Phylogenetic and ecological characterisation of the root nodule bacteria from legumes in the African genus Lessertia

Macarena Gerding González

This thesis is presented for the degree

of

Doctor of Philosophy of

Murdoch University

December 2011

DECLARATION

I declare that this thesis is my own account of my research and contains as its main content work which has not previously been submitted for a degree at any tertiary education institution.

______Macarena Gerding González

I ABSTRACT

Legumes of the genus Lessertia have recently been introduced to Western Australia in an attempt to increase the diversity of perennial legumes available to help remediate effects of climate change and dryland salinity. These species were introduced along with a collection of rhizobia isolated from Lessertia in different agro climatic areas of the Eastern and Western Cape, South Africa.

The aim of the thesis was to perform a phylogenetic and ecological characterisation of rhizobia isolated from the herbaceous legume Lessertia spp. The first specific aim was to characterize 73 isolates of rhizobia associated with Lessertia spp. Isolates were authenticated on their original hosts and diversity at strain level was determined by ERIC- and RPO1-PCR fingerprinting analysis. Forty three distinct authenticated strains showed diverse colony morphology and growth rate.

The diversity and phylogeny of the 43 strains was examined via dnaK, 16srRNA and nodA partial sequencing. Strains were identified as Mesorhizobium except one strain that was identified as Burkholderia sp. 16s rRNA phylogeny of 17 strains was overall congruent with the dnaK phylogeny. The topology of the housekeeping genes phylogram was independent of the original host and geographical origin of the strains. The nodA sequences formed a unique cluster separate from previously known Mesorhizobium nodA sequences, and when compared with that of the 16s rRNA and dnak genes, showed a wide dispersion of nodulation genes among the different Mesorhizobium clades indicating possible genetic transfer.

A glasshouse experiment was set up to assess the symbiotic interaction between six Lessertia species (L. diffusa, L. incana, L.excisa, L. herbacea, L. capitata and L. pauciflora) and 17 rhizobial strains, selected from differerent phylogenetic clusters. The strains showed marked differences in their nodulation patterns. L.diffusa, L.herbacea and L.excisa were nodulated by

II most of the strains tested, and fixed nitrogen with 10, 7 and 4 strains respectively, while the other three hosts were quite selective. Interestingly, strains belonging to the same cluster in the nodA phylogeny showed similar nodulation patterns in the glasshouse experiment. WSM3636, WSM3612, WSM3565 and WSM3898 were selected for field experiments.

L. capitata, L. diffusa, L. excisa L. incana and L. herbacea were sown, along with their selected rhizobial strains, at five different agroclimatic areas of the Western Australian Wheatbelt: Badgingarra, Buntine, Katanning, Newdegate and Muresk. However, surviving plant populations in the field were lower than expected and although these species are perennials, most plants showed no or little regrowth after summer. To assess whether these problems were related to the adaptation and saprophytic ability of their root nodule bacteria, two field and a glasshouse experiments were carried out. Trap plants were established around surviving Lessertia plants at the Badgingarra site. At Buntine and Merredin, L. capitata, L.diffusa, L.excisa L. incana and L. herbacea were sown and inoculated with their appropriate inoculant to assess plant establishment, summer survival and nodulation. In addition, soils were collected from Lessertia plots at the five sites where Lessertia had been sown a year earlier. The soil samples were used to set up a soil trapping experiment in the glasshouse using the same five species of Lessertia. Nodulation was assessed and the root nodule bacteria were re-isolated and PCR fingerprinted. None of the trap plants at Badgingarra had nodules on their roots, suggesting that the inoculant strains were not able to survive in or to colonize that soil. Plant populations at Buntine and Merredin were again very low and the plants sampled were poorly nodulated. Results from the glasshouse experiment showed that L. herbacea and L. diffusa were able to nodulate in most of the soils, L. capitata, and L. incana only in two of them and L. excisa in none. Original inoculants could not be isolated from nodules, and strains occupying the nodules were identified as Rhizobium leguminosarum rather than Mesorhizobium spp. Since Mesorhizobium is known to have the ability to transfer symbiotic genes through a symbiosis island, the nodA gene was sequenced for all the “new” strains. The nodA sequences were again clustered with R. leguminosarum sequences, thus

III discarding the prospect of lateral gene transference from Mesorhizobium, and suggesting a competition problem with indigenous rhizobia.

Two field experiments were set up at Karridale, Western Australia, a site with more benign conditions to assess the effect of increased doses of an effective inoculant strain (WSM3565) with L. herbacea, and to study the competitive ability and symbiotic performance of different Mesorhizobium strains nodulating L. diffusa. Increasing the inoculation dose of L. herbacea with WSM3565 did not improve establishment and survival of the legume. Although WSM3565 nodule occupancy improved from 28 to 54% with higher doses of inoculation, none of the treatments increased L. herbacea yield over the inoculated control.

The inoculation of L. diffusa with strains WSM3598, 3636, 3626 and 3565 resulted in greater biomass production than the uninoculated control. These strains were able to outcompete resident rhizobia and to occupy a high (>60%) proportion of lateral root nodules.

The high numbers of resident R. leguminosarum in Western Australian soils, and their ability to nodulate Lessertia spp. represent a barrier to the successful introduction of the exotic legume genus Lessertia to Western Australian soils.

IV ACKNOWLEDGEMENTS

I would like to take this opportunity to thank the people who supported me throughout my PhD.

I would like to thank my supervisors for their assistance throughout this project. I am grateful to Graham O’Hara for his encouragement and guidance. Thanks to John Howieson for his great ideas and enthusiasm. My sincere thanks to Lambert Bräu for his permanent support and his invaluable help in the edition of this thesis.

I acknowledge the financial support of Murdoch University through the International Postgraduate Research Scholarship (IPRS) and a Murdoch Studentship. I would also like to acknowledge the financial assistance of Universidad de Concepción and the Chilean Government through the Mecesup Studentship.

I am grateful to Daniel Real and Ron Yates from the Department of Agriculture and Food of Western Australia (DAFWA) for their support in the field studies in the Wheatbelt and Karridale.

I would like to thank all the CRS research team. In particular to Sharon Fox, Bec Swift, Rob Walker and Vanessa Melino for their friendship and good vibes. Thanks to Julie Ardley for her friendship and for our very productive writing sessions. To Regina Carr for her technical support in my glasshouse experiments and for providing me with very good photos of Lessertia. My deepest thanks to my friends Regina and Steve Carr, Jason Terpolilli, Lambert Bräu, William Lee, Helena Moneta and “my aussie sister” Yvette Hill for all the happy moments we shared and for always being there for me and my family.

V I would like to thank my parents Marcos and María Inés for their unconditional support, wise advice, and for being an inspiration to me. I would also like to thank my parents in law María Eliana and Sergio for their genuine interest and encouragement.

I wish to thank my wonderful sons Martín and Pedro for their unconditional love, and for always being happy and healthy boys. You two make it all worthwhile.

Finally, my deepest thanks to my husband and best friend Felipe, for his love, patience, confidence in me and his invaluable help in my field experiments. I would not have completed this thesis without you.

VI PUBLICATIONS ARISING FROM THIS THESIS

Gerding, M., O’Hara, G.W., Bräu, L., Nandasena, K.G. and Howieson, J.G. 2012. Diverse Mesorhizobium spp. with unique nodA nodulate the South African herbaceous legume species in the genus Lessertia. Plant and Soil. DOI: 10.1007/s11104-012-1153-3

Refereed Conference Abstracts:

Gerding, M., O’Hara, G.W, Bräu L., Nandasena K. and Howieson J.G. 2009. Symbiotic and taxonomic diversity of root nodule bacteria isolated from the South African forage legume Lessertia spp. 16th International Congress on Nitrogen Fixation. Big Sky, Montana, USA. 14–19th June, 2009.

Gerding, M., O’Hara, G.W, Bräu L., Nandasena K., Real, D. and Howieson J.G. 2009. The role of competitive inffective nodulation in the poor establishment of Lessertia spp. in Western Australia. 15th Australian Nitrogen Fixation Conference. Margaret River, Australia. 8-13th November, 2009.

Gerding, M., O’Hara, G.W, Bräu L., Nandasena K., Real, D. and Howieson J.G. 2011. Mesorhizobium strains from the South African herbaceous legume Lessertia spp. Differ in their competitive ability in Australian soils. 17th International Congress on Nitrogen Fixation. Fremantle, Australia. 27th November – 1st December, 2011.

VII TABLE OF CONTENTS Pages

DECLARATION ...... I

ABSTRACT ...... II

ACKNOWLEDGEMENTS ...... V

PUBLICATIONS ARISING FROM THIS THESIS ...... VII

TABLE OF CONTENTS ...... VIII

CHAPTER 1: Literature Review ...... 1

1.1 Challenges for agriculture in South Western Australia ...... 3

1.2 Impact of climate change in South Western Australian agriculture ...... 5

1.3 Inclusion of perennial species into agricultural systems...... 6

1.4 Legumes ...... 7

1.5 South African Western Cape as a source of perennial species for South Western Australia ...... 10

1.6 Lessertia spp...... 13

1.7 Root nodule bacteria ...... 20

1.7.1 Mesorhizobium spp...... 21

1.8 Rhizobia-legume symbiotic interaction ...... 22

1.8.1 Recognition and plant infection ...... 22

1.8.2 Bacterial invasion and release into cells ...... 24

1.9 Rhizobial inoculants ...... 28

1.10 Constraints to adaptation of introduced rhizobia ...... 29

1.11 Approaches and Aims of the thesis ...... 30

CHAPTER 2. Authentication, characterisation and diversity of Lessertia spp. root nodule bacteria...... 33

2.1 Introduction ...... 35

2.2 Material and Methods ...... 37

2.2.1 Bacterial culture conditions ...... 37

2.2.2 Molecular Fingerprinting ...... 37

2.2.3 Authentication of isolates as root nodule bacteria ...... 40

VIII 2.2.4 pH range ...... 41

2.2.5 Sequencing the dnaK gene ...... 42

2.2.6 Sequencing the 16s rRNA gene ...... 43

2.2.7 Phylogenetic analysis ...... 44

2.3 Results ...... 45

2.3.1 Molecular Fingerprinting ...... 45

2.3.2 Rhizobia authentication ...... 47

2.3.3 pH range ...... 49

2.3.4 Phylogenetic analysis dnaK ...... 52

2.3.5 16s rRNA phylogeny...... 58

2.4 Discussion ...... 62

CHAPTER 3. Symbiotic effectiveness and nodulation gene phylogeny ...... 69

3.1 Introduction ...... 71

3.2 Materials and Methods ...... 72

3.2.1 Study of nodulation gene nodA ...... 72

3.2.2 Nodulation specificity and symbiotic effectiveness ...... 73

3.2.3 Sequencing of the symbiosis island insertion region ...... 75

3.3 Results ...... 76

3.3.1 Study of nodulation gene nodA ...... 76

3.3.2 Nodulation specificity and symbiotic effectiveness ...... 79

3.3.3 Sequencing of the Symbiosis island insertion region ...... 82

3.4 Discussion ...... 83

CHAPTER 4. Field adaptation and symbiotic promiscuity ...... 89

4.1 Introduction ...... 91

4.2 Materials and Methods ...... 92

4.2.1 Establishment of Lessertia spp. in the field (June 2007–March 2008) ...... 92

4.2.2. Inoculant survival and soil colonization (August 2008) ...... 94

4.2.3 Recovering rhizobia from five different sites in the Wheatbelt (October 2008) ...... 97

4.2.4 Establishment of Lessertia spp. in the field (July 2008) ...... 99

IX 4.2.5 Molecular Fingerprinting ...... 99

4.2.6 Authentication and assessment of effectiveness of field strains...... 100

4.2.7 Sequencing of the genes dnaK and nodA for soil trapped strains ...... 101

4.2.8 Nodulation of Lessertia spp. by commercial inoculants...... 101

4.3 Results ...... 104

4.3.1 Establishment of Lessertia spp. in the field (June 2007–March 2008) ...... 104

4.3.2 Inoculant survival and soil colonization ...... 105

4.3.3 Recovering rhizobia from five different sites in the Wheatbelt ...... 107

4.3.4 Establishment of Lessertia spp. in the field (Year 2008) ...... 108

4.3.5 Molecular Fingerprinting ...... 110

4.3.6 Authentication and assessment of effectiveness of field strains...... 112

4.3.7 Sequencing of the genes dnaK and nodA for soil trapped strains ...... 113

4.3.8 Nodulation of Lessertia spp. by commercial inoculants...... 115

4.4 Discussion ...... 118

CHAPTER 5. Competitive ability of Mesorhizobium strains from Lessertia spp...... 125

5.1 Introduction ...... 127

5.2 Materials and Methods ...... 128

5.2.1 Experiment 1: Effect of rate of inoculation of WSM3565 on symbiotic performance with L. herbacea...... 129

5.2.2 Experiment 2: Competitive ability of different strains of Mesorhizobium for nodulation of Lessertia diffusa ...... 131

5.2.3 Statistical analyses ...... 133

5.3 Results ...... 134

5.3.1 Effect of rate of inoculation of WSM3565 on symbiotic performance with L. herbacea...... 134

5.3.2 Competitive ability of different strains of Mesorhizobium nodulating Lessertia diffusa 139

5.4 Discussion ...... 143

CHAPTER 6. General Discussion ...... 149

6.1 Potential of Lessertia spp. for Western Australian stressful environments...... 151

6.2 Rhizobial competition and legume promiscuity as constraints for establishment of Lessertia spp...... 153

X 6.3 Future research ...... 156

6.4 Concluding remarks ...... 157

References ...... 159

Appendices ...... 183

XI

XII

CHAPTER 1: Literature Review

1 CHAPTER 1

2 CHAPTER 1

1.1 Challenges for agriculture in South Western Australia The Mediterranean type-climate area of South Western Australia, the eastern section of which is commonly known as the “wheatbelt” (Figure 1.1), has been extensively cleared of its native vegetation and developed for broad scale wheat and sheep farming during the last 200 years (Hobbs, 2003). Approximately five million hectares of this area are planted to wheat each year (Ludwig et al., 2009). The remaining natural vegetation is rich in species diversity and considered one of the world’s biological “hotspots” (Hobbs, 2003); and is generally deep rooted and very effective at using rainfall (Dear & Ewing, 2008).

Figure 1.1. Western Australia’s Wheatbelt

This area is characterized by wet, cool winters and dry, hot summers (Hobbs, 2003; Ludwig & Asseng, 2006) with 75% of rainfall occurring between May and October (Ludwig et al., 2009) and a summer drought of variable length (Di Castri, 1991). The annual rainfall is typically 300 to 600 mm (Dear & Ewing,

3 CHAPTER 1

2008). Soils are mainly sandy and duplex, with a permeable “A” horizon overlaying an impermeable “B” horizon (Ludwig & Asseng, 2006; McKenzie et al., 2004). Soils are usually poor in nutrients, having been heavily leached over millenia, and between 2 to 3% of the agricultural lands herein are affected by soil acidity (

Large areas of South Western Australia have the additional latent stress of secondary dryland salinity, which refers to human induced salinity, considered to be one of the region’s main environmental problems (McFarlane & Ruprecht, 2005; Pannell & Ewing, 2006). This has been primarily caused by the replacement of native vegetation (deep-rooted trees, shrubs and perennial grasses) by annual crops and pastures (George et al., 1997; Pannell & Ewing, 2006). This change in vegetation has disturbed the water balance by reducing evapotranspiration and water use, and increasing the amount of water percolating beyond the root zone, thus contributing to groundwater recharge (Clarke et al., 1998; Cocks, 2001; Cransberg & McFarlane, 1994; Dear et al., 2003; Lambers, 2003; Pannell & Ewing, 2006). This has caused watertables to rise resulting in soil salinity through the mobilisation of stored salts (Cocks, 2001; Cransberg & McFarlane, 1994; George et al., 1997; Pannell & Ewing, 2006) that are present at high levels in many Australian agricultural subsoils (Pannell & Ewing, 2006).

Salinisation of soils and groundwater is considered a serious form of land- degradation that causes important loses in crop productivity, as plant growth + - 2+ 2- is inhibited by high concentrations of Na , Cl , Mg and SO4 (Lambers, 2003). Currently, dryland salinity affects about 10% of agricultural land in Western Australia and it is estimated that by 2050 it will exceed 30%, causing substantial economic loss of agricultural production (Clarke et al., 1998; Cocks, 2001; Dear et al., 2003; Pannell & Ewing, 2006).

4 CHAPTER 1

1.2 Impact of climate change in South Western Australian agriculture The challenges to agriculture in this region have been accentuated in the last two decades with the increase in greenhouse gas concentrations that are producing a rise in global temperature and a modification in weather patterns (Desjardins et al., 2005), commonly referred to as climate change.

Climate change is already evident in the world and will have subsequent impacts on agriculture (Howden et al., 2007; Porter & Semenov, 2005). While some agricultural areas will be positively affected by climate change due to increased temperatures, CO2 concentrations and rainfall (Gregory et al., 2005; van Ittersum et al., 2003), South Western Australia is predicted to face an increase in temperature and in potential evaporation, but a decrease in the amount of precipitation, which will possibly cause losses in crop production (Gregory et al., 2005; Nielsen & Brock, 2009; Pittock, 2003; van Ittersum et al., 2003). These changes have been clear lately, as average temperatures have risen by 0.7°C over the last century (Pittock, 2003); and a significant decline in rainfall has been observed since the 1970s (Ludwig et al., 2009; Pittock, 2003). This has caused a reduction of over 50% in rainfall collected in Perth’s dams (Power et al., 2005) and in agricultural areas within the Western Australian wheatbelt (Ludwig et al., 2009; Pittock, 2003), which are largely attributed to the enhanced greenhouse effect but also to natural variability (IOCI, 2005). A further decrease in rainfall is predicted for the coming years, with a 15% reduction by 2030 (IOCI, 2005) that will impose severe impacts on the grain industry and on forage species (Ludwig & Asseng, 2006).

High rainfall usually leads to higher groundwater recharge (Asseng et al., 2001b; Ludwig et al., 2009; McFarlane & Ruprecht, 2005; van Ittersum et al., 2003). Therefore, the rate of spread of dryland salinity is expected to decline with the reduction in annual rainfall due to climate change. However, the high year to year rainfall variability will still generate occasional wet seasons that will contribute to the progress of salinization (Asseng et al., 2001b; McFarlane & Ruprecht, 2005), particularly in soils with sandy texture that have low water

5 CHAPTER 1 retention (Asseng et al., 2001a). Consequently, farming systems in South Western Australia need to integrate strategies to adapt crops to reduced water availability, to reduce the buildup of greenhouse gases by enhancing carbon accumulation in soils (Desjardins et al., 2005; Porter & Semenov, 2005) and to aid in the reduction of the advance of dryland salinity (Pittock, 2003; van Ittersum et al., 2003).

1.3 Inclusion of perennial species into agricultural systems Most of the farming systems in Australia rely on annual crops and pastures (Cocks, 2001) and efforts to improve water use of annual crops and pastures by plant breeding have had minimal impact (Pannell & Ewing, 2006). Introduction of perennial species, particularly pastures (Cransberg & McFarlane, 1994), is one of the most promising practices for salinity management (Cocks, 2001; Dear et al., 2003; George et al., 1997; Pannell & Ewing, 2006; Ward, 2006). Deep rooted perennials extract water from greater depth than annuals and have a larger water holding capacity (Asseng et al., 2001a; George et al., 1997). Moreover, perennial species have growth cycles that more closely match the availability of water in the soil and which allows the soil to completely dry before autumn, reducing the chances of groundwater recharge from early winter rainfall (Cocks, 2001; Dear et al., 2003; Ward, 2006).

Further to this, some deep rooted perennial species could also fulfill the demand for plant cultivars with an improved water use efficiency and higher drought tolerance to adapt agriculture to climate change (Howden et al., 2007; Pittock, 2003). There are several plant traits that can affect crop response to drought and may permit adaptation to climate change: reduced canopy size through smaller leaves, slower rate of leaf appearance and an increased root length that will allow the plant to explore soil and increase the water supply (Gregory et al., 2005; Porter & Semenov, 2005) and to eventually tolerate drought (Cocks, 2001).

6 CHAPTER 1

Inclusion of perennial species can also have a positive impact on wind and water erosion, by preventing high-velocity winds reaching the soil surface. Added to this, the soil-binding effect of the perennial root systems and the potential for reducing rates of acidification of the soil because of a greater capacity to use soil nitrate and water, with reduced accumulation of nitrates in the soil, are cited beneficial outcomes (Cransberg & McFarlane, 1994; Peoples et al., 1995).

One of the strategies to mitigate greenhouse gas emissions is to enhance carbon sequestration in agricultural soils (Desjardins et al., 2005). Accumulation of soil organic carbon requires a positive balance between inputs to and outputs from soil organic matter stocks (Jastrow et al., 2007). Carbon sequestration can be promoted by increasing root:shoot ratios (Conant et al., 2001; Jastrow et al., 2007) and consequently by favouring the selection of species with a well-developed root system (Rees et al., 2005). Around 50% of the carbon fixed by photosynthesis is transferred below ground and partitioned between root growth, rhizosphere respiration and soil organic matter (Rees et al., 2005). Perennial species accumulate carbohydrates in their roots on which they rely to survive and regrow after grazing and drought (Cocks, 2001). Therefore, the inclusion of perennials, particularly pasture species, in farming systems not only improves water use efficiency but also contributes to carbon sequestration (Conant et al., 2001).

1.4 Legumes The Leguminosae, also called The Fabaceae (Sprent, 2008), is the third largest family of flowering plants, comprising more than 18,000 species distributed worldwide and show a great diversity in morphology and ecology (Lewis et al., 2005). Some legume species are of human and agricultural significance due to the high protein content of their grains (Duranti & Gius, 1997) and their ability to increase soil nitrogen fertility and yield of subsequent crops (Howieson et al., 2008; Peoples et al., 1995). In addition, the inclusion of legumes into rotations and farming systems has been shown to increase carbon accumulation in soils (Conant et al., 2001; Desjardins et al., 2005).

7 CHAPTER 1

The success of legumes in agriculture relies largely on the ability to form a mutualistic nitrogen-fixing symbiosis with soil bacteria, commonly named rhizobia or root nodule bacteria, which allows them to increase soil nitrogen and to adapt and colonize soils of low fertility with a low economic input (Brockwell et al., 1995; Peoples et al., 1995; Sessitsch et al., 2002). Soil nitrogen derived from symbiotic nitrogen fixation further reduces the requirements for nitrogen fertilisers and the carbon cost of their manufacture (Desjardins et al., 2005), which is 1.4 kg of emitted carbon per kilogram of nitrogen manufactured (Conant et al., 2001). With the increasing need of nitrogen fertiliser to increase global food production, and the detrimental consequences of increased amount of toxins and nutrients in surface and ground waters (Porter & Semenov, 2005; Tilman et al., 2002), the use of legumes emerges as an alternative to reduce the environmental and economic costs of nitrogen fertilisation (Herridge, 2008; Howieson et al., 2008).

South Western Australian agriculture is based mainly on extensive, low input cropping systems, where pasture legumes play a key role in contributing to nitrogen fertility (Loi et al., 2005). With pasture legumes, the increase of nitrogen in the soil is maximised, because mainly animal products and fibre are removed from the system and much of the fixed nitrogen is returned to the ecosystem (Brockwell et al., 1995; Peoples et al., 1995). The commercial use of perennial legumes is currently dominated by a small number of species, of which lucerne (Medicago sativa) is the most prominent (Dear et al., 2003; Pannell & Ewing, 2006). Lucerne has had an important role in salinity management in Western Australia due to a high water use efficiency (George et al., 1997) and a high nitrogen fixation efficiency (Peoples et al., 1995). When lucerne is well adapted and established it is be able to achieve high yields and demonstrates high tolerance to drought and persistence (Dear et al., 2003). Moreover, its incorporation into cropping rotations has resulted in an improvement in soil structure and in increases in yield in the following crop (Cocks, 2001; Dear et al., 2003; Holford & Crocker, 1997). Nevertheless, lucerne is poorly adapted to winter waterlogging (Hill, 1996; Howieson et al., 2008), especially in the duplex soils (Dear & Ewing, 2008) present in large

8 CHAPTER 1 areas of South Western Australia (McKenzie et al., 2004). It is also sensitive to acidity and aluminium and is unable to tolerate continuous grazing (Cocks, 2001; Dear et al., 2003; Dear & Ewing, 2008; Hill, 1996; Howieson et al., 2008).

Although genetic improvements have allowed lucerne to grow in areas where it was previously considered unsuited (Dear et al., 2003), relying on this species alone as a perennial forage would lead to an increase of diseases and insect pests, as has been widely demonstrated in single species agricultural systems (Cocks, 2001). There is thus an urgent need for suitable perennial legumes that would not only feed livestock but also replace the nitrogen-fixing capacity of annuals, improve the water balance of Western Australian farming systems (Cocks, 2001; Dear et al., 2003), and adapt to acidic infertile soils and low rainfall conditions (Howieson et al., 2008).

There is a range of available deep-rooted perennial legumes that could be suitable for use in Australian farming systems (Li et al., 2008; Pannell & Ewing, 2006). Failure to incorporate some of these has been mainly due to their temperate origin and their inability to survive summer drought and winter waterlogging (Dear & Ewing, 2008).

Besides the desirable attributes of high seed production (Dear & Ewing, 2008; Howieson et al., 2008), rapid seedling development, low weed threat (Dear & Ewing, 2008) and a broad compatibility with rhizobia (Howieson et al., 2008), an essential consideration when introducing new species is that they are adapted to soils that match the target area in edaphic characteristics such as: clay content, pH, cation exchange capacity and inorganic fertility (Howieson et al., 2008), and in climate (Dear & Ewing, 2008).

Australian perennial herbaceous legumes such as Glycine spp., Desmodium spp., Kennedia spp., Swainsona spp. and Lespedeza spp. have been considered but they are less productive than commercial species and most of them do not tolerate intensive grazing and are toxic (Dear et al., 2003). In general, perennials in Australia are more frequent in areas where rainfall

9 CHAPTER 1 exceeds 500mm (Cocks, 2001). Therefore the exploration for new legume species to fulfil the demand for alternative perennial species has focused on the collection of overseas perennial germplasm, particularly from similar climatic regions (Cocks, 2001; Dear et al., 2003; Howieson et al., 2008).

1.5 South African Western Cape as a source of perennial species for South Western Australia There are five regions of the world with a mediterranean-type climate: Mediterranean Sea basin (latitude 30°- 45° N), Central Chilean coast (latitude 32°- 41°S), the south-western Cape Region of South Africa (latitude 32°- 35°S), the pacific coast of North America (latitude 31°- 41°N) and Southern Australia (latitude 32°- 38°S) (Di Castri, 1991; Gasith & Resh, 1999) (Figure 1.2). The Mediterranean climate is considered a transitional regime between temperate and dry tropical climates (Di Castri, 1991). Within these regions, Southern Africa and Southwest Australia are very closely related in terms of soil parental material, soil fertility, species diversity, high fire hazard and rainfall patterns (Figure 1.2) (Cowling et al., 2009; Di Castri, 1991) and show relatively mild quaternary climates as a result of strong maritime influences (Cowling et al., 1996). The South Western Australian flora also shows remarkable affinities to the Cape flora (Cowling et al., 2009), first reported by Bews (1921).

The south Western Cape Region of South Africa, also known as the Cape Floristic Region, is a unique centre of plant diversity (Cowling et al., 2009; Sprent et al., 2010) and one of the world’s biodiversity hotspots (Myers et al., 2000). The environmental conditions that have produced this rich flora are apparently the relative climatic and geomorphic stability, the high diversification and the low extinction rates (Cowling et al., 2009). There are a high number of endemic species due to particular combinations of soil pH and nutrient status (Deacon, 1991; Sprent et al., 2010; Spriggs & Dakora, 2009). Most of the species in this region are short lived perennials (Sprent et al., 2010) and Fabaceae is the fourth largest family (644 species in 38 genera) (Cocks & Stock, 2001).

10 CHAPTER 1

Figure 1.2. Geographic location of the five regions of the world with a mediterranean-type climate. (Map from California Natural Gardens (2010). http://www.californianaturalgardens.com/mediterranean_climate.php)

The Cape Floristic region comprises different biomes or vegetation types: Fynbos, Nama Karoo and Succulent Karoo, Forest and Thicket (Figure 1.3) (Robelo, 1996). The Fynbos corresponds to Mediterranean heathland that is periodically destroyed by fire (Cocks & Stock, 2001) and represents a significant part of the vegetation of The Cape floristic region. It is characterized by sandy, acid and low nutrient soils (Cowling et al., 1997). The Fynbos biome is the main contributor to species richness (Robelo, 1996). Legumes are well represented in terms of numbers of species, where Aspalathus is one of the genera that normally dominates the fynbos after fire (Cocks & Stock, 2001).

11 CHAPTER 1

Figure 1.3. Biomes of South Africa. Adapted from: Enviro-info (2001). Accessed in March 2008. http://www.environment.gov.za/enviro-info/nat/biome.htm

Karoo vegetation is typically composed of scattered small bushes with a good deal of bare soil or rocks in between. Soils are typically lime-rich and weakly developed (Hoffmann, 1996b). The Succulent Karoo biome has the highest diversity of plant species recorded for semi-arid vegetation (Milton et al., 1997). This biome is dominated by dwarf, succulent shrubs and is primarily determined by the presence of low winter rainfall and extreme summer aridity (Hoffmann, 1996b; Milton et al., 1997). Rainfall varies between 20 and 290 mm per year (Hoffmann, 1996b). In the Nama Karoo, rain is concentrated in summer but it varies between 100 and 520 mm per year. The dominant vegetation is a grassy, dwarf shrubland (Hoffmann, 1996a).

These three biomes (Fynbos, Nama Karoo and Succulent Karoo) have been prospected in search of new plant material and diverse, palatable herbaceous legumes with potential forage value have been collected, particularly from

12 CHAPTER 1 areas with annual rainfall between 150 to 600mm, and where coarse textured, infertile and low pH soils are common (Howieson et al., 2008; Yates et al., 2007). Within the species of interest the legume genus Lessertia has emerged as a possible alternative for stressful environments in South Western Australia.

1.6 Lessertia spp. The genus Lessertia is named after Baron J. P. B. de Lessert (1773-1847) patron of botany; (Allen & Allen, 1981; Lock & Schrire, 2005). Lessertia belongs to the subfamily Papilionoideae and tribe Galegeae and to the subtribe Coluteinae (Lock & Schrire, 2005; Sanderson & Liston, 1995) and comprises 50 to 60 species (Allen & Allen, 1981; Balkwill & Balkwill, 1999; McVaugh, 1968). It is closely related to the genera Astragalus, Oxytropis, Biserrula and to the Colutineinae subtribe, which form part of the Astragalean clade of the Galegeae phylogeny (Sanderson & Liston, 1995).

Early reports described Lessertia as difficult to define as a genus and that probably too many species had been established (Harvey, 1862). It is still regarded as a genus in need of revision (Lock & Schrire, 2005). Initially the genera Sutherlandia and Lessertia were separated based on differences in their corollas, stipes and the form of the pods (Balkwill & Balkwill, 1999). Recently, Goldblatt and Manning (2000) included the two Sutherlandia spp.: S. frutescens and S. microphylla within the genus Lessertia. In spite of this, some authors still regard Lessertia and Sutherlandia as separate genera (Lock & Schrire, 2005; van Wyk & Albrecht, 2008).

Lessertia spp. are distributed mainly in South African Western and Northern Capes (Balkwill & Balkwill, 1999; Goldblatt & Manning, 2000; Lock & Schrire, 2005), with some species extending to Tropical Africa (Botswana, Namibia, Western Zambia, Angola, Tanzania and Zimbabwe), encompassing areas of Mediterranean shrubland (fynbos), semi-desert and subtropical shrubland, wooded grassland and montane grassland (Figure 1.4) (Lock & Schrire, 2005). In general, representatives of this genus can be found in rocky and

13 CHAPTER 1 sandy grassland in seasonally wet areas (Lock & Schrire, 2005), along stream banks and mountains (Nkonki, 2004) and in coastal areas (Allen & Allen, 1981).

Overall Lessertia spp., exhibit many of the attributes desirable in a new species for Western Australian stressful environments. Lessertia comprises mainly perennial species (Nkonki, 2004) that include prostrate to decumbent herbs, sub-shrubs (Allen & Allen, 1981; Harvey, 1862; Lock & Schrire, 2005) and erect shrubs up to 1.5m tall (Balkwill & Balkwill, 1999). They are self- pollinating and prolific seeders, with large seeds that germinate vigorously and with fruits that adhere relatively strongly to the stems (Howieson et al., 2008). Importantly, some Lessertia spp. grow in areas of acid and infertile soils (Anderson & Hoffmann, 2007; Nkonki, 2003), which would be highly likely to adapt to similar conditions in Australia.

