N-hydroxy-pipecolic acid is a mobile metabolite that induces systemic disease resistance in Arabidopsis

Significance

Plants lack circulating immune cells and instead rely on small molecule chemistry for local and long-distance defense signaling. Following pathogen attack, plants activate innate immune pathways at the site of infection to limit pathogen growth. Plants also possess the ability to prime similar immune responses in uninfected tissues to prevent the spread of pathogens or protect against new infections. Despite the importance of systemic immunity, the mechanism for signaling is not clear. In this study, we show that N-hydroxy-pipecolic acid metabolites are mobile defense signals produced at the site of bacterial infection and establish and amplify defense in uninfected, distal tissues. Our study illuminates the chemical nature of a mobile bioactive metabolite that confers pathogen resistance throughout the plant.

Abstract

Systemic acquired resistance (SAR) is a global response in plants induced at the site of infection that leads to long-lasting and broad-spectrum disease resistance at distal, uninfected tissues. Despite the importance of this priming mechanism, the identity and complexity of defense signals that are required to initiate SAR signaling is not well understood. In this paper, we describe a metabolite, N-hydroxy-pipecolic acid (N-OH-Pip) and provide evidence that this mobile molecule plays a role in initiating SAR signal transduction in Arabidopsis thaliana. We demonstrate that FLAVIN-DEPENDENT MONOOXYGENASE 1 (FMO1), a key regulator of SAR-associated defense priming, can synthesize N-OH-Pip from pipecolic acid in planta, and exogenously applied N-OH-Pip moves systemically in Arabidopsis and can rescue the SAR-deficiency of fmo1 mutants. We also demonstrate that N-OH-Pip treatment causes systemic changes in the expression of pathogenesis-related genes and metabolic pathways throughout the plant and enhances resistance to a bacterial pathogen. This work provides insight into the chemical nature of a signal for SAR and also suggests that the N-OH-Pip pathway is a promising target for metabolic engineering to enhance disease resistance.

Plants have developed a complex and dynamic innate immune system that relies on sensing and signaling using small molecules for defense against pathogens (1, 2). At a primary site of infection, plants respond to common molecular features of microbes (e.g., bacterial flagellin or fungal chitin, collectively known as microbial-associated molecular patterns or MAMPs) and pathogen-derived proteins, termed effectors, that enter the plant cell (3). Immune responses to both classes of molecules (effector-triggered or pattern-triggered; refs. 3 and 4) activate signal transduction networks through the action of hormones such as salicylic acid (SA), ethylene, and jasmonic acid, which cause changes in defense gene expression and production of antimicrobial metabolites at the site of infection (5). Interactions between these hormones function synergistically and antagonistically to tailor a specific immune response to different pathogens (2, 6). While local defense is critical for limiting pathogen growth, plants also possess the ability to prime and amplify immune responses at distal sites. This global response is termed systemic acquired resistance (SAR) (2, 7). SAR is critical for preventing the spread of pathogens and protecting against new infections (1, 2). Although much is understood about how the immune response is activated locally, the chemical nature of the plant-derived molecules that mediate long-distance communication between the site of infection (primary) and distal uninfected (secondary, systemic) sites remain elusive.

Genetic and mechanistic studies of the model plant Arabidopsis thaliana have led to the identification of key components of the SAR pathway, providing critical insight into the mechanisms controlling long-lasting and broad-spectrum disease resistance (7). Several plant-derived small molecules [e.g., SA (8), methyl salicylate (MeSA) (9), azelaic acid (AzA) (10), glycerol-3-phosphate (G3P) (11), dehydroabietinal (DA) (12), and pipecolic acid (Pip) (13)] are associated with long-distance communication and signal amplification during SAR (8, 14, 15). However, the onset of SAR signaling also requires the uncharacterized enzyme FLAVIN-DEPENDENT MONOOXYGENASE 1 (FMO1) (16–18). Remarkably, treatment of fmo1 mutants with AzA, DA, or Pip does not elicit systemic resistance, suggesting that a metabolite produced by FMO1 plays a key role in the establishment of SAR signaling (10, 12, 13). The biochemical function of FMO1 has remained unknown.

Several forward genetic screens searching for SAR-deficient mutants identified multiple alleles of fmo1, ald1 (AGD2-LIKE DEFENSE RESPONSE PROTEIN 1), and sard4 (SAR-DEFICIENT 4), highlighting the importance of metabolites produced by these enzymes (16, 17, 19, 20). ALD1 and SARD4 are involved in the biosynthesis of Pip (13, 21, 22). Irrigation of wild-type Arabidopsis plants with Pip induces SAR (13), which suggested that Pip might be a mobile SAR metabolite. Návarová et al. (13) reported, however, that Pip could not trigger SAR in fmo1 mutants. In addition, fmo1 plants accumulate high levels of Pip during a late stage of infection compared with wild-type plants (13). These findings feature FMO1 as a key missing link in the mechanism of Pip-associated SAR.

We and others have found untargeted metabolite analysis of Arabidopsis genetic mutants to be a powerful approach for the identification of small molecules associated with fitness phenotypes (either previously characterized or suggested by transcriptome analysis) (23). Examples include the identification and characterization of cytochromes P450 involved in phytoalexin production (24) and iron acquisition (25). Given that FMO1 is one of the genes most responsive to biotic stress (as indicated by analysis of previously reported Arabidopsis microarray data summarized in SI Appendix, Fig. S9) and that genetic data suggest molecules generated by FMO1 are required for initiating SAR (16–18), we sought to apply an untargeted metabolomics approach to determine the products of FMO1 and their function.

Here, we report the discovery of glycosylated N-hydroxypipecolic acid (N-OGlc-Pip) in Arabidopsis. We provide evidence that the aglycone, N-OH-Pip, is the direct product of FMO1 and has a central role in SAR signal transduction (26). We also demonstrate that exogenously applied N-OH-Pip can rescue the SAR-deficient response of fmo1 mutants, cause systemic changes in expression of pathogenesis-related genes and metabolic pathways throughout the plant (and distal to the site of application), and enhance resistance to the bacterial pathogen, Pseudomonas syringae. In addition, we provide biochemical evidence that Arabidopsis FMO1 catalyzes N-hydroxylation of the nonproteinogenic amino acid Pip to N-OH-Pip using transient expression assays in Nicotiana benthamiana. Taken together, our data indicate that N-OH-Pip is a key signaling molecule that is required to initiate SAR in Arabidopsis.

Results

Untargeted Metabolomics of Arabidopsis fmo1 Seedlings.

