Microbial Symbionts: Functions and Molecular Interactions on Host

By Dhanasekaran Dharumadurai

Microbial Symbionts: Functions and Molecular Interactions on Host focuses on microbial symbionts of plants, animals, insects and molecular methods in the identification of microbial symbionts. The book describes the molecular mechanism and interactions of symbiosis of microbiome in plants, animals and humans. It brings the latest techniques for identification, localization and functional characterization of host-associated microbes and explains the role/importance of microbial symbionts. This comprehensive reference covers a wide range of symbiotic microorganisms used for basic and advanced techniques associated with the isolation, characterization and identification of microbial symbiotic microorganisms and their functions and molecular interactions on the host.

The book will also helps users plan and execute experiments with appropriate knowledge rather than experimental trial and error in a wide range of disciplines, including Microbiology, Biotechnology, Botany and Zoology.

• Provides basic knowledge and working protocols for a wide range of disciplines like Microbiology, Biotechnology, Botany and Zoology
• Presents the most current information in symbiotic microbiome and holobiome
• Includes color photos pertaining to techniques

• Language: English
• Publisher: Academic Press
• Release date: Sep 25, 2022
• ISBN: 9780323993357

Book preview

Preface

The proposed book entitled "Microbial Symbionts: Functions and Molecular Interactions on Host" illustrates the techniques involved in the study of symbiotic microorganisms and their functional role in plants, animals, and insects. Microbial symbionts create organic material through carbon and nitrogen fixation, as well as via inorganic nitrogen and phosphorous assimilation processes. Microbial symbionts are primary producers in association with fungi, plants, and animals living in low-nutrient environments. This book covers a wide range of symbiotic microorganisms used for basic and advanced techniques associated with the isolation, characterization, and identification of microbial symbiotic microorganisms and also their functions and molecular interactions on host. Photosynthetic bacteria are primary producers as the carbon dioxide-fixing and methane-oxidizing symbionts of marine ciliates, bivalves, crustaceans, oligochaete and polychaete worms, and gastropods in habitats with high concentrations of chemical reductants. Bacterial symbionts fix inorganic nitrogen gas in association with photoautotrophs with high nitrogen needs (eukaryotic phytoplankton and plants), as well as with animals living in oligotrophic habitats or those subsisting on low-nitrogen food sources like wood (sponges, corals, bivalves, and insects).

The book consists of 48 chapters under the four broad sections under which different titles are prepared based on its content with main focus on microbial symbionts of plants, animals, insects, and molecular methods in the identification of microbial symbionts. The chapters are contributed from valued contributors worldwide including leading experts in Colombia, Egypt, Finland, Morocco, India, and the Netherlands. It also outlines the molecular mechanism, interactions of symbiosis of microbiome in lichen, sponges, rumen, root nodules, coralloid roots, insects, human gut, and other animals, latest techniques for identification, localization, and functional characterization of host-associated microbes, culture-independent and dependent approaches in symbionts analysis, key model systems in animal–microbe and plant–microbe beneficial interactions of symbionts, molecular symbiotic interactions of cyanobacterial association in nonvascular seedless plants, symbiotic functional molecules in endophytic actinobacteria in actinorhizal plants, in situ techniques to study noncultivable microbial symbionts, biosynthetic gene clusters of symbiotic gut microbiome in succession of human health, and functions and molecular interactions of the symbiotic microbiome in oral cavity of human. Information panel at the beginning of each chapter will detail the basic understanding as well as advancement on relevant topic. Postgraduate students, research scholars, postdoctoral fellows, and teachers belonging to different disciplines via life sciences including microbiology, biotechnology, botany, zoology, and agricultural sciences disciplines are the targeted readers of this book.

I am extremely thankful to all the authors who contributed chapters for their prompt and timely responses. I extend my earnest appreciation to Mrs. Linda Versteeg-Buschman, Sr. Acquisitions Editor, Elsevier/Academic Press, The Netherlands, and their team for their constant encouragement and help in bringing out the volume in the present form. I am also indebted to Elsevier/Academic Press, The Netherlands and the Authorities of Bharathidasan University, Tiruchirappalli, Tamil Nadu, India, for their support in the task of publishing this book.

Dr. Dhanasekaran Dharumadurai

Associate Professor

Department of Microbiology

Bharathidasan University,

Tiruchirappalli- 620 024, India

[e-mail]

[e-mail]

Section 1

Microbial symbionts of plants: functions and molecular interactions with the host

Outline

Chapter 1. Cyanobacterial symbiotic interaction in pteridophytes

Chapter 2. Cyanobacterial symbiosis with bryophytes

Chapter 3. Symbiotic cyanobacteria in gymnosperms

Chapter 4. Cyanobacterial symbionts from angiosperm

Chapter 5. Frankia from actinorhizal plants

Chapter 6. Conventional and unconventional symbiotic nitrogen fixing bacteria associated with legumes

Chapter 7. Ensifer meliloti is the main microsymbiont of Prosopis chilensis in arid soils of Eastern Morocco

Chapter 8. Fungal mycorrhizae from plants roots: functions and molecular interactions

Chapter 9. Photobiont symbiotic association in lichens

Chapter 10. Endolichenic actinobacterial association in fruticose, foliose, and crustose lichens

Chapter 11. Epibionts and endolichenic microbial communities

Chapter 12. Mycobionts interactions in lichen

Chapter 13. Symbiotic functional molecules in endophytic actinobacteria in actinorhizal plants: scientometric profile

Chapter 14. Bacterial symbiosis in edible mushrooms

Chapter 15. Symbiotic microbial interactions in medicinal mushroom: an insight of phenotypic and genotypic characterization methods

Chapter 16. Molecular symbiotic interactions of cyanobacterial association in nonvascular seedless plants

Chapter 17. Microbial symbionts from Algae

Chapter 18. Screening of symbiotic ability of Rhizobium under hydroponic conditions

Chapter 19. Arbuscular mycorrhizal community analysis from a grassland ecosystem

Chapter 20. Mass production of cyanobacterial symbiont

Chapter 21. Exploration of Rhizobium for its mass production and plant growth promoting properties

Chapter 22. Symbiotic associations of Frankia in actinorhizal plants

Chapter 1: Cyanobacterial symbiotic interaction in pteridophytes

Neelam Mishra     Department of Botany, St. Joseph's University, Bengaluru, Karnataka, India

Abstract

The Azolla–Anabaena symbiosis is a mutualistic association between the prokaryotic nitrogen-fixing cyanobacterium Anabaena and water fern Azolla. During symbiosis, RUBISCO is conserved by the vegetative cells of the cyanobacteria and are perhaps able to fix CO2 during the different stages of the association, maintaining their autotrophic condition. Leaf cavity of Azolla is considered to be a natural microcosm, as it behaves as both the physiological and dynamic interface unit of this symbiotic association, where the main metabolic and energetic flows occur. Obligatory nature of this symbiosis is indicated by the sustained symbiosis throughout the life cycle of water fern where the cyanobionts are always present and transferred from one generation to the next, either in the dorsal leaf cavities or in the megasporocarps of Azolla. This symbiotic association is considered as an alternative to chemical fertilizers. Moreover, due to the high amino acids and protein content, this fern can also be used as food, and it can be used as phytoremediator due to its ability to uptake heavy metals. This interesting association has been explored at a physiological, biochemical, and molecular level to understand the mechanistic details of the association.

