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8.14E: Psychrophilic Crenarchaeota - Biology

8.14E: Psychrophilic Crenarchaeota - Biology



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Psychrophiles crenarchaeotes are extremophilic organisms that are capable of growth and reproduction in cold temperatures.

Learning Objectives

  • Discuss the specific characteristics associated with psychrophilic crenarchaeotes

Key Points

  • Psychrophiles are characterized by lipid cell membranes chemically resistant to the stiffening caused by extreme cold, and often create protein ‘antifreezes’ to keep their internal space liquid and protect their DNA even in temperatures below water’s freezing point.
  • Crenarchaea are thought to be very abundant and one of the main contributors to the fixation of carbon.
  • Crenarchaeote are abundant in the ocean and some species have a 200 times greater affinity for ammonia than ammonia oxidizing bacteria, leading researchers to challenge the previous belief that ammonia oxidizing bacteria are primarily responsible for nitrification in the ocean.

Key Terms

  • crenarchaeota: Archae that have been recently identified to be present in marine environments where they responsible for nitrification.
  • psychrophile: An organism that can live and thrive at temperatures much lower than normal; a form of extremophile.

Psychrophiles or cryophiles (adj. cryophilic) are extremophilic organisms that are capable of growth and reproduction in cold temperatures, ranging from −15°C to +10°C. Temperatures as low as −15°C are found in pockets of very salty water (brine) surrounded by sea ice. They can be contrasted with thermophiles, which thrive at unusually hot temperatures. The environments they inhabit are ubiquitous on Earth, as a large fraction of our planetary surface experiences temperatures lower than 15°C. They are present in alpine and arctic soils, high-latitude and deep ocean waters, polar ice, glaciers, and snowfields. Most psychrophiles are bacteria or archaea, and psychrophily is present in widely diverse microbial lineages within those broad groups. Psychrophiles are characterized by lipid cell membranes chemically resistant to the stiffening caused by extreme cold, and often create protein ‘antifreezes’ to keep their internal space liquid and protect their DNA even in temperatures below water’s freezing point.

The Crenarchaeota (Greek for “spring old quality”) (also known as Crenarchaea or eocytes) are Archaea that have been classified as either a phylum of the Archaea kingdom or a kingdom of its own. Initially, the Crenarchaeota were thought to be sulfur-dependent extremophiles but recent studies have identified characteristic Crenarchaeota environmental rRNA indicating the organism may be the most abundant archaea in the marine environment. Originally, they were separated from the other archaea based on rRNA sequences. However, other physiological features, such as lack of histones have supported this division, although some crenarchaea were found to have histones. Until recently all cultured Crenarchaea had been thermophilic or hyperthermophilic organisms, some of which have the ability to grow at up to 113 °C. These organisms stain Gram negative and are morphologically diverse having rod, cocci, filamentous and oddly shaped cells. Beginning in 1992, data were published that reported sequences of genes belonging to the Crenarchaea in marine environments making these bacteria psychrophiles or cryophiles. Since then, analysis of the abundant lipids from the membranes of Crenarchaea taken from the open ocean have been used to determine the concentration of these “low temperature Crenarchaea.” Based on these measurements of their signature lipids, Crenarchaea are thought to be very abundant and one of the main contributors to the fixation of carbon. DNA sequences from Crenarchaea have also been found in soil and freshwater environments, suggesting that this phylum is ubiquitous to most environments.

Nitrification, as stated above, is formally a two-step process; in the first step ammonia is oxidized to nitrite, and in the second step nitrite is oxidized to nitrate. Different microbes are responsible for each step in the marine environment. Several groups of ammonia oxidizing bacteria (AOB) are known in the marine environment, including Nitrosomonas, Nitrospira, and Nitrosococcus. All contain the functional gene ammonia monooxygenase (AMO) which, as its name implies, is responsible for the oxidation of ammonia. More recent metagenomic studies have revealed that some Crenarchaeote Archaea possess AMO. Crenarchaeote are abundant in the ocean and some species have a 200 times greater affinity for ammonia than AOB, leading researchers to challenge the previous belief that AOB are primarily responsible for nitrification in the ocean.


Psychrophiles

Richard Y. Morita , Craig L. Moyer , in Encyclopedia of Biodiversity (Second Edition) , 2001

Biodiversity of Psychrophiles

The first known species of psychrophiles described taxonomically are Vibrio (Moritella gen. nov.) marinus MP-1 and Vibrio (Colwellia gen. nov.) psychroerythrus, both isolated in 1964. The biodiversity among psychrophiles in the various cold environments has yet to be studied extensively. Nevertheless, the various species within the genera Achromobacteria, Alcaligenes, Altermonas, Aquaspirillum, Arthrobacter, Bacillus, Bacteroides, Brevibacterium, Clostridium, Colwellia, Cytophaga, Flavobacterium, Gelidibacter, Methanococcoides, Methanogenium, Methanosarcina, Microbacterium, Micrococcus, Moritella, Octadecabacter, Phormidium, Photobacterium, Polaribacter, Polaromonas, Pseudomonas, Psychroserpens, Shewanella, and Vibrio have been found to be psychrophilic. Even a psychrophilic methanotroph (resembling Methylococcus) has been isolated from the tundra. Both the domains Bacteria and the Archaea are represented in the various genera listed. To date, only the genus Moritella appears to be composed of psychrophiles only. However, it should be noted that many species of reported psychrophiles do not meet the definition given in this article. Because of this situation, the cardinal temperatures of many psychrophiles need to be determined.

All psychrophilic and barophilic bacteria that have been cultivated belong to the subdivision γ-Proteobacteria: Shewanella, Photobacterium, Colwellia, Moritella, and a new group designate containing strain CNPT3 and Alteromonas haloplanktis. The indication is that the combined barophilic and psychrophilic phenotype evolved independently in the different γ-Proteobacteria genera ( DeLong et al., 1997 ).

Several psychrophilic, gas vacuolate bacteria have been isolated from the sea ice and water from both the Antarctic and the Arctic. These include representatives belonging to the α-, β-, and γ-Proteobacteria subdivisions as well as the Cytophaga–Flavobacterium–Bacteroides phylogenetic group ( Gosink et al., 1998 ). These results demonstrate a wide range of phyolgenetic diversity capable of psychrophily across the domain Bacteria.

Incredibly, as much as 30% of the marine picoplankton (planktonic organisms with an average diameter of 0.2–2.0 μm) from both polar and temperate coastal waters are also Archaea (i.e., archaeoplankton) and the majority of these are associated with the Crenarchaeota. Despite this cosmopolitan distribution in the world's oceans, little is known about the physiological properties of these archaeoplankton, with the exception that they are hypothesized to potentially be psychrophilic based on the marine sponge symbiont Cenarchaeum symbiosum ( Preston et al., 1996 ). However, similar archaeal phylotypes have been located at a deep-sea hydrothermal vent system ( Moyer et al., 1998 ) in waters with an environmental temperature of 15–30 °C. Many hyperthermophiles are members of the Archaea that can utilize H2 as an energy source, and recently Methanogenium frigidum, a psychrophilic, slightly halophilic, H2-using methanogen, was isolated from the perennially cold, anoxic hypolimnion of Ace Lake, Antarctica ( Franzmann et al., 1997 ). Although it may be too early to state that this may be an indiction of the evolutionary processes affecting both hyperthermophiles and psychrophiles, this does demonstrate that members of the domain Archaea are also capable of the psychrophilic lifestyle and that further research is needed. The main deterrent is the lack of isolation of psychrophiles, mainly because interest in this thermal group was lacking for many years. Yet to be isolated from the meltwater in the Antarctic are psychrophilic bacteria that are also alkaliphiles. Each body of water in the Antarctic has its own cations and anions and the salinity may be very high due to freezing of the water.

