
recombinant GDH fully active in vitro at 85 uC (DiRuggiero
& Robb, 1995). Sequencing of the Pyrococcus horikoshii
genome revealed the presence of a gdhA gene homologous
to the GDH-encoding genes of P. furiosus and T. litoralis.
Recently, this gdhA gene has been cloned and a recombinant His6-tagged GDH was overexpressed in E. coli and
purified. The enzyme is a hexamer of 290 kDa (subunit
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Nitrogen cycle in archaea
mass 48 kDa), requires NAD(P)H and has a temperature
optimum of 90 uC (Wang et al., 2003). This P. horikoshii
GDH shows a lower Km value for glutamate than for
ammonium, suggesting that the reaction is biased toward
oxidative deamination of glutamate. NADP is the preferred
coenzyme for this reaction. The enzyme subunit contains
many charged residues that at high temperature arrange
networks of ion pairs for proper folding and maintaining
the native conformation of the enzyme (Wang et al., 2003).
The GDH enzyme of Aeropyrum pernix has also been
characterized (Bhuiya et al., 2000; Helianti et al., 2001,
2002). The enzyme also has a hexameric structure with
a molecular mass of about 285 kDa, and is specific for
NADP. This GDH is thermostable, and the N-terminal
length seems to be important for its thermostability
(Helianti et al., 2002). The A. pernix enzyme has biphasic
kinetics for glutamate, but monophasic kinetics for NADP
at 50–90 uC, and it has also been crystallized and analysed
by X-ray diffraction (Bhuiya et al., 2002a, b).
regulators involved in bacterial nitrate reduction, such as
FNR, NNR, NorR, NarR, NarXL, ResDE and RegAB, are not
present in the sequenced archaeal genomes, thus suggesting
that completely novel regulatory systems could operate in
archaea, probably as a consequence of the fundamental
differences in the transcription machinery between bacteria
and archaea. However, although most archaeal helix–turn–
helix-containing transcription factors belong to archaeaspecific families, it has recently been described that the
molybdenum-responsive transcription factor ModE is
present in some archaea. Moreover, the upstream ModE
DNA-binding sequence seems to be conserved between
bacteria and archaea (Studholme & Pau, 2003). These results
indicate that Mo also regulates the expression of the genes
required for archaeal molybdoproteins, and suggest either
an ancient common origin for the ModE regulon, or more
likely, horizontal gene transfer probably between green
sulfur bacteria and methanogenic archaea (Studholme &
Pau, 2003).
Protein sequence analysis indicates that the enzymes from
Pyrococcus furiosus and Pyrococcus horikoshii are more
closely related to the mesophilic bacterial GDH than to
the thermophilic archaeon Sulfolobus solfataricus or to the
halophilic archaeon Halobacterium salinarum (DiRuggiero
& Robb, 1996), suggesting that lateral gene transfer between
bacterial and archaeal gdhA genes may have occurred during evolution. Some organisms have two or more GDHs,
which differ in coenzyme specificity. This is the case for H.
salinarum, which has both NADP- and NAD-dependent
enzymes (Bonete et al., 1987; Hayden et al., 2002). The
NAD-enzyme of H. salinarum has been characterized and
is subject to activation by amino acids and inhibition by
TCA cycle intermediates such as fumarate, oxalacetate,
succinate and malate (Bonete et al., 1996). The Haloferax
mediterranei GDH has been also purified and is an NADPspecific enzyme composed of six identical monomers of
55 kDa (Ferrer et al., 1996). In addition, a putative gdhA
gene is present in the Halobacterium sp. genome (Ng et al.,
2000; Table 2). It is also remarkable that the amino acid
sequence of the grapevine (Vitis vinifera) GDH is more
similar to the archaeal GDH than to those of eukaryotes. A
stress-related function of plant GDH has been invoked to
explain this resemblance and the thermostability of the
enzyme (Syntichaki et al., 1996).
On the other hand, ammonium is usually the preferred
nitrogen source in bacteria because it is easily assimilated
with low energy cost. As other inorganic nitrogen sources,
like nitrate, nitrite or N2, must be reduced before assimilation, inorganic nitrogen assimilation is often repressed by
ammonium.
Regulation of nitrogen metabolism in archaea
Denitrification and other anaerobic respiratory processes
are alternative metabolic pathways developed by facultative micro-organisms to obtain energy for growth under
anoxic conditions, and therefore they are usually repressed
by oxygen (Moreno-Vivián & Ferguson, 1998). Regulation
of anaerobic metabolism in E. coli and control of denitrification genes in Pseudomonas strains and other denitrifying
bacteria are well studied (Zumft, 1997), but the mechanisms controlling nitrate respiration and denitrification
in archaea are still not established. In addition, database
searches reveal that homologues of different potential
http://mic.sgmjournals.org
Glutamine synthetase (GS) is highly regulated at both
transcriptional and post-translational levels. In bacteria,
GS is reversibly modified by adenylylation in response
to the cellular nitrogen status, basically the glutamine/
2-oxoglutarate ratio. An adenylyltransferase catalyses the
adenylylation and deadenylylation of GS in response to
the PII protein, which in turn can be uridylylated or
deuridylylated by a uridylyltransferase. When glutamine
levels are high, deuridylylated PII stimulates GS inactivation by adenylylation, whereas PII-UMP promotes GS
deadenylylation for activation when 2-oxoglutarate levels
are high. PII is also involved in transcriptional regulation
of several nitrogen metabolism genes in response to the
nitrogen status, acting on the general Ntr regulatory system
or affecting specific regulators, such as the NifA protein
that activates nif gene expression. PII is encoded by the glnB
gene, and glnB homologues are known in a variety of
bacteria and archaea, and have been recently identified in
higher plants, indicating that the GlnB family plays a central
role in nitrogen regulatory processes in both prokaryotes
and eukaryotes (Ninfa & Atkinson, 2000; Arcondéguy et al.,
2001). However, PII proteins are not ubiquitous and several
bacterial and archaeal genomes do not encode glnB genes.
