Significance

Inoculation of cereals with diazotrophic (N2-fixing) bacteria offers a sustainable alternative to the application of nitrogen fertilizers in agriculture. While natural diazotrophs have evolved multilayered regulatory mechanisms that couple N2 fixation with assimilation of the product NH3 and prevent release to plants, genetic modifications can permit excess production and excretion of NH3. However, a lack of stringent host-specificity for root colonization by the bacteria would allow growth promotion of target and nontarget plants species alike. Here, we exploit synthetic transkingdom signaling to establish plant host-specific control of the N2-fixation catalyst nitrogenase in Azorhizobium caulinodans occupying barley roots. This work demonstrates how partner-specific interactions can be established to avoid potential growth promotion of nontarget plants.

Abstract

Engineering N2-fixing symbioses between cereals and diazotrophic bacteria represents a promising strategy to sustainably deliver biologically fixed nitrogen (N) in agriculture. We previously developed novel transkingdom signaling between plants and bacteria, through plant production of the bacterial signal rhizopine, allowing control of bacterial gene expression in association with the plant. Here, we have developed both a homozygous rhizopine producing (RhiP) barley line and a hybrid rhizopine uptake system that conveys upon our model bacterium Azorhizobium caulinodans ORS571 (Ac) 103-fold improved sensitivity for rhizopine perception. Using this improved genetic circuitry, we established tight rhizopine-dependent transcriptional control of the nitrogenase master regulator nifA and the N metabolism σ-factor rpoN, which drove nitrogenase expression and activity in vitro and in situ by bacteria colonizing RhiP barley roots. Although in situ nitrogenase activity was suboptimally effective relative to the wild-type strain, activation was specific to RhiP barley and was not observed on the roots of wild-type plants. This work represents a key milestone toward the development of a synthetic plant-controlled symbiosis in which the bacteria fix N2 only when in contact with the desired host plant and are prevented from interaction with nontarget plant species.
Exploitation of symbiotic and associative bacterial N2 fixation (the reduction of N2 → NH3) represents a key strategy to sustainably meet the rising nitrogen (N)-demands of modern-day agriculture (1). Maintaining productivity of major cereal crops—such as wheat, maize, barley, and rice—which constitute over 50% of global human caloric intake, is paramount (2, 3). In contrast to legumes, cereals lack the genetic components required to establish intimate associations with N2-fixing rhizobial bacteria, but their root compartments are naturally colonized by associative epiphytic and endophytic diazotrophic bacteria that are estimated to fix up to 5 Tg N ha−1 yr−1 in global agrosystems (36). In some instances, inoculation of cereals with associative diazotrophic bacteria has a positive influence on yield (7, 8), though responses are typically inconsistent largely because of failure of inoculants to competitively colonize root systems and reach quorum density (9). Furthermore, the costly energy demands of N2 fixation have driven the evolution of multilayered genetic mechanisms in bacteria that repress N2 fixation and favor NH3 assimilation over release to plants (10).
Delivery of biologically fixed N could be improved by engineering the N2-fixing catalyst nitrogenase or the capacity for nodule symbiosis directly into cereals (1114). Alternatively, preexisting associative bacterial N2-fixing interactions could be genetically manipulated for improved function (9, 1517). Overriding negative feedback regulation of N2 fixation to permit excess production and release of NH3 to plants represents a focal point for the latter strategy (10, 18). However, a lack of stringent host-specificity for root colonization by the bacteria (6, 9) would allow growth promotion of target and nontarget plants species alike. Establishing host-plant control of engineered bacterial traits would provide specificity to such interactions. Plant–microbe signaling circuitry has already been engineered into tobacco plants, conveying them with the ability to synthesize bacterial acylhomoserine lactones (AHL), which activated gene expression in bacterial AHL biosensors and restored biocontrol activity and pathogenicity of AHL synthase-deficient Pseudomonas aureofaciens and Erwinia carotovora mutants, respectively (19). Similarly, opine producing Arabidopsis plants generated by Agrobacterium-mediated transformation were shown to selectively enrich the rhizosphere for opine catabolizing bacteria (2022), with ratio and population density positively correlating with the input of opines (23). Critically, AHLs exhibit poor signaling specificity (24, 25), whereas opines induce pathogenic agrobacteria to form crown galls on plants (26); therefore, these signaling circuits may be suboptimal for control of N2-fixation and other plant–microbe interactions in agriculture.
To develop a more practical plant–microbe signaling circuit, we engineered into barley and Medicago a synthetic biosynthesis pathway for production of the rhizopine signaling molecule scyllo-inosamine (SI), which is exuded from the roots and can be perceived by bacteria carrying a rhizopine receiver plasmid on the root surface and rhizosphere (27, 28). The pathway requires myo-inositol as a substrate, which is converted to scyllo-inosose by an inositol dehydrogenase (IolG/IdhA) and then to SI by the aminotransferase (MosB), both of which are expressed constitutively in the plant (27). In nature, rhizopines including SI and 3-O-methyl scyllo-inosamine (3-O-MSI) are synthesized by rhizobia occupying nodules under control of the nitrogenase master regulator NifA (28). They are excreted from the nodule and enhance nodulation competitiveness in progeny strains carrying the catabolic moc genes, but do not increase the population size of these bacteria occupying the rhizosphere (29). As a signaling molecule, SI exhibits a high degree of specificity, with only a few species of Rhizobium and Sinorhizobium identified that catabolize it as a nutrient source (28, 30, 31). Furthermore, SI does not stimulate pathogenic behavior by resident bacteria, as is true of opines (32).
Here, we built upon our rhizopine signaling circuitry to establish host-plant control of associative N2 fixation by the model cereal endophyte Azorhizobium caulinodans ORS571 (here termed Ac). Previously, we established tight control over free-living N2 fixation in response to exogenous supplementation of Ac cultures with synthetic and plant-derived chemicals (15). This involved replacing the native master regulator of N2 fixation nifA with chemically inducible nifA or nifA-rpoN controller plasmids driving the expression of the large suite of nif and fix genes required for assembly and function of nitrogenase. Controlled expression of nifA circumvented natural transcriptional repression by N, whereas N-mediated posttranslational repression of NifA was partially overcome by expressing a NifAL94Q/D95Q variant protein that is defective in sensing N status (15). Because mechanisms such as the DraT-DraG system that directly inhibit nitrogenase activity in response to N availability are absent in Ac (3335), regulated NifAL94Q/D95Q expression permits control of N2 fixation that is partially resistant to feedback inhibition by the product NH3 (15).
In this study, we develop a homozygous rhizopine-producing (RhiP) barley line and bacterial SI receiver plasmid that conveys markedly improved sensitivity to rhizopine perception in Ac. With this improved sensitivity, we established tight rhizopine-dependent transcriptional control of nifA and a variant gene encoding NifAL94Q/D95Q (15) that when coexpressed with the N metabolism σ-factor, drove partially NH3-resistant, rhizopine-inducible nitrogenase activity in vitro. We also demonstrated that nitrogenase activity is specifically activated in situ by bacteria colonizing RhiP barley roots, but not in bacteria colonizing wild-type roots. Although this activity was suboptimal compared to the wild-type system, this work provides a proof-of-concept for the exclusive coupling of plant–microbe interactions in the field.

