Some arbuscular mycorrhizal fungi contain endocellular bacteria. In Gigaspora margarita BEG 34, a homogenous population of β-Proteobacteria is hosted inside the fungal spore. The bacteria, named Candidatus Glomeribacter gigasporarum, are vertically transmitted through fungal spore generations. Here we report how a protocol based on repeated passages through single-spore inocula caused dilution of the initial bacterial population eventually leading to cured spores. Spores of this line had a distinct phenotype regarding cytoplasm organization, vacuole morphology, cell wall organization, lipid bodies and pigment granules. The absence of bacteria severely affected presymbiotic fungal growth such as hyphal elongation and branching after root exudate treatment, suggesting that Ca. Glomeribacter gigasporarum is important for optimal development of its fungal host. Under laboratory conditions, the cured fungus could be propagated, i.e. could form mycorrhizae and sporulate, and can therefore be considered as a stable variant of the wild type. The results demonstrated that – at least for the G. margarita BEG 34 isolate – the absence of endobacteria affects the spore phenotype of the fungal host, and causes delays in the growth of germinating mycelium, possibly affecting its ecological fitness. This cured line is the first manipulated and stable isolate of an arbuscular mycorrhizal fungus.
Arbuscular mycorrhizal (AM) fungi colonize the root cortex of the majority of land plants, supplying them with nutrients, including phosphate, and conferring resistance against a variety of biotic and abiotic stresses (Smith and Read, 1997). In addition to the strong emphasis which is currently placed on the ecological roles played by AM fungi in nature (Van der Heijden and Sanders, 2002), there are other aspects of their biology which make them unique microbes: they are obligate biotrophs, meaning they get carbon exclusively through a symbiotic relationship with a host plant, lack host specificity (being present in approximately 80% of land plants), and are multinucleate, opening the question whether they are heterokaryotic (Hijri and Sanders, 2005) or homocaryotic polyploid organisms (Pawlowska and Taylor, 2004). In addition and as a further element of their genomic complexity, AM fungi offer the best-known example of an association between fungi and bacteria (Bonfante, 2003). Many bacteria complete their life cycle within eukaryotic cells (Moran, 2001), but the Fungal Kingdom offers a limited number of examples of endobacteria (Partida-Martinez and Hertweck, 2005; Lumini et al., 2006). Glomeromycota represent a niche for a largely undescribed population of bacteria (Scannerini and Bonfante, 1991; Schüßler, 2002). Until a few years ago, the biology of these bacterial populations – with only their morphology described – was a black box. Recently we have shown that an isolate of Gigaspora margarita hosts a homogenous population of bacteria (at least 20 000 per fungal spore) (Bianciotto et al., 2003; Jargeat et al., 2004; Lumini et al., 2006). The spores of G. margarita seem to be a specialized niche for rod-shaped bacteria which have also been consistently found in many other Gigasporaceae in all stages of their life cycle (Bianciotto et al., 1996; 2000). On the basis of their 16S RNA sequence, the bacteria have been grouped into a new taxon, which – due to their current unculturable status – has been named as Candidatus Glomeribacter gigasporarum (Bianciotto et al., 2003), clustering within the large group of β-Proteobacteria related to the Burkholderia, Pandorea and Ralstonia genera. Their genome is highly reduced: 1.4 Mb consisting of a chromosome and a plasmid (Jargeat et al., 2004). The status of uncultivable microbe and this small genome strongly suggest that Ca. Glomeribacter gigasporarum is an obligate endosymbiont, like other comparable endosymbionts living in aphids (Moran, 2003), and justifies fundamental questions regarding the exact nature of a potentially intimate fungus–bacterium symbiotic relationship.
As antibiotic treatments to cure G. margarita from its endobacteria were unsuccessful (P. Jargeat and G. Bécard, unpubl. results), biological material appropriate to understand the role of Ca. Glomeribacter gigasporarum was not available. Here, we report how a protocol based on the use of successive in vitro single-spore inocula (Bianciotto et al., 2004) caused a dilution of the microbial population eventually leading to bacteria-cured spores. Morphological analyses as well as mycorrhizal experiments showed that the cured spore line (B–) was a stable variant of the original wild type (WT) (B+), showing clear-cut morphological differences and important changes in the growth of the germinating mycelium.
Production of G. margarita cured spores
Successive spore generations (SGs) were obtained after inoculating transformed carrot roots with single-spore inocula originating from two separate batch pots. Spores originated from a single SG0 spore were identified as a lot (Fig. 1). Six lots were considered during the experiments. As previously described (Bianciotto et al., 2004), the endobacterial population in successive spore generations was found to rapidly decrease from SG0 to SG4. Here (irrespectively of the starting lot), 68% of the SG4 spores were found free of bacteria when observed by confocal microscopy (Table 1), while conventional PCR showed that only 19% of SG4 spores were indeed free of bacteria (Table 1). The discrepancy between the two methods suggests a different level of sensitivity. Single SG4 spores were used to inoculate new root cultures in order to obtain SG5 and SG6 spores. No endobacteria were detected by confocal microscopy and by conventional PCR in SG5 and SG6 spores (Table 1).