In any cold environment in which microbes have been isolated, many of the isolates are psychrotrophs. When all things are equal, the psychrophiles will outgrow the psychrotrophs at low temperature.


Examples of extremophile in the following topics:

Thermoplasmatales, Thermocaccales, and Methanopyrus

  • There are many classes in the phylum Euryarchaeota, many of which are extremophiles.
  • There are many classes in the phylum Euryarchaeota, many of which are extremophiles, surviving in extreme conditions that are uninhabitable for most other organisms.
  • A thermophile is an extremophile that thrives at relatively high temperatures, between 45 and 122 °C.

Extremophiles and Biofilms

  • Other bacteria and archaea are adapted to grow under extreme conditions and are called extremophiles, meaning "lovers of extremes."
  • Because they have specialized adaptations that allow them to live in extreme conditions, many extremophiles cannot survive in moderate environments.
  • There are many different groups of extremophiles.
  • Other extremophiles, like radioresistant organisms, do not prefer an extreme environment (in this case, one with high levels of radiation), but have adapted to survive in it.
  • Discuss the distinguishing features of extremophiles and the environments that produce biofilms

Psychrophilic Crenarchaeota

  • Psychrophiles crenarchaeotes are extremophilic organisms that are capable of growth and reproduction in cold temperatures.
  • Psychrophiles or cryophiles (adj. cryophilic) are extremophilic organisms that are capable of growth and reproduction in cold temperatures, ranging from −15°C to +10°C.
  • Initially, the Crenarchaeota were thought to be sulfur-dependent extremophiles but recent studies have identified characteristic Crenarchaeota environmental rRNA indicating the organism may be the most abundant archaea in the marine environment.

Industrial Microorganisms

  • Those suriving in the most hostile and extreme settings are known as extremophile archaea.
  • The isolation and identification of various types of Archaea, particularly the extremophile archaea, have allowed for analysis of their metabolic processes, which have then been manipulated and utilized for industrial purposes.
  • Extremophile archaea species are of particular interest due to the enzymes and molecules they produce that allow them to sustain life in extreme climates, including very high or low temperatures, extremely acid or base solutions, or when exposed to other harmful factors, including radiation.

Extremely Halophilic Archaea

  • Halophiles are extremophiles that thrive in environments with very high concentrations of salt.
  • Halophiles are extremophiles that thrive in environments with very high concentrations of salt.

Nongenetic Categories for Medicine and Ecology

  • An extremophile is an organism that thrives in physically or geochemically extreme conditions, detrimental to most life on Earth.
  • Most known extremophiles are microbes.
  • There are many different classes of extremophiles, each corresponding to the way its environmental niche differs from mesophilic conditions.
  • Many extremophiles fall under multiple categories and are termed polyextremophiles.
  • Some examples of types of extremophiles:

Oligotrophs

Microbes and the Origin of Life on Earth

Ocean Floor

  • Ocean floor extremophile chemosynthetic microbes provide energy and carbon to the other organisms in these environments.

Growth Rate and Temperature

  • Organisms that prefer extreme environments are known as extremophiles: those that prefer cold environments are termed psychrophilic, those preferring warmer temperatures are termed thermophilic or thermotrophs and those thriving in extremely hot environments are hyperthermophilic.
  • These colorful microorganisms are called extremophiles—these in particular are thermophiles.
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Psychrophilic Crenarchaeota

Psychrophiles crenarchaeotes are extremophilic organisms that are capable of growth and reproduction in cold temperatures.

Learning Objective

Discuss the specific characteristics associated with psychrophilic crenarchaeotes

Key Points

      are characterized by lipid cell membranes chemically resistant to the stiffening caused by extreme cold, and often create protein 'antifreezes' to keep their internal space liquid and protect their DNA even in temperatures below water's freezing point.
    • Crenarchaea are thought to be very abundant and one of the main contributors to the fixation of carbon.
    • Crenarchaeote are abundant in the ocean and some species have a 200 times greater affinity for ammonia than ammonia oxidizing bacteria, leading researchers to challenge the previous belief that ammonia oxidizing bacteria are primarily responsible for nitrification in the ocean.

    Terms

    Archae that have been recently identified to be present in marine environments where they responsible for nitrification.

    An organism that can live and thrive at temperatures much lower than normal a form of extremophile.

    Full Text

    Psychrophiles or cryophiles (adj. cryophilic) are extremophilic organisms that are capable of growth and reproduction in cold temperatures, ranging from −15°C to +10°C. Temperatures as low as −15°C are found in pockets of very salty water (brine) surrounded by sea ice. They can be contrasted with thermophiles, which thrive at unusually hot temperatures. The environments they inhabit are ubiquitous on Earth, as a large fraction of our planetary surface experiences temperatures lower than 15°C. They are present in alpine and arctic soils, high-latitude and deep ocean waters, polar ice, glaciers, and snowfields. Most psychrophiles are bacteria or archaea, and psychrophily is present in widely diverse microbial lineages within those broad groups. Psychrophiles are characterized by lipid cell membranes chemically resistant to the stiffening caused by extreme cold, and often create protein 'antifreezes' to keep their internal space liquid and protect their DNA even in temperatures below water's freezing point.

    The Crenarchaeota (Greek for "spring old quality") (also known as Crenarchaea or eocytes) are Archaea that have been classified as either a phylum of the Archaea kingdom or a kingdom of its own. Initially, the Crenarchaeota were thought to be sulfur-dependent extremophiles but recent studies have identified characteristic Crenarchaeota environmental rRNA indicating the organism may be the most abundant archaea in the marine environment. Originally, they were separated from the other archaea based on rRNA sequences. However, other physiological features, such as lack of histones have supported this division, although some crenarchaea were found to have histones. Until recently all cultured Crenarchaea had been thermophilic or hyperthermophilic organisms, some of which have the ability to grow at up to 113 °C. These organisms stain Gram negative and are morphologically diverse having rod, cocci, filamentous and oddly shaped cells. Beginning in 1992, data were published that reported sequences of genes belonging to the Crenarchaea in marine environments making these bacteria psychrophiles or cryophiles. Since then, analysis of the abundant lipids from the membranes of Crenarchaea taken from the open ocean have been used to determine the concentration of these "low temperature Crenarchaea." Based on these measurements of their signature lipids, Crenarchaea are thought to be very abundant and one of the main contributors to the fixation of carbon. DNA sequences from Crenarchaea have also been found in soil and freshwater environments, suggesting that this phylum is ubiquitous to most environments.

    Nitrification , as stated above, is formally a two-step process in the first step ammonia is oxidized to nitrite, and in the second step nitrite is oxidized to nitrate. Different microbes are responsible for each step in the marine environment. Several groups of ammonia oxidizing bacteria (AOB) are known in the marine environment, including Nitrosomonas, Nitrospira, and Nitrosococcus. All contain the functional gene ammonia monooxygenase (AMO) which, as its name implies, is responsible for the oxidation of ammonia. More recent metagenomic studies have revealed that some Crenarchaeote Archaea possess AMO. Crenarchaeote are abundant in the ocean and some species have a 200 times greater affinity for ammonia than AOB, leading researchers to challenge the previous belief that AOB are primarily responsible for nitrification in the ocean.


