JXB Advance Access originally published online on February 10, 2008
Journal of Experimental Botany 2008 59(5):1047-1058; doi:10.1093/jxb/ern005
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REVIEW-ARTICLE |
Cyanobacteria–bryophyte symbioses
Institute of Integrative and Comparative Biology, Faculty of Biological Sciences, Garstang Building, University of Leeds, Leeds LS2 9JT, UK
* To whom correspondence should be addressed. E-mail: d.g.adams{at}leeds.ac.uk
Received 1 August 2007; Revised 23 December 2007 Accepted 7 January 2008
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Abstract |
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 Top Abstract IntroductionThe symbiotic partnersThe infection processPost-infection changesCyanobacterial motilityHormogonia and piliConcluding remarksReferences |
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Cyanobacteria are a large group of photosynthetic prokaryotes of enormous environmental importance, being responsible for a large proportion of global CO2 and N2 fixation. They form symbiotic associations with a wide range of eukaryotic hosts including plants, fungi, sponges, and protists. The cyanobacterial symbionts are often filamentous and fix N2 in specialized cells known as heterocysts, enabling them to provide the host with fixed nitrogen and, in the case of non-photosynthetic hosts, with fixed carbon. The best studied cyanobacterial symbioses are those with plants, in which the cyanobacteria can infect the roots, stems, leaves, and, in the case of the liverworts and hornworts, the subject of this review, the thallus. The symbionts are usually Nostoc spp. that gain entry to the host by means of specialized motile filaments known as hormogonia. The host plant releases chemical signals that stimulate hormogonia formation and, by chemoattraction, guide the hormogonia to the point of entry into the plant tissue. Inside the symbiotic cavity, host signals inhibit further hormogonia formation and stimulate heterocyst development and dinitrogen fixation. The cyanobionts undergo morphological and physiological changes, including reduced growth rate and CO2 fixation, and enhanced N2 fixation, and release to the plant much of the dinitrogen fixed. This short review summarizes knowledge of the cyanobacterial symbioses with liverworts and hornworts, with particular emphasis on the importance of pili and gliding motility for the symbiotic competence of hormogonia.
Key words: Bryophyte, cyanobacteria, gliding motility, pili, symbiosis
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Introduction |
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TopAbstractIntroductionThe symbiotic partnersThe infection processPost-infection changesCyanobacterial motilityHormogonia and piliConcluding remarksReferences |
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Cyanobacteria form a wide variety of symbiotic associations with eukaryotic hosts including plants, fungi, sponges, and protists (for reviews, see Adams, 2000; Rai et al., 2000, 2002; Adams et al., 2006; Bergman et al., 2007). The cyanobacterial symbionts (cyanobionts) generally supply their hosts with fixed nitrogen, although they can also provide fixed carbon to non-photosynthetic hosts. The major plant hosts are bryophytes, cycads, the angiosperm Gunnera, the water-fern Azolla, and fungi (to form lichens). Although all cyanobacteria are photoautotrophs, many are also facultative heterotrophs and so are not restricted to the areas of the plant that receive light, and can be found in roots, stems, leaves, and thalli. This review will concentrate on the cyanobacteria–bryophyte symbioses, focusing in particular on the importance of pili and gliding motility in plant infection. For comprehensive reviews on the cyanobacteria–bryophyte symbiosis, see Adams (2002), Meeks and Elhai (2002), and Meeks (2003). Space constraints have meant that, in reviewing the literature, we have had to concentrate on the more recent articles and reviews, rather than referring to the original literature.
Plant cyanobionts all have two major characteristics in common: (i) the ability to differentiate both specialized nitrogen-fixing cells known as heterocysts (for reviews, see Adams and Duggan, 1999; Meeks and Elhai, 2002; Golden and Yoon, 2003; Zhang et al., 2006); and (ii) short, motile filaments known as hormogonia, which lack heterocysts and provide a means of dispersal for otherwise immotile cyanobacteria (Campbell and Meeks, 1989; Meeks, 1990, 1998; Johansson and Bergman, 1994; Bergman et al., 1996). Heterocysts usually occur singly in a semi-regular spacing within filaments of vegetative cells (Golden and Yoon, 2003; Zhang et al., 2006). The infective agents in most plant symbioses are hormogonia and some, perhaps all, plants produce chemical signals that trigger their formation and chemoattractants that guide them into the plant tissue.
