The Biodiversity of Hydrogenases in Frankia
Characterization, regulation and phylogeny
Melakeselam Leul Zerihun
DOCTORAL DISSERTATION
To be defended on Friday 7th December 2007, 10:00 AM at
the Lecture Hall KB3A9, KBC, Umeå University
Faculty opponent
Kornel Kovacs, Professor, University of Szeged, Hungary
Department of Plant Physiology
Umeå Plant Science Center
Umeå University, Sweden
©Melakeselam Leul Zerihun, 2007
Department of Plant Physiology
Umeå Plant Science Center
Umeå University
SE-901 87 Umeå
Sweden
Doctoral Dissertation, Umeå 2007
ISBN 978-91-7264-444-1
Printed by VMC, KBC, Umeå University, Umeå.
“Life is available to anyone no matter what age. All you have to do is grab it”- Art Carney
Dedicated to
my beloved wife Genet F. Shawl
my beloved daughter Abigail
my beloved parents
The Biodiversity of Hydrogenases in Frankia: Characterization, regulation and
phylogeny
Melakeselam Leul Zerihun (2007) ISBN 978-91-7264-444-1
Department of Plant Physiology, Umeå Plant Science Center, Umeå University, Sweden
Dissertation abstract
All the eighteen Frankia strains isolated from ten different actinorhizal host plants
showed uptake hydrogenase activity. The activity of this enzyme is further increased by
addition of nickel. Nickel also enhanced the degree of hydrogenase transfer into the
membranes of Frankia, indicating the role of this metal in the processing of this enzyme.
The uptake hydrogenase of Frankia is most probably a Ni-Fe hydrogenase.
Genome characterization revealed the presence of two hydrogenase genes
(syntons) in Frankia, which are distinctively separated in all the three available Frankia
genomes. Both hydrogenase syntons are also commonly found in other Frankia strains.
The structural, regulatory and accessory genes of both hydrogenase synton #1 and #2 are
arranged closely together, but in a clearly contrasting organization. Hydrogenase synton
#1 and #2 of Frankia are phylogenetically divergent and that hydrogenase synton #1 is
probably ancestral among the actinobacteria. Hydrogenase synton #1 (or synton #2) of
Frankia sp. CcI3 and F. alni ACN14a are similar in gene arrangement, content and
orientation, while the syntons are both reduced and rearranged in Frankia sp. EANpec.
The hydrogenases of Frankia sp. CcI3 and F. alni ACN14a are phylogenetically grouped
together but never with the Frankia sp. EAN1pec, which is more closely related to the
non-Frankia bacteria than Frankia itself. The tree topology is indicative of a probable
gene transfer to or from Frankia that occurred before the emergence of Frankia. All of
the available evidence points to hydrogenase gene duplication having occurred long
before development of the three Frankia lineages. The uptake hydrogenase synton #1 of
Frankia is more expressed under free-living conditions whereas hydrogenases synton #2
is mainly involved in symbiotic interactions. The uptake hydrogenase of Frankia can also
be manipulated to play a larger role in increasing the efficiency of nitrogen fixation in the
root nodules of the host plants, there by minimizing the need for environmentally
unfriendly and costly fertilizers.
The hydrogen-evolving hydrogenase activity was recorded in only four Frankia
strains: F. alni UGL011101, UGL140102, Frankia sp. CcI3 and R43. After addition of
15mM Nicl2, activity was also detected in F. alni UGL011103, Frankia sp. UGL020602,
UGL020603 and 013105. Nickel also increased the activity of hydrogen-evolving
hydrogenases in Frankia, indicating that Frankia may have different types of hydrogen-
evolving hydrogenases, or that the hydrogen-evolving hydrogenases may at least be
regulated differently in different Frankia strains. The fact that Frankia can produce
hydrogen is reported only recently. The knowledge of the molecular biology of Frankia
hydrogenase is, therefore, of a paramount importance to optimize the system in favor of
hydrogen production. Frankia is an attractive candidate in search for an organism
efficient in biological hydrogen production since it can produce a considerable amount of
hydrogen.
Key words: Biodiversity, Frankia, immunoblotting, gene expression, uptake
hydrogenase, hydrogen-evolving hydrogenase, nickel, phylogeny
CONTENTS
PAGE
LIST OF PAPERS 9
ABBREVIATIONS 10
PREFACE 11
INTRODUCTION 12
Hydrogen 12
Biodiversity of hydrogenases 13
Physiological Regulation of Hydrogenases 16
Biotechnology of hydrogenases 17
Hydrogen metabolism in nitrogen-fixing organisms 18
Nitrogenases 18
Uptake hydrogenases 18
Bidirectional/reversible hydrogenases 20
Frankia and their host plants 21
SUMMARY OF MATERIALS AND METHODS 24
Frankia strains and growth conditions 24
Seeds and inoculation of host plants 25
Enzyme activity assays 25
Protein extraction, determination and electrophoretic analysis 26
Immunoblotting and immunolabeling 27
Phylogenetic analysis of Frankia hydrogenases 28
Transcriptional analysis of Frankia hydrogenases 29
THE AIM OF THIS THESIS 30
7
RESULTS AND DISCUSSION 30
UPTAKE HYDROGENASES IN FRANKIA 30
The molecular characterization of uptake hydrogenases in Frankia 31
The structure of uptake hydrogenases in Frankia 33
The phylogeny of uptake hydrogenases in Frankia 35
The regulation of uptake hydrogenases in Frankia 37
Hydrogenase gene expression in free-living vs. symbiotic condition 37
Ni-dependent regulation of uptake hydrogenases in Frankia 38
Effects of nitrogenase and hydrogen on the uptake hydrogenase of Frankia 39
HYDROGEN-EVOLVING ENZYMES IN FRANKIA 39
Molecular characterization of hydrogen-evolving enzymes in Frankia 40
Gel and peptide analysis 40
Localization of the hydrogen evolving enzyme in Frankia 41
Regulation of the enzymes in Frankia 41
Nitrogenase and the hydrogen-evolving hydrogenases of Frankia 41
Ni-dependent regulation of the hydrogen-evolving hydrogenases of Frankia 42
DOES FRANKIA HAVE OTHER HYDROGENASES? 43
CONCLUSIONS 43
FUTURE PERSPECTIVES 45
Frankia – bakterien som pruducerar både kväve gödsel och vätgas! 46
ACKNOWLEDGEMENTS 47
REFERENCE LIST 49
8
List of papers The thesis is based on the publications listed below, which will be referred to in the text
by their corresponding Roman numerals.
I. Leul M, Mohapatra A and Sellstedt A (2005) Biodiversity of hydrogenases in
Frankia. Curr Microbiol 50(1): 17-23.
II. Leul M, Mattsson U and Sellstedt A (2005) Molecular characterization of uptake
hydrogenase in Frankia. Biochem Soc Trans 33: 64–66.
III. Leul M, Normand P and Sellstedt A (2007) The organization, regulation and
phylogeny of uptake hydrogenase genes in Frankia. Physiologia Plantarum
130(3): 464-70.
IV. Mohapatra A, Leul M, Mattsson U and Sellstedt A (2004) A hydrogen-evolving
enzyme is present in Frankia sp. R43. FEMS Microbiol Lett 236(2): 235-40.
V. Mohapatra A, Leul M, Sandström G and Sellstedt A (2006) Occurrence and
characterization of the hydrogen-evolving enzyme in Frankia sp. Int J Hydrogen
Energy 31: 1445-51.
VI. Leul M and Sellstedt A (2007) The phylogeny of uptake hydrogenases in
Frankia. Manuscript.
Papers I-V are reproduced with the kind permission of the publishers.
9
Abbreviations
Ct control LGT lateral gene transfer
PCR polymerase chain reaction
REST relative expression software tool
RT-PCR reverse transcription–polymerase chain reaction
REST relative expression software tool
ARA acetylene reduction activity
GC gas chromatograph
MALDI-TOF matrix-assisted laser-desorption-ionization
-time-of-flight mass spectrometry
NAD nicotinamide adenine dinucleotide
NADH nicotinamide adenine dinucleotide hydride
GTPase guanosine triphosphatase.
hup hydrogen uptake
shc squalene hopane cyclase
nif genes encoding nitrogenase
10
Preface
The increased awareness of global environmental crises and the depletion of fossil fuels
have prompted researchers to seek alternative, renewable energy sources. One of the
obvious options is hydrogen, which could potentially be used as an extremely clean
energy source, producing only water on burning. Hydrogen can be produced biologically
by microorganisms, thanks to the special group of their enzymes called hydrogenases.
Hydrogenases also increase the efficiency of nitrogen the fixation process and have other
other biotechnological applications such as wastewater treatment etc.
Hydrogenases have been characterized in detail in some organisms. In Frankia,
the research work progressed specially over the last decade as a new ways of growing
Frankia was being adopted. The recent availability of the three Frankia genomes did
definitely contributed to this study. The knowledge of the molecular biology of Frankia
hydrogenase is of a paramount importance to optimize the system in favor of hydrogen
production or nitrogen fixation. In this thesis the characterization, regulation and
phylogeny of Frankia hydrogenases have been studied, which I hope, will increase
knowledge of hydrogenases.
Melakeselam Leul Zerihun
Department of Plant Physiology
December 2007, Umeå
11
Introduction
There are more than 25 published definitions of biodiversity, which is short for
biological diversity. The simplest is "variation of life at all levels of biological
organization". Since the variety of life can be expressed in various ways, there is no
overall measure of biodiversity; rather there are multiple measures of different facets of
it (Gaston and Spicer, 2004). Biodiversity has traditionally been identified at three
levels: (i) genetic diversity (diversity of genes within a species); (ii) species diversity
(diversity among species in an ecosystem); and (iii) ecosystem diversity (diversity at a
higher level of organization, the ecosystem). Thus, biodiversity for geneticists is the
diversity of genes and organisms, but for ecologists it applies to the diversity of species
in the context of their immediate environments and ecosystems.
In the project this thesis is based upon the diversity of hydrogenases within the
genus Frankia was examined, rather than the diversity of the organisms per se. In
addition, Frankia hydrogenases were compared with those of other organisms, and the
regulation of Frankia hydrogenases under various environmental conditions was
investigated. Various Frankia strains were used that were originally isolated by
screening a wide range of actinorhizal host plants from diverse parts of the world. Hydrogen
Hydrogen, the simplest naturally occurring atom, is the most abundant of all the
elements, accounting for three-fourths of the mass of the universe. The abundance of
gaseous hydrogen at the earth’s surface is generally low, because it is less dense than air.
However, hydrogen is a major component of myriads of compounds, and it is found in
all organisms (biomass) in both a huge range of molecules and in the ionic form, H+ (or,
more precisely, protonated water complexes). Hydrogen also accounts for ca. 70% of the
sun’s current mass (helium accounting for a further ca. 28% and all other constituents for
<2%). The fusion of the nuclei of hydrogen atoms into helium atoms releases radiant
energy that sustains life on earth, and some of this energy is eventually stored as
12
chemical energy in fossil fuels. Most of the energy we use today originates from the
sun's radiant energy. However, hydrogen is a promising energy carrier that has potential
use as an extremely clean energy source, producing only water on burning.
Hydrogen is involved in fundamental aspects of microbial physiology and it plays
a central role in life forms that inhabit anaerobic environments. It has been estimated that
about 200 million tones of hydrogen are produced and consumed per year in anoxic
habitats (Thauer et al., 1996). In spite of the high turnover rates, the steady-state
concentration of H2 in most anoxic habitats is very low (1-10 Pa), indicating that H2
formation rather than H2 consumption is the rate-limiting step in the overall process.
Hydrogen-consuming anaerobes obtain energy by using the electrons from hydrogen to
produce methane (methanogens) or acetate from carbon dioxide (acetogens), sulfide
from sulfate (sulfate reducers), ferrous from ferric iron (iron reducers), or nitrogen and
nitrite from nitrate (denitrifying bacteria) depending on the environment (Adams and
Stiefel, 1998). These and other organisms can metabolize hydrogen since they produce a
special group of enzymes called hydrogenases.
Biodiversity of hydrogenases
Hydrogenases are microbial enzymes that catalyze the reversible oxidation of molecular
hydrogen. Hydrogenase activity has been reported in a large number of anaerobic and
aerobic prokaryotes, as well as some eukaryotes, including various algae, green plants
(such as barley), and protozoa (Adams et al., 1981; Lindmark and Muller, 1973; Torres
et al., 1986).
Hydrogenases differ in the type of electron carriers they use, and associated
differences in their structures and, more importantly, redox potentials (Mertens and
Liese, 2004). Some hydrogenases reduce electron acceptors like quinones, while others
that are hydrogen producers have electron donors such as ferredoxins and cytochromes
(Cammack, 2001). Analysis of the physiological diversity of hydrogenases has revealed
three phylogenetically distinct classes: the [Fe] hydrogenases, the [NiFe] hydrogenases
and the metal-free hydrogenases (Vignais et al., 2001). The [NiFe] hydrogenases include
a subgroup of hydrogenases containing selenium called [NiFeSe]-hydrogenases (Vignais
13
et al., 2001) that are probably ancient, and are only found in Archaea (Robson, 2001).