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    A psychrophilic crenarchaeon inhabits a marine sponge: Cenarchaeum symbiosum gen. nov., sp. nov

    Archaea, one of the three major domains of extant life, was thought to comprise predominantly microorganisms that inhabit extreme environments, inhospitable to most Eucarya and Bacteria. However, molecular phylogenetic surveys of native microbial assemblages are beginning to indicate that the evolutionary and physiological diversity of Archaea is far greater than previously supposed. We report here the discovery and preliminary characterization of a marine archaeon that inhabits the tissues of a temperate water sponge. The association was specific, with a single crenarchaeal phylotype inhabiting a single sponge host species. To our knowledge, this partnership represents the first described symbiosis involving Crenarchaeota. The symbiotic archaeon grows well at temperatures of 10 degrees C, over 60 degrees C below the growth temperature optimum of any cultivated species of Crenarchaeota. Archaea have been generally characterized as microorganisms that inhabit relatively circumscribed niches, largely high-temperature anaerobic environments. In contrast, data from molecular phylogenetic surveys, including this report, suggest that some crenarchaeotes have diversified considerably and are found in a wide variety of lifestyles and habitats. We present here the identification and initial description of Cenarchaeum symbiosum gen. nov., sp. nov., a symbiotic archaeon closely related to other nonthermophilic crenarchaeotes that inhabit diverse marine and terrestrial environments.


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    Abstract

    Eight anaerobic granular sludges were surveyed for Crenarchaeota using rRNA gene cloning. Microbial arrangement and substrate uptake patterns were elucidated by fluorescent in situ hybridization and beta imaging. Group 1.3 Crenarchaeota represented up to 50% of Archaea and 25% of the total microbiota in five sludges. Crenarchaeota were localized in close association with methanogenic Archaea.

    Crenarchaeal phylogeny has been fundamentally changed by the isolation of novel small-subunit rRNA gene clones from mesophilic and psychrophilic habitats. Indeed, rRNA gene sequences representing nonthermophilic clades of Crenarchaeota have been reported from, for example, mesophilic soils in disparate locations (e.g., the United States [3], Japan [14], and Finland [12]) and granular biofilms from psychrophilic anaerobic bioreactors treating various wastewaters (7, 8, 9, 16).

    Notwithstanding the many reports documenting their presence in submesophilic environmental samples, the absence of nonthermophilic isolates has hindered a biochemical and physiological characterization of these organisms. Thus, the ecological roles and physiological functions of these abundant and cosmopolitan Crenarchaeota remain a mystery. While the unusual properties of thermophilic Crenarchaeota have attracted the attention of both exobiologists wishing to study microbial evolution and biotechnology companies wishing to exploit the hyperthermotolerance of crenarchaeal cellular enzymes, the widespread occurrence of nonthermophilic Crenarchaeota, particularly in engineered environments such as anaerobic digesters, means that the biotechnological significance and potential of these organisms should now be explored.

    This paper describes observations regarding the prevalence and spatial distribution of Crenarchaeota in anaerobic granules from wastewater treatment reactors. The microbial community structure was determined by using 16S rRNA clone library analysis. Fluorescent in situ hybridization (FISH) was applied in conjunction with radioactive tracer techniques and microbeta imaging to investigate substrate uptake patterns. This is the first application of this technology to anaerobic granular biofilms and represents an advancement in terms of both microbial ecology and efforts to describe the role of microorganisms involved in wastewater treatment.

    Eight anaerobic granular sludges, S1 to S8, were obtained from various full (37ଌ)- and lab (15ଌ)-scale anaerobic biological wastewater treatment plants. S1 was from a full-scale upflow anaerobic sludge bed treatment plant treating citric acid production wastewater at Archer Daniels Midland, Ringaskiddy, County Cork, Ireland S2 was from a lab-scale expanded granular sludge bed-anaerobic filter (EGSB-AF) reactor treating volatile fatty acid-based wastewater S3 and S4 were from lab-scale upflow anaerobic sludge bed reactors used to treat acetate-based and propionate/butyrate/ethanol-based wastewaters, respectively S5 and S6 were from full-scale internal circulation (IC) reactors at Archer Daniels Midland, County Cork, and Carbery Milk Products, Ballineen, County Cork, respectively and S7 and S8 were from lab-scale EGSB-AF reactors treating high- and low-strength whey-based wastewater, respectively. All lab-scale reactors had been operated for extended trials of 250 to 500 days at 12 to 15ଌ (9).

    Total genomic DNAs were extracted from all eight samples (S1 to S8) as described previously (7). Briefly, various methods were examined for the isolation of nucleic acids, the merits of which were assessed by determining the cell lysis efficiency, i.e., the best protocol for DNA collection from biomass was that which resulted in the smallest amount of unlysed cells while maintaining a high yield of good-quality DNA. Microorganisms in sludge samples before and after DNA extraction were observed according to the method of Bitton and coworkers (4) and as described in detail by Collins et al. (7). Sludge granules were initially disassociated by grinding or by sonication prior to microbial cell lysis using a chemical approach, as described by Zhou et al. (25), or a method using mechanical disruption by bead beating combined with chemical lysis. Different combinations of the above were tested, and it was found that gently crushing sludge granules with a pestle and mortar before passing the biomass, according to the manufacturer's instructions, through a soil DNA kit (MoBio Laboratories, Inc.) provided an optimal DNA yield with little shearing and provided the highest cell lysis efficiency (7). Although some Archaea possess a particularly thick and rigid outer layer named methanochondroitin (15), this DNA recovery procedure was sufficiently robust and efficient to retrieve representative DNA yields for community structure analyses.

    Archaeal 16S rRNA genes were amplified with the forward primer 21F (5′-TTCCGGTTGATCCYGCCGGA-3′ [21]) and the reverse primer 958R (5′-YCCGGCGTTGAMTCCAATT-3′ [10]) sequences were obtained from 16S rRNA gene clone libraries, and phylogenetic reconstruction was carried out as described in detail previously (7). No Crenarchaeota-like clones were detected in S2, S3, or S4, while high levels of uncultured crenarchaeotes were found in S1, S5, S6, S7, and S8 (69%, 55%, 59%, 14%, and 78% of all archaeal clones, respectively). While Crenarchaeota from sediment and soils group into several phylogenetic subclusters (11), there appears to be a well-defined phylogenetic coherence among all crenarchaeal clones recovered from this panel of anaerobic sludges (Fig. ​ (Fig.1). 1 ). Our clones fall into group 1.3 of the Crenarchaeota (11, 13), or group 1.3b proposed by Ochsenreiter et al. (17).


    Introduction

    Crenarchaeota are abundant in the world's oceans, comprising an estimated 20% of all planktonic prokaryotes [1, 2]. They are distributed over a wide depth range, spanning both euphotic and aphotic zones [3–5], and at least one species, Cenarchaeum symbiosum, has a symbiotic association with the marine sponge Axinella mexicana [6]. Given their numerical abundance and cosmopolitan nature, Crenarchaeota represent an important constituent of marine ecosystems around the globe.

    Organic geochemical biomarkers in the form of lipid biomass have proven successful for tracking Archaea in marine sediments and in the water column. Unlike the fatty acid–derived membranes of bacteria and eukarya, the major components of archaeal cell membranes are isoprenoid ether-linked glyceroldiethers or tetraethers [7, 8]. Planktonic Archaea produce glycerol dialkyl glycerol tetraethers [9–11], including one unique structure crenarchaeol [7, 12, 13] often considered a diagnostic biomarker for planktonic archaeal species. It remains unclear if this compound is unique to marine Crenarchaeota, or is found in both euryarchaeotal lineages, too. Radiocarbon analyses of 14 C in lipid biomarkers associated with marine plankton [14], and 13 C-labeled bicarbonate tracer studies [15] suggest that marine Crenarchaeota are capable of light-independent autotrophic carbon assimilation into membrane lipid biomass, an hypothesis further strengthened by recent single cell phylogenetic identification and autoradiographic verification of carbon dioxide incorporation [16]. The direct incorporation of dissolved inorganic carbon by marine Crenarchaeota may be homologous to metabolic properties of more distantly related thermophilic Crenarchaeota that utilize a modified 3-hydroxypropionate [17–21] or reductive tricarboxylic acid [22–24] cycle for autotrophic carbon assimilation.