Keywords

Anabaena azollae; Azolla; Biofertilizer; Cyanobiont; Symbiosis

Introduction

Cyanobacterial pteridophyte symbiosis

Biogeography of Azolla

Morphology of Azolla

Reproduction of Azolla and maintenance of symbiosis

Physiology, biochemistry associated with symbiotic interaction

Photosynthesis and factors affecting rate of photosynthesis

Photosynthetic pigments

Nitrogen metabolism

Molecular studies on AnabaenaAzolla symbiosis

Economic importance of Azolla

References

Introduction

Cyanobacteria represent a phylogenetically rational group of ancient, morphologically diverse phototrophic bacteria with enormous ecological importance. They are defined by their ability to carry out oxygenic photosynthesis (water-oxidizing and oxygen-evolving). Cyanobacteria are unique in the wide range of symbiotic associations they form with eukaryotic hosts including plants, fungi, sponges, and protists (Adams, 2000; Adams & Duggan, 2012; Adams et al., 2012). Cyanobacteria are photoautotrophs, nitrogen fixers, and can provide nonphotosynthetic hosts with both nitrogen and carbon. The most common cyanobacteria (called cyanobionts) that form those symbioses belong to the genus Anabaena and Nostoc, are able to fix the atmospheric nitrogen in specialized cells (called heterocyst) through the nitrogenase complex enzyme, and thus convert it into the ammonium ion (NH4 +). As a result of symbiosis, the prokaryote gets a continuous supply of nutrients, protection against herbivores and environmental extremes such as high light intensity and desiccation, while the host gets all or almost all of the nitrogen needed for its development, allowing the successful colonization of nitrogen-poor environments (aquatic or terrestrial) (Bergman et al., 2003, 2007, 2008; Rai et al., 2000, 2002). The focus of this chapter is to emphasize the basic characteristics of symbiotic association between cyanobacteria and pteridophytes through biochemical, physiological, and molecular biology studies. Lastly a note on economic importance of the pteridophyte genus, i.e., Azolla, involved in symbiotic interaction is added.

Cyanobacterial pteridophyte symbiosis

The only pteridophyte known to exist in symbiosis with a N2 fixing cyanobacteria is the heterosporous aquatic fern comprising the genus Azolla Lam. Azolla occurs naturally in freshwater ditches, ponds, lakes, and sluggish rivers of warm-temperate and tropical regions. The name Azolla (azo: to dry, ollyo: to kill) implies that the plant dies under dry conditions (Lumpkin & Plucknett, 1980; Moore, 1969). Till date seven species of genus Azolla such as Azolla caroliniana, Azolla filiculoides, Azolla mexicana, Azolla microphylla and Azolla rubra in the section Euazolla (consists of three megaspore floats) and Azolla pinnata and Azolla nilotica in the section Rhizosperma (consists of nine megaspore floats) are known to harbor a symbiotic cyanobacterium, called as Anabaena azollae Strass (Pereira et al., 2011). Initially the genus Azolla was placed in a separate family called as Azollaceae (Eichler, 1965; Konar & Kapoor, 1974; Melchior & Werdermann, 1954; Sculthorpe, 1967). However, its close phylogenetic relationship with genus Salvinia made it to include Azolla along with Salvinia in family Salviniaceae (Bailey, 1949; Benson, 1957; Black, 1948; Lawrence, 1951; Smith, 1938).

Biogeography of Azolla

Azolla has an overall distribution from mild to heat and humidities; seven species assembled in two sections are classically accepted. The two segments (Rhizosperma and Euazolla) are recognized by distinctiveness of the megasporocarps and glochidia. The Rhizosperma segment includes Azolla nilotica, local to East Africa, and A. pinnata, of which two assortments are by and large perceived: A. pinnata var. pinnata, present all through Africa south of the Sahel, in Madagascar, and in Australia, and A. pinnata var. imbricata from subtropical and tropical Asia (Lawrence, 1951; Smith, 1938). In Euazolla section, A. caroliniana, A. Mexicana, and A. microphylla belong to temperate, subtropical, and tropical America, respectively, and A. filiculoides and A. rubra from America and the Far East, respectively.

Morphology of Azolla

Azolla are heterosporous species. On the first ventral leaf lobe, sporocarps arise in pairs with two microsporocarps/two megasporocarps/one of each (Lumpkin & Plucknett, 1980). Azolla plant floating on the surface of the water is roughly triangular or circular in shape and rarely exceeds 3–4cm except in the species Azolla nilotica that can reach upto 40cm in length (Lumpkin, 1985). The extensively branched horizontal floating stems are covered and hidden by overlapping, small, alternate, scalelike imbricate leaves. Adventitious roots are formed on the ventral part of the stem, are chlorophyllous, and grow vertically in the water (Lumpkin & Plucknett, 1980; Peters, 1978). Each leaf is deeply bilobed; the lower, thin, convex, and transparent achlorophyllous lobe ensuring flotation and the upper, aerial, chlorophyllous one developing an ellipsoid cavity that remains in contact with the external environment through a structurally sophisticated pore. Each ellipsoid cavity contains heterocyst forming nitrogen fixing filamentous cyanobacterium Anabaena azollae throughout its development (Pieterse et al., 1977). It was first described as Nostoc by Strasburger in 1873 and later renamed as Anabaena azollae in 1884 (Carrapiço, 2010). However, identity of cyanobiont was a controversy and was believed that it involves the genera Anabaena, Nostoc, and Trichormus (Adams, 2000; Adams et al., 2012; Brouwer et al., 2014; Papaefthimiou et al., 2008; Pereira & Vasconcelos, 2014; Peters & Meeks, 1989; Ran et al., 2010). The symbiotic association is permanent and it occurs at all stages in the life cycle of the pteridophyte. Cyanobacteria are transferred to next generation via dorsal leaf cavities or the megasporocarps, which confirms the obligatory nature of symbiosis (Adams et al., 2012; Brouwer et al., 2014; Carrapiço, 1991). The cyanobacterium Anabaena azollae occurs as filaments on the plant stem apexes and inside the leaf cavities, which are inoculated during their formation with few Anabaena from the apex. Filaments of Anabaena azollae associated with stem apex of Azolla are actively dividing, undifferentiated, and lack nitrogenase activity (Hill, 1975, 1977; Peter et al., 1976). Vegetative cells are uniform in size and contain numerous carboxysomes (Hill, 1977). Vegetative cells possess a bilayered cell wall typical to that of gram negative bacteria (Neumüller & Bergman, 1981). Up to 30% of the Anabaena cells differentiate into heterocysts as the leaves mature (Adams et al., 2012; Carrapiço & Tavares, 1989; Lechno-Yossef & Nierzwicki-Bauer, 2002). Heterocysts differentiation from vegetative cells are manifested due to observable structural changes, viz. formation of extra envelope layers, narrowing between the junctions of heterocysts and adjacent vegetative cells, and the formation of large electron dense polar plugs (Lang, 1965; Neumüller & Bergman, 1981). Vegetative cells of some strain of Anabaena azollae may also form akinetes or resting cells. Isolated resting cells called akinetes may play a role in the persistence of the symbiosis during sexual reproduction of the plant; they also sometimes appear in leaf cavities (Miller & Lang, 1968).

