University of Groningen
Ecology of Acidobacteria and Verrucomicrobia in the plant-soil system Nunes da Rocha, Ulisses
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Download date: 29-09-2021 Ecology of Acidobacteria and Verrucomicrobia in the plant-soil ecosystem
Proefschrift
ter verkrijging van het doctoraat in de Wiskunde en Natuurwetenschappen aan de Rijksuniversiteit Groningen op gezag van de Rector Magnificus, dr. F. Zwarts, in het openbaar te verdedigen op vrijdag 22 oktober 2010 om 11.00 uur
door
Ulisses Nunes da Rocha
geboren op 29 mei 1981 te Rio de Janeiro, Brazilië
Promotor: Prof. dr. J.D van Elsas
Copromotor: Dr. L. S. van Overbeek
Beoordelingscommissie: Prof. dr. O. Kuipers
Prof. dr. S. Agathos
Prof. dr. G. Kowalchuk
ISBN (Book): 978-90-367-4577-2
ISBN (Digital): 978-90-367-4576-5
This research was part of the Ecogenomics program which is sponsored by the Dutch National Genomics Initiative and the basic research program on sustainable agriculture (KB4) sponsored by the Dutch ministry of agriculture, nature and food safety.
Table of content
Summary 7
Chapter 1: General Introduction 9
The great plate count anomaly (GPCA) 10 Hitherto-uncultured bacteria – The unseen majority 11 Acidobacteria 13 Verrucomicrobia 17 Implications of the GPCA for rhizosphere ecology 18 Objectives 19 Approach 19 Working hypothesis 20 Research questions 20 Outline of the thesis 21 References 24
Chapter 2: Exploration of hitherto-uncultured bacteria from the rhizosphere 29
Abstract 29 Introduction 30 Soils, plant roots and microbial communities – an intricate triplet 34 Dominant hitherto-uncultured bacteria in the rhizosphere 36 Ecology of hitherto-unculturable bacteria in the rhizosphere 39 Methods to improve bacterial and Archaeal culturability 44 Towards the as-yet-uncultured microbial populations in the rhizosphere and their basic ecology 48 Conclusions 49 References 51
Chapter 3: Cultivation of hitherto uncultured bacteria belonging to the Verrucomicrobia subdivision 1 from the potato (Solanum tuberosum L) rhizosphere 57
Abstract 57 Introduction 58 Materials and methods 59 Results 64 Discussion 75 Conclusions 78 Recommendations and perspectives 78 Acknowledgements 78 References 79
Chapter 4: Isolation and partial characterization of Holophaga, Luteolibacter, unclassified Verrucomicrobia and Verrucomicrobium spp. from the leek (Allium porrum) rhizosphere 82
Abstract 82 Introduction 83 Material and methods 84 Results 87 Discussion 95 Acknowledgements 99 References 100
Chapter 5: Real-time PCR detection of Holophagae (Acidobacteria) and Verrucomicrobia subdivision 1 groups in bulk and leek (Allium porrum) rhizosphere soils 103
Abstract 103 Introduction 104 Material and methods 105 Results 111 Discussion 115 Acknowledgements 118 References 119
Chapter 6: Rhizocompetence of culturable Holophaga (Acidobacteria) sp. in the leek (Allium porrum) rhizosphere 121
Abstract 121 Introduction 122 Material and methods 123 Results 127 Discussion 132 Acknowledgements 134 References 135 chapter 7: Different rhizosphere competence in two Verrucomicrobium subdivision 1 strains previously isolated from the leek (Allium porrum) rhizosphere 137
Abstract 137 Introduction 138 Material and methods 139 Results 143 Discussion 149 Acknowledgements 151 References 152 Chapter 8: Distribution of different Acidobacteria and Verrucomicrobia subdivision 1 groups over compartments of different plant species 154
Abstract 154 Introduction 155 Material and methods 156 Results 160 Discussion 168 Acknowledgments 171 References 172
Chapter 9: General Discussion 174
Dominant hitherto-uncultured bacteria in the rhizosphere 174 Diminishing the gap between cultured and total bacteria in rhizosphere samples 175 Phylogeny and phenotypic analysis of culturable Acidobacteria and Verrucomicrobia from the rhizosphere 176 Phenotypic analysis of culturable Holophaga and Verrucomicrobia subdivision 1 177 Real-time PCR detection of particular Acidobacterium and Verrucomicrobium types in bulk and rhizosphere soil 178 Holophaga and Verrucomicrobium subdivision 1 spp. show shifts in abundance from bulk to rhizosphere soil 180 Environmental factors that affect the distribution of Acidobacterium and Verrucomicrobium spp. over rhizosphere and bulk soil 182 Future perspectives 184 References 188
Appendix 191
Samenvatting 195
Aknowledgments 197
About the Author 199
Summary
It is generally accepted that less than 1% of the total bacterial diversity can be cultured in the laboratory. This complicates microbial ecological research in different ecosystems, like the plant-soil ecosystem. Bacteria belonging to the phyla of Acidobacteria and Verrucomicrobia were successfully isolated from the rhizosphere during the research done for this thesis. Members from both phyla had never been cultured before from soil samples taken in the neighborhood of plant roots. Bacterial isolations may facilitate the study of the ecology of both phyla in the plant-soil ecosystem.
From the rhizosphere of potato and/ or leek the following strains were isolated: four Luteolibacter spp., four Candidatus genus Rhizospheria spp. and one Verrucomicrobia sp. (all affiliating with Verrucomicrobia subdivision 1) and two Holophagae spp., both affiliating to the Acidobacteria. The taxonomy of these strains were determined on basis of their 16S rRNA genes and further the potential of these strains to interact with plants was determined, like the possibilities for growing on root exudates. It appeared that the Holophagae sp. strains taxonomically and phenotypically closely resembled each other and that Verrucomicrobia subdivision 1 strains were more diverse in their taxonomies and phenotypes.
Real time PCR was applied to describe the fate of Acidobacteria and Verrucomicrobia subdivision 1 in plant-soil ecosystems and this technique appeared to be an excellent tool for this purpose. In vitro colonization assays and plant-soil microcosms gave insights in the ecology of Holophagae and Candidatus genus Rhizospheria (proposed new taxonomical group within Verrucomicrobia subdivivsion 1 species). It was presumed that both groups would associate with different plant species. On the other hand, higher numbers of Luteolibacter (determined by real time PCR) were found in bulk soil when compared with respective rhizosphere soil layers in the plant- soil microcosms. In the field experiment, Acidobacteria subgroups 1, 3, 4 were found to be present in higher numbers in bulk soil than in their respective rhizospheres. On their turn, Acidobacteria subgroup 6 cell numbers were higher number in the rhizosphere than in surrounding bulk soil. The distribution of the Holophagae was different over
7 rhizosphere and bulk soils and it appeared that this Acidobacteria group occupied a specific niche in the leek rhizosphere.
Successful culturing of representatives of Acidobacteria and Verrucomicrobia subdivision 1 groups facilitated experimentation in the neighborhood of plants; studies that are needed to develop new ecological theories on microbial interactions in the plant-soil ecosystem. Ecological research of Acidobacteria and Verrucomicrobia, respectively Holophagae and Candidatus genus Rhizospheria, in plant-soil ecosystems opened new directions towards future research. This will focus on possible interactions with plants and other microorganisms living in association with plants. This research contributes to our current knowledge on the relevance of both groups in the plant soil ecosystem and their relevance to plant growth.
8 Chapter 1: General Introduction
Prokaryotic organisms on Earth are estimated to range in population size between 4 and 6 x 1030 cells, amounting to 350-550 x 109 tons of C biomass (Witman et al., 1998). Thus, the estimated amount of prokaryote-bound C is estimated to be up to 60-100% of the carbon residing in plants. The first bacteria, as cellular units, were observed by Antonie van Leeuwenhoek in the 17th century (Madigan & Martigo, 2006). More than 300 years have passed since the first bacteria were observed and, due to the positive as well as negative impacts of microorganisms on humans and their ecological importance on Earth, different disciplines have evolved in microbiology. A first discipline, bacteriology, was founded in the 19th century by Ferdinand Cohn, who described bacteria such as Bacillus and Beggiatoa while studying microalgae and photosynthetic bacteria (Encyclopædia Britannica. 2010). Next to that, environmental and microbial ecology branched of general microbiology by the pioneers Martinus Beijerinck and Sergei Winogradsky. These pioneers were the first to study microorganisms without medical relevance as from the late 19th century (Paustian & Roberts, 2009). On the other hand, the first bacteria had already been isolated by Robert Koch, a contemporary of Cohn, who was the first to demonstrate that specific bacterial isolates could be assigned as causative agents of particular diseases. For that, the so- called postulates of Koch were invoked. In particular, he found that Mycobacterium tuberculosis was the causative agent of tuberculosis (Madigan & Martigo, 2006). Up to the 1980’s, a major part of the knowledge in bacteriology has been achieved on the basis of isolation procedures and the use of Koch’s postulates to demonstrate bacterial involvement in pathogenesis and other functions (Grimes, 2006). In spite of this progress, it has become widespread among the microbiologists that most bacteria in nature have so far escaped isolation by culturing (Rothschild, 2006). The phenomenon that most microbiota in natural systems is difficult to culture has been coined the great plate count anomaly (Staley & Konopka, 1985).
9 General Introduction
The great plate count anomaly (GPCA)
The large discrepancy between the bacterial numbers calculated from viable plate counts and those from direct microscopic cell counts was probably first observed and described by Ramuzov (1932), while studying oligotrophic to mesotrophic aquatic habitats. A contemporary illustration of this anomaly was provided by Staley & Konopka (1985) while studying non photosynthetic microorganisms in aquatic and terrestrial habitats. The question arises: why does this discrepancy happen? As an easy explanation, it might be cogitated that most bacteria from nature would not be able to grow under laboratory conditions and are thus nonviable. Nevertheless, cells could be alive under field conditions, i.e. perfectly adapted to their environmental niche (Staley & Konopka, 1985). If the explanation is correct, then it is important to understand how such bacteria might be active and increase their population sizes in nature and still be recalcitrant to culturing in the laboratory. A model acid mine drainage system recently demonstrated that low-abundance bacteria and Archeae, that had not yet been isolated, were active in situ (Belnap et al., 2010). In the light of these results, it is likely that, although in principle capable of activity in the environment, most bacteria may remain uncultured (and thus non-isolated) due to laboratory cultivation procedures. This may lead us to the question “Which factors inhibit most environmental bacteria to grow under laboratory conditions?” The traditional cultivation techniques often lacked the precise factors of the environment, such as the adequate levels of micronutrients and growth substrates, which may have impeded bacterial growth. However, in recent work improved bacterial culturability - as a result of incubation under conditions better-tuned to the habitat - has been achieved for bulk soil (Janssen et al, 2002; Sait et al, 2002; Schoenborn et al, 2004; Davis et al, 2005) as well as freshwater bacteria (Bruns et al, 2003). In these studies, factors relevant for the improved cultivation were: i) reduced nutrient availability, ii) prolonged incubation times and iii) reduction of oxidative stress by the addition of protective agents. Recovery percentages from soils and other environments can thus be increased by simple modifications of existing protocols. However, we still ignore to what extent this improvement holds true for all extant bacterial diversity. And, a major part of the extant bacterial diversity in many ecosystems has remained cryptic so far (Rothschild, 2006). Our inability to grow all organisms on agar media under the conditions used may be related to the inability of some bacteria to grow on agar (Kamagata & Tamaki, 2005; Bollman et al, 2007); this may even come down to a general absence of the capacity to grow on solid media (Schoenborn et al, 2004). In addition, some bacteria may need particular signal molecules to initiate cell division
(Nichols et al, 2008), or particular conditions for growth, like specific CO2 : O2 ratio’s (Stevenson et al, 2004). Alternatively, bacteria may only grow when in consortia
10 Chapter 1
together with other microorganisms (Erkel et al, 2005; Lo et al, 2007; Ettwig et al, 2008). Finally, it cannot be excluded that bacteria residing in nature are permanently injured, resulting in a loss of culturability. Concluding, we may say that, although advances in research have shown that members of bacterial groups that were previously considered to be unculturable can now be isolated on media under particular laboratory conditions, a major part of the extant bacteria diversity still remains cryptic due to their recalcitrance to culturing.
Hitherto-uncultured bacteria – The unseen majority
Prior to the introduction of modern DNA-based methods for the phylogenetic (based on 16S rRNA genes) and ecological analysis of bacterial communities, the investigation of bacterial diversity was severely hampered by the need to obtain a pure culture of each bacterial type from the environment (Amann et al., 1995). As the sequencing of rRNA genes directly retrieved from complex environmental samples by polymerase chain reaction (PCR) and cloning has become a routine technique in microbial ecology (Cardenas & Tiedje, 2008), our knowledge of the extant bacterial diversity has greatly expanded. Indeed, environment-derived 16S rRNA gene sequences nowadays represent a sizeable fraction of the rRNA sequence data deposited in public databases, such as the Ribosomal Database Project (Cole et al., 2005). Although it is broadly accepted that the majority of environmental bacteria have so far escaped culturing, there is currently no recent general oversight of dominant and widespread bacterial groups that have never been cultured or those that have few culturable representatives. Therefore, the Ribosomal Database Project, Release 10 update 20 (May 19, http://rdp.cme.msu.edu/index.jsp) was used here to pinpoint those bacterial groups that dominate sequence databases but are sparsely present in isolate collections (Table 1). I found that a subset of the 35-odd currently-known bacterial phyla, e.g. OP11, OD1, BRC1, SR1 and WS3, fall into this class. Moreover, a set of eight bacterial phyla (or “candidatus phyla”) had more than 95% of their 16S rRNA gene sequences retrieved from culture independent studies, as follows: candidatus phylum OP10 (0.6% of sequences associated with cultured strains), candidatus phylum TM7 (0.8%), Gemmatimonadetes (1.0%), Acidobacteria (1.5%), Caldiserica (2.6%), Chloroflexi (2.9%), Verrucomicrobia (3.0%) and Planctomycetes (4.3%). This analysis may be biased due to the different methods used to acquire the sequences, but it gives a glimpse of the state-of-the-art at this point in time: a sizeable fraction of the bacterial phyla remains either completely uncultured or encompass just a few cultured representatives. Among the groups with just a few culturable strains are the Acidobacteria and Verrucomicrobia. These bacterial phyla are widespread in many environments, in which
11 General Introduction
they sometimes dominate. They are also diverse. Reports that describe the ecology of these groups in different environments are sparse, in spite of their ubiquity and diversity. Therefore, I decided for the purpose of this thesis to place a focus on the advancement of our ecological knowledge of the Acidobacteria and Verrucomicrobia.
Table 1 Number of 16S rRNA genes for each bacterial phylum present in the Ribosomal database (RDP) release 10, update 20, May 19, 2010. Phyluma,b Number of sequencesc Culture-independent studies Culture-dependent studies Acidobacteria 5,348 85 Firmicutes 127,859 23,332 Bacteroidetes 49,870 4,866 Actinobacteria 71,639 18,579 Aquificae 707 97 Caldiserica 39 1 Chlamydiae 51 209 Chlorobi 165 122 Chloroflexi 3,088 95 Chrysiogenetes 0 6 Deferribacteres 232 30 Deinococcus-Thermus 301 348 Dictyoglomi 5 9 Fibrobacteres 160 62 Fusobacteria 986 211 Gemmatimonadetes 579 6 Lentisphaerae 151 7 Nitrospira 563 91 Planctomycetes 3,150 140 Proteobacteria 99,344 48,336 Spirochaetes 1,714 1,357 Synergistetes 655 72 Tenericutes 313 1,605 Thermodesulfobacteria 77 10 Thermotogae 307 112 Verrucomicrobia 3,537 110 Bacteria incertae sedis 231 16 OP10 157 1
12 Chapter 1
OP11 57 0 OD1 93 0 BRC1 51 0 Cyanobacteria 4,717 2,700 SR1 2 0 WS3 96 0 TM7 487 4 Unclassified Bacteria 7,931 253 a In bold, bacterial groups with no 16S rRNA gene sequences retrieved from culturable-dependent studies b Underlined, bacterial groups with more than 95% of 16S rRNA gene sequences retrieved from culture-independent studies c Number of sequences with more than 1200 bp in length and good quality according to RDP standards
Acidobacteria
In terms of understanding bacterial behaviour, the phylum Acidobacterium has remained enigmatic for a long time. Only in the half-1990-ies were the first three acidobacterial species, i.e. Acidobacterium capsulatum (Hirashi et al., 1995), Geothrix fermentans (Lonergan et al., 1996) and Holophagae foetida (Liesack et al., 1994) obtained and described. The suggestion to create a new phylum for these three species was made by Ludwig et al. (1997). This was based on the observation that particular 16S rRNA gene sequences derived from different environments, e.g. soil (Stackebrandt et al., 1993), sediment (Wise et al., 1997) and sludge (Bond et al., 1995), were phylogenetically related to Acidobacterium capsulatum. By 1999, Acidobacterium was proposed as a new bacterial kingdom (Barns et al., 1999). In the past 12 years, the singling out of Acidobacterium as a novel phylum has been substantiated. The phylum seems robust, and at the same time exhibits a remarkable diversity. Quaiser et al. (2003) first described five different phylogenetic groups of Acidobacteria, but shortly following this work, eleven different groups were proposed by Zimmermann et al. (2005). More recently, the number of acidobacterial subgroups was updated to an astonishing 26 (Barns et al., 2007). Concerning isolation, by 1999 only three strains of Acidobacteria had been isolated. This number (type strains) has increased to eight (Table 2). At the same time, the number of 16S rRNA gene sequences based on cultured Acidobacteria in public databases has increased (http://rdp.cme.msu.edu/) to a total of 85 at the time of writing of this thesis. The culturable Acidobacteria affiliate the following subgroups of Acidobacteria: Holophagae (6 strains), subgroup 1 (61), subgroup 2 (2), subgroup 3 (12) subgroup 4 (4). Up to the completion of this thesis, no study concerning the
13 General Introduction
phylogenetic relationship among these strains had been done. Most of the improvement in the cultivation of the Acidobacteria was achieved in the last few years and was mostly brought about by simple modification of traditional cultural techniques (detailed information on this subject is provided in chapter 2). The phylum Acidobacterium is not only highly diverse but its members are broadly distributed. They have been detected in marine sponges (Radwan et al., 2010), soil (Liles et al., 2010), the rhizosphere of pioneer plants (Navarro-Noya et al., 2010), domestic toilets (Egert et al., 2010), healthy coral reef (de Castro et al., 2010), fire- induced grassland soil (Lin et al., 2010b), among many other terrestrial and aquatic environments. Interestingly, Acidobacteria are often among of the most dominant bacterial groups in soils. They may contribute with 20-52% to the 16S rRNA gene abundance present in this environment, as evidenced from the analysis of 16S rRNA gene based clone libraries (Janssen, 2006; Sait et al., 2002). Although they make part of the dominant community of soil and related environments, little information is available about their ecology. A good example of inconclusive ecological information is the distribution of Acidobacteria in plant-soil environments. In fact, contradictory information has been provided on the occurrence of different members of the Acidobacteria in bulk and rhizosphere soil (Chow et al., 2002; Sanguin et al., 2006; Zul et al., 2007; Kielak et al., 2008). Therefore, many horizons are still to be explored in the ecology of Acidobacteria in the plant-soil ecosystem.
Table 2 Recognized species of Acidobacteria and Verrucomicrobia. Taxa References Phylum Acidobacteria Class Acidobacteria T. Cavalier-Smith (2002) Order Acidobacteriales T. Cavalier-Smith (2002)
Family Acidobacteriaceae T. Cavalier-Smith (2002)
Genus Acidobacterium Kishimoto et al., 1991 Acidobacterium capsulatum Kishimoto et al., 1991 Genus Edaphobacter Koch et al., 2008 Edaphobacter aggregans Koch et al., 2008 Edaphobacter modestus Koch et al., 2008 Genus Terriglobus Eichorst et al., 2007 Terriglobus roseus Eichorst et al., 2007 Order Acanthopleuribacterales Fukunaga et al., 2008 Family Acanthopleuribacteraceae Fukunaga et al., 2008 Genus Acanthopleuribacter Fukunaga et al., 2008
14 Chapter 1
Acanthopleuribacter pedis Order Holophagales Fukunaga et al., 2008 Family Holophagaceae Fukunaga et al., 2008 Genus Geothrix Coates et al., 1999 Geothrix fermentans Coates et al., 1999 Genus Holophaga Liesack et al., 1994 Holophaga foetida Liesack et al., 1994 Subdivision 3 Barns et al. (2007) Genus Bryobacter Kulichevskaya et al. (2010) Bryobacter aggregatus Kulichevskaya et al. (2010) Phylum Verrucomicrobia Hedlung et al. (1997) Class Verrucomicrobia Hedlund et al. (1997) Order Verrucomicrobiales Ward-Rainey et al. (1995)
Family Rubritaleaceae Genus Rubritalea Scheuemayer et al. (2006) Rubritalea marina Scheuemayer et al. (2006)
Rubritalea sabuli Yoon et al. (2008b) Rubritalea spongiae Yoon et al. (2007b) Rubritalea squalenifaciens Kasai et al. (2007) Rubritalea tangerina Yoon et al. (2007b) Family Verrucomicrobiaceae Ward-Rainey et al. (1995) Genus Akkermansia Derrien et al. (2004)
Akkermansia muciniphila Derrien et al. (2004) Genus Haloferula Yoon et al. (2008c) Haloferula harenae Yoon et al. (2008c) Haloferula helveola Yoon et al. (2008c) Haloferula phyci Yoon et al. (2008c) Haloferula rosea Yoon et al. (2008c) Haloferula sargassicola Yoon et al. (2008c) Genus Luteolibacter Yoon et al. (2008a) Luteolibacter algae Yoon et al. (2008a) Luteolibacter pohnpeiensis Yoon et al. (2008a) Genus Persicirhabdus Yoon et al. (2008a) Persicirhabdus sediminis Yoon et al. (2008a) Genus Prosthecobacter Hedlung et al. (1996)
15 General Introduction
Prosthecobacter debontii Hedlung et al. (1997) Prosthecobacter dejongeii Hedlung et al. (1997) Prosthecobacter fluviatilis Takeda et al. (2008) Prosthecobacter fusiformis Staley et al. (1980) Prosthecobacter vaneervenii Hedlung et al. (1997) Genus Roseibacillus Yoon et al. (2008a) Roseibacillus ishigakijimensis Yoon et al. (2008a) Roseibacillus persicicus Yoon et al. (2008a) Roseibacillus ponti Yoon et al. (2008a) Genus Verrucomicrobium Schlesner (1987) Verrucomicrobium spinosum Schlesner (1987) Class Opitutae Choo et al. (2007) Order Opitutales Choo et al. (2007) Family Opitutaceae Choo et al. (2007) Genus Alterococcus Shieh & Jean (1998) Alterococcus agorolyticus Shieh & Jean (1998) Genus Opitutus Chin et al. (2001) Opitutus terrae Chin et al. (2001) Order Puniceicoccales Cho et al. (2007) Family Puniceicoccaceae Cho et al. (2007) Genus Cerasicoccus Yoon et al. (2007a) Cerasicoccus arenae Yoon et al. (2007a) Genus Coraliomargarita Yoon et al. (2007d) Coraliomargarita akajimensis Yoon et al. (2007d) Genus Pelagicoccus Yoon et al. (2007e) Pelagicoccus abus Yoon et al. (2007e) Pelagicoccus croceus Yoon et al. (2007c) Pelagicoccus litoralis Yoon et al. (2007e) Pelagicoccus mobilis Yoon et al. (2007e) Genus Puniceicoccus Choo et al. (2007) Puniceicoccus vermicola Choo et al. (2007)
16 Chapter 1
Verrucomicrobia
Verrucomicrobium spinosum, described by Schlesner (1987), was the first identified species of the phylum Verrucomicrobium. This strain, together with five others, had initially been grouped together with other bacteria, such as Prosthecomicrobium, Ancalomicrobium and Stella, on the basis of morphological criteria (shape, number, length and location of prosthecae). Nowadays, based on their 16S rRNA gene sequences, the genus Prosthecomicrobium is classified as part of the order Rhizobiales (Alphaproteobacteria); it encompasses both Prostecomicrobium and Analomicrobium spp, according to RDP classification (release 10, update 20, May 19, 2010, http://rdp.cme.msu.edu/). According to the same identifier, the genus Stella now belongs to the order Rhodospirillales (Alphaproteobacteria). Eight years after Verrucomicrobium was described as a genus, Verrucomicrobium-like sequences were defined as a separate group within the domain Bacteria (Ward-Rainey et al., 1995). In that study, the almost-complete 16S rRNA gene sequence of V. spinosum, isolated from an alkaline lake, and those of so-called Cluster III clones, isolated from acid soil (Liesack & Stackebrandt, 1992) were grouped in the family Verrucomicrobiaceae from the order Verrucomicrobiales (Ward-Rainey, 1995). One year later, members of Prosthecobacter, that had previously been affiliated to the genus Caulobacter (order Caulobacterales, Alphaproteobacteria), were demonstrated to be phylogenetically related to the then defined Verrucomicrobiaceae (Hedlund et al., 1996). Finally, 10 years after the description of V. spinosum, Verrucomicrobium was proposed as a new division within the domain Bacteria (Hedlund et al., 1997). Up to the current moment, seven different subdivisions of Verrucomicrobium have been described based on 16S rRNA gene sequences of cultured and yet-to-be cultured organisms (Schlesner et al., 2006). Much like for the Acidobacteria, recent advances in culture-dependent techniques (reviewed in chapter 2) have increased the numbers of cultured strains affiliated to Verrucomicrobium. In the RDP database (release 10, update 20, May 19, 2010, http://rdp.cme.msu.edu/), over one hundred 16S rRNA gene sequences belonging to cultured members of Verrucomicrobium are available (Table 1). To the best of our knowledge, two valid classes have been described, i.e. Verrucomicrobiae (Hedlund et al., 1998) and Opitutae (Choo et al., 2007), along with three orders, four families and fourteen genera. Moreover, at least 30 type strains (i.e. recognized species) have been described for this phylum (Table 2), twenty one of them by one group (Yoon et al. 2007a, 2007b, 2007c, 2008a, 2008b, 2008c). Reports on the detection of Verrucomicrobium in the environment demonstrate that this phylum is not only diverse but also broadly distributed among different aquatic
17 General Introduction
and terrestrial environments. Thus, members of this division were among the dominant bacterial groups in soils and rhizospheres (Rosenberg et al., 2009), drinking water reservoirs (Lymperopoulou et al., 2010), human intestinal tract systems (Wang et al., 2005), contaminated groundwater (Herrmann et al., 2008), animal (gorilla) feces (Frey et al., 2006) and swine waste lagoons (Goh et al., 2009). However, akin to the Acidobacteria, little conclusive information can be found on the ecology of Acidobacteria in soil and related environments. Some reports concern the distribution of members of Verrucomicrobium over bulk and rhizosphere soil (Chow et al., 2002; Sanguin et al., 2006; Zul et al., 2007; Kielak et al., 2008), however this information was judged to be very crude. This highlights the necessity to further explore the ecology of Verrucomicrobium in plant-soil systems.
Implications of the GPCA for rhizosphere ecology
Based on the fact that plants are key primary producers in most terrestrial ecosystems (Dennis et al., 2010), it is reasonable to suppose that they are essential for soil life. Since plants colonized terrestrial environments as from around 700 million years ago, co-evolution between plants and soil bacteria has occurred (Phillips et al., 2003). During this co-evolution, interactions of many kinds have been created. For example, symbiosis between Rhizobium and legumes (Kempel et al., 2009), parasitism of Agrobacterium forming tumors on plants (Hooykaas & Beijersbergen, 1994), pathogenicity of Ralstonia solanacearum causing wilting disease of potato or tomato (Thoquet et al., 1996; Van Elsas et al., 2000; Tans-Kersten et al., 2001). Many times, such interactions are dependent on soil-borne bacteria; therefore, the main habitat linking bacteria and plants is the soil under the influence of plant roots, so called rhizosphere (Hiltner, 1904). The picture of grand unculturability of microbiota (Sharma et al. 2005) apears to hold for the rhizosphere environment. Molecular techniques have demonstrated an enormous unexplored reservoir of microorganisms in the rhizosphere (Duineveld et al., 2001; Buée et al., 2009). Acidobacteria and Verrucomicrobia may be part of dominant rhizosphere populations in different plant species, soil types and geographic locations, e.g. in natural forest (Lin et al., 2010a; Chan et al., 2006) and agricultural systems (Kielak et al., 2008; Ulrick & Becker, 2006), and even plant-soil microcosm experiments (De Cárcer et al., 2007; DeAngelis et al., 2009). Besides, it has been shown that these two groups can also be active in their habitat. For instance, members of Acidobacterium were shown to be metabolically active in the rhizosphere of C. crenata, suggesting an ecological role for these in this habitat (Lee et al., 2008). Members of Verrucomicrobium were shown to be active as part of the microbial community
18 Chapter 1
assimilating rape root exudates (Haichar et al., 2008), indicating that these bacteria may find their niche in the rhizosphere and possibly interact with plants. These reports pointed to a dedicated role of members of both Acidobacterium and Verrucomicrobium in the rhizosphere.
Objectives
The general aim of this study was to advance the understanding of the prevalence and ecological role of hitherto poorly-culturable bacterial groups in the rhizosphere, with a special focus on members of Acidobacterium and Verrucomicrobium. As providers of the rhizosphere, we selected the monocotyledon leek and the dicotyledon potato. Given the fact that the availability of isolates would enable to do behavioral studies, a prime focus was placed on the acquisition of isolated strains. The focused objective was then to determine if the selected strains, representatives of particular Acidobacterium and Verrucomicrobium subgroups, could be shown to occur in the rhizosphere and whether they would show preference for the rhizosphere over bulk soil. To achieve the objectives, a combination of culturing and culture-independent techniques was developed and applied.
Approach
A debate about which techniques needed to investigate the functioning of natural microbial communities currently challenges the community of microbial ecologists (Donachie et al., 2007; Giovannoni and Stingl, 2007; Nichols, 2007). After more than a century of successful cultivation-based studies, the limitations of these approaches have become apparent. This has coincided with the emergence of the now called ‘omics’ era, which has evolved from both the rRNA gene analyses and the direct DNA/RNA based microbial community analyses, both developed 20-30 years ago. In spite of the fact that technical hurdles actually still hinder the application of different ‘omics’ techniques, such as metagenomics, much has been achieved; on the other hand, significant gaps in microbial community diversity data still persist (DeLong, 2005; DeLong e al., 2006; Schirmer et al., 2005; Tringe et al., 2005; Tyson et al., 2004; Venter et al., 2004; ). For instance, although we can assess the distribution of different ‘as-yet-uncultured’ bacteria, we can not easily infer the ecological behaviour of these organisms from culture-independent studies (Sharma et al., 2005). In this thesis, I used an initial culture- dependent step to first explore those representatives of Acidobacterium and Verrucomicrobium that were culturable from the rhizosphere habitat. Later, a range of
19 General Introduction
culture-independent techniques (i.e. group-specific real time PCR systems) were developed, validated and used to describe the population dynamics of selected strains in the selected habitat. Fore the latter, I used a plethora of experimental systems, from the laboratory to the open field.
Working hypothesis
The following hypothesis were used as base line for this thesis: (i) hitherto-uncultured Acidobacteria and Verrucomicrobia are part of the dominant bacterial community in the rhizosphere; (ii) these hitherto-uncultured bacteria play an important role in the plant- soil ecosystem; (iii) due to their recalcitrance to grow in laboratory conditions, a combination of innovative culture-dependent and -independent techniques will facilitate the study of Acidobacteria and Verrucomicrobia in the rhizosphere environment.
Research questions
The following research questions were formulated to reach the aforementioned thesis objectives:
1. What hitherto-unculturable bacteria dominate in the rhizosphere environment?
2. What strategy would allow to significantly enhance the culturability of bacteria from rhizosphere samples?
3. Can members of Acidobacterium and Verrucomicrobium be isolated from the rhizosphere of distantly-related plants (the monocotyledon leek and the dicotyledon potato)?
4. Would members of Acidobacterium and Verrucomicrobium that are recovered from different plant species and soils be phylogeneticaly related?
5. Do different subgroups of Acidobacterium and Verrucomicrobium exhibit differences in their preference of bulk and rhizosphere soil?
6. Which environmental factors influence the distribution of the different Acidobacterium and Verrucomicrobium subgroups over rhizosphere and bulk soil?
7. Is the ecology of Acidobacterium and Verrucomicrobium related to plants?
20 Chapter 1
Outline of the thesis
Chapter 1 briefly introduces the following concepts: great plate count anomaly, poorly- uncultured bacteria groups (i.e. Acidobacteria and Verrucomicrobia) and their implications on the rhizosphere ecology. In chapter 2, I review the state of the art concerning the ecology of hitherto- uncultured bacteria of the rhizosphere environment, focusing on Acidobacteria, Verrucomicrobia and Planctomycetes. These groups were among the most dominant ones in the rhizosphere, at the same time revealing the lowest numbers of cultured strains. Furthermore, I show that culture-independent techniques still have technological hurdles that hamper a thorough analysis of the rhizosphere microbiota. Therefore, a strategy was proposed to recover bacteria from the rhizosphere that affiliate with Acidobacterium and Verrucomicrobium. This strategy was based on the use of low- nutrient media in combination with long incubation periods and the addition of oxidative stress protective agents or even substances that might better mimic the rhizosphere, such as rhizosphere extract. The approach to recover hitherto-uncultured bacteria from the potato rhizosphere was evaluated in chapter 3. The use of low-carbon-availability media led to the recovery of up to 33.6% of the total estimated cell numbers, which is at least seven- fold the recovery on R2A. Also, this approach led to the isolation of the first Verrucomicrobia subdivision 1 strains from the potato rhizosphere. Following this success, in Chapter 4 it was demonstrated that members of Acidobacterium as well as Verrucomicrobium can be obtained from the leek rhizosphere. In this chapter, I describe the isolation of two Holophaga spp. (Acidobacterium) and nine Verrucomicrobium subdivision 1 strains. Following isolation, these Acidobacterium and Verrucomicrobium strains, next to those described in chapter 3, were characterized using a suite of biochemical, physiological, genetic and cell structural approaches. The Holophaga spp. strains demonstrated similar phenotypic characteristics, indicating that they may occupy the same ecological niche. The phenotypic diversity within the Verrucomicrobium subdivision 1 strains isolated from the leek (chapter 4) and potato rhizospheres (chapter 3) indicates that these isolates occupy different ecological niches in the soil- plant system. Finally, Candidatus genus Rhizospheria was proposed as a novel taxonomic denotation of our strains that affiliated with (as yet) unclassified Verrucomicrobiaceae. The novel Holophaga (Acidobacterium) and Verrucomicrobium subdivision 1 strains, although in culture, were shown to be fastidious (or hard to culture) under laboratory growth conditions. Therefore, in chapter 5, specific real time PCR systems were developed for detection of (each of) Holophaga, Luteolibacter and Candidatus genus Rhizospheria. This would facilitate subsequent ecological studies of these
21 General Introduction
bacteria in the plant-soil environment. The specificity of the primers designed was evaluated in three steps, i.e. in silico, following PCR amplification on genomic DNA extracts from target and non-target bacteria and similarly from soil DNA extracts. Furthermore, real-time PCR quantification of Holophaga, Luteolibacter and Candidatus genus Rhizospheria numbers was applied to leek rhizosphere compartments and bulk soil. A higher preference for leek rhizosphere compartments above bulk soil was found for all three bacterial groups. To evaluate if the novel Holophaga and Verrucomicrobium subdivision 1 strains have their ecological niche in the rhizosphere, their population dynamics was studied in plant-soil microcosms. In Chapter 6, we describe such experiments. The two Holophaga sp. strains CHC25 and ORAC were shown to colonize in vitro roots. Furthermore, Holophaga sp. CHC25 was selected for studies on dynamics and migration in the plant-soil microcosm. Both Holophaga sp. strain CHC25 and the Holophaga natural populations consistently revealed raised numbers at the leek roots compared to bulk soil. This phenomenon occurred both locally due to growth, but was also due to migration of strain CHC25 towards the roots. In chapter 7, Luteolibacter sp. strain CHC12 and Candidatus genus Rhizospheria (Verrucomicrobium subdivision 1) were tested for their ability to grow in the rhizosphere in comparison with the bulk soil. The data shown in this chapter indicated that the natural populations of Luteolibacter (akin to strain CHC12) had lower numbers in the rhizosphere than in corresponding bulk soil. On the other hand, Candidatus genus Rhizospheria (akin to strain CHC8) showed higher numbers at leek roots than in bulk soil. As observed for Holophaga, the raised numbers were not only the result of in situ cell multiplication but also to migration of cells from distant sites in the soil towards the roots. Therefore, these Holophaga and Luteolibacter strains may represent rhizosphere-competent bacteria, opposing what was observed for Candidatus genus Rhizospheria in plant soil microcosms. To evaluate the environmental factors that influence the distribution of different Acidobacterium and Verrucomicrobium types, a field experiment was performed, as described in chapter 8. In this experiment, potato and leek were planted in a spot in pasture land. The most dominant Acidobacterium subgroups, i.e. 1, 3, 4 and 6 (Jones et al., 2009), and Holophaga and Verrucomicrobium subdivision 1 groups (isolated in chapter 3 and 4) were targets in this study. For the detection of the latter, the specific real time PCR detection systems developed in chapter 5 were used. Novel group- specific real time PCR systems targeting Acidobacteria subgroups 1, 3, 4 and 6 were designed like in chapter 5. I found that the differences between rhizosphere and bulk soils were the main driving forces that differentiated the numerical distributions of Acidobacterium and Verrucomicrobium subdivision 1. This was followed by plant type and, to a minor extent, by pH. Moreover, different subgroups of Acidobacterium are
22 Chapter 1
differentially influenced by the rhizosphere effect. Different from Acidobacterium, Verrucomicrobium subdivision 1 numbers were higher in the rhizosphere compartments than in the respective bulk soil independent of the plant type. Chapter 9 presents a general discussion of the findings of this thesis, integrates the different chapters and proposes an outlook to future studies.
23 General Introduction
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26 Chapter 1
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27 General Introduction
Tringe SG, Von Mering C, Kobayashi A et al. (2005) Comparative metagenomics of microbial communities. Science 308: 554-557. Tyson GW, Chapman J, Hugenholtz P et al. (2004) Community structure and metabolism through reconstruction of microbial genomes from the environment. Nature 428: 37-43. Ulrich A & Becker R (2006) Soil parent material is a key determinant of the bacterial community structure in arable soils. FEMS Microbiol Ecol 56: 430-443. Van Elsas JD, Kastelein P, van Bekkum P, van der Wolf J M, de Vries PM & van Overbeek LS (2000) Survival of Ralstonia solanacearum biovar 2, the causative agent of potato brown rot, in field and microcosm soils in temperate climates. Phytopathology 90: 1358–1366. Venter JC, Remington K, Heidelberg JF et al. (2004) Environmental Genome Shotgun Sequencing of the Sargasso Sea. Science 304: 66-74. , Jeppsson B & Molin G (2005) Comparison of bacterial diversity along the human intestinal tract by direct cloning and sequencing of 16S rRNA genes. FEMS Microbiol Ecol 54: 219- 231. Ward-Rainey N, Rainey FA, Schlesner H & Stackebrandt E (1995) Assignment of hitherto unidentified 16S rDNA species to a main line of descent within the domain Bacteria. Microbiol 141: 3247-3250. Whitman et al. (1998) Proc Natl Acad Sci USA 95: 6578-6583. Wise et al. (1997) Bacterial diversity of a Carolina Bay as determined by 16S rRNA gene analysis: confirmation of novel taxa. Appl Environ Microbiol 63: 1505-1514. Yoon J, Matsuo Y, Adachi K, Nozawa M, Matsuda S, Kasai H & Yokota A (2008a) Description of Persicirhabdus sediminis gen. nov., sp. nov., Roseibacillus ishigakijimensis gen. nov., sp. nov., Roseibacillus ponti sp. nov., Roseibacillus persicicus sp. nov., Luteolibacter pohnpeiensis gen. nov., sp. nov. and Luteolibacter algae sp. nov., six marine members of the phylum ‘Verrucomicrobia’, and emended descriptions of the class Verrucomicrobiae, the order Verrucomicrobiales and the family Verrucomicrobiaceae. Int J Sys Evol Microbiol 58: 998-1007. Yoon J, Matsuo Y, Matsuda S, Adachi K, Kasai H & Yokota A (2007a) Cerasicoccus arenae gen. nov., sp. nov., a carotenoid-producing marine representative of the family Puniceicoccaceae within the phylum 'Verrucomicrobia', isolated from marine sand. Int J Sys Evol Microbiol 57: 2067-2072. Yoon J, Matsuo Y, Matsuda S, Adachi K, Kasai H & Yokota A (2007b) Rubritalea spongiae sp. nov. and Rubritalea tangerina sp. nov., two carotenoid- and squalene-producing marine bacteria of the family Verrucomicrobiaceae within the phylum Verrucomicrobia, isolated from marine animals. Int J Sys Evol Microbiol 57: 2337-2343. Yoon J, Matsuo Y, Matsuda S, Adachi K, Kasai H & Yokota A (2008b) Rubritalea sabuli sp. nov., a carotenoid- and squalene-producing member of the family Verrucomicrobiaceae, isolated from marine sediment. Int J Sys Evol Microbiol 58: 992-997. Yoon J, Matsuo Y, Katsuta A, Jang J-H, Matsuda S, Adachi K, Kasai H & Yokota A (2008c) Haloferula rosea gen. nov., sp. nov., Haloferula harenae sp. nov., Haloferula phyci sp. nov., Haloferula helveola sp. nov. and Haloferula sargassicola sp. nov., five marine representatives of the family Verrucomicrobiaceae within the phylum 'Verrucomicrobia'. Int J Sys Evol Microbiol 58: 2491-2500. Yoon J, Oku N, Matsuda S, Kasai H & Yokota A (2007c) Pelagicoccus croceus sp. nov., a novel marine member of the family Puniceicoccaceae within the phylum 'Verrucomicrobia' isolated from seagrass. Int J Sys Evol Microbiol 57: 2874-2880. Yoon J, Yasumoto-Hirose M, Katsuta A, Sekiguchi, H, Matsuda S, Kasai H & Yokota A (2007d) Coraliomargarita akajimensis gen. nov., sp. nov., a novel member of the phylum 'Verrucomicrobia' isolated from seawater in Japan. Int J Sys Evol Microbiol 57: 959-963. Yoon J, Yasumoto-Hirose M, Matsuo Y, Nozawa M, Matsuda S, Kasai H & Yokota A (2007e) Pelagicoccus mobilis gen. nov., sp. nov., Pelagicoccus albus sp. nov. and Pelagicoccus litoralis sp. nov., three novel members of subdivision 4 within the phylum 'Verrucomicrobia', isolated from seawater by in situ cultivation. Int J Sys Evol Microbiol 57: 1377-1385. Zimmermann J, Gonzalez JM, Saiz-Jimenez C & Ludwig W. (2005) Detection and phylogenetic relationships of highly diverse uncultured acidobacterial communities in Altamira cave using 23S rRNA sequence analyses. Geomicrobiol J 22: 379–388. Zul D, Denzel S, Kotz A & Overmann J (2007) Effects of plant biomass, plant diversity, and water content on bacterial communities in soil lysimeters: Implications for the determinants of bacterial diversity. Appl Environ Microbiol 73: 6916-6929.
28
Chapter 2: Exploration of hitherto-uncultured bacteria from the rhizosphere*
Abstract
The rhizosphere environment selects a particular microbial community that arises from the one present in bulk soil due to the release of particular compounds in exudates and different opportunities for microbial colonization. During plant-microbe co-evolution, microbial functions supporting plant health and productivity have developed, of which most are described in cultured plant-associated bacteria. This chapter discusses the state-of-the-art concerning the ecology of hitherto-uncultured bacteria of the rhizosphere environment, focusing on Acidobacteria, Verrucomicrobia and Planctomycetes. Furthermore, a strategy is proposed to recover bacterial isolates from these taxa from the rhizosphere environment.
* Authored by:Ulisses Nunes da Rocha, Leonard Simon van Overbeek & Jan Dirk van Elsas Published in: FEMS Microbiology Ecology (2009) 69: 313-328 Hitherto-uncultured bacteria from the rhizosphere
Introduction
The rhizosphere, here defined as the volume of soil over which plant roots exert an influence (Hiltner, 1904), differs from bulk soil due to the biophysicochemical processes that occur as a consequence of root growth, water and nutrient uptake, respiration and rhizodeposition (Hinsinger, 1998). The rhizosphere extends from the surface of the plant root to a position in soil which depends on the diffusion rate of exudates and the roots’ biochemistry and development (Hinsinger et al., 2005). This extension also varies with plant type and microbial community composition (Huisman, 1982; Watt et al., 2006; Watt et al., 2006). Besides, as the rhizosphere is known to contain strong gradients of compounds from the root surface into the soil, any rhizosphere sample will inevitably average out the effects of such gradients. In fact, the further away from the root the soil is sampled, the lower the rhizosphere effect on the sample will be. Thus, plant roots create selective pressures that influence the local microbial communities, resulting in effects on their abundance and composition. These selective pressures do not depend only on biological processes, but also on abiotic ones, such as temperature and water content. For instance, seasonal and daily temperature changes have been found to affect microbial activities (Turpault et al., 2007; Gaumont- Guay et al., 2008) and community composition (Pandey et al., 2001). Water content is also important as it directly influences the microbiota in soils due to the high correlation of microbial activity and soil moisture (Krivtsov et al., 2007). Different studies demonstrated that soil structure, as well as different granulometric fractions determine the presence of different bacterial populations (Sessitsch et al., 2001; Kotani-Tanoi et al., 2007); these factors might influence the microbial communities present in the rhizosphere depending on the soil where the root developed. Also, pH and CO2/O2 tensions will influence the rhizosphere microbial community functional diversity and activity (Jossi et al., 2006; Nelson & Mele, 2006). The organisms thriving in the rhizosphere encompass a range of different taxa, including prokaryotic and eukaryotic microorganisms. Most abundant among these groups are the bacteria and fungi. Usually, the bacterial fraction ranges from 109-1010 cells g-1 of soil and approximately 106 cells mm-3 of rhizosphere biofilm in most cases (Watt et al., 2006). The description of the bacterial taxa present in the rhizosphere will be discussed later. Key ecological functions can be ascribed to the different bacterial populations that thrive in the rhizosphere environment. For instance, bacteria with nitrogen-fixing and phosphorus-dissolving activities provide nitrogenous compounds and available phosphorus to plant roots, thus increasing the growth of plants, for instance in organic agriculture (Canbolat et al., 2006; Hameeda et al., 2008). Also, the production of particular phytohormones by rhizosphere bacteria can enhance plant growth (Lee &
30 Chapter 2
Song, 2007). The activity of the enzyme 1-aminocyclopropane-1-carboxylate (ACC) deaminase in particular plant-associated bacteria such as Burkholderia phytofirmans has important implications for rhizosphere functioning as well (Hardoim et al., 2008). In addition, the rhizosphere microbiota is important, as a wide range of bacteria isolated from this environment has already shown to act as biocontrol agents of plant pathogens; for instance those producing antimicrobial compounds or eliciting plant defence reactions (Sfalanga et al., 1999; Földes et al., 2000; Jiménez-Esquilín & Roane, 2005; Flores-Vargas & O'Hara, 2006; Romanenko et al., 2007). The ‘great plate count anomaly’ states that 95 to 99% of the microbial community present in the environment is not readily accessible by traditional culture techniques (Nichols, 2007). This fraction of the microbial community has been coined the uncultured microbiota. Given the fact that a greater fraction of the total soil microbiota may be culturable than thought before, we will here address this fraction as the hitherto-unculturable bacteria. Most often, the biogeochemical transformations observed in soil have been ascribed to well-characterized culturable bacteria, simply because their ecological relevance has driven a plethora of studies based on culturable bacteria. The large gap between what we know about the culturable microbial diversity versus the hitherto-unculturable members of these communities is illustrated in Fig. 1. The fraction of culturable with regard to total representatives differs for each prokaryotic group. This gap also demonstrates our lack of knowledge about the putative functions hidden in the hitherto-uncultured microbial fractions of particular groups that are involved in rhizospheric microbial-microbial and microbial-plant interactions and pinpoints the need to invest in this basic area of research. One key question is whether members of the uncultured majority are actually involved in key rhizosphere processes. Recently, using stable isotope probing-RNA analysis, a broad new range of inhabitants of the rhizosphere was found, that were active under the experimental conditions, but did not have their specific role in the system identified (Vandenkoornhuyse et al., 2007). Moreover, the current ‘omics’ techniques (here defined as integrative studies of biological systems, including genomics, transcriptomics, proteomics and metabolomics) offer good perspective in describing function or potential function in environmental studies. For instance, proteomics-based studies have already revealed the abundance of particular proteins in the rhizosphere, such as glycoside hydrolases, trypsin/protease inhibitors, plastocyanin-like domains, copper-zinc superoxide dismutases and plant basic secretory proteins (Kiely et al., 2006). Also, metagenomics studies of the rhizospheres of plants growing in acid mine drainage revealed several novel microbial genes that are possibly involved in heavy metal resistance to their bacterial hosts (Mirete et al., 2007). By using a metabolomics approach, it was shown that the
31 Hitherto-uncultured bacteria from the rhizosphere different udwig et Bacteroidetes-Chlorobi Planctomyces Verrucomicrobia Acidobacteria Firmicutes Fibrobacteres nces of total (culturable and nces of total (culturable and Deltaproteobacteria Epsilonproteobacteria irs in length), derived from Actinobacteria Unclassified Alphaproteobacteria Cyanobacteria s were constructed in the ARB program (L 0.10 Crenarchaeota Euryarchaeota
B represented by 16S rRNA gene seque ected sequences (1200 base pa Beta_Gammaproteobacteria Acidobacteria Planctomyces Verrucomicrobia Bacteroidetes-Chlorobi Firmicutes TM6 OP8 OP3 OP9 Fibrobacteres base, release 94 (Pruesse et al., 2007). Tree type strains only (B). Sel Deltaproteobacteria between selected prokaryotic groups ity distance of 10% between sequences. Actinobacteria Epsilonproteobacteria OP2 OP10 OP5 OPB7 Cyanobacteria Alphaproteobacteria OD1 -OP11 -WS6 -TM7 -WS6 -OP11 OD1 0.10 Euryarchaeota ta A Phylogenetic relationship Beta_Gammaproteobacteria renarchaeo C Figure 1 al., 2004) and bars indicate a similar uncultured) bacteria (A) and of culturable ecosystems, originated from the SILVA data
32 Chapter 2
microbial populations in the rhizosphere were able to metabolize uncommon nutrients exuded by plants (Narasimhan et al., 2003). Even though the omics techniques offer great advances in our capabilities to unravel the identity of genes present in the rhizosphere microbiota, there still are technological hurdles that hamper a thorough analysis. For instance, full genomes can only be assembled from dominant species in natural habitats, and only ‘mosaic’, instead of single-species genomes, may be assembled due to the presumed occurrence of closely-resembling species in the same habitat (Tyson et al., 2004; Venter et al., 2004). In rhizosphere research, time course data are of utmost importance due to the drastic changes occurring in associated microbial communities during plant growth. Thus, in cases where shotgun sequencing is used, the sequence information only will relate to a momentary snap shot of the dominant sequences present in the DNA extract, and information on community development in the rhizosphere is often lacking. Also, given the heterogeneity occurring in all natural environments, the interpretation of metagenomics data will be complicated due to the expected variations within residing communities (Tyson et al., 2004; Tringe et al., 2005). Moreover, it is questionable what knowledge of the genome of particular species populations actually means given the expected genomic heterogeneity within a species, where genomic rearrangements, deletions or insertions may abound (Tringe et al., 2005). Metagenomics will thus provide, at best, information at the level of the prevalence of genes and will depend on the availability of information about the functions of such genes in cultured species (DeLong, 2005; Schirmer et al., 2005; DeLong et al., 2006). The increase in the number of genes with as-yet-unknown functions is keeping pace with the amount of metagenomic information that is becoming available over time. As long as no attempts are made to unravel the function of the many ‘unidentified’ or ‘hypothetical’ proteins commonly found in metagenomic databases, progress in the elucidation of the roles of uncultured microorganisms in ecosystem functioning will be hampered.