Figure 1.4. Distribution of Lessertia spp. in Africa. Source: African Plant database. Conservatoire et Jardin Botaniques (CJB), Ville de Geneva. Accessed in March 2010 http://www.ville-ge.ch/cjb/

Although several species are suspected of being poisonous to stock (Allen & Allen, 1981; Lock & Schrire, 2005), there are some species that form part of

14 CHAPTER 1 natural pastures in the Western and Northern cape that have been reported to be palatable for sheep and cattle (Breebaart, 2003; Howieson et al., 2008; Nkonki, 2004) and have shown tolerance to grazing (Anderson & Hoffmann, 2007). Moreover, they become prostrate under high-grazing pressure (Howieson et al., 2008), which may allow them to withstand continuous grazing (Cocks, 2001). These species are: L. capitata, L. diffusa, L. excisa, L. herbacea, L. incana and L. pauciflora.

These six Lessertia spp. have been recently introduced to Western Australia as potential perennial pasture legumes (Howieson et al., 2008) and included among the permitted species by the Australian Quarantine amendment proclamations 2006 and 2007 (AQIS, 2006). Therefore, the focus of this research will be on these six species, and descriptions of each are provided in the following sections.

1.6.1 Lessertia capitata E. Mey. A perennial decumbent species of 3 to 25 cm height (Nkonki, 2003) (Figure 1.5). The stem petioles and peduncles have short patent hairs. Pubescent leaflets with long peduncles, 5-8 flowered; high number of seeds per pod (Harvey, 1862). L. capitata is distributed along the Northern and Western cape and in Free State (Nkonki, 2003) (Appendix 1).

15 CHAPTER 1

Figure 1.5. Lessertia capitata showing prostrate growth and purple flower racemes.

1.6.2 Lessertia diffusa R.Br. Perennial procumbent subshrub of 20-45 cm height. Its stems, petioles and peduncles are pubescent, and flowers are pink and densely congested on inflorescences (Goldblatt & Manning, 2000; Harvey, 1862). Pods are obliquely ovate, semilunate, compressed and prominently veined. (Goldblatt & Manning, 2000) (Figure 1.6)

Figure 1.6. L. diffusa showing maturing seed pods.

16 CHAPTER 1

L. diffusa is distributed mainly in the semi-arid area of Namaqualand which falls within the Succulent Karoo biome in the Northern Cape (Anderson & Hoffmann, 2007; Rösch et al., 1997); and has also been reported in the Fynbos in the Western Cape (Goldblatt & Manning, 2000; Nkonki, 2003) and Namibia (Nkonki, 2003) (Appendix 2).

Although L. diffusa has been regarded as a poor competitor in its natural environment (Rösch et al., 1997), it has shown tolerance to grazing in natural grasslands where it adopts a low prostrate growth (Anderson & Hoffmann, 2007).

1.6.3 Lessertia excisa DC. Perennial procumbent species of 20 to 40 cm height (Harvey, 1862; Nkonki, 2003). The stems, petioles and peduncules are albopubescent with short patent hairs (Harvey, 1862). It can be distinguished from L. diffusa by its half- moon shaped pods (Goldblatt & Manning, 2000; Nkonki, 2004), by its loosely flowered raceme and by a calyx slightly covered with black, soft, short hairs (Nkonki, 2004) (Figure 1.7). It is distributed in Northern and Western Capes along sandstone slopes and flats (Goldblatt & Manning, 2000; Nkonki, 2003) (Appendix 3).

Figure 1.7. Lessertia excisa flower raceme, showing calyx covered in black short hair.

17 CHAPTER 1

1.6.4 Lessertia herbacea (L.) Druce. (basionym Colutea herbacea L. (CJB,2010) http://www.ville-ge.ch/cjb/). This species has been reported as a perennial (Nkonki, 2003), although it has occasionally been regarded as an annual species (Nkonki, 2004). It is a non- climbing herb that can grow from 30 to 100 cm tall (Nkonki, 2003). Its inflorescence is a loose raceme with many purple flowers (Figure 1.8). The pod is compressed, often veiny, hairless, shortly stipitate, obliquely ovoid- oblong with 2 to 6 seeds (Nkonki, 2004). It is distributed in the Western Cape, extending sparsely to the Northern and Eastern Cape at altitude from 30 to 961 m (Nkonki, 2003; Nkonki, 2004) (Appendix 4).

Figure 1.8. Lessertia herbacea flower.

1.6.5. Lessertia incana Schinz. This species is a perennial dwarf shrub of 2 to 70 cm height. Flowers are pink to purple, disposed in racemes (Figure 1.9). L. incana distribution ranges from Namibia to South Africa’s Northern Cape (Nkonki, 2003) (Appendix 5).

18 CHAPTER 1

Figure 1.9. Lessertia incana flower raceme. 1.6.6. Lessertia pauciflora Harv. var. pauciflora. This species has been described as a many stemmed, herbaceous, suberect or prostrate species (Harvey, 1862) with a height of 20 to 120 cm (Nkonki, 2003). Harvey (1862) reported it as a very variable plant, that has usually straight legumes, that can vary to curved or even falcate ones. Flowers are dull purple and white (Nkonki, 2003) (Figure 1.10). It has been reported in Namibia, Botswana, Limpopo, Free state, Northern, Western and Eastern Cape (Nkonki, 2003) (Appendix 6).

Figure 1.10. Lessertia pauciflora flowers.

19 CHAPTER 1

1.7 Root nodule bacteria Root nodule bacteria are a large and diverse group of soil bacteria that are able to live in symbiosis with legumes and fix atmospheric nitrogen (Sprent, 2001). The genus Rhizobium was initially defined by the ability of these organisms to induce nodule formation in legumes. In recent years the species have been taxonomically classified through polyphasic approaches including sequencing of housekeeping genes, DNA:DNA hybridization and morphological and biochemical characterizations (Sawada et al., 2003; Tindall et al., 2010). This has resulted in the emergence of several different genera and many new species of root nodule bacteria in recent years (Graham, 2008; Sawada et al., 2003; Sprent, 2009).

There are currently 92 symbiotic root nodule bacteria species (Weir, 2011), which are grouped in the following genera: Azorhizobium, Bradyrhizobium, Burkholderia, Cupriavidus, Devosia, Mesorhizobium, Methylobacterium, Ochrobactrum, Phyllobacterium, Rhizobium, Sinorhizobium (Ensifer), and Shinella (Chen et al., 2003; Graham, 2008; Moulin et al., 2001; Sawada et al., 2003; Weir, 2011; Willems, 2006).

Although Lessertia is a genus widely distributed in the South African Eastern and Western Capes, its root nodule bacteria have not been thoroughly investigated. Species of the tribe Galegeae are characterised by being usually temperate or boreal, all have indeterminate nodules, usually nodulated by fast growing α-rhizobia, with large variations in degrees of specificity (Heenan, 1998; Sprent, 2009). Nodulation of the genus Lessertia has been reported for 26 of the 50 species (Sprent, 2009) including L. diffusa, L. capitata and L. excisa (Grobbelaar & van Rooyen, 1979), L. herbacea (L.), L. incana and L. pauciflora (Allen & Allen, 1981). Yates et al (2004) reported L. (Sutherlandia) microphylla as able to nodulate with Australian native rhizobia.

Preliminary studies of some strains of root nodule bacteria isolated from Lessertia have identified them as Mesorhizobium spp. (J. Ardley pers.

20 CHAPTER 1 comm.). It has also been reported that the Mesorhizobium strain ICMP12690 isolated from New Zealand native legumes was able to nodulate Lessertia spp. (Weir, 2006). Therefore, the following section will focus on the genus Mesorhizobium.

1.7.1 Mesorhizobium spp. The genus Mesorhizobium belongs to the order Rhizobiales and the family Phyllobacteriaceae (Chen et al., 2005; Sawada et al., 2003). It was formerly included in the Rhizobium genus (Chen et al., 1995), but since a number of species formed a closely related group based on 16S rRNA gene sequence similarities that clustered separately from the other Rhizobium spp., and also showed slow or moderately slow growth rates they were renamed Mesorhizobium (Jarvis et al., 1997). It has been described as an acid producer and an intermediate in terms of colony growth rate between Rhizobium and Bradyrhizobium (Jarvis et al., 1997), although variations in growth rate within the genus have been reported (Gao et al., 2004; Nour et al., 1995; Sessitsch et al., 2002). Mesorhizobium are aerobic, Gram-negative non-spore-forming rods with one polar or subpolar flagellum (Chen et al., 2005; Jarvis et al., 1997).

The nodulation genes of Mesorhizobium spp. are mainly chromosome encoded (Kaneko et al., 2000), but for some strains of the species M. amorphae and M. huakuii they have also been found on megaplasmids (Wang et al., 1999; Zou et al., 1997). Other Mesorhizobium species have been reported to carry nodulating and nitrogen fixing genes on a symbiosis island (Kaneko et al., 2000; Nandasena et al., 2006; Sullivan & Ronson, 1998). This is described as a large (500kb) transmissible element that can be transferred to resident soil bacteria (Sullivan et al., 2002). Recent studies have described this transfer as a mechanism that may convert existing saprophytes into new nodulating organisms. Unfortunately, these new nodulating forms often have variable nitrogen fixation efficiency, which can lead to them becoming unfavourable competitors (Nandasena et al., 2007a).

21 CHAPTER 1

The genus Mesorhizobium comprises 19 recognized species that are able to nodulate a wide variety of legumes (Table 1.1) (Sprent, 2009; Weir, 2011).

Table 1.1. List of described species of the genus Mesorhizobium Species Hosts Source M. albiziae Albizia, Glycina max, Wang et al. (2007) Leucaena, Phaseolus vulgaris M. amorphae Amorpha fruticosa Wang et al. (1999) M. australicum Biserrula pelecinus Nandasena et al. (2009) Astragalus membranaceus M. camelthorni Alhagi sparsifolia Chen et al. (2011) M. caraganae Astragalus adsurgens, Guan et al. (2008) Caragana spp., Glycyrrhiza uralensis, Phaseolus vulgaris M. chacoense Prosopis alba Velázquez et al. (2001) M. ciceri Cicer arietinum Nour et al. (1994) M. ciceri bv. biserrulae Biserrula pelecinus Nandasena et al. (2007b) M. huakuii Astragalus sinicus Chen et al. (1991) M. loti Lotus spp. Jarvis et al. (1982) M. mediterraneum Cicer arietinum Nour et al. (1995) M. metallidurans Anthyllis vulneraria Vidal et al. (2009) M. opportunistum Biserrula pelecinus, Astragalus Nandasena et al. (2009) spp., Lotus peregrinus M. plurifarium Acacia senegal, Chamaecrista, de Lajudie et al. (1998) Prosopis, Leucaena M. robiniae Robinia pseudoacacia Zhou et al. (2010) M. septentrionale Astragalus adsurgens Gao et al. (2004) M. shangrilense Caragana, Vigna, P. vulgaris Lu et al. (2009) M. temperatum Astragalus adsurgens Gao et al. (2004) M. tianshanense Caragana sp., Glycyrrhiza Chen et al. (1995) spp., Sophora spp., Glycine max

1.8 Rhizobia-legume symbiotic interaction

1.8.1 Recognition and plant infection In general, there is strict specificity between host legumes and rhizobial symbionts (Hirsch et al., 2001; Perret et al., 2000). Even if host plants and rhizobia have the ability to enter into symbiosis with more than one partner, only certain combinations of symbionts result in nitrogen fixation (Perret et al., 2000). For example Rhizobium leguminosarum bv. trifolii is specific to the legume genus Trifolium (Broughton & Perret, 1999; Hirsch et al., 2001; Pueppke & Broughton, 1999), R. leguminosarum bv. vicieae to the genera Vicia, Pisum, Lathyrus and Lens (López-Lara et al., 1996) and Mesorhizobium ciceri bv. biserrulae is specific to Biserrula pelecinus (Nandasena et al.,

22 CHAPTER 1

2007b). However, the exact mechanisms responsible for this specificity are not clear.

Symbiotic infection comprises a series of complex coordinated events that require signal exchanges between legume and rhizobia (Gibson et al., 2008; Masson-Boivin et al., 2009; Oldroyd & Downie, 2008). Host specificity is determined even before there is a physical interaction between symbionts. It starts when seeds and then roots release flavonoids and isoflavonoids that are perceived by the appropriate bacteria, and which assist them to reach the root by chemotaxis (Gibson et al., 2008; Hirsch et al., 2001). At the same time, the bacterial transcription factor NodD is stimulated, triggering the co- ordinated expression of nod genes required for Nod Factor synthesis (Gibson et al., 2008; Hirsch et al., 2001; Perret et al., 2000).

Nod factor is a signalling molecule that consists of a chitin backbone that varies in size and saturation state with a N-linked fatty acid moiety attached to the non-reducing terminal sugar (Gibson et al., 2008; Oldroyd & Downie, 2008). Nod Factors are the signal required for entry of most rhizobia into the legume root and are therefore essential for nodulation (Gibson et al., 2008; Oldroyd & Downie, 2008; Perret et al., 2000). However, a compatible Nod factor is not the only requirement for effective nodulation. Bacterial cell surface polysaccharides such as exopolysaccharides (EPS), lipopolysaccharides (LPS) and cyclic-β-glucans, are also recognized by the plant and play a role in protecting rhizobia from host defence responses (Gibson et al., 2008; Martínez-Romero, 2003; Masson-Boivin et al., 2009).

Lectins, that are located in root hairs near the growing tip, are believed to facilitate bacterial attachment causing a localized increase in Nod factor concentration (Hirsch et al., 2001) enhancing nodulation capacity of rhizobia that do not produce the optimal Nod Factor for a particular legume (Oldroyd & Downie, 2008). Nevertheless, lectins do not bind Nod factors and are questioned as essential for nodulation (Hirsch et al., 2001). The presumed Nod factor receptors in legumes belong to the lysine –motif (LysM) receptor-

23 CHAPTER 1 like kinase family which will respond to specific Nod Factor structures (Gibson et al., 2008; Oldroyd & Downie, 2008; Radutiou et al., 2007). In the initial stages of plant infection, Nod Factors induce several morphological changes on the host. Root epidermal responses such as alkalinisation of the cytosol lead to root hair curling that traps intimately associated bacteria (Gibson et al., 2008; Oldroyd & Downie, 2008). At the same time, Nod factors induce calcium oscillations in nucleoplasm and nucleum associated cytoplasm, in a process known as calcium spiking (Gibson et al., 2008; Hirsch et al., 2001; Oldroyd & Downie, 2008). The latter gives rise to changes in host gene expression and modifies the plant hormone balance, inducing the dedifferentiation of root cortical cells into an actively dividing meristem which forms the initial nodule primordium (Gibson et al., 2008; Oldroyd & Downie, 2008).

1.8.2 Bacterial invasion and release into cells Developmental processes in the cortex during nodulation are different from those in the epidermis, and responses can be independent. Therefore, nodule organogenesis can occur without bacterial infection (Gleason et al., 2006) leading to formation of empty nodules or ineffective nodulation (Gibson et al., 2008; Perret et al., 2000). To produce bacterially infected nodules, both processes should be coordinated (Oldroyd & Downie, 2008), and after infection thread formation and propagation, bacteria should be released into the cytoplasm of cortical enlarged cells (Gibson et al., 2008; Perret et al., 2000).

For infection thread initiation there must be an intimate connection between the rhizobial cell surface and the plant cell wall (“Lectin recognition hypothesis”) (Hirsch et al., 2001). This process is considered the major “checkpoint” for the host in terms of whether to allow bacterial invasion to proceed (Gibson et al., 2008). In the process of infection thread growth, a high level of Nod Factor specificity is required as well as the presence of appropriate polysaccharides on the surface of the bacteria (Oldroyd & Downie, 2008). The infection thread will then allow growth of bacteria from the

24 CHAPTER 1 root surface to newly created endoploid cells, where it undergoes ramification (Gibson et al., 2008).

Infection threads can be intracellular or intercellular. The former requires significant cytoskeletal changes, whereas intercellular infection threads are organized lines of infecting rhizobia between cells with an associated lignification of adjacent walls, process that requires less stringency in the Nodfactor structure (Oldroyd & Downie, 2008). The infection thread starts at the tip of the root hair and is formed by hydrolysis of the cell walls and continues growth via deposition of new cell material (Gibson et al., 2008). Nod Factors and a variety of polysaccharides play a determinant role in permitting rhizobial invasion through infection thread by protecting rhizobia from host defence responses (Gibson et al., 2008; Oldroyd & Downie, 2008). The complete process of rhizobial infection from recognition to development of a nodule can be summarized in Figure 1.11.

Bacteria are deposited into the host cell cytoplasm in a process that resembles endocytosis (Gibson et al., 2008; Masson-Boivin et al., 2009). However, the internalized bacteria must first adapt to their new environment and establish a persistent nodule infection (Masson-Boivin et al., 2009). For the release and adaptation of rhizobia in the cytoplasm, polysaccharides and surface components of the cell wall such as (EPS) LPS (Lipopolysaccharides) CPS (Capsular polysaccharides), are required (Gibson et al., 2008; Perret et al., 2000). At this point the structural specificity of Nod Factors is less stringent (Goormachtig et al., 2004; Oldroyd & Downie, 2008).

25 CHAPTER 1

Figure 1.11. Schematic model of nodule development through root hair infection. Host flavonoids exuded into the soil trigger bacterial Nod Factor production, which elicits root hair curling and root hair invasion. Root hair invasion also requires bacteria EPS and host ROS production. Nod factors induce mitotic cell division in the root cortex leading to formation of the nodule meristem. Adapted from (Gibson et al., 2008).

Once bacteria colonize the cytoplasm they go through a differentiation process that results in the formation of a symbiosome, that corresponds to bacteria that have differentiated into bacteroids enclosed in a plant cell membrane (Gibson et al., 2008; Prell & Poole, 2006) (Figure 1.12). Symbiosomes are the site of nitrogen fixation (Perret et al., 2000).

26 CHAPTER 1

Figure 1.12. (a) Entry of bacteria through infection thread (IT) that elongates towards the nodule meristem. (b) Endocytosis of bacteria into the cytoplasm of host cells. Each bacterium is surrounded by a host-derived peribacteroid membrane (PBM) and differentiates into a bacteroid. Adapted from (Gibson et al., 2008).

The induction of nitrogenase gene expression is triggered by the microaerobic conditions inside the nodule, which creates an oxygen diffusion barrier in combination with high expression levels of plant leg-hemoglobin (Gibson et al., 2008). The host supports nitrogenase activity by providing bacteroids with constant flux of oxygen for aerobic respiration and by providing energy (Gibson et al., 2008). The metabolic product of nitrogenase activity is ammonia, which is (generally) secreted from the bacteroid and incorporated into the aminoacids glutamine and asparagine for assimilation by the host (Prell & Poole, 2006).

Two main types of nodules can be formed: indeterminate and determinate (Figure 1.13). In determinate nodulation, the nodule basal meristem undergoes limited cell division, therefore further nodule growth depends on cell expansion (Gibson et al., 2008; Sprent, 2009). In the indeterminate nodules there is a persistent meristem that is continuously generating new cells that are invaded by bacteria. Therefore each developmental stage required to establish symbiosis is present within a single mature indeterminate nodule (Gibson et al., 2008; Sprent, 2009).

27 CHAPTER 1

Figure 1.13. a) Indeterminate nodule with a persistent meristem (Zone I), an invasion zone (Zone II), a nitrogen fixing zone (Zone III) and a senescent zone (Zone IV). b) Determinate nodule (Determinate meristematic activity). Adapted from (Gibson et al., 2008).

1.9 Rhizobial inoculants Inoculation of legumes with rhizobia is one of the success stories of world agriculture (Herridge, 2008). In fact, most of the fixed nitrogen in Australia can be attributed to present and past inoculation of legume crops (Sprent, 2008). There is a need to inoculate when soil nitrogen is insufficient to meet the plant’s requirements or when the legume cannot establish effective symbiosis with the native and naturalised rhizobia. Normally, when a new legume is introduced into an environment with no record of a similar species, inoculation is needed (Herridge, 2008).

An inoculant strain should be selected according to its ability to fix nitrogen and increase plant biomass (Howieson et al., 2000a) and, ideally, with a wide range of host legume species (O'Hara et al., 2002). To ensure the inoculant effectiveness in the field, it should be genetically stable, able to adapt to the edaphic environment targeted for the host (Brockwell et al., 1982; Howieson et al., 2000b; O'Hara et al., 2002), able to compete with indigenous or naturalised rhizobia and be able to reach and attach to roots and initiate nodule formation (Denton et al., 2003; Thies et al., 1992). Particularly for perennial legumes, it is important that the rhizobial inoculant shows initial high

28 CHAPTER 1 nodule occupancy and is able to colonise the host rhizosphere and persist in the soil for long periods to ensure second year nodulation (Brockwell et al., 1982; Graham, 2008; Sessitsch et al., 2002).

1.10 Constraints to adaptation of introduced rhizobia Root nodule bacteria, just as their legume symbionts, have to cope with several environmental and edaphic stresses, which may affect their symbiotic performance in the field (Howieson & Ballard, 2004; Slattery et al., 2001). One of the primary stresses for legume symbiosis are extremes of soil pH (<5.0 or >8.5) (Howieson & Ballard, 2004), which can affect soil bacteria diversity and persistence (Fierer & Jackson, 2006) and reduce soil rhizobial populations (Brockwell et al., 1991). Low soil pH has proved detrimental for the establishment of the host-rhizobia symbiosis in numerous species (Ballard et al., 2003; Cocks, 2001; Richardson et al., 1988) mainly due to a disruption in the signal exchange between symbiotic partners (Hungria & Stacey, 1997; McKay & Djordjevic, 1993), and to alterations of rhizobial external motility mechanisms and EPS ornamentations, affecting the ability of rhizobia to reach the plant rhizosphere and establish symbiosis (Dilworth et al., 2001). Low soil pH is also associated with an increase in the availability of elements that are toxic for rhizobia such as aluminium, copper, zinc and manganese (Dilworth et al., 2001; Hungria & Vargas, 2000). Low clay content in the soil is also considered a primary stress for rhizobia, which makes low pH effects even more dramatic on cell structural integrity (Howieson & Ballard, 2004).

Secondary stress factors are those that are seasonally dependent (Howieson & Ballard, 2004), some examples of which are extreme temperatures, water stress and low phosphorous levels. High soil temperatures can reduce soil rhizobial populations and disrupt nodulation by altering the exchange of molecular signals between legume and rhizobia (Hungria & Vargas, 2000). Low water availability is considered a major cause of nodulation failure and reduced nitrogen fixation as it can affect rhizobial growth and survival in soils and the longevity of the nodules (Hungria & Vargas, 2000). Low soil

29 CHAPTER 1 phosphorous levels have a detrimental effect in production and excretion of Nod metabolites (McKay & Djordjevic, 1993).

Another important constraint for root nodule bacteria adaptation in a new environment is the presence of high populations of indigenous rhizobia, which may compete for nodulation (Denton et al., 2002; Dowling & Broughton, 1986; Thies et al., 1991; Thies et al., 1992). The chances of managing competition are scarce, it can be reduced by increasing application rates (Dowling & Broughton, 1986; Mårtensson, 1990) or by enhancing the inoculant competitive abilities through genetic engineering (Martínez-Romero & Rosenblueth, 1990; Oresnik et al., 1999; Sessitsch et al., 2002). An understanding of which phases of nodulation are exposed to competition is crucial to managing this phenomenon (Sessitsch et al., 2002; Yates et al., 2011). Howieson et al. (2008) suggest that it could be mostly avoided by introducing novel genera or species of rhizobia into environments.

Overall, the fundamental approach to overcome biotic and abiotic stresses is to select symbiotic partners that are more competent under stressful soil conditions (Graham, 2008; Howieson & Ballard, 2004). For example, to select strains that can overcome stress better than others (Hungria & Vargas, 2000) or by identifying a pool of effective rhizobia for the target legume and then select based on the impact of the environment in the symbiont performance (Sessitsch et al., 2002).

1.11 Approaches and Aims of the thesis The legumes L. capitata, L. diffusa, L. excisa, L. herbacea, L. incana and L. pauciflora have been introduced in Australia along with a collection of 73 root nodule bacteria strains isolated from diverse sites in South Africa. It is widely known that nitrogen fixation through the symbiosis with root nodule bacteria is one of the main purposes of the inclusion of legumes in Australian farming systems. Moreover, the adaptation of legumes to target environments will rely, largely, upon an optimal performance of the N-fixing symbiosis with an appropriate strain of root nodule bacteria (Howieson & Ballard, 2004;

30 CHAPTER 1

Howieson et al., 2008; Richardson et al., 2000). Therefore, the study of the diversity and symbiotic properties of these strains and the selection of the most effective ones is a key step in Lessertia spp. establishment in South Western Australia. The evaluation of their field performance is particularly important considering that many of the biotic and abiotic stresses that rhizobia can be subject to, are present in the target area (section 1.1). The specific aims of this thesis are,

I. To establish the identity and diversity of the rhizobial strains associated with the South African legumes Lessertia capitata, L. diffusa, L. excisa, L. herbacea, L. incana and L. pauciflora. II. To determine the specificity and nitrogen fixation effectiveness of the symbiotic associations of Lessertia spp. with different root nodule bacteria. III. To investigate the phylogenetic relationship of Lessertia spp. rhizobia nodulation genes and the possible exchange of transmissible genetic elements. IV. To assess Lessertia spp. rhizobial strains for adaptation, competitive ability and symbiotic performance in the target environments.

31 CHAPTER 1

32

CHAPTER 2. Authentication, characterisation and diversity of Lessertia spp. root nodule bacteria.

33 CHAPTER 2

34 CHAPTER 2

2.1 Introduction The South African species Lessertia capitata, L. diffusa, L. excisa, L. herbacea, L. incana and L. pauciflora are perennial legumes (Balkwill & Balkwill, 1999; Nkonki, 2003) that are part of natural pastures and may have forage potential (Breebaart, 2003). They have recently been introduced into Western Australia (Howieson et al., 2008) to increase the diversity of perennial legumes to overcome climate and agricultural changes (Cocks, 2001; Cransberg & McFarlane, 1994; Dear et al., 2003).

Successful growth of a legume is dependent upon rhizobial symbionts (Brockwell & Bottomley, 1995), therefore the study of root nodule bacteria is a key step in introducing the plant into a new environment. The root nodule bacteria associated with Lessertia spp. have not been studied in detail. To date, the only reports of Lessertia symbionts have been those of de Faria et al. (1989) and Allen & Allen (1981) who reported the presence of nodules in Lessertia spp. in situ and that of the ICMP database (International Collection of Micro-organisms from Plants, Landcare Research, Auckland, New Zealand) which includes Lessertia as one of the genera able to nodulate with one of the Mesorhizobium sp. strains listed (ICMP 12690). However, there is no evidence of a detailed study of the genetic and phenotypic diversity and phylogeny of the micro-symbiont of the Lessertia spp.

The isolates used in this study come from root nodules of Lessertia spp. collected from different geographic areas of the Northern, Eastern and Western Cape of South Africa (Table 2.1). Hence, the study of genetic diversity of root nodule bacteria isolated from Lessertia spp. and the investigation of the relationship between genetic diversity and geographical origin of these isolates may increase the likelihood of identifying a pool of different efficient rhizobial strains for Lessertia that would suit diverse environmental conditions.

The phylogenetic relationships and genetic diversity within different bacterial strains can be assessed through several molecular and phenotypic methods.

35 CHAPTER 2

Traditionally, phylogenetic relatedness in the order Rhizobiales has been estimated through 16s rRNA sequence comparison (Brenner et al., 2005; Jaspers & Overmann, 2004; Laguerre et al., 1997; McArthur, 2006; Olsen et al., 1994). Nevertheless, the use of alternative genes as phylogenetic markers, particularly protein encoding genes, is becoming a common approach (Chelo et al., 2007; Gaunt et al., 2001; Konstantinidis & Tiedje, 2005; Zeigler, 2003). The variable 3’ end fragment of the protein coding gene dnaK has a strong phylogenetic signal and is congruent with other housekeeping genes (Martens et al., 2007; Stępkowski et al., 2003a). Furthermore, the dnaK sequences provide a higher level of sequence divergence than 16S rRNA and have therefore proven to be useful to distinguish closely related bacteria (Chelo et al., 2007; Martens et al., 2007). Therefore dnaK was chosen in this study to estimate phylogenetic relationship among the root nodule isolates of Lessertia spp.

This chapter reports on authentication, strain diversity and the phylogenetic relationship among 73 isolates of root nodule bacteria from eight different species: L. pauciflora, L. incana, L. diffusa, L. capitata, L. herbacea L. frutescens, L. excisa and L. microphylla, isolated from 17 different sites (Table 2.1). The pH tolerance and colony growth rate were also assessed.

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2.2 Material and Methods 2.2.1 Bacterial culture conditions The 73 isolates from root nodules of Lessertia spp. (Table 2.1) were obtained from the Western Australian Soil Microbiology (WSM) culture collection. Bacterial cultures were grown on half lupin-agar (½ LA) plates [1.25 gL-1 Yeast extract; 26.7mM D-mannitol; 27.7mM D-glucose; 3.24mM MgSO4.7H2O;

1.71mM NaCl; 180µM FeSO4.7H2O; 100mM K2HPO4; 100mM KH2PO4;

11.6µM Na2B4O7; 9.10 µM MnSO4 4H2O; 760nM ZnSO4 7H2O; 320nM CuSO4

5H2O; 520nM Na2MoO4 2H2O; 0.15% agar (Howieson et al., 1988)] for 5 to 10 days at 28°C. The isolates were suspended in saline solution (0.89% w v-1 NaCl) and preserved in long-term storage at -80°C in 2.0 mL cryo-vials containing 30% (v v-1) glycerol.

The template for molecular studies was prepared using whole cells that were suspended in sterile saline solution (0.89% w v-1 NaCl). The suspension was centrifuged (5000 rpm/2 min); the supernatant was discarded and then the pellet was resuspended in fresh sterile saline solution. This was repeated three times and cell suspensions were standardized to an optical density (OD) of 6.0 at 600nm. Cell suspensions were stored at -20°C.

2.2.2 Molecular Fingerprinting The genetic diversity of the 73 isolates (Table 2.1) was assessed at the strain level by molecular fingerprinting using the primers RPO1 (5’ AATTTTCAAGCGTCGTGCCA 3’) (Richardson et al., 1995), EricF and EricR. (ERIC1R: 5’ ATGTAAGCTCCTGGGGATTCAC 3’; ERIC2F: 5’ AAGTAAGTGACTGGGGTGAGCG 3’) (Versalovic et al., 1991).

The PCR reaction mix for RPO1-PCR contained 1.0 µl of cell suspension, 0.5 µl of Taq DNA polymerase (Invitrogen Life Technologies), 1.0 µl of RPO1 primer (50nM) (Geneworks), 2.4 µl of 25mM MgCl2, 4.0 µl 5x Fisher-Biotech polymerization buffer [composition of 1x buffer: 67mM Tris-HCl (pH 8.8), -1 16.6mM (NH4)2SO4, 0.45% (v/v) Triton X-100, 0.2 mg mL gelatin and 0.2mM

37 CHAPTER 2 dNTP] (Fisher Biotech, Adelaide, Australia) and 11.1 µl of UltraPure grade water (Fisher Biotech) to total 20 µl (modified from Richardson et al. (1995)).

PCR was conducted on an iCycler (BIORAD) and the cycling conditions were a cell lysis step of 5 min at 95C, followed by 5 cycles at 94C for 30 s; 50C for 10 s and 72C for 1:30 min; and then 35 cycles at 94°C for 30 s, 55°C for 25 s and 72°C for 1:30 min and a final extension at 72°C for 5 min.

The ERIC PCR reaction mixture contained 2.0 µl of cell suspension, 0.5 µl of Taq polymerase (Invitrogen Life Technologies), 1.0 µl of forward and reverse primers 50µM (Geneworks), 5.0 µl of 5x Gitschier buffer (83.0 mM (NH4)2SO4 ;

0.335 M Tris-HCl (pH 8.8); 33.5 mM MgCl2 ;0.032 mM EDTA (pH 8.8); 0.15 M β-mercapto-ethanol), 0.4 µl of BSA (10 mg/ml), 2.5 µl of DMSO, 1.25 µl of dNTPs 100 µM and 11.35 µl of UltraPure grade water (Fisher Biotech) to total 20 µl. The reaction mixture was held at 95°C for 7 minutes, followed by 35 cycles at 94°C for 30s, 50°C for 1 min and 65°C for 8 min with a final hold at 65°C for 16 min (Modified from (Versalovic et al., 1991).

The reproducibility of the RPO1 and ERIC PCRs was checked by repeating the procedures at least twice per strain, and every time with fresh cultures from the original glycerol stocks.

The PCR products were electrophoresed in 2% (w/v) agarose gels pre-stained with 1:10,000 SYBR ® Safe DNA gel stain (Invitrogen). The marker used was 1Kb DNA ladder (Promega G5711). Electrophoresis was carried out in tanks buffered with 1xTAE (40mM Tris-Acetate, 1mM EDTA, pH 8.0) at 100V for 3 hours. Bands were visualized in a UV transilluminator.