Each group is characterized by a distinctive functional core that is conserved within each
class. The widely spread and most thoroughly studied [NiFe] hydrogenases are less
active than their Fe-only counterparts, and their physiological role is usually the
oxidation of H2. The [Fe] hydrogenases are found in few microorganisms and are
difficult to study due to their sensitivity to oxygen. Their usual function is H2 evolution, and they have higher specific activities than the [NiFe] hydrogenases (Adams, 1990). It
has been noted that the term “[FeS] cluster-free hydrogenases” is more appropriate for
the third group of hydrogenases “metal-free hydrogenases”, since this group contains
functional iron, although it is not catalytically active (Lyon et al., 2004). The proteins of
the so-called metal-free hydrogenases are encoded by hmd genes and they may play
important roles in methanogenesis in nickel-deficient conditions, since their specific
activities increase in cells growing in nickel-limiting conditions (Afting et al., 1998).
Hydrogenases vary substantially in terms of subunit composition, metal content,
structure and size between different organisms (Adams, 1990). For example, the [Fe]
hydrogenase of the anaerobic bacterium Megasphaera elsdenii has only a single
polypeptide chain of 58 kDa while that of Desulfovibrio desulfuricans ATCC 7757 has
two different subunits of 42.5 kDa and 11 kDa (Filipiak et al. 1989). Hydrogenases may
also vary in the size of their structural subunits. The hydrogenase of Thermotoga
maritima contains iron as the only metal and consists of three subunits, with masses of
73 (α), 68 (β) and 19 (γ) kDa (Verhagen et al., 1999).
Further functional analyses of hydrogenases have identified 13 families to date
(Table 1), all of which, but one, are directly or indirectly involved in energy metabolism
(Robson, 2001). The main physiological functions of hydrogenases are the oxidation of
H2 or reduction of protons. The oxidation of hydrogen is linked to energy conservation,
via coupling to energy-conserving electron transfer chain reactions, allowing energy to
be obtained either from H2 or the oxidation of substrates of lower potential. The
evolution of hydrogen (H+ reduction) is linked to the disposal of excess reducing
potential. Other hydrogenases, e.g. bidirectional NAD(P)-reactive hydrogenases of
cyanobacteria, may interact with respiratory electron transport chains and provide
electron "valves" that control the redox poise of the respiratory chain at the level of the
14
Hydrogenase family Occurrence Function . Fe-only hydrogenases Obligately anaerobic Fermentation/ bacteria and Eubacteria Energy conservation? NAD(P)-reactive Obligately anaerobic, Fermentation hydrogenases Archaea NiFe-hydrogenases associated Facultative and obligate Fermentation with the formate hydrogen anaerobes, Archaea lyases complex NiFe(Se) membrane-bound Aerobes, facultative anaerobes, Energy conservation periplasmic hydrogenases Proteobacteria NAD(P)-reactive Facultative and obligately Energy conservation hydrogenases anaerobic Eubacteria F420-non-reactive Methanogens Energy conservation hydrogenases F420-reactive Methanogens Energy conservation hydrogenases Non metal hydrogenases Methanogens Energy conservation NiFe (thylakoid) uptake Cyanobacteria Energy conservation? hydrogenases Bidirectional NAD (P)-reactive Cyanobacteria Energy conservation, hydrogenases Eedox poising? NiFe-sensor hydrogenases Chemolithotrophic/ Hydrogen sensing Phototrophic proteobacteria components in genetic regulation of hydrogenase expression Ech hydrogenases Methanogenesis pathway Methanogenesis pathway .
Table 1. Families of hydrogenases: their occurrences and functions (Redrawn from Robson,
2001).
quinone pool and ensure the correct functioning of the respiratory chain in the presence of
excess reducing equivalents (Vignais and Colbeau, 2004; Robson, 2001). Soluble [NiFe]
hydrogenases, HupUVs, which have been identified in organisms like Rhodobacter
capsulatus (Elsen et al., 1996) and B. japonicum (Black et al., 1994) may participate in the
regulation of gene expression by acting as hydrogen sensors. The facultative
chemolithoautotroph R. eutropha also harbors a regulatory hydrogenase, HoxBC, which
enables the organism to sense hydrogen in its environment (Kleihues et al., 2000).
15
Physiological Regulation of Hydrogenases
Organisms may have one or more types of hydrogenases. For instance, five sets of
structural genes that code for active hydrogenases have been identified in Thiocapsa
roseopersicina (Kovács and Rákhely, 2007). The presence of multiple isofunctional
hydrogenases in some microorganisms indicates the importance of hydrogen in their
metabolism and their ability to modify their metabolism in adaptive responses to
different environments. Different hydrogenase isoenzymes may be localized in different
cell compartments: the cytosol, cell membrane or periplasm. Hydrogenases are also
known to be differentially expressed under different environmental conditions and to
have differing functions (Robson, 2001).
The ability of microbes to either take up or evolve H2 is usually a facultative trait.
A variety of factors, including the concentration of hydrogen, oxygen, nickel ions,
molybdenum, nitrate, formate, carbon monoxide, nitrogen/phosphate, carbon and energy
sources can affect hydrogenase gene expression (Friedrich et al., 2001; Vignais and
Colbeau, 2004). Molecular hydrogen, which is also the substrate, activates hydrogenase
expression in aerobic bacteria, photosynthetic bacteria and free-living Rhizobia, whereas
molecular oxygen is inhibitory for most hydrogenases. Hydrogenase synthesis in
facultatively H2-oxidizing bacteria like Azotobacter vinelandii (Kennedy and
Toukdarian, 1987) and Bradyrhizobium japonicum (Hanus et al., 1979) depends on the
availability of H2. In facultative and (especially) obligately anaerobic bacteria, the
availability of O2 and the redox state of the cells are important regulatory variables for
hydrogenase gene expression (Kovács et al., 2005). Several carbon sources, such as
pyruvate and propionate, have also been shown to affect the regulation of hydrogenase
activity in three strains isolated from Casuarina sp. (Sellstedt et al., 1994). The
heterotrophic and strictly anaerobic archaeon Methanococcus voltae harbors four
hydrogenase operons, including two encoding [NiFe] hydrogenases that are expressed
under selenium depletion conditions when [NiFeSe] hydrogenases cannot be made in
sufficient amounts (Berghofer et al., 1994). Hydrogenase gene expression in A.
cylindrica sp. strain PCC7120 requires genome re-arrangements that occur during the
cellular differentiation process leading to heterocyst formation (Friedrich et al., 2001).
16
Biotechnology of hydrogenases
Knowledge about hydrogenases in microorganisms has greatly increased in the last
decade, and our enhanced understanding of the structure and function of the active sites
of hydrogenases has led to the synthesis of a close analogue of the hydrogen-producing
active centre of hydrogenases, the H-cluster (Tard et al., 2005). The availability of an
active, free-standing analogue of the H-cluster has enabled scientists to develop useful
electrocatalytic materials for applications in, inter alia, reversible hydrogen fuel cells.
The precious metal platinum (Pt) is the currently preferred electrocatalyst for such
applications, but it is very expensive (costing more than $17 per gram), its availability is
limited and its use is unsustainable in the long term. Thus, such alternatives could be
extremely valuable. Hydrogenases can also be used in various biotechnological
applications such as biohydrogen production, wastewater treatment, the prevention of
microbial-induced corrosion and the generation/regeneration of NADP cofactors
(Mertens and Liese, 2004).
In addition, increasing awareness of global environmental crises and the depletion
of fossil fuels has prompted researchers to seek alternative, renewable energy sources.
An obvious option is hydrogen, which could potentially be used as an extremely clean
energy source, producing only water on burning. Biological hydrogen production by
photosynthetic prokaryotic and eukaryotic organisms (e.g. cyanobacteria and the green
alga Chlamydomonas reinhardtii) or by fermentation in anaerobic bacteria (e.g.
Clostridium butyricum) has been reported (Melis et al., 2000; Karube et al., 1976).
Several strategies for boosting their hydrogen production are being explored, such as
genetic modification of the light-harvesting antennae complexes, sulfur deprivation of
the cultures, screening for mutants that produce more hydrogen than wild type strains,
optimization of conditions for the hydrogenase enzyme and investigation of naturally
occurring hydrogen production. Biological hydrogen production has several advantages
over conventional means of hydrogen production, such as photoelectrochemical or
thermochemical processes.
17
Hydrogen metabolism in nitrogen-fixing organisms
Three key classes of enzymes that are directly involved in hydrogen metabolism have
been identified in nitrogen-fixing organisms to date: nitrogenases, uptake hydrogenases
and bidirectional hydrogenases (Fig. 1). More than 10 million tons of H2 are globally
generated in oxic habitats (e.g. oxic soils and fresh water) by aerobic and
microaerophilic microorganisms as side-products of nitrogen fixation (Thauer et al.,
1996).
Nitrogenases: are oxygen-labile enzymes that catalyze the reduction of nitrogen to
ammonia in the highly energy-demanding process of nitrogen fixation, which requires
metabolic energy in the form of ATP as shown in the reaction:
N2 + 8H+ + 8e- + 16ATP → 2NH3 + H2 + 16ADP + 16Pi
Since two ATP molecules are required for each electron transferred from dinitrogenase
reductase to dinitrogenase, a total of 16 ATP molecules are needed to reduce dinitrogen
(N2) to ammonia (NH3), a form in which the nitrogen is available for further biological
reactions. During the nitrogen fixation process, substantial amounts of hydrogen are
produced via the reduction of protons, catalyzed by the nitrogenase enzyme. In fact, it
has been shown that in most symbionts only 40-60% of the electron flows to the
nitrogenase are transferred to nitrogen, and the remainder is lost through hydrogen
evolution (Schubert and Evans, 1976).
Uptake hydrogenases: Uptake hydrogenases catalyze the consumption of hydrogen (H2
oxidation) produced by nitrogenases during nitrogen fixation. Hydrogen oxidation is
coupled to the reduction of electron acceptors such as oxygen, nitrate, sulfate, carbon
dioxide, and fumarate. The hup (hydrogen uptake) systems have been studied in detail in
two species of root nodule rhizobia, Rhizobium leguminosarum bv. viciae and B.
japonicum, from which a multigenic (18–24 genes) cluster responsible for the synthesis
18
of an active hydrogenase has been isolated (Ruiz- Argüeso et al., 2000). Uptake
hydrogenase is considered beneficial to the nitrogen-fixing organisms in both the free-
living and symbiotic states (Dixon, 1976) since the hydrogen produced during the
nitrogen fixation can be consumed and the reductant generated can be used by the cells
in various ways. Hydrogen recycling has been shown to reduce energy losses associated
with nitrogen fixation (Schubert and Evans, 1976). In addition to the provision of an
additional source of energy, other possible functions such as prevention of H2 inhibition
of the nitrogenase reaction and protection of oxygen-sensitive nitrogenase from O2
damage have been proposed (Dixon, 1972). All strains of cyanobacteria (Tamagnini et
al., 2002) and Frankia (Sellstedt, 1989) investigated to date have uptake hydrogenases,
but only a few examined strains of rhizobia have this enzyme.
Fig. 1. Enzymes directly involved in hydrogen metabolism in cyanobacteria (Redrawn
from Tamagnini et al., 2002). The bidirectional hydrogenase of cyanobacteria has five
subunits: HoxEFUYH (Schmitz et al., 2002).
Uptake hydrogenase and nitrogenase encoding genes of Rhi. leguminosarum,
which are induced together, are controlled by the nitrogen fixation regulatory protein
19
NifA (Brito et al., 1997). A strong correlation between activities of nitrogenase and
uptake hydrogenase has been reported in Frankia, although they might not be
coregulated (Mattsson and Sellstedt, 2000). Also, uptake hydrogenase was localized in
vesicles and hyphae (Sellstedt and Lindblad, 1990).
Bidirectional/reversible hydrogenases: this group of hydrogenases has the capacity to
metabolize hydrogen both directions. They have been characterized by their sensitivity
to oxygen, thermotolerance, and high affinity to hydrogen, and are widely distributed
among cyanobacteria, including nitrogen fixing, non-nitrogen-fixing, unicellular, non-
heterocystous, and heterocystous strains (Houchins, 1984). The physiological functions
of the bidirectional hydrogenases are still unclear but it has been suggested that they may
mediate the release of excess reducing power in anaerobic environments (Tamagnini et
al., 2002), acting as electron valves during the light reactions of photosynthesis and thus
preventing retardation of the electron transport chain under stress conditions (Appel et
al., 2000), and/or be involved in fermentation (Troshina et al., 2002) and respiratory
complex I (Appel et al., 1996). Since the activity of the enzymes is not strongly affected
by combined hydrogen, it has also been suggested that these hydrogenases may function
independently of nitrogen fixation (Tamagnini et al., 2002). Furthermore, it has been
shown that the biosynthesis of nitrogenase is not essential for biosynthesis of the
bidirectional hydrogenase and hydrogen evolution in several unicellular strains (Howarth
and Codd, 1985).
The crystal structures of the [Ni-Fe] hydrogenases of five sulfate-reducing bacteria
(Desulfovibrio gigas, D. vulgaris, D. desulfuricans, D. fructosovorans and
Desulfomicrobium baculatum) have been reported so far (Volbeda et al., 1995; Higuchi
et al., 1997; Matias et al., 2001; Montet et al., 1997; Garcin et al., 1999). The crystal
structures of the [FeFe] hydrogenases of two organisms (Clostridium pasteurianum and
D. desulfuricans) have also been resolved (Peters et al., 1998; Nicolet et al. 1999). Both
NiFe and Fe-hydrogenases share a common active site low-spin Fe center with CO and
CN coordination, although these hydrogenases are evolutionarily unrelated. From the
studies undertaken to date, [Fe] hydrogenases appear to have simpler structures than the
[NiFe] hydrogenases (Nicolet et al., 2002).