Table 1. Bacterial presence in spores of Gigaspora margarita produced in vitro with monosporal inoculum.
Conventional PCR analysis
Presence of endobacteria (%)
Presence of endobacteria (%)
To assess the sensitivity of the PCR technique to detect the endobacteria in our biological system serial dilutions of DNA template from the WT spores (B+) were amplified with the specific 23S rDNA bacterial primers in conventional and hemi-nested PCR (Fig. 2). With conventional PCR the amplification product was visible until 10−3 sporal DNA dilution, while the hemi-nested technique detected the presence of the endobacteria until the 10−4 dilution. Real-time quantitative PCR assays confirmed the data obtained by hemi-nested technique, allowing us to detect a minimum of 10 bacterial genomes for each PCR mixture in the same 10−4 dilution (A. Salvioli, E. Lumini and V. Bianciotto, unpubl. results). DNA extracts from randomly selected SG5 and SG6 spores did not give any amplification, even in undiluted samples (Fig. 2). The fungal DNA was detected in all the dilutions tested, demonstrating the good quality of the DNA extract and the absence of inhibition. The results show that the hemi-nested PCR technique is a suitable tool to detect endobacteria, increasing the level of sensitivity of conventional PCR at least 10-fold.
Cured spores of G. margarita can still complete their life cycle
To examine the stability of the bacteria-free status of cured spores, and to obtain them in quantity, sorghum and clover were grown in pots and inoculated with a multisporal inoculum consisting of 40–60 cured spores.
The spores produced in 10 pots of sorghum inoculated with cured spores were analysed by PCR and confocal microscopy. All were still free of bacteria (Table 2). In order to confirm this result with a different host plant, the spores produced by 10 clover plants inoculated with cured spores were also analysed. PCR and confocal microscopy confirmed that they were free of bacteria (Table 2).
Table 2. Bacterial presence in spores of Gigaspora margarita produced in pot cultures with multisporal inoculum of cured spores.
Origin of starting B– spores
Production of B– spores
Conventional PCR analysis
Presence of endobacteria (%)
Presence of endobacteria (%)
Gigaspora margarita-cured spores have maintained their genetic diversity
Random amplified polymorphic DNA (RAPD) and inter-simple sequence repeat (ISSR) amplifications revealed 33 bands among which 19 were polymorphic (57.6%). When considering the polymorphic bands, the same level of diversity was found for both spore populations [percentage of polymorphic bands (p.p.b.) = 54.5 and 51.5 for B+ and B– populations respectively].
The genetic diversity [Shannon–Weaver index (H)] in B+ and B– spore populations was calculated as H = 4.06 and H = 3.07 respectively [the average diversity within populations (Hpop) = 3.57 and the diversity among all spores (Hsp) = 3.71 for total spore populations]. This result suggests that: (i) G. margarita BEG 34 contains genetic variability, and (ii) B+ spores show a slightly higher genetic diversity than B– spores. However, only 3.8% [(Hsp − Hpop)/Hsp] of the total genetic variation can be attributed to divergence between B+ and B– populations whereas a large proportion of diversity was recorded among spores within populations (Hpop/Hsp = 96.2%). Accordingly, amova partitioned 98.1% of the total variation within populations and only 1.9% between the two populations.
The patterns of genetic relationships among spores have been drawn by principal co-ordinate analysis (PCA) based on the pairwise Euclidean distances between spores. The first two axes account for 45.4% of the total variation ( Fig. 3). PCA analysis reveals a complete overlap among B+ and B– spores, without any clustering.
Gigaspora margarita-cured spores do not appear different in mycorrhizal phenotype
To assess more carefully the infectivity of cured spores, mycorrhization experiments were set up in pots of sorghum and clover. Sorghum pots were inoculated with B+ spores and four lots of B– spores, identified as B-1, B-2, B-3 and B-4 (i.e. groups of cured spores originated from independent SG0 spores). Clover pots were inoculated with B+ spores and one additional lot of B– spores (lot 5). The frequency of mycorrhization (F), the intensity of mycorrhization (M), the percentage of arbuscules per infected areas (a) and per root apparatus (A) were not different between plants inoculated with B+ spores or B– spores (Table 3). Similar results showing no difference between B+ and B– mycorrhization were obtained with clover as well as with the in vitro system using transformed carrot roots (Table 3).
Table 3. Mycorrhizal colonization intensity.
Different letters indicate significantly different values according to the Krukall–Wallis test of variance (P < 0.05). Statistical analyses of clover and sorghum data were treated separately.
Transformed carrot root WT
Transformed carrot root cured
No cytological difference was noted, using light microscopy and electron microscopy, in the respective mycorrhizal structures observed in clover roots. The integrity of important biotrophic structures such as the plant membrane surrounding the arbuscules looked normal in B– mycorrhizae (not shown).
The experiment with sorghum allowed us to evaluate sporulation of the cured line versus that of the original line. No significant difference was observed between the lines. After 3 months the mean number of spores produced was 980.2 and 875.4 for WT and cured spores respectively (Table 3). Similarly, no significant difference was found after 3 months in the spore production of the in vitro system using carrot hairy roots (Table 3).