Results

Coexpression of myo-Inositol Uptake Genes Enhances SI Uptake and SI-Inducible Expression.

Rhizopine perception in bacteria is dependent on activation of MocR, which drives expression from the rhizopine-inducible promoter PmocB in the presence of rhizopine (27). Initial tests of rhizopine perception in Ac carrying the first-generation rhizopine receiver plasmid pOPS0761 revealed that PmocB::GFP induction was poor, requiring 1 mM of SI to invoke a less than twofold induction (SI Appendix, Fig. S1). We reasoned that coexpression of a high-affinity rhizopine transporter might improve the sensitivity and dynamic range of rhizopine-inducible expression. Although the solute binding protein (SBP) MocB was encoded on our first-generation rhizopine-receiver plasmids, transmembrane components required for transport of rhizopine into the cytoplasm had not yet been identified. An ATP-binding cassette (ABC) transporter (IntABC) that accumulates the structurally related molecule myo-inositol is present in the model pea nodule symbiont Rhizobium leguminosarum bv. viceae 3841 (here termed Rlv) (36, 37). Given close homology of the myo-inositol SBP IntA with MocB, and the structural relatedness of their substrates (SI Appendix, Fig. S2), we explored whether the transmembrane components IntBC act in concert with MocB to transport SI. When Rlv carrying the rhizopine-receiver plasmid pOPS0046 was grown on myo-inositol to induce transcription of the native intABC genes, PmocB::lux expression was increased compared to bacteria grown on pyruvate at all concentrations of SI tested (SI Appendix, Fig S3A). We also compared PmocB::lux expression between a polar intA::mini-Tn5 mutant of Rlv carrying pOPS0046 where intABC expression was abolished, against an identical strain additionally carrying plasmid pOPS0385, where intBC was constitutively expressed from the Plac promoter, and found that PmocB::lux expression was increased by intBC expression when 10 μM or more SI was supplemented into the growth media (SI Appendix, Fig. S3B). Thus, the intBC genes are required for a sensitive transcriptional response to SI.
To test whether coexpression of intBC would improve rhizopine-inducible expression in our model strain Ac, we assembled a “second-generation” rhizopine-receiver plasmid pSIR02 where mocB-intBC was expressed from the weak constitutive Anderson promoter PJ23115 (http://parts.igem.org/Promoters/Catalog/Anderson) and assembled the plasmid pSIR01 devoid of the intBC genes as a control (Fig. 1A). PmocB::GFP expression in Ac carrying pSIR02 was increased across all levels of SI-tested, with minimum induction achieved at 100 nM SI and maximum induction achieved with less than 10 μM SI supplemented into the growth media (Fig. 1B). In comparison, Ac carrying pSIR01 required 1 mM SI to elicit a less than twofold induction of PmocB::GFP (Fig. 1B and SI Appendix, Fig. S1B). The sensitivity of SI-perception by Ac carrying pSIR02 was therefore improved at least 103-fold. We also analyzed SI-inducible expression by flow-cytometry in the mCherry-labeled strain AcCherry carrying pSIR02 and found that ∼36% (±SEM 0.3%) of cells exhibited GFP fluorescence above the mean 99th percentile of the noninduced cells (defined here as GFP+) (Fig. 1C and Dataset S1). Lastly, we bio-assayed the intracellular contents of the Ac strains after 16-h incubation with 100 μM SI and found that cells carrying the pSIR02 plasmid harbored more SI relative to Ac carrying pSIR01 and wild-type Ac (Fig. 1D and Dataset S2). Overall, the data support IntBC functioning as transmembrane and ABC components, respectively, of a hybrid SI-uptake system utilizing MocB as the SBP.
Fig. 1.
Coexpression of the myo-inositol uptake genes intBC enhances rhizopine uptake and sensitivity of the transcriptional response. (A) Genetic schematics (not to scale) for the second-generation rhizopine receiver plasmid pSIR02 and a near identical plasmid lacking intBC. (B) PmocB promoter activity (RFU, defined as GFP fluorescence/ODλ600 nm) was measured in populations of Ac carrying pSIR01 or pSIR02. (C) Flow-cytometry analysis of AcCherry (pSIR02) cells induced with 10 μM SI or not induced after 18-h incubation (n = 4). Cells exhibiting GFP fluorescence above 20,000 a.u. were considered as GFP+. Supplementary statistics supporting flow cytometry are provided in Dataset S1. (D) The intracellular contents of thrice washed Ac cells grown in the presence of 100 μM SI was bio-assayed for SI using S. meliloti carrying pOPS0046 and pOPS0385 as a sensor. Error bars represent one SEM (n = 3). Independent two-tailed Student’s t tests were used to compare means. Exact values for P > 0.05 are given, ***P < 0.001.

