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Nitric oxide is required for an optimal establishment of the Medicago truncatula–Sinorhizobium meliloti symbiosis
Article first published online: 1 APR 2011
© 2011 The Authors. New Phytologist © 2011 New Phytologist Trust
Volume 191, Issue 2, pages 405–417, July 2011
How to Cite
del Giudice, J., Cam, Y., Damiani, I., Fung-Chat, F., Meilhoc, E., Bruand, C., Brouquisse, R., Puppo, A. and Boscari, A. (2011), Nitric oxide is required for an optimal establishment of the Medicago truncatula–Sinorhizobium meliloti symbiosis. New Phytologist, 191: 405–417. doi: 10.1111/j.1469-8137.2011.03693.x
- Issue published online: 29 JUN 2011
- Article first published online: 1 APR 2011
- Received: 23 November 2010, Accepted: 3 February 2011
- nitric oxide;
- nitrogen fixation;
- nodule initiation and organogenesis;
- Rhizobium legumes symbiosis
- Top of page
- Materials and Methods
- Supporting Information
- •Nitric oxide (NO) is a gaseous molecule that participates in numerous plant signalling pathways. It is involved in plant responses to pathogens and development processes such as seed germination, flowering and stomatal closure.
- •Using a permeable NO-specific fluorescent probe and a bacterial reporter strain expressing the lacZ gene under the control of a NO-responsive promoter, we detected NO production in the first steps, during infection threads growth, of the Medicago truncatula–Sinorhizobium meliloti symbiotic interaction. Nitric oxide was also detected, by confocal microscopy, in nodule primordia.
- •Depletion of NO caused by cPTIO (2-(4-carboxyphenyl)-4,4,5,5-tetramethyl imidazoline-1-oxyl-3-oxide), an NO scavenger, resulted in a significant delay in nodule appearance. The overexpression of a bacterial hmp gene, encoding a flavohaemoglobin able to scavenge NO, under the control of a nodule-specific promoter (pENOD20) in transgenic roots, led to the same phenotype. The NO scavenging resulting from these approaches provoked the downregulation of plant genes involved in nodule development, such as MtCRE1 and MtCCS52A. Furthermore, an Hmp-overexpressing S. meliloti mutant strain was found to be less competitive than the wild type in the nodulation process.
- •Taken together, these results indicate that NO is required for an optimal establishment of the M. truncatula–S. meliloti symbiotic interaction.
- Top of page
- Materials and Methods
- Supporting Information
Nitric oxide (NO) is a gaseous signalling molecule that has been established as a major signal in mammals (Ignarro, 2000). The participation of NO in a large number of plant signalling pathways is now well established (Grun et al., 2006) and there is increasing evidence of its role in plant growth and development (del Rio et al., 2004; Delledonne, 2005). Its involvement in seed dormancy (Bethke et al., 2006), germination and hypocotyl elongation (Beligni & Lamattina, 2000), flowering (He et al., 2004) and regulation of stomatal closure (Neill et al., 2002) has also been demonstrated, although the pathway of its synthesis in plants is still a matter of debate (Moreau et al., 2008).
The importance of NO in plants emerged from pioneering works on plant responses to pathogens (Delledonne et al., 1998; Durner et al., 1998). Nitric oxide signalling in the induction of cell death, defence genes and interaction with reactive oxygen species (ROS) during plant defence against pathogen attack is well documented (Neill et al., 2003; Delledonne, 2005; Mur et al., 2006; Besson-Bard et al., 2008). Nitric oxide not only plays an important role in the plant hypersensitive response (HR), a localized programmed cell death (PCD) that confines the pathogen to the site of attempted infection, but also in non PCD defence (Mur et al., 2006). Conversely, there is also evidence of its potential importance within pathogens. Detoxification of NO in bacteria has been intensively investigated. Under aerobic conditions, it occurs through the nitric oxide dioxygenase activity of the flavohaemoglobin, which converts it to nitrate (Gardner et al., 1998; Poole & Hughes, 2000). Bacterial mutants lacking flavohaemoglobin are hypersensitive to NO (Poole & Hughes, 2000) and flavohaemoglobins are important for animal pathogens to survive host-derived NO (Stevanin et al., 2002; Gilberthorpe et al., 2007; Svensson et al., 2010). Examining the genomes of several bacterial phytopathogens has revealed the conservation of the hmp gene (Mur et al., 2006). Furthermore, mutation of the hmpX gene from Dickeya dadantii (formerly Erwinia chrysantemi) resulted in a significant increase in NO concentrations and led to a resistance response of the HR type. Conversely, expression of hmpX into Pseudomonas syringae pv. tomato avrB suppressed the HR elicited in Arabidopsis (Boccara et al., 2005). Such data demonstrate that the scavenging of NO by certain pathogens is a requirement for successful infection.
Contrary to pathogenic situations, the interaction between legumes and soil bacteria of the Rhizobiaceae leads, after extensive recognition by both partners, to the establishment of a symbiotic relationship characterized by the formation of new differentiated organs called nodules, which provide a niche for bacterial nitrogen fixation. Functional nodules result from the combination of developmental and infectious processes; bacteria released in plant differentiate into bacteroids with the unique ability to fix atmospheric nitrogen via nitrogenase activity (Long, 2001; Oldroyd & Downie, 2008).