20
Frankia and their host plants
Frankia is a genus of nitrogen-fixing filamentous, heterotrophic, Gram-positive,
actinomycetous soil bacteria. Frankia resemble fungi and are phylogenetically and
morphologically distinct from the rhizobial bacteria that are responsible for nitrogen
fixation in legumes (Binkley et al., 1994). Frankia can fix nitrogen in both free-living
aerobic conditions and in symbiosis, unlike other soil microsymbionts, such as some
species of rhizobia (Zhang et al., 1984). Frankia can differentiate into three cell types:
hyphae, vesicles and spores. The vesicles are formed under nitrogen-limiting conditions
from the swollen tips of hyphae in most free-living Frankia. Vesicles, in which the
oxygen-sensitive nitrogenase is localized (Meesters, 1987; Huss-Danell and Bergman,
1990), are surrounded by a multi-layer lipid membrane that maintains a low internal
oxygen tension (Parsons et al., 1987). In a liquid medium, an exponentially growing
culture forms spherical or ellipsoidal mycelial colonies, while overgrown cultures are
like huge, uniform mycelia (Schwencke, 2001).
The first attempt to classify members of the genus Frankia was by Baker (1987),
who proposed that there were four “infectivity groups”, based on the results of
infectivity studies using pure cultures in cross-inoculation tests, although it is
questionable whether such tests reflect host specificity under normal conditions. A more
sophisticated approach based on phenotypic characteristics was subsequently adopted,
which differentiated two Frankia species; F. alni and F. elaeagni (Lalonde et al., 1988).
In the following years, phylogenetic studies were performed, based on analyses of the
widely used 16S rDNA and 16S rRNA sequences (Nazaret et al., 1991; Normand et al.,
1996; Clawson et al., 2004), arbitrary primers (Sellstedt et al., 1992), nitrogen fixation
genes (Jeong et al., 1999) and glutamine synthetase (Clawson et al., 2004). The studies
conducted to date indicate that Frankia strains can be generally divided into three
clusters. Cluster 1 includes strains that nodulate plants of the Fagales, Betulaceae and
Myricaceae and are often referred to as “Alnus strains” (Normand et al., 1996) and a
subclade of “Casuarina strains”, which only nodulate Casuarina and Allocasuarina
species of the Casuarinaceae under natural conditions (Benson et al., 2004). Cluster 2 is
comprised of unisolated strains of Frankia that only infect members of the Coriariaceae,
21
Datiscaceae, Rosaceae and Ceanothus of the Rhamnaceae. Cluster 3 strains form
effective nodules on members of the Myricaceae, Rhamnaceae, Elaeagnaceae and
Gymnostoma of the Casuarinaceae. In the studies underlying this thesis the phylogenetic
relationships and characteristics of Frankia hydrogenases (and their relationships with
hydrogenases of other organisms), rather than those of Frankia organisms, were
investigated. In all cases in the following text the term “hydrogenase synton” will be
used instead of “hydrogenase cluster” to avoid possible confusion with the term
“Frankia cluster” described in this paragraph.
Frankia can interact and form symbiotic relationships with a diverse, globally
distributed group of dicotyledonous plants called actinorhizal plants that are classified
into four subclasses, eight families, and 25 genera of plants comprising more than 240
species of dicotyledonous angiosperms (Wall, 2000). Actinorhizal plants are widespread
(Fig. 2) and grow in all types of climate, although they are mainly found in temperate
climates (Silvester, 1977). They inhabit diverse ecosystems, including arctic tundra
(Dryas species), coastal dunes (Casuarina, Hippophae, Myrica, and Elaeagnus species),
riparian (Alnus and Myrica species), glacial till (Alnus and Dryas species), forest (Alnus,
Casuarina, Coriaria, and Shepherdia species), chapparal and xeric (Casuarina, Purshia,
Ceanothus, Cercocarpus, Comptonia, and Cowania species), and alpine (Alnus species)
systems (Benson and Silvester, 1993). Actinorhizal plants often serve as pioneer species
in early successional plant communities since they thrive on marginal soils. They
contribute considerable amounts of fixed nitrogen, especially in cool, temperate areas
where indigenous legumes are rare or absent (Silvester, 1976). They are economically
important in forestry programs, such as land reclamation and reforestation programs, and
have high potential for introduction in areas with problem (arid, saline or waterlogged)
soils, for timber, pulp and fuel wood production, and for acting as windbreaks and/or
ornamental plants (Chaudhary and Mirza, 1987; Diem and Dommergues, 1990).
22
Fig. 2. Present-day native distribution of actinorhizal plant hosts: (I) Betulaceae (B) and
Myricaceae (M) and their overlap (M+B); (II) Elaeagnaceae (E), Myricaceae (M),
Rhamnaceae (R). Elaeagnaceae and Myriceae (E+M) overlap in some areas. The
Casuarinaceae (not shown in the figure) are distributed in Indo-Malaysia, Australia, and
the Pacific islands (Redrawn from Normand et al., 2007a)
23
Summary of materials and methods
Frankia strains and growth conditions
Twenty Frankia strains isolated from 12 different actinorhizal host plants native to
different parts of the world (Table 2) were grown at 27ºC, as described in Mattsson and
Sellstedt (2000). Cells were successively transferred to fresh PUM medium containing
0.1 g/L NH4Cl on a weekly basis to obtain actively growing cultures. In the experiments
Frankia strains Source or references Location Host plant . F. alni ACN14a Normand and Lalonde, 1982 Canada A. viridis subsp. crispa
F. alni AvCI1 Baker and Torry, 1980 USA A. viridis subsp. crispa
Frankia sp. KB5 Sellstedt et al., 1991 Australia C. equisetifolia
Frankia sp. UGL020603 Vel´azquez et al., 1998 Egypt C. equisetifolia
Frankia sp. UGL020602 Wheeler C Brazil C. equisetifolia
F. alni UGL011103 Wheeler C Sweden A. incana
F. alni UGL011102 Wheeler C Sweden A. incana
F. alni ArI3 Berry and Torrey, 1979 USA A. rubra
Frankia sp. 013105 Wheeler C USA A. rubra
F. alni 010701 Wheeler C Scotland A. glutinosa
F. alni 010702 Wheeler C Scotland A. glutinosa
F. alni UGL011301 Sayed et al., 1997 S. Korea A. inokumai
Frankia sp. UGL161101 Wheeler C Scotland M. gale
Frankia sp. UGL161102 Wheeler C Scotland M. gale
Frankia sp. HFPCcI3 Zhang et al., 1984 USA C. cunninghamiana
Frankia sp. R43 Zhang et al., 1984 USA C. cunninghamiana
Frankia sp. EAN1pec Lalonde et al., 1981 USA E. angustifolia
Frankia sp. BCU110501 Chaia, 1998 Argentina D. Trinervis
Frankia sp. UGL140104 Wheeler C Scotland H. rhamnoides
Frankia sp. UGL140102 Lumini et al., 1996 Scotland H. rhamnoides
Frankia sp. CpI1 Callaham et al., 1978 USA C. peregrina .
Table 2. Frankia sp. strains used in the study.
24
cells were placed in portions of 50 mL growth medium in 100 mL flasks, at 5 μg/mL
total protein concentration, without nitrogen to induce vesicle formation and nitrogen
fixation, to which nickel (II) chloride was added at various concentrations, where
appropriate, to assess the effects of nickel on hydrogenase expression.
Seeds and inoculation of host plants
Alnus glutinosa seeds were sterilized, imbibed in water overnight and germinated before
being transferred to a plastic pot containing a sterile soil and vermiculite mix (5:1 ratio),
which was supplemented with Evans solution (Evans et al., 1972) twice a week. When
the seedlings were six weeks old they were inoculated with 5 ml (from a 5 mg protein
per ml bacterial culture) of F. alni ACN14a. The plants were grown in a growth chamber
with metal halide lamps (HQI-T, 400 W, daylight) providing light with an irradiance of
300 μmol m-2s-2 for 17 h day-1. The day/night temperature was kept at 20ºC/17ºC and the
relative humidity at 70%.
Enzyme activity assays
Nitrogenase assays: nitrogenase activity in cultures of free-living Frankia cells was
determined as acetylene reduction activity (ARA) using a gas chromatograph (GC-8AIF,
Shimadzu Scientific Instruments Inc., Columbia, MD), as described earlier (Mattsson
and Sellstedt, 2000).
Uptake hydrogenase: Eight-day-old Frankia cultures were collected in 6.5-mL flasks
containing 1.8 mL 50 mM Tris-HCl and sealed with a gas-tight rubber membrane.
Uptake hydrogen activity was analyzed immediately after the addition of hydrogen gas
(1% v/v) into the gas phase of the flasks and then at 1 h intervals while the strains were
incubated at room temperature with shaking, using a gas chromatograph (GC8AIT,
Shimadzu Scientific Instruments, Colombia, MD) according to Mattsson and Sellstedt
(2000).
25
Hydrogen-evolving hydrogenase: Frankia cultures were incubated for 24 hours under
anaerobic conditions, at 27ºC with shaking in 6.5 mL glass vials sealed with gas-tight
rubber membranes. At the start of induction of hydrogen evolution by argonization,
ammonium chloride was added to the cultures to a final concentration of 10 mM to block
hydrogen evolution from nitrogenase. Hydrogen evolution was then measured in 2 mL
reaction mixtures, each containing 1.8 mL of Frankia culture resuspended in 50 mM
Tris-HCl (pH 7.0), to which 2 mM of freshly prepared methyl viologen and 20 mM
sodium dithionite were added (Tamagnini et al., 1997). Measurements began after an
incubation period of 90 min and continued until a linear increase in hydrogen was
recorded, using gas chromatography as outlined above.
NAD-reducing hydrogenase: Hydrogen evolution from NAD-reducing hydrogenase in
Frankia sp. R43 was measured by adding NAD to cultures of the organism, and
measuring NADH formation at 340 nm as previously described (Friedrich et al., 1980).
Protein extraction, determination and electrophoretic analysis
Total protein determination: To measure their protein concentrations, Frankia cells
were collected by centrifugation, treated as described by Mattsson and Sellstedt (2000),
and the protein contents of the resulting suspensions were determined using the
bicinchonic acid (BCA) assay and BSA as a standard.
One and two-dimensional gel electrophoresis: For one-dimensional analyses, Frankia
protein extracts containing 30 μg membrane (total) proteins were electrophoretically
separated on NuPage 12% Bis-Tris gels. For two-dimensional analyses Frankia proteins
were initially precipitated by acetone, centrifuged and the resulting pellets were air-
dried. Immobilized pH gradient gels (ZOOMTM Strips) were rehydrated at room
temperature, and a ZOOMTM IPGRunner System (Invitrogen) was then used for
isoelectric focusing by gradually increasing the voltage and maintaining the final
focusing voltage for approx. 2 h. The electro-focused IPG strip was incubated in a
reducing solution for 15 min, then in an alkylating solution for 15 min before
26
electrophoretic separation of the proteins in it on a NuPAGETM 4–12% Bis-Tris gel at
200 V for 50 min.
Immunoblotting and immunolabeling
Western blots analysis: The polypeptides on 1D or 2D gels were electrotransferred to
nylon transfer membrane and western blots were performed using a Western Breeze kit
(Invitrogen), according to the manufacturer’s instructions, except that the membrane was
incubated for 1-1.5 h with the primary antibody. The primary antibodies used in these
experiments were raised in rabbit against the large subunit of Ni-Fe hydrogenase (HoxG)
of the MB hydrogenase and HoxH of the SH-hydrogenase HY of R. eutropha, the small
hydrogenase subunit of B. japonicum (Hup S), and [Fe]-hydrogenase from D.
desulfuricans ATCC 7757, and were used at dilutions of 1:1000.
Southern blot analyses: Frankia DNA was digested with BamHI or Sal1, transferred to
a membrane and hybridized at 52ºC with a P32-labeled PCR fragment from part of the
small hydrogenase subunit originating from Frankia local source.
Preparation and immunolabeling of cryosections: Fixation, embedding, sectioning and
immunogold labeling were all performed as described earlier (Wheeler et al., 1998;
Mattsson et al., 2001), except that in the studies underlying this thesis we used primary
antiserum raised against HoxH and HoxG of R. eutropha, followed by secondary goat-
anti-rabbit IgG conjugated with 5 nm colloidal gold particles, before viewing the
samples under a Philips CM 10 transmission electron microscope operating at 60 kV.
Cells labeled with only the secondary antibody were used as controls.
Peptide analysis: Protein spots were excised from two-dimensional gels and cleaved in-
gel by trypsin. Peptide analysis was performed by electrospray ionization mass
spectrometry (Wilm et al., 1996) using a quadrupole-time-of-flight instrument and
Masslynx software or matrix-assisted laser-desorption ionization–time-of-flight mass
27
spectrometry, in which the resulting peptide ‘fingerprints’ were analyzed using Protein
Prospector software.
Phylogenetic analysis of Frankia hydrogenase
DNA extraction: Genomic DNA was extracted from seven-day-old Frankia cultures
using the bacterial protocol supplied with the Blood and Tissue Genomic DNA
Extraction Kit (Viogene, USA).
RNA isolation and cDNA synthesis: RNA was extracted using a RNeasy Mini Kit
according to the manufacturer’s protocol. F. alni cells (4.5 days old) grown in nitrogen-
fixing conditions and fresh nodules collected from a 6-month-old plant were treated
immediately after harvest with RNA ProtectTM Bacteria Reagent (Qiagen) to stabilize the
RNA in the bacterial cells. To remove the DNA, the extracted RNA was treated with
DNA-freeTM prior to cDNA synthesis using an iScriptTM cDNA Synthesis Kit according
to the manufacturer’s (Ambion) protocol.
