University of Groningen The dynamics of root microbiomes ......salt marsh salt marsh plants rhizosphere community structure of diazotrophs major major (Davis et al., 2011) crop land

University of Groningen

The dynamics of root microbiomes along a salt marsh primary successionWang, Miao

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CHAPTER 1

General Introduction

9

1Studies on microbial diversity have revealed soils as being among the most biologically diverse habitats in our planet (Curtis et al., 2002; Nannipieri et al., 2003; Gans et al., 2005; Hinsinger et al., 2009; Philippot et al., 2013; Saleem and Moe, 2014). Soil microbes represent thus an unseen majority that account for a great proportion of the ge-netic diversity on Earth (Whitman et al., 1998; Torsvik et al., 2002; Garbeva et al., 2004b; Gams, 2007; van der Heijden et al., 2008; Buée et al., 2009; McGuire and Treseder, 2010; Bever et al., 2012), playing key roles in ecosystems and influencing a large number of important eco-system processes, such as nutrient acquisition and cycling and soil for-mation (Högberg et al., 2001; Kowalchuk and Stephen, 2001; Torsvik and Øvreås, 2002; Rillig and Mummey, 2006; Delgado-Baquerizo et al., 2016).

Plants live in close association with the soil microbes and both influ-ence each other (Berendsen et al., 2012). From the perspective of plants, the presence and composition of soil microbial communities have large impacts on plant health, stress tolerance, plant productivity and diver-sity (Saleem et al., 2007; van der Heijden et al., 2008; Buée et al., 2009; Faure et al., 2009; Lambers et al., 2009; Lugtenberg and Kamilova, 2009; Segarra et al., 2009; Chaparro et al., 2012; Zamioudis and Pieterse, 2012; Vogelsang et al., 2013). The final outcome of these plant-microbial in-teractions will depend on the nature of the association — i.e. neutral, deleterious or positive — which might also be dependent on the physi-ological state of the plant or plant species (van Loon and Bakker, 2005; Lugtenberg and Kamilova, 2009; Raaijmakers et al., 2009; Hardoim et al., 2015). In other words, microbial species that are beneficial for a certain plant species might be deleterious or neutral to others, and have their positive influence associated with other factors such as nutrition. From the microbial perspective, the amount and diversity of compounds released by plants as root exudates create a selective environment in the rhizosphere, boosting the activity of surrounding bacteria (Kuzyakov, 2002; Berendsen et al., 2012; Cibichakravarthy et al., 2012), the so-called rhizosphere effect (Smalla et al., 2006; Hartmann et al., 2008; Faure et al., 2009; Compant et al., 2010). The rhizosphere community also represents the source of endophytic bacteria, which cross the root bar-rier and colonize the plant tissues, the root endosphere (Sessitsch et al., 2002; Compant et al., 2005; Hardoim et al., 2008, 2015; Long et al., 2010).

Given the importance of plant-microbial interactions for ecosys-tem functioning — i.e. primary productivity, carbon sequestration and nutrient cycling — in natural ecosystems as well as in agricultural

CHAPTER 1 General Introduction 10

systems (Singh et al. , 2004; Hussain et al. , 2011; Ribeiro and Cardoso, 2012; Ke et al. , 2015; Rodríguez-Blanco et al. , 2015; Yang et al. , 2015; Delgado-Baquerizo et al. , 2016; Thion et al. , 2016), it is crucial to un-derstand the drivers of the plant-associated microbiome (Garbeva et al. , 2004b; Berg and Smalla, 2009; Vandenkoornhuyse et al. , 2015; Thijs et al. , 2016). In this context, soil properties such as texture and pH as well as plant host are known to play important roles in determining the composition and function of root-associated microbial communi-ties (Garbeva et al. , 2004b; Jousset et al. , 2006; Rasche et al. , 2006; Garbeva et al. , 2008; Berg and Smalla, 2009; Sessitsch et al. , 2013; Saleem et al. , 2015; Lareen et al. , 2016).

Plants shape the composition and function of the microbiome in their rhizosphere by releasing root exudates with different composition de-pending on plant species, genotypes, cultivars, growth stages and root structures (Miethling et al., 2000; Berg et al., 2002, 2006; Garbeva et al., 2008; Haichar et al., 2008; Micallef et al., 2009; Inceoglu et al., 2011; Mendes et al., 2013; Pérez-Jaramillo et al., 2016). However, this selec-tive force exerted by plants is different at different locations (Costa et al., 2006a; Salles et al., 2006; Inceoǧlu et al., 2010), given that soil properties such as pH or organic carbon — recognized as major drivers of soil bacte-rial community assembly (Fierer and Jackson, 2006; Lauber et al., 2009; Dini-Andreote et al., 2015; Li et al., 2015) — vary according to soil type (Marschner et al., 2001; Gottel et al., 2011) or land use (Lauber et al., 2008; Shen et al., 2013a). Therefore, the observed differences in plant-associated microbial communities could be due to varying microbial biogeographi-cal patterns across different locations (Martiny et al., 2006; Geremia et al., 2016). Thus, an important but unanswered question is whether plants ex-ert a stronger effect on the selection of plant-associated microbial com-munities than soil physical and chemical properties (Table 1). In this thesis, I will address this question by sampling the same plant species in a gra-dient of soil types, while controlling for environmental conditions and meta-communities. For that I will focus on the salt marsh soil primary succession located on the island of Schiermonnikoog, the Netherlands. Finally, although the soil microbiome encompasses bacteria, fungi, ar-chaea, as well as a suite of non-fungal microbial eukaryotes, all equally important in regulating plant heath, in this thesis I will focus mostly on the bacterial communities. In the following pages, I will briefly review some of the issues introduced above, followed by the general aims of this thesis as well as a short description of the topics studied in each chapter.