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The symbiotic partners |
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TopAbstractIntroductionThe symbiotic partnersThe infection processPost-infection changesCyanobacterial motilityHormogonia and piliConcluding remarksReferences |
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Bryophytes
Bryophytes are small, non-vascular land plants encompassing the liverworts (Hepaticae), the hornworts (Anthocerotae), and the mosses (Musci), a relatively small number of which are able to form epiphytic or endophytic associations with cyanobacteria (Adams, 2002; Meeks, 2003). In the case of the mosses, the cyanobacteria grow mostly epiphytically (Solheim and Zielke, 2002; Gentili et al., 2005), the exception being two Sphagnum species in which the cyanobacteria occupy water-filled, hyaline cells, which are thought to provide some protection from the acidic bog environment (Solheim and Zielke, 2002). Similar protection for cyanobacteria on the moss leaf surface is provided by the secretion of alkaline substances (Belnap, 2001). These nitrogen-fixing cyanobacterial associations with mosses often supply most of the combined nitrogen in local ecosystems in the Arctic, the Antarctic, and boreal forest regions (Zielke et al., 2002, 2005). This may be particularly significant in boreal forests where the feather moss Pleurozium schreberi (with its epiphytic cyanobacteria) can provide 80% of the ground cover (DeLuca et al., 2002, 2007; Zackrisson et al., 2004; Gentili et al., 2005).
Cyanobacterial associations with liverworts are rare, being found in only four of the >340 liverwort genera, two of the associations (Marchantia and Porella) being epiphytic and two (Blasia and Cavicularia) endophytic (Meeks, 1990). By contrast, in the hornworts, of which there are presently 13 genera described (Duff et al., 2007), endophytic associations are ubiquitous (Renzaglia et al., 2007). In nature, liverworts and hornworts exist as a flattened gametophyte thallus a few centimetres in length and symbiotic colonies are visible as small, dark spots (Fig. 1A). The endophytic associations with hornworts, such as Anthoceros and Phaeoceros, and liverworts such as Blasia, are particularly suited to laboratory experimentation because the host plant can be freed of its symbiotic cyanobacteria and grown in shaken liquid culture, and the symbiosis re-established with the original or with novel cyanobacteria (Fig. 1B; Adams, 2002; Meeks, 1998, 2003).
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Bryophyte ‘symbiotic’ structures
Mature cyanobacterial colonies are up to 0.5 mm in diameter and occupy dome-shaped structures known as auricles on the ventral surface of the thallus of the liverwort Blasia (Fig. 1C), or slime cavities within the thallus of hornworts such as Anthoceros and Phaeoceros (Fig. 1D). These structures are not unique to the symbiotic state, but are also present in plants not infected with cyanobacteria. The hornwort slime cavity is connected to the ventral surface of the thallus via stomata-like pores (Meeks, 1998, 2003; Adams, 2002). By contrast, in the hornwort Leiosporoceros dussii, the cyanobacteria are found in mucillage-filled ‘canals’ which run parallel to the main axis of the thallus (Villarreal et al., 2005; Villarreal and Renzaglia, 2006). These canals are formed by the separation of plant cell walls along their middle lamellae and, as the canals elongate they bifurcate to form an integrated network throughout the plant (Villarreal and Renzaglia, 2006). This unique arrangement of canals enables the Nostoc to spread throughout the Leiosporoceros dussii thallus without the repeated infections required for the discrete colonies in other hornworts and liverworts.
The cyanobacteria
In the great majority of cases, plant cyanobionts are members of the genus Nostoc, which is commonly found free-living in nature (Dodds et al., 1995; Rai et al., 2002). This is equally true of the liverworts and hornworts in which the major cyanobionts are Nostoc spp. (West and Adams, 1997; Costa et al., 2001). However, in the laboratory, other hormogonium-developing cyanobacterial genera, such as Calothrix and Chlorogloeopsis, may infect liverworts (West and Adams, 1997).