To discover metabolites whose production is dependent on FMO1, we used liquid chromatography-mass spectrometry (LC-MS)-based untargeted metabolomics to compare the composition of methanolic extracts from two sets of 12-d-old seedlings grown hydroponically: A. thaliana Col-0 WT and a T-DNA insertion mutant of FMO1, fmo1-1 (16), herein referred to as fmo1. The bacterial pathogen P. syringae pathovar tomato DC3000 (Pst) was added to the seedling media 24 h before metabolite analysis to elicit expression of enzymes associated with defense and SAR (including FMO1). This analysis revealed a major mass signal that is present in WT plants in response to Pst treatment but not Mock treatment (10 mM MgCl₂) and was absent from all fmo1 plants (Fig. 1A and SI Appendix, Fig. S1 A and B). We propose this signal corresponds to the O-glycosylated form of the metabolite N-OH-Pip, based on comparison of the MS/MS spectrum to an authentic synthetic standard of the N-OH-Pip aglycone (SI Appendix, Figs. S2 and S3). Although N-OH-Pip has not previously been observed as a naturally occurring metabolite in plants or other organisms, pipecolic acid has been observed in Arabidopsis and tomato (13, 27) and was shown to be associated with the SAR response (13). Therefore, we hypothesized that FMO1 catalyzes the N-hydroxylation of Pip to produce N-OH-Pip. Glycosylated N-OH-Pip (N-OGlc-Pip) detectable in Arabidopsis plant extracts is likely the product of unknown UDP glycosyltransferases that further processes N-OH-Pip (Fig. 1C). We did not detect free N-OH-Pip in WT seedlings elicited with Pst by LC-MS analysis (SI Appendix, Fig. S1B). These data suggest that the N-OH-Pip aglycone does not accumulate in cells and/or is an unstable metabolite.

Fig. 1.

Untargeted metabolomics of Arabidopsis seedlings elicited with Pst implicates FMO1 in the production of pipecolic acid derivatives. (A) Ion abundances for N-OGlc-Pip (gray bars) detected in extracts isolated from Arabidopsis Col-0 WT and fmo1 seedlings grown hydroponically and elicited with P. syringae pv. tomato DC3000 (Pst). Levels represent the mean ± STD of six biological replicates. Levels reported as zero indicate no detection of metabolites. (B) Ion abundances for N-OGlc-Pip (gray bars) in lower (L) and upper (U) leaves of adult Arabidopsis Col-0 plants 48 hpi with a 5 × 10⁶ cfu/mL suspension of Pst expressing the T3S effector avrRpt2 (Pst avrRpt2). Levels represent the mean ± STD of six biological replicates. Levels reported as zero indicate no detection of metabolites. (C) Proposed biosynthetic activity for Arabidopsis FMO1. The Arabidopsis enzymes ALD1 and SARD4 convert lysine to pipecolic acid (Pip) (22). FMO1 is proposed to hydroxylate Pip to N-OH-Pip. Unknown enzymes are proposed to glycosylate N-OH-Pip to produce N-OGlc-Pip, the molecule identified in the untargeted metabolite analysis.

Measurement of N-OH-Pip Derivatives in Adult Plants.

We also measured the level of N-OGlc-Pip in adult WT and fmo1 plants (both in lower and upper leaves), after treatment of lower leaves with 10 mM MgCl₂ (Mock) or Pst expressing the type III secretion (T3S) effector gene avrRpt2 (Pst avrRpt2). Pst avrRpt2 is an avirulent strain that induces effector-triggered immunity (ETI) and SAR signaling in resistant RPS2 Arabidopsis plants (28–30). N-OGlc-Pip was detected in the lower and upper leaves of WT only after infection of lower leaves with Pst avrRpt2, but not in fmo1 plants (Fig. 1B). As in seedlings, we were not able to detect free N-OH-Pip by LC-MS. Collectively, these data indicate that FMO1 is required for the production of N-OGlc-Pip in Arabidopsis.

Biochemical Activity of Arabidopsis FMO1.

Arabidopsis FMO1 expressed and purified as a 6×-His fusion protein (FMO1-6×-His) from Escherichia coli lacked the cofactor FAD and was not catalytically active in any of our in vitro biochemical assays. However, we were able to detect FMO1-dependent activity when Arabidopsis FMO1 was transiently expressed in N. benthamiana only after feeding pipecolic acid and then analyzing products present in leaf methanolic extracts (Fig. 2 and SI Appendix, Fig. S1 C–E). As shown in Fig. 2, FMO1-expressing leaves accumulated free N-OH-Pip. They also contained a second metabolite with an m/z of 100 and a putative structure of N-hydroxy piperideine (Fig. 2 and SI Appendix, Fig. S4), which we propose is the result of oxidative decarboxylation of N-OH-Pip. We noted that the mass signal m/z 100 can be produced in samples of synthetic N-OH-Pip after heating in buffer. It is also present in Col-0 seedlings after Pst treatment and absent from fmo1 seedlings (SI Appendix, Fig. S1B), suggesting that the m/z 100 metabolite is likely FMO1-derived. In addition, N-OH-Pip levels in N. benthamiana extracts decreased between 28 and 48 h after infiltration (hpi) while the abundance of m/z 100 increased (SI Appendix, Fig. S1C), further suggesting that N-OH-Pip is unstable in planta and may convert to m/z 100 over time. N. benthamiana leaves expressing two FMO1 active-site mutants, FMO1 G17A/G19A or FMO1 G215A, did not produce N-OH-Pip or m/z 100 at 28 hpi (Fig. 2 and SI Appendix, Fig. S1 D and E). The abundance of the mutant proteins was less than WT FMO1 at this time point (SI Appendix, Fig. S1F). At 48 hpi, protein abundance was similar (SI Appendix, Fig. S1F); however, N-OH-Pip was not detected in extracts expressing FMO1 G17A/G19A or FMO1 G215A (SI Appendix, Fig. S1E). Taken together, these data demonstrate that FMO1 can catalyze the hydroxylation of Pip in planta (Fig. 2) and support the requirement of the putative FAD and NADP⁺ domains for FMO1 catalytic activity (18). N-OGlc-Pip was not detected in N. benthamiana leaf extracts when Arabidopsis FMO1 was expressed, suggesting that N. benthamiana either does not have the necessary glycosyl transferases or that they are not expressed under our experimental conditions.

Fig. 2.

Overexpression of Arabidopsis FMO1 in N. benthamiana converts Pip to N-OH-Pip. Ion abundance of Pip (green bars), N-OH-Pip (blue bars), and an unknown metabolite, m/z = 100.075 (orange bars) in leaves of N. benthamiana expressing Arabidopsis Col-0 WT FMO1-6×-His or two mutants, FMO1-G17A/G19A-6×-His (a FAD binding mutant) or FMO1-G215A-6×-His (a NADP⁺ binding mutant). One millimolar Pip was coinfiltrated with an A. tumefaciens strain carrying FMO1-6×-His, FMO1-G17A/G19A-6×-His, or FMO1-G215A-6×-His, and then leaves were harvested 28 hpi. N-OH-Pip and the unknown metabolite were identified as FMO1-dependent signals using an untargeted analysis. Levels represent the mean ± STD of three biological replicates. Levels reported as zero indicate no detection of metabolites. Pathway represents proposed reactions occurring in N. benthamiana.