11

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1. The plant microbiome

1.1. Definition

Soils are complex assemblages of extremely diverse microhabitats, har-bouring a vast diversity of organisms (Zhou et al., 2004; Hinsinger et al., 2009). It has been estimated that one gram of soil contains as many as 10 billion bacteria (Rossello-Mora and Amann, 2001; Horner-Devine et al., 2003). One of the most fascinating hot spots of activity and di-versity in soils is the rhizosphere (Sorensen and Sessitsch, 2007; Jones and Hinsinger, 2008; Hinsinger et al., 2009; Mendes et al., 2013), best defined as the volume of soil around living roots, which is influenced by root activity according to the “rhizosphere effect” first described by Hiltner (1904). The rhizosphere represents a highly dynamic front for interactions between roots and pathogenic and beneficial soil microbes, invertebrates, and root systems of competitors (Hirsch et al., 2003).

The majority of rhizobacteria derives from the soil environment (Compant et al., 2010). Rhizosphere-associated microbes can also pene-trate plant roots, thus becoming endophytic bacteria, which colonize the plant tissue in lower cell density than the rhizosphere (Compant et al., 2005; Hardoim et al., 2008; Faure et al., 2009; Compant et al., 2010; Gottel et al., 2011). The term endophyte refers to the microflora, com-posed mostly of bacteria and fungi (Hallmann et al., 1997; Faeth and Fagan, 2002; Rosenblueth and Martínez-Romero, 2006), which actively or pas-sively colonizes the internal plant parts (endosphere) without conferring deleterious effect to the host plant (Gray and Smith, 2005). Distributions of plant-root associated bacteria were described in Figure 1.

1.2. Colonization mechanisms

Microbial colonization of the root surface is initiated by the microbial response to the compounds released by the root (Smalla et al., 2006; Sorensen and Sessitsch, 2007; Hartmann et al., 2008; Faure et al., 2009). Root exudation includes the secretion of ions, free oxygen, water, en-zymes, mucilage, and a diverse array of carbon-containing primary and secondary metabolites (Bertin et al., 2003). The amount and composi-tion of these exudates varies according to ecosystem type, plant species and growth stage of the plant (Hartmann et al., 2008; Inceoğlu et al.,

13

1


2011). The roots can also produce chemical signals that attract microbes and induce chemotaxis (Bais et al., 2006), creating a very selective envi-ronment with lower biodiversity but higher activity compared with the bulk soil (Berendsen et al., 2012; Cibichakravarthy et al., 2012). These selection mechanisms make the rhizosphere an excellent source of spe-cialized rhizobacteria (Barriuso et al., 2005).

Following rhizosphere and rhizoplane (the surface of plant root) colo-nization, soil-borne microorganisms harbouring specific traits required for endophytic competence can enter roots — those are defined as oppor-tunistic or competent endophytes, each showing particular root coloni-zation characteristics (e.g. chemotactic response). For instance, whereas the competent endophytes can be well adapted to the internal plant en-vironment and maintain beneficial associations with plant hosts, the opportunistic endophytes might accidentally become endophytic by en-tering root tissue, even though they lack genes that are key to their eco-logical success inside the plant (Hardoim et al., 2008). Additionally, some soil bacteria might become endophytic by stochasticity (e.g. via coloniza-tion of natural wounds), being considered as passenger endophytes. The colonization of the interior of plant roots by microbes is more advanta-geous, because plant nutrient resources can be explored even more effec-tively without the tough competition often observed in the rhizosphere, due to the high microbial density (Rosenblueth and Martínez-Romero, 2006). Moreover, endophytes are better buffered against abiotic stresses, common in the complex soil environment (Hallmann et al., 1997). The microbiome within plant roots can differ significantly from that in the

Figure 1 Distributions of plant-root associated bacteria


rhizosphere, suggesting that plants exert a selective force on the micro-bial communities found inside their roots (Germida et al., 1998; Garbeva et al., 2001; Gottel et al., 2011), which is expected to be stronger than the selective force exerted by the rhizosphere.

1.3. Plant growth promoting rhizobacteria (PGPR)

Root exudates and mucilage-derived nutrients attract deleterious as well as beneficial and neutral bacteria, fungi and other soil organisms (Walker et al. , 2003). Negative interactions mediated by root exudates involve secretion of antimicrobials, phytotoxins, nematicidal, and in-secticidal compounds; meanwhile, rhizosphere bacteria can also have detrimental effects on plant health and survival through pathogen or parasite infection (Dias et al. , 2008). Contrastingly, it has been widely documented that, in the case of rhizobial and mycorrhizal host inter-actions, the roots secrete secondary metabolites that act as messengers to attract these microorganisms (Besserer et al. , 2006). Positive interac-tions also involve root exudate-mediated interactions with plant growth promoting rhizobacteria (PGPR), which are free-living non-pathogenic beneficial soil rhizobacteria often found near, on or even in the plant tissues, which play a key role in plant health and nutrition (Kloepper et al. , 1980; Lynch, 1990).

PGPR can impact plant growth and development through indirect or direct effects. In the one hand, through indirect mechanisms, they protect the plant from the deleterious effect from plant pathogens via siderophore- mediated competition for iron, antibiosis, or the induction of systemic resistance in the plant host (Garbeva et al., 2004a, 2004b; Sgroy et al., 2009; Ahmed and Holmstrom, 2014). On the other hand, they might be directly involved in promoting plant growth through phos-phate solubilisation, the production of molecules associated with stress signalling such as bacterial 1-aminocyclopropane-1-carboxylate (ACC) deaminase, the production of siderophores and indole acetic acid (IAA) (Glick, 1995; Glick et al., 1998; de-Bashan and Bashan, 2004; Kuklinsky-Sobral et al., 2004; Lugtenberg and Kamilova, 2009; Yang et al., 2009; Bal et al., 2013) and supply of biologically fixed nitrogen (Glick, 1995; Glick et al., 1999; Kuklinsky-Sobral et al., 2004; de-Bashan and Bashan, 2004; Rosenblueth and Martínez-Romero, 2006; Lugtenberg and Kamilova, 2009; Yang et al., 2009; Compant et al., 2010; Bal et al., 2013).

