Discovery of endogenous nitroxyl as a new redox player in Arabidopsis thaliana

Nitroxyl (HNO) is the one-electron reduced and protonated congener of nitric oxide (•NO), owning a distinct chemical profile. Based on real-time detection, we demonstrate that HNO is endogenously formed in Arabidopsis. Senescence and hypoxia induce shifts in the redox balance, triggering HNO decay or formation mediated by non-enzymatic •NO/HNO interconversion with cellular reductants. The stimuli-dependent HNO generation supports or competes with •NO signalling, depending on the local redox environment.

On the other hand, the HNO relative nitric oxide (•NO) was identified as a physiological mediator of endothelial cell relaxation in mammalian systems in the late 1980s 6 , and has since proven to be a master regulator of numerous physiological and pathophysiological processes in all kingdoms. In land plants, •NO is synthesized endogenously via either reductive or oxidative routes 7 . Its generation fluctuates with various developmental and stress stimuli, tightly balanced by the formation of other reactive nitrogen species (RNS), highlighting •NO complex biology in cells 7,8 . Most of the impact of •NO in plants is attributed to its uncharged state or peroxynitrite. However, pharmacological studies in mammalian systems also emphasize the potential functionality of HNO 1,3 .
Despite intense research on HNO's biological effects, its endogenous formation had not been detected in mammalian cells 9,10 , and in plants, HNO-generating systems and nitroxyl involvement in triggering biological responses were virtually terra incognita. Here, we present the first experimental evidence that HNO is formed endogenously in living cells of the model plant Arabidopsis thaliana (L.). Further, using precise methods to detect HNO and specific modulators of nitroxyl formation or scavenging, we show that the cellular redox environment tightly balances HNO formation.
In real time we used an electrochemical microsensor to measure HNO concentrations up to low nanomolar levels (1 nM-1 µM). Our previously described method is based on a three-electrode system consisting of a platinum counter electrode, an Ag/AgCl reference electrode and a gold working electrode modified with a cobalt porphyrin covalently attached via a thiol moiety 11,12 . Note that this method targets the HNO molecule; no interference or spurious signal arises from the presence of •NO, O 2 , NO 2 − or other RNS [11][12][13] . More specifically, we first verified the effectiveness of the microsensor's HNO electrodetection in three structurally distinct donor solutions: Angeli's salt (AS 14 ), 4-NO 2 -Piloty's acid (NPA 15 ) and Cimlanod (CM, formerly BMS-986231 or CXL-1427 (ref. 16)) in the presence or absence of the HNO scavenger, phosphine tris(4,6-dimethyl -3-sulfonatophenyl)phosphine trisodium salt hydrate (TXPTS; Fig. 1a and Supplementary Table 1). Next, we recorded concentration/time traces of endogenous HNO generation in extracts from 21-day-old, wild-type (WT) Arabidopsis leaves (Fig. 1b). We detected a high current reaching 0.240 ± 0.005 µA and calibrated it against one measured in a known concentration of HNO in solution; both were 150 ± 3 nM (Extended Data Fig. 1). WT 0.5 g ml -1 WT 0.5 g ml -   To confirm that the sensor selectively detects HNO in the plant cellular environment under in vitro conditions, we added donors (10 µM AS or 10 µM CM) to release the molecule at physiological pH ( Fig. 1b). Leaves pretreated with HNO-releasing agents (AS, NPA, CM) showed significant nitroxyl enrichment, but the electrochemical signal was dependent on donor type, concentration, time and temperature (Extended Data Fig. 2). To prove the presence of HNO in plant tissues, we selected a specific concentration of the water-soluble TXPTS as the trapping agent because it does not produce nitroxyl in reacting with other components of the extract media, especially NO and thiols (Extended Data Fig. 3).