Rhizopine Perception by Ac Colonizing RhiP Barley Roots.

We previously developed a T1 rhizopine-producing (RhiP) barley line expressing codon optimized R. leguminosarum idhA and Sinorhizobium meliloti L5-30 mosB genes in multicopy under the control of Zea mays ubiquitin1 promoter and Oryza sativa ubiquitin1 promoter, respectively. Here, we generated a homozygous T2 RhiP line which, based on GC-MS analysis of metabolite extracts from 10-d-old seedlings (n = 2, 10 technical replicates each), produced 0.93 (±SEM 0.10) ng mg−1 root dry weight (Datasets S3 and S4), equating to approximately sevenfold less than the T1 RhiP line (27). To track rhizopine-inducible expression on the root surface, we inoculated plants with the constitutive mCherry-labeled strain AcCherry (pSIR02) and analyzed PmocB::GFP induction in single cells colonizing the root surface by confocal microscopy at 7 d postinoculation (dpi) (Fig. 2 A and B and SI Appendix, Fig. S4). GFP fluorescence above background levels was never observed in bacteria-colonizing wild-type plants, but was observed in a subpopulation of cells colonizing T2 RhiP barley, indicating PmocB was active.
Fig. 2.
Rhizopine-inducible expression on RhiP barley roots and in the rhizosphere. (A) Schematic representation of the experimental design to monitor PmocB expression on barley roots and in the rhizosphere. AcCherry cells are marked with a constitutively expressed mCherry gene allowing tracking. The strain additionally carries plasmid pSIR02. (B) Maximum-intensity projections showing mCherry and GFP fluorescence by AcCherry (pSIR02) cells colonizing the rhizoplane of wild-type and T2 RhiP barley at 9 dpi. Projections are representative of 15 images acquired from 3 whole lateral roots excised from 5 plants per treatment. (C) Flow-cytometry analysis of AcCherry (pSIR02) cells recovered from the rhizoplane and endosphere (RA) or rhizosphere (RS) of wild-type (blue), T1 RhiP (green), and T2 RhiP (brown) barley plants at 9 dpi (n = 5). Cells exhibiting mCherry fluorescence above 5,000 a.u. were analyzed. GFP+ cells are defined as those exhibiting GFP fluorescence above the mean 99th percentile of the populations colonizing wild-type barley. Supplementary statistics supporting flow cytometry are provided in Dataset S5. (D) In situ PmocB promoter activity (median GFP fluorescence intensity) was measured for R2 GFP+ bacteria in C. Error bars represent one SEM. Independent two-tailed Student’s t tests were used to compare means. Exact values for P > 0.05 are given, *P < 0.05, **P < 0.01.
AcCherry (pSIR02) cells were subsequently recovered from the rhizoplane and endosphere (here termed root-associated, RA) and from the rhizosphere (RS) of 9-dpi barley plants for quantification of SI-induction by flow-cytometry (38). Using the same experimental design, we also analyzed SI-induction by AcCherry (pSIR02) cells recovered from our previously reported T1 RhiP barley line. We found that 3.60% of cells isolated from the RA and 4.56% of cells isolated from the RS fraction of T2 RhiP plants exhibited GFP fluorescence higher than the mean 99th percentile of that detected in bacteria recovered from wild-type plants (here defined as GFP+) (Fig. 2C), confirming these subpopulations were induced for PmocB::GFP. On T1 barley roots, 4.96% of cells isolated from the RA fraction and 10.70% of cells from the RS fraction were GFP+, suggesting that the percentage of AcCherry (pSIR02) cells activated for GFP increases with increasing supply of rhizopine from the plant. Moreover, higher rhizopine production by the T1 RhiP barley translated to stronger induction of PmocB::GFP by AcCherry (pSIR02) cells (Fig. 2D).
We checked whether loss or silencing of the plasmid pSIR02 had occurred in bacterial populations recovered from wild-type and T2 RhiP barley roots. After regrowing bacteria recovered from three plants on nonselective universal minimal salts (UMS) agar, 60 of 60 (n = 3) of the colonies grew when patched onto UMS agar supplemented with kanamycin, indicating the plasmid pSIR02 was retained in all cases. Moreover, between 58% and 72% of colonies recovered from both fractions of the wild-type and T2 RhiP barley plants were induced for PmocB::GFP expression when regrown on UMS agar supplemented with 1 μM SI (SI Appendix, Fig. S5), indicating that the majority retained the capacity for functional rhizopine signaling.