In Azolla–Anabaena symbiosis, cyanobiont helps the plant with its nitrogen requirements by playing a crucial role in performing nitrogen fixation (Adams et al., 2012). As in many plant cyanobacterial symbioses, in this association also, cyanobiont greatly reduces or loses its photosynthetic capacity and is solely dependent on the host plant for its carbon requirements (Adams et al., 2012; Peters & Calvert, 1983).

Reproduction of Azolla and maintenance of symbiosis

Azolla undergoes either asexual reproduction via vegetative fragmentation or sexual reproduction via sporocarp formation. Most strains of Azolla reproduce exclusively by vegetative fragmentation (Watanabe, 1982). During vegetative fragmentation, abscission layers formed at the base of lateral branches facilitate separation of branches from the main rhizome. Sexual reproduction via sporocarp formation is less common. Azolla being a heterosporous fern produces two different types of sporocarp on the same plant, i.e., megasporocarp containing megasporangium (megaspore and megaspore apparatus) and microsporocarp containing microspore. Both mega and microsporocarp contain large amounts of sporopollenin that makes them very resistant to UV radiation, dessication, biodegradation, and nonoxidative chemical attack (Toia et al., 1985). Following maturation, sporocarps detach from the mother sporophyte and zygote formation occurs within megaspore apparatus at the bottom of the water surface (Konar & Kapoor, 1974).

The symbiotic association between Azolla and Anabaena is maintained throughout both sexual and asexual modes of reproduction. Each new Azolla plant produced by fragmentation carries its own inoculums, since vegetative sporophyte contains a generative Anabaena colony at each branch apex. The source of inoculums and mechanism of inoculation of Anabaena in megaspore during sporogenesis is not yet understood. It is well known that apex of sporophyte emerges from megaspore apparatus; it is inoculated with Anabaena. The nature of cyanobacterial inoculums may be different in different species of Azolla, for example, in A. filiculoides, the inoculums consist mainly of resting cells (Becking, 1978). The mechanism by which the cyanobacteria cells become associated with the developing sporocarps involves branched epidermal trichomes, which arise on the ventral surface of the apical meristem and probably includes chemical mediators (Adams et al., 2012). On entering the megasporocarp, the hormogonia differentiate into akinetes in the indusium chamber above the megaspore, remaining dormant until the plant germinates (Adams et al., 2012), initiating a new generation of the life cycle of Azolla.

Physiology, biochemistry associated with symbiotic interaction

Various biochemical, physiological, and molecular studies have been carried out in intact association, symbiont-free Azolla, Anabaena azollae separated from the entire plant.

Photosynthesis and factors affecting rate of photosynthesis

Pulse chase studies have shown that both partners in Azolla and Anabaena symbiotic association are to fix atmospheric CO2 and sucrose is major product of photosynthesis in both endophyte-free Azolla and its association. It has been observed that cyanobacterial populations isolated from Azolla leaf cavities are able to fix atmospheric CO2 (Peters, 1975; Ray et al., 1979). Additionally, it has also been demonstrated through autoradiographic studies that photosynthetic CO2 fixation takes place exclusively within the vegetative cells of endophyte (Peters, 1975). ¹⁴C sucrose was not detected as a major product of photosynthesis when ¹⁴CO2 was added to cyanobacteria isolated from leaf cavities. However ¹⁴CO2 was accumulated in a time-dependent manner when it is associated with endophyte. This clearly demonstrates that the carbon synthesized by Azolla is cross-fed and transferred to endosymbiont (Peter et al., 1985). Vegetative cells located at the apical meristem of Azolla are the primary site of cyanobacterial CO2 fixation (Hill, 1977) and Anabaena residing in older leaf cavities changes to mixotrophic/photohetrotrophic mode of nutrition due to increase in the number of heterocysts (Hill, 1975, 1977). Heterocysts lack photosystem II activity and are not capable of carbon fixation and larger amounts of reductants required by heterocysts for nitrogen fixation can be well supported (Peters, 1975).

A number of studies have been conducted to assess the factors affecting rate of photosynthesis such as light intensity, photoperiod, and temperature. Temperature of 35 °C was shown to inhibit rates of photosynthesis in A. pinnata (Tinh & Faludi-Daniel, 1981); however, maximum rates of photosynthesis were observed at temperature of 36 °C in A. caroliniana endophyte (Ray et al., 1979). Variations in photoperiod did not have significant effects on the rates of photosynthesis, for example, rates of photosynthesis were not affected in A. caroliniana when day lengths were varied as 12, 16, or 24h (Peters et al., 1981). Also it was reported that rates of photosynthesis were saturated at light intensity of 8 klx in A. caroliniana.

Photosynthetic pigments

Isolation and characterization of cyanobacterial pigments such as phycobiliproteins, c-phycocyanin, allophycocyanin, and phycoerythrocyanin have been done in A. caroliniana symbiosis (Tyagi et al., 1980 ). After analyzing the absorption and fluorescence emission spectra, it has been reported that there is some amount of chromatic adaptations in the photosynthetic pigments of the endophyte (Ray et al., 1978; Tung et al., 1981; Tyagi et al., 1980). The composition and pigment content of Anabaena and Azolla is influenced by environmental conditions, for example, anthocyanins such as luteolinidin-5-glucoside are produced by Azolla under low temperature and high light conditions (Holst and Yopp, 1979; Martin, 1976; Newton & Cavins, 1976; Watanabe, 1982).

In addition to above studies, photorespiration has also been discussed in detail in A. caroliniana symbiosis. It was found that CO2 compensation point was as high as 40μL/L in intact association or endophyte-free plants. Also, rate of CO2 fixation decreased to about 40% as a result of increase in O2 concentration from 2% to 20% (Ray et al., 1979). Therefore, it can be said that Azolla carries out photorespiratory metabolism. Contrary to this, cyanobacterial compensation point was reported to be less than 5μL/L and moreover rates of CO2 fixation were not affected by increase in O2 concentration. Therefore as in other cyanobacterial species, photorespiratory metabolism was not detected in Anabaena azollae (Bidwell, 1977; Lloyd et al., 1977). Apart from this, photochemical activity was also estimated for symbiotic, endophyte-free A. caroliniana and in cell-free extracts of the endosymbiont, and it was found that PSI and PSII activities of vegetative cells were higher than heterocysts (Haselkorn, 1978; Peters & Mayne, 1974a, 1974b). Phycobiliproteins exclusively serve as light harvesting pigments for PS II in free living, heterocystous cyanobacteria (Haselkorn, 1978). Fluorescence emission microspectrofluorometry has confirmed the presence of high phycocyanin contents in heterocysts of Anabaena isolated from leaf cavities of A. pinnata and A. caroliniana (Kaplan et al., 1986).