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Table 2.1. Origin of root nodule bacteria used in this study

Strain Host plant Geographic origin Lat Long Source (year isolated) WSM2211 Lessertia frutescens Graaf Reinet, Eastern cape Yates and Real (2002) WSM2559 Lessertia microphylla Klawer, Western Cape 31º41'512''S 18º41'640''E Yates and Real (2002) WSM2561 Lessertia diffusa Nieuwoudtville, Northern Cape 31º25'684''S 19º10'616''E Yates and Real (2002) WSM2562 Lessertia diffusa Nieuwoudtville, Northern Cape 31º25'684''S 19º10'616''E Yates and Real (2002) WSM2566 Lessertia pauciflora Rooiberg pass, Western Cape 32º29'997''S 21º37'158''E Yates and Real (2002) WSM2607 Lessertia microphylla Beaufort West, Western Cape 32º10'394''S 22º32'598''E Yates and Real (2002) WSM2622 Lessertia microphylla Beaufort West, Western Cape 32º10'394''S 22º32'598''E Yates and Real (2002) WSM2623 Lessertia annularis Beaufort West, Western Cape 32º21'109''S 22º36'127'' Yates and Real (2002) WSM2628 Lessertia diffusa Nieuwoudtville, Northern Cape 31º25'684''S 19º10'616''E Yates and Real (2002) WSM2801 Lessertia frutescens Nama Karoo, Eastern Cape Yates and Real (2002) WSM2804 Lessertia frutescens Nama Karoo, Eastern Cape Yates and Real (2002) WSM2805 Lessertia frutescens Nama Karoo, Eastern Cape Yates and Real (2002) WSM2808 Lessertia frutescens Nama Karoo, Eastern Cape Yates and Real (2002) WSM2809 Lessertia frutescens Nama Karoo, Eastern Cape Yates and Real (2002) WSM2814 Lessertia microphylla Beaufort West, Western Cape 32º10'394''S 22º32'598''E Yates and Real (2002) WSM2991 Lessertia microphylla Calvinia, Northern Cape 31º28'172''S 19º49'088''E Law (2003) WSM2995 Lessertia inflata Beaufort West, Western Cape 32º14'082''S 22º55'081''E Law (2003) WSM3059 Lessertia annularis Beaufort West, Western Cape 32º21'109''S 22º36'127'' Law (2003) WSM3060 Lessertia annularis Beaufort West, Western Cape 32º21'109''S 22º36'127'' Law (2003) WSM3061 Lessertia annularis Beaufort West, Western Cape 32º21'109''S 22º36'127'' Law (2003) WSM3062 Lessertia annularis Beaufort West, Western Cape 32º21'109''S 22º36'127'' Law (2003) WSM3265 Lessertia microphylla Middelburg, Eastern Cape Law (2003) WSM3270 Lessertia annularis Middelburg, Eastern Cape Law (2004) WSM3273 Lessertia annularis Beaufort West, Western Cape Law (2004) WSM3274 Lessertia annularis Beaufort West, Western Cape Law (2004) WSM3275 Lessertia annularis Beaufort West, Western Cape Law (2004) WSM3310 Lessertia frutescens Stellenbosch, Western Cape Law (2003) WSM3311 Lessertia frutescens Stellenbosch, Western Cape Law (2003) WSM3326 Lessertia microphylla Stellenbosch, Western Cape Law (2003) WSM3472 Lessertia excisa Lamberts Bay, Western Cape 32º03' 562''S 18º18' 965''E Howieson et al . (2004) WSM3492 Lessertia diffusa Garies, Northern Cape 30º32'419'' S 17º57'031''E Howieson et al . (2004) WSM3493 Lessertia diffusa Springbok, Northern Cape 29º39'473''S 17º54'309''E Howieson et al . (2004) WSM3495 Lessertia capitata Komkans, Western Cape 31º13'359''S 18º03' 07''E Howieson et al . (2004) WSM3501 Lessertia diffusa Springbok, Northern Cape 29º39'473''S 17º54'309''E Howieson et al . (2004) WSM3502 Lessertia diffusa Springbok, Northern Cape 29º39'473''S 17º54'309''E Howieson et al . (2004) WSM3503 Lessertia diffusa Springbok, Northern Cape 29º34'506''S 18º01'611'' Howieson et al . (2004) WSM3507 Lessertia capitata Komkans, Western Cape 31º13'359''S 18º03' 07''E Howieson et al . (2004) WSM3513 Lessertia diffusa Springbok, Northern Cape 29º39'473''S 17º54'309''E Howieson et al . (2004) WSM3564 Lessertia diffusa Grotto Bay, Western Cape 33º29'21''S 18º19'36''E Howieson et al . (2004) WSM3565 Lessertia diffusa Grotto Bay, Western Cape 33º29'21''S 18º19'36''E Howieson et al . (2004) WSM3571 Lessertia tomentosa Geelbek, Western Cape 33º13'85''S 18º12'23''E Howieson et al . (2004) WSM3592 Lessertia herbacea Citrusdal, Western Cape 32º43'92''S 19º02'42''E Howieson et al . (2004) WSM3595 Lessertia herbacea Citrusdal, Western Cape 32º43'92''S 19º02'42''E Howieson et al . (2004) WSM3596 Lessertia herbacea Citrusdal, Western Cape 32º43'92''S 19º02'42''E Howieson et al . (2004) WSM3597 Lessertia herbacea Citrusdal, Western Cape 32º43'92''S 19º02'42''E Howieson et al . (2004) WSM3598 Lessertia herbacea Citrusdal, Western Cape 32º43'92''S 19º02'42''E Howieson et al . (2004) WSM3602 Lessertia herbacea Citrusdal, Western Cape 32º38'99''S 19º11'937''E Howieson et al . (2004) WSM3605 Lessertia herbacea Marcuskraal, Western Cape 32º25'43''S 18º57' 60''E Howieson et al . (2004) WSM3606 Lessertia herbacea Marcuskraal, Western Cape 32º25'43''S 18º57' 60''E Howieson et al . (2004) WSM3612 Lessertia excisa Lamberts Bay, Western Cape 32º03' 562''S 18º18' 965''E Howieson et al . (2004) WSM3620 Lessertia diffusa Garies, Northern Cape 30º32'419'' S 17º57'031''E Howieson et al . (2004) WSM3626 Lessertia diffusa Kamieskroon, Northern Cape 30º11'261''S 17º59'350''E Howieson et al . (2004) WSM3628 Lessertia diffusa Kamieskroon, Northern Cape 30º11'261''S 17º59'350''E Howieson et al . (2004) WSM3635 Lessertia capitata Komkans, Western Cape 31º13'359''S 18º03' 07''E Howieson et al . (2004) WSM3636 Lessertia capitata Komkans, Western Cape 31º13'359''S 18º03' 07''E Howieson et al . (2004) WSM3803 Lessertia herbacea Citrusdal, Western Cape 32º43'92''S 19º02'42''E Law (2004) WSM3804 Lessertia herbacea Citrusdal, Western Cape 32º43'92''S 19º02'42''E Law (2004) WSM3805 Lessertia herbacea Citrusdal, Western Cape 32º43'92''S 19º02'42''E Law (2004) WSM3806 Lessertia herbacea Citrusdal, Western Cape 32º43'92''S 19º02'42''E Law (2004) WSM3807 Lessertia herbacea Citrusdal, Western Cape 32º43'92''S 19º02'42''E Law (2004) WSM3822 Lessertia rigida Springbok, Northern Cape 29º34'506''S 18º01'611'' Howieson et al . (2004) WSM3891 Lessertia excisa Lamberts Bay, Western Cape 32º03' 562''S 18º18' 965''E Howieson et al . (2004) WSM3892 Lessertia excisa Lamberts Bay, Western Cape 32º03' 562''S 18º18' 965''E Howieson et al . (2004) WSM3893 Lessertia excisa Lamberts Bay, Western Cape 32º03' 562''S 18º18' 965''E Howieson et al . (2004) WSM3894 Lessertia excisa Lamberts Bay, Western Cape 32º03' 562''S 18º18' 965''E Howieson et al . (2004) WSM3895 Lessertia excisa Lamberts Bay, Western Cape 32º03' 562''S 18º18' 965''E Howieson et al . (2004) WSM3897 Lessertia excisa Lamberts Bay, Western Cape 32º03' 562''S 18º18' 965''E Howieson et al . (2004) WSM3898 Lessertia excisa Lamberts Bay, Western Cape 32º03' 562''S 18º18' 965''E Howieson et al . (2004) WSM3900 Lessertia excisa Lamberts Bay, Western Cape 32º03' 562''S 18º18' 965''E Howieson et al . (2004) WSM3917 Lessertia diffusa Springbok,Northern Cape 29º34'506''S 18º01'611'' Howieson et al . (2004) WSM3918 Lessertia diffusa Springbok,Northern Cape 29º34'506''S 18º01'611'' Howieson et al . (2004) WSM3919 Lessertia incana Springbok,Northern Cape 29º34'506''S 18º01'611'' Howieson et al . (2004) WSM3920 Lessertia incana Springbok,Northern Cape 29º34'506''S 18º01'611'' Howieson et al . (2004)

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The banding patterns were analysed using the software BIOCapt (ALYS Technologies, Lausanne) to scan for presence of PCR product of specific molecular sizes. A binary matrix was constructed with the scores 1 or 0 for the presence or absence of a band at each molecular size. The matrix was then analysed with AFPL SURV version 1.0 (Vekemans, 2002) to calculate the genetic distance among isolates. The distance matrix was then subjected to UPGMA cluster analyses using the NEIGHBOR application from the PHYLIP software package. The cladograms were visualized in MEGA4 (Tamura et al., 2007).

2.2.3 Authentication of isolates as root nodule bacteria A glasshouse experiment using a closed vial system was set up to confirm the ability of the 61 distinct strains (Appendix 7) to nodulate Lessertia spp.

Seeds of: Lessertia capitata, L. diffusa, L. excisa, L. frutescens, L. herbacea, L. incana, L. microphylla, L. pauciflora and Macroptilium atropurpureum (Siratro), were scarified with sand paper (P240) and surface sterilised by immersion in 70% (v/v) ethanol for 1 minute, followed by one soak in 3% (v/v) sodium hypochlorite for 3 minutes and six washes in sterile DI water. Sterilised seeds were placed on 1.5% (w/v) water agar plates and incubated for 48 hours at room temperature in the dark. Once the radicle had emerged the seeds were transferred to the vials.

Seedlings were grown in 250ml plastic vials, containing 125ml of the substrate that consisted of 2 parts washed river sand and 3 parts washed yellow sand, which were previously autoclaved at 121°C for 30 minutes. Each vial contained one Lessertia plant that corresponded to the original host of each of the strains and one of the highly promiscuous legume M. atropurpureum. For those strains whose original hosts were L. inflata, L. rigida and L. tomentosa the vials contained seedlings of L. diffusa, L. herbacea and M. atropurpureum, as seeds of the original host species were not available. There were three replicates per strain. After planting the seedlings, 10 ml of sterile nutrient solution devoid of nitrogen was added to each vial. The nutrient solution -1 -1 -1 - contained 0.31 gL MgSO4.7H2O; 0.21 gL KH2PO4; 0.44 gL K2SO4; 0.06 gL

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1 -1 -1 -1 FeEDTA; 0.05 gL CaSO4; 0.116 mgL H3BO3 0.116; 0.0045 mgL -1 -1 Na2MoO4.2H2O; 0.134 mgL ZnSO4.7H2O; 0.01 mgL MnSO4,4H2O and 0.06 -1 mgL CoSO4.7H2O (Howieson, 1995).

The bacterial strains were recovered from the glycerol stocks and grown on ½ LA plates. At planting, each seedling was inoculated with 1 ml of a cell suspension aseptically washed from fresh colonies and suspended in 1%

(w/v) sucrose solution to an OD600nm of 7.0. The plants were grown in a temperature controlled glasshouse at 22°C, under axenic conditions for 8 weeks.

At harvest, plants were carefully removed from the vials, washed in tap water and nodulation was recorded. Where nodulation occurred two nodules were sampled from each plant. These were disinfected using the same protocol as for the seed disinfection but reducing the time in hypochlorite to 90 seconds and in 70% (v/v) ethanol to 30 seconds. Surface sterile nodules were crushed in a sterile petri dish and the nodule contents streaked on ½ LA agar plates. After incubation at 28°C for 3 to 9 days, depending on the rate of growth of the strain, the bacteria were re-streaked from individual colonies onto fresh ½ LA plates. The re-isolated bacteria were fingerprinted with the primer RPO1 to confirm their identity.

2.2.4 pH range The ability of 43 authenticated strains to grow at different pH was assessed. Starter cultures of the strains were streaked from glycerol stocks onto ½ LA plates at pH 7. The media ½ LA was prepared and adjusted to 8 different pH (4.0, 4.5, 5.0, 5.5, 6.0, 7.0, 8.0 and 9.0). To maintain each of the pH, the following buffers were added into the media:10 mM Homopipes for pH 4.0, .4.5 and 5.0; 0.5M Hepes for 7.0 and 8.0; and Ches for pH 9. For pH 4.0, 4.5 and 5.0 media, agarose was used instead of agar, and a concentrated agarose mixture [6% (w v-1)] was autoclaved separately from the buffered nutrient solution. Each of the strains was inoculated from the starter culture into each pH media with a sterile inoculating stick. Colony growth was assessed during 10 days.

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2.2.5 Sequencing the dnaK gene To investigate phylogenetic relationships among the 43 authenticated Lessertia rhizobial strains (Appendix 7) and to evaluate the RNB diversity at species level, the variable 3’ end fragment of the protein encoding gene dnaK (330 bp) was amplified using the primers TSdnak2 and TSdnak3 (Stępkowski et al., 2003a) (Table 2.2).

Cell templates were prepared as described in 2.2.1 but this time the cells were concentrated to an OD600 of 2.0. The PCR reaction mixture comprised 2 µl of cell suspension, 0.5 µl of Taq polymerase (invitrogen Life Technologies), 1 µl of forward and reverse primers (Geneworks), 6.0 µl of 25mM MgCl2, 20.0 µl 5x Fisher-Biotech polymerization buffer and 69.5 µl of UltraPure grade water (Fisher Biotech) to total 100 µl for a single reaction. The cycling parameters were: 7 minutes at 95°C followed by 35 cycles of 94°C for 30 s, 62°C for 30 s and 72°C for 45s and a final hold at 72°C for 5 min.

The amplification of the 330bp dnaK sequence was verified by agarose gel electrophoresis as described above (2.2.2). The PCR products were column- purified using the Qia-quick purification kit (QIAGEN) as recommended by the manufacturer, with a final elution in 50 µl of buffer EB (10mM Tris-Cl, pH8.5).

In a 96 well plate, purified PCR products were sequenced in both directions with the primers TSdnak2 and TSdnak3. For each 4 µl of purified PCR product, 4 µl BIG-dye terminator (version 3.1), 1 µl of a primer and 1 µl of Ultrapure PCR water were added. The cycling conditions were: 2 min at 96°C and 25 cycles of 96°C for 10 s, 50°C for 5 s and 60°C for 4 min.

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Table 2.2. Sequences of primers for housekeeping genes used in this work. Name Target Gene Sequence (5’ 3’) SOURCE 20F 16S rRNA AGTTTGATCCTGGCTCA* Yanagi & Yamasato, 1993

420F 16S rRNA GATGAAGGCCTTAGGGTTGT Yanagi & Yamasato, 1993

800F 16S rRNA GTAGTCCACGCCGTAAACGA Yanagi & Yamasato, 1993

1540R 16S rRNA AAGGAGGTGATCCAGCCGCA Yanagi & Yamasato, 1993

1190R 16S rRNA GACGTCATCCCCACCTTCCT Yanagi & Yamasato, 1993

820R 16S rRNA CATCGTTTACGGCGTGGACT Yanagi & Yamasato, 1993

520R 16S rRNA GCGGCTGCTGGCACGAAGTT Yanagi & Yamasato, 1993

TSdnak2 dnaK GTACATGGCCTCGCCGAGCTTCA Stępkowski et al., 2003a

TSdnak3 dnak AAGGAGCAGCAGATCCGCATCCA Stępkowski et al., 2003a

*Symbols: A, C, G, T – standard nucleotides

The Sequencing PCR products were ethanol/EDTA/Sodium acetate precipitated in a 96 well plate following the protocol recommended by Applied Biosystems 3730 DNA Analyzers sequencing chemistry guide. The DNA sequence chromatograms were obtained using an automated ABI Prism ® 377 DNA sequencer (Applied Biosystems).

2.2.6 Sequencing the 16s rRNA gene The 16s rRNA gene was sequenced for a subset of 17 strains (Appendix 7) chosen from each of the clusters of the phylogenetic tree developed with dnaK and including strains isolated from every Lessertia sp.. The procedure was similar to the one described for the amplification of dnaK partial sequences (2.2.5), with the exception that the concentration of cells in the template was adjusted to an OD600 of 10. The primers 20F and 1540R (Table 2.2) were initially used to amplify a 1500 bp internal region of the 16S rRNA and this was followed by sequencing with the other 16S rRNA primers listed in table 2.2.

The cycling parameters were: 5 minutes at 95°C followed by 30 cycles of 94°C for 30 s, 55°C for 30 s and 30°C for 45s and a final hold at 72°C for 7 min. The annealing temperature had to be adjusted for some strains to increase or reduce stringency when needed. For the strains WSM2559 and

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WSM3270 the temperature was reduced to 54.5°C; for the strain WSM3602 it was reduced to 53.5°C; for the strains WSM2622 and WSM2623 it was increased to 56.5°C; and for the strains WSM3598 and WSM3626 to 58°C.

2.2.7 Phylogenetic analysis The chromatograms for partial dnaK and 16s rRNA were analysed and edited in the Gene Tool Lite 1.0 (2000) software (Doubletwist, Inc., Oakland, CA, USA). Sequences alignments and phylogenetic analyses were conducted in MEGA4 (Tamura et al., 2007). The alignments were made by ClustalX and were edited manually. The phylogenetic tree was inferred by the Neighbor- Joining method (Saitou & Nei, 1987) with a bootstrap analysis based on 1,000 replicates to evaluate the confidence of the nodes. The genetic distances were computed using the Maximum Composite Likelihood method (Tamura et al., 2004).

The BLAST search tool from the National Centre for Biotechnological Information (NCBI) was used to find species closely related to Lessertia spp. strains and to download type strain’s genetic sequences to be included in the phylogenetic trees.

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2.3 Results 2.3.1 Molecular Fingerprinting There was a high diversity of strains even for isolates coming from the same host and from the same geographical origin. Appendices 8 and 9 show the complete cladograms obtained with RPO1-PCR and ERIC-PCR respectively.

In general, isolates that were clustered together by RPO1 also clustered together, with minor differences, in the ERIC PCR cladogram. Most of the strains that had identical RPO1-PCR fingerprinting patterns also had identical or very similar fingerprinting patterns when using ERIC primers. However isolate WSM2607 from L. microphylla and isolates WSM3897, 3898, 3895 and 3900 from L. excisa had different fingerprints with ERIC primers although they all had identical fingerprinting patterns with RPO1. On the other hand, isolates WSM2801, 3595, 3472, 3493 and 3516 shared an identical fingerprint through ERIC-PCR but their RPO1 fingerprinting patterns were different.

Identical fingerprints were only detected for isolates coming from the same host and same geographical sites. In spite of this, there were strains that were originally isolated from the same host and same site but their fingerprints were distantly related. This is evident in the cases of L. herbacea that was nodulating with 7 distinct strains (WSM3592, 3596, 3602, 3595, 3597, 3598, 3803 and 3804) (Figure 2.1) all isolated from a site in Citrusdal (Western Cape) and L. excisa that had 7 different symbionts (WSM3472, 3612, 3900, 3894, 3893, 3891 and 3898) (Figure 2.2) isolated from one single site (Lamberts Bay, Western Cape).

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Figure 2.1. Cladogram showing the relationships among Lessertia herbacea root nodule bacteria according to fingerprints obtained with RPO1-PCR.

Figure 2.2. Cladogram showing the relationships among Lessertia excisa root nodule bacteria according to fingerprints obtained with RPO1-PCR.

From the 73 original isolates (Table 2.1), 12 strains (WSM2562, 2805, 2809, 2814, 3274, 3605, 3628, 3635, 3807, 3892, 3895 and 3897) were excluded from future experiments as they revealed identical fingerprints to other isolates with RPO1 and ERIC PCR. The 61 isolates that had unique banding patterns with at least one of the sets of primers were considered as individual strains and were selected for further studies.

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2.3.2 Rhizobia authentication From the 61 strains used in this study, 43 were able to nodulate Lessertia spp. forming pink, either fan-shaped or bifurcate indeterminate nodules in most of the cases (Figure 2.3). Only 22 strains were able to nodulate M. atropurpureum forming small white nodules (Table 2.3). Of the 9 strains from L. annularis only WSM2623, 3059 and 3270 were able to nodulate L.diffusa and L. herbacea. None of the L. tomentosa, L. inflata and L. rigida strains was able to nodulate L.diffusa, L. herbacea or M. atropurpureum. Numbers of nodules per plant were in general low, ranging from 2 to 7 nodules per root system; the only exceptions were the strains WSM3602 and 3620 that formed more than 30 round white nodules on their original hosts: L .herbacea and L. diffusa.

Figure 2.3. Nodules on Lessertia capitata (a), L. herbacea (b), L. excisa (c) and L. diffusa (d) inoculated with strains WSM3636, WSM3592, WSM3900 and WSM3626 respectively. Bars: 500µm.

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When re-isolated, the majority of the strains produced white mucoid colonies, whereas WSM3602 and WSM3270 produced dry colonies. There were variations in the colony growth rate within strains at 28ºC on ½LA and TY agar. Twelve strains were able to form 1mm colonies within 2-4 days and were classified as fast-growing strains; another group of 14 strains took 5-7 days and were thus categorized as medium-slow. Seventeen isolates were considered slow-growing strains, as there was visible growth only after 8-10 days (Table 2.3).

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Table 2.3. Colony growth rate of Lessertia spp. strains and ability to nodulate their original host and Macroptilium atropurpureum. Nodulation Colony Strain Host plant Lessertia sp. M. atropurpureum Growth rate WSM2211 Lessertia frutescens + (P) - Slow WSM2559 Lessertia microphylla + (P) - Slow WSM2561 Lessertia diffusa + (P) - Slow WSM2566 Lessertia pauciflora + (P) - Slow WSM2607 Lessertia microphylla + (P) - Slow WSM2622 Lessertia microphylla + (P) + (W) Slow WSM2623 Lessertia annularis + (P) - Slow WSM2628 Lessertia diffusa + (P) + (W) Slow WSM2808 Lessertia frutescens + (P) + (W) Fast WSM3059 Lessertia annularis + (P) + (W) Fast WSM3270 Lessertia annularis + (P) + (W) Medium-slow WSM3310 Lessertia frutescens + (P) - Slow WSM3472 Lessertia excisa + (P) - Medium-slow WSM3492 Lessertia diffusa + (P) - Slow WSM3493 Lessertia diffusa + (P) - Slow WSM3495 Lessertia capitata + (P) + (W) Medium-slow WSM3502 Lessertia diffusa + (P) - Slow WSM3503 Lessertia diffusa + (P) + (W) Slow WSM3507 Lessertia capitata + (P) + (W) Medium-slow WSM3513 Lessertia diffusa + (P) - Slow WSM3564 Lessertia diffusa + (P) + (W) Fast WSM3565 Lessertia diffusa + (P) + (W) Medium-slow WSM3592 Lessertia herbacea + (P) - Fast WSM3596 Lessertia herbacea + (P) - Fast WSM3598 Lessertia herbacea + (P) - Medium-slow WSM3602 Lessertia herbacea + (W) + (W) Fast WSM3606 Lessertia herbacea + (P) - Fast WSM3612 Lessertia excisa + (P) + (W) Medium-slow WSM3620 Lessertia diffusa + (W) - Medium-slow WSM3626 Lessertia diffusa + (P) + (W) Medium-slow WSM3636 Lessertia capitata + (P) + (W) Medium-slow WSM3803 Lessertia herbacea + (P) - Fast WSM3804 Lessertia herbacea + (P) - Medium-slow WSM3805 Lessertia herbacea + (P) + (W) Medium-slow WSM3806 Lessertia herbacea + (P) - Medium-slow WSM3891 Lessertia excisa + (W) + (W) Fast WSM3893 Lessertia excisa + (P) + (W) Fast WSM3894 Lessertia excisa + (P) + (W) Fast WSM3898 Lessertia excisa + (P) + (W) Medium-slow WSM3900 Lessertia excisa + (P) + (W) Fast WSM3917 Lessertia diffusa + (P) - Slow WSM3919 Lessertia incana + (P) - Slow WSM3920 Lessertia incana + (P) - Slow

+: all plants nodulated, -: no nodulation, P: pink inside nodules, W: white inside nodules

2.3.3 pH range The strains showed differences in low pH tolerance. All of them were able to grow between pH 6.0 to 9.0, but only three strains (WSM2808, 3592 and

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3602) were able to grow from pH 4.0 (Table 2.4) and 13 strains (WSM3495, 3564, 3565, 3596, 3598, 3612, 3626, 3891, 3893, 3894, 3898 and 3900) from 4.5. Strains WSM3270, 3472, 3507, 3606, 3803, 3804, 3805 and 3806 grew from pH 5.0 to 9.0. The strains WSM2211, 2607, 2622, 2623, 2628, 3059, 3310, 3492, 3493 and 3513 formed colonies at pH 5.5 and higher. The strains WSM2559, 2561, 2566, 3502, 3503, 3620, 3917, 3919 and 3920 were low pH intolerant and could only form colonies from pH 6.0 to 9.0 (Table 2.4).

Table 2.4. Growth of Mesorhizobium strains in media at different pH pH

WSM 4 4.5 5 5.5 6 7 8 9

2211 - - - + + + + +

2559 - - - - + + + +

2561 - - - - + + + +

2566 - - - - + + + +

2607 - - - + + + + +

2622 - - - + + + + +

2623 - - - + + + + +

2628 - - - + + + + +

2808 + + + + + + + +

3059 - - - + + + + +

3270 - - + + + + + +

3310 - - - + + + + +

3472 - - + + + + + +

3492 - - - + + + + +

3493 - - - + + + + +

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Table 2.4. Continued

3495 - + + + + + + +

3502 - - - - + + + +

3503 - - - - + + + +

3507 - - + + + + + +

3513 - - - + + + + +

3564 - + + + + + + +

3565 - + + + + + + +

3592 + + + + + + + +

3596 - + + + + + + +

3598 - + + + + + + +

3602 + + + + + + + +

3606 - - + + + + + +

3612 - + + + + + + +

3620 - - - - + + + +

3626 - + + + + + + +

3636 - - + + + + + +

3803 - - + + + + + +

3804 - - + + + + + +

3805 - - + + + + + +

3806 - - + + + + + +

3891 - + + + + + + +

3893 - + + + + + + +

3894 - + + + + + + +

3898 - + + + + + + +

3900 - + + + + + + +

3917 - - - - + + + +

3919 - - - - + + + +

3920 - - - - + + + +

+: colony growth, -: no growth observed

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2.3.4 Phylogenetic analysis dnaK A single DNA fragment of 318bp representing the variable 3’ end fragment of the dnaK gene was amplified in 42 of the 43 strains. It was not possible to amplify the region in the strain WSM3602 isolated from L. herbacea although several attempts were made to optimise the PCR protocol by testing a range of lower annealing temperatures from 55°C to 60°C, and by increasing the

MgCl2 concentrations from 1.5mM to 2.5mM.

All the 42 strains formed a well supported cluster (88% bootstrap support) together with the Mesorhizobium type strains (Figure 2.4). The sequences were distributed in 10 clades (A to J) within the Mesorhizobium genus. Clades A, D and H were well defined with a bootstrap support higher that 85%. All the other clades showed a bootstrap support lower than 70%.

Clade A in the tree (Figure 2.4) comprised 12 strains that clustered with the strain WSM3872 isolated from a Biserrula sp. from Eritrea, Africa. This clade included strains isolated from L. excisa (WSM3472, 3900, 3898, 3893, 3891 and 3894), L. herbacea (WSM3596 and 3592), L. capitata (WSM3495 and 3507), WSM3059 from L. annularis and the strain WSM2808 isolated from L. frutescens.

Each of the clades B, C, F and G comprised a sole strain [WSM3805 (L. herbacea), 3612 (L. excisa), 3804 (L. herbacea) and 3270 (L. annularis) respectively] that was sufficiently well separated from other Lessertia strains and from any published Mesorhizobium spp.

Clade D included three L. herbacea strains (WSM 3606, 3806 and 3803). Clade E comprised three strains from L. diffusa (WSM3564, 3565 and 3626), one strain isolated from L. capitata (WSM3636) and the strains WSM 3598, isolated from L. herbacea. Clades D and E were closely grouped and not related to any known Mesorhizobium species.

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Figure 2.4. Neighbor joining phylogenetic tree based on partial dnaK gene sequencing of 43 strains of Lessertia spp. root nodule bacteria. M.: Mesorhizobium; S.: Sinorhizobium; R.: Rhizobium and B.: Bradyrhizobium. * Initials in parentheses after the strain number indicate original host: L. annularis (La); L. capitata (Lc); L. diffusa (Ld); L.excisa (Le); L.frutescens (Lf); L.herbacea (Lh); Li: L.incana; L.microphylla (Lm); L.pauciflora(Lp). The type strains sequences in the phylogram were obtained from GenBank (accession number).

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Clade H included two strains from L. diffusa (WSM2561 and 2628) that showed identical dnaK sequences. This clade was closely related to clade I and J, but as it had more than 6 nucleotide mismatches with the strains included in those clades and had a high bootstrap support (85%) it was considered a sole cluster.

Clade I included two strains isolated from L. microphylla (WSM2607 and 2622) that showed identical dnaK sequences, and one strain from L. frutescens (WSM2211) that had only 2 mismatches with the other two. These three strains were intermingled with the type strains M. mediterraneum, M. tianshanense and M. temperatum.

Clade J comprised 13 strains. Most of the strains in this group had been isolated from L. diffusa (WSM3492, 3502, 3503, 3513, 3620, 3493 and 3917), two strains from L. incana (WSM 3919 and 3920), WSM3310 from L. frutescens, WSM2559 from L. microphylla, WSM2623 from L. annularis and WSM2566 from L. pauciflora.

The diversity of sequences varied depending on the plant host from which the strains had been isolated. There were strains isolated from a sole plant species that tended to group in one or two clusters. For example, the strains from L. capitata were distributed in groups A and E; and six of the strains from L. excisa were clustered in clade A. In contrast, 9 strains isolated from L. herbacea were distributed in five different clades: A, B, D, E and F; and the 12 strains from L. diffusa were distributed in clades E, H and J.

Representatives of groups of strains with identical or nearly identical (2bp mismatches) dnaK sequences were selected to be compared with the type strains of other Mesorhizobium species. The strains included in table 2.5 that are representing other strains are the following: WSM2559 (representing WSM2623, 3492, 3493, 3502, 3503, 3513, 3620 and 2566); WSM2561 (WSM2628); WSM2622 (WSM2607 and 2211); WSM3495 (WSM2808, 3059, 3472, 3507, 3592, 3596, 3891, 3893, 3894, 3898 and 3900); WSM3606 (WSM3806), WSM3626 (WSM3565 and 3598) and WSM3919 (WSM3917

54 CHAPTER 2 and WS3920). The strains WSM3270, 3564, 3612, 3636, 3803, 3804 and 3805 had unique dnaK sequences.

Strains WSM2561, WSM2622 and WSM3310 had more than 98% sequence similarity to Mesorhizobium temperatum (less than 5 nucleotide mismatches) and were also very similar to M. mediterraneum with less than 10 nucleotide mismatches. The dnaK sequences of WSM2559 and WSM3919 were also similar to M. temperatum and M. mediterraneum showing a sequence similarity higher than 96.8% and less than 10 nucleotide mismatches.

All the other strains (WSM3495, WSM3564, WSM3606, WSM3626, WSM3636, WSM3803, WSM3804 and WSM3805) showed less than 94% sequence similarity and more than 16 nucleotide mismatches with any of the Mesorhizobium type strains.

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Table 2.5. Partial dnaK gene sequence similarity of Lessertia symbionts against some Mesorhizobium type strains.

