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Fungi and mycorrhizal symbiosis
The fungi, belonging to the kingdom of fungi, are eukaryotic and can be either unicellular (yeasts) or multicellular microorganisms. When multicellular, fungi form most of the time hyphae that are in average a few micrometers (< 10) wide and few tens of micrometers long. The main characteristics of fungi are that they contain chitin in their cell wall instead of cellulose and accumulate glycogene instead of amidon. As they are heterotrophic, they constitute the other way to channel energy in the soil from carbohydrates produced by photosynthesis. Fungal energy channels are considered as slow cycles, because they are favored by acidic soils low in available nutrients, recalcitrant organic materials and high C/N ratio in soil, leading to relatively long generation times (Blagodatskaya and Anderson, 1998; Högberg et al., 2007). Fungi have been shown to use organic substrates more efficiently (Holland and Coleman, 1987; Sakamoto and Oba, 1994), i.e. they form more biomass from the same amount of substrate than bacteria (Joergensen and Wichern, 2008). Indeed, estimations of microbial biomass based on biochemical markers derived from cell-wall compounds such as glucosamine (derived from fungal chitin) and muramic acid (derived from bacteria) showed that fungal C constitutes in average 70% of the total microbial C (Joergensen and Wichern, 2008) in most of the situations studied (Table 1).
Formation of mycorrhizal roots
The process of mycorrhization can be divided into several steps (Martin et al., 1997) but the starting point is always the recognition between plants and mycorrhizal fungi to allow the early contacts between the two partners of the symbiosis. As underlined by Sanders ³the problem for both plants and mycorrhizal fungi is how to tell each other to HVWDEOLVK D V\PELRVLV DQG KRZ WR GR WKDW ZLWKRXW VHWWLQJ RII VRPH RI WKH SODQW¶V FRPSOH[ GHIHQVH UHDFWLRQV´ 2QH RI WKH ODVW DGYDQFHPHQW RI our knowledge in mycorrhizal signaling comes from the demonstration of the role of small protein produced produced by ECM (Plett et al., 2011) and AM (Kloppholz et al., 2011) fungi. These mutualistic fungal effector proteins allow symbiosis formation and aOORZ WKH IXQJXV WR PDQLSXODWH WKH SODQW¶V GHIHQVH response, respectively. In the case of ECM symbiosis, this achievement was facilitated by the whole genome sequencing of the ECM fungus Laccaria bicolor and genome-wide transcriptome studies on L. bicolor/poplar ECM (Martin et al., 2008). These studies revealed that a protein of 68 amino acids, known as Mycorrhizal induced Small Secreted Protein7 (MiSSP7), highly accumulated in ECM root tips (Martin et al., 2008) is produced only when the fungus makes the symbiosis and not when the fungus is in a free-living state (Plett et al., 2011). In addition, the fungus released this protein without direct contact with its host-plant (Poplar) or in presence of Arabidopsis thaliana which is not capable of forming any type of mycorrhizal symbiosis, indicating that diffusible plant-produced signals must be released by the root but that they are not specific. Finally, Plett et al. (2011) demonstrate that MiSSP7 enters plant cells, probably by endocytosis, and accumulates in the plant nucleus as described in figure 6. In addition to early signalling, MiSSP7 also controls Hartig net formation by inducing transcripts involved in cell wall remodeling and auxin homeostasis. Taken as a whole, it seems that MiSSP7 is really a fungal effector necessary for ectomycorrhiza formation (Sanders, 2011).