Nitrogen metabolism

Nitrogen, though abundantly present in atmosphere, cannot be utilized directly by most organisms. In microorganisms, the enzyme that is responsible for nitrogen fixation is nitrogenase. The enzyme consists of two proteins. The iron protein (Fe-protein, also called dinitrogenase reductase) is encoded by the nifH gene and is a α2 homodimer of about 64kDa. The molybdenum-iron protein (MoFe-protein, also called dinitrogenase) consists of two alpha and two beta subunits. Therefore, it is a α2β2 heterotetramer of about 240kDa, in which the α subunits are encoded by nifD and the β subunits by nifK (Burgess & Lowe, 1996).

Nitrogenase enzyme reduces atmospheric nitrogen into ammonia, utilizable form. Azolla can successfully grow in areas with low nitrogen content due to its symbiotic association with Anabaena azollae that has the ability to fix atmospheric nitrogen. This association makes Azolla an important biofertilizer in many parts of the world. Nitrogen requirements of both Azolla and Anabaena are met as a result of nitrogen fixation carried out in the heterocysts of endophytes. In A. caroliniana, A. pinnata, and A. filiculoides, it has been found that nitrogenase activity increases from zero at shoot apex to almost maximum at leaf number seven (Becking, 1978). Also it has been reported that number of heterocysts has been shown to increase with age of leaf (Hill, 1977; Kaplan et al., 1986). Estimation of nitrogen fixation rates has been done using assays for acetylene reduction, ¹⁵N fixation, and ATP dependent H2 production (Lumpkin & Plucknett, 1982; Watanabe, 1982). Rates of nitrogen fixation are affected by several environmental conditions such as temperature, quantity and quality of light, water turbulence, etc. (Becking & Donze, 1981; Peters & Mayne, 1974; Vincenzini et al., 1985). Also, hydrogenase functions in the recycling of electrons and ATP by oxidizing H2 produced during nitrogen fixation (Peters 1976, 1977). It has been clearly demonstrated using inhibitor DCMU that nitrogen fixation is indirectly dependent on activity of photosystem II, and cyclic photophosphorylation associated with PSI supplies most of ATP for nitrogenase activity (Peters & Mayne, 1974; Tyagi et al., 1980).

Superoxide dismutase (SOD) protects nitrogenase enzyme from damage by harmful reactive oxygen species. Three forms of SOD were found in Anabaena azollae from A. filiculoides of which the Fe-SOD was the major form in both vegetative cells and heterocysts. In vegetative cells, manganese-SOD (Mn-SOD), iron-SOD (Fe- SOD), and hybrid iron-manganese (Hy-SOD) isozymes were found. Of these, the Mn-SOD was not found in heterocysts. The total SOD activity in the heterocysts was found to be 15% lower than that in the vegetative cells. The activity of Fe-SOD in heterocysts was 25% of its activity measured in vegetative cells, while the activity of Hy-SOD in heterocysts was comparable to that in vegetative cells (Canini et al., 1991).

Exposure of Azolla pinnata–Anabaena azollae to NaCl induced a sharp decline in nitrogenase activity. A lethal dose of NaCl (40mM) for the growth of A. pinnata–A. azollae complex could inhibit the nitrogenase activity by 39%, revealing for higher sensitivity of general growth to NaCl than the nitrogenase activity (Rai et al., 1999, 2001). Overall, salt had a higher effect on the growth of the system than on nitrogenase activity, which indicates that there is some protection of the cyanobiont from salt stress within the host leaf cavity. Symbiotically associated Azolla had 5–10 times greater pools of intracellular ammonia as compared to endophyte-free Azolla (Newton & Cavins, 1976). Glutamate, arginine, and aspartate were found to be major amino acid constituents in the nitrogenous pools of the endophyte (Peters et al., 1985). In the intact A. caroliniana association, major constituents were ammonia, glutamine, glutamate, and an unknown amino acid N-γ-L-glutamyl-β-D-aminophenylpropanoic acid (Corbin et al., 1986). In addition to this, Anabaena azollae, isolated from leaf cavities excreted majority of nitrogen as ammonia. Three major ammonia assimilating enzymes such as glutamine synthetase (GS), glutamate dehydrogenase (GDH), and glutamate synthase (GOGAT) have been studied in detail to understand nitrogen metabolism in Azolla–Anabaena symbiosis. It has been reported that GS activity is much lower in symbiotic association when compared to free-living Anabaena (Ray et al., 1978). In another independent study, it was found that activities of GS and both NADH and NADPH-dependent GDH was found to be higher in cavity hairs than in Azolla leaves (Uheda, 1986). Through use of ¹³N and inhibition studies using L-methionine-DL-sulfoxime (MSX), it has been concluded that ammonia assimilation occurs with the help of both GS and GOGAT enzymes without much of significant contribution from GDH or alanine dehydrogenase (Peters et al., 1985). Therefore it can be said that in Anabaena–Azolla association, nitrogen fixed by Anabaena is transferred to Azolla which is then metabolized by GS-GOGAT (the glutamine synthetase-L-glutamine-2-oxoglutarate aminotransferase) pathway. Pulse chase studies using ¹⁵N have clearly shown that fixed nitrogen is transported from mature leaf cavities to stem apex in a time-dependent manner (Kaplan & Peters, 1981). In A caroliniana, glutamate, glutamine, ammonia, and a glutamate derivative are few of those major compounds that are transported from mature leaf cavities to stem apex (Peters et al., 1985). It was also reported that in A. pinnata, nitrate-derived nitrogen was primarily distributed in older branches; however, ammonia or dinitrogen-derived nitrogen was distributed in younger branches (Ito & Watanabe, 1983).

Molecular studies on Anabaena–Azolla symbiosis

In most of the studies, cyanobacterial partner of this symbiotic association has been studied at molecular level to assess the details of the molecular interactions between Anabaena and Azolla. Southern blotting was used to study the arrangement of nifHDK genes of endosymbionts extracted from four different Azolla species and one free living Anabaena azollae isolate, and it was found that arrangement of nifHDK genes are conserved among all Azolla species (Franche & Cohen-Bazire, 1985) and that it is transcribed from one nifHDK operon in the endosymbiont (Nierzwicki-Bauer & Haselkorn, 1986 ). The nif genes are genes encoding enzymes involved in the fixation of atmospheric nitrogen into a form of available nitrogen to living organisms and are found in both free-living nitrogen-fixing bacteria and in symbiotic bacteria associated with various plants.