    Assuming marine Crenarchaeota do assimilate carbon autotrophically, the energy source utilized for growth must be consistent with the surrounding chemical environment. Ammonia (NH3) produced by the decomposition of organic matter has the potential to provide both nutrient nitrogen and a source of reducing power. The seasonal distribution of marine Crenarchaeota in the oxic and ammonia-rich surface waters off Palmer Station, Antarctica [4], as well as a correlation of increasing crenarchaeal abundance with a nitrite (NO2 − ) maximum are both consistent with the hypothesis that marine Crenarchaeota are capable of ammonia oxidation [25]. Whole genome shotgun (WGS) analysis of DNA sequences derived from the Sargasso Sea (SAR) identified potential ammonia monooxygenase (amo) genes associated with presumptive archaeal contigs supporting this hypothesis [26]. Another study of fosmids derived from complex soil libraries identified amo gene sequences related to those observed in the SAR, linked to a crenarchaeal ribosomal RNA operon, and a recent PCR survey has confirmed the widespread occurance of archaeal amoA genes in marine water columns and sediments [27], reinforcing a potential and widespread role for ammonia oxidation in varied archaeal lineages [28]. Recent isolation of a marine crenarchaeote in pure culture using bicarbonate and ammonia as sole carbon and energy sources strongly supports this hypothesis [29]. However, the specific biochemical pathways mediating ammonia oxidation and carbon assimilation in this isolate remain unknown.

    C. symbiosum, a mesophilic crenarchaeal symbiont of the marine sponge Axinella mexicana [6] provides a useful system for modeling the physiology and genetics of marine Crenarchaeota. Previous studies have determined that C. symbiosum constitutes the sole archaeal phylotype associated with A. mexicana, reaching up to 65% of the total prokaryotic cell population within a given host [6]. The purity and abundance of C. symbiosum cells in host tissue has enabled the construction of fosmid DNA libraries containing its full genomic repertoire ([30, 31] and Hallam et al., unpublished data). Although C. symbiosum is a sponge symbiont and therefore at some level adapted to life within its host environment, it remains phylogenetically clustered with the planktonic Crenarchaeota [6]. Given this relationship, sequence data obtained from C. symbiosum genomic libraries can be used to query and interpret crenarchaeal sequences recovered from the planktonic environment [32, 33]. We report here analyses of unassembled C. symbiosum genomic DNA sequences, combined with environmental gene and database surveys, to determine the presence and distribution of highly conserved genes with the potential to mediate carbon assimilation and ammonia oxidation in marine Crenarchaeota.


    3 Results

    3.1 DNA extraction and detection of Archaea populations

    Image analysis revealed that the three different soil sample types, i.e. preferential water flow path soil, matrix soil, and bulk soil, all displayed the same depth-dependent changes in the Archaea HaeIII RFLP fingerprints [ 31] ( Table 2). Therefore the bulk soil samples from each depth layer, representing the actual soil type including preferential flow paths and matrix soil, were chosen for detailed cloning and sequencing analyses of archaeal SSU rDNA PCR products.

    Relative band intensities a of Archaea community RFLP fingerprints along the depth gradient

    Band label b Depth layer [cm]
    0–9 9–20 20–50 50–100
    I 1.00±0.05 1.17±0.02 0.82±0.01 0.47±0.05
    II 1.00±0.25 ND c ND c ND c
    III 1.00±0.14 1.41±0.07 1.65±0.05 1.70±0.01
    IV 1.00±0.07 0.94±0.01 1.05±0.01 1.17±0.03
    V 1.00±0.04 1.04±0.01 1.10±0.01 1.24±0.02
    Band label b Depth layer [cm]
    0–9 9–20 20–50 50–100
    I 1.00±0.05 1.17±0.02 0.82±0.01 0.47±0.05
    II 1.00±0.25 ND c ND c ND c
    III 1.00±0.14 1.41±0.07 1.65±0.05 1.70±0.01
    IV 1.00±0.07 0.94±0.01 1.05±0.01 1.17±0.03
    V 1.00±0.04 1.04±0.01 1.10±0.01 1.24±0.02

    a Intensities of bands were determined densitometrically and were expressed relative to the value determined for the respective bands detected in the surface soil layer. Standard deviations were calculated using the three soil sample types from each depth layer as replicates.

    b RFLP bands labeled ‘I’–‘V’ according to labeling indicated in Fig. 1.

    Relative band intensities a of Archaea community RFLP fingerprints along the depth gradient

    Band label b Depth layer [cm]
    0–9 9–20 20–50 50–100
    I 1.00±0.05 1.17±0.02 0.82±0.01 0.47±0.05
    II 1.00±0.25 ND c ND c ND c
    III 1.00±0.14 1.41±0.07 1.65±0.05 1.70±0.01
    IV 1.00±0.07 0.94±0.01 1.05±0.01 1.17±0.03
    V 1.00±0.04 1.04±0.01 1.10±0.01 1.24±0.02
    Band label b Depth layer [cm]
    0–9 9–20 20–50 50–100
    I 1.00±0.05 1.17±0.02 0.82±0.01 0.47±0.05
    II 1.00±0.25 ND c ND c ND c
    III 1.00±0.14 1.41±0.07 1.65±0.05 1.70±0.01
    IV 1.00±0.07 0.94±0.01 1.05±0.01 1.17±0.03
    V 1.00±0.04 1.04±0.01 1.10±0.01 1.24±0.02

    a Intensities of bands were determined densitometrically and were expressed relative to the value determined for the respective bands detected in the surface soil layer. Standard deviations were calculated using the three soil sample types from each depth layer as replicates.

    b RFLP bands labeled ‘I’–‘V’ according to labeling indicated in Fig. 1.

    The quantities of DNA extracted from fresh bulk soil markedly decreased with increasing depth in the profile. In the surface soil layer (0–9 cm), DNA quantity was highest with 30 μg g −1 . Between 9 and 20 cm, 24 μg g −1 DNA was extracted. DNA quantity dropped to 13 μg g −1 in the soil layer between 20 and 50 cm and was lowest in the bottom soil layer (50–100 cm) with 3 μg g −1 . PCR amplification of target rRNA gene fragments was performed on the same quantity (2 ng) of bulk soil DNA to allow for a direct comparison of relative band intensities in the HaeIII RFLP fingerprints ( Fig. 1). High-resolution agarose gel analysis resolved five prominent bands within the Archaea community HaeIII RFLP fingerprints ( Fig. 1, bands I–V). Densitometric quantification and statistical analysis of band intensities from bulk soil, preferential water flow path soil, and matrix soil revealed significant changes in the HaeIII RFLP fingerprints between the four depth layers ([ 31] Table 2). Intensity of Band I decreased from the surface soil layer (0–9 cm) to the bottom soil layer (50–100 cm) to 47% (**P<0.01). Band III displayed the opposite trend as it increased in intensity from the surface to deeper soil layers with a maximum of 170% in the bottom relative to the surface soil layer (*P<0.05). Band II was only detected in the surface, but not in deeper soil layers. Band IV revealed no significant changes along the entire depth profile, whereas band V showed a slight increase to 124% intensity relative to the surface soil layer (*P<0.05). No significant differences were observed when comparing total lane intensities between the four depth layers.

