Author’s Accepted Manuscript
The ability of plants to produce strigolactones affects rhizosphere community composition of fungi but not bacteria
Lilia Costa Carvalhais, Vivian A. Rincon-Florez, Philip B. Brewer, Christine A. Beveridge, Paul G. Dennis, Peer M. Schenk www.elsevier.com
PII: S2452-2198(18)30116-2 DOI: https://doi.org/10.1016/j.rhisph.2018.10.002 Reference: RHISPH128 To appear in: Rhizosphere Received date: 23 September 2018 Revised date: 23 October 2018 Accepted date: 23 October 2018 Cite this article as: Lilia Costa Carvalhais, Vivian A. Rincon-Florez, Philip B. Brewer, Christine A. Beveridge, Paul G. Dennis and Peer M. Schenk, The ability of plants to produce strigolactones affects rhizosphere community composition of fungi but not bacteria, Rhizosphere, https://doi.org/10.1016/j.rhisph.2018.10.002 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting galley proof before it is published in its final citable form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain. The ability of plants to produce strigolactones affects rhizosphere community composition of fungi but not bacteria
Lilia Costa Carvalhais1,2*, Vivian A. Rincon-Florez1,2, Philip B. Brewer3, Christine A.
Beveridge3, Paul G. Dennis4, Peer M. Schenk1
1School of Agriculture and Food Sciences, The University of Queensland, Brisbane, QLD
4072, Australia; 2Centre for Horticultural Science, Queensland Alliance for Agriculture Food and Innovation, Ecosciences Precinct, Dutton Park, 4001, Australia; 3School of Biological
Sciences, The University of Queensland, Brisbane, QLD 4072, Australia; and 4School of
Earth and Environmental Sciences, The University of Queensland, Brisbane, QLD 4072,
Australia
*Corresponding author: [email protected]
Abstract
Strigolactones are an important group of plant hormones. When released from roots, they act as signalling molecules that induce branching of arbuscular mycorrhizal hyphae. However, the extent to which they affect the rhizosphere microbiome is unknown. Filling this knowledge gap is important because the diversity and composition of the root-associated microbiome influence plant fitness. In this study, we hypothesised that strigolactone- producing plants harbour a different community of rhizosphere bacteria and fungi compared to plants whose strigolactone synthesis is impaired. To test this hypothesis, we compared the diversity of rhizosphere bacterial and fungal communities associated with wild-type
Arabidopsis thaliana and a mutant impaired in the production of strigolactones due to a disruption of the MORE AXILLARY GROWTH 4 (MAX4) gene. Our results indicate that the plant’s ability to produce strigolactone is significantly correlated with changes in the composition (beta diversity) of rhizosphere fungal but not bacterial communities. No differences in alpha diversity (richness and evenness) were observed for either bacterial or fungal communities between the rhizospheres of max4 and wild-type. Epicoccum nigrum,
Penicillium, Fibulochlamys chilensis, Herpotrichiellaceae, Mycosphaerella and
Mycosphaerellaceae were among the fungal taxa possibly attracted to or mostly influenced by strigolactones given that they were present at higher abundances in the rhizosphere of the wild-type compared to the mutant. Our study provides evidence that rhizosphere fungal diversity are more strongly affected than bacterial diversity by the plant’s ability to produce strigolactones.
Keywords: terpenoid lactones, strigolactones, phytohormones, fungi, bacteria, rhizosphere, diversity Introduction
Plants are sessile organisms that use a wide-range of signalling molecules to interact with other organisms and coordinate responses to environmental changes. For instance, root- released compounds can influence soil-microbe composition and soil microbes can have a promotive effect on plant growth (Carvalhais et al., 2015; Sasse et al., 2018). However, the extent to which plants interact with and derive various benefits from soil microbes may be broader than previously thought. To test this hypothesis further, we used a non-mycorrhizal plant species to explore the impact of strigolactones exuded from plant roots on the diversity of the rhizosphere microbiome. Strigolactones are carotenoid-derived molecules that play important roles in the regulation of plant development and chemical communication during biotic interactions (Brewer et al., 2013; Smith, 2014). While their first discovered role was to induce seed germination of parasitic plants (Cook et al., 1966), it was later found that they initiate and trigger symbiotic interactions between plants and arbuscular mycorrhizal fungi –
AMF (Akiyama et al., 2005; Parniske, 2008). The mutual relationship between AMF occurs with c. 80% of land plants and date from approx. 450 million years ago (Parniske, 2008).