PCR: Fragments of genes (800 bp long) encoding structural subunits of hydrogenases
from several Frankia strains were amplified by touchdown PCR using designed primers
listed in Table 3. The PCR was performed with 25 ng of DNA in 20 µl mixtures (0.6 µM
of each primer, 3 mM of either MgCl2 or Q solution, and 1 U Taq from Qiagen, with
temperature programs of 3 min at 95 oC followed by 35 cycles of 30 s at 95 oC, 15 s at 58
oC, 15 s at 54 oC and 1 min at 72 oC, then a final elongation step of 10 min at 72 oC. PCR
products were electrophoretically separated on agarose gels, purified using a QIAquick
Gel Extraction Kit (Qiagen) and sequenced by an ABI377 sequencer (Applied
Biosystems).
Sequence alignments and phylogenetic analysis: Multiple alignments of hydrogenase
sequences of several Frankia strains and other related organisms were constructed using
ClustalX (Thompson et al., 1997). Matrix pair-wise comparisons were corrected for
multiple-base substitutions according to the method of Kimura (1980), followed by a
28
phylogenetic analysis using Neighbor-Joining (Saitou and Nei, 1987) with standard
parameters. Bootstrap confidence analysis was performed using 1000 replicates to
determine the reliability of the distance tree topologies obtained (Felsenstein, 1985).
Tree representations were constructed by Tree-View (Page, 1996).
Product Designation Forward (5’- 3’) Reverse (5’ to 3’) .
HupL1 HupL 20/HupL 21 cctcgttgacccagtccttg cgcatcatcggcaacctc
HupL1 HupL 20/HupL-13B cctcgttgacccagtccttg aaggggaaggatccacgcgacgcc
HupS1 HupS-6F/HupS-6B gttgtgccgccacctcggctc tgcgacggcgacacggtctcg
HupS1 HupS-6F/HupL 24 gttgtgccgccacctcggctc acggtccacctgcacaacaa
HupL2 HupL-F1/HupL33-2-(b) gacgtcacccactcgttctac cgttgatgacgaacctgct
HupL2 HupL32-2-(f)/HupL33-2-(b) tcacccactcgttctacgc cgttgatgacgaacctgct
HupS2 HupS34-2-(f)/HupS-B2 gatgtcatccgtgctctgg agccgaactcgtagaacagg
HupS2 HupS-F1/HupS35-2-(b) tcatccgtgctctggtttc gtgggtgaacgtggtgaag
HupS2 HupS34-2-(f)/HupSB1 gatgtcatccgtgctctgg gtcggtgatcaggtcgatg
HupL2 HupL-F1/HupL(II)_B3 gacgtcacccactcgttctac gacttggcccagctgtactt
HupL1 HupS(I)-27F/HupS(I)-27B acaccaggttgtcctggaag gtgttcatgaaggggaagga
HupS1 HupS(I)_26_F3/HupL(I)-26B caccgttgatgttctcgttg ctgcacaacaaggtgctctc
16S rDNA 16S-F134/16S-B134 gatttatcggctcgggatg gtaggagtctgggccgtgt .
Table 3. List of primers used in the study.
Transcriptional analysis of Frankia hydrogenases
Real-time PCR and conventional RT-PCR: Gene transcripts were measured by
amplification using primers specific for the structural subunits of F. alni strain ACN14a
hydrogenase. The primers (Table 3) were designed using Primer Premier 5 software.
Triplicate amplification of all standards, unknowns and controls was performed using a
multicolour iCycler iQ Real-Time PCR Detection System. Unknowns were compared
with cDNA standards covering four dynamic ranges obtained by serial dilution of
quantified starting concentrations. Expression levels were calculated from standard
curves generated during each run and with each primer pair. The acquired real-time PCR
data were analyzed by the relative expression software tool REST© (Pfaffi et al., 2002).
29
Conventional PCR was performed in varying conditions, as optimized for the sequences
and primers concerned, and in some cases by lowering the annealing temperature
gradually from 59ºC to 56ºC.
The aim of this thesis
• To study the diversity of hydrogenases in Frankia isolated from different
actinorhizal species growing in different regions.
• To characterize the hydrogenases of Frankia at the molecular level (gene/peptide
sequencing etc).
• To study the evolutionary relationships between uptake hydrogenases of Frankia
isolated from a various different actinorhizal plants from growing in different
regions, and also in comparison with hydrogenases of other organisms.
• To study the regulation of Frankia hydrogenases under different physiological
conditions.
Results and discussion
Uptake hydrogenases in Frankia
The biodiversity project this thesis is based upon was started by screening 18 Frankia
strains originally isolated from ten different actinorhizal host plants for physiological
activity of uptake hydrogenases at day eight of their growth in a medium with no
nitrogen (nitrogen-fixing conditions), in which the activity was expected to be maximal
(Mattsson and Sellstedt, 2000). The physiological activity data were subsequently
confirmed by molecular biology techniques, including blotting (Western, Southern),
30
mass spectrometry and PCR as discussed in the following sections. Hydrogenase activity
was detected in all strains investigated (Paper I), corroborating earlier studies in which
activity was found in Frankia capable of infecting Alnus in northern Sweden (Sellstedt,
1989) and Casuarina (Sellstedt et al., 1991).
The molecular characterization of uptake hydrogenases in Frankia
Western analysis: To confirm the physiological findings, several Frankia strains were
screened to investigate whether their uptake hydrogenases are immunologically related
to hydrogenases from other organisms. Antibodies rose against the large subunit of Ni-
Fe hydrogenase of R. eutropha (HoxG) recognized a polypeptide at about 60 kDa,
corresponding to the large hydrogenase subunit, in Frankia UGL020603, KB5 and
AvCI1. The HoxG antibody also recognized hydrogenases of other strains such as
Frankia sp. UGL140102 (Paper I). These results do not prove the absence of uptake
hydrogenases in other Frankia strains, but rather indicate the diversity of uptake
hydrogenases in Frankia. For example, antibodies raised against the large subunit of the
[NiFe] hydrogenase of R. eutropha recognized the large subunit of hydrogenase of
Frankia sp. KB5, but antibodies raised against the large subunit of [NiFe] hydrogenase
of A. vinelandii did not (Mattsson et al., 2001), and no hydrogenase subunits in any of
the Frankia strains investigated to date appear to be immunologically related to the small
subunit of the Ni-Fe hydrogenase (Hup S) in B. japonicum (Paper I). In the study of the
interspecies immunological cross-reactivity of hydrogenases, in seven cases the
immunological tests showed between-strain cross-reactivity with the large hydrogenase
subunits but not with the small subunits, suggesting that at least one conserved protein
region is present among the large subunits of these enzymes, while their small subunits
are less conserved (Kovács et al., 1989).
Peptide analysis: Protein spots of Frankia sp. KB5, corresponding to 60 kDa of the large
subunit of uptake hydrogenase, were excised from Coomassie-stained 2-D mini gels and
analyzed by matrix-assisted laser-desorption ionization-time-of-flight mass spectrometry
(MALDI-TOF). The resulting peptide ‘fingerprint’ showed identity (20% matches) with
31
the membrane-bound hydrogenase 2 large subunit (HYD2) in Escherichia coli (Acc.
P37181). The matched peptides covered 83% (476/567 amino acids) of the protein
(Paper II).
PCR and southern blot analysis: An NCBI blastx search with the translated sequence of
a 127-bp PCR-amplified gene fragment from F. alni AvCI1yielded up to 76% similarity
with the large hydrogenase subunit of various other organisms, e.g. Azotobacter
chrococcum (Paper II).
Uptake hydrogenase activity has been recorded from all Frankia strains
investigated but one, namely Frankia “local source” (Sellstedt et al., 1986; Paper I).
However, interestingly, a partial sequence of 500 bp could be amplified from DNA
isolated from nodules of the symbiosis between Frankia ‘local source’ and A. incana
(Mattsson, 2001; Paper VI), which was analyzed using the NCBI-translated query versus
the protein database (blastx), yielding 69% and 67% identity with the small subunits of
hydrogenases of B. japonicum and Rhi. leguminosarum, respectively. In addition, using
Southern-blot analysis, the hupS fragment of Frankia ‘local source’ hybridized with
DNA isolated from Frankia sp. KB5 (Paper VI). However, the Frankia DNA used was
extracted from nodules in which several different Frankia strains may be present. It is
possible that the uptake hydrogenase of Frankia “local source” may be active only under
specific environmental conditions, which are not yet known, or that it has an uptake
hydrogenase system but some regulatory genes are missing that are required to make an
active enzyme. It has been reported that Rhi. leguminosarum bv. viciae is unable to
express its uptake hydrogenase in free-living conditions because it contains a defective
hoxA gene (Brito et al., 1997). A hup-specific transcriptional activator encoded by the
hoxA gene is known to control hydrogenase gene expression in A. eutrophus (Friedrich
and Schwartz, 1993) and B. japonicum (Van Soom et al., 1993).
The recent release of the sequences of three Frankia genomes: F. alni ACN14a
(GenBank accession no. CT573213), Frankia sp. HFPCCi3 (GenBank accession no.
CP000249) and Frankia sp. EAN1pec (GenBank accession no. ZP_00571168) by
Genoscope (France), the National Science Foundation (USA) and the U.S. Department
of Energy Joint Genome Institute, respectively, have provided a wealth of information
32
and opportunities for studying Frankia hydrogenases (Normand et al., 2007a). Although
some standard genetic techniques cannot be applied to Frankia as yet (since Frankia has
never been genetically transformed), rapidly developing molecular biology techniques,
such as gene and protein arrays, could be exploited to make use of the available data and
study the molecular biology of Frankia. Further characterization of Frankia
hydrogenases (following the availability of the Frankia genome sequences) will be
addressed in the following sections.
The structure of uptake hydrogenase genes in Frankia
The genome analyses have shown the presence of two hydrogenase syntons in Frankia,
which are distinctly separated in all three genomes (Paper III). The structural, regulatory
and accessory genes of the hydrogenases are arranged closely together in each synton. In
F. alni ACN14a, hydrogenase synton #1 corresponds to GI:111221817–111221829 and
is situated at co-ordinates 2614407–2627969, whereas hydrogenase synton #2,
corresponds to GI:111221263–11221273 and is situated at co-ordinates 1959070–
1971271. Hydrogenase syntons #1 and #2 of Frankia sp. CcI3 and Frankia sp. EAN1pec
correspond to GI:86740641–86740652 and GI:86739780–86739790 and GI:68199305–
68199315 and GI:68232162–68232167, respectively (Fig. 3; Paper III). The gene
encoding the small subunit of the hydrogenase is located upstream of the large subunit in
Frankia, in accordance with several other organisms like Nostoc sp., Rhi.
leguminosarum and B. japonicum (Przybyla et al., 1992; Vignais and Toussaint, 1994;
Voordouw, 1992; Wu and Mandrand, 1993).
All of the available evidence indicates that hydrogenase syntons #1 and #2 are
both uptake hydrogenases that have many features in common with uptake hydrogenases
of other Frankia strains or other organisms. For example, the gene structures of
hydrogenase syntons #1 and #2 of F. alni ACN14a are very similar to those of
hydrogenase syntons #1 and #2 of Frankia sp. CcI3, respectively. The uptake
hydrogenases of Frankia may also have some dissimilarity, even in the same Frankia
strain. For example, the physical position and orientation of the uptake hydrogenase
genes varies very clearly between hydrogenase syntons #1 and #2 in F. alni ACN14a,
33
Frankia sp. CcI3 and EAN1pec. Interestingly, other symbiotic gene clusters that have
been found in the Frankia genome: the nitrogen fixation (nif) and squalene hopane
cyclase (shc), have similar numbers of genes in similar arrangement in F. alni ANC14a,
Frankia sp. CcI3, and EAN1pec (Normand et al., 2007b). In addition, a simple sequence
comparison indicated that the sequence conservation between the structural subunits of
hydrogenase syntons #1 and #2 in F. alni ACN14a itself (e.g. HupL1 of synton #1 vs.
HupL2 of synton #2) was as low as 27%, whereas that between F. alni ACN14a and
non-Frankia bacteria such as S. avermitilis was as high as 73%. Clearly, one of the
syntons is not simply a duplicate of the other, but rather both are required for hydrogen
metabolism under different circumstances in Frankia (Paper III).
Fig. 3. Genome maps of Frankia sp. EAN1pec and CcI3. Circles, from the outside in,
show (1) gene regions related to symbiosis including shc1, hup2, hup1, and nif; (2) the
coordinates in Mb beginning at 0 = oriC; (3) regions of synteny (syntons) calculated as a
minimum of five contiguous genes present in all strains with an identity >30% over 80%
of the length of the shortest gene. Redrawn from Normand et al., 2007a.
34
The phylogeny of uptake hydrogenases in Frankia
The phylogenetic analysis of the structural subunits hydrogenase syntons #1 and #2 of F.
alni ACN14a, Frankia sp. CcI3 and EAN1pec has shown that the two syntons are
distinctly different. This is not surprising considering the degree of sequence divergence
between them, even in the same Frankia strain. The phylogenetic and sequence
similarity of one of the hydrogenase syntons to hydrogenases of other organisms, on the
other hand, is remarkable.