In Vitro Rhizopine Control of N2 Fixation.

To establish rhizopine-dependent control of N2 fixation in Ac, we used pSIR02 as a backbone to generate nifA-rpoN and nifAL94Q/D95Q-rpoN controller plasmids pSIN01 and pSIN02, respectively (Fig. 3A). These controllers were tested for functionality in the markerless nifA deletion mutant AcΔnifA grown under diazotrophic conditions (N-free liquid UMS with a starting atmosphere of 3% O2 and ∼97% N2), by using a PnifH::mCherry reporter construct (pOPS1218) designed to monitor expression of the nitrogenase structural gene nifH and by performing acetylene reduction assays (ARAs) to assess nitrogenase activity. In AcΔnifA carrying pSIN01 or pSIN02 and pOPS1218, basal levels of PnifH::mCherry expression were detected in the absence of SI, while supplementation of 10 μM SI into the growth media resulted in above wild-type levels of PnifH::mCherry expression (Fig. 3B). No nitrogenase activity was detected for the same bacteria (devoid of pOPS1218) in the absence of SI, while supplementation of 10 μM SI into the growth media resulted in wild-type rates of nitrogenase activity, demonstrating tight binary control (Fig. 3C). As previously reported (15), induction of either nifA-rpoN or nifAL94Q/D95Q-rpoN cassettes resulted in partial PnifH expression and nitrogenase activity in the presence of 10 mM NH4Cl, with strains carrying nifAL94Q/D95Q-rpoN exhibiting the highest partial activity at ∼50% that of wild-type strains grown in N-free conditions (Fig. 3).
Fig. 3.
In vitro rhizopine control of N2-fixation. (A) Genetic schematics (not to scale) for nifA-rpoN and nifAL94Q/D95Q rhizopine controller plasmids. (B) PnifH promoter activity (RFU, defined as mCherry fluorescence/ODλ600 nm) was measured in populations of Ac and AcΔnifA (n = 3) carrying rhizopine controller plasmids and the reporter pOPS1218 grown in N2-fixing conditions (N-free UMS supplemented with 20 mM succinate with a starting headspace of 3% O2). SI and NH4Cl was added to a final concentration of 10 μM and 10 mM, respectively, where indicated. (C) The same conditions were used to assess specific nitrogenase activity in Ac and AcΔnifA carrying rhizopine controller plasmids. Error bars represent one SEM (n = 3). Independent two-tailed Student’s t tests were used to compare means. Exact values for P > 0.05 are given, *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.001.

In Situ Plant Host-Dependent Control of Associative N2 Fixation.