In this chapter, the following questions are posed: x What are the key interactions among soil, plant and the local microbial communities? x Which are the dominant hitherto-unculturable bacteria in the rhizosphere environment? x Can rhizosphere-relevant functions of these organisms in their natural habitat be revealed by the actual knowledge in literature? x How to culture the hitherto-unculturable bacteria from rhizosphere environment? x How to reveal the basic ecology of the hitherto-unculturable populations in the rhizosphere?
33 Hitherto-uncultured bacteria from the rhizosphere
Soils, plant roots and microbial communities – an intricate triplet
Plant-soil-microbial community systems have evolved ever since plants started to colonize the terrestrial environment, around 700 million years ago. The first interactions between soil microbial communities and plant root systems may not have been beneficial to plants, as they may have mainly consisted of attacks to the roots by microorganisms, especially bacteria (Phillips et al., 2003). However, plants colonizing terrestrial environments, being “bathed” in microorganisms, have also initiated evolutionarily-useful interactions with microorganisms, for instance to optimize growth and reproduction. Such - initially primitive - interactions are thought to have resulted in the evolution of all currently known mutually beneficial relationships (Phillips et al., 2003). Classical examples of such interactions are the intricate associations of both rhizobia and mycorrhizal fungi with plant roots. Fossilized remains of primordial root systems also provide evidence that epiphytic and endophytic microbial populations already existed on/in the primitive land plants, although the nature of these associations has not been elucidated (Phillips et al., 2003). We here postulate that, in particular highly-selected cases, the co-evolution of plants and microbes in soil has also resulted in firm and dedicated relationships between particular players in the rhizosphere. For one, the belowground competition for soil nutrients between plants certainly has exerted selective pressure that has driven the emergence of plants with wide variations in root structure and function, adapted to perform well under the different conditions in soil. As a consequence, the plant- associated microbial communities may have evolved to the actual scenery of positive, negative and neutral microbe-microbe and microbe-plant interactions. It is known that current-day plant roots and their associated microorganisms are highly affected by soil conditions, which determine the distribution, density and depth of roots in various soils (Passioura, 1991; Jackson et al., 1996; Stewart et al., 1999; Jobbágy & Jackson, 2000). Thus, firstly, rhizosphere microbial community structure and diversity, in different soil textural types, are strongly determined by the types of plant roots (Garbeva et al., 2004b; Kotani-Tanoi et al., 2007). It is known that during root growth, local soil characteristics, for instance gas composition and water flow, are increasingly affected by the roots (Colmer, 2003; Lipiec et al., 2007; Benedict & Frelich, 2008; Whalley et al., 2008). In addition, carbonaceous compounds are often abundantly released from roots into the rhizosphere. These factors, in turn, will influence the root-associated microbial communities. Moreover, effects in the opposite way also occur, e.g. the modulation of root architecture by microorganisms that are locally present, for instance via the production of phytohormones (Belimov et al., 2009). Some of the relevant interactions between plants and microorganisms, as well as among different microorganisms in the rhizosphere are summarized in Fig. 2.
34 Chapter 2
Plant (roots)
Growth Secondary Mass metabolites Density (exudates) Depth
Influence
Influence Nutrient uptake Degradation of complex polymers
Antimicrobial compounds Soil structure & texture Production of Exopolissacharides Influence Nutrient availability Phytohormones Gas composition & exchange Rhizosphere Soil microbial community composition
Figure 2 Interrelationship between plant, soil and microbial communities. Arrows demonstrate the most relevant interactions occurring between the different players in the rhizosphere.
Many transformations in nutrient cycles that commonly occur in the rhizosphere are still unresolved, whereas others are attributed to microbial activities. Examples of microbial activities in nutrient cycles are for instance: nitrogen fixation and demineralisation (Canbolat et al., 2006) and solubilisation of phosphorus (Canbolat et al., 2006) and carbohydrates (Kohler et al., 2006). Plants and other soil micro-organisms profit from these transformations occurring in the rhizosphere. Examples of other interactions commonly occurring in the rhizosphere are: commensalism, e.g. by creation of new niches for microorganisms via the secretion of exopolysaccharides (Kaci et al., 2005; Haggag, 2007) or via production of phytohormones (Lee & Song, 2007); antagonism via production of secondary metabolites with antibiotic activity (Garbeva et al., 2004b; Berg et al., 2005; Costa et al., 2007) and syntrophism, e.g. via degradation of toxic compounds like recalcitrant hydrocarbons and herbicides (Biryukova et al., 2007; Vaishampayan et al., 2007). Often, a multitude of microbial species is responsible for particular conversions in the rhizosphere and hence a particular degree of functional redundancy may exist for important processes in the rhizosphere. For instance, several different microbial groups are known to be involved in ammonia oxidation in the rhizosphere (Nicolaisen et al., 2004; Herrmann et al., 2008).
35 Hitherto-uncultured bacteria from the rhizosphere
However, all of these studies have so far ignored the role of the major part of the hitherto-unculturable soil microbiota. Exploring the hitherto-unculturable bacteria by improvement of culturability presumably will reveal a range of novel functions in the rhizosphere, including functions involved in enhancing rhizosphere competence. An examination of the bacterial groups that are most commonly found in rhizosphere environments and our understanding about their presumed roles in these environments will be discussed further.
Dominant hitherto-uncultured bacteria in the rhizosphere
Soil bacteria with comparable metabolic capacities will respond in similar fashions to the emergence of different plants, provided that (nutritional) conditions are similar. However, it is still unclear to what extent commonality exists in the microbial communities that associate with plant roots across plant genera, species or cultivars. This is mostly due to the fact that little attempts were made to synthesize data available in the literature into a systematic evaluation on the relationships between microbial communities, plants, soil types and geographical locations. Given this limitation, we performed such a limited study on data in the literature to indicate the most dominant bacterial groups in the rhizosphere using the following procedure: (i) Collation of an inventory of culture-independent studies that addresses the rhizosphere bacterial diversity across a broad array of plant types, (ii) Identification of the most dominant phyla in the rhizosphere as found in each study, (iii) Singling out the most dominant groups as the ‘most commonly found’ taxa in the rhizosphere by frequency of detection across all studies. Recent studies that fit all the aforementioned procedures were selected using three different databases, i.e. Scopus (www.scopus.com), Web of Science (isiknowledge.com) and Pubmed (www.pubmedcentral.nih.gov) (Kaiser et al., 2001; Kuske et al., 2002; Gremion et al., 2003; Schmalenberger & Tebbe, 2003; Graff & Conrad, 2005; Sharma et al., 2005; Stafford et al., 2005; Jossi et al., 2006; Poonguzhali et al., 2006; Sanguin et al., 2006; Fierer et al., 2007; Villadas et al., 2007; Wang et al., 2007; Zhang et al., 2007; Andreote et al., 2008; Bharathkumar et al., 2008; Elliott et al., 2008; Garbeva et al., 2008). From these studies, it was concluded that in total seven bacterial phyla were most dominant in the rhizospheres examined. These were the following: Proteobacteria, in particular the Alfa, Geta and Gamma-proteobacteria, Actinobacteria, Acidobacteria, Verrucomicrobia, Planctomycetes, Bacteriodetes and Firmicutes (Fig. 3).
36 Chapter 2
Culture-dependent Culture-independent approaches approaches
Į-Proteobacteria (16013)
ȕ-Proteobacteria (11100)
Ȗ-Proteobacteria (30689)
Actinobacteria (15733)
Acidobacteria (2473)
Verrucomicrobia (1674)
Planctomycetes (1821)
Bacterioidetes (22901)
Firmicutes (59132)
100 75 50 25 0 25 50 75 100 Percentage of total sequences in the database
Figure 3 Percentage of 16S rRNA gene sequences from bacterial groups common to the rhizosphere and obtained via culture-dependent and culture-independent approaches. All sequences (1200 base pairs in length), derived from different ecosystems, were obtained from the RDP Query database (Cole et al., 2007), release 9.60 (latest accession, April 22nd, 2008). Between brackets; total number of sequences used per bacterial group.
Within these phyla, the origin of the 16S rRNA gene sequences, i.e. whether these were obtained from isolates or uncultured organisms, was taken into account; it was thus found that the contribution of isolates to the total number of sequences (redundant or not) per bacterial group was variable. For Proteobacteria, the taxonomical information available from isolates was about the same as that from uncultured organisms. However, for Actinobacteria the available taxonomical information was even higher for isolates, reflecting better adaptation of this group for growth in pure culture. For the other five groups, by far the highest amount of phylogenetic information was derived from uncultured organisms, i.e. most organisms occurring in these groups are regarded as as-yet-uncultured or plainly uncultured bacteria. Among these groups,
37 Hitherto-uncultured bacteria from the rhizosphere
the only knowledge in respect of rhizospheric Acidobacteria, Verrucomicrobia and Planctomycetes was obtained on the basis of culture-independent studies. Hence, our understanding of these taxa in the rhizosphere is still limited, as it concerns mainly their 16S rRNA gene fluctuations in the rhizosphere (Sanguin et al., 2006; Zul et al., 2007). From the three groups, the Acidobacteria and Verrucomicrobia have a low number of species recognized in the literature. To date, the Acidobacteria contain three early-described species, i.e. Acidobacterium capsulatum, Geothrix fermentans and Holophaga foetida (Garrity et al., 2005), next to four recently-recognized cultured species (Edaphobacter aggregans, E. modestus, Chloracidobacterium thermophilum and Terriglobus roseus (Bryant et al., 2007; Eichorst et al., 2007; Koch et al., 2008). The Verrucomicrobia show a similar picture, with only ten defined species, i.e. Verrucomicrobium spinosum, four Prosthecobacter spp., Opitutus terrae Rubritalea marina and three Xiphinematobacter sp. (Ward-Rainey et al., 1995; Garrity et al., 2005). Although some representatives of the Acidobacteria and Verrucomicrobia have now been cultured from soil (Janssen et al., 2002; Sait et al., 2002; Stevenson et al., 2004; Davis et al., 2005), no isolates have as yet been obtained from the rhizosphere. Isolation directly from the rhizosphere is clearly needed to provide supportive data that these isolates are really rhizosphere-relevant organisms. Their availability would allow an assessment of their influences on plant growth, as well as of their interaction with plants and other soil microorganisms. Also, their involvement in important biogeochemical conversions, in the degradation of particular compounds and in the mobilization of nutrients would be facilitated. The members of the Acidobacteria and Verrucomicrobia that have previously been isolated from soil were slow growing, aerobic and heterotrophic bacteria (Janssen et al., 2002; Sait et al., 2002; Stevenson et al., 2004; Davis et al., 2005). Such characteristics are broadly distributed among rhizosphere isolates, which indicates that such isolates may thrive in the rhizosphere environment. The abundance of hitherto-unculturable bacteria recognized solely by their phylogeny in the rhizosphere (Kaiser et al., 2001; Kuske et al., 2002; Gremion et al., 2003; Schmalenberger & Tebbe, 2003; Graff & Conrad, 2005 Sharma et al., 2005; Stafford et al., 2005; Sanguin et al., 2006; Wang et al., 2007; Zhang et al., 2007; van Elsas et al., 2008) urges us to address their functional roles. However, information about their putative roles or interactions with the plant is as yet very sparse. Each of the organisms involved might, for instance, occupy a particular niche in the rhizosphere. It is a challenge to develop strategies – based on cultivation dependent and independent studies – that allow the exploration of the putative roles and function of the hitherto- unculturable microbiota in the rhizosphere, in particular analyzing their modes of interactions with plants and/or rhizosphere microorganisms.
38 Chapter 2
Ecology of hitherto-unculturable bacteria in the rhizosphere
If particular niches exist for the hitherto-unculturable bacteria in the rhizosphere, it is logical to assume that selective forces present within such niches act on these rhizosphere populations. Such a selection might be nutritional, dependent on the availability of oxygen or other electron acceptors, or otherwise. To our knowledge, no conclusive data have extensively shown the factors that influence the fate and behaviour of these populations in the rhizosphere. Therefore, the following points should be addressed to elucidate the roles of these organisms in the rhizosphere: i) their general occurrence and numerical dominance, ii) the ecological conditions driving these populations, iii) in situ (local) metabolic activities and iv) their functional roles in the soil-plant ecosystem. When comparing bulk and rhizosphere soil, the general occurrence and numerical abundances of the Acidobacteria, Verrucomicrobia and Planctomycetes are often variable, as demonstrated in culture-independent studies based on 16S rRNA gene detection (Dunbar et al., 1999; Kuske et al., 2002; Gremion et al., 2003; Filion et al., 2004; Stafford et al., 2005; Sanguin et al., 2006; De Cárcer et al., 2007; Singh et al., 2007; Zul et al., 2007; Kielak et al., 2008). Specifically, a higher prevalence of 16S rRNA gene clones affiliated with particular groups of Acidobacteria, especially those of subgroups 1, 2 and 3 were found in rhizospheres of lodgepole pine (Chow et al., 2002), Proteacea sp. (Stafford et al., 2005) and grass (Chow et al., 2002; Stafford et al., 2005; Singh et al., 2007) as compared to corresponding bulk soils. Also, a higher number of 16S rRNA gene clones of Verrucomicrobia was observed in rhizospheres of lodgepole pine, in particular of subdivisions 2, 3 and 4 (Chow et al., 2002), Thalaspi caerulescens, subdivision 2 (Gremion et al., 2003) and Proteacea sp., subdivision 3 (Stafford et al., 2005) in comparison with the corresponding bulk soils. Besides, Planctomycete representatives were found in higher numbers in the rhizospheres of Thlaspi caerulescens than in the bulk soil with 16S rRNA gene clones affiliating only with the Nostocoida limilula III cluster (Gremion et al., 2003) and Proteacea sp. (Stafford et al., 2005) (Table 1). In contrast, a study performed with different plants demonstrated higher numbers of Acidobacteria in a 16S rRNA gene clone libraries made from bulk soil DNA than in the ones made from rhizosphere DNA, although no information was given about the dominant groups in both soil compartments (Kielak et al., 2008). Also, the rhizosphere of maize plants showed lower numbers of Acidobacteria, Verrucomicrobia and Planctomycetes than corresponding bulk soil; and again no information was given about the dominant subdivisions (Sanguin et al., 2006). In Thlaspi caerulescens (Gremion et al., 2003) different Acidobacteria groups were found in bulk and rhizosphere soil, although the number of detected 16S rRNA genes was the same in both soil compartments. In this study, more clones were found to be affiliated
39 Hitherto-uncultured bacteria from the rhizosphere , 2003 , 2003
et a.l Chow et al., 2002 2002 al., et Chow Reference Kuske et al., 2002 2002 al., et Kuske Gremion ,
and and
, Acidobacteria Planctomycetes and and representatives. Verrucomicrobia , Planctomycetes Acidobacteria and and Planctomycetes representatives. No significant effects of soil disturbance effects of soil disturbance No significant location. or geographical selective a strong roots exerted Plant on pressure Verrucomicrobia The composition of Acidobacterium of Acidobacterium The composition soil with differed members division of or the presence plants and with depth top soil layer. on crust cyanobacterial Plant roots exerted a strong selective selective a strong roots exerted Plant of diversity the on pressure Acidobacteria
x x x x
communities
Presumed factors influencing the the influencing factors Presumed Verrucomicrobia
and and and presumed factors sizes influencing community in the Major observations made: Major observations Planctomycetes ND Verrucomicrobia and Acidobacteria Verrucomicrobia Planctomycetes and
b Higher abundance of hitherto-uncultured groups found at bulk soil rhizosphere
a Verrucomicrobia , B ND Acidobacteria Calcareous loamy loamy Calcareous sand Loam soil A Sandy loam Sandy A Abundance of ) Stipa Stipa ); and and ) rhizosphere 1 Table Bromus Bromus Pinus contorta Pinus grass ( tectorum Hilaria jamesii Plant speciesPlant pine Lodgepole ( Soil type Approach Bunchgrasses ( hymenoides Thlaspi caerulescens Thlaspi 40 Chapter 2
De Cárcer et al., De Cárcer 2007 2007 al., et Singh Filion et al., 2004 et al., 2004 Filion Stafford et al., 2005 et al., 2005 Stafford 2006 al., et Sanguin
Acidobacteria representatives were were representatives representatives. l communities were were higher in number number in higher were , Verucomicrobia and and , Verucomicrobia had no effect in the the in effect no had c Representatives of the division of of division the of Representatives Acidobacteria of rhizosphere the in diversity and black plants than plants with of healthy spruce. Endemic communitiesEndemic in the rhizosphere soil and location per geographic differed covering vegetation type Plant roots exerted a strong selective selective a strong roots exerted Plant of diversity the on pressure Acidobacteria Planctomycetes Acidobacteria favoured in rhizosphere soil, but the ofmajor community determinant composition was soil type. Verrucomicrobia diseased of rhizosphere the in found only plants. spruce black PCB diversity diversity in the rhizosphere of Willow.
x x x x x x
,
Planctomycetes Acidobacteria Acidobacteria Verrucomicrobia and
and and
, Planctomycetes Acidobacteria Verrucomicrobia A Sand-silt soil A Not described Not described C ND ND Acid sand-stone- soil derived Sand-loam soil D Not described E ND ND cv. cv. and and ) ) Picea Salix
L.) ) Zea mays Lolium Leucospermun Leucospermun Grass ( perenne mariana Conifer ( Proteacea species ( truncatulum Lecadendron xanthoconus vimanalis x schewerinii Maize ( PR38a24) Willow (
41 Hitherto-uncultured bacteria from the rhizosphere
Kielak et al., 2008 2008 al., et Kielak 2008 al., et Hao DeAngelis et al., DeAngelis 2008 Zul et al., 2007 2007 al., et Zul l and and , Acidobacteria l population diversity and Plancomycetes Plancomycetes and member in rhizosphere in member Verrucomicrobia Acidobacteria , Verrucomicrobia Planctomycetes the respective compared soil with when bulk soil. plant diversity or content water were observed. and structure were observed in response response in observed were structure and to different vegetation coverages. or root hairs and mature root. root. and mature hairs root Geographic originGeographic important is more in the bacterial community shaping species. plant than composition differed also differed among root tip, root tip, among differed also differed Verrucomicrobia The number of of The number Rhizosphere effect increased numbers of of increased numbers effect Rhizosphere Acidobacteria Plant species strongly influenced the influenced strongly the species Plant composition; bacterial community as a second identified was season by caused No effects factor. important in No detectable changes
x x x x x
,
and and ,
Planctomycetes Acidobacteria Verrucomicrobia and Verrucomicrobia Verrucomicrobia Planctomycetes Acidobacteria
Acidobacteria F G Not described Not described A ND ND Medium-texture Medium-texture loam Endostagnic Endostagnic cambisol soil Loamy sand soil A Applied approaches for studying bacterial community composition in described studies: library clone gene rRNA – 16S A (T-RFLP) polymorphism length restriction Terminal B – (ARDRA) analysis restriction DNA ribosomal – Amplified C microarray taxonomic rRNA-based D – 16S (TGGE) electrophoresis reaction-thermal chain gel gradient E – Polymerase PCB, Polychlorinated biphenyls F – Polymerase chain reaction-denaturating gradient electrophoresis (DGGE) gel gradient reaction-denaturating chain F – Polymerase G – Quantitative PCR ) Avena Avena media media x ND, not determined not ND, a b c Taxus ) Taxus mairei Taxus and and Yew ( Yew Wild oat root ( fatua Different plant Different communities plant Different diversity treatments 42 Chapter 2
with Acidobacteria subgroup 6 in the rhizosphere, whereas in the bulk soil more clones were found to be affiliated with Acidobacteria subdivision 1. Furthermore, the diversities of the Acidobacteria and Verrucomicrobia in the rhizosphere of Lolium perenne L. were large. Such diversities even approximated those of the total Proteobacteria (Singh et al., 2007). The numerical abundance of members of the Acidobacteria, Verrucomicrobia and Planctomycetes is not only dependent on differences between bulk and rhizosphere soil, but also changes are incited upon root development. In a microcosm experiment made with wild oat roots, members of the Acidobacteria, Verrucomicrobia and Planctomycetes occurred at higher number, as demonstrated by the higher number of 16S rRNA gene copies in the rhizosphere of adult plants than in the bulk soil (DeAngelis et al., 2008). Besides, there was a greater abundance of Acidobacteria 16S rRNA genes at the surface of mature roots than at root hairs and tips; the Planctomycetes showed higher 16S rRNA gene abundance at the surface of mature roots and root tips, than at the root hairs; and the Verrucomicrobia, although having lower abundance at the root tips, had greater abundance at the surface of mature roots and root hairs than in the corresponding bulk soil (DeAngelis et al., 2008).Clearly, the high diversity of Acidobacterial and Verrucomicrobial species indicates that different bacterial populations are favoured by different selective forces exerted in the rhizosphere. At the microhabitat level, such forces can be different across space and time, thus allowing the emergence of diverse populations across each phylum. An important question here is which selective forces influence the Acidobacterial, Verrucomicrobial and Planctomycetales communities in the rhizosphere. On the basis of incidental data, this question has been difficult to answer. Thus, no consensus was found about which ecological factors would select for these organisms in the rhizosphere. Recent studies demonstrated that different factors select distinct members of these groups (Table 1). For instance, plant species (Chow et al., 2002; Gremion et al., 2003; Sanguin et al., 2006), soil type (Singh et al., 2007), field history (Kielak et al., 2008) and different abiotic factors (Stafford et al., 2005; Hao et al., 2008) have all been implicated as selective factors that differentially affect these groups (Table 1). Moreover, another key question is whether we can actually show to what extent hitherto-uncultured organisms, e.g. members of the Acidobacteria, are part of the metabolically-active community (derived from the higher abundances in RNA versus DNA extracts) in the rhizosphere and how this is temporally organized. In the metal- hyperaccumulating plant Thlaspi caurulescens, a major discrepancy in abundance between the metabolically-active and the total Acidobacteria was found (Gremion et al., 2003). In this study, it was indicated that only a subset of the Acidobacteria may be active in the rhizosphere, whereas the majority may be inactive or dormant. On the other hand, in a study using a culture-independent approach to search for 16S rRNA gene
43 Hitherto-uncultured bacteria from the rhizosphere
sequences in DNA and RNA extracts from the chestnut tree (Castanea crenata) rhizosphere (Sang-Hoon Lee, 2008), Acidobacteria were found to be not only numerically dominant but also metabolically active, implying that these Acidobacteria may play a key role this habitat. Additionally, in a metagenomic study done on the rhizosphere of Erica andevalenis adapted to acid mine drainage (AMD), it was demonstrated that in this acid environment, Acidobacteria of subdivision 1 were metabolically active (Mirete et al., 2007). The authors also found that four of 13 clones with nickel resistance genes most likely were from representatives of Acidobacteria subgroup 1 (Mirete et al., 2007). However, it is unclear whether these genes were actually expressed in the environment and whether they play key roles in the heavy metal resistance of E. andevalensis.
Methods to improve bacterial and Archaeal culturability
Recently, several successes in the culturing of hitherto-uncultured bacteria were reported (Table 2). Key factors that presumably influence bacterial colony formation on solid media were found to be the type of growth medium, the incubation conditions
(temperature, CO2 concentration and time of incubation) and several factors that are known to reduce oxidative stress (e.g. the presence of catalase or pyruvate) (Stevenson et al., 2004; Van Overbeek et al., 2004; Sangwan et al., 2005; Eichorst et al., 2007). It is a well known fact that medium composition will determine which fraction of the microbial community is recoverable from the environment. Often, the medium itself limits bacterial growth due to the presence of excessive amounts of nutrients that do not reflect environmental conditions (Janssen et al., 2002; Sait et al. 2002). This phenomenon, known as ‘substrate-accelerated death’, has first been observed when organisms growing under oligotrophic conditions are transferred to high substrate concentrations (Postgate & Hunter, 1964; Straskrabová, 1983). It is a key issue, given the roughly low substrate concentrations in the rhizosphere compartment. Therefore, to overcome this limitation, novel media are designed to match conditions in the environment in which some of the key facets (e.g. amount of nutrients, nutrient composition, presence of trace elements and pH) are adjusted for optimized bacterial growth. For instance, the use of soil-extract agar medium (Pouchon & Tardieu, 1962; Hamaki et al., 2005) or of dilute standard media, both aiming to reduce nutrient levels in order to avoid substrate-accelerated death, have allowed the isolation of bacterial groups that had been regarded as uncultured only a short while ago (Joseph et al., 2003; Davis et al., 2005). Also, the presence of fast-growing bacteria that inhibit colony formation of slower growing ones, especially on nutrient-rich media, affect the recovery of hard-to- culture organisms. Media with low nutrient levels, in combination with long incubation
44 Chapter 2
periods at relatively low temperatures, will allow colony formation by bacteria that have low growth rates. The approach has been successful in revealing a range of novel organisms in various recent studies (Janssen et al., 2002; Rappe et al., 2002; Sait et al., 2002; Davis et al., 2005; Miteva & Brenchley, 2005; Sangwan et al., 2005; Eichorst et al., 2007). Hence, this allowed the cultivation from soil of a range of novel organisms such as bacteria belonging to the following groups: Chloroflexi, Planctomycetes, Acidobacteria, Verrucomicrobia; and some Archaea representatives (Table 2). However, such approaches seldom have been used with rhizosphere samples, and only one reference was found reporting cultivation of hard-to-culture microorganisms from rhizosphere samples (Simon et al., 2005) Besides, the inadvertent presence of growth-inhibitory compounds in agar media may prevent the formation of colonies by particular bacteria. The use of gellan gum as an alternative solidifying agent was shown to allow the cultivation of many hitherto- unculturable bacteria (Tamaki et al., 2005), as agar might actually be inhibitory to some groups of microorganisms, e.g. those from freshwater. Also, incubation of samples (in this case from marine sediments) in diffusion chambers, which allows dilution of inhibitory compounds produced by bacteria during growth, permitted the growth of novel bacterial groups (Bollmann et al., 2007) (Table 2). Another reason why many bacteria cannot be cultured is the obligate growth of bacteria in consortia (Sakai et al., 2007). Thus, the growth of individual species in these consortia rely on growth of others, e.g. when bacteria feed on compounds produced by others (Simon et al., 2005) or require growth factors produced by other bacteria (Bollmann et al., 2007). Therefore, it will be relevant to screen for unknown species in clone libraries made from bacterial mixtures, e.g. made by plating of undiluted rhizosphere samples.
45 Hitherto-uncultured bacteria from the rhizosphere
et al.,
et al.,
et al.,
et al., 2007 2007 al., et et al., 2007 2007 al., et
et al., 2006 2006 al., et
et al., 2005 2005 al., et et al., 2002; Sait 2002; al., et et al., 2005 2005 al., et
et al., 2003; 2003; al., et et al., 2005 2005 al., et et al., 2002; 2002; al., et
et al., 2005; 2005; al., et
et al., 2007 2007 al., et
References References Janssen Davis 2002; al., et Sangwan 2005; Joseph Davis Hamaki Bollmann Tamaki Sakai Bollmann Cavaletti 2005; Eichorst 2005; Simon 2007 2007 Rappe Brenchley, Miteva & 2005; Rhizosphere Marine sediment Fresh watersediment Bulk soil Rice paddy field paddy Rice Fresh watersediment Bulk soil Seawater Bulk soil
Environments where where Environments novel isolates were from obtained x x x x x
x x x , , nd proposed approaches to improve culturability , , ,
, Verrucomicrobia Verrucomicrobia Planctomycetes Acidobacteria Verrucomicrobia , Proteobacteria, , Proteobacteria, , CFB group group , CFB , Gemmatimonadetes, Acidobacteria Acidobacteria
Phylogeny of novel isolates that were obtained Actinobacteria, Actinobacteria, Acidobacteria Chloroflexi, Alphaproteobacteria, Betaproteobacteria, Gammaproteobacteria, Actinobacteria, Actinobacteria, Gammaproteobacteria, Deltaproteobacteria, Deltaproteobacteria, Spirochaetes, group methanogen Cluster-I Rice Deltaproteobacteria, Deltaproteobacteria, Spirochaetes, Planctomycetes Bacteroidetes Proteobacteria, Gammaproteobacteria, SAR11 clade, SAR11 Gammaproteobacteria, Gemmatimonadetes, Chloroflexi, Planctomycetes Archaea th upon isolation from natural environments a Diffusion or dilution of of or dilution Diffusion growth-inhibiting Reduce nutrient nutrient Reduce availability in growth medium; incubation longer Apply time Development of media select a different that for of spectrum or broader microorganisms Incubation of environmental samplesin diffusion chambers; Addition of syntrophic enrichment bacteria in cultures Application of alternative solidifying agents, e.g. gellan gun instead of agar compounds, e.g. by by e.g. compounds, diffusion of use making chambers;
x x x x x x x Proposed approaches to approaches Proposed improve culturability Factors hampering bacterial Factors grow hitherto-uncultured bacteria Table 2. Factors that limit growth of Inability to at grow high nutrient concentrations and growing faster by overgrowth species for Media selectiveness of groups particular microorganisms Syntrophic growth or factors growth for requirement micro- other by produced Compounds inhibiting inhibiting Compounds bacterial growth organisms
46 Chapter 2
et al., 2004) 2004) al., et et al., 2007 2007 al., et et al., 2005 2005 al., et
et al., 2004; 2004; al., et 2004; al., et 2004 al., et
et al., 2005; 2005; al., et et al., 2005) 2005) al., et
et al., 2007 2007 al., et et al., 2005 2005 al., et
et al., 2007 et 2007 al.,
et al., 2005; 2005; al., et et al., 2002 et al., 2002 et al., 2002 et al., 2002
Stevenson Sangwan Zengler Rappe Ingham Stevenson Ferrari Stevenson Shigematsu Brenchley, Miteva & Simu 2005; Eichorst Schoenborn Sangwan Rappe processed food, raw raw food, processed vegetables Bulk soil soil Bulk soil Bulk salt- Sea food, Seawater Bulk soil Seawater Sea water Bulk soil Fresh water
x x x x x x x x x x
,
Acidobacteria , , Candidate division division , Candidate Verrucomicrobia ,
Acidobacteria Verrucomicrobia Gemmatimonadetes, Proteobacteria and and Proteobacteria Gemmatimonadetes, group CFB Acidobacteria Verrucomicrobia TM7, Acidobacteria tension 2
Increase CO Increase during incubation. incubation. during Single cell detection parallel with combined cultivation;microbial in isolates for Screening solid and liquid both media Colony identification by FISH in plates;microtitre High through put dilution dilution put through High to extinction cultivation method ; Micro-Petri dish Colony identification by hybridization on nylon membranes; Micro cultivation. colony Reduction of oxidative oxidative of Reduction stress by addition of oxidation protective agents.
x x x x x x x x x tension during incubation 2 CO in abundance Low environmental samples Lack the ability to grow under circumstances; i.e. specific growth on solid or in liquid substrate Formation of colonies that are of Formation unarmed the by undetectable eye Restriction bacterial of growth reactive of presence upon oxygen species
47 Hitherto-uncultured bacteria from the rhizosphere
Towards the as-yet-uncultured microbial populations in the rhizosphere and their basic ecology
The recovery of the broadest spectrum of species diversity from the rhizosphere will offer new insight in hitherto unculturable populations and their functions, as demonstrated with the recovery of novel nickel resistance genes from heather (Erica andevalensis) rhizosphere metagenome (Mirete et al., 2007). A major focus should be placed on those bacterial phyla that are presumably abundant in the rhizosphere and that have so far only been described via culture-independent approaches, i.e. the Acidobacteria, Verrucomicrobia and Planctomycetes. Several studies have indicated strategies to improve culturability in natural ecosystems (Table 2), but only one so far has been addressed the rhizosphere, in this case by improving culturability of Archaea species (Simon et al., 2005). Improving cultivation of Acidobacteria and Verrucomicrobia from the rhizosphere. The rhizosphere clearly is a selective habitat for different groups of microorganisms in soil and this selectivity comes about as a result of a plethora of beneficial and stress conditions imposed on its inhabitants. Analysis of the factors that shape rhizosphere conditions might help to design methods for the recovery of a broad range of novel isolates. The following factors should be considered as relevant for the design of such methods: (a) the provision of the (solid) substrate for bacterial growth; (b) the nutritional status of the rhizosphere which is often oligotrophic, although pulse-wise receiving nutrients; (c) the high partial CO2 and low partial O2 pressures (due to bacterial and root respiration); (d) the temporary and fluctuating lowering of the pH, and (e) the presence of signalling and other (eventually toxic) molecules of root and bacterial origin. Given the fact that no single medium or technique is likely to solve the culturability issue all-at-once, here a polyphasic approach is proposed, which should apply parallel cultivation systems in an attempt to yield as many hitherto-uncultured bacteria from the rhizosphere as possible. Key to such a polyphasic approach will be:
- Use of low-nutrient media in combination with long incubation periods to avoid substrate-induced death and overgrowth of fast growers; thus supporting the growth of slower ones; - Inclusion of incubation at elevated carbon dioxide tension levels; - Amendment with agents in the growth medium that protect cells from reactive oxygen species produced during their metabolism or present as a result of autoclaving;
48 Chapter 2
- Amendment of media with rhizosphere extract to mimic the rhizosphere environment, with emphasis on the presence of available nutrients and signalling molecules for growth; - Parallel use of solid and liquid media – the rhizosphere offers both a solid substrate (root surface and soil matrix) and an aqueous phase (water film surrounding soil particles and water-filled soil pores) for bacterial growth, thus the use of a combination of solid and liquid media will support outgrowth of micro-organisms with sessile and benthic lifestyles;
However, the successful isolation of hitherto-uncultured bacteria will be no a priori guarantee that key bacteria with defined roles in the rhizosphere will be found. Indeed, so far, the efforts to increase the culturability of microorganisms from natural habitats have recovered a broad range of novel bacteria (see references in Table 2). However, their involvement in key ecosystem processes is often not yet clearly understood, which is mainly due to the lack of further in-depth studies concerning their ecological roles. Hence, we propose that - following their isolation - the novel taxa are thoroughly investigated with respect to their metabolism to indicate their ecological niches and functions in the soil-plant system. In such studies, the conditions reigning in the rhizosphere should be mimicked as accurately as possible and particular attention should be paid on the occurrence of signalling between plant roots and rhizosphere micro-organisms or even between populations of rhizosphere micro-organisms. Functions can be assigned to genes or operons in newly recovered isolates by knock-out mutations and testing derived mutants in realistic plant-soil settings. Following the fate of knock out mutants upon introduction into the rhizosphere should reveal their fitness, activities and/ or interactions with other species. Here, the omics tools are a welcome addition to the toolbox that is currently available for the molecular ecologist. From a genomics-based analysis of potential function, an analysis of transcriptomics and/or proteomics under different (soil, rhizosphere) conditions can quickly indicate to what extent particular functions are operational under rhizosphere conditions.
Conclusions
Rhizosphere communities are important for key plant processes such as nutrient acquisition, protection against soil-borne diseases and plant development, as the rhizosphere is the transitional zone in between bulk soil and plant roots. Key functions of
49 Hitherto-uncultured bacteria from the rhizosphere
rhizosphere bacterial communities, for instance nitrogen fixation (Canbolat et al., 2006), phosphate solubilisation (Kohler et al., 2006) and plant health protection (Garbeva et al., 2004a) have been described, mainly for well-characterized culturable species. In this chapter, we show that among the groups of bacteria highest in density in the rhizosphere environment, the Acidobacteria, Verrucomicrobia and Planctomycetes are the ones whose functions remain to be unrevealed. So far, these three groups have not been cultivated from this environment. Therefore, most of the information that came about on these taxa in the rhizosphere was from measurements on the 16S rRNA gene level. The only exception is a metagenomics study done in the rhizosphere of plants adapted to acid mine drainage (Mirete et al., 2007). However, in this study not all the modes of action of the novel genes found could be elucidated, neither was it shown that these genes were actually active in the rhizosphere environment. One way to circumvent the hurdles of culture-independent techniques (Van Elsas et al., 2008) is the development of approaches that improve culturability, here with special focus on the rhizosphere environment. Several studies already aimed at the isolation of novel species from bulk soil, with variable successes in the recovery of Acidobacteria and Verrucomicrobia. Although these two groups have shown their recalcitrance to cultivation from the rhizosphere environment, it is likely to presume that they can be isolated using approaches directed to recover the widest possible range of bacteria. We propose the use of a polyphasic approach comprising low-nutrient solid and liquid media with the inclusion of oxidative stress protective agents and/ or rhizosphere extract, long incubation periods at elevated carbon dioxide concentrations. The exploration of hitherto-uncultured organisms - following their growth to purity - will clearly simplify an assessment of their interactions with other organisms, which cannot be achieved via culture-independent approaches. Moreover, there is a lack of studies that merge culture-dependent with advanced culture-independent studies. Such a combination of approaches will greatly increase our analytical power in order to answer one of the ultimate questions in microbial ecology – ‘Which are the ecological niches, putative roles, functions and modes of interactions of the hitherto-uncultured microbiota?’
50 Chapter 2
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Chapter 3: Cultivation of hitherto uncultured bacteria belonging to the Verrucomicrobia subdivision 1 from the potato (Solanum tuberosum L) rhizosphere*
Abstract
The role of dominant bacterial groups in the plant rhizosphere, e.g. those belonging to the phyla Acidobacteria and Verrucomicrobia, has, so far, not been elucidated and this is mainly due to the lack of culturable representatives. This study aimed to isolate hitherto-uncultured bacteria from the potato rhizosphere by a combination of cultivation approaches. An agar medium low in carbon availability (oligotrophic agar medium) and either amended with potato root exudates or catalase or left unamended was used with the aim to improve the culturability of bacteria from the potato rhizosphere. The CFU numbers based on colonies and microcolonies were compared with microscopically- determined fluorescence-stained cell numbers. Taxonomical diversity of the colonies was compared with that of library clones made from rhizosphere DNA, on the basis of 16S rRNA gene comparisons. The oligotrophic media amended or not with catalase or rhizosphere extract recovered up to 33.6% of the total bacterial numbers, at least seven times more than the recovery observed on R2A. Four hitherto-uncultured Verrucomicrobia subdivision 1 representatives were recovered on agar, but representatives of this group were not found in the clone library. The use of oligotrophic medium and its modifications enabled the growth of colony numbers, exceeding those on classical agar media. Also, it led to the isolation of hitherto-uncultured bacteria from the potato rhizosphere. Further improvement in cultivation will certainly result in the recovery of other as-yet-unexplored bacteria from the rhizosphere, making these groups accessible for further investigation, e.g. with respect to their possible interactions with plants. Introduction
The rhizosphere is a complex environment in which many different microbial species coexist. Some species interact intimately with plant roots, thereby supporting
* Authored by: Ulisses Nunes da Rocha, Fernando Dini Andreote, João Lúcio de Azevedo, Jan Dirk van Elsas & Leonard Simon van Overbeek Published in: J Soils Sediments (2010) 10:326-339 Isolation of Verrucomicrobium spp. subdivision 1 from potato
important processes for plant growth, like nutrient cycling and acquisition, disease suppression and growth stimulation (Curl & Truelove 1986). The microbial community structure in the potato rhizosphere was shown to be influenced by different biotic and abiotic factors like plant growth stage, plant genotype and local environmental conditions (Rasche et al., 2006; van Overbeek & van Elsas 2008). The high microbial diversity in the rhizosphere makes it difficult to pinpoint and eventually explore those functions that are important for in situ stimulation of plant health. Exploration of these functions is further complicated because the vast majority of bacterial species is not able to grow on standard agar media (Stevenson et al., 2004). In fact, the taxonomical status of soil isolates obtained by plating on such media will often hardly reflect the composition of the extant bacterial community because the isolation method used only accesses a small subset of this community (Eilers et al., 2000; Felske et al., 1999; Stevenson et al., 2004). Different approaches have been used to access the microbiota of natural systems, aiming to enhance the culturability of cells extracted from such habitats (Barer & Harwood 1999; Bogosian & Bourneuf 2001; Bruns et al., 2003; Guan & Kamino 2001; Ingham et al., 2007; Janssen et al., 2002; Kamagata & Tamaki 2005; Stevenson et al., 2004; Ueda et al., 2008; van Overbeek et al., 2004). Such approaches were, for instance, a decrease in nutrient concentrations by the use of dilute media and long incubation periods (Janssen et al., 2002), addition of bacterial signalling molecules (Bruns et al., 2003; Guan & Kamino 2001), replacement of agar by gellan gum as the solidifying agent (Kamagata & Tamaki 2005) and modification of the gas composition during incubation (Stevenson et al., 2004; Ueda et al., 2008). Compounds that reduce the stress imposed by the culture medium can improve culturability, as it was shown that addition of anti-oxidative stress compounds like pyruvate and catalase can lead to a higher recovery of cells from soil and water systems (Barer et al., 1999; Bogosian & Bourneuf 2001; van Overbeek et al., 2004). Also, mimicking environmental nutrient levels rather than using concentrations that are typically used in bacterial growth media can result in the growth of enhanced cell numbers (Hamaki et al., 2005; Kaeberlein et al., 2002). Thus, different hitherto-undescribed species were isolated from soil on low-nutrient media to which soil extract had been added (Hamaki et al., 2005). Clearly, to sensibly enhance bacterial culturability, it is important to know which parameters restrict or prevent growth of bacteria when these are taken out of their natural habitat. Two factors that affect the bacterial response upon transfer from the environment to culture media are exposure to oxidative stress and sudden upshifts in nutrient composition and availability (Barer & Harwood 1999). On low-nutrient media, soil isolates belonging to the unexplored Verrucomicrobia and Acidobacteria have been obtained (Janssen et al., 2002). Members of both groups also prevail in rhizospheres, although culturable representatives are hardly found in this habitat (Buée et al., 2009; Nunes da Rocha et al.,
58 Chapter 3
2009). Parameters used to establish the culturability levels of the soil microbiota are (1) the ratio of the colony numbers over the total cells and (2) the diversity of the isolates. A key consideration here is whether the enhanced CFU numbers go hand-in-hand with higher diversities of culturable species. The use of parallel and possibly complementary culturing techniques may offer an avenue to achieve a higher diversity of cultured species. The objective of this chapter was to evaluate different approaches that may enhance the culturability of bacterial cells from the potato rhizosphere, establishing their effects on colony yields, bacterial species diversity and cultivation of new species. To reach this objective, culturable cell numbers and species diversities were compared on different agar media. The diversity of the culturable bacterial community from the potato rhizosphere was also compared with that determined by a cultivation- independent technique. A selection of isolates from all agar media was identified by 16S rRNA gene sequencing.
Materials and methods
Rhizosphere sampling and processing
Seven potato plants (Solanum tuberosum L) cv. Agria, healthy in their appearances, were randomly collected from an agricultural field (250 m2 in size) near Wageningen, the Netherlands (51q59’N, 5q39’E) at flowering stage, growth stage 6 (Hack et al., 1993). Minimal distances between replicate plants was 1 m and from each individual plant, roots with soil were taken, stored on moisture paper towels in closed boxes for transport to the laboratory where samples were further processed (maximum time between sampling and processing was 1 h). The soil from the field plot was typed as loamy sand containing 2% organic matter, a water holding capacity of 25% and a pH (KCl) of 4.8. Soil adhering to the roots of each plant after shaking by hand was considered to represent the rhizosphere. This soil, approximately 1 g, was washed from the roots by shaking for 10 min in 200 ml of 0.1% sodium pyrophosphate solution. The thus obtained rhizosphere soil suspensions were blended for 1 min at 12,000 RPM, and blending was repeated 2 times - with 30 s intervals - to release as many bacterial cells as possible. Rhizosphere suspensions were used for dilution plating, cell counting and total DNA extraction followed by PCR amplification and construction of the rhizosphere DNA library.
Agar medium preparation and plating
59 Isolation of Verrucomicrobium spp. subdivision 1 from potato
R2A (Difco, France) was prepared according to the procedure described by the manufacturer. Oligotrophic agar medium was prepared in accordance with Semenov et al. (1999), with the following modifications: purified agar was used instead of Noble agar and the total carbon content was ten times lower in our agar medium. The following ingredients were dissolved in 1 L of Millipore-membrane filtered ultrapure water: MgSO4, 0.5 g; KNO3, 0.5 g; K2HPO4, 1.3 g; Ca(NO3)2, 0.06 g; glucose, 0.05 g; hydrolyzed casein 0.004 g; purified agar (Oxoid, Basingstoke, UK), 12g (pH 7.5); followed by sterilization at 121oC for 20 min. Molten oligotrophic agar was first allowed to cool down to 47ºC after which it was either mixed with catalase (100 U per ml, Sigma, The Netherlands) (CAT) or left unmixed (OLI). Potato rhizosphere extract agar was prepared by mixing molten oligotrophic agar without glucose (47oC) with potato rhizosphere extract reaching a final carbon concentration of 0.05 g per L (PEX). Potato rhizosphere extract was made from potato rhizosphere soil by incubation of 20 g of potato roots with tightly-adhering soil in 1 L of sterilized Millipore-membrane- filtered ultrapure water overnight at 4ºC. Then, soil particles were removed by centrifugation for 10 min at 200 g and the remaining supernatant was sterilized by filtration through a 0.22 Pm filter (MILLEX® GP, Millipore, Ireland) (Da Rocha et al., 2009). Sterility of thus obtained filtrate was checked by spreading 100 µl onto OLI plates and incubation in moisturized boxes at 25oC for 15 d; this resulted in the absence of any colony formation.
Bacterial cell enumerations
For total bacterial cell enumeration in the potato rhizosphere, cells in soil suspensions were stained with 5-([dichlorotriazinyl]amino) fluorescein (DTAF; Sigma-Aldrich, St Louis, MO) followed by automated counting using confocal microscopy, performed according to the procedure of Bloem et al. (1995). For CFU enumeration from the potato rhizosphere, the rhizosphere suspensions were serially tenfold diluted in sterile distilled water, after which (per dilution) 0.1 ml was plated onto R2A, OLI, CAT or PEX media. Plates kept in moisturized boxes were incubated at 25ºC and colony formation was followed in time by counting the number of colony forming units (CFU) between 3 and 15 days. Colonies were distinguished upon visibility by the unarmed eye (colonies, CFU, > 250 µm in diameter) or at 50 times magnification (microcolonies, mCFU, between 50 – 250 µm in diameter). mCFUs were enumerated by scanning entire agar plates under aseptic conditions in a laminar flow hood to circumvent contamination from the air. Total DTAF-stainable (fluorescence-stained) cell and CFU numbers were expressed per g of dry rhizosphere soil and log-transformed prior to statistical comparisons.
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Colony selection, genomic comparison by BOX-PCR fingerprinting and storage of isolates
After 15 days of incubation, a total of 180 (45 from each agar medium) CFUs were randomly picked from plates that had received the two highest dilutions of the rhizosphere soil suspensions. CFUs were streaked to purity on the same agar medium and allowed to grow out to new colonies after 5 d of incubation. mCFUs were picked from the agar surfaces by making use of sterile dissecting needles under 50 times magnification in a sterile environment. A total of 12 (4 from each agar medium: OLI, CAT and PEX) mCFUs were selected and cell material from single colonies were transferred to fresh agar media of their respective composition to allow formation of new colonies after 150 d of incubation. A total of 180 CFUs plus one resulting from a mCFU were compared by BOX- PCR (Rademakers et al., 1998) fingerprinting. Volumes of 1 Pl cell lysate, prepared from material from single colonies suspended in 100 Pl of sterile demineralised water and heated to 100oC for 5 min were used as DNA templates for PCR reactions. After PCR amplificationsYROXPHVRIȝORf each mixture were loaded on 1.2 % agarose gels, 40 cm in sizes, and run for 16h at 4oC and 1.2V/cm. Each individual gel included WKUHHODQHVHDFKORDGHGZLWKȝORINEODGGHU ,QYLWURJHQ&DUOVEDG &D IRUODWHU normalization. After running, gels stained with ethidium bromide were photographed under UV and digitized fingerprints were compared using Gelcompare II software (Applied Maths, Belgium). BOX-PCR fingerprints were compared at 100, 95 and 90 % similarity levels. For storage of isolates, cells from single colonies were suspended in oligotrophic broth (OLI with omission of agar) amended with glycerol (final concentration 20 %) and placed at -80°C.
Construction of a 16S rRNA gene clone library made from rhizosphere DNA
A 16S rRNA gene clone library was made from potato rhizosphere soil. Therefore, DNA was extracted from a pooled suspension from all seven rhizospheres using the UltraClean Soil DNA Isolation Kit (MO BIO laboratories, Inc., Carlsbad, CA). Then, one Pl (20 ng) of purified DNA extract was PCR amplified using primers 27F and 1492R. The PCR product with the expected fragment size was purified from non-incorporated dNTPs and primers using the QIAquick PCR purification kit (Qiagen, Hilden, Germany). Purified PCR product was cloned into the pCR2.1-TOPO vector from the TOPO-TA PCR cloning kit (Invitrogen) and introduced into Escherichia coli Top10 cells (Invitrogen) by transformation according to the protocol provided by the manufacturer. White colonies, indicating insertional inactivation of the lacZ gene, were
61 Isolation of Verrucomicrobium spp. subdivision 1 from potato
PCR-amplified with primers annealing with the M13F and M13R sites in the pCR2.1- TOPO vector using the procedure provided by the manufacturer. In total, 93 clones with the expected fragment size were selected for later sequencing.
Identification of isolates and clone inserts from the rhizosphere DNA library by 16S rRNA gene sequencing
For PCR amplification of the 16S rRNA gene, cell lysates from all 180 isolates recovered as CFUs and 12 as mCFUs (192 in total) were used as DNA templates. For that, cell material was used from purified colonies with the exception of 11 mCFUs that were not able to grow upon transfer to fresh media and from these, cell material taken from the original colonies were added to the PCR mix. Per cell lysate, 1 PL was added to 49 PL PCR reaction mixtures consisting of: Tris-HCl (pH 8.3), 10mM; KCl, 10mM;
MgCl2, 2.5mM; each deoxyribonucleoside triphosphate, 200mM; 400mM of each primer, 27F (Lane et al., 1985) and 1492R (Rochelle et al., 1992), and 5U of SuperTaq DNA Polymerase (HT Biotechnology, UK). PCR amplifications were run in a PTC-200 thermal cycler (MJ Research, The Netherlands) programmed at: one cycle of 95ºC, 5 min; 30 cycles at 95ºC for 60 s, 62ºC for 60 s, 72ºC for 90 s; and one cycle 72ºC for 10 min. The PCR products obtained were used for later sequencing. PCR products of all 192 isolates from the four agar media and the 93 clones from the rhizosphere DNA library were sequenced. For that purpose, purified PCR products were added to reaction mixtures containing five Pl of sequencing reaction mixture, one Pl of DETT Dye (Dyenamic ET Terminator Cycle Sequencing Kit, Healthcare, GE), three Pl of dilution buffer and one Pl (0.5 PM) of primer 1492R. Linear amplifications were performed for 25 cycles at 94ºC, 20s; 50 ºC, 15s; 60 ºC, 60 s. The amplified products, approximating 600 bp in size, were sequenced in an ABI prism automatic sequencer by making use of the services of Greenomics (Plant Research International, Wageningen, The Netherlands). Sequence data from all isolates and clones were first checked for chimeras using the Check Chimera tool (http://www.ncbi.nlm.nih.gov) and nonchimeric sequences were assessed for similarity, using the Sim_Identity index (SI) with sequences of type strains in the Ribosomal Database Project (RDP) (Cole et al., 2009) using the bioinformatic analysis tools of the Sapelo Island Microbial Observatory (last accession 30 September 2009; http://simo.marsci.uga.edu/). The identities of colonies and clones, expressed in operational taxonomic units (OTUs), were provided according to their nearest matches at levels of 99 % (strain), 95 % (genus) and 90 % (class) with sequences present in the database. OTUs of 12 isolates showed < 95% similarity with sequences of type strains in the RDP and these isolates were considered as non-readily identifiable. Sequencing of a larger part of the 16S rRNA genes of these isolates was performed for better
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comparisons with SILVA release 94 (Pruesse et al., 2007) database entries. The following primers were used for further sequencing of these 16S rRNA genes: P027F (Lane et al., 1985), R530 (Muyzer et al., 1993) and 968R (Heuer et al., 1997). Assembled contiguous fragments were compared for nearest matches with SILVA database entries using ARB software (Ludwig et al., 2004).