15

11.3.1. An exceptional PGPR: the Pseudomonas species

The genus Pseudomonas belongs to the Gamma subclass of the Proteo-bacteria and includes fluorescent pseudomonads as well as a few non- fluorescent species. Pseudomonas is a diverse genus that occupies many different niches and exhibits versatile metabolic capacity (Clarke, 1982; Lugtenberg and Kamilova, 2009; Jain and Das, 2016). A number of pseu-domonad strains function as PGPR, which can protect plants from vari-ous soil borne pathogens and/or stimulate plant growth (Haas and Défago, 2005; Hayat et al., 2010; Jin et al., 2014). Among them, some Pseudomonas putida and Pseudomonas fluorescens strains — also known as fluorescent Pseudomonas species — play a significant role in plant growth- promotion activity by various mechanisms, such as stimulating plant growth by phosphate solubilisation, ACC deaminase synthesis and IAA production (Shaharoona et al., 2008; Ahmad et al., 2013; Timm et al., 2015), or inhib-iting the growth of bacteria and fungi by producing potent iron- binding siderophores, such as the diffusible pigment called pyoverdin (Pvd) or pseudobactin, in addition to a large range of molecules with antimicro-bial activity (Vlassak et al., 1992; Haas and Défago, 2005; Botelho and Mendonça-Hagler, 2006; Bakker et al., 2007; Jain and Das, 2016).

The prevalence of these organisms in the rhizosphere is related to their fast colonization ability (Kamilova et al., 2005) and their capacity to produce secondary metabolites (Thomashow et al., 1990; Gross and Loper, 2009). For example, some P. fluorescens strains can produce pro-teins and secondary metabolites that function as biocontrol factors that kill bacteria, fungi and nematodes, such as 2,4-diacetylphloroglucinol (2,4-DAPG) with antifungal, anthelminthic, and phytotoxic properties (Abbas et al., 2002; Siddiqui and Shahid Shaukat, 2003; Neidig et al., 2011), pyrrolnitrin (Prn) against Rhizoctonia solani and Caenorhabditis ele-gans (Garbeva et al., 2004a; Solanki et al., 2014; Nandi et al., 2015), and hydrogen cyanide (HCN) against Pythium ultimum, Fusarium oxysporum and Caenorhabditis elegans (Ramette et al., 2003; Nandi et al., 2015). For instance, phenotypic characterization of P. fluorescens PCL1751 revealed that it could provide protection against the disease foot and root rot in to-mato caused by the fungal pathogen Fusarium oxysporum (Kamilova et al., 2005) and in cucumber caused by F. solani (Egamberdieva et al., 2011). Moreover, root-associated fluorescent Pseudomonas spp. (e.g. P. fluorescens CHA0) are the key components in biological control of Gaeummanomyces graminis tritici (Ggt), which cause ‘take-all’ disease (TAD) of wheat root



(Raaijmakers and Weller, 1998), being naturally selected by the plant from the pool of soil microorganisms. P. fluorescens CHA0’s effectiveness as a biocontrol agent is mainly determined by the biosynthesis of anti-fungal compounds — 2,4-DAPG, pyoluteorin, and hydrogen cyanide (HCN) (Schnider-Keel and Seematter, 2000; Jousset et al., 2006; Rochat et al., 2010; Raaijmakers and Mazzola, 2012).

The community structure of Pseudomonas and their antagonistic prop-erties against the soil-borne pathogens were reported to be influenced by soil type or management, by showing different community compo-sition and antagonistic activity under different agricultural regimes (Garbeva et al., 2004a). In addition to the soil effect, plant selectivity also exerts a significant influence on the predominance of the rhizo-sphere competence of species within the Pseudomonas genus (Glandorf et al., 1993; Lemanceau et al., 1995; Berg et al., 2002, 2006). By investi-gating sugar beet seedling colonization by Pseudomonas in five different soil types, Schmidt et al. (2004) indicated that the population density of P. fluorescens B5 on the root was rather stable than differing according to soil types. Costa et al. (2006b) revealed that the community structure of Pseudomonas in rhizosphere soils was significantly influenced by multi-ple factors including sampling site, plant species and year-to-year varia-tion. To take this one step further, by developing a PCR-DGGE system to target Pseudomonas-specific gacA gene fragments, the rhizosphere effects on Pseudomonas gacA gene composition and both plant- and site-specific gacA gene structure in rhizosphere and bulk soils were respectively de-tected (Costa et al., 2007).

Specifically, in salt marsh ecosystems, Pseudomonas spp. associ-ated with plants have shown the potential to assist phytoremediation by acting as plant growth promoters and as environmental detoxifiers, through mechanisms of promoting plant growth, enhancing the poly-cyclic aromatic hydrocarbon (PAH) degradation, increasing disease re-sistance and metal tolerance (Aguilar-Barajas et al., 2010; Diab and Din, 2013; Rocha et al., 2016).

1.4. The plant microbiome and nitrogen cycling

Plant-associated microbes play an important role in nutrient cycling in agricultural and natural ecosystems (Walker et al., 2008; Wartiainen et al., 2008; Hayden et al., 2010; Levy-Booth and Winder, 2010; Romero

17

1et al., 2012; Chaudhary et al., 2015). Microbial communities associated with plant roots are responsible for key nitrogen transformations such as N2 fixation, nitrification and denitrification (Figure 2). In this study, we will focus on nitrogen fixation and nitrification processes. The biolog-ical fixation of nitrogen (BNF) is an important source of ‘new’ nitrogen in oligotrophic ecosystems, such as mangrove forests and salt marshes (Lee and Joye, 2006; Purvaja et al., 2008; Fan et al., 2015). Nitrification is a major pathway in the N cycle, playing an important role in plant up-take of inorganic N, emission of nitrous oxide, and nitrate leaching to surface and groundwater, particularly in coastal and estuarine ecosys-tems (Wankel et al., 2011; Zhalnina et al., 2012).