Several studies have demonstrated the occurrence of NO production during legume–Rhizobium symbiosis. A nitric oxide synthase-like activity has been measured in lupine nodules (Cueto et al., 1996). Nitric oxide complexed to leghaemoglobin (Lb), the haemoprotein ensuring O2 fluxes to bacteroids, has been detected in soybean nodules using electron paramagnetic resonance (EPR) (Mathieu et al., 1998; Meakin et al., 2007). This nitrosylleghaemoglobin (LbNO) was mainly observed in young nodules, which led the authors to propose a relationship between LbNO abundance and nodule development and/or functioning (Mathieu et al., 1998). Production of NO during the early steps of legume–rhizobia symbiosis has also been reported by Shimoda et al. (2005) and NO, together with auxin, has been shown to control indeterminate nodule formation (Pii et al., 2007). In the same way, gene expression of non-symbiotic haemoglobin (Hb) in Lotus japonicus has been shown to be transiently induced by the inoculation with the symbiotic bacteria and by an NO donor (Shimoda et al., 2005; Nagata et al., 2008). Although the functions of class 1 Hbs are not fully understood, they appear to play some role in relation to NO metabolism (Igamberdiev & Hill, 2004) and in the establishment of the symbiosis (Nagata et al., 2008). Furthermore, modulation of the NO level by overexpression of class 1 plant Hb genes appeared to enhance symbiotic nitrogen fixation activity between L. japonicus and Mesorhizobium loti (Shimoda et al., 2009). Based on the use of a permeable NO-sensitive fluorescent probe, NO formation has been detected in bacteroid-containing cells of the nodule fixation zone in Medicago truncatula–Sinorhizobium meliloti functional nodules (Baudouin et al., 2006). A wide modulation of NO-responsive genes has been detected during the establishment of a M. truncatula–S. meliloti functioning nodule, pointing to a possible contribution of NO to nodule metabolism (Ferrarini et al., 2008). Moreover, the response to NO of S. meliloti has been studied recently via a transcriptomic approach. Among genes responding to NO, was a flavohaemoglobin-encoding gene. An hmp mutant displayed a higher sensitivity toward NO in culture and led to reduced nitrogen fixation efficiency in planta (Meilhoc et al., 2010). Together, these data indicate that NO is produced at different steps of the symbiotic process, where it may play different roles.
In this work, using pharmacological and genetic approaches, we investigated the presence of NO and its possible role(s) in the first steps of the M. truncatula–S. meliloti symbiosis. Our data suggest that NO is required for an optimal establishment of the symbiotic interaction.
Materials and Methods
- Top of page
- Materials and Methods
- Supporting Information
Bacterial strains and growth conditions
Bacterial strains and plasmids are listed in Table 1. Escherichia coli strains were propagated in Luria–Bertani (LB) medium. Sinorhizobium meliloti strains were constructed and grown in LB medium supplemented with 2.5 mM CaCl2 and 2.5 mM MgSO4 (LBMC). Antibiotics, when required, were added at the following concentrations: streptomycin 100–300 μg ml−1, tetracycline 5–10 μg ml−1, gentamycin 40 μg ml−1, carbenicillin 50 μg ml−1, kanamycin 50 μg ml−1.
|Strain or plasmid||Description||Reference or source|
|GMI11495||wt strain, SU-47 derived, SmR||Pobigaylo et al. (2008)|
|RCR2011||wt strain, SU-47 derived||Rosenberg et al. (1981)|
|RCR2011-DsRed||RCR2011 containing pDG77||E. Andrio, unpublished|
|GMI11549||GMI11495 hmp :: mTn5-STM-2.12.A01||A. Becker, see Pobigaylo et al. (2008)|
|CBT515||Rm1021 nnrR :: pVO155||Meilhoc et al. (2010)|
|GMI11545||GMI11495 containing pXLGD4||D. Capela, unpublished|
|CBT602||GMI11495 containing pBBR-hmp||Meilhoc et al. (2010)|
|DH5α||supE44ΔlacU169 (Φ80 lacZΔM15)||Sambrook et al. (1989)|
|hsdR17 recA1 endA1 gyrA96 thi-1 relA1|
|pRK2013||Helper plasmid for triparental matings||Figurski & Helinski (1979)|
|pXLGD4||Ditta et al. (1985)|
|pGEM-T||Cloning vector, AmpR||Promega|
|pBBR1MCS-5||Cloning vector, GmR derivative of pBBR1||Kovach et al. (1995)|
|pVO155||Integrational plasmid with promoterless uidA gene, KanR||Oke & Long (1999)|
|pBBR-hmp||pBBR1MCS-5 + hmp||Meilhoc et al. (2010)|
|pDG77||Expresses DsRed under Salmonella typhimurium pTrp promoter||Bringhurst et al. (2001)|
|pCZ962||Derived from pCZ917 cloning vector by insertion of a terminator upstream from lacZ TcR AmpR||C. Zischek et al., unpublished|
|psma1289-lacZ||sma1289 promoter in pCZ962 cloning vector with promoterless lacZ gene||This work|
To test the bacterial response to NO, S. meliloti was grown in Vincent minimal medium (VMM: 7.35 mM KH2PO4, 5.74 mM K2HPO4, 1 mM MgSO4, 18.7 mM NH4Cl, 10 mM Na2 succinate, 456 μM CaCl2, 35 μM FeCl3, 4 μM biotin, 48.5 μM H3BO3, 10 μM MnSO4, 1 μM ZnSO4, 0.5 μM CuSO4, 0.27 μM CoCl2, 0.5 μM NaMoO4, pH = 7) at 28°C. The NO donor Spermine NONOate (SpNN) was purchased from Cayman Chemicals Coger (CAY-82150-M100, Paris, France). A 100 mM stock solution was prepared, just before use, in Na-phosphate buffer (0.1 M) pH 6.9 and then diluted in the cell culture medium at the appropriate concentration.