    HaeIII RFLP patterns of archaeal 16S rDNA fragments amplified from bulk soil DNA extracts from depth layers 0–9, 9–20, 20–50 and 50–100 cm of a Swiss forest soil. MW: 1 kb molecular mass marker (Promega) (−): negative control. The arrowhead indicates the migration position of the undigested Archaea PCR-product at approximately 500 bp.

    HaeIII RFLP patterns of archaeal 16S rDNA fragments amplified from bulk soil DNA extracts from depth layers 0–9, 9–20, 20–50 and 50–100 cm of a Swiss forest soil. MW: 1 kb molecular mass marker (Promega) (−): negative control. The arrowhead indicates the migration position of the undigested Archaea PCR-product at approximately 500 bp.

    3.2 Characterization of changing Archaea populations

    In order to gain detailed information on the Archaea populations represented by the changing HaeIII RFLP fingerprints in the different soil layers, gene libraries of Archaea SSU rDNA amplified from the surface soil layer (0–9 cm) and the bottom soil layer (50–100 cm) were constructed. The two libraries were screened by use of Archaea-specific PCR and HaeIII RFLP analysis performed on single clones. Among the 104 clones screened (39 from the surface soil library and 65 from the bottom soil library), eight different HaeIII RFLP patterns were observed ( Fig. 2). The abundance of each pattern was quantified in both gene libraries, which allowed for a numeric comparison of the relative clone compositions of the top and the bottom soil libraries ( Table 3). Richness estimations indicated that from the surface soil library 91% (5 of 5.5) and from the bottom soil library 89% (5 of 5.6) of the patterns were recovered (data not shown). HaeIII RFLP pattern ‘a’ was 1.4-fold more abundant in the bottom soil library while patterns ‘b’, ‘c’ and ‘e’ were exclusively found in the surface soil library. Pattern ‘d’ was 7.7-fold more abundant in the bottom soil library. Patterns ‘f’–‘h’ were infrequent and restricted to the bottom soil library. Further analysis revealed that each of the five prominent bands observed in the complex Archaea community RFLP fingerprints ( Fig. 1) was also identified in the RFLP fingerprint of at least one clone ( Fig. 2 and Table 4). Band I was exclusively found in pattern ‘b’ while band II was detected in pattern ‘e’ only. Band III was mainly attributed to pattern ‘d’ with a minor contribution of pattern ‘f’, while band IV mainly originated from patterns ‘a’ and ‘c’ with minor contributions of patterns ‘g’ and ‘h’. Band V was composed by patterns ‘a’–‘e’ with minor contributions of patterns ‘f’ and ‘g’. The R 2 -value of the linear correlation between band intensities in the community HaeIII RFLP fingerprints and the band occurrences in the gene libraries was 0.54. These analyses revealed that each of the five HaeIII RFLP bands displayed the same trend of abundance in the two clone libraries as indicated by the band intensities of the Archaea community HaeIII RFLP fingerprints.

    Calculated HaeIII RFLP patterns that occurred among 104 cloned archaeal 16S rDNA fragments from surface (0–9 cm) and bottom (50–100 cm) soil layers from a Swiss forest soil. Fragment sizes were inferred from DNA sequences of representative clones. MW: 1 kb molecular mass marker (Promega) RFLP patterns were labeled ‘a’–‘h’. Bands I–V were detected in the complex RFLP fingerprints (see also Fig. 1a and Table 3).

    Calculated HaeIII RFLP patterns that occurred among 104 cloned archaeal 16S rDNA fragments from surface (0–9 cm) and bottom (50–100 cm) soil layers from a Swiss forest soil. Fragment sizes were inferred from DNA sequences of representative clones. MW: 1 kb molecular mass marker (Promega) RFLP patterns were labeled ‘a’–‘h’. Bands I–V were detected in the complex RFLP fingerprints (see also Fig. 1a and Table 3).

    RFLP pattern frequencies in archaeal SSU rDNA fragment libraries

    RFLP pattern labels a Archaea SSU rDNA libraries
    surface soil b bottom soil b
    a c 48.7 (19) 67.8 (44)
    b 17.9 (7)
    c 23.1 (9)
    d c 2.6 (1) 20.0 (13)
    e 7.6 (3)
    f 3.1 (2)
    g 1.5 (1)
    h 1.5 (1)
    x 6.2 (4)
    Total 100.0 (39 clones) 100.0 (65 clones)
    RFLP pattern labels a Archaea SSU rDNA libraries
    surface soil b bottom soil b
    a c 48.7 (19) 67.8 (44)
    b 17.9 (7)
    c 23.1 (9)
    d c 2.6 (1) 20.0 (13)
    e 7.6 (3)
    f 3.1 (2)
    g 1.5 (1)
    h 1.5 (1)
    x 6.2 (4)
    Total 100.0 (39 clones) 100.0 (65 clones)

    a RFLP pattern labels as defined in Fig. 2 ‘x’ represents cloning artifacts.

    b RFLP pattern frequencies are presented in percent for each library. The number of clones identified is given in parentheses.

    c Sequence and phylogenetic analyses ( Fig. 3) revealed that clones from the surface and bottom soil layers, which shared the same HaeIII RFLP pattern, belonged to different phylotypes.

    RFLP pattern frequencies in archaeal SSU rDNA fragment libraries

    RFLP pattern labels a Archaea SSU rDNA libraries
    surface soil b bottom soil b
    a c 48.7 (19) 67.8 (44)
    b 17.9 (7)
    c 23.1 (9)
    d c 2.6 (1) 20.0 (13)
    e 7.6 (3)
    f 3.1 (2)
    g 1.5 (1)
    h 1.5 (1)
    x 6.2 (4)
    Total 100.0 (39 clones) 100.0 (65 clones)
    RFLP pattern labels a Archaea SSU rDNA libraries
    surface soil b bottom soil b
    a c 48.7 (19) 67.8 (44)
    b 17.9 (7)
    c 23.1 (9)
    d c 2.6 (1) 20.0 (13)
    e 7.6 (3)
    f 3.1 (2)
    g 1.5 (1)
    h 1.5 (1)
    x 6.2 (4)
    Total 100.0 (39 clones) 100.0 (65 clones)

    a RFLP pattern labels as defined in Fig. 2 ‘x’ represents cloning artifacts.

    b RFLP pattern frequencies are presented in percent for each library. The number of clones identified is given in parentheses.

    c Sequence and phylogenetic analyses ( Fig. 3) revealed that clones from the surface and bottom soil layers, which shared the same HaeIII RFLP pattern, belonged to different phylotypes.

    Assignment of individual RFLP bands in Archaea community fingerprints to specific RFLP patterns

    Band labels as defined in Fig. 1.

    Fragment sizes (in bp) of bands I–V. Sizes were determined from sequence data as shown in Fig. 2.

    RFLP pattern labels as defined in Fig. 2.

    Patterns shown in parentheses revealed a minor contribution.

    Assignment of individual RFLP bands in Archaea community fingerprints to specific RFLP patterns

    Band labels as defined in Fig. 1.

    Fragment sizes (in bp) of bands I–V. Sizes were determined from sequence data as shown in Fig. 2.

    RFLP pattern labels as defined in Fig. 2.

    Patterns shown in parentheses revealed a minor contribution.