In another study, transcript levels of enzymes either involved in nitrogen fixation or heterocyst differentiation were quantified using Northern blot and dot blot hybridization. mRNA levels of corresponding genes in Anabaena endosymbiont were identified by using cloned genes from Anabaena 7120 for enzymes such as glutamine synthetase, RUBISCO, nitrogenase and 32 kd protein of photosystem II (Nierzwicki-Bauer & Haselkorn, 1986). Glutamine synthetase is one of key enzymes in Anabaena–Azolla association and is coded by Anabaena glnA gene. It was found that GS mRNA transcript levels were tenfold lower in symbiotic versus free living Anabaena (Nierzwicki-Bauer & Haselkorn, 1986). Another 32kDa protein required for PS II quinone–mediated electron transfer is encoded by psbA gene. Transcript levels of 32kDa protein were approximately 7–10 times more abundant in endosymbiont than in free-living Anabaena (Arntzen et al., 1982; Nierzwicki-Bauer & Haselkorn, 1986). Transcript level of RUBP carboxylase was also measured using carboxylase-specific probe. Transcript levels were five to seven times less abundant in symbiont than in free living Anabaena which is consistent with the reduced CO2 fixation reported for the symbiont (Nierzwicki-Bauer & Haselkorn, 1986).

Economic importance of Azolla

Azolla was mostly ignored or neglected for centuries, except in some parts of China and Vietnam where it was utilized as green manure and as animal feed. However after the first world energy crisis, Azolla has gained importance among the international scientific community and as a consequence during the 1970s and 1980s there had been a tremendous increase in research effort and publications in this field. Azolla is indeed rich in protein and has a reasonably good amino-acid balance (Sanginea & Van Hove, 1989) and a large number of mineral elements, vitamins, chlorophylls, carotenoids, and amino acids (Kamalasanana Pillai et al., 2002) and is therefore consumed as fresh, dried or ensiled material by animals. Research has shown that Azolla has good potential in controlling mosquito proliferation (Amerasinghe & Kulasooriya, 1986; Rajendran & Reuben, 1991; Smith, 1910). Azolla has been reported to be used for reclaiming saline soils (Ge et al., 1980; Gu, 1983), waste water treatment (Shiomi & Kitoh, 1986). Azolla has been shown to accumulate phosphorus (Reddy & De Busk, 1984; Shiomi & Kitoh, 1987) as well as some heavy metals (de Wet et al., 1990; Sarkar & Jana, 1986) and has therefore been proposed as a water purifier. Azolla has been extensively used as nitrogen fertilizer (Kumarasinghe & Eskew, 1993; Vlek et al., 1992) in especially paddy cultivation thus reducing cost of cultivation and increased paddy yield. Azolla is a great feed for livestock such as poultry, pigs, sheep, goat, fish, etc. Azolla when incorporated in fields decomposes and forms humuslike compound. Humus increases the water holding capacity and promotes aeration. It also helps in suppression of growth of certain aquatic weeds.

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Chapter 2: Cyanobacterial symbiosis with bryophytes

D. Jayanthi ¹ , S.J. Meghana ² , Harita G. Rao ³ , and S. Shravya ⁴       ¹ Department of Botany, Mount Carmel College (Autonomous), Bengaluru, Karnataka, India      ² Department of Biology, Sindhi PU College, Bengaluru, Karnataka, India      ³ Department of Biology, Abheek Academy, Bengaluru, Karnataka, India      ⁴ Clinical research, IQVIA, Bengaluru, Karnataka, India

Abstract

Various eukaryotic organisms have been host to cyanobacteria which fulfill the activity of fixing nitrogen for their hosts. Their adaptability to various conditions in the host and ability to change the host metabolism makes them one of the most efficient symbionts in the living world. Various cyanobacteria like Nostoc, which is the predominant symbiotic genus, produce special nitrogen-fixing cells called heterocysts and motile filaments called hormogonia which give them the ability to fix nitrogen in their hosts. With regards to bryophytes, cyanobacteria like Nostoc, Blasia, and Cavicularia exhibit symbiotic associations. Efficient infection processes and molecular interactions between the host and the cyanobacteria are seen in liverworts, hornworts, and Mosses. Several changes have been noticed in the thalli of the bryophytes during and post the infection processes. Deficiency of nitrogen, production of hormogonia inducing factors, cell signaling and production of plant exudates, entry of the cyanobiont, regulation of cell division and production of hormogonia repressing factor, followed by assimilation of nitrogen are intricately designed molecular processes. Study of several genes and operons like the Hrm operon, sigH and ctpH genes, tprN gene, etc., has given insights to understand these molecular processes. These processes have widely been studied in various bryophytes but specifically more so in the Anthoceros–Nostoc association. Although a complete conclusion is yet to be obtained on several factors regarding molecular interactions, the available information has been used to summarize in the following chapter.

Keywords

Bryophyte; Cyanobacteria; Gene; Hormogonia; Infection; Nitrogen-fixation; Operon

Introduction to cyanobacterial symbiosis

Cyanobacterial symbiosis in the plant kingdom

Cyanobacterial symbiosis in bryophytes

Cyanobacterial symbiosis in liverworts

Infection process of cyanobacteria in liverwort and molecular interaction

Conclusion

Cyanobacterial symbiosis in hornworts

Infection of symbionts

Signaling between hornworts and cyanobionts

Molecular basis of interaction

The hrm operon

sigH and ctpH

tprN

ntcA, hetR, and hetF

Cyanobacterial symbiosis in mosses

Introduction

Cyanobacteria and mosses

Role of mosses and cyanobacterial communities in their habitat

Molecular interactions

References

Further reading

Introduction to cyanobacterial symbiosis

Plants, fungi, sponges, and protists have been eukaryotic hosts to various species of cyanobionts. Cyanobionts generally provide the function of nitrogen fixation in photosynthetic hosts, while they also perform the function of fixing carbon in nonphotosynthetic hosts (Adams & Duggan, 2008). Cyanobacteria, though a large group of microorganisms, only a few genera show symbiotic capacity. Cyanobacteria must have a feature of extreme adaptability if they have to be symbiotically capable. They must be able to adapt to the conditions of the host while being able to modify their metabolism so that they can exist in a symbiotic state, actively exchanging metabolites with the host (Rasmussen & Nilsson, 2002).

Cyanobionts in association with plants have two characteristic features: one—to form specialized nitrogen fixing cells called heterocysts and two—to form motile filaments called+hormogonia. These hormogonia do not have heterocysts and act as a means of movement for immobile cyanobacteria. Also, they must exist in a free-living state, accessible within the territory of the host (Adams & Duggan, 2008). Hormogonia are the most common infective agents, and perhaps eukaryotic host plants produce certain chemical signals that cause hormogonia to form. They perhaps secrete chemoattractants to guide the cyanobiont to the plant (Adams & Duggan, 2008). Although many cyanobacteria produce hormogonia, only a few genera have the capability to share symbiotic relationship with the host (Rasmussen & Nilsson, 2002).

In terrestrial ecosystems, Nostoc is the most predominant genus that exhibits a symbiotic relationship with fungi, bryophytes, and some genus like Azolla, Gunnera, and also cycads (Rasmussen & Nilsson, 2002). In aquatic ecosystems, diatoms, sponges, and dinoflagellates exhibit symbiotic associations with cyanobacteria (Carpenter & Foster, 2002).