Data analysis
For statistical analyses, comparisons were made between total fluorescence-stained cell and total CFU numbers from all four agar media, between total CFU numbers from the different agar media after 15 days of incubation. Differences were calculated by Tukey’s test with seven replicates per sample and were considered to be significant at a level of P Rarefaction analysis was performed on isolates of the four media and clones of the rhizosphere DNA library to determine OTU richness at 99 and 95% similarity levels. The rarefaction modus of the DOTUR software (http://www.plantpath.wisc.edu/fac/joh/dotur.html) was used (Schloss & Handelsman 2005). Rarefaction diagrams were made by plotting the number of OTUs as a function of the number of individual colonies or clones sampled from, respectively, the different media or the rhizosphere DNA clone library. Correspondence between pools of (V6-V8) rRNA gene sequences from library clones and isolates was made by pair wise comparisons using -LIBSHUFF (Schloss et al., 2004). Therefore, sequences from different pools were aligned and a matrix of similarity was generated by the application of DNADist which was used as input for analysis by -LIBSHUFF after which corrected P values were calculated. Phylogenetic distances were calculated using the ARB software package between 16S rRNA gene sequences of over 1000 bp in size from four selected isolates and ninety other Verrucomicrobia sequences and ten representatives from the phylum of Chlamydiae as an out group from the Silva database. Aligned sequences were manually edited taking 16S rRNA sequence secondary structures into consideration. Reconstruction of phylogenetic relationships was based on neighbour joining (Ludwig et al., 1998). The branches were tested with bootstrap analysis (1000 iterations).
Nucleotide sequence accession numbers
The DNA sequences of the partial 16S rRNA genes of 12 isolates (between 804 and 1492 bp in sizes) were deposited in the EMBL Nucleotide Sequence Database (Cochrane et al., 2009) under accession numbers FN394501 to FN394512.
63 Isolation of Verrucomicrobium spp. subdivision 1 from potato
Results
Bacterial numbers in the potato rhizosphere
The log-transformed fluorescence-stained cell numbers from seven potato rhizospheres, expressed per g of dry rhizosphere soil were between 9.19 and 9.58 (Table 1). CFUs from the same samples appeared on the R2A, OLI, CAT and PEX media within seven days of incubation (Fig. 1). This stood in contrast to the microcolonies (mCFU), which in general appeared later, i.e. between seven and 15 days of incubation (Fig. 1). After 15 days of incubation, the average log CFU per g of dry soil was 8.09 on R2A, 8.27 on OLI, 8.55 on CAT and 8.36 on PEX (Table 1). The log mCFU per g of dry soil were 8.21 on OLI, 8.39 on CAT and 8.45 on PEX. No mCFU were found on R2A. Log total CFU (CFU + mCFU) per g of dry soil after 15 days of incubation were 8.09 on R2A, 8.54 on OLI, 8.78 on CAT and 8.71 on PEX. These numbers were roughly between 0.47 and 1.58 log units lower than the corresponding total cell counts as determined by fluorescence-stained cell numbers. Expressed as percentages of the fluorescence-stained cell numbers, the total CFUs on day 15 were 2.6 - 6.6 % on R2A, 4.5 - 33.9 % on OLI, 14.4 - 33.6 % on CAT and 12.3 - 28.1% on PEX. The sizes of the culturable fractions thus differed per medium and incubation time and occasionally comprised (highest value found on OLI medium) almost one third of the total fluorescence-stained cell fraction. Considering variability over the different plants, the isolate fractions across the different agar media were lowest in plant 7 and highest in plant 6. Hence, the sizes of these fractions in the rhizosphere varied among the replicate plants. The medium used for the recovery of bacterial cells from the potato rhizosphere was determinative for the level of culturability. The highest isolate numbers were found on CAT, PEX and OLI media, whereas such numbers were significantly lower on R2A (Table 1).
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800 800 R2A OLI ) ) 6 6 600 600
400 400
200 200 CFU/g soil (x10 CFU/g soil (x10 0 0 3 d 7 d 9 d 12 d 15 d 3 d 7 d 9 d 12 d 15 d 800 800 CAT PEX ) ) 6 6 600 600
400 400
200 200 CFU/g soil (x10 CFU/g soil (x10 0 0 3 d 7 d 9 d 12 d 15 d 3 d 7 d 9 d 12 d 15 d
Colonies Microcolonies
Figure 1 Colony appearance over time on different agar media (i.e. R2A, OLI, CAT and PEX). Bars represent standard deviations. Colonies; colonies with > 250 µm in diameter, Microcolonies; colonies between 50 and 250 µm in diameter.
Successive growth of isolates on agar media
A total of 192 isolates obtained from both CFUs and mCFUs were successively streaked to purity and transferred to fresh media in order to determine their capacity to show continued growth under medium conditions. Isolation of cells from mCFU is a laborious process and hence we included a maximum of 12 such isolates. Thus, 180 isolates (45 from each agar medium) were obtained from CFUs and 12 (four from each agar medium: OLI, CAT and PEX) from mCFUs. All isolates recovered as CFUs grew into CFUs upon transfer to the same agar medium, whereas 11 of the isolates recovered as mCFUs did not form mCFUs- or CFUs, not even after 150 d of incubation. The one mCFU (from CAT, denoted as CR28) that did grow formed a CFU after transfer to the same agar medium within 5 days of incubation. Hence, whereas all CFUs maintained culturability, the majority of mCFU did not.
65 Isolation of Verrucomicrobium spp. subdivision 1 from potato 1) (20.6) B (11.3) 8.71 (11.3)
PEX PEX l cell: total DTAF-stained umbers of CFU and mCFUs and mCFUs CFU of umbers a, b (24.7) (24.7) 8.36(9.28) 8.45 B agar media after 15 d of incubation. FU CFU Total CFU mCFU CFU Total 3) 3) (11.4) 8.33 8) (26.6) 8.70 (16.2) 8.42 (31.0) 8.70 (9.9) 8.27 (11.8) 1) 8.34 (9.3) 8.18 (21.7) 8.61 (9.3) 8.46 (8.6) 8.15 (22.4) 8.84 (17.9) 8.46 (11.6) 8.56 (14.0) 8.64 (25.6) 8.90 6) 6) (11.6) 8.38 (28.1) 8.77 (9.8) 8.31 (13.9) 8.46 (23.7) 8.69 .9) .9) (5.5) 7.93 (14.4) 8.35 (5.4) 7.92 (6.9) 8.03 (12.3) 8.28 12.0) 12.0) (5.0) 8.09 (17.0) 8.62 (6.2) 8.18 (8.4) 8.31 (14.6) 8.55 19.8) 19.8) (13.8) 8.72 (33.6) 9.11 (12.8) 8.69 (15.3) 8.77 (28. 9.03 (14.3) 8.39(10.4) 8.78 8.39(10.4) (14.3)
(12.3) 8.55 (12.3) B m potato rhizospheres recovered on different (5.66) 8.54 (5.66)
Log cells / CFU per gram of dry rhizosphere soil (CFU fraction of total cell number, %) number, cell total of fraction soil (CFU rhizosphere dry of gram / CFU per cells Log OLI CAT CAT OLI c (4.8) (4.8) 8.27(6.6) 8.21 A CFU CFU mCFU Total CFU CFU R2A mC CFU CFU CFU mCFU Total 8.09 Total bacterial cell and CFU numbers Total bacterial cell and CFU fro C Total cells cells Total < B < C A different, where Significantly tota extract; rhizosphere or potato catalase with or amended non-amended respectively agar medium, oligotrophic PEX: CAT, OLI, R2A on found was No mCFU 1 Table a b c cells; CFU, colonies with > 250 µm in diameter; mCFU, colonies between 50 and 250 µm in diameter; Total CFU, summation of the n of the summation CFU, Total diameter; in 250 µm 50 and between colonies mCFU, diameter; in > 250 µm with colonies CFU, cells; 1 2 9.27 9.21 4 7.87 (4.0) 9.39 7.76 (3.5) 7.94 (4.7) 7 7.79 (3.8) (5.9) 8.16 7.83 (3.7) 9.19 7.78 (3.7) 8.19 (8.4) (6.1) 8.17 8.08 (7.5) 1(2.6) 7.6 (4.4) 8.03 8.45 (15. (10.5) 8.41 8.38 (14. (2.5) 7.58 ( 8.47 (2.1) 7.50 (4.5) 7.84 (8 8.14 3 9.32 5 6 (6.3) 8.12 9.49 9.58 (5.8) 8.09 8.19 (5.0) (6.6) 8.40 (5.5) 8.06 8.21 (5.3) (11.4) 8.38 (18.2) 8.84 8.14 (4.5) (16. 8.54 (15.8) 8.78 8.48 (9.8) (33.9) 9.11 ( 8.61 8.88 (13. plant plant Mean 9.35 number number
Replicate 66 Chapter 3
Comparison of BOX-PCR fingerprints from culturable isolates
Cell lysates prepared from 181 isolates (i.e. the ones that grew upon transfer) were analysed by BOX-PCR to cross-compare their genomes (data not shown). Similarity levels of 100, 95 and 90% were used in the groupings. At 100 and 95 % of similarity, all isolates differed from each other, indicating that a huge diversity of culturable forms was indeed sampled. At 90% similarity, three clusters, consisting of respectively 2, 5 and 6 isolates, were distinguished, whereas all other 168 fingerprints formed singletons that differed from each other. No commonalities among isolates from the same agar medium were found at this level. This indicates that all isolates differed substantially from each other, irrespective of their initial growth on different agar media.
Comparison of partial 16S rRNA gene sequences of isolates with database entries
The sequences of the variable regions V6 - V8 of the 16S rRNA genes of 180 isolates from all four agar media were affiliated with, in total, five different phyla. Representatives of the Proteobacteria, Bacteroidetes, Actinobacteria and Firmicutes were obtained from all four media, but those of the Verrucomicrobia only from CAT and PEX (Table 2). Differences in the dominant groups on the four media were found at the class level. On R2A, the group of Į-Proteobacteria was the largest, on OLI the Flavobacteria and Į-Proteobacteria, on CAT the Flavobacteria and on PEX the Į- Proteobacteria and Actinobacteria. The Į-Proteobacteria was the most dominant class among isolates from R2A and PEX, whereas Flavobacteria were dominant among the ones from OLI and CAT. In total, 169 (of 180) sequences of isolates recovered as CFUs from all four agar media matched database entries of type strains at 95% or higher levels, as follows: 44 from R2A, 41 from OLI, 41 from CAT and 42 from PEX. Eleven isolates showed similarities at levels < 95% with type strains of the RDP. These isolates were regarded as non-readily-identifiable and so larger stretches of their 16S rRNA genes were sequenced for more accurate comparisons with database entries (Table 2).
67 Isolation of Verrucomicrobium spp. subdivision 1 from potato
Table 2 Taxonomical distribution of library clones and isolates from the potato rhizosphere. Phylum Number of library clones/ isolates a Class/ group/ Clone library R2A OLI CAT LEX subdivision 93 45 45 / 4 45 / 4 45 / 4 Acidobacteria group 3 1 - - - - group 5 2 - - - - group 6 14 - - - - group 7 5 - - - - group 10 1 - - - - Actinobacteria 14 6 5 / 2 7 / 1 12 Bacteroidetes Flavobacteria - 5 11 / 1 15 / 2 9 / 2 Sphingobacteria 8 3 4 3 1 Firmicutes Clostridia 2 - - - - Bacilli 1 5 4 3 1 Proteobacteria Į-Proteobacteria 8 14 10 / 1 7 14 / 2 ȕ-Proteobacteria 13 5 8 4 4 į-Proteobacteria 3 - - - - Ȗ-Proteobacteria 4 7 3 5 2 Planctomycetes 2 - - - - Chloroflex 1 - - - - Gemmatimonadetes 8 - - - - OP10 1 - - - - Verrucomicrobia Subdivision 1 - - - 1 / 1 2 Subdivision 3 2 - - - - Unclassified 3 - - - - a OLI, CAT, PEX: oligotrophic agar medium, respectively non-amended or amended with catalase or potato rhizosphere extract; numbers in bold represent mCFUs
Partial 16S rRNA gene sequences of the 12 isolates recovered as mCFUs were affiliated with four phyla, i.e. the Bacterioidetes (class Flavobacteria), Actinobacteria (class Actinobacteria), Proteobacteria (class Į-Proteobacteria) and Verrucomicrobia (subdivision 1) (Table 3). The 16S rRNA gene sequence of the isolate that showed stable growth when streaked on the same medium, CR28, closely matched those of the subdivision 1 of Verrucomicrobia. The aforementioned four taxonomic groups were not all recovered from different agar media, indicating that unique selection of particular bacterial groups by the different agar media had not occurred.
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EU135492 (97.9) (97.9) EU135492 FJ516766 (97.9) (97.9) FJ516766 EU979055 (98.8) (98.8) EU979055 EU979055 (98.2) (98.2) EU979055 bacterium bacterium
bacterium bacterium
AF467106 (93.3) (93.3) AF467106 AF130951 (90.0) (90.0) AF130951 sp. FJ380996 (98.0) (98.0) FJ380996 sp. sp. Enf62 DQ339596 (94.0) (94.0) DQ339596 Enf62 sp. sp. As1-3 AY367025 (97.0) (97.0) AY367025 As1-3 sp. Verrucomicrobiales Verrucomicrobiales Verrucomicrobia Verrucomicrobia sp. EF488749 (92.6) (92.6) EF488749 sp. unidentified bacterium EF154188 (86.1) (86.1) EF154188 bacterium unidentified unidentified bacterium EF154188 (95.2) (95.2) EF154188 bacterium unidentified Arthrobacter aurescens aurescens Arthrobacter Sphingomonas Similarity with type or non-type strain from the SILVA SILVA the from strain non-type or type with Similarity database Bradyrhizobium Lysobacter Pantoea agglomerans Pantoea Flavobacterium
AJ512504 (93.2) (93.2) AJ512504 D32229 (93.2) D32229 X94098 (93.3) (93.3) X94098 AB019582 (92.1) AB019582 AF130953 (90.0) AF130953 DQ112353 (85.0) DQ112353 DQ302104 (89.6) uncultured DQ112353 (92.3) DQ112353 DQ112353 (94.2) DQ112353 DQ302104 (88.4) uncultured DQ302104 (88.6) uncultured DQ302104 (88.9) uncultured ates on different grown agar media. Rubritalea marina Rubritalea Lysobacter antibioticus antibioticus Lysobacter roseus Pedobacter marina Rubritalea
Rubritalea marina Rubritalea
Pedobacter roseus Pedobacter
Methylobacterium rhodinum rhodinum Methylobacterium Similarity with type strain strain type with Similarity Nearest match (% identity) Nearest match (% identity) Nearest match -Proteobacteria Sphingobium xenophagum xenophagum Sphingobium -Proteobacteria -Proteobacteria Pantoea agglomerans agglomerans Pantoea -Proteobacteria Sphingobacteria Pedobacter roseus roseus Pedobacter Sphingobacteria Verrucomicrobia Rubritalea marina marina Rubritalea Verrucomicrobia Ȗ Actinobacteria Arthrobacter nitroguajacolicus nitroguajacolicus Arthrobacter Actinobacteria Į bp Class Class bp a Identity of non-readily-identifiable isol Identity Agar Agar media PEX 1098 1098 PEX
extract. rhizosphere or potato catalase with or amended non-amended respectively agar medium, oligotrophic PEX: CAT, OLI, 3 Table a ZNBB5 C48 CAT 720 720 CAT C48 962 O53 OLI 1085 CAT C20 804 O33 OLI Z35 PEX 1100 1100 PEX Z35 CAT CR28 1114 Z70 PEX 940 940 PEX Z70 960 O77 OLI CH7 CAT 1085 1085 CAT CH7 Isolate Isolate OAG4 OLI 847 952 R66 R2A
69 Isolation of Verrucomicrobium spp. subdivision 1 from potato
Only one isolate recovered as a mCFU closely matched the 16S rRNA gene sequence of a type strain (Flavobacterium hydatis) in the RDP database at a similarity level > 99% (Table 4). At similarity levels between 99 and 95%, the 16S rRNA genes of in total four isolates were affiliated with those of type strains and eight with those of non-type strains. At similarity levels < 95%, another four isolates (amongst which CR28) were affiliated with non-type strains, indicating that these were only distantly related to representative species present in the RDP.
Potato rhizosphere DNA library analysis and comparison between cultured and uncultured bacteria
A total of 93 PCR-amplified 16S rRNA gene sequences from uncultured bacteria was obtained from the potato rhizosphere. DNA sequence analysis of all clones revealed that there were no distinguishable groups at a 97% similarity level (meaning all sequences were singletons), whereas at a 95% similarity level only two clusters, both consisting of two sequences, were distinguished. Hence, most of the clones in the rhizosphere 16S rRNA gene library were indeed singletons, indicating a high diversity within the bacterial community that was sampled. In total, only ten of the 93 sequences matched type strains in the RDP database at similarity levels > 95%, as follows: Streptomyces ciscaucasicus (99.2 %), Arthrobacter ramosus (99.1 %), Variovorax paradoxus (2 isolates: 98.6 and 98.2 %), Aquabacterium parvum (97.6 %), Aquabacterium commune (97.1 %), Agrococcus jenensis (96.5 %), Terrimonas ferruginae (95.6 %), Paenibacillus alkaliterrae (95.2 %) and Sphingosinicella microcystinivoran (95.0 %). The other 83 sequences showed similarity values <95% to RDP database entries, revealing best matches with the Acidobacteria, Proteobacteria, Actinobacteria, Bacterioidetes, Gemmatimonadetes, Firmicutes, Verrucomicrobia, Planktomycetes, Chloroflexi, division OP10 and unclassified bacteria (Table 2). Of the two groups that were most abundant in the clone library, i.e. the Acidobacteria and Proteobacteria, nearest matches with particular groups or classes were found: for the Acidobacteria groups 3, 5, 6 and 7 and for the Proteobacteria, Į-Proteobacteria, ȕ-Proteobacteria, į-Proteobacteria and Ȗ- Proteobacteria.
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IFO 15499-T (93.2) (93.2) 15499-T IFO DQ513326 (97.2) (97.2) DQ513326 sp. F4b AB220167 (93.2) (93.2) AB220167 F4b sp. sp. AF514796 (90.4) (90.4) AF514796 sp. FUs, but lost their capacity to upon grow sp. WB 2.1-78 AM167559 (96.6) (96.6) AM167559 2.1-78 WB sp. sp. 35/19 AY571807 (96.8) (96.8) AY571807 35/19 sp. (94.0) AY571807 35/19 sp. extreme arid zone bacterium HX-IIF54 AY378195 (99.7) (99.7) AY378195 HX-IIF54 bacterium zone arid extreme Bradyrhizobium Nocardioides Nocardioides luteus Micrococcus asaccharolytica Sphingomonas Bradyrhizobium Similarity with type or non-type strain from the SILVA SILVA the from strain non-type or type with Similarity database Flavobacterium
ended or amended with catalase or potato rhizosphere extract extract rhizosphere or potato catalase with or amended ended D12671 (93.9) (93.9) D12671 (96.2) AY994036 bacterium uncultured IFO 15499-T (93.2) (93.2) 15499-T IFO AY423718 (96.0) (96.0) AY423718 (93.2) AY423718 AJ557888 (94.4) (94.4) AJ557888 (98.0) AY994036 bacterium uncultured (93.7) AJ557888 (97.5) AY994036 bacterium uncultured isolates. Isolates wereisolates. recoveredIsolates as mC AB075230 (95.0) AB075230 U35000 (89.8) (89.8) U35000 M58764 (99.6) M58764 U35000 (92.9) AJ536198 (96.4) (96.4) AJ536198 Nearest match (% identity) Nearest match ganghwensis Nocardioides ganghwensis Nocardioides elkanii Bradyrhizobium limicola Flavobacterium luteus Micrococcus micromati Flavobacterium hydatis Flavobacterium (% identity) Nearest match elkani Bradyrhizobium asaccharolytica Sphingomonas saccharophilum Flavobacterium micromati Flavobacterium Similarity with type strain strain type with Similarity
bp bp 387 411 382 870 405 843 313 421 420 737 788 a Agar Agar media Identity of unculturable potato rhizosphereIdentity
ZPM37 ZPM3 CPM25 CAT ZPM39 PEX CPM27 Isolate Isolate OLIOPM24 OPM40 CPM2 OPM36 OPM36 OPM1 ZPM4 non-am respectively agar medium, oligotrophic PEX: CAT, OLI, successive transfers. a 4 Table
71 Isolation of Verrucomicrobium spp. subdivision 1 from potato
A comparison between the clone library sequences and those from the 192 isolates by -LIBSHUFF analyses revealed that the two libraries differed more than 5% from each other (P < 0.0001), indicating that the correlation between the two was low. Thus, the sequences present in the directly-obtained pool from the potato rhizosphere differed substantially from the ones present in the isolates. Rarefaction analysis performed at similarity levels of 99 and 95% on the basis of the clone library sequences (Fig. 2) showed an almost linear relationship between clone number and type. Hence, an increased sampling from the rhizosphere DNA library would have yielded additional novelty at both similarity levels, and the rhizosphere DNA was not fully explored for diversity. Similarly, rarefaction curves (99% similarity level) based on the 16S rRNA gene sequences of isolates obtained from the OLI and PEX media showed almost linear relationships, whereas those from R2A and CAT tended to level off. This observation indicated that progressively lower numbers of new species are expected to be recovered from the latter media upon further sampling. At 95% similarity levels, the rarefaction curves of 16S rRNA gene sequences of isolates from all four agar media tended to level off.
Library R2A OLI CAT PEX 90 90
80 (A)80 (B)
70 70
60 60
50 50
40 40
30 30
20 20
10 10 Number of bacterial groups 0 0 0 1020304050607080900 102030405060708090 Number of analyzed sequences
Figure 2 Rarefaction curves, based on partial 16S rRNA gene sequences from isolates grown on different agar media and clones from the rhizosphere DNA library. Similarities were calculated at two different levels: 99% (A) and 95% (B).
The sequences obtained in the clone library were aligned with those of the 192 isolates to examine the putative matches between the two sequence pools. At six occasions, matches between the sequences of clones and isolates were found (similarity levels between 95 and 98%). Three library clones showed matches with isolates from OLI medium. Specifically, these clones matched the 16S rRNA gene sequences of Arthrobacter ramosus, Variovorax paradoxus and Ramlibacter henchirensis. Two
72 Chapter 3
clones showed matches with isolates from R2A, affiliating with Stenotrophomonas acidaminiphila and Sphingobium yanoikuyae, whereas one matched an isolate from PEX medium with closest affiliation to Agrococcus jenensis. No sequences that were identical between the clone library and any of the isolates from the four media were found. This emphasizes the difference between the cultured and uncultured fractions obtained from the potato rhizosphere.
Identity of culturable bacteria with less than 95% similarity to RDP database entries
The sequences of, in total, 12 isolates (11 recovered as CFUs and one as a mCFU) showed <95% similarity with sequences of type strains of the RDP database. These isolates should hence be considered as novel taxa. The isolates had been obtained from R2A (1), OLI (4), CAT (4) and PEX (3). Larger (804-1114 bp) 16S rRNA gene sequences of these isolates were obtained and analysed using sequences from the RDP, thus allowing more accurate identifications. Based on the larger fragment sizes, the 12 isolates were different from any validated database entry. However, rather distant nearest matches were found with (type strains) Rubritalea marina (4 isolates; nearest matches between 88.4 and 89.6%), Pedobacter roseus (3; 85.1 and 94.2%), Arthrobacter nitroguajacolicus (1; 93.1%), Lysobacter antibioticus (1; 92.1), Pantoea agglomerans (1; 90.0%), Sphingobium xenophagum (1; 93.3%) and Methylobacterium rhodinum (1; 93.2%) (Table 3). All matches to type strains were clearly below 95%. However, some sequences matched those of non-type strains from the Silva database. Remarkable were the matches of the four isolates CR28, C20, Z35 and ZNBB5 (which matched that of the R. marina type strain at 88.9 % or below) with unidentified/uncultured bacteria at 96.5 % similarity or higher. Furthermore, the four isolates clustered with four uncultured representatives of subdivision 1 of the Verrucomicrobia, as shown in a dendrogram constructed with sequences of representatives of the seven subdivisions of Verrucomicrobia (Fig. 3). This indicates that the isolates are affiliated with members of subdivision 1 of the Verrucomicrobia. Given the fact that the isolates originated from CAT and PEX, these two media may allow the enhanced isolation of representatives of subdivision 1 of Verrucomicrobia. On the other hand, no 16S rRNA gene sequences were found in the clone library that matched representatives of subdivision 1 of Verrucomicrobia (Silva database). However, two clone library sequences showed nearest matches with representatives of subdivision 3 of the Verrucomicrobia. One matched an uncultured soil bacterium affiliated with unclassified Verrucomicrobia (accession number AY493907) and the other one an uncultured Verrucomicrobium sp. (AY921923). The similarity levels were, respectively, 98.7 and 99.0%. Representatives of subdivision 3 of the Verrucomicrobia
73 Isolation of Verrucomicrobium spp. subdivision 1 from potato
may be more dominant in the potato rhizosphere than those of subdivision 1, whereas only the latter could be recovered as isolates on CAT and PEX.
6 Chlamydiae
Verrucomicrobiaceae bacterium CR28 (FN394505) uncultured Verrucomicrobia bacterium (EU979055) uncultured bacterium (EU135492) Verrucomicrobiaceae bacterium Z35 (FN394511) 98 Verrucomicrobiaceae bacterium ZNBB5 (FN394508) 95 uncultured Verrucomicrobia bacterium (EF188433) uncultured Verrucomicrobiales bacterium (FJ516766) Subdivision 1 94 Verrucomicrobiaceae bacterium C20 (FN394506) Luteolibacter pohnpeiensis (AB331895) 99 97 Luteolibacter algae (AB331893)
86 5 Rubriaitela
6 Prosthecobacter 91 Verrucomicrobium spinosum (X90515)
6 Xiphinematobacter Subdivision 2 97 13 99 Unclassified Subdivision 2
5 Subdivision 3 72 97 6 Subdivision 6
11 Subdivision 4 85
5 Subdivision 5 75
4 Subdivision 7
0.10 Figure 3 Phylogenetic relationship between four non-readily-identifiable isolates, R28, C20, Z35, ZNBB5 and representatives from the phylum Verrucomicrobia. Distances between partial 16S rRNA gene sequences over 1100 bp in length were calculated using ARB software. A cluster made of rRNA gene sequences of 10 different species from the phylum Chlamydiae was used as an out group. Bar at the bottom indicate 10% divergence among sequences and boot strap values (%) are presented near each junction in the tree.
74 Chapter 3
Discussion
Improvement of culturability
A comparative study on the recovery of bacteria from the potato rhizosphere on different agar media was conducted. The total CFU numbers that were found occasionally contributed to substantial fractions (up to 33.6%) of the total fluorescence- stained cell counts, depending on the type of medium used, incubation time and the presence of mCFU and plant replicate. Our data thus show that culturability from the rhizosphere can be substantially improved by using oligotrophic agar media with or without modifications in comparison with R2A, representing a standard laboratory medium. However, culturability may be overestimated when certain bacteria have smaller than expected cell volumes (Iizuka et al., 1998), thus remaining undetectable by DTAF-assisted confocal microscopy, which has a cut-off cell volume of approximately 0.3 - 0.ȝP3 (Bloem et al., 1995). The agar media in this study were employed with the rationale to enhance the bacterial culturability by reducing cellular stresses upon exposure to oxygen (catalase) or high-nutrient conditions, the latter by mimicking the (low) nutritional status of the rhizosphere (rhizosphere extract). The presence of catalase reduces oxidative stress at the agar surface resulting in better growth and higher CFU yields (Barer & Harwood 1999; Bogosian and Bourneurf 2001; Wagner and Horn 2006). We indeed found higher CFU numbers on catalase-containing agar medium, however at the expense of a lowered diversity, both compared to those on the corresponding unamended agar. The lowered diversity may have been caused by a faster outgrowth of colonies on CAT, which was apparent from the larger colony sizes, than on OLI medium (not shown). Faster-growing colonies may suppress the growth of slower ones on the same plate (Kamagata & Tamaki 2005), which is consistent with our observations. Mimicking the nutritional status of the rhizosphere was indeed successful concerning the recovery of novel bacterial types, as previously also found for bulk soil (Hamaki et al., 2005). Rhizosphere soil is distinguished from bulk soil as an environment differing in species composition (Van Overbeek & Van Elsas 2008), which incites a higher microbial activity as a result of the availability of plant root exudates (van Elsas and van Overbeek 2008; Wagner and Horn 2006). By replacing glucose as the main carbon source by rhizosphere extract (which in reality is a combination of root exudate compounds and soil-extractable compounds), the growth of rhizosphere bacteria was expected to be favoured. Higher total CFU numbers were indeed found on PEX than on OLI medium. Moreover, the slight difference in composition of bacterial groups between the two media reflected the variation in the preference for carbon sources among bacteria present in the potato rhizosphere.
75 Isolation of Verrucomicrobium spp. subdivision 1 from potato
The presence of the mCFUs on the media made up large part of the observed higher CFU numbers. Concerning the physiological status of the bacterial cells that grew in these mCFUs, it becomes apparent that the majority only divided for just the number of generations it took to form the mCFU. Possibly, these cells suffered from substrate-accelerated death in the trajectory from rhizosphere soil via one medium to the next medium. In previous work, microcolonies have been obtained from water and soil (Kaeberlein et al., 2002; Zengler et al., 2002; Ferrari et al., 2005) and the bacteria forming these colonies represented, for the greatest part, as-yet-uncultured groups. For instance, a representative of candidate division TM7 was found in soil (Ferrari et al., 2005). mCFUs may represent an untapped resource for many novel species and this stresses their importance as sources of unexplored bacteria from soils. However, the lack of stable growth upon transfer awaits a future solution. The large variation in the recovered culturable fractions per plant indicates that the circumstances for efficient growth of different bacterial groups vary per plant, even under almost identical conditions regarding soil type, plant genotype and growth conditions. Stochasticity in bacterial colonization of the rhizosphere is one factor that may explain this large variation in uncultured fractions (66-96%) in the rhizosphere. Cells of these uncultured bacteria may have been either viable but unable to grow outside the rhizosphere, or moribund and thereby unable to divide. Using PCR amplification on the basis of rhizosphere DNA extracts, such fractions may have been mainly accessed if they dominated the community. Overall, the cultured and total bacterial fractions from the potato rhizosphere were clearly different and neither of the two types of communities was exhaustively sampled, which indicates a continued need to explore the potato rhizopsphere for novelty.
Cultivation of a member of the Verrucomicrobia subdivision 1, a hitherto-uncultured group from the rhizosphere
Improvements of culturability have been described before for soil (Janssen et al., 2002; Schoenborn et al., 2004) and also freshwater (Bruns et al., 2003). These have resulted in the isolation of bacteria that were previously known for their recalcitrance for growth, like those belonging to the Acidobacteria (Janssen et al., 2002). In our study, the approaches used did not result in the cultivation of members of the Acidobacteria, but our sampling was obviously limited. On the other hand, the recovery of six different acidobacterial groups in the clone library indicates that representatives of this phylum are quite abundant in the potato rhizosphere. Acidobacteria, especially members of group 6, have been found in rhizospheres of other plant species (Schmalenberger & Tebbe 2003; Sharma et al., 2005; Wang et al., 2007; Hao et al., 2008; Kielak et al., 2009). Acidobacteria, albeit quite diverse, as-a-group has been
76 Chapter 3
shown to thrive in bulk soils and also in the rhizosphere (Buée et al., 2009; Nunes da Rocha et al., 2009). Representatives of another bacterial phylum recalcitrant for cultivation, the Verrucomicrobia, were also found. The organisms found among the isolates differed from those found in the total fraction, i.e. at the DNA level. The isolates belonged to subdivision 1 whereas the DNA-based evidence pointed to subdivision 3 verrucomicrobia. The isolates matched R. marina but showed substantially higher affiliations with either an unidentified or an uncultured bacterium. R. marina is often found in sponges (Scheuermayer et al., 2006), the unidentified bacterium in the Agrostis stolonifera (creeping bentgrass) rhizosphere (http://turf.lib.msu.edu/tero/v03/n24.pdf; September 28, 2009) and an uncultured bacterium in sediment (Nelson et al., 2007). They thus resemble hitherto-undescribed verrucomicrobial species one of which also originated from rhizosphere soil. The four isolates grouped together in a cluster of species belonging to the Verrucomicrobia, indicating that they belong to a separate branch within this phylum. So far, the Verrucomicrobia encompass cultured as well as uncultured species (Hedlund et al., 1997; Stevenson et al., 2004; Sanguin et al., 2006; Wagner and Horn 2006). Most observations of verrucomicrobial species in soils are from direct molecular studies. Verrumicrobial species often show relative dominance in soils, as up to 9.8 % of the total extractable RNA from e.g. arable soil can consist of verrucomicrobial rRNA (Buckley & Schmidt 2001; Wagner and Horn 2006). It is unclear whether Verrucomicrobia prefer bulk or rhizosphere soils and reports on their prevalence are occasionally contradictory (Buckley & Schmidt 2001; Sanguin et al., 2006; Kielak et al., 2008). It was thus not expected on beforehand that subdivision 1 Verrucomicrobia were present among the isolates in such high frequency. To the best of our knowledge, subdivision 1 Verrucomicrobia isolates have never been obtained from the rhizosphere. Their abundance among potato rhizosphere isolates indicates that they may thrive in this habitat, although their relationship with the plant roots is so far unexplored. Their isolation on CAT and PEX media indicates that both media offer conditions propitious for growth. Although we ignore to which extent this organism dominates the bacterial community in the potato rhizosphere, it can be concluded that subdivisions 1 and 3 of the Verrucomicrobia are present in the potato rhizosphere. Contradicting information about the prevalence of Verrucomicrobia in this habitat may be based on general statements about the entire phylum and not on the individual subdivisions within this phylum.
77 Isolation of Verrucomicrobium spp. subdivision 1 from potato
Conclusions
In conclusion, cultivation was improved and isolate fractions as high as one third of the total microscopically discernable bacterial cells from the potato rhizosphere were obtained. Application of oligotrophic media (either or not amended with potato root exudates or catalase) in combination with selection for mCFUs resulted in raised CFU numbers. However, improved cultivation did not always yield a higher taxonomical diversity of culturable bacteria. Comparison between the isolate and directly detectable bacterial fractions from the potato rhizosphere revealed that there was still a large difference between both. A major finding in this chapter was the isolation of subdivision 1 Verrucomicrobia from the potato rhizosphere. Isolates of this group have not yet been recovered from rhizosphere soil. Considering the relatively high number of isolates belonging to this group, we conclude that the Verrucomicrobia subdivision 1 are important members of the bacterial community in the potato rhizosphere. Their occurrence in the highest-dilution soil suspensions indicates that they numerically dominate the communities of the potato rhizosphere.
Recommendations and perspectives
New isolates of Verrucomicrobia subdivision 1 will be further investigated for their ecophysiological roles in the rhizosphere. Important questions that will be answered in follow-up studies are: 1) to which extent do members of this group prevail in the rhizosphere and 2) what is their relationship with the plant roots and their associated communities. These studies will provide new insight in the roles of newly-cultured bacteria that are thought to dominate plant/soil ecosystems. Thus, new mechanistic effects on plant growth and development may be found, which is difficult to achieve with culture-independent approaches alone.
Acknowledgements This work was supported by the Netherlands Genomics Initiative (Ecogenomics program) and the Dutch Ministry of Agriculture, Nature and Food quality (KB4, research program on sustainable agriculture). We thank CAPES for the fellowship provided to F.D.A (process 2990/05-9). We also thank An Vos, Meint Veninga and Jaap Bloem for their help with total bacterial counts. References
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Barer MR & Harwood CR (1999) Bacterial viability and culturability. p. 93-137. In R. Poole (Ed), Advances in Microbial Physiology 41, Academic Press, New York. Bloem J, Veninga M & Shepherd J (1995) Fully-automatic determination of soil bacterium numbers, cell volumes, and frequencies of dividing cells by confocal laser-scanning microscopy and image- analysis. Appl Environ Microbiol 61: 926-936. Bogosian G & Bourneuf EV (2001) A matter of bacterial life and death. EMBO Reports 2: 770-774. Bruns A, Nubel U, Cypionka H & Overmann J (2003) Effect of signal compounds and incubation conditions on the culturability of freshwater bacterioplankton. Appl Environ Microbiol 69: 1980- 1989. Buckley DH & Schmidt TM (2001) Environmental factors influencing the distribution of rRNA from Verrucomicrobia in soil. FEMS Microbiol Ecol 35: 105-112. Buée M, De Boer W, Martin F, van Overbeek L & Jurkevitch E (2009) The rhizosphere zoo: An overview of plant-associated communities of microorganisms, including phages, bacteria, archaea, and fungi, and of some of their structuring factors. Plant soil 321: 189-212. Cochrane G, Akhtar R, Bonfield J et al (2009) Petabyte-scale innovations at the European Nucleotide Archive. Nucl Acids Res 37: D19-D25. Cole JR, Wang Q, Cardenas E et al (2009) The Ribosomal Database Project: improved alignments and new tools for rRNA analysis. Nucl Acids Res 37: D141-D145. Curl AE & Truelove B (1986) The rhizosphere. Springer-Verlag, Berlin. Da Rocha UN, Tótola MR, Pessoa DMM, Júnior JTA, Neves JCL & Borges AC (2009) Mobilisation of bacteria in a fine-grained residual soil by electrophoresis. J Hazard Mat 161: 485-491. Eilers H, Pernthaler J, Glockner FO & Amann R (2000) Culturability and in situ abundance of pelagic bacteria from the North Sea. Appl Environ Microbiol 66: 3044-3051. Felske A, Wolterink A, van Lis R, de Vos WM & Akkermans ADL (1999) Searching for predominant soil bacteria: 16S rDNA cloning versus strain cultivation. FEMS Microbiol Ecol 30: 137-145. Ferrari BC, Binnerup SJ & Gillings M (2005) Microcolony cultivation on a soil substrate membrane system selects for previously uncultured soil bacteria. Appl Environ Microbiol 71: 8714-8720. Guan LL & Kamino K (2001) Bacterial response to siderophore and quorum-sensing chemical signals in the seawater microbial community. BMC Microbiol 1: 1-11. Hack H, Gall H, Klemke T, Klose R, Meier U, Stauss R & Witzenberger A (1993) Phänologische Entwicklungsstadien der Kartoffel (Solanum tuberosum L.). Nachrichtenbl Deut Pflanzenschutzd 45: 11-19. Hamaki T, Suzuki M, Fudou R, Jojima Y, Kajiura T, Tabuchi A, Sen K & Shibai H (2005). Isolation of novel bacteria and actinomycetes using soil-extract agar medium. J Bioscience Bioeng 99: 485- 492. Hao DC, Ge GB & Yang L (2008) Bacterial diversity of Taxus rhizosphere: Culture-independent and culture-dependent approaches. FEMS Microbiol Lett 284: 204-212. Hedlund BP, Gosink JJ & Staley JT (1997). Verrucomicrobia div. nov., a new division of the bacteria containing three new species of Prosthecobacter. Antonie Leewenhoek 72: 29-38. Heuer H, Krsek M, Baker P, Smalla K & Wellington EMH (1997) Analysis of actinomycete communities by specific amplification of genes encoding 16S rRNA and gel-electrophoretic separation in denaturing gradients. Appl Environ Microbiol 63: 3233-3241. Iizuka T, Yamanaka S, Nishiyama T & Hiraishi A (1998) Isolation and phylogenetic analysis of aerobic copiotrophic ultramicrobacteria from urban soil. J Gen Appl Microbiol 44: 75-84. Ingham CJ, Sprenkels A, Bomer J, Molenaar D, van den Berg A, van Hylckama Vlieg JET & de Vos WM (2007) The micro-petri dish, a million-well growth chip for the culture and high-throughput screening of microorganisms. Proc Natl Acad Sci USA 104: 18217-18222. Janssen PH, Yates PS, Grinton BE, Taylor PM & Sait M (2002) Improved culturability of soil bacteria and isolation in pure culture of novel members of the divisions Acidobacteria, Actinobacteria, Proteobacteria, and Verrucomicrobia. Appl Environ Microbiol 68: 2391-2396. Kaeberlein T, Lewis K & Epstein SS (2002). Isolating “uncultivable” microorganisms in pure culture in a simulated natural environment. Science 296: 1127-1129. Kamagata Y & Tamaki H (2005) Cultivation of uncultured fastidious microbes. Microbes Environ 20: 85-91. Kielak A, Pijl AS, van Veen JA & Kowalchuk GA (2008) Differences in vegetation composition and plant species identity lead to only minor changes in soil-borne microbial communities in a former arable field. FEMS Microbiol Ecol 63: 372-382.
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Kielak A, Pijl AS, van Veen JA& Kowalchuk GA (2009) Phylogenetic diversity of Acidobacteria in a former agricultural soil. ISME J 3: 378-382. Lane DJ, Pace B, Olsen GJ, Stahl DA, Sogin ML & Pace NR (1985) Rapid determination of 16S ribosomal RNA sequences for phylogenetic analyses. Proc Natl Acad Sci USA 82: 6955-6959. Ludwig W, Strunk O, Klugbauer S, Klugbauer N, Weizenegger M, Neumaier J, Bachleitner M & Schleifer KH (1998) Bacterial phylogeny based on comparative sequence analysis. Electrophoresis 19: 554-568. Ludwig W, Strunk O, Westram R et al (2004) ARB: a software environment for sequence data. Nucl Acids Res 32: 1363-1371. Muyzer G, Dewaal EC & Uitterlinden AG (1993) Profiling of complex microbial populations by denaturing gradient gel electrophoresis analysis of polymerase chain reaction-amplified genes coding for 16S ribosomal RNA. Appl Environ Microbiol 59: 695-700. Nelson DM, Ohene-Adjei S, Hu FS, Cann IKO & Mackie RI (2007) Bacterial diversity and distribution in the Holocene sediments of a northern temperate lake. Microbiol Ecol 54: 252-263. Nunes da Rocha U, van Overbeek L & van Elsas JD (2009) Exploration of hitherto uncultured bacteria from the rhizosphere. FEMS Microbiol Ecol 69: 313-328. Pruesse, E, Quast C, Knittel K, Fuchs BM, Ludwig W, Peplies J & Glockner FO (2007) SILVA: a comprehensive online resource for quality checked and aligned ribosomal RNA sequence data compatible with ARB. Nucl Acids Res 35: 7188-7196. Rademakers JLW, Louws FJ & de Bruijn FJ (1998) Characterization of the diversity of ecologically important microbes by rep-PCR genomic fingerprinting. In:. Akkermans ADL, van Elsas JD, de Bruijn FJ (Eds.), Molecular Microbial Ecology Manual, Kluwer Academic Publishers, Dordrecht chapter 3.4.2 pp.1- 27. Rasche F, Hodl V, Poll C, Kandeler E, Gerzabek MH, van Elsas JD & Sessitsch A (2006) Rhizosphere bacteria affected by transgenic potatoes with antibacterial activities compared with the effects of soil, wild-type potatoes, vegetation stage and pathogen exposure. FEMS Microbiol Ecol 56: 219- 235. Rochelle PA, Fry JC, Parkes RJ & Weightman AJ (1992) DNA extraction for 16S ribosomal RNA gene analysis to determine genetic diversity in deep sediment communities. FEMS Microbiol Lett 100: 59-65. Sanguin H, Remenant B, Dechesne A, Thioulouse J, Vogel TM, Nesme X, Moënne-Loccoz Y & Grundmann GL (2006) Potential of a 16S rRNA-based taxonomic microarray for analyzing the rhizosphere effects of maize on Agrobacterium spp. and bacterial communities. Appl Environ Microbiol 72: 4302-4312. Scheuermayer M, Gulder TAM, Bringmann G & Hentschel U (2006). Rubritalea marina gen. nov., sp. nov., a marine representative of the phylum 'Verrucomicrobia', isolated from a sponge (Porifera). Int J Syst Evol Microbiol 56: 2723-2723. Schmalenberger A & Tebbe CC (2003) Bacterial diversity in maize rhizospheres: Conclusions on the use of genetic profiles based on PCR-amplified partial small subunit rRNA genes in ecological studies. Mol Ecol 12: 251-261. Schloss PD & Handelsman J (2005) Introducing DOTUR, a computer program for defining operational taxonomic units and estimating species richness. Appl Environ Microbiol 71: 1501-1506. Schloss PD, Larget BR & Handelsman J (2004). Integration of microbial ecology and statistics: a test to compare gene libraries. Appl Environ Microbiol 70: 5485-5492. Schoenborn L, Yates PS, Grinton BE, Hugenholtz P & Janssen PH (2004) Liquid serial dilution is inferior to solid media for isolation of cultures representative of the phylum-level diversity of soil bacteria. Appl Environ Microbiol 70: 4363-4366. Semenov AM, van Bruggen AHC & Zelenev VV (1999) Moving waves of bacterial populations and total organic carbon along roots of wheat. Microb Ecol 37: 116-128. Sharma S, Aneja MK, Mayer J, Munch JC & Schloter M (2005) Characterization of bacterial community structure in rhizosphere soil of grain legumes. Microb Ecol 49: 407-415. Stevenson BS, Eichorst SA, Wertz JT, Schmidt TM & Breznak JA (2004). New strategies for cultivation and detection of previously uncultured microbes. Appl Environ Microbiol 70: 4748-4755. Ueda K, Tagami Y, Kamihara Y, Shiratori H, Takano H & Beppu T (2008) Isolation of bacteria whose growth is dependent on high levels of CO2 and implications of their potential diversity. Appl Environ Microbiol 74: 4535-4538.
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Van Overbeek LS, Bergervoet JHH, Jacobs F & van Elsas JD (2004) The low-temperature-induced viable-but-nonculturable state affects the virulence in Ralstonia solanacearum biovar 2. Phytopathol 94: 463-469. Van Overbeek LS & van Elsas JD (2008) Effects of plant genotype and growth stage on the structure of bacterial communities associated with potato (Solanum tuberosum L.). FEMS Microbiol Ecol 64: 283-296. Wagner M & Horn M (2006) The Planctomycetes, Verrucomicrobia, Chlamydiae and sister phyla comprise a superphylum with biotechnological and medical relevance. Curr Opinion Biotechnol 17: 241-249. Wang M, Chen JK & Li B (2007) Characterization of bacterial community structure and diversity in rhizosphere soils of three plants in rapidly changing salt marshes using 16S rDNA. Pedosphere 17: 545-556. Zengler K, Toledo G, Rappé M, Elkins J, Mathur EJ, Short JM & Keller M (2002) Cultivating the uncultured. Proc Natl Acad Sci USA 99: 15681-15686.
81 Chapter 4: Isolation and partial characterization of Holophaga, Luteolibacter, unclassified Verrucomicrobia and Verrucomicrobium spp. from the leek (Allium porrum) rhizosphere*
Abstract
Strains affiliated with Acidobacteria (2) and Verrucomicrobia (5) were newly cultured from the leek rhizosphere. Phylogenetic analysis of these isolates plus four other verrucomicrobial strains - previously isolated from potato rhizosphere - were performed. The two acidobacterial strains isolated from leek were affiliated with the class Holophagae (former subgroup 8). All nine Verrucomicrobia belonged to subdivision 1 of this phylum, being that three of these resembled Luteolibacter, five unclassified Verrucomicrobiaceae and one Verrucomicrobium. Strains falling in the same group (Holophaga, Luteolibacter and unclassified Verrucomicrobiaceae) had >97% similarity on the basis of their 16S rRNA gene. They were therefore considered as the same species, but none of them was clonal (as determined by BOX-PCR). Also, a new name for the group of unclassified Verrucomicrobiaceae (Candidatus genus Rhizospheria) to be included in the family Rubritaleaceae (class Verrucomicrobiae, phylum Verrucomicrobia) was suggested. Holophaga isolates had similar phenotypic characteristics, indicating that they may occupy the same ecological niche. The phenotypic diversity within Luteolibacter and Candidatus Rhizospheria isolated from the rhizosphere of leek (this chapter) and those previously isolated from the potato rhizosphere (chapter 3) indicates that these isolates occupy different ecological niches in the soil-plant system.
*Authored by: Ulisses Nunes da Rocha, Jan Dirk van Elsas, Caroline Plugge, Fernando Dini Andreoti, Ines Mandiü-Mulec, Luka Ausec & Leonard Simon van Overbeek Submitted for publication Chapter 4
Introduction
Different reports have been published on the occurrence of Acidobacteria and Verrucomicrobia in the rhizosphere over the past decade (Sanguin et al., 2009; Kielak et al., 2008; Zul et al., 2007; Sanguin et al. 2006; Fillion et al., 2004; Gremion et al., 2003, Chow et al., 2002). Acidobacteria and Verrucomicrobia are bacterial phyla that are widely distributed over different ecosystems. Both represent phylogenetically very diverse groups. The phylum Acidobacteria is considered to be one of the most dominant bacterial groups present across soils (George et al., 2009), sediments (Ben Said et al., 2010), freshwater systems (Hardoim et al., 2009), lichen-associated bacterial communities (Hodkinson & Lutzoni, 2009), groundwater (Spain et al., 2007) and even in domestic toilets (Egert et al., 2010). The phylum Verrucomicrobia was also found to be among the dominant bacterial groups in communities present in soils and rhizospheres (Rosenberg et al., 2009), drinking water reservoirs (Lymperopoulou et al., 2010), human intestinal tract systems (Wang et al., 2005), contaminated groundwater (Herrmann et al., 2008), animal (gorilla) feces (Frey et al., 2006) and swine waste lagoons (Goh et al., 2009). A total of twenty six different phylogenetic groups has been defined so far within the Acidobacteria (Barns et al., 2007), whereas seven subdivisions were found for the Verrucomicrobia (Schlesner et al., 2006). Information on the relative abundance of the individual groups within these phyla in different ecosystems is sparse. Therefore, the ecological niches of the different Acidobacteria and Verrucomicrobia are hardly characterized so far and so is their involvement in ecological processes. Moreover, contradictory information was provided on the occurrence of members of the two phyla in rhizosphere and bulk soils (Chow et al., 2002; Sanguin et al., 2006; Zul et al., 2007; Kielak et al., 2008). The general lack of knowledge is mainly due to the fact that many members of the two groups are recalcitrant to growth in pure culture (Nunes da Rocha et al., 2009). The availability of culturable strains would truly facilitate studies on their behavior in different ecosystems, including the putative involvement in key processes that can be predicted from biochemical, physiological, genetic and cell structural measurements in pure cultures (Zengler, 2009). Although Acidobacteria and Verrucomicrobia are difficult to grow under confined conditions in the laboratory (Jones et al., 2009; da Rocha et al., 2010), procedures for their isolation, mostly from soil, have been described. In fact, successful isolations have been based on standard procedures with small modifications such as: (i) use of media low in nutrient availability (Janssen et al., 2002), (ii) increased levels of
CO2 in the surrounding atmosphere (Stevenson et al., 2004), (iii) selection of microcolonies (Ferrari et al., 2005), (iv) reduction of oxidative stress in the growth medium (Stevenson et al., 2004), (v) application of elongated incubation times (Janssen et al., 2002). Recently, an approach that included several of these modifications simultaneously was applied by us with the aim to improve bacterial isolation from the
83 Isolation of Acidobacteria and Verrucomicrobia from leek
potato rhizosphere. A total of four strains of subdivision 1 of the Verrucomicrobia (da Rocha et al., 2010) was successfully recovered upon plating of rhizosphere samples on agar media that allowed three- to ten-fold higher bacterial recoveries than on standard medium R2A. Questions about the prevalence and ecology of these strains in the rhizosphere still remain open. Also, the potential to obtain other hitherto uncultured Verrucomicrobia and Acidobacteria awaits further work. For our understanding of the ecology of Verrucomicrobia subdivision 1 strains, it is interesting to assess the culturability of organisms from this group from a rhizosphere different from that of the dicotyledonous potato and whether these are distinct from those from potato. In this chapter, we hypothesized that novel Acidobacteria and Verrucomicrobia can be obtained by isolation from the rhizosphere of the monocot leek (Allium porrum). We thus searched for organisms of both groups defined on the basis of 16S rRNA gene sequences. Following isolation, selected strains from both phyla were characterized using a suite of biochemical, physiological, genetic and cell structural approaches. The final goal was to compare strains of the same group from different rhizospheres in order to infer their potential ecological roles in the plant-soil environment.