Spores from both B+ and B– lines had the same mean size, with a diameter ranging from 333 to 325 μm (Table 4). In agreement with previous descriptions (Bonfante et al., 1994), the B+ spores of G. margarita BEG 34 had a laminated wall and complex protoplasm (Fig. 4A–C), containing many nuclei, membranous secretion structures, abundant melanin-like granules, storage structures (lipid droplets, protein bodies and glycogen particles) as well as pleiomorphic vacuoles. Some of them contained bacteria, while others contained electron-dense globules. Thin sections of cured spores (Fig. 4B–D) showed cytoplasm organization different from B+ spores mostly regarding vacuole populations. The bacteria-containing vacuoles disappeared, as well as the conspicuous protein bodies, leaving electron-transparent vacuoles, often containing small electron-dense granules of uncertain composition (Fig. 4B).
Table 4. Main features of WT and cured spores of Gigaspora margarita.
Different letters indicate significantly different values according to the Krukall–Wallis test of variance (P < 0.01).
Spore diameter (μm)
Wall thickness (μm)
Nuclear diameter (μm)
By light microscopy, Coomassie Blue stained many minute structures in WT spores (Fig. 5A), suggesting the presence of protein bodies that may correspond to those seen by transmission electron microscopy (Bonfante et al., 1994). In contrast, there were no protein-positive structures in cured spores (Fig. 5B). The periodic acid-Schiff (PAS) reaction detecting polysaccharides showed abundant masses of glycogen in WT spores (Bonfante et al., 1994) (Fig. 5C), absent in cured spores (Fig. 5D).
The morphology of lipid bodies (stained by Sudan Black) was strikingly different. In WT spores, structures of different size were identified: some small 2–3 μm globules (inset in Fig. 5E) were strongly stained, while the large (14–16 μm) globules filling up most of the spore cytoplasm were not stained (Fig. 5E). As these structures are usually identified as the characteristic oil droplets of AM fungal spores (Grandmougin-Ferjani et al., 2005), the result suggests that these were extracted during fixation and dehydration, as already suggested by Sward (1981). The same staining on cured spores revealed very few Sudan black-positive bodies, while the dominant lipid droplets were consistently smaller (diameter about 4 μm; Fig. 5F). This led to a different organization of the cytoplasmic network in the two spore lines.
Other differences, perhaps reflecting changes in lipid and membrane composition, were also suggested by confocal microscopy experiments aimed to reveal the presence of bacteria (Fig. 6). For these observations, spores were squashed in order to liberate their cytoplasm and to stain the bacteria. While the cytoplasm of WT spores spread over the slide, forming a strip where nuclei and endobacteria were mostly present in an external halo (Fig. 6A, arrowheads and inset), cytoplasm from the cured spores remained attached to the spore wall, forming a viscous drop with a sharp border. Nuclei were all grouped at the periphery of this drop (Fig. 6B). Mean nuclear sizes (6.5 and 6.3 μm for WT and cured spores respectively) were not affected (Table 4).
Melanin-like granules are an important component of Glomeromycota spore cytoplasm, as melanin is often incorporated into the cell wall, contributing to its strength against biotic and abiotic stresses (Bonfante and Vian, 1984). The presence of these electron-dense granules was decreased in the cured spores. In contrast with the WT spores (Fig. 4C), there was no evidence of their incorporation into the fungal wall (Fig. 4D).
The lamination of the spore wall appeared the same in both fungal populations, as also confirmed by PAS staining for polysaccharides, but walls of cured spores were significantly thicker than those of WT spores (16.1 μm versus 12.15) (Fig. 7 and Table 4).
More detailed differences in the cell wall of the two lines could be seen by electron microscopy. The cell wall inner layers (Iw) were comparable in organization but not in thickness. The middle layer (Mw) always showed the architecture described as helicoidal (Bonfante and Vian, 1984), and contained many chitin-rich sheets (Fig. 7C and D). Surprisingly, additional sheets were present in the middle layer of cured spores, and also in the outer layer (Ow), which was consistently thicker (Fig. 7D).
Presymbiotic growth of cured G. margarita is severely affected
In order to compare growth capacities of B+ and B–G. margarita during the presymbiotic stage, the length of the main hyphae produced by germinating spores was recorded over time. Only main hyphae were considered here because of their importance in long-distance connections to host roots. After 30 days, hyphal length produced by WT germinating spores was twice as great as that produced by B– spores (Fig. 8). During the first 10 days, there was no significant difference in growth rate, but B+ hyphal growth lasted more than 25 days while B– hyphal growth stopped after 15 days. The experiment was repeated three times with different lots of B– spores and gave similar results, sometimes with an even greater difference between B+ and B– spores (not shown). In the presence of root exudates, significant hyphal length stimulation (twice) was observed for both spore lines (Fig. 8). Stimulation of hyphal branching by GR24, a strigolactone analogue known to strongly stimulate presymbiotic growth of AM fungi (Akiyama et al., 2005; Besserer et al., 2006), was also analysed. Hyphal branching typically occurs when the fungus grows close to a root, and is believed to increase the probability of fungus–root contacts. After 15 days the number of hyphal branches produced by B+ and B– germinating spores was recorded (Fig. 9). Branching was more strongly activated in B+ spores (18.5 times) than in B– spores (5.2 times). The experiment was repeated five times to be confirmed with different lots of B– spores and gave similar results.