Brief Communication
Results confirm that under physiological (non-stress) conditions, the plant endogenously produces low (nanomolar) concentrations of HNO. The fainter amperometric signal reflecting a lower HNO concentration was detected in both WT Arabidopsis cell suspension and leaves of Arabidopsis mutants with a lower •NO content-noa1-2 and nia1nia2 (Fig. 1c). These results demonstrate that endogenous •NO determines HNO content in biological systems, possibly via nitric oxide one-electron reduction to nitroxyl 4 . Plant cells probably have other chemical sources of HNO because in the triple-mutant nia1nia2noa1-2 we detected only trace amounts of •NO, whereas HNO content constituted over 10% of the value recorded in WT plants (Fig. 1c).
To track HNO formation in plant cells, we used a fluorescent triarylphosphine-based probe that relies on the reductive Staudinger ligation of HNO with an aromatic phosphine 17 (Extended Data Fig. 4a). By confirming the fluorophore's ability to detect and estimate the HNO concentration in the plant cellular environment we performed quantification and bio-imaging of HNO in vivo in Arabidopsis leaves and a cell suspension (Fig. 1d,e and Extended Data Fig. 4b,c). Epidermal and leaf vascular bundles showed faint intracellular fluorescence. Adding CM enriched HNO production to enhance the signal. Co-treating the leaves with the scavenger TXPTS reduced, but did not quench, HNO-dependent fluorescence (Fig. 1d,e).
To determine whether the cellular redox status tightly regulates in vivo HNO formation, we shifted the redox balance in WT Arabidopsis leaves towards an oxidative or reductive environment by applying 0.1 mM menadione (MN) or 1 mM N-acetylcysteine (NAC), respectively (Extended Data Fig. 5). Electrochemical quantification revealed that MN pretreatment decreased the HNO level by ~20% and NAC pretreatment showed the opposite effect: the nitroxyl level increased by ~15% (Extended Data Fig. 5a,b).
Based on our observations, we hypothesized that developmental or environmental stimuli alter the cellular redox balance and provoke HNO fluctuations. To test this assumption, we subjected WT Arabidopsis plants to dark-induced leaf senescence (DILS) as a model for the perturbation of redox homeostasis 18 . As expected, senescence promoted an unfavourable oxidative environment for HNO (Extended Data Fig. 6a). Electrodetection in extracts of individually darkened WT Arabidopsis leaves (leaf seven of the rosette) revealed a significant (~50%) decrease in HNO signal from day 1 to day 7 (Fig. 2a). In vivo, a needle-type electrode (Extended Data Fig. 1f) indicated that HNO-dependent current was weakest in the leaf zone where chlorophyll content decreased most sharply (zone c in Fig. 2b and Supplementary Table 2). Notably, reversing the DILS programme by restoring light access on day 3 recovered the HNO pool and increased chlorophyll content ( Fig. 2a and Supplementary Table 2). Nitroxyl-pretreated leaves showed significantly less accumulation of senescence-associated gene transcripts, including SAG12 encoding cysteine protease, and less chlorophyll degradation, indicating that from day 3, senescence was delayed (Extended Data Fig. 6b,c). We conclude that fluctuation in HNO production is a part of the redox pathway that modulates dark-induced senescence.
To further support our hypothesis that cellular redox status is decisive for HNO kinetics, WT Arabidopsis plants experienced hypoxia associated with reductive conditions. Hypoxia encouraged a significant (~25%) increase in HNO formation mainly during the first 24 h (Fig. 3a-c). One day after the stress was removed, HNO decreased sharply (Fig. 3a). These results indicate that a switch in nitroxyl kinetics toward HNO formation is an early reductive stress-related response in plant cells. Because hypoxia can also occur in plant tissues and organs under normoxia 19 , it creates (micro)hot-spots of HNO bioavailability and bioactivity.
Searching for physiologically relevant HNO sources under a reductive environment, we also confirmed that non-enzymatic •NO/HNO interconversion mediated by cellular reductants such as ascorbic acid, salicylic acid or hydrogen sulfide (H 2 S) 4 might constitute an important in vivo route to HNO formation in plant cells. Enrichment of leaves with pseudo-aromatic alcohols and H 2 S resulted in a time-dependent rise in the HNO level (Extended Data Fig. 7 and Supplementary Table 3). All the three compounds are essential players in plant cells involved, for example, in signal transduction at physiological concentrations from µM (H 2 S, salicylic acid) to mM (ascorbic acid) [20][21][22] and may provide ubiquitous HNO bioavailability.
Finally, to discover the physiological significance of the redox-dependent HNO fluctuations in WT Arabidopsis leaves, profiling of the HNO-dependent transcriptome (TXPTS-treated) was  Table 2). Data are presented as the mean ± s.d. of three biologically independent replicates (n = 3). *Values differ significantly (P ≤ 0.05) from day 0 of DILS, or day 3 in the case of recovery experiment. Statistical significance was assessed using two-tailed t-tests.