According to the phylogenetic trees of uptake hydrogenase syntons #1 and #2, the
structural subunits of F. alni ACN14a and Frankia sp. CcI3, which belong to
phylogenetic Frankia cluster 1, group together but not with those of Frankia sp.
EAN1pec, which belongs to Frankia cluster 3 (Normand et al., 1996). HupL2 of
Frankia sp. EAN1pec appears to be most closely related to the hydrogenases of the non-
Frankia bacteria Geobacter Sulfurreducens, providing strong evidence for the
occurrence of lateral gene transfer (LGT) between these organisms. Clearly, neither of
the syntons is simply a recent duplicate of the other, and functional complementarities
are less likely, given their apparent sequence divergence. The tree topology is indicative
of probable gene transfer to or from ancestral organisms that occurred before the
emergence of Frankia. All of the available evidence points to hydrogenase gene
duplication having occurred long before emergence of the three Frankia lineages (Paper
III).
Phylogenetic analysis of the structural subunits of hydrogenase syntons #1 and #2
(analyzed separately) showed distinct clustering among hydrogenases of various Frankia
strains (Paper VI). The large subunits of hydrogenase synton #1 of Frankia sp. CpI1, F.
alni ACN14a and AvCI1 (isolated from A. viridis subsp. crispa) grouped together, while
those of Frankia sp. CcI3, KB5, UGL140104 and UGL011102 (isolated from C.
cunninghamiana, C. equisetifolia, H. rhamnoides and A.incana, respectively) formed
another group (Table 4). The large subunits of hydrogenase synton #2 of F. alni
ACN14a, Frankia sp. CcI3 and BCU110501 (isolated from D. trinervis) also grouped
together, while those of Frankia sp. KB5, CpI1, F. alni AvCI1 and ArI3 (isolated from
35
Hydrogenase synton Phylogenetic groups Frankia strains . # 1 A Frankia sp. CpI1, Frankia sp. AvCI1,
F. alni ACN14a
B Frankia sp. EAN1pec, R. eutropha*,
S. erythraea*, S. avermitilis*
C Frankia sp. CcI3, Frankia sp. KB5,
Frankia sp. UGL011102, UGL140104
# 2 A Frankia sp. EAN1pec, Anaeromyxobacter sp.*
D. ethenogenes*, C. hydrogenoformans*
B F. alni ACN14a, Frankia sp. CcI3,
Frankia sp. BCU110501
C Frankia sp. KB5, Frankia sp. CpI1,
. F. alni AvCI1 and ArI3 .
Table 4. Summary of the phylogenetic relationships between the large subunits (HupL)
of hydrogenase synton #1 (and #2) of various Frankia strains (also in comparison with
hydrogenases of other organisms). *Non-Frankia bacteria.
Alnus rubra) formed another group (Table 4). The Frankia strains which grouped
together might probably have related ancestors. Interestingly, the structural subunits of
both hydrogenase syntons #1 and #2 of Frankia sp. EAN1pec appear to be more closely
related to the hydrogenases of non-Frankia bacteria than they are to hydrogenases of
Frankia and thus it may have acquired its hydrogenase in a different way from the
Frankia strains in the two subgroups. We were unable to establish a connection between
the phylogenetic relationships of Frankia uptake hydrogenases and the geographical
distribution of Frankia strains or their hosts. However, a link between the biogeographic
history of the actinorhizal plants and the genome evolution of the bacterial symbionts
36
has been proposed from a comparative study of the whole genomes of F. alni ACN14a,
Frankia sp. CcI3 and EAN1pec (Normand et al., 2007a).
The Hyp genes of Frankia hydrogenases are closer to those of bacteria than
archaea, and the Hyp genes of hydrogenases of F. alni ACN14a and Frankia sp. CcI3
are more closely related to each other than they are to those of Frankia sp. EAN1pec (for
both hydrogenase syntons #1 and #2). HypD1 and HypF1 (synton #1) and HypB2 and
HypE2 (synton #2) of F. alni ACN14a and Frankia sp. CcI3 are more strongly related to
those of non-Frankia bacteria than those of Frankia sp. EAN1pec. Frankia sp. CcI3 and
F. alni ACN14a have similar contents and orientation of genes in their uptake
hydrogenase synton #1 , while the syntons are both reduced and rearranged in Frankia
sp. EANpec, although Frankia EAN1pec has the largest genome (9.0 Mb), followed by
F. alni ACN14a (7.5 Mb) and Frankia sp. CcI3 (5.4 Mb) (Normand et al., 2007a; Fig.
3). Similarly, the content and orientation of genes in uptake hydrogenase synton #2 in F.
alni ACN14a and Frankia sp. CcI3 are very similar, but quite different in Frankia
EAN1pec. Frankia EAN1pec may have acquired its uptake hydrogenase by lateral gene
transfer from other non-Frankia bacteria in a different way from Frankia sp. CcI3 and F.
alni ACN14.
The regulation of uptake hydrogenases in Frankia
From the structural and phylogenetic studies outlined above, the uptake hydrogenase
syntons of Frankia are clearly not gene duplicates but different isozymes, which might
be required for hydrogen metabolism under different circumstances (Paper III). In
accordance with this hypothesis, there are indications that several microorganisms may
express different types of hydrogenases with differing specific functions under different
environmental conditions (Laurinavichene et al., 2002).
Hydrogenase gene expression in free-living vs. symbiotic condition: The transcript
levels of the structural subunit genes of uptake hydrogenase synton #1 (hupS1 and
hupL1) were higher than those of hydrogenase synton #2 (hupS2 and hupL2) in F. alni
ACN14a grown under free-living conditions. In contrast, the transcript levels of uptake
37
hydrogenase synton #2 were higher than those of hydrogenase synton #1 under
symbiotic conditions, with observed hupL1:hupL2 expression ratios of 2:1 under free-
living conditions and 34:1 under symbiotic conditions. Therefore, synton #1 uptake
hydrogenases of Frankia are expressed more strongly under free-living conditions, and
synton #2 hydrogenases are mainly involved in symbiotic interactions (Paper III). It was
not possible to further confirm this result at a protein level since we have not as yet been
able to raise antibodies against the structural subunits of Frankia hydrogenases.
Ni-dependent regulation of uptake hydrogenases in Frankia: Nickel is an essential
micronutrient for many microorganisms, which is incorporated into at least four
microbial enzymes that participate in important reactions of hydrogen metabolism,
ureolysis, methane biogenesis, and acetogenesis (Hausinger, 1987). The presence of
nickel had previously been shown to have positive effects on the activity of uptake
hydrogenase in free-living Frankia strains (Sellstedt and Smith, 1990; Mattsson and
Sellstedt, 2002). In the studies underlying this thesis, the activity of the uptake
hydrogenases in Frankia UGL011301 and R43 were also found to be significantly
enhanced by the addition of nickel to the growth medium (Paper I). Since it is important
for microorganisms to maintain precise homeostasis of nickel ions in their cells, various
organisms have been shown to have evolved nickel-specific sensing and transport
systems (Eitinger and Mandrand-Berthelot, 2000), allowing them to take up nickel when
it is required, and to avoid doing so when it is not needed, via an inducible nickel-
resistance mechanism (Grass et al., 2003). Frankia may also have evolved a mechanism
to take up nickel when needed and avoid doing so when it is in excess, although there are
no data to support this hypothesis as yet
Thin cryosections of free-living Frankia strains treated with antiserum raised
against the hydrogenase of A. latus (the holoenzyme) showed that there were higher
degrees of hydrogenase transfer into the membranes of nickel-treated cells than in
nickel-free controls. The relative abundance of gold particles was much higher in
membranes when nickel had been added than in cells lacking nickel, where the gold
particles were localized mainly in the cytoplasm. As in various other organisms, the
38
processing and correct positioning of the protein in the membrane of Frankia is essential
for the enzyme to be biologically active (Paper I).
Effects of nitrogenase and hydrogen on the uptake hydrogenase of Frankia: Frankia
sp. KB5 produces its uptake hydrogenase only when grown in nitrogen-free media.
Frankia sp. R43, on the other hand, produces its uptake hydrogenase when cells are
grown in both nitrogen-fixing and non-nitrogen-fixing conditions (Paper IV). Frankia
sp. R43, unlike other Frankia strains, can produce vescicles in both nitrogen-fixing and
non-nitrogen-fixing conditions (Fernando, 1991). A strong correlation between uptake
hydrogenase and nitrogenase activity in Frankia KB5 grown in nitrogen-fixing
conditions has previously been reported (Mattsson and Sellstedt, 2000). Some uptake
hydrogenases of Frankia may be produced independently of nitrogenase, but
dependently on the hydrogen produced as an inevitable byproduct of the nitrogen
fixation process. In accordance with this hypothesis, Frankia KB5 grown in non-
nitrogen-fixing conditions, but in the presence of exogenous hydrogen, can produce an
uptake hydrogenase (Mattsson and Sellstedt, 2000). In addition, Frankia sp. R43 may
have another type of uptake hydrogenase, which can be produced independently of both
nitrogenase and hydrogen. In contrast, uptake hydrogenase and nitrogenase synthesis are
coregulated in Rhizobium leguminosarum bv. viciae by the nitrogen fixation regulator
NifA (Brito et al., 1997) and two FnrN proteins (Gutiérrez et al., 1997). Coregulation of
hydrogenase and nitrogenase synthesis in R. capsulatus (by the RegB-RegA two-
component regulatory system) has also been reported (Dischert et al., 2000).
A hydrogen-evolving enzyme in Frankia
A broad range of Frankia strains grown in a medium without nitrogen (and without
nickel) were screened to investigate the presence of hydrogen-evolving hydrogenases in
the genus. The activity of the enzymes was measured as H2 production by gas
chromatography (Mattsson and Sellstedt, 2000). Ammonium chloride was added at the
start of induction of hydrogen evolution to block hydrogen production from the
39
nitrogenase-catalyzed nitrogen-fixation process, and ARA measurements were taken to
check that all nitrogenase activity was eliminated.
Methyl viologen-mediated hydrogen evolution was recorded in only four Frankia
strains: F. alni UGL011101, UGL140102, Frankia sp. CcI3 and R43, originally isolated
from three different host plants. The highest rate of hydrogen evolution recorded was
obtained from Frankia UGL140102 (Paper IV). This is the first time, to our knowledge,
that evidence of methyl viologen-mediated hydrogen evolution in the actinomycete
Frankia has been published. The hydrogen recorded from Frankia sp. R43 did not
originate from an uptake hydrogenase acting in the reverse direction, since this protein
was not present in anaerobic conditions, as confirmed by the measurements of enzyme
activity and western blot analysis.
Molecular characterization of the hydrogen-evolving enzyme in Frankia
The presence of a hydrogen-evolving enzyme in Frankia was confirmed by Western
immunoblot analysis using antisera raised against HoxF of R. eutropha, which
recognized a 60 kDA protein in Frankia sp. R43 extracts (Paper IV).
Gel and peptide analysis: Frankia R43 proteins extracted from cells grown in anaerobic
conditions were electrophoretically separated on replicate 2-D gels. Protein spots of a
molecular weight of approx. 47 kDa identified by immunoblots were excised, digested
and analyzed using ESI-MS/MS Q-TOF. Short sequences in the resulting peptide
fingerprint showed nearly exact matches in searches of the NCBI protein database to
protein sequences of the bidirectional hydrogenase hoxH subunit of Anabaena siamensis
TISTR8012 (GenBank accession no. AAN65267). Thus, the hydrogen-evolving enzyme
in Frankia has strong similarity to this bidirectional cyanobacterial hydrogenase.
However, the metabolic function of the hydrogen-evolving hydrogenase in Frankia does
not appear to involve NAD-reduction, as proposed for the bidirectional hydrogenase in
cyanobacteria (Appel and Schulz, 1996), since no NAD-reducing activity was detected
in Frankia cells in which hydrogen evolution had been induced (Paper V). The
hydrogen-evolving hydrogenase in Frankia may act as an electron scavenger under
40
anaerobic conditions, especially since Frankia is commonly found in
microaerophilic/anaerobic environments, e.g. in root nodules in its symbiotic form and in
river and lake sediments in its free-living form (Huss-Danell et al., 1997).
Localization of the hydrogen evolving enzyme in Frankia: Cryosectioning in
combination with immuno-gold labeling techniques using antibodies raised against
HoxH of the SH-hydrogenase HY of R. eutropha was performed to study the subcellular
localization of the hydrogenase in Frankia cells grown in nitrogen-limiting conditions
(and without nickel) for seven days and then kept anaerobically for 24 h (the conditions
used for the physiological measurements). A polypeptide was recognized by the
antibody in both hyphae and vesicles of Frankia sp. R43 (Paper IV) and Frankia sp.
UGL140102 (Paper V), the labeling being evenly distributed in these organelles,
indicating that the hydrogen-evolving hydrogenase is a soluble enzyme.