Plants and growth conditions
Medicago truncatula cv Jemalong A17 was used, and was cultivated as follows. Seeds of M. truncatula cv Jemalong A17 were scarified with H2SO4, surface-sterilized in a bleach solution, rinsed with sterile distilled water, germinated on agar plates in the dark and allowed to grow on nitrogen-free Farhäeus medium in test tubes or plates (covered with pouch paper) for 2 to 7 d before inoculation in a culture room (22–25°C) with a 16 h light/8 h dark photocycle (all details and medium used are described in the Medicago Handbook: http://www.noble.org/MedicagoHandbook/). For NO detection with 4,5-diaminofluorescein (DAF-2, Sigma-Aldrich) (Figs 1, 4), transgenic roots (Fig. 6), and quantitative reverse-transcription polymerase chain reaction (qRT-PCR) experiments (Fig. 7), S. meliloti strains were grown in liquid LBMC medium, cells were pelleted at 2500 g, washed twice with sterile distilled water, resuspended in sterile distilled water to a final optical density at 600 nm (OD600) of 0.01, and plants were inoculated with 200 μl of bacterial suspension per root. In all other experiments, S. meliloti strains were grown on LBMC plates, cells were resuspended in sterile distilled water to a final OD600 of 0.001, and plants were inoculated with 100 μl of bacterial suspension per root. When 2-(4-carboxyphenyl)-4,4,5,5-tetramethyl imidazoline-1-oxyl-3-oxide (cPTIO; Sigma) was used, 200 μl of a 1 mM solution prepared in sterile water was added along the whole length of the roots, 2 h before, and 2 h after inoculation, and then, every 24 h during 4 d.
For competition assays, plants were inoculated with a 1 : 1 mixture of wild-type (wt) and mutant strains (OD600 = 0.001). In order to verify that the number of cells used for inoculation with each strain were identical, CFUs were determined for each strain. To determine the relative proportions of nodules occupied by mutant or wt bacteria, the wt strain contained a plasmid (pXLGD4) constitutively expressing the β-galactosidase enzyme. After 3 wk, nodules were wounded and incubated (1 h) in Z′ buffer (0.1 M Na2HPO4/NaH2PO4 pH 7.4, 10 mM KCl, 1 mM MgSO4) containing 1.5% of glutaraldehyde under vacuum. Nodules were then washed three times in Z′ buffer and incubated in the same buffer containing potassium ferricyanide (5 mM) and X-gal (0.8% in dimethylformamide). Incubation was performed under vacuum at room temperature (90 min) and then at 37°C (90 min). Blue-stained nodules corresponding to nodules infected by GMI11545 (wt) and pink nodules corresponding to nodules infected by the mutant strain were counted. For each competition test, 50 –100 nodules from 10–20 plants were analysed, and the mean value of the percentage of nodule occupancy was calculated for each strain. When cPTIO was used in the test, nodules were taken at 4 wk instead of 3 wk after inoculation because of the delayed nodulation.
The binary vector pKm43GW-rolD::GFP (Van de Velde et al., 2006) was modified by replacing the green fluorescent protein (GFP) with a red fluorescent protein (RFP) reporter cassette under the control of the rolD promoter to facilitate the analysis by confocal microscopy with the cell permeable 4,5-Diaminofluorescein diacetate (DAF-2DA) fluorescent probe (excitation 495 nm, emission max c. 515 nm; Sigma-Aldrich) and the selection of transgenic roots. The RFP cassette was amplified by PCR from the pK7RWG2 vector (http://gateway.psb.ugent.be) with the primers DsRedX and DsRedS containing, respectively, a XhoI and SacI restriction site and cloned in the pKm43GW-rolD by using these two restriction sites to get the pKm43GW-rolD::RFP.
The promoter from MtENOD20 (Accession: AC136953.17; GenBank accession no. X99467) (2050 bp) was amplified with the specific primers a74F and a74R containing the corresponding attB recombination sites. The promoter region was recombined into pDONR-P4-P1 (Invitrogen). The Multisite Gateway Three-Fragment Vector Construct kit (Invitrogen) was used to fuse the promoter region with the GUS gene (from pDONR207-GUS) or the hmp gene (from pDONR207-hmp) and the T35S terminator (from pENTR-R2-T35S-L3) in the pKm43GW-rolD-RFP vector. The hmp gene was amplified from S. meliloti genomic DNA with the specific primers a76F and a76R.