Brief Communication
https://doi.org/10.1038/s41477-022-01301-z performed. The functional analysis of gene set enrichment revealed that the most significantly enriched Gene Ontology (GO) categories were ethylene receptor activity (molecular function, GO:0038199) and negative regulation of the ethylene-activated signalling pathway (biological process, GO:0010105). Of all the above genes, two met the strict criteria for the significance of differential expression with the adjusted P-value <0.05: EIN3-binding F-box protein 2 (EBF2 -AT5G25350) and ethylene response sensor 2 (ERS2 -AT1G04310; Fig. 3d and Supplementary Table 4). The observed implication of HNO in regulation of the ethylene response could be critical for responses Statistical significance was assessed using two-tailed t-tests.

Brief Communication
https://doi.org/10.1038/s41477-022-01301-z to environmental changes with elevated HNO formation. ERS2 belongs to a group of membrane-located receptor proteins, whereas EBF2 governs ethylene signalling diminishing and/or resetting ethylene responses in cells with initiated transduction of the hormone signal 23 .
In confirmation, hypoxia-induced HNO generation coincides with EBF2 and ERS2 upregulation mainly in the first 48 h of the stress. No change or downregulation of EBF2 and ERS2 expression coinciding with significantly reduced cell viability was observed when TXPTS trapped HNO during hypoxia stress (Fig. 3e,f and Extended Data Fig. 8). The results shed light on potential HNO implication in a well proved NO/ethylene signalling pathway for example Hartman et al. 24 These studies are the first to confirm endogenous HNO production in living cells and point toward a novel regulatory role of the molecule in the ethylene signalling pathway in plants. Added to our previous findings on HNO chemistry 4,11,13,15 , our revelation of HNO's effects on plant •NO signalling and metabolism mandate the inclusion of nitroxyl in the group of gasotransmitters now composed of •NO, CO and H 2 S. The ubiquitous bioavailability of the HNO molecule provided by non-enzymatic •NO/HNO interconversion allows it to support or compete with •NO signalling, depending on the local redox environment. Nitroxyl's contribution to the reactive species interactome engaged in cell signalling requires re-evaluation of the consensus on •NO biology. Our results provide the impetus and a scaffold to explore the biological functions of HNO in living cells.

Plant material and growth conditions
The Columbia (Col-0) ecotype of Arabidopsis thaliana was used as the WT. T-DNA insertion lines, nia1nia2 (SALK_ N2356) and noa1-2 (SAIL_507_E11) were obtained from the SIGnAL collection of the Nottingham Arabidopsis Stock Centre 25 . The triple nia1nia2noa1-2 mutant (impaired in nitrate reductase and nitric oxide-associated 1 (NOA1)-mediated NO biosynthetic pathways) was generated by crossing, and the putative triple homozygous mutant plants were confirmed by polymerase chain reaction (PCR) and sequence analyses, following Lozano-Juste and León 26 . The PCR primers (Genomed) used for genotyping SALK lines and triple mutants are listed in Supplementary  Table 5.
After 3 d of cold stratification, all seeds were planted in a growth chamber under a 16:8 h light/dark cycle at a photon fluency rate of 110 µmol m −2 s −1 at 22 °C and 60% relative humidity. nia1nia2noa1-2 was cultivated with Murashige and Skoog medium supplemented with nitrites to promote growth. The experiments were performed on leaves from 21-day-old plants.
Arabidopsis thaliana (Col-0) cell suspension cultures from leaf-derived callus were grown following Encina et al. 27 in a modified Murashige and Skoog 28 medium (including vitamins, Duchefa Biochemie) enriched with 30 g l −1 sucrose. They were subcultured every 21 d when 2 ml were transferred into 50 ml of fresh medium in 500 ml flasks. The cultures were grown on a shaker at 120 rpm under a 16:8 h light/dark cycle at a photon fluency rate of 110 µmol m −2 s −1 at 22 °C. Experiments were carried out on d21 of the cultivation cycle.

DILS
To initiate DILS, rosette leaf seven from a 21-day-old WT Arabidopsis plant was darkened by gently covering it with aluminium foil; the rest of the rosette remained under the normal light/dark cycle. Senescence progression was monitored on d1, d3, d5 and d7. Control (non-darkened) samples were also harvested from leaf seven.