Regulation of the hydrogen-evolving enzyme in Frankia
Nitrogenase and the hydrogen-evolving hydrogenases of Frankia: Frankia sp. R43
showed methyl-viologen-mediated hydrogen evolution when grown in either nitrogen-
fixing or non-nitrogen fixing conditions. On the other hand, Frankia sp. KB5 showed
methyl-viologen-mediated hydrogen evolution only when grown in nitrogen-fixing
conditions (Paper III). Although it is premature to conclude that there is any correlation
between the expression of the nitrogenase and hydrogen-evolving enzymes, the
possibility that the nitrogenase enzyme affects the regulation of the hydrogen-evolving
enzyme directly or indirectly, at least in some Frankia strains, cannot be excluded. In
cyanobacteria, the level of activity of the bidirectional hydrogenase enzyme has been
found to be unaffected by exposing cells to hydrogen during their growth, and by the
addition of nitrogen to the growth medium, indicating that the expression this enzyme is
not dependent on, or even related to diazotrophic growth conditions, unlike their uptake
hydrogenase, the expression of which is linked to nitrogenase expression (Schütz et al.,
2004). The biosynthesis of nitrogenase is not essential for the biosynthesis of
bidirectional hydrogenase and hydrogen evolution in several unicellular strains (Howarth
41
and Codd, 1985). Hydrogenases of cyanobacteria are expressed independently of
nitrogenase synthesis (Houchins, 1984).
Ni-dependent regulation of the hydrogen-evolving hydrogenases of Frankia: nickel
appears to play a role in the regulation of the hydrogen-evolving enzyme in Frankia.
Methyl viologen-mediated hydrogen evolution was recorded from F. alni UGL011103,
Frankia sp. UGL020602, UGL020603, and 013105 when grown in a medium without
nitrogen, but only in the presence of nickel (II) chloride (Paper V). These Frankia strains
did not display methyl viologen-mediated hydrogen evolution when grown in nitrogen-
fixing conditions without nickel (Paper I). Frankia may have different types of
hydrogen-evolving hydrogenases, or the hydrogen-evolving hydrogenases of Frankia
may at least be regulated differently in different Frankia strains. HypB genes encode
proteins that are highly conserved between different organisms and consist of at least
four domains (Robson, 2001). The presence of multiple histidinyl residues, which
characterizes the second domain of the proteins in several microorganisms like B.
japonicum, R. capsulatus, Azotobacter sp., B. leguminosarum (but which are known to
be missing in Archaea), suggests that this protein plays a role in binding divalent metals,
especially nickel (Rey et al., 1994; Robson, 2001). HypB, the nickel-binding GTPase, is
involved in nickel storage/sequestering and incorporation of nickel into the hydrogenase
in B. japonicum (Olson et al., 1997; Olson and Maier, 2000). The hydrogen sensor
hydrogenase HupUV, which regulates hydrogenase synthesis (e.g. in B. japonicum),
requires nickel to be active. The HypB2 of Frankia sp. CcI3 has seven histidinyl
residues in this domain while that of F. alni ACN14a has only one, and Frankia sp.
EAN1pec has none at all (Table 5). The mechanism whereby nickel regulates
hydrogenase transcription in Frankia is not known, but the available sequence
information indicates that, as in other organisms like B. japonicum, it may be via the
HypB, although this needs to be experimentally confirmed. HypB protein lacking the
nickel-binding polyhistidine region near the N terminus lacks the ability to store nickel,
but it is still able to bind a single nickel ion and also retains complete GTPase activity
(Rey et al., 1994).
42
F. alni ACN14a*1 ------------------------------------------------MG 2 Frankia sp. CcI3*2 ----MGRFHPHPEGAHPHPEGAPHEYSGPHPPAGVS------------VG 34 Mycobacterium vanbaalenii3 ----MGRFHRHDDG-----TAHTHDHDG---SPSHD------------HG 26 Frankia sp. EAN1pec*4 ----MRRSGSPPSRCADLRAGWTSRRVGRPGR--LD--VPAGWSGGGLMC 42 Nodularia spumigena*5 MCVTCGCSDDAESTITNLETGEVEHNHHDHTHTLLDGTVISHSHNHDTQH 50 Methanosarcina mazei*6 -------------------------------------------------- Archaeoglobus fulgidus*7 -------------------------------------------------- F. alni ACN14a DHSGYRTGAE--RVEVLERILGENEKVARANRAAFDAAGVTVVNLMSAPG 50 Frankia sp. CcI3 DHAGYGTGPE--RVEVLERILGENERVALANRAAFDAAGVTVVNLMSAPG 82 Mycobacterium vanbaalenii DHSGYHTAAE--RVDVLEAIFSENDLRAAANRNAFEENGIRALNLMSSPG 74 Frankia sp. EAN1pec GTCGCEAETR--TLRLEMDVLARNEESADDNRAWLAARRASAVNLMSSPG 90 Nodularia spumigena EASQVHAKIHNTTISLEQDILAKNNLIAAQNRGWFKGRNILALNLMSSPG 100 Methanosarcina mazei -----MMFMLMHVIHMGHDVYKANDKIAEKNRKTLDKHGVFSVNVMGAIG 45 Archaeoglobus fulgidus ----------MHEYELNQDLLAENKRLAEKNREALRESGTVAVNIMGAIG 40 : : *. * ** : :*:*.: *
Table 5. Multiple sequence alignments of N-terminal sequence of the HypB proteins
from Frankia (hydrogenase synton #2), non-Frankia bacteria and Archaea (produced by
ClustalW). *1-*7 Protein_IDs: *1 = 111221273, *2 = 86739790, *3 = 120403348, *4 =
68233089 *5 = 119509235, *6 = 21228420 and *7 = 11498964. The Domain 2 of HypB
proteins of many Eubacteria are known to have multiple histidinylresidues that are known
to bind divalent metals, especially nickel.
Does Frankia have other hydrogenases?
Frankia may also produce Fe-only hydrogenases, since hydrogenases of Frankia
UGL011102 and Frankia KB5 were found to be immunologically related to the [Fe]-
hydrogenase of D. desulfuricans ATCC 7757. However, more experiments are required
to confirm this finding.
Conclusions
Frankia has at least three types of hydrogenases.
Uptake hydrogenases are common in Frankia and are probably Ni-Fe hydrogenases.
Frankia commonly has two uptake hydrogenase syntons, which are distinctively
separated in their genome. The structural, accessory and regulatory genes of these
43
hydrogenases syntons are organised tightly together. These hydrogenase syntons differ
in many ways.
Hydrogenase synton #1 and #2 in a Frankia are phylogenetically divergent and
hydrogenase synton #1 is probably ancestral among the actinobacteria.
The hydrogenases genes of F. alni ACN14a and Frankia sp. CcI3 are closely related but
relatively distant from those of Frankia sp. EAN1pec, which was more related to the
hydrogenases of non-Frankia bacteria than Frankia. The tree topology is indicative of
probable gene transfer to or from ancestral organisms that occurred before the
emergence of Frankia. All of the available evidence points to hydrogenase gene
duplication having occurred long before development of the three Frankia lineages.
Uptake hydrogenase synton #1 of Frankia is expressed more strongly under free-living
conditions but hydrogenase synton #2 is mainly involved in symbiotic interactions.
Nickel enhances the degree of hydrogenase transfer into the membranes and the activity
of uptake hydrogenases in Frankia. The processing and correct positioning of the protein
in the membrane of Frankia is essential for the enzyme to be biologically active.
Some Frankia strains have a hydrogen-evolving enzyme. This hydrogenase is a soluble
enzyme as it is localized in both hyphae and vesicles, and is related to the bidirectional
cyanobacterial hydrogenases.
Unlike other Frankia strains, Frankia sp. R43 showed a methyl-viologen-mediated
hydrogen evolution and uptake hydrogenase activity in both nitrogen-fixing and non-
nitrogen-fixing conditions.
Nickel appears to play a role in the regulation of the hydrogen-evolving enzyme in
Frankia as some Frankia strains showed activity of this enzyme only in the presence of
this metal. Frankia may have different types of hydrogen-evolving hydrogenases, or the
44
hydrogen-evolving hydrogenases of Frankia may at least be regulated differently in
different Frankia strains.
Hydrogen-evolving enzyme of Frankia evolves a considerable amount of hydrogen that
can be used as a clean energy source.
Future perspectives
It would be interesting to further study:
• The activities of uptake hydrogenases synton #1 and #2 of F. alni ACN14a in
symbiotic vs. free-living conditions to in order to confirm the present result of
their differential expression under these conditions. It would also be interesting to
see if this difference also exists in the uptake hydrogenase synton #1 and #2 of
other Frankia strains.
• The effect of nickel and selenium on uptake hydrogenase synton #1 and #2 of
Frankia at both mRNA and protein levels.
• The Fe only hydrogenases Frankia strains.
• The effect of different physiological conditions on the activity of the hydrogen-
evolving hydrogenases in Frankia. This helps to know the optimum condition in
which maximum biological hydrogen production can be obtained from Frankia.
45
Frankia – bakterien som pruducerar både kväve gödsel och vätgas!
Alla 18 Frankia stammar som isolerats från tio aktinorhiza värdväxter härstammande
från olika delar av världen, hade det väte-förbrukande enzymet hydrogenas. Tillsättning
av nickel till odlingsmediet ökade enzymets aktivitet och mängd i membran, vilket tyder
på att nickel behövs för ett aktivt enzym och att enzymet sannolikt är ett Ni-Fe
hydrogenas.
Karakterisering av tre Frankia genom v
isade närvaro av två helt skilda hydrogenas kluster. De två hydrogenasen är även vanliga
hos andra Frankia stammar. De strukturella, reglerande och accessoriska generna i
hydrogenas kluster#1 och #2 ligger intill varandra, men har olika struktur. Medans
hydrogenas kluster #1 (eller kluster #2) av F. alni ACN14a och Frankia sp. CcI3 har
likande gen sammansättning, innehåll och orientering har Frankia sp. EAN1pec minskat
antal gener och är annorlunda organiserat. Hydrogenas från Frankia sp. ACN14a och
CcI3 är fylogenetiskt mer relaterad till varandra än till Frankia sp. EAN1pec, som i sin
tur är mer relaterad till en icke-Frankia bakterie. Fylogenetiska och topologiska studier
tyder på en sannolik genöverföring till eller från Frankia långt innan Frankias
uppkomst. De väte-förbrukande hydrogenasets kluster #1 är mer uttryckt i frilevande
Frankia medans hydrogenas kluster #2 är huvudsakligen involverad i rotknölar. Detta
hydrogenas kan spela en betydande roll för att öka effektiviteten av kväve-fixeringen i
rotknölar hos värdväxter med Frankia och därmed kan behoven av miljöovänligt och
dyrt gödningsmedel minskas.
De väte-producerande hydrogenaset återfanns endast hos fyra Frankia stammar,
F. alni UGL011101, UGL140102, Frankia sp. CcI3 och R43. Efter tillsättning av nickel
visade även F. alni UGL011103, Frankia sp. UGL020602, UGL020603, och 013105
aktivitet. Nickel bidrog också här till en ökning, vilket indikerar att Frankia möjligen
har olika typer av väte-producerande hydrogenas, eller att enzymet regleras på olika sätt
i olika Frankia stammar. Det faktum att Frankia producerar väte blev nyligen känt.
Därför är molekylärbiologiska kunskaper om hydrogenaser av yttersta vikt för att
optimera ett system som kan bli en biologisk producent av väte.
46
Acknowledgements
First and foremost, praise and thanks goes to my heavenly father, who loved me and gave
me His only begotton Son, my Lord Jesus Christ, and in Him, everything else. May His
Name be glorified for ever and ever, Amen!
I am privileged to do my PhD studies at Fys-Bot, which is for sure the nicest working
environment I have ever experienced. I am very thankful to all senior scientists, PhD
students, technical staff, post-docs for contributing to this work, directly or indirectly.
Special thanks to:
My Parents – you didn’t have a lot for your self, but you gave me all you have. You
believed in me, you encouraged me, you loved me. Thank you very much.
My better half – Genni, you are a special gift to me! Thank you for being patient when
we lived apart. Thank you for your love, understanding, encouragement and prayers.
Anita Sellstedt – my supervisor, for introducing me to molecular biology and laboratory
work, for providing me with all I needed for my study (for allowing me to attend
international conference etc), for the courage you gave me by believing in me. You have
been very understanding, especially in times of need, thank you! Thank you for helping
me with, among many other things, correcting the thesis.
Berhanu, Barbro, Rut and Hanna – Thank you for kindness. You have been a real
blessing to our life. Thank you for your concern, advice and prayers. You have a special
place in our heart.
Franscesco (Sir) – for being very nice to me. Thanks for the company, all the story we
shared together, for the ideas at meetings. Marie – thanks for your kindness and concern.
You have been very helpful. You are so organized, thank you for being a good example.
47
Anasuya, Prabha, Lars, Mats-Jerry, João, Jenny, Catarina and others - my Lab mates (of
the past), thank you for helpful ideas and your contribution. You have been so nice.
Stefan Jansson (Prof) and Per Gardeström (Prof) – my reference group, thank you for
your valuable comments.
All PhD students (of the last 5 years) – Jacob, Stefan, Andreas, Henrik, Maribel, Junko,
Charlene and many others – for helpful interaction and contribution to my work.
Philippe Normand (Prof) – for being available for advice. You have been very helpful.
John Blackwell – for correcting my English.
Slim, Inger, Monika, Siv, Karin, Brit-Marie, Gunilla, Ulrika, Eva, Susie, Rupali, Per-
Ingvar, Leszek, Laszlo, Catherine, Roland, Paavo, Lars, Janne, Hannele, Benedicte,
Simon, Jan, Thomas, Frank, Arsenio, Estelle, Luis and others – you have been very
helpful in the many ways I needed you, thank you!
Ale, Tesfu, Ben, Kalu, the king, Bezi – for your kindness and support; Wube, Aseged,
Abebaw, Chuni, Baby – for you kindness and support; Bekele, Tsehay, Betty, Eyerus,
Hanna – for your love and concern and prayers.