The bacterial NO biosensor was constructed as follows. The promoter of the S. meliloti gene sma1289 was amplified by PCR with the following primers: OCB744 and OCB745 containing the PstI and XbaI restriction sites, respectively. The PCR amplification was performed using genomic DNA of strain GMI11495 as a template. The amplified fragment (410 nucleotides) was ligated in pGEM-T (Promega). The cloned S. meliloti DNA region was verified by DNA sequencing. The pGEM-T plasmid containing the sma1289 promoter was then digested with XbaI and PstI, and ligated into the XbaI and PstI digested pCZ962 plasmid containing the promoterless lacZ gene. Plasmids (pCZ962 with or without the sma1289 promoter) were introduced in S. meliloti strains (GMI11495, CBT515, CBT602) by tri-parental mating using pRK2013 as a helper, and subsequent selection for antibiotic resistance.
β-galactosidase activity assay
Sinorhizobium meliloti cells were grown to exponential phase at 28°C in VMM. At OD600 = 0.2, Spermine NONOate (0 or 25 μM) was added to the cultures which were then incubated at 28°C. After 1.5 h incubation, OD600 was measured and aliquots (1 ml) of the cultures were centrifuged. The pellets were immediately frozen in liquid nitrogen and stored at −20°C. β-Galactosidase assays were performed on the thawed samples as described previously (Miller, 1972).
β-galactosidase and β-glucuronidase detection in planta
For β-galactosidase detection, plants were inoculated with strains GMI11495 (wt), CBT515 (nnrR), or CBT602 (hmp++) carrying the plasmid pSma1289–lacZ or with strain GMI11495 carrying the empty plasmid. Six days after inoculation, plant roots were incubated (1 h) under vacuum, in Z′ buffer containing 1.5% glutaraldehyde. Roots were then washed three times in Z′ buffer and incubated (room temperature) in Z′ buffer containing potassium ferricyanide (5 mM) and X-gal (0.8% in dimethylformamide) for 90 min or for a longer period when the staining was too weak.
For β-glucuronidase detection, roots 5 d post-inoculation (dpi) and 13 dpi transformed with pKm43W-pENOD20-GUS were incubated for 1 h in acetone 90% (diluted in a phosphate buffer: Na2HPO4, NaH2PO4 0.1M pH = 7.4) at −20°C. Roots were then washed twice in the phosphate buffer and incubated at room temperature, protected from light, in phosphate buffer containing potassium ferricyanide (0.5 mM) and X-gluc (0.5 ng ml−1) for 3 h and 16 h.
The roots were observed by optical microscopy (Axioplan imagin2; Zeiss). Pictures were taken with an AxioCam (Zeiss) and acquired with the corresponding software.
Agrobacterium rhizogenes root transformation and inoculation
The construct pKm43GW-pENOD20-hmp was introduced into Agrobacterium rhizogenes strain Arqua1 (Quandt et al., 1993). Medicago truncatula plants were transformed with Agrobacterium rhizogenes, as described by Boisson-Dernier et al. (2001). Control plants were transformed with A. rhizogenes containing the pKm43GW-pENOD20-GUS. Selection of hairy roots took place 21 d after transformation. Plants were screened for transgenic roots by RFP with a MZFLII stereomicroscope (Leica, Wetzlar, Germany). One transgenic root per plant was retained, and the composite plants were transferred onto agar plates containing modified Fahräeus medium without nitrogen. Plants were inoculated 3 d after transfer.
Measurement of NO production
The NO produced by both the wt and transgenic hairy roots, and released into the detection medium, was measured using the fluorescent probe DAF-2. At various times, the fluorescence of DAF-2T, the reaction product formed from DAF-2 and NO, was measured using a microplate reader spectrofluorimeter (Cary Eclipse; Varian, Les Ulis, France), excitation wavelength 495 nm/emission wavelength 515 nm. Plants were pooled into groups of seven to nine plants, and the roots were put into 3 ml of detection medium: 10 mM Tris-KCl detection buffer (DB: 10 mM Tris-HCl pH 7.4, 10 mM KCl) in the presence of 10 μM DAF-2 fluorescent probe. The roots were protected from light with tinfoil while the leaves were left in contact with air and light, allowing the photosynthesis to take place. Production of NO was measured at 10, 30, 60 and 90 min after the addition of the DAF-2 probe into the detection medium. Blank assays contained detection medium without plants.
Experiments were performed on young entire roots and fresh nodule slices (100 μm) obtained with a vibratome 1000 Plus (Labonord, Templemars, France). Plant material was then incubated for 30 min in the dark in 100–500 μl of DB containing 10 μM DAF-2DA the cell permeable analog of DAF-2 probe (Sigma-Aldrich). Plant tissues were subsequently washed for 30 min with DB and then mounted between slide and coverslip in DB for observation. When cPTIO was used, plant tissues were preincubated for 45 min in DB containing 1 mM of cPTIO, subsequently labelled with DAF-2DA and washed three times during 20 min with DB. The formation of DAF-2T following the NO reaction with DAF-2DA was visualized using a Zeiss LSM 500 confocal laser microscope upon excitation at 488 nm with an argon 2 laser. Dye emission was recorded using a 505–530 nm band-pass filter coupled with a 515 nm long-pass filter. Autofluorescence was judged negligible at all steps before acquisition with the DAF-2DA probe. Data acquisition settings (laser power, pinhole size, filters, detectors settings and scan conditions) were identical in all experiments. Images were processed and analysed using the Zeiss LSM510 meta software and Adobe Photoshop.