Nutrient exchange in mycorrhizal roots
The fungus is heterotroph for carbon, the plant supplies carbohydrates derived from photosynthesis at the level of common interfaces in mycorrhizal roots, ie in the Hartig net for ECM and arbuscules for AM symbiosis. Mycorrhizal symbiosis constitutes therefore a privileged pathway to drive plant C to the soil through the hyphae network. Data from a range of microcosm-based labeling studies suggest that 7-30% of net C fixation is allocated to ectomycorrhizal mycelium and that 16-71% of this C is lost by respiration (Leake et al., 2004; Jones et al., 2009). Of the nutrient exchanged between fungal and root cells, those of N and P are the most extensively studied. Regarding N, evidence have been gained in AM symbiosis that a net transfer of N taken up by the extraradical hyphae to carrot roots occurs at the level of intraradical hyphae (Govindarajulu et al., 2005). However, in the field, direct and indirect evidence indicate high variability of N transfer among fungal species (Gobert and Plassard, 2008; Chalot and Plassard, 2010). In contrast to N, the positive effect of mycorrhizal symbiosis on P nutrition of the host plant has been repeatedly demonstrated (Smith and Read, 2008). In AM plants, the occurrence of two pathways to take up P from the soil solution has been proposed (Fig. 8) in which direct plant P uptake can be replaced by fungal uptake at different degrees (Smith et al., 2003; 2004; 2011, Facelli et al., 2010).
Such pattern is strongly supported by the discovery of mycorrhiza-inducible Pi transporters specifically expressed at the level of arbuscules in herbaceous plants (as for example StPT3 in potato, Rausch et al., 2001; MtPT4 in Medicago, Harrisson et al., 2002) or perennial plants (PtPt10 in poplar, Loth-Pereda et al., 2011) among others plant Pi transporters reviewed by Javot et al. (2007a) and Bucher (2007).
Role of soil food web in root architecture and growth
Root architecture is a fundamental aspect of plant productivity. Therefore, breeding attempts have focused on developing larger root systems in crop plants. In addition to abiotic factors, such as patchy supply of different nutrients (Drew, 1975; Zhang and Forde, 2000), water stress (Fukai and Cooper, 1995) and soil compaction (Iijima et al., 1991), also biotic components of the root environment strongly influence the architecture of the root system.
For example, rhizosphere bacteria have been shown to affect root growth by the release of signal molecules, such as hormones, toxins or other metabolites (Arshad and Frankenberger, 1998; Phillips et al., 2004; Matiru and Dakora, 2005). These complex interactions are further complicated by the fact that protozoan and nematodes grazers are a strong selective force that affects bacterial activity in soils and in the rhizosphere (Blanc et al., 2006; Griffiths et al., 1999; Ronn et al., 2002). Jentschke et al. (1995) found that the protozoa increased N uptake because hormonal effects resulted in the changes in the root morphology and plant growth; the significant changes in plant roots in the presence of protozoa might be mostly due to the effects of protozoa grazing on the rhizosphere bacteria. Despite the positive effect of protozoa on root architecture mycorrhiza had opposite effects on root architecture.
Table of contents :
CHAPTER 1. REVIEW OF LITERATURE
1. Importance of soil food web in soil plant nutrient cycling
2. Components of soil food web
2.1. Bacteria
2.2. Fungi and mycorrhizal symbiosis
2.2.1. Classification of mycorrhizal roots
2.2.2. Formation of mycorrhizal roots
2.2.3. Nutrient exchange in mycorrhizal roots
2.3. Nematodes
2.4. Protozoa
3. Functionning of soil food web in rhizosphere
3.1. Importance of rhizosphere in microbial populations
3.2. Role of soil food web in root architecture and growth
3.3. Role of soil food web in rhizosphere N cycle
3.4. Role of ectomycorrhizal symbiosis on N cycling
3.5. Role of soil food web in rhizosphere P cycle
3.6. Role of mycorrhizal symbiosis on P cycling
4. Hypothesis and objectives of thesis
5. References
CHAPTER 2. MATERIAL AND METHODS
1. Biological material and culture method in routine conditions
1.1. Plant and fungus
1.1.1. Seed germination
1.1.2. Fungal cultures
1.1.3. Mycorrhizal synthesis