As salt marshes are often limited in available nitrogen, phosphorus and sulfur compounds, microbial activities in nutrient stabilization and transformation (e.g. biological nitrogen fixation, phosphate solubiliza-tion and sulphate reduction) have been thought to influence the devel-opment of plant zonation (Gayathri et al., 2010). Moreover, microbial communities associated with salt marsh plants are known to modu-late the above- and belowground interactions by increasing the cycling of organic matter and nutrients, thus promoting the succession of salt

Figure 2 N-cycling processes of plant-root associated bacteria



marshes (e.g. Piceno and Lovell, 2000a, 2000b; Caravaca et al., 2005; Davis et al., 2011; Lovell and Davis, 2012; Zhang et al., 2013). Therefore, understanding the microbial community structure, diversity and func-tions is necessary to assess the ecosystem situation and provides useful information of predictions on the development of salt marshes.

1.4.1. Nitrogen fixation:

Performed by either free-living or symbiotic microbes, nitrogen fixa-tion contributes to a significant portion of N increments across diverse environments, such as marine, alpine, and agricultural ecosystems (Galloway et al. , 1995; Karl et al. , 2002; Zhang et al. , 2006; Roesch et al. , 2008; Reardon et al. , 2014). The capacity for N-fixation is unique to cer-tain groups of bacteria and archaea that contain the highly-conserved gene nifH, which encodes the iron protein subunit of nitrogenase (Zehr et al. , 2003; Raymond et al. , 2004). Nitrogen fixation by diazotrophic endophytes has also been observed in a wide variety of plants (e.g. James, 2000; Gupta et al. , 2013; Madhaiyan et al. , 2013; Videira et al. , 2013). In terms of benefiting plants by fixing atmospheric nitrogen, en-dophytic bacteria are considered to be ‘better’ than rhizosphere bacte-ria, as they provide fixed nitrogen directly to host plants (Sturz et al. , 2000), and are better placed to exploit carbon substrates supplied by the plant (Taylor et al. , 2010). Moreover, they encounter the microaero-philic conditions needed for nitrogenase activity. By isolation followed by phylogenetic analysis and nifH gene examination, many nitrogen- fixing bacterial strains in the endosphere have been characterized (Tejera et al. , 2003; Terakado-Tonooka et al. , 2012; Gupta et al. , 2013; Madhaiyan et al. , 2013).

Given that diazotrophs are regulated by the concentration of nitro-gen forms and their enzyme activities are influenced by soil character-istics, the diazotrophic community activity, size and structure in soils have been reported to relate to soil physical and chemical properties, such as water content, texture, pH, carbon (C) and N quantity and avail-ability (Hsu and Buckley 2009; Hayden et al. 2010; Wakelin et al. 2010). By characterizing the structure and dynamic changes in diazotrophic communities, based on the nifH gene, across eight different representa-tive Dutch soils, Pereira e Silva et al. (2011) indicated that soil type (sandy and clay soils) was the main factor influencing the N-fixing communities,

19

1being more abundant and diverse in the clay than in the sandy soils. They also suggested that the diazotrophic communities associated with clay soils might be more sensitive to fluctuations associated with the season and agricultural practices. However, in some cases nitrogen fertilization might have no impact on the abundance of root-associated diazotrophs, as observed for sugarcane roots (Yeoh et al., 2015). Overall, both plant species and phenology significantly influence the size and structure of diazotrophs (Engelhard et al. 2000; Tan et al. 2003; Knauth et al. 2005; Roesch et al. 2006; Dias et al. 2012).

BNF in salt marshes is thought to be more important than in most other ecosystems due to its significant contribution to the nitrogen budget in these systems (Capone and Carpenter, 1982; Fan et al., 2015). Diazotrophs associated with salt marsh grasses are present in high numbers on the roots of these plants, where they are highly active (Whiting et al., 1986), being supported by root exudates, as well as by products of plant decomposition (Lovell et al., 2000; Piceno and Lovell, 2000a, 2000b; Larocque et al., 2004). The salt marsh diazotroph assem-blages include organisms that are quite specific for a given marsh mi-croenvironment as well as for specific marsh plant hosts on a local scale (Berg and Smalla, 2009; Lovell and Davis, 2012). Along the salt marsh succession, the dynamics of diazotrophic communities and the struc-ture of the population of nifH genes were mostly driven by shifts in the soil physical structure, as well as variations in pH and salinity (Dini-Andreote et al., 2014, 2016). Moreover, the community structure of ni-trogen fixers was also observed to follow the variation of plant diversity across the succession (Davis et al., 2011; Salles et al., 2017). This proved that rhizosphere-associated diazotrophs were also driven by host speci-ficity, besides the effects of seasonal and elevational changes. Strong ev-idence thus suggests that salt marsh diazotrophs are affected by plant host and that the biogeography of the diazotrophs can be well predicted by plant hosts and edaphic factors varying in response to elevation, as reviewed by Lovell and Davis (2012).

1.4.2. Nitrification:

A rate-limiting step in soil nitrification is the oxidation of ammonia to nitrite (Leininger et al., 2006; Nicol and Schleper, 2006; Jackson et al., 2008; Prosser and Nicol, 2008). This step is performed by organisms



known as ammonia-oxidizing bacteria (AOB) and archaea (AOA) (Prosser and Nicol, 2012). The chemolithotrophic AOB commonly belong to the Beta- and Gammaproteobacteria, including Nitrosomonas (Beta), Nitrosospira (Beta), and Nitrosococcus (Gamma) (Madigan et al., 2000). The discovery of AOA in natural and engineered systems demonstrates that members of the kingdom Crenarchaeota, within the archaeal do-main, play an important role in nitrification in soils and aquatic systems (Könneke et al., 2005; Hansel et al., 2008; Tourna et al., 2008).