N-OH-Pip Treatment of Arabidopsis Leaves Is Sufficient to Induce SAR.

To test if N-OH-Pip treatment alone is sufficient to induce SAR and rescue the SAR deficiency of fmo1 mutants, we infiltrated three lower leaves of Arabidopsis WT and fmo1 plants with 10 mM MgCl₂ (Mock) or 10 mM MgCl₂ containing 1 mM Pip or 1 mM N-OH-Pip. We used 1 mM of each metabolite because it was previously established that watering plants with 1 mM Pip elicits SAR in the leaves of WT but not fmo1 mutants (13). After chemical incubation for 24 h, one upper leaf of each plant was inoculated with a 1 × 10⁵ cfu/mL suspension of P. syringae pathovar maculicola strain ES4326 (Psm ES4326), a virulent bacterium (Fig. 3A). At 3 d post inoculation (dpi), bacterial growth was quantified (Fig. 3B), and leaf symptoms were photographed (Fig. 3C). Both WT and fmo1 plants treated with N-OH-Pip contained significantly less Psm ES4326 in infected upper leaves compared with plants treated with Mock or Pip (Fig. 3B). N-OH-Pip treatment also reduced symptom development (i.e., leaf yellowing and tissue collapse) (Fig. 3C). These results suggest that treatment of leaves with N-OH-Pip, but not Pip, is sufficient to induce SAR and this does not require FMO1.

Fig. 3.

Infiltration of N-OH-Pip into Arabidopsis Col-0 WT and fmo1 leaves inhibits growth of virulent Psm ES4326 in distal leaves. (A) Photograph showing a representative plant and leaves used for SAR experiments. Typically, leaf numbers 7, 8, and 9 were used for chemical treatments (treated lower leaf) and leaf numbers 11, 12, or 13 was challenged with bacteria (upper leaf). (B) Titer of Psm ES4326 in challenged upper leaves of Arabidopsis WT and fmo1 plants. Three lower leaves of Col-0 WT and fmo1 were infiltrated with 10 mM MgCl₂ (Mock) or 10 mM MgCl₂ containing 1 mM Pip or 1 mM N-OH-Pip. After chemical incubation for 24 h, one upper leaf of each WT and fmo1 plant was inoculated with a 1 × 10⁵ cfu/mL suspension of Psm ES4326. The number of Psm ES4326 was quantified 3 d later to measure resistance of leaves. Bars (black, WT; gray, fmo1) represent the mean ± STD of four biological replicates. Asterisks denote the significant differences between indicated samples using a one-tailed t test (**P < 0.01). The experiment was repeated twice with similar results. (C) Symptoms of Arabidopsis WT and fmo1 upper leaves infected with Psm ES4326 following the chemical treatment described in B. Leaves were photographed 3 dpi. The SAR experiment was repeated twice with similar results.

N-OH-Pip Treatment of Arabidopsis Leaves Is Sufficient To Induce Metabolite Production and Gene Transcription Associated with SAR.

Next, we examined the ability of N-OH-Pip to systemically induce the production of SAR-associated metabolites and mRNAs in Arabidopsis leaves. Three lower leaves of WT and fmo1 plants were infiltrated with 10 mM MgCl₂ (Mock) or 10 mM MgCl₂ containing 1 mM Pip or 1 mM N-OH-Pip. After 48 h, the three treated lower leaves and three untreated upper leaves were pooled independently, and then analyzed by LC-MS and quantitative real-time PCR (qRT-PCR) for measurement of metabolites and mRNAs, respectively.

For metabolite profiling, we quantified Pip, N-OH-Pip, N-OGlc-Pip, m/z 100, and two canonical defense metabolites, the phytoalexin camalexin and SA-Glucoside (SA-Glc) (Fig. 4 A and B). After infiltration of the lower leaves of both WT and fmo1 plants with N-OH-Pip, we observed accumulation of camalexin and SA-Glc in both lower and upper leaves (Fig. 4B). Our findings are consistent with reports showing that FMO1 is required to stimulate camalexin and SA-Glc synthesis and accumulation in distal, uninfected leaves during SAR (31).

Fig. 4.

N-OH-Pip induces expression of SAR marker genes and metabolites in treated lower leaves and untreated upper leaves of Arabidopsis Col-0 WT and fmo1 plants. (A) Proposed pathway of metabolite and gene induction in N-OH-Pip–associated SAR. In this study, FMO1 is hypothesized to convert Pip to N-OH-Pip and unknown enzymes convert N-OH-Pip to N-OGlc-Pip and m/z = 100.075. Pip biosynthesis requires ALD1 activity and is partially dependent on SARD4 activity (13, 21, 22). N-OH-Pip treatment leads to the accumulation of camalexin and ALD1, SARD4, FMO1, ICS1, and PR2 mRNAs. ICS1 converts chorismic acid to isochorismic acid, leading to SA production. SA is required for camalexin production in during SAR (31), and SA induces the expression of NPR1. NPR1 is a transcription factor that regulates expression of PR1 and other defense-associated genes. In plants, SA-Glucoside (SA-Glc) is thought to be nontoxic storage form of SA (40). (B) Ion abundance of Pip, N-OGlc-Pip, m/z = 100.075, camalexin and SA-Glc in leaves. Three lower leaves of Col-0 WT and fmo1 were infiltrated with 10 mM MgCl₂ (Mock) or 10 mM MgCl₂ containing 1 mM Pip or 1 mM N-OH-Pip. After chemical incubation for 48 h, three treated lower (L) leaves and three untreated upper (U) leaves of WT and fmo1 were analyzed by metabolite profiling and qPCR (C). Bars (black, WT; gray, fmo1) represent the mean ± STD of three biological replicates. (C) Relative expression of SAR marker genes in leaves: ALD1 (AT2G13810), AGD2-like defense response protein 1; SARD4 (AT5G52810), SAR deficient 4; FMO1 (At1g19250), flavin-dependent monooxygenase 1; ICS1 (At1g74710), isochorismate synthase 1; PR1 (At2g14610), pathogenesis-related protein 1 and PR2 (At3g57260), β-1,3-glucanase 2. Bars (black, WT; gray, fmo1) represent the mean ± STD of three biological replicates. The experiment was repeated twice with similar results.

Although we did not detect free N-OH-Pip, we observed accumulation of N-OGlc-Pip and m/z 100 in both lower and upper leaves of all plants (Fig. 4B), suggesting rapid metabolism to these derivatives. These data show that local treatment of purified N-OH-Pip alone, even in the absence of infection, can induce systemic changes in defense metabolite production across the plant and this does not require FMO1. In contrast, the infiltration of lower leaves of WT or fmo1 plants with Pip did not induce changes in N-OH-Pip metabolites, camalexin, or SA-Glc. Given that fmo1 plants are unable to produce N-OH-Pip, the detection of N-OH-Pip metabolites (N-OGlc-Pip and m/z 100) in the upper leaves after N-OH-Pip treatment of lower leaves (Fig. 4B) provides evidence that one or more of these metabolites are mobile. Taken together, these data indicate that N-OH-Pip or rapidly formed derivatives move systemically through the plant to induce metabolic changes during SAR.