Hormogonia
For free-living cyanobacteria to locate and enter the symbiotic cavities of host plants they must be motile. Cyanobacteria move by a process known as gliding, which requires contact with a surface. The mechanism of gliding is unknown, but as motility is so vital to plant infection it will be dealt with separately in the final two sections of this review. Members of the genus Nostoc are primarily non-motile, but a characteristic of the genus is the ability to produce specialized motile filaments known as hormogonia which serve as a means of dispersal as well as plant infection (Bergman et al., 1996; Meeks, 1998, 2003; Adams, 2000; Meeks and Elhai, 2002). Hormogonia development is triggered by a variety of environmental factors, including plant-derived chemical signals (see the section on hormogonia induction). For a successful symbiosis to be established with the plant host the cyanobiont must also have the capacity to develop nitrogen-fixing heterocysts, although mutants capable of forming hormogonia, but unable to form heterocysts, are still able to infect Anthoceros punctatus but cannot establish a fully functional, nitrogen-fixing symbiosis (Wong and Meeks, 2002).
The development of hormogonia in heterocystous cyanobacteria results from a round of rapid, synchronous cell divisions which result in a decrease in cell size (Meeks and Elhai, 2002; Meeks et al., 2002). This is followed by fragmentation of the filament at the heterocyst–vegetative cell junctions, releasing short, motile hormogonia. Hormogonia lack heterocysts and are a temporary stage in the Nostoc life-cycle, soon returning to vegetative growth and developing heterocysts once more. For hormogonia to locate the symbiotic tissue of a plant host they must attach to the surface and both extracellular polysaccharides and pili (fimbriae) are thought to play a role in this process (Adams, 2000). Type IV pili are required for gliding in some unicellular cyanobacteria (Bhaya, 2004), and the cell surface of hormogonia of the symbiotically competent Nostoc punctiforme is covered with pili (Duggan et al., 2007). The importance of pili in the symbiotic competence and motility of hormogonia will be discussed in detail in the final section of this review.
Recent microarray data have revealed over 1800 differentially transcribed genes in hormogonia 24 h post-induction, some 5-fold greater than the number transcriptionally active in akinete (spore)-forming or dinitrogen-fixing cultures (Campbell et al., 2007). Of these, 944 genes are up-regulated, with the majority of these encoding proteins for signal transduction and transcriptional regulation. Genes encoding putative chemotaxis proteins and pilus biogenesis genes are also up-regulated in hormogonia (Meeks et al., 2001; Klint et al., 2006; Campbell et al., 2007). The hormogonium appears genetically rich in terms of potential for sensory and regulatory perception and has evolved to sense and respond to a variety of environmental signals, such as light quality and quantity, chemicals released from potential host plants, or favourable environments that will support subsequent growth. The genes involved in these sensory pathways remain to be characterized and genes central to the hormogonia differentiation process itself remain to be identified.
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The infection process |
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TopAbstractIntroductionThe symbiotic partnersThe infection processPost-infection changesCyanobacterial motilityHormogonia and piliConcluding remarksReferences |
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Hormogonia induction
Plant hosts increase the likelihood of infection by cyanobacteria by both stimulating the formation of hormogonia in potential cyanobionts and by guiding the hormogonia to the symbiotic tissues by chemotaxis. Hormogonia formation is stimulated by hormogonia-inducing factors (HIFs). HIF production has been found in the hornwort Anthoceros punctatus (Meeks, 2003), as well as cycads and the angiosperm Gunnera (Rasmussen et al., 1994; Ow et al., 1999). Anthoceros punctatus HIF is a small, heat-labile product released by the hornwort when starved of combined nitrogen (Meeks and Elhai, 2002; Meeks, 2003). The liverwort Blasia also releases HIF when nitrogen-starved (Adams, 2002). Nostoc punctiforme mutants with increased sensitivity to Anthoceros HIF, also show a greater initial frequency of infection of the hornwort than the wild type (Cohen and Meeks, 1997). Mutation of ntcA, which encodes the global transcriptional regulator, NtcA (Herrero et al., 2004) results in a reduction in the frequency of hormogonia induced by HIF in N. punctiforme, and the hormogonia that are produced do not infect Anthoceros (Wong and Meeks, 2002).