Rhizosphere oxygenation can creates a microaerobic environment around plant roots in otherwise anaerobic soils to promote nitrifica-tion, next to denitrification (You et al. , 2009). Studies have demon-strated that AOA dominate the ammonia-oxidizing community in the rhizosphere of Littorella uniflora (Herrmann et al. , 2008). The pre-dominance of amoA-AOA over amoA-AOB has also been reported in a range of environments, including terrestrial (Leininger et al. , 2006), semi-arid agricultural soil (Banning et al. , 2015), hot spring sediment (Hatzenpichler et al. , 2008), coastal ecosystems (Wuchter et al. , 2006) and marine ecosystems (Beman et al. , 2008). However, from other es-tuarine and coastal studies, amoA-AOB outnumbered amoA-AOA (e.g. Caffrey et al. , 2010; Wankel et al. , 2011; Dini-Andreote et al. , 2016). Moreover, in response to N fertilizer amendment, AOA have exhibited less sensitivity compared to AOB, by showing relatively stable com-munity dynamics irrespective of ammonia availability (Santoro et al. , 2008; Glaser et al. , 2010).

Soil properties and environmental conditions, including soil pH, or-ganic carbon content, nutrient availability, water content, and oxygen concentration, CO2 concentration, N fertilization, ammonium and sa-linity levels greatly affect the abundance and diversity of ammonia oxi-dizers (Horz et al., 2004; Nicol et al., 2008; Tourna et al., 2008; Erguder et al., 2009; Prosser and Nicol, 2012). Pereira e Silva et al. (2012b) indi-cated that the nitrifying activity was driven by soil pH, mostly related to its effect on AOA but not on AOB abundances. In addition, clay content was the main soil factor shaping the structure of both the AOA and AOB communities. Moreover, plant species and phenology are also likely to influence the size and structure of ammonia-oxidizers in rhizosphere soils (Fan et al., 2011; Dias et al., 2012). It has been shown that the com-munity structures of AOB and AOA in the rhizosphere vary according to the potato cultivars, which could due to different exudation patterns among potato cultivars (Dias et al., 2012).

21

1Ammonia oxidizers play an importance role in nitrogen cycling in marine ecosystems, and the wide distribution of AOA and AOB has been reported by a number of studies in salt marshes, estuaries, and marine habitats (Wuchter et al., 2006; Mincer et al., 2007; Beman et al., 2008; Brochier-Armanet et al., 2008; Mosier and Francis, 2008; Dini-Andreote et al., 2016; Salles et al., 2017). Previous studies in our system demonstrated that the community structure of ammonia oxidizers fol-lowed mainly N availability (Salles et al., 2017), besides the potential ef-fects of shifts in the soil physical structure, as well as variations in pH and salinity (Dini-Andreote et al., 2014, 2016).

1.5. Brief overview of the methods used for assessing the plant microbiome

The analytical methods for assessment of the community structure, di-versity and functions of rhizospheric and endophytic microbes have de-veloped significantly in recent decades. Culture-dependent techniques are biased due to the Great Plate Count Anomaly (Staley, 1985). Easily isolated organisms are the ‘weeds’ of the microbial world and are esti-mated to constitute less than 1% of all microbial species (Hugenholtz, 2002). Culture-dependent methods, however, are very important to un-derstand the physiology of microorganisms and thus culture- dependent and culture-independent methods are usually combined when studying the microbial properties (Palaniappan et al., 2010; Souza et al., 2012; Lucas et al., 2013).

Culture independent techniques based on DNA or RNA targeted methods are very diverse and have significantly changed over the past 25 years due to the advances in sequencing technologies. The first technological jump that refreshed microbial ecology was driven by PCR-based fingerprinting techniques such as terminal restriction fragment length polymorphism (T-RFLP) analysis, denaturing gradi-ent gel electrophoresis (DGGE) analysis and temperature gradient gel electrophoresis (TGGE), which were initially employed to overcome the limitations of classical isolation procedures (Smalla et al. , 2001; Lodewyckx et al. , 2002; Zhang and Xu, 2008; Dias et al. , 2011; Ali et al. , 2014). These methods, however, were limited in the amount of infor-mation generated — restricted to a small percentage of the dominant groups — and the phylogenetic affiliations of the taxa, usually provided



by sequencing of relatively few clone libraries using the Sanger method. Alternative approaches based on microarray analyses — such as Phylochip based on 16S rRNA gene sequence (DeSantis et al. , 2007) and Geochip based on functional gene sequence (Zhang et al. , 2007) — have over-come some of these limitations by improving the number of processed samples and the depth of analyses, thus providing a more comprehen-sive information about overall microbial diversity. In the past 10 years we have experienced another technological jump, led by the Next-Generation Sequencing approaches, which provide the possibility of processing thousands of samples simultaneously by massively paral-lel high-throughput sequencing methods, such as amplicon pyrose-quencing, metagenome sequencing, meta-transcriptomics, and whole genome sequencing (Fullwood et al. , 2009; Cummings et al. , 2010; Teeling and Glöckner, 2012; Zhou et al. , 2015; Aguiar-Pulido et al. , 2016).