For transcript profiling, we quantified the mRNA abundance for known SAR marker genes, which were reported to be induced in untreated upper leaves of WT plants but not fmo1 plants in response to bacterial infection (i.e., Psm ES4326) of lower leaves (32). These included genes dependent on SA [pathogenesis-related protein (PR) 1], partially dependent on SA (PR2 and PR5), or independent of SA (ALD1; AGD2-like defense response protein 1; SAG13, senescence-associated gene 13; and ICS1, isochorismate synthase 1). We also monitored ALD1, SARD4 (SAR deficient 4 protein), and FMO1, three genes in the proposed N-OH-Pip metabolic pathway (Figs. 1C and 4A).

WT and fmo1 plants treated with N-OH-Pip accumulated the highest levels of ICS1 and PR1 mRNA in treated lower and untreated upper leaves, compared with plants treated with Mock or Pip (Fig. 4C). Similar trends were observed for ALD1, SARD4, PR2, PR5, and SAG13 mRNAs (Fig. 4C and SI Appendix, Fig. S5). N-OH-Pip–triggered accumulation of ALD1 and SARD4 mRNA suggests that N-OH-Pip regulates ALD1 and SARD4 transcription via a positive feedback loop. Accumulation of Pip in leaves of WT and fmo1 plants treated with N-OH-Pip implies that N-OH-Pip induction of ALD1 and SARD4 transcription leads to increased synthesis of Pip in leaves. In addition, N-OH-Pip treatment increased FMO1 mRNA levels in WT leaves but not the fmo1 mutant (Fig. 4C). Our inability to detect transcript for this fmo1 mutant is consistent with it being a null mutant (16, 18). This implies that N-OH-Pip also positively regulates FMO1 transcription via a positive feedback loop. Notably, induction of SAR gene expression was greater in N-OH-Pip–treated plants compared with Pip-treated plants (Fig. 4C and SI Appendix, Fig. S5), demonstrating that N-OH-Pip is a more bioactive SAR molecule compared with Pip.

Collectively, these data demonstrate that exogenous treatment of N-OH-Pip by leaf infiltration is sufficient to activate metabolite production and SAR-associated gene transcription.

N-OH-Pip Irrigation Induces SAR in Arabidopsis WT and fmo1 Plants.

Previously, Návarová et al. (13) showed that irrigation with Pip was sufficient to recover SAR activity in ald1 plants but not in fmo1 plants. We tested if irrigation with N-OH-Pip could similarly induce SAR in WT and fmo1 plants. SAR assays were performed as described previously (13) with slight modification. WT and fmo1 plants were drenched with water, 1 mM Pip, or 1 mM N-OH-Pip by root application. One day later, three lower (local) leaves per plant were inoculated with 10 mM MgCl₂ (Mock) or a 5 × 10⁶ cfu/mL suspension of Pst avrRpt2, an avirulent strain. Two days later, the untreated upper leaves of each plant were challenged with Mock or a 1 × 10⁵ cfu/mL suspension of Psm ES4326, a virulent strain. The titer of Psm ES4326 in the infected upper leaves was determined at 3 dpi to quantify the impact of Pip and N-OH-Pip treatment on the level of disease resistance. An overview of the SAR experiment is shown in Fig. 5.

Fig. 5.

Root application of N-OH-Pip elicits local defense and SAR in Arabidopsis Col-0 WT and fmo1 plants. (A) Experimental design of SAR assay. Each Col-0 WT and fmo1 plant was treated with 10 mL of water, 1 mM Pip, or 1 mM N-OH-Pip by root application. One day later, three lower leaves of each plant were infiltrated with 10 mM MgCl₂ or a 5 × 10⁶ cfu/mL suspension of Pst DC3000 expressing avrRpt2 (Pst avrRpt2) in 10 mM MgCl₂. Two days later, one upper leaf of each plant was inoculated with a 1 × 10⁵ cfu/mL suspension of Psm ES4326. The number of Psm ES4326 in upper leaves was quantified 3 d later. (B) Phenotype of lower leaves inoculated with Pst avrRpt2 after chemical treatment. Phenotypes of four biological replicates were recorded 2 dpi with Pst avrRpt2. Similar phenotypes were observed in three independent experiments. (C) Titer of Psm ES4326 in upper leaves of Col-0 WT (black bars) and fmo1 (gray bars). Bars represent the mean ± STD of three biological replicates. Asterisks denote the significant differences between indicated samples using a one-tailed t test (**P < 0.01; *P < 0.05; ns, not significant). The experiment was repeated three times with similar results.

Before challenging upper leaves with Psm ES4326, lower leaves infected with Pst avrRpt2 were yellow (chlorotic) and collapsed by 2 dpi (Fig. 5B). We noticed that these symptoms were significantly delayed in WT and fmo1 plants irrigated with N-OH-Pip (Fig. 5B). Pip irrigation reduced leaf symptom development in WT plants to a lesser degree, but not in fmo1 plants (Fig. 5B). Metabolic profiling revealed that N-OGlc-Pip and m/z 100 were present in Pst avrRpt2-infected leaves of WT plants irrigated with Pip or N-OH-Pip, and fmo1 plants irrigated with N-OH-Pip (SI Appendix, Fig. S6). These results indicate that N-OH-Pip treatment reduces symptom development of Arabidopsis leaves in an FMO1-independent manner. The delay of symptom development caused by N-OH-Pip and Pip could be attributed to metabolite-induced defense priming in infected leaves or inherent bactericidal activity of the metabolites.

After challenging upper leaves with Psm ES4326, the titer of Psm ES4326 in water-treated WT plants infected with Pst avrRpt2 was lower than that in water-treated WT plants inoculated with Mock (Fig. 5C). These data indicate that Pst avrRpt2 induces SAR under the conditions tested. By contrast, Psm ES4326 titers were similar in water-treated fmo1 plants inoculated with Mock or Pst avrRpt2 (Fig. 5C). These data are consistent with previous findings that fmo1 mutants are deficient in SAR signaling (16, 18).

Irrigating plants with Pip and N-OH-Pip enhanced SAR, albeit to different extents. Pip absorption by roots provided a moderate but significant reduction in Psm ES4326 titer in Mock and Pst avrRpt2-inoculated fmo1 plants (Fig. 5C). However, the level of protection was less than SAR-induced protection elicited by Pst avrRpt2 in water-treated WT plants. Interestingly, N-OH-Pip absorption by roots provided the strongest level of protection (Fig. 5C). Psm ES4326 titers in Mock and Pst avrRpt2-inoculated fmo1 plants were significantly less than those detected in water- or Pip-treated fmo1 plants. Moreover, N-OH-Pip-induced protection in fmo1 plants was similar to that observed for Pst avrRpt2-induced protection in WT plants (Fig. 5C). Collectively, these data indicate that N-OH-Pip is a potent SAR-inducing metabolite, and irrigation of plants with N-OH-Pip is also sufficient to establish SAR in WT plants and complement the SAR defect of fmo1 plants.