The efficiency of infection of the liverwort Blasia by N. punctiforme is influenced by mutations in the gene cyaC, which encodes adenylate cyclase, the enzyme responsible for the biosynthesis of the intracellular messenger adenosine 3',5'-cyclic monophosphate (cAMP). Cyanobacteria possess multiple adenylate cyclases, of which CyaC is the most complex, consisting of seven domains (Cann, 2003). A N. punctiforme mutant containing an antibiotic resistance cassette towards the 5' region of cyaC (within the second of the two GAF domains encoded by the gene), infects Blasia at 
25% of the wild-type frequency (Chapman et al., 2004). By contrast, a mutant carrying an insertion of the same cassette in the 5' region of cyaC, encoding the putative catalytic domain, infects Blasia at a 3–4-fold higher frequency than the wild type (Chapman et al., 2004; Chapman, 2006). In the presence of Blasia, both mutants form similar frequencies of hormogonia to the wild type, but 12 h earlier than the wild type. Transfer of wild-type cells from light growth conditions to the dark produced a rapid increase in cAMP, consistent with its role as a signalling molecule. By contrast, the cyaC mutants do not show any corresponding increase in cellular cAMP levels following transfer from the light to the dark and cellular cAMP levels are only 25% of the wild type. The Nostoc genome contains additional putative adenylate cyclases which may account for the small amounts of cAMP present in the mutant strains. The similar cAMP levels in these two mutants with contrasting infection phenotypes, implies that cAMP per se is not involved in symbiotic competency. The reason for these different phenotypes remains unclear; both mutants are motile and there is no apparent difference in hormogonia piliation compared with the wild type. It may be that unrecognized differences in the behaviour of the mutant hormogonia, in response to plant signals, are responsible for their infection phenotypes.
Further evidence that hormogonia behaviour, as well as frequency, has an influence on plant infection comes from work on the N. punctiforme gene sigH, which encodes an alternative sigma subunit of RNA polymerase, and transcription of which is induced by Anthoceros HIF (Campbell et al., 1998; Meeks, 2003). Inactivation of sigH has no effect on the frequency of HIF-induced hormogonia, but the hormogonia are 3–5-fold more efficient than the wild type at infecting A. punctatus (Meeks and Elhai, 2002; Meeks, 2003).
Chemotaxis
Hormogonia production alone is not sufficient to guarantee establishment of symbiosis since non-infective Nostoc strains are also capable of forming motile hormogonia in the presence of the angiosperm Gunnera (Johansson and Bergman, 1994). Clearly, hormogonia must possess specific characteristics that enable them to infect a host successfully. One such characteristic is likely to be the ability to respond chemotactically to plant-derived chemicals and this is especially important in plants such as Gunnera and cycads in which the symbiotic tissue does not receive light and, as a consequence, the hormogonia must overcome their natural positive phototaxis. The importance of chemotaxis is reflected in the genome of the symbiotically competent cyanobacterium N. punctiforme which has three to five copies each of the genes encoding homologues of the chemotaxis-related proteins CheA, CheB, CheW, CheD, and CheR (Meeks et al., 2001). When the liverwort Blasia is starved of combined nitrogen it releases both HIF (Babic, 1996) and a very effective chemoattractant (Knight and Adams, 1996; Watts et al., 1999). Non-host plants such as Trifolium repens (Nilsson et al., 2006) and germinating wheat seeds (Knight and Adams, 1996; Watts et al., 1999) can also release hormogonia chemoattractants.
Invasion of the plant
Once hormogonia enter Blasia auricles (and presumably the slime cavities of hornworts), they lose motility and differentiate heterocysts (Kimura and Nakano, 1990). The infection of hornworts via the stomata-like opening to the slime cavity has interesting parallels with the likely method of entry of cyanobacteria into the primitive, extinct land plant Aglaophyton major. This symbiosis is only known from fossil evidence, but an Archaeothrix-type filamentous cyanobacterium is thought to have entered the plant via stomatal pores (Taylor and Krings, 2005). The cyanobacteria are thought to have initially colonized the substomatal chambers and then spread throughout the outer cortical tissue, where they can be seen in fossil specimens of the plant. This is somewhat similar to the infection process in the extant hornwort Leiosporoceros dussii in which the cyanobacteria are found in mucilage-filled ‘canals’ (see section on bryophyte symbiotic structures; Villarreal et al., 2005; Villarreal and Renzaglia, 2006).