Strigolactones have been linked to nodule initiation and rhizobacterial swarming in legumes, and interactions with bacterial, fungal and oomycetes pathogens (Akiyama and Hayashi,
2006; McAdam et al., 2017). These findings suggest that the diversity of plant-microbe interactions mediated by strigolactones is much more extensive than previously appreciated.
Functions of strigolactones in symbiotic interactions have been mostly reported for plants and
AMF. When secreted by roots, these compounds not only trigger AMF hyphal branching during the pre-symbiotic stage, but also induce spore germination and metabolism (Besserer et al., 2006; Mori et al., 2016). The hyphal network of AMF outreaches the rhizosphere, accesses nutrients and water from a greater volume of soil, and transfers these nutrients to the roots in exchange of photosynthates (Smith and Read, 2010; Yoneyama et al., 2007a). This helps plants access nutrients like phosphorus, which have limited mobility in the soil
(Yoneyama et al., 2007b). Indeed, depending on the species, strigolactone secretion from roots increases to enhance AMF symbiosis, particularly under phosphorus (P) deficiency
(Yoneyama et al., 2012). Furthermore, AMF require a component of strigolactone signalling to penetrate into roots in rice and pea and, in the case of rice, the karrikin receptor complex is also needed for perception of AMF (Yoshida et al., 2012; Foo et al., 2013; ). In contrast, high exogenous phosphate supply systemically suppresses AM colonization and symbiotic gene expression (Breuillin et al., 2010). This may be a mechanism for plants to limit symbiosis with AMF unless nutrients are low. Nonetheless, comparative transcriptomic analysis in petunia, rice, Medicago and Lotus suggested that the repressed genes encode not only carotenoid and strigolactone biosynthesis, but also proteases and phosphate transporters
(Breuillin et al., 2010). This suggests that strigolactones do not solely regulate P responses in plants. Moreover, pea mutants that were impaired in strigolactone biosynthesis and sensing were poorly colonised by AM compared to the wild-type, which also indicates that phosphate supply is not the only factor controlling mycorrhizal symbiosis (Foo et al., 2013).
Non-mycotrophic plants including Lupinus spp. (Yoneyama et al., 2008) and Arabidopsis thaliana (Goldwasser et al., 2008) also release strigolactones from roots, which indicates that these signals may be involved in processes other than the ones mediating mycorrhizal associations. For example, in legumes, strigolactones are involved in symbiosis with rhizobial bacteria. They enhance bacterial motility on surfaces, known as swarming, which facilitate the establishment of the interaction (Pelaez-Vico et al., 2016). Furthermore, endogenous strigolactones positively control nodulation of pea, alfalfa (Medicago sativa), Medicago truncatula, soybean (Glycine max) and Lotus japonicus (Foo and Davies, 2011; Liu et al.,
2013; McAdam et al., 2017; Rehman et al., 2018; Soto et al., 2010; van Zeijl et al., 2015). It is also important to note that although canonical strigolactones have been reported in Arabidopsis root exudates (Goldwasser et al., 2008), these results were never confirmed with more modern analytical chemistry systems (Yoneyama et al., 2018). Non-canonical rather than canonical strigolactones have then been proposed to act more frequently as rhizosphere signals as they seem to be more common in root exudates of various plants (Yoneyama et al.,
2018).
The involvement of strigolactones in plant defence has also been reported (Marzec, 2016;
Torres-Vera et al., 2014). Strigolactone-deficient tomato plants exhibited higher susceptibility to the necrotrophic fungi Botrytis cinerea and Alternaria alternata (Torres-Vera et al., 2014).
However, when infected with the hemibiotrophic oomycete Pythium irregulare and fungus
Fusarium oxysporum, strigolactone deficient pea showed no differences in susceptibility to disease compared with the wild-type (Abe et al., 2014; Blake et al., 2016). When using an in vitro system to examine the effect of the synthetic strigolactone GR24 on a range of phytopathogenic fungi, growth inhibition was observed for Fusarium oxysporum f. sp. melonis, Fusarium solani f. sp. mango, Sclerotinia sclerotiorum, Macrophomina phaseolina,
Alternaria alternata, Colletotrichum acutatum and Botrytis cinerea; while intense branching was observed for S. sclerotiorum, C. acutatum and F. oxysporum f. sp. melonis (Dor et al.,
2011). Despite that not all these results could be repeated (Lopez-Raez et al., 2017), these findings suggest that the feeding behaviour of the pathogen, the plant or the pathogen species affect the role of strigolactones. In regards to plant interactions with pathogenic bacteria, knowledge on the function of strigolactones is still limited. Nonetheless, strigolactone mutants of A. thaliana are hypersensive to the infections by the pathogens Pectobacterium carotovorum, Pseudomonas syringae and Rhodococcus fascians (Piisila et al., 2015; Stes et al., 2015).