Hypoxia stress and recovery procedure
In hypoxia treatments, 21-day-old WT plants were subjected to a low-O 2 air mix in an airtight chamber under the same light/dark cycle. Air-flux conditions were 3% O 2 , 0.03% CO 2 and 97% N 2 gas. Control plants were maintained at a normal oxygen level (normoxia). Analyses were performed 1, 3, 6, 12, 24, 48 and 72 h after treatment. One group of plants was removed from the chamber at 24 h and recovered under normal growth conditions for 24 h.

HNO-modulator treatments
To test the efficiency of our modulators (Extended Data Figs. 2 and 3) at enriching or quenching HNO in plant tissues, rosette Arabidopsis leaves were sprayed with different concentrations of HNO donors, such as AS (0.25, 0.5, 1, 1.5 and 10 mM), NPA (0.5 mM) and CM (0.5 and 1.5 mM) (MedChemExpress), or the HNO scavenger TXPTS (0, 1, 5 and 10 mM) and incubated under various time and temperature conditions (Extended Data Figs. 2 and 3) in an airtight chamber. For electrochemical measurements of HNO concentration, a crude extract was prepared from leaves five to seven as described in the section 'Crude leaf extract preparation for electrochemical measurements'.
To study the effect of CM on physiological and molecular parameters during DILS, the seventh leaf of the rosette of the 21-day-old plants was sprayed once with 1.5 mM CM (MedChemExpress), incubated for 1 h in an airtight chamber, then subjected to DILS as described above. Control plants, also collected from leaf seven, were treated with distilled water and incubated for 1 h in an airtight chamber. To study the effect of TXPTS on transcriptomic changes and physiological parameters, leaves of the rosette of 21-day-old plants were sprayed once with 5 mM TXPTS and incubated for 2 h in an airtight chamber under normal growth conditions; control plants were treated with distilled water, and incubated as above. All collected samples were immediately used for experiments (chlorophyll measurement) or frozen in liquid nitrogen and stored at −80 °C until further use (RNA extraction). Each sample consisted of three leaves (leaf seven) pooled from different plants.

HNO quantification by electrochemical method
A three-electrode system consisting of a platinum counter electrode, Ag/AgCl quasi-reference electrode and a gold working electrode modified with a cobalt porphyrin covalently attached via a thiol moiety was used to detect HNO, following a previously described method 11,12,29,30 . Briefly, in the presence of HNO, a CoIII(P)NO-adduct forms and oxidizes. The resulting CoIII(P)NO is unstable, thus completing the catalytic cycle. The current intensity is proportional to the amount of HNO that binds to Co(P) 12 (Extended Data Fig. 1a). TEQ_HNO software v.2.0 was used for HNO data collection.
For in vivo measurement, electrochemical etching was used to form the gold working electrode into a needle (Extended Data Fig. 1f). The method has demonstrated specificity for HNO, with no interference or spurious signal arising from •NO, O 2 , NO 2 − and other RNS 11,13,31,32 . The calibration curve measures current responses at a potential of 0.8 V with the addition of freshly prepared AS, the HNO donor (Merck) 3,14,33 and aqueous solutions of 30-900 nM nitroxyl (Extended Data Fig. 1c,d).
Electrochemical monitoring of HNO generation in vivo used a setup (microelectrode) previously described by Floryszak-Wieczorek et al. 34,35 for •NO electrodetection in plant leaves. Briefly, a leaf blade was placed on an agar layer in which a Pt wire was introduced as a counter electrode. An AgCl-coated Ag needle was then introduced into the leaf tissue close to the area of HNO monitoring to serve as a quasi-reference electrode, and finally the HNO-selective microelectrode was introduced into the leaf.