Cyanobacterial symbiosis in the plant kingdom

All plant parts like roots, stems, leaves are involved in cyanobacterial–plant symbiosis. Among the bryophytes, 6 genera out of about 340 show endosymbiotic cyanobacteria, mainly the hornworts and liverworts. Among the pteridophytes, Azolla shows evidence of cyanobacterial symbiosis with 7 species. About 150 cycads show the presence of diazotrophic cyanobacteria, making them a unique division among the gymnosperms. Gunneraceae, a monogeneric family, is the only one to show cyanobacterial symbiosis among the angiosperms. About 7 species of sub-marine lichens exhibit cyanobacterial symbiosis, e.g., Chroococcus sp. in association with ascolichen Halographis runica; Hyella caespitosa with Arthropyrenia halodytes. In the cycads, bryophytes, and Gunnera, Nostoc is found to be the most predominant genus. Several cyanobacterial genera have been found in lichen symbionts. Anabaena and Calothrix have been found in cycads in a few investigations. Calothrix and Chlorogloeopsis have shown their presence in a few bryophytes in a few investigations (Table 2.1) (Bergman et al., 1992; Rasmussen & Nilsson, 2002) .

The fact that Nostoc is the most widely distributed cyanobacterial genus prevalent in terrestrial ecosystems could be the reason for several plants to prefer it as a symbiotic partner (Rasmussen & Nilsson, 2002).

Cyanobacterial symbiosis in bryophytes

Of the bryophytes, which are composed of liverworts, hornworts, and mosses, relatively, a very small number of genera show epiphytic or endophytic cyanobacterial symbiotic relationships. Bryophytes are the only plant group where cyanobacterial symbiosis occurs in temperate climates. Here, the symbionts infect existing plant structures. For example, in Blasia, the symbiont occupies the auricles on the ventral surface of the thallus (Rasmussen & Nilsson, 2002).

Table 2.1

Usually, nitrogen fixing bryophytes form small gametophytic thalli to a few centimeters in length. The cyanobionts are seen as small dark dots about 1mm in diameter, spread all over the thalli (Rasmussen & Nilsson, 2002).

Both epiphytic and endophytic associations require the cyanobacteria to attach to the host surface. In some associations, pili and extracellular polysaccharides have triggered the cyanobacterial association, though this might not be the case in bryophytes (Rasmussen & Nilsson, 2002).

In liverworts, cyanobacterial associations are rare, being restricted to only 4 out of more than 340 genera. Out of these, 2 associations are epiphytic and 2 are endophytic. In hornworts, about 13 genera show endophytic cyanobacterial associations. In mosses, cyanobacteria grow epiphytically barring two Sphagnum species in which they occupy water-filled hyaline cells, which may provide protection from the acidic environment (Rasmussen & Nilsson, 2002).

Among the liverworts Marchantia and Porella show epiphytic associations, Blasia and Cavicularia show endophytic association (Rasmussen & Nilsson, 2002).

Endophytic associations have been well investigated due to their better adaptability to laboratory conditions. Epiphytic associations remain relatively poorly understood. These associations may be more common than thought (Rasmussen & Nilsson, 2002).

Anthoceros and Phaeceros can be successfully grown in axenic shaken liquid cultures and be reinfected with pure cyanobacterial cultures. Results obtained by Costa et al. (2001) showed that two bryophytes were infected by different Nostoc strains and a single thallus could be infected by a single or multiple strains, after tRNA Leu (UAA) intron sequence analysis (Rasmussen & Nilsson, 2002).

Cyanobacterial symbiosis in liverworts

The liverworts like Jungermanniaceae species have leaflike appendages which are photosynthetically active. They arise from the root hair like rhizoids to stem, whereas they gain nutrients, minerals, and water from the substrate (Carella & Schornack, 2018).

In some liverworts which pose a thallus showcase dense morphology which generally composes the whole plant body. The dorsal surface of the thallus has a cell layer which is photosynthetically active, a nonphotosynthetic storage space in the center, and the rhizoids will arise from the ventral surface which is accompanied by the other structures. Marchantia polymorpha and other liverworts contain an air chamber on the dorsal surface where the interchange/trading of gas takes place through the intercellular space present between synthetic filament cells and moves out from the open pore (Carella & Schornack, 2018).

Liverworts exhibit an endophytic symbiosis; whereas the colonies of Nostoc punctiforme cyanobacterium is noted to be located on the ventral surface of Blasia and Cavicularia species (Carella & Schornack, 2018). .

From more than 340 genera, liverworts’ association with cyanobacteria is much lesser. The four main genera are Marchantia, Porella exhibiting epiphytic association and Blasia, Cavicularia exhibiting endophytic association. Hormogonia formation and symbionts is same as hornworts (Rai et al., 2002).

The most observed cyanobacterial symbionts are the members of the genus Nostoc in liverworts. But other cyanobacteria like Calothrix and Chlorogloeopsis develop hormogonia and may infect the liverworts while experimenting in laboratory (West & Adams, 1997).

Infection process of cyanobacteria in liverwort and molecular interaction

Cyanobacteria infects the host by producing the hormogonia on the host through chemotaxis. During the chemotaxis the HIF is released when the liverwort Blasia is deprived of combined nitrogen and also it releases a chemoattractant which is effective. After the formation of hormogonia, it invades the host Blasia through auricles losing the motility and heterocyst is differentiated once it enters (Adams & Duggan, 2008).

In Blasia, the mutation in gene cyaC influences the efficiency of infection by N. punctiforme. A mutant of N. punctiforme which is antibiotic-resistant infects Blasia at approximately 25% of wild-type frequency. The branched multicellular filaments are elaborated in the walls of symbiotic cavity in Blasia post-infection (Adams & Duggan, 2008).

The molecular interactions in liverworts are observed to be similar to that of Hornworts. However, the detailed information on the molecular interaction in liverworts is not specified or not defined.

Conclusion

From the information collected from the past years reports on cyanobacterial symbiosis in liverworts, it is known that the symbiosis process is very likely to be similar to hornworts. The only exception is that the site of infection in both is different. To further report on the interactions at molecular level, activities of the molecules in liverworts are not defined in the works done previously. It explains the immediate need of exploration in this area.

Cyanobacterial symbiosis in hornworts

Four hornwort genera, Anthoceros, Phaeceros, Notothylas, and Dendroceros show endophytic cyanobacterial symbiotic association (Adams, 2002).

Infection of symbionts

In hornworts Anthoceros and Phaeceros, which have a thicker thallus in comparison to the liverworts, the symbionts are found within the thallus, in slime cavities which open to the ventral side via narrow, slitlike pores. The symbionts enter the slime cavities as hormogonia, after which they lose their motility and differentiate to form heterocysts (Adams, 2002).

The colonies are 0.5–1.0mm in diameter. In Anthoceros, studies have suggested that these cavities are microaerobic, because of which mutant Nostoc ATCC29133 with a defective heterocyst is unable to fix nitrogen under aerobic conditions (Adams, 2002).

Several morphological changes have been observed in the thallus of Anthoceros post-infection. Several, branched multicellular filaments grow from the wall of the host and ramify within the cyanobacterial colony, causing increase in surface area between both the partners (Adams, 2002).

Physiological changes include weakening of cell junctions. In Nostocaceae and Stignomataceae, rapid cell division occurs in the absence of DNA replication and cell growth, leading to heterocysts and hormogonia formation. The formation of hormogonia can be triggered by unidentified plant exudates (Adams, 2002).