    3.3 Phylogenetic analyses of Archaea clones

    The question of which phylotypes were represented in the Archaea community HaeIII RFLP fingerprints was addressed by sequencing Archaea SSU rDNA clones, which represented the different HaeIII RFLP types. The 20 new sequences isolated from the surface and the bottom layers of the forest soil depth profile were aligned to 118 defined control sequences derived from public databases. Only control sequences that covered the entire SSU rDNA region defined by the PCR primers used in this study were included in the analysis. The phylogeny of the aligned clone sequences was inferred by using standard distance estimation and cluster analysis routines (data not shown) as well as a maximum likelihood routine ( Fig. 3). The resulting phylogenetic trees revealed similar tree topologies and in particular identical clustering of the distinct clusters ‘A’–‘E’ indicated in Fig. 3.

    Phylogenetic tree based on maximum likelihood calculation. Each of the archaeal SSU rDNA sequences is identified by its clone name, by its source, and by its sequence accession number. Sequences isolated in the present study are printed in bold. Clone names indicate the origin of the different clones, i.e. layer (surface soil: 03 bottom soil: 12) followed by the clone number (01–58) and the HaeIII RFLP type (‘a’ to ‘h’). Cluster (I) defines the archaeal kingdom of Korarchaeota, cluster (II) the kingdom of Euryarchaeota, and cluster (III) the kingdom of Crenarchaeota. Shaded clusters ‘A’–‘E’ mark novel archaeal forest soil clusters. Numbers at condensed clusters indicate the number of sequences included. Brackets at the right margin mark phylogenetic clusters defined in the literature [ 4]. The scale bar indicates the average substitution rate per nucleotide position.

    Phylogenetic tree based on maximum likelihood calculation. Each of the archaeal SSU rDNA sequences is identified by its clone name, by its source, and by its sequence accession number. Sequences isolated in the present study are printed in bold. Clone names indicate the origin of the different clones, i.e. layer (surface soil: 03 bottom soil: 12) followed by the clone number (01–58) and the HaeIII RFLP type (‘a’ to ‘h’). Cluster (I) defines the archaeal kingdom of Korarchaeota, cluster (II) the kingdom of Euryarchaeota, and cluster (III) the kingdom of Crenarchaeota. Shaded clusters ‘A’–‘E’ mark novel archaeal forest soil clusters. Numbers at condensed clusters indicate the number of sequences included. Brackets at the right margin mark phylogenetic clusters defined in the literature [ 4]. The scale bar indicates the average substitution rate per nucleotide position.

    Of the 20 sequences that were derived from the soil depth profile described in this study, 16 grouped with Crenarchaeota and four with Euryarchaeota. The crenarchaeal sequences were associated with four distinct clusters (‘A’–‘D’). Cluster ‘A’ included clones 12 11d and 12 19d from the bottom soil layer and displayed highest similarity to two sequences from deep subsurface acidic mine water (clones SAGMA-D and SAGMA-X [ 48]) and to one sequence from groundwater (clone SRS62DAR03). Cluster ‘B’ included four sequences from the surface soil (clones 03 01a, 03 02a, 03 03a, and 03 12a) and two sequences from the bottom soil layer (clones 12 28g and 12 58h). They grouped with four clones isolated from a wetland soil in Japan (clones OS-19, OS-25, OS-31, and WSB-6). Two sequences from the bottom soil layer (clones 12 45f and 12 47f) formed cluster ‘C’, which was only weakly associated with clone pJP96 from a Yellowstone hot spring. Three sequences from the bottom soil layer (clones 12 01a, 12 02a, and 12 30a) formed cluster ‘D’, which branched closely from a group containing eight clones from the Japanese wetland soil mentioned above (clones OS-6, OS-14, OS-21, OS-22, OS-26, OS-27, AM-11, and WSB-20). Two sequences from surface soil (clones 03 06c and 03 17c) were allocated to the terrestrial cluster of uncultured Crenarchaeota (Group I.1b) in close vicinity to sequences cloned from plant roots and agricultural bulk soil (several of the TRC-clones and clone SCA1145). One sequence retrieved from surface soil (clone 03 21d) formed a separate branch without close association to any sequences reported to date. Euryarchaeal sequences derived from this study clustered exclusively with Thermoplasmales and relatives (clones 03 11b, 03 15b, 03 14e, and 03 27e) and formed a cluster termed ‘E’ within this diverse order. One sequence from Japanese wetland soil (clone OS-10) and one sequence from a hydrocarbon-contaminated aquifer (clone WCHD3–16 [ 49]) supported this cluster.


    Results

    Environmental Genomic Analysis of a Fosmid Library Enriched for C. symbiosum DNA

    C. symbiosum cells were enriched from host tissue using differential density centrifugation (see Materials and Methods). High molecular weight DNA purified from this cell enrichment was used to construct two 32–45 Kb insert fosmid DNA libraries composed of 10,236 and 2,100 clones, respectively [31]. Seven C. symbiosum SSU rRNA genes were identified in the first library, representing approximately 0.07% of the total clone population. Eight C. symbiosum SSU rRNA genes were identified in the second library, representing approximately 0.38% of the total clone population [31]. For a 2 million base pair (Mb) genome containing one copy of the SSU rRNA gene, approximately 50 fosmids arranged in a linear tiling path have the potential to cover the entire genomic sequence. Given this estimate and the percentage of C. symbiosum SSU rRNA genes identified in the second library, approximately 400 of the 2,100 clones in the library should be derived from C. symbiosum donors (̃8-fold coverage of a 2-Mb genome). Paired-end sequencing of the smaller fosmid library generated 1.8 Mb of DNA sequence from 2,779 nonredundant reads greater than 200 bp (base pair) per read. A total of 168 fosmids encompassing more than 6.4 Mb of genomic DNA were selected for subcloning and sequencing based on the following criteria: (1) paired ends predicted to contain open reading frames most similar to archaeal genes, (2) linkage with previously reported fosmids harboring crenarchaeal phylogenetic anchors, and (3) sets of paired ends, assembled in opposing orientations and predicted to contain open reading frames homologous to two or more archaeal genes. On average, selected fosmids had a G+C content of 58%. In general, the identity of completed fosmids could be verified on the combined basis of G+C content, SSU rRNA, or functional gene linkage, and the taxonomic distribution of predicted open reading frames contained on individual fosmid sequences. Based on this set of criteria, 155 of the completed fosmids were derived from C. symbiosum, ten from unspecified bacteria and three from host donors.

    Overall genomic representation of the dataset was determined based on the identification of complete or redundant sets of genes encoding ribosomal proteins, amino-acyl tRNA synthases, and conserved components of several core processes including but not limited to the transcription, translation, and replication machinery identified within the set of 155 archaeal fosmids. On average, two copies of each gene within a given category were identified (unpublished data), consistent with 2- to 3-fold coverage of an estimated 2-Mb genome. Previous studies of C. symbiosum population structure identified two coexisting ribotypic variants, a and b, exhibiting approximately 99% nucleotide identity at the ribosomal level but ranging between 70% and 90% nucleotide identity in genomic intervals adjacent to the ribosomal operon [30]. Despite this variation in nucleotide identity, gene content and order appeared to be conserved between the two ribotypes [30]. The present study focuses on pathways of autotrophic carbon assimilation and ammonia oxidation, based on analyses of individual fosmid sequences, and does not attempt to discriminate between sequences derived from a or btypes. The issue of heterogeneity and the assembly of a-type and b-type genomic scaffolds will be the subject of future work exploring the complete genome sequence of C. symbiosum (Hallam et al., unpublished data).