N-OH-Pip Is Not Bactericidal.

To rule out that N-OH-Pip is toxic to bacterial growth in planta, we performed a minimum inhibitory concentration (MIC) assay (33) to measure potential bactericidal activity. Cultures of Psm ES4326, Pst DC3000, and Pst avrRpt2 were incubated with different concentrations (0–1 mM) of Pip, N-OH-Pip, or SA and then grown at 28 °C for 36 h. SA was included as a positive control because it inhibits multiplication of Pseudomonas aeruginosa and Agrobacterium tumefaciens in vitro (34, 35). Similar to previous studies, 1 mM SA inhibited the multiplication of Psm ES4326, Pst DC3000, and Pst avrRpt2 in vitro (SI Appendix, Fig. S7). However, neither Pip nor N-OH-Pip inhibited Psm ES4326, Pst DC3000, and Pst avrRpt2 growth in vitro. These data indicate that Pip and N-OH-Pip are not bactericidal.

N-OH-Pip–Treated Arabidopsis Plants Exhibit a Faster Hypersensitive Response.

Next, we investigated if N-OH-Pip enhances ETI in local, infected leaves by performing electrolyte leakage and HR assays. WT and fmo1 plants were irrigated with water, 1 mM Pip, or 1 mM N-OH-Pip. One day later, six to seven leaves of each plant were inoculated with a 3 × 10⁸ cells per mL suspension of Pst DC3000 carrying an empty vector (Pst vector) or expressing avrRpt2 (Pst avrRpt2). Ion leakage was quantified at 5 hpi (Fig. 6A), and HR symptoms were recorded at 8 hpi (Fig. 6B).

Fig. 6.

AvrRpt2-elicited HR phenotype occurs earlier in Arabidopsis Col-0 WT and fmo1 plants treated with N-OH-Pip compared with water or Pip. Col-0 WT and fmo1 plant pots were treated with 10 mL of water, 1 mM Pip, or 1 mM N-OH-Pip by root application. Twenty-four hours later, leaves of Col-0 WT and fmo1 plants were infiltrated with a 3 × 10⁸ cells per mL suspension of Pst DC3000 with an empty vector (Pst vector) or avrRpt2 (Pst avrRpt2). (A) Percent ion leakage of inoculated leaves at 5 hpi. Bars (black, WT; gray, fmo1) represent the mean ± STD of nine randomly inoculated leaves of three WT or fmo1 plants. Asterisks denote the significant differences between indicated samples using a one-tailed t test (**P < 0.01; *P < 0.05). (B) HR phenotypes of inoculated leaves at 8 hpi. Fraction refers to number of leaves showing HR of 10 randomly inoculated leaves. Experiments were repeated three times with similar results.

WT and fmo1 leaves infected with Pst vector exhibited similar electrolyte leakage (below 20%) when irrigated with water, Pip, and N-OH-Pip (Fig. 6A), and no HR phenotypes were observed for these infected plants at 5 hpi (Fig. 6B). WT plants irrigated with water and infected with Pst avrRpt2 exhibited more electrolyte leakage at 5 hpi than similarly treated fmo1 plants (Fig. 6A). HR symptoms were not observed at 8 hpi (Fig. 6B); however, full leaf collapse occurred by 10 hpi. Compared with water irrigation, Pip irrigation resulted in a significant increase in electrolyte leakage from WT leaves infected with Pst avrRpt2 (Fig. 6A), with most (7/10) of the leaves undergoing HR by 8 hpi (Fig. 6B). No significant changes were detected for similarly treated fmo1 leaves. Notably, N-OH-Pip irrigation resulted in a significant increase in electrolyte leakage from both WT and fmo1 leaves infected with Pst avrRpt2 (Fig. 6A). HR phenotypes for the N-OH-Pip–irrigated plants appeared faster and were more severe than those observed for the Pip-irrigated plants (Fig. 6B). Taken together, these results show that N-OH-Pip treatment stimulates AvrRpt2-elicited HR in plants compared with water and Pip treatment, demonstrating that N-OH-Pip treatment also affects ETI.

Discussion

FMO1 Produced Metabolite N-OH-Pip Activates SAR.

FMO1-dependent regulation of defense signaling has emerged as a critical determinant of the establishment and amplification of SAR in Arabidopsis (16, 31, 32). The SAR deficiency of fmo1 mutants could be due to a failure to produce a priming signal, to recognize defense signal(s), or to transmit defense signal(s) from infected tissue to uninfected tissue. Our discovery that FMO1 produces N-OH-Pip, a bioactive metabolite that is mobile in plants and confers pathogen resistance in systemic tissues, reveals that the key biological function of FMO1 is to produce the priming signal for SAR. The accumulation of N-OH-Pip–derived metabolites (N-OGlc-Pip and m/z 100) in untreated, distal leaves of fmo1 plants following infiltration of N-OH-Pip in local leaves indicates that one or more of these metabolites are transported during SAR.

N-OH-Pip establishment of SAR in Arabidopsis is characterized by the induction of transcriptional and metabolic programs in systemic plant tissue. Application of N-OH-Pip to local leaves resulted in the robust activation of several SA-dependent and SA-independent SAR marker genes in distal leaves (Fig. 4C and SI Appendix, Fig. S5). This is consistent with reports showing that FMO1 is required for SA-dependent and SA-independent defenses (31, 32). N-OH-Pip also activated transcription of the N-OH-Pip biosynthetic genes (ALD1, SARD4, and FMO1) in both local and systemic leaves (Fig. 4C), highlighting the importance of transcriptional feedback regulation for amplifying N-OH-Pip–dependent signaling during SAR.

Remarkably, disease symptoms in Arabidopsis leaves were dramatically reduced in leaves of bacterially infected plants treated with N-OH-Pip (Figs. 3C and 5B). We speculated that N-OH-Pip might directly affect bacterial growth and/or the ability of bacteria to deliver T3S effectors to plant cells. However, N-OH-Pip was not toxic to pathogen growth in culture (SI Appendix, Fig. S7) and did not inhibit the activation of ETI in resistant Arabidopsis plants (Fig. 6). These data suggest that N-OH-Pip stimulates or enhances plant-specific processes in infected leaves to prevent their collapse and yellowing. What these physiological processes are remains to be determined.

We also found that N-OH-Pip–treated leaves displayed enhanced resistance in SAR assays (Fig. 5C) and are highly sensitized for ETI (Fig. 6). This suggests that the level of N-OH-Pip in tissues plays a critical role in establishing the magnitude and timing of the disease resistance response. Consistent with our findings, overexpression of FMO1 in Arabidopsis resulted in enhanced basal resistance to the oomycete Hyaloperonospora parasitica and P. syringae pv. DC3000 (Pst), as well as enhanced resistance to Pst avrRpt2 (17). In contrast, loss of FMO1 function resulted in enhanced susceptibility to Pst and H. parasitica (17, 18).