We next sought to demonstrate rhizopine control of nifH expression and nitrogenase activity by bacteria occupying the root systems of RhiP barley plants. Plasmid pSIN02 was modified by replacing the GFP gene downstream of PmocB with a divergently oriented PnifH::GFP reporter fusion (SI Appendix, Fig. S6) and the resulting plasmid pSIN03 was conjugated into AcCherryΔnifA. This strain was inoculated onto wild-type and T2 RhiP barley and induction of PnifH::GFP in single cells on the root surface was assessed by confocal microscopy at 9 dpi after 24-h exposure to a headspace atmosphere of 1% O2. As previously observed for SI-induction of PmocB::GFP on barley roots (Fig. 2 A–C and SI Appendix, Fig. S4), GFP fluorescence above background levels was never observed in bacteria colonizing wild-type plants, but was observed in a subpopulation of cells colonizing T2 RhiP barley (Fig. 4 A and B and SI Appendix, Fig. S7), indicating that nifH expression was induced. We also used flow-cytometry to quantify PnifH::GFP activation and found the results relatively mirrored those obtained for activation of PmocB::GFP (Fig. 2 C and D). On T1 barley roots, 22.92% and 22.32% of bacteria occupying the RA and RS fractions, respectively, were GFP+, whereas on T2 barley roots 5.26% and 5.10% of bacteria occupying the RA and RS fractions, respectively, were GFP+ (Fig. 4C). The mean GFP fluorescence was slightly higher in bacteria colonizing T1 RhiP barley compared to the T2 line, though this was not statistically significant (Fig. 4D).
Fig. 4.
In situ plant-dependent control of associative N2-fixation (A) Schematic representation of the experimental design to monitor rhizopine-dependent PnifH expression on barley roots. AcCherry cells were marked with a constitutively expressed mCherry gene allowing tracking. The strain additionally carries plasmid pSIN03. (B) Maximum-intensity projections showing mCherry and GFP fluorescence by AcCherryΔnifA (pSIN03) cells colonizing the rhizoplane of wild-type and T2 RhiP barley at 9 dpi after 24-h exposure to 1% O2 in the headspace. Projections are representative of 15 images acquired from 3 whole lateral roots excised from 5 plants per treatment. (C) Flow-cytometry analysis of AcCherry (pSIN03) cells recovered from the rhizopine and endosphere (RA) or rhizosphere (RS) of wild-type (blue), T1 RhiP (green), and T2 RhiP (brown) barley plants at 9 dpi after 24-h exposure to 1% O2 in the headspace (n = 5). Cells exhibiting mCherry fluorescence above 1,500 a.u. were analyzed. GFP+ cells are defined as those exhibiting GFP fluorescence above the mean 99th percentile of the populations colonizing wild-type barley. Supplementary statistics supporting flow-cytometry are provided in Dataset S7. (D) In situ PnifH promoter activity (median GFP fluorescence intensity) was measured for R2 GFP+ bacteria in C. (E) In situ ARAs performed at 9 dpi after 48-h exposure to 1% O2 in the headspace. Error bars represent one SEM. Independent two-tailed Student’s t tests with Holm–Bonferroni adjusted P values were used to compare means. Exact values for P > 0.05 are given, **P < 0.01.
To assess activation of N2 fixation, in situ ARAs (38) were performed on wild-type and RhiP barley inoculated with AcΔnifA carrying pSIN02 at 9 dpi after adjusting the headspace of vessels to 1% O2. Partially effective nitrogenase activity was induced on both T1 and T2 RhiP barley, whereas it was not detected on wild-type plants (Fig. 4E). Compared to the wild-type Ac, which produced ∼70 nmol C2H4 h−1 plant−1 when colonizing wild-type or RhiP barley roots, nitrogenase in AcΔnifA carrying pSIN02 produced 10.24 nmols C2H4 h−1 plant−1 when colonizing T1 RhiP barley roots and 3.63 nmols C2H4 h−1 plant−1 when colonizing T2 RhiP barley roots, equating to ∼15% and 5% of wild-type levels, respectively. Importantly, the relative difference in nitrogenase activity detected for AcΔnifA carrying pSIN02 colonizing T1 RhiP plants compared to T2 RhiP plants was consistent with in situ PmocB::GFP (Fig. 2 C and D) and PnifH expression (Fig. 4 C and D) in the same plant lines, suggesting that in situ rhizopine-inducible nitrogenase activity is also dependent on the concentration of SI supplied by the plant. Upon recovering the bacteria and performing viable counts, it was apparent that root colonization by AcΔnifA bacteria carrying pSIN02 was mildly defective compared to Ac (SI Appendix, Fig. S8), which may partially contribute to the suboptimal nitrogenase activity observed in this strain. Nevertheless, the results reported here provide a proof-of-concept for the establishment of synthetic host-specific control over bacterial nitrogenase activity.

Discussion

In this work, we have stably engineered a homozygous rhizopine biosynthesis pathway into barley and concurrently improved rhizopine signaling circuitry in Ac to establish plant host-dependent control of nifA-rpoN cassettes driving expression and activity of nitrogenase. Despite the 103-fold improved sensitivity of in vitro rhizopine perception, our confocal microscopy and flow-cytometry experiments indicated that only a small fraction of the AcCherry (pSIR02) population occupying the RhiP barley roots responded to rhizopine signaling by expressing PmocB::GFP (Fig. 2). Likewise, in situ nitrogenase activity by AcΔnifA (pSIN02) colonizing RhiP barley was suboptimal compared to wild-type Ac colonizing wild-type or RhiP barley (Fig. 4E). It seems unlikely that plasmid loss was responsible for these results given that all the AcCherry (pSIR02) colonies recovered from RhiP barley tested positive for kanamycin resistance conveyed by pSIR02. Silencing may have impeded efficient in situ rhizopine signaling and nitrogenase activity, although 58 to 72% of colonies recovered from T2 RhiP barley roots retained the capacity for SI induction of PmocB::GFP on agar plates (SI Appendix, Fig. S5). Finally, we found that AcΔnifA (pSIN02) are mildly defective in their ability to colonize barley roots, which may reduce nitrogenase activity per plant compared to the wild-type system. Our results overall suggested that increased rhizopine production by RhiP plants translated to stronger individual and population level SI perception and more effective nitrogenase expression and activity by bacteria (Figs. 2 and 4). Thus, further optimization of the rhizopine sensor and tuning of rhizopine biosynthesis gene expression may be necessary to generate a synthetic interaction that is better suited for practical use. It is crucial to note that excessive overexpression of SI biosynthesis genes would likely cause defects in plant metabolism that might influence biomass accumulation and increase the potential for enrichment of specialized bacteria capable of catabolizing the signal as a source of carbon or N in the field (2830), as was previously reported for transgenic opine-producing plants (2023). Thus, retaining SI biosynthesis at the minimum level required for a functional bacterial response will be crucial to optimize this circuitry.
Building on rhizopine control of N2 fixation demonstrated here, our future aim is to engineer a more complete “synthetic symbiosis” where bacteria are also prompted to release fixed N to the host plant. This will require rewiring of nitrogen metabolism as Ac does not naturally release large quantities of fixed N into the growth media (39) but can be engineered to do so by deletion of the PII genes glnB and glnK (4042) or potentially by other strategies (18, 4345). In addition, rhizopine signaling could be deployed by bacteria to initiate response signals to plants, establish biocontainment of bacteria in the root system, and develop relay signaling circuitry that will allow genetically incompatible bacteria to indirectly respond to rhizopine (9). Ultimately, we intend to utilize rhizopine signaling circuitry for control of bacterial traits within engineered barley nodules or “pseudo-nodules” where more fine-tuned signal and nutrient exchange would presumably exist (11, 12, 46). The biotechnological achievements made in this study represent a significant milestone toward the development of efficient partner-specific synthetic symbioses between cereals and bacteria.