WSM 2559* 2561* 2622* 3270 3310* 3495 3564 3606

M. albiziae 88.9 88.6 88.2 85.7 88.6 86.8 85.7 83.6 T CCBAU 61158 (37) (38) (39) (50) (38) (45) (49) (53)

M. amorphae 91.1 91.8 91.8 90.9 92.1 89.3 88.5 87.1 T LMG 18977 (26) (24) (25) (24) (22) (32) (35) (38)

M. australicum 85.4 85.7 86.1 86.8 86.4 87.1 93.2 92.9 T WSM2073 (42) (40) (39) (39) (38) (38) (20) (21)

M. chacoense 86.1 87.1 87.1 87.2 87.5 87.1 85.0 85.0 T LMG19008 (41) (38) (38) (39) (37) (39) (47) (44)

M. ciceri bv. biserrulae 90.9 91.3 92.1 83.9 92.5 89.4 89.0 89.8 T UPM-Ca7 (26) (24) (21) (32) (21) (32) (31) (29)

M. huakuii 89.3 90.4 91.1 89.1 91.1 88.2 91.8 92.5 T IAM14158 (31) (27) (25) (33) (25) (35) (25) (22)

M. loti 88.9 89.6 90.0 88.7 90.7 87.5 89.6 90.0 T LMG6125 (33) (30) (28) (34) (27) (38) (32) (30)

M. mediterraneum 96.8 97.9 96.8 91.7 98.2 87.1 87.8 86.8 T UPM-Ca36 (9) (6) (9) (24) (5) (36) (36) (38)

M. metallidurans 93.2 93.9 92.9 91.3 94.6 88.6 88.2 88.2 T LMG24485 (19) (17) (20) (24) (15) (33) (36) (35)

M. opportunistum 88.2 89.3 88.6 88.3 89.3 87.9 91.0 91.1 T WSM2075 (34) (30) (32) (36) (30) (35) (27) (26)

M. plurifarium LMG 84.6 85.0 84.6 83.8 85.7 90.4 86.4 86.1 T 11892 (44) (43) (44) (43) (41) (29) (40) (40)

M. septentrionale 91.8 92.5 92.5 91.7 92.9 90.0 89.3 87.9 T SDW014 (24) (22) (23) (22) (20) (30) (33) (36)

M. temperatum 97.1 98.2 98.2 93.2 98.6 87.9 88.2 87.1 T SDW018 (8) (5) (5) (21) (4) (34) (35) (37)

M. thiogangeticum 82.9 83.9 83.6 82.3 83.9 82.5 81.0 81.1 T SJT (48) (45) (47) (48) (45) (49) (55) (54)

M. tianshanense 95.0 95.4 94.6 90.9 96.4 88.9 87.1 86.8 T USDA3592 (14) (13) (15) (26) (10) (32) (39) (39) *Strains representing a clade or groups of strains: WSM2559 (Clades L and J); 2561 (I); 2622 (K); 3495 (E); 3606 (3806); 3626 (3565 and 3598) **number of nucleotide mismatches are given in parenthesis.

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Table 2.5 continued.

WSM 3612* 3626* 3636* 3803 3804* 3805 3919

M. albiziae 89.9 85.4 85.3 86.0 84.4 86.6 88.2 T CCBAU 61158 (27) (49) (50) (38) (46) (38) (39)

M. amorphae 90.8 89.6 89.3 88.5 87.7 89.1 90.7 T LMG 18977 (20) (31) (33) (30) (32) (28) (27)

M. australicum 91.2 94.3 93.6 91.0 93.9 90.3 84.3 T WSM2073 (19) (16) (19) (23) (16) (24) (45)

M. chacoense 88.0 86.1 85.7 85.2 84.8 87.4 85.7 T LMG19008 (28) (43) (45) (37) (41) (31) (42)

M. ciceri bv. biserrulae 85.7 90.6 90.6 80.7 90.6 88.2 90.6 T UPM-Ca7 (21) (27) (27) (25) (25) (25) (25)

M. huakuii 90.3 92.9 92.1 92.2 92.2 90.3 88.9 T IAM14158 (21) (21) (24) (20) (20) (24) (32)

M. loti 91.7 90.7 90.0 90.5 88.5 90.3 88.6 T LMG6125 (19) (27) (31) (25) (30) (25) (32)

M. mediterraneum 89.9 88.2 88.2 88.5 88.1 88.7 96.4 T UPM-Ca36 (22) (34) (35) (28) (30) (27) (10)

M. metallidurans 92.2 89.3 88.9 88.9 88.5 89.5 92.9 T LMG24485 (17) (32) (34) (28) (30) (26) (20)

M. opportunistum 89.4 91.8 92.1 91.0 91.0 90.3 87.9 T WSM2075 (23) (24) (24) (22) (23) (23) (35)

M. plurifarium LMG 89.4 86.8 87.8 87.7 86.5 88.7 84.3 T 11892 (23) (38) (36) (30) (35) (28) (45)

M. septentrionale 91.7 90.4 90.0 89.3 88.5 89.9 91.4 T SDW014 (18) (29) (31) (28) (30) (26) (25)

M. temperatum 91.2 88.6 88.5 88.9 88.5 89.1 96.8 T SDW018 (19) (33) (34) (27) (29) (26) (9) 84.3 82.1 81.0 82.3 80.3 81.9 81.8 T M. thiogangeticum SJT (34) (51) (55) (43) (48) (43) (51)

M. tianshanense 90.8 87.9 87.5 87.2 88.1 87.8 94.6 T USDA3592 (20) (36) (38) (32) (31) (30) (15) *Strains representing a clade or groups of strains: WSM2559 (Clades L and J); 2561 (I); 2622 (K); 3495 (E); 3606 (3806); 3626 (3565 and 3598) **number of nucleotide mismatches are given in parenthesis.

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2.3.5 16s rRNA phylogeny An internal fragment of the 16S rRNA gene of 1210 bp was successfully amplified for the 17 strains. Consistent with dnaK phylogeny, 16 of the strains clustered within the Mesorhizobium clade (Figure 2.5). However, strain WSM3602 (which failed to give amplification with dnaK as described in section 2.3.4) clustered with the genus Burkholderia which belongs to β-proteobacteria, and very closely related to the species B. tuberum.

Strains were distributed in four major clades within the genus Mesorhizobium in the phylogenetic tree developed with 16S rRNA sequences. Clade 1 consisted of strains isolated from L. annularis (WSM2623), L. frutescens (WSM3310), L. incana (WSM3919), L. microphylla (WSM2559 and 2622) and L. pauciflora (WSM2566). The strains WSM2559, 2623 and 3919 shared identical 16s rRNA sequences and had 3 or less nucleotide mismatches with the other strains included in this group. Clade 1 was congruent with the grouping and position of the strains in the dnaK tree, as they were in closely related clades in the dnaK phylogeny and intermingled with the type strains M. temperatum and M. mediterraneum in both phylogenies.

Clade 2 comprised only WSM3270 from L. annularis which was closely related to four Mesorhizobium type strains: M. metallidurans, M. tarimense, M. gobiense and M. tianshanense. In the dnaK phylogenetic tree, this strain was not close to any known Mesorhizobium species.

There were seven Lessertia strains with identical 16s rRNA sequences in clade 3 that clustered close to the type strains of M. australicum and M. shangrilense. These strains had been originally isolated from L. herbacea (WSM3598), L. diffusa (WSM3626 and 3565), L. capitata (WSM3495 and 3636) and L. excisa (WSM3898 and 3612). There was no congruence between the position of these strains in 16srRNA and dnaK phylogenies, as they were included in different clades in the dnaK tree.

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Figure 2.5. Neighbor joining phylogenetic tree based on 16S rRNA gene sequencing of 17 strains of Lessertia spp. root nodule bacteria. The type strain sequences in the phylogram were obtained from GenBank (accession number). M.: Mesorhizobium; S.: Sinorhizobium and R.: Rhizobium. * Initials in parentheses after the strain number indicate original host: L. annularis (La); L. capitata (Lc); L. diffusa (Ld); L.excisa (Le); L.frutescens (Lf); L.herbacea (Lh); Li: L.incana; L.microphylla (Lm); L.pauciflora(Lp). Capital letters in bold indicate grouping in dnak phylogeny.

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Clade 4 comprised the strain WSM3804 from L. herbacea that clustered with the type strain of M. opportunistum.

Table 2.6 shows 16s rRNA sequence similarity percentages between Lessertia symbionts and 14 Mesorhizobium spp. type strains. Two of the strains in the table represent a group of strains that had identical or nearly identical (1 nucleotide mismatch) 16S rRNA sequences; these were WSM2559 (representing WSM2566, 3310, 2623 and 3919) and WSM3495 (representing WSM2598, 3612, 3626, 3636, 3565 and 3898). All Lessertia strains showed more than 97% 16S rRNA sequence similarities with each Mesorhizobium type strain.

Strains WSM2559, 2561 and 2622 shared 99.9%, 99.8% and 99.7% similarities with M. temperatum respectively and had only 1, 2 and 3 nucleotide mismatches with this type strain (Table 2.6). WSM2559, 2561 and 2622 showed also high similarities (>99.7%) and less than 4 mismatches to the type strain M. mediterraneum.

WSM3270 shared 99.9% 16s rRNA sequence similarities with the species M. gobiense, M. metallidurans, M. tarimense and M. tianshanense, and had only 2 nucleotide mismatches with each of these type strains.

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Table 2.6. 16S rRNA gene sequence similarity of Lessertia RNB against the type strains of some of the RNB in the genus Mesorhizobium WSM strains Type strains 2559** 2561 2622 3270 3495** 3804 M.albiziae 98.8 (14)* 98.7 (15) 98.6 (16) 98.7 (16) 98.6 (19) 98.9 (15) CCBAU 61158T M. alhagi 97.5 (33) 97.4 (34) 97.1 (35) 98.0 (27) 97.9 (30) 97.4 (36) HAMBI3019T M.amorphae LMG 99.1 (13) 99.0 (14) 98.9 (15) 99.5 (9) 98.6 (20) 99.5 (6) 18977T M.australicum 98.3 (23) 98.5 (21) 98.4 (21) 98.4 (23) 99.8 (3) 98.5 (22) WSM2073T M.caraganae 99.5 (7) 99.4 (8) 99.3 (9) 99.8 (4) 98.6 (20) 99.0 (15) CCBAU11299T M.ciceri 98.4 (22) 98.3 (23) 98.2 (24) 98.6 (20) 99.2 (10) 98.0 (28) UPM-Ca7 T M.chacoense 98.2 (22) 98.4 (21) 98.4 (18) 98.8 (16) 98.3 (22) 98.6 (21) LMG19008T M.gobiense 99.5 (6) 99.4 (7) 99.3 (8) 99.9 (2) 98.7 (18) 99.1 (13) CCBAU83330T M.huakuii 98.8 (16) 98.7 (17) 98.6 (18) 98.9 (16) 98.2 (25) 99.2 (9) IAM14158T M.loti 98.6 (20) 98.5 (21) 98.4 (22) 99.0 (16) 99.3 (8) 98.1 (26) LMG6125T M.mediterraneum 99.8 (2) 99.8 (3) 99.7 (4) 99.4 (8) 98.6 (20) 99.1 (13) UPM-Ca36T M. metallidurans 99.5 (6) 99.4 (7) 99.3 (8) 99.9 (2) 98.7 (18) 99.1 (13) LMG24485T M.opportunistum 99.2 (12) 99.3 (10) 99.1 (12) 99.0 (15) 98.6 (20) 99.8 (3) WSM2075T M.plurifarium LMG 98.7 (17) 98.6 (18) 98.5 (19) 99.0 (15) 98.1 (26) 99.2 (10) 11892T M. septentrionale 99.0 (16) 98.9 (17) 98.8 (18) 99.4 (12) 98.6 (23) 99.4 (9) SDW014T M. shangrilense 98.6 (20) 98.5 (21) 98.4 (22) 98.6 (20) 99.7 (4) 98.5 (22) CCBAU65327T M. tarimense 99.5 (6) 99.4 (7) 99.3 (8) 99.9 (2) 98.7 (18) 99.1 (13) CCBAU83306T M.temperatum 99.9 (1) 99.8 (2) 99.7 (3) 99.5 (7) 98.6 (19) 99.2 (12) SDW018T M.thiogangeticum 97.8 (38) 98.0 (36) 98.0 (31) 97.7 (39) 97.5 (40) 98.4 (30) SJTT M.tianshanense 99.5 (6) 99.4 (7) 99.3 (8) 99.9 (2) 98.7 (18) 99.1 (13) USDA3592T * number of nucleotide mismatches are given in parenthesis ** Strains representing a clade or groups of strains: WSM2559 (Clade 1) and 3626 (Clade 4).

The group of strains represented by WSM3495 in table 2.5, shared 99.8% sequence similarities with M. australicum (3 nucleotide mismatches) and 99.7% similarities with M. shangrilense (4 nucleotide mismatches). The strain WSM3804 showed 99.8% sequence similarities and 3 mismatches with M. opportunistum.

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2.4 Discussion

In this chapter the species and strain diversity of rhizobia nodulating Lessertia spp. was assessed. The genus Lessertia is nodulated by Mesorhizobium spp. and by one strain identified as a Burkholderia sp.

There were clear differences in strain and species diversity depending on the host the isolates originated from. Each of the species L. capitata, L. frutescens, L. microphylla, L. excisa and L. incana were nodulated by closely related or identical strains and only with one or two species. In contrast, L. diffusa formed symbiosis with diverse strains and with 3 different Mesorhizobium species, and L. herbacea nodulated with 5 different species even though most of the isolates came from a common site.

Legumes able to form symbiosis with a broad range of species have been widely reported, even within the same plant and geographic area (Ba et al., 2002; Bala et al., 2002; Gao et al., 2001; Laguerre et al., 1997; Rincón-Rosales et al., 2009). Although it has been said that the ability to nodulate with different rhizobia species is beneficial for the establishment of the legume (Baraibar et al., 1999; Beauregard et al., 2004; Hungria & Vargas, 2000; Musiyiwa et al., 2005), it highly depends on the effectiveness of the symbiotic relationships, as there have been cases where the legume tends to nodulate with ineffective competitive soil rhizobia (Aouani et al., 2001; Ballard & Charman, 2000; Ben Romdhane et al., 2007; Nandasena et al., 2006). When the isolates from Lessertia were authenticated, all of them except strains WSM3602, 3620 and 3891, formed pink nodules in symbiosis with Lessertia spp. (Table 2.3) indicating that nitrogen was being fixed in most of the cases. Whether these strains effectively increase plant dryweight and are advantageous for the establishment of Lessertia spp. in the field is something that will be further analysed.

Although there have been rhizobial strains isolated from low pH soils that succeed in acid environments (Howieson et al., 1988), low pH tolerance cannot be strictly predicted from the pH at the site of isolation (Hungria & Vargas, 2000). These data

62 CHAPTER 2 showed no relationship between pH of the site of origin and the strains pH growth range evaluated in media. For example strains WSM3592 and WSM3620 were both isolated from a site with a soil pH of 6.5 (Appendix 10), but WSM3592 was able to grow from pH 4.0 to 9.0 whereas WSM3620 grew only at pH 6.0 or higher. Interestingly, the strains that showed low pH intolerance (growth only at pH 5.5 or higher) were all clustered together in clades H, I and J in the dnaK phylogenetic tree (Figure 2.4) and in clade 1 in the 16S rRNA tree (Figure 2.5) close to the species M. mediterraneum and M. temperatum, suggesting that the ability of some strains to tolerate low pH may be associated to their phylogenetic position.

The techniques used for fingerprinting and determining strain diversity in this study were ERIC-PCR and RPO1-PCR. It has been suggested ERIC-PCR is a more discriminatory fingerprinting tool for rhizobia, being able to detect differences between strains that are not evident through RPO1-PCR fingerprints or other fingerprinting techniques (Jaspers & Overmann, 2004; Yates, 2008). Other authors have reported RPO1 being more divergent than ERIC-PCR fingerprints between different strains (Garau et al., 2005). In this work, ERIC and RPO1-PCR fingerprints were congruent with minor differences. However, the strain typing with the RPO1 primer generated more bands per isolate and made the analysis and clustering of the fingerprinting patterns more reliable. Therefore, RPO1-PCR was selected as the preferred fingerprinting technique to be used for Lessertia strains in future experiments.

The species diversity of Lessertia isolates was inferred from the partial sequencing of the genes dnaK and 16S rRNA. The dnaK 330 bp variable 3’ fragment was chosen as a first approach to describe the identity of the 44 strains as it is a taxonomic marker that has shown good correlation with other housekeeping genes (Chelo et al., 2007; Martens et al., 2007; Nandasena et al., 2009; Ormeño-Orrillo et al., 2006; Stępkowski et al., 2003a). The partial gene 16S rRNA was sequenced for some strains selected from the different dnaK tree clusters, to confirm their taxonomic position, given its value in intrageneric phylogenetic evaluation and because of the large quantity of data available for this gene (Graham et al., 1991; Stackebrandt & Goebel, 1994).

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The overall taxonomic position of Lessertia spp. strains in the dnaK phylogeny was within the genus Mesorhizobium, a position that was further confirmed with the sequencing of 16S rRNA. However, the position of the strains at the intra-genus level was incongruent between phylogenies. Some of these discordances were due to a lower sequence divergence between strains in the 16S rRNA analysis than with the dnaK sequences. Clear examples of this were the strains in clade 3 in the 16s rRNA phylogenetic tree (Figure 2.5) that showed identical 16s rRNA sequences but in the dnaK phylogeny were split into three well supported clusters (clades A, C and E) (Figure 2.4). Furthermore, the strain WSM3270 clustered on its own in dnaK phylogenetic tree, whereas its 16s rRNA sequence was nearly identical to the type strains M. metallidurans, M. tarimense, M. gobiense and M. tianshanense. This is not surprising as it has long been discussed that the use of 16S rRNA as a tool to estimate bacterial phylogeny is not able to determine differences between some closely related species and therefore provides a poor resolution below the genus level (Brenner et al., 2005; Martens et al., 2007; McArthur, 2006; Ormeño-Orrillo et al., 2006). dnaK has shown a better resolution of internal branches specially within the genus Mesorhizobium and Bradyrhizobium (Alexandre et al., 2008; Eardly et al., 2005; Kwon et al., 2005; Stępkowski et al., 2003a). This low sequence divergence could be in part attributed to the differential rates of evolution between core genes; as 16S rRNA has shown to be a slowly evolving molecule in comparison to other housekeeping genes and therefore lacks the required level of resolution to distinguish similar species (Chelo et al., 2007; Eardly et al., 2005; Hanage et al., 2006; Stępkowski et al., 2003b)

There were obvious incongruities in the positioning of some strains between both phylogenies. The closest neighbouring strain for WSM3804 in the dnaK phylogenetic tree was M. australicum (Figure 2.4), while it was closely related to M. opportunistum in the 16s rRNA phylogeny and included in a clearly separated cluster from M. australicum (Figure 2.5). Another example were the strains comprised in clade A of the dnaK tree that showed identical dnaK sequences to that of WSM3872 isolated from Biserrula, whereas in the 16s phylogeny they were not related to this strain but to M. australicum and M. shangrilense. The inconsistency between the array of 16S rRNA sequences and those of other housekeeping genes has been widely reported within the order Rhizobiales (Ba et al., 2002; Eardly et al., 2005; Gao et al., 2004;

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Gaunt et al., 2001; Nandasena et al., 2009; Ormeño-Orrillo et al., 2006; van Berkum et al., 2003). There is evidence that 16s rRNA can occasionally undergo lateral transference and genetic recombination between similar species distorting some of the relationships inferred from its sequencing (Gaunt et al., 2001; Hanage et al., 2006; Stępkowski et al., 2003b; van Berkum et al., 2003). There is no evidence so far of transfer of the dnaK gene, but there have been discordances between that loci and groups of other housekeeping genes that are well correlated between them, suggesting that dnaK transfer may as well be possible (Parker, 2004; Stępkowski et al., 2007).

In spite of the discordances between both phylogenies, there were strains that were consistently clustered together in the dnaK and 16S rRNA trees. This group was composed of seven strains (WSM 2559, 2623, 3919, 3310, 2566, 2561 and 2622) that were closely related to M. temperatum and M. mediterraneum in both phylogenetic trees (Figures 2.4 and 2.5). Interestingly, all the strains grouped in this clade, including the other 12 strains that were clustered with these species in the dnaK phylogeny, were characterised as slow growing bacteria (Table 2.3). Coincidently with these results, Gao et al. (2004) described M. temperatum as able to form a single 1mm colony within 7 to 10 days, whereas M. mediterraneum forms colonies in 4 to 5 days (Nour et al., 1995). The similarities in rate of growth and the high sequence similarity between this group of slow growing Lessertia strains and M. temperatum, suggests that these strains are more closely related to the latter than to M. mediterraneum. Nevertheless, to fully resolve the taxonomic position of this group it is necessary to carry out additional phylogenetic analyses.

Instances like the close relationship between M. mediterraneum and M. temperatum, where the boundaries between species have not been clear enough through phylogenetic studies, have occurred with other species within the genus Mesorhizobium (Alexandre et al., 2008) and in every one of these cases species identity has been ultimately defined by DNA:DNA hybridization (Chen et al., 2008; Gao et al., 2004; Vidal et al., 2009). Wei and collaborators (2009) believe that cases like these need re-examination, as although DNA:DNA hybridization is a valuable technique to define species taxonomic position, it involves pairwise comparison of the whole genome including “transferable” accessory genes, that may separate

65 CHAPTER 2 species that have similar or identical core genome (Doolittle & Papke, 2006; Hanage et al., 2006; Turner et al., 2002; Young et al., 2006).

Although the dnaK gene for the strain WSM3602 could not be amplified, its 16S rRNA gene clustered, with 100% bootstrap support, with the type strain Burkholderia tuberum STM678. Nodulation with β-proteobacteria by plants that normally nodulate with α-rhizobia has been previously reported and in most of the cases it has been ineffective (Chen et al., 2003; Yan et al., 2007) and attributed to the ability of Burkholderia spp. to compete particularly under poor soil conditions and low N availability (Elliott et al., 2009; Lindeque, 2005). Coincidently with this, the strain WSM3602 formed white and apparently ineffective nodules on its original host L. herbacea

The use of 16S rRNA, in spite of its limitations, is still a standard in the description of new species, especially because of the high number of sequences available for comparison (Tindall et al., 2010). It has been suggested that a 3% difference in 16S rRNA sequences should be used to place strains into different species (Brenner et al., 2005; Cohan, 2002; Tindall et al., 2010) which means that strains with less than 97% 16s rRNA sequence similarities can be assigned confidently into different species, whereas strains with more than 97% similarities should be studied further (Hanage et al., 2006). Results from this study show that all the 16S rRNA sequences from Lessertia isolates, excluding the Burkholderia strain WSM3602, showed more than 97% similarity with Mesorhizobium type strains (Table 2.6).

In view of the high 16S rRNA sequence similarities between strains and the discordance of 16s phylogeny with that of dnaK gene, the exact taxonomic position of these strains should be verified by additional approaches. There are several available techniques that used in combination may provide a clear species delineation. DNA-DNA hybridization, despite the before mentioned drawbacks, is still acknowledged as a standard for species delineation (Stackebrandt & Goebel, 1994; Stackebrandt et al., 2002; Tindall et al., 2010; Wayne et al., 1987). An alternative and relatively rapid and inexpensive approach is to sequence several housekeeping genes to provide more informative nucleotide sites to resolve similar species (Stackebrandt et al., 2002; Tindall et al., 2010; Zeigler, 2003). Recently, a multi locus

66 CHAPTER 2 sequence analysis (MLSA) approach was developed for this purpose, which consists of analysing the phylogeny and clustering of concatenated sequences of multiple housekeeping genes (Hanage et al., 2006; Mignard & Flandrois, 2008). This technique has resulted in well supported phylogenies and seems to be one of the most promising approaches to clarify rhizobial systematics and to define Mesorhizobium spp. (Alexandre et al., 2008; Martens et al., 2008; Nzoué et al., 2009; Vinuesa et al., 2005b).

There was no apparent effect of the geographic origin of the Lessertia strains and the way they clustered in dnaK and 16S rRNA phylogenies, as strains collected from the different regions of South Africa were distributed indistinctly in different clades. A relationship between geographic distribution of the strains and their phylogeny was expected as, although the strains assessed were not collected from widely separated areas, they did come from different ecological areas within South Africa’s Western and Northern Cape, and environment has proved to play a determinant role in microbial distribution (Martiny et al., 2006).

From a broader perspective no phylogeographic array was detected. Most of the clades in the dnaK and in the 16s rRNA phylogenies comprised strains isolated from different continents. Other authors have also reported little correspondence between geographic origin of rhizobia and their 16S rRNA phylogeny and attributed this to intercontinental migration and to the slow evolution rate of 16S rRNA gene (Donate- Correa et al., 2007; Parker et al., 2002; Parker & Kennedy, 2006). Stępkowski et al. (2007) suggest that the geographic structure of housekeeping genes has been probably erased not only by a series of dispersals but also through gene recombination events.

There was a group of clades in the dnaK phylogeny, though, that could be considered a geographic cluster. The strains included in clades A to E, including WSM3872 isolated from Biserrula sp., and the next neighbouring clade comprising M. plurifarium (de Lajudie et al., 1998) and WSM3869 from Biserrula sp. in Eritrea, were all originally isolated from legumes growing in Africa and therefore would be conforming to an African clade. M. plurifarium strains have subsequently been isolated from South America as well (Vinuesa et al., 2005b).

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Another possible geographic cluster would be the one comprising WSM3602 and B. tuberum STM678 isolated from Aspalathus carnosa (Moulin et al., 2001; Vandamme et al., 2002) in the 16S rRNA phylogenetic tree. The strain DUS833 from Aspalathus callosa (Muofhe & Dakora, 1998) (not included in this phylogenetic tree) has shown identical 16s rRNA sequences with STM678 and high sequence similarity with symbionts from Cyclopia spp. (Elliott et al., 2007; Kock, 2004). Both Aspalathus spp. and Cyclopia spp. are native to the fynbos biome in the South Africa’s Western Cape (Cocks & Stock, 2001; Elliott et al., 2007; Spriggs & Dakora, 2007) that matches with the area where WSM3602 was collected. Therefore, this seems to be a 16S rRNA sequence cluster restricted to a well delimited area in South Africa. However, studies of additional wild legumes in South Africa and their symbionts, as well as the study of more housekeeping genes would be necessary to further support this finding.

While there is still work to be done regarding the exact taxonomic position of Lessertia rhizobial symbionts, based on these results it can confidently be said that their preferred symbiont under natural conditions are Mesorhizobium spp.

Furthermore, some Mesorhizobium spp. have the ability to transfer symbiotic genes through a symbiosis island (Nandasena et al., 2006; Sullivan et al., 2002). Hence, symbiotic gene transfer within Mesorhizobium species could explain the diversity of species nodulating Lessertia spp. Moreover, the fact that Lessertia herbacea and other legume species from similar geographic origins are establishing symbiosis with the species B. tuberum could be suggesting either that B. tuberum is a highly promiscuous root nodule bacterium or that there has been transfer of symbiotic genes that allow this to happen.

In order to interrogate these possibilities, in the next chapter the phylogeny of nodulation genes will be studied and compared with the chromosomal background of the strains. It will also comprise studies of nodulation and nitrogen fixing abilities of Lessertia strains as they should be key steps before introducing this species into the field.

68

CHAPTER 3. Symbiotic effectiveness and nodulation gene phylogeny

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70 CHAPTER 3

3.1 Introduction Compatibility between a legume and its rhizobial symbiont is essential for nodulation and nitrogen fixation (Graham, 2008). Nodule formation is a complex multi-step process that involves signal exchanges between both symbionts. One group of host- specific signalling molecules are the Nod factors, which are produced by rhizobia in response to root exudates and are determinant in initial root hair infection (Kobayashi & Broughton, 2008; Maunoury et al., 2008). The nodulation genes are responsible for the synthesis and regulation of Nod-factors. Of these, the common nodulation gene nodA is one of the most frequently sequenced and studied, because it plays a critical role in determining the structure of the Nod-factor and therefore in determining host specificity (Broughton & Perret, 1999; Kobayashi & Broughton, 2008; Roche et al., 1996). The characterisation and phylogenetic classification of symbiotic genes have become an important basis for the understanding of the rhizobium-legume symbiosis (Laguerre et al., 2001).

The study of the interaction between plant and symbionts and the selection of effective nitrogen fixing rhizobia are critical steps to optimize the symbiotic relationship and to maximise legume productivity once a legume is introduced into a new environment (Herridge, 2008; Howieson et al., 2000a; Lindström et al., 2010; O'Hara et al., 2002). In the particular case of Lessertia spp. there is a further important issue that should be taken into consideration. The species L. capitata, L. diffusa, L. excisa, L. herbacea, L. incana and L. pauciflora are associated predominantly with root nodule bacteria of the genus Mesorhizobium (see Chapter 2). Some species of Mesorhizobium have been reported to be able to transfer symbiotic genes to soil bacteria via a ‘symbiosis island’ (Nandasena et al., 2007a; Sullivan & Ronson, 1998). This could eventually become a major threat for any legume with agricultural potential, as non-symbiotic bacteria can become ineffective competitors in the field.

This chapter seeks to sequence the nodulation gene nodA from Lessertia nodulating strains to determine and analyse nodA phylogeny. Further, the relationship between nodA phylogeny and geographical region of isolation of the strains will be studied. In addition, the presence of symbiosis islands amongst the Mesorhizobium strains from

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Lessertia will be investigated. Given the importance of understanding the relationship between a legume and its symbionts before their establishment in field, this chapter also seeks to study the interactions between the recently introduced South African legumes L. capitata, L. diffusa, L. excisa, L. herbacea, L. incana and L. pauciflora and their diverse microsymbionts, and to select effective nitrogen fixing bacteria.

3.2 Materials and Methods 3.2.1 Study of nodulation gene nodA The nodA gene was amplified and sequenced in order to determine the symbiotic phylogeny of 34 strains (Appendix 7). The primers used in this study were nodA1 and nodA2 which are detailed in Table 3.1.

Table 3.1. Sequences of PCR primers used in this work Name Sequence (5’ 3’)* Target Source gene nodA1 TGCRGTGGAARNTRNNCTGGGAAA nodA Haukka et al. (1998) nodA2 GGNCCGTCRTCRAAWGTCARGTA nodA Haukka et al. (1998) phetRNAf GCCCAGATAGCTCAGTTGGT Phet- Nandasena et al. (2006) intS intS522r ATTGCATATCGAAACACG Phet- Nandasena et al. (2006) intS msi001-KF GTGCAGCCTACCGGATCTCG intS Nandasena et al. (2004) msi001-KR CCAGATAATCGGCCCACCAT intS Nandasena et al. (2004) *Symbols: A, C, G, T – standard nucleotides; R: A,G; W: A,T; N: all.

Cell templates were prepared as described in section 2.2.1 with the exception that the cells where concentrated to an OD600 of 2.0. The PCR reaction mixture comprised 2 µl of cell suspension, 0.5 µl of Taq polymerase (Invitrogen Life Technologies), 1 µl of forward and reverse primers (50μM) (Geneworks), 6.0 µl of

25mM MgCl2, 20.0 µl 5x Fisher-Biotech polymerization buffer and 69.5 µl of UltraPure grade water (Fisher Biotech) to total 100 µl for a single reaction. The cycling conditions were: 5 minutes at 94°C followed by 5 cycles of 94°C for 30 s, 52°C for 30 s and 72°C for 1min; 30 cycles of 94°C for 30s, 55°C for 30s and 72°C for 1 min; and a final hold at 72°C for 5 min. The annealing temperature was reduced to 52°C for the strain WSM3310.

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The amplification of the 600bp partial nodA sequence was verified by agarose gel electrophoresis as described in section 2.2.2. The PCR products were purified and sequenced following the procedure described in section 2.2.5.

The chromatograms were analyzed and edited in Gene Tool Lite 1.0 (2000) software and the phylogenetic analysis were conducted in MEGA4 (Tamura et al., 2007) (section 2.2.7). The BLAST search tool from the National Centre for Biotechnological Information (NCBI) was used to find nodA sequences closely related to those from Lessertia spp. strains and to download known nodA sequences to be included in the phylogenetic tree.

3.2.2 Nodulation specificity and symbiotic effectiveness 3.2.2.1 Experimental design A subset of 17 Lessertia strains (Appendix 3), selected from different dnak (section 2.3.4) and nodA phylogenetic clusters, were examined for their nodulating and nitrogen fixing abilities with six Lessertia species: L. capitata, L. diffusa, L.excisa, L. herbacea, L. incana and L. pauciflora.

The experiment was designed as a split-plot system with one rhizobial strain per pot as a main treatment, two Lessertia spp. as sub-treatments and three replicates per treatment. Two sets of uninoculated controls, with added nitrogen (N+) and nitrogen free (N-), were included for all species.

3.2.2.2 Experimental preparation, planting and inoculation Seeds of the six Lessertia species were scarified, sterilized and germinated in 1.5% (w/v) water agar plates (section 2.2.3). Following radicle emergence, the seedlings were planted in 500ml free-draining plastic pots filled with a steamed sand mix consisting of 2 parts of river sand and 3 parts of leached yellow sand [modified from Howieson et al.(1995)]. The substrate was flushed with boiling sterile DI water to remove inorganic N before sowing.

The 17 bacterial cultures were grown on ½ LA plates for 5 to 10 days at 28°C. Inoculation was performed three days after planting with 1 ml of bacterial suspension

73 CHAPTER 3 aseptically washed from fresh colonies and suspended in 1% (w/v) sucrose solution to an OD600nm of 1.0.

The pots were initially covered with plastic film to avoid contamination with air-borne rhizobia. Once the plants had emerged and were 2 cm in height, the plastic film was removed and the exposed soil surface was covered with sterile alkathene beads. A sterile Polyvinyl chloride (PVC) watering tube with cap was pushed in the centre of each pot to a depth of 5 cm for supply of water and nutrient solution.