Material and Methods
Site description and leek rhizosphere soil collection
The site chosen for sampling in this study is an agricultural field located at the experimental farm ‘De Vredepeel’, The Netherlands (51o 32’ 27.10” N and 5o 51’ 14.86” E) where leek was grown. The soil was characterized as a sand (pH 5.4 and 2.2% of organic matter). Leek (Allium porrum) cultivar Kenton (Nunhems Zaden BV, The Netherlands) plants were collected by the end of January 2007 (experiment 1) and the beginning of February 2008 (experiment 2). At both sampling occasions, seven leek plants, including the entire root system and the soil adhering to the roots, were randomly collected from the field with minimal distances of 5 m between the plants. The plants were transported to the lab and processed within 4 hours after collection (da Rocha et al., 2010). Following process, rhizosphere soil suspensions, made from the soil adhering to roots were prepared as previously described (da Rocha et al., 2010) and used for dilution plating, cell counting and total DNA extraction.
Bacterial recovery from the leek rhizosphere on agar media low in available carbon
Bacterial cells in the rhizosphere were enumerated by direct microscopy, as described by Bloem et al. (1995). Preparation of agar media for isolation, i.e. oligotrophic agar medium (OLI) amended or not with catalase (CAT) or leek rhizosphere extract (LEX) was performed as described by da Rocha et al. (2010). R2A (Difco, France) was used as
84 Chapter 4
the reference medium. The following modifications to the incubation conditions used before (da Rocha et al., 2010) were applied, in addition to the previously used ones, for
CAT and LEX plates: these plates were also incubated in an atmosphere of 5% CO2 (elevated CO2) and 16% O2 (further denoted as CAT H and LEX H, respectively). Different plates were used in experiment 1 and 2, as follows: Experiment 1 - R2A, OLI, CAT, LEX, CAT H and LEX H; Experiment 2 - CAT H plates were used. Significance of differences between cell and CFU numbers on different agar media were calculated by analysis of variance (ANOVA) using GenStat 12th edition (VSN International Ltd., UK). Differences were considered to be significant at levels of P = 0.05 and lower. After 15 days of incubation, 360 (60 from each agar medium/incubation condition) colonies next to 225 (45 each from OLI, CAT and LEX incubated at normal and elevated CO2) microcolonies (mCFU, colonies between 80-250 µm and only visible under 66 x magnification) from experiment 1 and 150 colonies from experiment 2 were randomly picked from plates that had received the two highest dilutions of the rhizosphere soil suspensions. Material from all colonies grew when streaked to purity on their respective agar media under the adequate CO2 incubation conditions after 5 days of incubation for colonies and up to 150 days for mCFUs. Upon colony formation, material from single separate colonies was again transferred to fresh agar media of the same composition and incubated under the respective CO2 conditions to allow the formation of new colonies. All pure isolates were taken up and stored in stock solutions containing oligotrophic broth (OLI medium with omission of agar) supplemented with glycerol (final concentration 20%, w/w) at -80oC.
Screening for representatives of Acidobacteria and Verrucomicrobia via 16S rRNA gene-assisted identification
Genomic DNA was extracted from all (in total 735) isolates using the MasterPure DNA purification kit (Epicentre, WI, USA) following instruction of the manufacturer. PCR amplification using primers 27F (Lane et al., 1985) and 1492R (Rochelle et al., 1992) was then performed on all DNA extracts, followed by single-strand sequencing of all amplicons using primer 1492R. Almost-complete 16S rRNA genes (> 1200 bp) were then sequenced for those isolates that showed closest matches with Acidobacteria and Verrucomicrobia (SILVA database, release 102 NR - Pruesse et al., 2007) as described in da Rocha et al. (2010). These larger sequences were again compared to those of the SILVA database using ARB software (Ludwig et al. 2004). Strains that showed closest matches with 16S rRNA gene sequences of Acidobacteria and Verrucomicrobia were selected for further analyses. These were supplemented with four strains from the potato rhizosphere identified as Verrucomicrobia subdivision 1 isolates (da Rocha et al. 2010).
85 Isolation of Acidobacteria and Verrucomicrobia from leek
Strains from leek and potato rhizospheres were different at the genome level as evidenced using BOX-PCR
BOX-PCR genomic profiling (Rademakers et al., 1998) of the 11 selected strains was performed to further distinguish these. DNA extraction was performed using the MasterPure DNA purification kit (Epicentre, WI, USA) following instruction of the manufacturer. Aliquots (1 µL) containing 1 ng of DNA were used as the templates for PCR reactions. After PCR amplification, 10 ȝ/ of each mixture was loaded onto 1.5 % agarose gel (20 cm in size), and gels were run at 24 V for 16h, 4oC. Each gel contained three lanes, located at different places, loaded with 5 ȝ/ of 1-kb ladder (Invitrogen, Carlsbad, Ca) for normalization of the fingerprints. After staining with ethidium bromide, gel images were digitized using a digital camera and the digitized fingerprints were used for analysis with Gelcompar II software (Applied Maths, Belgium).
Morphological and physiological characterization of selected strains from leek and potato rhizospheres
Morphological and physiological parameters of selected isolates on individual cell and colony levels were determined. Suspensions with grown cells of all stains were Gram- stained. Then, the morphologies (cell morphological type, length and width) of at least 100 individual cells (at 1,000 x magnification) were determined according to Doetsch (1981). Further, cell motility was determined (Smibert & Krieg, 1981), assessing at least 50 cells per suspension, using the same magnification level. Also, colony size and morphology on the same colonies were described on at least 10 colonies per isolate (Doetsch, 1981). To test for growth in liquid medium, suspensions were made of cells from single colonies on OLI agar. Optical densities of the resulting suspensions (OD600) were set at 0.05 before introduction into liquid OLI, 0.1 strength trypticase soy broth (1/10 TSB, BD, France), R2A, King’s B (Roitman et al., 1990) and Luria-Bertani (LB) media (Sambrook et al., 1989). Cultures were incubated at 28o C, with shaking, and the increase of OD600 values was measured. To test for growth on solid media, 5 ȝ/ cell suspensions in OLI were spread over the surfaces of R2A, 1/10 TSA, King’s B and LB agar media (purified agar, 12.5 g L-1 used). Plates were incubated at 28o C in the dark and monitored daily for colony growth to up to 3 months after inoculation. Growth rates at colony level were determined on R2A in accordance with Wimpenny & Lewis (1977). Escherichia coli K12 was used as a reference strain for growth rate measurements at colony level. Colony growth at different pH was tested on OLI agar with modified pH. The pH buffer composition described in Costilow (1981) was used, i.e. pH 4.0 was set with 0.1 M acetate buffer, pH 5.0 - 0.1M acetate buffer, pH 6.0 - 0.1M citrate-phosphate buffer, pH 7.0 - 0.1M phosphate buffer and pH 8.0 - Tris-Cl buffer. To test for growth
86 Chapter 4
on organic acids and amino acids, we used OLI agar without glucose and casein hydrolysate supplemented with 54 mg L-1 of either oxalic acid, DL-(-)-malic acid, succinic acid, citric acid, L-glutamine or DL-alanine (Sigma-Aldrich Company Ltd., UK). Following inoculation, plates were incubated for up to 2 months at 25oC and colony growth was determined regularly. As controls, OLI (da Rocha et al., 2010) without any carbon source was also inoculated and these plates were incubated and examined similarly. Growth on cellulose and cellulolytic activity measurements near the colonies were performed on OLI agar, with the glucose and casein hydrolysate substituted by 54 mg L-1 of cellulose (Sigmacell®, Sigma Chemical Co., USA). Colony formation and eventual formation of haloes surrounding individual colonies were recorded over time as in Smibert & Krieg (1981). The presence of putative laccase genes in the genomes of the selected isolates was determined on genomic DNA extracts as templates using the PCR primers and conditions described by Ausec & Mandic-Mulec (2010).
Nucleotide sequence accession numbers
DNA sequences of the almost-complete 16S rRNA genes of the Acidobacteria and Verrucomicrobia strains recovered in this study were deposited in the EMBL Nucleotide Sequence Database under accession numbers FN554388 to FN554392 and FN689719 to FN689720. DNA sequences of putative laccase gene were deposited in the EMBL Nucleotide Sequence Database under accession numbers HM453207 to HM453211
Results
Bacterial numbers in the leek rhizospheres
Experiment 1. Seven healthy mature leek plants sampled from the V field soil revealed the presence of between Log 9.71 and 9.97 DTAF-stainable cells per g of dry soil (Table 1). The total Log colony numbers (including macro- and microcolonies) per g of dry soil from the same samples were in the range 7.87 - 8.61 on R2A; 8.91 - 9.39 on OLI; 9.10 - 9.47 on CAT; 9.16 - 9.48 on CAT H; 9.13 - 9.49 on LEX; and 9.12 - 9.39 on LEX H. This yielded colony recovery percentages (expressed as fraction of the number of DTAF-stainable cells) of 1.3 -4.4 on R2A, 15.9-26.1 on OLI, 24.6-31.9 on CAT, 25.5-32.3 on CAT H, 23.3-33 on LEX and 23.4-28.8 on LEX H. The culturability levels were clearly raised on OLI, CAT, CAT H, LEX and LEX H as compared to R2A. In fact, the differences between the CFU numbers on R2A versus those on OLI, CAT,
CAT H, LEX and LEX H , irrespective of CO2 level, were significant (P<0.05), whereas the CFU numbers on the latter media were statistically similar. We decided to focus on
87 Isolation of Acidobacteria and Verrucomicrobia from leek CAT H Experiment 2 2 Experiment 9.68 9.09 (25.5) 9.09 9.68 ct; H for medium H for medium ct; stainable cells cells stainable
C izosphere extra 9.23 (24.0) number to percentage and DTAF stainable cells cells DTAF and stainable to percentage number -1 ase or leek rh C,D,E dry soil -1 fferent leek plants sampled in two different 9.27 (26.4) 9.27 E CFU the ratio (%) g of 9.30 (28.4) non-amended or catal amended with D,E D,E CAT CAT H LEX LEX H DTAF < 0.05) < 0.05) p Experiment 1 1 Experiment 9.28 (27.0) analysis made was with B a 8.91 (15.9) 8.91 (15.9) 9.10 (24.6) 9.16 (28.6) 9.13 (26.3) 9.12 (25.9) 9.65 9.11 (28.8) ia and CFU (normal plus microcolonies) in 7 di 9.11 (18.6) b A concentration concentration R2A OLI 2 and 16% O 16% and 2 9.71 7.87 (1.5) 7.87 9.71 9.77 9.80 9.83 7.87 (1.3) 9.90 8.09 (2.0) 9.01 (17.6) (2.7) 9.96 8.27 (2.7) 9.02 (16.7) 9.18 (25.8) 9.97 8.38 (3.0) 9.08 (17.6) 9.22 (26.3) 9.18 (26.1) 8.24 9.85 8.55 (3.9) 9.16 (18.4) 9.25 (26.1) 9.20 (25.5) 9.18 (25.9) 0.10 8.61 (4.4) 9.20 (17.5) 9.29 (24.9) 9.31 (30.0) 9.22 (26.3) 9.15 (24.3) 9.39 (26.1) 9.43 (29.6) 9.36 (29.0) 9.25 (26.1) 9.19 (24.4) 0.30 (1.2) 9.47 (31.9) 9.40 (27.5) 9.28 (24.1) 9.21 (23.7) 9.48 (32.3) 0.16 (3.4) 9.62 9.33 (23.3) 9.23 (21.7) 9.67 9.49 (33.0) 9.29 (21.5) 0.14 (2.7) 9.72 9.03 (25.7) 9.39 (26.2) 0.12 (2.3) 9.78 9.07 (25.1) 9.59 0.12 (3.1) 9.09 (23.4) 9.74 9.17 (24.5) 0.09 (1.8) 8.96 (23.4) 9.18 (27.5) 0.07 0.08 (2.0) DTAF Total (DTAF-stainable) bacter stainable cells cells stainable OLI, CAT, lowOLI, LEX: carbon availability agar respectivelymedium, >Significantly different, where A B > StatisticalC.
cells CFU stainable and DTAF (%) between Ratio c occasions. 1. Table a b c (arcsine transformation) and calculated with one way ( ANOVA one way calculated and with transformation) (arcsine incubated at 5% CO 5% at incubated
88 Plant Plant 2 4 6 Avarage Standard deviation Chapter 4
CAT H for further isolations in experiment 2, as this medium yielded highest fractional colony recoveries. Experiment 2. The DTAF-stainable cell numbers in seven leek rhizosphere samples were between Log 9.59 and 9.78 cells per g dry soil and CFU numbers on CAT H were between Log 8.96 and 9.18 CFU per g of dry soil, resulting in recovery percentages of between 23.4 and 28.8. CFU recovery on CAT H was again significantly higher than that on R2A in the same experiment (1.8-3.1%; not shown).
Selection of isolates belonging to the Acidobacteria and Verrucomicrobia
A total of 735 isolates (585 from experiment 1 and 150 from experiment 2) were screened for the presence of isolates belonging to the Acidobacteria or Verrucomicrobia by partial sequencing of their 16S rRNA genes (amplicons approximately 350 bp in size). Among the isolates of experiment 1, a total of five in 585 (0.9%) presumptively fell in the Acidobacteria/Verrucomicrobia. Specifically, one isolate (CHC25) from CAT H showed >90% similarity (sim) to Geothrix fermentans (Acidobacteria), and four to Rubritalea marina (Verrucomicrobia subdivision 1). Of the latter, one (ONA9) was obtained from OLI, one (CNC16) from CAT, and two (CHC8 and CHC12) from CAT H (Table 2). In experiment 2, two of 150 (1.2%) isolates fell in the target phyla, being one (ORAC) from CAT H. This organism showed >90% similarity to Geothrix fermentans (Acidobacteria). The other one (IRVE), obtained from CAT H, affiliated to Rubritaea marina (Verrucomicrobia subdivision 1) (Table 2). All isolates were isolated from media and conditions that allowed higher colony recoveries and even so appeared at a prevalence of roughly 1%. This strongly indicated that increased culturability on specific media is required to efficiently obtain representatives of these phyla from the leek rhizosphere. There was no preference for growth on any agar medium or CO2 level in particular. The strains obtained from the leek rhizosphere were pooled with four strains isolated from the potato rhizosphere, belonging to Verrucomicrobia subdivision 1 (da Rocha et al. 2010). On the basis of their almost-complete 16S rRNA genes, the new leek rhizosphere strains were subjected to further phylogenetic analyses. Strains CHC25 and ORAC, which were tentatively found to affiliate with Acidobacteria in the first run, again affiliated with this phylum, showing closest matches with Geothrix fermentans (accession number U41563, 94.2% sim), class Holophagae (previously known as Acidobacteria group 8). However, the highest similarity (97.5%) of both was with an uncultured bacterium (accession number GU169059) recovered from “synthetic river water with humic substances” (Fig 1A). The five strains isolated from leek rhizosphere
89 Isolation of Acidobacteria and Verrucomicrobia from leek
A 91 Acidobacterium bacterium CHC25, FN554392 Acidobacterium sp. ORAC, FN689719 58 Uncultured bacterium, GU169059 Holophagae Uncultured bacterium, EU331375 63 Geothrix fermentans, U41563 96 Holophaga foetida, X77215 Acanthopleuribacter pedis, AB303221
74 20 Group 6
80 20 Group 4
22 Group 1
81 20 Group 2 74 19 Group 3
30 Candidate division TM6
uncultured bacterium, FJ716054 89 Verrucomicrobiaceae bacterium CHC12, FN554390 uncultured bacterium, FJ716011 90 Verrucomicrobiaceae bacterium C20*, FN394506 Luteolibacter Luteilibacter pohnpeiensis, AB331895 B 97 88 Verrucomicrobiaceae bacterium ONA9, FN554388 99 uncultured bacterium, FJ264561 99 Luteilibacter algae, AB331893 Verrucomicrobiaceae bacterium CHC8, FN554389 uncultured bacterium, AB240273 uncultured bacterium, EF188433 Unclassified uncultured bacterium, FJ542838 Verrucomicrobium sp. IRVE, FN689720 Verrucomicrobiaceae 93 uncultured bacterium, EU979055 Verrucomicrobiaceae bacterium CR28*, FN394505 (Candidatus genus Subdivision 1 Verrucomicrobiaceae bacterium Z35*, FN394511 Rhizosphereae) 86 uncultured bacterium, DQ814962 uncultured bacterium, AY345492 98 uncultured bacterium, EU135492 97 Verrucomicrobiaceae bacterium ZNBB5*, FN394508
83 Haloferula
84 Rubritalea Verrucomicrobiaceae bacterium CNC16, FN554391 Verrucomicrobium spinosum, X90515 Prosthecobacter Subdivision 2 Subdivision 5
79 94 Subdivision 6
93 39 Subdivision 3
24 Subdivision 4 75 Subdivision 7 Chlamydiae
Figure 1 Taxonomic affiliation of Verrucomicrobia (A) and Acidobacteria (B) isolates obtained from the leek and/or potato rhizosphere. Distances between partial 16S rRNA gene sequences over 1100 bp in length were calculated using ARB software, the topology was reconstructed using ARB neighbour joining. A cluster made of 16S rRNA gene sequences of 10 different species from the Chlamydiae (A) or Candidate division TM6 (B) were used as out groups. Isolates followed by an asterisk (*) were recovered by Nunes da Rocha et al. (2010); and isolates in bold were recovered in this study. Bar at the bottom indicate 10% divergence among sequences and bootstrap values (calculated from 1000 iterations, %) are presented near each junction in the tree.
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(2010) (2010) (2010) (2010) (2010) (2010) (2010) (2010) d/collection Reference 2007 Vredepeel This study study This Vredepeel 2007 2008 Vredepeel This study study This Vredepeel 2008 2007 Vredepeel This study study This Vredepeel 2007 2006 2006 Droevendaal al. et da Rocha Nunes 2006 2006 Droevendaal al. et da Rocha Nunes 2006 2006 Droevendaal al. et da Rocha Nunes 2007 Vredepeel This study study This Vredepeel 2007 2007 Vredepeel This study study This Vredepeel 2007 2006 2006 Droevendaal al. et da Rocha Nunes 2007 Vredepeel This study study This Vredepeel 2007 2008 Vredepeel This study study This Vredepeel 2008 Year of isolation isolation Year of Fiel
Solanum Solanum Solanum Solanum Solanum Solanum Solanum Solanum tuberosum tuberosum tuberosum tuberosum Allium porrum Allium porrum Allium porrum Allium porrum Allium porrum Allium porrum Allium porrum
Plant type Plant a pohnpeiensis
Verrucomicrobia Verrucomicrobia Verrucomicrobia (98.7%) (98.7%) bacterium EU979055 (98.5%) EU979055 (98.5%) bacterium DQ815271 (97.3%) (97.3%) DQ815271 (97.9%) (97.9%) bacterium EU979055 (98.8%) EU979055 (98.8%) bacterium bacterium EU979055 (98.2%) EU979055 (98.2%) bacterium Luteolibacter AB331895 (98.0%) (98.0%) AB331895 (97.6%) (97.6%) (98.6%) (98.6%) Nearest match Nearest (94.7%) (94.1%) (94.1%) CNC16 uncultured bacterium FJ230907 bacterium CNC16 uncultured IRVE uncultured CHC8 uncultured bacterium CHC8 uncultured ZNBB5 ZNBB5 EU135492 bacterium uncultured Z35 uncultured CR28 uncultured ONA9 CHC12 uncultured bacterium FJ716054 bacterium CHC12 uncultured C20 uncultured bacterium FJ715972 bacterium C20 uncultured olate CHC25 EU937971 bacterium uncultured ORAC EU937971 bacterium uncultured ) genus Bacterial isolates used in this study. Bacterial isolates used in this study. Canditatus Verrucomicrobium Verrucomicrobium Unclassified Unclassified Verrucomicrobiaceae ( Rhizospheria Luteolibacter Affiliation Isolate/is Holophagae (http://simo.marsci.uga.edu/public_db/analysis_login.asp) tools analysis SIMO RDP using accordance with In a 2. Table
91 Isolation of Acidobacteria and Verrucomicrobia from leek
that fell in the Verrucomicrobia were shown to affiliate with different groups within this phylum. Strains CHC12, C20 and ONA9 showed nearest matches with Luteolibacter pohnpeiensis (accession number AB331895, 97.1 and 98.0% sim), genus Luteolibacter, subdivision 1 Verrucomicrobia (Fig. 1B). Strains CHC8 and IRVE grouped in a separate cluster denoted in the RDP classification as ‘unclassified Verrucomicrobiaceae’ of subdivision 1 (Fig. 1B). Finally, strain CNC16 showed the closest match with Verrucomicrobium spinosum (accession number X90515, 98.6% sim), genus Verrucomicrobium, subdivision 1 (Fig. 1B). Of the four potato rhizosphere strains, one, i.e. C20, grouped with Luteolibacter pohnpeiensis (accession number AB331895, 97.3% sim), whereas the remaining strains, i.e., CR28, Z235 and ZNBB5, clustered with unclassified Verrucomicrobiaceae (Fig. 1B). Reports on the internal matches between the novel strains on the basis of their almost-complete 16S rRNA gene sequences, Holophaga CHC25 and ORAC were 98.9% similar, Luteolibacter CHC12, C20 and ONA9 97.4 - 99.0%, whereas unclassified Verrucomicrobiaceae CHC8, IRVE, CR28, Z35 and ZNBB5 were 97.6 - 99.1% similar (Table 3). In contrast, the BOX-PCR profiles (see Appendix Fig. A1) never showed similarities exceeding 91% Pearson correlation between strains (the cut- off limit set to 100 % similarity) (see Appendix Fig. A1), indicating that none of the isolates was identical to any other one.
Mophological and physiological characterization of the novel strains
The 11 novel Holophagae, Luteolibacter and unclassified Verrucomicrobiaceae strains were characterized at cell and colony levels (Table 4). In all cases, strains turned out to be Gram-negative, motile and able to grow on OLI agar, and even R2A media under atmospheric conditions. They were, therefore, considered to be aerobic and heterotrophic bacteria. Luteolibacter CHC12 and ONA9, unclassified Verrucomicrobiaceae CHC8 and Verrucomicrobium CNC16 were able to grow on 1/10 TSA, whereas none of the isolates grew on King’s B or LB agar. Unclassified Verrucomicrobiaceae CHC8 and Verrucomicrobium CNC16 were the only strains that grew in liquid 1/10 TSB. Holophagae CHC25 and ORAC formed small rough colonies (1.0 mm in diameter), whereas those of Verrucomicrobia were larger (between 1.4 - 4.8 mm in diameter), isolate CHC8 being the largest for the unclassified Verrucomicrobiaceae. The colonies of all Verrucomicrobia isolates have a smooth shiny surface.
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Verrucomicrobium
) subdivision 1 isolates Rhizospheria 84.7 84.3 97.6 99.0 98.2 80.9 72.6 73.4 73.8 72.6 73.4 73.8 71.1 73.0 73.5 74.2 70.0 93.9 93.8 94.4 83.7 94.0 94.5 84.9 94.4 94.3 94.6 83.8 97.6 97.5 97.4 84.0 99.1 99.0 97.6 82.9 CR28 Z35 ZNBB5 ZNBB5 CNC16 CR28 Z35 genus genus Verrucomicrobiaceae Verrucomicrobiaceae Verrucomicrobia
71.2 70.3 93.0 93.5 93.5 97.1 IRVE and Canditatus Canditatus ( Unclassified Holophagae
Luteolibacter Luteolibacter
(%) of the 16S rDNA ofgene the different a Holophagae Holophagae Similarity
Table 3. Table
2004) al., et (Ludwig software ARB using determined were levels Similarity a Isolate CHC25 ORAC CHC12 C20 ONA9 CHC8 ONA9 C20 CHC12 Taxa ORAC CHC25 Isolate CHC25 ORAC 98.9 71.1 73.9 70.5 70.8 70.5 74.5 70.5 70.5 CHC12 94.4 C20 ONA9 97.8 CHC8 99.0 97.4 IRVE 93.3 CR28 Z35 93.4 ZNBB5 CNC16
93 Isolation of Acidobacteria and Verrucomicrobia from leek
Holophaga foetida CHC25 and ORAC only grew on OLI agar set at pH 4.0 and 5.0 and not on those with higher pH. Luteolibacter CHC12 and ONA9 grew on OLI agar with pH 6.0, 7.0 and 8.0 and not on those with lower pH. C20 only grew on OLI with pH 7.0 and 8.0. Unclassified Verrucomicrobiaceae strain CHC8 only grew on OLI agar at pH 5.0, 6.0, 7.0 and 8.0, CR28 only at pH 6.0 - 8.0 and IRVE, Z35 and ZNBB5 only at pH 7.0 and 8.0. Verrucomicrobium CNC16 grew on OLI agar at pH 8.0 and not at lower pH levels. The Holophagae strains thus grew under more acidic circumstances, whereas subdivision 1 Verrucomicrobia preferred more neutral to basic circumstances. All strains were then examined for growth on oxalic acid, malic acid, succinic acid and citric acid, as well as the amino acids glutamine and alanine, as sole carbon sources. After 60 days of incubation, Holophaga CHC25 and ORAC grew on OLI agar containing either malic acid, succinic acid, citric acid, glutamine or alanine, but not on that containing oxalic acid. Luteolibacter CHC12, C20 and ONA9 differed from each other in carbon source utilization. C20 grew on all six OLI agar media, ONA9 only on the ones with glutamine and alanine and CHC12 did not form colonies at all on any of the six agar media. The five unclassified Verrucomicrobiaceae also showed different carbon source utilization patterns; CHC8 grew on all six carbon sources, IRVE only on oxalic acid, malic acid, glutamine and alanine, CR28 on succinic acid, citric acid and glutamine, Z35 on citric acid, glutamine and alanine and ZNBB5 did not form colonies with organic acids but grew on glutamine and alanine. Verrucomicrobium CNC16 only grew on OLI agar with glutamine and alanine. The estimated growth rates of Holophaga strains CHC25 and ORAC were 12.3h and 13.8h, respectively, per cell division. The subdivision 1 Verrucomicrobia strains varied in estimated doubling times, i.e. between 9.6 and 68.8 h per cell division. The Luteolibacter CHC12, C20 and ONA9 had growth rates between 9.6h and 65.5h per cell division, the unclassified Verrucomicrobiaceae CHC8, IRVE, CR28, Z35 and ZNBB5 between 15.6h and 68.8h and Verrucomicrobium CNC16 14.6h. All strains grew significantly slower than the reference strain E. coli K12 (37 min) under the same conditions. Growth on OLI agar with cellulose, cellulose hydrolysis activity and presence of putative laccase genes were measured. Only unclassified Verrucomicrobiaceae CHC8 showed growth and halo formation on OLI agar containing cellulose as sole carbon source. Putative laccase genes, using laccase gene-specific PCR amplification, were found to be present in five subdivision 1 Verrucomicrobia, ie. Luteolibacter CHC12 and ONA9, unclassified Verrucomicrobiaceae IRVE and CR28 and Verrucomicrobium CNC16. Sequence analyses of amplified fragments from all five isolates by BLAST- assisted searches revealed that all five isolates contained a putative 3-domain laccase gene (see Appendix Table 1).
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Overall, the two Holophagae strains strongly resembled each other, whereas the nine subdivision 1 Verrucomicrobia substantially differed from each other.
Discussion
The major outcome of this chapter is the isolation of novel Acidobacteria and Verrucomicrobia strains from leek. Remarkably, all 11 strains belonged to single subgroups within each phylum, i.e. to the Holophagae (formerly known as group 8 Acidobacteria) and to the subdivision 1 Verrucomicrobia. The isolation of subdivision- 1 Verrucomicrobia from leek was in line with previous isolations from potato (da Rocha et al., 2010). This would indicate that subdivision-1 Verrucomicrobia are typical rhizosphere bacteria that abound in different plant species. However, the nine strains that we analyzed strongly differed from each other in phenotypic and genotypic terms, being classified in three distinct clades within the subdivision. Differences in their preferred niches in the rhizosphere are likely on the basis of the data; this aspect will be further investigated in a follow-up study. The close resemblance of the two leek Holophaga foetida strains indicates this monophyletic group within the Acidobacteria may particularly associate with leek. However, it is not possible to draw firm conclusions on the basis of the low number (two) of strains from this group obtained in the current study. All novel strains were isolated under conditions that allowed significantly raised recovery rates from rhizosphere soil. Bacterial counts rising up to 33% of the total bacterial cell fractions were reached, corroborating what we previously achieved for the potato rhizosphere using the same media (da Rocha et al., 2010). Improved bacterial culturability as a result of incubation under conditions better-tuned to the natural environment has been achieved before with bulk soil (Janssen et al., 2002; Sait et al., 2002; Schoenborn et al., 2004; Davis et al., 2005) and freshwater samples (Bruns et al., 2003). In these studies, key factors contributing to improved cultivation were: i) reduced nutrient availability, ii) prolonged incubation times and iii) reduction of oxidative stress by the addition of protective agents. Recoveries from natural environments can thus be increased by simple modifications of already existing protocols, yielding access to hitherto uncultured bacterial groups, as demonstrated in the current study as well as in others (Janssen et al., 2002). Strikingly, two Holophagae were obtained from the leek rhizosphere. To the best of our knowledge, this is the first report on isolation of an Acidobacterium species from the plant rhizosphere. The class Holophagae represents a small group within the Acidobacteria. Estimated population sizes may range from 0 and 3.4 % of the total Acidobacteria community present in (bulk) soils (Jones et al., 2009). Due to the fact that
95 Isolation of Acidobacteria and Verrucomicrobia from leek Staphylo rod Staphylo isolates isolates Verrucomicrobium rod rod neg neg Single
) Verrucomicrobia pos. pos. Single fusiform Rhizospheria rod rod Single subdivision 1 subdivision genus Verrucomicrobiaceae Verrucomicrobiaceae and subdivision 1 rod rod neg neg neg. neg neg Staphylo Staphylo Canditatus ( Unclassified Unclassified pos. pos. Single Acidobacteria fusiform cocci Staphylo Staphylo rod rod Single rod rod Single Verrucomicrobia short chains chains short Filamentous Holophagae Holophagae
- - + - + + - - - - + - - - - + + - - - - - + + - - - + - - + - - + + - - - - + + + - - + - - + - - CHC25 ORAC CHC12 C20 ONA9 CHC8 IRVE CR28 Z35 ZNBB5 Z35 CNC16 CR28 IRVE CHC8 ONA9 C20 CHC25 ORAC CHC12 short chains short Luteolibacter Acidobacteria Acidobacteria
Morphological, characteristics and physiological Morphological, phenotypical of
d e neg. neg. neg neg neg neg. neg neg neg neg. neg. a + + + + + + + + + + + + + + + + + + + + + b Alanine + + - + + + + - + + + + - + + + + + + + Glutamine + + + + + + + Alanine + - + - 7.0 - - + + + + + + + + - + + + + + + + + + + + + + + + + + 5.0 - 6.0 - - 7.0 - 8.0 + - + - - - + - - - - + + + - - + - - - Malic acid + + acid Malic + - - + Succinic + + - - - + acid Citric + - - + + - - + - - acid King’s B - - King’s - LBA ------OLI + + + + + + + + + + + + + + + + + + + + + + + + + + + + + + OLI + + R2A TSA 1/10 from leek and potato rhizospheres Table 4. Oxalic g f
c Agar Media Agar Liquid Media Liquid pH 4.0 4.0 pH + + ------OAs AAs Growth Colony morphology morphology Colony Cell motility Characteristic Gram staining Cell morphology Filamentous
96 Chapter 4
white bright King’s B and LB. Positive yellow acid .1% sodium azide sodium .1% old cultures (more than 30 days 30 days than (more cultures old sent absent sent absent absent bright curled curled curled curled entire yellow bright yellow bright yellow circular circular circular circular circular circular circular circular circular id viscous viscous viscous viscous viscous viscous id viscous viscous 2h 68.8h 68.8h 33.8h 15.6h 15.6h 33.8h 68.8h 68.8h 2h 14.6h rled curled curled curled rled curled sent present present absent absent absent absent present present sent present esent absent absent ab absent absent esent bright yellow spindle cultureyoung negative Gram were and bright yellow 2A addition2A without of agar), 1/10TSB broth), soy (trypticase to be motile be to motile when movements arrested upon administration 0 of s measured changing Glucose for the respective organic or amino organic or amino respective Glucose the for s changing measured yellow opaque opaque
pink pink white / white stained positive; Gram neg. pos., cells of yellow / brownish brownish 1.0 1.0 1.0 1.7 1.9 2.3 4.8 1.4 1.8 1.8 2.0 1.9 12.3h 13.8h 32.7h 65.5h 9.6h 18. 9.6h 65.5h 32.7h 13.8h 12.3h brownish brownish absent absent absent absent absent pr ab absent absent present absent absent absent absent present absent absent
AAs, amino acids acids amino AAs,
Form punctiform punctiform circular circular circular slightly circular circular circular punctiform Size (diameter, Form punctiform Texture dry dry viscous viscous viscous muco viscous viscous / viscous dry Surface Texture dry Color yellow rough rough shine shine shine shine shineshine shine shine shine Margin entire entire entire curled curled cu curled curled entire entire Elevation entire Margin flat flat flat convex convex convex convex convex convex convex flat Determined light by –microscopy bacterial cells consideredwere carbon Liquid used – Low media availability (OLI), R2Bmedium (R hours h, on R2A; phase growth exponential during size colony of the bases on estimated time Doubling neg., cell stained Gram negative; pos., cells +, positive growth; -; negative growth carbon low availability OLI, medium; TSA (trypticase soy agar broth) acids amino and wa acids acids. Growth in organic organic OAs, incubation days 14 after developed was colour Pink growth was only detected on 1/10 TSB TSB 1/10 on detected only was growth old) stained positive Gram a b c d e f h g i
h Cellulase activityCellulase Laccase gene Laccase mm) mm) Colony doubling time time doubling Colony (g)
97 Isolation of Acidobacteria and Verrucomicrobia from leek
other Acidobacteria, i.e. those of groups 1, 2, 3, 4 and 6, are present in much higher numbers in soils, Holophagae may have been overlooked in the past. Also, members of this group may prefer sites proximate to plant roots. Other Acidobacteria, especially group 6 ones, have been detected before in the rhizosphere of different plants via culture- independent approaches (Schmalenberger & Tebbe, 2003; Sharma et al., 2005; Wang et al., 2007; Hao et al., 2008). However, no culturable representatives from the group-6 Acidobacteria have ever been recovered from the rhizosphere. Many uncertainties still exist about the roles of the cultured fastidious bacteria in the rhizosphere. The Holophagae, strains CHC25 and ORAC, were motile, indicating the possibility of chemotaxis, adherence to substrates, biofilm formation and even swarming (Young, 2007). The filamentous short chains formed by CHC25 and ORAC cells may be an indication that these bacteria are adapted to resist bacterivory (Young, 2007). Also, strain CHC25 and ORAC cells could not grow in liquid media, a phenomenon observed for other Acidobacteria isolates (Valáková et al., 2009). Within the group of Holophagae, only three cultures have been described to date, i.e.Holophaga foetida (accession number X77215), Geothrix fermentans (accession number U41563) and Acanthopleuribacter pedis (accession number AB303221). Both Geothrix fermentans (Coates et al., 1999) and Holophaga foetida (Liesack et al., 1994) are strictly anaerobic bacteria isolated from hydrocarbon-contaminated areas, whereas Acanthopleuribacter pedis is a strictly aerobic bacterium isolated from a beach chiton (Fukunaga et al., 2008). This demonstrates that the physiology within this group may be diverse. Recently, Holophagae were detected in clone libraries made from the endophytic bacterial community in rice (Oryza sativa L.) roots (accession number DQ340903) (Sun et al., 2008), as well as from the rhizosphere of Phragmites (accession number AB240249). We hypothesize that particular members of Holophagae live in association with plants, either in the rhizosphere or even as endophytes. It may well be that the ones that live in association with plants, or eukaryotes in general, actually have an aerobic and heterotrophic lifestyle. A high diversity among the subdivision-1 Verrucomicrobia was further observed in the current study and in that of da Rocha et al. (2010). Although 16S rRNA gene sequences from this phylum may be more abundant in rhizosphere than in bulk soils (Chow et al. 2002; Zul et al. 2007), no information about their ecological roles in the rhizosphere is available. All unclassified Verrucomicrobiaceae were able to grow on organic acids and/or amino acids. These compounds are common in the rhizospheres of different plant species and our novel strains will likely be able to grow in the proximity of roots of plant species that exude such compounds (Jones, 1998; Baudoin et al., 2003). Cells of CHC8 showed cellulase activity, indicating that they may be able to grow on plant material in soil and
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even colonize living roots (Mostajeran et al., 2007) and/or the interior tissues of plants (Bischoff et al., 2009). Five of the nine Verrucomicrobia subdivision 1 isolates yielded evidence for the presence of laccase genes, which is interesting as these enzymes may specifically oxidize phenolic and non-phenolic lignin-related compounds (Kunamneni et al., 2008). Subdivision-1 Verrucomicrobia have been isolated before from other environments, e.g. from soil, freshwater (Schlesner, 1987; Hedlund et al., 1996) and marine environments (Hedlund et al., 1996; Scheuermayer et al., 2006; Yoon et al., 2008). The strains found here fell in three groups, Luteolibacter, unclassified Verrucomicrobiaceae and Verrucomicrobium (Verrucomicrobia subdivision 1). This is the first report on cultivation of members of this “unclassified Verrucomicrobiaceae” group, which is a tight hitherto- unnamed, group in the verrucomicrobia. We propose to coin this group “Candidatus genus Rhizospheria”. It is naturally included in the family Rubritaleaceae (class Verrucomicrobiae, phylum Verrucomicrobia). The name of the proposed group is based on the repeated isolation of the cultured representatives of this group from the leek rhizosphere.
Acknowledgements This research was part of the Ecogenomics program which is sponsored by the Dutch National Genomics Initiative and the basic research program on sustainable agriculture (KB4) sponsored by the Dutch ministry of agriculture, nature and food safety. We would like to thank Johnny Visser and field workers of ‘De Vredepeel’ experimental farm for their assistance in growth and sampling of leek plants. We also thank An Vos, Meint Veninga and Jaap Bloem for their help with total bacterial counts.
99 Isolation of Acidobacteria and Verrucomicrobia from leek
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Sanguin H, Remenant B, Dechesne A, Thioulouse J et al. (2006) Potential of a 16S rRNA-based taxonomic microarray for analyzing the rhizosphere effects of maize on Agrobacterium spp. and bacterial communities. Appl Environ Microbiol 72: 4302-4312. Sanguin H, Sarniguet A, Gazengel K, Moënne-Loccoz Y & Grundmann GL (2009) Rhizosphere bacterial communities associated with disease suppressiveness stages of take-all decline in wheat monoculture. New Phytol 184: 694-707. Scheuermayer M, Gulder TAM, Bringmann G & Hentschel U (2006) Rubritalea marina gen. nov., sp. nov., a marine representative of the phylum 'Verrucomicrobia', isolated from a sponge (Porifera). Int J Syst Evol Microbiol 56: 2119-2124. Schlesner H (1987) Verrucomicrobium spinosum gen. nov., sp. nov.: a fimbriated prosthecate bacterium. System Appl Microbiol 10: 54-56. Schlesner H, Jenkins C & Staley JT (2006) The Phylum Verrucomicrobia: A Phylogenetically Heterogeneous Bacterial Group. Prokaryotes 7: 881-896. Schmalenberger A & Tebbe CC (2003) Bacterial diversity in maize rhizospheres: Conclusions on the use of genetic profiles based on PCR-amplified partial small subunit rRNA genes in ecological studies. Mol Ecol 12: 251-261. Schoenborn L, Yates PS, Grinton BE, Hugenholtz P & Janssen PH (2004) Liquid serial dilution is inferior to solid media for isolation of cultures representative of the phylum-level diversity of soil bacteria. Appl Environ Microbiol 70: 4363-4366. Sharma S, Aneja MK, Mayer J, Munch JC & Schloter M (2005) Characterization of bacterial community structure in rhizosphere soil of grain legumes. Microb Ecol 49: 407-415. Smibert RM & Krieg NR (1981) General characterization. In: Gerhardt, P. (ed-in-chief) Manual of methods for general bacteriology, 3rd edn, American Society for Microbiology, Washington, USA, pp 409-443. Spain AM, Peacock AD, Istok JD, Elshahed MS, Najar FZ, Roe BA, White DC & Krumholz LR (2007) Identification and isolation of a Castellaniella species important during biostimulation of an acidic nitrate- and uranium-contaminated aquifer. Appl Environ Microbiol 73: 4892-4904. Stevenson BS, Eichorst SA, Wertz JT, Schmidt TM & Breznak JA (2004) New strategies for cultivation and detection of previously uncultured microbes. Appl Environ Microbiol 70: 4748-4755. Sun L, Qiu F, Zhang X, Dai X, Dong X & Song W (2008) Endophytic bacterial diversity in rice (Oryza sativa L.) roots estimated by 16S rDNA sequence analysis. Microb Ecol 55: 415-424. Valáková V, De Boer W, Klein Gunnewiek PJA, Pospíek M & Baldrian P (2009) Phylogenetic composition and properties of bacteria coexisting with the fungus Hypholoma fasciculare in decaying wood. ISME J 3: 1218-1221. , Jeppsson B & Molin G (2005) Comparison of bacterial diversity along the human intestinal tract by direct cloning and sequencing of 16S rRNA genes. FEMS Microbiol Ecol 54: 219-231. Wang M, Chen JK & Li B (2007) Characterization of bacterial community structure and diversity in rhizosphere soils of three plants in rapidly changing salt marshes using 16S rDNA. Pedosphere 17: 545- 556. Wimpenny JWT & Lewis MWA (1977) The growth and respiration of bacterial colonies. J Gen Microbiol 103: 9-18. Yoon J, Matsuo Y, Adachi K, Nozawa M, Matsuda S, Kasai H & Yokota A (2008) Description of Persicirhabdus sediminis gen. nov., sp. nov., Roseibacillus ishigakijimensis gen. nov., sp. nov., Roseibacillus ponti sp. nov., Roseibacillus persicicus sp. nov., Luteolibacter pohnpeiensis gen. nov., sp. nov. and Luteolibacter algae sp. nov., six marine members of the phylum 'Verrucomicrobia', and emended descriptions of the class Verrucomicrobiae, the order Verrucomicrobiales and the family Verrucomicrobiaceae. Int J Syst Evol Microbiol 58: 998-1007. Young KD (2007) Bacterial morphology: why have different shapes? Curr Opin Microbiol 10: 596-600. Zengler K (2009) Central role of the cell in microbial ecology. Microbiol Mol Biol Rev 73: 712-729. Zul D, Denzel S, Kotz A & Overmann J (2007) Effects of plant biomass, plant diversity, and water content on bacterial communities in soil lysimeters: Implications for the determinants of bacterial diversity. Appl Environ Microbiol 73: 6916-6929.
102 Chapter 5: Real-time PCR detection of Holophagae (Acidobacteria) and Verrucomicrobia subdivision 1 groups in bulk and leek (Allium porrum) rhizosphere soils*
Abstract
In the light of the poor culturability of Acidobacteria and Verrucomicrobia species, group-specific real-time (qPCR) systems were developed based on the 16S rRNA gene sequences from culturable representatives of both groups. The number of DNA targets from three different groups, i.e. Holophagae (Acidobacteria group 8) and Luteolibacter and Candidatus genus Rhizospheria (both from Verrucomicrobia subdivision 1), were determined in DNA extracts from different leek (Allium porrum) rhizosphere soil compartments and from bulk soil with the aim to determine the distribution of the three bacterial groups in the plant-soil ecosystem. The specificity of the designed primers was evaluated in three steps. First, in silico tests were performed which demonstrated that all designed primers 100% matched with database sequences of their respective groups, whereas lower matches with other non-target bacterial groups were found. Second, PCR amplification with the different primer sets was performed on genomic DNA extracts from target and from non-target bacteria. This test demonstrated specificity of the designed primers for the target groups, as single amplicons of expected sizes were found only for the target bacteria. Third, the qPCR systems were tested for specific amplifications from soil DNA extracts and 48 amplicons from each primer system were sequenced. All sequences were > 97% similar to database sequences of the respective target groups. Estimated cell numbers based on Holophagae-, Luteolibacter- and Candidatus genus Rhizospheria-specific qPCRs from leek rhizosphere compartments and bulk soils demonstrated higher preference for one or both rhizosphere compartments above bulk soil for all three bacterial groups.
* Authored by: Ulisses Nunes da Rocha, Jan Dirk van Elsas & Leo Simon van Overbeek Published in: J Microbiol Methods (accepted) Detection of Holophagae and Verrucomicrobia subdivision 1 in soil
Introduction
Acidobacteria and Verrucomicrobia are diverse bacterial phyla that are widely distributed and highly abundant in the soil environment (Hugenholtz et al., 1998; Barns et al., 1999; Lee & Cho, 2009). In spite of their high abundance and diversity, little information is available on their ecology, which is mainly due to the lack of culturable representatives in bacterial collections (Nunes da Rocha et al., 2009). Therefore, most of the ecological information about these taxa is based on the Acidobacteria and Verrucomicrobia 16S rRNA gene sequence distribution over different ecosystems. Analyses of 16S rRNA genes of Acidobacteria and Verrucomicrobia in samples from terrestrial habitats often revealed contradictory information about their preferred sites, especially in the distinction between bulk or rhizosphere soils (Chow et al., 2002; Sanguin et al., 2006; Zul et al., 2007; Kielak et al., 2008). The phylogenetic diversity of Acidobacteria and Verrucomicrobia is high. In fact, both are deep-branching groups within the bacterial ‘tree of life’. Twenty six different phylogenetic groups have been defined so far for Acidobacteria (Barns et al., 2007) and seven subdivisions for Verrucomicrobia (Schlesner et al., 2006). Most studies on the numeric distribution of Acidobacteria and Verrucomicrobia in natural ecosystems have been performed with primers or probes that cover multiple groups or subdivisions within both phyla. Minority groups present among both phyla may have been overlooked and their behavior may be contrary to what is currently assumed to be true about the distribution of Verrucomicrobia and Acidobacteria in plant-soil ecosystems. In spite of the fact that members of the Acidobacteria and Verrucomicrobia are considered to be ‘hard to culture’ (Jones et al., 2009; da Rocha et al., 2010), an increasing number of strains can currently be found in public databases. For instance, 209 16S rRNA gene sequences of culturable representatives of Acidobacteria next to 134 sequences of culturable Verrucomicrobia are currently available in the RDP Release 10 (update 19, March 31, 2010 - http://rdp.cme.msu.edu/index.jsp). The availability of culturable representatives of the two bacterial groups would allow the determination of metabolic, morphological and genetic parameters, which is impossible to accomplish with molecular techniques alone (Zengler, 2009). Further, such cultures would allow experimentation under controlled conditions mimicking the situation in the environment. The vast majority of these strains likely consists of slow growers, which require specific conditions that favor their growth in pure culture (Nunes da Rocha et al., 2009). Because of their recalcitrance to growth under laboratory conditions, combinations of culture-dependent and -independent approaches will further improve detection at finer resolution levels. This is important to allow distinction between
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separate groups within both phyla, something required to gain understanding about the ecology of particular representatives within both groups. Recently, two close related culturable representatives of the Holophagae, previously known as Acidobacteria group 8, were found in the leek rhizosphere at two independent occasions (Nunes da Rocha et al., 2010). Also, nine culturable members of three phylogenetically distinguishable groups of subdivision 1 of the Verrucomicrobia were isolated from leek and potato rhizospheres, sampled at different occasions (da Rocha et al., 2010; Nunes da Rocha et al., 2010). To the best of our knowledge, no information about their presence in soil compartments proximate to plants has been reported in literature (Nunes da Rocha et al., 2009). As these cultured Holophagae and Verrucomicrobia were recovered from soil adhering to roots (rhizosphere), it was hypothesized that both groups might show preferences for rhizosphere over bulk soil. To test this hypothesis, bulk and rhizosphere soil samples taken from field- grown leek plants were analyzed for the presence of Holophagae and Verrucomicrobia subdivision 1 groups. Analyses were done with newly developed group-specific real- time PCR (qPCR) systems, based on the 16S rRNA gene sequences of culturable representatives of Holophagae and Verrucomcirobia subdivision 1. Three systems (one from the Holophagae and two from Verrucomicrobia subdivision 1) were evaluated and tested for their functionality in the detection of both groups in the plant-soil ecosystem.
Material and Methods
Identity and growth of bacterial strains
Isolation and identification of strains belonging to the Holophagae (2), Luteolibacter (3) and candidatus Rhizospheria (5) are described in da Rocha et al. (2010) and Nunes da Rocha et al. (2010) (Table 1). The taxonomic relationship, based on the16S rRNA gene sequences of these strains and uncultured members of the Acidobacteria and Verrucomicrobia (SILVA database - Pruesse et al., 2007) is depicted in Fig. 1 of chapter 4 (page 111). R2A (Difco, France) was used for routine cultivation of all strains at 25o C.