Despite the aforementioned insights into the role of strigolactones in plant interactions with specific microbes, a more comprehensive analysis involving collective assemblages of plant- associated microbes still remains to be undertaken. There is a growing body of evidence that microbial communities associated with a plant improve its overall fitness through a variety of processes pertaining to nutrient acquisition and protection against biotic and abiotic stresses
(Berendsen et al., 2012; Berg et al., 2017; Lugtenberg and Kamilova, 2009). For this reason, presently plants and their associated microbiomes are often referred to as a holobiont, a single dynamic evolutionary entity which behaves as a unit throughout the process of selection and adaptation (Vandenkoornhuyse et al., 2015; Zilber-Rosenberg and Rosenberg, 2008). Here, we tested the hypothesis that strigolactone-producing plants affect the diversity of the root- associated microbiome. To this end, we used phylogenetic marker gene sequencing to characterise rhizosphere bacterial and fungal communities associated with A. thaliana wild- type Col-0 and its mutant which has the MORE AXILLARY GROWTH 4 (MAX4) gene disrupted. This gene is essential in the biosynthesis pathway of strigolactones, therefore the max4 mutant is impaired near the start of the pathway in the production of strigolactones
(Abe et al., 2014; Alder et al., 2012; Brewer et al., 2016; Sorefan et al., 2003). Because A. thaliana is a non-mycorrhizal plant, it was chosen to limit the development of a
‘mycorrhizosphere’ (the soil environment immediately surrounding AMF hyphae) and therefore exclude an additional factor influencing the rhizosphere microbiome. Arbuscular mycorrhizal mycelial exudates influence soil bacterial growth and community structure
(Toljander et al., 2007). Our results indicate that the plant ability to produce strigolactones have a more pronounced effect on the diversity of fungi compared to bacteria.
Material and Methods
Plant growth conditions Arabidopsis thaliana Col-0 and the max4 mutant plants (Sorefan et al., 2003) were grown in unfertilised free-draining 1:1 mix of medium-coarse sand and peatmoss in a controlled environment chamber (Percival Scientific, Boone, IA, USA) at 24 ºC with a light intensity of
150 µmol m-2 s-1. Elements analysis of this sand-peatmoss mix was previously described and the sufficiency rating of P confirmed to be very low to low (Loch and Zhou, 2013). Similarly to mycorrhizal hosts, P deprivation may also enhance the release of strigolactones in
Arabidopsis roots (Kohlen et al., 2011), therefore we applied P deficiency to all plants to potentiate the release of strigolactones by the wild-type plants.
Fertilization was applied using an Arabidopsis ‘standard’ nutrient solution as described in
(Tocquin et al., 2003), but phosphate was omitted from the composition. A sand-peatmoss mix was chosen as a growth substrate for plants so that we could better control that the only introduced stress was phosphate limitation and soil-borne pathogens were less likely to be present. Watering of the plants with the nutrient solution was conducted simultaneously when needed. At the 8-12 leaf stage, rhizosphere soil was collected, which was considered here as the soil attached to the roots after up-rooting. The rhizosphere soil was separated from roots by washing them off with saline water (NaCl 0.90 % w/v). Each biological replicate was composed of pooled rhizosphere soil of 40 plants. Treatments were composed of three biological replicates. Root-free soil subjected to the same nutrient regime, here referred as
‘bulk soil’, was also sampled for comparison.
Analysis of microbial diversity
DNA was extracted using the PowerSoil DNA isolation kit (MO BIO Laboratories) according with the manufacturers’ recommendations except for the following modifications: the bead beating step using a vortex was extended to 30 min, and the 4ºC incubation step after addition of Solution C3 was increased to 30 min. To verify DNA integrity and purity, electrophoresis of the extracted DNA was performed in a 1% agarose gel and 260/280 as well as 260/230 ratios were obtained by undertaking absorbance measurements at 230, 260 and 280 nm using a Thermo Scientific™ NanoDrop. DNA concentrations were measured using the Quant-iT dsDNA HS/BR assay kit in the Qubit fluorometer (Invitrogen, Waltham, MA, U.S.A.).