Total RNA isolation, reverse transcription and gene expression analysis
For the cPTIO treatment used to identify NO-regulated genes, plantlets were grown on plates and then, at 4 d post-inoculation, were treated with 200 μl of 1 mM of cPTIO for 8 h. A control assay was not treated with cPTIO. Furthermore, in order to obtain a higher concentration of RNA from nodule primordia, a 2 cm region of the root in which most of the nodule primordia appear was harvested. To localize this root infection zone (IZ), we marked the position of the primary root apex on the day of the inoculation. Four days later, the IZ was mainly located around this mark.
Aproximately 100–200 mg of plant material (root) were ground in liquid nitrogen and total RNA was isolated using Trizol Reagent (Invitrogen). The integrity of total RNA was checked on agarose gels and its quantity as well as purity was determined spectrophotometrically. About 500–1500 ng of RNA was used as a template for reverse transcription reaction in 20 μl reaction volume using the Omniscript RT Kit (Qiagen). Quantitative real-time RT-PCR was carried out using the qPCR Mastermix Plus for SYBR Green I reagent (Eurogentec, http://www.eurogentec.com). Reactions were run on the Chromo4 Real-Time PCR Detection System (Bio-Rad) and quantification was performed with the Opticon Monitor analysis software v. 3.1 (Bio-Rad). Each reaction was set up in three technical replicates. For each reaction, 5 μl of 100-fold-diluted cDNA and 0.3 μM primers were used. The initial denaturing time was 10 min, followed by 40 PCR cycles at 95°C for 10 s, and 60°C for 1 min. The specificity of the amplification was confirmed by a single peak in a dissociation curve at the end of the PCR reaction. Data were quantified using Opticon Monitor 2 (MJ Research, Waltham, Massachusetts, USA) and analysed with RqPCRBase, an R package working on R computing environment for analysis of quantitative real-time PCR data (T. Tran & F. Hilliou, unpublished). The mRNA levels were normalized against three constitutively expressed endogenous genes, Mtc27 (Mtr.27459.1.S1_s_at) (Van de Velde et al., 2006) and two genes (Mtr.10324.1.S1_at; Mtr.31250.1.S1_at) selected as reference genes because of their stable expression profile in different microarray results (Benedito et al., 2008). The PCR reactions for each biological replicate were performed in triplicate. For each experiment, the stability of the reference genes across samples was tested using genorm software (Vandesompele et al., 2002). The absence of genomic DNA contaminations in the RNA samples was tested by PCR analysis of all samples using oligonucleotides bordering an intron in the glutathione synthetase gene of M. truncatula. The gene-specific primers used are listed in the Supporting Information, Table S1.
- Top of page
- Materials and Methods
- Supporting Information
NO is produced in infection threads and nodule primordia
To detect NO during the early steps of the M. truncatula–S. meliloti interaction, we used the cell-permeable NO-specific fluorescent probe, DAF2-DA (Kojima et al., 1998). Between 2 dpi and 6 dpi, the formation of the fluorescent triazol derivative, DAF-2T, was visualized in infection threads (ITs). For all these experiments, roots were inoculated with a S. meliloti DsRed strain harbouring the pDG77 vector, which exhibits a constitutive red fluorescence. A yellow colour from the merging of the two fluorescent markers occurred (Fig. 1a–d) in the infection pockets, where rhizobia are enclosed within an apoplastic space created by root hair tip curling (the so-called ‘shepherd’s crooks’) upstream of the IT. Interestingly, the red fluorescence from the bacteria and the green fluorescence from the DAF probe did not overlap along the ITs (Fig. 1c–e). On a higher magnification (Fig. 1e), the DsRed-labelled bacteria can be visualized in the centre of the IT surrounded by the green fluorescence. In Fig. 1(c) and (d), we can observe the position of the nucleus close to the IT tip indicating that the IT is growing as mentioned by Fournier et al. (2008). To validate the specificity of the green fluorescence observed, the roots were treated with the NO scavenger (cPTIO). The cPTIO treatment extinguished the fluorescence and further established that it was representative of NO production (Fig. 1f).
To confirm the presence of NO in the infection threads of M. truncatula root hairs, and to ascertain whether bacteria respond to the presence of NO in this environment, we constructed a reporter plasmid (psma1289-lacZ) carrying a transcriptional lacZ fusion to the S. meliloti sma1289 gene promoter. The sma1289 gene was chosen since it is known to be upregulated by NO via the NO-specific regulator NnrR (de Bruijn et al., 2006; Meilhoc et al., 2010). As expected, in free-living conditions, expression of the reporter fusion, assessed by measuring β-galactosidase activity, was upregulated approx. 15-fold in the presence of the NO donor spermine NONOate (25 μM) in the wt strain. By contrast, no significant upregulation of the reporter fusion was observed in an nnrR mutant, or in a wt strain carrying the empty vector pCZ962 (Fig. 2). Thus, this strain was used as a NO biosensor strain. The bacteria carrying the reporter fusion were used to inoculate M. truncatula plantlets, and expression of the fusion was assessed, 6 d post-inoculation, by X-Gal staining and microscopic analysis of the roots. Blue-stained bacteria entrapped in ‘shepherd’s crooks’ of root hairs (Fig. 3) were detected on roots inoculated with the wt strain, whereas no staining was observed with the nnrR mutant or with the wt strain carrying the empty vector pCZ962 (data not shown). Also, no staining was detected when we used a S. meliloti strain overexpressing the flavohaemoglobin Hmp from plasmid pBBR-MCS5 (Meilhoc et al., 2010). Together, these results confirm that NO is present in the infection pockets and infection threads, and indicate that bacteria respond to NO in infection pockets.