Although the development of heterocysts is essential for functional, nitrogen-fixing symbiotic colonies, Nostoc mutants unable to form heterocysts can still infect the hornwort A. punctatus. Strains carrying mutations in hetR (Wong and Meeks, 2002), which is the primary activator of heterocyst development (Golden and Yoon, 2003; Zhang et al., 2006), and hetF (Wong and Meeks, 2002), which is involved in the control of hetR transcription and the localization of HetR to developing heterocysts (Wong and Meeks, 2001), infect Anthoceros at wild-type frequency but fail to support the growth of the hornwort (Wong and Meeks, 2002). By contrast, a strain carrying a mutation in ntcA, which encodes the global nitrogen regulator NtcA (Flores and Herrero, 2005), does not infect the hornwort, despite producing motile hormogonia, albeit at a reduced level to the wild type (Wong and Meeks, 2002).
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Post-infection changes |
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TopAbstractIntroductionThe symbiotic partnersThe infection processPost-infection changesCyanobacterial motilityHormogonia and piliConcluding remarksReferences |
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Once the cyanobacterium has entered the host plant a number of morphological, developmental, and physiological changes occur (Table 1). The development of hormogonia is repressed, whereas the development of heterocysts is greatly stimulated. The rate of cell division is reduced, ensuring that the cyanobiont does not outgrow the host. The rate of CO2 fixation is greatly reduced, whereas nitrogen fixation is stimulated and ammonium assimilation down-regulated.
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Hormogonia repression
Once inside the host plant the hormogonia revert to vegetative growth, differentiating heterocysts and fixing nitrogen. In the hornwort A. punctatus (and presumably also liverworts) further hormogonia formation within the symbiotic colony is prevented by a water-soluble hormogonium repressing factor (HRF) released into the colony by the plant (Meeks and Elhai, 2002; Meeks, 2003). HRFs induce expression of the hormogonium-repressing genes located in the hrmUA locus of N. punctiforme (Cohen et al., 1994; Cohen and Meeks, 1997). Regulation is achieved through the sugar-binding transcriptional repressor HrmR, which prevents any new rounds of hormogonium development (Campbell et al., 2003).
CO2 fixation
Light-dependent CO2 fixation of the Nostoc cyanobiont of Anthoceros, immediately after isolation from the hornwort, is only 12% that of the free-living cyanobacterium (Table 1). However, the level of ribulose-1,5-bisphosphate carboxylase/oxygenase (Rubisco) protein, the major enzyme involved in the fixation of CO2 in cyanobacteria, is little different in free-living or Anthoceros-associated Nostoc, implying that activity is regulated by an unknown post-translational modification of the protein (Meeks and Elhai, 2002; Meeks, 2003). The reduced carbon-fixing capacity of the cyanobiont is compensated by sugars derived from the host (Meeks and Elhai, 2002; Adams, 2002; Meeks, 2003).
N2 fixation
The nitrogen fixation rates for cyanobacteria symbiotically associated with bryophytes are several-fold higher than for the same free-living cyanobacteria (Table 1). This increase is due to a greatly elevated heterocyst frequency, which may be 6–10-fold higher than in the free-living state (Table 1). As little as 20% of the nitrogen fixed is retained by the cyanobiont, the remainder being transferred as ammonia to the host (Adams, 2002; Meeks and Elhai, 2002). The primary route of ammonia assimilation in cyanobacteria is the GS-GOGAT (glutamine synthetase–glutamate synthase) pathway (Meeks and Elhai, 2002). The level of GS protein in Anthoceros-associated Noctoc is similar to that in free-living cyanobacteria, but GS activity is reduced (Table 1) implying that activity is regulated by an unknown, and presumably plant-regulated, post-translational modification of the enzyme (Meeks and Elhai, 2002; Meeks, 2003).
Morphological changes to cyanobiont and host
Cyanobiont growth rate must match that of the host plant if the association is to be stable. Indeed, the growth of Nostoc symbiotically associated with Anthoceros can be 5–10-fold slower than in the free-living state (Meeks, 2003; see Table 1). The host can also regulate the biomass and nitrogen fixation of the symbiotic colonies to maintain a constant rate of nitrogen fixation per unit of plant tissue, but the mechanism by which this is achieved is unknown (Meeks, 2003).