3.2.2.3 Growth conditions The plants were grown for 8 weeks in a naturally-lit phytotron, maintained at 22ºC during the day time and under axenic conditions. The pots were arranged on the bench in a randomized design.

Plants were supplied weekly with a nutrient solution devoid of nitrogen (section 2.2.3) at a rate of 20 ml per pot. The nitrogen fed controls were additionally supplied with 10 ml of 0.1 M KNO3 solution on a weekly basis. All plants were watered every two days with 20 ml of sterile DI water.

3.2.2.4 Evaluations and Statistical analysis At harvest plants were carefully removed from the pots and the roots were washed under running tap water. The colour and general appearance of the plant was recorded as well as the number, morphology and distribution of the nodules. Shoot tops were dried at 60°C for 48 hours and weighed. Two nodules were collected from each of the replicates and the bacteria were re-isolated to verify strain identity through RPO1-PCR fingerprinting (sections 2.2.1 and 2.2.2).

Dry weight data were analysed and presented as percentage of the nitrogen fed control. The analysis of variance of the yields of each of the plant hosts across the different inoculants was carried out with the statistics programme SPSS 13.0 for Windows (2004). The mean separation test used was the Fisher’s least significant difference (LSD) test with α=0.05.

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Strains were classified into three groups based on the average plant yield across replicates; effective= >75% of +N control (E), partially effective= >20% but <75% of +N control (PE) and ineffective= <20% of +N control (I) (Yates, 2008). This classification was modified for L. herbacea increasing the limit between PE and I strains from 20% to 40%, due to the high dry weight of the uninoculated plants caused by a larger seed size.

3.2.3 Sequencing of the symbiosis island insertion region The intS gene codes for the protein responsible for the excision and insertion of the symbiosis island in Mesorhizobium which is then inserted into a Phe-tRNA gene (Sullivan & Ronson, 1998). To test for the presence of symbiosis islands amongst 16 Mesorhizobium strains isolated from Lessertia spp., the amplification of the insertion regions phetRNA-intS and intS was attempted.

Cell templates for each of the 16 strains were prepared as previously described and adjusted to an OD600 of 2.0 (section 2.2.3). The PCR reaction mixture was the same as in section 2.2.3. Initially the primers phetRNAf and intS522r (Table 3.1) were used to try to amplify the phetRNA-intS region. When amplification failed, the intS region was amplified using the primers msiOO1-KF and msi001-FR (Table 3.1). The PCR cycling conditions were: 5 min at 95°C followed by 30 cycles of 94°C for 30s, 50°C for 30s and 72°C for 30s; and a final hold at 72°C for 5 min. Annealing temperatures ranging from 45 to 50° C were tested for those strains that did not amplify the region with the standard PCR conditions.

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3.3 Results 3.3.1 Study of nodulation gene nodA The partial nodA gene was successfully amplified for 34 strains and the assembled sequence was approximately 600bp long. Most of the strains’ nodA genes were grouped in a large monophyletic clade (Figure 3.1), that was subdivided into 6 well supported clades (A to F).

All the sequences included in the Lessertia nodA clade shared few similarities with any other published nodA sequences on GenBank. The only exceptions were the L. herbacea strains in clade E that clustered and had more than 96% similarity with the nodA sequence of the Mesorhizobium sp. strain DUS835. This strain was originally isolated from the South African species Aspalathus linearis (rooibos tea) in the Western Cape (Elliott et al., 2007).

Clade A comprised nodA sequences from L. diffusa strains WSM3503, 3917, 3513, 3492, 3502 and 3620, and two strains from L. incana: WSM3920 and 3919 (Figure 3.1). Clade B included four strains: WSM2559 (L. microphylla), WSM2566 (L. pauciflora), WSM2561 (L. diffusa) and WSM3270 (L. annularis). The nodA sequences of the L. microphylla strains WSM2607 and WSM2622 were identical and formed clade C. Clade D comprised three strains from L. capitata: WSM3636, 3507 and 3495, and WSM3626 from L. diffusa. Clade E comprised the strains WSM3804, 3803, 3805 and 3806 all of them isolated from L. herbacea and the Mesorhizobium strain DUS835. Clade F included strains WSM3891, 3898, 3612, 3472, 3893, 3894 and 3900 isolated from L. excisa, WSM3565 and 3564 from L. diffusa and the L. herbacea strain WSM3598.

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Figure 3.1. Neighbour joining tree based on partial nodA sequencing of 34 strains of Lessertia spp. root nodule bacteria. The type strain sequences in the phylogram were obtained from GenBank (accession number). Initials after WSM strain number indicate original host (L. annularis (La); L. capitata (Lc); L. diffusa (Ld); L. excisa (Le); L. frutescens (Lf); L. herbacea (Lh); Li: L. incana; L. microphylla (Lm); L. pauciflora(Lp). Different colours indicate different geographic origin (Figure 3.2).

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The two strains that were not included in the large clade described above were WSM3602 which clustered with Burkholderia tuberum DUS833 (Elliott et al., 2007) and Burkholderia sp. WSM3937 (Garau et al., 2009) nodA sequences and WSM3310 from L. frutescens which had a 100% nodA sequence similarities to that of Sinorhizobium medicae WSM419 (Reeve et al., 2010).

Although the nodA sequences from strains isolated from L. incana, L. excisa and L. capitata tended to cluster together and clade E comprised only L. herbacea strains, there is no evidence of a relationship between the original host species and the grouping of the strains in the nodA phylogeny. The nodA sequences from L. diffusa are distributed in 4 of the clades and those from L. herbacea’s symbionts in three different clades.

Figure 3.2. Map of South Africa showing the location of sites from where isolates from Lessertia spp. were collected. Colours relate to Figure 3.1.

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The nodA sequences included in each of the clades appear to relate to rhizobia having a common or close geographic origin. Strains included in clade A (Figure 3.1) had all been isolated from three sites located in the western area of the Northern Cape (Figure 3.2) and Clades C and E comprised strains isolated from the same sites in the Western Cape. Likewise, strains included in clade F had all been isolated from a nearby area in the Western Cape and in clade D, three of the strains had been isolated from the same site in the Western Cape. The other strain included in this clade (WSM3626), although located in the Northern Cape, was less than 100 km away.

The only exception was cluster B that includes strains that were originally distributed along the southern area of the Northern Cape, in the Western Cape and in the Eastern Cape.

3.3.2 Nodulation specificity and symbiotic effectiveness

There were differences in nodulation patterns and symbiotic effectiveness depending on the strain used and the plant host combination. Some hosts (L. herbacea, L. excisa and L. diffusa) were able to nodulate with most of the strains, whereas L. pauciflora and L. incana could only nodulate with a small number of strains.

L. diffusa was able to form nodules with 14 strains and 10 of these produced significantly higher dry weights than the uninoculated control (Figure 3.3). Plants of L. diffusa inoculated with two strains, WSM3636 and WSM3898, achieved statistically similar dry weights to the nitrogen control and with WSM3598 L. diffusa produced higher dry weight than the nitrogen control (P≤0.05).

L. herbacea was able to nodulate with 16 strains, of which WSM2559, WSM2566, WSM3636, WSM3598, WSM3612 and WSM3565 resulted in larger dry weights than the uninoculated control. The strains WSM3636, WSM3598 and WSM3612 increased dry weights of L. herbacea to the level of the nitrogen fed control (P≤0.05).

Ten of the strains formed nodules with L. capitata, albeit only six of these (WSM2559 WSM3636, WSM3612, WSM3898, WSM3565 and WSM3626) significantly increased

79 CHAPTER 3 dry weights in comparison to the uninoculated plants (P≤0.05) and none of them to the level of the nitrogen fed control (Figure 3.3). L. incana was able to nodulate with 10 strains, and only WSM3598 and WSM3612 exceeded the uninoculated control dry weight, but these strains achieved lower yields than the nitrogen fed control.

L. diffusa L. herbacea

120 * 120 * 100 * 100 * * 80 80 * * * * * * * 60 * * * 60 * * 40 40

20 20

0 0

120 120 L. capitata L. incana

WS M3919WS M2622WS M3804WS M2566WS M3310WS M2623WS M3602WS M2559WS M3270WS M2561WS M3626WS M3565WS M3612WS M3495WS M3898WS M3636WS M3598

WS M2622 WS M3804 WS M2566 WS M3310 WS M2623 WS M3602 WS M2559 WS M3270 WS M2561 WS M3626 WS M3565 WS M3612 WS M3495 WS M3898 WS M3636 WS M3598

100 WS M3919 Uninoculated100 Uninoculated

80 80

60 60 * * * 40 * 40 * * * * 20 20

0 0

Shoot dry weight (% +N control) control) +N +N (% (% ShootShoot drydryweight weight

120 L. excisa 120 L. pauciflora WS M3919WS M2622WS M3804WS M2566WS M3310WS M2623WS M3602WS M2559WS M3270WS M2561WS M3626WS M3565WS M3612WS M3495WS M3898WS M3636WS M3598 WS M3919WS M2622WS M3804WS M2566WS M3310WS M2623WS M3602WS M2559WS M3270WS M2561WS M3626WS M3565WS M3612WS M3495WS M3898WS M3636WS M3598 Uninoculated100 Uninoculated100

80 80

60 60

40 40 * * 20 * * 20

0 0

WS M3919WS M2622WS M3804WS M2566WS M3310WS M2623WS M3602WS M2559WS M3270WS M2561WS M3626WS M3565WS M3612WS M3495WS M3898WS M3636WS M3598 WS M3919WS M2622WS M3804WS M2566WS M3310WS M2623WS M3602WS M2559WS M3270WS M2561WS M3626WS M3565WS M3612WS M3495WS M3898WS M3636WS M3598 Uninoculated Uninoculated

Figure 3.3. Shoot dry weights for L. diffusa, L. herbacea, L. capitata, L. incana, L. excisa and L. pauciflora inoculated with 17 different rhizobial strains. Solid green bars indicate nodulation, bars with diagonal stripes correspond to non-nodulated treatments. The red line indicates 70% of the nitrogen control dry weight, which corresponds to the level above which nodulation is considered effective. * Significant difference from the uninoculated control (P0.05)

L. excisa formed nodules with 13 of the 17 inoculated strains, of which WSM3612, 3898, 3565 and 3626 were able to increase shoot dry weight (P≤0.05) in comparison to the uninoculated control (Figure 3.3). However, none of these treatments increased dry weight to the level of the nitrogen fed control. L. pauciflora nodulated

80 CHAPTER 3 only with the strains WSM3495, 2623 and 3270 and in every case nodulation was ineffective.

Some strains were able to effectively nodulate more than one Lessertia species (Table 3.2). One of these is WSM3636 which effectively nodulated L. diffusa and L. herbacea, and while only partially effective with L. capitata, it was the strain that reached the highest dry weight with that host. Similarly, WSM3598 was effective with the same species and partially effective when inoculated onto L. incana, where it also achieved the highest dry weight. The strains WSM3612 and 3898 effectively nodulated L. herbacea and L. diffusa respectively, but also were able to increase dry weights in all the other species, with the exception of L. pauciflora.

Table 3.2. Host-strain interaction for 6 Lessertia spp. and 17 root nodule bacteria. N2 fixation categories are: E, effective; PE, partially effective; I, Ineffective; X, no nodulation. Strain Original Host L.capitata L.diffusa L.excisa L.herbacea L.incana L.pauciflora WSM2559 L. microphylla PE PE I PE PE X WSM2561 L. diffusa X PE I I X X WSM2566 L. pauciflora PE PE I PE X X WSM2622 L. microphylla X X I I X X WSM2623 L. annularis X PE X X I I WSM3270 L. annularis I PE I I X PE WSM3310 L. frutescens X PE X PE X X WSM3495 L. capitata I PE I I PE I WSM3565 L. diffusa PE PE PE E I X WSM3598 L. herbacea X E I E PE X WSM3602 L.herbacea X X X I X X WSM3612 L. excisa PE PE PE E PE X WSM3626 L. diffusa PE PE PE PE I X WSM3636 L. capitata PE E I E I X WSM3804 L. herbacea X I X I X X WSM3898 L. excisa PE E PE PE PE X WSM3919 L. incana I X I I I X

There was no evidence of a relationship between the original host species of origin of the strains and the plant nodulation preferences. Lessertia spp. nodulated indiscriminately with strains isolated from other hosts as with those isolated from their own species. For example, L. herbacea nodulated effectively with strains isolated from L. capitata (WSM3636), L. excisa (WSM3612), L. diffusa (WSM3565) and only one isolated from itself (WSM3598). Moreover, L. diffusa nodulated effectively with strains from L. herbacea (WSM3598), L. capitata (WSM3636) and L. excisa (WSM3898).

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There were strains that had been originally isolated from hosts that were not included in this study, such as WSM2559 and WSM2622 (L. microphylla), WSM2623 and WSM3270 (L. annularis) and WSM3310 (L. frutescens). These strains were all capable of forming symbiosis with at least two of the hosts included in the study (Table 3.2). The strains isolated from L. microphylla were quite different in their nodulation patterns; WSM2559 was able to nodulate and increase dry weight in four species (L. capitata, L. diffusa, L. herbacea and L. incana) in comparison to the uninoculated controls (P≤0.05) (Figure 3.3, Table 3.2), whereas WSM2622 nodulated only L. excisa and L. herbacea and in both cases ineffectively.

3.3.3 Sequencing of the Symbiosis island insertion region An 800-bp fragment of the symbiosis island was successfully amplified for the Mesorhizobium strains WSM2561 (L. diffusa) and WSM3495 (L. capitata) using standard PCR procedures with an annealing temperature of 50°C. The region amplified from WSM3495 showed 82 % similarity to the phe-tRNA-intS region of M. loti strain MAFF303099 (Kaneko et al., 2000) and 81% to that of M.ciceri bv. biserrulae WSM1271 (Nandasena et al., 2007a) (Appendix 11). No sequence similarity to the target gene was found for the region of WSM2561. Three other strains (WSM2566, 2623 and 3310) amplified a 1,800bp fragment with these primers but the sequence of this product did not correspond to the target gene. For all other 13 strains no amplification was possible, even when using the alternate primers msi001-KF and msi001-KR and reducing the annealing temperature to 45°C.

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3.4 Discussion The first aim of this chapter was to study the phylogeny of the common nodulation gene nodA for 34 strains of Mesorhizobium spp. isolated from root nodules on Lessertia spp. The nodA genes were shown to be clustered in a robust (100% bootstrap support) monophyletic clade separated from other known Mesorhizobium spp. nodA sequences. This indicates that nodA has diverged from a common ancestor for this group of strains (Lecointre & Le Guyader, 2006) and has had a long history of separate evolution (Ochman et al., 2000; Wei et al., 2009).

The overall topology of the nodA tree was not congruent with 16S rRNA and dnaK phylogenies (Chapter 2), as some strains that had clustered with distinct Mesorhizobium spp. in the core gene phylogenies shared similar nodA genes. For example the group of strains in clades A, B and C (Figure 3.1) had previously shown nearly identical 16S rRNA and dnaK genes to those of M. mediterraneum and M. temperatum (Chapter 2), but their nodA sequences are closer to other Lessertia strains than to M. mediterraneum and M. temperatum.

Several research groups have reported the discordance of symbiotic and core loci phylogenies in rhizobial strains (Chen et al., 2003; Donate-Correa et al., 2007; Haukka et al., 1998; Laguerre et al., 2001; Moulin et al., 2004; Ormeño-Orrillo et al., 2006; Parker et al., 2002; Parker & Kennedy, 2006; Stępkowski et al., 2003b; Stępkowski et al., 2007; Suominen et al., 2001; Wernegreen & Riley, 1999) as well as the tendency of symbiotic genes from strains from a particular legume to cluster in separate clades despite their chromosomal background (Andam et al., 2007; Ba et al., 2002; Chen et al., 2008; Donate-Correa et al., 2007; Han et al., 2010; Haukka et al., 1998; Kalita et al., 2006; Moulin et al., 2004; Stępkowski et al., 2005; Vinuesa et al., 2005c; Wei et al., 2009). Both phylogenetic discordances and monophyletic clustering are often attributed to lateral gene transfer during the diversification of the family (Ochman et al., 2000; Sullivan et al., 1995; van Berkum et al., 2003) causing the separation of populations into subgroups that differ in symbiotic properties but share a common core gene pool (Doolittle & Papke, 2006).

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The discordance between nodA and housekeeping genes from Lessertia strains could imply that there has been lateral transfer of symbiotic elements. However, when a gene transfer event takes place it would be expected that strains would share a high degree of similarity for the gene in question (Ochman et al., 2000). This does not seem to be the case, as the nodA sequences are divided in deeply divergent branches. The only cases of high sequence similarities were observed within the clades A, B, C and F (Figure 3.1). However, these strains also clustered in the same groups in core gene phylogenies, suggesting that sequence similarities are not due to lateral transfer of nodA but to co-evolution of chromosomal and symbiotic genes. Therefore, symbiotic gene transfer within different Mesorhizobium spp. associated with Lessertia spp. seems to have happened early in the symbionts’ evolution and recent lateral transfer events do not seem to have played an important role.

Legume symbionts from undisturbed environments often show congruence between symbiotic and housekeeping genes suggesting that vertical gene transmission is a more significant event than symbiotic gene transfer during rhizobial evolution (Aguilar et al., 2006; Chen et al., 2003; Chen et al., 2008; Elliott et al., 2009; Han et al., 2010; Wei et al., 2009; Wernegreen et al., 1997; Wernegreen & Riley, 1999; Zhang et al., 2001). Lateral transfer of symbiotic genes is more evident in agricultural settings, where strains are subject to human selective pressures (Han et al., 2010; Laguerre et al., 2001; Wernegreen et al., 1997; Wernegreen & Riley, 1999).

In this work, the only possible evidence of recent lateral symbiotic gene transfer is the case of the L. frutescens strain WSM3310, that had showed nearly identical 16S rRNA and dnaK sequences to M. mediterraneum, M. temperatum and to at least five other Lessertia symbionts (Chapter 2) but its nodA gene matched with 100% similarity to that of Sinorhizobium medicae strain WSM419 (Reeve et al., 2010). Transfer of genes from Sinorhizobium to Mesorhizobium species has previously been demonstrated (van Berkum et al., 2003) and the close phylogenetic proximity between both genera (Jarvis et al., 1997) as well as the fact that plasmids are quite promiscuous makes genetic transfer among them possible (Doolittle & Papke, 2006).

One of the aims of this chapter was to establish the relationship between geographical region of isolation and nodA phylogeny of Lessertia root nodule

84 CHAPTER 3 bacteria. The divergence of nodA clades A to F may be explained by the geographical origin of the symbionts (Figure 3.1 and 3.2), as in most cases strains that grouped in a particular clade were isolated from plants collected from closely related sites or from a common biome. For example, strains from clade A were all collected from Succulent Karoo biome in the Northern Cape, whereas the strains in clade C and F had been isolated from the Nama Karoo and from the Fynbos biomes (Western Cape) respectively. Furthermore, the only nodA sequence from a different host included in the nodA cluster of Lessertia symbionts was that of the Mesorhizobium strain DUS835 (Elliott et al., 2007). This strain was originally isolated from the South African legume Aspalathus linearis (Boone et al., 1999; Muofhe & Dakora, 1999) from the fynbos biome of the Western Cape Province (Joubert et al., 2008). The biome from where this legume was collected matches with the geographic origin of the Lessertia strains included in that clade (WSM3804, 3803, 3805 and 3806), suggesting either that these strains together with DUS835 have evolved together, adapting to a similar legume flora, or that at some point in time there was transfer of symbiotic genes between symbionts.

The clustering of rhizobial symbiotic genes following geographic origin has been reported by other research groups (Gao et al., 2004; Han et al., 2010; Parker et al., 2002). It seems that geographic distribution of the host plant is consistently influencing symbiotic genes phylogenetic structure rather than geographic barriers (Andam et al., 2007; Ba et al., 2002; Han et al., 2010; Ormeño-Orrillo et al., 2006; Parker et al., 2002) or the host range associated with them (Chen et al., 2008; Donate-Correa et al., 2007; Haukka et al., 1998; Jarabo-Lorenzo et al., 2003; Laguerre et al., 2001; Perret et al., 2000; Vinuesa et al., 2005a; Vinuesa et al., 2005c; Wernegreen & Riley, 1999). This supports the hypothesis that the biogeography of symbiotic microorganisms should be similar to that of the host with which it is associated (Martiny et al., 2006). While this may be true when examining plant host and symbiotic genes it is not the case when looking at phylogeny based on 16S rRNA.

Roche et al. (1996) suggested that nodABC genes have evolved to contribute to the adaptation to specific legume hosts. Consequently, symbiotic gene classification is often associated with host specificity (Laguerre et al., 2001; Perret et al., 2000). It is

85 CHAPTER 3 thus likely that based on nodA sequences, symbionts from Lessertia spp. diverged forming distinct clades to adapt to the legumes present within each different biome, rather than by the effect of the geographic distance or environmental conditions.

One of the nodA sequences that was not located in the Lessertia spp. strains nodA clade was that of WSM3602 (L. herbacea), which clustered with nodA from B. tuberum STM678 (Elliott et al., 2007) and Burkholderia sp. WSM3937 (Garau et al., 2009). This is not surprising as WSM3602 had been formerly identified as a Burkholderia spp (Chapter 2). It is notable that STM678 was originally isolated from Aspalathus carnosa, and WSM3937 from Rhynchosia ferulifolia, both native to the fynbos vegetation in South Africa’s Western Cape (Elliott et al., 2007; Garau et al., 2009) which corresponds to the area from where WSM3602 was collected (Figure 3.2), reinforcing the idea that nodA clusters are based on adaptation to a particular biome.

Although nodA is the nod gene with the most available sequences in the data bases, most of these are derived from rhizobia associated with crops or forage legumes and only a small number from non-commercial legumes (Doyle & Luckow, 2003; Hirsch et al., 2001; Moschetti et al., 2005; Willems, 2006). Further studies of root nodule bacteria from legumes in different African habitats may reveal new rhizobia lineages, increasing the chances of the nodA clade of Lessertia symbionts being intermingled with those from other not yet studied hosts.

The genus Mesorhizobium differs from many other root nodule bacteria genera in that its symbiotic genes are located on the chromosome (Sullivan et al., 1995; Sullivan & Ronson, 1998). Some Mesorhizobium spp. carry genes required for nodulation and nitrogen fixation on large transmissible elements called symbiosis islands (Nandasena et al., 2007a; Sullivan & Ronson, 1998). The presence of the symbiosis island insertion region was studied for Lessertia spp. symbionts. From 17 strains, only WSM3495 successfully amplified the region, suggesting that the symbiotic genes of this strain are located on a symbiosis island. The potential location of Lessertia spp. strains symbiotic genes on a symbiosis island represents a threat to the genetic stability of the inoculant in the environment and thus to the establishment of the legume in the field, as symbiotic genes can be easily and rapidly

86 CHAPTER 3 transferred to competitive and ineffective soil mesorhizobia (Finan, 2002; Nandasena et al., 2004; Nandasena et al., 2006; Sullivan et al., 1995; Sullivan et al., 2002).

The second major aim of this chapter was to determine the nitrogen fixing abilities of 17 of Lessertia root nodule bacteria on the six plant host species. Some hosts like L. herbacea, L. excisa and L. diffusa were able to form effective or partially effective symbiosis with a number of strains, while for the others this number was much smaller. All the species apart from L. pauciflora were able to nodulate with many of the same strains and hence might belong to the same cross inoculation group. This further suggests that nodA topology reflects host range (Laguerre et al., 2001; Perret et al., 2000) as all the strains that were able to nodulate the five Lessertia species were clustered together in the nodA phylogram (clades A, B, D and F; Figure 3.1). L. pauciflora, seems not to belong to this cross inoculation group as it could only nodulate with three of the strains tested and only one was partially effective. Further it was not able to nodulate consistently even with the strain isolated from it (WSM2566), so there may be other abiotic factors that are affecting its successful nodulation in the substrate used to grow the plants.

Overall, the highest dry weights in comparison to uninoculated controls were recorded for the species L. herbacea (in association with WSM3565 and 3612) and L. diffusa (with WSM3636 and 3598) (Figure 3.3). None of the strains inoculated was able to achieve more than 50% of the N control dry weight with the species L. incana, L. excisa or L. pauciflora, although some of the strains increased dry weight significantly in comparison to uninoculated control. Some of the most promising strains are WSM3636, WSM3612, WSM3565 and WSM3898 that achieved dry weights as high as in the nitrogen control with at least one of the species and were able to nodulate and fix nitrogen with more than three other Lessertia spp. Having identified effective legume-rhizobia matches, the next step will be to assess these in the field across a range of edaphic conditions.

There were wide differences in effectiveness and in the number of symbiotic partners within Lessertia strains, which is expected for legumes that associate with different rhizobia lineages under natural conditions (Moschetti et al., 2005; Parker & Kennedy, 2006). L. herbacea followed by L. diffusa were able to associate with most of the

87 CHAPTER 3 strains tested. Both species were even able to form nodules with the Mesorhizobium strain WSM3310 that contains nod genes from Sinorhizobium medicae. Moreover, Burkholderia sp. WSM3602 formed ineffective nodules on L. herbacea. These two strains showed wide differences between their nodA sequences and with those of most of the Lessertia spp. strains (Figure 3.1). It is thus apparent that L. herbacea and L. diffusa are able to recognize widely different Nod factors. Future studies might reveal the extent of this apparent symbiotic “promiscuity” and the impacts this could have in the establishment of these species in the field.

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CHAPTER 4. Field adaptation and symbiotic promiscuity

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4.1 Introduction The inclusion of perennial plants, particularly deep-rooted herbaceous legumes, into Western Australian agricultural systems is likely to play a key role in managing salinity as well as contributing to plant biodiversity (Cocks, 2001; Dear et al., 2003; Pannell & Ewing, 2006; Ward, 2006). Perennial plants are also likely to be more resilient to changing weather patterns (Howieson et al., 2008). However, in areas like the Western Australian wheatbelt, where the edaphic stresses of aridity, infertility and acidity are present (Howieson & Ballard, 2004), it is particularly important to introduce legume species originating from similar edaphic conditions (Howieson et al., 2008). Cocks (2001) and Dear et al. (2003) both acknowledge the necessity of developing new agricultural species from regions with similar climate to the target.

The perennial herbaceous legumes of the genus Lessertia were targeted for inclusion in the Western Australian wheatbelt, as they are native to areas of South Africa that are subject to dry periods, high summer temperatures, low soil pH and low soil organic matter (Hoffmann, 1996a; Hoffmann, 1996b; Robelo, 1996), thus matching the edaphic and climatic conditions of the wheatbelt (Howieson & Ballard, 2004).

When introducing a legume into a new environment it is important to include an effective symbiotic partner (Brockwell & Bottomley, 1995; Howieson et al., 2008; Lindström et al., 2010). In the previous chapter strains of Mesorhizobium spp. were assessed for nitrogen fixation effectiveness with Lessertia spp. and those able to increase the biomass of L. diffusa, L. capitata, L. herbacea, L. excisa and L. incana were selected for further studies (section 3.3.2). However, glasshouse results cannot always be extrapolated to the field, as adaptation of a legume and its microsymbiont are subject to diverse soil and environmental factors (Howieson & Ballard, 2004; Zahran, 1999). Moreover, the selected inoculant strain has to be able compete with the native microflora, survive saprophytically in the soil and maintain genetic stability (Herridge, 2008; Howieson et al., 2000b; O'Hara et al., 2002). It is thus essential to assess plant and legume symbiont performance in the environment before selecting the appropriate inoculant and to determine the capacity of strains to survive in and colonize soils (Graham, 2008; O'Hara et al., 2002). This is particularly important

91 CHAPTER 4 considering that the Western Australian wheatbelt is subject to diverse stresses, which are known to be synergistic when combined (Howieson & Ballard, 2004), creating a highly stressful environment for symbiosis.

In this chapter the species L. diffusa, L.capitata, L.herbacea, L. excisa and L. incana were assessed for their potential to establish at five different sites in the Western Australian wheatbelt with varying soil properties, to gauge the effect of edaphic factors upon nodulation. The ability of Lessertia spp. root nodule bacteria to persist and colonize these soils was also evaluated. Finally a pristine soil that had minimal exposure to agricultural influences was assessed for the presence of Lessertia nodulating bacteria.

The phylogenetic studies (sections 2.3.4 and 2.3.5) showed that in its natural environment, L. herbacea forms symbiosis with at least two genera of rhizobia, and with different species within the genus Mesorhizobium. Also, the glasshouse effectiveness experiment revealed that L. herbacea and L. diffusa are able to nodulate with strains showing diverse nodA gene sequences (sections 3.3.1 and 3.3.2). Symbiotic promiscuity can be beneficial when legumes associate effectively with different rhizobia species allowing their adaptation to different environments (Lafay & Burdon, 2006; Mierzwa et al., 2010; Ulrich & Zaspel, 2000), but it can become a serious threat when nodulation is ineffective (Ben Romdhane et al., 2007). Therefore, the assessment of nodulation of Lessertia spp. with inoculants of current commercial application in Western Australia representing seven genera of root nodule bacteria, was also included in this study.

4.2 Materials and Methods 4.2.1 Establishment of Lessertia spp. in the field (June 2007–March 2008) With the aim of assessing plant establishment, adaptation and summer survival, L. capitata, L. diffusa, L. excisa, L. herbacea and L. incana were sown at five sites in the low-rainfall zone of the Western Australian wheat-belt in June 2007. This part of the project was performed in collaboration with the Department of Agriculture and Food, Western Australia (DAFWA).

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The sites were the DAFWA Research Stations at Badgingarra and Newdegate, the Research Station of the Great Southern Agricultural Research Institute in Katanning, LIEBE group long-term research site 20 km west of Buntine and Muresk Institute (Curtin University of Technology) at Northam. The location and main characteristics of each site are detailed in Table 4.1. Monthly rainfall for each site is shown in Appendices 12 to16.

Table 4.1. Experimental site characteristics. Place Badgingarra Buntine Katanning Newdegate Muresk Location 30°20’S 30°00’30” S 33°42’S 33°6’S 31°73’S 115°30’E 116°20’40”E 117°37’E 118°49’E 116°70’E Mean annual 547 357 478 354 429 rainfall (mm) Soil texture Shallow sand Sandy yellow Clayey sand Gravelly Loamy sand over gravel earth (yellow overlying a loamy sand over sandy over clay sandplain) clay subsoil. over clay. clay loam Soil pH 4.9 4.94 4.6 4.72 4.9 (CaCl2)

4.2.1.1 Seed inoculation Lessertia spp. seeds were inoculated with the appropriate inoculant, selected from section 3.3.2. The strains were recovered from glycerol stocks, streaked onto ½ LA plates and incubated at 28°C for five days. Single colonies from each strain were inoculated into conical flasks containing 50 ml of ½ LA broth, and were incubated for

5 days at 28°C on a shaker at 140rpm. After the cultures had reached an OD600 of 1.0 (approximately 109 cells ml-1) , 30 ml of the broth cultures were inoculated into 50g sterile peat bags (Biocare Technology Pty, Ltd. Somersby, NSW, Australia) using a sterile syringe. The peat was mixed and incubated for 10 days at 28°C and then kept at 4°C until used.

Seeds of L. capitata, L. diffusa, L. excisa, L. herbacea and L. incana were surface- sterilised as described in section 2.2.3 and air-dried in a laminar flow cabinet. The surface sterilised seeds (5 g per species) were inoculated with a mixture of 2 mL of autoclaved 1.5% (W/V) Methocel adhesive solution and 0.1g of peat of the correspondent inoculant and left to dry. Inoculated seeds were stored at 4°C overnight and sown the next day.

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4.2.1.2 Experimental design and sowing The same plot designs were used in each of the sites. The experimental design was a randomized Complete Block with 4 replicates. Each plot consisted of a single row of 1 m with 0.5 m spacing between rows and between replicates. The experimental units were:

1. L. capitata inoculated with WSM3636 2. L. diffusa inoculated with WSM3598 3. L. excisa inoculated with WSM3626 4. L. herbacea inoculated with WSM3612 5. L. incana inoculated with WSM3598

The experiments were set up during June, 2007. Each plot was sown with 20 inoculated seeds. Sowing equipment was disinfected or discarded between treatments to avoid cross contamination. During growth, the plants were not watered or fertilized and depended up on natural rainfall.

4.2.1.3 Evaluations The plant population was monitored and recorded every 4 weeks. The number of plants recorded 8 weeks after sowing, was used to calculate the percentage establishment from the number of seeds sown. The plant population present in March 2008 was used to calculate the summer survival, which corresponded to the percentage of plants that survived summer from the established plants

4.2.2. Inoculant survival and soil colonization (August 2008) The saprophytic ability and colonisation of Lessertia spp. inoculant strains were assessed at the DAFWA Research Station at Badgingarra, where Lessertia spp. with their appropriate inoculants had been sown 14 months previously. This site was chosen because there were surviving individuals of each of the sown species and there was enough separation between plants to establish trap hosts at different distances.