Biochemistry of FMO1 and Chemistry of Pip Hydroxylation.

Flavoprotein monooxygenases (FMOs) catalyze incorporation of one atom of molecular oxygen onto a nucleophilic substrate (36). Similar to plant cytochromes P450 that activate and transfer oxygen, the FMO family of genes has greatly expanded in plants, suggesting they play critical roles in metabolism and fitness. In Arabidopsis, 29 genes encode FMO-like proteins, compared with only 5 in animals (37). These proteins have been divided into three clades: Clade 1 includes FMO1 and a pseudogene. Clade 2 includes the YUCCA group of 11 enzymes, which have been well-studied and are associated with auxin biogenesis. Clade 3 has 16 enzymes, including an enzyme that S-oxygenates glucosinolates, molecules critical for pathogen defense (37). Of these, FMO1 is notably one of the most overexpressed metabolic genes after pathogen challenge (SI Appendix, Fig. S9). Our discovery that the FMO1 product N-OH-Pip is a bioactive SAR metabolite is further evidence that FMOs have a privileged role in plant defense metabolism. Moreover, it predicts that direct products of FMO1 homologs in other plants may be novel natural products involved in defense priming.

Numerous nonproteinogenic amino acids have been described in biological systems (38), yet N-OH-Pip has not previously been reported. It remains a mystery why N-oxidation of pipecolic acid, a metabolite produced in both plants and bacteria, has evolved as a key step in generating an active signaling molecule for SAR. The hydroxylamine functionality bears resemblance to the hydroxamic acids required for binding iron in bacterial siderophores, and intriguingly, some bacterial siderophores are now thought to be involved in signaling (39). Further studies are required to determine if the hydroxylamine substituent of N-OH-Pip has a functional role in its transport, perception, and/or metabolism.

Results from our transient expression experiments in N. benthamiana indicate that N-OH-Pip may be unstable in planta as FMO1-dependent levels of N-OH-Pip decreased from 28 to 48 h while levels of m/z 100 increased (SI Appendix, Fig. S1D). We speculate that O-glycosylation of N-OH-Pip could serve as a stabilization mechanism and that N-OGlc-Pip may be the primary storage form of the molecule in Arabidopsis. This proposed process is reminiscent of SA glycosylation and storage of SA-Glc in response to pathogen infection (40). While we cannot definitively rule out that m/z 100 is a product of extraction, its presence could indicate a passive or active attenuation strategy. For example, the level of N-OH-Pip could be controlled via conversion to N-OGlc-Pip for storage or m/z 100 for deactivation. Future work should investigate the precise roles of N-OH-Pip and its derivatives in SAR.

Presence of N-OH-Pip Biosynthesis Pathway in Other Plant Species.

Because SAR is a broad-spectrum defense strategy shared among many plants, we mined available plant genomes to determine the prevalence of enzymes involved in N-OH-Pip biosynthesis. We tabulated the best BLAST hit between the Arabidopsis SARD4, ALD1, and FMO1 proteins and those of sequenced plant genomes, and found that >94% of species analyzed had FMO1 homologs with greater than 50% amino acid identity (SI Appendix, Fig. S8). All plants within the Brassicaceae that we evaluated have enzymes with >88% amino acid identity to Arabidopsis SARD4, ALD1, and FMO1, and these plants contained the top five homologs to the Arabidopsis FMO1. In Arabidopsis, the closest homolog to FMO1 has only a 74% amino acid identity, and this protein originates from a likely pseudogene of FMO1 that is missing a large internal coding region. The next best BLAST hit has only a 26% amino acid identity to FMO1. The scarcity of FMO1 homologs in Arabidopsis suggests that the homologs in other plants (and especially those of the Brassicaceae) may have the same function. While the N-OH-Pip pathway may be broadly conserved, it is possible that this pathway may have arisen independently in the Brassicaceae and other metabolites may mediate defense signaling outside this plant family.

Is N-OH-Pip a Long-Distance Mobile Signal for SAR?

The molecule that initiates systemic defense signaling is still up for debate (7, 14). It is not clear if there is one molecule or a variety of molecules that orchestrate broad-spectrum plant disease resistance. Several mobile metabolites (MeSA, AzA, G3P, and DA) are known to prime resistance in uninfected tissues (7, 14) and some require DIR1 (a lipid transport protein) (10–12, 41), suggesting that DIR1 is required for the accumulation and/or transport of molecules that induce SAR. Notably, AzA, DA, and Pip require FMO1 for the initiation of SAR (10, 12, 13), highlighting the possibility that FMO1 may produce one of the elusive long-distance mobile signals critical for SAR. Our work demonstrates that the product of FMO1, N-OH-Pip, is necessary and sufficient to initiate SAR in local and systemic tissue, making it a prime candidate for a long-distance SAR signal. It remains to be determined how and where in the plant the N-OH-Pip signal is perceived to initiate SAR.

In conclusion, the discovery of N-OH-Pip as a mobile signal provides critical insight into how plants use small molecules to resist the spread of infection. This work provides an opportunity to address fundamental open questions in SAR biology, including the mechanism of transport, signal perception, and signal attenuation once a heightened defense response is no longer required. The sensitivity of plants to N-OH-Pip treatment also highlights the possibility for translating a chemical or metabolic engineering approach to prime or enhance disease resistance in plants under pathogen pressure.

Materials and Methods

Bacterial Strains and Growth Conditions.

P. syringae pv. maculicola strain ES4326 (Psm ES4326), P. syringae pv. tomato DC3000 (Pst DC3000), P. syringae pv. tomato carrying pVSP61 + avrRpt2 (Pst avrRpt2), P. syringae pv. tomato carrying pVSP61 vector (Pst vector), E. coli strain DH5 alpha, and A. tumefaciens strain C58C1 pCH32 were used in this study. Growth conditions for all bacterial strains are described in SI Appendix, Table S1.

Plant Materials and Growth Conditions.

A. thaliana ecotype Col-0 (WT) and fmo1 (SALK_026163) plants were grown in a controlled growth chamber (22 °C, 80% relative humidity, 103 µmol/m² per s of light intensity) on a 10-h light/14-h dark cycle. Genotypes were confirmed by PCR using FMO1 gene-specific primers and a T-DNA–specific primer (SI Appendix, Table S2). Four- to 5-wk-old adult plants were used for SAR assays, qRT-PCR, and metabolite profiling. For seedling hydroponics experiments, Arabidopsis seeds were sterilized by suspending seeds in 50% ethanol for 1 min, washing three times in sterile water, suspending in 50% bleach for 10 min, and washing three more times in sterile water. Seeds were resuspended in 1× Murashige–Skoog (MS) medium with vitamins (PhytoTechnology Laboratories) (pH 5.7) and vernalized for 48 h at 4 °C. Seeds (15 ± 1) were placed into 3 mL of MS medium + 5 g/L sucrose in wells of a six-well microtiter plate. Plates were sealed with Micropore tape (3M) and grown in a chamber at 50% humidity, 22 °C, and 100 µmol/m² per s photon flux under a 16-h light/8-h dark cycle. After 1 wk of growth, spent medium was replaced with 3 mL of fresh MS medium + 5 g/L sucrose and plants were grown for one additional week. N. benthamiana plants were grown in soil on a growth shelf with a 16-h light cycle for 5 wk.