Bacterial profiling: DNA samples were normalised to 1 ng µL-1. Universal 16S rRNA primers, 803F (5’-CTACCRGGGTATCTAATCC-3’) and 1392wR (5’-
ACGGGCGGTGWGTRC-3’) were used to amplify a partial sequence of the bacterial phylogenetic marker gene encoding the 16S rRNA. Each PCR reaction included 2 ng of
DNA, 50 nM of each of the dNTPs (Invitrogen), 1.5 mM MgCl2 (Invitrogen), 0.3 mg bovine serum albumin (New England Biolabs, Ipswich, MA, U.S.A.), 0.02 U Taq DNA polymerase
(Invitrogen), 8 μM each of the primers, molecular biology grade water, 1× MgCL2-free PCR buffer (Invitrogen) in a total volume reaction of 50 µL. The 5’ end of the primers 1392wR and 803F included 454 FLX Titanium Lib L adapters A and B, respectively. A unique 5-6 barcode sequence was added between the reverse primer sequence and the adapter for each sample. PCR cycles comprised of 95oC for 3 min, 30 cycles of 95oC for 30 s, 55oC for 45 s,
72oC for 90 s, and a final extension step of 72oC for 1 min using a Veriti 96-well thermocycler (Applied Biosystems, Foster City, CA, USA). A QIA-quick PCR purification kit (Qiagen, Hilden, Germany) was used to purify amplification products. Amplicon concentrations were measured using a Qubit fluorometer with a Quant-iT dsDNA BR assay
-1 kit. All samples were normalised to 25 ng µL and were pooled for 454 pyrosequencing.
Sequencing was performed by the Australian Centre for Ecogenomics at the University of
Queensland (Brisbane, Australia).
Processing of bacterial sequence data: Sequences were processed as previously described
(Newsham et al., 2015). Briefly, sequences were quality-filtered and de-replicated using the
QIIME script split_libraries.py with the homopolymer filter deactivated (Caporaso et al., 2010) and were then checked for chimeras against the GreenGenes database using UCHIME ver. 3.0.617 (Edgar et al., 2011). Homopolymer errors were corrected using Acacia (Bragg et al., 2012). Sequences were then subjected to the following procedures using QIIME scripts with the default settings: i) sequences were clustered at 97% similarity, ii) cluster representatives were selected, iii) Silva taxonomy (https://www.arb-silva.de/aligner/, SILVA release 131) was assigned to the cluster representatives using BLAST, and iv) tables with the abundance of different OTU and their taxonomic assignments in each sample were generated.
The number of reads was then rarefied to 1,550 per sample.
Fungal profiling: The primers gITS7 (5′-GTGARTCATCGARTCTTTG-3′) (Ihrmark et al.,
2012) and ITS4 (5′-TCCTCCGCTTATTGATATGC-3′) (White et al., 1990) were used to amplify the internal transcribed spacer 2 (ITS2) region of the ribosomal DNA. These primers anneal to sites in the 5.8S rRNA gene and the ribosomal large subunit, respectively. A sample-specific barcode sequence was added together with the 454 FLX sequencing adaptor to the 5’ end of the primer ITS4 and the adaptor B sequence was added to the 5’end of the primer gITS7. Amplifications were carried out in duplicate in 20 µL reactions. Each reaction included 0.5 µM of the gITS7 primer, 0.3 µM of the ITS4 primer, 1X Phire Reaction Buffer
(provides a MgCl2 final concentration of 1.5 mM) (New England Biolabs), 1 U of Phire Hot
Start II polymerase (New England Biolabs), 0.2 mM of each dNTP (Invitrogen) and 0.8 ng of template DNA. The PCR cycles comprised 98 °C for 30 s, 27 cycles of 98°C for 5 s, 56°C for
20 s, 72°C for 20 s and a final extension at 72°C for 1 min. The Wizard SV Gel and PCR
Clean-Up System (Promega) was used to purify the amplicons. A Qubit fluorometer with a
Quant-iT dsDNA HF assay kit were used to measure the concentration of the amplicons after purification and 1.47 ng of each sample was pooled. A QIAquick PCR Purification Kit
(Qiagen) was used to purify the pooled sample, and an aliquot of 50 µL at 10.6 ng µL-1 was sent to Macrogen (South Korea) for 454 pyrosequencing (Roche). Processing of fungal sequence data: Sequences were processed as previously described
(Newsham et al., 2015). Briefly, quality-filtering and de-replication of sequences were performed using the QIIME script split_libraries.py with the homopolymer filter deactivated
(Caporaso et al., 2010). Correction of homopolymer errors was done using Acacia v. 1.48
(Bragg et al., 2012). Extraction of ITS2 sequences from fungi was performed with ITSx v.