    Autotrophic Carbon Assimilation Genes in C. symbiosum

    In order to identify potential effectors of autotrophic metabolism in the unassembled C. symbiosum genomic sequences, systematic searches for each of the four known pathways of autotrophic CO2 fixation were conducted (see Materials and Methods). These include: (1) the unidirectional reductive pentose phosphate cycle [34], (2) the bidirectional reductive acetyl coenzyme A (CoA) pathway [35], the (3) the reductive tricarboxylic acid pathway [36], and (4) the unidirectional 3-hydroxypropionate cycle [37]. Each pathway is defined by a subset of diagnostic enzymes. The reductive pentose phosphate cycle requires the activities of ribulose 1,5-bisphosphate carboxylase, phosphoribulokinase, and sedoheptulose bisphosphatase [38]. The reductive acetyl-CoA pathway requires the activity of carbon monoxide dehydrogenase [39]. The reductive TCA (tricarboxylic acid) cycle requires the activities of citrate lyase, 2-oxoglutarate:ferredoxin oxidoreductase, and fumarate reductase [40]. Finally, the 3-hydroxypropionate cycle requires the activities of acetyl-CoA/propionyl-CoA carboxylase [37], malonyl-CoA reductase [41], and propionyl-CoA synthase [42]. Search results identified numerous components of the 3-hydroxypropionate cycle and the citric acid cycles (Tables 1, 2, and S1). Homologues for 1,5-bisphosphate carboxylase/oxygenase (RubisCO), phosphoribulokinase, sedoheptulose bisphosphatase, and carbon monoxide dehydrogenase representing the reductive pentose phosphate cycle and reductive acetyl-CoA cycle, respectively, were not identified.

    3-Hydroxypropionate Cycle Components Identified in C. symbiosum

    The 3-hydroxypropionate cycle was first identified in the phototrophic green nonsulfur bacterium Chloroflexus aurantiacus [37, 43] and more recently recognized in several thermophilic crenarchaeotes within the Sulfolobales [17, 18, 20, 44]. This pathway employs several enzymes typically associated with bacterial fatty acid biosynthesis, including the biotin-dependent enzyme acetyl-CoA/propionyl-CoA carboxylase [45]. Because archaeal lipids are typically devoid of fatty acids, the presence of acetyl-CoA/propionyl-CoA carboxylase in Crenarchaeota is necessary but not sufficient evidence for autotrophic carbon assimilation by the 3-hydroxypropionate cycle.

    Genes predicted to encode components of eight steps mediating the 3-hydroxypropionate cycle, including acetyl-CoA/propionyl-CoA carboxylase, were unambiguously identified in the C. symbiosum fosmid sequences (Tables 1 and S1 and Figure 1). Fosmids harboring gene sequences predicted to encode acetyl-CoA/propionyl-CoA carboxylase subunits could be assembled into a single contig approximately 1.29 million bp in length containing the C. symbiosum SSU-LSU ribosomal RNA operon, further reinforcing this identification scheme (Hallam et al., unpublished data). Four additional steps represented by malonyl-CoA reductase [41] and propionyl-CoA synthase [42] could not be definitively identified, in part because the specific enzymes mediating these steps remain uncharacterized throughout the archaeal domain, including those archaeal groups known to express a fully catalytic form of the 3-hydroxypropionate cycle [18, 20]. Despite this challenge, candidates for both enzymes could be tentatively assigned based on the identification of open reading frames containing conserved domains with putative pathway-related functions. In addition to core components of the 3-hydroxypropionate cycle, a biotin ligase (birA) required for assembly and activation of the carboxylase complex in vivo was also identified (Tables 1 and S1). Consistent with biochemical observations in M. sedula [18, 20], homologous genes encoding succinyl-CoA/malate transferase or L-malyl-CoA lyase, required for the regeneration of acetyl-CoA from glyoxylate, were not identified in the C. symbiosum fosmid sequences.

    Each step is mediated by the following enzymes: (1) acetyl-CoA carboxylase, (2–3) malonyl-CoA reductase, (4–6) propionyl-CoA synthase, (7) propionyl-CoA carboxylase, (8) methylmalonyl-CoA epimerase, (9) methylmalonyl-CoA mutase, (10) succinate dehydrogenase, (11) fumarase, (12) succinyl-CoA/malate CoA transferase, and (13) malyl-CoA lyase (B) TCA cycle. Each step is mediated by the following enzymes: (1) citrate synthase, (2–3) aconitase, (4) isocitrate dehydrogenase, (5) 2-oxoacid ferredoxin oxidoreductase or 2-oxoglutarate dehydrogenase, (6) succinyl-CoA synthase, (7) succinate dehydrogenase, (8) fumarase, and (9) malate dehydrogenase. In the reductive direction, the steps are reversed and citrate synthase is replaced by citrate lyase in (1). Diagrams are based on KEGG pathway maps and include enzyme classification numbers (EC) for each step in boxes when available. Box color indicates the identification status of genes encoding a particular step. See Tables 1 and 2 for more information.

    In C. aurantiacus, malonyl-CoA reductase activity is associated with a bifunctional enzyme containing alcohol dehydrogenase and aldehyde dehydrogenase domains, mediating the conversion of malonyl-CoA to 3-hydroxypropionate via malonate semialdehyde [41]. In the C. symbiosum fosmid sequences, 14 genes predicted to encode short-chain alcohol dehydrogenase domains with the potential to mediate conversion of malonyl-CoA to malonate semialdehyde were identified (Tables 1 and S1). One candidate in particular, identified on three fosmids (101G10, C03A05, and C04H09), was found to contain an N-terminal alcohol dehydrogenase domain 32% identical and 46% similar to the corresponding interval of malonyl-CoA reductase from C. aurantiacus . Adjacent to this domain, a 142–amino acid interval, 27% identical and 40% similar to an aldehyde dehydrogenase from Mus musculus, was also identified as potentially involved in the conversion of malonate semialdehyde to 3-hydroxypropionate.

    In C. aurantiacus, propionyl-CoA synthase activity is associated with a trifunctional enzyme containing CoA ligase, enoyl-CoA hydratase, and enoyl-CoA reductase domains [42]. Three copies of a gene predicted to encode enoyl-CoA hydratase, one of three enzymatic steps associated with propionyl-CoA synthase activity, were identified in the C. symbiosum fosmid sequences (Tables 1 and S1). One copy of enoyl-CoA hydratase, identified on fosmid C13E07 contained a 182–amino acid interval 27% identical and 40% similar to propionyl-CoA synthase from C. aurantiacus . Immediately upstream of this open reading frame, a second gene predicted to encode a CoA binding protein related to acyl-CoA synthase, a potential effector of the indeterminate CoA ligase step, was also identified. In C. aurantiacus the enoyl-CoA reductase domain of propionyl-CoA synthase belongs to the family of zinc-binding dehydrogenases that includes NAD(P)H-dependent crotonyl-CoA reductases [42]. Seven copies of genes predicted to encode zinc-binding alcohol dehydrogenases with potential to mediate propionyl-CoA synthase activity were identified in the C. symbiosum fosmid sequences (Tables 1 and S1). One candidate in particular, identified on three fosmids (C04F04, C05C02, and C13G08) was 29% identical and 48% similar to NAD(P)H-dependent crotonyl-CoA reductase from Silicibacter sp. TM1040. In all three instances, this gene was found in an operon with a second gene predicted to encode the beta subunit of citrate lyase (see below).

    Taken together, these observations suggest that C. symbiosum has the genetic potential to encode a modified version of the 3-hydroxypropionate cycle for CO2 incorporation into biomass. Biochemical and physiological studies remain necessary, however, to fully validate this hypothesis and determine the specific pathway of acetyl-CoA regeneration in the absence of a glyoxylate shunt.