Altogether these data suggest that the overall growth potential of B– germinating spores, during both asymbiotic and presymbiotic stages, is significantly smaller than with WT spores.
To check whether such growth differences were maintained in the presence of the plant, an additional experiment was set up using in vitro carrot roots. When the spores were put at 1.5 and 3 cm from the root fragments, germinated hyphae from cured and WT spores both reached the root surface in 4/5 days, showing comparable development rates, as already found with growth in the absence of the plant (see Fig. 8, first 5 days). However, when the spores were put at 10 cm, growth differences between B+ and B– spores were already significant after 5–6 days and became more and more important after 10–15 days; after 20 days mycelium from B+ spores produced conspicuous branching, established root contacts and formed appressoria. At that time, mycelium B– spores were 4 cm long, but still active (Fig. 10).
Endobacteria or bacteria-like organisms are widespread in the cytoplasm of AM fungi, as integrated components which add new pieces to the AMF genome puzzle (Bonfante, 2003). Here we have presented evidence that G. margarita can be cured from the endobacteria Ca. Glomeribacter gigasporarum living in the spore cytoplasm, giving rise to a new line. To our knowledge this line is the first manipulated and stable isolate of an AM fungus, as – differently from the original isolate – it lacks part of its natural genetic information. Organisms which in laboratory conditions lose their endobacteria are defined as aposymbiotic (Claes and Dunlap, 2000). We have shown that aposymbiotic G. margarita is affected in its phenotype, showing important modifications in spore reserves (glycogen, proteins, lipids), in wall structure and in growth of the germinating mycelium, though not in its ability to mycorrhize plant roots. These features were consistently found in six independent lots of cured spores suggesting that the new traits do not result from the accumulation of mutations related to the in vitro curing process. Finger print analyses comparing genetic polymorphism in populations of WT and cured spores suggest that the new traits of the cured spores are not either the result of a loss of genetic diversity.
How does G. margarita lose its endobacteria under laboratory conditions?
Antibiotic treatments do not affect endobacterial viability (P. Jargeat and G. Bécard, unpubl. results), whereas endobacteria were lost after four to six generations in root organ culture (ROC). Some ideas are suggested by the following observations: (i) G. margarita has been maintained in pot cultures for years using multispore inoculation (Bonfante et al., 1994), and under these conditions the endobacteria are always found, even if in variable numbers (Bianciotto et al., 2004; Jargeat et al., 2004), (ii) bacteria are present in the spores produced in ROCs following multispore inoculation, and (iii) spores without bacteria can easily be found after several single-spore-inoculated ROCs. Thus single-spore inoculation can be identified as the condition affecting maintenance of the endobacterium. If we speculate that spores containing the highest population of endobacteria are more infective (better fitness, see below) and therefore more competitive, they will be positively selected within a multispore inoculum and will transmit large endobacterial populations to the next generation. With single-spore inoculation, there is no selection, and even spores with low endobacterial content may establish mycorrhizae, and produce new spores with a similar low bacterial content, increasing the chance of losing bacteria along the generations.
The progressive decrease in ROC conditions (bacterial numbers fall by one-fifth at each ROC generation according to Bianciotto et al., 2004) could also be the result of a reduced capacity of the bacteria to multiply in this system. Identification of a bacterial cell division marker for Ca. Glomeribacter, i.e. the FtsZ gene belonging to the dcw cluster and differentially expressed during the fungal life cycle (I. Anca et al., unpubl. results) should provide a molecular tool to test the hypothesis.
The endobacteria affect the spore phenotype
A detailed ultrastructural study of the B+ and B– spores of G. margarita revealed that the B– spores had a thicker cell wall. Not only did the bacteria disappear in cured spores, but the protein bodies also became less abundant (or less detectable with Coomassie blue). The small electron-dense granules, identified as melanin-like pigment granules (Grippiolo and Bonfante, 1984), are usually incorporated in the cell wall of WT spores but were rarely detectable in cured spores. The glycogen particles also seemed to be affected in the cured spores, but the most striking morphological difference involved lipids, the lipid bodies appearing smaller in cured spores. Spore morphology thus shows that the presence of bacteria modifies some metabolic aspects involving protein, lipids and melanin; cell wall changes can be a consequence of the nutritional balance between the fungus and its endobacteria.
All these important differences, likely reflecting significant metabolic modifications, suggest that the bacterial and fungal partners strongly and intimately interact at the level of their C metabolism. To address the question whether the bacterial presence represents a carbon cost for the fungus, we compared the multiple germination capacities of B– and B+ spores. Both types of spores exhibited the same capacity as they could germinate successively up to 35 times (not shown data). Simple sugars must be sequestered by Ca. Glomeribacter but without causing any serious quantitative carbon deprivation for the fungal host. We hypothesize that both the partners exploit common resources, such as sugars, in a cooperative manner for fuelling their own metabolism. Examples of such functional integration are available from the well-known insect endosymbionts. Studies of endobacteria such as Buchnera and Wigglesworthia show that notwithstanding their reduced genome these bacteria provide important metabolites to their host, like essential amino acids (Wernegreen, 2004).