This is the case for Aeropyrum pernix, Pyrococcus horikoshii
and Pyrococcus abyssi (Arcondéguy et al., 2001). In addition,
as mentioned above, archaeal GSs are not regulated by
adenylylation, and control of GS activity in response to PII
proteins could be via a different mechanism than that in
most bacteria. Many organisms contain a second PII gene
called glnK, which is located upstream from the amtB gene
encoding an ammonium transporter in most prokaryotes,
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3541
P. Cabello, M. D. Roldán and C. Moreno-Vivián
including archaea, where this gene pair is found in multiple
copies. This conserved gene organization indicates that the
two genes were associated before the divergence of bacteria
and archaea, and suggests that the GlnK regulatory protein
and the ammonium transporter interact directly, probably
for regulation of ammonium transport by GlnK (Thomas
et al., 2000). It has recently been demonstrated that GlnK
binds to AmtB in the membrane and acts as a negative
regulator of the transport activity of AmtB. This binding is
dependent on GlnK uridylylation and is modulated according to the cellular nitrogen status, being maximal at high
nitrogen (Coutts et al., 2002; Javelle et al., 2004). Therefore,
this regulatory mechanism could be conserved throughout
bacteria and archaea. Some archaeal genomes contain other
glnB-like genes in addition to glnB and glnK. Thus, two
different GlnB homologues are encoded in the nif gene
clusters of diazotrophic methanogens (Chien & Zinder,
1996; Kessler & Leigh, 1999; Kessler et al., 2001). In
Methanococcus maripaludis, these glnB-like genes (termed
nifI1 and nifI2) are both required for ammonia switch-off
of nitrogenase, probably acting through a novel reversible
mechanism which does not require ADP-ribosylation of
NifH protein and does not affect transcription of nif genes
or stability of the NifH protein (Kessler et al., 2001).
Transcription in archaea resembles the eukaryotic process
more than the bacterial, and could be considered as a
simplified version of the eukaryotic transcription complex
(Bell & Jackson, 2001). In addition, although bacteria
and archaea contain homologous regulatory proteins, the
possibility that some regulators could be present only in
archaea should not be excluded. In Methanosarcina barkeri,
a putative TATA box is located in the nifHDK2 promoter region that is transcribed like typical archaeal genes.
Expression of nitrogenase structural genes is repressed by
ammonium (Lobo & Zinder, 1990; Chien & Zinder, 1996)
and ammonium-grown cells contain a putative protein
that inhibits binding of transcription initiation proteins
at the TATA box promoter region (Chien et al., 1998). In
Methanococcus maripaludis, expression of nif (nitrogenase)
and glnA (glutamine synthetase) operons is repressed by
ammonium (Kessler et al., 1998; Cohen-Kupiec et al., 1999)
and a palindromic consensus sequence has been identified
as a nitrogen operator for binding of a putative repressor
which negatively regulates the nif and glnA operons
in response to ammonia (Kessler & Leigh, 1999). Very
recently, this nitrogen repressor has been identified in
M. maripaludis and called NrpR (Lie & Leigh, 2003).
NrpR is a tetramer of 60 kDa subunits with a putative
N-terminal winged helix–turn–helix motif, which represents a new type of regulator exclusive to euryarchaeota
(Lie & Leigh, 2003). C/N balance could regulate binding
of the NrpR repressor to the nif promoter, because 2oxoglutarate decreases the binding of NrpR to the nitrogen
operator sequence. Therefore, archaea could use a different
mechanism than bacteria for the control of nitrogen gene
expression in response to the C/N balance (Lie & Leigh,
2003).
3542
Concluding remarks
Archaea can sustain the N-cycle in the extreme environments where they live because they are able to drive both
dissimilatory and assimilatory processes of the cycle. In
spite of this, archaeal inorganic nitrogen metabolism is
much less known than bacterial nitrogen metabolism.
Emerging data from recent functional studies and genome
sequencing projects reveal important differences in the
structure and regulation of enzymes and transport systems
involved in the denitrification and nitrogen assimilation
pathways, with interesting evolutionary implications. In
particular, different regulatory mechanisms may operate
in the control of nitrogen assimilation in archaea. Of
special interest is whether or not transcriptional regulation
in archaea is unique or possesses similarities to bacterial or
eukaryotic regulation. Major goals of future studies will
be to characterize at the molecular level the regulatory
mechanisms acting on nitrogen metabolism in archaea,
especially those for nitrate assimilation and denitrification.
Functional genomic approaches should be also performed
in order to identify the proteins and enzymes involved in
the N-cycle pathways in the different archaea and their
biochemical properties.
Acknowledgements
The authors are grateful for the financial support of Ministerio de
Ciencia y Tecnologı́a (grant BMC2002-04126-CO3-03) and Junta
de Andalucı́a (CVI 0117), Spain, and Alexander von Humboldt
Foundation, Germany.
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