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4.2.2.1 Experimental set up This field experiment was established in August 2008. Prior to sowing, seeds of L. herbacea, L. capitata, L. excisa and L. diffusa were scarified, surface sterilised (section 2.2.3) and dried in the laminar flow cabinet for an hour. Six trap plants (2 plants of L. diffusa, 1 L. herbacea, 1 L. capitata, 1 L. excisa and 1 L. incana) were chosen and marked. Trap plants were sown in 30cm rows at three distances (30, 70 and 120cm) from each surviving plant in a semi-radial array as shown in Figure 4.1.

Figure 4.1. Diagram of the sowing design of trap plants surrounding a surviving Lessertia spp. plant. Each x corresponds to a L. capitata plant; x to L. diffusa; x to L. herbacea and x to L. excisa

At sowing, all equipment used in each row was disinfected or replaced, to avoid cross contamination of the soil. A starter fertiliser solution [Poly-feed, Haifa

Chemicals (15% available N; 30% P2O5, 15% K2O)] was supplied to the plants at a rate of 2 L m-2. During the first month the plants were watered with boiled DI water at a weekly basis to ensure their establishment in the field. After the first month, water supplies came exclusively from rain (Appendix 12). Due to the high presence of weeds in the plots, glyphosate (0.5 L ha-1) was applied to suppress weeds nearby. To avoid disturbing the soil no weeds were mechanically removed.

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Trap plants were excavated 8 weeks after sowing. The roots were washed and assessed for the presence of nodules.

4.2.2.2 Estimation of the population of Lessertia nodulating rhizobia in the soil Soil samples were taken from each of the plots sown with Lessertia spp. at Badgingarra 14 months earlier (Section 4.2.1) and from non-inoculated plots. Three sub samples (10 cm deep) were taken from each plot and were kept in polyethylene bags which later constituted one bulk sample per plot. Soil samples were kept in a cooler until used in the laboratory.

In the laboratory, the Most Probable Number (MPN) of rhizobia able to nodulate Lessertia spp. was estimated for each soil sample under controlled glasshouse conditions (Brockwell, 1982). Seeds of L. excisa, L. diffusa, L. herbacea, L. capitata and L. incana were sterilized and pre-germinated as described in section 2.2.3 and were then planted in 150ml pots with a sterile sand mix (2:3 of washed river sand and washed yellow sand).

Soil samples were well mixed. From each sample (corresponding to a plot) 10g of soil was diluted in 90ml of 1% sterile sucrose solution and shaken on a wrist-action shaker for 10 min at 200-300 cycles per minute. This initial suspension was subsequently diluted 6 times in 1% sucrose at a rate of 1:5. From each dilution, including the initial suspension, four 1 ml aliquots were used to inoculate 4 test plants. The test plant species chosen for each sample corresponded to the Lessertia spp. that had been sown on each plot in the field. Plants were inoculated with the dilutions three days after planting and the surface of the pots was then covered with sterile alkathene beads to avoid aerial contamination. Plants were supplied weekly with 10 ml of nutrient solution devoid of nitrogen (section 2.2.3), and watered with sterile DI water every 2 days. Plants were assessed for presence of nodules 8 weeks after inoculation.

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4.2.3 Recovering rhizobia from five different sites in the Wheatbelt (October 2008) Soil samples were taken from the five sites at which Lessertia spp. had been sown in year 2007, to set up a soil trapping experiment under glasshouse conditions. The sites were Badgingarra, Katanning, Muresk, Buntine and Newdegate (Table 4.1). The main purpose of this experiment was to assess whether plant establishment in the different areas of the wheat belt was affected by poor survival of their inoculants and to evaluate presence of soil rhizobia able to nodulate Lessertia.

4.2.3.1 Soil collection Three soil samples were taken from each plot previously sown with Lessertia spp. and subsequently bulked. If plants were still present the sample was taken at 10 cm depth and within a 20 cm radius from the crown of the plant. If there were no surviving plants, three samples were taken from the row where plants had been originally sown. Additionally, soil samples were taken from Medicago sativa subsoil to be used as control samples to ensure that handling did not affect the rhizobial population. M. sativa had been established at every site (except Badgingarra) as a buffer and when excavated the plants were all nodulated. Samples were also taken from un-inoculated soil and from a pristine area at the Jandawanning National Park (30º34’35.90’’S; 115º27’30.85’’E). All the samples were stored in plastic bags and placed in a cool box for transporting and once in the laboratory were stored at 4°C.

4.2.3.2 Trap plant experiment set-up Each of the soil samples was used to establish a rhizobial trapping experiment in the glasshouse. Free draining plastic pots (500ml) were filled to one third of their capacity with steamed sand mix (2 parts river sand: 3 parts leached yellow sand). In parallel, sand mix was autoclaved in plastic containers, to be used to cover each of the pots. After steam sterilization of the sand mix, 200 g of the field soil were layered on top, and later covered with the autoclaved sand mix as shown in figure 4.2. The layering of the soil sample between sterilized sand mix was established to avoid fungal diseases in the earlier stages of development of the plantlets, as from the field it was observed that this was one of the common causes of plant death.

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A

B

A

Figure 4.2. Rhizobia trapping system. Bottom and top layers correspond to sterilized sand mix (A). The middle layer (B) corresponds to 200 g of soil collected from trial sites.

Seeds of L. excisa, L. diffusa, L. herbacea, L. capitata and L. incana were surface sterilized and germinated as described in section 2.2.3. The pots were watered with sterile DI water and planted with the host that corresponded to the species sown in the plot from where the soil sample was taken. Following radicle emergence, two seedlings were planted per pot. Once plants were approximately 4cm in height the soil surface was covered with sterile alkathene beads and a PVC watering tube was installed in each pot.

Plants were watered every 48 hours and 20 ml of nutrient solution devoid of nitrogen was provided every week (section 2.2.3). Control pots were provided for each of the species and consisted of sand mix only as a substrate.

4.2.3.3 Harvest and evaluations At harvest, plants were carefully removed from the pots. The roots were washed in tap water and were assessed for the presence of nodules. Where nodules were present, they were kept in microcentrifuge tubes at 4°C for a maximum of four days. The nodules were surface sterilized and the symbionts were re-isolated (section 2.2.3). Pure cultures of the isolates were re-suspended in saline solution (0.89% w v-1

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NaCl) and placed in 2.0mL cryo-vials containing 30% (v v-1) glycerol for long-term storage at -80°C.

4.2.4 Establishment of Lessertia spp. in the field (July 2008) In July 2008, the species L. capitata, L. diffusa, L. excisa, L. herbacea and L. incana were established at Buntine and Newdegate (Table 4.1). These experiments were essentially a repetition of those established in 2007. The main purpose was to assess establishment and nodulation in situ of Lessertia spp. and to discard the possible effect that the low rainfall conditions in 2007 may have had on Lessertia spp. establishment that year.

4.2.4.1 Experimental design and sowing The plots were established in a randomized complete block design with 4 replicates. Each plot consisted of five, 5 m long rows (10 m2). The seeding rate was 20 seeds per m. The strains and the seed inoculation methods were as for season 2007 (section 4.2.1).

4.2.4.2 Evaluations The plant population was recorded at weeks 6, 8 and 12 after sowing. Plant samples were taken in the 12th week to assess nodulation. Ten plants (if present) were excavated from each plot. The root system was washed and the nodules were carefully removed and stored in microcentrifuge tubes with silica gel. In the laboratory, the nodules were re-hydrated and surface sterilized and the bacteria re- isolated (section 2.2.3). The isolates were fingerprinted with the primer RPO1 and the banding patterns were analysed and compared with the inoculant bacteria as described in section 2.2.2. Pure cultures of the isolates were stored at -80°C in 30% glycerol as described in 4.2.3.3.

4.2.5 Molecular Fingerprinting The isolates obtained from the soil trapping experiment (section 4.2.3) and those isolated from field plants during Lessertia spp. establishment in 2008 (section 4.2.4) were RPO1-PCR fingerprinted (section 2.2.2). Banding patterns were analyzed using the software BIOCapt (ALYS Technologies, Lausanne) and compared with those of the original inoculants by performing UPGMA cluster analyses using the NEIGHBOR

99 CHAPTER 4 application from the PHYLIP software package. A cladogram was constructed in MEGA4 (Tamura et al., 2007), to select strains with unique banding patterns.

4.2.6 Authentication and assessment of effectiveness of field strains In a glasshouse experiment, strains collected from soils (4.2.3) and from the field (4.2.4) were authenticated and assessed for nitrogen fixation effectiveness with four Lessertia species (L. capitata, L. diffusa, L.incana and L. herbacea).

4.2.6.1 Experimental design The experiment was a split-pot design with one rhizobial strain per pot as the main treatment, two Lessertia spp. as sub-treatments and four replicates per treatment. Treatments consisted of 8 strains collected from soils and field (N1-N8) and an effective Mesorhizobium strain (WSM3636). Two sets of uninoculated controls, with added nitrogen (N+) and nitrogen free (N-), were included for all species.

4.2.6.2 Experimental preparation, planting and inoculation Seeds were scarified, surface sterilised and germinated as described in section 2.2.3. The seedlings were planted in 500 ml free-draining plastic pots filled with steamed sand mix (section 3.2.2).

The rhizobial strains were grown on ½ LA plates for 3 to 8 days at 28°C. Pot preparation and inoculation was performed as in 3.2.2.

Plants were supplied weekly with a nutrient solution devoid of nitrogen (section 2.2.3) at a rate of 20 ml per pot. The nitrogen-fed control pots were additionally supplied with 10 ml of 0.25M KNO3 solution on a weekly basis. Plants were watered every two days with 20 ml of sterile DI water.

4.2.6.3 Harvest and evaluations Plants were harvested after 8 weeks, by carefully removing them from the pots and washing the roots. Nodulation was assessed and shoot tops were dried at 60°C for 48 hours and weighed. Two nodules were collected from each of the replicates and the bacteria were re-isolated to verify strain identity through RPO1-PCR fingerprinting (sections 2.2.2 and 2.2.3).

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4.2.7 Sequencing of the genes dnaK and nodA for soil trapped strains Eight soil trapped and field isolated strains (N1-N8) from experiments 4.2.3 and 4.2.4 were identified to the species level through the sequencing of the dnak gene. The primers used were TSdnak2 and TS dnak3 (Stępkowski et al., 2003a).

The nodulation gene nodA was sequenced for the eight strains to investigate if there had been transference of symbiotic genes from the original inoculant to soil rhizobia. The primers used were nodA1 and nodA2 (Table 3.1, Chapter 3)

Cell preparation was performed as described in section 2.2.1 but the cells were concentrated to an OD600 of 3.0. Cycling conditions and gene amplification procedures for both genes are described in 2.2.5 and 3.2.1. dnaK and nodA sequence assembly was carried out using Gene Tool Lite 1.0 (2000) software (Doubletwist, Inc., Oakland, CA, USA). Phylogenetic analyses and phylogram construction were conducted using MEGA4 (Tamura et al., 2007) (section 2.2.7).

4.2.8 Nodulation of Lessertia spp. by commercial inoculants. The ability of L. diffusa, L. capitata, L. herbacea and L. excisa to nodulate with 17 inoculants of current use in Western Australia was assessed. The inoculants used for this experiment are listed in table 4.2. Although the strains from Lebeckia sp. (WSM3937) and Rhynchosia sp. (WSM4187) are not commercial inoculants, they were included in the experiment as these plants have been introduced into Western Australia with the same purpose as Lessertia spp. and are likely to be sown in the same agro-ecological areas.

The inoculant strains were recovered from glycerol stocks and were grown on ½ LA plates. The cultures were incubated at 28°C for 3 to 7 days. Seeds of the Lessertia spp. were sterilized and scarified as described in section 2.2.3.

The plants were grown in a closed 250ml vial system with 125ml of sand substrate (section 2.2.3). Each vial contained two Lessertia species. Initially, two seedlings of

101 CHAPTER 4 each species were transplanted into the vial and after three weeks were thinned to one plant per species. After planting the seedlings, 10 ml of sterile nutrient solution devoid of nitrogen and 20 ml of sterile deionized water were added to each vial.

Three days after planting each plant was inoculated with 1 ml (OD600nm = 1.0) of a cell suspension obtained from fresh colonies and suspended in 1% (w/v) sucrose solution. There were three replicates for each of the inoculants, and an uninoculated control was provided.

This experiment was carried out under axenic conditions in a temperature controlled phytotron at 22°C for eight weeks. At harvest, plants were carefully removed and washed with tap water. Nodulation was assessed and nodule characteristics were recorded. To verify strain identity, the bacteria were re-isolated and RPO1-PCR fingerprinted (sections 2.2.1 and 2.2.2).

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Table 4.2. Legume rhizobial inoculants under current use in Western Australia and evaluated in this study

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4.3 Results 4.3.1 Establishment of Lessertia spp. in the field (June 2007–March 2008) The percentage of established plants varied between species and among sites. L. diffusa was present in greater proportion 8 weeks after sowing at Buntine and Muresk, with 80% and 47.5% of plants established; followed by L. capitata with 35% and 25% of plants established at the same sites respectively (Figure 4.3). The species L. excisa, L. incana and L. herbacea performed poorly at every site, achieving less than 20% establishment. Overall, Muresk and Buntine were the sites that supported better establishment. Katanning, Newdegate and Badgingarra supported less than 10% of plants established during the winter for every species.

Figure 4.3. Percentage of plants of L. capitata, L. diffusa, L. excisa, L. herbacea and L. incana successfully established 8 weeks after sowing at 5 sites in the Western Australian wheat belt: Muresk, Badgingarra, Buntine, Katanning and Newdegate (100%= 20 plants m-2). Vertical bars correspond to standard error of means.

Muresk supported the best over-summer survival (Figure 4.4), especially for L. capitata, L. diffusa and L. excisa, where 81.2, 86.7 and 87.5% of the established plants persisted. Few individuals of the poorly established species L. incana and L. herbacea managed to survive after summer, and L. incana showed weak re-growth (Figure 4.5). The summer survival at Badgingarra, Newdegate and Katanning was very poor, due to a combination of drought and soil-borne diseases.

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100 L. capitata L. diffusa 80 L. excisa L herbacea 60 L. incana

40

summer (%) survival 20

0 Muresk Badgingarra Buntine Katanning Newdegate

Figure 4.4. Percentage of established plants of L. capitata, L. diffusa, L. excisa, L. herbacea and L. incana that survived after summer at 5 sites in the Western Australian wheat belt: Muresk, Badgingarra, Buntine, Katanning and Newdegate. Vertical bars correspond to standard error of means.

Figure 4.5. L. incana re-sprouting after summer at Muresk (May 2008).

4.3.2 Inoculant survival and soil colonization 4.3.2.1 In situ soil trapping experiment (August – October, 2008) The number of trap plants successfully established after 10 weeks in this experiment was very low, probably due to the small amount of rainfall during the second month after sowing (Appendix 12). The percentage of established plants was 31.7% for L. excisa, 18.3% for L. diffusa, 14.4% for L. capitata and 0 % for L. herbacea. Plants of all species present looked stressed, were small and showed symptoms of nitrogen deficiency. None of the trap plants presented nodules on their roots after 8 weeks, suggesting that the inoculant strains were unable to survive in the soil or to colonize

105 CHAPTER 4 it, even in the presence of the host plant. There were a high number of plants that appeared to be affected by hypocotyl rot, particularly within the species L. excisa.

4.3.2.2 Glasshouse-based MPN estimation of the population of Lessertia-nodulating rhizobia in the soil No nodules were detected on plants inoculated with any of the soil samples indicating that the population of soil rhizobia able to nodulate Lessertia spp. in Badgingarra was lower than the detectable limit of 1 cell g-1 of soil. A hypocotyl fungal disease affected some of the L. capitata and L. diffusa plants inoculated with the two highest soil concentrations. L. excisa was highly susceptible to the hypocotyl rot (Figure 4.6), which affected every plant inoculated with the four highest soil concentrations.

Figure 4.6. Plants of L. excisa affected by hypocotyl rot.

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4.3.3 Recovering rhizobia from five different sites in the Wheatbelt There was very little impact on nodulation of Lessertia spp. according to whether the soil was collected from a previously inoculated plot or from the uninoculated sites (Table 4.3). This indicated that the plants were nodulating either with indigenous soil bacteria rather than the inoculant, or that inoculants had strongly colonized the region. The exception was L. capitata in the Newdegate soil sample, which was able to nodulate only with inoculated soil and not with the uninoculated sample. L. herbacea formed nodules when growing in every soil except in those from Badgingarra. L. excisa failed to nodulate in soils from any site, and nodulation by L. capitata and L. incana was rare. None of the species formed nodules in the pristine soil from the Jandawanning National Park. M. sativa (lucerne) nodulated with every soil sample taken from M. sativa plots. Soil samples from Badgingarra were not assessed for nodulation with lucerne, as this species was not present at that site.

Table 4.3. Nodulation of Lessertia spp. with rhizobia recovered from inoculated and uninoculated plots from different sites in the Western Australian wheat-belt Origin of soil sample

HOST Muresk Buntine Katanning Badgingarra Newdegate Pristine soil

a* b a b a b a b a b

L. diffusa + + - - + + - - + + -

L. herbacea + + + + + + - - + + -

L. excisa ------

L. capitata - - - - + + - - + - -

L. incana - - + + ------

M. sativa + + + +

*a: Inoculated plot; b: uninoculated soil; +: nodulated plant; -: no nodules

Most nodule isolates grew faster than the original inoculants, taking only 2- 3 days to form 1 mm colonies (Table 4.4). The only exceptions were two isolates from L. capitata inoculated with Newdegate soils, which formed 1mm colonies in 6 days.

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Colony growth had been previously assessed for the inoculants and on average they took between 5-7 days to form 1mm colonies (Chapter 2, section 2.3.2).

Table 4.4. Rhizobia isolates from Lessertia spp. and Medicago sativa potted in soils from Buntine, Katanning, Muresk and Newdegate (Western Australia) Isolate Host Site Growth Rate* 1 L. herbacea Buntine Fast 2 L. herbacea Buntine Fast 3 L. herbacea Buntine Fast 4 L. incana Buntine Fast 5 L. incana Buntine Fast 6 M. sativa Buntine Fast 7 L. capitata Katanning Fast 8 L. capitata Katanning Fast 9 L. diffusa Katanning Fast 10 L. diffusa Katanning Fast 11 L. diffusa Katanning Fast 12 L. diffusa Muresk Fast 13 L. diffusa Muresk Fast 14 L. diffusa Muresk Fast 15 L. diffusa Muresk Fast 16 L. herbacea Muresk Fast 17 L. herbacea Muresk Fast 18 M. sativa Muresk Medium-slow 19 L. capitata Newdegate Medium-slow 20 L. capitata Newdegate Medium-slow 21 L. herbacea Newdegate Fast 22 L. herbacea Newdegate Fast 23 L. herbacea Newdegate Fast 24 M. sativa Newdegate Medium-slow 25 M. sativa Newdegate Medium-slow *Fast: Forms 1mm colony within 2-4 days; Medium-slow: Forms 1mm colony within 5-7 days.

4.3.4 Establishment of Lessertia spp. in the field (Year 2008) Overall, plant populations were low at Buntine and Newdegate (Figure 4.7). There was an improvement in plant establishment at the Newdegate experimental site compared to that achieved in 2007. At Buntine, though, the populations were not improved and in some cases, such as for L. diffusa and L. capitata the number of plants were even lower. L. diffusa achieved the highest number of established plants at both sites: 50% at Newdegate and 48% at Buntine, followed by L. excisa with 38% (Newdegate) and 25% (Buntine). L. capitata showed 18% and 22% of plant establishment at Newdegate and Buntine respectively. Only 10% of L. herbacea plants established at both sites.

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Figure 4.7. Percentage of plants of L. capitata, L. diffusa, L. excisa and L. herbacea successfully established 8 weeks after sowing at two sites in the Western Australian wheat belt: Buntine and Newdegate.

The percentage of plants nodulated at these two sites was low. Only 33.3% of L. diffusa, 24.5% of L. excisa, 24% of L. herbacea and 19.5% of L. capitata plants were nodulated. Most of the nodules appeared desiccated and, when taken to the laboratory, it was not possible to isolate bacteria from them. In total, two isolates from the species L. capitata, two from L. diffusa and six from L. herbacea were obtained (Table 4.5).

Table 4.5. Rhizobia isolates from L. herbacea, L. capitata and L. diffusa growing in the field at Buntine and Newdegate. Isolate Host Site Growth Rate* 26 L. capitata Buntine Medium-slow 27 L. capitata Buntine Medium-slow 28 L. diffusa Buntine Fast 29 L. diffusa Buntine Fast 30 L. herbacea Buntine Fast 31 L. herbacea Buntine Fast 32 L. herbacea Buntine Fast 33 L. herbacea Newdegate Fast 34 L. herbacea Newdegate Fast 35 L. herbacea Newdegate Fast

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4.3.5 Molecular Fingerprinting All the isolates from the soil trapping experiment (Table 4.4) and from the established Lessertia spp. at Buntine and Newdegate (Table 4.5) were successfully fingerprinted using RPO1-PCR. The cladogram (Figure 4.8) shows that the RPO1 fingerprints of only four isolates from L. capitata [19 and 20 (Trapped from Newdegate soil) and 26 and 27 (Isolated from field plants at Buntine)] and three from L. herbacea from Newdegate (Isolates 33, 34 and 35) matched with original inoculant fingerprints (WSM3636 and WSM3612 respectively) (Figure 4.8).

The RPO1 fingerprint for most of the trapped isolates (in blue) and field isolates (in red) did not correspond to the original inoculants. The fingerprints of the isolates from M. sativa from every site, matched with that of the inoculant [S. meliloti (RR1128)] used for that species (Figure 4.8).

Two isolates trapped by L. incana from Buntine soils (4 and 5) and five isolates obtained from field plants of L. diffusa (28 and 29) and L. herbacea (30, 31 and 32) at Buntine showed identical banding patterns. These will be hereafter termed N1. The RPO1 banding pattern from isolates trapped by L. diffusa from Katanning (Isolate 9) and Muresk (Isolate 13) and two trapped by L. herbacea from Newdegate soil (22 and 23) clustered together representing strain N2. The patterns from four isolates from L. diffusa potted in soils from Muresk (12 and 14) and Katanning (10 and 11), and two trapped by L. capitata (7 and 8) were identical and are grouped as N3. Two isolates trapped by L. herbacea from soils from Buntine (1 and 2) and two from Muresk (15 and 16) were named N5 and N6 respectively. Three isolates with an unique banding pattern nodulated L. herbacea when growing in soils from Buntine (3), Newdegate (21) and Muresk (17). These isolates will hereafter be N4, N7 and N8 respectively (Figure 4.8).

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Isolate 33 (Lh) Isolate 34 (Lh) Isolate 35 (Lh) WSM3612 WSM3626 Isolate 30 (Lh) Isolate 31 (Lh) Isolate 32 (Lh) Isolate 28 (Ld) N1 Isolate 29 (Ld) Isolate 4 (Li) Isolate 5 (Li) Isolate 9 (Ld) Isolate 13 (Ld) N2 Isolate 22 (Lh) Isolate 23 (Lh) Isolate 12 (Ld) Isolate 14 (Ld) Isolate 7 (Lc) N3 Isolate 8 (Lc) Isolate 10 (Ld) Isolate 11 (Ld) Isolate 3 (Lh) N4 Isolate 1 (Lh) N5 Isolate 2 (Lh) Isolate 15 (Ld) N6 Isolate 16 (Lh) Isolate 21 (Lh) N7 Isolate 24 (Ms) Isolate 25 (Ms) Isolate 6 (Ms) Isolate 18 (Ms) RR1128 Isolate 17 (Lh) N8 Isolate 26 (Lc) Isolate 27 (Lc) Isolate 19 (Lc) Isolate 20 (Lc) WSM3636 WSM3598

0.8 0.6 0.4 0.2 0.0

Figure 4.8. Cladogram showing RPO1 fingerprints and relationships between soil-trapped isolates (in blue) and rhizobia re-isolated from field plants (in red) from different sites along the Western Australian wheatbelt with Lessertia spp. original inoculants are shown in black. Host initials are indicated in parenthesis: L. capitata (Lc); L. diffusa (Ld); L. herbacea (Lh); L. incana (Li) and M. sativa (Ms).

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4.3.6 Authentication and assessment of effectiveness of field strains Lessertia spp. showed variation in their ability to nodulate with soil-rhizobia collected in sections 4.3.3 and 4.3.4 (Table 4.6). L. herbacea was able to nodulate with every inoculant and L. diffusa with all but one (N6). In contrast L. capitata and L. incana formed nodules with two (N3 and N4) and one (N5) of the field strains respectively. L. excisa was not able to nodulate with any of the soil and field trapped strains. All the Lessertia spp. nodulated with the Mesorhizobium strain WSM3636, forming pink nodules in every case. Plants nodulated with the soil and field trapped strains always produced high numbers (>20) of white nodules.

Table 4.6. Nodulation compatibilities of Lessertia spp. with soil rhizobia (N1-N8) and a Mesorhizobium inoculant (WSM3636) Strains L. capitata L. diffusa L. incana L. herbacea L. excisa N1 - + - + - N2 - + - + - N3 + + - + - N4 + + - + - N5 - + + + - N6 - - - + - N7 - + - + - N8 - + - + - WSM3636 + + + + + +: nodulated plant; -: no nodules

No increase in shoot dry weights was observed for any of the Lessertia spp. nodulating with soil trapped strains (Figure 4.9). Dry weights in every case were similar to those of the uninoculated N- control. The only strain that increased shoot dry weights was WSM3636 for the species L. capitata, L. diffusa, L. excisa and L. herbacea. The highest dry weights were achieved in the nitrogen control.

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Figure 4.9. Shoot tops dry weights for L. capitata, L. diffusa, L. herbacea and L. incana, inoculated with 8 soil trapped strains (N1-N8), and an effective Mesorhizobium strain (WSM3636).

4.3.7 Sequencing of the genes dnaK and nodA for soil trapped strains A 320bp dnaK fragment was amplified for the eight strains recovered from soil. All clustered close to the species Rhizobium leguminosarum bv. trifolii and R. leguminosarum bv. viciae and separate from the Lessertia inoculants (WSM3612, 3636, 3598 and 3626). These latter clustered in the Mesorhizobium clade (Figure 4.10). Strain N4 showed identical dnaK sequence to that of strain WSM1325 of R. leguminosarum bv. trifolii, and most of the other strains (N1, N2, N3, N6, N7 and N8) were in the neighbouring clade. The strain N5 clustered apart from other field strains, but the closest neighbour was still R. leguminosarum.

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Figure 4.10. Neighbor joining phylogenetic tree based on partial dnaK gene sequencing of 8 strains (N1 to N8) nodulating Lessertia spp. The strain sequences in the phylogram were obtained from GenBank (accession number in parentheses). M.: Mesorhizobium; S.: Sinorhizobium; R.: Rhizobium and B.: Bradyrhizobium.

A 600bp long nodA sequence was successfully amplified for all the strains. The nodA phylogenetic tree for this gene (Figure 4.11) shows that the nodA sequences of all the trapped strains (N1-N8) clustered with Rhizobium leguminosarum bv. trifolii (XO3721) nodA sequences, showing high sequence similarities between them and a high bootstrap support for the clade (86%).

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Figure 4.11. Neighbour joining tree based on partial nodA sequencing of 8 strains (N1 to N8) nodulating Lessertia spp. The strain sequences in the phylogram were obtained from GenBank (accession number in parentheses). M.: Mesorhizobium; S.: Sinorhizobium; R.: Rhizobium and B.: Burkholderia.

4.3.8 Nodulation of Lessertia spp. by commercial inoculants. There were clear differences in promiscuity between the different Lessertia species (Table 4.7). L. excisa was not able to nodulate with any of the commercial inoculants of current use in Western Australia. L. incana nodulated with the strains TA1 and WSM1325, both being clover inoculants.

L. capitata and L. diffusa had similar nodulation patterns as they were both nodulated by the Sinorhizobium strains RR1128 and WSM1115 and with Rhizobium leguminosarum bv trifolii strains TA1 and WSM1325.

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Table 4.7. Nodulation of Lessertia spp. by commercial rhizobial inoculants.

Strain Species L.capitata L.diffusa L.excisa L.herbacea L. incana RR1128 Sinorhizobium meliloti + + - + - WSM1115 Sinorhizobium medicae + + - + - TA1 R. leguminosarum bv. trifolii + + - + + WSM1325 R. leguminosarum bv. trifolii - + - + + SU303 R. leguminosarum bv. viciae - - - + - WSM1455 R. leguminosarum bv. viciae - - - + - CC829 Mesorhizobium loti - - - - - WSM1497 M. ciceri bv. biserrulae - - - + - WSM1284 M. ciceri bv. biserrulae - - - + - CC1192 Mesorhizobium ciceri - - - + - WSM471 Bradyrhizobium sp. - - - - - WU425 Bradyrhizobium sp. - - - - - WSM2598 Methylobacterium sp. - - - - - WSM3693 Microvirga sp. - - - + - WSM3557 Microvirga sp. - - - + - WSM3937 Burkholderia sp. - - - - - WSM4187 Burkholderia sp. - - - - - +: All plants nodulated, -: No plants nodulated

Surprisingly, L. herbacea was able to nodulate with 13 of the 17 inoculants tested. It was able to nodulate with every Sinorhizobium spp. and Rhizobium leguminosarum inoculant. It formed nodules with most of the Mesorhizobium spp. strains except for Mesorhizobium loti strain CC829 whose original host is Lotus pedunculatus. None of the Bradyrhizobium sp, Methylobacterium sp and Burkholderia sp. strains were able to nodulate L. herbacea nor any of the other Lessertia spp.

In every case, nodules were indeterminate, white and apparently ineffective. There were some differences in nodule morphology (Figure 4.12). The strains RR1128 (S. meliloti), TA1 and WSM1325 (R. leguminosarum bv. trifolii) and strains WSM1497 and WSM1284 (M. ciceri bv. biserrulae) formed elongated nodules, whereas strains WSM1115 (S. medicae) and CC1192 (M. ciceri) formed fan shaped nodules.

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Figure 4.12. Nodules on L. herbacea inoculated with (a) Sinorhizobium meliloti strain RR1128; (b) S. medicae strain WSM1115; (c) Rhizobium leguminosarum bv. trifolii strain TA1; (d) Mesorhizobium ciceri bv. biserrulae strain WSM1497; (e) M. ciceri strain CC1192; (f) Microvirga sp. strain WSM3693. Bars: 200µm.

The identity of the inoculant strains was verified through RPO1-PCR fingerprints in most of the cases. The only case were isolation was not possible was from L. herbacea nodulating with the Lotononis spp. strains WSM3693 and WSM3557, which formed pseudo-nodules (Figure 4.12-f).

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4.4 Discussion The success (or otherwise) of the legume-rhizobia symbiosis can have a major impact on production, plant density and the contribution of fixed N to the farming system of interest (Ballard et al., 2003). One of the limiting factors in the introduction of new legumes is to find symbiotic partners i.e. legume and rhizobia, well adapted to the new edaphic and climatic conditions (Howieson et al., 2008; O'Hara et al., 2002). The first aim of this work was to assess the establishment of the species L. herbacea, L. diffusa, L. capitata, L. incana and L. excisa in low rainfall areas of the Western Australian wheat belt. This was attempted in years 2007 and 2008 and in both seasons there was poor plant establishment and poor over-summer survival. This is largely attributable to the formation of a sub-optimal symbiosis.

The Lessertia spp. were sown at five sites, differing mainly in soil texture and structure, in the Western Australian wheat belt (Table 4.1). In general, species showed the lowest rates of establishment at Katanning, on a poorly draining soil with low nutrient availability (McKenzie et al., 2004), followed by the sites of Newdegate and Badgingarra, that had clayey subsoils which limited water percolation. In soils with high permeability and without physical limitations for root and plant growth such as in Muresk and Buntine (McKenzie et al., 2004), Lessertia spp. produced better establishment. L. diffusa achieved 50 and 80% establishment respectively at these sites. These results suggest a preference of this legume for permeable sands over duplex and clayey soils, which are similar to the well-drained soils present on their natural environment in South Africa (Hoffmann, 1996b; Robelo, 1996).

The summer survival was higher than 80% for L. capitata, L. diffusa and L. excisa at Muresk. For all other sites and species, the number of established plants that survived over the summer was highly variable. It is unlikely that the high temperatures in summer had an important effect in the poor adaptation and survival of Lessertia spp., as temperatures in the wheat-belt are similar or lower to those of their original environment in South Africa (Hoffmann, 1996b; Robelo, 1996).

One of the most likely causes for low plant establishment in the sites was the low numbers of Lessertia rhizobia detected in soils. The inoculant strains for Lessertia

118 CHAPTER 4 were undetectable through the MPN method and through trap plants established in situ (in Badgingarra), and from all sites only one of the inoculants was recovered from soil traps (WSM3636). Moreover, there were few active nodules in the roots of plants excavated only eight weeks after sowing and not all of the rhizobia isolated from these corresponded to the inoculant.