The 16S rRNA gene and functional gene based amplicon sequencing allows a much deeper sampling of microbial communities by providing more sequence information than traditional Sanger sequencing; there-fore it has been applied to unravel the fluctuations of microbial com-munities across different ecosystems and along time series or seasonal changes (Engelbrektson et al., 2010; Pereira e Silva et al., 2011, 2012a; Dini-Andreote et al., 2014; Lema et al., 2014; Yan et al., 2016). On the one hand, amplicon sequencing studies of the root microbiome have mostly focused on phylogenetic composition, providing information on the presence of specific operational taxonomic units. For instance, by performing DNA-based pyrosequencing to characterize the structure of bacterial communities in both rhizosphere and bulk soils in a field cropped with six potato cultivars, Inceoglu et al. (2011) indicated that members of the Actinobacteria, Alphaproteobacteria, next to as-yet-unclas-sified bacteria, dominated the taxonomic composition. Moreover, rhizo-sphere samples were significantly different from corresponding bulk soil in each growth stage whereas cultivar effects were only observed in the young plant stage. Development of functional metagenomics, metapro-teomics and transcriptomics, on the other hand, can deliver insight into the activities and functions of the microbiome (Schoenfeld et al., 2011; Zhou et al., 2015; Aguiar-pulido et al., 2016). For example, a metapro-teomic approach was used to study microbial communities in the phyl-losphere and rhizosphere of rice (Knief et al., 2012). The results showed that despite the presence of nifH genes in both microenvironments, dini-trogenase reductase was exclusively identified in the rhizosphere. If such

23

1an approach could be applied to study the endosphere, more significant data regarding endophyte functionality can be collected.

Furthermore, complete microbial genome sequencing has made it possible to identify and characterize all genes present in a species, allow-ing the detection of novel metabolic pathways, gene regulatory elements, genes of unknown function, and genes for pathogenesis, virulence and drug resistance (Brown et al., 2012; Neupane et al., 2012; Gupta et al., 2014; Cho et al., 2015). This information also provides insights into the evolution of genes and species. Thus, the understanding of species diver-sity based on comparative genomics has led to a new epoch for biologi-cal investigations (Alföldi and Lindblad-Toh, 2013; Chen et al., 2015; Jun et al., 2015; Garrido-Sanz et al., 2016, 2017; Guo et al., 2016).

Comparative genomics has emerged as a powerful tool to identify functionally important genomic elements (Rodrguez-Palenzuela et al., 2010; Wu et al., 2011). By using Multilocus sequence typing ( MLST) and multilocus sequence analysis ( MLSA), Mulet et al. (2010) confirmed that the widest range of genomic diversity in the Pseudomonas genus is found in the P. fluorescens species complex. Comparisons among the genomes of strains within the P. fluorescens group highlight the tremendous di-versity of these bacteria (Loper et al., 2012; Cho et al., 2015; Garrido-Sanz et al., 2016; Guo et al., 2016). Moreover, the genes that were con-served among the different Pseudomonas species have provided clues to the common characteristics of pseudomonad PGPR, such as rhizo-sphere competence traits, while the strain-specific genes differentiated each strain on the basis of its lifestyle, specific ecological adaptations, and physiological role in the rhizosphere (Shen et al., 2013b). By analy-sis of each group-specific genome within the P. fluorescens complex and the search for key features, Garrido-Sanz et al. (2016) revealed congru-ence between the phylogenetic determination of these groups and their eco-physiology, showing the presence of proteins within the genetic clusters of biocontrol, siderophores, denitrification, toxins, bioremedi-ation and plant-bacteria interactions, providing insights into biocontrol and bioremediation applications of strains within the P. fluorescens com-plex as well as their role as PGPR.

The proper application of these techniques could assess the proper-ties of plant-associated microbial communities in heterogeneous en-vironments and enlighten the relative importance of soil and plant on the diversity, structure, and functions of plant-associated microbiomes. In the next section, I will discuss how I have made use of a salt marsh



primary succession to address whether plant or soil exerts a stronger effect on the selection of plant associated microbial communities, by applying a combination of culture-independent methods (16S rRNA sequencing, quantitative PCR (qPCR), molecular community finger-printing, and whole genome sequencing) and culture-dependent meth-ods (biochemical tests on bacterial isolates).

2. The study system

Salt marshes are boundary ecosystems between terrestrial uplands and the sea, forming the upper parts of the coastal intertidal zone in tem-perate regions and high latitudes. These ecosystems are strongly in-fluenced by physical, geographical and biological parameters, which correspondingly refer to tidal regime and wind-wave patterns, topo-graphical and hydrological differences, and interactions among species (Bertness, 1991a, 1991b; Gray, 1992). Surviving species on salt marshes have to face these environmental stresses, especially the frequent fluc-tuations in salinity and water level effected by predictable tidal regime (Lovell and Davis, 2012). These ecosystems, however, offer unique en-vironments for diverse species to exist, evolve and interact, forming an intricate network which provides researchers with perfect opportuni-ties to reveal the operating mechanisms and functions of each partici-pant in these ecosystems.

More than fifty percent of the North Sea beach of the Dutch barrier island Schiermonnikoog (53°30′, 6°10′), in the Netherlands, is covered by a salt marsh vegetation (Dijkman et al. , 2010). A special characteris-tic of this salt marsh is the fact that it displays a well-documented chro-nosequence crossing more than one hundred years of succession (Olff et al. , 1997). Previous work has revealed that the sedimentation caused by the tidal regime has led to modifications on the soil physicochemical conditions along the primary succession, promoting an accumulation of silt and clay particles. In addition, the salinity level also increased over time during succession, due to an accumulative effect. These ac-cretions led to higher elevations and subsequent development of vege-tation, whose diversity peaks at the intermediate to late stage, which in turn lead to an increase in organic matter (For detailed information, see Table 2). Along the primary successional gradients, the earliest stages are formed on the east side of the island, and later stages of succession

25

1

2. The study system

Tab

le 2

. Lo

cati

on

an

d s

oil

ph

ysic

och

emic

al p

aram

eter

s m

easu

red

at

the

sam

plin

g s

ites

alo

ng

th

e sa

lt m

arsh

ch

ron

ose

qu

ence

on

th

e is

lan

d

of

Sch

ierm

on

nik

oo

g, t

he

Net

her

lan

ds.