1.0.9 (Bengtsson-Palme et al., 2013), and the presence of chimeras in the ITS2 sequences with UNITE v. 6 (Nilsson et al., 2015) using UCHIME v. 3.0.617 (Edgar, 2010). A minimum of 30,000 non-chimeric quality-filtered ITS2 sequences were recovered for each sample.
UCLUST v. 1.2.22 (Koljalg et al., 2013) was used to cluster the sequences at 97% similarity and Unite version 7.2 to assign taxonomy to representative OTU sequences
(https://unite.ut.ee/). The number of reads was rarefied to 30,000 per sample and abundances of different OTUs together with their taxonomic assignments were summarised in tables. The estimated total number of OTUs (Chao 1) and the average number of observed OTUs were calculated using QIIME.
Microbial diversity analysis: The mean number of OTU (observed richness, Sobs) and
Simpson’s diversity index values (Simpson, 1949) corresponding to the normalised number of sequences per sample (1,550 for bacteria; 30,000 for fungi) were calculated using QIIME.
The values of the Sobs, and Simpson’s reflect the richness (number of ‘species’) and evenness (equitability of population abundances within a sample) of microbial communities, respectively. These metrics are core attributes of alpha diversity (within sample diversity).
Differences in alpha diversity metrics were investigated using linear regression. Differences in the composition of microbial communities were investigated using redundancy analysis with subsequent Monte-Carlo permutation tests (9,999 permutations) on Hellinger transformed OTU abundances. Comparisons of relative abundances between samples for each OTU were made using one-way Analysis of Variance (ANOVA). Data were checked for normality before comparisons. If the assumption of normality was violated data were either square root or log10 transformed. When normality could not be achieved through these transformations, a Kruskal-Wallis one-way analysis of variance was done. All analyses were implemented in R using the package vegan (Dixon, 2003), except for one-way ANOVA and
Krustal-Wallis, which were performed using Minitab v16.
Results
Universal bacterial and fungal phylogenetic marker gene sequencing was used to identify rhizosphere microorganisms that potentially respond to strigolactone signals. Overall fungal communities were dominated by members of Ascomycota (76.9%) and Basidiomycota
(11.5%) (Fig. 1). Interestingly, the composition of fungal (P = 0.03), but not bacterial (P =
0.10) communities differed significantly between A. thaliana wild type and max4 rhizospheres.
For fungi, the following OTUs were the most closely associated with the wild type as evidenced in the RDA (RDA1 axis): a Fibulochlamys chilensis (OTU 39), a Talaromyces
(OTU 54), a Mycosphaerella (OTU 58), a Mycosphaerellaceae (OTU 59) and a Peziza (OTU
54) population, as well as two Gjaerumia minor populations (OTU 45 and 46) (Fig. 2). In contrast, the OTUs most closely associated with the max4 strigolactone defective mutant were members of the Plectosphaerella cucumerina (OTU 25), Fusarium (OTUs 22 and 23),
Hypocreomycetidae (OTU 19), Leotiomycetes (OTU 55), Alternaria radicina (OTU 6) and
Phialemonium inflatum (OTU 32, Fig. 2).
The fungal OTUs that were present at significantly higher abundances in the rhizosphere of the wild-type plants relative to max4 and bulk soil, included a member of Herpotrichiellaceae
(OTU 9, Fig. 1), two members of Penicillium sp. (OTUs 12 and 13), two members of
Plectosphaerella cucumerina (OTUs 26 and 28), a member of Mycosphaerella (OTU 37), a member of Mycosphaerellaceae (OTU 33), a Epicoccum nigrum (OTU 44), and two members of Fibulochlamys chilensis (OTUs 39 and 42, Fig. 1). The OTUs which were present at significantly higher abundances in the rhizosphere of max4 compared to the wild type included Geomyces sp. (OTU 1, Fig. 1), Alternaria radicina (OTU 6), a member of
Pleosporaceae (OTU 8), a member of Chaetothyriales (OTU 10), a member of Talaromyces
(OTU 14), Plectosphaerella cucumerina (OTU 25), Simplicillium lamellicola (OTU 31), another member of Cordycipitaceae (OTU 43), two members of Hypocreomycetidae (OTU
19s and 20), two members of Fusarium (OTUs 22 and 23), a member of Ramularia (OTU
30), a member of Mycosphaerella (OTU 34), a member of Pleosporales (OTU 35), two members of Gjaerumia minor (OTUs 45 and 46), two members of Fibulochlamys chilensis
(OTUs 40 and 41), and three OTUs with no BLAST hit (OTUs 47, 48, and 52, Fig. 1).