Elicitation Methods for Seedling Hydroponics Experiments.

For bacterial elicitation, Pst was grown on LB agar plates at 30 °C. A single colony was picked and grown to exponential phase in liquid LB at 30 °C. Cells were then centrifuged at >13,000 × g and resuspended in sterile 1× MS media to an OD₆₀₀ of 0.2. Seventy microliters of the Pst suspension was added to each seedling-containing well for elicitation. Plants were elicited 24 h before sample harvest.

Plant Extraction for Metabolite Profiling.

Arabidopsis seedlings were lyophilized to dryness and ground using a ball mill (Retsch MM 400). A single 5-mm steel ball was added to each sample and samples were shaken at 25 Hz for 2 min. Samples were resuspended in 10 mL of 80:20 MeOH:H₂O per gram wet tissue and heated at 65 °C for 10 min. Samples were filtered through 0.45-µm polytetrafluoroethylene (PTFE) filters and analyzed via LC-MS. Adult leaf tissue was flash frozen and ground in liquid nitrogen using a mortar and pestle. Samples were resuspended in 10 mL of 80:20 MeOH:H₂O per gram wet tissue and heated at 65 °C for 10 min. Extracts were filtered through 0.45-µm PTFE filters and analyzed via LC-MS.

LC-MS Methods.

N-OGlc-Pip, SA-Glc, and camalexin were measured using reverse-phase chromatography on an Agilent 1260 HPLC coupled to an Agilent 6520 Q-TOF ESI mass spectrometer as previously described (24). A 5-μm, 2 × 100 mm Gemini NX-C18 column (Phenomenex) was used for separation. The mobile phases were A [water with 0.1% formic acid (FA)] and B [acetonitrile (ACN) with 0.1% FA] and the following gradient was implemented at a flow rate of 0.4 mL/min (percentages indicate percent B): 0–30 min (3–50%), 30–45 min (50–97%), 45–50 min (97%), 50–51 min (97–3%), 51–55 min (3%). The MS was run in positive ion mode with conditions as used previously (24). For MS/MS analysis, a fragmentor voltage of 10 V was used with an m/z window of 1.3.

Pip, N-OH-Pip, and hypothesized N-hydroxy piperideine were measured using hydrophilic interaction chromatography on an Agilent 1290 UHPLC coupled to an Agilent 6545 Q-TOF ESI mass spectrometer. A 130-Å, 1.7-µm, 2.1 mm × 50 mm Acquity UPLC BEH Amide column (Waters) was used for separation. The mobile phases were A (water, 10 mM ammonium formate, 0.125% FA) and B (95% ACN, 10 mM ammonium formate, 0.125% FA), and the following gradient was implemented at a flow rate of 0.6 mL/min (percentages indicate percent B): 0–1.5 min (100%), 1.5–6 min (100–40%), 6–8 min (40%), 8–8.5 min (40–30%), 8.5–9.5 min (30–100%), 9.5–12 min (100%). The MS was run in positive ion mode with the following parameters: mass range: 30–1,700 m/z; drying gas: 250 °C, 12 L/min; nebulizer: 10 psig; capillary: 3,500 V; fragmentor: 140 V; skimmer: 65 V; octupole 1 RF Vpp: 750 V; 333.3 ms/spectrum. For MS/MS analysis, a fragmentor voltage of 10 V was used with an m/z window of 1.3 for N-OH-Pip and a fragmentor voltage of 20 V with an m/z window of 1.3 was used for hypothesized N-hydroxy piperideine. The initial 0.5 min of each run was sent to waste to avoid salt contamination of the MS.

N-OH-Pip Chemical Synthesis.

N-OH-Pip was synthesized from l-pipecolic acid (98% purity; Oakwood Chemical) using a modified protocol (42). To begin, 10.1 g of l-pipecolic acid was added to a cooled solution of 4.93 g of 88% pure KOH (1 equivalent). Acrylonitrile (5.58 mL; 1.1 equivalents) was then added drop wise to the solution over 5 min. The solution was stirred for 1.5 h in an ice bath and another 1.5 h at room temperature. Then, the pH of the solution was adjusted to 6.6 with 12 M HCl to quench the KOH. A rotary evaporator was used to evaporate the solvent. Four hundred milliliters of acetone was added to the residue, and the solution was brought to a boil. After several minutes of boiling, the solution was filtered, and a rotary evaporator was used to remove approximately half of the solvent. Then, the residue from the filtration and fresh acetone was added back into the remaining filtrate, and the solution was brought to a boil, filtered, and set on the rotary evaporator. The cycle of heating the solution and filtering was repeated five times. Two hundred milliliters of filtrate from the fifth cycle was then stored at −20 °C overnight to recrystallize.

Crystallized product (2-cyanoethyl-pipecolic acid; 3.78 g) was mixed with 60 mL of MeOH and 4.80 g of 70% metachloroperoxybenzoic acid (mCPBA) (1 equivalent) was added to 20 mL of MeOH. The mCPBA solution was added drop wise to the cooled 2-cyanoethyl-pipecolic acid slurry over 30 min. The slurry was stirred for an additional hour in an ice bath, and as the reaction progressed, the slurry dissolved into solution. Then, 300 mL of precooled diethyl ether was added to the reaction and the mixture was stored at −20 °C overnight to recrystallize.

Crystallized product (2-cyanoethyl-pipecolic acid oxide; 0.5 g) was dissolved in 150 mL of acetone in a 250-mL flask with a short path distillation head. Acetone was slowly distilled drop by drop for 3 h, and fresh acetone was periodically added to keep the original volume. Then, a rotary evaporator was used to evaporate the majority of the solvent and the remainder was evaporated to dryness under reduced pressure.

¹H and ¹³C NMR spectra were taken of a 25 mM solution of N-OH-Pip in D₂O with a Varian Inova 500 NMR spectrometer. The following parameters were used for ¹H NMR spectra: temperature: ambient; probe: 5-mm PFG switchable; scan number: 16; receiver gain: 40; relaxation delay: 0; pulse width: 8; frequency: 499.75 Hz. The following parameters were used for ¹³C NMR spectra: temperature: ambient; probe: 5-mm PFG switchable; scan number: 120; receiver gain: 54; relaxation delay: 0.5; pulse width: 7; frequency: 125.67 Hz.

Chemical Treatment of Leaves for Bacterial Growth Assay.