DNA extraction from pure cultures and soils
For DNA extraction from all pure culture strains (Table 1), cells were scraped off from R2A agar plates with bacterial growth. The cells were then suspended in 500 µL of a sterile solution consisting of 0.85 % KCl in DNase/Rnase-free distilled water (Invitrogen, The Netherlands). DNA was extracted from the resulting cell suspensions
105 Detection of Holophagae and Verrucomicrobia subdivision 1 in soil
) rhizospheres. Reference (2010) al. et da Rocha Nunes (2010) al. et da Rocha Nunes et al. (2010) da Rocha (2010) al. et da Rocha Nunes (2010) al. et da Rocha Nunes et al. (2010) da Rocha et al. (2010) da Rocha et al. (2010) da Rocha (2010) al. et da Rocha Nunes (2010) al. et da Rocha Nunes (2010) al. et da Rocha Nunes Allium porrum ) or leek ( on Field/collection
2008 Vredepeel Vredepeel 2008 Vredepeel 2007 Vredepeel 2007 Droevendaal 2006 Droevendaal 2006 Vredepeel 2007 Vredepeel 2008 Year of isolati Year of 2007 Vredepeel Vredepeel 2007 Droevendaal 2006 Droevendaal 2006 Vredepeel 2007 Solanum tuberosum
Allium porrum Allium porrum Allium porrum tuberosum Solanum tuberosum Solanum Allium porrum Allium porrum Allium porrum tuberosum Solanum tuberosum Solanum Allium porrum isolated from potato (
strains
ORAC CHC12 ONA9 Z35 ZNBB5 CHC8 IRVE CHC25 C20 CNC16 CR28 Verrucomicrobia and Rhizospheria Rhizospheria genus Acidobacteria Holophagae Holophagae Luteolibacter Candidatus Verrucomicrobium Affiliation Strain Plant type Plant 1 Table Affiliation Strain Acidobacteria Verrucomicrobia
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using the MasterPureTM DNA purification kit (Epicentre Biotechnologies, WI, USA) following the instructions provided by the manufacturer. For extraction from soil, the PowerSoil Isolation Kit (MO BIO Laboratories, Inc., CA, USA) was used, following the instructions provided by the manufacturer.
Design and testing of Holophagae-, Luteolibacter- and Candidatus genus Rhizospheria- specific primers
Because the Holophaga (2), Luteolibacter (3) and Candidatus genus Rhizospheria (5) strains all formed separate clusters in the dendrogram (Fig 1), we decided to design three primer systems, one for each group. 16S rRNA gene sequences of these strains and of Holophagae (128 sequences), Luteolibacter (121 of Luteolibacter and 22 of Prosthecobacter) and Candidatus genus Rhizospheria (180 sequences) from the RDP Release 10, Update 19 (http://rdp.cme.msu.edu/) were separately aligned for each group using SINA Webaligner (http://www.arb-silva.de/aligner/). The alignments were checked for similarities in conserved regions among the sequences within each of the three groups using ARB software (Ludwig et al., 2004). Then, primers were designed on the basis of the conserved regions and these were checked for in silico specificity using Primer-BLAST software (http://www.ncbi.nlm.nih.gov/tools/primer-blast/). This recent software, last modified December 2009, enables the search for primer pairs that are specific for the intended PCR template and also allows to check for eventual occurrences of misprimed products and possible non-intended templates (http://www.ncbi.nlm.nih.gov/tools/primer- blast/primerinfo.html). The melting temperature of the primers was restricted to values between 59 and 61oC, to closely match the Taq polymerase optimal extension temperature of SYBR® Premix Ex Taq TM (TAKARA Bio Inc., Japan). The sequence, melting behavior and most relevant PCR amplification parameters of the designed primer systems are presented in Table 2. Luteolibacter and Prostecobacter are closely related genera of Verrucomicrobia subdivision 1, and only recently taxonomical distinction between both groups was made (Yoon et al., 2008). Although in silico analysis indicated Luteolibacter-specific amplification when using primers VS1Af and VS1Ar, sequencing of clones generated with this primer pair demonstrated that both Luteolibacter and Prosthecobacter members are actually amplified from soil DNA. The bacterial group distinguished by this specific qPCR system is therefore denoted as Luteolibacter. The primers were further tested for their specificity using target (Table 1) and non-target DNA by PCR amplification. Non-target DNA from the following strains were included in the evaluation assays of the qPCR systems: Agroabcterium tumefaciens strain UBAPF2 (Alphaproteobacteria), Burkholderia cepacia strain LMG 1222T
107 Detection of Holophagae and Verrucomicrobia subdivision 1 in soil
efficiency Amplification
c range Dynamic late concentrations in the soil ) b Amplicon Amplicon length (bp
a C) o ( Tm 59.19 59.19 60.41 470 60.04 59.98 7.54 2.54 to 199 59.96 60.06 2.01 8.26 2.26 to 83 1.98 8.45 2.45 to 1.95 name Acg8f Acg8r Primer Primer calculated efficiency, ; slope, slope amplification Ae, from where, VS1Bf VS1Br VS1Af VS1Ar (-1/slope) subdivision 1 qPCR primers. primers. subdivision 1 qPCR r gram of dry of soil),r gram which indicates the initial of range temp Verrucomicrobia and Holophagae Forward TGGGATGTTGATGGTGAAACForward Forward CAGCTCGTGTCGTGAGATGT Forward GCCCGACAGGGTTGATAGTAForward
genus e group-specific Candidatus Rhizospheria Description of th bp, base pairs pairs bp, base d of the reaction The efficiency calculated by equation: was Ae = 10 the following Tm, melting temperature melting Tm, Theoretical range (expressed dynamic in, Log cell equivalents pe
standard curve curve standard 2 Table a b c d DNA extracts over which reliable Ct values are obtained, assuming DNA extraction and qPCR reaction efficiencies of 100% 100% of efficiencies reaction and qPCR extraction DNA assuming are obtained, values Ct reliable over which extracts DNA Phylum Phylum AGTCTCGGATGCAGTTCCTG Holophagae Acidobacteria group Target Reverse Sense – 3’) (5’ sequence Primer Reverse TCTCGGTTCTCATTGTGCTG TCTCGGTTCTCATTGTGCTG CGCTTGGGACCTTCGTATTA Luteolibacter Verrucomicrobia Reverse Reverse
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(Betaproteobacteria), Escherichia coli strain E1 (Gammaproteobacteria), Streptomyces griseus strain IPO 857 (Actinobacteria), Flavobacterium columnar strain 2003/035 (Bacteroidetes) and Bacillus subtilis strain Bs4 (Firmicutes); all derived from the strain collection available at Plant Research International (Wageningen, The Netherlands). Each 25-µl reaction mixture contained the following ingredients: 12.5 µl of SYBR Premix Ex Taq 2x (TAKARA Bio Inc., Japan), 0.5 µl of each primer (10 µM; Biolegio, NL), 0.5 µl of ROX Reference
Dye II 50x (TAKARA Bio Inc., Japan), 6.0 µl H2O and 5.0 µl template DNA (containing 1ng of target or non-corresponding strains). All PCR reactions were run for one cycle at 50oC for 2 min; one cycle at 95oC for 10 s; 35 cycles at 95oC for 5 s and 60oC for 35 s. The number of cycles was limited to 35 to avoid occurrences of false positive signals (Sipos et al., 2007). Additionally, melting curves derived from each individual reaction were inspected in order to ascertain that the signals obtained originated from specific PCR reactions and not from primer dimer formation or any other artifact. Further, PCR amplicons made with all three primer systems with DNA from target and non-corresponding strains were checked in 1.5% agarose gels for presence of bands of expected sizes and absence of any secondary, or false-positve products. As a final check on the specificity of the selected primer systems, amplification from soil DNA extracts (Vredepeel [V] soil) was performed. Thus, bulk soil DNA was added as template to PCR reaction mixtures (1 – 5 ng per reaction). Then, reactions were run under the same conditions as described before. Subsequently, PCR products were purified using the QIAquick PCR purification kit (Qiagen, Hilden, GE) and the resulting products were cloned into the pGEM-T Easy Vector (Promega, WI, USA) using the protocol provided by the manufacturer. Totals of 48 clones from each primer system were selected for sequencing. All clones were checked for the presence of inserts of the correct sizes before sequencing. The resulting sequences were aligned using MEGA 4 software (Kumar et al. 2008) and then individually compared with database sequences by BlastN-assisted searches (http://blast.ncbi.nlm.nih.gov/Blast.cgi).
Calibration of real time PCR (qPCR) systems.
The three qPCR systems, aimed to detect Holophagae, Luteolibacter and Candidatus genus Rhizospheria, were calibrated on cell lysates made from pure culture strains, one from each group. For each reaction (performed as described above), 5 µl of tenfold serial dilutions (ranging from approximately 102 to approximately 1010 cells per mL) per target strain per group-specific qPCR system was used. Holophagae strain CHC25, Luteolibacter strain CHC12 and Candidatus genus Rhizospheria strain CHC8 were used to yield DNA templates for the respective group-specific qPCR systems. Standard curves were made in separate for each series of serially-diluted suspensions and all
109 Detection of Holophagae and Verrucomicrobia subdivision 1 in soil
dilutions were made in triplicate. Group-specific qPCR systems were employed on these series and the measured threshold cycle (Ct) values were plotted against the Log cell number for each reaction. Line slopes and intercepts from resulting graphs were calculated by regression analysis using GenStat 12th edition (VSN International Ltd., UK). The amplification efficiency (Ae) of the different primer systems was calculated using the formula Ae = 10(-1/slope), in which the slope represents the slope value calculated by regression analysis.
Leek plant growth and root and soil sampling procedures
The site chosen for sampling was an agricultural field located at the experimental farm ‘De Vredepeel’, The Netherlands (51o 32’ 27.10” N and 5o 51’ 14.86” E). Here, leek was grown in accordance with practices common for leek production, in particular concerning soil tillage, crop rotation, chemical fertilization and chemical pest (thrips) control. The V soil was characterized as sand, with pH 5.4 and 2.2% organic matter. Roots with adhering soil of seven leek (Allium porrum) plants (cultivar Kenton, Nunhems Zaden BV, The Netherlands) were collected, all at the same occasion, but at different sites in the fields (minimum distance between plants was 5 m). Three soil compartments were distinguished: bulk soil, outer rhizosphere and inner rhizosphere. The soil free of roots in the neighborhood of the sampled plants (at least at 1 m distance from each plant) was considered as bulk soil. The soil adhering to leek roots was separated into two fractions: the outer rhizosphere was represented by the soil adhering to the root surface after mild manual shaking. It was removed from the roots by scratching with a spatula. Then, roots devoid of the outer rhizosphere were shaken in a 1:10 ratio (v/v basis) in 0.1% sodium pyrophosphate solution. The resulting soil suspension was considered to represent the inner rhizosphere. The inner rhizosphere suspension was concentrated by centrifugation at 10,000 x g for 15 min followed by resuspension of the resulting pellet in 1 ml of 0.85% NaCl solution. qPCR detection of Holophagae, Luteolibacter and Candidatus genus Rhizospheria in different plant-soil compartments qPCR was conducted with the three selected primer systems using DNA from different soil compartments (bulk, outer and inner rhizosphere soils) of the seven individual leek plants. Three reaction mixtures per sample, each containing 5 ng of soil DNA , were subjected to either one of the qPCRs for detection of Holophagae, Luteolibacter and Candidatus genus Rhizospheria, as described above. One positive control (containing DNA from a corresponding strain as a template) and seven negative controls, including DNA from six non-corresponding strains (Table 1) and one with sterile demineralized
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water only, were used in each run. Differences between the average Ct values per bacterial group in each soil compartment were statistically compared by one-way analysis of variance (ANOVA) and differences were considered to be significant at levels of P < 0.05. Dynamic range here is considered as the range over which cell numbers can be reliably measured by qPCR. A theoretical dynamic range, using the extreme values of equivalent cell number determined by the calibration curves, was estimated for each -1 -1 primer system using the formula: Ev = eqcR × VPCR × VR × WS ; where, Ev is the estimated highest or lowest dynamic range value, eqcR is the equivalent cell number in the reaction mixture calculated from equations made by regression analysis, VPCR is the volume of DNA solution added to the qPCR reaction PL[WXUH ȝO 9R is the volume in
ZKLFK WKH '1$ SHOOHW ZDV UHVXVSHQGHG ȝO :S is the precise amount of soil, calculated on dry weight basis, that was used for DNA extraction.
Nucleotide sequence accession numbers Non-redundant sequences of the 144 partial 16S rRNA gene library clones constructed with the three primer systems (48 clones each) were deposited in the EMBL Nucleotide Sequence Database and are available under the accession numbers FN796785 to FN796791 and FN796793 to FN796795.
Results
Specificity of the Holophagae, Luteolibacter and Candidatus genus Rhizospheria primer systems
Three quantitative PCR primer systems aimed to specifically amplify 16S rRNA gene sequences of Holophagae, Luteolibacter and Candidatus genus Rhizospheria were designed (Table 2) and evaluated for quantification of members of these groups in soil. In silico comparisons of the sequences of individual primers with database sequences, using Primer-Blast, predicted that all primers (Holophagae, Luteolibacter and Candidatus genus Rhizospheria) would amplify 16S rRNA gene sequences that matched the sequences of their respective groups, yielding amplicons with expected sizes. Further, no matches with other bacterial groups were expected to be found. PCR amplifications performed with genomic DNA of the two Holophagae strains, CHC25 and ORAC, using the primer system specific for the Holophagae resulted in the generation of single amplicons of the expected size of 470 bp, in the absence of any visible primer dimer or other products (data not shown). PCR amplification with this primer system, using genomic DNA from six non-corresponding
111 Detection of Holophagae and Verrucomicrobia subdivision 1 in soil
strains as well as all nine Verrucomicrobia strains (Table 1) resulted in the absence of any band under these amplification conditions. Using the primers designed for Luteolibacter in PCR reactions with genomic DNA of Luteolibacter strains CHC12, C20 and ONA9 showed the emergence of single amplicons of the expected size, of199 bp in the absence of any primer dimer or other product after agarose gel electrophoresis. No PCR product was found from PCRs run with the same primers on genomic DNAs from six non-corresponding, five Candidatus genus Rhizospheria, two Holophagae and one Verrucomicrobium strains as templates (Table 1). Using the primers designed for the Candidatus genus Rhizospheria group with DNA from the five Candidatus genus Rhizospheria strains CHC8, IRVE, CR28, Z35 and ZNBB5 revealed the presence of single amplicons of the expected size of 83 bp in the absence of any primer dimer or other product in agarose gel. This band was absent when using genomic DNA from six non-corresponding strains and from all strains of Luteolibacter, Verrucomicrobium and Holophagae (Table 1) using the same PCR system and run under the same circumstances. DNA sequence analyses of library clones constructed with the three primer systems (48 clones per system), using V soil DNA as template, always revealed the presence of insert sequences that matched with database sequences belonging to the expected groups (Table 3). DNA sequence analysis of the inserts of the clones made with Holophagae-specific primers resulted in the detection of two sequence groups, i.e. those denoted clone lib1_1_36 (encompassing 36 clones) and those denoted lib1_2_12 (encompassing 12 clones). Both groups were closest affiliated with Acidobacterium bacterium CHC25, albeit at different similarity levels, i.e. 97% for lib1_1_36 and 99% for lib1_2_12 (Table 3). Sequencing of the inserts of the 48 Luteolibacter library clones resulted in five different sequence groups; those denoted as lib2_3_16 (16 clones), lib2_5_15 (15) lib2_1_7 (7), lib2_4_8 (8) and lib2_2_2 (2). DNA sequences belonging to lib2_1_7 and lib2_2_2 showed closest matches with those of Prosthecobacter species (99% similarity) and those of lib2_3_16, lib2_5_15, lib2_4_7 groups matched closely with sequences of Luteolibacter species (> 98% similarity). Sequencing of the inserts of the 48 Candidatus genus Rhizospheria library clones resulted in three sequence groups, denoted as lib3_1_33 (33 clones), lib3_2_10 (10) and lib3_3_5 (5). All showed closest matches to different sequences belonging to the Candidatus genus Rhizospheria (100% similarity). The primers designed within this study thus specifically targeted DNA sequences from Holophagae, Luteolibacter and Candidatus genus Rhizospheria species, both in pure culture as well as in complex soil DNA extracts.
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d efficiency Amplification
c range Dynamic late concentrations in the soil ) b Amplicon Amplicon length (bp
a C) o ( Tm 59.19 59.19 60.41 470 60.04 59.98 7.54 2.54 to 199 59.96 60.06 2.01 8.26 2.26 to 83 1.98 8.45 2.45 to 1.95 name Acg8f Acg8r Primer Primer calculated efficiency, ; slope, slope amplification Ae, from where, VS1Bf VS1Br VS1Af VS1Ar (-1/slope) subdivision 1 qPCR primers. primers. subdivision 1 qPCR r gram of dry of soil),r gram which indicates the initial of range temp Verrucomicrobia and Holophagae Forward TGGGATGTTGATGGTGAAACForward Forward CAGCTCGTGTCGTGAGATGT Forward GCCCGACAGGGTTGATAGTAForward
genus e group-specific Candidatus Rhizospheria Description of th bp, base pairs pairs bp, base
of the reaction The efficiency calculatedwas by equation: Ae = 10 the following Tm, melting temperature melting Tm, Theoretical range (expressed dynamic in, Log cell equivalents pe DNA extracts over which reliable Ct values are obtained, assuming DNA extraction and qPCR reaction efficiencies of 100% 100% of efficiencies reaction and qPCR extraction DNA assuming are obtained, values Ct reliable over which extracts DNA standard curve curve standard a b c d 2 Table Phylum Phylum AGTCTCGGATGCAGTTCCTG Holophagae Acidobacteria group Target Reverse Sense – 3’) (5’ sequence Primer Reverse TCTCGGTTCTCATTGTGCTG TCTCGGTTCTCATTGTGCTG CGCTTGGGACCTTCGTATTA Luteolibacter Verrucomicrobia Reverse Reverse
113 Detection of Holophagae and Verrucomicrobia subdivision 1 in soil
Amplification efficiency and theoretical dynamic range of qPCR systems that detect the Holophagae, Luteolibacter and Candidatus genus Rhizospheria
Standard curves of all three qPCR systems, constructed by plotting measured Ct values using cell lysates made from Acidobacterium bacterium strain CHC25 (Holophagae), Verrucomicrobiaceae bacterium strain CHC12 (Luteolibacter) and Verrucomicrobiaceae bacterium strain CHC8 (Candidatus genus Rhizospheria) against the respective log cell numbers revealed linear relationships between the two parameters for all three systems. For the Holophagae qPCR system, the slope of the curve, as calculated by regression analysis, was -3.296 (R2 value 0.9963) and the amplification efficiency (Ae) value was 2.01. For the Luteolibacter qPCR system, the slope of the curve was -3.375 (R2 value 0.9945) and the Ae value 1.98. For the Candidatus genus Rhizospheria qPCR system, the slope of the curve was -3.459 (R2 value 0.9863) and the Ae value 1.95. The theoretical dynamic range, in Log equivalent cell number per gram of dry soil, of the different qPCR systems were 2.54 - 7.54 for the Holophagae qPCR system, 2.26 - 8.26 for the Luteolibacter qPCR system and 2.45 - 8.45 for the Candidatus genus Rhizospheria qPCR system. qPCR enumeration of Holophagae, Luteolibacter and Candidatus genus Rhizospheria in bulk soil and in the outer and inner rhizosphere of leek plants
The distribution, in Log equivalent cell number per g of dry soil, of Holophagae, Luteolibacter and Candidatus genus Rhizospheria in three soil compartments, i.e. bulk soil and outer and inner rhizosphere of leek plants, is shown in Fig. 1. For the Holophagae, the highest numbers were found in the outer rhizosphere (6.25 - 6.41), followed by the bulk soil (6.08 - 6.20) and the inner rhizosphere (5.89 - 5.85). This stood in sharp contrast to the distribution of Luteolibacter, where the highest numbers were found in the inner rhizosphere (6.24 - 6.46), followed by the outer rhizosphere (5.91 - 6.21) and the bulk soil (4.11 - 4.39). For the Candidatus genus Rhizospheria, the highest numbers were found in both rhizosphere compartments: 5.33 - 5.39 in the outer rhizosphere, 5.25 - 5.35 in the inner rhizosphere and 4.31 - 4.48 in the bulk soil. Real-time PCR analysis of the natural V soil thus demonstrated higher numbers in log cell equivalents of Holophagae, Luteolibacter and Candidatus genus Rhizospheria in one or both leek rhizosphere compartments as compared to bulk soil.
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Bulk Soil Outer rhizosphere Inner rhizosphere 7
A A B B C 6 A A
5
B C
4 Log of equivalent cell number per g of dry soil dry equivalent g of Log of per cell number Holophagae Luteolibacter Candidatus genus Rhizospheria
Figure 1 Holophagae, Luteolibacter and Candidatus genus Rhizospheria cell estimates in bulk, outer rhizosphere and inner rhizosphere soils. Numbers were generateded by Holophagae-, Luteolibacter- and Candidatus genus Rhizospheria-specific qPCRs. Averages followed by different letters significantly differ from each other, as determined by one-way ANOVA (P<0.05).
Discussion
Although representatives of the Acidobacteria and Verrucomicrobia are known for their recalcitrance to grow in pure culture, members of both taxa can be cultured from different environments. Indeed, several reports on successful isolation have appeared, being the work of da Rocha et al. (2010) and Nunes da Rocha et al. (2010), the first reports on recovery of Verrucomicrobia and Acidobacteria members from rhizosphere. Because of these findings, it was hypothesized that members of these groups might prefer the rhizosphere over bulk soil. To test this hypothesis, it was necessary to design a culture-independent quantitative method to determine cell equivalent numbers of the groups in different plant-soil compartments. This would allow an estimation of the cell numbers of the distinctive groups given that cultivation from soil still is cumbersome due to the specific requirements for growth and the long incubation times needed. Here, specific qPCR systems for each of the
115 Detection of Holophagae and Verrucomicrobia subdivision 1 in soil
three groups (Holophagae, Luteolibacter and Candidatus genus Rhizospheria) are presented. These were all designed on the basis of the 16S rRNA gene and evaluated for their specificity of detection in soil. The new qPCR systems will help us to explore the members of the three groups in the plant-soil ecosystem. To the best of our knowledge, the approach to develop such qPCR-based methods, validated on the basis of actual cell numbers, have never been developed before for the groups of Acidobacteria and Verrucomicrobia. Two strains of the Holophagae (previously known as Acidobacteria group 8) were recovered at two different occasions from the leek rhizosphere (Nunes da Rocha et al., 2010). The Holophagae can be considered as a minority group in soil ecosystems, as they may comprise only up to 3.4% of total Acidobacteria sequences, as determined by pyrosequencing (Jones et al., 2009). More importantly, group 8 of Acidobacteria will not be included in Acidobacteria cell measurements in environmental samples when using primer Acd31F (Barns et al., 1999). Namely, it was recently reported that Holophagae- specific 16S rRNA sequences could not be amplified from soil with this primer (Kielak et al., 2009). The Verrucomicrobia subdivision 1 strains, obtained from leek and potato rhizospheres at different occasions, were affiliated with two distinct groups in this subdivision, i.e. the Luteolibacter and the Candidatus genus Rhizospheria groups (da Rocha et al., 2010; Nunes da Rocha et al., 2010). Species belonging to Luteolibacter, i.e. Luteolibacter algae and Luteolibacter pohnpeiensis, were so far all only isolated from marine environments, namely from red algae and driftwood, respectively (Yoon et al., 2008). Another Verrucomicrobia subdivision 1 strain, initially identified only at the phylum level, also originated from seawater (Stingl et al., 2007). Later database comparisons revealed this strain to belong to the so called unclassified Verrucomicrobiaceae group of subdivision 1 of Verrucomicrobia and later the name of this group was coined to Candidatus genus Rhizospheria in Nunes da Rocha et al. (2010). The isolation of subdivision 1 Verrucomicrobia is therefore not new, but isolation from the plant-soil ecosystem is. It is still unknown whether the two groups are minority groups within the Verrucomicrobia. So far, only one report on the detection of Verrucomicrobia subdivision 1 was made in rhizospheres (Haichar et al., 2008), and hence we surmised that also this group may have been overlooked in the rhizosphere so far. Quantitative PCR is currently a widely accepted approach in environmental microbiology. It is used to quantify bacterial gene or transcript numbers in environmental samples (Cardenas & Tiedje, 2008). Developing specific qPCR primers for detection in complex habitats like soil is a challenging endeavor because of the high bacterial diversity
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in soil (van Elsas et al., 2008). Therefore, we opted for the validation of the designed qPCR primer systems in respect of their specificity in three steps, i.e. an in silico step followed by PCR on target and non-target DNA templates and by analysis of clone libraries generated with DNA from the same V soil. Further, specificity was improved by careful selection of melting temperature (around 60o C), annealing time (short, maximally 5 sec) and number of PCR cycles (relatively low, i.e. 35), in accordance with Edwards (2004) and Sipos et al. (2007). Our approach resulted in three sets of primers that were specific for the detection and quantification of the target Holophagae, Luteolibacte/Prosthecobacter and Candidatus genus Rhizospheria groups in soil. Microarray- and qPCR-assisted approaches for the detection and quantification of Acidobacteria and Verrucomicrobia in soil and thermal springs have been reported and all of these approaches have their merits and caveats (Hall et al., 2008; Kuramae et al., 2010; Liles et al., 2010). Our approach is different from these approaches because we validated the quantification systems using cells obtained from our cultured representatives of the three groups. This approach has not been done before with Acidobacteria and Verrucomicrobia. Our endeavor thus yielded a set of dedicated qPCR systems that quantify the cell numbers of three target groups of organisms in the plant-soil system. The standard curves produced and the calculated theoretical dynamic ranges of all three qPCR systems revealed that cell numbers could be assessed over a broad spectrum (grossly, between Log 2 and Log 8 cell equivalents per g of dry soil) in soil. Quantification in soil was highly robust as demonstrated by the high R2 values (over 0.980) for all three systems. The amplification efficiencies were all above 1.9, values that are acceptable for Sybr green detection in qPCR (Ruijter et al., 2009). The estimated limit of detection was between 100 and 500 cells per g of dry soil for all three qPCR systems, amounts that allow reliability in studies on the ecology of the three groups in the plant-soil ecosystem. An important further outcome of this study was that the three groups, Holophagae, Luteolibacter and Candidatus genus Rhizospheria, revealed to be more abundant in one or even both rhizosphere compartments than in the corresponding bulk soil. Moreover, the distinction between outer and inner rhizosphere made it possible to demonstrate that the numbers of both Verrucomicrobia groups became higher when coming into closer contact with the leek roots. For the Holophagae, this was different: although higher numbers were found in the outer rhizosphere than in bulk soil, the numbers were again lower in the inner rhizosphere. Supposedly, Holophagae cells do not thrive in spheres very proximate to roots or on the root surface, which may be the case for the Verrucomicrobia groups. One can only speculate about the driver of the preference for the outer rhizosphere in Holophagae cells. For instance, these cells may utilize some root-released compounds, but may be
117 Detection of Holophagae and Verrucomicrobia subdivision 1 in soil
unable to compete with the root-associated bacteria typically occurring at root surfaces. We conclude that the members of the Holophagae, Luteolibacter and Candidatus genus Rhizospheria groups detected by our PCR systems appear to thrive in the rhizosphere. These groups may tentatively be considered ‘rhizosphere competent’ bacteria, as is the case for other well-known plant-associated bacteria like the fluorescent pseudomonads. Our data corroborate those of previous studies, such as those done in the rhizosphere of lodgepole pine (Chow et al., 2002) and different other plant communities (Zul et al., 2007). These studies, without specifying the subgroups, also demonstrated that Acidobacteria and Verrucomicrobia are numerically more abundant in the rhizosphere than in corresponding bulk soil. However, in this study we distinguish between bacterial groups at lower taxonomical resolution levels and demonstrated that particular groups of Acidobacteria and Verrucomicrobia may be considered as ‘rhizosphere competent’.
Acknowledgements This research was part of the Ecogenomics program which is sponsored by the Dutch National Genomics Initiative and the basic research program on sustainable agriculture (KB4) sponsored by the Dutch ministry of agriculture, nature and food safety. We would like to thank Johnny Visser and field workers of ‘De Vredepeel’ experimental farm for their assistance in growth and sampling of leek plants.
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References
Barns, S.M., Cain, E.C., Sommerville, L., Kuske C.R., 2007. Acidobacteria phylum sequences in uranium- contaminated subsurface sediments greatly expand the known diversity within the phylum. Appl. Environ. Microbiol. 73, 3113-3116. Barns, S.M., Takala, S.L., Kuske, C.R., 1999. Wide distribution and diversity of members of the bacterial kingdom Acidobacterium in the environment. Appl. Environm. Microbiol. 65, 1731-1737. Cardenas, E., Tiedje, J.M., 2008. New tools for discovering and characterizing microbial diversity. Curr. Opin. Biotechnol. 19, 544-549. Chow, M.L., Radomski, C.C., McDermott J.M., Davies, J., Axelrood, P.E., 2002. Molecular characterization of bacterial diversity in Lodgepole pine (Pinus contorta) rhizosphere soils from British Columbia forest soils differing in disturbance and geographic source. FEMS Microbiol. Ecol. 42, 347-357. da Rocha, U.N., Andreote, F.D., de Azevedo, J.L., van Elsas, J.D., van Overbeek, L.S., 2010. Cultivation of hitherto-uncultured bacteria belonging to the Verrucomicrobia subdivision 1 from the potato (Solanum tuberosum L.) rhizosphere. J. Soils Sediments 10, 326-339. Edwards, K.J., 2004. Performing real-time PCR. In: Edwards, K., Logan, J., Saunders, N. (Eds.), Real-time PCR: an Essential Guide. Horizon Bioscience, Norfolk, pp. 71–83. Haichar, F.E.Z., Marol, C., Berge, O., Rangel-Castro, J.I., Prosser, J.I., Balesdent, J., Heulin, T., Achouak, W., 2008. Plant host habitat and root exudates shape soil bacterial community structure. ISME J. 2, 1221-1230. Hall, J.R., Mitchell, K.R., Jackson-Weaver, O., Kooser, A.S., Cron, B.R., Crossey, L.J., Takacs-Vesbach, C.D., 2008. Molecular characterization of the diversity and distribution of a thermal spring microbial community by using rRNA and metabolic genes. Appl. Environ. Microbiol. 74, 4910-4922. Hugenholtz, P., Goebel, B.M., Pace, N.R., 1998. Impact of culture-independent studies on the emerging phylogenetic view of bacterial diversity. J. Bacteriol. 180, 4765-4774. Jones, R.T., Robeson, M.S., Lauber, C.L., Hamady, M., Knight, R., Fierer, N., 2009. A comprehensive survey of soil acidobacterial diversity using pyrosequencing and clone library analyses. ISME J. 3, 442-453. Kielak, A., Pijl, A.S., Van Veen, J.A., Kowalchuk, G.A., 2008. Differences in vegetation composition and plant species identity lead to only minor changes in soil-borne microbial communities in a former arable field. FEMS Microbiol. Ecol. 63, 372-382. Kielak, A., Pijl, A.S., Van Veen, J.A., Kowalchuk, G.A., 2009. Phylogenetic diversity of Acidobacteria in a former agricultural soil. ISME J. 3, 378-382. Kumar, S., Ne, M., Dudley, J. Tamura, K., 2008. MEGA: A biologist-centric software for evolutionary analysis of DNA and protein sequences. Brief. Bioinform. 9, 299-306. Kuramae, E.E., Gamper, H.A., Yergeau, E., Piceno, Y.M., Brodie, E.L. et al., 2010. Microbial secondary succession in a chronosequence of chalk grasslands. ISME J. 4, 711-715. Lee, S.-H., Cho, J.-C.,2009. Distribution patterns of the members of phylum Acidobacteria in global soil samples. J. Microbiol. Biotechnol. 19, 1281-1287. Liles, M.R., Turkmen, O., Manske, B.F., Zhang, M., Rouillard, J.-M. et al., (2010) A phylogenetic microarray targeting 16S rRNA genes from the bacterial division Acidobacteria reveals a lineage-specific distribution in a soil clay fraction. Soil. Biol. Biochem. 42, 739-747. Ludwig, W., Strunk, O., Westram, R., Richter, L., Meier, H., Yadhukumar, A. et al., 2004. ARB: a software environment for sequence data. Nucleic Acids Res. 32, 1363-1371. Nunes da Rocha, U., Andreote, F.D., Plugge, C., van Elsas, J.D., van Overbeek, L.S., 2010. Isolation of culturable Holophagae (Acidobacteria), Luteolibacter, unclassified Verrucomicrobiaceae and Verrucomicrobium (Verrucomicrobia) from the Allium porrum rhizosphere. Submitted. Nunes Da Rocha, U., Van Overbeek, L., Van Elsas, J.D., 2009. Exploration of hitherto-uncultured bacteria from the rhizosphere. FEMS Microbiol. Ecol. 69, 313-328. Pruesse, E., Quast, C., Knittel, K., Fuchs, B.M., Ludwig, W., Peplies, J., Glockner, F.O., 2007. SILVA: a comprehensive online resource for quality checked and aligned ribosomal RNA sequence data compatible with ARB. Nucleic Acids Res. 35, 7188-7196.
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Ruijter, J.M., Ramakers, C., Hoogaars, W.M.H., Karlen, Y., Bakker, O., van den Hoff, M.J.B., Moorman, A.F.M., 2009. Amplification efficiency: Linking baseline and bias in the analysis of quantitative PCR data. Nucleic Acids Res. 37, e45. Sanguin, H., Remenant, B., Dechesne, A., Thioulouse, J. et al., 2006, Potential of a 16S rRNA-based taxonomic microarray for analyzing the rhizosphere effects of maize on Agrobacterium spp. and bacterial communities. Appl. Environ. Microbiol. 72, 4302-4312. Schlesner, H., Jenkins, C. Staley, J.T., 2006, The Phylum Verrucomicrobia: A Phylogenetically Heterogeneous Bacterial Group. Prokaryotes 7, 881-896. , K., Nikolausz, M., 2007, Effect of primer mismatch, annealing temperature and PCR cycle number on 16S rRNA gene-targetting bacterial community analysis. FEMS Microbiol. Ecol, 60, 341-350. Stingl, U., Tripp, H.J., Giovannoni, S.J., 2007, Improvements of high-throughput culturing yielded novel SAR11 strains and other abundant marine bacteria from the Oregon coast and the Bermuda Atlantic Time Series study site. ISME J. 1, 361-371. van Elsas, J.D., Costa, R., Jansson, J., Sjöling, S., Bailey, M., Nalin, R., Vogel, T.M., van Overbeek, L., 2008, The metagenomics of disease-suppressive soils - experiences from the METACONTROL project. Trends Biotechnol. 26, 591-601. Yoon, J., Matsuo, Y., Adachi, K., Nozawa, M., Matsuda, S., Kasai, H., Yokota, A., 2008. Description of Persicirhabdus sediminis gen. nov., sp. nov., Roseibacillus ishigakijimensis gen. nov., sp. nov., Roseibacillus ponti sp. nov., Roseibacillus persicicus sp. nov., Luteolibacter pohnpeiensis gen. nov., sp. nov. and Luteolibacter algae sp. nov., six marine members of the phylum 'Verrucomicrobia', and emended descriptions of the class Verrucomicrobiae the order Verrucomicrobiales and the family Verrucomicrobiaceae. Int. J. Syst. Evol. Microbiol. 58, 998-1007. Zengler, K., 2009. Central role of the cell in microbial ecology. Microbiol. Mol. Biol. Rev. 73, 712-729. Zul, D., Denzel, S., Kotz, A., Overmann, J., 2007. Effects of plant biomass, plant diversity, and water content on bacterial communities in soil lysimeters: Implications for the determinants of bacterial diversity. Appl. Environ. Microbiol. 73, 6916-6926.
120 Chapter 6: Rhizocompetence of culturable Holophaga (Acidobacteria) sp. in the leek (Allium porrum) rhizosphere*
Abstract
Holophaga is a minority group of Acidobacteria found in many soils. Due to its reduced abundance, this bacterial group may have been overlooked in previous studies of bacterial diversity in the rhizosphere. In the current chapter, cells of the recently-cultured Holophaga sp. CHC25 were found to be capable of colonizing sterilized leek roots, where they probably grew on diffusible compounds released from the roots. In experiments with an artifical rhizosphere, Holophaga sp. strain CHC25 cells consistently revealed raised numbers, as determined by Holophaga-specific qPCR, in the vicinity of leek roots. Using uninoculated soil, a similar observation was made for naturally occurring Holophagae. Furthermore, strain CHC25 cells were shown to migrate through soil towards the roots, which indicated the attractiveness of this niche. It was concluded that Holophagae, exemplified by strain CHC25, are leek rhizosphere competent Acidobacteria. This study is the first reporting on the ecology of Holophagae bacteria in the plant-soil ecosystem.
* Authored by: Ulisses Nunes da Rocha, Jan Dirk van Elsas & Leonard Simon van Overbeek Submitted for publication Rhizosphere competence of Holophaga
Introduction
The rhizosphere is defined as the soil under the direct influence of plant roots (Hiltner 1904). The co-evolution of plants and microbes has created a complex net of interactions between these two partners, which may influence the growth and health status (fitness) of plants and therefore crop productivity (Phillips et al. 2003). For instance, members of the microbial populations that inhabit the rhizosphere may be used as stress “controllers” (Glick et al. 2007), biofertilizers (van Rhijn and Vanderleyden 1995), biocontrol agents (Compant et al. 2005) or phytostimulators (Ryu et al. 2003). Before characterizing the ecological interactions between specific rhizosphere bacteria and the host plant, it is necessary to know if the former are rhizosphere-competent, i.e. find a colonizable niche in the rhizosphere. Rhizosphere competence here is defined as the ability to compete with other rhizosphere microbes for the nutrients secreted by roots and for sites that can be occupied on the root surface or in rhizosphere soil (Lugtenberg and Kamilova 2009). The complexity of microbial communities in the plant-soil ecosystem hampers investigations of bacterial root colonization at the level of single populations. Hence, simplified systems, in which cells of one or a few bacterial strains are applied to plant-soil systems, are commonly used to study the efficiency of root colonization, e.g. as a response to root exudation (Chin-A-Woeng et al. 1997). Different substances can be exuded by plant roots (Uren 2007). These can be water-soluble (e.g. sugars, amino acids, organic acids, hormones and vitamins) or -insoluble (e.g. polymeric carbohydrates). Further, lysates
(e.g. products of cell autolysis such as cell walls) and gases (e.g. ethylene and CO2) can be released. Although the composition of root exudates is complex and not always known, amino acids and organic acids have consistently been shown to serve as chemoattractants to root colonizing bacteria like Pseudomonas fluorescens (De Weert et al. 2002) and Bacillus subtilis (Rudrappa et al. 2008). Most bacteria that occur in the rhizosphere have so far remained uncultured (Nunes da Rocha et al. 2009), which restricts our ecological knowledge of these bacterial groups to what can be gleaned from culture-independent (molecular) techniques. For instance, Acidobacterium represents a bacterial phylum that, although often described as part of the dominant groups in the rhizosphere, has few culturable representatives. This phylum can make up 15 - 52% of 16S rRNA gene sequences in bacterial clone libraries (Dunbar et al. 2002; Sait et al. 2002). On the other hand, information about the ecology of this phylum in the rhizosphere is as-yet sparse (Nunes da Rocha et al., 2009). The class Holophagae (formely known as group 8 Acidobacteria) is a minority group within the Acidobacteria,
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which may range from 0 to 3.4% of the total Acidobacteria in soils (Jones et al., 2009). Due to the fact that other subgroups of the Acidobacteria (i.e. 1, 2, 3, 4, and 6) are present in higher numbers in soils (Jones et al., 2009), Holophagae may have been overlooked, much like their behaviour inside or near plants. Two culturable Holophagae strains, CHC25 and ORAC, were recently obtained from the rhizosphere (Nunes da Rocha et al., 2010a), but their ecological behaviour remained unaddressed. We hypothesized that Holophagae strains CHC25 and ORAC may be rhizosphere competent. To test this hypothesis we studied the in vitro colonization of Allium porrum roots by Holophagae strains CHC25 and ORAC, followed by studies on the survival of introduced Holophaga strain CHC25 in bulk soil and their migration to leek roots. To achieve our objectives, an ingenious plant-soil microcosm set-up, the so-called Kuchenbuch system (Kuchenbuch and Jungk, 1982) was used.
Material and Methods
Plant, soil and Holophagae strains
Seeds of leek (Allium porrum) cultivar Kenton (Nunhems Zaden BV, The Netherlands) were used. Bulk soil (taken at least at 1 m distance from leek plant roots) was collected from leek fields at ‘De Vredepeel’ farm, The Netherlands (51o 32’ 27.10” N and 5o 51’ 14.86” E). The soil was characterized as a sand (pH 5.4 and 2.2% of organic matter). It was stored for up to two days, for later experimentation. Also, separate aliquots of the same soil (500 g each) were sterilized by gamma irradiation (minimum doses 25 kGray, Isotron Nederland B.V., The Netherlands). Holophagae spp. strains CHC25 (accession number FN554392) and ORAC (FN689719), which were recently isolated from the leek rhizosphere (Nunes da Rocha et al., 2010a), were used in this study.
Leek root colonization by cells of Holophagae spp. strains CHC25 and ORAC
A leek root colonization assay was performed with cells of strains CHC25 and ORAC, which were previously grown on oligotrophic agar medium as described before (da Rocha et al., 2010). After growth, cells were harvested from the agar surface using oligotrophic broth (without glucose and agar) and adjusted to approximately 103 cells per 10 µL. Seeds were surface-sterilized according to the procedure described in Van Overbeek and Van Elsas (1995) and the sterilized seeds placed on the surface of oligotrophic agar without
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glucose and casein amino acids (da Rocha et al., 2010) in sterile Wavin containers model 2/80 (LAB Associates B.V., NL), with a total of five seeds per container. Each seed received 10 µl (103 cells) of the respective strain. The containers were incubated at room temperature with a 12h photoperiod for 14 days. In total, 30 seeds were inoculated per strain and the experiment was performed in duplicate. Two controls were added, first, surface-sterilized seeds - without addition of strains – were allowed to germinate on the same medium to check for the presence of endogenous bacteria, and, second, 10 µl (103 cells) of each strain was placed on oligotrophic agar -without glucose- to check if it could grow on this agar medium in the absence of emerging seeds. After incubation, pictures of the root colonization patterns were made with a stereomicroscopy Leica Wild M32 FL4 (Leica Microsystems B.V., NL) and analyzed for the following criteria: (i) bacterial growth along the plant roots, (ii) localization of bacterial growth (tip or distal), and (iii) effect of inoculation on root growth and development. The identity of the cells colonizing the roots was checked by BOX-PCR (Rademakers et al. 1998) and the size of the root and shoot of each individual plant was measured. The definition of the zones of primary root tip architecture was used as in Lópes- Bucio et al. (2005) and the following regions were distinguished: apical meristem, elongation and maturation zone. Although Holophagae spp. strains CHC25 and ORAC are not identical (clonal), they share morphological, physiological and phenotypical characteristics (Nunes da Rocha et al., 2010a). Therefore, it was decided to perform the soil microcosm experiments with only one strain, namely CHC25. The same experiments were also performed with Escherichia coli K12 strain E2 (bacterial collection of PRI, The Netherlands), a non-rhizosphere-competent bacterium, as the reference.
Quantitative Holophaga-specific PCR detection at different distances from leek roots in soil
Using specific primers, Acg8f and Acg8r (Nunes da Rocha et al, 2010b), qPCR assays were conducted on DNA extracts from samples taken from the soil and plant-soil microcosms (see later). DNA from all soil samples was extracted using the PowerSoil Isolation Kit (MO BIO Laboratories, Inc., CA, USA) following instructions provided by the manufacturer. The assays were done in Optical 96-Well Reaction Plates (Applied Biosystems, UK) on a 7500 Real-Time PCR system (Applied Biosystems, CA, USA). Each 25-µL reaction contained the following ingredients: 12.5 µL of SYBR Premix Ex Taq 2x (TAKARA Bio Inc., Japan), 0.5 µL of each primer (10 µM; Biolegio, NL), 0.5 µL of ROX Reference Dye
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II 50x (TAKARA Bio Inc., Japan), 6.0 µL H2O and 5.0 µl template. PCR conditions were one cycle of 50oC, 2 min; one cycle at 95oC, 10 s; 35 cycles at 95oC for 5 s, 60oC for 35 s. Each plate included triplicate reactions per DNA sample. Melting behaviour measurements of the PCR products was conducted after each amplification to confirm that the fluorescence signal originated from specific PCR products and not from primer-dimers or other nonspecific products formed during PCR. Cell numbers were estimated using DNA extracts made from serial 10-fold diluted suspensions, starting at the highest density of 108 cells per 5 µl for each strain. DNA extracts were added to the qPCR plates and for each dilution qPCR reactions were run in triplicate. Standard curves correlating the Log of cell numbers and threshold cycle (Ct) values were calculated and the theoretical dynamic range was estimated from these data in accordance with Nunes da Rocha et al. (2010b).
Background quantification of Holophaga cell equivalents in sterilized soil
Uninoculated sterilized (gamma radiated) soil was used as the control for the plant-soil microcosms inoculated with Holophaga sp. strain CHC25 cells. Because DNA from indigenous cells may remain intact after irradiation, Holophaga-qPCR was applied on DNA extracts made from this soil to set the background level for quantification of strain CHC25 cells after introduction into sterilized soil.
Survival of Holophaga sp. strain CHC25 in bulk soil
Survival of Holophaga strain CHC25 cells was followed over time in soil contained in 24 polyvinyl chloride (PVC) tubes (4.2 cm diameter; 4.0 cm height) covered with a nylon gauze (48 µm mesh size, Sefar B.V., CH). Before filling with soil, the rings with gauze covers were surface-sterilized by submergence for 2 min in a 2 % hypochlorite solution followed by repeated washings in sterile water and drying to air in a laminar flow hood. The rings were filled with sterile soil inoculated at a level of 105 cells per g dry soil, setting the bulk density at 1.2. Microcosms filled with soil were placed on a moisture tension table set at pF 2. Within one day, soil moisture corresponding to pF 2 was established (40%) and this condition remained stable for 5 weeks. Sampling was performed at set times, i.e. at 0 (1 hour after inoculation), 1, 3, 7, 14, 21, 28 and 35 days after inoculation. At each time point, samples taken from the PVC tubes were homogenized before one g samples from each ring were taken for DNA extraction.
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Rhizosphere colonization and migration to leek roots in a plant-soil microcosm set up
The model rhizosphere microcosm set up, previously described on Dijkstra et al. (1987) was applied using the same surface sterilized gauze-covered PVC rings as described before. Rings were filled with uninoculated and/ or strain CHC25-inoculated sterile soils or with uninoculated non-sterile soils (see below) and the bulk density was always set at 1.2. Twenty five leek seeds, surface-sterilized as described in Van Overbeek and Van Elsas (1995), were allowed to germinate on a sterile water-soaked filter paper and placed on top of the nylon gauze. The microcosms were then placed on a moisture tension table set at pF 2. This model plant-soil microcosm was used in four different experimental set ups, as follows: (A) Microcosms filled with sterile soil inoculated at a level of 105 Holophaga strain CHC25 cells per g dry soil. (B) Microcosms first filled with a 1 cm layer of uninoculated sterile soil near the gauze (chamber topside down) and then with a second layer of sterilized soil inoculated at a level of 105 Holophaga sp. strain CHC25 cells per g dry soil. (C) Microcosms filled with uninoculated non-sterile soil only. (D) Microcosms filled with sterile uninoculated soil only. Treatment D was meant as a control for eventual occurrences of contamination with Holophaga cells from external sources, like from the surface-sterilized seeds or via water flow from the tension table). (E) Microcosms first filled with a 1 cm layer of uninoculated sterile soil near the gauze (chamber topside down) and then with a second layer of sterilized soil inoculated at a level of 105 Holophaga sp. strain CHC25 cells per g dry soil. Here, leek seeds were not introduced in the top of the nylon gauze. This treatment was meant as a control for eventual migration of Holophaga sp. CHC25 without the leek root mat. From all four microcosm systems, the soil layers most proximate to the gauze with adhering leek root mat (0-2 mm) and at a further distance from the roots (10-12 mm) were singled out for DNA extraction followed by Holophaga-specific qPCR analysis.
Statistical analysis of the data
All experiments were done in triplicate systems. One-way ANOVA was used to compare the average sizes of the aerial parts and roots of the leek plants from the in vitro colonization assay. One-way ANOVA was used to compare the average Ct values measured at different time points during Holophaga sp. strain CHC25 survival in bulk soils. Two-factor ANOVA was used to compare the average Ct values measured in the
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different layers of the plant-soil microcosm set ups. Differences were considered to be significant at levels of P < 0.05.
Results
In vitro colonization of leek roots by Holophagae spp. strains CHC25 and ORAC
The ability of Holophagae spp. strains CHC25 and ORAC to colonize leek roots was determined by inoculation of surface-sterilized leek seeds placed on an OLI (without C source) agar surface with 103 cells of each strain. Three days after inoculation, the majority (70%) of the seeds had emerged, whereas after 14 days all seeds had emerged, yielding fully developed plants containing shoots and roots. At the latter time point, no differences were observed across the replicates between either the lengths of the roots or the aerial parts of the leek plants. This included plants inoculated with each of the two strains, the E. coli K12 E2 strain and the uninoculated plants (Table 1). The average length of the aerial parts ranged between 43.1 and 54.6 mm and the root size between 6.3 and 10.2 mm.
Table 1 Size of leek plants after 14 days growing in vitro on oligotrophic agar with no C source and inoculated with 103 cells of each isolate in different seeds. Holophaga Experiment Aerial part Root size strain Repetitiona Avarage Standard deviation Avarage Standard deviation None A 44.7 13.6 7.84 3.81 (control) B 43.1 13.4 9.41 2.91 CHC25 A 50.9 10.6 7.67 3.82 B 48.9 10.8 8.46 2.76 ORAC A 49.7 10.0 7.97 2.93 B 47.3 10.0 8.54 2.66 a Each of the repetitions consist on the inoculation of 30 different seeds
Bacterial colonization was never observed near emerging leek plants grown from uninoculated seeds and also not near those inoculated with reference strain E. coli K12 E2. Holophaga strain CHC25 was found to visually colonize the roots of leek plants in 27 out
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of 55 occasions (49%) and strain ORAC in 29 of 57 occasions (51%), implying visible growth at the root surface. Bacterial cells retrieved from the roots showed BOX-PCR profiles identical to the strains they derived from (data not shown), demonstrating that the growth along the roots resulted from the inoculant Holophaga strains and not from eventual contaminating bacteria. Because no carbon source was added to the OLI medium, and no growth was observed when the single strains were placed on this medium, the growth along the leek roots was likely dependent on diffusible compounds released from the roots. Plants treated with the same strain always showed the same pattern of colonization, and these differed between the two strains (Fig. 1). Holophaga strain CHC25 showed dense and strain ORAC translucent colonization of the root maturation zone, in particular in the zone close to the aerial part, but no growth was observed on the apical meristem and the elongation zones (Fig. 1B and C).
Survival of Holophaga strain CHC25 cells in bulk soil
One hour after introduction of Holophaga sp. strain CHC25 cells into sterile soil, the Holophaga cell equivalents per g dry soil were (Log-transformed values) between 4.86 and 5.15 (Fig. 2). After one day, these numbers had decreased to 4.15 - 4.47, after which an increase was noted, to levels of 4.85 - 5.25 after 14 days. These numbers then remained virtually stable (4.98 - 5.21) over the entire monitoring period of 35 days.
Plant-soil microcosm experiments
Root and shoot development in leek plants emerging from surface-sterilized seeds became evident after four days and dense root mats were observed after 3 weeks in all plant-soil microcosms. There was no difference in growth between the leek plants in the four different microcosm set-ups (A –D) (data not shown). In the control plant-soil microcosm (D, with sterilized soil only), the Holophaga cell equivalents per g dry soil in the 0-2 mm and 10-12 mm zones were at background level, respectively, 3.43 - 3.64 and 3.46 - 3.65 (Log scale) (Fig. 3). In the A microcosms, in which the inoculated soil was in direct contact with the gauze and root mat, the Holophaga cell equivalents per g dry soil were (Log scale) between 5.86 and 5.93 in the 0-2 mm zone, whereas these numbers were significantly lower in the 10-12 mm zone (P<0.05), i.e. between 4.91 and 5.36 (Fig. 3). In the B microcosms, in which the inoculated soil was separated from the gauze and root mat by a 1-cm layer of uninoculated sterile soil, we found between (Log scale) 5.08 and 5.45 Holophaga cell equivalents per g dry soil in the 0-2 mm zones, and 5.55 and 5.90 in the 10-12 mm zones.