The DAF-2T fluorescence was also observed 4 dpi in the bumps of the root infection zone in the region of infection thread progression. To more precisely localize this NO production, we carried out confocal microscopic analysis on fresh root slices (100 μm) from the infection zone (Fig. 4). All the pictures showed the fluorescence in dividing cortical cells of the nodule primordia (Fig. 4a,b). At the same time, we also detected NO production in lateral root primordia (data not shown), as described in lateral root development in tomatoes (Correa-Aragunde et al., 2004). As expected, this green fluorescence was significantly diminished by treating root slices with cPTIO (Fig. 4c,d). These results show that NO is present in the early stages of the symbiotic interaction.
NO is necessary for efficient competitiveness of S. meliloti
To determine whether the NO detected at early steps of symbiosis is involved in the infection process, we used an approach to deplete the NO levels. For this, we first tested the ability of the S. meliloti hmp++ strain to compete with the wt strain for nodule occupancy. Medicago truncatula plants were coinoculated with a mixture (1 : 1) of the wt and hmp++ strain. To differentiate nodules occupied by each strain, the wt harboured plasmid pXLGD4, which displays a constitutive and strong β-gal activity. Nodules occupied by the wt and hmp++ strains could thus be numbered after addition of X-Gal by counting blue and white nodules, respectively. In control experiments, the wt strain harbouring pXLGD4 was mixed (1 : 1) with either the empty wt strain, or the wt strain carrying the empty vector pBBR-MCS5.
The competition experiment revealed a lower competitiveness of the hmp++ strain compared with the wt (31.6% vs 68.4% nodules occupancy, respectively; P < 0.05 according to Student’s t-test) whereas in control experiments, all strains were as competitive as the wt (Table 2). Even when the competition experiment was done with a 1 : 3 mixture of the wt vs hmp++, the hmp++ strain still remained significantly less competitive (36.5%, n = 1). It has been shown that overexpression of hmp in bacteria can lead to elevated concentrations of superoxide and peroxide (Gilberthorpe et al., 2007; McLean et al., 2010). To rule out the possibility that such an effect is responsible for the lower competitiveness of the hmp++strain, the coinoculation of wt and hmp++ strains was repeated in the presence of the NO scavenger (cPTIO). In these conditions, the wt and hmp++ strains recovered a similar competitiveness. This confirmed that the low competitivity of the hmp++ strain was indeed the result of its greater ability to degrade NO. These results therefore indicate that NO production has a positive effect on the early steps of bacterial infection.
|Strain testeda||cPTIO treatment (1 mM)||Per cent of nodule occupancy (± SE)||Total number of nodules (number of experiments)|
|wt||−||51.3 (±2.3)||136 (2)|
|wt (pBBR-MCS5)||−||47.0 (±0.3)||121 (2)|
|hmp++||−||31.6 (±4.9)||481 (6)|
|hmp++||+||51.2 (±2.1)||183 (3)|
NO is necessary for optimal nodule formation
To investigate the role of NO in the establishment of the symbiotic interaction, we first tested the effect of the NO scavenger cPTIO on the nodulation ability of M. truncatula plants inoculated by the S. meliloti wt strain. The plants grown in tubes were treated by addition of 1 mM cPTIO on the whole length of the root 2 h before and 2 h after inoculation, and then every 24 h for 4 d. Appearance of nodules was then monitored. The results show that the NO depletion caused by cPTIO treatment led to a significant delay (3–4 d) in nodule appearance (Fig. 5a). Similarly, plant inoculation with the hmp++ strain led to a slight but significant delay in nodule formation (Fig. 5b).
To corroborate these results, transgenic hairy roots overexpressing the bacterial hmp gene, encoding a flavohaemoglobin able to scavenge NO, were obtained. To avoid any effect on root development, the hmp gene was expressed under the control of a nodule specific promoter pMtENOD20. As described in the literature (Vernoud et al., 1999), we observed the expression of the pMtENOD20-GUS construction in dividing inner cortical cells corresponding to the site of nodule primordium from the fifth day of nodule development (see the Supporting Information, Fig. S1). However, compared with the results of Vernoud et al. (1999), we did not observe glucuronidase (GUS) staining in associated root hairs displaying shepherd’s crook curling. Detection of NO by fluorescent microscopy with DAF2-DA revealed a decrease of fluorescence on the Hmp-overexpressing transgenic nodule slices compared with the control transgenic nodules (Fig. 6a). Using a spectrofluorometric method, we estimated a 40% reduction of the NO production in hairy roots overexpressing the Hmp protein compared with the control hairy roots at 5 dpi (Fig. 6b) as well as at 13 dpi (data not shown). In addition, the overexpression of hmp under the control of this promoter significantly delayed the nodulation (Fig. 6c), with a 28% reduction in the number of nodules compared with the control transgenic roots. This result is in line with the results demonstrated with cPTIO treatment (Fig. 5a) or when plants were inoculated with the hmp++S. meliloti strain (Fig. 5b). These results show that decreasing the NO level results in a delay in the nodulation process.