HNO quantification by a triarylphosphine-based probe
Confocal laser scanning microscopy. Cross-sections taken from the middle of the fully developed seventh leaf of the rosette were incubated in 100 µl of a 16 µM phosphine-based fluorescent probe (synthesized in S. B. King's laboratory as described previously 17 ) in 10 mM Tris-HCl, pH 7.4, for 15 min in the dark at 25 °C. As verified previously, the probe reacts with HNO under physiological conditions without interference by other biological redox species 36 . After incubation, the buffer was removed, and cross-sections were washed three times with 10 mM Brief Communication https://doi.org/10.1038/s41477-022-01301-z Tris-HCl, pH 7.4. Sections were placed on glass slides and observed under a Zeiss Axiovert 200M inverted microscope equipped with a confocal laser scanner (Zeiss LSM 510, Carl Zeiss AG). Sections were excited at 488 nm using an argon laser. Dye emissions were recorded using a 505-530 nm bandpass filter, and chloroplast autofluorescence was captured with the 585 nm long-pass filter. Microscope, laser and photomultiplier settings were held constant to obtain comparable data. Images were processed and analysed using Zeiss LSM 510 software (v.3.2 SP2).
A portion of Arabidopsis cell culture (250 µl) was incubated with a 16 µM phosphine-based fluorescent probe in 10 mM Tris-HCl, pH 7.4 for 15 min in the dark at 25 °C, then centrifuged at 2,000g, washed twice in fresh Tris-HCl buffer, and immediately imaged by confocal laser scanning microscopy as described for leaf cross-sections.

Fluorescence quantification.
To measure the amount of HNO produced by Arabidopsis, 250 µl of cell culture sample or ten discs of 0.5 cm diameter cut from the middle of the fully developed seventh leaf were incubated in buffer containing 16 µM of phosphine-based fluorescent probe in 10 mM Tris-HCl, pH 7.4, for 1 h in darkness at 25 °C. Next, the discs were washed twice with 10 mM Tris-HCl, pH 7.4, finely homogenized in 1 ml Tris-HCl buffer and centrifuged at 900g at room temperature. Fluorescence was measured at 465 nm excitation and 520 nm emission wavelength (Fluorescence Spectrophotometer F-2500, Hitachi). Samples were normalized to the recorded autofluorescence of the plant material incubated without the fluorescent dye.

•NO quantification by electrochemical method
•NO generation in leaf tissues was monitored by constant potential amperometry with a NO-selective disc-type electrode 34,35,[37][38][39][40] . The electrode was prepared by electropolymerizing a poly-eugenol thin film on a cleaned Pt disc 37 and repeatedly scanning the potential between −0.2 and 0.6 V in a 10 mM solution of eugenol (Merck) in 0.1 M NaOH. The modified electrode was then conditioned at a constant potential of 0.9 V in a phosphate buffer (pH 7.4) until a stable background current was reached. Electrochemical monitoring of NO generation in leaf tissue extracts and cell suspensions was performed as described previously 37 . The current was recalculated into concentration units based on a calibration curve (ISO-NO Mark II instruction manual, World Precision Instruments) constructed by measuring current responses to the addition of freshly prepared •NO aqueous solutions generated in situ from the reaction of iodide with nitrite in acid solution within the range of 0.3-100 µM (Extended Data Fig. 1g,h; ISO-NO Mark II instruction manual, World Precision Instruments) 41 .

Crude leaf extract preparation for electrochemical measurements
Electrochemical monitoring of HNO and •NO generation in Arabidopsis leaf tissues was performed following Floryszak-Wieczorek et al. 34 for •NO electrodetection in plant leaves. Briefly, leaves five to seven were pooled from different plants to obtain 0.5 g of fresh weight and homogenized under limited access to oxygen (in a hypoxia chamber; Hypoxy-Lab) in 0.5 ml of 0.05 M phosphate buffer, pH 7.4, at 4 °C. The extract was centrifuged at 900g for 15 s at 4 °C and analysed immediately. All data were obtained following the same protocol, and the results were normalized to the same time.

Quantification of chlorophyll contents
Chlorophyll was extracted from leaf seven (100 mg fresh material), following Hiscox and Israelstam 42 , incubated with 5 ml of dimethylsulfoxide (Merck) at 65 °C for 2 h. Chlorophyll a content was measured by spectrophotometer (Shimadzu UV-Vis-160) with emission at 665 nm and chlorophyll b content was measured at 649 nm.

Pharmacological modulation of the cellular redox environment
To shift the cellular redox balance toward reduction, leaves of 21-day-old WT Arabidopsis plants were sprayed with 1 mM NAC (Merck), which depletes oxidized electron acceptors, such as glutathione and thioredoxin 43 , to induce reductive stress. To shift the balance toward oxidation, leaves of 21-day-old WT Arabidopsis plants were sprayed with 100 µM MN (Merck), a redox-active quinone that generates intracellular superoxide 44 . To identify HNO sources under reductive environment leaves of 21-day-old WT Arabidopsis plants were sprayed with 1 mM ascorbic acid (Merck), 1 mM salicylic acid (Merck) and 1 mM sodium hydrosulfide (as a H 2 S donor, Merck). In all treatments, control plants were sprayed with distilled water. Analyses were performed 0, 1, 3, 6 and 24 h after treatment (in the case of H 2 S, HNO formation was also monitored over a period of 1 h).