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Figure 1 In vitro root colonization of two-weeks-old leek plants emerged from surface- sterile seeds and either remained untreated (A) or were treated with 103 cells of Holophagae strains CHC25 (B) or ORAC (C).
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5.5
A 5.0 A A A A
A,B
B 4.5
c
Log equivalent cell number per gram of dry soil of dry per gram number cell equivalent Log 4.0 0 7 14 21 28 35 Days
Figure 2 Survival of Holophaga strain CHC25 in sterile soil. Soil was inoculated at a level of 105 cells per gram of dry sterile bulk soil and equivalent cell number per gram of dry soil were measured over time using a Holophagae-specific qPCR system (Nunes da Rocha et al., 2010b). Letters at each data point represent significance of difference, where A > B > C (P
In the E microcosms, similar to set up B but without the root mat on top of the nylon gauze, we found between (Log scale) 3.43 and 3.65 Holophaga cell equivalents per g dry soil in the 0-2mm zones, similar number to those found in the background control, and 5.33 and 5.76 in the 10-12 mm zones. This indicated that Holophaga strain CHC25 cells had migrated through the uninoculated soil zone towards the gauze containing the leek roots (Fig 3.). In the C microcosms, in which uninoculated non-sterile soil was used, the 0-2 mm zones contained between (Log) 4.87 and 4.95 Holophaga cell equivalents per g dry soil, whereas in the 10-12 mm soil zone these numbers were between 4.19 and 4.29. From the foregoing, it was concluded that significantly higher numbers of strain CHC25 or of total Holophaga cell equivalents (natural soil) were consistently present in the soil zones proximate to the leek roots in the A and C microcosms. The presence of strain CHC25 in the same soil zone in the B microcosms demonstrated that strain CHC25 cells had migrated towards the leek roots, as such migration was not found in the control setup.
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10.0 A
8.0 ** 6.0
4.0
2.0 CHC25 10.0 B
8.0 ns 6.0
4.0
2.0 CHC25 10.0 C 8.0
6.0 **
4.0
2.0 Natural population
0-2 mm zone
10-12 mm zone
Figure 3 Leek rhizosphere colonization of Holophaga strain CHC25 (A, B) and of Holophagae bacteria naturally occurring in soil (C) in different plant-soil microcosm settings. Sterilized soil inoculated with Holophagae isolate CHC25 was brought into direct contact with the nylon gauze and leek root mat (A), or was separated with a 1 cm layer of non-inoculated sterile soil layer (B). Non-sterile and non-inoculated soil was brought into contact with the nylon gauze with roots in C. Log equivalent cell numbers, expressed per g of dry soil, were determined by Holophagae-specific qPCR in 0-2 mm and 10-12 mm soil zones in all microcosm settings. Ns indicates not significantly different and ** significantly different at levels of p<0.01, arrow indicates presence of CHC25 cell equivalents in the non-inoculated soil layer in B and bar (B) represent the average Log equivalent cell numbers (per gram of dry soil) measured as the estimated background Holophagae cell level in sterile soil.
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Discussion
In this study, it was hypothesized that the two Holophaga spp. strains CHC25 and ORAC that were recently obtained from the leek rhizosphere, are rhizosphere-competent, i.e. show a growth/migration response to leek roots. From the in vitro experiments conducted in this study, it became evident that the two Holophaga spp. strains could thrive on plates where leek plants developed, whereas no growth was observed on plates without plants. This indicated the utilization of nutrients released by the leek roots in the form of exudates. The in vitro colonization of, and growth on, the sterile leek roots corroborates findings made before with strains CHC25 and ORAC (Nunes da Rocha et al., 2010a). Specifically, both strains were able to grow on media containing malic acid, succinic acid, citric acid, glutamine or alanine as the sole carbon sources. These compounds are considered as ‘common’ for rhizospheres of many different plant species (Baudoin et al., 2003; Jones 1998) and hence the two strains have potential to occupy niches proximate to roots that exude such compounds. If cells of these strains are able to proliferate in the neighborhood of sterile leek roots then the question arises whether they will behave in the same way as typical rhizosphere bacteria, i.e. are they able to survive in the complex soil matrix in the rhizosphere (Berg and Smalla, 2009) and are they able to compete with other soil microbes in there (Bueé et al., 2009)? A model plant-soil microcosm designed and evaluated about 30 years ago (Kuchenbuch and Jungk 1982) was applied in the original, and in a modified, version to investigate the putative rhizosphere competence of Holophaga sp. strain CHC25 in sterilized soil and of endogenous Holophagae in non-sterile soils. A gradient of root- released compounds is formed in the soil zone directly beneath the gauze that carries the root mat. Bacteria that are attracted to this zone may utilize the released compounds for growth and metabolic activities. Using this experimental set-up, rhizosphere competence was shown in the past with the typical rhizosphere dweller Enterobacter cloaceae (Dijkstra et al., 1987). Later, the Kuchenbuch system demonstrated its validity for measuring bacterial activities in the form of enhanced plasmid mobilization and transfer in soil- introduced Pseudomonas fluorescens strains (Van Elsas et al., 1988) and in the induction of a root exudate responsive operon in the same species (Van Overbeek and Van Elsas, 1995). The estimated Holophaga numbers found in the rhizosphere (expressed as number of cell equivalents per g dry soil) were about 105. This number is approximately one to two orders in magnitude lower than measured P. fluorescens CFU or cell numbers in different rhizospheres (Van Elsas et al., 1992; Hase et al., 2000). The Holophagae may not represent a numerically dominant group in the rhizosphere, but the fact that Holophaga equivalent
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cell numbers were significantly higher in the rhizosphere than in corresponding bulk soils indicated that the soil zone proximate to plants roots are preferred sites for this bacterial species. Moreover, cells of strain CHC25 had migrated towards the roots, indicating chemotaxis as a likely mechanism involved in rhizosphere competence. Apparently, Holophaga cells follow the nutrient gradient to the roots upwards and migratory activity towards plant roots can be considered as a first step in root colonization (De Weert et al., 2002; Rudrappa et al., 2008). Collectively, our observations demonstrate clear rhizosphere competence for Holophaga sp. CHC25. Holophagae may successfully compete for root-released nutrients in the rhizosphere, which is a remarkable finding, because the in vitro growth rates of the two Holophaga strains (between 12.3 h to 13.8 h per cell division; Nunes da Rocha et al., 2010b) were much lower than that of the rhizosphere-competent P. fluorescens, which is about 100 min per duplication (Compeau et al., 1988), and of Bacillus subtilis, approximately 60 min per duplication (Wimpenny and Lewis, 1977). In comparison with P. fluorescens and B. subtilis, species that must be considered as typical opportunists commonly occurring in the rhizosphere, the two Holophaga strains may have to be regarded as oligotrophs or K strategists. However, the fact that our Holophagae can grow at higher nutrient levels (Nunes da Rocha et al., 2010b) indicates that they are not fine tuned to scavenge scarcely-available nutrients, a feature unique to the obligate oligotrophs (Senechkin et al., 2010). The two Holophagae may thus be copiotrophic in nature, allowing an ecologically relevant growth rate. This in spite of the fact that their growth rates, estimated under ex situ circumstances, are very slow. The phylum Acidobacteria has been described as consisting of oligotrophs as a generalized statement for the entire phylum (Fierer et al., 2009); here, we indicate that this is not the case for Holophagae spp. in the leek rhizosphere environment. How these bacteria find their niche in the rhizosphere can only be speculated upon. Possibly, they occupy specific niches in which they are nutritionally or even physically shielded. Holophagae may thus thrive in soil proximate to plant roots. This finding contradicts reports made before on the occurrences of Acidobacteria in different plant-soil compartments. Acidobacteria are very often present in raised numbers in bulk soils (Jones et al., 2009), like the group 1, 2, 3, 4 and 6 Acidobacteria (Jones et al., 2009). These groups were even found to dominate in bulk soils, occupying 15% or more of the total bacterial community (Rousk et al., 2010). Holophaga supposedly represents a minority group, in terms of prevalence in diverse ecosystems, among the Acidobacteria. It may be considered as the ‘exception on the rule’ that Acidobacteria thrive in bulk but not in rhizosphere soils. The fact that Holophagae - among the Acidobacteria - might represent uniqueness in their
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interaction with higher plants will make future studies on the mechanisms of these interactions highly relevant. We conclude that the two Holophaga strains that we recently cultured from the leek rhizosphere are indeed rhizosphere-competent. The ecophysiology of Holophagae thus differs from what is commonly understood about Acidobacteria, namely that they prefer bulk over rhizosphere soils (Kielak et al., 2008) and that they are oligothrophic in nature (Fierer et al., 2007). Our qPCR-based assessment of the Holophaga sp. CHC25 dynamics enabled us to draw firm conclusions about strain behaviour in the rhizosphere, something that would have been impossible to perform with culture-independent approaches alone. Application of culture-based methods for studying representatives of the phylum of Acidobacteria opens new avenues to explore minority groups within this vast bacterial phylum in their natural settings.
Acknowledgements This research was part of the Ecogenomics program which is sponsored by the Dutch National Genomics Initiative and the basic research program on sustainable agriculture (KB4) sponsored by the Dutch ministry of agriculture, nature and food safety. We would like to thanks Pieter Kastelein and Vladimir Fediy for their assistance in the plant-soil microcosm experiments.
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References
Baudoin E, Benizri E, Gauckert A (2003) Impact of artificial root exudates on the bacterial community structure in bulk soil and maize rhizosphere. Soil Biol Biochem 35: 1183-1192. Berg G, Smalla K (2009) Plant species and soil type cooperatively shape the structure and function of microbial communities in the rhizosphere. FEMS Microbiol Ecol 68: 1-13. Buée M, De Boer W, Martin F, van Overbeek L, Jurkevitch E (2009) The rhizosphere zoo: An overview of plant-associated communities of microorganisms, including phages, bacteria, archaea, and fungi, and of some of their structuring factors. Plant Soil 321: 189-212. Chin-A-Woeng TFC, de Priester W, van der Bij AJ, Lugtenberg BJJ (1997) Description of the colonization of a gnotobiotic tomato rhizosphere by Pseudomonas fluorescens biocontrol isolate WCS365, using scanning electron microscopy. Mol Plant-Microbe Interact 10: 79-86. Compant S, Duffy B, Nowak J, Clément C, Barka EA (2005) Use of plant growth-promoting bacteria for biocontrol of plant diseases: principles, mechanisms of action, and future prospects. Appl Environ Microbiol 71: 4951-4959. Compeau G, Al-Achi BJ, Platsouka E, Levy SB (1988) Survival of rifampin-resistant mutants of Pseudomonas fluorescens and Pseudomonas putida in soil systems. Appl Environ Microbio 54: 2423- 2438. da Rocha UN, Andreote FD, de Azevedo JL, van Elsas JD, van Overbeek LS (2010) Cultivation of hitherto- uncultured bacteria belonging to the Verrucomicrobia subdivision 1 from the potato (Solanum tuberosum L.) rhizosphere. J Soils Sediments 10: 326-339. De Weert S, Vermeiren H, Mulders IHM, Kuiper I, Hendrickx N, et al. (2002) Flagella-driven chemotaxis towards exudate components is an important trait for tomato root colonization by Pseudomonas fluorescens. Mol Plant-Microbe Interact 15: 1173-1180. Dijkstra AF, Govaert JM, Scholten GHN, Van Elsas JD (1987) A soil chamber for studying the bacterial distribution in the vicinity of roots. Soil Biol Biochem 19: 351-352. Dunbar J, Barns SM, Tichnor LO, Kuske CR (2002) Empirical and theoretical bacterial diversity in four Arizona soils. Appl Environ Microbiol 68: 3035-3045. Fierer N, Bradford MA, Jackson RB (2007) Toward an ecological classification of soil bacteria. Ecology 88: 1354-1364. Glick BR, Cheng Z, Czarny J, Duan J (2007) Promotion of plant growth by ACC deaminase-producing soil bacteria. Eur J Plant Pathol 119: 329-339. Hase C; Hottinger M; Moënne-Loccoz y; Defago G (2000) Survival and cell culturability of biocontrol Pseudomonas fluorescens CHAO in the rhizosphere of cucumber grown in two soils of contrasting fertility status. Biol Fertil Soils 32: 217-221. Hiltner L (1904) Über neuere Erfahrungen und Probleme auf dem Gebiete der Bodenbakteriologie unter bessonderer Berüchsichtigung der Gründung und Brache. Arb Dtshc Landwirtsch Ges Berl 98: 59-78. Jones DL (1998) Organic acids in the rhizosphere – a critical review. Plant Soil 205: 25-44. Jones RT, Robeson MS, Lauber CL, Hamady M, Knight R, Fierer N (2009) A comprehensive survey of soil acidobacterial diversity using pyrosequencing and clone library analyses. ISME J 3: 442-453. Kielak A, Pijl AS, Van Veen JA, Kowalchuk GA (2008) Differences in vegetation composition and plant species identity lead to only minor changes in soil-borne microbial communities in a former arable field. FEMS Microbiol Ecol 63: 372-382. Kuchenbuch R, Jungk A (1982) A method for determining concentration profiles at the soil-root interface by thin slicing rhizospheric soil. Plant Soil 68: 391-394. Lópes-Bucio J, Cruz-Ramírez A, Pérez-Torres A., Ramírez-Pimente JG, Sánchez-Calderón L, Herrera- Estrella L (2005) Root Architecture. In: Turnbull CGN (Ed.), Plant Architecture and its manipulation, Blackwell Publishing, Oxford, UK. Chapter 7.4.1 pp. 189-191. Lugtenberg B, Kamilova F (2009) Plant-growth-promoting rhizobacteria. Annu Rev Microbiol 63: 541-556. Nunes da Rocha U, Andreoti FD, Plugge C, Ausek L, van Elsas JD, Overbeek L (2010a) Isolation and partial characterization of Holophagae, Luteolibacter, unclassified Verrucomicrobia and Verrucomicrobium from the leek (Allium porrum) rhizosphere. Submitted.
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Nunes da Rocha U., van Elsas JD, Overbeek L (2010b) Real-time PCR detection of Holophagae (Acidobacteria), Luteolibacter/Prosthecobacter and unclassified Verrucomicrobiaceae (Verrucomicrobia subdivision 1) in bulk and leek (Allium porrum) rhizosphere soils. Submitted. Nunes Da Rocha U, Van Overbeek L, Van Elsas JD (2009) Exploration of hitherto-uncultured bacteria from the rhizosphere. FEMS Microbiol Ecol 69: 313-328. Phillips DA, Ferris H, Cook DR, Strong DR (2003) Molecular control point in rhizosphere food webs. Ecology 84: 816-826. Rademaker JLW, Louws FJ, De Bruijn FJ (1998) Characterization of the diversity of ecologically important microbes by rep-PCR genomic fingerprinting. Molecular Microbial Ecology Manual (SUPPL. 3), pp. 1- 26. Rousk J, Bååth E, Brookes PC, Lauber CL, Lozupone C, Caporaso JG, Knight R, Fierer N (2010) Soil bacterial and fungal communities across a pH gradient in an arable soil. ISME J DOI: 10.1038/ismej.2010.58. Rudrappa T, Czymmek KJ, Paré PW, Bais HP (2008) Root-secreted malic acid recruits beneficial soil bacteria. Plant Physiol 148: 1547-1556. Ryu C-M, Farag MA, Hu C-H, Reddy MS, Wie H-X, et al. (2003) Bacterial volatiles promote growth of Arabidopsis. Proc Natl Acad Sci USA 100: 4927-4932. Sait M, Hugenholtz P, Janssen PH (2002) Cultivation of globally distributed soil bacteria from phylogenetic lineages previously only detected in cultivation-independent surveys. Environ Microbiol 4: 654-666. Senechkin IV, Speksnijder AGCL, Semenov AM, van Bruggen AHC, van Overbeek LS (2010) Isolation and Partial Characterization of Bacterial Strains on Low Organic Carbon Medium from Soils Fertilized with Different Organic Amendments. Antonie Van Leeuwenhoek DOI: 10.1007/s00248-010-9670-1. Uren NC (2007) Types, amounts, and possible functions of compounds released into the rhizosphere by soil- grown plants. In: Pinton R, Varanini Z, Nannipieri P (Eds), The rhizosphere. Biochemistry and Organic Substances at he Soil-Plant Interface, CRC Press/Taylor & Francis Group, Boca Raton pp 1-21. Van Elsas JD, Trevors JT, Jain D, Wolters AC, Heijnen CE, van Overbeek LS (1992) Survival of, and root colonization by, alginate-encapsulated Pseudomonas fluorescens cells following introduction into soil. Biol Feril Soils 14: 14-22. Van Elsas JD, Trevors JT, Starodub ME (1988) Bacterial conjugation between pseudomonads in the rhizosphere of wheat. FEMS Microbiol Ecol 53: 299-306. Van Overbeek LS, Van Elsas JD (1995) Root exudate-induced promoter activity in Pseudomonas fluorescens mutants in the wheat rhizosphere. Appl Environ Microbiol 61: 889-898. van Rhijn P, Vanderleyden J (1995) The Rhizobium-plant symbiosis. Microbiol Rev 59: 124-142. Wimpenny JWT, Lewis MWA (1977) The growth and respiration of bacterial colonies. J Gen Microbiol 103: 9-18.
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Chapter 7: Different rhizosphere competence in two Verrucomicrobium subdivision 1 strains previously isolated from the leek (Allium porrum) rhizosphere*
Abstract
In chapter 4, two new strains, CHC12 and CHC8, belonging to, respectively, Luteolibacter and Candidatus genus Rhizospheria (Verrucomicrobia subdivision 1) were isolated from the leek rhizosphere. To test their rhizosphere competence, we investigated the in vitro colonization of the roots of leek plants growing in oligotrophic medium without an added carbon source. Furthermore, experiments were performed in soil microcosms to assess the survival of the two strains in soil and in model plant-soil microcosms to assess the cell numbers in the vicinity of plant roots compared to bulk soil. As growth of these bacteria from soil is extremely fastidious, real-time qPCR detection with specific primers was used. The data shown in this chapter indicated that natural populations of Luteolibacter (akin to strain CHC12) had lower number in the rhizosphere than in the corresponding bulk soil. On the other hand, the populations of Candidatus genus Rhizospheria, i.e. strain CHC8, showed higher numbers in the leek rhizosphere than in the bulk soil. The raised numbers in the rhizosphere were not only the result of in situ cell multiplication but also of migration of cells from distant sites in the soil towards the roots.
* Authored by: Ulisses Nunes da Rocha, Jan Dirk van Elsas & Leonard Simon van Overbeek Submitted by publication Rhizosphere competence of different Verrucomicrobium subdivision 1 strains
Introduction
Verrucomicrobium was proposed as a new division of the bacterial domain in the late 1990-ies (Hedlund et al., 1997) and finally ranked as a phylum in the beginning of this decade (Garrity and Holt, 2001). Members of this phylum are widely distributed and abundant in the soil environment (Hugenholtz et al., 1998). In spite of their high abundance and diversity, little information is available on their ecology, which is mainly due to the lack of culturable representatives in current-day bacterial collections (Nunes da Rocha et al., 2009). Therefore, most of the ecological information about the taxon is based on Verrucomicrobium 16S rRNA gene sequence distribution over different ecosystems. For instance, the phylum was found to be among the dominant bacterial groups in communities present in soils and rhizospheres (Rosenberg et al., 2009), drinking water reservoirs (Lymperopoulou et al., 2010), human intestinal tract systems (Wang et al., 2005), contaminated groundwater (Herrmann et al., 2008), animal (gorilla) feces (Frey et al., 2006) and swine waste lagoons (Goh et al., 2009). Analyses of 16S rRNA genes of Verrucomicrobia in samples from terrestrial habitats often revealed contradictory information about their preferred sites, especially in the distinction between bulk and rhizosphere soils (Chow et al., 2002; Sanguin et al., 2006; Zul et al., 2007; Kielak et al., 2008). Next to being widely distributed and rather abundant, members of the Verrucomicrobia are also highly diverse. The phylum contains seven subdivisions (Schelsner et al., 2006). Previous studies on the distribution between bulk and rhizosphere soil considered all Verrucomicrobium subdivisions together. This approach makes a differentiation of those groups that are ecologically related to plants difficult. Furthermore, culturable members of the subdivision 1 Verrucomicrobia have recently been isolated from the rhizospheres of potato (da Rocha et al., 2010) and leek (Nunes da Rocha et al., 2010a). Strains of Verrucomicrobia that belong to the same genus were shown to exhibit different physiological profiles (Nunes da Rocha et al., 2010a). Therefore, two questions about these strains can be posed: (i) Are these bacteria rhizosphere competent? (ii) Do closely related strains of Verrucomicrobia subdivision 1 respond similarly to the rhizosphere? In the current chapter, we hypothesized that strains of subdivision 1 of the Verrucomicrobia that had been isolated from the rhizosphere were rhizosphere competent. Rhizosphere competence here is defined as the ability to compete with other rhizosphere micro-organisms for the nutrients secreted by roots and for sites that can be occupied on the root surface or in rhizosphere soil (Lugtenberg and Kamilova 2009). To test this hypothesis, the in vitro colonization of Allium porrum roots by Luteolibacter strain CHC12 and Candidatus genus Rhizosphere strain CHC8 (Nunes da Rocha et al., 2010a) was studied, using the methods described in Nunes et al (2010). To achieve our
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objectives, an ingenious plant-soil microcosm set-up, i.e. the so-called Kuchenbuch system (Kuchenbuch and Jungk, 1982) and real time qPCR detection of the strains was used (Nunes da Rocha et al., 2010c).
Material and Methods
Plant type and soil
Seeds of leek (Allium porrum) cultivar Kenton (Nunhems Zaden BV, The Netherlands) were used. Bulk soil (taken at least at 1 m distance from leek plant roots) was collected from leek fields at ‘De Vredepeel’ farm, The Netherlands (51o 32’ 27.10” N and 5o 51’ 14.86” E). The soil was characterized as a sand (pH 5.4 and 2.2% of organic matter). It was stored for up to two days, for later experimentation. Also, separate aliquots of the same soil (500 g each) were sterilized by gamma irradiation (minimum doses 25 kGray, Isotron Nederland B.V., The Netherlands).
Bacterial strains
Luteolibacter sp. strain CHC12 (accession number FN554390) and Candidatus genus Rhizospheria strain CHC8 (FN554389), which had both been isolated from the leek rhizosphere (Nunes da Rocha et al., 2010a), were used in this study. Strains were routinely grown on oligotrophic agar medium (Nunes da Rocha et al., 2010a).
In vitro root colonization assay
A leek root colonization assay was performed with freshly-grown cells of strains CHC12 and CHC8. Prior to the experiment, both strains were grown on oligotrophic agar, as described before (da Rocha et al., 2010). After growth, cells were washed from the agar surface using oligotrophic broth (without glucose and agar) and cell density was adjusted to approximately 103 cells per 10 µL. Leek seeds were surface-sterilized and rinsed thoroughly according to Van Overbeek and Van Elsas (1995). The sterilized seeds were then placed on the surface of oligotrophic agar without glucose and casein amino acids (da Rocha et al., 2010) in sterile Wavin containers model 2/80 (LAB Associates B.V., NL), placing a total of five seeds per container. Each seed received 10 µL (103 cells) of a suspension of one strain. The containers were then incubated at room temperature under a 12-h photoperiod for 14 days. In total, 30 seeds were inoculated per strain and the experiment was performed in duplicate. Two controls were added: (i) surface-sterilized seeds, without addition of strains, were allowed to germinate on the
139 Rhizosphere competence of different Verrucomicrobium subdivision 1 strains
same medium to check for the healthy outgrowth of seedlings and the possible presence of remaining (endogenous) bacteria, and (ii) 10 µL (103 cells) of each strain was placed on oligotrophic agar without glucose to check if growth would occur on this agar medium in the absence of emerging seedlings. After the incubation period, pictures were made of the root colonization patterns using a Leica Wild M32 FL4 stereomicroscope (Leica Microsystems B.V., NL). The pictures were then analyzed for the following criteria: (i) bacterial growth along the plant roots, (ii) localization of bacterial growth (tip or distal), and (iii) effect of bacterial inoculation on root growth and development. The identity of the cells that colonized the roots was checked by BOX-PCR (Rademakers et al. 1998) and the sizes of the roots and shoots of each individual plant were measured. The definition of the zones of primary root tip architecture was used as described in Lópes-Bucio et al. (2005). Using this, the following regions were distinguished: apical meristem, elongation and maturation zone. The same experiments were also performed with Escherichia coli K12 strain E2 (bacterial collection of PRI, The Netherlands), a non-rhizosphere-competent bacterium.
Quantitative Luteolibacter and Candidatus genus Rhizospheria specific PCR detection at different distances from leek roots in soil
Specific primers developed for Luteolibacter (VS1Af and VS1Ar) and for Candidatus genus Rhizospheria (VS1Bf and VS1Br) (Nunes da Rocha et al, 2010c) were used in qPCR assays on DNA extracts from samples taken from soil and plant-soil microcosms (see later). The DNA was extracted using the PowerSoil Isolation Kit (MO BIO Laboratories, Inc., CA, USA) following the instructions provided by the manufacturer. The assays were done in Optical 96-Well Reaction Plates (Applied Biosystems, UK) on a 7500 Real-Time PCR system (Applied Biosystems, CA, USA). Each 25 µL reaction contained: 12.5 µL of SYBR Premix Ex Taq 2x (TAKARA Bio Inc., Japan), 0.5 µL of each primer (10 µM; Biolegio, NL), 0.5 µL of ROX Reference Dye II 50x (TAKARA
Bio Inc., Japan), 6.0 µL H2O, and 5.0 µL (< 5 ng) of template. PCR conditions were: one cycle of 50 oC, 2 min; one cycle at 95 oC, 10 s; 35 cycles at 95 oC for 5 s, 60 oC for 35 s. Each plate included triplicate reactions per DNA sample. Melting behavior measurements of the PCR products were conducted after each amplification to confirm that the fluorescence signal originated from specific PCR products and not from aspecific products formed during PCR. Cell numbers were estimated using DNA extracts prepared from serial 10-fold diluted suspensions, starting at the highest density, i.e. 108 cells per 5 µL for each strain. DNA extracts were added to the qPCR plates and, for each dilution, qPCR reactions were run in triplicate. Standard curves correlating the
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Log cell numbers and threshold cycle (Ct) values were calculated. From these data, a theoretical dynamic range was estimated in accordance with Nunes da Rocha et al. (2010c).
Background quantification of sequences of Luteolibacter and Candidatus genus Rhizospheria cells in sterilized soil
Uninoculated gamma-irradiated soil was used as the control for the plant-soil microcosms inoculated with strains CHC12 and CHC8. Since DNA from indigenous cells may remain intact after irradiation and may thus be amplified in a subsequent PCR, Luteolibacter (and Prosthecobacter) and Candidatus genus Rhizospheria qPCRs were applied on DNA extracts from this soil. This set the background for quantification of strain CHC12 and CHC8 populations after introduction into the sterilized soil.
Survival of Luteolibacter sp. strain CHC12 and Candidatus genus Rhizospheria sp. CHC8 in bulk soil
Survival of Luteolibacter sp. strain CHC12 and Candidatus genus Rhizospheria sp.CHC8 cells were separately followed over time in soil contained in polyvinyl chloride (PVC) tubes (4.2 cm diameter; 4.0 cm height), each one covered with a nylon gauze (48 µm mesh size, Sefar B.V., CH). Before filling with soil, the rings with gauze covers were surface-sterilized by submergence for 2 min in a 2 % hypochlorite solution followed by repeated rinsing in sterile water and air drying in a laminar flow hood. The rings were filled with sterile soil inoculated at a level of 105 cells per g dry soil, setting the bulk density at 1.2. Microcosms filled with soil were placed on a moisture tension table set at pF 2. Within one day, soil moisture corresponding to pF 2 was established (40%) and this condition remained stable for 5 weeks. Sampling was performed at set times, i.e. at 0 (1 hour after inoculation), 1, 3, 7, 14, 21, 28 and 35 days after inoculation. At each time point, samples taken from the PVC tubes were homogenized, after which 0.5 g samples were taken for DNA extraction. Rhizosphere colonization and migration to leek roots in a plant-soil microcosm set up
The model rhizosphere microcosm set-up, previously described in Dijkstra et al. (1987) was applied using the surface-sterilized gauze-covered PVC rings described above. Rings were filled with uninoculated and/or strain CHC25-inoculated sterilized soils or with uninoculated non-sterile soils (see below), setting the bulk density at 1.2. Twenty- five leek seeds, surface-sterilized as described (Van Overbeek and Van Elsas 1995), were allowed to germinate on a sterile water-soaked filter paper, after which they were placed on top of the nylon gauze (see below). The microcosms were then placed on a
141 Rhizosphere competence of different Verrucomicrobium subdivision 1 strains
moisture tension table set at pF 2. This model plant-soil microcosm was used in four different experimental set-ups, as follows: (A) Microcosms filled with sterile soil inoculated with 106 Luteolibacter sp. CHC12 or Candidatus genus Rhizospheria sp. CHC8 cells per g dry soil. (B) Microcosms first filled with a 1 cm layer of uninoculated sterile soil near the gauze (chamber topside down) and then with a second layer of sterilized soil containing 106 Luteolibacter sp. CHC12 or Candidatus genus Rhizospheria sp. CHC8 cells per g dry soil. (C) Microcosms filled with uninoculated non-sterile soil only. (D) Microcosms filled with sterile uninoculated soil only. Treatment D was meant as a control for eventual occurrence of Luteolibacter or Candidatus genus Rhizospheria cells from external sources, e.g. from the surface-sterilized seeds or via water flow from the tension table. In treatment B, leek seeds were either introduced or left out of the top of the nylon gauze. The non-plant treatment (denoted B2) was meant as a control for eventual migration of Luteolibacter sp. CHC12 or Candidatus genus Rhizospheria sp. CHC8 without the presence of the leek root mat. From all microcosm systems, the soil layers most proximate to the gauze with an adhering leek root mat (0-2 mm) and at a further distance from the roots (10-12 mm) were singled out for DNA extraction and subsequent Luteolibacter or Candidatus genus Rhizospheria-specific qPCR analysis.
Statistical analysis of the data
All experiments were done in triplicate systems. One-way ANOVA was used to compare the average sizes of the aerial parts and roots of the leek plants from the in vitro colonization assay. One-way ANOVA was used to compare the average Ct values measured at different time points during Luteolibacter sp. CHC12 or Candidatus genus Rhizospheria sp. CHC8 survival in bulk soils. Two-factor ANOVA was used to compare the average Ct values measured in the different layers of the plant-soil microcosm set ups. Differences were considered to be significant at levels of P < 0.05.
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Results
In vitro colonization of leek roots by Luteolibacter sp. CHC12 and Candidatus genus Rhizospheria sp. CHC8
The ability of Luteolibacter sp. CHC12 and Candidatus genus Rhizospheria sp. CHC8 to colonize leek roots was first determined by inoculation of surface-sterilized leek seeds placed on an OLI agar surface with 103 cells of either of the two strains. Three days following inoculation, the majority (70%) of the seeds had emerged, whereas after 14 days all seeds had emerged, yielding fully developed plants containing shoots and roots. At the latter time point, no differences were observed across the treatments between the lengths of the roots or the aerial parts of the nascent leek plants. This included plants inoculated with each of the two Verrucomicrobium strains, the E. coli K12 E2 strain and the uninoculated plants (Table 1). The average length of the aerial parts ranged from 34.3 to 67.5 mm and the root lengths from 4.5 to 14.7 mm. Bacterial growth was never observed near the emerging leek plants grown from uninoculated seeds and also not at those inoculated with reference strain E. coli K12 E2. The latter observation indicated the absence of growth of this strain at the expense of root-exuded compounds. Both Verrucomicrobium subdivision 1 strains consistently colonized the roots of the leek plants. Luteolibacter sp. CHC12 colonized all 53 seeds that had emerged, whereas Candidatus genus Rhizospheria sp. CHC8 grew on all 51 emerged seeds. For both strains, visible growth at the root surface was noted. All bacterial cells retrieved from the root sites showed BOX-PCR profiles identical to the strains they derived from (data not shown), indicating that the growth along the roots indeed resulted from the introduced Luteolibacter sp. CHC12 or Candidatus genus Rhizospheria sp. CHC8. Because no carbon source had been added to the OLI medium and the single strains could not grow on this medium, the growth observed along the leek roots after 14 d was likely dependent on compounds that had been released from the roots. The patterns of root colonization observed differed between the two strains (Fig. 1), being that plants treated with the same strain always showed the same pattern of colonization. Strain CHC12 showed dense colonization of the maturation and elongation zones, but no colonization of the apical meristem, whereas CHC8 showed dense colonization of the maturation zone and the intersection between the aerial plant part and the root, but no colonization of the elongation zone and the apical meristem (Fig. 1B and 1C).
143 Rhizosphere competence of different Verrucomicrobium subdivision 1 strains
Table 1 Sizes of 14-days-old leek plants grown in oligotrophic agar (without amended carbon source). Initial bacterial densities were 103 cells per seed. Verrucomicrobia subdivision 1 strain Experiment Aerial part Root size Repetitiona Avarage Standard Avarage Standard deviation deviation Control A 44.5 11.6 9.37 2.81 B 46.9 13.7 8.84 3.91 Luteolibacter sp. CHC12 A 47.1 10.6 9.11 7.54 B 45.2 14.1 8.76 2.65 Candidatus genus Rhizospheria sp. A 54.6 8.4 6.92 3.26 CHC8 B 44.8 10.6 8.15 2.61
a Each repetition consisted of 30 different seeds
Survival of Luteolibacter sp. CHC12 and Candidatus genus Rhizospheria sp. CHC8 in Vredepeel bulk soil
One h after introduction of Luteolibacter sp. CHC12 into the sterile soil (level approx Log 5 g-1 dry soil), the cell equivalents per g dry soil determined via real time PCR were (Log-transformed values) between 5.81 and 6.07. Until the fourteenth day, these numbers increased to the range 7.17 - 7.62 and then remained virtually stable over the further monitoring period of 35 days (Fig. 2). Candidatus genus Rhizospheria sp. CHC8 showed stable cell equivalent numbers (Log-transformed values) of between 5.79 and 6.13 g-1 dry soil. From 3 to 7 days after introduction, these numbers increased to 6.95 - 7.27 g-1 dry soil. Subsequently, cell equivalent numbers of Candidatus genus Rhizospheria sp. strain CHC8 decreased to 6.46 - 6.86 until day 14, these numbers remaining stable over the entire monitoring period of 35 days
144 Chapter 7 strain CHC8. strain Rhizospheria genus cells of the different strains after 2 weeks of growth. 3 Candidatus ts inoculated with 10 strain CHC12; and (C) Luteolibacter Luteolibacter colonization of leek plan In vitro Figure 1 (A) Sterile control; (B)
145 Rhizosphere competence of different Verrucomicrobium subdivision 1 strains
7.75
A A A A a 7.00
b b b b B 6.25 BC
Cc C c c CHC8 CHC12
Log equivalent cell numberpergof drysoil 5.50 0 5 10 15 20 25 30 35 40
Days
Figure 2 Growth and survival of Luteolibacter strain CHC12 and Candidatus genus Rhizospheria strain CHC8 in sterilized soil. Sterile soils were inoculated at levels of 106 cells per gram of dry soil. Smoothed lines link the averages of replicate measured values, for CHC8 and for CHC12. Average values marked with different letters significantly differ between time point for the same strain, as determined by one-way ANOVA, where capital letter stands for strain CHC8 and small captions for strain CHC12.
Plant-soil microcosm experiments
Root and shoot development in the leek plants that emerged from surface-sterilized seeds became evident four days after the microcosm set-up. Dense root mats were observed after 3 weeks in all plant-soil microcosms. There was no difference in growth (evidenced by measuring shoot length and appearance) between the leek plants in the four different microcosm set-ups (A - D) (data not shown). Luteolibacter sp. CHC12. In the control microcosm (D, sterile soil only), the Luteolibacter cell equivalents per g dry soil in the 0-2 mm and 10-12 mm zones were at background levels, i.e. respectively, 2.20 - 2.44 and 2.16 - 2.71 (Log scale) (Fig. 3). In the A microcosms, in which the inoculated soil was in direct contact with gauze and root mat, the respective cell equivalents per g dry soil were (Log scale) between 6.08
146 Chapter 7
and 7.17 in the 0-2 mm zone, whereas these numbers were significantly higher in the 10-12 mm zone (P<0.05), i.e. between 7.38 and 7.65. In the B microcosms, in which the inoculated soil had been separated from the gauze and root mat by a 1 cm layer of uninoculated sterile soil, the Luteolibacter sp. CHC12 cell equivalents (Log scale) were at background level (between 2.36 and 2.83) in the 0-2 mm zones, and between 7.48 and 7.63 in the 10-12 mm zones. In the B2 microcosms (no root mat), the results were similar to those of the B microcosms, i.e. values (Log scale) between 2.19 and 2.74 cell equivalents per g dry soil were found in the 0-2 mm zones, whereas in the 10-12 mm zones, numbers between 7.35 and 7.67 were found. This indicates that Luteolibacter sp. CHC12 cells did not thrive in the soil close to the plant roots nor migrated towards the root mat. In the C microcosms, in which uninoculated non-sterile soil was used, the 0-2 mm zones contained (Log) 4.68 and 4.88 Luteolibacter cell equivalents per g dry soil, whereas in the 10-12 mm soil zone these numbers were between 5.08 and 5.12. Moreover, the averages of the natural population of Luteolibacter (C microcosm) were similar (P < 0.05) in rhizosphere and bulk soil (Fig. 3). Candidatus genus Rhizospheria sp. CHC8. In the control plant-soil microcosm D, with sterile soil only, the Candidatus genus Rhizospheria cell equivalents per g of dry soil in the 0-2 mm and 10-12 mm zones were at background level, i.e. (Log values) 3.34 - 4.04 and 3.39 - 4.12 (Fig. 3). In the A microcosms, the respective cell equivalents per g dry soil were between 7.57 and 7.70 in the 0-2 mm zone, whereas they were significantly lower in the 10-12 mm zone, i.e. 6.30 - 6.93. In the B microcosms, these numbers (Log) were 4.32 - 4.71 in the 0-2 mm zones, and 6.64 - 7.09 in the 10-12 mm zones. In the B2 microcosms (no plants), we found (Log) 3.42 - 3.95 in the 0-2 mm zones and 6.17 - 6.86 in the 10-12 mm zones. These data indicate that Candidatus genus Rhizospheria CHC8 occurred in significantly raised numbers in the soil layers closest to the root mats, indicating local growth. Also, considerable numbers of the organism were shown to occupy these layers following migration towards the root mat. In the C microcosms (uninoculated non-sterile soil), the 0-2 mm zones contained between (Log) 7.25 and 7.30 Candidatus genus Rhizospheria cell equivalents per g dry soil, whereas in the 10-12 mm soil zones these numbers were also lower, i.e. between 6.93 and 7.03 (the averages of the 0-2 mm and 10-12 mm zones were considered to be significantly different by ANOVA, P<0.05) (Fig. 3).
147 Rhizosphere competence of different Verrucomicrobium subdivision 1 strains s n
genu ns ns Rhizospheria Candidatus D 0-2 mm soil zone 10-12 mm soil zo strain CHC8 with strain ns ns il that was in closest il that Luteolibacter tly in one-way ANOVA; erilized soil inoculated with oots. (C) Non-inoculated non- Rhizospheria genus ** ** genus Rhizospheria Candidatus C of P<0.05 and P<0.01. Arrow in B indicate non-inoculated sterile so non-inoculated species in soil layers proximate to (0-2 mm), proximate in soil layers species ns ns do not differ significan do Candidatus Luteolibacter ns strain CHC8 in direct with the nylon gauze with leek with leek with the nylon gauze strain CHC8 in direct Rhizospheria nd the nylon gauze with leek r Kuchenbuch microcosms. (A) St ** ** CHC8 genus strain CHC12 and Rhizospheria B genus ** ** Candidatus Luteolibacter Luteolibacter and ffer significantly, respectively at levels erile soil. Averages marked with with marked soil. Averages erile Candidatus ** ** subdivision 1 cells through the 1 cm layer subdivision 1 cells of CHC8 Luteolibacter Luteolibacter A
* * CHC12 CHC12 strain CHC12 and Verrucomicrobia Distribution of 8 7 6 5 4 3 2 celleq numberLog per g of contact with the roots. and at a further distance (10-12 mm) from leek roots in Luteolibacter a 1 cm non-inoculated soil layer between inoculated soils a st soil. (D) Non-inoculated sterile averages with one or two asterisks di migration of Figure 3 roots. (B) Sterilized soil inoculated with soil inoculated roots. (B) Sterilized
148 Chapter 7
Discussion
In this chapter it was hypothesized that Luteolibacter sp. CHC12 and Candidatus genus Rhizospheria CHC8, which were recently isolated from the leek rhizosphere, are rhizosphere competent, i.e. they show a growth/migration response to leek roots. From the in vitro experiments that were conducted, it became evident that both Verrucomicrobium subdivision 1 strains could grow on plates on which leek plants developed from seeds, whereas no growth was observed on similar plates without plants. This indicated the utilization of nutrients released by the leek roots in the form of exudates or even sloughed-off cells. The in vitro colonization of, and growth on, sterilized leek roots corroborates findings made before with strain CHC8 and contradict those found for strain CHC12 (Nunes da Rocha et al., 2010a). Candidatus genus Rhizospheria CHC8 was able to grow with oxalic, malic, citric or succinic acid, as well as glutamine and alanine as the sole carbon sources. These compounds have been considered as ‘common’ for the rhizospheres of different plant species (Baudoin et al., 2003; Jones, 1998) and hence strain CHC8 clearly has the potential to occupy niches proximate to roots that exude such compounds. On the other hand, strain CHC12 did not grow on any of the organic or amino acids tested (Nunes da Rocha et al. 2010a), i.e. those in which strain CHC8 was able to grow. These clear findings are consistent with the hypothesis that, although closely related in their 16S rRNA gene, distinct strains of Verrucomicrobium subdivision 1 have different ecological roles in the plant-soil system. If cells of these strains occur in the neighborhood of leek roots then questions arise as to whether they will behave in ways similar to typical rhizosphere bacteria. In other words: will they be able to survive in the complex soil matrix in the rhizosphere (Berg and Smalla, 2009) and, are they able to compete with other soil microorganisms in there (Bueé et al., 2009)? Strikingly, contrasting behavioral patterns were observed for rhizosphere colonization of Luteolibacter sp. CHC12 and Candidatus genus Rhizospheria sp. CHC8 in sterilized soil and of endogenous Luteolibacter and Candidatus genus Rhizospheria in non-sterile soils. The estimated average numbers of Candidatus genus Rhizospheria sp. CHC8 in rhizosphere soil (0-2 mm layer) were approximately 1 Log unit per g dry soil higher than those found in the respective bulk soil (10-12 mm layer) in the Kuchenbuch system. A similar pattern was observed for the natural populations of Candidatus genus Rhizospheria, which had increased in the rhizosphere soil in comparison to the respective bulk soil. In contrast, the average numbers of Luteolibacter sp. CHC12 were approximately 1 Log unit per g dry soil lower in the rhizosphere (0-2 mm) than in the respective bulk soil (10-12 mm). The data shown in this study demonstrate that care should be taken when higher taxonomical rankings (e.g. phylum, class, order) are used to analyze the distribution of
149 Rhizosphere competence of different Verrucomicrobium subdivision 1 strains
bacterial groups over soil compartments. Especially when determining correlations between high-ranking taxonomical groups and environmental factors. For instance, if instead of discriminating among the different Verrucomicrobia subdivision 1 groups, the data of the current study would have sum up all the Verrucomicrobia subdivision 1, the distinct behavior observed for Luteolibacter and Candidatus genus Rhizospheria would have been overlooked. Similar observations have been made in Nunes da Rocha et al. (2010b), where it was demonstrated that a minor group of Acidobacteria, i.e. Holophagae, is rhizosphere competent while, in general, Acidobacteria have been characterized as normal inhabitants of bulk soil (Kielak et al., 2008). The estimated Candidatus genus Rhizospheria numbers found in the rhizosphere (expressed as number of cell equivalents per g of dry soil) were about 107. This number is approximately at the same level as measured Pseudomonas fluorescens CFU or cell numbers in different rhizospheres (Van Elsas et al., 1992; Hase et al., 2000). This indicates that Candidatus genus Rhizospheria members colonize the rhizosphere environment with approximately the same numbers as other rhizosphere competent bacteria, such as Pseudomonas spp. Moreover, cells of strain CHC8 had migrated towards the roots, thereby indicating chemotaxis as a likely mechanism involved with rhizosphere competence. Apparently, Candidatus genus Rhizospheria cells follow the nutrient gradient to the roots upwards and migratory activity towards plant roots can be considered as a first step in root colonization (De Weert et al., 2002; Rudrappa et al., 2008). Collectively, our observations demonstrate clear rhizosphere competence for Candidatus genus Rhizospheria sp. strain CHC8. On the other hand, as estimated numbers of Luteolibacter sp. strain CHC12 or natural populations of Luteolibacter did not increase in rhizosphere soil, this group seems to prefer bulk soil over rhizosphere, at least at the tested conditions. Thus, it was demonstrated that Candidatus genus Rhizospheria thrive in soil proximate to plant roots, but Luteolibacter do not. Recently, soil type was shown as one of the strongest influences on the Verrucomicrobia diversity (Kuramae et al., 2010). Therefore, the contradictory reports about distribution of Verrucomicrobia over bulk and rhizosphere soil (Sanguin et al., 2006; Kielak et al., 2008) could occur due to differences in the autochthones populations of Verrucomicrobia in the different soil types studied. Another factor that may influence this phenomenon is the differential selection of Verrucomicrobia groups according to plant type (Hartmann et al., 2009; Berg & Smalla, 2009). Further studies are necessary to elucidate if specific populations of Verrucomicrobia are differentially selected by soil type and plant species and other factors, such as pH or plant growth stage that may be involved in the rhizosphere competence of this group. In conclusion, Candidatus genus Rhizospheria sp. CHC8 recently cultured from the leek rhizosphere are indeed rhizosphere competent. On the other hand, Luteolibacter
150 Chapter 7
sp. CHC12 seems to be more adapted to survive in bulk soil. The last contradicts what is generally accepted for Verrucomicrobia, i.e. that these groups prefer rhizosphere over bulk soil. With the differentiation of Candidatus genus Rhizospheria and Luteolibacter groups of Verrucomicrobia subdivision 1 it was possible to draw firm conclusions about strain behaviour in the rhizosphere. Furthermore, qPCR-based assessment of these groups made it possible to perform assays that would be virtually ‘impossible’ with culture-dependent techniques alone. The combination of culture-dependent and independent techniques for studying members of Verrucomicrobia in the plant-soil system opens new avenues to explore minority groups within their natural settings.
Acknowledgements This research was part of the Ecogenomics program which is sponsored by the Dutch National Genomics Initiative and the basic research program on sustainable agriculture (KB4) sponsored by the Dutch ministry of agriculture, nature and food safety. We would like to thank Pieter Kastelein and Vladimir Fediy for their assistance in the plant- soil microcosm experiments.
151 Rhizosphere competence of different Verrucomicrobium subdivision 1 strains
References
Baudoin E, Benizri E & Gauckert A (2003). Impact of artificial root exudates on the bacterial community structure in bulk soil and maize rhizosphere. Soil Biol Biochem 35: 1183-1192. Berg G & Smalla K (2009). Plant species and soil type cooperatively shape the structure and function of microbial communities in the rhizosphere. FEMS Microbiol Ecol 68: 1-13. Buée M, De Boer W, Martin F, van Overbeek L & Jurkevitch E (2009). The rhizosphere zoo: An overview of plant-associated communities of microorganisms, including phages, bacteria, archaea, and fungi, and of some of their structuring factors. Plant Soil 321: 189-212. Chow ML, Radomski CC, McDermott JM, Davies J & Axelrood PE (2002) Molecular characterization of bacterial diversity in Lodgepole pine (Pinus contorta) rhizosphere soils from British Columbia forest soils differing in disturbance and geographic source. FEMS Microbiol Ecol 42: 347-357. da Rocha UN, Andreote FD, de Azevedo JL, van Elsas JD & van Overbeek LS (2010). Cultivation of hitherto-uncultured bacteria belonging to the Verrucomicrobia subdivision 1 from the potato (Solanum tuberosum L.) rhizosphere. J Soils Sediments 10: 326-339. De Weert S, Vermeiren H, Mulders IHM, Kuiper I, Hendrickx N, et al. (2002). Flagella-driven chemotaxis towards exudate components is an important trait for tomato root colonization by Pseudomonas fluorescens. Mol Plant-Microbe Interact 15: 1173-1180. Dijkstra AF, Govaert JM, Scholten GHN & Van Elsas JD (1987). A soil chamber for studying the bacterial distribution in the vicinity of roots. Soil Biol Biochem 19: 351-352. Frey JC, Rothman JM, Pell AN, Nizeyi JB, Cranfield MR & Angert ER (2006) Fecal bacterial diversity in a wild gorilla. Appl Environ Microbiol 72: 3788-3792. Garrity M & Holt JG (2001). The road map to the manual. In: D.R. Boone and R.W. Castenholz. Bergey’s Manual of Systematic Bacteriology, 2nd ed. Springer-Verlag. New York, NY. 119-162. Goh SHM, Mabbett AN, Welch JP, Hall SJ & McEwan AG (2009) Molecular ecology of a facultative swine waste lagoon. Lett Appl Microbiol 48: 486-492. Hase C, Hottinger M, Moënne-Loccoz y & Defago G (2000). Survival and cell culturability of biocontrol Pseudomonas fluorescens CHAO in the rhizosphere of cucumber grown in two soils of contrasting fertility status. Biol Fertil Soils 32: 217-221. Hartmann A, Schmid M, van Tuinen D & Berg G (2009). Plant-driven selection of microbies. Plant Soil 321: 235-257. Hedlund BP, Gosink JJ & Staley JT (1997). Verrucomicrobia div. nov., a new division of the bacteria containing three new species of Prosthecobacter. Antonie Van Leeuwenhoek 72: 29-38. Herrmann S, Kleinsteuber S, Neu TR, Richnow HH & Vogt C (2008) Enrichment of anaerobic benzene- degrading microorganisms by in situ microcosms. FEMS Microbiol Ecol 63: 94-106. Hugenholtz P, Goebel BM & Pace NR (1998) Impact of culture-independent studies on the emerging phylogenetic view of bacterial diversity. J Bacteriol 180: 4765-4774. Jones DL (1998). Organic acids in the rhizosphere – a critical review. Plant Soil 205: 25-44. Kielak A, Pijl AS, Van Veen JA & Kowalchuk GA (2008). Differences in vegetation composition and plant species identity lead to only minor changes in soil-borne microbial communities in a former arable field. FEMS Microbiol Ecol 63: 372-382. Kuchenbuch R & Jungk A (1982). A method for determining concentration profiles at the soil-root interface by thin slicing rhizospheric soil. Plant Soil 68: 391-394. Kuramae EE, Gamper HA, Yergeau E, Piceno YM, Brodie EL et al. (2010) Microbial secondary succession in a chronosequence of chalk grasslands. ISME J 4: 711-715. Lópes-Bucio J, Cruz-Ramírez A, Pérez-Torres A., Ramírez-Pimente JG, Sánchez-Calderón L & Herrera- Estrella L (2005). Root Architecture. In: Turnbull CGN (Ed.), Plant Architecture and its manipulation, Blackwell Publishing, Oxford, UK. Chapter 7.4.1 pp. 189-191. Lugtenberg B & Kamilova F (2009). Plant-growth-promoting rhizobacteria. Annu Rev Microbiol 63: 541- 556. Lymperopoulou DS, Kormas KAr, Moustaka-Gouni M & Karagouni AD (2010) Diversity of cyanobacterial phylotypes in a Mediterranean drinking water reservoir (Marathonas, Greece). Environ Monit Assess, DOI: 10.1007/s10661-010-1378-7. Nunes da Rocha U, Andreoti FD, Plugge C, Ausek L, van Elsas JD & Overbeek L (2010a). Isolation and partial characterization of Holophagae, Luteolibacter, unclassified Verrucomicrobia and Verrucomicrobium from the leek (Allium porrum) rhizosphere. Submitted.