As is the case with all cyanobacteria–plant symbioses, cyanobacteria in symbiosis with hornworts and liverworts exhibit greatly elevated heterocyst frequencies compared with the free-living state (Table 1; Meeks, 2003). This enhances the colony's dinitrogen-fixing capacity to the benefit of the host, although it seems likely that at least some of these extra heterocysts are non-functional (Meeks and Elhai, 2002). Within the symbiotic colony, heterocysts become difficult to recognize because they often lose some of the morphological characteristics that distinguish them from vegetative cells, including their regular shape and thickened cell walls (Adams, 2002; Meeks and Elhai, 2002). The vegetative cells also show altered morphology in symbiosis, being enlarged and irregular in shape compared with their free-living counterparts (Meeks and Elhai, 2002).
Although cyanobacteria occupy liverwort and hornwort structures present prior to the infection, limited morphological changes do occur to these structures post-infection. In both Blasia and Anthoceros, the walls of the symbiotic cavity elaborate branched, multicellular filaments that penetrate the colony, facilitating the exchange of nutrients between the colony and the plant (Adams, 2002). Indeed, the filaments in Blasia, which show ultrastructural differences to those in Anthoceros, possess some of the characteristics of transfer cells (Duckett et al., 1977). What induces these morphological changes is not known, although cyanobacteria do produce the phytohormone auxin, indole-3-acetic acid (Sergeeva et al., 2002) and other compounds known to be involved in plant development (Liaimer and Bergman, 2004). Plant host genes equivalent to the nodulation genes of legumes have so far not been identified in any of the plant hosts in cyanobacterial symbioses (Udvardi et al., 2005).
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Cyanobacterial motility |
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TopAbstractIntroductionThe symbiotic partnersThe infection processPost-infection changesCyanobacterial motilityHormogonia and piliConcluding remarksReferences |
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For cyanobacteria to locate and invade the relevant plant host tissue, the capacity for motility, coupled with chemotaxis, is essential. Of the many motile cyanobacteria the vast majority move by gliding, which requires attachment to a surface. The exception is a few unicellular Synechococcus strains that can swim by a mechanism in which thrust is thought to be generated, not by flagella, but by the extracellular glycoprotein SwmA (McCarren and Brahamsha, 2005; McCarren et al., 2005). The form of gliding employed by the unicellular Synechocystis sp. strain PCC 6803 is sometimes referred to as twitching motility, and is powered by type IV pili on the cell surface (Bhaya et al., 1999, 2000; Yoshihara and Ikeuchi, 2004). In filamentous strains, such as members of the Oscillatoriaceae, forward movement is often accompanied by rotation of the filament about its long axis (Halfen, 1973), although the mechanism of gliding is unknown. An early model for gliding in these filamentous strains was developed by Halfen and Castenholz (1971), based on both their own observations and the theories of Jarosch (1963). They proposed that the thrust for gliding might be provided by contraction of 6–9 nm diameter protein fibrils, situated between the peptidoglycan layer and the outer membrane. The helical arrangement of these fibrils could account for the rotation of filaments as they glide.
More recently, Adams et al. (1999) observed an array of parallel fibrils, in the same location as those observed by Halfen and Castenholz (1971), between the peptidoglycan and the outer membrane, but of a much greater diameter (25–30 nm, rather than 6–9 nm). Penetration of the outer membrane between the rows of fibrils results in corrugations in the outer surface of the cells. This corrugated appearance can be visualized by field emission scanning electron microscopy (FEGSEM; Fig. 2) and by atomic force microscopy (AFM; Fig. 3). In a re-iteration of the model of Halfen and Castenholz (Halfen and Castenholz, 1971; Halfen, 1973), Read et al. (2007) have proposed that rhythmical waves passing along the 25–30 nm fibrillar array could be transmitted via the outer membrane and, by interaction with the substrate on which the cyanobacterium rests, drive the filament in the opposite direction.
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An alternative model, which involves slime extrusion, has been developed by Hoiczyk and Baumeister (1995) who identified a layer of helically arranged 10–12 nm diameter fibrils of a calcium-binding protein, which they called oscillin, in several filamentous cyanobacteria (Hoiczyk and Baumeister, 1997). These fibrils are external to the outer membrane and rest on a tetragonal S-layer (Hoiczyk and Baumeister, 1995). Hoiczyk and Baumeister (1997, 1998) proposed that gliding was powered by the extrusion of slime from the junctional pores that form a ring around each cell septum. They concluded that the helically arranged oscillin fibrils acted like a screw thread, resulting in rotation of the trichome. A mutant of Phormidium uncinatum unable to extrude slime and lacking both the S-layer and oscillin fibrils is immotile, implying that one, or more, of these components is essential for motility (Hoiczyk and Baumeister, 1997). At present there is no conclusive evidence to support either the slime extrusion or the surface wave models for force generation in gliding. The development of AFM scanning of live cyanobacterial filaments under liquid may soon make it possible to visualize surface waves, should they exist.