Heterocysts enable nitrogen fixation in both symbiotic partners and motile hormogonia enable cyanobacterial entry into the plant (Adams, 2002).

Many cyanobacteria have shown higher rates of heterocyst formations in symbiotic associations than in a free-living state. Higher rates of formation of heterocysts in symbiotic associations with bryophytes have been difficult to understand, as they lose their shape and thick walls making them difficult to be differentiated from vegetative cells of the thallus, which are large and irregular. This maybe because the cells of the symbiont lack murein layer (Adams, 2002).

Hormogonia move by gliding, the mechanism of which remains unknown. In Nostocaceae, the hormogonia filaments are smaller than the vegetative filaments, do not contain heterocysts, and have a unique cell shape unlike members of Oscillatoraceae members where the hormogonia filaments resemble the original trichome (Adams, 2002).

Reductive cell division in Nostoc characterizing small-celled hormogonia might explain the fact that effectiveness of infection requires the hormogonial filament to be motile, be chemotactic, and form hormogonia in the presence of plant chemoattractant exudates (Adams, 2002).

Hormogonia may be the most effective cyanobacterial symbiosis inducing agents, although it has been studied that hormogonia formation does not necessarily induce symbiosis (Adams, 2002). Eg: Nostoc ATCC 27896, which produces hormogonia in the presence of Anthoceros plant exudates, rarely infects the thallus. If infection occurs, nitrogen fixation does occur but the infection does not persist and new tissues are not inhabited. This may be due to the failure to recognize Anthoceros exudates (Adams, 2002).

Signaling between hornworts and cyanobionts

For effective infection, it is necessary that the host must be able to induce hormogonia formation within its locally available cyanobacterial population. Host plants produce chemical exudates for the same purpose (Adams, 2002). For example: Anthoceros punctatus produces a low molecular mass product—a hormogonia inducing factor: HIF that induces hormogonia formation in Nostoc. This HIF is not produced in vitro when Anthoceros is cultured on a medium containing excess ammonium, indicating that HIFs are produced only when there is a deficiency of nitrogen. The composition of HIF remains unknown, although similar compounds have been found in exudates of Gunnera stem gland, wheat root, Blasia (Adams, 2002).

The fact that the entry into the slime cavity is narrow and that the cavities are small indicate that these infections are not obvious. A chemoattractant is required to induce symbiosis. It is seen that formation of fingerlike transfer cells in Anthoceros may result from the signals derived from cyanobacteria (Adams, 2002).

Arabinogalactan proteins (AGPs) in the slime cavity of Phaeoceros strain, flavonoids like naringin triggering hormogonia formation in Nostoc are few compounds that take part in signaling between bryophytes and cyanobacteria (Adams, 2002).

For establishing a stable symbiosis, it is important that the rate of cell division between the host and the cyanobiont must be at par. Evidence has however shown that the doubling time in symbiotic strains is much higher compared to the free-living strains of the cyanobacteria. The mechanism of this phenomenon is unknown although this might indicate that if primary cell division of the cyanobiont is controlled by the host, thereby reducing the nutrient availability to the cyanobacteria, the host can use the excess nutrients for itself (Adams, 2002).

The host also has shown to exhibit control over the biomass of the cyanobacteria. If fresh growth of Nostoc colonies is prevented in-vitro for a Anthoceros–Nostoc association, it is seen that existing colonies increase in size and show higher nitrogen fixing rates. However, if ammonium is added to the medium, there is decrease in the heterocyst formation and nitrogen fixing rate (Adams, 2002).

The presence of ammonia, however, does not affect the rate of infection. The effect is that colonies remain small, show less frequent heterocysts, and lower rates of dinitrogen fixation, indicating that the nitrogen status of Anthoceros regulates the rate of heterocyst formation and nitrogen fixation (Adams, 2002).

It has been studied that ammonium actually does not inhibit heterocyst formation. When methionine sulfoximine, an inhibitor to glutamine synthase, was added to a medium containing ammonium chloride growing Anabaena cylindrica, the inhibition of GS activity prevented assimilation of ammonia, resulting in nitrogen deficiency and heterocysts were formed even in the presence of high levels of ammonia (Adams, 2002).

Apart from regulating growth rate, the host should also control hormogonia formation. Post-infection, hormogonia formation must be prevented as they do not form symbiotically viable colonies. A hormone repressing factor (HRF) has been identified in the aqueous extracts of Anthoceros tissue which inhibits HIF-induced hormogonia formation in wild-type N. punctiforme. This ceases hormogonium formation probably by the production of an inhibitor or catabolism of an activator of the hrmUA operon, whose gene products were found in the above mentioned Anthoceros extract (Adams, 2002).

Dinitrogen fixation has been observed to be much higher in symbiotic cyanobacteria than in their free-living state. A 4-35-fold increase in nitrogen fixation has been observed in Anabaena–Nostoc relationship than in free living Nostoc. Nitrogen fixed by Nostoc is released as ammonia to Anthoceros. About 20% of the fixed nitrogen is retained by the cyanobiont. The initial assimilation of released ammonia takes place by the GS GOGAT pathway in the host. Further transport mechanism into the bryophyte tissue remains unknown (Adams, 2002).

The release of ammonia by the cyanobiont occurs due to decrease in activity of glutamine synthase (GS), the first enzyme of the GS GOGAT pathway. The amount of GS present in filaments of free-living cyanobacteria and their symbiotic counterparts is observed to be different (Table 2.2) (Adams, 2002).

In spite of supplying up to 80% of fixed nitrogen to the host, the cyanobacteria do not experience nitrogen starvation, but retain two nitrogen reserves, the phycobiliproteins and cyanophycin, which are broken down in the case of nitrogen starvation (Adams, 2002).

Carbon dioxide fixation primarily occurs by the Calvin cycle, using ribulose-1, 5-bisphosphate carboxylase/oxygenase (Rubisco) as the main carboxylating enzyme. Like dinitrogen fixation, carbon dioxide fixation is observed to be less in free-living state than in a symbiotic state. The amounts of Rubisco are also different in both the cases indicating that a post-translational modification takes place to Rubisco, the mechanism being unknown. The cyanobiont receives fixed carbon from its photosynthetic host, thus living photoheterotrophically. In Anthoceros–Nostoc associations, the cyanobiont is observed to have glycogen granules that are used in the case of carbon starvation (Adams, 2002) (Fig. 2.1).

Molecular basis of interaction

Development of recent genetic techniques, like transposon mutagenesis, has been used by Meeks and coworkers (2003) to identify genes involved in infection of Anthoceros–Nostoc association. Nostoc punctiforme strain ATCC 29133 has been analyzed to understand symbiotic associations with Anthoceros (Adams, 2002).

Table 2.2

a  While mostly Nostoc infects liverworts, evidences for Calothrix spp. have also been found.

Figure 2.1  Flow chart to depict the mechanism of infection and post-infection changes in cyanobacteria–bryophyte symbiosis: References to Anthoceros–Nostoc symbiosis.