The Assosa community – for the advice, encouragement and fun.
Emmanuelgruppen - Tsehay and Mike; Saba and Haile; Tenagne and Jhoni; Dawit and
Martha; Misgina and Yordi; Mike; Firewoyni; all the small lovely kids. Thank you for
the great fellowship. Misgina, thanks for the hug! Yeshewas, you have been very helpful.
Fremi – thank you for correctiong my Swedish, stories we shared, for being a nice friend.
Haidy and Mattias; Per-Olov and Habibe - you made our life in Umeå easier. Thank you
for your advice, generosity and friendship.
Kemiförrådet, Vaktmästeriet KBC and Caelum (catering) staffs – for the help and fun.
48
References
Adams MWW (1990) The structure and mechanism of iron hydrogenases. Biochim
Biophys Acta 1020: 115-45.
Adams MWW, Mortenson LE, Chen J-S (1981) Hydrogenase. Biochim Biophys Acta
594: 105-76.
Adams MWW and Stiefel EI (1998) Biological hydrogen production: Not so elementary.
Science 282: 1842-43.
Afting C, Hochheimer A and Thauer RK (1998) Function of H2-forming
methylenetetrahydrometha-nopterin dehydrogenase, a metal free hydrogenase in
methanogenic Archaea growing on H2 and CO2. Arch Microbiol 169: 206-10.
Appel J, Phunpruch S, Steinmüller K and Schulz R (2000) The bidirectional
hydrogenase of Synechocystis sp. PCC 6803 works as an electron valve during
photosynthesis. Arch Microbiol 173: 333-38.
Appel J and Schulz R (1996) Sequence analysis of an operon of a NAD(P)-reducing
nickel hydrogenase from the cyanobacterium Synechocystis sp. PCC 6803 gives
additional evidence for direct coupling of the enzyme to NAD(P)H-dehydrogenase
(complex I). Biochim Biophys Acta 1298 (2): 141-47.
Baker DD (1987) Relationships among pure cultured strains of Frankia based on host
specificity. Physiol Plantarum 70: 245-48.
Baker D and Torrey JG (1980) Characterization of an effective actinorhizal
microsymbiont, Frankia sp. AvCI1 (Actinomycetales). Can J Microbiol 26: 1066-71.
49
Benson DR, Vanden Heuvel BD and Potter D (2004) Actinorhizal symbioses: Diversity
and biogeography. In Gillings M (ed) Plant microbiology. BIOS Scientific Publishers
Ltd., Oxford.
Benson DR and Silvester WB (1993) Biology of Frankia Strains, actinomycete
symbionts of actinorhizal plants. Microbiol Rev 57: 293-319.
Berghofer Y, Agha-Amiri K and Klein A (1994) Selenium is involved in the negative
regulation of the expression of selenium-free [NiFe] hydrogenases in Methanococcus
voltae. Mol Gen Genet 242: 369-73.
Berry A and Torrey JG (1979) Isolation and characterization in vivo and in vitro of an
actinomycetous endophyte from Alnus rubra Bong, p. 69–83. In Gordon JC, Wheeler CT
and Perry DA (eds), Symbiotic nitrogen fixation in the management of temperate forests.
Forest Research Laboratory, Oregon State University, Corvallis.
Binkley D, Cromack K Jr and Baker DD (1994) N fixation by red alder: biology, rates
and controls, p. 57–72. In Hibbs D, DeBell D and Tarrant R (eds), The biology and
management of red alder. Oregon State University Press, Corvallis.
Black LK, Fu C and Maier RJ (1994) Sequence and characterization of hupU and hupV
genes of Bradyrhizobium japonicum encoding a possible nickel-sensing complex
involved in hydrogenase expression. J Bacteriol 176: 7102-06.
Brito B, Martínez M, Fernández D, Rey L, Cabrera E, Palacios JM, Imperial J and Ruiz-
Argüeso T (1997) Hydrogenase genes from Rhizobium leguminosarum bv. viciae are
controlled by the nitrogen fixation regulatory protein NifA. Proc Natl Acad Sci USA 94:
6019-24.
Callaham D, DelTredici P and Torrey JG (1978) Isolation and cultivation in vitro of the
actinomycete causing root nodulation in Comptonia. Science 199: 899–902.
50
Cammack R (2001) Hydrogenases and their activities, p. 9-32. In Cammack R and
Florence KY (eds), Hydrogen as fuel: Learning from nature. London: Taylor & Francis.
Chaia E (1998) Isolation of an effective strain of Frankia from nodules of Discaria
trinervis (Rhamnaceae) Plant and Soil 205: 99-102.
Chaudhary AH and Mirza MS (1987) Isolation and characterization of Frankia from
nodules of actinorhizal plants of Pakistan. Physiol Plantarum 70: 255-58.
Clawson ML, Bourret A and Benson DR (2004) Assessing the phylogeny of Frankia-
actinorhizal plant nitrogen-fixing root nodule symbioses with Frankia 16S rRNA and
glutamine synthetase gene sequences. Mol Phylogenet Evol 31: 131-38.
Diem HG and Dommergues YR (1990) Current and potential uses and management of
Casuarinaceae in the tropics and subtropics, p. 317-342. In Schwintzer CR and
Tjepkema JD (eds), The biology of Frankia and actinorhizal plants. Academic Press,
Inc. San Diego, California.
Dischert SW, Colbeau A and Bauer CE (2000) Expression of Uptake Hydrogenase and
Molybdenum Nitrogenase in Rhodobacter capsulatus Is Coregulated by the RegB-RegA
Two-Component Regulatory System. J Bacteriol 182: 2831-37.
Dixon ROD (1972) Hydrogenase in legume root nodule bacteroids: occurrence and
properties. Arch Microbiol 85: 193-201.
Dixon ROD (1976) Hydrogenases and efficiency of nitrogen fixation in aerobes. Nature
263: 173.
Eitinger T and Mandrand-Berthelot M-A (2000) Nickel transport systems in
microorganisms. Arch Microbiol 173: 1-9.
51
Elsen, S., Colbeau, A., Chabert, J. & Vignais, P. M. (1996) The hupTUV operon is
involved in negative control of hydrogenase synthesis in Rhodobacter capsulatus. J
Bacteriol 178: 5174-81
Evans HJ, Koch B and Klucas R (1972) Preparation of nitrogenase from nodules and
separation into components. Methods Enzymol 24: 470-76.
Fernando Tavares (1991). Género Frankia (Actinomycetales) Aspectos morfológicos e
ultraestruturais. Msc thesis.
Felsenstein J (1985) Confidence limits on phylogenies: An approach using the bootstrap.
Evolution 39: 783-91.
Filipiak M, Hagen WR and Veeger C (1989) Hydrodynamic, structural and magnetic
properties of Megasphaera elsdenii Fe hydrogenase reinvestigated. Eur J Biochem 185:
547-53.
Friedrich B, Heine E, Finck A and Friedrich CG (1980) Nickel requirement for active
hydrogenase formation in Alcaligenes eutrophus. J Bacteriol 145 (3): 1144-49.
Friedrich B and Schwartz E (1993) Molecular biology of hydrogen utilization in aerobic
chemolithotrophs. Annu Rev Microbiol 47: 351-83.
Friedrich B, Vignaise PM, Lenz O and Colbeau A (2001) Regulation of hydrogenase
gene expression, p. 33-56. In Cammack R and Florence KY (eds), Hydrogen as fuel:
Learning from nature. London: Taylor & Francis.
Garcin E, Vernede X, Hatchikian EC, Volbeda A, Frey M and Fontecilla-Camps JC
(1999) The crystal structure of a reduced [NiFeSe] hydrogenase provides an image of the
activated catalytic center. Structure 7: 557-66.
52
Gaston KJ and Spicer JI (2004) Biodiversity: an introduction. Oxford: Blackwell
Publishing, 2nd ed., ISBN 1-4051-1857-1
Grass G, Fan B, Rosen BP, Lemke K, Schlegel HG and Rensing V (2001) NreB from
Achromobacter xylosoxidans 31A is a nickel induced transporter conferring nickel
resistance. J Bacteriol 183: 2803-07.
Gutiérrez D, Hernando Y, Palacios JM, Imperial J and Ruiz-Argüeso T (1997) FnrN
controls symbiotic nitrogen fixation and hydrogenase activities in Rhizobium
leguminosarum biovar viciae UPM791. J Bacteriol 179: 5264-70.
Hanus F, Maier RJ and Evans H (1979) Autotrophic growth of H2-uptake positive strains
of Bradyrhizobium japonicum in an atmosphere supplied with hydrogen gas. Proc Natl
Acad Sci USA 76: 1788-92.
Hausinger RP (1987) Nickel Utilization by Microorganisms. Microbiol Rev 51: 22-42.
Higuchi Y, Yagi T and Yasouka N (1997) Unusual ligand structure in Ni-Fe active
center and an additional Mg site in hydrogenase revealed by high resolution X ray
structure analysis. Structure 5: 1671-80.
Houchins JP (1984) The physiology and biochemistry of hydrogen metabolism in
cyanobacteria. Biochim Biophys Acta 768: 227-55.
Howarth DC and Codd GA (1985) The uptake and production of molecular hydrogen by
unicellular cyanobacteria. J Gen Microbiol 131: 1561-69.
Huss-Danell K and Bergman B (1990) Nitrogenase in Frankia from Root Nodules of
Alnus incana (L.) Moench: immunolocalization of the Fe- and MoFe-proteins during
vesicle differentiation. New Phytologist 116: 443-455.
53
Huss-Danell K, Uliassi D and Renberg I (1997) River and lake sediments as sources of
infective Frankia (Alnus). Plant and Soil 197(1): 35-9.
Jeong SC, Ritchie NJ and Myrold DD (1999) Molecular Phylogenies of Plants and
Frankia Support Multiple Origins of Actinorhizal Symbioses. Mol Phylogenet Evol 13
(3): 493-503.
Karube I, Matsunaga, Tsuni S and Suzuki S (1976) Continuous hydrogen production by
immobilized whole cell of Clostridium butyricum. Biochem Biophys Acta 44: 338-45.
Kennedy C and Toukdarian A (1987) Genetics of azotobacters: applications to nitrogen
fixation and related aspects of metabolism. Annu Rev Microbiol 41: 227-58.
Kimura M (1980) A simple method for estimating evolutionary rates of base substitutions
through comparative studies of nucleotide sequences. J Mol Evol (16): 111-20.
Kleihues L, Lenz O, Bernhard M, Buhrke T and Friedrich B (2000) The H2 Sensor of
Ralstonia eutropha Is a Member of the Subclass of Regulatory [NiFe] Hydrogenases. J
Bacteriol 182: 2716-24.
Kovács AT, Rákhely G, Balogh J, Maróti G, Fülöp A and Kovács KL (2005) Anaerobic
regulation of hydrogenase transcription in different bacteria. Biochem Soc Transact 33:
36-38.
Kovács KL and Rákhely G (2007) Thiocapsa roseopersicina Hydrogenases: Why There
Are So Many and What Do They Do? Lecture Abstract. The 8th International
Hydrogenase Conference, August 5-10, Breckenridge, Colorado.
Kovács KL, Seefeldt LC, Tigyi G, Doyle CM, Mortenson LE and Arp DJ (1989)
Immunological relationship among hydrogenases. J Bacteriol 171: 430-35.
54
Laurinavichene TV, Zorin NA, Tsygankov AA (2002) Effect of redox potential on
activity of hydrogenase 1 and hydrogenase 2 in Escherichia coli. Arch Microbiol 178(6):
437-42.
Lalonde M, Calvert HE and Pine S (1981) Isolation and use of Frankia strains in
actinorhizae formation, p. 296-99. In Gibson AH and Newton WE (eds), Current
Perspectives in Nitrogen Fixation. Australian Academy of Science, Canberra.
Lalonde M, Simon L, Bousquet J and Séguin A (1988) Advances in the taxonomy of
Frankia: recognition of species alni and elaeagni and novel subspecies pommerii and
vandijkii, p. 671-80. In Bothe H, Bruijn FJ and Newton WE (eds), Nitrogen Fixation:
Hundred Years After. Fischer, Stuttgart.
Lindmark DG and Muller M (1973) Hydrogenosome, a cytoplasmic organelle of the
anaerobic flagellate, Trichomonas foetus, and its role in pyruvate metabolism. J Biol
Chem 248: 7724-28.
Lumini E, Bosco M and Fernandez MP (1996) PCR-RFLP and total DNA homology
revealed three related genomic species among broad-host-range Frankia strains. FEMS
Microbiol Ecol 21(4): 303-11.
Lyon EJ, Shima S, Buurman G, Chowdhuri S, Batschauer A, Steinbach K, Thauer RK
(2004) UV-A/blue-light inactivation of the ‘metal-free’ hydrogenase (Hmd) from
methanogenic archaea. Eur J Biochem 271: 195-204.
Matias PM, Soares CM, Saraiva LM, Coelho R, Morais J, Gall JL and Carrondo MA
(2001) [NiFe] Hydrogenase from Desulfovibrio desulfuricans ATCC 27774: Gene
sequencing, three-dimensional structure determination and refinement at 1.8 Å and
modeling studies of its interaction with the tetrahaem cytochrome c3. J Biol Inorg Chem
6: 63-81.
Mattsson U (2001) Hydrogenases in Frankia. Phd thesis. ISBN 91-7191-942-2
55
Mattsson U, Johansson L, Sandström G and Sellstedt A (2001) Frankia sp. KB5
possesses a hydrogenase immunologically related to membrane-bound [NiFe]-
hydrogenases. Curr Microbiol 42: 438-41.