To unambiguously confirm that the loss of bacteria is responsible for these phenotypic changes, a complementation experiment should be performed showing that reintroduction of the bacteria into a cured line restores the WT phenotype. As Ca. Glomeribacter gigasporarum cannot be grown in pure culture (Jargeat et al., 2004), such an experiment is not yet feasible. In this context, our analyses of B+ and B– spores were made by using several lines of B– spores, coming from independent curing (mother spores) runs. Thus, when the different B– lines show similar phenotypic traits, we can be more confident that the differences are the result of the loss of bacteria, and not the outcome of variability between spores.
Cured spores still keep their biotrophic capacities
The mycorrhization experiments using clover and sorghum showed that cured lines keep their biotrophic capacities. This trait, already suggested by our first in vitro experiments (Fig. 1, spore generations SG4, SG5 and SG6), allows us to conclude that production of cured spores is possible because they can still colonize roots and complete their life cycle; without mycorrhization, a new generation of spores is not possible.
Symbiotic status depends on many other physiological features; first the fungus has to perceive the plant in the rhizosphere in order to successfully interact (Harrison, 2005). Fungal branching after root exudate treatment is one phenotypic marker of this event (Giovannetti et al., 1996; Buée et al., 2000). Strigolactones have recently been identified to stimulate branching (Akiyama et al., 2005) and to activate respiration and mitochondrial activities in the fungus (Tamasloukht et al., 2003; Besserer et al., 2006). Interestingly, the growth rate of B+ and B– mycelium was comparable for the first 10 days, but after that the WT mycelium developed faster and was more abundantly branched. The B– mycelium was also less stimulated by the root exudates and the strigolactone analogue. The reduced lipid and protein reserves of the cured spores could offer a first explanation for this important physiological difference. The colonization experiment performed on using transformed roots showed that both cured and WT spores germinated and reached the root surface more or less at the same time (5 days), when distances were short (not longer than 3 cm). At least in this time and space frames, B– spores are vigorous enough to establish mycorrhizae. However, with spores placed 10 cm away from the root during long-time experiments (20 days), mycorrhization with B– spores was severely affected.
Taken as a whole, our results suggest that cured spores are disadvantaged at a specific moment of their life cycle (the presymbiotic phase), from the sixth day after germination. The germinating mycelium is less fitted in reaching its host, and this may cause the positive selection towards the B+ condition.
Symbiosis covers a wide range of situations, in which one of the partners may be more or less dispensable. For example, plants, which naturally live in association with AM fungi, can grow without AM fungi under controlled conditions. The reverse is not true: AM fungi cannot accomplish their life cycle without a plant host. In a similar way, AM fungi directly isolated from the field often contain bacteria or bacteria–like organisms (BLOs) (Scannerini and Bonfante, 1991), while AM fungi maintained under in vitro or laboratory conditions often do not contain bacteria. Here also if the endobacteria are, to some extent (this study), dispensable for the fungus, the reverse seems not to be true, at least if we recall the obligate biotrophic status of Ca. Glomeribacter gigasporarum (Jargeat et al., 2004). AM fungi are therefore dispensable for plants, and endobacteria for AM fungi. However, in ecological conditions natural selection has favoured mycorrhizal fungi as indispensable for plant success (Read, 1991). The same may be true for endobacteria.
Spores of G. margarita Becker and Hall (BEG 34; deposited at the European Bank of Glomeromycota) containing the Ca. Glomeribacter gigasporarum endobacteria and produced in pot culture by BIORIZE (Dijon, France) were used for all experiments. They are referred as WT or B+ (with bacteria) spores, while the cured spores are identified as B– (without bacteria). WT or cured spore populations are indicated as lines. A voucher of the cured spores is deposited with the number E32 HC/F at Herbarium Cryptogamicum Fungi, Department of Plant Biology, Torino, Italy.
Conditions for obtaining and cultivating G. margarita-cured spore
Figure 1 summarizes the main steps of the procedure.
Production. A clone of root-inducing T-DNA-transformed carrot roots, established by Bécard and Fortin (1988), was propagated on minimal (M) medium in 120 × 120 mm square Petri dishes. For use as the plant partner in interaction with the fungus, root explants were standardized and prepared as described by Bécard and Piché (1992). Spores of G. margarita were collected from pot cultures (each pot culture representing a batch), surface sterilized, stored at 4°C in sterile water and used as fungal inoculum (Bianciotto et al., 2004). A single starting spore (SG0) was used to inoculate the transformed root and to initiate mycorrhizal cultures in Petri dishes. The dishes were carefully sealed with Parafilm to confine the internal atmosphere and incubated in the dark at 26°C. They were held vertically so that the spore germ tubes elongated upward (as a result of negative geotropism) and contacted the transformed root. In each Petri dish, new spores (SG1) were produced within 2 months as a result of mycorrhizal colonization. These spores were used individually to inoculate new root cultures in order to obtain SG2 spores. Similar successive cycles were reproduced to obtain SG3, SG4, SG5 and SG6 spores. A decreasing number of bacteria were found in spores of each new generation (Bianciotto et al., 2004). These successive subcultures were therefore used to eventually produce cured spores. All the cured spores originated from an independent single SG0 spore were referred as belonging to the same lot.