Materials and Methods

Bacterial Strains and Growth Conditions.

Bacterial strains used in this study are described in SI Appendix, Table S1. Escherichia coli was cultured on LB media and incubated at 37 °C, whereas Rhizobium and Azorhizobium strains were cultured on TY or UMS media and incubated at 28 °C or 37 °C, respectively. Where appropriate, UMS media was supplemented with 20 mM succinate or 30 mM pyruvate as a sole carbon source, 10 mM NH4Cl as a sole nitrogen source, and 300 μM nicotinate (for Ac only). Antibiotics were added to media for E. coli and rhizobia, respectively, at the following concentrations: kanamycin 50, 100 μg mL−1; gentamycin 10 μg mL−1, 25 μg mL−1; tetracycline 10 μg mL−1, 5 μg mL−1; carbenicillin 100 μg mL−1; and spectinomycin 50 μg mL−1. Strains harboring markerless nifA deletions were constructed via homologous recombination and sucrose counter selection (38) of plasmid pOPS1565 (SI Appendix, Table S2).

Plasmids.

Plasmids were constructed using golden-gate assembly (47, 48), HiFi assembly (New England Biolabs), and yeast recombination (49), as described in SI Appendix, Supplementary Materials and Methods. All inserts were confirmed by Sanger sequencing. Plasmids were chemically transformed into E. coli strains (50) and mobilized into rhizobia via biparental conjugation with E. coli ST18 (51) or via triparental conjugation with a helper E. coli strain carrying plasmid pRK2013. Plasmids with R6K replicons were maintained at low copy number in E. coli EC100D Pir+ where possible.

Reporter Assays.

Single colonies were initially streaked onto 10 mL TY agar slopes in 28-mL universal vials and incubated for 2 to 3 d. Cells were washed from slopes in PBS, pelleted by centrifugation, and inoculated to an optical density (OD)λ600 nm of 0.1 into 96-deep-well culture plates, each well containing 500 μL UMS supplemented with the appropriate inducer. For PnifH induction assays, single colonies were initially streaked onto UMS agar slopes in 28-mL universal vials and incubated for 2 d prior to washing with PBS three times to remove residual nitrogen. Cells were finally resuspended in 5 mL N-free UMS liquid media with 20 mM succinate at ODλ600 nm 0.4 in 28-mL universal vials and the O2 concentration was adjusted to 3% by flushing the headspace with N2 gas for 1 h before sealing the vials with a rubber septum. Cultures were incubated for 18 h prior to 1:1 dilution with UMS and transfer into clear-bottom 96-well plates for quantification. Relative luminescence units (defined as RLU, luminescence/ODλ600 nm) were measured with a Promega GloMax multidetection system and relative fluorescence units (defined as RFU, mCherry/ODλ600 nm or GFP/ODλ600 nm) were measured with an Omega FLUOstar set at gain 1,000 for GFP and 1,500 for mCherry. GFP fluorescence was measured at excitation 485 nm and emission 520 nm, whereas mCherry fluorescence was measured at excitation 560 nm and emission 590 nm.
To bioassay intracellular SI in Ac, cells were washed from TY slopes after 2-d growth, pelleted by centrifugation, and inoculated at ODλ600 nm 0.4 into 28-mL universal vials containing 5 mL UMS supplemented with 100 μM SI. Cultures were incubated with vigorous shaking for 16 h, after which the culture ODλ600 nm was recorded, and 4 mL of cells were harvested by centrifugation. Cells were washed three-times in 5 mL PBS and resuspended in a final volume of 800 μL prior to mechanical lysis using a FastPrep-24 5G instrument. Cellular debris were removed by centrifugation and filter sterilization (0.22-μm pore size). Using S. meliloti 1021 carrying the SI receiver plasmid pOPS0046 (PmocB::luxCDABE) and plasmid pOPS0365 (Plac::intBC) as a biosensor, an SI standard curve was established and a four-parameter log-logistic model was fitted (Dataset S2) using the AAT Quest Graph Four Parameter Logistic (4PL) Curve Calculator (52, 53). SI was concurrently bio-assayed in 50-μL aliquots of the cleared intracellular lysates. Derived molar concentrations of SI in the total lysates were converted on a per cell basis using our experimentally derived conversion factor of ODλ600 nm 1.0 = 2 × 108 cells.