ROS measurement. The O 2
•− level was assayed spectrophotometrically based on the capacity of the superoxide anion-radical to reduce nitro blue tetrazolium (Merck) to diformazan 45 . H 2 O 2 concentration was assayed spectrophotometrically using the titanium (Ti 4+ ) method 46 .
Glutathione. Reduced glutathione (GSH) and oxidized glutathione (GSSH) contents were determined as described by Griffith 48 . GSH was oxidized by 5,5′-dithiobis-(2-nitrobenzoic acid) (Merck) to form GSSH and 5-thio-2-nitrobenzene. GSSH was then reduced to GSH by glutathione reductase and NADPH. Leaf tissues (200 mg), ground with a mortar and pestle in liquid nitrogen, were centrifuged at 15,000g for 15 min at 4 °C with 2.5 ml of 2.5% trichloroacetic acid. A supernatant (0.3 ml) was then used to assay total glutathione (GSH + GSSH). A further 0.3 ml of supernatant was pretreated with 6 µl of 2-vinylpyridine (Merck) for 60 min at 20 °C to mask GSH by derivatization. Both types of sample (0.1 ml each) were mixed with 0.7 ml of 0.3 mM NADPH, 0.1 ml of 6 mM 5,5′-dithiobis-(2-nitrobenzoic acid) and 0.1 ml of glutathione reductase (50 units ml −1 ). Absorbance at 412 nm was recorded after 5 min at room temperature. The total glutathione (GSH + GSSH) and GSSH contents were calculated using a standard curve and expressed as µmol per g (fresh weight). GSH content was calculated from the difference between total glutathione and GSSH.

Cell viability determination
Cell viability, defined by plasma membrane integrity, was measured spectrophotometrically as Evans blue uptake 49 .

Gene expression analysis
Arabidopsis leaves were frozen in liquid nitrogen and stored at −80 °C until use. RNA was isolated from 100 mg of a frozen sample using Tri-Reagent (Merck) and purified using a Deoxyribonuclease Kit (Merck). We processed 1 µg of RNA for reverse transcription using a Reverse Transcription Kit (Thermo Fisher Scientific) following the manufacturer's instructions.  Table 5 lists all the primers (Genomed) used. Relative gene expression was calculated using the Pfaffl mathematical model 51 .

Read mapping and identification of differentially expressed genes
TXPTS-treated versus control (untreated) samples were analysed. The complementary DNA library for RNA sequencing was prepared using a standard TruSeq Stranded messenger RNA kit from Illumina. Sequencing was performed on an Illumina NovaSeq machine with paired-end setting and a 150-nucleotide read length. The quality of raw sequencing reads was analysed using FastQC (v.0.11.9) software (https://www.bioinformatics.babraham.ac.uk/projects/fastqc/). Next, reads were subjected to mapping to the reference genome of Arabidopsis thaliana, obtained from the Ensembl Plants database 52 , using RNA STAR (v.2.7.10a) software 53 . The gene expression quantification was obtained from the STAR aligner using ARAPORT11 gene annotation 54 and subjected to differential expression analysis using the R (v.4.2.0) environment with the limma (v.3.52.0) 55 and EdgeR (v.3.38.0) 56 packages. Statistically significant differentially expressed genes were determined using an adjusted P-value cut-off of <0.05. Functional analysis of gene set enrichment was performed using g:Profiler web service (v. e105_eg52_p16_5d1f001) 57 .

Statistical analysis
All included experiments were replicated three times on independently grown plants. In addition, each sample was tested in three technical repetitions, and results from representative data sets are presented. Statistical differences were calculated using two-tailed t-tests (P ≤ 0.05). Related information is listed in the source data.