Cultivation. In pot culture. To maintain and propagate the cured spores, leek, sorghum and clover plants were inoculated with a multisporal inoculum of cured spores obtained as described above.
Plants were grown in pots in order to become colonized and to produce a new generation of cured spores within 3 months. The same protocol was used to maintain WT spores. The cycles of spore production continued from 2002 to 2006.
In vitro culture. To maintain and propagate the cured spores, T-DNA-transformed carrot roots were propagated on minimal medium in Petri dishes and inoculated with a multisporal inoculum of cured spores. The same protocol was used to maintain the WT spores.
Identification of batches and lots of cured spores. Six lots of cured spores were originated by two batches (two starting pots): batch 1 (lot 1, 2, 3, 4) and batch 2 (lot 5 and 6). In particular the different lots of cured spores, originating from independent in vitro curing treatments, were submitted to different culture cycles. Cured spores of lot 1 were propagated through three cycles of leek pot culture, lot 2 through one cycle of leek pot culture, lot 4 through one cycle of clover pot culture, lot 5 through four cycles of clover pot cultures, lot 6 through one cycle (of 2 years) of sorghum pot culture, while spores of lot 3 were maintained under in vitro conditions throughout.
Checking the presence of bacteria
Microscopical detection of bacteria. After each mycorrhizal cycle, the presence or absence of bacteria was monitored in randomly selected spores. These were placed on microscope slides in 20 μl of Bacteria Counting Kit component A (B-7277; Molecular Probes) diluted 1:1.000 according to the manufacturer's directions. The spores were then crushed with a coverslip, incubated in the dark for at least 5 min, and observed with an Olympus FluoView confocal microscope. Bacteria were observed directly using a mercury arc fluorescence lamp, an excitation band-pass (BP) filter at 490 nm (blue) and an emission BP filter at 520 nm (green). For confocal imaging, the 488 nm band of a Kr-Ar laser and a BP 510–540 nm emission filter were used.
Detection of Ca. Glomeribacter gigasporarum by conventional and hemi-nested PCR amplification. For the molecular analyses, DNA extracts either from SG0 (B+) or from SG6 (B–) spores (single or pooled in 10) were used in conventional and hemi-nested PCR assays. Miniprep DNA extraction was performed by crushing the spores in a volume of 30 μl (single spore) or 50 μl (pool of 10 spores) of 1:1 H2O:10X RedTaq PCR Buffer (100 mM Tris-HCl, pH 8.3, 500 mM KCl, 11 mM MgCl2 and 0.1% gelatine). After incubation at 95°C for 15 min, the crude extract was centrifuged at 10 000 g for 5 min, and the supernatant was used for 10-fold serial dilutions. Extreme care was taken to avoid external and cross-bacterial contamination, and all steps were carried out under a laminar flow hood. For the conventional PCR, DNA extracted from B+ and B– spores was amplified by specific bacterial primers BLOf-BLOr and GlomGIGf-GlomGIGr. Primers NS31-AM1 (Helgason et al., 1998) that amplify a portion of the fungal 18S ribosomal gene were used as positive controls to assess the quality of the DNA extraction and the absence of inhibition of the PCR assays. Conventional PCR was carried out as described in Bianciotto et al. (2004).
To confirm the results obtained by conventional PCR we performed a hemi-nested PCR assay.
For such reaction, 1 μl of the conventional PCR products obtained with GlomGIGf-GlomGIGr on DNA extracted from B+ and B– was added as DNA template to 24 μl of PCR mix. The reaction mixture contained 10 mM Tris-HCl, 50 mM KCl, 1.5 mM MgCl2, 0.1% gelatine, 0.2 mM of each dNTP, 0.5 μM primers GlomGIGf and GIGrA(5′-GTTGTTGCCCTCTTGACACC-3′), 1 U of Taq polymerase. Amplifications were carried out in 0.2 ml PCR tubes using a GeneAmp PCR System 9700 thermocycler. The programme consisted of 25 cycles of 45 s at 94°C, 1 min at 60°C and 45 s at 72°C and a final extension of 7 min at 72°C. The resulting 106 bp PCR products were visualized in a 2.5% high-resolution agarose gel using SYBR Safe™ DNA gel stain (Invitrogen) for visualization under UV light.
Estimation of genetic diversity among WT and cured spores by RAPD and ISSR amplification
PCR and electrophoretic conditions. To estimate and compare levels of genetic diversity among B+ and B–G. margarita spores, finger printing experiments were carried out. Forty B+ spores and 36 B– spores (belonging to lots 5 and 6) were analysed by RAPD and ISSR amplification. These two lots were also used for growth test (see Growth capacities of WT versus cured germinating spores).
DNA was extracted from individual spores by using the WIZARD Genomic DNA Purification kit (Promega – Lyon, France). The final pellet was re-suspended in 20 μl of sterile water. To avoid any contamination, DNA extraction and PCR preparation were always performed under sterile conditions.