Depletion of NO affects transcript level of genes involved in nodulation process
To further investigate the involvement of NO in this process, we measured the expression levels of genes known to be associated with nodule development and potentially regulated by NO. We first analysed the expression of MtCRE1, a gene encoding a cytokinin receptor involved in the early symbiotic interaction (Gonzalez-Rizzo et al., 2006) as it was shown to be NO regulated in M. truncatula (Ferrarini et al., 2008). The qRT-PCR experiments performed either on inoculated plants treated with cPTIO (during 8 h) or on transgenic hairy roots overexpressing the Hmp protein (Fig. 7a,b) showed that NO depletion downregulates the expression of MtCRE1. This result therefore confirms that NO upregulates the expression of MtCRE1, and further points to the presence of NO during the early steps of nodule development.
We then measured the expression levels of several well-studied genes associated with various infection and nodule developmental stages: MtRR4, MtNIN, MtN6, MtENOD40 and MtCCS52A (Pichon et al., 1992; Charon et al., 1997; Cebolla et al., 1999; Vinardell et al., 2003; Gonzalez-Rizzo et al., 2006). Only one of them, MtCCS52A, displayed a lower expression level upon NO depletion by either cPTIO or Hmp overexpression (Fig. 7 and data not shown) suggesting that the gene expression could be induced by the NO production observed at this stage. During nodule organogenesis, the cell cycle-switching gene MtCCS52A triggers selected cells within the primordium to switch from mitotic cycles into endoreduplicating cycles (Cebolla et al., 1999; Vinardell et al., 2003). These data therefore strengthen the role of NO in nodule development.
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In previous work, we showed that NO is present in late stages of nodule development (Baudouin et al., 2006). The results presented here clearly show that NO is also present in the first steps of the M. truncatula–S. meliloti symbiosis, that is, in the root hair infection pockets as well as in the nodule primordium, clearly pointing to a role of this reactive nitrogen species in the early exchange of signals between the two partners. They are in agreement with previous reports of NO detection on M. sativa or L. japonicus roots a few hours after inoculation by the respective microsymbionts and further indicate that NO production occurs at different steps during the nodulation process (Shimoda et al., 2005; Pii et al., 2007; Nagata et al., 2008). Furthermore, in each step the NO observed might have specific functions.
In order to evaluate the role of NO, a strategy could have been to diminish the NO production and analyse the consequences on the symbiotic steps. However, although numerous studies demonstrate the importance of NO in plants, NO synthesis pathway(s) are yet to be deciphered. Preliminary indications pointed out the possible implication of an NO synthase-like enzyme in M. truncatula–S. meliloti interaction (Baudouin et al., 2006). However, the MtNoa1 gene (the orthologue of AtNoa1) recently identified in M. truncatula is not involved in NO production (Pauly et al., 2010). In addition, recent evidence demonstrates that both plant and bacterial nitrate reductase and the electron transfer chains are responsible for NO production in M. truncatula–S. meliloti nodules (Horchani et al., 2011). However, the NO source(s) during the first steps of the interaction remain(s) to be identified. Therefore to reduce NO levels in both bacteria and plant cells, we first used an S. meliloti strain overexpressing a flavohaemoglobin (Hmp) (Meilhoc et al., 2010), a well-characterized NO-scavenging enzyme. Second, to assess the role of NO during the nodule primordium development, we generated transgenic hairy roots of M. truncatula expressing the S. meliloti flavohaemoglobin hmp gene under the control of a nodule-specific promoter. As reported by Mishina et al. (2007) in Arabidopsis, this strategy was efficient in reducing the NO levels during the nodulation process and allowed us to observe effects both on bacterial infection and nodule development. We first observed a lower competitive ability of the hmp++ strain compared with the wild type. Moreover, expression of Hmp from either the plant or bacterial side, or the early addition of the NO scavenger cPTIO to plant roots, led to delayed nodulation. In agreement with these observations, Pii et al. (2007) described that early addition of cPTIO reduced the M. truncatula nodule number. Together, these results provide convincing evidence that NO is involved in the establishment of the M. truncatula–S. meliloti symbiosis, where it plays a positive role in the nodulation process.