Fungal alpha diversity was measured using three different indexes: Sobs, Chao1 and
Simpson. Sobs simply accounts for the number of observed species, Chao1 is an estimator of richness and Simpson combines richness and evenness. All these indexes have shown that there were no differences in fungal alpha diversity in the rhizosphere of Arabidopsis wild- type and the mutant max4 (Supplementary Fig. 1).
Bacterial communities were dominated by representatives of Proteobacteria (54.8%),
Bacteroidetes (21.4%), Firmicutes (11.9%), Planctomycetes (4.8%), and Verrucomicrobia
(4.8%), Actinobacteria (2.4%, Fig. 3). In contrast to fungi, bacterial community composition did not differ significantly between the wild type and max4 (P = 0.10). Overall differences in composition of bacterial communities were only marginally significant (P = 0.10). Despite the fact that our 16S primers also target Archaea, archaeal populations were not detected.
Although all rhizosphere samples generated the expected amplicon, no amplification was obtained with the bacterial primers from ‘bulk soil’. Similarly to fungi, all alpha diversity indexes have shown no differences between the rhizospheres of wild type and max4
(Supplementary Fig. 2).
Discussion
Differential colonisation of the plant’s rhizosphere with fungi was observed for strigolactone compromised max4 mutants, but this was not evident for bacteria. Considering that strigolactones are involved in inducing mycorrhizal symbiosis with mycorrhizal plants under
P-limiting conditions, it seems plausible that other soil microorganisms that improve P acquisition (fungi, bacteria, archaea, protists) may be recruited via exuded strigolactones.
The ability to solubilise P is a common trait of many rhizosphere bacteria (Hayat et al., 2010;
Rodriguez and Fraga, 1999), but our data did not reveal a preferential presence of genera commonly associated with P-solubilising bacteria in wild-type compared to max4 plants (e.g.
Pseudomonas, Bacillus, Rhizobium, Burkholderia, Achromobacter, Agrobacterium,
Microccocus, Aereobacter, Flavobacterium and Erwinia).
Within fungi, an OTU affiliated to the family Herpotrichiellaceae was present at higher abundance in the rhizosphere of the wild type compared to max4. Fungi belonging to this family have been found as root endophytes in a range of plants other than Arabidopsis; however whether they confer any benefit still remains to be investigated (Narisawa et al.,
2007).
Other fungi that were present at different abundances between treatments have also been associated with plant protection. Two members of the genus Penicillum were more prevalent in the rhizosphere of the wild type compared to max4 plants and bulk soil. A range of bioactive compounds with insecticidal properties have been isolated from Penicillum strains, including okaramines, communesins, chrodrimanins (Hayashi, 2015). It is possible that strigolactones may be involved in plant defence by attracting microbes that play an antagonistic role against pests. Furthermore, three out of four OTUs affiliated to
Plectosphaerella cucumerina were present at higher abundance in the rhizosphere of the wild-type compared to max4 plants. This species of filamentous fungus can thrive on plant debris as free-living saprophytes; but can also be pathogenic to a range of plant species including Arabidopsis thaliana, tomato, and other horticultural crops (Alam et al., 2017;
Carlucci et al., 2012). Interestingly, P. cucumerina has also been isolated from healthy leaves of Orychophragmus violaceus, an edible medicinal plant from China (Zhou et al.,
2017). Furthermore, extracts of this fungus supress the formation and disrupt bacterial biofilms (Zhou et al., 2017). Intriguingly, four P. cucumerina were shown to have different abundances in rhizospheres of max4 compared to the wild-type plants. Three of them had decreased abundances in max4 compared to the wild-type and one of them displayed increased abundances in the mutant relative to the wild-type rhizospheres. This indicates that
OTUs with opposite relative abundance responses may be affiliated to distinct strains of P. cucumerina that respond differently to strigolactones. Most ascomycetes affiliated to the order Hypocreales (seven out of ten) were less abundant in the rhizosphere of the wildtype compared with max4. This order includes typical plant pathogens with wide host spectra
(Coleman, 2016; Kepler et al., 2017; Lombard et al., 2016).