Th

e so

il p

hys

ico

chem

ical

par

amet

ers

wer

e ca

lcu

late

d b

y co

mb

inin

g t

he

resu

lts

fro

m t

wo

sam

plin

gs

take

n in

20

14 (f

rom

5 y

ear t

o 1

05

year

) an

d t

he

resu

lt o

f 0-y

ear f

rom

th

e sa

mp

ling

per

form

ed in

20

12 b

y D

ini-

An

dre

ote

et

al. (

2014

). Th

e p

lan

t d

iver

sity

dat

a w

ere

mo

difi

ed f

rom

Sch

ram

a (2

012

).

Mea

sure

d P

aram

eter

s0

yea

r5

year

15 y

ear

35 y

ear

65 y

ear

105

year

pH

8.70

±0.1

08.

51±0

.07

7.74

±0.1

07.

57±0

.08

7.72

±0.1

17.

51±0

.16

soil

wat

er c

on

ten

t (%

)11

.00

±1.0

06.

64±2

.04

40

.24

±4.8

14

6.32

±6.5

663

.83±

0.9

257

.53±

4.96

Soil

org

anic

car

bo

n (%

)0

.06±

0.0

00

.51±

0.2

45.

28±1

.46

7.0

4±1

.74

13.2

2±0

.02

11.2

3±0

.36

Tota

l nit

rog

en (%

)0

.01±

0.0

00

.01±

0.0

00

.28±

0.0

60

.41±

0.1

30

.92±

0.0

30

.79±

0.0

3

Nit

rate

(mg

/kg

dry

so

il)3.

00

±0.0

09.

94±2

.13

13.8

3±4.

06

14.2

1±8.

6019

.33±

24.1

823

.82±

25.4

5

Am

mo

niu

m (m

g/k

g d

ry s

oil)

5.0

0±1

.00

1.0

6±0

.44

12.2

5±7.

41

21.5

8±16

.68

38.3

8±19

.80

29.7

0±1

0.6

8

Ca

(mg

/kg

dry

so

il)—

2096

.16±

107.

8129

72.6

3±4

59.0

036

76.9

6±16

6.78

3228

.29±

199.

2231

54.7

8±60

7.0

0

Mg

(mg

/kg

dry

so

il)—

75.7

1±15

.71

967.

49±

363.

9719

91.6

3±28

5.34

3134

.24

±162

.20

2536

.61±

228.

13

K (m

g/k

g d

ry s

oil)

—60

.14

±18.

6953

5.0

6±24

7.71

114

3.26

±213

.1617

78.0

8±72

.57

1557

.98±

176.

68

Na

(mg

/kg

dry

so

il)—

150

.82±

70.7

138

51.5

1±12

56.9

577

88.3

4±4

26.9

271

44.

92±1

144.

1472

68.5

5±12

30.7

2

Ava

ilab

le p

ho

sph

ate

(P2O

5; m

g/k

g d

ry s

oil)

—28

4.0

8±3.

04

810

.69±

37.9

913

08.

58±2

6.16

2226

.62±

15.6

321

16.1

8±50

.59

Soil

Typ

e

San

d (%

)92

.30

±0.5

02.

96±0

.00

—2.

73±0

.01

2.29

±0.0

22.

13±0

.02

Silt

(%)

2.70

±0.5

01.

48±

0.1

3—

2.35

±0.0

12.

64±0

.02

2.69

±0.0

1

Cla

y(%

)5.

00

±0.0

01.

71±0

.00

—2.

37±0

.02

2.56

±0.0

12.

58±0

.00

Pla

nt

div

ersi

ty (n

um

ber

of

spec

ies)

2.0

±0.7

—6.

4±3

.48.

6±1.

24.

4±1

.14.

2±1.

8

Soil

typ

e d

ata

was

no

t te

sted

in t

he

15-y

ear

stag

e; S

oil

mic

ron

utr

ien

ts d

ata

wer

e n

ot

test

ed in

th

e 0

-yea

r st

age;

Pla

nt

div

ersi

ty d

ata

was

no

t d

etec

ted

in t

he

5-ye

ar s

tag

e.


are situated up to 8 kilometres to the west (Olff et al. , 1997). Salt marsh age at each successional stage has been estimated from topographic maps, aerial photographs, and the thickness of the sediment layer ac-cumulated on top of the underlying sand layer (Olff et al. , 1997; van Wijnen and Bakker, 1999) and calibrated using long-term observations of permanent plots (van Wijnen et al. , 1997). This salt marsh primary succession provides, thus, a natural gradient of soil development, of-fering a perfect opportunity to unravel the relative importance of soil types and plant species on the structure and functions of plant associ-ated microbial communities.

Previous work on our study system on the island of Schiermonnikoog has quantified in detail the dynamics of soil and vegetation succession (Olff et al., 1997), the interactions between plants and vertebrate her-bivores (van De Koppel et al., 1996; Olff et al., 1997; van Wijnen and Bakker, 1999; Kuijper and Bakker, 2005; Schrama et al., 2013a), the com-position and variability of food web assembly along the primary succes-sion (Schrama et al., 2013b), and the soil microbial communities (Dini-Andreote et al., 2014, 2015, 2016). By performing 454 pyrosequencing for bacterial composition and diversity analyses, Dini-Andreote et al. (2014) revealed for the first time that the initial stages of soil develop-ment held higher phylogenetic diversities than the soil at late succes-sion, suggesting temporal niche partitioning as the dominant mech-anism of assembly, given the great dynamism imposed by the daily influence of the tide. Moreover, they indicated that the allogenic suc-cession of total bacterial communities and the structure of the popu-lations of genes involved in N cycling processes (nifH, amoA, nirK, nirS and nosZ) were mostly driven by shifts in the soil physical structure, as well as variations in pH and salinity (Dini-Andreote et al., 2014; 2016). However, there is still no illustration on the dynamics and characteris-tics of microbial communities associated with plants.