    TCA Cycle Components Identified in C. symbiosum

    The steps of the TCA cycle define a central clearinghouse in cellular metabolism, linking the oxidative breakdown of sugars, fats, and proteins with the production of precursor molecules for biosynthesis and energy metabolism. Alternatively, the TCA cycle can be reversed in certain prokaryotic organisms, resulting in the formation of oxaloacetate from two molecules of CO2, thereby providing an alternative route for autotrophic carbon assimilation. The enzyme citrate synthase is responsible for the conversion of oxaloacetate and acetyl-CoA to citrate in the oxidative branch of the TCA cycle. The TCA cycle operating in the reductive direction requires the activity of ATP citrate lyase (also known as ATP citrate synthase) first described in Chlorobium limicola [46, 47]. ATP citrate lyase is encoded by the genes aclA and aclB, which when expressed form a heteromeric enzyme complex capable of cleaving citrate to produce oxaloacetate and acetyl-CoA [48]. An evolutionarily related alternative to ATP citrate lyase has recently been described in the hydrogen-oxidizing thermophilic bacterium Hydrogenobacter thermophilus, mediating citrate cleavage in conjunction with the enzymes citryl-CoA synthase (ccs) and citryl-CoA lyase (ccl) [49, 50].

    Genes with the potential to encode components of nine steps mediating the oxidative or reductive TCA cycles were identified in the C. symbiosum fosmid sequences (Tables 2 and S1 and Figure 1B). In addition to core TCA components, genes encoding phosphoenolpyruvate carboxykinase and phosphoenolpyruvate synthase were identified, consistent with an anapleurotic role for these enzymes in balancing carbon transfer into and out of TCA-dependent pathways (Tables 2 and S1 and Figure 1B). No homologue for pyruvate carboxylase mediating the ATP-dependent carboxylation of pyruvate to oxaloacetate was recovered from the C. symbiosum fosmid sequences. A relatively small subset of TCA components was not identified or could not be readily assigned due to high levels of similarity between close relatives with varying substrate specificities. These include subunits of citrate lyase and ferredoxin oxidoreductase described below.

    In the case of the oxidative TCA cycle, a near-complete pathway for the conversion of citrate to oxaloacetate could be reconstructed based on the identification of at least one copy of genes predicted to encode citrate synthase, aconitase, isocitrate dehydrogenase, succinyl-CoA synthase, succinate dehydrogenase, fumarase, and malate dehydrogenase (Tables 2 and S1 and Figure 1B). Step 5 (see Figure 1B) mediating the conversion of 2-oxoglutarate to succinyl-CoA could not be unequivocally determined because of the high degree of amino acid conservation between the group of ferredoxin oxidoreductases responsible for synthesis or cleavage of pyruvate, 2-isoketovalerate, or 2-oxoglutarate (Tables 2 and S1). Although four fosmids containing operons predicted to encode 2-oxoacid:ferredoxin oxidoreductase subunits were identified in the C. symbiosum sequences (C01A08, C17D04, C05B02, and C05G02), close examination of overlapping and adjacent intervals found them to be syntenic, consistent with common coverage of an equivalent genomic interval derived from separate donors ( Table S1, unpublished data). In Clostridium thermoaceticum, pyruvate:ferredoxin oxidoreductase has been shown to function in both oxidative decarboxylation of pyruvate to CO2 and acetyl-CoA, and the carboxylation of acetyl-CoA to form pyruvate [51]. It is possible that C. symbiosum, like C. thermoaceticum, encodes a multifunctional enzyme complex mediating forward and reverse reactions in one or more of the TCA-dependent steps. Functional studies remain necessary, however, to determine the specificity of this complex towards pyruvate, 2-isoketovalerate, or 2-oxoglutarate.

    In the case of the reductive TCA cycle, no homologues for aclA, aclB, ccs, or ccl were identified in the C. symbiosum fosmid sequences. However, three fosmids (C04F04, C05C02, and C13G08) containing genes predicted to encode the beta subunit of citrate lyase (citE), mediating the conversion of citryl-CoA to acetyl-CoA and oxaloacetate were identified (Tables 2 and S1). In bacteria, citE exists as part of an operon containing genes encoding citrate-CoA transferase (citF), and an acyl carrier protein subunit (citD). Bacterial homologues of citrate lyase play defined roles in citrate fermentation pathways [52, 53]. However, little is known about the functional aspects of archaeal citrate lyases. A similar case to C. symbiosum has been described in the facultative heterotrophic crenarchaeon, Thermoproteus tenax, where a single gene predicted to encode the beta subunit of citrate lyase (citE) was identified in the draft genome sequence [54]. No direct homologues for citD or citF were identified. However, the authors speculate that two genes adjacent to citE, predicted to encode acetyl-CoA synthetase and acety-CoA transferase/carnitine dehydratase have the potential to fill in for the missing citrate-CoA transferase and acyl carrier subunits, respectively [54]. At present, ATP citrate lyase activity in T. tenax remains unmeasured, although 13 C-labeling studies in a close relative, Thermoproteus neutrophilus , successfully measured incorporation patterns consistent with the operation of a reductive TCA cycle in autotrophically grown cells [22, 24]. Similar to T. tenax, no direct homologues of citF or citD were identified in the C. symbiosum fosmid sequences, although unlinked genes predicted to encode acetyl-CoA synthetases were identified on four separate fosmids (C01C01, C17E03, C07D05, and C13E07). In contrast to T. tenax, no homologues for acety-CoA transferase/carnitine dehydratase were identified.

    Taken together, the data suggests that C. symbiosum utilizes either the oxidative TCA cycle in the consumption of organic carbon and in the production of intermediates for amino acid and cofactor biosynthesis, or a horseshoe version of the TCA cycle charting an oxidative branch between citrate and 2-oxoglutarate and a reductive branch between oxaloacetate and succinate for biosynthetic purposes alone.

    A Potential Mode of Ammonia Oxidation in C. symbiosum

    Chemolithoautotrophic ammonia oxidation produces energy and reducing equivalents used for cell growth, carbon assimilation, and the generation of a proton motive force. In bacteria such as Nitrosomonas europaea and Nitrosospira multiformis, ammonia monooxygenase composed of α, β, and γ membrane-bound subunits, encoded by the genes amoA, amoB, and amoC, respectively, catalyzes the conversion of ammonia (NH3) to hydroxylamine (NH2OH). In these, NH2OH is subsequently converted to nitrite through the activity of hydroxylamine oxidoreductase, a phylogenetically unique homotetramer containing eight heme groups [55]. Among ammonia-oxidizing bacteria, up to three copies of the amo operon may be present, with conserved gene order, amoCAB [55]. Two fosmids (C07D08 and C18D02) containing genes encoding putative α, β, and γ subunits, co-located over an approximately 6-Kb interval were identified in the C. symbiosum fosmid sequences (Tables 3 and S1 and Figures 2, 4, and S1). The C. symbiosum amoA, amoB, and amoC genes were predicted to encode proteins 26% identical and 40% similar, 25% identical and 44% similar, and 50% similar and 32% identical to the corresponding α, β and γ subunits from N. multiformis, N. oceani, and N. europaea, respectively. A second unlinked gene encoding a γ subunit, 31% identical and 51% similar to the γ3 subunit from N. europaea was also identified in the C. symbiosum fosmid sequences (Tables 1 and S1 and Figure 4B). Analysis of predicted transmembrane domains affiliated with C. symbiosum ammonia monooxygenase subunits identified 6, 2, and 4 membrane spanning intervals in the α, β, and γ subunits respectively, compared with 6, 2, and 6 membrane spanning intervals for related subunits in N. europaea (see Materials and Methods).