The hrm operon

Transposon mutagenesis was used to generate mutants of Nostoc ATCC 29133 which showed high rates of initial infection in Anthoceros. Nostoc ATCC 29133 UCD 328 showed a higher frequency of hormogonia in the presence of A. punctatus and a eightfold increase in infection.

These open reading frames—hrmU and hrmA—were identified flanking the site of transposition. hrmU was found to have close similarity to NAD(P)H oxidoreductases while hrmA showed no similarity to sequences in databases. hrmA was expressed in the presence of an aqueous extract of A. punctatus but not by the hormogonia inducing factor HIF. This extract also suppresses hormogonia formation induced by HIF in the wild type of the cyanobiont thus suggesting that it might contain a hormogonia repressing factor HRF (Adams, 2002).

The HRF is released into the cavity, post-infection to suppress hormogonia formation, thereby permitting the formation of heterocysts (Adams, 2002).

ORFs hrmI, hrmR, and orfK were also identified at the 5′ of hrmUA. All of these genes show similarity to genes encoding proteins for gluconate and glucuronate metabolism. However, the significance of these genes in hormogonia formation remains unknown (Adams, 2002).

sigH and ctpH

Gene sigH encoding an alternative sigma subunit of RNA polymerase was identified in Nostoc ATCC 29133 using Anabaena PCC 7120 sigB gene as a heterologous probe (Adams, 2002).

Transcription of this gene is induced by Anthoceros HIF. The 5′ of sigH is gene ctpH that encodes a protein similar to carboxy-terminal protease of Synechocystis PCC 6803, a cyanobacterium (Adams, 2002).

In Nostoc ATCC 29133 ctpH is not transcribed under vegetative conditions, and transcription occurs upon Anthoceros HIF induction only, unlike in Synechocystis 6803 where the gene is required for processing the D1 protein of carboxy-terminal portion of photosystem II. The implication of this, however, has not been well understood yet (Adams, 2002).

tprN

The tprN encodes proteins similar to tetratricopeptide repeat proteins (TPR). This gene lies 3′ of devR which is required for heterocyst maturation. When tprN is inactivated in Nostoc ATCC 29133, no phenotypic effect has been observed in a free-living state, but the mutant colonizes Anthoceros twice in effect when compared to the wild type. The significance of this process of infection is unknown (Adams, 2002).

This tprN gene although transcribed in the vegetative growth stage, the level of transcription increases on exposure to HIF and HRF (Adams, 2002).

ntcA, hetR, and hetF

NtcA regulates the transcription of several genes involved in heterocyst development including hetR which is the first gene, specific to heterocysts to be expressed in the case of nitrogen scarcity, also acting as a primary activator for the development of heterocysts in cyanobacteria. The hetR protein is found specifically gathered in developing heterocysts of Nostoc punctiforme, but accumulates in all cells in a hetF mutant, which cannot develop heterocysts (Adams, 2002).

HetF protein is hence observed to be an activator of differentiation of heterocysts, increasing hetR transcription leading to accumulation of hetR protein in developing heterocysts (Adams, 2002).

In Nostoc punctiforme (Nostoc ATCC 29133), in case of mutation of any of the above three genes, the cyanobacterium is unable to differentiate heterocysts. HetR and hetF mutants have a similar infection frequency in comparison to the wild type, but they cannot support plant growth due to the lack of capability to fix dinitrogen. The ntcA mutant forms hormogonia at about 05%–15% of the wild-type frequency but fails to infect Anthoceros (Adams, 2002).

Cyanobacterial symbiosis in mosses

Introduction

Bryophytes are recognized as the oldest living land plants evolved from 470 Mya. The class Bryophyta (mosses) is the largest group of bryophytes constituting about 10,000 species (Warshan, 2017). The class Bryophyta (Mosses) is further divided into sub-classes—(a) Sphagnidae (sphagnum or peat mosses) (b) Andreaeidae (granite or rock mosses) (c) Bryidae (true mosses). The classes Hepatophyta (liverworts) and Anthocerophyta (hornworts) have true symbiotic associations between cyanobacteria and bryophytes. The cyanobacterial associations with mosses were discovered but special symbiotic structures in comparison with liverworts and hornworts were not described (Solheim & Zielke, 2002).

Cyanobacteria form a rare broad range of symbiotic associations with hosts that include protists, sponges, fungi, and plants (Warshan, 2017). These cyanobacteria are photoautotrophs, nitrogen fixers, and facultative heterotrophs. They help providing the nonphotosynthetic hosts with carbon and nitrogen (Adams et al., 2013).

Cyanobacteria and mosses

Mosses in symbiotic associations with nitrogen fixing cyanobacteria are mostly distributed in all ranges from Arctic tundra to Antarctica and temperate to tropical ecosystems. In habitats where there is low atmospheric nitrogen, the two partners play a vital role in nitrogen and carbon cycles. In boreal ecosystems, the feathermoss and cyanobacterial symbiosis is responsible for the major biological input of the nitrogen into the ecosystem (Warshan, 2017). With the symbiosis between bryophytes and cyanobacteria, the cyanobacterial cells have an extracellular lifestyle and are commonly found as epiphytes on the moss gametophyte or endophytes in the cavities of liverworts and hornworts (Warshan, 2017).

In boreal forests, the common feathermoss species associated with high diversity of cyanobacteria are Pleurozium schreberi, Ptilium crista-castrensis, and Hylocomium splendens. In regions of arctic tundra and subarctic, the cyanobacterial symbiosis with Sphagnum (peat moss) is the main source of nitrogen for these types of ecosystems, where the nitrogen fixation rates exceed the atmospheric nitrogen deposition (Warshan, 2017). These forests receive low amount of nitrogen and adding to it the boreal forests soils have a characteristic low concentration of inorganic nitrogen, low temperature, and low pH that also contribute to the nitrogen limitations (Rousk et al., 2013).

Role of mosses and cyanobacterial communities in their habitat

Mosses play a vital role in the boreal ecosystems because of their contribution to diverse habitat; their effect on temperature, hydrology, and chemistry of soil in the boreal forests. For example, during summer the temperatures of soil below the moss carpets are lower compared to that of the soil without the moss carpets. This leads to the slower decomposition rate below the mosses. Mosses release generous amounts of nutrients like carbon, nitrogen, and phosphorous upon the rewetting of the dried tissue and microbial available nutrients into the soil. Mosses contribute to the biomass and productivity in boreal forests. Feathermoss Pleurozium schreberi accounts for about 70%–100% of the ground cover of the boreal forest ecosystem (Rousk et al., 2013). The abiotic factors include temperature, wind, and exhibiting a high-water retention capacity. Mosses can provide a stable and supportive habitat for the cyanobacteria, increasing the N2 fixation in sites where N fixation is limited (Rousk et al., 2013).

The associations between cyanobacterial colonies and mosses play a vital role for the N-cycle in nitrogen limited boreal ecosystems by contributing about >2kgN ha−1yr−1 by fixation of nitrogen in the N-pool in boreal forest ecosystem. This value is on par with the magnitude of the atmospheric nitrogen deposition