The involvement of strigolactones in plant defence has been previously proposed. The strigolactone-deficient tomato mutant Slccd8 was more susceptible to the foliar pathogens
Botrytis cinerea and Alternaria alternata (Torres-Vera et al., 2014). It was hypothesised that strigolactones are involved in hormone homeostasis, and especially their interaction with the jasmonic acid signalling pathway seem to be compromised (Torres-Vera et al., 2014). Two
OTUs affiliated to the genus Mycosphaerella have opposite relative abundances in the rhizospheres of max4 and the wild-type: OTU 37 is more predominant in the wild-type and the OTU 34 in the rhizosphere of max4. This genus includes various leaf infecting pathogens, but also has many non-pathogenic members (Crous et al., 2009). This indicates that members of the same genus are commonly diverse in their lifestyles, therefore interpreting behaviours at this level is inevitably speculative.
An OTU affiliated to the genus Ramularia (OTU 30) was present at higher abundance in the rhizosphere of max4 compared to the wild-type. The genus Ramularia has over 1,200 species, the majority of which are plant pathogens that cause chlorosis, necrosis or leaf spots (Videira et al., 2016). This genus includes agriculturally important pathogens such as Ramularia collo- cygni and Ramularia beticola, which affect barley and sugarbeet, respectively (Videira et al.,
2016). Most of the Ramularia seem to have a narrow host range. However, some Ramularia have also been reported to be even mycophylic or saprophytes (Videira et al., 2016). As previously mentioned, it has been hypothesised that strigolactones are involved in plant defence (Torres-Vera et al., 2014) and this may be the reason why a potential plant pathogen is present at higher relative abundance of the strigolactone-impaired mutant max4 in comparison to the wild-type.
A member affiliated to Epicoccum nigrum (OTU 44) was more predominant in the rhizosphere of wild-type compared to max4. E. nigrum has been commonly described as a saprophyte with ubiquitous distribution (Favaro et al., 2011). This fungal species has also been described as a biocontrol agent for various diseases which affect many crops including potato, sunflower and stone fruit (De Cal et al., 2009; Li et al., 2013; Pieckenstain et al.,
2001). Its mode of action is often associated with the production of pathogen-inhibiting secondary metabolites (Alcock et al., 2015; Favaro et al., 2012). It is possible that strigolactones attract, directly or indirectly, beneficial fungi that play a role in disease suppression.
Whereas our study represents a breakthrough into the potential broader effects of SL- producing plants on root-associated microbes, there are many questions still outstanding. Despite being non-mycorrhizal, Arabidopsis plants presumably still exude non-canonical SLs from roots, and exuded non-canonical SLs are active not only because they induce plant parasitism but also stimulate hyphal branching in arbuscular mycorrhizal fungi (Goldwasser et al., 2008; Mori et al., 2016). Therefore, it is not surprising that the ability of plants to produce SLs correlates with changes in the rhizosphere microbial composition. However, the changes may not be due specifically to SLs. Discovery of the causal factor(s) will require further investigation. For example, SL-defective plants have increased auxin transport and/or levels (Arite et al., 2007; Bennett et al., 2006), which could lead to changes in soil microbial composition (Boivin et al., 2016; Fu et al., 2015). In addition, levels of flavonoids have been reported to be lower in SL mutants [70] and may affect plant-microbe interactions in the rhizosphere [71]. The effect on flavonoids was not observed in pea (McAdam et al., 2017), so may be specific to Arabidopsis. Among others, isothiocyanates can also be included as a potential causal factor for changes in microbial composition , which may be especially implicated with Brassicaceae species such as Arabidopsis (Rask et al., 2000). However, there is currently no evidence that they are altered in SL mutants (Auger et al., 2012).
It will be important to try to mimic or phenocopy SL-producing plants by treating soil with
SLs. This could also be useful in understanding side-effects of the potential use of SLs in agriculture, as has been proposed for crop treatments and eradication of parasitic weeds
(Fukui et al., 2013; Vurro et al., 2016). However, to be specifically meaningful relating to this study, it would be best to test the non-canonical strigolactones that are produced by
Arabidopsis, when these become available (Yoneyama et al., 2018). The literature points that it is highly unlikely that exogenous application of non-canonical strigolactones would reverse the effect of the strigolactone-deficient mutant because these hormones are less stable than canonical strigolactones due to degradation during purification and storage (Yoneyama et al.,
2018). Moreover, microbial responses triggered by naturally occurring strigolactones may not be activated in the same way as racemic pools of artificially-synthesised strigolactones
(Scaffidi et al., 2014). Therefore, until efficient ways of synthesising artificially non- canonical strigolactones without degradation are developed, this experiment is currently not feasible. Ultimately, we wish to know whether the performance of plants is affected by SL- dependent changes in rhizosphere microbial communities, and whether this is an important strategy of non-mycorrhizal plants for adapting to soil conditions. This will require further investigations and could be very useful knowledge for agricultural applications.