Mattsson U and Sellstedt A (2000) Hydrogenase in Frankia sp. KB5: expression of and
relation to nitrogenase. Can J Microbiol 46: 1091-95.
Mattsson U and Sellstedt A (2002) Nickel affects activity more than expression of
hydrogenase protein in Frankia. Curr Microbiol 44: 88-93.
Meesters TM (1987) Localization of nitrogenase in vesicles of Frankia sp. Cc1.17 by
immunogoldlabelling on ultrathin cryosections. Arch Microbiol 146: 327-31.
Melis A, Zhang L, Forestier M, Ghirardi ML and Seibert M (2000) Sustained
photobiological hydrogen gas production upon reversible inactivation of oxygen
evolution in the green alga Chlamydomonas reinhardtii. Plant Physiology 122(1): 127-
36.
Mertens R and Liese A (2004) Biotechnological applications of hydrogenases. Curr
Opin Biotechnol 15(4): 343-48.
Montet Y, Amara P, Volbeda A, Vernede X, Hatchikian EC, Field MJ, Frey M and
FontecillaCamps JC (1997) Gas access to the active site of Ni-Fe hydrogenases probed
by Xray crystallography and molecular dynamics. Nature Struct Biol 4: 523-26.
Monz CA and Schwintzer CR (1989) The physiology of spore-negative and spore-
positive nodules of Myrica gale. Plant and Soil 118: 75-87.
Nazaret S, Cournoyer B, Normand P, and Simonet P (1991) Phylogenetic relationships
among Frankia genomic species determined by use of amplified 16S rDNA sequences. J
Bacteriol 173: 4072-78.
56
Nicolet Y, Cavazza C and Fontecilla-Camps JC (2002) Fe-only hydrogenases: structure,
function and evolution. J Inorg Biochem 91: 1-8.
Nicolet Y, Piras C, Legrand P, Hatchikian CE and Fontecilla-Camps JC (1999)
Desulfovibrio desulfuricans iron hydrogenase: the structure shows unusual coordination
to an active site Fe binuclear center. Structure Fold Des 7(1): 13-23.
Normand P and Lalonde M (1982) Evaluation of Frankia strains isolated from
provenances of two Alnus species. Can J Microbiol 28: 1133-42.
Normand P, Lapierre P, Tisa LS, Gogarten JP, Alloisio N, Bagnarol E, Bassi CA, Berry
AM, Bickhart DM, Choisne N, Couloux A, Cournoyer B, Cruveiller S, Daubin V,
Demange N, Francino MP, Goltsman E, Huang Y, Kopp OR, Labarre L, Lapidus A,
Lavire C, Marechal J, Martinez M, Mastronunzio JE, Mullin BC, Niemann J, Pujic P,
Rawnsley T, Rouy Z, Schenowitz C, Sellstedt A, Tavares F, Tomkins JP, Vallenet D,
Valverde C, Wall LG, Wang Y, Medigue C and Benson DR. (2007a) Genome
characteristics of facultatively symbiotic Frankia sp. strains reflect host range and host
plant biogeography. Genome Res 17: 7-15.
Normand P, Orso S, Cournoyer B, Jeannin P, Chapelon C, Dawson J, Evtushenko L and
Misra AK (1996) Molecular Phylogeny of the Genus Frankia and Related Genera and
Emendation of Family Frankiaceae. Int J Syst Bacteriol 46: 1-9.
Normand P, Queiroux C, Tisa LS, Benson DR, Rouy Z, Cruveiller S and Médigue C
(2007b) Exploring the genomes of Frankia. Physiol Plantarum 130: 331-43.
Olson JW, Fu C and Maier RJ (1997) The HypB protein from Bradyrhizobium
japonicum can store nickel and is required for the nickel-dependent transcriptional
regulation of hydrogenase. Mol Microbiol 24: 119-28.
57
Olson JW and Maier RJ (2000) Dual roles of Bradyrhizobium japonicum nickelin
protein in nickel storage and GTP-dependent Ni mobilization. J Bacteriol 182: 1702-05.
Page RDM (1996) TREE VIEW: an application to display phylogenetic trees on
personal computers. Comput Appl Biosci 12: 357-58.
Parsons R, Silvester WB, Harris S, Gruijters WTM and Bullivant S (1987) Frankia
Vesicles Provide Inducible and Absolute Oxygen Protection for Nitrogenase. Plant
Physiol 83(4): 728-31.
Peters JW, Lanzilotta WN, Lemon BJ and Seefeldt LC (1998) The X-ray Crystal
Structure of the Fe Hydrogenase (CpI) from Clostridium pasteurianum to 1.8 A
Resolution. Science 282: 1853-58.
Pfaffi MW, Horgan GW and Dempfle L (2002) Relative expression software tool
(REST©) for group-wise comparison and statistical analysis of relative expression results
in real-time PCR. Nucleic Acids Research 30(9): e36.
Przybyla AE, Robbins J, Menon N and Peck HD Jr (1992) Structure-function
relationships among the nickel-containing hydrogenases. FEMS Microbiol Rev 88: 109-
36.
Rey L, Imperial J, Palacios J-M, and Ruiz-Argüeso T (1994) Purification of Rhizobium
leguminosarum HypB, a Nickel-Binding Protein Required for Hydrogenase Synthesis. J
Bacteriol 176: 6066-73.
Robson R (2001) Biodiversity of hydrogenases, p. 73-92. In Cammack R and Florence
KY (eds), Hydrogen as fuel: Learning from nature. London: Taylor & Francis.
Saitou N and Nei M (1987) The neighbor-joining method: a new method for
reconstructing phylogenetic trees. Mol Biol Evol 4: 406-25.
58
Sayed WF, Wheeler CT, Zahran HH and Shoreit AAM (1997) The effect of temperature
and soil moisture on the survival and symbiotic effectivity of Frankia. Biol Fertil Soils
25: 349-53.
Schmitz O, Boison G, Salzmann H, Bothe H, Schütz K, Wang S-h and Happe T (2002)
HoxE-a subunit specific for the pentameric bidirectional hydrogenase complex
(HoxEFUYH) of cyanobacteria. Biochim Biophys Acta 1554: 66-74.
Schubert KR and Evans HJ (1976) Hydrogen evolution: a major factor affecting the
efficiency of nitrogen fixation in nodulated symbionts. Proc Natl Acad Sci USA 73:
1207-11.
Schwencke J (2001) Advances in Actinorhizal Symbiosis: Host Plant-Frankia
Interactions, Biology, and Applications in Arid Land Reclamation. Arid Land Res
Manag 15: 285-327.
Schütz K, Happe T, Troshina O, Lindblad P, Leitão E, Oliveira P and Tamagnini P
(2004) Cyanobacterial H2 production - a comparative analysis. Planta 218: 350-59.
Sellstedt A (1989) Occurrence and activity of hydrogenase in symbiotic Frankia from
field-collected Alnus incana. Physiol Plantarum 75: 304-8.
Sellstedt A, Huss-Danell K and Ahlqvist A-S (1986) Nitrogen fixation and biomass
production in symbiosis between Alnus incana and Frankia strains with different
hydrogenase metabolism. Physiol Plantarum 66: 99-107.
Sellstedt A and Lindblad P (1990) Activities, Occurrence, and Localization of
Hydrogenase in Free-Living and Symbiotic Frankia. Plant Physiol 92: 809-15.
59
Sellstedt A, Reddell P and Rosbrook P (1991) The Occurrence of Hemoglobin and
Hydrogenase in Nodules of 12 Casuarina-Frankia Symbiotic Associations. Physiol
Plantarum 82: 458-64.
Sellstedt A, Rosbrook PA, Kang L and Reddell P (1994) Effect of Carbon Source on
Growth, Nitrogenase and Uptake Hydrogenase Activities of Frankia Isolates from
Casuarina sp. Plant and Soil 158: 63-8.
Sellstedt A and Smith GD (1990) Nickel Is Essential for Active Hydrogenase in Free-
Living Frankia Isolated from Casuarina. FEMS Microbiol Lett 70: 137-40.
Sellstedt A, Wullings B, Nyström U and Gustafsson P (1992) Identifcation of
Casuarina-Frankia strains by use of polymerase chain reaction (PCR) with arbitrary
primers. FEMS Microbiol Lett 93: 1-6.
Silvester WB (1976) Ecological and economic significance of the non-legume
symbiosis, p. 489-506. In W. E. Newton and C. J. Nyman (eds), Proceedings of the First
International Symposium on Nitrogen Fixation. Washington University Press, Pullman.
Silvester WB (1977) Dinitrogen fixation by plant associations excluding legumes. In
Hardy R and Silvester W (eds), A treatise of dinitrogen fixation, Section IV: Agronomy
and Ecology, p. 141-90, John Wiley and Sons, New York.
Tamagnini P, Axelsson R, Lindberg P, Oxelfelt F, Wünschiers R and Lindblad P (2002)
Hydrogenases and hydrogen metabolism of cyanobacteria. Microbiol Mol Biol Rev
66(1): 1-20.
Tamagnini P, Troshina O, Oxelfelt F, Salema R and Lindblad P (1997) Hydrogenases in
Nostoc PCC 73102, a strain lacking a bidirectional enzyme. Appl Environ Microbiol 63:
1801-07.
60
Tard C, Liu X, Ibrahim SK, Bruschi M, Gioia LD, Davies SC, Yang X, Wang L-S,
Sawers G and Pickett CJ (2005) Synthesis of the H-cluster framework of iron-only
hydrogenase. Nature 433: 610-13.
Thauer RK, Klein AR and Hartmann GC (1996) Reactions with Molecular Hydrogen in
Microorganisms: Evidence for a Purely Organic Hydrogenation Catalyst. Chem Rev 96:
3031-42.
Thompson JD, Gibson TJ, Plewniak F, Jeanmougin F and Higgins DG (1997) The
CLUSTAL X windows interface: fexible strategies for multiple sequence alignment
aided by quality analysis tools. Nucleic Acids Res 25: 4876-82.
Torres V, Ballesteros A and Fernandez VM (1986) Expression of hydrogenase activity
in Barley (Hordeum vulgare L.) after anaerobic stress. Arch Biochem Biophys 245: 174-
78.
Troshina O, Serebryakova LT, Sheremetieva ME and Lindblad P (2002) Production of
H2 by the unicellular cyanobacterium Gloeocapsa alpicola CALU 743 during
fermentation. Int J Hydrogen Energy 27: 1283-9.
Ruiz-Argüeso T, Imperial J and Palacios JM (2000) Uptake hydrogenases in root nodule
bacteria. In Triplett EW (ed), Prokaryotic nitrogen fixation: A model system for the
analysis of a biological process. Madison, WI: Horizon Scientific Press.
Van Soom C, Verreth C, Sampaio MJ and Vanderleyden J (1993) Identification of a
potential transcriptional regulator of the hydrogenase activity in free-living
Bradyrhizobium japonicum strains. Mol Gen Genet 239: 235-40.
Velázquez E, Cervantes E, Igual J M, Peix A, Mateos PF, Benamar S, Moiroud
A,Wheeler CT, Dawson JO, Labeda D, Rodríguez-Barrueco C and Martínez-Molina E
61
(1998) Analysis of LMW RNA profiles of Frankia strains by staircase Electrophoresis.
Syst Appl Microbiol 21: 539-45.
Verhagen M-F, O'Rourke T and Adams MWW (1999) The hyperthermophilic
bacterium, Thermotoga maritima, contains an unusually complex iron-hydrogenase:
amino acid sequence analyses versus biochemical Characterization. Biochim Biophys
Acta 1412: 212-29.
Vignais P M, Billoud B and Meyer J (2001) Classification and phylogeny of
hydrogenases. FEMS Microbiol Rev 25: 455-501.
Vignais PM and Colbeau A (2004) Molecular Biology of Microbial Hydrogenases. Curr
Issues Mol Biol 6: 159-88.
Vignais PM and Toussaint B (1994) Molecular biology of membrane-bound H2 uptake
hydrogenases. Arch Microbiol 161: 1-10.
Volbeda A, Charon MH, Piras C, Hatchikian EC, Frey M and Fontecilla-Camps JC
(1995) Crystal structure of the nickel-iron hydrogenase from Desulfovibrio gigas. Nature
373: 580-87.
Voordouw G (1992) Evolution of hydrogenase genes. Adv Inorg Chem 38: 397-422.
Wall LG (2000) The actinorhizal symbiosis. J Plant Growth Regul 19: 167-82.
Wheeler CT, McEwan NR, Sellstedt A and Sandström G (1998) Application of
molecular techniques to ecological studies of symbioses in actinorhizal plants, p. 41-63.
In Warma A (ed), Mycorrhiza Manual, Springer-Verlag, Berlin.
Wilm M, Shevchenko A, Houthaeve T, Breit S, Schweigerer L, Fotsis T and Mann M
(1996) Femtomole sequencing of proteins from polyacrylamide gels by nano-
electrospray mass spectrometry. Nature 379: 466-9.
62
Wu L-F and Mandrand MA (1993) Microbial hydrogenases: primary structure,
classification, signatures and phylogeny. FEMS Microbiol Rev 10: 243-70.
Zhang Z, Lopez MF and Torrey JG (1984) A comparison of cultural characteristics and
infectivity of Frankia isolates from root nodules of Casuarina species. Plant and Soil
78: 79-90.
63
Top Related