Abiotic stresses like soil acidity, aridity, low clay content and high temperatures are common in large areas of southern Australia (Dear & Ewing, 2008; Howieson & Ballard, 2004) and in the areas where Lessertia spp. were sown. These stressful edaphic and climatic conditions to which plants and symbionts were subject may explain in part the overall low plant establishment, low survival and poor nodulation.

At all the experimental sites soil pH was 5.0 or lower. Low pH is known to limit plant growth and life span (Cocks, 2001; Dear & Ewing, 2008; Hill, 1996) and is considered one of the primary stresses for legume symbiosis (Howieson & Ballard, 2004) by limiting rhizobial survival, competitive ability and persistence in soils and by reducing nodulation (Ballard et al., 2003; Dilworth et al., 2001; Graham, 2008; Slattery et al., 2004; Zahran, 1999). However, the species that was collected from areas with the lowest soil pH in Southern Africa, L. herbacea (Appendix 5), had the lowest survival at every site, suggesting that soil pH was not the sole cause of the low plant populations.

Low soil moisture can also be a limiting factor for both plant and microbial symbionts (Zahran, 1999). The annual rainfall at the experimental sites ranged from 350 to 550mm, which was very similar to the rainfall patterns in the collection points of L. diffusa, L. incana and L. capitata. Therefore for these three species low soil moisture should have had little impact. Perhaps for L. herbacea and L. excisa low rainfall could be one of the causes of their poor establishment, as these species were collected exclusively from the Fynbos biome, which has slightly higher rainfall levels than the target environments in Australia, reaching 600mm (Robelo, 1996).

Given that other strains of the genus Mesorhizobium have been successfully introduced and able to proliferate and adapt to stressful soils in Western Australia (Howieson & Ballard, 2004; Loi et al., 2005) it was expected that the Mesorhizobium

119 CHAPTER 4 inoculants for Lessertia spp. would be able to survive and colonize these soils. The soils in the trial sites did not originally contain high levels of Mesorhizobium (Howieson et al., 2008) so the observed nodulation phenomenon was unlikely to be a result of inter-strain competition. Despite this, very low levels of the inoculants were detected in the soils collected from the trial sites. Moreover, in soil trapping experiments from these sites, L. capitata, L. diffusa, L. herbacea and L. incana formed nodules with R. leguminosarum and not the inoculant. It is unlikely that a transfer of symbiotic genes occurred between the Mesorhizobium inoculants and the R. leguminosarum recovered from field trials and soil traps (as confirmed by nodA sequences).

It is not unusual to find legume species nodulating with different symbionts when growing in a different region (Aouani et al., 2001; Graham, 2008; Han et al., 2009; Han et al., 2005; Liu et al., 2005). It was quite surprising though, that Lessertia spp. formed symbiosis with strains from a genus different to Mesorhizobium, as competition for nodulation is most commonly intraspecific (Howieson & Ballard, 2004). It is even more intriguing that it nodulated with R. leguminosarum a species that has traditionally been thought to be specific for clovers (Broughton & Perret, 1999; Hirsch et al., 2001; Pueppke & Broughton, 1999) and that is not known for nodulating other legume species (Perret et al., 2000; Trinick et al., 1991; Weir, 2006).

The isolation of R. leguminosarum from Lessertia nodules led to the study of the symbiotic interaction of Lessertia spp. with other inoculants under current use in Western Australia (Table 4.2), to which these legumes might be exposed once introduced into the field. Results showed that the species L. capitata, L. diffusa and L. herbacea were able to nodulate with Sinorhizobium meliloti (RR1128), S. medicae (WSM1115) and R. leguminosarum bv. trifolii (TA1 and WSM1325). L. herbacea also formed nodules with the inoculants for Biserrula pelecinus: M. ciceri bv. biserrulae (WSM1497 and WSM1284), inoculant for chickpeas: M. ciceri (CC1192), with R. leguminosarum bv. viciae (SU303 and WSM1455) and formed pseudo-nodules with two Microvirga sp. strains (WSM3693 and 3557), suggesting that this species is rather promiscuous. None of the Lessertia species formed nodules with Mesorhizobium loti, which is reported as able to form nodules on several Lotus species (Sullivan & Ronson, 1998) nor with the Burkholderia strains originating from

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South African Lebeckia sp. and Rhynchosia sp.. Nodulation with the Burkholderia sp. strain WSM3937 was expected; as one of the L. herbacea strains (WSM3602) nodA had matched with the nodA corresponding to that strain (Chapter 3, Section 3.3.1) and nodA genes are known to correlate with host range (Laguerre et al., 2001; Perret et al., 2000; Roche et al., 1996). This might be explained by the fact that nodA is one of the common genes that code for the NodFactor core, which can be modified by various chemical groups which also confer host specificity (López-Lara et al., 1996; Perret et al., 2000).

It is notable that L. diffusa, L. capitata and L. herbacea can nodulate with taxonomically and symbiotically different rhizobia not known for their broad host range. S. meliloti is compatible only with species of the genera Medicago, Melilotus and Trigonella (Gibson et al., 2008), M. ciceri is highly specific to Cicer arietinum (Nour et al., 1994; Slattery et al., 2004). R. leguminosarum bv. trifolii is generally thought to be specific to the legume genus Trifolium (Broughton & Perret, 1999; Hirsch et al., 2001; Pueppke & Broughton, 1999) and bv. vicieae to the genera Vicia, Pisum, Lathyrus and Lens (López-Lara et al., 1996).

There are various factors that contribute to host specificity (Perret et al., 2000), from which the amount and structure of the Nod Factor molecule are predominant (Gibson et al., 2008; Hirsch et al., 2001). The rhizobia species that were able to interact with L. capitata, L. diffusa and L. herbacea produce different Nod factor structures (Oldroyd & Downie, 2008; Perret et al., 2000). Although some studies report that there is no strict correlation between Nod Factor and the plants they nodulate (Perret et al., 2000), the length and degree of saturation of the fatty acid group of the Nod factor can have a function in defining specificity of interaction between host and bacteria and regulate nodule initiation and morphogenesis (Gibson et al., 2008; Oldroyd & Downie, 2008; Sessitsch et al., 2002). The results in this study suggest that L. diffusa, L. capitata and L. herbacea do not require specific Nod Factor structures to begin the process of nodulation.

Nod factor signalling is important at various stages of rhizobial infection (Hirsch et al., 2001; Oldroyd & Downie, 2008; Perret et al., 2000). However, the stringency of Nod factor perception is differentially important at each stage: The induction of root hair

121 CHAPTER 4 curling and the formation of nodule primordia have a lower stringency for the Nod factor structure than is required for bacterial infection (Oldroyd & Downie, 2008; Walker & Downie, 2000). This may explain the formation of empty nodules in L. herbacea when inoculated with Microvirga sp. strains WSM3693 and 3557, whose Nod Factor may have been recognized initially inducing the formation of nodule primordia without the development of an infection thread or the invasion of nodule cells.

The initiation of the infection thread marks a key point in bacterial invasion and it is influenced by a variety of bacterial polysaccharides as well as Nod Factor specific structures (Gibson et al., 2008; Oldroyd & Downie, 2008). When L. herbacea and L. diffusa formed nodules with the Rhizobium, Sinorhizobium and Mesorhizobium ciceri strains, the strains were easily isolated from the nodules following standard surface sterilization methods. This indicated that these rhizobia were able to form infection threads and reach the nodule cells (Trinick et al., 1991), most probably by forming an intercellular infection thread which requires less stringency in the NodFactor structure (Oldroyd & Downie, 2008).

It is important to consider that a suboptimal Nod Factor can still induce signalling if their concentrations are high enough (Oldroyd & Downie, 2008; Walker & Downie, 2000). Martínez-Romero (2003) showed that the promiscuous legume Phaseolus vulgaris is able to nodulate with many more rhizobium species under controlled axenic conditions, than when grown in the field. This could explain the variety of strains able to nodulate L. capitata, L. diffusa and L. herbacea under glasshouse conditions, where the plants were inoculated with a high number of bacteria, and therefore not representative of the ability of L. herbacea to nodulate with all these strains in field.

All the R. leguminosarum field strains were unable to increase Lessertia spp. dry weight, and all the inoculants in use in Western Australia formed white, ineffective nodules. Ineffectiveness is likely due to an inability of the rhizobia cells to gain access to the plant cytoplasm, as specific cell surface components are required at this stage (Perret et al., 2000). Only after the bacteria are in the host cell cytoplasm do they differentiate into the morphologically distinct bacteroid form that is capable of

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N fixation (Gibson et al., 2008; Perret et al., 2000). Even if rhizobia get access to the nodule cell cytoplasm, overcoming all plant responses and Nod-Factor stringency, nitrogen fixation can still fail due to the inability of rhizobia to adapt to their new environment and to persist for enough time in the plant cell (Masson-Boivin et al., 2009). To fully understand the basis of Lessertia spp. promiscuity and at what stage the infection process is interrupted, future studies of nodule structure will be needed.

Results from previous chapters showed that Lessertia spp. in their natural environment nodulate preferentially with Mesorhizobium. It is thus intriguing that L. capitata, L. diffusa, L. herbacea and L. incana choose to nodulate with strains that are ineffective in nitrogen fixation and that belong to a genus that is different to that of their natural symbionts. Moreover, the inoculants selected for L. herbacea and L. diffusa achieved higher dry weights under controlled glasshouse conditions than for any of the other species.

The fact that Lessertia spp. were not successfully established in Badgingarra, although no strains were recovered from this soil (Table 4.3), suggests that the poor establishment of plants and symbionts might be due to a combination of factors and not only to the presence of competitive background rhizobia.

If promiscuity and background competition were the sole reasons for Lessertia spp. poor establishment, then L. excisa should be considered a promising species, as it was unable to form symbiosis with any of the inoculants and did not trap any background strains from the soil. Unfortunately, this species was heavily affected by fungal hypocotyl rot at every site, and in the MPNs and soil trapping experiment, suggesting its successful introduction to Australia could be hampered by susceptibility to fungal pathogens.

Legume establishment and optimal nitrogen fixing symbiosis in a new environment depends upon the pre-selection of plant and microsymbiont for adaptation to the new conditions (Graham, 2008; Howieson & Ballard, 2004). The results from this chapter bring into sharp focus the uncertainties in achieving an optimal symbiosis when developing new legumes and rhizobia for agriculture. The fact that the inoculant strains used in this study were originally collected from sites with soil pH of >5.5

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(Appendix 10) suggests that low soil pH in the target areas could had been one of the causes for low plant adaptation and poor rhizobial survival. Therefore, in the next chapter plant survival, symbiotic performance and competitive ability of different strains will be assessed in a more benign environment, with higher rainfall levels and a slightly higher soil pH. The effect of inoculant dose on nodule occupancy will also be assessed.

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CHAPTER 5. Competitive ability of Mesorhizobium strains from Lessertia spp.

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5.1 Introduction Rhizobia inoculant strains are exposed to diverse environmental and biological factors that may constrain their ability to colonise new niches and therefore reduce overall symbiotic performance and plant productivity (Graham, 2008; Zahran, 1999). One well documented constraint is the presence of indigenous or naturalised soil rhizobia able to nodulate the introduced legume (Dowling & Broughton, 1986; Thies et al., 1991), which may compromise the inoculation outcome, and result in suboptimal nitrogen fixation, or in the worst scenario, a total displacement of the effective inoculant by ineffective soil bacteria (Denton et al., 2002; Dowling & Broughton, 1986; Slattery & Coventry, 1993; Slattery & Coventry, 1999; Thies et al., 1991).

Lessertia spp., along with their microsymbionts, were introduced into Western Australian where the target soils had no record of introduction of this genus, or of any related species. The introduced species had been collected from similar edaphic areas in South Africa. Quite unexpectedly, Lessertia spp. showed an overall poor establishment and survival in different sites of the Western Australian wheat belt (Chapter 4) caused in part by poor nodulation by the inoculant strains and by ineffective nodulation with naturalised R. leguminosarum.

Indigenous and naturalised soil rhizobia are difficult to displace (Brockwell et al., 1995; Howieson et al., 2008), especially in stressful environments, where they may hold the competitive advantage through adaptation to those harsh conditions (Dowling & Broughton, 1986; Howieson & Ballard, 2004; Sessitsch et al., 2002; Thies et al., 1992). Some approaches to overcome competition are to increase inocula concentration (Brockwell & Bottomley, 1995; Dowling & Broughton, 1986; Mårtensson, 1990; Roughley et al., 1993), to select the appropriate host genotype (Brockwell et al., 1982; Demezas & Bottomley, 1986; Drew & Ballard, 2009; Duodu et al., 2005; Yates et al., 2008). to select inoculant strains with a higher competitive ability (Aguilar et al., 2001; Amarger, 1981b; Hungria et al., 2003) and to choose symbioses that are selective (Yates et al., 2008).

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L. diffusa was the species that performed best in all the sites evaluated in 2007 and 2008 but still showed a high incidence of nodulation by background bacteria. This legume was also able to nodulate and increase dry weight with ten different Mesorhizobium strains in glasshouse experiments, which provided an opportunity to test the competitive ability and symbiotic performance of different strains in the field. L. herbacea, on the other hand, did not establish well at any of the sites. It proved to be highly promiscuous in the field and is nodulated ineffectively by several of the commercial inoculant strains currently in use in Western Australia. It was hypothesised that for L. herbacea an increase in the inoculation rate would enrich the number of live cells on the seed and rhizosphere, and therefore the chances of the inoculant to outcompete naturalised rhizobia for infection sites.

This chapter reports on field studies of the effect of increased doses of an effective inoculant strain with the legume L. herbacea, and on the competitive ability and symbiotic performance of different L. diffusa strains, at a site with higher soil pH, higher rainfall and with a presence of background rhizobia.

5.2 Materials and Methods The field experiments described in this chapter were carried out on Boathaugh Estate, Karridale, Western Australia (34º08’27.68’’S; 115º10’35.62’’E) in August 2009. The paddock history included pasture fallow in 2005 and 2006, rye grass- clover mix in 2007, with clover inoculated with WSM1325, and winter wheat and millet (summer) in 2008. Average and year 2009 monthly rainfall is listed in appendix

18. The soil pH of the site was 6.1 (CaCl2).

Prior to sowing into moist soil the site was mowed and sprayed with glyphosate (0.5 L ha-1). Soil samples were randomly collected to estimate the population of indigenous R. leguminosarum bv. trifolii, R. leguminosarum bv. viciae, Sinorhizobium meliloti and Mesorhizobium ciceri bv. biserrulae through the most-probable-number (MPN) technique (section 4.2.2.2) using Trifolium subterraneum (subterranean clover), Pisum sativum (Field pea), Medicago sativa (lucerne) and Biserrula pelecinus respectively as trap plants.

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5.2.1 Experiment 1: Effect of rate of inoculation of WSM3565 on symbiotic performance with L. herbacea. The host species chosen for this trial was L. herbacea, which proved to be a promiscuous species in previous experiments (Chapter 4; section 4.3.7). The chosen inoculant was WSM3565, a strain that was able to significantly increase L. herbacea biomass under glasshouse conditions (Chapter 3; section 3.3.2) and which had not been tested under field conditions.

5.2.1.1 Seed inoculation Seeds of L. herbacea were scarified with sand paper (P240) and surface-sterilised for three minutes in 4% (w/v) bleach, 1 minute in 70%(v/v) ethanol followed by six rinses in sterile de-ionised water. Seeds were aseptically dried in a laminar flow cabinet and stored in sterile paper bags until being slurry inoculated, lime pelletized and sown. The seed germination percentage was assessed by placing 3 replicates of 100 sterilised seeds on 1.5% (w/v) water agar plates and incubating them for 72h at room temperature.

The inoculant strain WSM3565 was recovered from glycerol stocks and grown in ½ LA agar plates at 28°C for five days. One loop full of fresh cells was inoculated into a 250ml conical flask containing 50ml of ½ LA broth which was incubated in a shaker at 28°C for 7 days until it contained approximately 109 cells ml-1 (estimated by optical density (OD600nm) of 1.0). Eight ml of the broth culture were inoculated onto 10 g of sterile peat contained in 30 ml sterile polycarbonate capped tubes. The inoculated peat was mixed and incubated at 28°C for another 10 days until the cell number was approximately 109 cells g-1. Cell number in the peat was assessed by colony count through the Miles and Misra drop plate method (Singleton, 1992) in ½ LA media and using 10-fold dilutions.

Inoculant rates using this peat culture ranged from 1 to 10 times the recommended peat based inoculation rate for medium size seeds (250 g of peat per 50 Kg of seeds), equivalent to be 104 rhizobial cells seed-1 (Deaker et al., 2004; Lupwayi et al., 2000) (Table 5.1).

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Twelve hours prior to sowing, the seeds were pelleted by mixing the amount of peat needed to achieve each of the target doses (Table 5.1), with 3 ml of 1% methocel adhesive solution. The mixture of methocel and peat was applied at a rate of 200 µl for 3 g of seeds and mixed until seeds were evenly covered. Then, in a sterile 150ml polycarbonate container they were mixed with 1 g of dolomite lime until evenly coated. For the maximum rate of inoculation (50 g of peat kg-1), the 3 g of seeds were inoculated with 300 µl and were carefully mixed with dolomite to avoid seed clustering. Each of the treatments were separated into replicates, packed in paper envelopes and stored at 4°C until sowing (14 h later).

Table 5.1. Inoculation rates and peat doses applied to the seed of L. herbacea. Inoculation Rate Proportion of Slurry concentration Potential rhizobia Standard Rate cells seed-1 peat (g) seed (Kg)-1 peat (g) ml-1

2.5 0.5 0.1 1x104

5.0 1.0 0.2 2x104

10.0 2.0 0.4 4x104

50.0 10.0 0.9 1x105

A sample of 100 peat inoculated seeds per treatment was randomly selected to perform cell counts immediately after inoculation. The peat inoculated seeds were suspended in 50mL solution of 0.1% Na4P2O7.10H2O and shaken on a rotary shaker at 200 rpm for 20 minutes (Pijnenborg et al., 1991). Samples were diluted in a 10 fold dilution series in sterile saline solution (0.89% w/v NaCl). Rhizobia cell counts were performed using the Miles and Misra drop plate count method. Plates were incubated for 5 days at 28°C.

5.2.1.2 Experimental design and sowing. The experiment was a complete randomized block design with 5 replicates (blocks) per treatment (Figure 5.1). Each plot consisted of a single row 1.5 m long and spaced 1m apart. The seeds were sown by hand at 1.5 cm depth at a rate of 1g of seed per m. Plots were hand fertilised after sowing with phosphate fertiliser (15% available P) at a rate of 12 g per plot (equivalent to 150kg ha-1).

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Figure 5.1. Experimental block design of treatments including 4 concentrations (0.5, 1, 2 and 10) of seed inoculation and an uninoculated control (UNINOC).

5.2.1.3 Evaluations Five plants (when present) from each plot were sampled at week 12 after sowing to assess plant dry weight and nodule occupancy. Three nodules were carefully removed from each plant, surface sterilised and the bacteria re-isolated in ½ LA media as described in chapter 2, section 2.2.3. The position of the nodules on the root was also recorded. The shoots of the plants sampled were dried at 60°C for 48 hours then weighed.

All the isolates were RPO1-PCR fingerprinted to confirm their identity (Chapter 2, section 2.2.2). In cases where fingerprints did not match that of the inoculant, the banding patterns were further analysed to construct a cladogram and to determine the number of distinct strains. The proportion of nodule occupancy was determined for each of the resident strains as well as for WSM3565.

The identity of the resident strains were estimated at the species level by sequencing their partial dnaK gene following the procedures described in Chapter 2 section 2.2.5.

5.2.2 Experiment 2: Competitive ability of different strains of Mesorhizobium for nodulation of Lessertia diffusa The plant chosen for this experiment was L. diffusa as it achieved the highest establishment percentages at some sites in the Wheatbelt (Chapter 4). The inoculant strains are indicated in table 5.2. The 10 strains had previously increased L. diffusa biomass in comparison to uninoculated plants in a glasshouse experiment (Chapter

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3, section 3.3.2). The possible species identity of each strain was inferred from the dnaK phylogeny (Section 2.3.4)

Table 5.2. Inoculant strains for Lessertia diffusa used in field trial 2. Strain Mesorhizobium species Original Host

WSM2566 M. mediterraneum L. pauciflora

WSM2623 M. mediterraneum L. annularis

WSM3270 Mesorhizobium sp. L. annularis

WSM3495 Mesorhizobium sp. L. capitata

WSM3626 Mesorhizobium sp. L. diffusa

WSM3636 Mesorhizobium sp. L. capitata

WSM3565 Mesorhizobium sp. L. diffusa

WSM3598 Mesorhizobium sp. L. herbacea

WSM3612 Mesorhizobium sp. L. excisa

WSM3898 Mesorhizobium sp. L. excisa

5.2.2.1 Seed inoculation Seed scarification, disinfection and coating were as described in 5.2.1.1. Seed germination percentage was calculated by incubating three subsamples of 100 sterilised seeds on 1.5% (w/v) water agar plates and incubating them for 48h at room temperature.

For each strain, 3 g of seeds were inoculated with 200 l of a mixture of peat in 3ml of 1% methocel. Due to variations in the number of rhizobial cell g-1 of peat within strains, the amount of peat was adjusted to achieve the standard inoculation rate for small seeded legumes of 103 cells seed-1 (Deaker et al., 2004). The inoculated seeds were then coated with 1g of dolomite lime. The replicates of each treatment were packed separately in sterile paper bags and sown 14h later. A sample of 50 peat inoculated seeds per treatment was taken to assess rhizobia survival in the seed as described in section 5.2.1.1.

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5.2.2.2 Experimental design and sowing The experiment was a complete randomized block design with 5 replicates (blocks) per treatment (Figure 5.2). Each plot consisted of a single row 1.5 m long, with a spacing of 1m between rows. The sowing was by hand at a rate of 1g of seed per m2. Plots were fertilised after sowing with 12g of phosphate fertiliser (15% available P).

PLOT 1 2 3 4 5 6 7 8 9 10 11 TREATMENT WSM3898 WSM3612 WSM3598 WSM3626 WSM3270 WSM3636 UNINOC WSM2623 WSM3495 WSM3565 WSM2566 PLOT 12 13 14 15 16 17 18 19 20 21 22 TREATMENT WSM3598 WSM3565 WSM2566 WSM3495 WSM3626 WSM2623 WSM3898 WSM3636 WSM3612 WSM3270 UNINOC PLOT 23 24 25 26 27 28 29 30 31 32 33 TREATMENT WSM3270 WSM3898 WSM3495 UNINOC WSM3565 WSM3598 WSM3636 WSM2566 WSM2623 WSM3626 WSM3612 PLOT 34 35 36 37 38 39 40 41 42 43 44 TREATMENT WSM3612 WSM2566 UNINOC WSM3898 WSM3495 WSM3626 WSM3270 WSM3565 WSM3636 WSM3598 WSM2623 Figure 5.2. Experimental block design of treatments including 10 different inoculant strains for L. diffusa and an uninoculated control (UNINOC).

5.2.2.3 Evaluations Five individual plants were sampled from each plot 12 weeks after sowing for nodulation and dry matter measurements. Shoots were dried at 60°C for 48 hours and weighed. Three nodules were removed from each plant, surface sterilised and the bacteria re-isolated in ½ LA media as described in chapter 2, section 2.2.3. The Mesorhizobium inoculant strains were initially differentiated from soil strains by their growth rate in ½ LA media, as all the inoculant strains used in this experiment required 5 or more days to form 1mm colonies (Chapter 2, section 2.3.2), whereas all soil rhizobia able to nodulate Lessertia in previous experiments (chapter 4, sections 4.3.3 and 4.3.4) and in Experiment 1 of this chapter, were fast growing rhizobia (2-3 days to form 1mm colonies). A sub-sample of 3 isolates from each of the plots were RPO1-PCR fingerprinted to confirm their identity. Nodule occupancy (%) was the criterion to assess competitiveness for each of the strains (Brockwell et al., 1982).

5.2.3 Statistical analyses To determine significant differences between treatments in the percentage of established plants and plant dry weight, a one-way analysis of variance was performed with the statistics programme SPSS 13.0 for Windows (2004). When differences were determined, the treatment means were compared using the Fishers’s Least significant difference (LSD) test (P0.05).

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The banding patterns for the isolates from section 5.2.1, were analysed using the software BIOCapt (ALYS Technologies, Lausanne). The genetic distance between isolates was calculated using AFPL SURV version 1.0 (Vekemans, 2002) and the UPGMA cluster analyses were performed using the NEIGHBOR application from the PHYLIP software package. The cladograms were visualized in MEGA4 (Tamura et al., 2007).

The dnaK sequences of the distinct strains from section 5.2.1 were analysed in Gene Tool Lite 1.0 (2000) software (Doubletwist, Inc., Oakland, CA, USA) and in MEGA4 (Tamura et al., 2007) as described in chapter 2, section 2.2.6.

5.3 Results The soil population of R. leguminosarum bv. trifolii nodulating T. subterraneum, estimated through the MPN technique, was 2 x 105 cells g-1of soil. No nodulation of V. sativa, M. sativa and B. pelecinus was recorded at any soil dilution.

5.3.1 Effect of rate of inoculation of WSM3565 on symbiotic performance with L. herbacea.

5.3.1.1 Viable rhizobia cells on seeds The germination (%) of L. herbacea seed was 72.2%. The population of rhizobia on the seed surface after inoculation was slightly lower than the potential number of cells per seed (Table 5.1). For the inoculation rate of 2.5 g of peat per Kg of seed the number of rhizobia cells per seed was of 7.2 x103, which corresponded to 72% of the -1 expected rhizobia population. For the inoculation rates of 5, 10 and 50 g Kg , the rhizobia cell counts per seed were 1.6x104 (80% cell survival), 3.7x104 (92.5% cell survival) and 8.7x104 (87% cell survival) respectively.

5.3.1.2 Plant population and biomass production The percentage of established plants ranged between 28 to 47%. There was no effect of the different concentrations of the inoculant strain (WSM3565) (P0.05) on the percentage of established L. herbacea plants at week 12 after sowing (Figure 5.3).

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There were no differences (P0.05) in shoot dry weight at different inoculation rates. The shoot dry weights of the uninoculated plants were equivalent to those inoculated with increasing doses of WSM3565 (Figure 5.4). It is important to note that there was high variability in plant size within plots and treatments, and that no clear signs of nitrogen deficiency (chlorotic foliage) were evident.

Figure 5.3. Established plants of L. herbacea (percent of potential) inoculated with WSM3565 at four inoculation rates (equivalent to 2.5, 5, 10 and 50 g of peat kg-1 seed), assessed 12 weeks after sowing. Vertical lines correspond to the standard error of means. No significant difference between treatments was detected through ANOVA (P0.05)

Figure 5.4. Shoot dry weights of L. herbacea inoculated at four inoculation rates of strain WSM3565 (equivalent to 2.5, 5, 10 and 50 g of peat kg-1 seed) Vertical lines correspond to the standard error of means. No difference between treatments was detected through ANOVA (P0.05)

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5.3.1.3 Nodule occupancy Of the 197 isolates obtained from L. herbacea nodules across all inoculation rates, only 78 isolates (39.5%) matched the inoculant strain WSM3565 RPO1-PCR fingerprint. The other 119 isolates were grouped as 12 distinct strains (A to L) based on their RPO1 banding patterns (Appendix 17).

The nodule occupancy achieved by the inoculant strain WSM3565 (indicated in red, Figure 5.5) ranged between 28 to 31% for the three lowest inoculation rates (7.2x103, 1.6x104 and 3.7x104 cells seed-1) and increased to 54% with the highest inoculation rate. The percentage of nodule occupancy of each of the soil naturalised strains (in shades of green and yellow, Figure 5.5) was different in the five treatments, with B dominating.

Figure 5.5. Percentage nodule occupancy by Mesorhizobium inoculant WSM3565 (In red) and 12 soil strains (A to L; in shades of green and yellow) in L. herbacea inoculated with: 1) 7x103 cells seed-1 ; 2) 1.6x104 cells seed-1; 3) 3.7x104 cells seed-1 4) 9.8x104 cells seed-1 and 5) Uninoculated control.

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Several soil strains were present in only some of the treatments, and in a low proportion of nodules (less than 3%) and were usually represented by 1 or 2 isolates (Strains from F to L). Strains A, B and C were the dominant resident strains and were present in every plot.

5.3.1.4 Nodule occupancy according to distribution of nodules Overall, the percentage of nodules on the tap root occupied by WSM3565 was 70% and only 17.65% of the nodules on lateral roots corresponded to WSM3565. It is important to note, that 45.7% of the nodules on the tap root were desiccated. Some of these dry nodules (25%) were rehydrated in laboratory and the rhizobia could be recovered, corresponding in every case to WSM3565.

5.3.1.5 Sequencing of dnaK gene A single DNA fragment of 283bp representing the partial dnaK gene was amplified for the 12 soil naturalised strains isolated from L. herbacea. The majority of the strains clustered in the Rhizobium genus clade with a high bootstrap support (87%) (Figure 5.6). From these, 9 of the strains (A, B, C, D, E, F, G, J and L) were closely related to the species R. leguminosarum bv. trifolii, and to strains isolated from sites from the Western Australian wheat belt (Buntine, Katanning, Muresk and Newdegate) in chapter 4, which are also included in this tree. Only strain I, representing two isolates, clustered apart from this genus and its dnaK sequence did not show similarities with any known species.

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Figure 5.6. Neighbor joining tree based on the sequencing of the partial dnaK gene. Strains in red correspond to naturalised soil rhizobia isolated from L. herbacea in Karridale. The number of isolates and the origin are indicated in brackets.

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5.3.2 Competitive ability of different strains of Mesorhizobium nodulating Lessertia diffusa 5.3.2.1 Viable rhizobia cells on seeds L. diffusa seed germination (%) in water agar media was 89.3%. The number of viable rhizobia cells per seed after inoculation was variable depending on the inoculant strain. The highest number was recorded for strain WSM3612 (960 cells seed-1). Most of the strains (WSM2566, 3565, 3636, 3598, 3898, 2623 and 3626) were present on the seed in numbers that ranged from 270 to 490 cells seed-1. The lowest numbers were registered for the strains WSM3270 and 3495: 191.9 and 51.1 viable rhizobia cells per seed which corresponded to 19.2 and 5.1% of the potential rhizobial population respectively (Table 5.3).

Table 5.3. Number of viable rhizobia cells on seeds of L. diffusa after inoculation. (Average number of cells ± SE)

5.3.2.2.Plant population The percentage of established plants in the field ranged from 38 to 63% (Figure 5.7). None of the inoculant treatments had a significant effect on the numbers of L. diffusa plants (P0.05). There was high variation between replicates.

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Figure 5.7. Percentage of established plants of L. diffusa inoculated with different Mesorhizobium strains and of uninoculated plants. Data from 12 weeks after sowing. Vertical lines correspond to the standard error of means. No significant difference between treatments was detected through ANOVA (P0.05)

5.3.2.3 Plant biomass production and nodule occupancy There were differences in L. diffusa dry weights for the different inoculant strains (solid blue bars in Figure 5.8). The strains WSM2566, 3495, 3612, 3898, 2623 and 3270 had no effect on plant biomass in comparison to the uninoculated control. The highest dry weights were achieved when inoculating with WSM3636, 3598, 3565 and 3626 all of which produced higher dry weights than the uninoculated control (P0.05).

The percentage of nodule occupancy (bars with diagonal stripes in Figure 5.8) was variable according to the inoculant strain. The highest nodule occupancy (%) was achieved by strains WSM3636 and WSM3598, with 81.8 and 84.6% of the nodules respectively. Two of the strains that significantly increased dry weights, WSM3565 and 3626, achieved only 40 and 55.5% of nodule occupancy respectively. Most of the strains that did not increase dry weights (WSM2566, 2623, 3898 and 3270) showed a percentage of nodule occupancy lower than 50% (ranging from 37.5 to 47.4%), the only exception was WSM3495 which occupied 52% of the nodules.

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Figure 5.8. L. diffusa shoot dry weights (solid bars) and percentage of nodules occupied by the inoculant strains (bars with stripes). Shoot dry weight of strains that share a letter are not significantly different according to Fisher’s LSD test (P0.05)

5.3.2.4 Nodule occupancy according to distribution of nodules The percentage of the inoculant nodule occupancy varied according to the position of the nodules. The amount of nodules colonized by the inoculant strains tended to be higher when the nodules assessed were located on the tap root, close to the crown, than in lateral root nodules (Figure 5.9). The strains WSM3612, 3598, 3270, 3636 and 3898 achieved tap root nodule occupancies >75%. However, only WSM3636 and 3598 maintained high nodule occupancy in lateral roots. Overall, strains that achieved over 50% nodule occupancy on both tap and lateral roots (WSM3598, 3636 and 3626) induced a significant increase in plant biomass in comparison to the uninoculated control (Figure 5.8).