Reporting summary
Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Data availability
All data generated or analysed during this study are included in the manuscript or as a Supplementary Information (Extended Data Figs. 1-9 and Supplementary Tables 1-5). For RNA-seq data analysis, the Arabidopsis thaliana TAIR10 reference genome assembly has been used (GenBank ACC: GCA_000001735.1). RNA-seq data have been deposited in the European Nucleotide Archive (ENA) under accession number LPRJEB53633. Source data are provided with this paper.  Fig. 1 | Nitroxyl (HNO) and nitric oxide (NO) biosensors calibration. a, Mechanism of specific electrochemical detection of HNO by cobalt porphyrin (CoP) covalently bonded to a gold surface. The developed sensor is an electrochemical HNO sensing device based on the covalent bonding of a cobalt porphyrin to a gold surface. A surface effect modulates the redox potentials allowing discrimination between HNO and NO. The electrode potential is set at a value of 0.8 V, where the porphyrin is stable as Co(III)P, observing a basal current. The reaction with HNO produces the Co(III)PNO¯ complex, which under the conditions described above is oxidized to Co(III)PNO. Due to the lability of the resulting Co(III)PNO complex, the system rapidly returns to Co(III)P, allowing the catalytic cycle to begin again. Since each electrode is only covered with a surface concentration, the total amount of trapped HNO is very small. Also, it is important to note that since a small amount of current can be detected, the 'real' amount of azanone reacting with the electrode is generally negligible compared to the total amount of HNO in the system (less than 1 %), and does not significantly perturb the production and consumption of HNO, making it a powerful tool in mechanism and kinetics studies where azanone plays a key role. 11,12 b-e, HNO biosensor calibration; b, Current response after adding different Angeli's salt (AS) concentrations to a stabilized sensor's baseline. One microliter of an AS stock solution was added to a stabilized sensor's baseline. A few seconds after, the signal starts to grow, and after 15 min, it begins to decrease.

Brief Communication
No signal was detected after adding SNAP, a nitric oxide donor. c, Current response after pretreatment of 10 min of different AS concentrations. One microliter of an AS stock solution was added to 1 ml of buffer and then was waited for 10 min. Next, the measurement with the sensor was started. In this case, as expected, the signal only decreased. d, Δcurrent vs. AS concentration.
e, Δcurrent vs. HNO concentration. The HNO concentration in the Co(P) electrode reaction solution was estimated by the following kinetic analysis 11,12 : (1) represents the donor decomposition characterized by a first-order rate constant k d , and (2) represents bimolecular HNO dimerization, characterized by rate constant k h . Reaction (1) was considered an irreversible reaction because its inverse rate is negligible concerning reactions (2) at the initial rate condition. Reaction (2) is irreversible. Calculating [HNO], all the HNO produced through reaction (1) must be consumed in reaction (2) https://doi.org/10.1038/s41477-022-01301-z Extended Data Fig. 2 | The treatment effect of Arabidopsis WT leaves with nitroxyl donors on HNO concentration. a, The impact of a one-time treatment of Arabidopsis WT leaves with Angeli's salt (AS) on HNO concentration. Leaves were sprayed with the following AS concentrations 0.25, 0.5, 1.0, and 10 mM, respectively, at 25 °C. HNO measurements with microsensor were carried out in the leaf extracts 10 min after spraying. HNO formation was comparable in AS concentration range of 0.5-10 mM indicating a leaf penetration limit for AS diffusion. This could be explained by the fact that plant epidermis is covered by an extracellular hydrophobic layer (cuticles), and AS is the sodium trioxodinitrate salt (Na 2 [N2O 3 ]). This difference between nonpolar and polar compounds, especially during multiple treatments, was optimized using an organic surfactant (see c). The treatment with the lowest AS concentration (0.25 mM) decreased (20%) the HNO level. b, The effect of a three-time treatment of Arabidopsis WT leaves with AS on HNO concentration. Leaves were threetime sprayed in 10 min intervals with the following AS concentrations 0.25, 0.5, 1.0 mM, respectively, at 25 °C. HNO measurements with microsensor were carried out in the leaf extracts 10 min after the third spraying The maximum HNO concentration in leaf tissues three-time treated with 0.5 mM AS increased by 40% compared with one-time pretreatment (for comparison see A). HNO concentrations measured in a group of leaves no. 5-7 or only leaf no. 7 (pooled from several plants) treated with AS (0.5 mM) were comparable. c, The effect of time, temperature, and multiple treatments of Arabidopsis WT leaves with AS on HNO concentration. Leaves were sprayed once, three, or five-times in 1 or 10 min intervals, respectively, with 0.5 mM AS at 25 °C or 37 °C. HNO measurements with microsensor were carried out in the leaf extracts after 1 or 10 min after the last spraying, respectively. Angeli's salt decomposition rate is temperaturedependent, so HNO production from 0.5 mM AS was almost 2-fold lower at 25   Enrichment of leaves with pseudo-aromatic alcohols such as AA and SA (a, b) resulted in a time-dependent rise in the HNO level measured in vivo by electrodetection in leaf extracts. However, ascorbate as a diol may react in vitro approximately twenty times faster than SA being a phenol 63,64 . The proposed mechanism involves a nucleophilic attack to •NO by the alcohols, coupled with proton transfer and subsequent decomposition of the thus-produced radical to yield HNO (Supplementary Table 3). Leaf pretreatment with H 2 S (c) also resulted in a significant increase in HNO production. The signal accelerated during the first minutes reflecting the fast nature of the reaction between H 2 S and •NO to produce HNO (k = 10 4 M −1 s −1 ) 65 . However, there is a more complicated mechanism than a simple bimolecular reaction between •NO and H 2 S/HS − (Supplementary For manuscripts utilizing custom algorithms or software that are central to the research but not yet described in published literature, software must be made available to editors and reviewers. We strongly encourage code deposition in a community repository (e.g. GitHub). See the Nature Portfolio guidelines for submitting code & software for further information.