In this context, the salt marsh successional gradients on the island of Schiermonnikoog provide a long enough time scale to study the plant- associated microbial community and the dynamics of plant-microbe in-teractions. This study could illustrate a general paradigm of salt marsh succession in similar regions and enlighten the conservation measures of coastal marshes in perspectives of plant-microbe interplays. In this study, successional ages of 5, 15, 35, 65 and 105 years are the sampling sites where our focus plant species consistently occurred (Figure 3).

27

1

3. General aim of this thesis


In the introduction of the root microbiome, I have touched upon a key question which has been discussed by many studies but without a con-sistent answer yet: what is the main driving force of plant-associated microbial communities? The aim of the work presented in this the-sis was to disentangle the relative importance of plant selectivity and soil type in reigning the root-associated bacterial community dynam-ics, both from phylogenetic and functional aspects. I selected typical salt marsh plants — Limonium vulgare and Artemesia maritima — as the focal plant species (Figure 4), as these are widely distributed across the suc-cessional stages. Hence, effects of plant-exerted selective force would be discernible from that exerted by local soil conditions. Thus, by mak-ing use of a well-established salt marsh chronosequence located at the island of Schiermonnikoog, the Netherlands, in this thesis I provide a comprehensive understanding of the driving force of community struc-ture, diversity and functions of root-associated bacteria associated with salt marsh plants.

Figure 3 The study system along the salt marsh chronosequence on the island

Schiermonnikoog.


3.1. Hypotheses

Based on the assumption that the selection by plants rather than soil regulates the dynamics of the root-associated bacterial assembly along the chronosequence, I posed the following hypotheses for further explo-ration in this thesis:

• The phylogenetic diversity of root-associated (rhizosphere and en-dosphere) bacterial communities is constant along the chronose-quence whereas that associated with the bulk soil will vary accord-ing to the different successional stages.

• N-cycling gene abundances are higher in rhizosphere than in bulk soil samples across all soil successional stages, and the gene abundances in bulk soil will consistently increase towards late successional stages.

• The functional diversities — as determined by metabolic traits as well as associated genome traits — of strains obtained from rhizospheres but not those from root endosphere will increase along succession.

Figure 4 Focal salt marsh plant

species (A) L. vulgare, (B) A.

maritima

29

1


• The sizes of the genomes of Pseudomonas isolates associated with the rhizosphere but not those associated with the root endosphere will increase along succession, while the latter will remain constant across all soil successional stages.

3.2. Thesis outline

The aforementioned hypotheses are investigated throughout chap-ters 2-5 of this thesis (Figure 5). I started by investigating the dis-tribution of the community structures of root-associated bacteria along the salt marsh chronosequence in Chapter 2, by performing 454- pyrosequencing. Specifically, the bacterial communities associ-ated with the soil, rhizosphere and the root endosphere of L. vulgare were examined. Here I explored the relative importance of soil type and plant selectivity on the bacterial diversity and structure, in dif-ferent plant compartments (rhizosphere and endosphere) as well as in the bulk soil. Thus, by using a gradient of plant-associated microbes — from bulk to endosphere, the latter being under stronger plant con-trol — I aimed at verifying if the plant selective force in the endosphere was indeed stronger than that in rhizosphere and bulk soil across soils that vary in physicochemical composition.

Following the study of the total root-associated bacterial community from phylogenetic prospective, I then investigated the root microbiome

Figure 5 General outline in this study


from the functional aspect by focusing on N-cycling communities, given their important role in maintaining the salt marsh ecosystem functioning. Using the same set up as in Chapter 2, in Chapter 3, I stud-ied the abundance and community structure of diazotrophs and am-monia oxidizers associated with the bulk soil and rhizosphere of the plant species Limonium vulgare, by quantitative PCR (qPCR) and molec-ular community fingerprinting methods, respectively, along the chro-nosequence. Here, the main question I tackled was whether plants exert an overriding driving force on the community dynamics of N-cycling microbes following the systematic changes of soil properties and plant diversity across the succession.

Although chapters 2 and 3 provided a general view (on the basis of cultivation-independent methods) of the effects of plant and soil on the diversity, structure and function of microbial communities along the primary succession, they were limited in the amount of in-formation regarding the physiological characteristics of these micro-bial communities. Therefore, in Chapter 4, I explored the bacterial species isolated from rhizosphere and endosphere of two salt marsh plants, L. vulgare and A. maritima, along the chronosequence. Given the significance of plant-growth promoting traits on plant health and de-velopment, in this chapter I used traditional biochemical and physio-logical microbiological tests to screen the functional traits associated with bacterial fitness and plant growth, thus providing a better under-standing of the root- associated bacterial functionalities in salt marsh plants, as well as unravelling how multiple factors such as soil type, plant species and plant micro-environment influence the functional diversity in the root microbiome.

For Chapter 5, I delved further into the potential genomic modi-fications that could lead to the physiological adaptations observed in chapter 4. Thus, by using the same set up as in the previous chapter and focusing on an important, well-known PGPR group, the fluores-cent pseudomonads, I used comparative genomics of 70 different root- associated Pseudomonas strains to explore their genome evolution, from both phylogenetic and functional perspectives along the salt marsh primary succession. The aim of this chapter was to potentially reveal core genes associated with the adaptation of Pseudomonas to soil type and plant. Finally, in Chapter 6 I provided a summary of the main find-ings and discussed future questions in the light of the results described in the previous chapters.

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University of Groningen The dynamics of root microbiomes ......salt marsh salt marsh plants rhizosphere community structure of diazotrophs major major (Davis et al., 2011) crop land