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Figure 5.9. Percentage of nodule occupancy of ten different Mesorhizobium strains inoculated on L. diffusa. The strain numbers are indicated in red, each of them are linked to two pie charts that correspond to tap root and lateral root nodule occupancy. Strains in figure A did not increase plant dry weight; strains in figure B significantly increased L. diffusa dry weight. .

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5.4 Discussion A primary aim of legume inoculation is to maximize rhizobial survival in the time between introduction to the soil and nodulation (Brockwell & Bottomley, 1995). The presence of naturalised or indigenous rhizobia capable of nodulating the target legume can become a threat to nodulation and nitrogen fixation by introduced strains (Denton et al., 2002; Slattery & Coventry, 1993; Slattery & Coventry, 1999; Thies et al., 1991). This study assessed the survival, competitive ability and symbiotic effectiveness of different Mesorhizobium strains nodulating L. diffusa. It also investigated whether an increase in the inoculation dose would improve the overall symbiotic performance of L. herbacea inoculant strain WSM 3565 in a field site with background competitive rhizobia.

Despite the interventions of increased inoculation doses and the improved soil and environmental conditions (higher average rainfall and a soil pH of 6.3), the percentage of plants of L. diffusa and L. herbacea that established did not improve in comparison to previous field trials in the Western Australian wheat belt (Chapter 4). The average establishment for L. herbacea was 37.4% (10 plants m-2) and for L. diffusa 46.6% (16 plants m-2). There were no differences in establishment (%) between inoculated treatments (i.e. different inoculant strains for L. diffusa and different inoculation doses for L. herbacea) and the uninoculated control, suggesting that the low plant establishment was due to factors including poor nitrogen availability. The seed germination for both species was previously assessed in laboratory and was satisfactory (72.2% for L. herbacea and 89.3% for L. diffusa), but in the field seeds can show a reduced vigour as well as reduced viability due to detrimental climatic and edaphic factors (Bewley & Black, 1985; Powell, 2006). Plant density might have also been affected by the below average rainfall conditions during October (2009)(Appendix 11).

Although WSM3565 had proved effective for L. herbacea under glasshouse conditions (Chapter 3), none of the inoculation doses increased shoot dry weight in comparison to the uninoculated control (P0.05). The nodule occupancy achieved by WSM3565 in L. herbacea, was also low and varied from 28 to 31% with the three

143 CHAPTER 5 lowest inoculation rates and increased to 54% with the highest rate, which was still not enough to increase dry weight.

The naturalised strains competing for nodulation in L. herbacea were identified as R. leguminosarum. This was no surprise, since it had been the dominant competing species in previous field experiments with Lessertia spp. (Chapter 4) and it is present in high populations in different environments of Australia (Ballard et al., 2002; Brockwell et al., 2008; Denton et al., 2002; Howieson & Ballard, 2004).

A great diversity of R. leguminosarum strains were identified through RPO1 PCR fingerprinting, with the most frequently isolated strain (B) (figures 5.5 and Appendix 17), reaching a nodule occupancy ranging from 27 to 67% in the uninoculated control. Several authors have indicated that within mixed populations of naturalised R. leguminosarum strains in the field, there are some that dominate nodule occupancy (Denton et al., 2002; Leung et al., 1994a; Leung et al., 1994b; Zhang et al., 2001). This can be related to their competitive ability (Handley et al., 1998; Leung et al., 1994a; Meade et al., 1985), or an effect of the legume host (Ballard & Charman, 2000; Dowling & Broughton, 1986; Leung et al., 1994b; Yang et al., 2001) but mainly to their long term adaptation to edaphic and environmental conditions (Denton et al., 2002; Dowling & Broughton, 1986; Leung et al., 1994a; Zhang et al., 2001).

When some of these competitive soil rhizobia are effective or partially effective with the target legume, an advisable strategy is to maximise nitrogen fixation by manipulating the rhizobial population (Aguilar et al., 2001; Brockwell & Bottomley, 1995; Brockwell et al., 1995; Denton et al., 2003; Hungria et al., 2003; Mpepereki et al., 2000; Thies et al., 1991). This recommendation cannot apply for L. herbacea, as it forms ineffective nodules with R. leguminosarum strains (Chapter 4). In cases like this, displacing the ineffective population by the effective inoculant is quite a challenge, and the most sensible choice seems to be to increase inoculant dose and to select for competitive strains to outcompete the resident soil rhizobia (Amarger, 1981a; Brockwell et al., 1995; Herridge, 2008; Meade et al., 1985) or in the worst case scenario to change the host (Sessitsch et al., 2002). While the approach of increasing inoculant dose with L. herbacea did not render the expected results, the

144 CHAPTER 5 outcomes achieved by L. diffusa inoculated with different microsymbionts were more promising.

A potential forage species should ideally be able to germinate and establish at high rates in the target environment (Dear & Ewing, 2008). Although the establishment (%) of L. diffusa was low, if the criteria for evaluating the success of inoculation are the proportion of occupied nodules and an increase in dry matter production (Brockwell et al., 1995; Thies et al., 1991), then the inoculation of L. diffusa with some of the inoculants could be considered a success. The strains WSM3636, 3598, 3626 and 3565 were able to increase L. diffusa dry weight, consistent with data obtained under controlled conditions (Chapter 3, section 3.3.2). However, contradictory results were obtained with strains WSM3898 and 3495, which increased plant biomass in glasshouse experiments but not in the field. Differences in symbiotic performance for similarly effective strains are likely to happen in the field, as their tolerance to environmental conditions and competitive ability may vary (Amarger, 1981b; Ames-Gottfred & Christie, 1989; Brockwell, 1982; Dowling & Broughton, 1986). The current data emphasize the importance of matching the legume host with its appropriate microsymbiont under field conditions (Brockwell et al., 1995; Graham, 2008; Howieson & Ballard, 2004; Sessitsch et al., 2002) and not just extrapolate results obtained under controlled conditions.

The higher yield achieved following inoculation of L. diffusa with WSM3598 and 3636 may be explained by their higher nodule occupancy (>80%). However, WSM3626 and 3565 occupied only 55.5 and 37.5% of the nodules respectively and plant dry weight was as high as with the two former strains. In contrast, strains like WSM2566, 3495, 2623 3270 and 3612 occupied between 45 and 55% of sampled nodules but did not increase plant biomass despite being effective strains under glasshouse conditions.

Nodule occupancy has frequently been correlated with an increase in plant yield (Amarger, 1981a; Demezas & Bottomley, 1986; Mårtensson, 1990; Thies et al., 1992). It is thus intriguing that the increase in nodule occupancy with the highest inoculant dose in L. herbacea, or with some strains in the case of L. diffusa, did not result in higher biomass accumulation by the plant. The literature is conflicting on the

145 CHAPTER 5 ideal proportion of inoculant nodule occupancy to induce a response to inoculation. There have been reports of low nodule occupancy (<20%) by the effective strain that still lead to an increase in plant yield (Aguilar et al., 2001); and there are also cases where the effective inoculants occupy more than 50% of the nodules resulting in suboptimal symbiotic performance (Ames-Gottfred & Christie, 1989; Demezas & Bottomley, 1986; Denton et al., 2002; Yates et al., 2008). Leung et al. (1994a) suggested that ineffective or partially effective soil strains may exert an antagonistic effect over other nodule occupying strains (in this case the inoculant) leading to reduced symbiotic performance. On the other hand, an increase in dry weight by strains that achieve only low nodule occupancy might be explained by the ability of legume to compensate for the low number of nodules by increasing nodule size and therefore enhancing nitrogen fixation (Coventry et al., 1985; Richardson et al., 1988). In the case of L. diffusa and L. herbacea data suggests that the plant is unable to compensate.

In general, nodules present on the tap root were occupied in a higher proportion by the inoculant strains (Table 5.5), which is likely due to the strategic placement of the inoculant on the seed (Brockwell et al., 1995; Denton et al., 2003; Hardarson et al., 1989; McDermott & Graham, 1989). This is an indication that the inoculation was successful in ensuring that the inoculant strain formed the first nodules, which are the ones that start contributing nitrogen in the earlier stages of development of the legume (Brockwell et al., 1982; Pijnenborg & Lie, 1990; Spriggs & Dakora, 2007). Yet, nodule formation on lateral roots is a strongly indicative of the ability of the strains to move and to colonise the rhizosphere (McDermott & Graham, 1989; Pijnenborg et al., 1991; Spriggs & Dakora, 2007), and become increasingly important at later stages in plant development (Dowling & Broughton, 1986; Hardarson et al., 1989; Sessitsch et al., 2002). Moreover, lateral root nodules may have played an important role in these experiments due to the low rainfall conditions during October (Appendix 11) that caused the top soil to dry and therefore the earlier senescence of a great proportion of the crown nodules. Thus, plants might have had to rely on nitrogen fixed by newly formed nodules, i.e. those formed on secondary roots. In L. diffusa the proportion of nodules formed by the different strains in lateral roots was variable. Most of the strains occupied less than 55% of lateral root nodules, and in L. herbacea the occupancy of WSM3565 did not exceed 20% with any of the

146 CHAPTER 5 inoculation doses. This is of concern considering that L. herbacea and L. diffusa are perennial species that should rely on the inoculant established with the seed in the first year during the following seasons (Ballard et al., 2003; Brockwell et al., 1982; Sessitsch et al., 2002) and that nodule occupancy is expected to decline in time (Brockwell et al., 1995; Denton et al., 2002; Svenning et al., 2001). Nevertheless, the strains WSM3636 and 3598 nodulating L. diffusa consistently increased plant dry weight and were present in 85.7 and 75.0 % of the lateral nodules respectively and appeared to have the capacity to colonise the rhizosphere. Therefore, these strains should be considered as prospective inoculants for this legume in the field.

The high numbers of resident rhizobia in the field are likely to be the main reason for the low nodule occupancy by some effective strains with L. diffusa and of WSM3565 with L. herbacea. The population of clover rhizobia (also able to nodulate L. herbacea and L. diffusa) was 2 x 105 cells g-1 of soil, which is considered high (Slattery et al., 2004) and if converted to number of cells per area were equivalent to 2 x 1010 cell m-2 (assuming a depth of 10cm and a bulk density of 1.0 (Brockwell et al., 1995)). The maximum inoculation rate was 8.7 x 104 cells seed-1 which corresponds to approximately 9.6 x 106 cells m-2 (at a sowing rate of 110 seed m-2). This indicates that the inoculant strain was outnumbered by naturalised soil rhizobia 2000 times.

This large numerical difference is applied before the loss of viability of the inoculant cells from the time they are exposed to seed inoculation until they are able to colonise the rhizosphere (Deaker et al., 2004; Dowling & Broughton, 1986; Lowther & Patrick, 1995; Roughley et al., 1993; Trotman & Weaver, 2000). The numbers of WSM3565 on the L. herbacea seed were reduced to 70-80% immediately after the inoculated seeds were air dried, and in L. diffusa inoculants varied from 5 to 96.8% (Table 5.3) cell survival. A further reduction in rhizobia numbers in the soil is highly likely to happen (Lowther & Patrick, 1995; Roughley et al., 1993). The ability of rhizobia to survive on seeds can be highly variable and have an impact on nodule occupancy and dry weights (Becerra Stiefel et al., 1998; Brockwell et al., 1982; Lowther & Patrick, 1995; Trotman & Weaver, 2000). However, the variation in L. diffusa strains survival after inoculation did not have a clear effect in the nodule occupancy achieved by each strain.

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Inoculating legumes in soils with the presence of a large population of competitive rhizobia is generally regarded as challenging (Ballard & Charman, 2000; Ballard et al., 2003; Brockwell et al., 1995; Denton et al., 2002; Slattery & Coventry, 1993; Thies et al., 1991). Moreover, the use of legumes that are prone to form ineffective symbioses with soil rhizobia is discouraged (Howieson & Ballard, 2004). However, this study showed that some L. diffusa strains are able to outcompete local populations of rhizobia (WSM3636 and 3898) and to colonise lateral roots, which makes them promising to develop as inoculants. There was also evidence of an overall higher performance of L. diffusa in comparison to L. herbacea, as had been observed in previous field experiments (Chapter 4) suggesting an effect of the host on strain performance in the field. It should be noted though, that L. diffusa establishment was still not up to standard (Li et al., 2008) and that a great disparity between plants was observed within plots, which may in part had had an impact in the poor plant establishment. This suggests a need for selection of L. diffusa for adaptation to these conditions and for symbiotic performance (Dowling & Broughton, 1986; Tripplett & Sadowsky, 1992) before embarking into new field inoculation trials.

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CHAPTER 6. General Discussion

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150 CHAPTER 6

The replacement of native perennial vegetation by annual agricultural crops and pastures in many areas of South Western Australia has led to an increase in dryland salinity associated with a slowly rising water table (George et al., 1997; Lambers, 2003; Pannell & Ewing, 2006; Ward, 2006) and to a reduction in plant diversity making production systems more vulnerable to diseases, pests and environmental stresses (Dear et al., 2003). This area is also facing a reduction in rainfall and an increase in average temperature due to climate change (Gregory et al., 2005; Nielsen & Brock, 2009). The inclusion of perennial plants, particularly deep-rooted herbaceous legumes, into agricultural systems is considered likely to play a key role in managing salinity and in facing changing weather patterns (Cocks, 2001; Dear et al., 2003; Howden et al., 2007; Howieson et al., 2008; Pannell & Ewing, 2006; Ward, 2006).

The main purposes of this work were to introduce the South African perennial legumes Lessertia capitata, L. diffusa, L. excisa, L. herbacea, L. incana and L. pauciflora into Western Australia, to characterize their root nodule bacteria and to study the impact of their nitrogen fixing symbionts on their establishment.

6.1 Potential of Lessertia spp. for Western Australian stressful environments. Lessertia spp. were thought to be eligible candidates for establishment in Western Australian stressful environments due to their inherent characteristics of deep rooting systems, perennial habit and high seed production (Balkwill & Balkwill, 1999; Goldblatt & Manning, 2000; Howieson et al., 2008). These species are native to areas of infertile soils and low rainfall conditions in South Africa (Anderson & Hoffmann, 2007; Goldblatt & Manning, 2000; Nkonki, 2003), which match the conditions of the Western Australian wheat belt. This is important as it is widely accepted that when introducing a plant species into a new environment, it should have ideally evolved under similar edaphic and climatic conditions (Francis, 1999; Howieson et al., 2008; Vogel et al., 2005).

Since legume performance relies to a great extent on the symbiotic relationship with root nodule bacteria (Brockwell & Bottomley, 1995; Howieson et al., 2000a; Parker,

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2001; Richardson et al., 2000), the first approach of this work was to study the diversity and phylogenetic relationships of micro-symbionts associated with Lessertia spp. aiming to shed some light on rhizobia identity, genetic stability and possible performance in the field. The RPO1 and ERIC PCR fingerprints showed a wide diversity of strains isolated from Lessertia spp. collected from different areas of South African Western, Northern and Eastern Cape. Sequencing of the housekeeping genes dnaK and 16S rRNA identified the great majority of the strains as Mesorhizobium spp. and one as a Burkholderia sp.

The sequencing of the nodulation gene nodA showed that Lessertia strains were closely related and separated from other known nodA sequences and that the sequences were clustered according to the ecological area from which they were isolated. The nodA genes are usually associated to host specific nodulation (Perret et al., 2000; Roche et al., 1996), therefore the close relationship between the strain’s nodA suggested specificity of Lessertia spp. for a particular rhizobial symbiotic genotype, which is common for plants growing in restricted ecological niches (Perret et al., 2000), as well as symbiotic genes adaptation to environment and to available hosts (Masson-Boivin et al., 2009; Rivas et al., 2007).

One of the risks in using Mesorhizobium spp. as inoculants is the possibility of transfer of genetic elements to soil bacteria via their symbiosis island (Nandasena et al., 2007a; Sullivan & Ronson, 1998). The discordance between housekeeping and symbiotic phylogenies in Lessertia strains suggests that lateral transfer events of symbiotic genes had occurred in the past (Ochman et al., 2000; Sullivan et al., 1995; van Berkum et al., 2003). This led to the investigation of the presence of the insertion region of the “symbiosis island” in Lessertia strains. While, the insertion region could only be amplified for one of the strains (WSM3495), it cannot be ruled out that the other strains may possess the ability to transfer and receive symbiotic genes, as identified in other Mesorhizobium spp.

The phylogenetic studies have shown that Lessertia microsymbionts belong to the genus Mesorhizobium. As this genus represents a small proportion of Australian native rhizobia (Lafay & Burdon, 1998; Lafay & Burdon, 2007; Yates et al., 2004) it is unlikely that competition will be a major obstacle in the introduction of these species.

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The nodA clustering also suggests that Lessertia spp. are selective with regards to the rhizobia able to nodulate them. Moreover, the absence of closely related legumes to Lessertia spp. in the target regions of Australia indicated a reduced risk of rhizobial competition and a greater likelihood of success in the establishment of this legume (Brockwell et al., 1995).

6.2 Rhizobial competition and legume promiscuity as constraints for establishment of Lessertia spp.

In the first two attempts to establish Lessertia spp. in areas with acid soils and low rainfall patterns in the Western Australian wheat belt (Chapter 4), the number of established plants was very low. The main cause for this was the low numbers of the inoculant in the soil and nodules, and the unexpected nodulation of Lessertia spp. with soil rhizobia of the genus Rhizobium spp.

Further field studies to assess plant and rhizobia symbiotic performance in a more benign environment (higher rainfall, higher soil pH) in Karridale, Western Australia (Chapter 5), included only the species L. diffusa and L. herbacea. The approach was to improve rhizobia survival in the soil by delivering higher numbers of rhizobia; and to assess inoculant strains competitive abilities by testing different strains with L. diffusa, which had shown the best establishment rates in previous trials.

In every field trial, L. herbacea failed to establish effective symbiosis with its inoculant strain and tended to form symbiosis with resident Rhizobium spp., even when inoculated at higher rates of inoculum and grown under improved environmental conditions. This species was expected to perform best in field experiments in the target areas due to its adaptation to low soil pH, as it was originally collected from acid and poor soils (Appendix 5). L. herbacea was also able to establish effective associations with several Lessertia strains (Chapter 3), and was the most promiscuous of the Lessertia spp., nodulating with Sinorhizobium meliloti, S. medicae, Rhizobium leguminosarum bv. trifolii, R. leguminosarum bv. viciae and Mesorhizobium ciceri (Chapter 4). The ability of some legumes to associate with different strains and species of rhizobia can allow them to establish and even invade

153 CHAPTER 6 different environments. Some examples are the widely distributed invasive legumes, Cytisus scoparius (scotch broom) (Lafay & Burdon, 2006), Robinia pseudoacacia (black locust) (Mierzwa et al., 2010; Ulrich & Zaspel, 2000) Macroptilium atropurpureum (Jesus et al., 2005; Trinick et al., 1991), some Mimosa spp. (Elliott et al., 2009) and several Acacia spp. (Ben Romdhane et al., 2006; de Lajudie et al., 1998; Marsudi et al., 1999) some of which are widely dispersed within Australia (Hussey et al., 2007) and worldwide (Weber, 2003). Why the promiscuity of L. herbacea has become its fate and not a means of improving its establishment in the field is something that still cannot be answered. Symbiotic promiscuity could perhaps be agriculturally managed if there were suitable native or naturalised strains present in the field (Aguilar et al., 2001; Brockwell et al., 1995; Graham, 2008; Hungria et al., 2003; Mpepereki et al., 2000). Nevertheless, L. herbacea in every case nodulated ineffectively with naturalised rhizobia, therefore reducing the chances of managing promiscuity.

The fact that Lessertia spp. nodulated ineffectively with strains of the genus Rhizobium is puzzling, especially considering that these legume species were nodulated with Mesorhizobium spp. in nearly 100% in their natural environment in South Africa, where they are equally exposed to diverse root nodule bacteria (Elliott et al., 2007; Sprent et al., 2010; Spriggs & Dakora, 2009; Yates et al., 2007). Several cases of rhizobial inter-strain competition have been reported, but in most of them legumes nodulate with rhizobia of the same genus or species of their original inoculant (Ames-Gottfred & Christie, 1989; Denton et al., 2002; Mårtensson, 1990; Slattery & Coventry, 1999; Yates et al., 2005). Cases of competition with other rhizobia species are less common (Aguilar et al., 2001; Ben Romdhane et al., 2007; Elliott et al., 2009) and the plant usually tends to nodulate with its appropriate inoculant if present or with naturalised rhizobia that effectively fix nitrogen (Graham, 2008; Martínez-Romero, 2003; Trinick et al., 1991).

L. diffusa was the species that performed best at all sites and particularly at the Karridale site (Chapter 5). Although it was affected by ineffective nodulation with naturalised rhizobia, WSM3636 and 3598 were able to overcome competition and significantly improve plant dry weight. Thies et al. (1992) suggested that highly competitive inoculant strains are able to perform well across a range of

154 CHAPTER 6 environments. This seems to be true for WSM3636 which was superior to other strains in the Karridale field trial but had also been the only strain recovered from plants in situ and from soil (one year after inoculation), in experiments in the Wheatbelt (Chapter 4). These results also emphasized the importance of the rhizosphere colonisation by the inoculant, and lateral root infection, in the L. diffusa response to inoculation. The strains that achieved highest plant dry weight were those that were present in a high proportion in lateral root nodules that were formed later during the growing season. Those strains that were present mainly in the tap root nodules but in a small proportion in newly formed lateral nodules (WSM3898 and 3495), did not have a significant effect on plant biomass in spite of their nitrogen fixing effectiveness in glasshouse experiments (Chapter 5).

The comparison between L. diffusa and L. herbacea field performances demonstrates the impact that legume host can have in the symbiotic relationship with rhizobia. The inoculant strain WSM3565 had proved effective with L. herbacea in glasshouse conditions and was only partially effective with L. diffusa (Chapter 3). However, in the field, WSM3565 produced an increase in L. diffusa dry weight but not in L. herbacea, although it was inoculated at 10 times the inoculant dose. The exact mechanisms that allowed this to happen are unknown. It could be due to an ability of L. diffusa to better discriminate for effective rhizobia, as has been reported for other legume species and cultivars (Ballard & Charman, 2000; Drew & Ballard, 2009; Trinick et al., 1991; Yates et al., 2005).

Low soil pH is known to be one of the primary stresses for legume and rhizobium symbiosis (Dilworth et al., 2001; Graham, 2008; Howieson & Ballard, 2004; Slattery et al., 2004). However, in this study plant establishment for L. diffusa and L. herbacea was not improved when soil pH increased from the wheatbelt sites (pH 4.6 - 4.9) to the Karridale site (pH 6.3). There was limited improvement in herbage production with L. diffusa and higher nodule occupancy at the Karridale site.

All inoculant strains’ pH tolerance range was established in the laboratory prior to the field trials (Chapter 2), however the observed pH tolerances did not translate to the field setting. Strains WSM3612 and 3898, which had shown a superior low pH tolerance to WSM3636 in the laboratory, failed to result in enhanced biomass

155 CHAPTER 6 production in the field and colonized only a small fraction of root nodules outside of the crown at the Karridale site. Indeed, at the Wheatbelt sites they were not recovered from soil or nodules.

Assessment of rhizobia pH tolerance in laboratory should suggest an ability of the bacteria to survive in acid soils (Dilworth et al., 2001; Graham et al., 1994; Slattery et al., 2004). However these results, coincidently with other studies (Howieson et al., 1988; Slattery & Coventry, 1995; Slattery & Coventry, 1999), show that competitive ability and symbiotic effectiveness is variable in the field no matter whether strains are low pH tolerant or sensitive in the laboratory.

6.3 Future research There is a wide diversity of native and naturalised rhizobial strains in Australian agricultural soils (Howieson & Ballard, 2004) and strains able to tolerate and persist under stressful conditions such as low pH and drought have been released in the last decades (Howieson & Ewing, 1986; Watkin et al., 2000) which makes them likely to persist in these soils (Slattery & Coventry, 1999). The main competitors for nodulation in Lessertia spp. are strains very closely related to inoculant strains TA1 and WSM1325 of R. leguminosarum bv. trifolii, which have had widespread use as inoculants for clover in Australia (Herridge, 2008). To establish L. diffusa as an alternative pasture legume in Australia, the saprophytic and competitive ability of the most promising strains WSM3636 and WSM3598 needs to be established. It will also need to be determined whether sufficient rhizobia are being released from decayed nodules to initiate new nodules (Brockwell et al., 1995; Roughley et al., 1993; Thies et al., 1995) and whether they are able to outcompete the high populations of background R. leguminosarum.

The production of bacteriocins by root nodule bacteria has been reported previously and is thought to confer a competitive advantage in the soil environment (Oresnik et al., 1999; Robleto et al., 1997; Sridevi & Mallaiah, 2008; Triplett & Barta, 1987; Tripplett & Sadowsky, 1992), and has been observed in particular for several naturalised R. leguminosarum strains (Joseph et al., 1983; Oresnik et al., 1999; Rodelas et al., 1998; Triplett & Barta, 1987; Wilson et al., 1998). Determining

156 CHAPTER 6 whether the R. leguminosarum strains collected from field trials are producing bacteriocins that are affecting Mesorhizobium strains could explain high levels of ineffective nodulation by the clover strains. This remains to be investigated.

High dry matter production is one of the main selection parameters for perennial legumes for future domestication (Howieson et al., 2008) and a high percentage of plant establishment and density are essential for a forage legume from an agricultural point of view (Dear & Ewing, 2008; Li et al., 2008). In this study, even with strains that allowed a significant increase in L. diffusa dry weights and were able to occupy a high proportion of the nodules, the plant did not reach the potential observed in glasshouse trials. Lessertia spp. showed overall low numbers of established plants and a wide variation in size and performance within plants from the same plots. The most likely cause for these could be that edaphic and nutritional factors, other than nitrogen availability, are affecting L. diffusa overall performance and yield (Brockwell et al., 1995; Thies et al., 1991). However, based on the variation observed, it cannot be discarded that within plant species there might be different ecotypes, which can show differences in tolerance to environmental conditions (Joshi et al., 2001; Vogel et al., 2005) and also in their ability to select rhizobia (Ballard et al., 2002; Charman & Ballard, 2004; Drew & Ballard, 2009). If the latter is true, then plant screening to select accessions that show better adaptation and also a higher symbiotic performance should be a next step (Brockwell et al., 1995; Sessitsch et al., 2002) in Lessertia spp. domestication.

6.4 Concluding remarks There has been a lack of studies of unutilized legume species and their micro- symbionts (Brockwell et al., 1995; Doyle & Luckow, 2003; Hirsch et al., 2001; Howieson et al., 2008; Laguerre et al., 2001; Masson-Boivin et al., 2009; Willems, 2006). This work contributes to the knowledge of a new legume genus and associated rhizobia as a potential symbiosis for pasture crop development. This work also highlights the importance of studying legume-rhizobia symbiotic relationships and their interaction with soil flora under field conditions to ensure their successful establishment in a new geographic region.

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158 REFERENCES

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182 APPENDIX

Appendix 1. Lessertia capitata distribution in Southern Africa Source: African Plant database. Conservatoire et Jardin Botaniques (CJB), Ville de Geneva. Accessed in March 2010 http://www.ville-ge.ch/cjb/

183 APPENDIX

Appendix 2. Distribution of Lessertia diffusa in Southern Africa. Source: African Plant database. Conservatoire et Jardin Botaniques (CJB), Ville de Geneva. Accessed in March 2010 http://www.ville-ge.ch/cjb/

184 APPENDIX

Appendix 3. Distribution of Lessertia excisa in Southern Africa. Source: African Plant database. Conservatoire et Jardin Botaniques (CJB), Ville de Geneva. Accessed in March 2010 http://www.ville-ge.ch/cjb/

185 APPENDIX

Appendix 4. Lessertia herbacea distribution in Southern Africa. Source: African Plant database. Conservatoire et Jardin Botaniques (CJB), Ville de Geneva. Accessed in March 2010 http://www.ville-ge.ch/cjb/

186 APPENDIX

Appendix 5. Lessertia incana distribution. Source: African Plant database. Conservatoire et Jardin Botaniques (CJB), Ville de Geneva. Accessed in March 2010 http://www.ville-ge.ch/cjb/

187 APPENDIX

Appendix 6. Distribution of Lessertia pauciflora. Source: African Plant database. Conservatoire et Jardin Botaniques (CJB), Ville de Geneva. Accessed in March 2010 http://www.ville- ge.ch/cjb/

188 APPENDIX

Appendix 7. Experiments performed with isolates from Lessertia spp. l

Experiment n° 2.2.2 2.2.3 2.2.4 2.2.5 2.2.6 3.2.1 Molecular Authentication pH range Sequencing Sequencing Sequencing Isolates Fingerprint dnaK 16S rRNA nodA WSM2211     WSM2559       WSM2561       WSM2562  WSM2566       WSM2607      WSM2622       WSM2623      WSM2628     WSM2801   WSM2804   WSM2805  WSM2808     WSM2809  WSM2814  WSM2991   WSM2995   WSM3059     WSM3060   WSM3061   WSM3062   WSM3265   WSM3270       WSM3273   WSM3274  WSM3275   WSM3310       WSM3311   WSM3326   WSM3472      WSM3492      WSM3493     WSM3495       WSM3501   WSM3502      WSM3503      WSM3507      WSM3513      WSM3564      WSM3565       WSM3571   WSM3592     WSM3595   WSM3596     WSM3597   WSM3598       WSM3602       WSM3605  WSM3606      WSM3612       WSM3620      WSM3626       WSM3628  WSM3635  WSM3636       WSM3803      WSM3804       WSM3805      WSM3806      WSM3807  WSM3822   WSM3891      WSM3892  WSM3893      WSM3894      WSM3895  WSM3897  WSM3898       WSM3900      WSM3917      WSM3918   WSM3919       WSM3920     

189 APPENDIX

Appendix 7. continued

Experiment n° 3.2.2 3.2.3 4.2.1 4.2.4 5.2.1 5.2.2 Nodulation Sequencing Establishment Establishment Inoculation rate Competition Isolates Specificity intS Field 2007 Field 2008 Field 2009 Field 2009 WSM2211 WSM2559   WSM2561   WSM2562 WSM2566    WSM2607 WSM2622   WSM2623    WSM2628 WSM2801 WSM2804 WSM2805 WSM2808 WSM2809 WSM2814 WSM2991 WSM2995 WSM3059 WSM3060 WSM3061 WSM3062 WSM3265 WSM3270    WSM3273 WSM3274 WSM3275 WSM3310   WSM3311 WSM3326 WSM3472 WSM3492 WSM3493 WSM3495    WSM3501 WSM3502 WSM3503 WSM3507 WSM3513 WSM3564 WSM3565     WSM3571 WSM3592 WSM3595 WSM3596 WSM3597 WSM3598      WSM3602   WSM3605 WSM3606 WSM3612      WSM3620 WSM3626      WSM3628 WSM3635 WSM3636      WSM3803 WSM3804   WSM3805 WSM3806 WSM3807 WSM3822 WSM3891 WSM3892 WSM3893 WSM3894 WSM3895 WSM3897 WSM3898    WSM3900 WSM3917 WSM3918 WSM3919   WSM3920

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Appendix 8. Cladogram showing the relationships among Lessertia spp. root nodule bacteria according to fingerprints obtained with RPO1-PCR. Letters after each strain correspond to the original host: L.annularis (La); L.capitata (Lc); L. diffusa (Ld); L. excisa (Le); L. frutescens (Lf); L. herbacea (Lh); L. incana (Li); L. inflata (Linf); L. microphylla (Lm); L. pauciflora (Lp); L. rigida (Lr); L. tomentosa (Lt)

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Appendix 9. Cladogram showing the relationships among Lessertia spp. root nodule bacteria according to fingerprints obtained with ERIC-PCR. Letters after each strain correspond to the original host: L.annularis (La); L.capitata (Lc); L. diffusa (Ld); L. excisa (Le); L. frutescens (Lf); L. herbacea (Lh); L. incana (Li); L. inflata (Linf); L. microphylla (Lm); L. pauciflora (Lp); L. rigida (Lr); L. tomentosa (Lt)

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Appendix 10. Some edaphic and climatic characteristics of the collection sites in South Africa

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Appendix 11. Sequence alignment for the intergenic region between phe-tRNA and intS for Lessertia capitata mesorhizobia strainWSM3495, Biserrula pelecinus mesorhizobia strain WSM1271 and M. loti strain MAFF303099.

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Appendix 12 . Average and 2007 monthly rainfall. Badgingarra, Western Australia.

Appendix 13 . Average, 2007 and 2008 monthly rainfall. Buntine, Western Australia

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Appendix 14. Average and 2007 monthly rainfall. Katanning, Western Australia

Appendix 15 . Average and 2007 monthly rainfall. Muresk, Western Australia

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Appendix 16 . Average, 2007 and 2008 monthly rainfall. Newdegate, Western Australia

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Appendix 17. Cladogram showing the relationships among soil rhizobia isolates nodulating L. herbacea in Karridale, Western Australia according to fingerprints obtained with RPO1-PCR.

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Appendix 17. continued

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Appendix 18. Monthly average and 2009 rainfall summary, Karridale, Western Australia.

200