March 2021
Data Policy information about availability of data All manuscripts must include a data availability statement. This statement should provide the following information, where applicable: -Accession codes, unique identifiers, or web links for publicly available datasets -A description of any restrictions on data availability -For clinical datasets or third party data, please ensure that the statement adheres to our policy The authore declare that all data supporting the findings of this study are available within the article and its Supplementary Information Files. For RNA-seq data analysis, the Arabidopsisn thaliana TAIR10 reference genome assembly has been used (GenBank ACC: GCA_000001735.1). RNA-seq data that support the findings of this study has been deposited in the European Nucleotide Archive (ENA) under accession code LPRJEB53633. Materials generated in this study are available from the corresponding author upon request.

Human research participants
Policy information about studies involving human research participants and Sex and Gender in Research.
Reporting on sex and gender Note that full information on the approval of the study protocol must also be provided in the manuscript.

Field-specific reporting
Please select the one below that is the best fit for your research. If you are not sure, read the appropriate sections before making your selection.

Life sciences Behavioural & social sciences Ecological, evolutionary & environmental sciences
For a reference copy of the document with all sections, see nature.com/documents/nr-reporting-summary-flat.pdf

Life sciences study design
All studies must disclose on these points even when the disclosure is negative.

Sample size
Sample size per biological replicate (per treatment) was at least 25 plants which made it possible to obtain fresh weight of leaves necessary for molecular and biochemical analyzes. All included experiments were replicated three times (n=3). Sample size used in experiments was sufficient to generate statistical significance.
Data exclusions No data was excluded from the analysis.

Replication
All included experiments were replicated three times on independently grown (and treated) plants. Additionally, each sample was tested in three technical repetitions. The number of replication is indicated in the figure legends. Similar results were obtained between independent experiments.
Randomization Plants were always randomly distributed during growth and treatment.
For the preparation of a leaf sample, leaves were pooled from randomly selected different plants of the same genotype and treatment to obtain fresh weight appropriate to the experiment. For confocal observation, the leaf cross-sections were randomly selected from ~ 20 slices pooled from leaves of randomly selected different plants of the same genotype and treatment.

Blinding
Blinding was not used in this study as our study did not involve animals and/or human research participants. All the experiments were performed without prior knowledge of the outcome.
Reporting for specific materials, systems and methods We require information from authors about some types of materials, experimental systems and methods used in many studies. Here, indicate whether each material, system or method listed is relevant to your study. If you are not sure if a list item applies to your research, read the appropriate section before selecting a response.

Authentication
Describe the authentication procedures for each cell line used OR declare that none of the cell lines used were authenticated.

Mycoplasma contamination
Confirm that all cell lines tested negative for mycoplasma contamination OR describe the results of the testing for mycoplasma contamination OR declare that the cell lines were not tested for mycoplasma contamination.

Commonly misidentified lines (See ICLAC register)
Name any commonly misidentified cell lines used in the study and provide a rationale for their use.