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Nunes da Rocha U., van Elsas JD & Overbeek L (2010b). Rhizocompetence of culturable Holophaga (Acidobacteria) sp. in the leek (Allium porrum) rhizosphere. Submitted. Nunes da Rocha U., van Elsas JD & Overbeek L (2010c). Real-time PCR detection of Holophagae (Acidobacteria), Luteolibacter and unclassified Verrucomicrobiaceae (Verrucomicrobia subdivision 1) in bulk and leek (Allium porrum) rhizosphere soils. Submitted. Nunes Da Rocha U, Van Overbeek L & Van Elsas JD (2009). Exploration of hitherto-uncultured bacteria from the rhizosphere. FEMS Microbiol Ecol 69: 313-328. Rademaker JLW, Louws FJ & De Bruijn FJ (1998). Characterization of the diversity of ecologically important microbes by rep-PCR genomic fingerprinting. Molecular Microbial Ecology Manual (SUPPL. 3), pp. 1-26. Rosenberg K, Bertaux J, Krome K, Hartmann A, Scheu S & Bonkowski M (2009) Soil amoebae rapidly change bacterial community composition in the rhizosphere of Arabidopsis thaliana. ISME J 3: 675- 684. Rudrappa T, Czymmek KJ, Paré PW & Bais HP (2008). Root-secreted malic acid recruits beneficial soil bacteria. Plant Physiol 148: 1547-1556. Sanguin H, Remenant B, Dechesne A, Thioulouse J et al. (2006) Potential of a 16S rRNA-based taxonomic microarray for analyzing the rhizosphere effects of maize on Agrobacterium spp. and bacterial communities. Appl Environ Microbiol 72: 4302-4312. Schlesner H, Jenkins C & Staley JT (2006) The Phylum Verrucomicrobia: A Phylogenetically Heterogeneous Bacterial Group. Prokaryotes 7: 881-896. Van Elsas JD, Trevors JT, Jain D, Wolters AC, Heijnen CE & van Overbeek LS (1992). Survival of, and root colonization by, alginate-encapsulated Pseudomonas fluorescens cells following introduction into soil. Biol Feril Soils 14: 14-22. Van Elsas JD, Trevors JT & Starodub ME (1988). Bacterial conjugation between pseudomonads in the rhizosphere of wheat. FEMS Microbiol Ecol 53: 299-306. Van Overbeek LS & Van Elsas JD (1995). Root exudate-induced promoter activity in Pseudomonas fluorescens mutants in the wheat rhizosphere. Appl Environ Microbiol 61: 889-898. , Jeppsson B & Molin G (2005) Comparison of bacterial diversity along the human intestinal tract by direct cloning and sequencing of 16S rRNA genes. FEMS Microbiol Ecol 54: 219- 231. Zul D, Denzel S, Kotz A & Overmann J (2007) Effects of plant biomass, plant diversity, and water content on bacterial communities in soil lysimeters: Implications for the determinants of bacterial diversity. Appl Environ Microbiol 73: 6916-6929.
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Chapter 8: Distribution of different Acidobacteria and Verrucomicrobia subdivision 1 groups over compartments of different plant species*
Abstract
Contradictory information about the numerical dominance of Acidobacteria and Verrucomicrobia over bulk and rhizosphere soil is found in literature. In a field experiment, potato and leek were planted in a former pasture land. Rhizosphere and bulk soil were sampled at different plant growth stages. The numbers of Acidobacteria subgroups 1, 3, 4 and 6, and Holophaga, Luteolibacter and Candidatus genus Rhizospheria were assessed by qPCR. Multivariate analyses were performed to determine which populations were affected by the rhizosphere and which factors influence their distribution. It was found that different subgroups of the Acidobacteria were differentially influenced by the rhizosphere. The numbers of subgroup 6 Acidobacteria mainly increased in the rhizosphere compartments as compared to those in bulk soil, whereas Holophaga numbers increased only in the leek rhizosphere. The numbers of Acidobacteria subgroups 1 and 4 occasionally increased in the rhizosphere, but Acidobacteria subgroup 3 estimated cell numbers were grossly lower in the rhizosphere than in the respective bulk soil. Differently from the Acidobacteria, Luteolibacter and Candidatus genus Rhizospheria numbers were higher in rhizosphere compartments than in their respective bulk soil independently of plant type.
* Authored by: Ulisses Nunes da Rocha, Jan Dirk van Elsas, Isabelle George, Spiros Nicolas Agathos & Leonard Simon van Overbeek Submitted for publication Chapter 8
Introduction
Acidobacteria and Verrucomicrobia often make part of the dominant bacterial groups in soil, contributing (grossly) between 20-52% (Janssen, 2006; Sait et al., 2002) and between 1-10% (Sangwan et al., 2005) of the 16S rRNA gene diversity present in the soil environment. These bacterial groups are not only dominant, but also diverse. Twenty six different phylogenetic groups have been assigned to the Acidobacteria (Barns et al., 2007), whereas the Verrucomicrobia are separated in seven different subdivisions (Schlesner et al., 2006). Despite their ubiquity and abundance, our actual knowledge about the ecology and, especially, the ecological niches of these bacterial groups in plant-soil systems are far from being understood. Regardless of the fact that members of the Acidobacteria and Verrucomicrobia are considered to be ‘hard-to-culture’ (Jones et al., 2009; Nunes da Rocha et al., 2010a), the numbers of cultured strains of both groups have steadily increased in recent years. The Ribosomal Database Project (RDP Release 10, Update 20, http://rdp.cme.msu.edu/) now contains 210 and 137 different sequences of cultured Acidobacteria and Verrucomicrobia, respectively. Mostly, this increase has been achieved with simple adaptation of traditional isolation methods, e.g. the use of low- carbon-availability media and long incubation periods (Jansen et al., 2002). Using a similar approach, different strains that affiliated with subgroups 1, 3, 4, and 6 of the Acidobacteria have recently been isolated from bulk soil (George et al., 2010). Also, Acidobacteria affiliating with Holophaga spp and members of Verrucomicrobium subdivision 1 have been cultured from rhizospheres (Nunes da Rocha et al., 2010a; Nunes da Rocha et al., 2010b). Due to the fact that most Acidobacteria and Verrucomicrobia are slow growers, being very fastidious in their growth under laboratory conditions, ecological studies with them using culturing techniques alone are difficult to perform. Plant-soil microcosm studies in which sterile soil had been inoculated with Holophaga sp. CHC25, Luteolibacter sp. CHC12 and Candidatus genus Rhizospheria sp. CHC8 (subdivision 1 of Verrucomicrobia) demonstrated that both the Holophaga and Candidatus genus Rhizospheria strains were competent in the rhizosphere of leek. Furthermore, Luteolibacter showed preference for bulk over rhizosphere soil under microcosm conditions (Nunes da Rocha et al., 2010c; Nunes da Rocha et al., 2010e). These conclusions were achieved by estimating cell numbers using group-specific real- time PCR detection (Nunes da Rocha et al., 2010d), being this technique preferred as it is most straightforward. Although it is generally accepted that the Acidobacteria are best adapted to life in bulk soil (Fierer et al., 2007) and Verrucomicrobia to that in the rhizosphere (Nunes da Rocha et al., 2009), contradictory information has been provided on the occurrence
155 Distribution of different Acidobacteria and Verrucomicrobia in plant-soil systems
of members of the two phyla in rhizosphere and bulk soils (Chow et al., 2002; Sanguin et al., 2006; Zul et al., 2007; Kielak et al., 2008). These conclusions, however, were based on analyses of the entire phyla, not paying attention to the different subgroups in these. In the current chapter, it was hypothesized that distinct groups of Acidobacteria and Verrucomicrobia would demonstrate different ecological behavior in plant-soil ecosystems. To evaluate this hypothesis, a field experiment with potato, leek and permanent grass was performed. Samples were taken at different plant growth stages and two different rhizosphere compartments, the outer and inner rhizosphere, and their respective bulk soil were analyzed. Acidobacteria subgroups 1, 3, 4 and 6, i.e. the most dominant subgroups of the Acidobacteria (Jones et al., 2009; Rousk et al., 2010), Holophaga sp., and two different Verrucomicrobium subdivision 1 groups previously cultured from the rhizosphere (Nunes da Rocha et al., 2010a; Nunes da Rocha et al., 2010b), were separately quantified by real-time PCR. To achieve this aim, novel real- time PCR systems were designed and evaluated.
Material and Methods
Bacterial strains and routine cultivation
Strains belonging to Acidobacterium groups 1, 3, 4 and 6 and Holophaga (group 8) and to Verrucomicrobium subdivision 1 subgroups Luteolibacter and Candidatus genus Rhizospheria were described in Nunes da Rocha et al. (2010a; 2010b) and George et al. (2010) (Table 1). R2A (Difco, France) was used for routine cultivation of all Holophaga Luteolibacter and Candidatus genus Rhizospheria strains at 25° C (Nunes da Rocha et al., 2010b), whereas for routine cultivation of Acidobacterium groups 1, 3, 4 and 6, 200- fold diluted nutrient agar was used (George et al., 2010).
DNA extraction from pure cultures
To extract DNA from all pure strains, cells were scraped from R2A or 200-fold diluted nutrient agar plates with bacterial growth. The cells were suspended in 500 µL of a sterile solution consisting of 0.85% KCl in DNase/RNase-free distilled water (Invitrogen, The Netherlands). Subsequently, DNA was extracted from these cell suspensions using the MasterPureTM DNA purification kit (Epicentre Biotechnologies, WI, USA) following the instructions provided by the manufacturer.
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Table 1 Acidobacteria and Verrucomicrobia isolated from bulk or rhizosphere soil samples used to validate qPCR primers. Affiliation Strain Isolation source Year of Field/collection Reference Soil Soil isolation type compartment Acidobacteria Subgroup 1 IGE012 sandy Surface bulk 2007 Louvain-la- George et al. silt soil Neuve/BE (2010) Subgroup 3 IGE015 sandy Surface bulk 2007 Louvain-la- George et al. silt soil Neuve/BE (2010) Subgroup 4 IGE017 sandy Surface bulk 2007 Louvain-la- George et al. silt soil Neuve/BE (2010) Subgroup 6 IGE001 sandy Surface bulk 2007 Louvain-la- George et al. silt soil Neuve/BE (2010) Holophagae CHC25 sand Leek 2007 Vredepeel/NL Nunes da Rocha rhizosphere et al. (2010b) ORAC sand Leek 2008 Vredepeel/NL Nunes da Rocha rhizosphere et al. (2010b) Verrucomicrobia Luteolibacter C20 loamy Potato 2006 Droevendaal/NL Nunes da Rocha sand rhizosphere et al. (2010a) CHC12 sand Leek 2007 Vredepeel/NL Nunes da Rocha rhizosphere et al. (2010b) ONA9 sand Leek 2007 Vredepeel/NL Nunes da Rocha rhizosphere et al. (2010b) Candiatus CR28 loamy Potato 2006 Droevendaal/NL Nunes da Rocha genus sand rhizosphere et al. (2010a) Rhizospheria Z35 loamy Potato 2006 Droevendaal/NL Nunes da Rocha sand rhizosphere et al. (2010a) ZNBB5 loamy Potato 2006 Droevendaal/NL Nunes da Rocha sand rhizosphere et al. (2010a) CHC8 sand Leek 2007 Vredepeel/NL Nunes da Rocha rhizosphere et al. (2010b) IRVE sand Leek 2008 Vredepeel/NL Nunes da Rocha rhizosphere et al. (2010b)
Design, evaluation and calibration of Acidobacterium group 1, 3, 4 and 6-specific qPCR systems
Primers specific for the Acidobacterium groups 1, 3, 4 and 6 were designed and evaluated according to the procedure described for Holophaga, Luteolibacter and Candidatus genus Rhizospheria (Nunes da Rocha et al. 2010d). In short, primers were validated in three steps. First, for each acidobacterial group, alignments were made between sequences of the almost- entire16S rRNA gene and those of strains from the same and related bacterial group retrieved from the SILVA database, release 102 (Pruesse et al., 2007). Following this, primers were designed based on conserved sequences specific for each group and checked in silico for the absence of formation of misprimed products using Primer-BLAST software (http://www.ncbi.nlm.nih.gov/tools/primer-blast/). In the second step, specificity of the
157 Distribution of different Acidobacteria and Verrucomicrobia in plant-soil systems
designed primer sets was tested using DNA from pure cultures of target (Table 1) and non-target strains by PCR amplification. As non-target strains, all non-corresponding Acidobacteria and Verrucomicrobia subdivision 1 strains were chosen, supplemented with Agroabcterium tumefaciens UBAPF2 (Alphaproteobacteria), Burkholderia cepacia LMG 1222T (Betaproteobacteria), Escherichia coli E1 (Gammaproteobacteria), Streptomyces griseus IPO 857 (Actinobacteria), Flavobacterium columnar 2003/035 (Bacteroidetes) and Bacillus subtilis Bs4 (Firmicutes). All these strains were derived from the strain collection of Plant Research International (Wageningen, The Netherlands). Standard and real-time (q)PCRs were run under the same conditions, as follows. Each 25-µl reaction mixture contained the following ingredients: 12.5 µL of SYBR Premix Ex Taq 2x (TAKARA Bio Inc., Japan), 0.5 µL of each primer (10 µM; Biolegio,
NL), 0.5 µL of ROX Reference Dye II 50x (TAKARA Bio Inc., Japan), 6.0 µL H2O and 5.0 µL template DNA (containing 1 ng of target and non-corresponding strains). All PCR reactions were run for one cycle at 50 oC for 2 min; one cycle at 95 oC for 10 s; 35 cycles at 95 oC for 5 s and 60 oC for 35 s. The number of cycles was limited to 35 to avoid formation of false positives products in PCRs (Sipos et al., 2007). Amplicons made with the four primer sets in standard PCR runs were routinely checked for the expected sizes and the absence of any other products by agarose (1.5%) gel electrophoresis. Additionally, melting curves derived from each individual reaction were routinely inspected for the absence of primer dimers or any other artifacts resulting from unintended amplifications. Calibration of the qPCR systems that aimed to detect Acidobacteria groups 1, 3, 4 and 6 was the same as for those aimed to detect Holophaga and Verrucomicrobium subdivision 1 groups Luteolibacter and Candidatus genus Rhizospheria, as described before (Nunes da Rocha et al. 2010d). In short, standard curves based on ranges between 10 and 108 cells per qPCR reaction mixture were made and run with the appropriate primers. The curves were made in triplate and the measured threshold cycle (Ct) values were plotted against the Log cell number for each system. Line slopes and intercepts were calculated by regression analysis using GenStat 12th edition (VSN International Ltd., UK). The amplification efficiency (Ae) of the different primer systems was calculated using the formula Ae = 10(-1/slope), in which ‘slope’ represents the slope value calculated by regression analysis. Bacterial qPCR with primers Eub338 (Lane, 1991) and Eub518 (Muyzer et al., 1993) were used to estimate the bacterial numbers in soil DNA extracts. This system was calibrated with Escherichia coli E1 cells, and qPCRs were performed according to the procedure described in Fierer et al. (2005). Theoretical dynamic ranges for all qPCR systems were determined according to Nunes da Rocha et al. (2010d).
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Field site, soil and plant sampling, sample processing and analyses procedures
The study site was a field of the experimental farm Droevendaal, Wageningen, The Netherlands (51q59’N, 5q39’E). The soil was a loamy sand containing 2% organic matter, a water holding capacity of 25% and a pH (KCl) of 4.8. The experiment was conducted in the summer 2009 and before onset of the experiment the field was covered with permanent grassland (commertial mix, containing Lolium perenne as the main plant species, Salles et al.; 2006). This field was divided into four plots of 4 x 25 m, with distances of 1.5 m between neighbouring plots, and permanent grassland was removed from three of the four plots. Subsequently, two fallow plots were planted with either potato (Solanum tuberosum L. cultivar Agria) or leek (Allium porrum cultivar Kenton, Nunhems Seeds BV, The Netherlands), whereas one plot remained fallow and the forth one was kept as permanent grassland. Potato seed tubers and nursery leek plants were planted in May 2009, according to agricultural management schedules common for production of these crops in the Netherlands. Before and during experimentation, the field was maintained under organic agricultural practices, i.e. no pesticides or chemical fertilizers were applied to the field and during the experimental period weed plants were removed by hand. Samples from each plot were taken in June (month 1), July (month 2) and September (month 4). During sampling, one sample was taken from each plot for each treatment, totaling four replicates per treatment. Samples from the potato and leek fields were taken as individual plants, whereas those from the grass fields consisted of plant cover with roots of 10 cm in size. Samples taken from the fallow plot consisted of 10 g portions from the 0-5 cm soil horizons. All samples were directly processed in the laboratory, where soil loosely adhering to the roots was removed by mild manual shaking. ‘Outer rhizosphere’ was represented by the soil adhering to the root surface after mild manual shaking. It was removed from the roots by scratching with a spatula. Then, roots devoid of soil aggregates were removed from the flasks, placed in other flasks and again shaken in 0.1% sodium pyrophosphate solution (1:10 v/v ratio). The resulting soil suspensions were considered to represent the ‘inner rhizosphere’. The inner rhizosphere suspensions were concentrated by centrifugation at 10,000 x g for 15 min followed by resuspension of the pellet in 1 ml of 0.85% NaCl solution. Bulk soil samples were processed according to the procedure applied for outer rhizosphere soil.
Soil pH was measured in all (bulk, inner and outer rhizosphere) soils in 0.01 M CaCl2 (1: 10 w/v ratio) according to the procedure of Houba & Novozamsky (1998). DNA was extracted from all soils using the PowerSoil Isolation Kit (MO BIO Laboratories, Inc., CA, USA) following the instructions provided by the manufacturer. For bacteria, group 1, 3, 4, 6 Acidobacteria and Holophaga and Verrucomicrobium subdivision 1 groups Luteolibacter and Candidatus genus
159 Distribution of different Acidobacteria and Verrucomicrobia in plant-soil systems
Rhizospheria qPCRs, 25 PL reaction mixtures containing 5 µL of diluted (at least 10- fold) soil DNA (approximately 5 ng) were applied, using the amplification conditions described before. All reactions were carried out in triplicate, and one positive (target DNA added) and two negative controls (one including DNA from non-target strains – not for total bacteria qPCR- and one with sterile demineralized water) were included for each primer system.
Statistics and multivariate analyses
Comparisons were made between the average Log equivalent cell numbers (derived from the estimated target gene numbers calibrated with cell from the respective target strains of each group(Table 1) and expressed per g dry soil) in three soil compartments (bulk, inner and outer rhizosphere) and three sampling periods. The analysis was performed for each of the eight primer systems, using analysis of variance (ANOVA) (GenStat 12th edition, VSN International Ltd., UK). Multivariate analysis (CANOCO for Windows version 4.55, Biometris, Plant Research International, The Netherlands) was performed on all soil samples using soil localization, sampling time, plant species and pH as the ‘environmental’ variables and Log equivalent cell numbers calculated for each of the eight qPCR systems as ‘species’ variables. The relationships between environmental and species variables were calculated by detrended correspondence analysis (DCA) in a first and by redundancy analysis (RDA) in a second step. Community similarities were placed in ordination plots with scaling focused on inter- sample differences (Marschner & Baumann, 2003).
Nucleotide sequence accession numbers
Non-redundant sequences of 192 partial 16S rRNA gene library clones constructed with the primer systems designed for Acidobacterium subgroups 1, 3, 4 and 6 (48 clones each) were deposited in the EMBL Nucleotide Sequence Database and are available under accession numbers FN994868 to FN994889.
Results
Specificity of Acidobacterium subgroups 1, 3, 4 and 6 qPCR primer systems
Of the seven qPCR primer systems applied to enumerate different subgroups of the Acidobacteria and Verrucomicrobia in rhizosphere soil, four are newly described in this study (Table 2). These primer sets aimed to specifically detect 16S rRNA gene
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Reference Lane (1991) (1991) Lane This chapter chapter This chapter This chapter This chapter This chapter This chapter This chapter This chapter This Muyzer et al. (1993) (1993) al. et Muyzer Nunes da Rocha et al. 2010 2010 al. et da Rocha Nunes 2010 al. et da Rocha Nunes 2010 al. et da Rocha Nunes 2010 al. et da Rocha Nunes 2010 al. et da Rocha Nunes 2010 al. et da Rocha Nunes
c rate Ct values are obtained. are obtained. rate Ct values 9.17 9.17 9.21 7.28 8.74 8.42 7.54 8.26 8.45 8.45 ver which these system are able to Dr b Ae ; where, Ae, amplification efficiency, amplification efficiency, ; Ae, where, ) a (-1/slope) Amplicon Amplicon length (bp concentrations accu over which C) o Tm Tm ( 1 qPCR primers used in this study. Melting temperature temperature Melting in this study. used 1 qPCR primers name Verrucomicrobia - the range of initial of - the range template and the dynamic range was grossly estimated to indicate o estimated range the dynamic the range grossly was Acidobacteria Sense Sense – 3’) (5’ sequence Primer Primer Reverse ATTACCGCGGCTGCTGG Eub518 54.4 54.4 ATTACCGCGGCTGCTGG Reverse Eub518 Reverse CCTTTGAGTTTCAGCCTTGC 60.0 Acg1r Reverse AGGAATTCCGCTTTCCTCTC 59.8 Acg3r Reverse CGCTGCATTATGCGGTATTA Acg4r 59.7 Reverse GTCCCGTTCGACAGGAGTT 60.1 Acg6r Reverse AGTCTCGGATGCAGTTCCTG Acg8r 60.4 Forward ACTCCTACGGGAGGCAGCAG ACTCCTACGGGAGGCAGCAG Forward Eub338 57.6 200 to 1.91 CAGGTACCCAATCCTGTCGT Forward 4.17 Acg1f 59.8 to 83 TAGGCGGTTGGGTAAGTTTGForward 1.95 4.21 Acg3f 60.0 to 100 GCACGGGTGAGTAACACGTAAForward 1.92 4.28 Acg4f 61.0 GAGGTAATGGCTCACCAAGGForward 86 Acg6f 59.6 to 1.92 193 to 3.74 TGGGATGTTGATGGTGAAAC Forward 1.92 Acg8f 4.42 59.2 to 470 CAGCTCGTGTCGTGAGATGT Forward 2.01 VS1Af 60.0 2.54 to 199 1.98 2.26 Forward GCCCGACAGGGTTGATAGTA GCCCGACAGGGTTGATAGTA Forward VS1Bf 60.0 83 to 1.95 2.45 genus A description of the group-specific Target group group Target groups All Subgroup 1 Subgroup 3 Subgroup 4 Subgroup 6 Holophagae Luteolibacter Candidatus Rhizospheria
= 10 equation: the Ae reaction offollowing calculated the was by efficiency. amplification The efficiency Ae, pairs. bp, base Theoretical (here, range cell per dry of Log dynamic gram soil) slope, mathematically calculated slope of standard curve curve standard of slope calculated mathematically slope, (Tm), approximate amplicon length theoretical dynamic range (Dr) and amplification efficiency of qPCR systems. qPCR of systems. efficiency amplification (Dr) and range dynamic theoretical length amplicon (Tm), approximate Table 2 Table a b c Assuming 100% efficiency of DNA extraction and of qPCR extraction of DNA efficiency 100% reaction Assuming in soil target groups detect the
Reverse TCTCGGTTCTCATTGTGCTG VS1Ar 60.0 60.0 60.06 VS1Ar VS1Br Taxa Bacteria Acidobacteria TCTCGGTTCTCATTGTGCTG CGCTTGGGACCTTCGTATTA Verrucomicrobia Reverse Reverse 161 Distribution of different Acidobacteria and Verrucomicrobia in plant-soil systems
sequences of the subgroups 1, 3 ,4 and 6 of Acidobacteria. Sequence and characteristics of the primers are shown in Table 2. In silico comparisons of the primer sequences with database sequences, using Primer-Blast, predicted that all primers targeting subgroup 1, 3, 4 and 6 of Acidobacteria would amplify 16S rRNA gene sequences that matched the sequences of these groups, yielding amplicons with expected sizes. PCR amplifications using the four primer sets on genomic DNA from their respective target strains (Table 1) invariably resulted in single amplicons of the expected sizes, in the absence of any visible primer dimer or other products (data not shown). Also, PCR amplification - using all primer sets - of genomic DNA from six non-target strains of the Firmicutes, Proteobacteria and Bacteriodetes, as well as of all non-corresponding Acidobacteria and/or Verrucomicrobia subdivision 1 strains, resulted in the absence of any band under the amplification conditions used. Finally, sequence analyses of amplicons made with subgroups 1, 3, 4 and 6 of Acidobacteria- specific primers on bulk soil DNA as the template (48 per system) consistently revealed matches (>96% similarity) with database sequences belonging to the expected groups, with the exception of the primer set designed for detection of group 6 species (Table 3). Using the latter primer set, 46 sequences turned out to be divided over four group-6 species, whereas two sequences showed 99% similarity with group-10 species. In contrast, the amplicons generated with the first three primer sets gave closest matches with between four and seven different species, indicating that these primers amplify multiple species within each Acidobacteria subgroup from soil. Based on specificity tests done on pure culture strains and on sequence analyses of PCR amplicons made from soil DNA, these primer sets were specific for their target Acidobacteria, with the slight exception of the group-6 primer set, which co-amplifies group-10 sequences. Standard curves of the four qPCR systems, constructed by plotting Ct values against log target cell numbers at different density levels in qPCR assays revealed linear relationships between the two parameters in all four cases (R2 > 0.9841) (Table 2). The amplification efficiency in all used qPCR systems ranged between 1.91 and 2.01 and the calculated dynamic ranges were between 2.54 to 7.54 and 4.21 to 9.21 (Log equivalent cell numbers per g dry soil) (Table 2).
Distribution of total bacteria, Acidobacteria and Verrucomicrobium subdivision 1 groups in bulk soil and two different rhizosphere compartments of different plant species.
Bacterial cell numbers, expressed as Log equivalent cell numbers per g dry soil, were between 8.29 and 9.39 in bulk soils taken from the three different field plots. Bacterial cell numbers in one or both rhizosphere soil compartments of potato, leek and grass were consistently higher than those in corresponding bulk soils (see Appendix Fig. A2).
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96% 96% 97% 96% 96% 97% 97% 97% 97% 97% 98% 98% 98% 98% 97% 98% 99% 100% 100% 100% 100% 100% 100% 100% similarity d group 1 1 group 1 group 1 group 1 group 1 group 3 group 3 group 3 group 3 group 3 group 3 group 3 group 4 group 4 group 4 group 4 group 6 group 6 group 6 group 6 group 10 group 10 group and unclassified Phylogeny of closest match Acidobacteria Acidobacteria Acidobacteria Acidobacteria Acidobacteria Acidobacteria Acidobacteria Acidobacteria Acidobacteria Acidobacteria Acidobacteria Acidobacteria Acidobacteria Acidobacteria Acidobacteria Acidobacteria Acidobacteria Acidobacteria Acidobacteria Acidobacteria Acidobacteria Acidobacteria Luteolibacter , (EU979057) (EU979057) (FJ569756) Holophagae bacterium bacterium
(DQ528761) (DQ528761) ), in parenthesis accession numbers of closest of matches numbers accession parenthesis ), in IGE-018 (GU187039) (GU187039) IGE-018 IGE-013 (GU187038) (GU187038) IGE-013 (GU187038) IGE-013 (FJ405886) YJF2-13 (GU187039) IGE-018 (GU187039) IGE-018 (DQ660895) (DQ660895) 16SrRNA farm library experimental gene from Droevendaal clone bacterium bacterium aggregans bacterium bacterium bacterium bacterium
c roseus Acidobacteria Acidobacteria
cteria a Terriglobus Acidobacteria Acidobacteria Edaphobacter Acidobacteria Acidobacteria Acidobacteria Closest match Closest b http://blast.ncbi.nlm.nih.gov/Blast.cgi d with the groups-specific primers for # a 5 (AY364074) bacterium Uncultured Acc. FN994868 3 FN994868 18 FN994869 3 FN994870 12 FN994871 12 FN994872 FN994873 FN994874 FN994875 7 FN994876 3 (GQ918887) bacterium soil Uncultured 15 FN994877 (GQ127801) bacterium Uncultured FN994879 4 (GQ918887) bacterium soil Uncultured FN994880 6 (FJ166826) bacterium Uncultured 6 (GQ127801) bacterium Uncultured 7 FN994881 7 (FJ166757) bacterium Uncultured 14 FN994882 Acidob (GQ918949) bacterium soil Uncultured 12 FN994883 Uncultured 15 FN994884 Uncultured 7 FN994885 FN994886 FN994887 11 FN994888 13 FN994889 (AY289390) bacterium soil Uncultured 1 (GU082782) bacterium soil Uncultured 1 (GQ495523) bacterium Uncultured (GQ495523) bacterium Uncultured . Clone Lib4_1_3x Lib4_2_18x Lib4_3_3x Lib4_4_12x Lib4_5_12x Lib5_1_7x Lib5_2_3x Lib5_3_15x Lib5_4_4x Lib5_5_6x Lib5_6_6x Lib5_7_7x Lib6_1_17x Lib6_2_14x Lib6_3_12x Lib6_4_5x Lib7_1_15x Lib7_2_7x Lib7_3_11x Lib7_4_13x Lib7_5_1x Lib7_6_1x Identities of clones (out of 48 for each library) recovered in of clones (out of 48 for recovered library) each Identities Number of clones with same sequence recovered out of a clone library of 48 clones 48 clones of library a clone of out recovered sequence same clones with of Number Subgroup 1 Subgroup 4 Subgroup 3 Subgroup 6
to RDP classification in accordance of closet determined was match Phylogeny number assession clones Closest match determinedwas Blastn by ( Table 3 Table group Target Acidobacteria a b c d soil. The cloned amplicons were generate Verrucomicrobiaceae
163 Distribution of different Acidobacteria and Verrucomicrobia in plant-soil systems
For potato, bacterial cell numbers ranged between 9.10 and 10.40 in the outer and between 9.48 and 11.19 in the inner rhizospheres. For leek, the values were 9.12 - 9.93 in the outer and 9.15 - 11.17 in the inner rhizosphere, whereas for grass they were 9.36 - 10.52 and 9.26 - 10.25, respectively. Over time, the bacterial cell numbers remained grossly stable in both potato rhizosphere compartments, in the outer rhizosphere of leek and in the inner rhizosphere of grass. However, these numbers significantly increased over time in the inner rhizosphere of leek and in the outer rhizosphere of grass. In grass, they diminished from the first to the third month and increased from the third to the fifth month (see Appendix Fig. A2). The bacterial cell numbers thus increased in the rhizosphere examined, although they fluctuated depending on location (inner, outer rhizosphere), plant species and period of sampling. The dynamics of the four Acidobacteria and two Verrucomicrobium subdivision 1 groups showed disparate patterns (Fig. 1). The Acidobacterium group 1 numbers (Log cell equivalents per g dry soil) in the rhizosphere compartments of both plant species were grossly similar to those in their respective bulk soils. The exceptions were formed by the leek inner and outer rhizospheres at four months (revealing higher numbers than in bulk soil), and the grass outer rhizospheres after two months (showing lower numbers than in bulk soil). In rhizosphere compartments of both plants, the Acidobacterium group 3 cell numbers were grossly lower than those in the corresponding bulk soils after one and two months. These numbers were similar between bulk soils and corresponding potato rhizosphere compartments, the leek inner rhizosphere and the grass outer rhizosphere after two months. After four months, the Acidobacteria group 3 cell numbers in the leek outer rhizosphere and the grass inner rhizosphere were higher than in corresponding bulk soils. The Acidobacterium group 4 numbers were the same or lower in inner and outer rhizosphere compartments than in corresponding bulk soils of both plant species, the exception being the inner and outer rhizospheres of leek and the outer rhizosphere of grass, both after four months, where numbers were higher. For the Acidobacterium group 6, cell numbers were grossly higher in the inner and outer rhizosphere soil compartments than in corresponding bulk soils of all three plant species, with the exception of the inner rhizosphere of leek after one month, in which numbers were lower than in corresponding bulk soil. For the Holophaga spp., cell numbers were consistently lower or similar in both rhizosphere compartments of potato and grass as compared to corresponding bulk soils. These numbers were consistently higher or the same in both rhizosphere compartments of leek than in bulk soils. The Verrucomicrobium subdivision 1 groups Luteolibacter and Candidatus genus Rhizospheria cell numbers all were consistently higher in the two rhizosphere compartments of all three plant species than in corresponding bulk soils. However, Candidatus genus Rhizospheria cell numbers were lower in potato rhizosphere compartments and grass outer rhizosphere than in bulk soil after one month. Moreover,
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the grass inner rhizosphere compartments in the same time period revealed numbers similar to those in bulk soil.
4 Potato 3 * 2 * * * * * * * * * * * * * 1 * * * * * *** 0 124124124124124124124124 -1 * General * * * * * * * * Candidatus genus bacteria Group 1 Group 3 Group 4 Group 6 Holophagae Luteolibacter *Rhizospheria -2
Log ratio rhizophere : bulk soil -3 4 * * Leek * 3 * * * * 2 * * * * *** * * * * * * * * 1 * * * * * * 0 124124124124124124124124 -1 * * * * * General * Candidatus genus bacteria Group 1 Group 3 Group 4 Group 6 Holophagae Luteolibacter Rhizospheria -2
Log ratioLog rhizophere : bulksoil -3 4 Grass
3
2 * * * * * * * * * * * * * * * * * * * 1 * * * 0 124124124124124124124124 -1 * * General * * * Candidatus genus Group 1* Group 3 Group 4 Group 6 Holophagae Luteolibacter bacteria * Rhizospheria -2 ** * Log ratio rhizophere :bulk soil * * -3 Outer rhizosphere Inner rhizosphere
1, 2, 4 – Sampling month
Figure 1 Ratio of Log of equivalent cell numbers, determined by qPCR, between rhizosphere and bulk soil in the different soil compartments, plant types and growth stages of general bacteria, Acidobacteria subgroups 1, 3, 4, 6 and Holophagae and Verrucomicrobia subdivision 1 Luteolibacter and Candidatus genus Rhizospheria.
In general terms, the numbers of Acidobacterium groups 1, 3 and 4 were grossly the same or lower in the two rhizosphere compartments than in corresponding bulk soils, being only incidentally higher. This contrasts with Acidobacterium group 6 and Verrucomicrobium subdivision 1 Luteolibacter and Candidatus genus Rhizospheria groups, which had shown generally higher cell numbers in the two rhizosphere compartments than in corresponding bulk soils. The distribution of Holophaga sp. numbers over rhizosphere compartments and bulk soil was different from that of all other groups. In particular, the numbers were higher in the leek inner and outer
165 Distribution of different Acidobacteria and Verrucomicrobia in plant-soil systems
rhizosphere compartments than in corresponding bulk soil, whereas in potato and grass, numbers were lower or similar. The five Acidobacteria and two Verrucomicrobia subdivision 1 groups did not follow the same trend with respect to their distribution over plant-soil compartments of the different plant species.
Environmental factors affecting the distribution of Acidobacterium and Verrucomicrobium subdivision 1 groups in plant-soil compartments.
The effect of the ‘factors’ plant species, time of sampling and location in the plant-soil ecosystem on the distribution of five Acidobacterium and two Verrucomicrobium subdivision 1 groups was determined by redundancy analysis (RDA). The pH values in the plant-soil compartments of the three different plant species over time were included in the analyses. The measured pH values varied between 4.74 and 5.27 and no general tendencies towards lower or higher pH values, with respect to plant soil compartment, plant species or sampling time was found (see Appendix Fig. A3). Analysis by DCA, as a first step, resulted in a linear relationship between environmental and species variables, justifying the use of RDA in a second step. A biplot constructed by RDA on all environmental and species variables revealed strongest correspondence of bulk soil in one direction and inner and outer rhizosphere soils in the opposite direction with the first (horizontal) axis (Fig. 2). The first axis, explaining 69.9% of the variation, thus separated bulk soil samples from those of the two rhizosphere soils. The vectors explaining the distribution of bacteria, Acidobacterium group 6 and the Verrucomicrobium subdivision 1 groups Luteolibacter and Candidatus genus Rhizospheria were in the same direction as the vectors explaining the variation caused by inner and outer rhizosphere compartments. These four groups are thus correlated with the rhizosphere compartments of all three plant species. The second (vertical) axis, explaining 21.4% of all variation, revealed strongest correlation with plant species, leek in one, and mainly grass in the opposite direction. The vector explaining the distribution of the Holophagae showed highest association with the second axis, revealing the same direction as the leek vector. This shows the existence of a strong correlation between Holophaga and leek plants, opposing the correlation of this taxon with grass and to a lesser extent with potato plants. The other Acidobacteria subgroups were not linked with one of both axes in particular, indicating that the distribution of these groups is not affected by location in plant-soil environment, plant species, pH or sampling time.
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Grass
Acg6 4 Acg3 BS GB Potato IR Acg4Acg1 pH 2 CgRhiz OR 1 Lut
Environmental Hol variables Bacterial groups Leek -1.0 1.0 -1.0 1.0 Figure 2 Ordination plots of ‘species’ (Log equivalent cell numbers of different bacterial groups determined by qPCR) and environmental variables (plant type, pH and growth stages). GB, general bacteria; Acg1, Acidobacteria subgroup 1; Acg3, Acidobacteria subgroup 3; Acg4, Acidobacteria subgroup 4; Acg6, Acidobacteria subgroup 6; Hol, Holophagae; Lut, Luteolibacter; CgRhiz, Candidatus genus Rhizospheria. BS, bulk soil; OR, outer rhizosphere; IR, inner rhizosphere. Plant types - leek, potato and grass. Values on the axes indicate the percentage of total variation explained by each axis.
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Distribution of different Acidobacteria and Verrucomicrobia in plant-soil systems
Discussion
The distribution of different Acidobacteria and Verrucomicrobia subdivision 1 subgroups over bulk soil and two rhizosphere soils was investigated in an experimental field. The field design allowed investigation of the effects of potato and leek plants on the bacterial soil community present in a former permanent grassland soil. Soils from a fallow plot and from a plot still covered by grass served as controls. The investigated bacterial groups responded differently to the plants with respect to plant species and growth stage. We hypothesized that different plant species would differentially affect Acidobacteria and Verrucomicrobia subdivision 1 subgroups in soils. On the basis of our results we conclude that, indeed, plant roots affect some – but not all – of these groups. Our approach differed from that in previous studies on Acidobacteria and Verrucomicrobia in rhizosphere soils (Chow et al., 2002; Sanguin et al., 2006; Zul et al., 2007; Kielak et al., 2008), as we distinguished different acidobacterial and verrucomicrobial subgroups and investigated these groups in different compartments in the plant-soil ecosystem. For that purpose, novel real time PCR systems were designed and evaluated to quantify dominant Acidobacteria in soil, i.e. groups 1, 3, 4 and 6 (Jones et al., 2009; Rousk et al., 2010). Acidobacterium group 6, Holophaga and Verrucomicrobium subdivision 1 subgroups Luteolibacter and Candidatus genus Rhizospheria were generally higher in rhizosphere soils than in corresponding bulk soils. This indicated the attractiveness of plant roots for these bacterial groups. Therefore, these bacterial groups can be considered to be ‘rhizosphere competent’. Rhizosphere competence for bacteria may be defined as the capacity to survive in the neighborhood of plant roots in comparison with the surrounding soil and/or to colonize root surfaces and interior parts of roots (Lugtenberg & Kamilova, 2009). Because we only studied the presence of these groups in the rhizosphere, we may conclude that they are capable of thriving at plant roots where they may utilize root-released compounds and compete with other microorganisms present. The rationale behind the examination of two different rhizosphere compartments was that a spatial distinction in the rhizosphere could be made. The inner rhizosphere consisted of soil particles that were separated from the roots in a second wash step and these particles were therefore considered to adhere more tightly to the root surface than those from the outer rhizosphere. The inner rhizosphere is the compartment in closest contact with roots. None of the studied bacterial groups showed any consistent preference for any of both compartments during all plant growth stages, suggesting that the two compartments are to be considered as offering a similar niche for all groups. This is an important finding, because it is impossible to standardize the ‘rhizosphere’ for
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different plant species over different growth stages. We thus conclude that the outcome of our real time PCR measurements in the two compartments is representative for the rhizosphere as a whole. Remarkable was the specificity of Holophaga species for the leek rhizosphere and - to a lesser extent - to those of potato and grass. Equivalent Holophaga species cell numbers were higher in leek rhizosphere soil than in surrounding bulk soil and this effect was independent of the growth stage of the plants. The preference for leek roots may explain why Holophaga species were only isolated from the leek rhizosphere (Nunes da Rocha et al., 2010b) and not from that of potato (Nunes da Rocha et al 2010a). One may assume that the cultured Holophaga species may have co-evolved with leek plants. This aspect needs further attention in later research. Candidatus genus Rhizospheria species was always higher in numbers in the rhizosphere of all studied plants than in corresponding bulk soils, irrespective of plant growth stage. This in contrast with the Acidobacterium group 6 and Luteolibacter targets, which were also higher in the rhizospheres of all studied plant species, but not in all growth stages. Candidatus genus Rhizospheria may occupy the rhizospheres of many different plant species and may therefore be considered a ‘rhizosphere generalist’. Representatives of this group have previously been isolated from potato and leek rhizospheres at different occasions (Nunes da Rocha et al., 2010a; 2010b). This study is the first report on their ecology in relation to plants. In addition, no ecological information on Verrucomicrobium subdivision 2 (i.e. Spartobacteria, Sangwan et al., 2004) or 4 (i.e. Opitutus, Chin et al., 2001) can be found in the literature. Verrucomicrobial groups probably represent relatively minor groups in soil, e.g. calculations based on total bacterial estimates from this study indicate that 0.005 – 0.11% of the 16S rRNA genes is of Verrucomicrobia subdivision 1 origin (here, Luteolibacter and Candidatus genus Rhizospheria). Our data show that subdivision 1 Verrucomicrobium species may occupy niches at roots of many different plant species. In spite of the fact that representatives of this phylum are often moderately abundant in soils, their ecological roles with respect to interaction with plants may be important. Key factors driving the distribution of the different Acidobacterium and Verrucomicrobium subdivision 1 groups in the plant-soil system were plant growth (rhizosphere) followed by plant species. These factors cooperatively shape the composition and functioning of different microbial populations in the rhizosphere (Garbeva et al, 2004; Van Overbeek and Van Elsas, 2008; Berg and & Smalla, 2009) and therefore should be regarded as responsible for the fate and activities of the target bacterial groups in soil. In many studies, pH was found to be a key factor affecting the distribution of Acidobacterium and Verrucomicrobium in soil (Jones et al., 2009; Kuramae et al., 2010; Rousk et al., 2010). Within the realms of our study, we did not find evidence that this was the case for the groups studied by us. Possibly, pH is of
169 Distribution of different Acidobacteria and Verrucomicrobia in plant-soil systems
lower importance for acidobacterial and verrucomicrobial groups that are rhizosphere competent. On the other hand, the difference in pH between the different soil samples may have been too low to find effects on community composition. Therefore, further investigation of the effect of pH on the distribution of these groups will be needed, e.g. by extending the pH gradient by using different soils or by application of different amendments to the same soil. Preference of rhizosphere over bulk soil is not commonly observed for Acidobacterium. The rhizosphere is a soil zone where, at least temporarily, nutrients become available for microorganisms at higher levels than in corresponding bulk soils, which is mainly due to exudates released by plant roots (Lynch & Whipps, 1990). The rhizosphere may therefore be more favorable for copiotrophs than for oligotrophs. Acidobacteria have classically been considered to favor an oligotrophic lifestyle (Fierer et al., 2007). This assumption implicitly regarded the Acidobacteria, nowadays comprehending a total of 26 different groups (Barns et al., 2007), as a single (homogeneous) group. Our current study revealed the importance of making distinctions between the different Acidobacterium groups in plant-soil systems. Acidobacterium groups 1, 3 and 4 probably possess lifestyles comparable with those of oligothrophs (Poindexter, 1981; Semenov, 1991; Senechkin et al., 2010). On the other hand, Acidobacterium subgroup 6, Holophaga, Luteolibacter and Candidatus genus Rhizospheria revealed glimpses of copiotrophic lifestyles, meaning that they might detectably respond with growth to substrate that becomes available in their habitat. These groups indeed showed preferences for rhizosphere over bulk soils. Strains of all these groups are now available (George et al, 2010; Nunes da Rocha et al, 2010a; Nunes da Rocha et al., 2010b) and this will enable the further study of their ecophysiologies in more detail. In conclusion, plant roots as well as type of plant species were found to be main factors that shape the selected acidobacterial and verrucomicrobial communities in soil. Acidobacterium subgroup 6, Holophaga and Verrucomicrobium subdivision 1 Luteolibacter and Candidatus genus Rhizospheria clearly showed preferences for rhizosphere over bulk soils. These groups are likely rhizosphere competent, revealing a glimpse of a copiotrophic lifestyle. This last aspect is new for Acidobacterium because this group is classically considered as typically oligotrophic. Selection by the plant of subgroup 6 Acidobacterium, Holophaga and of Luteolibacter and Candidatus genus Rhizospheria can be of great importance in terrestrial ecosystems. Therefore, the ecology of Acidobacteria and Verrucomicrobia in plant-soil ecosystems needs to be studied in this context, also opening avenues for their biotechnological exploration.
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Acknowledgments This research was part of the Ecogenomics program which is sponsored by the Dutch National Genomics Initiative and the basic research program on sustainable agriculture (KB4) sponsored by the Dutch ministry of agriculture, nature and food safety. We would like to thank Gerard Derks and field workers of ‘Droevendaal’ experimental farm (WUR, Wageningen, The Netherlands) for their assistance in growth and sampling of field experiments. We also thank Michel M. Klerks, and Ronald van Doorn for discussion about qPCR development and optimization and Vladimir Fediy for his assistance with qPCR assays.
171 Distribution of different Acidobacteria and Verrucomicrobia in plant-soil systems
References
Barns SM, Cain EC, Sommerville L & Kuske CR (2007) Acidobacteria phylum sequences in uranium- contaminated subsurface sediments greatly expand the known diversity within the phylum. Appl Environ Microbiol 73: 3113-3116. Berg G & Smalla K (2009) Plant species and soil type cooperatively shape the structure and function of microbial communities in the rhizosphere. FEMS Microbiol Ecol 68: 1-13. Chin K-J, Liesack W & Janssen PH (2001) Opitutus terrae gen. nov., sp. nov., to accommodate novel strains of the division 'Verrucomicrobia' isolated from rice paddy soil. Int J Syst Evol Microbiol 51: 1965-1968. Chow ML, Radomski CC, McDermott JM, Davies J & Axelrood PE (2002) Molecular characterization of bacterial diversity in Lodgepole pine (Pinus contorta) rhizosphere soils from British Columbia forest soils differing in disturbance and geographic source. FEMS Microbiol Ecol 42: 347-357. Fierer N., Jackson JA, Vilgalys R & Jackson RB (2005) Assessment of soil microbial community structure by use of taxon-specific quantitative PCR assays. Appl Environ Microbiol 71: 4117-4120. Fierer N, Bradford MA & Jackson RB (2007) Toward an ecological classification of soil bacteria. Ecology 88: 1354-1364. Garbeva P, Van Veen JA & Van Elsas JD (2004) Microbial diversity in soil: Selection of microbial populations by plant and soil type and implications for disease suppressiveness. Annu Rev Phytopathol 42: 243-270. George IF, Hartmann M, Liles MR & Agathos SN (2010) Recovery of as-yet uncultured soil Acidobacteria on dilute solid media. Submitted. Houba VJG & Novozamsky I (1998) Influence of storage time and temperature of dry soils on pH and extractable nutrients using 0.01 M CaCl2. Freseninus J Anal Chem 360: 362-365. Janssen PH (2006) Identifying the dominant soil bacterial taxa in libraries of 16S rRNA and 16S rRNA genes. Appl Environm Microbiol 72: 1719-1728. Janssen PH, Yates PS, Grinton BE, Taylor PM & Sait M (2002) Improved culturability of soil bacteria and isolation in pure culture of novel members of the divisions Acidobacteria, Actinobacteria, Proteobacteria, and Verrucomicrobia. Appl Environ Microbiol 68: 2391-2396. Jones RT, Robeson MS, Lauber CL, Hamady M, Knight R & Fierer N (2009) A comprehensive survey of soil acidobacterial diversity using pyrosequencing and clone library analyses. ISME J 3: 442-453. Kielak A, Pijl AS, Van Veen JA & Kowalchuk GA (2008) Differences in vegetation composition and plant species identity lead to only minor changes in soil-borne microbial communities in a former arable field. FEMS Microbiol Ecol 63: 372-382. Kuramae EE, Gamper HA, Yergeau E, Piceno YM, Brodie EL et al. (2010) Microbial secondary succession in a chronosequence of chalk grasslands. ISME J 4: 711-715. Lane D (1991) 16s/23s rRNA sequencing, p. 115-175. In E. stackebrandt and M. Goodfellow (ed.), Nucleic acid techniques in bacterial systematics. John Wiley & Sons, West Sussex, United Kindom. Lugtenberg B & Kamilova F (2009) Plant-growth-promoting rhizobacteria. Annu Rev Microbiol 63: 541- 556. Lynch JM & Whipps JM (1990) Substrate flow in the rhizosphere. Plant Soil 129: 1-10. Marschner P & Baumann K (2003) Changes in bacterial community structure induced by mycorrhizal colonization in split-root maize. Plant Soil 251: 279-289. Muyzer G, de Waal EC & Witterlinden AG (1993) Profiling of complex microbial populations by denaturating gradient gel electrophoresis analysis of polymerase chain reaction-amplified genes coding for 16S rRNA. Appl Environ Microbiol 70: 1008-1016. Nunes da Rocha U, Andreote FD, de Azevedo JL, van Elsas JD & van Overbeek LS (2010a) Cultivation of hitherto-uncultured bacteria belonging to the Verrucomicrobia subdivision 1 from the potato (Solanum tuberosum L.) rhizosphere. J Soils Sediments 10: 326-339. Nunes da Rocha U, Andreote FD, Plugge C, van Elsas JD & van Overbeek LS (2010b) Isolation of culturable Holophagae (Acidobacteria), Luteolibacter, unclassified Verrucomicrobiaceae and Verrucomicrobium (Verrucomicrobia) from the Allium porrum rhizosphere. Submitted. Nunes da Rocha U, van Elsas JD & Overbeek LS (2009) Exploration of hitherto-uncultured bacteria from the rhizosphere. FEMS Microbiol Ecol 69: 313-328. Nunes da Rocha U., van Elsas JD & Overbeek L (2010c) Different rhizosphere competence in two Verrucomicrobium subdivision 1 strains previously isolated from leek (Allium porrum) rhizosphere. In preparation.