Concluding remarks
Strigolactones play a dual role in plant physiology because they behave as signals outside the plants and as a regulator of plant development and growth inside (Lopez-Raez et al., 2017).
Within the plant, strigolactones influence root system architecture by stimulating lateral root formation under phosphate limitation; however, under phosphate sufficiency strigolactones supress this process in A. thaliana (Kapulnik et al., 2011; Ruyter-Spira et al., 2011). Outside the plant, strigolactones act as a rhizosphere signal by inducing the branching of pre- symbiotic hyphae in AMF or by triggering germination of seeds of parasitic plants (Akiyama and Hayashi, 2006; Tsuchiya et al., 2015). It is well documented that strigolactones affect the growth of AMF at a cellular level (Kapulnik et al., 2011). Although there are discrepancies in these studies, strigolactones affect hyphal branching and the growth of different pathogenic fungi and oomycetes (Belmondo et al., 2017; Blake et al., 2016; Dor et al., 2011; Foo et al.,
2016; Torres-Vera et al., 2014). However, a significant difference in the composition of whole fungal communities between the rhizosphere of plants which differ in their ability to synthesise strigolactones is, to our knowledge, a new finding. This indicates that the effect of strigolactones may not be specific to AMF and points to the need of more detailed studies on the influence of strigolactones on other fungi before recommending its use for field applications (Koltai, 2014).
The use of synthetic strigolactones analogues in agriculture has also been suggested in some reports for suicidal seed bank germination of weeds such as Striga hermonthica and Striga asiatica (Kgosi et al., 2012; Zwanenburg et al., 2016). Alternatively, other agricultural practices that alter levels of strigolactones in the soil include: i) weed management strategies which include treatments with chemical compounds or fungi that decompose germination stimulants such as strigolactones (Boari et al., 2016; Kannan et al., 2015; Kannan and
Zwanenburg, 2014); ii) treatment with strigolactone derivatives which could alleviate abiotic stresses including phosphate deficiency and drought (Screpanti et al., 2016). Although field applications of strigolactones sound very promising, it is important to consider that they may affect other organisms that play key ecological roles in the soil. Our results revealed that strigolactone-producing plants have a more pronounced effect on fungal diversity than bacterial diversity compared to strigolactone-deficient plants that are impaired in producing strigolactones. This may have important consequences for the provision of ecosystem services including decomposition, nutrient acquisition and biological control (Barrios, 2007).
Chemotaxis analyses and the investigation of potential strigolactone receptors in fungi may reveal the extent and mechanisms of attraction of various fungal species. Functional studies which include culture-independent approaches such as metatranscriptomics, metaproteomics and metabolomics may help to gain novel insights into the practical implications of agricultural practices that modulate strigolactone levels in the soil.
Acknowledgements The authors are grateful to Ms Susann Aue and Mr Falk Stuermann for the technical support on plant cultivation experiments and to the Australian Research Council for the financial support (DP1094749).
Declarations of interest: none
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Figure legends
Figure 1. Heatmap summarising variation in the composition of rhizosphere fungal communities associated with Arabidopsis thaliana wild type and the strigolactone-deficient mutant max4. Operational taxonomic units (OTUs) were assigned unique numbers that are also referred to in the text. The OTUs listed are those present at ≥ 1% average relative abundance in any plant genotype. Each column represent a biological replicate.
Figure 2. Redundancy analysis (RDA) ordination summarising the maximum variation between genotypes and bulk soil in two dimensions and depicting the most discriminating
OTUs along the primary and secondary axes.
Figure 3. Heatmap summarising variation in the composition of rhizosphere bacterial communities associated with Arabidopsis thaliana wild type and the strigolactone-deficient mutant max4. Operational taxonomic units (OTUs) were assigned unique numbers that are also referred to in the text. The OTUs listed are those present at ≥ 1% average relative abundance in any plant genotype. Each column represent a biological replicate. Bulk soil was not included in the heatmap because no PCR products were obtained for the bacterial 16S rRNA gene.
Captions Supplementary Figures:
Supplementary Figure 1. Alpha diversity indexes of fungal communities in bulk soil, and in the rhizosphere of Arabidopsis thaliana wild-type and the mutant max4
Supplementary Figure 2. Alpha diversity indexes of bacterial communities in bulk soil, and in the rhizosphere of Arabidopsis thaliana wild-type and the mutant max4