For RAPD analysis, primers 152 (Sélosse et al., 1998), cTB6 and cTB9 (http://plantbio.berkeley.edu/~bruns/primers.html) were used. Reactions were carried out in a 25 μl volume containing 5 μl of 5× GoTaq buffer (Promega – Lyon, France), 1 μM of primer, 250 μM of each dNTP, 1 unit of GoTaq Polymerase (Promega – Lyon, France) and 1 μl of DNA solution. PCR cycling conditions of the Mastercycler thermocycler (Eppendorf) were as follows: initial denaturation at 94°C for 3 min; 40 cycles at 94°C for 1 min, 40°C for 1 min, 72°C for 2 min and a final extension at 72°C for 10 min.
For ISSR analysis, reactions were carried out using primers R1 (DHB CGA CGA CGA CGA CGA; Hantula et al., 1996), R2 (DDB CCA CCA CCA CCA CCA; Hantula et al., 1996) and R3 (BDB ACA ACA ACA ACA ACA; Hantula et al., 1996) as described for RAPD procedure except amplifications were performed with an initial denaturation of 3 min at 94°C, followed by 37 repetitions of 45 s at 94°C, 45 s at 58°C (R1 and R2) or 50°C (R3) and 3 min at 72°C, and ended by a final extension of 10 min at 72°C.
Amplification products were separated by electrophoresis in 1× TAE buffer using a 1.5% agarose gel. Gels were ethidium bromide stained and numerical pictures of banding patterns were captured under UV light with a CDD camera supported by VISION-CAPT software (Vilber Lourmat, France) for later scoring.
Reproducibility of the patterns given by each primer was checked by comparing results of two independent PCR carried out with 10 subsamples. Once reproducibility was assessed for each primer, the 76 samples were amplified in one run of PCR. A negative control without template DNA was included in each run to test the absence of DNA contamination.
Scoring of RAPD and ISSR fragments and data analysis. Only unambiguously clear fragments that were present in at least two samples were used in the analysis of RAPD and ISSR patterns. Each fragment was scored as present (1) or absent (0) in a binary matrix.
Genetic diversity within each spore populations (B+ and B–) was estimated by calculating the percentage of polymorphic bands (p.p.b.) and the Shannon-Weaver diversity index (H) (Shannon and Weaver, 1949). This index is based on number and frequency of markers in fingerprints and calculated as: H = –Σpi·log2pi, where pi is the frequency of the ith band. H was also calculated at two levels: the diversity among all spores (Hsp) and the average diversity within populations (Hpop). Then, the proportion of genetic variation between B+ and B– spore populations could be estimated by calculating (Hsp − Hpop)/Hsp.
In order to estimate the genetic divergence between B+ and B– populations by another way, an amova analysis (Excoffier et al., 1992) was made using arlequin 2.000 software (Schneider et al., 2000). amova allowed (i) to apportion the total molecular variance in among- and in within-population components and (ii) to test the significance of the partitioning.
Pairwise genetic distances were estimated as Euclidean distance (Excoffier et al., 1992). PCA based on genetic distances was performed using genalex software (Peakall and Smouse, 2002) to provide a graphical representation of the genetic relationships between all the spores.
Evaluation of mycorrhizal capacities of the cured spores
To analyse the mycorrhizal phenotype of the cured spores, different host plants were used: a dicot, Trifolium repens grown in growth chamber (16 clover plants mycorrhized with cured spores and four plants mycorrhized with WT spores), a monocot, Sorghum bicolor grown in greenhouse (24 sorghum plants mycorrhized with the cured spores and six plants inoculated with WT spores) and transformed carrot roots grown in vitro. In the sorghum experiment, the cured spores we used belong to the lots 1, 2, 3 and 4. In the clover and in the transformed carrot roots experiments, an additional lot (lot 5) of cured spores was used.
Clover and sorghum plants were inoculated with 40–60 spores and grown under greenhouse conditions for 12 weeks. The entire root was then stained overnight at room temperature in 0.1% Cotton blue (w:v) in lactic acid, and washed several times in lactic acid. Root segments (1 cm long) were randomly selected and observed under an optical microscope (Nikon Eclipse E400) for scoring mycorrhizal colonization. Four mycorrhizal variables were considered as described by Trouvelot et al. (1986). Root segments were classified according to the percentage of their length occupied by mycelium and/or arbuscules. In total, 72 m and 18 m of sorghum roots inoculated, respectively, with cured and WT spores, 157 cm and 100 cm of clover roots inoculated, respectively, with cured and WT spores, and 90 cm and 90 cm of transformed carrot roots inoculated, respectively, with cured and WT spores were carefully examined. In order to quantify the sporulation capacity, the number of new spores generated during the sorghum and transformed carrot roots experiments was also counted.
Morphological characterization of the cured spores
Light microscopy on fresh material. To check the morphology and wall organization of spores, 69 B+ spores and 144 B– spores coming from pot cultures inoculated with the five lots (1, 2, 3, 4 and 5) described in Conditions for obtaining and cultivating G. margarita-cured spore were collected, sterilized with 3% (w:v) chloramine T, 0.03% (w:v) streptomycin and mounted in a solution of Polyvinyl-Lacto-Glycerol (PVLG) prepared with 100 ml of distilled water, 100 ml of lactic acid and 16.6 g of Polyvinyl alcohol (PVA).