Engineering Rhizopine Biosynthesis in Barley.

The rhizopine biosynthesis plasmid pEC12811 carried codon optimized R. leguminosarum idhA and S. meliloti L5-30 mosB genes expressed under the control of Z. mays ubiquitin1 promoter and O. sativa ubiquitin 1 promoter, respectively (27). The plasmid was integrated into the genome of Spring Golden Promise barley via Agrobacterium-mediated transformation (54) and plants were segregated to generate the homozygous T2 line RhiP (rhizopine-producing). Rhizopine production was confirmed in RhiP barley by metabolite extraction and GC-MS analysis of 10-d-old seedlings, as previously described (17). Raw data are provided in Datasets S3 and S4.

Confocal Microscopy.

Barley plants for confocal microscopy were propagated in fire sand as previously described (38) and were inoculated with 2 mL of a thrice-washed N- and C-free UMS Ac cell suspension standardized to ODλ600 nm 0.1. To assess PnifH induction, the headspace atmosphere of 9-dpi barley plants was adjusted to 1% O2 by flushing with N2 gas for 1 h and the vessels were sealed with a rubber septum and returned to the growth chamber for a further 24 h prior to analysis. Dual-channel confocal images were taken of whole lateral roots of barley using a ZEISS LSM 880 Axio Imager 2 with a C-Apochromat 40×/1.2W Korr FCS M27 objective, as previously described (38). Excitation of GFP and mCherry was achieved using 488-nm (3% power) and 561-nm (4% power) lasers, respectively, and fluorescence emissions were collected using photomultiplier tube detectors for GFP (493 to 598 nm, gain 520 for PmocB or 580 for PnifH induction) and mCherry (detection 598 to 735, gain 640 for PmocB or 700 for PnifH induction). The z-stack images were captured in 0.5-μm slices with the pinhole set to 1.39 AU using LineSequential unidirectional scan mode. Maximum-intensity projections were created from zstacks using the Zen 3.2 Blue software and are representative of 15 projections imaged from 5 plants (3 per plant) for each treatment.

Flow Cytometry.

For flow cytometry of bacteria grown in vitro, AcCherry (pSIR02) cells were induced with 10 μM SI as described above for reporter assays, diluted 1:100 in PBS, and analyzed using an Amnis Cellstream flow-cytometer equipped with an autosampler. GFP and mCherry was excited with 488- and 561-nm lasers, respectively. Singlets exhibiting mCherry fluorescence above 3,000 a.u. were considered as mCherry+ bacteria for analysis. mCherry+ bacteria exhibiting GFP fluorescence above the mean 99th percentile determined for noninduced cultures were considered as GFP+ cells. The supplementary statistics for this analysis are provided (Dataset S1).
Barley plants for flow-cytometry experiments were propagated and inoculated as described for confocal microscopy experiments. After 9 dpi, bacteria occupying the RA and RS fractions were recovered and analyzed by flow-cytometry, as previously described (38). Singlets exhibiting mCherry fluorescence above the 5,000 a.u. or 1,500 a.u. were considered as mCherry+ for analysis of PmocB and PnifH expression, respectively. mCherry+ cells exhibiting GFP fluorescence above the mean 99th percentile determined for bacterial populations colonizing wild-type barley roots were defined as GFP+. The supplementary statistics for these analyses are provided (Datasets S5 and S7).

Assessment of Plasmid Loss and Silencing in Bacteria Grown in Association with Barley Roots.

To assess plasmid loss in AcCherry (pSIR02) cells recovered from the RS and RA fractions of RhiP barley from flow-cytometry experiments, cell suspensions were plated on nonselective UMS agar and incubated 2 d. Sixty colonies recovered from each fraction of three RhiP plants were patched onto UMS agar supplemented with kanamycin, and onto nonselective UMS agar as a control. As the kanamycin resistance gene was expressed from pSIR02, resistant colonies were scored as having retained the plasmid. Silencing of pSIR02 was also assessed in the RS and RA fractions recovered from three wild-type and RhiP barley plants by plating cell suspensions on UMS agar supplemented with kanamycin or the same media additionally containing 1 μM SI. Plates were imaged using a Leica M165 FC fluorescent stereo microscope fitted with a CLS100x cold light source and DFC310 FX. To assess GFP induction, dual-channel images captured using dsRed and GFP filters were converted to 8-bit using ImageJ (55). mCherry+ colonies were demarcated for analysis in the dsRed channel using the thresholding tool (gray value > 20 and area > 100 pixel2) and these coordinates were superimposed over the GFP channel images. Colonies were scored as GFP+ if the maximum gray value exceeded 25. At least 32 colonies were analyzed per biological replicate. The raw data are provided (Dataset S6).

ARAs.