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Nunes da Rocha U., van Elsas JD & Overbeek L (2010d) Real-time PCR detection of Holophagae (Acidobacteria), Luteolibacter and unclassified Verrucomicrobiaceae (Verrucomicrobia subdivision 1) in bulk and leek (Allium porrum) rhizosphere soils. Submitted. Nunes da Rocha U., van Elsas JD & Overbeek L (2010e) Rhizocompetence of culturable Holophaga (Acidobacteria) sp. in the leek (Allium porrum) rhizosphere. Submitted. Poindexter JS (1981) Oligotrophy. Adv Microbiol Ecol 5: 63-89. Pruesse E, Quast C, Knittel K, Fuchs BM, Ludwig W, Peplies J & Glockner FO (2007) SILVA: a comprehensive online resource for quality checked and aligned ribosomal RNA sequence data compatible with ARB. Nucleic Acids Res 35: 7188-7196. Rousk J, Bååth E, Brookes PC, Lauber CL, Lozupone C, Caporaso JG, Knight R & Fierer N (2010) Soil bacterial and fungal communities across a pH gradient in an arable soil. ISME J DOI: 10.1038/ismej.2010.58. Sait M, Hugenholtz P & Janssen PH (2002) Cultivation of globally distributed soil bacteria from phylogenetic lineages previously only detected in cultivation-independent surveys. Environ Microbiol 4: 654-666. Sanguin H, Remenant B, Dechesne A, Thioulouse J et al. (2006) Potential of a 16S rRNA-based taxonomic microarray for analyzing the rhizosphere effects of maize on Agrobacterium spp. and bacterial communities. Appl Environ Microbiol 72: 4302-4312. Sangwan P, Chen X, Hugenholtz P & Janssen PH (2004) Chthoniobacter flavus gen. nov., sp. nov., the first pure-culture representative of subdivision two, Spartobacteria classis nov., of the phylum Verrucomicrobia. Appl Environm Microbiol 70: 5875-5881. Sangwan P, Kovac S, Davis KER, Sait M, Janssen PH (2005) Detection and cultivation of soil Verrucomicrobia. Appl Environm Microbiol 71: 8402-8410. Schlesner H, Jenkins C & Staley JT (2006) The Phylum Verrucomicrobia: A Phylogenetically Heterogeneous Bacterial Group. Prokaryotes 7: 881-896. Semenov AM (1991) Physiological bases of oligotrophy of microorganisms and the concept of microbial community. Microb Ecol 37: 116-128. Senechkin IV, Speksnijder AGCL, Semenov AM, van Bruggen AHC & van Overbeek LS (2010) Isolation and Partial Characterization of Bacterial Strains on Low Organic Carbon Medium from Soils Fertilized with Different Organic Amendments. Microb Ecol DOI: 10.1007/s00248-010-9670-1 Van Overbeek L. & Van Elsas JD (2008) Effects of plant genotype and growth stage on the structure of bacterial communities associated with potato (Solanum tuberosum L.). FEMS Microbiol Ecol 64: 283- 296. Van Rhijn P & Vanderleyden J (1995) The Rhizobium-plant symbiosis. Microbiol Rev 59: 124-142. Zul D, Denzel S, Kotz A & Overmann J (2007) Effects of plant biomass, plant diversity, and water content on bacterial communities in soil lysimeters: Implications for the determinants of bacterial diversity. Appl Environ Microbiol 73: 6916-6929.
173 General Discussion
Chapter 9: General Discussion
In this thesis, I intended to shed light on the ecology of hitherto poorly-culturable bacterial groups in the rhizosphere, focussing on members of the Acidobacteria and Verrucomicrobia. Special emphasis was placed on the distribution of selected members of these bacterial groups over rhizosphere versus bulk soil. To reach these goals, I carried out an initial step using elaborate isolation procedures, to isolate novel bacterial strains. Later, group-specific real time PCR systems for the detection and quantification of particular Acidobacteria and Verrucomicrobia were developed. These quantification systems allowed in-depth studies on some of the newly-cultured bacterial groups both in laboratory microcosms and in the field.
Dominant hitherto-uncultured bacteria in the rhizosphere
The rhizosphere, generally defined as the zone in soil that is influenced by plant roots (Hiltner, 1904), is key to the fate of root-associated bacteria in every plant species, soil type and geographic location. An important impediment to our understanding of the functioning of rhizosphere communities is the lack of culturability of a major fraction of the extant microbiota. It would be a major achievement if general rules could be discerned that govern culturability from the rhizosphere. However, the diversity of combinations created by the different soil, plant type, microbial community and environmental conditions on Earth make it difficult to establish a general concept of which bacterial groups are poorly culturable yet dominate the soil under the influence of plant roots. Hence, it appears that the “one at a time” paradigm will still be with us for some time to come. An attempt to shed some light on theories pertaining to culturability from the rhizosphere was made in chapter 2. For this, we used the following strategy: (i) collation of all available culture-independent studies that address the rhizosphere bacterial diversity across a broad array of plant types; (ii) identification of the most dominant phyla in the rhizosphere as found in each study; (iii) singling out of the most dominant groups as the ‘most commonly found’ taxa in the rhizosphere by frequency of detection across all studies. Thus, among the eighteen studies that address the diversity of the microbiota associated with different plant types between 2001 and 2008, we found as a major issue
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that rhizospheric Acidobacteria, Verrucomicrobia and Planctomycetes are often widely detected in the rhizosphere, however exclusively on the basis of culture-independent studies. We therefore set out to improve and apply methods for the isolation of new bacterial groups from the rhizosphere and placed a special emphasis on members of the aforementioned groups. The work performed in this part of the study (chapters 3 and 4) isolated and identified new members of Holophaga (Acidobacterium), Luteolibacter and Candidatus genus Rhizospheria (Verrucomicrobium) from rhizosphere samples. On the other hand, although an increasing number of reports detects particular members of the Planctomycetes in the rhizosphere of different plant types (Norstrom et al., 2008; Rosenberg et al., 2009; Sanguin et al., 2009; Yarwood et al., 2009), this taxon was not found among the isolates from our rhizospheres.
Diminishing the gap between cultured and total bacteria in rhizosphere samples
In the past decade, several reports have shown that it is possible to diminish the so- called culturability gap encountered with traditional cultivation techniques (Janssen et al. 2002; Sait et al.,2002; Bruns et al., 2003; Schoenborn et al., 2004; Davis et al., 2005; Yamada et al., 2006; Stingl et al., 2007; Ferrari et al., 2008; D'Onofrio et al., 2010). Although some of these studies reveal sophisticated techniques to isolate hitherto- uncultured bacteria, such as microcolony cultivation (Ferrari et al., 2005), substrate membrane systems (Ferrari et al., 2008) and high-throughput culturing (Stingl et al., 2007), most of the advances made in culturing were based on rather simple adaptations of traditional culturing techniques. Specifically, these techniques used reduced nutrient availability, prolonged incubation times and reduction of oxidative stress by the addition of protective agents. Moreover, natural nutrient conditions were mimicked by using additions of environmental (filter-sterilized) extracts, such as soil extracts (Hamaki et al., 2005). In chapters 3 and 4, we clearly show that use of the aforementioned culturing strategies indeed helped to diminish (but not abolish) the culturability gap in rhizosphere samples. Low- carbon-availability medium, modified or not, was compared to R2A, the standard medium for bacterial isolation from environmental samples. Furthermore, we screened for both “normal-sized” and microcolonies and assessed growth both after short and long incubation periods. Colony forming unit (CFU) counts were compared to counts of DTAF-stained cells by confocal microscopy to determine what fraction of the extant bacterial community was cultured. Up to an unprecented 33.6% and 33.0%, respectively (chapter 3 and 4), of the extant bacterial community was obtained in culture using the combined adaptations and searching from microcolonies. Indeed, the most important adaptations were the use of low-carbon-
175 General Discussion
availability media and long incubation periods. Moreover, the novel strategies yielded CFU counts that exceeded those on R2A by 12-fold, demonstrating great progress in culturability. Among the novelty found were presumptively new clades of Actinobacteria, Alphaptroteobacteria, Gammaptroteobacteria and Sphingobacteria. However, the rationale used in chapters 3 and 4 was to place a focus on newly- culturable Acidobacteria and Verrucomicrobia, and hence we further did not emphasize the remainder of the community.
Phylogeny and phenotypic analysis of culturable Acidobacteria and Verrucomicrobia from the rhizosphere
Two novel strains belonging to Acidobacterium were isolated from the rhizosphere of leek (chapter 4). Moreover, nine novel strains of Verrucomicrobium (chapters 3 and 4) were isolated from the rhizospheres of potato as well as leek growing in different soil types and geographic locations. To the best of my knowledge, these are the first reports on the cultivation of particular members of Acidobacterium and Verrucomicrobium from the rhizosphere. The two acidobacterial, strains CHC25 and ORAC, were affiliated with Holophaga (previously known as Acidobacterium subgroup 8). Holophaga represents a genus with few members within the phylum Acidobacterium. A pyrosequencing study focusing on the distribution of acidobacterial subgroups in (bulk) soil estimated that Holophaga population sizes amounted to up to 3.4% of the total Acidobacterium community (Jones et al., 2009). As other subgroups of the Acidobacteria, for instance 1, 2, 3, 4 and 6, are present in much higher numbers in (bulk) soils, Holophaga may have been overlooked in the past. Additionally, plant roots may offer preferred sites for members of this group, allowing increases in abundance. Previously, several other acidobacterial subgroups (especially 6), have been detected in the rhizosphere of different plants (Schmalenberger & Tebbe, 2003; Sharma et al., 2005; Wang et al., 2007; Hao et al., 2008). With respect to isolation, subgroup 6 Acidobacteria have only been obtained from surface bulk soil (George et al., 2010). The nine Verrucomicrobium strains, which all affiliated with subdivision 1 (chapters 3 and 4), were quite diverse. These strains were identified as Luteolibacter (strains CHC12, C20 and ONA9), Candidatus genus Rhizospheria (strains CHC8, IRVE, CR28, Z35 and ZNBB5) and Verrucomicrobium (strain CNC16). The subgroup Candidatus genus Rhizospheria coined by us was striking, as it appears to be a coherent group within the Verrucomicrobia that is typical for the rhizosphere. Members of subdivision 1 Verrucomicrobia have previously been isolated from, e.g., freshwater (Schlesner, 1987; Hedlund et al., 1996) and marine settings (Hedlund et al., 1996;
176 Chapter 9
Scheuermayer et al., 2006; Yoon et al., 2008). Prior to the current study, Candidatus genus Rhizospheria had not been cultured. They were thus denoted as “unclassified Verrucomicrobiaceae” group, a tight hitherto-unassigned group in the Verrucomicrobia. In chapter 4, I proposed to denote this group at the genus level as “Candidatus genus Rhizospheria”, to be included in the family Rubritaleaceae (class Verrucomicrobiae, phylum Verrucomicrobia). The proposed name for the group is based on its repeated isolation from the rhizosphere. The isolation of novel Holophaga, Luteolibacter, Candidatus genus Rhizospheria and Verrucomicrobium spp. brings new perspectives in the study of the ecology of Acidobacterium and Verrucomicrobium in soil. An analysis of the ecological potential of these novel strains based on partial phenotypic characterization was made in chapter 4.
Phenotypic analysis of culturable Holophaga and Verrucomicrobia subdivision 1
The isolation of Acidobacterium and Verrucomicrobium types from the rhizosphere inspired studies on their potential ecological role in this habitat. In chapter 4, we found that Holophaga strains CHC25 and ORAC were similar but not clonal. Both strains were motile, indicating the possibility of chemotaxis, adherence to substrates, potential biofilm formation and even swarming (Young, 2007). As both strains form short chains of cells, another ecological feature might be defence against bacterivory (Young, 2007). As protists can ingest only those bacteria that are ‘just right’ as far as size and shape are concerned, the short chains make these Holophaga strains ‘too long’ to be ingested by protozoa (Young, 2007). As observed for other Acidobacterium strains (Valáková et al., 2009), strains CHC25 and ORAC could not grow in liquid media, whereas their growth was always surface-associated on aerobically-incubated oligotrophic medium amended with catalase (in the first 10 transfers) and later in R2A (agar) plates. Holophaga foetida (accession number X77215), Geothrix fermentans (accession number U41563) and Acanthopleuribacter pedis (accession number AB303221), the only three cultured Holophagae described to date, showed different ecophysiological features. Both Holophaga foetida (Liesack et al., 1994) and Geothrix fermentans (Coates et al., 1999) are strictly anaerobic bacteria isolated from hydrocarbon-contaminated areas, whereas Acanthopleuribacter pedis is a strictly aerobic bacterium isolated from beach chiton (Fukunaga et al., 2008). The striking differences between the physiologies of these Holophagae indicate that different ecological niches may be occupied by different members of this acidobacterial group. That Holophaga spp. can indeed be plant- associated, was recently found. Thus, they were detected, by use of 16S rRNA gene clone libraries, in the endophytic bacterial community of rice roots (Sun et al., 2008) as
177 General Discussion
well as in the rhizosphere of Phragmites (accession number AB240249). Although these two sequences are related to those of strains CHC25 and ORAC, their closest match, 97.5% similarity, was with an uncultured bacterium (accession number GU169059) recovered from “synthetic river water with humic substances”. Our data, together with these recent literature data, indicate that particular Holophaga spp. may indeed find their niche in association with plants, either in the rhizosphere of even as endophytes. Quite diverse phenotypic profiles were observed among the subdivision 1 Verrucomicrobia as described in chapter 4. All Candidatus genus Rhizospheria spp. were able to grow on several organic acids and/or amino acids. These compounds are commonly found in the rhizospheres of different plant species. Hence, we posit that our novel strains will likely be able to grow at the roots of plants that exude such compounds (Jones, 1998; Baudoin et al., 2003). Moreover, cellulase activity was observed for strain CHC8, and this may indicate that it is able to grow on plant material in soil, or may colonize living roots (Mostajeran et al., 2007) and/or the interior tissues of plants (Bischoff et al., 2009). In addition, the putative presence of laccase genes in five of the nine Verrucomicrobium subdivision 1 strains demonstrated that these may specifically oxidize phenolic and non-phenolic lignin-related compounds (Kunamneni et al., 2008). Although the phenotypic characteristics of the culturable Holophaga and Verrucomicrobium subdivision 1 strains provided glimpses of possible ecological niches for these bacterial groups, more in-depth studies in soil-plant systems were deemed necessary. These were performed in chapters 5 to 8. The general objective was to determine if the strains would respond to a rhizosphere or would prefer persistence in bulk soil.
Real-time PCR detection of particular Acidobacterium and Verrucomicrobium types in bulk and rhizosphere soil.
Among the striking phenotypic characteristics of Holophaga and Verrucomicrobium subdivision 1 strains obtained in chapters 3 and 4 was the fact that only Candidatus genus Rhizospheria sp. strain CHC8 could grow in liquid media. Furthermore, all strains were very slow growers when compared to Escherichia coli (chapter 4). These features would hamper ecological studies based on growth-dependent methods (CFU counts). To circumvent this problem, real-time PCR systems for the specific detection of Holophaga, Luteolibacter and Candidatus genus Rhizospheria were developed and evaluated in chapter 5. Moreover, the same procedure to develop real time PCR systems was used to create group-specific systems for quantification of Acidobacterium subgroups 1, 3, 4 and 6 (chapter 8); these are the most dominant acidobacterial groups
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in the environment (Jones et al., 2009) and were recently cultured by George et al. (2010). Real time PCR has been promoted as a rapid, sensitive and specific method for quantification of specific bacteria in soil, water, air and plant samples (Hermansson & Lindgren, 2001; Ibekwe et al., 2002; Makino et al., 2001; Weller et al., 2000). Fluorescence is used to monitor the accumulation of PCR products after each PCR cycle. The fluorescence data are used to estimate the amount of target DNA present in the sample before amplification; thus, detection and quantification are achieved in a single assay (Wittwer et al., 1997). The use of target bacterial cells to establish calibration curves for the group-specific real-time PCR allows the estimation of so- called “equivalent cell numbers” of the target group in environmental samples. Although problems may occur, e.g. due to different bacteria possessing different copy numbers of the 16S rRNA gene in the genome, the method provides a glimpse of the numbers of cells of the target group that are present in environmental samples. Pertaining to amplification efficiency, validation of real time PCR is probably needed for each new detection system and habitat. I decided to validate our new group-specific real time PCR systems in three steps, i.e. in silico, testing with PCR amplifications of target and non-target strains in pure culture and in soil DNA extracts. All real time PCR systems designed turned out to be specific, with the exception of those designed for Luteolibacter (Verrucomicrobium subdivision 1; chapter 5), and subgroup 6 acidobacteria (chapter 8). The primers designed for Luteolibacter also amplified Prosthecobacter sequences. Prosthecobacter also belongs to Verrucomicrobium subdivision 1 and was only recently defined as a separate genus (Yoon et al., 2008). Although in silico analysis indicated Luteolibacter-specific amplification when using primers VS1Af and VS1Ar, sequencing of clones generated with this primer pair demonstrated that both Luteolibacter and Prosthecobacter members are actually amplified from soil DNA. The primers designed to specifically amplify subgroup 6 of Acidobacteria also amplified sequences of subgroup 10 of this phylum. Subgroup 6 acidobacteria may range from 0.6 to 43.3% of the Acidobacterium community in soils, whereas Acidobacterium subgroup 10 is often lower, i.e. 0 to 2.6% (Jones et al., 2009). However, the designed primer set thus recognized acidobacterial subgroups 6 and 10 and further differentiation awaits novel systems. For simplification, the real-time PCR systems targeting Luteolibacter/Prosthecobacter and subproup 6 and 10 acidobacteria were denoted respectively as specific for ‘Luteolibacter’ and ‘Acidobacterium subgroup 6’ throughout this thesis. I conclude that on the basis of the designed primer sets (specific forAcidobacterium subgroups 1, 3, 4 and Holophaga and Verrucomicrobium Candidatus genus Rhizospheria and semi-specific for Acidobacterium subgroup 6 and Verrucomicrobium Luteolibacter) it is possible to obtain reasonable estimates of target cell numbers across habitats, for instance bulk
179 General Discussion
versus rhizosphere soil. In the two cases where the PCR systems actually lumped groups together extra scrutiny is necessary.
Holophaga and Verrucomicrobium subdivision 1 spp. show shifts in abundance from bulk to rhizosphere soil
In this thesis, three different experimental approaches were used to study the numerical shift of Acidobacterium and Verrucomicrobium subdivision 1 target cells from bulk to rhizosphere soil. The first was the study of in vitro root colonization by Holophaga sp. (chapter 6), Luteolibacter and Candidatus genus Rhizospheria sp. strains (chapter 7). We specifically questioned whether bacterial growth would demand the utilization of root exudates as C sources. Later, a model plant-soil microcosm (Kuchenbuch and Jungk, 1982) was used to evaluate rhizosphere colonization by Holophaga sp. CHC25 (chapter 6) and Luteolibacter sp. and Candidatus genus Rhizospheria sp. (chapter 7). The so-called Kuchenbuch system has been used previously to measure bacterial activities in soil and model rhizosphere. For instance, colonization (Dijkstra et al., 1987), plasmid mobilization and transfer (Van Elsas et al., 1988) and the induction of a root exudate responsive operon (Van Overbeek & Van Elsas, 1995) have all been measured. In our studies, to investigate the rhizosphere relevance of our new strains, the respective equivalent cell numbers were determined by using the real time PCR systems developed in chapter 5. The in vitro studies conducted in chapter 6 revealed that the two Holophaga sp. strains CHC25 and ORAC could thrive on agar plates without added C in the presence of leek plantlets, whereas no growth was observed on plates without plants. A similar observation was made for Luteolibacter sp. CHC12 and Candidatus genus Rhizospheria sp. CHC8 (chapter 7). The in vitro colonization of, and growth on, sterile leek roots on ‘C-less’ agar are consistent with the phenotypes of strains CHC25, ORAC and CHC8 and contrast with that of strain CHC12 (chapter 4). Strains CHC25 and ORAC were able to grow on malic acid, succinic acid, citric acid, glutamine or alanine as the sole C sources. Strain CHC8 was able to grow on oxalic, malic, citric or succinic acid, as well as glutamine and alanine as the sole carbon sources. These compounds have been considered as ‘common’ for rhizospheres of different plant species (Baudoin et al., 2003; Jones, 1998). In contrast, strain CHC12 did not grow on any of the organic or amino acids tested (chapter 4). Hence, despite being closely related on the basis of the 16S rRNA gene, different subdivision 1 Verrucomicrobia may occupy different niches in the plant-soil systems. Given the propensity of Holophaga sp. strains CHC25 and ORAC and Candidatus genus Rhizospheria sp. CHC8 to grow on root exudates, further studies about their rhizosphere colonization were made, as described in chapters 6 and
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7. The purpose was to determine if these bacteria may thrive in the rhizosphere and if they are able to compete with other soil microorganism in there. Different to what is generally accepted for Acidobacterium, the Holophaga numbers in the leek rhizosphere were grossly one Log unit per gram of dry soil higher than those observed in the respective bulk soil, what reveals capacity to build up a substantive rhizosphere population. This observation hinted at a response of the bacterium to the rhizosphere and hence possible rhizosphere competence. Supporting this, we found that cells of Holophaga sp. CHC25 migrated towards the roots from a soil layer 10 mm away, suggesting that Holophaga cells follow nutrient gradients that emerge from roots in an upward fashion. Migratory activity towards plant roots (chemotaxis) can be considered as an initial step in root colonization (De Weert et al., 2002; Hozore and Alexander, 1991; Rudrappa et al., 2008). Contrasting with the fact that both Luteolibacter sp. CHC12 and Candidatus genus Rhizospheria sp. CHC8 were able to colonize leek roots on agar at the expense of C sources in root exudates, distinct rhizosphere colonization was observed for both strains in chapter 7. While the equivalent cell numbers g-1 dry soil of Candidatus genus Rhizospheria sp. CHC8 were approximately 1 Log unit higher than those found in bulk soil, those of Luteolibacter sp. CHC12 were approximately 1 Log unit per g dry soil lower. Similar patterns were observed for the natural populations of these Verrucomicrobium subdivision 1 groups using the same model plant-soil microcosm. We cannot explain this discrepancy. Indeed, Candidatus genus Rhizospheria is a presumptive rhizosphere-competent group, whereas more in-depth studies are necessary to affirm about the preference of Luteolibacter for bulk or rhizosphere soil. Possibly, Luteolibacter members may need direct contact with the root to demonstrate rhizosphere competence. Moreover, further development of tools to distinguish between Luteolibacter and Prosthecobacter is necessary to determine if one of both groups prefer rhizosphere over bulk soil. Taken together, the data described in chapters 6 and 7 demonstrate the clear rhizosphere competence of Holophaga sp. strain CHC25 and Candidatus genus Rhizospheria sp. strain CHC8. The data further indicate that interpretation of the distribution of bacterial groups over soil compartments, especially when dealing with high-ranking taxonomic groups (e.g. phylum, class, order) should be made with great care, as effects at lower taxonomic resolution levels may indicate quite contrasting phenomena. Clearly, if we would not have discriminated the different Verrucomicrobium subdivision 1 groups, or would have focused specifically on Acidobacterium, the distinct behavior observed for Holophaga, Luteolibacter and Candidatus genus Rhizospheria would have been completely overlooked.
181 General Discussion
Environmental factors that affect the distribution of Acidobacterium and Verrucomicrobium spp. over rhizosphere and bulk soil
To address a possible dichotomy between microcosm and field, an open field experiment with potato, leek and permanent grass was performed. The specific objectives were to evaluate the distribution of Acidobacterium subgroup 1, 3, 4 and 6, as well as Holophagae, and of Luteolibacter and Candidatus genus Rhizospheria (Verrucomicrobium subdivision 1) (chapter 8). To this end, group-specific real time PCR detection systems developed in chapters 5 and 8 were used. Moreover, the distribution of the target groups over plants, bulk soil and time was correlated with environmental (soil, plant) factors. We also considered our data in the context of other field experiments dealing with the distribution of Acidobacterium and Verrucomicrobium between rhizosphere and bulk soils (Chow et al., 2002; Kielak et al., 2008; Sanguin et al., 2006; Zul et al., 2007). These studies considered Acidobacterium and Verrucomicrobium, bacterial phyla with high taxonomic diversities, as single groups. In chapter 8, we designed group-specific real time PCR systems that target different subgroups within Acidobacterium and Verrucomicrobium subdivision 1. The distributions of subgroup 1, 3 and 4 acidobacteria corroborated the generally accepted role of Acidobacterium, i.e. these groups revealed lower or similar numbers in rhizospheres as compared to bulk soils (Kielak et al., 2008) (Fig. 1). This was independent of plant species or growth stage. On the other hand, subgroup 6 Acidobacteria showed higher equivalent cell numbers in the rhizosphere than in corresponding bulk soil. These organisms might be considered to be rhizosphere- competent (Fig. 1). The distribution of Holophaga over rhizosphere and bulk soil was distinct from that observed for the other acidobacterial groups. Holophaga equivalent cell numbers were higher in the rhizosphere of leek but lower in that of potato and grass as compared to the bulk soil (Fig. 1). This was in line with observations on Holophaga in the plant-soil microcosms made in chapter 6. Therefore, members of Holophaga may occupy a specific ecological niche in the leek rhizosphere.
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Figure 1 Preferred site in the plant-soil system for the different subgroups of Acidobacteria and Verrucomicrobia subdivision 1. Interrogation mark following a bacterial taxum (?) indicates that further research is necessary to evaluate the preferred site for the respective bacterial group. An asterisk (*) indicates that the respective bacterial taxum preferred the rhizosphere over the respective bulk soil only in the leek rhizosphere.
Luteolibacter and Candidatus genus Rhizospheria numbers were always higher in the rhizosphere than in corresponding bulk soil and this was independent of plant type and growth stage. Thus, the data described in chapter 8 about the distribution of Candidatus genus Rhizospheria corroborate those observed in chapter 7. Differently, Luteolibacter equivalent cell numbers were raised in the rhizosphere in the field (chapter 8) but reduced in the rhizosphere in the microcosm as compared to corresponding bulk soils (Fig. 1). One caveat here is the co-detection of Prosthecobacter sequences. Clearly, further studies on the rhizosphere competence of Luteolibacter should be performed to clarify these different observations. Multivariate analyses performed to determine what environmental factors influence the distribution of the selected Acidobacterium and Verrucomicrobium subdivision 1 types over the different soil compartments revealed that the most important factor was compartment, i.e. rhizosphere or bulk soil. The second factor shaping the rhizosphere community was plant type. The structure and function of microbial communities in soil are cooperatively shaped by these factors (Berg & Smalla, 2009, Garbeva et al., 2004). In bulk soil, the most important factor shaping the community make-up of Acidobacterium and Verrucomicrobium is often pH (Jones et al., 2009); Kuramae et al., 2010; Rousk et al., 2010). Multivariate analysis (chapter 8) demonstrated, in our case, that pH exerted a smaller influence on the distribution of the
183 General Discussion
different Acidobacterium and Verrucomicrobium groups than soil compartment and plant type. Indeed, the observed differences between bulk and rhizosphere soil pH were not large, possibly explaining why this factor was not as important as the differences between soil compartments and plant types. Bacteria that grow at extremely low nutrient availability are denoted as oligothrophs (Poindexter, 1981; Semenov, 1991; Senechkin et al., 2010). Given the general scarcity of readily-available nutrients, bulk soil is often regarded as propicious for oligotrophs. Compared to bulk soil, the rhizosphere may be considered as a soil zone with raised nutrient availability as a result of the release of exudates (Lynch & Wipps, 1990). This thesis (chapters 6 and 8) indicated that specific Acidobacterium groups deviate from the generally accepted concept that Acidobacteria are oligotrophs (Fierer et al., 2007). Subgroup 6 Acidobacteria showed preference for rhizosphere over bulk soil independently of plant type. Holophaga members were in higher abundance in the rhizosphere of leek. Also Luteolibacter and Candidatus genus Rhizospheria showed preference for rhizosphere over bulk soil independently of plant type and growth stage. Although Acidobacteria are generally regarded as oligotrophs (Fierer et al., 2007) this study indicated that specific acidobacterial groups, among them subgroup 6 (a dominant Acibacterium type in bulk soils), may deviate from this general concept. It is possible that Fierer et al. (2007) could not demonstrate this trend because they considered Acidobacterium as-a-whole, disregarding the high diversity of this phylum.
Future perspectives
A suite of novel strains of Acidobacterium and Verrucomicrobium were isolated in this study. Also, generally accepted theories about the ecology of these groups were challenged. However, much remains unknown, for instance: i) what are the ecological niches of the Acidobacterium and Verrucomicrobium subgroups related to plants; ii) what novel biotechnological applications may these novel microbial taxa hide in their genomic machinery; iii) to what extent are Acidobacteria actually oligothrophic; iv) may the observations on the Acidobacterium and Verrucomicrobium subgroups be generalized or extrapolated to other members of these bacterial groups; and v) what is the actual involvement of Acidobacterium and Verrucomicrobium types in soil and plant health. The following approaches are proposed to advance our knowledge of the ecology of these bacterial groups in the rhizosphere
Recognition of novel Acidobacteria and Verrucomicrobia strains as species The taxonomy of Acidobacterium and Verrucomicrobium has expanded in the past decade. Still, much has to be done to standardize the nomenclature of these two phyla.
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Both groups have increased in culture collections but most of their diversity derives from culture-independent studies based on the 16S rRNA gene. Although knowledge of the diversity within Acidobacterium and Verrucomicrobium has been increasing, no consensus about the taxonomy of these groups exists. Bergey’s Manual contains the official nomenclature of both phyla, but the complete diversity observed with the use of culture-independent studies goes far beyond that. Therefore, I suggest that a forum consisting of specialists of both phyla should be created to analyze and standardize the Acidobacterium and Verrucomicrobium taxonomic nomenclature. This effort would facilitate the comparison of data and management decisions on what specific acidobacterial and verrucomicrobial groups should be further explored
Exploration of phenotypic characteristics of novel Acidobacterium and Verrucomicrobium strains The novel strains of Acidobacterium and Verrucomicrobium represent previously hitherto-uncultured bacteria. The abundance of Acidobacterium subgroup 6 and Holophagae and Verrucomicrobium subdivision 1 in the rhizosphere suggest that they serve functions that are important in this environment. Therefore, in-depth studies digging into their physiologies to find interesting biotechnological applications should be performed. An initial step would be the confirmation and further exploration of the production of laccases observed for Luteolibacter spp. CHC12 and ONA9, Candidatus genus Rhizospheria spp. IRVE and CR28 and Verrucomicrobium sp. CNC16. Besides, the genome sequences of the novel Acidobacteria and Verrucomicrobia isolated in this thesis may identify potential traits that suggest ecological roles of these bacteria in plant-soil systems. Moreover, the search for transposons and plasmids in all novel Acidobacterium and Verrucomicrobium strains may reveal novel mobile genomic elements and plasmids with interesting biotechnological applications. Indeed, selected strains may be useful as hosts for vectors in the construction of metagenomic libraries. Due to their phylogenetic distance to the most commonly used host (Escherichia coli), this bacterium may be useful in the expression of genes that the genetic machinery of E. coli is not prepared to process.
Ecological niche of Holophaga sp. CHC25 in the leek rhizosphere The data from chapters 6 and 8 suggest that Holophaga sp. CHC25 has a specific ecological role in the rhizosphere of leek. I would suggest the use of the novel PM Biolog plates (Biolog, Inc., USA) that test the basic cellular nutritional pathways for C, N, P, and S metabolism, besides tolerance to osmotic stress, pH and different chemical agents. This analysis, coupled to the generation of a draft genome, may help elaborate further experiments to delineate the actual ecological niche of this bacterium in the leek
185 General Discussion
rhizosphere. Similar experiments may advance the ecological knowledge of the novel verrucomicrobial strains.
Oligotrophic status of Acidobacterium It has generally accepted that Acidobacterium encompasses exclusively oligotrophs (Fierer et al., 2007). Oligotrophy is a phenotypic characteristic of bacteria (Senechkin et al., 2010) and may change even with the number of transfers that specific strains are subjected to under laboratory conditions (Senechkin et al., 2010). Therefore, the oligotrophic status of Acidobacterium should be tested under laboratory conditions with regard to numbers of transfers. As the number of Acidobacterium strains grows in culture collections, an effort to analyze such phenotypic characteristic would be simple. The outcome of this approach, that might depend on strain-specific characteristics (Senechkin et al., 2010), would further expand the view on this matter in Acidobacterium.
Meta analysis of the distribution of Acidobacterium and Verrucomicrobium in bulk and rhizosphere soil As explained in chapter 2, the determination of which Acidobacterium and Verrucomicrobium spp. find their niche in the rhizosphere has been hampered by their recalcitrance to grow in pure culture. Furthermore, the huge number of combinations of soil, plant type and environmental condition in which these bacterial groups occur around the globe is blurring this facet. Sequencing technologies, such as pyrosequencing, have become cheaper and more accessible to a broad scientific community. For instance, this technique has already been used to study the distribution of acidobacterial communities in a broad range of soils with different pHs (Jones et al., 2009; Rousk et al., 2010). Similar studies comparing rhizosphere samples of different soils, plants and geographical locations may determine which Acidobacterium and Verrucomicrobium groups are actually related to the presence of plants.
Involvement of Acidobacteria and Verrucomicrobia in plant and soil health Acidobacterium and Verrucomicrobium are bacterial groups that were implicated to correlate with take-all decline in wheat monoculture (Sanguin et al., 2009). A microarray study demonstrated that Verrucomicrobium was correlated with the outbreak stage and Acidobacterium with the suppression of take-all decline. This was the first study relating these bacterial groups to suppressiveness. The novel Acidobacterium and Verrucomicrobium strains and detection system described in this thesis are excellent tools to further study the correlation of these two groups and soil/plant health. An initial step would be a screening - using real time PCR based systems - of different soil types and compartments (i.e. bulk and rhizosphere soil) known to be conducive or suppressive
186 Chapter 9
to different diseases. Such data would demonstrate to which ecological phenomena (e.g. disease suppressiveness) Acidobacteria and Verrucomicrobia would be correlated.
187 General Discussion
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190 Appendix
Figure A1
A ZNBB5 CHC25 CR28 IRVE CNC16 ORAC ONA9 CHC12 Z35 CHC8 C20
B Pearson correlation (%) 20 40 60 80 100 CNC16 Verrucomicrobium ONA9 Luteolibacter CHC8 unclassified Verrucomicrobiaceae C20 Luteolibacter ZNBB5 unclassified Verrucomicrobiaceae CR28 unclassified Verrucomicrobiaceae IRVE unclassified Verrucomicrobiaceae Z35 unclassified Verrucomicrobiaceae CHC12 Luteolibacter ORAC Holophagae CHC25 Holophagae
Legend. BOX-PCR profiling of the different Holophagae, Luteolibacter and Candidatus genus Rhizosphereae isolates. A, original gel; B, nomalized gel with Pearson correlation analysis.
191
orial/Altschul-1.html) ://www.ncbi.nlm.nih.gov/BLAST/tut (http chance by occurring nment alig the of probability the esent repr results blast in e-values
Similarity at protein sequence level (calculated from (calculated level sequence protein at Similarity the amino acid sequence alignment as 1 – p-distance) p-distance) – 1 as alignment sequence acid amino the
Similarity at DNA sequence level (calculated from from (calculated level sequence DNA at Similarity the nucleotide sequence alignment as 1 – p-distance) p-distance) – 1 as alignment sequence nucleotide the
Best match determined by BLASTx (translated nucleotide nucleotide (translated BLASTx by determined match Best sequence blasted against the nonredundant protein database) database) protein nonredundant the against blasted sequence
DSM 4136] - ZP_02928598 ZP_02928598 - 4136] DSM
CNC16 multicopper oxidase, type 3 [ 3 type oxidase, multicopper CNC16 89.2 96.0 0.0 0.0 96.0 89.2 spinosum Verrucomicrobium
Verrucomicrobium Verrucomicrobium
DSM 4136] - ZP_02928598 ZP_02928598 - 4136] DSM
IRVE multicopper oxidase, type 3 [ 3 type oxidase, multicopper IRVE 64.9 61.0 3.0 e-146 3.0 61.0 64.9 spinosum Verrucomicrobium
DSM 4136] - ZP_02928598 ZP_02928598 - 4136] DSM
CR28 multicopper oxidase, type 3 [ 3 type oxidase, multicopper CR28 64.9 61.0 3.0 e-146 3.0 61.0 64.9 spinosum Verrucomicrobium
) Rhizosphereae genus Canditatus
Verrucomicrobiaceae Unclassified
DSM 4136] - ZP_02928598 ZP_02928598 - 4136] DSM
ONA9 multicopper oxidase, type 3 [ 3 type oxidase, multicopper ONA9 62.4 60.0 1.2 e-139 1.2 60.0 62.4 spinosum Verrucomicrobium
DSM 4136] - ZP_02928598 ZP_02928598 - 4136] DSM
CHC12 multicopper oxidase, type 3 [ 3 type oxidase, multicopper CHC12 61.9 59.0 2.5 e-140 2.5 59.0 61.9 spinosum Verrucomicrobium
Luteolibacter Luteolibacter
number number (%) (%)
a (%)
Isolate Accession Accession Isolate Best match Best Sim Protein Protein Sim DNA Sim e-Value b
a a b c c isolates.
d Supplemetantary Table 1 1 Table Supplemetantary Best matches (at DNA and protein level) of putative laccase genes present in different different in present genes laccase putative of level) protein and DNA (at matches Best subdivision 1 1 subdivision Verrucomicrobia 192 Figure A2
Potato 12
10
8
6
4
2 1 241 241 241 241 241 241 241 24 Candidatus genus Log eq cell number per gram of dry soil General bacteria Group 1 Group 3 Group 4 Group 6 Holophagae Luteolibacter Rhizospheria
12 Leek
10
8
6
4
2 1 241 241 241 241 241 241 241 24 General Candidatus genus Log eq cell number per gramsoil of dry bacteria Group 1 Group 3 Group 4 Group 6 Holophagae Luteolibacter Rhizospheria
12 Grass 10
8
6
4
2 1 241 241 241 241 241 241 241 24 Candidatus genus
Log eq cell number per gram of soildry General bacteria Group 1 Group 3 Group 4 Group 6 Holophagae Luteolibacter Rhizospheria
Bulk soil Outer rhizosphere Inner rhizosphere
1, 2, 4 – Sampling month
Legend Log of equivalent cell numbers, determined by real time PCR, of the ‘general’ bacteria, Acidobacteria subgroups 1, 3, 4, 6 and Holophagae and Verrucomicrobia subdivision 1 Luteolibacter and Candidatus genus Rhizospheria in the different soil sompartments, plant types and growth stages.
193 Figure A3
Potato 5.5
5.0
pH 4.5
4.0 124
5.5 Leek
5.0 pH 4.5
4.0 124 Grass 5.5
5.0 pH 4.5
4.0 1 24
Bulk soil Outer rhizosphere Inner rhizosphere
1, 2, 4 – Sampling month
Legend pH measurements in the different soil compartments, plant types and growth stages.
194 Samenvatting
Het is algemeen aanvaard dat minder dan 1% van de totale bacteriële diversiteit kan worden opgekweekt in het laboratorium. Dit maakt microbieel ecologisch onderzoek in verschillende ecosystemen, zoals het plant-bodem systeem, ingewikkeld. In het onderzoek beschreven in dit proefschrift, is een geslaagde poging gedaan om bacteriën die behoren tot de phyla van Acidobacteria en Verrucomicrobia op te kweken vanuit de rhizosfeer. Leden van beide taxonomische groepen zijn nog nooit opgekweekt uit bodemmonsters die zijn genomen in de buurt van plantenwortels. Bacterie-isolatie is belangrijk omdat hiermee experimenteel onderzoek kan worden verricht waardoor er meer duidelijkheid kan worden verschaft over de ecologie van beide phyla in het plant- bodem ecosysteem.
Uit de rhizosfeer van aardappel en/of prei zijn de volgende stammen geïsoleerd: vier die behoren tot Luteolibacter, vier tot kandidaat geslacht Rhizospheria en één tot Verrucomicrobia (allen behorend tot Verrucomicrobia subdivisie 1) en twee Holophagae stammen, beiden behorend tot de Acidobacteria. De taxonomie van deze stammen is bepaald op basis 16S rRNA genen en verder zijn de mogelijke interacties van deze stammen met planten bepaald, zoals het vermogen om te groeien op wortelexudaten. Het bleek dat de Holophagae stammen taxonomisch en fenotypisch sterke gelijkenis met elkaar vertoonden en dat de Verrucomicrobia subdivisie 1 stammen taxonomisch en fenotypische juist meer uiteenlopend waren.
Real-time PCR werd gebruikt om het lot van Acidobacteria en Verrucomicrobia subdivisie 1 aan te tonen in plant-bodem ecosystemen en deze techniek bleek voor dit doel uitstekend geschikt te zijn. In vitro toetsen en kolonisatie studies in plant-bodem microkosmos systemen gaven inzicht in de ecologie van de soorten behorend tot de Holophagae en kandidaat geslacht Rhizospheria (voorgesteld als een niet eerder beschreven taxonomische groep binnen Verrucomicrobia subdivisie 1). Van beide groepen werd verondersteld dat ze associaties aan zouden gaan met diverse plantensoorten. Aan de andere kant werden juist hogere aantallen Luteolibacter cellen aangetroffen in bulk grond (geschat met behulp van real-time PCR) in vergelijking met overeenkomstige rhizosfeer grond in de plant-bodem microkosmos systemen. Uit onderzoek in het veld bleek dat Acidobacteria subgroepen 1, 3, 4 in hogere celaantallen
195 aanwezig waren in bulk grond dan in overeenkomstige rhizosfeer grond. Celaantallen van subgroep 6 van de Acidobacteria waren op hun beurt weer hoger in de rhizosfeer dan in de omliggende bulk grond. Uit de verdeling van Holophagae celaantallen over de verschillende rhizosfeer en bulk gronden bleek dat deze Acidobacteria groep een specifieke niche in de rhizosfeer van prei inneemt.
Succesvolle opkweek van vertegenwoordigers van Acidobacteria en Verrucomicrobia subdivisie 1 bleek experimenteel onderzoek in de nabijheid van planten te vereenvoudigen en deze studies waren nodig om nieuwe ecologische theorieën over mogelijke microbiële interacties in het plant-bodem ecosysteem te kunnen ontwikkelen. Er is een nieuwe richting gegeven aan het onderzoek over de ecologie van leden behorend tot de phyla van Acidobacteria en Verrucomicrobia, respectievelijk Holophagae en kandidaat geslacht Rhizospheria, in plant-bodem ecosystemen. Toekomstig onderzoek zal zich richten op mogelijke interacties met planten en andere micro-organismen die in associatie met planten leven. Dit onderzoek draagt bij aan de huidige kennis over de ecologie van beide groepen in het plant-bodem ecosysteem en hun relevantie tot plantengroei.
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Aknowledgements
First of all, I would like to thank my supervisor, Dick, for his amazing support, encouragement, ideas and challenges throughout these last four years. I learned from you that science is not only my work but also my hobby and passion. Our brain storms taught me to approach research with an open and critic mind and made possible the completion of my thesis. My other supervisor, Leo, it was a daily pleasure to work with you. Without your guidance I am sure the course of my PhD would not be as smooth. The fact that you would critically challenge my work not only prepared me to discuss my subject but pushed me to go further than I would. I would like to thank my colleagues at PRI. The Biointeractions cluster is a working environment that I will miss for the rest of my career. Your help, friendship, teachings, coffee and lunch breaks side by side with our scientific discussions and relaxed working environment made PRI a happy place to work. I would like also to thank my colleagues at the Microbial Ecology department of RUG for the friendly working discussions and advises. The work of my thesis did not happen only at PRI and RUG, it embraced bench and field work in other places. Therefore, I direct grateful thanks for my colleagues from Vredepeel experimental field (WUR) that helped me with part of my field experiments, Caroline Plugge from the Microbiology Department of WUR and Isabelle George from the Catholic University of Louvain (Belgium). Your help improved the quality of my thesis. I would like to thank the Ecogenomics program for support with my scholarship and for financing my research.
During my PhD I met other students and researchers that became my friends. Your advices, company, friendship and support helped during all the work I performed on my thesis. Thank you Rady, Cauca, Sarrah, Sara, Marhmood, Ronald, Barbara, Rahim, Fernando, Fedor, Ilya, Manoel, Rodrigo Costa, Katia, Goes, Vladmir, Oscar, Odette, Joeke, Els, Carin, Mirjam, Pieter, Lia, Vilma, Jan and many others. My paranimfen Isabela and Saulo thanks for our friendship during the past 4 years. In Wageningen, one learns that we don’t know as much about the world as we thought. We are so many from so many different nationalities, this teach us that cultural differences exist and must be respected. I made so many good friends that it would be unfair to name some and leave the others out. But I must say that meeting you all showed me that we live in an amazing and diverse world. Everyone should explore this diversity. During the period I lived in Wageningen I participated in different extracurricular activities that made my
197 life complete. I love to thank the International Club of Wageningen for offering me a place where I could meet many of the great friends I made in Wageninen and feel at home. I own the places where I practiced judo to keep my body healthy and my mind in peace, thank you. Many thanks for the friends I made while I was dancing salsa, especially in the International Student Organization of Wageningen, you are just amazing. I hope you perpetuate teaching newcomers that dancing relax the mind and free our souls. I also give thanks for all my dear friends that I met by chance during my life in the Netherlands. Concerning this aspect, I believe the Netherlands is a special place, where one learns to live, to the best of my knowledge, in the most tolerant place in the world. I learned and respect the Dutch way, I define it as the special way the Dutch are raised that create a common sense of unit, respect and friendly environment that I hope to find everywhere I will be in my life time. I thank the Netherlands for allowing me to have probably the best period of my life.
Obrigado a todos os meus amigos brasileiros que eu fiz em Wageningen. Eu não consegui por o nome de nenhum de vocês em meus agradecimentos porque quando cheguei a este paragráfo não resisti e chorei. Chorei por todos os bons momentos que vivi com vocês. Chorei porque sei que vou sentir saudades arrebatadoras. Chorei porque apesar de saber que nossa amizade vai estar sempre impressa na minha alma, provavelmente não vou passar tanto tempo com vocês novamente. Acredito que somente aqueles que nasceram brasileiros poderiam me entender. Nosso país, nossa cultura, nossa energia corre não somente numa raça, cor ou etnia, mas na simples, bela e forte energia que nos une sobre uma mesma nacionalidade. Agradeço a todos vocês e do fundo do meu coração digo que só consegui começar, sobreviver durante o percurso, terminar meu doutorado e seguir em frente para um novo início porque vocês estavam na minha vida. Tia De e Zé, amo vocês, sua figura maternal e paternal me ajudaram concluir mais esta etapa da minha vida. Agradecimentos e saudades incomensuráveis daqueles que eu gostaria que estivessem comigo nesse momento mas infelizmente não se encontram entre nós.
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About the Author
Ulisses Nunes da Rocha (Rio de Janeiro, 1981) started his scientific life in the first year of his high school in the Scientific Vocation program of the FIOCRUZ (Oswaldo Cruz Foundation, Rio de Janeiro, Brazil). After one year assisting in a Protozoology laboratory he carried out a small project (1.5 years) on the infection of Toxoplama gandii in mices. After that (1999) he decided to follow a bachelor in Biology where he developed experience in anatomy and cell morphology, general and inorganic chemistry and entomology. In August/2001 he gave his first steps in microbiology in the Pontifical Catholic University of Rio de Janeiro where he carried out his bachelor thesis entitled ‘Influence of electrical fields and nitrate addition by electrokinetics on tropical residual soil microorganisms’. After his bachelor he pursued microbiology as a career. In 2003, he was offered a scholarship to study agriculture microbiology at the Federal University of Viçosa (Brazil) where he managed to continue the studies of his bachelor thesis. While finishing his master he taught biophysics at Univiçosa (Minas Gerais, Brazil) and received his M.Sc degree at the end of 2005. At that time he made the first contacts with Prof. van Elsas and Dr. Leo van Overbeek and started his PhD in March 2006. During his PhD, completed in 2010, he studied the ecology of hitherto uncultured bacteria in the rhizosphere. His work was presented in different international and national conferences and he co-authored several scientific publications. After completion of his PhD Ulisses plans to move to the University of Florida (USA) and study the ecology of Candidatus Liberibacter asiaticus, a yet-to-be cultured bacteria that is correlated with citrus greening, an emerging disease considered to be the most destructive citrus disease worldwide.
Existing and accepted publications of Ulisses Nunes da Rocha *Denote a publication part of this thesis
8. *Nunes da Rocha UN, van Elsas JD, van Overbeek LS (2010) Real-time PCR detection of Holophagae (Acidobacteria) and Verrucomicrobia subdivision 1 groups in bulk and leek (Allium porrum) rhizosphere soils. J Microbiol Methods, accepted. 7. Andreote FD, Nunes da Rocha U, Araújo WL, Azevedo JL, Van Overbeek LS (2010) Effect of bacterial inoculation, plant genotype and developmental stage on root-
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associated and endophytic bacterial communities in potato (Solanum tuberosum). Antonie van Leeuwenhoek 97: 389-399. 6. *Nunes da Rocha U, Andreoti FD, Azevedo JL, van Elsas JD, van Overbeek L (2010) Cultivation of hitherto uncultured bacteria belonging to the Verrucomicrobia subdivision 1 from the potato (Solanum tuberosum) rhizosphere. J Soil Sediments (accepted). IF: 2.8 5. *Nunes da Rocha U, van Elsas JD, van Overbeek L (2009) Exploration of hitherto- uncultured bacteria from the rhizosphere. FEMS Microbiol Ecol 69: 313-328. IF: 3.4 4. Andreote FD, De Araújo WL, De Azevedo JL, van Elsas JD, Nunes da Rocha U, van Overbeek LS (2009) Endophytic colonization of potato (Solanum tuberosum L.) by a novel competent bacterial endophyte, Pseudomonas putida strain P9, and its effect on associated bacterial communities. Appl Environm Microbiol 75: 3396-3406. IF: 3.8 3. Nunes da Rocha U, Tótola MR, Pessoa DMM, Júnior JTA, Neves JCL, Borges AC (2009) Mobilisation of bacteria in a fine-grained residual soil by electrophoresis. J Hazard Mat 161: 485-491. IF: 3.0 2. van Overbeek L, Gassner F, van der Plas CL, Kastelein P, Nunes da Rocha U, Takken W (2008) Diversity of Ixodes ricinus tick-associated bacterial communities from different forests. FEMS Microbiol Ecol 66: 77-84. IF: 3.4 1. Borges MT, Nascimento AG, Nunes da Rocha U, Tótola MR (2008) Nitrogen starvation affects bacterial adhesion to soil. Braz J Microbiol 39: 457-463. IF: 0.6
Submitted publications
4. *Nunes da Rocha U, van Elsas JD, Plugge C, Andreoti FD0DQGLü-Mulec I, Ausec L, van Overbeek LS. Isolation and partial characterization of Holophaga, Luteolibacter, unclassified Verrucomicrobia and Verrucomicrobium spp. from the leek (Allium porrum) rhizosphere 3. *Nunes da Rocha U, van Elsas JD, van Overbeek L. Rhizocompetence of culturable Holophaga (Acidobacteria) sp. in the leek (Allium porrum) rhizosphere 2. *Nunes da Rocha U, van Elsas JD, van Overbeek L. Different rhizosphere competence in two Verrucomicrobium subdivision 1 strains previously isolated from the leek (Allium porrum) rhizosphere 1. *Nunes da Rocha U, van Elsas JD, van Overbeek L. Distribution of different Acidobacteria and Verrucomicrobia subdivision 1 groups over compartments of different plant species
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