What could be the function of NO in the nodulation process? It has been suggested that NO produced by bacterial pathogens could protect them against oxidative stress encountered during host infection (Gusarov & Nudler, 2005). Similarly, NO could thus be produced by rhizobia in the early stages of symbiosis in order to counteract the oxidative stress encountered during infection (Pauly et al., 2006). Interestingly, it appears that NO is produced very early in the infection process, as soon as infection pockets are formed. Both cell-permeable NO-specific fluorescent probe and NO biosensor bacterial strain indicate that bacteria respond to NO in infection pockets. However, in this study, we cannot conclude what from the plant or the bacteria is involved in this NO synthesis. Furthermore, detection of NO along the infection threads but not within bacteria during the IT growth suggests that NO production at this step is mainly from the plant partner, and/or that bacterial system(s), including the flavohaemoglobin Hmp, prevent(s) NO accumulation inside bacteria (Meilhoc et al., 2010). In the Vibrio fischeri–Euprymna scolopes symbiosis, host-derived NO has an important role during the initiation of symbiotic colonization where it is hypothesized to serve as a specificity determinant (Davidson et al., 2004). It has been suggested that NO in this case could be a signal that prepares the microsymbiont for stronger NO stress occurring later in the symbiotic process (Wang et al., 2010). Similarly, in the Rhizobium–legume symbiosis, early production of NO could serve to induce the expression of bacterial genes that could be necessary to adapt bacteria to NO encountered later on during symbiosis. The lower competitiveness of the hmp++ strain compared with the wt is an argument in favour of a role of this NO production during the infection process.
The question remains whether or not NO synthesis by the plant is directly induced by the microsymbiont. In L. japonicus and M. sativa, Nagata et al. (2008) observed a transient NO production on roots only when the plants were inoculated with their own specific symbiotic partners (Mesorhizobium loti and S. meliloti, respectively). Furthermore, this increase in NO was accompanied by the transient induction of the expression of a class 1 Hb-encoding gene which could be responsible for subsequent NO decrease. It was suggested that class 1 Hb might be involved in the development of symbiosis in L. japonicus (Shimoda et al., 2009). The same authors also reported that the number of nodules formed on transgenic hairy roots overexpressing LjHb1 was increased compared with those formed on untransformed hairy root from the same plant (Shimoda et al., 2009). These results appear contradictory to the present work where we observed a reduced number of nodules per plant when depleting NO concentrations using either hmp overexpression or the NO scavenger cPTIO. This could be caused by: (1) the different models and approaches used, including the expression of hmp from different promoters (i.e. a nodule-specific promoter in our work vs the strong and constitutive p35S promoter in Shimoda’s work); (2) a putative specific role played by the class I Hb to promote the initial infection process; and/or (3) the need for a set concentration range of NO for successful establishment of the symbiotic relationship.
The detection of NO in dividing cortical cells of the root, not yet invaded by the rhizobial cells within nodule primordia, shows that NO could also play a direct role in nodule organogenesis. Nitric oxide was also detected in lateral root primordia as previously described in tomato plants (Correa-Aragunde et al., 2004), which suggests that it could have a similar function in these two different developmental processes. The role of NO in nodule development is underlined by the effect of NO depletion on the expression of genes known to be involved in this process. A first target appears to be MtCRE1, which has been shown already as an NO responsive gene (Ferrarini et al., 2008). This gene encodes a cytokinin receptor, which regulates the early symbiotic interaction between M. truncatula and S. meliloti, by modulating both the progression of the infection and the formation of nodule primordia (Gonzalez-Rizzo et al., 2006); its expression was localized in nodule primordia (Lohar et al., 2006). It has been proposed that cytokinin, as a key differentiation signal for nodule organogenesis, might be a ‘secret agent’ of the symbiotic interaction (Frugier et al., 2008). Thus, in regulating cytokinin perception, NO may control the nodulation process. Moreover, as NO has been put forward as an intermediate in cytokinin signalling in Arabidopsis (Tun et al., 2008), an amplification loop may exist. The proposed event cascade involving the response regulator MtRR4 and the downstream MtNIN and MtENOD40 functions (Frugier et al., 2008) was not affected by the cPTIO treatment, pointing to the existence of other signalling routes functioning downstream of MtCRE1 and controlling nodule development. The effect of the decrease in the NO content on the expression of the MtCCS52A genes may also contribute to explain the positive role of this signalling molecule on the nodulation process. MtCCS52A, which is involved in the transition of mitotic cycles to endoreduplication cycles, is indeed required for symbiotic cell differentiation in M. truncatula nodules (Vinardell et al., 2003). These results are in line with a previous report on the NO-induced modulation of the expression of cell cycle regulatory genes during lateral root formation in tomato plants (Correa-Aragunde et al., 2006).
In conclusion, the results presented here show that NO is required for an optimal establishment of the M. truncatula–S. meliloti symbiosis, and suggest that it could have functions in both bacterial infection and nodule development. Indeed, enhancing NO scavenging activities, both at the plant and microsymbiont levels, resulted in a delay in nodule formation and a weaker bacterial infectivity. They point to a potential link between cytokinin and NO signalling during the nodulation process. In addition, they emphasize a difference with the role of NO in pathogenic interactions, where it mainly activates plant defence reactions.
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This work was supported by an Agence Nationale de la Recherche programme (grant BLAN 071_184783). EM was supported by the National Institute for Applied Sciences (INSA-Toulouse); YC was a recipient of a ‘Contrat Jeune Scientifique INRA’. We thank Aurore Poteau for her dedicated technical assistance and William Rhamey for the English proofreading of the manuscript.
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Fig. S1 Histochemical localization of β-glucuronidase (GUS) activity throughout various stages of root nodulation in transgenic hairy roots of Medicago truncatula expressing the pMtENOD20–GUS fusion.
Table S1 Primer sequences for cloning and quantitative reverse-transcription polymerase chain reaction (RT-PCR) analysis
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