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Hormogonia and pili |
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TopAbstractIntroductionThe symbiotic partnersThe infection processPost-infection changesCyanobacterial motilityHormogonia and piliConcluding remarksReferences |
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The mechanism of gliding in non-heterocystous filamentous cyanobacteria, such as members of the Oscillatoriaceae, remains obscure, but what of the heterocystous cyanobacteria that are potential cyanobionts of plants? These cyanobacteria are capable of transforming immotile filaments into motile hormogonia, but what is known about the mechanism of hormogonia motility?
The external surface of a wide range of bacteria is covered with filamentous structures known as Type IV pili (Tfp) which function in numerous processes including motility, adhesion, pathogenesis, and DNA uptake (Wolfgang et al., 2000; Mattick, 2002; Nudleman and Kaiser, 2004; Burrows, 2005). Nostoc punctiforme vegetative filaments are non-motile and appear to lack these structures (Fig. 4A), whereas the cell surface of motile N. punctiforme hormogonia has abundant Tfp-like appendages (Fig. 4B), produced during differentiation (Doherty and Adams, 1999). The pili are arranged peritrichously, extend up to 10 µm from the cell surface, and have diameters of 7–10 nm. Tfp are composed primarily of a single protein, pilin (PilA), which is synthesized as a precursor (prepilin) and undergoes endoproteolytic cleavage, followed by methylation of the N-terminal residue, prior to assembly into the filament. Both endoproteolytic cleavage and N-methylation are carried out by the prepilin peptidase (PilD), which is known to process a wide range of other prepilin-like proteins (Strom et al., 1991; Paranjpye et al., 1998; Lammertyn and Anne, 2004). Motility associated with Tfp involves cycles of pilus extension, surface attachment of the pilus tip, and retraction of the pilus filaments to transport the cell toward the point of adhesion (Merz et al., 2000; Skerker and Berg, 2001; Merz and Forest, 2002). Pilus retraction generates considerable force (Merz et al., 2000; Maier et al., 2002) and is thought to occur via filament disassembly mediated by the hexameric ATPase nucleotide-binding protein, PilT (Skerker and Berg, 2001; Morand et al., 2004). PilT mutants are unable to retract their pili, leading to hyperpiliation and loss of twitching motility (e.g. Whitchurch et al., 1991; Wolfgang et al., 1998; Merz et al., 2000).
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To date, cyanobacterial Tfp systems have been demonstrated in Synechocystis PCC 6803 (Bhaya et al., 1999, 2000, 2001; Yoshihara et al., 2001, 2002) and Microcystis aeruginosa PCC 7806 (Nakasugi and Neilan, 2005; Nakasugi et al., 2007), together with a report of Tfp-like genes in the genome of the thermophillic cyanobacterium Thermosynechococcus elongates BP-1 (Iwai et al., 2004). Additionally, Tfp-like genes can be found in the cyanobacterial genomes of Anabaena sp. PCC 7120, Gloeobacter violaceus PCC 7421, Prochlorococcus marinus MIT9313, Synechococcus sp. WH8102, Synechococcus elongatus PCC 6301, and Synechococcus sp. CC9311 (http://bacteria.kazusa.or.jp/cyano/). The genome of the symbiotically competent N. punctiforme (http://genome.jgi-psf.org/finished_microbes/nospu/nospu.home.html) also contains open reading frames (ORFs) that are predicted to encode proteins with similarity to Tfp biogenesis proteins, including two pilT-like genes. Inactivation of one of these pilT-like genes (NpR0117) results in a hyperpiliated (over-production of pili) phenotype (Fig. 5C) that is typical of mutations within pilT in other bacteria (Duggan et al., 2007). Mutation of the second Nostoc pilT-like NpF2507 results in slightly less-abundant pili than the wild type, although their length and diameter appear to be similar to those of the wild type (Fig. 5B). The NpR0117 (hyperpiliated) mutant infects the host plant Blasia pusilla at 9.5% of the frequency of the wild type, implying that normal surface piliation is required for effective establishment of symbiosis. The role of pilT-like NpF2507 is less clear. NpF2507 mostly resembles PilT2 from Synechocystis PCC 6803, which is dispensable for motility but is believed to function in a signalling pathway that regulates the positive phototaxis of this cyanobacterium (Bhaya et al., 2000). The NpF2507 mutant infects Blasia at an initial frequency 50% that of the wild type, but reaches a level equivalent to that of the wild type after 28 d, implying a possible role during the early stages of the infection process. Whether NpF2507 is required for the direction of hormogonia motility (possibly involving chemotaxis-like regulatory elements), as may be the case of the Synechocystis PilT2, remains to be investigated.