1.2. Bacteria
1.3. Nematodes
1.3.1. Isolation and multiplication of nematodes
1.3.2. Sterilization and testing of nematodes with Bacillus subtilis (111b)
2. Specific culture conditions
2.1. 15N labeling of bacteria
2.2. Phytate medium
2.3. Co-inoculation media
2.4. Co-inoculation experiment design
3. Quantitative assessment of microbial populations
3.1. Bacteria
3.2. Nematodes
3.3. Ectomycorrhizal degree
3.3.1. Acid hydrolysis
3.3.2. Colorimetric assay
4. Molecular methods
4.1. Bacteria
4.2. Nematodes
5. Plant and medium analysis
5.1. Root parameters and plant harvest
5.2. Solution extraction from solid medium
6. Chemical measurements
6.1.1. Nitrogen
6.1.1.1. 15N and total N
6.1.1.2. Free ammonium
6.1.1.3. Nitrate
6.1.2. Phosphorus
6.1.3. Sodium
7. Statistical analysis
8. References
CHAPTER 3 Grazing by nematodes on rhizosphere bacteria enhances nitrate and phosphorus availability to Pinus pinaster seedlings
1. Abstract
2. Introduction
3. Materials and methods
3.1. Plant production
3.2. Bacteria
3.3. Isolation, multiplication and identification of nematodes
3.4. Experimental design
3.5. Sampling and analytical procedures
3.6. Statistical analysis
4. Results
4.1. Nematode abundance
4.2. Shoot biomass and root parameters
4.3. 15N accumulation in shoot biomass
4.4. Nutrient accumulation in shoot biomass
5. Discussion
5.1. Nematode population
5.2. Effects of bacteria and bacteria plus nematodes on P. pinaster development
5.3. Effects of inoculation on the fate of bacterial 15N and N nutrition of P. pinaster seedlings
5.4. Effects of inoculation treatments on P nutrition of P. pinaster seedlings
6. Conclusion
7. Acknowledgements
8. References
CHAPTER 4 Phosphorus is required to the operation of microbial loop in the rhizosphere, independently of N availability
1. Abstract
2. Introduction
3. Material and methods
3.1. Fungal and plant material
3.2. Bacterial strain and nematode
3.3. Experimental design for co-inoculation
3.4. Medium nitrate measurement
3.5. Plant analysis
3.6. Bacterial and nematode populations
3.7. Statistical analysis
4. Results
4.1. Assimilation of 15NH4 + and phosphorus in bacterial biomass
4.2. Plant parameters
4.2.1. Root architecture
4.2.2. Plant biomass and N accumulation
4.2.3. P accumulation
4.2.4. 15N accumulation in root and shoot biomass
4.3. Nitrate concentration in medium
4.4. Abundance of bacteria and nematodes
5. Discussion
5.1. Root architecture and plant growth
5.2. Total N and 15N accumulation
5.3. P accumulation
5.4. Populations of bacteria, nematode grazers and 15N and P mineralization
6. Acknowledgement
7. References
CHAPTER 5 Phosphorus acquisition from phytate depends on efficient bacterial grazing, irrespective of the mycorrhizal status of Pinus pinaster
1. Abstract
2. Introduction
3. Material and methods
3.1. Fungal and plant material
3.2. Bacterial strain and nematodes
3.3. Experimental design for co-inoculation
3.4. Use of phytate by the fungus, the bacteria and the nematodes without plant
3.5. Plant analysis
3.6. Bacterial and nematode populations
3.7. Statistical analysis
4. Results
4.1. Use of phytate by microbial partners in pure culture
4.2. Effect of food web complexity on root development, plant growth and mineral nutrition
4.3. Evolution of bacterial and nematode populations in presence of P. pinaster seedlings
5. Discussion
5.1. Effect of plant and ectomycorrhizal symbiosis on bacterial and nematode populations
5.2. P accumulation and plant growth
5.3. Effect of food web on root growth and P acquisition
6. Acknowledgement
7. References
Supplimentary data
CHAPTER 6 Sodium toxicity and phytate use in the rhizosphere: a helping hand by microbial partners
1. Abstract
2. Introduction
3. References
GENERAL CONCLUSIONS AND PERSPECTIVES
1. Conclusions
1.1. Direct contribution of bacteria and nematode grazers in N & P mineralization and mineral nutrient acquisition by plants
1.2. Role of fungi, bacteria and their nematodes grazers on plant growth and root architecture
1.3. Evolution of bacteria and nematodes in different nutrient environment during plant growth
2. Perspectives
2.1. Nematode enzymatic activiy
2.2. Bacterial phosphorus labeling (32/33P)
2.3. Soil utilization as growth medium
2.4. Contribution of ectomycorrhizal symbiosis to N and P flow to the plant
3. References