Light and transmission electron microscopy on embedded material. Twenty WT and 20 cured spores (15 of them originating from pot cultures inoculated with the five lots previously described and five of them directly picked from a multisporal in vitro culture) (Fig. 1, Production step) were cryo- or chemically fixed and then resin-embedded. For cryo-fixation, spores were vacuum-infiltrated in 12% Dextran water solution; each spore was then placed in an aluminium holder and frozen at high pressure using the Balzers HPM 010 apparatus. Frozen samples were stored in liquid nitrogen until further processing. Freeze substitution (FS) was performed in two FS solutions; the spores were incubated in 1% glutaraldehyde, 1% tannic acid in dry acetone at −85°C for 48–72 h, rinsed in dry, cold acetone three times, then transferred to 1% osmium tetroxide in acetone at −85°C for 1 h. To warm it up slowly, the osmium solution was held for 2 h at −20°C, 2 h at 4°C and 1 h at room temperature. After three rinses in fresh acetone, samples were infiltrated with an Epon/Araldite-acetone mixture using 10% increase of resin each step for 1 h and then polymerized at 60°C for 10 h.
For chemical fixation, spores were perforated with a sterile needle and fixed under vacuum with 2.5% glutaraldehyde in 50 mM sodium cacodylate buffer pH 7.2 for 2 h at room temperature and overnight at 4°C. Samples were then post-fixed in 1% osmium tetroxide in cacodylate buffer for 2 h, dehydrated in an acetone series and embedded in Epon/Araldite.
Thin sections (70 nm thick) were stained with uranyl acetate and lead citrate and examined with a Philips CM10 transmission electron microscope, while semi-thin sections (1 μm thick) were stained for general observation with 1% toluidine blue in 1% aqueous sodium tetraborate and observed with a Nikon Eclipse E400 optical microscope. Some sections were stained for 10 min at 50°C in 0.25% Coomassie brilliant blue in 7% acetic acid (O'Brien and McCully, 1981) to visualize proteins. Others were treated with 70% ethanol for 2 min, stained with 0.3% Sudan Black B in 70% ethanol for 1 h at 60°C and rinsed with 70% ethanol for 1 min, for lipid detection. A third group of sections was treated with 1% periodic acid for 10 min, rinsed briefly with distilled water, stained with Schiff's reagent for 30 min and rinsed twice with distilled water to evaluate the presence of polysaccharides.
Growth capacities of WT versus cured germinating spores
Spores were germinated on solid medium (0.1% w:v MgSO4; 0.4% w:v Phytagel) in vertically incubated Petri dishes (90 mm diameter or 120 × 120 mm square plates), in a 2% CO2 incubator at 28°C. Eight to 10 Petri dishes with one spore per dish were used per treatment. The experiments were performed by using cured spores belonging to four lots (lots 1, 3, 5 and 6 as previously described).
Wild-type and cured germinating spores were treated 4 days after germination with an active fraction (branching factor) of carrot root exudates prepared as described by Buée et al. (2000). The branching factor was extracted with ethyl acetate from carrot root exudate in water, dried and re-suspended in (50:50) methanol:water (Buée et al., 2000). Ten microlitres (2 × 5 microlitres) of this solution, or of methanol:water for control, were injected near the growing germ tubes. Hyphal length from germinating spores was measured under a binocular microscope by using a 2 mm grid every 2–3 days from day 4 to 30.
Under the same conditions, WT and cured germinating spores were also treated with the strigolactone analogue GR 24 at 10−7 M. After 15 days of growth, the number of hyphal branches formed by each germinating spore was recorded. Differences were tested for significance by analysis of variance and Tukey's pairwise comparison.
To further check whether the growth capacities of WT and cured germinating spores were affected by the presence of the host plant, an additional small scale in vitro experiment was set up to follow the timing of infection. Carrot roots, as described inProduction, were inoculated with B+ and B– spores and infection was followed daily. In each plate, four segments of carrot roots were placed and two spores were put under each segment. The distance between spore and root was 1.5 cm, 3 cm and 10 cm, for each experiment. Five plates with cured spores and five plates with WT spores were prepared.
The authors thanks Bachar Blal (BIORIZE, Dijon, France) for the mycorrhization experiments on Sorghum, Sara Torrielli for spore collection, Santina Buonomo and Maria Teresa Della Beffa for maintaining ROC cultures and mycorrhizal plants respectively. The authors are particularly indebted to Dr Robert W. Roberson (University of Arizona, USA) for the freeze substitution experiments performed in his laboratory and to Dr Hervé Gryta (Toulouse 3 University) for genetic variability analyses. The facilities of the Laboratory of Advanced Microscopy (University of Torino) are acknowledged, in particular Andrea Genre for confocal observations and Silvano Panero for assistance with transmission electron microscopy. This research was funded by the European Union GENOMYCA project (Project No. QLK5-CT-2000-01319), CEBIOVEM programme (D.M. 193/2003) and IPP-CNR (Biodiversity Project AGP02 n. 371–Torino). E.L. was founded by Lagrange Project – CRT Foundation with a Post-Doc Lagrange Fellow 2004.