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Disruption of N. punctiforme pilD-like NpR2800 gives rise to much shorter pili (up to 2.5 µm in length) than those of the wild type (up to 10 µm in length), although pilus diameter (7–10 nm) is similar to that of the wild type (Fig. 5A; Duggan et al., 2007). In other bacteria, mutations in pilD abolish motility and produce a non-piliated phenotype (e.g. Lory and Strom, 1997; Bhaya et al., 2000; Friedrich et al., 2002). The truncated pilus phenotype observed in the Nostoc pilD mutant is unusual and may represent partially retracted or partially extended pili. With the pilD-like NpR2800 mutant the frequency of Blasia infection is only 2.4% of that of the wild type. Many of the symbiotic filaments are brown in colour, implying that they are dead or dying and that the mutant is unable to grow effectively within the host symbiotic cavity. The Nostoc pilD-like gene may therefore be required for some aspect of symbiotic growth.
Nostoc punctiforme has at least five ORFs whose predicted products contain features that characterize the pilus structural subunit (PilA). Of the three Nostoc pilA-like ORFs investigated by Duggan et al. (2007), complete segregation was only achieved with pilA-like ORF NpF0069. The pili expressed on the surface of the hormogonia produced by this mutant were indistinguishable from those of the wild type, yet the mutant infected plant tissue at a frequency only 38% that of the wild type. It is possible that pilA-like NpF0069 encodes a minor pilin (Alm and Mattick, 1996; Winther-Larsen et al., 2005) that may support the activity of the major pilin in some aspect of motility and/or adhesion and therefore contribute to symbiotic competency.
It is clear that mutation of some pil genes affects the symbiotic competence of hormogonia, but what effect do these mutations have on motility? The hormogonia of the wild-type strain employed in the study of Duggan et al. (2007) were inconsistently motile and when motility was present the hormogonia progressed slowly at rates of 0.7 µm s–1 to 1.7 µm s–1. Slight twitching of filaments, but without forward progression, was detected in the pilA-like NpF0069 and pilT-like NpF2507 mutants, whereas motility was not observed in strains inactivated in pilD-like NpR2800 and pilT-like NpR0117. Given the inconsistencies in the motile behaviour of the wild-type strain, the possibility that the lack of motility associated with the NpR2800 and NpR0117 mutants is the result of mutation in the pilD-like ORF and the pilT-like ORF, respectively, remains to be demonstrated. Nevertheless, the data confirm that Tfp are required for effective establishment of Nostoc–Blasia symbiosis and, most likely, other cyanobacteria–plant symbioses.
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Concluding remarks |
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TopAbstractIntroductionThe symbiotic partnersThe infection processPost-infection changesCyanobacterial motilityHormogonia and piliConcluding remarksReferences |
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Although current understanding of the molecular basis for many of the interactions involved in cyanobacteria–plant symbioses is poor, continuing advances in molecular genetic techniques for symbiotically competent cyanobacteria should lead to more rapid advances in this area. For example, recently obtained N. punctiforme microarray data has revealed the remarkable complexity of hormogonium differentiation, with over 1800 genes being differentially transcribed during the process (Campbell et al., 2007). Despite the importance of motility and chemotaxis in plant infection by hormogonia, neither process is well understood. However, the availability of the genome sequence of the symbiotically competent N. punctiforme (http://genome.jgi-psf.org/finished_microbes/nospu/nospu.home.html) and the N. punctiforme microarray, should lead to significant advances in this area. Certainly such advances will be needed if the goal of developing novel associations between cyanobacteria and crop plants is to be realized.
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References |
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