For ARAs of free-living bacteria in liquid cultures, cells were prepared and inoculated at ODλ600 nm 0.4 into 5 mL N-free UMS media supplemented with 20 mM succinate in 28-mL universal vials as described for PnifH induction assays. The starting headspace atmosphere was adjusted to 3% O2 by placing the vials on a rotary shaker in a controlled atmosphere of 97% N2 for 1 h. To initiate the assay, 10% of the total headspace atmosphere was replaced with C2H2 using a 20-mL syringe and needle and the cultures were incubated at 37 °C for 10 h or 24 h prior measuring the production of C2H4 using a PerkinElmer, Clarus 480 gas chromatograph equipped with a HayeSep N (80 to 100 MESH) 584 column. A second measurement was taken 2 h later and the rate of nitrogenase activity between these time points was calculated as previously described (38). Subsequently, 2 mL of the bacterial culture was harvested by centrifugation, washed in PBS, and mechanically lysed using a FastPrep-24 5G instrument. BCA assays were used to determine total protein content and specific nitrogenase activity calculated. In situ ARAs of bacteria inoculated onto barley plants were performed after 9 dpi with a headspace atmosphere of 1% O2 and 10% C2H4 as previously described (38).

Statistical Analyses.

Statistical analyses were carried out using the R package Rstatix (56). Dose–response curves were fitted using the R package drc (53). Details of each statistical analysis are outlined in the figure captions.

Data Availability

All study data are included in the main text and supporting information. All research materials supporting this publication can be accessed by contacting corresponding authors T.L.H. or P.S.P.

Acknowledgments

This work was supported by the Biotechnology and Biological Sciences Research Council (BB/T006722/1, BB/T001801/1, and BB/M011224/1), the Gatsby Foundation (GAT3395/GLH) and by the NSF (NSF-1331098). T.L.H. is the recipient of an 1851 Royal Commission for the Exhibition of 1851 Research Fellowship (RF-2019-100238) and Wolfson College, University of Oxford Junior Research Fellowship.

Supporting Information

Appendix 01 (PDF)
Dataset S01 (XLSX)
Dataset S02 (XLSX)
Dataset S03 (XLSX)
Dataset S04 (XLSX)
Dataset S05 (XLSX)
Dataset S06 (XLSX)
Dataset S07 (XLSX)

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Information & Authors

Information

Published in

Go to Proceedings of the National Academy of Sciences
Go to Proceedings of the National Academy of Sciences
Proceedings of the National Academy of Sciences
Vol. 119 | No. 16
April 19, 2022
PubMed: 35412890

Classifications

Data Availability

All study data are included in the main text and supporting information. All research materials supporting this publication can be accessed by contacting corresponding authors T.L.H. or P.S.P.

Submission history

Received: September 22, 2021
Accepted: March 11, 2022
Published online: April 11, 2022
Published in issue: April 19, 2022

Keywords

  1. rhizopine
  2. nitrogen fixation
  3. rhizobium
  4. barley
  5. symbiosis

Acknowledgments

This work was supported by the Biotechnology and Biological Sciences Research Council (BB/T006722/1, BB/T001801/1, and BB/M011224/1), the Gatsby Foundation (GAT3395/GLH) and by the NSF (NSF-1331098). T.L.H. is the recipient of an 1851 Royal Commission for the Exhibition of 1851 Research Fellowship (RF-2019-100238) and Wolfson College, University of Oxford Junior Research Fellowship.

Notes

This article is a PNAS Direct Submission.

Authors

Affiliations

Department of Plant Sciences, University of Oxford, Oxford OX1 3RB, United Kingdom
Sainsbury Laboratory, University of Cambridge, Cambridge CB2 1LR, United Kingdom
Marta D. Mendes
Department of Plant Sciences, University of Oxford, Oxford OX1 3RB, United Kingdom
Department of Plant Sciences, University of Oxford, Oxford OX1 3RB, United Kingdom
Department of Plant Sciences, University of Oxford, Oxford OX1 3RB, United Kingdom
Department of Microbiological Sciences, North Dakota State University, Fargo, ND 58105
Department of Plant Sciences, University of Oxford, Oxford OX1 3RB, United Kingdom
Department of Plant Sciences, University of Oxford, Oxford OX1 3RB, United Kingdom
Min-Hyung Ryu
Synthetic Biology Center, Department of Biological Engineering, Massachusetts Institute of Technology, Cambridge, MA 02139
Biochemistry and Metabolism, The John Innes Centre, Norwich NR4 7UH, United Kingdom
Synthetic Biology Center, Department of Biological Engineering, Massachusetts Institute of Technology, Cambridge, MA 02139
Giles E. D. Oldroyd
Sainsbury Laboratory, University of Cambridge, Cambridge CB2 1LR, United Kingdom
Department of Plant Sciences, University of Oxford, Oxford OX1 3RB, United Kingdom

Notes

1
To whom correspondence may be addressed. Email: [email protected] or [email protected].
Author contributions: T.L.H. and P.S.P. designed research; T.L.H., P.P., M.D.M., P.G., B.A.G., H.E.K., B.J., and P.B. performed research; T.L.H., P.P., P.G., B.A.G., H.E.K., B.J., M.-H.R., C.A.V., and G.E.D.O. contributed new reagents/analytic tools; T.L.H., P.P., M.D.M., P.G., B.A.G., H.E.K., B.J., P.B., and P.S.P. analyzed data; and T.L.H., P.P., C.A.V., G.E.D.O., and P.S.P. wrote the paper.

Competing Interests

The authors declare no competing interest.

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    Engineered plant control of associative nitrogen fixation
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