Leaf numbering was performed according to refs. 43 and 44. Three lower leaves (leaf nos. 7–9) of WT and fmo1 Arabidopsis plants (4- to 5-wk-old) were infiltrated with 10 mM MgCl₂, 1 mM Pip in 10 mM MgCl₂, or 1 mM N-OH-Pip in 10 mM MgCl₂. After 24 h, one untreated upper leaf (leaf nos. 11, 12, or 13) of each plant was inoculated with a 1 × 10⁵ cfu/mL suspension of Psm ES4326. The inoculated plants were then covered with a dome to maintain humidity. The titer of Psm ES4326 in the upper leaves was quantified at 3 dpi by homogenizing leaves discs in 1 mL of 10 mM MgCl₂, plating appropriate dilutions on nutrient yeast glycerol medium supplemented with 1.5% wt/vol agar (NYGA) with rifampicin (100 µg/mL), incubating plates at 28 °C for 2 d, and then counting bacterial colonies. Four biological repeats were performed per treatment in two independent experiments.

Chemical Treatment of Leaves for qRT-PCR and Metabolite Profiling.

Three lower leaves (leaf nos. 7–9) of WT and fmo1 Arabidopsis plants (4- to 5-wk-old) were infiltrated with 10 mM MgCl₂, 1 mM Pip in 10 mM MgCl₂, or 1 mM N-OH-Pip in 10 mM MgCl₂. After 48 h, the three treated lower leaves and three untreated upper leaves (leaf nos. 11–13) were harvested, pooled, respectively, and then frozen in liquid nitrogen. Frozen tissue was pulverized and divided into two aliquots: one for qRT-PCR and the other for metabolic profiling. Three biological repeats were performed per treatment in two independent experiments.

RNA Isolation and qRT-PCR.

Total RNA was isolated from leaves using TRIzol reagent (Invitrogen) according to the manufacturer’s instructions. Two micrograms of RNA were used to synthesize cDNA by oligo dT and reverse transcriptase. For qRT-PCR, each cDNA sample was amplified with gene-specific primers (SI Appendix, Table S2) using Green Taq DNA polymerase (GenScript) with EvaGreen Dye (Biotium) and the MJ Opticon 2 (Bio-Rad). UBC21 (Ubiquitin-Conjugating Enzyme 21; At5g25760) mRNA abundance was used to normalize the expression value in each sample. The comparative Ct method (2^−∆∆Ct) was performed to determine the relative expression. The fold change of each value was normalized to the value of MgCl₂-treated local leaves of WT.

SAR Assay.

SAR bacterial growth assays were performed as described (13) with slight modification. Each plant pot was drenched with 10 mL of water, 1 mM l-(-)-Pipecolinic acid (Pip) (Oakwood) or 1 mM N-OH-Pip. After 24 h, three lower leaves of each plant were infiltrated with 10 mM MgCl₂ or a 5 × 10⁶ cfu/mL suspension of Pst avrRpt2 in 10 mM MgCl₂. Two days later, one upper leaf of each plant was inoculated with a 1 × 10⁵ cfu/mL suspension of Psm ES4326, and then plants were covered with a dome to maintain humidity. The titer of Psm ES4326 in the upper leaves was quantified at 3 dpi by homogenizing leaves discs in 1 mL of 10 mM MgCl₂, plating appropriate dilutions on NYGA medium with rifampicin (100 µg/mL), incubating plates at 28 °C for 2 d, and then counting bacterial colonies. Three plants were used per condition, and the experiment was repeated more than three times.

Construction of FMO1 Mutants.

The ORF of FMO1 lacking the stop codon was amplified from Arabidopsis Col-0 WT cDNA by PCR using FMO1-specific primers (SI Appendix, Table S2) and cloned into the pCR8/GW/TOPO vector (Life Technologies). Two alanine substitution mutants, FMO1(G17A/G19A) and FMO1(G215A), were generated using pCR8/GW/TOPO-FMO1 as template and fmo1 mutant primers (SI Appendix, Table S2). All constructs were confirmed by DNA sequence analysis. WT and mutant FMO1 cDNAs were subcloned into pEAQ-HT-DEST3 (45) to create C-terminal 6× His-tagged fusion proteins. Plasmids were introduced into E. coli DH5 alpha and A. tumefaciens C58C1 by heat shock transformation.

Transient Expression in N. benthamiana.

Agrobacterium strains harboring the pEAQ-gene constructs were grown on LB agar plates with the appropriate antibiotics. After 48 h of growth, cells were removed from plates using an inoculation loop and resuspended in 1 mL of LB. Cells were centrifuged at 4,000 × g for 5 min, the supernatant was removed, and cells were resuspended in 1 mL of Agrobacterium induction medium (10 mM MES buffer, 10 mM MgCl₂, 150 µM acetosyringone, pH 5.7) and incubated at room temperature with shaking for 2 h. Cells were then diluted to a final OD₆₀₀ of 0.3 in induction medium. In tests with supplemented Pip, cells were diluted to a final OD₆₀₀ of 0.3 in induction medium + 1 mM Pip. These solutions were then infiltrated into the underside of N. benthamiana leaves (three leaves per plant) using a needleless 1-mL syringe. Plants were grown on a growth shelf with a 16-h light/8-h dark cycle for 28 or 48 h before sample harvest for metabolic analysis and immunoblot. Total protein of each sample was extracted from two leaf discs (1-cm diameter per disk) by using urea buffer (8 M urea, 15% β-mercaptoethanol, 3× Laemmli buffer). Proteins were separated by 12% SDS/PAGE analysis and transferred to a PVDF membrane and visualized by Ponceau S red staining before immunoblot analysis. FMO1-6×-His, FMO1(G17A/G19A)-6×-His, FMO1(G215A)-6×-His proteins were visualized by chemiluminescence using anti-His (Qiagen), peroxidase-conjugated secondary antibodies (Bio-Rad), and ECL reagent (GE Biosciences).

Electrolyte Leakage and Hypersensitive Reaction Assays.

Electrolyte leakage and hypersensitive reaction (HR) assays were performed according to ref. 46. Thirty- to 32-d-old WT and fmo1 plants were irrigated with 10 mL of water, 1 mM Pip, or 1 mM N-OH-Pip. One day later, six to seven random leaves of each plant were inoculated with a 3 × 10⁸ cells per mL suspension of Pst DC3000 (vector) or Pst DC3000 (avrRpt2) and then incubated at room temperature under lights. For electrolyte leakage assay, at 5 hpi, three leaf discs (7 mm diameter) of each plant were pooled and floated in 20 mL of water in Petri dishes. Five minutes later, the leaf discs were transferred to a 15-mL tube containing 3 mL of water and incubated at room temperature for 1 h with shaking. Conductivity of each sample before and after boiling was measured using an electrical conductivity meter (Spectrum Technologies). The percentage of electrolyte leakage was calculated as conductivity before boiling/conductivity after boiling. For HR assays, leaf phenotypes of WT and fmo1 for each condition were record at 8 hpi. Three plants were used per condition, and the experiment was repeated three times.